Cardiotoxin-III selectively enhances activation-induced apoptosis of human CD8+ T lymphocytes

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Toxicology and Applied Pharmacology 193 (2003) 97–105

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Cardiotoxin-III selectively enhances activation-induced apoptosis of human CD8⫹ T lymphocytes Shu-Hui Su,a Shu-Jem Su,b Shinne-Ren Lin,c and Kee-Lung Changd,* a

Department of Cosmetic Science, Chung Hwa College of Medical Technology, Tainan 717 Taiwan b Department of Medical Technology, FooYin University, Kaohsiung 831 Taiwan c School of Chemistry, Kaohsiung Medical University, Kaohsiung 807, Taiwan d Department of Biochemistry, Kaohsiung Medical University, Kaohsiung 807, Taiwan Received 20 December 2002; accepted 7 July 2003

Abstract Cardiotoxin-III (CTX-III), a major cardiotoxin isolated from the venom of the Taiwan cobra (Naja naja atra), is a highly basic, hydrophobic, toxic protein, which can induce lysis of mononuclear cells by an unknown mechanism. This study was undertaken to investigate the effects of CTX-III on untreated and PHA-activated peripheral blood mononuclear cells (PBMCs) in vitro. The results show that treatment of PHA-activated lymphocytes with CTX-III (10 ␮g/ml) induced apoptosis and depletion of the CD8⫹ population. In both untreated and PHA-treated lymphocytes, interferon-␥ production was dramatically reduced and interleukin-2 (IL-2) production was moderately reduced by CTX-III treatment. In PHA-activated lymphocytes, CD4 expression was increased, whereas CD8 and IL-2R ␤ chain (CD25) expression were decreased. In contrast, CTX-III had no effect on the viability of PHA-activated monocytes but significantly enhanced their tumor necrosis factor-␣ production. These results show that CTX-III selectively enhanced activation-induced apoptosis in CD8⫹ T cells. CTX-III was found to bind to the cell membrane of PHA-stimulated PBMCs, and three CTX-III-binding proteins, with molecular weights of 92, 77, and 68 kDa, were identified. We therefore propose that CTX-III interacts with one or more cell surface proteins and initiates a signal pathway causing functional changes. These findings provide an insight into the immunomodulatory properties of CTX-III and suggest a novel method for the selective induction of apoptosis in CD8⫹ T lymphocytes. © 2003 Elsevier Inc. All rights reserved. Keywords: Cardiotoxin; Apoptosis; CD8⫹ T lymphocyte; Cytokine

Introduction Cardiotoxins (CTXs), highly basic polypeptides from cobra venom (Lee, 1979), make up 60% of the total venom protein and have a wide variety of biological activities, including cardiac muscle cell contraction, erythrocyte lysis, skeletal muscle necrosis, selective toxicity for certain tumors, and inhibition of the activity of enzymes, such as the Na⫹, K⫹-ATPase and protein kinase C (Kumar et al., 1996, 1997; Lee et al., 1998; Ownby et al., 1993). CTXs may damage cells by their ability to interact with the cell mem* Corresponding author. Department of Biochemistry, Kaohsiung Medical University. No. 100 Shih-Chuan 1st Rd., Kaohsiung 807, Taiwan. Fax: ⫹886-7-397-2257. E-mail address: [email protected] (K.-L. Chang). 0041-008X/$ – see front matter © 2003 Elsevier Inc. All rights reserved. doi:10.1016/S0041-008X(03)00327-2

brane or oligosaccharides (Bougis et al., 1982; Chien et al., 1994; Ksenzhek et al., 1978; Patel et al., 1997; Sue et al., 1997). In addition, they are cytolytic for cells from various species and for both normal and transformed cells (Iwaguchi et al., 1985). Different cell types show differential susceptibility to CTXs (Hinman et al., 1987, 1990). More than 50 CTXs have been purified from different cobra venoms and their amino acid sequences determined. The functional heterogeneity of CTXs in terms of the lysis of different cell types may be due to relatively minor structural differences between CTXs (Menez et al. 1990). CTXs have been used to study muscle regeneration, to improve the efficiency of HIV-1 DNA vaccines by their myonecrotic effect (Fomsgaard, 1999; Rajnoch et al., 2001), and to target active compounds to the cell membrane in anticancer therapy, one example being the cardiotoxin– crotoxin conjugate,

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VRCTC-310 (DeTolla et al., 1995). Before CTXs can be used safely in the clinic, information about the toxic and biological effects of the individual variants is required. One of the CTXs, CTX-III from the Taiwan cobra (Naja naja atra) (Lee, 1991), is a 60-amino acid ␤-sheet protein (Sivaraman et al., 1998), which contains different binding and cytolytic domains and is cytolytic for heart cells and human leukemic T cells (Sivaraman et al., 1998; StevensTruss et al., 1996a, 1996b, 1997). CTX-III is known to cause lysis of human lymphocytes (Stevens-Truss et al., 1997), but there have been few reports on its effects on other immune cells (Stevens-Truss et al., 1996b; Xiao and Hinman, 1990). In the present study, we examined its effects on cytokine production and cell surface molecule expression by human peripheral blood mononuclear cells (PBMCs) and whether it induces apoptosis of these cells. The results show that CTX-III selectively enhances activation-induced apoptosis of CD8⫹ T lymphocytes.

Materials and methods Purification of CTX-III. CTX-III, purified from the venom of the Taiwan cobra (Naja naja atra), as previously described (Kaneda et al., 1977), was shown to be homogeneous by high-performance liquid chromatography (Kaneda and Hayashi, 1983). Cell isolation, culture, and treatment. Blood was obtained from six healthy volunteers (four men and two women) aged 25–34 years (mean ⫾ SD, 30 ⫾ 3.16 years) who were free of allergies and major organ diseases. None of the volunteers were smokers, had a history of drug or alcohol abuse or occupational exposure to chemicals, or were taking medication that might interfere with the evaluation of immune cells; in addition, the female subjects were not menstruating, taking contraceptive agents, pregnant, or lactating. Human PBMCs and monocyte-enriched and purified lymphocyte preparations were obtained from blood as described previously (Su et al., 1997). Briefly, blood was collected by veinpuncture in a heparinized tube (10 U/ml) and immediately centrifuged at 150g for 20 min at room temperature. The buffy coat was pooled, layered over Ficoll–sodium diatrizoate (Ficoll-Paque Plus; Amersham Bioscience, Uppsala, Sweden), and centrifuged at 250g for 15 min at room temperature. The PBMC layer was collected, and the cells were washed three times with Hank’s balanced salt solution (HBSS; GibcoBRL, Grand Island, NY) without phenol red. The cells were then resuspended in RPMI 1640 medium supplemented with 10% heated-inactivated fetal bovine serum (FBS; GibcoBRL) and incubated at 37°C in 95% air/5% CO2 for 2 h, then nonadherent cells were removed and the adherent cells washed with HBSS.

More than 90% of the adherent cells were monocytes, as assessed by staining with Diff-Quik (Baxter Healthcare Corp., Miami, FL). The adherent cells were recovered using a cell scraper (Becton Dickinson, Lincoln, NJ). The nonadherent cells (lymphocytes) were cultured at 37°C in 95% air/5% CO2 in RPMI 1640 medium supplemented with 10% heat-inactivated FBS, as was the human leukemia cell line, K-562, used to test the effect of CTX-III on cell proliferation. After isolation, lymphocytes (1 ⫻ 106 cells/ml) or monocytes (2 ⫻ 105 cells/ml) were incubated for 30 min at 37°C in RPMI/10% FBS in the presence or absence of 2 ␮g/ml of phytohemagglutinin (PHA; Sigma, St. Louis, MO) in 96well plates (100 ␮l/per well) for cytotoxicity assays or in 24-well plates (1 ml per well) for cytokine production and apoptosis induction assays. Cytotoxicity assay. Cell proliferation and DNA synthesis were measured, respectively, using an XTT kit (Boehringer Mannheim, Germany) (Scudiero et al., 1988) or [3H]thymidine incorporation. In the XTT test, PBMCs (2 ⫻ 105 cells per well) were incubated with different concentrations of CTX-III for 72 h at 37°C, 50 ␮l of the XTT labeling mixture was added to each well (final XTT concentration 0.3 mg/ml) and incubation was continued for 2– 4 h at 37°C, and the absorbance at 450 nm was measured using an ELISA reader (EL312e, Bio-Tek instruments). To measure DNA synthesis, the cells were treated with CTX-III for 24, 48, or 72 h, then 0.5 ␮Ci of [3H]thymidine (specific activity 6.7 Ci/ mmol; Amersham Pharmacia Biotech, UK) was added and incubation was continued for 4 h before the cells were harvested and cell-bound radioactivity was counted in a ␤-counter. Cell cycle analysis and measurement of apoptosis by flow cytometry. The distribution of cells at different stages in the cell cycle was estimated by flow cytometric DNA analysis, as described previously (Telford et al., 1994). Briefly, 1 ⫻ 106 untreated or PHA-activated cells in 10-cm plastic dishes were treated with various concentrations of CTX-III for 72 h. Cell suspensions (lymphocytes or trypsin/EDTA-detached monocytes) were centrifuged at 900g at 4°C, collected, and washed twice with cold phosphate-buffered saline (PBS), pH 7.2, and fixed at 4°C with 80% ethanol/20% PBS. The fixed cells were treated for 30 min at 4°C in the dark with fluorochrome DNA staining solution (1 ml) containing 40 ␮g of propidium iodide and 0.1 mg of RNase A, and then the stained cells were analyzed using a FACS cytometer (Becton Dickinson, San Jose, CA). The percentage of cells in each cell cycle phase (G0/G1, S, or G2/M) was calculated using Lysis II Software with a minimum of 1.5 ⫻ 104 cells per sample being evaluated in each case. In addition, apoptosis was detected by terminal deoxynucleotidyl transferase-mediated deoxyuridine triphosphate nick-end labeling (TUNEL) staining performed as described in the Boehringer kit (Boehringer). Briefly, 1 ⫻ 106 cells

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Fig. 1. CTX-III inhibits the proliferation of human blood PBMCs. Triplicate samples of untreated or PHA-treated human blood PBMCs were incubated for 3 days with different concentrations of CTX-III. The cells were then either labeled with XTT and measured by spectrophotometer at 450 nm (A) or incubated with [3H]thymidine and the cell-bound radioactivity was counted on a ␤-counter (B). Data are means ⫾ SD.

were washed, fixed for 30 min at room temperature in 4% paraformaldehyde in PBS, washed again, permeabilized by incubation for 2 min at 4°C with 0.1% Triton X-100 and 0.1% sodium citrate, washed twice, and then incubated for 60 min at 37°C with 50 ␮l of TUNEL mixture (FITC-12dUTP, dATP, CoCl2, terminal deoxynucleotidyltransferase, and TdT buffer; Boehringer), and the sample was analyzed by flow cytometry. Cytokine production assay. Cytokine levels were measured as described previously (Huang et al., 2001). After treatment with CTX-III, culture medium from suspensions containing 2 ⫻ 106/ml of monocytes or lymphocytes was centrifuged at 900g for 10 min at 4°C and the supernatants were collected and stored at ⫺20°C for no more then 1 month before levels of tumor necrosis factor-␣ (TNF-␣), interleukin-2 (IL-2), interferon-␥ (IFN-␥), and IL-4 were measured using ELISA kits (R&D, Minneapolis, MN) according to the manufacturer’s instructions. Detection of cell surface molecule expression by flow cytometry. After CTX-III treatment for 72 h, cells were harvested for evaluation of the expression of CD4, CD8, and IL-2R ␤ chain (CD25) by flow cytometry (Yu et al., 2002). The harvested cells were washed twice with HBSS and their concentration was adjusted to 2 ⫻ 106 cells/ml. They were then incubated for 1 h at 4°C with mouse monoclonal antibodies against human CD4, CD8, or CD25 (BD PharmMingen, San Diego, CA), washed twice with cold HBSS, and then incubated for 1 h at 4°C with FITC-labeled anti-mouse IgG antibody (BD PharMigen). After two washes with cold HBSS, the stained cells were analyzed on a FACScan flow cytometer (Becton Dickinson) and the

expression of surface molecules was estimated using WinMDI Version 2.8 Software. Binding of CTX-III to PBMCs. Binding of CTX-III to PHAstimulated PBMCs was determined by an immunofluorescent assay (Bueno et al., 2002). The PHA-stimulated PBMC suspension (1 ⫻ 106 cells/ml) was incubated for 30 min at 4°C with CTX-III (1 ␮g/ml) and then washed three times with HBSS. Mouse monoclonal anti-CTX-III antibody, produced in our laboratory (Chang et al., 1993), was added and incubation was continued for 40 min at 4°C. The cells were then washed with HBSS, incubated for 40 min at 4°C in the dark with Alexa Fluor 488-conjugated goat anti-mouse IgG (Molecular Probes, Eugene, OR) and the nuclear stain Hoechst 33342 (Molecular Probes), washed, smeared on a slide, and viewed using a fluorescent microscope. Bound CTX-III was seen as green fluorescence, while nuclear labeling was seen as blue fluorescence. Affinity isolation of PBMC surface proteins binding to CTX-III. PBMC surface proteins that bind to CTX-III were isolated using a CTX-III Sepharose 4B affinity column (Yang et al., 1977). CNBr–Sepharose (5 g) (Pharmacia Fine Chemicals, Uppsala, Sweden) was swollen in 1 mM HCl and then activated by coupling buffer (0.1 M NaHCO3, 0.5 M NaCl, pH 8.3). The beads were incubated for 3 h at room temperature with 15 mg of CTX-III in 1 ml of coupling buffer and then for 2 h at room temperature with 2.5 M glycine in 0.1 M Tris–HCl buffer, pH 8.0, to block remaining free binding sites. The CTX-III Sepharose 4B was washed with 0.1 M sodium acetate buffer containing 0.5 M NaCl, pH 4.0 and then with 0.1 M Tris–HCl buffer con-

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three times in HBSS, suspended in 3 ml of buffer A (20 mM Tris–HCl, 250 mM sucrose, 2 mM EDTA, 10 mM EGTA, 10 mM benzamidine, 1 mg/ml of leupeptin, and 2 mM phenylmethyl sulfonylfluoride), sonicated for 6 ⫻ 15 s in ice, and then centrifuged at 100,000g for 60 min at 4°C. The supernatant was discarded and the pellet incubated for 1 h at room temperature in buffer A containing 1% Nonidet P-40 (Chen, 1993). The membrane extract was then applied to the CTX-III-Sepharose 4B column and left in contact with the column for 1 h at 4°C. The column was washed at a flow rate of 20 ml/h with PBS, pH 7.2, until the absorbance at 280 nm returned to baseline, and then with 1 M Tris–HCl containing 0.15 M NaCl, pH 4.0, to elute bound protein, which was concentrated on an RCF-Conflict centrifugal concentrator (Bio-Molecular Dynamics, Beaverton, OR), and a sample was subjected to 12% SDS–PAGE on a Novex Xcell (San Diego, CA). Following denaturation in sample buffer (0.125 M Tris–HCl, 4% SDS, 20% glycerol, and 10% ␤-mercaptoethanol, pH 6.8), the samples were resolved on a 12% SDS gel with Tris– glycine running buffer, pH 8.3, and the gels stained with 0.1% Coomassie brilliant blue R-250. Fig. 2. Cytotoxicity of CTX-III for lymphocytes, monocytes, and K-562 cells. Triplicate samples of untreated or PHA-treated lymphocytes or untreated K-562 cells were incubated for 3 days with different concentrations of CTX-III. After treatment, cells number were counted by XTT labeling and expressed as a percentage of the control value. Data are means ⫾ SD (n ⫽ 3).

taining 0.5 M NaCl, pH 8.0, packed into a 1.0 ⫻ 10 cm column, and equilibrated with PBS, pH 7.2. PHA-stimulated PMBCs (1 ⫻ 108 cells) were washed

Statistical analysis. The results are expressed as the mean ⫾ SD. Differences in the cell cycle distribution were analyzed using the ␹2 test, while other differences between the control group and each treated group were analyzed using Student’s t test for unpaired data. Statistical analyses were performed using SAS (version 6.011; SAS Institute Inc, Cary, NC). A p value of ⬍0.05 was considered statistically significant.

Fig. 3. CTX-III induces time-dependent apoptosis of human lymphocytes. Apoptosis was evaluated by TUNEL staining followed by flow cytometry. (A) Untreated or PHA-treated human lymphocytes were incubated for 48 h with different concentrations of CTX-III, and then the percentage of apoptotic cells was measured. (B) Untreated or PHA-treated human lymphocytes were incubated with CTX-III (5 ␮g/ml) for different time intervals and the percentage of apoptotic cells was measured. Data are means ⫾ SD (n ⫽ 3).

S.-H. Su et al. / Toxicology and Applied Pharmacology 193 (2003) 97–105 Table 1 Percentage distribution of lymphocytes in different phases of the cell cycle following CTX-III treatment CTX-III concentration (␮g/ml) ⫺PHA 0 1 10 ⫹PHA 0 1 10

Phase

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Table 3 Effect of CTX-III treatment on interferon-␥ release by lymphocytes p

G1

S

G2/M

92.2 ⫾ 2.5 91.3 ⫾ 1.5 91.8 ⫾ 3.6

2.1 ⫾ 0.8 3.0 ⫾ 1.2 1.9 ⫾ 0.8

5.7 ⫾ 1.6 5.7 ⫾ 1.8 6.3 ⫾ 2.6

NS NS

89.6 ⫾ 2.4 88.6 ⫾ 2.5 87.9 ⫾ 3.5

4.5 ⫾ 2.8 3.7 ⫾ 2.8 4.6 ⫾ 0.8

5.9 ⫾ 2.6 7.7 ⫾ 2.8 7.6 ⫾ 3.2

NS NS

Note. Treatment was for 72 h. Data are means ⫾ SD (n ⫽ 3). p, distribution of cell cycle phase compared to the 0 toxin group by the ␹2 test. NS, nonsignificant.

Results Cytotoxicity of CTX-III for human PBMCs and the K-562 cell line When CTX-III was added to untreated or PHA-activated human PBMC, it caused dose-dependent inhibition of the proliferation of both untreated and PHA-activated cells (Fig. 1A) and of DNA synthesis in the PHA-activated cells (Fig. 1B); the effect on proliferation was more marked with PHA-activated cells. When the effects of CTX-III on separate lymphocyte and monocyte populations were investigated, it was found to be cytotoxic for PHA-treated, but not untreated, lymphocytes with a median effective dose (ED50) on PHA-treated cells of approximately 10 ␮g/ml (Fig. 2) but was not cytotoxic for either untreated or PHAtreated monocytes (data not shown), showing that its cytotoxic activity on human PBMCs was lymphocyte selective. Parallel experiments using the human leukemia cell line K-562 showed that it was even more effective against transformed lymphocytes (Fig. 2).

CTX-III concentration (␮g/ml)

⫺PHA

⫹PHA

IFN-␥ (pg/ml) 0 1 10

280 ⫾ 42 279 ⫾ 32 36 ⫾ 32***

476 ⫾ 89 290 ⫾ 53** 30 ⫾ 25***

Note. Treatment was for 72 h. Data are means ⫾ SD (n ⫽ 5). ** and *** p ⬍ 0.005 and 0.001, respectively, compared to the control (0 ␮g/ml) group using the unpaired Student’s t test.

3A) but not in monocytes (data not shown); again, the effect was more marked on PHA-activated lymphocytes. Induction of apoptosis in lymphocytes, either with or without PHA activation, was time dependent (Fig. 3B). Despite apoptosis being induced, the cell cycle distribution was not affected in either untreated or PHA-treated lymphocytes (Table 1). Effects of CTX-III on cytokine production To determine whether CTX-III-induced cytotoxicity was associated with altered cytokine expression, untreated or PHA-treated monocytes or lymphocytes were incubated with CTX-III for various periods and an incubation time was chosen for each cytokine (TNF-␣, IFN-␥, IL-2, and IL-4) to give maximal basal cytokine release. TNF-␣ release by both untreated and PHA-activated monocytes was significantly increased following 24 h exposure to 10 ␮g/ml of CTX-III (Table 2). In contrast, incubation of either untreated or PHA-treated lymphocytes with 10 ␮g/ml of CTXIII for 72 h resulted in reduced IFN-␥ release (Table 3), while incubation with the same concentration of toxin for 24 h resulted in a slight decrease in IL-2 release and no effect on IL-4 release (Table 4). Effect of CTX-III on lymphocyte cell surface molecule expression Since surface molecules, including CD4, CD8, and the IL-2R ␤ chain (CD25), play important roles in T lympho-

CTX-III induces apoptosis of lymphocytes After 48 h treatment with CTX-III, apoptosis was induced in both untreated and PHA-treated lymphocytes (Fig. Table 2 Effect of CTX-III treatment on tumor necrosis factor-␣ release by monocytes CTX-III concentration (␮g/ml)

⫺PHA

⫹PHA

TNF-␣ (pg/ml) 0 1 10

306 ⫾ 25 320 ⫾ 23 400 ⫾ 32***

410 ⫾ 36 420 ⫾ 14 486 ⫾ 23**

Note. Treatment was for 24 h. Data are means ⫾ SD (n ⫽ 5). ** and *** p ⬍ 0.005 and 0.001, respectively, compared to the control (0 ␮g/ml) group using the unpaired Student’s t test.

Table 4 Effects of CTX-III treatment on interleukin-2 and interleukin-4 release by lymphocytes CTX-III concentration (␮g/ml) IL-2 (pg/ml) 0 1 10 IL-4 (pg/ml) 0 1 10

⫺PHA

⫹PHA

42.2 ⫾ 9.2 38.6 ⫾ 11.3 22.7 ⫾ 8.4*

40.5 ⫾ 8.6 42.7 ⫾ 9.9 30.8 ⫾ 7.9*

4.5 ⫾ 0.6 4.3 ⫾ 0.5 3.9 ⫾ 0.8

10.8 ⫾ 1.3 10.6 ⫾ 0.6 11.5 ⫾ 1.0

Note. Treatment was for 24 h. Data are means ⫾ SD (n ⫽ 5). * p ⬍ 0.01 compared to the control (0 ␮g/ml) group using the unpaired Student’s t test.

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Table 5 Effects of CTX-III treatment on lymphocyte expression of surface molecules CTX-III concentration (␮g/ml) CD4 (intensity) 0 1 10 CD8(intensity) 0 1 10 CD25(intensity) 0 1 10

⫺PHA

⫹PHA

169.2 ⫾ 28.3 169.8 ⫾ 33.6 172.9 ⫾ 41.2

140.3 ⫾ 21.5 138.5 ⫾ 32.6 193.6 ⫾ 36.5*

176.3 ⫾ 30.6 179.6 ⫾ 41.5 217.6 ⫾ 45.9

329.7 ⫾ 35.1 296.9 ⫾ 45.3 230.2 ⫾ 29.6**

5.1 ⫾ 2.3 5.7 ⫾ 2.7 5.6 ⫾ 1.3

94.8 ⫾ 6.7 58.2 ⫾ 5.2*** 47.2 ⫾ 6.3***

Note. Treatment was for 72 h. Data are means ⫾ SD (n ⫽ 5). * , **, *** p ⬍ 0.05, ⬍0.001, and ⬍0.0001, respectively, compared to the control (0 ␮g/ml) group using the unpaired Student’s t test.

cyte function after activation, we examined the effect of CTX-III on the expression of these surface molecules by lymphocytes. Flow cytometric analysis showed that CTXIII significantly increased CD4 expression and decreased CD8 and CD25 expression in PHA-activated lymphocytes, but not in untreated lymphocytes (Table 5). As shown in Fig. 4, treatment with 0.1 or 10 ␮g/ml of CTX-III led to a marked reduction in the percentage of CD8⫹ lymphocytes, and the CD4⫺CD8⫹/CD4⫹CD8⫺ ratio showed a dramatic decrease from 0.512 in controls to 0.023 after treatment with 10 ␮g/ml of CTX-III. These results showed that CTXIII had a selective toxic effect on CD8⫹ cells. Binding of cell surface proteins to CTX-III To determine whether CTX-III interacted with cell surface proteins on PBMCs, CTX-III binding was measured in an immunofluorescent assay. As shown in Fig. 5A, CTX-III was seen as a diffuse staining of the membrane and as aggregates within the cell, indicating that it bound to and entered the cells. When CTX-III binding components were isolated from a PBMC membrane extract by affinity chromatography on CTX-III–Sepharose 4B beads, three proteins

Fig. 5. CTX-III binds to PHA-treated PBMC cell surface proteins. (A) Binding of CTX-III to the cell membrane of PHA-treated human PBMCs incubated with CTX-III (1 ␮g/ml) for 30 min at 4°C. Bound CTX-III was detected using anti-CTX-III monoclonal antibody plus Alexa Fluor 488conjugated goat anti-mouse IgG (Molecular Probes) and Hoechst 33342 (Molecular Probes), binding being determined by immunofluorescence microscopy. The green fluorescence shows bound CTX-III and the blue fluorescence shows the nucleus. Bar represent 5 ␮m. (B) Isolation and characterization of binding proteins. A PBMC membrane extract was applied to a CTX-III affinity column and the eluted proteins were subjected to electrophoresis on a 12% SDS–polyacrylamide gel and detected by Coomassie blue staining.

were eluted with apparent molecular weights on SDS– PAGE of 92, 77, and 68 kDa (Fig. 5B), suggesting that CTX-III binds either to more than one site on the cell membrane or to a trimolecular complex.

Discussion In this study, we found that CTX-III showed cell typeselective cytotoxicity for human PBMCs, being cytotoxic for, and inducing apoptosis in, lymphocytes, but not monocytes. Interestingly, CD8⫹ T lymphocytes were preferentially induced to undergo apoptosis by CTX-III treatment.

Fig. 4. The CD4⫹ and CD8⫹ cell distribution in PHA-activated PMBCs treated with CTX-III. Cells were treated for 72 h with 0, 0.1, or 10 ␮g/ml of CTX-III and the percentages of CD4⫹ and CD8⫹ cells were measured by flow cytometry. The dot plots are separated into four sections representing the CD4⫹CD8⫺, CD4⫹CD8⫹, CD4⫺CD8⫹, and CD4⫺CD8⫺ populations; the inset value for each section is the percentage of the corresponding cell population.

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Since CD8⫹ T lymphocytes are important contributors to IFN-␥ production, IFN-␥ levels were reduced following CTX-III treatment. Moreover, in lymphocytes, CTX-III decreased IL-2 production, increased CD4 expression, and reduced IL-2R ␤ chain (CD25) expression, whereas, in monocytes, it enhanced TNF-␣ production. In addition, CTX-III was shown to bind to the PBMC membrane, and three CTX-III-binding proteins with apparent molecular weights of 92, 77, and 68 kDa were isolated. Accordingly, we propose that CTX-III interacts with cell surface proteins and that this binding initiates a signaling pathway and causes functional changes that contribute to its cytotoxicity. Himnan et al. (1987) reported that macrophages and B lymphocytes are approximately 1000 times less susceptible than T lymphocytes to Naja naja siamensis cytotoxin D. The present study showed that CD8⫹ T lymphocytes were more sensitive than CD4⫹ lymphocytes to CTX-III. Comparison of its cytotoxicity for normal and transformed lymphocytes showed that the ED50 for the leukemia cell line K-562 was about 3 ␮g/ml, lower than that of 10 ␮g/ml seen with PHAactivated lymphocytes, indicating that transformed cells are more susceptible to this toxin. Previous studies have shown that the fluidity of membrane lipids plays an important role in the cytolytic action of CTX-III and that lipid fluidity is altered in the membranes of transformed cells, which may explain the modified susceptibility to the toxin (Chiou et al., 1993; Kaneda et al., 1985). Since CTX-III was more effective on lymphocyte-derived transformed cells, these results suggest that CTXIII may have potential clinical use in the treatment of abnormal lymphocytes. The immune response is mediated by cytokines, such as tumor necrosis factor-␣, released by macrophages/monocytes, and by interleukins (IL-2 and IL-4) and interferon-␥, released by T lymphocytes. After exposure to an effective dose of CTX-III, TNF-␣ release by monocytes was increased, while IFN-␥ and IL-2 release by lymphocytes was inhibited. In addition, following CTX-III treatment of lymphocytes, CD4 expression was increased, while CD8 and CD25 expression was reduced. In addition, the percentage of CD8⫹ T cells was markedly reduced by treatment with 0.1 or 10 ␮g/ml of CTX-III and the CD8⫹/CD4⫹ ratio was reduced from 0.512 to 0.023 after treatment with 10 ␮g/ml of CTX-III. This marked depletion of CD8⫹ cells probably explains the reduction in IFN-␥ levels, since these cells are important contributors to IFN-␥ production. The observed decrease in CD8⫹ expression is probably due to the high proportion of these cells being preferentially induced to undergo apoptosis. The reason why CTX-III selectively causes apoptotic death in CD8⫹ cells is not known but may be due to the preferential binding of CTX-III to CD8⫹ T cells or/and to some intrinsic properties of this cell subset, such as the CTX/lipid interactions in the membrane reported by Dubovskii et al. (2001). In this regard, it is interesting to note that a similar higher frequency of apoptotic death of CD8⫹ cells is seen in PBMCs from HIV-1 and EBVinfected patients (Meyaard et al., 1992; Akbar et al., 1993).

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Moreover, CD8⫹ T cells were recently shown to be more susceptible than CD4⫹ T cells to enterotoxin B-induced apoptosis (Salmond et al., 2002). Loss of CD28 expression on CD8⫹ T cells results in enhanced activation-induced apoptosis of these cells (Borthwick et al., 2000). The selectivity of CTX-III in triggering apoptosis of CD8⫹ T cells suggests potent clinical applications of CTX-III, such as the treatment of leukemias of CD8⫹ origin or preventing graft rejection following transplantation, since the CD8⫹ T cell population plays an important part in the immunopathology of this process. CTX-III caused dose- and time-dependent induction of apoptosis of lymphocytes, but not of monocytes. Since TNF-␣ is regarded as an apoptosis-preventing factor for monocytes (Flad et al., 1999), the lower sensitivity of monocytes to CTX-III-induced apoptotic death may be explained by the high TNF-␣ induction. We previously reported that the functional site on CTX-III responsible for cytotoxicity is different from that responsible for hemolytic activity (Chang et al., 1993), i.e., the binding of CTX-III to different target sites may result in different responses. Moreover, immunofluorescent staining demonstrated that CTX-III showed capping on monocyte membranes, but bound uniformly, without capping, to lymphocyte membranes (our unpublished data). In addition, CTX-III–Sepharose affinity chromatography, followed by SDS–PAGE analysis, showed that three PBMC membrane proteins with apparent molecular weights of 92, 77, and 68 kDa bound to CTX-III. On the basis of our results, we propose that CTX-III binds to receptor(s) on the cell membrane and sets in progress an intracellular cascade of enzymatic and genetic events, leading to its capping and endocytosis. It has been suggested that CTXs bind to membrane proteins, lipid receptors, Ca2⫹ binding sites, or other sites, and, as a consequence of this interaction, affect the cell membrane or intracellular processes that will eventually lead to cellular destruction. CTX-III is highly hydrophobic and acts as a protein kinase C inhibitor (Chiou et al., 1993), these two properties being shared with other naturally occurring bioactive amphiphilic polypeptides, such as mastoparan (Raynor et al., 1991) and melittin (Katoh et al., 1982). The diverse mechanisms involved in the effect of CTX-III on human immune cells deserve further clarification. One possibility would be to measure transforming growth factor-␤ production and/or to determine whether CTX-III binds to or interacts with the transforming growth factor-␤ receptor (De Crescenzo et al., 2003), which is a multimeric receptor similar, although not identical, to the CTX-III binding proteins.

Acknowledgments This study was supported by the National Science Council, Executive Yuan, Taiwan (Grant NSC 83-0412-B037055).

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References Akbar, A.N., Borthwick, N., Salmon, M., Gombert, W., Bofill, M., Hamsadeen, N., Pilling, D., Pett, S., Grundy, J.E., Janossy, G., 1993. The significance of low bcl-2 expression by CD45RO T cells in normal individuals and patients with acute viral infections: the role of apoptosis in T cell memory. J. Exp. Med. 178, 427– 438. Borthwick, N.J., Lowdell, M., Salmon, M., Akbar, A.N., 2000. Loss of CD 28 expression on CD8⫹ T cell is induced by IL-2 receptor ␥ chain signaling cytokines and type I IFN, and increases susceptibility to activation-induced apoptosis. Int. Immunol. 12, 1005–1013. Bougis, P., Rochat, H., Pieroni, G., Verger, R., 1982. A possible orientation change of cardiotoxin molecule during its interaction with phospholipid monolayer. Toxicon 20, 187–190. Bueno, C., Rodriguez-Caballero, A., Garcia-Montero, A., Pandiella, A., Almeida, J., Orfao, A., 2002. A new method for detecting TNF-alphasecreting cells using direct-immunofluorescence surface membrane staining. J. Immunol. Methods 264, 77– 87. Chang, K.L., Chen, Y.S., Lin, S.R., Chang, L.S., Chang, C.C., 1993. Probing the functional sites in Naja naja atra (Taiwan cobra) cardiotoxin III with monoclonal antibody. Biochem. Mol. Biol. Int. 29, 1015–1022. Chen, C.C., 1993. Protein kinase C ␣, ␦, ␧, and ␰ in C6 glioma cells: TPA induces translocation and down-regulation of conventional and new PKC isoforms but not atypical PKC ␰. FEBS Lett. 32, 169 –173. Chien, K.Y., Chiang, C.M., Hseu, Y.C., Vyas, A.A., Rule, G.S., Wu, W., 1994. Two distinct types of cardiotoxin as revealed by the structure and activity relationship of their interaction with zwitterionic phospholipid dispersions. J. Biol. Chem. 269, 14473–14483. Chiou, S.H., Raynor, R.L., Zheng, B., Chambers, T.C., Kuo, J.F., 1993. Cobra venom cardiotoxin (cytotoxin) isoforms and neurotoxin: comparative potency of protein kinase C inhibition and cancer cell cytotoxicity and models of enzyme inhibition. Biochemistry 32, 2062– 2067. De Crescenzo, G., Pham, P.L., Durocher, Y, O’Connor-McCourt, MD., 2003. Transforming growth factor-beta (TGF-beta) binding to the extracellular-domain of the type II TGF-beta receptor: receptor capture on a biosensor surface using a new coiled-coil capture system demonstrates that avidity contributes significantly to high affinity binding. J. Mol. Biol. 328, 1173–1183. DeTolla, L.J., Stump, K.C., Russell, R., Viskatis, L.J., Vidal, J.G., Newman, R.A., Etcheverry, M.A., 1995. Toxicity of the novel animalderived anticancer agent, VRCTC-310: acute and subchronic studies in beagle dogs. Toxicon 99, 31– 46. Dubovskii, P.V., Dementieva, D.V., Bocharov, E.V., Utkin, Y.N., Arseniev, A.S., 2001. Membrane binding motif of the p-type cardiotoxin. J. Mol. Biol. 305, 137–149. Flad, H.D., Grage-Griebenow, E., Petersen, F., Scheuerer, B., Brandt, E., Baran, J., Pryjma, J., Ernst, M., 1999. The role of cytokines in monocyte apoptosis. Pathobiology 67, 291–293. Fomsgaard, A, 1999. HIV-1 DNA vaccines. Immun. Lett. 65, 127–131. Hinman, C.L., Jiang, X.L., Tang, H.P., 1990. Selective cytolysis by a protein toxin as a consequence of direct interaction with the lymphocyte plasma membrane. Toxicol. Appl. Pharmacol. 104, 290 –300. Hinman, C.L., Lepisto, E., Stevens, R., Montgomery, I.N., Rauch, H.C., Hudson, R.A., 1987. Effects of cardiotoxin D from Naja naja siamensis snake venom upon murine splenic lymphocytes. Toxicon 25, 1011– 1014. Huang, L.W., Liu, H.W., Chang, K.L., 2001. Development of a sandwich ELISA test for arginase measurement based on monoclonal antibodies. Hybridoma 20, 53–57. Iwaguchi, T., Takechi, M., Hayashi, K., 1985. Cytolytic activity of cytotoxin isolated from Indian cobra venom against experimental tumor cells. Biochem. Int. 10, 343–349. Kaneda, N., Hamaguchi, M., Kojima, K., Kaneshima, H., Hayashi, K., 1985. Action of cobra venom cardiotoxin on chick embryonic fibro-

blasts transformed with a temperature-sensitive mutant of Rous sarcoma virus. FEBS Lett. 192, 313–316. Kaneda, N., Hayashi, K., 1983. Separation of cardiotoxins (cytotoxins) from the venoms of Naja naja atra by reversed-phase high-performance liquid chromatography. J. Chromatogr. 281, 389 –392. Kaneda, N., Sasaki, T., Hayashi, K., 1977. Primary structure of cardiotoxin analogues II and IV from the venom of Naja naja atra.. Biochem. Biophys. Acta 491, 53– 66. Katoh, N., Raynor, R.L., Wise, B.C., Schatzman, R.C., Turner, R.S., Helfman, D.M., Fain, J.N., Kuo, J.F., 1982. Inhibition by melittin of phospholipid-sensitive and calmodulin-sensitive Ca2⫹-dependent protein kinases. Biochem. J. 202, 217–224. Ksenzhek, O.S., Gedov, V.S., Omel’chenko, A.M., Semenov, S.N., Sotnichenko, A.I., Mirsoshnikov, A.I., 1978. Interaction of the cardiotoxin from the venom of the cobra Naja naja oxiana with phospholipid membrane model system. Mol. Biol. 12, 1057–1065. Kumar, T.K., Jayaraman, G., Lee, C.S., Arunkumar, A.I., Sivaraman, T., Samuel, D., Yu, C., 1997. Snake venom cardiotoxins: structure, dynamics, function and folding. J. Biomol. Struct. Dyn. 15, 431– 463. Kumar, T.K., Lee, C.S., Yu, C., 1996. A case study of cardiotoxin III from the Taiwan cobra (Naja naja atra). Solution structure and other physical properties. Adv. Exp. Med. Biol. 391, 115–129. Lee, C.S., Kumar, T.K., Lian, L.Y., Cheng, J.W., Yu, C., 1998. Main-chain dynamics of cardiotoxin II from Taiwan cobra (Naja naja atra) as studied by carbon-13 NMR at natural abundance: delineation of the role of functionally important residues. Biochemistry 37, 155–164. Lee, C.Y., 1979. Ceccarelli, B., Clementi, F. (Eds.), Advances in Cytopharmacology, Vol. 3. Raven Press, New York, pp. 1–16. Menez, A., Gatineau, E., Roumestand, C., Harvey, A.L., Mouawad, L., Gilquin, B., Toma, F., 1990. Do cardiotoxins possess a functional site? Structural and chemical modification studies reveal the functional site of the cardiotoxin from Naja nigricollis. Biochimie 72, 575–588. Meyaard, L., Otto, S.A., Jonker, R.R., Mijnster, M.J., Keet, R.P.M., Miedema, F., 1992. Programmed death of T cell in HIV-1 infection. Science 257, 217–219. Ownby, C.L., Fletcher, J.E., Colberg, T.R., 1993. Cardiotoxin 1 from cobra ((Naja naja atra) venom causes necrosis of skeletal muscle in vivo. Toxicon 31, 697–709. Patel, H.V., Vyas, A.A., Vyas, K.A., Liu, Y.S., Chiang, C.M., Chi, L.M., Wu, W., 1997. Heparin and heparan sulfate bind to snake cardiotoxin: sulfated oligosaccharides as a potential target for cardiotoxin action. J. Biol. Chem. 272, 1484 –1492. Rajnoch, C., Chachque, J.C., Berrebi, A., Bruneval, P., Benoit, M-O., Carpentier, A., 2001. Cellular therapy reverses myocardial dysfunction. J. Thorac. Cardiovas. Surg. 121, 871– 878. Raynor, R.L., Zheng, B., Kuo, J.F., 1991. Membrane interactions of amphiphilic polypeptides mastoparan, melittin, polymyxin B, and cardiotoxin: differential inhibition of protein kinase C, Ca2⫹/calmodulindependent protein kinase II and synaptosomal membrane Na,KATPase, and Na⫹ pump and differentiation of HL60 cells. J. Biol. Chem. 266, 2753–2758. Salmond, R.J., Pitman, R.S., Jimi, E., Sorina, M., Hirst, T.R., Ghosh, S., Rincon, M., Williams, N.A., 2002. CD8⫹ T cell apoptosis induced by Escherichia coli heat-labile enterotoxin B subunit occurs via a novel pathway involving NK-␬ B-dependent caspase activation. Eur. J. Immunol. 32, 1737–1747. Scudiero, D.A., Shoemaker, R.H., Paull, K.D., Monk, S.A., Tierney, S., Nofziger, T.H., Currens, M.J., Seniff, D., Boyd, M.R., 1988. Evaluation of a soluble tetrazolium/formazan assay for cell growth and drug sensitivity in culture using human and other tumor cell lines. Cancer Res. 48, 4827– 4833. Sivaraman, T., Kumar, T.K.S., Chang, D.K., Yu, C., 1998. The role of acetic acid in the prevention of salt-induced aggregation of snake venom cardiotoxins. Biochem. Mol. Biol. Int. 44, 29 –39.

S.-H. Su et al. / Toxicology and Applied Pharmacology 193 (2003) 97–105 Stevens-Truss, R., Hinman, C.L., 1996a. Chemical modification of methionines in a cobra venom cytotoxin differentiates between lytic and binding domains. Toxicol. Appl. Pharmacol. 139, 234 –242. Stevens-Truss, R., Hinman, C.L., 1997. Activities of cobra venom cytotoxins toward heart and leukemic T-cells depend on localized amino acid differences. Toxicon 35, 659 – 669. Stevens-Truss, R., Messer, W.S., Hinman, C.L., 1996b. Heart and Tlymphocyte cell surfaces both exhibit positive cooperativity in binding a membrane-lytic toxin. J. Membr. Biol. 150, 113–122. Su, S.J., Chang, K.L., Lin, T.M., Huang, Y.H., Yeh, T.M., 1997. Uromodulin and Tamm-Horsfall protein induce human monocytes to secrete TNF and express tissue factor. J. Immunol. 158, 3449 –3456. Sue, S.C., Rajan, P.K., Chen, T.S., Hsieh, C.H., Wu, W., 1997. Action of Taiwan cobra cardiotoxin on membranes: binding modes of a ␤-sheet

105

polypeptide with phosphatidylcholine bilayers. Biochemistry 36, 9826 –9836. Telford, W.G., King, L.E., Fraker, P.J., 1994. Rapid quantitation of apoptosis in pure and heterogeneous cell populations using flow cytometry. J. Immunol. Methods 172, 1–16. Xiao, L.J., Hinman, C.L., 1990. Ablation of natural killer cell function by soluble cardiotoxin. Int. J. Immunopharmacol. 12, 247–254. Yang, C.C., Lin, M.F., Chang, C.C., 1977. Purification of anti-cobratoxin antibody by affinity chromatography. Toxicon 15, 51–56. Yu, H.S., Liao, W.T., Chang, K.L., Yu, C.L., Chen, G.S., 2002. Arsenic induces tumor necrosis factor alpha release and tumor necrosis factor receptor 1 signaling in T helper cell apoptosis. J. Invest. Dermatol. 119, 812– 819.