Mutagenesis, Vol. 18, No. 3, 293-298,
May 2003
© 2003 UK Environmental Mutagen Society/Oxford University Press
Aneugenic potential of okadaic acid revealed by the micronucleus assay combined with the FISH technique in CHO-K1 cells
1 AFSSA, Laboratoire d’Etudes et de Recherches sur les Médicaments Vétérinaires et les Désinfectants, Unité de Toxicologie Alimentaire, BP 90203, F-35302 Fougères, France, 2 Bureau de la Recherche et des Laboratoires d’Analyses, Direction Générale de l’Alimentation, Ministère de l’Agriculture, de l’Alimentation, de la Pêche et des Affaires Rurales, 251 Rue de Vaugirard, F-75732 Paris Cedex 15, France and 3 AFSSA, Laboratoire d’Etudes et de Recherches sur l’Hygiène et la Qualité des Aliments, Unité Toxines Microbiennes, 10 Rue Pierre Curie, F-94704 Maisons Alfort Cedex, France
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Abstract |
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Okadaic acid (OA) is a major toxin involved in diarrhetic shellfish poisoning in humans and has been shown to be both a potent tumor promoter in rodent skin and stomach and an inhibitor of serine/threonine protein phosphatases, specifically PP1 and PP2A. The research on the genotoxic potential of OA amounts to only a few studies, which give conflicting results. In order to evaluate the ability of OA to induce DNA damage, the cytokinesis-block micronucleus assay was performed in the CHO-K1 cell line. A statistically significant induction of micronuclei without strong cytotoxicity was obtained after a 24 h treatment with 20 (~5-fold) and 30 nM (~10-fold) OA. Then, in order to discriminate between a clastogenic or aneugenic effect of OA, the micronucleus assay was carried out in combination with fluorescence in situ hybridization (FISH) using a (TTAGGG)n DNA probe for centromere detection. FISH analysis showed that OA mainly induced centromere-positive micronuclei (68.9% induction with 20 nM OA and 77.0% with 30 nM). Therefore, OA can be considered aneugenic. Using the same assay, biotransformation of OA was studied after a 4 h treatment with and without metabolic activation. The results show that reactive metabolites of OA were generated with a significant increase in genotoxic potential. The relationship between the different components involved in the mitotic process and OA inhibition of protein phosphatase is also discussed.
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Introduction |
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Okadaic acid (OA) is a polyether derivative fatty acid produced by marine dinoflagellates. Its isolation from the marine sponge Halicondria okadaii and its structural determination were performed by Tachibana et al. (1981)
. OA is one of the major causative agents of diarrhetic shellfish poisoning (DSP) for which the most prominent human symptoms are gastrointestinal, including diarrhea, nausea, vomiting and abdominal pain (Yasumoto et al., 1978). DSP is widely distributed throughout the world, particularly in Japan and Western Europe, and is a serious problem for public health and the shellfish industry.
OA is a potent tumor promoter (Fujiki and Suganuma, 1993) but it belongs to a new class of tumor promoters which do not act on protein kinase C, like the PMA (phorbol esters) class. OA has been found to inhibit serine/threonine protein phosphatases (PPs), particularly PP1 and PP2A (Takai et al., 1987), to different extents: PP2A was the most strongly inhibited, followed by PP1; PP2B was less sensitive and PP2C was not inhibited at all (Cohen, 1989). In fact, PP1 and PP2A account for >90% of total PP activity in mammalian cells (Cohen, 1989). OA has been widely used to elucidate the role of phosphorylation reactions on serine/threonine residues in various cell processes and cell lines (for a review see Fernandez et al., 2002).
The research on the genotoxic potential of OA amounts to only a few studies, with contradictory results. OA has been shown to: (i) inhibit sister chromatid separation in HeLa cells (Ghosh et al., 1992); (ii) increase mutation frequencies (Aonuma et al., 1991; Rogers et al., 1994); (iii) induce DNA adducts in BHK-21 and HESV cells (Fessard et al., 1996); (iv) induce DNA strand breaks in Caco-2 cells (Traoré et al., 2001); (v) induce apoptosis in many cell types (Boe et al., 1991; Rossini et al., 2001). However, no mutation induction was reported in the Ames test with the Salmonella typhimurium strains TA100 and TA98 (Aonuma et al., 1991) and in the CHO/HGPRT assay (Fessard et al., 2002), with or without a metabolic activation system. In the present study, the effects of OA on DNA were evaluated further, using the cytokinesis-block micronucleus (CBMN) test in CHO-K1 cells. This cell type has been used routinely in in vitro micronucleus (MN) screening assays for the detection of genotoxicants (Gibson et al., 1998; Erexson et al., 2001). The CBMN methodology is based on the inhibition of cytokinesis by cytochalasin B (cyt B), allowing distinction between cells which did not divide, mononucleated cells, and those which either divided once, binucleated cells (BN), or divided more that once, polynucleated cells (PN) (Fenech and Morley, 1985). It also allows evaluation of the induction of MN in cultured cells (Fenech, 2000) and facilitates scoring chromosome damage in cells undergoing one round of nuclear division after exposure to genotoxic agents. MN could originate from fragments or whole chromosomes excluded from the main nucleus during cell division. Clastogenic versus aneugenic events can be distinguished further by a combination of the CBMN test with a fluorescence in situ hybridization (FISH) technique using centromere DNA probes (Farooqi et al., 1993; Schuler et al., 1997). Aneuploidy refers to a change in chromosome number which may arise either spontaneously or by chemical induction. It also plays a significant role in disorders such as cancer (for reviews see Mitelman, 1994; Duesberg et al., 1999; Cowell, 2001).
In this study, we report the results obtained in the CBMN assay in CHO-K1 cells exposed to various concentrations of OA, in the presence and absence of metabolic activation. To discriminate between a clastogenic or an aneugenic effect of OA, the CBMN assay was coupled to the FISH detection of centromeres in MN. The present results suggest that OA has an aneugenic effect and is more toxic in vitro after metabolic activation by a rat liver post-mitochondrial fraction.
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Materials and methods |
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Cells
Chinese hamster ovary cells (CHO-K1) were obtained from Eurobio. Cells were routinely maintained in Ham’s nutrient F12 medium (Life Technologies) with L-glutamate supplemented with 10% fetal calf serum (FCS) (Life Technologies) and 100 U penicillin and 100 µg/ml streptomycin at 37°C in a 5% CO2 humidified atmosphere.
Chemicals
OA was provided by Calbiochem. Cyclophosphamide (CP), methyl methanesulfonate (MMS) and cyt B were purchased from Sigma. Griseofulvin (GF) was obtained from CEVA Santé Animal (Libourne, France). OA was dissolved in ethanol whereas CP and MMS were diluted directly with serum-free Ham’s F12. GF and cyt B were dissolved in dimethylsulfoxide (DMSO) (Merck). The final concentration of ethanol and DMSO represented 0.5% of the final volume.
Cell treatment
Exponentially growing cells were plated in a 6-well plate on glass coverslips (1.5x104 cells/cm2) and cultured for 24 h prior to drug treatment. Duplicate coverslips were established for each concentration, and at least two independent experiments were conducted. Cells were exposed to the chemicals for 4 or 24 h, the control cultures receiving an equivalent amount of solvent.
With S9 activation
The liver post-mitochondrial fraction S9 (IFFA-CREDO, Lyon, France) was obtained from a male rat (OFA Sprague Dawley) treated with a phenobarbital/ß-naphthoflavone mixture (50 and 20 mg/kg, respectively). The S9 mix composition was as follows: 10% S9 homogenate, 32 mM KCl, 8 µM MgCl2, 0.5 mM glucose 6-phosphate, 0.4 mM NADP and 0.1 M phosphate buffer. As described before, 24 h after seeding the cells were exposed to fresh FCS-free medium containing 20% S9 mix and the appropriate drugs for only 4 h.
In vitro CBMN assay
At the end of treatment, cells were washed twice with phosphate-buffered saline (PBS) prior to a 20 h incubation in fresh medium containing 10% FCS and 3 µg/ml cyt B. At the end of cyt B treatment, the cells were washed twice with PBS and allowed to recover for 1.5 h in fresh 10% FCS medium.
Slide preparation
For DAPI/phalloidin staining, cells were washed once with PBS and fixed with 3% paraformaldehyde for 10 min. Coverslips were incubated with 0.5% Triton X-100 for 5 min and rinsed again twice with PBS. Staining of cells with 0.5 µg/ml DAPI (ICN) for 10 min and with 1.2 µg/ml phalloidin (Sigma) for 30 min was followed by two washes with PBS. Glass coverslips were mounted with Mowiol (Calbiochem) and slides were observed under a Leica epifluorescence microscope.
Fluorescence in situ hybridization
For FISH analysis, cells were washed twice with PBS, fixed with 3% paraformaldehyde for 10 min and rinsed twice with PBS. The FISH procedure was performed immediately after. A fluorescein isothiocyanate isomer 1 (FITC)-conjugated peptide nucleic acid probe obtained from DAKO Corp. (DAKO Telomere FISH kit/FITC) was used. FISH was performed according to the instructions provided by the supplier. After pre-treatment with 3.7% formaldehyde and a solution containing proteinase K for 10 min, the DNA was denatured at 80°C for 4 min in the presence of the FITC-conjugated DNA probe. The 1 h hybridization at room temperature was followed by two washes, one with a rinsing solution for 1 min and another with a washing solution at 65°C for 5 min, and fixation with cold 96% ethanol solution. Afterwards, coverslips were stained with 0.25 µg/ml DAPI (ICN) and 0.5 µg/ml propidium iodide (Sigma) for 1 min and mounted with antifade reagent (Vectashield).
Slide scoring
All slides were coded prior to scoring. Scoring was carried out using a Leica epifluorescence microscope at 1000x magnification under oil immersion. Criteria for cell and MN scoring were as described by Fenech (2000). Briefly, the cells should be binucleated with an intact nuclear membrane and should be located within the same cytoplasmic boundary. The two nuclei should be approximately equal in size, staining pattern and intensity. MN are morphologically identical to, but smaller than, nuclei; their diameter usually varies between one-sixth and one-third of the mean diameter of the main nuclei. MN can be readily distinguished and are not linked or connected to the main nuclei. MN usually have the same staining intensity as the main nuclei. For each slide (two slides per treatment) 1000 BN were scored (when possible). The frequencies of BN with MN (MNBN) and of apoptotic, mitotic and PN were determined. Cytotoxicity was measured by the binucleated cell ratio between control and treated slides, the control being considered as presenting no cytotoxicity. Slides with cytotoxicity exceeding 50% were not taken into account.
In FISH experiments, 100 MNBN were scored (when possible) and MN were examined for the presence (CEN+) or absence (CEN-) of fluorescent signals. The positive staining of MN was considered as an indicator of the presence of a whole chromosome.
Statistical analysis
For each treatment, data were analyzed using the Mann–Whitney U-test (SYSTAT 9 for Windows) to determine a statistically significant difference between treated cells and the negative control. For FISH analysis, the data were analyzed using Pearson’s 
2 test to determine a significant difference between treated cells and the clastogen control (MMS).
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Results |
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Effects of OA in the CBMN assay in CHO-K1 cells
Table I
summarizes the micronucleus data after 4 h exposure to different OA concentrations. In the absence of metabolic activation, OA concentrations from 1 to 50 nM did not induce MN formation or cytotoxicity. The positive control MMS (30 µg/ml) induced the formation of 10.7% MNBN cells.
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If treatment was prolonged up to 24 h, a statistically significant increase in the frequency of MNBN was obtained for 20 to 50 nM OA (12.9 and 33.4%, respectively) (Table II
). The percentage of PN was also statistically increased from 0.27% in the control to 10.3% at 20 nM and 21.6% at 50 nM. Cytotoxicity increased along with OA concentration: 2.8% at 5 nM, 29.2% at 20 nM and 49.39% at 30 nM. The 50 nM data were not taken into account because cytotoxicity exceeded 50%. For all OA concentrations, the number of apoptotic cells remained very low and insignificant. The clastogenic chemical MMS (30 µg/ml) induced a potent increase in MNBN frequency (77.3%). With the aneugenic compound GF at 10 µg/ml, 13.13% of BN were micronucleated and 13.56% of cells were polynucleated.
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In the presence of metabolic activation, OA statistically increased the frequency of MN in BN cells at 30 and 50 nM, by 4.3 and 9.8%, respectively, after only 4 h exposure (Table III
). It also significantly induced the formation of PN at 30 (1.01%) and 50 nM (1.72%) compared with the control (0.44%). At the same time, cytotoxicity increased from 13.9% at 30 nM to 57.3% at 50 nM. CP (25 µg/ml), used as a positive control, induced a significant increase in DNA damage: 9.69% MNBN, with only a slight cytotoxic effect (26.6%). The data suggest that OA metabolites were formed, that they induced MN formation and that they were more toxic than the parent compound.
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Aneugenic potential of OA detected by the CBMN assay coupled with FISH
To determine the nature of the MN induced by OA, we carried out a CBMN test in combination with FISH using a centromeric DNA probe. The results are summarized in Table IV
. OA induced a statistically significant increase in the frequency of MN containing a centromeric signal, compared with the clastogen MMS. Frequencies were 54 and 77.0% for 20 µg/ml MMS and 30 nM OA, respectively. The results obtained with OA were similar to that obtained with GF, a well-known aneugen, inducing 77.2% of CEN+ MN at 10 µg/ml.
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Discussion |
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The CBMN assay has become one of the most commonly used methods for assessing chromosome breakage and loss in different cell types (Fenech and Morley, 1985
). The present results indicate a background level of MNBN in CHO cells between 1.5 and 3%, these data being comparable to those obtained by other authors (Wei et al., 1997; Erexson et al., 2001). The clastogen MMS showed a large induction of MN formation (78%) at 30 µg/ml after 24 h treatment, which was higher than those obtained previously with the same cell line (Eastmond and Tucker, 1989) and with other cell types (Elhajouji et al., 1995; Matsushima et al., 1999). After exposure to GF (10 µg/ml), a well-known aneugen, the frequency of MNBN cells (13.1%) confirmed the data obtained with primary human liver fibroblasts (Nesti et al., 2000) and human lymphocytes (Migliore and Nieri, 1991; Parry et al., 2000). CP (25 µg/ml) gave positive results only in the presence of S9 mix for 4 h, with 9.69% MNBN cells. These positive results were lower than those reported by Erexson et al. (2001) in CHO cells, but confirmed the data of Matsushima et al. (1999) in the CHL cell line.
We have shown that treatment with OA for 4 h without metabolic activation did not induce MN formation or cytotoxicity. However, a prolonged treatment for 24 h significantly increased the frequency of MNBN cells: about 5- to 10-fold at 20 and 30 nM, respectively. As the CBMN assay alone could not discriminate between an aneugenic and a clastogenic effect, it was coupled with FISH using a centromeric DNA probe enabling distinction of MN containing a whole chromosome (Becker et al., 1990; Miller et al., 1991). The (TTAGGG)n DNA probe selected recognized the centromeric regions of the CHO chromosomes except those of chromosomes 1 and 2 (Meyne et al., 1990; Balajee et al., 1994; Bolzan et al., 2001). The CBMN assay coupled to FISH showed that the percentage of CEN+ MN increased in a dose-dependent fashion from 68.9% at 20 nM through 77% at 30 nM to 81% at 50 nM OA. These frequencies are similar to those obtained with the aneugen GF and are comparable to previously published results with other aneugens (Eastmond and Tucker, 1989; Elhajouji et al., 1995). FISH analysis showed that the clastogen MMS induced 54% CEN+ MN at 20 µg/ml. This frequency is higher than that obtained by Eastmond and Tucker (1989) and Elhajouji et al. (1995). The relatively high frequency of CEN+ MN after MMS exposure could be explained by some signals corresponding to acentric fragments rather than a whole chromosome. It could be concluded that OA is an aneugenic compound. It is the first time that the aneugenic potential of OA has been demonstrated and the first time for a DSP toxin.
In the literature some evidence suggests that OA could act as an aneugen. The majority of aneugens are negative in the Ames Salmonella mutation assay (Aardema et al., 1998). This is the case for OA, which did not induce mutations, with or without metabolic activation, either in the Ames test (Aonuma et al., 1991) or in the CHO/HGPRT assay (Fessard et al., 2002). However, aneugens generally induce polyploidy in vitro (Matsuoka et al., 1999; Kirsch-Volders et al., 2002) and we observed that OA increased the number of PN after 24 h exposure in a dose-dependent manner. Moreover, an increase in karyotype instability was previously shown in cells treated with OA (Afshari et al., 1993; Kuwabara et al., 1995).
Aneuploidy occurs when replicated chromosomes fail to accurately segregate between the two daughter cells. The end result is the production of cells with an abnormal number of chromosomes. Aneugens could act on different cell targets, but disturbance of the mitotic spindle [kinetochores, centrosomes, microtubules and the anaphase promoting complex (APC)] is most often reported. Indeed, OA has been found to disturb the mitotic spindle in many cell types (Picard et al., 1989; Ghosh et al., 1992; Vandre and Wills, 1992; Larsen and Wolniak, 1993; Chaudhuri et al., 1997), to induce a multipolar spindle and to affect centrosome replication (Van Dolah and Ramsdell, 1992; Vandre and Wills, 1992; Meraldi and Nigg, 2001). All studies suggest that the inhibition of PP1 and PP2A by OA alters centrosome replication inducing aberrant spindle formation and leading to PN. Protein phosphorylation plays a central role in controlling many aspects of cell division and has been implicated in the regulation of microtubule dynamics (Johnston et al., 1994; Howell et al., 1997). For example, Drosophilia mutants for the catalytic subunit of PP2A died during embryogenesis around the time of cellularization (Snaith et al., 1996). These embryos possessed multiple centrosomes with disorganized microtubules and the authors suggested that PP2A was required for the attachment of microtubules to chromosomal DNA at the kinetochore.
Op18/stathmin is a major cytosolic phosphoprotein that plays an important role in the regulation of microtubule dynamics during cell cycle progression. When stathmin is inactivated by phosphorylation, it promotes tubulin polymerization and allows formation of the mitotic spindle (Larsson et al., 1997). In the late stage of mitosis stathmin is dephosphorylated, inducing depolymerization of the mitotic spindle, which allows the cells to complete cell division and enter a new G1 phase. Tournebize et al. (1997) have indicated that OA inhibits Op18/stathmin activity, resulting in constitutive stabilization of microtubules, abnormalities in the organization of the mitotic spindle and difficulty in completing mitosis. Mistry et al. (1998) noted that OA increased the level of phosphorylated stathmin in K562 cells.
The initiation of anaphase and exit from mitosis depend on activation of the APC. Two forms, APCCdc20 and APCCdh1, mediate the degradation of critical cell cycle regulators (for a review see Nasmyth, 2002). OA inhibited sister chromatid separation in HeLa cells (Ghosh and Paweletz, 1992; Chaudhuri et al., 1997) and in LLC-PK cells (Vandre and Wills, 1992). In Sf9 cells, Cdh1 was hyperphosphorylated after OA treatment (Kramer et al., 2000); this form was much less active in stimulating APC activity (Bembenek and Yu, 2001). Therefore, by inhibiting PP1 and PP2A, OA would modify the phosphorylation state of the APC or its cofactors, disturbing the initiation of anaphase and deregulating the segregation of chromosomes.
We conclude that OA is an aneugen even if direct DNA damage has been reported by some authors (Rogers et al., 1994; Fessard et al., 1996).
We have also shown that, after 4 h exposure, OA induced an increase in MNBN cell frequency only in the presence of metabolic activation by a liver post-mitochondrial fraction. These original results strongly suggest that OA could be biotransformed in vitro to metabolites that are more cytotoxic and genotoxic than the parent compound. Therefore, the potential metabolic activation of OA in vivo cannot be excluded. Further studies on the identification of OA metabolites and the enzymes involved in their formation are needed.
In conclusion, the CBMN assay in combination with FISH showed that OA induced chromosome loss during cell division. This aneugenic effect of OA could be induced by the inhibition of PP1 and PP2A, modifying the general phosphorylation status of components implicated in chromosome segregation, such as microtubules, the APC and cofactors (such as MAD 2, Cdh1 and Cdc20), centrosomes and cdks.
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Acknowledgments |
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We thank Dr Michel Laurentie for his help with the statistical analysis and Dr Michael Holzhauser-Alberti who reviewed the English editing.
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Notes |
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4 To whom correspondence should be addressed. Tel: +33 2 99 94 78 78; Fax: +33 2 99 94 78 80; Email: v.fessard{at}fougeres.afssa.fr
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Received on October 17, 2002; revised on February 3, 2003; accepted on February 10, 2003.
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