ToxSci Advance Access originally published online on January 6, 2007
Toxicological Sciences 2007 96(2):268-278; doi:10.1093/toxsci/kfl198
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In Vitro Toxicity of Tetrabromobisphenol-A on Cerebellar Granule Cells: Cell Death, Free Radical Formation, Calcium Influx and Extracellular Glutamate
* Norwegian Defence Research Establishment, Division for Protection, PO Box 25, N-2027 Kjeller, Norway
Norwegian Institute for Air Research, PO Box 100, N-2027 Kjeller, Norway
Department of Biochemistry, Institute of Basic Medical Science, University of Oslo, PO Box 1112, Blindern, N-0317 Oslo, Norway
1 To whom correspondence should be addressed. Fax: + 47 63807509. E-mail: trine.reistad{at}ffi.no.
Received November 7, 2006; accepted December 20, 2006
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Tetrabromobisphenol-A (TBBPA) is one of the worlds most widely used brominated flame retardant. The present study reports effects of TBBPA on primary cultures of cerebellar granule cells (CGC). Using the trypan blue exclusion assay, we show that TBBPA induces death of CGC at low micro molar concentrations. Cell death was reduced by the NMDA receptor antagonist MK-801 (3µM), the antioxidant vitamin E (50µM), and in calcium-free buffer. We further demonstrate that TBBPA's toxicity was accompanied by apoptosis-like nuclear shrinkage, chromatin condensation, and DNA fragmentation. Other hallmarks of apoptosis such as caspase activity were, however, absent, indicating an atypical form of apoptosis. TBBPA increased intracellular free calcium in a concentration-dependent manner. TBBPA also induced an increase in extracellular glutamate in a time-dependent manner. TBBPA gave a concentration-dependent increase information reactive oxygen species (ROS) of measured with 2,7-dichlorofluorescein diacetate. The ROS formation was inhibited by the extracellular signal-regulated protein kinase (ERK) inhibitor U0126 (10µM), the tyrosine kinase inhibitor erbstatin-A (25µM), eliminating calcium from the buffer and by the superoxide dismutase inhibitor diethyldithio-carbamic acid (DDC, 100µM). Further analysis with Western blot confirmed phosphorylation of ERK1/2 after exposure to TBBPA. We found that TBBPA induces ROS formation, increases intracellular calcium, extracellular glutamate, and death of CGC in vitro at concentrations comparable to those of polychlorinated biphenyl. These findings implicate TBBPA as a predicted environmental toxin and bring out the importance of awareness of its hazardous effects.
Key Words: tetrabromobisphenol-A (TBBPA); cerebellar granule cells (CGC); reactive oxygen species (ROS); cell death; glutamate; calcium.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Tetrabromobisphenol-A (TBBPA) is the most utilized brominated flame retardant (BFR) in the world today with an estimated annual demand of approximately 120,000 metric tons. This represents approximately half of the annual demand of BFRs (de Wit, 2002
). TBBPA is primarily used as a chemical bound flame retardant (
90%) and is therefore not expected to reach the environment in larger amounts. In spite of this, TBBPA is found in significant amounts in the environment and in human samples. TBBPA in concentrations of 67 pg/g lw and 0.3–1.8 ng/g lw was found in milk from Norwegian mothers (Thomsen et al., 2002) and in human plasma samples (Thomsen et al., 2001), respectively. Sellstrom and Jansson (1995) showed that this compound could leak from treated products and Osako et al. (2004) reported up to 620 ng/l TBBPA in leachate samples from landfills in Japan.
A few toxicological studies with TBBPA show that this compound exerts some interesting effects in vitro. TBBPA has some structural resemblance with thyroxin and is shown to possess a high affinity to transthyretin (Meerts et al., 2000) and thyroid hormone receptors (Kitamura et al., 2005). Mariussen and Fonnum (2003) found that TBBPA inhibits the vesicular and synaptosomal transport of the neurotransmitters dopamine and glutamate at similar concentration level as previously shown for polychlorinated biphenyls (PCBs). Recently, we showed that TBBPA induces a potent activation of extracellular signal-regulated protein kinase (ERK1/2) in human neutrophil granulocytes, an increase in cytosolic calcium, ß-nicotineamide adenine dinucleotide phosphate (NADPH)-oxidase activation and a potent formation of reactive oxygen species (ROS) (Reistad et al., 2005). In a preliminary work by Strack et al. (2004) it was reported that TBBPA influences ERK phosphorylation in different types of mammalian cells. Aquatic organisms are regarded the most susceptible for TBBPA exposure, and TBBPA has high acute toxicity to algae, molluscs, crustaceans, and fish. Canesi et al. (2005) have shown that TBBPA induces activation of MAP kinases followed by a stimulation of immune functions in mussel hemocytes. This indicates that the same transduction pathway is involved in mediating the effects of TBBPA in mammalian and invertebrate cells. A recent report shows that pre- and postnatal exposure of mice to TBBPA induces focal necrosis of hepatocytes and inflammatory cell infiltration in the liver of treated dams and offspring (Tada et al., 2006). Fukuda et al. (2004) observed nephrotoxicity after repeated dose exposure of TBBPA in newborn rats. These findings implicate TBBPA as a predicted environmental toxin, and due to its massive use it is of importance to bring out an awareness of its potential hazardous effects.
In this study, we examined the effects of TBBPA on cultured cerebellar granule cells (CGC) on a cellular level with focus on oxidative stress, calcium influx, extracellular glutamate, and cell death. CGC is a well-suited model to investigate cytotoxic effects and is extensively used to elucidate the mechanisms of action of a range of environmental toxicants such as ortho-chlorinated PCBs (Kodavanti et al., 1998; Mariussen et al., 2002), dioxins (Kim and Yang, 2005), and organic solvents (Dreiem et al., 2005). A preliminary report has previously been presented in a conference proceeding (Reistad et al., 2003).
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Chemicals.
TBBPA (Lot GL16) is the original technical material from Great Lake Company and was obtained from Promochem Kungsbacka (Sweden). The PCB mixture Aroclor 1254 (Lot 124-191-B) was purchased from AccuStandard Inc (New Haven, CT). Stock solutions were prepared by dissolving the compounds in dimethyl sulfoxide (DMSO). The final DMSO concentration was always less than or equal to 0.01%. Bisphenol-A, 2,7-dichlorofluorescein diacetate (DCFH-DA), cytosine ß-D-arabinofuranoside (ARA-C), DDC, D-glucose-6-phosphate (G6P), NADPH, Hoechst 33342 (±)
-tocopherol (vitamin E), SB203580, poly-L-lysine, deoxyribonuclease, Triton X-100, Tween 20, trypsin inhibitor, trypsin, bovine serum albumin, ionomycin, L-glutamine, methanol, DMSO, -aminoadipate (-aaa), o-phthaldialdehyde (OPA), and trypan blue were from Sigma-Aldrich Oslo, Norway. U0126 was obtained from Promega Corporation (Madison, WI). MK-801 was purchased from RBI (Natick, MA). Hanks' Balanced Salt Solution (HBSS) (CaCl2 1.26mM, KCl 5.37mM, KH2PO4 0.44mM, MgCl2 0.49mM, MgSO4 0.41mM, NaCl 140mM, NaHCO3 4.17mM, Na2HPO4 0.34mM, D-glucose 5.55mM), HEPES buffer, basal medium Eagle's (BME), heat-inactivated fetal bovine serum (FBS), and penicillin/streptomycin were purchased from GibcoBRL Paisley (UK). Enhanced chemiluminescence (ECL) reagent was from Amersham Biosciences, Oslo Norway. Monoclonal mouse anti-phospho-ERK antibody (Tyr204) and polyclonal rabbit anti-ERK2 antibody were from Santa Cruz Biotechnology (Santa Cruz, CA). Horseradish peroxidase (HRP)-conjugated rabbit-anti-mouse antibody and HRP-conjugated goat-anti-rabbit antibody were purchased from DAKO A/S (Glostrup, Denmark). 2,5-Dihydroxymethylcinnamate (erbstatin-A), FK-506 (tacrolimus), caspase inhibitor III, caspase-3 substrat IV (fluorigenic), and fura-2/AM were from Calbiochem Novabiochem Corp, Oslo, Norway. Fura-2/AM was from Molecular Probes Inc (Holland). Other reagents were from standard commercial suppliers.
Preparation of CGC.
CGC in culture provide a well-characterized neuronal cell population suited for morphological and biochemical studies (Drejer et al., 1983; Kingsbury et al., 1985). Primary cultured neurons from rat cerebellum were isolated mainly as described previously (Schousboe et al., 1989). Cerebella from 7-day-old pups were dissected under sterile conditions. The brain tissue was then mechanically dissected from the meninges, trypsinated and chopped in buffered solution. The cells were seeded (106/ml) in BME (containing 10% heat-inactivated FBS, 25mM KCl, 2mM L-glutamine, 100 IU/ml penicillin, and 100 µg/ml streptomycin) and plated into 50 mm–cell culture dishes that had been previously coated with poly-L-lysine (10 µg/ml). After 16–22 h, ARA-C was added (final concentration 2.5 µg/ml) to prevent growth of glial cells. The cells were grown for 6–8 days (37°C with 5% CO2) before exposure to TBBPA in culture media without FBS.
Assay for measuring survival of cerebellar granule cells.
The growth media were removed from the plated cells and replaced by 4 ml prewarmed culture media without FBS containing TBBPA. Thereafter, the granule cells were incubated for 24 h prior to determining neuron survival using the trypan blue exclusion assay. The cells were incubated with 100 µl of 1% trypan blue mixture at 37°C for 3 min followed by counting the relative number of dead cells (blue) using light microscopy. Failure to exclude trypan blue reflects a loss of membrane integrity associated with necrosis. The effects on cell death are presented as estimated LC50 (concentration causing 50% cell death) values from nonlinear regression analysis (Demo version of SlideWrite Plus Version 6, Advanced Graphics Software Inc) and represent the mean (± SEM) of five separate experiments assayed in duplicate.
Lactate dehydrogenase assay.
Leakage of lactate dehydrogenase (LDH) was assessed as an index of cell injury (Koh and Choi, 1987). The measurements were performed as described elsewhere (Dreiem et al., 2005). In brief, cells (2 x 106/ml) were exposed to the test compounds for 24 h. Supernatant from each sample was spun down and transferred to sample tubes and stored at 4°C until measured (within 2 h). LDH measurements were performed by transfer of 50 µl aliquots of the supernatant to the wells of a 96-well microplate with glass bottom, and the volume was adjusted to 200 µl with 0.1M KPO4 buffer (pH 7.5). The reactions were started by automated injection of 25 µl of a 0.6 mg/ml stock solution of NADH (final concentration 100 µM) followed by automated injection of 12 µl of a 6 mg/ml stock solution of pyruvate (final concentration 3 mM). The LDH activity was measured, using a BMG FLUOstar Optima fluorimeter, from the decay rate of NADH fluorescence for 30 min at 28°C. The LDH activity was calculated off line and is given as the rate constant of the decrease in fluorescence emission at 460 nm (excitation wavelength 340 nm). The LDH activity (fluorescence units/s) is not a direct measure of the number of dead cells but gives a qualitative measure of the relative amount of cell necrosis; 0.01% Triton was added 30 min prior to LDH measurement as a measure for total LDH release. In Table 1, the values are shown as percentage of Triton ± SEM.
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Biotransforming enzyme systems.
A rat liver postmitochondrial (S9) fraction was used as a source for biotransformation enzymes, as described by Dreiem and Fonnum (2004)
. The S9 fraction was prepared from the liver of a male Wistar rat (250g) given a single ip injection of a mixture of PCBs, Aroclor 1254 (500 mg/kg), to induce cytochrome P-450. As a control, S9 fraction was also made from one unexposed rat liver. Five days after injection, the rat was killed by decapitation. The liver was dissected out and homogenized in 3x vol 0.15M KCl in a Potter-Elvehjelm glass/teflon apparatus. The homogenate was centrifuged for 10 min at 9000g and aliquots of the supernatant (S9) were immediately frozen in liquid nitrogen and stored at – 75°C. S9 was added directly to the incubation medium to a concentration of 0.18 mg protein/ml. In most of the experiments, NADPH and G6P was added (final concentrations 300 and 250µM, respectively) as cofactors for the cytochrome P-450 monooxygenases.
Examination of nuclear morphology.
Cells were exposed to the test substances in high potassium HBSS (25mM KCl) for 24 h at 37°C, with 5% CO2 and constant humidity. Condensed and fragmented nuclei were evaluated by intercalation of the fluorescent probe Hoechst 33258 (final concentration 0.1 mg/ml) into nuclear DNA. Hoechst 33258, which is cell permeable and labels both intact and apoptotic nuclei, was visualized by fluorescence microscopy (Olympus IMT-2) and pictures were taken with an Olympus Camedia C-5060 digital camera. In these experiments, 50nM okadaic acid was used as a positive control. All experiments were repeated three to four times.
Internucleosomal DNA fragmentation.
Cells were exposed in high potassium HBSS as described for examination of nuclear morphology, and harvested in 400 µl PBS before DNA isolation. We used an apoptotic DNA ladder kit (Roche diagnostics Scandinavia Oslo, Norway AS) designed for purification of nucleic acids from different sample materials like whole blood and cultured cells, for detection of DNA ladder. After lysis of the cells in binding buffer, the lysate was applied to a filter tube with glass fiber fleece and passed through the glass fiber fleece by centrifugation (1 min at 8000 rpm). During the procedure, the DNA binds specifically to the surface of glass fibers. Residual impurities were removed by a wash step and subsequently DNA was eluted in elution buffer. After incubation for 30 min at 37°C with RNAse (final concentration 2 µg/ml), isolated DNA was mixed with loading buffer (final concentration 0.1% sodium dodecyl sulphate [SDS], 3% glycerol, bromphenol blue to color) and fragments were separated by electrophoresis on a 1% agarose gel with 0.008 µg/ml ethidiumbromide at 75 V for 90 min. DNA bands were visualized with a UV light source and the gel was photographed.
Measurement of caspase-3 activity.
Caspase-3 activity was determined by measuring the release of 7-amido-4-methylcoumarin (AFC) from the caspase substrate enyloxycarbonyl-asp-glu-val-asp-7-amido-4-methylcoumarin (Z-DEVD-AFC). Cells were exposed as described for examination of nuclear morphology. After indicated time, the medium was removed and 300 µl exposure buffer (255mM sucrose, 20mM HEPES, 50mM KCl, 2.5mM MgCl2, 0.01% Nonidet P40, 1mM DTT) was added to each dish. The cells were harvested and 150 µl was transferred in duplicate to a 96-well microtiter plate. Each well was supplemented with 33µM of the caspase-3 substrate Z-DEVD-AFC, a fluorigenic substrate preferentially cleaved by caspase-3 and -7. Production of fluorigenic AFC was assayed by monitoring fluorescence in a Perkin Elmer LW50B spectrofluorimeter for 30 min (excitation wavelength 400 nm, emission wavelength 505 nm, slit widths 10 nm). In these experiments, 50 nM okadaic acid was used as a positive control, and measurements were done after 6, 12, 18, and 24 h.
Intracellular Ca2+ measurement.
Changes in [Ca2+]i were measured with the aid of a plate reader (FLUOstar Optima, BMG Labtechnologies, Germany) using the membrane permeable Ca2+-binding fluorescent probe fura-2/AM. Primary cultures of CGC were cultured on 60 x 24 mm rectangular coverslips (Menzel-gläser purchased from VWR (Oslo, Norway) coated with poly-L-lysine. Coverslips with cells were incubated at 37°C in a humidified 5% CO2 atmosphere for 30 min with 5µM fura-2/AM in the standard saline solution containing 15mM Tris/HCl, 140mM NaCl, 3.5mM KCl, 1.25mM CaCl2, 1mM MgSO4, 1.2mM Na2HPO4 and 5mM glucose. Four coverslips were mounted in a custom-built chamber giving 12 isolated wells on each coverslip and a total of 48 wells for each experiment. Excitation, obtained from a UV lamp and a filter wheel, was alternated between 340 and 380 nm and emission was measured at 510 nm. The bandwidth was 10 nm for all filters. The system was calibrated with buffer solutions, fura-2 acid, and a calcium buffer kit obtained from Molecular Probes Inc. Background autofluorescence was determined from coverslips with cells (not preincubated with fura-2/AM). Cytosolic calcium was estimated from the 340/380 fluorescence emission ratio as described previously (Grynkiewicz et al., 1985). For single cell–imaging experiments, [Ca2+]i was measured with standard digital-imaging techniques with the aid of an inverted stage microscope (Olympus IMT 70). Briefly, cells were plated on 18 mm diameter round coverslips (Menzel). Cells were loaded with fura-2/AM similar to the protocol above. The UV lamp, monochromator, control hardware, air-cooled CCD camera, and image acquisition software were from TILL-photonics, Germany. The program allowed for simultaneous measurement of a number of regions (typically about 30) each covering one cell. One 340/380 ratiometric measurement of emission at 510 nm was obtained every 2–5 s. The imaging data were stored on a PC, and background subtraction, calcium estimation, and calibration were performed offline similar to that described for the plate reader experiments.
Assay for measuring released amino acids with HPLC.
CGC were washed twice in HBSS before incubation with 10µM TBBPA in high potassium HBSS for different time spans (0–180 min). After the indicated time, 450 µl incubation medium was mixed with 50 µl -aaa (final concentration 10µM). Amino acids were quantified by HPLC and fluorescence detection after precolumn derivatization with OPA, using -aaa as an internal concentration standard as described (Hassel et al., 1997).
Assay for measuring ROS formation.
Formation of ROS was measured by using the fluorescent probe DCFH-DA as previously described (Myhre et al., 2000). The DCF assay is an attractive and sensitive method as an overall index for oxidative stress in biological systems. It is reported to detect several types of reactive molecules such as H2O2 (in combination with cellular peroxidases), OONO- and OH (Myhre et al., 2003). The cells were preincubated with DCFH-DA, which is permeable across cell membranes, and inside the cell the acetate moiety is cleaved by cellular esterases. DCFH readily reacts with ROS to form the fluorescent DCF. CGC were loaded with 5µM DCFH-DA for 20 min at 37°C with 5% CO2 and constant humidity. The medium was then replaced with 1.5 ml prewarmed HBSS containing TBBPA and/or potentially neuroprotective substances. The cells were harvested and transferred in triplicate to 250 µl wells (microtiter plate, 96 wells) for ROS measurements. The formation of the fluorescent-oxidized derivate of DCFH, namely DCF, was measured with a luminescence spectrometer at 37°C for 180 min (excitation wavelength 485 nm, emission wavelength 530 nm). In each experiment, we used two cell culture dishes for each TBBPA concentration.
Western blotting.
The CGC were prepared as described above and incubated at 37°C with 5% CO2 for 7–8 days before stimulation. Prior to use, the culture medium was removed and replaced with fresh 1.5 ml HBSS, supplemented with 20mM HEPES and 10mM glucose. The CGC were incubated (5–60 min) in TBBPA (5–20µM), washed in 0.9% NaCl, and immediately lysed in 150 µl 20mM phosphate buffer (pH 7.4) with 0.1% Triton X-100. Before SDS-polyacrylamide gel electrophoresis (PAGE), aliquots of lysate were mixed with loading buffer (final concentration 3% SDS, 5% glycerol, 62.5mM Tris/HCl [pH 6.9], 0.1% bromphenol blue, 6% 2-mercaptoetanol). Total cell lysate was boiled for 2 min at 95°C and analyzed on a 3% stacking/12% separating SDS-PAGE gel (2 h at 90 V). The separated proteins were then electrophoretically (50 mA) transferred to nitrocellulose membranes (0.45 µm) overnight. The nitrocellulose blots were incubated in blocking buffer (Tris-buffered saline containing 0.05% Tween 20 [TBST] and 5% low-fat dry milk) for 1 h and probed with phospho-ERK1/2 mAb (1:200 dilution in blocking buffer) for 1 h. The blots were washed in TBST (6 x 5 min) and then incubated with polyclonal HRP-conjugated anti-mouse antibody (1:1000 dilution in blocking buffer) for 1 h. After washing in TBST (6 x 5 min), the blots were exposed to ECL reagent for 1 min. The signals were visualized on X-OmatBlue XB-1 film (Kodak). The experiments were repeated three times. The membranes were then stripped in 100mM ß-mercaptoethanol, 2% SDS, and 62.5mM Tris/HCl (pH 7.6) for 30 min at 50°C and proceeded again with a rabbit polyclonal anti-ERK2 primary antibody and peroxidase-conjugated goat anti-rabbit secondary antibody to confirm equal amounts of protein in each well.
Statistical analysis.
Differences between controls and treated groups were evaluated using a two-way Student's t-test (paired, two-tail distribution), or by one-way ANOVA followed by Dunnett's two-sided post hoc test. The calculations were performed using SPSS 11.5.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Cell Viability
The cytotoxic effects of different concentrations of TBBPA (2–20µM), on the CGC, were evaluated after 24 h exposure. Relatively low concentrations of TBBPA, LC50 values about 7µM, induced cell death as demonstrated by their failure to exclude trypan blue (Fig. 1). Cells treated with vehicle (DMSO as control) showed no sign of toxicity (10 ± 0.8%) compared to the control (9 ± 0.7%) after 24 h. The nonbrominated bisphenol-A did not induce cell death at the concentrations tested (5–15µM). This observation demonstrates that bromination is important for the cytotoxic effect in vitro. The release of LDH from the cells to the incubation medium was also used as a measure of cell death and supports the trypan blue assay. TBBPA gave a concentration-dependent increase in LDH release (Table 1). TBBPA increased LDH significantly compared to unexposed control cells (DMSO) at 10 and 20µM. Bisphenol-A did not induce cell death at the concentrations tested.
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0.01, ***p Inhibition of Cytotoxicity
MK-801 (3µM) reduced cell death from TBBPA by 86%. Calcium-free medium with EGTA reduced the cell death by 60%. An involvement of ROS was indicated because vitamin E (50µM) inhibited cell death by 46% (Fig. 1). Furthermore, S9 hepatic microsomal fraction was used to discern the mediating influence of metabolic bioactivation in TBBPA's cytotoxicity. In the presence of liver, S9 fraction from rats treated with Aroclor 1254, an inducer of mixed-function oxidase activity, and cofactors, TBBPA-induced cell death was reduced by 68%. This indicates that TBBPA itself is the ultimate toxicant after TBBPA exposure (Fig. 2). Cells were also exposed to TBBPA in combination with S9 fraction from an unexposed rat liver to ensure that the observed effect was not due to protein binding of TBBPA. Cell death after TBBPA exposure was not inhibited in these experiments (data not shown).
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The tyrosine kinase inhibitor erbstatin-A (25µM), the MAPK/ERK kinase (MEK) 1/2 inhibitor U0126 (10µM), and the superoxide dismutase (SOD) inhibitor DDC (100µM) did not inhibit cell death induced by TBBPA.
Nuclear Morphology
In order to attempt to discriminate between apoptotic and necrotic cell death, we screened cells for typical morphological features after exposure to TBBPA, with the aid of Hoechst 33342 nuclear staining; 5µM TBBPA induced changes in nuclear morphology associated with chromatin condensation giving 63% apoptotic cells after incubation for 24 h. Okadaic acid was used as a positive control for apoptosis and showed 94% apoptotic cell death (Fig. 3). Cells treated with vehicle alone (DMSO as control) showed no signs of toxicity.
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Internucleosomal DNA Fragmentation
DNA isolated from cells exposed to 5µM TBBPA for 24 h was subjected to electrophoresis on agarose gels to separate DNA fragments formed by internucleosomal DNA cleavage, a late hallmark of apoptosis. Okadaic acid is known to produce DNA fragments (Bezvenyuk et al., 2000
) and was used as a positive control. In cells treated with okadaic acid (50nM) or TBBPA (5µM), there was a distinct DNA ladder after 24 h exposure (Fig. 4).
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Caspase Activity
We investigated caspase activation, another hallmark of the apoptotic pathway, using a caspase-3 substrate IV (Z-DEVD-AFC) and caspase inhibitor III. No increase in caspase-3 activity was observed after 6, 12, 18, and 24 h incubation with 5µM of TBBPA compared with the controls. Okadaic acid (50nM) was used as a positive control and showed an increase in activity after 12 (49 ± 4
fluorescence/min), 18 (35 ± 2 fluorescence/min), and 24 h (24 ± 2 fluorescence/min) compared with the controls (12 ± 2, 11 ± 1, 8 ± 2 fluorescence/min, respectively). Caspase inhibitor III, which is a broad-spectrum caspase inhibitor, did not reduce the TBBPA-induced cell death (data not shown).
Intracellular-free Calcium
The CGC were exposed to different concentrations of TBBPA, and changes in intracellular-free calcium for 20 min were measured using the membrane permeable Ca2+-binding fluorescent probe fura-2/AM. As shown in Figure 5, TBBPA increased calcium in a concentration-dependent manner. There was an increase in calcium concentration at 10µM although the lowest dose that gave a significant increase in Ca2+ was 30µM. Figure 5A shows the Ca2+ increase of a representative experiment and the maximum response was 330 ± 38nM (mean ± SEM, six parallels). In two other experiments, the maximum intracellular calcium measured was 151 ± 35nM and 246 ± 40nM. The difference in intracellular calcium is probably due to variability in cell number from batch to batch and the loading efficiency with fura-2/AM.
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The imaging pictures (Figs. 5C–5G) visualize the calcium influx after exposure for TBBPA with the aid of a pseudocoloring scheme.
HPLC Measurements
CGC were exposed to 10µM TBBPA for different time spans (0–180 min), and TBBPA increased the extracellular concentration of glutamate and aspartate in a time-dependent manner (Fig. 6). After 3 h, the concentration of glutamate and aspartate was 8.8 ± 1.5µM and 3.8 ± 0.6µM, respectively, compared to not detectable in the controls.
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ROS Formation
CGC were exposed to different concentrations of TBBPA, and ROS formation was assessed by measuring DCFH oxidation after 3 h. TBBPA induced a concentration-dependent increase in ROS formation (Fig. 7). TBBPA increased ROS formation by about three times that of the control at 3µM. Experiments with bisphenol-A, a nonbrominated analog to TBBPA, did not elevate the ROS indicating a structure-activity relationship dependent on bromine substitution for ROS formation (Fig. 7). This is consistent with the previously reported effect of TBBPA and bisphenol-A on ROS formation in human neutrophil granulocytes (Reistad et al., 2005
).
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Inhibition of ROS Formation
Enzymatic inhibitors were used to elucidate involvement of different intracellular-signaling pathways in TBBPA-induced ROS formation. The SOD inhibitor DDC (Misra, 1979
) reduced the ROS formation by 83%. This indicates that at least some of the ROS formation is due to the formation of H2O2 by the enzyme SOD. The influence of ERK activation was investigated by treatment with the inhibitor of MEK activation, U0126 (Duncia et al., 1998; Favata et al., 1998), which reduced the fluorescent by 55% (Fig. 8). MEK lies immediately upstream of ERK in the signaling cascade and is responsible for its phosphorylation and activation. We also tested inhibitors of the p38 and Jun kinase branch of the MAP kinase pathway. The p38 inhibitor SB203580 (1µM) and the Jun kinase inhibitor FK506 (1µM) gave no significant decrease in free radical formation when the cells were exposed to 10µM TBBPA (data not shown). The response to TBBPA was strongly inhibited, 70% relative to the control (Fig. 8), by erbstatin-A, an inhibitor of tyrosine kinases (Kawada et al., 1993). Incubation of the cells in a calcium-free buffer containing 2mM EGTA showed a small but significant reduction in ROS formation (18%). The NMDA receptor blocker MK-801 and the antioxidant vitamin E induced no significant reduction in ROS formation (Fig. 8).
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Western Blotting Analysis
To confirm the ability of TBBPA to induce phosphorylation of ERK1/2, a phospho-specific antibody that recognizes phosphorylated Tyr204 of ERK1/2 was employed (Fig. 9). The results show that TBBPA induced phosphorylation of both ERK1 and -2 in a time-dependent manner (5–60 min) demonstrated by an increase in the relative intensities of the two immunodetectable bands, 44 and 42 kDa, respectively, compared with control. Phosphorylation of ERK1/2 was most pronounced after about 60 min (Fig. 9A). The results also show that TBBPA (5–25µM) induced a concentration-dependent increase in phosphorylation of ERK1/2 (Fig. 9B). Incubation with TBBPA (10µM) for 60 min in combination with U0126 (10µM) eliminated this response completely. In contrast, the tyrosine kinase inhibitor erbstatin-A and the SOD inhibitor DDC did not affect the phosphorylation of ERK. The NMDA receptor blocker MK-801 and the antioxidant vitamin E, which inhibit cell death but not ROS formation, did not influence the ERK phosphorylation. To compare the level of p-ERK with the total amount of ERK protein we reprobed the p-ERK filters with an anti-ERK2 antibody recognizing both ERK1 and ERK2. Only small variations in the amounts of protein were observed (lower panel, Figs. 9A and 9B).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The data presented here demonstrate that TBBPA is capable of inducing calcium influx, elevation of extracellular glutamate, ROS formation, and cell death in CGC in vitro. CGC were incubated with TBBPA for 24 h, which induced necrotic cell death with disruption of membrane integrity and failure to exclude trypan blue. The effect of TBBPA was comparable to what has been found for PCB. Mariussen et al. (2002)
estimated the LC50 value for the single PCB congener, PCB 153, to 8µM, while for TBBPA LC50 values were estimated to 7µM. The toxicity could result from the direct action of the compound or from toxic metabolites derived from the parent molecule. Earlier reports have demonstrated that the predominant metabolic pathways of TBBPA in vitro are cytochrome P450 dependent and result in different types of metabolites that may be reactive and able to bind macromolecules and trigger toxicological effects (Zalko et al., 2006). In this case, using hepatic S9 fraction in combination with TBBPA, cell death was reduced by 68%, indicating that TBBPA is the parent molecule responsible for the effect.
In cell cultures, the distinctions between apoptosis and necrosis can be confusing because of the lack of scavenging cells, and thus the phagocytic step after apoptosis may not occur. Instead, an apoptotic cell can undergo secondary necrosis and release its contents into the surrounding medium (Bonfoco et al., 1995). Several substances are known to induce apoptosis and necrosis depending on the dose. Therefore, we examined typical markers of apoptotic cell death in the CGC after exposure to lower concentrations of TBBPA. Apoptotic cells are characterized by condensed and fragmented nuclei, need for new synthesis of proteins and activation of caspases, whereas necrotic cells present loss of membrane integrity without apparent nuclear damage. TBBPA induced apoptotic morphology, characterized by chromatin condensation and fragmentation of nuclei (Fig. 3), whereas no caspase activity was detected. These results suggest that TBBPA at low concentration induces apoptotic cell death by a mechanism that is caspase independent. A caspase-independent mechanism of apoptotic cell death has been reported in various other models of neuronal cell death. Cell death induced by methylmercury in CGC (Castoldi et al., 2000), by serum deprivation in cortical neurons (Hamabe et al., 2000), by kainic acid in CGC (Verdaguer et al., 2002), and by H2O2 in PC12 cells (Jiang et al., 2001) has been reported to be independent of caspases. The lack of caspase activity may be explained by generation of ROS, which was reported to inhibit the activity of caspases (Baker et al., 2000; Hampton and Orrenius, 1997). Moreover, elevation of intracellular calcium can activate calpains, proteases that can cause inactivation of caspases (Lankiewicz et al., 2000).
Amino acid analysis of the growth media in TBBPA-exposed cells show that the levels of the excitatory amino acids glutamate and aspartate increase in a time-dependent manner (Fig. 6). The concentrations of glutamate reach a level that was previously shown to induce death of CGC (Eimerl and Schramm, 1991). TBBPA might induce release of neurotransmitters by a calcium-dependent mechanism or by an influence on transport and uptake mechanisms. In a study by Mariussen and Fonnum (2003), it was shown that TBBPA nonselectively inhibited uptake of glutamate, dopamine, and 
-amino-n-butyric acid in detached nerve terminals (synaptosomes).
MK-801 is known as a potent and selective noncompetitive NMDA receptor antagonist that acts at the NMDA receptor-operated ion channel as an open channel blocker (Wong et al., 1986). MK-801 protected against TBBPA-induced cell death in a 24-h exposure, indicating an involvement of the NMDA receptor. This is in accordance with the findings of elevated extracellular glutamate after TBBPA exposure, suggesting that TBBPA may lead to an increased stimulation of glutamate receptors. Stimulation of glutamate receptors is associated with excitotoxic injury as a consequence of elevated intracellular calcium (Choi, 1994).
The importance of calcium is reflected by the fact that cell death was reduced by 60% in calcium-free buffer. In addition, TBBPA provoked a concentration-dependent increase in intracellular-free calcium using Fura-2/AM (Fig. 5). Perturbations in intracellular calcium levels have been associated with cellular apoptosis. TBBPA has previously been shown to elevate intracellular calcium in human neutrophil granulocytes (Reistad et al., 2005). This is consistent with the view that disturbance of the calcium homeostasis is associated with the toxicity of TBBPA on CGC. It was surprising that calcium elevations were small for TBBPA concentrations as high as 10µM. Nevertheless, it should be noted that these are acute elevations of calcium and it is likely that calcium changes accumulates over 24 h. The TBBPA dose-dependent acute elevation of calcium therefore supports the suggestion that calcium dysregulation is involved in TBBPA toxicity. Disturbance of the calcium homeostasis and activation of glutamate receptors may lead to oxidative stress.
Reactive oxygen and nitrogen species have been implicated as an important causative factor in the mechanism of action of several environmental contaminants (Ali et al., 1992; Dreiem et al., 2005; Mariussen et al., 2002; Myhre et al., 2004). Vitamin E is a lipid-soluble antioxidant known to efficiently scavenge peroxyl radicals and is probably the most important inhibitor of the free radical chain reaction of lipid peroxidation in animals (Halliwell and Gutteridge, 1999). Vitamin E reduced cell death induced by TBBPA by 46% and this suggests that formation of lipid peroxyl radicals could be involved in the cytotoxicity after TBBPA exposure. TBBPA induced ROS formation in a concentration-dependent manner as measured with DCF fluorescence. Oxidation of DCFH was reduced by DDC, a potent inhibitor of the copper-zinc SOD (Heikkila et al., 1976; Misra, 1979), indicating H2O2 formation. Vitamin E did not, however, reduce the ROS formation induced by TBBPA. This might be due to the fact that the vitamin E effect is mediated in the membranes and not in the cytosol. DCFH primarily measures ROS in the cytosolic compartment, suggesting that the protective effect of vitamin E may be due to its property as a membrane-stabilizing compound as previously described (Urano et al., 1992).
MK-801 had no significant effect on ROS formation. This was surprising since blocking the NMDA receptor almost completely inhibited cell death, and it is known that glutamate receptor activation may lead to oxidative stress. Previously we have shown that MK-801 inhibited cell death as well as ROS formation in A1254-exposed CGC (Mariussen et al., 2002). The present findings indicate that TBBPA induces ROS formation independently of NMDA receptor activation. Calcium-free media, on the other hand, reduced cell death and inhibited ROS formation, strongly suggesting that disruption of the calcium homeostasis is involved in the observed effects on TBBPA-exposed cells.
Stimulation of glutamate receptors and influx of calcium are associated with exitotoxic injury (Choi, 1994) and lead to phosphorylation of ERK1/2 in neurons (Rosen et al., 1994; Xia et al., 1996). We have shown that TBBPA induces ERK1/2 activation in human granulocytes in vitro (Reistad et al., 2005). Canesi et al., (2005) subsequently reported similar effects on mussel hemocytes demonstrating TBBPA activation of ERK in different cell types. In agreement with these earlier reports, TBBPA-exposed CGC showed both time and concentration-dependent induction of ERK-phosphorylation. Phosphorylation of ERK, shown by Western blotting, was fully inhibited by pretreatment with U0126, which inhibits the ERK branch of the MAP-kinase pathway (Duncia et al., 1998; Favata et al., 1998). This indicates that TBBPA does not act directly on the kinase but must have a target upstream in the signaling cascade. ERK1/2 can be activated by various stimuli, such as oxidative stress (Aikawa et al., 1997), calcium influx (Rosen et al., 1994), stimulation of glutamate receptors (Sgambato et al., 1998) and growth factors (Boulton et al., 1991). The link between ERK activation and ROS formation is unclear. It was found that ERK activates NADPH oxidase and PLA2 in neurons (Myhre et al., 2004), but we did not find evidence for similar mechanism here. The role of ERK1/2 in neuronal cell death remains controversial. ERK activation is typically associated with neuronal survival (Jacobs et al., 2004; Xia et al., 1995). Nevertheless, activation of ERK has also been found to contribute to neuronal cell death in some in vitro models of neurotoxicity (Creedon et al., 1996; Murray et al., 1998; Runden et al., 1998).
This study has shown that TBBPA induces cell death, ROS formation, calcium influx, and elevation of extracellular glutamate in CGC in vitro. The results also show that different mechanisms are involved in formation of ROS and cell death, although disruption of calcium homeostasis seems to be involved in both. TBBPA is found in human plasma at 0.3–1.8 ng/g lw which corresponds to low nanomolar concentrations (Thomsen et al., 2001), which is considerably lower than that used in this investigation. TBBPA is previously shown to have a low bioaccumulation potential, which probably is due to its phenolic structure making it more attributed to metabolic breakdown. TBBPA has, however, some interesting mechanistic effects, as shown in vitro in this investigation and in previous studies. This makes the compound very interesting in a toxicological point of view, especially with regard to combination effects with other toxicants, and the toxicity and bioavailability of metabolic breakdown products. The fact that TBBPA is used in such large amount should therefore merit further investigations.
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ACKNOWLEDGMENTS |
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The authors wish to thank Dr Bjørnar Hassel for technical assistance with the HPLC experiments and Dr Yngvar Gundersen for proofreading the manuscript. This work was supported by grants from the Norwegian Research Council.
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