3-Amino-1,4-dimethyl-5H-pyrido[4,3-b]indole (Trp-P-1) triggers apoptosis by DNA double-strand breaks caused by inhibition of topoisomerase I

Bunsyo Shiotani and Hitoshi Ashida1,2

Division of Life Science, Graduate School of Science and Technology and 1 Department of Biofunctional Chemistry, Faculty of Agriculture, Kobe University, 1-1 Rokkodai-cho, Nada-ku, Kobe 657-8501, Japan

2 To whom correspondence should be addressed Email: ashida{at}kobe-u.ac.jp


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
3-Amino-1,4-dimethyl-5H-pyrido[4,3-b]indole (Trp-P-1) is one of the dietary carcinogens. At the initial step in the carcinogenic process, its exocyclic amino group is metabolically activated to the hydroxyamino derivative by the cytochrome P450 (CYP) 1A and 1B subfamily and then form DNA adducts, which are considered to be the main cause of DNA damage during the carcinogenic process. On the other hand, our previous study has shown that Trp-P-1 exhibits cytotoxicity to primary cultured rat hepatocytes, via induction of caspase-9-dependent apoptosis without being metabolized by CYP 1A1. In the present study, we investigated what type of DNA damage would be involved in the induction of apoptosis induced by Trp-P-1. When RL-34 cells derived from normal rat liver were treated with a high (30 µM) concentration of Trp-P-1, apoptotic events such as the loss of cell viability, nuclear condensation and the activation of caspase-3 were observed. In these apoptotic cells, intracellular topoisomerase I activity was inhibited and histone H2AX phosphorylation, which occurs after introduction of DNA double-strand breaks (DSBs), was observed in the early phase of the apoptosis. On the other hand, treatment with a non-apoptotic concentration (1 µM) of Trp-P-1 increased the formation of 8-hydroxy-2'-deoxyguanosine. The formation of DNA adducts was detected at almost the same level in both cells exposed to the apoptotic and non-apoptotic concentrations of Trp-P-1. These results indicate that Trp-P-1-induced apoptosis was triggered by DNA DSBs through the inhibition of topoisomerase I but not the formation of DNA adducts.

Abbreviations: CYP, cytochrome P450; DMSO, dimethyl sulfoxide; DSBs, double strand breaks; 8-OHdG, 8-hydroxy-2'-deoxyguanosine; PBS, phosphate buffered saline; Trp-P-1, 3-amino-1,4-dimethyl-5H-pyrido[4,3-b]indole; Trp-P-2, 3-amino-1-methyl-5H-pyrido[4,3-b]indole


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The heterocyclic amine 3-amino-1,4-dimethyl-5H-pyrido[4,3-b] indole (Trp-P-1) not only causes carcinogenesis (1) but also induces apoptosis (2). Several types of DNA damage cause genetic alterations, and the accumulation of such alterations leads normal cells to cancer cells. In the case of Trp-P-1, the initial step in the carcinogenic process is related to the oxidation of exocyclic amino group to the hydroxyamino derivative and is mainly catalyzed by the cytochrome P450 (CYP) 1A and 1B subfamily (37). It has been reported that N-hydroxy-Trp-P-1 is a direct-acting mutagen toward Salmonella typhimurium, being able to bind directly and covalently to DNA (8). The N-hydroxy form is further metabolically activated to N-O-acetyl-Trp-P-1, the ultimate form, which damages DNA by the formation of DNA adducts (9). It is known that DNA adducts formed by heterocyclic amines including Trp-P-1 cause genetic mutations (10). In addition, it has been reported that 3-amino-1-methyl-5H-pyrido[4,3-b]indole (Trp-P-2), the structure of which is similar to that of Trp-P-1, produces reactive oxygen species (ROS) that can cause oxidative DNA damage (11). The formation of 8-hydroxy-2'-deoxyguanosine (8-OHdG) in DNA is an important process in oxygen-radical induced mutagenesis and is a good marker of carcinogenesis (12). Furthermore, it has also been reported that Trp-P-1 inhibits DNA excision repair following UV-induced DNA damage (13), and this effect may be due to the inhibition of topoisomerases by Trp-P-1 (14). These DNA damages, the formation of DNA adducts and 8-OHdG and double strand breaks (DSBs) following the inhibition of topoisomerases, can be a trigger of apoptosis, if the damages were severe (1518).

The DNA-damaging carcinogen Trp-P-1 is cytotoxic to primary cultured rat hepatocytes, due to induction of apoptosis (2). Trp-P-1 induces apoptosis via caspase-9 without being metabolized by CYP 1A1 (19), indicating that DNA adduct formation is not involved in the induction of apoptosis. However, it is not clear what type of DNA damage is involved in apoptosis induced by Trp-P-1. In this study, RL-34 cells derived from normal rat liver were treated with apoptotic (30 µM) and non-apoptotic (1 µM) concentrations of Trp-P-1, and the DNA damages were examined. Intracellular topoisomerase I was inhibited and histone H2AX was phosphorylated after treatment with the apoptotic concentration of Trp-P-1 in the early phase of apoptosis. Formation of 8-OHdG was observed in the cells treated with the non-apoptotic concentration. The formation of DNA adducts was observed in the cells treated with either concentration of Trp-P-1 at almost the same level. These results indicate that apoptosis induced by Trp-P-1 was triggered by DNA DSBs through the inhibition of topoisomerase I.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Materials
Trp-P-1 (acetate form) was purchased from Wako Pure Chemical Industries (Osaka, Japan). A goat polyclonal antibody to caspase-3 was purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Mouse monoclonal antibodies to p53, phospho-p53 (ser-15) and phospho-histone H2AX (ser139) were purchased from Oncogene Research Products (Cambridge, MA), Cell Signaling Technology (Beverly, MA) and Upstate (Charlottesville, VA), respectively. Secondary antibodies to rabbit IgG and mouse IgG were purchased from Amersham Pharmacia Biotech (Tokyo, Japan). A secondary antibody to goat IgG was purchased from Wako Pure Chemical Industries. Micrococcal nuclease, phosphodiesterase II and nuclease P1 were purchased from Sigma Chemical (St Louis, MO). [{gamma}-32P]ATP was purchased from Amersham Pharmacia Biotech (Tokyo, Japan). pBR322 DNA and T4 polynucleotide kinase were purchased from Takara Shuzo (Shiga, Japan). All other chemicals were of the highest quality commercially available.

Cell culture and treatment
The rat liver cell line RL-34 (JCRB 0247) was obtained from the Health Science Research Resources Bank (Osaka, Japan) and cultured in DMEM supplemented with 10% heat-inactivated fetal bovine serum (Sigma, Irvine, UK), 4 mM L-glutamine, 100 µg/ml streptomycin and 100 U/ml penicillin. The cells were incubated at 37°C in a humidified atmosphere containing 5% CO2. RL-34 cells were treated with various concentrations of Trp-P-1 in dimethyl sulfoxide (DMSO) for various times as indicated in each figure. Parallel dishes were treated with vehicle alone to obtain control samples (maximum concentration of DMSO in the medium was 0.1% v/v).

Cell viability measurement
A cell viability test was performed using the MTT (3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide) test according to the method of Mosmann (20).

Fluorescent stain of nuclei
RL-34 cells were cultured on plastic coverslips and treated with or without Trp-P-1. The cells were washed twice with phosphate-buffered saline (PBS) and fixed to the coverslips using 1% glutaraldehyde in PBS for 30 min. After fixation, cells were stained with 1% 4',6-diamino-2-phenylindole (DAPI) for 30 min. Morphological microscopic analyses of nucleus structures were carried out with a fluorescence microscope.

Cell cycle analysis
Cell cycle analysis was performed after treatment with Trp-P-1 at various concentrations for the indicated times as shown in Figure 1. RL-34 cells were removed from culture dishes by trypsinization, collected by centrifugation, re-suspended in ice-cold PBS and then fixed in 70% ethanol at –20°C for 4 h. After fixation, cells were washed with PBS and re-suspended in PBS containing 100 µg/ml RNase and 10 µg/ml propidium iodide for 30 min at room temperature in the dark. Cells (1 x 104 cells/analysis) were analyzed by flow cytometry using a Coulter EPICS XL (Beckman Coulter, Tokyo, Japan) and cell cycle distribution was determined based on DNA contents. Cells distributed in sub-G1 phase were defined as apoptosis according to the criteria described previously (21).



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Fig. 1. Differential cell cycle regulation in Trp-P-1-treated cells. Cell cycle analysis was performed after treatment with Trp-P-1 at 1 or 30 µM for the indicated times. These cells were harvested and analyzed by the flow cytometry as described in Materials and methods. Cell cycle distribution was determined based on 2 and 4 N DNA contents. Cells with DNA contents <2 N are indicative of sub-G1 cells.

 
Fluorimetric assay of caspase-3-like protease activity
Proteolytic activity of caspase-3-like protease was measured spectrofluorometrically using the synthetic fluorogenic peptide substrate, Ac-DEVD-MCA, as described previously (19). The protease activity was calculated from the slope of the recording and calibrated with standard concentrations of 7-amino-4-methyl-coumarin.

Western blotting analysis
RL-34 cells were treated with Trp-P-1, and the nuclear protein extracts and cell lysates were prepared as described previously (19). For determination of caspase-3 and p53, aliquots of 30 µg of nuclear protein were separated on the 15 and 10% gels, respectively. In the case of histone H2AX, aliquots of 30 µg protein of cell lysates were separated on the 15% gels. After SDS–PAGE, proteins were transferred onto a PVDF membrane followed by blocking of the non-specific binding sites with 10% FBS in TBST buffer (10 mM Tris–HCl, pH 8.0, 150 mM NaCl and 0.06% Tween 20) at 4°C overnight. The membranes were washed with TBST buffer four times for 5 min each and incubated with respective primary antibodies for 1 h. After washing with TBST buffer under the same conditions, the membranes were incubated with the secondary antibodies conjugated with horseradish peroxidase for 30 min. Specific immune complexes were visualized with the ECL detection system (Amersham Pharmacia Biotech).

32P-Post-labeling method
For DNA adducts analysis, DNA was isolated from aliquots of 1 x 108 RL-34 cells as described by Gupta (22) with some modifications. Briefly, the RL-34 cells were scraped off with 500 µl of PBS and homogenized with a glass homogenizer (10 strokes). The homogenate was incubated with proteinase K (500 µg/ml) at 50°C for 30 min. After addition of 200 µl of 1 M Tris–HCl (pH 7.4), the homogenate was extracted successively with 1 vol each of phenol (5 min), 1:1 mixture of phenol/Sevag (chloroform/isoamyl alcohol) (3 min) and Sevag (3 min). The aqueous phases were separated by centrifugation (17 000 g at 4°C 10 min for the first extraction and 5 min for subsequent extractions). After addition of 0.1 vol of 5 M NaCl, DNA was precipitated by gradual addition of 1 vol of absolute ethanol pre-cooled to –20°C. After inverting several times, DNA precipitate was obtained by centrifugation at 17 000 g for 20 min, and washed with 70% ethanol. The DNA sample was dissolved in 600 µl of 0.01 x NaCl/Cit/1 mM EDTA (1x NaCl/Cit = 0.15 M NaCl/0.015 M Na citrate). Residual RNA was removed by incubation with RNase T1 (50 U/ml) and RNase A (500 µg/ml) at 38°C for 30 min. After extraction of this solution with Sevag, DNA was recovered from the aqueous phase as described above, dissolved in 75 µl of 0.01 x NaCl/Cit/0.1 mM EDTA and quantified by UV-spectroscopy. Aliquots of 10 µg of isolated DNA were digested with a mixture of micrococcal nuclease and phosphodiesterase II, and the digest was 32P-post-labeled under the modified adduct intensification conditions as described by Reddy and Randerath (23). Briefly, the digest of 5 µg of DNA was added 1.5 µl of 0.3 M sodium acetate (pH 5.3) and 1 µl of 0.1 mM ZnCl2 followed by treatment with 1 µl of 5 mg/ml nuclease P1. The resulting mixtures were subsequently mixed with 1.5 µl of 0.5 M Tris–HCl (pH 7.4) and labeled enzymaticaly with T4 polynucleotide kinase and [32P]ATP (7000 Ci/mmol). Another 2 µg of DNA was separated from the digest and 32P-labeled and used for measurement of the total nucleotide levels after treatment with apyrase. The 32P-post-labeled samples were applied to a polyethyleneimine (PEI)-cellulose TLC sheet (Polygram CEL 300 PEI; Macherey-Nagel, Duren, Germany) and developed with 2.3 M sodium phosphate buffer (pH 6.0) to remove normal nucleotides. Modified nucleotides, which remained at the origin, were contact transferred to another PEI-cellulose sheet and then subjected to two-dimensional thin layer chromatography. The solvent system for development consisted of buffer A (2.25 M lithium chloride, 4.25 M urea, pH 3.5) from bottom to top and buffer B (0.8 M lithium chloride, 0.4 M Tris–HCl, 6.8 M urea, pH 8.0) followed by 1.7 M sodium phosphate buffer (pH 6.0) from left to right, with a 3.5 cm wick. Adducts were detected with a Bio-Image Analyzer (BAS 3000; Fuji Photo Film, Tokyo, Japan) after exposing the thin layer chromatography sheet to a Fuji imaging plate. Relative adduct labeling (RAL) was determined by the method of Randerath et al. (24).

8-OHdG analysis
For 8-OHdG analysis, DNA was isolated by the method described previously (19) with some modifications. Briefly, aliquots of 2 x 107 cells were homogenized in TE buffer (10 mM Tris–HCl, pH 7.4 and 10 mM EDTA) containing 0.5% SDS. The homogenates were incubated with 500 µg/ml of RNase A at 50°C for 30 min and then with 500 µg/ml of proteinase K at 50°C for 30 min. After addition of 0.5 M NaCl (final concentration), DNA was precipitated in 50% pre-cooled isopropanol. DNA precipitate was obtained by centrifugation at 17 000 g for 20 min, and washed with 70% ethanol. The DNA samples were dissolved in a mixture of 200 µl of 1 mM EDTA and 10 µl of 0.5 M sodium acetate. Then, they were digested with Nuclease P1 (10 U, 30 min, 37°C) and further incubated with alkaline phosphatase (3 U, 1 h, 37°C) after addition of 80 µl of 0.4 M Tris–HCl (pH 7.4). The amounts of 8-OHdG present in DNA were detected using a Hitachi HPLC (L-7100) equipped with a UV detector (Hitachi L-7420) and electrochemical detector (ECD; IRICA {Sigma}875) connected in series. An Inertsil column, ODS (i.d. 4.6 x 250 mm), was maintained at 35°C. The mobile phase consisted of 20 mM KH2PO4 (pH 4.6)/methanol (88/12, v/v) containing 0.1 mM EDTA, and the flow rate was 1.0 ml/min. The UV detector was set at 254 nm, {lambda}max of deoxyguanosine. ECD with a glassy carbon electrode was set at +600 mV. The molar ratio of 8-OHdG to deoxyguanosine (dG) in each DNA sample was determined based on the peak area of authentic 8-OHdG with the ECD and UV absorbance at A254 of dG.

Topoisomerase I activity assay
Crude nuclear extracts were prepared according to the method described by Deffie et al. (25) with some modifications. RL-34 cells were treated with various concentrations of Trp-P-1 for 3 h. The cells were washed twice with a NB solution [2 mM K2HPO4, pH 6.5, 5 mM MgCl2, 150 mM NaCl, 1 mM EGTA and 0.1 mM dithiothreitol (DTT)], lysed in 1 ml of the cold NB solution containing 0.35% Triton X-100 and 1 mM phenylmethylsulfonyl fluoride on ice for 10 min, and then centrifuged at 1000 g for 10 min. The nuclear pellets were washed once with the Triton-free NB solution, re-suspended in 150 µl of NB solution containing 0.35 M NaCl and kept on ice for 60 min. The mixture was then centrifuged at 17 000 g for 15 min, and the protein concentration in the nuclear protein extracts was determined by the method of Bradford (26).

DNA relaxation activity of topoisomerase I was examined according to the method of Liu (27) with some modifications. Briefly, the reaction buffer (50 mM Tris–HCl, pH 7.8, 100 mM KCl, 10 mM MgCl2, 0.5 mM DTT, 0.5 mM EDTA, 30 µg/ml bovine serum albumin and 0.3 µg pBR322 plasmid DNA) and various protein concentrations of nuclear extracts (total 20 µl) were incubated at 37°C for 15 min. The reactions were terminated by addition of 5x stop solution (5% SDS, 0.05% BPB and 50% sucrose). Then, the samples (15 µl each) were electrophoresed on the 0.7% agarose gels at 50 V in TBE buffer (89 mM Tris–HCl, 89 mM borate and 2 mM EDTA). The gels were stained with ethidium bromide and visualized on a UV transilluminator.


    Results
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Trp-P-1 induces apoptosis to RL-34 cells
Trp-P-1, but not the metabolically activated form of Trp-P-1, induces apoptosis in primary cultured rat hepatocytes (19,28). In this study, RL-34 cells, an immortal and non-transformed cell line originally derived from rat liver (29) was used. Figure 1 shows representative cell cycle profiles of asynchronous RL-34 cells treated with or without Trp-P-1. The experiment was repeated on more than three separate occasions with similar results. Cell cycle analysis showed that the control cells at 0 h were distributed in G0–G1 (63.9%) and G2–M phase (17.6%). After 24 h of treatment with the low concentration of Trp-P-1 (1 µM), the cells were accumulated in G2–M phase (45.2%). The accumulation in G2–M phase was time-dependent and associated with a decrease in G0–G1 and S phase. These responses were observed in cells treated with only the lower concentrations of Trp-P-1 (<10 µM, data not shown). In contrast, after 3 h of treatment with the high concentration of Trp-P-1 (30 µM), a small increase in the number of cells in G0–G1 phase (73.0%) followed by an increase in sub-G1 phase was observed. It has been proposed that the presence of sub-G1 cell population is indicative of apoptotic cells (21). Little change was seen in the distribution of the control cells treated with DMSO (vehicle alone) during 24-h culture. These observations indicated that the low concentration of Trp-P-1 induced cell cycle arrest in G2–M phase, whereas the high concentration induced arrest in G0–G1 phase resulting in apoptosis. These results suggest that different types of DNA damages occurred in these cells. Therefore, 30 µM is defined as an apoptotic concentration, whereas 1 µM is a non-apoptotic concentration in the following experiments.

Trp-P-1 showed strong cytotoxicity against RL-34 cells in a time- and dose-dependent manner (Figure 2A). The cell viability was decreased to below 20% after the cells treated with the apoptotic concentration of Trp-P-1 (30 µM) for 24 h. The viability of the cells treated with the non-apoptotic concentration of Trp-P-1 (1 µM) remained almost 100% at 24 h and was over 80% by 96 h (data not shown). Moreover, after treatment with the apoptotic concentration of Trp-P-1 for 6 h, nuclear condensation, one of the morphological characteristics of apoptotic cells, was observed (Figure 2B). Nuclei of the cells treated with the non-apoptotic concentration of Trp-P-1 did not show any such changes (data not shown).



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Fig. 2. Trp-P-1 induces apoptosis in RL-34 cells. (A) RL-34 cells were treated with 30 µM Trp-P-1 for 6 (square) or 24 h (circle). Cell viability was measured by MTT test. Data are presented as the means ± SD (n = 3). (B) RL-34 cells were treated with or without 30 µM Trp-P-1 for 6 h. Control cells were treated with 0.1% DMSO (vehicle alone). Nuclei were stained with DAPI and fluorescence photomicrographs were taken. Solid arrows indicate typical apoptotic cells.

 
In RL-34 cells treated with the apoptotic concentration of Trp-P-1, caspase-3-like protease activity increased in a time- and dose-dependent manner (Figure 3A and B). Immunoblotting analysis showed those activated caspase-3 p20 subunits were also increased in a time-dependent manner by 6 h (Figure 3C). In the cells treated with DMSO (control) or the non-apoptotic concentration of Trp-P-1, there was no increase in the caspase-3-like protease activity throughout the experimental period. These results are consistent with our previous findings that Trp-P-1 induces caspase-dependent apoptosis in primary cultured hepatocytes at the high concentration (30 µM) (19).



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Fig. 3. Activation of caspase-3 in the RL-34 cells after treatment with Trp-P-1. Cell lysates were prepared from RL-34 cells treated with 1 (circle) or 30 µM (triangle) Trp-P-1 or 0.1% DMSO (vehicle alone, square) for the indicated times (A) and from the cells treated at various concentrations for 12 h (B). The activities of caspase-3-like protease in the lysates were determined using the peptidyl substrate, Ac-DEVD-MCA. Activation of caspase-3 was also detected in RL-34 cells after induction of apoptosis by Trp-P-1 by western blotting (C). Nuclear protein extract was prepared from cells treated with 30 µM Trp-P-1 at the indicated time points. Equal amounts of nuclear proteins (30 µg) were separated by 15% SDS–PAGE followed by transfer onto a PVDF membrane. The membrane was probed with primary antibody to p20-subunit of caspase-3, and then treated with secondary antibody. Signals on the blots were detected with an ECL chemiluminescence detection kit.

 
Trp-P-1 increases the p53 protein level and alters its phosphorylation status
The p53 protein appears to sense multiple types of DNA damages and coordinate with multiple options for cellular responses, and is rapidly induced by various types of DNA damages. In the cells treated with the non-apoptotic concentration of Trp-P-1 (1 µM), the p53 protein was gradually accumulated in a time-dependent manner by 96 h (Figure 4A). On the other hand, the p53 level increased rapidly by 6 h, and decreased thereafter, in cells treated with the apoptotic concentration of Trp-P-1 (30 µM) (Figure 4B). Phosphorylation of p53 at ser-15 promotes both accumulation and functional activation of p53 in response to DNA damages (30,31). After treatment with 30 µM Trp-P-1, p53 protein was rapidly phosphorylated at 3 and 6 h followed by the acute accumulation of p53 protein, and rapidly dephosphorylated with the decrease in p53 level at 12 h since an intracellular phosphatase might be activated in dying cells (Figures 1 and 4B and C). However, the cells treated with 1 µM Trp-P-1, p53 protein were barely phosphorylated at 24 h.



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Fig. 4. Immunoblotting of p53 and phospho-p53 (ser-15) in RL-34 cells treated with Trp-P-1. Nuclear protein extracts were prepared from the cells treated with 1 (A and C) or 30 µM (B and C) Trp-P-1 at the indicated time points. Equal amounts of nuclear proteins (30 µg) were separated by 10% SDS–PAGE followed by transfer onto the PVDF membranes. The membranes were probed with primary antibody to p53 (A and B) or phospho-p53 (ser-15) (C) and then treated with secondary antibody. Signals on the blots were detected with an ECL chemiluminescence detection kit.

 
DNA adduct formation dose not trigger apoptosis
The DNA adducts formed in the cells treated for 24 h with 1 or 30 µM Trp-P-1 or N-hydroxy-Trp-P-1 (30 µM equivalent of Trp-P-1), prepared by incubation with microsomes of a recombinant yeast strain, Saccharomyces cerevisiae, AH22/pAMR2 expressing rat CYP 1A1 gene as described previously (19), were analyzed by the 32P-post-labeling method under the modified adduct intensification conditions. In the control sample, adduct spots could not be detected (Figure 5A). One major and several minor adduct spots were detected in each sample with the following total RAL values: 1 µM Trp-P-1, 1.70/106 nt; 30 µM Trp-P-1, 2.25/106 nt; and N-hydroxy-Trp-P-1, 4.43/106 nt (Figure 5B, C and D). The DNA adduct pattern of these three samples were almost the same, and the major spot was considered to be the adduct formed with N-O-acetyl-Trp-P-1, which is known to be the ultimate metabolite of N-hydroxy-Trp-P-1. In agreement with this, the highest spot intensity was obtained in the sample from cells treated with CYP 1A1-activated Trp-P-1 metabolite. However, apoptosis was not induced in RL-34 cells by the CYP 1A1-activated Trp-P-1 derivatives (data not shown), as shown previously in primary cultured rat hepatocytes (19). From these results, the formation of DNA adducts, the main type of DNA damage contributing to mutagenesis or carcinogenesis induced by Trp-P-1, would not trigger apoptosis.



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Fig. 5. Autoradiograms of DNA adducts in RL-34 cells treated with Trp-P-1 or N-hydroxy-Trp-P-1. RL-34 cells were treated with 1 (B) and 30 µM (C) Trp-P-1 for 24 h. Trp-P-1 (30 µM) was incubated with CYP 1A1 for 1 h, and then treated to the cells for 24 h (D). Control cells were treated with DMSO as a vehicle (A). DNA adduct formation was analyzed under the modified adduct intensification conditions as described in Materials and methods. Exposure time of the imaging plates was 1 h.

 
The apoptotic concentration of Trp-P-1 does not produce 8-OHdG
Time-related changes in the formation of 8-OHdG in DNA of the cells treated with Trp-P-1 were analyzed (Figure 6). The experiment was repeated on more than three separate occasions with similar results. The 8-OHdG level was inclined to increase by ~2-fold as compared with the control level at 96 h, when the cells were treated with 1 µM Trp-P-1. The 8-OHdG level in the control cells remained at a constant level throughout the experimental period. When the cells were treated with 30 µM Trp-P-1, the level did not change during 24 h in culture, and the level was lower than that of the cells treated with 1 µM Trp-P-1 for 96 h. Because of drastic cell death (Figure 2), we did not analyze the production of 8-OHdG at longer time points. These results suggest that the increase in level of 8-OHdG would not trigger apoptosis induced by Trp-P-1.



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Fig. 6. Formation of 8-OHdG in RL-34 cells treated with Trp-P-1. RL-34 cells were treated with 1 or 30 µM Trp-P-1 for the indicated times. Control cells were treated with 0.1% DMSO (vehicle alone). The formation of 8-OHdG was analyzed as described in Materials and methods.

 
The apoptotic concentration of Trp-P-1 introduces DSBs to RL-34 cells through the inhibition of intracellular topoisomerase I
The effect of Trp-P-1 on topoisomerase I was determined by measuring relaxation of pBR322 DNA. A 3 h treatment of Trp-P-1 decreased the formation of relaxed DNA in a dose-dependent manner (Figure 7). Supercoiled pBR322 DNA was completely relaxed at a concentration of 1 µg/ml of nuclear protein extracts from the cells treated with Trp-P-1 at 1 µM or lower (Figure 7, lanes 2, 6 and 10) and partially relaxed at concentrations of 0.1 (lanes 3, 7 and 11) and 0.01 µg/ml (lanes 4, 8 and 12). This relaxation was inhibited in the nuclear extracts (1 µg protein) from the cells treated with 10–60 µM Trp-P-1 (Figure 7, lanes 14, 18 and 22).



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Fig. 7. Inhibition of topoisomerase I in RL-34 cells treated with Trp-P-1. Nuclear extracts were prepared from RL-34 cells treated with Trp-P-1 at indicated concentrations for 3 h. Supercoiled plasmid (pBR322) DNA was incubated with nuclear extracts at 10, 1, 0.1 or 0.01 µg protein/ml or without nuclear extracts as a control. Catalytic DNA relaxation assay was performed as described in Materials and methods.

 
One of the first cellular responses to the introduction of DSBs is the phosphorylation of histone H2AX in eukaryotic cells (32). After 3 h treatment of Trp-P-1, histone H2AX was phosphorylated in a dose-dependent manner (Figure 8A). This indicates that DSBs were introduced to the cells by the higher concentrations of Trp-P-1 (10 µM or higher), and those concentrations were enough to inhibit the topoisomerase I activity (Figures 7 and 8A). Moreover, the histone H2AX was transiently phosphorylated at 3 h, and de-phosphorylated thereafter, and then re-phosphorylated at 12 and 24 h in the cells treated with the apoptotic concentration of Trp-P-1 (Figure 8B). This suggests that the first phosphorylation resulted from DSBs and the second one was due to apoptotic DNA fragmentation. These results indicate that intracellular topoisomerase I was inhibited by 10 µM or higher concentrations of Trp-P-1 and this inhibition would cause DSBs followed by induction of apoptosis.



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Fig. 8. Phosphorylation of histone H2AX in RL-34 cells treated with Trp-P-1. Cell lysates were prepared from the cells treated with 0, 0.1, 1, 10, 30 and 60 µM Trp-P-1 for 3 h (A) or 30 µM Trp-P-1 for the indicated times (B). Equal amounts of proteins (30 µg) were separated by 15% SDS–PAGE followed by transfer onto the PVDF membranes. The membranes were probed with primary antibody to phospho-Histone H2AX (ser-139) and then treated with secondary antibody. Signals on the blots were detected with an ECL chemiluminescence detection kit.

 

    Discussion
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Cells respond to DNA damages by activating a complex DNA damage–response pathway that includes cell cycle arrest and apoptosis under certain circumstances. Many genotoxic agents perturb the G1 events that mediate cell cycle progression, thereby resulting in cell cycle arrest prior to entry into S-phase (3335). Functional p53 is required for the activation of G1 checkpoint, and the resulting growth arrest is thought to allow cells time to repair DNA prior to replication (30,36,37). It has been reported that high levels of DNA damage, which induce rapid accumulation of p53, evoke G1 arrest after treatment with high concentrations of camptothecin (CPT), a powerful inhibitor of topoisomerase I (38). In the cells treated with the apoptotic concentration of Trp-P-1 that inhibited topoisomerase I activity, rapid phosphorylation of p53 at ser-15 with subsequent p53 accumulation was observed resulting in a slight increase in G1 phase, suggesting that these cells might be arrested in G1 phase. After G1 arrest, the sub-G1 cell population increased at the apoptotic concentration of Trp-P-1. In these cells, nuclear condensation and activation of caspase-3, which are recognized as markers of apoptosis, were observed. These results indicate that treatment with the apoptotic concentration of Trp-P-1 caused p53- and caspase-dependent apoptosis. In contrast, the cells treated with the non-apoptotic concentration (1 µM) of Trp-P-1 were accumulated in G2–M phase, instead of G1 phase. In these cells, a gradual accumulation of p53 was observed with little phosphorylation. Recent studies show that diverse chemical carcinogens including benzo[a]pyrene and acetylaminofluorene, which are recognized to damage DNA by the formation of DNA adducts, fail to induce G1 arrest (39). Moreover, mere activation of p53 induced by low levels of DNA damage after low concentrations of CPT treatment is not sufficient to induce G1 arrest (38). Therefore, the degree of DNA damages elicited by the non-apoptotic concentration of Trp-P-1 under our experimental conditions may be insufficient to trigger p53-mediated cell cycle responses and apoptosis.

Trp-P-1 induces various types of DNA damages including the formation of DNA adducts, the production of ROS that causes oxidative DNA damages and the inhibition of topoisomerase I resulting in DSBs. In the cells treated with the non-apoptotic concentration of Trp-P-1 for 24 h, Trp-P-1 might be metabolized by endogenous CYP 1A or 1B, and the metabolically activated forms such as N-hydroxy-Trp-P-1 caused the formation of DNA adducts. In addition, the N-hydroxy form is highly electrophilic, and unstable. ROS might be produced during spontaneous degradation of N-hydroxy-Trp-P-1 with an increase in culture times. Eventually, the formation of 8-OHdG increased at 96 h. The inhibition of topoisomerase I could also occur, although the non-apoptotic concentration of Trp-P-1 did not inhibit this enzyme in the supercoiled DNA relaxation assay. On the other hand, in the cells treated with the apoptotic concentration of Trp-P-1 for 24 h, the formation of DNA adducts was also observed with almost the same level that was in the cells treated with the non-apoptotic concentration of Trp-P-1, although the production of 8-OHdG was not observed during 24 h in culture. Interestingly, treatment with the apoptotic concentration of Trp-P-1 inhibited intracellular topoisomerase I activity and caused apparent phosphorylation of histone H2AX, which occurs after introduction of DSBs to DNA. Intact Trp-P-1 might inhibit topoisomerase, because most of the Trp-P-1 would remain in the apoptotic cells, although a little Trp-P-1 was metabolized resulting in the formation of DNA adducts. These observations are consistent with our previous reports (19,28) that the apoptogenic potency of Trp-P-1 was due to the intact molecule and not to metabolically activated derivatives. Taken together, higher concentrations of Trp-P-1 are required to induce apoptosis via inhibition of topoisomerase I while lower concentrations induce considerable levels of DNA adduct leading to carcinogenesis. Therefore, it is suggested that the apoptotic effect of Trp-P-1 is unrelated to the induction of tumors, because exposure levels in carcinogenesis experiments are much lower and even more so the human dietary exposure levels.

The fate of cells, i.e. whether they undergo apoptosis or survive, seems to be dependent on the intensity of DNA damages and the ability to repair the damages. If some DNA damages cannot be repaired completely, the remaining damages may result in carcinogenesis, but severe damages may elicit apoptotic cell death. It has been reported that N-hydroxylated 2-amino-1-methyl-6-phenylimidazo [4,5-b]pyridine (PhIP) activates the S-phase checkpoint in TK6 cells, yet eludes G1- and G2–M-phase checkpoints, accompanied by increased apoptosis and gene mutations caused by the formation of DNA adducts (15). Although the DNA adduct formation-dependent apoptotic mechanism induced by PhIP may be different from the topoisomerase I inhibition-dependent mechanism induced by Trp-P-1, the determination to undergo apoptosis or not may be dependent on the degree of DNA damages such as DSBs. Thus, it is important in future to determine the number of DNA lesions in the apoptotic cells after treatment with various DNA-damaging agents.


    Acknowledgments
 
Supported by a Grant-in-Aid for Scientific Research (c) (No. 11660126) from the Ministry of Education, Culture, Sports, Science and Technology of Japan to H.A.


    References
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received November 9, 2003; revised January 28, 2004; accepted February 4, 2004.





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