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Originally published In Press as doi:10.1074/jbc.M610718200 on January 8, 2007

J. Biol. Chem., Vol. 282, Issue 11, 8188-8198, March 16, 2007
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Mechanism of Abasic Lesion Bypass Catalyzed by a Y-family DNA Polymerase*Formula

Kevin A. Fiala{ddagger}§1, Cameron D. Hypes{ddagger}, and Zucai Suo{ddagger}§||2

From the {ddagger}Department of Biochemistry, the §Ohio State Biochemistry Program, the Molecular, Cellular and Developmental Biology Program, and the ||Comprehensive Cancer Center, Ohio State University, Columbus, Ohio 43210

Received for publication, November 20, 2006 , and in revised form, January 3, 2007.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The 3 million-base pair genome of Sulfolobus solfataricus likely undergoes depurination/depyrimidination frequently in vivo. These unrepaired abasic lesions are expected to be bypassed by Dpo4, the only Y-family DNA polymerase from S. solfataricus. Interestingly, these error-prone Y-family enzymes have been shown to be physiologically vital in reducing the potentially negative consequences of DNA damage while paradoxically promoting carcinogenesis. Here we used Dpo4 as a model Y-family polymerase to establish the mechanistic basis for DNA lesion bypass. While showing efficient bypass, Dpo4 paused when incorporating nucleotides directly opposite and one position downstream from an abasic lesion because of a drop of several orders of magnitude in catalytic efficiency. Moreover, in disagreement with a previous structural report, Dpo4-catalyzed abasic bypass involves robust competition between the A-rule and the lesion loop-out mechanism and is governed by the local DNA sequence. Analysis of the strong pause sites revealed biphasic kinetics for incorporation indicating that Dpo4 primarily formed a nonproductive complex with DNA that converted slowly to a productive complex. These strong pause sites are mutational hot spots with the embedded lesion even affecting the efficiency of five to six downstream incorporations. Our results suggest that abasic lesion bypass requires tight regulation to maintain genomic stability.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Numerous DNA-damaging agents continuously attack the cellular genome and generate a myriad of DNA lesions. Although a majority of these lesions are repaired by DNA repair pathways, some damage evades repair. Unrepaired lesions arrest DNA replication by inhibiting replicative DNA polymerases. However, a recently identified class of low fidelity enzymes, known as the Y-family DNA polymerases, can replicate through DNA damage and rescue cells from apoptosis. In humans, four Y-family DNA polymerases have been identified (1). Because of its thermostability and ease of purification (2), we have decided to use Sulfolobus solfataricus DNA polymerase IV (Dpo4)3 as a model Y-family DNA polymerase to study the mechanistic basis of DNA synthesis. Dpo4, which retains catalytic activity at 37 °C, has been shown to exhibit a low fidelity in the range of 10-3 to 10-4, or one error for every 1,000–10,000 nucleotides incorporated on undamaged DNA as revealed by our pre-steady state kinetic studies (3) and by both steady state kinetics and a forward mutation assay performed by others (2, 4). Notably, Dpo4 and a human Y-family member, DNA polymerase {kappa} (human pol {kappa}), are both DinB homologs and have been shown to possess very similar error rates (5, 6), yet a previous study (2) suggests that the lesion bypass properties of Dpo4 are more similar to eukaryotic DNA polymerase {eta} (pol {eta}). As such, because of its functionally and structurally conserved features with respect to other Y-family enzymes, Dpo4 is considered to be the archetypal Y-family DNA polymerase for studies elucidating the mechanism and function of this novel class of enzymes.

Careful kinetic analysis of the mechanistic basis of nucleotide incorporation into undamaged DNA catalyzed by the Y-family DNA polymerases has been reported for yeast pol {eta} (7), Dpo4 (3, 8), and Dbh (9). However, besides studies reporting several kinetic parameters and mutation profiles for the bypass of 7,8-dihydro-8-oxodeoxyguanine (8-oxodG) by yeast pol {eta} (10), and for the bypass of a cis-syn-thymine-thymine (T-T) dimer catalyzed by yeast pol {eta} (11), thorough mechanistic analysis of a Y-family polymerase promoting lesion bypass has not been reported. In this study, we report the first comprehensive characterization of the mechanism of lesion bypass catalyzed by Dpo4 through rigorous pre-steady state kinetic analysis. It has been well established that apurinic/apyrmidinic (AP) sites resulting from the hydrolytic cleavage of the N-glycosidic bond are among the most abundant lesions encountered in a mammalian cell (12) with ~10,000 spontaneous AP sites generated in each cell every day (13, 14). It is plausible that AP sites will be generated at a similar or higher frequency in S. solfataricus because this aerobic crenarchaeon propagates at roughly 80 °C (15). Although replication past these noninstructive lesions is a mechanistic challenge, replicative and repair DNA polymerases have been shown to preferentially incorporate dATP, although inefficiently, opposite the AP lesion in a phenomenon known as the A-rule (16). Yet recent structural studies have concluded that instead of the A-rule, Dpo4, and thus other DinB homologs, almost exclusively uses the template base 5' to the lesion (referred to as the 5'-rule) to instruct nucleotide incorporation during AP bypass (17). However, our thorough examination of the kinetic effects of AP bypass has clearly illustrated a sequence-dependent competition between these two pathways in disagreement with the conclusions from these previous structural studies.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Reaction Buffer R—Buffer R contains 50 mM HEPES-NaOH (pH 7.5 at 37 °C), 5 mM MgCl2, 50 mM NaCl, 5 mM dithiothreitol, 10% glycerol, 0.1 mM EDTA, and 0.1 mg/ml bovine serum albumin (3). All concentrations reported here refer to the concentration of components after mixing. All reactions, unless noted, were carried out at 37 °C.

Running Start Assay—Experiments were carried out in a rapid chemical quench flow apparatus (KinTek) by rapidly mixing an aliquot (15 µl) of a solution containing 100 nM 5'-[32P]DNA and 100 nM Dpo4 preincubated in buffer R with an aliquot (15 µl) containing all four dNTPs (200 µM each) for times ranging from milliseconds to minutes followed by quenching with 0.37 M EDTA. The nucleotide incorporation pattern was resolved by sequencing gel analysis. Assays performed with pol B1 were carried out using the same buffer and reaction conditions used for Dpo4.

Determination of the kp and Kd of an Incoming Nucleotide—A solution containing Dpo4 (120 nM) and 5'-[32P] DNA (30 nM) preincubated in buffer R was mixed with increasing concentrations of a single nucleotide. Reactions were terminated by the addition of EDTA. Products were separated from substrate via sequencing gel electrophoresis (17% acrylamide, 8 M urea) and quantitated using a PhosphorImager 445 SI (Amersham Biosciences). The time course of product formation was fit to Equation 1,

Formula 1(Eq. 1)
for each concentration of dNTP to yield an observed rate constant (kobs) and a reaction amplitude (A). The extracted kobs values were then plotted as a function of the concentration of dNTP and fit to Equation 2,

Formula 2(Eq. 2)
to give kp and Kd. The substrate specificity (kp/Kd) was then calculated.

Electrophoretic Mobility Shift Assay—Mobility shift titrations were conducted using a 4.5% native polyacrylamide gel with a running buffer (50 mM Tris acetate, pH 7.5, 23 °C, 5.5 mM Mg(OAc)2, and 0.5 mM EDTA). Dpo4 (35–425 nM) was titrated into a solution of 5'-[32P]DNA (100 nM) in buffer R at 23 °C. Aliquots of the titration were loaded into the native gel and run at a constant voltage of 80 V for 30 min at 23 °C. The dried gels were exposed to a PhosphorImager and quantitated. The concentration of Dpo4·DNA was plotted against the concentration of Dpo4 and fit using Equation 3,

Formula 3(Eq. 3)
where Eo represents the active enzyme concentration and Do the DNA concentration.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Previously we exploited transient state kinetic analyses to measure the fidelity and establish a minimal kinetic mechanism for single nucleotide incorporation into an undamaged DNA substrate catalyzed by Dpo4 (3, 8). These results demonstrated that Dpo4 uses an induced-fit mechanism to select a correct nucleotide as observed for several replicative and repair polymerases (18) and another Y-family DNA polymerase, yeast pol {eta} (7). This mechanism served as a foundation to establish the mechanistic basis for bypassing DNA lesions such as AP sites, catalyzed by Dpo4. Notably, natural AP sites exist as an uneven mixture of four species, with the equilibrium favoring the two anomeric hemiacetals over the two open chain aldehydes (19, 20). However, the aldehydic forms are subject to beta-elimination and subsequent scission of the DNA backbone, producing a heterogeneous mixture of DNA substrates. Because of this instability, we have chosen to perform our studies with the stable AP analog tetrahydrofuran because a homogeneous population of DNA substrates is requisite for rigorous kinetic studies. Most crystal structure studies and kinetic assays involving AP-containing DNA have been carried out using tetrahydrofuran (17, 21–26). Moreover, tetrahydrofuran has been shown to retain the biological properties of the natural AP site in vivo (27).

Bypassing an AP Lesion—To investigate the response of Dpo4 to an AP lesion, we designed a "running start" assay (see under "Experimental Procedures") such that the effect of the AP lesion on DNA synthesis could be determined via observation of the polymerization pattern over time at 37 °C for reasons described previously (3). Elongation of 5'-32P-labeled 17-mer/41AP (Table 1) proceeded rapidly up until the template AP site where the incorporation pattern showed two consecutive strong pause sites (Fig. 1B), corresponding to incorporation opposite the AP site and its extension. At these two strong pause sites, the nucleotide incorporation pattern showed a significant accumulation of intermediate products 21-mer and 22-mer (Fig. 1B), suggesting slow turnover. Nonetheless, the AP site was bypassed relatively efficiently by Dpo4 (full-length product was observed at 40 s), and subsequent downstream incorporation was not significantly perturbed by the embedded AP lesion. In comparison, Dpo4 pausing was not observed, and the full-length product was formed after 10 s (Fig. 1A) in the control reactions with an undamaged 17-mer/41CTL (Table 1). Interestingly, Dpo4 was able to catalyze blunt-end addition to the full-length 41-mer to generate 42-mer as described previously (28). Notably, the same running start assay was performed with S. solfataricus replicative DNA polymerase pol B1 and showed no bypass of the AP site, instead stalling predominantly one nucleotide preceding the lesion (supplemental Fig. 1B).


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  		TABLE 1
DNA primers and templates

 


Figure 1
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  		FIGURE 1.
Running start nucleotide incorporation assay. A preincubated solution of Dpo4 (100 nM) and 17-mer/41-mer (100 nM) were mixed with all four dNTPs (200 µM each) for various reaction times before quenching with EDTA. The product lengths were labeled, and the AP was designated. A, reaction with the 17-mer/41CTL substrate; B, reaction with the 17-mer/41AP substrate.

 


Figure 2
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  		FIGURE 2.
Measurement of the binding of Dpo4 to 21-mer/41AP. A, reactions containing 100 nM 5'-32P-labeled 21-mer/41AP were incubated with increasing concentration of Dpo4 (35–425 nM) followed by native gel analysis to separate binary complex from unbound DNA substrate. B, solid line is a fit of the data (•) to a quadratic equation giving a Kd = 36 ± 2 nM.

 
Measurement of the Binding of Dpo4 to DNA Containing an AP Site—It is well established that AP sites have a destabilizing effect on the DNA duplex stability (29, 30) and conformation (31, 32). It is thus plausible that the strong pause sites observed proximal to the lesion (Fig. 1B) could be due to reduced binding of Dpo4 for the AP-containing DNA. To evaluate this hypothesis, the electrophoretic mobility shift assay was employed to measure overall binding affinity of Dpo4 to a series of lesion-containing DNA substrates. In these substrates, primers of increasing length were designed (Table 1) such that when annealed to the template 41AP they would structurally mimic the progression of DNA synthesis as Dpo4 incorporated one nucleotide at a time. These measurements allowed for the estimation of the equilibrium dissociation constant (Kd) of the Dpo4·DNA binary complex, as Dpo4 incrementally approached and proceeded downstream from the AP site. For example, a solution containing the estimated after gel analysis (Fig. 2A) to be 36 ± 2 nM (Fig. 2B). Similarly, the Kd value for the binding of Dpo4 to undamaged 21-mer/41CTL was estimated to be 10 ± 2 nM. This value was very similar to 11 ± 1 nM, which was measured previously using active site titration (8) suggesting that electrophoretic mobility shift assay was a reliable assay to measure the Kd value of the Dpo4·DNA binary complex.

Our kinetic analysis (see below) indicated that although incorporation opposite the AP site (first pause site) was ~210-fold slower than a matched incorporation, 90% of the incorporation events at this site were either dATP or dCTP, suggesting two dominant pathways to bypass the AP lesion. To observe the effect of an AP site on DNA binding to Dpo4, the DNA substrates assayed were designed to precisely represent the sequence contexts at the two strong pause sites as well as at both upstream and downstream nonpause sites (Fig. 1B) with either adenine or cytosine opposite the AP site when applicable (Table 2). Interestingly, the Dpo4-binding affinities at the two strong pause sites were less than 4-fold lower than their corresponding controls, whereas at nonpause sites all the Kd values were within 2-fold of their corresponding control substrates (Table 2). Thus, the flexibility from the AP lesion did not significantly perturb Dpo4 binding and therefore was not the major contributor to the observed strong pausing of Dpo4 in Fig. 1B. However, the local structural flexibility at the AP site may hinder nucleotide incorporation. We subsequently hypothesized that these strong pause sites were generated by a significant decrease in the incorporation efficiency of an incoming nucleotide.


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  		TABLE 2
Binding affinity of Dpo4 to control and damaged DNA substrates at 23 °C

 
Efficiency of Nucleotide Incorporation in the Vicinity of the AP Lesion—To test the aforementioned hypothesis, single turnover kinetic experiments used previously (3) were employed to measure the maximum incorporation rate constant (kp) and the Kd of an incoming dNTP, which are used to calculate catalytic efficiency (kp/Kd). Initially we measured the kp and Kd values for nucleotide incorporation into 21-mer/41AP, corresponding to the DNA substrate at the first pause site where the AP site was the first "templating" position. Notably, this substrate also contained a template guanine immediately 5' to the AP site (Table 1). Assuming that Dpo4 would follow the A-rule for incorporation opposite the AP site, a solution containing Dpo4 (120 nM) was preincubated with 5'-radiolabeled 21-mer/41AP (30 nM) and reacted with increasing concentrations of dATP for various times (see under "Experimental Procedures"). The resulting product concentrations were plotted against reaction time and fit to Equation 1 (see under "Experimental Procedures") to yield individual kobs values at each corresponding dATP concentration (Fig. 3A). The extracted kobs values were then plotted against the dATP concentration and fit to Equation 2 (see under "Experimental Procedures") to yield a kp of 0.015 ± 0.002 s-1 and a Kd of 490 ± 173 µM (Fig. 3B). The kp/Kd of dATP was then calculated to be 3.1 x 10-5 µM-1 s-1 (Table 3), which was ~2,520-fold slower than a matched nucleotide incorporation into the control substrate 21-mer/41CTL (3). Strikingly, we observed that the catalytic efficiency for this and the remaining three nucleotide incorporations (dCTP, dGTP, and dTTP) into the 21-mer/41AP substrate dropped to the level observed previously for incorrect nucleotides (kp/Kd ~ 10-5 µM-1 s-1) into 21-mer/41CTL (3). Similar results were observed for all incorporations at the second pause site (Table 3) where incorporation efficiencies were in the range of 10-4–10-6 µM-1 s-1. Because all the catalytic efficiencies for incorporations at these two strong pause sites were of similar magnitude (Table 3) and several orders of magnitude smaller than a correct incorporation into the undamaged DNA (3), these data kinetically justified our hypothesis that the strong pause sites were caused by a significant reduction in catalytic efficiency.


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  		TABLE 3
Kinetic parameters for each nucleotide incorporation into damaged DNA at the two pause sites

 


Figure 3
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  		FIGURE 3.
Concentration dependence of dATP on nucleotide incorporation. A, preincubated solution of Dpo4 (120 nM) and 5'-labeled 21-mer/41AP (30 nM) was rapidly mixed with increasing concentrations of dATP (100 µM, •; 300 µM, {circ}; 500 µM, {blacksquare}; 800 µM, {square}; 1100 µM, {blacktriangleup}; 1400 µM, {triangleup}; 1800 µM, {diamondsuit}) for various reaction times. The solid lines are best fits to the single exponential equation. B, extracted observed rate constants from the above data fitting were plotted as a function of dATP concentration and fit to Equation 2 to obtain a kp of 0.015 ± 0.002 s-1 and a Kd of 490 ± 174 µM.

 
Many previous studies have demonstrated a proclivity of DNA polymerases to preferentially incorporate dATP opposite an AP site (A-rule). Table 3 indicated that Dpo4 followed the A-rule by favoring dATP over dGTP and dTTP into 21-mer/41AP; however, Dpo4 failed to incorporate dATP (3.1 x 10-5 µM-1 s-1) with a higher efficiency than dCTP (5.4 x 10-5 µM-1 s-1) in this sequence context. In contrast, the kp, Kd, and calculated kp/Kd values for both dGTP and dTTP were more similar to those of the least efficient misincorporation into an undamaged 21-mer/41CTL (3) and were 39- and 6-fold less likely to be incorporated, respectively (Table 3). The kinetic preference (1.7-fold) for dCTP over dATP incorporation may be due to the formation of a Watson-Crick base pair between dCTP and the template guanine 5' to the AP site. To evaluate this hypothesis, we synthesized a template 22AP that was identical in sequence to 41AP, except that it lacked all the template bases 5' to the AP site (Table 1). Upon measuring the catalytic efficiency for each single nucleotide incorporation into 21-mer/22AP, we observed dCTP was incorporated 33-fold less efficiently than dATP, indicating that the template information 5' to the AP site significantly influenced nucleotide incorporation (Table 4). These results, coupled with structural analysis (17) and additional data from our laboratory (see "Discussion"), demonstrated the existence of two competing pathways for bypass of an AP site catalyzed by Dpo4, the A-rule and the previously described 5'-rule (17), which involves incorporation directed by the template base 5' to the lesion in the context of an extrahelical AP site (Scheme 1).


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  		TABLE 4
Comparison of the substrate specificity for nucleotide incorporation into 21-mer/22AP

 
Nucleotide Incorporation Downstream from the AP Site—Under single turnover conditions, we also determined the incorporation efficiency of each dNTP for several downstream incorporation events from the AP site (supplemental Tables 1 and 2). Our data revealed that the effect from the embedded AP site on nucleotide incorporation decreased with distance, regardless of the pathway (A-rule or 5'-rule) (Fig. 4). Interestingly, the relatively low kp/Kd values before normalization suggested that Dpo4 paused slightly at several downstream incorporation events. This observed "weak pausing" was because of a lingering effect on the kp values, which incrementally diminished as Dpo4 moved farther from the embedded lesion, whereas all Kd values varied as observed previously with undamaged DNA (3). For the pathway with adenine incorporated opposite the AP site, the kinetic effect of an embedded lesion normalized after six incorporations (Fig. 4A), whereas the pathway with cytosine opposite the AP site normalized after five incorporations (Fig. 4B). The "weak pause sites" were less noticeable (Fig. 1B) because of their reduced effect on catalysis coupled with longer reaction times.

Measurement of the Possibility of Primer Realignment after Bypassing the AP Lesion—After Dpo4 bypassed the AP lesion via the 5'-rule, the embedded AP lesion may remain looped out or the primer may realign with the DNA template to force the incorporated cytosine to be located opposite the AP site during downstream incorporation events (Scheme 1). To determine the dominant branch pathway in Scheme 1, we evaluated the kp, Kd, and kp/Kd values for all four possible nucleotide incorporations at each position downstream from the lesion (supplemental Tables 1 and 2). For each substrate listed in Table 5, only the two dNTPs relevant to the looped-out and realignment pathways were shown. The realignment percentage values indicated that the looped-out branch pathway was dominant to the realignment pathway especially when the embedded AP lesion was further from the nascent base pair.


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  		TABLE 5
Realignment possibilities of the looped out DNA intermediates

a Realignment % = (1/(1 + efficiency ratio)) x 100%.

 
Sequence Dependence of the 5'-rule—To determine whether the 5'-rule was sequence-dependent, we measured the kinetic parameters (supplemental Table 3) for nucleotide incorporation into two additional DNA substrates, AP-1 and AP-8 (Table 6), where the template base immediately 5' to the AP site was changed to an adenine and a cytosine respectively (thymine is irrelevant in this context). Interestingly, the competition between the A-rule and the 5'-rule pathways was similar in magnitude for both AP-1 and AP-8 as observed with the substrate containing a 5' template guanine (21-mer/41AP; Table 3). For the AP-1 substrate, the competition between the A-rule and 5'-rule was indistinguishable, whereas for the AP-8 substrate, the A-rule was favored by ~4-fold. Therefore, all other incorporations, incorporations other than dCTP (21-mer/41AP)/dTTP (AP-1)/dGTP (AP-8) or dATP into these three DNA substrates, were on average 18-fold less likely to occur (supplemental Table 3). Incidentally, incorporation of dTTP (16.5%) had a similar probability to the incorporation of dGTP (17.3%) into AP-8 (supplemental Table 3), yet could not be explained based on these two bypass pathways. We had reported previously a similar observation where Dpo4 incorporated nucleotides onto a blunt-end DNA substrate with the following preference: dATP > dTTP > dCTP > dGTP (28). In addition, a previous report demonstrated a nearly equal probability of incorporation of dATP and dTTP opposite this lesion in vivo (33). Ultimately, we do not know the reason for the increased probability of dTTP incorporation into AP-8, but we suspect this trend is likely polymerase-dependent. Intriguingly, in these three sequence contexts (21-mer/41AP, AP-1, and AP-8), the preference for the A-rule was only observed in the presence of a 5' template cytosine. Thus, because of the inherent ambiguity in the nomenclature for the 5'-rule (17), as revealed by our aforementioned sequence dependence studies, we have decided to rename the pathway the "lesion loop-out mechanism" henceforth.


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  		TABLE 6
Preference between the A-rule and the lesion loop-out mechanism for bypass of an AP site

 


Figure 4
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  		SCHEME 1
 


Figure 5
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  		FIGURE 4.
Quantitative effect of the AP site on nucleotide incorporation. To determine the kinetic influence on the bypass of an AP site, the catalytic efficiency for the favored nucleotide incorporation, (kp/Kd)AP, for each AP-containing DNA substrate was divided by the catalytic efficiency for the corresponding correct incorporation into each control substrate, (kp/Kd)CTL, and was then plotted for each substrate. A, ratios for AP site bypass following the A-rule normalized after six incorporations. B, ratios for AP site bypass following the lesion loop-out mechanism normalized after five incorporations.

 
Incorporation of a Nonhydrogen Bonding Nucleotide Analog Opposite an AP Lesion—Interestingly, previous studies have shown that yeast pol {eta} requires Watson-Crick hydrogen bonds for efficient incorporation, unlike the geometric discrimination that helps govern the fidelity at the active sites of replicative DNA polymerases (34). To determine the effect of a nucleotide analog that lacked the ability to hydrogen bond with a template base on the incorporation opposite an AP site by Dpo4, we measured the kp/Kd values of pyrene nucleoside 5'-triphosphate (dPTP, supplemental Fig. 2) with 21-mer/41AP. dPTP served as a bifunctional probe because of its inability to hydrogen bond and its robust base-stacking abilities, attributed to its resonance-stabilized aromatic ring system. The single turnover experiments were carried out as described above, and we determined the kp to be 0.26 ± 0.02 s-1, the Kd to be 24 ± 4 µM (supplemental Fig. 2), and the kp/Kd to be 1.1 x 10-2 µM-1 s-1 which was, strikingly, 358- and 205-fold higher than dATP and dCTP incorporation, respectively (Table 3). The 20-fold higher ground-state binding affinity of dPTP over dATP (Table 3) indicated base stacking was the main contributor to the binding affinity of a nucleotide opposite an AP lesion at the Dpo4 active site.

Biphasic Kinetics of Nucleotide Incorporation at the First Pause Site—To provide a more detailed analysis for the kinetics of nucleotide incorporation at the first strong pause site, a preincubated solution of Dpo4 (120 nM) and 5'-32P-labeled 21-mer/41AP (30 nM) was mixed with a solution containing a nonradiolabeled 21-mer/41CTL DNA trap (5 µM) and either 1.2 mM dATP or 1.2 mM dCTP for various times before being quenched with 0.37 M EDTA. The product concentrations were calculated and fit to Equation 4

Formula 4(Eq. 4)
where Eo represents the total enzyme concentration; A1 and A2 represent the fast and slow phase reaction amplitude, respectively, and k1 and k2 represent the rate constants for the fast phase and slow phase, respectively. The trap DNA (~170-fold excess) functioned to remove all Dpo4 molecules that dissociated from the lesion-containing substrate. Interestingly, the results for both dATP and dCTP incorporation revealed biphasic kinetics, showing both a low amplitude, fast phase of nucleotide incorporation preceding a higher amplitude, slow phase. For dATP incorporation, the fast phase was determined to proceed with k1 of 0.25 ± 0.10 s-1 and A1 of 1.7 ± 0.1 nM (or 5.7%), whereas the slow phase was characterized by a k2 of 0.00124 ± 0.00009 s-1 and A2 of 22.2 ± 0.8 nM (or 74%) (Fig. 5). For dCTP incorporation, the fast phase had k1 of 0.24 ± 0.06 s-1 and A1 of 7.1 ± 0.3 nM (or 23.7%), whereas the slow phase was determined to have k2 of 0.0005 ± 0.0004 s-1 and A2 of 18.2 ± 5.9 nM (or 60.7%) (data not shown). Although Dpo4 was in molar excess to initially ensure all DNA substrate was bound, neither time course reached full amplitude (80% for dATP, and 84% for dCTP), suggesting that a portion of Dpo4 bound to the DNA substrate either dissociated or was unable to change from a nonproductively bound state to a productively bound state competent for catalysis.


Figure 6
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  		FIGURE 5.
Incorporation of dATP into 21-mer/41AP catalyzed by Dpo4 in the presence of a DNA trap showed biphasic kinetics. The product concentration versus reaction time were fit to Equation 4 that gave a fast phase reaction amplitude and rate constant of 1.7 ± 0.1 nM and 0.25 ± 0.10 s-1, respectively, whereas the slow phase was characterized by a rate constant of 0.00124 ± 0.00009 s-1 and an amplitude of 22.2 ± 0.8 nM.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The evolution of DNA repair pathways has provided cells with several different mechanisms to detect and eliminate DNA damage. However robust these systems may be, a population of DNA lesions will persist, halting the progression of the replication machinery and jeopardizing cell survival. With the recent discovery of a group of polymerases that promiscuously replicate through various types of DNA damage, we now know that lesion tolerance is a bona fide mechanism to rescue stalled replication forks. Here we use the powerful techniques of presteady state kinetics to dissect the mechanism of lesion bypass by the Y-family member Dpo4 as it approaches, incorporates across from, and synthesizes downstream from an embedded AP site lesion. Bypass of an AP lesion by Dpo4 was accomplished very efficiently in comparison to the replicative DNA polymerase pol B1 from S. solfataricus, which was completely stalled by the same AP lesion even after 10 min of incubation under the same reaction conditions (supplemental Fig. 1). However, in the process of AP lesion bypass, intermediate product accumulation indicated that Dpo4 paused strongly at one nucleotide preceding the AP site, which has never been reported previously, and at the position opposite the lesion. This strong pausing (Fig. 1B) suggested a perturbation in the mechanism of nucleotide incorporation during bypass.

Kinetic Studies Reveal Competition between Two Bypass Pathways—Our kinetic data in Table 3 demonstrated that Dpo4 preferentially incorporated dATP and dCTP when it bypassed a template AP site containing a guanine 5' to the lesion (Table 1). Although dATP was likely incorporated via the A-rule, incorporation of dCTP was governed by a different mechanism. Because of the well established structural flexibility of AP site lesions in DNA (29, 30, 35), the preference for dCTP led us to hypothesize that this nucleotide could be incorporated via a mechanism directed by the downstream template guanine involving an extrahelical AP site. Further evidence showed the following: (i) loss of the preference of dCTP incorporation into 21-mer/22AP (Table 4) relative to 21-mer/41AP (Table 3); (ii) extension assays of Dpo4 with the 21-mer/41AP (Table 1) in the presence of dideoxynucleotides showing full-length products containing -1 deletions (data not shown); and (iii) crystal structure evidence showing the ability of Dpo4 to "loop-out" an AP site (17). This evidence led us to conclude that incorporation of dCTP was because of the lesion loop-out mechanism (Scheme 1). Structure analysis showed the looped-out AP lesion can be accommodated in the cavity between the finger and little finger domains (17). In contrast, replicative DNA polymerases have been found to only follow the A-rule to bypass an AP site. This is because of their lack of the aforementioned structural cavity to accommodate the extrahelical AP site. Intriguingly, this kinetically observable intermediate in the A-rule pathway for Dpo4 was not able to be crystallized (17). This could be due to its weak thermostability or transient lifetime, which makes it difficult to be captured by x-ray crystallography. Thus, we conclude that AP bypass by Dpo4 follows both the A-rule and the lesion loop-out mechanism (Scheme 1) as opposed strictly to the latter as concluded previously (17).

AP Lesion Bypass via Different Pathways—Interestingly, the preference of these two pathways in Scheme 1 depended on the identity of the template base immediately 5' to the AP site (Table 6). When the 5'-base to the AP site was cytosine, the A-rule was preferred by 3.7-fold, indicating the mechanism of AP bypass was sequence-dependent. Neither pathway was significantly favored when the 5'-base was adenine. Overall, our data in Table 6 indicated that Dpo4 and possibly all Y-family DNA polymerases use both the A-rule and the lesion loop-out mechanism to select an incoming nucleotide opposite an AP site.

After Dpo4 followed the lesion loop-out mechanism to incorporate dCTP into 21-mer/41AP to form 22-C-mer/41AP (Table 1), further elongation of this product could occur either retaining the looped-out AP site or after primer/template realignment (Scheme 1). The preferred nucleotide was expected to form a Watson-Crick base pair with the corresponding template base of each of these two intermediates, e.g. dGTP for the looped-out intermediate and dCTP for the realigned DNA substrate. Interestingly, such a realigned DNA substrate has been crystallized with Dpo4 and an incoming dCTP (see ternary structure Ab-3 (17)). From the measured catalytic efficiencies of the incorporation of these two nucleotides into 22-C-mer/41AP (Table 3), we calculated the efficiency ratio of 5.2 and a realignment percentage of 16.1% (Table 5). These values indicated that the looped-out branch pathway was preferred over the realignment branch pathway at this position. This preference was more significant for downstream nucleotide incorporations (Table 5), suggesting the looped-out AP lesion will remain extrahelical until the primer is fully extended. The A-rule intermediates, which possessed an intrahelical AP site, were also stabilized by downstream incorporation events (supplemental Table 1). On the basis of our kinetic data, we proposed several major pathways in Scheme 1 for the AP lesion bypass catalyzed by Dpo4. The pathways in Scheme 1 also apply to AP-1 and AP-8, although the pathway preference varied depending on the base 5' to the AP site (Table 6). Notably, many other possible pathways were not included in Scheme 1 because of their reduced kinetic partitioning. However, the totality of possible bypass pathways implied by the kinetic partitioning of each incorporation event strongly suggested that the AP lesion will cause numerous downstream mutations during primer elongation (see below).

Insignificant Effect of an AP Lesion on the Affinity of the Dpo4·DNA Binary Complex—A plausible explanation for the pausing observed in the vicinity of the AP lesion during processive synthesis by Dpo4 could involve a dramatically reduced affinity of Dpo4 for the AP-containing DNA substrate. Although an AP site does not affect the global B-form conformation of DNA (29, 36), numerous studies have shown significant structural aberrations in the immediate vicinity of an AP lesion (29, 31, 36, 37). The resulting bending and kinking of the duplex DNA and disruption of the hydrogen bonding network proximal to the lesion (38) may be significant enough to disrupt contacts within the Dpo4 binding pocket, potentially accounting for the observed pausing. However, the measured affinity for the Dpo4·DNA binary complex was only modestly affected (2–4-fold) at the two strong pause sites, whereas the Kd values at the flanking nonpause/weak pause sites were indistinguishable from those of control DNA substrates (Table 2). Interestingly, our study concluded that the mechanism of bypass of an AP site by Dpo4 competed between incorporation of dATP (A-rule) and incorporation directed by the template base 5' to the AP site (lesion loop-out mechanism). Notably, when bypass was directed by the latter mechanism, a looped-out AP site required accommodation in the Dpo4 active site. A series of ternary structures of Dpo4 showed that single-base bulges in the template can be inserted into the gap between the finger and little finger domain, whereas extrahelical bases in the primer can fit in the DNA minor groove because of the unusually small finger and thumb domains (17). These structural observations are consistent with our quantitative results and help justify why Dpo4 binding to the AP site was not the cause of intermediate product accumulation in Fig. 1B.

Kinetic Mechanism for Dpo4 Pausing—Our systematic presteady kinetic analysis demonstrated that Dpo4 pausing was because of a significantly reduced catalytic efficiency (kp/Kd) near the AP lesion. Comparisons of the kp/Kd values for the two preferred incorporations opposite the AP site versus matched nucleotide incorporation into undamaged DNA (3) revealed decreases in kp/Kd of 2,560-fold (dATP, the A-rule) and 2,020-fold (dCTP, the lesion loop-out mechanism), indicating the extension of the DNA substrate 21-mer/41AP was slow. In contrast, the production of this substrate from 20-mer/41AP was as efficient as for undamaged DNA (Fig. 4A) thus resulting in 21-mer accumulation (Fig. 1B). Similarly, the elongation of 22-mer/41AP was difficult because even the most efficient nucleotide (dGTP) was incorporated into the most probable 22-mer/41AP substrate with 230-fold lower efficiency than into undamaged control DNA (Table 3) (3), leading to strong accumulation of 22-mer in Fig. 1B. Following the two strong pause sites, the catalytic efficiencies of downstream incorporation events were also lowered, but gradually normalized, regardless of the bypass pathway (Fig. 4).

The aforementioned reduction of nucleotide incorporation efficiency at the two strong pause sites was mainly because of significantly slower kp values, with respective decreases in the kp values via the A-rule and the lesion loop-out mechanism pathways of ~1,070- and 100-fold (Table 3) at the first pause site and 50- and 130-fold (Table 3) at the second pause site relative to undamaged DNA (3). Considering the two proposed mechanisms for AP site bypass in Scheme 1, the slow kp values observed opposite the lesion were justified. Incorporation via the A-rule most likely proceeded by means of nontemplated incorporation of dATP that was stabilized in the ground state by stacking interactions with the primer 3'-terminal base (adenine in our case). The lack of viable hydrogen bonding contacts between the incoming nucleotide (dATP) and this noncoding lesion precluded turnover rates on the same order as observed for matched nucleotide incorporation into undamaged DNA. In this respect, extension from this incorporation event (i.e. 22-A-mer/41AP) was also relatively slow (Table 3) because, once incorporated, structure analysis shows the adenine at the primer terminus shifts toward the template strand and stacks with the preceding bases of both strands (17). This inward shift of adenine in the primer strand increases the distance between the 3'-OH and the {alpha}-phosphate of the next incoming nucleotide (dCTP in structure Ab-3 (17)) to 7 Å. This is beyond the 3.4 Å observed for an optimum catalytically active DNA polymerase ternary complex (39), therefore retarding catalysis. A similar rate reduction was observed for incorporation via the lesion loop-out mechanism, which involved a shift of the downstream template strand into the active site generating an extrahelical AP site in which structural studies suggest the distance between the 3'-OH and the {alpha}-phosphate of the incoming dCTP is >4Å (17).


Figure 7
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  		SCHEME 2
 
X-ray crystal structure studies reveal that Dpo4 can form different types of ternary structures when it bypasses an AP site (17), suggesting that damaged DNA may not be bound specifically at the Dpo4 active site. Further interrogation into this hypothesis revealed biphasic kinetics for the incorporation of both dATP and dCTP (21-mer/41AP DNA substrate) opposite the AP site. The biphasic kinetic trace (Fig. 5) showed a fast phase of nucleotide incorporation with a low reaction amplitude followed by a slow phase of nucleotide incorporation with a significantly higher amplitude, thus suggesting the existence of two Dpo4-bound species as follows: a productive ternary complex E·D Pn·dNTP (D Pn denotes productively bound DNA) that was competent for catalysis and a separate nonproductive ternary complex E·D Nn·dNTP that was slowly converted to E·D Pn·dNTP or simply dissociated (Scheme 2). In contrast, nucleotide incorporation into the nonpause site 20-mer/41AP substrate (data not shown) showed only a fast phase with a rate constant of 7.0 s-1, suggesting that the large slow phase and the significantly smaller "fast" phase observed opposite the AP site contributed to strong Dpo4 pausing in Fig. 1B. Interestingly, because the assay in Fig. 5 was performed in the presence of a large molar excess of nonradio-labeled DNA trap (>150-fold), the tightly bound ternary complexes E·D Pn·dNTP and E·D Pn·dNTP could only be converted from substrate Dn to product Dn + 1 in a single binding event. Thus, it was possible to calculate the individual contribution of each population of productive and nonproductively bound Dpo4 ternary complex to the overall reaction rate constant (kp) measured under single turnover conditions. For example, when considering dATP incorporation, the summation of the contributions from the fast phase of nucleotide incorporation ((0.25 s-1) x (5.7% reaction amplitude)) and the slow phase ((0.00124 s-1) x (74% reaction amplitude)) gave an overall rate constant for the reaction of 0.0152 s-1, which was roughly equivalent to the kp of 0.015 s-1 (Table 3). Similarly, the total contribution of the fast phase ((0.24 s-1) x (23.7% reaction amplitude)) and the slow phase ((0.0005 s-1) x (60.7% reaction amplitude)) generated a rate constant of 0.0571 s-1, which was similar to the kp of 0.079 s-1 (Table 3) for dCTP incorporation. There was inherently more error in the latter calculation because the assay was performed at a subsaturating concentration of dCTP (1,200 µM) and as such the fast phase reaction amplitude was likely underestimated (Kd = 1,453 µM for dCTP incorporation into 21-mer/41AP). Because the kp was the primary determinant in the overall contribution to the catalytic efficiency and the catalytic efficiency was responsible for Dpo4 pausing (see above), these data suggest that because the rate constant for the fast phase (~0.25 s-1) was similar for both dATP and dCTP incorporation, the relative population of productively bound complex (5.7 versus 23.7%) brought about the preference for dCTP over dATP incorporation opposite the AP site (Table 6). This preference for productive ternary complex containing dCTP was most likely because of the fact that although extrahelical and intrahelical AP sites are in equilibrium in solution, a bulged AP site is more thermodynamically favorable especially when flanked by purines (40). In this example, the AP site was flanked by one pyrimidine and one purine (3'-PydXPuo-5'), and in this sequence context, the lesion loop-out mechanism was preferred by ~2-fold. Interestingly, when we change the 5' template base to adenine (3'-PydXPuo-5') and cytosine (3'-PydXPyd-5'), we observed a preference for the A-rule (no AP collapse) only in the latter context (~4-fold), which contained two flanking pyrimidines that are known to poorly base stack. This suggested that the identity of the flanking template bases may play an important role in influencing the mechanism by which Dpo4 prefers to bypass an AP site. Although the second strong pause site was not assayed to determine whether it too was characterized by biphasic kinetics, similar results were expected because this strong pause site was also characterized by a significant accumulation of intermediate product, again suggesting the existence of nonproductive complex.

Interestingly, the effect of the AP site on the ground-state binding affinity (Kd) of the two preferred dNTPs at the first pause site (Table 3) only decreased 2.4–20.8-fold with respect to matched nucleotide incorporation (3). This decrease (on average, 11.6-fold) in Kd corresponded to a free energy change ({Delta}{Delta}G) of 1.51 kcal/mol. Whereas these Kd differences were similar to the differences on undamaged DNA (3), they were also similar in magnitude to the decreases observed for preferred incorporation catalyzed by yeast pol {eta} opposite a cis-syn T-T dimer (11) and an 8-oxodG (10) with respect to matched incorporation on undamaged DNA, where differences in Kd values varied from 1.8- to 4.6-fold ({Delta}{Delta}G = 0.41 kcal/mol and 0.82 kcal/mol) opposite the 5' thymine and 3' thymine, respectively, and 2.1-fold ({Delta}{Delta}G = 0.43 kcal/mol) for incorporation opposite 8-oxodG. On the other hand, high fidelity replicative polymerases like human DNA polymerase {gamma} have significantly higher values for {Delta}{Delta}G ({Delta}{Delta}G = 4.0 kcal/mol) because of the greater discrimination between matched and mismatched nucleotides in the ground state attributed to more stringent fidelity checking mechanisms (41). These results suggested that the contribution from the active sites of Dpo4, yeast pol {eta}, and most likely all Y-family polymerases, in selecting the incoming nucleotide in the ground state when bypassing a DNA lesion, is quite insignificant. The deficiency in specific interactions of Dpo4 with the incoming nucleotide in the ground state suggested that nucleotide selection depended on its ability to stack with the base at the 3' end of the primer (A-rule) or to form Watson-Crick base pairing with the downstream template base (lesion loop-out mechanism). The base stacking effect was justified based on a 20-fold decrease in the Kd value from dPTP to dATP (Table 3) incorporation because of the superior aromatic {pi}-stacking capabilities of dPTP relative to dATP, the best stacking natural nucleotide (26, 42). Additionally, the stacking power of dPTP also facilitated catalysis by significantly increasing its kp value over those of dNTPs (Table 3).

Strong Pause Sites Are Mutational Hot Spots—Before Dpo4 encountered the AP site, our kinetic data with 20-mer/41AP (supplemental Table 4) suggested that the fidelity and likely the mechanism of nucleotide incorporation were equivalent to that reported for undamaged DNA (3, 8), yet this mechanism vastly differed opposite the lesion (Scheme 2) and at several downstream bases. When opposite the AP site, our kinetic data (Table 3) predicted that Dpo4 can insert each of the four dNTPs, although the incorporation percentages of dCTP (56.8%) and dATP (32.6%) were significantly higher because of the lesion loop-out mechanism and the A-rule, respectively. These data further suggested that bypass of an AP lesion by Dpo4 will cause either a "-1 frame-shift" or randomized substitutions. Moreover, our kinetic data suggested that substitution events occurred when Dpo4 extended these bypassed intermediates, especially 22-C-mer/41AP and 22-A-mer/41AP in which two dNTPs were preferentially incorporated (Table 3). Thus, our pre-steady state kinetic analysis predicted that the two strong pause sites in Fig. 1B were mutational hot spots and that the embedded AP lesion lowered downstream incorporation preference but gradually normalized (supplemental Tables 1 and 2). If these kinetic predictions are confirmed in vivo, bypassing AP lesions by Dpo4 or any Y-family member will be detrimental to genetic stability and thus would require tight cellular regulation.


   FOOTNOTES
 
* This work was supported in part by the National Science Foundation Career Award MCB-0447899 (to Z. S.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

Formula The on-line version of this article (available at http://www.jbc.org) contains supplemental Figs. 1 and 2 and Tables I–IV. Back

1 Recipient of American Heart Association Predoctoral Fellowship 0415129B and the Herta Camerer Gross Graduate Research Fellowship. Back

2 To whom correspondence should be addressed: 740 Biological Sciences, 484 West 12th Ave., Columbus, OH 43210. Tel.: 614-688-3706; Fax: 614-292-6773; E-mail: suo.3{at}osu.edu.

3 The abbreviations used are: Dpo4, S. solfataricus DNA polymerase IV; AP, abasic; dNTP, deoxynucleoside 5'-triphosphate; dPTP, pyrene nucleoside 5'-triphosphate; pol, polymerase. Back


   ACKNOWLEDGMENTS
 
We thank John-Stephen Taylor for providing us dPTP.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Burgers, P. M., Koonin, E. V., Bruford, E., Blanco, L., Burtis, K. C., Christman, M. F., Copeland, W. C., Friedberg, E. C., Hanaoka, F., Hinkle, D. C., Lawrence, C. W., Nakanishi, M., Ohmori, H., Prakash, L., Prakash, S., Reynaud, C. A., Sugino, A., Todo, T., Wang, Z., Weill, J. C., and Woodgate, R. (2001) J. Biol. Chem. 276, 43487-43490[Free Full Text]
  2. Boudsocq, F., Iwai, S., Hanaoka, F., and Woodgate, R. (2001) Nucleic Acids Res. 29, 4607-4616[Abstract/Free Full Text]
  3. Fiala, K. A., and Suo, Z. (2004) Biochemistry 43, 2106-2115[CrossRef][Medline] [Order article via Infotrieve]
  4. Kokoska, R. J., Bebenek, K., Boudsocq, F., Woodgate, R., and Kunkel, T. A. (2002) J. Biol. Chem. 277, 19633-19638[Abstract/Free Full Text]
  5. Ohashi, E., Bebenek, K., Matsuda, T., Feaver, W. J., Gerlach, V. L., Friedberg, E. C., Ohmori, H., and Kunkel, T. A. (2000) J. Biol. Chem. 275, 39678-39684[Abstract/Free Full Text]
  6. Zhang, Y., Yuan, F., Xin, H., Wu, X., Rajpal, D. K., Yang, D., and Wang, Z. (2000) Nucleic Acids Res. 28, 4147-4156[Abstract/Free Full Text]
  7. Washington, M. T., Prakash, L., and Prakash, S. (2001) Cell 107, 917-927[CrossRef][Medline] [Order article via Infotrieve]
  8. Fiala, K. A., and Suo, Z. (2004) Biochemistry 43, 2116-2125[CrossRef][Medline] [Order article via Infotrieve]
  9. Cramer, J., and Restle, T. (2005) J. Biol. Chem. 280, 40552-40558[Abstract/Free Full Text]
  10. Carlson, K. D., and Washington, M. T. (2005) Mol. Cell. Biol. 25, 2169-2176[Abstract/Free Full Text]
  11. Washington, M. T., Prakash, L., and Prakash, S. (2003) Proc. Natl. Acad. Sci. U. S. A. 100, 12093-12098[Abstract/Free Full Text]
  12. Lindahl, T. (1982) Annu. Rev. Biochem. 51, 61-87[CrossRef][Medline] [Order article via Infotrieve]
  13. Lindahl, T., and Nyberg, B. (1972) Biochemistry 11, 3610-3618[CrossRef][Medline] [Order article via Infotrieve]
  14. Lindahl, T., and Andersson, A. (1972) Biochemistry 11, 3618-3623[CrossRef][Medline] [Order article via Infotrieve]
  15. She, Q., Singh, R. K., Confalonieri, F., Zivanovic, Y., Allard, G., Awayez, M. J., Chan-Weiher, C. C., Clausen, I. G., Curtis, B. A., De Moors, A., Erauso, G., Fletcher, C., Gordon, P. M., Heikamp-de Jong, I., Jeffries, A. C., Kozera, C. J., Medina, N., Peng, X., Thi-Ngoc, H. P., Redder, P., Schenk, M. E., Theriault, C., Tolstrup, N., Charlebois, R. L., Doolittle, W. F., Duguet, M., Gaasterland, T., Garrett, R. A., Ragan, M. A., Sensen, C. W., and Van der Oost, J. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 7835-7840[Abstract/Free Full Text]
  16. Strauss, B., Rabkin, S., Sagher, D., and Moore, P. (1982) Biochimie (Paris) 64, 829-838
  17. Ling, H., Boudsocq, F., Woodgate, R., and Yang, W. (2004) Mol. Cell 13, 751-762[CrossRef][Medline] [Order article via Infotrieve]
  18. Joyce, C. M., and Benkovic, S. J. (2004) Biochemistry 43, 14317-14324[CrossRef][Medline] [Order article via Infotrieve]
  19. Manoharan, M., Ransom, S. C., Mazumder, A., Gerlt, J. A., Wilde, J. A., Withka, J. A., and Bolton, P. H. (1988) J. Am. Chem. Soc. 110, 1620-1622
  20. Wilde, J. A., Bolton, P. H., Mazumder, A., Manoharan, M., and Gerlt, J. A. (1989) J. Am. Chem. Soc. 111, 1894-1896
  21. Kokoska, R. J., McCulloch, S. D., and Kunkel, T. A. (2003) J. Biol. Chem. 278, 50537-50545[Abstract/Free Full Text]
  22. Johnson, R. E., Washington, M. T., Haracska, L., Prakash, S., and Prakash, L. (2000) Nature 406, 1015-1019[CrossRef][Medline] [Order article via Infotrieve]
  23. Zhang, Y., Wu, X., Dongyu, G., Rechkoblit, O., Taylor, J. S., Geacintov, N. E., and Wang, Z. (2002) J. Biol. Chem. 277, 44582-44587[Abstract/Free Full Text]
  24. Masutani, C., Kusumoto, R., Iwai, S., and Hanaoka, F. (2000) EMBO J. 19, 3100-3109[CrossRef][Medline] [Order article via Infotrieve]
  25. Hogg, M., Wallace, S. S., and Doublie, S. (2004) EMBO J. 23, 1483-1493[CrossRef][Medline] [Order article via Infotrieve]
  26. Matray, T. J., and Kool, E. T. (1999) Nature 399, 704-708[CrossRef][Medline] [Order article via Infotrieve]
  27. Takeshita, M., Chang, C.-N., Johnson, F., Will, S., and Grollman, A. (1987) J. Biol. Chem. 262, 10171-10179[Abstract/Free Full Text]
  28. Fiala, K. A., Brown, J. A., Ling, H., Kshetry, A. K., Zhang, J., Taylor, J. S., Yang, W., and Suo, Z. (2007) J. Mol. Biol. 365, 590-602[CrossRef][Medline] [Order article via Infotrieve]
  29. Gelfand, C. A., Plum, G. E., Grollman, A. P., Johnson, F., and Breslauer, K. J. (1998) Biochemistry 37, 7321-7327[CrossRef][Medline] [Order article via Infotrieve]
  30. Ide, H., Shimizu, H., Kimura, Y., Sakamoto, S., Makino, K., Glackin, M., Wallace, S. S., Nakamuta, H., Sasaki, M., and Sugimoto, N. (1995) Biochemistry 34, 6947-6955[CrossRef][Medline] [Order article via Infotrieve]
  31. Wang, K. Y., Parker, S. A., Goljer, I., and Bolton, P. H. (1997) Biochemistry 36, 11629-11639[CrossRef][Medline] [Order article via Infotrieve]
  32. Beger, R. D., and Bolton, P. H. (1998) J. Biol. Chem. 273, 15565-15573[Abstract/Free Full Text]
  33. 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]
  34. Washington, M. T., Helquist, S. A., Kool, E. T., Prakash, L., and Prakash, S. (2003) Mol. Cell. Biol. 23, 5107-5112[Abstract/Free Full Text]
  35. Vesnaver, G., Chang, C.-N., Eisenberg, M., Grollman, A. P., and Breslauer, K. J. (1989) Proc. Natl. Acad. Sci. U. S. A. 86, 3614-3618[Abstract/Free Full Text]
  36. Groljer, I., Kumar, S., and Bolton, P. H. (1995) J. Biol. Chem. 270, 22980-22987[Abstract/Free Full Text]
  37. Ayadi, L., Coulombeau, C., and Lavery, R. (1999) Biophys. J. 77, 3218-3226
  38. Barsky, D., Foloppe, N., Ahmadia, S., Wilson, D. M., and MacKerell, A. D. (2000) Nucleic Acids Res. 28, 2613-2626[Abstract/Free Full Text]
  39. Batra, V. K., Beard, W. A., Shock, D. D., Krahn, J. M., Pedersen, L. C., and Wilson, S. H. (2006) Structure (Lond.) 14, 757-766[Medline] [Order article via Infotrieve]
  40. Cuniasse, P., Fazakerley, G. V., Guschlbauer, W., Kaplan, B. E., and Sowers, L. C. (1990) J. Mol. Biol. 213, 303-314[CrossRef]