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Plant-derived abrin-a induces apoptosis in cultured leukemic cell lines by different mechanisms
Toxicology and Applied Pharmacology 195 (2004) 182 – 193 www.elsevier.com/locate/ytaap
Plant-derived abrin-a induces apoptosis in cultured leukemic cell lines by different mechanisms Hideki Ohba, a,b,* Sawako Moriwaki, c Rumiana Bakalova, b Seiji Yasuda, a and Nobuyuki Yamasaki d a
Natural Substance-Composed Materials Group, National Institute for Advanced Industrial Science and Technology, AIST-Kyushu, Tosu, Saga-ken 841-0052, Japan b Single-Molecule Bioanalysis Laboratory, National Institute for Advanced Industrial Science and Technology, AIST-Shikoku, 2217-14 Hayashi-cho, Takamatsu, Kagawa 761-0395, Japan c National Institute of Vegetables and Tea Science, National Agricultural Research Organization, 2769 Kanaya, Shizuoka 428-8501, Japan d Laboratory of Biochemistry, Faculty of Agriculture, Kyushu University, 6-10-1 Hakozaki, Fukuoka 812-0053, Japan Received 3 July 2003; accepted 18 November 2003
Abstract Abrin-a consists of A-chain with N-glycosidase activity, which inhibits protein synthesis, and lectin-like B-chain responsible for binding with cell-surface receptors and penetrating of abrin-a molecule into the cells. As a lectin component, the B-chain can also participate in cell signal transduction. It has been reported that abrin induces apoptosis, but the molecular mechanism(s) of this induction have been obscure and several alternative variants have been discussed. The present study demonstrates that abrin-a induces apoptosis in human cultured cell lines, derived from acute lymphoblastic leukemia (ALL) (Jurkat, CCRF-CEM, MOLT-4, HPB-ALL). The apoptosis was estimated by: phosphatidylserine (PSer) exposure at the cell surface, activation of caspase cascade, and DNA fragmentation. The penetrating of abrin-a into the cells was detected by fluorescent confocal microscopy, using fluorescein isothiocyanate (FITC) as a fluorescent marker. It was established that the effect of abrin-a on the apoptosis induction in leukemic cells was dose- and time-dependent. The process was initiated 1 h after abrin-a application (before its penetrating into the cells) and was characterized with PSer translocation from the inner to the outer monolayer of plasma membrane, caspase activation on the first to second hour after beginning of treatment, with maximum on the third to fourth hour, and DNA fragmentation on the fourth to sixth hour, depending of the cell line. The exposure of PSer on the cell surface was detected in Jurkat, CCRF-CEM, and MOLT-4 cells. In HPB-ALL, no significant changes in PSer exposure on the cell surface was observed. Activation of caspase-3, -8, and -9 was detected in Jurkat, MOLT-4, and HPB-ALL. Surprisingly, the activity of caspase-3 increased on the first hour after beginning of treatment, while the activity of caspase-8 and -9 began to increase on the second hour. In CCRF-CEM, activation of caspases was not measured, but the apoptosis progressed to DNA fragmentation in a dose- and time-dependent manner. DNA fragmentation was also detected in Jurkat, but not in MOLT-4 and HPB-ALL cells. It seems that the mechanisms of abrin-a-induced apoptosis are different and the progress of apoptosis depends of the cell line. There was a very good positive correlation between the agglutinating activity of abrin-a and development of apoptosis to DNA fragmentation. The time-dependent effects of abrin-a on apoptosis as well as its time-dependent penetration into the cells suggest that the B-chain probably triggers the apoptosis, while the A-chain and breakage of the disulfide bond are responsible for its progress. D 2004 Elsevier Inc. All rights reserved. Keywords: Abrin-a; Apoptosis; Phosphatidylserine translocation; Caspase activity; DNA fragmentation; Acute lymphoblastic leukemia
Introduction Abbreviations: ALL, acute lymphoblastic leukemia; FITC, fluorescein isothiocyanate; PBS ( ), phosphate-buffered saline (Ca2+ and Mg2+ free); PSer, phosphatidylserine. * Corresponding author. Natural Substance-Composed Materials Group, National Institute for Advanced Industrial Science and Technology, AIST-Kyushu, 807-1 Shuku-machi, Tosu, Saga-ken 841-0052, Japan. Fax: +81-942-81-3690. E-mail address: [email protected] (H. Ohba). 0041-008X/$ - see front matter D 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.taap.2003.11.018
Abrin-a, obtained from Abrus precatorius seeds, belongs to the family of type II ribosome inactivating proteins— heterodimeric glycoproteins, consisting of two nonidentical polypeptide chains (A- and B-chain) cross-linked through a single disulfide bond (Liu et al., 2000; Wei et al., 1974). The A-chain affects the protein synthesis by modification of the
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ribosomal subunits of the eukaryotic cells (Endo et al., 1987; Hartley et al., 1991; Hegde et al., 1993). The B-chain is a carbohydrate-binding protein and serves to bind the abrin-a molecule specifically with D-galactosyl-containing cell-surface receptors (Hegde and Podder, 1997; Olsnes et al., 1974). It has been found that abrin-a is a promising tool for distinguishing of cancer cells from normal ones as a result of its higher affinity to the cancer cells. The drug attracts a great deal of interest because of its unique biological activities as cytoagglutination and cytotoxicity, accompanied with induction of apoptosis, and all these effects are predominantly expressed in cancer cells. (Geier et al., 1996; Keppler-Hafkemeyer et al., 1998; Moriwaki et al., 2000; Shin et al., 2001). The accent of the present study is on the abrin-a-mediated mechanism(s) of apoptosis induction in leukemic cells. The apoptotic cascade in mammalian cells is a multistep process, initiating in most cases by loss of integrity of the mitochondrial membrane. This usually involves the activation of permeability mitochondrial transition pores, loss of mitochondrial membrane potential (
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of apoptosis: phosphatidylserine (PSer) translocation from the inner to the outer monolayer of plasma membrane as an early event of apoptosis, activation of caspases as a moderate phase, and internucleosomal DNA fragmentation as a last phase (Adams and Cory, 2002; Beauparlant and Shore, 2003; Collins et al., 1997; Zhang and Xu, 2000).
Materials and methods Reagents. Abrin-a was purified from the seeds of A. precatorius, grown in Thailand, by the method of Lin et al. (1980) and was stored as ammonium sulfate precipitate at 4 jC until use. Apoptosis Ladder Detection Kit, Wako, and fetal calf serum (FCS) were purchased from Wako Pure Chemical Industries, Ltd., Osaka. CapACE Assay System, Colorimetric was purchased from Promega (Madison, WI). ApoAlert Caspase-8 Fluorometric Assay Kit and ApoAlert Caspase-9/6 Fluorescent Assay Kit were obtained from Clontech Laboratories, Inc., (Palo Alto, CA). Fluorescein isothiocyanate (FITC)– Annexin V was obtained from Pharmingen. Antibiotics were obtained from Gibco GRL (Grand Island, NY). All chemicals were of analytical grade. Abrin-a. Abrin-a solution was prepared by dialyzing of ammonium sulfate precipitate of abrin-a against phosphatebuffered saline [PBS ( ), Ca2+ and Mg2+ free] at 4 jC before use and filtered through a Millipore filter (0.2 Am). The concentration of abrin-a was measured spectrophotometrically, using an absorption coefficient A1cm1% = 15.9 at 280 nm (Endo et al., 1987). Cultured leukemic cells. The human leukemic cell lines [Jurkat, MOLT-4, HPB-acute lymphoblastic leukemia (ALL), and CCRF-CEM] were cultured in RPMI-1640 medium, supplemented with 10% heat-inactivated FCS, and 100 Ag/ml of streptomycin and 100 U/ml of penicillin in a humidified atmosphere at 37 jC with 5% CO2. These cells were generously provided by Dr. J. Minowada (Hayashibara Biochemical Laboratories, Inc., Okayama, Japan). Abrin-a uptake into the cells. The penetrating of abrin-a into the cells was detected by confocal fluorescent microscopy using fluorescein isothiocyanate (FITC) as a fluorescent marker. FITC was conjugated with abrin-a and then served as a fluorescent probe to detect the drug penetrating into the cells in a time-dependent manner. Cells (1.0 106 cells/well) were incubated with 0.3 AM FITC– abrin-a at different time intervals. The samples were analyzed by ‘‘Olympus IX70’’ confocal microscope. Flow cytometric assay. The viability of cells were analyzed by flow cytometer Beckman Coulter—Epics XL, operating in accordance with the manufacturer’s recommendations after fine adjustments for optimization. The forwardscatter (FS) and side-scatter (SS) parameters were adjusted to accommodate the inclusion of viable and dead leukemic cells within the acquisition data. No cells excluded from the
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analysis and 10,000 cells were counted. Data were collected and analyzed by ‘‘XL System II’’ software. The results were presented as a dot plot of cell fluorescence with quadrant markers drawn to distinguish viable and dead cells, as well as to distinguish leukemic and normal cells. Quadrant A in Fig. 1 contains viable normal cells, quadrant B contains viable leukemic cells, quadrant C contains all viable cells, and quadrant D contains all dead cells. The percentage of dead cells before and after abrin-a treatment was calculated and presented in the table. DNA-fragmentation assay. Cells (1.0 106 cells/well) were cultured as described above. After 24 h of cultivation, cells were incubated with 2-fold dilution series of abrin-a (maximum concentration 8 AM) in PBS ( ) for 24 h under
the same conditions. For the time-dependent analysis of DNA fragmentation, cells were incubated with abrin-a (final concentration 0.3 AM) and the DNA fragmentation was determined in the range of 0 – 24 h. At each time point, the cells were collected and cell lysis and DNA extraction were provided using commercially available kit for DNA-fragmentation assay (Apoptosis Ladder Detection Kit, Wako) based on the method of Ioannou and Chen (1996). The extracted DNA was analyzed by electrophoresis on agarose gels (1.5% gel density) using TBE buffer. The sample, in which PBS ( ) was added instead of abrin-a solution, was assumed to be a contrast. DNA from apoptotic cells forms a ladder on the agarose gel, while DNA from non-apoptotic cells appears as a single band or smeared on the agarose gel. Presence of random DNA fragmentation (smear) is a characteristic of necrosis.
Fig. 1. Agarose gel electrophoresis of DNA extracted from acute T-lymphoblastic leukemia cells after treatment with abrin-a in different concentrations. (A) DNA-fragmentation assay. The cells (2.22 104 cells/well) were cultured in the presence of 2-fold dilution series of abrin-a (8 AM maximal concentration) as it is described in Materials and methods. After 24 h incubation the cells were collected by centrifugation and DNA extraction was carried out using DNAfragmentation assay kit (Apoptosis Ladder Detection Kit, Wako). The DNA was analyzed by TBE electrophoresis on 1.5% agarose gel. The sample, in which PBS ( ) was added instead of abrin-a solution, was considered as a control. Blots from one typical experiment are presented in the figure. St, standard marker; lane 1, control (nontreated cells); lanes 2 – 11, cells treated with 0.016 – 8 AM abrin-a, respectively. (B) Flow cytometry. The percentage of dead cells before and after abrin-a treatment was determined by flow cytometry and the results are presented in the table (*P < 0.05, **P < 0.01, ***P < 0.001 vs. the respective control nontreated cells, C). A typical histogram as a dot plot of cell fluorescence with quadrant markers drawn to distinguish viable and dead cells, as well as to distinguish leukemic and normal cells, is also shown in the figure. Quadrant A contains viable normal cells, quadrant B contains viable leukemic cells, quadrant C contains all viable cells, and quadrant D contains all dead cells.
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PSer detection assay. Cells (1.0 106 cells/well) were incubated with abrin-a (final concentration 0.3 AM) in PBS ( ) for 0 –3 h under the same conditions as mentioned above. At each time point the cells were collected by centrifugation (450 g, 20 min), washed twice with PBS ( ) containing 2.5 mM CaCl2 (Annexin V-binding buffer), and resuspended in the same buffer. After that, 100 Al of cell suspension was incubated with 5 Al of FITC-conjugated Annexin V (FITC –Annexin V) for 10 min at room temperature (RT) in a dark place. The cells were washed twice with Annexin V-binding buffer and were resuspended in 100 Al of the same buffer. FITC – Annexin V, bound to PSer exposed on the cell surface, was detected spectro-
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fluorimetrically at kem = 535 nm (kex = 488 nm) using Fluoromark Fluorescent Microplate Leader (BioRad). Cell lysis. Cells (1.0 106 cells/well) were incubated with abrin-a (final concentration 0.3 AM) in the range 0 –4 h. At each time point, the cells were sedimented by centrifugation at 450 g for 10 min at 4 jC, washed three times with cold PBS ( ), and lysed by lysis buffer (20 mM HEPES/KOH, pH 7.5, 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, 1 mM EGTA, 1 mM DTT, 5 Ag/ml pepstatin A, 10 Ag/ml leupeptin, 2 Ag/ml aprotinin, 0.1% Triton X-100) and freezing – thawing. Freeze –thaw cycles were repeated three times to ensure complete cell lysis. After centrifugation at 15,000 g for 20
Fig. 2. Agarose gel electrophoresis of DNA extracted from acute T-lymphoblastic leukemia cells after treatment with abrin-a at different time intervals. (A) DNA-fragmentation assay: The cells (1 106 cells/well) were cultured in the presence of abrin-a (0.3 AM) as described in Materials and methods section. At each time point, the cells were collected by centrifugation and DNA analysis was carried out as it is described in Fig. 1. The sample, in which PBS ( ) was added instead of abrin-a solution, was considered as a control (C, nontreated cells). Blots from one typical experiment are presented in the figure. St, standard marker. (B) Flow cytometry. The percentage of dead cells before and after abrin-a treatment (0.3 AM) was determined by flow cytometry at different time intervals and the results are presented in the table (*P < 0.05, **P < 0.01, ***P < 0.001 vs. the respective control nontreated cells, C).
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min at 4 jC, the cell lysates were used for analysis of caspase isoenzyme activity.
FMK) in PBS ( ) were prepared analogous to described above.
Analysis of caspase isoenzymes. Cells, cultured in the absence or in the presence of abrin-a (3 AM), were washed twice by PBS ( ) and lysed in RIPA-buffer. Lysates were centrifuged at 16,000 g for 15 min at 4 jC. Protein concentrations were determined on supernatants using a protein microassay of the Bradford method (Bio-Rad). Cell lysates (obtained from viable, apoptotic, necrotic, and apocrotic cells) were analyzed for caspase activity by two procedures: (i) Western blot analysis as described in Andreeff et al. (1999) and Towbin et al. (1979), and (ii) spectrophometrically or spectrofluorimetrically, detecting the products of the caspase-catalyzed reactions. Enzyme activities and protein expression were determined in cell lysates, obtaining from all cell forms— viable, apoptotic, necrotic and apocrotic.
Spectrofluorimetric detection of caspase-9 activity. Fifty microliters of the cell lysates were incubated with 50 Al of 2 reaction buffer/DTT (ApoAlert Caspase-9/6 Fluorometric Assay Kit, Clontech), 5 Al of substrate solution (5 mM Leu-Glu-His-Asp-AMC) and 2 Al of DMSO for 1 h at 37 jC. The release of 7-amino-4-methyl coumarin (AMC), catalyzed by caspase-9, was estimated fluorimetrically at kem = 490 nm (kex = 390 nm) using Fluoromark Fluorescence Microplate Leader (BioRad). As a negative control, cells treated with abrin-a plus caspase inhibitor (0.5 mM Leu-Glu-Thr-Asp-CHO) in PBS ( ) were prepared analogous to described above.
Western blot. Briefly, 60 mg cell protein extract was dissolved 1:1 in 2 Laemmli sample buffer and the samples were subjected to electrophoresis on SDS-polyacrylamide gel (5% stacking, 4 –20% resolving). Electrophoresis was carried out in two steps: at 80 V for 15 min and 120 V for 2 h at room temperature (RT approximately 22 jC). The separated protein fractions were transferred to a Hybond-P PVDF membranes (Amersham Bioscience) during 18 h at 35 V, 4 jC. The transferred proteins were immunoblotted using adequate primary anti-caspase antibodies or anti-hactin antibody (for comparison), and subsequent secondary antibodies (Andreeff et al., 1999; Towbin et al., 1979). Five and seven independent experiments were carried out for caspase-3 and -9, respectively. Spectrophotometric detection of caspase-3 activity. Fiftyfour microliters of the cell lysates were incubated with 32 Al of caspase assay buffer (20 mM HEPES/KOH, pH 7.5, 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, 1 mM EGTA), 2 Al of DMSO, 10 Al of 100 mM DTT and 2 Al of substrate solution (1 mM Ac-Asp-Glu-Val-Asp-pNA) for 15 min at 37 jC. The release of chromophore pnitroaniline, catalyzed by caspase-3, was monitored spectrophotometrically at 405 nm, using an Inter Med Immunomini NJ-2300 multiwell scanning spectrophotometer. As a negative control, cells treated with abrin-a plus caspase inhibitor (50 AM Z-Val-Ala-Asp-FMK) in PBS ( ) were prepared at the same manner as described above. Spectrofluorimetric detection of caspase-8 activity. Fifty microliters of the cell lysates were incubated with 50 Al of 2 reaction buffer/DTT (ApoAlert Caspase-8 Fluorometric Assay Kit, Clontech) and 5 Al of substrate solution (1 mM Ile-Glu-Thr-Asp-AFC) for 1 h at 37 jC. The release of 7amino-4-trifluoromethyl coumarin (AFC), catalyzed by caspase-8, was estimated fluorimetrically at kem = 535 nm (kex = 390 nm) using Fluoromark Fluorescence Microplate Leader (BioRad). As a negative control, cells treated with abrin-a plus caspase inhibitor (5 AM Ile-Glu-Thr-Asp-
Measurement of cytotoxicity. The cytotoxicity of abrin-a against leukemic cells was measured by CellTiter-GloTM luminescent cell viability assay (Promega) (Crouch, 1993). The ATP bioluminescence was used as a marker of cell proliferation and viability. Briefly, the lectins were dissolved in PBS ( ) and were dispensed as 10-Al aliquots into 96-well microplates. The cells were suspended in the culture medium to a concentration 5 105 cells/ml and then were added in 90-Al aliquots to each patch. After 24
Fig. 3. Changes in the cell-surface phosphatidylserine (PSer) induced by abrin-a in acute T-lymphoblastic leukemia cells. The cells (1 106 cells/ well) were incubated with abrin-a (0.3 AM) for 0 – 3 h, collected by centrifugation, and incubated with FITC – Annexin V in PBS ( ), containing 2.5 mM CaCl2 (Annexin V-binding buffer) for 10 min at RT in a dark place. The cells were washed twice by Annexin V-binding buffer and were resuspended in the same buffer (the cell concentration was identical in all samples). The detection of FITC – Annexin V, bound to PSer exposed on a cell surface, was carried out spectrofluorimetrically at kex = 488 nm and kem = 525 nm. The intensity of fluorescence (in arbitrary units) is expressed in the figure. The results are mean F SD from six independent experiments (n = 6). In the case of MOLT-4, CCRF-CEM, and Jurkat, the increase of fluorescence after 1, 2 and 3 h treatment with abrin-a was statistically significant ( P < 0.001 vs. control nontreated cells). In the case of HPB-ALL, the changes in fluorescence intensity were statistically significant only at 3-h treatment with abrin-a ( P < 0.05 vs. control nontreated cells).
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h incubation at 36.9 jC in cell incubator (5.1% CO2), CellTiter 96 kit (Promega) was added in aliquots of 100 Al to each patch and incubated with cell suspensions for 1 h, following the procedure, recommended by the producer. The luminescence, produced by luciferase catalyzed lucif-
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erin + ATP reaction, was measured by an Inter Med Immunomini NJ-2300 multiwell scanning spectrophotometer. The data were normalized to the control group. The abrin-a cytotoxicity was calculated as percentage decrease of luminescence in comparison with the control.
Fig. 4. Caspase-3 activity in acute T-lymphoblastic leukemia cells after treatment with abrin-a. The cells (1.0 106 cells/well) were incubated with abrin-a (0.3 AM). At each time point, the cells were sedimented, washed with cold PBS ( ), lysed, and detection of caspase-3 activity was carried out as follows: (A) The release of chromophore p-nitroaniline, catalyzed by caspase-3, was monitored spectrophotometrically at 405 nm. As a negative control, cells treated with abrina in the presence of caspase-3 inhibitor (Z-Val-Ala-Asp-FMK) were used. Closed symbols, in the absence of caspase-3 inhibitor; open symbols, in the presence of caspase-3 inhibitor. The results are mean F SD from seven independent experiments (n = 7). In the case of Jurkat, HPB-ALL and MOLT-4, P < 0.01 vs. control group, at 1 h; P < 0.001 vs. control group, at 2 – 4 h. In the case of CCRF-CEM, nonsignificant (ns), at 1 h; P < 0.05 vs. control group, at 2 – 4 h. (B) Western blot analysis of caspase-3 was carried out after 1-h incubation of cells with abrin-a. Forty milligrams of protein was applied in electrophoresis. Blots from one typical experiment are presented in the figure. C, in the absence of abrin-a (control); 1, 1 h after addition of abrin-a (0.3 AM) to the cell suspensions. Densitograms correspond to the immunoblot analysis and the results are expressed as mean F SD from five independent experiments.
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Cytoagglutination assay. Cytoagglutination was evaluated by the method of Ohba et al. (1997) using a Jasco V-550 spectrophotometer equipped with a magnetic stirrer. Briefly, the cells were washed three times by PBS ( ) and resuspended in the same buffer to a concentration 2 106 cells/ ml. The cell suspension and PBS ( ) were placed into the sample and reference cuvettes, respectively, and were incubated until the baseline at 600 nm became constant. A solution of abrin-a was added to the cell suspension in different concentrations and the decrease in turbidity at 600 nm (OD600) was recorded under continuous stirring. The measurement was stopped when the OD600 reached to a plateau. The velocity of cytoagglutination (
and 6 h, respectively, after addition of abrin-a to the cell suspensions, and the density of the bands increased with the time of treatment (Figs. 2a, d). The same tendency was established in the increase of percentage of dead cells, estimated by flow cytometry (Fig. 2, table). It may also be speculated that necrosis (smear) proceeded apoptosis (ladder) and the cell death was a result of apoptosis development. In HPB-ALL and MOLT-4, the step ladder of DNA was not detected even 24 h after addition of abrin-a to the cell suspensions (Figs. 2b, c). Only a smear was detected and probably the cell death was a result of necrosis. However, the percentage of dead cells markedly increased after 24-h treatment.
Statistical analysis. One-way analysis of variance (ANOVA) was employed, following by Bonferroni’s test for significant differences. Statistical significance was defined at P < 0.05. The statistical procedures were performed with GraphPad InStat software. Data are expressed as mean F SD.
The fluorescence of FITC – Annexin V was detected in the target cells as an index for translocation of PSer from the inner to the upper monolayer of the cell-surface membrane (flip-flop), which happens during the first stage of apoptosis (Van Engeland et al., 1998; Vermes et al., 1995). FITC – Annexin V specifically recognizes PSer– Ca2+ complexes and is widely used to detect the early apoptotic cells. It was established that the fluorescence intensity began to increase in CCRF-CEM, Jurkat, and MOLT-4 1 h after addition of abrin-a (Fig. 3). On the other hand, in HPB-ALL, fluores-
Results
Change in cell-surface PSer after abrin-a application
Dose-dependent effects of abrin-a on the DNA fragmentation DNA fragmentation in abrin-a-treated cells was used as a conventional marker of the final step of apoptosis and as a major characteristic of apoptotic cell death. The results in Fig. 1 demonstrate that even low concentrations of abrin-a (0.01 –0.06 AM) induced DNA fragmentation in Jurkat and CCRF-CEM cells—a step ladder of DNA fragmentation was detected (Fig. 1d, lanes 2– 11; Fig. 1a, lanes 5– 11). The process of DNA fragmentation was better and clearly manifested in the higher concentrations of abrin-a. The strength of bands markedly enhanced and they were well defined at abrin-a concentrations >0.5 AM. In HPB-ALL and MOLT-4, a ladder of DNA fragmentation was not detected even at high concentrations of abrin-a (Figs. 1b, c). The percentage of dead cells, determined by flow cytometry, also increased with increasing abrin-a concentration (Fig. 1, table). The results were statistically significant at 0.12 AM abrin-a and higher, even in MOLT-4 and HPB-ALL, in which DNA fragmentation was not detected. Time-dependent effects of abrin-a on the DNA fragmentation The time-dependent effects of abrin-a on the different phases of apoptosis were determined at concentration 0.3 AM. In Jurkat and CCRF-CEM, the fragmentation of DNA was better and clearly manifested with the time of cell treatment by abrin-a. DNA fragmentation was detected 4
Fig. 5. Caspase-8 activity in acute T-lymphoblastic leukemia cells after treatment with abrin-a. The cells (1.0 106 cells/well) were incubated with abrin-a (final concentration 0.3 AM). At each time point, the cells were sedimented, washed with cold PBS ( ), lysed, and detection of caspase-8 activity was carried out as it is described in Materials and methods. The release of 7-amino-4-trifluoromethyl coumarin (AFC), catalyzed by caspase-8, was estimated fluorimetrically at kem = 535 nm (kex = 390 nm). As a negative control, cells treated with abrin-a in the presence of caspase-8 inhibitor (Ile-Glu-Thr-Asp-FMK) were used. Closed symbols, in the absence of caspase-8 inhibitor; open symbols, in the presence of caspase-8 inhibitor. The results are mean F SD from six independent experiments (n = 6). In the case of Jurkat and HPB-ALL, ns, at 1 h; P < 0.001 vs. control group, at 2 – 3 h. In the case of MOLT-4, ns, at 1 – 2 h; P < 0.01 vs. control group, at 3 h. In the case of CCRF-CEM, ns, at 1 – 3 h.
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1.5
0.5
Fig. 6. Caspase-9 activity in acute T-lymphoblastic leukemia cells after treatment with abrin-a. The cells (1.0 106 cells/well) were incubated with abrin-a (final concentration 0.3 AM). At each time point, the cells were sedimented, washed with cold PBS ( ), lysed, and detection of caspase-9 activity was carried out as follows: (A) The release of 7-amino-4-methyl coumarin (AMC), catalyzed by caspase-9, was estimated fluorimetrically at kem = 490 nm (kex = 390 nm). As a negative control, cells treated with abrin-a in the presence of caspase-9 inhibitor (Leu-Glu-Thr-Asp-CHO) were used. Closed symbols, in the absence of caspase-9 inhibitor; open symbols, in the presence of caspase-9 inhibitor. The results are mean F SD from eight independent experiments (n = 8). In the case of Jurkat and HPB-ALL, ns, at 1 h; P < 0.001 vs. control group, at 2 – 3 h. In the case of MOLT-4, ns, at 1 – 2 h; P < 0.001 vs. control group, at 3 h. In the case of CCRF-CEM, ns, at 1 – 3 h. (B) Western blot analysis of caspase-9 was carried out as described in Materials and methods. Forty milligrams of protein was applied in electrophoresis. Blots from one typical experiment are presented in the figure. C, in the absence of abrin-a (control); 1, 3 h after addition of abrin-a (0.3 AM) to the cell suspensions. No active caspase-9 was detected after 1-h treatment with abrin-a (data not shown). Densitograms correspond to the immunoblot analysis and the results are expressed as mean F SD from six independent experiments.
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cence intensity increased slightly at the third hour after abrin-a application. Activation of caspase isoenzymes During induction of apoptosis, several caspases (cysteine proteases) are activated by the signal transmission in the cells (Beauparlant and Shore, 2003). In caspase cascade, caspase-3 is a key enzyme in the apoptotic pathway. The activation of caspase-3 happens in the downstream of the signal transmission and is responsible for DNA cleavage. It was established that the caspase-3 activity increased after addition of abrin-a, and it reached a maximum 2 h after beginning of treatment in Jurkat and HPB-ALL, and 3 h after that in MOLT-4 (Fig. 4). In CCRF-CEM, the caspase-3 activity only increased gradually, but slightly, until fourth
hour after abrin-a application. An increase in the enzyme activity was not detected in the presence of caspase-3 inhibitor in all cell lines. To investigate in more details the signal transmission mechanism of apoptosis, induced by abrin-a, we analyzed the activation of caspase-8 and -9, usually proceeding the activation of caspase-3 in the cascade. The results are presented in Figs. 5 and 6. In Jurkat, the caspase-8 and -9 activity increased rapidly on the second hour after addition of abrin-a. Caspase-8 activity reached a plateau after 2h incubation of Jurkat with abrin-a, while caspase-9 activity gradually increased until the third hour. In HBPALL, the caspase-8 and -9 activity increased rapidly and reached a plateau on the second hour after addition of abrin-a. In CCRF-CEM and MOLT-4, the caspase-8 and -9 manifested a poor activity even after 3 h of cell treatment with abrin-a.
Fig. 7. Time-dependent penetrating of abrin-a into Jurkat (A) and CCRF-CEM (B) cells. The cells (1.0 106 cells/well) were incubated with FITC – abrin-a (final concentration 0.3 AM). At each time point, the cells were sedimented, washed with PBS ( ) and FITC – abrin-a, bound to the cell surface or penetrating into the cells, was measured by confocal fluorescent microscopy (green light). In parallel, propidium iodide (PI) staining was used as a marker of cell survival (red light). Note that in Jurkat cell aggregation takes place during abrin-a treatment.
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Penetrating of abrin-a into the cells and induction of apoptosis Using FITC-labeled abrin-a and propidium iodide (PI) staining, we detected in parallel the time-dependent penetrating of the drug into the cells and development of apoptosis. The green light in Fig. 7 is representative for FITC-labeled abrin-a and the red light is representative for penetrating of propidium iodide into the cells, which is a characteristic of apoptotic cell death. In Jurkat, it was observed that abrin-a (0.3 AM) penetrated into the cells after 2-h incubation, while propidium iodide was detected into the cells on the third hour. On the fourth to fifth hour, cell aggregation was also observed. The results suggests that Jurkat cells are very sensitive to abrin-a and induction of apoptosis takes place immediately after its cellular uptake. In CCRF-CEM, abrin-a penetrated into the cells on the second hour, while propidium iodide was detected into the cells on the sixth hour. It seems that CCRF-CEM cells are most resistant to abrin-a and several hours are necessary to detect the pro-apoptotic effect of the drug in this cell line.
Discussion Apoptosis is a programmed cell death, an essential mechanism for cell removal during embryonic development and elimination of lymphocytes during thymic development (Majno and Joris, 1995; Trump et al., 1997). To date, several endogenous physiological mediators of apoptosis have been identified. There are many data, demonstrating that some environmental factors are also responsible for induction of apoptosis, as radiation, viruses, drugs, etc., and all these candidates act by a similar route (Kopecky et al., 2001; Zhang et al., 2001). It is widely accepted that apoptosis is characterized by several biochemical events: (i) changes in the lipid composition of the cell-surface membrane resulted in translocation of PSer from the inner to the outer monolayer as an early event of apoptosis; (ii) disturbance of mitochondria and release of cytochrome c in cytosol, which serves as a cofactor, together with the apoptotic protease activating factor, to activate pro-caspase-9; (iii) pro-caspase-9 unlocks caspase cascade, resulting in caspase-3 activation, responsible for DNA fragmentation as a final stage. Recently, it has been reported that caspase-3 can be activated through two alternative pathways: from caspase-8 directly, or caspase8 activates pro-caspase-9, which serves as activator of caspase-3 (Hegde et al., 1998; Luo et al., 1998; Pan et al., 1998). However, there are no data confirming that both pathways exist and act in parallel in one and the same time. In this study, we established that abrin-a induces apoptosis in leukemic cells and the process goes by different mechanisms, depending on the cell line. In Jurkat, the apoptosis followed the conventional pathway—PSer trans-
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location on the cell surface, caspase activation, and DNA fragmentation. In CCRF-CEM, the apoptosis began with PSer translocation on the cell surface and went to DNA fragmentation without any induction of caspase activity. In MOLT-4, the apoptosis was initiated and characterized with PSer exposure and caspase activation, but does not progress to DNA fragmentation. In HPB-ALL, the translocation of PSer was negligible, the activity of caspases enhanced markedly, but DNA-fragmentation was not detected. The time-dependent effects of abrin-a on the development of apoptosis demonstrated that PSer translocation preceded the penetrating of drug into the cells (Fig. 7). Because the B-chain is responsible for the binding of abrin-a to the cell surface, the interaction between B-chain and carbohydrates can be considered as one of the trigger signals of early apoptotic events. Recently, it has been reported that PSer synthesis by serine-base exchange enzyme system, in the endoplasmic reticulum of mammalian cells, is able to regulate PSer exposure in cells undergoing the apoptotic process (Pelassy et al., 2000). PSer synthesis is modulated by alterations of the plasma membrane potential that modify the activity of the plasma membrane serine transporter (Breintmayer et al., 1996). The lectins, including abrin-a, are well-known substances, affecting the plasma membrane potential and therefore affecting PSer synthesis and its membrane translocation. However, in HPB-ALL, we did not observe significant changes in PSer exposure at the cell surface. The caspase-3 activation also preceded the penetrating of abrin-a into the cells, as well as the activation of caspase8 and -9, which is in contrast with the commonly accepted hypothesis that abrin-a crosses the plasmatic membrane of tumor cells by endocytosis, activates directly caspase-9, followed by activation of caspase-3. In our study, we established that caspase-3 activity increased on the first hour after abrin-a application and reached a maximum on the second hour in Jurkat and HPB-ALL, and on the third hour in MOLT-4. In contrast, caspase-8 and -9 began to increase on the second hour after abrin-a application and reached a maximum on the third hour in Jurkat and HPBALL. In MOLT-4, caspase-8 and -9 activity increased on the third hour, however the effect was lower that in Jurkat and HPB-ALL. In CCRF-CEM, the activities of caspase-8 and 9 were negligible and only the activity of caspase-3 increased slightly. The results suggest that probably B-chain-dependent signaling is responsible for caspase-3 direct activation by abrin-a in Jurkat, MOLT-4, and HPB-ALL. On the second hour after the beginning of abrin-a application, the drug was detected into the cells and it is impossible to clarify the role of both B- and A-chains in induction of caspase-8 and-9 activity, as well as in the continuously increase of caspase-3 activity. Moreover, it was found that the activation of caspase cascade and particularly caspase-3 is not enough for induction of DNA fragmentation, as well as it is not obligatory
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event, proceeding DNA fragmentation. DNA fragmentation was detected only in Jurkat and CCRF-CEM, but not in MOLT-4 and HPB-ALL cells. In Jurkat and CCRF-CEM, the abrin-a caused the apoptosis at low drug concentration and in a short time of treatment (4 h in Jurkat and 6 h in CCRF-CEM). In CCRF-CEM, abrin-a induced DNA fragmentation without caspase activation—a mechanism different from an already known apoptotic route. Presumably, apoptosis-inducing factor (AIF), which has been characterized as a caspase-independent death effector, participates in the development of apoptosis in CCRF-CEM cells. On our knowledge, there are no data about the effect of abrin-a on the translocation of AIF from mitochondria into the nucleus and this hypothesis is speculative, but it seems to be likely and its proof is in our future efforts. The pro-apoptotic effect of abrin-a can also explain its possibility to induce indirectly oxidative stress in leukemic cells. Separation of A-chain and B-chain of abrin-a molecule, which requires a breakage of –S – S– bond, has a potential to destroy the balance of antioxidant thiol-containing substances and to provoke free radical production in the cells. Free radicals are well-known apoptotic signals. Moreover, recently, it has been reported that abrin-a inactivates 30-kDa thiol-specific antioxidant protein and thus can decrease antioxidant defense potential of the cells (Shin et al., 2001). The differences in caspase activation and DNA fragmentation between the leukemic cell lines used can also explain the different amount and type of abrin-a-binding receptors on the cell surface. Different signals can be triggered by abrin-a, depending on the degree of abrin – cell binding and the type of this bound, analogous to other lectins (Danguy et al., 2002). In our previous paper (Moriwaki et al., 2000), we demonstrated that abrin-a manifests target cytoagglutinating and cytotoxic activity against leukemic cells and the drug is available as a marker of malignancy. Comparing the present results with previously published data, an interesting fact was mentioned: DNA fragmentation was observed only in cell lines in which abrin-a manifested a high cytoagglutinating activity (Fig. 1, Table 1). Probably, the cytoagglutination and changes in the charge of the cell surface by abrin-a are trigger signals, programming the apoptosis for development to the final stage. Abrin-a also manifested a high cytotoxicity against Jurkat and CCRF-CEM, followed by HPBALL and MOLT-4. In conclusion, the results shown that the abrin-a induces apoptosis in leukemic cells by different mechanisms, depending of the cell line. The time-dependent effects of abrin-a on apoptosis, as well as its time-dependent penetrating into the cells suggest that the B-chain is probably responsible for induction of apoptosis, while the A-chain and breakage of the disulfide bond defined its progress (Shin et al., 2001). Because a very good positive correlation between the cytoagglutinating activity of abrin-a and development of apoptosis to DNA fragmentation was found, it
Table 1 Effect of abrin-a on the velocity of cytoagglutination, cell viability, and induction of apoptosis in acute lymphoblastic leukemia T-cell lines Cell line
Velocity of cytoagglutination (DOD600 nm/min)a
Cell viability (% from control)a
DNA fragmentation
Jurkat CCRF-CEM MOLT-5 HPB-ALL
1.46 0.90 0.28 0.18
20 28 68 53
++ +
a
F F F F
0.09 0.03 0.01 0.01
F F F F
4 7 12 11
According to Moriwaki et al. (2000).
seems that the B-chain-mediated signaling is also involved in the programming of the development of apoptosis to the last phase or its termination to the phase of caspase activation.
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Plant-derived abrin-a induces apoptosis in cultured leukemic cell lines by different mechanisms Hideki Ohba, a,b,* Sawako Moriwaki, c Rumiana Bakalova, b Seiji Yasuda, a and Nobuyuki Yamasaki d a
Natural Substance-Composed Materials Group, National Institute for Advanced Industrial Science and Technology, AIST-Kyushu, Tosu, Saga-ken 841-0052, Japan b Single-Molecule Bioanalysis Laboratory, National Institute for Advanced Industrial Science and Technology, AIST-Shikoku, 2217-14 Hayashi-cho, Takamatsu, Kagawa 761-0395, Japan c National Institute of Vegetables and Tea Science, National Agricultural Research Organization, 2769 Kanaya, Shizuoka 428-8501, Japan d Laboratory of Biochemistry, Faculty of Agriculture, Kyushu University, 6-10-1 Hakozaki, Fukuoka 812-0053, Japan Received 3 July 2003; accepted 18 November 2003
Abstract Abrin-a consists of A-chain with N-glycosidase activity, which inhibits protein synthesis, and lectin-like B-chain responsible for binding with cell-surface receptors and penetrating of abrin-a molecule into the cells. As a lectin component, the B-chain can also participate in cell signal transduction. It has been reported that abrin induces apoptosis, but the molecular mechanism(s) of this induction have been obscure and several alternative variants have been discussed. The present study demonstrates that abrin-a induces apoptosis in human cultured cell lines, derived from acute lymphoblastic leukemia (ALL) (Jurkat, CCRF-CEM, MOLT-4, HPB-ALL). The apoptosis was estimated by: phosphatidylserine (PSer) exposure at the cell surface, activation of caspase cascade, and DNA fragmentation. The penetrating of abrin-a into the cells was detected by fluorescent confocal microscopy, using fluorescein isothiocyanate (FITC) as a fluorescent marker. It was established that the effect of abrin-a on the apoptosis induction in leukemic cells was dose- and time-dependent. The process was initiated 1 h after abrin-a application (before its penetrating into the cells) and was characterized with PSer translocation from the inner to the outer monolayer of plasma membrane, caspase activation on the first to second hour after beginning of treatment, with maximum on the third to fourth hour, and DNA fragmentation on the fourth to sixth hour, depending of the cell line. The exposure of PSer on the cell surface was detected in Jurkat, CCRF-CEM, and MOLT-4 cells. In HPB-ALL, no significant changes in PSer exposure on the cell surface was observed. Activation of caspase-3, -8, and -9 was detected in Jurkat, MOLT-4, and HPB-ALL. Surprisingly, the activity of caspase-3 increased on the first hour after beginning of treatment, while the activity of caspase-8 and -9 began to increase on the second hour. In CCRF-CEM, activation of caspases was not measured, but the apoptosis progressed to DNA fragmentation in a dose- and time-dependent manner. DNA fragmentation was also detected in Jurkat, but not in MOLT-4 and HPB-ALL cells. It seems that the mechanisms of abrin-a-induced apoptosis are different and the progress of apoptosis depends of the cell line. There was a very good positive correlation between the agglutinating activity of abrin-a and development of apoptosis to DNA fragmentation. The time-dependent effects of abrin-a on apoptosis as well as its time-dependent penetration into the cells suggest that the B-chain probably triggers the apoptosis, while the A-chain and breakage of the disulfide bond are responsible for its progress. D 2004 Elsevier Inc. All rights reserved. Keywords: Abrin-a; Apoptosis; Phosphatidylserine translocation; Caspase activity; DNA fragmentation; Acute lymphoblastic leukemia
Introduction Abbreviations: ALL, acute lymphoblastic leukemia; FITC, fluorescein isothiocyanate; PBS ( ), phosphate-buffered saline (Ca2+ and Mg2+ free); PSer, phosphatidylserine. * Corresponding author. Natural Substance-Composed Materials Group, National Institute for Advanced Industrial Science and Technology, AIST-Kyushu, 807-1 Shuku-machi, Tosu, Saga-ken 841-0052, Japan. Fax: +81-942-81-3690. E-mail address: [email protected] (H. Ohba). 0041-008X/$ - see front matter D 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.taap.2003.11.018
Abrin-a, obtained from Abrus precatorius seeds, belongs to the family of type II ribosome inactivating proteins— heterodimeric glycoproteins, consisting of two nonidentical polypeptide chains (A- and B-chain) cross-linked through a single disulfide bond (Liu et al., 2000; Wei et al., 1974). The A-chain affects the protein synthesis by modification of the
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ribosomal subunits of the eukaryotic cells (Endo et al., 1987; Hartley et al., 1991; Hegde et al., 1993). The B-chain is a carbohydrate-binding protein and serves to bind the abrin-a molecule specifically with D-galactosyl-containing cell-surface receptors (Hegde and Podder, 1997; Olsnes et al., 1974). It has been found that abrin-a is a promising tool for distinguishing of cancer cells from normal ones as a result of its higher affinity to the cancer cells. The drug attracts a great deal of interest because of its unique biological activities as cytoagglutination and cytotoxicity, accompanied with induction of apoptosis, and all these effects are predominantly expressed in cancer cells. (Geier et al., 1996; Keppler-Hafkemeyer et al., 1998; Moriwaki et al., 2000; Shin et al., 2001). The accent of the present study is on the abrin-a-mediated mechanism(s) of apoptosis induction in leukemic cells. The apoptotic cascade in mammalian cells is a multistep process, initiating in most cases by loss of integrity of the mitochondrial membrane. This usually involves the activation of permeability mitochondrial transition pores, loss of mitochondrial membrane potential (
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of apoptosis: phosphatidylserine (PSer) translocation from the inner to the outer monolayer of plasma membrane as an early event of apoptosis, activation of caspases as a moderate phase, and internucleosomal DNA fragmentation as a last phase (Adams and Cory, 2002; Beauparlant and Shore, 2003; Collins et al., 1997; Zhang and Xu, 2000).
Materials and methods Reagents. Abrin-a was purified from the seeds of A. precatorius, grown in Thailand, by the method of Lin et al. (1980) and was stored as ammonium sulfate precipitate at 4 jC until use. Apoptosis Ladder Detection Kit, Wako, and fetal calf serum (FCS) were purchased from Wako Pure Chemical Industries, Ltd., Osaka. CapACE Assay System, Colorimetric was purchased from Promega (Madison, WI). ApoAlert Caspase-8 Fluorometric Assay Kit and ApoAlert Caspase-9/6 Fluorescent Assay Kit were obtained from Clontech Laboratories, Inc., (Palo Alto, CA). Fluorescein isothiocyanate (FITC)– Annexin V was obtained from Pharmingen. Antibiotics were obtained from Gibco GRL (Grand Island, NY). All chemicals were of analytical grade. Abrin-a. Abrin-a solution was prepared by dialyzing of ammonium sulfate precipitate of abrin-a against phosphatebuffered saline [PBS ( ), Ca2+ and Mg2+ free] at 4 jC before use and filtered through a Millipore filter (0.2 Am). The concentration of abrin-a was measured spectrophotometrically, using an absorption coefficient A1cm1% = 15.9 at 280 nm (Endo et al., 1987). Cultured leukemic cells. The human leukemic cell lines [Jurkat, MOLT-4, HPB-acute lymphoblastic leukemia (ALL), and CCRF-CEM] were cultured in RPMI-1640 medium, supplemented with 10% heat-inactivated FCS, and 100 Ag/ml of streptomycin and 100 U/ml of penicillin in a humidified atmosphere at 37 jC with 5% CO2. These cells were generously provided by Dr. J. Minowada (Hayashibara Biochemical Laboratories, Inc., Okayama, Japan). Abrin-a uptake into the cells. The penetrating of abrin-a into the cells was detected by confocal fluorescent microscopy using fluorescein isothiocyanate (FITC) as a fluorescent marker. FITC was conjugated with abrin-a and then served as a fluorescent probe to detect the drug penetrating into the cells in a time-dependent manner. Cells (1.0 106 cells/well) were incubated with 0.3 AM FITC– abrin-a at different time intervals. The samples were analyzed by ‘‘Olympus IX70’’ confocal microscope. Flow cytometric assay. The viability of cells were analyzed by flow cytometer Beckman Coulter—Epics XL, operating in accordance with the manufacturer’s recommendations after fine adjustments for optimization. The forwardscatter (FS) and side-scatter (SS) parameters were adjusted to accommodate the inclusion of viable and dead leukemic cells within the acquisition data. No cells excluded from the
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analysis and 10,000 cells were counted. Data were collected and analyzed by ‘‘XL System II’’ software. The results were presented as a dot plot of cell fluorescence with quadrant markers drawn to distinguish viable and dead cells, as well as to distinguish leukemic and normal cells. Quadrant A in Fig. 1 contains viable normal cells, quadrant B contains viable leukemic cells, quadrant C contains all viable cells, and quadrant D contains all dead cells. The percentage of dead cells before and after abrin-a treatment was calculated and presented in the table. DNA-fragmentation assay. Cells (1.0 106 cells/well) were cultured as described above. After 24 h of cultivation, cells were incubated with 2-fold dilution series of abrin-a (maximum concentration 8 AM) in PBS ( ) for 24 h under
the same conditions. For the time-dependent analysis of DNA fragmentation, cells were incubated with abrin-a (final concentration 0.3 AM) and the DNA fragmentation was determined in the range of 0 – 24 h. At each time point, the cells were collected and cell lysis and DNA extraction were provided using commercially available kit for DNA-fragmentation assay (Apoptosis Ladder Detection Kit, Wako) based on the method of Ioannou and Chen (1996). The extracted DNA was analyzed by electrophoresis on agarose gels (1.5% gel density) using TBE buffer. The sample, in which PBS ( ) was added instead of abrin-a solution, was assumed to be a contrast. DNA from apoptotic cells forms a ladder on the agarose gel, while DNA from non-apoptotic cells appears as a single band or smeared on the agarose gel. Presence of random DNA fragmentation (smear) is a characteristic of necrosis.
Fig. 1. Agarose gel electrophoresis of DNA extracted from acute T-lymphoblastic leukemia cells after treatment with abrin-a in different concentrations. (A) DNA-fragmentation assay. The cells (2.22 104 cells/well) were cultured in the presence of 2-fold dilution series of abrin-a (8 AM maximal concentration) as it is described in Materials and methods. After 24 h incubation the cells were collected by centrifugation and DNA extraction was carried out using DNAfragmentation assay kit (Apoptosis Ladder Detection Kit, Wako). The DNA was analyzed by TBE electrophoresis on 1.5% agarose gel. The sample, in which PBS ( ) was added instead of abrin-a solution, was considered as a control. Blots from one typical experiment are presented in the figure. St, standard marker; lane 1, control (nontreated cells); lanes 2 – 11, cells treated with 0.016 – 8 AM abrin-a, respectively. (B) Flow cytometry. The percentage of dead cells before and after abrin-a treatment was determined by flow cytometry and the results are presented in the table (*P < 0.05, **P < 0.01, ***P < 0.001 vs. the respective control nontreated cells, C). A typical histogram as a dot plot of cell fluorescence with quadrant markers drawn to distinguish viable and dead cells, as well as to distinguish leukemic and normal cells, is also shown in the figure. Quadrant A contains viable normal cells, quadrant B contains viable leukemic cells, quadrant C contains all viable cells, and quadrant D contains all dead cells.
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PSer detection assay. Cells (1.0 106 cells/well) were incubated with abrin-a (final concentration 0.3 AM) in PBS ( ) for 0 –3 h under the same conditions as mentioned above. At each time point the cells were collected by centrifugation (450 g, 20 min), washed twice with PBS ( ) containing 2.5 mM CaCl2 (Annexin V-binding buffer), and resuspended in the same buffer. After that, 100 Al of cell suspension was incubated with 5 Al of FITC-conjugated Annexin V (FITC –Annexin V) for 10 min at room temperature (RT) in a dark place. The cells were washed twice with Annexin V-binding buffer and were resuspended in 100 Al of the same buffer. FITC – Annexin V, bound to PSer exposed on the cell surface, was detected spectro-
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fluorimetrically at kem = 535 nm (kex = 488 nm) using Fluoromark Fluorescent Microplate Leader (BioRad). Cell lysis. Cells (1.0 106 cells/well) were incubated with abrin-a (final concentration 0.3 AM) in the range 0 –4 h. At each time point, the cells were sedimented by centrifugation at 450 g for 10 min at 4 jC, washed three times with cold PBS ( ), and lysed by lysis buffer (20 mM HEPES/KOH, pH 7.5, 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, 1 mM EGTA, 1 mM DTT, 5 Ag/ml pepstatin A, 10 Ag/ml leupeptin, 2 Ag/ml aprotinin, 0.1% Triton X-100) and freezing – thawing. Freeze –thaw cycles were repeated three times to ensure complete cell lysis. After centrifugation at 15,000 g for 20
Fig. 2. Agarose gel electrophoresis of DNA extracted from acute T-lymphoblastic leukemia cells after treatment with abrin-a at different time intervals. (A) DNA-fragmentation assay: The cells (1 106 cells/well) were cultured in the presence of abrin-a (0.3 AM) as described in Materials and methods section. At each time point, the cells were collected by centrifugation and DNA analysis was carried out as it is described in Fig. 1. The sample, in which PBS ( ) was added instead of abrin-a solution, was considered as a control (C, nontreated cells). Blots from one typical experiment are presented in the figure. St, standard marker. (B) Flow cytometry. The percentage of dead cells before and after abrin-a treatment (0.3 AM) was determined by flow cytometry at different time intervals and the results are presented in the table (*P < 0.05, **P < 0.01, ***P < 0.001 vs. the respective control nontreated cells, C).
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min at 4 jC, the cell lysates were used for analysis of caspase isoenzyme activity.
FMK) in PBS ( ) were prepared analogous to described above.
Analysis of caspase isoenzymes. Cells, cultured in the absence or in the presence of abrin-a (3 AM), were washed twice by PBS ( ) and lysed in RIPA-buffer. Lysates were centrifuged at 16,000 g for 15 min at 4 jC. Protein concentrations were determined on supernatants using a protein microassay of the Bradford method (Bio-Rad). Cell lysates (obtained from viable, apoptotic, necrotic, and apocrotic cells) were analyzed for caspase activity by two procedures: (i) Western blot analysis as described in Andreeff et al. (1999) and Towbin et al. (1979), and (ii) spectrophometrically or spectrofluorimetrically, detecting the products of the caspase-catalyzed reactions. Enzyme activities and protein expression were determined in cell lysates, obtaining from all cell forms— viable, apoptotic, necrotic and apocrotic.
Spectrofluorimetric detection of caspase-9 activity. Fifty microliters of the cell lysates were incubated with 50 Al of 2 reaction buffer/DTT (ApoAlert Caspase-9/6 Fluorometric Assay Kit, Clontech), 5 Al of substrate solution (5 mM Leu-Glu-His-Asp-AMC) and 2 Al of DMSO for 1 h at 37 jC. The release of 7-amino-4-methyl coumarin (AMC), catalyzed by caspase-9, was estimated fluorimetrically at kem = 490 nm (kex = 390 nm) using Fluoromark Fluorescence Microplate Leader (BioRad). As a negative control, cells treated with abrin-a plus caspase inhibitor (0.5 mM Leu-Glu-Thr-Asp-CHO) in PBS ( ) were prepared analogous to described above.
Western blot. Briefly, 60 mg cell protein extract was dissolved 1:1 in 2 Laemmli sample buffer and the samples were subjected to electrophoresis on SDS-polyacrylamide gel (5% stacking, 4 –20% resolving). Electrophoresis was carried out in two steps: at 80 V for 15 min and 120 V for 2 h at room temperature (RT approximately 22 jC). The separated protein fractions were transferred to a Hybond-P PVDF membranes (Amersham Bioscience) during 18 h at 35 V, 4 jC. The transferred proteins were immunoblotted using adequate primary anti-caspase antibodies or anti-hactin antibody (for comparison), and subsequent secondary antibodies (Andreeff et al., 1999; Towbin et al., 1979). Five and seven independent experiments were carried out for caspase-3 and -9, respectively. Spectrophotometric detection of caspase-3 activity. Fiftyfour microliters of the cell lysates were incubated with 32 Al of caspase assay buffer (20 mM HEPES/KOH, pH 7.5, 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, 1 mM EGTA), 2 Al of DMSO, 10 Al of 100 mM DTT and 2 Al of substrate solution (1 mM Ac-Asp-Glu-Val-Asp-pNA) for 15 min at 37 jC. The release of chromophore pnitroaniline, catalyzed by caspase-3, was monitored spectrophotometrically at 405 nm, using an Inter Med Immunomini NJ-2300 multiwell scanning spectrophotometer. As a negative control, cells treated with abrin-a plus caspase inhibitor (50 AM Z-Val-Ala-Asp-FMK) in PBS ( ) were prepared at the same manner as described above. Spectrofluorimetric detection of caspase-8 activity. Fifty microliters of the cell lysates were incubated with 50 Al of 2 reaction buffer/DTT (ApoAlert Caspase-8 Fluorometric Assay Kit, Clontech) and 5 Al of substrate solution (1 mM Ile-Glu-Thr-Asp-AFC) for 1 h at 37 jC. The release of 7amino-4-trifluoromethyl coumarin (AFC), catalyzed by caspase-8, was estimated fluorimetrically at kem = 535 nm (kex = 390 nm) using Fluoromark Fluorescence Microplate Leader (BioRad). As a negative control, cells treated with abrin-a plus caspase inhibitor (5 AM Ile-Glu-Thr-Asp-
Measurement of cytotoxicity. The cytotoxicity of abrin-a against leukemic cells was measured by CellTiter-GloTM luminescent cell viability assay (Promega) (Crouch, 1993). The ATP bioluminescence was used as a marker of cell proliferation and viability. Briefly, the lectins were dissolved in PBS ( ) and were dispensed as 10-Al aliquots into 96-well microplates. The cells were suspended in the culture medium to a concentration 5 105 cells/ml and then were added in 90-Al aliquots to each patch. After 24
Fig. 3. Changes in the cell-surface phosphatidylserine (PSer) induced by abrin-a in acute T-lymphoblastic leukemia cells. The cells (1 106 cells/ well) were incubated with abrin-a (0.3 AM) for 0 – 3 h, collected by centrifugation, and incubated with FITC – Annexin V in PBS ( ), containing 2.5 mM CaCl2 (Annexin V-binding buffer) for 10 min at RT in a dark place. The cells were washed twice by Annexin V-binding buffer and were resuspended in the same buffer (the cell concentration was identical in all samples). The detection of FITC – Annexin V, bound to PSer exposed on a cell surface, was carried out spectrofluorimetrically at kex = 488 nm and kem = 525 nm. The intensity of fluorescence (in arbitrary units) is expressed in the figure. The results are mean F SD from six independent experiments (n = 6). In the case of MOLT-4, CCRF-CEM, and Jurkat, the increase of fluorescence after 1, 2 and 3 h treatment with abrin-a was statistically significant ( P < 0.001 vs. control nontreated cells). In the case of HPB-ALL, the changes in fluorescence intensity were statistically significant only at 3-h treatment with abrin-a ( P < 0.05 vs. control nontreated cells).
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h incubation at 36.9 jC in cell incubator (5.1% CO2), CellTiter 96 kit (Promega) was added in aliquots of 100 Al to each patch and incubated with cell suspensions for 1 h, following the procedure, recommended by the producer. The luminescence, produced by luciferase catalyzed lucif-
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erin + ATP reaction, was measured by an Inter Med Immunomini NJ-2300 multiwell scanning spectrophotometer. The data were normalized to the control group. The abrin-a cytotoxicity was calculated as percentage decrease of luminescence in comparison with the control.
Fig. 4. Caspase-3 activity in acute T-lymphoblastic leukemia cells after treatment with abrin-a. The cells (1.0 106 cells/well) were incubated with abrin-a (0.3 AM). At each time point, the cells were sedimented, washed with cold PBS ( ), lysed, and detection of caspase-3 activity was carried out as follows: (A) The release of chromophore p-nitroaniline, catalyzed by caspase-3, was monitored spectrophotometrically at 405 nm. As a negative control, cells treated with abrina in the presence of caspase-3 inhibitor (Z-Val-Ala-Asp-FMK) were used. Closed symbols, in the absence of caspase-3 inhibitor; open symbols, in the presence of caspase-3 inhibitor. The results are mean F SD from seven independent experiments (n = 7). In the case of Jurkat, HPB-ALL and MOLT-4, P < 0.01 vs. control group, at 1 h; P < 0.001 vs. control group, at 2 – 4 h. In the case of CCRF-CEM, nonsignificant (ns), at 1 h; P < 0.05 vs. control group, at 2 – 4 h. (B) Western blot analysis of caspase-3 was carried out after 1-h incubation of cells with abrin-a. Forty milligrams of protein was applied in electrophoresis. Blots from one typical experiment are presented in the figure. C, in the absence of abrin-a (control); 1, 1 h after addition of abrin-a (0.3 AM) to the cell suspensions. Densitograms correspond to the immunoblot analysis and the results are expressed as mean F SD from five independent experiments.
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Cytoagglutination assay. Cytoagglutination was evaluated by the method of Ohba et al. (1997) using a Jasco V-550 spectrophotometer equipped with a magnetic stirrer. Briefly, the cells were washed three times by PBS ( ) and resuspended in the same buffer to a concentration 2 106 cells/ ml. The cell suspension and PBS ( ) were placed into the sample and reference cuvettes, respectively, and were incubated until the baseline at 600 nm became constant. A solution of abrin-a was added to the cell suspension in different concentrations and the decrease in turbidity at 600 nm (OD600) was recorded under continuous stirring. The measurement was stopped when the OD600 reached to a plateau. The velocity of cytoagglutination (
and 6 h, respectively, after addition of abrin-a to the cell suspensions, and the density of the bands increased with the time of treatment (Figs. 2a, d). The same tendency was established in the increase of percentage of dead cells, estimated by flow cytometry (Fig. 2, table). It may also be speculated that necrosis (smear) proceeded apoptosis (ladder) and the cell death was a result of apoptosis development. In HPB-ALL and MOLT-4, the step ladder of DNA was not detected even 24 h after addition of abrin-a to the cell suspensions (Figs. 2b, c). Only a smear was detected and probably the cell death was a result of necrosis. However, the percentage of dead cells markedly increased after 24-h treatment.
Statistical analysis. One-way analysis of variance (ANOVA) was employed, following by Bonferroni’s test for significant differences. Statistical significance was defined at P < 0.05. The statistical procedures were performed with GraphPad InStat software. Data are expressed as mean F SD.
The fluorescence of FITC – Annexin V was detected in the target cells as an index for translocation of PSer from the inner to the upper monolayer of the cell-surface membrane (flip-flop), which happens during the first stage of apoptosis (Van Engeland et al., 1998; Vermes et al., 1995). FITC – Annexin V specifically recognizes PSer– Ca2+ complexes and is widely used to detect the early apoptotic cells. It was established that the fluorescence intensity began to increase in CCRF-CEM, Jurkat, and MOLT-4 1 h after addition of abrin-a (Fig. 3). On the other hand, in HPB-ALL, fluores-
Results
Change in cell-surface PSer after abrin-a application
Dose-dependent effects of abrin-a on the DNA fragmentation DNA fragmentation in abrin-a-treated cells was used as a conventional marker of the final step of apoptosis and as a major characteristic of apoptotic cell death. The results in Fig. 1 demonstrate that even low concentrations of abrin-a (0.01 –0.06 AM) induced DNA fragmentation in Jurkat and CCRF-CEM cells—a step ladder of DNA fragmentation was detected (Fig. 1d, lanes 2– 11; Fig. 1a, lanes 5– 11). The process of DNA fragmentation was better and clearly manifested in the higher concentrations of abrin-a. The strength of bands markedly enhanced and they were well defined at abrin-a concentrations >0.5 AM. In HPB-ALL and MOLT-4, a ladder of DNA fragmentation was not detected even at high concentrations of abrin-a (Figs. 1b, c). The percentage of dead cells, determined by flow cytometry, also increased with increasing abrin-a concentration (Fig. 1, table). The results were statistically significant at 0.12 AM abrin-a and higher, even in MOLT-4 and HPB-ALL, in which DNA fragmentation was not detected. Time-dependent effects of abrin-a on the DNA fragmentation The time-dependent effects of abrin-a on the different phases of apoptosis were determined at concentration 0.3 AM. In Jurkat and CCRF-CEM, the fragmentation of DNA was better and clearly manifested with the time of cell treatment by abrin-a. DNA fragmentation was detected 4
Fig. 5. Caspase-8 activity in acute T-lymphoblastic leukemia cells after treatment with abrin-a. The cells (1.0 106 cells/well) were incubated with abrin-a (final concentration 0.3 AM). At each time point, the cells were sedimented, washed with cold PBS ( ), lysed, and detection of caspase-8 activity was carried out as it is described in Materials and methods. The release of 7-amino-4-trifluoromethyl coumarin (AFC), catalyzed by caspase-8, was estimated fluorimetrically at kem = 535 nm (kex = 390 nm). As a negative control, cells treated with abrin-a in the presence of caspase-8 inhibitor (Ile-Glu-Thr-Asp-FMK) were used. Closed symbols, in the absence of caspase-8 inhibitor; open symbols, in the presence of caspase-8 inhibitor. The results are mean F SD from six independent experiments (n = 6). In the case of Jurkat and HPB-ALL, ns, at 1 h; P < 0.001 vs. control group, at 2 – 3 h. In the case of MOLT-4, ns, at 1 – 2 h; P < 0.01 vs. control group, at 3 h. In the case of CCRF-CEM, ns, at 1 – 3 h.
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1.5
0.5
Fig. 6. Caspase-9 activity in acute T-lymphoblastic leukemia cells after treatment with abrin-a. The cells (1.0 106 cells/well) were incubated with abrin-a (final concentration 0.3 AM). At each time point, the cells were sedimented, washed with cold PBS ( ), lysed, and detection of caspase-9 activity was carried out as follows: (A) The release of 7-amino-4-methyl coumarin (AMC), catalyzed by caspase-9, was estimated fluorimetrically at kem = 490 nm (kex = 390 nm). As a negative control, cells treated with abrin-a in the presence of caspase-9 inhibitor (Leu-Glu-Thr-Asp-CHO) were used. Closed symbols, in the absence of caspase-9 inhibitor; open symbols, in the presence of caspase-9 inhibitor. The results are mean F SD from eight independent experiments (n = 8). In the case of Jurkat and HPB-ALL, ns, at 1 h; P < 0.001 vs. control group, at 2 – 3 h. In the case of MOLT-4, ns, at 1 – 2 h; P < 0.001 vs. control group, at 3 h. In the case of CCRF-CEM, ns, at 1 – 3 h. (B) Western blot analysis of caspase-9 was carried out as described in Materials and methods. Forty milligrams of protein was applied in electrophoresis. Blots from one typical experiment are presented in the figure. C, in the absence of abrin-a (control); 1, 3 h after addition of abrin-a (0.3 AM) to the cell suspensions. No active caspase-9 was detected after 1-h treatment with abrin-a (data not shown). Densitograms correspond to the immunoblot analysis and the results are expressed as mean F SD from six independent experiments.
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cence intensity increased slightly at the third hour after abrin-a application. Activation of caspase isoenzymes During induction of apoptosis, several caspases (cysteine proteases) are activated by the signal transmission in the cells (Beauparlant and Shore, 2003). In caspase cascade, caspase-3 is a key enzyme in the apoptotic pathway. The activation of caspase-3 happens in the downstream of the signal transmission and is responsible for DNA cleavage. It was established that the caspase-3 activity increased after addition of abrin-a, and it reached a maximum 2 h after beginning of treatment in Jurkat and HPB-ALL, and 3 h after that in MOLT-4 (Fig. 4). In CCRF-CEM, the caspase-3 activity only increased gradually, but slightly, until fourth
hour after abrin-a application. An increase in the enzyme activity was not detected in the presence of caspase-3 inhibitor in all cell lines. To investigate in more details the signal transmission mechanism of apoptosis, induced by abrin-a, we analyzed the activation of caspase-8 and -9, usually proceeding the activation of caspase-3 in the cascade. The results are presented in Figs. 5 and 6. In Jurkat, the caspase-8 and -9 activity increased rapidly on the second hour after addition of abrin-a. Caspase-8 activity reached a plateau after 2h incubation of Jurkat with abrin-a, while caspase-9 activity gradually increased until the third hour. In HBPALL, the caspase-8 and -9 activity increased rapidly and reached a plateau on the second hour after addition of abrin-a. In CCRF-CEM and MOLT-4, the caspase-8 and -9 manifested a poor activity even after 3 h of cell treatment with abrin-a.
Fig. 7. Time-dependent penetrating of abrin-a into Jurkat (A) and CCRF-CEM (B) cells. The cells (1.0 106 cells/well) were incubated with FITC – abrin-a (final concentration 0.3 AM). At each time point, the cells were sedimented, washed with PBS ( ) and FITC – abrin-a, bound to the cell surface or penetrating into the cells, was measured by confocal fluorescent microscopy (green light). In parallel, propidium iodide (PI) staining was used as a marker of cell survival (red light). Note that in Jurkat cell aggregation takes place during abrin-a treatment.
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Penetrating of abrin-a into the cells and induction of apoptosis Using FITC-labeled abrin-a and propidium iodide (PI) staining, we detected in parallel the time-dependent penetrating of the drug into the cells and development of apoptosis. The green light in Fig. 7 is representative for FITC-labeled abrin-a and the red light is representative for penetrating of propidium iodide into the cells, which is a characteristic of apoptotic cell death. In Jurkat, it was observed that abrin-a (0.3 AM) penetrated into the cells after 2-h incubation, while propidium iodide was detected into the cells on the third hour. On the fourth to fifth hour, cell aggregation was also observed. The results suggests that Jurkat cells are very sensitive to abrin-a and induction of apoptosis takes place immediately after its cellular uptake. In CCRF-CEM, abrin-a penetrated into the cells on the second hour, while propidium iodide was detected into the cells on the sixth hour. It seems that CCRF-CEM cells are most resistant to abrin-a and several hours are necessary to detect the pro-apoptotic effect of the drug in this cell line.
Discussion Apoptosis is a programmed cell death, an essential mechanism for cell removal during embryonic development and elimination of lymphocytes during thymic development (Majno and Joris, 1995; Trump et al., 1997). To date, several endogenous physiological mediators of apoptosis have been identified. There are many data, demonstrating that some environmental factors are also responsible for induction of apoptosis, as radiation, viruses, drugs, etc., and all these candidates act by a similar route (Kopecky et al., 2001; Zhang et al., 2001). It is widely accepted that apoptosis is characterized by several biochemical events: (i) changes in the lipid composition of the cell-surface membrane resulted in translocation of PSer from the inner to the outer monolayer as an early event of apoptosis; (ii) disturbance of mitochondria and release of cytochrome c in cytosol, which serves as a cofactor, together with the apoptotic protease activating factor, to activate pro-caspase-9; (iii) pro-caspase-9 unlocks caspase cascade, resulting in caspase-3 activation, responsible for DNA fragmentation as a final stage. Recently, it has been reported that caspase-3 can be activated through two alternative pathways: from caspase-8 directly, or caspase8 activates pro-caspase-9, which serves as activator of caspase-3 (Hegde et al., 1998; Luo et al., 1998; Pan et al., 1998). However, there are no data confirming that both pathways exist and act in parallel in one and the same time. In this study, we established that abrin-a induces apoptosis in leukemic cells and the process goes by different mechanisms, depending on the cell line. In Jurkat, the apoptosis followed the conventional pathway—PSer trans-
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location on the cell surface, caspase activation, and DNA fragmentation. In CCRF-CEM, the apoptosis began with PSer translocation on the cell surface and went to DNA fragmentation without any induction of caspase activity. In MOLT-4, the apoptosis was initiated and characterized with PSer exposure and caspase activation, but does not progress to DNA fragmentation. In HPB-ALL, the translocation of PSer was negligible, the activity of caspases enhanced markedly, but DNA-fragmentation was not detected. The time-dependent effects of abrin-a on the development of apoptosis demonstrated that PSer translocation preceded the penetrating of drug into the cells (Fig. 7). Because the B-chain is responsible for the binding of abrin-a to the cell surface, the interaction between B-chain and carbohydrates can be considered as one of the trigger signals of early apoptotic events. Recently, it has been reported that PSer synthesis by serine-base exchange enzyme system, in the endoplasmic reticulum of mammalian cells, is able to regulate PSer exposure in cells undergoing the apoptotic process (Pelassy et al., 2000). PSer synthesis is modulated by alterations of the plasma membrane potential that modify the activity of the plasma membrane serine transporter (Breintmayer et al., 1996). The lectins, including abrin-a, are well-known substances, affecting the plasma membrane potential and therefore affecting PSer synthesis and its membrane translocation. However, in HPB-ALL, we did not observe significant changes in PSer exposure at the cell surface. The caspase-3 activation also preceded the penetrating of abrin-a into the cells, as well as the activation of caspase8 and -9, which is in contrast with the commonly accepted hypothesis that abrin-a crosses the plasmatic membrane of tumor cells by endocytosis, activates directly caspase-9, followed by activation of caspase-3. In our study, we established that caspase-3 activity increased on the first hour after abrin-a application and reached a maximum on the second hour in Jurkat and HPB-ALL, and on the third hour in MOLT-4. In contrast, caspase-8 and -9 began to increase on the second hour after abrin-a application and reached a maximum on the third hour in Jurkat and HPBALL. In MOLT-4, caspase-8 and -9 activity increased on the third hour, however the effect was lower that in Jurkat and HPB-ALL. In CCRF-CEM, the activities of caspase-8 and 9 were negligible and only the activity of caspase-3 increased slightly. The results suggest that probably B-chain-dependent signaling is responsible for caspase-3 direct activation by abrin-a in Jurkat, MOLT-4, and HPB-ALL. On the second hour after the beginning of abrin-a application, the drug was detected into the cells and it is impossible to clarify the role of both B- and A-chains in induction of caspase-8 and-9 activity, as well as in the continuously increase of caspase-3 activity. Moreover, it was found that the activation of caspase cascade and particularly caspase-3 is not enough for induction of DNA fragmentation, as well as it is not obligatory
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event, proceeding DNA fragmentation. DNA fragmentation was detected only in Jurkat and CCRF-CEM, but not in MOLT-4 and HPB-ALL cells. In Jurkat and CCRF-CEM, the abrin-a caused the apoptosis at low drug concentration and in a short time of treatment (4 h in Jurkat and 6 h in CCRF-CEM). In CCRF-CEM, abrin-a induced DNA fragmentation without caspase activation—a mechanism different from an already known apoptotic route. Presumably, apoptosis-inducing factor (AIF), which has been characterized as a caspase-independent death effector, participates in the development of apoptosis in CCRF-CEM cells. On our knowledge, there are no data about the effect of abrin-a on the translocation of AIF from mitochondria into the nucleus and this hypothesis is speculative, but it seems to be likely and its proof is in our future efforts. The pro-apoptotic effect of abrin-a can also explain its possibility to induce indirectly oxidative stress in leukemic cells. Separation of A-chain and B-chain of abrin-a molecule, which requires a breakage of –S – S– bond, has a potential to destroy the balance of antioxidant thiol-containing substances and to provoke free radical production in the cells. Free radicals are well-known apoptotic signals. Moreover, recently, it has been reported that abrin-a inactivates 30-kDa thiol-specific antioxidant protein and thus can decrease antioxidant defense potential of the cells (Shin et al., 2001). The differences in caspase activation and DNA fragmentation between the leukemic cell lines used can also explain the different amount and type of abrin-a-binding receptors on the cell surface. Different signals can be triggered by abrin-a, depending on the degree of abrin – cell binding and the type of this bound, analogous to other lectins (Danguy et al., 2002). In our previous paper (Moriwaki et al., 2000), we demonstrated that abrin-a manifests target cytoagglutinating and cytotoxic activity against leukemic cells and the drug is available as a marker of malignancy. Comparing the present results with previously published data, an interesting fact was mentioned: DNA fragmentation was observed only in cell lines in which abrin-a manifested a high cytoagglutinating activity (Fig. 1, Table 1). Probably, the cytoagglutination and changes in the charge of the cell surface by abrin-a are trigger signals, programming the apoptosis for development to the final stage. Abrin-a also manifested a high cytotoxicity against Jurkat and CCRF-CEM, followed by HPBALL and MOLT-4. In conclusion, the results shown that the abrin-a induces apoptosis in leukemic cells by different mechanisms, depending of the cell line. The time-dependent effects of abrin-a on apoptosis, as well as its time-dependent penetrating into the cells suggest that the B-chain is probably responsible for induction of apoptosis, while the A-chain and breakage of the disulfide bond defined its progress (Shin et al., 2001). Because a very good positive correlation between the cytoagglutinating activity of abrin-a and development of apoptosis to DNA fragmentation was found, it
Table 1 Effect of abrin-a on the velocity of cytoagglutination, cell viability, and induction of apoptosis in acute lymphoblastic leukemia T-cell lines Cell line
Velocity of cytoagglutination (DOD600 nm/min)a
Cell viability (% from control)a
DNA fragmentation
Jurkat CCRF-CEM MOLT-5 HPB-ALL
1.46 0.90 0.28 0.18
20 28 68 53
++ +
a
F F F F
0.09 0.03 0.01 0.01
F F F F
4 7 12 11
According to Moriwaki et al. (2000).
seems that the B-chain-mediated signaling is also involved in the programming of the development of apoptosis to the last phase or its termination to the phase of caspase activation.
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