Chem. Res. Toxicol. 2003, 16, 113-122
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Articles Mechanism of Ascididemin-Induced Cytotoxicity Sandra S. Matsumoto,† Jason Biggs,† Brent R. Copp,‡ Joseph A. Holden,§ and Louis R. Barrows*,† Department of Pharmacology and Toxicology, University of Utah, Salt Lake City, Utah 84112, Department of Pathology, University of Utah, Salt Lake City, Utah 84112, and Department of Chemistry, University of Auckland, Auckland, New Zealand Received September 16, 2002
Some marine animals are rich sources of unique polycyclic aromatic alkaloids that are cytotoxic against tumor cell lines and effective in mouse tumor xenograft models. Ascididemin is a pyridoacridine alkaloid originally derived from a Didemnum sp. tunicate. It has potent cytotoxicity against tumor cells in vitro and in vivo. Preclinical screening at NCI revealed the antineoplastic activities of ascididemin and a synthetic analogue 48. Ascididemin has been reported to inhibit topoisomerase II and induce topoisomerase II-mediated DNA cleavage. This study, however, focuses on the unique ability of ascididemin and two synthetic analogues (48 and 109) to cleave DNA in the absence of topoisomerase I or II. An in vitro assay revealed their concentration-dependent ability to cleave DNA and identified dithiothreitol as the sole requirement for maximal activity. On the basis of shared structural features of the three analogues, a double N-bay region and iminoquinone heterocyclic ring, two possible mechanisms of action were hypothesized: (1) generation of reactive oxygen species facilitated by metal binding to the common phenanthroline bay region, and (2) production of reactive oxygen species by direct reduction of the iminoquinone moiety. Experimental results supported direct iminoquinone reduction and ROS generation as the mechanism of ascididemin cytotoxicity. Antioxidants protected against DNA cleavage in vitro and protected cultured Chinese hamster ovary cells from toxicity. Additionally, it was shown that cells deficient in the ability to repair reactive oxygen species damage to their DNA were more susceptible to ascididemin and analogues than repair competent cells. Ascididemin-treated cells were also shown to induce oxygen-stress related proteins, further implicating the production of reactive oxygen species as the mechanism of cytotoxicity for these molecules.
Introduction Several marine alkaloids have been recognized as potential antineoplastic agents because of their potent cytotoxicity, structural diversity, and mechanistic complexity. In 1983, amphimedine was the first pyridoacridine alkaloid reported, isolated from an Amphimedon sp. sponge collected near Guam (10). Pyridoacridines are marine alkaloids based on the 11H-pyrido[4,3,2-mn]acridine skeleton. Since that initial discovery, pyridoacridine alkaloids from sponges and ascidians have become an established class of molecules with significant biological activity including antifungal, antimicrobial, cytotoxic, and DNA binding properties (11). Ascididemin (51; Figure 1) has been shown to be cytotoxic to a number of tumor cell lines in vitro and to inhibit topoisomerase II [top 21 (1, 9, 12, 13)]. Ascididemin was cytotoxic at submicromolar levels to mouse leukemia (P388), human colon (HCT 116), and human breast (MCF-7) tumor cell lines, and showed enhanced cytotox* To whom correspondence should be addressed. Phone: (801) 5814547. Fax: (801) 585-5111. E-mail:
[email protected]. † Department of Pharmacology and Toxicology, University of Utah. ‡ University of Auckland. § Department of Pathology, University of Utah.
icity against mutant CHO (Chinese hamster ovary) cell lines that were deficient in either single- or double-strand break repair (14). Bonnard and colleagues (15) evaluated the biological activity of ascididemin and reported (1) only minor effects on top 2 catalytic activity; (2) marked cytotoxicity to human leukemia cells; and (3) DNA intercalation at GCrich sequences. From their findings, Bonnard et al. concluded that DNA, not topoisomerases, is the likely target of ascididemin cytotoxicity. More recently, this group has reported the potent ability of ascididemin to induce apoptosis in cultured cells (16). Our study provides further evidence of the molecular mechanism by which ascididemin targets cellular DNA. Reactive oxygen species (ROS), although produced during normal aerobic metabolism, have been implicated 1 Abbreviations: top 2, topoisomerase II; ROS, reactive oxygen species; BA, benzoic acid; BHA, butylated hydroxyanisole; SOD, superoxide dismutase; NAC, N-acetylcysteine; 109, BC-109-1, 7Hpyrido[4,3,2-de][1,10]phenanthrolin-7-one (1); 51, ascididemin or asc; 48, BCMH-1-48, 1,10-phenanthroline-5,6-dione (1); BC-1-21, 21, sampangine (2); BC-1-31, 31, 9H-benzo[b]pyrido[4,3,2-mn]acridin-9-one (3); BC1 29, 3-bromoascididemin (unpublished); BC1 55.1, diplamine (4); BC1 55.2, varamine A (5); NP164, kuanoniamine A (6); BC1 14.2, kuanoniamine D (7); BC1 14.3, shermilamine B (8); BC1 55.3 (9).
10.1021/tx025618w CCC: $25.00 © 2003 American Chemical Society Published on Web 01/28/2003
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Figure 1. Structures of ascididemin, 109, 48, 31, and 21.
in cell and tissue damage (17). Free radicals produced by oxidative processes can attack DNA at bases or sugars, causing primarily single-, but secondarily, double-strand breaks as well as abasic sites (18). Quinone functions are capable of redox cycling. One electron reduction of the quinone forms reactive semiquinones, allowing electron transfer to other species (19). Higher redox potential of quinone molecules has been correlated with increased DNA cleavage. Moreover, evidence suggests that doxorubicin and daunorubicin, which contain quinone moieties, owe some of their clinical activity to ROS generation mechanisms (20). This study reports on the ability of ascididemin, and two structurally related synthetic alkaloids 109 and 48, to generate ROS and damage DNA independently of enzymatic catalysts. The two common structural motifs shared by these molecules are a double nitrogen bay region and a reducible, iminoquinone heterocyclic ring (Figure 1). Structure-activity analysis showed that both features were required for DNA cleaving activity. The observation that DTT greatly stimulated drug-DNA cleavage in vitro, promoted the hypothesis that cleavage stimulation was accomplished via an ROS mechanism. Further, it was hypothesized that either a metalcatalyzed reaction or direct reduction of the pyridoacridine iminoquinone, generated the ROS responsible for the observed DNA damage. The Fenton-like mechanism would require metal binding to the common phenanthroline double N-bay region and metal-facilitated redox cycling to generate ROS, while direct-reduction would involve reduction of the iminoquinone pharmacophore to produce ROS. The results presented here support the latter (direct) mechanism.
Materials and Methods Chemicals and Reagents. Ascididemin was synthesized according to published procedures (6). Additional synthetic schemes have recently been published (21, 22). The synthetic analogues 109 and 48 (1), as well as 14 additional analogues, were synthesized or isolated as pure compounds. Analogues 21 (2) and 31 (3) served as controls for the bay nitrogen moieties of 109 and ascididemin, respectively. Chemical purity was verified by chromatography and NMR. All other chemicals were purchased from Sigma Chemical Co., St. Louis, MO, or Baker Chemical Co., Springfield, NJ. Radiolabeled (4.4 × 103 cpm/mg) 3H replicative form (rf) of M13 mp 19 plasmid DNA was isolated by the alkaline lysis method as described by Holden et al. (23). Human top 2R was also isolated as described by Holden et al. (23).
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Figure 2. Quantitative DNA cleavage gel of etoposide, ascididemin, 21, 31, 48, and 109 in the presence of top 2. Lane C1, DNA only; lane C2, DNA + top 2; lane 1, DNA + top 2 + 1% DMSO; lane 2, DNA + etoposide; lane 3, DNA + etoposide + top 2; lane 4, DNA + 21 + top 2; lane 5, DNA + 31 + top 2; lane 6, DNA + 48 + top 2; lane 7, DNA + ascididemin + top 2; lane 8, DNA + 109 + top 2. Lane 3 shows cleavage by etoposide, a prototypic top 2 poison. Top 2 concentrations were 44 ng/ reaction. All drug concentrations in the reactions were 91 µM. Compounds 109, 48, and ascididemin strongly stimulate DNA cleavage above controls; 21 and 31 did not. Abbreviations: n ) nicked, l ) linear, sc ) super coiled, and r ) relaxed plasmid DNA, respectively. Quantitation of in Vitro DNA Cleavage. DNA cleavage assays were performed in 20 µL volumes of 50 mM Tris-HCl buffer (pH 7.5), containing 100 mM KCl, 10 mM MgCl2, 0.5 mM EDTA, 0.2 mg/mL bovine serum albumin, 500 µM ATP, 5 mM DTT, 500 ng of radiolabeled (3H thymidine), and supercoiled rf M13 mp 19 DNA in reactions containing top 2 (20-44 ng). In the absence of topoisomerase, reactions contained ascididemin or analogues at concentrations of 100 µM in H2O and 500 ng of radiolabeled mp 19 DNA ([3H]thymidine) both with and without 5 mM DTT. All reactions were incubated for 30 min at 30 °C. Reactions containing top 2 were treated with 0.5% SDS and 1 mg/mL proteinase K for an additional 45 min at 37 °C to convert drug-topoisomerase-DNA cleavage complexes to nicked or cut DNA prior to electrophoresis (24) (Figure 2). The reactions were stopped and DNA topological isoforms fractionated by electrophoresis in a 0.8% agarose gel containing 50 ng/mL ethidium bromide. Nicked, cut, relaxed, and supercoiled DNA isoforms were visualized under UV light and excised from the gel. Gel slices were placed in 20 mL scintillation vials containing 1 mL of H2O, melted in a microwave then mixed with 10 mL of OptiFluor (Packer, Meridien, CT) while molten. Radioactivity was determined by standard scintillation counting (23). The fractions of total radioactivity from endogenously nicked DNA were subtracted from radioactivity detected in treated samples to normalize cleavage stimulated by test compounds. Percent DNA-cleavage was calculated as (nicked - endogenous nicked)/ (supercoiled + relaxed) × 100. Each experiment was repeated at least three times (n g 3). Time and Concentration Dependence of DNA Cleavage. To determine the concentration dependency of DNA cleavage in vitro the DNA cleavage assay described above was performed using increasing concentrations of ascididemin, 109, 48, and the inactive analogues 21 and 31. Final analogue concentrations were 10 µM, 50 µM, 100 µM, 500 µM, and 1 mM. It is customary to use micromolar to millimolar concentrations of drug in such assays in order to provide sufficient DNA damage for purposes of visualizing cleavage products in an agarose gel. Mechanisms identified in this manner reflect actions in the cell, where the lesser cleavage accomplished by lower drug concentrations is sufficient to induce cytotoxicity (25). To determine time dependence, each reaction was performed using 100 µM concentration of drug, but with varied times of incubation including 5, 10, 15, 20, 25, 30, 45, and 60 min. Effects of Metals, Radical Scavengers, and Chemoprotective Agents. The quantitative DNA cleavage assay described above was repeated for ascididemin, 109, 48, 21, and 31, but with the addition of either (1) metal salts (i.e., FeSO4, NiCl2, CuSO4, MgCl2, and MnCl2); (2) metal ion chelators (EDTA and desferoxamine); (3) antioxidant enzymes (i.e., SOD and catalase); and (4) radical scavengers [glutathione, BHA and
Mechanism of Ascididemin Cytotoxicity BHT, ascorbic acid and N-acetylcysteine (NAC)] in order to further define the potential mechanisms of DNA cleavage. The effects of hydroxyl radical scavengers, ethanol, salicyclic acid, benzoic acid and DMSO, on ascididemin and analogue induced cleavage were also determined. Assessment of DNA Cleavage under Anoxic Conditions. Quantitative DNA cleavage reactions were carried out as described above; however, the reactions were performed under anoxic conditions based on the method described by Radisky et al. (26). To ensure anoxic conditions, each component (H2O, DTT, drug, DNA) of the reaction was degassed for 5 min under vacuum followed by 5 min under bubbling argon three times. The reactions were mixed under argon, capped, and incubated at 30 °C for 30 min before being quantified as above. Determination of Reduction-Oxidation Potential. Cyclic voltammetry was conducted using methods of Kohen (27) and Radisky et al. (26). Briefly, a flow of electrons was introduced to a voltammetric cell containing 100 µM drug in H2O. The smooth platinum working electrode repetitively scanned the reaction at a speed of 2 V/s with a potential scan range from -0.4 to 2.2 V. Hyperpolarized current corresponded to oxidative reactions. Ag/AgCl served as the reference electrode. Electron Paramagnetic Resonance (EPR) Spectroscopy. EPR spectra were obtained using a Bruker model EMX EPR (Billerica, MA) spectrometer. Reactions were made up in 280 µL total volume and analyzed at room temperature. Each tube contained H2O as solvent plus one or more of the following reagents: (1) 100-500 µM drug; (2) 5-10 mM DTT; (3) 100 µM FeSO4; (4) 100 µM H2O2; and (5) 100 µM N-t-butyl-R-phenylnitrone (PBN). Immediately before EPR analysis, reactions were added individually to quartz EPR flat cells as suggested for aqueous solutions (28). Electromagnetic radiation was applied to the sample and parameters (i.e., receiver gain, time constant, sweep time, modulation amplitude, modulation frequency) were optimized for each reaction. Because free radicals formed in such reactions have very short half-lives, PBN was employed as a spin trap for ‚OH and H2O2. The Fenton reaction of H2O2 with 100 µM ferrous sulfate (29.4 mM), in the presence of 50 mM PBN, served as the positive control. Assessment of DNA Intercalation. DNA intercalation was measured as the ability of drug to displace ethidium bromide from DNA. On the basis of the protocol of McDonald et al. (29), 10 serial dilutions (1 nM-70 µM) of ascididemin, 109, 48, 21, or 31 were reacted with 0.5 µM ethidium bromide and 0.5 µM salmon testes DNA in a 96-well microtiter plate. Samples were allowed to react for 30 min at room temperature. A fluorimeter was used to quantify the fluorescence of each reaction (530 nm excitation wavelength; displacement of ethidium bromide was measured as a decreased fluorescence at 600 nm). Cell Culture. Chinese hamster ovary (CHO) cell lines were used for many of the in vitro assays. Particularly informative were the wild-type AA8 cell line, the single strand break repairdeficient EM9 cell line, and the double strand break repairdeficient xrs-6 cell line. In addition, the human colon carcinoma HCT 116 and the human nasopharyngeal carcinoma KB cell lines were used. AA8, EM9, KB, and HCT 116 cells were purchased from ATCC (Rockville, MD). The xrs-6 cell line was a generous gift from Dr. P. Jeggo and co-workers (ref 30; University of Sussex, Brighton, U.K.). AA8, EM9, and xrs-6 cells were maintained in R-Minimal Essential Medium (R-MEM) supplemented with 10% fetal bovine serum (Atlanta Biologicals, Atlanta, GA), 100 units/mL penicillin, and 100 µg/mL streptomycin. HCT 116 cells were maintained in McCoy’s medium similarly enriched with 2% fetal bovine serum, 8% newborn bovine serum, 100 units/mL penicillin, 100 µg/mL streptomycin, and 240 units/mL nystatin. All cells were maintained at 37 °C in a humidified 5% CO2 atmosphere. Cells were grown as monolayers in 75-cm2 culture flasks and detached using trypsin prior to drug treatment. Cytotoxicity Assays. Ascididemin and analogues were tested for cytotoxicity using the MTT-microtiter plate tetrazolium cytotoxicity assay (1, 31). Cells were plated in 96-well
Chem. Res. Toxicol., Vol. 16, No. 2, 2003 115 microtiter plates at 20 000 cells/well (AA8, EM9), 20 000 cells/ well (HCT 116, KB), or 40 000 cells/well (xrs-6) in 200 µL growth medium and allowed to adhere to the wells for 4 h. For treatment of CHO cells, 10 serial dilutions of drug (8.6 nM to 86 µM final concentration) dissolved in 100% DMSO (1% final concentration) were added to the wells and cells incubated overnight at 37 °C. Four replicate wells of each drug dose, as well as eight replicate control wells containing an equivalent volume of DMSO only, were included in each plate. Cells were exposed to drug for 18 h before the media were removed and the cells re-fed. KB and HCT cells were treated with half the volume of drug as the CHO cells without removing drug or refeeding. Medium was aspirated from the wells after 72 h and 100 µL of McCoy’s medium with 11 µL of MTT was added to each well. The plates were then incubated for 4 h at 37 °C. Medium was aspirated, and 100 µL of DMSO was added to each well. A Bio-Rad MP450 plate reader (Bio-Rad Laboratories, Hercules, CA) was used to measure the absorbance (540 nm) of each well. The average absorbance for the drug-treated cells was divided by that of control cells to determine fractional survival at each concentration. In the glutathione (GSH) protection assays, cells were plated as described above but pretreated with GSH following adhesion and incubated an additional 2 h prior to drug treatment. All experiments were performed at least twice (n g 2). Determination of Intracellular Glutathione Concentration. Total intracellular glutathione was measured using a modification of the methods of Tietze (32). Approximately 2.5 × 105 AA8 cells were plated on a 6-well culture plate and grown to confluence in R-MEM. Treatment groups included drugtreated cells with or without 400 µM GSH. Control groups included untreated cultures and cells treated with 2 mM diethylmaleate (DEM). Cells were treated with GSH for 2 h before addition of other drugs. All replicates were carried out in 4 mL medium for 4 h. Medium was then aspirated, and the cells were washed three times with 200 µL of 1× phosphate buffered saline (PBS), pH 7.5. Cells were transferred to 15 mL conical tubes, counted with a coulter counter, and centrifuged at 5000g for 5 min. Supernatants were discarded and cell pellets resuspended in 1× PBS. Cells were lysed by 10 s sonication and centrifuged at 10 000 rpm for 10 min to pellet debris. Supernatants were then stored at -80 °C for later analyses. All analyses employed a modification of the Lin and Miller method (33). All reagents for the assay were prepared in 1× PBS (pH 7.5) containing 5 mM EDTA, DTNB (236 ng/µL), reduced NADPH (170 ng/µL), and baker’s yeast glutathione reductase (9.4 milliunits/µL). The analyses were performed in a 96-well microtiter plate with triplicate wells for each sample. GSH standards were prepared at concentrations of 0, 60, and 120 ng/mL in 1× PBS (pH 7.5) with 5 mM EDTA. Added to each well were 140 µL of DTNB, 20 µL of cell extract, 20 µL of GSH reductase, and 20 µL of NADPH. The plate was immediately inserted into a Molecular Devices (Sunnyvale, CA) kinetic microplate reader and the absorbance (405 nm) of the wells determined. The plate was read every minute for 9 min. Values were plotted against the standard curve to determine intracellular GSH concentration. All experiments were performed at least twice (n g 2). Western Blot Analysis of Heme Oxygenase and GST. Approximately 5 × 105 AA8 cells were plated in 60 mm diameter culture dishes and grown to confluence. Cells were treated with IC50 or IC80 concentrations of drugs and incubated overnight at 37 °C. Cells were harvested 18 h later and transferred to 15 mL conical tubes. Additional experiments were carried out at incubation intervals of 8 and 28 h to ensure that peak induction levels were not overlooked. Cells were spun down in a centrifuge at 5000g for 5 min. Cell pellets were suspended in 1.5 mL of 1× PBS and centrifuged again. Ice-cold HNE extraction buffer (20 mM Tris-HCl, pH 7.4, 0.25 M sucrose, 1 mM EDTA, 1 mM PMSF, 100 µg/mL aprotinin) was added to the cell pellets, and cells were lysed by repeated passage through a 23 gauge needle. Further centrifugation at 5000g (5 min) removed debris to yield
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Figure 3. Gel showing DNA cleavage by etoposide, 48 and 109, in H2O containing 5 mM DTT. Duplicate reactions were run side by side. Lanes C1 and C2, DNA only; lanes 1 and 2, etoposide; lanes 3 and 4, 48; lanes 5 and 6, ascididemin; lanes 7 and 8, 109. Ascididemin and its analogues produced significant DNA cleavage. Upper bands represent nicked plasmid DNA, lower bands represent supercoiled plasmid DNA. Etoposide requires top 2 to produce DNA cleavage. All drugs were tested at 91 µM. the lysate of interest. Protein concentration was quantified by the Bradford protein determination assay (34). Oxidative stress protein extraction followed the method of Kutty et al. (35). Each lysate (10 µg of protein) was subjected to western blot analysis. Proteins were fractionated on a 12% denaturing polyacrylamide gel. Samples (15 µL protein dilution and 15 µL of 2× SDS loading buffer) were heated in boiling H2O for 3 min and loaded onto the gel. Molecular weight standards were run alongside the test samples. PAGE was carried out at 40 V for 4 h followed by electroblot transfer of proteins to nitrocellulose membrane at 10 V overnight (4 °C). Nitrocellulose membranes were stained with Ponceau S to confirm transfer of proteins. Lanes and standards were marked with a ballpoint pen. Membranes were destained in TBST (150 mM NaCl, 20 mM Tris, pH 8.0, with Tween 20), blocked for 2 h at 37 °C in a milk protein blocking solution (3% BSA, 5% nonfat milk in TBST), and washed with TBST, three times for 5 min. Membranes were transferred to plastic pouches to which primary rabbit anti-human HO-1 (StressGen, Victoria BC, Canada) or primary rabbit anti-human GST (Oxford Biomedical Research, Oxford, MI) were added in a 1:400 or 1:750 dilution, respectively, in a 10 mg/mL BSA/TBST solution. Membranes were incubated with primary antibody overnight (37 °C). Membranes were then washed with TBST, three times for 5 min, and treated with secondary anti-rabbit antibody conjugated to alkaline phosphatase for 1 h at room temperature. BCIP/NBT, substrates for alkaline phosphatase, were used in colorimetric detection of HO-1 or GST. Sodium arsenite was used as a positive control for HO induction and analyzed concurrently. A liver cytosolic extract, containing induced GST, from a polychlorinated biphenyl (PCB)-treated rat (generous gift from Dr. Michael R. Franklin, University of Utah) served as a GST positive control. Titrations of purified mouse hepatic GST were performed to determine the detection limit of this system. The primary GST antibody was able to detect as little as 0.02 µg of purified GST (or 0.2% of the loaded 10 µg of protein extract).
Results Quantitation of in Vitro DNA Cleavage. The potent DNA cleaving abilities of ascididemin, 109, and 48 were first observed in a quantitative DNA cleavage assay as they were being tested for top 2 inhibitory activity. The compounds were found to significantly inhibit top 2 catalytic activity and to generate some minor top 2 dependent DNA cleavage (Figure 2). However, under closer examination, it was discovered that only 5 mM DTT and 91 µM ascididemin, 109, or 48 in H2O was required to stimulate maximal DNA cleavage (Figure 3), independent of topoisomerases. Concentration and Time Dependency of DNA Cleavage. DNA cleavage by ascididemin, 109, and 48 was found to be both concentration and time dependent. Plasmid DNA treated with increasing concentrations of drug, in the presence of 5 mM DTT, showed concentration-dependent cleavage up to 100 µM final concentration
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Figure 4. Concentration-dependent DNA cleavage by ascididemin, 48 and 109. Drug concentrations used were 10 µM, 50 µM, 100 µM, 500 µM, and 1 mM. The concentration of DTT in the reactions was 5 mM where indicated. Lane C1, DNA only; lane C2, DNA + DTT; lane 1, 1 mM 109; lane 2, 10 µM 109 + DTT; lane 3, 50 µM 109 + DTT; lane 4, 100 µM 109 + DTT, lane 5, 500 µM 109 + DTT; lane 6, 1 mM 109 + DTT; lane 7, 1 mM 48; lane 8, 10 µM 48 + DTT; lane 9, 50 µM 48 + DTT; lane 10, 100 µM 48 + DTT; lane 11, 500 µM 48 + DTT; lane 12, 1 mM 48 + DTT; lane 13, 1 mM ascididemin; lane 14, 10 µM ascididemin + DTT; lane 15, 50 µM ascididemin + DTT; lane 16, 100 µM ascididemin + DTT; lane 17, 500 µM ascididemin + DTT; lane 18, 1 mM ascididemin + DTT. Maximal cleavage was reached by 50 µM for 109 and 48, ascididemin was more potent, maximal cleavage obtained by 10 µM Abbreviations: n ) nicked, l ) linear, and sc ) super coiled plasmid DNA, respectively.
(Figure 4). High concentrations (1 mM) of drug were sufficient to fully cleave the DNA, even in the absence of DTT. In some lanes (i.e., 17 and 18), the total amount of DNA in the bands appears to be less than in the other reactions. This can be accounted for by the loss of prominent DNA bands resulting from extreme fragmentation at higher drug concentrations, as well as the competitive displacement of ethidium bromide from the remaining DNA by the pyridoacridines. As shown in Figure 4, 109 was most potent at stimulating DNA cleavage followed by similar cleavage abilities of 48 and ascididemin. The pyridoacridine analogues 21 and 31 did not cleave DNA significantly. The DNA cleavage by ascididemin, 109, and 48 yielded predominantly single strand breaks, resulting in DNA with nicked conformation. Very little linear DNA (the product of DNA double strand breakage) was detected. Similarly, drug-stimulated DNA cleavage was discovered to be time dependent. DNA cleavage reactions that contained 500 ng of plasmid DNA, 100 µM drug concentration, and 5 mM DTT were quantitated after incubation times of 5, 10, 15, 20, 25, 30, 45, and 60 min. Near maximal cleavage was observed around 30 min for ascidemin, 109, and 48 (Figure 5). In comparison, DNA cleavage by 48 was only 80% of ascididemin and 109, although it appeared to plateau earlier. Therefore, on the basis of this time dependence, compounds were routinely tested using 30 min 30 °C incubations. Very little DNA cleavage was detected with other pyridoacridines tested (Table 1). Effect of Antioxidants, Antioxidant Enzymes, Metals, Chelators, and Radical Scavengers on DrugInduced DNA Cleavage. A variety of metals, radical scavengers and chemoprotective agents were employed to determine which reactive species were important for DNA cleavage by ascididemin, 109, and 48. Each of the metals was titrated to determine the maximum concentration at which their effects on DNA equaled controls. Similarly, serial dilutions of chelators, antioxidants, and scavengers were assayed to ensure inclusion of effective concentrations. None of the metals
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Figure 5. Time-dependent DNA cleavage by ascididemin, 109, and 48. DNA cleavage by the three compounds increased with time of incubation (incubation temp ) 30 °C). Maximal cleavage was reached approximately 45 min under these conditions. Table 1. Comparison of in Vitro DNA Cleavage Abilities of Ascididemin and Its Analoguesa compd
% cleavageb
compd
% cleavageb
ascididemin 109 48 21 31 BC1 29 BC1 55.1
73.2 97.3 75.3 3.5 8.0 11.0 3.0
BC1 55.2 BC1 55.3 BC1 14.2 BC1 14.3 NP-164 amphimedine neoamphimedine
8.5 2.0 5.3 3.3 3.0 3.0 6.0
a All reactions repeated three times. b Values are % cleavage above untreated controls.
Figure 6. In vitro DNA cleavage protection by catalase against ascididemin. Lane 1, 91 µM ascididemin; lane 2, 91 µM ascididemin + 5 mM DTT; lane 3, 91 µM ascididemin + 5 mM DTT + 32 nM SOD; lane 4, 91 µM ascididemin + 5 mM DTT + 28 nM catalase. Catalase reversed drug-DTT-stimulated DNA cleavage. SOD had no effect. Upper bands represent nicked plasmid DNA, lower bands represent supercoiled plasmid DNA.
(e.g., FeSO4 to a concentration of 0.43 µM with 23 µM drug, or NiCl to a concentration of 23 mM with 91 µM drug), facilitated drug cleavage. Nickel actually decreased DNA cleavage, possibly because of drug-Ni interaction reducing the effective drug concentration. Addition of chelators (up to a concentration of 23 mM) did not protect against cleavage. In contrast, catalase (28 nM in 100 µM drug) provided extensive protection from ascidemin, 109, or 48 induced DNA cleavage (Figure 6). Such protection was not seen with SOD. Of the antioxidants, only glutathione (227 µM in 93 µM drug) afforded strong protection from druginduced cleavage (see cell data, below). BHA (93 µM in 93 µM drug) showed some protection, as did N-acetylcysteine. The radical scavengers, ethanol, and benzoic acid (BA) also exhibited some protection against DNA cleavage (Figure 7).
Figure 7. In vitro DNA cleavage protection against ascididemin, 48, and 109 with benzoic acid (BA). Lane C1, DNA only; lane C2, DNA + 5 mM DTT; lane C3, DNA + 45.4 mM benzoic acid (BA); lane C4, DNA + DTT + 45.4 mM BA; lane 1, DNA + 91 µM 109; lane 2, DNA + DTT + 91 µM 109; lane 3, DNA + DTT + 91 µM 109 + 4.54 mM BA; lane 4, DNA + DTT + 91 µM 109 + 22.7 mM BA; 5, DNA + DTT + 91 µM 109 + 45.4 mM BA; lane 6, DNA + 91 µM 48; lane 7, DNA + DTT + 91 µM 48; lane 8, DNA + DTT + 91 µM 48 + 4.54 mM BA; lane 9, DNA + DTT + 91 µM 48 + 22.7 mM BA; lane 10, DNA + DTT + 91 µM 48 + 45.4 mM BA; lane 11, DNA + 91 µM ascididemin; lane 12, DNA + DTT + 91 µM ascididemin; lane 13, DNA + DTT + 91 µM ascididemin + 4.54 mM BA; lane 14, DNA + DTT + 91 µM ascididemin + 22.7 mM BA; lane 15, DNA + DTT + 91 µM ascididemin + 45.4 mM BA. Drugs were tested at 91 µM, DTT was 5 mM when added. Protection can be seen against 48 from 4.54 mM benzoic acid and against ascididemin and 109 with 22.7 mM BA. The designations n, l, and sc stand for nicked, linear, and supercoiled DNA, respectively.
Assessment of DNA Cleavage under Anoxic Conditions. Redox cycling requires a source of oxygen. If, in fact, ascididemin, 109, and 48 induce DNA cleavage through the production of ROS, this ability would cease under anoxic conditions. As hypothesized, ascididemin, 109, and 48 failed to stimulate DNA cleavage above controls under anoxic conditions. A positive control reaction of 95 µM 109 with 5 mM DTT under aerobic conditions produced DNA cleavage as shown before. The results indicated that oxygen is a necessary component for in vitro DNA cleavage stimulation by these drugs. Reduction-Oxidation Potential. To quantify chemical characteristics of ascididemin, 109, and 48 that influence the degree of DNA cleavage, their reductive cleavage potentials were compared to those of the noncleaving analogues 21 and 31. Using Kohen’s (27) method for measuring reductive potential by cyclic voltammetry, both hyperpolarized (oxidative) and depolarized (reductive) currents were quantified. The redox potentials determined for 48, ascididemin, 109, 31, and 21, were -0.157, -0.226, -0.251, -0.284, and -0.318 V, respectively (Figure 8). The three cleaving analogues were marginally more easily reduced than the two noncleaving compounds. Electron Paramagnetic Resonance (EPR) Spectroscopy. Because each discrete radical state carries its own corresponding energy, electron paramagnetic resonance (EPR) spectrometry was used to measure energy differences between the DNA cleaving and noncleaving molecules. EPR utilizes electromagnetic radiation in the gigahertz range to create a magnetic field to interact with and excite unpaired electrons to a higher energy state. Theoretically, EPR, could confirm the presence or absence of electrochemically reactive molecules under these experimental conditions. In EPR, the integrated intensity, or area under the curve, of the signal is proportional to the concentration of the active species in the sample. Reactions containing 100-500 µM drug, 5-10 mM DTT,
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Figure 8. Redox potential values of 48, ascididemin, 109, 31, and 21. Cyclic voltammetry revealed slightly less energy necessary to reduce ascididemin, 48, and 109 than the two inactive analogues, 21 and 31. The redox curve for 109 is shown.
Figure 9. Electron paramagnetic resonance spectra. Panel A shows the classical triplet signal of the positive control reaction H2O2 + FeSO4 + PBN + H2O. The spiked signal pattern was observed with a large signal intensity of -36 × 103 to 46 × 103 mV. Panel B shows the background signal obtained with 10 mM DTT + 100 mM PBN (spin trap agent N-tert-butyl-R-phenyl-nitrone) + H2O. Panel C shows the electron paramagnetic resonance spectrum of 1 mM 109 + 10 mM DTT + 100 mM PBN + H2O. A complicated spectrum was detected with an increased intensity over baseline of -8.6 × 103 to 8 × 103 mV. Panel D shows the electron paramagnetic resonance spectrum of 1 mM 21 + 10 mM DTT + 100 mM PBN + H2O. The signals observed with the non-DNA cleaving analogues were no different than baseline with a signal intensity of -1.75 × 103 to 3 × 103 mV. Experimental parameters were microwave frequency ) 9.678 GHz, receiver gain ) 7.96 × 105, modulation frequency ) 100 kHz, modulation amplitude ) 1 G, time constant ) 163.840 ms, and sweep time ) 41.943 s.
and 50 mM PBN in 300 µL of H2O final volumes were dispensed into quartz flat cells and scanned in the EPR. With the test reactions a signal calibration test was performed as well as a signal-to-noise ratio test to confirm optimal sensitivity and functioning of the spectrometer. The experimental parameters of the EPR system were optimized for each sample but were similar among the compounds tested. The parameter settings were fre-
quency, 9.785 Ghz; receiver gain, 7.10 × 105; modulation frequency, 100 kHz; modulation amplitude, 1 G; time constant, 163.84 ms; and scan time, 41.943 s. A hyperfine triplet spiked EPR signal (Figure 9b) was recorded for the positive control reaction (H2O2, FeSO4, PBN in H2O), which produced •OH radicals through a Fenton reaction. The half-lives of hydroxyl radicals were prolonged by use of a spin trap reagent (PBN). Baseline signal was
Mechanism of Ascididemin Cytotoxicity
Chem. Res. Toxicol., Vol. 16, No. 2, 2003 119
Table 2. IC50 Values in Culture (µM)a drug
AA8
EM9
xrs-6
HCT 116
KB
109 48 ascididemin
8.3 28.5 3.1
0.3 0.5 0.4
1.4 8.7 0.7
1.8 6.6 0.1
8.2 6.7 0.6
21 31
25.8 3.9
34.5 7.1
ND ND
ND ND
ND ND
a All cell treatments were run at least twice. Mean values are given. ND ) not determined.
determined for DTT + PBN in H2O (Figure 9a). Hyperfine splitting was observed for ascididemin, 109, (Figure 9c) and 48, but the signal was complex, indicating detection of many radical species. Nevertheless, spectra of all three DNA-cleaving drugs were greater in magnitude than spectra from untreated controls, with signal excursions of -8 mV to 8 mV. Reactions containing noncleaving analogues (21 and 31; Figure 9d) produced only baseline signals. Therefore, it was concluded that only the active drugs were capable of radical production. DNA Binding Affinity/Intercalation. Intercalation is one mechanism of drug interaction with DNA and is thought to be important for the cytotoxocity and reductive cleavage potential of several marine pyrroloiminoquinones and pyridoacridines (15). An ethidium bromide displacement assay was used to measure intercalation. Eight dilutions (50 µM-100 nM) of ascididemin, 109, 48, 21, and 31 were reacted with 0.5 µM ethidium bromide and 0.5 µM salmon testes DNA. Displaced ethidium bromide was quantified by measuring a decrease in fluorescence at 600 nm. A loss of fluorescence was detected for all compounds at 50 µM, consistent with DNA binding which displaced ethidium bromide (i.e., intercalation). Ethidium bromide was completely displaced by ascididemin and 21 at 10 and 20 µM concentrations, respectively. In contrast, displacement of ethidium bromide was not complete at 20 µM concentrations for 109, 48, or 31. Although all these compounds have the ability to intercalate DNA, this capacity did not correlate with DNA cleavage activity. Cytotoxicity Assays. Ascididemin, 109, 48, 21, and 31 were tested for cytotoxicity in three Chinese hamster ovary (CHO) cell lines. The repair competent AA8 line was compared to the DNA strand break repair deficient lines, EM9 and xrs-6. Enhanced toxicity to the repair deficient lines implicated production of DNA strand breaks as a mechanism of cytotoxicity. Two human tumor cell lines, HCT 116 (colon) and KB (nasopharyngeal) were also tested. All the pyridoacridines tested were found to be cytotoxic at varying concentrations. More importantly, enhanced cytotoxicity was observed in the DNA damage repair lines when compared to wild-type (Table 2). EM9 cells were much more sensitive than AA8 cells to the DNA cleaving drugs. This effect was less dramatic for xrs-6 cells; however, they still exhibited enhanced sensitivity to the DNA cleaving analogues. These data implicate DNA strand breakage as the mechanism of cytotoxicity (Figure 10). The two noncleaving analogues, 21 and 31, did not show enhanced toxicity to the DNA damage sensitive cells. Ascididemin was toxic to human colon carcinoma cells (HCT 116) with a sub-micromolar IC50. Glutathione Protection against Cytotoxic Drug Effects. Pretreatment of AA8 cells with GSH protected from the cytotoxicity of ascididemin, 109, and 48. Cells were treated with IC90 concentrations of drug, and
Figure 10. Enhanced cytotoxicity of 109 in single strand break repair deficient (EM9) CHO cells. The cytotoxicity curves for 109 in the AA8 and EM9 cell lines are shown. Data shown are the average of two experiments.
Figure 11. Protection of CHO AA8 cells from 109 cytotoxicity by glutathione. Pretreatment of cells with 25-200 µM glutathione 2 h before drug treatment significantly protected cells against the IC90 concentration (8.5 µM) of 109. Error bars equal standard deviation
increasing concentrations of GSH (25-200 µM). Addition of GSH to drug-treated cells restored growth to control levels in a dose dependent fashion (Figure 11). Similar protection was not observed against 21 or 31. Determination of Intracellular Glutathione Concentration. Our results are consistent with the finding that GSH administered externally is actively taken up by CHO cells in culture (33). AA8 cells were treated with either 2 mM DEM or 400 µM GSH. Cells receiving DEM had a 2 h pretreatment with GSH. GSH standards and treated samples were run concurrently to provide a standard curve to ensure accurate quantification of GSH levels. The data revealed a 3-fold increase in intracellular GSH with external GSH treatment. Diethylmaleate (DEM), a GSH depletor, was able to reverse the effect of exogenous GSH on intracellular GSH pools. DEM also decreased intracellular GSH levels of untreated AA8 cells, though this effect was slight because levels were low to begin with. Thus, intracellular scavenging molecules such as GSH can protect from ROS mediated cytotoxicity of ascididemin, 109, and 48.
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Figure 12. Western blot analysis of heme oxygenase-1 (HO-1) induction by ascididemin, 48, and 109 in CHO AA8 cells. Lane 1, IC80 ascidemin; lane 2, IC50 ascididemin; lane 3, IC80 48; lane 4, IC50 48; lane 5, IC80 109; lane 6, IC50 109; lane 7, 25 µM sodium arsenite; lane 8, 6.25 µM sodium arsenite; lane 9, untreated; lane S, molecular weight standards from top to bottom: myosin heavy chain (205 kDa), phosphorylase B (97 kDa), bovine serum albumin (66 kDa), and ovalbumin (40 kDa). Cells were treated for 18 h. Strong induction of HO-1 (32 kDa) was seen in the positive control (sodium arsenite) reaction and in all drug treatments at both concentrations. Only slight induction was seen with 21, none with 31 (Table 3). Table 3. HO-1 (Heme oxygenase-1) Induction by Ascididemin and Analogues in Culturea treatment
AA8
sodium arsenite 6.25 µM sodium arsenite 25 µM 109 IC50 109 IC80 48 IC50 48 IC80 ascididemin IC50 ascididemin IC80 21 IC50 21 IC80 31 IC50 31 IC80
strong strong strong strong strong strong strong strong weak weak none none
a The data represent a summary of two western blots per treatment.
Determination of Heme Oxygenase-1 Induction. HO-1 is a 32 kDa oxidative stress response protein present in mammalian cells that acts as part of a general oxidative stress mechanism (33). Western blot analysis for HO-1 induction was used to test whether AA8 cells treated with vehicle, ascididemin, 109, 48, 21, or 31 responded in this manner. Sodium arsenite (a potent inducer of HO-1) was used as a positive control for both cell lines. The results of the western blot analysis are depicted in Table 3. Sodium arsenite and cleaving-drug treatments produced strong HO-1 induction in the AA8 cells (Figure 12), which supports the hypothesis that these agents provoke an oxidative stress response. The noncleaving agent 21 induced a weak response while 31 did not induce HO-1. Induction of glutathione S-transferase (GST) was detected in pyridoacridine-treated cells as well, but the overall resulting induction was low.
Discussion Ascididemin, 109, and 48, were the only three pyridoacridines (of the 14 marine analogues tested) found to have the in vitro ability to cleave plasmid DNA. Oxygen and the reducing agent DTT proved to be the sole requirements for full activity. Several published studies have supported a top 2-mediated mechanism for ascididemin DNA cleavage (6, 13). However, the mechanism is controversial. For example, Bonnard and co-workers (15) found that ascididemin had little effect on topoisomerase I or II catalytic activity. They concluded that
ascididemin’s mechanism of action was dependent on its ability to intercalate into DNA rather than its selective targeting of topoisomerases. These conflicting results illustrate the uncertainty regarding the mode of action responsible for ascididemin cytotoxicity. The in vitro characterization of the DNA-cleaving abilities of ascididemin, 109, and 48 presented here resolves these uncertainties. Increases in drug concentration lead to increases in DNA cleavage. DNA damage was found to be both concentration and time dependent. Antioxidants and antioxidant enzymes protected DNA against ascididemininduced cleavage, indicating a ROS-mediated mechanism. Extensive protection against DNA cleavage by the antioxidant enzyme catalase, suggested H2O2 as the possible reactive species initially generated. In addition, the antioxidants glutathione, BHA, NAC, and the radical scavenger benzoic acid all protected against DNA damage by ascididemin, 109, and 48. Redox cycling and ROS generation require a source of oxygen. Under anaerobic conditions, ascididemin, 109, and 48 lost their ability to cleave DNA, even in the presence of DTT and reaction conditions that were otherwise sufficient to induce significant DNA damage. The lack of stimulation of DNA cleavage by metal ions and only slight protection from cleavage by chelators (at very high concentrations) excluded a Fenton-type reaction as the primary mechanism of ROS generation. Overall, these data strongly implicate an oxidative mechanism in ascididemin DNA cleavage. We propose that the production of ROS likely occurs through reduction of the iminoquinone moiety. Reduction of the drugs to semiquinone species likely facilitates the production of H2O2 and DNA damaging free radicals. This hypothesis was supported by experiments that revealed that ascididemin, 109, and 48 are reducible at potentials consistent with bioreduction in the cell (∼-200 mV), similar to the conclusion of previous work (26), which correlated higher reductive potentials of marine metabolites with increased cytotoxicity and oxidative DNA cleavage. The reductive potentials of the ascididemin analogues compare to that of NADPH (-280 mV). It is likely that reduction of the ascididemin analogues is essential for their toxicity. This is similar to doxorubicin and daunomycin, for which reduction of the C-11 hydroxyl group has been reported to be indispensable for • OH formation, DNA cleavage, and ultimately, cytotoxicity (27).
Mechanism of Ascididemin Cytotoxicity
Free-radical production suggested by EPR spectrometry further supports a ROS-mediated mechanism of DNA cleavage. ROS have been reported to react indiscriminately with the closest neighboring molecules (37). ROS are thought to abstract H+ from the deoxyribose sugars of either purine or pyrimidine bases of DNA forming random single strand breaks. DNA sequence analysis of ascididemin-treated DNA revealed such random DNA cleavage, yielding a uniform smear of fragments in a DNA sequencing gel (unpublished results). Mitra et al. (38) reported similar nonspecific DNA cleavage by a natural antibiotic, leinamycin, via an oxidative mechanism. The ethidium bromide displacement revealed DNA intercalation of ascididemin, 109, 48, 21, and 31 would be likely. Several other marine pyridoacridines with similar structures (e.g., dercitin, cystodytin J, and diplamine) have been previously shown to intercalate into DNA (29, 39). The ability of ascididemin and its analogues to interact closely with DNA increases the likelihood that ROS formed as a result of drug reduction would directly affect DNA. Although the analogues 21 and 31 can intercalate DNA, their inability to produce ROS fully explains their inactivity. Mechanism-based cytotoxicity screening revealed the capacity of ascididemin, 109, and 48 to predominantly cause single strand DNA breaks. The mutant CHO cell line EM9 was particularly sensitive to the lethality of these molecules as compared to the wild-type AA8 cell line. The EM9 cells have a defective XRCC1 protein, which plays a critical role in DNA ligase III activity, DNA polymerization and repair of DNA single strand breaks (40). This deficiency makes them hypersensitive to drugs that cause ROS (41, 42), or single strand breaks via top 1 (43). The xrs-6 line has a defect in the Ku 80 protein, important in the repair of double strand breaks (44). The xrs-6 cells have decreased recombinatorial repair versus wild-type cells. In combination, these cell lines provide mechanistic insight into a drug’s effects. The enhanced cytotoxicity of ascididemin and other DNA cleaving analogues in these two mutant lines correlated well with DNA cleavage activity in vitro. All three drugs exhibited enhanced toxicity to EM9 cells and, to a lesser extent, the xrs-6 cells. This activity was distinct from that of the DNA noncleaving compounds 21 and 31, which showed no enhanced cytotoxicity to the repair deficient lines. GSH protected cells in culture from ascididemin toxicity and mimicked GSH protection from DNA cleavage in vitro. Determination of intracellular GSH levels confirmed that external GSH treatment increased intracellular GSH levels. Again, these data suggest a ROSdependent DNA cleavage mechanism of cytotoxicity. Induction of oxidative stress genes is a common effect of ROS-generating drugs. The role of oxidative stress proteins and oxidative stress gene products in physiological response to oxidative stress is well documented (36). Heme oxygenase-1 and GST are two oxidative stress proteins that may participate in such responses. HO-1 and GST were both induced in our test models effectively linking cellular oxidative stress with cell damage by ascididemin, 109, and 48. HO-1 and GST induction by these cytotoxic drugs was consistent with ROS being formed in sufficient quantities to warrant an aggressive defense. Weak induction observed with the noncleaving analogues may be due to other effects of their cytotoxicity (i.e., an artifact of testing all compounds at equitoxic
Chem. Res. Toxicol., Vol. 16, No. 2, 2003 121 Table 4. In Vivo Anti-Tumor Activity of Ascididemin in Six Different Tumor Cell Lines Using the Hollow Fiber Assay (NCI)a %T/C (12 mg/kg) cell line
i.p. fiberb
s.c. fiberc
OVCAR-3 (ovarian) SF-295 (CNS) MDA-MB-435 (breast) MDA-MB-231 (breast) NCI-H23 (nonsmall cell lung) LOX IMVI (melanoma)
10d
76 81 80 100 44* 29*
22* 30* 40* 77 97
a MTD ) 24 mg/kg ascididemin. Six other cell lines were tested with insignificant results, data generated at NCI. b i.p. ) intraperitoneally located fiber. c s.c. ) subcutaneously located fiber. d Significant tumor growth suppression based on %TC < 50.
concentrations). Overall, induction of these classical oxidative stress markers, when combined with the preceding data, argues strongly in favor of oxidative stress as the predominant mechanism of ascididemin cytotoxicity. Studies performed at the NCI revealed the significant activity of ascididemin in vivo in a hollow fiber assay (9, 45; Table 4). Because of its anticancer potential, ascididemin was advanced to human tumor xenograft studies in nude mice, but tests showed only weak activity. MDAMB-435 and -231 breast, U251 CNS, LOX IMVI melanoma, OVCAR-3 ovarian, and HCT-116 colon cancers were tested. A maximal response of 58% T/C was seen in the colon cancer xenograft when treated ip with 8 mg/ kg on days 5 and 9. The potent ability of these molecules to cleave DNA via ROS generation mechanism is unprecedented among the marine pyridoacridines and is likely a strongly pro-apoptotic stimulus to many cell types (16).
Acknowledgment. We thank Dr. M. R. Franklin and Dr. C. M. Ireland for their suggestions and comments; Dr. C. Mayne, D. Chamberlin, and R. Jameton for assistance with EPR; B. Bathe for assistance with the cyclic voltammetry experiments; and the Utah Cancer Center Support Grant for support of core facilities. This work was supported by California Sea Grant R/MP 68 (L.R.B.) and Auckland Medical Research Foundation (B.R.C.).
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