Morphological Transformation and Oxidative Stress Induced by ...

TOXICOLOGICAL SCIENCES 68, 437– 443 (2002) Copyright © 2002 by the Society of Toxicology

Morphological Transformation and Oxidative Stress Induced by Cyanide in Syrian Hamster Embryo (SHE) Cells Lisa M. Kamendulis, Haizhou Zhang, Yanhong Wang, and James E. Klaunig 1 Division of Toxicology, Department of Pharmacology and Toxicology, Indiana University School of Medicine, 635 Barnhill Dr., MS 1021, Indianapolis, Indiana 46202 Received February 2, 2002; accepted April 5, 2002

Cyanide is a well-established poison known for its rapid lethal action and toxicity. Although long-term mammalian studies examining the carcinogenic potential of cyanide have not been previously reported, cyanide was reported to be positive in Salmonella typhimurium mutagenesis assay and induced aneuploidy in Drosophila. To further evaluate the carcinogenic potential of cyanide, the ability of cyanide to induce morphological transformation in Syrian hamster embryo (SHE) cells was studied. Cyanide induced a dose-dependent increase in morphological transformation in SHE cells following a 7-day continuous treatment. A significant increase in transformation was observed at potassium cyanide doses of 200 ␮M and greater. Transformation induced by cyanide was inhibited in a dose-related manner by vitamin E, suggesting a role of oxidative stress in the induction of morphological transformation by cyanide. Further, it was shown that 500 ␮M cyanide induced oxidative DNA damage in SHE cells, evidenced by the formation of 8-hydroxy-2ⴕ-deoxyguanosine (50 – 66% increase over control). The induction of oxidative stress by cyanide involved an early and temporal inhibition of antioxidant enzymes (catalase and superoxide dismutase) as well as an increased production of reactive oxygen species (1.5- to 2.0-fold over control). Key Words: catalase; cyanide; morphological transformation; oxidative stress; SHE cells; superoxide dismutase.

Cyanide is a potent neurotoxicant that rapidly produces lethality by depleting cellular adenosine triphosphate as a result of inhibition of mitochondrial cytochrome oxidase, the terminal enzyme in the electron transport chain (Isom and Way, 1984). Because of the high energy requirement and low anaerobic capacity, the central nervous system, in particular, is one of the primary targets of cyanide toxicity (Way, 1984). Cyanide produces several effects on the central nervous system, ranging from central nervous system dysfunction to degeneration of selective brain areas (Blanc et al., 1985; Carella et al., 1988; Chandra et al., 1980; Funata et al., 1984; Kales et al., 1997). In addition to inhibition of oxidative phosphorylation, cyanide produces central nervous system toxicity through activation of NMDA receptors, disruption of neurotransmitter 1

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function, and elevation of cellular free calcium (Isom et al., 1999; Patel et al., 1994). Several short-term in vitro and in vivo test systems have been used to assess the mutagenicity and carcinogenicity of chemicals. Cell transformation in Syrian hamster embryo (SHE) cells measures the carcinogenic potential of xenobiotics by assessing transformed colonies based on morphological criterion (Berwald and Sachs, 1963). This in vitro system exhibits a multistage transformation process similar to that seen in carcinogenesis in vivo (Barrett and Ts’o, 1978; Isfort et al, 1994; LeBoeuf et al., 1990). Morphological transformation is the first identifiable stage during the transformation process and shows a ⬎ 85% concordance rate with the 2-year rodent bioassays (Isfort et al., 1996; LeBoeuf et al., 1996). Oxidative stress resulting from either overproduction of prooxidants or deficiency of antioxidants has been suggested as an important factor in many degenerative diseases, such as rheumatoid (Mapp et al., 1995), cardiovascular, and neurodegenerative diseases (Giacosa and Filiberti, 1996; Simonian and Coyle, 1996). Several studies have shown that the induction of oxidative stress is involved in cyanide-induced neurodegenerative disease (Akira et al., 1994; Ardelt et al., 1989; Gunasekar et al., 1996). Resulting from oxidative stress, cyanide induces lipid peroxidation in vitro that is reversible by coincubation with N-acetyl cysteine and curcumin (Bhattacharya et al., 1999). The induction of oxidative stress by cyanide may involve increases in reactive oxygen species and nitric oxide (Gunasekar et al., 1996; Mills et al., 1996), inhibition of antioxidant systems (Ardelt et al., 1989), and inhibition of mitochondrial function (Way, 1984). Inhibition of mitochondrial function can result in the production of additional reactive oxygen species, principally the superoxide anion. In addition, the basal ganglia are affected by cyanide exposure, resulting in neurological sequelae similar to those seen in individuals with Parkinson’s disease (Carella et al., 1988). Oxidative stress has been shown to participate in the carcinogenesis process (Guyton and Kensler, 1993; Trush and Kensler, 1991; Vuillaume, 1987). Although no direct evidence exists to link chronic exposure to low doses of cyanide with cancer development, cyanide was shown to induce aneuploidy

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in Drosophila. Several groups have identified a causal link between the induction of aneuploidy and cancer development (Dellarco et al., 1986; Holliday, 1989; Li et al., 2000; Osgood and Sterling, 1991). Cyanide has also tested positive in the Salmonella typhimurium mutagenicity assay (Kushi et al., 1983). In the present study, cyanide was evaluated for its ability to induce morphological transformation in the SHE cell system. In addition, the examination of oxidative stress end points (evaluation of oxidative damage and antioxidant status) following cyanide exposure were examined in SHE cells in an attempt to elucidate the potential mechanism(s) by which cyanide induces cellular transformation. MATERIALS AND METHODS Chemicals. Potassium cyanide, L-glutamine, dimethylsulfoxide (DMSO), citric acid, sodium acetate, cacodylic acid, diethylenetriamine pentaacetic acid, KMnO 4, pyrogallol, 30% H 2O 2, and Hanks balanced salt solution of highest purity available were obtained from Sigma (St. Louis, MO). Fetal bovine serum was purchased from HyClone (Logan, UT). DMEM LeBoeuf’s modification (DMEM-L) was from Quality Biological (Gaithersburg, OH). SHE cell transformation assay. SHE cell transformation was conducted as described previously (Kerckaert et al., 1996). Briefly, feeder cells were prepared by reconstituting cryopreserved SHE cells in DMEM-L and culturing at 37°C, 10% CO 2, 95% humidity for 72 h. The cells were then rinsed with CMF-HBSS, detached (0.05% trypsin-EDTA, 5 ml), resuspended in 30 ml culture medium, and exposed to X-ray (⬃ 5000 rad) such that the cells remained viable but not capable of replication. The cells were plated at a density of 2 ⫻ 10 4/ml in 60-mm culture dishes (2 ml/dish) and incubated for 24 h. Target cells (cells to be used for clonal growth) were prepared as described above for feeder cells. After 24 h, target cells were plated on feeder cells at a density of 85 cells/dish in 2 ml complete medium. Following an additional 24-h incubation, cells were treated and allowed to grow for 7 days. Colonies were then fixed with methanol, stained with Giemsa, and scored for morphological transformation as described (Kerckaert et al., 1996). Cell culture and treatment. For assays other than the examination of morphological transformation, cells were cultured as follows. Cells were isolated from the embryos of Syrian golden hamsters at day 13 of gestation (Charles River, Portage, MI) and cryopreserved as described (Kerckaert et al., 1996). For each experiment, cryopreserved SHE cells (2 ⫻ 10 6) were thawed and cultured in DMEM-L at 37°C, 10% CO 2, and 95% humidity. At 90% confluency, cells were trypsinized and plated at a density of 1 ⫻ 10 6 cells per 60-mm culture dish. Cell cultures were then treated when approximately 75% confluent. Analysis of 8-hydroxydeoxyguanosine (OH8dG). OH8dG was measured as described previously (Shigenaga et al., 1994). Briefly, DNA was isolated from nuclei using a sodium iodide choatropic method (Wang et al., 1994). DNA was dissolved in Tris-HCl buffer (10 mM, pH 7.0) containing 2 mM butylated hydroxytoluene (BHT) and 0.1 mM desferral. DNA (100 –200 ␮g) was digested sequentially with nuclease P 1 (10 units, 37°C, 30 min) followed by alkaline phosphatase (14 units, 37°C, 60 min). After centrifugation (10,000 ⫻ g, 10 min, 4°C), the supernatant was removed for analysis. OH8dG was measured by HPLC and detected electrochemically (set at E1: 100 mV, 1 ␮A; E2: 400 mV, 5 ␮A; esa, Inc., Chelmsford, MA). 2⬘-Deoxyguanosine (2⬘-dG) was detected using an ultraviolet detector (Waters PDA, 260 nm; Waters Inc., Milford, MA). OH8dG and 2⬘-dG were quantified from standard curves and expressed as the ratio of OH8dG to 2⬘-dG. Detection of hydroxyl radicals. Hydroxyl radical formation in SHE cells was determined by monitoring the nonenzymatic aromatic hydroxylation of

salicylic acid, a reaction that reflects hydroxyl radical production (Floyd et al., 1984). Four hours prior to harvest, salicylic acid (10 mM) was added to SHE cell cultures. After treatment, trichloroacetic acid was added (5% v/v final concentration) and the cell cultures were incubated on ice for 15 min prior to collection. The cell extracts were centrifuged (10,000 ⫻ g, 10 min). The resultant supernatant was injected into HPLC and 2,3-DHBA resolved on a Nova-Pak C 18 reversed-phase analytical column and eluted with sodium citrate buffer (30 mM, pH to 4.65) at a flow rate of 1.3 ml/min. 2,3-DHBA was detected electrochemically (ESA Colouchem II, set at 0.5 ␮A, 350 mV; esa, Inc., Chelmsford, MA). Salicylic acid was measured at 296 nm (Waters PDA, Waters, Inc. Milford, MA). 2,3-DHBA and salicylic acid were quantitated from standard curves, and the results expressed as the ratio of 2,3-DHBA relative to salicylic acid. Enzymatic antioxidant activity assays. After treatment, SHE cells were washed twice with PBS (0°C) and collected. Cell pellets were resuspended in lysis buffer (0.1% triton X-100, 10 mM potassium phosphate buffer, pH 7.2). Cell membranes were disrupted by sonication (10 s; Fisher model 550 sonic dismembranetor; Fisher Scientific, Pittsburgh, PA). Cell lysates were centrifuged (13,000 ⫻ g; 15 min), and the supernatants were removed for enzyme measurement. Protein content in supernatants was determined using the BioRad DC protein assay, based on the method of Lowry et al. (1951), with bovine serum albumin serving as a standard. Superoxide dismutase activity was measured using the inhibition of the autooxidation of pyrogallol (Marklund and Marklund, 1974). Briefly, an aliquot of cell extract was mixed with buffer (50 mM Tris HCl, 50 mM cacodylic acid, 1 mM diethylenetriamine pentaacetic acid, pH 8.2) containing 2 mM pyrogallol. The autooxidation of pyrogallol and the inhibition of this reaction were monitored spectrophotometrically. Under the conditions of this assay, one unit of superoxide dismutase activity was equivalent to the amount of enzyme that resulted in a 50% inhibition of pyrogallol autooxidation. Catalase activity was determined by monitoring the enzyme-catalyzed decomposition of hydrogen peroxide by potassium permanganate (Cohen et al., 1970). Samples (50 ␮l) were added to test tubes, followed by the addition of H 2O 2, then allowed to incubate on ice for 3 min. The reaction was stopped by the addition of H 2SO 4. KMnO 4 was then added, and absorbance was recorded at 480 nm. Under the conditions of this assay, one unit of enzyme activity equals k/(0.00693) (Aebi, 1974), where k ⫽ log (S 0/S 2) ⫻ (2.3/t), S 0 ⫽ absorbance of standard – absorbance of blank, S 2 ⫽ absorbance of standard – absorbance of sample, and t ⫽ time interval. The measured activities were normalized to the protein content of each sample. Statistics. For studies examining morphological transformation, data were analyzed using the Fisher exact test (Kerckaert et al., 1996). For all other studies, the data were analyzed by one-way ANOVA followed by Duncan’s test. For all studies, treatment groups were considered statistically different from controls at p ⬍ 0.05.

RESULTS

Morphological Transformation Induced by Cyanide An initial study evaluating the cytolethality of cyanide in SHE cells was performed to determine the concentration of cyanide to be used in subsequent studies. SHE cells were treated with potassium cyanide for 7 days at concentrations ranging from 10 to 5000 ␮M. A reduction in relative plating efficiency of ⬎ 50% was considered cytolethal (Kerckaert et al., 1996). Cyanide treatment reduced relative plating efficiency in a concentration-related manner (Table 1). Cyanide (1000 ␮M) reduced relative plating efficiency to 54%. Because concentrations above 1000 ␮M were cytolethal, this concentration was selected as the highest concentration utilized in cellular transformation studies.

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TABLE 1 Cytotoxicity of Cyanide (KCN) in SHE Cells following a 7-Day Treatment Dose (␮M)

Total no. colonies

Average no. colonies

RPE (%)

0 20 50 200 500 1000 2500 5000

505 518 512 405 347 275 204 129

33.7 34.5 34.1 27.0 23.1 18.3 13.6 8.6

100.0 102.0 101.0 80.0 68.5 54.3 40.3 25.5

Note. Average number of colonies ⫽ total number of colonies/number of dishes counted. RPE, relative plating efficiency (treated group average number of colonies/dish ⫼ untreated control number of colonies/dish) ⫻ 100.

Morphological transformation was assessed in SHE cells treated with 20 –1000 ␮M potassium cyanide for 7 days (Fig. 1). Cyanide exposure resulted in a concentration-related increase in morphological transformation. Significant increases in morphological transformation were observed at cyanide concentrations of 200, 500, and 1000 ␮M (Fig. 1). Benzo(a)pyrene was used as a positive control, which showed increases in morphological transformation consistent with historical controls. Effect of Vitamin E on Cyanide-Induced Morphological Transformation The effect of vitamin E (50 –200 ␮M) on cyanide-induced (500 mM) SHE cell morphological transformation was examined (Fig. 2). Vitamin E alone had no apparent effect of morphological transformation. Cyanide (500 ␮M), as shown previously, significantly increased morphological transforma-

FIG. 2. Effect of Vitamin E on potassium cyanide-induced morphological transformation in SHE cells. Morphological transformation frequency (no. of transformed colonies/total no. of colonies) was determined in SHE cells treated with 500 ␮M potassium cyanide in the presence or absence of vitamin E (50, 100, or 200 ␮M) for 7 days. *Significantly different from control (p ⬍ 0.05; one-sided Fisher Exact Test).

tion. The induction of morphological transformation by 500 ␮M cyanide was reduced in a concentration-dependent manner by vitamin E supplementation (Fig. 2). All the vitamin E concentrations examined significantly reduced cyanide-induced cell transformation. The positive control [benzo(a)pyrene] increased cellular transformation. Effect of Cyanide on Hydroxyl Radical Formation The formation of the reactive oxygen free radical (the hydroxyl radical), assessed by 2,3-DHBA formation (Floyd et al., 1984), was measured in SHE cells after treatment with 20 ␮M or 500 ␮M potassium cyanide for 4 h, 1 day, 2 days, and 7 days. No significant change in hydroxyl radical production was observed after treatment with 20 ␮M potassium cyanide at any of the time points examined. In contrast, 500 ␮M potassium cyanide induced a significant and consistent increase in 2,3 DHBA formation following 1, 2, or 7 days of treatment (37.4, 43.5, and 83.6% increase over controls, respectively; Fig. 3). Effect of Cyanide on OH8dG Production

FIG. 1. Effect of potassium cyanide on morphological transformation in SHE cells. Morphological transformation frequency (no. of transformed colonies/total no. of colonies) was calculated in SHE cells incubated with 10 – 1000 ␮M KCN for 7 days. *Significantly different from control (p ⬍ 0.05; one-sided Fisher Exact Test).

The ability of cyanide (20 and 500 ␮M) to induce oxidative DNA damage was examined in SHE cells. OH8dG formation in SHE cells was used as a marker for oxidative DNA damage. No significant increase in OH8dG was seen in SHE cells treated with 20 ␮M cyanide at any time point examined (Fig. 4). Cyanide (500 ␮M) produced a significant increase in OH8dG following treatment for 1 and 2 days (78% and 76% increase over controls, respectively). OH8dG levels, however, returned to control levels after 7 days of continuous treatment with 500 ␮M potassium cyanide (Fig. 4).

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FIG. 3. Effect of potassium cyanide on 2,3-DHBA formation in SHE cells. SHE cells were plated in 60-mm dishes at a density of 1 ⫻ 10 6 cells/dish. When cells were 75% confluent, the cells were treated with 20 mM or 500 ␮M potassium cyanide for 4 h, 1, 2, and 7 days. 2,3-DHBA was determined by HPLC with EC detection. Data are reported as mean ⫾ SD (n ⫽ 4). *Significantly different from control (p ⬍ 0.05; ANOVA followed by Duncan’s test).

Effect of Cyanide on Catalase and Superoxide Dismutase Activities Catalase activity after treatment with 500 ␮M potassium cyanide was decreased by 42% and 68% from controls at 4 h and 1 day, respectively (Fig. 5). A minor inhibition of catalase activity was also seen after treatment with 20 ␮M cyanide at 4 h and 1 day (17% and 12% decrease from controls, respec-

FIG. 5. Effect of potassium cyanide on catalase activity in SHE cells. SHE cells were plated in 60-mm dishes at a density of 1 ⫻ 10 6 cells/dish. When cells were 75% confluent, the cells were treated with 20 ␮M or 500 ␮M potassium cyanide for 4 h, 1, 2, and 7 days. Catalase activity was determined spectrophotometrically and reported as mean ⫾ SD (n ⫽ 4). *Significantly different from control (p ⬍ 0.05; ANOVA followed by Duncan’s test).

tively). However, after 2 days of treatment, both 20 and 500 ␮M potassium cyanide resulted in a significant increase in catalase activity (80% and 94% over controls, respectively). The increase in catalase activity by 20 and 500 mM cyanide remained through 7 days of treatment (32% and 36% over controls, respectively). Similarly, cyanide produced a time-related inhibition of superoxide dismutase activity (Fig. 6). A significant decrease in superoxide dismutase activity was seen in SHE cells treated with 500 ␮M potassium cyanide for 4 h and 1 day (57.2% and 35.5% decrease from control, respectively). However, after treatment with 500 ␮M cyanide for 2 days or longer, superoxide dismutase activity returned to control levels (Fig. 6). No significant decrease in superoxide dismutase activity was seen in SHE cells treated with 20 ␮M potassium cyanide at any time point examined (Fig. 6). DISCUSSION

FIG. 4. Effect of potassium cyanide on 8-hydroxydeoxyguanosine (OH8dG) formation in SHE cells. SHE cells were plated in 60-mm dishes at a density of 1 ⫻ 10 6 cells/dish. When cells were 75% confluent, the cells were treated with 20 ␮M or 500 ␮M potassium cyanide for 4 h, 1, 2, and 7 days. After treatment, DNA was isolated and OH8dG was determined using HPLC with EC detection. Data are reported as mean ⫾ SD (n ⫽ 4). *Significantly different from control (p ⬍ 0.05; ANOVA followed by Duncan’s test).

Results from the present study showed that cyanide induced a significant increase in morphological transformation in SHE cells at doses of 500 and 1000 ␮M. This is the first report showing that cyanide induces cellular transformation, an event that correlates with carcinogenicity in vivo (Kerckaert, 1996). Previous studies have shown that cyanide was mutagenic in Salmonella typhimurium (Kushi et al., 1983) and induced aneuploidy in Drosophila (Osgood and Sterling, 1991). Human exposure to cyanide occurs occupationally through groundwater contamination and consumption of foods containing cyanide. Neuropathological effects associated with chronic cyanide exposure have been recognized and well studied (Isom et

MORPHOLOGICAL TRANSFORMATION AND OXIDATIVE STRESS BY CYANIDE

FIG. 6. Effect of potassium cyanide on superoxide dismutase activity in SHE cells. SHE cells were plated in 60-mm dishes at a density of 1 ⫻ 10 6 cells/dish. When cells were 75% confluent, the cells were treated with 20 ␮M or 500 ␮M potassium cyanide for 4 h, 1, 2, and 7 days. Superoxide dismutase activity was determined spectrophotometrically and reported as mean ⫾ SD (n ⫽ 4). *Significantly different from control (p ⬍ 0.05 vs. control; ANOVA followed by Duncan’s test).

al., 1999). However, the carcinogenic effect of cyanide has not been adequately addressed. Cyanide has been previously shown to induce oxidative stress and damage in a number of biological systems. Administration of sublethal doses of potassium cyanide to CF1 mice induced lipid peroxidation in brain (Johnson et al., 1987). This action has been ascribed to the activation of xanthine oxidase (and subsequent increase in reactive oxygen species) as well as altered regulation of neuronal calcium homeostasis. The induction of lipid peroxidation by cyanide was also observed in cultured neurons from chick embryo telencephalon (Muller and Krieglstein, 1995) and in mouse brain cortical slices incubated with potassium cyanide in vitro (Ardelt et al., 1994). Several studies document an increase in reactive oxygen species following cyanide exposure. Through the inhibition of the mitochondrial respiratory chain at the ubiquinone-cytochrome b site, cyanide produces the superoxide anion. In addition, the generation of hydroxyperoxide was also seen in cyanide-treated PC12 cells (Ardelt et al., 1994; Mills et al., 1996), rat leukemia 2H3 cells (Arai et al., 1999), and cerebellar granule cells (Gunasekar et al., 1996). The formation of the superoxide anion was also shown to be produced nonenzymatically via interaction of cyanide with flavonoids (Hodnick et al., 1994). In addition to the generation of reactive oxygen species, cyanide is a potent inhibitor of the enzymatic antioxidants catalase, superoxide dismutase, and glutathione peroxidase (Ardelt et al., 1989; Kanthasamy et al., 1997). These mechanisms may function in concert to produce the oxidative stress and damage seen after cyanide exposure. The induction of oxidative stress by cyanide in SHE cells

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appears to involve both the production of reactive oxygen species and the depletion of the cellular antioxidants catalase and superoxide dismutase. Inhibition of these antioxidants was seen within a short duration after cyanide exposure (4-h and 1-day treatments). The inhibition of these enzymatic antioxidants may have accounted for or contributed to the increase in reactive oxygen species seen in SHE cells treated with cyanide at the early time points (4 h and 1 day). Although the cellular activity of superoxide dismutase had returned to control level and catalase activity was increased significantly after 2 days and 7 days of treatment with cyanide, the level of reactive oxygen species continued to be elevated over control throughout 7 days of study. This suggests that additional mechanisms must be functioning to maintain the level of reactive oxygen species seen in these studies. One such possibility is the inhibition of mitochondrial respiration and the subsequent generation of superoxide anion by cyanide. It is not clear whether cyanide remains bioavailable at the later time points or if, because of irreversible binding to the mitochondria, cyanide produces a sustained production of the superoxide anion through the time points examined in the present study. Additional studies are being performed to address this issue. The induction of oxidative stress by cyanide in SHE cells was further evidenced by the formation of the oxidative DNA adduct OH8dG. OH8dG is the most prevalent form of oxidative DNA damage and is commonly used as a biomarker for oxidative stress (Kasai, 1997; Grollman and Moriya, 1993; Shigenaga et al., 1994). The results of the present study showed that 500 ␮M cyanide induced significant increase of OH8dG in SHE cells after 1 and 2 days of exposure, whereas no increase in oxidative DNA damage was seen after 4 h of treatment. Thus, it appears that the induction of oxidative DNA damage occurs subsequent to an increase in hydroxyl radical formation by cyanide. Additionally, treatment with a lower nontransforming concentration of cyanide also failed to induce an increase in either hydroxyl radical formation or oxidative DNA damage, further supporting a link between the induction of oxidative stress by cyanide and cellular transformation. The formation of OH8dG has been associated with cancer development (Floyd, 1990). OH8dG is mutagenic in several in vitro systems (Kamiya et al., 1992; Moriya et al., 1991; Shibutani et al., 1991; Wood et al., 1990) and is known to inhibit human DNA methyltransferase in vitro (Turk et al., 1995). This suggests that OH8dG could be involved in carcinogenesis through both genotoxic and nongenotoxic mechanisms. Recent results from our laboratory have demonstrated a causal relationship between the level of OH8dG in SHE cells and the induction of morphological transformation (Zhang et al., 2000a). Collectively, the results from the present studies showing the inhibition of cyanide-induced morphological transformation by antioxidant cotreatment and the formation of OH8dG in SHE cells by cyanide provide direct evidence link-

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ing the induction of oxidative stress to cyanide-induced cellular transformation. The finding of increased catalase activity in SHE cells may be a result of gene regulation by reactive oxygen species. Low levels of reactive oxygen intermediates have been shown to modulate gene expression (Schulze-Osthoff et al., 1995). Several groups have demonstrated that the catalase and superoxide dismutase gene expression levels were upregulated by hydrogen peroxide and other agents that produce active oxygen species (Cornelissen et al., 1997; Diez-Fernandez et al., 1998; Rohrdanz and Kahl, 1998). Thus, the increase in hydroxyl radicals produced by cyanide at the early time points in the present study may serve as second messengers to regulate the expression of catalase and/or superoxide dismutase genes, resulting in the observed increases in the activity of these antioxidants seen at the later treatment points. Besides gene upregulation, catalase is regulated at both the posttranscriptional and posttranslational levels (Clerch, 1995; Reimer et al., 1994) as well as by gene amplification (Yamada et al., 1991). Further studies are needed to determine whether the increased catalase and superoxide dismutase activity is regulated at the transcriptional level, posttranscriptional level, or posttranslational level, as well as the signaling pathways involved in their regulation. In summary, the present study showed that cyanide induced morphological transformation in SHE cells after 7 days of continuous treatment. These results also demonstrated that oxidative stress (both the inhibition of antioxidant enzyme activity and production of reactive oxygen species) is involved in the induction of morphological transformation by cyanide. Results of the present study, together with findings demonstrating the induction aneuploidy in Drosophila and mutagenicity in Salmonella typhimurium by cyanide, point to the need for chronic bioassays to evaluate the carcinogenic potential of cyanide. REFERENCES

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