Steric and Electrostatic Effects at the C2 Atom Substituent Influence Replication and Miscoding of...

doi:10.1016/j.jmb.2009.07.019 J. Mol. Biol. (2009) 392, 251–269

Available online at www.sciencedirect.com

Steric and Electrostatic Effects at the C2 AtomSubstituent Influence Replication and Miscodingof the DNA Deamination Product Deoxyxanthosineand Analogs by DNA Polymerases

Huidong Zhang1,3, Urban Bren4, Ivan D. Kozekov2,3,Carmelo J. Rizzo2,3, Donald F. Stec2 and F. Peter Guengerich1,3⁎
1Department of BiochemistryVanderbilt University Schoolof Medicine, Nashville,TN 37232-0146, USA2Department of Chemistry,Vanderbilt University Schoolof Medicine, Nashville,TN 37232-0146, USA3Center in MolecularToxicology, VanderbiltUniversity School of Medicine,Nashville, TN 37232-0146,USA4National Institute ofChemistry, SI-1001 Ljubljana,Slovenia
Received 4 May 2009;received in revised form2 July 2009;accepted 7 July 2009Available online14 July 2009

*Corresponding author. DepartmentCenter in Molecular Toxicology, VanSchool of Medicine, Nashville, 638 RBuilding, 2200 Pierce Avenue, NashUSA. E-mail address: f.guengerich@Abbreviations used: 2-BrdI, 2-bro

CD, circular dichroism; CID, collisiodissociation; DBU, 1,8-diazabicyclo[Dpo4, S. solfataricus P2 DNA polymfluoro-2′-deoxyinosine; HMBC, hetebond coherence; HRMS, high-resoluspectrometry; LC, liquid chromatogspectrometry; MS/MS, tandem mas(DNA) polymerase; pol T7–, bacterioT7/exonuclease–.

0022-2836/$ - see front matter © 2009 E

Deoxyinosine (dI) and deoxyxanthosine (dX) are both formed in DNA atappreciable levels in vivo by deamination of deoxyadenosine (dA) anddeoxyguanosine (dG), respectively, and can miscode. Structure–activityrelationships for dA pairing have been examined extensively using analogsbut relatively few studies have probed the roles of the individual hydrogen-bonding atoms of dG in DNA replication. The replicative bacteriophageT7 DNA polymerase/exonuclease and the translesion DNA polymeraseSulfolobus solfataricus pol IV were used as models to discern themechanisms of miscoding by DNA polymerases. Removal of the 2-aminogroup from the template dG (i.e., dI) had little impact on the catalyticefficiency of either polymerase, as judged by either steady-state or pre-steady-state kinetic analysis, although the misincorporation frequency wasincreased by an order of magnitude. dX was highly miscoding with bothpolymerases, and incorporation of several bases was observed. Theaddition of an electronegative fluorine atom at the 2-position of dI loweredthe oligonucleotide Tm and strongly inhibited incorporation of dCTP. Theaddition of bromine or oxygen (dX) at C2 lowered the Tm further, stronglyinhibited both polymerases, and increased the frequency of misincorpora-tion. Linear activity models show the effects of oxygen (dX) and thehalogens at C2 on both DNA polymerases as mainly due to a combinationof both steric and electrostatic factors, producing a clash with the pairedcytosine O2 atom, as opposed to either bulk or perturbation of purine ringelectron density alone.

© 2009 Elsevier Ltd. All rights reserved.

Keywords: DNA polymerase; translesion DNA synthesis; DNA deamina-tion; DNA fidelity; steady-state enzyme kinetics
Edited by J. Karn
of Biochemistry andderbilt Universityobinson Researchville, TN 37232-0146,vanderbilt.edu.mo-2′-deoxyinosine;n-induced5.4.0]undec-7-ene;erase IV; 2-FdI, 2-ronuclear multipletion massraphy; MS, masss spectrometry; pol,phage polymerase

lsevier Ltd. All rights reserve

Introduction

The preservation and transmittal of accurategenetic information is critically dependent uponthe fidelity of DNA replication.1 Most DNA poly-merases (pol) are highly accurate, and N100 humangenes are known to be involved in DNA repair,emphasizing the importance of genetic integrity inbiological systems. Even with the presence of all ofthese DNA repair systems, a finite level of DNAmodification exists and challenges the DNA poly-merases.2 The issue of exactly what drives the highfidelity of DNA replication has been the subjectof considerable interest for the past 55 years.

d.

252 Guanine Substitutions and DNA Polymerases

The roles of Watson–Crick hydrogen bonds havebeen recognized since the elucidation of the struc-ture of DNA,3 but energetic considerations arguethat the specificity of dG:dC and dA:dT pairingis not sufficient to account for the selectivity.4Moreover, fidelity varies considerably among DNApolymerases. One view is that DNA polyme-rases have steric restraints and “search” for pairsthat have the overall molecular dimensions of dG:dC and dA:dT pairs.5,6 Consistent with this hypo-thesis are observations with isosteric pairs of bases,where polymerization activity is observed.2,7–14

With such base isosteres there is evidence thatstacking forces are involved, arguing that these andother forces may also be at work with the fourcanonical bases.2Oxidative deamination of the purine bases is

known to occur during inflammation and otherbiological processes, e.g. with nitric oxide.15–17 Thisprocess converts dA to dI and dG to dX, res-pectively (Fig. 1), and can change the coding pro-perties at individual bases in the genome.15,18–20Levels of ∼5 dX molecules (measured as the basehypoxanthine) in 107 nucleotides have beendetected in mouse tissues (a level intermediatebetween those found for the well studied 7,8-dihydro-8-oxo dG and the major malondialdehyde-dG adduct).21 Treatment of TK6 human lympho-blast cells with nitric oxide led to a 47-fold increasein DNA hypoxanthine (i.e., dX) and 15- to 18-foldincreases in hprt and tk6 mutations.16 Nitric oxide isproduced during inflammation, in host defense,18

and in thermal injury to tissues.22 The mutationsgenerated by treatment with nitric oxide were pre-dominantly dA:dT to dG:dC and dG:dC to dA:dTtransitions, 23 being consistent with what is knownabout in vitro miscoding properties of dI and dX,respectively, in some systems.19,20 The biologicalrelevance of dX also has some support in theexistence of a repair pathway system, Escherichiacoli endonuclease V.24

Yasui et al.20 recently reported that human pol α,pol η, and a shortened version of pol κ all insertedpredominantly dCTP opposite dI, and the corre-sponding dA:dT to dG:dC transition was seen ex-clusively in a site-specific mutagenesis study with ac-Ha-ras gene in a mammalian cell system.25 The

Fig. 1. Structures of purines. (a) Tautomers of 2-susbstitute

same group19 showed that these three human poly-merases incorporated dTTP opposite dX, but pol βpreferentially incorporated dCTP. Pol α paireddCTP and dTTP opposite dX in other studies.26,27

Wuenschell et al.28 reported that E. coli DNApolymerase I (Klenow fragment) incorporateddCTP opposite dX,29 but that HIV-1 reverse tran-scriptase inserted dTTP at a frequency one-quarterthat of dCTP. The molecular basis for the effects onthese polymerases is still unclear. Almost all of theliterature involving artificial DNA bases and iso-stere questions has been focused on derivatives ofthe purine dA, as opposed to dG.9–14 Literaturesearches for DNA polymerases plus dI or substi-tuted dI (or dX) found few examples, other thanthose already cited. Poly(dI) was a poor templatefor Moloney murine leukemia virus reverse trans-criptase.30 2-Bromodeoxyinosine triphosphate wasreported to be a substrate for E. coli pol I but fewdetails were provided.31

We utilized the replicative DNA polymerasebacteriophage polymerase T7/exonuclease– (polT7–) and translesion DNA polymerase IV fromSulfolobus solfataricus P2 (Dpo4) (A- and Y-familyDNA polymerases, respectively) with an oligo-nucleotide containing dI, two halogen substituents(2-fluoro-2′-deoxyinosine (2-FdI) and 2-bromo-2′-deoxyinosine (2-BrdI)), and a 2-oxo substituent (dX)(Figs. 1 and 2). Pol T7– is a kinetically well-behavedDNA polymerase for which crystal structures havebeen determined.33–35 The kinetics of its mechanismhave been studied extensively,36,37 including bypassof DNA adducts.38–43 Dpo444 is capable of bypass ofvarious DNA adducts and has been investigated inconsiderable detail in terms of both its function45–49

and structure.50–52 In addition to these C2 modifica-tions, we changed the C6 oxo group of dG to sulfur(6-SdG; the nucleoside is a drug and has beenstudied previously regarding interactions with someDNA polymerases53–56). Both steady-state and pre-steady-state kinetic approaches were used. Bothpolymerases utilized template dI well, while 2-FdI,2-BrdI, and dX attenuated the catalytic efficiency(kcat/Km) of both significantly. The parameterkcat/Km,57 which should be considered a parameteritself and not a derivative of separate terms,58 isrelated to the highest (rate-determining) activation

d purines. (b) Tautomerization and ionization of dX.

Fig. 2. Structures of dG, dA, dI,6SdG, 2-FdI, 2-BrdI, dX, and N2,N2-diMedG and their correspondingConnolly molecular surfaces inaqueous solution.32

253Guanine Substitutions and DNA Polymerases

free energy along the polymerization pathway(ΔG≠) by transition state theory:59,60

kcatKm

=kBTh

e�DGp

RT ð1Þ

where kB is Boltzmann's constant, h is Planck'sconstant, R is the universal gas constant, and T is thethermodynamic temperature (derivation of equa-tions are given in the Supplementary Data). Thisrate-determining barrier may generally correspondto the substrate binding/release and the relatedconformational change or to the chemical (phos-phodiester bond formation) step,59 which wehypothesize would be hindered significantly by acombination of both steric and electrostatic clashesbetween the C2 substituent of the template and theO2 atom of the incoming dCTP, as depicted in Fig. 3.This hindrance would be caused by a combinationof enthalpic (steric and electronic repulsion) as wellas entropic (non-optimal substrate orientation)factors. To address the validity of this hypothesis,a linear quantitative structure–activity relationshipbased on AMBER force field61 partial atomic chargeand van der Waals radius of the C2 substituent wasdeveloped. Consistent with other work, Dpo4 was

Fig. 3. Steric and electrostatic clash of the purine C2atom substituent and deoxycytidine O2 atom.

shown to be somewhat less sensitive than pol T7– tothe substitution effect. The results allow discern-ment of some elements of nucleotide selectivity byDNA polymerases and insight into why the ubiqui-tous lesion dX miscodes.

Results

Primer extension in the presence of all fourdNTPs

Pol T7– extended the primer (Table 1) to full length(Fig. 4a) in the order dG (not shown) NdI N2-FdIN6-SdGNdX≫2BrdI. The 2-BrdI extension reactionshowed mainly single-base incorporation. The pat-tern was very similar for Dpo4 (Fig. 4b).One concern was that 2-FI could partially react

with the amino group in Tris buffer,62,63 althoughthe incubation conditions (37 °C) were only forminutes, not hours. Analysis of the 2-FdI oligonu-cleotide after prolonged incubation in Tris did notproduce detectable amounts of a modified oligonu-cleotide (Supplementary Data Fig. S2), and subs-tituting Mops buffer (which is not reactive becauseit contains a hindered tertiary amino group asopposed to the primary amine in Tris) did not pro-

Table 1. Oligodeoxynucleotides used in kinetic studies

24-mer 5′-GCCTCGAGCCAGCCGCAGACGCAG25-mer 5′-GCCTCGAGCCAGCCGCAGACGCAG(T/C/G/A)36-mer 3′-CGGAGCTCGGTCGGCGTCTGCGTCXCTCCTGC-

GGCT

X: dG, dA, dC, dT, dI, dA, 2-FdI, 2-BrdI, dX, 6-SdG, or N2,N2-Me2dG.

Fig. 4. Extension of 32P-labeled primers. (a) Extension with all four dNTPs present opposite dI, 2-FdI, 6-SdG, dX, and2-BrdI by pol T7–. (b) Extension opposite dG, dI, 2-FdI, 6-SdG, dX, and 2-BrdI by Dpo4. Some of the lanes in b (dX and 2-BrdI) were run in a different set of experiments and are superimposed into the figure.


duce results different from those obtained using Trisin subsequent kinetic analyses (Supplementary DataFig. S3).

Analysis of full-length extension products

In order to establish the identities of the basesinserted in the extension reactions, the extendedprimers were subjected to liquid chromatography-mass spectrometry (LC-MS) using a method dev-eloped in this laboratory.64 pol T7– showed onlydCTP incorporation opposite dI, 2-FdI, and 6-SdG(Table 2). Dpo4 also inserted dCTP opposite dG andopposite dI, 2-FdI, and 6-SdG (Table 2). Consider-able incorporation of dATP was observed oppositedX with both polymerases, and small amounts ofdTTP and dGTP incorporation were detected in thecases of pol T7– and Dpo4, respectively (Table 2, seealso Supplementary Data Figs. S4 and S5, and TablesS1–S5). No frameshift product was found in anycase. The limited full-length extension with thetemplate containing 2-BrdI (Fig. 4) did not permitapplication of this technique.

Steady-state kinetics of single-baseincorporation

With both pol T7– and Dpo4 (Tables 3 and 4), thecatalytic efficiency (kcat/Km) for incorporation invol-ving the normal Watson–Crick base pairs weresimilar. The greater efficiency for dCTP insertionopposite dG obtained with Dpo4 was reproducible;however, an efficiency similar to the other canonicalbase pairs (∼0.03 μM−1 s−1) was observed in othersequence contexts (results not shown). Similar para-meters were obtained with dI:dCTP or dC:dITPpairing (Table 3), although the frequency of mis-insertion was an order of magnitude greater than fordG:dCTP pairing. Substitution at the C2 position ofdI was very disruptive to the function of bothpolymerases, with catalytic efficiency in the order2-FdINdXN2-BrdI, although the relative decreasefrom dI to 2-FdI was less in the case of Dpo4 (Table 4)than pol T7– (Table 3). 6-SdG produced only a smallattenuation in the catalytic efficiency of pol T7–

(∼75% decrease) but a larger difference was seenwith Dpo4 (Table 4), even considering the unusually

Table 2. LC-MS/MS analysis of primer extension products

5′ – GCCUCGAGUCAGCCGUAGACGUAG3′ – CGGAGCTCGGTCGGCGTCTGCGTCGCTCCTGCGGCTa % of total product

pol T7–b dI AGCGAGGACGCCGA 9AGCGAGGACGCCGAA 26AGCGAGGACGCCGAC 19AGCGAGGACGCCGAT 45

2-FdI AGCGAGGACGCCGAA 25AGCGAGGACGCCGAC 18AGCGAGGACGCCGAG 11AGCGAGGACGCCGAT 47

dX AGCGAGGACGC 19AGCGAGGACGCCG 4AGCGAGGACGCCGA 21AGCGAGGACGCCGAC 16

AGAGAGGACGC 13AGAGAGGACGCCG 12AGTGAGGACGCCGA 16

6-SdG AGCGAGGACGCCGA 24AGCGAGGACGCCGAA 20AGCGAGGACGCCGAC 19AGCGAGGACGCCGAT 37

Dpo4c dG AGCGAGGACGCCGA 17AGCGAGGACGCCGAC 24AGCGAGGACGCCGAA 59

dI AGCGAGGACGCCGA 44AGCGAGGACGCCGAC 17AGCGAGGACGCCGAA 39

2-FdI AGCGAGGACGCCGA 38AGCGAGGACGCCGAC 20AGCGAGGACGCCGAA 42

dX AGCGAGGACGCCGA 24AGCGAGGACGCCGAC 38AGAGAGGACGCCG 13AGAGAGGACGCCGA 22AGGGAGGACGCCGA 4

6-SdG AGCGAGGACGCCGA 40AGCGAGGACGCCGAC 20AGCGAGGACGCCGAA 40%

a G: dG, dI 2-FdI, dX, 6-SdG, as indicated.b Reaction conditions: 5 μM pol T7–, 100 μM thioredoxin, 10 μM indicated 24/36-mer oligonucleotide complex, 1 mM each dNTP,

12.5 mM MgCl2, 4 h at 25 °C. LC-MS: tR, 2.90–2.96 min; charge, –4; area, m/z value±0.2.c Reaction conditions: 5 μMDpo4, 10 μM indicated 24/36 -mer oligonucleotide complex, 1 mM each dNTP, 5 mMMgCl2, 4 h at 37 °C.

LC-MS: tR, 2.90–2.96 min; charge, –4; area, m/z value±0.2.


high value measured for dG:dCTP pairing in thissequence context with this enzyme (see alsoSupplementary Data Table S4). In the case ofincorporation opposite dX, pol T7– preferentiallyinserted dCTP but also inserted the other threecanonical dNTPs, at frequencies of 0.17–0.42(Table 3). Dpo4 inserted all four dNTPs, withdTTP and dATP being most preferred.The catalytic parameters indicated very limited

misincorporation for dI, 2-FdI, and 6-SdG, consis-tent with full-length extension results reported inTable 2. dX and 2-BrdI produced significant mis-coding with both polymerases. For both dX and 2-BrdI, the preferences for inserting dCTP versus dTTPwere reversed between pol T7– and Dpo4.Steady-state parameters were collected for “next-

base” extension past dG, dI, 2-FdI, and 6-SdG pairswith dCTP and dTTP (Table 1) with pol T7– andDpo4 (Supplementary Data Table S6). In all casesexamined, extension beyond a dT paired oppositethe modified base was highly unfavorable (seeTable 2). These frequencies, along with the single-dNTP misinsertion results (Table 3), can explain the

results observed in the LC-MS analysis of fullyextended products. With pol T7— and dX, bothinsertion (opposite dX) and extension were veryslow, and the preference for extension beyond a dX:dC pair explains the dominance of dC in the LC-MSresults (Table 2). The pattern with Dpo4 and dX wasless clear. Insertion opposite dX was slow for alldNTPs but somewhat more efficient with dTTP(Table 4). Extension past all dX:dNpairswas slowandthe extension past dX:dTwas preferred (Supplemen-taryData Table S6) but the LC-MS analysis showednoproduct with dTTP incorporation (Table 2). Theapparent incongruity cannot be readily explained byback-folding/slippage,64 in that the template wasdevoid of dA 5′ of the dX, andwe presume that dTTPcan be inserted opposite dX and extended slowly byone base but extension stops for other reasons.

Pre-steady-state kinetics of single-baseincorporation

With pol T7–, sharp kinetic bursts were observedfor all of the canonical incorporations (dG:dC and

Table 3. Steady-state kinetic parameters for one-base incorporation by pol T7

Template base dNTP kcat (s−1×10−3) Km (μM)

kcat/Km(μM−1 s−1)

Decrease(fold relativeto dG:dCTP)

Misinsertionratioa

dA T 73±3 0.17±0.04 0.43 — —C 18±4 3160±830 5.7×10−6 — 1.3×10−5

dT A 620±40 4.1±1.0 0.15 — —G 32±3 2400±340 1.3×10−5 — 8.7×10−5

I 113±5 890±100 1.3×10−4 — 8.7×10−4

dC G 230±10 1.1±0.4 0.21 1.9 —I 150±6 0.62±0.10 0.24 — 1.1A 32±2 3000±380 1.1×10−5 — 5.1×10−5

dG C 1200±100 3.1±0.6 0.39 — —T 18±1 800±160 2.3×10−5 — 5.9×10−5

dI C 150±4 0.69±0.10 0.22 1.8 —T 52±2 374±26 1.4×10−4 — 6.4×10−4

2-FdI C 44±4 33±6 1.3×10−3 300 —T 20±3 2040±500 9.8×10−6 — 7.5×10−3

2-BrdI C 3.3±1.2 7100±3300 4.7×10−7 830,000 —T 3.0±0.6 1800±600 1.7×10−6 — 3.5

dX C 13±1 200±20 6.5×10−5 6000 —G 21±3 770±250 2.7×10−5 — 0.42A 38±9 3500±1200 1.1×10−5 — 0.17T 24±3 1800±350 1.3×10−5 — 0.21

6-SdG C 84±2 1.5±0.2 5.6×10–2 7 —T 4.0±0.3 680±120 5.8×10−6 — 1.1×10−4

Conversion of primer to product was kept b 20% by adjusting the enzyme concentration and reaction time.a Misinsertion frequency=(kcat/Km)incorrect/(kcat/Km)correct.

Table 4. Steady-state kinetic parameters for one-base incorporation by Dpo4

Template base dNTP kcat s−1, ×10−3 Km μM kcat/Km μM−1 s−1

Decrease(fold relativeto dG:dCTP)

Misinsertionratioa

dA T 160±10 6.1±1.6 0.026 — —A 22±10 840±230 2.6×10−5 — 1.0×10−3

C 5.4±0.8 1000±300 5.4×10−6 — 2.1×10−4

G 4.6±0.3 770±110 6.0×10−6 — 2.3×10−4

dT A 360±5 11±0.8 0.033 — —C 15±1 940±120 1.6×10−5 — 4.8×10−4

G 42±2 580±60 7.2×10−5 — 2.2×10−3

T 31±1 750±80 4.1×10−5 — 1.3×10−3

dG C 930±60 2.5±0.8 0.37b — —A 5.7±0.2 490±60 1.2×10−5 — 3.1×10−5

G 6.3±0.1 190±20 3.3×10−5 — 9.0×10−5

T 34±2 780±140 4.4×10−5 — 1.2×10−4

dC G 540±20 17±2 0.032 — —A 11±2 1900±540 5.8×10−6 — 1.8×10−4

C 11±2 1800±430 6.1×10−6 — 1.9×10−4

T 17±2 1100±200 1.5×10−5 — 4.7×10−4

dI C 410±20 17±3 0.024 15 —A 12±1 200±30 6.0×10−5 — 2.5×10−3

G 1.6±0.1 200±30 8.0×10−6 — 3.3×10−4

T 14±3 820±260 1.7×10−5 — 7.1×10−4

2-FdI C 44±2 6.8±1.5 6.5×10−3 57 —T 4.7±0.4 200±60 2.4×10−5 — 3.7×10−3

2-BrdI C 19±2 2000±320 9.5×10−6 39,000 —T 4.6±0.8 990±270 4.6×10−6 — 0.49

dX C 11±1 730±200 1.5×10−5 25,000 —G 10±1 480±140 2.1×10−5 — 1.4A 20±1 720±70 2.8×10−5 — 1.9T 35±3 560±120 6.3×10−5 — 4.2

6-SdG C 160±10 22±4 0.0073 51 —A 2.0±0.2 490±100 4.1×10−6 — 5.6×10−4

G 1.3±0.1 250±40 5.2×10−6 — 7.1×10−4

T 1.6±0.1 390±90 4.1×10−6 — 5.6×10−4

Conversion of primer to product was kept b20% by adjusting the enzyme concentration and the reaction time.a Misinsertion frequency=(kcat/Km)incorrect/(kcat/Km)correct.b See footnote 3.


Fig. 5. Pre-steady-state kinetics of nucleotide incorporation of dNTP or (Sp)-dNTPαS by pol T7–. Pol T7– (70 nM) wasincubated with 120 nM 24-mer/36-mer primer:template complexes (32P-labeled primer) in a rapid quench-flowinstrument and mixed with 1 mM dNTP (■) (or (Sp)-dNTPαS (●)) plus 12.5 mM MgCl2 to initiate reactions.


dA:dT pairs), the dI:dC pairs (Fig. 5b and e), anddCTP:6-SdG (Fig. 5j). A weaker burst was observedfor the 2-FdI:dCTP pair (Fig. 5g) but none was seenwith dX (Fig. 5h) or 2-BrdI (Fig. 5i). Dpo4 extensionalso showed (Fig. 6) burst kinetics for all of thenormal incorporations and dI:dCTP (Fig. 6c) anddCTP:6-SdG (Fig. 6d). As with pol T7–, a weakerburst was seen with 2-FdI (Fig. 6f) but none with 2-BrdI or dX (Figs. 6g, 5h) (all with dCTP). In mostcases, the substitution of (Sp)-dCTPαS for dCTP (or(Sp)-dTTPαS for dTTP) did not change the kineticsappreciably, except in the case of dCTP incorpora-tion opposite 2-FdI (with both polymerases) (Figs.5g and 6f), suggesting that the phosphodiester bondformation step is not rate-limiting in most cases.The assays were repeated at various concentra-

tions of dCTP (Supplementary Data Figs. S5-S7) andthe kpol and Kd,dCTP values were determined (Tables5 and 6). Misincorporation was also analyzed, butkinetic bursts were observed only in the cases of pol

T7– with dI:dTTP and dT:dITP pairing (Supplemen-tary Data Figs. S8 and S9).

Spectroscopic measurements

Each of the template (36-mer) oligonucleotides(Table 1) was paired with an 11-mer overlapping themodified site and used for circular dichroism (CD)spectroscopy and Tm measurements. The CD spec-tra of oligonucleotides containing G or any of theother modified bases examined (I, 2-FI, 2-BrI) werevery similar (at 23 °C and pH 7.25) (See Supple-mentary Data Fig. S10).The Tm value was lowered by substitution of dI for

dG (Table 7; Supplementary Data Fig. S11), and add-ition of a 2-fluoro, -oxo, or -bromo group lowered theTm even more. The Tm value for 2-FdI was similar tothatmeasured forN2,N2-Me2G, a lesion that stronglydisrupts polymerization by pol T7–, Dpo4, andhuman Y-family DNA polymerases.65–68

Fig. 6. Pre-steady-state kinetics of incorporation of dNTP or (Sp)-dNTPαS by Dpo4. Dpo4 (70 nM) was incubated with 120 nM 24-mer/36-mer primer:template complexes(32P-labeled primer) in a rapid quench-flow instrument and mixed with 1 mM dNTP plus 5 mM MgCl2 to initiate reactions.

258Guanine

Substitutions

andDNAPolym

erases

Table 7. Tm values for 11-mer containing C hybridized toG or other residue

5′-CGCAGCGAGGA3′-CGGAGCTCGGTCGGCGTCTGCGTCXCTCCTGCGGCT

X Tm (°C)

dG 49.5±0.2dI 45.4±0.22-FdI 35.5±0.32-BrdI 25.3±1.1dX 32.0±0.46-SdG 44.7±0.4N2,N2-Me2dG 34.1±0.2

Table 5. Pre-steady-state kinetic parameters for one-baseincorporation by pol T7–

Templatebase dNTP kpol (s

−1)Kd,dCTP(μM)

kpol/Kd,dNTP(μM−1 s−1)

dG dCTP 28±1.8 31±6.6 0.90dA dTTP 260±40 160±40 1.6dI dCTP 230±10 70±9 3.3

dTTP 0.25±0.01 44±7 5.7×10−3

dC dGTP 130±8 44±7 3.0dITP 210±20 105±20 2.0

dT dATP 200±20 84±21 2.4dITP 0.57±0.04 64±13 8.9×10−3

6-SdG dCTP 45±4 90±20 0.52-FdI dCTP 1.3±0.1 130±30 0.010

See also Supplementary Data Fig. S17.


The pKa values of the free nucleosides weredetermined using UV titration experiments (Table8; Supplementary Data Fig. S12). The pKa valuesobtained for dG, dI, 2-FdI, and 2-BrdI were 9.2, 8.9,4.7, and 6.0, respectively, and reflect the keto–enolequilibrium of the C6 carbonyl, except in the case ofdX (the C6 NMR chemical shifts verify the keto asopposed to the enol form, Fig. 1, vide infra). There-fore, 2-FdI exists in the enol form at neutral pH, atleast in the free nucleoside. dX had a measured pKaof 6.1 (Table 8), consistent with the literature.69–73 Inthis case, the equilibrium is more complex due to theionization of the N3 proton as well as the N1 proton(Fig. 1b); previous work indicates that initialdissociation is for the N3 proton of dX rather thanN1.69,71–75 Therefore, at physiological pH (the pH ofthe DNA polymerization buffer is 7.5) dX existslargely as a stable oxy anion at C2 (Fig. 1b).28,70

NMR spectroscopy

The natural abundance 13C NMR spectra wereassigned using literature precedents76 and with theaid of heteronuclear multiple bond coherence(HMBC) data (Table 8). The 13C chemical shifts atthe C6 atoms (all obtained at pH 7.5) are consistentwith the pKa values (Table 8), in that the 2-FdI and 2-BrdI values are ∼167 ppm, apparently reflectingenol character, as opposed to the range of 159.6–161.7 ppm for dG, dI, and dX, which are in the keto(lactam) form. The 13C chemical shifts of the C2atom varied considerably, from 145.0 to 160.3 ppm,

Table 6. Pre-steady-state kinetic parameters for one-baseincorporation by Dpo4

Templatebase dNTP kpol (s

−1)Kd,dCTP(μM)

kpol/Kd,dCTP(μM−1 s−1)

dA dTTP 2.2±0.2 58±11 0.038dT dATP 3.0±0.1 64±7 0.047dG dCTP 3.0±0.1 10±1 0.30dI dCTP 2.8±0.2 36±6 0.0782-FdI dCTP 0.89±0.04 16±2 0.0566-SdG dCTP 2.4±0.1 33±5 0.073

See also Supplementary Data Fig. S18.

but did not show any clear trend with regard to theelectronegativity of the substituent atom. Neitherchemical shift (C2 or C6) showed a correlation withthe catalytic efficiency of either DNA polymerase(Tables 3, 4, and 8). Attempts to measure naturalabundance (N1) 15N NMR shifts were abandonedbecause of the low signal intensity.

Linear activity models

Experimental activation free energies (ΔGexp≠ ) of

pol T7– and Dpo4 were calculated on the basis of Eq.(1) and collected in Table 9 with regard to the C2substituent of the templating base. Merz-Kollmanpartial atomic charges of C2 substituents H, F, O,and Br calculated at the HF/6-31G(d) level of theoryare reported together with the corresponding vander Waals radii taken from the AMBER force fieldatom types HA, F, O2, and Br, respectively.61

A linear activity model was developed on thebasis of the steric and electronic clash of the C2substituent with the O2 atom of the incoming dCTPdepicted in Fig. 3. The model:

DGpmod = a� rð Þ + b� qð Þ + n

where a=9.86 kcal (mol Å)−1, b=–3.37 kcal (mole0)

−1, and n=–4.00 kcal mol−1, provided the bestagreement with the experiments for pol T7– and wasable to reproduce the experimentally observed orderof substituent effects on the pol T7– catalyticefficiency. As hypothesized, the positive coefficienta shows that the free energy barrier increases withthe bulkiness of the substituent due to a steric clash,while the negative coefficient b indicates that theactivation barrier increases with the negative charge

Table 8. pKa values and NMR chemical shift data forpurines

PurineC2

substituent pKa

13C Chemical shift

C2 (ppm) C6 (ppm)

dG NH2 9.2±0.1 154.8 159.9dI H 8.9±0.0 147.0 159.62-FdI F 4.7±0.1 160.3 166.62-BrdI Br 6.0±0.1 145.0 167.4dX OH 6.1±0.1 153.2 161.7

The NMR chemical shifts were determined in 0.3 M potassiumphosphate buffer (pH 7.5) in H2O/2H2O (95:5, v/v) at 25 °C. Thecarbon chemical shifts are referenced externally to TMS (0.0 ppm).

Table 9. Linear quantitative structure–activity relation-ship based on partial atomic charge and van der Waalsradius of the C2 substituent

Parameter H F O Br

ra (Å) 1.46 1.75 1.66 2.22qb (e0) 0.094 –0.227 –0.776 –0.009

A. pol T7–

ΔGexp≠ c (kcal mol−1) 10.3 13.4 15.2 18.1

ΔGmod≠ , kcal mol−1)ar+bq+nd 10.1 14.0 15.0 17.9ar+ne 11.4 14.0 13.2 18.4bq+nf 13.6 14.3 15.4 13.8

B. Dpo4ΔGexp

≠ (kcal mol−1)c 12.1 12.9 16.7 17.0ΔGmod

≠ (kcal mol−1)ar+bq+ng 11.5 14.5 16.1 16.5ar+nh 13.0 14.6 14.1 17.1bq+ni 13.6 14.6 16.4 13.9a van der Waals radii.b Merz-Kollman partial atomic charges.c Experimental activation free energies.d Linear activity model ΔGmod

≠ =ar+bq+n, where a=9.86 kcal(mol Å)−1, b=–3.37 kcal (mol e0)

−1, and n=-4.00 kcal mol−1.e Linear activity model ΔGmod

≠ =ar+n, where a=9.23 kcal (molÅ)−1 and n=–2.11 kcal mol−1.

f Linear activity model ΔGmod≠ =bq+n, where b=–2.10 kcal

(mol e0)−1 and n=13.8 kcal mol−1.

g Linear activity model ΔGmod≠ =ar+bq+n, where a=6.07 kcal

(mol Å)−1, b=–3.95 kcal (mol e0)−1, and n=3.00 kcal mol−1.

h Linear activity model ΔGmod≠ =ar+n, where a=5.34 kcal (mol

Å)−1 and n=5.21 kcal mol−1.i Linear activitymodelΔGmod

≠ =bq+n, where b=–3.17 kcal (mole0)

−1 and n=13.9 kcal mol−1.


of the substituent due to electrostatic repulsion bythe adjacent cytosine oxygen. The abbreviatedmodelbased only on the steric contribution:

DGpmod = a� rð Þ + n

where a=9.23 kcal (mol Å)−1 and n=–2.11 kcalmol−1, was not able to reproduce the experimentallyobserved order of substituent effects on the pol T7–

catalytic efficiency. The truncated model based onlyon the electrostatic contribution

DGpmod = b� qð Þ + n

where b=–2.10 kcal (mol e0)−1 and n=13.8 kcal

mol−1, was also not able to reproduce the expe-rimentally observed order of substituent effects onthe pol T7– catalytic efficiency.The linear activity model

DGpmod = a� rð Þ + b� qð Þ + n

where a=6.07 kcal (mol Å)−1, b=–3.95 kcal (mole0)

−1, and n=3.00 kcal mol−1, provided the bestagreement with the experiment for Dpo4 and wasable to reproduce the experimentally observed orderof substituent effects on Dpo4 catalytic efficiency. Asin the case of pol T7–, the positive coefficient a showsthat the free energy barrier increases with increasingsteric demands of the substituent, while the negativecoefficient b indicates that the same barrier increaseswith the negative charge of the substituent due to

electrostatic repulsion by the adjacent cytosineoxygen. The abbreviated model:

DGpmod = a� rð Þ + n

where a=5.34 kcal (mol Å)−1 and n=5.21 kcal mol−1

was not able to reproduce the experimentallyobserved order of substituent effects on the Dpo4catalytic efficiency. The truncated model:

DGpmod = b� qð Þ + n

where b=–3.17 kcal (mol e0)−1 and n=13.9 kcal

mol−1 was also not able to reproduce the experi-mentally observed order of substituent effects on theDpo4 catalytic efficiency.

Discussion

The focus of this investigation was to examine theeffects of two naturally-occurring DNA bases, dIand dX, on two model DNA polymerases, bacter-iophage pol T7– and S. solfataricus Dpo4. dI isderived from deamination of dA and acts as arelatively high-fidelity mimic of G despite missingone hydrogen bond. Rates of dNTP insertion werenot attenuated (relative to insertion of dCTP oppo-site dG or dTTP opposite dA).3 dX is a deaminationproduct of G, known to be present at relatively highlevels in mammalian cells,21,77 and stronglyimpeded polymerization and produced consider-able misinsertion with both polymerases (Tables 3and 4). The molecular basis of the effect of dX wasinvestigated further by substituting halogens (foroxygen) at the C2 position. These electronegativeatoms had effects similar to the oxyanion of dX inthat insertion of dCTP, but not dTTP, was attenu-ated. Little misincorporation was observed with 2-FdI with either DNA polymerase, as judged byeither steady-state kinetics (Tables 3 and 4) or LC-MS analysis of primer extension products (Table 2).None of the perturbations of the purine ring electrondensity (as manifested in the pKa determinationsand NMR chemical shifts, Table 8) explained thetrends in enzymatic activity, nor did either the stericbulk at the C2 position or the substituent atompartial charge alone (Table 9). The most satisfactoryexplanation for the effect was a combination of boththe size and the charge of the C2 substituent atom(Table 9); i.e. how much charge is “pushed” how faragainst the O2 atom of the pairing dCTP (Fig. 3). Thesame trend in the physicochemical parameters isalso reflected in the Tm changes (Table 7), whichindicates a similar role of steric-electronic clashbetween the C2 substituent of the template and theO2 atom of the C nucleotide in determining thethermal DNA duplex stability.Halogens were selected as mimics of the anionic

oxygen of dX because of their electronegativity, andthe resulting attenuation of polymerase function(Tables 3 and 4) has already been discussed.Previous studies indicated that the purine N3 protonof dX is largely ionized (i.e., the O2 enolate) at


neutral pH (pKa∼6) but the N1 proton is not (Fig.1b),69–73 and our own results are consistent with thisfinding (Table 8). Interestingly, the low pKa of 2-FdI(4.8) and the NMR spectra suggest that the N1 atomis in the ionized enol form (C6 oxyanion) at neutralpH (Table 8), yet activity opposite this base is betterthan with dX (Tables 3 and 4; Fig. 7a). 2-BrdI had ameasured pKa of 6.0 and the N1 proton should belargely dissociated at neutral pH; however, theselectivity for base insertion opposite 2-BrdI issimilar to dX (Tables 3 and 4). Overall, the pKavalues of the template bases and patterns ofionization are not well correlated to the order ofcatalytic efficiency (kcat/Km).Linear activity models (Table 9) demonstrated

that a combination of both van der Waals radii(steric clash) and Merz-Kollman partial atomiccharges (electrostatic repulsion) was able to repro-duce the experimentally observed order of sub-stituent effects on the pol T7– and Dpo4 catalyticefficiencies. These results implicate both steric andelectrostatic clashes between the C2 substituent ofthe template base and the O2 atom of the incomingdCTP, depicted in Fig. 3, as the predominant factorcontrolling the insertion efficiency for the DNApolymerases examined here. The free energy barrieris increased by the negative charge of the C2 sub-stituent in a similar fashion for both DNA poly-merases, as evidenced by nearly identical values ofthe linear coefficient b. This result can be rationa-lized by almost identical electrostatic repulsionbetween individual C2 substituent and the adjacentcytosine O2 atom obtained in both polymerases. Onthe other hand, pol T7– is significantly more inhi-bited by the increasing van der Waals radius thanDpo4, as evidenced by a larger linear coefficient a.Because the van der Waals radius represents a directmeasure of the bulkiness of the C2 substituent aswell as of its distance from the C2 atom, theobserved trend correlates well with the smallerenzyme active site of pol T7–.A caveat, however, has to be issued against

exceeding the limited scope of the described linear

Fig. 7. Comparison of effects of modifications at C2 on pol(■) and Dpo4 (❑); (b) Dpo4 pre-steady-state.

activity model. The model was developed solely forthe C2 substituent of the templating purine in thecase of the incoming dCTP. Moreover, C2 substitu-ents forming H-bonds with the O2 atom of theincoming dCTP should be handled with care due toincreased level of complexity. Furthermore, thevalidity of this linear activity model will bechallenged by investigation of additional C2 sub-stituents or DNA polymerases and is by definitionsubject to change. Finally, the exact physical mechan-ism of polymerase inhibition is out of the grasp ofthis linear activity model, because the initial stereo-electronic clash may result in impeded substratebinding (and related conformational change), non-optimal substrate orientation (entropic effect), and/or distortion of the preorganized electrostatics60 ofthe enzyme active site. To that end, one has to relyon more elaborate all-atom molecular dynamicssimulations of correct and incorrect dNTP bindingto pol T7¯78 as well as on QM/MM simulations ofalternative chemical mechanisms79 and the overallfidelity59 of pol T7¯. In particular, direct interactionof Lys522 (which was normally found to stabilizethe chemical transition state)79 with the templatingpurine might be responsible for the observedattenuation of pol T7¯.A dI:dC pair has only two hydrogen bonds but

the Tm was only slightly lower than that of a dG:dC pair (Table 7), and the steady-state (Tables 3and 4) and pre-steady-state catalytic efficiencies(Tables 5 and 6) were very similar to those seenwith dG:dC and dA:dT pairs. The only disadvan-tage of a dI versus dG template base was an orderof magnitude decrease in fidelity for both enzymes(Tables 3 and 4). In a similar vein, 6-SdG wasrather similar to dG in its Tm (Table 7) andpolymerization properties (Tables 3–6). We con-clude that the two Watson-Crick hydrogen bondsformed in dI:dC pairs (which are effective, at leastwith pol T7–, regardless of the “polarity” of whichbase is in the template and the dNTP) aresufficient for typical catalytic activity. The 2-amino group of dG does appear to provide some

T7– and Dpo4 kinetic parameters. (a) Steady-state, pol T7–


additional selectivity for dC in terms of the extrahydrogen bond (to the O2 atom of dC).The steady-state and pre-steady-state kinetic

analysis for the Dpo4 insertion reaction indicatedthat this polymerase was somewhat less sensitive tothe C2 substituent of the template purine than polT7– (Tables 3 and 4; and Fig. 7a). The differencescaused by the fluorine and bromine substitutions aremost dramatic (Fig. 7a and b). It is interesting to notethat both polymerases still showed burst kineticswith 2-FI (but not dX or 2-BrdI) (Figs. 4 and 5). Theexact basis of the mechanism cannot be known in theabsence of a crystal structure, but the larger size ofthe active site of Dpo443,50 may accommodate anenolized 2-FdI paired with dCTP. One possibility isthat dX (and the halide analogs) disrupts the minorgroove contacts between pol T7– and the nascentbase pair; these contacts are not present in Dpo4 andthus the attenuation is lower.Very recently Patro et al. reported a study of

purine C2 substituent atoms with human pol α,using steady-state kinetics.80 As in our own workwith pol T7– and Dpo4, dI:dC pairing was similar todG:dC pairing in terms of enzyme efficiency and theonly disadvantage of dI was an enhanced miscodingfrequency. (dX was not examined in that study.) It isof interest that the addition of a C2 amino group todATP or 6-chloropurine deoxyribonucleoside tri-phosphate increased the tendency to insert oppositea template dC (by pol α), possibly by increasing thenumber of hydrogen bonds from one to two.However, even in those cases where the formationof these two hydrogen bonds is possible, thecatalytic efficiency was much less than in the caseof dG:dC (and dI:dC), probably because the hydro-gen bonding schemes were not Watson-Crick due tothe lack of a proton on the N3 atom.80 The results ofanother recent study81 with Herpes Simplex Virus 1DNA polymerase are also similar.Our present findings have biological relevance in

light of the relatively high levels of dX and dIfound in vivo.21 These deaminated residues are notthe only base damage products resulting fromnitric oxide18,21,77 but the levels are of the sameorder of magnitude as the extensively studied 7,8-dihydro-8-oxo G and major malondialdehyde-dGadducts. The levels are considerably higher thanthose measured for the etheno adducts of dA anddG, although the latter showed a greater increasein level due to the production of nitric oxide.21

Several other potential DNA adducts were notfound at levels N1 per 107 bases in vivo in a mouseinflammation model,21 and are assumed to makeless contribution.In summary, studies with these model replicative

and translesion DNA polymerases use dI withhigh catalytic efficiency, as judged by both steady-state and pre-steady-state kinetic analyses. Theonly deficiency is a relatively small enhancementof the misincorporation frequency, but on the basisof dG (not dA). Thus, deamination of dA to formdI should yield an almost complete dA:dT to dG:dC transition mutation, as reported in one cellular

experiment.25 The conversion of dG to dX bydeamination yields a variable degree of dG:dC todA:dT transition mutations depending upon theparticular DNA polymerase, in line with theliterature.15,19,28 Our studies with electronegativeanalogs show that the strong attenuation ofbinding and of catalytic activity (dCTP incorpora-tion) can be explained by a combination of thesteric and electrostatic clash of the charged oxygenatom (of dX, or a halogen in 2-FdI or 2-BrdI) withthe O2 atom of dCTP (Fig. 3). The Tm differencesare also consistent with this explanation (Table 7),and both a replicative and a translesion DNApolymerase show rather similar patterns (Fig. 7;Table 9). Accordingly, we expect that the mechan-isms proposed here will apply to many DNApolymerases. Overall, these base structure–activityrelationships provide a mechanistic understandingof the miscoding properties of two major DNAadducts derived from deamination.

Materials and Methods

Materials

Pol T7– and E. coli thioredoxin were expressed in E.coli and purified as described.36,38,42 Dpo4 wasexpressed in E. coli and purified to electrophoretichomogeneity as described.64 Unlabeled dNTPs wereobtained from Amersham Biosciences (Piscataway, NJ),(Sp)-dCTPαS and (Sp)-dTTPαS were from Biolog LifeScience Institute (Bremen, Germany), and [γ-32P]ATP(specific activity 3×103 Ci mmol−1) was from PerkinEl-mer Life Sciences (Boston, MA). Phage T4 polynucleo-tide kinase and restriction endonucleases werepurchased from New England Biolabs (Beverly, MA).Bio-spin columns were obtained from Bio-Rad (Her-cules, CA). The (deoxy)nucleoside dX was purchasedfrom Sigma-Aldrich (St. Louis, MO), and (deoxynucleo-sides) dG and dI were purchased from Berry andAssociates (Dexter, MI).

Synthesis of 2-FdI

2-FdI was synthesized as the protected nucleoside,82

deprotected using 1,8-diazabicyclo[5.4.0]undec-7-ene(DBU),83 and purified by HPLC using a Beckman ODSoctadecylsilane (C18) semi-preparative column (5 μmparticle size, 10 mm diam.×250 mm). The followinggradient program was used with a flow rate of 1.5 mlmin−1 (all percentages are v/v): 0–15 min, linear gradientfrom 1% to 10%CH3CN inH2O; 15–20min, linear gradientfrom 10% to 20% CH3CN; 20–35 min, hold at 20% CH3CN;35-45 min, linear gradient from 20% to 100% CH3CN; 45–65 min, hold at 100% CH3CN; 65–70 min, linear gradientfrom 100% to 1% CH3CN; 70–75 min, hold at 1% CH3CN.The recovered product (84%) yieldwas characterized by itsMS and NMR spectra. MS: m/z calc for C10H11FN4O4 [M+H]+ 271.2, found 270.9; [M+Na]+ 293.2, found 293.1; calcfor purine ring fragment C5H3FN4O 155.1, found 155.1. 1HNMR (2H2O) δ 8.55 (s, 1H,N1-H), 8.19 (s, 1H,C8-H), 6.37 (s,1H, C-1′); 13C NMR (2H 2O) δ 158.85 and 160.25 (C2),149.65 (C4), 121.75 (C5), 168.24 (C6), 138.16 (C8). 19F NMR(2H 2O) δ -52.95 (2-F).


Synthesis of 2-BrdI and its phosphoramidite

2-Bromo-O6-(2-(4-nitrophenyl)ethyl)-2′-dI

3′,5′-O-[1,1,3,3-(Tetrakis-isopropyl)-1,3-disiloxanediyl]-6-O-[2-(4-nitrophenyl)ethyl]-2′-dG (170 mg, 0.26 mmol)84and SbBr5 (140 mg, 0.39 mmol) were dried by co-evaporation of the solvent from solutions in anhydroustoluene (3×3 ml). Anhydrous CH2Br2 was added to thesolution and cooled to –30 °C; tert-butylnitrite (108 μl,93 mg, 0.90 mmol) was then added slowly. The reactionwas stirred at –30 °C for 2 h and then poured into asolution of NaHCO3 and ice and extracted (3×) withCH2Cl2. The combined extracts were dried with Na2SO4,filtered, and concentrated in vacuo. The product was usedfor the next step without further purification.85

Tetrabutylammonium fluoride (0.5 ml, 1.0 M in tetra-hydrofuran) was added to a stirred solution of this fullyprotected 2-BrdI derivative (180 mg, 0.25 mmol) intetrahydrofuran (5 ml) The reaction was stirred for 2 h atroom temperature and the solvent was removed in vacuo.Purification on a Biotage SP1 medium-pressure chroma-tography apparatus using an Si-25M silica gel column (1-12% CH3OH gradient in CH2Cl2, v/v) afforded thedesired product (92 mg, 77% yield): 1H NMR (300 MHz,DMSO-d6): δ 8.58 (s, 1H, C8-H), 8.18 (d, 2H, J=8.6 Hz,aromatic), 7.64 (d, 2H, J=8.6 Hz, aromatic), 6.34 (t, 1H,J=6.6 Hz, H-1′), 5.35 (d, 1H, J=4.3 Hz, 3′-OH), 4.90 (t, 1H,J=5.4 Hz, 5′-OH), 4.82 (t, 2H, J=6.45 Hz, CH2), 4.40 (m,1H, H-3′), 3.86 (m, 1H, H-4′), 3.61-3.30 (m, 2H, H-5′, H-5″),3.30 (t, 2H, J=6.45 Hz, CH2), 2.66 (m, 1H, H-2′), 2.34 (m,1H, H-2″); high-resolution mass spectrometry (HRMS):calcd for C18H18 N5O6BrNa [M+H]+,m/z 502.0338; found,m/z 502.0338 (+0.0 ppm).

2-Bromo-5′-O-(4,4′-dimethoxytrityl)-O6-[2-(4-nitrophenyl)ethyl]-2′-dI

2-Bromo-O6-(2-(4-nitrophenyl)ethyl)-2′-dI (vide supra,92 mg, 0.19 mmol) was dried by evaporation of anhydrouspyridine (3×3 ml) and then dissolved in anhydrouspyridine (5 ml). 4,4′-Dimethoxytrityl chloride (98 mg,0.29 mmol) was added and the reaction was stirred for 5 hat room temperature under an Ar atmosphere. The solventwas removed in vacuo, and the residue was purified on aBiotage SP1 apparatus using an Si-25M silica gel column(0% to 6% CH3OH gradient in CH2Cl2, with 1% (C2H5)3N,v/v) to afford the desired product (110 mg, 74% yield): 1HNMR (300 MHz, DMSO-d6): δ 8.49 (s, 1H, C8-H), 8.17 (d,2H, J=8.6 Hz, aromatic), 7.63 (d, 2H, J=8.6 Hz, aromatic),7.28 (m, 2H, aromatic), 7.18 (m, 7H, aromatic), 6.77 (d, 2H,J=8.8 Hz, aromatic), 6.71 (d, 2H, J=8.8 Hz, aromatic), 6.39(t, 1H, J=6.1 Hz, H-1′), 5.39 (d, 1H, J1=4.7 Hz, 3′-OH), 4.80(m, 2H, CH2- CH2), 4.45 (m, 1H, H-3′), 3.99 (m, 1H, H-4′),3.70 and 3.68 (s, 6H, CH3O), 3.30 (m, 2H, CH2- CH2), 3.22-3.26 (m, 1H, H-5′), 3.08-3.12 (m, 1H, H-5″), 2.80 (m, 1H, H-2′), 2.37 (m, 1H, H-2″). HRMS: calc for C39H36N5O8BrNa[M+H] +, m/z 804.1645; found, m/z 804.1660 (+1.9 ppm).

2-Bromo-3′-O-[(N,N-diisopropylamino)-2-cyanoethoxy-phosphinyl]-5′-O-(4,4′-dimethoxytrityl)-O6-(2-(4-nitrophenyl)ethyl)-2′-dI

2-Bromo-5′-O-(4,4′-dimethoxytrityl)-O6-[2-(4-nitrophe-nyl)ethyl]-2′-dI (vide supra, 110mg, 0.14mmol)was dried byevaporation of anhydrous pyridine (3×3 mL) and thendissolved in freshly distilled CH2Cl2 (3 ml). A solution ofanhydrous 1H-tetrazole (460 μl of a 0.45 M solution inCH3CN, 14 mg, 0.20 mmol) and 2-cyanoethyl-N,N,N′,N′-

tetraisopropylphosphoramidite (67 μl, 64 mg, 0.21 mmol)were added and the reaction was stirred for 2 h at roomtemperature under an Ar atmosphere. The solvent wasremoved in vacuo, and the residue was purified on BiotageSP1 apparatus using an Si-25M silica gel column (10% to90% gradient of ethyl acetate in hexane, plus 1% (v/v)(C2H5)3N) to afford the phosphoramidite reagent (75 mg,54% yield): 1H NMR (300 MHz, DMSO-d6): δ 8.50 (m, 1H,C8-H), 8.19 (m, 2H, aromatic), 7.64 (m, 2H, aromatic), 7.29(m, 2H, aromatic), 7.16 (m, 7H, aromatic), 6.74 (m, 4H,aromatic), 6.43 (m, 1H,H-1′), 5.02 (m, 1H,H-3′), 4.78 (m, 2H,Ar-CH2-CH2), 4.28 (m, 1H, H-3′), 4.14 (m, 1H, POCH2), 4.02(m, 2H, H-4′, POCH2), 3.69 (s, 6H, CH3O), 3.62 (m, 2H,POCH2), 3.51 (m, 2H, CH(CH3)2), 3.30 (m, 4H,Ar-CH2- CH2,H-5′, H-5″), 2.87 (m, 2H, CH2CN), 2.66 (m, 2H, CH2CN),2.56 (m, 1H, H-2′), 2.27 (m, 1H,H-2″), 1.21 and 1.00 (m, 12H,CH(CH3)2).

31P NMR (DMSO-d6, 121 MHz): δ 147.9, 147.2.HRMS: calcd for C48H54N7O9PBr [M+H]+, m/z 982.2904;found, m/z 982.2919 (+1.9 ppm).

2-BrdI

DBU (11 μl, 17 mg, 0.11 mmol) was added to a solutionof 2-bromo-O6-(2-(4-nitrophenyl)ethyl)-2′-dI (52 mg,0.11 mmol) in 95:5 CH3CN/H2O (v/v) (3 ml) and thereaction was heated under reflux for 24 h. The solvent wasremoved in vacuo and the residue was purified with aBiotage SP1 apparatus using a Si-25M silica gel column(5% to 30% (v/v) CH3OH gradient in CH2Cl2) to afford theproduct (23 mg, 64% yield): 1H NMR (300 MHz, DMSO-d6): δ 7.92 (s, 1H, C8-H), 6.16 (t, 1H, J=6.1 Hz, H-1′), 5.27 (d,1H, J=3.6 Hz, 3′-OH), 5.14 (t, 1H, J=5.3 Hz, 5′-OH), 4.35(m, 1H, H-3′), 3.83 (m, 1H, H-4′), 3.60-3.48 (m, 2H, H-5′, H-5″), 2.65-2.56 (m, 1H, H-2′), 2.21-2.14 (m, 1H, H-2″). 13CNMR (0.3 M potassium phosphate in H2O/2H2O (95:5,v/v), pH 7.5): δ 144.26 (C2), 149.80 (C4), 123.24 (C5), 166.87(C6), 138.39 (C8).HRMS: calc forC10H11N4O4BrNa [M+H] +,m/z 352.9861; found, m/z 352.9865 (+1.1 ppm).

Oligonucleotide synthesis

Unmodified 24-mer, 25-mers, and 36-mer (Table 1) werepurchased from Midland Certified Reagent Co. (Midland,TX). Template 36-mers (HPLC-purified) containing dI, 2-FdI, dX, and 6-SdG were also purchased from Midland.The 36-mer oligonucleotide containing N2,N2-Me2G wassynthesized earlier.65 The template 36-mer with 2-BrdIwas synthesized using modified phosphoramiditereagents62,86 with a Perseptive Biosystems model 8909DNA synthesizer, on a 1 μmol scale. Standard syntheticprotocols using Glen Research reagents with p-(tert-butylphenoxy)acetyl-protected phosphoramidites wereemployed. The modified oligodeoxynucleotide wascleaved from the solid support and the exocyclic aminogroups were deprotected in a single step using 0.1 Maqueous NaOH. DBU was used to remove the O6-(4-nitrophenethyl) group. For a capillary gel electrophoreto-gram and the mass spectrum of the 2-BrdI oligonucleotide(Supplementary Data Fig. S1).

Reaction and analysis conditions for enzyme assays

Standard DNA polymerase reactions were carried outin 50 mM Tris–HCl buffer (pH 7.5) containing 50 mMNaCl, 5 mM DTT, 100 μg of bovine serum albumin ml−1,and 5% (v/v) glycerol. Pol T7– was used in reactions with12.5 mM MgCl2 at 25 °C and Dpo4 with 5 mM MgCl2 at


37 °C.42,64,83,87 In the case of pol T7–, the enzyme wascombined with 20-fold molar excess E. coli thioredoxinbefore incubations.36,38,83

Primers were 5′ end-labeled using T4 polynucleotidekinase/[γ-32P]ATP and annealed to different templates byheating a 1:1 solution of oligonucleotide to 95 °C for 5 minand then cooling slowly to room temperature. All reactionswere initiated by the addition of dNTP solutions contain-ing MgCl2 (at the indicated final concentration) topreincubated enzyme/DNA mixtures. After reaction,5 μl aliquots of the reaction mixture were quenched withEDTA/formamide (50 μl of 20 mM EDTA in 95% (v/v)formamide, 0.5% (w/v) bromphenol blue and 0.05% (w/v)xylene cyanol). Products were resolved with denaturinggel electrophoresis (20% (w/v) polyacrylamide gel) con-taining 8 M urea, visualized using a Bio-Rad MolecularImager FX instrument (Bio-Rad), and quantified byphosphorimaging analysis using the Molecular ImagerFX instrument with Quantity One software.

Primer extension assay with all four dNTPs

Each of the DNA substrates (100 nM) was incubatedwith pol T7– or Dpo4 (0, 0.5, 2, 10, or 50 nM) in Tris–HClbuffer and reactions were initiated by adding dNTP·Mg2+

solution (100 μM each dNTP and MgCl2, conditions asdescribed above) (Fig. 3). At the indicated time points, thereactions were quenched with EDTA/formamide andanalyzed by gel electrophoresis and phosphorimaging.

Steady-state kinetic analyses

A 32P-labeled primer, annealed to either an unmodifiedor adducted template, was extended in the presence ofvarious concentrations of a single dNTP. The molar ratioof primer/template complex to pol T7– or Dpo4 was≥10:1. Polymerase concentrations and reaction times wereadjusted to limit primer conversion to b20%.5 Reactions(20 μl) were done with 12 different concentrations ofdNTP, quenched by the addition of EDTA/formamide,and analyzed by gel electrophoresis, with subsequentquantification of the results. Plots of product formationversus dNTP concentration were fit using nonlinearregression (hyperbolic fits) in GraphPad Prism version3.0 (San Diego, CA) for the estimation of kcat and Km.

Pre-steady-state reactions

Rapid chemical quench experiments were performedusing a model RQF-3 KinTek Quench Flow Apparatus(KinTek Corp., Austin, TX) with 50 mM Tris–HCl buffer(pH 7.4) in the drive syringes. Reactions were initiated byrapid mixing of 32P-labeled primer/template/pol T7– orDpo4 mixtures (12.5 μl) with the dNTP·Mg2+ complex(10.9 μl) and then quenched with 0.6 M EDTA afterincubation for various lengths of time. The reactionproducts (20 μl) were mixed with 100 μl of EDTA/formamide, separated using denaturing gel electrophor-esis, and quantified. Pre-steady-state experiments withexcess DNA or with excess pol T7– or Dpo4 were fit withEqs. (2) and (3), respectively, using nonlinear regressionanalysis (GraphPad Prism version 3.0):

y =A 1� exp �kpt� ��

+ ksst ð2Þ

y =A 1� exp �kpt� �� ð3Þ

where y is the concentration of the product, A is the burstamplitude, kp is the pre-steady-state rate of nucleotideincorporation, t is time, and kss is a steady-state velocity ofnucleotide incorporation.36,88

kpol and Kd,dCTP were estimated by performing pre-steady-state reactions with different concentrations ofdNTP and reaction for various lengths of time. Graphs ofburst rates (kobs) versus the concentration of dNTP were fitto the hyperbolic equation:

kobs = kpol dNTP½ �= dNTP½ � + Kd;dCTP� � ð4Þ

where kpol is the maximal rate of nucleotide incorporationand Kd,dCTP is the equilibrium dissociation constant fordNTP.36,88

Phosphorothioate analysis

With the 32P-labeled primer annealed to a template,reactions were initiated by rapid mixing of 32P-labeledprimer/template/polymerase mixtures (12.5 μl) with (Sp)-dNTPαS·Mg2+ complex (or dNTP·Mg2+) (10.9 μl) and thenquenched with 0.6 M EDTA after reaction for variouslengths of time. Products were analyzed as describedabove for the pre-steady-state reactions.

LC-tandem mass spectrometry (MS/MS) analysis ofoligonucleotide products from DNA polymerasereactions64

Dpo4 (5 μM) or pol T7– (5 μM, plus 100 μM E. colithioredoxin) was incubated with uracil-containing primer/template DNA (10 μM, see Table 2), all four dNTPs (1 mMeach), and MgCl2 (5 mM for Dpo4, 12.5 mM for pol T7–) ina final volume of 100 μl at 37 °C for Dpo4 or 25 °C for polT7– for 4 h in 50 mM Tris–HCl buffer (pH 7.4) containing5 mM DTT. Reactions were terminated by extraction of theremaining dNTPs using size-exclusion chromatography(Bio-Spin 6 chromatography column, Bio-Rad). Concen-trated stocks of Tris–HCl, DTT, and EDTA were added toadjust the concentrations to 50 mM, 5 mM, and 1 mM,respectively. Next, E. coli uracil DNA glycosylase (20 units,Sigma-Aldrich) was added and the solution was incubatedat 37 °C for 6 h to hydrolyze the uracil residues on theextended primer. The reaction mixture was then heated at95 °C for 1 h in the presence of 0.25 M piperidine (to breakthe primer chain at the abasic sites), followed by removalof the solvent by centrifugation in vacuo. The dried samplewas resuspended in 100 μl of water for MS analysis.LC-MS/MS analysis was performed on a Waters

Acquity UPLC system (Waters, Milford, MA) connectedto a Finnigan LTQ mass spectrometer (Thermo FisherScientific, Waltham, MA), operating in the ESI negativeion mode. An Acquity UPLC BEH octadecylsilane (C18)column (1.7 mm, 1.0 mm×100 mm) was used with thefollowing LC conditions: Buffer A was 10 mM NH4CH3-CO2, 2% (v/v) CH3CN and buffer B was 10 mM NH4-CH3CO2, 95% (v/v) CH3CN. The following gradientprogram was used with a flow rate of 150 μl min−1(allpercentages are v/v): 0–2.5 min, linear gradient from100%A to 95%A/5% B; 2.5–6.0 min, linear gradient to 75%A/25% B; 6-6.5 min, linear gradient to 100% B; 6.5–8.0 min, hold at 100% B; 8.0–9.0 min, linear gradient to100% A; 9.0–12.0 min, hold at 100% A. The temperature ofthe column was maintained at 50 °C. Samples wereinjected with an autosampler system. The ESI conditionswere as follow: source voltage 4 kV, source current100 mA, auxiliary gas flow rate setting 20, sweep gasflow rate setting 5, sheath gas flow setting 34, capillary


voltage –49 V, capillary temperature 350 °C, tube lensvoltage –90 V. The MS/MS conditions were: normalizedcollision energy 35%, activation Q 0.250, activation time30 ms. The doubly (negatively) charged species weregenerally used for CID analysis. The calculations of theCID fragmentations of oligonucleotide sequences weredone using a program linked to the Mass SpectrometryGroup (Medicinal Chemistry) at the University of Utah†.

pKa determinations

Aliquots (10 μl) of stock aqueous solutions containingthe nucleosides dG, dI, 2-FdI, 2-BrdI, or dX (1.2–1.5 mM,based on ɛ254 of 10.2 mM−1 cm−1) were diluted with 400 μlof 50 mM potassium phosphate buffer adjusted to theindicated pH value (2.0–12.0). The UV spectra wererecorded at 23 °C using a modified OLIS-Hewlett Packard8452 diode array spectrophotometer (On-Line InstrumentSystems, Bogart, GA) from 240 nm to 340 nm, with thecorresponding buffer as reference. The absorbance differ-ence A254–A320 was plotted versus pH. Data points were fitwith Graphpad Prism (Graphpad, San Diego, CA) usingnonlinear regression, fitting to the equation:

Y = Ymin 10�X� �+ Ymax 10�pKa� ��

= 10�X + 10�pKa� �

with SE calculated using the software program.89,90

NMR spectroscopy

13C NMR experiments were acquired using a 14.0 TBruker magnet (Bruker, Billerica, MA) equipped with aBruker AV-III console, operating at 600.13 MHz, and a5 mm TCI cryogenically-cooled NMR probe. 1H and 19FNMR experiments were acquired using a 9.3 T Oxfordmagnet equipped with a Bruker AV console, operating at400.13MHz, and a 5mmBBFONMRprobe. Samples weredissolved in either H2O/2H2O (95/5, v/v) or, for the 13Cwork, 0.3 M potassium phosphate buffer (pH 7.5) inH2O/2H2O (95:5, v/v). 1H, 13C, and 1H-15N HSQC and1H-13C HMBC NMR spectra were acquired in 3.0-mmNMR tubes at 25 °C while spectra 19F NMR spectra wereacquired in standard 5 mmNMR tubes at 25 °C. Chemicalshifts for 19F and 13C were referenced externally tohexafluorobenzene (–164.9 ppm) and TMS (0.0 ppm)respectively. For one-dimensional 1H NMR, solventsuppression using the WATERGATE91,92 pulse sequencewas necessary to reduce the intensity of the H2Oresonance. Experimental conditions for 1H NMR spectraincluded 32 K data points, a 13 ppm sweep width, arecycle delay of 1.5 s, and 32–256 scans. For one-dimensional 19F NMR spectra, experimental conditionsincluded 65 K data points, 200 ppm sweep width, a recycledelay of 2.0 s, and 64 scans; for one-dimensional 13C NMR,experimental conditions included 32 K data points, a260 ppm sweep width, a recycle delay of 2.0 s, and 11 Kscans. 1H-13C HMBC experiments were acquired using a2048×64 data matrix, a J(C-H) value of 9 Hz for detection oflong-range couplings (resulting in an evolution delay of55 ms), a J1(C-H) filter delay of 145 Hz (34 ms) for thesuppression of one-bond couplings, a recycle delay of1.5 s, and 64 scans per increment. The HMBC data werealso processed using a π/2-shifted squared sine windowfunction and displayed in magnitude mode.

†www.medlib.med.utah.edu/massspec

Tm determination

UV melting experiments were carried with a VarianCary 4E spectrophotometer (Varian, Walnut Creek, CA).The 11-mer oligonucleotide:

5′-CGCAGCGAGGA-3′

and the 36-mer:

5′-TCGGCGTCCTCG⁎CTGCGTCTGCGGCTGGCTC-GAGGC-3′

form an 11-mer duplex containing G⁎ in the middle of theduplex, where G⁎ indicates dG, dI, 2-FdI, 2-BrdI, dX, 6-SdG, or N2,N2-Me2dG. The oligonucleotides (final duplexconcentration ∼1.7 μM) were dissolved in 1.0 ml of10 mM sodium phosphate buffer (pH 7.0), 100 mM NaCl,1 mM EDTA in a 1.0 cm cuvette. The UVabsorbance (A2600.7–1.0) was monitored at 1 min intervals with a 1 °Cmin−1 temperature gradient. The temperature was cycledbetween 15 °C and 80 °C. The error was estimated fromthe fit of the determined curve (increasing temperaturedata) to the sigmodal equation:

Y =A2 + A1 � A2ð Þ= 1 + exp X � Tmð Þ=A3½ �ð Þ

where A1, A2, and A3 are three independent parameters.The Tm values were matched with the estimates from thefirst derivatives (dA/dT) of the melting curves.

CD Analysis of duplexes93-95

CD spectra were recorded in 1 mm cuvettes with aJASCO J-810 instrument (JASCO, Easton, MD) in theVanderbilt facility. A solution of the duplex of 11-mer/36-mer (6 μM) in 10 mM potassium phosphate buffer (pH7.25), 100 mM NaCl was placed in the cell and spectrawere recorded from 350–200 nm at 20 °C. The 11-meroligonucleotide:

5′-CGCAGCGAGGA-3′

and 36-mer oligonucleotide:

5′-TCGGCGTCCTCG⁎CTGCGTCTGCGGCTGGCTCG-AGGC-3′

were annealed to form a duplex (with G⁎ indicating dG,dI, 2-FdI, or 2-BrdI). The absorbance in all CD studies wasb0.20. CD output (in mdeg) was converted to standardmean residue ellipticity, [θ] (in mdeg μM−1 mm−1), usingthe relationship:

u½ � = 100� signal=cl ð5Þwhere signal is the CD output in mdeg, c is the DNAconcentration (in μM, based on total oligonucleotideduplex), and l is the cell pathlength (in mm).

Ab initio calculations

In order to reduce the computational costs, the 2′-deoxyribose moiety of dI, 2-FdI, 2-BrdI, and dX wasreplaced by a methyl group. The structures underconsideration were subjected to full geometry optimiza-tion and subsequent vibrational analysis in the harmonicapproximation at the Hartree-Fock (HF) level of theory


using 6-31G(d) basis set encoded in the Gaussian 03program.96 The absence of imaginary vibrational frequen-cies indicated correctly performed minimizations.97 Merz-Kollman partial atomic charges were finally calculatedfrom the electrostatic potential surrounding the optimizedstructures.

Acknowledgements

We thank M. J. Stone for the use of the spectro-photometer for the Tm experiments, R. L. Eoff forhelpful discussions and comments on the manu-script, and K. Trisler for assistance in preparation ofthe manuscript. This research was supported, inpart, byNational Institutes of Health grants NIHR01ES010375 (to F.P.G.), P01 ES00535 (to I.D.K., C.J.R.),andP30 ES000267 (to F.P.G., C.J.R.) and the SlovenianResearch Agency Grants (P1-0002, Z1-2000) and aWorld Federation of Scientists Scholarship (to U.B.).NMR measurements were made using a 600 MHzinstrument purchased with partial support fromNIH grant S10 RR019022 (D.F.S.).

Supplementary Data

Supplementary data associated with this articlecan be found, in the online version, at doi:10.1016/j.jmb.2009.07.019

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