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Originally published In Press as doi:10.1074/jbc.M607479200 on September 27, 2006

J. Biol. Chem., Vol. 281, Issue 47, 35649-35655, November 24, 2006
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Kinetic Effect of a Downstream Strand and Its 5'-Terminal Moieties on Single Nucleotide Gap-filling Synthesis Catalyzed by Human DNA Polymerase {lambda}*

Wade W. Duym{ddagger}, Kevin A. Fiala{ddagger}§1, Nikunj Bhatt{ddagger}, and Zucai Suo{ddagger}§||**2

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

Received for publication, August 7, 2006 , and in revised form, September 21, 2006.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
During short-patch base excision repair, the excision of a 5'-terminal 2-deoxyribose-5-phosphate moiety of the downstream strand by the 5'-2-deoxyribose-5-phosphate lyase activity of either DNA polymerase beta or {lambda} is believed to occur after each respective enzyme catalyzes gap-filling DNA synthesis. Yet the effects of this 5'-terminal 2-deoxyribose-5-phosphate moiety on the polymerase activities of these two enzymes have never been quantitatively determined. Moreover, x-ray crystal structures of truncated polymerase {lambda} have revealed that the downstream strand and its 5'-phosphate group of gapped DNA interact intensely with the dRPase domain, but the kinetic effect of these interactions is unclear. Here, we utilized pre-steady state kinetic methods to systematically investigate the effect of a downstream strand and its 5'-moieties on the polymerase activity of the full-length human polymerase {lambda}. The downstream strand and its 5'-phosphate were both found to increase nucleotide incorporation efficiency (kp/Kd) by 15and 11-fold, respectively, with the increase procured by the effect on the nucleotide incorporation rate constant kp rather than the ground state nucleotide binding affinity Kd. With 4 single nucleotide-gapped DNA substrates containing a 1,2-dideoxyribose-5-phosphate moiety, a 2-deoxyribose-5-phosphate mimic, we measured the incorporation efficiencies of 16 possible nucleotides. Our results demonstrate that although this 5'-terminal 2-deoxyribose-5-phosphate mimic does not affect the fidelity of polymerase {lambda}, it moderately decreased the polymerase efficiency by 3.4-fold. Moreover, this decrease in polymerase efficiency is due to a drop of similar magnitude in kp rather than Kd. The implication of the downstream strand and its 5'-moieties on the kinetics of gap-filling synthesis is discussed.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
In mammalian cells, single base lesions are the most common form of DNA damage that arises either from exogenous DNA-damaging agents (1) or from endogenous biological processes resulting in base alkylation (2, 3), base oxidation (4), spontaneous cytosine deamination (3), and hydrolytic base loss (3, 5, 6). Base excision repair (BER)3 is the major pathway to repair single base lesions (7). Short-patch and long-patch BER are the two subpathways of BER that remove and replace 1 (8–10) and 2–11 nucleotides (11–13), respectively. Short-patch BER starts with the excision of a modified base by a DNA glycosylase, leaving a noncoding apurinic or apyrimidinic site in DNA. This lesion is further processed and repaired by a 5'-acting apurinic or apyrimidinic endonuclease, a DNA polymerase, a 5'-2-deoxyribose-5-phosphate lyase (dRPase), and a DNA ligase (12, 14–16). It has been established that in mammalian systems DNA polymerase beta (Fig. 1, Polbeta), an X-family DNA polymerase, plays a critical role in short-patch BER (8, 9). The polymerase activity of Polbeta catalyzes single nucleotide gap-filling synthesis (17), while its dRPase activity removes the 5'-terminal 2-deoxyribose-5-phosphate moiety (dRP) of a downstream strand (18). The uracil-initiated short-patch BER has been reconstituted in vitro by using purified recombinant human enzymes (19–21), and its reaction sequences (Scheme 1) have been established from steady state kinetic studies (21).

Like Polbeta, the full-length DNA polymerase {lambda} (fPol{lambda}), a recently discovered member of the X-family DNA polymerases, also contains a dRPase (8 kDa) and a DNA polymerase (31 kDa) domain on its C terminus (Fig. 1) (22–25). In addition, fPol{lambda} possesses a nuclear localization signal motif, a breast cancer susceptibility gene 1 C-terminal (BRCT) domain, and a prolinerich domain. Although the biological role of fPol{lambda} has not been clearly identified, it is plausible that fPol{lambda} contributes to BER because it shares 33% sequence identity to Polbeta and possesses two key enzymatic activities required by BER. This hypothesis is directly supported by the following observations: (i) recombinant human fPol{lambda} purified from Escherichia coli can replace human Polbeta in an in vitro reconstituted short-patch BER assay (26); (ii) mouse embryonic fibroblast Polbeta–/– cell extract contains a substantial amount of active fPol{lambda} that can also replace Polbeta in a similar in vitro reconstituted BER assay, and monoclonal antibodies against fPol{lambda} in this cell extract strongly reduce in vitro BER (27). fPol{lambda}, like Polbeta, lacks 3' -> 5' exonuclease activity (22–24) and has low processivity when copying non-gapped or large gap DNA (25). With short gap DNA, the downstream strand, especially one with a 5'-phosphate, has been shown to increase polymerase processivity of both Polbeta (28) and fPol{lambda} (25, 29). Moreover, Polbeta has been found to incorporate a single nucleotide ~10-fold more efficiently with single nucleotide-gapped DNA than non-gapped DNA (30, 31), but the polymerase fidelity is not altered (31). Structural evidence suggests that the increase in polymerase processivity and efficiency is due to additional contacts that are established in a gapped DNA substrate between the dRPase domains of these two enzymes and the downstream strand (29, 32). The terminal 5'-phosphate on the downstream strand is buried in a positively charged pocket of the dRPase active site (29, 32, 33). So far, the effects of a downstream strand and its 5'-phosphate on the efficiency and fidelity of gap-filling synthesis catalyzed by fPol{lambda} has not been quantitatively determined. Here, these effects associated with human fPol{lambda} will be investigated through presteady state kinetic studies.


Figure 1
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  		FIGURE 1.
Schematic representations of the domain organizations of human DNA polymerases beta and {lambda} (the full-length and the C-terminal fragment). Each domain, with amino acid residue numbers indicated above, is shown as a rectangle.

 


Figure 2
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  		SCHEME 1.
Reaction sequence of uracil-initiated short-patch BER.

 
Moreover, steady state kinetic studies have identified the rate-limiting step in the reconstituted and uracil-initiated BER system (Scheme 1) as the removal of the dRP moiety by Polbeta (21). fPol{lambda} is estimated to possess a 4-fold slower dRPase activity than Polbeta (26), while these two enzymes catalyze single nucleotide gap-filling DNA synthesis with <2-fold difference in catalytic efficiency (34). These kinetic data strongly indicate that the 5'-dRP moiety in the downstream strand is removed after Polbeta or fPol{lambda} fills single nucleotide-gapped DNA in Scheme 1. However, the effect of the 5'-dRP moiety on the fidelity and efficiency of either Polbeta or fPol{lambda} has never been kinetically evaluated. In this report, we will use single nucleotide-gapped DNA substrates containing a dRP mimic on the 5'-terminus of the downstream strand to determine the kinetic effect of the 5'-dRP moiety on gap-filling DNA synthesis catalyzed by human fPol{lambda} through detailed pre-steady state kinetic analysis.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—These chemicals were purchased from the following companies: [{gamma}-32P]ATP, GE Healthcare; Biospin columns, Bio-Rad Laboratories; dNTPs, Invitrogen; T4 polynucleotide kinase, USB (Cleveland, OH). The full-length human Pol{lambda} was cloned, expressed, and purified as described previously (34).

Synthetic Oligonucleotides—The DNA substrates shown in Fig. 2 were purchased from Integrated DNA Technologies (Coralville, IA) and purified by denaturing polyacrylamide gel electrophoresis (18% acrylamide, 8 M urea). Their concentrations were determined by measuring UV absorbance at 260 nm with calculated molar extinction coefficients. The primer strand 21-mer was 5'-32P-labeled by incubation with T4 polynucleotide kinase and [{gamma}-32P]ATP for 1 h at 37 °C. The unreacted [{gamma}-32P]ATP was subsequently removed by centrifugation through a Biospin-6 column (Bio-Rad). The 5'-32P-labeled primer 21-mer was then annealed with the corresponding non-radio-labeled downstream strand 19-mer to the template 41-mer at a molar ratio of 1.0:1.25:1.15, respectively, to form a 21–19/41-mer single nucleotide-gapped substrate (the top strand was composed of two oligonucleotides with a single nucleotide gap). Mixtures to be annealed were denatured at 95 °C for 8 min and then cooled slowly to room temperature over several hours.

Optimized Reaction Buffer L—All kinetic experiments were performed in buffer L containing 50 mM Tris-Cl (pH 8.4 at 37 °C), 5 mM MgCl2, 100 mM NaCl, 0.1 mM EDTA, 5 mM dithiothreitol, 10% glycerol, and 0.1 mg/ml bovine serum albumin. All reactions were carried out at 37 °C.

Measurement of Equilibrium Dissociation Constant of Next Incoming Nucleotide—A preincubated solution of 150 nM human fPol{lambda} and 30 nM 32P-labeled DNA was rapidly mixed with increasing concentrations of dNTP (0.25–120 µM) in buffer L in a rapid chemical quench-flow apparatus (KinTek) to initiate the reaction. The reactions at each concentration of dNTP were terminated with 0.37 M EDTA at varying times from milliseconds to minutes. Reaction products were analyzed by sequencing gel electrophoresis (17% acrylamide, 8 M urea, 1x Tris borate-EDTA buffer) and quantitated with a PhosphorImager 445 SI (GE Healthcare). The time course of product formation was fit to a single exponential equation (Equation 1) for each concentration of dNTP to give the observed rate constant of nucleotide incorporation (kobs). "A" represents the reaction amplitude, which is equal to the initial concentration of the enzyme and DNA binary complex. The observed rate constants extracted from these time courses of product formation were then plotted against the concentrations of dNTP, and these data were fit via hyperbolic regression (Equation 2) to give the equilibrium dissociation constant of dNTP, Kd, and the maximum rate constant for incorporation of dNTP, kp. The substrate specificity and polymerase fidelity were calculated as (kp/Kd) and (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect], respectively, as shown in Equations 1 and 2.

Formula 1(Eq. 1)

Formula 2(Eq. 2)


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Model DNA Substrates—Based on the steady state kinetic results (21), the natural DNA substrate for the polymerase activity of fPol{lambda} in BER is a dRP-DNA-type substrate (Fig. 2A) that exists as a mixture of different chemical species in solution (see "Discussion") (35). To simplify kinetic analysis, we prepared four H-DNA substrates, shown in Fig. 2B, whose downstream strands contained a 5'-terminal 1,2-dideoxyribose-5-phosphate moiety that mimicked 5'-dRP. The H-DNA substrates are stable and exist in one chemical form in solution due to the lack of C1-hydroxyl group in their dRP-mimic moiety. To examine the kinetic effect of the downstream strand and its 5'-phosphate, we designed another gapped substrate, D-1OH (Fig. 2D). This substrate contained a downstream 19-mer that was not 5'-phosphorylated as in D-DNA (D-1) (Fig. 2C) and non-gapped D-1N (Fig. 2E), which lacked a downstream strand.

Kinetic Effect of a dRP Mimic—Previously, we have established a minimal kinetic mechanism (Scheme 2) for dTTP incorporation onto D-DNA (D-1) (Fig. 2C) catalyzed by the C-terminal Polbeta-like domain of human Pol{lambda} (Fig. 1, tPol{lambda}) (36). This scheme shows that an incoming dNTP binds to the tPol{lambda}·D-1 binary complex to establish a rapid equilibrium prior to nucleotide incorporation. We have also demonstrated that human fPol{lambda} follows the same minimal mechanism shown in Scheme 2 (34). This mechanism allows us to measure the apparent affinity of dNTP (Kd) for the Pol{lambda}·DNA binary complex via the dNTP concentration dependence of the observed single turnover rate constant (kobs). The polymerase efficiency and fidelity of both tPol{lambda} (36) and the fPol{lambda} (34) have thus been determined. Here, we utilized similar pre-steady state kinetic methods to determine the kinetics of all 16 possible single nucleotide incorporations onto the 4 DNA substrates shown in Fig. 2B.


Figure 3
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  		FIGURE 2.
DNA substrates. A, dRP-DNA. B, H-DNA. C, D-DNA. D, D-1OH. E, D-1N. The 5' terminus of the downstream strand 19-mer contains either a dRP (A), a 1,2-dideoxyribose-5-phosphate (B), a phosphate moiety (C), or a hydroxyl moiety (D). X represents one of the four natural bases (e.g. X=A, H-1; X=G, H-6; X=T, H-7; X=C, H-8), while the circled P denotes phosphate. The upstream primer 21-mer, rather than the template 41-mer, was 5'-32P-labeled.

 


Figure 4
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  		SCHEME 2.
Minimal kinetic mechanism for dNTP incorporation onto D-DNA.

 
Under single turnover conditions, a preincubated solution of 30 nM 5'-32P-labeled H-7 (Fig. 2B) and 150 nM human fPol{lambda} was reacted with increasing concentration of an incoming dATP in buffer L (see "Experimental Procedures"). The single turnover method was employed because DNA dissociates from fPol{lambda} with a dissociation rate constant (k1) that is only ~2- to 3-fold slower than the maximum nucleotide incorporation rate constant kp (36), rendering the burst phase insignificant. Thus, the experiments were performed with fPol{lambda} in molar excess over DNA to allow the direct observation of nucleotide incorporation in a single pass of the reactants through the enzymatic pathway without complications resulting from the steady state formation of products (37). The DNA product 22-mer and remaining primer 21-mer were separated by gel electrophoresis and quantitated with a PhosphorImager. The product concentration was plotted against reaction time intervals. These data were subsequently fit to Equation 1 (see "Experimental Procedures") to yield an observed single turnover rate constant (kobs) at each concentration of dATP (Fig. 3A). The observed single turnover rate constants were then plotted against dATP concentration (Fig. 3B). These data were subsequently fit to Equation 2 (see "Experimental Procedures") to yield a kp of 0.40 ± 0.01 s–1 for the maximum dATP incorporation rate constant and a Kd of 0.78 ± 0.08 µM for dATP binding. The substrate specificity or polymerase efficiency (kp/Kd) of dATP incorporation onto H-7 was calculated to be 0.485 µM–1 s–1 (Table 1).


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  		TABLE 1
Kinetic parameters of nucleotide incorporation onto single nucleotide-gapped H-DNA containing a dRP mimic catalyzed by human fPol{lambda} at 37°C

 


Figure 5
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  		FIGURE 3.
Concentration dependence on the pre-steady state rate constant of correct nucleotide incorporation. A, a preincubated solution of fPol{lambda} (150 nM) and 5'-32P-labeled H-7 (30 nM) was rapidly mixed with increasing concentrations of Mg2+-dATP (0.5 µM, {blacktriangleup};1 µM, {square};2 µM, X;4 µM, {diamondsuit};8 µM, +;12 µM, {triangleup};25 µM, •;48 µM, {blacksquare}) for various time intervals. The solid lines are the best fits to Equation 1. B, the observed rate constants obtained from the above data fitting were plotted as a function of dATP concentration. The data (•) were then fit to Equation 2, yielding a k of 0.40 ± 0.01 s–1p and a Kd of 0.78 ± 0.08 µM.

 
The kinetic parameters (Table 1) for the incorporation of each of the remaining three correct nucleotides (dTTP onto H-1, dCTP onto H-6, and dGTP onto H-8) and for the incorporation of each of the 12 possible misincorporations onto H-1, H-6, H-7, and H-8 (Fig. 2B) were carried out in the same manner as described above. Notably, the ground state binding affinity of all nucleotides to fPol{lambda}·(H-DNA) (Table 1) were within 10-fold, but the kp values of correct nucleotides were three to five orders of magnitude higher than those of incorrect nucleotides. The differences in kp and Kd led to significantly different nucleotide incorporation efficiencies (kp/Kd) between correct and incorrect nucleotide incorporations and resulted in a fidelity of gap-filling synthesis in the range of 10–4 to 10–5 (Table 1).

Kinetic Effect of the 5'-Phosphate of a Downstream Strand—The kinetic parameters of nucleotide incorporation onto D-1OH (Fig. 2D) were measured with fPol{lambda} under the single turnover conditions described above and listed in Table 2. The apparent binding affinity of all four nucleotides was unaffected by the absence of the 5'-phosphate of the 19-mer in D-1OH, but the kp values were lowered by 10- to 100-fold (Table 2). Thus, the substrate specificity (kp/Kd) of matched and mismatched nucleotides with D-1OH was 10- and 100-fold smaller, respectively, than those with D-DNA (D-1) (Table 2). Notably, dATP incorporation onto D-1OH was too slow to be measured. A slightly larger effect on misincorporations compared with correct dTTP incorporation resulted in a slightly higher fidelity with D-1OH than with D-DNA (D-1).


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  		TABLE 2
Pre-steady state kinetic parameters with D-DNA (D-1), D-1OH, and D-1N

 
Kinetic Effect of a Downstream Strand—The significant kinetic effect of the 5'-phosphate moiety suggested the downstream strand itself could have a dramatic influence on the single nucleotide gap-filling efficiency of fPol{lambda}. To examine this hypothesis, we measured the pre-steady state kinetic parameters (Table 2) of single nucleotide incorporation onto D-1N that lacked a downstream strand (Fig. 1E) by employing single turnover experiments as described above. The correct dTTP was incorporated at a rate constant of 0.025 s–1 and a ground state binding affinity of 2.6 µM (Table 2). In comparison with the kinetic data with D-1OH (Table 2), the lack of a downstream strand further decreased the incorporation efficiency of matched dTTP and mismatched dCTP by approximately an additional 10-fold each. Incorporations of mismatched dGTP and dATP onto D-1N were too slow to be observed in several hours.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The short-patch base excision repair pathway involves at least five enzymes that catalyze six reaction steps, including two catalyzed by Polbeta (Scheme 1). Many lines of biochemical and in vitro evidence (22–27, 38, 39) indicate that the polymerase and dRPase activities of fPol{lambda} are able to catalyze these two BER steps in vitro and may do so in vivo. Under the in vitro reconstituted BER reaction conditions, the velocity values for the uracil-DNA glycosylase, apurinic or apyrimidinic endonuclease, the DNA polymerase and dRPase activities of Polbeta, and DNA ligase I have been measured to be 420, 100, 4.5, 0.75, 4.0 nM/s, respectively (21). The rate-limiting step is identified as the dRP excision reaction that occurs at a similar velocity as the overall reconstituted BER system (0.6 nM/s) (21). In addition, the polymerase activity of Polbeta that catalyzes the single nucleotide gap-filling synthesis is 6-fold faster than its dRPase activity which cleaves the dRP moiety on the 5' terminus of the downstream strand. This suggests that the natural single nucleotide-gapped DNA for the polymerase activity of Polbeta should contain a dRP moiety on its downstream strand. Thus, the best model DNA substrates are the dRP-DNA substrates (Fig. 2A) rather than the D-DNA substrates (Fig. 2C). However, because the 2-deoxyribose moiety in dRP-DNA is an equilibrium mixture of {alpha}- and beta-hemiacetals (2-deoxy-D-erythro-pentofuranoses), an aldehyde, and a hydrated aldehyde in solution (35), rigorous kinetic analysis becomes overly complicated; as such, D-DNA substrates have been predominantly used as the gapped DNA substrates to examine the kinetics of nucleotide incorporation catalyzed by both Polbeta (30, 31, 40–44) and Pol{lambda} (34, 36). To overcome the instability of dRP-DNA in solution, we decided to use the H-DNA substrates (Fig. 2B) in our presteady state kinetic analysis with human fPol{lambda}.

Under single turnover conditions, the kinetic parameters of all 16 possible nucleotide incorporations were measured using the H-DNA substrates (Fig. 2B), and the fidelity of human fPol{lambda} was determined to be in the range of 10–4 to 10–5 (Table 1). This range was identical to the fidelity with D-DNA (34), suggesting the presence of the dRP mimic did not affect the gap-filling error frequency of fPol{lambda}. However, the substrate specificity values (Table 1) are 1.6- to 6.7-fold lower than those corresponding values for nucleotide incorporation onto D-DNA (34), resulting in an average efficiency ratio, (kp/Kd)D-DNA/(kp/Kd)H-DNA, of 3.4 (Table 1). The difference in substrate specificity (kp/Kd) can be mathematically attributed to either kp, Kd, or both. However, the ground state binding affinities (Kd) of both correct and incorrect nucleotides with H-DNA (Table 1) are very similar to those corresponding Kd values obtained with D-DNA (34), indicating that the presence of the dRP mimic on the 5' terminus of the downstream strand did not affect the binding affinity of an incoming nucleotide. In contrast, the 16 maximum incorporation rate constants (kp) with H-DNA on average were 3.5-fold lower than those corresponding kp values with D-DNA, suggesting the presence of the dRP mimic moderately affected catalysis during nucleotide incorporation. However, what is the catalytic efficiency of fPol{lambda} with the natural dRP-DNA substrates (Fig. 2A)? Because of the close chemical similarity between dRP-DNA and H-DNA (Fig. 2), it is reasonable to speculate that fPol{lambda} will incorporate nucleotides onto dRP-DNA with efficiencies closer to those observed with H-DNA than with D-DNA. This hypothesis goes against the qualitative experiments of Srivastava et al. (21) in which they have observed similar nucleotide incorporation efficiency with Polbeta in the presence or absence of the dRP moiety. It is known that the dRP excision catalyzed by the dRPase domain occurs via a Schiff-base formation and beta-elimination (45–47). If the Schiff-base formation between the C1 of the dRP group and Lys-312 of fPol{lambda} (29, 48) is faster than nucleotide incorporation while the beta-elimination limits both dRP excision and BER, it is possible that fPol{lambda} will be more efficient with dRP-DNA than with H-DNA because the covalent anchoring of the downstream strand of dRP-DNA may facilitate gap-filling DNA synthesis catalyzed by fPol{lambda}. More kinetic experiments are required to examine the aforementioned hypothesis. From a structural perspective, the effect of the dRP moiety or its mimic on DNA, dNTP binding, and the polymerase active site conformation remains unclear, because all ternary crystal structures of both tPol{lambda} (33, 49) and Polbeta (32, 50, 51) are solved in the presence of "D-DNA," rather than "dRP-DNA" or "H-DNA." However, these structures do reveal intimate contacts between the dRPase domains of both Polbeta and tPol{lambda} (Fig. 1) and the downstream strand (29, 32). The terminal 5'-phosphate of the downstream strand is buried in a positively charged pocket of the dRPase active site (29, 32), e.g. Tyr-267, Arg-275, Tyr-279, Lys-307, and Arg-308 in tPol{lambda} (29, 48). Thus, the dRP mimic likely interacts with the dRPase domain, which in turn may affect the active site conformation of fPol{lambda} and thus the kp value of nucleotide incorporation.

Consistently, the intimate interactions between the dRPase domain, the downstream strand, and its 5'-phosphate moiety did impact catalysis significantly. Table 3 indicates that fPol{lambda} incorporated a matched dTTP most efficiently with single nucleotide-gapped D-DNA (D-1) (Fig. 2C). The absence of the 5'-phosphate group in D-1OH (Fig. 2D) caused an 11-fold decrease in dTTP incorporation efficiency (kp/Kd), whereas lack of the entire downstream strand in D-1N (Fig. 2E) led to an additional 15-fold decrease. As a consequence, the catalytic efficiency decreased by 160-fold from D-DNA (D-1) to D-1N. Similarly, Polbeta is found to have 6- to 40-fold higher nucleotide incorporation efficiency with single nucleotide-gapped DNA than with non-gapped DNA (52, 53). The low substrate specificity of matched dTTP with D-1N further suggested that fPol{lambda} is too inefficient to be a primer/template-dependent polymerase. In contrast, the dTTP incorporation efficiency of fPol{lambda} (1.5 µM–1s–1) is close to the range of Polbeta (1.9–8.5 µM–1s–1) in the presence of single nucleotide-gapped DNA (12, 14–16). Moreover, our recent fidelity studies suggest that human fPol{lambda} has similar single nucleotide gap-filling fidelity (10–4-10–5) as Polbeta (34). Taken together, these results strongly suggested that fPol{lambda}, like Polbeta, preferred short gapped DNA over non-gapped DNA and was likely a gap-filling polymerase involved in BER.


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  		TABLE 3
Pre-steady state kinetic parameters of matched dTTP incorporation

 
Interestingly, from D-1N to D-1OH and to D-DNA (D-1), the change in dTTP incorporation efficiency was due to a 100-fold variation in kp while the binding of an incoming nucleotide (Kd) was similarly tight (Table 2). In contrast, the increase of catalytic efficiency from non-gapped to gapped DNA with Polbeta was due to a considerable change in Kd rather than kp (12, 14–16). These suggested that fPol{lambda} and Polbeta achieve higher catalytic efficiency through different mechanisms and these two enzymes have evolved divergently. Similar Kd values of dTTP with three different DNA substrates (Table 3) indicated that the unprecedentedly high nucleotide binding affinity was due to the intimate interactions between an incoming nucleotide and the active site residues of fPol{lambda} as revealed by the ternary crystal structures of tPol{lambda} (33), rather than the presence of a downstream strand and its 5'-phosphate moiety. The significant decrease in kp from D-DNA (D-1) to D-1OH and to D-1N (Table 3) suggested the intense interactions between the downstream strand, including its 5'-phosphate and the dRPase domain, as demonstrated by the binary and ternary crystal structures of tPol{lambda} (29, 33), directed and anchored the productive binding of DNA and dNTP at the active site of fPol{lambda}. Consequently, the absence of these interactions will either improperly align the 3'-OH of the upstream primer 21-mer and the {alpha}-phosphate of the dNTP for in-line attack or affect the local protein conformational change, leading to slower catalysis. In the meantime, however, the fidelity of fPol{lambda} (Table 2) was slightly increased. This trend is consistent with what we have observed with different truncated fragments of Pol{lambda} (34) but disagrees with the general trend summarized from a survey of the A-, B-, X-, and Y-families by Beard et al. (54) that a more catalytically efficient DNA polymerase has a higher polymerization fidelity.

In conclusion, our pre-steady state kinetic data have demonstrated that the downstream strand and its 5'-phosphate moiety are critical to the polymerase efficiency of fPol{lambda}. For the first time, we have quantitatively evaluated the kinetic effect of a dRP mimic on the 5' terminus of a downstream strand. Because this dRP mimic only moderately affected the incorporation efficiency of both correct and incorrect nucleotides while having insignificant effect on the fidelity of human DNA polymerase {lambda}, gapped substrates like D-DNA in Fig. 2C, which have been predominantly used in the literature, are reasonable model substrates.


   FOOTNOTES
 
* This work was supported in part by startup funds from The Ohio State University (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

1 Supported by a Herta Camerer Gross graduate research fellowship and American Heart Association Predoctoral Fellowship Grant 0415129B. Back

2 To whom correspondence should be addressed: 740 Biological Sciences, 484 W. 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: BER, base excision repair; dRP, 2-deoxyribose-5-phosphate; dRPase, 5'-2-deoxyribose-5-phosphate lyase; dRP mimic, 1,2-dideoxyribose-5-phosphate; Pol, DNA polymerase; fPol{lambda}, full-length DNA polymerase {lambda}; tPol{lambda}, truncated DNA polymerase {lambda}. Back



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Lawley, P. D. (1966) Prog. Nucleic Acids Res. Mol. Biol. 5, 89–131[Medline] [Order article via Infotrieve]
  2. Lutz, W. K. (1990) Mutat. Res. 238, 287–295[Medline] [Order article via Infotrieve]
  3. Lindahl, T. (1993) Nature 362, 709–715[CrossRef][Medline] [Order article via Infotrieve]
  4. Ames, B. N., and Gold, L. S. (1991) Mutat. Res. 250, 3–16[Medline] [Order article via Infotrieve]
  5. Lindahl, T. (1979) Prog. Nucleic Acids Res. Mol. Biol. 22, 135–192[Medline] [Order article via Infotrieve]
  6. Lindahl, T., and Ljungquist, S. (1975) Basic Life Sci. 5A, 31–38[Medline] [Order article via Infotrieve]
  7. Lindahl, T., and Wood, R. D. (1999) Science 286, 1897–1905[Abstract/Free Full Text]
  8. Sobol, R. W., Horton, J. K., Kuhn, R., Gu, H., Singhal, R. K., Prasad, R., Rajewsky, K., and Wilson, S. H. (1996) Nature 379, 183–186[CrossRef][Medline] [Order article via Infotrieve]
  9. Singhal, R. K., Prasad, R., and Wilson, S. H. (1995) J. Biol. Chem. 270, 949–957[Abstract/Free Full Text]
  10. Dianov, G., and Lindahl, T. (1994) Curr. Biol. 4, 1069–1076[CrossRef][Medline] [Order article via Infotrieve]
  11. Liu, Y., Beard, W. A., Shock, D. D., Prasad, R., Hou, E. W., and Wilson, S. H. (2005) J. Biol. Chem. 280, 3665–3674[Abstract/Free Full Text]
  12. Klungland, A., and Lindahl, T. (1997) EMBO J. 16, 3341–3348[CrossRef][Medline] [Order article via Infotrieve]
  13. Zhou, J., Ahn, J., Wilson, S. H., and Prives, C. (2001) EMBO J. 20, 914–923[CrossRef][Medline] [Order article via Infotrieve]
  14. Burgers, P. M. (1998) Chromosoma 107, 218–227[CrossRef][Medline] [Order article via Infotrieve]
  15. Frosina, G., Fortini, P., Rossi, O., Carrozzino, F., Raspaglio, G., Cox, L. S., Lane, D. P., Abbondandolo, A., and Dogliotti, E. (1996) J. Biol. Chem. 271, 9573–9578[Abstract/Free Full Text]
  16. Wilson, S. H. (1998) Mutat. Res. 407, 203–215[Medline] [Order article via Infotrieve]
  17. Beard, W. A., and Wilson, S. H. (2000) Mutat. Res. 460, 231–244[Medline] [Order article via Infotrieve]
  18. Matsumoto, Y., and Kim, K. (1995) Science 269, 699–702[Abstract/Free Full Text]
  19. Kubota, Y., Nash, R. A., Klungland, A., Schar, P., Barnes, D. E., and Lindahl, T. (1996) EMBO J. 15, 6662–6670[Medline] [Order article via Infotrieve]
  20. Nicholl, I. D., Nealon, K., and Kenny, M. K. (1997) Biochemistry 36, 7557–7566[CrossRef][Medline] [Order article via Infotrieve]
  21. Srivastava, D. K., Berg, B. J., Prasad, R., Molina, J. T., Beard, W. A., Tomkinson, A. E., and Wilson, S. H. (1998) J. Biol. Chem. 273, 21203–21209[Abstract/Free Full Text]
  22. Aoufouchi, S., Flatter, E., Dahan, A., Faili, A., Bertocci, B., Storck, S., Delbos, F., Cocea, L., Gupta, N., Weill, J. C., and Reynaud, C. A. (2000) Nucleic Acids Res. 28, 3684–3693[Abstract/Free Full Text]
  23. Garcia-Diaz, M., Dominguez, O., Lopez-Fernandez, L. A., de Lera, L. T., Saniger, M. L., Ruiz, J. F., Parraga, M., Garcia-Ortiz, M. J., Kirchhoff, T., del Mazo, J., Bernad, A., and Blanco, L. (2000) J. Mol. Biol. 301, 851–867[CrossRef][Medline] [Order article via Infotrieve]
  24. Nagasawa, K., Kitamura, K., Yasui, A., Nimura, Y., Ikeda, K., Hirai, M., Matsukage, A., and Nakanishi, M. (2000) J. Biol. Chem. 275, 31233–31238[Abstract/Free Full Text]
  25. Garcia-Diaz, M., Bebenek, K., Sabariegos, R., Dominguez, O., Rodriguez, J., Kirchhoff, T., Garcia-Palomero, E., Picher, A. J., Juarez, R., Ruiz, J. F., Kunkel, T. A., and Blanco, L. (2002) J. Biol. Chem. 277, 13184–13191[Abstract/Free Full Text]
  26. Garcia-Diaz, M., Bebenek, K., Kunkel, T. A., and Blanco, L. (2001) J. Biol. Chem. 276, 34659–34663[Abstract/Free Full Text]
  27. Braithwaite, E. K., Prasad, R., Shock, D. D., Hou, E. W., Beard, W. A., and Wilson, S. H. (2005) J. Biol. Chem.
  28. Singhal, R. K., and Wilson, S. H. (1993) J. Biol. Chem. 268, 15906–15911[Abstract/Free Full Text]
  29. Garcia-Diaz, M., Bebenek, K., Krahn, J. M., Blanco, L., Kunkel, T. A., and Pedersen, L. C. (2004) Mol. Cell 13, 561–572[CrossRef][Medline] [Order article via Infotrieve]
  30. Vande Berg, B. J., Beard, W. A., and Wilson, S. H. (2001) J. Biol. Chem. 276, 3408–3416[Abstract/Free Full Text]
  31. Ahn, J., Kraynov, V. S., Zhong, X., Werneburg, B. G., and Tsai, M. D. (1998) Biochem. J. 331, Pt. 1, 79–87[Medline] [Order article via Infotrieve]
  32. Sawaya, M. R., Prasad, R., Wilson, S. H., Kraut, J., and Pelletier, H. (1997) Biochemistry 36, 11205–11215[CrossRef][Medline] [Order article via Infotrieve]
  33. Garcia-Diaz, M., Bebenek, K., Krahn, J. M., Kunkel, T. A., and Pedersen, L. C. (2005) Nat. Struct. Mol. Biol. 12, 97–98[CrossRef][Medline] [Order article via Infotrieve]
  34. Fiala, K. A., Duym, W. W., Zhang, J., and Suo, Z. (2006) J. Biol. Chem. 281, 19038–19044[Abstract/Free Full Text]
  35. Wang, K. Y., Parker, S. A., Goljer, I., and Bolton, P. H. (1997) Biochemistry 36, 11629–11639[CrossRef][Medline] [Order article via Infotrieve]
  36. Fiala, K. A., Abdel-Gawad, W., and Suo, Z. (2004) Biochemistry 43, 6751–6762[CrossRef][Medline] [Order article via Infotrieve]
  37. Johnson, K. A. (1992) Enzymes 20, 1–61
  38. Uchiyama, Y., Kimura, S., Yamamoto, T., Ishibashi, T., and Sakaguchi, K. (2004) Eur. J. Biochem. 271, 2799–2807[Medline] [Order article via Infotrieve]
  39. Hirose, F., Hotta, Y., Yamaguchi, M., and Matsukage, A. (1989) Exp. Cell Res. 181, 169–180[CrossRef][Medline] [Order article via Infotrieve]
  40. Chagovetz, A. M., Sweasy, J. B., and Preston, B. D. (1997) J. Biol. Chem. 272, 27501–27504[Abstract/Free Full Text]
  41. Liu, J., and Tsai, M. D. (2001) Biochemistry 40, 9014–9022[CrossRef][Medline] [Order article via Infotrieve]
  42. Li, S. X., Vaccaro, J. A., and Sweasy, J. B. (1999) Biochemistry 38, 4800–4808[CrossRef][Medline] [Order article via Infotrieve]
  43. Shah, A. M., Li, S. X., Anderson, K. S., and Sweasy, J. B. (2001) J. Biol. Chem. 276, 10824–10831[Abstract/Free Full Text]
  44. Beard, W. A., Shock, D. D., Yang, X. P., DeLauder, S. F., and Wilson, S. H. (2002) J. Biol. Chem. 277, 8235–8242[Abstract/Free Full Text]
  45. Matsumoto, Y., Kim, K., Katz, D. S., and Feng, J. A. (1998) Biochemistry 37, 6456–6464[CrossRef][Medline] [Order article via Infotrieve]
  46. Piersen, C. E., Prasad, R., Wilson, S. H., and Lloyd, R. S. (1996) J. Biol. Chem. 271, 17811–17815[Abstract/Free Full Text]
  47. Deterding, L. J., Prasad, R., Mullen, G. P., Wilson, S. H., and Tomer, K. B. (2000) J. Biol. Chem. 275, 10463–10471[Abstract/Free Full Text]
  48. DeRose, E. F., Kirby, T. W., Mueller, G. A., Bebenek, K., Garcia-Diaz, M., Blanco, L., Kunkel, T. A., and London, R. E. (2003) Biochemistry 42, 9564–9574[CrossRef][Medline] [Order article via Infotrieve]
  49. Garcia-Diaz, M., Bebenek, K., Krahn, J. M., Pedersen, L. C., and Kunkel, T. A. (2006) Cell 124, 331–342[CrossRef][Medline] [Order article via Infotrieve]
  50. Batra, V. K., Beard, W. A., Shock, D. D., Krahn, J. M., Pedersen, L. C., and Wilson, S. H. (2006) Structure 14, 757–766[Medline] [Order article via Infotrieve]
  51. Pelletier, H., Sawaya, M. R., Kumar, A., Wilson, S. H., and Kraut, J. (1994) Science 264, 1891–1903[Abstract/Free Full Text]
  52. Kraynov, V. S., Werneburg, B. G., Zhong, X., Lee, H., Ahn, J., and Tsai, M. D. (1997) Biochem. J. 323, Pt. 1, 103–111[Medline] [Order article via Infotrieve]
  53. Ahn, J., Werneburg, B. G., and Tsai, M. D. (1997) Biochemistry 36, 1100–1107[CrossRef][Medline] [Order article via Infotrieve]
  54. Beard, W. A., Shock, D. D., Vande Berg, B. J., and Wilson, S. H. (2002) J. Biol. Chem. 277, 47393–47398[Abstract/Free Full Text]

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