Given the information described above, zinc deficiency‐treated male rat model was constructed in order to determine the potential value of FoxO3a and p27 kip1 . Through this model system, we investigated the expression and distribution of FoxO3a in brain hippocampus exposure chronic zinc deficiency. In addition, we established an in vitro model to further illuminate the effects of FoxO3a on cell proliferation and p27 kip1 expression.

Forkhead box O3 (FOXO3a) is a key transcription factor that regulates a wide range of cellular events, including metabolism, differentiation, proliferation, survival, stress resistance, and aging (Weigel et al . 1989 ; Accili and Arden 2004 ; Barthelemy et al . 2004 ). FOXO3a plays a pivotal role in neurobiology, and is critically implicated in the development of various neurological diseases. Under stress conditions, FOXO3a may exert neuroprotective role through inducing stress resistance, thus preventing the onset of motor neuron diseases (Mojsilovic‐Petrovic et al . 2009 ). However, during many pathological conditions, FOXO3a facilitates neuronal death via the transcription of pro‐apoptotic proteins, such as p27 kip1 , p21 cip1 and bim (Li et al . 2009 ; Xu et al . 2014 ). In addition to neurons, FOXO3a also elicits an important role in the regulation of NSCs physiology. For example, Wu et al . ( 2009 ) reported that CXCL12 stimulated the proliferation of NSCs through Akt‐mediated phosphorylation and inactivation of FOXO3a. FOXO3a‐deficient mice exhibited enhanced proliferation and renewal of NSCs during early postnatal life (Paik et al . 2009 ; Renault et al . 2009 ). In mammalian neurogenesis, peripheral nerve injury induces down‐regulation of FoxO3a and p27 kip1 in rat dorsal root ganglia (Wang et al . 2009 ). p27 kip1 serves as a target gene for FoxO3a, which regulates the transcription of p27 kip1 gene expression (Dijkers et al . 2000 ; Medema et al . 2000 ). Active FoxO3a control the cell cycle progression through activating p27 kip1 expression, acting as cyclin‐dependent‐kinase (CDK) inhibitor, which resulted in cell cycle arrest in the G1 phase of cell cycle (Medema et al . 2000 ). However, whether FoxO3a exerts a similar effect in zinc deficiency‐induced aberrant NSCs regulation remains largely obscure.

Mounting evidence indicates that the hippocampus plays an integral role in cognitive functions, such as memory formation and spatial processing as well as mood regulation (Squire et al . 1992 ; Ekstrom et al . 2003 ). Hippocampal neurons initiate long‐term potential, serving as the molecular basis underlying declarative memory and other cognitive functions (Vila‐Luna et al . 2012 ). In addition, recent findings pointed to a crucial involvement of hippocampal neurogenesis in the regulation of cognitive performance (Monje and Dietrich 2012 ). In the adult mammalian brain, continuous neurogenesis occurred in two discrete areas including the subgranular zone (SGZ) of the dentate gyrus (DG) in the hippocampus and the subventricular zone of the lateral ventricles, in which neurogenesis persists throughout life (Eriksson et al . 1998 ; Gage 2000 ; Kernie and Parent 2010 ). Hippocampal neural stem cells (NSCs) resided in the SGZ are significant in the maintenance of normal cognition (Deng et al . 2010 ). SGZ NSCs generate excitatory neurons that integrate into the DG, whose processing is important for certain types of hippocampus‐dependent learning and memory (Abrous et al . 2005 ; Leuner et al . 2006 ). Inhibition of neurogenesis impairs hippocampus‐dependent memory function (Winocur et al . 2006 ). Moreover, nutritional and environmental factors may trigger the alteration of NSC physiology, leading to cognitive and memory deficits. For example, chronic manganese exposure results in sustained abnormality of neurogenesis in the hippocampal DG of mice, contributing to manganese‐induced cognitive disorders (Wang et al . 2012 ). Besides, Dioxins have been reported to impair spatial learning and memory in rats through the attenuation of neurogenesis (Markowski et al . 2002 ). Interestingly, the hippocampus is one of the richest zinc‐containing areas in the brain, and cheletable zinc is enriched inside the synaptic vesicle of mossy fiber terminals of the DG and CA3 areas (Perez‐Clausell and Danscher 1985 ; Frederickson et al . 2005 ). Therefore, a potential involvement of zinc ion in the regulation of hippocampal neurogenesis is a matter of considerable concern (Suh et al . 2009 ). Indeed, Corniola et al . ( 2008 ) reported that zinc deficiency led to impaired proliferation and enhanced apoptosis of NSCs. However, the molecular mechanisms underlying zinc deficiency‐induced NSC alterations remain largely elusive.

Zinc is one of the most abundant essential elements in humans and animals for normal brain development. Dietary zinc deficiency brings about abnormal change in the central nervous system and leads to behavioral abnormalities. For example, zinc deprivation during prenatal and postnatal results in learning and memory impairment (Halas et al . 1983 , 1986 ). In animals, zinc deficiency leads to behavioral changes including reduced exploratory activity compared to the pair‐fed control, anorexia, lethargy, and anxiety (Caldwell et al . 1970 ; Golub et al . 1995 ; Evans et al . 2004 ; Takeda et al . 2008 ). Moreover, zinc deficiency in adolescents impaired growth and development, motor functioning and cognitive function (Black 2003a , b ; de Moura et al . 2013 ). Attention‐deficit/hyperactivity disorder was tightly associated with zinc‐deprived school‐age children (Toren et al . 1996 ). The evidence described above suggests that zinc plays a pivotal role in neurodevelopment and neurobehavior.

The number of FoxO3a positive cells in the hippocampus was counted at 40× magnifications. Three separate hippocampus regions were examined for each section. Every three section was analyzed to count the total number of FoxO3a/p27 kip1 positive cells per square millimeter. Cells double labeled for FoxO3a/p27 kip1 and nestin in the experiment were quantified. To identify the proportion of nestin‐positive cells expressing FoxO3a/p27 kip1 , a minimum of 200 cells expressing a cell specific marker were counted. For all animal experiment, we calculated and analyzed at least three sections per animals.

C17.2 cells were transfected with FoxO3a siRNA before exposed to TPEN for 24 h. For cell cycle analysis, cells were trypsinized and fixed with 70% ethanol/PBS at 4°C. After washing with PBS, the cells were incubated with 1 mg/mL RNase A for 30 min at 37°C. Cells were examined by the staining DNA with propidium iodide (50 μg/mL) in PBS with 0.5% Tween‐20, and detected via a Becton–Dickinson Fascine flow cytometry and Cell Quest acquisition and analysis software (Becton–Dickinson, Franklin Lakes, NJ, USA).

C17.2 cells were seeded into 96‐well plates at a density of 3.0 × 10 3 cells/well, and cultured for 24 h to allow for exponential cell growth. After reaching 20–30% confluence, cells were transfected with FoxO3a siRNA, and then exposed to 1000 nM TPEN for various time periods (0, 12, 24, 48, 72 h). Next, 10 μL Cell counting kit‐8 (CCK‐8) solution was applied into each well of the plates and incubated for 2 h in 37°C incubator. The viability of C17.2 cells was assessed by the CCK‐8 assay (Bio‐Tek) according to the manufacturer's instruction. Assay was performed using triplicate independent cell cultures.

Rats were anesthetized via chloral hydrate (10% solution) and perfused pericardially with 0.9% saline followed by 4% paraformaldehyde. The brains were removed, fixed in the same fixative solution for 12 h, and then cryoprotected with 20% sucrose for 2–3 days, followed by 30% sucrose for 2–3 days. Thereafter, the brain tissue was embedded in O.C.T. compound (Sakura, Sapporo, Japan). Then 7‐μm frozen cross‐sections were prepared and stored at −20°C before use. The sections were blocked with blocking solution consisting of 10% donkey serum, 1% (w/v) bovine serum albumin, 0.3% Triton X‐100 and 0.15% Tween‐20 for 2 h at 25°C and incubated overnight at 4°C with rabbit anti‐FoxO3a (1 : 100; Cell Signaling Technology), anti‐p27 kip1 (1 : 100; Sigma). Control sections were same with experimental sections without primary antibody. After washing PBS, the sections were incubated with biotinylated secondary antibody for 30 min at 37°C, and then were color‐reacted with 0.02% diaminobenzidine tetrahydrochloride and 3% H 2 O 2 in PBS. Finally, the sections were dehydrated and coverslipped. All sections were examined on Leica light microscope (Heidelberg, Germany). Cells with strong or moderate brown straining were regarded as positive, whereas cells with weak or no staining were considered negative.

The brain area of hippocampus was dissected out and immediately frozen at −80°C until use. The frozen hippocampus samples were weighed and minced in ice. Then, the samples were homogenized in lysis buffer containing 1% Nonidet P‐40 (NP‐40), 100 mM Tris‐HCl, 0.5 mM ethylene diamine tetraacetic acid, 0.1% sodium dodecyl sulfate, 10 μg/mL aprotinin, 1 μg/mL leupeptin, and 1 mM phenyl methylsulfonyl fluoride (Sigma‐Aldrich, St. Louis, Missouri, USA), 1% Triton X‐100. The lysates were centrifuged at 400 g , 4°C for 20 min to collect the supernatant. Protein samples were quantified using a bicinchoninic acid protein assay kit (Bio‐Rad Laboratories, Hercules, CA, USA). The samples were separated by sodium dodecyl sulfate‐polyacrylamide gel electrophoresis and then transferred to a polyvinylidene difluoride filter membrane. The filters were blocked with 5% non‐fat skim milk in Phosphate Buffered Saline with Tween‐20 (PBST) for 2 h, followed by incubation overnight at 4°C with primary antibodies against rabbit anti‐p27 kip1 (1 : 1000; Santa Cruz Biotechnology), rabbit anti‐foxO3a (1 : 1000; Cell Signaling Technology, Beverly, MA, USA), rabbit anti‐pSer253‐FoxO3a (1 : 1000; Cell Signaling Technology) mouse anti‐β‐actin (1 : 1000; Santa Cruz Biotechnology), rabbit anti‐lamin A/C (1 : 1000; Santa Cruz Biotechnology) mouse anti‐proliferating cell nuclear antigen (PCNA) (1 : 1000; Santa Cruz Biotechnology), rabbit anti‐Cyclin A (1 : 1000; Santa Cruz Biotechnology). Finally, the filters were incubated with secondary antibody using horseradish peroxidase‐conjugated anti‐rabbit or anti‐mouse antibodies (ZhongShan Biotechnology, Beijing, China) at 25°C for 2 h. Immunocomplex was detected by using an enhanced chemiluminescence system (Santa Cruz Biotechnology).

Cells were briefly washed with ice‐cold PBS, and cell pellets were suspended in 200 μL of hypotonic buffer (10 mM HEPES pH 7.9, 10 mM KCl, 0.1 mM EGTA, 1 mM dithiothreitol, 1 mM phenylmethylsulfonyl fluoride, and a protease inhibitor cocktail) and 2.5 μL of 10% NP‐40 for 15 min on ice. The cell suspension was violently shocked on a vortex for 10 s and then stood on the ice about 10 min before centrifugation at 400 g for 5 min. The supernatant was represented as the cytoplasmic extract for western blotting. The pellets were resuspended in high salt nuclear extraction buffer (20 mM HEPES pH 7.9, 400 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1 mM dithiothreitol, 0.5 mM phenylmethylsulfonyl fluoride and a protease inhibitor cocktail) for 1 h on ice with intermittent vortexing. The supernatant containing nuclear proteins was isolated from the reminder at 400 g for 10 min.

C17.2 neural stem cells were cultured in Dulbecco's modified Eagle's medium (Sigma, St Louis, MO, USA) supplemented with 10% fetal bovine serum (HyClone, Logan, UT, USA), 2 mM glutamine, 5% horse serum, and 1% penicillin‐streptomycin at 37°C in a 5% CO 2 humidified incubator. Zinc chelator, N , N , N ’, N ’‐tetrakis (2‐pyridylmethy) ethylenediamine (TPEN; Sigma) was dissolved in dimethylsulfoxide (DMSO) and stored in −20°C before use. Cells were treated with various concentrations of TPEN (0, 200, 500, 1000, 1500, 2000, 2500 nM) or with dimethylsulfoxide (control) for 24 h before subjected to subsequent experiments. C17.2 cells were transiently transfected with Foxo3a siRNA before TPEN exposure. For all experiments, C17.2 cells were plated at a confluence of 50–70%.

EdU staining was conducted using Click‐iT ™ EdU imaging kit according to the manufacturer's instruments (Invitrogen, Carlsbad, CA, USA). Briefly, frozen brain sections were placed at 25°C for 30 min, fixed with 4% paraformaldehyde in phosphate‐buffered saline (PBS) for 10 min and then permeablized with 0.5% Triton X‐100 in PBS for 15 min. After washing twice in PBS, the sections were incubated with a Click‐iT ™ reaction cocktail for 30 min without light. Next, the sections were washed once with PBS, and incubated with 5 μg/mL Hoechst 33342 for 30 min. The sections were then washed twice with PBS and incubated overnight with rabbit anti‐nestin (1 : 100; Santa Cruz Biotechnology, Santa Cruz, CA, USA) at 4°C. Lastly, slides were incubated with FITC‐ and TRITC‐conjugated secondary antibodies for 2 h at 4°C. The stained sections were detected with a Leica fluorescence microscope (Heidelberg, Germany).

Thirty male Sprague–Dawley rats (6/group, 4‐week old, weighing approximately 60 g) were obtained from the Experimental Animal Center of Nantong University, and maintained in stainless steel cages, which were pre‐treated with 0.5% EDTA. [Correction added on 30 July 2015, after first online publication: sentence was revised, giving the source of experimental animals.] Rats were acclimated for 1 week to the temperature‐ and humidity‐controlled environment with a 12‐h‐dark/12‐h‐light cycle. Experimental diets were prepared in the same way as AIN‐93G (Reeves et al . 1993 ). Rats were randomly divided into five groups: control group (CO, 38 mg Zn/kg/day diet), pair‐fed group (PF, 38 mg Zn/kg/day diet), marginally zinc‐deficient group (5 mg Zn/kg/day diet), and severely zinc‐deficient group (SZD, 1 mg Zn/kg/day diet). For the PF groups, each rat was fed with the PF diet daily in amounts equal to that consumed in the previous 24 h by its SZD/marginally zinc‐deficient group paired mate. [Correction added on 30 July 2015, after first online publication: “mouse” has been corrected to “rat”.] All the five groups were supplied with double‐distilled drinking water. The weight of rats was monitored throughout the experiments. When the rats were fed after 2 months, rats ( n = 3 per group) were killed for sections. 24 h before all rats were killed, three rats per group were injected with EdU 1 mg/kg. According to the National of Health Guideline for the Care and Use of Laboratory Animals, animal care was executed and all experimental procedures have been approved by the Animal Care and Use Committee of Nantong University, China.

Our results revealed that FoxO3a and p27 kip1 expression potentially contributed to TPEN‐induced NSCs growth impairment. Next, we analyzed whether application of FoxO3a siRNA could lessen TPEN‐induced proliferating impairment. As shown in Fig. 7 (a), FoxO3a siRNA restored TPEN‐induced growth in C17.2 cells following TPEN exposure. Additionally, the cell cycle distribution of C17.2 cells was analyzed using flow cytometric analysis, which confirmed that the cells in S phase were significantly decreased after TPEN treatment, and FoxO3a siRNA partly restored the proportion of cells in S phase, compared with TPEN‐exposed cells (Fig. 7 b). Overall, our results demonstrated that FoxO3a played a key role in zinc deficiency‐induced growth impairment of C17.2 cells.

To further clarify the regulatory role of FoxO3a in zinc deficiency‐induced NSCs dysfunction, we next examined the effect of depleting FoxO3a expression on NSCs physiology using FoxO3a siRNA. C17.2 cells were transfected with FoxO3a siRNA after 24 h transfection, and cells were exposed to 1500 nM TPEN for an additional 24 h. After transfecting C17.2 NSCs with FoxO3a‐2541 siRNA, TPEN‐induced up‐regulation of FoxO3a was markedly attenuated (Fig. 6 c and d). Additionally, knockdown of FoxO3a significantly attenuated zinc deficiency‐induced p27 kip1 increasing (Fig. 6 c and d). Therefore, our results implicated that FoxO3a activation played an important role in zinc deficiency‐induced p27 kip1 expression in C17.2 NSCs.

Western blot analysis of the level of FoxO3a and p27 kip1 p‐foxO3a, proliferating cell nuclear antigen (PCNA) and CyclinA following zinc deficiency exposure in C17.2 cells. Cells were treated for different concentration (a, b, e, and f). Samples of cytoplasmic or nuclear extracts isolated at indicated N , N , N ’, N ’‐tetrakis‐(2‐pyridylmethy) ethylenediamine (TPEN) exposure and were examined to western blot analysis. The cells were treated 1500 nM TPEN for 24 h compared to the control (c and d). Values are means ± SEM ( n = 3). * p < 0.05 significantly different from control group.

Since our aforementioned data proved that severe dietary zinc deficiency increased FoxO3a and p27 kip1 expression and induced the impairment of NSCs proliferation in dietary zinc‐deficient rat hippocampus, we analyzed whether similar phenomenon could be observed in C17.2 NSCs cultures. Numerous studies have showed that activated FoxO3a induced cell cycle arrest cooperating with regulating p27 kip1 expression (Medema et al . 2000 ). As shown in Fig. 4 (a) and (b), TPEN exposure induced the expression of FoxO3a and p27 kip1 in C17.2 cells in a dose‐dependent manner. Because FoxO3a is inactivated via phosphorylation on three conserved serine/threonine residues by phosphatidylinositol 3‐kinase‐Akt pathway, we detected the level of phosphorylated form of FoxO3a following TPEN exposure (Burgering and Kops 2002 ). Western blot analysis revealed that the level of phosphorylated FoxO3a was down‐regulated under same conditions (Fig. 4 a and b). Moreover, it was reported that Akt‐mediated phosphorylation led to FoxO3a translocation from the nucleus into the cytoplasm (Brunet et al . 1999 ; Burgering and Kops 2002 ). We further analyzed the subcellular distribution of FoxO3a and found that FoxO3a was mainly localized in the nucleus when the cells were treated with 1500 nM TPEN for 24 h (Fig. 4 c and d). As expected, the expression of p27 kip1 had a similar alternation under same circumstance (Fig. 4 c and d). Morphological experiment also demonstrated that TPEN could elevate nuclear accumulation of FoxO3a and p27 kip1 compared with the control (Fig. 5 a–l). Next, we detected the expression of pro‐proliferation proteins, including PCNA and Cyclin A. After treatment with different concentrations of TPEN, the expression of PCNA and Cyclin A was examined using western blot analysis, which showed that TPEN exposure resulted in the up‐regulation of PCNA and Cyclin A in a dose‐dependent manner (Fig. 4 e and f). Taken together, consistent with our in vivo model, these findings validated that zinc deficiency facilitated the expression of FoxO3a and p27 kip1 , which coincided with impaired proliferation of C17.2 NSCs.

Our experiments showed that dietary zinc deficiency led to impaired NSCs proliferation and up‐regulation of FoxO3a and p27 kip1 . Under this circumstance, we further investigated PCNA, which has been regarded as a general regulator of DNA replication and cell cycle control (Strzalka and Ziemienowicz 2011 ). Western blot analysis showed PCNA expression was evidently decreased in severely dietary zinc‐deficient rat hippocampus (Fig. 3 j and k). Furthermore, double immunofluorescent staining results showed that the number of nestin‐positive cells expressing PCNA significantly declined after severe dietary zinc deficiency (Fig. 3 l–r). Thus, our in vivo model demonstrated that severe dietary zinc deficiency could induce the impairment of NSCs proliferation via an unrevealed role of up‐regulating FoxO3a/p27 kip1 expression.

We next investigated whether dietary zinc restriction had any effect on the impairment of NSC proliferation in the male rat brain hippocampus. To detect NSCs, proliferation was performed on brain sections after EdU injection for 7 days exposure to 2‐month dietary zinc deprivation. The staining images showed that the number of EDU‐positive NSCs was obviously decreased in the severely zinc‐deficient group compared with control group (Fig. 3 a–i). Thus, severe dietary zinc deficiency leads to significant impairment of NSCs proliferation in adult rat hippocampus.

To verify whether severe dietary zinc deficiency could alter the expression and distribution of FoxO3a and p27 kip1 in rat brain hippocampus, immunohistochemistry on transverse cryosections was performed. Compared with the control group, immunopositive cells of FoxO3a and p27 kip1 in the brain hippocampus were significantly increased (Fig. 1 c–h). At the same time, double‐immunofluorescent labeling was conducted to analyze the co‐localization of FoxO3a and p27 kip1 (green) with different phenotype‐specific markers, such as nestin (neural stem cell marker; red), GFAP (astrocyte marker). It was found that FoxO3a and p27 kip1 were mainly distributed in nestin‐positive cells (Fig. 2 ), and were less found in GFAP (data not shown). Notably, severe dietary zinc deficiency also resulted in markedly increased number of Foxo3a and p27 kip1 ‐positive NSCs in severely dietary zinc‐deficient rat hippocampus. Therefore, coinciding with western blot results, the findings showed that severe dietary zinc deficiency could lead to significant alterations in FoxO3a and p27 kip1 expression in severely dietary zinc‐deficient rat hippocampus, which potentially contributed to functional deficits of hippocampal NSCs.

In order to determine the aberrant changes of zinc deficiency‐induced neurological function, we established a dietary zinc deficiency model using Sprague–Dawley rats. After fed with control or zinc‐deficient diet for 2 months significant growth retardation was observed in severely zinc‐deficient rats, compared with control and maginally zinc‐deficient rats (Data not Shown). Next, the level of zinc in the hippocampus of the rats was examined. As shown in Figure S1, zinc content was evidently decreased in the hippocampus of severely zinc‐deficient rats, while zinc content in the hippocampus of all other groups remains largely unchanged. This piece of data is consistent with previous reports indicating that the alteration of zinc homeostasis in the hippocampus requires chronic zinc deficiency at a relatively severely zinc‐deficient condition. Then, the expression of various proteins was detected after dietary zinc deficiency. Western blot results showed that FoxO3a protein level was up‐regulated in severe zinc deficiency group, compared to pair‐fed control and control group. Meanwhile, the expression of p27 kip1 was also increased in the model (Fig. 1 a and b). Therefore, these data revealed that severe dietary zinc deficiency increased FoxO3a and p27 kip1 expression in the brain hippocampus.

Discussion

Zinc deficiency leads to reduced hippocampus neurogenesis in mammalian brain, whose underlying mechanism remains poorly understood. The use of dietary zinc‐deficient rat models and cell cultures to describe the pathological symptoms is an effective way to investigate the molecular and cellular mechanisms involved in zinc deficiency‐induced abnormal changes in the rat hippocampus. In the present study, we demonstrated that zinc deficiency up‐regulates the protein levels of FoxO3a and p27kip1, leading to consequent impairment of NSC proliferation using animals models and NSCs cultures. Moreover, we showed that depletion of FoxO3a expression suppressed p27kip1 transcription and largely restored NSCs viability following zinc deficiency. Taken together, these findings validated FoxO3a activation's contribution to aberrant p27kip1 regulation and NSCs growth impairment during zinc deficiency‐induced abnormal neurogenesis.

Zinc deficiency has been an important public health concern affecting the development of human intelligence, especially during pregnancy and childhood. Various studies have linked chronic zinc deficiency to the impairment of learning and memory capacities. Because hippocampal function plays a central role in cognitive processing and spatial learning, many researchers have focused the physiological alterations of hippocampus after zinc deprivation. These researches established that hippocampal neuron functions, including synaptic responses, the release of neurotransmitters as well as the expression of neuronal receptors, were heavily affected after zinc deficiency (Hesse 1979, Takeda et al. 2003, Doboszewska et al. 2015). Notably, studies implicated that NSCs might be also involved in zinc deficiency‐induced hippocampal disorders. Wang et al. reported that the expression of nestin, a marker protein of NSCs, declined in mice following maternal zinc deficiency (Wang et al. 2001). Later, Corniola et al. (2008) found that zinc deficiency impairs the proliferation of NSCs and induced the apoptosis of NSCs through the modulation of p53 signaling (Seth et al. 2015, Corniola et al. 2008). Zinc deficiency may also affect the differentiation of neuronal precursor cells, and thus played a role in neurogenesis in this regard (Gower‐Winter et al. 2013). We also recently found that Wnt/β‐catenin signaling was altered in TPEN‐exposed NSCs in vitro (Zhao et al. 2015). The current study verified that severe zinc deficiency for 60 days significantly attenuated the growth of hippocampal NSCs in vitro, supporting the notion that reduced neurogenesis may be an important mechanism underlying zinc deficiency‐induced hippocampal malfunction.

Recent evidence indicates that FoxO3a plays a significant role in governing NSCs biology including self‐renewal, proliferation, survival, and oxidative stress (Paik et al. 2009; Wu et al. 2009). Renault et al. (2009) have shown that deficit of FoxO3a alone in the mammalian brain results in significantly decreased number of NSCs in vivo, which elucidates the dominant role of FoxO3a in maintaining the brain NSCs function. Besides, the microRNA cluster miR‐106b~25‐binding FoxO3a has been reported to regulate NSCs/NPCs proliferation and neural differentiation in adult mammals (Brett et al. 2011). Consistent with functional redundancy among FoxO3a in other tissues (Paik et al. 2007), FoxO3a drives proliferation in anaplastic thyroid carcinoma via transcription of cyclin A1 (Marlow et al. 2012). In addition, FoxO3a is essential for maintaining the hematopoietic stem cell pool (Miyamoto 2008). These findings implicated a key role of FoxO3a in regulating NSCs biology. FoxO3a has also been manifested to bind the Foxo‐binding sites in the promoter of p27kip1, a key negative regulator of G1 to S cell cycle transition involved in cell quiescence of NSCs (Kops et al. 2002; Nakao et al. 2008; Renault et al. 2009). It has been reported that zinc deprivation impairs neuronal precursor cell proliferation through p53‐mediated mechanism using in vitro NSCs cultures (Corniola et al. 2008). However, whether zinc deficiency can affect FoxO3a and p27kip1 gene expression and NSCs physiology function remains unclear. In line with the hypothesis, our findings demonstrated that FoxO3a is evidently involved in dietary zinc deficiency‐exposed hippocampal NSCs growth impairment via regulating the target gene, p27kip1, which is similar to the findings of the in vitro model of TPEN‐exposed C17.2 cells. Furthermore, we proved that the marker of S phase progression, Cyclin A, was significantly decreased in a FoxO3a‐directed manner in C17.2 cells. Critically, interference of FoxO3a effectively restored NSCs proliferation after zinc deficiency exposure in C17.2 cells accompanying decreased expression of p27kip1. Additionally, FoxO3a, serving as down‐stream target of Akt, was inactivated through Akt phosphorylation on T32, S253, and S315 residues, particularly on S253, which hinders FoxO3a‐binding to DNA (Zhang et al. 2002; Liu et al. 2009). From our series of study, we found the level of p‐FoxO3a (Ser253) deceased after being exposed to zinc deficiency in a dose‐dependent manner. Taken together, these findings inferred that the regulation of Foxo3a and p27kip1 might play a significant part in zinc deficiency‐induced NSCs growth impairment.

Our data showed that the aberrant alternation of FoxO3a plays a vital role in zinc deficiency‐induced NSCs growth impairment. While various studies have also been done with regard to the mechanisms triggering FoxO3a expression, the role of FoxO3a regulatory pathways in the process remains largely obscure. FoxO3a is tightly regulated by ubiquitination, acetylation and phosphorylation of Akt, SGK1, AMP‐activated protein kinase (AMPK), Inhibition of IkappaB kinase β (IKKβ) and c‐Jun N‐terminal kinase (JNK) in order to adjust cell cycle, oxidative resistance, and apoptosis (Huang and Tindall 2007; Storz 2011). For example, in human mesenchymal stem cells, resveratrol promoted osteogenesis of stem cells through activating RUNX2 gene transcription via Sirt1/FoxO3a axis (Tseng et al. 2011). FoxO‐null NSCs regulation of intracellular Reactive Oxidative Species (ROS) level affects NSCs self‐renewal capacity in postnatal brain (Paik et al. 2009). In the central nervous system, caspase, and p53 temporally modulate FoxO3a/1d1 signaling during mouse NSCs differentiation (Aranha et al. 2009). In addition, there is evidence that FoxO3a‐mediated gene transcription induces oxidative stress, generating neuronal cell death (Peng et al. 2015). For example, in the hippocampus of rats, oxidative stress activated the transcriptions FoxO1a and FoxO3a exposed to low dosed of ozone, which induced neuronal reentry in to the cell cycle and apoptotic death (Gomez‐Crisostomo et al. 2014). However, zinc deficiency in neuronal precursor cells also resulted in the translocation of p53 to the mitochondria leading to mitochondrial alternations and apoptosis, along with increase in the production of ROS (Seth et al. 2015). Further, these data could provide insight into a novel molecular pathway that zinc deficiency‐induced aberrant expression of FoxO3a might involve the alternation of NSCs fates. However, the precise molecular mechanisms of FoxO3a underlying zinc deficiency‐exposed NSCs growth impairment still need to be further elucidated.

In conclusion, we present evidence that dietary zinc deficiency causes significantly increased expression of FoxO3a and p27kip1 in adult rat hippocampus and C17.2 cells, which might play a key role in regulation of FoxO3a in NSCs growth impairment. These results suggest that zinc deficiency has adverse effects on adult rat brain hippocampal NSCs proliferation. Taken together, our findings shed light on the mechanisms of NSCs growth impairment in zinc deficiency exposure, and may aid the prevention or treatment of zinc deficiency‐induced neurological disorders.