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Translesion DNA Synthesis across the Heptanone-Etheno-2′-Deoxycytidine Adduct in Cells Michael Pollack,† In-Young Yang,‡ Hye-Young H. Kim,†,§ Ian A. Blair,† and Masaaki Moriya*,‡ Center for Cancer Pharmacology, UniVersity of PennsylVania, Philadelphia, PennsylVania 19104, and the Laboratory of Chemical Biology, Department of Pharmacological Sciences, State UniVersity of New York, Stony Brook, New York 11794 ReceiVed March 6, 2006
4-Oxo-2(E)-nonenal, a lipid peroxidation-derived product, reacts with dG, dA, and dC in DNA to form heptanone (H)-etheno () adducts. Among the three adducts, H-dC is formed in the greatest abundance in in vitro reactions, and it has been detected in the C57BL/6JAPCmin mouse model of colorectal cancer. To establish the genotoxic properties of this adduct, a site-specifically modified oligonucleotide was synthesized and incorporated into a shuttle vector. The modified vector was replicated in Escherichia coli and human cells. Analysis of the progeny plasmid has revealed that H-dC strongly blocks DNA synthesis and markedly miscodes in both hosts. The miscoding frequency was 40-50% in bacteria and more than 90% in three human cell lines (xeroderma pigmentosum A and variant cells, and DNA repair wild-type cells). There was a drastic difference in coding events in these two hosts: dG and dC were almost exclusively inserted opposite the lesion in E. coli, while dA and dT were the preferential choices in human cells. These results indicate that this endogenous DNA adduct is very genotoxic to both organisms. Introduction The generation of reactive products by oxidative degradation of lipids is implicated in a number of age-related diseases such as cancer, cardiovascular disease, and neurodegenerative diseases. Polyunsaturated fatty acids can be oxidized to lipid hydroperoxides by reactive oxygen species, lipoxygenases (1), or cyclooxygenases (COXs)1 (2). Lipid hydroperoxides can subsequently undergo FeII, CuI, or vitamin C-mediated homolytic decomposition (3-5) to form R,β-unsaturated aldehydes, such as 4-hydroperoxy-2(E)-nonenal, 4-oxo-2(E)-nonenal (ONE), 4-hydroxy-2(E)-nonenal, and 4,5-epoxy-2(E)-decenal (Figure 1) (5-7). These aldehydes act as bifunctional electrophiles that can covalently modify nucleosides, DNA, amino acids, and proteins (8-11). Covalent modifications to DNA by these electrophiles have been unequivocally identified and quantified in mammalian DNA (12-14). DNA adducts are biologically very significant due to their potential to block DNA replication and transcription, induce DNA strand breaks, trigger apoptosis, and cause gene mutations and chromosomal aberrations. If they lead to mutations in the genome stability maintenance genes, tumor suppressor genes, and/or protooncogenes, they could directly contribute to carcinogenesis (8, 15). The novel lipid peroxidation product, ONE, reacts with free nucleosides of dA, dC, and dG, as well as those in double* To whom correspondence should be addressed. Tel: +1-631-444-3082. Fax: +1-631-444-7641. E-mail:
[email protected]. † University of Pennsylvania. ‡ State University of New York. § Present address: Department of Chemistry, Vanderbilt University, Nashville, TN 37235. 1 Abbreviations: COX, cyclooxygenase; , etheno; ESI, electrospray ionization; H, heptanone; H-dC, H-3,N4-dC; ONE, 4-oxo-2(E)-nonenal; pol, DNA polymerase; XPA, xeroderma pigmentosum complementation group A; XPV, xeroderma pigmentosum variant.
Figure 1. Lipid peroxidation of linoleic acid and arachidonic acid resulting in the production of bifunctional electrophiles and the formation of the heptanone-etheno-DNA base adducts from ONE. EDE, 4,5-epoxy-2(E)-decenal; HNE, 4-hydroxy-2(E)-nonenal; HPNE, 4-hydroperoxy-2(E)-nonenal; LOX, lipoxygenase; 13-HPODE, 13(R,S)hydroperoxy-9Z,11E-octadecadienoic acid; 15-HPETE, 15(R,S)-hydroperoxy-5Z, 8Z, 11Z, 13E-eicosatetraenoic acid.
stranded DNA, to form heptanone (H)-etheno () DNA adducts: H-1,N6-dA (16, 17), H-1,N2-dG (18), and H-3,N4dC (H-dC) (19) (Figure 1). When ONE was reacted in vitro with individual nucleosides, H-dC was produced in the
10.1021/tx0600503 CCC: $33.50 © 2006 American Chemical Society Published on Web 07/14/2006
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Figure 2. Incorporation of H-dC into oligonucleotide: (i) reaction of dC (1) with ONE to generate H-dC (2); (ii) reaction of 2 with DMT-Cl (4,4′-dimethoxytrityl chloride) and DMAP [4-(dimethylamino)pyridine] to yield the DMT-capped adduct 3; (iii) reaction of 3 with 1H-tetrazole and 2-cyanoethyl-tetraisopropylphosphoramidite to produce DMT-capped phosphoramidite (4) for use in DNA synthesizer.
greatest amount (20). Similarly, when DNA was incubated with oxidized linoleic acid or ONE, H-dC was present in the greatest amount, indicating that ONE has the strongest electrophilic potential to dC (20). H-dC was detected in the colon of the Min mouse (C57BL/6JAPCmin), a widely used mouse model of colorectal cancer (21). This mouse strain has a mutation in the adenomatous polyposis coli gene, which causes a large number of colon polyps that frequently have upregulated COX-2 expression (22). Recently, H-dC was also detected in human colorectal cancer tissue samples (Williams, M. V., and Blair, I. A., unpublished observation). In addition, H-dG was detected immunohistochemically in human colonic cancer HCT8 cells undergoing H2O2 treatment (23) and quantified in rat intestinal epithelial cells overexpressing the COX-2 gene (24). Since cells undergoing increased lipid peroxidation could have increased amounts of H--DNA adducts, it is important to characterize their genotoxicity. Toward the goal of revealing the genotoxic mechanism of this adduct, we first synthesized the phosphoramidite of H-dC and incorporated it into an oligonucleotide. Following extensive characterization, this oligonucleotide was inserted site-specifically into a shuttle plasmid for genotoxic studies in cells. The H-dC adduct has been found to show unique genotoxic properties in bacteria and human cells.
Materials and Methods Synthetic Methods. All reagents were purchased from SigmaAldrich (St. Louis, MO) and used without further purification or distillation. Glassware and syringes were dried in an oven overnight and kept in a desiccator until used. Thin-layer chromatography was performed on silica gel glass plates (Silica Gel 60 F254, layer thickness 250 µm; Merck, Aston, PA). The plates were visualized under UV light (254 nm) or by staining with an anisaldehyde/H2SO4 solution, followed by heating. Column chromatography was conducted using silica gel (130-270 mesh, Sigma-Aldrich). DNA synthesis reagents were purchased from Glen Research (Sterling, VA). Synthesis of Heptanone-3, N4-Etheno-dC Phosphoramidite. The synthetic scheme is shown in Figure 2. 2′-Deoxycytidine (1) (185 mg, 0.82 mmol) was reacted with ONE (308 mg, 2 mmol) in a mixture of ethanol and H2O (500 µL/2 mL) at 65 °C for 24 h. The reaction mixture was extracted with CH2Cl2 (1 mL, 3 times) and dried over Na2SO4. CH2Cl2 was evaporated and applied onto a silica gel column (1.5 cm × 10 cm) to collect H-dC (2) (85 mg, yield: 27%). The adduct 2 (85 mg, 0.23 mmol) was
coevaporated with anhydrous pyridine (1 mL, 3 times) and kept under vacuum overnight. 4,4′-Dimethoxytrityl chloride (0.276 mmol) and 4-(dimethylamino)pyridine (0.14 mg) were added to adduct 2 and dissolved in 2 mL of anhydrous pyridine. The mixture was stirred for 3 h at room temperature and then evaporated to dryness. The residue was applied to a silica gel column (2 cm × 13 cm). Excess 4,4′-Dimethoxytrityl chloride was eluted with CH2Cl2, and the column was then washed with 3% methanol/97% CH2Cl2. Gradual elution of the column with CH2Cl2 followed by 5% methanol/95% CH2Cl2 provided product 3, 5′-O-(4,4′-dimethoxytrityl)-4-oxo-2-nonenal-dC (124 mg, yield: 81%). Product 3 (46 mg, 0.069 mmol) was coevaporated with anhydrous pyridine (1 mL, 3 times) for phosphitylation. 1H-Tetrazole (82.8 µmol) and 2-cyanoethyl tetraisopropylphosphoramidite (89.7 µmol) were added to product 3 dissolved in CH2Cl2 and stirred for 3 h at room temperature. The solvent was evaporated, and the residue was purified by flash chromatography under nitrogen (1 cm × 10 cm, 20% dimethyl chloride in ethyl acetate containing 0.1% triethylamine) to collect product 4, 3′-O-[(N,N-diisopropylamino)-(2cyanoethyl)phosphinyl]-5′-O-(dimethoxytrityl)-H-dC (27 mg, yield: 45%). NMR assignment data for the products 2-4 are provided in Figure S1 of the Supporting Information. Synthesis of Modified Oligonucleotides. An oligonucleotide, 5′d(CTCCTCXATACCT) where X represents H-dC, was synthesized by an Applied Biosystems 394 DNA/RNA synthesizer (Foster City, CA) using standard phosphoramidite chemistry with mildly protected bases. The synthesized oligomer was deprotected and released from the glass support by treating with 50 mM K2CO3 in methanol at room temperature for 4-6 h. Following removal of the solvent by evaporation, the 4,4′-dimethoxytrityl-capped oligonucleotide was purified on a Waters HPLC system (Milford, MA) consisting of a 600E Multisolvent Delivery system, a U6K injector, and a 996 Photodiode Array Detector. A Luna 5 µm Phenyl-Hexyl (250 × 10 mm) column (Phenomenex, Torrance, CA) run at a flow rate of 4 mL/min and a gradient of 16 f 36% acetonitrile over 35 min in a 0.1 M triethylammonium acetate buffer, pH 6.8, was used. The 4,4′-Dimethoxytrityl group was removed with 80% acetic acid, and the decapped oligonucleotide was purified with a gradient of 5 f 15% acetonitrile over 30 min. The main peaks were collected as a single fraction. The modified 13-mer eluted as a single peak at 16.1 min with λmax at 265.8 nm (Supporting Information Figure S2). The modified and unmodified oligonucleotides were finally purified by polyacrylamide gel electrophoresis in a denaturing 20% gel. Subsequent analyses confirmed that they migrated as single bands in the gel. This combination of HPLC and gel purification showed that the purity of the modified oligonucleotide was >99.5%.
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Figure 3. Negative ESI analysis of H-dC-containing 13-mer.
The purified modified 13-mer was characterized by a combination of negative electrospray ionization (ESI) mass spectrometry and enzymatic digestion to nucleosides followed by an HPLC analysis. Mass spectrometry analyses were performed using a Micromass LCZ Platform mass spectrometer (Waters). Samples were diluted with 40% acetonitrile/water to a concentration of ∼10 pmol/µL, then introduced into the ESI source. The source was coupled to a syringe pump (Harvard Apparatus, Inc., South Natick, MA) supplying 40% acetonitrile/water containing 1% triethylamine at 12 µL/min. Nitrogen was used both as a drying gas and for nebulization. Deconvolution of the molecular negative ion envelopes corresponding to [M - H]- was performed using proprietary Mass Lynx v 3.0 software (Waters). [M - H]- for the modified 13-mer was determined to be 3941.34 ( 0.73 Da from the multiply charged ions in the mass spectrum (Figure 3). The calculated mass of [M - H]- was 3940.72 Da, indicating a mass accuracy of 0.016%. For the enzymatic digestion, 2.8 nmol of modified 13-mer was sequentially digested as follows at 37 °C for 20 min per step: (1) 280 units of DNase I (Sigma-Aldrich) in 25 µL of 10 mM TrisHCl/100 mM MgCl2, pH 7.4; (2) addition of 0.5 units of phosphodiesterase (Roche; Nutley, NJ) in 25 µL of 0.2 M glycine, pH 10; and (3) addition of 5-10 units of shrimp alkaline phosphatase (Roche) in 50 µL of 50 mM Tris-Cl/50 mM MgCl2. The solution was filtered through a Costar Spin-X filter (0.22 µm;
Pollack et al. Corning, Corning, NY) and analyzed by HPLC with a Phenomenex Jupiter (250 × 4.6 mm, 5 µm, C5) column. HPLC was performed using a Hitachi L-7100 pump with 5 mM NH4OAc in H2O as solvent A and 5 mM NH4OAc in CH3CN as solvent B. A flow rate of 1 mL/min was used, and solvent B was increased from 0 to 10% over 25 min. Peaks were monitored by UV-visible spectrophotometry. H-dC was observed at a retention time of 17.2 min (Supporting Information, Figure S3), which was identical with that of an authentic standard. Cell Lines. SV40-transformed human DNA repair-proficient (GM637) and xeroderma pigmentosum complementation group A (XPA) cells (GM04429) were obtained from the Coriell Institute (Camden, NJ). XP variant, CTag (25), was obtained from M. Cordeiro-Stone (University of North Carolina, Chapel Hill, NC). Cells were cultured in Dulbecco’s modified Eagle’s medium supplemented with 10% fetal bovine serum, penicillin (100 units/ mL), and streptomycin (100 µg/mL) at 37 °C under 5% CO2. Escherichia coli DH10B [F-, mcrA, ∆(mrr-hsdRMS-mcrBC), φ80lacZ∆M15, ∆lacX74, deoR, recA1, endA1, ara∆139, ∆(ara, leu)7697, galU, galK, λ-, rpsL, nupG, tonA)] (Invitrogen, Carlsbad, CA) was also used. Genotoxic Studies. The shuttle vector pBTE has been described previously (26). This vector is stably maintained in human cells and confers blasticidin S resistance on human and E. coli cells. The expression of this resistance gene is driven by the SV40 early promoter in human cells and the EM7 bacterial promoter in E. coli. Construction of the double-stranded plasmid containing a sitespecific DNA adduct has been described in detail previously (27). The procedure is shown schematically in Figure 4 together with the experimental strategy. H-dC was incorporated into the leading strand template. An important feature of this construct is that the adduct was inserted opposite a unique SnaBI site (5′ TACGTA) with mismatches on both sides of the adduct (highlighted, Figure 4B); therefore, only the unmodified complementary strand contained the restriction enzyme site. Progeny plasmid derived from the unmodified strand was sensitive to SnaBI digestion, whereas that derived from translesion synthesis events was resistant. Hence,
Figure 4. Site-specific experiment. Outline of experimental procedure (A), construction of modified plasmid (B), and oligonucleotide probes used for sequence analysis of progeny plasmid (C). In panels B and C, note mismatches at the sequence of 3′-AXC (highlighted). X represents H-dC. Probe S (overscored) hybridizes only to unmodified strand. Probes L and R detect plasmid containing a 13-mer insert. Probes G, T, A, C, and D determine targeted events.
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Table 1. Translesion Events in E. coli and Human Cells
host E. coli DH10B XPA GM637 XPV
exp. no.
no. of transformants or transfectantsa
progeny (%)b
C
1 2 1 2 1 1
∼9 × 107 ∼9 × 107 2 × 104 4 × 104 3 × 104 1 × 104
92%) A or T in XPA cells (Table 1). The same preference was observed in the wild-type GM637 and XPV human cells. Accordingly, the miscoding frequency was extremely high in human cells, reaching more than 90%, and it was 40-50% in E. coli. In E. coli, the correct dG insertion was somewhat more frequent than was dC insertion. In human cells, dT insertion was more dominant over dA insertion. Thus, the coding
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Figure 5. Efficiency of translesion synthesis in XPA cells for dG, γ-OH-PdG, R-OH-PdG, PdG, 1,N6-dA, and H-dC. Progeny plasmid was recovered from transfected cells, digested with Dpn I, and used to transform E. coli. E. coli transformants were hybridized to oligonucleotide probes shown in Figure 4C to determine the strand from which progeny was generated. The y-axis shows percent of progeny derived from the adduct-containing strand. The result for dA is unpublished, whereas results for the other adducts have been published (27). PdG is an abbreviation for 1,N2-(1,3-propano)-2′-deoxyguanosine, and refer to ref 27 for their chemical names.
specificity of this adduct is totally different in the two hosts, and the miscoding events were mostly H-dC f dA or dT in human cells and H-dC f dG in E. coli. The preferential insertion of dG or dC was also observed in other E. coli strains of MM1933 (As MV1932 (32), but mutS201::Tn5) and MM1934 (As MM1933, but uVrA6, malE3::Tn10). This analysis was conducted on a small scale and data are not presented.
Discussion H-dC has an exocyclic ring and a bulky heptanone side chain (Figure 1), both of which disrupt Watson-Crick hydrogen bonding to dG, and hence, DNA synthesis was expected to be blocked strongly. We also expected the adduct to be miscoding like the unsubstituted dC (33). These expectations proved to be true. H-dC is the strongest blocking lesion among various exocyclic lesions thus far examined by this system (Figure 5). H-dC markedly miscodes in both E. coli and human cells. Since the preferential insertion of dG or dC in E. coli and of dA or dT in human cells was observed in different host strains, this phenomenon may be true. We have also examined the effects of SOS induction in E. coli, but the enhancing effects on the efficiency of translesion synthesis were not clearly observed when double-stranded DNA was used (data not shown). A study with single-stranded DNA will be necessary to reveal the effects. The drastic difference in the coding specificities in the two hosts is likely due to the difference of DNA polymerases involved in the translesion synthesis. Since the host E. coli was not SOS-induced, the preferential insertion of dG or dC might be catalyzed by pol III during its processive synthesis mediated by β clamps or SOS polymerases, such as pol II, pol IV, and pol V, present in a residual amount in cells. The experiments using single- or multiple-SOS pol-defective E. coli hosts will reveal the role for those SOS pols in this coding specificity. Among mammalian translesion-specialized pols, the polymerase activities of pol η, pol κ, and pol ζ have proven to be important in cells (34-39). In the present study, we also used XPV pol η-deficient cells. XPV cells showed similar translesion synthesis efficiency and coding specificity to those observed in the wild-type GM637 cells, suggesting that this polymerase
Pollack et al.
is not critical to this translesion synthesis. Unfortunately, human cells defective in the other translesion-specialized polymerases are not available, inhibiting a systematic study. In this regard, we are modifying the current vector so that gene-knockout mouse cells can be used as hosts. We had better yields of progeny derived from the modified strand in the excision repairwild XPV and GM637 than in XPA, indicating that replication took place before DNA repair operated. The repair converts the sequence of the adducted region to that complementary to the unmodified strand. Unexpectedly, the fractions of progeny derived from the modified stand in these two cell lines were higher than that in the repair-deficient XPA cells (Table 1). We do not have a good explanation for this finding at present, but the same phenomenon has been observed with a different DNA adduct (submitted). The ability to conduct translesion synthesis might be reduced in this XPA cell line. H-dC has recently been quantified in mouse (21) and human colorectal tissue (unpublished results) that overexpresses the COX-2 enzyme, an enzyme known to be upregulated in colorectal cancer (40) and some breast cancers (41, 42). The tumor-promoting properties of COX-2 are thought to derive from increased prostaglandin synthesis (43, 44); however, we have shown that COX-2 may promote tumorigenesis through an alternative pathway, that is, DNA adduct formation via COX2-mediated lipid peroxidation (24). Upregulation of COX-2 in colon and breast cancers would increase lipid peroxidation, resulting in an increased formation of DNA adducts such as the H--DNA adducts. As shown here, H-dC is a strong block to DNA synthesis, which likely leads to the formation of double-strand breaks and thereby chromosomal aberrations. When this adduct is bypassed, it directs misinsertion at a very high frequency. Thus, H-dC exerts such significant deleterious genotoxic effects in human cells that removal of the lesion from the genome is critical before DNA replication takes place. Therefore, the detection of unrepaired H-dC adducts in genomic DNA has significance in considering a carcinogenic role of this lesion in COX-2-upregulated cancers. Acknowledgment. We thank Dr. Sivaprasad Attaluri (SUNY, Stony Brook) for incorporating the H-dC adduct into oligonucleotides. This work was supported by the Solomon Erulkar Traveling Fellowship and NIH grants CA91096, CA76163, and CA47995. Note Added after ASAP Publication. There were errors in Figures 1 and 2 in the version published ASAP July 14, 2006. The correct versions were published July 18, 2006. Supporting Information Available: Characterization of H-dC phosphoramidite and modified oligonucleotides. This material is available free of charge via the Internet at http://pubs.acs.org.
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