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apoptosis induced by intracellular zinc deficiency associated with changes in amino-acid neurotransmitters and glutamate receptor subtypes
Journal of Inorganic Biochemistry 179 (2018) 54–59
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Journal of Inorganic Biochemistry journal homepage: www.elsevier.com/locate/jinorgbio
Neuronal death/apoptosis induced by intracellular zinc deficiency associated with changes in amino-acid neurotransmitters and glutamate receptor subtypes Kun Tian, Yu-xiang Wang, Li-xia Li, Yan-qiang Liu
T
⁎
College of Life Sciences, Nankai University, Tianjin 300071, China
A R T I C L E I N F O
A B S T R A C T
Keywords: Zinc deficiency Neuronal apoptosis Amino acid neurotransmitters GluR2 NR2B
In the present study, a model of zinc deficiency was developed by exposing primary neurons to an N,N,N′,N′Tetrakis (2-pyridylmethyl) ethylenediamine (TPEN)-containing medium. The cell survival rate, apoptosis rate, intracellular and extracellular concentrations of 4 amino acids, and the expression of 2 glutamate receptor subtypes α-amino-3-hydroxy-5-methyl-4-isoxazolepropionate receptor (GluR2)and N-methyl-D-aspartate receptor subtype 2B (NR2B) were evaluated in zinc-deficient cells. The results revealed that zinc deficiency led to a decrease in cell viability and an increase in the apoptosis rate. Additionally, in cultured neurons, zinc deficiency led to an increase in the concentration of aspartic acid (Asp) and a decrease in the concentrations of glutamate (Glu), glycine (Gly), and gamma-aminobutyric acid (GABA). These changes were reversed by concurrent zinc supplementation. Furthermore, zinc deficiency led to an increase in the secreted amounts of Glu, Gly, and Asp but a decrease in secreted amounts of GABA, as measured using the concentrations of these amino acids in the cell-culture medium. These changes were partially reversed by zinc supplementation. Finally, zinc deficiency led to a significant decrease in GluR2 expression and an increase in NR2B expression in cultured neurons, whereas simultaneous treatment with zinc sulfate (ZnSO4) prevented these changes. These results suggest that zinc deficiency-induced neuronal death/apoptosis involves changes in the concentrations of 4 amino acid neurotransmitters and the expression of 2 glutamate receptor subtypes.
1. Introduction Zinc has long been regarded as an important nutrient, and is involved in several biochemical pathways [1]. Zinc is essential for the activity of multiple enzymes involved in cell survival pathways and metabolic homeostasis [2,3]. In addition, some studies have reported that zinc plays an important role in hippocampal neurogenesis in rats [4]. Under normal conditions, a dynamic balance of zinc is maintained throughout the body in multiple ways. Perturbations in zinc homeostasis can cause several physiological disorders and neurological diseases, and result in brain dysfunction. Previous studies have reported that intracellular zinc deficiency induces apoptosis and cell death in cultured neurons [5,6]. The exact mechanism underlying zinc deficiency-induced neuronal apoptosis remains to be elucidated. Neurotransmitters play an important role in the transmission of signals across chemical synapses. They are released from pre-synaptic neurons and bind to receptors on post-synaptic neurons, resulting in a change in the excitatory state of these neurons [7]. Glutamate, an excitatory neurotransmitter, is released into the synapse [8,9]. Changes in ⁎
glutamate metabolism can cause the release of large amounts of glutamate, resulting in cellular excitotoxicity and diseases of the nervous system [10]. Glutamate is released from neurons and binds to specific receptors, thus regulating cellular excitability [8]. Ionic glutamate receptors (iGluRs) are important glutamate receptors and include αamino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA), N-methyl-D-aspartate (NMDA), and kainite receptor subtypes [11,12]. Nmethyl-D-aspartate (NMDA) receptor subtype 2B (NR2B) is the regulatory subunit of the functional NMDA receptor [13], and glutamate receptor 2 (GluR2) is an important component of the AMPA receptor. Previous studies have reported that impairment of signaling via both the AMPA and NMDA receptors is involved in multiple brain pathologies. Overactivation of NMDA receptors may cause calcium overload within a cell, and this may lead to cytotoxicity and subsequently, to neuropathy [14–16]. Glycine (Gly) and γ-aminobutyric acid (GABA) are the main inhibitory neurotransmitters in the central nervous system, and thus regulate neuronal excitation. Gly triggers the opening of the ionic channels coupled to Gly receptors, which induces hyperpolarization in the postsynaptic neuron and inhibits neuronal excitation
Corresponding author. E-mail address: [email protected] (Y.-q. Liu).
https://doi.org/10.1016/j.jinorgbio.2017.11.014 Received 18 July 2017; Received in revised form 7 November 2017; Accepted 17 November 2017 Available online 21 November 2017 0162-0134/ © 2017 Elsevier Inc. All rights reserved.
Journal of Inorganic Biochemistry 179 (2018) 54–59
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5 μM of TPEN to the culture medium. To assess whether Zn2 + supplementation can rescue the effects of zinc deficiency, in the TPEN + ZnSO4 condition, 5 μM of TPEN and 10 μM of ZnSO4 were added to the culture medium.
[17,18]. GABA also inhibits neuronal firing, and has been implicated in the regulation of cognitive functions [19,20]. Reports suggest that in some diseases, cellular apoptosis is closely related to changes in the concentrations of amino acid neurotransmitters [21]. In the present study, we attempted to elucidate the mechanism underlying zinc deficiency-induced neuronal death/apoptosis by evaluating the effects of zinc deficiency and concomitant Zn2 + supplementation on the concentration of amino acid neurotransmitters and the expression of glutamate receptors in cultured neurons.
2.4. Measurement of cell viability using the MTT assay Viability of the cultured primary hippocampal neurons was determined using an MTT assay, which measures the activity of mitochondrial succinate dehydrogenase within viable cells by the reduction of exogenous MTT to insoluble purple formazan crystals. Neurons were plated in 96-well plates at a density of 1 × 105 cells/mL. The volume of medium added to each well was 200 mL. For the assay, MTT solution was added to each well. The final concentration of the MTT solution was 1 mg/mL. Neurons were incubated in this solution at 37 °C for 4 h, following which the medium was discarded and the formazan crystals were dissolved in 150 μL of DMSO. The absorbance of this solution at 570 nm was measured using a Beauty Diagnostic Microplate Reader (Molecular Devices, Sunnyvale, California, USA). The results were expressed in terms of percentage of the absorbance value obtained from the control group.
2. Materials and methods 2.1. Chemicals and animals 4-(2-Hydroxyethyl)-1-Piperazine ethane sulfonic acid (HEPES), Lglutamic acid, and glycine (purity > 99%) were obtained from Genview Scientific Inc. (Tallahassee, USA). Antibiotic solution, poly-Llysine, γ-aminobutyric acid (purity ≥ 99.0%), and N,N,N′,N′-Tetrakis (2-pyridylmethyl) ethylenediamine (TPEN) were obtained from SigmaAldrich (St. Louis, MO, USA). L-Aspartic acid (purity = 99%) was obtained from KisanBiotech Co. Ltd. (Seoul. Korea). 2,4Dinitrofluorobenzene(DNFB)was obtained from Yolne Chemical Co. Ltd. (Shanghai, China). 3-(4,5-Dimethylthiazol-2-yl)-2,5Diphenyltetrazolium bromide (MTT) was obtained from Amresco (Solon, OH, USA). Dulbecco's Modified Eagle's Medium (DMEM)/nutrient mixture F12 with Glutamax™-1 and B27 supplements were obtained from Gibco (Gran Island, NY, USA). Dimethyl sulfoxide was obtained from Dingguo Changsheng Biotechology Co. Ltd. (Beijing, China). The terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling (TUNEL) assay kit and the fixative used in immunohistochemistry were purchased from Biyuntian Co. (Beijing, China). The GluR2 immunohistochemical assay kit [polyclonal rabbitanti GluR2 (N-19), donkey anti-rabbit IgG-TR, etc.] and the NR2B immunohistochemical assay kit [polyclonal goat anti-NMDA NR2B (C-20), donkey anti-goat IgG-FITC, etc.] were purchased from Santa Cruz Biotechnology, Inc. (CA, USA). All animals used for experiments in the present study were obtained from the experimental animal center of the Academy of Military Medical Sciences, and were cared for in accordance with the institutional guidelines for the health and care of experimental animals. The experimental protocols were approved by the Committee on the Ethics of Animal Experiments of Nankai University.
2.5. Measurement of cell apoptosis rate using the TUNEL assay Neurons, at a density of 1 × 105 cells/mL, were grown on sixchamber well slides pre-coated with poly-L-lysine (PLL). After 24 h of treatment, cells were washed thrice with phosphate buffered saline (PBS), following which 2 mL of TUNEL assay solution was added to each well. The cells were then fixed for 30 min, and washed thrice with PBS, then permeabilized with 0.1% Triton X-100 PBS on an ice bath. According to the manufacturer's instructions, cells processed for TUNEL staining were co-stained with 4, 6-diamidino-2-phenylindole (DAPI) for 2 min. The number of TUNEL-positive neurons and the total number of neurons were counted. The percentage of TUNEL-positive cells was calculated and averaged. The results were expressed in terms of the percentage of TUNEL-positive cells. 2.6. Determination of amino acid concentrations using high-performance liquid chromatography (HPLC) The concentrations of 4 free amino acid neurotransmitters (Glu, Asp, Gly, and GABA) were measured in primary neurons and the medium they were cultured in using a CoM6000 HPLC System consisting of two 6000 LDS pumps, a 6000 UV–Vis detector (360 nm), and a Comatex C18 (5 μm, 250 mm × 4.6 mm pore size) (CoMetro Technology, USA). Neurons were cultured at a density of 1.12 × 105 cells/cm2. After treatment with the respective chemical agents, culture medium was separately collected from neurons of all three experimental groups and centrifuged for 10 min at 1000 rpm to remove any remaining cells. Subsequently, acetonitrile was added to the supernatant (1:1, v:v) in order to precipitate proteins, which were separated out after centrifugation at 14,000 rpm for 20 min at 4 °C. The same process was carried out using blank culture media (culture media that was not used to grow any cells). To obtain amino acids from cultured neurons, 100 μL RIPA lysis buffer (50 mM Tris–HCl [pH 7.4], 150 mM NaCl, 1% NP-40) was added to fully lyse the cells. A 5-μL sample of the lysis solution was used for protein quantification using a bicinchoninic acid-protein quantification assay kit by following the manufacturer's instructions. Subsequently, 80 μL of acetonitrile was added to 80 μL of the lysis solution for protein precipitation, and the mixture was centrifuged at 14,000 rpm for 20 min at 4 °C. The sample solutions were derivatized by the addition of 0.1 M carbonate buffer and 2,4-dinitrochlorobenzene (DNCB)-acetonitrile (1:1000, v/v) followed by incubation in a 60 °C water-bath for 60 min in the dark. The derivatization reaction was stopped by the addition of mobile phase B. The mobile phase consisted
2.2. Primary neuronal culture Primary hippocampal neurons were collected from brains of newborn Wistar rats (approximately 1 day old). The hippocampus was isolated from the brain of each rat and treated with 0.125% trypsin for 20 min at 37 °C, triturated in a solution of DMEM/F12 with Glutamax™1 and 15% fetal bovine serum, and centrifuged for 15 min. The cells were suspended in DMEM/F12 medium containing Glutamax™-1 and supplemented with 2% B27, and 1% antibiotic solution. The cells were plated onto cover glasses pre-coated with poly-L-lysine (0.1 mg/mL) in multiwall cell-culture plates at a density of 1.0–5.0 × 105 cells/cm2. Cells were cultured in a humidified incubator (Sanyo, Japan) at 37 °C with 5% CO2 for 7 days, following which they subjected to treatment and further experiments. The medium was changed twice a week by replacing half the volume of culture medium with serum-free DMEM/ F12 medium with Glutamax™-1. 2.3. Experimental groups and TPEN treatment There were 3 experimental groups in the present study: the control group, the TPEN exposure group, and the TPEN exposure plus ZnSO4 treatment (TPEN + ZnSO4) group. Treatment lasted for 24 h. The model of zinc deficiency in primary neurons was established by adding 55
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of two parts: A (an isocratic mixture of acetonitrile and ultrapure water) and B (50 mM sodium acetate buffer solution containing 0.1% trimethylamine, pH 6.4). The elution program was selected to maintain a flow rate of 1.0 mL/min, starting with 75% B for 26 min, then 75% to 32% B for 8 min, 32% to 10% B for 8 min, 10% to 0% B for 5 min, 0% to 75% B for 5 min, and a final retention of 75% B for 2 min. Twenty microliters of the derivative were injected into the HPLC system and separation was performed at 40 °C. 2.7. Measurement of GluR2 and NR2B expression using immunofluorescence Expression of neuronal GluR2 and NR2B was measured using an immunofluorescence assay. After undergoing treatment for 24 h, neurons were fixed for 30 min using an immuno-staining fixation solution and then incubated with 2 mL of 2% bovine serum albumin (BSA) solution for 20 min. After this step, the cells were incubated in 80 μL (per cover glass) of solution containing rabbit-anti GluR2 primary antibody (1:50) at 4 °C overnight. Antibodies were detected using 80 μL (per cover glass) of fluorescein isothiocyanate (FITC)-conjugated donkey anti-goat IgG (1:100) in which the cells were incubated at 37 °C 1 h in a water bath. Finally, the neurons were mounted in fluorescence mounting medium. The prepared slides were viewed using a fluorescence microscope (20 × field). For each experimental group, 6 fieldviews were randomly selected. The sample was stimulated with green light to obtain red fluorescence, and the resulting images were recorded. Similarly, to measure the expression of NR2B, treated neurons were incubated in a solution containing anti-NMDAƐ2 (C-20) (1:50) antibody at room temperature, followed by an incubation in a solution containing FITC-donkey anti-goat IgG secondary antibody (1:100). Unlike in the method used to detect GluR2, a blue light was selected to stimulate the neurons and obtain green fluorescence. Image-Pro Plus6.0 (IPP) software was used to calculate the average optical density (OD).
Fig. 1. Effects of N, N, N′, N′-Tetrakis (2-pyridylmethyl) ethylenediamine (TPEN) and zinc supplement on the neuronal survival rate (n = 8). **P < 0.01 compared with the control group; ##P < 0.01 compared with the TPEN group.
percentage of apoptotic neurons. 3.3. Zinc deficiency leads to changes in amino acid neurotransmitter levels in cultured neurons and the culture media Standard curves for 4 amino acids were plotted using a series of different concentrations (5, 25, 50, 100, 200, and 500 μM for L-glutamate [y = 0.0004x − 1.7566, R2 = 0.9991] and Gly [y = 0.0004x + 1.4421, R2 = 0.9984]; 5, 10, 25, 50, 100, and 200 μM for Asp [y = 0.0005x − 2.4202, R2 = 0.9947] and GABA [y = 0.0004x − 0.8948, R2 = 0.9999]). The HPLC peaks obtained for each of the four amino acids were used to determine the corresponding concentrations of those amino acids based on their respective standard curves. The concentration of the amino acids present in the treatment medium subtracted from that in blank culture medium represents the quantity of amino acids released from the cultured neurons. The concentrations of intracellular and extracellular amino acids were normalized to the concentration of amino acids per gram of cellular protein. Fig. 3A depicts the effect of 24 h of TPEN-treatment (zinc deficiency) on intracellular concentrations of Asp, Glu, Gly, and GABA. Concentrations of Glu, Gly, and GABA were significantly lower in the TPEN-treated cells than in the controls, but the TPEN-treated cells showed a tendency to have a higher concentration of Asp. Zn2 + supplementation could rescue the observed TPEN-induced decrease in Glu, Gly, and GABA concentration, but the concentration of Asp did not show much change. Fig. 3B depicts the effect of 24 h of zinc deficiency on the secreted amounts of Asp, Glu, Gly, and GABA. TPEN treatment led to a significant increase in the concentrations of Gly and Glu in medium used to culture neurons (P < 0.05), whereas the levels of Asp and GABA (P < 0.05) showed a significant decrease. Concurrent treatment with Zn2 + could rescue the TPEN-induced change in levels of Gly and GABA, but had obvious effects on Asp and Glu levels.
2.8. Data analysis SPSS 20 and Origin 8.5 were used for data treatment and analysis, and the results were presented as the mean ± standard error of the mean (SEM). A one-way ANOVA was performed for the comparison of multiple groups, and a value of P < 0.05 was considered as statistically significant; a value of P < 0.01 was considered as extremely statistically significant. 3. Results 3.1. Zinc deficiency reduces neuronal viability Following pilot experiments using MTT, 5 μM TPEN and 10 μM ZnSO4 were determined as optimum concentrations to induce zinc deficiency and provide Zn2 + supplementation, respectively, and were used in all subsequent experiments. The viability of neurons following 24 h of treatment was determined. As shown in Fig. 1, the viability rate of the cultured cells after TPEN treatment was significantly lower (73.7 ± 3.0%, P < 0.01) than that of the controls. In contrast, application of ZnSO4 could rescue the TPEN-induced reduction of viability (TPEN + ZnSO4 vs. TPEN, P < 0.01), and this rescue was statistically significant.
3.4. Zinc deficiency alters GluR2 and NR2B expression in cultured neurons
3.2. Zinc deficiency causes an increase in neuronal apoptosis
Fig. 4 depicts the effects of zinc deficiency on GluR2 expression. Treatment with TPEN caused a significant reduction in GluR2 expression (P < 0.01, compared with control), whereas TPEN + ZnSO4 treatment rescued this TPEN-induced reduction (P < 0.01, compared with TPEN). The effects of TPEN on NR2B expression are shown in Fig. 5. TPEN treatment led to a significant increase in neuronal NR2B expression (P < 0.05, compared with control). Concurrent ZnSO4 treatment could bring the NR2B expression back to the levels observed
Fig. 2 reveals the effect of zinc deficiency on neuronal apoptosis. Results show that the apoptosis rate was higher in neurons that underwent TPEN treatment than in those that did not (P < 0.01). In neurons treated with TPEN + ZnSO4, the apoptosis rate was lower than in neurons treated with only TPEN (P < 0.01). These results suggest that TPEN-induced zinc deficiency causes a significant increase in the 56
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Fig. 2. Effects of N, N, N′, N′-Tetrakis (2-pyridylmethyl) ethylenediamine (TPEN) and zinc supplement on the neuronal apoptosis rate (n = 8). (A). Representative images of immunofluorescence stained neurons after different treatments; scale bars = 50 μm. (B). The neuronal apoptosis rate following different treatments **P < 0.01 compared with the control group; ##P < 0.01 compared with the TPEN group.
changes in the concentration of amino acid neurotransmitters, and altered expression of two glutamate receptor subtypes, GluR2 and NR2B. These results also show that Zn2 + supplementation reversed these changes. A previous study has demonstrated that zinc deficiency causes intracellular oxidative stress, neuronal damage, and apoptosis [6]. It has been demonstrated that zinc deficiency can induce apoptosis of MC3T3E1 cells through a mitochondria-mediated pathway [22]. The zinc deficiency-induced apoptosis of cells may involve several pathways. Zinc is involved in the metabolism of free amino acid neurotransmitters, and zinc deficiency causes an increase in the glutamate concentration in hippocampal neurons, which increases the excitability of these cells
in the control (P < 0.01, compared with TPEN), thus reversing the change caused by TPEN treatment. 4. Discussion Zinc plays an important role in regulating the physiological functions of neurons. As an important neuromodulator, it can regulate neurotransmitter release. Perturbation of zinc homeostasis in the brain leads to disturbances in cognitive function and may cause neurological disorders. In the present study, TPEN was used to establish a model of zinc deficiency in cultured neurons. These results show that zinc deficiency led to lower rates of neuronal survival, higher rates of apoptosis,
Fig. 3. Effects of N, N, N′, N′-Tetrakis (2-pyridylmethyl) ethylenediamine (TPEN) and zinc supplement on the concentration of amino acids concentration in neurons and culture medium (n = 6). (A) The free amino acid concentration in neurons following different treatments. (B) The free amino acid concentration in the culture medium following different treatments. *P < 0.05, **P < 0.01 compared with the control group; #P < 0.05, ##P < 0.01 compared with the TPEN group.
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Fig. 4. Effects of N, N, N′, N′-Tetrakis (2-pyridylmethyl) ethylenediamine (TPEN) and zinc supplement on GluR2 expression in neurons (n = 8). (A). Representative images of GluR2 immunofluorescence staining in neurons following different treatments; scale bars = 50 μm. (B). The optical density (OD) of GluR2 immunofluorescence in neurons following different treatments. **P < 0.01 compared with the control group; ##P < 0.01 compared with the TPEN group.
Fig. 5. Effects of N, N, N′, N′-Tetrakis (2-pyridylmethyl) ethylenediamine (TPEN) and zinc supplement on NR2B expression in neurons (n = 8). (A). Representative images of NR2B immunofluorescence staining in neurons following different treatments; scale bars = 50 μm. (B). The optical density (OD) of NR2B immunofluorescence in neurons following different treatments. **P < 0.01 compared with the control group; ##P < 0.01 compared with the TPEN group.
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DMEM TUNEL
Dulbecco's Modified Eagle's Medium terminal deoxynucleotidyl transferase-mediated dUTP nickend labeling DMSO dimethyl sulfoxide PLL poly-L-lysine PBS phosphate buffered saline DAPI 4,6-diamidino-2-phenylindole HPLC high-performance liquid chromatography RIPA lysis buffer Radio Immunoprecipitation Assay BSA bovine serum albumin FITC fluorescein isothiocyanate IgG Immunoglobulin G
[23]. Our experimental results show that exposure to TPEN causes a significant reduction in the glutamate concentration and an increase in aspartic acid levels in cultured primary neurons, whereas Zn2 + supplementation could reverse these changes to a certain extent. In addition, TPEN-treatment caused a significant increase in the Glu concentration of the culture medium, which suggests that large amounts of glutamate were released from the neurons. Furthermore, providing additional Zn2 + to these neurons partially reversed the change induced by zinc deficiency, indicating that zinc deficiency might influence Glu metabolism, resulting in toxic neuronal excitability. In addition, TPENtreatment led to a significant reduction in Gly and GABA levels in neurons, and simultaneous addition of Zn2 + significantly reversed these changes. This could be a result of TPEN reducing the synthesis of GABA in the culture medium; the implications of these findings warrant further studies for validation. Amino acid neurotransmitters usually bind to specific receptors to perform their physiological functions. Previous studies have shown that glutamate induces cell excitability by binding to ionic glutamate receptors [24], such as NMDA and AMPA receptors [25,26]. The NR2 subunit plays an important role in regulating physiological functions of the NMDA receptor [11,27]. The GluR2 subunit of the AMPA receptor determines the intracellular calcium permeability of the AMPA receptor [2]. The results of the present study indicate that TPEN-treatment leads to an increase in the expression of the NR2B subunit of the NMDA receptor. In addition, TPEN-treatment leads to a reduction in the expression of the GluR2 subunit of the AMPA receptor. We know that the activation of the NMDA receptor and the depression of the AMPA receptor together cause a large Ca2 + influx, excitatory cytotoxicity and neuronal apoptosis. In addition, we know that the role of redox active zinc in neurons is tightly linked to the role of redox active copper via the enzyme copper–zinc superoxide dismutase [28,29], a homodimeric metalloprotein that binds copper and zinc ions and forms an intramolecular disulfide bond. It has been reported that copper–zinc superoxide dismutase functions as a molecular switch that activates the endoplasmic reticulum stress response, which plays an important role in cellular homeostasis under zinc-deficient conditions [29]. Therefore, neuronal death/apoptosis induced by zinc deficiency might involve changes in copper–zinc superoxide dismutase and cupric ion concentrations. However, this has not been addressed in the present study and will need further investigation to be validated. In conclusion, neuronal death induced by zinc deficiency may be associated with increased neuronal glutamate release, down-regulation of GluR2, and up-regulation of NR2B.
Author contributions Conception and design: Yan-qiang Liu, Kun Tian. Acquisition of data: Kun Tian, Yu-xiang Wang, Li-xia Li. Analysis and interpretation of data: Kun Tian. Writing, review of the manuscript: Kun Tian, Yan-qiang Liu. Conflict of interest The authors declare that they have no conflict of interest. Acknowledgements This work was supported by the National Natural Science Foundation of China (No. 31272317) and the Natural Science Foundation of Tianjin City (15JCYBJC24500) sanctioned to YQL. References [1] R.J. Radford, S.J. Lippard, Curr. Opin. Chem. Biol. 17 (2013) 129–136. [2] Z. Liu, Y.Y. Huang, Y.X. Wang, H.G. Wang, F. Deng, B. Heng, L.H. Xie, Y.Q. Liu, J. Trace Elem. Med. Biol. 31 (2015) 45–52. [3] Y. Song, S.W. Leonard, M.G. Traber, E. Ho, J. Nutr. 139 (2009) 1626–1631. [4] S.W. Suh, S.J. Won, A.M. Hamby, B.H. Yoo, Y. Fan, C.T. Sheline, H. Tamano, A. Takeda, J. Liu, J. Cerebr. Blood F. Met. 29 (2009) 1579. [5] W. Pang, H. Lu, Y.D. Hu, H.P. Yang, X. Leng, Y.G. Jiang, Nutr. Neurosci. 15 (2013) 18–24. [6] M.P. Zago, G.G. Mackenzie, A.M. Adamo, C.L. Keen, P.I. Oteiza, Antioxid. Redox Signal. 7 (2005) 1773–1782. [7] A. Miszke, K. Rapacz, Otolaryngol. Pol. 45 (1991) 374–380. [8] R.S. Sundaram, L. Gowtham, B.S. Nayak, Asian J. Pharm. Clin. Res. 5 (2012) 1–7. [9] M.V. Johnston, MRDD. Res. Rev. 7 (2001) 229–234. [10] D.W. Choi, J. Neurobiol. 23 (1992) 1261–1276. [11] X.L. Ma, F. Zhang, Y.X. Wang, C.C. He, K. Tian, H.G. Wang, D. An, B. Heng, Y.Q. Liu, Chem. Biol. Interact. 254 (2016) 73. [12] M. Hollmann, M. Hartley, S. Heinemann, Science 252 (1991) 851–853. [13] S. Cull-Candy, S. Brickley, M. Farrant, Curr. Opin. Neurobiol. 11 (2001) 327–335. [14] R.P. Simon, J.H. Swan, T. Griffiths, B.S. Meldrum, Science 226 (1984) 850–852. [15] B. Liu, M. Liao, J.G. Mielke, K. Ning, Y. Chen, L. Li, Y.H. El-Hayek, E. Gomez, R.S. Zukin, M.G. Fehlings, J. Neurosci. 26 (2006) 5309–5319. [16] M.S. Beattie, A.R. Ferguson, J.C. Bresnahan, Eur. J. Neurosci. 32 (2010) 290–297. [17] M.S. Hernandes, L.R.P. Troncone, J. Neural Transm. 116 (2009) 1551–1560. [18] M.S. Hernandes, M.L. De, L.R. Troncone, Brain Res. 1168 (2007) 32–37. [19] J.L. Barker, Science 240 (1988) 548–549. [20] J.E. Walker, Neurochem. Res. 8 (1983) 521–550. [21] Y.H. Sun, Y. Yue, Y. Wang, J. Apoplexy Nervous Dis. 22 (2005) 344–347. [22] B. Guo, M. Yang, D. Liang, L. Yang, J. Cao, L. Zhang, Mol. Cell. Biochem. 361 (2012) 209–216. [23] J.C. Wallwork, D.B. Milne, R.L. Sims, H.H. Sandstead, J. Nutr. 113 (1983) 1895–1905. [24] L. Kiedrowski, J. Neurochem. 130 (2014) 87–96. [25] N. Armstrong, E. Gouaux, Neuron 28 (2000) 165–181. [26] R. Gill, A.A. Alanine, B. Buttelmann, G. Fischer, M.P. Heitz, J.N. Kew, T.B. Levet, H.P. Lorez, P. Malherbe, M.T. Miss, J. Pharmacol. Exp. Ther. 302 (2002) 940–948. [27] G. Köhr, Cell Tissue Res. 326 (2006) 439–446. [28] E. Tokuda, I. Anzai, T. Nomura, K. Toichi, M. Watanabe, S. Ohara, S. Watanabe, K. Yamanaka, Y. Morisaki, H. Misawa, Y. Furukawa, Mol. Neurodegener. 12 (2017) 2–18. [29] K. Homma, T. Fujisawa, N. Tsuburaya, N. Yamaguchi, H. Kadowaki, K. Takeda, H. Nishitoh, A. Matsuzawa, I. Naguro, H. Ichijo, Mol. Cell 52 (2013) 75–86.
Abbreviations TPEN AMPA GluR2
N,N,N′,N′-Tetrakis (2-pyridylmethyl) ethylenediamine α-amino-3-hydroxy-5-methyl-4-isoxazolepropionate α-amino-3-hydroxy-5-methyl-4-isoxazolepropionate (AMPA) receptor NMDA N-methyl-D-aspartate NMDAR N-methyl-D-aspartate receptor NR2B N-methyl-D-aspartate (NMDA) receptor subtype 2B Asp aspartic acid Glu glutamate Gly glycine GABA gamma-aminobutyric acid ZnSO4 zinc sulfate iGluR ionic glutamate receptors HEPES 4-(2-hydroxyethyl)-1-piperazine ethane sulfonic acid DNFB 2,4-dinitrofluorobenzene MTT 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide
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Neuronal death/apoptosis induced by intracellular zinc deficiency associated with changes in amino-acid neurotransmitters and glutamate receptor subtypes Kun Tian, Yu-xiang Wang, Li-xia Li, Yan-qiang Liu
T
⁎
College of Life Sciences, Nankai University, Tianjin 300071, China
A R T I C L E I N F O
A B S T R A C T
Keywords: Zinc deficiency Neuronal apoptosis Amino acid neurotransmitters GluR2 NR2B
In the present study, a model of zinc deficiency was developed by exposing primary neurons to an N,N,N′,N′Tetrakis (2-pyridylmethyl) ethylenediamine (TPEN)-containing medium. The cell survival rate, apoptosis rate, intracellular and extracellular concentrations of 4 amino acids, and the expression of 2 glutamate receptor subtypes α-amino-3-hydroxy-5-methyl-4-isoxazolepropionate receptor (GluR2)and N-methyl-D-aspartate receptor subtype 2B (NR2B) were evaluated in zinc-deficient cells. The results revealed that zinc deficiency led to a decrease in cell viability and an increase in the apoptosis rate. Additionally, in cultured neurons, zinc deficiency led to an increase in the concentration of aspartic acid (Asp) and a decrease in the concentrations of glutamate (Glu), glycine (Gly), and gamma-aminobutyric acid (GABA). These changes were reversed by concurrent zinc supplementation. Furthermore, zinc deficiency led to an increase in the secreted amounts of Glu, Gly, and Asp but a decrease in secreted amounts of GABA, as measured using the concentrations of these amino acids in the cell-culture medium. These changes were partially reversed by zinc supplementation. Finally, zinc deficiency led to a significant decrease in GluR2 expression and an increase in NR2B expression in cultured neurons, whereas simultaneous treatment with zinc sulfate (ZnSO4) prevented these changes. These results suggest that zinc deficiency-induced neuronal death/apoptosis involves changes in the concentrations of 4 amino acid neurotransmitters and the expression of 2 glutamate receptor subtypes.
1. Introduction Zinc has long been regarded as an important nutrient, and is involved in several biochemical pathways [1]. Zinc is essential for the activity of multiple enzymes involved in cell survival pathways and metabolic homeostasis [2,3]. In addition, some studies have reported that zinc plays an important role in hippocampal neurogenesis in rats [4]. Under normal conditions, a dynamic balance of zinc is maintained throughout the body in multiple ways. Perturbations in zinc homeostasis can cause several physiological disorders and neurological diseases, and result in brain dysfunction. Previous studies have reported that intracellular zinc deficiency induces apoptosis and cell death in cultured neurons [5,6]. The exact mechanism underlying zinc deficiency-induced neuronal apoptosis remains to be elucidated. Neurotransmitters play an important role in the transmission of signals across chemical synapses. They are released from pre-synaptic neurons and bind to receptors on post-synaptic neurons, resulting in a change in the excitatory state of these neurons [7]. Glutamate, an excitatory neurotransmitter, is released into the synapse [8,9]. Changes in ⁎
glutamate metabolism can cause the release of large amounts of glutamate, resulting in cellular excitotoxicity and diseases of the nervous system [10]. Glutamate is released from neurons and binds to specific receptors, thus regulating cellular excitability [8]. Ionic glutamate receptors (iGluRs) are important glutamate receptors and include αamino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA), N-methyl-D-aspartate (NMDA), and kainite receptor subtypes [11,12]. Nmethyl-D-aspartate (NMDA) receptor subtype 2B (NR2B) is the regulatory subunit of the functional NMDA receptor [13], and glutamate receptor 2 (GluR2) is an important component of the AMPA receptor. Previous studies have reported that impairment of signaling via both the AMPA and NMDA receptors is involved in multiple brain pathologies. Overactivation of NMDA receptors may cause calcium overload within a cell, and this may lead to cytotoxicity and subsequently, to neuropathy [14–16]. Glycine (Gly) and γ-aminobutyric acid (GABA) are the main inhibitory neurotransmitters in the central nervous system, and thus regulate neuronal excitation. Gly triggers the opening of the ionic channels coupled to Gly receptors, which induces hyperpolarization in the postsynaptic neuron and inhibits neuronal excitation
Corresponding author. E-mail address: [email protected] (Y.-q. Liu).
https://doi.org/10.1016/j.jinorgbio.2017.11.014 Received 18 July 2017; Received in revised form 7 November 2017; Accepted 17 November 2017 Available online 21 November 2017 0162-0134/ © 2017 Elsevier Inc. All rights reserved.
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5 μM of TPEN to the culture medium. To assess whether Zn2 + supplementation can rescue the effects of zinc deficiency, in the TPEN + ZnSO4 condition, 5 μM of TPEN and 10 μM of ZnSO4 were added to the culture medium.
[17,18]. GABA also inhibits neuronal firing, and has been implicated in the regulation of cognitive functions [19,20]. Reports suggest that in some diseases, cellular apoptosis is closely related to changes in the concentrations of amino acid neurotransmitters [21]. In the present study, we attempted to elucidate the mechanism underlying zinc deficiency-induced neuronal death/apoptosis by evaluating the effects of zinc deficiency and concomitant Zn2 + supplementation on the concentration of amino acid neurotransmitters and the expression of glutamate receptors in cultured neurons.
2.4. Measurement of cell viability using the MTT assay Viability of the cultured primary hippocampal neurons was determined using an MTT assay, which measures the activity of mitochondrial succinate dehydrogenase within viable cells by the reduction of exogenous MTT to insoluble purple formazan crystals. Neurons were plated in 96-well plates at a density of 1 × 105 cells/mL. The volume of medium added to each well was 200 mL. For the assay, MTT solution was added to each well. The final concentration of the MTT solution was 1 mg/mL. Neurons were incubated in this solution at 37 °C for 4 h, following which the medium was discarded and the formazan crystals were dissolved in 150 μL of DMSO. The absorbance of this solution at 570 nm was measured using a Beauty Diagnostic Microplate Reader (Molecular Devices, Sunnyvale, California, USA). The results were expressed in terms of percentage of the absorbance value obtained from the control group.
2. Materials and methods 2.1. Chemicals and animals 4-(2-Hydroxyethyl)-1-Piperazine ethane sulfonic acid (HEPES), Lglutamic acid, and glycine (purity > 99%) were obtained from Genview Scientific Inc. (Tallahassee, USA). Antibiotic solution, poly-Llysine, γ-aminobutyric acid (purity ≥ 99.0%), and N,N,N′,N′-Tetrakis (2-pyridylmethyl) ethylenediamine (TPEN) were obtained from SigmaAldrich (St. Louis, MO, USA). L-Aspartic acid (purity = 99%) was obtained from KisanBiotech Co. Ltd. (Seoul. Korea). 2,4Dinitrofluorobenzene(DNFB)was obtained from Yolne Chemical Co. Ltd. (Shanghai, China). 3-(4,5-Dimethylthiazol-2-yl)-2,5Diphenyltetrazolium bromide (MTT) was obtained from Amresco (Solon, OH, USA). Dulbecco's Modified Eagle's Medium (DMEM)/nutrient mixture F12 with Glutamax™-1 and B27 supplements were obtained from Gibco (Gran Island, NY, USA). Dimethyl sulfoxide was obtained from Dingguo Changsheng Biotechology Co. Ltd. (Beijing, China). The terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling (TUNEL) assay kit and the fixative used in immunohistochemistry were purchased from Biyuntian Co. (Beijing, China). The GluR2 immunohistochemical assay kit [polyclonal rabbitanti GluR2 (N-19), donkey anti-rabbit IgG-TR, etc.] and the NR2B immunohistochemical assay kit [polyclonal goat anti-NMDA NR2B (C-20), donkey anti-goat IgG-FITC, etc.] were purchased from Santa Cruz Biotechnology, Inc. (CA, USA). All animals used for experiments in the present study were obtained from the experimental animal center of the Academy of Military Medical Sciences, and were cared for in accordance with the institutional guidelines for the health and care of experimental animals. The experimental protocols were approved by the Committee on the Ethics of Animal Experiments of Nankai University.
2.5. Measurement of cell apoptosis rate using the TUNEL assay Neurons, at a density of 1 × 105 cells/mL, were grown on sixchamber well slides pre-coated with poly-L-lysine (PLL). After 24 h of treatment, cells were washed thrice with phosphate buffered saline (PBS), following which 2 mL of TUNEL assay solution was added to each well. The cells were then fixed for 30 min, and washed thrice with PBS, then permeabilized with 0.1% Triton X-100 PBS on an ice bath. According to the manufacturer's instructions, cells processed for TUNEL staining were co-stained with 4, 6-diamidino-2-phenylindole (DAPI) for 2 min. The number of TUNEL-positive neurons and the total number of neurons were counted. The percentage of TUNEL-positive cells was calculated and averaged. The results were expressed in terms of the percentage of TUNEL-positive cells. 2.6. Determination of amino acid concentrations using high-performance liquid chromatography (HPLC) The concentrations of 4 free amino acid neurotransmitters (Glu, Asp, Gly, and GABA) were measured in primary neurons and the medium they were cultured in using a CoM6000 HPLC System consisting of two 6000 LDS pumps, a 6000 UV–Vis detector (360 nm), and a Comatex C18 (5 μm, 250 mm × 4.6 mm pore size) (CoMetro Technology, USA). Neurons were cultured at a density of 1.12 × 105 cells/cm2. After treatment with the respective chemical agents, culture medium was separately collected from neurons of all three experimental groups and centrifuged for 10 min at 1000 rpm to remove any remaining cells. Subsequently, acetonitrile was added to the supernatant (1:1, v:v) in order to precipitate proteins, which were separated out after centrifugation at 14,000 rpm for 20 min at 4 °C. The same process was carried out using blank culture media (culture media that was not used to grow any cells). To obtain amino acids from cultured neurons, 100 μL RIPA lysis buffer (50 mM Tris–HCl [pH 7.4], 150 mM NaCl, 1% NP-40) was added to fully lyse the cells. A 5-μL sample of the lysis solution was used for protein quantification using a bicinchoninic acid-protein quantification assay kit by following the manufacturer's instructions. Subsequently, 80 μL of acetonitrile was added to 80 μL of the lysis solution for protein precipitation, and the mixture was centrifuged at 14,000 rpm for 20 min at 4 °C. The sample solutions were derivatized by the addition of 0.1 M carbonate buffer and 2,4-dinitrochlorobenzene (DNCB)-acetonitrile (1:1000, v/v) followed by incubation in a 60 °C water-bath for 60 min in the dark. The derivatization reaction was stopped by the addition of mobile phase B. The mobile phase consisted
2.2. Primary neuronal culture Primary hippocampal neurons were collected from brains of newborn Wistar rats (approximately 1 day old). The hippocampus was isolated from the brain of each rat and treated with 0.125% trypsin for 20 min at 37 °C, triturated in a solution of DMEM/F12 with Glutamax™1 and 15% fetal bovine serum, and centrifuged for 15 min. The cells were suspended in DMEM/F12 medium containing Glutamax™-1 and supplemented with 2% B27, and 1% antibiotic solution. The cells were plated onto cover glasses pre-coated with poly-L-lysine (0.1 mg/mL) in multiwall cell-culture plates at a density of 1.0–5.0 × 105 cells/cm2. Cells were cultured in a humidified incubator (Sanyo, Japan) at 37 °C with 5% CO2 for 7 days, following which they subjected to treatment and further experiments. The medium was changed twice a week by replacing half the volume of culture medium with serum-free DMEM/ F12 medium with Glutamax™-1. 2.3. Experimental groups and TPEN treatment There were 3 experimental groups in the present study: the control group, the TPEN exposure group, and the TPEN exposure plus ZnSO4 treatment (TPEN + ZnSO4) group. Treatment lasted for 24 h. The model of zinc deficiency in primary neurons was established by adding 55
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of two parts: A (an isocratic mixture of acetonitrile and ultrapure water) and B (50 mM sodium acetate buffer solution containing 0.1% trimethylamine, pH 6.4). The elution program was selected to maintain a flow rate of 1.0 mL/min, starting with 75% B for 26 min, then 75% to 32% B for 8 min, 32% to 10% B for 8 min, 10% to 0% B for 5 min, 0% to 75% B for 5 min, and a final retention of 75% B for 2 min. Twenty microliters of the derivative were injected into the HPLC system and separation was performed at 40 °C. 2.7. Measurement of GluR2 and NR2B expression using immunofluorescence Expression of neuronal GluR2 and NR2B was measured using an immunofluorescence assay. After undergoing treatment for 24 h, neurons were fixed for 30 min using an immuno-staining fixation solution and then incubated with 2 mL of 2% bovine serum albumin (BSA) solution for 20 min. After this step, the cells were incubated in 80 μL (per cover glass) of solution containing rabbit-anti GluR2 primary antibody (1:50) at 4 °C overnight. Antibodies were detected using 80 μL (per cover glass) of fluorescein isothiocyanate (FITC)-conjugated donkey anti-goat IgG (1:100) in which the cells were incubated at 37 °C 1 h in a water bath. Finally, the neurons were mounted in fluorescence mounting medium. The prepared slides were viewed using a fluorescence microscope (20 × field). For each experimental group, 6 fieldviews were randomly selected. The sample was stimulated with green light to obtain red fluorescence, and the resulting images were recorded. Similarly, to measure the expression of NR2B, treated neurons were incubated in a solution containing anti-NMDAƐ2 (C-20) (1:50) antibody at room temperature, followed by an incubation in a solution containing FITC-donkey anti-goat IgG secondary antibody (1:100). Unlike in the method used to detect GluR2, a blue light was selected to stimulate the neurons and obtain green fluorescence. Image-Pro Plus6.0 (IPP) software was used to calculate the average optical density (OD).
Fig. 1. Effects of N, N, N′, N′-Tetrakis (2-pyridylmethyl) ethylenediamine (TPEN) and zinc supplement on the neuronal survival rate (n = 8). **P < 0.01 compared with the control group; ##P < 0.01 compared with the TPEN group.
percentage of apoptotic neurons. 3.3. Zinc deficiency leads to changes in amino acid neurotransmitter levels in cultured neurons and the culture media Standard curves for 4 amino acids were plotted using a series of different concentrations (5, 25, 50, 100, 200, and 500 μM for L-glutamate [y = 0.0004x − 1.7566, R2 = 0.9991] and Gly [y = 0.0004x + 1.4421, R2 = 0.9984]; 5, 10, 25, 50, 100, and 200 μM for Asp [y = 0.0005x − 2.4202, R2 = 0.9947] and GABA [y = 0.0004x − 0.8948, R2 = 0.9999]). The HPLC peaks obtained for each of the four amino acids were used to determine the corresponding concentrations of those amino acids based on their respective standard curves. The concentration of the amino acids present in the treatment medium subtracted from that in blank culture medium represents the quantity of amino acids released from the cultured neurons. The concentrations of intracellular and extracellular amino acids were normalized to the concentration of amino acids per gram of cellular protein. Fig. 3A depicts the effect of 24 h of TPEN-treatment (zinc deficiency) on intracellular concentrations of Asp, Glu, Gly, and GABA. Concentrations of Glu, Gly, and GABA were significantly lower in the TPEN-treated cells than in the controls, but the TPEN-treated cells showed a tendency to have a higher concentration of Asp. Zn2 + supplementation could rescue the observed TPEN-induced decrease in Glu, Gly, and GABA concentration, but the concentration of Asp did not show much change. Fig. 3B depicts the effect of 24 h of zinc deficiency on the secreted amounts of Asp, Glu, Gly, and GABA. TPEN treatment led to a significant increase in the concentrations of Gly and Glu in medium used to culture neurons (P < 0.05), whereas the levels of Asp and GABA (P < 0.05) showed a significant decrease. Concurrent treatment with Zn2 + could rescue the TPEN-induced change in levels of Gly and GABA, but had obvious effects on Asp and Glu levels.
2.8. Data analysis SPSS 20 and Origin 8.5 were used for data treatment and analysis, and the results were presented as the mean ± standard error of the mean (SEM). A one-way ANOVA was performed for the comparison of multiple groups, and a value of P < 0.05 was considered as statistically significant; a value of P < 0.01 was considered as extremely statistically significant. 3. Results 3.1. Zinc deficiency reduces neuronal viability Following pilot experiments using MTT, 5 μM TPEN and 10 μM ZnSO4 were determined as optimum concentrations to induce zinc deficiency and provide Zn2 + supplementation, respectively, and were used in all subsequent experiments. The viability of neurons following 24 h of treatment was determined. As shown in Fig. 1, the viability rate of the cultured cells after TPEN treatment was significantly lower (73.7 ± 3.0%, P < 0.01) than that of the controls. In contrast, application of ZnSO4 could rescue the TPEN-induced reduction of viability (TPEN + ZnSO4 vs. TPEN, P < 0.01), and this rescue was statistically significant.
3.4. Zinc deficiency alters GluR2 and NR2B expression in cultured neurons
3.2. Zinc deficiency causes an increase in neuronal apoptosis
Fig. 4 depicts the effects of zinc deficiency on GluR2 expression. Treatment with TPEN caused a significant reduction in GluR2 expression (P < 0.01, compared with control), whereas TPEN + ZnSO4 treatment rescued this TPEN-induced reduction (P < 0.01, compared with TPEN). The effects of TPEN on NR2B expression are shown in Fig. 5. TPEN treatment led to a significant increase in neuronal NR2B expression (P < 0.05, compared with control). Concurrent ZnSO4 treatment could bring the NR2B expression back to the levels observed
Fig. 2 reveals the effect of zinc deficiency on neuronal apoptosis. Results show that the apoptosis rate was higher in neurons that underwent TPEN treatment than in those that did not (P < 0.01). In neurons treated with TPEN + ZnSO4, the apoptosis rate was lower than in neurons treated with only TPEN (P < 0.01). These results suggest that TPEN-induced zinc deficiency causes a significant increase in the 56
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Fig. 2. Effects of N, N, N′, N′-Tetrakis (2-pyridylmethyl) ethylenediamine (TPEN) and zinc supplement on the neuronal apoptosis rate (n = 8). (A). Representative images of immunofluorescence stained neurons after different treatments; scale bars = 50 μm. (B). The neuronal apoptosis rate following different treatments **P < 0.01 compared with the control group; ##P < 0.01 compared with the TPEN group.
changes in the concentration of amino acid neurotransmitters, and altered expression of two glutamate receptor subtypes, GluR2 and NR2B. These results also show that Zn2 + supplementation reversed these changes. A previous study has demonstrated that zinc deficiency causes intracellular oxidative stress, neuronal damage, and apoptosis [6]. It has been demonstrated that zinc deficiency can induce apoptosis of MC3T3E1 cells through a mitochondria-mediated pathway [22]. The zinc deficiency-induced apoptosis of cells may involve several pathways. Zinc is involved in the metabolism of free amino acid neurotransmitters, and zinc deficiency causes an increase in the glutamate concentration in hippocampal neurons, which increases the excitability of these cells
in the control (P < 0.01, compared with TPEN), thus reversing the change caused by TPEN treatment. 4. Discussion Zinc plays an important role in regulating the physiological functions of neurons. As an important neuromodulator, it can regulate neurotransmitter release. Perturbation of zinc homeostasis in the brain leads to disturbances in cognitive function and may cause neurological disorders. In the present study, TPEN was used to establish a model of zinc deficiency in cultured neurons. These results show that zinc deficiency led to lower rates of neuronal survival, higher rates of apoptosis,
Fig. 3. Effects of N, N, N′, N′-Tetrakis (2-pyridylmethyl) ethylenediamine (TPEN) and zinc supplement on the concentration of amino acids concentration in neurons and culture medium (n = 6). (A) The free amino acid concentration in neurons following different treatments. (B) The free amino acid concentration in the culture medium following different treatments. *P < 0.05, **P < 0.01 compared with the control group; #P < 0.05, ##P < 0.01 compared with the TPEN group.
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Fig. 4. Effects of N, N, N′, N′-Tetrakis (2-pyridylmethyl) ethylenediamine (TPEN) and zinc supplement on GluR2 expression in neurons (n = 8). (A). Representative images of GluR2 immunofluorescence staining in neurons following different treatments; scale bars = 50 μm. (B). The optical density (OD) of GluR2 immunofluorescence in neurons following different treatments. **P < 0.01 compared with the control group; ##P < 0.01 compared with the TPEN group.
Fig. 5. Effects of N, N, N′, N′-Tetrakis (2-pyridylmethyl) ethylenediamine (TPEN) and zinc supplement on NR2B expression in neurons (n = 8). (A). Representative images of NR2B immunofluorescence staining in neurons following different treatments; scale bars = 50 μm. (B). The optical density (OD) of NR2B immunofluorescence in neurons following different treatments. **P < 0.01 compared with the control group; ##P < 0.01 compared with the TPEN group.
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DMEM TUNEL
Dulbecco's Modified Eagle's Medium terminal deoxynucleotidyl transferase-mediated dUTP nickend labeling DMSO dimethyl sulfoxide PLL poly-L-lysine PBS phosphate buffered saline DAPI 4,6-diamidino-2-phenylindole HPLC high-performance liquid chromatography RIPA lysis buffer Radio Immunoprecipitation Assay BSA bovine serum albumin FITC fluorescein isothiocyanate IgG Immunoglobulin G
[23]. Our experimental results show that exposure to TPEN causes a significant reduction in the glutamate concentration and an increase in aspartic acid levels in cultured primary neurons, whereas Zn2 + supplementation could reverse these changes to a certain extent. In addition, TPEN-treatment caused a significant increase in the Glu concentration of the culture medium, which suggests that large amounts of glutamate were released from the neurons. Furthermore, providing additional Zn2 + to these neurons partially reversed the change induced by zinc deficiency, indicating that zinc deficiency might influence Glu metabolism, resulting in toxic neuronal excitability. In addition, TPENtreatment led to a significant reduction in Gly and GABA levels in neurons, and simultaneous addition of Zn2 + significantly reversed these changes. This could be a result of TPEN reducing the synthesis of GABA in the culture medium; the implications of these findings warrant further studies for validation. Amino acid neurotransmitters usually bind to specific receptors to perform their physiological functions. Previous studies have shown that glutamate induces cell excitability by binding to ionic glutamate receptors [24], such as NMDA and AMPA receptors [25,26]. The NR2 subunit plays an important role in regulating physiological functions of the NMDA receptor [11,27]. The GluR2 subunit of the AMPA receptor determines the intracellular calcium permeability of the AMPA receptor [2]. The results of the present study indicate that TPEN-treatment leads to an increase in the expression of the NR2B subunit of the NMDA receptor. In addition, TPEN-treatment leads to a reduction in the expression of the GluR2 subunit of the AMPA receptor. We know that the activation of the NMDA receptor and the depression of the AMPA receptor together cause a large Ca2 + influx, excitatory cytotoxicity and neuronal apoptosis. In addition, we know that the role of redox active zinc in neurons is tightly linked to the role of redox active copper via the enzyme copper–zinc superoxide dismutase [28,29], a homodimeric metalloprotein that binds copper and zinc ions and forms an intramolecular disulfide bond. It has been reported that copper–zinc superoxide dismutase functions as a molecular switch that activates the endoplasmic reticulum stress response, which plays an important role in cellular homeostasis under zinc-deficient conditions [29]. Therefore, neuronal death/apoptosis induced by zinc deficiency might involve changes in copper–zinc superoxide dismutase and cupric ion concentrations. However, this has not been addressed in the present study and will need further investigation to be validated. In conclusion, neuronal death induced by zinc deficiency may be associated with increased neuronal glutamate release, down-regulation of GluR2, and up-regulation of NR2B.
Author contributions Conception and design: Yan-qiang Liu, Kun Tian. Acquisition of data: Kun Tian, Yu-xiang Wang, Li-xia Li. Analysis and interpretation of data: Kun Tian. Writing, review of the manuscript: Kun Tian, Yan-qiang Liu. Conflict of interest The authors declare that they have no conflict of interest. Acknowledgements This work was supported by the National Natural Science Foundation of China (No. 31272317) and the Natural Science Foundation of Tianjin City (15JCYBJC24500) sanctioned to YQL. References [1] R.J. Radford, S.J. Lippard, Curr. Opin. Chem. Biol. 17 (2013) 129–136. [2] Z. Liu, Y.Y. Huang, Y.X. Wang, H.G. Wang, F. Deng, B. Heng, L.H. Xie, Y.Q. Liu, J. Trace Elem. Med. Biol. 31 (2015) 45–52. [3] Y. Song, S.W. Leonard, M.G. Traber, E. Ho, J. Nutr. 139 (2009) 1626–1631. [4] S.W. Suh, S.J. Won, A.M. Hamby, B.H. Yoo, Y. Fan, C.T. Sheline, H. Tamano, A. Takeda, J. Liu, J. Cerebr. Blood F. Met. 29 (2009) 1579. [5] W. Pang, H. Lu, Y.D. Hu, H.P. Yang, X. Leng, Y.G. Jiang, Nutr. Neurosci. 15 (2013) 18–24. [6] M.P. Zago, G.G. Mackenzie, A.M. Adamo, C.L. Keen, P.I. Oteiza, Antioxid. Redox Signal. 7 (2005) 1773–1782. [7] A. Miszke, K. Rapacz, Otolaryngol. Pol. 45 (1991) 374–380. [8] R.S. Sundaram, L. Gowtham, B.S. Nayak, Asian J. Pharm. Clin. Res. 5 (2012) 1–7. [9] M.V. Johnston, MRDD. Res. Rev. 7 (2001) 229–234. [10] D.W. Choi, J. Neurobiol. 23 (1992) 1261–1276. [11] X.L. Ma, F. Zhang, Y.X. Wang, C.C. He, K. Tian, H.G. Wang, D. An, B. Heng, Y.Q. Liu, Chem. Biol. Interact. 254 (2016) 73. [12] M. Hollmann, M. Hartley, S. Heinemann, Science 252 (1991) 851–853. [13] S. Cull-Candy, S. Brickley, M. Farrant, Curr. Opin. Neurobiol. 11 (2001) 327–335. [14] R.P. Simon, J.H. Swan, T. Griffiths, B.S. Meldrum, Science 226 (1984) 850–852. [15] B. Liu, M. Liao, J.G. Mielke, K. Ning, Y. Chen, L. Li, Y.H. El-Hayek, E. Gomez, R.S. Zukin, M.G. Fehlings, J. Neurosci. 26 (2006) 5309–5319. [16] M.S. Beattie, A.R. Ferguson, J.C. Bresnahan, Eur. J. Neurosci. 32 (2010) 290–297. [17] M.S. Hernandes, L.R.P. Troncone, J. Neural Transm. 116 (2009) 1551–1560. [18] M.S. Hernandes, M.L. De, L.R. Troncone, Brain Res. 1168 (2007) 32–37. [19] J.L. Barker, Science 240 (1988) 548–549. [20] J.E. Walker, Neurochem. Res. 8 (1983) 521–550. [21] Y.H. Sun, Y. Yue, Y. Wang, J. Apoplexy Nervous Dis. 22 (2005) 344–347. [22] B. Guo, M. Yang, D. Liang, L. Yang, J. Cao, L. Zhang, Mol. Cell. Biochem. 361 (2012) 209–216. [23] J.C. Wallwork, D.B. Milne, R.L. Sims, H.H. Sandstead, J. Nutr. 113 (1983) 1895–1905. [24] L. Kiedrowski, J. Neurochem. 130 (2014) 87–96. [25] N. Armstrong, E. Gouaux, Neuron 28 (2000) 165–181. [26] R. Gill, A.A. Alanine, B. Buttelmann, G. Fischer, M.P. Heitz, J.N. Kew, T.B. Levet, H.P. Lorez, P. Malherbe, M.T. Miss, J. Pharmacol. Exp. Ther. 302 (2002) 940–948. [27] G. Köhr, Cell Tissue Res. 326 (2006) 439–446. [28] E. Tokuda, I. Anzai, T. Nomura, K. Toichi, M. Watanabe, S. Ohara, S. Watanabe, K. Yamanaka, Y. Morisaki, H. Misawa, Y. Furukawa, Mol. Neurodegener. 12 (2017) 2–18. [29] K. Homma, T. Fujisawa, N. Tsuburaya, N. Yamaguchi, H. Kadowaki, K. Takeda, H. Nishitoh, A. Matsuzawa, I. Naguro, H. Ichijo, Mol. Cell 52 (2013) 75–86.
Abbreviations TPEN AMPA GluR2
N,N,N′,N′-Tetrakis (2-pyridylmethyl) ethylenediamine α-amino-3-hydroxy-5-methyl-4-isoxazolepropionate α-amino-3-hydroxy-5-methyl-4-isoxazolepropionate (AMPA) receptor NMDA N-methyl-D-aspartate NMDAR N-methyl-D-aspartate receptor NR2B N-methyl-D-aspartate (NMDA) receptor subtype 2B Asp aspartic acid Glu glutamate Gly glycine GABA gamma-aminobutyric acid ZnSO4 zinc sulfate iGluR ionic glutamate receptors HEPES 4-(2-hydroxyethyl)-1-piperazine ethane sulfonic acid DNFB 2,4-dinitrofluorobenzene MTT 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide
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