Division of Pharmacology and Toxicology, College of Pharmacy, The University of Texas at Austin, Austin, Texas 78712-1074
Received June 1, 1999; accepted August 23, 1999
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ABSTRACT |
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Key Words: caspase-3; nordihydroguaiaretic acid; apoptosis; lipoxygenase; bcl-xL; mitochondrial potential; N-acetylcysteine; glutathione.
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INTRODUCTION |
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Lipoxygenase (LOX) products, including hydroperoxy (HPETEs) or hydroxy (HETEs) fatty acids, have been implicated as important elements in the regulation of tumor cell growth by modulating cell proliferation and apoptosis (Tang et al., 1996). Blocking the activity of LOXs will induce apoptosis in some cell lines. For example, nordihydroguaiaretic acid (NDGA), a general LOX inhibitor, induces apoptosis in some, but not all cell lines tested, with W256 cells, a monocytic cell line (Tang et al., 1996
; 1997), being the most sensitive. NDGA also potently induces apoptosis in FL5.12 cells, a pro-B lymphocytic cell line (Datta et al., 1998
) but blocks apoptosis in nerve, glioma, neuroblastoma, melanoma, and erythroleukemia cells (Li et al., 1997
; Maccarrone et al., 1997
; Wagenknecht et al., 1997
).
Because of the discrepant effects of NDGA in terms of apoptosis, it is possible that the inhibition of LOX by this compound is not related to its ability to induce this form of cell death. NDGA-induced apoptosis is associated with lipid peroxidation and the depletion of glutathione (GSH) (Tang and Honn, 1997). Although NDGA is usually considered an antioxidant, this suggests some pro-oxidant activity in at least some biological systems. Since oxidants are capable of inducing apoptosis (Payne et al., 1995
), this effect could explain the pro-apoptotic activity of NDGA.
The concept that NDGA-induced apoptosis is unrelated to LOX activity is also supported by our finding that NDGA induces apoptosis in murine FL5.12 cells that lack LOX protein and activity (Datta et al., 1998). The current study documents the pro-oxidant activity of NDGA by showing increases in cellular levels of glutathione disulfide (GSSG) as well as lipid peroxidation products. Pretreatment with N-acetylcysteine (NAC) blocked apoptosis indicating an oxidative mechanism related to thiols. Interestingly, there was no evidence of an increase in free levels of reactive oxygen species (ROS). There was, however, a complete loss of mitochondrial membrane potential that could be blocked by NAC in NDGA-treated cells. Compared to the dramatic decrease of bcl-xL expression after treatment of these cells with the 5-lipoxygenase-activating protein inhibitor, MK886, which also induces apoptosis (Datta et al., 1998
), the expression of this anti-apoptotic protein did not change after treatment with NDGA. Although activation of caspase-3 was evident after treatment with NDGA, blocking this activity with only the most general caspase inhibitor affected apoptosis.
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MATERIALS AND METHODS |
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Measurements of apoptosis.
A characteristic event in apoptosis is DNA fragmentation and release of nucleosomes into the cytoplasm. These can be detected by an ELISA assay that is quite sensitive and specific for apoptosis relative to necrosis (Salgame et al., 1997). Cells were gently lysed releasing mono- and oligonucleosomes from apoptotic cells. A 1-step ELISA kit (Roche Molecular, Indianapolis, IN) using streptavidin-coated microplates, biotinylated antihistone, and peroxidase conjugated anti-DNA antibodies completes the assay. Results are expressed as the
A405490, which indicates the enrichment of nucleosomes in the cytoplasm.
Apoptosis was quantitated by visualizing distinctive nuclear and cytoplasmic fluorescence, chromatin condensation and formation of multiple apoptotic bodies following staining of cells with acridine orange and ethidium bromide (Duke and Cohen, 1992). The validity of the acridine orange/ethidium bromide assay for detecting apoptosis in this cell line has been previously documented (Bojes et al., 1997
). Treated and control cells were pelleted at 300 x g for 5 min at 4°C. The cells were then resuspended in 20 µl of fresh medium containing 4 µl of ethidium bromide/acridine orange mixture (1:1 v/v, stock solution 100 µg/ml each). This suspension (10 µl) was examined under a fluorescent microscope in order to differentiate early from late apoptotic cells and normal and necrotic cells. A minimum of 400 cells per sample was counted (magnification, 40x).
Additional confirmation of apoptosis was by quantitation of viable, early and late apoptotic (secondary necrotic) cells using a Coulter EPICS-XL flow cytometer with the Annexin V FITC kit (Coulter, Miami, FL), which utilizes phosphatidylserine externalization as a marker of apoptosis.
Glutathione assay.
Total glutathione (GSH + GSSG) and GSSG were determined using a modified method of Neuschwander-Tetri and Roll (1989). Cells (5 x 106/ml) were collected following incubation with NDGA and centrifuged for 10 min at 300 x g at 4°C. The pellet was resuspended in 1 ml EDTA (2 mM) and sonicated for 2 min. Total glutathione was determined in the cell lysate by adding 83 µl of 25 mM NaH2PO4 (pH 7.0) to 250 µl of lysate. An additional 250 µl of cell lysate was mixed with 83 µl of N-ethylmaleimide for GSSG measurements. All samples (200 µl) were then mixed with 200 µl 25 mM dithiothreitol (in 25 mM NaH2PO4, pH 7.0) followed by 100 µl Tris buffer (pH 8.5), and were incubated on ice for 30 min. Following addition of 0.5 ml of 2.5% (w/v) sulfosalicylic acid, the samples were centrifuged for 10 min at 600 x g at 4°C. An aliquot (200 µl) of supernatant was mixed with 200 µl of o-phthalaldehyde (5 mg/ml in 0.4 M potassium borate solution, pH 9.9) and incubated for 2 min at 25°C. The mixture was neutralized and samples were either kept on ice in the dark and used immediately or were stored at 80°C. Derivatized samples were analyzed by HPLC as described previously (Liu and Kehrer, 1996).
Lipid peroxidation assay.
Lipid peroxidation was assessed by the thiobarbituric acid assay (Ohkawa et al., 1979). Cells were washed in phosphate-buffered saline before undergoing 3 cycles of freeze-thawing in 200 µl of water. A 20 µl aliquot was then removed for protein determination (Bradford, 1976
). The assay mix (0.4% (w/v) thiobarbituric acid, 0.5% (w/v) SDS, 9.4% (v/v) acetic acid, pH 3.5; 980 µl) was added to the remaining sample. Samples were incubated for 60 min at 95°C, cooled to room temperature, and centrifuged at 14,000 x g for 10 min. A532 of the supernatants was determined.
Detection of ROS.
The generation of ROS was assessed using 2 different cell-permeable flurogenic probes, dihydorhodamine 123 (DHR123) and dichlorofluorescin diacetate (DCHFDA). Cells were preloaded with either DHR123 (1 µM) or DCHFDA (10 µM) for 30 min, respectively. Cells were then treated with 10 µM NDGA and the samples analyzed after 30 min, 1 h, and 3 h in an Epics XL flow cytometer. H2O2 (0.01%) was used as a positive control.
Measurement of mitochondrial membrane potential.
Changes in mitochondrial membrane potential were detected using the fluorescence-based ApoAlertTM mitochondrial membrane sensor kit (Clontech Laboratories, Palo Alto, CA). Cells (106/ml) were treated with 10 µM NDGA for 2 and 4 h. After treatment, cells were pelleted, washed with PBS, and resuspended in 1 ml MitoSensorTM reagent. Following incubation at 37°C in 5% CO2 for 20 min, cells were diluted with 1 ml of Clontech incubation buffer and then pelleted. The final cell pellet was resuspended in 30 µl fresh incubation buffer and cells were observed by fluorescence microscopy, using a Nikon E800 microscope equipped with a FITC/Texas Red, dual-band filter set (#51006, Chroma Technology Corp., Brattleboro VT; wavelengths in nm: excitation 497, 571; dichroic mirror 375410, 513552; barrier filter 531, 627). The MitoSensorTM reagent, cationic by nature, is taken up by normal mitochondria where it forms aggregates that exhibit red fluorescence. A loss in mitochondrial membrane potential prevents this aggregation and MitoSensor remains in the cytoplasm as a monomer where it exhibits green fluorescence.
Immunoblotting.
Cells were pelleted and lysed by repeated pipetting in 15 µl/106 cells ice-cold buffer containing 10 mM Tris (pH 7.4), 10 mM NaCl, 3 mM MgCl2, 1 mM EDTA, 0.1% (v/v) NP-40, 1 mM phenylmethylsulfonyl fluoride, 1 mM sodium orthovanadate, and 30 µl/ml aprotinin. Cell lysates were centrifuged at 16,000 x g for 10 min at 4°C and total protein content in the supernatant was measured (Bradford, 1976). Supernatants were run on 15% reducing SDSpolyacrylamide gels. Protein was transferred to PVDF membranes (Millipore, Bedford, MA) and blocked for at least 1 h in 5% (w/v) non-fat dry milk (BioRad, Hercules, CA). The membrane was then incubated for 1 h with anti-bcl-xL rabbit polyclonal antibody (1:1500), or a goat polyclonal antibody (1:1500) that recognizes both the pro-caspase-3 (32-KD) and the p20 cleavage product subunit (both antibodies were from Santa Cruz Biotechnology, Santa Cruz, CA). The proteins were visualized using an enhanced chemiluminescence detection kit (Amersham Pharmacia Biotech, Arlington Heights, IL).
Measurement of caspase-3 activity.
Ac-DEVD-7-amino-4-methyl-coumarin (Ac-DEVD.AMC) was used as a fluorometric substrate for determining caspase-3 activity (Thornberry et al., 1992). Cells (0.7 x 106) were pelleted and lysed in 200 µl of 50 mM Tris (pH 7.5) containing 150 mM NaCl, 0.5 mM EDTA, and 0.5% v/v NP-40. Aliquots (50 µl) of the lysate were incubated with tetrapeptide substrate (40 µM), 10 mM HEPES (pH 7.5), 50 mM NaCl and 2.5 mM DTT in a final volume of 200 µl for 2 h at 37°C. The fluorescence of the released 7-amino-4-methyl coumarin was measured at excitation and emission wavelengths of 360 nm and 460 nm.
Statistical analyses.
Data are expressed as the means ± SE. Comparisons between groups were carried out using ANOVA followed by post hoc analyses with Student Newman-Keul's test. A p value of less than 0.05 was considered significant.
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RESULTS |
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Boc-asp.FMK, but not Ac-DEVD.CHO, was able to inhibit NDGA-induced cell death. This was evident using flow cytometry to assess changes in phosphatidylserine distribution (Table 5), acridine orange/ethidium bromide staining to assess nuclear morphology and membrane permeability, and an ELISA assay to assess DNA fragmentation (Table 6
). Protection by boc-asp.FMK was virtually complete at 8 h. At 18 h, the protection provided by boc-asp.FMK remained complete as assessed by ELISA (Table 7
), but was no longer complete in terms of phosphatidylserine externalization as measured by flow cytometry (Table 5
) and was absent using acridine orange/ethidium bromide staining (63% apoptosis with NDGA and 61% with NDGA + boc-asp.FMK). These data suggest that boc-asp.FMK delays, but does not prevent NDGA-induced apoptosis. However, since NDGA was present throughout the time period examined, the delay may simply reflect the gradual loss of boc-asp.FMK caspase inhibition activity in the presence of continued levels of NDGA.
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DISCUSSION |
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A report that NDGA leads to lipid peroxidation and a loss of GSH, but that 12-LOX was involved (Tang and Honn, 1997), made it of interest to determine whether oxidative processes could be detected following induction of apoptosis with NDGA in FL5.12 cells, which lack LOX. A substantial increase in GSSG was observed in FL5.12 cells, confirming and extending this prior report. The occurrence of lipid peroxidation in FL5.12 cells is also consistent with the conclusion that NDGA stimulates oxidative reactions. The failure to find changes in DHR123 or DCFH fluorescence after treatment with NDGA was, therefore, surprising. A previous study in W256 cells treated with NDGA and loaded with DHR123 reported the same result (Tang and Honn, 1997
). These authors concluded, based on this finding and other data, that NDGA causes oxidative stress through non-conventional mechanisms. The current data are consistent with this conclusion and suggest that rather than stimulating the production of ROS, NDGA is perhaps directly oxidizing GSH leading to mitochondrial depolarization and apoptosis.
Support for the idea that NDGA acts through a thiol oxidation mechanism comes both from prior work showing that the apoptosis induced by NDGA can be effectively inhibited by GSH-elevating thiol agents and lipid peroxidation inhibitors (Tang and Honn, 1997) and the current study demonstrating that NAC protects against the NDGA-induced loss of mitochondrial potential and apoptosis in FL5.12 cells. Numerous studies have implicated mitochondria in apoptosis (Green and Reed, 1998
), and NDGA is known to have some mitochondrial effects. For example, NDGA inhibits succinate cytochrome c reductase in EMT6 mouse mammary carcinoma cells while sulfhydryl compounds, including GSH and cysteine, prevent the inhibition of succinoxidase activity by NDGA (Shi and Pardini, 1995
). This suggests that NDGA exerts its biological effects by oxidation of sulfhydryl groups in key mitochondrial systems. The mechanism of such oxidation is unclear, but it could be a direct effect of NDGA or occur indirectly following the formation of GSSG, perhaps as a result of some sort of NDGA-dependent redox cycle.
The disruption of the glutathione redox state through the formation of GSSG may be the critical factor in the induction of apoptosis, since the simple depletion of GSH by BSO is unable to cause apoptosis in this (Bojes et al., 1999) or other (Tang and Honn, 1997
) cell lines. The oxidation of mitochondrial GSH may explain the profound loss in mitochondrial membrane potential by 2 to 4 h after treating cells with NDGA. Although this effect cannot yet be directly linked to the apoptosis observed in this system, the temporal data are suggestive. Specifically, NDGA causes lipid peroxidation by 1 h and GSSG oxidation at 4 h, while no mitochondrial depolarization was evident at 2 h but was complete at 4 h. Since NAC prevents both mitochondrial depolarization and apoptosis, and since other data demonstrate that changes in mitochondrial permeability and thus mitochondrial potential are critical steps in apoptosis (Susin et al., 1998
), the theory that a relationship between NDGA-induced thiol oxidation, mitochondrial depolarization, and apoptosis is at least consistent with the current data. It is also possible that the NDGA-catalyzed formation of lipid peroxides represents an important apoptosis-signaling mechanism since such species are known to induce apoptosis (Sandstrom et al., 1994
).
Multiple lines of evidence indicate that caspases are important effectors of apoptosis (Thornberry and Lazebnik, 1998). Based on structural homology, caspases can be broken into three subfamilies: (1) Interleukin-converting enzyme (ICE) subfamily, (caspase-1,-4 and 5); (2) CED-3/CPP32 subfamily, (caspases-3, -6, -7, -8, -9 and -10); and (3) ICH-1/Nedd2 subfamily (Cohen, 1997
). The activation of caspase-3 in particular is considered essential for the progression of apoptosis (Kuida et al., 1998
). The current study provides the first report that NDGA treatment activates caspase-3. Blocking general caspase activity with boc-asp.FMK largely prevented NDGA-induced apoptosis. In contrast, a more specific caspase-3 inhibitor, Ac-DEVD.CHO, was ineffective. These results suggest that while apoptosis induced by NDGA may require caspase activity, a specific requirement for caspase-3 is unclear. Since not all apoptotic cell deaths are inhibited by caspase inhibitors (Bojes et al., 1999
; Borner and Monney, 1999
; Xiang et al., 1996
), and since the specificity of these inhibitors is not absolute, it is likely that other caspase family members, particularly caspase-9, are involved.
In summary, a low dose of NDGA induced rapid and extensive apoptosis in FL5.12 cells. This effect was accompanied by an increase in GSSG and lipid peroxidation, and could be blocked by NAC. The massive loss of mitochondrial membrane potential evident at the time apoptosis could first be detected could also be blocked with NAC. A significant increase in caspase-3-like activity was seen after NDGA. However, the specific inhibition of caspase-3 did not protect cells from NDGA-induced apoptosis, while the more general inhibition of caspase activities delayed this effect. Thus, caspases other than caspase-3 may be important. The toxicity of NDGA appears to be caused through its pro-oxidant effects on thiols that generate pro-apoptotic signals and lead to a loss of mitochondrial membrane potential.
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ACKNOWLEDGMENTS |
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NOTES |
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2 To whom correspondence should be addressed. Fax: (512) 471-5002. E-mail: KehrerJim{at}mail.utexas.edu.
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