ToxSci Advance Access originally published online on February 16, 2006
Toxicological Sciences 2006 91(1):184-191; doi:10.1093/toxsci/kfj137
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Zinc Deficiency Increases the Susceptibility of Human Neuroblastoma Cells to Lead-Induced Activator Protein-1 Activation
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* Department of Nutrition, University of California, Davis, Davis, California 95616; and Department of Environmental Toxicology, University of California, Davis, Davis, California 95616
1 To whom correspondence should be addressed at Departments of Nutrition, University of California, Davis, One Shields Avenue, Davis, CA 95616. Fax: (530) 752-8966. E-mail: poteiza{at}ucdavis.edu.
Received January 10, 2006; accepted February 11, 2006
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Lead (Pb2+) is a major environmental pollutant that has severe adverse effects on the nervous system. Similar human populations are at risk of suffering both Pb2+ toxicity and zinc (Zn) deficiency. Thus, in the present study we investigated whether Zn deficiency can increase the susceptibility of human neuroblastoma IMR-32 cells to Pb2+-induced oxidative stress which could trigger the activation of the mitogen-activated protein kinases (MAPKs) c-Jun-N-terminal kinase (JNK) and p38 and subsequently activate transcription factor activator protein-1 (AP-1). After 24 h of incubation, 5–50µM Pb2+ caused a decrease in cell viability that was markedly higher in the Zn-deficient cells compared to controls. Pb caused a time (2–24 h) and dose (5–50µM)–dependent increase of cell oxidants, with a significantly higher effect in the Zn-deficient cells. Pb2+ treatment triggered the activation of JNK and p38, measured as the phosphorylation of JNK and p38, only in cells incubated in the Zn-deficient media. The exposure to Pb2+ (2–24 h) led to a higher AP-1 DNA-binding activity and AP-1–dependent gene transactivation, only in the Zn-deficient cells. Results show that Zn deficiency can increase the cytotoxicity of Pb2+ and the susceptibility of neurons to Pb2+-induced oxidative stress, leading to JNK and p38 phosphorylation and, subsequently, AP-1 activation.
Key Words: lead; zinc; AP-1; oxidative stress; MAPK.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The susceptibility to lead (Pb2+) toxicity is known to be influenced by several factors such as age, nutritional status, and environmental exposure. Pb2+ causes many adverse health effects, including toxicity of the hematopoietic, renal, and skeletal systems, with the central nervous system as the primary target organ (Peraza et al., 1998
; Pounds and Cory-Slechta, 1993). Pb2+ is recognized as a major global environmental health hazard, affecting sensitive populations including pregnant women (Hertz-Picciotto et al., 2000), young children (Muller and Krawinkel, 2005; Needham et al., 2005; Plunkett, 1999), and the elderly (Peraza et al., 1998). Similar populations are also at significant risk for poor zinc (Zn) nutritional status (Scheplyagina, 2005), and a condition of Zn deficiency could turn an individual more susceptible to Pb2+ neurotoxicity.
Several studies showed that Zn deficiency can increase the susceptibility to the toxicity of metals such as cadmium and iron (Brzoska and Moniuszko-Jakoniuk, 2001; Peraza et al., 1998; Sreedhar et al., 2004). Zn deficiency–induced oxidative stress could be one factor facilitating the deleterious action of pro-oxidant metals. In this regard, Zn-deficient rats were particularly sensitive to cadmium-induced oxidative damage to the testes (Oteiza et al., 1999). Human neuroblastoma IMR-32 cells exposed to Zn deficiency showed increased susceptibility to iron-induced oxidative stress and apoptotic cell death (Mackenzie et al., 2002).
Some of the mechanisms proposed to contribute to Pb2+ toxicity are the induction of oxidative stress (Adonaylo and Oteiza, 1999; Chen et al., 2002; Gurer-Orhan et al., 2004; Hermes-Lima et al., 1991; Monteiro et al., 1985) and the triggering of oxidant-sensitive transcription factors, such as activator protein-1 (AP-1) (Chakraborti et al., 1999; Hossain et al., 2000; Kim et al., 2000; Ramesh et al., 2001). AP-1 is composed of dimeric basic region leucine zipper proteins that belong to the Jun, Fos, Maf, and ATF subfamilies. AP-1 is activated by both physiological stimuli and stress conditions. The structural and regulatory complexities of this transcription factor underlie its capacity to regulate multiple and diverse cellular functions (Karin, 1995; Karin et al., 1997; Shaulian and Karin, 2002). AP-1 regulates genes involved in critical cell processes including cell proliferation, differentiation, and survival (Shaulian and Karin, 2002). Mitogen-activated protein kinases (MAPKs) belong to a family of Ser/Thr protein kinases that transmit extracellular signals into the nucleus. The MAPKs c-Jun-N-terminal kinase (JNK) and p38 are sensitive to oxidative stress (Karin and Shaulian, 2001; Karin et al., 2001; Kim and Sharma, 2004) and can activate AP-1 (Toone et al., 2001). Accordingly, JNK and AP-1 activities were higher in the brain from rats exposed to Pb2+ (Ramesh et al., 2001).
A condition of Zn deficiency can induce cellular oxidative stress, promoting an increased production of oxygen and nitrogen species (Ho and Ames, 2002; Mackenzie et al., 2006) and triggering the activation of AP-1 (Oteiza et al., 2000). In human neuroblastoma cells, Zn deficiency caused an increased production of H2O2 that led to JNK and p38 activation and subsequently to an increased transactivation of AP-1–regulated genes (Zago et al., 2005).
Previous findings of a pro-oxidant action of Pb2+ combined with the observation that Zn deficiency can induce cell oxidative stress and AP-1 activation lead us to hypothesize that a condition of Zn deficiency would potentiate the deleterious effects of Pb2+ on the nervous system. Therefore, the influence of neuronal Zn status on cell susceptibility to Pb2+-induced oxidative stress and MAPK and AP-1 activation was investigated.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Materials.
IMR-32 cells were obtained from the American Type Culture Collection (Rockville, MA). Cell culture media and reagents, and LipofectAMINE 2000, were obtained from Invitrogen Life Technologies (Carlsbad, CA). The oligonucleotide containing the consensus sequences for AP-1 (5'-CGC TTG ATG AGT CAG CCG GAA-3') and octamer binding transcription factor-1 (OCT-1), the reagents for the electrophoretic mobility shift assay (EMSA), the CellTiter 96 nonradioactive cell proliferation assay (MTT), and the luciferase assay system were obtained from Promega (Madison, WI). The PathDetect AP-1 cis-reporting system was obtained from Stratagene (La Jolla, CA). Phosphospecific polyclonal antibodies for JNK and p38 and antibodies for the nonphosphorylated form of JNK were obtained from Cell Signaling Technology (Beverly, MA). Antibodies for the nonphosphorylated p38, c-Jun, JunB, c-Fos, and ß-tubulin were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). PVDF membranes were obtained from BIO-RAD (Hercules, CA), and Chroma Spin-10 columns were obtained from Clontech (Palo Alto, CA). 5-(and-6)-carboxy-2'7'-dichlorodihydrofluorescein diacetate (DCF) and propidium iodide (PI) were obtained from Molecular Probes (Eugene, OR). The ECL Western blotting system was obtained from Amersham Pharmacia Biotech Inc. (Piscataway, NJ). Lead acetate and all other reagents were of the highest quality available and were purchased from Sigma (St. Louis, MO).
Cell culture and incubations.
IMR-32 cells were cultured at 37°C in complex medium (55% [vol/vol] Dulbecco's modified Eagle medium [DMEM] high glucose, 30% [vol/vol] Ham F-12, 5% [vol/vol] 
-MEM) supplemented with 10% (vol/vol) fetal bovine serum (FBS) and an antibiotic-antimycotic (50 U/ml penicillin, 50 µg/ml streptomycin, and 0.125 µg/ml amphotericin B).
Zn-deficient FBS was prepared as previously described (Oteiza et al., 2000). The Zn-deficient FBS was subsequently diluted with complex medium to a final concentration of 3 mg protein/ml to match the protein concentration of the control nondeficient media. The Zn concentration of the deficient medium was 1.5µM (1.5 Zn). An aliquot of the Zn-deficient media was supplemented with ZnCl2 to obtain a final concentration of 15µM (15 Zn).
Cells were grown in control medium (complex medium containing 10% [vol/vol] FBS) until 90% confluence, after which the media were removed and replaced with control medium or media containing 1.5µM Zn or 15µM Zn with or without the addition of (5–50µM) lead acetate. The lead acetate solution was freshly prepared in sterile water before each experiment. Cells were harvested at 2, 12, or 24 h.
Cell viability was measured using the MTT assay according to the manufacturer's protocol (Promega).
Evaluation of the global concentration of intracellular oxidants.
The global concentration of intracellular oxidants was estimated using the probe DCF (Molecular Probes). DCF is a nonfluorescent probe which is oxidized to a fluorescent derivative by endogenous oxidants. The fluorescence intensity is considered to be a parameter of global intracellular oxidant levels. Cells were incubated in control, 1.5 Zn, or 15 Zn media in the absence or presence of 5–50µM Pb2+. After 2, 12, or 24 h of treatment, the media were changed to DMEM high glucose containing DCF at a final concentration of 10µM. Cells were incubated in the dark at 37°C for 30 min. The media were then removed, and the cells were rinsed with phosphate-buffered saline (PBS), pH 7.4. Cells were scraped and resuspended in 0.5 ml of PBS, pH 7.4, containing 0.1% (vol/vol) Igepal. After brief sonication, fluorescence at 525 nm (
exc = 475 nm) was measured. To evaluate the DNA content, samples were incubated with 50µM PI. After incubation for 20 min at room temperature, fluorescence at 590 nm (exc = 538 nm) was measured. Results are expressed as a ratio of DCF to PI relative fluorescence.
Electrophoretic mobility shift assay.
Nuclear fractions were isolated as previously described (Dignam et al., 1983; Osborn et al., 1989). At the corresponding time points, the media were discarded; cells were rinsed with PBS and scraped. After centrifugation at 800 x g for 10 min, the pellet was resuspended in 150 µl of buffer A (10mM Hepes buffer, pH 7.9, containing 1.5mM MgCl2, 10mM KCl, 0.5mM dithiotreitol [DTT], and 0.1% [vol/vol] Igepal), incubated for 10 min, and centrifuged for 1 min at 12,000 x g at 4°C. The supernatant fraction was removed (cytosolic fraction), and the nuclear pellet was resuspended in 75 µl of buffer B (10mM Hepes buffer, pH 7.9, containing 1.5mM MgCl2, 420mM NaCl, 0.5mM DTT, 0.2mM ethylenediaminetetraacetic acid (EDTA), 25% [vol/vol] glycerol and 0.5mM PMSF, 5 µg/ml leupeptin, 1 mg/l pepstatin, and 10 µg/ml aprotinin). Samples were incubated for 20 min at 4°C and then centrifuged at 10,000 x g for 10 min at 4°C. The supernatant was transferred to a new tube and diluted with buffer C (20mM Hepes buffer, pH 7.9, containing 50mM KCl, 0.5mM DTT, 0.2mM EDTA, and 0.5mM PMSF). Samples were stored at –80°C, and protein concentration was determined (Bradford, 1976) immediately before starting the assay.
For the EMSA, the oligonucleotide containing the consensus sequence for AP-1 was end labeled with 
-32P-ATP using T4 polynucleotide kinase and purified using Chroma Spin-10 columns. Samples were incubated with the labeled oligonucleotide (20,000–30,000 cpm) for 20 min at room temperature in binding buffer [5 x binding buffer: 50mM Tris-HCl buffer, pH 7.5, containing 20% (vol/vol) glycerol, 5mM MgCl2, 2.5mM EDTA, 2.5mM DTT, 250mM NaCl, and 0.25 mg/ml poly(dI-dC)]. For the supershift assays, prior to the addition of the labeled nucleotide, samples were incubated in the presence of the corresponding antibodies (c-Jun, c-Fos, and JunB). The products were then separated by electrophoresis on a 4% (wt/vol) nondenaturing polyacrylamide gel using 0.5x TBE (45mM Tris/borate, 1mM EDTA) as the running buffer. The gels were dried, and the radioactivity was quantitated in a Phosphoimager 640 (Amersham Pharmacia Biotech Inc.).
Transfections.
Cells were seeded at 2.5 x 106 cells per well in six-well plates. After 24 h in culture, cells were transfected with the pAP-1-Luc plasmid (1 µg DNA) using LipofectAMINE 2000 according to the manufacturer's protocols (Invitrogen Life Technologies). After 6 h of initiation of the transfection, cells were treated with the control, 1.5 Zn, or 15 Zn media and exposed to 0–50µM Pb2+. Cells were harvested and lysed after 12 and 24 h; their protein content and luciferase activity were determined following the manufacturer's protocols (Promega).
Western blot.
To evaluate the phosphorylation of JNK and p38, we used antibodies recognizing the phosphorylated/activated forms of JNK1/2 (Thr183/Tyr185) and p38 (Thr180/Tyr182). For these experiments, cell lysates were prepared in sodium dodecyl sulfate (SDS)-sample buffer (62.5mM Tris-HCl buffer, pH 6.8, containing 2% [wt/vol] SDS, 10% [vol/vol] glycerol, 50mM DTT, and 0.01% [wt/vol] bromophenol blue), sonicated for 10 s to shear DNA and reduce sample viscosity, and then heated at 95°C for 5 min. Twenty-five microliters of the lysate was loaded onto sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE). Proteins were resolved by SDS-PAGE and transferred to PVDF membranes. Molecular weight standards (Cell Signaling Technology) were run simultaneously. Membranes were immunoblotted with the corresponding primary antibody overnight at 4°C and the following day for 60 min at room temperature in the presence of the secondary antibody (HRP conjugated). The conjugates were detected by enhanced chemiluminescence in a Phosphoimager 640. The membranes were normalized by reblotting with the antibody for the corresponding nonphosphorylated form of each protein. Equal loading of cell lysate was controlled by ß-tubulin content.
Statistical analysis.
One-way ANOVA test, followed by Fisher's protected least-squares difference test and correlations, was performed using the routines available in Statview 5.0 (SAS Institute, Cary, NC). A p value < 0.05 was considered statistically significant. Values are given as means ± SEM.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Pb2+ Differentially Affects Cell Viability in Zn-Deficient Cells
After 24 h, only the Zn-deficient cells exposed to 50µM Pb2+ showed evident morphological alterations including rounded cell body with a decreased number of cell projections (Fig. 1). Cell viability was evaluated with the MTT assay after 2, 12, and 24 h of incubation of the cells in control, 1.5 Zn or 15 Zn media in the absence or presence of 5–50µM Pb2+ (Fig. 2). After 2 h, cell viability was similar among the different groups (Fig. 2A). Cell viability was significantly lower (p < 0.05) in the Zn-deficient cells after 12 h of exposure to 5–50µM Pb2+ (Fig. 2B). After 24 h of incubation, cell viability was significantly lower (p < 0.02) in the control group treated with 50µM Pb2+ (29%). In the zinc-deficient (1.5 Zn) cells, Pb2+ treatment caused a marked decrease (25–50%) in cell viability in all the concentration ranges tested (Fig. 2C).
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Pb2+ Increases Zn Deficiency–Induced Oxidative Stress
Incubation of IMR-32 cells for 2, 12, or 24 h in the Zn-deficient media caused a significant increase (p < 0.005) in global cell oxidants compared to control and 15 Zn cells (Fig. 3). When exposed to Pb2+ during 2 h, no significant differences were observed in any of the groups evaluated (Fig. 3A). In the Zn-deficient cells, a significant increase in DCF relative fluorescence (corrected by the DNA content) was observed after 12 h of exposure to 10 and 50µM Pb2+ (Fig. 3B), while no significant differences were observed in the control and 15 Zn cells. After 24 h of treatment with Pb2+, all groups (control, 1.5 Zn, and 15 Zn) showed a dose-dependent increase in DCF relative fluorescence (Fig. 3C). However, at 50µM Pb2+, a significantly higher increase in total oxidants was observed in the 1.5 Zn cells (63%) compared to control and 15 Zn cells (42 and 25%, respectively).
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Pb2+ Activates MAPKs in Zn-Deficient Cells
The MAPKs p38 and JNK are mainly stress responsive and are known to be activated by H2O2. To investigate the possible activation of these MAPKs in the Zn-deficient cells exposed to Pb2+, the levels of phosphorylation of p38 and JNK were measured by Western blotting (Figs. 4 and 5). Pb2+ treatment induced a significant increase in p38 phosphorylation in 1.5 Zn cells (14 and 25% at 10 and 50µM Pb2+, respectively). While Pb2+ treatment had no effect in the control group, a 12% increase in p38 phosphorylation was observed at 50µM Pb2+ in the 15 Zn cells. After 24 h of incubation in the presence of 50µM Pb2+, JNK phosphorylation increased significantly in the 1.5 Zn cells (30%) compared to control and 15 Zn cells (7 and 20%, respectively) (Fig. 4).
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Pb2+ Activates AP-1 in Zn-Deficient Cells
JNK and p38 are involved in the downstream activation of transcription factor AP-1. The activation of AP-1 was evaluated by measuring the AP-1 DNA-binding activity in nuclear cell fractions by EMSA and the transactivation of an AP-1–driven reporter gene (pAP-1-Luc).
For the EMSAs, the specificity of the AP-1–DNA complex was assessed by competition with a 100-fold molar excess of unlabeled oligonucleotide containing the consensus sequence for either AP-1 or OCT-1 (Fig. 6A). The major components of AP-1 were determined by supershift assays with specific antibodies corresponding to the Jun (c-Jun and JunB) and Fos (c-Fos) families of proteins. As previously observed (Zago et al., 2005), the main AP-1 protein present in IMR-32 cells is c-Jun, with a minor contribution of JunB (Fig. 6A). The AP-1 DNA-binding activity was measured in control, 1.5 Zn, and 15 Zn cells, after 2, 12, and 24 h of exposure to 0–50µM Pb2+. When measured after a 2-h treatment (Fig. 6B), AP-1 DNA-binding activity was significantly higher only in the Zn-deficient cells exposed to 10 and 50µM Pb2+ (16 and 28 %, respectively). Similarly, after 12 and 24 h of incubation with Pb2+ (Figs. 6C and 6D), AP-1 DNA-binding activity was significantly higher (p < 0.05) only in the Zn-deficient cells exposed to Pb2+.
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A reporter gene assay (luciferase) was conducted to evaluate AP-1–driven gene transactivation. Cells were cotransfected with a vector expressing ß-galactosidase (as a control for the transfection efficiency) and a pAP-1-Luc plasmid. Since the transfection efficiency was similar among all groups and the simultaneous transfection of both plasmids resulted in cell toxicity, cells were only transfected with the pAP-1-Luc plasmid. Results were expressed in terms of the protein content of the samples. After 12 h of incubation, Pb2+ treatment caused a significantly higher luciferase activity (p < 0.05) in 1.5 Zn cells compared to control and 15 Zn cells (Fig. 7A). After 24 h of treatment, luciferase activity measured in 1.5 Zn cells exposed to 50µM Pb2+ remained significantly higher (p < 0.05) (Fig. 7B).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The populations at risk of suffering both Zn deficiency and Pb2+ toxicity are strikingly similar. These populations are not only restricted to developing countries but also affect select groups, such as children and the elderly, in urban areas of developed countries. Pb2+ causes long-term adverse effects to the nervous system affecting both learning and behavior (White et al., 1993
). One mechanism potentially involved in Pb2+ neurotoxicity is the capacity of the metal to induce oxidative stress, thereby triggering oxidant-responsive transcription factors, such as AP-1. Given the findings that Zn deficiency also affects oxidant-driven cell signals (AP-1, NF-
B) (Mackenzie et al., 2006) critical to neuronal survival and plasticity, it is reasonable to suggest that a combination of marginal Zn deficiency and Pb2+ exposure during infancy could have a serious impact on neurobehavioral and learning capacities later in life. This study investigated the hypothesis that a condition of Zn deficiency could increase the susceptibility of neuroblastoma cells to Pb2+-induced oxidative stress and AP-1 activation, affecting cell survival.
In human neuroblastoma IMR-32 cells, Pb2+ affected cell viability in both Zn-deficient and Zn-adequate cells. However, cells grown in the Zn-deficient media were particularly affected by Pb2+. Accordingly, Zn-deficient cells exposed to Pb2+ showed morphological alterations including rounded cell body with a decreased number of cell projections.
Evidence of oxidative stress has been consistently found secondary to high Pb2+ exposure in humans and in different experimental conditions. Chronic exposure of rats to high Pb2+ levels leads to the accumulation of Pb2+ in the brain in association with high levels of lipid oxidation products and changes in components of the oxidant defense system (Adonaylo and Oteiza, 1999; Villeda-Hernandez et al., 2001). Different mechanisms, including direct or indirect effects, could underlie the pro-oxidant action of Pb2+ (Loikkanen et al., 1998; Naarala et al., 1995). Pb2+ could promote oxidative stress through the activation of protein kinase C (PKC) or by binding thiol groups, which can disturb the cell thiol redox status. As previously observed (Oteiza et al., 2000), Zn deficiency alone caused an increase in the cellular levels of oxidants. The actual reasons why Zn deficiency can induce oxidative stress are still unknown. It could be either due to its role in oxidant defense systems (Zago and Oteiza, 2001) or due to secondary changes in cell physiology (mitochondrial alterations, changes in enzyme activities that generate oxidant species) that occur as a consequence of such deficiency. We currently observed that Pb2+ also induced a dose-dependent increase in cell oxidants in IMR-32 cells. This effect was significantly higher in the Zn-deficient cells, indicating that a condition of Zn deficiency can increase the susceptibility of neurons to Pb2+-induced oxidative stress. Whether Zn deficiency and Pb2+ toxicity elicit oxidative stress through the same molecular mechanism is still unknown.
JNK has been described as being activated by Pb2+ in rat pheochromocytoma PC12 cells (Ramesh et al., 1999). JNK and the upstream MAPK kinase are also markedly activated in the frontal cortex, brain stem, striatum, and hippocampus of rats chronically exposed to Pb2+ (Ramesh et al., 2001). H2O2 is one major signal in the activation of MAPKs JNK and p38. It is important to consider that H2O2 can also cause inhibition of MAPK phosphatases by oxidation of their catalytic cysteine, which results in sustained activation of p38 and JNK MAPKs (Kamata et al., 2005). It has been recently demonstrated that H2O2 is involved in the activation of both MAPKs p38 and JNK in Zn-deficient IMR-32 cells (Zago et al., 2005). In the present experimental conditions, JNK and p38 were activated by Pb2+ exposure in the Zn-deficient cells. This is in agreement with the rapid increase in cell oxidants observed in the same group.
Both oxidant-responsive MAPKs, p38 and JNK, can lead to the activation of transcription factor AP-1. This transcription factor has been found to be activated by both Pb2+ exposure and Zn deficiency. We previously described that Zn deficiency is associated with a high AP-1 DNA-binding activity in 3T3 and IMR-32 cells and with an increase in the AP-1–dependent transactivation of endogenous and reporter genes (Oteiza et al., 2000; Zago et al., 2005). Pb2+ also activates AP-1 in cell and animal models (Chakraborti et al., 1999; Hossain et al., 2000; Ramesh et al., 2001; Scortegagna and Hanbauer, 2000) partially through the activation of PKC). In the present study, Pb2+ activated AP-1 in the Zn-deficient cells, as evidenced by both high AP-1 DNA-binding activity in nuclear fractions and an increased transactivation of an AP-1–driven gene. This indicates that Zn-deficient neuroblastoma cells are particularly susceptible to Pb2+-triggered MAPK and AP-1 activation.
The present results suggest a possible role for oxidants in Pb2+-induced activation of AP-1 in Zn-deficient cells. Besides oxidants, other mechanisms could be involved in Pb2+-induced AP-1 activation. For instance, it has been shown that PKC is involved in AP-1 activation (Chakraborti et al., 1999; Kim et al., 2000). One of the major factors in the activation of PKC is calcium (Nishizuka, 1986). It still remains unclear whether Pb2+ exposure modifies calcium concentration, but one possibility is that Pb2+ activates PKC most likely by mimicking calcium (Long et al., 1994; Markovac and Goldstein, 1988).
The concentrations used in this study are severalfold higher than the blood levels found in affected individuals. In cell culture, the exposure of cells to Pb2+ depends on both the time of exposure and the free concentration of Pb2+, which, in turn, is determined by the various chelators and competing ions in the media. Thus, although the in vitro models help us predict a possible mechanism to explain Pb2+ toxicity in humans, its physiological relevance needs to be explored in other models.
In summary, a condition of Zn deficiency facilitated the action of Pb2+ promoting oxidative stress, MAPK and AP-1 activation, and decreased neuronal cell viability. The effects of this activation on cell function and/or decisions to proliferate, differentiate, or undergo apoptosis need further investigation. We propose that populations at risk of Zn deficiency could be more susceptible to the long-term deleterious effects of Pb2+ on the nervous system.
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ACKNOWLEDGMENTS |
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This study was supported by grants from the National Institute of Environmental Health Services Center for Environmental Health Sciences, University of California, Davis, and from the University of California, Davis.
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REFERENCES |
|---|
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Adonaylo, V. N., and Oteiza, P. I. (1999). Lead intoxication: Antioxidant defenses and oxidative damage in rat brain. Toxicology 135, 77–85.[CrossRef][ISI][Medline]
Bradford, M. M. (1976). A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72, 248–254.[CrossRef][ISI][Medline]
Brzoska, M. M., and Moniuszko-Jakoniuk, J. (2001). Interactions between cadmium and zinc in the organism. Food Chem. Toxicol. 39, 967–980.[CrossRef][ISI][Medline]
Chakraborti, T., Kim, K. A., Goldstein, G. G., and Bressler, J. P. (1999). Increased AP-1 DNA binding activity in PC12 cells treated with lead. J. Neurochem. 73, 187–194.[CrossRef][ISI][Medline]
Chen, L., Yang, X., Jiao, H., and Zhao, B. (2002). Tea catechins protect against lead-induced cytotoxicity, lipid peroxidation, and membrane fluidity in HepG2 cells. Toxicol. Sci. 69, 149–156.
Dignam, J. D., Lebovitz, R. M., and Roeder, R. G. (1983). Accurate transcription initiation by RNA polymerase II in a soluble extract from isolated mammalian nuclei. Nucleic Acids Res. 11, 1475–1489.
Gurer-Orhan, H., Sabir, H. U., and Ozgunes, H. (2004). Correlation between clinical indicators of lead poisoning and oxidative stress parameters in controls and lead-exposed workers. Toxicology 195, 147–154.[CrossRef][ISI][Medline]
Hermes-Lima, M., Pereira, B., and Bechara, E. J. (1991). Are free radicals involved in lead poisoning? Xenobiotica 21, 1085–1090.[ISI][Medline]
Hertz-Picciotto, I., Schramm, M., Watt-Morse, M., Chantala, K., Anderson, J., and Osterloh, J. (2000). Patterns and determinants of blood lead during pregnancy. Am. J. Epidemiol. 152, 829–837.
Ho, E., and Ames, B. N. (2002). Low intracellular zinc induces oxidative DNA damage, disrupts p53, NFkappa B, and AP1 DNA binding, and affects DNA repair in a rat glioma cell line. Proc. Natl. Acad. Sci. U.S.A. 99, 16770–16775.
Hossain, M. A., Bouton, C. M., Pevsner, J., and Laterra, J. (2000). Induction of vascular endothelial growth factor in human astrocytes by lead. Involvement of a protein kinase C/activator protein-1 complex-dependent and hypoxia-inducible factor 1-independent signaling pathway. J. Biol. Chem. 275, 27874–27882.
Kamata, H., Honda, S., Maeda, S., Chang, L., Hirata, H., and Karin, M. (2005). Reactive oxygen species promote TNFalpha-induced death and sustained JNK activation by inhibiting MAP kinase phosphatases. Cell 120, 649–661.[CrossRef][ISI][Medline]
Karin, M. (1995). The regulation of AP-1 activity by mitogen-activated protein kinases. J. Biol. Chem. 270, 16483–16486.
Karin, M., Liu, Z., and Zandi, E. (1997). AP-1 function and regulation. Curr. Opin. Cell Biol. 9, 240–246.[CrossRef][ISI][Medline]
Karin, M., and Shaulian, E. (2001). AP-1: Linking hydrogen peroxide and oxidative stress to the control of cell proliferation and death. IUBMB Life 52, 17–24.[ISI][Medline]
Karin, M., Takahashi, T., Kapahi, P., Delhase, M., Chen, Y., Makris, C., Rothwarf, D., Baud, V., Natoli, G., Guido, F., et al. (2001). Oxidative stress and gene expression: The AP-1 and NF-kappaB connections. Biofactors 15, 87–89.[ISI][Medline]
Kim, J., and Sharma, R. P. (2004). Calcium-mediated activation of c-Jun NH2-terminal kinase (JNK) and apoptosis in response to cadmium in murine macrophages. Toxicol. Sci. 81, 518–527.
Kim, K. A., Chakraborti, T., Goldstein, G. W., and Bressler, J. P. (2000). Immediate early gene expression in PC12 cells exposed to lead: Requirement for protein kinase C. J. Neurochem. 74, 1140–1146.[CrossRef][ISI][Medline]
Loikkanen, J. J., Naarala, J., and Savolainen, K. M. (1998). Modification of glutamate-induced oxidative stress by lead: The role of extracellular calcium. Free Radic. Biol. Med. 24, 377–384.[CrossRef][ISI][Medline]
Long, G. J., Rosen, J. F., and Schanne, F. A. (1994). Lead activation of protein kinase C from rat brain. Determination of free calcium, lead, and zinc by 19F NMR. J. Biol. Chem. 269, 834–837.
Mackenzie, G. G., Keen, C. L., and Oteiza, P. I. (2002). Zinc status of human IMR-32 neuroblastoma cells influences their susceptibility to iron-induced oxidative stress. Dev. Neurosci. 24, 125–133.[CrossRef][ISI][Medline]
Mackenzie, G. G., Zago, M. P., Erlejman, A. G., Aimo, L., Keen, C. L., and Oteiza, P. I. (2006). Alpha-lipoic acid and N-acetyl cysteine prevent zinc deficiency-induced activation of NF-kappaB and AP-1 transcription factors in human neuroblastoma IMR-32 cells. Free Radic. Res. 40, 75–84.[CrossRef][ISI][Medline]
Markovac, J., and Goldstein, G. W. (1988). Picomolar concentrations of lead stimulate brain protein kinase C. Nature 334, 71–73.[CrossRef][Medline]
Monteiro, H. P., Abdalla, D. S., Arcuri, A. S., and Bechara, E. J. (1985). Oxygen toxicity related to exposure to lead. Clin. Chem. 31, 1673–1676.[Abstract]
Muller, O., and Krawinkel, M. (2005). Malnutrition and health in developing countries. CMAJ 173, 279–286.
Naarala, J. T., Loikkanen, J. J., Ruotsalainen, M. H., and Savolainen, K. M. (1995). Lead amplifies glutamate-induced oxidative stress. Free Radic. Biol. Med. 19, 689–693.[CrossRef][ISI][Medline]
Needham, L. L., Barr, D. B., Caudill, S. P., Pirkle, J. L., Turner, W. E., Osterloh, J., Jones, R. L., and Sampson, E. J. (2005). Concentrations of environmental chemicals associated with neurodevelopmental effects in U.S. population. Neurotoxicology 26, 531–545.[CrossRef][ISI][Medline]
Nishizuka, Y. (1986). Studies and perspectives of protein kinase C. Science 233, 305–312.
Osborn, L., Kunkel, S., and Nabel, G. J. (1989). Tumor necrosis factor alpha and interleukin 1 stimulate the human immunodeficiency virus enhancer by activation of the nuclear factor kappa B. Proc. Natl. Acad. Sci. U.S.A. 86, 2336–2340.
Oteiza, P. I., Adonaylo, V. N., and Keen, C. L. (1999). Cadmium-induced testes oxidative damage in rats can be influenced by dietary zinc intake. Toxicology 137, 13–22.[CrossRef][ISI][Medline]
Oteiza, P. I., Clegg, M. S., Zago, M. P., and Keen, C. L. (2000). Zinc deficiency induces oxidative stress and AP-1 activation in 3T3 cells. Free Radic. Biol. Med. 28, 1091–1099.[CrossRef][ISI][Medline]
Peraza, M. A., Ayala-Fierro, F., Barber, D. S., Casarez, E., and Rael, L. T. (1998). Effects of micronutrients on metal toxicity. Environ. Health Perspect. 106(Suppl. 1), 203–216.
Plunkett, L. M. (1999). Do current FIFRA testing guidelines protect infants and children? Lead as a case study. Federal Insecticide, Fungicide, and Rodenticide Act. Regul. Toxicol. Pharmacol. 29, 80–87.[CrossRef][ISI][Medline]
Pounds, J. G., and Cory-Slechta, D. A. (1993). New dimensions of lead neurotoxicity: Redefining mechanisms and effects. Neurotoxicology 14, 4–6.[ISI][Medline]
Ramesh, G. T., Manna, S. K., Aggarwal, B. B., and Jadhav, A. L. (1999). Lead activates nuclear transcription factor-kappaB, activator protein-1, and amino-terminal c-Jun kinase in pheochromocytoma cells. Toxicol. Appl. Pharmacol. 155, 280–286.[CrossRef][ISI][Medline]
Ramesh, G. T., Manna, S. K., Aggarwal, B. B., and Jadhav, A. L. (2001). Lead exposure activates nuclear factor kappa B, activator protein-1, c-Jun N-terminal kinase and caspases in the rat brain. Toxicol. Lett. 123, 195–207.[CrossRef][ISI][Medline]
Scheplyagina, L. A. (2005). Impact of the mother's zinc deficiency on the woman's and newborn's health status. J. Trace Elem. Med. Biol. 19, 29–35.[CrossRef][ISI][Medline]
Scortegagna, M., and Hanbauer, I. (2000). Increased AP-1 DNA-binding activity and nuclear REF-1 accumulation in lead-exposed primary cultures of astrocytes. Neurochem. Res. 25, 861–866.[CrossRef][ISI][Medline]
Shaulian, E., and Karin, M. (2002). AP-1 as a regulator of cell life and death. Nat. Cell Biol. 4, E131–E136.[CrossRef][ISI][Medline]
Sreedhar, B., Subramaniyan, R., and Nair, K. M. (2004). A protective role for zinc on intestinal peroxidative damage during oral iron repletion. Biochem. Biophys. Res. Commun. 318, 992–997.[CrossRef][ISI][Medline]
Toone, W. M., Morgan, B. A., and Jones, N. (2001). Redox control of AP-1-like factors in yeast and beyond. Oncogene 20, 2336–2346.[CrossRef][ISI][Medline]
Villeda-Hernandez, J., Barroso-Moguel, R., Mendez-Armenta, M., Nava-Ruiz, C., Huerta-Romero, R., and Rios, C. (2001). Enhanced brain regional lipid peroxidation in developing rats exposed to low level lead acetate. Brain Res. Bull. 55, 247–251.[CrossRef][ISI][Medline]
White, R. F., Diamond, R., Proctor, S., Morey, C., and Hu, H. (1993). Residual cognitive deficits 50 years after lead poisoning during childhood. Br. J. Ind. Med. 50, 613–622.[ISI][Medline]
Zago, M. P., Mackenzie, G. G., Adamo, A. M., Keen, C. L., and Oteiza, P. I. (2005). Differential modulation of MAP kinases by zinc deficiency in IMR-32 cells: Role of H(2)O(2). Antioxid. Redox Signal. 7, 1773–1782.[CrossRef][ISI][Medline]
Zago, M. P., and Oteiza, P. I. (2001). The antioxidant properties of zinc: Interactions with iron and antioxidants. Free Radic. Biol. Med. 31, 266–274.[CrossRef][ISI][Medline]
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