DNA polymerases were named for their function of catalysing DNA replication, a process that is necessary for growth and propagation of life. DNA involving Watson–Crick base-pairing can be synthesized with high fidelity, the structural and mechanistic origins of which have been investigated for many decades. Despite this, new chemical insights continue to be uncovered, including recent findings that may explain newly discovered functions for many DNA polymerases in DNA repair and mutation. Some of these reactions involve non-Watson–Crick base-pairing. In addition, certain DNA polymerases have been engineered for a wide variety of applications in biotechnology and biomedicine. This Review describes the molecular basis for the diverse and contrasting functions of different DNA polymerases, providing an up-to-date understanding of how these tasks are accomplished and the means by which we can benefit from them.
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Lehman, I. R., Bessman, M. J., Simms, E. S. & Kornberg, A. Enzymatic synthesis of deoxyribonucleic acid. I. Preparation of substrates and partial purification of an enzyme from Escherichia coli.. J. Biol. Chem. 233, 163–170 (1958).
Bessman, M. J., Lehman, I. R., Simms, E. S. & Kornberg, A. Enzymatic synthesis of deoxyribonucleic acid. II. General properties of the reaction. J. Biol. Chem. 233, 171–177 (1958).
Lehman, I. R. et al. Enzymatic synthesis of deoxyribonucleic acid. V. Chemical composition of enzymatically synthesized deoxyribonucleic acid. Proc. Natl Acad. Sci. USA 44, 1191–1196 (1958).
Hübscher, U., Spadari, S., Villani, G. & Maga, G. DNA Polymerases: Discovery, Characterization, and Functions in Cellular DNA Transactions 1st edn (World Scientific, 2010). A comprehensive reference in which detailed biochemical and functional information can be found.
Jaszczur, M. et al. Mutations for worse or better: low-fidelity DNA synthesis by SOS DNA polymerase V is a tightly regulated double-edged sword. Biochemistry 55, 2309–2318 (2016).
Zhao, Y. et al. Mechanism of somatic hypermutation at the WA motif by human DNA polymerase η. Proc. Natl Acad. Sci. USA 110, 8146–8151 (2013).
Kunkel, T. A. DNA replication fidelity. J. Biol. Chem. 279, 16895–16898 (2004).
Showalter, A. K. & Tsai, M.-D. A. DNA polymerase with specificity for five base pairs. J. Am. Chem. Soc. 123, 1776–1777 (2001).
Ahn, J., Werneburg, B. G. & Tsai, M.-D. DNA polymerase β: structure−fidelity relationship from pre-steady-state kinetic analyses of all possible correct and incorrect base pairs for wild type and R283A mutant. Biochemistry 36, 1100–1107 (1997).
Steitz, T. A. DNA polymerases: structural diversity and common mechanisms. J. Biol. Chem. 274, 17395–17398 (1999).
Beard, W. A. & Wilson, S. H. Structure and mechanism of DNA polymerase β. Biochemistry 53, 2768–2780 (2014). Reviews the research on the structure and mechanism of Pol β up until 2014.
Sawaya, M. R., Pelletier, H., Kumar, A., Wilson, S. H. & Kraut, J. Crystal structure of rat DNA polymerase β: evidence for a common polymerase mechanism. Science 264, 1930–1935 (1994).
Gridley, C. L. et al. Structural changes in the hydrophobic hinge region adversely affect the activity and fidelity of the I260Q mutator DNA polymerase β. Biochemistry 52, 4422–4432 (2013).
Sawaya, M. R., Prasad, R., Wilson, S. H., Kraut, J. & Pelletier, H. Crystal structures of human DNA polymerase β complexed with gapped and nicked DNA: evidence for an induced fit mechanism. Biochemistry 36, 11205–11215 (1997).
Beard, W. A., Shock, D. D., Batra, V. K., Pedersen, L. C. & Wilson, S. H. DNA polymerase β substrate specificity: side chain modulation of the “A-rule”. J. Biol. Chem. 284, 31680–31689 (2009).
Freudenthal, B. D., Beard, W. A. & Wilson, S. H. Structures of dNTP intermediate states during DNA polymerase active site assembly. Structure 20, 1829–1837 (2012).
Pelletier, H., Sawaya, M. R., Kumar, A., Wilson, S. H. & Kraut, J. Structures of ternary complexes of rat DNA polymerase β, a DNA template-primer and ddCTP. Science 264, 1891–1903 (1994).
Batra, V. K. et al. Magnesium-induced assembly of a complete DNA polymerase catalytic complex. Structure 14, 757–766 (2006).
Freudenthal, B. D., Beard, W. A., Shock, D. D. & Wilson, S. H. Observing a DNA polymerase choose right from wrong. Cell 154, 157–168 (2013). Time-resolved crystallography enabled the first direct comparison of structural intermediates involved in the incorporation of matched and mismatched dNTP.
Arndt, J. W. et al. Insight into the catalytic mechanism of DNA polymerase β: structures of intermediate complexes. Biochemistry 40, 5368–5375 (2001). The authors of this study show that it is metal B (in this case Cr(III)dNTP) alone that, in addition to inducing rapid conformational change, also induces closure of the N subdomain. This work provides the first direct evidence against conformational closure being rate limiting.
Batra, V. K., Beard, W. A., Shock, D. D., Pedersen, L. C. & Wilson, S. H. Structures of DNA polymerase β with active-site mismatches suggest a transient abasic site intermediate during misincorporation. Mol. Cell 30, 315–324 (2008).
Krahn, J. M., Beard, W. A. & Wilson, S. H. Structural insights into DNA polymerase β deterrents for misincorporation support an induced-fit mechanism for fidelity. Structure 12, 1823–1832 (2004).
Reed, A. J., Vyas, R., Raper, A. T. & Suo, Z. Structural insights into the post-chemistry steps of nucleotide incorporation catalyzed by a DNA polymerase. J. Am. Chem. Soc. 139, 465–471 (2017).
Kim, K. H. et al. Direct observation of bond formation in solution with femtosecond X-ray scattering. Nature 518, 385–389 (2015).
Florián, J., Goodman, M. F. & Warshel, A. Computer simulations of protein functions: searching for the molecular origin of the replication fidelity of DNA polymerases. Proc. Natl Acad. Sci. USA 102, 6819–6824 (2005).
Lin, P. et al. Energy analysis of chemistry for correct insertion by DNA polymerase β. Proc. Natl Acad. Sci. USA 103, 13294–13299 (2006).
Wang, L., Broyde, S. & Zhang, Y. Polymerase-tailored variations in the water-mediated and substrate-assisted mechanism for nucleotidyl transfer: insights from a study of T7 DNA polymerase. J. Mol. Biol. 389, 787–796 (2009).
Lior-Hoffmann L. et al. Preferred WMSA catalytic mechanism of the nucleotidyl transfer reaction in human DNA polymerase κ elucidates error-free bypass of a bulky DNA lesion. Nucleic Acids Res. 40, 9193–9205 (2012).
Li, Y., Freudenthal, B. D., Beard, W. A., Wilson, S. H. & Schlick, T. Optimal and variant metal-ion routes in DNA polymerase β's conformational pathways. J. Am. Chem. Soc. 136, 3630–3639 (2014).
Nakamura, T., Zhao, Y., Yamagata, Y., Hua, Y. J. & Yang, W. Watching DNA polymerase η make a phosphodiester bond. Nature 487, 196–201 (2012). A breakthrough in studying phosphodiester bond formation was achieved with the use of a revised method of time-resolved X-ray crystallography. Monitoring this reaction revealed the role of a third metal ion in DNA synthesis.
Brautigam, C. A. & Steitz, T. A. Structural and functional insights provided by crystal structures of DNA polymerases and their substrate complexes. Curr. Opin. Struct. Biol. 8, 54–63 (1998).
Biertümpfel, C. et al. Structure and mechanism of human DNA polymerase η. Nature 465, 1044–1048 (2010).
Batra, V. K. et al. Amino acid substitution in the active site of DNA polymerase β explains the energy barrier of the nucleotidyl transfer reaction. J. Am. Chem. Soc. 135, 8078–8088 (2013).
Yang, W., Lee, J. Y. & Nowotny, M. Making and breaking nucleic acids: two-Mg2+-ion catalysis and substrate specificity. Mol. Cell 22, 5–13 (2006).
Gao, Y. & Yang, W. Capture of a third Mg2+ is essential for catalyzing DNA synthesis. Science 352, 1334–1337 (2016). This work used time-resolved crystallography to observe an intermediate, possibly near-TS structure involving a third metal ion during DNA synthesis.
Yang, W., Weng, P. J. & Gao, Y. A new paradigm of DNA synthesis: three-metal-ion catalysis. Cell Biosci. 6, 51 (2016).
Genna, V., Vidossich, P., Ippoliti, E., Carloni, P. & De Vivo, M. A self-activated mechanism for nucleic acid polymerization catalyzed by DNA/RNA polymerases. J. Am. Chem. Soc. 138, 14592–14598 (2016). Analysis of crystal structures enabled identification of a common intramolecular hydrogen bond between the 3′-OH and β-phosphate of incoming dNTP or rNTP. This interaction is proposed to assist activation of the nucleophilic 3′-OH.
Freudenthal, B. D. et al. Uncovering the polymerase-induced cytotoxicity of an oxidized nucleotide. Nature 517, 635–639 (2015). Describes a time-resolved crystallographic study showing that 8-oxo-dGTP is preferably incorporated opposite to a template dA. This reaction, which leads to mutation and cancer, is facilitated by anti–syn Hoogsteen base-pairing and the presence of a third metal ion.
Vyas, R., Reed, A. J., Tokarsky, E. J. & Suo, Z. Viewing human DNA polymerase β faithfully and unfaithfully bypass an oxidative lesion by time-dependent crystallography. J. Am. Chem. Soc. 137, 5225–5230 (2015). This study mirrors the study undertaken in reference 38 by showing that the preferred incorporation of dATP opposite to a 8-oxo-dG lesion is facilitated by 8-oxo-dG–dATP syn–anti Hoogsteen base-pairing as well as the third metal ion.
Perera, L. et al. Requirement for transient metal ions revealed through computational analysis for DNA polymerase going in reverse. Proc. Natl Acad. Sci. USA 112, E5228–E5236 (2015).
Perera, L., Freudenthal, B. D., Beard, W. A., Pedersen, L. G. & Wilson, S. H. Revealing the role of the product metal in DNA polymerase β catalysis. Nucleic Acids Res. 45, 2736–2745 (2017).
Zhong, X., Patel, S. S. & Tsai, M.-D. DNA polymerase β. 5. Dissecting the functional roles of the two metal ions with Cr(III)dTTP1. J. Am. Chem. Soc. 120, 235–236 (1998).
Zhong, X., Patel, S. S., Werneburg, B. G. & Tsai, M.-D. DNA polymerase β: multiple conformational changes in the mechanism of catalysis. Biochemistry 36, 11891–11900 (1997).
Dunlap, C. A. & Tsai, M.-D. Use of 2-aminopurine and tryptophan fluorescence as probes in kinetic analyses of DNA polymerase β. Biochemistry 41, 11226–11235 (2002). The activation constant of Mg2+ is found to be substantially higher than that of MgdNTP; this work thus provides functional support for the additional low-affinity metal-binding site now known to be occupied by metal C.
Bakhtina, M. et al. Use of viscogens, dNTPαS, and rhodium(III) as probes in stopped-flow experiments to obtain new evidence for the mechanism of catalysis by DNA polymerase β. Biochemistry 44, 5177–5187 (2005).
Garcia-Diaz, M., Bebenek, K., Kunkel, T. A. & Blanco, L. Identification of an intrinsic 5′-deoxyribose-5-phosphate lyase activity in human DNA polymerase λ: a possible role in base excision repair. J. Biol. Chem. 276, 34659–34663 (2001).
Garcia-Diaz, M., Bebenek, K., Krahn, J. M., Pedersen, L. C. & Kunkel, T. A. Role of the catalytic metal during polymerization by DNA polymerase lambda. DNA Repair 6, 1333–1340 (2007).
Dominguez, O. et al. DNA polymerase μ (Pol μ), homologous to TdT, could act as a DNA mutator in eukaryotic cells. EMBO J. 19, 1731–1742 (2000).
Moon, A. F. et al. Structural insight into the substrate specificity of DNA Polymerase μ. Nat. Struct. Mol. Biol. 14, 45–53 (2007).
Andrade, P., Martín, M. J., Juárez, R., López de Saro, F. & Blanco, L. Limited terminal transferase in human DNA polymerase μ defines the required balance between accuracy and efficiency in NHEJ. Proc. Natl Acad. Sci. USA 106, 16203–16208 (2009).
Moon, A. F. et al. Sustained active site rigidity during synthesis by human DNA polymerase μ. Nat. Struct. Mol. Biol. 21, 253–260 (2014).
Liu, M.-S. et al. Structural mechanism for the fidelity modulation of DNA polymerase λ. J. Am. Chem. Soc. 138, 2389–2398 (2016).
Vashishtha, A. K., Wang, J. & Konigsberg, W. H. Different divalent cations alter the kinetics and fidelity of DNA polymerases. J. Biol. Chem. 291, 20869–20875 (2016).
Gardner, A. F., Joyce, C. M. & Jack, W. E. Comparative kinetics of nucleotide analog incorporation by vent DNA polymerase. J. Biol. Chem. 279, 11834–11842 (2004).
Vyas, R., Zahurancik, W. J. & Suo, Z. Structural basis for the binding and incorporation of nucleotide analogs with l-stereochemistry by human DNA polymerase λ. Proc. Natl Acad. Sci. USA 111, E3033–E3042 (2014).
Arana, M. E., Potapova, O., Kunkel, T. A. & Joyce, C. M. Kinetic analysis of the unique error signature of human DNA polymerase ν. Biochemistry 50, 10126–10135 (2011).
Gowda, A. S. P., Moldovan, G.-L. & Spratt, T. E. Human DNA polymerase ν catalyzes correct and incorrect DNA synthesis with high catalytic efficiency. J. Biol. Chem. 290, 16292–16303 (2015).
Lee, Y.-S., Gao, Y. & Yang, W. How a homolog of high-fidelity replicases conducts mutagenic DNA synthesis. Nat. Struct. Mol. Biol. 22, 298–303 (2015).
Arana, M. E., Seki, M., Wood, R. D., Rogozin, I. B. & Kunkel, T. A. Low-fidelity DNA synthesis by human DNA polymerase theta. Nucleic Acids Res. 36, 3847–3856 (2008).
Takata, K.-i., Shimizu, T., Iwai, S. & Wood, R. D. Human DNA polymerase N (POLN) is a low fidelity enzyme capable of error-free bypass of 5S-thymine glycol. J. Biol. Chem. 281, 23445–23455 (2006).
Wood, R. D. & Doublié, S. DNA polymerase θ (POLQ), double-strand break repair, and cancer. DNA Repair 44, 22–32 (2016).
Loeb, L. A. & Monnat, R. J. DNA polymerases and human disease. Nat. Rev. Genet. 9, 594–604 (2008).
Tsai, M.-D. How DNA polymerases catalyze DNA replication, repair, and mutation. Biochemistry 53, 2749–2751 (2014).
Lee, H. R., Helquist, S. A., Kool, E. T. & Johnson, K. A. Importance of hydrogen bonding for efficiency and specificity of the human mitochondrial DNA polymerase. J. Biol. Chem. 283, 14402–14410 (2008).
Winnacker, M. & Kool, E. T. Artificial genetic sets composed of size-expanded base pairs. Angew. Chem. Int. Ed. 52, 12498–12508 (2013).
Chen, T., Hongdilokkul, N., Liu, Z., Thirunavukarasu, D. & Romesberg, F. E. The expanding world of DNA and RNA. Curr. Opin. Chem. Biol. 34, 80–87 (2016).
Hirao, I. & Kimoto, M. Unnatural base pair systems toward the expansion of the genetic alphabet in the central dogma. Proc. Jpn Acad. Ser. B 88, 345–367 (2012).
Benner, S. A. et al. Alternative Watson–Crick synthetic genetic systems. Cold Spring Harb. Perspect. Biol.http://dx.doi.org/10.1101/cshperspect.a023770 (2016).
Oertell, K. et al. Kinetic selection versus free energy of DNA base pairing in control of polymerase fidelity. Proc. Natl Acad. Sci. USA 113, E2277–E2285 (2016).
Showalter, A. K. & Tsai, M.-D. A reexamination of the nucleotide incorporation fidelity of DNA polymerases. Biochemistry 41, 10571–10576 (2002). The authors of this article posit, for the first time, that the main step that controls the fidelity of DNA polymerase catalysis should be the chemical step.
Showalter, A. K. et al. Mechanistic comparison of high-fidelity and error-prone DNA polymerases and ligases involved in DNA repair. Chem. Rev. 106, 340–360 (2006).
Bakhtina, M., Roettger, M. P., Kumar, S. & Tsai, M.-D. A unified kinetic mechanism applicable to multiple DNA polymerases. Biochemistry 46, 5463–5472 (2007).
Sucato, C. A. et al. Modifying the β, γ leaving-group bridging oxygen alters nucleotide incorporation efficiency, fidelity, and the catalytic mechanism of DNA polymerase β. Biochemistry 46, 461–471 (2007).
Sucato, C. A. et al. DNA polymerase β fidelity: halomethylene-modified leaving groups in pre-steady-state kinetic analysis reveal differences at the chemical transition state. Biochemistry 47, 870–879 (2008).
Bakhtina, M., Roettger, M. P. & Tsai, M.-D. Contribution of the reverse rate of the conformational step to polymerase β fidelity. Biochemistry 48, 3197–3208 (2009).
Balbo, P. B., Wang, E. C.-W. & Tsai, M.-D. Kinetic mechanism of active site assembly and chemical catalysis of DNA polymerase β. Biochemistry 50, 9865–9875 (2011).
Oertell, K. et al. Transition state in DNA polymerase β catalysis: rate-limiting chemistry altered by base-pair configuration. Biochemistry 53, 1842–1848 (2014).
Kellinger, M. W. & Johnson, K. A. Nucleotide-dependent conformational change governs specificity and analog discrimination by HIV reverse transcriptase. Proc. Natl Acad. Sci. USA 107, 7734–7739 (2010).
Fiala, K. A. & Suo, Z. Mechanism of DNA polymerization catalyzed by Sulfolobus solfataricus P2 DNA polymerase IV. Biochemistry 43, 2116–2125 (2004).
Rothwell, P. J., Mitaksov, V. & Waksman, G. Motions of the fingers subdomain of Klentaq1 are fast and not rate limiting: implications for the molecular basis of fidelity in DNA polymerases. Mol. Cell 19, 345–355 (2005).
Johnson, K. A. The kinetic and chemical mechanism of high-fidelity DNA polymerases. Biochim. Biophys. Acta 1804, 1041–1048 (2010).
Watson, J. D. & Crick, F. H. C. Genetical implications of the structure of deoxyribonucleic acid. Nature 171, 964–967 (1953).
Watson, J. D. & Crick, F. H. C. Molecular structure of nucleic acids: a structure for deoxyribose nucleic acid. Nature 171, 737–738 (1953).
Wang, W., Hellinga, H. W. & Beese, L. S. Structural evidence for the rare tautomer hypothesis of spontaneous mutagenesis. Proc. Natl Acad. Sci. USA 108, 17644–17648 (2011).
Yu, H., Eritja, R., Bloom, L. B. & Goodman, M. F. Ionization of bromouracil and fluorouracil stimulates base mispairing frequencies with guanine. J. Biol. Chem. 268, 15935–15943 (1993).
Bebenek, K., Pedersen, L. C. & Kunkel, T. A. Replication infidelity via a mismatch with Watson–Crick geometry. Proc. Natl Acad. Sci. USA 108, 1862–1867 (2011).
Nikolova, E. N. et al. Transient Hoogsteen base pairs in canonical duplex DNA. Nature 470, 498–502 (2011).
Alvey, H. S., Gottardo, F. L., Nikolova, E. N. & Al-Hashimi, H. M. Widespread transient Hoogsteen base pairs in canonical duplex DNA with variable energetics. Nat. Commum. 5, 4786 (2014).
Zhou, H. et al. New insights into Hoogsteen base pairs in DNA duplexes from a structure-based survey. Nucleic Acids Res. 43, 3420–3433 (2015).
Kimsey, I. J., Petzold, K., Sathyamoorthy, B., Stein, Z. W. & Al-Hashimi, H. M. Visualizing transient Watson–Crick-like mispairs in DNA and RNA duplexes. Nature 519, 315–320 (2015).
Sirover, M. & Loeb, L. Infidelity of DNA synthesis in vitro: screening for potential metal mutagens or carcinogens. Science 194, 1434–1436 (1976).
Weymouth, L. A. & Loeb, L. A. Mutagenesis during in vitro DNA synthesis. Proc. Natl Acad. Sci. USA 75, 1924–1928 (1978).
Werneburg, B. G. et al. DNA polymerase β: pre-steady-state kinetic analysis and roles of arginine-283 in catalysis and fidelity. Biochemistry 35, 7041–7050 (1996).
Vaisman, A., Ling, H., Woodgate, R. & Yang, W. Fidelity of Dpo4: effect of metal ions, nucleotide selection and pyrophosphorolysis. EMBO J. 24, 2957–2967 (2005).
Frank, E. G. & Woodgate, R. Increased catalytic activity and altered fidelity of human DNA polymerase ι in the presence of manganese. J. Biol. Chem. 282, 24689–24696 (2007).
Bebenek, K. et al. Substrate-induced DNA strand misalignment during catalytic cycling by DNA polymerase λ. EMBO Rep. 9, 459–464 (2008).
Koag, M.-C. & Lee, S. Metal-dependent conformational activation explains highly promutagenic replication across O6-methylguanine by human DNA polymerase β. J. Am. Chem. Soc. 136, 5709–5721 (2014).
Choi, J.-Y. et al. Kinetic and structural impact of metal ions and genetic variations on human DNA polymerase ι. J. Biol. Chem. 291, 21063–21073 (2016).
Vashishtha, A. K. & Konigsberg, W. H. Effect of different divalent cations on the kinetics and fidelity of RB69 DNA polymerase. Biochemistry 55, 2661–2670 (2016).
Xia, S. & Konigsberg, W. H. RB69 DNA polymerase structure, kinetics, and fidelity. Biochemistry 53, 2752–2767 (2014).
Xia, S., Wang, J. & Konigsberg, W. H. DNA mismatch synthesis complexes provide insights into base selectivity of a B family DNA polymerase. J. Am. Chem. Soc. 135, 193–202 (2013).
Oliveros, M. et al. Characterization of an African swine fever virus 20-kDa DNA polymerase involved in DNA repair. J. Biol. Chem. 272, 30899–30910 (1997).
Maciejewski, M. W. et al. Solution structure of a viral DNA repair polymerase. Nat. Struct. Mol. Biol. 8, 936–941 (2001).
Showalter, A. K., Byeon, I.-J. L., Su, M.-I. & Tsai, M.-D. Solution structure of a viral DNA polymerase X and evidence for a mutagenic function. Nat. Struct. Biol. 8, 942–946 (2001).
Kumar, S., Bakhtina, M. & Tsai, M.-D. Altered order of substrate binding by DNA polymerase X from African swine fever virus. Biochemistry 47, 7875–7887 (2008).
Wu, W.-J. et al. How a low-fidelity DNA polymerase chooses non-Watson–Crick from Watson–Crick incorporation. J. Am. Chem. Soc. 136, 4927–4937 (2014). This study uses NMR structural determination to demonstrate how a mutagenic DNA polymerase achieves its low fidelity by overcoming the forces that govern Watson–Crick base-pairing.
Chen Y. et al. Unique 5′-P recognition and basis for dG:dGTP misincorporation of ASFV DNA polymerase X. PLoS Biol. 15, http://dx.doi.org/10.1371/journal.pbio.1002599 (2017).
García-Escudero, R., García-Díaz, M., Salas, M. L., Blanco, L. & Salas, J. DNA polymerase X of African swine fever virus: insertion fidelity on gapped DNA substrates and AP lyase activity support a role in base excision repair of viral DNA. J. Mol. Biol. 326, 1403–1412 (2003).
Bebenek, K., Pedersen, L. C. & Kunkel, T. A. Structure–function studies of DNA polymerase λ. Biochemistry 53, 2781–2792 (2014).
Nelson, J. R., Lawrence, C. W. & Hinkle, D. C. Deoxycytidyl transferase activity of yeast Rev1 protein. Nature 382, 729–731 (1996).
Haracska, L., Prakash, S. & Prakash, L. Yeast Rev1 protein is a G template-specific DNA polymerase. J. Biol. Chem. 277, 15546–15551 (2002).
Nair, D. T., Johnson, R. E., Prakash, L., Prakash, S. & Aggarwal, A. K. Rev1 employs a novel mechanism of DNA synthesis using a protein template. Science 309, 2219–2222 (2005).
Fiala, K. A., Abdel-Gawad, W. & Suo, Z. Pre-steady-state kinetic studies of the fidelity and mechanism of polymerization catalyzed by truncated human DNA polymerase λ. Biochemistry 43, 6751–6762 (2004).
Ahn, J., Kraynov, V. S., Zhong, X., Werneburg, B. G. & Tsai, M.-D. DNA polymerase β: effects of gapped DNA substrates on dNTP specificity, fidelity, processivity and conformational changes. Biochem. J. 331, 79–87 (1998).
Johnson, K. A. Conformational coupling in DNA polymerase fidelity. Annu. Rev. Biochem. 62, 685–713 (1993).
Johnson, S. J. & Beese, L. S. Structures of mismatch replication errors observed in a DNA polymerase. Cell 116, 803–816 (2004).
Tang, K.-H. et al. Mismatched dNTP incorporation by DNA polymerase β does not proceed via globally different conformational pathways. Nucleic Acids Res. 36, 2948–2957 (2008).
Moscato, B., Swain, M. & Loria, J. P. Induced fit in the selection of correct versus incorrect nucleotides by DNA polymerase β. Biochemistry 55, 382–395 (2016).
Roettger, M. P., Bakhtina, M. & Tsai, M.-D. Mismatched and matched dNTP incorporation by DNA polymerase β proceed via analogous kinetic pathways. Biochemistry 47, 9718–9727 (2008).
Santoso, Y. et al. Conformational transitions in DNA polymerase I revealed by single-molecule FRET. Proc. Natl Acad. Sci. USA 107, 715–720 (2010).
Rothwell, P. J. et al. dNTP-dependent conformational transitions in the fingers subdomain of Klentaq1 DNA polymerase: insights into the role of the “nucleotide-binding” state. J. Biol. Chem. 288, 13575–13591 (2013).
Delarue, M. et al. Crystal structures of a template-independent DNA polymerase: murine terminal deoxynucleotidyltransferase. EMBO J. 21, 427–439 (2002).
Gouge, J., Rosario, S., Romain, F., Beguin, P. & Delarue, M. Structures of intermediates along the catalytic cycle of terminal deoxynucleotidyltransferase: dynamical aspects of the two-metal ion mechanism. J. Mol. Biol. 425, 4334–4352 (2013).
Ummat, A. et al. Human DNA polymerase η is pre-aligned for dNTP binding and catalysis. J. Mol. Biol. 415, 627–634 (2012).
Starcevic, D., Dalal, S. & Sweasy, J. Hinge residue Ile260 of DNA polymerase β is important for enzyme activity and fidelity. Biochemistry 44, 3775–3784 (2005).
Yamtich, J. & Sweasy, J. B. DNA polymerase family X: function, structure, and cellular roles. Biochim. Biophys. Acta 1804, 1136–1150 (2010).
Lindahl, T. & Barnes, D. E. Repair of endogenous DNA damage. Cold Spring Harbor Symp. Quant. Biol. 65, 127–134 (2000).
Tubbs, A. & Nussenzweig, A. Endogenous DNA damage as a source of genomic instability in cancer. Cell 168, 644–656 (2017).
Yang, W. An overview of Y-family DNA polymerases and a case study of human DNA polymerase η. Biochemistry 53, 2793–2803 (2014).
Maxwell, B. A. & Suo, Z. Recent insight into the kinetic mechanisms and conformational dynamics of Y-family DNA polymerases. Biochemistry 53, 2804–2814 (2014).
Williams, J. S., Lujan, S. A. & Kunkel, T. A. Processing ribonucleotides incorporated during eukaryotic DNA replication. Nat. Rev. Mol. Cell. Biol. 17, 350–363 (2016).
Fang, E. F. et al. Nuclear DNA damage signalling to mitochondria in ageing. Nat. Rev. Mol. Cell. Biol. 17, 308–321 (2016).
Marteijn, J. A., Lans, H., Vermeulen, W. & Hoeijmakers, J. H. J. Understanding nucleotide excision repair and its roles in cancer and ageing. Nat. Rev. Mol. Cell. Biol. 15, 465–481 (2014).
Kunkel, T. A. & Soni, A. Exonucleolytic proofreading enhances the fidelity of DNA synthesis by chick embryo DNA polymerase-γ. J. Biol. Chem. 263, 4450–4459 (1988).
Longley, M. J., Nguyen, D., Kunkel, T. A. & Copeland, W. C. The fidelity of human DNA polymerase γ with and without exonucleolytic proofreading and the p55 accessory subunit. J. Biol. Chem. 276, 38555–38562 (2001).
Yousefzadeh, M. J. et al. Mechanism of suppression of chromosomal instability by DNA polymerase POLQ. PLoS Genet. 10, e1004654 (2014).
Zahn, K. E., Averill, A. M., Aller, P., Wood, R. D. & Doublié, S. Human DNA polymerase θ grasps the primer terminus to mediate DNA repair. Nat. Struct. Mol. Biol. 22, 304–311 (2015).
Mateos-Gomez, P. A. et al. Mammalian polymerase θ promotes alternative NHEJ and suppresses recombination. Nature 518, 254–257 (2015).
Yang, W. & Lee, Y.-S. A. DNA-hairpin model for repeat-addition processivity in telomere synthesis. Nat. Struct. Mol. Biol. 22, 844–847 (2015).
Hedglin, M., Pandey, B. & Benkovic, S. J. Stability of the human polymerase δ holoenzyme and its implications in lagging strand DNA synthesis. Proc. Natl Acad. Sci. USA 113, E1777–E1786 (2016).
Hedglin, M., Pandey, B. & Benkovic, S. J. Characterization of human translesion DNA synthesis across a UV-induced DNA lesion. eLife 5, e19788 (2016).
Dilley, R. L. et al. Break-induced telomere synthesis underlies alternative telomere maintenance. Nature 539, 54–58 (2016).
Johnson, R. E., Klassen, R., Prakash, L. & Prakash, S. A major role of DNA polymerase δ in replication of both the leading and lagging DNA strands. Mol. Cell 59, 163–175 (2015).
Burgers, P. M. J., Gordenin, D. & Kunkel, T. A. Who is leading the replication fork, Pol ε or Pol δ? Mol. Cell 61, 492–493 (2016).
Johnson, R. E., Klassen, R., Prakash, L. & Prakash, S. Response to Burgers et. al. Mol. Cell 61, 494–495 (2016).
Moon, A. F. et al. The X family portrait: structural insights into biological functions of X family polymerases. DNA Repair 6, 1709–1725 (2007).
García-Díaz, M. et al. DNA polymerase λ, a novel DNA repair enzyme in human cells. J. Biol. Chem. 277, 13184–13191 (2002).
García-Díaz, M., Bebenek, K., Krahn, J. M., Kunkel, T. A. & Pedersen, L. C. A closed conformation for the Pol λ catalytic cycle. Nat. Struct. Mol. Biol. 12, 97–98 (2005).
Wei, Y. Portraits of a Y-family DNA polymerase. FEBS Lett. 579, 868–872 (2005).
Prakash, S., Johnson, R. E. & Prakash, L. Eukaryotic translesion synthesis DNA polymerases: specificity of structure and function. Annu. Rev. Biochem. 74, 317–353 (2005).
Nugent, C. I. & Lundblad, V. The telomerase reverse transcriptase: components and regulation. Genes Dev. 12, 1073–1085 (1998).
Cong, Y.-S., Wright, W. E. & Shay, J. W. Human telomerase and its regulation. Microbiol. Mol. Biol. Rev. 66, 407–425 (2002).
García-Gómez, S. et al. PrimPol, an archaic primase/polymerase operating in human cells. Mol. Cell 52, 541–553 (2013).
Bianchi, J. et al. PrimPol bypasses UV photoproducts during eukaryotic chromosomal DNA replication. Mol. Cell 52, 566–573 (2013).
Guilliam, T. A. et al. Human PrimPol is a highly error-prone polymerase regulated by single-stranded DNA binding proteins. Nucleic Acids Res. 43, 1056–1068 (2015).
Kamath-Loeb, A. S., Hizi, A., Kasai, H. & Loeb, L. A. Incorporation of the guanosine triphosphate analogs 8-oxo-dGTP and 8-NH2-dGTP by reverse transcriptases and mammalian DNA polymerases. J. Biol. Chem. 272, 5892–5898 (1997).
Pursell, Z. F., McDonald, J. T., Mathews, C. K. & Kunkel, T. A. Trace amounts of 8-oxo-dGTP in mitochondrial dNTP pools reduce DNA polymerase γ replication fidelity. Nucleic Acids Res. 36, 2174–2181 (2008).
Batra, V. K. et al. Mutagenic conformation of 8-oxo-7,8-dihydro-2′-dGTP in the confines of a DNA polymerase active site. Nat. Struct. Mol. Biol. 17, 889–890 (2010).
Çag˘layan, M., Horton, J. K., Dai, D.-P., Stefanick, D. F. & Wilson, S. H. Oxidized nucleotide insertion by pol β confounds ligation during base excision repair. Nat. Commun. 8 14045 (2017).
Burak, M. J., Guja, K. E. & Garcia-Diaz, M. Nucleotide binding interactions modulate dNTP selectivity and facilitate 8-oxo-dGTP incorporation by DNA polymerase lambda. Nucleic Acids Res. 43, 8089–8099 (2015).
Hsu, G. W., Ober, M., Carell, T. & Beese, L. S. Error-prone replication of oxidatively damaged DNA by a high-fidelity DNA polymerase. Nature 431, 217–221 (2004).
Fouquerel, E. et al. Oxidative guanine base damage regulates human telomerase activity. Nat. Struct. Mol. Biol. 23, 1092–1100 (2016).
Kirouac, K. N. & Ling, H. Unique active site promotes error-free replication opposite an 8-oxo-guanine lesion by human DNA polymerase iota. Proc. Natl Acad. Sci. USA 108, 3210–3215 (2011).
Petta, T. B. et al. Human DNA polymerase iota protects cells against oxidative stress. EMBO J. 27, 2883–2895 (2008).
Burak, M. J., Guja, K. E., Hambardjieva, E., Derkunt, B. & Garcia-Diaz, M. A fidelity mechanism in DNA polymerase lambda promotes error-free bypass of 8-oxo-dG. EMBO J. 35, 2045–2059 (2016).
Reha-Krantz, L. J., Nonay, R. L., Day, R. S. III & Wilson, S. H. Replication of O6-methylguanine-containing DNA by repair and replicative DNA polymerases. J. Biol. Chem. 271, 20088–20095 (1996).
Koag, M.-C., Nam, K. & Lee, S. The spontaneous replication error and the mismatch discrimination mechanisms of human DNA polymerase β. Nucleic Acids Res. 42, 11233–11245 (2014).
Lindahl, T. Instability and decay of the primary structure of DNA. Nature 362, 709–715 (1993).
Greenberg, M. M. Looking beneath the surface to determine what makes DNA damage deleterious. Curr. Opin. Chem. Biol. 21, 48–55 (2014).
Schaaper, R. M., Kunkel, T. A. & Loeb, L. A. Infidelity of DNA synthesis associated with bypass of apurinic sites. Proc. Natl Acad. Sci. USA 80, 487–491 (1983).
Sagher, D. & Strauss, B. Insertion of nucleotides opposite apurinic apyrimidinic sites in deoxyribonucleic acid during in vitro synthesis: uniqueness of adenine nucleotides. Biochemistry 22, 4518–4526 (1983).
Hoeijmakers, J. H. Genome maintenance mechanisms for preventing cancer. Nature 411, 366–374 (2001).
Obeid, S. et al. Replication through an abasic DNA lesion: structural basis for adenine selectivity. EMBO J. 29, 1738–1747 (2010).
Arian, D. et al. Irreversible inhibition of DNA polymerase β by small-molecule mimics of a DNA lesion. J. Am. Chem. Soc. 136, 3176–3183 (2014).
Setlow, R. B. Cyclobutane-type pyrimidine dimers in polynucleotides. Science 153, 379–386 (1966).
Todo, T. et al. A new photoreactivating enzyme that specifically repairs ultraviolet light-induced (6–4) photoproducts. Nature 361, 371–374 (1993).
Essen, L. O. & Klar, T. Light-driven DNA repair by photolyases. Cell. Mol. Life Sci. 63, 1266–1277 (2006).
Tan, C. et al. The molecular origin of high DNA-repair efficiency by photolyase. Nat. Comm. 6, 7302 (2015).
Faraji, S. & Dreuw, A. Insights into light-driven DNA repair by photolyases: challenges and opportunities for electronic structure theory. Photochem. Photobiol. 93, 37–50 (2017).
Trincao, J. et al. Structure of the catalytic core of S. cerevisiae DNA polymerase η: implications for translesion DNA synthesis. Mol. Cell 8, 417–426 (2001).
Masutani, C. et al. The XPV (xeroderma pigmentosum variant) gene encodes human DNA polymerase η. Nature 399, 700–704 (1999).
Johnson, R. E., Kondratick, C. M., Prakash, S. & Prakash, L. hRAD30 Mutations in the variant form of xeroderma pigmentosum. Science 285, 263–265 (1999).
Joyce, C. M. Choosing the right sugar: how polymerases select a nucleotide substrate. Proc. Natl Acad. Sci. USA 94, 1619–1622 (1997).
Astatke, M., Ng, K., Grindley, N. D. F. & Joyce, C. M. A single side chain prevents Escherichia coli DNA polymerase I (Klenow fragment) from incorporating ribonucleotides. Proc. Natl Acad. Sci. USA 95, 3402–3407 (1998).
Traut, T. W. Physiological concentrations of purines and pyrimidines. Mol. Cell. Biochem. 140, 1–22 (1994).
Nick McElhinny, S. A. et al. Abundant ribonucleotide incorporation into DNA by yeast replicative polymerases. Proc. Natl Acad. Sci. USA 107, 4949–4954 (2010).
Donigan, K. A., McLenigan, M. P., Yang, W., Goodman, M. F. & Woodgate, R. The steric gate of DNA polymerase ι regulates ribonucleotide incorporation and deoxyribonucleotide fidelity. J. Biol. Chem. 289, 9136–9145 (2014).
Brown, J. A. & Suo, Z. Unlocking the sugar “steric gate” of DNA polymerases. Biochemistry 50, 1135–1142 (2011).
Crespan, E. et al. Impact of ribonucleotide incorporation by DNA polymerases β and λ on oxidative base excision repair. Nat. Commun. 7 10805 (2016).
Lamarche, B. J., Showalter, A. K. & Tsai, M.-D. An error-prone viral DNA ligase. Biochemistry 44, 8408–8417 (2005).
Zanotti, K. J. & Gearhart, P. J. Antibody diversification caused by disrupted mismatch repair and promiscuous DNA polymerases. DNA Repair 38, 110–116 (2016).
Bartlett, J. M. S. & Stirling, D. in PCR Protocols (eds Bartlett, J. M. S. & Stirling, D. ) 3–6 (Humana, 2003).
Mullis, K. B. et al. Process for amplifying, detecting, and/or-cloning nucleic acid sequences. US Patent 4683195 A (1986).
Hottin, A. & Marx, A. Structural insights into the processing of nucleobase-modified nucleotides by DNA polymerases. Acc. Chem. Res. 49, 418–427 (2016).
Zhang, L., Kang, M., Xu, J. & Huang, Y. Archaeal DNA polymerases in biotechnology. Appl. Microbiol. Biotechnol. 99, 6585–6597 (2015).
Sanger, F. & Coulson, A. R. A rapid method for determining sequences in DNA by primed synthesis with DNA polymerase. J. Mol. Biol. 94, 441–448 (1975).
Sanger, F., Nicklen, S. & Coulson, A. R. DNA sequencing with chain-terminating inhibitors. Proc. Natl Acad. Sci. USA 74, 5463–5467 (1977).
Bentley, D. R. et al. Accurate whole human genome sequencing using reversible terminator chemistry. Nature 456, 53–59 (2008).
Rhoads, A. & Au, K. F. PacBio sequencing and its applications. Genomics Proteomics Bioinformatics 13, 278–289 (2015).
Goodwin, S., McPherson, J. D. & McCombie, W. R. Coming of age: ten years of next-generation sequencing technologies. Nat. Rev. Genet. 17, 333–351 (2016). A useful Review on next-generation sequencing technology.
Deamer, D., Akeson, M. & Branton, D. Three decades of nanopore sequencing. Nat. Biotechnol. 34, 518–524 (2016).
Stranges, P. B. et al. Design and characterization of a nanopore-coupled polymerase for single-molecule DNA sequencing by synthesis on an electrode array. Proc. Natl Acad. Sci. USA 113, E6749–E6756 (2016).
Lander, E. S. Initial impact of the sequencing of the human genome. Nature 470, 187–197 (2011).
Soon, W. W., Hariharan, M. & Snyder, M. P. High-throughput sequencing for biology and medicine. Mol. Syst. Biol. 9, 640 (2013).
Regalado, A. For $999, Veritas Genetics will put your genome on a smartphone app. MIT Technology Reviewhttps://www.technologyreview.com/s/600950/for-999-veritas-genetics-will-put-your-genome-on-a-smartphone-app/ (2016).
Malyshev, D. A. & Romesberg, F. E. The expanded genetic alphabet. Angew. Chem. Int. Ed. 54, 11930–11944 (2015).
McMinn, D. L. et al. Efforts toward expansion of the genetic alphabet: DNA polymerase recognition of a highly stable, self-pairing hydrophobic base. J. Am. Chem. Soc. 121, 11585–11586 (1999).
Matsuda, S., Henry, A. A. & Romesberg, F. E. Optimization of unnatural base pair packing for polymerase recognition. J. Am. Chem. Soc. 128, 6369–6375 (2006).
Hirao, I. Unnatural base pair systems for DNA/RNA-based biotechnology. Curr. Opin. Chem. Biol. 10, 622–627 (2006).
Clever, G. H., Kaul, C. & Carell, T. DNA–metal base pairs. Angew. Chem. Int. Ed. 46, 6226–6236 (2007).
Zhang, L. et al. Evolution of functional six-nucleotide DNA. J. Am. Chem. Soc. 137, 6734–6737 (2015).
Georgiadis, M. M. et al. Structural basis for a six nucleotide genetic alphabet. J. Am. Chem. Soc. 137, 6947–6955 (2015).
Hirao, I. et al. An unnatural base pair for incorporating amino acid analogs into proteins. Nat. Biotechnol. 20, 177–182 (2002).
Matsunaga, K.-i., Kimoto, M. & Hirao, I. High-affinity DNA aptamer generation targeting von Willebrand factor A1-domain by genetic alphabet expansion for systematic evolution of ligands by exponential enrichment using two types of libraries composed of five different bases. J. Am. Chem. Soc. 139, 324–334 (2017).
Kimoto, M., Yamashige, R., Matsunaga, K.-i., Yokoyama, S. & Hirao, I. Generation of high-affinity DNA aptamers using an expanded genetic alphabet. Nat. Biotechnol. 31, 453–457 (2013).
Malyshev, D. A. et al. Efficient and sequence-independent replication of DNA containing a third base pair establishes a functional six-letter genetic alphabet. Proc. Natl Acad. Sci. USA 109, 12005–12010 (2012).
Li, L. et al. Natural-like replication of an unnatural base pair for the expansion of the genetic alphabet and biotechnology applications. J. Am. Chem. Soc. 136, 826–829 (2014).
Delaney, J. C. et al. Efficient replication bypass of size-expanded DNA base pairs in bacterial cells. Angew. Chem. Int. Ed. 48, 4524–4527 (2009).
Malyshev, D. A. et al. A semi-synthetic organism with an expanded genetic alphabet. Nature 509, 385–388 (2014). A semi-synthetic organism can be constructed by engineering DNA polymerases and other proteins, and exposing these to unnatural dNTP substrates.
Zhang, Y. et al. A semisynthetic organism engineered for the stable expansion of the genetic alphabet. Proc. Natl Acad. Sci. USA 114, 1317–1322 (2017).
Packer, M. S. & Liu, D. R. Methods for the directed evolution of proteins. Nat. Rev. Genet. 16, 379–394 (2015).
Ellefson, J. W. et al. Synthetic evolutionary origin of a proofreading reverse transcriptase. Science 352, 1590–1593 (2016).
Sauter, K. B. M. & Marx, A. Evolving thermostable reverse transcriptase activity in a DNA polymerase scaffold. Angew. Chem. Int. Ed. 45, 7633–7635 (2006).
Blatter, N. et al. Structure and function of an RNA-reading thermostable DNA polymerase. Angew. Chem. Int. Ed. 52, 11935–11939 (2013).
Aschenbrenner, J. & Marx, A. Direct and site-specific quantification of RNA 2′-O-methylation by PCR with an engineered DNA polymerase. Nucleic Acids Res. 44, 3495–3502 (2016).
Huber, C., von Watzdorf, J. & Marx, A. 5-Methylcytosine-sensitive variants of Thermococcus kodakaraensis DNA polymerase. Nucleic Acids Res. 44, 9881–9890 (2016).
Xia, G. et al. Directed evolution of novel polymerase activities: mutation of a DNA polymerase into an efficient RNA polymerase. Proc. Natl Acad. Sci. USA 99, 6597–6602 (2002).
Chen, T. et al. Evolution of thermophilic DNA polymerases for the recognition and amplification of C2′-modified DNA. Nat. Chem. 8, 556–562 (2016). The authors demonstrate that a polymerase evolution system can produce thermostable enzymes that efficiently interconvert C2′-OMe-modified oligonucleotides and their DNA counterparts through transcription and reverse transcription. This seems to be a powerful tool for tailoring polymerases to have other types of novel functions.
Reha-Krantz, L. J., Woodgate, S. & Goodman, M. F. Engineering processive DNA polymerases with maximum benefit at minimum cost. Front. Microbiol. 5 380 (2014).
Wong, I., Patel, S. S. & Johnson, K. A. An induced-fit kinetic mechanism for DNA replication fidelity: direct measurement by single-turnover kinetics. Biochemistry 30, 526–537 (1991).
Johnson, K. A. 1 Transient-state kinetic analysis of enzyme reaction pathways. Enzymes 20, 1–61 (1992).
Fersht, A. Enzyme Structure and Mechanism 2nd edn, 350 (W. H. Freeman, 1985).
Bertram, J. G., Oertell, K., Petruska, J. & Goodman, M. F. DNA polymerase fidelity: comparing direct competition of right and wrong dNTP substrates with steady state and pre-steady state kinetics. Biochemistry 49, 20–28 (2010).
Kohlstaedt, L. A., Wang, J., Friedman, J. M., Rice, P. A. & Steitz, T. A. Crystal structure at 3.5 Å resolution of HIV-1 reverse transcriptase complexed with an inhibitor. Science 256, 1783–1790 (1992).
Steitz, T. A., Smerdon, S. J., Jäger, J. & Joyce, C. M. A unified polymerase mechanism for nonhomologous DNA and RNA polymerase. Science 266, 2022–2025 (1994).
Knowles, J. R. & Albery, W. J. Perfection in enzyme catalysis: the energetics of triosephosphate isomerase. Acc. Chem. Res. 10, 105–111 (1977).
Kravchuk, A. V., Zhao, L., Kubiak, R. J., Bruzik, K. S. & Tsai, M.-D. Mechanism of phosphatidylinositol-specific phospholipase C: origin of unusually high nonbridging thio effects. Biochemistry 40, 5433–5439 (2001).
Tsai, M.-D. & Yan, H. Mechanism of adenylate kinase: site-directed mutagenesis versus X-ray and NMR. Biochemistry 30, 6806–6818 (1991).
Matute, R. A., Yoon, H. & Warshel, A. Exploring the mechanism of DNA polymerases by analyzing the effect of mutations of active site acidic groups in Polymerase β. Proteins 84, 1644–1657 (2016).
The authors acknowledge financial support from the Ministry of Science and Technology (Grant Nos MOST103-2113-M-001-016-MY3, MOST105-0210-01-12-01 and MOST106-0210-01-15-04) to M.-D.T. and a US National Institutes of Health intramural grant (DK036146-08) to W.Y.
The authors declare no competing interests.
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Wu, WJ., Yang, W. & Tsai, MD. How DNA polymerases catalyse replication and repair with contrasting fidelity. Nat Rev Chem 1, 0068 (2017). https://doi.org/10.1038/s41570-017-0068
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