Journal List J Biol Chem 2018 May 4 PMC5936803 J Biol Chem. 2018 May 4; 29318 6824–6843. Andrey G. Baranovskiy,‡,1 Vincent N. Duong,§,1 Nigar D. Babayeva,‡ Yinbo Zhang,‡,2 Youri I. Pavlov,‡¶ Karen S. Anderson,§,3 and Tahir H. Tahirov‡,4AbstractDNA polymerase α Polα plays an important role in genome replication. In a complex with primase, Polα synthesizes chimeric RNA–DNA primers necessary for replication of both chromosomal DNA strands. During RNA primer extension with deoxyribonucleotides, Polα needs to use double-stranded helical substrates having different structures. Here, we provide a detailed structure–function analysis of human Polα's interaction with dNTPs and DNA templates primed with RNA, chimeric RNA–DNA, or DNA. We report the crystal structures of two ternary complexes of the Polα catalytic domain containing dCTP, a DNA template, and either a DNA or an RNA primer. Unexpectedly, in the ternary complex with a DNADNA duplex and dCTP, the “fingers” subdomain of Polα is in the open conformation. Polα induces conformational changes in the DNA and hybrid duplexes to produce the universal double helix form. Pre-steady-state kinetic studies indicated for both duplex types that chemical catalysis rather than product release is the rate-limiting step. Moreover, human Polα extended DNA primers with higher efficiency but lower processivity than it did with RNA and chimeric primers. Polα has a substantial propensity to make errors during DNA synthesis, and we observed that its fidelity depends on the type of sugar at the primer 3′-end. A detailed structural comparison of Polα with other replicative DNA polymerases disclosed common features and some differences, which may reflect the specialization of each polymerase in genome DNA replication, DNA polymerase, human, crystal structure, DNA-protein interaction, RNA, conformational change, chimeric RNA-DNA primer, Polα, Polδ, Polε, pre-steady-state kineticsIntroductionThree DNA polymerases are required for genome replication in eukaryotes DNA polymerase α Polα,5 Polϵ, and Polδ 1. All of them belong to the B-family and fulfill the different tasks. Polϵ synthesizes most of the leading strand, Polδ is mainly involved in synthesis of the lagging strand, and Polα generates the primers for both polymerases 2, 3. Polα alone cannot synthesize DNA primers de novo and relies on RNA primers created by primase. Polα works in a tight complex with primase, called the primosome 4, 5. Synthesis of the chimeric RNA–DNA primers by the primosome is highly coordinated by autoregulation through the alternating activation/inhibition of two catalytic centers, which is mediated by the C-terminal domain of the primase accessory subunit 6. Relatively low fidelity of Polα, which does not possess a proofreading activity, results in mutational hot spots predominantly on the lagging strand 7. In addition to the established role of primosome in nuclear replication, it is involved in formation of hybrid DNARNA duplexes in the cytosol, which are important for regulation of the type I interferon response 8. Polα is a direct target of an anti-tumor toxin CD437, an attractive anti-cancer lead molecule, which induces apoptosis selectively in cancer cells 9.Human Polα hPolα is composed of two polypeptides the catalytic subunit p180 and the accessory B-subunit p70, with calculated molecular masses of 166 and 66 kDa, respectively. p180 contains two domains, the catalytic residues 338–1250 and the C-terminal Polα CTD, residues 1266–1462 domains, which are flexibly connected by a 15-residue-long linker 4. The catalytic domain possesses DNA-polymerizing activity but has no proofreading exonuclease activity, in contrast to other replicative DNA Pols, δ and ϵ 10. Polα CTD connects the catalytic domain with p70 and primase and contains two conserved zinc-binding modules, where each zinc ion is coordinated by four cysteines 6, 11, 12. The N terminus of p180 residues 1–337 is predicted to be poorly folded and does not participate in primer synthesis. The structural information for this region is limited to a small peptide in the catalytic subunit of yeast Polα yPolα; residues 140–147 that interacts with the replisome 13.In this work, we use the structural and kinetic approaches to analyze hPolα interaction with the templateprimer and dNTP and the effect of the primer structure on hPolα catalysis, processivity, and structure of the catalytic domain of human DNA polymerase α in complex with a DNA template, RNA, or DNA primer, incoming dCTP, and divalent metal ionsThe structures of the ternary complexes of p180core the part of hPolα containing the catalytic domain, residues 335–1257 with dCTP and DNARNA or DNADNA duplexes have been determined at and Å resolution, respectively Table 1. The p180core adopts the universal “right-hand” DNA polymerase fold consisting of five subdomains 14, which encircle the active site upper schematic in Fig. 1A N-terminal residues 338–534 and 761–808, exonuclease Exo; residues 535–760; inactive, palm residues 834–908 and 968–1076, fingers residues 909–967, and thumb residues 1077–1250 domains. The palm is the most conserved subdomain in DNA polymerases and responsible for catalytically proficient positioning in the active site of two substrates to be polymerized a templateprimer and incoming dNTP. The palm harbors two catalytic aspartates Asp860 and Asp1004, which coordinate two divalent metal ions important for dNTP binding and nucleophilic attack on its α-phosphate by the primer's 3′-OH 15. The thumb makes additional contacts with a templateprimer, and their pattern significantly differs between B-family Pols see below. The L-shaped thumb has two parts the all-helical base adjacent to the palm and the tip with mixed α/β-fold residues 1135–1207, which packs against the Exo. Polα lost proofreading exonuclease activity by substitution of the catalytic residues in the Exo subdomain during evolution 10. The fingers are composed of two antiparallel α-helices Fig. 1A, which make additional contacts with the triphosphate moiety of dNTP and oscillate between the open and closed conformations. The thumb encloses the circle of p180core subdomains around the active site by connecting the palm with Exo Fig. 1A. The interactions at the interfaces of the thumb with the palm and Exo have a lower amount of hydrophobic contacts in Polα compared with Polδ and Polϵ, which allows the thumb to rotate relative to the palm. Such rotation of the thumb away from Exo was observed in apo-forms of the human primosome and yPolα 6, 16.Table 1Data collection and refinement statisticsp180core/DNARNA/dCTP PDB code 4QCLp180core/DNADNA/dCTP PDB code 6AS7Data collection Space groupP 3221P 42212 Cell dimensions a = b, c Å Resolution Å40– 3– I/ Completeness %91 81 Unique reflections95,874 419628,565 1406 Resolution Å No. of reflections Working set88,726 12,00226,970 4410 Test set4683 6431380 209 Rwork/ No. of atoms Protein69476996 DNA and RNA496487 Ligands/ions48/528/4 Water4141 Mean B-factors Å2 RMSDs Bond lengths Å Bond angles degrees Ramachandran plot % Generously complexes of p180core containing dCTP and DNADNA or DNARNA duplexes. A, overall view of the p180core/DNARNA/dCTP complex. The carbons of dCTP, DNA template, and RNA primer are colored yellow, marine, and purple, respectively. B, alignment of p180core ternary complexes containing RNA or DNA primers PDB codes 4QCL and 6AS7, respectively. p180core, templateprimer, and dCTP are colored gray in the complex containing DNADNA. In the bottom panel, DNADNA from the binary complex PDB code 5IUD, which is superimposed onto p180core/DNARNA/dCTP with RMSD of Å for 762 Cα atoms, is colored orange for clarity, double helices are shown separately from p180core.The alignment of hPolα ternary complexes containing DNADNA and DNARNA duplexes with a root mean square deviation RMSD of Å for 648 Cα atoms shows good superposition for the first four bp from the primer 3′-end and for p180core subdomains except the thumb and fingers Fig. 1B. It is likely that the crystal packing affected the conformation of these subdomains in the hPolα/DNADNA/dCTP complex. The thumb interacts with the palm and the N-terminal subdomain of one neighboring molecule and with Exo of the other neighboring molecule. The fingers are stabilized in the open conformation due to interaction with the N-terminal and Exo subdomains of the neighboring molecule. Significant difference in the position of the templateprimer for bp 5–11 from the primer 3′-end the bottom part of Fig. 1B is due to steric hindrance with the thumb tip, which moves closer to the template upon the outward rotation of the thumb. In addition, the blunt end of the DNADNA duplex is packed against the N-terminal domain of the other molecule. In support of the idea that crystal packing affected the conformation of the most flexible subdomains in the structure of p180core/DNADNA/dCTP, the thumb of hPolα in the binary complex with DNADNA has a very similar position as in the hPolα/DNARNA/dCTP complex alignment was provided in Ref. 17. Moreover, DNADNA from the binary complex superimposes well with DNARNA from the ternary complex the bottom part of Fig. 1B. Accordingly, hPolα has similar affinity for DNA and hybrid duplexes 17.In the ternary complex containing DNADNA and dCTP, the fingers have the open conformation as in the binary complex hPolα/DNADNA Fig. 2A and in the complex hPolα/DNARNA/aphidicolin Fig. 2B. In all structures of yeast and human Polα, only two conformations of fingers were observed open and closed 16,–18. This indicates that intermediate positions of fingers are energetically much less favorable. Upon closing, one of the fingers α-helices slides along the N-terminal subdomain toward to Exo Fig. 2C. Analysis of intersubdomain interfaces in the open and closed forms revealed the switching of Ile944 of the fingers between two adjacent pockets. One pocket is formed by Asn760, Pro763, Leu764, and Gln767 of the N-terminal subdomain. The other is formed by Leu764 of the N-terminal subdomain and by Leu700, Ile701, and Leu759 of Exo. The open and closed conformations are also stabilized by a hydrogen bond between the side chain of Gln941 and the main-chain oxygens of Leu570 and Cys757, of the fingers in hPolα. A, the open-fingers conformation in the aligned ternary hPolα/DNADNA/dCTP and binary hPolα/DNADNA complexes PDB codes 6AS7 and 5IUD, respectively. B, the open-fingers conformation in the aligned complexes hPolα/DNADNA/dCTP and hPolα/DNARNA/aphidicolin PDB codes 6AS7 and 5Q4V, respectively. C, switching of Ile944 between two sites upon fingers moving. In the aligned ternary complexes, the subdomains of hPolα in the complex with DNARNA and DNADNA PDB codes 4QCL and 6AS7, respectively are colored according to Fig. 1 and gray, respectively. Ile944 and the residues interacting with it in different finger conformations are shown as sticks. Leu764 interacts with Ile944 in either conformation and is shown on both aligned molecules. The arrow depicts the distance between Cα atoms of Leu934, located in the loop connecting two helices of the fingers. The substrates and aphidicolin are not shown for the structures of both ternary complexes, all three disordered regions residues 674–677, 810–833, and 883–895 are situated near the templateprimer. The loop 878–903 originates from the same site of the palm as the processivity domain P-domain, which is found in Polϵ and provides additional contacts with a templateprimer Fig. 3. The linker 808–837 between the N-terminal and palm subdomains of p180core corresponds to the shorter linker in Polϵ, which adds two β-strands to the stem of the P-domain. In Polδ, the corresponding region is also shorter than in Polα and well-structured in the crystal shown in blue in Fig. 3.Analysis of two disordered regions in p180core. Ternary complexes of yPolδ and yPolϵ PDB codes 3IAY and 4M8O, respectively were superimposed onto p180core/DNARNA/dCTP with RMSD of and Å for 524 and 483 Cα atoms, respectively. Structured p180core regions are colored according to Fig. 1; unstructured regions are depicted by dashed gray lines. The yPolϵ regions 537–557 and 677–764 form the of hPolα with dNTPThe dNTP-binding site of B-family Pols is located at the junction between the palm and the fingers Fig. 4A. Upon dNTP binding, the latter subdomain is able to change its conformation from open to the closed one, which results in stabilization of the precatalytic ternary complex Pol/template primer/dNTP. Despite the open conformation of fingers in the complex of hPolα with a DNA duplex, dCTP occupies the dNTP-binding site Fig. 4A. To our knowledge, this is the first structure of the ternary complex of a B-family DNA polymerase with a DNA duplex and dNTP where the fingers are open. This structure catches the transient step of dNTP binding followed by fingers closing and phosphodiester bond formation. It revealed that fingers interact with dNTP even in the open conformation. Asn954, which establishes the water-mediated contact with the β-phosphate in the closed-fingers conformation, directly interacts with this phosphate in the open-fingers state Table 2. Interaction of Lys950 with dCTP switched from the α- to γ-phosphate upon fingers opening. Interestingly, in the two reported structures of Polα in the complex with DNADNA in all cases, dNTP was added to the crystallization mix, the fingers are open this work; also, see Ref. 17, where dCTP was not detected in the active site. This may indicate a weak stabilization of Polα fingers in the closed conformation. Indeed, analysis of dNTP-finger interactions in replicative DNA Pols shows that two conservative residues of hPolα, Lys926 and Asn954, do not directly interact with the triphosphate Table 2. In yPolδ, Asn705 makes an H-bond with the β-phosphate. In yPolϵ, both water-mediated contacts observed in the hPolα ternary complex are changed to direct interactions. Thereby, only two residues of Polα fingers Arg922 and Lys950 in hPolα directly interact with dNTP in the closed state, whereas yeast Polδ and -ϵ use three and four residues, of Polα with incoming dNTP. A, hydrophilic interactions between hPolα and the triphosphate moiety of dNTP. In aligned ternary complexes containing DNARNA and DNADNA duplexes PDB codes 4QCL and 6AS7, respectively, there is a good superposition of dCTP-interacting residues except the fingers. The subdomains of hPolα in the complex with DNARNA are colored according to Fig. 1. Polα in the complex with DNADNA is colored gray only α-helices of fingers are shown for clarity; dCTP from this complex is shown as lines and colored dark brown. In both complexes, the side or main chains of the hPolα residues involved in hydrophilic interactions with dCTP and metals are shown as sticks. Potential H-bonds between Polα in complex with DNARNA, dCTP, and catalytic metals are depicted by pink dashed lines. The A and B metal-binding sites of hPolα in complex with DNARNA or DNADNA contain Zn2+ and Mg2+ or only Mg2+, respectively. Asp860 and Asp1004 each interact with both divalent metals; the metal-B interacts with all phosphates of dNTP, whereas the metal-A makes a contact only with the α-phosphate. B, comparison of dNTP-binding pockets in hPolα and yPolδ. The ternary complexes hPolα/DNARNA/dCTP and yPolδ/DNADNA/dCTP PDB codes 4QCL and 3IAY, respectively were aligned with RMSD of Å for 524 Cα atoms. The carbons of hPolα and yPolδ are colored according to Fig. 1 and gray, respectively. hPolα Phe861 and yPolδ Phe609, as well as two water molecules coordinated by zinc are not shown for clarity. dCTP from Polδ complex is shown as lines and colored dark brown. Atoms of magnesium, calcium, and zinc are presented as spheres at reduced scale and colored green, light orange, and slate, respectively. A different conformation of Asp608 in yPolδ is probably due to the coordination of a third calcium ion. Accordingly, Asp860 of hPolα is oriented similarly to Asp640 in yPolϵ where Mg2+ occupies the site B 22. In both panels, DNADNA and DNARNA duplexes are not shown for 2Amino acids of replicative DNA Pols making hydrophilic contacts with incoming dNTPThe residues that coordinate the divalent metals in the catalytic site are not listed. The residues marked by an asterisk use a main-chain nitrogen to establish an H-bond with the triphosphate. The residues making a water-mediated contact with the triphosphate are italicized. Residues shown in parentheses do not interact with dNTP in the specified complexFinger conformationPDB codeResidues of the palmResidues of the fingershPolα/DNARNA/dCTPClosed4QCLSer863*Leu864*Arg922Lys926Lys950Asn954hPolα/DNADNA/dCTPaOpen6AS7Ser863*Leu864*Arg922Lys926Lys950Asn954yPolα/DNARNA/dGTPbClosed4FYDSer867*Leu868*Arg917Lys921Lys944Asn948yPolδ/DNADNA/dCTPClosed3IAYSer611*Leu612*Arg674Lys678Lys701Asn705yPolϵ/DNADNA/dATPClosed4M8OSer643*Met644*Arg781Lys785Lys824Asn828In hPolα, the sugar and the base of dNTP fit into the hydrophobic pocket formed by Leu864, Tyr865, Pro866, Tyr957, and Thr1003 Fig. 4B. The conserved Tyr865 defines a steric gate for rNTPs due to steric hindrance with the ribose 2′-OH, which is facilitated by a C3′-endo pucker. Upon fingers closing, the base of dCTP is fixed between Asn954 and the 3′-nucleotide of the primer in the position that is optimal for pairing with guanine from the DNA template. The triphosphate of dCTP is secured in the active site by 11 bonds Fig. 4A and Table 2, including two H-bonds with the main-chain nitrogens of Ser863 and Leu864, four H-bonds with the side chains of Arg922 and Lys950, and five coordinate bonds with two divalent ions, which in turn are coordinated by Asp860, Asp1004, and the main-chain oxygen of Phe861. hPolα also makes two water-mediated contacts with the phosphates of dCTP, using the side chains of Lys926 and Asn954. The presence of Zn2+ at the metal-binding site A in the hPolα/DNARNA/dCTP complex is probably due to the missing hydroxyl at the primer 3′-end, which disrupts the octahedral coordination of Mg2+. In contrast to Mg2+, Zn2+ is flexible with respect to the number of ligands that it can adopt in its first coordination shell 19. Moreover, Zn2+ forms a stronger complex with carboxylates and the triphosphate group of nucleotides than Mg2+ 20, 21. Accordingly, Mg2+ was missing at the site A in the crystal structure of yPolϵ ternary complex, where the primer 3′-end also contained dideoxycytidine 22. Alignment of the hPolα/DNARNA/dCTP complex with the ternary complexes of hPolα and yPolδ containing a DNA primer shows good superposition for incoming dCTP and the residues coordinating dNTP and the catalytic metals Fig. 4. This indicates a high structural conservation of dNTP-binding pocket in replicative Pols and that the type of the primer does not affect of the conserved fingers residues, Arg922 and Lys950, which interact with dNTP, dramatically reduced hPolα activity in the presence of Mg2+ Fig. 5. Consistent with that, mutation of Lys944 in yPolα is lethal 23. Substitution of Asn954, which makes a water-mediated contact with the triphosphate in the closed conformation, resulted in a modest effect on DNA polymerase activity. Kinetic studies have demonstrated that changes of Lys950 primarily affect affinity for the incoming dNTP and the processivity of DNA synthesis, whereas their effect on catalysis was much less 24. Interestingly, activity of p180core mutants R922Q and K950S was almost completely restored in the presence of manganese, whereas the activity of the WT protein was weakly affected Fig. 5. These data are consistent with studies of metal dependence of the activity of entire hPolα and its mutants with different substitutions of Lys950 24. Manganese provides a tighter complex with dNTP versus magnesium 25, allowing rescue of the compromised interaction of dNTP with the fingers in these mutants. The restoration of DNA-polymerizing activity of the finger mutants in the presence of Mn2+ explains the well-known mutagenic effect of this metal, which reduces Polα fidelity by 2 orders of magnitude 24, 25. Briefly, Mn2+-mediated binding of dNTP becomes less dependent on finger closure, an important selectivity step preventing effect of finger mutations on hPolα activity depends on the divalent metal. Extension of dA15 primer annealed to dT70 was carried out in the presence of 30 nm p180core, μm templateprimer, mm dATP, and specified divalent metal ions. The dashed line indicates splicing of the original of human Polα with DNADNA and RNADNA double helicesLike other B-family DNA polymerases, Polα makes continuous contact with the minor groove of the double helix of a templateprimer 16, 22, 26, 27. The palm interacts with phosphates of P1, P2, and T2–T4 Fig. 6B, which provides the optimal positioning of the primer 3′-end for catalysis. The flexible thumb secures the polymerase grip on the double helix by interacting with phosphates of T7, T9, and P3–P5. Two thumb residues, Arg1082 and Thr1140, make double contacts with primer phosphates, using both the side- and main-chain atoms. Each of the N-terminal and Exo subdomains make only one contact with a templateprimer. Interestingly, the position of T−1 phosphate is not fixed in Polα Fig. 6, A–D, whereas Polδ and Polϵ make a number of H-bonds with this phosphate Fig. 6, E and F. In both ternary complexes of hPolα, the side chain of Arg834 extends into the major groove and packs against the base of T3. On the other side of the double helix, the side chains of conserved Arg1081 and Lys1053 protrude into the minor groove and together with Arg834 serve as rails for templateprimer diagrams of the interactions of eukaryotic replicative Pols with the sugar-phosphate backbone of a templateprimer. The crystal structures of the following complexes were analyzed hPolα/DNADNA/dCTP A, hPolα/DNARNA/dCTP B, hPolα/DNADNA C, yPolα/DNARNA/dGTP D, yPolδ/DNADNA/dCTP E, and yPolϵ/DNADNA/dATP F. Among residues making van der Waals interactions with a templateprimer, only the arginines contacting the P4 sugar are shown for clarity. Residues highlighted in italic type can make a hydrogen bond with a templateprimer in their alternative conformation. Ribose, deoxyribose, and dideoxyribose rings are colored red, gray, and olive, respectively. The residues interacting with a templateprimer are color-coded according to the subdomains they belong to see Fig. 1 for color-coding. The conservative amino acids are underlined. The residues marked by an asterisk use a main-chain atom to establish H-bond with a sugar-phosphate backbone. The pound sign indicates that the residue makes double contact with a templateprimer, using both the side- and main-chain atoms. The type of the double helix A, B, or intermediate AB was determined for each dinucleotide step using Web 3DNA software Rutgers University 69. This software requires the presence of two consecutive dinucleotide steps of the same type to confidently determine the helix type, so the third dinucleotide step of DNARNA was downgraded by the program to unclassified. We assume that in locally distorted double helices, the presence of a single isolated step may take place, which is in line with data on yPolα 16. In the p180core/DNARNA/aphidicolin complex PDB code 4Q5V, the pattern of interactions between hPolα and the templateprimer is the same as shown in B except for the loss of the H-bond between T−1 and reduced number of hPolα-DNA contacts in the hPolα/DNADNA/dCTP complex Fig. 6A is probably due to the crystal packing, which affected the thumb position see above. This explanation is supported by the crystal structure of the hPolα/DNADNA binary complex showing the same network of DNA-protein interactions as in the hPolα/DNARNA/dCTP ternary complex Fig. 6, B and C, with the additional contact between the P1 phosphate and Tyr1055. The binding constants for p180core complexes with both duplexes are comparable Kd = 58 ± 2 nm for DNADNA and Kd = 101 ± 6 nm for DNARNA; Table 3; the former value is in good agreement with a Kd of 44 nm for the hPolα/DNADNA complex reported by Copeland et al. 25. The discrepancy with data reported by Coloma et al. 17 Table 3 might be due to different methods and experimental settings. Regardless, in both cases, the DNA and hybrid duplexes were compared at the same conditions, and their binding characteristics are 3Affinity of hPolα for DNADNA and DNARNADuplexKdThis studyRef. 17Ref. 25nmDNADNA58 ± 243044DNARNA101 ± 6320As was observed in yeast Pols α, δ, and ϵ Fig. 6, D–F, the invariant arginines 1081 and 1082 of hPolα trap the sugar-phosphate backbone of the primer near P4. The side chain of Arg1081 extends into the minor groove and packs its guanidinium moiety against the sugar Figs. 6 A–C and ​7A. Such a position of Arg1081 is fixed by two H-bonds with Asp1083. The side chain of Arg1082 is located between the phosphates of P3 and P4 and can interact with any of them. Binding of a hybrid duplex results in potential steric hindrance between Arg1081 and the 2′-OH of the P4 ribose Fig. 7A and in a local change of RNA conformation Fig. 7B, which includes the unusual syn-conformation of P4 torsion angle χ = − 2′-endo pucker of its ribose, and unstacking between the bases of P4 and P5. This bending of the RNA primer was also observed in the yPolα/DNARNA/dGTP complex 16. It is notable that the orientation of the P3 phosphate in the RNA primer is more favorable for hydrogen bonding with Arg1082 and Arg702, compared with the P3 phosphate of the DNA primer. The same position of Arg1081 analog in ternary complexes of yeast Pols δ and ϵ indicates a common mechanism of RNA sensing in replicative DNA primer bending. A, primer conformation in the complexes of hPolα with DNARNA and DNADNA. The p180core/DNARNA/dCTP and p180core/DNADNA complexes PDB codes 4QCL and 5IUD, respectively were aligned with RMSD of Å using the palm and thumb. In the complex containing DNARNA, RNA primer and Arg1082 are colored according to Fig. 1. In the complex containing DNADNA, the carbons of a DNA primer and amino acids are colored gray and green, respectively. In the complexes containing RNA or DNA primer, amino acids are represented as sticks or lines, respectively. The modeled ribose with the 3′-endo pucker was superimposed on the P4 sugar of the DNA primer and has the same position of carbons colored yellow except C2. The arrow depicts the difference in position of P4 2′-OH in the RNA primer and the modeled ribose. Pink dashed lines and double arrows depict H-bonds and distances between atoms, respectively. B, overall view of RNA primer in the complex p180core/DNARNA/ three reported complexes of Polα with a hybrid duplex 16, 18, the conformation of the P1 phosphate is different compared with the complexes of hPolα, yPolδ, and yPolϵ with a DNA duplex Fig. 8. The conformation of the P1 phosphate was not affected by the addition of two deoxyribonucleotides to the 3′-end of the RNA primer, as seen in the yPolα ternary complex Fig. 8B, which indicates that it depends on primer bending due to the presence of ribose at P4 Fig. 7. Thus, accommodation of RNA and chimeric primers is accompanied by dislocation of the P1 phosphorus, which results in disruption of the H-bond between this phosphate and the conserved Tyr1055 Fig. 6B; this H-bond is replaced by the weaker water-mediated interaction in the case of a DNA duplex.Conformation of the P1 phosphate in the DNA and chimeric RNA–DNA primers. A, conformation of P1-P2 in the complexes of hPolα with DNA and chimeric primers. The p180core/DNARNA/dCTP and p180core/DNADNA complexes PDB codes 4QCL and 5IUD, respectively were aligned with RMSD of Å using the palm. In the former complex, the RNA primer may be considered chimeric due to the presence of dideoxycytidine at the 3′-end. The carbons are colored gray in the complex of p180core with DNADNA. B, conformation of P1-P2 in the complexes of yPolδ and yPolα with DNA and chimeric primers, respectively. The ternary complexes of yPolα and yPolδ PDB codes 4FYD and 3IAY, respectively were aligned with RMSD of Å using the palm. The carbons are colored gray in the complex of yPolδ with DNADNA. Blue dashed lines depict the does not exhibit typical polymerase burst kineticsA pre-steady-state burst experiment was performed to provide insight into the relative rates of DNA polymerization steps in the kinetic mechanism of Polα. It has been frequently observed that under burst conditions slight excess of templateprimer over enzyme, many polymerases exhibit biphasic kinetics 28,–30. This biphasic pattern indicates that in the overall mechanism, the release of products is rate-limiting and slow compared with steps governing chemical catalysis. p180core 3 μm was incubated with either a 15-nucleotide DNA D15 or RNA primer R15 annealed to a 25-nucleotide DNA template T25; 9 μm to allow for the formation of the binary complex Fig. 9A. The preincubated solution was rapidly mixed with dATP 200 μm using an RQF-3 rapid chemical quench apparatus. When the concentrations of products were plotted versus reaction time, an apparent biphasic curve was not observed but rather a linear formation of product for the elongation of both the DNA kss = ± s−1 and RNA kss = ± s−1 primer substrates Fig. 9, B and C. Therefore, it appears that the Polα catalytic core does not exhibit typical biphasic kinetics observed in reactions with main replicative polymerases, Polδ and Polϵ 31, 32. This kinetic behavior indicates that for Polα, the rate of product release is equal to or faster than the rate of chemistry. These data are consistent with structural information showing weaker stabilization of Polα fingers in the closed state compared with Pols δ and ϵ Table 2, which should increase the rate of finger opening and dissociation of a pyrophosphate/templateprimer from a polymerase after catalysis. Conversely, the increased rate of finger opening may slow down the overall rate of chemistry because finger closure stabilizes the triphosphate moiety of dNTP and, therefore, should facilitate the formation of a new phosphodiester bond. These results establish a similarity between Polα and Polλ, a polymerase implicated in base excision repair, which does not exhibit significant subdomain movements in response to nucleotide binding 33, 34.Single-incorporation kinetics of hPolα. A, sequence of templateprimers used for transient kinetic assays. B and C, burst kinetics assay on DNA and RNA primers, respectively. p180core 3 μm was incubated with radiolabeled DNA B or RNA C primer 9 μm annealed to a DNA template and rapidly mixed with dATP 200 μm. Product formation was plotted versus time, and a linear equation was fit to the data red dashed line. D, single-nucleotide incorporation assay. p180core 3 μm was incubated with radiolabeled DNA or RNA primer 100 nm annealed to a DNA template and rapidly mixed with various concentrations of dATP 1–300 μm. Product formation was plotted versus time and fit to a single-exponential equation to obtain the observed rates of product formation. The rates were plotted against concentration of dATP and fit to a hyperbolic equation for both DNA primer blue and RNA primer red elongation to obtain Kd and kpol incorporates the first and second nucleotides into a DNA primer more efficiently than into an RNASingle-nucleotide incorporation experiments were done under single-turnover conditions excess Polα over templateprimer to quantify the maximal rate of catalysis, kpol, and the apparent binding constant for the incoming nucleotide, Kd. p180core 3 μm was incubated with a templateprimer 100 nm and rapidly mixed with varying concentrations of dATP 1–300 μm under rapid chemical quench conditions Fig. 9D. For the DNA primer substrate, the kpol is ± s−1, the Kd is ± μm, and the incorporation efficiency kpol/Kd is μm−1 s−1 Table 4. For the RNA primer substrate, the kpol, Kd, and kpol/Kd values are ± s−1, ± μm, and μm−1 s−1, respectively. It appears that Polα incorporates a nucleotide more efficiently to a DNA primer by approximately a factor of Strikingly, this discrepancy is primarily due to differences in the Kd for the incoming nucleotide. This difference might be explained by reduced stability of the 3′-end of an RNA primer due to lack of interaction between Tyr1055 and the P1 phosphate Fig. 6, B and D. As a result, increased dynamics of the primer 3′-end may affect dNTP loading to the active site through steric hindrance. In addition, the position of the primer 3′-end and dNTP binding might be affected by the presence of ribose at P1 see “Discussion”.Table 4Incorporation efficiencies of dATP from single-incorporation assaysPrimerkpolKdEfficiencyas−1μmμm−1 s− ± ± ± ± were conducted to examine whether subsequent incorporations to a DNA primer were more efficient as compared with an RNA primer after the initial dNTP incorporation event. During the second incorporation, a 3′-hydroxyl of deoxyribonucleotide attacks the α-phosphate of incoming dNTP in both the DNA and RNA primer extension assays. Double-incorporation assays were conducted to estimate the kinetic parameters of a subsequent incorporation after the first incorporation. Experiments were designed to observe a dATP followed by a dTTP incorporation under single-turnover conditions. Preincubated solutions of p180core 3 μm and templateprimer 100 nm were mixed with a solution containing both dATP and dTTP. Because the kinetic parameters of the second incorporation were desired, concentrations of dATP were held at saturating amounts 100 μm for DNA primer and 300 μm for RNA, based on single-incorporation experiments, and concentrations of dTTP were varied from 1 to 300 μm Fig. 10A.Sequential incorporation modeling of hPolα kinetics. A, a representative gel of the elongation of the DNA primer substrate in the presence of 100 μm dATP and 2 μm dTTP. The n band represents the 15-mer primer; n + 1, the primer elongated by dATP across template dT; n + 2, the primer elongated first by dATP followed by dTTP across template dA. p180core 3 μm was incubated with radiolabeled DNA templateprimer 100 nm and rapidly mixed with saturating concentrations of dATP 100 μm and various concentrations of dTTP 1–100 μm. B, the time course of DNA primer elongation. p180core 3 μm was incubated with radiolabeled DNA templateprimer 100 nm and rapidly mixed with saturating concentrations of dNTPs 240 μm. C, time course of RNA primer elongation. D, processivity modeling for DNA primer elongation. Product formation was plotted versus time, and the incorporations were fit to a processive kinetic model using KinTek Global Explorer to provide estimates of the kpol and koff values. The first five incorporations are shown. The red line represents the unelongated primer 15-mer. The inset shows a zoomed in view of the third through fifth incorporations. E, processivity modeling for RNA primer elongation. The dashed line shown in the inset represents the model if the rate of the fourth incorporation of the RNA assay was simulated to be identical to the rate of the fourth incorporation from the DNA processivity appearance of the second incorporation product is complex due to its dependence on the first product being made; product concentration cannot simply be fit to a single-exponential curve like the single-incorporation assays. Thus, the KinTek Global Explorer program 35 was utilized to fit the data and provide an estimate for the kpol and Kd of the dTTP incorporation Fig. S1. For the second incorporation to a DNA primer, the kpol is estimated to be s−1, and the Kd is μm. The estimates for the RNA second incorporation are kpol = s−1 and Kd = μm. This leads to incorporation efficiencies of and μm−1 s−1 in favor of DNA primer extension Table 5. This 6-fold difference in incorporation efficiency for DNA versus RNA for sequential dNTPs is comparable with the single-incorporation results. Contrary to the single-incorporation experiments, a major difference in kpol values is observed in the double-incorporation experiments. We hypothesize that the ∼3-fold difference in kpol values may stem from the primer bending at P4 in the hybrid duplex. The altered helix structure possibly affects the rate of any step in subsequent incorporations after the initial incorporation, such as translocation or necessary protein conformational changes. The reason for similar kpol values during the first incorporation is the preformation of the enzyme/templateprimer complex before the incorporation reaction is 5Incorporation efficiencies of dTTP from double-incorporation assaysPrimerkpolKdEfficiencyas−1μmμm−1 s− ± ± of Polα processivity on DNADNA and DNARNA duplexesA processivity experiment was employed to examine multiple rounds of dNTP incorporation and look at the full extension of the primers and determine whether the rates of each incorporation were different over time between DNA and RNA primers. This analysis observes the kpol for each incorporation event as well as koff, which characterizes the dissociation rate of the Polα/templateprimer complex. The assay was conducted under single-turnover conditions and saturating concentrations 240 μm of all dNTPs Fig. 10, B and C. The products were plotted against time and fit to a processive mechanism using KinTek Global Explorer Fig. 10, D and E; for visual clarity, only the first five incorporations are shown. The kpol and koff values for the first seven incorporations are shown in Table 6. Although this experiment provides only estimates of these rates, it is clear that there is a higher average kpol for the second to fourth incorporations during DNA primer extension, with an average value of s−1 compared with s−1 for RNA. As was shown above, the first incorporation does not show the real kpol values due to preformation of Polα/templateprimer complex; that is why it was omitted from 6Kinetic parameters from processivity assaysIncorporationkpolkoffDNARNADNARNAs−1s− ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± second to fourth fifth to seventh we observe an increase in the rate of incorporation after the fourth incorporation into the RNA primer, with an average kpol for the fifth to seventh incorporations of s−1, which is similar to the value obtained on a DNA primer s−1. This is particularly interesting in light of our structural results pertaining to the kink at P4 in the hybrid duplex structure. This observation can be explained in structural terms by the fact that after four incorporations, the rigid and conserved DNA-binding cleft of Polα no longer makes contacts with the ribonucleotides in the hybrid duplex. This increase in kpol agrees with our previous experiments demonstrating that DNA primer elongation is more efficient. Furthermore, to confirm that the increase from to s−1 after the fourth incorporation is significant, we simulated how the time course of processive elongation of an RNA primer would change if the fourth incorporation rate was modified to match the rate observed in the DNA experiment Fig. 10E, dotted lines. Changing s−1 to s−1 results in a drastically different product distribution. The most prominent effects are the decreased accumulation of R18 and faster production of R20 because the rate of R19 formation is no longer significantly was also observed that the average koff value for DNA primer elongation is s−1 compared to s−1 for RNA for the second to fourth incorporations. The higher koff value for the DNA duplex may be due to the tendency for it to adopt the B-form, which is slightly narrower than the templateprimer-binding cleft. In contrast, the hybrid duplex is squeezed between Gly841 and Gly1076 Fig. 6B; see “Discussion”, which may potentially slow the dissociation and association of the duplex. Indeed, the estimates for kon values and nm−1 s−1 for DNA and the hybrid duplex, as a first approximation calculated by dividing the average koff for the first four incorporations by the Kd show a higher rate of complex formation with the DNA duplex. It is noteworthy that the interaction of the thumb with the distant part of a templateprimer T5P5–T9P9 can also contribute to slower dissociation of Polα from the hybrid duplex because this DNA-binding region is optimal for recognition of the double helix of mixed AB-form Fig. 6B. Accordingly, two dinucleotide steps in this region of the DNA duplex have the mixed AB-form Fig. 6C.The ratio kpol/koff provides an estimate of processivity and indicates that on average for the second to fourth incorporations hPolα incorporates approximately three or four and six consecutive nucleotides before complex dissociation in the case of DNA and RNA primer, respectively. Using a DNA-trap, it was shown that hPolα incorporates six consecutive nucleotides before dissociation from the DNA duplex 36. Slightly higher processivity observed by Copeland and Wang's assay may be due to the absence of the monovalent salt in reaction m NaCl was used in our experiments. In extension of the DNA primer, yPolδ has the same processivity as hPolα, whereas yPolϵ catalytic core containing an unique processivity domain incorporates 27 nucleotides in one binding event, in the absence of monovalent salt in reaction 22. Interestingly, koff values are comparable for hPolα and yPolϵ, and the difference in processivity is mainly due to the higher incorporation rate of the latter. Higher processivity of Polα on the hybrid duplex can prevent the enzyme from premature the processivity of a polymerase can be qualitatively assessed by inspection of separated products on a gel. Comparing the processivity gels from Polα Fig. 10, B and C and T7 DNA polymerase 30, the striking difference is the presence of primers that have not been fully extended with dNTPs at later points in the experimental time course. Through our experiments, it is observed that there is a significant population of primers that have been only singly elongated D16 at later time points even when the final product is finally produced D25. In contrast, once T7 DNA polymerase incorporates a dNTP into a primer, it continues to incorporate nucleotides into the same primer. In the direction of increasing time, the pattern on the gel for the Polα can be described as extending a ladder upward, whereas T7 can be described as shifting a ladder upward. Processivity experiments have also been done with Polϵ 31, and the pattern of processive polymerization is similar to Polα. Although Polϵ is considered more processive than Polα, the striking difference between T7 and the former polymerases highlights the highly variable kinetic characteristics of sugar at the primer 3′-end affects Polα fidelityThe potential of Polα to generate 12 possible mismatches during the extension of a DNA or RNA primer was compared. This study revealed that p180core has pronounced difference in nucleotide selectivity during the addition of the first dNMP to the RNA and DNA primers Fig. 11. It is relatively accurate on RNA but easily generates various mismatches when extending the DNA primer. The most prominent of them are AdA, TdG, GdT, GdG, and CdT the left letter indicates a nucleotide of the template. Moreover, p180core efficiently extends some mismatches Fig. 11B, lanes 6, 7, 11, 13, and 16, even with noncognate nucleotides lanes 6, 7, and particularly lane 16 showing two consecutive AdC mismatches and their extension. Interestingly, Polα is not able to extend the most prominent mismatch, AdA, whereas it easily extends CdA and GdA by generating the next consecutive mismatch, AdA Fig. 11B, compare lanes 6–8. The results are consistent with kinetic studies of human and yeast Polα, showing that they discriminate against noncognate dNTPs and the mismatched primer terminus by a factor of 103 to 104 and 500, respectively 25, 37, 38. The discrimination mainly depends on the difference in affinity for a correct versus incorrect incoming nucleotide, thus explaining reduced selectivity in the presence of mm dNTP in our experiments this concentration is ∼20-fold higher than concentration of dNTPs in mitotic cells 39, 40.Comparison of Polα fidelity in extension of DNA and RNA primers. A, template and primer sequences. DNA B and RNA C primers were extended with cognate and noncognate dNTPs. Reactions were carried out at 35 °C for 20 min and contained 20 nm p180core except lane 1, which is a control reaction without enzyme, 1 μm templateprimer, 2 mm MgCl2, and mm Polα becomes less accurate after incorporation of just one dNMP to an RNA primer. For example, no product was detected during the first step of RNA primer extension with dATP opposite the template adenine Fig. 11C, lane 8, whereas in the same time frame, a significant amount of AdA mismatch was generated during the second step of RNA primer extension, following incorporation of the first dAMP opposite the template thymine lane 9. More efficient mismatch generation during the second step is not due to formation of the weak TdA bp because the same effect was observed using other dNTPs Fig. 11C, comparison of lanes 2 and 4 dGTP, 11 and 12 dTTP, and 15 and 16 dCTP; appearance of the second and/or the following bands above the product indicates mismatch formation. Dramatic accuracy reduction after the insertion of just one dNMP indicates that Polα fidelity depends on the presence of ribose at P1 but not on the primer bending at P4 Fig. 7.Initial burst experiments, performed on a broad time scale to establish the relative rate of dNTP incorporation by the p180core, revealed that Polα incorporates an incorrect dNTP at a notable rate Fig. 12. This is illustrated at later time points by the appearance of a second band that corresponds to a dATP incorporation across from a template adenine. This observation was unexpected due to the prevalence of misincorporation within a time frame relatively close to correct incorporation. Although previous studies 25, 41 have indicated that Polα is a low-fidelity polymerase, to the best of our knowledge, this is the first direct observation of significant Polα misincorporation in a millisecond time scale. These results further emphasize the benefits of utilizing a pre-steady-state kinetic displays misincorporation on a rapid time scale. p180core 3 μm was incubated with radiolabeled DNA templateprimer substrate 10 μm and rapidly mixed with saturating dATP 500 μm. The gel shows the elongation of the DNA primer in the presence of only dATP. The n + 1 band corresponds to a correct incorporation of dATP across from the templating dT. The n + 2 band corresponds to a misincorporation of dATP across from a templating dA. The sequence shows where the incoming dATP is incorporated onto the the structure of Polα catalytic domain is thoroughly studied. Eight crystal structures have been reported for yeast and human orthologues, which include the apo-form, binary, and ternary complexes, containing DNA or RNA primer, as well as the structures of human primosome and the complex of hPolα with the natural inhibitor aphidicolin 6, 16,–18. In addition, the crystal structures of the CTD–B-subunit complexes are known for yeast and human Polα 11, 12. With regard to the catalytic subunits of Polδ and Polϵ, only the structures of the ternary complexes have been determined for their catalytic domains 22, 26, 42 as well as the structure of the Polϵ CTD–B-subunit complex 43.The role of conformational dynamics in DNA synthesis by PolαAccumulated structural information for the catalytic domain of Polα indicates that the N-terminal and exonuclease subdomains are firmly associated, whereas positions of other subdomains relative to the N-terminal part of p180core and each other are quite flexible this work; also, see Refs. 6 and 16. Such flexibility of the subdomains responsible for substrate binding and catalysis defines the conformational dynamics of the polymerase during DNA synthesis. For example, significant mobility of the thumb might be important for accommodation and translocation of duplexes containing lesions, mismatches, and different types of primers RNA, DNA, and chimeric RNA–DNA. Accordingly, Polα has notable translesion activity and the ability to extend the mismatched 3′ terminus of a primer 25, 44.Relatively weak interactions of the mobile part of the fingers with the other subdomains allow them to switch quickly between the open and closed states. High dynamics of the fingers is important for rapid closing after dNTP binding and opening after catalysis to release the products. The binding of dNTP results in the shift of equilibrium to the closed conformation, which in hPolα is primarily due to formation of two H-bonds between Arg922 and the γ-phosphate Table 2. The dNTP becomes trapped after fingers closing, promoting effective catalysis. Crystal packing may have helped to stabilize the open conformation of the fingers in the hPolα/DNADNA/dCTP complex because, in free energy terms, the open state should be less favorable versus the closed one in the presence of bound dNTP. The other known example of a DNA polymerase crystallized in the open ternary complex is Polβ, which belongs to the X-family 45. That structure was obtained after mutating the conserved Arg283 important for stabilization of the closed state, which shifted the equilibrium between two conformations of the helix N to the open one. Whereas the helix N fulfills a similar task as the fingers in Polα, it does not interact with a triphosphate of an incoming study revealed that Polα fingers sense the triphosphate moiety of dNTP even in the open state Fig. 4A and Table 2. Interaction of open fingers with dNTP might be important for stabilization of the ternary complex during the first step of its formation and could trigger fingers closing. Currently, there is a lack of information on how the fingers of yeast Polδ and Polϵ interact with dNTP in the open conformation. Furthermore, the structures of these Pols in the apo-form are not reported, which precludes the possibility to model potential interactions between the residues of open fingers and the triphosphate. The absence of a rate-limiting step for product release revealed from pre-steady-state burst experiments reflects fast dynamics of Polα fingers caused by weak stabilization of the closed conformation. This structural peculiarity of Polα results in higher probability of fingers opening, facilitating mismatch accommodation at the nascent bp-binding pocket see below. This finding may provide a key insight into the mechanism of the generation of mutation hot spots in the genome 7. Interestingly, all amino acid changes in hPolα conferring resistance to CD437, a potential anticancer molecule recently discovered to target Polα, are located at or near the interface between the fingers and the N-terminal/Exo subdomains 9.Comparison of templateprimer binding by replicative DNA PolsAll invariant residues of replicative DNA polymerases contacting the templateprimer interact with the first five bp of the duplex Fig. 6. The most interesting feature of these polymerases is the interaction of T3 and P2 phosphates with two conservative glycines of the palm Gly841 and Gly1076 in hPolα. These glycines determine the width of the double helix between the second and third bp from the primer 3′-end, with similar distances between their main-chain nitrogens for hPolα in complex with DNARNA Å and yeast Polδ and -ϵ in complex with DNADNA and Å, respectively. The other common feature of replicative polymerases is the absence of interaction with a T1 phosphate Fig. 6, which might be important for adjusting the position of the templating nucleotide to obtain optimal pairing with an incoming dNTP. In contrast to Polϵ, the thumbs of Polα and Polδ are engaged almost exclusively with the primer and do not interact with T5 and T6. The exonuclease subdomains of Polα and Polϵ interact with the P3 phosphate, whereas there is no such interaction in Polδ. The catalytic domain of yPolδ has the smallest footprint on the double helix eight bp, and that of yPolϵ has the largest one 10 bp, with a potential up to 12 bp 22. The low footprint of Polδ on the template is compensated by the double contact with the P6 phosphate Fig. 6E. In contrast, Polα and Polϵ sense only five continuous nucleotides of the primer. The recently discovered P-domain of Polϵ allows it to make three additional contacts with a DNA duplex Fig. 6F, one with a primer P8 and two with a template T2 and T3, and two extra contacts are possible 22. These additional contacts and the extensive network of polar contacts between the thumb and the template explain the higher processivity of Polϵ catalytic domain compared with Polδ and Polα. In both ternary complexes of hPolα, the side chain of Arg834 located on the flexible loop extends into the major groove and packs against the base of T3. For comparison, yeast Polα and Polϵ have no contacts with the major groove, whereas the β-hairpin of yPolδ is anchored in it 26.In solution, the RNA, DNA, and hybrid duplexes DNARNA prefer to be in the A-, B-, and mixed AB-form, respectively 46,–48. The A-form is wider and shorter than the B-form, with a diameter of 23 Å versus 20 Å and a rise per base of Å versus Å, respectively. DNA-protein interactions can affect the shape of the double helix, locally or globally 49. The conserved character of interaction with the first four bp determines the universal form of the double helix for this region in the ternary complexes of Polα, Polδ, and Polϵ, independent of the primer nature Fig. 6. This is also true for the chimeric RNA–DNA primer Fig. 6D, which reflects the initial steps of RNA primer extension by Polα. Replicative DNA polymerases transform the first two dinucleotide steps of DNADNA into the mixed AB-form and the third dinucleotide step of DNARNA into the B-like form the latter is currently shown only for ternary complexes of Polα. Interestingly, in all ternary complexes of eukaryotic replicative DNA polymerases, the sugars of T0, T1, P1, and incoming dNTP have a 3′-endo pucker or closely C2′-exo and C4′-exo puckers, which might be important for alignment of hPolα/DNARNA/dCTP and yPolδ/DNADNA/dCTP complexes RMSD of Å for 524 Cα atoms shows good superposition for the first four bp of the double helix, which is consistent with the same duplex form in this region as well as with high structural similarity for the palm and the N-terminal part of the thumb Fig. 13. In contrast to Polα and Polϵ, Polδ has a short loop in the tip of the thumb residues 895PNYTNP900; highlighted in blue, which allows it to interact with the P6 phosphate using the main-chain nitrogens of Thr898 and Asn899 Fig. 13. This loop also contains the conserved Tyr897, which interacts with P5 and, together with Thr898 and Asn899, would clash with P6 and P7 upon DNARNA binding by Polδ. The additional steric gate residues for the wider AB-form of the hybrid duplex in yPolδ are Leu935 and Tyr936, which are located on the 310-helix of the thumb tip and correspond to Ala1188 and Ser1189 of hPolα, respectively Fig. 13. This region may have a minor role in hybrid duplex sensing by Polδ due to potential flexibility of the thumb. Consistent with this structural analysis, Polδ extends the DNA primer with significantly higher efficiency versus RNA 50. Similar to Polα, Polϵ does not interact with P6 Fig. 6, B and F and efficiently extends primers containing up to five consecutive ribonucleotides at the 3′-end 51. Polδ also differs from Polα and Polϵ 22 by the presence of a β-hairpin near the unprimed region of the template Fig. 13.Alignment of hPolα/DNARNA/dCTP and yPolδ/DNADNA/dCTP. The ternary complexes were aligned as described in the legend to Fig. 4B. hPolα and DNARNA are colored according to Fig. 1. yPolδ and DNADNA are colored pale yellow and gray, respectively; the region spanning residues 895–900 of yPolδ is highlighted in blue. Two potential clash areas between Polδ and the RNA primer are depicted by red stars. The distant part of the 5′-overhang of DNA template complexed with yPolδ is not shown for summary, all replicative DNA Pols have the same rigid templateprimer-binding cleft for the first four bp from T1P1 to T4P4, which is mainly formed by the palm residues Fig. 6. The DNA-binding site distant from the catalytic center for the duplex region spanning 5–10 bp from the primer 3′-end significantly differs in B-family Pols and probably reflects the role of each polymerase in DNA replication and repair. For example, the low efficiency of RNA primer extension by Polδ as well as increased processivity of the Polϵ catalytic domain are mainly due to polymerase-specific structural peculiarities of this part of the DNA-binding basis of Polα fidelity and infidelityPolα demonstrates relatively high fidelity during RNA primer extension with the first dNTP, whereas the incorporation of the following nucleotides is less accurate Fig. 11. Surprisingly, the bulky purinepurine mismatches are among the most prominent, which is probably due to the ability of Polα to easily fit them into the nascent bp-binding pocket. Current structural information suggests that at least four factors may assist in formation of the purinepurine mismatches fingers opening Fig. 4A, syn-conformation of T0 stabilized by Arg784 17, 18, absence of the β-hairpin Fig. 13; in Polδ, the T−1 sugar and the T0 phosphate are secured between the β-hairpin and the N-terminal subdomain, and weak interaction of Polα with the template 5′-overhang Fig. 6B, which allows more conformational freedom for T1–T−1 during mismatch accommodation. The exact mechanism of Polα low fidelity is not known due to the absence of structural information for the WT B-family DNA polymerases in the mismatch-containing ternary the structure of the p180core/DNARNA/aphidicolin complex 18 provides a clue for how the most prominent AdATP mispair fits the active site of Polα. Indeed, all four factors listed above play a role in the accommodation of the bulky aphidicolin in place of the incoming dCTP, where the local change of template conformation Fig. 14A resulted in disruption of interaction between T−1 and Ser785. The modeling shows that Polα can easily accommodate all purinepurine mispairs when the T0 purine is in syn-conformation Fig. 14B. In addition, AdATP can fit the nascent bp-binding pocket when the templating adenine is in the anti-conformation. Small steric hindrance between two purines in this case might be relieved by the additional minor adjustments in the active site. This model of AdATP mispair accommodation is applicable to the other mismatches. Kinetic studies of yeast Pols δ and ϵ are consistent with the role of the β-hairpin in the fixation of the template position at T0, which should significantly reduce the chance to generate the bulky purinepurine mismatches. Among replicative Pols of eukaryotes, only Polδ has the β-hairpin and generates the purinepurine mismatches with relatively low efficiency 32. Accordingly, exonuclease-deficient yPolδ and yPolϵ discriminate against the GdG mismatch by a factor of 6 × 104 32 and 5 × 103 52, of structural factors affecting Polα selectivity for dNTP. A, DNA template has significant conformational flexibility near T0 in the complex with hPolα. Aphidicolin binding by Polα results in rotation of the T0 base and the T−1 sugar by 112 and 168°, respectively. The p180core/DNARNA/dCTP and p180core/DNARNA/aphidicolin complexes PDB codes 4QCL and 5Q4V, respectively were aligned with RMSD of Å using the palm. B, modeling of the AdATP mispair in the hPolα active site. DNA carbons are colored slate and gray in the case of the ternary complexes with dCTP and aphidicolin, respectively. The carbons of modeled adenines at T0 and in place of dCTP are colored green and cyan, respectively. C, the modeled 2′-OH has steric hindrance with Thr1003. The coordinates of the p180core/DNARNA/dCTP complex PDB code 4QCL were used for modeling the sugar of the 3′-dideoxycytidine has a 3′-endo pucker.The apparent low fidelity of Polα during incorporation of the second and following dNMPs allows us to compare the enzyme with other well-studied polymerases. HIV-1 reverse transcriptase has been established as a low-fidelity polymerase 53. Its activity is beneficial for the virus, allowing for rapid mutation and development of resistance against antivirals. Although poor discrimination between nucleotides may be desirable for viruses, this trait is not particularly advantageous for host polymerases, which are responsible for perfectly replicating the genome. The low accuracy of Polα might be important for the evolution of regulatory elements in eukaryotic genomes 7. Polα has a significant contribution to the mutation rate in vivo, despite the fact that only of Polα-synthesized DNA has been shown to be retained in the mature genome 7. Poor fidelity in host polymerases is not limited to Polα and translesion polymerases, including the other member of the B-family, Polζ. Recently, PrimPol, a primase-polymerase that is implicated to reinitiate replication after mitochondrial DNA damage, has been characterized to have low fidelity 54, 55. It is interesting that we observe a prominent purinepurine mismatch in our experiments with Polα AdA as well as with PrimPol GdA.High accuracy during the addition of the first dNMP to the RNA primer, which Polα receives from primase during intramolecular transfer 6, might be important for a smooth switch from the RNA- to DNA-polymerizing mode of the primosome. The structure of the p180core/DNARNA/dCTP complex can provide a key to the effect of the 3′-ribose on Polα fidelity. The 2′-OH modeled at the primer 3′-end has steric hindrance with the conserved Thr1003 Fig. 14C. The P1 ribose shall move from Thr1003 by ∼ Å to avoid steric clashing, which will affect the position of the T1–T−1 and can alter their ability to change the conformation upon mispair structures of the catalytic domain of replicative DNA Pols provide a rationale for the mutagenic effect of substitutions of the conservative leucine 37, 44, 56, 57, whose main chain interacts with the dNTP β-phosphate Leu864 in hPolα; Fig. 4. This leucine is located in the hydrophobic pocket formed in hPolα by Phe861, Tyr865, Ile868, Ile983, and Leu1036. Bulky side chains of methionine and phenylalanine in place of Leu864 cannot fit into this pocket, which would result in the distortion of the upper part of the α-helix Fig. 4, affecting the position of Ser863, Leu864, and Tyr865. Among these residues, Tyr865 is the most important for Pol selectivity. Such distortion of the dNTP-binding pocket can result in dNTP wobbling and reduced discrimination against overall mechanism of RNA primer extension by PolαAccumulated information from structural and functional studies allows us to understand the mechanism of chimeric RNA–DNA primer synthesis by the primosome, including the ability of Polα to efficiently extend primers with varying DNA/RNA ratio and bind double helices with different helical parameters. The following structural peculiarities of the Polα catalytic domain are important to perform this task small primer-binding interface, thumb flexibility, and the universal templateprimer binding site, which is able to accommodate the substrates with different structures. Perhaps during evolution, Polα developed the ability to fit the templateprimer into a universal form of the double helix by deforming both DNADNA and DNARNA studies of the human primosome point out that Polα receives a 9-mer RNA primer from primase 6. The data presented here establish the four main steps of RNA primer extension by Polα incorporation of the first dNMP first step, second to fourth second step, fifth to ninth third step, and tenth and further fourth step. The main features of the first step are the enhanced accuracy Fig. 11 and the relatively low affinity for dNTP Table 3. The increased Polα selectivity for a cognate dNTP is probably due to the presence of ribose at P1 Fig. 14C, whereas dNTP binding is also affected by the primer bending at P4 Fig. 7. The second step of RNA primer extension is characterized by reduced polymerization rate and increased processivity compared with Polα activity on a DNA primer Tables 5 and ​6. For this step, primer bending at P4 and the squeezing of a hybrid duplex between Gly841 and Gly1076 may affect templateprimer translocation along the DNA binding cleft and dissociation from the polymerase. The third step of extension is similar to the fourth step, which corresponds to extension of a DNA primer Table 6. The hybrid part of the templateprimer leaves the rigid and relatively narrow section of the DNA binding cleft and interacts only with the thumb, which is slightly more selective for a mixed AB-form of the double helix versus a B-form Fig. 6, B and C. During the fourth step, the Polα catalytic domain does not sense the hybrid part of a templateprimer, and the characteristics of DNA synthesis should be identical to that on a DNADNA size of DNA tracks synthesized by Polα on the leading and lagging strands has not been determined yet. It is possible that the length of chimeric RNA–DNA primers is precisely regulated, as was recently shown for the RNA primer 6. Only the structural properties of the primosome define the mechanism of RNA primer switching between primase and Polα, whereas additional factors should play a role in chimeric primer switching between Polα and Polϵ or Polδ on the leading or lagging strand, respectively. In 2013, the mechanism of primer synthesis termination by Polα, based on its low activity in extension of homopolymeric DNA primers, had been proposed 16. As was shown later, the low activity of Polα in primer synthesis on polydT template is due to formation of unconventional DNA structures, which depends on the type of the divalent ion and its concentration 58. Moreover, comparable activity was observed during extension of DNA and RNA heteroprimers 59, 60. The data presented here indicate that Polα works on the DNA duplex with higher efficiency than on the hybrid one Tables 4 and ​5. On the other hand, Polα has low processivity on the DNA duplex, which can favor its displacement from the templateprimer by Polϵ on the leading strand and by the complex of the replication factor C and proliferating cell nuclear antigen on the lagging strand replication factor C is then exchanged for Polδ 61.It is possible that additional factors play a role in determination of the size of DNA track synthesized by Polα. For example, the primase accessory subunit p58 in humans has higher affinity for the templateprimer Kd is 33 nm 62 than Polα Kd is 58 nm; Table 3 and can hold the 5′-end of RNA primer during its extension with dNTPs and promote Polα reloading on the DNA primer 6. In this case, the linkers in primosome would allow Polα to generate a ∼20-mer DNA track. The primosome structure also predicts significant steric hindrance after synthesis of a ∼10-mer DNA track due to ∼360° rotation of the catalytic domain of Polα relative to the C-terminal domain of p58; this would require a temporary dissociation of Polα from the templateprimer to untwist the linkers. In addition, it might be the signal from the replisome, leading to primosome dissociation from the templateprimer. For example, as was shown in yeast, Polα is tethered to the CMG helicase through the interaction between the N-terminal domain of the catalytic subunit and the replication factor Ctf4 13. In humans, however, the N-terminal domain of the accessory B-subunit connects Polα to a replisome 63, similar to the mode of Polϵ recruitment to a replication fork 64. In both cases the CMG-interacting domain of Polα is connected with the rest of the primosome by a long flexible linker 4. During Okazaki fragment synthesis, Polα moves along the parental DNA strand in the direction opposite to the replication fork, and at some point, the tension in the linker, connecting the catalytic core with the replisome, will result in Polα displacement from the primer 3′-end. Conducting structural and functional studies with the reconstituted eukaryotic replisome will shed more light on the mechanism of the templateprimer switch from Polα to Polδ and proceduresOligonucleotides and reagentsOligonucleotides were manufactured by IDT Inc. Deoxyribonucleotides used to extend the primers were obtained from Roche Life Sciences. Reagents for crystallization were obtained from Hampton Research. [γ-32P]ATP was obtained from PerkinElmer Life expression and purificationCloning, expression, and purification to homogeneity of p180core residues 335–1257, which contains the catalytic domain of hPolα, have been described elsewhere 18. Peak fractions obtained from Heparin HP HiTrap column GE Healthcare were combined and dialyzed to the buffer specific for each application see below.Analysis of DNA-polymerizing activity in extension of fluorophore-labeled primersPurified p180core was dialyzed to 10 mm Tris-HCl, pH 100 mm KCl, 1% glycerol, and 1 mm DTT, and flash-frozen in aliquots. The primers had a Cy5 label at the 5′-end. Reactions were carried out at 35 °C in buffer containing 30 mm Hepes-KOH, pH 50 mm KCl, and 1 mm DTT. Reactions were started by mixing two solutions; one of them contained p180core and the template primer, and the other one contained dNTP and the divalent metal their concentrations are indicated in the figure legends; other components were the same in both stocks.CrystallizationDNARNA and DNADNA duplexes were obtained at mm concentration by annealing at 43 °C for 30 min after heating at 70 °C for 1 min in buffer containing 10 mm Tris-HCl, pH and 70 mm KCl. In the case of DNARNA duplex, the sequences for DNA template and RNA primer were 5′-ATTACTATAGGCGCTCCAGGC the region complementary to a primer is underlined and 5′-rGrCrCrUrGrGrArGrCrG/ddC/, respectively /ddC/is a dideoxycytidine, which prevents polymerization during crystallization. In the case of DNADNA duplex, the template and primer were 5′-ATAGGCGCTCCAGGC and 5′-GCCTGGAGCG/ddC/, respectively. Dialyzed protein sample 15 μm p180core was diluted with dialysis buffer 10 mm Tris-HCl, pH 100 mm KCl, 1% glycerol, and 1 mm DTT containing 36 μm templateprimer, mm MgCl2, and 12 mm dCTP. The ternary complex was concentrated 10-fold and flash-frozen in liquid nitrogen. Before crystallization experiments, the aliquots of protein/templateprimer/dCTP solutions were defrosted and centrifuged to remove the precipitate, and the sample monodispersity was verified with the dynamic light scattering. The screening of crystallization conditions was performed with the sitting-drop vapor diffusion method at 295 K by mixing 1 μl of ternary complex solution with 1 μl of reservoir solution. Initial screen solutions producing tiny crystals were optimized to produce well-shaped crystals at 295 K with reservoir solutions containing mm zinc sulfate, v/v PEG MME 550, and 50 mm MES, pH for p180core/DNARNA/dCTP and mm cobalt chloride, 2 mm tris2-carboxyethyl phosphine TCEP, pH 250 mm 1,6-hexanediol, and 50 mm sodium acetate, pH for p180core/DNADNA/ collectionFor data collection, each crystal was soaked in cryoprotectant solution 100 mm potassium chloride, mm zinc sulfate, mm magnesium chloride, 15% v/v PEG MME 550, 15% v/v ethylene glycol, and 50 mm MES, pH for p180core/DNARNA/dCTP crystals, and mm cobalt chloride, 2 mm TCEP, pH 230 mm 1,6-hexanediol, 26% v/v ethylene glycol, and 50 mm sodium acetate, pH for p180core-DNADNA-dCTP crystals for a few seconds, scooped in a nylon-fiber loop, and flash-cooled in a dry nitrogen stream at 100 K. All initial diffraction data were obtained on a Rigaku R-AXIS IV imaging plate using Osmic VariMaxTM HR mirror-focused CuKα radiation from a Rigaku FR-E rotating anode operated at 45 kV and 45 mA. Complete diffraction data sets were collected using synchrotron X-rays on the Argonne National Laboratory Advanced Photon source beamline 24-ID-E using ADSC Quantum 315 detector. All intensity data were indexed, integrated, and scaled with DENZO and SCALEPACK from the HKL-2000 program package 65. The crystals of p180core/DNARNA/dCTP belong to trigonal space group P3221 and diffract up to Å resolution, and the crystals of p180core/DNADNA/dCTP belong to tetragonal space group P42211 and diffract up to Å resolution. Both crystals contained only one copy of the ternary complex in the asymmetric unit. The crystal parameters and data-processing statistics are summarized in Table structure determinationInitial phases for p180core/DNARNA/dCTP structure were determined by the molecular replacement method using the coordinates of backbone atoms of the yeast Polδ catalytic core derived from its ternary complex with templateprimer and dCTP PDB code 3IAY as a search model. The positions of magnesium and zinc ions were determined using an anomalous difference Fourier map. Molecular replacement and initial automated model rebuilding with Phenix 66 revealed over 70% of correctly built protein structure. The model building was continued and completed manually with Turbo-Frodo, and the structure was refined using standard protocols of CNS version 67. After the addition of solvent molecules, the model was refined at Å resolution to an Rcryst of 21% and an Rfree of The structured region of protein starts with Glu338 and ends with Thr1244. The electron density maps were poor or were missing also for the internal regions 674–677, 809–833, and 883–895; therefore, these regions were excluded from the p180core/DNADNA/dCTP structure was determined by the molecular replacement method using the coordinates of p180core in the complex with DNARNA and aphidicolin PDB code 5Q5V as a search model. The model was adjusted manually with Turbo-Frodo. The positions of magnesium and cobalt ions were determined using an anomalous difference Fourier map. The structure was refined at Å resolution using standard protocols of CNS version 67 to an Rcryst of and an Rfree of The traceable electron density starts with Glu338 and ends with Val1248. The regions 674–677, 810–833, and 883–895 with missing electron density were excluded from the model. The electron density was relatively weak for the portion of the thumb closer to the thumb tip, especially for residues 1137–1154, indicating their partial disorder and/or elevated refinement statistics for both structures are summarized in Table 1. The figures containing molecular structures were prepared with the PyMOL Molecular Graphics System version Schrödinger, LLC.Analysis of DNA-binding activityPurified p180core was dialyzed to 10 mm Tris-Hepes, pH 200 mm NaCl, 2% glycerol, and 2 mm DTT, concentrated to 100 μm, and flash-frozen in aliquots. The 15-mer primers 5′-GGACTCCGAGCTGCC; DNA and RNA were labeled using [γ-32P]ATP by T4 polynucleotide kinase from New England Biosciences. The 15-mer primers was were then annealed to the 25-mer template 5′-AATGTTTCATGGCAGCTCGGAGTCC; the complementary region is underlined at 95 °C for 5 min, 55 °C for 15 min, and 37 °C for 10 min. The annealed substrates were prepared to contain 50,000 cpm/μl and were used within approximately 1 p180core 4 μm was mixed with reaction buffer 10 mm Tris-HEPES pH m NaCl, 5 mm MgCl2, 1 mm DTT and 1 mg/ml BSA. Radiolabeled templateprimer 6 nm, DNA or RNA primers was mixed with the reaction buffer and 1 mg/ml BSA. Serial dilutions of the p180core solution were made so that solutions contained a range of 20 nm to 4 μm enzyme. The diluted enzyme solutions were mixed with an equal volume of the templateprimer solutions to obtain a final concentration of 10 nm to 2 μm p180core and 3 nm templateprimer. The solutions were allowed to incubate for 15 min at 25 °C, and glycerol was added for a final percentage of 2%. The unbound and enzyme-bound radiolabeled templateprimers were separated by 5% native PAGE in TBE for 33 min at 150 V at 4 °C. The gels were visualized by the Molecular Imager FX phosphor imager Bio-Rad and quantified by Quantity One, version Bio-Rad.Pre-steady-state kinetic assaysPre-steady-state kinetic assays were performed using the RQF-3 rapid chemical quench apparatus KinTek at 37 °C. All kinetics assays were carried out using reaction buffer. Polα was incubated with templateprimer and rapidly mixed with dNTPs before quenching with m EDTA. Products were collected in a tube with formamide dye bromphenol blue w/v, xylene cyanol w/v and separated by denaturing urea PAGE. The radiolabeled products were visualized by the Molecular Imager FX phosphor imager Bio-Rad and quantified by Quantity One, version Bio-Rad.For optimal separation of RNA primer-elongated products, a modified denaturing urea PAGE protocol was employed. A solution of acrylamide mix was made with the addition of 10% formamide 19% acrylamide, 1% bisacrylamide, 10% formamide, 7 m urea, 1× TBE buffer pH The acrylamide polymerization reaction was initiated with ammonium persulfate and TEMED. Before loading the gel with sample, the gel was run at 3000 V for at least 2 h to allow the gel to heat up. The elongated products containing equal parts of quenched product and formamide dye were then loaded onto the gel <10 μl and allowed to run for ∼6 incorporation kineticsBurst incorporation kinetics were conducted with excess templateprimer 9 μm to Polα 3 μm. To ensure that binding of the incoming nucleotide was not rate-limiting, a high concentration of dATP 200 μm was used in the polymerization reaction. The reaction products were quantified, and the concentration of product was plotted against time and fit to both a linear and a burst equation, [product] = A × 1 − e−kobst + A × kss + t1 where A is the burst phase amplitude corresponding to the active site concentration, kobs is the observed rate, kss is the steady-state rate, and t is the time. In the event of misincorporation after a correct incorporation, 500 μm dATP was used to ensure nucleotide and double-nucleotide incorporation assaysSingle- and double-nucleotide incorporation assays were performed with excess Polα 3 μm to templateprimer 100 nm. In the case of single-nucleotide incorporation assays, the concentration of dATP was varied 1–300 μm. For each experiment at a particular dATP concentration, the concentration of product was plotted against time and fit to a single-exponential equation, [product] = A × 1 − e−kobst2 where A is the amplitude, kobs is the observed rate for the incorporation of the incoming dNTP, and t is the time. The kobs was then plotted against dATP concentration and then fit to a quadratic equation, kobs=kpol×[dNTP]KD×[dNTP]3 to obtain the Kd, the apparent binding constant for the incoming nucleotide for the Polα/templateprimer binary complex, and kpol, the maximum rate of nucleotide the case of double-nucleotide incorporation assays, Polα first incorporates a dATP followed by a subsequent dTTP incorporation. The concentration of dATP was held constant at saturating amounts based on single-turnover experiments 100 μm for DNA primer and 300 μm for RNA primer, and the concentration of dTTP was varied 1–300 μm. The data were fit using KinTek Global Explorer 35 to provide an estimate for the Kd and kpol for the second incorporation assaysProcessive polymerization assays were performed under similar single-turnover conditions as the single- and double-nucleotide incorporation assays with excess Polα 3 μm to templateprimer 100 nm. The Polα/templateprimer complex was mixed with saturating concentrations of all dNTPs 240 μm each to allow for complete extension of the primer substrate. The processivity data were modeled in KinTek Global Explorer to determine the kpol and koff, the rate of dissociation of the templateprimer substrate from Polα, for each incorporation fitting the data to the models, the following assumptions were made 1 all incorporation events were irreversible, 2 the Kd and kpol values obtained from the single-incorporation experiments were manually fixed for the first incorporation in the double-incorporation modeling, 3 the kon values for templateprimer binding to Polα were assumed to be the same, disregarding primer length analysis and kinetic modelsAnalysis of the radiolabeled products through the phosphor imager provides raw data in count values. For each time point, the counts of each band present were summed to calculate the total amount of counts and subsequently the percentage of each product length present. These values were then normalized to add up to 100% and then converted into concentration amounts based on the total amount of nucleic acid substrate used in the KinTek Global Explorer program requires user-inputted models to fit kinetic data for the estimation of rates Fig. S2 35, 68. Briefly, the data from double-incorporation assays were fit to a two-step mechanism of dNTP binding followed by incorporation that was modified to account for two incorporations. The processivity data were fit using a model that allows for binding or dissociation of each DNA product and combines the dNTP binding and incorporation event into one contributionsA. G. B. expressed and purified the catalytic domain of hPolα and prepared the ternary complexes. N. D. B. and T. H. T. crystallized the ternary complexes and collected the diffraction data. V. N. D. performed kinetic studies under the supervision of K. S. A. Y. Z. analyzed polymerase activity under the supervision of A. G. B. and Y. I. P. T. H. T. initiated the project and determined the crystal structures. A. G. B. and V. N. D. wrote the manuscript, with contributions and critical comments from the other MaterialAcknowledgmentsThe Eppley Institute's X-ray Crystallography Core Facility is supported by NCI, National Institutes of Health NIH, Cancer Center Support Grant P30CA036727. This work is also based upon research conducted at the Northeastern Collaborative Access Team NE-CAT beamline at the Advanced Photon Source at Argonne National Laboratory, which is supported by NIGMS, NIH, Grant P41 GM103403. Use of the Advanced Photon Source, an Office of Science User Facility operated for the United States Department of Energy Office of Science by Argonne National Laboratory, was supported by Department of Energy Contract DE-AC02-06CH11357. The University of Nebraska DNA Sequencing Core receives partial support from NIGMS, NIH, Grants INBRE-P20GM103427-14 and COBRE-1P30GM110768-01 as well as Fred and Pamela Buffett Cancer Center Support Grant work was supported by NIGMS, National Institutes of Health, Grants GM101167 to T. H. T. and GM49551 to K. S. A. and by National Institutes of Health training Grant 5T32GM06754 3-12 to V. N. D.. The authors declare that they have no conflicts of interest with the contents of this article. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of article contains Figs. S1 and atomic coordinates and structure factors codes 4QCL and 6AS7 have been deposited in the Protein Data Bank abbreviations used are PolDNA polymerasehPolhuman PolyPolyeast Polp180corecatalytic domain of hPolαExoexonuclease subdomain of PolαH-bondhydrogen bondRMSDroot mean square deviationTCEPtris2-carboxyethylphosphinebpbase pairsTEMEDN,N,N′,N′-tetramethylethylenediaminePDBProtein Data Burgers P. M. J., and Kunkel T. A. 2017 Eukaryotic DNA replication fork. Annu. Rev. Biochem. 86, 417–438 [PMC free article] [PubMed] [CrossRef] [Google Scholar]2. Pavlov Y. I., and Shcherbakova P. V. 2010 DNA polymerases at the eukaryotic fork 20 years later. Mutat. Res. 685, 45–53 [PMC free article] [PubMed] [CrossRef] [Google Scholar]3. Lujan S. A., Williams J. S., and Kunkel T. A. 2016 DNA polymerases divide the labor of genome replication. Trends Cell Biol. 26, 640–654 [PMC free article] [PubMed] [CrossRef] [Google Scholar]4. Baranovskiy A. G., and Tahirov T. H. 2017 Elaborated action of the human primosome. Genes Basel 8, E62 [PMC free article] [PubMed] [CrossRef] [Google Scholar]5. Núñez-Ramírez R., Klinge S., Sauguet L., Melero R., Recuero-Checa M. A., Kilkenny M., Perera R. L., García-Alvarez B., Hall R. J., Nogales E., Pellegrini L., and Llorca O. 2011 Flexible tethering of primase and DNA Pol α in the eukaryotic primosome. Nucleic Acids Res. 39, 8187–8199 [PMC free article] [PubMed] [CrossRef] [Google Scholar]6. Baranovskiy A. G., Babayeva N. D., Zhang Y., Gu J., Suwa Y., Pavlov Y. I., and Tahirov T. H. 2016 Mechanism of concerted RNA-DNA primer synthesis by the human primosome. J. Biol. Chem. 291, 10006–10020 [PMC free article] [PubMed] [CrossRef] [Google Scholar]7. Reijns M. A. M., Kemp H., Ding J., de Procé S. M., Jackson A. P., and Taylor M. S. 2015 Lagging-strand replication shapes the mutational landscape of the genome. Nature 518, 502–506 [PMC free article] [PubMed] [CrossRef] [Google Scholar]8. Starokadomskyy P., Gemelli T., Rios J. J., Xing C., Wang R. C., Li H., Pokatayev V., Dozmorov I., Khan S., Miyata N., Fraile G., Raj P., Xu Z., Xu Z., Ma L., et al. 2016 DNA polymerase-α regulates the activation of type I interferons through cytosolic RNADNA synthesis. Nat. Immunol. 17, 495–504 [PMC free article] [PubMed] [CrossRef] [Google Scholar]9. Han T., Goralski M., Capota E., Padrick S. B., Kim J., Xie Y., and Nijhawan D. 2016 The antitumor toxin CD437 is a direct inhibitor of DNA polymerase α. Nat. Chem. Biol. 12, 511–515 [PMC free article] [PubMed] [CrossRef] [Google Scholar]10. Pavlov Y. I., Shcherbakova P. V., and Rogozin I. B. 2006 Roles of DNA polymerases in replication, repair, and recombination in eukaryotes. Int. Rev. Cytol. 255, 41–132 [PubMed] [CrossRef] [Google Scholar]11. Klinge S., Núñez-Ramírez R., Llorca O., and Pellegrini L. 2009 3D architecture of DNA Pol α reveals the functional core of multi-subunit replicative polymerases. EMBO J. 28, 1978–1987 [PMC free article] [PubMed] [CrossRef] [Google Scholar]12. Suwa Y., Gu J., Baranovskiy A. G., Babayeva N. D., Pavlov Y. I., and Tahirov T. H. 2015 Crystal structure of the human Pol α B subunit in complex with the C-terminal domain of the catalytic subunit. J. Biol. Chem. 290, 14328–14337 [PMC free article] [PubMed] [CrossRef] [Google Scholar]13. Simon A. C., Zhou J. C., Perera R. L., van Deursen F., Evrin C., Ivanova M. E., Kilkenny M. L., Renault L., Kjaer S., Matak-Vinković D., Labib K., Costa A., and Pellegrini L. 2014 A Ctf4 trimer couples the CMG helicase to DNA polymerase α in the eukaryotic replisome. Nature 510, 293–297 [PMC free article] [PubMed] [CrossRef] [Google Scholar]14. Wang J., Sattar A. K., Wang C. C., Karam J. D., Konigsberg W. H., and Steitz T. A. 1997 Crystal structure of a pol α family replication DNA polymerase from bacteriophage RB69. Cell 89, 1087–1099 [PubMed] [CrossRef] [Google Scholar]15. Yang W., Lee J. Y., and Nowotny M. 2006 Making and breaking nucleic acids two-Mg2+-ion catalysis and substrate specificity. Mol. Cell 22, 5–13 [PubMed] [CrossRef] [Google Scholar]16. Perera R. L., Torella R., Klinge S., Kilkenny M. L., Maman J. D., and Pellegrini L. 2013 Mechanism for priming DNA synthesis by yeast DNA polymerase α. Elife 2, e00482 [PMC free article] [PubMed] [Google Scholar]17. Coloma J., Johnson R. E., Prakash L., Prakash S., and Aggarwal A. K. 2016 Human DNA polymerase α in binary complex with a DNADNA template-primer. Sci. Rep. 6, 23784 [PMC free article] [PubMed] [CrossRef] [Google Scholar]18. Baranovskiy A. G., Babayeva N. D., Suwa Y., Gu J., Pavlov Y. I., and Tahirov T. H. 2014 Structural basis for inhibition of DNA replication by aphidicolin. Nucleic Acids Res. 42, 14013–14021 [PMC free article] [PubMed] [CrossRef] [Google Scholar]19. Alberts I. L., Nadassy K., and Wodak S. J. 1998 Analysis of zinc binding sites in protein crystal structures. Protein Sci. 7, 1700–1716 [PMC free article] [PubMed] [CrossRef] [Google Scholar]20. Smith R. M., Martell A. E., and Chen Y. 1991 Critical evaluation of stability constants for nucleotide complexes with protons and metal ions and the accompanying enthalpy changes. Pure Appl. Chem. 63, 1015–1080 [CrossRef] [Google Scholar]21. Dawson R. M. C., Elliott D. C., Elliott W. H., and Jones K. M. 1986 Stability constants for metal complexes. in Data for Biochemical Research, 3rd Ed., pp. 399–415, Oxford Science Publications, Oxford, UK [Google Scholar]22. Hogg M., Osterman P., Bylund G. O., Ganai R. A., Lundström E. B., Sauer-Eriksson A. E., and Johansson E. 2014 Structural basis for processive DNA synthesis by yeast DNA polymerase varepsilon. Nat. Struct. Mol. Biol. 21, 49–55 [PubMed] [CrossRef] [Google Scholar]23. Ogawa M., Limsirichaikul S., Niimi A., Iwai S., Yoshida S., and Suzuki M. 2003 Distinct function of conserved amino acids in the fingers of Saccharomyces cerevisiae DNA polymerase α. J. Biol. Chem. 278, 19071–19078 [PubMed] [CrossRef] [Google Scholar]24. Dong Q., and Wang T. S. 1995 Mutational studies of human DNA polymerase α. Lysine 950 in the third most conserved region of α-like DNA polymerases is involved in binding the deoxynucleoside triphosphate. J. Biol. Chem. 270, 21563–21570 [PubMed] [CrossRef] [Google Scholar]25. Copeland W. C., Lam N. K., and Wang T. S. 1993 Fidelity studies of the human DNA polymerase α. The most conserved region among α-like DNA polymerases is responsible for metal-induced infidelity in DNA synthesis. J. Biol. Chem. 268, 11041–11049 [PubMed] [Google Scholar]26. Swan M. K., Johnson R. E., Prakash L., Prakash S., and Aggarwal A. K. 2009 Structural basis of high-fidelity DNA synthesis by yeast DNA polymerase delta. Nat. Struct. Mol. Biol. 16, 979–986 [PMC free article] [PubMed] [CrossRef] [Google Scholar]27. Franklin M. C., Wang J., and Steitz T. A. 2001 Structure of the replicating complex of a pol α family DNA polymerase. Cell 105, 657–667 [PubMed] [CrossRef] [Google Scholar]28. Kati W. M., Johnson K. A., Jerva L. F., and Anderson K. S. 1992 Mechanism and fidelity of HIV reverse transcriptase. J. Biol. Chem. 267, 25988–25997 [PubMed] [Google Scholar]29. Graves S. W., Johnson A. A., and Johnson K. A. 1998 Expression, purification, and initial kinetic characterization of the large subunit of the human mitochondrial DNA polymerase. Biochemistry 37, 6050–6058 [PubMed] [CrossRef] [Google Scholar]30. Patel S. S., Wong I., and Johnson K. A. 1991 Pre-steady-state kinetic analysis of processive DNA replication including complete characterization of an exonuclease-deficient mutant. Biochemistry 30, 511–525 [PubMed] [CrossRef] [Google Scholar]31. Zahurancik W. J., Klein S. J., and Suo Z. 2013 Kinetic mechanism of DNA polymerization catalyzed by human DNA polymerase epsilon. Biochemistry 52, 7041–7049 [PMC free article] [PubMed] [CrossRef] [Google Scholar]32. Dieckman L. M., Johnson R. E., Prakash S., and Washington M. T. 2010 Pre-steady state kinetic studies of the fidelity of nucleotide incorporation by yeast DNA polymerase delta. Biochemistry 49, 7344–7350 [PMC free article] [PubMed] [CrossRef] [Google Scholar]33. Garcia-Diaz M., Bebenek K., Krahn J. M., Kunkel T. A., and Pedersen L. C. 2005 A closed conformation for the Pol lambda catalytic cycle. Nat. Struct. Mol. Biol. 12, 97–98 [PubMed] [CrossRef] [Google Scholar]34. Fiala K. A., Abdel-Gawad W., and Suo Z. 2004 Pre-steady-state kinetic studies of the fidelity and mechanism of polymerization catalyzed by truncated human DNA polymerase λ. Biochemistry 43, 6751–6762 [PubMed] [CrossRef] [Google Scholar]35. Johnson K. A., Simpson Z. B., and Blom T. 2009 Global kinetic explorer a new computer program for dynamic simulation and fitting of kinetic data. Anal. Biochem. 387, 20–29 [PubMed] [CrossRef] [Google Scholar]36. Copeland W. C., and Wang T. S. 1993 Mutational analysis of the human DNA polymerase α the most conserved region in α-like DNA polymerases is involved in metal-specific catalysis. J. Biol. Chem. 268, 11028–11040 [PubMed] [Google Scholar]37. Pavlov Y. I., Frahm C., Nick McElhinny S. A., Niimi A., Suzuki M., and Kunkel T. A. 2006 Evidence that errors made by DNA polymerase α are corrected by DNA polymerase δ. Curr. Biol. 16, 202–207 [PubMed] [CrossRef] [Google Scholar]38. Dong Q., Copeland W. C., and Wang T. S. 1993 Mutational studies of human DNA polymerase α identification of residues critical for deoxynucleotide binding and misinsertion fidelity of DNA synthesis. J. Biol. Chem. 268, 24163–24174 [PubMed] [Google Scholar]39. Sabouri N., Viberg J., Goyal D. K., Johansson E., and Chabes A. 2008 Evidence for lesion bypass by yeast replicative DNA polymerases during DNA damage. Nucleic Acids Res. 36, 5660–5667 [PMC free article] [PubMed] [CrossRef] [Google Scholar]40. Gandhi V. V., and Samuels D. C. 2011 A review comparing deoxyribonucleoside triphosphate dNTP concentrations in the mitochondrial and cytoplasmic compartments of normal and transformed cells. Nucleosides Nucleotides Nucleic Acids 30, 317–339 [PMC free article] [PubMed] [CrossRef] [Google Scholar]41. Kunkel T. A., Roberts J. D., and Sugino A. 1991 The fidelity of DNA synthesis by the catalytic subunit of yeast DNA polymerase α alone and with accessory proteins. Mutat. Res. 250, 175–182 [PubMed] [CrossRef] [Google Scholar]42. Jain R., Rajashankar K. R., Buku A., Johnson R. E., Prakash L., Prakash S., and Aggarwal A. K. 2014 Crystal structure of yeast DNA polymerase ϵ catalytic domain. PLoS One 9, e94835 [PMC free article] [PubMed] [CrossRef] [Google Scholar]43. Baranovskiy A. G., Gu J., Babayeva N. D., Kurinov I., Pavlov Y. I., and Tahirov T. H. 2017 Crystal structure of the human Pol B-subunit in complex with the C-terminal domain of the catalytic subunit. J. Biol. Chem. 292, 15717–15730 [PMC free article] [PubMed] [CrossRef] [Google Scholar]44. Tanaka S., Cao K., Niimi A., Limsirichaikul S., Miao H. Q., Nakamura N., Murate T., Hasegawa Y., Takahashi T., and Suzuki M. 2010 Functions of base selection step in human DNA polymerase α. DNA Repair 9, 534–541 [PubMed] [CrossRef] [Google Scholar]45. Freudenthal B. D., Beard W. A., and Wilson S. H. 2012 Structures of dNTP intermediate states during DNA polymerase active site assembly. Structure 20, 1829–1837 [PMC free article] [PubMed] [CrossRef] [Google Scholar]46. Fujiwara T., and Shindo H. 1985 Phosphorus-31 nuclear magnetic resonance of highly oriented DNA fibers. 2. Molecular motions in hydrated DNA. Biochemistry 24, 896–902 [PubMed] [CrossRef] [Google Scholar]47. Arnott S., Chandrasekaran R., Millane R. P., and Park H. S. 1986 DNA-RNA hybrid secondary structures. J. Mol. Biol. 188, 631–640 [PubMed] [CrossRef] [Google Scholar]48. Priyakumar U. D., and Mackerell A. D. Jr. 2008 Atomic detail investigation of the structure and dynamics of hybrids a molecular dynamics study. J. Phys. Chem. B 112, 1515–1524 [PMC free article] [PubMed] [CrossRef] [Google Scholar]49. Lu X. J., Shakked Z., and Olson W. K. 2000 A-form conformational motifs in ligand-bound DNA structures. J. Mol. Biol. 300, 819–840 [PubMed] [CrossRef] [Google Scholar]50. Lee S. H., Eki T., and Hurwitz J. 1989 Synthesis of DNA containing the simian virus 40 origin of replication by the combined action of DNA polymerases α and δ. Proc. Natl. Acad. Sci. 86, 7361–7365 [PMC free article] [PubMed] [CrossRef] [Google Scholar]51. Göksenin A. Y., Zahurancik W., LeCompte K. G., Taggart D. J., Suo Z., and Pursell Z. F. 2012 Human DNA polymerase ϵ is able to efficiently extend from multiple consecutive ribonucleotides. J. Biol. Chem. 287, 42675–42684 [PMC free article] [PubMed] [CrossRef] [Google Scholar]52. Shimizu K., Hashimoto K., Kirchner J. M., Nakai W., Nishikawa H., Resnick M. A., and Sugino A. 2002 Fidelity of DNA polymerase epsilon holoenzyme from budding yeast Saccharomyces cerevisiae. J. Biol. Chem. 277, 37422–37429 [PubMed] [CrossRef] [Google Scholar]53. Zinnen S., Hsieh J. C., and Modrich P. 1994 Misincorporation and mispaired primer extension by human immunodeficiency virus reverse transcriptase. J. Biol. Chem. 269, 24195–24202 [PubMed] [Google Scholar]54. Mislak A. C., and Anderson K. S. 2016 Insights into the molecular mechanism of polymerization and nucleoside reverse transcriptase inhibitor incorporation by human PrimPol. Antimicrob. Agents Chemother. 60, 561–569 [PMC free article] [PubMed] [CrossRef] [Google Scholar]55. Zafar M. K., Ketkar A., Lodeiro M. F., Cameron C. E., and Eoff R. L. 2014 Kinetic analysis of human PrimPol DNA polymerase activity reveals a generally error-prone enzyme capable of accurately bypassing 7,8-dihydro-8-oxo-2′-deoxyguanosine. Biochemistry 53, 6584–6594 [PMC free article] [PubMed] [CrossRef] [Google Scholar]56. Niimi A., Limsirichaikul S., Yoshida S., Iwai S., Masutani C., Hanaoka F., Kool E. T., Nishiyama Y., and Suzuki M. 2004 Palm mutants in DNA polymerases α and η alter DNA replication fidelity and translesion activity. Mol. Cell Biol. 24, 2734–2746 [PMC free article] [PubMed] [CrossRef] [Google Scholar]57. Venkatesan R. N., Hsu J. J., Lawrence N. A., Preston B. D., and Loeb L. A. 2006 Mutator phenotypes caused by substitution at a conserved motif A residue in eukaryotic DNA polymerase δ. J. Biol. Chem. 281, 4486–4494 [PubMed] [CrossRef] [Google Scholar]58. Zhang Y., Baranovskiy A. G., Tahirov E. T., Tahirov T. H., and Pavlov Y. I. 2016 Divalent ions attenuate DNA synthesis by human DNA polymerase α by changing the structure of the template/primer or by perturbing the polymerase reaction. DNA Repair Amst. 43, 24–33 [PMC free article] [PubMed] [CrossRef] [Google Scholar]59. Zhang Y., Baranovskiy A. G., Tahirov T. H., and Pavlov Y. I. 2014 The C-terminal domain of the DNA polymerase catalytic subunit regulates the primase and polymerase activities of the human DNA polymerase α-primase complex. J. Biol. Chem. 289, 22021–22034 [PMC free article] [PubMed] [CrossRef] [Google Scholar]60. Thompson H. C., and Kuchta R. D. 1995 Arabinofuranosyl nucleotides are not chain-terminators during initiation of new strands of DNA by DNA polymerase α-primase. Biochemistry 34, 11198–11203 [PubMed] [CrossRef] [Google Scholar]61. Maga G., Stucki M., Spadari S., and Hübscher U. 2000 DNA polymerase switching I. Replication factor C displaces DNA polymerase alpha prior to PCNA loading. J. Mol. Biol. 295, 791–801 [PubMed] [CrossRef] [Google Scholar]62. Baranovskiy A. G., Zhang Y., Suwa Y., Gu J., Babayeva N. D., Pavlov Y. I., and Tahirov T. H. 2016 Insight into the human DNA primase interaction with template-primer. J. Biol. Chem. 291, 4793–4802 [PMC free article] [PubMed] [CrossRef] [Google Scholar]63. Kilkenny M. L., Simon A. C., Mainwaring J., Wirthensohn D., Holzer S., and Pellegrini L. 2017 The human CTF4-orthologue AND-1 interacts with DNA polymerase α/primase via its unique C-terminal HMG box. Open Biol. 7, 170217 [PMC free article] [PubMed] [CrossRef] [Google Scholar]64. Sengupta S., van Deursen F., de Piccoli G., and Labib K. 2013 Dpb2 integrates the leading-strand DNA polymerase into the eukaryotic replisome. Curr. Biol. 23, 543–552 [PubMed] [CrossRef] [Google Scholar]65. Otwinowski Z., and Minor W. 1997 Processing of x-ray diffraction data collected in oscillation mode. Methods Enzymol. 276, 307–326 [PubMed] [CrossRef] [Google Scholar]66. Adams P. D., Afonine P. V., Bunkóczi G., Chen V. B., Davis I. W., Echols N., Headd J. J., Hung L. W., Kapral G. J., Grosse-Kunstleve R. W., McCoy A. J., Moriarty N. W., Oeffner R., Read R. J., Richardson D. C., et al. 2010 PHENIX a comprehensive Python-based system for macromolecular structure solution. Acta Crystallogr. D Biol. Crystallogr. 66, 213–221 [PMC free article] [PubMed] [CrossRef] [Google Scholar]67. Brünger A. T., Adams P. D., Clore G. M., DeLano W. L., Gros P., Grosse-Kunstleve R. W., Jiang J. S., Kuszewski J., Nilges M., Pannu N. S., Read R. J., Rice L. M., Simonson T., and Warren G. L. 1998 Crystallography & NMR system a new software suite for macromolecular structure determination. Acta Crystallogr. D Biol. Crystallogr. 54, 905–921 [PubMed] [CrossRef] [Google Scholar]68. Johnson K. A., Simpson Z. B., and Blom T. 2009 FitSpace explorer an algorithm to evaluate multidimensional parameter space in fitting kinetic data. Anal. Biochem. 387, 30–41 [PubMed] [CrossRef] [Google Scholar]69. Lu X. J., and Olson W. K. 2003 3DNA a software package for the analysis, rebuilding and visualization of three-dimensional nucleic acid structures. Nucleic Acids Res. 31, 5108–5121 [PMC free article] [PubMed] [CrossRef] [Google Scholar]Articles from The Journal of Biological Chemistry are provided here courtesy of American Society for Biochemistry and Molecular Biology