Oxidation of Thymine to 5-Formyluracil in DNA Promotes Misincorporation of dGMP and Subsequent Elongation of a Mismatched Primer Terminus by DNA Polymerase*

Aya Masaoka, Hiroaki Terato, Mutsumi Kobayashi, Yoshihiko Ohyama, and Hiroshi IdeDagger

From the Department of Mathematical and Life Sciences, Graduate School of Science, Hiroshima University, Higashi-Hiroshima 739-8526, Japan

Received for publication, September 20, 2000, and in revised form, December 11, 2000


    ABSTRACT
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
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DISCUSSION
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5-Formyluracil (fU) is a major oxidative thymine lesion generated by ionizing radiation and reactive oxygen species. In the present study, we have assessed the influence of fU on DNA replication to elucidate its genotoxic potential. Oligonucleotide templates containing fU at defined sites were replicated in vitro by Escherichia coli DNA polymerase I Klenow fragment deficient in 3'-5'-exonuclease. Gel electrophoretic analysis of the reaction products showed that fU constituted very weak replication blocks to DNA synthesis, suggesting a weak to negligible cytotoxic effect of this lesion. However, primer extension assays with a single dNTP revealed that fU directed incorporation of not only correct dAMP but also incorrect dGMP, although much less efficiently. No incorporation of dCMP and dTMP was observed. When fU was substituted for T in templates, the incorporation efficiency of dAMP (fA = Vmax/Km) decreased to 1/4 to 1/2, depending on the nearest neighbor base pair, and that of dGMP (fG) increased 1.1-5.6-fold. Thus, the increase in the replication error frequency (fG/fA for fU versus T) was 3.1-14.3-fold. The misincorporation rate of dGMP opposite fU (pKa = 8.6) but not T (pKa = 10.0) increased with pH (7.2-8.6) of the reaction mixture, indicating the participation of the ionized (or enolate) form of fU in the mispairing with G. The resulting mismatched fU:G primer terminus was more efficiently extended than the T:G terminus (8.2-11.3-fold). These results show that when T is oxidized to fU in DNA, fU promotes both misincorporation of dGMP at this site and subsequent elongation of the mismatched primer, hence potentially mutagenic.


    INTRODUCTION
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Faithful replication of DNA is essential for maintaining genetic integrity of living organisms. High fidelity of DNA replication is achieved by two cellular functions that involve discrimination of correct versus incorrect nucleotides by DNA polymerases (1, 2) and postreplication mismatch repair (3). The overall error frequency of DNA replication is one in 108 to 1010 base pairs when they function properly. Fidelity of DNA replication also relies on the structural integrity of DNA itself that serves as a template for the newly synthesized strand. A number of endogenous and exogenous agents have been identified to induce structural deterioration of DNA (4). Among them, reactive oxygen species generate a very complicated spectrum of DNA damage (5, 6). These lesions are mostly restored by the base excision repair pathway both in prokaryotic and eukaryotic cells, but if left unrepaired, they arrest DNA synthesis or direct misincorporation of nucleotides during DNA replication, hence exerting deleterious effects on cells (7, 8). Replication blocks and nucleotide misincorporation have been related to lethality and mutation of cells, respectively, until recently. However, this concept is now challenged by the discovery of numerous error-prone and error-free DNA polymerases that can bypass the blocking lesions (9).

Although the past several years have witnessed the discovery of novel lesion replicating DNA polymerases (mentioned above) as well as remarkable progress in understanding the molecular basis for the nucleotide discrimination mechanism by DNA polymerases (10, 11), the assessment of genotoxic effects of structurally diverse oxidative DNA damage largely relies on experimental data obtained from defined lesions (7, 8, 12, 13). We have been studying the response of DNA polymerases to the encountered oxidative thymine (13-17) and other base (18-21) lesions. Oxidative thymine lesions formed by ionizing radiation, Fenton-type reactions, and photosensitized reaction have been best characterized among the four DNA bases and can be classified into four subgroups depending on their structural features. The first group includes C-5---C-6 saturation products such as thymine glycol (5,6-dihydroxy-5,6-dihydrothymine) and 5,6-dihydro-5-hydroxythymine. 5,6-Dihydrothymine belongs to this group, although it is a reduction product formed by ionizing radiation. The second group is ring fragmentation products such as a urea residue and its analogues. The response of DNA polymerases to the first and second groups has been clarified fairly well by in vitro and in vivo studies (13-15, 22, 23). The third group includes 5-hydroxy-5-methylhydantoin, a ring contraction product. The ability of this lesion to block DNA replication has been demonstrated recently (24) by in vitro DNA polymerase reactions using a defined oligonucleotide template. The fourth group contains methyl oxidation products such as 5-hydroxymethyluracil and 5-formyluracil (fU).1 Several lines of evidence indicate that 5-hydroxymethyluracil is neither a replicative block nor mutagenic (25, 26) and hence is an innocuous lesion.

The genotoxic potential of fU belonging to the fourth group has been assessed in this (27, 28) and other (29, 30) laboratories. In our previous approach, we synthesized 5-formyl-2'-deoxyuridine 5'-triphosphate (fdUTP) and studied its incorporation into DNA by DNA polymerases. fdUTP efficiently substituted for dTTP and to a much less extent for dCTP. Moreover, the pH-dependent variation of the substitution efficiency for dCTP suggested involvement of an ionized (or enolate) form of fU as a key intermediate responsible for the mispairing with template G. Such a mutation mechanism involving ionized bases (thymine and 5-bromouracil (BrU)) was originally suggested by Lawley and Brookes (31). Later, the pH-dependent variation of the replication error frequency due to ionization of BrU and 5-fluorouracil was experimentally demonstrated by Yu et al. (32), providing the conceptional basis of the previous studies (27, 28). Translesion bypass and nucleotide incorporation at the site of template fU were also studied (29, 30) using oligonucleotides containing the site-specific lesion (33). Consistent with very efficient substitution of fdUTP for dTTP in our study, DNA polymerase readily passed through the fU site in the template. However, to our surprise, the primer extension study showed that fU directed incorporation of dCMP as well as correct dAMP, implying the formation of an fU:C mispair during DNA replication. The discrepancy between the two studies concerning the base pairing capacity of fU might have originated from several reasons. First, although overall Watson-Crick geometry of a newly formed base pair plays a dominant role in nucleotide selection by DNA polymerases (1, 2), this process can also be affected by base pairing symmetry whether an X:Y base pair is formed from X (template):Y (dNTP) or X (dNTP):Y (template) (34-36). Second, the sequence context can affect the selection of dNTP opposite the template lesion (7). Third, the base ionization mechanism somehow does not hold when fU is present in template DNA.

In view of the potential influences of base pairing asymmetry, the sequence context, and deviations from the base ionization mechanism mentioned above, we have prepared oligonucleotide templates containing site-specific fU following the previously reported phosphoramidite method (33), and we reexamined the base pairing capacity of template fU with the four possible nearest neighbor base pairs. The results show that fU in the template directs misincorporation of dGTP in a pH-dependent manner, supporting our previous results obtained by the analysis of fdUTP incorporation.

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INTRODUCTION
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Chemicals and Enzymes-- Ultra-pure dATP, dGTP, dCTP, and dTTP (purity >99.3%) were purchased from Amersham Pharmacia Biotech. 5-Formyl-2'-deoxyuridine (fdU) was synthesized as described previously (27). The phosphoramidite monomer of protected 5-(1,2-dihydroxyethyl)-2'-deoxyuridine was synthesized following the reported procedure (33). [gamma -32P]ATP (110 TBq/mmol) was purchased from Amersham Pharmacia Biotech. Escherichia coli DNA polymerase I Klenow fragment (Pol I Kf), Pol I Kf deficient in 3'-5'-exonuclease [Pol I Kf (exo-)], and T4 polynucleotide kinase were obtained from New England Biolabs, and Penicillium citrium nuclease P1 and calf intestine alkaline phosphatase were from Roche Molecular Biochemicals.

Oligonucleotides-- Oligonucleotides comprising normal components were synthesized by the standard phosphoramidite method and purified by reversed phase HPLC. Oligonucleotides containing fU were prepared following the reported procedure (33). First, oligonucleotides containing 5-(1,2-dihydroxyethyl)uracil, a precursor of fU, were synthesized by the standard phosphoramidite chemistry using a phosphoramidite monomer of protected 5-(1,2-dihydroxyethyl)-2'-deoxyuridine and purified by reversed phase HPLC. The oligonucleotides containing 5-(1,2-dihydroxyethyl)uracil was treated by sodium periodate to convert 5-(1,2-dihydroxyethyl)uracil to fU. After treatment, crude oligonucleotides were desalted by passing through a Sephadex G-10 column and further purified by reversed phase HPLC. The sequences of oligonucleotides used in the present study are listed in Table I.

                              
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Table I
List of oligonucleotides used in this study

Composition Analysis of Oligonucleotides-- 25T and 25F (0.2 OD) were incubated with nuclease P1 (1 unit) in a reaction buffer (60 µl) containing 21 mM sodium acetate (pH 5.3) and 1 mM ZnSO4 at 37 °C for 1 h. To this solution, alkaline phosphatase buffer (30 µl) comprising 0.5 M Tris-HCl (pH 9.0) and 10 mM MgCl2 and alkaline phosphatase (3 units) were added, and the reaction mixture was further incubated at 37 °C for 2 h. The sample was passed through a molecular weight cut-off filter (Mr = 10,000), and an aliquot of the filtrate was analyzed by HPLC equipped with a C18 WS-DNA column (4.6 × 150 mm, Wako). The sample was eluted by a gradient of methanol in 10 mM sodium phosphate buffer (pH 7.4) at a flow rate 0.8 ml/min. The concentration of methanol was 0% for 0-5 min and 0-5% linear gradient for 5-35 min. The column temperature was maintained at 40 °C by a column oven, and eluents were monitored at 280 nm.

Treatments with Repair Enzymes-- 25F was 5'-end-labeled as described for the primers (see below) and annealed to the complementary strand (25COM). 25F/25COM (0.02 pmol) was incubated with AlkA (1 pmol) followed by endonuclease (Endo) IV (0.03 pmol) in a reaction buffer (10 µl) at 37 °C. Alternatively, the substrate was incubated with Endo III (0.3 pmol), formamidopyrimidine glycosylase (Fpg, 0.3 pmol), or Endo IV (0.03 pmol) in a similar manner. The procedures of the enzymatic treatments were essentially similar to those reported previously for AlkA (28, 37), Endo III (38), Fpg, and Endo IV (39). After incubation, products were analyzed by 16% denaturing PAGE.

Preparation of Template-Primer-- The primers were 5'-end-labeled using T4 polynucleotide kinase and [gamma -32P]ATP. Appropriate template and primer (molar ratio = 2:1) were annealed in 20 mM Tris-HCl (pH 7.5) and 50 mM NaCl by heating the solution at 90 °C for 5 min and allowing to cool slowly to room temperature.

Analysis of Translesion DNA Synthesis-- The annealed template-primer (0.5 pmol) containing T (24AT/P10 and 24CT/P10) or fU (24AF/P10 and 24CF/P10) at the same position was incubated with Pol I Kf (exo-) (0.05 unit) and four dNTPs (50 µM each) in a polymerase reaction buffer (15 µl) at 25 °C for 3-10 min. The polymerase reaction buffer consisted of 66 mM Tris-HCl (pH 7.5), 1.5 mM 2-mercaptoethanol, and 6.6 mM MgCl2.

Analysis of Nucleotides Incorporated Opposite fU-- To determine the nucleotide incorporated opposite fU, primer extension reactions were performed in a manner essentially similar to that described for translesion DNA synthesis except that the reaction mixture contained a single dNTP (50 µM) and incubation time was 5 min. The template-primer used in the analysis was 24AF/13T, 24GF/13C, 24CF/13G, and 24TF/13A that contained fU at the same position and different nearest base pairs next to fU (i.e. primer terminus base pairs). For control reactions, primer extension assays were also performed using template-primers (24AT/13T, 24GT/13C, 24CT/13G, and 24TT/13A) that contained T in place of fU. By using these template-primers containing fU or T, kinetic parameters of nucleotide incorporation opposite fU and T were also determined. For dATP incorporation, the dATP concentration was 0.05-1 µM, and the amount of Pol I Kf (exo-) was 0.002 unit, whereas for dGTP incorporation, those were 10-80 µM and 0.03 unit. The incubation time was 5 min for both dATP and dGTP incorporation assays. Under these conditions, the extent of primer elongation was essentially proportional to the reaction time and the unit of Pol I Kf (exo-) used. The initial velocity of the reaction (V) (average of two experiments) was calculated as the percentage of the extended primer per min per 0.03 unit of Pol I Kf (exo-). Km and Vmax values were evaluated from a hyperbolic curve fitting program.

The pH effect on the incorporation of dATP and dGTP opposite template fU or T was determined by varying the pH (pH 7.2-8.6) of the polymerase reaction buffer as described above. The template-primer (24AF/13T and 24AT/13T, 0.5 pmol) was incubated with Pol I Kf (exo-) (0.03 unit) and dATP or dGTP (20 µM) at 25 °C for 5 min. For a wide pH range (pH 6.9-9.3), GTA buffer (buffering capacity pH 3.5-10) was used in place of the Tris buffer for the DNA polymerase reaction. The composition of GTA buffer was 3,3-dimethylglutaric acid, Tris, and 2-amino-2-methyl-1,3-propanediol (17.3 mM each), and pH was adjusted by adding HCl or NaOH.

Extension of Mismatched Primer Termini-- Template-primers (0.5 pmol) containing a correctly paired (24AT/14TA and 24AF/14TA) or mismatched (24AT/14TG and 24AF/14TG) primer terminus were incubated with Pol I Kf (exo-) (0.05 unit) and dCTP (10 µM) in the polymerase reaction buffer (15 µl) at 25 °C for 5 min. Alternatively, kinetic parameters of the reaction (average of two experiments) were determined by varying the dCTP concentration between 0.01 and 1 µM for the correctly paired primer terminus (24AT/14TA and 24AF/14TA) or 0.01-20 µM for the mispaired primer terminus (24AT/14TG and 24AF/14TG). The reaction was performed as described above except that the amount of Pol I Kf (exo-) was 0.015 unit. The parameters for mismatch extension with templates 24CT and 24CF were also determined in the same manner.

Electrophoresis-- DNA polymerase reactions were terminated by adding loading buffer containing 0.1% xylene cyanol, 0.1% bromphenol blue, 20 mM EDTA, and 95% formamide. The sample was boiled and subjected to 16% denaturing polyacrylamide gel electrophoresis (PAGE). Electrophoresis was performed at 1800 V, and gel was autoradiographed at -80 °C overnight. The radioactivity of the separated bands was quantified on a PhosphorImager Fuji Bas 2000.

    RESULTS
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
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Nucleoside Composition of Oligonucleotides-- To ensure the validity of phosphoramidite method used in the present preparation of oligonucleotides containing fU, pilot oligonucleotides containing T (25T) and fU (25F) in the same sequence were prepared. 25T and 25F were digested by nuclease P1 and alkaline phosphatase, and the nucleoside composition was analyzed by HPLC. Digestion of 25T resulted in the HPLC peaks of dC, dG, dT, and dA with an expected molar ratio (8:6:4:7) (data not shown). In the HPLC analysis of the digested 25F, two extra peaks were observed in addition to the four normal nucleosides (Fig. 1A). The first peak eluted at 15.1 min was readily identified as 5-formyl-2'-deoxyuridine (fdU) by comparison with the retention time of authentic fdU. The peak at 32.4 min (indicated by *) was an unknown product. When authentic fdU was incubated under the same conditions as those used for oligonucleotide digestion, fdU was partially converted to this product (Fig. 1B). The formyl group of fdU is fairly reactive and forms adducts with nucleophiles (40). Accordingly, this product is most likely an adduct between fdU and a nucleophilic molecule present in the reaction buffer or enzyme preparations. The long HPLC retention time of the product relative to fdU and retention of the UV absorption around 280 nm were also consistent with the adduct formation of the exocyclic formyl group of fdU. The fU moiety of fdU is known to be degraded by strong base and oxidizing reagents, giving rise to ring fragmentation products (27, 41) and 5-carboxyuracil (42), respectively. If such products were formed during the preparation of 25F, they might be present as contaminated lesions in 25F. However, the retention of the UV absorption (around 280 nm) of the product was inconsistent with the ring fragmentation products bearing no chromophores. That retention time of the product (32.4 min) was much longer than fdU (15.1 min) in the reversed phase HPLC column also contradicted the expected very short retention time of 5-carboxy-2'-deoxyuridine bearing a negative charge of a carboxylate ion. Thus, the product was not the ring fragmentation products or 5-carboxy-2'-deoxyuridine. Moreover, when the amount of authentic fdU converted to the putative adduct was taken into account, the corrected molar ratio of nucleosides in 25F agreed with the expected value (dC:dG:dT:dA:fdU = 8:6:3:7:1).


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Fig. 1.   HPLC analysis of nucleosides in 25F. A, 25F containing fU was digested by nuclease P1 and alkaline phosphatase as described under "Experimental Procedures." An aliquot of the reaction mixture was analyzed by reversed phase HPLC equipped with a C18 WS-DNA column (4.6 × 150 mm). The sample was eluted by a gradient of methanol (0% for 0-5 min and 0-5% linear gradient for 5-35 min) in 10 mM sodium phosphate buffer (pH 7.4). The flow rate was 0.8 ml/min, and the monitoring wavelength was 280 nm. The attribution of the elution peak is indicated above the peak. The elution peak with an asterisk was a putative fdU adduct formed by the enzymatic treatment. B, authentic fdU was treated by nuclease P1 and alkaline phosphatase and subjected to HPLC analysis as described above. Note that fdU before incubation was eluted as a single peak (not shown) and no adduct (*) was observed.

We also attempted to identify the structure of the unknown product by mass spectrometric analysis after isolating the product by HPLC. However, the attempt was unsuccessful because of the lack of the apparent molecular ion (M+) in the spectrum. In an alternative approach, 25F (as a duplex) was digested by several DNA repair enzymes with different damage specificities. Consistent with the previous reports (28, 37), the treatment with AlkA followed by Endo IV resulted in incision of 25F at the fU site (data not shown). In contrast, neither Endo III, Fpg, nor Endo IV incised 25F,2 supporting the absence of base damage other than fU in 25F. On the basis of the results from the composition analysis and the treatment with repair enzymes, we concluded that fU was successfully incorporated into oligonucleotides in the present procedure of synthesis.

Translesion DNA Synthesis at the fU Site-- To clarify whether fU present in template DNA constitutes a replication block, 24AF and 24AT containing fU and T, respectively, at the same site (4 nucleotides beyond the primer terminus) were primed by 32P-labeled P10, and the templates were replicated by Pol I Kf (exo-) for up to 10 min. The resulting products were analyzed by denaturing PAGE (Fig. 2). After 3 min of incubation, the primer annealed to the undamaged template 24AT was almost completely extended to a fully replicated product (lane 2). Similarly, the primer annealed to 24AF containing fU was mostly extended to fully replicated and 1 nucleotide shorter products after 3 min of incubation (lane 6). In addition, very weak bands also appeared at and 1 nucleotide before the fU site, indicating a pause of DNA synthesis at these sites. Quantification of the arrested and bypassed products showed that 91% of the original primer was extended beyond the fU site at 3 min. This result indicates that fU in template DNA allows efficient translesion DNA synthesis. Similar results were obtained with templates 24CF and 24CT (data not shown).


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Fig. 2.   Translesion DNA synthesis at the fU site. DNA templates containing T (24AT) and fU (24AF) at the same site were primed with 32P-labeled P10. 24AT/P10 and 24AF/P10 (0.5 pmol) was incubated with Pol I Kf (exo-) (0.05 unit) and four dNTPs (50 µM each) at 25 °C, and products were analyzed by 16% denaturing PAGE. Templates (24AT and 24AF) and incubation times (3, 5, and 10 min) are indicated on the top. Lanes 1 and 5, template-primer without the polymerase reaction; lane 9, 14-mer marker (32P-labeled 14TA in Table I) showing the position of fU.

Nucleotides Incorporated Opposite fU-- Since efficient translesion DNA synthesis occurred at the fU site, the nucleotide incorporated opposite this lesion was analyzed by a primer extension assay. In this assay, primers (13T, 13C, 13G, and 13A) that were 1 nucleotide shorter than the template fU site were annealed to appropriate templates (24AF, 24GF, 24CF, and 24TF), and the primers were extended by Pol I Kf (exo-) in the presence of a single dNTP (50 µM) at 25 °C for 5 min. These template-primers (24AF/13T, 24GF/13C, 24CF/13G, and 24TF/13A) contained four different nearest neighbor base pairs in the primer terminus. Control experiments were also performed under the same conditions using template-primers (24AT/13T, 24GT/13C, 24CT/13G, and 24TT/13A) that contained T instead of fU. The reaction products were analyzed by denaturing PAGE. Fig. 3A shows PAGE data obtained for 24AF/13T and 24AT/13T containing an A (template):T (primer) pair at the primer terminus. According to the band intensity of the extended product (14-mer), dAMP was most efficiently incorporated opposite fU (lane 6) as well as T (lane 1), with a preference for T. The bands indicative of misincorporation of dGMP opposite fU (lane 7) and T (lane 2) were also observed, but the incorporation was more efficient for fU than T. In the presence of dCTP and dTTP, extended products were not observed over the background for both templates containing fU (lanes 8 and 9) and T (lanes 3 and 4), showing that the misincorporation frequency of dCMP and dTMP was below the detection limit under these conditions. Essentially similar results were obtained with other template-primers containing G:C (Fig. 3B), C:G (Fig. 3C), and T:A (Fig. 3D) as the nearest neighbor base pairs. Accordingly, fU directed incorporation of correct dAMP and to a less extent incorrect dGMP but not pyrimidine nucleotides (dCMP and dTMP).


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Fig. 3.   PAGE analysis of the nucleotide incorporated opposite fU. Template-primers (0.5 pmol) containing four different primer terminus base pairs were incubated with the indicated dNTP (50 µM) and Pol I Kf (exo-) (0.05 unit) at 25 °C for 5 min, and the incorporated nucleotide opposite X (= T or fU) was analyzed by 16% denaturing PAGE. Template-primer used was 24AT/13T (lanes 1-4) and 24AF/13T (lanes 6-9) (A), 24GT/13C (lanes 1-4) and 24GF/13C (lanes 6-9) (B), 24CT/13G (lanes 1-4) and 24CF/13G (lanes 6-9) (C), 24TT/13A (lanes 1-4) and 24TF/13A (lanes 6-9) (D). A-D, lane 5 shows template-primer without the polymerase reaction. The arrow indicates the extended primer. The sequence surrounding T or fU (indicated by X) is shown above the gel.

Parameters of dAMP and dGMP Incorporation-- For quantitative analysis of the nucleotide incorporation efficiency and the sequence context effect, kinetic parameters of dAMP and dGMP incorporation opposite fU and T were determined by the gel fidelity assay under standing start conditions (43). The experiments were performed as described under "Experimental Procedures" using a set of template-primer employed above (see under "Nucleotides Incorporated Opposite fU"). Table II summarizes the parameters (Vmax and Km) and efficiencies (fA Vmax/Km) of dAMP incorporation (average of two experiments). Although there were variations depending on the nearest neighbor base pair, the Vmax values for dAMP incorporation were consistently higher for T than fU. Conversely, the Km values for T were consistently lower than for fU. Consequently, the incorporation efficiency of dAMP (fA) opposite fU was reduced to 1/4 to 1/2 of that opposite T. The efficiency difference between T and fU was not large but significant, showing that conversion of T to fU in template DNA slows down incorporation of the correct nucleotide dAMP. Table III summarizes the parameters (Vmax and Km) and efficiencies (fG = Vmax/Km) of dGMP misincorporation together with the replication error frequencies (fRE = fG/fA) (average of two experiments). With the same nearest neighbor base pair, the Vmax values for dGMP misincorporation were consistently higher for fU than T. However, the Km values for fU showed no systematic variations. Despite these variations, misincorporation of dGMP was favored for fU over T by 1.1-5.6-fold as judged from relative fG. Granting competitive incorporation of dAMP and dGMP at the same site (T or fU) of DNA, the replication error frequency (fRE fG/fA) was calculated for individual template-primers using the data in Tables II (fA) and III (fG). The values of fRE for fU were consistently in a 10-4 range, whereas those for T were in a 10-5 range. Thus, the increase in fRE due to the substitution of fU for T was 3.1-14.3-fold (Fig. 4A). These increases arose from reduced fA (Table II) and increased fG (Table III). Comparison of the parameters (Vmax and Km) in Tables II and III also indicated that discrimination of the nucleotide at the fU site originated from Vmax and Km. The averaged discrimination factors for Vmax and Km were 16 and 360, respectively, which were calculated from the ratio of the parameters for dAMP versus dGMP incorporation. Therefore, the contribution of Km was much greater than that of Vmax.

                              
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Table II
Parameters (Vmax and Km) and efficiencies (fA) of dAMP incorporation opposite template T and fU

                              
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Table III
Parameters (Vmax and Km) and efficiencies (fG) of dGMP misincorporation opposite template T and fU, and replication error frequencies (fRE)


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Fig. 4.   Sequence context effects on fA, fG, fRE, and increases in fRE and fEX associated with conversion of T to fU. A, increases in the replication error frequency (fRE) associated with conversion of T to fU. The ratio of the replication error frequencies (fRE (X = fU)/fRE (X = T)) was calculated for the same 3'-nearest neighbor base using the data in Table III. The ratio was plotted against the template sequence (3'-NX-5', N = A, G, C, and T, X = T and fU). B, increases in the mismatch extension frequency (fEX) associated with conversion of T to fU. The ratio of the misextension frequencies (fEX (X = fU)/fEX (X = T)) was calculated for the corresponding templates (i.e. 24AF versus 24AT and 24CF versus 24CT) using the data in Table IV. The ratio was plotted against the template sequence (3'-NXG-5', N = A and C, X = T and fU). C, sequence context effects on the incorporation efficiencies of dAMP (fA). D, sequence context effects on the misincorporation efficiencies of dGMP (fG). E, sequence context effects on the replication error frequency (fRE). C-E, the values of fA, fG, and fRE were taken from Tables II and III and plotted against the template sequence (3'-NT-5' and 3'-NF-5', N = A, G, C, and T, F = fU).

pH Effects of dAMP and dGMP Incorporation Opposite fU-- We have previously reported pH-dependent misincorporation of fdUTP opposite template G by DNA polymerase, and we have pointed out the importance of an ionized (or enolate) form of fU in fU:G mispair formation (27). To ask whether this mispairing scheme also held for fU in template DNA, the pH effect on the dGMP misincorporation was analyzed. Template-primers containing fU (24AF/13T) or T (24AT/13T) were incubated with Pol I Kf (exo-) and a single dNTP (dGTP or dATP, 20 µM) at pH 7.2-8.6. The percentage of the extended primer resulting from incorporation of dAMP or dGMP was determined by PAGE analysis (Fig. 5). Incorporation of dAMP was virtually unaffected by the pH change and was less efficient for fU than T (Fig. 5, A and C). In contrast, incorporation of dGMP opposite fU showed a clear pH dependence and the amount of dGMP increased with increasing pH (Fig. 5, B and D). Although dGMP incorporation opposite T was also pH-dependent, the increase with pH was extremely small (Fig. 5, B and D). The pH effect on dGMP misincorporation opposite fU was further analyzed in a wider pH range (pH 6.9-9.3) using GTA buffer in place of Tris buffer (Fig. 5D, inset). The plot of the efficiency of dGMP misincorporation against pH showed a sigmoidal curve reminiscent of a pH titration, although the efficiency was somewhat different between the GTA and Tris buffer systems. Since the pKa values of fU and T were 8.6 and 10.0, respectively, these results strongly suggest that an acid-base equilibrium of fU (Fig. 6A) is involved in the misincorporation of dGMP, and the ionized (or enolate) form of fU forms a mispair with incoming dGTP during DNA synthesis (Fig. 6B, left).


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Fig. 5.   pH effects on the incorporation of dAMP and dGMP opposite fU. Template-primers (24AF/13T and 24AT/13T, 0.5 pmol) were incubated with Pol I Kf (exo-) (0.03 unit) in the presence of dATP or dGTP (20 µM) at pH 7.2-8.6 (Tris buffer) or 6.9-9.3 (GTA buffer). Incubation was performed at 25 °C for 5 min. Incorporation of dAMP (A) and dGMP (B) opposite template fU and T (indicated above the gel) was analyzed by 16% denaturing PAGE. A and B, pH of the reaction mixture was 7.2 (lanes 1 and 6), 7.7 (lanes 2 and 7), 8.0 (lanes 3 and 8), 8.3 (lanes 4 and 9), and 8.6 (lanes 5 and 10). The extended primer is indicated by the arrow. The percentage of the extended primer resulting from incorporation of dAMP (C) and dGMP (D) was plotted against pH based on the product analysis in A and B. dGMP misincorporation opposite fU was also measured in a wide pH range (pH 6.9-9.3) using GTA buffer, and the result is shown in the inset of D. C and D, incorporation opposite template T and fU is represented by open and closed symbols, respectively.


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Fig. 6.   A proposed mechanism for mispair formation between fU and G. A, an acid-base equilibrium of fU (right) involving keto and ionized (enolate) forms and a tautomeric equilibrium of fU (left) involving keto and enol forms. B, base pairing schemes for the fU:G mispair involving ionized (or enolate) and keto forms of fU.

Extension of Matched and Mismatched Primer Termini-- The primer extension assay described above revealed that fU in template DNA directed incorporation not only of correct dAMP but also of incorrect dGMP, although less efficiently, during DNA synthesis, thereby giving rise to matched (fU:A) and mismatched (fU:G) primer termini. It is known that mismatched primer termini are extended less efficiently than matched termini by DNA polymerases and constitute a barrier for erroneous replication of DNA. Thus, the extension of primer termini containing fU:A and fU:G pairs was examined, and the results were compared with those of T:A and T:G pairs. Fig. 7 shows gel data when matched (24AT/14TA (T:A pair) and 24AF/14TA (fU:A pair)) and mismatched (24AT/14TG (T:G pair) and 24AF/14TG (fU:G pair)) primer termini were extended by Pol I Kf (exo-) in the presence of dCTP, a nucleotide to be incorporated following the primer termini. The matched primer termini containing T:A and fU:A pairs were elongated with comparable efficiencies (lanes 2 and 4). Although extension of the mismatched primer termini containing T:G (lane 6) and fU:G (lane 8) pairs were less efficient than matched ones, the extension of the fU:G terminus was clearly preferred over T:G. For quantitative comparison of the extension efficiencies, kinetic parameters (Vmax and Km) for dCMP incorporation were determined by the gel fidelity assay under standing start conditions (43) using 24AT, 24AF, 24CT, and 24CF (average of two experiments) (Table IV). Irrespective of the undamaged (24AT and 24CT) or damaged (24AF and 24CF) templates, the matched primer termini containing fU:A or T:A pairs were efficiently extended, with a slight preference of T:A (~1.2-fold as the extension efficiency (fC)). With the matched termini, Km values of extension (i.e. dCMP incorporation) were comparable to those of dAMP at the previous site (Table II), whereas the corresponding Vmax values were severalfold lower, presumably due to the difference in the incorporated nucleotide (C or A) or the sequence context. The extensions of the mismatched primer termini containing fU:G and T:G were inefficient, and the extension efficiency (fC) was 2 or 3 orders of magnitude lower than that of the corresponding matched termini (Table IV). For both T and fU, discrimination of the matched and mismatched termini exclusively originated from Km. Interestingly, the mismatched primer termini containing the fU:G pair was extended with significantly higher efficiencies than those containing the T:G pair. The mismatch extension frequency (fEX = fC(mismatched terminus)/fC(matched terminus) for the same template) was in a 10-2 range for fU, whereas that for T was in a 10-3 range. The increase in fEX associated with the substitution of fU for T was 8.2- (24AF versus 24AT) and 11.3-fold (24CF versus 24CT) (Fig. 4B). Accordingly, conversion of T to fU in template DNA promotes not only misincorporation of dGMP (Fig. 4A) but also elongation of the resulting mismatched primer termini (Fig. 4B).


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Fig. 7.   PAGE analysis of the extension of mismatched primer termini containing T:G and fU:G pairs. Template-primers (0.5 pmol) containing correctly paired (24AT/14TA (T:A pair) and 24AF/14TA (fU:A pair)) and mismatched (24AT/14TG (T:G pair) and 24AF/14TG (fU:G pair)) primer termini were incubated with Pol I Kf (exo-) (0.05 unit) in the presence of dCTP (10 µM) at 25 °C for 5 min. Products were analyzed by 16% denaturing PAGE. Lanes 1, 3, 5, and 7 show template-primers without the polymerase reaction, and lanes 2, 4, 6, and 8 show the extended products formed by the polymerase reaction. The base pair in the primer terminus is indicated above the gel.

                              
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Table IV
Parameters (Vmax and Km) and efficiencies (fC) of extension of matched and mismatched primer termini, and mismatch extension frequencies (fEX)


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

fU is one of the major oxidative thymine lesions found in DNA and nucleoside that were exposed to ionizing radiation (44-46), Fenton-type reactions (46, 47), photosensitized reactions (48, 49), and peroxy radicals (50). The yield of fU in Fenton-type reactions and gamma -irradiation is comparable to those of 8-oxoG (47) and 5-hydroxypyrimidines (45) that are known as major mutagenic oxidative base lesions (51-55). Bacterial (28, 37, 56) and mammalian (57, 58) cells contain repair enzyme or activity that excises fU from damaged DNA, implying potential genotoxic influences of this lesion in vivo. Direct incorporation of fdUTP into permeated E. coli cells resulted in a small but significant increase in chromosomal lacI mutation with G:C right-arrow A:T transitions being most preferred (59). fdU was also mutagenic to Salmonella typhimurium when added to the culture medium (44). In the present study, we have assessed the genotoxic potential of fU in template DNA by utilizing in vitro DNA replication reactions. The product analysis of translesion synthesis revealed that fU constituted very weak blocks to DNA synthesis (Fig. 2). Thus, unlike other thymine lesions such as thymine glycol, urea residues, and 5-hydroxy-5-methylhydantoin (13-15, 24), fU will exert a weak to negligible cytotoxic effect due to inhibition of DNA replication in vivo. Conversely, fU was shown to be a potentially mutagenic lesion based on the following results. First, the substitution of T by fU promoted misincorporation of incorrect dGMP (1.1-5.6 times as fG) and at the same time retarded incorporation of correct dAMP (1/4 to 1/2 as fA), hence leading to 3.3-14.3-fold increases in the replication error frequency (fRE) relative to T (Fig. 4A). Second, the resulting mismatched primer terminus containing an fU:G pair was more readily extended (8.2-11.3 times as fEX) than that containing a T:G pair (Fig. 4B) (see also the discussion below on fEX). This step will affect the probability that a genome DNA molecule is replicated to completion, and thereby scored as mutation. According to the present data, fU is moderately mutagenic, but for more quantitative estimation of the mutation frequency of fU, it is necessary to consider the influence of repair and the property of replicative DNA polymerases.

The mismatch extension frequency (fEX) in the absence of proofreading depends explicitly on the binding constant of DNA polymerase to matched versus mismatched template-primer DNA as well as on the concentrations of the template-primer DNA and next correct dNTP (60). Accordingly, to evaluate the intrinsic mismatch extension frequency (fEX(int)), possible differential binding of DNA polymerase to matched and mismatched primer termini needs to be taken into account under standing start conditions, i.e. differential binding to T:A versus T:G and fU:A versus fU:G termini in this study. The relationship between fEX and fEX(int) has been formulated in Equation 1 (60), where [Dr] and [Dw] are the concentrations of template-primer DNA having correctly and incorrectly paired primer termini (Dr and Dw), respectively, and Kr and Kw are the equilibrium constants for dissociation of polymerase-Dr and polymerase-Dw complexes, respectively.
f<SUB><UP>EX</UP></SUB>/f<SUB><UP>EX</UP>(<UP>int</UP>)</SUB>=(1+[D<SUB>r</SUB>]/K<SUB>r</SUB>)/(1+[D<SUB>w</SUB>]/K<SUB>w</SUB>) (Eq. 1)
In Equation 1, it is generally assumed that the affinity of DNA polymerase for a correctly paired terminus is similar to or higher than that for an incorrectly paired terminus (Kw >=  Kr). The values of [Dr] and [Dw] are both 33 nM in this study. When Pol I (exo-) has comparable affinities (Kr approx  Kw) for the matched and mismatched termini (i.e. T:A and T:G, and fU:A and fU:G), fEX in Table IV approximately represents the intrinsic value. According to Equation 1, the largest discrepancy between fEX and fEX(int) occurs when Kw is much higher than [Dw] (Kw 33 nM). In this case, Equation 1 can be transformed into Equation 2 by approximation.
f<SUB><UP>EX</UP></SUB>/f<SUB><UP>EX</UP>(<UP>int</UP>)</SUB>=(1+[D<SUB>r</SUB>]/K<SUB>r</SUB>) (Eq. 2)
Although the Kr values of Pol I (exo-) are not known, those for Avian myeloblastosis reverse transcriptase and Drosophila melanogaster DNA polymerase alpha  have been estimated as 5 and 20-50 nM, respectively (60). Granted that the Kr value of Pol I (exo-) is in a similar range (5-50 nM), the fEX values in Table IV are subjected to a 7.6-fold reduction. However, the correction factor given by Equation 2 is presumably similar for the T and fU templates since the affinities (Kr) of Pol I (exo-) for the structurally resembling T:A and fU:A termini are likely comparable. These considerations suggest that the relative difference in fEX(int) for the T versus fU templates remains similar to that shown in Fig. 4B, although the absolute value of fEX(int) may be lower than that obtained experimentally in this study (fEX).

Concerning the mispairing mechanism of fU, we have previously suggested participation of the ionized (or enolate) form of fU based on the pH-dependent misincorporation of fdUTP opposite template G (27). Privat and Sowers (61) also proposed a similar mispairing scheme on the basis of pKa measurement of fdU and related nucleosides. Consistent with this mechanism, the efficiency of dGMP misincorporation opposite template fU increased around the pKa value of fU (pKa = 8.6), whereas the corresponding increase for T (pKa = 10.0) was much smaller than that for fU (Fig. 5, B and D). The result obtained for T also agrees with a small increase in the base substitution frequency of Pol I (exo-) in this pH range (62). Unlike thymine bearing an electron donating methyl group, fU has an electron withdrawing formyl group that promotes ionization of fU in an acid-base equilibrium (Fig. 6A). According to the pKa values, the fraction of the ionized form of fU increases from 4 to 50% in the pH range of 7.2-8.6 but that of thymine is virtually negligible (0.2-4%). Ionized fU can form a base pair with incoming dGTP through two hydrogen bonds (Fig. 6B, left). The base pair formed between ionized fU and dGTP essentially assume Watson-Crick geometry (or B form geometry) and can fit into the active site of DNA polymerase. Since the geometric recognition is key to discrimination of correct versus incorrect nucleotides by DNA polymerases (1), this geometry probably promoted misincorporation of dGMP opposite fU. Participation of base ionization promoted by electron withdrawing substituents has been demonstrated in the mispairing of 5-halogenated uracils (BrU and 5-fluorouracil) with G (32). Thus, fU and 5-halogenated uracils share a common mutation mechanism. Although participation of a rare enol tautomer of fU (Fig. 6A) in the mispairing with G cannot be fully ruled out, recent NMR studies show that the tautomeric equilibrium between keto and enol forms of fU is not significantly affected by oxidation of the methyl group of T to the formyl group (63, 64). Therefore, involvement of the enol form of fU is unlikely in the mispairing with G. It is assumed that after dGMP incorporation, the resulting fU(ionized):G pair in Watson-Crick geometry shifts to wobble geometry (fU(keto):G) due to the acid-base equilibrium (Fig. 6B, right). However, a certain fraction of the base pair will still exist as an fU(ionized):G pair whose geometry can again promote incorporation of the next nucleotide. Probably this is the reason why the mismatched primer terminus containing an fU:G pair was more efficiently extended than that containing a T:G pair. Although there is no experimental evidence that directly shows the equilibrium between fU(keto):G and fU(ionized):G base pairs in duplex DNA, the presence of such a equilibrium has been demonstrated for BrU(keto):G and BrU (ionized):G pairs in a duplex oligonucleotide by the NMR study (65). According to the proposed mutation mechanism for fU, it is reasonable that 5-hydroxymethyluracil, another methyl oxidation product of T, does not direct misincorporation (25, 26) since the hydroxymethyl group has electron donating nature and cannot promote ionization of the base. Although a mutation mechanism involving an altered acid-base equilibrium has been previously demonstrated for 5-halogenated uracils (32), to our knowledge, fU is the first example adapting to this mechanism among oxidative DNA base lesions.

To assess the sequence context effect on the base pairing property of fU, the 3'-nearest neighbor base of template fU and the paired base (i.e. the primer terminus base pair) was systematically changed, and the nucleotide incorporated opposite fU was analyzed. fU with the four possible nearest neighbor base pairs directed incorporation of dAMP and to a less extent dGMP but not dCMP and dTMP (Fig. 3). Thus, 3'-nearest neighbor base exhibited no influence concerning the type of base pairs formed from template fU. Combining the result with fdUTP (27, 28) and template fU (this study), it follows that fundamental base pairing symmetry is retained whether the fU base pairs are formed from incoming fdUTP or template fU during DNA polymerase reactions. Although fundamental base pairing symmetry held with respect to the formation of the fU base pairs, the nearest neighbor base pair showed quantitative effects on the incorporation efficiency of dAMP and dGMP. Regardless of the correct or incorrect incorporation (Fig. 4, C and D), the sequence context effect was less pronounced for fU than T. For example, the difference in fG for T was 9.4-fold for the highest (TT) and lowest (GT) sequences, whereas the corresponding value (TF versus AF, F = fU) was only 2.2-fold for fU (Fig. 4D). Another notable sequence context effect was a tendency of preferred incorporation of dAMP and dGMP for the sequences containing 3'-pyrimidines over 3'-purines, although the difference was not so large. In other words, incorporation of the nucleotides was favored for the primer terminus containing purines. This was common to T and fU. Presumably, stabilization of the incoming purine nucleotides through a favored purine-purine stacking interaction (purine-purine > pyrimidine-purine > pyrimidine-pyrimidine) (66) promoted their incorporation over pyrimidine nucleotides. Despite the sequence context dependent variations of fA and fG, the fRE (=fG/fA) values of fU were consistently higher than those of T and were virtually independent of the nearest neighbor base pair (Fig. 4E). This result suggests that the distribution of T right-arrow C transitions induced by fU will be relatively uniform with respect to the variation of the 3'-nearest neighbor base unless heterogeneous formation or repair of fU occurs in cells. Finally, the relative increase in fRE associated with the conversion of T to fU was calculated. The order of the increase was A > G > C > T with respect to the 3'-nearest neighbor base (Fig. 4A), showing an inverse correlation with that (T > C > G > A) ranged by fRE of T (Fig. 4E). Thus, the sequence with a low replication error frequency before conversion to fU gave rise to a relatively large increase in the replication error frequency after conversion to fU.

Previously, Zhang et al. (29, 30) assessed the mutagenic potential of fU using an in vitro system similar to that used in this study. According to their experiments with Pol I Kf, Pol I Kf (exo-), and thermophilic DNA polymerases, fU directed incorporation of dCMP as well as dAMP in all cases but not dGMP and dTMP at all. The incorporation efficiency of dCMP relative to dAMP was 0.09-0.15 for Pol I Kf, 0.06 for Pol I Kf (exo-), and 0.23-0.27 for Tth DNA polymerase. These values are unusually high as a frequency of pyrimidine:pyrimidine mispair formation by DNA polymerases. Generally, pyrimidine:pyrimidine pairs are too small to fit into the B form helix. For this reason, formation of these mispairs is not favored by prokaryotic and eukaryotic DNA polymerases. The most common mispairs are G:T mispairs with observed frequencies between 10-2 and 10-4, and the least common ones are pyrimidine:pyrimidine mispairs with frequencies between 10-4 and 10-5 (1). The rate of misincorporation of dNMP opposite T by Pol I Kf and Pol I Kf (exo-) also follows this rule (dGMP dCMP >=  dTMP) (62, 67). The measurement of the melting temperature (Tm) of duplex oligonucleotides containing an fU:N pair (N = A, G, C, T) revealed that Tm decreased in the following order: fU:A > fU:G > fU:T >=  fU:C.3 This order indicates that the fU:C pair exerts the largest destabilization effect on DNA among the four possible base pairs and further suggests the least favored formation of this pair during DNA replication. In addition, Zhang et al. (29) found significant incorporation of dCMP but not dGMP opposite normal T (Table V), suggesting a fundamental discrepancy in the experimental set up used in the present and their studies. We repeated this experiment using the same template-primer (i.e. template 1 and primer 3 shown in Table V). However, the reported result was not reproduced, and misincorporation of only dGMP was detected (Table V). Thus, T in this sequence context was not particularly prone to incorporate dCMP. We also repeated another control experiment with Tth DNA polymerase under the reported conditions (29, 30). Template 1 (see Table V for the sequence) was annealed to a 9-mer primer (5'-TGCAGGTCG) and primer extension assays were performed in the presence of a single dNTP at 74 °C for 5 or 10 min. Although they observed incorporation of dAMP opposite T under these conditions, we did not see incorporation of any nucleotides. We believe the present result is reasonable in light of the expected low melting temperature of the 9-mer primer (Tm = 30 °C). It is very likely that the template-primer dissociated at 74 °C and was unable to serve as a substrate for Tth DNA polymerase in the present experiment. For the same reason, the reported incorporation of dCMP with the 9-mer primer by Tth DNA polymerase (29, 30) is very unlikely to occur at such a high temperature. In view of the inconsistencies such as the contradiction against the general preference of mispair formation by DNA polymerases and the lack of reproducibility of the certain experimental results, we believe fU directs misincorporation of dGMP but not dCMP. Therefore, whether fU is in incoming dNTP or a template, base pairing symmetry in the nucleotide selection by DNA polymerase and the mutagenesis mechanism involving ionized fU hold, although the incorporation efficiency of dAMP and dGMP varies depending on the nearest neighbor base pair.

                              
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Table V
Specificity of nucleotide incorporation opposite template T


    ACKNOWLEDGEMENTS

We thank Akira Matsuda and Naoko Karino (Hokkaido University) for communicating the Tm data for the duplexes containing fU.

    Note Added in Proof

Recently, Miyabe et al. (69) have reported that the fU site-specifically incorporated into plasmid vectors induces mutations but does not direct misincorporation of dCMP when the plasmids are replicated in Escherichia coli.

    FOOTNOTES

* This work was supported by grants-in-aid from the Ministry of Education, Science, and Culture of Japan (to H. I.) and by JSPS Research Fellowships for Young Scientists (to A. M.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Dagger To whom correspondence should be addressed. Tel./Fax: 81-824-24-7457; E-mail: ideh@hiroshima-u.ac.jp.

Published, JBC Papers in Press, January 29, 2001, DOI 10.1074/jbc.M008598200

2 Recently, the activity of Endo III and Fpg for fU was reported by Zhang et al. (68). Our study on the kinetic parameter revealed that their activity for fU was 2-3 orders of magnitude lower than their intrinsic substrates (thymine glycol for Endo III and 7,8-dihydro-8-oxoguanine for Fpg). Thus, the activity of the two enzymes for fU was negligible under standard assay conditions, and extremely large excesses of enzymes were required to detect the activity for fU (A. Masaoka, H. Terato, Y. Ohyama, and H. Ide, manuscript in preparation).

3 The UV melting curves (plots of A260 against temperature) were measured using the duplexes (total strand concentration 3 µM) of 25F and its complementary strand containing A, G, C, or T as the base opposite fU in 10 mM NaCl and 10 mM sodium cacodylate (pH 7.0). The Tm value was evaluated from the inflection point of the melting curve (A. Masaoka, H., Terato, Y. Ohyama, N. Karino, A. Matsuda, and H. Ide, manuscript in preparation).

    ABBREVIATIONS

The abbreviations used are: fU, 5-formyluracil; fdUTP, 5-formyl-2'-deoxyuridine 5'-triphosphate; fdU, 5-formyl-2'-deoxyuridine; BrU, 5- bromouracil; Pol I Kf, Escherichia coli DNA polymerase I Klenow fragment; Pol I Kf (exo-), Pol I Kf deficient in 3'-5'-exonuclease; PAGE, polyacrylamide gel electrophoresis; Endo, endonuclease; Fpg, formamidopyrimidine glycosylase; HPLC, high-performance liquid chromatography.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

1. Echols, H., and Goodman, M. F. (1991) Annu. Rev. Biochem. 60, 477-511[CrossRef][Medline] [Order article via Infotrieve]
2. Johnson, K. A. (1993) Annu. Rev. Biochem. 62, 685-713[CrossRef][Medline] [Order article via Infotrieve]
3. Modrich, P., and Lahue, R. (1996) Annu. Rev. Biochem. 65, 101-133[CrossRef][Medline] [Order article via Infotrieve]
4. Friedberg, E. C., Walker, G. C., and Siede, W. (1995) DNA Repair and Mutagenesis , American Society for Microbiology, Washington, D. C.
5. von Sonntag, C. (1987) Chemical Basis of Radiation Biology , Taylor & Francis, New York
6. Breen, A. P., and Murphy, J. A. (1995) Free Radic. Biol. Med. 18, 1033-1077[CrossRef][Medline] [Order article via Infotrieve]
7. Hatahet, Z., and Wallace, S. S. (1998) in DNA Damage and Repair (Nickoloff, J. A. , and Hoekstra, M. F., eds), Vol. 1 , pp. 229-262, Humana Press Inc., Totowa, NJ
8. Wang, D., Kreutzer, D. A., and Essigmann, J. M. (1998) Mutat. Res. 400, 99-115[CrossRef][Medline] [Order article via Infotrieve]
9. Friedberg, E. C., Feaver, W. J., and Gerlach, V. L. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 5681-5683[Free Full Text]
10. Lewis, D. A., Bebenek, K., Beard, W. A., Wilson, S. H., and Kunkel, T. A. (1999) J. Biol. Chem. 274, 32924-32930[Abstract/Free Full Text]
11. Steiz, T. A. (1999) J. Biol. Chem. 274, 17395-17398[Free Full Text]
12. Gasparutto, D., Bourdat, A.-G., D'Ham, C., Duarte, V., Romieu, A., and Cadet, J. (2000) Biochimie (Paris) 82, 19-24[CrossRef][Medline] [Order article via Infotrieve]
13. Evans, J., Maccabee, M., Hatahet, Z., Courcelle, J., Bockrath, R., Ide, H., and Wallace, S. S. (1993) Mutat. Res. 299, 147-156[Medline] [Order article via Infotrieve]
14. Ide, H., Kow, Y. W., and Wallace, S. S. (1985) Nucleic Acids Res. 13, 8035-8052[Abstract]
15. Ide, H., Petrullo, L. A., Hatahet, Z., and Wallace, S. S. (1991) J. Biol. Chem. 266, 1469-1477[Abstract/Free Full Text]
16. Ide, H., Melamede, R. J., and Wallace, S. S. (1987) Biochemistry 26, 964-969[Medline] [Order article via Infotrieve]
17. Ide, H., and Wallace, S. S. (1988) Nucleic Acids Res. 16, 11339-11354[Abstract]
18. Ide, H., Yamaoka, T., and Kimura, Y. (1994) Biochemistry 33, 7127-7133[Medline] [Order article via Infotrieve]
19. Ide, H., Murayama, H., Sakamoto, S., Makino, K., Honda, K., Nakamuta, H., Sasaki, M., and Sugimoto, N. (1995) Nucleic Acids Res. 23, 123-129[Abstract]
20. Shimizu, H., Yagi, R., Kimura, Y., Makino, K., Terato, H., Ohyama, Y., and Ide, H. (1997) Nucleic Acids Res. 25, 597-603[Abstract/Free Full Text]
21. Suzuki, T., Yoshida, M., Yamada, M., Ide, H., Kobayashi, M., Kanaori, K., Tajima, K., and Makino, K. (1998) Biochemistry 37, 11592-11598[CrossRef][Medline] [Order article via Infotrieve]
22. Hayes, R. C., Petrullo, L. A., Huang, H. M., Wallace, S. S., and LeClerc, J. E. (1988) J. Mol. Biol. 201, 239-246[CrossRef][Medline] [Order article via Infotrieve]
23. Maccabee, M., Evans, J. S., Glackin, M. P., Hatahet, Z., and Wallace, S. S. (1994) J. Mol. Biol. 236, 514-530[CrossRef][Medline] [Order article via Infotrieve]
24. Gasparutto, D., Ait-Abbas, M., Jaquinod, M., Boiteux, S., and Cadet, J. (2000) Chem. Res. Toxicol. 13, 575-584[CrossRef][Medline] [Order article via Infotrieve]
25. Levy, D. D., and Teebor, G. W. (1991) Nucleic Acids Res. 19, 3337-3343[Abstract]
26. Mi, L.-J., Mahl, E., Chaung, W., and Boorstein, R. J. (1997) Mutat. Res. 374, 287-295[CrossRef][Medline] [Order article via Infotrieve]
27. Yoshida, M., Makino, K., Morita, H., Terato, H., Ohyama, Y., and Ide, H. (1997) Nucleic Acids Res. 25, 1570-1577[Abstract/Free Full Text]
28. Terato, H., Masaoka, A., Kobayashi, M., Fukushima, S., Ohyama, Y., Yoshida, M., and Ide, H. (1999) J. Biol. Chem. 274, 25144-25150[Abstract/Free Full Text]
29. Zhang, Q.-M., Sugiyama, H., Miyabe, I., Matsuda, S., Saito, I., and Yonei, S. (1997) Nucleic Acids Res. 25, 3969-3973[Abstract/Free Full Text]
30. Zhang, Q.-M., Sugiyama, H., Miyabe, I., Matsuda, S., Kino, K., Saito, I., and Yonei, S. (1999) Int. J. Radiat. Biol. 75, 59-65[CrossRef][Medline] [Order article via Infotrieve]
31. Lawley, P. D., and Brookes, P. (1962) J. Mol. Biol. 4, 216-219[Medline] [Order article via Infotrieve]
32. Yu, H., Eritja, R., Bloom, L. B., and Goodman, M. F. (1993) J. Biol. Chem. 268, 15935-15943[Abstract/Free Full Text]
33. Sugiyama, H., Matsuda, S., Kino, K., Zhang, Q.-M., Yonei, S., and Saito, I. (1996) Tetrahedron Lett. 37, 9067-9070[CrossRef]
34. Carroll, S. S., Cowart, M., and Benkovic, S. J. (1991) Biochemistry 30, 804-813[Medline] [Order article via Infotrieve]
35. Ahn, J., Werneburg, B. G., and Tsai, M.-D. (1997) Biochemistry 36, 1100-1107[CrossRef][Medline] [Order article via Infotrieve]
36. Einolf, H. J., Schnetz-Boutaud, N., and Guengerich, F. P. (1998) Biochemistry 37, 13300-13312[CrossRef][Medline] [Order article via Infotrieve]
37. Masaoka, A., Terato, H., Kobayashi, M., Honsho, A., Ohyama, Y., and Ide, H. (1999) J. Biol. Chem. 274, 25136-25143[Abstract/Free Full Text]
38. Asagoshi, K., Yamada, T., Okada, Y., Terato, H., Ohyama, Y., Seki, S., and Ide, H. (2000) J. Biol. Chem. 275, 24781-24786[Abstract/Free Full Text]
39. Asagoshi, K., Yamada, T., Terato, H., Ohyama, Y., Monden, Y., Arai, T., Nishimura, S., Aburatani, H., Lindahl, T., and Ide, H. (2000) J. Biol. Chem. 275, 4956-4964[Abstract/Free Full Text]
40. Terato, H., Morita, H., Ohyama, Y., and Ide, H. (1998) Nucleosides & Nucleotides 17, 131-141[Medline] [Order article via Infotrieve]
41. Armstrong, V. W., Dattagupta, J. K., Eckstein, F., and Saenger, W. (1976) Nucleic Acids Res. 3, 1791-1810[Medline] [Order article via Infotrieve]
42. Cline, R. E., Fink, R. M., and Fink, K. (1959) J. Am. Chem. Soc. 81, 2521-2527
43. Creighton, S., Bloom, L. B., and Goodman, M. F. (1995) Methods Enzymol. 262, 232-256[Medline] [Order article via Infotrieve]
44. Kasai, H., Iida, A., Yamaizumi, Z., Nishimura, S., and Tanooka, H. (1990) Mutat. Res. 243, 249-253[CrossRef][Medline] [Order article via Infotrieve]
45. Douki, T., Delatour, T., Paganon, F., and Cadet, J. (1996) Chem. Res. Toxicol. 9, 1145-1151[CrossRef][Medline] [Order article via Infotrieve]
46. Murata-Kamiya, N., Kamiya, H., Karino, N., Ueno, Y., Kaji, H., Matsuda, A., and Kasai, H. (1999) Nucleic Acids Res. 27, 4385-4390[Abstract/Free Full Text]
47. Murata-Kamiya, N., Kamiya, H., Muraoka, M., Kaji, H., and Kasai, H. (1997) J. Radiat. Res. 38, 121-131[Medline] [Order article via Infotrieve]
48. Decarroz, C., Wagner, J. R., Van Lier, J. E., Krishna, C. M., Riesz, P., and Cadet, J. (1986) Int. J. Radiat. Biol. 50, 491-505
49. Saito, I., Takayama, M., and Kawanishi, S. (1995) J. Am. Chem. Soc. 117, 5590-5591
50. Martini, M., and Termini, J. (1997) Chem. Res. Toxicol. 10, 234-241[CrossRef][Medline] [Order article via Infotrieve]
51. Shibutani, S., Takeshita, M., and Grollman, A. P. (1991) Nature 349, 431-434[CrossRef][Medline] [Order article via Infotrieve]
52. Maki, H., and Sekiguchi, M. (1992) Nature 355, 273-275[CrossRef][Medline] [Order article via Infotrieve]
53. Purmal, A. A., Kow, Y. W., and Wallace, S. S. (1994) Nucleic Acids Res. 22, 72-78[Abstract]
54. Purmal, A. A., Kow, Y. W., and Wallace, S. S. (1994) Nucleic Acids Res. 22, 3930-3935[Abstract]
55. Kreutzer, D. A., and Essigmann, J. M. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 3578-3582[Abstract/Free Full Text]
56. Bjelland, S., Birkeland, N., Benneche, T., Volden, G., and Seeberg, E. (1994) J. Biol. Chem. 269, 30489-30495[Abstract/Free Full Text]
57. Bjelland, S., Eide, L., Time, R. W., Stote, R., Eftedal, I., Volden, G., and Seeberg, E. (1995) Biochemistry 34, 14758-14764[Medline] [Order article via Infotrieve]
58. Zhang, Q.-M., Fujimoto, J., and Yonei, S. (1995) Int. J. Radiat. Biol. 68, 603-607[Medline] [Order article via Infotrieve]
59. Fujikawa, K., Kamiya, H., and Kasai, H. (1998) Nucleic Acids Res. 26, 4582-4587[Abstract/Free Full Text]
60. Mendelman, L. V., Petruska, J., and Goodman, M. F. (1990) J. Biol. Chem. 265, 2338-2346[Abstract/Free Full Text]
61. Privat, E. J., and Sowers, L. C. (1996) Mutat. Res. 354, 151-156[CrossRef][Medline] [Order article via Infotrieve]
62. Eckert, K. A., and Kunkel, T. A. (1993) J. Biol. Chem. 268, 13462-13471[Abstract/Free Full Text]
63. La Francois, C. J., Fujimoto, J., and Sowers, L. C. (1998) Chem. Res. Toxicol. 11, 75-83[CrossRef][Medline] [Order article via Infotrieve]
64. La Francois, C. J., Jang, Y. H., Cagin, T., Goddard, W. A. I., II, and Sowers, L. C. (2000) Chem. Res. Toxicol. 13, 462-470[CrossRef][Medline] [Order article via Infotrieve]
65. Sowers, L. C., Goodman, M. F., Eritja, R., Kaplan, B., and Fazakerley, G. V. (1989) J. Mol. Biol. 205, 437-447[Medline] [Order article via Infotrieve]
66. Saenger, N. (1984) Principles of Nucleic Acid Structure , Springer-Verlag Inc., New York
67. Joyce, C. M., Sun, X. C., and Grindley, N. D. F. (1992) J. Biol. Chem. 267, 24485-24500[Abstract/Free Full Text]
68. Zhang, Q.-M., Miyabe, I., Matsumoto, Y., Kino, K., Sugiyama, H., and Yonei, S. (2000) J. Biol. Chem. 275, 35471-35477[Abstract/Free Full Text]
69. Miyabe, I., Zhang, Q.-M., Sugiyama, H., Kino, K., and Yonei, S. (2001) Int. J. Radiat. Biol. 77, 53-58[CrossRef][Medline] [Order article via Infotrieve]


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