Cu complex

European Journal of Pharmacology 594 (2008) 9–17

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European Journal of Pharmacology j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / e j p h a r

Involvement of activating transcription factors JNK, NF-κB, and AP-1 in apoptosis induced by pyrrolidine dithiocarbamate/Cu complex Sung-Ho Chen a, Jen-Kun Lin c, Yu-Chih Liang c, Min-Hsiung Pan c, Shing-Hwa Liu a, Shoei-Yn Lin-Shiau b,⁎ a b c

Institute of Toxicology, College of Medicine, National Taiwan University, Taipei, Taiwan Institute of Pharmacology, College of Medicine, National Taiwan University, Taipei, Taiwan Institute of Biochemistry, College of Medicine, National Taiwan University, Taipei, Taiwan

a r t i c l e

i n f o

Article history: Received 29 February 2008 Received in revised form 3 July 2008 Accepted 10 July 2008 Available online 16 July 2008 Keywords: PDTC Copper Oxidative stress JNK NF-κB AP-1 Caspase Apoptosis BCPS

a b s t r a c t Pyrrolidine dithiocarbamate (PDTC) is a metal chelator. Biologically, slight toxic affects EC50, 100 ± 5.9 µM are observed when added to cultured HL-60 cells. CuCl2 at a physiological concentration (1 µM), but not FeCl2, Pb potentiated the cytotoxic effect of PDTC by 700 fold (EC50, 0.14 ± 0.02 µM). Furthermore, results indicated that the PDTC/Cu complex induced an apoptotic process, evidenced by apoptotic bodies, DNA ladder and hypodiploidy cells. Additional studies showed that PDTC/Cu complex significantly decreased mitochondrial membrane potential, increased cytochrome c release, and reactive oxygen species production, and depleted reduced non-protein thiols in a time-dependent manner. Following oxidative stress, the PDTC/Cu complex sequentially activated JNK, NF-κB and AP-1 signaling pathways while IκB kinase activity was enhanced. The apoptotic process was eventually induced by caspase 3 activation and PARP degradation. The non-permeable copper-specific chelator-bathocuproine disulfonate (BCPS) and vitamin C were able to inhibit apoptosis and the elevation of intracellular Cu. Based on these findings; we conclude that PDTC/Cu complex-induced apoptosis is mediated by activation of JNK, NF-κB, AP-1 and caspase 3. Due to its high potency, PDTC may be useful as a therapeutic anti-cancer drug. © 2008 Published by Elsevier B.V.

1. Introduction Pyrrolidine dithiocarbamate (PDTC) is a thiol compound derived from dithiocarbamates (DCs), a highly characterized class of antioxidants in both free cells and biological systems (Hayes, 1982). Consequently, PDTC is often used as a potent inhibitor of apoptosis in different cell types (Nobel et al., 1997; Bessho et al., 1994) as it inhibits the function of nuclear factor κB (NF-κB, Kim et al., 1999) and NO synthase (Sherman et al., 1993). While the cellular and molecular mechanisms of PDTC-induced apoptosis remain to be elucidated, recent studies have demonstrated that the induction of PDTC mediated apoptosis occurs through a pro-oxidative pathway (Nobel et al., 1995; Chen et al., 2000). Copper is an essential transition metal ion that modulates many biological processes (Camakaris et al., 1999) including angiogenesis (Brewer, 2001). However, when in excess, Cu can significantly increase oxidative stress. Copper containing protein-chaperons are abundant in peripheral system and brain (Rothstein et al., 1999). However, it is not only an essential metal in our body, but also a major contaminant

⁎ Corresponding author. Institute of Pharmacology, College of Medicine, National Taiwan University, Section 1, Jen-Ai Road, No. 1, Taipei, 10043, Taiwan. Tel.: +886 2 23123456x8313; fax: +886 2 23915297. E-mail address: [email protected] (S.-Y. Lin-Shiau). 0014-2999/$ – see front matter © 2008 Published by Elsevier B.V. doi:10.1016/j.ejphar.2008.07.024

in our environment (Talbot, 1986). Blood is an important target of exposure to drugs and environmental chemicals. An interesting study by Zuo et al., (2006) showed that serum copper concentration was higher in leukemia patients than that of controls(Zuo et al., 2006) and hence in the present study, HL-60 cells (human promyelocytic leukemia) are used to investigate the action mechanisms of PDTC, Cu and their complex forms. Apoptosis contributes to the pathogenesis of several neurodegenerative disorders and immune deficiencies (Thompson, 1995). Morphological and biochemical events of the apoptotic process include mitochondrial membrane potential changes, activation of caspases, cellular shrinkage, chromatin condensation, and nucleosomal DNA fragmentation (Susin et al., 1997). In response to the extracellular stimuli, the mitogen-activated protein (MAP) kinase family is activated. Additional evidence suggests that the JNK/SAPK pathway may play an important role in triggering apoptosis in response to inflammatory cytokines (Zhang et al., 1996), free radicals generated by UV-C and gamma radiation (Alder et al., 1995). Moreover, direct application of hydrogen peroxide (Yu et al., 1996), cisplatin, a DNAdamaging agent (Potapova et al., 1997) or an alkylating agent such as N-nitrosoguanidine also produced apoptosis. Conversely, both AP-1 (activator protein-1) and NF-κB transcription factors have been suggested as regulators of cell death and survival (Alder et al., 1995). Supershift analysis of nuclear extracts indicated that the AP-1 complex

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consisted of c-Jun, c-Fos, JunD, and possibly JunB proteins, while the NF-κB complex contained p50, p65, and c-Rel proteins. Activation of AP-1 and NF-κB is involved in hydrogen peroxide-induced apoptotic cell death of oligodendrocytes (Derijard et al., 1994). Caspase-3 is the downstream signaling pathway of activated JNK (Kim et al., 2005). A number of ICE/CED-3 protease targets have been identified, including the nuclear enzyme poly (ADP-ribose) polymerase (PARP, Zhu et al., 1997), we examined whether this signaling pathway involved in the apoptotic process induced by PDTC/Cu complex. Here we present data showing that a low concentration of PDTC plus Cu exerts a toxic pro-oxidant effect on HL-60 cells by increasing free radical production and depleting reduced non-protein thiols. Furthermore, our findings suggest that the activation of JNKs, NF-κB, AP-1 and caspase 3 followed by PARP degradation is involved in PDTC/ Cu complex-induced apoptosis.

2.4. Flow cytometry The apoptotic cells containing hypodiploidy DNA could be measured by flow cytometric technique. After centrifugation, the pelleted cells were washed with ice-cold phosphate-buffered saline (PBS), and fixed in 70% ethanol at −20 °C for at least 1 h. The fixed cells were washed twice with PBS and incubated at 37 ± 0.5 °C for 30 min with 1 mg/ml of RNase A dissolved in 0.5 ml of 0.5% Triton X-100/PBS solution. Following the incubation, cells were stained with 0.5 ml of 50 µg/ml propidium iodide (PI) for 10 min, during which time the PI bound to the intracellular DNA. Upon excitation of the fluorescent dye by a FACScan flow cytometer (Becton Dickinson), the PI–DNA complex emitted a fluorescent signal that could be quantified. 2.5. Determination of copper contents using atomic absorption spectrophotometric technique

2. Materials and methods 2.1. Materials 3-4, 5-dimethyl thiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT), PDTC, CuCl2, vitamin C, superoxide dismutase, glutathione (GSH), catalase and N-acetyl-cysteine were purchased from Sigma (St. Louis, Mo, U.S.A.). BCPS (Sigma), Benzyloxycarbonyl-Asp-Glu-Val-Asp (Ome)-fluoromethyl ketone (Z-DEVD-FMK) (Calbiochem, La Jolla, CA, U.S.A), Höechst 33258, 3,3′-dihexyloxacarbocyanine iodide (DiOC6(3)) and 2′, 7′-dichlorofluorescein diacetate (DCFH-DA) (Molecular Probes, Eugene, OR, U.S.A.) were dissolved in DMSO. [γ32P] ATP was obtained from Amersham Life Science Ltd. Antibody to JNK1, p38, Bcl-2, and PARP were purchased from Santa Cruz (Santa Cruz, California, U.S.A.). Double-stranded oligonucleotide probe containing the sequences of NF-κB and AP-1 as well as the T4 polynucleotide kinase were from Promega (Madison, WI). The sequences of the oligonucleotide were as follows: NF-κB, 5′-AGT TGA GGG GAC TTT CCC AGG C-3′, and AP-1, 5′CGC TTG ATG AGT CAG CCG GAA-3′. Poly(dI–dC) and protein GSepharode obtained from Pharmacia Biotech (Piscataway, NJ). 2.2. Cell cultures and cell viability assay The HL-60 cell line was obtained from American Type Culture Collection. The cells were grown in RPMI 1640 media (Falcon) supplemented with 10% heat-inactivated fetal calf serum (Falcon) and penicillin–streptomycin in a 5% CO2 incubator at 37 ± 0.5 °C. The cytotoxic effects of applied reagents were estimated after incubation with HL-60 cells for various times using the MTT test. The viability of HL-60 cells was measured by their ability to reduce the dye MTT to blue purple formazan crystal (Harada and Sugimoto, 1999). The reduced blue purple formazan was dissolved in glycine buffer containing DMSO for quantification by measuring O.D. at 570 nm using an ELISA Reader (Dynatech MR-7000), which was proportional to the viability of HL-60 cells. 2.3. Morphological features and DNA fragmentation of HL-60 cells HL-60 cells were incubated with applied reagents for various times. Photomicrographs were obtained from 40× objective lens using a cooled CCD camera OlymPix 50 2500) adapted to a Zeiss Axiovert 135-TV microscope. Cells were lysed in solution containing 5 mM Tris buffer (pH 7.5) 0.5% Triton X-100 and 20 mM EDTA. Supernatants were separated by centrifugation and incubated with the same lysis buffer containing 0.1% RNase A for 30 min at 37 °C. Lysate were then further treated with 1 mg/ml of protease K for 30 min at 37 °C. DNA extracted from a phenol chloroform precipitation was then resuspended in TE buffer (Tris buffer, pH 7.2, plus 1 mM EDTA and 0.5% SDS) and subjected to electrophoresis on 1.2% agarose gel containing 0.5 µg/ml of ethidium bromide for 2 h at 50 V.

After being stimulated according to experimental protocols, the cells (3 × 106) were centrifuged to collect the pellets and washed three times with 15 mM HEPES in 0.9% NaCl (w/v), pH 7.3, to minimize external contamination of copper. Cells were resuspended in the HEPES buffer. Cellular fractions or DNA were extracted using different lysis buffers (Fernandes and Cotter , 1994). An aliquot was removed for protein or DNA determination, and then the remaining part was resuspended in 0.1 N nitric acid for the analysis of copper. The standard solution was commercially available of 1000 ppm (1000 mg/l or L) for atomic absorption spectrophotometer. It was diluted to 10 ppm by adding 0.1 N nitric acid. The contents of Cu elements were measured on a model Z-8200 atomic absorption spectrophotometer (Hitachi, Tokyo) using a graphite furnace (flameless mode). 2.6. Measurement of mitochondrial membrane potential Mitochondrial membrane potential was indicated by retention of the cationic lipophilic fluorochrome 3, 3′-dihexyloxacarbocyanine DiOC6(3) (Decaudin et al., 1997). It is positively charged, the intracellular distribution of DiOC6(3) is determined by the electric potentials across various membranes and follows the Nernst equation. The higher the Δψm, the more dye is sequestered in mitochondrial matrix. After treatment with cytotoxic agents, the cells were collected and washed with PBS. Cells (1 × 106 cells in 500 µl of PBS) were loaded with 50 nM DiOC6(3) and incubated at 37 ± 0.5 °C for 15 min. Fluorescence intensity of the DiOC6(3) dye was determined by FACScan flow cytometry (excitation at 475 nm and emission at 525 nm). 2.7. Determination of non-protein thiols Total concentration of glutathione (GSH) was monitored by a modification of the GSH reductase method (Griffith, 1980). Cells were harvested and washed with ice-cold PBS and lysed with RIPA solution. An aliquot was removed and 100% trichloroacetic acid (TCA) was added to make a final concentration of 5%. The aliquot was centrifuged at 12,000 ×g for 10 min and reaction buffer containing 0.15 M imidazole (pH 7.4) was added to 50 µl of the supernatant. To the initiate the reaction, 5, 5-dithiobis (2-nitrobenzoic acid)-DTNB was added. Concentration of GSH was determined with a spectrophotometer at 412 nm, respective to the GSH standard. 2.8. Determination of free radical production The production of free radicals post exposure of the HL-60 cells to the PDTC/Cu complex was measured with flow cytometry (Sasada et al., 1996). Cells (1 × 106) were incubated for various lengths of time at 37 ± 0.5 °C with the Cu/NCP, in the presence of 30 µM of 2′, 7′dichlorofluorescein diacetate (DCFH-DA). Upon entering the cells,

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DCFH-DA deesterased and turned into a nonfluorescent polar derivative, 2′, 7′-dichlorofluorescein (DCFH). In the presence of H2O2 and other peroxides, this derivative became oxidized, forming a fluorescent compound, 2′, 7′-dichlorofluorescein (DCF) that emitted a fluorescent signal at 525 nm. A flow cytometer (Beckton Dickinson) was employed to measure cellular fluorescence intensity (excitation at 475 nm), which directly reflected the concentration of intracellular peroxides. 2.9. Measurement of JNK, p38 and IκB kinase activity Extracted cells were centrifuged to remove cellular debris, and the protein contents of the supernatants were determined by biocinchoninic acid (BCA) protein assay. JNK1, p38 and IκB kinase were immunoprecipitated and kinase activity was measured using an immunokinase complex assay with the substrates GST-c-Jun, GSTATF2 or GST-IκBα respectively (Ninomiya-Tsuji et al., 1999). Briefly, cell lysates (200 µg of protein) were incubated overnight at 4 °C with 10 µg of polyclonal anti-JNK1, anti-p38, and anti-IKK1 antibodies. Cell lysates were then incubated with 20 µl of Sepharose A-conjugated protein A for an additional 1 h. The beads were pelleted and washed three times with cold PBS containing 1% Nonidet P-40 and 2 mM sodium orthovanadate., once with cold 100 mM Tris–HCl (pH 7.5) buffer containing 0.5 M LiCl, and once with cold kinase reaction buffer (12.5 mM morpholinepropanesulfonic acid pH7.5, 12.5 mM βglycerophosphate, 7.5 mM MgCl2, 0.5 mM EGTA, 0.5 mM NaF, 0.5 mM sodium orthovanadate). The kinase reaction was performed in the presence of 1 µCi of [γ-32 P] ATP, 20 µM of ATP, 3.3 µM of DTT, 3 µg of the substrate GST-c-Jun-(1–135), GST-ATF2 and GST-IκBα in kinase reaction buffer for 30 min at 30 °C and stopped by addition of 10 µl of 5× Laemmli loading buffer. The samples were heated for 5 min at 95 °C and analyzed by SDS-PAGE (12% polyacrylamide). Phosphorylated substrates (GST-c-Jun, GST-ATF2 and GST-IκBα) were visualized by autoradiography. The optical density of autoradiogram was determined with the NIH Image program. The kinase activity was expressed as a fold of the control. 2.10. Electrophoretic mobility shift assay (EMSA) Cells were collected after treatment, and processed for nuclear extraction. Cells were resuspended in hypotonic lysis buffer and incubated on ice for 15 min, then centrifuged at 13,000 rpm for 1 min. The pellet was resuspended in high salt extraction buffer (20 mM HEPES, pH 7.9, 420 mM NaCl, 1.5 mM MgCl2, 0.2 mM EDTA, 25% V/V glycerol, 0.5 mM PMSF, 0.5 mM DTT, 1µg/ml PMSF, 1µg/ml aprotinin, 1µg/ml leupeptin) and incubated on ice for 15 min. The suspension was then centrifuged at 10,000 ×g for 20 min and the supernatant was collected. The protein contents of supernatants were determined using biocinchoninic acid protein assay. Nuclear NF-κB and AP-1 were assessed by electrophoretic mobility shift assay using doublestranded oligonucleotide probe containing the sequences of NF-κB and AP-1. The sequences of the oligonucleotide were as follows: NFκB, 5′-AGT TGA GGG GAC TTT CCC AGG C-3′, and AP-1, 5′-CGC TTG ATG AGT CAG CCG GAA-3′ containing the human κ-light chain enhancer motif, which had previously been end-labeled with [γ-32P] ATP as described (Mahon and O'Neill, 1995). Typically, 7.5 µg of nuclear extract protein was incubated with radiolabeled oligonucleotide (10,000 cpm) at room temperature for 30 min. NF-κB and AP-1 complexes were resolved in 5% acrylamide gels and identified following autoradiography. 2.11. Assessment of cytochrome c release Mitochondria and cytosolic fractions were prepared by resuspending cells in ice-cold buffer A (250 mM sucrose, 20 mM HEPES, 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, 1 mM EGTA, 1 mM DTT, 17 µg/ml

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phenylmethylsulfonyl fluoride, 8 µg/ml aprotinin, and 2 µg/ml leupeptin [pH 7.4]). Cells were passed through a 26G × 1/2” needle ten times. Unlysed cells and nuclei were pelleted by centrifuging at 750 ×g for 10 min. The supernatant was further spun at 100,000 × g for 15 min. This pellet was resuspended in buffer A and the supernatant represented the mitochondrial fraction. This supernatant was then centrifuged at 100,000 ×g for 1 h and the supernatant obtained from this step represented the cytosolic fraction. A 50 µg aliquot of protein from each sample was subjected to SDS-PAGE. After transferring to the nitrocellulose membrane, samples were probed with monoclonal anti-cytochrome c antibody coupled with HRP-mouse secondary antibody. Immunoreactivity was detected by the enhanced chemiluminescence detection system (NEN; Life Science Products). Immunoblots were quantified by densitometry (Bio-Rad GS-700 Imaging Densitometer equipped with Molecular Analysis software, version 2.1; Bio-Rad Laboratories Inc., Hercules, California, USA). 2.12. Caspase activity Cells were harvested and treated with the cytotoxic reagents. They were then washed with PBS and lysed in a solution containing 25 mM HEPES (pH 7.5), 5 mM MgCl2, 5 mM EDTA, 5 mM dithiothreitol (DTT), 2 mM PMSF, 10 µg/ml pepstatin A and 10 µg/ml leupeptin. The Promega CaspACE kit (Fluorometric Assay System; Madison, Wisconsin, U.S.A) was used to measure activity of caspases-1 and -3. Cells were lysed and centrifuged at 12,000 ×g for 5 min. Cell lysates containing 50 µg of protein were incubated with either 50 µM of AcDEVD-AMC (the substrate of caspase-3) or 50 µM of Ac-YVAD-AMC (the substrate of caspase-1) at 30 °C for 1 h. To measure caspase activity, levels of cleaved substrate were monitored using a spectrofluorometer (Hitachi F-4500) with excitation at 360 nm and emission at 460 nm. Caspase activity was expressed as a fold of the control. 2.13. Immunoblotting against JNK1, p38, IKK1, PARP, and Bcl-2 Equal amounts of lysate protein (50 µg/lane) were subjected to SDS-PAGE with 10% polyacrylamide gels and electrophoretically transferred to nitrocellulose membranes. Nitrocellulose blots were first blocked with 3% BSA in PBST buffer (PBS with 0.01% Tween 20, pH7.4) and incubated overnight at 4 °C with primary antibodies, (JNK1, P38, IKK1, PARP, and Bcl-2,) in PBST containing 1% BSA. After washing three times with PBST buffer, the mmunoreactivity was detected by sequential incubation with horseradish peroxidase-conjugated secondary antibodies, and detected by the enhanced chemiluminescence technique. 2.14. Statistic analysis of the data Data was expressed as mean values± standard error of mean (S.E.M.). Statistical analysis was carried out using one-way ANOVA followed by Dunnett's test to assess statistical significance (⁎P b 0.05) between treated and untreated groups in all the experiments. 3. Results 3.1. Effects of PDTC/Cu complex on the cell viability of HL-60 cells A MTT reduction assay was used to evaluate the cytotoxic effects of either native or complexed forms of PDTC and Cu on HL-60 cells. Our data shows that high concentrations of either PDTC or CuCl2 could induce cell death on HL-60 cells; EC50 were 100 ± 5.9 µM and 80±6.5 µM, respectively (Fig. 1A). In the serum-free medium however, 100 µM PDTC was almost nontoxic (9 ± 4.18% of control, n = 3) as compared with that in 10% serum containing cultured medium (49.5 ±5.9% of control, n = 4) on human HL-60 cells. As Cu may be essential for PDTC-induced cytotoxicity, we determined 0.11 ± 0.015 µM of Cu was present in the

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BCPS completely prevented the complex-induced cytotoxicity (Fig. 1C). In addition, we found that the antioxidant N-acetyl-cysteine and vitamin C (0.3–1 mM) but not vitamin E (1–20 µM) could also inhibit PDTC/Cu-induced cytotoxicity (Fig. 1C). 3.2. Morphological changes and DNA fragmentation PDTC (30–300 µM) or PDTC/Cu complex-induced dose-dependent changes to HL-60 cellular morphology (Fig. 2A). Non-treated cells exhibit typical non-adherent, fairly round morphology until 48 h of culture (Fig. 2Aa). After 6–24 h incubation with PDTC alone or PDTC/ Cu complex, some of the cells exhibited characteristics of apoptosis and numerous apoptotic bodies were observed. Apoptotic cells, as well as other intact cells, excluding trypan blue dye, suggested that the

Fig. 1. Concentration- and time-dependent cytotoxicity induced by PDTC/Cu complex and its prevention by antioxidants and BCPS in HL-60 cells. The viability of HL-60 cells was determined by MTT assay after incubation with PDTC, CuCl2 alone or combination without (A) or with 100 µM BCPS, 1 mM N-acetyl cystein, or 1 mM vitamin C (C) at 37 ± 0.5 °C for 24 h or for the other various times (B). Data are presented as mean ± S.E.M. (n = 5). ⁎: P b 0.05 as compared to the PDTC (A), vehicle (B) or treated with PDTC plus 1 µM CuCl2.

culture medium. Furthermore, we found that only Cu (0.3–3 µM) and not Fe or Pb (data not shown) significantly potentiated the cytotoxicity induced by low concentrations of PDTC (0.1–10 µM, Fig. 1A). The EC50 for 0.14 ± 0.02 µM, suggests that Cu (1 µM) potentiated the cytotoxic effect of PDTC by about 700-fold (Fig. 1A, B). The late marker of cell death lactate dehydrogenase (LDH) released to extracellular medium was also analyzed and the results were correlated with those of MTT assay. As compared to vehicle control PDTC (0.3 µM)/Cu (0.3 µM) resulted in 5-fold increase of LDH released in the extracellular medium after 24 hr incubation (P = 0.0006, n = 3). However, as the concentration of PDTC increased higher than 1 µM, the potentiating effect of PDTC for 1 µM of Cu sustained and slightly declined (Fig. 1A) Nevertheless, we showed that all of the compounds (100 µM CuCl2 alone, 100 µM PDTC alone, 0.1 µM PDTC+ 1 µM CuCl2 or 1 µM PDTC+ 1 µM CuCl2) induced cytotoxicity in time-dependent manner and reached a plateau phase in the incubation period of 24–36 h (Fig. 1B).

Fig. 2. Induction of morphological changes and DNA ladder by PDTC alone or PDTC/Cu complex in the HL-60 cells. (A) Cells were treated for 12 h with vehicle (a), PDTC 30 µM (b), PDTC 100 µM (c), PDTC 1 mM (d) or PDTC 1 µM plus Cu (1 µM) without (e) or with BCPS 100 µM (f). Note that BCPS prevents the morphological changes induced by PDTC/ Cu2+ complex examined under phase contrast microscope. Scale bar =10 µm. (B) Agarose gel electrophoresis of DNA fragments. HL-60 cells were treated with various concentrations of PDTC or PDTC/Cu complex for 6 h except that for 3-h (lane 5). Lane 1: vehicle, lane 2: PDTC 0.3 µM, lane 3: Cu 1 µM, lane 4: PDTC 100 µM, lane 5: PDTC 0.1 µM+ Cu 1 µM, lane 6: PDTC 0.3 µM+ Cu 1 µM, lane 7: BCPS + PDTC 0.3 µM+ Cu 1 µM. The results represent one of three independent experiments.

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cells were not in necrosis process. Surprisingly, PDTC at concentrations higher than 300 µM gradually reduced the cytotoxic effect on HL-60 cells (Fig. 2Aa–2Ae and data not shown). Furthermore, all of the morphological changes caused by PDTC/Cu complex were prevented by the addition of a non-permeable specific Cu chelator BCPS (Fig. 2Af). Fig. 2B showed that PDTC 100 µM (lane 4) and PDTC/Cu complex (lanes 5–6) induced DNA ladder in HL-60 cells in a time- and concentration-dependent manner which happened as early as 3 h (lane 5) while neither of vehicle control (lane 1), nor PDTC 0.3 µM (lane 2) nor CuCl2 1 µM (lane 3) showed DNA ladder. There is a decrease in DNA fragments induced by PDTC/Cu complex by BCPS application (lane 7). 3.3. The PDTC/Cu complex caused hypodiploidy in HL60 cells Subdiploid quantities of DNA were measured by staining with propidium iodide and analyzed by flow cytometry. While PDTC/Cu complex-induced hypodiploidy cells in time- and concentrationdependent manner (Fig. 3A, B, neither 0.3 µM of PDTC, nor 1 µM of Cu induced the hypodiploidy cells during 36 h incubation (Fig. 3A). The trend of this subdiploid effect corresponded with the cytotoxicity detected by MTT assay. Furthermore, PDTC (0.1–10 µM) with 1 µM of Cu-induced hypodiploidy DNA in a concentration-dependent manner

Fig. 4. Changes of intracellular Cu contents of HL-60 cells after treatment with PDTC/Cu complex. Cells were treated with PDTC and Cu either alone or complex at 37 ± 0.5 °C for 3 h (A) or those treated with 0.3 µM PDTC plus 1 µM CuCl2 for various times as indicated (B). The Cu contents of the subcellular fractions were assayed by atomic absorption spectrophotometer as described in Materials and methods. The Cu contents of control membrane, cytosol, and nuclear fractions are 2.54 ±0.39, 12.50 ± 1.60, and 1.19 ± 0.19 ng/mg protein respectively. Data are expressed as fold of control, mean± S.E.M. (n = 3). ⁎: P b 0.05 compared with respective control.

(Fig. 3B). This subdiploid effect was successfully inhibited by BCPS, N-acetyl-cysteine and vitamin C (Fig. 3B). 3.4. The PDTC/Cu increased complex on the concentration of intracellular Cu

Fig. 3. Time course of hypodiploidy cells induced by PDTC/Cu complex and its prevention by BCPS, N-acetyl cystein, and vitamin C in HL-60 cells. Cells were treated with PDTC/Cu complex for the indicated times without (A) or with 100 µM BCPS, 1 mM NAC and 1 mM vitamin C (B) for 24 h, and then analyzed DNA strand after staining with propidium iodide by the flow cytometric technique. Data are presented as mean ± S.E.M. (n = 3). ⁎: P b 0.05 compared with respective control.

While PDTC (1 µM) or Cu (1 µM) did not significantly alter the intracellular Cu contents, a higher concentration of 100 µM of PDTC or complex of PDTC (0.3–1 µM) plus Cu (1 µM) markedly increased the Cu contents in membrane, cytosolic, and nuclear fractions after 3 h incubation (Fig. 4A). Cu (1 µM) efficiently potentiated PDTC in transporting Cu into the cells. The kinetics of increasing Cu contents in time-dependent manner was similar in the subcellular fractions (Fig. 4B). The Cu contents in DNA fraction began to increase by 5.7 ± 0.8-fold within 30 min, and reach a maximum of 18.1 ± 2.4 at 1 h. The elevation profile of Cu contents closely correlated with the toxic effects detected by the MTT reduction test, hypodiploidy cells, DNA ladder and the apoptotic morphological changes. The non-permeable specific copper chelator BCPS 100 µM also abolished all of these effects of PDTC (0.1–10 µM)/Cu (0.3–3 µM) complex (Figs. 2Af, B, 3B and 4A). These findings indicate that PDTC/Cu complex penetrated into cells

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and dramatically increased the Cu contents in the cytosolic and nuclear fractions, thus inducing cytotoxicity. 3.5. PDTC/Cu complex increased the production of reactive oxygen species, reduced the mitochondrial membrane potential, and decreased the concentration of GSH contents PDTC/Cu complex increased free radical production in concentration- and time-dependent manners with a rapid onset (within 1 h) and reached a peak at 3–6 h (Fig. 5A). We also found that PDTC (0.3 µM)/Cu (1 µM) complex decreased non-protein thiols to 74 ± 6.4% of control within 30 min and reached 31 ± 4.8% at 6 h (Fig. 5C). We monitored the effects of PDTC/Cu complex on the mitochondrial transmembrane potential (Δψm) using fluorescent probe DiOC6(3) coupled with flow cytometry analysis. The fluorescent dye DiOC6(3) was localized to mitochondria, and the reduced accumulation of

Fig. 6. PDTC/Cu complex-induced JNK kinase activation in time-dependent manner and bathocuproine disulfonate (BCPS) reverses the activation. Cells were treated with PDTC/ Cu complex for various times. The kinase reaction was performed by the procedures as described in Materials and methods. The top panels represent the JNK phosphorylation shown as autoradiogram of [γ32P] ATP incorporation into exogenous GST-c-Jun (1–135). The bottom panel is Western blots performed with antibody specific to either JNK1 protein. The fold induction in this figure is presented as the ratio of JNK activity to JNK1 protein against time. Data are presented as mean ± S.E.M. (n = 3). The results represent one of three independent experiments.

DiOC6(3) reflected the loss of mitochondrial transition potential (Vavssiäre et al., 1994). Treatment of HL-60 cells with PDTC (0.1 µM)/Cu (1 µM) complex produced a 33.6 ± 5.5% decline in the mitochondria membrane potential within 1 h. The onset and extent of decreasing mitochondrial membrane potential was dependent upon PDTC concentration (0.03–0.1 µM,). BCPS also abolished this effect of the PDTC/Cu complex (Fig. 5B). 3.6. PDTC/Cu complex differentially activate the JNK1and P38 kinase To examine our working hypothesis that JNK activation may be involved in cell death induced by PDTC/Cu complex, we assessed the effects of PDTC/Cu complex on JNK activity by detecting the level of JNK protein expression and the extent of phosphorylation of its substrate, c-Jun. While the kinase activity was significantly increased by 3-fold at 1 h after treatment, reached a plateau by 19.5-fold at 6 h (Fig. 6), and sustained activation till 12 h at least, JNK1 protein remained unaltered after treatment with PDTC/Cu complex. After treatment of PDTC/Cu complex, p38 kinase was slightly activated at both 6 h without degradation and 12 h with partial degradation of p38 protein and cell cytotoxicity (data not shown). The non-cell-permeable specific Cu chelating agent-BCPS (Fig. 6) completely prevented the activation of JNK kinase. 3.7. Activation of NF-κB and AP-1

Fig. 5. PDTC/Cu complex reduced the mitochondrial membrane potential, increased free radicals and decreased non-protein thiols in the HL-60 cells. Cells were treated with PDTC/Cu complex for various intervals and then an aliquot of the cells was labeled with the fluorescent dye either DiOC6 (3) (A) or DCFH-DA (B) for analyzing the fluorescent intensity by flow cytometry as described in Materials and methods. The fluorescent intensity produced by retention in mitochondrial membrane or free radicals of the control cells was defined as 100%. a. control, b. BCPS 100 µM plus PDTC 0.1 µM and CuCl2 1 µM, c. PDTC 0.1 µM plus CuCl2 1 µM. Another portion of cells was used for measuring the reduced glutathione (GSH) content as described in Materials and methods (C). The GSH content of control cells (16.2 ± 1.8 nmol GSH/mg protein) was defined as 100%. Data are presented as mean ± S.E. M. (n = 3). ⁎: P b 0.05 as compared with the respective control.

As both NF-κB and AP-1 have been implied in triggering apoptosis, we determined whether PDTC/Cu complex activated NF-κB and AP-1 in HL60 cells. We performed EMSA's analysis to examine the extent of DNA binding of the nuclear extract isolated from HL-60 cells to the consensus sequence of either NF-κB or AP-1. Our results indicate that the DNA binding activity of NF-κB in the nuclear fraction of HL-60 cells was about 2-fold increased after 2–6 h incubation of PDTC/Cu complex (Fig. 7A). AP-1 DNA binding activity in the HL-60 cells treated with the PDTC alone or PDTC/Cu complex for 2–6 h was also increased by about two fold, and then restored to normal level in 24 h (Fig. 7B). No appreciable binding of nuclear extracts to the AP-1 consensus was detected in the absence of nuclear extracts or in the presence of 100-fold molar excess of unlabeled oligonucleotide probe (Fig. 7B lanes 9 and 10). 3.8. PDTC/Cu complex stimulated cytochrome c release and IκB activation in a time-dependent manner The process of cell death may involve the release of cytochrome c from mitochondria, which subsequently activates caspase and

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ultimately results in apoptosis. PDTC/Cu complex caused cytochrome c to release into the cytosol as fast as 1 h after treatment on HL-60 cells (Fig. 8A). The result was consistent with the decrease of mitochondrial membrane potential. In addition, PDTC/Cu complex were found to be capable of activating NF-κB. This finding is in contrast to the conventional inhibitory effect of PDTC on NFκB. We further determined the IKK1 activity of HL-60. PDTC/Cu complex-induced IκB kinase activation, and while IKK1 protein remained unaffected. IκB kinase was activated within 30 min, reaching a maximal level (3 fold) at 1 h prior to the NF-κB activation. BCPS also inhibited the IKK1 activity induced by PDTC/Cu complex (Figs. 7A, 8B). This finding further provides evidence that PDTC/Cu complex can activate the transcription factor NF-κB through increasing IKK1 activity. 3.9. PDTC/Cu complex stimulated caspase 3 Activity in a time-dependent manner and induced PARP degradation Since caspases are believed to play a central role in mediating various apoptotic responses, we have detected the enzymatic activity of caspase 3 during the induction of PDTC/Cu complex mediated apoptosis. The specific substrate of caspase 3, fluorogenic peptide

Fig. 8. BCPS inhibited cytochrome c release and IKK1 activation induced by PDTC/Cu complex in HL-60 cells. Cells were treated with PDTC/Cu complex for various time and then harvested. (A) Cytochrome c was released from mitochondria into the cytosol by PDTC/Cu complex in a time-dependent manner. (B) IKK1 activity in the cytosol fraction was analyzed by immune complex kinase assays as described in Materials and methods. IKK1 was immunoprecipitated with the anti-IKK1 antibody, and then the activity in the immune complex was assayed by using GST-IκBα (1–317) as a substrate. The results represent one of three independent experiments.

substrate (Ac-DEVD-AMC) was used in this experiment. As shown on Fig. 9A, addition of the PDTC/Cu complex caused a 14-fold induction in enzymatic activity of caspase 3 in HL-60 cells while neither the

Fig. 7. Representative autoradiogram of EMSA showing changes in DNA binding activity of NF-κB and AP-1 induced by PDTC alone or PDTC/Cu complex in HL-60 cells. Cells were treated with indicated concentration of PDTC or PDTC/Cu complex for the indicated times. Nuclear extract obtained after stimulation were used to measure NF-κB (A) and AP-1 (B) DNA binding activity by EMSA using a 32P-labeled oligonucleotide probes containing consensus sequence of NF-κB or AP-1. In competition experiments (lane 10), the unlabeled oligonucleotide at 100-fold molar excess was added to the binding reaction mixture. Reactions were carried out with or without (lane 9) 7.5 µg of nuclear extract, 1 µg labeled oligonucleotide, and 1 µg of poly(dI–dC) for 20 min at 30 °C. Protein–DNA complexes were resolved by electrophoresis on 5% polyacrylamide gels and visualized by autoradiography. Similar results were obtained in other two separate experiments.

Fig. 9. PDTC/Cu complex activated caspase 3 and cleaved PARP in HL-60 cells and its blockade by BCPS. Cells were treated with PDTC (0.1, 0.3, or 1 µM) and Cu (1 µM) complex for various times as indicated, and then caspase 3 activities (A) were analyzed as described in Materials and methods. Data are presented as mean ± S.E.M. (n = 3). ⁎: P b 0.05 as compared with respective control. The cleavage of PARP from 116 kDa to 85 kDa was assayed by Western blots (B) performed with antibody specific to anti-PARP (left). Bcl-2 was determined with its specific antibody (right, as negative control). The amount of cell lysates used was 50 µg of protein in each lane. Similar results were obtained in the other two separate experiments.

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vehicle, PDTC 0.3 µM nor Cu 1 µM had such effect on caspase 3 activity. By contrast, the enzymatic activity of caspase 1 was scarcely activated by the PDTC/Cu complex. BCPS also abolished this activation of caspase 3 by PDTC/Cu complex (Fig. 9A). A number of ICE/CED-3 protease targets have been identified, including the nuclear enzyme PARP. In many cellular systems undergoing apoptosis, the endogenous PARP 116-kDa protein, is cleaved to generate an 85-kDa fragment. Therefore, we evaluated the changes of the PARP protein and our results show that PDTC/Cu complex causes the degradation of 116-kDa PARP into 85-kDa fragment (Fig. 8B). In addition, we also found that BCPS effectively abolished the degradation of PARP by PDTC/Cu complex (Fig. 9B). 4. Discussion In this study, we examined the molecular mechanisms of cytotoxic effects induced by a PDTC/Cu complex in HL-60 cells. Our results indicated that PDTC/Cu complex increased intracellular Cu levels and reduced the mitochondrial membrane potential, generation of free radical generation and depletion of non-protein thiols. In addition, activation of JNK, NF-κB, AP-1 and caspase 3 signaling pathways was observed prior to PARP degradation and cell apoptosis. Although dithiocarbamates can chelate various metals, resulting in the formation of a lipophilic dithiocarbamate–metal complexes (Thorn and Ludwig, 1962), our data suggested that only Cu but not Fe, Pb could potentiate the toxicity of PDTC in HL-60 cells. This finding indicates that PDTC was more selective to chelate Cu than other metals. As Cu is an essential metal we tested to see whether physiological concentrations (0.3–3 µM) could potentiate the toxic effects of PDTC. The results obtained indicated that Cu can potentiate PDTC-toxicity. Consequently, these findings suggest that exposure to PDTC in medicinal or agricultural environments may be hazardous. PDTC (0.1–10 µM) combined with a fixed concentration of Cu 1 µM (0.06 ppm) induced cytotoxic effects on HL-60 cells in a concentrationand time-dependent manner. The non-permeable specific copper chelator BCPS, at a concentration of 100 µM, blocked all of the effects of PDTC/Cu complex (Figs. 2Af, B, 3B and 4A), suggesting an essential role of Cu involved in PDTC-induced cytotoxicity. In other studies, it has been shown that a ratio of PDTC/Cu in the complex form below 1.0 is crucial for high potency of inducing cytotoxicity. This observation is further supported by experiments conducted on thymocytes (Nobel et al., 1995) and cortical astrocytes (Chen et al., 2000). Measurement of intracellular Cu contents indicated that PDTC increased the transport and uptake of Cu into HL-60 cells. Although PDTC may chelate Cu with its thiol groups to form complex that enter the cells, whether the chemical form acted as the PDTC/Cu complex form or as the dissociated free Cu is still unclear. We propose that the redox state of Cu increases free radical production and subsequently reduces the mitochondrial membrane potential. In contrast, when the PDTC concentration was over 1 mM (Fig. 2Ad), the cytotoxicity was remarkably reduced, perhaps due to the formation of larger nonpermeable complex with Cu in medium. Mitochondria play a crucial role in inducing apoptosis (Mahon and O'Neill, 1995; Nieminen et al., 1995) as membrane depolarization is an early irreversible event during apoptosis (Zamzami et al., 1995). Here, we showed that PDTC/Cu complex significantly decreased the mitochondrial membrane potential, stimulated the release of cytochrome c and free radicals while caused non-protein thiols depletion. While the phenomena of depleted non-protein thiols occurred more rapidly than previously reported (Nobel et al., 1995), we could not exclude the possibility that the decreased content of non-protein thiols reflected decreased synthesis and not degradation. Evidently, the oxidative stress produced by PDTC-Cu is in contrast to the antioxidant property of PDTC. One might speculate that higher concentrations of PDTC, Cu or PDTC/Cu complex may induce apoptosis

at the early stage with lower oxidative stress and then produce an irreversible necrotic lysis at later stage. Consistent with this hypothesis, our data showed that both N-acetyl cystein and vitamin C could partially inhibit the cytotoxicity induced by the PDTC/Cu. The onset of GSH reduction corresponded to PDTC concentration. Since both the production of free radicals and the depletion of non-protein thiols occurred prior to the morphological changes and cytotoxicity, we concluded that the mechanism of PDTC/Cu complex-induced cytotoxic effects probably mediated these processes. As the activation of the JNK pathway has also been implicated in the cell death-signaling pathway in response to several stimuli (Zhang et al., 1996; Ninomiya-Tsuji et al., 1999; Verheij et al., 1996), we sought to determine whether the PDTC/Cu complex affected the JNK pathway. Our results showed that PDTC/Cu complex caused activation of JNK and caspase 3, and degradation of PARP, which preceded the onset of apoptosis. Furthermore, the PDTC/Cu complex-induced JNK activation as early as 3 h which persisted for more than 12 h. This elevation of JNK activity followed a transient increase of ROS production, and the cytotoxicity induced by PDTC/Cu, suggesting a hit and run phenomenon. The importance of Cu for PDTC/Cu mediated apoptosis was demonstrated through the use of a selective Cu chelator BCPS. BCPS chelated the extracellular Cu and inhibited both JNK activation and apoptosis. The antioxidant vitamin C could also block JNK activation and cytotoxicity induced by PDTC/Cu complex. Therefore, we suggest that JNK activation occurs following free radical production as previously described (Chen et al., 2000; Xia et al., 1995). However, whether the activation of p38 kinase (data not shown) plays a role in PDTC/Cu mediated apoptosis is still unclear. Although PDTC is an inhibitor of NF-κB activation in various cell types (Nobel et al., 1997) that ultimately results in the inhibition of apoptosis (Nobel et al., 1997; Sherman et al., 1993; Boland et al., 1997), PDTC is also a pro-oxidant capable of inducing cell death in PC12 cells. In order to understand the relation between NF-κB activation and PDTC/Cu complex-induced apoptosis, we examined whether PDTC alone or PDTC/Cu complex could activate NF-κB in HL-60 cells. Our results showed that both of NF-κB and AP-1 were activated which preceded IκB kinase activation. While the activation of NF-κB is consistent experiments conducted on smooth muscle cells (Erl et al., 2000) but not RAW cells (Sohn et al., 2005) treated with PDTC alone. Furthermore, as NF-κB has a role in inducing apoptosis by daunorubicin (Boland et al., 1997), we considered whether the observed activation of NF-κB might be also involved in the PDTC/Cu complexinduced apoptosis. The IκB kinase activation induced by PDTC/Cu complex in HL-60 cells is consistent with hippocampal progenitor cells in response to PDTC (Min et al., 2003). The downstream of JNK activation signaling could be activation of Bid which translocated to the mitochondria and induced cytochrome c release and apoptosis (Luo et al., 1998), or directly activated caspase 3, or mediated by AP1 activation and caspase 8 activation (Lauricella et al., 2006). AP-1 activation induced by PDTC/Cu complex was first demonstrated in HL-60 cells in this paper. Although AP-1 is known to play a role in cell proliferation, there are an increasing number of studies that demonstrate that AP-1 is also involved in apoptosis in other cell types (Domanska-Janik et al., 1999). In addition, it has been shown that AP-1 is induced by cyclosporin A (Pyrzynska et al., 2000) and that AP-1 activation induced by PDTC is attenuated in JNK1 deficient-human umbilical vein endothelial cells (Liao et al., 2000). Consequently, we suggest that the activation of AP-1 may be downstream to JNK activation induced by PDTC/Cu complex in HL-60 cells. Intriguingly, in contrast to normal cells, high levels of copper have been found in cells extracted from leukemia, breast, prostate and colon cancer (Kuo et al., 2002; Nayak et al., 2003; Zuo et al., 2006). Consequently, PDTC may form more active complexes with copper in these tumor tissues rather than normal cells and thereby specifically inhibit angiogenesis in tumor cells. PDTC might therefore be a useful anti-cancer drug and even more potent than D-penicillamine (Gupte and Mumper, 2007).

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