insight into mechanisms of 3 -5 exonuclease activity and ...providing insight into the mechanism for...

Insight into mechanisms of 3′-5′ exonuclease activityand removal of bulky 8,5′-cyclopurine adducts byapurinic/apyrimidinic endonucleasesAbdelghani Mazouzia, Armelle Vigourouxb, Bulat Aikesheva,c, Philip J. Brooksd,e, Murat K. Saparbaeva,Solange Morerab,1, and Alexander A. Ishchenkoa,1

aUniversité Paris-Sud, Laboratoire “Stabilité Génétique et Oncogenèse”, Centre National de la Recherche Scientifique (CNRS), Unité Mixte de Recherche 8200,Institut de Cancérologie Gustave-Roussy, F-94805 Villejuif Cedex, France; bLaboratoire d’Enzymologie et Biochimie Structurales, CNRS, F-91198 Gif-sur-YvetteCedex, France; cL.N. Gumilev Eurasian National University, Astana, Republic of Kazakhstan, 010008; and dLaboratory of Neurogenetics and Division ofMetabolism and Health Effects, National Institute on Alcohol Abuse and Alcoholism, eOffice of Rare Diseases Research, National Center for AdvancingTranslational Sciences, National Institutes of Health, Bethesda, MD 20892

Edited by Philip C. Hanawalt, Stanford University, Stanford, CA, and approved June 28, 2013 (received for review March 19, 2013)

8,5′-cyclo-2’-deoxyadenosine (cdA) and 8,5′-cyclo-2’-deoxyguano-sine generated in DNA by both endogenous oxidative stress andionizing radiation are helix-distorting lesions and strong blocks forDNA replication and transcription. In duplex DNA, these lesions arerepaired in the nucleotide excision repair (NER) pathway. How-ever, lesions at DNA strand breaks are most likely poor substratesfor NER. Here we report that the apurinic/apyrimidinic (AP) endo-nucleases—Escherichia coli Xth and human APE1—can remove 5′ScdA (S-cdA) at 3′ termini of duplex DNA. In contrast, E. coli Nfo andyeast Apn1 are unable to carry out this reaction. None of theseenzymes can remove S-cdA adduct located at 1 or more nt awayfrom the 3′ end. To understand the structural basis of 3′ repairactivity, we determined a high-resolution crystal structure of E. coliNfo-H69Amutant bound to a duplex DNA containing an α-anomeric2′-deoxyadenosine:T base pair. Surprisingly, the structure reveals abound nucleotide incision repair (NIR) product with an abortive 3′-terminal dC close to the scissile position in the enzyme active site,providing insight into the mechanism for Nfo-catalyzed 3′→5′ exo-nuclease function and its inhibition by 3′-terminal S-cdA residue.This structure was used as a template to model 3′-terminal residuesin the APE1 active site and to explain biochemical data on APE1-catalyzed 3′ repair activities. We propose that Xth and APE1 mayact as a complementary repair pathway to NER to remove S-cdAadducts from 3′ DNA termini in E. coli and human cells, respectively.

oxidative DNA damage | endonuclease IV | DNA glycosylase |base excision repair | damage specific endonuclease

Oxidative damage to DNA caused by reactive oxygen speciesis believed to be a major type of endogenous cellular damage.If unrepaired, the damage will tend to accumulate and lead topremature aging, neurodegenerative disorders, and cancer (1).More than 80 different oxidative modifications of DNA basesand sugar backbone have been identified to date (2). Diaste-reoisomeric (5′S)- and (5′R)-8,5′-cyclo-2′-deoxyadenosine (cdA)and 8,5′-cyclo-2′-deoxyguanosine (cdG) are generated by endog-enous oxidative stress and ionizing radiation among other oxidizedbases (Fig. 1A). 8,5′-cyclo-2′-deoxypurines (cdPu) are generated byhydroxyl radical attack at C5′ sugar by H-abstraction resulting information of C5′-centered sugar radical, which then reacts in theabsence of oxygen with the C8 of the purine. Subsequent oxidationof the resulting N7-centered radical leads to intramolecular cy-clization with the formation of a covalent bond between the C5′-and C8-positions of the purine nucleoside. When present in DNAduplex cdA causes large changes in backbone torsion angles,which leads to weakening of base pair hydrogen bonds andstrong perturbations of the helix conformation near the lesionfor both diastereoisomers. Interestingly, the glycosidic bond inS-cdA is approximately 40-fold more resistant to acid hydrolysis

compared with regular dA, implying that this base lesion wouldbe resistant to DNA glycosylase action (3).The cdA adducts in DNA are a strong block to various DNA

polymerases, such as T7, δ, and η (4). Interestingly, translesionDNA polymerase η can perform lesion bypass synthesis on theR-cdA but not on S-cdA (5). Both diastereomers of cdA alsoinhibit DNA transcription by blocking primer extension by T7DNA polymerase, and S-cdA inhibits binding of the TATA boxprotein in vitro and strongly reduces gene expression in vivo (6).In addition, in vivo human RNA polymerase II generates mutatedRNA transcripts when using DNA template containing S-cdA (7).Given the strong genotoxic effect of cdA adducts on DNA me-tabolism, cells should have a repair mechanism to remove thesehelix-distorting DNA adducts. Indeed, it was shown that the nu-cleotide excision repair (NER) pathway can remove cdA adductswith efficiency comparable to that of T = T cyclobutane dimersand exhibits higher activity in excising the R-isomer (4, 8). Inagreement with the biochemical data, it was shown that cdPuadducts accumulate in keratinocytes from xeroderma pigmento-sum group C and Cockayne syndrome (CS) group A patients ex-posed to X-rays and potassium bromate (KBrO3) (9, 10) and alsoin organs of CS group B knockout mice (11). Importantly, cdA

Significance

Oxidative DNA damage has been postulated to play an im-portant role in human neurodegenerative disorders and cancer.8,5′-cyclo-2′-deoxyadenosine (cdA) is generated in DNA byhydroxyl radical attack and strongly blocks DNA replicationand transcription. Here we demonstrate that cdA adducts at3′ termini of DNA can be removed by 3′-5′ exonuclease activityof the apurinic/apyrimidinic (AP) endonucleases: Escherichiacoli Xth and human APE1. The crystal structure of bacterial APendonuclease in complex with DNA duplex provides insightinto the mechanism of this activity. This new repair functionprovides an alternative pathway to counteract genotoxic effectof helix-distorting DNA lesions.

Author contributions: A.M., A.V., M.K.S., S.M., and A.A.I. designed research; A.M., A.V.,B.A., S.M., and A.A.I. performed research; P.J.B. contributed new reagents/analytic tools;A.M., B.A., P.J.B., M.K.S., S.M., and A.A.I. analyzed data; and P.J.B., M.K.S., S.M., and A.A.I.wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

Freely available online through the PNAS open access option.

Data deposition: The atomic coordinates and structure factors of H69A Endo IV:DNA havebeen deposited in the Protein Data Bank, www.pdb.org (PDB ID code 4K1G).1To whom correspondence may be addressed. E-mail: [email protected] [email protected].

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1305281110/-/DCSupplemental.

www.pnas.org/cgi/doi/10.1073/pnas.1305281110 PNAS | Published online July 29, 2013 | E3071–E3080

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and cdG lesions accumulate spontaneously in nuclear DNA ofWT mice with age, suggesting that DNA repair is unable to keepthe steady-state level of these complex DNA lesions over thelifespan of an organism (12). Interestingly, S-cdA diastereoisomersare removed in the NER pathway much less efficiently than thecorresponding 5′R-cdA ones and are also present at a higherlevel in nontreated mice organs (4, 12).At present, NER is the only known DNA repair pathway to

remove cyclopurine adducts in duplex DNA. However, removalof lesions located in close proximity to DNA strand breaks byNER has not been reported except in one study by Plunkett andcoworkers (13). These authors demonstrated that a 3′-terminalcytosine analog present at a single-strand break is not a substratefor the human global genome NER pathway, but transcription-

coupled NER may participate in cleansing single-strand breaksfrom this 3′ adduct (13). Therefore, it is unlikely that NER couldbe able to efficiently remove cdPu located at 3′ termini of a sin-gle-strand break. Recently it has been demonstrated that 5′R and5′S isomers of cdATP could be incorporated with low efficiencyby replicative DNA polymerases and then inhibit further DNAsynthesis, thus potentially generating gapped DNA with a cdAadduct at the 3′ end (14). Furthermore, in the absence of ion-izing radiation and/or drugs, cdPu could arise at 3′ termini asa result of 3′→5′ exonuclease degradation of DNA, for exampleby TREX1 (5).The majority of oxidatively damaged DNA bases are sub-

strates for two overlapping pathways: DNA glycosylase-initiatedbase excision repair (BER) and apurinic/apyrimidinic (AP) en-donuclease-mediated nucleotide incision repair (NIR) (15). Inthe NIR pathway, an AP endonuclease makes an incision 5′ to adamaged nucleotide and then extends the resulting single-strandbreak to a gap by a nonspecific 3′→5′ exonuclease activity (16,17). AP endonucleases are multifunctional DNA repair enzymesthat possess AP site nicking, 3′ repair diesterase, NIR, and 3′→5′exonuclease activities and are divided into two distinct familiesbased on amino acid sequence identity to either Escherichia coliexonuclease III (Xth) or endonuclease IV (Nfo) (18). HumanAPE1 is homologous to Xth, whereas Saccharomyces cerevisiaeApn1 is homologous to Nfo. Previously it was shown that APendonuclease-catalyzed 3′→5′ exonuclease activity could serve asa 3′ editing function for removing mismatched and oxidizedbases at 3′ termini of DNA duplex (19–21). However, the de-tailed mechanisms for those 3′ editing repair activities are not yetclearly understood. Although Xth and Nfo AP endonucleasefamilies share common DNA substrate specificities, they aredistinguished by their modes of DNA damage recognition. In-deed, cocrystal structures of Nfo bound to an AP site analog,tetrahydrofuran (THF), showed that the enzyme drastically dis-torts the DNA helix by ∼90° bending and flips out not only thetarget AP site but also its opposing nucleotide out of the DNAbase stack (22, 23). Interestingly, the Nfo active site pocketsterically excludes binding of normal β-configuration nucleotides,but it can fit α-anomeric nucleotides. In contrast, cocrystal struc-tures of APE1 bound to abasic site-containing DNA show that theenzyme kinks the DNA helix by only 35° and binds a flipped-outAP site in a pocket that excludes DNA bases, whereas the op-posite base remains stacked in the duplex (24). Importantly, theDNA substrate specificity of APE1 but not that of Xth variesdepending on reaction condition (25).Here we investigate whether S-cdA adducts in DNA can be

removed in alternative to NER pathways by AP endonucleases.Our results show that in contrast to Nfo and Apn1, Xth andAPE1 remove S-cdA adducts when present at 3′ termini ofa recessed, nicked, or a gapped DNA duplex. However, S-cdAadducts located at 1 or more nt away from the 3′ end are notsubstrates for AP endonucleases. Using the high-resolution crystalstructure of Nfo-H69A:DNA complex, which provides a pictureof the position of the DNA for exonuclease activity, we were ableto model the DNA exonuclease conformation into APE1’s activesite and provide insight into the mechanism for repair of S-cdAadducts. The structural basis and potential biological importanceof the reported substrate specificity of Xth and APE1 incleansing genomic DNA of highly cytotoxic lesions are discussed.

ResultsXth/APE1 but Not Nfo/Apn1 Remove Bulky S-cdA Adducts at 3′ Terminiof Recessed, Nicked, and Gapped Duplex DNA. First we examinedwhether an S-cdA nucleotide in duplex DNA is a substrate forNIR-AP endonucleases E. coli Nfo, yeast Apn1, human APE1,or the human endonuclease VIII-like 1 (NEIL1) DNA glyco-sylase. For this, we incubated a 3′-[32P]-labeled 42-mer duplexoligonucleotide referred to as cdA42•T42 with an excess of en-

Fig. 1. Repair of 3′-blocking bulky adducts by AP endonucleases-catalyzed3′→5′ exonuclease activity. (A) Chemical structures of 5′S- (S-cdA) and 5′R-(R-cdA) diastereoisomers of cdA. (B) Denaturing PAGE analysis of repairproducts generated by the AP endonucleases when acting upon 3′-terminalS-cdA nucleotide. 5′-[32P]-labeled A•Trec, cdA•Trec, and cdAA•Trec duplexoligonucleotides were incubated with various AP endonucleases of differentorigins (5 nM of APE1, Nfo, and Apn1 or 10 pM of Xth) for 10 min at 37 °C.(C) APE1-catalyzed 3′ repair activities toward recessed, gapped, and nickedduplex DNA. 5′-[32P]-labeled oligonucleotide duplexes were incubated in thepresence of 5 nM Ape1 for 10 min at 37 °C. The “X” denotes the position ofS-cdA nucleotide. Details in Materials and Methods.

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zyme and then analyzed the reaction products by denaturingPAGE. No cleavage of cdA42•T42 by any of these enzymes wasobserved, indicating that bulky S-cdA adduct is not a substratefor either NIR or BER pathways (Fig. S1).It was previously shown that S-cdA nucleotide in duplex DNA

is a strong block to 3′→5′ exonuclease activities of T4 DNApolymerase, mammalian 3′ repair exonuclease 1 (TREX1), E. coliXth, snake venom phosphodiesterase, and nuclease P1 (5). Be-cause AP endonucleases contain robust 3′→5′ exonuclease ac-tivity that removes regular and modified nucleotides in duplexDNA (20), we examined whether APE1, Nfo, Apn1, and Xthcould remove S-cdA monophosphate (S-cdAMP) when placed inclose proximity to 3′ termini of the recessed duplex oligonucle-otide cdA•Trec and cdAA•Trec (Table 1). As shown in Fig. 1, allAP endonucleases tested efficiently degrade regular dA•Trecduplex oligonucleotide containing normal dA nucleotide at the3′ termini of a gap (Fig. 1B, lanes 2, 7, 10, and 14). Intriguingly,APE1 and Xth, but not Nfo and Apn1, can remove 3′-terminalS-cdA nucleotide in cdA•Trec with high efficiency (lanes 4 and15 vs. 8 and 11) and then continue to degrade DNA further ina processive manner by their nonspecific 3′→5′ exonuclease ac-tivity (Fig. 1B, lanes 2, 7, 10, and 14). Interestingly, the recessedduplex cdAA•Trec with the S-cdA adduct placed at the secondposition from the 3′ terminus completely blocks the 3′→5′exonuclease activity of APE1, Nfo, and Xth (lanes 6, 9, and 16).However, Apn1 was able to remove 3′-terminal dAMP incdAA•Trec duplex with very low efficiency but then was com-pletely blocked by the remaining S-cdA adduct (lane 12).To precisely determine the distance from the 3′ end to S-cdA

nucleotide at which the adduct starts to inhibit 3′→5′ exo-nuclease activity of AP endonucleases, we constructed recessedDNA duplex cdA42•T55 containing S-cdA nucleotide on the re-cessed strand 14 nt away from the 3′ end. As expected, the pres-ence of S-cdA adduct strongly blocks the 3′→5′ exonucleaseactivities of all AP endonucleases tested (Fig. S2). APE1 and Xthexonuclease activities stop 2 nt before the S-cdA adduct (in thecontext 5′-DNA-cdA-A-T-3′), whereas Nfo exonuclease slowsdown 3 nt and completely blocked 1 nt before the lesion, andApn1 is blocked at the lesion site (Fig. S2). Taken together, theseresults suggest that a bulky S-cdA adduct placed in the middle ofa DNA duplex cannot be removed by AP endonuclease-catalyzedNIR or by exonuclease activities. Nevertheless, human APE1 and

E. coli Xth can efficiently eliminate S-cdA adduct at the 3′ ter-mini of a recessed DNA duplex.Next we examined APE1 activities on the recessed, nicked, and

gapped DNA duplexes containing either regular dA or S-cdAnucleotide in close proximity to the 3′ end. As shown in Fig. 1C,whereas APE1 exhibits highly processive 3′→5′ exonucleaseactivity on nondamaged recessed A•Trec duplex (lane 2), it isstrongly inhibited on the gapped A•Tgap duplex and almostblocked on the nicked A•Tnick duplex (lanes 3 and 4). Theseresults indicate that APE1 requires an extended single-strandedDNA region for its processive exonuclease function. It should benoted that previous studies showed either similar efficiency ofAPE1-catalyzed exonuclease toward the recessed, nicked, andgapped DNA (26, 27) or even lowest efficiency on the recessedDNA duplex (28). These apparent discrepancies between ourdata and the previous studies could be explained by severalfactors, such as (i) different reaction conditions used, (ii) dif-ferent sequence context of DNA substrates used, and (iii) theuse of histidine-tagged APE1 instead of native form (19, 28).Interestingly, APE1 efficiently eliminates S-cdA nucleotide at the3′ end of all recessed, nicked, and gapped DNA duplexes (lanes6–8), suggesting that a single-strand break at the 3′ side of theS-cdA adduct in duplex DNA is sufficient for its removal. Afterremoving the 3′-terminal S-cdA nucleotide in the recessed DNAduplex, APE1-catalyzed 3′→5′ exonuclease activity continues todegrade DNA in a nonspecific manner (lane 6), whereas it isinhibited on the gapped and nicked DNA duplexes (lanes 7 and8). Strikingly, when acting upon nicked DNA duplexes APE1efficiently removes a 3′-terminal S-cdAMP (lane 8) but not a3′-terminal regular dAMP (lane 4). This result strongly suggeststhat APE1 recognizes S-cdA adduct with a high degree of speci-ficity when present at the 3′ end. However, APE1 exonucleasefunction is totally inhibited on the recessed cdAA•Trec, nickedcdAA•Tnick, and gapped cdAA•Tgap DNA duplexes whereS-cdA is located 1 nt before the 3′ end (lanes 10–12), in agree-ment with the results described above.

Characterization of APE1 Interactions with DNA Duplex Containing3′-Terminal S-cdA Adduct. To characterize the specificity of APE1interactions with S-cdA adducts, we measured the kinetic param-eters of excision of 3′-terminal S-cdA adduct by APE1 understeady-state conditions. Comparison of the kinetic constants for

Table 1. Sequence of the oligonucleotides used in the study, where X is S-cdA

Oligonucleotide name Sequence 5′→3′ Source

Upstream strandsExo20A, 21 mer d(GTGGCGCGGAGACTTAGAGAA) 20Exo20THF, 21 mer d(GTGGCGCGGAGACTTAGAGA-THF) 20, 30, 35Exo20PG, 21 mer d(GTGGCGCGGAGACTTAGAGA-PG) This studyExo20-ScdA, 21 mer d(GTGGCGCGGAGACTTAGAGAX) This studyExo19-ScdAA, 21 mer d(GTGGCGCGGAGACTTAGAGXA) This studycdA42, 42 mer d(AGAAACAACAGCACTACTGTACTCATGXATTCTATTCCAGCA) This study

Downstream strands3′-pExo19, 19 mer pd(ATTTGGCGCGGGGAATTCC) 16, 17, 20, 28, 303′-pExo18, 18 mer pd(TTTGGCGCGGGGAATTCC) 16

Template strandsRex-T, 40 mer d(GGAATTCCCCGCGCCAAATTTCTCTAAGTCTCCGCGCCAC) 20, 35T42, 42 mer d(TGCTGGAATAGAATTCATGAGTACAGTAGTGCTGTTGTTTCT) This studyT55, 55 mer d(CGAGGACAGACACTGCTGGAATAGAATTCATGAGTACAGTAGTGCTGTTGTTTCT) This study

Duplex name Oligonucleotides hybridizedA•Trec (nick/gap) Exo20A, Rex-T and 3′-pExo19 (nick) or 3′-pExo18 (gap) 20, 35THF•Trec Exo20THF and Rex-T 17, 35PG•Trec Exo20PG and Rex-T This studycdA•Trec (nick/gap) Exo20-ScdA, Rex-T and 3′-pExo19 (nick) or 3′-pExo18 (gap) This studycdAA•Trec (nick/gap) Exo19-ScdAA, Rex-T and 3′-pExo19 (nick) or 3′-pExo18 (gap) This study

Mazouzi et al. PNAS | Published online July 29, 2013 | E3073

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recessed DNA substrates containing different DNA lesions on 3′termini (Table 2) showed that the kcat/KM value for the WTAPE1-catalyzed excision of S-cdA adduct (1,700 min−1 μM−1)was two- and fivefold higher compared with that for 3′-terminalTHF residue (780 min−1 μM−1) and regular 3′-terminal dA nu-cleotide (360 min−1 μM−1), respectively. These results indicatethat the efficiency of APE1 3′ cleansing activity for the S-cdAadduct was comparable to that for 3′ sugar-phosphate group andsignificantly higher than for a regular deoxynucleotide in therecessed DNA duplex.Next, to understand the mechanism of inhibition of APE1

3′→5′ exonuclease activity on the recessed DNA duplex con-taining an S-cdA adduct located 1 nt away from the 3′ end, westudied the interactions between APE1 and 5′-[32P]-labeled A•Trec,cdA•Trec, and cdAA•Trec duplexes using an EMSA. APE1 bindsto recessed duplex cdA•Trec with 3′-terminal S-cdA more effi-ciently than to a regular A•Trec duplex and essentially fails toform stable DNA–protein complexes with cdAA•Trec duplex inwhich S-cdA nucleotide is located at the second position fromthe 3′ end of a gap (Fig. S3A). These results suggest that the lackof APE1 activity on recessed DNA duplexes containing an S-cdAadduct 1 or more nt away from the 3′ end might be due to theloss of enzyme affinity to the DNA substrate.

Structure of E. coli Nfo-H69A Mutant Bound to a Cleaved DNA DuplexReveals the Mechanism of Exonuclease Activity. To gain insight intothe structural basis of substrate specificity of E. coli Nfo andhuman APE1, we performed crystallographic studies using a15-mer DNA duplex containing a single α-anomeric 2′-deoxy-adenosine (αdA) nucleotide. The sequence context was takenfrom previous study by Garcin et al. (23) [Protein Data Bank(PDB) code 2NQJ] of the catalytically inactive mutant E261Qbut having an αdA•T pair instead of a AP•G pair at position 7.Unfortunately, all attempts to cocrystallize these two WT APendonucleases with αdA•T oligonucleotide duplex were un-successful because both enzymes contain a nonspecific 3′→5′exonuclease activity (29). Previously we have isolated Nfo-H69Amutant that contains reduced metal content and lacks NIR and3′→5′ exonuclease activities in the absence of divalent cations. Italso contains lower AP endonuclease/3′-diesterase activitiescompared with WT Nfo (30). Therefore, we decided to use thisNfo mutant for cocrystallization with DNA because it should notdegrade DNA in a nonspecific manner (17).We succeeded in obtaining a 1.9-Å resolution X-ray structure

of Nfo-H69A 15-mer αdA•T complex (Table 3). The electrondensity maps are of very good quality, and the asymmetric unitcontains two identical DNA-Nfo complexes with an rmsd valueof 0.13 Å between 279 Cα atoms. The small differences concernfour additional residues at the C terminus of molecule A (283residues) and the disordered terminal DNA base pair in mole-cule B, whereas the full DNA fragment is well defined in mol-ecule A. Additionally, this structure resembles previouslypublished Nfo:DNA containing AP site structures, such as1QUM (22) and 2NQJ (23), with respective rmsd values of only0.32 and 0.27 Å for all defined Cα atoms. Noticeably, DNAfragments present the same large distortion (Fig. 2A), meaning

that the WT enzyme, E261Q, and H69A mutants bend the DNAidentically by making similar protein–DNA interactions (Fig.2E). However, the state of the bound DNA in the Nfo-H69Astructure was unexpected. Instead of showing the base of αdAplaced in the solvent-accessible pocket on the enzyme surface toaccommodate its 5′ phosphate in the active site for a catalyticallycompetent complex, as proposed by Hosfield et al. (22), elec-tron density maps show that the αdA site 5′ phosphate is notconnected to the 3′ hydroxyl of the preceding nucleotide but is13.4 Å away from the cytosine C6 (Fig. 2B and Fig. S4). Thephosphodiester bond cleavage proves that the Nfo-H69A mutantwas able to cleave the DNA backbone 5′ of αdA in the crystal-lization solution, and thus it can recognize the αdA nucleotide asa target base (Fig. S5A). This cleavage occurred before the for-mation of the crystal because the structure represents a boundNIR product in a catalytically abortive complex for exonucleaseactivity (Fig. S5B).

Table 2. Steady-state kinetic parameters of the WT and D308A mutant APE1 proteins

Substrates

APE1 WT APE1 D308AFold decrease ofkcat/KM value,WT/D308A SourceKM, nM kcat, min

−1kcat/KM, min

−1

M−6 KM, nM kcat, min−1

kcat/KM, min−1

M−6

cdA•Trec 5.4 9.3 1700 15 5.5 360 4.7 This studyA•Trec 2.4 0.86 360 21 0.027 1.3 280.0 35THF•Trec 8.2 6.4 780 5.4 0.56 104 7.5 35

SDs for KM and kcat values varied within 20–40%. Details in Materials and Methods.

Table 3. Crystallographic and refinement data for NfoH69A–DNA complex

PDB code 4K1G

Data collectionBeamline PROXIMA 1 (SOLEIL)Wavelength (Å) 0.98Space group C2221Cell parameters (Å) a = 117.9, b = 136.6, c = 112.4Resolution (Å) 40–1.9 (2.02–1.9)No. of observed reflections 429,728 (68,483)No. of unique reflections 71,003 (11,229)Rsym (%)* 9.5 (68.1)Completeness (%) 99.7 (98.8)I/σ 13 (2.5)Redundancy 6Wilson B factor 26.58

Refinement statisticsRcryst (%)

† 17.6Rfree (%)

‡ 21.2rms bond deviation (Å) 0.01rms angle deviation (°) 1.1

Average B (Å2), molecules A; B (no. of atoms)Protein 24.5; 26.8 (4,366)Zn ions 20; 23.6 (4)DNA 46.8; 48.9 (1,182)Solvent 36.3 (610)

Ramachandran statistics (%)Preferred regions 98.92Allowed regions 1.08Outliers 0

Numbers in parentheses are for the highest-resolution range.*Rsym = Σhkl ΣijIi(hkl) - j/Σhkl Σi Ii (hkl), where Ii(hkl) is the i th observedamplitude of reflection hkl and is the mean amplitude for all obser-vations i of reflection hkl.†Rcryst = Σ jjFobsj − jFcalcjj/Σ jFobsj.‡5% of the data were set aside for free R-factor calculation.


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The base pair αdA7•T24 is well stacked in the DNA (Fig. 2B),and the phosphate group of αdA points toward the solvent. TheαdA base, which is 180° rotated around the C1’-N9 axis comparedwith an adenine base makes modified Watson–Crick contacts. ItsN6 atom interacts with the O2 and not O4 atom of the partnerthymine (Fig. 2B). Its N6 and N7 atoms make hydrogen bondswith Asn35 side chain. In contrast, the preceding base pair (C6:G25) is now unpaired, with both bases being extrahelical. Al-though G occupies the previously observed position of the flip-ped-out AP site opposite base in previously published Nfo•DNAcomplexes, the cytosine C6 occupies the position of the flipped-out AP site in a compact conformation (Fig. 2B). A 2-Å rear-rangement of Tyr72 side chain coupled to that of Gln36 allows C6to adopt this particular position. The base is maintained by twohydrogen bonds; its N4 atom interacts with the main chain car-bonyl group of the catalytic E261 residue and with a 5′ phosphateoxygen of the base upstream C5 (Fig. 2C). The C6 ribose O3′atom, rotated by 180° around the phosphate group, moving it 6.4 Åaway from its intrahelical position, points toward the mutatedresidue H69A and interacts with the E261 side chain (Fig. 2C).The phosphate group is 1.6 Å away from its position to becleaved. Moving this phosphate group to a position amenable forcatalysis leads to a displacement of the whole cytosine compatiblewith the presence of the His69 (Fig. S6). Thus, in the WT enzyme,we expect that the ribose O3′ atom would make an additionalinteraction with the His7 side chain, whereas the N4 atom wouldpreserve the interactions observed in the H69A mutant (Fig. S6).Replacing the C6 base by dT, dA, or dG shows that any naturalDNA base would be in contact with the 5′ phosphate group of thepreceding nucleotide and the E261 CO group.To assess the effect of S-cdA, we used data from a recently

published NMR structure (PDB 2LSF) of DNA containing thelesion (31). In contrast to natural DNA bases, replacing the C6nucleotide with a S-cdA adduct (referred to as 02I in PDB 2LSFof the NMR in structure) shows that the base would be less than

2 Å from the phosphate group of the preceding nucleotide,resulting in a clash (Fig. 2D). The coordinates of all models arepart of the Supporting Information, where Nfo coordinates forFig. 2D and Fig. S6 presented as Dataset S1; APE1 coordinatesfor Fig. 3A and Fig. 3C as Dataset S2; S-cdA model for Fig. 2Das Dataset S3; Nfo DNA model for Fig. 3A as Dataset S4;S-cdA model for Fig. 3C as Dataset S5; C6 model for Fig. S6as Dataset S6.These structural considerations can explain why E. coli Nfo-

H69A mutant cannot bind 3′-terminal S-cdAMP nucleotide incdA•Trec duplex contrary to nondamaged 3′-terminal dAMPnucleotide in A•Trec duplex. To test this prediction, we performedan EMSA using the mutant Nfo-H69A protein and 5′-[32P]-labeled A•Trec and cdA•Trec duplexes. The results showed thatunder the conditions used Nfo-H69A forms a stable DNA–proteincomplex only with regular A•Trec duplex but not with a damagedcdA•Trec one (Fig. S3B). These results are in agreement with ourstructural model demonstrating that the Nfo-H69Amutant canbind regular nicked DNA duplex in a nonproductive enzyme/substrate complex but loses its affinity to DNA duplex containing3′-terminal S-cdA adduct at a nick site.

Superimposition of APE1 and Nfo Active Site Structures: Model ofHuman APE1–3′-Terminal S-cdA Adduct Interactions and Effect ofD308A Mutation. Although Nfo and APE1 have distinct struc-tures, comparison of their active site conformations revealsstrong geometric conservation of the catalytic reaction, sup-porting a unified mechanism for AP site removal from DNA(32). Indeed, both enzymes flip-out the AP site in a similarconformation into the active site pocket. To determine whetherthe conformation and the position of the 3′-terminal nucleotidefor exonuclease activity were the same in Nfo and APE1, we useda structural analysis similar to that described by Tsutakawa et al.(32). Using the common AP site substrate, we superimposedtertiary structures of APE1 (1DEW) (24) and Nfo (2NQJ) (22)in complex with DNA. As described, the superposition based on

Fig. 2. Crystal structure of Nfo:αdA•T-DNA complex. (A) Superposition of the DNA fragment shown in ribbon and bound to Nfo: in magenta the 15 mercontaining an αdA•T site in the H69A structure with the 3′-terminal cytosine is cyan, in orange the same 15 mer containing an AP•G site in the inactive E261Qstructure (2NQJ.pdb). Nfo is shown in gray surface representation. (B) Close-up view showing the two unpaired C6:G25 in cyan. The αdA•T pair shown in pinkis well stacked, making Watson–Crick-like interactions. The cleavage separates the O3′ atom of the cytosine C6 13.4 Å away from its phosphodiester bond.Hydrogen bonds are shown as black dashes. (C) Close-up view of the cytosine C6 in an abortive complex for the intrinsic exonuclease activity of Nfo. Su-perposition around the DNA cleavage site of the H69A:DNA complex (in magenta except the cytosine C6 in cyan) and the E261Q:DNA complex containing anAP site (in orange). The ribose O3′ atom of C6 rotates by 180° around the phosphate group, moving it 6.4 Å away from its intrahelical position, and pointstoward the mutated residue H69A. The two present Zn ions are shown in green spheres and that lost in the H69A mutant in a red sphere. Hydrogen bonds areshown as black dashes. (D) Model of S-cdA at the position of the C6 shows steric hindrance: it is in close contact (less than 2 Å) with the phosphate group ofthe preceding nucleotide. (E) Schematic diagram of NfoH69A–DNA interactions. Polar interactions of DNA–protein side chains and DNA–backbone atoms areshown in blue and black arrows, respectively. The active site Zn2+ ions, which bind the phosphate of the C6 nucleotide, are shown as green spheres.


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only the THF moiety in the DNA oriented the respective scissile5′ phosphates and 3′ ribose oxygen atoms to overlay on top ofeach other. A portion of the DNA in Nfo:DNA complex (thestrand from the 5′-αdA nucleotide to the downstream end and itspartner strand) can be overlaid with that in APE1, and the scissilebond superimposes well (Fig. 3A and Fig. S7A). The superpositionof our present DNA structure describing the exonuclease activityof Nfo on DNA in APE1 was then straightforward (Fig. S7A). Thissuperposition reveals that the 3′-terminal cytosine C6 cannot beorientated and positioned in APE1 as it is in Nfo because ofsteric hindrances (Fig. S7B). Indeed, the ribose O3′ atom clasheswith the phenol ring of Phe266, the O2 atom is less than 2 Å tothe Trp280 side chain, and the N4 atom is at similar distances(less than 2 Å) from the carbonyl of N212 and the Cα of Gly231.Rotating the cytosine around the phosphate group (180° alongthe P-O5′ axis) leads to a unique position of C6 for APE1 exo-nuclease activity (Fig. 1A and Fig. S7C). To optimize this posi-tion, the Phe266 side chain needs to slightly reorient. Our modelconfirms the biochemical data showing that Phe266 mutationsincrease APE1-catalyzed 3′→5′ exonuclease activity (33). In ourmodel, the ribose O3′ atom is shifted by 3.3 Å away from itsintrahelical position and interacts with the Asn174 side chain andthe carbonyl group of Ala130 (Fig. 3B). The N4 and O2 atoms ofC6 hydrogen bond the Asp308 and Arg177 side chains, re-spectively. Replacing the C6 base by dA or dG shows thata regular purine could be adapted with a rearrangement of theArg177 side chain and would be involved in a contact withThr268. An S-cdA nucleotide at this position can also be ac-commodated making the same protein interactions (Fig. 3C).However, a close contact (around 2 Å) with Asp308 would forcethe side chain to slightly move. Therefore, Asp308 would havelittle or no effect on the removal of S-cdAMP. In agreement withthis, the D308A mutant is capable of removing the 3′-terminalS-cdA, THF, and phosphoglycolate (PG) adducts almost as ef-ficiently as WT APE1 (Table 2 and Fig. S8). As shown in Table 2,the kcat/KM values for the APE1 D308A-catalyzed excision of3′-terminal S-cdA, THF, and dA nucleotides were 5-, 7-, and280-fold lower compared with that of WT APE1, respectively,indicating that D308 residue is very important for the efficientremoval of 3′-terminal regular nucleotides but not that of S-cdAin agreement with our structure-based model of APE1-DNAexo interactions.

In Vitro Reconstitution and Repair of DNA Containing S-cdA Adductsin E. coli, Yeast, and Human Cell-Free Extracts.Removal of 3′-terminalS-cdAMP by AP endonuclease-catalyzed hydrolysis of the phos-phodiester bond on the 5′ side of the damaged nucleotide shouldgenerate 3′-OH termini and enable DNA synthesis. To examinethis, we reconstituted the repair of 5′-[32P]-labeled A•Trec andcdA•Trec oligonucleotide duplexes in the presence of the purifiedAPE1 and DNA polymerase β (POLβ) and dNTPs. Incubation ofA•Trec with POLβ generated elongated products up to the full-length 40-mer product after 30 min (Fig. S9). Interestingly, addi-tion of APE1 resulted in the appearance of degradation productswith size less than 21 mer owing to its processive 3′→5′ exo-nuclease activity. The presence of 3′-terminal S-cdA adduct incdA•Trec duplex strongly inhibited the first nucleotide insertionand completely blocked POLβ-catalyzed strand elongation from3′-terminal S-cdAA. As expected, addition of APE1 resulted inthe removal of 3′-blocking S-cdAMP and allowed the elongationand restoration of the damaged recessed DNA duplex by POLβ(Fig. S9). These results indicate that efficient elongation of DNAprimers containing 3′-blocking bulky S-cdA nucleotide can bereactivated by the 3′-repair cleansing function APE1.Data obtained with the purified AP endonucleases show that

S-cdA nucleotide at the 3′ end are efficiently repaired by E. coliXth and human APE1 enzymes but not by E. coli Nfo and yeastApn1. To address the role of the AP endonuclease-catalyzed 3′repair cleansing activity in a more physiological context, we ex-amined repair of 5′-[32P]-labeled cdA•Trec duplex in cell-free ex-tracts from E. coli, S. cerevisiae, and HeLa cells lacking Nfo/Xth,Apn1, and APE1, respectively (Fig. 4). As expected, we observedrobust nonspecific 3′→5′ exonuclease activities on regular A•Trecduplex in cell-free extracts from HeLa (Fig. 4, lane 2), S. cer-evisiae WT (lane 4), and E. coli WT (lane 17). The nonspecificexonuclease activity is strongly decreased in S. cerevisiae apn1(lane 5) and E. coli xth (lane 19) but not in siRNA-APE1 silencedHeLa cells (lane 3) and nfo (lane 18) compared with extracts

Fig. 3. Model of APE1 with 3′-terminal nucleotide. (A) Superposition of theDNA fragment of APE1:substrate (1DEW.pdb in green) and Nfo-H69A:exo (inmagenta). Only the part of the DNA fragment of Nfo-H69A that overlays isshown. APE1 is shown in gray surface representation. Because the cytosineC6 (shown in cyan) in exonuclease position in Nfo makes steric clashes inAPE1, the cytosine has been rotated around its phosphate group to find anadequate position in APE1 shown in the figure. (B) Close-up view of themodeled cytosine in exonuclease conformation in APE1. Hydrogen bonds areshown as black dashes. (C) Close-up view of a modeled S-cdA adduct in exo-nuclease position in APE1. Hydrogen bonds are shown as black dashes.

Fig. 4. 3′ repair activities in cell-free extracts of human, yeast, and E. colicells. 5′-[32P]-labeled recessed oligonucleotide duplexes (5 nM) were in-cubated with 1 μg of cell-free extract for 10 min at 37 °C. The “X” denotesthe position of S-cdA nucleotide. Details in Materials and Methods.


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from WT cells. As expected, cell-free extracts from HeLa andE. coli WT and nfo efficiently remove 3′-terminal S-cdA nucleotidein cdA•Trec duplex (lanes 7, 21, and 22). Importantly, extractsfrom siRNA-APE1 silenced HeLa cells and E. coli xth still containexonuclease activity on A•Trec (lanes 3 and 19) but have lost theactivity on cdA•Trec duplexes (lanes 8 and 23). No activity wasobserved on cdAA•Trec duplex in the extracts from HeLa andE. coli cells (lanes 12–13 and 25–27). Interestingly, the extractsfrom S. cerevisiae WT can remove with very low efficiencyS-cdAMP in cdA•Trec duplex (lane 9) and 3′-terminal dAMP incdAA•Trec duplex (lane 14), and this weak activity was not ob-served in apn1 extracts (lanes 10 and 15), thus confirming resultsobtained with the purified protein. These results suggest that inyeast extracts S-cdA adducts present in DNA duplex could beremoved, albeit inefficiently, by combined action of Apn1 andnonidentified factors. Taken together the results obtained withcell-free systems confirm the role of APE1, Apn1, and Xthproteins in the repair of bulky S-cdA adduct when present nextto a DNA strand break in human, yeast, and E. coli cells.

DiscussionStructurally unusual helix-distorting cdA and cdG adducts inDNA are biologically important owing to their strong inhibitoryeffect on DNA replication and transcription. The presence ofC8-C5′ covalent bond prevents repair of cdPu by DNA glyco-sylase-mediated excision and direct damage reversal. Therefore,cells use the NER system to remove cdPu adducts in vitro and invivo (4, 8, 9). However, ionizing radiation, breakage of replica-tion forks stalled at bulky DNA lesions, and/or misincorporationof oxidatively damaged precursors during DNA synthesis cangenerate cdPu lesions located in close proximity to strand breaks,making them resistant to NER. Therefore, alternative repairpathways may exist to remove these endogenous helix-distortingDNA lesions at DNA termini.Here, we investigated the mechanism of 3′→5′ exonuclease

activity of AP endonucleases involved in the BER and NIRpathways and their ability to recognize S-cdA adducts in DNAduplex. We have shown that the S-cdA nucleotide, when presentin fully duplex DNA, is not a substrate of the AP endonuclease-catalyzed NIR activity, but it can be a substrate of AP endonu-clease-catalyzed 3′→5′ exonuclease activity when present at the3′ end of single-stranded DNA break. E. coli Xth and humanAPE1, but not E. coli Nfo and yeast Apn1, remove S-cdAadducts at 3′ termini of recessed DNA duplex with high effi-ciency. However, when the S-cdA nucleotide is located 1 or morent away from the 3′ termini of a DNA duplex, it strongly blocksthe 3′→5′ exonuclease activity of all AP endonucleases tested.Our EMSA data suggest that APE1 fails to bind DNA duplexeswith S-cdA adduct located at the second position from the 3′ endof a gap. Taken together these results establish that 3′-terminalS-cdA adduct in gapped DNA duplex can be removed by analternative mechanism distinct from NER that involves 3′→5′exonuclease function of the Xth family AP endonucleases.It should be stressed that Xth and APE1 contain two distinct

activities toward DNA strand breaks: a 3′ repair diesterasefunction that catalyzes removal of 3′-dRP groups (34), and 3′→5′exonuclease, which catalyzes nonspecific removal of regular andoxidized bases (19, 20). Comparison of APE1 exonuclease ac-tivities on recessed, gapped, and nicked DNA duplexes as well askinetic data for recessed DNA substrates revealed that APE1was much more efficient on 3′-terminal S-cdA compared witha regular dA nucleotide. This strongly suggests that APE1-cat-alyzed 3′ cleansing activities are highly specific to damagedDNA. Interestingly, APE1 also removes mismatched nucleotidesfrom the 3′ terminus of DNA much more efficiency thannucleotides from matched pairs (26–28). It was suggested thatthe efficiency of APE1’s proof-reading exonuclease activitydepends primarily on the melted conformation of the 3′ end of

DNA duplex at the site of mismatch (26). Therefore, we proposethat helix-distorting S-cdA adduct at the 3′ end would resemblea melted duplex conformation similar to that of a 3′-terminalmismatch base pair and thus would facilitate the recognition ofthe adduct by APE1.At present there are no structural data available on AP en-

donuclease-catalyzed 3′→5′ exonuclease function. Here, we solvedthe crystal structure of Nfo mutant with nicked DNA duplex.Initially, the crystal structure of the Nfo-H69A mutant in com-plex with a 15 mer containing a αdA•T base pair at position 7was expected to reveal the molecular mechanism of the NIRactivity because the H69A mutant was previously shown to beinactive for both NIR and 3′→5′ exonuclease functions owing toa loss of one Zn atom (30). Surprisingly, the cocrystal structureshowed a cleaved αdA•T duplex (the expected product of WTNfo). The NIR product with the 3′-terminal cytosine C6 boundto the enzyme’s active site forms a nonproductive complex withthe Nfo-H69A mutant, which is unable to perform exonucleaseactivity. This structure of the Nfo-H69A:DNA complex with the3′-terminal cytosine C6 bound to the enzyme active site is veryinteresting and informative for understanding the mechanism of3′→5′ exonuclease activity of the WT Nfo, because both mutantand WT enzymes behave identically. Indeed, both Nfo proteinsrecognize an AP site and 3′-terminal nucleotide by binding anddistorting two DNA substrates by 90° and flipping-out the baseopposite the target AP site or the 3′-terminal base (Fig. 2 andFig. S6). Because the 3′-end of C6 nucleotide overlaps with theAP site, a slight shifting of the C6 nucleotide to bring it in anamenable position for catalysis was accomplished by super-imposing the complexed H69A and WT Nfo structures. It isimportant to note that the catalytic 5′-phosphate group in theresulting model stays in the same scissile position for both APendonuclease and 3′-exonuclease activities of Nfo, ensuring thehydrolysis of phosphodiester bond. Replacing C6 by either A, G,or T nucleotide was also straightforward. All these regular nucleo-tides can be accommodated in the WT Nfo active site, thusexplaining the structural basis of nonspecific 3′ exonucleasefunction of Nfo. It should be stressed that the compact andcurved position of the 3′-terminal C6 nucleotide, which interactswith the phosphate group of an upstream C5 nucleotide, is notcompatible with the presence of a S-cdA, which will generatesteric hindrance in the enzyme’s active site. Therefore, the in-ability of Nfo to remove 3′-terminal S-cdA adduct comes from aninadequate accommodation of this nucleotide in the enzymeactive site pocket. In agreement with previous data (30), thecomplex of Nfo with cleaved DNA well illustrates that after in-cision of the duplex 5′ next to the lesion site by NIR activity, Nfoproceeds further to degrade DNA by its 3′→5′ exonuclease ac-tivity at the site of the nick and that this latter function fails whenthe imidazole group of His69 and Zinc-1 atom are absent.Recent work by Tsutakawa et a. (32) demonstrated that ter-

tiary structures of APE1 and Nfo in complex with DNA can besuperimposed. Therefore, the structure of Nfo-H69A:DNA com-plex can serve as an excellent template for modeling of interactionsbetween active site of APE1 and DNA to provide the structuralbasis for the enzyme’s 3′ repair activities. Using superimposedactive site structures of APE1:AP site-DNA and Nfo:AP site-DNA complexes, we have been able to observe how the C6nucleotide in Nfo would be positioned in APE1. The confor-mation of C6 nucleotide adopted in the Nfo’s active site results insteric clashes with F266, W280, G231, and CO Asn212 amino acidresidues of APE1. To avoid steric hindrance and to accommodateC6 into the APE1 active site, we rotated the nucleotide around its5′ phosphate group, fixing it in catalytic position. In the resultingmodel, N174 interacts with the ribose O3′ atom of C6, whereasD308 interacts with the C6 base moiety and maintains the 3′-ter-minal nucleotide in a position favorable for exonuclease activityof APE1. Most interestingly, replacing the 3′ end C6 by an S-cdA


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nucleotide creates no steric hindrance except a small rear-rangement of D308 side chain to accommodate the bulky adduct.The present model of APE1 interactions with 3′-terminal nu-cleotide adducts provides the structural basis for the observed 3′repair exonuclease activity toward regular nucleotides and S-cdAadduct and reveals the role of D308 residue. In agreementwith this model, we showed that APE1-D308A mutant canremove S-cdA adduct in cdA•Trec duplex with good efficiency,similar to that of WT APE1 (Table 2), indicating that APE1removes the damaged bulky nucleotide by its 3′ repair cleansingfunction. Furthermore, we have previously demonstrated thatD308A mutant has dramatically decreased 3′→5′ exonucleaseactivity compared with WT APE1 (35).The lack of exonuclease activity toward S-cdA adduct in

cdAA•Trec duplex in all AP endonuclease tested observed in ourstudies is very intriguing. To examine the structural basis of thisexonuclease resistance, we used the recently published NMRstructure of duplex DNA containing S-cdA nucleotide (31) tosuperimpose it onto DNA bound to APE1. As described, thepresence of S-cdA nucleotide at position 6 in DNA duplex re-sults in the decrease of the phosphate oxygen atom (from residueat position 7)-O5′ (from S-cdA) bond distance that shortens byapproximately 1 Å (31), which could explain the lack of exo-nuclease activity toward S-cdA nucleotide in cdAA•Trec duplexin all AP endonucleases tested. Therefore, the shorter distanceof 6.3 Å between the phosphate group of S-cdA and that of the3′ nucleotide instead of 7.05 Å between two regular nucleotidesprobably leads to an incorrect positioning of the scissile phos-phodiester bond in the AP endonuclease active site and conse-quently to the loss of both 3′ repair activities and stable bindingto cdAA•Trec duplex. Finally, although the conserved structuralchemistry of active sites of APE1 and Nfo supports a unifiedmechanism for the AP site cleavage in DNA, our structuralmodels reveal that this mechanism cannot be extended to APE1and Nfo exonuclease activities. The conformation and position ofthe 3′-terminal nucleotide for exonuclease activity differs betweenthese two enzymes and thus involves different protein interactions.This also leads to a different DNA substrate specificity of theirrespective exonuclease functions. In conclusion, the structuraldata described in the present work provide insight into themechanism of 3′ cleansing activity of the Nfo and Xth families ofAP endonucleases.Ionizing radiation and certain anticancer drugs generate

complex or “dirty” DNA strand breaks containing 3′ end proxi-mal damaged bases, which are poorly repaired by classic BERand NER pathways (21, 36). Here we demonstrate that APE1-catalyzed removal of 3′-terminal S-cdA nucleotide in recessedDNA duplex enables otherwise blocked DNA polymerase syn-thesis in vitro, pointing to a possible role of APE1 in cleansing ofcomplex and/or dirty DNA strand breaks (Fig. 2). To furthersubstantiate physiological relevance of this specific repair func-tion of AP endonucleases, we demonstrated the presence of the3′ repair activities in cell-free extracts toward DNA duplexescontaining a 3′-terminal S-cdA adduct (Fig. 4). Importantly, underthe experimental condition used we did not observe significantrepair of cdAA•Trec duplex in any cell-free extracts tested. In-terestingly, we have identified a weak Apn1-independent activityon cdA•Trec duplex in yeast extracts and also found that extractsfrom WT cells and the purified Apn1 can remove a regular dAnucleotide in cdAA•Trec duplex, albeit with low efficiency. Thus,S. cerevisiae contains two enzymes: Apn1 and unknown exo-nuclease that can remove S-cdA adducts when located 1 or morent away from the 3′ end of strand breaks.Tirapazamine is an experimental bioreductively activated an-

ticancer drug that selectively kills cells under hypoxia (37). It hasbeen demonstrated that in vitro tirapazamine mediates forma-tion of 8,5′-cyclopurine-2’-deoxynucleosides in DNA underhypoxic condition, suggesting that these lesions may contribute

to the drug’s cytotoxicity (38). It is tempting to speculate thattirapazamine treatment may generate DNA strand breaks con-taining cdPu adducts and that this specific APE1 activity on the3′-terminal lesions described in the present work could providea rationale for the use of APE1 inhibitors to enhance effective-ness of tirapazamine and perhaps other anticancer drugs. Pre-viously, it was shown that APE1 removes β-L-Dioxolane-cytidine(L-OddC), a nonnatural stereochemical L-nucleoside analog,when incorporated at the 3′ terminus of duplex DNA by DNApolymerases (39). APE1 was also shown to remove 3′-tyrosylresidues from the recessed and nicked DNA duplexes, suggestingits potential role in the processing of covalent topoisomerase I:DNA complexes generated by anticancer drugs (27). Recently ithas been demonstrated that 5′S isomer of cdATP could be in-corporated more efficiently than the 5′R isomer by replicativeDNA polymerases during DNA synthesis (14). On the basis ofthe results reported here, we propose that APE1 may act ina similar manner on above adducts and also on 3′-terminalS-cdA nucleotides resulting from DNA polymerase catalyzedmisincorporation. Such an activity would be of relevance to theproposal that S-cdA could be used as an anticancer and/or an-tiviral drug (14).Finally, recent studies from the Dizdaroglu laboratory (40)

have found that both free R-cdA and S-cdA nucleosides can bedetected in human urine. Whether this material derived fromnuclease digestion of cdA-containing oligonucleotides resultingfrom NER, and/or APE1-catalyzed repair, and/or digestion ofDNA from dead cells remains to be determined. However, onthe basis of the current results, an intriguing possibility is thatAP endonucleases play a key role in the generation of urinarycdA, which could be a biomarker of endogenous oxidative stressto DNA.

Materials and MethodsOligonucleotides, Proteins, and Antibodies. Sequences of the oligonucleotidesand their duplexes used in the present work are shown in Table 1. All oli-gonucleotides were purchased from Eurogentec, including regular oligo-nucleotides and those containing S-cdA, αdA, THF, and PG. Before enzymaticassays oligonucleotides were either 5′-end-labeled by T4 polynucleotide ki-nase (New England Biolabs, Ozyme) in the presence of [γ-32P]-ATP (3,000 Ci/mmol-1) (PerkinElmer), or 3′-end-labeled by terminal deoxynucleotidyl trans-ferase (New England Biolabs) in the presence of [α-32P]-3′-dATP (Cordycepin5′-triphosphate, 5,000 Ci/mmol-1) (PerkinElmer) as recommended by themanufacturers. Radioactively labeled oligonucleotides were desalted witha Sephadex G-25 column equilibrated in water and then annealed withcorresponding complementary strands for 3 min at 65 °C in a buffer con-taining 20 mM Hepes-KOH (pH 7.6) and 50 mM KCl.

The sequence of the 15-mer DNA duplex used for crystallization assays isd(GCGTCCXCGACGACG)/d(CGTCGTCGTGGACGC), where X is αdA. The oli-gonucleotides were hybridized by mixing equal concentrations (10 mM) in2 mM Tris·HCl (pH 7.0) heated to 65 °C for 3 min and cooled down to roomtemperature over 2 h. The MALDI-TOF mass spectrometry analysis of theoligonucleotides performed by the manufacturer validated their size andhomogeneity. In addition the purity and integrity of the oligonucleotidepreparations were verified by denaturing PAGE. The siRNA sequences usedto decrease APE1 in HeLa cells have been taken from previously describedstudies (41). The siRNA specific to mouse major AP endonuclease, mAPEX,was used as negative control in both cases.

All AP endonucleases, their mutants, and human DNA glycosylase Neil1used in the present study were expressed and purified in their native formwithout tags or other modifications as described previously (16, 17). Thepurified human DNA POLβ and T4 DNA polymerase were purchased fromTrevigen and New England Biolabs, respectively.

Strains, Extract Preparation, Cell Culture, and Silencing of APE1 Expression.AB1157 [IeuB6 thr-1 Δ(gpt-proA2) hisG4 argE3 lacY1 gaIK2 ara-14 mtl-1xyl-5 thi-1 tsx-33 rpsL31 supE44 rac] (WT) and its isogenic derivatives BH130(nfo::kanR) and BH110 (nfo::kanR [Δ(xth-pncA)90 X::Tn10]) were from thelaboratory stock (42). S. cerevisiae FF18733 WT strain (MATa his7-3 leu2-1,112 lys1-1 trp1-289 ura3-52) and its isogenic derivative BG1 (apn1Δ::HIS3)


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were kindly provided by S. Boiteux (French Alternative Energies and AtomicEnergy Commission, France).

Crude cellular extracts from E. coli, S. cerevisiae, and HeLa cells with down-regulated APE1 expression were prepared as described previously (16, 20, 43).

DNA Repair Assays. The 3′-phosphodiesterase/exonuclease activity assay ofAPE1 was performed in the standard reaction mixture (20 μL) containing5 nM of [32P]-labeled oligonucleotide duplexes, 50 mM KCl, 20 mM Hepe-s·KOH (pH 6.8), 0.1 mg/mL BSA, 1 mM DTT, 1 mM MgCl2, and a limitedamount of pure protein or extract; when measuring repair activities in hu-man cell-free extracts BSA and DTT were omitted. For bacterial cell-freeextracts and the purified Xth and Apn1 proteins, the standard reactionmixture (20 μL) contained 5 nM of [32P]-labeled DNA substrate, 50 mM KCl,20 mM Hepes·KOH (pH 7.6), 0.1 mg/mL BSA, and 5 mM MgCl2, except whenincubating with Nfo, when MgCl2 was omitted from the buffer.

The reactions were stopped by adding 10 μL of a solution containing 0.5%SDS and 20 mM EDTA and then desalted by hand-made spin columns filledwith Sephadex G25 (Amersham Biosciences) equilibrated in 7.5 M urea.Purified reaction products were separated by electrophoresis in denaturing20% (wt/vol) polyacrylamide gels (7 M urea, 0.5× tris-borate-EDTA buffer,42 °C). Gels were exposed to a Fuji FLA-3000 Phosphor Screen and analyzedusing Image Gauge V3.12 software.

The kinetic parameters for exonuclease activity of APE1 were measured asdescribed previously (35). Briefly, 2–1,000 nM of duplex oligonucleotidesubstrate was incubated with 0.2 nM APE1 under standard reaction con-ditions, the linear velocity was measured, and the KM and kcat constantswere determined from Lineweaver-Burk plots. All biochemical experimentswere performed independently and repeated at least three times.

Crystallographic Analysis. The E. coli Nfo-H69A mutant was cloned, expressed,and purified as previously described (17). In crystallization trials, the 15-merDNA duplex was mixed with Nfo-H69A used at a concentration of 18 mg/mL ina buffer containing 50 mM KCl and 20 mM Hepes·KOH (pH 7.6) in a 2:1 stoi-chiometry. Commercial crystallization solutions (Qiagen kits) were screened in

sitting-drop vapor diffusion experiments using a nanodrop Cartesian robot(Proteomic Solutions) at 293 K. One condition [number 88: 30% (wt/vol) PEG4000, 0.1 M Tris·HCl (pH 8.5), and 0.2 M MgCl2] in the Classics suite wasmanually optimized with home-made solutions in hanging drops composed of1:1 volume ratio of crystallization solution and of Nfo-H69A:DNA complex.Crystals obtained in 0.1 M Tris·HCl (pH 8.0), 12% (wt/vol) PEG 4000, and 200mM MgCl2 were flash-frozen in a cryo-protecting solution consisting of themother solution supplemented with 20% (wt/vol) PEG 400.

X-ray diffraction data were collected at 100 K on beamline PROXIMA I atSOLEIL, and intensities were integrated using XDS19. The asymmetric unit cancontain two complexes of Nfo-H69A:DNA, corresponding to a Matthews co-efficient (44) of 2.78Å3Da−1 and a solvent content of 55.8%. The phase problemwas solved by molecular replacement using the program PHASER (45) and anNfo mutant:DNA structure (PDB code 2NQJ) as a search model. The resultingatomic model was refined using BUSTER (46) and manually improved usingCOOT23. Data collection and refinement statistics are given in Table 3. His-109 isnot well defined in the electron density owing to the absence of the Zn1 ion.

ACKNOWLEDGMENTS. We thank Dr. Jacques Laval for critical reading of themanuscript and thoughtful discussions, and Dr. Beatriz Guimaraes for help indata collection on PROXIMA I at SOLEIL. This work was supported by Fondationde France Grant 2012 00029161 (to A.A.I.) (www.fondationdefrance.org);Russian Federal Program “Scientific and education personnel for innovativeRussia” for 2009–2013 No. 8473 (to A.A.I.) (www.fcpk.ru); Centre Nationalde la Recherche Scientifique funds to S.M. and Grant PICS N5479-Russie (toM.K.S.) (www.cnrs.fr); Agence Nationale pour la Recherche Blanc 2010 ProjetANR-09-GENO-000 (to M.K.S.) (www.agence-nationale-recherche.fr); and Elec-tricité de France Contrat Radioprotection RB 2012 (to M.K.S.) (www.edf.fr). Thecrystallization work has benefited from the Laboratoire d’Enzymologie et Bio-chimie Structurales (LEBS) facilities of the IMAGIF Structural Biology and Pro-teomic Unit in the “Centre de Recherche de Gif” (www.imagif.cnrs.fr). A.M.and B.A. were supported by the student scholarships from Institut de Cancér-ologie Gustave-Roussy (www.igr.fr) and the Bolashak International Program,Kazakhstan (www.bolashak.gov.kz), respectively. Funding for open access chargewas provided by Agence Nationale pour la Recherche.

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insight into mechanisms of 3 -5 exonuclease activity and ...providing insight into the mechanism for...

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