apyrimidinic endonuclease 1 on specific DNA binding and catalysis

Biochimie 163 (2019) 73e83

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Research paper

The impact of single-nucleotide polymorphisms of human apurinic/ apyrimidinic endonuclease 1 on specific DNA binding and catalysis Irina V. Alekseeva a, Anastasiia T. Davletgildeeva a, b, Olga V. Arkova c, Nikita A. Kuznetsov a, b, **, Olga S. Fedorova a, b, * a b c

Institute of Chemical Biology and Fundamental Medicine, 8 Lavrentyev Ave., Novosibirsk, 630090, Russia Department of Natural Sciences, Novosibirsk State University, 2 Pirogova St., Novosibirsk, 630090, Russia Institute of Cytology and Genetics, Siberian Branch of Russian Academy of Sciences, 10 Lavrentyev Ave., Novosibirsk, 630090, Russia

a r t i c l e i n f o

a b s t r a c t

Article history: Received 29 March 2019 Accepted 23 May 2019 Available online 28 May 2019

Human apurinic/apyrimidinic (AP) endonuclease APE1 is a crucial enzyme of the base excision repair (BER) pathway, which is in charge of recognition and initiation of removal of AP-sites in DNA. It is known that some single-nucleotide polymorphism (SNP) variants of APE1 have a reduced activity as compared to wild-type APE1. It has been hypothesized that genetic variation in APE1 might be responsible for an increased risk of some types of cancer. In the present work, analysis of SNPs of the APE1 gene was performed to select the set of variants having substitutions of amino acid residues on the surface of the enzyme globule and in the DNA-binding site, thereby affecting proteineprotein interactions or the catalytic reaction, respectively. For seven APE1 variants (R221C, N222H, R237A, G241R, M270T, R274Q, and P311S), conformational dynamics and catalytic activities were examined. The conformational changes in the molecules of APE1 variants and in a DNA substrate were recorded as fluorescence changes of Trp and 2-aminopurine residues, respectively, using the stopped-flow technique. The results made it possible to determine the kinetic mechanism underlying the interactions of the APE1 variants with DNA substrates, to calculate the rate constants of the elementary stages, and to identify the stages of the process affected by mutation. © 2019 Elsevier B.V. and Société Française de Biochimie et Biologie Moléculaire (SFBBM). All rights reserved.

Keywords: DNA repair Human apurinic/apyrimidinic endonuclease Abasic site Stopped-flow enzyme kinetics Fluorescence

1. Introduction Cellular DNA during its functioning is constantly exposed to various endo- and exogenous factors, including highly reactive cell metabolites, alkylating compounds, and UV and ionizing irradiation, which can result in a chemical modification of nucleotides [1e4]. Such lesions in genomic DNA can lead to cardiovascular, neurodegenerative, and oncological diseases [5e8]. Furthermore,

Abbreviations: AP-site, apurinic/apyrimidinic site; APE1, AP endonuclease; aPu, 2-aminopurine; BER, base excision repair; DTT, dithiothreitol; F-site, (3hydroxytetrahydrofuran-2-yl)-methyl phosphate; ODN, oligodeoxyribonucleotide; PAGE, polyacrylamide gel electrophoresis; WT, wild type. * Corresponding author. Institute of Chemical Biology and Fundamental Medicine, 8 Lavrentyev Ave., Novosibirsk, 630090, Russia. ** Corresponding author. Institute of Chemical Biology and Fundamental Medicine, 8 Lavrentyev Ave., Novosibirsk, 630090, Russia. E-mail addresses: [email protected] (N.A. Kuznetsov), fedorova@ niboch.nsc.ru (O.S. Fedorova).

oxidative stress inducing accumulation of DNA lesions causes accelerated development of degenerative processes and premature aging [9e12]. Recognition and removal of nonbulky damaged nucleobases is mediated by the base excision repair pathway (BER), involving, in general, consecutive actions of DNA glycosylases, AP endonucleases, DNA polymerases, and DNA ligases [13]. Apurinic/apyrimidinic sites (AP-sites) are considered some of the most common lesions that occur in DNA spontaneously or owing to hydrolysis of N-glycosidic bonds catalyzed by DNA glycosylases. It has been estimated that more than 10000 AP-sites are formed in each mammalian cell daily [14] resulting in both cytotoxic and mutagenic effects [15]. Of note, cells and knockout animals lacking the activity of various DNA glycosylases are more sensitive to the influence of factors leading to DNA damage [16e18]. In contrast, geneticengineering removal of AP endonuclease from the cell results in its death, indicating the crucial role of this enzyme in DNA repair

https://doi.org/10.1016/j.biochi.2019.05.015 0300-9084/© 2019 Elsevier B.V. and Société Française de Biochimie et Biologie Moléculaire (SFBBM). All rights reserved.

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[19]. Human AP endonuclease APE1 indeed is a key enzyme in BER, being responsible for the processing of AP-sites and a number of other DNA lesions [20e25]. The main physiological function of this enzyme is thought to be the hydrolysis of the phosphodiester bond 50 to an AP-site, leading to a break in the ribose-phosphate backbone and formation of 30 -hydroxyl and 20 -deoxyribose-50 -phosphate termini [26e28]. Analysis of crystal structures of the APE1 enzyme in a free form [29e31] and in complex with DNA [32e34] has shown that in the catalytically competent conformation, the AP-site is flipped out from the DNA double helix (Fig. 1). Amino acid residues of the DNAbinding site interact predominantly with the damaged strand and form hydrogen bonds and electrostatic contacts with DNA phosphate groups. Residues Asp70, Glu96, Tyr171, Asn174, Asp210, Asn212, Asp308, and His309 form an active site of the enzyme. Stabilization of the “flipped-out” conformation of the AP-site is implemented by residues Met270 and Arg177. The base opposite to the AP-site is displaced by Met270 inserted into the minor groove of DNA. At the same time, Arg177 is integrated into the major groove, forming a hydrogen bond 30 to the AP-site. The phosphate group located 50 to the AP-site forms coordination bonds with Asn174, Asn212, and His309. The catalytic reaction is initiated by a nucleophilic attack of the oxygen atom of an H2O moleculedcoordinated directly or through a Mg2þ ion with Asp210don the phosphorus atom 50 of the AP-site [32,34]. Amino acid substitutions in BER enzymes associated with

single-nucleotide polymorphisms (SNPs) are widespread in the human population [35,36]. The relation between SNP variants of BER proteins and etiology of some human diseases requires further investigation. Moreover, some polymorphisms of the human APendonuclease gene (APE1) have been reported to decrease the enzymatic activity [37e41]. Such variants of APE1 in the human population can be associated with an increased risk of diseases as demonstrated for other BER enzymes [42,43]. The association of SNPs of BER genes with some diseases was recently discussed in a number of reviews [44,45]. It should be noted that a decrease in the functional activity of individual BER enzymes and disruption of coordination between them or with protein factors such as XRCC1 (which acts as a scaffold protein in the BER pathway) can have severe negative consequences for the human body [46e48]. Therefore, our aim was to determine the influence of known SNPs on the conformation and activity of human APE1. Here, seven SNP variants of APE1 (R221C, N222H, R237A, G241R, M270T, R274Q, and P311S) were chosen for the stopped-flow analyses of conformational dynamics and kinetics of interaction with DNA substrates containing F-site as a stable analog of an abasic nucleotide. Conformational changes of APE1 and of the DNA substrate were recorded by changes in fluorescence intensity of Trp residues and of a 2-aminopurine (aPu) base located opposite the abasic site, respectively. Analysis of the kinetic data uncovered the stages of the APE1 enzymatic process most affected by an SNP. 2. Methods 2.1. Choosing potentially important SNPs Using the NCBI dbSNP database (http://www.ncbi.nlm.nih.gov/ SNP/), exonic polymorphic variants of the human APE1 gene were selected. Subsequent analysis of potential importance of an SNP was based on a principle of a maximal change in the chemical nature of the side group of the amino acid encoded by a corresponding nucleotide triplet. Spatial location of the amino acid residue close to the DNA-binding site and/or active site of the enzyme was taken into account as well, on the basis of http://www. ncbi.nlm.nih.gov/Structure/(Conserved Domains and Protein Classification) data. According to these principles, seven SNPs were chosen for the study of their influence on APE1 activity (Table 1, Fig. 2). Potential importance of these SNPs was additionally analyzed in the PolyPhen-2 software (Polymorphism Phenotyping v2, http://genetics.bwh.harvard.edu/pph2/). 2.2. Site-directed mutagenesis and protein purification Mutations Arg221Cys, Asn222His, Arg237Ala, Gly241Arg Met270Thr, Arg274Gln, or Pro311Ser within the APE1 coding sequence were generated using a site-directed mutagenesis kit (QuikChange XL, Stratagene). For expression of the recombinant proteins, 1 L of Escherichia coli strain Rosetta II(DE3) culture (Invitrogen, France) (in LuriaeBertani [LB] broth) carrying the pET11aAPE1 construct was grown at 100 mg/mL ampicillin and 37  C until absorbance at 600 nm (A600) reached 0.6e0.7; APE1 variant expression was induced overnight with 0.2 mM isopropyl-b-Dthiogalactopyranoside. The isolation and purification of the enzymes were performed as in Refs. [49,50]. The protein concentration was measured by the Bradford method; stock solutions were stored at 20  C. 2.3. DNA substrates

Fig. 1. Schematic representation of APE1 complexed with F-siteecontaining DNA (PDB ID 1DE8). Amino acid residues of the active site are highlighted in red. “Void-filling” residues Met270 and Arg177 are blue.

The sequences of the DNA substrates used in this work are presented in Table 2. Oligodeoxyribonucleotides (ODNs) were

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Table 1 Amino acid substitutions analyzed in this work. Amino acid Location substitution R221C N222H R237A G241R M270T R274Q P311S

PolyPhen-2 prediction

DNA-binding site (distance 8.0 Å between Nh1 of Arg221 and O atom of 30 -phosphate group of second nucleotide on 30 side of F-site). DNA-binding site (distances 4.5 and 5.0 Å between Nd2 atom of Ans222 and two O atoms of 50 - and 30 -phosphate groups of second nucleotide on 30 side of F-site). Internal coordination of a-helixes (distances 3.9 and 7.5 Å between Nh1 of Arg237 and Oε1 atom of Glu217 and Od1 atom of Asp219, respectively). Exterior of protein globule.

benign probably damaging probably damaging probably damaging Active site. Met270 is embedded into DNA minor groove, thereby displacing base opposite to AP site. possibly damaging Internal coordination near active site (distances 2.6 and 3.0 Å between Nh1/Nh2 of Arg274 and carbonyl O atoms of Ala304 and Ser307, probably respectively). damaging probably Structural element at end of b-sheet, bordering catalytic loop. damaging

presence of 20 mM triethylammonium acetate, pH 7.0, for 30 min at a flow rate of 2 mL/min. Fractions containing ODNs were dried in vacuum, dissolved in water, and precipitated with 2% LiClO4 in acetone. After washing with pure acetone and drying, the ODN precipitates were dissolved in water and stored at 20  C until use. Concentrations of the ODNs were determined by means of A260. Homogeneity of the purified ODNs was evaluated by electrophoresis in a 20% denaturing polyacrylamide gel, containing 8 M urea and 0.1 M Tris-borate buffer (pH 8.3). The ODNs were visualized with the Stains-All dye (Sigma, USA). DNA duplexes were prepared by annealing of modified and complementary strands at the 1:1 M ratio in reaction buffer (50 mM Tris-HCl pH 7.5, 50 mM KCl, 1.0 mM EDTA, 1.0 mM DTT, 5.0 mM MgCl2, and 7% of glycerol). 2.4. Stopped-flow fluorescence measurements

Fig. 2. Structure of the APE1 complex with F-DNA (PDB ID 1DE8). Amino acids chosen for analysis are blue.

Table 2 Oligonucleotide substrates. Shorthand

Sequence

50 -TCTCTCFCCTTCC-30 30 -AGAGAGGGGAAGG-50 F/aPu-substrate 50 -TCTCTC F CCTTCC-30 30 -AGAGAG (aPu)GGAAGG-50

Stopped-flow measurements with fluorescence detection were carried out essentially as described previously [51e53] using a model SX.18 MV stopped-flow spectrometer (Applied Photophysics Ltd., Leatherhead, UK). To detect intrinsic Trp fluorescence only, the wavelength lex ¼ 290 nm was chosen for excitation, and emission at lem > 320 nm was detected as transmitted by a Schott filter WG320 (Schott, Mainz, Germany). If aPu was present in the ODNs, lex of 310 nm was used to excite aPu fluorescence, and its emission was monitored at lem > 370 nm (Corion filter LG-370). The concentration of the enzyme in all the experiments with Trp fluorescence detection was 1.0 mM, and concentrations of F/G-substrate were varied from 0.5 to 2.0 mM. The concentration of F/aPusubstrate in the experiments with aPu fluorescence detection was 1.0 mM, and the enzyme concentration was varied from 0.5 to 2.0 mM. Typically, each trace shown is the average of four or more individual experiments. All the experiments were carried out at 25  C in a buffer consisting of 50 mM Tris-HCl pH 7.5, 50 mM KCl, 1.0 mM EDTA, 1.0 mM DTT, 5.0 mM MgCl2, and 7% of glycerol (v/v).

F/G-substrate

2.5. Analysis of the DNA cleavage time course The damaged DNA strand (50 -TCTCTCFCCTTCC-30 ) was 50 -end P-labeled and annealed to the complementary strand. To start the reaction, the solutions of F/G-substrate and enzyme in the buffer were mixed to obtain the final concentrations 1.0 mM and 0.1 mM, respectively. Aliquots (2 mL) of the reaction mixture were taken at certain time points, immediately quenched with 3 mL of a gelloading dye containing 7 M urea and 50 mM EDTA, and loaded on a 20% (w/v) polyacrylamide/7 M urea gel. The gels were imaged by autoradiography and quantified by scanning densitometry in GelPro Analyzer 4.0 software (Media Cybernetics, Silver Spring, MD). The kinetic traces of product accumulation were fitted to a single 32

synthesized by standard phosphoramidite methods on an ASM-700 synthesizer (BIOSSET Ltd., Novosibirsk, Russia) using phosphoramidites purchased from Glen Research (Sterling, VA). The synthetic oligonucleotides were purified with HPLC using an Agilent 1200 chromatograph (USA) and a Zorbax SB-C18 column (5 mm), 4.6  150 mm, via a linear gradient of acetonitrile (0 / 50%) in the

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exponential curve by means of the Origin software (Originlab Corp., USA; Eq. (1)). [Product] ¼ A  [1  exp(kobst)]

(1)

where A is the amplitude, kobs is the observed rate constant, and t is reaction time (s). 2.6. Circular dichroism (CD) spectra CD spectra were recorded on a Jasco J-600 spectropolarimeter (Jacso, Japan), at 5  C in quartz cells with 1 cm light path length. The concentration of APE1 in the device cell was 1.0 mM. The experiments were carried out in a buffer consisting of 50 mM Tris-HCl pH 7.5, 50 mM KCl, 1.0 mM EDTA, and 5.0 mM MgCl2. The spectra were recorded at bandwidth 1.0 nm and resolution 1.0 nm with a scan speed of 50 nm/min. The scans were accumulated and automatically averaged. 2.7. Global fitting of stopped-flow data The sets of kinetic curves obtained at different concentrations of the reactants were analyzed in the DynaFit software (BioKin, Pullman, WA) [54] as described elsewhere [55e58]. The kinetic curves represent fluorescence intensity variations in the course of the reaction owing to the sequential formation and subsequent transformation of the DNAeenzyme complex and its conformers. The stopped-flow fluorescence traces were directly fitted to fluorescence intensity (F) at any reaction time point (t) as the sum of the background fluorescence and fluorescence intensity values of each intermediate complex formed by the enzyme with DNA:

F ¼ Fb þ

n X

fi  ½ESi

(2)

i¼0

where Fb is background fluorescence or the equipment-related photomultiplier parameter (“noise”), fi is the molar response coefficient of the i-th intermediate ESi (i ¼ 0 corresponds to the free protein and i > 0 to the enzymeeDNA complexes). The concentrations of each species in the mechanisms are described by a set of differential equations according to a kinetic scheme (see Results). The software performs numerical integration of a system of ordinary differential equations with subsequent nonlinear least-squares regression analysis. In the fits, the values of all relevant rate constants for the forward and reverse reactions are optimized, as are the specific molar “response factors” for all intermediate complexes. 3. Results 3.1. Rationale Earlier [59e61], the stopped-flow preesteady-state kinetic

analysis, combined with measurements of protein Trp and DNA aPu fluorescence, has been applied to detect and quantitatively describe conformational transitions of the enzymeeDNA complex corresponding to specific steps of the APE1 reaction. DNA duplexes containing a native AP-site or F-site analog served as substrates. The Trp fluorescence traces of APE1's reaction with abasic DNA comprised a few characteristic phases, which were assigned to DNA-binding steps and catalysis in our previous reports [59e61]. The initial double-phase decrease in the Trp fluorescence intensity could be attributed to the process of formation of a specific enzymeesubstrate complex that is responsible for catalytic hydrolysis of the internucleotide phosphodiester bond. After the catalytic reaction, the enzymeeproduct complex dissociates, resulting in an increase of fluorescence intensity. These conclusions were also proved by the finding that in the case of the interaction of catalytically inactive apo-APE1 (without Mg2þ in the active site) with abasic DNA in the presence of EDTA, the Trp fluorescence intensity was not increased at the ends of kinetic traces [61]. Besides, the Trp fluorescence time-course recorded during the preesteadystate stopped-flow experiments correlated with the kinetics of the product accumulation revealed by PAGE. It was also shown that the increase phase in the Trp fluorescence curves coincided with the emergence of a product in a chemical quench assay [60]. All these findings allowed us to attribute the definite phases of the fluorescence curves to binding or catalytic steps of the kinetic mechanism. As a result, a minimal kinetic mechanism was proposed (see Scheme 1). According to Scheme 1, the interaction of APE1 with a substrate involves at least two reversible steps, corresponding to binding to the DNA substrate and AP-site recognition, followed by transformation of the initial enzymeesubstrate complex in a catalytically competent conformation. Then, the irreversible catalytic step of AP-site 50 -phosphodiester bond hydrolysis proceeds, followed by the final step of the catalytic mechanism: equilibrium dissociation of the enzymeeproduct complex. Here, the same approach was used for the analyses of catalytic properties of APE1 SNP variants. The conformational dynamics of APE1 and model DNA substrates were studied at short time intervals (from 1 ms up to 20 s), under the conditions close to singleenzyme turnover. Conformational changes of the enzyme were recorded as Trp fluorescence intensity changes, and those of the DNA as changes in fluorescence of the aPu residue located opposite the F-site in the complementary strand. 3.2. CD analysis of APE1 variant structures The extent of changes in the conformation of APE1 variants after an amino acid substitution was evaluated by the CD method. As shown in Fig. 3, CD spectra of APE1 mutants contain two negative bands, near 208 and 222 nm, which are characteristics of the ahelical structure of a protein. The band near 222 nm is observed due to the strong hydrogen-bonding interior of this conformation. This transition is relatively independent of the length of the helix. On the other hand, the intensity of the negative band near 208 nm is lower in short helices. It is seen on Fig. 3 that the negative band

Scheme 1. The kinetic mechanism of APE1 processing of the F/G-substrate. Е: enzyme, S: substrate, (ES)1 and (ES)2: enzymeesubstrate complexes, Р: substrate transformation product, EP: enzymeeproduct complex, ki and k-i: rate constants of the equilibrium stage's forward and reverse reactions, kcat: rate constant of a catalytic reaction, Kp: equilibrium constant of ЕР complex dissociation.

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decrease in Trp fluorescence intensity in the initial part of kinetic curves [59e61] reveals the binding of an enzyme to DNA followed by formation of the catalytically competent complex. The catalytic stage of the process leading to product formation and subsequent dissociation of the enzymeeproduct complex is followed by an increase in Trp fluorescence intensity at later time points, starting from ~1 s. Analysis of the kinetic curves characterizing the interactions of APE1 variants revealed that the minimal kinetic mechanism corresponds to the one proposed earlier [59e61] (Scheme 1) for WT APE1. The rate constants for forward and reverse reactions and the Kp equilibrium constant corresponding to Scheme 1 are given in Table 3. 3.5. Conformational changes of damaged DNA induced by WT and SNP variants of APE1 Fig. 3. CD spectra of WT APE1 and its SNP variants.

near 208 nm is slightly affected in variants P311S, R237A, G241R, and N222H. These data suggested that the tested mutations do not induce significant structural rearrangements of the enzyme. 3.3. Activity of SNP variants of APE1 The activity of APE1 variants was studied by direct PAGE analysis of the kinetics of accumulation of products derived from the 32Plabeled F/G-substrate (Fig. 4A and B). Multiple turnover conditions allowed to use eq [1]. to calculate the values of observed rate constants kobs, which are presented in the form of a histogram in Fig. 4C. As shown in this figure, the strongest effect on enzymatic activity is caused by SNP R237A. Notably, substitution G241R leads to a ~50% increase in the observed rate constant of DNA cleavage. All other APE1 variants possess enzymatic activities 20e30% lower than that of the wild type (WT). 3.4. Conformational dynamics of WT APE1 and its SNP variants in the course of interaction with DNA The conformational transitions in the molecules of WT APE1 and its SNP variants in the course of interaction with F/G-substrate were monitored as changes in the intrinsic Trp fluorescence. The fluorescence traces presented in Fig. 5 for different concentrations of F/ G-substrate display complex behavior. As shown previously, the

To determine the effect of amino acid substitutions in APE1 (caused by SNPs) on the conformational changes in the DNA substrate, the changes in fluorescence intensity of aPu residues placed opposite to F-site in F/aPu-substrate were monitored next. As depicted in Fig. 6, in the initial phase of the process up to 50 ms, aPu fluorescence intensity decreased, with a subsequent growth phase up to 5 s (Fig. 6). It is known that the fluorescence intensity of aPu in DNA is strongly dependent on a fluorophore microenvironment [62e64], specifically, an increase of hydrophobicity in close proximity to the fluorophore decreases aPu fluorescence intensity. Thus, the decrease in fluorescence intensity occurring in the initial part of the kinetic curves may represent the displacement of residues Arg177 and Met270 (which are embedded in the DNA duplex) from the major and minor grooves. At the later time points of the reaction, formation of a catalytic complex (resulting in 50 -phosphodiester bond hydrolysis and subsequent dissociation of the enzymeeproduct complex) increases aPu fluorescence intensity (Fig. 6). The analysis of kinetic curves characterizing interactions of WT APE1 or its SNP variants R237A, N222H, G241R, R274Q, R221C, M270T, and P311S with F/aPu-substrate yielded a minimal kinetic mechanism (Scheme 2). This scheme contains one reversible step (corresponding to the binding of the enzyme to DNA substrate and leading to formation of a catalytically active complex), an irreversible catalytic step, and an equilibrium step of enzymeeproduct complex dissociation. Rate constants of forward and reverse reactions in Scheme 2 as well as the equilibrium constant Kp are presented in Table 4.

Fig. 4. Enzymatic activity of WT APE1 and its SNP variants toward F/G-substrate. PAGE analysis of reaction product accumulation in the reaction with (A) WT APE1, kobs ¼ 0.08 s1 and (B) R237A APE1, kobs ¼ 0.013 s1. (C) Relative cleavage activity of APE1 and its SNP variants. Concentration of enzyme was 0.1 mM, DNA concentration was 1.0 mM.

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Fig. 5. Changes in Trp fluorescence intensity during the interaction of WT APE1 (А), G241R APE1 (B), R237A APE1 (C), N222H APE1 (D), R274Q APE1 (E), R221C APE1 (F), M270T APE1 (G), and P311S APE1 (H) with F/G-substrate. APE1 concentration was 1.0 mM, concentration of F/G-substrate varied from 0.5 to 2.0 mM (a.u. is arbitrary units).

Table 3 The kinetic parameters of F/G-substrate cleavage by WT APE1 or its SNP variants. Constants

WT

R221C

N222H

R237A

G241R

M270T

R274Q

P311S

k1, M1s1,  106 k-1, s1 K1, M1,  106 k2, s1 k-2, s1 K2 Kass, M1s kcat, s1 Kp, M,  106

450 ± 80 490 ± 50 0.9 ± 0.3 150 ± 30 16 ± 3 9.4 ± 3.6 8.5  106 1.0 ± 0.1 50 ± 10

280 ± 100 260 ± 80 1.1 ± 0.7 26 ± 2 13 ± 1 2.0 ± 0.3 2.2  106 1.1 ± 0.2 25 ± 10

520 ± 30 230 ± 40 2.3 ± 0.5 70 ± 25 13 ± 1 5.4 ± 2.3 12.4  106 1.2 ± 0.1 8.9 ± 1.3

560 ± 150 390 ± 80 1.4 ± 0.7 70 ± 20 15 ± 4 4.7 ± 2.6 6.6  106 0.6 ± 0.1 580 ± 170

730 ± 180 250 ± 100 2.9 ± 1.8 70 ± 25 45 ± 10 1.6 ± 0.9 4.6  106 1.6 ± 0.3 110 ± 40

360 ± 90 500 ± 180 0.7 ± 0.4 30 ± 4 7.5 ± 1.7 4.0 ± 1.4 2.8  106 1.2 ± 0.2 2.0 ± 0.2

510 ± 60 380 ± 50 1.3 ± 0.3 8.2 ± 1.4 28 ± 10 0.3 ± 0.1 0.4  106 1.0 ± 0.4 480 ± 80

300 ± 50 450 ± 70 0.7 ± 0.2 23 ± 9 19 ± 4 1.2 ± 0.7 0.8  106 0.9 ± 0.2 130 ± 80

Ki ¼ ki/k-i. Kass ¼ K1  K2.

4. Discussion The comparison of activities of WT APE1 and SNP variants in PAGE analysis (Fig. 4C) reveals that amino acid substitutions R221C,

N222H, M270T, R274Q, and P311S moderately decrease the activity of human AP endonuclease (by 70e80% of the WT enzymatic activity). Variant R237A (the affected residue controls the location of two a-helixes in the protein globule) manifested a substantial loss

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Fig. 6. Changes in aPu fluorescence intensity during interactions of WT APE1 (А), G241R APE1 (B), R237A APE1 (C), N222H APE1 (D), R274Q APE1 (E), R221C APE1 (F), M270T APE1 (G), and P311S APE1 (H) with F/aPu-substrate. F/aPu-substrate concentration was 1.0 mM, enzyme concentration varied from 0.5 to 2.0 mM (a.u. is arbitrary units).

Table 4 The kinetic parameters of F/aPu-substrate cleavage by WT APE1 or its SNP variants.* Constants

WT

R221C

N222H

R237A

G241R

M270T

R274Q

P311S

k1, M1s1,  106 k-1, s1 Kass, M1 kcat, s1 Kp, M,  106

55 5.9 9.3  106 1.49 0.80

12 26 0.5  106 1.2 50

42 9.6 4.4  106 1.8 2.8

17 4.3 3.9  106 1.1 220

36 16 2.2  106 2.0 2.2

15 2.4 6.2  106 1.0 9

6 20 0.3  106 1.8 1.4

5 19 0.26  106 1.3 60

Kass ¼ k1/k-1. * Standard deviations in the values of constants are 30e50%.

of activity: kobs ¼ 0.013 s1 in comparison with kobs ¼ 0.08 s1 for the WT (see Fig. 4B). The G241R substitution was found (Fig. 4C) to cause a ~50% increase in the enzymatic activity. This increase in activity may be associated with the appearance of an additional positive charge on the enzyme surface and accelerated formation of the primary collisional complex between the protein and DNA. Previously [37], only a ~40e60% reduction in activity was registered for the R237A variant in comparison with WT APE1, and

enhancement of specific endonuclease activity was noted in the G241R variant. To elucidate which steps of the enzymatic process are more influenced by an amino acid substitution, the conformational changes of the enzyme and DNA substrate during their interactions were monitored, allowing us to separately follow the stages of damaged DNA site binding and the subsequent catalytic stage of phosphodiester bond hydrolysis. A comparison of kinetic curves

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obtained by recording the changes in Trp and aPu fluorescence intensity for all APE1 variants at the same concentrations of the enzyme and substrate is shown in Fig. 7. The figure indicates that during interactions of WT APE1 or its variants with F/G-substrate, Trp fluorescence decreased in the initial part of kinetic curves, thus revealing a similar way of formation of the catalytically active complex. An increase in Trp fluorescence intensity occurring at later time points corresponds to the catalytic stage of the process and to the subsequent dissociation of the enzymeeproduct complex. Such changes in fluorescence intensity for APE1 SNP variants take place in a time range close to the one observed for the WT enzyme. Even though the kinetic curves characterizing the interactions of the WT enzyme or its SNP variants with DNA look similar, the stepwise analysis suggests that amino acid substitutions can influence both DNA binding stages and the catalytic stage of the enzymatic process separately (Table 3). A comparison of rate constants of binding and catalytic steps obtained by recording aPu fluorescence in the DNA substrate (Fig. 7A, Table 4) indicates that the R237A substitution affects DNA binding stages only slightly (Kass is 9.4  106 and 8.0  106 M1 for WT and R237A, respectively). Meanwhile, this substitution results in a 1.7-fold decrease in catalytic rate constant kcat. Analysis of Trp conformational dynamics (Fig. 7A, Table 3) revealed that the G241R substitution raises the rate constant for the formation of the primary complex between APE1 and a DNA substrate, in agreement with the data obtained by the PAGE method. Kass is lower for the G241R enzyme in comparison with the WT (Table 3, 4.6  106 and 8.5  106 M1, respectively). Calculation of the concentration of the G241R enzyme complex with DNA, at 1.0 mM concentration of the enzyme and DNA-substrate (stopped-flow conditions) revealed that this decrease in Kass does not affect the binding equilibrium and that 100% of the enzyme is in the complex with DNA. Moreover, in the case of the G241R variant, the acceleration of the catalytic stage was noted too: kcat is 1.0 s1 and 1.6 s1 for the WT and G241R, respectively (see Table 3). Therefore, the acceleration in the case of the G241R enzyme is mostly provided by the increase in the catalytic rate constant. Substitutions R274Q and P311S (affecting an amino acid residue located in the active site region) exerted a significant effect on the second stage of DNA substrate binding (K2 being 30- and 8-fold less than that for the WT, respectively), which characterizes formation of a catalytically competent complex, according to earlier data [50,61]. Calculation of the concentration of the enzymeesubstrate complex under PAGE analysis conditions ([enzyme] ¼ 0.1 mM, [DNA] ¼ 1.0 mM) reveals that no more than 30% of the enzyme is in the complex with DNA. Thus, destabilization of the protein globule

in the active site region has a significant influence on the step of formation of the catalytically competent complex and leads to decrease of enzyme activity of these mutants. To some extent, this conclusion can be drawn for the R221C variant, which manifests a reduction in Kass approximately 4-fold in comparison with the WT (Table 3). Moreover, the R221C substitution also has a large effect on the second stage, which leads to the catalytic-complex formation (K2 is 4.7-fold smaller than that for the WT) as in the case of R274Q and P311S variants. The conformational changes in the enzyme also indicate insignificance of Asn222 (located close to DNA binding site) for the overall enzyme efficiency. Nevertheless, the N222H substitution causes a small increase in the association constant K1 for primarycomplex formation and in total substrate binding constant Kass. Nonetheless, this change of Kass does not affect the binding equilibrium because the enzyme is fully bound to DNA under stoppedflow conditions. The reason for the deceleration of N222H variant activity under the conditions of PAGE analysis ([enzyme] ¼ 0.1 mM, [DNA] ¼ 1.0 mM) most likely is a decrease in Kp (Table 3, 8.9  106 M and 50  106 M for N222H and WT, respectively), characterizing dissociation of the complex of the enzyme with the DNA product. Given that amino acid residue Asn222 is located in the DNA-binding site, the substitution of Asn222 with a positively charged His residue leads to stabilization of the complex with the DNA product and decreases the enzyme efficacy under multipleturnover conditions. The substitution of Met270 embedded into the double helix to stabilize the flipped out state of the damaged nucleotide does not significantly affect either DNA binding or catalysis. Nevertheless, M270T variant has a reduced dissociation constant toward the DNA product (Tables 3, 2  106 M) which leads to a decrease in enzyme efficacy under multiple-turnover conditions as in the case of N222H variant. Analysis of DNA substrate conformational changes during its interaction with APE1 was conducted by means of a fluorescent analog of adenine: 2-aminopurine (Fig. 7B). It has to be mentioned that the aPu residue located in the complementary strand opposite to F-site does not show high sensitivity to conformational changes of the DNA substrate. Nevertheless, a series of kinetic curves, which include those corresponding to higher concentrations of mutant forms, enabled estimation of the rate constants for the step of formation of the enzymeesubstrate complex and a subsequent catalytic reaction (Scheme 2). Therefore, some discrepancy seen during the comparison of rate and equilibrium constants obtained in Trp and aPu fluorescence experiments (for example, for N222H, Kass is higher than that of the WT in Table 3 and lower than that of

Fig. 7. Changes in Trp (A) and aPu (B) fluorescence intensity during the interaction of WT APE1 or its SNP variants with DNA substrates. The concentrations of enzymes and DNA substrate were 1.0 mM. The kinetic traces are manually offset for clarity (a.u. is arbitrary units).

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Scheme 2. The kinetic mechanism of APE1 processing of the F/aPu-substrate. Е: enzyme, S: substrate, (ES): enzymeesubstrate complex, Р: substrate transformation product, EP: enzymeeproduct complex, k1 and k-1: rate constants of the equilibrium stage's forward and reverse reactions, kcat: rate constant of a catalytic reaction, Kp: equilibrium constant of ЕР complex dissociation.

the WT in Table 4) may be due to the low accuracy of fluorescence detection of the aPu reporter. If we compare Schemes 1 and 2, it is possible to conclude that the two reversible binding steps in Scheme 1 and one binding step in Scheme 2 must result in formation of the same catalytically competent complex. Therefore, the equation (K1  K2)Trp ¼ KaPu 1 should be valid. The values presented in Tables 3 and 4 show that this is true almost always. Both sets of quantitative data obtained by monitoring of Trp and aPu fluorescence lead to similar conclusions. As in the case of Trp fluorescence detection, the aPu fluorescence data reveal a decrease in the rate of the catalytic stage for variant R237A and its increase for variant G241R. Therefore, for variants R237A and G241R, the results of both Trp and aPu fluorescence analysis (Tables 3 and 4) indicate the opposite effect of these substitutions on the catalytic stage of the enzymatic process. A comparison of the rate and the equilibrium constants (Table 4) suggests that substitutions R274Q and P311S (in the active site region) lead to a >10-fold decrease in total association constant Kass relative to the WT enzyme. The R221C substitution also results in a slowdown of a phase of the decrease in aPu fluorescence intensity (Fig. 7B). As illustrated in Table 4, this substitution decreases the binding constant ~20-fold as compared to the WT enzyme. A smaller difference was found during Trp fluorescence detection (Table 3), but this discrepancy can be explained by greater experimental error in the case of aPu. It also seems that the apparent slowdown of aPu fluorescence intensity in this case is associated with a disruption of specific contacts between Arg221 and DNA. The absence of the Arg221 residue in the enzyme molecule changes the polarity in the aPu region but does not affect much the general efficacy of the enzyme, as suggested by PAGE analysis (Fig. 4) and Trp fluorescence data (Figs. 5 and 7). 5. Conclusions In this report, kinetic analysis of DNA excision by WT human AP endonuclease APE1 and its natural SNP variants R221C, N222H, R237A, G241R, M270T, R274Q, and P311S was conducted. The influence of these amino acid substitutions on the conformational changes in APE1 and in DNA substrates at the stages of catalyticcomplex formation and catalysis was studied by the stopped-flow fluorescence method. Monitoring the fluorescence intensities of Trp residues in proteins or of an aPu residue in the substrate allows us to analyze changes (in the microenvironments of the fluorescent groups) occurring in the enzyme or substrate molecules owing to changes in their conformation in the course of the enzymatic process. Our data revealed multiple effects of an amino acid substitution on APE1 catalysis. It was demonstrated that substitutions R221C, N222H, M270T, R274Q, and P311S yield only a 20e30% decrease in the enzymatic activity. Substitution R237A leads to ~6fold reduction in the rate of the catalytic stage. In contrast, the other natural variant G241R shows increased enzymatic activity, caused mainly by an increase in the catalytic-stage rate. Variants N222H and G241R having greater positive charge on the protein surface

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have a tendency for accelerated formation and stabilization of the primary collisional complex. Besides, amino acid substitutions R274Q and P311S in the region of the catalytic site result in a reliable decrease in the rates and equilibrium constants of the stages leading to formation of the catalytically active complex. In general, the obtained data indicate that these natural SNP variants of human AP endonuclease do not show any dramatic changes in their catalytic activities, except for R237A. Nevertheless, a genetic knockout of APE1 is embryonically lethal [65], and even a knockdown of APE1 activity raises mutation rates, sensitivity to oxidative stress, and the incidence of tumor formation [66e68]. Therefore, it seems that all existing natural APE1 variants must possess a catalytic activity not less than some threshold value, which is sufficient for rigorous enzymatic activity in the cell. Nevertheless, because AP endonuclease is a multifunctional enzyme, the amino acid substitutions induced by SNPs may have a major influence on different functions of APE1, for instance proteineprotein interactions with other base excision enzymes or a nucleosome, ribonuclease activity, or the redox function. Most likely, to predict a possible disease in a person as a consequence of DNA repair dysfunction, it is necessary to take into consideration a combination of various polymorphisms of BER proteins rather than one SNP of a single protein [44,45]. Author contributions NAK and OSF designed the study and contributed reagents/ materials. IVA and ATD performed the experiments. OVA performed SNP database analysis. IVA, ATD, NAK and OSF drafted the manuscript. All authors contributed to data analysis, read, corrected and approved the final version of the manuscript. Declaration of interests The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. Acknowledgments This work was supported partially by a Russian Governmentefunded project (No. АААА-А17-117020210022-4) and a grant from the Russian Foundation for Basic Research (19-0400012). The part of the study dealing with Trp and aPu detection combined with stopped-flow kinetics was specifically funded by Russian Science Foundation grant 18-14-00135. References [1] S.S. Wallace, Biological consequences of free radical-damaged DNA bases, Free Radic. Biol. Med. 33 (2002) 1e14. [2] L.J. Marnett, Oxyradicals and DNA damage, Carcinogenesis 21 (2000) 361e370. [3] M. Dizdaroglu, P. Jaruga, M. Birincioglu, H. Rodriguez, Free radical-induced damage to DNA: mechanisms and measurement, Free Radic. Biol. Med. 32 (2002) 1102e1115. [4] S. Boiteux, M. Guillet, Abasic sites in DNA: repair and biological consequences in Saccharomyces cerevisiae, DNA Repair 3 (2004) 1e12. [5] M.S. Cooke, M.D. Evans, M. Dizdaroglu, J. Lunec, Oxidative DNA damage: mechanisms, mutation, and disease, FASEB J. 17 (2003) 1195e1214. [6] M.D. Evans, M. Dizdaroglu, M.S. Cooke, Oxidative DNA damage and disease: induction, repair and significance, Mutat. Res. 567 (2004) 1e61. [7] Y. Xie, H. Yang, C. Cunanan, K. Okamoto, D. Shibata, J. Pan, D.E. Barnes, T. Lindahl, M. McIlhatton, R. Fishel, J.H. Miller, Deficiencies in mouse Myh and Ogg1 result in tumor predisposition and G to T mutations in codon 12 of the K-Ras oncogene in lung tumors, Cancer Res. 64 (2004) 3096e3102. [8] S. Maynard, S.H. Schurman, C. Harboe, N.C. de Souza-Pinto, V.A. Bohr, Base excision repair of oxidative DNA damage and association with cancer and aging, Carcinogenesis (2009), https://doi.org/10.1093/carcin/bgn250. [9] S. Raha, B.H. Robinson, Mitochondria, oxygen free radicals, disease and ageing,

82

I.V. Alekseeva et al. / Biochimie 163 (2019) 73e83

Trends Biochem. Sci. 25 (2000) 502e508. [10] Z. Radak, Z. Bori, E. Koltai, I.G. Fatouros, A.Z. Jamurtas, I.I. Douroudos, G. Terzis, M.G. Nikolaidis, A. Chatzinikolaou, A. Sovatzidis, S. Kumagai, H. Naito, I. Boldogh, Age-dependent changes in 8-oxoguanine-DNA glycosylase activity are modulated by adaptive responses to physical exercise in human skeletal muscle, Free Radic. Biol. Med. 51 (2011) 417e423. [11] G.S. Leandro, P. Sykora, V.A. Bohr, The impact of base excision DNA repair in age-related neurodegenerative diseases, Mutat. Res. Mol. Mech. Mutagen. 776 (2015) 31e39. [12] F. Coppede, L. Migliore, DNA damage in neurodegenerative diseases, Mutat. Res. Mol. Mech. Mutagen. 776 (2015) 84e97. [13] E.C. Friedberg, G.C. Walker, W. Siede, R.D. Wood, R.A. Schultz, T. Ellenberger, DNA Repair and Mutagenesis, ASM Press, Washington, 2006. [14] T. Lindahl, Instability and decay of the primary structure of DNA, Nature 362 (1993) 709e715. [15] D.M. Wilson III, D. Barsky, The major human abasic endonuclease: formation, consequences and repair of abasic lesions in DNA, Mutat. Res. 485 (2001) 283e307. [16] A. Klungland, I. Rosewell, S. Hollenbach, E. Larsen, G. Daly, B. Epe, E. Seeberg, T. Lindahl, D.E. Barnes, Accumulation of premutagenic DNA lesions in mice defective in removal of oxidative base damage, Proc. Natl. Acad. Sci. U.S.A. 96 (1999) 13300e13305. [17] J.L. Parsons, R.H. Elder, DNA N-glycosylase deficient mice: a tale of redundancy, Mutat. Res. 531 (2003) 165e175. [18] H. Nilsen, I. Rosewell, P. Robins, C.F. Skjelbred, S. Andersen, G. Slupphaug, G. Daly, H.E. Krokan, T. Lindahl, D.E. Barnes, Uracil-DNA glycosylase (UNG)deficient mice reveal a primary role of the enzyme during DNA replication, Mol. Cell 5 (2000) 1059e1065. [19] H. Fung, B. Demple, A vital role for Ape1/Ref1 protein in repairing spontaneous DNA damage in human cells, Mol. Cell 17 (2005) 463e470. [20] C.D. Mol, S.S. Parikh, C.D. Putnam, T.P. Lo, J.A. Tainer, DNA repair mechanisms for the recognition and removal of damaged DNA bases, Annu. Rev. Biophys. Biomol. Struct. 28 (1999) 101e128. [21] S.S. David, S.D. Williams, Chemistry of glycosylases and endonucleases involved in base-excision repair, Chem. Rev. 98 (1998) 1221e1261. [22] A.A. Ischenko, M.K. Saparbaev, Alternative nucleotide incision repair pathway for oxidative DNA damage, Nature 415 (2002) 183e187. [23] L. Gros, A.A. Ishchenko, H. Ide, R.H. Elder, M.K. Saparbaev, The major human AP endonuclease (Ape1) is involved in the nucleotide incision repair pathway, Nucleic Acids Res. 32 (2004) 73e81. [24] P. Prorok, D. Alili, C. Saint-Pierre, D. Gasparutto, D.O. Zharkov, A.A. Ishchenko, B. Tudek, M.K. Saparbaev, Uracil in duplex DNA is a substrate for the nucleotide incision repair pathway in human cells, Proc. Natl. Acad. Sci. U S A. 110 (2013) E3695eE3703. [25] A.A. Kuznetsova, A.G. Matveeva, A.D. Milov, Y.N. Vorobjev, S.A. Dzuba, O.S. Fedorova, N.A. Kuznetsov, Substrate specificity of human apurinic/apyrimidinic endonuclease APE1 in the nucleotide incision repair pathway, Nucleic Acids Res. 46 (2018) 11454e11465. [26] B. Demple, J.-S. Sung, Molecular and biological roles of Ape1 protein in mammalian base excision repair, DNA Repair 4 (2005) 1442e1449. [27] N.S. Dyrkheeva, S.N. Khodyreva, O.I. Lavrik, Multifunctional human apurinic/ apyrimidinic endonuclease 1: the role of additional functions, Mol. Biol. 41 (2007) 450e466. [28] A. Kuznetsova, O. Fedorova, N. Kuznetsov, Kinetic features of 30 -50 exonuclease activity of human AP-endonuclease APE1, Molecules 23 (2018) 2101. [29] M.A. Gorman, S. Morera, D.G. Rothwell, E. de La Fortelle, C.D. Mol, J.A. Tainer, I.D. Hickson, P.S. Freemont, The crystal structure of the human DNA repair endonuclease HAP1 suggests the recognition of extra-helical deoxyribose at DNA abasic sites, EMBO J. 16 (1997) 6548e6558. [30] P.T. Beernink, B.W. Segelke, M.Z. Hadi, J.P. Erzberger, D.M. Wilson 3rd, B. Rupp, Two divalent metal ions in the active site of a new crystal form of human apurinic/apyrimidinic endonuclease, Ape1: implications for the catalytic mechanism, J. Mol. Biol. 307 (2001) 1023e1034. [31] B.A. Manvilla, E. Pozharski, E.A. Toth, A.C. Drohat, Structure of human apurinic/apyrimidinic endonuclease 1 with the essential Mg2þ cofactor, Acta Crystallogr. D Biol. Crystallogr. 69 (2013) 2555e2562. [32] C.D. Mol, T. Izumi, S. Mitra, J.A. Talner, J.A. Tainer, DNA-bound structures and mutants reveal abasic DNA binding by APE1 and DNA repair coordination, Nature 403 (2000) 451e456. [33] C.D. Mol, D.J. Hosfield, J.A. Tainer, Abasic site recognition by two apurinic/ apyrimidinic endonuclease families in DNA base excision repair: the 3’ ends justify the means, Mutat. Res. 460 (2000) 211e229. [34] S.E. Tsutakawa, D.S. Shin, C.D. Mol, T. Izumi, A.S. Arvai, A.K. Mantha, B. Szczesny, I.N. Ivanov, D.J. Hosfield, B. Maiti, M.E. Pique, K.A. Frankel, K. Hitomi, R.P. Cunningham, S. Mitra, J.A. Tainer, Conserved structural chemistry for incision activity in structurally non-homologous apurinic/ apyrimidinic endonuclease APE1 and endonuclease IV DNA repair enzymes, J. Biol. Chem. 288 (2013) 8445e8455. [35] M.R. Shen, I.M. Jones, H. Mohrenweiser, Nonconservative amino acid substitution variants exist at polymorphic frequency in DNA repair genes in healthy humans, Cancer Res. (1998), https://doi.org/10.1046/j.13652656.1999.00279.x. [36] H.W. Mohrenweiser, I.M. Jones, Variation in DNA repair is a factor in cancer susceptibility: a paradigm for the promises and perils of individual and population risk estimation? Mutat. Res. Fund Mol. Mech. Mutagen (1998)

https://doi.org/10.1016/S0027-5107(98)00059-1. [37] M.Z. Hadi, M.A. Coleman, K. Fidelis, H.W. Mohrenweiser, D.M. Wilson III, Functional characterization of Ape1 variants identified in the human population, Nucleic Acids Res. (2000), https://doi.org/10.1093/nar/28.20.3871. [38] W.C. Kim, C. Ma, W.M. Li, M. Chohan, D.M. Wilson, C.H. Lee, Altered endoribonuclease activity of apurinic/apyrimidinic endonuclease 1 variants identified in the human population, PLoS One (2014), https://doi.org/10.1371/ journal.pone.0090837. [39] J.L. Illuzzi, N.A. Harris, B.A. Manvilla, D. Kim, M. Li, A.C. Drohat, D.M. Wilson, Functional assessment of population and tumor-associated APE1 protein variants, PLoS One 8 (2013), e65922. [40] J.M. Hinz, P. Mao, D.R. McNeill, D.M. Wilson, Reduced Nuclease activity of apurinic/apyrimidinic endonuclease (APE1) variants on nucleosomes: identification of access residues, J. Biol. Chem. 290 (2015) 21067e21075. [41] J.L. Illuzzi, D.R. McNeill, P. Bastian, B. Brenerman, R. Wersto, H.R. Russell, F. Bunz, P.J. McKinnon, K.G. Becker, D.M. Wilson, Tumor-associated APE1 variant exhibits reduced complementation efficiency but does not promote cancer cell phenotypes, Environ. Mol. Mutagen. 58 (2017) 84e98. [42] B. Karahalil, V.A. Bohr, D.M. Wilson, Impact of DNA polymorphisms in key DNA base excision repair proteins on cancer risk, Hum. Exp. Toxicol. (2012), https://doi.org/10.1177/0960327112444476. [43] J.B. Sweasy, T.M. Lang, D. DiMaio, Is base excision repair a tumor suppressor mechanism? Cell Cycle 5 (2006) 250e259. [44] S.S. Wallace, D.L. Murphy, J.B. Sweasy, Base excision repair and cancer, Cancer Lett. (2012), https://doi.org/10.1016/j.canlet.2011.12.038. [45] C.G. Marsden, J.A. Dragon, S.S. Wallace, J.B. Sweasy, Base excision repair variants in cancer, Methods Enzymol. 591 (2017) 119e157. [46] R. Abbotts, D.M. Wilson, Coordination of DNA single strand break repair, Free Radic. Biol. Med. 107 (2017) 228e244. [47] D. Kwiatkowski, P. Czarny, P. Galecki, A. Bachurska, M. Talarowska, A. Orzechowska, K. Bobinska, A. Bielecka-Kowalska, T. Pietras, J. Szemraj, M. Maes, T. Sliwinski, Variants of base excision repair genes MUTYH, PARP1 and XRCC1 in Alzheimer's disease risk, Neuropsychobiology 71 (2015) 176e186. [48] P. Czarny, D. Kwiatkowski, M. Toma, J. Kubiak, A. Sliwinska, M. Talarowska, J. Szemraj, M. Maes, P. Galecki, T. Sliwinski, Impact of single nucleotide polymorphisms of base excision repair genes on DNA damage and efficiency of DNA repair in recurrent depression disorder, Mol. Neurobiol. 54 (2017) 4150e4159. [49] A.A. Kuznetsova, N.A. Kuznetsov, A.A. Ishchenko, M.K. Saparbaev, O.S. Fedorova, Pre-steady-state fluorescence analysis of damaged DNA transfer from human DNA glycosylases to AP endonuclease APE1, Biochim. Biophys. Acta 1840 (2014) 3042e3051. [50] A.D. Miroshnikova, A.A. Kuznetsova, N.A. Kuznetsov, O.S. Fedorova, Thermodynamics of damaged DNA binding and catalysis by human AP endonuclease 1, Acta Naturae 8 (2016) 103e110. [51] D.A. Yakovlev, A.A. Kuznetsova, O.S. Fedorova, N.A. Kuznetsov, Search for modified DNA sites with the human Methyl-CpG-binding enzyme MBD4, Acta Naturae 9 (2017) 88e98. [52] A.A. Kuznetsova, D.A. Iakovlev, I.V. Misovets, A.A. Ishchenko, M.K. Saparbaev, N.A. Kuznetsov, O.S. Fedorova, Pre-steady-state kinetic analysis of damage recognition by human single-strand selective monofunctional uracil-DNA glycosylase SMUG1, Mol. Biosyst. 13 (2017) 2638e2649. [53] O.A. Kladova, A.A. Kuznetsova, O.S. Fedorova, N.A. Kuznetsov, Mutational and kinetic analysis of lesion recognition by Escherichia coli endonuclease VIII, Genes 8 (2017) 1e13. [54] P. Kuzmic, Program DYNAFIT for the analysis of enzyme kinetic data: application to HIV proteinase, Anal. Biochem. 237 (1996) 260e273. [55] O.A. Kladova, L.N. Krasnoperov, N.A. Kuznetsov, O.S. Fedorova, Kinetics and thermodynamics of DNA processing by wild type DNA-glycosylase endo III and its catalytically inactive mutant forms, Genes (2018), https://doi.org/ 10.3390/genes9040190. [56] N.A. Kuznetsov, A.A. Kuznetsova, Y.N. Vorobjev, L.N. Krasnoperov, O.S. Fedorova, Thermodynamics of the DNA damage repair steps of human 8oxoguanine DNA glycosylase, PLoS One 9 (2014), e98495. [57] N.A. Kuznetsov, Y.N. Vorobjev, L.N. Krasnoperov, O.S. Fedorova, Thermodynamics of the multi-stage DNA lesion recognition and repair by formamidopyrimidine-DNA glycosylase using pyrrolocytosine fluorescence– stopped-flow pre-steady-state kinetics, Nucleic Acids Res. 40 (2012) 7384e7392. [58] N.A. Kuznetsov, V.V. Koval, D.O. Zharkov, O.S. Fedorova, Conformational dynamics of the interaction of Escherichia coli endonuclease VIII with DNA substrates, DNA Repair 11 (2012) 884e891. [59] N.A. Timofeyeva, V.V. Koval, D.G. Knorre, D.O. Zharkov, M.K. Saparbaev, A.A. Ishchenko, O.S. Fedorova, Conformational dynamics of human AP endonuclease in base excision and nucleotide incision repair pathways, J. Biomol. Struct. Dyn. 26 (2009) 637e652. [60] L.Y. Kanazhevskaya, V.V. Koval, D.O. Zharkov, P.R. Strauss, O.S. Fedorova, Conformational transitions in human AP endonuclease 1 and its active site mutant during abasic site repair, Biochemistry 49 (2010) 6451e6461. [61] A.D. Miroshnikova, A.A. Kuznetsova, Y.N. Vorobjev, N.A. Kuznetsov, O.S. Fedorova, Effects of mono- and divalent metal ions on DNA binding and catalysis of human apurinic/apyrimidinic endonuclease 1, Mol. Biosyst. 12 (2016) 1527e1539. [62] D.C. Ward, E. Reich, L. Stryer, Fluorescence studies of nucleotides and

I.V. Alekseeva et al. / Biochimie 163 (2019) 73e83

[63] [64]

[65]

[66]

polynucleotides. I. Formycin, 2-aminopurine riboside, 2,6-diaminopurine riboside, and their derivatives, J. Biol. Chem. 244 (1969) 1228e1237. J.M. Jean, K.B. Hall, 2-Aminopurine fluorescence quenching and lifetimes: role of base stacking, Proc. Natl. Acad. Sci. U.S.A. 98 (2001) 37e41. E.L. Rachofsky, R. Osman, J.B.A. Ross, Probing structure and dynamics of DNA with 2-aminopurine: effects of local environment on fluorescence, Biochemistry 40 (2001) 946e956. S. Xanthoudakis, The redox/DNA repair protein, Ref-1, is essential for early embryonic development in mice, Proc. Natl. Acad. Sci. Unit. States Am. (1996), https://doi.org/10.1073/Pnas.93.17.8919. L.B. Meira, S. Devaraj, G.E. Kisby, D.K. Burns, R.L. Daniel, R.E. Hammer, S. Grundy, I. Jialal, E.C. Friedberg, Heterozygosity for the mouse Apex gene

83

results in phenotypes associated with oxidative stress, Cancer Res. 61 (2001) 5552e5557. [67] J. Huamani, C.A. McMahan, D.C. Herbert, R. Reddick, J.R. McCarrey, M.I. MacInnes, D.J. Chen, C.A. Walter, Spontaneous mutagenesis is enhanced in Apex heterozygous mice, Mol. Cell. Biol. (2004), https://doi.org/10.1128/ MCB.24.18.8145-8153.2004. [68] A. Unnikrishnan, J.J. Raffoul, H.V. Patel, T.M. Prychitko, N. Anyangwe, L.B. Meira, E.C. Friedberg, D.C. Cabelof, A.R. Heydari, Oxidative stress alters base excision repair pathway and increases apoptotic response in apurinic/ apyrimidinic endonuclease 1/redox factor-1 haploinsufficient mice, Free Radic. Biol. Med. (2009), https://doi.org/10.1016/j.freeradbiomed.2009.02.021.