Mutagenesis, Vol. 14, No. 6, 541-546,
November 1999
© 1999 UK Environmental Mutagen Society/Oxford University Press
Evaluation of micronuclei and chromosomal breakage in the 1cen–q12 region by the butadiene metabolites epoxybutene and diepoxybutane in cultured human lymphocytes
Environmental Toxicology Graduate Program, University of California, Riverside, CA 92521, USA
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1,3-Butadiene is a widely used industrial chemical and common environmental pollutant that has been associated with increased risks of leukemias and lymphomas. Butadiene and its metabolites, 1,2-epoxybutene (EB) and diepoxybutane (DEB), have been shown to be genotoxic in a wide variety of test organisms. The objective of this research was to evaluate techniques for the rapid detection of chromosomal alterations occurring in humans exposed to butadiene. We have used a multicolored fluorescence in situ hybridization (FISH) method and the CREST-modified micronucleus assay to detect chromosomal breakage induced by EB (10–300 µM) and DEB (0.5–10 µM) in cultured human lymphocytes. A significant dose-related increase in the formation of micronuclei was seen in lymphocytes treated with DEB at concentrations as low as 2.5 µM, but not with EB over the dose range tested. Over 80% of the micronuclei induced by DEB were CREST-negative, indicating their origin from chromosomal breakage. Multicolor FISH using two adjacent chromosome-specific probes showed a significant increase in chromosomal breakage in the 1cen–q12 region induced by DEB at concentrations as low as 2.5 µM, but not by EB. Since DEB is likely to be one of the metabolites contributing to the genotoxic effects of butadiene, the sensitivity of the tandem FISH approach to detect breakage induced by diepoxybutane indicates that this technique may be useful for monitoring chromosomal alterations in butadiene-exposed workers.
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Introduction |
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1,3-Butadiene (BD) is an industrial monomer used in the production of synthetic rubber and plastics (Csanady et al., 1992
; Boogaard et al., 1996; Boogaard and Bond, 1996). It is also a widely used industrial chemical that, with an annual production of ~1 700 000 tons in 1995, is ranked twentieth among synthetic organic chemicals produced in the USA (Chemical Week, 1996). BD has also been classified as one of the 189 hazardous air pollutants in the 1990 Clean Air Act Amendment (Environmental Protection Agency, 1990) and is emitted into the atmosphere at low levels from common sources, such as cigarette smoke and automobile exhaust (Melnick and Kohn, 1995; Spano et al., 1996). The most significant exposure to BD occurs in the industrial setting, where the National Institute for Occupational Safety and Health has estimated that 69 555 US workers are potentially exposed to this chemical (Recio et al., 1997; National Toxicology Program, 1998).
The finding that inhaled BD is carcinogenic at multiple sites in rats (Owen et al., 1987) and mice (Huff et al., 1985; Melnick et al., 1990) has raised concern about human health effects that may be associated with exposure to this chemical. However, its potential to induce cancer in humans remains controversial (Bond et al., 1995; Melnick and Kohn, 1995). Recent epidemiological studies of workers exposed to BD have reported an increase in lymphatic and hematopoietic cancers (Divine, 1990; Matanoski et al., 1990; Ward et al., 1995; Delzell et al., 1996). The International Agency for Research on Cancer (IARC) has classified BD as a probable human carcinogen (Group 2A) based on the results of chronic rodent bioassays, mechanistic studies and limited evidence of human carcinogenicity (IARC, 1992).
BD undergoes metabolic activation in vitro and in vivo to form a number of reactive and genotoxic metabolites, including 1,2-epoxybutene (EB) and diepoxybutane (DEB) (Himmelstein et al., 1997). The oxidation of BD to the primary metabolite EB is mediated by the cytochrome P450 isozymes CYP2E1 and, at high concentrations, CYP2A6. EB is subjected to further oxidation by CYP2E1 and CYP3A4 to form DEB (Seaton et al., 1995). These epoxide metabolites are potent alkylating agents that bind to nucleophilic sites on DNA and are likely to play a role in the carcinogenicity of this compound (IARC, 1986). EB is a monofunctional alkylating agent, while DEB is a bifunctional alkylating agent that has been shown to induce interstrand cross-links between adjacent guanine bases (Lawley and Brookes, 1963, 1967; IARC, 1992; Recio et al., 1997). Both metabolites have been observed to alkylate guanine at the N7 position (Lawley and Brookes, 1967; Citti et al., 1984). BD (with metabolic activation), EB and DEB have been shown to cause genotoxicity in a large number of test organisms (IARC, 1992). For example, BD was shown to be genotoxic to bone marrow cells of exposed mice, causing an increase in the frequency of chromosomal aberrations, sister chromatid exchanges and micronucleated erythrocytes (Tice et al., 1987; Melnick et al., 1990).
In addition to bioactivation by cytochrome P450, detoxification of EB and DEB by glutathione S-transferases (GST) and epoxide hydrolases (EH) appear to play an important role in modifying the genotoxicity of BD (Csanady et al., 1992; Boogaard and Bond, 1996; Boogaard et al., 1996). GSTs are a family of enzymes that show genetic polymorphisms which have been shown to be responsible for individual differences in metabolism that may profoundly modulate the effects of BD and other chemical carcinogens (Norppa, 1997). For example, previous studies have shown that after treatment with EB, lymphocytes of GSTM1- and GSTT1-null donors exhibited higher frequencies of sister chromatid exchanges (SCEs) than GSTT1-positive donors (Uuskula et al., 1995; Bernardini et al., 1998). Similar studies have shown that after treatment with DEB, lymphocytes of GSTT1-null donors were more sensitive to the induction of micronuclei (MN) and SCEs compared with the GSTT1-positive donors (Wiencke et al., 1991; Norppa et al., 1995; Vlachodimitropoulos et al., 1997).
The objective of our research is to develop molecular cytogenetic techniques for the rapid detection of chromosomal alterations occurring in humans exposed to BD. Specifically, our goal was to determine the feasibility of using a recently developed multicolor fluorescence in situ hybridization (FISH) method, as well as the CREST-modified MN assay, to detect chromosomal breakage induced by the BD metabolites EB and DEB in human cells. This FISH technique uses two adjacent or tandem DNA probes and allows a more confident detection of hyperploidy in interphase human cells as well as allowing chromosomal breakage affecting the targeted region to be detected. Chromosomes 1 and 9 have large characteristic blocks of centromeric and pericentric heterochromatin and have been found to be particularly sensitive to breakage induced by cross-linking agents (Brogger, 1977; Aurias, 1993).
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Materials and methods |
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Cell culture and chemical treatments
Peripheral blood was collected from healthy male volunteers in heparinized vacutainers. Lymphocytes were isolated using Leucoprep tubes (Gibco BRL, Grand Island, NY) and cultures were established at an initial cell density of 0.5x106 cells/ml using RPMI 1640 medium (Mediatech, Herndon, VA) supplemented with 5% heat-inactivated fetal calf serum (Irvine Scientific, Santa Ana, CA), 5% heat-inactivated iron-supplemented calf serum (Hyclone, Logan, UT), 2 mM L-glutamine, 100 IU/ml penicillin and 100 µg/ml streptomycin (all from Mediatech) and 2.36% phytohemagglutinin (type M; Gibco BRL) at 37°C in a 5% CO2 atmosphere. Twenty-four hours after culture initiation, the cells were exposed to EB (10–300 µM in DMSO; CAS-930-22-3; Aldrich, Milwaukee, WI) and DEB (0–20 µM; CAS-298-18-0; Aldrich). For 20 h following chemical addition, the lids were tightly sealed. Control cultures were treated with DMSO (0.1% v/v; Sigma Chemical Co., St Louis, MO). For the MN assay, cytochalasin B (final concentration 3 µg/ml) was added to the cultures 44 h after culture initiation and cells were harvested 28 h later by centrifuging aliquots of the cell suspension directly onto slides using a Shandon (Pittsburgh, PA) cytospin centrifuge (600 r.p.m., 5 min). Slides were briefly air dried, fixed in 100% methanol for 10 min at room temperature and stored desiccated in a N2 atmosphere at –20°C until further use. To investigate cultures for interphase/metaphase analysis, colcemid (Sigma) at a final concentration of 50 ng/ml was added at 47 h and harvested at 49 h following initiation of the culture. The cells were harvested using standard methods (Eastmond et al., 1994
). Briefly, the cells were treated with 0.075 M KCl hypotonic solution for 30 min, fixed three times with methanol:acetic acid (3:1) and dropped onto slides. Due to the low frequency of mitotic cells, analysis was restricted to interphase cells. The slides were stored at –20°C under a N2 atmosphere until use. Each experiment was repeated once, using blood from two male donors under the age of 32.
CREST antibody labeling
The staining procedure for the CREST antibody was performed as previously described (Eastmond and Tucker, 1989a,b).
Fluorescence in situ hybridization
Two different chromosome 1-specific DNA probes were used in the present study: a classical satellite probe specific for the pericentric heterochromatin of chromosome 1 (pUC1.77) (Cooke and Hindley, 1979) and an 
-satellite probe specific for a small centromeric region adjacent to the pericentric heterochromatin region (Wade et al., 1979). The classical satellite probe was labeled with Cy3-dUTP (Amersham Life Science, Arlington Heights, IL) by nick translation in our laboratory according to the protocol of the manufacturer (Bethesda Research Laboratories, Gaithersburg, MD) and the digoxigenin-labeled -satellite probe was purchased from Oncor (Gaithersburg, MD).
Previously described methods were used to perform the multicolor FISH experiments (Trask and Pinkel, 1990; Rupa et al., 1995). Briefly, cells were denatured in 70% formamide/2x SSC and hybridized overnight at 37°C with the denatured hybridization cocktail consisting of 1 µl Cy3-labeled classical satellite probe (20–100 ng), 0.5 µl digoxgenin-labeled -satellite probe (5–20 ng), 0.5 µl ddH2O, 1 µl sonicated herring sperm DNA (1 mg/ml) and 7 µl MM 2.1 hybridization mix (to give a final concentration of 55% formamide/1x SSC/10% dextran sulfate). Post-hybridization washes were performed in 60% formamide/2x SSC, all at 44°C. The slides were then rinsed twice in PX buffer (0.1 M phosphate buffer, pH 8.0, containing 0.2% Triton-X-100) at room temperature. The digoxigenin-labeled -satellite probe was detected using a mouse anti-digoxigenin antibody [3.2 µg/ml in PX buffer with 5% non-fat dry milk supernatant (PXM); Boehringer-Mannheim, Indianapolis, IN] followed by a digoxigenin-conjugated sheep anti-mouse antibody (20 µg/ml in PXM; Boehringer-Mannheim) and FITC sheep anti-digoxigenin antibody (20 µg/ml; Boehringer Mannheim). 4',6-Diamidino-2-phenylindole (1 µg/ml in diphenylenediamine antifade) was used to counterstain the DNA.
Multiplex PCR method
A multiplex PCR method was used to detect the presence or absence of the GST 
1 (GSTT1; ~450 bp) and GST µ1 (GSTM1; 215 bp) genes in genomic DNA samples. Previously described methods were used with slight modifications (Chen et al., 1996). DNA was extracted from the blood of donors A and B using S&S Iscode stix PCR template preparation dipsticks (Schleicher & Schuell, Keene, NH). The modified method had combined GST primer sets in the same PCR, included a third primer set for ß-globin (268 bp) and used an annealing temperature of 61°C. PCR products from co-amplification of GSTT1, GSTM1 and ß-globin were resolved on a 0.5 µg/ml ethidium bromide-stained prepoured 10% TBE gel (Bio-Rad, Hercules, CA). ß-Globin was co-amplified as an internal positive control in the PCR reaction. The visual presence or absence of the 215 and ~450 bp bands was used to classify the donors as having the presence (+/+, two copies or +/0, one copy) or absence (0/0, deletion of both copies, `null') of the GSTM1 and the GSTT1 genes.
Scoring criteria
The scoring criteria for the MN assay were described earlier (Eastmond and Tucker, 1989a,b). The number of MN at each dose level was determined by scoring 500 binucleated cells for each of two separate experiments. The multicolor FISH slides were observed using a Nikon Optiphot II microscope equipped with a fluorescence attachment, a triple band-pass filter (Chroma Technology, Brattleboro, VT), a fluorescein filter (B2A; Nikon, Melville, NY) and a Texas Red filter (Chroma). For each test chemical or corresponding control, 1000 cells were scored from coded slides for aberrations involving the 1cen–q12 region based on hybridization patterns in each nucleus. The hybridization strategy using -satellite and classical satellite probe regions to detect breakage and hyperdiploidy has been illustrated previously (Eastmond et al., 1994; Rupa et al., 1995, 1997). Hybridization regions containing both the -satellite (green/yellow) and classical satellite (red) signals were scored as one intact copy of chromosome 1. A nucleus containing three or more intact copies of chromosome 1 was considered as a hyperdiploid cell. A nucleus containing only a red classical satellite signal in addition to two intact copies of chromosome 1 was scored as a break within the classical satellite region. A clear separation (generally more than the width of the probe region) between the -satellite and classical satellite regions was scored as a break between these two regions. Clumps of cells or cells with broken nuclei were eliminated from scoring. Whenever signals were weak, a fluorescein filter (Nikon B2A) or a Texas Red filter (Chroma) was used to verify the signal.
Statistical analysis
Two statistical tests were used to analyze the data. The Cochran–Armitage test for trend in binomial proportions was performed to determine if the frequency of micronucleated cells and of nuclei containing three or more hybridization regions exhibited a significant dose-related trend (Margolin and Risko, 1988). Following a positive response in the trend test, a one-tailed Fisher exact test was utilized to compare each treatment with the control. Critical values were determined using a 0.05 probability of type 1 error.
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Multiplex PCR method
To test whether the genotypes of donor A and B altered their sensitivity to EB and DEB, a multiplex PCR method was used to detect the presence or absence of the GSTT1 and the GSTM1 genes in genomic DNA. It was shown that both donors exhibited the presence of both the GSTT1 and GSTM1 genes. The results from the two assays for each donor were similar (Tables I and II
) and, consequently, were combined for presentation. It is likely that their sensitivities to EB and DEB are similar since each donor had at least one functional copy of both the GSTT1 and GSTM1 genes.
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Antikinetochore-modified MN assay
The induction of MN observed in the treated human lymphocytes of donors A and B is shown in Figures 1 and 2
. There was little or no increase in the formation of MN seen in the cultures treated with EB (Figure 1). In contrast, a significant dose-related increase in the formation of MN was seen in the lymphocytes treated with DEB (Figure 2; Cochran–Armitage binomial trend test, P < 0.05). In the DEB-treated cultures, significant increases in the induction of MN were seen at concentrations as low as 2.5 µM (one-tailed Fisher exact test, P < 0.05). The frequency of MN increased from 0.005 in the control cultures to 0.042 in cultures treated with 10 µM DEB. Over 80% of the MN were CREST-negative, indicating that the majority of the MN originated from chromosomal breakage rather than chromosome loss. Although a consistent dose-related increase in the formation of CREST-positive MN was not seen, at 10 µM DEB the induction of CREST-positive MN was significantly increased (one-tailed Fisher exact test, P < 0.05), indicating that at higher concentrations DEB may cause alterations in chromosome number.
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Multicolor FISH assay
The ability of multicolor FISH to detect structural aberrations in cultured interphase cells treated with EB and DEB was determined using tandem probes on chromosome 1. The frequencies of hyperdiploidy and breaks affecting the 1cen–q12 region observed in the interphase nuclei of human lymphocytes treated with EB and DEB are shown in Figures 3 and 4
. Similar to the findings in the MN assay, there was little or no increase in breakage affecting the 1cen–q12 seen in the cultures treated with EB. However, significant increases in heterochromatin breakage were seen at DEB concentrations as low as 2.5 µM (Figure 4; one-tailed Fisher exact test, P < 0.05). When comparing the induction of chromosomal breakage at 1 with 2.5 µM the difference is minimal. At 1 µM there appears to be an induction of chromosomal breakage, but the effect is of borderline significance (Figure 4; one-tailed Fisher exact test, P = 0.0572).
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Discussion |
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In this study we determined the ability of the antikinetochore antibody (CREST) and tandem-labeling FISH approaches to detect chromosome alterations in human lymphocytes treated with two BD metabolites. Using the CREST-modified MN assay, exposure of human lymphocytes to DEB, but not to EB, was shown to produce a significant dose-related induction of micronucleated cells. In addition, similar results were seen with the tandem DNA probes, indicating that multicolor FISH can be used to detect chromosomal breakage in the 1cen–q12 region in cultured interphase lymphocytes following in vitro exposure to DEB, but not to EB.
The induction of chromosomal breakage by DEB was expected and, in general, our results are consistent with previous reports in which DEB has been shown to be highly genotoxic and more effective than the EB metabolite (IARC, 1992). For example, studies by Sasiadek et al. (1991) on the in vitro induction of SCEs in human lymphocytes by EB and DEB reported that the lowest effective concentration for EB was 25 µM and for DEB 0.5 µM. Cochrane and Skopek (1994) also reported that DEB exhibited activity at concentrations ~100-fold lower than EB when the mutagenic potential of the metabolites were measured at the tk and hprt loci in TK6 human lymphoblastoid cells. Additional studies have shown that the concentrations of DEB that are mutagenic in vitro (1.2–2.6 µM) are similar to the levels of DEB measured in blood and tissues of mice (0.6–2.5 µM) exposed by inhalation to carcinogenic concentrations (62.5–1250 p.p.m.) of BD (Himmelstein et al., 1994, 1995; Seaton et al., 1995). For various cells and species, consistent positive results for the induction of MN by DEB have been reported, while mixed positive and negative results have been seen with EB (Sjoblom and Lahdetie, 1996; Xi et al., 1997; Saranko and Recio, 1998). In male rat spermatids in vitro exposure to DEB resulted in an increase in MN using the meiotic MN assay, but no increase was seen with exposure to EB (Sjoblom and Lahdetie, 1996). Saranko and Recio (1998) reported that the induction of MN by DEB revealed a concentration-dependent increase in MN in Rat2 cells following in vitro exposure to DEB. Recently Xi et al. (1997) reported a significant increase in MN by both EB and DEB in human lymphocytes. However, the effect of EB was weak, with the majority of the MN being kinetochore-positive rather than kinetochore-negative.
The differential response observed in the various genotoxicity tests, including the tandem FISH and MN assays, is likely related to the bifunctional alkylating ability of DEB compared with the monofunctional alkylating ability of EB. The resulting DEB-induced DNA intrastrand and interstrand cross-links are believed to inhibit DNA replication and result in large scale chromosome aberrations (Vogel and Natarajan, 1995). In contrast, the predominant effect of DNA monoadducts is the formation of small intragenic or point mutations (Vogel and Natarajan, 1995). Bifunctional alkylating agents have been shown to be 55- to 630-fold more effective in inducing SCEs and 300- to 2400-fold more effective in inducing cytotoxicity than similar monofunctional agents (Bodell, 1990). In addition, the low clastogenic effectiveness of the monofunctional agents such as EB has been proposed to be due to the efficient, error-free repair of DNA monoadducts by excision repair enzymes (Vogel and Natarajan, 1995).
It has been reported that there is a potential for large interindividual variation among humans in susceptibility to the potential genotoxic effects of BD (Norppa et al., 1995; Seaton et al., 1995; Norppa, 1997; Recio et al., 1997). GST is one of the enzymes involved in the metabolism of BD that may have a significant impact on individual susceptibility to the genotoxic effects of BD metabolites. In earlier studies, the sensitivity of human cells to EB and DEB has been shown to be dependent on GST genotype (Wiencke et al., 1991; Norppa et al., 1995; Norppa, 1997; Vlachodimitropoulos et al., 1997). To see if our data were influenced by the donors' genotypes, the genotypes of both donors were determined for GST and µ using a PCR-based method. Donors A and B were determined to have both the GSTT1 and GSTM1 genes present. Therefore, both donors had at least one copy of the GSTT1 and GSTM1 genes, indicating that they are not highly sensitive to the genotoxic effects of the BD metabolites EB and DEB. Although the donors may exhibit intermediate sensitivity, there was a significant induction of chromosomal breakage seen by DEB in both donors at relatively low concentrations in both assays. In particular, this study shows that FISH with centromeric and pericentromeric DNA probes is a sensitive method for detecting DEB-induced chromosomal breakage in the 1cen–q12 region. Since earlier studies have shown that sensitivity of individuals is influenced by their GST genotype (Kelsey et al., 1991; Wiencke et al., 1991), it would be useful to test these assays on individuals with other GST genotypes, particularly those with the more sensitive `null' genotype.
Recent studies suggest that EB and DEB are not only clastogenic, but also have weak aneugenic activity. Xiao et al. (1996) used centromeric DNA probes to analyze MN in mouse bone marrow and observed a weak, but significant, induction of aneuploidy and Xi et al. (1997) reported that EB and DEB cause chromosome-specific aneuploidy in human cells. In our experiments induction of aneuploidy was not seen with the multicolor FISH assay and only a weak increase was seen in the MN assay. These differences could be due to the weak aneuploidy-inducing effects of the metabolites, chromosome-specific effects or differences in donors, such as their GST genotype or treatment conditions. However, our experiments were optimized to detect chromosomal breakage and not hyperdiploidy. By harvesting the interphase cultures only 25 h after addition of the chemical, many of the cells may not have had adequate time to pass through mitosis and become aneuploid. In contrast, more recent experiments with lymphoblastoid cells show that in longer term cultures, modest, but significant, increases in hyperdiploidy can be detected in DEB-treated cultures (M.N.Murg and D.A.Eastmond, unpublished data).
In summary, these results indicate that the tandem FISH assay is capable of detecting chromosomal alterations following in vitro exposure of human lymphocytes to DEB. Tandem labeling of regions where breakage is likely to occur appears to be a powerful tool for detection of genetic alterations in human populations, particularly in cell types that have not previously been amenable to conventional cytogenetic analysis (Rupa et al., 1995). The sensitivity of these initial studies to detect breakage induced by DEB shows that tandem FISH may be a promising technique for monitoring chromosomal alterations in the interphase cells of BD-exposed workers. Currently additional studies are underway in our laboratory to characterize the alterations induced by DEB and to determine the persistence of these genetic alterations.
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Acknowledgments |
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We would like to thank Leslie Hasegawa and Robert Parks for their excellent technical assistance. The authors also wish to thank the UC Systemwide Toxic Substance Research & Teaching Program Air Toxics and Health Effects of Modern Technology Components for their support.
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1 To whom correspondence should be addressed. Tel: +1 909 787 4497; Fax: +1 909 787 3087; Email: david.eastmond{at}ucr.edu
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Received on January 25, 1999; accepted on June 22, 1999.
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