Carcinogenesis vol.19 no.8 pp.1451–1458, 1998
Identification of differentially expressed genes in aflatoxin B1-treated cultured primary rat hepatocytes and Fischer 344 rats
Angela J.Harris1, Joseph G.Shaddock, Mugimane G.Manjanatha, Jeffery A.Lisenbey and Daniel A.Casciano Division of Genetic and Reproductive Toxicology, National Center for Toxicological Research, Jefferson, AR 72079, USA 1To
whom correspondence should be addressed Email:
[email protected]
Aflatoxin B1 (AFB1), a mutagen and hepatocarcinogen in rats and humans, is a contaminant of the human food supply, particularly in parts of Africa and Asia. AFB1induced changes in gene expression may play a part in the development of the toxic, immunosuppressive and carcinogenic properties of this fungal metabolite. An understanding of the role of AFB1 in modulating gene regulation should provide insight regarding mechanisms of AFB1induced carcinogenesis. We used three PCR-based subtractive techniques to identify AFB1-responsive genes in cultured primary rat hepatocyte RNA: differential display PCR (DD-PCR), representational difference analysis (RDA) and suppression subtractive hybridization (SSH). Each of the three techniques identified AFB1-responsive genes, although no individual cDNA was isolated by more than one technique. Nine cDNAs isolated using DD-PCR, RDA or SSH were found to represent eight genes that are differentially expressed as a result of AFB1 exposure. Genes whose mRNA levels were increased in cultured primary rat hepatocytes after AFB1 treatment were corticosteroid binding globulin (CBG), cytochrome P450 4F1 (CYP4F1), alpha-2 microglobulin, C4b-binding protein (C4BP), serum amyloid A-2 and glutathione S-transferase Yb2 (GST). Transferrin and a small CYP3A-like cDNA had reduced mRNA levels after AFB1 exposure. Full-length CYP3A mRNA levels were increased. When liver RNA from AFB1treated male F344 rats was evaluated for transferrin, CBG, GST, CYP3A and CYP4F1 expression, a decrease in transferrin mRNA and an increase in CBG, GST, CYP3A and CYP4F1 mRNA levels was also seen. Analysis of the potential function of these genes in maintaining cellular homeostasis suggests that their differential expression could contribute to the toxicity associated with AFB1 exposure. Introduction Aflatoxin B1 (AFB1*), a potent mycotoxin of Aspergillus, contaminates many human and animal food sources. Exposure to AFB1 is prevalent in parts of Africa and Asia where the hot, humid conditions favor mold growth in stored grains. *Abbreviations: AFB1, aflatoxin B1; AFB1-dG, 8,9-dihydro-8-N-(N7-guanyl)9-hydroxy aflatoxin B1; C4BP, C4b-binding protein; CBG, corticosteroid binding globulin; CYP, cytochrome P450; DD-PCR, differential display PCR; DFO, deferoxamine; DMSO, dimethyl sulfoxide; GAPDH, glyceraldehyde 3phosphate dehydrogenase; GST, glutathione S-transferase; RDA, representational difference analysis; ROS, reactive oxygen species; SSH, suppression subtractive hybridization. © Oxford University Press
AFB1 is a hepatocarcinogen in rats (1) and humans (2), and may induce extrahepatic cancers as well (3,4). In addition, AFB1 causes aflatoxicosis, a toxic response to acute AFB1 poisoning that results in gastrointestinal bleeding and liver failure in humans (5,6). Aflatoxicosis also occurs in livestock from ingestion of AFB1-contaminated feed (7–10). The systemic toxicity of AFB1 is clearly demonstrated by the range of aflatoxicosis symptoms reported in animals, which include carcinogenesis, hepatic failure, pulmonary edema, immunosuppression and various hematological disorders. Metabolic conversion of AFB1 to aflatoxin 8,9-epoxide is required for carcinogenesis (11). This activation step is mediated by a number of P450 enzymes, including members of the CYP1A, 2B, 2C and 3A sub-families in rodents (12–14); and the CYP3A and 1A sub-families in humans (15,16). Lipooxygenases have also been shown to activate AFB1 in humans and guinea pigs (17,18). Aflatoxin 8,9-epoxide is able to form stable adducts with both DNA and protein. The major DNA adduct formed by covalent binding of the reactive 8,9-epoxide is 8,9-dihydro-8-N-(N7-guanyl)-9-hydroxy AFB1 (AFB1–dG) (19,20), and a number of studies have correlated carcinogenesis with increasing AFB1–dG adduct formation (reviewed in ref. 21). It has been proposed that a primary cause of AFB1mediated carcinogenesis is an accumulation of activating mutations in critical proto-oncogenes and/or mutations that inactivate tumor suppressor genes. Indeed a hot spot for G→T transversion at codon 249 of the human p53 gene has been identified in liver tumors from individuals who live in regions with high AFB1 levels in foods (22,23). AFB1-induced mutations have also been reported in codon 12 of the K-ras proto-oncogenes in rat liver tumors (24,25). The cytotoxicity of AFB1 is thought to play a role in the development of hepatocellular carcinoma. Using the nomenclature of the Solt–Farber model of chemical carcinogenesis (26), AFB1 appears to act as a complete carcinogen. AFB1 can promote carcinogenesis and this effect can be abrogated by compounds that protect against cytotoxicity (27). The appearance of hepatocellular foci resistant to AFB1 cytotoxicity may enhance the carcinogenicity of AFB1 (28). In order to identify additional mechanisms of AFB1-induced carcinogenesis and toxicity, we have isolated mRNAs that represent AFB1-responsive genes. In addition, we compared the utility of three different methods for the identification of differentially expressed genes: differential display PCR (DDPCR) (29), representational difference analysis (RDA) (30) and suppression subtractive hybridization (SSH) (31). Each of the techniques resulted in the identification of a different set of AFB1-responsive genes. We report here the identity of the cDNAs isolated and discuss possible roles these genes could play in AFB1-induced carcinogenesis and toxicity. Materials and methods Hepatocyte isolation and culture Hepatocyte primary cultures were established from the livers of 250–350 g adult male Fischer 344 rats via in situ collagenase perfusion (32,33). Following
1451
A.J.Harris et al. perfusion, the liver was removed to a beaker of cold Williams’ Medium E (WE; Life Technologies, Gaithersburg, MD) and single cells were isolated by mincing and stirring. The cellular mixture was filtered to remove stromal tissue and the resulting hepatocyte suspension was centrifuged to pellet. The cells were resuspended in cold medium. Monolayer cultures were established by plating 10–153106 viable hepatocytes in 20 ml of WE containing 17% fetal bovine serum (Life Technologies) in a Vitrogen-coated (Collagen Biomedical, Fremont, CA) 150 mm tissue culture dish. Following a 1–2 h attachment period, primary cultures were incubated in serum-free WE containing 1.0 µM AFB1 (Sigma, St Louis, MO) with dimethyl sulfoxide (DMSO, ø1%; Burdick and Jackson, Morristown, NJ) as the solvent, for 16 h at 37°C in a 5% CO2 humidified atmosphere. This dose was chosen because of its limited toxicity to primary hepatocytes. Control cultures were incubated with DMSO alone. AFB1 treatment of Fischer 344 rats Livers from AFB1-treated male Fischer 344 rats were a gift from Dr Ming Chou (Division of Biochemical Toxicology, NCTR). Six-week-old male Fischer 344 rats were fed AFB1 ad libitum for 4, 8 or 12 weeks on an intermittent 4-week schedule. Animals were fed 0.01, 0.04, 0.40 or 1.6 p.p.m. doses. The 4-week treatment group was treated with AFB1 for 4 weeks then killed. The 8-week treatment group was fed AFB1 ad libitum for 4 weeks, then was off AFB1 for 4 weeks before being killed. The 12-week treatment group was fed AFB1 ad libitum for 4 weeks, then were off AFB1 for 4 weeks, then on AFB1 for 4 weeks before being killed. Isolated rat livers were snap frozen in liquid nitrogen and then stored at –80°C until RNA extraction. Total and poly(A)1 RNA isolation Hepatocytes were lysed in TRIzol reagent (Life Technologies) in the culture dish. Total RNA was isolated from the hepatocyte lysate according to the manufacturer’s protocol. Livers from control or AFB1-treated Fischer 344 rats were homogenized in TRIzol and total RNA was isolated. Poly(A)1 RNA was purified from total RNA using the Mini Oligo (dT) Cellulose Spin Column Kit (5 Prime→3 Prime, Boulder, CO). DD-PCR Total RNA was treated with 1.0 U RQ1 DNase/µg RNA (Promega, Madison, WI) for 30 min at 37°C. DD-PCR was performed using a modification of the original protocol, which utilizes the Stoffel fragment of Taq DNA polymerase (29,34). Total RNA from control and AFB1-treated rat hepatocytes was used in 30 DD-PCR reactions using the dT12GT 39 primer and 30 different arbitrary 59 primers (arbitrary 10-mer primer kits GEN3-50, GEN4-50, GEN6-50; Genosys, The Woodlands, TX). Total RNA (500 ng) was reverse transcribed in the presence of 2.5 µM oligo dT12GT 39 primer (Cruachem, Dulles, VA) using Superscript™ II RNase H– RT (Life Technologies) and reaction conditions recommended by the manufacturer. The reaction was incubated at 37°C for 60 min, then 95°C for 5 min to heat-inactivate the reverse transcriptase. One-tenth of the cDNA reaction was added to a 20 µl PCR reaction that contained 2.5 µM dT12GT 39 primer, 2.5 µM 59 arbitrary primer, 25 µM dNTPs and 1 µCi of [α-35S]dATP (.1000 Ci/mmol; Amersham, Arlington Heights, IL). The PCR reaction was incubated at 98°C for 1 min, cooled to 72°C, then 2.5 U Stoffel fragment (Applied Biosystems/Perkin Elmer, Foster City, CA) was added under the mineral oil. The cycling parameters were 40°C for 2 min, 72°C for 30 s for one cycle; followed by 94°C for 30 s, 40°C for 2 min, and 72°C for 30 s for 40 cycles. DD-PCR fragments were resolved on a 6% denaturing polyacrylamide gel. The gel was then transferred to Whatman 3MM paper (Whatman, Maidstone), dried and exposed to film for 24–48 h. Bands representing cDNAs, which appeared to be differentially expressed, were excised from the dried gel and reamplified as described previously (29). Primers and unincorporated dNTPs were removed from the PCR reaction using a TE Midi column (5 Prime→3 Prime). The PCR product was ethanol precipitated then resuspended in 16 µl dH2O. One microliter of 103 kinase buffer (New England Biolabs, Beverly, MA), 1 µl 10 mM ATP (Life Technologies) and 1 µl T4 polynucleotide kinase (NEB) were added to the reaction. After incubation at 37°C for 30 min, the kinase reaction was placed briefly on ice and 1 µl of 10 mM dNTPs (Life Technologies) and 1 µl T7 DNA polymerase (NEB) was added. This reaction was incubated for 15 min at 16°C. The correct PCR fragments were purified on a 6% TBE acrylamide gel and subcloned into the EcoRV site of pBlueScript™ II (Stratagene, La Jolla, CA). Reverse Northern blot analysis (35) of each isolated cDNA fragment was used to identify false positive clones. The remaining cDNAs were tested by Northern blot analysis of hepatocyte RNA for differential expression. Corticosteroid binding globulin (CBG) RT-PCR An aliquot of 1 µg of total RNA from control hepatocytes was used to prepare oligo dT-primed first strand cDNA (36). PCR primers CBG-5 (59ATGTCACTCGCCCTGTAT) and CBG-3 (59-TTCTTAGGCTGGATTGAC)
1452
(Cruachem, Dulles, VA) were used to amplify the 1194 bp full-length CBG cDNA using standard PCR reaction conditions (36). The amplification was performed at 94°C for 1 min, 48°C for 1 min and 72°C for 2 min for 30 cycles. Representational difference analysis Poly(A)1 RNA (2 µg) from control and AFB1-treated hepatocytes was used to construct double-stranded cDNA by standard methods (36). This cDNA was subjected to three rounds of subtractive hybridization as described by Hubank and Schatz (30). Primers and small cDNAs were removed from the subtracted, amplified cDNA population using a TE Midi Select column. The mixture of subtracted cDNAs was treated to prepare blunt ends and subcloned into EcoRV digested pBlueScript™ II as previously described for DD-PCR fragments. This library of subtracted, subcloned cDNAs was designated RDAI. Reverse Northern blot analysis of individual cDNAs isolated from the RDAI library showed that most of the clones were from highly expressed mRNAs that were not differentially expressed. Therefore plasmid DNA was prepared from the entire plated RDAI library and used in a second round of RDA to subtract these sequences from the tester cDNA population. The second round of RDA included a total of four subtractive hybridization steps. An aliquot of 5 µg of RDAI plasmid DNA was used as driver DNA in the second subtractive hybridization step of the second round of RDA to remove these highly expressed cDNAs from the tester population. After the fourth hybridization step, the subtracted cDNAs were prepared for blunt end subcloning as described for DD-PCR fragments and subcloned into EcoRV digest pBlueScript™ to generate the RDAII library. Individual transformants from the library were picked and used to inoculate each well of a 96-well dish, containing 200 µl LB with 150 µg/ml ampicillin in each well. The dishes were incubated at 37°C with gentle agitation for 4 h. The library was replica-plated onto LB ampicillin plates (150 µg/ml ampicillin) and incubated overnight at 37°C. The colonies were transferred to Duralon membranes (Stratagene) and processed for colony hybridization (36). Individual cDNAs were isolated from colonies and used as probes to classify members of the library that cross-hybridized. Unique members of the library were identified and plasmid DNA was prepared from each isolate using Plasmid Select-100 kits (5 Prime→3 Prime). Suppression subtractive hybridization Double-stranded cDNA was synthesized from 2 µg poly(A)1 RNA using the Great Lengths cDNA Synthesis Kit (Clontech, Palo Alto, CA) and used in SSH as previously described (31). Primers and unincorporated nucleotides were removed from the subtracted cDNAs using TE Midi Select column chromatography. Blunt-ended cDNA was prepared as previously described and the cDNA population was subcloned into EcoRV digested pLitmus™ (New England Biolabs, Beverly, MA). Unique members of the SSH library were identified by colony hybridization. Northern blot analysis using isolated cDNAs was used to verify differential expression. Northern blot analysis Total RNA (20 µg) or 4 µg poly(A)1 RNA was size-fractionated on a formaldehyde/MOPS gel (36) and transferred by passive capillary blot to Duralon-UV™ according to the manufacturer’s specifications. Northern blot analysis was performed as described previously (37). Gel-purified cDNAs were radiolabeled using [α-32P]dCTP by random primer labeling using the Prime-It II kit (Stratagene). Unincorporated nucleotides were removed using a TE Midi Select column. A rat cyclophilin probe was used as a normalization control for RNA loading. Quantification and comparison of signal intensity on Northern blots was performed using ImageQuant™ software on a PhosphorImager™ (Molecular Dynamics). Sequence analysis Plasmid DNA was sequenced using conventional sequencing primers and the Fidelity™ (Oncor, Gaithersburg, MD) or Prism™ Ready Reaction (ABI) kits according to the manufacturer’s specifications. DNA sequences were identified by comparison to those in DNA sequence databases, including GenBank and EMBL. These searches utilized the FastA software package (Frederick Biomedical Supercomputing Center).
Results Evaluation of subtraction techniques Genes differentially expressed after AFB1 exposure were identified using each of the three techniques: DD-PCR, RDA and SSH. The AFB1-responsive cDNAs and the technique used for each cDNA isolation are listed in Table I. Most of the AFB1-responsive genes were isolated using the RDA protocol, which identified both AFB1-induced and AFB1-
Genes in aflatoxin B1-treated hepatocytes
Table I. Identity of cDNAs based on database search homology Clone
GenBank matcha
Region clonedb
Accession no.
Method
AFB 4 AFB 6 AFB 12 AFB 23 AFB 24 AFB 35 ATT 5
Serum amyloid A Cytochrome P450 4F1 Corticosteroid binding globulin Alpha-2-microglobulin Glutathione S-transferase, Yb2 subunit C4 binding protein alpha chain Transferrin Cytochrome P450 3A1
L22190 M94548 M74776 M26835 M13590 Z50051 D38380 D38380 X64401
RDAII RDAII DD-PCR RDAII RDAII RDAII RDAII
ATT 7
30–130 bp 1120–1538 bp 1279–1429 bp 538–810 bp 424–845 bp 506–994 bp 1741–2174 bp 344–574 bp 191–395 bp
SSH
homology of ù95% with indicated GenBank match, except for 68% sequence homology of serum amyloid A with mouse serum amyloid A and 75% homology of corticosteroid binding globulin (CBG) with hamster CBG. of cloned cDNA sequence where base 1 is the initiation codon.
aSequence bPosition
Table II. Summary of Northern blot data of aflatoxin B1 responsive genes in primary cultured rat hepatocytes cDNA
Response to AFB1a
Estimated RNA size (kb)
Serum amyloid A-2 Cytochrome P450 4F1 Corticosteroid binding globulin Alpha-2-microglobulin Glutathione S-transferase Yb2 subunit C4 binding protein alpha chain Transferrin Cytochrome P450 3Alike Cytochrome P450 3A
↑2.43 ↑2.93 ↑8.33
1.0 2.3 1.4
↑2.93 ↑2.63
1.0 1.0
↑1.63
2.1
↓2.03 ↓60.03
2.1 0.5
↑2.03
3.8
aAverage
of two Northern blots after normalization to an internal standard.
Table III. Comparison of false positive rate of DD-PCR, RDA and SSH Technique
False positive cDNAs/ total no. cDNAs
% false positives
DD-PCR RDA SSH
14/15 3/10 0/1
93 30 0
repressed genes (Table I). No up-regulated genes were identified using the SSH protocol. However, the use of SSH did identify an AFB1-repressed gene that exhibited a much higher level of differential expression (.60 times; Table II) than seen with any cDNA isolated using RDA or DD-PCR. Corticosteroid binding globulin (CBG) cDNA was isolated using DD-PCR. The fact that a single AFB1-responsive gene was identified using this technique is probably because of the limited number of DD-PCR reactions performed in this experiment. Liang and Pardee (29) originally estimated that 240 DDPCR reactions per sample would be necessary to generate representative PCR bands of the majority of mRNA species present in a cell. We performed 30 DD-PCR reactions, which should amplify only 12.5% of the estimated 15 000 mRNA species. The false positive rate of each technique is shown in Table III. In our hands, the DD-PCR technique had an extremely high false positive rate (14/15), which we attribute in part to
running only single DD-PCR reactions instead of duplicate reactions. There was also heterogeneity in the re-amplified PCR fragments, which increased the false positive rate. Finally, a third criterion for a false positive was failure to demonstrate differential expression in hepatocyte RNA isolated from two independent experiments, and this factor contributes to the false positive rate of both DD-PCR and RDA. There were no false positive clones isolated using the SSH technique. Northern blot analysis Northern blot analysis was used to confirm differential gene expression. Total or poly(A)1 RNA was isolated from control and AFB1-treated hepatocytes. These hepatocytes were derived from a Fischer 344 male rat that was separate from the animals used in order to generate the hepatocyte RNA for the subtractive hybridization experiments. A second Northern blot from a third independent experiment was also performed to verify differential expression of each clone reported in this study (Table II). The alpha chain of C4b-binding protein (C4BP) showed a 1.6-fold increase in mRNA levels after AFB1 treatment (Table II, Figure 1). Northern blot analysis revealed a single mRNA of ~2.1 kb from the control and AFB1-treated hepatocytes, which is in agreement with the reported size for the mature rat C4BP alpha chain mRNA transcript (38). Alpha-2-microglobulin and serum amyloid protein A-2 mRNA levels were increased ~2.9-fold and 2.4-fold, respectively, in mRNA isolated from AFB1-treated hepatocytes (Table II, Figure 1). Expression of glutathione S-transferase Yb2 (GST) mRNA levels was increased almost 3-fold in AFB1-treated hepatocytes (Table II, Figure 1). In addition, GST mRNA expression was increased ~2-fold, 5-fold and 3-fold in liver mRNA from rats fed 0.01, 0.04 and 0.40 p.p.m. AFB1, respectively (Figure 2). Although the GST mRNA levels at the 0.40 p.p.m. dose are significantly higher than the levels in control animals, they are reduced ~2-fold from the GST levels in animals fed 0.04 p.p.m. AFB1. This decrease may be caused by the toxic effects of AFB1 on RNA polymerase II and other macromolecules that influence gene transcription at the higher dose. CYP4F1 (AFB6) expression levels were examined in control and AFB1-treated hepatocyte poly(A)1 RNA derived from two independent experiments. In both experiments, a 2.3-kb band was found to have increased ~3-fold after AFB1 exposure (Table II, Figure 1). Expression of CYP4F1 mRNA was also up-regulated over 2-fold in total liver RNA from male Fischer 344 rats fed 0.40 p.p.m. AFB1 on an intermittent dosing regime for 12 weeks (Figure 3). 1453
A.J.Harris et al.
Fig. 2. GST mRNA levels in control and AFB1-treated Fischer 344 rat liver. Northern blot analysis of 4 µg poly(A)1 RNA from male rats fed 0, 0.01, 0.04 or 0.40 p.p.m. AFB1 ad libitum for 4 weeks, then killed. Quantitation of band intensity was performed using ImageQuant™ on a PhosphorImager™ (Molecular Dynamics). Values represent the means 6 SE (n 5 3 for all doses) after normalization of RNA loading using cyclophilin expression as the standard. **Statistically significant (P , 0.01) from control animals using Student’s t-test.
Fig. 1. Northern blot analysis of mRNA from control (CTL) and AFB1treated (AFB1) rat primary hepatocyte cultures. Poly(A)1 RNA (5 µg) or total RNA (20 µg)a was probed with 32P-labeled isolated cDNAs as described in Materials and methods. All blots were stripped after analysis by incubation in boiling 0.13 SSC and 0.1% SDS, then re-hybridized with a probe for cyclophilin or glyceraldehyde 3-phosphate dehydrogenase (GAPDH) for normalization of RNA loading. The images were generated using ImageQuant™ software and a PhosphorImager™ (Molecular Dynamics).
Our initial results using the AFB6 cDNA as a probe in Northern blot analysis showed that there were four hybridizing bands in hepatocyte RNA, rather than the unique band around 2.0 kb that was expected for CYP4F1 (39). Analysis of the AFB6 sequence revealed a 45 bp cDNA attached to the CYP4F1 cDNA, which had no sequence homology with CYP4F1 or any other sequence in GenBank. This unknown sequence (UK-1) was separated from the CYP4F1 cDNA by restriction digest and used as a probe in Northern blot analysis. The UK-1 probe hybridized with three bands in the hepatocyte RNA that correlated in size with the extra bands seen in our initial Northern blot studies using the AFB6 cDNA probe (data not shown). Initial Northern blots using UK-1 as a probe suggested that one of the mRNAs it recognizes is also differentially expressed; however, use of this probe has been inconsistent, possibly because of its small size. We are currently 1454
Fig. 3. CYP4F1 mRNA levels in control and AFB1-treated Fischer 344 rat liver. Northern blot analysis of 20 µg total RNA from male rats fed 0, 0.04 or 0.4 p.p.m. AFB1 ad libitum intermittently for 12 weeks (4 weeks on, 4 off, 4 on). Quantitation of band intensity was performed using ImageQuant™ on a PhosphorImager™ (Molecular Dynamics). Values represent the means 6 SE (n 5 3 for controls and 0.4 p.p.m., n 5 1 for 0.04 p.p.m.) after normalization of RNA loading using cyclophilin expression as the standard. *Statistically significant (P , 0.05) from control animals using Student’s t-test.
working to isolate larger UK-1 cDNAs in order to resolve this inconsistency. Sequence analysis of AFB12 showed it to be identical to the 39 untranslated open reading frame found upstream of the poly(A)1 signal in the corticosteroid binding globulin (CBG)
Genes in aflatoxin B1-treated hepatocytes
Fig. 4. CBG mRNA levels in control and AFB1-treated Fischer 344 rat liver. Northern blot analysis of 20 µg total RNA from male rats fed 0 or 0.4 p.p.m. AFB1 ad libitum intermittently for 12 weeks (4 weeks on, 4 off, 4 on). Quantitation of band intensity was performed using ImageQuant™ on a PhosphorImager™ (Molecular Dynamics). Values represent the means 6 SE (n 5 3) after normalization of RNA loading using cyclophilin expression as the standard. *Statistically significant (P , 0.05) from control animals using Student’s t-test.
cDNA sequence (Table I) (40). There was no signal after Northern blot analysis of total hepatocyte RNA using the 180 bp AFB12 cDNA. Full-length rat CBG was cloned by RTPCR and used for Northern blot analysis of poly(A)1 RNA isolated from control and AFB1-treated hepatocytes. CBG expression was analyzed in two independently derived Northern blots using poly(A)1 RNA isolated from AFB1-treated rat hepatocyte primary cultures. In both cases, CBG mRNA levels were higher in the AFB1 samples than in control hepatocytes by ~6-fold. A 53% increase in CBG mRNA levels was also demonstrated in vivo in the RNA isolated from the livers of male Fischer 344 rats fed 0.40 p.p.m. AFB1 on an intermittent schedule for 12 weeks (Figure 4). It has been reported that dexamethasone can decrease CBG mRNA levels in male rats, presumably by altering the mRNA stability (41). Since the hepatocytes used in this study were cultured in the presence of dexamethasone, the expression of CBG was evaluated in freshly isolated hepatocytes that contained no dexamethasone and in those that were incubated for 16 h on the culture dish in the presence of dexamethasone. We found that CBG mRNA levels had decreased by ~12fold in the hepatocytes after incubation in the presence of dexamethasone (data not shown), although CBG mRNA levels were increased 6-fold in hepatocytes treated with dexamethasone and AFB1. A CYP3A-like cDNA was isolated using SSH (ATT 7 probe; Table I). Northern blot analysis using this cDNA revealed a 450 bp mRNA transcript that was decreased .60-fold in the AFB1-treated hepatocytes (Figure 1). Two transcripts, 3.8 kb and 1.9 kb in size, were detected in a Northern blot analysis of poly(A)1 RNA after an extended exposure (data not shown). These transcripts increased in abundance 2- and 1.5-fold, respectively after AFB1 exposure. The cDNA region cloned (Table I) is identical in both CYP3A1
Fig. 5. CYP3A mRNA levels in control and AFB1-treated Fischer 344 rat liver. Northern blot analysis of 4 µg poly(A)1 RNA from male rats fed 0, 0.01, 0.04 or 0.40 p.p.m. AFB1 ad libitum for 4 weeks, then killed. Quantitation of band intensity was performed using ImageQuant™ on a PhosphorImager™ (Molecular Dynamics). Values represent the means 6 SE (n 5 3 for all doses) after normalization of RNA loading using cyclophilin expression as the standard. **Statistically significant (P , 0.01) from control animals using Student’s t-test
and CYP3A23 (42,43) and it is probable that the larger 1.9 kb and 3.8 kb signals were caused by hybridization to the CYP3A mRNAs. However, the identity of the 450 bp mRNA is unclear. It is unlikely to be a degradation product of CYP3A1 or CYP3A23 since the same mRNA species was detected in two Northern blots using RNA derived from independent experiments. It is possible that the 450 bp band represents an alternate splice of CYP3A1 or CYP3A23 that is differentially regulated by AFB1; or it may be a different CYP3A cDNA with a region homologous to the CYP3A1/CYP3A23 genes; or, alternatively, it is a unique cDNA. It may also be an artifact of the primary hepatocyte culture since the 450 bp mRNA species was not seen in liver mRNA from AFB1-treated rats (data not shown). Northern blot analysis of mRNA from livers of AFB1-treated rats using the ATT 7 cDNA as a probe revealed a hybridizing band of ~4 kb, which is probably CYP3A. This mRNA was increased .2-fold in rats fed 0.04 p.p.m. AFB1 for 4 weeks (Figure 5), which is similar to the increase seen in AFB1-treated primary rat hepatocytes (data not shown). Transferrin RNA levels were decreased ~2-fold in RNA from AFB1-treated hepatocytes (Table II, Figure 1). Northern blot analysis of total RNA from livers of rats fed 1.6 p.p.m. AFB1 on the 8-week intermittent dosing regime also demonstrated a 1.5-fold reduction in transferrin mRNA levels relative to control animals (Figure 6). Discussion We have identified several genes that could potentially play important roles in the carcinogenicity and toxicity of AFB1 in rat liver. In addition, we have evaluated three techniques for their utility in identification of differentially expressed genes, DD-PCR, RDA and SSH. DD-PCR had the highest false positive rate of the three techniques. However, there have been 1455
A.J.Harris et al.
Fig. 6. Transferrin mRNA levels in control and AFB1-treated Fischer 344 rat liver. Northern blot analysis of 20 µg total RNA from male rats fed 0 or 1.6 p.p.m. AFB1 ad libitum intermittently for 8 weeks (4 weeks on, 4 off). Quantitation of band intensity was performed using ImageQuant™ on a PhosphorImager™ (Molecular Dynamics). Values represent the means 6 SE (n 5 3 for controls, n 5 5 for 1.6 p.p.m.) after normalization of RNA loading using cyclophilin expression as the standard. **Statistically significant (P , 0.01) from control animals using Student’s t-test.
many improvements reported in the literature to significantly reduce this problem (44), such as the inclusion of duplicate DD-PCR reactions to control for reproducibility and the use of gel-systems, which enhance band resolution and reduce the heterogeneity problem (45). The two greatest advantages of DD-PCR over RDA and SSH are the ability to compare multiple samples simultaneously and the greater probability of identifying differentially expressed mRNAs present in low copy number. RDA and SSH allow the comparison of only two samples. RDA enabled us to identify six cDNAs that demonstrated modest changes in mRNA expression. In contrast, only one cDNA was isolated using SSH but the reduction in gene expression was striking (.60-fold; Figure 1). There were no false positive clones isolated using SSH. Although we have not done an exhaustive analysis of these three techniques, it may be that the use of SSH biased the isolation of genes toward those that have a larger differential in gene expression. This would explain in part why such a low number of cDNAs were isolated using SSH. Each technique generated a different set of cDNAs, which underscores the fact that the choice of technique will preferentially isolate a unique subset of genes based on the characteristics of the subtraction and PCR reactions, and that the use of one technique (or all three techniques) is unlikely to allow identification of all responsive genes. Representatives of both phase I and phase II metabolism enzymes were found to be AFB1-responsive. Phase I enzymes identified include two members of the cytochrome P450 superfamily, CYP3A and CYP4F1. CYP3A enzymes have been reported to metabolically activate AFB1 in both human (15,16,46) and mouse (14). Also, AFB1 activation is enhanced in rats pre-treated with phenobarbital, which acts as an inducer of CYP3A enzymes (42,43,47). It is intriguing that AFB1 can cause an increase in CYP3A mRNA levels in both primary 1456
rat hepatocytes and AFB1-treated rats, since CYP3A P450s apparently play a major role in AFB1 activation. CYP4F1 is a recently identified member of the CYP4 superfamily (39). We found that CYP4F1 mRNA levels were increased in AFB1-treated primary rat hepatocytes as well as in the livers of rats fed 0.4 p.p.m. AFB1 for 12 weeks on an intermittent dosing schedule. Previously, CYP4F1 mRNA was shown to be constitutively expressed in rat liver and upregulated in an AFB1-induced transplantable rat tumor (39). In the same study, CYP4F1 expression was apparent but not increased in tumors induced by ciprofibrate, an analog of the peroxisome proliferator clofibrate, or by the genotoxic hepatocarcinogens 2-acetylaminofluorene and diethylnitrosamine. The biological role of CYP4F1 is not known; however, it is intriguing that up-regulation of this mRNA has been observed in response to acute AFB1 exposure in cultured primary rat hepatocytes and after chronic exposure in Fischer 344 male rats. Furthermore, increased CYP4F1 expression was also found in AFB1-induced tumors, but not in tumors produced after exposure to other liver carcinogens (39). Perhaps AFB1 is a hitherto unidentified substrate of this isozyme and contributes to metabolic activation of this potent mutagenic carcinogen. Phase II glutathione S-transferases (GSTs) play a vital role in the detoxification of aflatoxin 8,9-expoxide by conjugation with glutathione. GST Yb2 is a constitutively expressed member of the mu class GSTs. Gopalan et al. (48) demonstrated that mu class GSTs were primarily responsible for microsome mediated aflatoxin 8,9-epoxide conjugation in rat cytosol, therefore AFB1-induced increases in GST Yb2 expression could play a role in the response of a cell to AFB1 exposure by enhancing the detoxification of reactive AFB1 metabolites. GST Yb2 mRNA levels have also been reported to be increased ~5-fold in the livers of rats treated with phenobarbital (49) and thiazole (50). That increase in GST mRNA is very similar to the induction levels we report here in AFB1-treated rats. CBG, C4bp, serum amyloid A (SAA) and transferrin mRNAs were also found to be AFB1-responsive in this study. Each of these genes is an acute phase response gene. CBG is a negative acute response reactant while C4bp, SAA and transferrin are all positive acute response reactants. However, contrary to their normal expression during an acute phase response, we found an increase in CBG mRNA levels and a decrease in transferrin mRNA levels after AFB1 exposure in the hepatocytes, which suggests the effect of AFB1-induced changes in expression of these mRNAs may be modulated through mechanisms other than the acute phase response. Transferrin mRNA levels decreased 2-fold in AFB1-treated rat hepatocyte cultures and 1.5-fold in the livers of rats exposed to 1.6 p.p.m. AFB1 in an intermittent 8-week dosing regime. Reduced levels of transferrin mRNA in N-methyl-N-nitrosourea induced rat mammary tumors (51) and lowered transferrin protein levels in human fibroblasts immortalized by 4-nitroquinoline 1-oxide and 60Co gamma rays (52) have also been reported. A previous study found that AFB1 did not alter transferrin gene expression in the chick embryo (53). This apparent discrepancy may be attributable to metabolic and/or gene regulation differences in species (chickens versus rats), developmental stage (embryonic versus adult) or AFB1 dosing levels. Decreased serum transferrin levels were proposed by Harvey et al. (7) to be responsible for the lowered unsaturated and total iron binding capacities seen in pigs exposed to AFB1
Genes in aflatoxin B1-treated hepatocytes
(7,9,10). Effects on iron levels are symptoms of AFB1 toxicosis observed in many agricultural animals after exposure through feed. A reduction in serum transferrin levels is usually associated with increases in hepatic iron levels. One consequence of iron overload could be the generation of reactive oxygen species (ROS), particularly the hydroxyl radical formed via the Fenton reaction. AFB1 has been reported by Shen et al. (54) to induce a dose-dependent increase in formation of the hydroxy radical DNA adduct, 8-hydroxydeoxyguanosine (8OH-dG) in liver DNA isolated from AFB1-dosed rats. Levels of 8-OH-dG were significantly reduced by pre-treatment with deferoxamine (DFO), a specific iron chelator, which indicates a role for iron in this mechanism. AFB1-mediated cell damage, lipid peroxidation and enhanced ROS production also have been reported in cultured rat hepatocytes (55,56). These effects were reduced significantly after pre-treatment with DFO or DMSO, a hydroxyl radical scavenger. C4BP alpha chain mRNA levels increased 2-fold in AFB1treated rat hepatocytes. C4BP plays an important role in the negative regulation of the soluble phase of the classical complement cascade by binding C4b in the bloodstream, and prevents the formation of C3 convertase (57,58) and ultimately of serum complement. It should be noted that one of the immunosuppressive effects of AFB1 is a reduction in serum complement levels (59–61). CBG is a 50–60 kDa monomeric glycoprotein produced primarily in the liver, which binds glucocorticoids with high affinity (62,63). Up to 90% of circulating steroids are bound to CBG in the sera (64) and CBG receptors have been identified and partially characterized in rat spleen and liver (65–67). AFB1 has not been reported to bind to CBG; however, AFB1 has a steroid-like structure, which makes the idea plausible. It is possible that elevated CBG levels act either as a detoxification mechanism by clearing AFB1 from the tissues, or conversely, they increase AFB1 intracellular levels through AFB1–CBG complex entry via CBG receptors into the cell and eventual transport into the nucleus, to increase its genotoxic potential. It is not known how AFB1 modulates gene transcription. Promoter analysis of the AFB1-responsive mdr1b gene in rats revealed no carcinogen specific response elements (68). The promoter sequences responsible for AFB1-induced increases in mdr1b transcription overlap those responsible for basal promoter activity. Based on this, it has been proposed that AFB1 and other carcinogens that produce bulky adducts affect gene expression by increasing the level of transcription factors responding to DNA damage (53). It is also possible that AFB1induced increases in ROS (56) could be responsible for alterations in gene expression (reviewed in ref. 69). Reactive AFB1 metabolites could bind to and thus alter the availability of basal transcription factors, and so affect transcription rates. In this way, negative regulators of transcription would be sequestered, promoting increased transcription levels from genes. Conversely, a decrease in basal transcription factor levels through AFB1 metabolite binding could cause reductions in transcription of many genes. AFB1-induced decreases in gene transcription are not surprising since AFB1 is known to have an inhibitory effect on both nuclear and nucleolar RNA synthesis, and RNA polymerase II activity (70). In summary, we have isolated several AFB1-responsive genes from rat hepatocytes using DD-PCR, RDA and SSH techniques. Exposure to AFB1 increased mRNA levels in rat hepatocytes for CBG, CYP4F1, CYP3A, alpha-2-microglobulin, C4b-binding protein, serum amyloid A-2 and GST Yb2.
mRNA levels were decreased for transferrin and a small CYP3A-like mRNA species. Furthermore, when male Fischer 344 rats were exposed to AFB1, CBG CYP3A, CYP4F1 and GST, mRNA levels increased and transferrin mRNA levels decreased, which is consistent with results from AFB1-exposed hepatocytes and demonstrates the utility of hepatocyte studies for studying genes important for in vivo effects. Further studies on how the AFB1-responsive genes identified here, individually as well as collectively, regulate biological processes in a cell may provide more clues to understand the toxic, immunosuppressive and carcinogenic actions of AFB1. Acknowledgements The authors thank Dr Robert Heflich and Dr William Tolleson for critical evaluation of the manuscript, and Dr Barbara Parsons for helpful discussions. We also thank Dr Ming Chou for providing tissues samples.
References 1. Busby,W.F. and Wogan,G.N. (1984) Aflatoxin. Chem. Carcinog., 2, 945– 1136. 2. Groopman,J.D., Cain,L.G. and Kensler,T. (1988) Aflatoxin exposure in human populations; measurements and relationship to cancer. CRC Crit. Rev. Toxicol., 19, 113–143. 3. Dvorackova,I. (1976) Aflatoxin inhalation and alveolar cell carcinoma. Br. Med. J., 1, 691. 4. Wieder,R, Wogan,G.N. and Shimkin,M.B. (1968) Pulmonary tumors in strain A mice given injections of aflatoxin B1. J. Natl Cancer Inst., 40, 1195–1197. 5. Krishnamachari,K., Bhar,R., Nagarajan,V. and Tilac,T. (1975) Hepatitis due to aflatoxicosis. Lancet, 1, 1061–1063. 6. Robens,J.F. and Richard,J.L. (1992) Aflatoxins in animal and human health. Rev. Environ. Contam. Toxicol., 127, 69–94. 7. Harvey,R.B., Clark,D.E., Huff,W.E., Kubena,L.F., Corrier,D.E. and Phillips,T.D. (1988) Suppression of serum iron-binding capacity and bone marrow cellularity in pigs fed aflatoxin. Bull. Environ. Contam. Toxicol., 40, 576–583. 8. Stewart,R.G., Skeeles,J.K., Wyatt,R.D., Brown,J., Page,R.K., Russell,I.D. and Lukert,P.D. (1985) The effect of aflatoxin on complement activity in broiler chickens. Poult. Sci., 64, 616–619. 9. Thurston,J.R., Baetz,A.L. and Cheville,N.F. (1980) Acute aflatoxicosis in guinea pigs; sequential changes in serum proteins, complement, C4, liver enzymes and histopathologic changes. Am. J. Vet. Res., 41, 1272–1276. 10. Harvey,R.B., Kubena,L.F., Huff,W.E., Corrier,D.E., Rottinghaus,G.E. and Phillips,T.D. (1990) Effects of treatment of growing swine with aflatoxin and T-2 toxin. Am. J. Vet. Res., 51, 1688–1693. 11. Swenson,D.H., Miller,E.C. and Miller,J.A. (1974) Aflatoxin B1-2,3-oxide: evidence for its formation in rat liver in vivo and by human liver microsomes in vitro. Biochem. Biophys. Res. Commun., 60, 1036–1043. 12. Robertson,I.G., Zeiger,E. and Goldstein,J.A. (1983) Specificity of rat liver cytochrome P-450 isozymes in the mutagenic activation of benzo[a]pyrene, aromatic amines and aflatoxin B1. Carcinogenesis, 4, 93–96. 13. Metcalfe,S.A., Colley,P.J. and Neal,G.E. (1981) A comparison of the effects of pretreatment with phenobartital and 3-methylcholanthrene on the metabolism of aflatoxin B1 by liver rat microsomes and isolated hepatocytes in vitro. Chem.-Biol. Interact., 35, 145–157. 14. Yanagimoto,T., Susumu,I., Sawada,M., Hashimoto,H. and Kamataki,T. (1994) Molecular cloning and functional expression of a mouse cytochrome P-450 (Cyp3a-13): examination of Cyp3a-13 enzyme to activate aflatoxin B1 (AFB1). Biochim. Biophys. Acta, 1201, 405–410. 15. Aoyama,T., Yamano,S., Guzelian,P.S., Gelboin,H.V. and Gonzalez,F.J. (1990) Five of 12 forms of vaccine virus-expressed human hepatic cytochrome P450 metabolically activate aflatoxin B1. Proc. Natl Acad. Sci. USA, 87, 4790–4793. 16. Crespi,C.L., Penman,B.W., Steimel,D.T., Gelboin,H.V. and Gonzalez,F.J. (1991) The development of a human cell line stably expressing human CYP3A4: Role in the metabolic activation of aflatoxin B1 and comparison to CYP1A2 and CYP2A3. Carcinogenesis, 12, 355–359. 17. Liu,L. and Massey,T.E. (1992) Bioactivation of aflatoxin B1 by lipoxygenases, prostaglandin H synthase and cytochrome P450 monooxygenase in guinea-pig tissues. Carcinogenesis, 13, 533–539. 18. Roy,S.K. and Kulkarni,A.P. (1997) Aflatoxin B1 epoxidation catalysed by partially purified human liver lipoxygenase. Xenobiotica, 27, 231–241. 19. Lin,J.K., Miller,J.A. and Miller,E.C. (1977) 2,3-Dihydro-2-(guanyl-7-yl)-3hydroxy-aflatoxin B1—a major acid hydrolysis product of aflatoxin Be–DNA
1457
A.J.Harris et al. or ribosomal RNA adducts formed in hepatic microsome-mediated reactions and in rat liver in vivo. Cancer Res., 37, 4430–4438. 20. Essigmann,J.M., Broy,R.G., Nadzan,A.M., Busby,W.F.Jr, Reinhold,V.N., Buchi,G. and Wogan,G.N. (1977) Structure identification of the major DNA adduct formed by aflatoxin B1 in vitro. Proc. Natl Acad. Sci. USA, 74, 1870–1874. 21. Choy,W.N. (1993) A review of the dose-response induction of DNA adducts by aflatoxin B1 and its implications to quantitative cancer-risk assessment. Mutat. Res., 296, 181–198. 22. Bressac,B., Kew,M., Wands,J. and Ozturk,M. (1991) Selective G to T mutations of p53 gene in hepatocellular carcinoma from Southern Africa. Nature, 350, 429–431. 23. Hsu,I.C., Metcalf,R.A., Sun,T., Welsh,J.A. and Wang,N.J. (1991) Mutational hotspot in the p53 gene in human hepatocellular carcinomas. Nature, 350, 427–428. 24. McMahon,G., Davis,E.F., Huber,L.J., KimY. and Wogan,G.N. (1987) Characterization of c-Ki-ras and N-ras oncogenes in aflatoxin B1 in rat liver tumors. Proc. Natl Acad. Sci. USA, 84, 4974–4978. 25. Soman,N.R. and Wogan,G.N. (1993) Activation of the c-Ki-ras oncogene in aflatoxin B1-induced hepatocellular carcinoma and adenoma in the rat: Detection by denaturing gradient gel electrophoresis. Proc. Natl Acad. Sci. USA, 90, 2045–2049. 26. Solt,D. and Farber,E. (1976) New principle for the analysis of chemical carcinogenesis. Nature, 263, 701–703. 27. Davis,E., Maxuitenko,Y., Macmillan,D., Dolan,P., Roebuck,B. and Kensler,T. (1993) Oltipraz protects rats from the autopromoting effects of aflatoxin. Proc. Am. Assoc. Cancer Res., 34, 550. 28. Roebuck,B.D. and Maxuitenko,Y.Y. (1994) Biochemical mechanisms and biological implications of the toxicity of aflatoxins as related to aflatoxin carcinogenesis. In Eaton, D.L. and Groopman, J.D. (eds) The Toxicology of Aflatoxins: Human Health, Veterinary and Agricultural Significants. Academic Press, San Diego, CA, pp. 27–43. 29. Liang,P. and Pardee,A.B. (1992) Differential display of eukaryotic messenger RNA by means of the polymerase chain reaction. Science, 257, 967–971. 30. Hubank,M. and Schatz,D.G. (1994) Identifying differences in mRNA expression by representational difference analysis of cDNA. Nucleic Acids Res., 22, 5640–5648. 31. Diatchenko,L., Lau,Y.C., Campbell,A.P. et al. (1996) Suppression subtractive hybridization: A method for generating differentially regulated or tissue-specific cDNA probes and libraries. Proc. Natl Acad. Sci. USA, 93, 6025–6030. 32. Oldham,J.W., Casciano,D.A. and Farr,J.A. (1979) The isolation and primary culture of viable, nonproliferating rat hepatocytes. RCA Manual, 5, 1047– 1050. 33. Shaddock,J.G., Heflich,R.H., McMillan,D.C., Hinson,J.A. and Casciano,D.A. (1989) Pretreatment with mixed-function oxidase inducers increases the sensitivity of the hepatocyte/DNA repair assay. Environ. Mutagen., 13, 281–288. 34. Ponte,P. and Liu,H. Unpublished communication, Berlex Biosciences, Richmond, CA. 35. Mou,L. Miller,H., Li,J., Wang,E. and Chalifour,L. (1994) Improvements to the differential display methods for gene analysis. Biochem. Biophys. Res. Commun., 199, 564–569. 36. Current Protocols in Molecular Biology (vols 1 and 2) (1994) John Wiley & Sons, UK. 37. Virca,G. D., Northemann,W., Shiels,B.R., Widera,G. and Broome,S. (1990) Simplified Northern blot hybridization using 5% sodium dodecyl sulfate. Biotechniques, 8, 370–371. 38. Hillarp,A., Wiklund,H., Thern,A. and Dahlba¨ck,B. (1997) Molecular cloning of rat C4b binding protein α- and β-chains. J. Immun., 158, 1315–1323. 39. Chen,L. and Hardwick,J.P. (1993) Identification of a new P450 subfamily, CYP4F1, expressed in rat hepatic tumors. Arch. Biochem. Biophys., 300, 18–23. 40. Smith,C.L. and Hammond,G.L. (1989) Rat corticosteroid binding globulin: Primary structure and messenger ribonucleic acid levels in the liver under different physiological conditions. Mol. Endrocinol., 1989, 420–426. 41. Smith,C.L. and Hammond,G.L. (1992) Hormonal regulation of corticosteroid-binding globulin biosynthesis in the male rat. Endocrinology, 130, 2245–2251. 42. Kirita,S. and Matsubara,T. (1993) cDNA cloning and characterization of a novel member of steroid-induced cytochrome P450 3A in rats. Arch. Biochem. Biophys., 307, 253–258. 43. Komori,M. and Oda,Y. (1994) A major glucocorticoid-inducible P450 in rat liver is not P450 3A1. J. Biochem., 116, 114–120. 44. Liang,P. and Pardee,A.B. (1995) Recent advances in differential display. Curr. Opinion Immunol., 7, 274–280. 45. Averboukh,L., Douglas,S.A., Zhao,S., Lowe,K., Maher,J. and Pardee,A.B. (1996) Better gel resolution and longer cDNAs increase the precision of differential display. BioTechniques, 20, 918–921.
1458
46. Shimada,T. and Guengerich,F.P. (1989) Evidence for cytochrome P-450NF, the nifedipine oxidase, being the principal enzyme involved in the bioactivation of aflatoxins in human liver. Proc. Natl Acad. Sci., USA, 86, 462–465. 47. Metcalfe,S.A., Colley,P.J. and Neal,G.E. (1981) A comparison of the effects of pretreatment with phenobarbital and 3-methylcholanthrene on the metabolism of aflatoxin B1 by liver rat microsomes and isolated hepatocytes in vitro. Chem. Biol. Interact., 35, 145–157. 48. Gopalan,P., Jensen,D.E. and Lotlikar,P.D. (1992) Glutathione conjugation of microsome-mediated and synthetic aflatoxin B1-8,9-oxide by purified glutathione S-transferases from rats. Cancer Lett., 64, 225–233. 49. Ding,G.J.F., Ding,V.D.H., Rodkey,J.A., Bennett,C.D., Lu,A.Y.H. and Pickett,C.B. (1996) DNA sequence analysis of a Yb2 cDNA clone and regulation of the Yb1 and Yb2 mRNAs by phenobarbital. J. Biol. Chem., 261, 7952–7957. 50. Kim,S.G. and Cho,M.K. (1996) Expression of glutathione S-transferases Ya, Yb1, Yb2, Yc1 and Yc2 and microsomal epoxide hydrolase genes by thiazole, benzothiazole and benzothiadiazole. Biochem. Pharmacol., 52, 1831–1841. 51. Young,L.H., Yang,X. and Voigt,J.M. (1996) Alteration of gene expression in rat mammary tumors induced by N-methyl-N-nitrosourea. Mol. Carcinogenesis, 15, 251–260. 52. Kondo,T., Mihara,K., Inoue,Y. and Masayosi,N. (1996) Two-dimensional electrophoretic studies on down-regulated intracellular transferrin in human fibroblasts immortalized by treatment with either 4-nitroquinoline 1-oxide or 60Co gamma rays. Electrophoresis, 17, 1638–1642. 53. Hamilton,J.W., Louis,C.A., Doherty,K.A., Hunt,S.R., Reed,M.J. and Treadwell,M.D. (1993) Preferential alteration of inducible gene expression in vivo by carcinogens that induce bulky DNA lesions. Mol. Carcinog., 8, 34–43. 54. Shen,H.M., Ong,C.N., Lee,B.L. and Shi,C.Y. (1995) Aflatoxin B1-induced 8-hydroxydeoxyguanosine formation in rat hepatic DNA. Carcinogenesis, 16, 419–422. 55. Shen,H.M., Ong,C.N. and Shi,C.Y. (1995) Involvement of reactive oxygen species in aflatoxin B1-induced cell injury in cultured rat hepatocytes. Toxicology, 99, 115–123. 56. Shen,H.M., Shi,C.Y., Shen,Y. and Ong,C.N. (1996) Detection of elevated reactive oxygen species levels in cultured rat hepatocytes treated with aflatoxin B1. Free Radical Biol. Med., 21, 139–146. 57. Fujita,T. and Nussenzweig,V. (1979) The role of C4-binding protein and beta 1H in proteolysis of C4b and C3b. J. Exp. Med., 150, 267–276. 58. Gigli,I., Fujita,T. and Nussenzweig,V. (1979) Modulation of the classical pathway C3 convertase by plasma proteins C4 binding protein and C3b inactivator. Proc. Natl Acad. Sci. USA, 76, 6596–6600. 59. Thurston,J.R., Baetz,A.L. and Cheville,N.F. (1980) Acute aflatoxicosis in guinea pigs; sequential changes in serum proteins, complement, C4, liver enzymes and histopathologic changes. Am. J. Vet. Res., 41, 1272–1276. 60. Stewart,R.G., Skeeles,J.K. Wyatt,R.D., Brown,J., Page,R.K., Russell, I.D. and Lukert,P.D. (1985) The effect of aflatoxin on complement activity in broiler chickens. Poult. Sci., 64, 616–619. 61. Thurston,J.R., Sacks,J.M., Richard,J.L., Peden,W.M. and Driftmier,K. (1989) Complement, bacteriostaticand enzymatic activities in sera from guinea pigs given aflatoxin and/or rubratoxin. Am. J. Vet. Res., 50, 356–358. 62. Westphal,U. (1971) Steroid–Protein Interactions. Springer Verlag, New York. 63. Weiser,J.N., Do,Y.S. and Feldman,D. (1979) Synthesis and secretion of corticosteroid-binding globulin by rat liver: a source of heterogeneity of hepatic corticosteroid-binders. J. Clin. Invest., 63, 461–467. 64. Siiteri,P.K., Murai,J.T., Hammond,G.L., Nisker,J.A., Raymoure,W.J. and Kuhn,R.W. (1982) The serum transport of steroid hormones. Recent Prog. Horm. Res., 38, 457–510. 65. Singer,C.J., Khan,M.S. and Rosner,W. (1988) Characteristics of the binding of corticosteroid-binding globulin to rat cell membranes. Endocrinology, 122, 89–96. 66. Maitra,U.S., Khan,M.S., Zhang,X.H. and Rosner,W. (1990) The rat hepatic corticosteroid-binding globulin receptor: distinction from the asialoglycoprotein receptor. Endocrinology, 127, 278–284. 67. Maitra,U.S., Khan,M.S. and Rosner,W. (1993) Corticosteroid-binding globulin receptor of the rat hepatic membrane: solubilization, partial characterization and the effect of steroids on binding. Endocrinology, 133, 1817–1822. 68. Silverman,J.A. and Hill,B.A. (1995) Characterization of the basal and carcinogen regulatory elements of the rat mdr1b promoter. Mol. Carcinog., 13, 50–59. 69. Sen,C.K. and Packer,L. (1996) Antioxidant and redox regulation of gene transcription. FASEB J., 10, 709–720. 70. Yu,F.L. (1977) Mechanism of aflatoxin B1 inhibition of rat hepatic nuclear RNA synthesis. J. Biol. Chem., 252, 3245–3251. Received on February 5, 1998; revised on April 22, 1998; accepted on April 28, 1998
