Contribution of Zinc-Dependent Delayed Calcium Influx via TRPC5 in Oxidative Neuronal Death and its Prevention by Novel TRPC Antagonist
Oxidative stress is a key mediator of neuronal death in acute brain injuries, such as epilepsy, trauma, and stroke. Although it is accompanied by diverse cellular changes, increases in levels of intracellular zinc ion (Zn2+) and calcium ion (Ca2+) may play a critical causative role in oxidative neuronal death. However, the mechanistic link between Zn2+ and Ca2+ dyshomeostasis in neurons during oxidative stress is not well-understood. Here, we show that the exposure of cortical neurons to H2O2 led to a zinc-triggered calcium influx, which resulted in neuronal death. The cyclin-dependent kinase inhibitor, NU6027, inhibited H2O2-induced Ca2+ increases and subsequent cell death in cortical neurons, without affecting the early increase in Zn2+. Therefore, we attempted to identify the zinc-regulated Ca2+ pathway that was inhibited by NU6027. The expression profile in cortical neurons identified transient receptor potential cation channel 5 (TRPC5) as a candidate that is known to involve in the generation of epileptiform burst firing and epileptic neuronal death (Phelan KD et al. 2012a; Phelan KD et al. 2013b). NU6027 inhibited basal and zinc-augmented TRPC5 currents in TRPC5-overexpressing HEK293 cells. Consistently, cortical neurons from TRPC5 knockout mice were highly resistant to H2O2-induced death. Moreover, NU6027 is neuroprotective in kainate-treated epileptic rats. Our results demonstrate that TRPC5 is a novel therapeutic target against oxidative neuronal injury in prolonged seizures and that NU6027 is a potent inhibitor of TRPC5.
KeywordsCa2+ H2O2 NU6027 Seizure TRPC Zn2+
Reactive oxygen species (ROS) play important pathological roles in numerous neurological disorders, such as seizure, ischemic stroke, and brain and spinal cord trauma [1, 2, 3]. Because the brain consumes a considerable amount of oxygen and contains a high concentration of polyunsaturated fatty acid that is easily oxidized, it is particularly susceptible to oxidative stress . During oxidative stress, the concentration of intracellular calcium ions ([Ca2+]i) gradually increases, leading to neuronal death. A variety of Ca2+ channels are involved in the elevation of [Ca2+]i during neuronal injuries, including ionic glutamate receptors [N-methyl-d-aspartate (NMDA), alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA), and kainic acid (KA) receptors], metabotropic glutamate receptors, voltage-dependent calcium channels, and the transient receptor potential (TRP) channel family in plasma membrane as well as inositol trisphosphate (IP3) receptors and ryanodine receptors in endoplasmic reticulum . Although the overactivation of glutamate receptors induces increases in [Ca2+]i during neuronal death, TRP channels may also mediate oxidative stress-induced increases in [Ca2+]i in case of brain injuries .
Mammalian TRP channels belong to a family of Ca2+-permeable nonselective cationic channels . TRP channels are further grouped into seven subfamilies: TRPC, TRPM, TRPV, TRPA, TRPP, TRPML, and TRPN. These TRP channels have been implicated in many physiological events, including development and neuroplasticity . TRP channels also play important roles in neuronal death, such as capsaicin-triggered TRPV1 activation in mesencephalic dopaminergic neuronal death , amyloid β- and H2O2-induced TRPM2 activation in striatal cell death , and ROS-mediated TRPM7 activation in ischemic neuronal injury . Recently, there has been an increasing interest in TRPC1/4/5 in epileptogenesis and neuronal death. TRPC channels comprise seven isotypes (TRPC1–TRPC7). Of these, TRPC4 and TRPC5 are abundantly expressed in the brain  and may be involved in epileptiform burst firing and epileptic neuronal death [13, 14]. In addition, S-glutathionylation of TRPC5 and downregulation of TRPC1 during oxidative stress may be involved in neuronal death in Huntington’s disease . Despite some scattered evidence, the role of TRPCs in neuronal death has not drawn much attention.
Zinc ions (Zn2+) also play physiological and pathological roles in the central nervous system. Under physiological conditions, intracellular Zn2+ is tightly regulated by zinc transporters (ZnTs), ZRT, IRT-like proteins, and buffering proteins, such as metallothioneins . However, excessively high levels of intracellular free Zn2+ in seizures, stroke, or trauma trigger neuronal death . Interestingly, the two potentially toxic events, Ca2+ and Zn2+ dyshomeostasis, are correlated. Several papers have suggested that increases in intracellular Zn2+ contribute to the subsequent increase in Ca2+ [18, 19]. Moreover, increasing intracellular Zn2+ by clioquinol and pyrithione, Zn2+-ionophores can activate TRPA1 channels . Hence, initial Zn2+ dyshomeostasis may trigger Ca2+ dyshomeostasis, which together causes neuronal death under injuries.
Although ample evidence supports that oxidative stress is a key mechanism contributing to neuronal death in acute brain injury, a variety of clinical trials with drugs targeting ROS have been unsuccessful . For instance, potent antioxidants, such as N-acetyl cysteine and NXY-059, were not beneficial in patients with epilepsy or ischemic stroke [22, 23]. There are many possible reasons for these failures, including weak antioxidant capacity, poor blood–brain barrier penetration, and rapid clearance in vivo . Despite these failures, ROS are major targets for neuroprotective drugs, and new insights into the toxic mechanisms of oxidative injury are required.
While searching for neuroprotective drugs effective against oxidative stress-induced cell death, we found that NU6027 showed marked protective effects. NU6027 blocks neuronal death induced by H2O2 and KA in primary cortical cultures and in a rat seizure model, respectively, via a novel mechanism, the inhibition of TRPC5.
Materials and Methods
Primary Mouse Cortical Cell Cultures
Pure astrocyte cultures were prepared from postnatal day 3 (P3) ICR mice and maintained in Dulbecco’s modified Eagle’s medium (Gibco) supplemented with 5% fetal bovine serum (Hyclone), 5% horse serum (Gibco), 2 mM glutamine (Sigma), and 1% penicillin/streptomycin (Cambrex). These astrocyte cultures were used for experiments or as feeder cells for mixed cortical cultures. Mixed cortical cultures were prepared by plating cortical neurons from embryonic day 14 (E14) ICR mice onto pure astrocyte cultures and growing them in growth media. Pure neuronal cultures were prepared from cortices of E14 ICR mice or age-matched wild-type (WT) or TRPC5 knock-out (KO) 129/SvImJ mice and were grown in neurobasal media (Gibco) containing B27 supplement (Gibco), 2 mM glutamine, and antibiotics.
Exposure to H2O2 and Other Reagents
Cells were exposed to glutamate, H2O2 (Sigma), sodium nitroprusside (SNP, a donor of nitric oxide, Sigma), and ZnCl2 (Sigma) in minimum essential media (Gibco) for 24 h at the indicated concentrations to induce cell death. Anthranilic acid, clotrimazole, dantrolene, flufenamic acid, kenpaullone, NU6027, olomoucine, roscovitine, ruthenium red, SU9516, N,N,N,N′-tetrakis (2-pyridylmethyl) ethylenediamine (TPEN), and 2-aminoethyl diphenylborinate (2-APB) were purchased from Sigma. Capsaicin, MK-801, ML204, Pyr3, and (−)-Xestospongin C (XeC) were purchased from Tocris. SB216763 was purchased from Enzo Life Science, and 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX) was purchased from RBI. All reagents were added 1 h prior to H2O2 exposure, unless otherwise stated.
Assessment of Cell Death
Lactate dehydrogenase (LDH) released into culture media from damaged cells was measured to evaluate cell death . The mean background value in control sister cultures that only received a sham wash (0% cell death) was subtracted from the LDH value in each test condition, and the LDH value was scaled to the mean value of sister cultures after 24 h of exposure to 200 μM glutamate, which resulted in nearly complete neuronal death without astrocytic damage (100%). Cell death was also detected by staining with 2 μg/ml propidium iodide (PI, Sigma) at 37 °C in a CO2 incubator for 10 min. Images were obtained using a fluorescent microscope (Olympus, IX71) equipped with a CCD camera (Olympus, IX-10) at a wavelength of 535/617 nm (ex/em).
Live Cell Imaging for Zn2+ and Ca2+
To label Zn2+, pure neuronal cultures were treated with 2.5 μM FluoZin-3 AM (Invitrogen) for 30 min before imaging. To detect Ca2+, cultures were transfected 2 days before experiments with 10 μg of pCMV-RGECO1 plasmid (R-GECO1, a genetically encoded Ca2+ indicator) using Lipofectamine 2000 (Invitrogen). Live cell images were obtained using an inverted fluorescent microscope (Ti-E; Nikon) equipped with a Cascade 212B camera (Roper Scientific), and images were acquired every 1 min at wavelengths of 494/516 nm and 570/595 nm (ex/em) for FluoZin-3 AM and R-GECO1, respectively. Images of Ca2+ staining were also obtained by staining cells with 2 μM Fluo-4 AM (Invitrogen) for 30 min. The images were obtained with an inverted fluorescent microscope (Olympus, IX71) equipped with a CCD camera (Olympus, IX-10) at a wavelength of 494/506 nm (ex/em). Fluorescence intensity was analyzed by Image J software and represented as the fold increase compared with control.
Reverse Transcription-Polymerase Chain Reaction (RT-PCR)
List of primers used for RT-PCR
Forward primer (5′ → 3′)
Reverse primer (5′ → 3′)
Cultured pure neurons and astrocytes were fixed with 4% paraformaldehyde for 15 min, permeabilized with 0.1% Triton X-100 for 5 min, and blocked with 1% bovine serum albumin for 30 min. For staining, cells were incubated with antibodies for glial fibrillary acidic protein (Millipore, AB5807), MAP2 (Abcam, AB32454), and TRPC5 (Neuromab, 75-104) at 4 °C overnight, followed by incubation with fluorescence-labeled secondary antibodies for 2 h. For nuclear staining, cells were incubated with 5 μg/ml Hoechst 33342 (Sigma). Cells were mounted and visualized under the EVOS Cell Imaging System (Thermo Fisher Scientific).
Membrane proteins from cultured pure neurons and astrocytes were extracted with lysis buffer (20 mM Tris-HCl pH 7.4, 150 mM NaCl, 1 mM EDTA, 0.5% SDS, 2.5 mM sodium pyrophosphate, 1 μM NaVO4, and protease inhibitors). Proteins were separated by 8% SDS-polyacrylamide gel electrophoresis and transferred to polyvinylidene difluoride membranes (Millipore). Membranes were probed with antibodies to TRPC5 (Neuromab, 75-104) and β-actin (Sigma, A0560), followed by incubation with the appropriate secondary antibody conjugated to horseradish peroxidase (Thermo Fisher Scientific). Immunoreactivity was visualized using Immunobilon™ Western Chemiluminescent HRP Substrate (Millipore) and the Kodak Image Station 4000MM (Kodak).
For TRPC5 current recordings, human embryonic kidney (HEK) 293 cells (ATCC) were maintained according to the manufacturer’s recommendations and transfected with plasmid DNA expressing mouse TPRC5 (pIRES-mTRPC5-GFP) using FuGENE6 (Roche). Whole-cell currents were recorded using an Axopatch 200B amplifier (Axon Instruments). Currents were filtered at 5 kHz (3 dB, 4-pole Bessel), digitized using a Digidata 1440A Interface (Axon Instruments), and analyzed using a personal computer equipped with pClamp 10.2 software (Axon Instruments) and Origin software (Microcal Origin v. 8.0). Patch pipettes were made from borosilicate glass and had resistances of 2–4 MΩ when filled with standard intracellular solutions. We used an external bath medium (normal Tyrode solution) of the following composition: 135 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 10 mM glucose, and 10 mM N-[2-hydroxyethyl]piperazine-N′-[2-ethanesulfonic acid] (HEPES), with pH adjusted to 7.4 using NaOH. A Cs+-rich external solution was made by replacing NaCl and KCl with equimolar CsCl. The standard pipette solution contained 140 mM CsCl, 10 mM HEPES, 0.2 mM Tris-GTP, 0.5 mM EGTA, and 3 mM Mg-ATP, with the pH adjusted to 7.3 using CsOH. After TRPC5 activation in a Cs+-rich solution, 10 μM NU6027 was externally applied at the time indicated by the bars. ZnCl2 was intracellularly applied via the pipette solution. Voltage ramp pulses were applied from + 100 to − 100 mV for 500 ms at a holding potential of − 60 mV. The junction potential between the pipette and bath solutions used for all cells during sealing was calculated to be 5 mV (pipette negative) using pClamp 10.2 software. No junction potential correction was applied. Experiments were performed at room temperature (18 °C–22 °C). Cells were continuously perfused at a rate of 0.5 ml/min. The inward current amplitudes of all bar graphs and current traces were taken during the ramp pulses at a holding potential of − 60 mV.
Breeding and Genotyping of TRPC5 KO Mice and Pure Neuronal Cultures
List of primers for genotyping WT and TRPC5 KO mice
Forward primer (5′ → 3′)
Reverse primer (5′ → 3′)
Animal Care and Seizure Induction
All animal experiments were approved by the Institutional Animal Care and Use Committee and followed a protocol approved by the Asan Medical Center. Adult male Sprague Dawley (SD) rats (8-week old, 240–270 g) were maintained under 12 h light/dark cycles. Seizures were induced by intraperitoneal injection of 10 mg/kg KA (Tocris) dissolved in saline. Animals were intraperitoneally injected with 100 μg/kg NU6027 or vehicle (10% DMSO in normal saline) 30 min after KA injection. Two hours after KA injection, seizure behavior was staged according to the classification system described by Zheng et al. . After 2.5 h, seizures were stopped by an intraperitoneal injection of 50 mg/kg sodium phenytoin. Body weight and mortality were determined 24 h later.
Tissue Preparation and Cresyl Violet and Fluoro-Jade B Staining
Brains were harvested 24 h after KA injection and immediately frozen in dry ice. Coronal brain sections were cut using a cryostat microtome and fixed with 4% paraformaldehyde for 30 min. Neurons were stained with a 1% cresyl violet solution at room temperature for 10 min. To determine cell death, sections were immersed in 6% potassium permanganate for 5 min, followed by 30 min of incubation with 0.001% Fluoro-Jade B (FJB) solution (Histo-Chem Inc.). The numbers of cresyl violet and Fluoro-Jade B positive cells in the hippocampus, pyriform cortex, and thalamus were counted from both hemispheres in a total of 5 coronal sections, every 150 μm starting 2.8 mm from the bregma. Images were obtained using a fluorescent microscope (BX60; Olympus) equipped with a DP70 CCD camera (Olympus) at a wavelength of 480/525 nm (ex/em) under × 10 objective.
Experimental Design and Statistical Analysis
For all in vivo experiments, at least eight male animals were used. Two-tailed Student’s t test was used for statistical analysis. For all in vitro experiments, data analysis was performed using the Sigmaplot version 13.0 statistical package programs (SIGMASOFT). The number of replicates is three unless others state. Data is represented as mean ± SEM from three independent experiments performed in triplicate. Statistical analyses were performed using the unpaired 2-tailed Student’s t test for comparisons between two groups, and one-way ANOVA was used for comparisons of multiple groups. Data were considered significant at a p value of < 0.05. The degrees of freedom and p values are reported in the results section for each experiment.
Cyclin-Dependent Kinase (CDK) Inhibitor, NU6027, Reduces Oxidative Stress-Induced Cell Death in Neurons
NU6027 Prevents Zinc-Dependent Elevation of Calcium by H2O2
NU6027 Inhibits H2O2-Triggered Ca2+ Influx and Death by Antagonizing TRPC5
NU6027 Ameliorates Cell Death Induced by Kainate in Rat Seizure Model
The main findings of this study are that neuronal death induced by oxidative stress, such as H2O2, is mediated by early transient increases in Zn2+ and delayed prolonged increases in Ca2+ and that these events are mechanistically linked. The increase in free Zn2+ levels appears to be necessary for the delayed accumulation of Ca2+ because the addition of a Zn2+-specific chelator, TPEN, while Zn2+ levels are increasing, completely blocks the Ca2+ influx. Hence, it is likely that delayed and prolonged Ca2+ influx is the final effector of cell death in cases of oxidative neuronal injury, whereas transient Zn2+ increases are critical for switching cells to Ca2+ entry.
Several lines of evidences from our results suggest that the delayed and prolonged influx of Ca2+ into cultured cortical neurons is mediated by TRPC5. First, among tested inhibitors of intracellular Ca2+ increases, only 2-APB and ML204 were effective in curtailing Ca2+ increases as well as cell death following H2O2 exposure. Although 2-APB is a broad-spectrum inhibitor of TRP class channels, ML204 is a specific inhibitor of TRPC4 and TRPC5 channels. Furthermore, neither inhibitors of TRPM channel, anthranilic acid, flufenamic acid, and clotrimazole, nor an inhibitor of TRPV, ruthenium red, showed any effect at all, making it unlikely that these TRP channels are involved. Second, NU6027, a CDK inhibitor, which blocked the delayed increase in Ca2+ and neuronal death induced by H2O2, directly reduced TRPC5-mediated channel currents in the channel-transfected HEK293 cells. It is unlikely that CDK inhibition is the underlying mechanism for this effect because other more potent CDK inhibitors did not have similar effects. Third, TRPC5 was selectively expressed in neurons but not in astrocytes in primary cortical cultures, which is consistent with the selective protection of zinc-exposed neurons by NU6027. Finally, knockout of TRPC5 was sufficient to reduce the delayed Ca2+ influx and cell death induced by H2O2 in cortical neurons.
TRPC channels belong to a family of TRP channels. TRP channels have many cysteine and histidine residues of which modifications activate TRP channels. For example, nitrosylation of Cys553/Cys558 residues in pore-forming region activates TRPC5, TRPV1, TRPV3, and TRPV4 . Glutathionylation of intracellular N-terminal Cys176/Cys178 activates TRPC5 . Another notable feature of TRP channels is that they can be regulated by metal ions, including Zn2+ . Intracellular Zn2+ activates TRPA1 by modulating specific intracellular Cys641, Cys1021, and His983 . It was also reported that extracellular Zn2+ activates TRPV1, though precise sites of modification are not elucidated in this report . Although detailed information regarding the gating mechanism for TRPC5 is not yet available, our results suggest the intriguing possibility that increase in intracellular Zn2+ contributes to activation of TRPC5. It should be examined in the future whether Zn2+ directly participates in the gating of TRPC5, as occurs in that of TRPA1 in which the binding of Zn2+ to cytosolic cysteine and histidine residues is responsible for gating.
Mechanisms involved in oxidative neuronal cell death have been investigated intensively over the last several decades. Although Ca2+ overload is a major ionic mechanism mediating cell death, Zn2+ dyshomeostasis has been proposed as an additional mechanism later. During oxidative stress, Zn2+ binding proteins, most notably metallothioneins, can release Zn2+ [39, 40]. Prolonged Zn2+ dyshomeostasis can activate cellular processes, such as mitochondrial damage, NADPH oxidase and nitric oxide synthase activation, PARP activation, and lysosomal membrane permeabilization, which eventually lead to cell death [41, 42, 43]. In the present study, we observed that even transient increases in free Zn2+ levels were a prerequisite for delayed and prolonged Ca2+ influx, as discussed above. Hence, even in cases where Zn2+ dyshomeostasis is not severe enough to cause cell death by itself, Zn2+ may still play a large role in oxidative neuronal cell death by permitting a large Ca2+ influx though TRPC channels.
TRPC channels are subdivided into two groups based on sequence homology and functional properties. One is TRPC1/4/5 and the other is TRPC3/6/7 of which homo- or hetero-tetramer can be regulated by receptor stimulation . Homomeric TRPC5 channel has been implicated in pathological roles for seizure and excitotoxicity . In TRPC5 KO mice, seizure and neuronal cell death induced by pilocarpine was significantly reduced . In cortical lesions of the focal cortical dysplasia, common intractable epilepsy in both pediatric and adult patients, the expression of TRPC5 is significantly increased in glutamatergic and GABAergic neurons . Consistently, in our results, TRPC5 mRNA and protein were predominantly expressed in neurons than astrocytes, cell death was dependent to the expression of TRPC5, and NU6027, an inhibitor of TRPC5, reduced neuronal death in cortical cultures. Our results also showed that NU6027 decreased cell death and mortality in a kainate model of epileptic brain damage. TRPC5 plays pathophysiological roles in other diseases, for example pain and anxiety, diabetic nephropathy, cardiovascular disease, rheumatoid arthritis, and cancer [26, 47, 48, 49, 50]. The pharmacological tools available to unveil its pathophysiological activities are limited. Small-molecular inhibitors, such as SKF-93635 and 2-APB, nonspecifically inhibit all TRPC channels and other ion channels . ML204 and the anti-histamine clemizole hydrochloride have a higher selectivity for TRPC4 than TRPC5 and inhibit channels at micromolar concentrations . Pico145, a recently reported TRPC1/4/5-specific inhibitor with picomolar range of potency, has not been verified its inhibitory activity in in vivo systems . Therefore, there is a pressing need for potent and specific inhibitory tool compounds. Our results suggest that NU6027 is a useful template in designing an effective inhibitor of TRPC5.
In conclusion, our results demonstrate that oxidative stress-induced neuronal cell death involves Zn2+-triggered delayed Ca2+ increases in neurons through TRPC5. NU6027 may directly block TRPC5-mediated Ca2+ influx in a CDK-independent manner. The time course of increase in Ca2+ suggests that inhibitors of TRPC5 have neuroprotective effects even when administered at later stages of acute neuronal injuries, such as epilepsy.
We thank the optical imaging core facility at the ConveRgence mEDIcine research cenTer (CREDIT), Asan Medical Center for support and instrumentation.
This research was supported by a Grant from the Korea Health Technology R&D Project through the Korea Health Industry Development Institute funded by the Ministry of Health & Welfare (HI14C1913, HI15C0527); Basic Science Research Program (2017R1A2B2005633) and Global PhD Fellowship Program (2015H1A2A1034032) through the National Research Foundation of Korea funded by the Ministry of Science, ICT, & Future Planning; and the Ministry of Education.
Compliance with Ethical Standards
Conflict of Interest
The authors declare that they have no conflict of interests.
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