Introduction

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1,4-Benzothiazine (1,4-B) derivatives have been shown to exert a number of effects in various in vivo and in vitro experimental systems. In particular, a series of 1,4-B analogs show good antifungal and immunostimulating activity (Fringuelli et al., 1998; Pitzurra et al., 1999; Schiaffella et al., 1999). 1,4-Bs are also active on the cardiovascular system, and vasorelaxant, antiarrhythmic, and antihypertensive effects have been reported in part associated with Na⁺ and Ca²⁺-blocking activity (Kajino et al., 1991; Hirasawa et al., 1992; Campiani et al., 1995; Matsumoto et al., 2000; Okuyama et al., 2000). Moreover, 1,4-B-induced neurotoxic or neuroprotective effects have been described, and a possible role in neurodegenerative diseases, such as Parkinson's and Alzheimer's disease, has been hypothesized (Shen and Dryhurst, 1996a,b; Li and Dryhurst, 1997; Kobayashi et al., 1997; Shen et al., 1997; Okuyama et al., 2000). In particular, whereas the neuroprotective effect has been attributed in part to the Ca²⁺-blocking activity (Campiani et al., 1995; Okuyama et al., 2000), the neurotoxic effect has been also associated with the interference with mitochondrial metabolism (Li et al., 1998a). Finally, in vivo antitumor efficacy of 1,4-B has been described and attributed to a direct cytotoxic activity against neoplastic cells (Coughlin et al., 1995; Hasegawa et al., 1997; Inoue et al., 1998). All those different effects may be relevant in terms of further development of 1,4-B pharmacology aimed at suggesting possible therapeutic applications. In particular, the cytotoxic activity reported in different experimental systems requires further characterization.

In the present study, we performed experiments to evaluate cell death induction by a number of 1,4-B analogs using mouse thymocytes, a cell population well sensitive to apoptosis induction that has been widely used to study various apoptotic agents (McConkey et al., 1990; Suzuki et al., 1991; Fehsel et al., 1995; Hartel et al., 1998; Cifone et al., 1999; Murshedul et al., 2000; Sharetskii et al., 2000). Results of our experiments indicate that some but not all 1,4-B analogs tested induce thymocyte apoptosis. In particular, preliminary structure-activity relationship (SAR) analysis, based on the evaluation of apoptotic activity of a number of 1,4-B analogs, indicate that the sulfur (S) oxidation state, the presence of the carbonyl group, and the nature and position of the side chain modulate the apoptotic activity.

Moreover, results indicate that 1,4-B-induced apoptosis requires a sequence of events including a rapid activation of phosphatidylcholine-specific phospholipase C (PC-PLC) and acidic sphingomyelinase (aSMase), loss of mitochondrial membrane potential (m), cytochrome c release, and caspase activation. These observations indicate that some of 1,4-B analogs may activate apoptosis through a complex signaling pathway that requires the activation of different biochemical events.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

Cell System and Treatments. Thymocytes from 4- to 6-week-old C3H/HeN mice were enriched by cell passage through nylon wool columns. The effect of several agents on 1,4-B-induced apoptosis, ceramide generation, and phospholipase activity was evaluated. The following agents were used: U73122 (Calbiochem, San Diego, CA), a PI-PLC inhibitor; D609 (Kamiya Biomedical, Thousand Oaks, CA), a PC-PLC inhibitor; monensin (Sigma-Aldrich, St. Louis, MO) and bafilomycin A1 (Calbiochem) to inhibit aSMase activity; and Z-DEVD-FMK (Calbiochem), a caspase-3 inhibitor (Cifone et al., 1999). In general, cells were incubated with inhibitors for 30 min prior to the addition of 1,4-B at the concentrations indicated in the figure legends. In all of the experiments dexamethasone (DEX; 10⁷ M) was used as a positive control (Cifone et al., 1999).

Apoptosis Evaluation by Propidium Iodide Solution. Apoptosis was measured by flow cytometry as described elsewhere (Cifone et al., 1999). After culturing, cells were centrifuged, and the pellets were gently resuspended in 1.5 ml of hypotonic propidium iodide (PI) solution (50 µg/ml in 0.1% sodium citrate plus 0.1% Triton X-100; Sigma). Tubes were kept at 4°C in the dark overnight. The PI fluorescence of individual nuclei was measured by flow cytometry with standard FACScan equipment (BD Biosciences, San Jose, CA). The nuclei traversed the light beam of a 488-nm argon laser. A 560-nm dichroid mirror (DM 570) and 600-nm band pass filter (bandwidth 35 nm) were used to collect the red fluorescence due to PI DNA staining, and data were recorded in logarithmic scale in a Hewlett Packard (HP 9000, model 310) computer. The percentage of apoptotic cell nuclei (sub-diploid DNA peak in the DNA fluorescence histogram) was calculated with specific FACScan research software (Lysis II).

Apoptosis Evaluation by Acridine Orange. To identify apoptotic nuclei, thymocytes (10⁶ cells/ml), untreated or treated with DEX or 6FS5, were labeled with acridine orange. Briefly, after treatment, cells were washed, gently resuspended in 100 µl of phosphate-buffered saline (PBS) with acridine orange (40 µg/ml), and left at room temperature for approximately 2 min. Finally, 10 µl of cell suspension were placed on a glass slide and analyzed with a fluorescence microscope.

Fluorimetric DNA-Fragmentation Assay and DNA Electrophoresis. After treatment, cells (2 × 10⁷) were washed and dissolved in a lysing buffer (5 mM Tris-HCl, 10 mM EDTA, and 0.5% Triton X-100) for 30 min on ice. After centrifugation for 15 min at 13,000g, RNase A (10 mg/ml, 2 h at 37°C) and proteinase K (10 mg/ml, 3 h at 50°C) were added to each sample, and DNA was extracted twice with phenol plus chloroform and recovered by centrifugation after overnight precipitation at 20°C in 2 volumes of ethanol in the presence of 0.3 M sodium acetate. Pellets were air dried, dissolved in 10 mM Tris-HCl, 1 mM EDTA, and 20 µg DNA, and loaded into the wells of a 1.5% agarose gel. Electrophoresis was carried out in TBE buffer (2 mM EDTA, 89 mM boric acid, and 89 mM Tris, pH 8.4), and DNA was visualized by ethidium bromide staining.

Cytofluorimetric Analysis of Loss of Mitochondrial Membrane Potential (m). Variations of the mitochondrial membrane potential (m) during thymocyte apoptosis was studied by using a cationic lipophilic fluorochrome, 5,5',6,6'-tetraethylbenzimidazolylcarbocyanine iodide (JC-1; Molecular Probes, Eugene, OR), which is able to selectively enter into mitochondria. JC-1 exists in a monomeric form emitting at 527 nm after excitation at 490 nm; however, depending on m, JC-1 is able to form J-aggregates that are associated with a large shift in emission (590 nm). Thus, the color of the dye changes reversibly from green to greenish orange as the mitochondria membrane becomes more polarized. The cell staining was performed as follows: cell suspensions were adjusted to a density of 10⁶ cells per ml and incubated in complete medium with JC-1 (10 µg/ml) for 10 min at room temperature in the dark. At the end of the incubation period, the cells were washed twice in cold PBS, resuspended in a total volume of 400 µl, and analyzed. Flow cytometry was performed by using standard FACScan equipment (BD Biosciences). For the analysis of cells stained with JC-1, the photomultiplier value of the detector in FL1 was set at 310 V and FL2 at 259. Data were acquired in list mode and analyzed with Lysis II software (BD Biosciences).

Determination of Cytochrome c Release. Thymus cells (3 × 10⁷) treated with 6FS5 (10 µg/ml) were washed in PBS and resuspended in 500 µl of ice-cold buffer containing 20 mM HEPES, 250 mM sucrose, 2 mM EDTA, 20 µg/ml phenylmethylsulfonyl fluoride (PMSF), 2 µg/ml leupeptin, and 10 µg/ml aprotinin, pH 7.1. Cells were disrupted on ice by using a Teflon homogenizer and centrifuged for 5 min at 3,000g to remove nuclei and unbroken cells. The supernatants were then centrifuged for 20 min at 12,000g to isolate mitochondrial fractions. For Western blot detection of cytochrome c, the supernatant from the last centrifugation and the mitochondrial fractions were subjected to 15% SDS-polyacrylamide gel electrophoresis. After protein transfer, the membrane was blocked with a 5% solution of bovine serum albumin in TBST buffer (25 mM Tris-HCl, 137 mM NaCl, and 0.05% Tween 20, pH 7.4) and incubated for 1 h at room temperature with monoclonal antibody against cytochrome c, diluted 1:1,000. The membrane was then incubated with goat anti-mouse monoclonal antibody (1:100,000) coupled to peroxidase. The specific protein complexes were identified using the SuperSignal substrate chemiluminescence reagent (Pierce, Rockford, IL).

Ceramide Mass Measurement [Diacylglycerol (DAG) Kinase Assay]. Aliquots of 5 × 10⁶ cells were suspended in 1 ml of RPMI 1640 medium supplemented with 10% fetal calf serum and 2 mM glutamine and antibiotics and treated for the indicated times with 1,4-B in the presence or absence of inhibitors. Treatment was stopped by immersion of samples in methanol/dry ice (70°C) for 10 s followed by centrifugation at 4°C in a microcentrifuge. To measure ceramide levels, pellets were dissolved in a buffer containing 20 mM Tris-HCl (pH 7.4), 1 µM PMSF, 10 µM leupeptin, 10 µM pepstatin, and 1 µM aprotinin. After incubation for 5 min at 4°C, the cells were sonicated (5 W, 80% output, 1 min and 50 s, and alternating 10-s sonication and 10-s pause) with a VibraCell sonicator (Sonics & Materials, Inc., Danbury, CT) and centrifuged for 30 min at 14,000 rpm at 4°C. The supernatants were then collected, and protein concentration was determined through the Pierce Micro BCA assay kit with bovine serum albumin standards. Lipids were extracted by the sequential addition of 400 µl of methanol, 500 µl of chloroform, and 200 µl of water. Samples were stirred for 2 min on a vortex mixer and centrifuged at 13,000 rpm for 10 min. The extraction and centrifugation steps were repeated twice. The organic phases, obtained from different extraction steps, were collected, washed once with 1 ml of solvent system containing chloroform/methanol/water (3:48:47 by volume), dried under nitrogen, and then incubated with Escherichia coli DAG kinase assay kit and [-³²P]ATP (specific activity, 3 Ci/mmol; Amersham Biosciences, Piscataway, NJ) as previously described (Cifone et al., 1999). Ceramide phosphate was then isolated by thin layer chromatography using chloroform/methanol/acetic acid (65:15:5, v/v) as solvent. Authentic ceramide from bovine brain (ceramide type III, nonhydroxy fatty acid ceramides; Sigma) was identified by autoradiography at rf = 0.25. Specific radioactivity of ceramide-1 phosphate was determined by scintillation counting of corresponding spots scraped off the gel. Quantitative results for ceramide production were obtained from comparing the experimental values with a linear curve of the ceramide standards and are expressed as picomoles of ceramide-1 phosphate per 10⁶ cells.

In Vitro SMase Analysis. Aliquots of 6 × 10⁶ cells/ml were treated for the indicated times with 1,4-B. Treatment was stopped by immersion of samples in methanol/dry ice (70°C) for 10 s, followed by centrifugation at 4°C in a microcentrifuge. To measure neutral SMase, pellets were dissolved in a buffer containing 20 mM HEPES (pH 7.4), 10 mM MgC1₂, 2 mM EDTA, 5 mM dithiothreitol, 0.1 mM Na₃VO₄, 0.1 mM Na₂MoO₄, 30 mM p-nitrophenylphosphate, 10 mM -glycerophosphate, 750 mM ATP, 1 µM PMSF, 10 µM leupeptin, 10 µM pepstatin, and 0.2% Triton X-100. After incubation for 5 min at 4°C, the cells were sonicated as described above and centrifuged for 30 min at 14,000 rpm at 4°C. The supernatants were then collected, and protein concentration was determined through the Pierce Micro BCA assay kit with bovine serum albumin standards. Proteins (50-100 µg) were incubated for 2 h at 37°C in a buffer containing 20 mM HEPES (pH 7.4), 1 mM MgCl₂, and 0.32 µl of N-methyl-¹⁴C-SM (0.04 µCi/ml; specific activity, 56.6 mCi/mmol; Amersham). To measure aSMase, after treatment the cells were washed, and the pellet was resuspended in 200 µl of 0.2% Triton X-100 and incubated for 15 min at 4°C. The cells were sonicated, and the protein concentration was assayed. Protein (50-100 µg) was incubated for 2 h at 37°C in a buffer (50 µl final volume) containing 250 mM sodium acetate, 1 mM EDTA (pH 5.0), and 0.32 µl of N-methyl-¹⁴C-SM (0.04 µCi/ml, specific activity, 56.6 mCi/mmol; Amersham). The reaction was stopped by the addition of 250 µl of chloroform/methanol (2:1 by volume). The lipids were extracted as described above. The organic phase, obtained in the different extraction steps, was collected and washed once with 1 ml of chloroform/methanol/water (3:48:47 by volume) to completely remove free radioactive phosphorylcholine. The aqueous phases were collected, transferred to scintillation vials, and routinely counted by liquid scintillation counting. The counts per minute represented the choline phosphate generated from SM hydrolysis. The organic phase was analyzed on TLC plates by using chloroform/methanol/ammonia hydroxide (7 N):water (85:15:0.5:0.5 by volume). The hydrolysis of SM was quantitated by autoradiography and liquid scintillation and expressed as picomoles of SM hydrolyzed per 10⁶ cells.

PI-PLC and PC-PLC Activity Assay. PI-PLC and PC-PLC activities were determined in vitro by their ability to hydrolyze [¹⁴C]PC or [¹⁴C]PI vesicles, respectively, to generate DAG. Cells were treated for the indicated times with 1,4-B in the presence or absence of the PC-PLC inhibitor, D609 (50 µg/ml), or the PI-PLC inhibitor, U73122 (2.5 µM). Treatment was stopped by immersion of samples in methanol/dry ice (70°C) for 10 s, followed by centrifugation at 4°C in a microcentrifuge. The pellets were then resuspended in 250 mM Tris-HCl buffer (pH 7.4) containing 10 µM PMSF, 100 µM bacitracin, 1 mM benzamidine, 1 µM aprotinin, 10 µM leupeptin, 10 µM pepstatin, and 5 µg/ml soybean trypsin inhibitor. Cells were lysed by sonication with a cell sonifier, and radiolabeled PC or PI vesicles were prepared by sonicating (5 min, 5 W, and 80% output) L-3-phosphatidyl [N-methyl-¹⁴C]choline-1,2 dipalmitoyl (specific activity, 56 mCi/mmol; Amersham) or L-3-phosphatidylinositol-1-stearoyl-2[¹⁴C]arachidonoyl for the detection of released DAG through PC-PLC or PI-PLC, respectively. Vesicles were resuspended at 10 µM in the reaction buffer [50 mM Tris-HCl (pH 7.4), 5 mM CaCl₂, 5 mM MgCl₂, and 0.01% fatty acid-free bovine serum albumin]. Whole-cell lysate (50-100 µg protein) was added to a 250-µl reaction buffer containing the vesicles, incubated at 37°C for 1 h, and the reaction buffer was stopped by the addition of 250 µl of chloroform/methanol/acetic acid (4:2:1 by volume). To separate the organic from the aqueous phase, 250 µl of H₂O, 250 µl of CHCl₃, and 100 µl of KCl were added, and the mixture was centrifuged at 4000 rpm in a microcentrifuge for 5 min. The organic phase was removed, dried under nitrogen, resuspended in 200 µl of chloroform, and applied to a silica gel TLC plate (Merck, Darmstadt, Germany) with an automatic applicator (Linomat IV, Camag, Muttenz, Switzerland). Samples were chromatographed in chloroform/methanol/acetic acid/water (100:60:16:8) to separate the parent phospholipids from the product of PC-PLC and PI-PLC, i.e., DAG. Authentic standards were chromatographed with the lipid extracts to locate the compounds of interest by exposure to iodine vapor. Radioactive spots, visualized by autoradiography and corresponding to standards, were scraped from the plate and counted by liquid scintillation. Radioactive measurements were converted to picomoles of product by using the specific activity of substrate. Blank values obtained from controls lacking cell proteins were subtracted from the experimental values. PC-PLC or PI-PLC activities were expressed as picomoles of DAG produced per 10⁶ cells.

Western Blotting for Evaluation of Activation of Caspase -3, -8, and -9. Cells were washed once with ice-cold PBS and lysed by incubating for 30 min on ice in 100 µl of lysis buffer (20 mM Tris-HCl, 0.15 M NaCl, 5 mM EDTA, 100 mM PMSF, 2.5 mM leupeptin, and 2.5 mM aprotinin). After centrifugation at 15,000 rpm for 15 min, extracted proteins were separated on a 12 or 15% SDS-polyacrylamide gel and electrophoretically transferred to a nitrocellulose transfer membrane (Schleicher & Schuell, Keene, NH). The membrane was blocked with TBST [20 mM Tris-HCl (pH 7.5), 150 mM NaCl, and 0.1% Tween 20] containing 5% skim milk for 1 h at room temperature, and each antibody was applied overnight at 4°C in the same blocking solution. Anti-caspase-3 antibody and anti-caspase-8 antibody were purchased from Santa Cruz Biochemicals (Santa Cruz, CA), and anti-caspase-9 antibody was purchased from New England Biolabs (Beverly, MA). After incubation, membranes were washed with TBST and incubated for 1 h with horseradish peroxidase-labeled goat anti-rabbit (for anti-caspase-8 and -9) or anti-mouse (for caspase-3) IgG (Pierce). The antigen-antibody complexes were revealed by enhanced chemiluminescence following the manufacturer's instructions (SuperSignal; Pierce).

Protease Activity Assay. For the caspase-3 activity assay, we used the colorimetric assay kit (CLONTECH Laboratories, Inc., Palo Alto, CA) based on spectrophotometric detection at 405 nm of the chromophore p-nitroanilide (pNA) after cleavage from the labeled specific substrates. The assay is based on the ability of the active enzyme to cleave the pNA from the enzyme substrate, DEVD-pNA. Briefly, the cells (6 × 10⁶ cells/ml), after treatment with 6FS5 (10 µg/ml) for 6 h in the presence or absence of inhibitor, were collected and resuspended in a lysis buffer (100 mM HEPES, 10 µM leupeptin, 10 µM aprotinin, 1 mM EDTA, 5 mM dithiothreitol, and 0.15% PMSF). Cellular lysates were then incubated in a microplate in the presence of conjugated protease substrates for 1 h at 37°C and then analyzed using a 96-well plate reader. Caspase-3 activity is expressed as nanomoles of pNA per 10⁶ cells.

Statistical Analysis. For data analysis, Student's t test was performed by the STATPAC Inc. (Minneapolis, MN) computerized program, and a p value <0.05 was used as the significance criterion.

	Results

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1,4-Benzothiazine (1,4-B) Analogs Induce Thymocyte Apoptosis: SARs. We performed experiments to evaluate the possible apoptotic effect of 1,4-B. In particular, in the attempt to analyze SAR we tested 16 1,4-B analogs looking at the possible role of sulfur (S), the carbonyl group, and the side chain. Figure 1, A and B, shows the structural differences of various 1,4-B analogs, whereas Fig. 1C shows the apoptosis effect as evaluated by the PI assay. Results shown in Fig. 1C were obtained at 18 h, when 1,4-B-induced apoptosis reaches the maximum level compared with untreated control (not shown).

View larger version (19K): [in this window] [in a new window]   Fig. 1.   1,4-Benzothiazine analogs structure and apoptosis activity. A, structures of 1,4-benzothiazine derivatives. B, structures of 1,4-benzothiazine synthetic intermediates. C, in vitro apoptotic activity. Apoptosis was tested as DNA fragmentation by PI assay, using thymocytes treated with different 1,4-B analogs (10 µg/ml) for 18 h. *, p < 0.05; **, p < 0.001 compared with untreated thymocytes.

1,4-Benzothiazine Analog, 6FS5, Induces Apoptosis and Thymus Cell Number Loss in Vivo. To better evaluate the characteristics of 1,4-B-induced cell death, we performed experiments to further characterize the molecular events associated with in vitro apoptosis induction and the possible in vivo effect on thymocytes. In the figures are shown results of representative experiments obtained with the compound 6FS5, one of the most active analogs (see Fig. 1).

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View larger version (32K): [in this window] [in a new window]   Fig. 2.   In vitro and in vivo thymocyte apoptosis. A, in vitro apoptotic activity of DEX (107 M) or 6FS5 (24 × 106 M) was evaluated by the DNA fragmentation PI assay. B, in vivo activity was measured by evaluating the thymus (six animals/group) cell numbers 24 h after i.p. injection. C, agarose gel electrophoresis of DNA extracted from mouse thymocytes after 18 h of incubation with DEX or 6FS5. D, fluorescence microscopy: orange acridine-stained thymocyte nuclei after 18 h of incubation in medium alone (a), medium plus DEX (b), or medium plus 6FS5 (c). , p < 0.001.

6FS5 Treatment Induces a Dose-Related Thymocyte Apoptosis and Ceramide Generation. 6FS5-induced apoptosis was dose-dependent (Fig. 3A) and paralleled the increase in the concentration of endogenous ceramide level (Fig. 3B). In particular, in Fig. 3B, results are reported showing thymocyte ceramide generation, obtained by treating lipid extracts with DAG kinase for quantitation of ceramide-1 phosphate amounts. As shown in Fig. 3, a basal level of ceramide was evident in untreated control and significantly increased after 6FS5 treatment (p < 0.001, 10, and 1 µg/ml, and p < 0.05 with 0.1 µg/ml compared with untreated control). Moreover, the dose able to induce maximal ceramide generation was 10 µg/ml and corresponded to the concentration able to induce maximal level of apoptosis (Fig. 3A). As for apoptosis, levels of ceramide declined almost to the basal level at the concentrations of 0.001 µg/ml, not being significantly different from the control values. Similar results were obtained with other 1,4-B analogs such as FS17 and FS5 (data not shown).

View larger version (20K): [in this window] [in a new window]   Fig. 3.   Dose-related induction of apoptosis parallels induction of ceramide generation. A, 6FS5-induced apoptosis. Mouse thymocytes incubated for 18 h with or without different doses of 6FS5 (10 to 0.001 µg/ml or 24 to 0.0024 × 106 M) were processed for DNA content analysis by PI staining. B, ceramide generation after 6FS5 treatment. Thymocytes were treated with 6FS5 at indicated doses for 5 min. Quantitative results for ceramide-1 phosphate levels are expressed as picomoles per 106 cells. C, kinetics of ceramide generation induced by 6FS5 treatment. Thymocytes were treated with 6FS5 (10 µg/ml) for the indicated times. Quantitative results for ceramide-1 phosphate levels are expressed as picomoles per 106 cells. , untreated control; , 6FS5-treated. Mean values ± S.E. of three different experiments in duplicate are reported. S.E. values were lower than 10% of mean values. , p < 0.05; , p < 0.001.

6FS5 Induces Activation of aSMase. We have previously shown that DEX-induced ceramide generation is due to activation of both aSMase, responsible for early ceramide generation (3-15 min), and neutral SMase, responsible for late ceramide generation (60 min) (Cifone et al., 1999). The contribution of aSMase to 1,4-B-induced ceramide generation in thymocytes was investigated by the specific in vitro aSMase analysis. Cellular extracts from mouse thymocytes treated for different times (1-30 min) with 6FS5 (10 µg/ml) or DEX (10⁷ M) were incubated with radiolabeled SM vesicles to detect aSMase. Results of a representative experiment presented in Fig. 4B show that aSMase activity was detected in extracts from untreated cells and significantly increased after 6FS5 or DEX treatment. In particular, the augmentation of aSMase activity was evident, statistically significant 1 to 15 min after treatment, and returned close to basal levels within 60 min (Fig. 4B).

View larger version (18K): [in this window] [in a new window]   Fig. 4.   Role of aSMase in 6FS5-induced thymocyte apoptosis. A, effect of monensin (10 µg/ml) or bafilomycin (2 µM) on DEX (107 M) or 6FS5 (10 µg/ml)-induced thymocyte apoptosis at 18 h. B, effect of DEX and 6FS5 on aSMase activity. Hydrolyzed SM was quantitated and expressed as picomoles per 106 cells. , untreated control; , DEX-treated; and , 6FS5-treated. Mean values ± S.E. of two different experiments in duplicate are reported. S.E. values were lower than 10% of mean values. C, effect of monensin on aSMase activity in control thymocytes and thymocytes treated for 5 min with DEX (107 M) or 6FS5 (10 µg/ml). , without monensin; , with monensin. Mean values ± S.E. of three different experiments in duplicate are reported. *, p < 0.05; **, p < 0.001.

6FS5-Induced Activation of aSMase and Apoptosis are Dependent on PC-PLC Activity. It has been previously reported that aSMase, associated with the lysosomal intracellular compartment or the caveolae, requires DAG for activation (Kolesnick, 1987; Riboni et al., 1997). The possible effect of 6FS5 on the activity of DAG-generating enzymes, such as PI-PLC or PC-PLC, was directly addressed by evaluating the enzymatic activities in vitro using appropriate radiolabeled substrates and TLC analysis of the reaction products. In Fig. 5, results are reported showing that 6FS5 induced DAG generation in mouse thymocytes (Fig. 5B). In particular, augmentation of PLC activity was evident and statistically significant at 1 min, peaked at 3 min, and then declined within 5 min after 6FS5 treatment. Furthermore, 6FS5-induced apoptosis was completely blocked by pretreatment of cells with D609, a selective PC-PLC inhibitor, but not with U9237, a selective PI-PLC inhibitor able to inhibit the PI-PLC-dependent, DEX-induced apoptosis (Fig. 5A) (Kolesnick, 1987). Moreover, the PC-PLC inhibitor D609 but not the PI-PLC inhibitor U73122 countered the 6FS5-induced SM hydrolysis (Fig. 5C) and the consequent ceramide generation (not shown), further suggesting that the 6FS5-induced aSMase activation is consequent to PC-PLC activity.

View larger version (22K): [in this window] [in a new window]   Fig. 5.   Role of PC-PLC activation in 6FS5-induced apoptosis. A, effect of U73122 (2.5 µM) or D609 (50 µg/ml) on DEX (107 M) or 6FS5 (10 µg/ml)-induced thymocyte apoptosis, as evaluated by 18 h after treatment. B, PLC activity involvement in DEX and 6FS5-induced thymocyte apoptosis. PLC activity is expressed as picomoles of DAG per 106 cells; , untreated control; , DEX-treated; and , 6FS5-treated. Mean values ± S.E. of two different experiments in duplicate are reported. S.E. values were lower than 10% of mean values. C, effect of U73122 () and D609 () on aSMase activity induced by DEX (107 M) or 6FS5 (10 µg/ml) treatment for 5 min. Mean values ± S.E. of three different experiments in duplicate are reported. , p < 0.05; , p < 0.001.

6FS5 induces Mitochondria Membrane Potential Transition (m) and Cytochrome C Release. It has been reported that m and consequent cytochrome c release are involved in apoptosis induction (Owens et al., 1991; Kroemer, 1995). We evaluated the possible effect of 1,4-B treatment on thymocyte m. Results in Fig. 6 indicate that 6FS5-induced m was evident at 1 h after treatment, peaked at 7 h (Fig. 6, see percentage (%), lower left panel), and was comparable to that induced by valinomycin, used as a positive control. Moreover, 6FS5 induced cytochrome c release in the cytoplasm that paralleled the loss of mitochondrial cytochrome c (Fig. 6B).

View larger version (23K): [in this window] [in a new window]   Fig. 6.   6FS5 induces loss of mitochondrial membrane transition (m) and cytochrome c release. A, thymocytes were treated with 6FS5 (10 µg/ml for 1, 3, 5, or 7 h) or with valinomycin (1 µM for 10 min) and then stained with JC-1 (10 µg/ml). FL1, green fluorescence intensity; FL2, red fluorescence intensity. UL, upper left quadrant; UR, upper right quadrant; LL, lower left quadrant; and LR, lower right quadrant. B, cytochrome c release from mitochondria: lane 1, time 0; lane 2, 2 h; lane 3, 3 h; and lane 4, 6 h after treatment. C, thymocytes were preincubated for 1 with different inhibitors, treated for 3 h with 6FS5 (10 µg/ml), and then stained with JC-1 (10 µg/ml). , untreated control; , 6FS5-treated. Mean values ± S.E. of three different experiments in duplicate are reported. **, p < 0.001.

6FS5 Induces Caspase -3, -8, and -9 Activation. It has been previously suggested that several interleukin-1-converting enzyme family cysteine proteases (caspases) play a prominent role in T-cell and thymocyte apoptosis (Martin et al., 1996; Thornberry et al., 1997). In particular, sequential pathways of caspase activation may exist, including those, activated by death receptors, of caspase-8 and caspase-3 and those, activated by nonreceptor signals, that include the cytochrome c-dependent caspase-9 activation and the consequent activation of caspase-3 (Krammer, 2000).

View larger version (48K): [in this window] [in a new window]   Fig. 7.   6FS5-induced caspase. Time-dependent 6FS5-induced activation of caspase-3, -8, and -9 as evaluated by Western blotting analysis. Numbers (from 0-6) indicate time (h) after treatment.

View larger version (22K): [in this window] [in a new window]   Fig. 8.   6FS5-induced apoptosis is dependent on PC-PLC and aSMase activation. A, effect of caspase-3 inhibitor on 6FS5-activated apoptosis. B, effect of D609 (50 µg/ml), monensin (10 µg/ml), bafilomycin (2 µM), and caspase-3 inhibitor (Z-DEVD-FMK; 75 µM) on caspase-3 activity. Thymocytes were preincubated for 30 min with the inhibitors and then treated with 6FS5 (10 µg/ml) for 6 h. Caspase activity is expressed in nanomoles of pNA per 106 cells. Mean values ± S.E. of three different experiments in duplicate are reported. , p < 0.001.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

The purpose of the present report was to investigate whether 1,4-B could induce thymocyte apoptosis and activate intracellular signals previously shown to be involved in cell death regulation. Results indicate that 1,4-B analogs induce apoptosis and activate a number of apoptotic signals.

Development and maintenance of healthy tissues involves apoptosis [or programmed cell death, (PCD)], a physiologically regulated cell death implicated in the control of normal cell turnover (Krammer, 2000). Dysregulated PCD contributes to many pathologies, including tumor promotion, neurodegenerative disorders, and autoimmune and immunodeficiency diseases (France-Lanord et al., 1997; Puranam et al., 1997; Taniwaki et al., 1999; Krammer, 2000; Yuan and Yankner, 2000). Therefore, the identification of agents and stimuli able to induce apoptosis, and the definition of activated signaling pathways, are of enormous interest.

A number of stimuli have been described to participate in the induction and inhibition of apoptosis so that the survival of a specific cell is under the control of a wide complex of signals. As an example, of the enzymatic activities, phospholipase and SMase activities have been shown to be important regulators of PCD (Kolesnick, 1987; Martin et al., 1996; Riboni et al., 1997; Thornberry et al., 1997; Cifone et al., 1999). Molecules, including ceramides, regulate apoptosis, and it has been shown that their effect is in part due to the modulation of mitochondrial activity, including cytochrome c release and the consequent caspase-9 activation (Kroesen et al., 2001). Moreover, the possible role of caspase activation has been described in a variety of experimental systems (Martin et al., 1996; Thornberry et al., 1997; Krammer, 2000).

To evaluate the possible effect of 1,4-B derivatives on apoptosis, we used mouse thymocytes, a cell population highly sensitive to PCD induction that has been widely used to study various apoptotic agents (McConkey et al., 1990; Suzuki et al., 1991; Fehsel et al., 1995; Hartel et al., 1998; Murshedul et al., 2000; Sharetskii et al., 2000). Results indicate that 1,4-B-treated thymocytes show the morphological characteristics of apoptotic cells as evaluated by microscopy analysis (Fig. 2). Furthermore, 1,4-B treatment induces DNA cleavage as indicated by the typical fragmentation pattern obtained by the agarose gel assay and the flow cytometry PI assay. Of interest, in vivo treatment with 1,4-B analogs induces significant thymocyte loss, an event that has been associated with other apoptosis inducers such as corticosteroids. Moreover, this apoptotic effect correlates with specific structural characteristics of 1,4-B, and preliminary SAR analysis, based on results obtained by testing 16 compounds, indicates that the S oxidation state, the carbonyl group, and the nature, position, and length of the side chain modulate the apoptotic efficacy. Based on these observations, future studies will be devoted to the design and synthesis of more active molecules and identification of the putative molecular targets responsible for 1,4-B binding and apoptosis activation.

We also performed experiments to evaluate the biochemical events associated with 1,4-B-activated apoptosis. Results indicate that thymocyte treatment with one of the most active compounds tested (6FS5, see Fig. 1) activates a number of signals previously described to correlate with apoptosis induction. In particular, drug treatment activates a sequence of biochemical events including PC-PLC (but not PI-PLC) activation, aSMase activation and ceramide generation, m and cytochrome c release, and caspase-3, -8, and -9 activation. Similar events have been previously described in different experimental models with apparent discrepancies as, for example, the temporal sequence of events. In particular, ceramide generation has been described as either upstream or downstream caspase activation and/or m and cytochrome c release, depending on the experimental system analyzed (Rodriguez-Lafrasse et al., 2001).

Our results clearly indicate that in the case of 1,4-B-induced thymocyte apoptosis, PC-PLC and aSMase activation precede caspase (Fig. 8) and mitochondrial activity (Fig. 6). Most of those molecular events, including aSMase activation and ceramide generation, have been previously associated with other cell death systems including DEX or Fas-induced apoptosis (Cifone et al., 1999; Kroesen et al., 2001). Of interest, aSMase activation and ceramide generation have been shown to be involved in neurodegenerative diseases, such as Parkinson's Disease, in which a role of 1,4-B has been proposed (France-Lanord et al., 1997; Puranam et al., 1997; Ariga et al., 1998; Taniwaki et al., 1999; Yuan and Yankner, 2000).

Caspase activation has been shown in most apoptosis models and two main pathways: a receptor-activated caspase-8 and -3 or a cytochrome c release-dependent caspase-9 and -3 pathway have been proposed (Krammer, 2000). In both pathways, caspase-3 activation is the final event of caspase cascade (Krammer, 2000). We show here that 1,4-B activates caspase-8, -9, and -3, and that caspase-3 inhibition does inhibit apoptosis. These results also correlate with the 1,4-B-induced m, release of cytochrome c and the consequent caspase-9 activation. However, like other apoptosis inducers such as DEX, 1,4-B also activates caspase-8, which has been recently described to regulate cytochrome c release and decrease in m (Li et al., 1998b; Luo et al., 1998).

In conclusion, these results indicate that, depending on the chemical structure, some of the tested 1,4-B analogs induce cell death through the activation of a complex cascade of intracellular biochemical events that have been previously correlated to a number of experimental models of apoptosis activation. Those events could contribute to some of the pharmacological effects associated with 1,4-B treatment, including the previously described neurotoxicity and antitumor activity. Future studies will be devoted to analyzing the possible role of 1,4-B-induced apoptosis in other experimental systems, including in vivo neurotoxicity and antitumor activity.