Critical Sites of DNA Backbone Integrity for Damaged Base Removal

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Article Cite This: Biochemistry 2019, 58, 2740−2749

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Critical Sites of DNA Backbone Integrity for Damaged Base Removal by Formamidopyrimidine−DNA Glycosylase Anton V. Endutkin†,‡ and Dmitry O. Zharkov*,†,‡ †

SB RAS Institute of Chemical Biology and Fundamental Medicine, 8 Lavrentieva Avenue, Novosibirsk 630090, Russia Novosibirsk State University, 2 Pirogova Street, Novosibirsk 630090, Russia

‡

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S Supporting Information *

ABSTRACT: DNA glycosylases, the enzymes that initiate base excision DNA repair, recognize damaged bases through a series of precisely orchestrated movements. Most glycosylases sharply kink the DNA axis at the lesion site and extrude the target base from the DNA double helix into the enzyme’s active site. Little attention has been paid so far to the role of the physical continuity of the DNA backbone in allowing the required conformational distortion. Here, we analyze base excision by formamidopyrimidine−DNA glycosylase (Fpg) from substrates keeping all phosphates but containing a nick within three nucleotides of the lesion in either DNA strand. Four phosphoester linkages at the damaged nucleotide and two nucleotides 3′ to it were essential for Fpg activity, while the breakage of the others, even at the same critical phosphates, had no effect or even stimulated the reaction. Reduction of the likelihood of hydrogen bonding at the nicks by using dideoxynucleotides as their 3′-terminal groups was more detrimental for the activity. All phosphoester bonds in the complementary strand were dispensable for base excision, but nicks close to the orphaned nucleotide caused early termination of damaged strand cleavage. Elastic network analysis of Fpg−DNA structures showed that the vibrational motions of the critical phosphates are strongly correlated, in part due to the presence of the protein. Overall, our results suggest that mechanical forces propagating along the DNA backbone play a critical role in the correct conformational distortion of DNA by Fpg and possibly by other target base-everting DNA glycosylases.

residues into the DNA duplex. Structural and computational studies indicate that the eversion not only is necessary to achieve the catalytically competent conformation but also provides several transient conformational intermediates that are important for distinguishing between the damaged and normal bases.7−10 One of the best-studied DNA glycosylases is formamidopyrimidine-DNA glycosylase (Fpg, or MutM; EC 3.2.2.23), which cleanses bacterial DNA of 8-oxoguanine (oxoG) and several other oxidatively damaged purines, the miscoding or blocking lesions that are abundantly produced in DNA both by normal aerobic metabolism byproducts and under oxidative stress.11,12 Inactivation of Fpg in bacteria increases the mutation rate13 and the sensitivity toward many widely used drugs.14,15 As a typical DNA glycosylase, Fpg dynamically extrudes oxoG from the base stack, with several kinetic gates on its way from fully intrahelical to fully extrahelical serving to discriminate against normal G9,16−24 (Figure 1A,B). The large, highly precise, and coordinated conformational change in substrate DNA upon damage recognition likely requires exquisitely located and timed interactions between protein and DNA residues. Given that DNA glycosylases

DNA glycosylases, a structurally heterogeneous group of enzymes, are central for protecting the genome against widespread base damage, initiating the base excision repair pathway.1,2 Chemically, all of these enzymes hydrolyze the Nglycosidic bond of damaged or undamaged but inappropriately placed nucleotides, with some also capable of cleaving the DNA backbone by β-elimination.1,3 Most of the DNA glycosylases known thus far, on the basis of their domain structure and the presence of conserved structural motifs, are classified into one of three structurally unrelated groups: uracilDNA glycosylase superfamily, helix−hairpin−helix superfamily, and helix−two-turn−helix superfamily.2,4 Despite the structural differences, a majority of DNA glycosylases have one common feature dictated by the reaction chemistry and canonical DNA conformation. Glycosidic bond hydrolysis requires a nucleophilic attack at C1′ of the damaged nucleotide, which can be carried out either by an activated water molecule or by a deprotonated amine in the enzyme’s active site.1,3 However, the C1′ atom is inaccessible in regular B-DNA, thus necessitating some conformational change to expose it to the reacting group.5,6 To achieve this, DNA glycosylases from all three major superfamilies sharply kink DNA at the site of the lesion and extrude the damaged nucleotide into the lesion-binding pocket, stabilizing this highly strained conformation by multiple contacts with the phosphodiester backbone and inserting several protein’s © 2019 American Chemical Society

Received: February 19, 2019 Revised: May 17, 2019 Published: May 23, 2019 2740

DOI: 10.1021/acs.biochem.9b00134 Biochemistry 2019, 58, 2740−2749

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Biochemistry

Figure 1. Fpg−DNA interactions. (A) Scheme of interactions of Fpg with DNA backbone phosphates observed in the crystal of the NaBH4reduced Fpg−DNA conjugate.27 (B) Enlarged view of DNA (shown as sticks) bound in the active center of Fpg (shown as semitransparent cartoons). The phosphoester linkages disrupted in this study are colored; P atoms in the damaged strand are colored dark orange, and those in the complementary strand are colored light orange. The protein’s residues interacting with the backbone phosphates are shown as colored sticks (C, green; N, blue; O, red); dashed lines show the interactions deduced from the crystal structure. The view is oriented for the best presentation of the interactions in the damaged strand. (C) Design of the nicked duplexes used in the experiments. All P−O bonds shown by dashes were disrupted one by one in the set of substrates. The numeration of the damaged strand used in the text starts with oxoG (X0), and its phosphate is accordingly p0. Nucleotides and their phosphates 5′ to X0 have positive numbers, while those 3′ to X0 have negative numbers. The complementary strand is numbered in parentheses so that its phosphates correspond to the phosphates of the damaged strand; hence, the phosphate of C opposite oxoG is p(−1) rather than p(0), etc. The positions where dideoxynucleotides were inserted are doubly underlined. The shaded bars map the footprint of Fpg inferred from the crystal structure;27 the heights of the bars correspond to the areas of the respective nucleotide occluded by Fpg (see the scale to the left). The arrows mark the positions of the breaks studied in ref 29 with their area approximately proportional to the observed effect (note that the sequence used in ref 29 was different from ours).

in the opposite direction.29 Human and yeast 8-oxoguanine− DNA glycosylases OGG1, which are structurally unrelated to Fpg but share its lesion recognition mechanism, show the same tendency.30,31 If the break is in the same strand, OGG1 poorly excises oxoG from substrates with a break between the second and third nucleotides 3′ to oxoG, if an abasic site is at the 3′ side of the break.32 However, placing a break one nucleotide farther in the 3′ direction or at an equidistant position to the 5′ direction from oxoG barely affects the activity.32 Because these studies were addressing the repair of substrates generated from ionizing radiation, their mechanistic utility is limited: the breaks used were in fact gaps or lacked a phosphate or a base, and only a limited number of positions were sampled. To the best of our knowledge, no report in the literature addressed the physical continuity of the phosphodiester backbone as the necessary requisite for efficient catalysis by DNA glycosylases. Here we report that catalysis by Fpg is critically dependent upon the integrity of several P− O bonds in the damaged strand, which likely reflects the need for coordinated conformational change in DNA involving the adjacent groups.

should remove damaged bases from any sequence context, such contacts are expected to occur mostly with the DNA backbone. Indeed, multiple structural and molecular dynamic studies reveal that these enzymes rarely interact with DNA bases except for the excised one and the base opposite to it but form an extensive network of direct and water-mediated bonds with the internucleoside phosphates.7−10,25−28 Thus, one could expect that molecular forces guiding the correct eversion of the damaged nucleotide will be strongly dependent on the continuity of the sugar−phosphate backbone near oxoG in the substrate DNA. Combinations of single-strand breaks and base lesions have mostly been studied in the context of repair of clustered damage, which is predominantly produced by ionizing radiation in aqueous solutions. For Fpg, it has been shown that if the nondamaged strand contains a one-nucleoside gap or a break with the adjacent nucleotide lacking its base, the activity of the enzyme decreases 2−16-fold (in terms of ksp; later in this paper we refer to ksp as the measure of activity unless specified otherwise), when the break is immediately 3′ or 5′ to the base opposite oxoG or three to five nucleotides from oxoG to the 5′ direction of the nondamaged strand (Figure 1C).29 No effect was observed if the break was moved 2741

DOI: 10.1021/acs.biochem.9b00134 Biochemistry 2019, 58, 2740−2749

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Biochemistry Table 1. Kinetic Parameters of Cleavage of Break-Containing Substrates by Fpg strand

broken linka

no break damaged damaged damaged damaged damaged damaged damaged damaged damaged damaged damaged damaged damaged damaged damaged damaged complementary complementary complementary complementary complementary complementary complementary complementary complementary complementary complementary complementary

T −p p+2−T+2 T+2−p+1 ddT+2−p+1 p+1−C+1 C+1−p0 ddC+1−p0 p0−oxoG0 oxoG0−p−1 p−1−C−1 C−1−p−2 ddC−1−p−2 p−2−T−2 T−2−p−3 ddT−2−p−3 p−3−C−3 p(+2)−A(+3) A(+2)−p(+2) p(+1)−A(+2) G(+1)−p(+1) p(0)−G(+1) C(0)−p(0) p(−1)−C(0) G(−1)−p(−1) p(−2)−G(−1) A(−2)−p(−2) p(−3)−A(−2) G(−3)−p(−3) +3

+2

KM (nM) 9.7 7.0 7.5 48

± ± ± ±

1.4 1.3 1.6 5

kcat (min−1) 0.33 1.3 1.4 1.2

± ± ± ±

23 ± 5 14 ± 1

2.8 ± 2.7 ±

8.2 ± 2.1

0.22 ±

11 4.0 12 18 21 14 27 36 30 28 28 12 27 23 17

± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

3 1.0 4 5 5 3 6 11 5 4 6 2 5 8 3

0.88 0.55 0.83 1.3 1.5 0.68 0.84 1.0 0.65 0.80 1.7 0.60 0.83 0.90 1.1

± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.03 0.1 0.1 0.1 n/cb 0.2 0.1 n/cb n/cb n/cb n/cb 0.02 n/cb n/cb 0.06 0.03 0.06 0.1 0.1 0.03 0.06 0.1 0.04 0.04 0.1 0.03 0.05 0.10 0.5

ksp (nM−1 min−1) (3.4 (1.9 (1.9 (2.5

± ± ± ±

0.5) 0.5) 0.5) 0.5)

× × × ×

10−2 10−1 10−1 10−2

(1.2 ± 0.3) × 10−1 (1.9 ± 0.4) × 10−1

(2.7 ± 0.9) × 10−2

(8.0 (1.4 (6.9 (7.2 (7.1 (4.9 (3.1 (2.8 (2.2 (2.9 (6.1 (5.0 (3.1 (3.9 (6.5

± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

2.6) 0.4) 2.6) 2.4) 2.1) 1.3) 0.8) 1.0) 0.5) 0.6) 1.7) 1.2) 0.8) 1.5) 3.3)

× × × × × × × × × × × × × × ×

10−2 10−1 10−2 10−2 10−2 10−2 10−2 10−2 10−2 10−2 10−2 10−2 10−2 10−2 10−2

a

dd, dideoxy end at the break. bNot cleaved.

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MATERIALS AND METHODS Enzymes. Phage T4 polynucleotide kinase and recombinant calf thymus terminal transferase were purchased from New England Biolabs (Beverly, MA) and were used according to the manufacturer’s protocols. Recombinant Escherichia coli Fpg and human OGG1 were expressed and purified as described previously.27,33 The fraction of active Fpg (∼90%) was determined by NaBH4 cross-linking to the reference oxoG:C-containing duplex,34 and the reported enzyme concentrations are those of the active Fpg. Oligonucleotides. The sequences of oligonucleotides used in this work are shown in Table S1. The substrates were designed to contain the 23-mer pyrimidine-rich sequence that was used as a standard for several investigations of the substrate specificity of Fpg9,24,35−39 and additional flanking sequences to stabilize the duplex with breaks. The oligonucleotides were synthesized in house from commercially available phosphoramidites (Glen Research, Sterling, VA) and purified by sequential ion-exchange and reversed-phase chromatography followed by gel electrophoresis in a 20% polyacrylamide/8 M urea mixture. Phosphorylation of 5′ and 3′ termini, when necessary, was done synthetically. OxoGcontaining strands were either 5′-labeled using T4 polynucleotide kinase and [γ-32P]ATP (PerkinElmer, Waltham, MA) if oxoG was located 5′ to the nick in the assembled substrate or 3′-labeled using terminal transferase and [α-32P]cordycepin triphosphate (PerkinElmer) if oxoG was located 3′ to the nick (Figure 1). Oligonucleotides containing a 3′-terminal dideoxy moiety were made from the corresponding primers one

nucleotide shorter using terminal transferase and the required ddNTP (Biosan, Novosibirsk, Russia). The labeled oligonucleotides were annealed with a 2-fold molar excess of the appropriate complementary and coaxial strands to obtain the nicked substrates. Enzyme Kinetics. The reaction mixtures (10 μL) contained 50 mM Tris-HCl (pH 7.5), 100 mM NaCl, 1 mM EDTA, 1 mM dithiothreitol, 0.5−4 nM Fpg, and 5−200 nM DNA substrate. All measurements were performed in the linear regions of the time courses and with enzyme concentration dependencies of the reaction rate (Figure S1). The reactions were initiated by adding Fpg, mixtures incubated at 30 °C for 5 min, and reactions quenched by adding 5 μL of the gel loading dye (20 mM EDTA, 0.1% xylene cyanol, and 0.1% bromophenol blue in deionized formamide) followed by heating for 3 min at 95 °C. Reaction products were separated by gel electrophoresis in a 20% polyacrylamide/8 M urea mixture and quantified by phosphorimaging (Storm 860 Molecular Imager, GE Healthcare, Chicago, IL). The KM and Vmax values were calculated from three to five independent experiments by least-squares nonlinear fitting using SigmaPlot version 9.0 software (Systat Software, Chicago, IL) (Figure S2). Normal Mode Analysis of Vibrational Motions in the Fpg−DNA Complex. The analysis of Fpg−DNA large-scale motions was done using the elastic network approach40,41 using Anisotropic Network Model Web Server version 2.1.42 Structures of Fpg from E. coli [Protein Data Bank (PDB) entry 1K82] and Geobacillus stearothermophilus (PDB entries 1R2Y 2742

DOI: 10.1021/acs.biochem.9b00134 Biochemistry 2019, 58, 2740−2749

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Figure 2. Effect of breaks on Fpg kinetics: (A−C) breaks in the damaged strand and (D−F) breaks in the nondamaged strand. Positions of the breaks are indicated by dashes in the sequences by the axes; the damaged strand is in the 5′-to-3′ direction. (A and D) KM, (B and E) kcat, and (C and F) ksp. “Effect” is the difference between the constants for substrates with and without the break normalized by the smaller of these values; therefore, positive values correspond to an increase in the shown constant, and negative values to a decrease.

and 3GQ3) bound to DNA served as models.17,19,27 The .pdb files were edited to remove cross-links introduced for crystallization. Twenty lowest-frequency modes were calculated with a 15 Å cutoff for interaction between the network nodes. Structural measurements were taken using MDTRA.43

eversion of the damaged base.27,45,46 Thus, it is not surprising that the integrity of at least some links involving p0, p−1, and p−2 is important for the enzyme’s function. Much less expectedly, the disruption of other backbone links, even the second links formed by critically important p0 and p−2, either did not affect Fpg activity or even improved it. Severing the bonds between C−1 and p−2 or T+2 and p+1 had a very minor negative effect on ksp (∼25%), whereas disrupting other links actually increased the activity 2−5-fold. In most cases, this was due to an increased kcat, whereas KM did not change significantly (Table 1). This suggests that in certain positions, breaks in the substrate may relieve conformational strain in the precatalytic complex or better stabilize the transition state rather than affecting the affinity of Fpg for the respective substrate. Discontinuity in the Complementary Strand Has Little Effect on Fpg Activity. To assess the need for backbone integrity of the nondamaged strand for Fpg activity, we have introduced breaks between the nucleotides complementary to the six-nucleotide region around oxoG. Although Fpg buries a longer stretch in the complementary strand compared with the damaged strand (Figure 1), it forms scant contacts with the phosphates in this strand, mostly p(0), p(+1), and p(+2).27 Also, Phe110 is wedged between the orphaned C and the base 5′ to it. In line with fewer direct contacts of the phosphates with Fpg, severing the links in the complementary strand had few consequences for the enzyme activity in terms of ksp, which was in the range of 0.6−2.1 of ksp for the continuous duplex (Table 1 and Figure 2). Interestingly, whereas ksp remained mostly unchanged, both KM and kcat increased in all cases, suggesting that the lower affinity of Fpg for the broken duplex is compensated by a faster cleavage rate. Of all positions, severing the bonds around the orphaned C had the most pronounced

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RESULTS Backbone Links in the Damaged Strand Critical for Cleavage by Fpg. The crystal structures of DNA-bound Fpg from different species reveal most contacts of the protein are formed with the phosphates of the damaged DNA strand.16,27,44 To assess the importance of the backbone integrity for damaged DNA cleavage by Fpg, we have systematically eliminated each phosphoester link from the oxoG-containing strand in six phosphodiester bonds around oxoG (Figure 1) and determined the kinetic constants for such substrates. Importantly, unlike in the previous reports on Fpg acting on oxoG near a break, our substrates still possessed the phosphate at the nick, thus maintaining all of the possibilities of hydrogen and electrostatic bond formation with this moiety. However, the lack of the P−O connection would presumably disturb the synchronous movement of the phosphate group and the adjacent sugar separated by the missing bond. Four of the links were absolutely necessary for Fpg activity. They include the covalent bonds of both 5′ and 3′ hydroxyls of oxoG, the bond between C−1 and p−1 and the bond between T−2 and p−2 (Table 1 and Figure 2). Disruption of any of these links totally abolished the Fpg activity on the respective substrate. In the structure of E. coli Fpg bound to DNA, these phosphates, p0, p−1, and p−2, are the only three phosphates in the damaged strand forming a tight network of bonds with the protein residues.27 Moreover, Fpg and the homologous protein endonuclease VIII pinch the damaged nucleotide, reducing the distance between p0 and p−1 phosphates, to assist in the 2743

DOI: 10.1021/acs.biochem.9b00134 Biochemistry 2019, 58, 2740−2749

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Figure 3. Generation of β-elimination products from substrates with breaks in the nondamaged strand. (A) Gel image showing the migration of cleavage products from several substrates. Arrows show the migration of β-elimination (40-mer substrate without breaks treated by OGG1, lane 8) and β/δ-elimination (40-mer substrate without breaks treated by Fpg, lane 7). Lane 1 contained the untreated substrate. Positions of the breaks in the nondamaged strand (lanes 2−6) are indicated in the sequence inset. (B) Fraction of the β-elimination products generated from the substrates with various positions of the break in the nondamaged strand.

the size marker of β/δ-elimination (Figure 3A). A systematic analysis of product mobility revealed that chain discontinuity near the orphaned C and two adjacent nucleotides caused Fpg to terminate the reaction prematurely, producing a sizable fraction of the β-elimination product (Figure 3B). This fraction sharply dropped when moving outward from the lesion, with breaks disrupting the fourth phosphoester bond from the orphaned C resulting in almost fully β/δ-products. This observed shift toward β-elimination is likely due to formation of a double-strand break after product release and the inability of Fpg to bind or process substrates located near DNA termini. Thus, we have inquired whether the δ-elimination step normally requires substrate release after β-elimination and the second enzyme binding event, as was shown for phage T4 endonuclease V.47 However, with a control substrate, δelimination could not be outcompeted by adding an unlabeled DNA trap 10 s after initiation of the reaction, in line with our earlier finding that the Fpg β-elimination intermediate is not detectable by a quench flow assay.48 We conclude that normally β/δ-elimination in Fpg is concerted but a break in the opposite strand may uncouple these two steps. Elastic Network Analysis of the Fpg−DNA Complex. In an attempt to rationalize the observed effects of damaged strand discontinuity on Fpg activity, we have analyzed vibrational motions in the Fpg−DNA complex using the elastic network method.40,41 In this approach, protein and DNA are represented as sets of point masses (nodes; normally Cα for proteins and P, C2, and C4′ for nucleic acids) connected by springs within some cutoff radius. Fluctuations of the whole system are decomposed into normal modes of increasing frequency; usually, the slowest frequencies reflect global movements of the system, while the high-frequency modes are local and do not propagate through the structure. It is noteworthy that the elastic network approach does not require molecular dynamics but rather relies on a static structure and spring force constants derived empirically from a database of crystallographic B-factors. This instrument is well suited to exploring force propagation in biomolecules and had been used to address the properties of a variety of systems such as cooperativity in protein dimers and tetramers,49,50 signal transduction in ion channels,51 and transcription factors bound to their DNA targets.52 Naked 12-mer DNA (PDB entry 355D53) demonstrated good correlations of vibrational movements propagating along each strand for approximately two or three nucleotides except

suppressing effect, although, in absolute terms, ksp decreased only 1.2−1.6-fold. The Nature of the Break Affects Fpg Activity. Although nicks flanked by a phosphate and a hydroxyl are better suited to studying the effects of covalent strand continuity than onephosphate gaps, our substrates still contain 5′ or 3′ hydroxyls at one side of the nick. This effectively adds a heavy oxygen atom and the possibility of donating or accepting extra hydrogen bonds at the nick. We have tried to eliminate this possibility at four positions (5′ to phosphates p+1, p0, p−2, and p−3) surrounding the damaged site in the cleaved strand, replacing nucleotides bearing a 3′-terminal hydroxyl with their 2′,3′-dideoxy analogues. As 8-oxo-2′,3′-dideoxyGTP is not available, the substitution 5′ to p−1 was not studied. The dideoxy-flanked nicks had more pronounced consequences for enzyme activity than hydroxyl-flanked ones (Table 1). Only in the farthest position, p−3, was the processing of the dideoxy substrate comparable with that of its deoxy counterpart. In three other positions, the dideoxy substitution abolished Fpg activity; it should be noted that even deoxy nicks 5′ to p+1 and p−2 are among the worst substrates in the whole series (Table 1 and Figure 2). On the contrary, a deoxy nick 5′ to p0 was tolerable and enhanced the activity whereas the dideoxy nick at this position was inactivating. In the X-ray structure and molecular dynamics of the E. coli Fpg−DNA complex,27,39 a catalytically important Tyr236 phenyl moiety resides within 3.5 Å of both O3′ and O1P of the p0 phosphate, and its interactions might be disrupted upon a full loss of O3′. Products Generated by Fpg Depend on the Continuity of the Complementary Strand. Normally, Fpg is a bifunctional DNA glycosylase capable of base excision followed by breakage of the sugar−phosphate backbone at the lesion site. Chemically, this breakage comprises two sequential reactions of β-elimination, usually termed β- and δ-elimination (elimination of the phosphate at C3′ and C5′, respectively). The ability to catalyze β/δ-elimination is a characteristic feature of glycosylases belonging to the helix−two-turn−helix superfamily, such as Fpg, endonuclease VIII, and NEIL proteins, whereas many other glycosylases, such as human 8oxoguanine−DNA glycosylase OGG1, can perform only βelimination. While investigating the effect of discontinuity in the complementary strand on Fpg activity, we have noticed anomalous mobility of the bands in some cases relative to 2744

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Figure 4. Heat map of vibrational movement correlations in the Fpg−DNA complex (structure of PDB entry 1K8227). Covariation was calculated over the first 20 normal modes. A dashed line marks the position of the lesion. The blue to red heat scale is for the cross correlation between different residues. (A) Correlations in the whole Fpg−DNA complex: (i) correlations within the DNA duplex, (ii) correlations within the Nterminal domain, (iii) correlations within the C-terminal domain, (iv) cross-correlations between the protein domains, (v) correlations between DNA and the N-terminal domain, and (vi) correlations between DNA and the C-terminal domain. The black arrowhead points to protein peaks (red in v and vi) correlated with the damaged strand, and the gray arrowhead to peaks correlated with the nondamaged strand. (B) Enlarged heat map of correlations in DNA: (i) correlations within the damaged strand, (ii) correlations within the nondamaged strand, and (iii) correlations between the strands. X, pentane-3,4-diol-5-phosphate (the lesion in PDB entry 1K82). (C) Differential DNA heat map, showing the change in correlation due to the presence of the protein (i−iii same as in panel B). (D) Structure of the Fpg−DNA complex (PDB entry 1K82) with the protein regions most strongly correlated with DNA colored red. The segment of the damaged strand most strongly correlated with the protein is colored magenta, and that of the nondamaged strand is colored blue.

in the full complex and the isolated DNA from the complex revealed that the presence of the protein strengthens the alongstrand correlations of the damaged nucleotide with two nucleotides 3′ to it (red spots in Figure 4C) and uncouples the movement 5′ to the site of the lesion. In the opposite strand, the correlations are strengthened 3′ to the orphaned C(0). Several segments of the polypeptide chain showed correlations with the DNA parts around the lesion in both damaged and nondamaged strands (Figure 4A,D). Interestingly, while the vibrations in the catalytic N-terminal domain were correlated with both DNA strands, the C-terminal domain was better correlated with the damaged strand

for the terminal parts (Figure S1). Correlations with the complementary strand were also evident albeit weaker. Introduction of a break in the middle of one strand (PDB entries 1NDN and 1VTE54) caused local decoupling of movements in the broken strand but had little effect on cross-strand correlations (Figure S1). In the Fpg−DNA complex (PDB entry 1K82, E. coli Fpg27), despite the highly distorted DNA structure, the correlations in the nondamaged strand of the DNA molecule were similar to those in free DNA (Figure 4A,B). In the damaged strand, however, the correlations 5′ to the lesion were notably weaker than those 3′ to the lesion (Figure 4B). Comparison of vibrational modes 2745

DOI: 10.1021/acs.biochem.9b00134 Biochemistry 2019, 58, 2740−2749

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p0 and p−1.25,26,28,67 Presumably, just the presence of a phosphate moiety at p0 and p−1 is not enough for proper nucleotide extrusion or positioning within the active site, and mechanical forces acting via the unbroken bond are required. In addition to positioning the nucleotide in the active site, disconnection of the adjacent phosphate may affect the sugar pucker, which is suggested to influence damage recognition and catalysis by Fpg.19,68,69 Another factor possibly affecting the interactions at the break is the effective negative charge at the phosphate monoester, because internucleoside phosphates have the total charge of 1.29 e, whereas the charge at a terminal phosphate is 1.98 e.70 By itself, a negative charge close to C1′ seems to assist rather than suppress the N-glycosidic bond breakage.1,3 However, the larger negative charge could strengthen the interactions of the phosphate with the positively charged Lys56 (phosphates p−1 and p−2) and Arg258 [p0 and p−1 (Figure 1A)] and interfere with their precise dynamics; Arg258 was shown to be important for oxoG eversion stabilizing the extrahelical nucleotide during the late stages.9 The p−2 phosphate also interacts with His70, a part of a “saddle motif” suggested to dynamically bend DNA during scanning in search of a lesion.24,37 In addition to the disruption of mechanical coupling, a nick could interfere with productive enzyme binding, either decreasing the general affinity of Fpg for nicked DNA or competing with oxoG as a binding site. Because oxoG is rapidly degraded by Fpg, the affinity for oligonucleotides containing a tetrahydrofuran residue, a noncleavable analogue of the AP site, is often measured in the literature as a proxy for Fpg affinity for damaged DNA.71−73 However, structural, modeling, and kinetic data suggest that abasic sites in DNA are partially extrahelical and that they do not need to proceed through the same eversion path as oxoG does,24,74,75 thus making general affinity for tetrahydrofuran ligands hard to interpret if eversion-related processes are involved. Although the affinity of Fpg for nicked DNA has not been measured, it should not be much lower than its affinity for undamaged DNA (∼250 nM vs ∼10 nM for damaged DNA36), singlestranded damaged DNA, or the reaction product, a singlenucleoside gap without the lesion (first micromolar).65,76,77 A single-nucleoside gap in the opposite strand one position 5′ or 3′ to the lesion was reported to decrease Fpg binding by an order of magnitude,29 which may be an upper estimate of the binding effect for our substrates with all phosphates in place. Overall, the affinity factor may contribute to the observed KM effects but is unlikely to be the primary reason for the apparent inability of Fpg to cleave some nicked substrates. Dideoxy-flanked 5′ breaks in the damaged strand were even more prohibitive for the reaction and seemed to behave more like 3′ breaks at the respective phosphate, although the number of sampled positions was limited. Studies with DNA polymerases revealed that, despite being very close in the conformation to their 2′-deoxy counterparts,78 2′,3′-dideoxynucleotides induce notable readjustment of the enzyme’s active site, which translates as a ≤3 order of magnitude loss of catalytic efficiency.79,80 In the hydroxyl- and phosphate-flanked break, there is a possibility of hydrogen bonding between these moieties, which could partially compensate for the loss of a covalent bond. The lack of this possibility in dideoxy-flanked breaks might account for their more pronounced effects. The lack of detrimental consequences of breaks in other positions, including everywhere in the nondamaged strand, came as a surprise, especially considering that phosphate- or

(marked by arrowheads in Figure 4A). Structures of Fpg from G. stearothermophilus bound to DNA with intra- or extrahelical oxoG (PDB entries 3GQ319 and 1R2Y17) showed vibrational modes closely resembling those of E. coli Fpg (Figure S2).

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DISCUSSION DNA glycosylases must efficiently discriminate their cognate bases from a vast excess of normal DNA. Many years of kinetic, structural, and computational studies of DNA glycosylases significantly advanced our understanding of the specificity and catalytic mechanism of these enzymes (reviewed in refs 1, 3, and 55−58). A key common factor emerging from these studies is the existence of multiple precatalytic transient conformers of the enzyme−DNA complex, which efficiently channel the damaged base into the active site while rejecting the normal ones or diverting them to dead-end catalytically incompetent conformations. The highly and precisely distorted DNA structure in the precatalytic complex also seems to be very important for the following electrostatic stabilization of the transition state in the glycosidic bond breakage step.59−61 An exception is presented by the enzymes belonging to the HEAT repeat family (AlkC and AlkD) as well as phage T4 endonuclease V, which access the damaged nucleotide without its eversion,62−64 but they represent only a minor fraction of DNA glycosylases. We reasoned that this importance of conformational changes in DNA should dictate the necessity of its backbone continuity, at least in certain critical positions, for the efficient substrate processing by DNA glycosylases. In particular, we addressed these requirements for Fpg, a DNA glycosylase that excises 8oxoguanine passing through several structurally and kinetically distinguishable precatalytic intermediates.9,16−24,34,48,65,66 Unlike previous studies of the removal of oxoG clustered with DNA breaks by Fpg29 and several other DNA glycosylases,29−32 we employed a system in which only the covalent linkage along the DNA backbone was interrupted at any time but all nucleosides and phosphates were in place, thus excluding the possible effects due to a loss of interactions with these moieties. In the positions that could be directly compared with the results of ref 29, keeping these interactions was enough to maintain Fpg activity and even improve it. Most notably, breaks at positions (+1) and (−1) in ref 29 caused, in terms of ksp, a 2−16-fold inhibition of Fpg depending on the nature of the break (products of abasic site hydrolysis or β- or β,δ-elimination), while clean nicks around the G(+1) and G(−1) in our experiments had almost no effect on ksp. However, the general tendency of better cleavage of substrates with opposite strand breaks farther from the lesion, observed in ref 29, also showed up in our results, albeit at a lower magnitude (Figure 2F). The effect of nicks in the damage-containing strand on Fpg activity has not previously been investigated. The integrity of four links in the damaged strand, namely, p0−X0, X0−p−1, p−1−C−1, and p−2−T−2, proved to be essential for Fpg activity. In line with the biochemical observations, elastic network analysis also showed that Fpg binding strengthens the mechanical coupling of X0, C−1, and T−2 nucleotides. Structures of Fpg reveal that p 0 and p −1 phosphates are closer to each other than in regular B-DNA and are held in place by multiple interactions with the protein residues. DNA glycosylases from other structural families, such as uracil−DNA glycosylase, MutY, OGG1, and endonuclease III, extrude the damaged nucleotides using a similar backbone compression mechanism but pinching p+1 and p−1 rather than 2746

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base-lacking breaks in several positions in the nondamaged strand were reported to suppress Fpg activity several-fold.29 Backbone compression around C(0) is also evident in Fpg structures. However, the contacts of the protein with the phosphates in the nondamaged strand are much scarcer, and the solvent-occluded surface is smaller (Figure 1). The energy of interaction of the nondamaged strand with Fpg is ∼2.5-fold smaller in absolute value than the binding energy for the damaged strand.77 It is quite possible that the conformational changes in the nondamaged strand follow those in the damaged strand rather than being actively induced by the protein. In the damaged strand, breaks 5′ to the lesion, if not detrimental, generally improved the activity while those in the permissive positions 3′ to the lesion had marginal effects. Interestingly, the area 5′ to the lesion in the damaged strand, even under the protein’s footprint, showed less correlation in the vibrational modes in the elastic network model compared to the nucleotides 3′ to the lesion, where the protein forms more contacts with DNA (Figure 4B), suggesting that this part of the duplex is partially mechanically decoupled from the rest of the Fpg−DNA complex. Perhaps breaks do not significantly affect the already less-coordinated movements in the 5′ part of the damaged strand. While the conformational changes in DNA upon binding many DNA glycosylases, as well as DNA methyltransferases, oxidative demethylases, and methylated base readers, are firmly established6,81 and highly detailed nucleotide extrusion models exist, the contribution of different energy terms to the transition between conformers has received very little attention. In free DNA, the decomposition of energy suggests that the stress built up in the backbone upon bending is released through base pair opening, suggesting force propagation along the covalent contour.82 In a mechanistic study of uracil−DNA glycosylase, which has another fold but extrudes the lesion in a manner similar to that of Fpg, it was found that the enzyme channels the energy from binding p−2 to push rigid cyclic sugars of +1 and −1 nucleotides into positions appropriate for catalysis with a p+1−p−1 pinch.83 Our results add weight to a view of DNA glycosylase action in which mechanical coupling within the DNA substrate is crucial for the induced fit. Although highly likely, how common the requirement is for backbone continuity in other glycosylases remains to be seen.

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This research was supported by the Russian Science Foundation (17-14-01190, the biochemical part) and the Russian Foundation for Basic Research (17-04-01761-a, the modeling part). Partial salary support from the Russian Ministry of Science and Education (AAAA-A17117020210023-1 for A.V.E. and D.O.Z. and 6.5773.2017/6.7 for D.O.Z.) is acknowledged. Notes

The authors declare no competing financial interest.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.biochem.9b00134. A table with sequences of oligonucleotides used in this work and four figures with additional data on enzyme kinetics and results of elastic network analysis of free DNA and Fpg from G. stearothermophilus (PDF) Accession Codes

Fpg, P05523.

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REFERENCES

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Telephone: +7(383) 3635187. Fax: +7(383)3635153. ORCID

Dmitry O. Zharkov: 0000-0001-5013-0194 2747

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