Antioxidant and Antiapoptotic Function of Metallothioneins in HL-60

Chem. Res. Toxicol. 2000, 13, 1275-1286

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Antioxidant and Antiapoptotic Function of Metallothioneins in HL-60 Cells Challenged with Copper Nitrilotriacetate Kazuaki Kawai,† Shang-Xi Liu,† Vladimir A. Tyurin,†,‡ Yulia Y. Tyurina,†,‡ Grigory G. Borisenko,† Jian Fei Jiang,† Claudette M. St. Croix,†,§ James P. Fabisiak,† Bruce R. Pitt,†,§ and Valerian E. Kagan*,†,§ Departments of Environmental and Occupational Health and Pharmacology, University of Pittsburgh, Pittsburgh, Pennsylvania 15238 Received May 30, 2000

Antioxidant activity is believed to be an important intracellular function of metallothioneins (MT), yet the specific mechanisms of their antioxidant action are not known. Under conditions when cells are challenged with elevated concentrations of free copper as a result of metabolic disturbances or environmental and occupational exposures, MTs may be ideally suited for antioxidant function as effective copper chelators. In the study presented here, we tested this hypothesis using a recently established model of copper nitrilotriacetate-induced oxidative stress in HL-60 cells. Since copper-induced oxidative stress triggers apoptosis, we further investigated antiapoptotic function of MTs in HL-60 cells. Using a Sephadex G-75 chromatographic partial purification of MTs from cell homogenates with subsequent immuno-dot-blot assay, we showed that zinc pretreatment yielded a pronounced induction of MTs in HL-60 cells. We report that zinc-induced MTs were able to (i) completely bind intracellular copper, (ii) completely quench redox-cycling activity of copper, (iii) significantly inhibit copper-dependent oxidative stress in membrane phospholipids, and (iv) prevent copper-dependent apoptosis and its characteristic biochemical features (cytochrome c release from mitochondria into cytosol, caspase-3 activation, and externalization of phosphatidylserine in plasma membranes). In separate experiments, we used lung fibroblasts derived from MT1, MT2 knockout mice (MT-/-) and MT wild-type (MT+/+) mice. ZnCl2 pretreatment resulted in a more than 10-fold induction of MTs in MT+/+ cells, whereas the MT content in MT-/- cells remained low, at levels ≈100-fold lower than in their MT wild-type counterparts. MT-/- cells were very sensitive to Cu-NTA and, most importantly, showed no response to ZnCl2 pretreatment. In contrast, MT+/+ cells were relatively more resistant to Cu-NTA, and this resistance was remarkably enhanced by ZnCl2 pretreatment. Combined, our results demonstrate that metallothioneins function as effective antioxidants and an antiapoptotic mechanism in copper-challenged HL-60 cells.

Introduction Metallothioneins (MTs)1 are low-molecular mass (6-7 kDa) ubiquitous proteins unusually rich in cysteines (1). Because the cysteines in MTs are organized in thiolate clusters and universally conserved across different species, it is accepted that the cysteines are necessary for MT function (2). While multiple functions for MTs have been proposed, such as a “storehouse” for metals, an antioxidant (free radical scavenger), and a scavenger of * To whom correspondence should be addressed: Department of Environmental and Occupational Health, University of Pittsburgh, 260 Kappa Dr., Pittsburgh, PA 15238. Telephone: (412) 967-6516. Fax: (412) 624-1020. E-mail: [email protected]. † Department of Environmental and Occupational Health. ‡ On leave from Institute of Evolutionary Physiology and Biochemistry, Russian Academy of Sciences, St. Petersburg, 194223 Russia. § Department of Pharmacology. 1 Abbreviations: AMC, 7-amino-4-methylcoumarin; BCS, bathocuprione sulfonate (4,7-phenylsulfonyl-2,9-dimethyl-1,10-phenanthroline); Cu-NTA, copper nitrilotriacetate; DTDP, 2,2′-dithiodipyridine; DTT, dithiothreitol; EGTA, ethylenebis(oxyethylenenitrilo)-N,N,N′,N′tetraacetic acid; FBS, fetal bovine serum; HMW protein, high-molecular weight protein; MTs, metallothioneins; PBS, phosphate-buffered saline; PS, phosphatidylserine; PnA, cis-parinaric acid [(9Z,11E,13E,15Z)-octadecatetraenoic acid].

toxic electrophiles (2, 3), none of them has been unambiguously proven. As has been recently pointed out by Palmiter (4), “the evolutionary forces that led to the initial appearance, gene duplications, and nearly ubiquitous expression of MTs remain enigmatic.” Conditions associated with Cu overload due to metabolic disorders (e.g., Wilson’s disease) or environmental and occupational factors may result in Cu-induced genoand cytotoxicity (5). The latter includes underground and river water contaminated by Cu from surrounding mines and smelting facilities, Cu in particulate fly ashes, Cu in waste byproducts generated by the poultry industry as a result of including excess trace elements in feed, and Cu dissolved from domestic plumbing utilizing Cu pipes (6-9). It is generally accepted that Cu toxicity results from the ability of Cu to function as a transition metal with redox cycling capacity in the presence of molecular oxygen, ultimately giving rise to the generation of a variety of reactive oxygen species capable of producing oxidative stress (10). The notion that MTs function as antioxidants, first suggested in the mid-1980s (11), is becoming increasingly attractive (2, 3, 12). Mechanisms underlying this function

10.1021/tx000119l CCC: $19.00 © 2000 American Chemical Society Published on Web 11/11/2000

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may include direct scavenging of free radicals, complexation of redox active transition metals, altered zinc (Zn) homeostasis, or interaction with GSH (3). In vitro studies have revealed that multiple cysteines in MT may react directly with reactive oxygen and nitrogen species, including superoxide, hydroxyl radicals, hydrogen peroxide (11-13), and nitric oxide (14, 15). While this mechanism of MT’s antioxidant action seems plausible, it is not clear if MT can protect against Cu-dependent cytotoxicity by virtue of its direct antioxidant quenching of reactive oxygen species or primarily by its ability to bind Cu with high affinity. In addition, the potential of MT-sequestered Cu to exert redox cycling activity in vivo has received little study. Last, since oxidative stress is considered a component in the execution of apoptotic cell death, MT may modulate the programmed cell death pathway initiated by multiple diverse stimuli. MTs have high affinity for copper (Cu) [stability constant of 1019-1017 based on pH stability and ligand substitution experiments (1)]. In addition, MT-bound Cu is redox inactive in the absence of oxidative or nitrosative stress (16). Therefore, MTs may be ideally suited for antioxidant function as effective Cu chelators under conditions when cells are challenged with elevated free Cu concentrations as a result of metabolic disturbances or environmental and occupational exposures. Results from in vitro experiments, however, do not provide unequivocal evidence for antioxidant activity of MT against Cu-induced oxidative stress. For example, Suzuki et al. (17) demonstrated production of hydroxyl radicals by Cu-containing MTs. Furthermore, Oikawa et al. (18) reported that Cu-MT itself could induce DNA damage. How relevant these in vitro findings are to actual intracellular environments is still not clear. With this in mind, we used a recently established model of copper nitrilotriacetate (Cu-NTA)-induced oxidative stress and apoptosis in HL-60 cells to establish whether MT binding of Cu may provide antioxidant protection and prevent apoptosis. We report that ZnCl2 pretreatment of HL-60 cells yielded induction of MT that was able to (i) completely bind intracellular Cu, (ii) completely quench redox-cycling activity of Cu, (iii) significantly inhibit Cu-dependent oxidative stress in membrane phospholipids, and (iv) prevent Cu-dependent apoptosis and its characteristic biochemical features [cytochrome c release, caspase-3 activation, and phosphatidylserine (PS) externalization]. Thus, we demonstrate that MTs function as effective antioxidants and an antiapoptotic mechanism in Cu-challenged HL-60 cells.

Experimental Procedures Materials and Reagents. All tissue culture media and additives were obtained from GIBCO BRL (Gaithersburg, MD) except fetal bovine serum (FBS), which was from Sigma Chemical Co. (St. Louis, MO). CuSO4, disodium nitrilotriacetic acid, 2,2′-dithiodipyridine (DTDP), Ponceau S, and Hoechst 33342 were also obtained from Sigma. Chloroform, methanol, hexane, 2-propanol (HPLC grade), and Tween 20 were purchased from Aldrich Chemical Co. (Milwaukee, WI). Alamar Blue and camptothecin were purchased from BioSource International (Sacramento, CA) and Alexis (San Diego, CA), respectively. Bathocuprione sulfonate (BCS; 4,7-phenylsulfonyl-2,9-dimethyl1,10-phenanthroline) was from Fisher Scientific (Pittsburgh, PA). cis-Parinaric acid [(9Z,11E,13E,15Z)-octadecatetraenoic acid] was obtained from Molecular Probes (Eugene, OR). Mono-

Kawai et al. clonal mouse antibody (E9) recognizing both MT1 and MT2 was purchased from DAKO Corp. (Carpinteria, CA). Mouse anticytochrome c monoclonal antibody and horseradish peroxidaseconjugated goat anti-mouse IgG specific polyclonal antibody were obtained from PharMingen International (San Diego, CA). SuperSignal West Pico Chemiluminescent Substrate was purchased from Pierce (Rockford, IL). Fuji X-ray film was purchased from Fisher Scientific. All other chemicals and reagents that were used were molecular biology grade. The cupric nitrilotriacetate solution (Cu-NTA) was prepared according to the method described by Toyokuni et al. (19), and the CuSO4 to nitrilotriacetic acid disodium molar ratio used was 1:2. The pH was adjusted to 7.4 with sodium bicarbonate. Cell Culture and Treatments. HL-60 cells were grown in RPMI medium 1640 supplemented with 12% fetal bovine serum (FBS) at 37 °C under a 5% CO2 atmosphere. Cells from passages 25-40 were used for the experiments. Some cells (5 × 105 cells/ mL) were first pretreated with ZnCl2 (150 µM) in medium containing 12% FBS under the same conditions described above for 24 h. ZnCl2 was then removed by centrifugation and washing of cells with serum-free medium. The cells were resuspended in FBS-containing RPMI medium 1640 at a density of 106 cells/ mL and cultured for 4 or 14 h in the presence or absence of Cu-NTA. Cu-NTA was removed from cells by repeated washing before cells were utilized for various assays. MT1 and MT2 null and wild-type mice were obtained by mating heterozygous mice (20) of 129 Ola and C57Bl/6 backgrounds. MT mutants were identified through a genotyping protocol using the polymerase chain reaction strategy to detect a neomyocin sequence inserted within the MT2 altered gene. Mouse lung fibroblasts were isolated essentially as described by Harvey et al. (21) except that lung tissue was utilized instead of dissected embryos. Tissue was first dispersed by passage through a 1 cm3 syringe using a 19 gauge needle, and cells were isolated by collagenase digestion. The dissociated cells were grown in DMEM supplemented with 20% FCS, penicillin (100 µg/mL), and streptomycin (100 µg/mL). Cells were split at a ratio of 1:3 or 1:4 every 48 h until they were used in experiments. MT levels in knockout (MT-/-) and wild-type (MT+/+) cells with and without ZnCl2 pretreatment were determined using the 109Cd binding assay as described by Eaton and Toal (22). The cellular MT content was calculated on the basis of the assumption that 7 mol of Cd is bound to 1 mol of MT, and was normalized to total cellular protein content. Viability Assay. Cell viability was measured by quantifying the reduction of the fluorogenic indicator Alamar Blue. Briefly, cells were resuspended in medium containing 12% FBS at a density of 5 × 105 cells/mL and applied to 96-well plates (200 µL/well). Twenty microliters of an Alamar Blue solution was added to each well. The cells were incubated at 37 °C for 4 h, and the fluorescence was determined in a Cytofluor 2350 fluorescence plate reader (Millipore, Bedford, MA) using an excitation filter at 530 ( 25 nm and an emission filter at 590 ( 35 nm. Apoptosis Assay. Nuclear morphology was assessed as previously described in our laboratory using Hoechst 33342 fluorescence staining (23, 24). The percentage of apoptotic cells was determined by counting the number of nuclei exhibiting chromatin condensation and fragmentation characteristic of apoptosis after observing at least 300 total cells under fluorescence microscopy. Separation of MT from High-Molecular Weight (HMW) Protein Thiols and GSH in Cell Homogenates. Cells (108) with or without ZnCl2 pretreatment were washed twice with cold PBS and resuspended in N2-saturated potassium phosphate buffer (10 mM, pH 7.5) containing 2.0 mM EDTA and sonicated using a 4710 series Ultrasonic homogenizer for 1 min. The resulting suspension was centrifuged at 40000g for 20 min. The supernatant was collected, applied to a 1 cm × 40 cm Sephadex G-75 column for size-exclusion chromatography, and eluted with 10 mM Tris-HCl and 2.0 mM EDTA (pH 7.5) (saturated with N2) at a rate of 0.5 mL/min at 4 °C, and fractions (0.5 mL) were

Antioxidant and Antiapoptotic Metallothioneins collected. The thiol content in each fraction was measured by adding 5 µL of a 5× concentrated DTDP solution [0.4 mM DTDP in 1.0 M sodium acetate (pH 4.0)] to a 20 µL aliquot of each fraction. After incubation for 30 min at room temperature, the absorbance at 343 nm was recorded using a SpectroMate micro spectrophotometer with a 2 µL cuvette micropipet (World Precision Instruments, Inc., Sarasota, FL). Measurement of MT Content by Immuno-Dot-Blot Assay. MT content in the fractions was measured by immunodot-blot assay with a modification of the method described by Garrett et al (25). Purified MT1 (Sigma) was used as a standard. A 5 µL aliquot from each Sephadex G-75 column fraction was diluted to 50 µL with 10 mM Tris-HCl buffer (pH 7.5), mixed with an equal volume of 3% glutaraldehyde, and then applied to a nitrocellulose membrane using a Bio-Dot Microfiltration Apparatus (Bio-Rad, Hercules, CA). The membrane was blocked with 5% fat-free milk in 50 mM Tris-HCl buffer (pH 7.5) containing 200 mM NaCl and 0.05% Tween 20 (TBST) for 1 h and then incubated with mouse monoclonal anti-MT (E9) antibody for 1 h at room temperature. Nonbinding primary antibody was removed by washing with TBST (6 times, 5 min each). The membrane was then incubated with horseradish peroxidase-conjugated polyclonal anti-mouse IgG (1:5000) in TBST for 1 h. Nonbinding antibody was removed by the same washing as described above. The membrane was finally incubated with SuperSignal West Pico Chemiluminescent Substrate and exposed to Fuji X-ray Fuji film. BCS/Ascorbate Assay of “Loosely Bound” and Total Cu in Chromatographic Fractions of HL-60 Cell Lysates. Lysates were prepared from HL-60 cells by suspending 107 cells in 100 µL of 10 mM Tris-HCl buffer (pH 7.4) and sonicating them for 1 min using a 4710 series Ultrasonic homogenizer. The resulting suspension was centrifuged at 40000g for 20 min. The cleared lysate was either assayed directly for Cu content (as described below) or applied to a Sephadex G-75 column. Column fractions corresponding to HMW proteins, MT, and GSH were identified by a DTDP assay of thiols and an immuno-dot-blot assay and separately pooled. These pooled fractions (≈5 mL) were evaporated using a SpeedVac (Savant Instruments, Inc., Farmingdale, NY) and concentrated 15-fold. The resulting samples were added to a BCS (100 µM) and ascorbate (400 µM) solution in 10 mM Tris-HCl buffer (pH 7.4). The amount of Cu available to BCS chelation was evaluated by measuring the optical density of the BCS-Cu(I) complex using a molar absorbance coefficient of 13 500 M-1 cm-1 at 480 nm (26) using a SpectroMate UV/vis fiber-optic micro spectrophotometer (World Precision Instruments) with a capillary microcuvette (sample volume of 15 µL and optical path length of 10 cm). Loosely bound Cu was defined as the amount accessible to BCS binding after incubation for 30 min at room temperature with BCS and ascorbate. The total amount of Cu was taken as the amount of the Cu-BCS complex formed after complete oxidation of thiols in the sample by treatment for 20 min with H2O2 (10 mM). Addition of catalase (80 units/mL) was used to stop the oxidative reaction prior to measurement of Cu-BCS absorbance. Caspase-3 Activity. Caspase-3 activity in HL-60 cell lysates was measured using a fluorometric assay kit obtained from Molecular Probes essentially as described in the manufacturer’s instructions. Briefly, 106 cells from various treatments were collected and lysed in 50 µL of ice-cold lysis buffer supplied with the kit. Fifty microliters of the 2× substrate working solution with 10 mM DTT and 0.2 mM fluorescent substrate, DEVD-7amino-4-methylcoumarin conjugate, was added to each lysate and kept on ice. Samples were transferred to individual wells of a Corning 96-well plate (Acton, MA) and incubated at room temperature for 30 min. The free 7-amino-4-methylcoumarin (AMC) fluorescence was determined at zero time and following incubation for 30 min at room temperature using a CytoFluor 2350 (Millipore) fluorescence plate reader using an excitation filter at 360 ( 40 nm and an emission filter at 460 ( 40 nm. The amount of product formed was calculated using a standard curve obtained for various dilutions of AMC.

Chem. Res. Toxicol., Vol. 13, No. 12, 2000 1277 Cytochrome c Release from Mitochondria. Cells (5 × 107) from various treatments were harvested by centrifugation at 700g for 10 min. The cell pellets were resuspended in buffer containing 20 mM HEPES/KOH (pH 7.5), 10 mM KCl, 1.5 mM MgCl2, 1.0 mM sodium EDTA, 1.0 mM sodium EGTA, 1.0 mM DTT, 0.1 mM phenylmethanesulfonyl fluoride, and 250 mM sucrose (27). The cells were homogenized with 10 strokes of a Potter-Elvehjem homogenizer, and the homogenates were centrifuged at 700g for 10 min. The supernatants were centrifuged at 100000g for 1 h at 4 °C, and the resulting pellets and supernatants were assayed for cytochrome c content by Western blotting. Samples containing 10 µg of protein were resolved on 12% SDS-PAGE gels and transferred to 0.2 µm nitrocellulose membranes (Bio-Rad). Ponceau S staining was applied to verify that equal amounts of protein were present in each lane. The membranes were blocked with 5% nonfat milk in TBST for 1 h at room temperature and subsequently probed with primary anti-cytochrome c antibody (1:100). Membranes were washed and then incubated with horseradish peroxidase-linked antimouse IgG secondary antibody (1:2500) for 1 h. Cytochrome c bands were visualized using the chemiluminescence assay system (Pierce) on Fuji X-ray film. Image capture and subsequent analysis were performed using a Fluor-S MultiImager and Multi-Analyst Software (Bio-Rad). Determination of the Extent of Cu-NTA-Induced Lipid Peroxidation in HL-60 Cells. cis-Parinaric acid (PnA) was incorporated into naı¨ve or ZnCl2-pretreated HL-60 cells (106 cells/mL) by addition of the PnA-human serum albumin complex which gave a final concentration of 2.0 µg of PnA/106 cells in serum-free RPMI medium 1640 without phenol red as described previously (28). PnA-labeled cells were treated with Cu-NTA (2.0 mM) for 1 h at 37 °C in RPMI medium 1640 with 10% FBS. At the end of the incubation period, cells were centrifuged and washed twice with PBS, and the total lipids were extracted using the Folch procedure (29) in the presence of butylated hydroxytoluene to retard subsequent oxidations. The lipid extract was dried under N2, dissolved in 0.2 mL of a 2-propanol/hexane/water mixture (4:3:0.16, v:v:v), and separated by normal phase HPLC using a 5 µm Microsorb-MV Si column (4.6 mm × 250 mm) and an ammonium acetate gradient as described previously (28). The separations were performed using a Shimadzu HPLC system (LC-600) (Kyoto, Japan) equipped with an in-line RF-551 fluorescence detector. The fluorescence of PnA was measured at 420 nm (emission) after excitation at 324 nm. Data were processed and stored in digital form with Shimadzu EZChrom software. The amount of lipid phosphorus was determined using a micro method (30). Annexin V Binding. Annexin V binding to cells was performed using flow cytometry essentially as previously described (17, 18) with a commercially available staining kit (R&D Systems, Minneapolis, MN). Briefly, 1 mL of HL-60 cells (106 cells/mL) was taken after treatments and washed twice with cold PBS. Cells were incubated with the annexin V-fluorescein conjugate (1 µg/mL) and propidium iodide (5 µg/mL) for 20 min at room temperature. Cells were analyzed with a FACScan flow cytometer (Becton-Dickenson, San Jose, CA) with simultaneous monitoring of green fluorescence (530 nm, 30 nm band-pass filter) for the annexin V-fluorescein conjugate and red fluorescence (long-pass emission filter that transmits >650 nm light) associated with propidium iodide. EPR of Ascorbate Radical. Ascorbate (60 µM) was added to a suspension of HL-60 cells (5 × 106 cells) in 60 µL of 50 mM disodium phosphate buffer (pH 7.4), and EPR spectra of ascorbate radicals were recorded immediately. EPR measurements were performed on a JEOL-RE1X spectrometer (Tokyo, Japan) at 25 °C using gas-permeable Teflon tubing (0.8 mm internal diameter, 0.013 mm thickness) obtained from Alpha Wire Corp. (Elizabeth, NJ). The tube (≈8 cm in length) was filled with 60 µL of mixed sample, folded into quarters, and placed in an open 3 mm internal diameter EPR quartz tube so that the entire sample was within the effective microwave area. Spectra

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Figure 2. Annexin V binding to naı¨ve and ZnCl2-pretreated HL-60 cells exposed to Cu-NTA. Cells were assessed for PS externalization by flow cytometry using annexin V binding. Data represent the means ( the SEM of the percentage of annexin V positive/propidium iodide negative cells obtained from three observations at each point. Statistically significant difference relative to naı¨ve cells in the absence of Cu (one asterisk) and relative to naı¨ve Cu-exposed cells (two asterisks) (p < 0.05).

Figure 1. ZnCl2 pretreatment maintains HL-60 cell viability and inhibits apoptosis during Cu-NTA exposure. HL-60 cells (105 cells/mL) were pretreated with 150 µM ZnCl2 at 37 °C for 24 h, and ZnCl2 was removed by washing with serum-free medium. These cells and naı¨ve cells (non-ZnCl2-pretreated) were then exposed to different concentrations of Cu-NTA (as indicated) at 37 °C for 14 h. Preliminary experiments revealed that both naı¨ve and ZnCl2-pretreated cells in the absence of Cu-NTA were >95% viable as determined by trypan blue exclusion. Also shown are the cell viability as determined by the alamar blue assay 4 h after Cu-NTA exposure (b) and the percentage of apoptotic cells as determined by changes in nuclear morphology (O). Values represent means ( the SEM normalized to control values for naı¨ve (A, n ) 3) and ZnCl2-pretreated (B, n ) 3) HL-60 cells. of ascorbate radicals were recorded with the following conditions: 335.5 mT center field, 20 mW power, 0.050 mT field modulation, 5 mT sweep width, 4000 receiver gain, and 0.1 s time constant. Quantitation of the Protein. The protein concentration in the HL-60 cell homogenates was determined with the Bio-Rad protein assay kit. A standard curve was established by addition of bovine serum albumin to the Bio-Rad assay kit, and the protein content was calculated. Statistical Analysis. The results are presented as means ( the SEM values of at least three experiments, and statistical analysis was performed by a paired Student’s t test or one-way analysis of variance (ANOVA). The statistical significance of differences was set at p < 0.05.

Results Pretreatment of HL-60 Cells with ZnCl2 Protects against Cu-NTA-Induced Cytotoxicity and Apoptosis. Cu-NTA caused a concentration-dependent cytotoxicity in naı¨ve (non-ZnCl2-pretreated) HL-60 cells [Figure 1A (b)]. A 50% loss of cell viability was observed after incubation for 14 h with 1.0 mM Cu-NTA. A more than 90% decrease in cell viability was induced by 2.0 mM Cu-

NTA. This is in line with previously reported results on cytotoxicity of Cu-NTA to HL-60 cells (25). Pretreatment of HL-60 cells with ZnCl2 (150 µM, 24 h), however, provided substantial protection of cells against Cu-NTA toxicity [Figure 1B (b)]. Neither 1.0 nor 2.0 mM Cu-NTA produced any cytotoxicity in ZnCl2-pretreated cells. More than 80% of the cells were still viable after exposure to 4.0 mM Cu-NTA. As has been reported by Ma et al. (31), Cu-NTA induced apoptosis in HL-60 cells. Similarly, we found a concentration-dependent increase in the amount of apoptotic cells after incubation of naı¨ve HL-60 cells with different concentrations of Cu-NTA [Figure 1A (O)]. Pretreatment of cells with 150 µM ZnCl2 for 24 h before Cu-NTA challenge completely inhibited apoptosis induced by Cu-NTA [Figure 1B (O)]. Even at the highest Cu-NTA concentration that was tested (4.0 mM), the number of cells containing apoptotic nuclei was not different from that in ZnCl2-pretreated cells in the absence of Cu challenge. Since the loss of cell viability correlates well with the number of apoptotic cells (Figure 1A), it appears that the primary mode of death following exposure of HL60 cells to Cu-NTA is apoptosis. Effects of Cu-NTA and ZnCl2 on PS Externalization. PS externalization, as assessed by annexin V binding, is an early marker of apoptosis (32). The function of PS exposure on the cell surface is to provide a signal for recognition and elimination of apoptotic cells by phagocytic macrophages (33). We next determined the effects of Cu-NTA and ZnCl2 on PS externalization, another early biomarker of apoptosis. As shown in Figure 2, exposure of naı¨ve HL-60 cells to 1.0 mM Cu-NTA for 14 h significantly increased the percentage of cells that were positive for annexin V binding (reflecting externalized PS) and negative for propidium iodide uptake (reflecting maintenance of membrane integrity). Pretreatment of HL-60 cells with 150 µM ZnCl2 for 24 h prior to Cu-NTA exposure completely prevented externalization of PS caused by Cu-NTA. Thus, ZnCl2 pretreatment of HL-60 cells provides substantial protection of HL-60 cells against the apoptosis-inducing effects of Cu-NTA. Effects of Cu-NTA and ZnCl2 on Cytochrome c Release from Mitochondria. Permeability transition (34) and cytochrome c release from mitochondria (27, 35)

Antioxidant and Antiapoptotic Metallothioneins

Figure 3. ZnCl2 pretreatment prevents Cu-NTA-induced cytochrome c release into cytosol and its loss from mitochondria. The inset shows typical Western blots of cytochrome c in cytosolic fractions (A) and in mitochondria (B). The numbers 1-3 represent naı¨ve, Cu-NTA-loaded, and ZnCl2-pretreated/CuNTA-loaded cells, respectively. Quantification of the optical density for the cytochrome c signals was carried out using a Fluor-S MultiImager and Multi-Analyst Software (Bio-Rad). Data are means ( the SEM from three independent experiments. Statistically significant difference relative to non-Cuexposed naı¨ve (one asterisk) and relative to naı¨ve Cu-exposed cells (two asterisks) (p < 0.05).

are two early events in the common pathway preceding caspase-3 activation, the key step in the execution of the apoptotic program induced by different stimuli (36, 37). Therefore, we next determined whether Cu-NTA caused release of cytochrome c from mitochondria and whether ZnCl2 pretreatment was able to protect against this effect. Figure 3 shows that after exposure of naı¨ve HL-60 cells to 2.0 mM Cu-NTA for 4 h, the concentration of cytochrome c in the cytosol fraction was increased almost 2-fold. The appearance of cytochrome c in cytosol (Figure 3A) occurred simultaneously with a significant loss of cytochrome c from mitchondria (Figure 3B). In ZnCl2pretreated HL-60 cells, however, elevated levels of cytochrome c in the cytosolic fraction could not be detected following Cu-NTA treatment and the cytosolic content of cytochrome c was essentially the same as that observed in both naı¨ve or ZnCl2-pretreated cells in the absence of Cu-NTA exposure. The content of cytochrome c in mitochondria of cells pretreated with ZnCl2 and exposed to Cu-NTA was not different from that in naı¨ve cells (Figure 3B). Thus, the protective action of ZnCl2 pretreatment is exerted early in the course of Cu-dependent apoptosis at a point upstream from the mechanisms that signal the mitochondrial changes necessary for the execution of apoptosis. Effects of Cu-NTA and ZnCl2 on Caspase-3 Activity in HL-60 Cells. We were next interested in studying the effects of ZnCl2 pretreatment on caspase-3 activity in HL-60 cells. Figure 4 shows the effects of ZnCl2 pretreatment on Cu-NTA-dependent activation of caspase3. When naı¨ve (non-ZnCl2-pretreated) HL-60 cells were challenged with 1.0 mM Cu-NTA, there were statistically significant 200 and 450% increases in caspase activity after exposure for 4 and 14 h, respectively (Figure 4). A similar activation was also observed when cells were

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Figure 4. ZnCl2 pretreatment prevents Cu-NTA-dependent caspase-3 activation. Naı¨ve or ZnCl2-pretreated HL-60 cells were exposed for 4 or 14 h to indicated concentrations of Cu-NTA in RPMI 1640 medium with 12% FBS. Control cells were left untreated in the same medium. Extracts from 106 cells were prepared and assayed for caspase-3 activity using 7-amino-4methylcoumarin (AMC) formed by the caspase-dependent cleavage from the substrate, DEVD-AMC, after incubation for 30 min as described in Experimental Procedures. Data represent means ( the SEM obtained from three separate experiments. One asterisk denotes a statistically significant difference compared to naı¨ve cells not exposed to Cu (p < 0.05). Two asterisks denote a statistically significant difference compared to naı¨ve cells not exposed to Cu (p < 0.01). A plus denotes a significant difference compared to similar Cu-exposed cells not receiving ZnCl2 pretreatment (p < 0.05).

exposed to 2.0 mM Cu-NTA. Caspase activation after exposure for 4 h to 2.0 mM Cu-NTA was statistically significant; however, activity measured after exposure for 14 h failed to obtain statistical significance. This most likely reflects the fact that exposure of naı¨ve cells to 2 mM Cu-NTA produced a much more rapid and robust apoptosis than 1 mM Cu-NTA. Measurement of caspase-3 activity in late stage apoptotic cells may not accurately reflect the degree of prior activation. Indeed, caspase-3 activation may not be detected at these late stages as a result of protein degradation or loss of enzymatic activity. In line with this, the recovery of total protein from cells treated for 14 h with 2 mM Cu-NTA was substantially less than in 4 h or in cells treated with 1 mM Cu-NTA. This fact indicates that significant protein degradation, dissolution of cells into apoptotic bodies, or onset of secondary necrosis could serve to reduce the level of caspase-3 activation after exposure to high Cu concentrations for prolonged periods of time. As shown in Figure 4, exposure of ZnCl2-pretreated HL-60 cells to similar concentrations of Cu-NTA failed to produce any significant activation of caspase-3 after incubation for both 4 and 14 h. Effects of ZnCl2 on Camptothecin-Induced Apoptosis and Caspase-3 Activation in HL-60 Cells. We employed ZnCl2 pretreatment as a method of inducing MT overexpression in HL-60 cells. Since Zn2+ is known to inhibit caspase-3 at extremely low (nanomolar) concentrations (38), however, we decided to test to what extent the protective effects of Zn against Cu-NTAmediated apoptosis could be attributed to MT induction or a direct inhibitory effect of Zn upon apoptotic execution signaling pathways. To this end, we determined the effects of ZnCl2 pretreatment on apoptosis induced by another stimulus besides Cu, namely, camptothecin (an inhibitor of topoisomerase I) (39). In line with its potent ability to produce apoptosis, we found that 10 µM

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Figure 5. Effects of camptothecin on caspase-3 and apoptosis in HL-60 cells. Caspase-3 activity (A) and apoptosis as assessed by changes in nuclear morphology (B) induced by camptothecin (10 µM, 2 h) in naı¨ve HL-60 cells, ZnCl2-pretreated HL-60 cells (150 µM ZnCl2 for 24 h), and ZnCl2-pretreated/Cu-NTA-loaded HL-60 cells (150 µM ZnCl2 for 24 h followed by 2.0 mM CuNTA for 14 h). Cells were washed free of exogenous metal ions after each exposure prior to camptothecin treatment. Data represent means ( the SEM from three independent experiments. One asterisk denotes a statistically significant difference compared to cells not exposed to camptothecin (p < 0.05).

camptothecin effectively activated caspase-3 (Figure 5A) and induced apopotosis as assessed by changes in nuclear morphology (Figure 5B) in HL-60 cells. Most importantly, ZnCl2 pretreatment did not protect HL-60 cells against camptothecin-induced caspase-3 activation (Figure 5A), nor did it prevent a camptothecin-induced increase in the amount of apoptotic HL-60 cells (Figure 5B). This was true for ZnCl2 pretreatment alone, as well as during the Cu exposure after ZnCl2 pretreatment when MT-bound Zn could have been displaced by preferential binding of Cu to MT. Therefore, it is apparent that the protective effects of ZnCl2 pretreatment are specific for Cu-dependent apoptosis and not generalized to apoptosis induced by other agents. The most likely mechanism of protection is via the induction of MT, an important metal-binding protein thought to regulate metal ion homeostasis and sequester free metal ions possessing toxic redox-cycling potential. Effect of ZnCl2 on the Distribution of Thiols and MT Assayed by Sephadex G-75 Chromatography. Our results shown above demonstrated that ZnCl2 pretreatment protected HL-60 cells against Cu-NTA-induced apoptosis. Since binding of Cu by thiolate clusters in MTs represents one of the major intracellular Cu sequestration mechanisms, we next studied the effects of Zn on the MT content of HL-60 cells. To this end, we used Sephadex G-75 size exclusion chromatography to separate MT from HMW proteins and GSH in cell lysates. Figure 6 shows a typical Sephadex G-75 profile of thiol distribution in lysates obtained from naı¨ve cells and Znpretreated cells. On the basis of our DTDP assay of SH groups, HL-60 lysates contained three major peaks. Early fractions contained a peak of HMW proteins with relatively large amounts of sulfhydryl groups. Intermediate fractions composed the middle peak which has been previously characterized as containing mainly MTs (13) followed by fractions corresponding to low-molecular weight compounds, most abundantly, GSH. The thiol content in the HMW protein and GSH fractions was not significantly altered after ZnCl2 pretreatment. The immuno-dot-blot assay using anti-MT antibody demonstrated the presence of MT in the middle peak from ZnCl2-pretreated cells but not from naı¨ve cells. No MT immunoreactivity could ever be detected in HMW protein

Figure 6. Sephadex G-75 chromatographic profiles of HMW proteins, MT, and GSH from control and ZnCl2-pretreated cells. Naı¨ve HL-60 cells (105 cells/mL) were pretreated with or without 150 µM ZnCl2 at 37 °C for 24 h. Cells (108) from each group were then washed with cold PBS and cytosolic fractions prepared and analyzed for SH content using DTDP and immunoreactive MT. Shown are the distribution of SH groups upon reaction with DTDP and the resultant absorbance at 343 nm. The inset shows the results of the immuno-dot-blot assay of fractions 17-31, which correspond to the expected migration of MT, obtained from lysates prepared from control and ZnCl2pretreated cells.

or GSH fractions obtained from naı¨ve or ZnCl2-pretreated cells (data not shown). Therefore, ZnCl2 pretreatment as applied here effectively induced the expression of thiolrich MTs as measured by both immunological and biochemical means. Using serial dilutions of column fractions and purified MT standards to assess the limit of detection of our assay, the induction of MT by Zn represents an at least 50-fold increase in the level of MT protein above that observed in naı¨ve cells. Cu Content and Distribution in HL-60 Cells. Since cell survival was strictly dependent on the concentration of Cu added to the medium and MTs are potent binders of intracellular Cu, we were eager to determine the Cu content in HL-60 cells. We used the BCS/ascorbate assay to evaluate the amount of intracellular Cu that is available to the chelator (26). When homogenates of Cuexposed naı¨ve cells (not pretreated with ZnCl2) were incubated with BCS and ascorbate, the characteristic spectrum of the BCS-Cu(I) complex was detectable with the maximum at 480 nm (Figure 7, trace A). Expectedly, no absorbance at 480 nm could be obtained from naı¨ve cells grown in the absence of added Cu (Figure 7, trace B). The amounts of intracellular Cu available to the chelator were dependent on the concentration of Cu-NTA added to the medium. Furthermore, when compared to that in naı¨ve cells, the availability of Cu for the chelator was remarkably lower in ZnCl2-pretreated cells exposed to similar concentrations of Cu (Figure 7, inset). Almost 10-fold higher concentrations of Cu-NTA in the medium were required to produce the same response to the BCS/ ascorbate mixture in ZnCl2-pretreated cells (≈5.0 mM Cu-NTA) as in the cells not pretreated with ZnCl2 (≈0.5 mM Cu-NTA) (Figure 7, inset). Thus, ZnCl2 pretreatment dramatically reduced the availability of intracellular Cu



Figure 7. BCS-available Cu in HL-60 cells exposed to Cu-NTA. Lysates from naı¨ve and ZnCl2-pretreated HL-60 cells were analyzed for the amount of Cu that is available for chelation by the Cu chelator, BCS. This amount of Cu was defined as free and/or “loosely bound” since Cu/BCS optical spectra were obtained after incubation of cell lysates for 20 min with BCS (0.1 mM) and ascorbate (0.4 mM) at room temperature. Trace A shows the amount of BCS-Cu complex that can be detected as peak absorbance at 480 nm in naı¨ve cells exposed to 1.0 mM Cu-NTA for 14 h. Trace B shows the optical spectrum obtained from similar naı¨ve cells in the absence of exogenous Cu. The inset compares the amount of BCS-available Cu measured in naı¨ve and ZnCl2-pretreated cells challenged with varying amounts of Cu-NTA.

to the chelator, presumably via high-affinity binding of Cu to MT. We further used the BCS/ascorbate assay to determine the Cu distribution among the three fractions (GSH, MTs, and HMW proteins) in ZnCl2-pretreated cells. Figure 8 shows optical spectra of pooled fractions of HMW proteins (panel A) (fractions 1-9), MTs (panel B) (fractions 20-29), and the GSH fraction (panel C) (fractions 39-48). Individual traces 1 and 2 in each panel represent spectra recorded before and after the addition of the BCS/ ascorbate mixture, respectively. One can see that a pronounced spectrum of the BCS-Cu complex (maximal absorbance at 480 nm) was observed only in the MT fraction. Note that the traces obtained from the MT fraction (Figure 8B) were diluted 3-fold relative to the others to achieve similar scale absorbance measurements. This demonstrates that only the MT fraction contained Cu available for BCS chelation in the presence of ascorbate. To assess the total amount of Cu in the fractions, we preincubated them in the presence of high concentrations of H2O2 to oxidize cysteines and release Cu from SH-dependent binding sites (e.g., GSH and thiolate clusters in MTs). After oxidation, we decomposed excess H2O2 by addition of catalase and then performed the BCS/ ascorbate assay. Trace 3 in Figure 8A-C represents the BCS-Cu response after oxidation. Once again, the most notable difference was observed in the MT fraction (Figure 8B, trace 3) where oxidation produced a substantial increase in the BCS-Cu signal, indicating release of Cu from the MT thiolate clusters. Only a small increment in absorbance was detected in the HMW protein fraction (Figure 8A, curve 3), and essentially no increase in the BCS-Cu peak was found in the GSH

Figure 8. Cu content in HMW protein, MT, and GSH fractions of ZnCl2-pretreated HL-60 cells. ZnCl2-pretreated HL-60 cells were exposed to Cu-NTA and lysates prepared and subjected to Sephadex G-75 chromatography. Cu was detected by optical spectroscopy of BCS-Cu(I) complexes in the pooled fractions corresponding to HMW proteins (A), MT (B), and GSH (C). Trace 1 shows the background spectra obtained without the addition of BCS and ascorbate. Trace 2 represents the amount of “loosely bound” Cu measured 20 min after the addition of BCS (0.1 mM) and ascorbate (0.4 mM). Trace 3 represents the total Cu content measured after oxidation of SH groups by 10 mM H2O2 for 20 min followed by addition of catalase prior to the ascorbate/BCS mixture. Note that the samples in the MT fraction (B) were diluted 3-fold relative to the others to keep spectra on the same relative scale.

fraction (Figure 8C, curve 3). These results indicate that ≈90% of the total Cu (Cu available after H2O2 treatment) was confined to the MT fraction and ≈10% Cu was located in other proteins (one should bear in mind, however, that H2O2 treatment may not necessarily release Cu from non-thiolate-binding sites). Only about 15% of the total Cu was available for the BCS/ascorbate assay without H2O2 oxidation, and two-thirds of this was detected in the MT fraction (Table 1). Redox-Cycling Activity of Cu in HL-60 Cells Assayed by Ascorbate Radical Production. Since Cu can catalyze one-electron oxidation of ascorbate to produce ascorbate radicals, we took advantage of EPR measurements of ascorbate radicals to evaluate the redox-cycling activity of intracellular Cu (16). Figure 9 (inset) shows the typical EPR spectra of ascorbate radicals obtained from naı¨ve (spectrum a) and ZnCl2-

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Figure 9. Effect of ZnCl2 pretreatment on the redox-cycling activity of Cu in HL-60 cells assayed by ascorbate radical production. Incubations of cell lysates (5 × 106 cells) were performed at room temperature in 50 mM sodium phosphate buffer (pH 7.4) in the presence of ascorbate (60 µM). The inset shows typical EPR spectra of ascorbate radical measured after exposure of naı¨ve (a) and ZnCl2-pretreated (b) HL-60 cells to 1.0 mM Cu-NTA for 14 h. Shown is the average magnitude of the ascorbate radical signal measured over 8 min beginning 1 min after the addition of ascorbate. White bars represent data obtained in the absence of Cu-NTA treatment and hatched bars after exposure to Cu-NTA. The black bar shows the effect of inclusion of 700 µM BCS with Cu-exposed cell lysates of naı¨ve cells exposed to Cu along with ascorbate. The data represent the magnitude of the EPR ascorbate radical signal (means ( the SEM, n ) 6). Statistically significant difference relative to naı¨ve cells in the absence of Cu (one asterisk) and relative to naı¨ve Cu-exposed cells (two asterisks) (p < 0.05).

pretreated (spectrum b) HL-60 cells after exposure to CuNTA. Quantitatively, the data are summarized in Figure 9 as the magnitude of ascorbate radical EPR signals. The ascorbate radical EPR signal was not detectable in naı¨ve or ZnCl2-pretreated cells in the absence of exogenously added Cu. Exposure of naı¨ve HL-60 cells to 1.0 mM CuNTA yielded a pronounced EPR signal of ascorbate radicals (1 min after ascorbate addition), which was almost completely suppressed by the addition of the Cu(I) chelator, BCS. No significant production of ascorbate radical was observed from ZnCl2-pretreated HL-60 cells exposed to the same concentration of Cu-NTA. Thus, Cudependent redox cycling activity in HL-60 cells was completely suppressed by ZnCl2 pretreatment.

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Effects of Cu-NTA and ZnCl2 on Lipid Peroxidation in HL-60 Cells. Proapoptotic effects of Cu-NTA may be associated with its ability to induce oxidative stress in cells (31). To determine whether Cu-NTA induced oxidative stress in membrane phospholipids of HL-60 cells, we utilized our sensitive HPLC fluorescent technique based on metabolic integration of oxidation-sensitive fluorescent fatty acid, PnA, in membrane phospholipids of the cells (28). No significant difference in incorporation of PnA into phospholipids of naı¨ve and ZnCl2-pretreated HL-60 cells was detected. Figure 10 shows that exposure of naı¨ve HL-60 cells to Cu-NTA (2.0 mM, 1 h) resulted in more than 50% oxidation of four major classes of membrane phospholipids [phosphatidylcholine (PC), phosphatidylethanolamine (PE), PS, and phosphatidylinositol (PI)] in these cells. Similar exposure of ZnCl2-pretreated HL-60 cells to Cu-NTA (2.0 mM, 1 h) produced significantly less peroxidation of all phospholipids than in naı¨ve cells (Figure 10). Thus, pretreatment of HL-60 cells with ZnCl2 protected all major phospholipids against oxidation induced by Cu-NTA. Interestingly, the Cu-NTA-dependent oxidation of PC, PE, and PI in ZnCl2-pretreated cells was attenuated by approximately 50% compared to that in naı¨ve cells. ZnCl2 pretreatment, however, provided essentially complete protection to PS such that PnA content of PS after CuNTA was not significantly different for cells in the absence of Cu. ZnCl2 Pretreatment Protects Wild-Type MT but Not MT Null Cells against Cu-NTA-Induced Apoptosis. To specifically assign the effects of ZnCl2 pretreatment to MT induction, we sought to compare the effects of Zn in cells with intact MT and similar cells derived from MT knockout animals. For these studies, we used lung fibroblasts derived from MT null (-/-) mice and their control counterparts expressing wild-type levels of MT. In this way, the contribution of Zn-inducible MT to protection against Cu-NTA toxicity can be assessed since any non-MT-mediated effects of Zn would be expected to remain intact in the MT null cells and should be observed as Zn-induced protection in these cells.

Figure 10. Effect of ZnCl2 pretreatment on Cu-NTA-dependent oxidation of PnA-labeled phospholipids in HL-60 cells. Naı¨ve HL60 cells were incubated in the presence and in the absence of ZnCl2 (150 µM) in RPMI medium 1640 with 12% FBS for 24 h at 37 °C and washed with RPMI medium 1640 without FBS and metabolically labeled by PnA. PnA-loaded cells (106 cells/mL) were exposed to Cu-NTA (2.0 mM) in RPMI 1640 medium with 10% FBS for 1 h at 37 °C. At the end of the incubation, cells were centrifuged and washed twice with PBS, and lipids were extracted and resolved by HPLC. PI, phosphatidylinositol; PS, phosphatidylserine; PE, phosphatidylethanolamine; PC, phosphatidylcholine. Data represent means ( the SEM (n ) 3). One asterisk denotes a significant difference vs naı¨ve cells not exposed to Cu (p < 0.03). Two asterisks denote a significant difference compared to naı¨ve Cu-exposed cells. Three asterisks denote a significant difference compared to ZnCl2-pretreated cells not exposed to Cu.



Table 1. Distribution of Copper in Different Fractions of ZnCl2-Pretreated and Cu-NTA-Loaded HL-60 Cell Lysates Obtained after Sephadex G-75 Filtrationa copper (nmol/106 cells) loosely bound total

HMW proteins

MT

GSH

0.17 ( 0.05 0.28 ( 0.06

2.17 ( 0.18 37.59 ( 0.97

NDb NDb

a ZnCl -pretreated (150 µM, 24 h) HL-60 cells were exposed to 2 Cu-NTA (2 mM) for 14 h, and lysates were prepared and subjected to Sephadex G-75 chromatography. Loosely bound and total Cu in HMW proteins, MT, and GSH fractions were detected by optical spectroscopy of BCS-Cu(I) complexes as described in Experimental Procedures. Data are means ( the SEM (n ) 3). b ND means that the Cu concentration was lower than 0.04 nmol/106 cells (the limit of the measurement).

Table 2. Content of MT in MT Knockout and Wild-Type Mouse Lung Fibroblasts with and without ZnCl2 Pretreatment As Determined by the 109Cd Assaya MT content (µg/mg of protein) MT+/+ MT-/-

ZnCl2-nontreated cells

ZnCl2-treated cells

0.105 ( 0.021 0.007 ( 0.002b

1.130 ( 0.035b 0.015 ( 0.001b

a Data are means ( the SD. b Statistically significant difference relative to MT+/+ cells not treated with ZnCl2 (p < 0.001).

Table 2 verifies the MT genotype and/or phenotype and Zn response in the measurements of the MT content in fibroblasts derived from the lungs of MT1, MT2 knockout mice (MT-/-) and MT wild-type (MT+/+) control cells by the Cd109 binding assay (22). In the absence of Zn, MT+/+ cells contained approximately 0.10 µg of MT/mg of protein while MT null cells expressed 10-fold less MT (essentially below the limits of detection of this assay). As expected, ZnCl2 pretreatment resulted in a more than 10-fold induction of MT in MT+/+ cells whereas MT null cells still contained MT at levels ≈100-fold lower than that in their MT wild-type counterparts. Figure 11 shows the apoptotic response of these cell lines after exposure to Cu-NTA with and without ZnCl2 pretreatment (compare with our Figure 1 where the data with HL-60 cells are presented). It is clear that MT-/cells were the most sensitive to Cu-NTA and, most importantly, show no response to ZnCl2 pretreatment. MT wild-type cells were moderately protected against low concentrations of Cu-NTA, while at higher Cu-NTA doses (g1.2 mM), their sensitivity was high and not significantly different from that of MT-/- cells (Figure 11). Notably, ZnCl2-pretreated MT wild-type cells exhibited only very low sensitivity to Cu-NTA at both low and high concentrations. Essentially, a slight increase in the number of apoptotic cells was observed only at Cu-NTA concentrations of >0.8 mM.

Discussion Cu is one of the transition metals essential for life as its catalytic redox properties are utilized in a number of enzymatic oxidase and oxygenase metabolic pathways (40). In these enzymatic reactions, Cu is a part of the enzyme’s catalytic site and its interactions with oxygen are strictly controlled by the surrounding parts of protein molecules (41). The same redox reactions, however, render Cu extremely toxic when Cu is released from this protein matrix and in its reduced form can directly interact with molecular oxygen to produce superoxide radical and other reactive oxygen species, i.e., act as a

Figure 11. Apoptotic response of MT knockout and wild-type mouse lung fibroblasts after exposure to Cu-NTA with and without ZnCl2 pretreatment. Mouse lung fibroblasts (5 × 105 cells/well) were pretreated with 100 µM ZnCl2 at 37 °C for 24 h, and ZnCl2 was removed by washing with PBS. The cells were then loaded with different concentrations of Cu-NTA at 37 °C for 14 h. Apoptosis was assessed by changes in nuclear morphology using Hoechst 33342 staining. Values represent means ( the SD (n ) 4). One asterisk denotes a statistically significant difference from MT+/+ cells without ZnCl2 pretreatment exposed to the same concentration of Cu-NTA (p < 0.05). Two asterisks denote a statistically significant difference from MT-/- cells with and without ZnCl2 pretreatment and exposed to the same concentration of Cu-NTA (p < 0.01). A pound sign denotes a significant difference from MT-/- cells with and without ZnCl2 treatment and MT+/+ cells without ZnCl2 treatment and exposed to the same concentration of Cu-NTA (p < 0.01). Two pound signs denote a significant difference from MT+/+ cells with ZnCl2 treatment and exposed to 0-0.8 mM Cu-NTA (p < 0.01).

Fenton-type catalyst (10, 42). Therefore, Cu is always protein-bound in biological fluids (mainly by ceruloplasmin, albumin, and transcuprein) as well as in cells (primarily by MTs). Moreover, intracellular delivery of Cu to its catalytic sites is performed by special highaffinity chaperones capable of maintaining extremely low concentrations of free Cu [less than one molecule per cell (43)]. Additionally, Cu-translocating ATPases contribute to elimination of excess Cu from cells (41). It has been known for some time that MT can confer resistance to toxicity resulting from exposure of cells or animals to such heavy metals as cadmium (2). Studies have also suggested a similar effect of MT on Cu toxicity; however, much of the definitive data has been accumulated using yeast systems (44, 45) or indirect associations of MT expression with sensitivity to Cu toxicity (46-48). In addition, the exact mechanisms of protection remain to be described. This is important since protection could arise from the ability of MT to bind Cu and render it redox inactive (16, 49) and/or because MT’s reported antioxidant activity (3, 6, 50) could mitigate oxidative stress following metal exposure without direct interaction with metal ions. In addition, since oxidative stress is an important signaling component in the pathway of apoptotic execution, MTs could inhibit Cudependent programmed cell death by modulation of apoptosis pathways in general. A recent study demonstrated that excess Cu produced by incubation of HL-60 cells with Cu-NTA induced oxidative damage to DNA and apoptosis (31, 51). NTA alone does not induce oxidative DNA damage and apoptosis in HL-60 cells (31). In the work presented here, we

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used HL-60 cells to show that induction of MTs (by pretreatment of cells with ZnCl2) can protect cells against apoptosis caused by Cu-NTA. On the basis of our immuno-dot-blot measurements, we found that ZnCl2pretreated HL-60 cells expressed at least 50-fold higher levels of MTs than nontreated cells. In fact, MTs could not be detected in naı¨ve HL-60 cells. This is in line with previous reports on MT induction by Zn in HL-60 cells (13). We further established that ZnCl2-pretreated cells were protected against Cu-induced apoptosis. This was directly demonstrated by the ability of ZnCl2 pretreatment to prevent a Cu-NTA-induced increase in the amount of apoptotic cells (assayed by changes in nuclear morphology and PS externalization using the annexin V assay) as well as activation of caspase-3 and release of cytochrome c from mitochondria. Moreover, we found that the high redox-cycling activity of Cu in Cu-exposed cells was completely suppressed in ZnCl2-pretreated cells challenged with the same concentration of Cu. Finally, ZnCl2-pretreated cells were protected against Cu-NTAinduced peroxidation of major classes of membrane phospholipids observed in naı¨ve cells after exposure to Cu. Importantly, MT induction did not appear to affect apoptotic execution pathways in general since the apoptosis response following camptothecin remained intact. Therefore, it is unlikely that MT antioxidant activity can appreciably modulate the oxidative events associated with apoptosis execution. Many antitumor drugs exert their cytotoxic effects via induction of apoptosis, and MT overexpression has been postulated as one potential mechanism to account for resistance of various tumors to chemotherapy (3). One may assume, given our results here where induction of MT failed to modulate camptothecin-induced apoptosis, that MT-dependent chemoresistance may be restricted to those agents that induce DNA damage as reactive electrophiles and not through other mechanisms. We hypothesized that the Zn-mediated protection was due to enhanced Cu-binding capacity following induction of MT expression. We realize that ZnCl2 pretreatment may have pleiotropic targets of which MT induction is only one. Therefore, one may assume that specific genetic manipulation of MT in a cell line (52, 53) may be a more precise model; however, our BCS/ascorbate assay showed that, indeed, negligible amounts of Cu were found in GSH and HMW protein fractions of ZnCl2-pretreated HL-60 cells. The majority of Cu was confined to the MT fraction. In addition, this bound Cu was apparently redox inactive since Cu-dependent ascorbate radical generation was not observed in ZnCl2-pretreated cells. This is in keeping with our previous observations using chemically defined cellfree model systems (16). This indicates that binding of Cu by MTs might be an important factor in the protection against the toxic effects of free Cu. Further, our experiments with MT knockout and wildtype mouse lung fibroblasts directly demonstrated that ZnCl2 pretreatment did not cause any protection against Cu-NTA-induced apoptosis in MT-/- cells. In contrast, ZnCl2 pretreatment was remarkably protective in MT+/+ cells. These observations rule out the contribution of nonMT effects mediating protection of Cu-NTA-induced apoptosis. If ZnCl2 pretreatment produced protection by other mechanisms, we would expect these to be intact in MT-/- cells and ZnCl2 protection to be observed; however, it is not. While MT wild-type cells show moderate

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resistance to Cu-NTA (most likely reflecting their basal level of MT), they are clearly more protected by prior ZnCl2 pretreatment. Thus, we are confident that our paradigm of ZnCl2 induction of MT in HL-60 cells is indeed responsible for the observed protection against Cu cytoxicity, redox activity, and apoptosis. The fact that Cu-dependent lipid peroxidation was attenuated following MT induction by ZnCl2 pretreatment further supports the idea that Cu-dependent oxidative stress is inhibited when Cu is complexed to MT. It is of interest that we observed a difference between the degree protection of PS (100% protection) and other phospholipids (50% protection). We have recently observed that selective oxidation of PS occurs as part of the apoptotic execution program and probably arises from a unique mechanism in contrast to random lipid peroxidation (23, 24). Since Zn-mediated induction of MT completely prevents Cu-dependent apoptosis, we would expect that selective oxidation of PS would also be completely blocked despite some continued low-level Cudependent oxidative stress in membranes. There is, however, an alternative interpretation of these results. It has been recently reported that Zn can act directly as a potent inhibitor of caspase-3 (38). Pretreatment of HL-60 cells with ZnCl2 is known to yield high concentrations of Zn MTs (13). Since MTs have a higher affinity for Cu than for Zn (54), one can assume that upon exposure to Cu-NTA Zn might be released and exert its direct antiapoptotic effects in HL-60 cells. In addition, residual high-affinity interaction of Zn with caspase after the pretreatment phase could serve to lower the amount of activatable caspase. To determine whether direct effects of Zn were engaged in the protection against Cu-induced apoptosis, we performed separate experiments with camptothecin that induces apoptosis via a Cu-independent mechanism, namely, inhibition of topoisomerase I (39). As discussed above, we established that, in contrast to its effects on Cu-induced apoptosis, ZnCl2 pretreatment did not render any protection against camptothecin-induced caspase-3 activation or an increased number of cells with apoptotic nuclear morphology. These results prove that direct antiapoptotic effects of Zn were minimal in the protective action of ZnCl2 pretreatment against Cu-induced apoptosis. In summary, MTs have been reported to have antioxidant activity functioning as endogenous scavengers of oxygen radicals and other reactive oxygen and nitrogen species (3, 55). The results of our study demonstrate that the enhanced redox-cycling activity of Cu in Cu-exposed cells could be completely eliminated by both ZnCl2 pretreatment and the Cu(I) chelator, BCS. Thus, we conclude that MT acts as a potent antioxidant primarily by sequestering free Cu and preventing its redox-cycling activity in HL-60 cells and thus protecting them against Cu-induced oxidative stress and apoptosis.

Acknowledgment. This work was supported by AICR Grants 97-B128, HL32154, and HL64145-01A1, EPA STAR Grant R827151, the NCI Oncology Research Faculty Development Program and Magee-Womens Research Institute, the Leukemia Research Foundation, and the International Neurological Science Fellowship Program (F05 NS 10669) administered by NIH/NINDS in collaboration with the World Health Organization, Unit of Neuroscience, Division of Mental Health and Prevention of Substance Abuse.


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Antioxidant and Antiapoptotic Function of Metallothioneins in HL-60

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