Redox-regulated fate of neural stem progenitor cells

Redox-regulated fate of neural stem progenitor cells

Biochimica et Biophysica Acta 1850 (2015) 1543–1554 Contents lists available at ScienceDirect Biochimica et Biophysica Acta journal homepage: www.el...

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Biochimica et Biophysica Acta 1850 (2015) 1543–1554

Contents lists available at ScienceDirect

Biochimica et Biophysica Acta journal homepage:


Redox-regulated fate of neural stem progenitor cells☆ Tim Prozorovski ⁎, Reiner Schneider, Carsten Berndt, Hans-Peter Hartung, Orhan Aktas Department of Neurology, Medical Faculty, Heinrich-Heine-University, Düsseldorf, Germany

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Article history: Received 5 January 2015 Accepted 29 January 2015 Available online 7 February 2015 Keywords: Neural stem progenitor cells Self-renewal Redox NAD NADP Differentiation

a b s t r a c t Background: Accumulated data indicate that self-renewal, multipotency, and differentiation of neural stem cells are under an intrinsic control mediated by alterations in the redox homeostasis. These dynamic redox changes not only reflect and support the ongoing metabolic and energetic processes, but also serve to coordinate redox-signaling cascades. Controlling particular redox couples seems to have a relevant impact on cell fate decision during development, adult neurogenesis and regeneration. Scope of review: Our own research provided initial evidence for the importance of NAD+-dependent enzymes in neural stem cell fate decision. In this review, we summarize recent knowledge on the active role of reactive oxygen species, redox couples and redox-signaling mechanisms on plasticity and function of neural stem and progenitor cells focusing on NAD(P)+/NAD(P)H-mediated processes. Major conclusions: The compartmentalized subcellular sources and availability of oxidizing/reducing molecules in particular microenvironment define the specificity of redox regulation in modulating the delicate balance between stemness and differentiation of neural progenitors. The generalization of “reactive oxygen species” as well as the ambiguity of their origin might explain the diametrically-opposed findings in the field of redoxdependent cell fate reflected by the literature. General significance: Increasing knowledge of temporary and spatially defined redox regulation is of high relevance for the development of novel approaches in the field of cell-based regeneration of nervous tissue in various pathological states. This article is part of a special issue entitled Redox regulation of differentiation and de-differentiation. © 2015 Elsevier B.V. All rights reserved.

Abbreviations: ARE, antioxidant response element; bHLH, basic helix-loop-helix; BMI1, B lymphoma Mo-MLV insertion region 1 homolog; CtBPs, C-terminal binding proteins; DLL4, delta-like protein 4; DVL, disheveled; DOUX, dual oxidases; ES, embryonic stem cells; EGFR, epidermal growth factor receptor; ERK1/2, extracellular-signal-regulated kinase 1/2; FGF2, fibroblast growth factor 2; FOXO, forkhead Box O; GFAP, glial fibrillary acidic protein; GCLC, glutamate cysteine ligase catalytic; GSH, glutathione; GSSG, glutathione disulfide; GPX1, glutathioneperoxidase 1;GST, glutathione-S-transferase;GRX,glutaredoxin; GDPD, glycerophosphodiester phosphodiesterase domain; GSK-3β, glycogen synthase kinase-3 beta; HES1, hairy and enhancer of split homolog-1; HO-1, heme oxygenase-1; HGF, hepatocyte growth factor; HIF-1α, hypoxia inducible factor-1 alpha; FIP200, interacting Protein of 200 kDa; MAPK, mitogen-activated protein kinases; MASH1, mouse achaete-scute complex homolog 1; QR, NAD(P)H:quinone oxidoreductase; PARP-1, poly(ADP-ribose) polymerase-1; NOX, NADPH oxidase; NSPCs, neural stem and progenitor cells; NSCs, neural stem cells; NEUROD1, neurogenic differentiation 1; NRF2, nuclear factor erythroid 2-related factor 2; NRX, nucleoredoxin; OCT4, octamer-binding transcription factor 4; PAX, paired box gene; PRX, peroxiredoxin; PI3K, Phosphoinositide-3-kinase; PDGFR, platelet-derived growth factor receptor; PcG, polycomb group; PRDM16, PR domain containing 16; RNS, reactive nitrogen species; ROS, reactive oxygen species; RMS, rostral migratory stream; SIRT, sirtuin; SOX2, sex determining region Y-box 2; SGZ, subgranular zone; SVZ, subventricular zone; SOD, superoxide dismutase; TBX3, T-box transcription factor; TRX, thioredoxin; TGFβ1, transforming growth factor beta 1; VCAM1, vascular cell adhesion molecule 1 ☆ This article is part of a special issue entitled Redox regulation of differentiation and dedifferentiation. ⁎ Corresponding author at: Department of Neurology, Medical Faculty, Heinrich-HeineUniversity, Merowingerplatz 1a, 40225 Düsseldorf, Germany. Tel.: +49 211 302039220; fax: +49 211 302039227. E-mail address: [email protected] (T. Prozorovski). 0304-4165/© 2015 Elsevier B.V. All rights reserved.

1. Neural stem cells Two specialized niches in the adult mammalian forebrain home stem cells that are primarily involved in generating neurons throughout life: the subgranular zone (SGZ) of the dentate gyrus in the hippocampus and the ventricular–subventricular zone (SVZ) in the lateral wall of the lateral ventricles [1–4] (Fig. 1). Although there are several structural differences, e.g. the unique “pinwheel” organization of SVZ stem niche [5], these two areas share a common architecture that allows close contact between neural stem cells (NSCs) with the extensive blood vessel network as well as the expansion of NSC processes into adjacent cell layers [6]. Both niches play a supportive role by assisting the essential functions of NSCs: multipotency and the ability to self-renew in response to mitotic stimuli. Furthermore, combination of various niche factors defines the neurogenic environment for differentiating progeny in order to support neuronal cell fate. Notably, NSCs transplanted into other areas of the brain predominantly differentiate into glial cells indicating the high degree of neural plasticity and the pro-neuronal role of stem niches [7]. 1.1. SVZ niche The SVZ is the largest neurogenic niche in the adult mammalian brain that maintains NSCs and progenitor cells, lying close to ependymal


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Subventrical zone LV

Subgranular zone GCL




Type B EGFRlow SOX2+ GFAP+

Type A EGFRhigh ASCL+ DLX+

Type C

Type 1

Type 2

Neuronal Progenitor





Fig. 1. Schematic representation of regional organization of NSC niches in the adult brain. The specific markers of NSCs and their progeny are shown for the ventricular–subventricular zone (SVZ) of the lateral ventricles (LV) and the subgranular zone (SGZ) of the dentate gyrus in the hippocampus. Immature neurons born in the SVZ migrate along the rostral migratory stream (RMS) into the olfactory bulb. Immature neurons derived from the SGZ of the hippocampus migrate a short distance into the granular cell layer (GCL).

cells of the ventricle. NSCs (Type B cells) are long-lived [8], slowly proliferating [9] and resistant to anti-mitotic agents [10]. These cells express glial fibrillary acidic protein (GFAP) and are characterized by the lack of a surface receptor for epidermal growth factor (EGF) [9]. When isolated from the SVZ and cultured in vitro, cells with characteristics of Type B cells are not able to form colonies under commonly used culture conditions and remain in a relatively quiescent state [11,12]. Upon activation, Type B cells up-regulate EGFR and generate rapidly dividing, short-lived, multipotent Type C cells [12,13] that retain high mitotic activity in vitro [11]. This finding is high relevant as the majority of in vivo and in vitro experiments directed to analyze the proliferation capacity do not consider the role of NSCs (Type B), but rather reflect the mitotic property of their activated progeny. For this reason, we will use the definition of neural stem and progenitor cells (NSPCs) to describe the population of uncharacterized cells possessing both selfrenewal and multipotent capacities. The Type C cells give rise primarily to immature neurons (Type A cells or neuroblasts) that migrate anteriorly over a long distance along the rostral migratory stream (RMS) to the olfactory bulb and generate interneurons [14]. These cells express immature neuronal markers, e.g. doublecortin (DCX) and polysialylated neural cell adhesion molecule (PSA-NCAM). In addition to neuronal fate, NSCs of the SVZ generate astrocytes in the olfactory bulb and oligodendrocytes in the cortex and the corpus callosum [15,16]. 1.2. SGZ niche Two putative types of NSCs are located in the SGZ and generate neurons in the dentate gyrus [17]. Type 1 cells divide slowly, have a radial structure and expand their processes into the adjacent granular cell layer. These cells express the stem cell marker sex determining region Y-box 2 (SOX2) and the glial protein GFAP. Type 2 progenitors are GFAP-negative, have short processes and proliferate more extensively as compared to Type 1. Both populations generate DCX-positive

immature neurons which migrate and integrate into the adjacent granular cell layer [18,19]. Stem cells of the SGZ are also able to generate astrocytes [18]. 2. Redox state of NSPCs The redox state within cells is defined by dynamic changes in the ratio of the interconvertible oxidized and reduced form of specific redox couples such as the nicotinamide adenine dinucleotide (NAD+/ NADH), NAD phosphate (NADP+/NADPH), glutathione (GSSG/2GSH), thioredoxin (TrxSS/Trx(SH)2), superoxide (O− 2 •)/oxygen, hydrogen peroxide (H2O2)/water, and many other redox pairs. Accumulation of oxidizing molecules such as reactive oxygen (ROS) and nitrogen species (RNS), due to their increased production or inefficient clearance, shifts the intracellular redox environment to a more oxidized state and promotes oxidation reactions. The subcellular origin of endogenous ROS has become more increasingly considered as the response of NSPCs to different oxidants and their amount can vary [20–26]. It became obvious that dynamic in the activity of endogenous sources of ROS are essential for both sustained self-renewal capacity [27,28] and differentiation of NSPCs [29]. Further data indicated that modulation of ROS may impact cell fate commitment towards glial or neuronal lineages [27,30]. This review aims to summarize and evaluate the existing data on the specificity in ROS signaling in the context of NSPC biology focusing on i) tempo-spatial generation of ROS, ii) specific redox-regulated signaling pathways, and iii) involvement of NAD(P)+/ NAD(P)H redox couples. 2.1. Hypoxic microenvironment of stem cell niches The oxygen tension in the adult brain among different mammalian species is estimated in the range of 0.1–5.3% [31,32] and low levels of O2 of around 1.3% have been observed in the regions with high cellular

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density such as the hippocampal dentate gyrus [33]. Based on in vitro culture conditions in which atmospheric oxygen levels are traditionally maintained at 20%, it has been accepted to consider O2 levels below 5% as hypoxic. The propagation of embryonic stem (ES) cells and NSPC cultures under lowered oxygen concentrations confirmed a crucial homeostatic role of “physiological hypoxia” for survival and selfrenewal of stem cells [34–38]. The hypoxic microenvironment of neurogenic niches may exert effects on NSC function in part by differential regulation of endogenous ROS levels, particularly in sub-cellular compartments. Thus, hypoxia promotes stemness by supporting the generation of ROS via upregulation of membrane-bond NADPH oxidases (NOX, discussed below) [27]. On the other hand, oxygen availability may also be a major factor in regulation of mitochondrial activity and therefore mitochondrial ROS production in stem cells [39–42]. As depicted below, in striking contrast to NOX system, mitochondria-derived ROS serves as a signal to inhibit proliferation of NSPCs. Mitotic activity of ES cells is supported by glycolytic flux and low mitochondrial oxygen consumption [43]. With respect to this, inhibition of mitochondrial respiration enhances the pluripotency of ES cells [44]. Moreover, reprogramming of somatic cells towards pluripotent stem cells is associated with the acquirement of an “anaerobic” metabolic profile and mitochondrial properties reminiscent to an ES cell-like phenotype (in regard to morphology, distribution, low ROS and low ATP production) [45]. Collectively, these data indicate that low oxygen tension in stem cell niches maintains the proliferating undifferentiated state of stem cells by differentially regulating the activity of different sources of ROS and compartmentalization of redox signaling. 2.2. Mitochondria-derived ROS Mitochondrial oxidative phosphorylation (OXPHOs) produces high amounts of ATP and endogenous ROS, which are mainly generated by complex I (NADH:ubiquinone oxidoreductase) and complex III (ubiquinol:cytochrome c oxidoreductase) of the electron transport chain [46,47]. Recent studies, in which the mitochondrial ROS sensitive probe mtcpYFP was claimed to detect superoxide production, allowed characterization of the mitochondrial input in oxidative state of proliferating [48] and differentiating NSPCs [29]. It has been shown, despite the weak basal superoxide signal, cultured proliferating cortical NSPCs exhibit spontaneous burst-like generation of mitochondrial superoxide anions (superoxide flashes) without global changes in intracellular ROS levels [48]. The supposed superoxide oscillation requires the opening of the mitochondrial permeability transition pore (mPTP) and activity of the electron transport chain. The frequency of this oscillation responds to environmental changes in oxygen concentration and Ca2 + fluxes. In parallel, these mitochondrial flashes negatively regulate NSPC proliferation via inhibition of the mitogen-activated protein (MAP) kinases ERK1 and ERK2 (extracellular-signal-regulated kinases 1 and 2) [48]. However, these data are under debate since cpYFP probe may rather reflect the alterations in pH than being an exclusive superoxide sensor [49]. Nevertheless, the mitochondrial flashes are linked to redox changes in the mitochondria [50], while the molecular explanation for this interesting physiological phenomenon remains unclear. According to the in vitro experiments, embryonic brains of superoxide dismutase 2 (SOD2)-deficient mice display increased superoxide levels and are characterized by reduced neurogenesis in the ventricular zone, related to diminished proliferating capacity of NSPCs [48]. Interestingly, deficiency in superoxide metabolism in SOD1 and SOD2 knockout mice has an impact on cell lineage decision favoring the glial fate of SGZ NSPCs [51]. These results are consistent with our own findings showing that depletion of intracellular GSH favors the glial lineage specification of NSPCs in expense of the neuronal fate [30]. The shift from anaerobic glycolysis to OXPHOs is an essential requirement to support the energy demand of differentiating cells.


Therefore, treatment of NSPC cultures with the mitochondrial complex IV inhibitor antimycin predominantly affects the survival of immature differentiating neuronal progenitors with less impact on dividing cells [52]. Additionally to ATP supply, it was shown that metabolic oxidation favors the differentiation of ES cells [53]. Early steps of neuronal differentiation, which can be synchronized by growth factor withdrawal, are associated with progressively increased mitochondrial biogenesis, mitochondrial mass, ATP production and concomitant rise in ROS formation [39,54–56]. The experiments involving the modulation of mitochondrial activity have revealed the key role of “superoxide” flashes, as depicted by application of mt-cpYFP probe, in neuronal differentiation. Pharmacological inhibition of superoxide anions has been shown to favor the maintenance of immature state of progenitors [29]. In line with this, elevated ROS levels are mostly found in newly generated neurons in vivo [57,58]. Obviously, redox changes in the mitochondria are of great importance for differentiation processes, although their molecular mechanisms and therein associated targets still remain largely unknown. The proper control of mitochondrial ROS generation is essential for NSPC fate decision. Several stem niche factors have been implicated in the control of mitochondrial ROS production. B lymphoma Mo-MLV insertion region 1 homolog (BMI1), a member of the polycomb group (PcG) of transcriptional repressors, is a key factor for the maintenance of both hematopoietic stem cells and NSCs [59–61]. Bmi1−/− mice are characterized by reduced proliferation and neurogenesis in the SVZ [59,62,63]. Importantly, treatment with antioxidant N-acetyl-cysteine rescues some developmental defects in Bmi1−/− mice such as the thymus size and the overall number of thymocytes [64]. This can argue that in addition to well-known function in the repression of the cell cycle inhibitor INK4a/ARF locus, BMI1 plays a role in oxidative metabolism. BMI1-deficient cells exhibit elevated mitochondrial ROS levels and impaired mitochondrial function [64]. In neurons, BMI1 represses prooxidant activity of tumor suppressor p53 [65]. Thus, it is likely that BMI1 may support the stemness of NSPCs at least in part through the control of ROS production. Hepatocyte growth factor (HGF) [66] reduces ROS levels in various cell types including NSPCs [67]. Expression of HFG and its tyrosine kinase receptor c-MET has been found in NSPCs of the SVZ [68]. In vitro and in vivo experiments with exogenous HGF demonstrated the positive effect on proliferation and self-renewal of NSPCs [67,68]. Moreover, mice deficient for the transcription factor PR domain containing 16 (PRDM16), an activator of HGF expression, display elevated levels of ROS [67]. The abnormal ROS levels as well as decreased proliferation of Prdm16−/− NSPCs can be reversed by treatment with exogenous HGF or N-acetyl-cysteine [67]. The molecular mechanism by which HGF controls ROS is still elusive. In epithelial Mv1Lu cell line HGF prevents the disruption of the mitochondrial respiratory function and ameliorates antimitotic effect of mitochondrial ROS overproduction induced upon prolonged treatment with transforming growth factor β1 (TGFβ1) [69]. Thus, it will be important to determine whether HGF-mediated control of mitochondrial activity is involved to support the self-renewal state of NSPCs. More recently, it has been shown that mitophagy, the autophagosomal removal of severely damaged mitochondria, is another mechanism to protect NSPCs from aberrant ROS production [70]. Conditional deletion of focal adhesion kinase (FAK)-family interacting protein of 200 kDa (FIP200), a protein essential for induction of autophagy [71], leads to p53-dependent apoptotic response and cell cycle arrest with subsequent depletion of the NSPC pool in both germinative zones, the SVZ and the SGZ of the postnatal brain [70]. Disturbed mitophagy was associated with increased mitochondrial number and aberrant ROS production as a result of mitochondrial damage. Treatment with N-acetyl-cysteine rescued the abnormalities in self-renewal of FIP200-deficient NSPCs, indicating the detrimental role of mitochondrial ROS overproduction. In addition, the decreased neurogenesis observed in Fip200−/− mice is paralleled with enhanced generation of astrocytes. This shift towards astrogenesis was


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blocked by N-acetyl-cysteine, demonstrating that a more oxidative state of NSPCs in the adult brain defines the glial lineage commitment [70]. This findings extend our own observations for the instructive role of redox state in the commitment of embryonic NSPCs [30]. In summary, the long-term maintenance of NSPC functions such as multipotency and self-renewal require a rigid control mechanism for mitochondrial metabolism. 2.3. Non-mitochondrial sources of ROS Apart from the mitochondrial electron transport chain, other sources of ROS formation localized in distinct subcellular compartments were implicated in NSPC biology (e.g. cyclooxygenases [53,72,73], lipoxygenases [53,74,75], nitric oxide synthases [76–79], and cytochrome P450 enzymes [80,81]). Among these, the family of membrane-associated NAD(P)H oxidases (NOX) is the best-characterized for its role in NSPCs. The NOX proteins generate ROS (superoxide as a primary product) from NADPH to molecular oxygen through electron transport across membranes [82–84]. These evolutionary conserved oxidases can be considered as “professional” ROS producers for various cellular functions related to the host defense mechanism against pathogens (innate immunity), signal transduction, and modification of the extracellular matrix [85,86]. The NOX family consists of seven homologues as follows: NOX1, NOX2 (gp91phox), NOX3, NOX4, NOX5 and dual oxidases (DOUX) 1 and 2. Enzymatic activity of NOX1-3 is regulated by complex formation with the membrane-bound p22phox and the cytosolic regulatory subunits, such as p40phox, p47phox, p67phox, and the small GTPase RAC. DUOX's activity is regulated by the maturation factors DUOXa1 and DUOXa2 [87–89]. The interaction of NOXs with various targeting proteins defines their specific subcellular localization (e.g. lamellipodial leading edge, focal adhesions, lipid rafts, the endosomes, the endoplasmic reticulum, or the nucleus) [86,87,90,91]. Therefore, in diverse cellular context, NOXs are responsible for localized ROS production and compartmentalization of redox signaling in response to growth factors, cytokines, and G-protein coupled receptor activators [90]. Expression of NOX2 is enriched in the SVZ of adult mice as compared to adjacent cortical tissue [27]. In striking contrast to mitochondriaderived superoxide (see above), several studies suggest the crucial physiological role of NOX2-generated superoxide in support of selfrenewal and multipotency of NSPCs of both niches, the SVZ [27] and the SGZ [92]. Consistent with results observed in NOX2-deficient mice [27,92], administration of apocynin, a pharmacological inhibitor of NOX, lowers the levels of superoxide and diminishes the self-renewal of SVZ NSPCs in vivo [27] and in cultured embryonic hippocampal progenitors [93]. Interestingly, NOX-deficiency affects the multipotency and favors the glial fate of SVZ-derived NSPCs [27], highlighting the role of NOXs in neuronal commitment. The beneficial effect of NOX-derived ROS on NSPC homeostasis can be explained by its spatially confined, compartment-specific production and immediate coupling into signaling cascades. Supporting this hypothesis, it was shown that downstream signaling involves ROSdependent inactivation of phosphatase and tensin homolog (PTEN) [94], the negative regulator of NSC proliferation [95–98], and the subsequent activation of phosphoinositide-3-kinase (Pi3K)/AKT pathway [27, 99]. Regulation of PTEN and AKT activity occurs upon their recruitment to the plasma membrane and mediated by the reversible oxidation of active site cysteines [100,101]. Thus, it is likely, that co-locolization of NOX, PTEN and AKT on the plasma membrane is required for proper signal transduction. Activation of NOXs is involved in signaling cascades initiated by several niche factors known to be essential for ability of NSPC to selfrenew. For example, fibroblast growth factor 2 (FGF2), a potent mitogen for NSPCs, requires the activation of NOX2 and generation of H2O2 in cultured adult hippocampal progenitors [92]. Silencing of NOX2 abrogates FGF2-induced AKT phosphorylation and inhibits proliferation [92]. Inhibition of NOX or treatment with N-acetyl-cysteine abrogates

the brain-derived neurotrophic factor (BDNF)-induced mitotic activity of NSPCs [27]. Similar to non-neural cells, it is likely that other receptors present on the surface of NSPCs may involve NOX activity for signal transduction (e.g. epidermal growth factor receptor (EGFR) and platelet-derived growth factor receptor (PDGFR) [102]). NOX2-generated ROS have been identified as the important mediators of vascular cell adhesion molecule 1 (VCAM1)-dependent maintenance of Type B NSCs in the SVZ niche [103]. VCAM1 and NOX2 are co-expressed on the end feet of Type B cells. Inhibition of VCAM1 by intra-ventricular administration of blocking antibodies leads to premature differentiation and subsequent loss of NSC pool. VCAM1 may support the response of NSCs to various niche factors via induced expression of NOX proteins. Consistent with this, silencing of VCAM1 by shRNA decreases the level of NOX2 transcripts [28]. Similarly, the positive effect of Angiotensin II on proliferation of the neural stem cell line C17.2 is linked to the induction of NOX4 and generation of superoxide [104]. NSPCs express Angiotensin II type-1 and type-2 receptors [105]. The latter is involved in Angiotensin II-induced proliferation of embryonic hippocampal NSPCs via activation of MAPK signaling [105], known target of NOX4 activity [106]. The plasma membrane-targeted DUOXs may also be involved in regulation of the NSPC fate. DUOX maturation factor, DUOXa1 (also known as a Numb-interacting protein 1) was described as an intrinsic regulator of neuronal fate in stem cell differentiation [107]. DUOXa1 is highly expressed in undifferentiated NSPCs and plays a role in DUOX1-dependent superoxide and H2O2 generation. Ectopic expression of DUOXa1 in the pluripotent embryonic carcinoma cell line P9 is associated with increased ROS levels and transient up-regulation of proneuronal genes Neurogenin1 and 2 followed by neuronal differentiation [107]. Accordingly, silencing of DUOXa1 diminishes ROS and partially inhibits retinoic acid-induced neuronal differentiation in this cell line [107]. In summary, hypoxic conditions in stem cell niches are likely required to up-regulate NOX proteins [27] in order to promote signaling cascades downstream from distinct niche signals. Of note, generation of NAD+ upon NOX activation may support glycolysis [108,109], the main cellular energy source in hypoxic microenvironment. In this regard, the bioavailability and the ratio of NAD(P)+/NAD(P)H redox couple is an important prerequisite for the maintenance of stem cell homeostasis. 3. Redox sensitive pathways in NSPCs The homeostatic self-renewal, lineage commitment and differentiation of NSCs are coordinated by several signaling pathways such as WNT/β-catenin [110] and NOTCH [111]. A growing area of interest has arisen in redox-dependent modulation of these pathways. Accumulated evidence suggests the crucial role of NOX-derived ROS, reducing NADPH-dependent thioredoxin systems and redox-sensitive molecules such as NRF2 and NAD+/NADH-dependent enzymes in spatiotemporal regulation of these signaling cascades for appropriate NSPC fate decision. 3.1. NRF2 signaling One of the emerging targets to counteract ROS and to increase intracellular oxidative defense is nuclear factor erythroid 2-related factor 2 (NRF2), a basic leucine zipper transcription factor featuring a cap ‘n’ collar structure [112]. Under basal conditions, NRF2 is retained in the cytoplasm by interaction with Kelch-like ECH-associated protein 1 (KEAP1), a cysteine rich protein, which can also serve as an adaptor protein for ubiquitin ligase E3 complex [113]. Oxidative modifications of reactive cysteine residues of KEAP1 in response to ROS lead to dissociation of the KEAP1/NRF2 complex and translocation of NRF2 into the nucleus [114]. NRF2 contains the multiple redox-sensitive nuclear export and localization signal (NES/NLS) motifs, indicating that in addition to interaction with KEAP1, the cellular distribution of NRF2

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may be per se regulated by the intracellular redox environment [115]. Nuclear heterodimerization of NRF2 with the small Maf proteins initiates transcription of a battery of cytoprotective genes harboring the antioxidant response element (ARE) in their promoter region [116,117]. Intriguingly, among the genes associated with antioxidant defense, NRF2 up-regulates transcription of genes involved in redox signaling (e.g. thioredoxin 1 [118]) and genes encoding for the key proteins of NOTCH pathway (NOTCH1 [119]; Jagged 1 [120]). The functional role of NRF2-mediated NOTCH signaling was demonstrated in a model of liver regeneration [119] and reconstitution of hematopoietic stem cell niche following myelo-suppressive irradiation [120]. Activation of NRF2 by low levels of ROS is required for NOTCH-mediated selfrenewal of airway basal stem cells [121]. Notably, NRF2 expression itself is regulated by active NOTCH signaling [122], indicating a more complex interaction between these two pathways. Taken together, these results provide evidence that ROS-sensing ability of NRF2 is integrated in NOTCH functioning in the distinct stem cell niches. Similar to other organs, NRF2 supports the stem cell-based regeneration of the neural tissue. In this regard, it was shown that NRF2 is essential for ischemia-induced neurogenesis in the SGZ niche [123]. In vitro, ectopic expression of NRF2 or treatment with a NRF2 activating compound, pyrrolidine dithiocarbamate, increases NSPCs self-renewal under proliferating conditions and supports neuronal differentiation upon mitogen withdrawal [123]. This indicates that beside its role in antioxidant defense NRF2 may be involved in the mechanisms coordinating the stem cell-fate decision. Particularly, it will be interesting to elucidate whether NRF2-dependent regulation of NOTCH contributes to NSPC homeostasis. 3.2. (NADP+)/(NADPH)-dependent signaling NADPH is generated by the pentose phosphate pathway and provides the reducing equivalents for regeneration of a variety of redox-regulating enzymes, especially members of the thioredoxin family. Oxidoreductases of this protein family are the key players in redox-dependent signaling. These proteins possess the ability to reverse oxidative modifications of protein thiols and to detoxify H2O2 [124], an important second messenger in diverse cellular processes [125]. Oxidative modifications, e.g. formation of disulphides and sulfenic acids, Sglutathionylation, or S-nitrosylation, occur at specific cysteine residues and modulate activities of an emerging number of proteins involved in signal transduction pathways. Reduction of oxidative thiol modifications is catalyzed by oxidoreductases in the expense of NADPH. 3.2.1. Thioredoxin system The thioredoxin system (thioredoxin (TRX), thioredoxin reductase and NADPH) reduces protein disulfides and S-nitrosylated thiols. Several lines of evidence suggest a role of TRXs in proliferating cells. Consistent with early stage lethality of Trx−/− embryos, ES cells isolated from TRX-deficient blastocysts fail to proliferate [126]. TRX enhances transcriptional activity of the key factor for ES cell totipotency, octamerbinding transcription factor 4 (OCT4), by increasing its DNA-binding capacity via reduction of oxidized cysteines in the DNA binding POU domain [127]. Treatment of ischemic mice with recombinant human TRX1 promotes neurogenesis via increasing NSPC proliferation in the SGZ [128,129]. Although, the precise mechanism underlying the capacity of TRX to support proliferation of NSPCs is unknown, it might be linked to its facilitating role in PTEN/PI3K/AKT signaling. Thus, TRX1 binds to PTEN in a redox-dependent manner and inhibits its lipid phosphatase activity resulting in activation of AKT [130], the well-known mediator of NSPC survival and proliferation [98,131]. Consistent with the role of TRX, mice with specific deletion of cytosolic TRX reductase 1 in the nervous system develop cerebellar dysfunction associated with impaired proliferation of cerebellar progenitors in the external granular layer [132].


3.2.2. Glutaredoxin system In addition to the reduction of protein disulfides, glutaredoxins (GRXs) has the capacity to reduce glutathionylated thiols [133]. Oxidized GRXs are re-reduced by GSH, glutathione reductase, and NADPH. GRXs 1 and 2 were shown to de-glutathionylate SIRT1, the crucial regulator of NSPC fate (see 3.3.3). However, the precise role of these molecules in NSPC biology is remained to be elucidated. Using the zebrafish model and retinoic acid-induced differentiation of SHSY5Y neuroblastoma cell line, we demonstrated that Grx2 promotes neuronal differentiation, in part by enhancing axonal outgrowth and survival [134]. 3.2.3. Peroxiredoxin system Peroxiredoxins (PRXs) constitute a family of enzymes (PRX1-6 in mammals) that catalyze the reduction of H2O2 with the use of reducing equivalents provided by the thiol-containing proteins such as TRX [124]. Notably, PRXs can be reversibly inactivated by H2O2 and their slow-rate reduction is catalyzed by sulfiredoxin or sestrin proteins. The sensitivity of PRXs to oxidation may be considered as an important “switch-off” mechanism to insure the availability of H2O2 for redox signaling [135,136]. Furthermore, the activity of cytosolic PRX1 is tightly regulated by the cyclin-dependent kinase Cdc2 during cell-cycle progression [137]. PRXs have been implicated in the maintenance of stemness in cultured cells via lowering of the intracellular levels of H2O2. Prx1−/− and Prx2−/− ES cells exhibit aberrant ROS levels and rapid loss of stemness associated with premature differentiation, a phenotype which could be rescued by N-acetyl-cysteine administration [138]. Enhancement of PRX activity by overexpression of Sestrin 3 reduces the elevated ROS levels and enhances the self-renewal capacity of FOXO-deficient NSPCs [96]. Prx1−/− embryos are characterized by deficit in post-mitotic motor neurons in the ventral spinal cord. However, this failure is not associated with either survival of motor neuron progenitors or altered neuronal commitment, but rather related to impaired capacity of proliferating Prx1−/− progenitors to exit the cell cycle [139]. This finding is consistent with the notion that active form of PRX1 is sustained in G1 [137], the critical phase for the cell cycle exit decision and differentiation. The mechanism, by which PRX1 promotes the differentiation of motor neurons relies on its ability to reduce disulphide bonds in the cytosolic part of pro-neurogenic factor GDE2 [139,140]. Whether GDE2-dependent timing of cortical neuronal differentiation [141] may similarly require the PRX activity is remained to be elucidated. The lack of the phenotype in developing spinal cord of Prx1−/− mice related to diminished self-renewal capacity [139], as it would be expected from in vitro studies [96,138], may rely on the differences in the compensatory mechanisms to detoxify H2O2. Further studies are needed to dissect the neural role of particular PRXs in ROS detoxification and in modulation of specific redox signaling. 3.2.4. Nucleoredoxin system Nucleoredoxin (NRX) is composed of three TRX-like modules and possess oxidoreductase activity [142]. In several publications, Funato et al. demonstrated the crucial role of nucleoredoxin (NRX) in regulating the activity of the WNT/β-catenin pathway [143–145]. Binding of WNT ligands to their cognate receptors activates disheveled (DVL) protein leading to inhibition of β-catenin phosphorylation by GSK-3β. Non-phosphorylated active β-catenin translocates into the nucleus and serves as a cofactor for the TCF/LEF family of transcription factors. Interaction of NRX with DVL protects the protein pool of DVL from proteasome-dependent degradation [145], but on the other hand suppresses the WNT/β-catenin signaling [143]. This interaction is primarily dependent on the redox-catalytic motif (WCPPC) of NRX and abrogated by elevated hydrogen peroxide levels [143,144]. Therefore, it is reasonable to assume, that WNT signal transduction may primarily depends on the permissive oxidative environment. Using an immortalized neural progenitor cell line ReNcell VM197, Rharass et al. demonstrated that dissociation of the NRX/DVL2 complex correlates


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with increased mitochondrial generation of ROS and activation of the WNT signaling axis in initial steps of differentiation [55]. Additionally to its role in neuronal differentiation [146], the WNT/β-catenin pathway supports self-renewal capacity of NSPCs [110,147,148]. Does the activity of WNT pathway in undifferentiated cells depend on ROS? Yoneyama et al. showed that treatment of embryonic hippocampal NSPCs with the selective superoxide scavenger, TEMPOL, reduces the level of nuclear β-catenin [93], indicating that regulation of the signaling cascade may involve endogenous sources of ROS. Accordingly, it was shown that NOX1-mediated superoxide generation is required for dissociation of the NRX/DVL complex and subsequent activation of signaling in colon epithelial cells [149,150]. In contrast to endogenous superoxide, treatment with exogenously applied H2O2 negatively regulates the WNT pathway by promoting de-phosphorylation of GSK-3β at Ser9 and therefore diminishing the pool of active β-catenin [151,152]. This difference may rely on the specific type of ROS and/or dependency on oxidative status at a particular site for signal transduction rather than overall cellular oxidation. Taken together, these data indicate that redox-dependent activity of non-nuclear NRX seems to be an important mechanism involved in spatiotemporal regulation of the WNT signaling in NSPCs. 3.3. (NAD+)/(NADH)-dependent signaling The (NAD+)/(NADH) ratio is one of the most important redox pairs due to the strong reducing capacity of NADH [153]. This ratio is mainly defined by the activity of complex I of the mitochondrial respiratory chain, making NAD+ a signal of mitochondrial OXPHOs [154]. Additionally to its function as electron carrier, this pyridine nucleotide has emerged as a critical cofactor in the signaling pathways providing a link between redox state and gene expression. Hydrolysis of NAD+ by several glycohydrolases is associated with post-translational protein modifications, such as ADP-ribosylation or deacetylation, and generation of messenger molecules (ADP-ribose, cyclic ADP-ribose or O-acetyl-ADP ribose). Several NAD+-consuming enzymes connected to signaling networks are involved in regulation of fundamental biological processes in NSPCs. Depletion of NAD+ in the adult murine brain is associated with loss of proliferating NSPCs [155]. Consistently, pharmacological inhibition or ablation of nicotinamide monophosphoribosyltransferase (NAMPT), the rate-limiting enzyme in NAD+ biosynthesis, affects G1/S cell cycle transition, self-renewal capacity and maintenance of the NSPC pool. Moreover, NAMPT plays a role in lineage commitment and differentiation of NSPCs into oligodendrocytes via activities of NAD+-dependent enzymes SIRT1 and SIRT2. 3.3.1. C-terminal binding proteins Mammalian C-terminal binding proteins (CtBP1 and CtBP2) were initially described as evolutionary conserved transcriptional corepressors [156]. Although, CtBPs share sequence homology with NAD+-dependent 2-hydroxy acid dehydrogenases [157], it is currently unclear whether this intrinsic enzymatic activity is involved in their repressor function [158]. In fact, CtBPs display high sensitivity to free nuclear NADH and alterations in the nuclear (NAD+)/(NADH) ratio modulate their binding to a large number of transcriptional repressors containing the specific PxDLS consensus motif [159,160]. Furthermore, NADH is required for dimerization of CtBP2, comprising the unique nuclear localization signal, with other CtBPs and therefore guiding their nuclear localization [161]. Consistent with the (NAD+)/(NADH)-dependency it is not unexpected that CtBP-mediated gene silencing is tightly regulated by oxygen availability and cellular metabolism [159,162,163]. Two recent studies demonstrate how the spatially and temporary defined differences in the oxygen levels may direct distinct NSPC fates by involving fundamental CtBP repressor activity [163,164]. Developing chick's neural tube exhibits a region-specific pattern in the oxygen gradients with highest oxygen level in the dorsal roof plate and decreasing in a graded manner towards the intermediate regions. High oxygen

concentrations in the non-neurogenic roof plate inhibit neuronal fate via establishing the CtBP/hairy and enhancer of split homolog 1 (HES1) repressor complex binding to the promoter region of proneuronal factor mouse atonal homolog 1 (MATH1) [163]. For comparison, in other parts of the dorsal neural tube, neurogenesis is permitted by more hypoxic environment [163]. In these regions CtBP promotes cell-cycle exit and neuronal differentiation through repression of transcriptional activities of Wnt-target genes by forming a repressor complex with β-catenin/T cell factor [164]. Interestingly, lowering (NAD+)/(NADH) ratio in cultured NSPCs by decreasing oxygen tension enhances CtBP-mediated transcriptional repression of HES1 [165]. Taken together, these data suggest that CtBP act as a master regulator of oxygen-dependent switch in activity of signaling pathways during stem cell fate decision. Consistent with this notion, CtBP2-deficient mouse embryos exhibit severe developmental defects related to changes in embryonic patterning [166]. It is reasonable to hypothesize that redox/metabolic state will determine the mode of CtBP action in particular cellular system. Therefore, differences in metabolic processes could explain the either promoting [167–169] or inhibiting [169,170] effects of CtBP on the WNT signaling axis reported in literature. 3.3.2. Poly(ADP-ribose) polymerase NAD+-dependent poly(ADP-ribose) polymerase 1 (PARP1) is another example of redox sensing enzymes that modulate activity of the signaling pathways and define the gene transcription profile. Initially identified as a DNA single-strand break repairing enzyme [171,172], PARP1 has been further characterized as a co-transcriptional regulator localized at the promoters of actively transcribed genes [173]. The capacity of PARP1 to modulate chromatin structures and the activity of transcription factors is dependent on the amount of NAD+ and its poly-ADP-ribosylation activity [173–176]. PARP1 contributes to the maintenance of transcriptional activity of a wide range of genes in ES cells [177]. Upon differentiation of ES cells, PARP1 enhances FGF4 transcription by poly-ADPribosylating SOX2 at the FGF4 promoter region [178]. In NSPCs, PARP1 enzymatic activity is required for dissociation of groucho/TLE co-repressor complex from the promoter of pro-neuronal mouse achaete-scute complex homolog 1 (MASH1) gene [175]. In the postnatal SVZ, deficiency of PARP1 directs cells towards glial fate, further supporting the proneuronal role of PARP1 [179]. Therefore, NAD+-dependent PARP1 activity may serve as a mechanism to promote the differentiating program by regulating repressor/co-repressor dissociation from the promoters of specific genes [175]. 3.3.3. Sirtuins SIRT1, the mammalian ortholog of yeast silent information regulator 2 (SIR2), belongs to an evolutionary conserved family of proteins (sirtuins; 7 members in mammals) with NAD+-dependent de-acetylating and mono-ribosyltransferase activity [180]. SIR2 was first identified as a histone deacetylase that suppresses genome-wide transcription and extends replicative lifespan [181]. Since then, numerous target genes and non-histone substrates of SIRT1 de-acetylating activity have been identified. SIRT1 expression and activity is tightly regulated by changes in the (NAD+)/(NADH) redox state [182,183]. Thus, the availability of the metabolic cofactor NAD+ determines the catalytic activity of SIRT1 and other sirtuins [184–187]. SIRT1 transcription is controlled by the redox-sensing ability of the transcriptional co-repressor CtBP [188]. Increase in the (NAD+)/(NADH) ratio affects the binding of the CtBP/ hypermethylated in cancer 1 (HIC1) complex to the SIRT1 promoter and leads to de-repression of SIRT1 transcription. This regulation seems to be involved in low oxygen-mediated SIRT1 suppression [188], since hypoxic conditions lead to a drastic increase in the levels of NADH and activation of the repressor function of CtBP [159]. Depletion of NADH levels by 2-deoxyglucose, an inhibitor of glycolysis, or by treatment with pyruvate restores SIRT1 expression under hypoxic conditions [188]. In addition, the activity and expression of SIRT1 are modulated by the GSH/GSSG redox couple in cell context-dependent manner. Although

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Oxygen, nutrients, growth factors

Stem cell niche

increasing evidence suggests that redox regulation via GSH is enzymedependent, the ratio between reduced GSH and the oxidized glutathione disulfide (GSSG) is still considered as an important regulator of cellular redox events [189]. Depletion of GSH in NSPCs sustains the transcriptional and translational level of SIRT1 upon differentiating conditions [30]. Regulation of SIRT1's enzymatic activity is connected to reversible oxidative thiol modifications. SIRT1 activity is inhibited following GAPDHdependent S-nitrosylation [190] or S-glutathionylation of critical cysteinyl groups [134,186,191,192]. Reduction of S-glutathionylated cysteines by GRXs re-activates SIRT1 [134,186,192]. Irreversible oxidative modifications, i.e. carbonylation, mark SIRT1 for proteasomal degradation [186]. Sirtuins are associated with a variety of cellular functions connected to redox regulation (for recent reviews see [193,194]). High levels of sirtuins are associated with more oxidized state of cells [195,196], suggesting their role in cellular anti-oxidative defense mechanisms. Sirtuins 1, 2, and 3 reduce cellular ROS damage by FOXO3-mediated activation of both, SOD2 and catalase [195–198]. The expression of SOD2 and other antioxidant proteins upon activation of NRF2 is enhanced via SIRT1-dependent deacetylation of p53 [199,200]. SIRT1 knockout mice are smaller at birth, exhibit several developmental defects, and show elevated postnatal lethality [201,202]. Consistent with an early expression of SIRT1 during CNS development [203], three independently generated Sirt1−/− mouse strains exhibit frequent CNS abnormalities characterized by neural tube closure defects (exencephaly) and disturbed neuroretinal morphogenesis [201,202,204]. These animals display cognitive deficits associated with reduced synaptic plasticity and altered activity of hippocampal genes encoding for proteins involved in synaptic function, lipid metabolism and myelination [205]. Similar to CtBP, SIRT1-mediated transcriptional regulation is implicated in the self-renewal as well as in differentiation processes of NSPCs. The ability of SIRT1 to modulate distinct NSPC fates is at least in part dependent on its interaction with specific transcription factors at particular differentiation state of the cell. For example, Ichi et al. showed that SIRT1 supports NSPC maintenance in the developing brain by deacetylating the transcription factor paired-box 3 (PAX3) and subsequent induction of the basic helix-loop-helix (bHLH) transcriptional factor HES1 [206]. In addition, SIRT1-mediated interaction of PAX3




with another bHLH transcription factor, the pro-neuronal factor Neurogenin 2, impairs its activity to initiate differentiation [206]. Consistent with these findings, down-regulation of SIRT1 in human and mouse ES cells is required for transcriptional de-repression of a set of pro-neuronal genes and the establishment of the neuroectodermal lineage differentiation program [207]. Further, upon pro-oxidative challenge (such as GSH depletion), sustained expression of SIRT1 in NSPCs inhibits neuronal fate by promoting HES1-mediated repression of the pro-neuronal gene MASH1 [30]. However, in differentiating NSPCs SIRT1 functions in opposite way to suppress the promoter activity of specific stemness genes [208,209]. This is consistent with its predicted role in cell differentiation as a component of the polycomb repressive complex 4 (PRC4) [210]. Thus, upon neuronal differentiation SIRT1 transiently translocates to the nucleus [208] to form a complex with nuclear receptor co-repressor (N-coR) and B-cell lymphoma 6 (BCL6) thereby suppressing HES1 [208] and HES5 [209] genes, respectively. The roles of SIRT1 in promoting neuronal differentiation under normal conditions [208,209] and inhibition of neuronal commitment upon oxidative challenge [30] coincide with a data showing that oxidative conditions cause a relocation of chromatin-associated SIRT1 across the genome leading to changes in the expression of individual genes [211]. Altogether these findings support the key role of SIRT1 in redox/metabolism-dependent coordination of NSPC fate. Such broad spectrum of SIRT1 functions in proliferating and differentiating cells requires tight control of SIRT1 activity to ensure proper NSPC differentiation. In this regard, identification of factors that regulate SIRT1 nuclearcytoplasmic shuttling [212] and defining the subcellular alterations in the NAD+/NADH ratio might lead to a better understanding of specific SIRT1-driven mechanisms. 4. Concluding remarks The intracellular redox state is emerging as an intrinsic regulatory mechanism for the long-term maintenance of the stem cell pool and directing the differentiation of essential cell lineages for regeneration and neural tissue homeostasis (Fig. 2). As demonstrated in this review, considerable attention needs to be paid to the source of ROS and the

Self-renewal Multipotency








Metabolism Glycolosis PPP










Biogenesis OXPHOS

Fig. 2. Schematic summary of the main neural stem cell (NSC) functions and their dependency on ROS and redox couples. Anion superoxide (O− 2 ) is the principal form of ROS produced by membrane-bound NAD(P)H oxidase (NOX) complex or mitochondria electron-transport chain. During the activation of NSCs NOX enzymes shift redox homeostasis towards more oxidized state and trigger self-renewal. In contrast, mitochondria-derived ROS are implicated in NSPC differentiation. To maintain redox signaling, the living cells engage NADPH-dependent oxidoreductases (TRX1, NRX, PRX1, GRX2) to reduce oxidative thiol modifications in redox-sensitive cysteine residues. NAD+/NADH-dependent transcriptional co-repressors (CtBP, SIRT1, PARP1) further modulate the specific functions of signaling pathways to promote stemness or differentiation of NSPCs.


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compartmentalization of redox signaling for NSC fate determination. Notably, experimental approaches based on the analysis of exogenous application of ROS, such as treatment with H2O2, generates conflicting results, particularly regarding the ability of NSPCs to self-renew. Therefore, these types of experimental data were not included within the review. Obviously, modulation of ROS levels in defined cellular compartment by using specific pharmacological agents or genetic modifications represents a far more reliable method. In this review, we mainly used the term “NSPCs” to describe the uncharacterized populations of multipotent stem cells with the capacity to self-renew. In fact, these cells encompass the population of quiescent NSCs (qNSCs) retaining non-proliferating state in typical culture conditions (10 and 11) and their mitotically-active progenies. Due to technical challenge, the most approaches are used to characterize the actively dividing progenitors and do not address the role of redox state in the biology and plasticity of qNSCs, the major population of NSCs in the adult mammalian brain. Thus, the development of new strategies (e.g. recently described in [11,12]) that allow to distinguish between these two population is of great importance for NSC research and may advance our knowledge about redox-dependent mechanisms in the long-term maintenance of endogenous stem cell pool. Therapeutic interventions altering the redox/metabolic state of CNS tissue may have a profound impact on the vital NSC functions and has to be considered with caution. On the other hand, the increasing knowledge of redox-dependent mechanisms will help to establish novel approaches to elicit NSCs-based regenerative capacity of CNS tissue in various pathological states and normal aging. Authors' contributions TP, RS, CB, and OA wrote the manuscript. HPH provided critical input. All authors read and approved the final manuscript. Competing interests The authors declare that they have no competing interests. Acknowledgements We would like to thank Jason Cline (Department of Neurology, Düsseldorf) for proofreading the manuscript. We apologize to the researchers whose work is not cited. Our research is supported by the grants from the “Forschungskommission” of the Medical Faculty (Heinrich Heine University Düsseldorf) to T.P., C.B. and O.A. (06/2012), the Hertie Foundation to C.B. and O.A. (P1120028), the DFG (German research foundation) priority program SPP1710 “Dynamics of thiolbased redox switches in cellular physiology” to C.B. (BE 3259/5-1) and the Walter and Ilse Rose Foundation (to O.A.). References [1] F. Doetsch, I. Caille, D.A. Lim, J.M. Garcia-Verdugo, A. Alvarez-Buylla, Subventricular zone astrocytes are neural stem cells in the adult mammalian brain, Cell 97 (1999) 703–716. [2] A. Kriegstein, A. Alvarez-Buylla, The glial nature of embryonic and adult neural stem cells, Annu. Rev. Neurosci. 32 (2009) 149–184. [3] F.H. Gage, Mammalian neural stem cells, Science 287 (2000) 1433–1438. [4] G.L. Ming, H.J. Song, Adult neurogenesis in the mammalian brain: significant answers and significant questions, Neuron 70 (2011) 687–702. [5] Z. Mirzadeh, F.T. Merkle, M. Soriano-Navarro, J.M. Garcia-Verdugo, A. AlvarezBuylla, Neural stem cells confer unique pinwheel architecture to the ventricular surface in neurogenic regions of the adult brain, Cell Stem Cell 3 (2008) 265–278. [6] F.D. Miller, A. Gauthier-Fisher, Home at last: neural stem cell niches defined, Cell Stem Cell 4 (2009) 507–510. [7] D.K. Ma, M.A. Bonaguidi, G.L. Ming, H.J. Song, Adult neural stem cells in the mammalian central nervous system, Cell Res. 19 (2009) 672–682. [8] S. Ahn, A.L. Joyner, In vivo analysis of quiescent adult neural stem cells responding to Sonic hedgehog, Nature 437 (2005) 894–897.

[9] E. Pastrana, L.C. Cheng, F. Doetsch, Simultaneous prospective purification of adult subventricular zone neural stem cells and their progeny, Proc. Natl. Acad. Sci. U. S. A. 106 (2009) 6387–6392. [10] C. Giachino, V. Taylor, Lineage analysis of quiescent regenerative stem cells in the adult brain by genetic labelling reveals spatially restricted neurogenic niches in the olfactory bulb, Eur. J. Neurosci. 30 (2009) 9–24. [11] J.K. Mich, R.A.J. Signer, D. Nakada, A. Pineda, R.J. Burgess, T.Y. Vue, J.E. Johnson, S.J. Morrison, Prospective identification of functionally distinct stem cells and neurosphere-initiating cells in adult mouse forebrain, Elife 3 (2014). [12] P. Codega, V. Silva-Vargas, A. Paul, A.R. Maldonado-Soto, A.M. Deleo, E. Pastrana, F. Doetsch, Prospective identification and purification of quiescent adult neural stem cells from their in vivo niche, Neuron 82 (2014) 545–559. [13] C.M. Morshead, B.A. Reynolds, C.G. Craig, M.W. McBurney, W.A. Staines, D. Morassutti, S. Weiss, D. Vanderkooy, Neural stem-cells in the adult mammalian forebrain — a relatively quiescent subpopulation of subependymal cells, Neuron 13 (1994) 1071–1082. [14] C. Lois, A. Alvarezbuylla, Long-distance neuronal migration in the adult mammalian brain, Science 264 (1994) 1145–1148. [15] B. Nait-Oumesmar, L. Decker, F. Lachapelle, V. Avellana-Adalid, C. Bachelin, A. Baron-Van Evercooren, Progenitor cells of the adult mouse subventricular zone proliferate, migrate and differentiate into oligodendrocytes after demyelination, Eur. J. Neurosci. 11 (1999) 4357–4366. [16] B. Menn, J.M. Garcia-Verdugo, C. Yaschine, O. Gonzalez-Perez, D. Rowitch, A. Alvarez-Buylla, Origin of oligodendrocytes in the subventricular zone of the adult brain, J. Neurosci. 26 (2006) 7907–7918. [17] G. Kempermann, S. Jessberger, B. Steiner, G. Kronenberg, Milestones of neuronal development in the adult hippocampus, Trends Neurosci. 27 (2004) 447–452. [18] H. Suh, A. Consiglio, J. Ray, T. Sawai, K.A. D'Amour, F.H. Gage, In vivo fate analysis reveals the multipotent and self-renewal capacities of Sox2(+) neural stem cells in the adult hippocampus, Cell Stem Cell 1 (2007) 515–528. [19] C.M. Zhao, W. Deng, F.H. Gage, Mechanisms and functional implications of adult neurogenesis, Cell 132 (2008) 645–660. [20] J.R. Fike, R. Rola, C.L. Limoli, Radiation response of neural precursor cells, Neurosurg. Clin. N. Am. 18 (2007) 115-+. [21] J.R. Fike, S. Rosi, C.L. Limoli, Neural precursor cells and central nervous system radiation sensitivity, Semin. Radiat. Oncol. 19 (2009) 122–132. [22] M. Noble, C. Proschel, M. Mayer-Proschel, Oxidative-reductionist approaches to stem and progenitor cell function, Cell Stem Cell 8 (2011) 1–2. [23] J. Smith, E. Ladi, M. Mayer-Proschel, M. Noble, Redox state is a central modulator of the balance between self-renewal and differentiation in a dividing glial precursor cell, Proc. Natl. Acad. Sci. U. S. A. 97 (2000) 10032–10037. [24] K. Suzukawa, K. Miura, J. Mitsushita, J. Resau, K. Hirose, R. Crystal, T. Kamata, Nerve growth factor-induced neuronal differentiation requires generation of Rac1regulated reactive oxygen species, J. Biol. Chem. 275 (2000) 13175–13178. [25] Y. Goldsmit, S. Erlich, R. Pinkas-Kramarski, Neuregulin induces sustained reactive oxygen species generation to mediate neuronal differentiation, Cell. Mol. Neurobiol. 21 (2001) 753–769. [26] M. Noble, J. Smith, J. Power, M. Mayer-Proschel, Redox state as a central modulator of precursor cell function, Ann. N. Y. Acad. Sci. 991 (2003) 251–271. [27] J.E. Le Belle, N.M. Orozco, A.A. Paucar, J.P. Saxe, J. Mottahedeh, A.D. Pyle, H. Wu, H.I. Kornblum, Proliferative neural stem cells have high endogenous ROS levels that regulate self-renewal and neurogenesis in a PI3K/Akt-dependant manner, Cell Stem Cell 8 (2011) 59–71. [28] E. Kokovay, Y. Wang, G. Kusek, R. Wurster, P. Lederman, N. Lowry, Q. Shen, S. Temple, VCAM1 is essential to maintain the structure of the SVZ niche and acts as an environmental sensor to regulate SVZ lineage progression, Cell Stem Cell 11 (2012) 220–230. [29] Y. Hou, M.P. Mattson, A.W. Cheng, Permeability transition pore-mediated mitochondrial superoxide flashes regulate cortical neural progenitor differentiation, PLos One 8 (2013). [30] T. Prozorovski, U. Schulze-Topphoff, R. Glumm, J. Baumgart, F. Schroter, O. Ninnemann, E. Siegert, I. Bendix, O. Brustle, R. Nitsch, F. Zipp, O. Aktas, Sirt1 contributes critically to the redox-dependent fate of neural progenitors, Nat. Cell Biol. 10 (2008) 385–394. [31] I. Silver, M. Erecinska, Oxygen and ion concentrations in normoxic and hypoxic brain cells, Adv. Exp. Med. Biol. 454 (1998) 7–16. [32] L.L. Zhu, L.Y. Wu, D.T. Yew, M. Fan, Effects of hypoxia on the proliferation and differentiation of NSCs, Mol. Neurobiol. 31 (2005) 231–242. [33] J. Mazumdar, W.T. O'Brien, R.S. Johnson, J.C. LaManna, J.C. Chavez, P.S. Klein, M.C. Simon, O2 regulates stem cells through Wnt/beta-catenin signalling, Nat. Cell Biol. 12 (2010) 1007–1013. [34] G. Santilli, G. Lamorte, L. Carlessi, D. Ferrari, N.L. Rota, E. Binda, D. Delia, A.L. Vescovi, F.L. De, Mild hypoxia enhances proliferation and multipotency of human neural stem cells, PLoS One 5 (2010) e8575. [35] H.L. Chen, F. Pistollato, D.J. Hoeppner, H.T. Ni, R.D. McKay, D.M. Panchision, Oxygen tension regulates survival and fate of mouse central nervous system precursors at multiple levels, Stem Cells 25 (2007) 2291–2301. [36] A. Mohyeldin, T. Garzon-Muvdi, A. Quinones-Hinojosa, Oxygen in stem cell biology: a critical component of the stem cell niche, Cell Stem Cell 7 (2010) 150–161. [37] K. Zhang, L. Zhu, M. Fan, Oxygen, a key factor regulating cell behavior during neurogenesis and cerebral diseases, Front. Mol. Neurosci. 4 (2011) 5. [38] C.A.V. Rodrigues, M.M. Diogo, C.L. da Silva, J.M.S. Cabral, Hypoxia enhances proliferation of mouse embryonic stem cell-derived neural stem cells, Biotechnol. Bioeng. 106 (2010) 260–270.

T. Prozorovski et al. / Biochimica et Biophysica Acta 1850 (2015) 1543–1554 [39] A.W. Cheng, R.Q. Wan, J.L. Yang, N. Kamimura, T.G. Son, X. Ouyang, Y.Q. Luo, E. Okun, M.P. Mattson, Involvement of PGC-1 alpha in the formation and maintenance of neuronal dendritic spines, Nat. Commun. 3 (2012). [40] K. Ito, T. Suda, Metabolic requirements for the maintenance of self-renewing stem cells, Nat. Rev. Mol. Cell Biol. 15 (2014) 243–256. [41] K. Ito, A. Hirao, F. Arai, S. Matsuoka, K. Takubo, I. Hamaguchi, K. Nomiyama, K. Hosokawa, K. Sakurada, N. Nakagata, Y. Ikeda, T.W. Mak, T. Suda, Regulation of oxidative stress by ATM is required for self-renewal of haematopoietic stem cells, Nature 431 (2004) 997–1002. [42] K. Ito, A. Hirao, F. Arai, K. Takubo, S. Matsuoka, K. Miyamoto, M. Ohmura, K. Naka, K. Hosokawa, Y. Ikeda, T. Suda, Reactive oxygen species act through p38 MAPK to limit the lifespan of hematopoietic stem cells, Nat. Med. 12 (2006) 446–451. [43] H. Kondoh, M.E. Lleonart, Y. Nakashima, M. Yokode, M. Tanaka, D. Bernard, J. Gil, D. Beach, A high glycolytic flux supports the proliferative potential of murine embryonic stem cells, Antioxid. Redox Signal. 9 (2007) 293–299. [44] S. Varum, O. Momcilovic, C. Castro, A. Ben-Yehudah, J. Ramalho-Santos, C.S. Navara, Enhancement of human embryonic stem cell pluripotency through inhibition of the mitochondrial respiratory chain, Stem Cell Res. 3 (2009) 142–156. [45] A. Prigione, B. Fauler, R. Lurz, H. Lehrach, J. Adjaye, The senescence-related mitochondrial/oxidative stress pathway is repressed in human induced pluripotent stem cells, Stem Cells 28 (2010) 721–733. [46] S. Orrenius, A. Gogvadze, B. Zhivotovsky, Mitochondrial oxidative stress: implications for cell death, Annu. Rev. Pharmacol. Toxicol. 47 (2007) 143–183. [47] W.J.H. Koopman, L.G.J. Nijtmans, C.E.J. Dieteren, P. Roestenberg, F. Valsecchi, J.A.M. Smeitink, P.H.G.M. Willems, Mammalian mitochondrial complex I: biogenesis regulation, and reactive oxygen species generation, Antioxid. Redox Signal. 12 (2010) 1431–1470. [48] Y. Hou, X. Ouyang, R.Q. Wan, H.P. Cheng, M.P. Mattson, A.W. Cheng, Mitochondrial superoxide production negatively regulates neural progenitor proliferation and cerebral cortical development, Stem Cells 30 (2012) 2535–2547. [49] M. Schwarzlander, S. Wagner, Y.G. Ermakova, V.V. Belousov, R. Radi, J.S. Beckman, G.R. Buettner, N. Demaurex, M.R. Duchen, H.J. Forman, M.D. Fricker, D. Gems, A.P. Halestrap, B. Halliwell, U. Jakob, I.G. Johnstonn, N.S. Jones, D.C. Logan, B. Morgan, F.L. Muller, D.G. Nicholls, S.J. Remington, P.T. Schumacker, C.C. Winterbourn, L.J. Sweetlove, A.J. Meyer, T.P. Dick, M.P. Murphy, The ‘mitoflash’ probe cpYFP does not respond to superoxide, Nature 514 (2014) E12–E14. [50] M. Breckwoldt, F.M.J. Pfister, P.M. Bradley, P. Marinkovic, P.R. Williams, M.S. Brilll, B. Plomer, A. Schmalz, D.K. St Clair, R. Naumann, O. Griesbeck, M. Schwarzlander, L. Godinho, F.M. Bareyre, T.P. Dick, M. Kerschensteiner, T. Misgeld, Multiparametric optical analysis of mitochondrial redox signals during neuronal physiology and pathology in vivo, Nat. Med. 20 (2014) (559-122). [51] T.T. Huang, Y. Zou, R. Corniola, Oxidative stress and adult neurogenesis—effects of radiation and superoxide dismutase deficiency, Semin. Cell Dev. Biol. 23 (2012) 738–744. [52] L.A. Voloboueva, S.W. Lee, J.F. Emery, T.D. Palmer, R.G. Giffard, Mitochondrial protection attenuates inflammation-induced impairment of neurogenesis in vitro and in vivo, J. Neurosci. 30 (2010) 12242–12251. [53] O. Yanes, J. Clark, D.M. Wong, G.J. Patti, A. Sanchez-Ruiz, H.P. Benton, S.A. Trauger, C. Desponts, S. Ding, G. Siuzdak, Metabolic oxidation regulates embryonic stem cell differentiation, Nat. Chem. Biol. 6 (2010) 411–417. [54] M. Tsatmali, E.C. Walcott, K.L. Crossin, Newborn neurons acquire high levels of reactive oxygen species and increased mitochondrial proteins upon differentiation from progenitors, Brain Res. 1040 (2005) 137–150. [55] T. Rharass, H. Lemcke, M. Lantow, S.A. Kuznetsov, D.G. Weiss, D. Panakova, Ca2+mediated mitochondrial ROS metabolism augments Wnt/beta-catenin pathway activation to facilitate cell differentiation, J. Biol. Chem. 289 (2014) 27937–27951. [56] D.T.W. Chang, I.J. Reynolds, Differences in mitochondrial movement and morphology in young and mature primary cortical neurons in culture, Neuroscience 141 (2006) 727–736. [57] M. Tsatmali, E.C. Walcott, H. Makarenkova, K.L. Crossin, Reactive oxygen species modulate the differentiation of neurons in clonal cortical cultures, Mol. Cell. Neurosci. 33 (2006) 345–357. [58] K. Forsberg, A. Wuttke, G. Quadrato, P.M. Chumakov, A. Wizenmann, S. Di Giovanni, The tumor suppressor p53 fine-tunes reactive oxygen species levels and neurogenesis via PI3 kinase signaling, J. Neurosci. 33 (2013) 14318–14330. [59] A.V. Molofsky, R. Pardal, T. Iwashita, I.K. Park, M.F. Clarke, S.J. Morrison, Bmi-1 dependence distinguishes neural stem cell self-renewal from progenitor proliferation, Nature 425 (2003) 962–967. [60] I.K. Park, D.L. Qian, M. Kiel, M.W. Becker, M. Pihalja, I.L. Weissman, S.J. Morrison, M.F. Clarke, Bmi-1 is required for maintenance of adult self-renewing haematopoietic stem cells, Nature 423 (2003) 302–305. [61] A.V. Molofsky, S. He, M. Bydon, S.J. Morrison, R. Pardal, Bmi-1 promotes neural stem cell self-renewal and neural development but not mouse growth and survival by repressing the p16Ink4a and p19Arf senescence pathways, Genes Dev. 19 (2005) 1432–1437. [62] S.W. Bruggeman, M.E. Valk-Lingbeek, P.P. van der Stoop, J.J. Jacobs, K. Kieboom, E. Tanger, D. Hulsman, C. Leung, Y. Arsenijevic, S. Marino, L.M. van, Ink4a and Arf differentially affect cell proliferation and neural stem cell self-renewal in Bmi1deficient mice, Genes Dev. 19 (2005) 1438–1443. [63] C.A. Fasano, T.N. Phoenix, E. Kokovay, N. Lowry, Y. Elkabetz, J.T. Dimos, I.R. Lemischka, L. Studer, S. Temple, Bmi-1 cooperates with Foxg1 to maintain neural stem cell self-renewal in the forebrain, Genes Dev. 23 (2009) 561–574. [64] J. Liu, L. Cao, J.C. Chen, S.W. Song, I.H. Lee, C. Quijano, H.J. Liu, K. Keyvanfar, H.Q. Chen, L.Y. Cao, B.H. Ahn, N.G. Kumar, I.I. Rovira, X.L. Xu, M. van Lohuizen, N. Motoyama, C.X. Deng, T. Finkel, Bmi1 regulates mitochondrial function and the DNA damage response pathway, Nature 459 (2009) 387-U100.


[65] W. Chatoo, M. Abdouh, R.H. Duparc, G. Bernier, Bmi1 distinguishes immature retinal progenitor/stem cells from the main progenitor cell population and is required for normal retinal development, Stem Cells 28 (2010) 1412–1423. [66] T. Nakamura, T. Nishizawa, M. Hagiya, T. Seki, M. Shimonishi, A. Sugimura, K. Tashiro, S. Shimizu, Molecular-cloning and expression of human hepatocyte growth-factor, Nature 342 (1989) 440–443. [67] S. Chuikov, B.P. Levi, M.L. Smith, S.J. Morrison, Prdm16 promotes stem cell maintenance in multiple tissues, partly by regulating oxidative stress, Nat. Cell Biol. 12 (2010) 999–1006. [68] C. Nicoleau, O. Benzakour, F. Agasse, N. Thiriet, J. Petit, L. Prestoz, M. Roger, M. Jaber, V. Coronas, Endogenous hepatocyte growth factor is a niche signal for subventricular zone neural stem cell amplification and self-renewal, Stem Cells 27 (2009) 408–419. [69] Y.S. Yoon, J.H. Lee, S.C. Hwang, K.S. Choi, G. Yoon, TGF beta 1 induces prolonged mitochondrial ROS generation through decreased complex IV activity with senescent arrest in Mv1Lu cells, Oncogene 24 (2005) 1895–1903. [70] C.R. Wang, C.C. Liang, Z.C. Bian, Y. Zhu, J.L. Guan, FIP200 is required for maintenance and differentiation of postnatal neural stem cells, Nat. Neurosci. 16 (2013) 532-+. [71] T. Hara, A. Takamura, C. Kishi, S.I. Iemura, T. Natsume, J.L. Guan, N. Mizushima, FIP200, a ULK-interacting protein, is required for autophagosome formation in mammalian cells, J. Cell Biol. 181 (2008) 497–510. [72] S.M. Nam, J.W. Kim, D.Y. Yoo, J.H. Choi, W. Kim, H.Y. Jung, M.H. Won, I.K. Hwang, J.K. Seong, Y.S. Yoon, Effects of treadmill exercise on neural stem cells, cell proliferation, and neuroblast differentiation in the subgranular zone of the dentate gyrus in cyclooxygenase-2 knockout mice, Neurochem. Res. 38 (2013) 2559–2569. [73] T. Sasaki, K. Kitagawa, S. Sugiura, E. Omura-Matsuoka, S. Tanaka, Y. Yagita, H. Okano, M. Matsumoto, M. Hori, Implication of cyclooxygenase-2 on enhanced proliferation of neural progenitor cells in the adult mouse hippocampus after ischemia, J. Neurosci. Res. 72 (2003) 461–471. [74] K. Wada, M. Arita, A. Nakajima, K. Katayama, C. Kudo, Y. Kamisaki, C.N. Serhan, Leukotriene B-4 and lipoxin A(4) are regulatory signals for neural stem cell proliferation and differentiation, FASEB J. 20 (2006) 1785–1792. [75] C. Huber, J. Marschallinger, H. Tempfer, T. Furtner, S. Couillard-Despres, H.C. Bauer, F.J. Rivera, L. Aigner, Inhibition of leukotriene receptors boosts neural progenitor proliferation, Cell. Physiol. Biochem. 28 (2011) 793–804. [76] E.R. Matarredona, M. Murillo-Carretero, B. Moreno-Lopez, C. Estrada, Role of nitric oxide in subventricular zone neurogenesis, Brain Res. Rev. 49 (2005) 355–366. [77] A. Torroglosa, M. Murillo-Carretero, C. Romero-Grimaldi, E.R. Matarredona, A. Campos-Caro, C. Estrada, Nitric oxide decreases subventricular zone stem cell proliferation by inhibition of epidermal growth factor receptor and phosphoinositide3-kinase/Akt pathway, Stem Cells 25 (2007) 88–97. [78] S.J. Kim, M.S. Lim, S.K. Kang, Y.S. Lee, K.S. Kang, Impaired functions of neural stem cells by abnormal nitric oxide-mediated signaling in an in vitro model of Niemann– Pick type C disease, Cell Res. 18 (2008) 686–694. [79] C.X. Luo, X. Jin, C.C. Cao, M.M. Zhu, B. Wang, L. Chang, Q.G. Zhou, H.Y. Wu, D.Y. Zhu, Bidirectional regulation of neurogenesis by neuronal nitric oxide synthase derived from neurons and neural stem cells, Stem Cells 28 (2010) 2041–2052. [80] C.T. Lee, J. Chen, T. Hayashi, S.Y. Tsai, J.F. Sanchez, S.L. Errico, R. Amable, T.P. Su, R.H. Lowe, M.A. Huestis, J. Shen, K.G. Becker, H.M. Geller, W.J. Freed, A mechanism for the inhibition of neural progenitor cell proliferation by cocaine, PLoS Med. 5 (2008) 987–1004. [81] M.J. Ricard, L.J. Gudas, Cytochrome P450 Cyp26a1 alters spinal motor neuron subtype identity in differentiating embryonic stem cells, J. Biol. Chem. 288 (2013) 28801–28813. [82] K. Bedard, K.H. Krause, The NOX family of ROS-generating NADPH oxidases: physiology and pathophysiology, Physiol. Rev. 87 (2007) 245–313. [83] K. Bedard, B. Lardy, K.H. Krause, NOX family NADPH oxidases: not just in mammals, Biochimie 89 (2007) 1107–1112. [84] H. Sumimoto, Structure, regulation and evolution of Nox-family NADPH oxidases that produce reactive oxygen species, FEBS J. 275 (2008) 3249–3277. [85] T.L. Leto, M. Geiszt, Role of Nox family NADPH oxidases in host defense, Antioxid. Redox Signal. 8 (2006) 1549–1561. [86] J.D. Lambeth, Nox enzymes and the biology of reactive oxygen, Nat. Rev. Immunol. 4 (2004) 181–189. [87] J.D. Lambeth, T. Kawahara, B. Diebold, Regulation of Nox and Duox enzymatic activity and expression, Free Radic. Biol. Med. 43 (2007) 319–331. [88] I. Al Ghouleh, N.K.H. Khoo, U.G. Knaus, K.K. Griendling, R.M. Touyz, V.J. Thannickal, A. Barchowsky, W.M. Nauseef, E.E. Kelley, P.M. Bauer, V. Darley-Usmar, S. Shiva, E. Cifuentes-Pagano, B.A. Freeman, M.T. Gladwin, P.J. Pagano, Oxidases and peroxidases in cardiovascular and lung disease: new concepts in reactive oxygen species signaling, Free Radic. Biol. Med. 51 (2011) 1271–1288. [89] D.I. Brown, K.K. Griendling, Nox proteins in signal transduction, Free Radic. Biol. Med. 47 (2009) 1239–1253. [90] M. Ushio-Fukai, Compartmentalization of redox signaling through NADPH oxidase-derived ROS, Antioxid. Redox Signal. 11 (2009) 1289–1299. [91] D.I. Brown, K.K. Griendling, Nox proteins in signal transduction, Free Radic. Biol. Med. 47 (2009) 1239–1253. [92] B.C. Dickinson, J. Peltier, D. Stone, D.V. Schaffer, C.J. Chang, Nox2 redox signaling maintains essential cell populations in the brain, Nat. Chem. Biol. 7 (2011) 106–112. [93] M. Yoneyama, K. Kawada, Y. Gotoh, T. Shiba, K. Ogita, Endogenous reactive oxygen species are essential for proliferation of neural stem/progenitor cells, Neurochem. Int. 56 (2010) 740–746. [94] W.H. Cui, K. Matsuno, K. Iwata, M. Ibi, M. Matsumoto, J. Zhang, K. Zhu, M. Katsuyama, N.J. Torok, C. Yabe-Nishimura, NOX1/nicotinamide adenine dinucleotide phosphate, reduced form (NADPH) oxidase promotes proliferation of stellate









[102] [103]


[105] [106]





[111] [112] [113]

[114] [115] [116]

[117] [118] [119]



T. Prozorovski et al. / Biochimica et Biophysica Acta 1850 (2015) 1543–1554 cells and aggravates liver fibrosis induced by bile duct ligation, Hepatology 54 (2011) 949–958. H. Sun, R. Lesche, D.M. Li, J. Liliental, H. Zhang, J. Gao, N. Gavrilova, B. Mueller, X. Liu, H. Wu, PTEN modulates cell cycle progression and cell survival by regulating phosphatidylinositol 3,4,5,-trisphosphate and Akt protein kinase B signaling pathway, Proc. Natl. Acad. Sci. U. S. A. 96 (1999) 6199–6204. J.H. Paik, Z.H. Ding, R. Narurkar, S. Ramkissoon, F. Muller, W.S. Kamoun, S.S. Chae, H.W. Zheng, H.Q. Ying, J. Mahoney, D. Hiller, S. Jiang, A. Protopopov, W.H. Wong, L. Chin, K.L. Ligon, R.A. DePinho, FoxOs cooperatively regulate diverse pathways governing neural stem cell homeostasis, Cell Stem Cell 5 (2009) 540–553. M. Groszer, R. Erickson, D.D. Scripture-Adams, R. Lesche, A. Trumpp, J.A. Zack, H.I. Kornblum, X. Liu, H. Wu, Negative regulation of neural stem/progenitor cell proliferation by the Pten tumor suppressor gene in vivo, Science 294 (2001) 2186–2189. M. Groszer, R. Erickson, D.D. Scripture-Adams, J.D. Dougherty, J. Le Belle, J.A. Zack, D.H. Geschwind, X. Liu, H.I. Kornblum, H. Wu, PTEN negatively regulates neural stem cell self-renewal by modulating G(0)–G(1) cell cycle entry, Proc. Natl. Acad. Sci. U. S. A. 103 (2006) 111–116. W.H. Cui, K. Matsuno, K. Iwata, M. Ibi, M. Matsumoto, J. Zhang, K. Zhu, M. Katsuyama, N.J. Torok, C. Yabe-Nishimura, NOX1/nicotinamide adenine dinucleotide phosphate, reduced form (NADPH) oxidase promotes proliferation of stellate cells and aggravates liver fibrosis induced by bile duct ligation, Hepatology 54 (2011) 949–958. H. Murata, Y. Ihara, H. Nakamura, J. Yodoi, K. Sumikawa, T. Kondo, Glutaredoxin exerts an antiapoptotic effect by regulating the redox state of Akt, J. Biol. Chem. 278 (2003) 50226–50233. N.R. Leslie, D. Bennett, Y.E. Lindsay, H. Stewart, A. Gray, C.P. Downes, Redox regulation of PI 3-kinase signalling via inactivation of PTEN, EMBO J. 22 (2003) 5501–5510. A. Petry, M. Weitnauer, A. Gorlach, Receptor activation of NADPH oxidases, Antioxid. Redox Signal. 13 (2010) 467–487. E. Kokovay, Y. Wang, G. Kusek, R. Wurster, P. Lederman, N. Lowry, Q. Shen, S. Temple, VCAM1 is essential to maintain the structure of the SVZ niche and acts as an environmental sensor to regulate SVZ lineage progression, Cell Stem Cell 11 (2012) 220–230. E. Topchiy, E. Panzhinskiy, W.S.T. Griffin, S.W. Barger, M. Das, W.M. Zawada, Nox4generated superoxide drives Angiotensin II-induced neural stem cell proliferation, Dev. Neurosci. 35 (2013) 293–305. J. Chao, L. Yang, S. Buch, L. Gao, Angiotensin II increased neuronal stem cell proliferation: role of AT2R, PLos One 8 (2013). N. Anilkumar, R. Weber, M. Zhang, A. Brewer, A.M. Shah, Nox4 and Nox2 NADPH oxidases mediate distinct cellular redox signaling responses to agonist stimulation, Arterioscler. Thromb. Vasc. Biol. 28 (2008) 1347–1354. K.A.M. Kennedy, E.A. Ostrakhovitch, S.D.E. Sandiford, T. Dayarathna, X.J. Xie, E.Y.L. Waese, W.Y. Chang, Q.P. Feng, I.S. Skerjanc, W.L. Stanford, S.S.C. Li, Mammalian numb-interacting protein 1/dual oxidase maturation factor 1 directs neuronal fate in stem cells, J. Biol. Chem. 285 (2010) 17974–17985. D.J. Morre, Preferential inhibition of the plasma membrane NADH oxidase (NOX) activity by diphenyleneiodonium chloride with NADPH as donor, Antioxid. Redox Signal. 4 (2002) 207–212. W.Q. Lu, Y.M. Hu, G. Chen, Z. Chen, H. Zhang, F. Wang, L. Feng, H. Pelicano, H. Wang, M.J. Keating, J.S. Liu, W. McKeehan, H.M. Wang, Y.D. Luo, P. Huang, Novel role of NOX in supporting aerobic glycolysis in cancer cells with mitochondrial dysfunction and as a potential target for cancer therapy, PLoS Biol. 10 (2012). R. Nusse, C. Fuerer, W. Ching, K. Harnish, C. Logan, A. Zeng, D. ten Berge, Y. Kalani, Wnt signaling and stem cell control, Cold Spring Harb. Symp. Quant. Biol. 73 (2008) 59–66. U. Koch, R. Lehal, F. Radtke, Stem cells living with a Notch, Development 140 (2013) 689–704. H. Motohashi, M. Yamamoto, Nrf2-Keap1 defines a physiologically important stress response mechanism, Trends Mol. Med. 10 (2004) 549–557. S.B. Cullinan, J.D. Gordan, J.O. Jin, J.W. Harper, J.A. Diehl, The Keap1-BTB protein is an adaptor that bridges Nrf2 to a Cul3-based E3 ligase: oxidative stress sensing by a Cul3-Keap1 ligase, Mol. Cell. Biol. 24 (2004) 8477–8486. A.K. Jain, D.A. Bloom, A.K. Jaiswal, Nuclear import and export signals in control of Nrf2, J. Biol. Chem. 280 (2005) 29158–29168. W.G. Li, A.N. Kong, Molecular mechanisms of Nrf2-mediated antioxidant response, Mol. Carcinog. 48 (2009) 91–104. K. Itoh, T. Chiba, S. Takahashi, T. Ishii, K. Igarashi, Y. Katoh, T. Oyake, N. Hayashi, K. Satoh, I. Hatayama, M. Yamamoto, Y. Nabeshima, An Nrf2 small Maf heterodimer mediates the induction of phase II detoxifying enzyme genes through antioxidant response elements, Biochem. Biophys. Res. Commun. 236 (1997) 313–322. A.K. Jaiswal, Nrf2 signaling in coordinated activation of antioxidant gene expression, Free Radic. Biol. Med. 36 (2004) 1199–1207. J.Y. Im, K.W. Lee, J.M. Woo, E. Junn, M.M. Mouradian, DJ-1 induces thioredoxin 1 expression through the Nrf2 pathway, Hum. Mol. Genet. 21 (2012) 3013–3024. N. Wakabayashi, S. Shin, S.L. Slocum, E.S. Agoston, J. Wakabayashi, M.K. Kwak, V. Misra, S. Biswal, M. Yamamoto, T.W. Kensler, Regulation of Notch1 signaling by Nrf2: implications for tissue regeneration, Sci. Signal. 3 (2010). J.H. Kim, R.K. Thimmulappa, V. Kumar, W.C. Cui, S. Kumar, P. Kombairaju, H. Zhang, J. Margolick, W. Matsui, T. Macvittie, S.V. Malhotra, S. Biswal, NRF2-mediated Notch pathway activation enhances hematopoietic reconstitution following myelosuppressive radiation, J. Clin. Investig. 124 (2014) 730–741. M.K. Paul, B. Bisht, D.O. Darmawan, R. Chiou, V.L. Ha, W.D. Wallace, A.T. Chon, A.E. Hegab, T. Grogan, D.A. Elashoff, J.A. Alva-Ornelas, B.N. Gomperts, Dynamic changes in intracellular ROS levels regulate airway basal stem cell homeostasis through Nrf2-dependent Notch signaling, Cell Stem Cell 15 (2014) 199–214.

[122] N. Wakabayashi, J.J. Skoko, D.V. Chartoumpekis, S. Kimura, S.L. Slocum, K. Noda, D.L. Palliyaguru, M. Fujimuro, P.A. Boley, Y. Tanaka, N. Shigemura, S. Biswal, M. Yamamoto, T.W. Kensler, Notch-Nrf2 axis: regulation of Nrf2 gene expression and cytoprotection by Notch signaling, Mol. Cell. Biol. 34 (2014) 653–663. [123] V. Karkkainen, Y. Pomeshchik, E. Savchenko, H. Dhungana, A. Kurronen, S. Lehtonen, N. Naumenko, P. Tavi, A.L. Levonen, M. Yamamoto, T. Malm, J. Magga, K.M. Kanninen, J. Koistinaho, Nrf2 regulates neurogenesis and protects neural progenitor cells against a beta toxicity, Stem Cells 32 (2014) 1904–1916. [124] E.M. Hanschmann, J.R. Godoy, C. Berndt, C. Hudemann, C.H. Lillig, Thioredoxins glutaredoxins, and peroxiredoxins-molecular mechanisms and health significance: from cofactors to antioxidants to redox signaling, Antioxid. Redox Signal. 19 (2013) 1539–1605. [125] H. Sies, Role of metabolic H2O2 generation, J. Biol. Chem. 289 (2014) 8735–8741. [126] M. Matsui, M. Oshima, H. Oshima, K. Takaku, T. Maruyama, J. Yodoi, M.M. Taketo, Early embryonic lethality caused by targeted disruption of the mouse thioredoxin gene, Dev. Biol. 178 (1996) 179–185. [127] Y. Guo, L. Einhorn, M. Kelley, K. Hirota, J. Yodoi, R. Reinbold, H. Scholer, H. Ramsey, R. Hromas, Redox regulation of the embryonic stem cell transcription factor Oct-4 by thioredoxin, Stem Cells 22 (2004) 259–264. [128] F. Zhou, P.P. Liu, G.Y. Ying, X.D. Zhu, H. Shen, G. Chen, Effects of thioredoxin-1 on neurogenesis after brain ischemia/reperfusion injury, CNS Neurosci. Ther. 19 (2013) 204–205. [129] L. Tian, H. Nie, Y. Zhang, Y. Chen, Z.W. Peng, M. Cai, H.D. Wei, P. Qin, H.L. Dong, L.Z. Xiong, Recombinant human thioredoxin-1 promotes neurogenesis and facilitates cognitive recovery following cerebral ischemia in mice, Neuropharmacology 77 (2014) 453–464. [130] E.J. Meuillet, D. Mahadevan, M. Berggren, A. Coon, G. Powis, Thioredoxin-1 binds to the C2 domain of PTEN inhibiting PTEN's lipid phosphatase activity and membrane binding: a mechanism for the functional loss of PTEN's tumor suppressor activity, Arch. Biochem. Biophys. 429 (2004) 123–133. [131] M. Groszer, R. Erickson, D.D. Scripture-Adams, R. Lesche, A. Trumpp, J.A. Zack, H.I. Kornblum, X. Liu, H. Wu, Negative regulation of neural stem/progenitor cell proliferation by the Pten tumor suppressor gene in vivo, Science 294 (2001) 2186–2189. [132] J. Soerensen, C. Jakupoglu, H. Beck, H. Forster, J. Schmidt, W. Schmahl, U. Schweizer, M. Conrad, M. Brielmeier, The role of thioredoxin reductases in brain development, PLos One 3 (2008). [133] C.H. Lillig, C. Berndt, Glutaredoxins in thiol/disulfide exchange, Antioxid. Redox Signal. 18 (2013) 1654–1665. [134] L. Brautigam, L.D.E. Jensen, G. Poschmann, S. Nystrom, S. Bannenberg, K. Dreij, K. Lepka, T. Prozorovski, S.J. Montano, O. Aktas, P. Uhlen, K. Stuhler, Y.H. Cao, A. Holmgren, C. Berndt, Glutaredoxin regulates vascular development by reversible glutathionylation of sirtuin 1, Proc. Natl. Acad. Sci. U. S. A. 110 (2013) 20057–20062. [135] A.V. Budanov, A.A. Sablina, E. Feinstein, E.V. Koonin, P.M. Chumakov, Regeneration of peroxiredoxins by p53-regulated sestrins, homologs of bacterial AhpD, Science 304 (2004) 596–600. [136] S.G. Rhee, S.W. Kang, W. Jeong, T.S. Chang, K.S. Yang, H.A. Woo, Intracellular messenger function of hydrogen peroxide and its regulation by peroxiredoxins, Curr. Opin. Cell Biol. 17 (2005) 183–189. [137] T.S. Chang, W. Jeong, S.Y. Choi, S.Q. Yu, S.W. Kang, S.G. Rhee, Regulation of peroxiredoxin I activity by Cdc2-mediated phosphorylation, J. Biol. Chem. 277 (2002) 25370–25376. [138] S.U. Kim, Y.H. Park, J.M. Kim, H.N. Sun, I.S. Song, S.M. Huang, S.H. Lee, J.I. Chae, S. Hong, S. Choi, S.C. Choi, T.H. Lee, S.W. Kang, S.G. Rhee, K.T. Chang, S.H. Lee, D.Y. Yu, D.S. Lee, Dominant role of peroxiredoxin/JNK axis in stemness regulation during neurogenesis from embryonic stem cells, Stem Cells 32 (2014) 998–1011. [139] Y. Yan, P. Sabharwal, M. Rao, S. Sockanathan, The antioxidant enzyme Prdx1 controls neuronal differentiation by thiol-redox-dependent activation of GDE2, Cell 138 (2009) 1209–1221. [140] M. Rao, S. Sockanathan, Transmembrane protein GDE2 induces motor neuron differentiation in vivo, Science 309 (2005) 2212–2215. [141] M. Rodriguez, J. Choi, S. Park, S. Sockanathan, Gde2 regulates cortical neuronal identity by controlling the timing of cortical progenitor differentiation, Development 139 (2012) 3870–3879. [142] B.J. Laughner, P.C. Sehnke, R.J. Ferl, A novel nuclear member of the thioredoxin superfamily, Plant Physiol. 118 (1998) 987–996. [143] Y. Funato, T. Michiue, M. Asashima, H. Miki, The thioredoxin-related redoxregulating protein nucleoredoxin inhibits Wnt-beta-catenin signalling through dishevelled, Nat. Cell Biol. 8 (2006) 501-U135. [144] Y. Funato, T. Michiue, T. Terabayashi, A. Yukita, H. Danno, M. Asashima, H. Miki, Nucleoredoxin regulates the Wnt/planar cell polarity pathway in Xenopus, Genes to Cells 13 (2008) 965–975. [145] Y. Funato, T. Terabayashi, R. Sakamoto, D. Okuzaki, H. Ichise, H. Nojima, N. Yoshida, H. Miki, Nucleoredoxin sustains Wnt/beta-catenin signaling by retaining a pool of inactive dishevelled protein, Curr. Biol. 20 (2010) 1945–1952. [146] Y. Hirabayashi, Y. Itoh, H. Tabata, K. Nakajima, T. Akiyama, N. Masuyama, Y. Gotoh, The Wnt/beta-catenin pathway directs neuronal differentiation of cortical neural precursor cells, Development 131 (2004) 2791–2801. [147] M.Y. Kalani, S.H. Cheshier, B.J. Cord, S.R. Bababeygy, H. Vogel, I.L. Weissman, T.D. Palmer, R. Nusse, Wnt-mediated self-renewal of neural stem/progenitor cells, Proc. Natl. Acad. Sci. U. S. A. 105 (2008) 16970–16975. [148] Q.H. Qu, G.Q. Sun, W.W. Li, S. Yang, P. Ye, C.N.A. Zhao, R.T. Yu, F.H. Gage, R.M. Evans, Y.H. Shi, Orphan nuclear receptor TLX activates Wnt/beta-catenin signalling to stimulate neural stem cell proliferation and self-renewal, Nat. Cell Biol. 12 (2010) 31-U80.

T. Prozorovski et al. / Biochimica et Biophysica Acta 1850 (2015) 1543–1554 [149] N. Coant, S. Ben Mkaddem, E. Pedruzzi, C. Guichard, X. Treton, R. Ducroc, J.N. Freund, D. Cazals-Hatem, Y. Bouhnik, P.L. Woerther, D. Skurnik, A. Grodet, M. Fay, D. Biard, T. Lesuffleur, C. Deffert, R. Moreau, A. Groyer, K.H. Krause, F. Daniel, E. Ogier-Denis, NADPH oxidase 1 modulates WNT and NOTCH1 signaling to control the fate of proliferative progenitor cells in the colon, Mol. Cell. Biol. 30 (2010) 2636–2650. [150] S. Kajla, A.S. Mondol, A. Nagasawa, Y.G. Zhang, M. Kato, K. Matsuno, C. YabeNishimura, T. Kamata, A crucial role for Nox 1 in redox-dependent regulation of Wnt-beta-catenin signaling, FASEB J. 26 (2012) 2049–2059. [151] S.Y. Shin, C.G. Kim, E.H. Jho, M.S. Rho, Y.S. Kim, Y.H. Kim, Y.H. Lee, Hydrogen peroxide negatively modulates Wnt signaling through downregulation of beta-catenin, Cancer Lett. 212 (2004) 225–231. [152] S.Y. Shin, B.R. Chin, Y.H. Lee, J.H. Kim, Involvement of glycogen synthase kinase-3 beta in hydrogen peroxide-induced suppression of Tcf/Lef-dependent transcriptional activity, Cell. Signal. 18 (2006) 601–607. [153] G. Unden, J. Bongaerts, Alternative respiratory pathways of Escherichia coli: energetics and transcriptional regulation in response to electron acceptors, Biochim. Biophys. Acta Bioenerg. 1320 (1997) 217–234. [154] M. Mimaki, X.N. Wang, M. McKenzie, D.R. Thorburn, M.T. Ryan, Understanding mitochondrial complex I assembly in health and disease, Biochim. Biophys. Acta Bioenerg. 1817 (2012) 851–862. [155] L.R. Stein, S. Imai, Specific ablation of Nampt in adult neural stem cells recapitulates their functional defects during aging, EMBO J. 33 (2014) 1321–1340. [156] G. Chinnadurai, Transcriptional regulation by C-terminal binding proteins, Int. J. Biochem. Cell Biol. 39 (2007) 1593–1607. [157] U. Schaeper, J.M. Boyd, S. Verma, E. Uhlmann, T. Subramanian, G. Chinadurai, Molecular-cloning and characterization of a cellular phosphoprotein that interacts with a conserved C-terminal domain of adenovirus E1A involved in negative modulation of oncogenic transformation, Proc. Natl. Acad. Sci. U. S. A. 92 (1995) 10467–10471. [158] V. Kumar, J.E. Carlson, K.A. Ohgi, T.A. Edwards, D.W. Rose, C.R. Escalante, M.G. Rosenfeld, A.K. Aggarwal, Transcription corepressor CtBP is an NAD(+)-regulated dehydrogenase, Mol. Cell 10 (2002) 857–869. [159] Q.H. Zhang, D.W. Piston, R.H. Goodman, Regulation of corepressor function by nuclear NADH, Science 295 (2002) 1895–1897. [160] K.G.R. Quinlan, A. Verger, A. Kwok, S.H.Y. Lee, J. Perdomo, M. Nardini, M. Bolognesi, M. Crossley, Role of the C-terminal binding protein PXDLS motif binding cleft in protein interactions and transcriptional repression, Mol. Cell. Biol. 26 (2006) 8202–8213. [161] A. Verger, K.G. Quinlan, L.A. Crofts, S. Spano, D. Corda, E.P. Kable, F. Braet, M. Crossley, Mechanisms directing the nuclear localization of the CtBP family proteins, Mol. Cell. Biol. 26 (2006) 4882–4894. [162] L.J. Di, J.S. Byun, M.M. Wong, C. Wakano, T. Taylor, S. Bilke, S. Baek, K. Hunter, H. Yang, M. Lee, C. Zvosec, G. Khramtsova, F. Cheng, C.M. Perou, C.R. Miller, R. Raab, O.I. Olopade, K. Gardner, Genome-wide profiles of CtBP link metabolism with genome stability and epithelial reprogramming in breast cancer, Nat. Commun. 4 (2013). [163] J.M. Dias, S. Ilkhanizadeh, E. Karaca, J.K. Duckworth, V. Lundin, M.G. Rosenfeld, J. Ericson, O. Hermanson, A.I. Teixeira, CtBPs sense microenvironmental oxygen levels to regulate neural stem cell state, Cell Rep. 8 (2014) 665–670. [164] Z.H. Xie, Y.F. Chen, Z.F. Li, G. Bai, Y. Zhu, R. Yan, F.Z. Tan, Y.G. Chen, F. Guillemot, L. Li, N.H. Jing, Smad6 promotes neuronal differentiation in the intermediate zone of the dorsal neural tube by inhibition of the Wnt/beta-catenin pathway, Proc. Natl. Acad. Sci. U. S. A. 108 (2011) 12119–12124. [165] J.M. Dias, S. Ilkhanizadeh, E. Karaca, J.K. Duckworth, V. Lundin, M.G. Rosenfeld, J. Ericson, O. Hermanson, A.I. Teixeira, CtBPs sense microenvironmental oxygen levels to regulate neural stem cell state, Cell Rep. 8 (2014) 665–670. [166] J.D. Hildebrand, P. Soriano, Overlapping and unique roles for C-terminal binding protein 1 (CtBP1) and CtBP2 during mouse development, Mol. Cell. Biol. 22 (2002) 5296–5307. [167] T. Valenta, J. Lukas, V. Korinek, HMG box transcription factor TCF-4's interaction with CtBP1 controls the expression of the Wnt target Axin2/Conductin in human embryonic kidney cells, Nucleic Acids Res. 31 (2003) 2369–2380. [168] M. Brannon, J.D. Brown, R. Bates, D. Kimelman, R.T. Moon, XCtBP is a XTcf-3 corepressor with roles throughout Xenopus development, Development 126 (1999) 3159–3170. [169] M. Fang, J. Li, T. Blauwkamp, C. Bhambhani, N. Campbell, K.M. Cadigan, C-terminalbinding protein directly activates and represses Wnt transcriptional targets in Drosophila, EMBO J. 25 (2006) 2735–2745. [170] F. Hamada, M. Bienz, The APC tumor suppressor binds to C-terminal binding protein to divert nuclear beta-catenin from TCF, Dev. Cell 7 (2004) 677–685. [171] F.R. Althaus, Poly(ADP-ribose) and chromatin organization in DNA excision repair, Br. J. Cancer 56 (1987) 176. [172] M. Masson, C. Niedergang, V. Schreiber, S. Muller, J. Menissier-de Murcia, G. de Murcia, XRCC1 is specifically associated with poly(ADP-ribose) polymerase and negatively regulates its activity following DNA damage, Mol. Cell. Biol. 18 (1998) 3563–3571. [173] W.L. Kraus, J.T. Lis, PARP goes transcription, Cell 113 (2003) 677–683. [174] M.Y. Kim, S. Mauro, N. Gevry, J.T. Lis, W.L. Kraus, NAD(+)-dependent modulation of chromatin structure and transcription by nucleosome binding properties of PARP-1, Cell 119 (2004) 803–814. [175] B.G. Ju, D. Solum, E.J. Song, K.J. Lee, D.W. Rose, C.K. Glass, M.G. Rosenfeld, Activating the PARP-1 sensor component of the Groucho/TLE1 corepressor complex mediates a CaMKinase II delta-dependent neurogenic gene activation pathway, Cell 119 (2004) 815–829. [176] M. Kanai, K. Hanashiro, S.H. Kim, S. Hanai, A.H. Boulares, M. Miwa, K. Fukasawa, Inhibition of Crm1-p53 interaction and nuclear export of p53 by poly(ADPribosyl)ation, Nat. Cell Biol. 9 (2007) 1175–1183.


[177] H. Ogino, T. Nozaki, A. Gunji, M. Maeda, H. Suzuki, T. Ohta, Y. Murakami, H. Nakagama, T. Sugimura, M. Masutani, Loss of Parp-1 affects gene expression profile in a genome-wide manner in ES cells and liver cells, BMC Genomics 8 (2007). [178] F.R. Gao, S.W. Kwon, Y.M. Zhao, Y. Jin, PARP1 poly(ADP-ribosyl)ates Sox2 to control Sox2 protein levels and FGF4 expression during embryonic stem cell differentiation, J. Biol. Chem. 284 (2009) 22263–22273. [179] J.M. Plane, S.K. Grossenbacher, W.B. Deng, PARP-1 deletion promotes subventricular zone neural stem cells toward a glial fate, J. Neurosci. Res. 90 (2012) 1489–1506. [180] M.C. Haigis, D.A. Sinclair, Mammalian sirtuins: biological insights and disease relevance, Annu. Rev. Pathol. Mech. Dis. 5 (2010) 253–295. [181] M. Kaeberlein, M. McVey, L. Guarente, The SIR2/3/4 complex and SIR2 alone promote longevity in Saccharomyces cerevisiae by two different mechanisms, Genes Dev. 13 (1999) 2570–2580. [182] S.J. Lin, M. Kaeberlein, A.A. Andalis, L.A. Sturtz, P.A. Defossez, V.C. Culotta, G.R. Fink, L. Guarente, Calorie restriction extends Saccharomyces cerevisiae lifespan by increasing respiration, Nature 418 (2002) 344–348. [183] S.J. Lin, E. Ford, M. Haigis, G. Liszt, L. Guarente, Calorie restriction extends yeast life span by lowering the level of NADH, Genes Dev. 18 (2004) 12–16. [184] J.M. Denu, Linking chromatin function with metabolic networks: Sir2 family of NAD(+)-dependent deacetylases, Trends Biochem. Sci. 28 (2003) 41–48. [185] E. Koltai, Z. Szabo, M. Atalay, I. Boldogh, H. Naito, S. Goto, C. Nyakas, Z. Radak, Exercise alters SIRT1, SIRT6 NAD and NAMPT levels in skeletal muscle of aged rats, Mech. Ageing Dev. 131 (2010) 21–28. [186] S. Caito, S. Rajendrasozhan, S. Cook, S. Chung, H.W. Yao, A.E. Friedman, P.S. Brookes, I. Rahman, SIRT1 is a redox-sensitive deacetylase that is post-translationally modified by oxidants and carbonyl stress, FASEB J. 24 (2010) 3145–3159. [187] N. Braidy, G.J. Guillemin, H. Mansour, T. Chan-Ling, A. Poljak, R. Grant, Age related changes in NAD plus metabolism oxidative stress and Sirt1 activity in Wistar rats, PLos One 6 (2011). [188] Q. Zhang, S.Y. Wang, C. Fleuriel, D. Leprince, J.V. Rocheleau, D.W. Piston, R.H. Goodman, Metabolic regulation of SIRT1 transcription via a HIC1:CtBP corepressor complex, Proc. Natl. Acad. Sci. U. S. A. 104 (2007) 829–833. [189] C. Berndt, C.H. Lillig, L. Flohe, Redox regulation by glutathione needs enzymes, Front. Pharmacol. 5 (2014) 168. [190] M.D. Kornberg, N. Sen, M.R. Hara, K.R. Juluri, J.V.K. Nguyen, A.M. Snowman, L. Law, L.D. Hester, S.H. Snyder, GAPDH mediates nitrosylation of nuclear proteins, Nat. Cell Biol. 12 (2010) 1094-1U89. [191] R.S. Zee, C.B. Yoo, D.R. Pimentel, D.H. Perlman, J.R. Burgoyne, X.Y. Hou, M.E. Mccomb, C.E. Costello, R.A. Cohen, M.M. Bachschmid, Redox regulation of sirtuin-1 by S-glutathiolation, Antioxid. Redox Signal. 13 (2010) 1023–1032. [192] D. Shao, J.L. Fry, J.Y. Han, X.Y. Hou, D.R. Pimentel, R. Matsui, R.A. Cohen, M.M. Bachschmid, A redox-resistant sirtuin-1 mutant protects against hepatic metabolic and oxidant stress, J. Biol. Chem. 289 (2014) 7293–7306. [193] B.R. Webster, Z.P. Lu, M.N. Sack, I. Scott, The role of sirtuins in modulating redox stressors, Free Radic. Biol. Med. 52 (2012) 281–290. [194] Z. Radak, E. Koltai, A.W. Taylor, M. Higuchi, S. Kumagai, H. Ohno, S. Goto, I. Boldogh, Redox-regulating sirtuins in aging, caloric restriction, and exercise, Free Radic. Biol. Med. 58 (2013) 87–97. [195] R.R. Alcendor, S.M. Gao, P.Y. Zhai, D. Zablocki, E. Holle, X.Z. Yu, B. Tian, T. Wagner, S.F. Vatner, J. Sadoshima, Sirt1 regulates aging and resistance to oxidative stress in the heart, Circ. Res. 100 (2007) 1512–1521. [196] F. Wang, M. Nguyen, F.X.F. Qin, Q. Tong, SIRT2 deacetylates FOXO3a in response to oxidative stress and caloric restriction, Aging Cell 6 (2007) 505–514. [197] A. Brunet, L.B. Sweeney, J.F. Sturgill, K.F. Chua, P.L. Greer, Y. Lin, H. Tran, S.E. Ross, R. Mostoslavsky, H.Y. Cohen, L.S. Hu, H.L. Cheng, M.P. Jedrychowski, S.P. Gygi, D.A. Sinclair, F.W. Alt, M.E. Greenberg, Stress-dependent regulation of FOXO transcription factors by the SIRT1 deacetylase, Science 303 (2004) 2011–2015. [198] N.R. Sundaresan, M. Gupta, G. Kim, S.B. Rajamohan, A. Isbatan, M.P. Gupta, Sirt3 blocks the cardiac hypertrophic response by augmenting Foxo3a-dependent antioxidant defense mechanisms in mice, J. Clin. Investig. 119 (2009) 2758–2771. [199] H. Vaziri, S.K. Dessain, E.E. Ng, S.I. Imai, R.A. Frye, T.K. Pandita, L. Guarente, R.A. Weinberg, hSIR2(SIRT1) functions as an NAD-dependent p53 deacetylase, Cell 107 (2001) 149–159. [200] S. Kume, M. Haneda, K. Kanasaki, T. Sugimoto, S.I. Araki, M. Isono, K. Isshiki, T. Uzu, A. Kashiwagi, D. Koya, Silent information regulator 2 (SIRT1) attenuates oxidative stress-induced mesangial cell apoptosis via p53 deacetylation, Free Radic. Biol. Med. 40 (2006) 2175–2182. [201] H.L. Cheng, R. Mostoslavsky, S. Saito, J.P. Manis, Y. Gu, P. Patel, R. Bronson, E. Appella, F.W. Alt, K.F. Chua, Developmental defects and p53 hyperacetylation in Sir2 homolog (SIRT1)-deficient mice, Proc. Natl. Acad. Sci. U. S. A. 100 (2003) 10794–10799. [202] M.W. McBurney, X. Yang, K. Jardine, M. Hixon, K. Boekelheide, J.R. Webb, P.M. Lansdorp, M. Lemieux, The mammalian SIR2alpha protein has a role in embryogenesis and gametogenesis, Mol. Cell. Biol. 23 (2003) 38–54. [203] J. Sakamoto, T. Miura, K. Shimamoto, Y. Horio, Predominant expression of Sir2alpha, an NAD-dependent histone deacetylase, in the embryonic mouse heart and brain, FEBS Lett. 556 (2004) 281–286. [204] R.H. Wang, K. Sengupta, C. Li, H.S. Kim, L. Cao, C. Xiao, S. Kim, X. Xu, Y. Zheng, B. Chilton, R. Jia, Z.M. Zheng, E. Appella, X.W. Wang, T. Ried, C.X. Deng, Impaired DNA damage response, genome instability, and tumorigenesis in SIRT1 mutant mice, Cancer Cell 14 (2008) 312–323. [205] S. Michan, Y. Li, M.M. Chou, E. Parrella, H. Ge, J.M. Long, J.S. Allard, K. Lewis, M. Miller, W. Xu, R.F. Mervis, J. Chen, K.I. Guerin, L.E. Smith, M.W. McBurney, D.A. Sinclair, M. Baudry, R. de Cabo, V.D. Longo, SIRT1 is essential for normal cognitive function and synaptic plasticity, J. Neurosci. 30 (2010) 9695–9707.


T. Prozorovski et al. / Biochimica et Biophysica Acta 1850 (2015) 1543–1554

[206] S. Ichi, V. Boshnjaku, Y.W. Shen, B. Mania-Farnell, S. Ahlgren, S. Sapru, N. Mansukhani, D.G. McLone, T. Tomita, C.S. Mayanil, Role of Pax3 acetylation in the regulation of Hes1 and Neurog2, Mol. Biol. Cell 22 (2011) 503–512. [207] V. Calvanese, E. Lara, B. Suarez-Alvarez, D.R. Abu, M. Vazquez-Chantada, M.L. Martinez-Chantar, N. Embade, P. Lopez-Nieva, A. Horrillo, A. Hmadcha, B. Soria, D. Piazzolla, D. Herranz, M. Serrano, J.M. Mato, P.W. Andrews, C. Lopez-Larrea, M. Esteller, M.F. Fraga, Sirtuin 1 regulation of developmental genes during differentiation of stem cells, Proc. Natl. Acad. Sci. U. S. A. 107 (2010) 13736–13741. [208] S. Hisahara, S. Chiba, H. Matsumoto, M. Tanno, H. Yagi, S. Shimohama, M. Sato, Y. Horio, Histone deacetylase SIRT1 modulates neuronal differentiation by its nuclear translocation, Proc. Natl. Acad. Sci. U. S. A. 105 (2008) 15599–15604. [209] L. Tiberi, J. van den Ameele, J. Dimidschstein, J. Piccirilli, D. Gall, A. Herpoel, A. Bilheu, J. Bonnefont, M. Iacovino, M. Kyba, T. Bouschet, P. Vanderhaeghen, BCL6

controls neurogenesis through Sirt1-dependent epigenetic repression of selective Notch targets, Nat. Neurosci. 15 (2012) 1627-+. [210] D.W. Huang, B.T. Sherman, R.A. Lempicki, Systematic and integrative analysis of large gene lists using DAVID bioinformatics resources, Nat. Protoc. 4 (2009) 44–57. [211] P. Oberdoerffer, S. Michan, M. McVay, R. Mostoslavsky, J. Vann, S.K. Park, A. Hartlerode, J. Stegmuller, A. Hafner, P. Loerch, S.M. Wright, K.D. Mills, A. Bonni, B.A. Yankner, R. Scully, T.A. Prolla, F.W. Alt, D.A. Sinclair, SIRT1 redistribution on chromatin promotes genomic stability but alters gene expression during aging, Cell 135 (2008) 907–918. [212] M. Tanno, J. Sakamoto, T. Miura, K. Shimamoto, Y. Horio, Nucleocytoplasmic shuttling of the NAD+-dependent histone deacetylase SIRT1, J. Biol. Chem. 282 (2007) 6823–6832.