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Journal of Nucleic Acids
Volume 2010 (2010), Article ID 456487, 9 pages
Research Article

Aflatoxin B1-Associated DNA Adducts Stall S Phase and Stimulate Rad51 foci in Saccharomyces cerevisiae

1Ordway Research Institute, Center for Medical Sciences, 150 New Scotland Avenue, Albany, NY 12209, USA
2Department of Biomedical Sciences, State University of New York at Albany, 150 New Scotland Avenue, Albany, NY 12209, USA
3Albany Medical College, 47 New Scotland Avenue, Albany, NY 12208, USA
4Bloomberg School of Public Health, Johns Hopkins University, 615 North Wolfe Street, Baltimore, MD 21205, USA

Received 15 August 2010; Accepted 9 September 2010

Academic Editor: Ashis Basu

Copyright © 2010 Michael Fasullo et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


AFB1 is a potent recombinagen in budding yeast. AFB1 exposure induces RAD51 expression and triggers Rad53 activation in yeast cells that express human CYP1A2. It was unknown, however, when and if Rad51 foci appear. Herein, we show that Rad53 activation correlates with cell-cycle delay in yeast and the subsequent formation of Rad51 foci. In contrast to cells exposed to X-rays, in which Rad51 foci appear exclusively in G2 cells, Rad51 foci in AFB1-exposed cells can appear as soon as cells enter S phase. Although rad51 and rad4 mutants are mildly sensitive to AFB1, chronic exposure of the NER deficient rad4 cells to AFB1 leads to increased lag times, while rad4 rad51 double mutants exhibit synergistic sensitivity and do not grow when exposed to 50 μM AFB1. We suggest RAD51 functions to facilitate DNA replication after replication fork stalling or collapse in AFB1-exposed cells.

1. Introduction

Hepatocellular carcinoma (HCC) ranks fifth in worldwide cancer mortality (for review, see [1]) and sixth in the United States [2]. High-risk factors for HCC include exposure to genotoxins, such as the mycotoxin aflatoxin B1 (AFB1), and infection with hepatitis B and C viruses [3]. Exposure to AFB1 is endemic in particular areas of China and sub-Saharan Africa due to Aspergillus flavus (mold) contamination of food and water [3]. A current hypothesis is that regeneration of liver cells following chronic liver injury renders liver cells susceptible to AFB1-associated carcinogenesis [4].

HCC pathogenesis is correlated with the accumulation of mutations and chromosomal rearrangements leading to either an inactivation of tumor suppressor genes or activation of oncogenes (for review, see [5]). MicroRNA-221 (MiR-221) overexpression contributes to liver tumorigenesis [6] and correlates with downregulation of cyclin dependent kinase inhibitors p21 and p57 [7]; however, there is no known correlation with AFB1 exposure. The p53(Ser)249 substitution mutation frequently occurs in liver cancer, where AFB1 exposure is highest [810]; however, there are conflicting reports whether the p53 249 codon is a direct hot spot for AFB1-associated mutagenesis [11]. Gross chromosomal translocations and gene amplifications have also been observed [12], and 10%–20% of HCCs contain cyclin D amplifications [13]. Although HCC associated with AFB1 exposure exhibits more genetic instability compared to HCC in nonendemic regions [14], it is unclear which types of genetic instability are directly caused by AFB1-associated DNA damage.

AFB1 is not genotoxic per se but requires metabolic activation. In humans, AFB1 metabolic activation in the liver is catalyzed by CYP1A2 and CYP3A4 [15] to form the highly unstable AFB1-8,9-exo-epoxide, which reacts primarily with the N7 position of guanine, present in the major groove of DNA [16]. The resulting adduct, 8,9-dihydro-8-(N7-guanyl)-9-hydroxyaflatoxin B1 (AFB1-N7-Guanine) is unstable and converts to either formamidopyrimidine (FAPY) derivatives or an apurinic site [16], both potentially mutagenic [17]. Both the FAPY derivatives and AFB1-N7-Guanine adducts are repaired by the nucleotide excision repair (NER) genes [18, 19]. The FAPY adducts also hinder DNA replication [20], which could lead to chromosomal breaks and require DNA repair genes that function in double-strand break and X-ray repair (XRCC). Thus, repair of AFB1-associated DNA damage may require both NER and XRCC genes.

Interestingly, a subset of known polymorphisms [21] in both NER gene XPD and the X-ray repair gene XRCC3 correlate with higher incidence in liver cancer in endemic areas of AFB1 exposure [22, 23]. Defective NER could lead to an increase in DNA adducts, while XRCC3 polymorphisms could confer defective repair of double-strand breaks (for review, see [24]). However, the polymorphism in XRCC3, Thr241Met, which is correlated with higher levels of AFB1-associated HCC [23], has not been correlated with a defect in double-strand break repair [25], suggesting that other functions in DNA damage or repair may be defective in cells containing this allele. Considering that recombinational repair may also participate in DNA damage-tolerance pathways, it is important to elucidate whether there are competing DNA repair pathways for AFB1-associated adducts.

Saccharomyces cerevisiae (budding yeast) is useful in elucidating the genetics of DNA repair of AFB1-DNA adducts. Yeast strains that express human CYP1A1 or CYP1A2 cDNAs on multicopy 2 μ plasmids can measure the genotoxicity of metabolically active carcinogens [2630]. Interestingly, AFB1 increases recombination frequencies more than mutation frequencies in cells expressing these cDNAs [26, 29].

The genotoxicity of AFB1 in yeast [2629] correlates with the transcription of DNA repair genes involved in recombination, including RAD51 [27, 30] and RAD54 [30]. RAD51 induction has been observed in log phase cells exposed to AFB1 [30], and RAD51 overexpression partially suppresses recombination defects in the mec1 null checkpoint mutant [27]. RAD51 is also required for AFB1-associated sister chromatid exchange (SCE) [29]. These results indicate that increased expression of RAD51 likely stimulates recombination when cells are exposed to AFB1-associated DNA adducts.

In log-phase yeast cultures, AFB1 is a mutagen [28]. Microarray analysis reveals not only a strong induction of RAD51 and RAD54 but also a downregulation of gene transcripts encoding histones [30]. rad51 mutants exhibit an increase in AFB1 -associated mutations [28]. Both AFB1-associated mutations and recombination events require checkpoint genes [28, 29]. Considering that AFB1 exposure triggers both checkpoint activation and S phase delay, it seems possible that the genotoxic responses to AFB1 result from stalled replication forks.

In this paper, we show that Rad53 activation correlates with S phase delay and subsequent Rad51 foci formation in yeast. A RAD51 deletion in NER defective strains results in high levels of lethality. We suggest that Rad51 foci formation may occur in S phase and be associated with error-free bypass of DNA lesions.

2. Materials and Methods

2.1. Strains and Media

Standard media were used for the culture of yeast cells. YPD (yeast extract, peptone, and dextrose), SC-TRP (synthetic complete lacking tryptophan), and SC-URA (synthetic complete lacking uracil) and FOA (5-fluro-orotic acid) were described in Burke et al. [31]. The genotypes of yeast strains used in this study are listed in Table 1. rad4, rad51 and rad4 rad51 strains for measuring SCE and AFB1 sensitivity are derived from YB163, which contains his3 recombination substrates in tandem at TRP1 [32]. Ura derivatives of rad4, and rad4 rad51 strains were selected on FOA medium. LSY1957 was a gift of Fung et al. [33]. pRS424-CYP1A2 was constructed by inserting the SacI CYP1A2 fragment from pCS316 into pRS424 and introduced into yeast strains by selecting for Trp+ transformants.

Table 1: Yeast strains.
2.2. Measuring DNA AFB1-Associated Recombination and Rad51 foci

To measure AFB1-associated genotoxic events, log-phase yeast cells ( ) were centrifuged and concentrated five-fold in selective media (SC-URA or SC-TRP). Cells were exposed to 50 μM AFB1, previously dissolved in DMSO. To synchronize cells in G1, log-phase cells were exposed to 10-4 M alpha factor (Sigma Co.) for two hours, and G1 cells were visualized in the light microscope. Cells were maintained in selective media (SC-URA or SC-TRP) during the carcinogen exposure and then washed twice in H2O. To measure SCE frequencies, recombinants were selected on SC-HIS, and viability was determined by plating an appropriate dilution on YPD. To visualize Rad51 foci formation, cells were resuspended in selective media (SC-TRP or SC-URA) and immobilized on glass slides.

To determine whether ionizing radiation stimulates the formation of Rad51 foci, cells were washed once in H2O, resuspended in 10 ml of H2O, and placed in a 81 mm diameter Petri dish. Cells were irradiated at 6 krad using a Nordion 1.8 kCi 137Cs irradiator (6 krad/hr). After irradiation cells were concentrated in YPD medium and immobilized on glass slides.

2.3. Live Cell Epifluorecence and Microscopy Analysis

Cells for microscopic analysis were grown to early-mid-log phase overnight in synthetic medium. After exposure to the genotoxin, cells were harvested by centrifugation, washed twice, and resuspended in YPD. Immobilization of cells was performed by mixing equal volumes of cell suspension and 1.4% low-melt agarose. Cover slips were sealed with a wax mixture as described by Lisby et al. [35]. Slides were visualized using a Zeiss LSM 510 META confocal microscope.

2.4. FACS Analyses

Cells were visualized by the fluorescent activated cell sorter (FACS) to directly correlate their DNA content with their cell-cycle phases. After AFB1 exposure, cells were washed, resuspended in 0.1 M sodium citrate, and treated with 1 mg/ml RNase A at 50°C for 1 hr. The cells were incubated at 50°C for 5 hr in 8 μg/ml proteinase K. An equal volume of 25 μg/ml propidium iodide (PI) diluted in 0.1 M sodium citrate was then added to the cells prior to the FACS analysis, to a final concentration of 12.5 μg/ml PI, and analyzed for fluorescence content with the use of a BD LSR II Flow cytometer.

2.5. Detection and Quantification of DNA Adducts

To measure the AFB1-associated DNA adducts in yeast, we used liquid chromatography-electrospray ionization tandem mass spectrometry (LC-ESI/MS/MS) [36]. Log-phase cultures of yeast-expressing human CYP1A2 (pCS316) were exposed to 50 μM AFB1 for 4 h. Because standard protocols for isolating yeast DNA involve alkaline buffers, rendering the highly unstable AFB1-N7-Guanine DNA adducts labile, we have modified the “smash-and-grab” protocol [37] so that we are using a neutral buffer containing 10 mM Tris-HCl, 1 mM EDTA, 100 mM NaCl, 2% Triton X-100, and 1% SDS, pH 7. DNA was isolated from two independent samples of yeast cells. The DNA adducts were identified and measured by high-performance liquid chromatography and tandem mass spectroscopy (LC-ESI/MS/MS) after acid hydrolysis [36].

2.6. Determining Rad53 Activation

Activation of Rad53 was determined by Western blots. Cells were inoculated in SC-URA medium. Log-phase cells ( ) were concentrated three fold in SC-URA and exposed to 50 μM AFB1 for 4 h. After washing cells twice in H2O, aliquots were plated directly on SC-HIS to measure recombination, appropriately diluted and plated on YPD to measure viability. Protein extracts were prepared as previously described by Foiani et al. [38], separated on 10% acrylamide/0.266% bis-acrylamide gels for Rad53 detection and transferred to nitrocellulose membranes. Rad53 was detected by Western blotting using goat anti-Rad53 (yC-19, Santa Cruz, Biotechnology, Santa Cruz, CA). The secondary antibody was antigoat IgG-HRP (Santa Cruz).

3. Results

3.1. Delay in Cell-Cycle Progression Correlated with Rad53 Activation

We previously observed that log-phase cells exposed to AFB1 exhibit Rad53 activation [29], which can result from replication blockage or delay. We measured Rad53 activation and cell-cycle delay in log-phase cells after continuous exposure to 50 μM AFB1. The data show that there is a peak activation of Rad53 after 3 hrs exposure to AFB1. Three hrs of exposure was also sufficient time to observe SCE recombinants (Figure 1). After 4 hrs of AFB1 exposure, there was less Rad53 activation and cell-cycle progression continued. The transient delay in the cell cycle is consistent with a previous study that indicated that AFB1-exposed yeast exhibit a transient S phase delay [30]. The data suggest that there is a correlation between AFB1-associated Rad53 activation and S phase delay. Because AFB1 adducts are detected in cycling cells after the S phase delay [29], we speculate that cells actively tolerate AFB1-associated DNA lesions during DNA replication.

Figure 1: Rad53 checkpoint activation, cell cycle progression, and recombination after log-phase cells were exposed to 50 μM AFB1. At indicated times, cells were collected for FACS analysis to measure frequencies of SCE and to make extracts to measure Rad53 activations. (a) Rad53 checkpoint activation was monitored after AFB1 exposure at the indicated times. Rad53 and the activated checkpoint protein, Rad53p, are indicated by arrows. (b) FACS analysis was performed at indicated time periods after exposure. The G1 and G2 cells are indicated by P4 and P5. The peak to the right of the G2 peak indicates bloated cells due to enlarged vacuoles. (c) AFB1-associated SCE were measured by selecting for His+ recombinants that result from unequal recombination between two truncated his3 fragments. Net recombination frequencies = recombination frequency after AFB1 exposure—spontaneous recombination frequency.

Rad53 not only delays cell-cycle progression but is also required for the phosphorylation of the Rad51 paralogs, Rad55, and Rad57, which may facilitate replication restart at stalled replication forks [39]. Rad55 and Rad57 facilitate the formation of DNA repair foci at double-strand breaks [40]. Previous data indicate that RAD53 is required for DNA damage associated SCE [41], including AFB1-associated SCE [29]. We, therefore, determined whether AFB1 exposure also stimulates Rad51 foci formation in yeast.

3.2. Exposure to AFB1 Results in Rad51 foci Formation in Cells that Are Entering S Phase

To visualize Rad51 foci that result from AFB1-associated DNA damage, we introduced pRS424-CYP1A2 into strain LSY1957 [33] to metabolically activate AFB1. This strain (YB405) contains rad51-I345T, an allele of RAD51, which when tagged with YFP, is still capable of conferring radiation resistance [33]. As a positive control, cells were exposed to either X rays (2 krad) or gamma rays (6 krads). After exposure, cells were returned to growth medium (YPD), and live cells were imaged for Rad51 foci. After 2 hrs of growth in YPD, most irradiated cells arrested in G2 and exhibited the dumb-bell shape (Figure 2). Cells were then visualized with the confocal microscope. Nearly 90% of the G2-arrested cells contain Rad51 foci, in agreement with Lisby et al. [40].

Figure 2: Confocal imaging of Rad51-YFP foci in live AFB1-exposed yeast cells. (a) Control, no AFB1 exposure, (b) Rad51-foci after exposure to X-rays, (c) Rad51 foci in logarithmically growing cells exposed to 50 μM AFB1, wide view, and (d) Rad51 focus in a synchronized cell after alpha factor arrest and exposure to AFB1.

Similarly, we determined whether Rad51 foci appear in cells after exposure to 50 μM AFB1 for 3 hrs. To first confirm the presence of AFB1 adducts, we extracted DNA from LSY197 cells after AFB1 exposure and observed approximately the same number of DNA adducts (Table 2) as previously observed in strains used to measure recombination [29]. After 3 hr of AFB1 exposure, we also observed that nearly 90% of the cells exhibited Rad51 foci. However, the difference with irradiated cells was that many AFB1-exposed cells that exhibited Rad51 foci were not G2 arrested. In addition, many cells exhibited Rad51 foci in both mother and daughter bud (Figure 2). These data indicate that AFB1-associated Rad51 foci are not restricted to the G2 phase of the cell cycle. Because daughter buds are not always visible in the confocal microscope, we decided to synchronize cells in G1, expose the cells to AFB1 and then determine when Rad51 foci could be detected.

Table 2: AFB1-N7 Guanine adducts in wild type and the rad4 mutant.

To determine whether cells can express Rad51 foci in S phase, cells were first arrested in G1 with alpha factor, and then exposed to AFB1 for three hours. Cells were then washed and returned to growth medium without AFB1. We observed that newly budded cells (90%) contain Rad51 foci. Rad51 foci were not evident after 30 minutes or 1 hr incubation time, but were evident after 1.5 hrs of incubation; after three hours of incubation, there were few Rad51 foci. These data indicate that Rad51 foci can be observed in cycling cells that are entering S phase.

3.3. rad4 Cells Are Defective in the Excision of AFB1 DNA Adducts

NER and recombinational repair mutants are modestly sensitive to AFB1 [27, 28]. Both rad4 and rad51 mutants exhibit higher rates of AFB1-associated mutagenesis [29, 30]. Measurements of DNA adducts indicated there are about three-fold higher levels of AFB1-N7-Guanine adducts in rad4, compared to wild type (Table 2). Consistent with the notion that AFB1 adducts persist longer in rad4 mutants, we observed by FACS analysis a delayed S phase after exposure to 10 μM AFB1. We asked whether wild type, rad4, and rad51 mutants could tolerate DNA adducts.

We used a growth assay in microtiter dishes [42] to determine differences in growth curves of wild type (YB401), rad4 (YB403), rad51(YB402), and rad4 rad51 (YB404) strains during chronic exposure to AFB1. Approximately 105 cells were inoculated in 96 well plates and exposed to 0 μM, 25 μM and 50 μM of AFB1; the experiments were done in triplicate. Growth was measured by (Figure 3). The lag time [42] for wild type was ~3 hrs and similar to rad51. Whereas the rad4 mutants exhibited a longer lag period, ~13 hrs, the rad4 rad51 mutant exhibited little, if any, growth. The data suggest that RAD51 function is critical in conferring AFB1 resistance in the rad4 mutant. This result is consistent with previous results that rad14 rad51 cells are also synergistically more sensitive to AFB1 [30].

Figure 3: Growth of wild type (a), rad51 (b), rad4 (e), and rad4 rad51 (d) cells after exposure to AFB1. (Left) Growth of cells containing pCS316 and expressing CYP1A2 after chronic exposure to 0 μM (black), 25 μM (blue), and 50 μM (red) AFB1. The relevant genotype is given below the panel (see Table 1, for complete genotype). 105 log-phase cells were inoculated in each well, . is plotted against time (h). (Right) rad4 cells synchronized in G1 were exposed to 50 μM AFB1 for 3 hrs, washed, and inoculated on YPD. Cell growth was monitored from 0–18 hrs. Bars indicate the actual range of measurements.

To further investigate whether rad4 cells can progress through S phase in the presence of DNA adducts, we arrested rad4 cells in G1 with alpha factor and exposed cells to AFB1 for 3 hrs. Cells were then washed, diluted, and inoculated on YPD plates to visualize the growth of single colonies every three hours under the microscope. After 12 hrs, ~70% (46/67) of cells that were not exposed to AFB1 formed colonies. However, only ~16% (46/268) of cells exposed to AFB1 (10 μΜ or 50 μΜ) formed colonies. 60% ( ) of either wild type or rad4 cells that do form colonies retain the URA3-containing plasmid expressing CYP1A2, indicating that colony formation occurred in cells that could still metabolically activate AFB1. Many of the rad4 cells that do not form colonies were arrested in early S phase, as evident by the presence of small daughter buds (Figure 3(f)). These data indicate that a few NER-deficient cells can progress through the cell cycle in the presence of AFB1-associated DNA adducts.

4. Discussion

AFB1 is a very potent liver carcinogen. The metabolic activation of AFB1 generates AFB1-associated DNA adducts which both stimulate mutagenesis and recombination in a variety of organisms. Polymorphisms in both XPD and XRCC3 are correlated with greater HCC risk [22, 23], thus underscoring the need to elucidate the role of recombinational repair in AFB1 metabolism. In budding yeast, well-conserved RAD and checkpoint genes are required for AFB1-associated mutation and recombination [28, 29]. Higher frequencies of AFB1-associated mutations in rad51 mutants [28, 29] and synergistic increase in the AFB1 sensitivity of double mutants defective in both recombinational and NER suggest that there are redundant pathways in conferring resistance to AFB1. AFB1 is highly recombinogenic and induces RAD51 in yeast [27]; however, it was unclear when and if Rad51 foci form. The salient conclusions from this study are (1) cells are delayed in S phase after AFB1 exposure, and there is a correlation with Rad53 checkpoint activation and cell cycle delay; (2) Rad51 foci form after AFB1-exposed cells enter S phase; (3) rad4 mutants accumulate AFB1-N7-Guanine adducts, and many exposed cells arrest or are delayed in S phase. This is the first study to show that AFB1 exposure leads to Rad51 foci formation.

Previous studies indicated that RAD53 is required for both AFB1-associated recombination and mutagenesis in yeast [29]. The coincident timing of recombination, Rad53 checkpoint activation, and S phase delay suggest that checkpoint activation is required to trigger AFB1-associated recombination in yeast. A possible link is that Rad53 is required for the DNA damage-associated phosphorylation of the Rad51 paralogs Rad55 and Rad57 [39], which participate in recombinational repair between sister chromatids [34]. We do not yet know how the checkpoint activation would trigger mutagenesis, but several studies suggest a role for checkpoint genes in DNA damage tolerance and translesion synthesis [4346].

The identification of AFB1-associated Rad51 foci was performed by detecting YFP fluorescence in the confocal microscope. Rad51 is not known to bind to specific DNA adducts, so we presume that the AFB1 N7-Guanine adducts and resulting FAPY and apurinic sites are further converted into recombinogenic lesions, including double-strand breaks or single-strand gaps. It is unlikely that all the lesions that initiate AFB1-associated Rad51 foci are the same as for X-ray-associated Rad51 foci, since we observed Rad51 foci in newly cycling cells entering S phase whereas we only observed X-ray-initiated Rad51-foci in G2-arrested cells. However, double-strand breaks or single-strand gaps could also result after replication forks stall or collapse in S phase, and AFB1 lesions have been reported to stall or hinder DNA replication in Escherichia coli [17]. Thus, an attractive model is that Rad51 foci form as AFB1-exposed cells transition through S phase.

We suggest that RAD51 confers AFB1 resistance in NER defective mutants by two possible functions. First, RAD51 would function in repairing double-strand breaks that indirectly results from AFB1-associated DNA damage. Considering that one double-strand break could confer lethality [47], one would estimate that at least one break occurs in every rad4 rad51 double mutant cell during chronic exposure to 50 μM AFB1. Second, RAD51 could actively participate in tolerating AFB1-associated DNA lesions; previous studies have indicated that the RAD52 pathway is involved in tolerating UV-associated damage [48]. Growth curves of wild-type cells indicate that some AFB1-associated DNA adducts can be tolerated without affecting doubling time. This second possibility is supported by observations that rad14 rad51 mutants exhibit extremely high frequencies of AFB1-associated mutagenesis [28]. Further speculation would suggest that the Rad51-paralog, XRCC3, has a similar function.

5. Conclusions

AFB1-associated DNA adducts stimulate both checkpoint activation and Rad51 focus formation. The timing of the Rad51 foci during the cell cycle in early S phase suggests a different mechanism of foci formation, compared to that of ionizing radiation. Understanding the function of these Rad51 foci will elucidate how polymorphisms in XRCC3 correlate with higher rates of liver cancer in endemic areas exposed to AFB1. It will thus be interesting if Rad51 foci also occur in mammalian cells after AFB1 exposure.


This work was supported by Grants nos. ES015954 and PO1ES006052 from the National Institutes of Health. The authors are thankful to Gary Schools for his expertise on the confocal microscope, Autumn Smith and Cinzia Cera for assistance with the microplate reader, and Cinzia Cera for critically reading this paper. We thank Chang-Uk Lim for assistance with the flow cytometer.


  1. S. A. Hussain, D. R. Ferry, G. El-Gazzaz et al., “Hepatocellular carcinoma,” Annals of Oncology, vol. 12, no. 2, pp. 161–172, 2001. View at Publisher · View at Google Scholar · View at Scopus
  2. A. Jemal, R. Siegael, J. Xu, and E. Ward, “Cancer statistics,” CA A Cancer Journal for Clinicians, vol. 60, no. 5, pp. 277–300, 2010. View at Google Scholar
  3. K. A. McGlynn and W. T. London, “Epidemiology and natural history of hepatocellular carcinoma,” Best Practice and Research, vol. 19, no. 1, pp. 3–23, 2005. View at Publisher · View at Google Scholar · View at Scopus
  4. M. C. Kew, “Synergistic interaction between aflatoxin B1 and hepatitis B virus in hepatocarcinogenesis,” Liver International, vol. 23, no. 6, pp. 405–409, 2003. View at Google Scholar · View at Scopus
  5. D. Moradpour and H. E. Blum, “Pathogenesis of hepatocellular carcinoma,” European Journal of Gastroenterology and Hepatology, vol. 17, no. 5, pp. 477–483, 2005. View at Publisher · View at Google Scholar · View at Scopus
  6. P. Pineau, S. Volinia, K. McJunkin et al., “miR-221 overexpression contributes to liver tumorigenesis,” Proceedings of the National Academy of Sciences of the United States of America, vol. 107, no. 1, pp. 264–269, 2010. View at Google Scholar
  7. F. Fornari, L. Gramantieri, M. Ferracin et al., “MiR-221 controls CDKN1C/p57 and CDKN1B/p27 expression in human hepatocellular carcinoma,” Oncogene, vol. 27, no. 43, pp. 5651–5661, 2008. View at Publisher · View at Google Scholar · View at Scopus
  8. I. C. Hsu, R. A. Metcalf, T. Sun, J. A. Welsh, N. J. Wang, and C. C. Harris, “Mutational hotspot in the p53 gene in human hepatocellular carcinomas,” Nature, vol. 350, no. 6317, pp. 427–428, 1991. View at Publisher · View at Google Scholar · View at Scopus
  9. H.-M. Shen and C.-N. Ong, “Mutations of the p53 tumor suppressor gene and ras oncogenes in aflatoxin hepatocarcinogenesis,” Mutation Research, vol. 366, no. 1, pp. 23–44, 1996. View at Publisher · View at Google Scholar · View at Scopus
  10. G. N. Wogan, “Aflatoxin as a human carcinogen,” Hepatology, vol. 30, no. 2, pp. 573–575, 1999. View at Publisher · View at Google Scholar · View at Scopus
  11. V. Paget, F. Sichel, D. Garon, and M. Lechevrel, “Aflatoxin B1-induced TP53 mutational pattern in normal human cells using the FASAY (Functional Analysis of Separated Alleles in Yeast),” Mutation Research, vol. 656, no. 1-2, pp. 55–61, 2008. View at Publisher · View at Google Scholar · View at Scopus
  12. D. B. Zimonjic, M. E. Durkin, C. L. Keck-Waggoner, S.-W. Park, S. S. Thorgeirsson, and N. C. Popescu, “SMAD5 gene expression, rearrangements, copy number, and amplification at fragile site FRA5C in human hepatocellular carcinoma,” Neoplasia, vol. 5, no. 5, pp. 390–396, 2003. View at Google Scholar · View at Scopus
  13. E. Pang, N. Wong, P. B.-S. Lai, K.-F. To, W.-Y. Lau, and P. J. Johnson, “Consistent chromosome 10 rearrangements in four newly established human hepatocellular carcinoma cell lines,” Genes Chromosomes and Cancer, vol. 33, no. 2, pp. 150–159, 2002. View at Publisher · View at Google Scholar · View at Scopus
  14. P. Pineau, A. Marchio, C. Battiston et al., “Chromosome instability in human hepatocellular carcinoma depends on p53 status and aflatoxin exposure,” Mutation Research, vol. 653, no. 1-2, pp. 6–13, 2008. View at Publisher · View at Google Scholar · View at Scopus
  15. D. L. Eaton and E. P. Gallagher, “Mechanisms of aflatoxin carcinogenesis,” Annual Review of Pharmacology and Toxicology, vol. 34, pp. 135–172, 1994. View at Google Scholar · View at Scopus
  16. C. N. Martin and R. C. Garner, “Aflatoxin B oxide generated by chemical or enzymic oxidation of aflatoxin B1 causes guanine substitution in nucleic acids,” Nature, vol. 267, no. 5614, pp. 863–865, 1977. View at Google Scholar · View at Scopus
  17. M. E. Smela, M. L. Hamm, P. T. Henderson, C. M. Harris, T. M. Harris, and J. M. Essigmann, “The aflatoxin B1 formamidopyrimidine adduct plays a major role in causing the types of mutations observed in human hepatocellular carcinoma,” Proceedings of the National Academy of Sciences of the United States of America, vol. 99, no. 10, pp. 6655–6660, 2002. View at Publisher · View at Google Scholar · View at Scopus
  18. Y. O. Alekseyev, M. L. Hamm, and J. M. Essigmann, “Aflatoxin B1 formamidopyrimidine adducts are preferentially repaired by the nucleotide excision repair pathway in vivo,” Carcinogenesis, vol. 25, no. 6, pp. 1045–1051, 2004. View at Publisher · View at Google Scholar · View at Scopus
  19. L. L. Bedard and T. E. Massey, “Aflatoxin B1-induced DNA damage and its repair,” Cancer Letters, vol. 241, no. 2, pp. 174–183, 2006. View at Publisher · View at Google Scholar · View at Scopus
  20. K. L. Brown, M. W. Voehler, S. M. Magee, C. M. Harris, T. M. Harris, and M. P. Stone, “Structural perturbations induced by the α-anomer of the aflatoxin B1 formamidopyrimidine adduct in duplex and single-strand DNA,” Journal of the American Chemical Society, vol. 131, no. 44, pp. 16096–16107, 2009. View at Publisher · View at Google Scholar · View at Scopus
  21. M. Manuguerra, F. Saletta, M. R. Karagas et al., “XRCC3 and XPD/ERCC2 single nucleotide polymorphisms and the risk of cancer: a HuGE review,” American Journal of Epidemiology, vol. 164, no. 4, pp. 297–302, 2006. View at Publisher · View at Google Scholar · View at Scopus
  22. X. D. Long, Y. Ma, D. Y. Qu et al., “The polymorphism of XRCC3 codon 241 and AFB1-related hepatocellular carcinoma in Guangxi population, China,” Annals of Epidemiology, vol. 18, no. 7, pp. 572–578, 2008. View at Publisher · View at Google Scholar · View at Scopus
  23. X. D. Long, Y. Ma, Y. F. Zhou et al., “XPD codon 312 and 751 polymorphisms, and AFB1 exposure, and hepatocellular carcinoma risk,” BMC Cancer, vol. 17, no. 9, article no. 400, 2009. View at Publisher · View at Google Scholar · View at Scopus
  24. R. D. Johnson and M. Jasin, “Double-strand-break-induced homologous recombination in mammalian cells,” Biochemical Society Transactions, vol. 29, no. 2, pp. 196–201, 2001. View at Publisher · View at Google Scholar · View at Scopus
  25. F. D. Araujo, A. J. Pierce, J. M. Stark, and M. Jasin, “Variant XRCC3 implicated in cancer is functional in homology-directed repair of double-strand breaks,” Oncogene, vol. 21, no. 26, pp. 4176–4180, 2002. View at Publisher · View at Google Scholar · View at Scopus
  26. C. Sengstag, B. Weibel, and M. Fasullo, “Genotoxicity of aflatoxin B1: evidence for a recombination-mediated mechanism in Saccharomyces cerevisiae,” Cancer Research, vol. 56, no. 23, pp. 5457–5465, 1996. View at Google Scholar · View at Scopus
  27. M. U. Keller-Seitz, U. Certa, C. Sengstag, F. E. Würgler, M. Sun, and M. Fasullo, “Transcriptional response of yeast to aflatoxin B1: recombinational repair involving RAD51 and RAD1,” Molecular Biology of the Cell, vol. 15, no. 9, pp. 4321–4336, 2004. View at Publisher · View at Google Scholar · View at Scopus
  28. Y. Guo, L. L. Breeden, H. Zarbl, B. D. Preston, and D. L. Eaton, “Expression of a human cytochrome P450 in yeast permits analysis of pathways for response to and repair of aflatoxin-induced DNA damage,” Molecular and Cellular Biology, vol. 25, no. 14, pp. 5823–5833, 2005. View at Publisher · View at Google Scholar · View at Scopus
  29. M. Fasullo, M. Sun, and P. Egner, “Stimulation of sister chromatid exchanges and mutation by aflatoxin B1-DNA adducts in Saccharomyces cerevisiae requires MEC1 (ATR), RAD53, and DUN1,” Molecular Carcinogenesis, vol. 47, no. 8, pp. 608–615, 2008. View at Publisher · View at Google Scholar · View at Scopus
  30. Y. Guo, L. L. Breeden, W. Fan, L. P. Zhao, D. L. Eaton, and H. Zarbl, “Analysis of cellular responses to aflatoxin B1 in yeast expressing human cytochrome P450 1A2 using cDNA microarrays,” Mutation Research, vol. 593, no. 1-2, pp. 121–142, 2006. View at Publisher · View at Google Scholar · View at Scopus
  31. D. Burke, D. Dawson, and T. Stearns, A Cold Spring Harbor Laboratory Course Manual, Methods in Yeast Genetics, Cold Spring Harbor Press, New York, NY, USA, 2000.
  32. M. T. Fasullo and R. W. Davis, “Recombinational substrates designed to study recombination between unique and repetitive sequences in vivo,” Proceedings of the National Academy of Sciences of the United States of America, vol. 84, no. 17, pp. 6215–6219, 1987. View at Google Scholar · View at Scopus
  33. C. W. Fung, A. M. Mozlin, and L. S. Symington, “Suppression of the double-strand-break-repair defect of the Saccharomyces cerevisiae rad57 mutant,” Genetics, vol. 181, no. 4, pp. 1195–1206, 2009. View at Publisher · View at Google Scholar · View at Scopus
  34. Z. Dong and M. Fasullo, “Multiple recombination pathways for sister chromatid exchange in Saccharomyces cerevisiae: role of RAD1 and the RAD52 epistasis group genes,” Nucleic Acids Research, vol. 31, no. 10, pp. 2576–2585, 2003. View at Publisher · View at Google Scholar · View at Scopus
  35. M. Lisby, U. H. Mortensen, and R. Rothstein, “Colocalization of multiple DNA double-strand breaks at a single Rad52 repair center,” Nature Cell Biology, vol. 5, no. 6, pp. 572–577, 2003. View at Publisher · View at Google Scholar · View at Scopus
  36. P. A. Egner, J. D. Groopman, J.-S. Wang, T. W. Kensler, and M. D. Friesen, “Quantification of aflatoxin-B1-N7-guanine in human urine by high-performance liquid chromatography and isotope dilution tandem mass spectrometry,” Chemical Research in Toxicology, vol. 19, no. 9, pp. 1191–1195, 2006. View at Publisher · View at Google Scholar · View at Scopus
  37. C. S. Hoffman and F. Winston, “A ten-minute DNA preparation from yeast efficiently releases automomous plasmids for transformation of Escherichia coli,” Gene, vol. 57, no. 2-3, pp. 267–272, 1987. View at Google Scholar · View at Scopus
  38. M. Foiani, F. Marini, D. Gamba, G. Lucchini, and P. Plevani, “The B subunit of the DNA polymerase α-primase complex in Saccharomyces cerevisiae executes an essential function at the initial stage of DNA replication,” Molecular and Cellular Biology, vol. 14, no. 2, pp. 923–933, 1994. View at Google Scholar · View at Scopus
  39. K. Herzberg, V. I. Bashkirov, M. Rolfsmeier et al., “Phosphorylation of Rad55 on serines 2, 8, and 14 is required for efficient homologous recombination in the recovery of stalled replication forks,” Molecular and Cellular Biology, vol. 26, no. 22, pp. 8396–8409, 2006. View at Publisher · View at Google Scholar · View at Scopus
  40. M. Lisby, J. H. Barlow, R. C. Burgess, and R. Rothstein, “Choreography of the DNA damage response: spatiotemporal relationships among checkpoint and repair proteins,” Cell, vol. 118, no. 6, pp. 699–713, 2004. View at Publisher · View at Google Scholar · View at Scopus
  41. M. Fasullo, Z. Dong, M. Sun, and L. Zeng, “Saccharomyces cerevisiae RAD53 (CHK2) but not CHK1 is required for double-strand break-initiated SCE and DNA damage-associated SCE after exposure to X rays and chemical agents,” DNA Repair, vol. 4, no. 11, pp. 1240–1251, 2005. View at Publisher · View at Google Scholar · View at Scopus
  42. M. Toussaint and A. Conconi, “High-throughput and sensitive assay to measure yeast cell growth: a bench protocol for testing genotoxic agents,” Nature Protocols, vol. 1, no. 4, pp. 1922–1928, 2006. View at Publisher · View at Google Scholar · View at Scopus
  43. H. Neecke, G. Lucchini, and M. P. Longhese, “Cell cycle progression in the presence of irreparable DNA damage is controlled by a Mec1- and Rad53-dependent checkpoint in budding yeast,” EMBO Journal, vol. 18, no. 16, pp. 4485–4497, 1999. View at Publisher · View at Google Scholar
  44. S. Sabbioneda, I. Bortolomai, M. Giannattasio, P. Plevani, and M. Muzi-Falconi, “Yeast Rev1 is cell cycle regulated, phosphorylated in response to DNA damage and its binding to chromosomes is dependent upon MEC1,” DNA Repair, vol. 6, no. 1, pp. 121–127, 2007. View at Publisher · View at Google Scholar · View at Scopus
  45. V. Pagès, S. R. Santa Maria, L. Prakash, and S. Prakash, “Role of DNA damage-induced replication checkpoint in promoting lesion bypass by translesion synthesis in yeast,” Genes and Development, vol. 23, no. 12, pp. 1438–1449, 2009. View at Publisher · View at Google Scholar · View at Scopus
  46. F. Conde, D. Ontoso, I. Acosta, A. Gallego-Sánchez,, A. Bueno, and P. A. San-Segundo, “Regulation of tolerance to DNA alkylating damage by Dot1 and Rad53 in Saccharomyces cerevisiae,” DNA Repair, vol. 9, no. 10, pp. 1038–1049, 2010. View at Google Scholar
  47. C. B. Bennett, A. L. Lewis, K. K. Baldwin, and M. A. Resnick, “Lethality induced by a single site-specific double-strand break in a dispensable yeast plasmid,” Proceedings of the National Academy of Sciences of the United States of America, vol. 90, no. 12, pp. 5613–5617, 1993. View at Google Scholar · View at Scopus
  48. V. Gangavarapu, S. Prakash, and L. Prakash, “Requirement of RAD52 group genes for postreplication repair of UV-damaged DNA in Saccharomyces cerevisiae,” Molecular and Cellular Biology, vol. 27, no. 21, pp. 7758–7764, 2007. View at Publisher · View at Google Scholar · View at Scopus