Glutamate Excitotoxicity Is Involved in Cell Death Caused by Tributylt
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《毒物学科学杂志》
Graduate School of Biomedical Sciences, Hiroshima University, 1–2–3, Kasumi, Minami-ku, Hiroshima, 734-8551, Japan
Center for Quantum Life Sciences, Hiroshima University, 1–2–3, Kasumi, Minami-ku, Hiroshima, 734-8551, Japan
Graduate School of Pharmaceutical Sciences, Kyoto University, 46–29, Yoshida-Shimoadachi-cho, Sakyo-ku, Kyoto, 606-8501, Japan
ABSTRACT
Tributyltin, an endocrine-disrupting chemical, has been used as a heat stabilizer, agricultural pesticide, and component of antifouling paints. In this study, the neurotoxicity of tributyltin was investigated in cultured rat cortical neurons. Tributyltin caused marked time- and dose-dependent increases in the number of trypan blue–stained cells. Measurement of extracellular glutamate concentration showed that glutamate release was induced by tributyltin. Application of the glutamate receptor antagonists MK-801 and CNQX decreased the neurotoxicity. These results suggest that released glutamate and glutamate receptors are involved in tributyltin toxicity. Next, we examined whether various factors, believed to be involved in glutamate excitotoxicity also influence tributyltin toxicity. Cell death induced by tributyltin was found to be reduced by -tocopherol (a membrane-permeable antioxidant), SB202190 (a p38 mitogen-activated protein kinase inhibitor), and U-0126 (an extracellular signal-regulated protein kinase kinase inhibitor). MK-801 and CNQX decreased the phosphorylation of ERK, but not that of p38. A caspase-3 inhibitor had no effect on tributyltin toxicity, and tributyltin did not change the nuclear morphology. These results suggest that the glutamate excitotoxicity caused by tributyltin is unrelated to apoptosis. In conclusion, we demonstrated that tributyltin induced glutamate release and subsequent activation of glutamate receptors, leading to neuronal death. We propose two independent neuronal death pathways by tributyltin; one is glutamate receptor–dependent cell death via ERK phosphorylation, and the other may be glutamate receptor–independent cell death via p38 activation.
Key Words: tributyltin-glutamate release-NMDA receptors; reactive oxygen species; mitogen-activated protein kinase; caspase.
INTRODUCTION
Organotin compounds have been widely used as heat stabilizers in vinyl chloride polymers, wood preservatives, and agricultural pesticides. Among them, tributyltin chloride (TBT) has been most widely used in marine antifouling paints, even though its residues represent an environmental and health hazard. The use of TBT in antifouling paints has been restricted recently (van Wezel et al., 2004), but TBT and its degradation products dibutyltin and monobutyltin will remain in marine sediments for some time to come, because the half-life of TBT is long, about 2 years (Sarradin et al., 1995). Organotin compounds contaminate human food and water through contributions from industrial effluents and leaching from polyvinyl chloride water pipes (Snoeij et al., 1987). Indeed, Tsuda et al. (1995) reported that the daily intake of TBT in Japan was 2.2–6.9 μg, and Whalen et al. (1999) reported the presence of butyltin compounds, including TBT, at concentrations between 50 nM and 400 nM in human blood.
Tributyltin chloride has been reported to show toxicity to mammals in addition to marine organisms. Ema et al. (1991) reported that TBT caused significant changes in rat behavior. Exposure of mammals to organotin compounds can induce epilepsy, amnesia, and memory defects (Feldman et al., 1993); it is toxic to the developing central nervous system (CNS) (O'Callaghan and Miller, 1988). Little is known about the mechanism of TBT toxicity, but several lines of evidence suggest that TBT causes an increase in intracellular calcium in various cells, including thymocytes (Chow et al., 1992), hepatocytes (Kawanishi et al., 2001), and PC12 cells (Viviani et al., 1995).
In the CNS, glutamate is an important factor for maintaining calcium homeostasis; it is the most abundant excitatory neurotransmitter, and it is widely distributed. Glutamate is associated with various brain functions, such as synaptic plasticity, learning, and long-term potentiation (Collingridge and Laster, 1989). Its physiological and pathological effects in the CNS are mediated mainly via two types of ionotropic glutamate receptors, the NMDA receptor and the non-NMDA receptor. Activation of NMDA receptors allows influx of extracellular Ca2+, but excessive release of glutamate causes prolonged stimulation of NMDA receptors, inducing calcium overload and neuronal death (so-called excitotoxicity). Excitotoxicity has been speculated to be involved in many neurodegenerative diseases, such as Alzheimer's disease (Hynd et al., 2004), Parkinson's disease (Blandini et al., 1996), and amyotrophic lateral sclerosis (ALS) (Rao and Weiss., 2004). Moreover, glutamate neurotoxicity has been reported to play an important role in the neurotoxicity of environmental pollutants, such as methyl mercury (Farina et al., 2003).
In the present study, we examined whether glutamate is involved in TBT toxicity in rat cortical cultures. We investigated whether TBT induces glutamate release and whether NMDA receptor antagonists reduce TBT toxicity. Moreover, we examined whether several factors involved in glutamate excitotoxicity also influence TBT toxicity in cortical neurons.
MATERIALS AND METHODS
Chemicals.
Eagle's minimal essential salt medium (Eagle's MEM) was purchased from Nissui Pharmaceutical (Tokyo, Japan). Fetal calf serum (FCS) and horse serum (HS) were purchased from JRH Biosciences (Lenexa, KS). Tributyltin chloride, trypan blue, nifedipine, -tocopherol, ascorbic acid, NaHCO3, and glucose were purchased from Wako (Osaka, Japan). Catalase and glutamine were purchased from Sigma (St. Louis, MO). MK-801 was purchased from Funakoshi (Osaka, Japan). CNQX was purchased from Tocris (Bristol, UK). HEPES was purchased from DOJINDO (Kumamoto, Japan). U-0126, SB202190, and caspase-3 inhibitor (Ac-DEVD-CMK) were purchased from Calbiochem (Darmstadt, Germany). SP600125 was purchased from Biomol Research Laboratories Inc. (Plymouth Meeting, PA). Primary antibodies and horseradish peroxidase–conjugated anti-rabbit antibody were purchased from Cell Signaling Technology (Beverly, MA).
Cell culture.
The following procedures were performed under sterile conditions. The animals were handled in accordance with the guidelines for the care and use of experimental animals published by the Japanese Association for Laboratory Animal Science in 1987. Primary cultures were obtained from the cerebral cortex of fetal rats (at 18 days of gestation) as described elsewhere (Taguchi et al., 2003). Fetuses were taken from pregnant Slc; Wistar/ST rats under pentobarbital anesthesia. A part of the cerebral cortex was dissected with a razor blade, and cells were dissociated by gentle pipetting. Dissociated cells were plated on plastic cover slips or culture plates (4 x 105 cells/cm2). Cultures were incubated in Eagle's MEM supplemented with 10% heat-inactivated FCS, L-glutamine (2 mM), D-(+)-glucose (11 mM), NaHCO3 (24 mM), and HEPES (10 mM). Cultures were maintained at 37°C in a humidified 5% CO2 atmosphere. The cultures were incubated in MEM containing 10% FCS (1–7 days) or 10% HS (8–11 days). The medium was exchanged every 2–3 days. Arabinocylcytosine (10 μM) was added to inhibit the proliferation of non-neuronal cells after 6 days of plating. Cultures were used for experiments at 10–14 days in vitro. This protocol has been reported to produce cultures containing about 90% neurons (Taguchi et al., 2003).
Drug treatment.
MK-801, ascorbic acid, and catalase were dissolved in water, and other chemicals were dissolved in dimethylsulfoxide (DMSO). Cell cultures were exposed to various drugs in MEM containing 10% HS. Tributyltin chloride was added to the culture medium 30 min after the addition of various inhibitors. The concentration of DMSO was 0.1% or 0.2%.
Measurement of neurotoxicity.
Neurotoxicity was quantified by trypan blue assay, referring to a previous report (Taguchi et al., 2003). After exposure to TBT, cell cultures were immediately stained with 1.5 % trypan blue for 10 min, fixed with formalin, and rinsed with physiological saline. Unstained cells were regarded as viable and stained cells were regarded as dead. The viability of the cultures was calculated as the percentage ratio of the number of unstained cells relative to the total cells counted. Over 200 cells per cover slip were randomly counted.
Measurement of released glutamate.
Released glutamate was measured with an Amplex Red Glutamic Acid/Glutamate Oxidase assay kit (Molecular Probes, Eugene, OR) according to the manufacturer's protocol. Culture medium was removed and cell cultures were incubated in HEPES-buffered salt solution containing 125 mM NaCl, 5 mM KCl, 1.2 mM KH2PO4, 1.2 mM MgSO4, 1.2 mM CaCl2, 6 mM glucose, and 25 mM HEPES. After the TBT stimulation, 50 μl of buffer was transferred to a 96-well microplate and mixed with 50 μl of reaction buffer containing Amplex red, alanine, glutamate oxidase, glutamate pyruvate transaminase, and horseradish peroxidase. The mixture was incubated for 30 min in the dark and the fluorescence was measured with the use of a fluorescence microplate reader (excitation at 538 nm and emission at 594 nm).
Western blotting.
After TBT treatment, cells were washed with PBS buffer and lysed in TNE buffer containing 50 mM Tris-HCl, 1% NP-40, 20 mM EDTA, 0.2 mM sodium orthovanadate, and 1 mM PMSF. The mixture was centrifuged at 13,500 rpm for 3 min, after which the supernatant was transferred to an Eppendorf tube. The supernatant was added to sample buffer containing 125 mM Tris-HCl, 4% SDS, 10% sucrose, 0.004% BPB, and 10% mercaptoethanol, then denatured at 95°C for 3 min. Protein was separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to a polyvinylidene difluoride membrane. The membrane was blocked with blocking buffer containing 5% skim milk for 1 h, then incubated with primary antibody (1:1000) overnight and secondary antibody (1:2000) for 1 h. The protein was detected with an enhanced chemiluminescence detection system.
Hoechst 33258 staining.
Cultured cells were stained with Hoechst 33258 to examine nuclear morphological changes. Cells were fixed with neutral formaldehyde for 30 min and then stained with 0.1 mg/ml Hoechst 33258 for 30 min. Cells were visualized and photographed under ultraviolet illumination, with a fluorescence microscope.
Statistics.
Data are expressed as mean ± SEM. Statistical evaluation of the data was performed by Student's t-test (Fig. 1 and Fig. 2A) or by analysis of variance (ANOVA) followed by the Newman-Keul's test (except Fig. 1 and Fig. 2A). A value of p < 0.05 was considered to be indicative of significance.
RESULTS
Neurotoxicity of TBT in Cortical Neurons
Toxicity of TBT in rat cortical neurons was examined by the trypan blue method (dead cells stained with trypan blue). Cortical neurons were exposed to various concentrations of TBT, and exposure of the cells to 500 nM TBT for 24 h resulted in a remarkable degree of neurotoxicity (survival of 49.8 ± 0.4%) (Fig. 1A), with increased numbers of trypan blue–stained cells (Fig. 2E, F). Next, cell viability was examined at 1, 3, 6, 12, and 24 h after 500 nM TBT exposure. Cell viability decreased when the cells were exposed to TBT for longer than 12 h (Fig. 1B).
Involvement of Glutamate in TBT Neurotoxicity
We examined whether glutamate is involved in TBT toxicity, because glutamate is believed to be a factor in neuronal death. First, the amount of glutamate in the medium was measured. The concentration of extracellular glutamate was increased significantly by exposure of the cells to 500 nM TBT for 30 min, and that increase was exposure time–dependent (Fig. 2A). Next, to investigate the involvement of glutamate release in TBT toxicity, we used NMDA receptor antagonists. Application of 5 μM MK-801, a noncompetitive NMDA receptor antagonist, alleviated TBT neurotoxicity (survival with TBT: 45.7 ± 3.3%; +MK-801: 87.8 ± 7.1%) (Fig. 2B). Further, 20 μM CNQX, a non-NMDA receptor antagonist, showed almost the same effect (survival with TBT: 52.1 ± 2.4%; +CNQX: 83.7 ± 5.3%) (Fig. 2C). These inhibitors greatly decreased the number of trypan blue–stained cells (Fig. 2G, H). Moreover, we investigated the involvement of voltage-dependent calcium channels (VDCC), because it has been reported that VDCC are involved in the toxicity of NMDA agonists (Fryer, 1999). Neuronal death caused by TBT was blocked by 10 μM nifedipine, a VDCC inhibitor (survival with TBT: 58.5 ± 2.7; +nifedipine: 84.7 ± 1.7%) (Fig. 2D).
Effect of Antioxidants on TBT Toxicity
It is known that oxidative stress contributes to glutamate toxicity (Campisi et al., 2004). Therefore, we examined the effect of antioxidants on TBT toxicity. We used water-soluble antioxidants (ascorbic acid and catalase) and a lipophilic antioxidant (-tocopherol). Co-administration of 100 μM ascorbic acid or 1000 U/ml catalase during TBT exposure failed to reduce TBT toxicity. In contrast, co-administration of 100 μM -tocopherol remarkably inhibited TBT-induced neurotoxicity (survival with TBT: 49.3 ± 3.7%; with -tocopherol 82.7 ± 4.5%) (Fig. 3).
Effect of MAPK Inhibitors on TBT Toxicity
Mitogen-activated protein kinase (MAPK) has been implicated in glutamate-induced neuronal death (Cao et al., 2004). Therefore we used inhibitors of the MAP kinase cascade to examine whether MAPKs are involved in TBT neurotoxicity. We found that 10 μM U-0126, an extracellular signal-regulated protein kinase (ERK) kinase inhibitor, reduced TBT neurotoxicity (survival with TBT: 46.4 ± 2.2%; +U-0126: 66.7 ± 1%) and that 10 μM SB202190, a p38 inhibitor, also significantly alleviated the neurotoxicity (survival with +SB202190: 76.7 ± 6.8%). However, 10 μM SP600125, a c-jun-N-terminal kinase (JNK) inhibitor, was ineffective against TBT-induced cell death (Fig. 4A). Further, we investigated whether TBT phosphorylated ERK and p38. As shown in Figure 4B and 4C, at 3 h after application of 500 nM TBT to the culture, p38 was phosphorylated, and at 6 h, ERK was activated. Phosphorylation of ERK and p38 was sustained until 24 h after exposure. We also examined the effects of glutamate receptor antagonists on ERK and p38 phosphorylation and found that MK-801 and CNQX decreased the phosphorylation of ERK, but not that of p38 (Fig. 4D and 4E).
Effect of Caspase-3 Inhibitor on TBT Toxicity
Because it has been reported that TBT activates caspase-3 in various cells (Nopp et al., 2002; Jurkiewicz et al., 2004), we used a caspase-3 inhibitor to confirm the involvement of caspase in TBT toxicity in cortical neurons. However, 10 μM of the caspase-3 inhibitor Ac-DEVD-CMK did not reduce TBT toxicity (Fig. 5A). Moreover, we investigated whether 500 nM TBT alters the nuclear morphology (an indication of apoptosis) of cortical neurons, and we found that it did not (Fig. 5C), although 2 μM TBT did induce nuclear fragmentation in PC12 cells, in agreement with a previous report (Fig. 5D) (Viviani et al., 1995).
DISCUSSION
Among organotin compounds, trimethyltin is neurotoxic, whereas tributyltin is primarily known as an immunotoxin (Snoeij et al., 1985). However, neurotoxicity of TBT has also been reported (O'Callaghan and Miller, 1988; Tsunoda et al., 2005), although its mechanism has been little investigated. Trimethyltin and tributyltin are thought to have different mechanisms of toxicity. For example, TMT is reported to exert neurotoxicity at least in part by binding specifically to the protein stannin, whereas TBT does not bind to stannin (Thompson et al., 1996, Buck et al., 2003).
Organotin compounds are important environmental pollutants, in part because of their toxicity. Braman et al. (1979) reported that methyltin compounds, including TMT, were detected at concentrations between 0.5 pM and 4 nM in human urine. Further, the butyltin concentration in human blood ranges from 50 nM to 400 nM (Whalen et al., 1999), and that in human liver is about 400 nM (Kannan and Falandysz, 1997; Mizuhashi et al., 2000). The TBT concentration in rock shells caught in polluted sea areas around Japan was about 5 μM (Horiguchi et al., 1994; Mizuhashi et al., 2000). The concentration of 500 nM TBT used in our experiments thus appears to be similar to levels of human exposure.
O'Callaghan and Miller (1988) reported that TBT decreased brain weight at dose levels that had no influence on body weight. This finding suggests that the brain may be particularly susceptible to injury by TBT. We found that exposure of cortical neurons to 500 nM TBT for 24 h significantly reduced their viability (Fig. 1). Thompson et al. (1996) reported that the 50%-toxic concentration of TBT for 48 h of exposure ranges from 5.5 μM to 9 μM for non-neuronal cells, such as NIH-3T3, TC7, and B cells, and that primary neuronal cells were more sensitive to TBT than non-neuronal cells. These results suggest that TBT exerts neuronal toxicity via a specific mechanism, at lower concentrations than the levels causing toxicity in other organs.
We examined the involvement of glutamate in TBT-induced neurotoxicity because glutamate is believed to play important roles in various types of neuronal death. We found that TBT increased extracellular glutamate concentration. The mechanism of glutamate release is not clear, but it is unlikely to be due to leakage from dead cells, because significant numbers of dead cells were not detected for up to 6 h after exposure to TBT. In our study glutamate release was not inhibited by NMDA receptor antagonists or a VDCC inhibitor (data not shown). Thus TBT appears to induce glutamate release via a Ca2+-independent pathway. Application of the glutamate receptor antagonists MK-801 and CNQX reduced the level of TBT toxicity (Fig. 2). These results support the idea that release of glutamate and activation of glutamate receptors are involved in TBT toxicity. It was previously reported that non-NMDA receptors were involved in TBT neurotoxicity in hippocampal slice cultures, but NMDA receptors and VDCC were not involved (Mizuhashi et al., 2000). Activation of both NMDA receptors and non-NMDA receptors may not necessarily be required for excitotoxicity (Kubo et al., 2001; Hakuba et al., 2003). The difference in the involvement of NMDA receptors may reflect differences in experimental conditions, such as brain region or exposure conditions. Also, VDCC plays important roles in other effects of glutamate excitotoxicity, such as ischemia (Hou et al., 2002). In our experimental model, nifedipine blocked neuronal death caused by TBT, and this result indicates that VDCC is involved in TBT toxicity in our system.
Moreover, we investigated the effects of combinations of inhibitors on TBT neurotoxicity, but we found that the combinations did not show potentiated inhibition (data not shown). Non-NMDA receptors are supposed to open after glutamate release, and their activation induces a change in the membrane potential. Glutamate and cancellation of Mg2+ blockade (caused by the change of membrane potential) are needed for NMDA receptor activation. Therefore, non-NMDA receptors should be activated before NMDA receptors and VDCC. This chain of activation is presumably why the effects of inhibitor combinations are the same as those of a single inhibitor.
Because reactive oxygen species (ROS) are generated by activation of the glutamate receptor (Campisi et al., 2004), we also examined whether ROS are involved in TBT toxicity. Application of -tocopherol blocked cell death induced by TBT, but catalase and ascorbic acid had no effect (Fig. 3). These results suggest that intracellular generation of ROS is important, because -tocopherol is more lipophilic and cell-permeable than the other two antioxidants. Ahlemeyer and Krieglstein (2000) showed that -tocopherol inhibited mitochondrial reactive ROS production, indicating that -tocopherol can decrease intracellular ROS. Ciani et al. (1996) reported that -tocopherol, but not ascorbic acid, decreased glutamate neurotoxicity. Kim et al. (1998) reported that glutamate exerts toxicity via lipid peroxidation. Several investigators have described that the most effective agents against glutamate toxicity appear to be chain-breaking antioxidants that block lipid peroxidation, such as -tocopherol (Ciani et al., 1996; Halliwell, 1992; Siesjo et al., 1989). These findings are consistent with our results.
Next, the effects of MAPK inhibitors were examined, because MAPKs play important roles in glutamate excitotoxicity (Mao et al., 2004). Yu et al. (2000) reported that TBT phosphorylated ERK, JNK, and p38 in human T lymphoblastoid cells CCRF-CEM. In our experimental model, a MEK inhibitor, U-0126, and a p38 MAPK inhibitor, SB202190, decreased TBT toxicity, but a JNK inhibitor did not (Fig. 4). Glutamate receptor antagonists inhibited phosphorylation of ERK, but not that of p38. These results indicate that ERK phosphorylation depends on glutamate receptors, and that there may also be another pathway involving p38 activation. It has been reported that glutamate activates ERK (Jiang et al., 2000). Activation of ERK is supposed to be characteristic of glutamate toxicity, and our results are consistent with this idea.
It is well known that caspase family members are the central executors of apoptosis, and among them, caspase-3 is the most important. Tributyltin activates caspase-3 in various cells, such as Jurkat T cells, eosinophils, neutrophils, and hepatocytes (Nopp et al., 2002, Jurkiewicz et al., 2004). Caspase-3 inhibitor did not protect neurons from TBT toxicity (Fig. 5A), and TBT did not induce any change in nuclear morphology (Fig. 5C). There have been many reports that glutamate-induced neurotoxicity is apoptotic (Kawasaki et al., 1997; Du et al., 1997) or necrotic (Sohn et al., 1998; Dessi et al., 1993). Our results indicate that glutamate neurotoxicity caused by TBT is not apoptotic, at least in our experimental system.
In summary, we have partially clarified the mechanism of TBT neurotoxicity in cultured rat cortical neurons (Fig. 6). Tributyltin at low concentration was toxic to cortical neurons through glutamate release and subsequent activation of glutamate receptors. The mechanism of the glutamate release induced by TBT remains unclear, but it is not simply leakage from dead cells. We demonstrated that intracellular ROS, ERK, and p38 MAPK were also involved in TBT neurotoxicity. It was shown that ERK phosphorylation depended on glutamate receptor activation, but its activation was not involved in p38 phosphorylation. These results suggest that there is a glutamate receptor–dependent pathway and a glutamate receptor–independent pathway. Moreover, the glutamate toxicity induced by TBT was not apoptotic. Thus, glutamate excitotoxicity seems to account for the high sensitivity of cortical neurons to TBT.
ACKNOWLEDGMENTS
This study was supported in part by a Grant-in Aid for Scientific Research from the Japan Society for the Promotion of Science.
REFERENCES
Ahlemeyer, B., and Krieglstein, J. (2000). Inhibition of glutathione depletion by retinoic acid and tocopherol protects cultured neurons from staurosporine-induced oxidative stress and apoptosis. Neurochem. Int. 36, 1–5.
Blandini, F., Porter, R. H., and Greenamyre, J. T. (1996). Glutamate and Parkinson's disease. Mol. Neurobiol. 12, 73–94.
Braman, R. S., and Tompkins, M. A. (1979). Separation and determination of nanogram amounts of inorganic tin and methyltin compounds in the environment. Anal. Chem. 51, 12–19.
Buck, B., Mascioni, A., Que, L., Jr., and Veglia, G. (2003). Dealkylation of organotin compounds by biological dithiols: Toward the chemistry of organotin toxicity. J. Am. Chem. Soc. 125, 13316–13317.
Campisi, A., Caccamo, D., Li Volti, G., Curro, M., Parisi, G., Avola, R., Vanella, A., and Ientile, R. (2004). Glutamate-evoked redox state alterations are involved in tissue transglutaminase upregulation in primary astrocyte cultures. FEBS Lett. 578, 80–84.
Cao, J., Semenova, M. M., Solovyan, V. T., Han, J., Coffey, E. T., and Courtney, M. J. (2004). Distinct requirements for p38alpha and c-Jun N-terminal kinase stress-activated protein kinases in different forms of apoptotic neuronal death. J. Biol. Chem. 279, 35903–35913.
Chow, S. C., Kass, G. E., McCabe, M. J., Jr., and Orrenius, S. (1992). Tributyltin increases cytosolic free Ca2+ concentration in thymocytes by mobilizing intracellular Ca2+, activating a Ca2+ entry pathway, and inhibiting Ca2+ efflux. Arch. Biochem. Biophys. 298, 143–149.
Ciani, E., Groneng, L., Voltattorni, M., Rolseth, V., Contestabile, A., and Paulsen, R. E. (1996). Inhibition of free radical production or free radical scavenging protects from the excitotoxic cell death mediated by glutamate in cultures of cerebellar granule neurons. Brain Res. 728, 1–6.
Collingridge, G. L., and Laster, R. A. (1989). Excitatory amino acid receptors in the vertebrate central nervous system. Pharmacol. Rev. 41, 143–210.
Dessi, F., Charriaut-Marlangue, C., Khrestchatisky, M., and Ben-Ari, Y. (1993). Glutamate-induced neuronal death is not a programmed cell death in cerebellar culture. J. Neurochem. 60, 1953–1955.
Du, Y., Bales, K. R., Dodel, R. C., Hamilton-Byrd, E., Horn, J. W., Czilli, D. L., Simmons, L. K., Ni, B., and Paul, S. M. (1997). Activation of a caspase 3-related cysteine protease is required for glutamate-mediated apoptosis of cultured cerebellar granule neurons. Proc. Natl. Acad. Sci. U.S.A. 94, 11657–11662.
Ema, M., Itami, T., and Kawasaki, H. (1991). Behavioral effects of acute exposure to tributyltin chloride in rats. Neurotoxicol. Teratol. 13, 489–493.
Farina, M., Dahm, K. C., Schwalm, F. D., Brusque, A. M., Frizzo, M. E., Zeni, G., Souza, D. O., and Rocha, J. B. (2003). Methylmercury increases glutamate release from brain synaptosomes and glutamate uptake by cortical slices from suckling rat pups: Modulatory effect of ebselen. Toxicol. Sci. 73, 135–140.
Feldman, R. G., White, R. F., and Eriator, I. I. (1993). Trimethyltin encephalopathy. Arch. Neurol. 50, 1320–1324.
Fryer, H. J., Knox, R. J., Strittmatter, S. M., and Kalb, R. G. (1999). Excitotoxic death of a subset of embryonic rat motor neurons in vitro. J. Neurochem. 72, 500–513.
Hakuba, N., Matsubara, A., Hyodo, J., Taniguchi, M., Maetani, T., Shimizu, Y., Tsujiuchi, Y., Shudou, M., and Gyo, K. (2003). AMPA/kainate-type glutamate receptor antagonist reduces progressive inner hair cell loss after transient cochlear ischemia Brain Res. 979, 194–202.
Halliwell, B. (1992). Reactive oxygen species and the central nervous system. J. Neurochem. 59, 1609–1623.
Horiguchi, T., Shiraishi, H., Shimizu, M., and Morita, M. (1994). Imposex and organotin compounds in Thais clavigera and T. bronni in Japan. J. Mar. Biol. Assoc. U.K. 74, 651–669.
Hou, X. Y., Zhang, G. Y., Yan, J. Z., Chen, M., and Liu, Y. (2002). Activation of NMDA receptors and L-type voltage-gated calcium channels mediates enhanced formation of Fyn-PSD95-NR2A complex after transient brain ischemia. Brain Res. 955, 123–132.
Hynd, M. R., Scott, H. L., and Dodd, P. R. (2004). Glutamate-mediated excitotoxicity and neurodegeneration in Alzheimer's disease. Neurochem. Int. 45, 583–595.
Jiang, Q., Gu, Z., Zhang, G., and Jing, G. (2000). N-methyl-D-aspartate receptor activation results in regulation of extracellular signal-regulated kinases by protein kinases and phosphatases in glutamate-induced neuronal apototic-like death. Brain Res. 887, 285–292.
Jurkiewicz, M., Averill-Bates, D. A., Marion, M., and Denizeau, F. (2004). Involvement of mitochondrial and death receptor pathways in tributyltin-induced apoptosis in rat hepatocytes. Biochim. Biophys. Acta 1693, 15–27.
Kannan, K., and Falandysz, Z. (1997). Butyltin residues in sediment, fish, fish eating birds, harbour porpoise and human tissues from the Polish coast of the Baltic Sea. Mar. Pollut. Bull. 34, 203–207.
Kawanishi, T., Kiuchi, T., Asoh, H., Shibayama, R., Kawai, H., Ohata, H., Momose, K., and Hayakawa, T. (2001). Effect of tributyltin chloride on the release of calcium ion from intracellular calcium stores in rat hepatocytes. Biochem. Pharmacol. 62, 863–872.
Kawasaki, H., Morooka, T., Shimohama, S., Kimura, J., Hirano, T., Gotoh, Y., and Nishida, E. (1997). Activation and involvement of p38 mitogen-activated protein kinase in glutamate-induced apoptosis in rat cerebellar granule cells. J. Biol. Chem. 272, 18518–18521.
Kim, Y. C., Kim, S. R., Markelonis, G. J., and Oh, T. H. (1998). Ginsenosides Rb1 and Rg3 protect cultured rat cortical cells from glutamate-induced neurodegeneration. J. Neurosci. Res. 53, 426–432.
Kubo, T., Yokoi, T., Hagiwara, Y., Fukumori, R., Goshima, Y., and Misu, Y. (2001). Characteristics of protective effects of NMDA antagonist and calcium channel antagonist on ischemic calcium accumulation in rat hippocampal CA1 region. Brain Res. Bull. 54, 413–419.
Mao, L., Tang, Q., Samdani, S., Liu, Z., and Wang, J. Q. (2004). Regulation of MAPK/ERK phosphorylation via ionotropic glutamate receptors in cultured rat striatal neurons. Eur. J. Neurosci. 19, 1207–1216.
Mizuhashi, S., Ikegaya, Y., and Matsuki, N., (2000). Cytotoxicity of tributyltin in rat hippocampal slice cultures. Neurosci. Res. 38, 35–42.
Nopp, A., Lundahl, J., and Stridh, H. (2002). Caspase activation in the absence of mitochondrial changes in granulocyte apoptosis. Clin. Exp. Immunol. 128, 267–274.
O'Callaghan, J. P., and Miller, D. B. (1988). Acute exposure of the neonatal rat to tributyltin results in decreases in biochemical indicators of synaptogenesis and myelinogenesis J. Pharmacol. Exp. Ther. 246, 394–402.
Rao, S. D., and Weiss, J. H. (2004). Excitotoxic and oxidative cross-talk between motor neurons and glia in ALS pathogenesis Trends Neurosci. 27, 17–23.
Sarradin, P. M., Lapaquellerie, Y., Astruc, A., Latouche, C., and Astruc, M. (1995). Long term behaviour and degradation kinetics of tributyltin in a marina sediment. Sci. Total Environ. 170, 59–70.
Siesjo, B. K., Agardh, C. D., and Bengtsson, F. (1989). Free radicals and brain damage. Cerebrovasc. Brain Metab. Rev. 1, 165–211.
Snoeij, N. J., van Iersel, A. A., Penninks, A. H., and Seinen, W. (1985). Toxicity of triorganotin compounds: Comparative in vivo studies with a series of trialkyltin compounds and triphenyltin chloride in male rats. Toxicol. Appl. Pharmacol. 81, 274–286.
Snoeij, N. J., Penninks, A. H., and Seinen, W. (1987). Biological activity of organotin compounds—An overview. Environ. Res. 44, 335–353.
Sohn, S., Kim, E. Y., and Gwag, B. J. (1998). Glutamate neurotoxicity in mouse cortical neurons: Atypical necrosis with DNA ladders and chromatin condensation. Neurosci. Lett. 240, 147–150.
Taguchi, R., Nishikawa, H., Kume, T., Terauchi, T., Kaneko, S., Katsuki, H., Yonaga, M, Sugimoto, H., and Akaike, A. (2003). Serofendic acid prevents acute glutamate neurotoxicity in cultured cortical neurons. Eur. J. Pharmacol. 477, 195–203.
Thompson, T. A., Lewis, J. M., Dejneka, N. S., Severs, W. B., Polavarapu, R., and Billingsley, M. L. (1996). Induction of apoptosis by organotin compounds in vitro: Neuronal protection with antisense oligonucleotides directed against stannin. J. Pharmacol. Exp. Ther. 276, 1201–1216.
Tsuda, T., Inoue, T., Kojima, M., and Aoki, S. (1995). Daily intakes of tributyltin and triphenyltin compounds from meals. J. AOAC Int. 78, 941–943.
Tsunoda, M., Konno, N., Nakano, K., and Liu, Y. (2005). Altered metabolism of dopamine in the midbrain of mice treated with tributyltin chloride via subacute oral exposure. Environ. Sci. 11, 209–219.
Whalen, M. M., Loganathan, B. G., and Kannan, K. (1999) Immunotoxicity of environmentally relevant concentrations of butyltins on human natural killer cells in vitro. Environ. Res. 81, 108–116.
van Wezel, A. P., and van Vlaardingen, P. (2004). Environmental risk limits for antifouling substances. Aquat. Toxicol. 66, 427–444.
Viviani, B., Rossi, A. D., Chow, S. C., and Nicotera, P. (1995). Organotin compounds induce calcium overload and apoptosis in PC12 cells. Neurotoxicology 16, 19–25.
Yu, Z. P., Matsuoka, M., Wispriyono, B., Iryo, Y., and Igisu, H. (2000). Activation of mitogen-activated protein kinases by tributyltin in CCRF-CEM cells: Role of intracellular Ca2+. Toxicol. Appl. Pharmacol. 168, 200–207.(Yusuke Nakatsu, Yaichiro Kotake, Kazuya )
Center for Quantum Life Sciences, Hiroshima University, 1–2–3, Kasumi, Minami-ku, Hiroshima, 734-8551, Japan
Graduate School of Pharmaceutical Sciences, Kyoto University, 46–29, Yoshida-Shimoadachi-cho, Sakyo-ku, Kyoto, 606-8501, Japan
ABSTRACT
Tributyltin, an endocrine-disrupting chemical, has been used as a heat stabilizer, agricultural pesticide, and component of antifouling paints. In this study, the neurotoxicity of tributyltin was investigated in cultured rat cortical neurons. Tributyltin caused marked time- and dose-dependent increases in the number of trypan blue–stained cells. Measurement of extracellular glutamate concentration showed that glutamate release was induced by tributyltin. Application of the glutamate receptor antagonists MK-801 and CNQX decreased the neurotoxicity. These results suggest that released glutamate and glutamate receptors are involved in tributyltin toxicity. Next, we examined whether various factors, believed to be involved in glutamate excitotoxicity also influence tributyltin toxicity. Cell death induced by tributyltin was found to be reduced by -tocopherol (a membrane-permeable antioxidant), SB202190 (a p38 mitogen-activated protein kinase inhibitor), and U-0126 (an extracellular signal-regulated protein kinase kinase inhibitor). MK-801 and CNQX decreased the phosphorylation of ERK, but not that of p38. A caspase-3 inhibitor had no effect on tributyltin toxicity, and tributyltin did not change the nuclear morphology. These results suggest that the glutamate excitotoxicity caused by tributyltin is unrelated to apoptosis. In conclusion, we demonstrated that tributyltin induced glutamate release and subsequent activation of glutamate receptors, leading to neuronal death. We propose two independent neuronal death pathways by tributyltin; one is glutamate receptor–dependent cell death via ERK phosphorylation, and the other may be glutamate receptor–independent cell death via p38 activation.
Key Words: tributyltin-glutamate release-NMDA receptors; reactive oxygen species; mitogen-activated protein kinase; caspase.
INTRODUCTION
Organotin compounds have been widely used as heat stabilizers in vinyl chloride polymers, wood preservatives, and agricultural pesticides. Among them, tributyltin chloride (TBT) has been most widely used in marine antifouling paints, even though its residues represent an environmental and health hazard. The use of TBT in antifouling paints has been restricted recently (van Wezel et al., 2004), but TBT and its degradation products dibutyltin and monobutyltin will remain in marine sediments for some time to come, because the half-life of TBT is long, about 2 years (Sarradin et al., 1995). Organotin compounds contaminate human food and water through contributions from industrial effluents and leaching from polyvinyl chloride water pipes (Snoeij et al., 1987). Indeed, Tsuda et al. (1995) reported that the daily intake of TBT in Japan was 2.2–6.9 μg, and Whalen et al. (1999) reported the presence of butyltin compounds, including TBT, at concentrations between 50 nM and 400 nM in human blood.
Tributyltin chloride has been reported to show toxicity to mammals in addition to marine organisms. Ema et al. (1991) reported that TBT caused significant changes in rat behavior. Exposure of mammals to organotin compounds can induce epilepsy, amnesia, and memory defects (Feldman et al., 1993); it is toxic to the developing central nervous system (CNS) (O'Callaghan and Miller, 1988). Little is known about the mechanism of TBT toxicity, but several lines of evidence suggest that TBT causes an increase in intracellular calcium in various cells, including thymocytes (Chow et al., 1992), hepatocytes (Kawanishi et al., 2001), and PC12 cells (Viviani et al., 1995).
In the CNS, glutamate is an important factor for maintaining calcium homeostasis; it is the most abundant excitatory neurotransmitter, and it is widely distributed. Glutamate is associated with various brain functions, such as synaptic plasticity, learning, and long-term potentiation (Collingridge and Laster, 1989). Its physiological and pathological effects in the CNS are mediated mainly via two types of ionotropic glutamate receptors, the NMDA receptor and the non-NMDA receptor. Activation of NMDA receptors allows influx of extracellular Ca2+, but excessive release of glutamate causes prolonged stimulation of NMDA receptors, inducing calcium overload and neuronal death (so-called excitotoxicity). Excitotoxicity has been speculated to be involved in many neurodegenerative diseases, such as Alzheimer's disease (Hynd et al., 2004), Parkinson's disease (Blandini et al., 1996), and amyotrophic lateral sclerosis (ALS) (Rao and Weiss., 2004). Moreover, glutamate neurotoxicity has been reported to play an important role in the neurotoxicity of environmental pollutants, such as methyl mercury (Farina et al., 2003).
In the present study, we examined whether glutamate is involved in TBT toxicity in rat cortical cultures. We investigated whether TBT induces glutamate release and whether NMDA receptor antagonists reduce TBT toxicity. Moreover, we examined whether several factors involved in glutamate excitotoxicity also influence TBT toxicity in cortical neurons.
MATERIALS AND METHODS
Chemicals.
Eagle's minimal essential salt medium (Eagle's MEM) was purchased from Nissui Pharmaceutical (Tokyo, Japan). Fetal calf serum (FCS) and horse serum (HS) were purchased from JRH Biosciences (Lenexa, KS). Tributyltin chloride, trypan blue, nifedipine, -tocopherol, ascorbic acid, NaHCO3, and glucose were purchased from Wako (Osaka, Japan). Catalase and glutamine were purchased from Sigma (St. Louis, MO). MK-801 was purchased from Funakoshi (Osaka, Japan). CNQX was purchased from Tocris (Bristol, UK). HEPES was purchased from DOJINDO (Kumamoto, Japan). U-0126, SB202190, and caspase-3 inhibitor (Ac-DEVD-CMK) were purchased from Calbiochem (Darmstadt, Germany). SP600125 was purchased from Biomol Research Laboratories Inc. (Plymouth Meeting, PA). Primary antibodies and horseradish peroxidase–conjugated anti-rabbit antibody were purchased from Cell Signaling Technology (Beverly, MA).
Cell culture.
The following procedures were performed under sterile conditions. The animals were handled in accordance with the guidelines for the care and use of experimental animals published by the Japanese Association for Laboratory Animal Science in 1987. Primary cultures were obtained from the cerebral cortex of fetal rats (at 18 days of gestation) as described elsewhere (Taguchi et al., 2003). Fetuses were taken from pregnant Slc; Wistar/ST rats under pentobarbital anesthesia. A part of the cerebral cortex was dissected with a razor blade, and cells were dissociated by gentle pipetting. Dissociated cells were plated on plastic cover slips or culture plates (4 x 105 cells/cm2). Cultures were incubated in Eagle's MEM supplemented with 10% heat-inactivated FCS, L-glutamine (2 mM), D-(+)-glucose (11 mM), NaHCO3 (24 mM), and HEPES (10 mM). Cultures were maintained at 37°C in a humidified 5% CO2 atmosphere. The cultures were incubated in MEM containing 10% FCS (1–7 days) or 10% HS (8–11 days). The medium was exchanged every 2–3 days. Arabinocylcytosine (10 μM) was added to inhibit the proliferation of non-neuronal cells after 6 days of plating. Cultures were used for experiments at 10–14 days in vitro. This protocol has been reported to produce cultures containing about 90% neurons (Taguchi et al., 2003).
Drug treatment.
MK-801, ascorbic acid, and catalase were dissolved in water, and other chemicals were dissolved in dimethylsulfoxide (DMSO). Cell cultures were exposed to various drugs in MEM containing 10% HS. Tributyltin chloride was added to the culture medium 30 min after the addition of various inhibitors. The concentration of DMSO was 0.1% or 0.2%.
Measurement of neurotoxicity.
Neurotoxicity was quantified by trypan blue assay, referring to a previous report (Taguchi et al., 2003). After exposure to TBT, cell cultures were immediately stained with 1.5 % trypan blue for 10 min, fixed with formalin, and rinsed with physiological saline. Unstained cells were regarded as viable and stained cells were regarded as dead. The viability of the cultures was calculated as the percentage ratio of the number of unstained cells relative to the total cells counted. Over 200 cells per cover slip were randomly counted.
Measurement of released glutamate.
Released glutamate was measured with an Amplex Red Glutamic Acid/Glutamate Oxidase assay kit (Molecular Probes, Eugene, OR) according to the manufacturer's protocol. Culture medium was removed and cell cultures were incubated in HEPES-buffered salt solution containing 125 mM NaCl, 5 mM KCl, 1.2 mM KH2PO4, 1.2 mM MgSO4, 1.2 mM CaCl2, 6 mM glucose, and 25 mM HEPES. After the TBT stimulation, 50 μl of buffer was transferred to a 96-well microplate and mixed with 50 μl of reaction buffer containing Amplex red, alanine, glutamate oxidase, glutamate pyruvate transaminase, and horseradish peroxidase. The mixture was incubated for 30 min in the dark and the fluorescence was measured with the use of a fluorescence microplate reader (excitation at 538 nm and emission at 594 nm).
Western blotting.
After TBT treatment, cells were washed with PBS buffer and lysed in TNE buffer containing 50 mM Tris-HCl, 1% NP-40, 20 mM EDTA, 0.2 mM sodium orthovanadate, and 1 mM PMSF. The mixture was centrifuged at 13,500 rpm for 3 min, after which the supernatant was transferred to an Eppendorf tube. The supernatant was added to sample buffer containing 125 mM Tris-HCl, 4% SDS, 10% sucrose, 0.004% BPB, and 10% mercaptoethanol, then denatured at 95°C for 3 min. Protein was separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to a polyvinylidene difluoride membrane. The membrane was blocked with blocking buffer containing 5% skim milk for 1 h, then incubated with primary antibody (1:1000) overnight and secondary antibody (1:2000) for 1 h. The protein was detected with an enhanced chemiluminescence detection system.
Hoechst 33258 staining.
Cultured cells were stained with Hoechst 33258 to examine nuclear morphological changes. Cells were fixed with neutral formaldehyde for 30 min and then stained with 0.1 mg/ml Hoechst 33258 for 30 min. Cells were visualized and photographed under ultraviolet illumination, with a fluorescence microscope.
Statistics.
Data are expressed as mean ± SEM. Statistical evaluation of the data was performed by Student's t-test (Fig. 1 and Fig. 2A) or by analysis of variance (ANOVA) followed by the Newman-Keul's test (except Fig. 1 and Fig. 2A). A value of p < 0.05 was considered to be indicative of significance.
RESULTS
Neurotoxicity of TBT in Cortical Neurons
Toxicity of TBT in rat cortical neurons was examined by the trypan blue method (dead cells stained with trypan blue). Cortical neurons were exposed to various concentrations of TBT, and exposure of the cells to 500 nM TBT for 24 h resulted in a remarkable degree of neurotoxicity (survival of 49.8 ± 0.4%) (Fig. 1A), with increased numbers of trypan blue–stained cells (Fig. 2E, F). Next, cell viability was examined at 1, 3, 6, 12, and 24 h after 500 nM TBT exposure. Cell viability decreased when the cells were exposed to TBT for longer than 12 h (Fig. 1B).
Involvement of Glutamate in TBT Neurotoxicity
We examined whether glutamate is involved in TBT toxicity, because glutamate is believed to be a factor in neuronal death. First, the amount of glutamate in the medium was measured. The concentration of extracellular glutamate was increased significantly by exposure of the cells to 500 nM TBT for 30 min, and that increase was exposure time–dependent (Fig. 2A). Next, to investigate the involvement of glutamate release in TBT toxicity, we used NMDA receptor antagonists. Application of 5 μM MK-801, a noncompetitive NMDA receptor antagonist, alleviated TBT neurotoxicity (survival with TBT: 45.7 ± 3.3%; +MK-801: 87.8 ± 7.1%) (Fig. 2B). Further, 20 μM CNQX, a non-NMDA receptor antagonist, showed almost the same effect (survival with TBT: 52.1 ± 2.4%; +CNQX: 83.7 ± 5.3%) (Fig. 2C). These inhibitors greatly decreased the number of trypan blue–stained cells (Fig. 2G, H). Moreover, we investigated the involvement of voltage-dependent calcium channels (VDCC), because it has been reported that VDCC are involved in the toxicity of NMDA agonists (Fryer, 1999). Neuronal death caused by TBT was blocked by 10 μM nifedipine, a VDCC inhibitor (survival with TBT: 58.5 ± 2.7; +nifedipine: 84.7 ± 1.7%) (Fig. 2D).
Effect of Antioxidants on TBT Toxicity
It is known that oxidative stress contributes to glutamate toxicity (Campisi et al., 2004). Therefore, we examined the effect of antioxidants on TBT toxicity. We used water-soluble antioxidants (ascorbic acid and catalase) and a lipophilic antioxidant (-tocopherol). Co-administration of 100 μM ascorbic acid or 1000 U/ml catalase during TBT exposure failed to reduce TBT toxicity. In contrast, co-administration of 100 μM -tocopherol remarkably inhibited TBT-induced neurotoxicity (survival with TBT: 49.3 ± 3.7%; with -tocopherol 82.7 ± 4.5%) (Fig. 3).
Effect of MAPK Inhibitors on TBT Toxicity
Mitogen-activated protein kinase (MAPK) has been implicated in glutamate-induced neuronal death (Cao et al., 2004). Therefore we used inhibitors of the MAP kinase cascade to examine whether MAPKs are involved in TBT neurotoxicity. We found that 10 μM U-0126, an extracellular signal-regulated protein kinase (ERK) kinase inhibitor, reduced TBT neurotoxicity (survival with TBT: 46.4 ± 2.2%; +U-0126: 66.7 ± 1%) and that 10 μM SB202190, a p38 inhibitor, also significantly alleviated the neurotoxicity (survival with +SB202190: 76.7 ± 6.8%). However, 10 μM SP600125, a c-jun-N-terminal kinase (JNK) inhibitor, was ineffective against TBT-induced cell death (Fig. 4A). Further, we investigated whether TBT phosphorylated ERK and p38. As shown in Figure 4B and 4C, at 3 h after application of 500 nM TBT to the culture, p38 was phosphorylated, and at 6 h, ERK was activated. Phosphorylation of ERK and p38 was sustained until 24 h after exposure. We also examined the effects of glutamate receptor antagonists on ERK and p38 phosphorylation and found that MK-801 and CNQX decreased the phosphorylation of ERK, but not that of p38 (Fig. 4D and 4E).
Effect of Caspase-3 Inhibitor on TBT Toxicity
Because it has been reported that TBT activates caspase-3 in various cells (Nopp et al., 2002; Jurkiewicz et al., 2004), we used a caspase-3 inhibitor to confirm the involvement of caspase in TBT toxicity in cortical neurons. However, 10 μM of the caspase-3 inhibitor Ac-DEVD-CMK did not reduce TBT toxicity (Fig. 5A). Moreover, we investigated whether 500 nM TBT alters the nuclear morphology (an indication of apoptosis) of cortical neurons, and we found that it did not (Fig. 5C), although 2 μM TBT did induce nuclear fragmentation in PC12 cells, in agreement with a previous report (Fig. 5D) (Viviani et al., 1995).
DISCUSSION
Among organotin compounds, trimethyltin is neurotoxic, whereas tributyltin is primarily known as an immunotoxin (Snoeij et al., 1985). However, neurotoxicity of TBT has also been reported (O'Callaghan and Miller, 1988; Tsunoda et al., 2005), although its mechanism has been little investigated. Trimethyltin and tributyltin are thought to have different mechanisms of toxicity. For example, TMT is reported to exert neurotoxicity at least in part by binding specifically to the protein stannin, whereas TBT does not bind to stannin (Thompson et al., 1996, Buck et al., 2003).
Organotin compounds are important environmental pollutants, in part because of their toxicity. Braman et al. (1979) reported that methyltin compounds, including TMT, were detected at concentrations between 0.5 pM and 4 nM in human urine. Further, the butyltin concentration in human blood ranges from 50 nM to 400 nM (Whalen et al., 1999), and that in human liver is about 400 nM (Kannan and Falandysz, 1997; Mizuhashi et al., 2000). The TBT concentration in rock shells caught in polluted sea areas around Japan was about 5 μM (Horiguchi et al., 1994; Mizuhashi et al., 2000). The concentration of 500 nM TBT used in our experiments thus appears to be similar to levels of human exposure.
O'Callaghan and Miller (1988) reported that TBT decreased brain weight at dose levels that had no influence on body weight. This finding suggests that the brain may be particularly susceptible to injury by TBT. We found that exposure of cortical neurons to 500 nM TBT for 24 h significantly reduced their viability (Fig. 1). Thompson et al. (1996) reported that the 50%-toxic concentration of TBT for 48 h of exposure ranges from 5.5 μM to 9 μM for non-neuronal cells, such as NIH-3T3, TC7, and B cells, and that primary neuronal cells were more sensitive to TBT than non-neuronal cells. These results suggest that TBT exerts neuronal toxicity via a specific mechanism, at lower concentrations than the levels causing toxicity in other organs.
We examined the involvement of glutamate in TBT-induced neurotoxicity because glutamate is believed to play important roles in various types of neuronal death. We found that TBT increased extracellular glutamate concentration. The mechanism of glutamate release is not clear, but it is unlikely to be due to leakage from dead cells, because significant numbers of dead cells were not detected for up to 6 h after exposure to TBT. In our study glutamate release was not inhibited by NMDA receptor antagonists or a VDCC inhibitor (data not shown). Thus TBT appears to induce glutamate release via a Ca2+-independent pathway. Application of the glutamate receptor antagonists MK-801 and CNQX reduced the level of TBT toxicity (Fig. 2). These results support the idea that release of glutamate and activation of glutamate receptors are involved in TBT toxicity. It was previously reported that non-NMDA receptors were involved in TBT neurotoxicity in hippocampal slice cultures, but NMDA receptors and VDCC were not involved (Mizuhashi et al., 2000). Activation of both NMDA receptors and non-NMDA receptors may not necessarily be required for excitotoxicity (Kubo et al., 2001; Hakuba et al., 2003). The difference in the involvement of NMDA receptors may reflect differences in experimental conditions, such as brain region or exposure conditions. Also, VDCC plays important roles in other effects of glutamate excitotoxicity, such as ischemia (Hou et al., 2002). In our experimental model, nifedipine blocked neuronal death caused by TBT, and this result indicates that VDCC is involved in TBT toxicity in our system.
Moreover, we investigated the effects of combinations of inhibitors on TBT neurotoxicity, but we found that the combinations did not show potentiated inhibition (data not shown). Non-NMDA receptors are supposed to open after glutamate release, and their activation induces a change in the membrane potential. Glutamate and cancellation of Mg2+ blockade (caused by the change of membrane potential) are needed for NMDA receptor activation. Therefore, non-NMDA receptors should be activated before NMDA receptors and VDCC. This chain of activation is presumably why the effects of inhibitor combinations are the same as those of a single inhibitor.
Because reactive oxygen species (ROS) are generated by activation of the glutamate receptor (Campisi et al., 2004), we also examined whether ROS are involved in TBT toxicity. Application of -tocopherol blocked cell death induced by TBT, but catalase and ascorbic acid had no effect (Fig. 3). These results suggest that intracellular generation of ROS is important, because -tocopherol is more lipophilic and cell-permeable than the other two antioxidants. Ahlemeyer and Krieglstein (2000) showed that -tocopherol inhibited mitochondrial reactive ROS production, indicating that -tocopherol can decrease intracellular ROS. Ciani et al. (1996) reported that -tocopherol, but not ascorbic acid, decreased glutamate neurotoxicity. Kim et al. (1998) reported that glutamate exerts toxicity via lipid peroxidation. Several investigators have described that the most effective agents against glutamate toxicity appear to be chain-breaking antioxidants that block lipid peroxidation, such as -tocopherol (Ciani et al., 1996; Halliwell, 1992; Siesjo et al., 1989). These findings are consistent with our results.
Next, the effects of MAPK inhibitors were examined, because MAPKs play important roles in glutamate excitotoxicity (Mao et al., 2004). Yu et al. (2000) reported that TBT phosphorylated ERK, JNK, and p38 in human T lymphoblastoid cells CCRF-CEM. In our experimental model, a MEK inhibitor, U-0126, and a p38 MAPK inhibitor, SB202190, decreased TBT toxicity, but a JNK inhibitor did not (Fig. 4). Glutamate receptor antagonists inhibited phosphorylation of ERK, but not that of p38. These results indicate that ERK phosphorylation depends on glutamate receptors, and that there may also be another pathway involving p38 activation. It has been reported that glutamate activates ERK (Jiang et al., 2000). Activation of ERK is supposed to be characteristic of glutamate toxicity, and our results are consistent with this idea.
It is well known that caspase family members are the central executors of apoptosis, and among them, caspase-3 is the most important. Tributyltin activates caspase-3 in various cells, such as Jurkat T cells, eosinophils, neutrophils, and hepatocytes (Nopp et al., 2002, Jurkiewicz et al., 2004). Caspase-3 inhibitor did not protect neurons from TBT toxicity (Fig. 5A), and TBT did not induce any change in nuclear morphology (Fig. 5C). There have been many reports that glutamate-induced neurotoxicity is apoptotic (Kawasaki et al., 1997; Du et al., 1997) or necrotic (Sohn et al., 1998; Dessi et al., 1993). Our results indicate that glutamate neurotoxicity caused by TBT is not apoptotic, at least in our experimental system.
In summary, we have partially clarified the mechanism of TBT neurotoxicity in cultured rat cortical neurons (Fig. 6). Tributyltin at low concentration was toxic to cortical neurons through glutamate release and subsequent activation of glutamate receptors. The mechanism of the glutamate release induced by TBT remains unclear, but it is not simply leakage from dead cells. We demonstrated that intracellular ROS, ERK, and p38 MAPK were also involved in TBT neurotoxicity. It was shown that ERK phosphorylation depended on glutamate receptor activation, but its activation was not involved in p38 phosphorylation. These results suggest that there is a glutamate receptor–dependent pathway and a glutamate receptor–independent pathway. Moreover, the glutamate toxicity induced by TBT was not apoptotic. Thus, glutamate excitotoxicity seems to account for the high sensitivity of cortical neurons to TBT.
ACKNOWLEDGMENTS
This study was supported in part by a Grant-in Aid for Scientific Research from the Japan Society for the Promotion of Science.
REFERENCES
Ahlemeyer, B., and Krieglstein, J. (2000). Inhibition of glutathione depletion by retinoic acid and tocopherol protects cultured neurons from staurosporine-induced oxidative stress and apoptosis. Neurochem. Int. 36, 1–5.
Blandini, F., Porter, R. H., and Greenamyre, J. T. (1996). Glutamate and Parkinson's disease. Mol. Neurobiol. 12, 73–94.
Braman, R. S., and Tompkins, M. A. (1979). Separation and determination of nanogram amounts of inorganic tin and methyltin compounds in the environment. Anal. Chem. 51, 12–19.
Buck, B., Mascioni, A., Que, L., Jr., and Veglia, G. (2003). Dealkylation of organotin compounds by biological dithiols: Toward the chemistry of organotin toxicity. J. Am. Chem. Soc. 125, 13316–13317.
Campisi, A., Caccamo, D., Li Volti, G., Curro, M., Parisi, G., Avola, R., Vanella, A., and Ientile, R. (2004). Glutamate-evoked redox state alterations are involved in tissue transglutaminase upregulation in primary astrocyte cultures. FEBS Lett. 578, 80–84.
Cao, J., Semenova, M. M., Solovyan, V. T., Han, J., Coffey, E. T., and Courtney, M. J. (2004). Distinct requirements for p38alpha and c-Jun N-terminal kinase stress-activated protein kinases in different forms of apoptotic neuronal death. J. Biol. Chem. 279, 35903–35913.
Chow, S. C., Kass, G. E., McCabe, M. J., Jr., and Orrenius, S. (1992). Tributyltin increases cytosolic free Ca2+ concentration in thymocytes by mobilizing intracellular Ca2+, activating a Ca2+ entry pathway, and inhibiting Ca2+ efflux. Arch. Biochem. Biophys. 298, 143–149.
Ciani, E., Groneng, L., Voltattorni, M., Rolseth, V., Contestabile, A., and Paulsen, R. E. (1996). Inhibition of free radical production or free radical scavenging protects from the excitotoxic cell death mediated by glutamate in cultures of cerebellar granule neurons. Brain Res. 728, 1–6.
Collingridge, G. L., and Laster, R. A. (1989). Excitatory amino acid receptors in the vertebrate central nervous system. Pharmacol. Rev. 41, 143–210.
Dessi, F., Charriaut-Marlangue, C., Khrestchatisky, M., and Ben-Ari, Y. (1993). Glutamate-induced neuronal death is not a programmed cell death in cerebellar culture. J. Neurochem. 60, 1953–1955.
Du, Y., Bales, K. R., Dodel, R. C., Hamilton-Byrd, E., Horn, J. W., Czilli, D. L., Simmons, L. K., Ni, B., and Paul, S. M. (1997). Activation of a caspase 3-related cysteine protease is required for glutamate-mediated apoptosis of cultured cerebellar granule neurons. Proc. Natl. Acad. Sci. U.S.A. 94, 11657–11662.
Ema, M., Itami, T., and Kawasaki, H. (1991). Behavioral effects of acute exposure to tributyltin chloride in rats. Neurotoxicol. Teratol. 13, 489–493.
Farina, M., Dahm, K. C., Schwalm, F. D., Brusque, A. M., Frizzo, M. E., Zeni, G., Souza, D. O., and Rocha, J. B. (2003). Methylmercury increases glutamate release from brain synaptosomes and glutamate uptake by cortical slices from suckling rat pups: Modulatory effect of ebselen. Toxicol. Sci. 73, 135–140.
Feldman, R. G., White, R. F., and Eriator, I. I. (1993). Trimethyltin encephalopathy. Arch. Neurol. 50, 1320–1324.
Fryer, H. J., Knox, R. J., Strittmatter, S. M., and Kalb, R. G. (1999). Excitotoxic death of a subset of embryonic rat motor neurons in vitro. J. Neurochem. 72, 500–513.
Hakuba, N., Matsubara, A., Hyodo, J., Taniguchi, M., Maetani, T., Shimizu, Y., Tsujiuchi, Y., Shudou, M., and Gyo, K. (2003). AMPA/kainate-type glutamate receptor antagonist reduces progressive inner hair cell loss after transient cochlear ischemia Brain Res. 979, 194–202.
Halliwell, B. (1992). Reactive oxygen species and the central nervous system. J. Neurochem. 59, 1609–1623.
Horiguchi, T., Shiraishi, H., Shimizu, M., and Morita, M. (1994). Imposex and organotin compounds in Thais clavigera and T. bronni in Japan. J. Mar. Biol. Assoc. U.K. 74, 651–669.
Hou, X. Y., Zhang, G. Y., Yan, J. Z., Chen, M., and Liu, Y. (2002). Activation of NMDA receptors and L-type voltage-gated calcium channels mediates enhanced formation of Fyn-PSD95-NR2A complex after transient brain ischemia. Brain Res. 955, 123–132.
Hynd, M. R., Scott, H. L., and Dodd, P. R. (2004). Glutamate-mediated excitotoxicity and neurodegeneration in Alzheimer's disease. Neurochem. Int. 45, 583–595.
Jiang, Q., Gu, Z., Zhang, G., and Jing, G. (2000). N-methyl-D-aspartate receptor activation results in regulation of extracellular signal-regulated kinases by protein kinases and phosphatases in glutamate-induced neuronal apototic-like death. Brain Res. 887, 285–292.
Jurkiewicz, M., Averill-Bates, D. A., Marion, M., and Denizeau, F. (2004). Involvement of mitochondrial and death receptor pathways in tributyltin-induced apoptosis in rat hepatocytes. Biochim. Biophys. Acta 1693, 15–27.
Kannan, K., and Falandysz, Z. (1997). Butyltin residues in sediment, fish, fish eating birds, harbour porpoise and human tissues from the Polish coast of the Baltic Sea. Mar. Pollut. Bull. 34, 203–207.
Kawanishi, T., Kiuchi, T., Asoh, H., Shibayama, R., Kawai, H., Ohata, H., Momose, K., and Hayakawa, T. (2001). Effect of tributyltin chloride on the release of calcium ion from intracellular calcium stores in rat hepatocytes. Biochem. Pharmacol. 62, 863–872.
Kawasaki, H., Morooka, T., Shimohama, S., Kimura, J., Hirano, T., Gotoh, Y., and Nishida, E. (1997). Activation and involvement of p38 mitogen-activated protein kinase in glutamate-induced apoptosis in rat cerebellar granule cells. J. Biol. Chem. 272, 18518–18521.
Kim, Y. C., Kim, S. R., Markelonis, G. J., and Oh, T. H. (1998). Ginsenosides Rb1 and Rg3 protect cultured rat cortical cells from glutamate-induced neurodegeneration. J. Neurosci. Res. 53, 426–432.
Kubo, T., Yokoi, T., Hagiwara, Y., Fukumori, R., Goshima, Y., and Misu, Y. (2001). Characteristics of protective effects of NMDA antagonist and calcium channel antagonist on ischemic calcium accumulation in rat hippocampal CA1 region. Brain Res. Bull. 54, 413–419.
Mao, L., Tang, Q., Samdani, S., Liu, Z., and Wang, J. Q. (2004). Regulation of MAPK/ERK phosphorylation via ionotropic glutamate receptors in cultured rat striatal neurons. Eur. J. Neurosci. 19, 1207–1216.
Mizuhashi, S., Ikegaya, Y., and Matsuki, N., (2000). Cytotoxicity of tributyltin in rat hippocampal slice cultures. Neurosci. Res. 38, 35–42.
Nopp, A., Lundahl, J., and Stridh, H. (2002). Caspase activation in the absence of mitochondrial changes in granulocyte apoptosis. Clin. Exp. Immunol. 128, 267–274.
O'Callaghan, J. P., and Miller, D. B. (1988). Acute exposure of the neonatal rat to tributyltin results in decreases in biochemical indicators of synaptogenesis and myelinogenesis J. Pharmacol. Exp. Ther. 246, 394–402.
Rao, S. D., and Weiss, J. H. (2004). Excitotoxic and oxidative cross-talk between motor neurons and glia in ALS pathogenesis Trends Neurosci. 27, 17–23.
Sarradin, P. M., Lapaquellerie, Y., Astruc, A., Latouche, C., and Astruc, M. (1995). Long term behaviour and degradation kinetics of tributyltin in a marina sediment. Sci. Total Environ. 170, 59–70.
Siesjo, B. K., Agardh, C. D., and Bengtsson, F. (1989). Free radicals and brain damage. Cerebrovasc. Brain Metab. Rev. 1, 165–211.
Snoeij, N. J., van Iersel, A. A., Penninks, A. H., and Seinen, W. (1985). Toxicity of triorganotin compounds: Comparative in vivo studies with a series of trialkyltin compounds and triphenyltin chloride in male rats. Toxicol. Appl. Pharmacol. 81, 274–286.
Snoeij, N. J., Penninks, A. H., and Seinen, W. (1987). Biological activity of organotin compounds—An overview. Environ. Res. 44, 335–353.
Sohn, S., Kim, E. Y., and Gwag, B. J. (1998). Glutamate neurotoxicity in mouse cortical neurons: Atypical necrosis with DNA ladders and chromatin condensation. Neurosci. Lett. 240, 147–150.
Taguchi, R., Nishikawa, H., Kume, T., Terauchi, T., Kaneko, S., Katsuki, H., Yonaga, M, Sugimoto, H., and Akaike, A. (2003). Serofendic acid prevents acute glutamate neurotoxicity in cultured cortical neurons. Eur. J. Pharmacol. 477, 195–203.
Thompson, T. A., Lewis, J. M., Dejneka, N. S., Severs, W. B., Polavarapu, R., and Billingsley, M. L. (1996). Induction of apoptosis by organotin compounds in vitro: Neuronal protection with antisense oligonucleotides directed against stannin. J. Pharmacol. Exp. Ther. 276, 1201–1216.
Tsuda, T., Inoue, T., Kojima, M., and Aoki, S. (1995). Daily intakes of tributyltin and triphenyltin compounds from meals. J. AOAC Int. 78, 941–943.
Tsunoda, M., Konno, N., Nakano, K., and Liu, Y. (2005). Altered metabolism of dopamine in the midbrain of mice treated with tributyltin chloride via subacute oral exposure. Environ. Sci. 11, 209–219.
Whalen, M. M., Loganathan, B. G., and Kannan, K. (1999) Immunotoxicity of environmentally relevant concentrations of butyltins on human natural killer cells in vitro. Environ. Res. 81, 108–116.
van Wezel, A. P., and van Vlaardingen, P. (2004). Environmental risk limits for antifouling substances. Aquat. Toxicol. 66, 427–444.
Viviani, B., Rossi, A. D., Chow, S. C., and Nicotera, P. (1995). Organotin compounds induce calcium overload and apoptosis in PC12 cells. Neurotoxicology 16, 19–25.
Yu, Z. P., Matsuoka, M., Wispriyono, B., Iryo, Y., and Igisu, H. (2000). Activation of mitogen-activated protein kinases by tributyltin in CCRF-CEM cells: Role of intracellular Ca2+. Toxicol. Appl. Pharmacol. 168, 200–207.(Yusuke Nakatsu, Yaichiro Kotake, Kazuya )