Justicidin A decreases the level of cytosolic Ku70 leading to apoptosis in human colorectal cancer cells

Jenq-Chang Lee, Chao-Hung Lee 2, Chun-Li Su 3, {dagger}, Chung-Wei Huang 1, Hsiao-Sheng Liu 1, {dagger}, Chun-Nan Lin 4 and Shen-Jeu Won 1, *

Department of Surgery and 1 Department of Microbiology, College of Medicine, National Cheng Kung University, Tainan, Taiwan, 2 Department of Pathology and Laboratory Medicine, Indiana University School of Medicine, Indianapolis, IN, USA, 3 Department of Nursing, Chang Jung Christian University, Tainan, Taiwan and 4 School of Pharmacy, Kaohsiung Medical University, Kaohsiung, Taiwan

* To whom correspondence should be addressed. Tel: +886 6 2744435; Fax: +886 6 2082705; Email: a725{at}mail.ncku.edu.tw


    Abstract
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 Abstract
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 Materials and methods
 Results
 Discussion
 References
 
The natural product justicidin A, an arylnaphthalide lignan isolated from Justicia procumbens, significantly inhibited the growth of human colorectal cancer cells HT-29 and HCT 116 at day 6 post-treatment. Further study revealed that justicidin A-treated HT-29 and HCT 116 colorectal cancer cells died of apoptosis. Justicidin A treatment caused DNA fragmentation and an increase in phosphatidylserine exposure of the cells. The number of cells in the sub-G1 phase was also increased upon justicidin A treatment. Caspase-9 but not caspase-8 was activated, suggesting that justicidin A treatment damaged mitochondria. The mitochondrial membrane potential was altered and cytochrome c and Smac were released from mitochondria to the cytoplasm upon justicidin A treatment. The level of Ku70 in the cytoplasm was decreased, but that of Bax in mitochondria was increased by justicidin A. Since Ku70 normally binds and sequesters Bax, these results suggest that justicidin A decreases the level of Ku70 leading to translocation of Bax from the cytosol to mitochondria to induce apoptosis. Oral administration of justicidin A was shown to suppress the growth of HT-29 cells transplanted into NOD-SCID mice, suggesting chemotherapeutic potential of justicidin A on colorectal cancer cells.

Abbreviations: cyto c, cytochrome c; DFF, DNA fragmentation factor; {Delta}{psi}m, mitochondrial membrane potential; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide; PARP, poly(ADP-ribose) polymerase; PBMCs, human peripheral blood mononuclear cells; PI, propidium iodide; RACK1, receptors for activated C-kinase; s.c., subcutaneously; Smac, second mitochondria-derived activator of caspase/direct IAP binding protein with low pI; XIAP, X-linked apoptosis-inhibiting protein


    Introduction
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 Abstract
 Introduction
 Materials and methods
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Colorectal cancer is a leading cause of death in both men and women (1). Chemotherapeutic agents such as 5-fluorouracil and leucovorin have been widely used postoperatively in colorectal cancer patients with regional lymph node metastasis to reduce the risk of distant metastasis (2). Oxaliplatin, irinotecan, or a combination of these drugs with 5-fluorouracil have been used to control distant metastasis (2). Unfortunately, the death rate of patients with colorectal cancer remains very high (3). Therefore, a continual search for potential anticancer agents is necessary. The crude extract of the plant Justicia procumbens is commonly used in Taiwan to treat pain, fever and inflammation (4,5). Recently, five 2,3-naphthalide lignans including justicidin A, justicidin E, neojusticin A, neojusticin B and diphyllin have been isolated from the plant (6). Among these, justicidin A has been shown to have an anti-cancer activity (79), but its mode of action is unknown. One possibility is that justicidin A induces apoptosis.

Apoptosis is characterized by morphological changes, DNA fragmentation, phosphatidylserine externalization, and generation of apoptotic bodies (10). Apoptosis can be induced by a variety of stimuli including radiation, tumor necrosis factor, and certain chemotherapeutic agents (11). Mitochondria are a major target of chemotherapy-induced apoptosis in tumors (10). Upon stimulation, death-promoting factors such as cytochrome c (cyto c), second mitochondria-derived activator of caspase/direct IAP binding protein with low pI (Smac), apoptosis inducing factor (AIF) and endonuclease G are released into the cytosol (12). Regulators such as Bcl-2 and Bcl-XL are anti-apoptotic (1315); they bind to mitochondria and inhibit the release of cyto c and Smac (16,17). Ku70 is a 70 kDa subunit of the Ku complex, which plays a crucial role in DNA double-strand break repair (18). Ku70 has been shown to inhibit the translocation of the pro-apoptotic factor Bax from cytosol to mitochondria by sequestering Bax (19,20).

In this study, we investigated effects of justicidin A on the growth of two human colorectal cancer cell lines. Our results revealed that justicidin A induces apoptosis by altering the balance between Ku70 and pro- and anti-apoptotic Bcl-2 family members in the cell. We also demonstrated that justicidin A is effective against colorectal cancer cells both in vitro and in vivo.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Isolation of justicidin A
Justicia procumbens plants were collected from Chu-Shan, Nantu Hsein, Taiwan, air-dried and then chipped. Justicidin A was isolated and purified from the chipped Justicia procumbens plants, as described previously (9).

Cell lines and cell culture
Human colorectal cancer HT-29 and HCT 116, cervix carcinoma SiHa, breast adenocarcinoma MCF7, bladder carcinoma T24 and human embryonic kidney epithelial HEK293 cells were obtained from ATCC (Rockville, MD) and maintained in Dulbecco's modified Eagle's medium (DMEM; GIBCO BRL, Grand Island, NY) containing 10% heat-inactivated fetal calf serum (FCS; HyClone, Logan, UT), 2 mM L-glutamine, 100 U/ml penicillin, and 100 µg/ml streptomycin. The cells were incubated in a humidified atmosphere with 5% CO2 at 37°C. Human peripheral blood mononuclear cells (PBMCs) were isolated from freshly collected buffy coat fraction of whole blood obtained from the Tainan Blood Bank Center (Tainan City, Taiwan) by Ficoll-Paque (Farmacia, Uppsala, Sweden) density gradient, as described previously (21). The isolated PBMCs were washed three times with serum-free RPMI-1640 medium (GIBCO BRL) and then resuspended in complete DMEM culture medium (GIBCO BRL) containing antibiotics.

Animal experiments
NOD.CB17-PRKDC<SCID>/J (NOD-SCID) mice were obtained from the Animal Center of the National Cheng Kung University (NCKU, Tainan, Taiwan). They were bred and housed at the Animal Center in a pathogen-free, temperature-controlled and air-conditioned environment with a 10/14 h light/dark cycle. Mice, 6–7 weeks old, were used. Food and water were provided ad libitum. All animal experiments were approved by the Animal Research Committee of NCKU and were performed under the guidelines of the National Research Council, Taiwan. Tumor cells were implanted subcutaneously (s.c.) to the flank of mice. Tumor growth was measured with a caliper 2–3 times per week. Tumor volume was calculated by using the formula (L x W2)/2, where L (length) and W (width) are in millimeters and L > W (22).

Reagents
Most chemicals were obtained from Sigma Chemical Co. (St Louis, MO) unless otherwise indicated. The caspase-9 inhibitor Z-LEHD-fmk was purchased from Santa Cruz Biotech (Santa Cruz, CA). Glycine and protein assay reagents were obtained from Bio-Rad Laboratories (Hercules, CA). Polyvinylidene fluoride membrane for western blot was purchased from Millipore (Bedford, MA). DiOC6(3) was purchased from Molecular Probes (Eugene, OR). Antibodies against various proteins were acquired from the following vendors: anti-caspase-3 mouse monoclonal and anti-Smac rabbit polyclonal antibodies, IMGENEX (San Diego, CA); anti-caspase-9 mouse monoclonal antibody, Upstate (Lake Placid, NY); anti-cyto c mouse monoclonal antibody, BD Pharmingen (San Diego, CA); anti-X-linked apoptosis-inhibiting protein (anti-XIAP) and anti-receptors for activated C-kinase (anti-RACK1) mouse monoclonal antibodies, BD Transduction Laboratories (Lexington, KY); anti-poly(ADP-ribose) polymerase (anti-PARP) rabbit polyclonal antibody, Cell Signaling (Beverly, MA); anti-Ku70, anti-Bcl-2, anti-Bax and anti-Bcl-XL mouse monoclonal antibodies, anti-caspase-8, Anti-DNA fragmentation factor (DFF)-45 and anti-DFF-40 rabbit polyclonal antibodies and goat anti-mouse conjugated HRP secondary antibody, Santa Cruz Biotech; goat anti-rabbit conjugated HRP secondary antibody, Amersham Pharmacia Biotech (Piscataway, NJ). Chemiluminescence reagents were obtained from NEN Life Science (Boston, MA).

Cell viability assay
Cytotoxicity of justicidin A was determined by a modified MTT [3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide] colorimetric assay, as described previously (23,24). Briefly, cells (1 x 103 cells/well) were grown overnight in 96-well plates (Nunc, Denmark) in 100 µl culture medium and then treated with different concentrations of justicidin A dissolved in 100% DMSO. Control cells were treated with DMEM alone or DMEM containing 0.1% DMSO. At different time points, 10 µl of MTT was added to each well to a final concentration of 0.5 mg/ml. The water-soluble MTT was taken up by live cells and converted to an insoluble purple formazan. After 4 h of incubation at 37°C in the dark, 100 µl of 10% SDS/0.01 N HCl was added to each well and incubated at 37°C overnight to dissolve the formazan. The amount of formazan in each sample thus produced was quantified by measuring the absorbance of light at 590 nm with a Multiscan photometer (MRX II, Dynatech, McLean, VA). All experiments were repeated three times and each drug concentration was tested in octuplicate. Viability of drug-treated cells was expressed as a percentage of population growth with standard error of the mean (SEM) relative to that of untreated control cells. Cell death caused by justicidin A was calculated as a percentage of inhibition as follows: % inhibition = (1 – mean experimental absorbance/mean control absorbance) x 100.

Colony formation assay
Cells (6 x 104 cells/well) in 0.1 ml medium were mixed with 0.9 ml of 0.33% agar in DMEM (GIBCO BRL) containing 10% FCS (HyClone) in the presence or absence of various concentrations of justicidin A at 37°C and then layered on top of 1 ml of 0.6% solid basal agar in six-well trays (Nunc). Colonies with a diameter >1 mm were counted 14 days later (25).

Analysis of DNA fragmentation
Justicidin A-treated or untreated cells were lysed by incubation in 200 µl lysis buffer [10 mM Tris–HCl (pH 7.6), 20 mM EDTA and 1% NP-40] for 20 min at 37°C. The cell lysates were centrifuged to remove cell debris, and the supernatants were incubated with 4 mg/ml of RNase A and 1% SDS at 56°C for 2 h, followed by incubation with 200 µg/ml proteinase K at 37oC for 2 h. DNA fragments were precipitated by the addition of 150 µl ammonium acetate (10 M) and 1.2 ml ethanol (100%). After an overnight incubation at –20°C, the DNA was pelleted, dried and then dissolved in 15 µl Tris–EDTA buffer. The sample was then electrophoresed on a 1% (w/v) agarose gel in TBE buffer at 50 V for 1 h, and the gel was stained with ethidium bromide to visualize DNA ladders under UV light (26).

Determination of apoptosis by flow cytometry
Apoptotic cells with externalized phosphatidylserine were enumerated using a FACScan flow cytometer (Becton Dickinson, Mountain View, CA). Briefly, cells (2 x 105 cells/well) in 2 ml medium were grown in six-well plates and then treated with different concentrations of justicidin A. At different time intervals, cells were trypsinized, washed with HEPES buffer solution (HBS), resuspended in 400 µl HBS containing 5 µl Annexin V-FITC (BD Pharmingen), and then subjected to flow cytometric analysis as described previously (27). To assess apoptosis by measuring DNA contents, justicidin A treated cells were fixed with 70% ethanol at 4°C and then stained with propidium iodide (PI) by incubating the cells in HBS containing 40 µg/ml PI and 100 mg/ml RNase A for 30 min at 37°C in the dark. The PI-stained cells were sorted by flow cytometry based on their DNA contents. Results were analyzed with the Windows Multiple Document Interface software for Flow Cytometry (WinMDI 2.8, Scripps Research Institute, San Diego, CA). Another event in apoptosis is change in {Delta}{psi}m. To assess {Delta}{psi}m, justicidin A treated and non-treated cells were incubated with 40 nM of the lipophilic cationic dye DiOC6(3) and 2.5 µg/ml PI in the dark for 30 min. Since the binding of DiOC6(3) to cells varies with the {Delta}{psi}m, the intensity of DiOC6(3) staining is directly proportional to the integrity of mitochondrial membrane. DiOC6(3) stained cells were analyzed by flow cytometry with an excitation wavelength of 488 nm and emission wavelength of 529 nm as described previously (28,29).

Confocal microscopy
Measurement of {Delta}{psi}m was also performed by confocal microscopy as described previously (30). Briefly, cells (2 x 105 cells/well) in 2 ml of medium were grown in six-well plates containing sterilized glass cover slips overnight, treated with or without various concentrations of justicidin A, and then stained with 5 µM rhodamine 123 at 37°C. Rhodamine 123 is a lipophilic, positively-charged, membrane-permeable dye. It can be taken up by living cells and preferentially stains mitochondria with increasing intensity proportional to the integrity of mitochondria (31). After washing twice with PBS at room temperature to remove excess rhodamine 123, cells were fixed for 15 min in 4% paraformaldehyde and then mounted on poly-L-lysine-coated glass microscope slides. Samples were examined with a Leica TCSNT laser scanning confocal imaging system coupled to a Leica DMRBE microscope with a 630 fluotar objective. Rhodamine 123-stained cells were excited with 488-nm lines of a 25 mW laser, and the fluorescence emitted was visualized with a BF530/30 filter combination. Optical sections close to the middle of the cell were evaluated.

Preparation of subcellular fractions and western blot analysis
Whole cell lysate, cytoplasmic, mitochondrial and nuclear fractions of cells were separately analysed. Whole cell lysate was obtained by resuspending cells (1 x 106 cells) in 200 µl of lysis buffer containing 1 mM EDTA, 0.5% (w/v) SDS, 10 mM Tris–HCl (pH 7.4), 0.15 M NaCl, 1 mM EGTA, 5 µg/ml leupeptin, 5 µg/ml aprotinin, 2 mM Na3VO4, 0.5 mM phenylmethylsulfonyl fluoride (PMSF) and 1% (v/v) Triton X-100 at 4°C for 30 min. The cell debris was removed by centrifugation at 15 000 g for 10 min. To prepare cytoplasmic fractions, 1 x 107 cells were washed with ice-cold PBS and then homogenized with a Dounce homogenizer (Glas-Col, Terre Haute, IN) in 500 µl of TSE buffer [10 mM Tris, 0.25 M sucrose, 0.1 mM EDTA (pH 7.4)]. After removal of cell debris by centrifugation at 750 g for 30 min, the samples were centrifuged at 100 000 g for 1 h. The resulting supernatants were used as the cytosolic fractions, and the pellets were lysed in 100 µl lysis buffer containing 10 mM Tris (pH 7.4), 1 mM EDTA, 10 mM NaF, 1 mM Na3VO4, 2.5 mM PMSF, 1 µg/ml leupeptin, 1 µg/ml pepstatin A and 0.25 mM sucrose and used as the mitochondrial fraction as described previously (32,33). To prepare nuclear extracts, 1 x 107 cells were washed twice with ice-cold PBS and then lysed in 400 µl of buffer A [10 mM HEPES (pH 7.9), 5 mM MgCl2, 10 mM KCl, 3 mM Na3VO4, 10 mM NaF, 0.5 mM dithiothreitol (DTT), 0.5 mM PMSF, and 2 µg/ml of leupeptin, antipain, aprotinin, and pepstatin A] on ice for 20 min. The nuclei were pelleted by centrifugation at 11 000 g for 20 s at 4°C and then resuspended in 60 µl of buffer B (20 mM HEPES, pH 7.9, 1.5 mM MgC12, 420 mM NaCl, 0.2 mM EDTA, 25% glycerol, 1 mM Na3VO4, 10 mM NaF, 0.5 mM DTT, 0.5 mM PMSF and 1 µg/ml each of leupeptin, antipain, aprotinin, and pepstatin A) for 15 min on ice with occasional mixing. Nuclear debris was removed by centrifugation at 12 000 g for 15 min at 4°C. Western blotting was performed as described previously (34).

Statistical analysis
Significant difference in tumor volume was determined by the Student's t-test using the Minitab (version 10.2) software package. A difference was considered significant if P < 0.05.


    Results
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 Materials and methods
 Results
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 References
 
Growth inhibition of human colorectal cancer cells by justicidin A
The inhibitory activity of justicidin A on the proliferation of human colorectal cancer cells (HT-29 and HCT 116), cervix carcinoma cells (SiHa), breast adenocarcinoma cells (MCF7) and bladder carcinoma cells (T24) was investigated. Cells were grown in the absence or presence of various concentrations (0.001–40 µM) of justicidin A for 6 days. MTT assays were then performed, and the 50% inhibitory concentrations (IC50) for HT-29, HCT 116, SiHa, MCF7 and T24 cells were determined to be 0.110, 0.400, 0.020, 1.540 and 0.004 µM, respectively (Table I). The effect of justicidin A on colony formation of these cancer cells in soft agar was also assessed. At day 14 post-treatment, justicidin A suppressed colony formation of HT-29, HCT 116, SiHa, MCF7 and T24 cells by 50% at concentrations of 0.030, 0.100, 0.080, 1.060 and 0.003 µM, respectively (Table I). The IC50 of justicidin A for human embryonic kidney cells HEK293 was ~100-fold higher (IC50 = 10.6 µM) than that for HT-29 cells. PBMCs were much more resistant to justicidin A with an IC50 of 25 µM. These results suggest that justicidin A preferentially inhibits the growth of cancer cells. Since colorectal cancers are one of the most common cancers, subsequent studies were focused on HT-29 and HCT 116 cells. A daily evaluation of justicidin A on growth inhibition of these cancer cells revealed that justicidin A inhibited cell growth in a dosage- and time-dependent manner. At a concentration of 2.5 µM, it inhibited the proliferation of HT-29 cells ~25% at day 1 and 82% at day 6 post-treatment (Figure 1A). A similar pattern of growth inhibition was observed in HCT 116 cells (Figure 1B). However, lower concentration (0.625 µM) of justicidin A had a better cytotoxic effect on HT-29 (Figure 1A) than HCT 116 (Figure 1B).


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Table I. Effects of justicidin A response for 50% growth inhibition (IC50), colony formation on human tumor cell lines, and human PBMCsa

 


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Fig. 1. Growth inhibition of justicidin A on colorectal cancer cells. (A) HT-29 or (B) HCT 116 cells (1 x 103 cells/well) in 0.1 ml medium were grown in 96-well plates and then treated with justicidin A at the indicated concentrations for various lengths of time. Growth inhibition was determined by the MTT assay every day for 6 days.

 
Induction of apoptosis in colorectal cancer cells by justicidin A
To determine whether justicidin A induced apoptosis, cells were treated with the drug (0.75 µM for HT-29 and 5 µM for HCT 116) and then assayed for signs of apoptosis at 0, 12, 16, 20, 24, 48 and 72 h post-treatment. The treated cells were stained with annexin V-FITC and then analysed by flow cytometry. As seen in Figure 2A, the percentage of annexin V positive HT-29 cells was increased with time from 2.8% at time 0 to 6.3% at 12 h and reached 75.4% at 72 h after treatment of justicidin A (0.75 µM). A similar pattern of increase (2.5% at time 0 to 10.4% at 12 h, and 78.2% at 72 h post-treatment) in annexin V positive HCT 116 cells was observed when they were treated with 5 µM of justicidin A. To confirm apoptosis, cells were stained with PI, and the relative number of cells with a sub-G1 DNA content was determined by flow cytometry. As seen in Figure 2B, treatment of HT-29 cells with 0.75 µM of justicidin A increased the population of cells with a sub-G1 DNA content from 1% at time 0 to 37.2% at the 72 h time point. The population of sub-G1 HCT 116 cells increased with time from 1.1% at time 0 to 19.8% at 96 h after treatment of 5 µM of justicidin A (Figure 2C). This increase in the population of sub-G1 cells was also dosage dependent, ranging from 2.2% in non-treated to 9.5% in those treated with 0.25 µM of justicidin A and to 14.5, 17.7 and 19.2% in those treated with 0.5, 0.75 and 1 µM, respectively, for 48 h (Figure 2B). Treatment of HCT 116 cells with 0–10 µM of justicidin A showed a similar dosage-dependent response pattern (Figure 2C). The population of sub-G1 HCT 116 cells was increased from 2.4% in non-treated to 20% in those treated with 10 µM of justicidin A for 72 h (Figure 2C). To further confirm that justicidin A-treated cells undergo apoptosis, DNA fragmentation in these cells was examined and all drug-treated cells were found to have a certain degree of DNA fragmentation (Figure 2D).




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Fig. 2. Justicidin A-induced apoptosis in colorectal cancer cells. HT-29 or HCT 116 cells (2 x 105 cells/well) in 2 ml medium were grown in 6-well dishes and then treated with justicidin A (0.75 µM for HT-29 cells and 5 µM for HCT 116 cells) for the indicated time periods, or with indicated concentrations of justicidin A for 48 h (HT-29 cells) or 72 h (HCT 116 cells). Cells were then harvested and stained with annexin V-FITC (A) or PI (B and C) and analysed by flow cytometry. In (A), the majority of non-treated cells were normal with no or very weak annexin V-FITC staining (intensity units <5 x 101). When the cells were exposed to justicidin A, the number of annexin V-FITC-stained cells with a staining intensity ranging from 5 x 101 to 3 x 103 units was increased with time. In (B and C), cells that had a PI-staining intensity ~200 units were in the G1 phase, and those with an intensity ~400 were in the G2–M phase. Cells with a PI-staining intensity <200 units were in sub-G1 phase and were apoptotic. The number of apoptotic HT-29 (B) or HCT 116 (C) cells was increased in a time- and dose-dependent manner upon justicidin A treatment. DNA fragmentation of justicidin A treated cells are shown in panel (D). Total DNA of vehicle- (0.1% DMSO in culture medium) or justicidin A-treated cells was extracted and electrophoresed on a 1% agarose gel. Lanes 1 and 7: molecular weight marker (M); lane 2: DNA from vehicle-treated HT-29 cells; lane 8: DNA from vehicle-treated HCT 116 cells; lanes 3–6 and 9–12: DNA from HT-29 or HCT 116 cells treated with indicated concentrations of justicidin A. Justicidin A was diluted with culture medium containing 0.1% DMSO. Results are representative of three independent experiments. JA, justicidin A.

 
Effects on caspases and its target proteins by justicidin A
Since the major enzyme involved in apoptosis is caspase-3, activation of caspase-3 in justicidin A-treated cells was examined. HT-29 and HCT 116 cells were treated with 0.75 and 5 µM of justicidin A, respectively. At different time points after treatment, cells were assessed for the occurrence of activated caspase-3 by western blotting. Both the 12 and 17 kDa forms of activated caspase-3 were first detected 24 h after justicidin A treatment in both cell lines, and more activated caspase-3 was detected with a longer drug treatment (Figure 3A).



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Fig. 3. Justicidin A induced the cleavage of caspases and their target proteins in human colorectal cancer cells. Caspase-3 (A), caspase-9 (B), PARP (D), and DFF (E) in HT-29 and HCT 116 cells treated with justicidin A (0.75 µM for HT-29 cells and 5.0 µM for HCT 116 cells) were assayed by western blot analysis. At indicated time points, whole cell, cytosolic and nuclear lysates were prepared and electrophoresed on 12% SDS–PAGE gels. Protein bands on the gels were transferred to polyvinylidene fluoride membranes. Anti-caspase-3, anti-caspase-9, anti-89 kDa PARP, anti-DFF-45 or DFF-40 antibodies were then used to detect respective proteins. (C) Inhibitory effect of the caspase-9 inhibitor Z-LEHD-fmk on justicidin A-induced apoptosis in HT-29 and HCT 116 cells was also examined. Cells were pretreated without or with 20 µM Z-LEHD-fmk for 4 h prior to exposure to justicidin A (0.75 µM for HT-29 cells and 5.0 µM for HCT 116 cells) for 30 h. Cells were then stained with PI and analysed by flow cytometry. RACK1 (receptors for activated C-kinase) was similarly assessed to serve as a loading control. Justicidin A was diluted in culture medium containing 0.1% DMSO. Control cells were treated with medium containing 0.1% DMSO. Results are representative of three independent experiments. JA, justicidin A; Casp, caspase.

 
To determine whether justicidin A-induced apoptosis was mediated by the extrinsic or intrinsic pathway, activation of caspase-8 or caspase-9 was examined. Caspase-9 activation would implicate an intrinsic pathway, whereas caspase-8 activation would suggest the extrinsic pathway. HT-29 and HCT 116 cells were again treated with 0.75 and 5 µM of justicidin A, respectively. If the caspase-9 is activated, the pro-caspase-9 is cleaved to yield two peptides of 35 and 37 kDa. As shown in Figure 3B, these two activated forms became visible 6 h after treatment in HT-29 cells and 12 h in HCT 116 cells and reached the highest level at the 72 h time point (Figure 3B). In contrast, activated caspase-8 was not detected in justicidin A-treated HT-29 or HCT 116 cells (data not shown). To confirm the involvement of caspase-9 in justicidin A-induced apoptosis, the caspase-9 inhibitor Z-LEHD-fmk was used. HT-29 and HCT 116 cells were treated with 0.75 and 5 µM of justicidin A, respectively, with or without pretreatment with 20 µM of Z-LEHD-fmk for 4 h. The cells were then stained with PI and assayed for those arrested in the sub-G1 phase by flow cytometry. As seen in Figure 3C, only 3.8% of HT-29 cells were found to undergo apoptosis without justicidin A treatment. The population of apoptotic HT-29 cells was increased to 35.3% upon justicidin A treatment. Pre-treatment of the cells with Z-LEHD-fmk reduced justicidin A-induced apoptosis rate to 8.8%. Z-LEHD-fmk treatment alone had no effect on the apoptosis of HT-29 cells (Figure 3C). Similar results were observed in HCT 116 cells; Z-LEHD-fmk was found to reduce the apoptosis rate caused by justicidin A from 25.5 to 6.5% in HCT 116 cells (Figure 3C).

Since activated caspase-3 cleaves PARP, experiments were performed to detect the cleaved form of PARP which has a molecular weight of 89 kDa. The cleaved PARP was first detected in both total cell lysate and nuclear fraction at 12 h after justicidin A treatment in both HT-29 and HCT 116 cells, and the amount of the 89 kDa PARP was increased in a time-dependent manner (Figure 3D). Activated caspase-3 also digests DFF-45 and DFF-35, both of which bind to DFF-40 and prevent it from entering the nucleus to cause fragmentation of nuclear DNA (35,36). Therefore, reduced levels of DFF-45 and DFF-35 are an indication of caspase-3 activation. As shown in Figure 3E, treatment of HT-29 cells with 0.75 µM of justicidin A caused a decrease in DFF-45 and DFF-35 levels at 12 h. The cytosolic levels of DFF-45 and DFF-35 continued to decrease and reached the lowest level at 72 h. In HCT 116 cells, the decrease in the level of DFF-45 was not as profound as that seen in HT-29 cells, but a dramatic decrease in DFF-35 level was observed 72 h after treatment with 5 µM of justicidin A (Figure 3E). While the levels of cytosolic DFF-45 and DFF-35 were decreased, the level of DFF-40 in the nuclear fraction of justicidin A-treated cells was increased in a time dependent manner. The increase in the level of nuclear DFF-40 was first observed at 6 h in HT-29 cells and 12 h in HCT 116 cells and peaked at 72 h in HT 29 cells and at 24–48 h in HCT 116 cells upon justicidin A treatment (Figure 3E).

Damage in mitochondria caused by justicidin A treatment
Activation of caspase-9 suggests that justicidin A triggers apoptosis by the intrinsic pathway which affects mitochondria. Therefore, the change in the membrane potential of mitochondria ({Delta}{psi}m) in response to justicidin A treatment was examined by flow cytometry after staining the cells with mitochondrial dye DiOC6(3) and PI. A time-dependent decrease in the intensity of DiOC6(3) staining was observed in the mitochondria of justicidin A-treated HT-29 (0.75 µM justicidin A) and HCT 116 (5 µM justicidin A) cells. The decrease in {Delta}{psi}m was first observed at 12 h and reached the lowest level at 72 h after justicidin A treatment in both cell lines (Figure 4A). Treatment of HT-29 cells with various concentrations of justicidin A (0.25–1 µM) for 48 h resulted in a dose-dependent loss of {Delta}{psi}m (Figure 4A). The decrease in the intensity of DiOC6(3) staining was found to be accompanied by an increase in PI staining (Figure 4A). Since PI does not penetrate the cell membrane of live cells, the increase in PI staining indicates an increase in the number of dead or apoptotic cells. The loss in {Delta}{psi}m upon justicidin A treatment was also examined by confocal microscopy and rhodamine 123 which preferentially stains intact mitochondria. Consistent with the data obtained from flow cytometry, a significant decrease in the intensity of rhodamine 123 staining was observed at 12 h. The staining intensity continued to decrease and reached the lowest level at 72 h after justicidin A treatment in both cell lines (Figure 4B and C). A justicidin A dose-dependent loss in rhodamine 123 staining in both cell lines was also observed (Figure 4B and C). There was no change in {Delta}{psi}m in vehicle-treated cells (Figure 4). Interestingly, addition of 10 µM cyclosporin A to the cell cultures 4 h prior to justicidin A treatment prevented the decrease in the intensity of rhodamine 123 staining and JA-induced apoptosis measuring by PI flow cytometry (Figure 4D). Since cyclosporine A hyperpolarizes the mitochondrial membrane potential ({Delta}{psi}m) by binding to cyclophilin D thus preventing the conversing of adenine nucleotide translocase (ANT) to permeability transition (PT) pores, these results indicate that justicidin A indeed causes a loss in {Delta}{psi}m.




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Fig. 4. Effect of justicidin A on {Delta}{psi}m in human colorectal cancer cells. HT-29 and HCT 116 cells (2 x 105 cells/well) in 2 ml medium were grown in six-well and then treated with justicidin A (0.75 µM for HT-29 cells and 5.0 µM for HCT 116 cells) as described elsewhere. At indicated time points, cells were stained with 40 nM of DiOC6(3) and then analysed by flow cytometry (A) or imaged by confocal microscopy after staining with rhodamine 123 (B and C). Normal cells had a DiOC6(3) staining intensity of 103–104 units (lower right quadrant). Justicidin A treatment damaged mitochondria and reduced the intensity of DiOC6(3) staining, thus shifting more cells to the lower left quadrant. Apoptotic cells had a PI staining intensity of ~102 units. More cells with this staining intensity (upper two quadrants) were seen upon justicidin A treatment. Inhibitory effects of cyclosporin A on justicidin A-induced change in {Delta}{psi}m and apoptosis in HT-29 and HCT 116 cells were also examined (D). Cells were pretreated with 10 µM cyclosporin A for 4 h prior to exposure to justicidin A (0.75 µM for HT-29 cells and 5.0 µM for HCT 116 cells) for 30 h. Cells were then stained with rhodamine 123 and examined by confocal microspcopy or stained with PI and then analysed by flow cytometry. Justicidin A was diluted in culture medium containing 0.1% DMSO. Control cells were treated with medium containing 0.1% DMSO. Results are representative of three independent experiments. JA, justicidin A.

 
Release of cyto c and Smac from mitochondria by justicidin A treatment
To further confirm that justicidin A induces apoptosis by affecting mitochondria, release of cyto c and Smac from mitochondria to the cytoplasm was examined by immunoblotting. The cytosolic levels of both cyto c and Smac were found to be profoundly increased (1.7- and 1.4-fold, respectively) in HT-29 cells at 24 h and reached peak levels (2.3–2.7 and 2.6-fold, respectively) at 48–72 h following the treatment with justicidin A (0.75 µM) (Figure 5A–D). Conversely, levels of both cyto c and Smac in the mitochondria-enriched fractions of HT-29 cells were decreased within 24 h (Figure 5A–D) and became undetectable at 96 and 72 h, respectively, after justicidin A treatment (Figure 5A–D). Treatment of HCT 116 cells with 5 µM of justicidin A resulted in similar changes in cyto c and Smac levels. Cytosolic cyto c level was greatly increased (7.2-fold) 96 h and that of Smac was increased (2.7-fold) 24 h after drug treatment (Figure 5A–D). The mitochondrial cyto c and Smac levels were decreased starting 24 and 48 h, respectively, after treatment (Figure 5A–D). Since Smac antagonizes the anti-apoptotic function of XIAP (37), the effect of justicidin A on the total level of XIAP was also examined. As shown in Figure 5E and F, XIAP levels were profoundly decreased at 24 h in both HT-29 (0.75 µM of justicidin A) and HCT 116 cell lines (5 µM of justicidin A) and reached the lowest levels at 48 h in HT-29 cells and at 96 h in HCT 116 cells. In addition, XIAP levels were reduced in a justicidin A dose-dependent manner (0.25–10 µM) in both cell lines (Figure 5E).




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Fig. 5. Distribution of cyto c and Smac and levels of XIAP in justicidin A-treated cells. Cyto c (A), Smac (B) and XIAP (E) levels in HT-29 and HCT 116 cells after justicidin A treatment were determined by western blot analysis. HT-29 or HCT 116 cells were grown and treated with justicidin A as described in the Figure 2 legend. At indicated time points, whole cell lysate, cytosolic and mitochondria-enriched fraction were prepared and electrophoresed on 12% SDS–PAGE gels. Protein bands on the gel were then transferred to polyvinylidene fluoride membranes. The intensity of each protein band was quantified by desitometry normalizing to that of RACK1 (receptors for activated C-kinase). The density values of cyto c, Smac and XIAP from control conditions were designated as 1. The levels of these proteins in the remaining samples were expressed as fold of the control (C), (D) and (F). Anti-cyto c, anti-Smac and anti-XIAP antibodies were used to detect respective proteins on the blots. RACK1 (receptors for activated C-kinase) served as the loading control. Justicidin A was diluted in culture medium containing 0.1% DMSO. Control cells were treated with medium containing 0.1% DMSO. Results are representative of three independent experiments. JA, justicidin A.

 
Translocation of Bax from the cytosol to mitochondria
A characteristic feature of mitochondria-mediated apoptosis is the translocation of the pro-apoptotic factor Bax from the cytosol to mitochondria. To confirm the involvement of mitochondria in this justicidin A-caused apoptosis, the levels of Bax in different cellular fractions were determined. The total cellular level of Bax was found to be slightly increased (1.34-fold) 6 h after justicidin A treatment and reached the peak level (2-fold) at 12 h in HT-29 cells (Figure 6A and B). In HCT 116 cells, the total Bax level was greatly increased (2-fold) at 24 h and reached peak level (4.5-fold) at 72 h post-treatment (Figure 6A and B). The level of Bax in mitochondria-enriched fraction was greatly increased (3.4-fold) within 6 h and peaked (4-fold) at 24 h in justicidin A-treated HT-29 cells (Figure 6A and B). A dramatic increase (4.34-fold) in mitochondrial Bax level was observed after 72 h of treatment in HCT 116 cells (Figure 6A and B). In contrast, the amount of cytosolic Bax in both cell lines was decreased upon justicidin A treatment in a time-dependent manner (Figure 6A). Contrary to the increase of the pro-apoptotic Bax in mitochondria, the level of the anti-apoptotic factor Bcl-XL in mitochondria was decreased upon justicidin A treatment in both HT-29 and HCT 116 cells, and a longer treatment resulted in a more severe decrease in the level of Bcl-XL in both cell lines (Figure 6C). The total cellular level of Bcl-XL was also found to be decreased by justicidin A treatment in a time-dependent manner (Figure 6C).



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Fig. 6. Effect of justicidin A on the levels of pro- and anti-apoptotic proteins of the Bcl-2 family. The levels of Bax (A) and Bcl-XL (C) in HT-29 and HCT 116 cells treated with justicidin A (0.75 µM for HT-29 cells and 5.0 µM for HCT 116 cells) were determined by western blot analysis. At indicated time points, whole cell, cytosolic and mitochondria-enriched lysates were prepared and electrophoresed on 12% SDS–PAGE gels. Protein bands on the gels were then transferred to polyvinylidene fluoride membranes. The intensity of each protein band was quantified by desitometry normalizing to that of RACK1 (receptors for activated C-kinase). The density value of Bax from control conditions was designated as 1. The levels of Bax in the remaining samples were expressed as fold of control (B).

 
Decrease in the level of Ku70 caused by justicidin A
To investigate whether Ku70 was involved in justicidin A-induced mitochondrial dysfunction in human colorectal cancer cells, the level of the Ku70 protein was determined by immunoblot analysis. As seen in Figure 7A and B, the level of cytosolic Ku70 was decreased (0.62-fold) as early as 2 h and reached the lowest level (0.37-fold) at 12 h in HT-29 cells after treatment with 0.75 µM of justicidin A. Similarly, the amount of cytosolic Ku70 in HCT 116 cells was profoundly decreased (0.6-fold) at 48 h and reached the lowest level (0.33-fold) at 96 h after justicidin A (5 µM) treatment in HCT 116 cells (Figure 7A and B). In contrast, no change in the level of Ku70 was observed in nuclear fractions of both untreated- and justicidin A-treated cells (Figure 7A).



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Fig. 7. Effect of justicidin A on the levels of the Ku70 protein. The levels of Ku70 (A) in HT-29 and HCT 116 cells treated with justicidin A (0.75 µM for HT-29 cells and 5.0 µM for HCT 116 cells) were determined by western blot analysis. At indicated time points, cytosolic and nuclear-enriched lysates were prepared and electrophoresed on 12% SDS–PAGE gels. Protein bands on the gels were then transferred to polyvinylidene fluoride membranes. The intensity of each protein band was quantified by desitometry normalizing to that of RACK1 (receptors for activated C-kinase). The density value of Ku70 from control conditions was designated as 1. The levels of Ku70 in the remaining samples were expressed as fold of control (B). Anti-Ku70 antibody was used to detect the proteins. RACK1 served as a loading control. Justicidin A was diluted with culture medium containing 0.1% DMSO. Control cells were treated with medium containing 0.1% DMSO. Results are representative of three independent experiments. JA, justicidin A.

 
Suppression of justicidin A on the growth of human colorectal cancer cells in mice
The possibility that justicidin A can inhibit the growth of HT-29 cells implanted in NOD-SCID mice was also examined. Mice were inoculated with HT-29 cells s.c. into the flank on day 0 and then randomly divided into three groups on day 4. One group (n = 5) of mice received oral administration of justicidin A (6.2 mg/kg) once a day for 56 consecutive days, and the other group (n = 5) received 10.6 mg/kg of justicidin A on the same schedule. The control group (n = 5) received the vehicle (0.05% DMSO). Tumor volume and body weight were recorded daily from day 4 (treatment start) until day 60 (treatment stop). Tumor-bearing mice treated with 10.6 or 6.2 mg/kg justicidin A showed a dramatic suppression of tumor growth and decrease in tumor weight (P < 0.05) as compared with vehicle-treated mice (Figure 8A and B and Table II). None of the justicidin A-treated or vehicle-treated mice showed significant changes in spleen or liver weight throughout the experiment (Table II).



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Fig. 8. Inhibitory effect of justicidin A on the growth of HT-29 cells xenografted into NOD-SCID mice. (A) HT-29 cells were implanted s.c. to the flank of mice. Four days after implantation, justicidin A (10.6 mg/kg, inverted triangle; 6.2 mg/kg, open circle) was orally administrated every day until day 60. (B) Photograph of mice bearing HT-29 human colorectal cancer cells at day 60 after oral administration of vehicle or justicidin A (10.6 mg/kg). Arrow indicates the time when justicidin A treatment was initiated. Data are presented as means ± SEM from two separate experiments (n = 5 in each group). Data points marked with asterisks indicate significant difference between experimental and control groups with a P-value < 0.05. JA, justicidin A.

 

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Table II. Effect of dietary justicidin A on tumor weight, body weight, liver weight and spleen weighta

 

    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
In this study, we found that justicidin A is toxic to many different types of cells, including human colorectal cancer (HT-29 and HCT 116), cervix carcinoma (SiHa), breast adenocarcinoma (MCF7), bladder carcinoma (T24), human embryonic kidney epithelial cells (HEK293) and PBMCs. Interestingly, all cancer cells tested in this study were particularly sensitive to justicidin A. For example, the IC50 for T24 bladder carcinoma cells was 0.004 µM which is 6250-fold lower than that for PBMCs (25 µM) (Table I). This result suggests that justicidin A can be used for chemotherapy for cancers. Since colorectal cancers are one of the most prevalent cancers, we examined the effects of justicidin A on two different colorectal cancer cell lines (HT-29 and HCT 116). IC50's for HT-29 and HCT 116 were determined to be 0.11 and 0.4 µM, respectively. Justicidin A is believed to kill HT-29 and HCT 116 cells by inducing apoptosis based on the observation that justicidin A treatment caused caspase-3 and caspase-9 to become activated (Figure 3A and B). Changes in {Delta}{psi}m and release of apoptogenic proteins such as cyto c and Smac from mitochondria upon justicidin A treatment suggest that this apoptosis is mediated by the intrinsic pathway in which mitochondria are affected.

Although both HT-29 and HCT 116 are colorectal cancer cells, their sensitivity to justicidin A is different. HT-29 cells are ~4-fold more sensitive than HCT 116 cells to justicidin A. However, the effects of justicidin A on these two cell lines appear to be similar. All parameters related to apoptosis in these two cell lines are similarly affected by justicidin A including activation of caspase-3 and caspase-9 (Figure 3), loss of {Delta}{psi}m (Figure 4), release of cyto c and Smac from mitochondria to the cytosol (Figure 5A and B), reduction in XIAP level (Figure 5E and F), and the translocation of Bax from the cytosol to mitochondrial (Figure 6A). Interestingly, the level of cytosolic Ku70 was found to be dramatically decreased upon treatment with justicidin A in both HT-29 and HCT 116 cells (Figure 7A). Ku70 is known to play a crucial role in apoptosis. Down regulation of Ku70 has been shown to enhance Bax-mediated apoptosis (19), whereas overexpression of Ku70 has been shown to inhibit apoptosis (38). Based on the results of this study, we propose the following mechanism of action of justicidin A in the induction of apoptosis in colorectal cancer cells: Justicidin A causes a decrease in the level of cytosolic Ku70 allowing Bax to enter mitochondria. Bax then oligomerizes and perturbs the outer mitochondrial membrane causing release of apoptogens such as Smac and cyto c as shown in Figure 5A. Justicidin A also causes a decrease in Bcl-XL level (Figure 6C). Since Bcl-XL may bind to Bax and prevent Bax from inserting into the outer membrane of mitochondria (10), the decrease in Bcl-XL level would favor Bax activation and thus the loss of {Delta}{psi}m and release of apoptogen from the mitochondria. Smac, also know as DIABLO, promotes caspase-9 activation by neutralizing members of the inhibitor of apoptosis protein family such as XIAP which was found to be decreased in justicidin A-treated cells (Figure 5E). This decrease in XIAP level enables cyto c, dATP, apoptotic protease activating factor-1 (Apaf-1), and pro-caspase-9 to form apoptosomes to activate caspase-9 (12). Activated caspase-9 in turn activates caspase-3 to cleave the 116 kDa PARP to two peptides of 89 and 24 kDa, rendering PARP unable to carry out poly-ADP-ribosylation of various proteins involved in DNA repair. In this study, the 89 kDa form of PARP was found to be increased with time upon justicidin A treatment (Figure 3D). Activated caspase-3 also digests DFF-45 and DFF-35 that are natural inhibitors of DFF-40. Degradation of DFF-45 and DFF-35 renders DFF-40 free to enter the nucleus as shown in Figure 3E to digest nuclear DNA leading to apoptosis of the cell. Treatment of HT-29 and HCT 116 cells with justicidin A caused a dose- and time-dependent increase in cytosolic cyto c and Smac (Figure 5). However, the cytosolic cyto c level in HT-29 cells treated with justicidin A for 96 h was decreased as compared with those treated for 72 h (Figure 5C). A similar unexpected decrease in cytosolic Smac was also seen in HCT 116 cells treated with justicidin A for 72 h (Figure 5D). The reason of this sudden decrease in cytosolic cyto c and Smac is not known. A possibility is that cyto c and Smac are degraded upon prolonged exposure in the cytoplasm or that they are consumed during apoptosis.

Justicidin A also has been shown to inhibit the release of tumor necrosis factor-alpha (TNF-{alpha}) from lipopolysaccharide-treated RAW 264.7 macrophages in a concentration- and time-dependent manner (39). In addition, justicidin A can inhibit the transport of TNF-{alpha} (39) and has antiviral activity (40). These observations suggest that justicidin A has multiple cellular targets. Justicidin A was shown to cause hepatocellular carcinoma Hep 3B and Hep G2 cells to die without decreasing the level of Ku70 in these cells (our unpublished data). It is possible that Ku70 is a target of justicidin A in colorectal cancer cells, but not in hepatocellular carcinoma. In contrast to cytosolic Ku70, the level of nuclear Ku70 in HT-29 and HCT 116 cells was not affected by justicidin A (Figure 7A), suggesting that justicidin A treatment activates a mechanism which degrades cytosolic Ku70. Whether justicidin A affects other factors involved in apoptosis remains to be determined.

Oral administrations of 6.2 or 10.6 mg/kg of justicidin A once a day for 56 consecutive days were found to suppress the growth of HT-29 cells transplanted in NOD-SCID mice but did not have a significant effect on spleen or liver weight (Table II). This result suggests that justicidin A at the dosage of 6.2 or 10.6 mg/kg/day has no effect on normal cells. Although more careful analyses of effects of justicidin A on various organs remain to be done, results of this study show the potential of using justicidin A for chemotherapy of colorectal cancers.


    Notes
 
{dagger} These authors contributed equally to this work. Back


    Acknowledgments
 
This study was supported by grants (NSC 92-2320-B-006-085 and NSC 91-2314-B-006-097) from the National Science Council, Taiwan, Republic of China.

Conflict of Interest Statement: None declared.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

  1. Pfister,D.G., Benson,A.B. III and Somerfield,M.R. (2004) Clinical practice. Surveillance strategies after curative treatment of colorectal cancer. N. Engl. J. Med., 350, 2375–2382.[Free Full Text]
  2. Mayer,R.J. (2004) Two steps forward in the treatment of colorectal cancer. N. Engl. J. Med., 350, 2406–2408.[Free Full Text]
  3. Hawk,E.T., Umar,A. and Viner,J.L. (2004) Colorectal cancer chemoprevention—an overview of the science. Gastroenterology, 126, 1423–1447.[CrossRef][ISI][Medline]
  4. Kan,W.S. (1981) Pharmaceutical Botany. National Research Institute of Chinese Medicine, Taipei, Taiwan.
  5. Hsu,H.Y. (1982) Treating Cancer with Chinese Herbs. Oriental Healing Arts Institute, Los Angeles, USA.
  6. Okigawa,M., Maeda,T. and Kawano,N. (1970) The isolation and structure of three new lignans from Justicia procumbens Linn var Leucantha Honda. Tetrahedron, 26, 4301–4305.[CrossRef][ISI]
  7. Fukamija,N. and Lee,K. (1986) Antitumor agents, justicidin-A and diphyllin, two cytotoxic principles from Justicia procumbens. J. Nat. Prod., 49, 348–350.[CrossRef][ISI][Medline]
  8. Day,S.H., Chiu,N.Y., Won,S.J. and Lin,C.N. (1999) Cytotoxic lignans of Justicia ciliata. J. Nat. Prod., 62, 1056–1058.[CrossRef][ISI][Medline]
  9. Day,S.H., Lin,Y.C., Tsai,M.L., Tsao,L.T., Ko,H.H., Chung,M.I., Lee,J.C., Wang,J.P., Won,S.J. and Lin,C.N. (2002) Potent cytotoxic lignans from Justicia procumbens and their effects on nitric oxide and tumor necrosis factor-alpha production in mouse macrophages. J. Nat. Prod., 65, 379–381.[CrossRef][ISI][Medline]
  10. Desagher,S. and Martinou,J.C. (2000) Mitochondria as the central control point of apoptosis. Trends Cell Biol., 10, 369–377.[CrossRef][ISI][Medline]
  11. Amarante-Mendes,G.P., Finucane,D.M., Martin,S.J., Cotter,T.G., Salvesen,G.S. and Green,D.R. (1998) Anti-apoptotic oncogenes prevent caspase-dependent and independent commitment for cell death. Cell Death Differ., 5, 298–306.[CrossRef][ISI][Medline]
  12. Danial,N.N. and Korsmeyer,S.J. (2004) Cell death: critical control points. Cell, 116, 205–219.[CrossRef][ISI][Medline]
  13. Adams,J.M. and Cory,S. (1998) The Bcl-2 protein family: arbiters of cell survival. Science, 281, 1322–1326.[Abstract/Free Full Text]
  14. Evan,G. and Littlewood,T. (1998) A matter of life and cell death. Science, 281, 1317–1322.[Abstract/Free Full Text]
  15. Thornberry,N.A. and Lazebnik,Y. (1998) Caspases: enemies within. Science, 281, 1312–1316.[Abstract/Free Full Text]
  16. Kluck,R.M., Bossy-Wetzel,E., Green,D.R. and Newmeyer,D.D. (1997) The release of cytochrome c from mitochondria: a primary site for Bcl-2 regulation of apoptosis. Science, 275, 1132–1136.[Abstract/Free Full Text]
  17. Shi,Y. (2001) A structural view of mitochondria-mediated apoptosis. Nat. Struct. Biol., 8, 394–401.[CrossRef][ISI][Medline]
  18. Bratton,S.B., Walker,G., Srinivasula,S.M., Sun,X.M., Butterworth,M., Alnemri,E.S. and Cohen,G.M. (2001) Recruitment, activation and retention of caspases-9 and -3 by Apaf-1 apoptosome and associated XIAP complexes. EMBO J., 20, 998–1009.[Abstract/Free Full Text]
  19. Nothwehr,S.F. and Martinou,J.C. (2003) A retention factor keeps death at bay. Nat. Cell Biol., 5, 281–283.[CrossRef][ISI][Medline]
  20. Cohen,H.Y., Lavu,S., Bitterman,K.J., Hekking,B., Imahiyerobo,T.A., Miller,C., Frye,R., Ploegh,H., Kessler,B.M. and Sinclair,D.A. (2004) Acetylation of the C terminus of Ku70 by CBP and PCAF controls Bax-mediated apoptosis. Mol. Cell, 13, 627–638.[CrossRef][ISI][Medline]
  21. Stadler,J., Stefanovic-Racic,M., Billiar,T.R., Curran,R.D., McIntyre,L.A., Georgescu,H.I., Simmons,R.L. and Evans,C.H. (1991) Articular chondrocytes synthesize nitric oxide in response to cytokines and lipopolysaccharide. J. Immunol., 147, 3915–3920.[Abstract/Free Full Text]
  22. Chang,M.J., Yu,W.D., Reyno,L.M., Modzelewski,R.A., Egorin,M.J., Erkmen,K., Vlock,D.R., Furmanski,P. and Johnson,C.S. (1994) Potentiation by interleukin 1 alpha of cisplatin and carboplatin antitumor activity: schedule-dependent and pharmacokinetic effects in the RIF-1 tumor model. Cancer Res., 54, 5380–5386.[Abstract]
  23. Mosmann,T. (1983) Rapid colorimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays. J. Immunol. Methods, 65, 55–63.[CrossRef][ISI][Medline]
  24. Gamet-Payrastre,L., Li,P., Lumeau,S., Cassar,G., Dupont,M.A., Chevolleau,S., Gasc,N., Tulliez,J. and Terce,F. (2000) Sulforaphane, a naturally occurring isothiocyanate, induces cell cycle arrest and apoptosis in HT29 human colon cancer cells. Cancer Res., 60, 1426–1433.[Abstract/Free Full Text]
  25. Liu,H.S., Scrable,H., Villaret,D.B., Lieberman,M.A. and Stambrook,P.J. (1992) Control of Haras-mediated mammalian cell transformation by Escherichia coli regulatory elements. Cancer Res., 52, 983–989.[Abstract]
  26. Chang,M.Y., Won,S.J., Yang,B.C., Jan,M.S. and Liu,H.S. (1999) Selective activation of Ha-ras(val12) oncogene increases susceptibility of NIH/3T3 cells to TNF-alpha. Exp. Cell Res., 248, 589–598.[CrossRef][ISI][Medline]
  27. Perkins,C.L., Fang,G., Kim,C.N. and Bhalla,K.N. (2000) The role of Apaf-1, caspase-9, and bid proteins in etoposide- or paclitaxel-induced mitochondrial events during apoptosis. Cancer Res., 60, 1645–1653.[Abstract/Free Full Text]
  28. Troiano,L., Granata,A.R., Cossarizza,A., Kalashnikova,G., Bianchi,R., Pini,G., Tropea,F., Carani,C. and Franceschi,C. (1998) Mitochondrial membrane potential and DNA stainability in human sperm cells: a flow cytometry analysis with implications for male infertility. Exp. Cell Res., 241, 384–393.[CrossRef][ISI][Medline]
  29. Mathur,A., Hong,Y., Kemp,B.K., Barrientos,A.A. and Erusalimsky,J.D. (2000) Evaluation of fluorescent dyes for the detection of mitochondrial membrane potential changes in cultured cardiomyocytes. Cardiovasc. Res., 46, 126–138.[CrossRef][ISI][Medline]
  30. Yang,J., Liu,X., Bhalla,K., Kim,C.N., Ibrado,A.M., Cai,J., Peng,T.I., Jones,D.P. and Wang,X. (1997) Prevention of apoptosis by Bcl-2: release of cytochrome c from mitochondria blocked. Science, 275, 1129–1132.[Abstract/Free Full Text]
  31. Nadakavukaren,K.K., Nadakavukaren,J.J. and Chen,L.B. (1985) Increased rhodamine 123 uptake by carcinoma cells. Cancer Res., 45, 6093–6099.[Abstract]
  32. Watabe,M., Machida,K. and Osada,H. (2000) MT-21 is a synthetic apoptosis inducer that directly induces cytochrome c release from mitochondria. Cancer Res., 60, 5214–5222.[Abstract/Free Full Text]
  33. Earnshaw,W.C., Martins,L.M. and Kaufmann,S.H. (1999) Mammalian caspases: structure, activation, substrates, and functions during apoptosis. Annu. Rev. Biochem., 68, 383–424.[CrossRef][ISI][Medline]
  34. Tseng,Y.S., Tzeng,C.C., Chiu,A.W., Lin,C.H., Won,S.J., Wu,I.C. and Liu,H.S. (2003) Ha-ras overexpression mediated cell apoptosis in the presence of 5-fluorouracil. Exp. Cell Res., 288, 403–414.[CrossRef][ISI][Medline]
  35. Enari,M., Sakahira,H., Yokoyama,H., Okawa,K., Iwamatsu,A. and Nagata,S. (1998) A caspase-activated DNase that degrades DNA during apoptosis, and its inhibitor ICAD. Nature, 391, 43–50.[CrossRef][ISI][Medline]
  36. Jayanthi,S., Deng,X., Noailles,P.A., Ladenheim,B. and Cadet,J.L. (2004) Methamphetamine induces neuronal apoptosis via cross-talks between endoplasmic reticulum and mitochondria-dependent death cascades. FASEB J., 18, 238–251.[Abstract/Free Full Text]
  37. Kumar,S. and Vaux,D.L. (2002) Apoptosis. A cinderella caspase takes center stage. Science, 297, 1290–1291.[Abstract/Free Full Text]
  38. Sawada,M., Sun,W., Hayes,P., Leskov,K., Boothman,D.A. and Matsuyama,S. (2003) Ku70 suppresses the apoptotic translocation of Bax to mitochondria. Nat. Cell Biol., 5, 320–329.[CrossRef][ISI][Medline]
  39. Tsao,L.T., Lin,C.N. and Wang,J.P. (2004) Justicidin A inhibits the transport of tumor necrosis factor-alpha to cell surface in lipopolysaccharide-stimulated RAW 264.7 macrophages. Mol. Pharmacol., 65, 1063–1069.[Abstract/Free Full Text]
  40. Asano,J., Chiba,K., Tada,M. and Yoshii,T. (1996) Antiviral activity of lignans and their glycosides from Justicia procumbens. Phytochemistry, 42, 713–717.[CrossRef][ISI][Medline]
Received March 29, 2005; revised May 10, 2005; accepted May 13, 2005.





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