Mutagenesis, Vol. 15, No. 1, 91-97,
January 2000
© 2000 UK Environmental Mutagen Society/Oxford University Press
Aflatoxin B1-induced mitotic recombination in L5178Y mouse lymphoma cells
Department of Oto-Rhinolaryngology and 1 Department of Toxicology, University of Würzburg, Versbacher Straße 9, 97078 Würzburg, Germany and 2 National Institutes of Health, Research Triangle Park, NC 27709, USA
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Abstract |
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Aflatoxin B1 is a human hepatocarcinogen. It is also a known point mutagen in bacteria and mammalian cells. This mutagenic activity may be at least partly responsible for its carcinogenic activity. However, recent studies show that aflatoxin B1 induces mitotic recombination in the yeast Saccharomyces cerevisiae. Because numerous reports have implicated mitotic recombination in mechanisms leading to carcinogenesis and because no one has shown that aflatoxin B1 induces recombination in mammalian cells, we decided to examine the ability of aflatoxin B1 to induce recombination in a mammalian cell line. We used a combination of methods, analysis for loss of heterozygosity and whole chromosome in situ hybridization, to identify mechanisms of chromosome mutation, including mitotic recombination in the mammalian L5178Y mouse lymphoma cell system. Our experiments revealed that mitotic recombination caused ~60% or more of the aflatoxin B1-induced mutagenic lesions in this cell system. Thus, mitotic recombination plays an important role in aflatoxin B1-induced mutagenesis in mammalian cells and possibly in chemically induced mutagenesis and carcinogenesis. This work suggests that multiple genetic lesions may be involved in aflatoxin B1-induced pathology.
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Introduction |
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Aflatoxins are mycotoxins produced by a group of common fungal molds including Aspergillus parasiticus and Aspergillus flavus. These molds are ubiquitous in areas of the world with hot, humid climates. They are found in animal feed and contaminate human dietary staples in these climates (Sargeant et al., 1961
). Since countries in colder climates import food supplies from these regions, the potential toxic effects of aflatoxins are of concern world wide.
Aflatoxin B1 is a human hepatocarcinogen. It is also a liver carcinogen when fed to certain rodent species (Wogan and Newberne, 1967; Wogan et al., 1973; IARC, 1993).
A case has been made that the mechanism leading to aflatoxin B1-induced cancer involves DNA adducts which are the precursors of guanosine to thymidine (G
T) transversions which are frequently observed in human liver (Garner et al., 1988; Hsieh et al., 1988; Lee et al., 1989; Zhang et al., 1991; Aguilar et al., 1993; Eaton and Gallagher, 1994; Riley et al., 1997). The following facts support this hypothesis: the urine of people exposed to aflatoxins contains aflatoxin B1–guanine adducts (Groopman et al., 1992; Wild et al., 1992) and most tumors from aflatoxin B1-contaminated regions harbor a GT transversion at codon 249 in the p53 gene (Bressac et al., 1991; Hsu et al., 1991; Murakami et al., 1991; Scorsone, 1992 et al.; Li,D. et al., 1993). It has also been shown that aflatoxin B1 induces point mutations in c-ras oncogenes although the role of these oncogenes in hepatocellular carcinoma has not been documented (McMahon et al., 1986; Sinha et al., 1988; Bauer Hofmann et al., 1990; Soman and Wogan, 1993).
Sengstag et al. (1996) proposed that recombination might be involved in aflatoxin-induced carcinogenesis. Aflatoxin B1 strongly induced mitotic recombination, resulting in chromosome translocation and gene conversion events in a set of metabolically competent Saccharomyces cerevisiae yeast strains (Sengstag et al., 1996; Sengstag, 1997). In contrast, this compound was only weakly mutagenic in a yeast strain sensitive to gene mutations. They concluded that metabolically activated aflatoxin B1 is much more prone to inducing mitotic recombination than gene mutation in yeast.
Although aflatoxin B1-induced recombinogenic activity has been shown to occur in yeast, its role in mammalian cell mutagenesis is not known. We decided to use L5178Y mouse lymphoma cells to examine the role of recombination in aflatoxin B1-induced mutagenesis (Clive et al., 1972; Caspary et al., 1997; Liechty et al., 1998). These cells are heterozygous at the selectable thymidine kinase locus (tk) on chromosome 11 (Liechty et al., 1993). If the active tk gene is inactivated or lost, the cells become resistant to the selective agent trifluorothymidine (TFT). This cell line detects a wide variety of mutagenic lesions, including point mutations, deletions and various types of chromosomal aberrations (Hozier et al., 1992; Caspary et al., 1997; Liechty et al., 1998). The in situ protocol that we used captures essentially all the viable mutants, including the slowly growing mutants, for analysis. It also ensures the independence of the induced mutants (Rudd et al., 1990).
To examine the possible role of mitotic recombination in aflatoxin B1 mammalian cell mutagenesis, we treated L5178Y mouse lymphoma cells with aflatoxin B1 and selected for mutants. We chose a dose that had the largest relative mutation fraction with little or no toxicity. We then isolated 41 mutant colonies, analyzed them for loss of heterozygosity (LOH) on chromosome 11 and, using fluorescent in situ hybridization, examined their metaphase spreads for their morphological characteristics. Our conclusion that mitotic recombination plays a role in aflatoxin B1-induced mutation shifts the emphasis of the mutational specificity of aflatoxin B1 away from point mutation toward large scale chromosome-type alterations.
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Materials and methods |
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Cell culture
Mouse L5178Y cells, clone 3.7.2c (5), were cultured in suspension in RPMI-1640 supplemented with 95 U/ml penicillin, 95 µg/ml streptomycin, 0.25 mg/ml L-glutamine, 107 µg/ml sodium pyruvate and 10% heat-inactivated horse serum (Sigma Chemie GmbH, Deisenhofen, Germany). Cell cultures were grown in a humidified atmosphere with 5% CO2 in air at 37°C.
Mutation assay
We used the in situ procedure to obtain dose–response curves for mutation induction and for mutant isolation (Rudd et al., 1990; Spencer and Caspary, 1994; Spencer et al., 1994). Cultures of mouse L5178Y cells were treated with methotrexate before each experiment to kill pre-existing TFT-resistant (TFTr) cells. To accomplish this, cells were incubated for 24 h in culture medium plus methotrexate (0.3 µg/ml), thymidine (9 µg/ml), hypoxanthine (15 µg/ml) and glycine (22.5 µg/ml). The cells were then incubated for at least 24 h in the same medium without methotrexate. To measure chemically induced mutations using the in situ procedure, cultures containing 1 000 000 cells in 5 ml medium were treated. For metabolic activation, S9 mix (uninduced rat liver cell homogenate, supplemented with 1.5% citrate and 0.8% w/v NADPH) was added to the cells at a concentration of 100 µg protein/ml cell suspension.
Ethylmethanesulfonate (EMS) was used as a positive control without metabolic activation, and aminoacetylfluorene (AAF) was used (with S9 mix) as a positive control for metabolic activation. Two solvent controls (with and without S9 mix) containing ethanol (final concentration 1%) were used. We treated the cells with the chemicals for 3 h, then washed the cells twice with fresh medium. After that, 500 000 cells from each culture were added to 50 ml of semi-solid culture medium (containing 0.25% granulated agar; Baltimore Biological Laboratories), plated into two plastic 100 mm culture dishes and allowed to solidify at room temperature. TFTr cells were selected by adding an overlay of TFT to a final concentration of 8 µg/ml in 10 ml semi-solid medium after an expression time of 40 h. The cloning efficiency was determined by adding 600 cells to 100 ml of semi-solid medium and pouring into three plastic 100 mm dishes and allowed to solidify at room temperature. For picking mutants, two additional mutant plates were prepared as described above for 60 ng/ml aflatoxin B1. Mutants were picked from these two plates at day 7 after the start of the experiment under a microscope with a sterile pasteur pipette. Cells from each colony were dispersed into 1 ml of medium for culture and subcultured as required for a duration of 1–3 weeks. All other plates were incubated for a total of 9 (viability plates) or 12 days (mutant plates) at 37°C in 5% CO2 for colony growth and the number of surviving colonies (viability plates) and TFTr colonies (mutant plates) were counted using an automatic colony counter.
Preparation of DNA for LOH analysis
DNA from 10 000 000 cells was isolated using a Qiagen kit (Qiagen, Hilden, Germany). Briefly, cells were lysed (AL buffer; Qiagen) and treated with proteinase K for 10 min at 70°C. DNA was precipitated with ethanol and purified using spin columns by washing twice with AW buffer (Qiagen). Elution of DNA was performed with AE buffer (Qiagen) at 70°C. DNA concentration and purity were determined photometrically at 260/280 nm.
LOH analysis
PCR reactions (20 µl) were prepared by mixing 10 µl of 2x PCR Master (Boehringer, Mannheim, Germany) containing 20 mM Tris–HCl, 100 mM KCl, 3 mM MgCl2, 0.05 U/µl Taq DNA polymerase, 400 µM each dNTP and 0.01% Brij 35, pH 8.3, with 11.6 pmol of each primer, 3.5 µl of H2O and 3 µl (30 ng) of template DNA. Reactions were performed in a Biozym PTC 100 thermal cycler (Hess, Oldendorf, Germany) by `touchdown' PCR, in which the optimal annealing temperature was gradually approached from above to decrease the quantity of spurious PCR products resulting from some primer pairs. For primer Agl2, an initial 94°C denaturation for 2.5 min was followed by two cycles of denaturation at 94°C for 20 s, high temperature annealing at 72°C for 30 s and extension at 72°C for 20 s. The annealing temperature was decreased by 1°C for each additional two-cycle set until the final annealing temperature (66°C) was reached. Then 12 additional cycles were performed at 65°C, followed by extension at 72°C for 5 min. For all other primers, the procedure was essentially the same, except that the highest annealing temperature was 61°C, the lowest annealing temperature was 52°C and instead of the 12 cycles at 65°C, 20 cycles with an annealing temperature of 50°C were performed.
Gel electrophoresis
PCR products were analyzed on 10% non-denaturing polyacrylamide gels in 1x TBE buffer. Gel dimensions were 170 cm (w)x150 cm (h)x0.5 mm. Electrophoresis was performed for 4–6 h at 200 V and gels were stained with SybrGreen (15 µl/150 ml 1x TBE for 30 min) (Molecular Probes, OR) for visualization.
Preparation of metaphases
Cells from mutant colonies were subcultured daily for at least 5 days. Then colcemid (Gibco BRL, Eggenstein, Germany) was added at a concentration of 0.05 µg/ml. After 20 min the cell culture medium was replaced by hypotonic solution (0.075 M KCl at 37°C). After 15 min five drops of methanol/acetic acid (3:1) were added to the hypotonic solution to begin cell fixation. Then the cell suspension was centrifuged, carefully resuspended in methanol/acetic acid and incubated for 20 min at –20°C. This was repeated twice. Finally, cells were dropped onto glass slides and the slides were quickly air dried.
In situ hybridization
In situ hybridization was performed using a biotinylated mouse chromosome 11-specific paint (Caspary et al., 1997; Liechty et al., 1998). Slides were aged for 10–20 days. The painting probe was prepared by placing it in a thermocycler at 37°C for 5 min, 80°C for 5 min and 37°C for 2.5–3 h. The slides were denatured at 70°C for 75 s in formamide solution (70% in 2x SSC, pH 7.0). After dehydration in an ethanol series (70, 85, 90 and 100%, –20°C, 2 min each), slides were air dried. Next, slides were put on a slide warmer at 42°C and 10 µl of chromosome painting probe were added for overnight incubation in a humid chamber at 37°C. Slides were then washed for 3x5 min in formamide wash solution (55% in 2x SSC, pH 7.0) at 45°C, 3x5 min in 1x SSC at 45°C and then in 4x SSC at room temperature for at least 5 min. Detection and signal amplification were performed using materials and the protocol of Clontech Laboratories (Palo Alto, CA) for FITC detection of biotinylated DNA. Counterstaining of whole DNA was achieved with bisbenzimide 33258 (5 µg/ml, 2.5 min). In the final step, a second counterstaining of whole DNA was achieved by adding mounting medium containing 0.3 µg/ml propidium iodide. We limited the analysis to mutants that showed LOH for at least 25% of the total chromosome 11 length because mutants with smaller losses would be difficult to differentiate from mutants with normal chromosomes using FISH. Metaphases from each mutant colony were analyzed for the number of chromosome 11 homologs and for chromosome 11 centromere sizes. The centromere size was also analyzed using bizbenzimide stain, which shows the C band regions equivalent to the centromere. The lengths of chromosomes 11 were visualized under the microscope. We required 10 good metaphases showing consistent results before drawing conclusions.
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Results |
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Aflatoxin B1-induced mutations in the L5178Y mouse lymphoma mutation assay at the tk locus
The mutation fraction was dose dependent (Table I
). We prepared replicate plates for all the doses and chose a concentration of 60 ng/ml for mutant isolation because of its high relative mutation fraction and high cloning efficiency (no or little toxicity). The relative mutation fraction of 6.6 for this concentration implies that 84% of the mutant colonies are aflatoxin B1 induced (Table I).
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Loss of heterozygosity
To determine the types of lesions present in these mutants, we examined them for LOH. We isolated 41 aflatoxin B1-treated mutant clones. The in situ protocol that we used to obtain and isolate the mutants ensured that the induced mutant colonies were independent (Rudd et al., 1990
). We PCR amplified up to six heteromorphic microsatellite sites along chromosome 11 in each of these clones. These loci were from 2 (Mit2) to 78 cM (Agl2) from the centromere (Figure 1). The total length of chromosome 11 is ~80 cM. The most distal probe was Agl2, which resides at a heteromorphic site within the tk gene between exons 6 and 7 (Liechty et al., 1996b). Figure 2 shows an example of our results using the probe Mit59.
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We categorized the mutants as group 1, 2 or 3 mutants. Into group 1 we placed 10 of the 41 mutants (24%) showing no LOH. Group 2 comprised 26 mutants (63%) showing LOH at some of the loci. Group 3 was composed of five mutants (12%) showing LOH at all analyzed sites (Figure 2
and Table II). In all mutants showing LOH, the losses included Agl2, which resides within the tk gene.
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The LOH data suggests that all the group 2 and 3 mutants possessed chromosomal lesions. The 10 mutants from group 1 showed no LOH with any of the probes. These mutants were potential point mutants. However, some of them might have small chromosomal lesions encompassing sequences distal to Agl2 down to the distal end of the chromosome or sequences proximal to Agl2 and distal to Mit103.
In situ hybridization
A limitation of the LOH analysis is that it is not always possible to identify the mechanisms leading to LOH. Specifically, translocations and changes in chromosome number are not detectable and mitotic recombination is not distinguishable from deletion. For example, if LOH occurs at all the loci we examined, mitotic recombination that includes those loci is not distinguishable from chromosome loss. To overcome these shortcomings, we combined the results from LOH analysis with those from whole chromosome 11 in situ hybridization. This technique allows ready identification of chromosome 11 sequences in these cells (Liechty et al., 1996a; Caspary et al., 1997).
The sizes of the two centromeres from the homologous chromosomes are different. This difference has been used to distinguish the two homologous chromosomes (Hozier et al., 1982; Sawyer et al., 1985; Blazak et al., 1986). The homolog with the larger centromere contains the functional tk allele (Hozier et al., 1982). We used this difference to identify the origin of the centromeric regions of these two homologous chromosomes cytogenetically (e.g. bisbenzimide stain) and by FISH. All conclusions about centromere size were based on the examination of at least 10 good metaphases from each mutant.
We limited FISH analysis to mutants that showed LOH for at least 25% of the total chromosome 11 length, i.e. mutants that show loss from Agl2 (located at 81 cM) to Mit59 (located at 61 cM). Mutants with smaller losses would be difficult to differentiate from mutants with normal chromosomes using FISH. Fifteen mutants fulfilled this requirement: 10 from group 2.2 and all five from group 3 (see Figure 3 for an example).
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We did not analyze any of the group 1 mutants by in situ hybridization because these lesions were either point mutations or small intragenic or intergenic lesions. Although mutants in this class that were due to intragenic or intergenic events could result from small recombinations, they would look normal by in situ hybridization because of the small size of the losses.
All five mutants from group 3 (mutants 6, 18, 52, 56 and 57) were analyzed by in situ hybridization (Table III). Three of these mutants had two chromosomes 11 of apparently equal length and different centromere sizes. Since the two original centromeres were present, recombination was responsible for the TFT resistance expressed by these mutants. The two remaining mutants showed LOH at all tested sites. As shown in Table III, these mutants were monosomic for chromosome 11. Therefore, analysis of centromere size and chromosome lengths was not possible. Ten mutants in group 2 had damage large enough to have interpretable data after chromosome painting (Table III). Four of these (25, 13, 21 and 58) were trisomic for chromosome 11. Because not all the sites showed LOH, we suggest that recombination was the primary event causing the mutation. It appears that in all four cases, a secondary event was the duplication of chromosome 11 containing the tk– allele since the centromere size of two of the chromosomes was small.
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In five of these mutants (9, 11, 24, 26 and 22), two chromosomes 11 were present and had different centromere sizes. The chromosome 11 with the large centromere was elongated (~30–50% additional length). Since not all the sites showed LOH, recombination must have been the mechanism leading to these mutations. One mutant in this group was monosomic for chromosome 11. Therefore, analysis of centromere size and chromosome lengths for this mutant was not possible.
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Discussion |
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Interest in the expression of recessive genes is due to the importance of tumor suppressor genes in carcinogenesis (Müller and Scott, 1992
). Loss of the second allele of a tumor suppressor gene in a cell that has already sustained a mutation (often a point mutation) in the first allele is a critical step in carcinogenesis. Tumor suppressor genes require the loss of function of both of the homologous alleles to express their phenotypes and it is believed that the second step is often a recombination event (Cavenee et al., 1983).
A powerful approach to assessing the role of recombination in mammalian cells is the use of LOH analysis with fluorescent in situ hybridization. Recent reports demonstrated the successful use of these techniques in mouse lymphoma cells (Caspary et al., 1997; Liechty et al., 1998).
There are many reports showing that aflatoxin B1 causes point mutations (Swenson et al., 1977; Aguila et al., 1993; Gerbes and Caselmann, 1993; Eaton and Gallagher, 1994; Bailey et al., 1996; Choi et al., 1996). The analysis to be presented here shows that aflatoxin B1 also causes the expression of mutations mediated by recombinogenic events in mammalian cells. This extends previous reports that aflatoxin B1 causes recombination in yeast (Sengstag and Wurgler, 1994; Sengstag et al., 1996) and lends credence to the idea that aflatoxin B1-induced chromosomal mutations manifested by LOH may lead to hepatocarcinogenesis.
Our analysis shows that the majority of aflatoxin B1-treated mutants were recombinants. In this analysis we exclude the three monosomic mutants because the mechanism of their formation is not known. Exclusion of these three mutants will not qualitatively affect our conclusions. Our sampling of the mutants from groups 2 and 3 indicated that the 12 mutants from groups 2.2 and 3 resulted from recombination. We would expect that the remaining mutants from group 2 (those with LOH too small to be analyzed by in situ hybridization) were also due to recombination. This assumption is justified because the only difference between group 2.1 and 2.2 (Figure 2) mutants was the size of the LOH observed. There is no evidence that the mechanisms of formation are different. Of the 41 mutants isolated, it appears that at least 28 were due to recombination. This number could be higher if some of the 10 mutants from group 1 were not point mutations but small recombinogenic lesions.
To estimate the percentage of aflatoxin B1-induced mutants that were recombinants, we have to consider the effect of spontaneous mutants. The relative mutation fraction of 6.6 tells us that 16% of the 41 isolated mutants in this study may be of spontaneous origin. Previous studies have shown that ~70% of spontaneous mutants contain recombinations with the remainder harboring gene mutations or small partially intragenic deletions (Liechty et al., 1996a, 1998; Caspary et al., 1997). Thus, of the 41 mutants examined here, seven (16%) could have been of spontaneous origin, of which approximately five (70%) could have harbored recombinogenic events. Thus, of the 28 mutants in this study harboring recombinations, aflatoxin B1 induced at least 23. Since 34 mutants were induced (41–7), 68% (23 of 34) of the aflatoxin B1-induced mutants were due to recombination. Even if all seven spontaneous mutants in our investigation were recombinants, 21 of the recombinants were aflatoxin B1 induced and, therefore, 62% (21 of 34) of the induced mutants were due to recombination. Thus, the majority of aflatoxin B1-induced mutants were recombinants.
The high recombinogenic activity of aflatoxin B1 is in agreement with the findings of Sengstag and co-workers (Senstag and Wurgler, 1994; Sengstag et al., 1996; Sengstag, 1997) in yeast and those of Zhang et al. (1991), who used a mammalian system for intrachromosomal recombination. Removal of an inserted sequence by recombination reactivates the HPRT gene in a V79 Chinese hamster cell line (SP5) and is detectable using a reversion mutation assay. They observed statistically significant enhancements in the frequency of reversion in SP5 cells after treatment with aflatoxin B1.
These data extend the list of chemicals that are known to induce mitotic recombination in mammalian cells (Hozier et al., 1992; Li,C.Y. et al., 1992; Smith and Grosovsky, 1993; Zhu et al., 1993; Xia et al., 1994). It also shows that mitotic recombination plays an important role in aflatoxin B1-induced mutagenesis in mammalian cells and possibly in chemically induced mutagenesis and carcinogenesis.
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Acknowledgments |
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We thank Mrs H. Raabe for her expert technical assistance. We thank Professor Wolfgang Dekant and Dr Jeff French for their comments on this manuscript. This work was supported by a NATO CRG program grant (no. 940684) to W.J.C. and H.S. and by SFB 172 of the Deutsche Forschungsgemeinschaft to H.S.
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Notes |
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3 To whom correspondence should be addressed. Tel: +1 919 541 2150; Fax: +1 919 541 2242; Email: caspary{at}niehs.nih.gov
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Received on August 9, 1999; accepted on September 24, 1999.
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