Kinetic Mechanisms of DNA Polymerases
Dissertation
Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University
By
Jessica Ann Brown, B.S.
Ohio State Biochemistry Program
The Ohio State University
2010
Dissertation Committee:
Zucai Suo, Advisor
Juan D. Alfonzo
James E. Hopper
Jennifer J. Ottesen
Copyright by
Jessica Ann Brown
2010
Abstract
High-fidelity DNA polymerases accurately replicate an organism’s genomic DNA while
low-fidelity DNA polymerases are specialized to function in DNA repair and DNA lesion
bypass, two processes that are necessary to overcome the DNA damage induced by
endogenous and exogenous sources. Therefore, understanding the molecular basis of
polymerase nucleotide selectivity and fidelity is an important objective towards
ascertaining the overall stability of an organism’s genome. Transient state kinetic
techniques were used to elucidate the mechanisms of DNA polymerization catalyzed by high- and low-fidelity enzymes.
Here, we established that the fidelity of Sulfolobus solfataricus DNA polymerase B1 synthesizing undamaged DNA to be in the range of 10-6 to 10-8 or one error per 1,000,000 to 100,000,000 nucleotide incorporations. PolB1 used an induced-fit mechanism to incorporate a correct nucleotide with a tight nucleotide binding affinity and fast rate of incorporation. In contrast, Saccharomyces cerevisiae DNA polymerase η and human
Rev1, two enzymes that function in DNA lesion bypass, synthesized undamaged DNA with a fidelity of 10-2 to 10-4 and 100 to 10-5, respectively. The extremely low fidelity of hRev1 was due to the preferred misincorporation of dCTP with templating bases dA, dT,
and dC.
ii Human DNA polymerase λ (Pol λ), a low-fidelity enzyme involved in gap-filling DNA
synthesis during DNA repair, utilizes unique mechanisms to select nucleotides and was
shown to be potentially mutagenic in different situations. Pol λ prefered to insert
deoxyribonucleotides over ribonucleotides by 3,000- to 50,000-fold due to a steric clash between the ribose 2′-hydroxyl group of a ribonucleotide and a backbone carbonyl group of Y505 in Pol λ’s active site. In addition, the unprecedented tight nucleotide binding affinity of both correct and incorrect nucleotides to the Pol λ•DNA complex was manifested in cooperative interactions with multiple active site residues. Furthermore, the fidelity of Pol λ was governed mostly by R517, a residue that interacts with the minor groove of the DNA template. During long gap-filling DNA synthesis, the fidelity of Pol λ
dropped two orders of magnitude, and this downregulation of fidelity was controlled by
Pol λ’s non-enzymatic N-terminal domains. Pol λ was error-prone when it encountered an
8-oxo-7,8-dihydro-2′-deoxyguanosine lesion in the DNA template, as dCTP and dATP
incorporation proceeded with essentially equal efficiency and probability.
A comprehensive mechanism for the bypass of cis-[Pt(NH3)2{d(GpG)-N7(1),-N7(2)}]
intrastrand cross-links was established for Sulfolobus solfataricus DNA polymerase IV
(Dpo4), an enzyme involved in DNA lesion bypass. Dpo4 was able to bypass this double-
base lesion, although, the incorporation efficiency of dCTP opposite the first and second
cross-linked guanine bases was reduced by 72- and 860-fold, respectively. Moreover, the
fidelity of Dpo4 at the lesion decreased up to two orders of magnitude.
iii Lastly, antiviral nucleotide analogs were determined to be substrates for six human DNA polymerases (Pols β, λ, η, ι, κ, and Rev1) involved in DNA repair and lesion bypass. The
kinetic results suggested that nucleotide analog incorporation catalyzed by these six
human enzymes may represent a potential mechanism of drug toxicity and also
established a structure-function relationship for designing improved analogs.
iv Dedication
Dedicated to the marginalized people throughout the world
v Acknowledgements
My journey through graduate school has introduced me to several special individuals who
have influenced the work presented in this dissertation. First, thank you to my advisor,
Dr. Zucai Suo, for awakening my scientific potential. Dr. Suo was the first person to
stimulate within me a deep passion and excitement for the quest of scientific discovery
and knowledge. Furthermore, his strong commitment to training me and unwavering
confidence in my abilities has allowed me to achieve at a high level.
Thank you to my committee members: Dr. Juan Alfonzo, Dr. James Hopper, and Dr.
Jennifer Ottesen. All of whom monitored my progress and advanced my scientific growth
by providing advice, challenging questions, and constructive criticism. Their positive
words and support were crucial for making me a competitive applicant for fellowships
and awards.
Thank you to the current and former members of the Suo laboratory. Many of the
following have made significant contributions toward advancing my research projects:
David Beyer, Nikunj Bhatt, Eric Bolin, Joseph Dunbar, Wade Duym, Dr. Kevin Fiala,
Dr. Jason Fowler, Dr. Sonja Fraas, Rebecca Frankel, Brian Maxwell, Sean Newmister,
Lindsey Pack, John Pryor, Laura Sanman, Shanen Sherrer, Joshua Wagner, Xin Xia, Dr.
vi Cuiling Xu, Paul Yourik, and Dr. Likui Zhang. The early success of my graduate career
was due to the strong mentorship and training imparted by Dr. Kevin Fiala and Dr. Jason
Fowler—thank you. In addition, Dr. Jason Fowler has helped me to overcome numerous
computer- and equipment-related problems. On many occasions, he would immediately
quit his task and focus solely on solving my problem. Thank you to Shanen Sherrer for her friendship and for extending her generous and invaluable assistance. Thank you to
Dr. Likui Zhang for collaborating on projects and for sharing co-first authorship with me
on two publications. Thank you to the undergraduate researchers, especially Sean
Newmister and Lindsey Pack, for their help with an array of tasks: from making
acrylamide solutions to measuring kinetic parameters.
Thank you to my undergraduate mentor, Dr. Steven Berberich, for giving me the
opportunity to undertake an honors research project in his laboratory at Wright State
University. That research experience encouraged me to pursue a doctoral degree. In
addition, thank you to Dr. Keven Huang, a former graduate student in the Berberich
laboratory. He taught me many techniques and continues to offer advice.
Thank you to our collaborators and others in the DNA polymerase field who contributed
important reagents and thoughtful suggestions. Notably, thank you to Dr. Hong Ling
(University of Western Ontario) and her former graduate student, Dr. Jimson Wong, for
their perseverance in solving several crystal structures of Dpo4•cisplatin-DNA•dNTP.
Thank you to Dr. John-Stephen Taylor (Washington University) and his former graduate
student, Dr. Ajay Kshetry, for synthesizing three non-natural nucleotide analogs. Thank
vii you to Dr. Michael Miller and Dr. Joy Feng at Gilead Sciences, Inc. for generously
providing antiviral nucleotide analogs and for providing information about the chemical
and pharmacological nature of the drugs.
Thank you to the former director, Dr. Ross Dalbey, and current director, Dr. Jill Rafael-
Fortney, of the Ohio State Biochemistry Program. Thank you to Dr. Dehua Pei, director
of the Chemistry-Biology Interface Program at Ohio State. All three of these individuals
have provided guidance and support for me. Moreover, the programs that they oversee were instrumental in my scientific development. Thank you to the faculty and staff in the
Department of Biochemistry for their helpful instruction.
Thank you to my family and friends for their patience, prayerful support, and
unconditional love. Thank you to my mother and father for knowing that I have the
potential to do something great. Thank you to my sister for listening to my good and not-
so-good moments. Thank you to Bethany Couts, Sister Susan Fraser, and Sister Miriam
Krusling for giving me the opportunities to better understand the needs of the poor in
Malawi, Jamaica, and other developing countries.
Thank you to God, the maker of heaven and earth, of all that is seen and unseen. It has
been a joy to learn about the seen and unseen.
This work was supported by grants awarded to my advisor from the National Science
Foundation, National Institutes of Health, and start-up fund from The Ohio State
viii University. Thank you to these generous funding agencies who provided financial support for me during my graduate career: National Institutes of Health Chemistry-
Biology Interface Program, P.E.O. Scholar Award, American Heart Association, and
Presidential Fellowship from The Ohio State University.
ix Vita
Education 2000...... Memorial High School 2005...... B.S. Biological Sciences and Chemistry, Wright State University 2005-present...... Graduate Fellow, Ohio State Biochemistry Program, The Ohio State University
Awards and Honors 2000-2005 ...... Valedictorian Scholarship from Wright State University 2000-2005 ...... Dean’s List 2002...... Inducted into Alpha Lambda Delta National Honor Society 2002...... Inducted into National Society of Collegiate Scholars 2002-2005 ...... College of Science and Mathematics Scholarship from Wright State University 2004-2005 ...... Fred White Scholarship from Wright State University 2005...... Departmental Honors Scholar in Biological Sciences at Wright State University 2005...... Graduated Summa Cum Laude 2005-2007 ...... National Institutes of Health Chemistry-Biology Interface Program Predoctoral Fellowship 2007...... Inducted into Phi Kappa Phi Honor Society 2008...... Outstanding Oral Student Presentation Award for OSU Molecular Life Sciences Interdisciplinary Graduate Programs Symposium 2008&2009 ...... Burrell Memorial Fund from the Department of Biochemistry at The Ohio State University 2008-2009 ...... International P.E.O. Scholar Award for Women 2008-2010 ...... American Heart Association Predoctoral Fellowship 2010...... Presidential Fellowship from The Ohio State University 2010...... Edward J. Ray Travel Award for Scholarship and Service from The Ohio State University
Presentations 2007...... Poster at Annual Chemistry-Biology Interface Program Symposium in Columbus, OH 2008...... Poster at Annual Chemistry-Biology Interface Program Symposium in Columbus, OH 2008...... Talk at monthly Chemistry-Biology Interface Program luncheon in Columbus, OH
x 2008...... Talk and poster at 2nd Annual Molecular Life Sciences Interdisciplinary Graduate Programs Symposium in Columbus, OH 2008...... Talk and poster at Annual Gordon Research Conference on Nucleic Acids in Newport, RI 2009...... Poster at Annual Chemistry-Biology Interface Program Symposium in Columbus, OH 2009...... Poster at 3rd American Society for Microbiology Conference on DNA Repair and Mutagenesis in Whistler, British Columbia, Canada 2010...... Talk at Hayes Graduate Research Forum in Columbus, OH 2010...... Poster at Annual Chemistry-Biology Interface Program Symposium in Columbus, OH 2010...... Plenary talk at 4th Annual Molecular Life Sciences Interdisciplinary Graduate Programs Symposium in Columbus, OH 2010...... Talk at American Society for Virology 29th Annual Meeting in Bozeman, MT
Publications
1. Fiala, K.A., Brown, J.A., Ling, H., Kshetry, A.K., Zhang, J., Taylor, J.-S., Yang, W. & Suo, Z. (2007) Mechanism of Template-Independent Nucleotide Incorporation Catalyzed by a Template-Dependent DNA Polymerase. Journal of Molecular Biology 365, 590-602.
2. Chen, C.-L., Hsieh, F.-C., Lieblein, J.D., Brown, J., Chan, C., Wallace, J.A., Cheng, G., Hall, B.M. & Lin, J. (2007) Stat3 Activation in Human Endometrial and Cervical Cancers. British Journal of Cancer 96, 591-599.
3. Brown, J.A., Duym, W.W., Fowler, J.D. & Suo, Z. (2007) Single-Turnover Kinetic Analysis of the Mutagenic Potential of 8-Oxo-7,8-dihydro-2'-deoxyguanosine during Gap-Filling Synthesis Catalyzed by Human DNA Polymerases λ and β. Journal of Molecular Biology 367, 1258-1269.
4. Fiala, K.A, Sherrer, S.M., Brown, J.A. & Suo, Z. (2008) Mechanistic Consequences of Temperature on DNA Polymerization Catalyzed by a Y-family DNA Polymerase. Nucleic Acids Research 6,1990-2001.
5. Fowler, J.D., Brown, J.A., Johnson, K.A. & Suo, Z. (2008) Kinetic Investigation of the Inhibitory Effect of Gemcitabine on DNA Polymerization Catalyzed by Human Mitochondrial DNA Polymerase. Journal of Biological Chemistry 283, 15339-15348.
6. Brown, J.A., Newmister, S.A., Fiala, K.A. & Suo, Z. (2008) Mechanism of Double- Base Lesion Bypass Catalyzed by a Y-family DNA Polymerase. Nucleic Acids Research 36, 3867-3878.
xi
7. Sherrer, S.M., Brown, J.A., Pack, L.R., Jasti, V.P., Fowler, J.D., Basu, A.K. & Suo, Z. (2009) Mechanistic Studies of the Bypass of a Bulky Single-Base Lesion Catalyzed by a Y-family DNA Polymerase. Journal of Biological Chemistry 284, 6379-6388.
8. Fowler, J.D., Brown, J.A., Kvaratskhelia, M. & Suo, Z. (2009) Probing Protein Conformational Changes of a Human DNA Polymerase Using Mass Spectrometry. Journal of Molecular Biology 390, 368-379.
9. Zhang, L.*, Brown, J.A.*, Newmister, S.A. & Suo, Z. (2009) Polymerization Fidelity of a Replicative DNA Polymerase from the Hyperthermophilic Archaeon Sulfolobus solfataricus P2. Biochemistry 48, 7492-7501.
10. Brown, J.A. & Suo, Z. (2009) Elucidating the Kinetic Mechanism of DNA Polymerization Catalyzed by Sulfolobus solfataricus P2 DNA Polymerase B1. Biochemistry 48, 7502-7511.
11. Xu, C., Maxwell, B.A., Brown, J.A., Zhang, L. & Suo, Z. (2009) Global Conformational Dynamics of a Y-family DNA Polymerase during Catalysis. PLoS Biol. 7(10): e1000225.
12. Brown, J.A., Fiala, K.A., Fowler, J.D., Sherrer, S.M., Newmister, S.A., Zhang, L. & Suo, Z. (2010) A Novel Mechanism of Sugar Selection Utilized by a Human X- family DNA Polymerase. Journal of Molecular Biology 395, 282-290.
13. Wong, J.H.Y., Brown, J.A., Suo, Z., Blum, P., Nohmi, T. & Ling, H. (2010) Structural Insight into Dynamic Bypass of the Major Cisplatin-DNA Adduct by Y- family Polymerase Dpo4. EMBO Journal 29, 2059-2069.
14. Brown, J.A.*, Zhang, L.*, Sherrer, S.M., Burgers, P.M.J., Taylor, J.-S. & Suo, Z. (2010) Pre-Steady State Kinetic Analysis of Truncated and Full-Length Saccharomyces cerevisiae DNA Polymerase η. Journal of Nucleic Acids 2010, 11 pages.
15. Brown, J.A., Fowler, J.D. & Suo, Z. (2010) Kinetic Basis of Nucleotide Selection Employed by a Protein Template-Dependent DNA Polymerase. Biochemistry 49, 5504-5510.
16. Brown, J.A., Pack, L.R., Sherrer, S.M., Kshetry, A.K., Newmister, S.A., Fowler, J.D., Taylor, J.-S. & Suo, Z. (2010) Identification of Critical Residues for the Tight Binding of Both Correct and Incorrect Nucleotides to Human DNA Polymerase λ. Journal of Molecular Biology 403, 505-515.
*indicates co-first author
xii Fields of Study
Major Field: Ohio State Biochemistry Program
xiii Table of Contents
Abstract...... ii
Dedication...... v
Acknowledgements...... vi
Vita...... x
List of Tables ...... xx
List of Figures...... xxvi
List of Schemes...... xxxi
Abbreviations...... xxxii
Chapter 1: Introduction...... 1
1.1 Families of DNA Polymerases...... 1
1.2 Structural, Chemical, and Kinetic Mechanisms...... 4
1.3 Kinetic Basis of Nucleotide Selectivity and Fidelity...... 6
1.4 Focus of Dissertation ...... 8
1.5 Table...... 14
1.6 Figures...... 15
1.7 Scheme...... 17
Chapter 2: Polymerization Fidelity of Sulfolobus solfataricus P2 DNA
Polymerase B1 ...... 18
xiv 2.1 Introduction...... 18
2.2 Materials and Methods...... 19
2.3 Results...... 23
2.4 Discussion...... 28
2.5 Tables...... 35
2.6 Figures...... 41
Chapter 3: Kinetic Mechanism of DNA Polymerization Catalyzed by Sulfolobus
solfataricus P2 DNA Polymerase B1 at 37 °C ...... 47
3.1 Introduction...... 47
3.2 Materials and Methods...... 48
3.3 Results...... 52
3.4 Discussion...... 59
3.5 Future Directions ...... 64
3.6 Table ...... 65
3.7 Figures...... 66
3.8 Scheme...... 78
Chapter 4: A Novel Mechanism of Sugar Selection Utilized by Human DNA
Polymerase λ...... 79
4.1 Introduction...... 79
4.2 Materials and Methods...... 80
4.3 Results...... 82
4.4 Discussion...... 87
4.5 Tables...... 92
xv 4.6 Figures...... 96
Chapter 5: Identification of Critical Residues for the Tight Binding of Both
Correct and Incorrect Nucleotides to Human DNA Polymerase λ ...... 101
5.1 Introduction...... 101
5.2 Materials and Methods...... 102
5.3 Results...... 104
5.4 Discussion...... 109
5.5 Tables...... 113
5.6 Figures...... 118
Chapter 6: Efficiency and Fidelity of Human DNA Polymerases λ and β
during Gap-Filling DNA Synthesis ...... 125
6.1 Introduction...... 125
6.2 Materials and Methods...... 127
6.3 Results...... 128
6.4 Discussion...... 132
6.5 Future Directions ...... 137
6.6 Tables...... 139
6.7 Figures...... 149
Chapter 7: Single-Turnover Kinetic Analysis of the Mutagenic Potential of
8-Oxo-7,8-dihydro-2′-deoxyguanosine during Gap-Filling Synthesis
Catalyzed by Human DNA Polymerase λ and β...... 156
7.1 Introduction...... 156
7.2 Materials and Methods...... 158
xvi 7.3 Results...... 160
7.4 Discussion...... 164
7.5 Future Directions ...... 170
7.6 Tables...... 172
7.7 Figures...... 177
Chapter 8: Pre-Steady State Kinetic Analysis of Truncated and Full-Length
Saccharomyces cerevisiae DNA Polymerase Eta...... 182
8.1 Introduction...... 182
8.2 Materials and Methods...... 183
8.3 Results and Discussion ...... 186
8.4 Tables...... 195
8.5 Figures...... 201
8.6 Scheme...... 205
Chapter 9: Kinetic Basis of Nucleotide Selection Employed by a Protein
Template-Dependent DNA Polymerase ...... 206
9.1 Introduction...... 206
9.2 Materials and Methods...... 207
9.3 Results...... 209
9.4 Discussion...... 212
9.5 Future Directions ...... 216
9.6 Tables...... 217
9.7 Figures...... 221
xvii Chapter 10: Mechanism of Double-Base Lesion Bypass Catalyzed by a
Y-family DNA Polymerase...... 224
10.1 Introduction...... 224
10.2 Materials and Methods...... 226
10.3 Results...... 228
10.4 Discussion...... 233
10.5 Future Directions ...... 240
10.6 Tables...... 241
10.7 Figures...... 247
10.8 Schemes ...... 254
Chapter 11: Pre-Steady State Kinetic Analysis of the Incorporation of Anti-HIV
Nucleotide Analogs Catalyzed by Human X- and Y-family DNA Polymerases ...... 256
11.1 Introduction...... 256
11.2 Materials and Methods...... 258
11.3 Results...... 260
11.4 Discussion...... 263
11.5 Future Directions ...... 267
11.6 Tables...... 268
11.7 Figures...... 272
Chapter 12: Kinetic Investigation of the Incorporation of Anti-HBV Nucleotide
Analogs Catalyzed by Non-Canonical Human DNA Polymerases ...... 274
12.1 Introduction...... 274
12.2 Materials and Methods...... 275
xviii 12.3 Results...... 278
12.4 Discussion...... 281
12.5 Future Directions ...... 285
12.6 Tables...... 286
12.7 Figures...... 291
Chapter 13: Additional Kinetic Studies on the Mechanism of Nucleotide
Incorporation Catalyzed by Human DNA Polymerases λ and β ...... 296
13.1 Introduction...... 296
13.2 Materials and Methods...... 297
13.3 Results, Discussion, and Future Studies ...... 300
13.4 Tables...... 309
13.5 Figures...... 314
13.6 Scheme...... 320
References...... 321
xix List of Tables
Table 1.1 Classification and function of human DNA polymerases ...... 14
Table 2.1 Sequences of synthetic double-stranded DNA substrates...... 35
Table 2.2 Kinetic parameters of nucleotide incorporation into DNA catalyzed
by PolB1 exo- at 37 °C ...... 36
Table 2.3 Temperature dependence of the nucleotide incorporation fidelity
catalyzed by PolB1 exo- ...... 37
Table 2.4 pH dependence of the nucleotide incorporation fidelity of PolB1 exo-
at 37 °C ...... 38
Table 2.5 Fidelity of the extension of a mismatched dA:dA terminus catalyzed
by PolB1 exo- at 37 °C ...... 39
Table 2.6 Comparison of the base substitution fidelity of PolB1 exo- and four
other DNA polymerases and HIV-1 reverse transcriptase...... 40
Table 3.1 Estimated kinetic constants...... 65
Table 4.1 Kinetic parameters of matched rNTP incorporation into single-
nucleotide gapped DNA catalyzed by Pol λ at 37 °C ...... 92
Table 4.2 Kinetic parameters of mismatched rNTP incorporation into
single-nucleotide gapped D-1 DNA catalyzed by Pol λ at 37 °C...... 93
xx Table 4.3 Kinetic parameters of dTTP or UTP incorporation into single-
nucleotide gapped D-1 DNA catalyzed by Pol λ at 37 °C...... 94
Table 4.4 Kinetic parameters of rCTP analogs incorporated into single-
nucleotide gapped D-6 DNA catalyzed by Pol λ at 37 °C...... 95
Table 5.1 DNA substrates...... 113
Table 5.2 Equilibrium dissociation constants of the Pol λ•DNA complex
at 37 °C ...... 114
Table 5.3 Kinetic parameters for incorrect nucleotide incorporation (dGTP) into
single-nucleotide gapped DNA (D-1) catalyzed by human Pol λ at 37 °C ...... 115
Table 5.4 Kinetic parameters for correct nucleotide incorporation (dTTP) into
single-nucleotide gapped DNA (D-1) catalyzed by human Pol λ at 37 °C ...... 116
Table 5.5 Kinetic parameters for non-natural nucleotide analog incorporation into
single-nucleotide gapped D-7 DNA catalyzed by WT human Pol λ at 37 °C...... 117
Table 6.1 DNA substrates...... 139
Table 6.2 Kinetic parameters for nucleotide incorporation into gapped or
recessed DNA catalyzed by Pol λ at 37 °C...... 140
Table 6.3 Kinetic parameters for nucleotide incorporation into gapped or
recessed DNA catalyzed by Pol β at 37 °C...... 142
Table 6.4 Kinetic parameters for nucleotide incorporation into gapped or
recessed DNA catalyzed by dPol λ at 37 °C...... 143
Table 6.5 Kinetic parameters for nucleotide incorporation into gapped or
recessed DNA catalyzed by tPol λ at 37 °C...... 144
xxi Table 6.6 Kinetic parameters for nucleotide incorporation into gapped DNA
catalyzed by Pol λ at 37 °C...... 145
Table 6.7 Kinetic parameters for nucleotide incorporation into gapped DNA
catalyzed by dPol λ at 37 °C...... 146
Table 6.8 Kinetic parameters for nucleotide incorporation into gapped DNA
catalyzed by tPol λ at 37 °C...... 147
Table 6.9 Kinetic parameters for nucleotide incorporation into gapped DNA
catalyzed by Pol β at 37 °C...... 148
Table 7.1 Kinetic parameters of nucleotide incorporation into single-
nucleotide gapped DNA catalyzed by human Pol λ at 37 °C ...... 172
Table 7.2 Kinetic parameters of nucleotide incorporation into single-
nucleotide gapped DNA catalyzed by human Pol β at 37 °C ...... 173
Table 7.3 Kinetic parameters of nucleotide incorporation into single-
nucleotide gapped DNA catalyzed by human Pol λ at 37 °C ...... 174
Table 7.4 Kinetic parameters of nucleotide incorporation into single-
nucleotide gapped DNA catalyzed by human Pol β at 37 °C ...... 175
Table 7.5 Class assignments for DNA polymerases based upon dCTP:dATP
incorporation ratios opposite template base 8-oxodG ...... 176
Table 8.1 Sequences of DNA substrates...... 195
Table 8.2 Rate and equilibrium dissociation constants for the binary
complex yPol η•DNA at 23 °C ...... 196
Table 8.3 Kinetic parameters of nucleotide incorporation into D-DNA
catalyzed by truncated yPol η at 23 °C...... 197
xxii Table 8.4 Kinetic parameters for nucleotide incorporation onto blunt-end
DNA catalyzed by truncated yeast Pol η at 23 °C ...... 198
Table 8.5 Kinetic parameters of nucleotide incorporation into D-1 DNA
catalyzed by full-length yPol η at 23 °C...... 199
Table 8.6 Comparison of base substitution fidelity for various DNA polymerases ...... 200
Table 9.1 Sequences of the D-DNA substrates...... 217
Table 9.2 Kinetic parameters for nucleotide incorporation into D-DNA catalyzed
by hRev1 at 37 °C...... 218
Table 9.3 Kinetic parameters for non-natural nucleotide analog incorporation
into D-G DNA catalyzed by hRev1 at 37 °C...... 219
Table 9.4 Kinetic parameters for CTP analog incorporation into D-G DNA
catalyzed by hRev1 at 37 °C...... 220
Table 10.1 DNA sequences of primers and templates...... 241
Table 10.2 Binding affinity of Dpo4 to control and damaged DNA substrates
at 23 °C ...... 242
Table 10.3 Kinetic parameters of nucleotide incorporation into cisplatin-
modified DNA ...... 243
Table 10.4 Kinetic parameters of correct nucleotide incorporation into the
44CTL DNA template...... 245
Table 10.5 Biphasic kinetic parameters of dCTP incorporation into cisplatin-
modified DNA ...... 246
Table 11.1 Sequences of oligonucleotides...... 268
xxiii Table 11.2 Kinetic parameters for nucleotide incorporation opposite template
base dG at 37 °C ...... 269
Table 11.3 Kinetic parameters for nucleotide incorporation opposite template
base dT at 37 °C...... 270
Table 11.4 Kinetic parameters for nucleotide incorporation opposite template
base dA at 37 °C ...... 271
Table 12.1 Sequences of oligonucleotides...... 286
Table 12.2 Kinetic parameters for nucleotide incorporation opposite template
dT at 37 °C...... 287
Table 12.3 Kinetic parameters for nucleotide incorporation opposite template
dG at 37 °C ...... 288
Table 12.4 Kinetic parameters for nucleotide incorporation opposite template
dA at 37 °C ...... 289
Table 12.5 Kinetic parameters for nucleotide incorporation opposite template
dC at 37 °C...... 290
Table 13.1 Sequences of the D-DNA substrates...... 309
Table 13.2 Kinetic parameters for nucleotide incorporation into D-T DNA
catalyzed by Pol λ at varying reaction temperatures ...... 310
Table 13.3 Kinetic parameters for nucleotide incorporation into D-T DNA
catalyzed by Pol β at varying reaction temperatures ...... 311
Table 13.4 Arrhenius activation energy barriers (kcal/mol) for nucleotide
incorporation opposite template dT ...... 312
xxiv Table 13.5 Arrhenius activation energy barriers (kcal/mol) for correct
nucleotide incorporations catalyzed by Pol λ ...... 313
xxv List of Figures
Figure 1.1 X-ray crystal structures of DNA polymerases...... 15
Figure 1.2 The two metal-ion mechanism of nucleotidyl transfer...... 16
Figure 2.1 Comparison of polymerase and 3′ → 5′ exonuclease activity for
PolB1 enzymes ...... 41
2+ Figure 2.2 Effect of Mg concentration, NaCl concentration, and pH on the
enzymatic activity of PolB1 exo-...... 42
Figure 2.3 Concentration dependence on the pre-steady-state rate of correct
and incorrect nucleotide incorporation catalyzed by PolB1 exo- ...... 43
Figure 2.4 Temperature dependence of the nucleotide incorporation fidelity
of PolB1exo- ...... 44
Figure 2.5 pH dependence of the nucleotide incorporation fidelity of PolB1 exo-...... 45
Figure 2.6 Rate of 3′ → 5′ exonuclease activity catalyzed by PolB1 ...... 46
Figure 3.1 Pre-steady state and steady-state kinetics of dTTP incorporation
into D-1 DNA by PolB1 exo- ...... 66
Figure 3.2 Active site titration of PolB1 exo-...... 68
Figure 3.3 PolB1 exo- displayed low processivity ...... 69
Figure 3.4 Kinsim and Fitsim analysis ...... 71
xxvi Figure 3.5 Elemental effect on the rate of correct and incorrect nucleotide
incorporation...... 72
Figure 3.6 Measurement of the dissociation rate for the E•DNA•dNTP complex ...... 73
Figure 3.7 Measurement of the dissociation rate for the E•DNA•dNTP complex
at 35 °C ...... 74
Figure 3.8 Measurement of the dissociation rate for the E′•DNA•dNTP complex ...... 75
Figure 3.9 Pulse-chase and pulse-quench experiment ...... 76
Figure 3.10 Activation energy for nucleotide incorporation ...... 77
Figure 4.1 Sequence alignment of human X-family DNA polymerases and the
active site of Pol λ...... 96
Figure 4.2 Concentration dependence on the pre-steady state rate constant of
matched ribonucleotide incorporation ...... 97
Figure 4.3 CD spectra of wild-type Pol λ and its mutants...... 98
Figure 4.4 Chemical structures of CTP analogs ...... 99
Figure 4.5 Model of a ribonucleotide bound at the active site of Pol λ...... 100
Figure 5.1 Active site of truncated Pol λ ...... 118
Figure 5.2 Chemical structure of nucleotide analogs...... 119
Figure 5.3 Circular dichroism spectra for WT and mutant Pol λ...... 120
Figure 5.4 Equilibrium dissociation constant for the dissociation of the binary
complex Pol λ•DNA ...... 122
Figure 5.5 Concentration dependence on the pre-steady state rate constant
of nucleotide misincorporation ...... 123
Figure 5.6 Fidelity of mutant enzymes versus WT...... 124
xxvii Figure 6.1 Domains of Pol λ, dPol λ, tPol λ, and Pol β ...... 149
Figure 6.2 Single-turnover kinetic parameters of nucleotide incorporation...... 150
Figure 6.3 Effect of gap size on polymerization efficiency...... 151
Figure 6.4 Effect of gap size on polymerase fidelity...... 152
Figure 6.5 Effect of DNA sequence on polymerization efficiency ...... 153
Figure 6.6 Effect of DNA sequence on polymerization fidelity...... 154
Figure 6.7 Model for long gap-filling DNA synthesis catalyzed by tPol λ ...... 155
Figure 7.1 Schematic representations of human Pol λ and Pol β...... 177
Figure 7.2 Single-nucleotide gapped-DNA substrates 21-19/41-mer (D-DNA)
and 21-18/40-mer (O-6)...... 178
Figure 7.3 Autoradiographed gel image depicting the ligation of incorporated
8-oxodG to a downstream primer of single-nucleotide gapped DNA ...... 179
Figure 7.4 Concentration dependence on the pre-steady state rate constant of
Modified nucleotide incorporation ...... 180
Figure 7.5 Model of 8-oxodG in the polymerase active site of human Pol λ ...... 181
Figure 8.1 Schematic illustration of yPol η ...... 201
Figure 8.2 Equilibrium dissociation constant for full-length yPol η ...... 202
Figure 8.3 Concentration dependence on the pre-steady state rate constant
of nucleotide incorporation catalyzed by truncated yPol η...... 203
Figure 8.4 Chemical structure of a non-natural nucleotide analog, dPTP...... 204
Figure 9.1 Active site of hRev1 ...... 221
Figure 9.2 Concentration dependence on the pre-steady state rate constant of
deoxycytidyl transferase catalyzed by hRev1...... 222
xxviii Figure 9.3 Chemical structures of nucleotide analogs...... 223
Figure 10.1 Running start nucleotide incorporation assay...... 247
Figure 10.2 Purity of template 44DDP ...... 248
Figure 10.3 Measurement of Dpo4 binding to 24-mer/44DDP ...... 249
Figure 10.4 Concentration dependence on the pre-steady state rate constant
of dCTP incorporation ...... 250
Figure 10.5 Quantitative effect of the cisplatin-DNA adduct on nucleotide
incorporation and fidelity...... 251
Figure 10.6 Effectiveness of the D-1 DNA trap ...... 252
Figure 10.7 Biphasic kinetics observed in the presence of a DNA trap when
Dpo4 incorporated dCTP opposite the cisplatin lesion...... 253
Figure 11.1 Chemical structures of NRTIs investigated in this study and
their natural counterpart...... 272
Figure 11.2 Concentration dependence on the pre-steady state rate constant
of L-3TC-TP incorporation catalyzed by Pol η ...... 273
Figure 12.1 Chemical structures of anti-HBV nucleoside analogs and their
natural counterpart ...... 291
Figure 12.2 Concentration dependence on the pre-steady state rate constant
of PMEA-DP incorporation catalyzed by Pol λ...... 292
Figure 12.3 Extension of ETV-MP and L-TBV-MP catalyzed by Pol β...... 293
Figure 12.4 Extension of ETV-MP and L-TBV-MP catalyzed by Pol η...... 294
Figure 12.5 Ligation of ETV-MP and L-TBV-MP catalyzed by human
DNA ligase I ...... 295
xxix Figure 13.1 Alpha-thiol elemental effect...... 314
Figure 13.2 Pulse-chase and pulse-quench experiment ...... 315
Figure 13.3 Concentration dependence on the pre-steady state rate constant
of nucleotide incorporation...... 316
Figure 13.4 Activation energy barrier...... 317
Figure 13.5 Domains of Pol λ, dPol λ, tPol λ, and Pol β ...... 318
Figure 13.6 Fidelity as a function of temperature...... 319
xxx List of Schemes
Scheme 1.1...... 17
Scheme 3.1...... 78
Scheme 8.1...... 205
Scheme 10.1...... 254
Scheme 10.2...... 255
Scheme 13.1...... 320
xxxi Abbreviations
1-dNATP...... 1-naphthalene 5′-triphosphate 2′-F-CTP ...... 2′-fluoro-2′-deoxycytidine-5′-triphosphate 2′-NH2-CTP...... 2′-amino-2′-deoxycytidine-5′-triphosphate 2′-OCH3-CTP...... 2′-O-methylcytidine-5′-triphosphate 5-dNITP ...... 5-nitroindole 5′-triphosphate 8-oxodG ...... 8-oxo-7,8-dihydro-2′-deoxyguanosine 8-oxodGTP...... 8-oxo-7,8-dihydro-2′-deoxyguanosine 5′-triphosphate araCTP ...... 2′-aracytidine-5′-triphosphate AZT...... 3′-azido-3′-deoxythymidine or zidovudine BER...... base excision repair BF...... Bacillus stearothermophilus DNA polymerase I fragment BRCT ...... breast cancer susceptibility gene 1 C-terminal BSA...... bovine serum albumin CD...... circular dichroism cisplatin...... cis-diamminedichloroplatinum(II) cisplatin-d(GpG) ...... cis-[Pt(NH3)2{d(GpG)-N7(1),-N7-(2)}] intrastrand cross-link Dbh...... Sulfolobus solfataricus P1 DinB homolog ddNTP...... 2′,3′-dideoxyribonucleotide-5′-triphosphate DDP...... cis-diamminedichloroplatinum(II) ddTTP ...... 2′,3′-dideoxythymidine 5′-triphosphate dNITP...... 5-nitroindole 5′-triphosphate dNTP...... 2′-deoxynucleoside 5′-triphosphate DP ...... diphosphate Dpo4 ...... Sulfolobus solfataricus P2 DNA polymerase IV dPTP...... pyrene 5′-triphosphate dRPase...... 5′-deoxyribose-5-phosphate lyase DTT...... dithiothreitol EDTA...... ethylenediaminetetraacetic acid EMSA ...... electrophoretic mobility shift assay ETV...... entecavir FDA...... Food and Drug Administration GemCTP ...... 2′-deoxy-2′,2′-difluorocytidine-5′-triphosphate HBV ...... hepatitis B virus HhH...... helix-hairpin-helix HIV-1 RT...... human immunodeficiency virus type 1 reverse transcriptase
xxxii hMYH ...... human MutY homolog hPol γ ...... human mitochondrial DNA polymerase gamma hRev1 ...... human Rev1 L-3TC...... β-L-2′,3′-dideoxy-3′-thiacytidine or lamivudine L-FTC ...... β-L-2′,3′-dideoxy-5-fluoro-3′-thiacytidine or emtricitabine LP...... long patch L-TBV...... β-L-2′-deoxythymidine 5′-triphosphate or telbivudine MP...... monophosphate MTH1...... MutT homolog-1 NHEJ...... non-homologous end joining NLS...... nuclear localization signal NMR ...... nuclear magnetic resonance NRTI ...... nucleoside reverse transcriptase inhibitors PAGE ...... polyacrylamide gel electrophoresis PBMC ...... peripheral blood mononuclear cells PCNA...... proliferating cell nuclear antigen PHA...... phytohemagglutinin PMEA ...... 9-[2-(phosphonomethoxy)ethyl]adenine or adefovir PMPA...... 9-[2-(phosphonomethoxy)propyl]adenine or tenofovir PIP...... PCNA-interacting peptide Pol ...... DNA polymerase Pol β ...... DNA polymerase beta PolB1...... Sulfolobus solfataricus P2 DNA polymerase B1 PolB2...... Sulfolobus solfataricus P2 DNA polymerase B2 PolB3...... Sulfolobus solfataricus P2 DNA polymerase B3 PolB1 exo- ...... exonuclease-deficient DNA polymerase B1 Pol γ ...... DNA polymerase gamma Pol δ ...... DNA polymerase delta Pol ε ...... DNA polymerase epsilon Pol η ...... DNA polymerase eta Pol ι ...... DNA polymerase iota Pol κ ...... DNA polymerase kappa Pol λ ...... DNA polymerase lambda Pol µ...... DNA polymerase mu rNTP...... ribonucleotide-5′-triphosphate rPol β...... rat DNA polymerase beta RT ...... reverse transcriptase SP ...... short patch Sp-dNTPαS ...... Sp-isomer of 2′-deoxynucleoside 5′-O-(1-thiotriphosphate) Sso...... Sulfolobus solfataricus T4 DNA Pol ...... T4 DNA polymerase T7 DNA Pol ...... T7 DNA polymerase Taq ...... Thermus aquaticus DNA polymerase I TBE...... tris/boric acid/EDTA TdT...... terminal deoxynucleotidyltransferase
xxxiii TLS ...... translesion DNA synthesis TP...... triphosphate tPol λ ...... truncated human DNA polymerase λ TT...... cis-syn thymine-thymine dimer UBZ...... ubiquitin-binding zinc finger WT ...... wild-type yPol η ...... Saccharomyces cerevisiae or yeast DNA polymerase eta yRev1 ...... Saccharomyces cerevisiae or yeast Rev1
xxxiv Chapter 1: Introduction
1.1 Families of DNA Polymerases
The synthesis of nucleic acid polymers is likely one of the earliest enzymatic activities to appear in the evolution of life. In 1958, Arthur Kornberg and colleagues reported the discovery of a protein enzyme, Escherichia coli DNA polymerase I, capable of creating an elongated DNA polymer using a DNA substrate and the initial, unreacted monomeric units of four deoxyribonucleotide 5′-triphosphates (dNTPs) [1]. Since then, a plethora of DNA polymerases (Pol) have been identified and characterized. The unique properties of DNA polymerases enable them to replicate and to maintain an organism’s genomic DNA through three basic processes: DNA replication, DNA repair, and DNA lesion bypass. It has been estimated that endogenous DNA damage accounts for approximately 30,000 lesions in a mammalian cell per day [2]. Thus, it is critical for an organism to have DNA polymerases proficient in DNA replication, repair, and lesion bypass. In all domains of life, the genomes of many organisms (excluding viruses) encode multiple DNA polymerases to perform these diverse DNA transactions: four in Sulfolobus solfataricus (archaea), five in E. coli (bacteria), ten in Saccharomyces cerevisiae (eukaryote, fungi), 12 in Arabidopsis thaliana (eukaryote, plant), and at least 16 in Homo sapiens (eukaryote, animal). Using phylogenetic relationships and primary sequence information, DNA polymerases are classified into one of six families: A, B, C, D, X, or Y [3-6].
Family A Members of the A family share similarity to E. coli Pol I. In addition to polymerase activity, these pols may also contain 5′ → 3′ exonuclease activity or 3′ → 5′ exonuclease activity which is a proofreading mechanism to correct incorporation mistakes by the
1 polymerase. Extensive biochemical characterization of E. coli Pol I and bacteriophage T7 DNA polymerase have provided significant insights into the mechanisms of DNA polymerization. In general, A-family DNA polymerases from bacterial organisms are more involved in DNA repair and Okazaki fragment maturation while A-family DNA polymerases from other organisms have a primary role in genomic replication [7]. The three A-family DNA polymerases encoded in the human genome (i.e. Pols γ, θ, and ν) exemplify the diverse functionality of this family (Table 1.1). For example, Pol γ is responsible for the replication and repair of the mitochondrial genome [8] while Pol θ and Pol ν have been implicated in DNA repair, translesion synthesis, and antibody generation [9-17].
Family B Many of the B-family members are high-fidelity replicative DNA polymerases that possess 3′ → 5′ exonuclease activity. The best characterized are DNA polymerases from bacteriophage Phi29, RB69, and T4. Human Pols α, δ, ε, and ζ are multisubunit enzymes. The subunits of Pol α confer both primase and polymerase functions to initiate replication [18]. Then, Pols δ and ε replicate the large genomes in higher eukaryotes, whereby Pol ε synthesizes most of the leading strand while Pol δ synthesizes most of the lagging strand (Table 1.1) [18, 19]. In contrast, Pol ζ is a low-fidelity polymerase with the ability to efficiently extend beyond mismatches and sites of DNA damage, therefore, its major cellular role is proposed to be the extension step during translesion synthesis [20-24].
Family C Bacterial replicative DNA polymerases, which are endowed with an additional 3′ → 5′ exonuclease activity, are the exclusive members of family C, and E. coli Pol III serves as the prototype [25, 26].
Family D Family D is comprised of replicative DNA polymerases from archaea [27]. Like family C, these polymerases have both 5′ → 3′ polymerase activity and 3′ → 5′ exonuclease
2 activity. The most prominent member is Pyrococcus furiosus (Pfu) DNA polymerase due to its widespread use in PCR.
Family X X-family DNA polymerases belong to a large superfamily of nucleotidyl transferses [25]. The human genome encodes five X-family DNA polymerases: Pols β, λ, μ, σ, and terminal deoxynucleotidyl transferase (TdT) (Table 1.1). Most of these enzymes participate in gap-filling DNA synthesis, lack a 3′ → 5′ exonuclease proofreading domain, and have an 8 kDa domain connected to the N-terminus of the polymerase domain. This 8 kDa domain is important for binding to gapped DNA substrates and, in Pol β and Pol λ, it contains 5′-deoxyribose-5-phosphate lyase (dRPase) activity which is required for these enzymes to complete dRP removal during short-patch base excision repair [28]. Interestingly, TdT and other select X-family members have the unique capacity to perform template-independent DNA synthesis [29]. Most of these polymerases play critical roles in DNA repair and antibody generation processes [28, 30]. Lastly, Pol σ is involved in sister chromatid cohesion [31].
Family Y Y-family DNA polymerases are characterized by their low polymerase fidelity, lack of 3′ → 5′ exonuclease activity, poor processivity, and loose active sites. Such properties are important for their role in translesion synthesis. It is hypothesized that replicative DNA polymerases stall upon encountering unrepaired DNA lesions. To rescue a stalled replication fork, a Y-family DNA polymerase incorporates nucleotides opposite and downstream of the lesion site before a replicative DNA polymerase can resume synthesis [32]. The four Y-family members in humans are Pol η, Pol ι, Pol κ, and Rev1 (Table 1.1). Pol η is one of the few Y-family DNA polymerases with a confirmed biological role, i.e. to bypass sunlight-induced lesions in an efficient and error-free manner [33-35]. Alterations in the Pol η gene lead to a disorder known as a variant of Xeroderma pigmentosum which is associated with increased photosensitivity and an early
3 predisposition to skin cancer [36-38]. Y-family DNA polymerases have also been suggested to participate in DNA repair and antibody generation pathways [39, 40].
1.2 Structural, Chemical, and Kinetic Mechanisms
Structural analysis X-ray crystal structures have been solved for at least one member of each DNA polymerase family (Figure 1.1). Among the six families, the overall structural architecture of DNA polymerases remains remarkably consistent despite the lack of sequence similarity [41]. The polymerase core is organized into the fingers, palm, and thumb subdomains [42, 43]. This nomenclature system originated from the topology of the polymerase resembling a right hand as it grips the DNA substrate. In general, the structure-function relationship for each subdomain is as follows: the thumb subdomain contributes to binding the DNA substrate, the fingers subdomain binds to an incoming dNTP, and the palm subdomain contains the conserved catalytic residues required for the nucleotidyl transfer reaction. Please note, X-family DNA polymerases follow a left-hand structure, therefore, the aforementioned roles of the fingers and thumb subdomains are reversed (Figure 1.1) [44]. The presence of DNA and nucleotide substrates can induce significant conformational changes to the polymerase structure as revealed by the superimposition of apo (Pol), binary (Pol•DNA), and ternary (Pol•DNA•dNTP) crystal structures [45, 46]. These conformational changes are crucial for nucleotide discrimination and to poise the active site for catalysis (see below).
In addition to the polymerase core, most DNA polymerases are linked to an additional domain or subdomain. For example, replicative DNA polymerases in the A-, B-, C-, and D-families contain a 3′ → 5′ exonuclease domain, X-family DNA polymerases have an 8 kDa domain, and Y-family DNA polymerases have a little finger subdomain (Figure 1.1). Each of these domains or subdomains contributes to the catalytic function and biological role of DNA polymerases. The exonuclease domain can detect and remove mispairs,
4 thereby enhancing polymerase fidelity by two orders of magnitude [47]. Some DNA polymerases, especially those in higher eukaryotes, have non-enzymatic domains or motifs which are important for protein-protein interactions. Currently, it is not clear whether or how many of these peptide regions affect polymerase activity. Moreover, the presence of additional protein co-factors can sometimes alter the mechanism and fidelity of DNA polymerization [48].
Chemical reaction for nucleotidyl transfer All known DNA polymerases catalyze phosphodiester bond formation using the two metal-ion mechanism in which magnesium is proposed to be the metal ion in vivo due to its cellular abundance (Figure 1.2) [41, 49]. The two catalytic magnesium ions are designated A and B. Two or three conserved carboxylate residues in the palm subdomain are essential for the coordination and orientation of the magnesium ions. Magnesium A lowers the pKa of the 3′-hydroxyl on the primer terminus so that deprotonation leads to in-line nucleophilic attack on the α-phosphate of an incoming nucleotide. The nucleotidyl transfer reaction proceeds through a pentacoordinated intermediate in an associative-like transition state [50, 51]. The pentacovalent transition state is stablized by both magnesium ions A and B. Then, magnesium B stabilizes the negatively-charged pyrophosphate leaving group. Once the pyrophosphate and magnesium ions dissociate, the DNA polymerase prepares for the next catalytic cycle. The crystal structure of Pol β in complex with gapped DNA, a non-hydrolyzable nucleotide analog [i.e. 2′- deoxyuridine-5′-(α,β)-imido triphosphate (dUPNPP)], and two magnesium ions is the best structural representation of the requisite substrates being aligned before execution of the chemical reaction within a DNA polymerase active site (Figure 1.2) [52]. Please note, due to the left-hand nature of Pol β, the assignments of magnesium ions A and B are reversed in Figure 1.2 part (B) [53].
Kinetic mechanism of DNA polymerization DNA polymerases follow a minimal kinetic mechanism that consists of 6 steps (Scheme 1.1) [54]. First, the DNA polymerase binds to the DNA substrate (Step 1) and then to a
5 nucleotide substrate (Step 2). Nucleotide binding induces a conformational change (Step 3) so that the ternary complex is distinct from the one in the ground state. This conformational change may involve the major movement of a polymerase subdomain (known as the “open” to “closed” transition), the minor repositioning of active site residues, or a combination of both [45, 46, 55]. Such conformational rearrangments configure the active site for the chemical step of nucleotidyl transfer (Step 4). Following phosphodiester bond formation, a conformational change occurs (Step 5) to rapidly release the pyrophosphate product (Step 6). The catalytic cycle can continue via polymerase translocation or the DNA polymerase can dissociate from the extended DNA product.
To elucidate the kinetic pathway, kinetic assays have been designed to measure the rate and equilibrium dissociation constants of the elementary steps of DNA polymerization. These kinetic parameters vary for each DNA polymerase. Overall, the slowest step in the reaction pathway is the release of the extended DNA product from the polymerase. However, there is a long-standing debate about the identity of the rate-limiting step of nucleotide incorporation, i.e. a conformational change (Step 3) versus phosphodiester bond formation (Step 4) [54, 56, 57]. Recent kinetic and structure-based evidence suggests the rate-limiting step is a conformational change involving local active site rearrangements rather than a dramatic shift in the positioning of the fingers subdomain [58-62].
1.3 Kinetic Basis of Nucleotide Selectivity and Fidelity
During DNA synthesis, DNA polymerases select the correct nucleotide substrate from a pool of four similar dNTPs and most of these enzymes insert the nucleotide in a template- dependent manner. To understand the kinetic basis of nucleotide selectivity, transient state kinetic assays have been the best kinetic tool for measuring the two kinetic parameters that define polymerase specificity (kp/Kd): the equilibrium dissociation
6 constant (Kd) and the maximum rate of nucleotide incorporation (kp) of an incoming nucleotide [63]. In general, DNA polymerases preferentially incorporate a correct nucleotide via tighter nucleotide binding affinity and faster rate of incorporation while misincorporations are disfavored due to weaker nucleotide binding affinity and slower rate of incorporation. These kinetic steps serve as fidelity checkpoints for the DNA polymerase. More specificially, this nucleotide substrate specificity is manifested in complex polymerase-substrate interactions and DNA-nucleotide interactions that are established during the nucleotide binding and incorporation steps. Important interactions for dNTP substrate recognition include the following: (i) Watson-Crick hydrogen bonding between the base of the incoming nucleotide and DNA template, (ii) base stacking between the base of the incoming nucleotide and primer terminus, (iii) minor groove hydrogen bonds, electrostatic interactions, and van der Waals interactions between active site residues and the substrates, (iv) steric clashes or an empty cavity in the active site due to sub-optimal geometry of the base pair, and (v) the effects of water molecules [64, 65]. Many of these interactions are critical for efficient nucleotide incorporation but are not essential for catalysis. Establishing such intimate contacts among a network of molecules likely play a role in altering the conformational dynamics of a polymerase as it achieves proper substrate alignment.
High nucleotide selectivity enforced by the polymerase domain provides the greatest contribution to the overall fidelity of DNA replication, thereby maintaining the integrity of an organism’s genome. DNA polymerases with high fidelity are the replicative polymerases in families A, B, C, and D. The typical error rate for these enzymes is one misincorporation for every 10,000 to 1,000,000 nucleotide incorporations which equates to a fidelity range of 10-4 to 10-6 [66]. The exonuclease domain can further enhance fidelity to 10-6 to 10-8. In contrast, X- and Y-family DNA polymerases exhibit a low fidelity of 100 to 10-5 [66]. For the Y-family enzymes, the fingers and thumb subdomains are smaller than those in the A- and B-family polymerases, therefore, the polymerase active site is more solvent exposed and there are fewer contacts with the DNA and dNTP substrates [67]. When fidelity is one, this indicates that an incorrect base pair is favored
7 over the canonical Watson-Crick base pair. Pol ι and Rev1, both Y-family members, exhibit such low fidelity since they do not favor the formation of a canonical Watson- Crick base pair. Pol ι prefers a dGTP:dT base pair over dATP:dT while Rev1 uses an arginine residue to direct dCTP incorporation regardless of the template base [68-72]. Thus, Y-family enzymes have the capacity to be potentially mutagenic inside a cell.
1.4 Focus of Dissertation
This dissertation focuses on one B-family member [Sulfolobus solfataricus P2 DNA polymerase B1 (PolB1)], two X-family members (human Pols β and λ), and six Y-family members [human Pol η, Pol ι, Pol κ, Rev1, Saccharomyces cerevisiae Pol η, and Sulfolobus solfataricus P2 DNA polymerase IV (Dpo4)]. These selected DNA polymerases have different primary roles in preserving the integrity of genomic DNA. Due to the relatively recent discovery of these selected B-, X-, and Y-family DNA polymerases, these enzymes have not undergone rigorous biochemical characterization. In addition, the precise biological functions of human Pols λ, ι, κ, and Rev1 are uncertain. Using a transient state kinetic approach, various mechanistic questions about nucleotide selectivity and fidelity of these nine DNA polymerases have been examined which has led to a better understanding of their structure-function relationships. The divergent kinetic properties observed for these enzymes support the notion by Joyce and Benkovic that DNA polymerases do not conform to a unified kinetic and structural mechanism of DNA polymerization [54].
PolB1 replicates the 2,992,245 base pair genome of S. solfataricus, an aerobic crenarchaeon that grows optimally at 80 °C and pH 2-4 [73]. The overall fidelity of PolB1 was estimated to be 10-6 to 10-8, whereby most of the discrimination between a correct and incorrect nucleotide occurred during the nucleotide binding step (370-fold on average) rather than the incorporation step (10-fold on average). In addition, four lines of
8 kinetic evidence supported the existence of a conformational change preceding nucleotidyl transfer during the incorporation of a correct dNTP.
Pol β and Pol λ share 34% sequence similarity, and their main role is in DNA repair. Both enzymes possess two enzymatic functions: dRPase and polymerase activity [74, 75]. Pol β is the central DNA polymerase in base excision repair in vivo while Pol λ is postulated to serve as a backup for Pol β [76-78]. However, Pol λ has unique N-terminal domains that expand Pol λ’s participation into other cellular pathways: non-homologous end joining and V(D)J recombination [30].
During DNA synthesis, most DNA polymerases select against ribonucleotides via a steric clash between the ribose 2′-hydroxyl group and the bulky side chain of an active site residue. Human Pol λ prefered to incorporate deoxyribonucleotides over ribonucleotides by 3,000- to 50,000-fold. Moreover, Pol λ used a novel sugar selection mechanism to discriminate against ribonucleotides, whereby the ribose 2′-hydroxyl group was excluded mostly by a backbone segment and slightly by the side chain of Y505. Such a steric clash was further demonstrated to be dependent upon the size and orientation of the substituent covalently attached at the ribonucleotide C2′ position.
Pre-steady-state kinetic studies have shown that the Pol λ•DNA complex binds both correct and incorrect nucleotides 130-fold tighter on average than the DNA polymerase β•DNA complex, although, the base substitution fidelity of both polymerases is 10-4 to 10-5. To better understand Pol λ's tight nucleotide binding affinity, we created single- substitution and double-substitution mutants of Pol λ to disrupt the interactions between active site residues and an incoming nucleotide or a template base. Single-turnover kinetic assays showed that Pol λ binds to an incoming nucleotide via cooperative interactions with active site residues (R386, R420, K422, Y505, F506, A510, and R514). Disrupting protein interactions with an incoming correct or incorrect nucleotide impacted binding to each of the common structural moieties in the following order: triphosphate >> base > ribose. In addition, the loss of Watson-Crick hydrogen bonding between the 9 nucleotide and the template base led to a moderate increase in Kd. The fidelity of Pol λ was maintained predominantly by a single residue, R517, which has minor groove interactions with the DNA template.
The base excision repair pathway coordinates the replacement of 1 to 10 nucleotides at sites of single-base lesions. This process generates DNA substrates with various gap sizes which can alter the catalytic efficiency and fidelity of a DNA polymerase during gap- filling DNA synthesis. Here, we quantitatively determined the substrate specificity and base substitution fidelity of human Pol λ and human Pol β, as each enzyme filled nucleotide gaps of varying size. Pol λ incorporated a correct nucleotide with relatively high efficiency until the gap size exceeded 9 nucleotides. Unlike Pol λ, Pol β did not have an absolute threshold on gap size, as the catalytic efficiency for a correct dNTP gradually decreased as the gap size increased from 2 to 10 nucleotides and then recovered for non- gapped DNA. Surprisingly, an increase in gap size resulted in lower polymerase fidelity for Pol λ, and this downregulation of fidelity was controlled by its non-enzymatic N- terminal domains. Overall, Pol λ was up to 160-fold more error-prone than Pol β, thereby suggesting Pol λ would be more mutagenic during long gap-filling DNA synthesis. In addition, dCTP was the preferred misincorporation for Pol λ. This nucleotide preference was shown to be dependent upon the identity of the adjacent 5′-template base. Our results suggested that both Pol λ and Pol β would catalyze nucleotide incorporation with the highest combination of efficiency and accuracy when the DNA substrate contains a single-nucleotide gap.
In the presence of 8-oxo-7,8-dihydro-2′-deoxyguanosine (8-oxodG) damage, many DNA polymerases exhibit a dual coding potential which facilitates efficient incorporation of dCTP via Watson-Crick base pairing or dATP via Hoogsteen base pairing. This also holds true for the insertion of 8-oxodGTP opposite template bases dC and dA. Employing single-turnover kinetic methods, we determined which nucleotide and template base was preferred when human Pol λ and human Pol β encounter 8-oxodG and 8-oxodGTP, respectively. While Pol β preferentially incorporated dCTP over dATP, Pol λ
10 incorporated dCTP or dATP with essentially equal efficiency and probability when opposite 8-oxodG in single-nucleotide gapped DNA. Insertion of 8-oxodGTP by both DNA polymerases λ and β occurred predominantly against template dA.
For the Y-family polymerases, their major function is translesion DNA synthesis. These low-fidelity enzymes gain access to the replication fork by “switching” with a replicative DNA polymerase at sites of DNA damage [79]. They can bypass a DNA lesion in an error-free or error-prone manner. Human and yeast Pol η specifically bypass sunlight- induced pyrimidine-pyrimidine dimers [37]. Unfortunately, the bypass of other DNA lesions by a specific DNA polymerase or combination of DNA polymerases remains elusive. Besides translesion synthesis, human Pol η, Pol ι, Pol κ, and Rev1 have been implicated in somatic hypermutation, homologous recombination, base excision repair, and/or nucleotide excision repair [39, 80].
Full-length (1-632) and truncated (1-513) Saccharomyces cerevisiae Pol η were kinetically characterized in order to determine whether the C-terminal residues (514-632) alter the activity of the polymerase domain. The DNA binding affinity of full-length Pol η was 2-fold tighter than the truncated construct, and the fidelity was up to 3-fold higher for full-length Pol η. These mild kinetic effects suggested the C-terminus does not have a significant impact on Pol η when synthesizing undamaged DNA. Overall, truncated Pol η discriminated between a correct and incorrect nucleotide more during the incorporation step (50-fold on average) than the nucleotide binding step (18-fold on average). Lastly, blunt end additions of dATP or pyrene 5′-triphosphate indicated that base stacking is important for binding an incorrect dNTP.
Human Rev1 is a unique nucleotidyl transferase that uses a protein template-directed mechanism to preferentially instruct dCTP incorporation. Nonetheless, this high substrate specificity is dependent upon both nucleic acid substrates: an incoming dCTP and a template base dG. The order of nucleotide incorporation with a template dG corresponded to the number of hydrogen bonds that can form between the incoming
11 dNTP and R357. The extremely low base substitution fidelity of human Rev1 (100 to 10-5) was due to the preferred misincorporation of dCTP with templating bases dA, dT, and dC over the complementary dNTP. Lastly, human Rev1 discriminated between a ribonucleotide (rCTP) and a deoxyribonucleotide (dCTP) by 280-fold, and the sugar selectivity of human Rev1 was sensitive to both the size and orientation of the 2′- substituent of a ribonucleotide.
Dpo4 is the only Y-family polymerase encoded in the genome of S. solfataricus [73]. It has served as a model Y-family DNA polymerase for elucidating the mechanistic basis of translesion synthesis. Here, Dpo4 was used to establish a model for the bypass of a
double-base lesion, cis-[Pt(NH3)2{d(GpG)-N7(1),-N7(2)}] intrastrand cross-link. Dpo4 was able to bypass this cisplatin-DNA adduct, although, significant accumulation of intermediate DNA products were observed at the site of the lesion. This pausing of Dpo4 was due to a weaker DNA binding affinity (up to 3-fold), reduced nucleotide incorporation efficiency (up to 860-fold), and less than 7% of the Dpo4•cisplatin-DNA complexes being bound in a productive mode. In addition, the bypass process was described as error-prone, since the fidelity of Dpo4 dropped up to two orders of magnitude.
Nucleoside analogs are an important class of antiviral drugs used to manage the infections of the hepatits B virus and human immunodeficiency virus. Unfortunately, these drugs cause unwanted side effects, and the molecular basis of drug toxicity is not fully understood. Putative routes of drug toxicity include the inhibition of human nuclear and mitochondrial DNA polymerases. However, it remains to be determined whether the phosphorylated nucleotde analogs are substrates for the recently-discovered human X- and Y-family DNA polymerases. Using pre-steady state kinetic techniques, we have measured the substrate specificity constants for human DNA polymerases β, λ, η, ι, κ, and Rev1 incorporating the active, 5′-phosphorylated forms of adefovir, emtricitabine, entecavir, lamivudine, tenofovir, telbivudine, and zidovudine. For the six enzymes, all of the drug analogs were incorporated less efficiently (2- to >126,000-fold) than the
12 corresponding natural nucleotides, usually due to a weaker binding affinity and a slower rate of incorporation for the incoming nucleotide analog. Our kinetic results suggested nucleotide analog insertion catalyzed by human X- and Y-family DNA polymerases is a potential mechanism of drug toxicity and established a structure-function relationship for designing improved nucleoside analogs.
Extensive kinetic, structural, and computational characterization of DNA polymerases has sparked a debate about the rate-limiting step of nucleotide incorporation catalyzed by these enzymes. Possible rate-limiting steps include a pre-chemistry conformational change, local active site rearrangements, or the chemical reaction. To help resolve the “rate-limiting step” debate, the mechanism of nucleotide incorporation catalyzed Pol β or Pol λ was examined using three kinetic probes: (i) alpha-thio elemental effect, (ii) pulse- chase and pulse-quench experiments, and (iii) Arrhenius activation energy barrier. Our preliminary results supported a rate-limiting active site rearrangement for a correct nucleotide incorporation.
13 1.5 Table
Table 1.1 Classification and function of human DNA polymerases. DNA Polymerase Functiona polymerase family Pol γ A Mitochondrial DNA replication and repair Pol ν A DNA repair and translesion synthesis? Pol θ A DNA repair, translesion synthesis, and antibody diversity? Pol α B DNA replication and repair Pol δ B DNA replication and repair Pol ε B DNA replication and repair Pol ζ B DNA repair, translesion synthesis, and antibody diversity? Pol β X DNA repair Pol λ X DNA repair and antibody diversity? Pol μ X DNA repair and antibody diversity? TdT X Antibody diversity Pol σ X Sister chromatid cohesion Pol η Y DNA repair, translesion synthesis, and antibody diversity? Pol ι Y DNA repair and translesion synthesis? Pol κ Y DNA repair and translesion synthesis? Rev1 Y Translesion synthesis and antibody diversity? aA “?” indicates proposed biological functions.
14 1.6 Figures
A B C
D E F
Figure 1.1 X-ray crystal structures of DNA polymerases. A representative member is shown for each family: (A) Klenow fragment of Thermus aquaticus DNA polymerase I (PDB 3KTQ) for family A, (B) RB69 DNA polymerase (PDB 1IG9) for family B, (C) E. coli DNA Pol III (PDB 2HNH) for family C, (D) Pfu (PDB 2JGU) for family D, (E) human Pol β (PDB 1BPY) for family X, and (F) human Pol η (PDB 3MR2) for family Y. Important structures, if present, are color coded as follows: N-terminal domain is a light gray ribbon, fingers subdomain is a blue ribbon, palm subdomain is a red ribbon, thumb subdomain is a purple ribbon, the DNA primer is light gray, the DNA template is dark gray, the incoming nucleotide is in gray sticks, the metal ions are orange spheres, and the purple ribbon is the 3′ → 5′ exonuclease domain in parts (A, B, C, and D), the 8 kDa domain in part (E), and the little finger subdomain in part (F).
15 A B
Figure 1.2 The two metal-ion mechanism of nucleotidyl transfer. (A) The chemical reaction that occurs in a polymerase active site is shown with residue numbers for E. coli Pol I. This figure is from references [41, 81]. (B) The two catalytic magnesium ions (green spheres) are coordinated with the 3′-OH of the primer terminus (gray sticks), triphosphate moiety of the incoming nucleotide dUPNPP (gray sticks), three catalytic aspartate residues (lavendar sticks), and water molecules (purple spheres) based on the crystal structure of human Pol β•gap DNA•dUPNPP (PDB 2FMS).
16 1.7 Scheme
Scheme 1.1
17 Chapter 2: Polymerization Fidelity of Sulfolobus solfataricus P2 DNA Polymerase B1
2.1 Introduction
DNA polymerases are known to catalyze genomic replication, DNA damage repair, and DNA lesion bypass. Since the discovery of Escherichia coli DNA polymerase I in the 1950s, six families (A, B, C, D, X, and Y) of DNA polymerases have been discovered in all three domains of life [82, 83]. The B-family is mainly composed of eukaryotic replicative DNA polymerases, although, some have been identified in archaea, bacteria, phages, and viruses [83]. Among the B-family members, archaeal DNA polymerases have been studied as model enzymes in order to understand DNA replication in the more complicated eukaryotic systems. In addition to template-dependent polymerase activity, most B-family enzymes also possess 3′ → 5′ exonuclease activity which detects and removes errors catalyzed by the polymerase domain. Such an editing function enhances the fidelity of replicative DNA polymerases by about 200-fold [47, 84]. The overall fidelity of numerous replicative DNA polymerases, estimated using either steady state kinetics [85, 86], pre-steady state kinetics [85, 87-92], or M13mp2-based forward and reversion mutation assays [93-96], is in the range of 10−6 to 10−8, i.e. one error per 106 to 108 nucleotide incorporations. This high polymerization fidelity is essential for maintaining genomic stability from generation to generation [97].
Sulfolobus solfataricus is an aerobic crenarchaeon that metabolizes sulfur and grows optimally at 80 °C and pH 2-4 [73]. The genome of S. solfataricus P2 has been completely sequenced and contains 2,992,245 base pairs [73]. This hyperthermophile encodes only four DNA polymerases: three in the B-family (DNA polymerases B1 18 (PolB1), B2, and B3) and one in the Y-family (DNA polymerase IV, Dpo4). Among these four DNA polymerases, DNA polymerases B2 and B3, which are identified based on sequence homology, have not been shown to be active in vitro [73, 98, 99]. In contrast, Dpo4 catalyzes DNA polymerization and bypasses a myriad of DNA lesions [100]. Its polymerase active site is flexible, solvent accessible, and can accommodate bulky DNA lesions [101-103]. The polymerization fidelity of Dpo4 with undamaged DNA has been measured to be in the range of 10−3 to 10−4, or one error per 103 to 104 nucleotide incorporations, by utilizing both kinetic and M13mp2-based fidelity assays [100, 104, 105]. PolB1, a replicative DNA polymerase [106-108], has been reported to possess both DNA polymerase and 3' → 5' exonuclease activities in vitro [108]. The crystal structure of apo PolB1 shows that this enzyme, like A- and B-family DNA polymerases, contains an N-terminal proofreading 3' → 5' exonuclease domain and a C- terminal “right-hand” DNA polymerase domain [109]. Thus, the polymerization fidelity of PolB1 is expected to be high but has not been reported. In this paper, we employed pre-steady state kinetic methods to determine the base substitution fidelity and mismatch extension fidelity of PolB1. The effect of reaction temperature and pH on the polymerase selectivity and the contribution of the 3′ → 5′ exonuclease activity to the polymerization fidelity of PolB1 were also investigated.
2.2 Materials and Methods
Materials Materials were purchased from the following companies: [γ-32P]ATP, PerkinElmer (Boston, MA); dNTPs, GE Healthcare (Piscataway, NJ); Biospin columns, Bio-Rad Laboratories (Herclues, CA); OptiKinase, USB (Cleveland, OH); Quikchange XL Site- Directed Mutagenesis kit, Stratagene (La Jolla, CA); Wizard Plus SV Miniprep kit, Promega (Madison, WI).
19 Mutagenesis, overexpression, and purification of wild-type PolB1 and the exonuclease- deficient mutant The plasmid pET33b-PolB1, a generous gift from Dr. Hong Ling at the University of
Western Ontario, Canada, encodes S. solfataricus P2 PolB1 fused to a His6-tag at the C- terminus. Two rounds of site-directed mutagenesis were performed to generate the exonuclease-deficient triple-point mutant (D231A, E233A, and D318A). First, a double mutant (D231A and E233A) was constructed using the wild-type PolB1 plasmid as a template. Then, the resulting plasmid was used to introduce the D318A mutation. The DNA sequence of the mutant plasmids was confirmed by sequencing (OSU Plant Microbe Genomics Facility).
Wild-type plasmid pET33b-PolB1 was transformed into E. coli expression strain BL21(DE3) Rosetta cells. An overnight culture of the Rosetta cells carrying the wild-type PolB1 plasmid was used to inoculate Luria-Bertani medium containing 50 μg/mL kanamycin and 25 µg/mL chloramphenicol. Cells were grown at 37 °C. Once the OD600 of the cells reached 0.6, cultures were induced with 0.15 mM IPTG and incubated at 30
°C until the OD600 reached 1.7. Induced cells were harvested and resuspended in buffer A
[10 mM KH2PO4 (pH 7.0), 50 mM NaCl, 10 mM MgCl2, 10% glycerol, 0.1% 2- mercaptoethanol, and 1 mM imidazole]. A protease inhibitor cocktail tablet (Roche) and PMSF (1 mM) were added to the cell paste. The cells were then lysed by passing them through a French Press cell three times at 16,000 psi. The cell lysate was clarified by ultracentrifugation (35,000 rpm for 40 min). Clear lysate was subjected to heat denaturation at 74 °C for 14 min to precipitate thermolabile E. coli proteins which were subsequently removed by ultracentrifugation. The supernatant was incubated with IMAC Fastflow6 Ni2+-charged resin (GE Healthcare) for 3 hours in the presence of 10 mM imidazole and the PolB1-bound resin was packed into a column. After washing with buffer B [10 mM KH2PO4 (pH 7.0), 350 mM NaCl, 35 mM imidazole, 10% glycerol, 0.05% 2-mercaptoethanol], PolB1 was eluted using a linear gradient of 35–500 mM imidazole in buffer B. Fractions containing PolB1 were pooled and then dialyzed against buffer C [50 mM Tris-HCl (pH 7.5 at 25 °C), 50 mM NaCl, 1 mM EDTA, 10% glycerol,
20 0.1% 2-mercaptoethanol). The dialyzed protein solution was passed through a HiTrap Q anion exchange column (GE Healthcare). The loading elute was applied to a HiTrap SP cation exchange column (GE Healthcare). The column-bound PolB1 was eluted using a gradient of 100-1000 mM NaCl in buffer C. The fractions containing PolB1 were pooled and dialyzed against buffer D [25 mM HEPES (pH 7.0 at 4 °C), 150 mM NaCl, 1 mM EDTA, 1 mM DTT, 50% glycerol]. The dialyzed protein solution was then concentrated and stored at –80 °C. The apparent homogeneity of purified PolB1 (>95%) was assessed based on Coomassie Blue-stained SDS-PAGE gels. The concentration of the purified wild-type PolB1 was measured spectrophotometrically at 280 nm using the calculated extinction coefficient of 124,785 M-1cm-1. Similarly, the exonuclease-deficient triple- point mutant (D231A, E233A, and D318A) of PolB1 was overexpressed, purified, and quantitated.
DNA substrates The synthetic oligonucleotides listed in Table 2.1 were purchased from Integrated DNA Technologies, Inc. (Coralville, IA) and purified as described previously [105]. The primer strand 21-mer was 5′-radiolabeled with [γ-32P]ATP and Optikinase and was annealed to the appropriate 41mer (Table 2.1) as described previously [105].
Reaction buffers All experiments, if not specified, were performed in buffer A which contains 50 mM
HEPES (pH 7.5 at all temperatures), 15 mM MgCl2, 75 mM NaCl, 5 mM DTT, 10% glycerol, and 0.1 mg/ml BSA. Reactions were carried out at temperatures ranging from 19 to 50 °C using either a rapid chemical-quench flow apparatus (KinTek, PA) or via manual quench. For buffer optimization and pH-dependent fidelity studies at 37 °C, assays were carried out in 25 mM MES buffer for pH between 6.0-7.0, 50 mM HEPES for pH of 7.5, 25 mM Tris-HCl for pH of 8.0 and 8.5, and 25 mM Glycine for pH of 9.0 and 10.0. For both DNA polymerization and excision experiments, the enzyme•DNA
complex was not pre-equilibrated in the presence of MgCl2.
21 Polymerase and exonuclease single-turnover assays For polymerization reaction assays, PolB1 (120 nM) was pre-incubated with D-DNA (30 nM) in buffer A before the reaction was initiated with varying concentrations of a single dNTP•Mg2+ (1-2400 µM). For DNA excision assays, PolB1 (400 nM) was pre- equilibrated with DNA (100 nM), and the reaction was initiated with the addition of Mg2+ (15 mM) in buffer A.
Product analysis Reaction products were analyzed by sequencing gel electrophoresis (17% acrylamide, 8M urea, 1×TBE running buffer) and quantitated using a Typhoon TRIO (GE Healthcare) and ImageQuant software (Molecular Dynamics).
Data analysis All kinetic data were fit by nonlinear regression using KaleidaGraph (Synergy Software). Data from single nucleotide incorporation experiments under single-turnover conditions were fit to equation 1,
[Product] = A[1 – exp(–kobst)] (1)
where A and kobs represent the reaction amplitude and observed rate, respectively. Data
from the plot of kobs against nucleotide concentrations were fit to equation 2,
kobs = kp[dNTP]/{Kd + [dNTP]} (2)
where kp is the maximum rate of nucleotide incorporation and Kd is the equilibrium
dissociation constant of a dNTP. When Kd is too large, the plot of kobs versus nucleotide concentrations was fit to equation 3,
kobs = (kp/Kd)[dNTP] (3) which extracted the substrate specificity constant, kp/Kd. Data from DNA excision experiments under single-turnover conditions were fit to equation 4,
[Product] = Aexp(–kexot) (4) where A and kexo represent the reaction amplitude and observed DNA excision rate, respectively.
22 2.3 Results
Generation of the 3′ → 5′ exonuclease-deficient mutant of PolB1 It has been demonstrated previously that PolB1 (882 amino acid residues and 101.2 kDa) from S. solfataricus P2 possesses a strong 3′ → 5′ exonuclease activity [108], which would complicate the interpretation of the kinetic data of nucleotide incorporation. To eliminate this problem, we constructed an exonuclease-deficient mutant of PolB1 (PolB1 exo-) by substituting an alanine for conserved residues D231, E233, and D318, which are located in the conserved sequence motifs Exo I-II among the B-family DNA polymerases [106]. On the basis of X-ray crystal structural analysis [109] and primary sequence alignment [106], these three residues and D413 bind two Mg2+ ions required for catalysis. Following the successful site-directed mutagenesis and purification processes, we examined the activity of both the polymerase and exonuclease functions. A pre-incubated solution of PolB1 (wild-type or exo-) and 5′-[32P]-labeled D-1 DNA (Table 2.1) was mixed with either 50 µM dTTP•Mg2+, 50 µM dCTP•Mg2+, or Mg2+. Our data confirmed that wild-type PolB1 displayed efficient exonuclease activity, for significant product degradation (<21-mer) was observed under all three reaction conditions (Figure 2.1). In contrast, elongated DNA products accumulated for PolB1 exo- in the presence of both correct (dTTP) and incorrect (dCTP) nucleotides (Figure 2.1). Furthermore, in the absence of nucleotides, only substrate (21-mer) was present for PolB1 exo- for reaction times of 15 sec to 3 hours (data not shown). Together, these results showed that PolB1 exo- lacked 3′ → 5′ exonuclease activity and its polymerase activity remained intact. Additionally, if one or two of the three residues (D231, E233, and D318) was mutated to alanine, the 3′ → 5′ exonuclease activity of the mutants decreased but was not completely eliminated (data not shown).
Optimization of the reaction conditions for the polymerase activity of PolB1 To optimize the reaction conditions for the polymerase activity of PolB1, all reaction components were kept constant while the concentrations of Mg2+ and NaCl and buffer pH were individually altered. A pre-incubated solution of 120 nM PolB1 exo- and 30 nM 5'-
23 [32P] D-1 (Table 2.1) was reacted with 100 μM dTTP•Mg2+ at 37 oC for various times before being quenched with 0.37 M EDTA. The time dependence of product
concentration was fit to equation 1 (see 2.2) to obtain the observed reaction rate, kobs, and reaction amplitude (data not shown). Figure 2.2 shows that the kobs values varied depending on the concentrations of Mg2+ and NaCl and with the buffer pH. In comparison, the reaction amplitude was close to 30 nM under all conditions except at 70 2+ mM Mg , 250 mM NaCl, and pH 10. On the basis of the kobs values and reaction amplitudes for single correct nucleotide incorporation, we concluded that the optimal
reaction conditions for PolB1 exo- were 15 mM MgCl2 (Figure 2.2), 75 mM NaCl (Figure 2.2), and pH 7.5 (Figure 2.2). Under non-optimum conditions, dTTP
incorporation occurred at significantly lower kobs values. Therefore, all kinetic assays presented herein, if not specified, were performed in the optimized buffer A which o contains 50 mM HEPES (pH 7.5 at 37 C), 15 mM MgCl2, 75 mM NaCl, 5 mM DTT, 10% glycerol, and 0.1 mg/mL BSA. Please note, most of the assays shown in this work and our accompanying paper [110] were performed at 37 °C, rather than 80 °C which is the in vivo temperature for PolB1. The reaction temperature of 37 °C was selected because (i) the rapid chemical-quench apparatus can function properly, (ii) the rates of correct nucleotide incorporation are within the range that can be accurately measured by the rapid chemical-quench apparatus, (iii) most of the DNA substrate 21/41-mer exist as a duplex, rather than single-stranded oligomers, during polymerization, and (iv) our fidelity and mechanism studies of Dpo4 revealed similar findings at both 37 and 56 °C [60, 105, 111].
Substrate specificity and base substitution fidelity of PolB1 exo- at 37 oC It was expected that PolB1, like other replicative DNA polymerases, incorporates correct nucleotides with much higher substrate specificity than it incorporates incorrect
nucleotides. To determine the substrate specificity constant (kp/Kd) of an incoming
nucleotide, we measured the maximum incorporation rate constant (kp) and the apparent
equilibrium dissociation constant (Kd) of an incoming nucleotide [112]. These two pre- steady state kinetic parameters can be determined through the nucleotide concentration
24 dependence of the observed incorporation rate (kobs). Figure 2.3 is a representative example of these kinetic measurements determined under single-turnover reaction conditions whereby the concentration of the enzyme was 4-fold greater than the DNA concentration in order to ensure that almost all of the DNA molecules were bound by PolB1 exo-. In Figure 2.3, a pre-incubated solution of 120 nM PolB1 exo- and 30 nM 5′- [32P]-labeled D-6 (Table 2.1) was reacted with increasing concentrations of correct dCTP•Mg2+ in buffer A for various times prior to being quenched with 0.37 M EDTA. At each dCTP concentration, the product concentration was plotted as a function of the
reaction time, and the data were fit to equation 1 (see 2.2) to yield a kobs (Figure 2.3). In
Figure 2.3, the extracted kobs values were plotted versus the corresponding dCTP concentration, and the data were subsequently fit to equation 2 (see 2.2) to yield a kp of -1 5.7 ± 0.1 s and a Kd of 8.2 ± 0.5 μM for dCTP incorporation. The substrate specificity constant of dCTP incorporation into D-6 was then calculated to be 0.70 μM-1s-1.
Similarly, we measured the kp, Kd, and kp/Kd of the other fifteen possible single nucleotide incorporations (correct and incorrect) into the four DNA substrates in Table 2.1 (data not shown), and the kinetic parameters are listed in Table 2.2. For several
incorrect nucleotides, we could not determine the individual kp and Kd values due to their
extremely weak apparent ground-state binding affinity (1/Kd) for an incoming dNTP.
Instead, we determined the kp/Kd values by fitting the plots of kobs versus nucleotide concentration to equation 3 in 2.2 (data not shown). On average, incorrect nucleotides had ~370-fold weaker apparent ground-state binding affinities and were incorporated
with 11-fold slower kp values than correct nucleotides. The calculated substrate specificity constants indicated that correct nucleotides were incorporated with 103 to 105- fold higher efficiency than incorrect nucleotides. In addition, we calculated the base substitution fidelity of PolB1 exo- to be in the range of 10-4 to 10-6 (Table 2.2), or one mistake per 104 to 106 nucleotide incorporations at 37 oC.
Effect of temperature on the base substitution fidelity of PolB1 exo- at pH 7.5 Since S. solfataricus P2 grows optimally at 80 °C [73], it is important to kinetically determine the fidelity of PolB1 at its physiological temperature. However, this cannot be
25 accomplished here because the rapid chemical-quench apparatus cannot be operated at 80 oC, and correct nucleotide incorporations catalyzed by PolB1 exo- at such a high temperature are predicted to be finished within the mixing dead time (1-3 ms) of the apparatus. To estimate the fidelity of PolB1 at 80 oC, we decided to determine the temperature dependence of the fidelity. Under single-turnover conditions, PolB1 exo- (120 nM) and 30 nM 5′-[32P] D-1 (Table 2.1) were first pre-incubated on ice and then equilibrated at 19, 26, 32, 44, or 50 °C. Such a gradual increase of the incubation temperature helped to stabilize the DNA duplex at the polymerase active site of PolB1 exo-, especially at temperatures exceeding 37 oC. The pre-incubated solutions of D-1 and PolB1 were then reacted with increasing concentrations of correct dTTP•Mg2+ or incorrect dATP•Mg2+ for various times at the appropriate temperature. The reactions were quenched and analyzed as described above. The kinetic parameters for the incorporations of correct dTTP and incorrect dATP into D-1 are listed in Table 2.3. The
kp for both dTTP and dATP increased dramatically with reaction temperature while the
apparent Kd changed slightly. Interestingly, the kp/Kd for both correct and incorrect nucleotide incorporations increased with a similar magnitude at each temperature, leading to a modest change of 5-fold in the base substitution fidelity of PolB1 exo- from 19 to 50 ºC (Figure 2.4).
Effect of pH on the base substitution fidelity of PolB1 exo- at 37 oC Although S. solfataricus P2 grows optimally at pH 2-4 [73], its cytoplasmic pH, which is unknown at present, is expected to be near neutral so that genomic stability is maintained. Because the cytoplasmic pH of S. solfataricus P2 may not be 7.5 as in our optimized reaction buffer, it is important to determine the effect of pH on the polymerase fidelity of PolB1. Using similar kinetic assays described above, we measured the kinetic parameters (Table 2.4) for the incorporations of correct dTTP and incorrect dATP into D-1 (Table 2.1) at 37 °C catalyzed by PolB1 exo- under different buffer pH values (6.0, 6.5, and 8.5). -1 For dTTP incorporation, the kp (5.8 to 8.9 s ) changed insignificantly while the apparent
Kd increased from 3.2 to 21 μM when the buffer pH changed from 6.0 to 8.5. In contrast, -1 the kp for the incorrect dATP incorporation changed significantly from 0.19 to 2.0 s with
26 an increase of buffer pH from 6.0 to 8.5. However, the pH did not affect the apparent Kd of dATP until the pH was 8.5, whereby a 6-fold increase was observed from a pH of 6.5 to 8.5. Interestingly, when the buffer pH changed from 6.0 to 8.5, the calculated substrate specificity constant for correct dTTP incorporation decreased by ~10-fold while the kp/Kd for dATP misincorporation increased by ~4-fold, leading to a 36-fold decrease in the base substitution fidelity of PolB1 exo- (Table 2.4 and Figure 2.5). Thus, PolB1 exo- had higher polymerase efficiency and fidelity at a relatively low pH.
Mismatch extension fidelity of PolB1 exo- at 37 oC When PolB1 exo- misincorporates a nucleotide, it may continue polymerization to bury the misincoporation by either a correct or incorrect nucleotide. Under similar single- turnover conditions as described above, we measured the kinetic parameters (Table 2.5) for each of the four possible incorporations into M-1 which contains a mismatched base pair dA:dA at its primer-template junction (Table 2.1). Among the four nucleotides, correct dCTP was incorporated by PolB1 exo- with the highest substrate specificity
constant (Table 2.5). Notably, the kp, apparent Kd, and kp/Kd values of dCTP are comparable to the corresponding values of several misincorporations in Table 2.2, e.g. incorrect dATP into D-8. While the incorporations of incorrect nucleotides dATP and dGTP were not observed even after 3 hours (data not shown), dTTP misincorporation into M-1 was readily detected, and the substrate specificity constant was determined to be -7 3.8 x 10 (Table 2.5). However, the kp and apparent Kd values for dTTP misincorporation were not determined due to an extremely weak apparent ground-state binding affinity (see above). Based on the substrate specificity constants, the extension fidelity was calculated to be 2.9 x 10-3 for dTTP or higher for dATP and dGTP (Table 2.5).
Exonuclease cleavage of double-stranded DNA substrates containing matched and mismatched 3′-terminal nucleotides at 37 oC The 3′→5′ exonuclease activity of a replicative DNA polymerase is known to proofread polymerization products and excise misincorporated nucleotides. To examine if the 3′→5′
27 exonuclease activity of PolB1 has similar function, we measured the excision rate (kexo) of the wild-type PolB1 with both D-1 and M-1 (Table 2.1) under single-turnover conditions. The wild-type PolB1 (400 nM) and 5′-[32P]-labeled D-1 or M-1 (100 nM) in buffer A (see 2.2) were pre-equilibrated before adding Mg2+ (15 mM) to initiate the exonuclease reaction. The molar amount of remaining substrate was plotted as a function
of time (Figure 2.6) and the data were fit to equation 4 (see 2.2) to yield kexo and reaction -1 amplitudes (A). The kexo and A values are 0.44 ± 0.02 s and 91 nM for matched D-1 DNA and 1.86 ± 0.08 s-1 and 94 nM with mismatched M-1 DNA. Thus, at the primer- template junction, a mismatched base pair dA:dA was excised with a 3-fold faster rate than a matched base pair, dA:dT.
2.4 Discussion
Pre-steady state kinetic methods have been used to determine the fidelity of several DNA polymerases and to establish a kinetic basis for the fidelity of each of them [47, 60, 89- 92, 105, 111, 113-117]. By employing these methods, we first measured the base substitution fidelity of the exonuclease-deficient PolB1 from S. solfataricus P2 at 37 °C. The kinetic parameters for all sixteen possible single nucleotide incorporations under single-turnover reaction conditions were individually determined. The base substitution -4 -6 fidelity (Fpol) of PolB1 exo- was determined to be in the range of 10 to 10 (Table 2.2). Additionally, PolB1 incorporated four correct incorporations with 6-60-fold higher substrate specificity than Dpo4 [105], a DNA lesion bypass Y-family polymerase from the same archaeon S. solfataricus P2. Both the high substrate specificity for correct incorporations and the low incorporation efficiency for misincorporations contributed to - the Fpol of PolB1 exo-. Because the average kp/Kd for four correct incorporations (1.3 μM 1 -1 -4 s ) was 6,800-fold higher than the average kp/Kd for twelve misincorporations (1.9 x 10 -1 -1 μM s ), the calculation formula of Fpol can be rearranged as shown below:
Fpol = (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect] ≈ -1 -1 (kp/Kd)incorrect/(kp/Kd)correct = [(Kd)incorrect/(Kd)correct] [(kp)correct/(kp)incorrect] =
28 (affinity difference)-1(rate difference)-1 (5)
In equation 5, Fpol is approximately equal to the ratio of substrate specificities of correct and incorrect nucleotides and is inversely proportional to both the apparent affinity
difference (defined as (Kd)incorrect/(Kd)correct) and to the rate difference (defined as
(kp)correct/(kp)incorrect). For correct incorporations, the Kd values were in the range of 4-11 -1 μM while the kp values were in the range of 5-12 s (Table 2.2). In comparison, the measurable apparent Kd values were in the range of 900-4500 µM while the measurable -1 kp values were in the range of 0.009-1.7 s for incorrect incorporations (Table 2.2). For each D-DNA substrate (Table 2.1), we then calculated the apparent affinity difference and the rate difference which respectively provided 109-918-fold and 4-589-fold
contributions to the Fpol of PolB1 exo- (Table 2.6). Interestingly, the apparent affinity difference was much larger than that observed with Dpo4 [105] but similar to those observed with the exonuclease-deficient T7 DNA polymerase (T7 DNA Pol) [89], human mitochondrial DNA polymerase γ (hPolγ) [91], rat DNA polymerase β (rPolβ) [114], and human immunodeficiency virus type 1 reverse transcriptase (HIV-1 RT) [113] (Table 2.6). The selection provided by nucleotide ground-state binding affinity reflects the tightness of a replicative polymerase’s active site and the strength of the interactions between active site residues and an incoming nucleotide [105]. For example, Dpo4, a low-fidelity Y-family enzyme which binds both correct and incorrect nucleotides weakly [105], possesses a loose active site which interacts minimally with an incoming nucleotide [101]. Conversely, more intimate contacts between the enzyme and dNTP have been observed in the ternary crystal structures (E•DNA•dNTP) of high-fidelity DNA polymerases [118-122] and HIV-1 RT [123], which possesses tighter active sites than the Y-family DNA polymerases. A tight polymerase active site can select a correct against incorrect nascent base pair by shape, leading to high base selectivity [65]. Therefore, we hypothesize that the active site of PolB1 is relatively tight and interacts extensively with an incoming nucleotide. Thermodynamically, this hypothesis is reasonable since the average apparent affinity difference (374) with PolB1 corresponds to a ΔΔG (=RTln[(Kd)incorrect/(Kd)correct]) of 3.8 kcal/mol. This value is larger than the free energy differences (0.3-1.0 kcal/mol at 37 °C) between correct and incorrect base pairs at
29 the primer terminus measured through DNA melting experiments [124]. Thus, the differential interactions between correct and incorrect nucleotides with the polymerase active site residues of PolB1 provided a free energy difference of 2.6-3.3 kcal/mol for nucleotide selection. To further evaluate our above hypothesis, we are currently attempting to solve the ternary crystal structure of PolB1 (PolB1•DNA•dNTP). Interestingly, Table 2.6 also shows that the rate difference, although varying in a large
range, does contribute to the Fpol of all five DNA polymerases from four different families. This suggests that the nucleotide incorporation step provides a common and important fidelity checkpoint during polymerization [54]. Thus, the polymerase activity of PolB1 uses both the nucleotide binding step and the nucleotide incorporation step to discriminate against incorrect nucleotides (Table 2.6).
Cost of proofreading During proofreading, the 3′ → 5′ exonuclease activity selectively excises mismatched bases on a primer’s 3′-terminus. Although rare, this activity can also cleave matched bases and slow down DNA synthesis. The “cost” of utilizing this editing function has
been previously defined as the ratio of the excision rate of matched DNA (kexo) and the
incorporation rate of the next correct nucleotide (kp) [84]. PolB1 excised matched D-1 (Table 2.1) at a rate of 0.44 s-1 (Figure 2.6) while its polymerase domain incorporated correct dTTP into D-1 at a rate of 8.2 s-1 (Table 2.2). Thus, the “cost” of proofreading at 37 °C is calculated to be 5.4% which is 38-fold higher than 0.14% observed with the human mitochondrial DNA polymerase complex [84]. Such costly proofreading activity could be caused by either non-physiological reaction conditions (e.g. temperature, pH) or the absence of a potential accessory protein to PolB1 (see below). These possibilities are being investigated in our laboratory.
Contribution of the 3′ → 5′ exonuclease activity to the fidelity of PolB1 at 37 oC When the polymerase activity of PolB1 incorporates an incorrect nucleotide under single- turnover conditions, one of the following competing pathways will occur if the DNA substrate does not dissociate from PolB1: the misincorporated nucleotide will be excised
30 with a rate of kexo by the 3′ → 5′ exonuclease activity or polymerization will proceed with a rate of kp to bury the misincorporated nucleotide by a correct nucleotide. Based on the kinetic partitioning between these two pathways, we define the mismatch removal
probability as the ratio of kexo/(kexo + kp) and the mismatch extension probability as the
ratio of kp/(kexo + kp). If the first pathway dominates, then the mismatch removal probability and the mismatch extension probability will approach 100% and 0%, respectively. Therefore, the 3′ → 5′ exonuclease activity will significantly increase the
polymerization fidelity of PolB1. Thus, we further define the component (Fexo) contributed by the 3′ → 5′ exonuclease to the overall fidelity of PolB1 as
Fexo = mismatch removal probability/mismatch extension probability = kexo/kp For M-1 (Table 2.1), the single mismatched dA:dA base pair at the junction of the primer -1 and template was excised with a kexo of 1.86 s (Figure 2.6) while it was extended by a -1 correct dCTP with a kp of 0.13 s (Table 2.5), resulting in a removal probability of 93%,
a mismatch extension probability of 7%, and an Fexo of 14. Because the mismatch extension fidelity (Table 2.5) was high, we did not consider the possibility of -4 misincorporations during mismatch extension. Considering that Fpol is in the range of 10 -6 to 10 (Table 2.2), the overall fidelity of PolB1, which is the ratio of Fpol and Fexo, was calculated to be in the range of 10-5 to 10-7. In addition to reaction conditions (pH and temperature), this fidelity can be affected by the following factors: (i) faster intramolecular transfer of mismatched than matched DNA from the polymerase to the exonuclease active site; (ii) mismatched DNA dissociates faster than matched DNA from the polymerase active site; and (iii) mismatched DNA binds faster than matched DNA to the 3′→5′ exonuclease active site. These factors have been proven to contribute to the fidelity of T7 DNA Pol [47, 89] and hPolγ [84] in which the 3′→5′ exonuclease activity enhances their overall fidelity by ~200-fold. We are currently investigating the potential contributions of these factors to the overall fidelity of PolB1. Moreover, the enzymatic properties, including fidelity of several replicative DNA polymerases, have been found to be altered by their accessory proteins. For example, the fidelity of hPolγ is increased by 14-fold with its accessory subunit [90]. With regard to PolB1, a similar accessory protein
31 has not been identified, although, the genome of S. solfataricus P2 encodes a heterotrimer of proliferating cell nuclear antigen.
Insignificant effect of temperature on the base substitution fidelity of the polymerase activity of PolB1 When reaction temperature increased from 19 to 50 oC, Table 2.3 shows that the base substitution fidelity of PolB1 exo- only decreased by 5-fold. A similar modest increase in base substitution rates with temperature has been observed with S. solfataricus Dpo4 [60, 104], the exonuclease-deficient Thermus aquaticus (Taq) DNA polymerase [94], VentTM DNA polymerase [95], and T4 DNA polymerase [93, 96]. Therefore, we project the base substitution fidelity of PolB1 exo- in vivo (80 oC) will be slightly higher than the fidelity value calculated at 50 oC (Table 2.3). As in our previous kinetic analysis of the temperature effect on the fidelity of Dpo4 [60], the kinetic basis for the small decrease in the base substitution fidelity of PolB1 exo- was due to a similar change in magnitude of
the kp/Kd with temperature for both correct and incorrect dNTP incorporations (Table 2.3). Also similar to Dpo4 [60], the apparent ground-state binding affinities of both
correct and incorrect nucleotides and their ratio Kd, incorrect/Kd, correct were altered slightly over a temperature range of 31 oC. In stark contrast, PolB1 exo- catalyzed the o incorporations of both correct and incorrect nucleotides with a much faster kp at 50 C than at 19 oC. This indicated that PolB1, like Dpo4 [60], becomes more dynamic at higher temperatures. The contribution of increased protein dynamics of PolB1 to catalysis is discussed in detail in Chapter 3.
Higher polymerization fidelity of PolB1 at lower pH Using M13mp2-based fidelity assays, the base substitution fidelity of several DNA polymerases, including the exonuclease-deficient Taq DNA polymerase [125] and E. coli DNA polymerase I [126], has been found to be about 10-60-fold higher when the reaction pH is lowered by three units from 8–9 to 5–6 [125-127]. Consistently, the base substitution fidelity of PolB1 exo- increased by 36-fold when reaction pH was decreased from 8.5 to 6.0 (Table 2.4). Thus, both kinetic and M13mp2-based fidelity assays indicate
32 that these DNA polymerases are less faithful at higher pH. Mechanistically, the 36-fold drop in the polymerase fidelity of PolB1 exo- when reaction pH was raised from 6.0 to 8.5 was contributed by both a 10-fold lower incorporation efficiency of correct dTTP and
a 3.6-fold higher kp/Kd for incorrect dATP incorporation (Table 2.4). Moreover, these changes in the incorporation efficiency were due mostly to a 7-fold increase in the
apparent Kd for correct dTTP incorporation and a 10-fold increase in the kp for dATP misincorporation. Interestingly, the cytoplasmic pH of another Sulfolobus species, S. acidocaldarius, has been estimated to be about 6.0 [128], although, this organism grows optimally under harsh environmental conditions (80 °C and pH 2-4) similar to S. solfataricus [73]. Based on the physiological similarity between these two Sulfolobus species, we hypothesize that the cytoplasmic pH of S. solfataricus, which has not been measured, is approximately 6.0. At this pH, the polymerase activity of PolB1 will have a 25-fold higher base substitution fidelity than the fidelity at pH 7.5 (Table 2.2). After offsetting the opposing effects of temperature and pH on the Fpol, we further estimate the overall fidelity of PolB1 to be in the range of 10-6 to 10-8 in vivo.
Biological relevance of the kinetically estimated fidelity of PolB1 Although inhabiting environments at extremely high temperature and low pH, hyperthermophilic archaea have much lower genetic mutation rates than other DNA microbes and eukaryotes [129]. For example, the base substitution frequency of S. acidocaldarius when growing at pH 3.5 and 75 °C is 3.2 × 10-10 per base [129]. Although such a frequency for S. solfataricus has not been reported, we estimated it to be either 3.2 × 10-10 or 100-fold higher based on the following in vivo observation: the average frequency of spontaneous mutations in S. solfataricus, which are dominated by transposon mutagenesis, is at least 50-60-fold higher than the one in S. acidocaldarius [130]. Taken together, we estimate the base substitution frequency in S. solfataricus to be in the range of 10-8 to 10-10 per base. This frequency is about 100-fold lower than the estimated error rate (10-6 to 10-8) of PolB1 in vitro (see above discussion). Although the effects of several kinetic factors and a potential accessory protein may further decrease the error rate of PolB1, other cellular proteins or pathways clearly play a major role in
33 lowering the substitution frequency in S. solfataricus. For example, active DNA repair pathways have been proposed to maintain the overall genetic fidelity in S. acidocaldarius [129]. Similar pathways in S. solfataricus likely correct most misincorporations generated by PolB1 or other polymerases in vivo.
34 2.5 Tables
Table 2.1 Sequences of synthetic double-stranded DNA substratesa D-1 5′-CGCAGCCGTCCAACCAACTCA-3′ 3′-GCGTCGGCAGGTTGGTTGAGTAGCAGCTAGGTTACGGCAGG-5′ D-6 5′-CGCAGCCGTCCAACCAACTCA-3′ 3′-GCGTCGGCAGGTTGGTTGAGTGGCAGCTAGGTTACGGCAGG-5′ D-7 5′-CGCAGCCGTCCAACCAACTCA-3′ 3′-GCGTCGGCAGGTTGGTTGAGTTGCAGCTAGGTTACGGCAGG-5′ D-8 5′-CGCAGCCGTCCAACCAACTCA-3′ 3′-GCGTCGGCAGGTTGGTTGAGTCGCAGCTAGGTTACGGCAGG-5′ M-1 5′-CGCAGCCGTCCAACCAACTCAA-3′ 3′-GCGTCGGCAGGTTGGTTGAGTAGCAGCTAGGTTACGGCAGG-5′ aImportant changes in the DNA sequence are highlighted in bold.
35
Table 2.2 Kinetic parameters of nucleotide incorporation into DNA catalyzed by PolB1 exo- at 37 °C.
k K k /K dNTP p d p d F a (s-1) (μM) (μM-1s-1) pol Template dA (D-1) dTTP 8.2 ± 0.6 11 ± 2 7.5 × 10-1 dATP 1.3 ± 0.1 (2.8 ± 0.3) × 103 4.6 × 10-4 6.2 × 10-4 dCTP - - 1.9 × 10-4 2.5 × 10-4 dGTP 0.05 ± 0.01 (1.2 ± 0.2) × 103 4.2 × 10-5 5.6 × 10-5 Template dT (D-7) dATP 11.5 ± 0.2 4.9 ± 0.3 2.3 dCTP 0.7 ± 0.2 (3.3 ± 1.5) × 103 2.1 × 10-4 9.0 × 10-5 dGTP 0.9 ± 0.1 (3.2 ± 0.8) × 103 2.8 × 10-4 1.2 × 10-4 dTTP 1.7 ± 0.3 (4.5 ± 1.0) × 103 3.8 × 10-4 1.6 × 10-4 Template dG (D-6) dCTP 5.7 ± 0.1 8.2 ± 0.5 7.0 × 10-1 dATP - - 7.5 × 10-6 1.1 × 10-5 0.032 ± -5 -5 dGTP (0.9 ± 0.1) × 103 3.3 × 10 4.8 × 10 0.002 dTTP 1.3 ± 0.2 (3.5 ± 0.7) × 103 3.7 × 10-4 5.3 × 10-4 Template dC (D-8) dGTP 5.3 ± 0.1 4.1 ± 0.3 1.3 dATP 0.34 ± 0.05 (2.3 ± 0.6) × 103 1.3 × 10-4 1.0 × 10-4 dCTP 0.009 ± 0.002 (2.0 ± 0.9) × 103 4.5 × 10-6 3.5 × 10-6 -4 -4 dTTP - - 1.6 × 10 1.2 × 10 a Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
36
Table 2.3 Temperature dependence of the nucleotide incorporation fidelity catalyzed by PolB1 exo-.
kp Kd kp/Kd a dNTP -1 -1 -1 Fpol (s ) (μM) (μM s ) 19 oC dTTP 0.14 ± 0.01 38 ± 6 3.7 × 10-3 dATP 0.0033 ± 0.0008 (4.0 ± 1.3) × 103 8.3 × 10-7 2.2 × 10-4 26 oC dTTP 1.6 ± 0.01 18 ± 0.4 8.9 × 10-2 dATP 0.018 ± 0.008 (2.5 ± 1.5) × 103 7.2 × 10-5 8.1 × 10-4 32 oC dTTP 2.6 ± 0.1 6.9 ± 0.5 3.8 × 10-1 dATP 0.19 ± 0.06 (2.3 ± 0.8) × 103 8.3 × 10-5 2.2 × 10-4 37 oC dTTP 8.2 ± 0.6 11 ± 2 7.5 × 10-1 dATP 1.3 ± 0.1 (2.8 ± 0.3) × 103 4.6 × 10-4 6.1 × 10-4 44 oC dTTP 38 ± 1 13 ± 1 2.9 dATP 5.4 ± 1.3 (2.5 ± 0.8) × 103 2.2 × 10-3 7.6 × 10-4 50 oC dTTP 92 ± 2 20 ± 1 4.6 -3 dATP 22 ± 5 (4.2 ± 1.5) × 103 5.2 × 10-3 1.1 × 10 a Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
37
Table 2.4 pH dependence of the nucleotide incorporation fidelity of PolB1 exo- at 37 °C. k K k /K dNTP p d p d F a (s-1) (μM) (μM-1s-1) pol 6.0 dTTP 8.9 ± 0.3 3.2 ± 0.4 2.8 dATP 0.19 ± 0.01 (2.9 ± 0.2) × 103 6.6 × 10-5 2.4 × 10-5 6.5 dTTP 8.5 ± 0.2 4.5 ± 0.3 1.9 dATP 0.32 ± 0.02 (2.2 ± 0.2) × 103 1.5 × 10-4 7.9 × 10-5 7.5 dTTP 8.2 ± 0.6 11 ± 2 7.5 × 10-1 dATP 1.3 ± 0.1 (2.8 ± 0.3) × 103 4.6 × 10-4 6.1 × 10-4 8.5 dTTP 5.8 ± 0.1 21 ± 0.4 2.8 × 10-1 dATP 2.0 ± 0.5 (8.4 ± 2.3) × 103 2.4 × 10-4 8.6 × 10-4 a Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
38
Table 2.5 Fidelity of the extension of a mismatched dA:dA terminus catalyzed by PolB1 exo- at 37 °C.
kp Kd kp/Kd a dNTP -1 -1 -1 Fext (s ) (μM) (μM s ) dCTP 0.13 ± 0.01 (1.0 ± 0.2) × 103 1.3 × 10-4 dATP No observed incorporation dGTP No observed incorporation dTTP - - 3.8 × 10-7 2.9 × 10-3 a Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
39 Table 2.6 Comparison of the base substitution fidelity of PolB1 exo- and four other DNA polymerases and HIV-1 reverse transcriptase.
a Polymerase Fpol Affinity difference Rate difference Polymerase family [(Kd)incorrect/(Kd)correct] [(kp)correct/(kp)incorrect]
PolB1 exo-b B 3.5 × 10-6 to 1.2 × 10-4 109 to 918 4 to 589
Dpo4c Y 1.5 × 10-4 to 3.2 × 10-3 1 to 18 2.4 × 102 to 1.7 × 103
rPolβd X 1.1 × 10-5 to 5.9 × 10-4 35 to 342 28 to 708
hPolγe A 4.6 × 10-7 to 2.9 × 10-4 42 to 900 39 to 1.2 × 104
T7 DNA Polf A 2.6 × 10-7 to 6.7 × 10-6 200 to 400 2.0 × 103 to 4.0 × 103
HIV-1 RTg RT 5.9 × 10-5 to 5.8 × 10-4 210 to 310 7 to 80 a b o c o d Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect]. At 37 C (this work). At 37 C [105]. At 37 oC [114]. eAt 37 oC, excluding the fidelity contribution from the 3′→5′ exonuclease activity [91]. fAt 20 oC, excluding the fidelity contribution from the 3′→5′ exonuclease activity [89]. gAt 37 oC and with a DNA substrate [113].
40 2.6 Figures
A
B
Figure 2.1 Comparison of polymerase and 3′→5′ exonuclease activity for PolB1 enzymes. The enzymatic activities of (A) wild-type PolB1 and (B) PolB1 exo- were examined by pre-incubating the appropriate enzyme (120 nM) with D-1 DNA. The polymerase reaction was initiated with 50 µM dTTP•Mg2+ (correct) or 50 µM dCTP•Mg2+ (incorrect) in independent reactions while the exonuclease reaction was initiated with 5 mM Mg2+. All reactions were quenched using 0.37 M EDTA at the designated time. Strong exonuclease activity was observed at all time points for wild-type PolB1 except for the 6 s time point in the presence of correct dTTP. In sharp contrast, no degradation was observed for the exonuclease-deficient PolB1 mutant under all reaction conditions tested here. 41 A B 10 30 8 30
7 25 25 Amplitude (nM,
8 Amplitude (nM, 6 20 20 ) ) 5 z z 6 , , -1 -1 15 4 15 (s
(s 4 obs obs 3 k k 10 10 )
) 2 2 5 5 1
0 0 0 0 0 10203040506070 0 50 100 150 200 250 2+ Mg (mM) NaCl (mM)
C 12 30
10 25 Amplitude (nM,
8 20 ) z , -1 6 15 (s obs
k 4 10 ) 2 5
0 0 6 6.5 7 7.5 8 8.5 9 9.5 10 pH
Figure 2.2 Effect of Mg2+ concentration, NaCl concentration, and pH on the enzymatic activity of PolB1 exo-. A pre-incubated solution of D-1 5'-labeled with 32P (30 nM) and a 3-fold excess of PolB1 exo- (120 nM) was rapidly mixed with the correct nucleotide (100 µM dTTP) for various time intervals under single-turnover conditions. Each plot shows 2+ the kobs values (●) and reaction amplitudes (■) as (A) Mg concentration, (B) NaCl concentration, or (C) buffer pH was varied. Activity in panel C was assayed in 25 mM MES-NaOH buffer between pH 6.0 and 7.0, in 25 mM Tris-HCl buffer for pH 8.0 and 8.5, and in 25 mM glycine-NaOH buffer for pH 9.0 and 10.0.
42 A 30
25
20
15
10 Product (nM)
5
0 0 1 2 3 4 5 6 Time (s)
B 6
5
) 4 -1
(s 3 obs k 2
1
0 0 20 40 60 80 100 dCTP (μM)
Figure 2.3 Concentration dependence on the pre-steady-state rate of correct and incorrect nucleotide incorporation catalyzed by PolB1 exo-. (A) A pre-incubated solution of PolB1 exo- (120 nM) and 5'-[32P]-labeled-D-6 (30 nM) was mixed with various concentrations of dCTP•Mg2+ (1 µM, ●; 2.5 µM, ○; 5 µM, ■; 10 µM, □; 20 µM, ▲; 45 µM, Δ; 90 µM, ♦) for various time intervals. The solid lines are the best fits to the single-exponential equation. (B) The single-turnover rates (kobs) were plotted as a function of dCTP concentration. These data were then fit to a hyperbolic equation to yield a kp of 5.7 ± 0.1 –1 s and a Kd of 8.2 ± 0.5 µM.
43 0.01
0.001
0.0001
Fidelity 10-5
10-6
10-7 15 20 25 30 35 40 45 50 55 Temperature (oC)
Figure 2.4 Temperature dependence of the nucleotide incorporation fidelity of PolB1 exo-.
44 0.01
0.001
0.0001 Fidelity
10-5
10-6 5.5 6 6.5 7 7.5 8 8.5 9 pH
Figure 2.5 pH dependence of the nucleotide incorporation fidelity of PolB1 exo-.
45 100
80
60
40
20 Remaining Substrate (nM) Substrate Remaining
0 0 2 4 6 8 10 12 14 16
Time (s)
Figure 2.6 Rate of 3′→5′ exonuclease activity catalyzed by PolB1. A pre-incubated solution of PolB1 (400 nM) and DNA (100 nM) was rapidly mixed with Mg2+ (15 mM) for various reaction times. Remaining substrate was plotted as a function of time. The solid lines represent the best fit to a single-exponential decay. PolB1 catalyzed the excision of a nucleotide for a matched (D-1 DNA, ●) or mismatched (M-1 DNA, ■) DNA terminus at a rate of 0.44 ± 0.02 s-1 and 1.86 ± 0.08 s-1, respectively.
46 Chapter 3: Kinetic Mechanism of DNA Polymerization Catalyzed by Sulfolobus solfataricus P2 DNA Polymerase B1 at 37 °C
3.1 Introduction
Elucidating the reaction pathway is an important goal towards understanding the catalytic events performed by an enzyme. To achieve this objective, a reaction pathway is established by identifying the chemical and conformational intermediates of each step and by measuring the rate and equilibrium constants for the elementary steps of converting substrate(s) into product(s). This knowledge subsequently provides the foundation for defining enzymatic specificity and the rate-limiting step. In pursuit of this goal, we have employed transient state kinetic techniques to define the minimal kinetic mechanism of DNA polymerization catalyzed by an exonuclease-deficient mutant of Sulfolobus solfataricus P2 DNA polymerase B1 (PolB1 exo-).
PolB1, a B-family DNA polymerase, is one of four DNA polymerases encoded within the S. solfataricus P2 genome: three are B-family polymerases and one is a DNA lesion bypass Y-family polymerase, i.e. DNA polymerase IV (Dpo4) [73]. Thus far, biochemical characterization of the isolated gene products has deemed only two of these polymerases functional in vitro: PolB1 and Dpo4 [100, 108, 131]. Furthermore, a direct physical interaction between PolB1 and Dpo4 has been demonstrated [131], and surprisingly, protein levels are similar for both polymerases during normal growth conditions [132].
Although a myriad of DNA polymerases have been biochemically characterized, detailed kinetic mechanisms of their 5′ → 3′ polymerase activity have been established for only a 47 handful of DNA polymerases. Some of the more prominent examples include Escherichia coli DNA polymerase I [61, 85, 87, 133-135], bacteriophage T7 DNA polymerase (T7 DNA Pol) [88, 89, 112], human immunodeficiency virus type I reverse transcriptase (HIV-1 RT) [113, 136, 137], bacteriophage T4 DNA polymerase (T4 DNA Pol) [86, 135, 138-141], Saccharomyces cerevisiae DNA polymerase η [115], and S. solfataricus P2 Dpo4 [60, 105, 111]. Of these and others, T4 DNA polymerase is the only B-family member to have undergone rigorous kinetic studies. Herein, we report the first kinetic mechanism of an archaeal B-family polymerase. S. solfataricus P2 PolB1 was selected for several reasons: (i) the crystal structure has been solved for S. solfataricus MT4 PolB1 in which the amino acid sequence differs from strain P2 by only two residues [109], (ii) it is an ideal model system for DNA replication because the replication machinery is a simplified version of that in eukaryotes (see [142] for a review), (iii) the kinetic mechanism of Dpo4 has been studied in detail which could facilitate future studies on the mechanism of polymerase switching between PolB1 and Dpo4 [60, 111], (iv) biochemical data are available for interactions among PolB1, Dpo4, and accessory proteins [131, 143-145], and (v) in Chapter 2, we showed that the polymerase active site confers a fidelity of 10-4 to 10-6 while the exonuclease active site increases the fidelity by 14-fold for an overall fidelity of 10-6 to 10-8 at pH 6.0 . Using an exonuclease-deficient mutant of PolB1, we have dissected the minimal kinetic pathway for DNA polymerization by measuring the elementary steps in a single catalytic cycle. Surprisingly, several of the reported parameters are similar to Dpo4, and we provide kinetic evidence that PolB1, like Dpo4, utilizes an induced-fit mechanism during correct nucleotide incorporation.
3.2 Materials and Methods
Materials 32 These chemicals were purchased from the following companies: [α- P]dTTP, MP 32 Biomedicals (Solon, OH); [γ- P]ATP, PerkinElmer (Boston, MA); Biospin columns,
48 Bio-Rad Laboratories (Herclues, CA); dNTPs, GE Healthcare (Piscataway, NJ); OptiKinase, USB (Cleveland, OH); Sp-dTTPαS and Sp-dGTPαS, Biolog – Life Science Institute (Bremen, Germany); ddTTP, Trilink Biotechnologies (San Diego, CA).
Pre-steady state kinetic assays
S. solfataricus PolB1 exo- (D231A/E233A/D318A) fused to a C-terminal His6 tag was overexpressed in E. coli and purified as described previously [146]. The D-1 DNA substrate shown in Figure 3.1 was prepared as described previously [105, 146]. All experiments using PolB1 exo- were performed at 37 °C, if not specified, and in optimized reaction buffer A which contained 50 mM HEPES (pH 7.5), 15 mM MgCl2, 75 mM NaCl, 5 mM DTT, 10% glycerol, and 0.1 mg/ml BSA [146]. Please note, although the PolB1 enzyme utilized in these studies is an exonuclease-deficient mutant, PolB1 exo- was never pre-incubated in the presence of magnesium. As in the case of HIV-1 RT, the kinetic results may be slightly different when the enzyme is incubated with or without magnesium [136]. Fast reactions were carried out at 37 °C using a rapid chemical-quench flow apparatus (KinTek, PA). All reported concentrations are the final reaction values.
Active site titration assay DNA The equilibrium dissociation constant (Kd ) of the binary complex was determined by mixing a pre-incubated solution of PolB1 exo- (45 nM) and increasing concentrations of 5′-radiolabeled D-1 DNA (2.5 - 100 nM) with dTTP (100 µM). All reactions were
quenched at 1.8 s (equivalent to 8 half times, t1/2) in order to achieve the maximal first- turnover amplitude.
Measurement of the phosphorothioate elemental effect A pre-incubated solution of PolB1 exo- (120 nM) and 5′-radiolabeled 21/41-mer D-1 (30 nM) was rapidly mixed with either dTTP (100 µM) or Sp-dTTPαS (100 µM, >95% purity) in buffer A and terminated with 0.37 M EDTA. For incorporation of an incorrect nucleotide, the reaction was initiated with either dGTP (500 µM) or Sp-dGTPαS (500
49 µM, >95% purity). The Sp isomers were used, opposed to the Rp isomers, due to the stereoselectivity observed previously [147, 148].
Pulse-chase and pulse-quench experiments Pulse-chase and pulse-quench experiments were performed in buffer A using a rapid chemical-quench flow instrument. A pre-incubated solution containing PolB1 exo- (100 nM) and unlabeled D-1 DNA substrate (100 nM) was loaded into one sample loop and 32 rapidly mixed with buffer containing [α- P]dTTP (50 μM) from a second sample loop for reaction times ranging from 22 ms to 2.0 s. In the pulse-quench experiment, reactions were immediately quenched with 1 M HCl. In the pulse-chase experiment, reactions were immediately chased with 1.0 mM unlabeled dTTP for 30 s, followed by quenching with 1 M HCl. In both cases, quenched reactions were treated with chloroform and neutralized with 1 M NaOH.
Product analysis Reaction products were analyzed by sequencing gel electrophoresis (17% acrylamide, 8 M urea, 1× TBE running buffer) and quantitated using a Typhoon TRIO (GE Healthcare) and ImageQuant software (Molecular Dynamics). For the pulse-chase and pulse-quench assays, DNA products were resolved using a 20% highly crosslinked polyacrylamide gel matrix as described previously [149].
Data analysis Data were fit by nonlinear regression using KaleidaGraph (Synergy Software). Data from the burst and pulse-chase experiments were fit to equation 1
[Product] = A[1 – exp(–k1t) + k2t] (1)
where A is the fraction of active enzyme, k1 is the observed burst rate, and k2 is the observed steady-state rate. Data from the experiments performed under steady-state conditions were fit to equation 2
[Product] = kssE0t + E0 (2)
50 where kss is the steady-state rate constant of dNTP incorporation at the initial active enzyme concentration of E0. Data from the assay measuring the DNA dissociation rate of the binary complex were fit to equation 3
[Product] = A[exp(–k1t)] + C (3) 32 where A is the reaction amplitude, k1 is the observed rate constant, and C is the [ P]- labeled product concentration in the presence of a DNA trap for unlimited time. For the active site titration assay, resulting product concentration (equivalent to burst amplitudes) was plotted versus the concentration of D-1 DNA and fit via quadratic regression to equation 4 DNA DNA 2 1/2 [E·DNA] = 0.5(Kd + E0 + D0) – 0.5[(Kd + E0 + D0) – 4E0D0] (4) DNA where Kd represents the equilibrium dissociation constant for the binary complex
(E•DNA), E0 is the active enzyme concentration, and D0 is the DNA concentration. Data from the experiments performed under single-turnover conditions were fit to equation 5
[Product] = A[1 – exp(–kobst)] (5)
where A is the reaction amplitude and kobs is the observed single-turnover rate. When the
assumption of k1 >> k2 is not valid for burst experimental data (refer to equation 1), equation 6 was applied. 2 [Product] = A{(k1k2/k1 + k2)t + (k1/k1 + k2) [1 – exp(–k1 + k2)t]} (6)
Using kp and temperature data from the previous chapter (Table 2.3), the activation energy was determined using the Arrhenius equation
[kp] = Ar[exp(–Ea/RT)] (7)
where kp is the maximum rate of nucleotide incorporation, Ar is the proportionality
constant, Ea is the activation energy, R is the universal gas constant, and T is the reaction temperature in Kelvin.
Data from the processive elongation of 21/41-mer to 27/41-mer were modeled using an improved personal computer version of Kinsim provided by Carl Frieden, Washington University (St. Louis, MO) [150]. Final fitting of the data was accomplished by nonlinear regression using an improved personal computer version of Fitsim [151].
51 3.3 Results
Biphasic kinetics observed for PolB1 exo- To begin characterizing S. solfataricus P2 PolB1 exo-, we first performed a burst experiment at 37 °C. The rationalization for studying PolB1 exo- at a sub-optimal temperature of 37 °C was explained in the Chapter (Section 2.3). Results from a burst assay can yield insight into (i) the rate of nucleotide incorporation occurring at the enzyme’s active site during the first turnover and (ii) whether product release from the enzyme during subsequent turnovers is rate-limiting. A pre-incubated solution of PolB1 exo- (30 nM based on UV absorbance measurements) and radiolabeled D-1 DNA (120 nM) was rapidly mixed with dTTP (100 µM) and quenched with EDTA at various reaction times. The time course of product formation indicated that PolB1 exo- followed biphasic or “burst” kinetics (Figure 3.1 part (A)). Applying equation 1 (Section 3.2) to the plot resolved an observed burst rate of 9 ± 1 s-1 (exponential phase) and an observed linear-phase rate of 0.074 ± 0.05 s-1. As observed with Dpo4 [111], the burst phase was likely limited by nucleotide incorporation in the first turnover while the linear phase could be the steady-state phase of product formation during subsequent enzyme turnovers (Scheme 3.1). To evaluate this possibility, we performed the following steady-state kinetic assay: PolB1 exo- (1 nM) was pre-equilibrated with a large excess of substrate (i.e. 250 nM of radiolabeled D-1 DNA) prior to initiating the reaction with dTTP (100 µM). A plot of product concentration versus time (Figure 3.1 part (B)) was linear and was fit to equation 2 (Section 3.2) which yielded a steady-state rate of 0.038 s-1. This value was within two-fold of 0.074 ± 0.05 s-1 determined in Figure 3.1 part (A), thereby confirming that the linear phase under burst conditions was indeed a steady-state phase. In this phase, production formation is usually limited by the slow dissociation of the binary complex (E•DNA). To confirm this kinetic trend, we directly measured the DNA dissociation rate. PolB1 exo- (50 nM) was pre-incubated with 5′-radiolabeled D-1 DNA (100 nM) and mixed with an excess of unlabeled D-1 DNA (2.5 µM) from 10 sec to 20 min. Then, dTTP (133 µM) was added for 15 sec so that any DNA substrate in complex with PolB1 exo- could be extended. The concentration of the radioactive DNA product
52 was plotted versus time and fit to a single-exponential equation (Equation 3 in Section -1 3.2) (Figure 3.1 part (C)). The measured rate of 0.043 ± 0.003 s (k-1 in Scheme 3.1) was similar to 0.038 and 0.074 s-1 determined in Figure 3.1 parts (A) and (B), substantiating the observation that multiple enzyme turnovers were limited by the dissociation of E•DNA.
Active site titration Since PolB1 exo- displayed “burst” kinetics, this implied a stable binary complex facilitated the rapid incorporation of dTTP so that the equilibrium dissociation constant DNA (Kd ) of the binary complex can be measured by titrating the active site of PolB1 exo- with varying concentrations of DNA. This active site titration assay was conducted by pre-incubating PolB1 exo- (45 nM based on UV absorbance measurements) with increasing concentrations of 5′-radiolabeled D-1 DNA (2.5-100 nM), starting the reaction by adding dTTP (100 µM), and quenching all reactions at 1.8 s with EDTA. For each DNA concentration, the reaction was performed in triplicate. The average product concentration was plotted for each concentration of DNA (Figure 3.2) and fit to a DNA quadratic equation (Equation 4 in Section 3.2). A Kd of 1.8 ± 0.4 nM and an E0 of
18.9 ± 0.4 nM were extracted. This E0 equates to 42% enzyme activity which is similar to that determined for HIV-1 RT [113]. Henceforth, all experiments described below were performed using the active site enzyme concentration. Lastly, the apparent second-order
binding rate constant (k1 in Scheme 3.1) of the binary complex can be calculated by DNA substituting the measured values for k-1 and Kd into the following equation: k1 = k- DNA -1 -9 7 -1 -1 -1 -1 1/Kd = (0.043 s )/(1.8 × 10 M) = 2.4 × 10 M s = 24 μM s .
Rates of association and dissociation of an incoming nucleotide In the Chapter 2 (Table 2.2), we measured the apparent equilibrium dissociation constant dNTP -1 (Kd = 11 ± 2 µM) and maximum rate (kp = 8.2 ± 0.6 s ) of dTTP incorporation into D-1 DNA catalyzed by PolB1 exo- under single-turnover conditions. Assuming the association rate (k2 in Scheme 3.1) of dTTP, a small molecule, approaches the diffusion limit of 1.0 × 108 M-1s-1, we estimated the upper limit of the dTTP dissociation rate to be
53 -1 dNTP 1,100 s (k-2 = k2 × Kd ) which is too fast to be accurately measured by current transient kinetic techniques.
PolB1 exo- exhibited low processivity For a DNA polymerase, the term processivity describes the number of nucleotides incorporated per DNA binding event, and this can be quantitatively defined as the ratio of the polymerization rate divided by the DNA dissociation rate. To kinetically investigate the processivity of PolB1 exo-, a pre-incubated solution of PolB1 exo- (35 nM) and 5′- radiolabeled D-1 DNA (100 nM) were rapidly mixed with dCTP, dGTP, and dTTP (100 µM each) in reaction buffer A before being quenched with EDTA after 8 ms to 1.75 s. These reaction conditions allowed PolB1 to elongate 21/41-mer to 27/41-mer in a single binding event, since the DNA substrate was in molar excess over the enzyme. Each DNA species was resolved using polyacrylamide gel electrophoresis and quantitated using ImageQuant software (Figure 3.3 part (A)). Then, the computer simulation programs, Kinsim [150] and Fitsim [151], were used to fit the data to a mechanism considering six consecutive nucleotide incorporations and seven DNA dissociation rates for the possible binary complexes in the reaction pathway (Figure 3.4). Fitsim generated the following rates of polymerization for the formation of each designated product: 4.8 ± 0.2 s-1 (22- mer), 7.5 ± 0.5 s-1 (23-mer), 12 ± 1 s-1 (24-mer), 8.2 ± 0.8 s-1 (25-mer), 4.9 ± 0.4 s-1 (26- mer), and 15 ± 3 s-1 (27-mer) (Figure 3.3 part (B)). Meanwhile, the rates of DNA dissociation were 0.6 ± 0.1 s-1 for E•21/41-mer, 0.7 ± 0.1 s-1 for E•22/41-mer, 0.7 ± 0.2 s- 1 for E•23/41-mer, 0.4 ± 0.2 s-1 for E•24/41-mer, 0.2 ± 0.1 s-1 for E•25/41-mer, 0.7 ± 0.5 s-1 for E•26/41-mer, and 0.8 ± 0.2 s-1 for E•27/41-mer. These DNA dissociation rates (0.2-0.8 s-1) are greater than the dissociation rates (~0.043 s-1) of PolB1 exo-•DNA measured in Figure 3.1 which suggested that DNA rapidly dissociated from an unidentified intermediate during processive synthesis. Using the average rate of polymerization (8.7 s-1) and DNA dissociation (0.6 s-1), the processivity of PolB1 exo- was calculated to be 15.
54 Elemental effect of nucleotide incorporation The presence of a “burst” (Figure 3.1 part (A)) suggested that the rate of the first turnover is limited by the chemistry step (Step 4 in Scheme 3.1), a protein conformational change preceding chemistry (Step 3 in Scheme 3.1), or a combination of these two steps. To discern the rate-limiting step of correct nucleotide incorporation, the observed rate constants for the incorporations of dTTP and Sp-dTTPαS, an analogue with a phosphothioate group at the α-phosphate position, were measured under single-turnover conditions: PolB1 exo- (120 nM) and 5′-[32P]-labeled D-1 DNA (30 nM) was mixed with 100 µM of dTTP or Sp-dTTPαS in independent reactions. After fitting the curves in Figure 3.5 part (A) to a single-exponential equation (Equation 5 in Section 3.2), the -1 -1 extracted kobs values were 10.3 ± 0.6 s and 7.0 ± 0.4 s for dTTP and Sp-dTTPαS,
respectively. Thus, the elemental effect, a ratio of kobs,dTTP/kobs,Sp-dTTPαS, was calculated to be 1.5. Previously, an elemental effect of 4-11 has been considered evidence for a rate- limiting chemistry step [50], therefore, these results suggested that correct nucleotide incorporation may not be limited by the rate of phosphodiester bond formation (Step 4 in Scheme 3.1).
Next, we performed an analogous experiment to deduce the rate-limiting step of incorrect nucleotide incorporation. For the incorporation of dGTP or Sp-dGTPαS (500 µM) into the same D-1 DNA substrate, the respective observed rate constants were 0.010 ± 0.001 s-1 and 0.00028 ± 0.00002 s-1 which translated into an α-thio elemental effect of 36 (Figure 3.5 part (B)). Relative to what was observed with correct dTTP, this larger elemental effect suggested that phosphodiester bond formation may be partially rate- limiting.
Determination of the dissociation rate from the E•DNA•dNTP complex Following formation of the E•DNA•dNTP complex, DNA can either dissociate (Step 7 in Scheme 3.1) or continue along the reaction pathway as part of a ternary species which undergoes a protein conformational change (Step 3 in Scheme 3.1) leading to the formation of the E′•DNA•dNTP complex. To assess the former possibility, the DNA
55 dissociation rate (k7) of the E•DNA•dNTP complex was measured by reacting a pre- incubated solution of PolB1 exo- (55 nM) and 5′-[32P]-labeled D-1 DNA (60 nM) with Sp-dTTPαS (100 µM) in the absence or presence of an unlabeled DNA trap (2.5 µM D-1 DNA) at 37 °C. Please note, rather than dTTP, Sp-dTTPαS was used since it is incorporated at a slower rate (Figure 3.5 part (A)). This will enhance the kinetic partitioning from E•DNA•dNTP → E + DNA + dNTP. Also, the purpose of the DNA trap is to sequester any of the PolB1 exo- molecules which dissociate (Step 7 of Scheme 3.1) so that the reverse reaction is attenuated. After fitting the data points to equation 1 (Section 3.2), identical burst rates of 7.8 ± 0.4 s-1 were determined, although, the burst amplitudes were 43.3 ± 0.5 nM and 42.3 ± 0.5 nM in the absence and presence of the DNA trap, respectively (Figure 3.6). The amplitude difference increased (~7 nM) when the reaction was performed at a lower temperature of 35 °C which enhanced partitioning from E•DNA•dNTP → E + DNA + dNTP (Figure 3.7). The kinetic parameter k7 was estimated from the following relationship: (Atrap/Ano trap) = kobs/(kobs + k7) = 42.3/43.3 = -1 -1 7.8 s /(7.8 s + k7). Solving this equation yielded a DNA dissociation rate of 0.18 ± 0.01 s-1 for the E•DNA•dNTP complex at 37 °C. This result suggested that DNA dissociation from the ternary complex (Step 7 in Scheme 3.1, k7) is faster relative to the dissociation
of the binary complex (Step 1 in Scheme 3.1, k-1). Interestingly, the value of k7 at 37 °C is similar to the lower limit of the dissociation rates (0.2-0.8 s-1) determined from the global
fitting analysis (Figure 3.3) while the value of k7 at 35 °C is similar to the upper limit (Figure 3.7). Thus, the unidentified intermediate during processive polymerization in Figure 3.3 is likely to be E•DNA•dNTP.
Measurement of the DNA dissociation rate of the E′•DNA•dNTP complex
To determine k8, a steady-state kinetic assay was employed, whereby a pre-incubated solution of PolB1 exo- (1 nM) and 5′-[32P]-labeled D-1 DNA (250 nM) was rapidly mixed with ddTTP (100 µM) in the absence or presence of dCTP (100 µM), the next correct nucleotide. For the former reaction (i.e. without dCTP), the steady-state rate should correspond to the rate of DNA dissociation from the binary complex as measured in Figure 3.1. For the latter reaction (i.e. with dCTP), PolB1 exo- should extend D-1
56 DNA by first incorporating ddTTP (PolB1 exo-•DNAn+1) and then form a ternary complex with dCTP for the second correct incorporation during the first enzyme binding event. However, due to the initially incorporated ddTMP which lacks the 3′-OH, the second incorporation can only proceed up to Step 3 in Scheme 3.1. Thus, following
ddTTP incorporation, PolB1 exo- has to dissociate from E′•DNAn+1•dCTP and then bind to other DNA molecules. Moreover, the multiple turnover rate in the presence of dCTP is
limited by the off rate of DNA from E′•DNAn+1•dCTP. The data points for both reactions were fit to equation 2 (Section 3.2), and the steady-state rates were 0.029 s-1 and 0.0071 s- 1 in the absence and presence of dCTP, respectively (Figure 3.8 part (A)). These steady- state rates differed by 4.1-fold, thereby indicating that (i) the presence of the next correct nucleotide inhibited DNA dissociation and (ii) a tightly bound ternary complex
(E′•DNAn+1•dCTP) was formed.
Since ddTTP was used for the above assay, it is possible that the steady-state rate measured using ddTTP may not be limited by product dissociation from the enzyme. To validate the steady-state rate of 0.029 s-1 measured above, a burst experiment was performed with ddTTP under identical reaction conditions as described in Figure 3.1 part (A). The biphasic curve was fit to equation 6 (Section 3.2) which yielded an observed burst and steady-state rate of 0.3 ± 0.2 s-1 and 0.04 ± 0.01 s-1, respectively (Figure 3.8 part (B)). The similar steady-state rates of 0.029 s-1 and 0.04 ± 0.01 s-1 obtained under steady-state and burst conditions, respectively, confirmed that these rates were limited by DNA dissociation from the E•DNA binary complex.
Pulse-chase and pulse-quench experiments Pulse-chase and pulse-quench experiments were performed to further validate the occurrence of a conformational step (Step 3) preceding the chemistry step (Step 4). For this assay, a pre-incubated solution of PolB1 exo- (100 nM) and unlabeled D-1 DNA (100 nM) was rapidly mixed with [α-32P]dTTP (50 µM). The pulse-quench reactions were immediately quenched with 1 M of HCl, denatured with chloroform, and neutralized with 1 M NaOH. In contrast, the pulse-chase reactions were chased using an
57 excess of unlabeled dTTP (1 mM) for 30 s prior to initiating the quenching, denaturation, and neutralization steps. During the chase time period, the PolB1 exo-•D-1•[α-32P]dTTP complexes, which were partitioning between both forward and reverse directions, would be chased in the forward direction to form more radiolabeled products. If more product formation occurs during the pulse-chase reaction, then this implies that (i) a ternary complex intermediate, which is distinguishable from the ground-state ternary complex PolB1 exo-•D-1•[α-32P]dTTP, occurred along the reaction pathway and (ii) the chemistry step (Step 4 in Scheme 3.1) is flanked immediately before and after by slower steps (Step 3 and Step 5 in Scheme 3.1). Equation 1 (Section 3.2) was applied to the time courses, and amplitudes of 75 ± 3 nM and 61 ± 2 nM for the pulse-chase and pulse-quench assays, respectively, were determined (Figure 3.9). This amplitude difference of 14 nM (~19%) provided evidence for the existence of a stably-bound ternary complex that accumulated prior to the chemical reaction. Since nucleotide binding was rapid for PolB1 exo-, partitioning in the forward direction likely represented an accumulation of E′•DNA•dNTP. Similarly as what we have discussed in the kinetic studies of Dpo4 [111], the accumulated intermediate cannot be either E•DNA•dNTP or E•DNA.
Activation energy barrier for nucleotide incorporation
Arrhenius activation energy (Ea) is the minimum amount of energy required for reactants
to be converted into products during a chemical reaction. To determine the Ea required for PolB1 exo- to elongate a DNA substrate during a single nucleotide incorporation, we
determined the kp values of both correct and incorrect dNTP incorporations at various temperatures (Table 2.3 in Chapter 2). These kp values were then used to construct an
Arrhenius plot [ln(kp) versus temperature (1000/T)] which was subsequently fit to
equation 7 (Section 3.2). This generated an Ea of 38 ± 3 kcal/mol for correct dTTP
incorporation and an Ea of 55 ± 3 kcal/mol for incorrect dATP incorporation (Figure 3.10). Thus, 17 kcal/mol of additional energy is required for PolB1 exo- to catalyze a misincorporation.
58 3.4 Discussion
Minimal kinetic mechanism for PolB1 exo- The kinetic studies of several A-, B-, X-, and Y-family DNA polymerases reveal that these enzymes share a minimal kinetic mechanism of DNA synthesis shown in Scheme 3.1 [54]. Our pre-steady state kinetic analysis of PolB1 exo- suggested that this B-family member followed the same kinetic mechanism and binds substrates sequentially: DNA first followed by an incoming nucleotide. As indicated by the active site titration assay (Figure 3.2), PolB1 exo- binds to a 21/41-mer (primer/template) DNA substrate with a DNA dNTP Kd value of 1.8 nM. The apparent ground-state binding affinity (1/Kd ) of an incoming nucleotide depended on whether a canonical base pair was formed. In general, the apparent binding affinity of a correct Watson-Crick base pair was about 1,000-fold tighter than the weak apparent affinities of incorrect nucleotides (Table 2.2 in Chapter 2). Furthermore, the kinetics of correct dTTP incorporation catalyzed by PolB1 exo- was biphasic (Figure 3.1 part (A)) which suggested that the first turnover was limited by either Step 3, Step 4, or both in Scheme 3.1 (see discussion below) while multiple turnovers were likely limited by dissociation of the elongated DNA product from the enzyme (Step 1). The latter observation was corroborated by the fact that the steady-state turnover rate was similar to the dissociation rate of the PolB1•DNA complex (Figure 3.1). The existence of Step 3 and its role as the rate-limiting step of correct nucleotide addition during the first turnover were supported by the following lines of kinetic evidence. (i) A small, α-thio effect of 1.5 for the rate of dTTP incorporation relative to its α-thio analog, dTTPαS, (Figure 3.5 part (A)) suggested that the chemistry step (Step 4) was unlikely the rate-limiting step. However, other evidences are required to support this conclusion because the intrinsic α-thio effect for any nucleotide incorporation is unknown, it is sensitive to the steric configuration at the α-phosphorus of a nucleotide bound at the active site of a polymerase, and it is influenced by the transition state structure [50, 148]. (ii) Different DNA dissociation rates for k-1, k7, and k8 (Figures 3.1, 3.6, and 3.8 part (A)) suggested the presence of E•DNA, E•DNA•dNTP, and E′•DNA•dNTP complexes, respectively, along the reaction pathway. (iii) A 14 nM (19%)
59 higher reaction amplitude in the pulse-chase reaction over that observed in the pulse- quench reaction (Figure 3.9) suggested the existence of E′•DNA•dNTP which did not yield products under pulse-quench reaction conditions but was chased to products by a large molar excess of dTTP under the pulse-chase conditions. The accumulation of E′•DNA•dNTP also suggested that Step 5 is slow [133]. Notably, the rates for Steps 5 and 6 in Scheme 3.1 were not measured in this paper. (iv) An activation energy of 38 ± 3
kcal/mol for dTTP incorporation was determined from the dependence of lnkp versus 1/T
(Figure 3.10). This Ea value may represent the energy barrier for either Step 3, Step 4, or
both because the kp values are potentially limited by these steps. For uncatalyzed
phosphodiester bond formation in solution, the Ea for its rate-limiting chemistry step is calculated to be 21.1 kcal/mol [152]. If this reaction occurs at a polymerase active site, its
Ea value should be significantly lower than 21.1 kcal/mol because enzymes are known to stabilize transition states and lower the activation energy barriers [153]. Consistently, the activation energy barriers for the chemistry step of phosphodiester bond formation catalyzed by T7 DNA polymerase [152] and human DNA polymerase β [154] are
predicted to be 12.3 and 17 kcal/mol, respectively. In comparison, our measured Ea value of dTTP incorporation was 21-25.7 kcal/mol greater than these predicted values and cannot represent the energy barrier for Step 4. Thus, Step 3 was the rate-limiting step of the first turnover during DNA synthesis catalyzed by PolB1 exo-.
Interestingly, the Ea for correct dTTP incorporation catalyzed by Dpo4 has been determined to be 32.9 kcal/mol and similarly used as one of the four lines of kinetic evidence to identify Step 3 as the rate-limiting step [60, 111]. Previously, Step 3 has also been recognized as the rate-limiting step for the kinetic mechanisms of correct nucleotide incorporation catalyzed by several other enzymes including E. coli DNA polymerase I [133], T7 DNA polymerase [88, 112], HIV-1 reverse transcriptase [113], and yeast DNA polymerase η [115]. Although these enzymes share a rate-limiting protein conformational change, the exact physical nature of Step 3 is unclear. Numerous X-ray crystallographic studies suggest that Step 3 may reflect the open-to-close transition of the finger domain for the A-, B-, and X-family DNA polymerases [119, 121, 122, 155, 156]. In contrast,
60 nucleotide binding only induces the subtle repositioning of select active site residues of the X- and Y-family members [46, 55]. Recently, this assignment of the rate-limiting step has been further questioned due to fluorescence resonance energy transfer-based evidence for two A-family DNA polymerases, E. coli DNA polymerase I [61] and Thermus aquaticus DNA polymerase I [59] which shows that the closure rate of the finger domain is too fast to limit catalysis. Thus, Step 3 may correspond to the subtle repositioning of active site residues, substrates, and the network of water molecules in the active site pocket [157] prior to catalysis. This step may be more dramatic and difficult with high- fidelity PolB1 exo- than with error-prone Dpo4 because PolB1 exo- has a 5.1 kcal/mol
larger Ea value. PolB1 exo- contains a tighter polymerase active site than Dpo4 and interacts more intensely with a nascent base pair than Dpo4, leading to a higher energy cost in Step 3.
For incorrect nucleotide incorporation catalyzed by T7 DNA polymerase [89, 112], E. coli DNA polymerase I [134], and Dpo4 [111], Step 4 has been suggested to be the rate- limiting step. Unfortunately, most of these studies [89, 111, 134] examined the α-thio effect which is largely considered an unreliable criterion (see discussion above) [54]. Based on the α-thio effect of 36 (Figure 3.5 part (B)), Step 4 could be rate-limiting for dGTP misincorporation. However, this conclusion is not supported by the large Ea for dATP misincorporation (55 kcal/mol) which indicated Step 3, rather than Step 4, as rate- limiting. More studies are required to unambiguously identify whether Step 3, Step 4, or both are rate-limiting during a misincorporation.
Kinetic implications of PolB1 interacting with a processivity cofactor According to the rates generated by computer simulation (Figure 3.3), the processivity of PolB1, a replicative B-family enzyme, was estimated to be 15 at 37 °C which is almost identical to the value (i.e. 16) obtained for Dpo4, a lesion-bypass Y-family enzyme notorious for exhibiting poor processivity [111]. The processivity of both PolB1 exo- and Dpo4 was reduced by an unexpectedly fast DNA dissociation rate that was subsequently identified to be the E•DNA•dNTP intermediate; a similar finding has been observed for
61 HIV-1 RT [111, 113]. Furthermore, the two active DNA polymerases of S. solfataricus P2 are not inherently processive. To compensate for this functional shortcoming, PolB1 likely associates with a processivity cofactor to increase the enzyme’s processivity during genomic replication. Such an effect of accessory proteins on the processivity of several replicative DNA polymerases including T4 DNA Pol [158], T7 DNA Pol [159, 160], and human DNA polymerase γ [161] has been reported. For example, when the 140-kDa catalytic subunit of human DNA polymerase γ associates with its 54-kDa accessory subunit, the processivity value increases from 290 to 2250 [161]. Interestingly, PolB1 has been shown to interact with both the PCNA2 and PCNA3 subunits of S. solfataricus P2 PCNA heterotrimer [143]. Through its N-terminal domain, PolB1 can be stimulated by PCNA, as processive DNA synthesis is greatly enhanced [143]. Thus, it will be interesting to determine the kinetic effect of PCNA on DNA polymerization catalyzed by PolB1.
Comparison of kinetic parameters of PolB1 and other DNA polymerases Despite the contrasting active sites of PolB1 and Dpo4 [101, 109], the kinetic parameters DNA discerned for these two polymerases are strikingly similar (Table 3.1) [111]. The Kd value for PolB1 differs by approximately 6-fold relative to Dpo4. Moreover, the most dNTP significant difference is the ground-state binding affinity (1/Kd ) of an incoming nucleotide. For the incorporation of dTTP into D-1 DNA, PolB1 catalyzes this reaction with a 21-fold tighter dNTP binding affinity compared to Dpo4, although, their incorporation rates were close (8.2 s-1 versus 9.4 s-1). The cellular levels of PolB1 and Dpo4 are similar according to Western blot analysis [132], therefore, maintaining low nucleotide concentrations may control the error-prone polymerase activity of Dpo4 on an undamaged DNA template during DNA replication [105].
So far, T4 DNA polymerase is the only B-family DNA polymerase with a well-studied kinetic mechanism [86, 138, 139]. To serve as another basis for comparison, bacteriophage T7 DNA polymerase is an A-family replicative polymerase with a detailed kinetic mechanism [88, 89]. Please note, the kinetic mechanism of T7 DNA Pol was
62 elucidated in the presence of its processivity cofactor, E. coli thioredoxin, which is in contrast to the T4 DNA Pol and PolB1 exo- studies. Interestingly, PolB1 exo- binds to DNA with a tighter binding affinity (~39- and 10-fold) than T4 DNA Pol and T7 DNA Pol under the specified reaction conditions (Table 3.1) [86, 88]. Also noteworthy is that
T4 DNA Pol has a dramatically faster k-1, however, this may be in part due to the use of a short primer-template substrate that is not optimal for the enzyme to bind (Table 3.1) [86]. Additionally, the fast dissociation rate of T4 DNA Pol may be attenuated by the accessory proteins, gp44, 62, 45, and 32. In regards to dNTP binding, all three replicative dNTP polymerases follow the same trend, whereby the Kd values of correct nucleotides are in the low micromolar range and mismatched nucleotides are in the millimolar range, a difference of almost three orders of magnitude [86, 88, 89, 146]. Thus, regulating the cellular concentrations of nucleotide pools is a natural way to limit mistakes by PolB1 in vivo. The rates of polymerization catalyzed by PolB1 exo- versus the bacteriophages vary widely, but this is largely a function of the temperature profile for each enzyme (Table -1 3.1). In the Chapter 2 (Table 2.3), k3 for PolB1 exo- was measured to be 92 s at a reaction temperature of 50 °C, therefore, it is plausible that catalysis by PolB1 is even faster at the physiological temperature of 80 °C [146]. In summary, these three replicative polymerases reveal nuances in their kinetic parameters for various elementary steps which stress the importance of defining these numerical values and how related polymerases defy a unified mechanism.
In summary, the forward polymerization pathway of PolB1 exo- has been investigated. Our pre-steady state kinetic results supported a rate-limiting conformational change during nucleotide incorporation. Together with our kinetic studies on Dpo4 [111], these kinetic mechanisms provide a foundation for investigating DNA polymerase switching and other complex DNA transactions that occur at the replication fork.
63 3.5 Future Directions
DNA replication and lesion bypass are processes performed in the presence of accessory proteins such as the processivity factor, PCNA. Both PolB1 and Dpo4 interact with PCNA [143, 145]. PCNA is likely an important structural factor when PolB1 and Dpo4 switch at sites of DNA damage. Therefore, it is important to understand how PCNA affects the mechanism of polymerization catalyzed by PolB1 and Dpo4 on both damaged and undamaged DNA. Important steps that may be altered in the kinetic pathway include DNA binding and dissociation (Step 1), nucleotide binding (Step 2), and nucleotide incorporation (Steps 3 and 4) (Scheme 3.1). Understranding these basic steps will be important for elucidating the molecular basis of polymerase switching.
64 3.6 Table
Table 3.1 Estimated kinetic constants. Polymerase PolB1 exo- Dpo4 T4 DNA T7 DNA polymerase polymerase Polymerase B Y B A family Parameter Value -1 -1 -1 -1 -1 -1 -1 -1 k1 24 µM s 1.9 µM s 85 µM s 11 µM s -1 -1 -1 -1 k-1 0.043 s 0.02 s 6 s 0.2 s DNA Kd 1.8 nM 10.6 nM 70 nM 17.8 nM -1 -1 -1 -1 -1 -1 k2 100 µM s 100 µM s ND ≥50 µM s -1 -1 -1 k-2 1,100 s 23,000 s ND ≥1,000 s dNTP Kd 11 µM 230 µM 20 µM 18 µM -1 -1 -1 -1 k3 8.2 s 9.4 s 400 s 287 s -1 -1 k7 0.18 s 0.41 s ND ND -1 -1 k8 0.0071 s 0.004 s ND ND Reaction 37 37 20 20 temperature (°C) Reference This work [111] [86] [88]
65 3.7 Figures
Figure 3.1 Pre-steady state and steady-state kinetics of dTTP incorporation into D-1 DNA by PolB1 exo-. (A) A pre-incubated solution of PolB1 exo- (30 nM) and 5′-[32P]- labeled D-1 DNA (120 nM) was rapidly mixed with dTTP (100 µM) and quenched at various times with 0.37 M EDTA. The data were fit by nonlinear regression to a biphasic equation (equation 1, see 3.2) with an observed burst and steady-state rate of 9 ± 1 s-1 and 0.074 ± 0.05 s-1, respectively. (B) Incorporation of dTTP into D-1 was independently measured under steady-state conditions whereby the addition of 100 µM dTTP with a pre-equilibrated solution of PolB1 exo- (1 nM) and 5′-[32P]-labeled D-1 DNA (250 nM) initiated the reaction. At various times, the reaction was terminated using 0.37 M EDTA. The data were fit to a linear relationship (Equation 2 in Section 3.2) which yielded a steady-state rate constant of 0.038 s-1. (C) The DNA dissociation rate constant from the binary complex was measured by mixing a pre-equilibrated solution of PolB1 exo- (50 nM) and 5′-[32P]-labeled D-1 DNA (100 nM) with unlabeled D-1 DNA (2.5 µM) for various time intervals. Then, dTTP (133 µM) was added for 15 sec before quenching the reaction. Data were fit to a single-exponential equation (Equation 3 in Section 3.2) to obtain a rate constant of 0.043 ± 0.003 s-1.
66
5’-CGCAGCCGTCCAACCAACTCA-3’ D-1 3’-GCGTCGGCAGGTTGGTTGAGTAGCAGCTAGGTTACGGCAGG-5’
A B 30 14
25 12
10 20 8 15 6 10 Product (nM) Product (nM) 4
5 2
0 0 0 5 10 15 20 0 50 100 150 200 250 Time (s) Time (s)
C 100
80
60
40 Product (nM)
20
0 0 200 400 600 800 1000 1200 Time (s)
Figure 3.1
67 20
15
10
5 PolB1:21/41mer (nM) PolB1:21/41mer
0 020406080100
DNA (nM)
Figure 3.2 Active site titration of PolB1 exo-. PolB1 exo- (45 nM) and increasing concentrations of 5′-radiolabeled D-1 DNA (2.5-150 nM) were mixed with dTTP (100 µM). All reactions were quenched at 1.8 s using EDTA. The product concentration was plotted as a function of substrate concentration and fit to equation 4 (Section 3.2). An equilibrium dissociation constant of 1.8 ± 0.4 nM and an active enzyme concentration of 18.9 ± 0.4 nM were measured.
68
Figure 3.3 PolB1 exo- displayed low processivity. A pre-incubated solution of PolB1 exo- (35 nM) and 5′-radiolabeled D-1 DNA (100 nM) was reacted with dCTP, dGTP, and dTTP (200 µM each) for various times until being quenched with 0.37 M EDTA. Products were analyzed using sequencing gel analysis and the gel image is shown in (A). (B) The amount of remaining substrate (21-mer, ■) and each product (22-mer, ●; 23- mer, ▲; 24-mer, ♦; 25-mer, □; 26-mer, ○; 27-mer, ) were plotted as a function of time. The solid lines represent the best fits generated by computer simulation using a mechanism that defines the rates for six consecutive nucleotide incorporations and seven DNA dissociations from the possible PolB1•DNA species. The rates of polymerization for the formation of each product were 4.8 ± 0.2 s-1 for 22-mer, 7.5 ± 0.5 s-1 for 23-mer, 12 ± 1 s-1 for 24-mer, 8.2 ± 0.8 s-1 for 25-mer, 4.9 ± 0.4 s-1 for 26-mer, and 15 ± 3 s-1 for 27-mer while the rates of DNA dissociation were 0.6 ± 0.1 s-1 for E•21/41-mer, 0.7 ± 0.1 s-1 for E•22/41-mer, 0.7 ± 0.2 s-1 for E•23/41-mer, 0.4 ± 0.2 s-1 for E•24/41-mer, 0.2 ± 0.1 s-1 for E•25/41-mer, 0.7 ± 0.5 s-1 for E•26/41-mer, and 0.8 ± 0.2 s-1 for E•27/41-mer.
69 A
B
Figure 3.3
70
Figure 3.4 Kinsim and Fitsim analysis. The above reaction scheme represents the mechanism entered into Kinsim. There were six consecutive polymerization reactions (k1-k6) and seven DNA dissociation events (k7-k13). Please note, three nucleotides (dCTP, dGTP, and dTTP) were present during the reaction, although, this mechanism does not explicitly account for them. For the Fitsim analysis, all thirteen rate constants were varied -1 -1 -1 to obtain polymerization rates of k1 = 4.8 ± 0.2 s , k2 = 7.5 ± 0.5 s , k3 = 12 ± 1 s , k4 = -1 -1 -1 8.2 ± 0.8 s , k5 = 4.9 ± 0.4 s , k6 = 15 ± 3 s and DNA dissociation rates of k7 = 0.6 ± 0.1 -1 -1 -1 -1 -1 s , k8 = 0.7 ± 0.1 s , k9 = 0.7 ± 0.2 s , k10 = 0.4 ± 0.2 s , k11 = 0.2 ± 0.1 s , k12 = 0.7 ± -1 -1 0.5 s , and k13 = 0.8 ± 0.2 s .
71 A 30
25
20
15
10 Product (nM)
5
0 00.511.52 Time (s)
B 30
25
20
15
10 Product (nM) Product
5
0 050001 104 1.5 104 Time (s)
Figure 3.5 Elemental effect on the rate of correct and incorrect nucleotide incorporation. PolB1 exo- (120 nM) and 5′-[32P]-labeled D-1 DNA (30 nM) was rapidly mixed with (A) 100 µM dTTP (●) or Sp-dTTPαS (○) in parallel time courses. The data were fit to a single-exponential equation (Equation 5 in Section 3.2) to extract kobs values of 10.3 ± 0.6 s-1 and 7.0 ± 0.4 s-1 for dTTP and Sp-dTTPαS, respectively, for a calculated elemental effect of 1.5. (B) The same PolB1•D-1 solution was mixed with 500 µM dGTP (●) or Sp- dGTPαS (○) in parallel time courses. The data were fit to a single-exponential equation -1 (Equation 5 in Section 3.2) to extract kobs values of 0.010 ± 0.001 s and 0.00028 ± 0.00002 s-1 for dGTP and Sp-dGTPαS, respectively, for a calculated elemental effect of 36.
72 50
40
30
20 Product (nM) Product
10
0 0 5 10 15 20
Time (s)
Figure 3.6 Measurement of the dissociation rate for the E•DNA•dNTP complex. PolB1 exo- (55 nM) and 5′-[32P]-labeled D-1 DNA (60 nM) was rapidly mixed with 100 µM Sp- dTTPαS in the absence (●) or presence of (○) an unlabeled DNA trap (2.5 µM of D-1). An identical burst rate of 7.8 ± 0.4 s-1 was determined for both time courses while the burst amplitudes were 43.3 ± 0.5 nM and 42.3 ± 0.5 nM in the absence and presence of the DNA trap, respectively. Thus, an amplitude reduction of 2.3% equates to a k7 of 0.18 ± 0.1 s-1.
73 60
50
40
30
20 Product (nM) Product
10
0 0 1020304050 Time (s)
Figure 3.7 Measurement of the dissociation rate for the E•DNA•dNTP complex at 35 °C. PolB1 exo- (55 nM) and 5′-[32P]-labeled D-1 DNA (60 nM) was rapidly mixed with 100 µM dTTP in the absence (●) or presence of (○) an unlabeled DNA trap (2.5 µM of D-1). An identical burst rate of 5.5 ± 0.6 s-1 was determined for both time courses. The burst amplitudes were 41 ± 1 nM and 34.3 ± 0.9 nM in the absence and presence of the DNA -1 trap, respectively. An amplitude reduction of 16% equates to a k7 of 1.1 ± 0.1 s .
74 A 12
10
8
6
4 Product (nM)
2
0 050100150200250 Time (s)
B 60
50
40
30
20 Product (nM) Product
10
0 0 102030405060 Time (s)
Figure 3.8 Measurement of the dissociation rate for the E′•DNA•dNTP complex. (A) PolB1 exo- (1 nM) and 5′-[32P]-labeled D-1 DNA (250 nM) was rapidly mixed with 100 µM ddTTP in the absence (●) or presence of (○) the next correct nucleotide (100 µM of dCTP). Fitting the data to equation 2 (Section 3.2) generated a steady-state rate of 0.029 s-1 and 0.0071 s-1 in the absence and presence of dCTP, respectively. (B) A pre-incubated solution of PolB1 exo- (30 nM) and 5′-[32P]-labeled D-1 DNA (120 nM) was rapidly mixed with 100 µM of ddTTP. Data were fit to equation 6 (Section 3.2) which yielded an observed burst and steady-state rate of 0.3 ± 0.2 s-1 and 0.04 ± 0.01 s-1, respectively.
75 100
80
60
40 Product (nM) Product
20
0 00.511.52 Time (s)
Figure 3.9 Pulse-chase and pulse-quench experiment. A pre-incubated solution of PolB1 exo- (100 nM) and unlabeled D-1 DNA (100 nM) was rapidly mixed with 50 µM [α- 32P]dTTP. The pulse-quench (○) reactions were immediately quenched with HCl (1 M) while the pulse-chase (●) reactions were chased with 1.0 mM of non-radioactive dTTP for 30 s prior to acid quenching. Fitting the data to equation 1 (Section 3.2) yielded amplitudes of 75 ± 3 nM and 61 ± 2 nM, for the pulse-chase and pulse-quench reactions, respectively.
76 6
4
2 ) -1
) (s 0 p k
ln( -2
-4
-6 3.1 3.2 3.3 3.4 -1 1000/T (K )
Figure 3.10 Activation energy for nucleotide incorporation. The kp values [extracted from Table 2.3] for dTTP (●) and dATP (○) were plotted as a function of reaction temperature and subsequently fit to equation 7 (Section 3.2). An activation energy of 38 ± 3 kcal/mol was determined for correct (dTTP:dA) nucleotide incorporation while an activation energy of 55 ± 3 kcal/mol was determined for incorrect (dATP:dA) nucleotide incorporation.
77 3.8 Scheme
Scheme 3.1
78 Chapter 4: A Novel Mechanism of Sugar Selection Utilized by Human DNA Polymerase λ
4.1 Introduction
DNA polymerases are classified into one of six families: A, B, C, D, X, or Y. In addition, the closely-related reverse transcriptases (RT), which catalyze DNA synthesis using both DNA and RNA templates, comprise a distinct family: the RT family. Most DNA polymerases and RTs are endowed with a stringent nucleotide recognition mechanism which prevents the aberrant incorporation of ribonucleotides (rNTP), the building block of RNA. Sugar specificity at the 2′ position occurs through a steric exclusion model, whereby a bulky side chain residue clashes with the 2′-hydroxyl of an incoming ribonucleotide. Depending upon the polymerase family, this residue has been identified as a Glu for the A-family [162] and a Tyr/Phe for the B-, Y- and/or RT-families [163- 168]. A wide range of discrimination factors have been measured using kinetic methodologies. Most polymerases and RTs exhibit sugar selectivity greater than 1000- fold [162, 164-167]. However, two X-family members, DNA polymerase µ (Pol µ) [169, 170] and terminal deoxynucleotidyltransferase (TdT) [171], select deoxyribonucleotides (dNTPs) over ribonucleotides by a maximum of 11-fold. Interestingly, both of these enzymes possess a Gly (Figure 4.1 part (A)) at the predicted ‘steric gate’ position, thereby providing a structural basis for their relaxed sugar specificities. In contrast, DNA polymerase λ (Pol λ) and DNA polymerase β (Pol β), both of which belong to the X- family, have Tyr and Phe residues at and nearby the predicted ‘steric gate’ position (Figure 4.1).
79 Pol λ, which shares 34% sequence identity with Pol β, is postulated to function in DNA repair pathways such as base excision repair (BER) and non-homologous end joining (NHEJ) [78, 172-177]. The cellular concentration of ribonucleotide pools has been found to be 10- to 200-fold greater than deoxyribonucleotide pools, and the level of rNTPs remains relatively high throughout the cell cycle [178, 179]. Since DNA repair occurs
when dNTP pools are low (e.g. NHEJ occurs during the G1 phase [180] and BER during
G0-G1 checkpoint [181]), it would be imperative that Pol λ displays high sugar selectivity in order to maintain genomic integrity. Using pre-steady state kinetic methods, protein engineering techniques, and rCTP analogs, we measured the sugar selectivity of Pol λ and elucidated a novel sugar selection mechanism utilized by Pol λ.
4.2 Materials and Methods
Materials 32 These chemicals were purchased from the following companies: [γ- P]ATP, MP Biomedicals; deoxyribonucleotide-5′-triphosphates, GE Healthcare; ribonucleotide-5′- triphosphates, MBI Fermentas; 2′-aracytidine-5′-triphosphate (araCTP), 2′-deoxy-2′,2′- difluorocytidine-5′-triphosphate (GemCTP), 2′-fluoro-2′-deoxycytidine-5′-triphosphate
(2′-F-CTP), and 2′-amino-2′-deoxycytidine-5′-triphosphate (2′-NH2-CTP), and 2′-O- methylcytidine-5′-triphosphate (2′-OCH3-CTP), TriLink Biotechnologies; Biospin columns, Bio-Rad Laboratories; OptiKinase™, USB Corporation; Microcon centrifugal filter devices, Millipore; synthetic oligodeoxyribonucleotides 21-mer, 5′-phosphorylated 19-mer, and 41-mers, Integrated DNA Technologies. The structural model was completed using software created by Swiss PDB and PyMol.
Mutagenesis, expression and purification of wild-type and mutant forms of Pol λ Mutations and deletions were introduced into the pET28b-Pol λ plasmid encoding full- length human DNA polymerase λ (1-575) using a QuikChange XL site-directed mutagenesis kit (Stratagene). The mutated or deleted regions of the Pol λ gene were
80 confirmed via DNA sequencing (Plant-Microbe Genomics Facility at The Ohio State University). The expression and purification of wild-type Pol λ and its variants was performed as previously described [117].
Circular dichroism spectroscopic studies Each Pol λ variant (11.3 µM) was added to a degassed buffer (25 mM sodium phosphate
at pH 7.5, 50 mM NaCl, 5 mM MgCl2, and 10% glycerol) by performing a series of buffer exchanges. The CD spectra were collected in a 1-mm path-length cuvette at 37 °C using an AVIV CD spectrometer model 62A DS (Lakewood, NJ, USA). Data points were recorded at 1-nm intervals from 270 to 200 nm. Spectra of the buffer were subtracted from the sample spectra and the molar ellipticity (degree cm2 dmol-1) was plotted as a function of wavelength.
Single-nucleotide gapped DNA substrates Commercially-synthesized oligomers in Figure 4.2 part (A) were purified using polyacrylamide gel electrophoresis. The 21-mer primer was radiolabeled with [γ-32P]ATP and OptiKinase™ as previously described [117, 182]. The single-nucleotide gapped DNA substrates were prepared by mixing the 5′-[32P]-radiolabeled 21-mer, the appropriate non-radiolabeled downstream strand 19-mer, and the appropriate 41-mer template at a 1:1.25:1.15 molar ratio, respectively. Then, the annealing mixture was denatured at 95 °C for 6 minutes and slowly cooled to room temperature over several hours.
Measurement of the kp and Kd for single-nucleotide incorporation Kinetic assays were completed using optimized buffer L (50 mM Tris-Cl, pH 8.4 at 37
°C, 5 mM MgCl2, 100 mM NaCl, 0.1 mM EDTA, 5 mM DTT, 10% glycerol, and 0.1 mg/ml BSA) as previously described [182]. All kinetic experiments described herein were performed at 37 °C and the reported concentrations were final after mixing all the components. A pre-incubated solution containing wild-type Pol λ (120 nM) or a Pol λ mutant (300 nM) and a single-nucleotide gapped DNA substrate (30 nM) was mixed with
81 increasing concentrations (0.25-1000 μM) of nucleotide in buffer L at 37 °C. Aliquots of the reaction mixtures were quenched at various times using 0.37 M EDTA. A rapid chemical-quench flow apparatus (KinTek) was utilized for fast nucleotide incorporations. Reaction products were resolved using sequencing gel electrophoresis (17% acrylamide, 8 M urea) and quantitated with a Typhoon TRIO (GE Healthcare). The time course of product formation at each nucleotide concentration was fit to a single-exponential equation (Equation 1)
[Product] = A[1 – exp(–kobst)] (1) using a nonlinear regression program KaleidaGraph (Synergy Software) to yield an observed rate constant of nucleotide incorporation (kobs). The kobs values were then plotted as a function of nucleotide concentration and fit using the hyperbolic equation (Equation 2)
kobs = kp[dNTP]/{[dNTP] + Kd} (2)
which resolved the kp and Kd values for nucleotide incorporation catalyzed by wild-type Pol λ or a mutant. When nucleotide binding was too weak to reach saturation, the plot of
kobs versus nucleotide concentration was fit to equation 3
kobs = (kp/Kd)[dNTP] (3)
which extracted the substrate specificity constant, kp/Kd.
4.3 Results
Ribonucleotide incorporation catalyzed by human DNA polymerase λ Previously, we used pre-steady state kinetic techniques to define the catalytic efficiency and fidelity of all 16 possible dNTP:dN base pair combinations catalyzed by the full- length human Pol λ [117]. To measure the sugar selectivity of Pol λ, we employed the
same kinetic methodology. The maximum rate of incorporation (kp) and the equilibrium
dissociation constant (Kd) of a matched incoming ribonucleotide was measured by mixing a pre-incubated solution of Pol λ and single-nucleotide gapped D-1 DNA (Figure 4.2 part (A)) with increasing concentrations of a ribonucleotide, UTP (Section 4.2). At discrete
82 times, aliquots of this reaction mixture were quenched using EDTA and subsequently resolved using denaturing polyacrylamide gel electrophoresis. A plot of product concentration versus time (Figure 4.2 part (B)) was fit to a single-exponential equation
(Equation 1) to extract the observed rate constant (kobs). Then, these kobs values were plotted as a function of UTP concentration (Figure 4.2 part (C)) and fit to a hyperbolic -1 equation (Equation 2) which yielded a kp of 0.053 ± 0.001 s and a Kd of 114 ± 8 µM (Table 4.1). The pre-steady state kinetic parameters were determined for the remaining, matched rNTPs and were used to calculate the catalytic efficiency, fidelity, and sugar selectivity as defined in Table 4.1. Pol λ displayed a high sugar selectivity range (3,000 to 50,000) and a moderate fidelity (10-4 to 10-5) for all four matched rNTPs. Thus, during gap-filling DNA synthesis, the probability of Pol λ inserting a matched rNTP is relatively low.
We also examined the possibility of Pol λ inserting a mismatched rNTP opposite a DNA template. The kinetic parameters (Table 4.2) were determined for rCTP and rGTP incorporation into D-1 DNA (Figure 4.2 part (A)); no incorporation was observed for rATP. Surprisingly, there was only a 13- and 21-fold decrease in the catalytic efficiency of Pol λ incorporating a mismatched rGTP or rCTP versus a matched UTP, therefore, the fidelity of Pol λ was higher (10-5 to 10-6) for the incorporation of mismatched rNTPs over matched rNTPs. These differences in substrate specificity between matched (UTP) and
mismatched rNTP incorporation originated mostly from a reduced kp (5- and 8-fold) for
rCTP and rGTP, although, the binding affinity (1/Kd) was weakened slightly by 2.6-fold.
Kinetic analysis of Pol λ mutants Next, our objective was to determine the mechanistic basis of Pol λ’s high sugar selectivity. Pol λ possesses two bulky amino acid residues, Y505 and F506 (Figure 4.1 part (A)), which are in close proximity (~3-8 Å) to the C2′-position of an incoming ddTTP (Figure 4.1 part (B)). To determine if these side chains clash with the 2′-hydroxyl of an incoming rNTP, we created three single-point Pol λ mutants: Y505G, Y505A, and F506A. Although Gly is considered a risky substitution due to its conformational
83 flexibility, we chose this amino acid residue because the other two X-family polymerases with low sugar selectivity, i.e. Pol µ and TdT, encode a Gly residue at this position (Figure 4.1 part (A)). First, we used circular dichroism (CD) spectroscopy to determine if the fold of these mutants was significantly different compared to wild-type Pol λ (Section 4.2). The CD spectra were resolved and overlaid for Y505A, F506A, and wild-type Pol λ (Figure 4.3). The spectra indicated that these mutants were folded similar to wild-type Pol λ.
Under single-turnover conditions, the kinetic parameters were measured for dTTP and UTP incorporation into D-1 DNA catalyzed by Y505G and F506A (Table 4.3). Since Pol λ exhibited the lowest sugar selectivity with UTP, the dTTP/UTP substrate pair was chosen as the sugar selectivity probe so that fluctuations, whether it is higher or lower, in the sugar selectivity of the mutants could be measured accurately. The efficiency of dTTP incorporation catalyzed by Y505G and F506A was on par with wild-type Pol λ. Interestingly, the sugar selectivity was 700 and 3,000 for Y505G and F506A, respectively. Relative to wild-type Pol λ, Y505G had 4-fold lower sugar selectivity while mutating F506A did not affect the sugar selection. The most notable kinetic change was a 3-fold rate enhancement for F506A inserting UTP, but the other kinetic parameters displayed a ~2-fold difference from wild-type Pol λ. Based on the sugar selectivity values in Table 4.3, the side chain of F506 did not act as a steric gate while the side chain of Y505 played a role in excluding ribonucleotides by Pol λ. However, the flexibility of Gly at position 505 may either allow residues near the Y505G-G508 segment to reposition so that another residue (e.g. L504 and F506) can act as a weak steric gate or increase the flexibility of the backbone segment. Therefore, we determined the kinetic parameters for dTTP and UTP incorporation into D-1 DNA catalyzed by Y505A (Table 4.3). This single amino acid substitution was predicted to have a minimal effect on the position and structure of the Y505A-G508 segment. The sugar selectivity for Y505A was 360 which is 9- and 2-fold less selective than wild-type Pol λ and the Y505G mutant, respectively. Most notably, the binding of UTP to the Y505A•D-1 complex was 16- and 7-fold tighter than wild-type Pol λ and the Y505G mutant, respectively. The UTP binding affinity
84 difference suggested that our abovementioned concern in regards to the Y505G mutation may be problematic in assessing its role in rNTP selection. However, the tighter UTP binding did not lead to a rate enhancement of UTP incorporation catalyzed by Y505A. Collectively, these kinetic data revealed that the side chain and positioning of Y505 contribute to Pol λ’s sugar selectivity, although, the reduced side chain volume of Pol λ Y505G and Y505A did not relax the sugar selectivity to a similar level as Pol µ and TdT [169, 170]. These relatively high sugar selectivity values suggested that Pol λ does not exclusively use the side chain of Y505 to discriminate against ribonucleotides. Based on the ternary crystal structure of truncated Pol λ•gapped DNA•matched ddNTP [55], Pol λ may also utilize the backbone segment of the peptide Y505-G508 to maintain high sugar selectivity (Figure 4.1 part (B)). To examine this possibility, we created two Pol λ deletion mutants: Y505d and Y505d-T507d. By removing these 1 or 3 amino acid residues, the objective was to create a larger space that would eliminate the steric clash between the 2′-OH of an rNTP and the side chain or backbone segment of Y505-T507 within the active site of Pol λ. The Y505d mutant could not be expressed in E. coli as a soluble and active enzyme, however, a small amount of the Y505-T507d mutant was purified despite a low level of expression. Since the CD spectra in Figure 4.3 suggested that the overall protein fold of Y505-T507d was similar to wild-type Pol λ, we further assayed the sugar selectivity of this triple deletion mutant. A pre-incubated solution of Y505-T507d (120 nM) and 5′-radiolabeled D-8 (30 nM, Figure 4.2 part (A)) was reacted with either correct dGTP (64 μM) or matched rGTP (64 μM) for various times before quenching the polymerization reaction. After fitting the two time courses of product concentration to Equation 1 (Section 4.2), the nucleotide incorporation rates were determined to be 2.0 × 10-4 s-1 (dGTP) and 2.0 × 10-5 s-1 (rGTP) (data not shown). Based
on the incorporation rate ratio (kdGTP/krGTP) under the same reaction conditions, the sugar selectivity of Y505-T507d was estimated to be 10, which is much lower than 14,000 of wild-type Pol λ (Table 4.1) and 360 of Y505A (Table 4.2). These results suggest that the backbone segment of Y505-G508 may function as a steric gate. However, further structural and biochemical studies are needed to validate this conclusion since the deletion of three residues caused a dramatic drop in polymerase efficiency which may
85 either significantly attenuate the sugar selectivity or make it difficult to be measured. Based on a 12,500-fold decrease in polymerase activity, the deletion may have repositioned other residues in the helix-loop-helix segment or have caused other subtle to mild structural perturbations at the active site which could not be revealed using CD spectroscopy (Figure 4.3).
Incorporation of rCTP-based analogs catalyzed by Pol λ To better understand the steric interactions between Pol λ and an incoming ribonucleotide, we assayed wild-type Pol λ in the presence of several rCTP analogs which have various chemical groups at the 2′-positions (Figure 4.4). rCTP was selected due to the physiological relevance of commercially available analogs. As the size and number of functional groups at the 2′-position increased, the binding affinity (1/Kd), the incorporation rate (kp), and the substrate specificity decreased while the sugar selectivity increased. For example, the presence of one (2′-F-CTP) versus two (GemCTP) fluorine atoms increased the sugar selectivity by 6- and 23-fold, respectively (Table 4.4). Moreover, the orientation of the 2′-hydroxyl was another important factor, since araCTP, which is a steric isomer of rCTP with a 2′-hydroxyl group directed above the ribose ring (OH↑), was incorporated 1,800-fold faster than rCTP with a 2′-OH pointing down (OH↓), and its substrate specificity was only 2-fold lower than dCTP (Table 4.4). Pol λ showed high sugar discrimination when an amine or methoxy group was at the 2′ position, for the sugar selectivity was respectively 37,000 and 1,000,000 for 2′-NH2-CTP and 2′-OCH3- CTP which is almost 10- and 250-fold greater than rCTP (Table 4.4). These data suggested that the larger size of an amine and methoxy over a hydroxyl group created more steric hindrance during incorporation. Overall, the sugar selectivity was correlated with the size and orientation of the 2′-group: araCTP (OH↑) < 2′-F-CTP (F) < GemCTP
(F2) < rCTP (OH↓) < 2′-NH2-CTP (NH2) < 2′-OCH3-CTP (OCH3).
86 4.4 Discussion
Human DNA polymerase λ exhibited high sugar selectivity Pol λ is an enzyme proposed to participate in DNA repair pathways which may occur when cellular dNTP pools are low and rNTP pools are high. Therefore, using pre-steady state kinetic techniques, we measured the sugar selectivity of Pol λ to assess the potential of Pol λ aberrantly inserting an rNTP into a single-nucleotide gapped DNA substrate. These data revealed that Pol λ exhibited a relatively high sugar selectivity (3,000 – 50,000), although, it depended on the base identity of the rNTP (Table 4.1). The order of sugar selectivity was UTP < rCTP < rGTP < rATP. In comparison to a correct dNTP, Pol λ displayed a weaker binding affinity (53-fold on average) and a slower rate of incorporation (117-fold on average) with matched rNTPs (Table 4.1) [117]. Furthermore, the fidelity (10-4 to 10-5) of matched rNTP incorporation was comparable to a mismatched dNTP incorporation [117], thereby suggesting the insertion of matched rNTPs would be more likely than dNTP misinsertions during DNA repair in vivo due to higher cellular concentrations of rNTPs than dNTPs. DNA repair pathways (e.g. NHEJ occurs during the G1 phase [180] and BER during G0-G1 checkpoint [181]) proceed outside of S phase when dNTP pools are low but rNTP pools are high. In regards to mismatched rNTP incorporation into DNA, the sugar selectivity was 9- to 20-fold higher compared to matched rNTP insertion (Tables 4.1 and 4.2). Overall, Pol λ displayed almost 460-fold less discrimination for matched versus mismatched rNTP (16-fold on average) compared to dNTP (7,100 on average) incorporations into D-1 DNA [117].
For other DNA polymerase and RT families, the sugar selectivity, as measured using kinetic techniques, spans from 1 to over 1,000,000. The sugar selectivity of Pol λ is similar to A-, B-, X-, and Y-family DNA polymerases such as the Klenow fragment of Escherichia coli DNA polymerase I (3,400 to 1,700,000) [162], the exonuclease-deficient RB69 DNA polymerase (64,000) [166], Pol β (2,000 to 6,000) [169], and the DinB homolog DNA polymerase from Sulfolobus solfataricus (3,400) [167]. In contrast, Pol µ (1 to 11) and TdT (3 to 9) have extremely low sugar selectivity values with gapped DNA
87 [169]. Although most DNA polymerases can incorporate rNTPs, some of them are inhibited by an RNA-terminated primer [162, 164, 170, 171]. However, a previous study showed that Pol λ can efficiently extend an RNA/DNA substrate, although, extension using RNA/RNA and DNA/RNA substrates was less efficient [183].
Mechanism of sugar selectivity employed by Pol λ The mechanism of sugar selectivity has been examined for several DNA polymerases and reverse transcriptases. From these works, a general mechanism for nucleotide selection emerged: the 2′-hydroxyl of a rNTP clashes with a bulky side chain of an active site residue (i.e. Glu, Phe, or Tyr) which has been termed the ‘steric gate’ [184]. Although Pol λ encodes potential ‘steric gate’ residues based on the sequence alignment analysis (Figure 4.1 part (A)), this enzyme displays high sugar discrimination (Table 4.1), and the side chains do not appear to provide a structural basis for sugar selectivity, as the backbone segment Y505-G508 is in closer proximity to C2′ (Figure 4.1 part (B)) [53, 55]. As presented herein, Pol λ mutants Y505G, Y505A, and F506A maintained relatively high sugar selectivity (360 to 3,000) compared to wild-type Pol λ (3,000) (Table 4.3). These data suggested that the side chains of Y505 and F506 respectively contributed ~9- fold and none to Pol λ’s sugar selectivity. This was in stark contrast to other DNA polymerases and RTs, whereby mutating the ‘steric gate’ residues (i.e. Glu, Tyr, or Phe) to Ala or Val resulted in mutants with sugar selectivity values that were reduced from 765 to 50,000-fold based upon kinetic measurements [162, 165-167]. Thus, if the Y505 side chain of Pol λ served as the predominant ‘steric gate’, then the sugar selectivity of Y505A and Y505G should have been approximately 20- to 100-fold less than the measured values of 360 and 700. Therefore, other structural component(s) played a major role as the ‘steric gate’ in controlling the sugar selectivity of Pol λ.
Our pre-steady state kinetic studies showed that the sugar selectivity of Pol λ is 3,000 to 50,000 for a matched rNTP (Table 4.1) while McElhinny and Ramsden have shown that the sugar selectivity of Pol β is 2,000 to 6,000 [169]. Based on the ternary crystal structure of Pol β [53], Pelletier et al. proposed that the backbone segment, Y271-G274,
88 is the structural basis for this enzyme’s sugar selectivity. Their hypothesis prompted us to propose the following mechanism for the sugar selectivity of Pol λ: the ribose 2′- hydroxyl clashes with both the side chain of Y505 (9-fold) and a backbone segment of the peptide Y505-G508 (360-fold). Please note, the exact contribution of the Y505 side chain and the backbone segment likely depends on the base identity of the rNTP (Table 4.1). To provide insight into the major ‘steric gate’ role of the Y505-G508 backbone segment, we generated a triple deletion mutant by removing the Y505-T507 tripeptide segment in order to create more space to accommodate the ribose 2′-hydroxyl group and to decrease the sugar selectivity of Pol λ. Fortunately, we purified a small amount of the triple-deletion mutant. Although about 12,500-fold less active than wild-type Pol λ, the triple-deletion mutant exhibited a low sugar selectivity of 10 (Section 4.3). Thus, these kinetic data indicated that residues Y505-T507, especially Y505, were critical for catalytic activity and sugar selectivity. Interestingly, the G433Y mutant of human Pol µ displays decreased polymerization activity [170]. Although Pol λ and Pol µ respectively encode Tyr and Gly at this position, both of these residues contribute significantly to catalysis.
To illustrate how the backbone segment of Y505-G508 played the major ‘steric gate’ role, we modeled ATP into the active site of Polλ bound to a gapped DNA substrate (Figure 4.5). To create this model, the structure of ATP, which was bound by HIV-1 RT (PDB 2IAJ), was superimposed with the ddTTP substrate in complex with Pol λ and gapped DNA (PDB 1XSN) [55, 185]. When a 2′-hydroxyl group was attached to a ribose in the C3′-endo conformation, there was an unfavorable steric clash between the hydroxyl and the backbone segment of Y505-G508, especially the carbonyl group of Y505 (1.88 Å). If this model is correct in solution, then Pol λ employs an effective mechanism of sugar selection, whereby the backbone segment of Y505-G508 and the side chain of Y505 respectively play a major and minor role in steric exclusion of the ribose 2′- hydroxyl group. When comparing the kinetic data for the incorporation of UTP and dTTP catalyzed by Y505A (Table 4.3), the backbone segment contributed to the nucleotide incorporation rate and affinity by 52- and 7-fold, respectively. Thus, steric exclusion by
89 the backbone segment of Y505-G508 affected both the kp and Kd. In contrast, comparison of the kinetic data for UTP incorporation catalyzed by wild-type Pol λ and the Y505A mutant indicated that the side chain of Y505 led to a 16-fold tighter ribonucleotide binding affinity (Table 4.3). Thus, the mobility of the side chain of Y505 is more dynamic than what has been extrapolated from the superimposed binary and ternary crystal structures of truncated Pol λ [55], especially when the side chain of Y505 is predicted to be >5 Å from the 2′-hydroxyl (Figure 4.5). Together, our kinetic and modeling studies indicated that Y505 of Pol λ dynamically interacts with an incoming nucleotide during catalysis.
Steric hindrance confers high sugar selectivity Since Pol λ utilized a novel sugar selection mechanism, we were interested in understanding how various factors (e.g. size and stereochemistry) may influence the sugar selectivity of Pol λ. These factors were examined using rCTP analogs (Figure 4.4) containing different chemical moieties at ribose 2′-positions. With the exception of rCTP,
2′-NH2-CTP, and 2′-OCH3-CTP, Pol λ bound tightly to these substrates, like all dNTPs, despite the altered ribose structure. The size of the 2′-atom(s) was correlated with substrate specificity, for polymerization catalyzed by Pol λ was more efficient when the substituent at the 2′-position was small (Table 4.4). Based on structural studies and our model (Figure 4.5), the sugar selectivity of Pol λ remained low for 2′-F-CTP and GemCTP because the small hydrogen and fluorine atoms did not clash with the Y505- G508 backbone segment nor the Y505 side chain (Figure 4.1 part (B)). Moreover, altering the stereochemistry of the 2′-hydroxyl group dramatically relaxed the sugar selectivity of Pol λ. This result supported our model which showed that there is space to accommodate the 2′-hydroxyl directed above the ribose. Additionally, a similar kinetic finding has also been observed for RB69 DNA polymerase with araCTP. Surprisingly, an amine group, which is slightly larger than the hydroxyl, led to a higher sugar selectivity value due to the ~190-fold slower rate of incorporation and 230-fold weaker binding affinity (Table 4.4). Collectively, these different rCTP analogs showed how the steric bulkiness of the ribose 2′-substituent influenced the degree of sugar selection by Pol λ.
90 In summary, Pol λ can incorporate ribonucleotides, albeit with greatly reduced catalytic efficiency compared to dNTPs. More importantly, this work demonstrated that Pol λ utilized primarily a backbone segment, especially the carbonyl group of Y505, and secondarily the bulky and dynamic side chain of Y505 to discriminate between dNTPs and rNTPs unlike other DNA polymerases and RTs examined to date.
91 4.5 Tables
Table 4.1 Kinetic parameters of matched rNTP incorporation into single-nucleotide gapped DNA catalyzed by Pol λ at 37 °C. a,b rNTP DNA kp Kd (µM) kp/Kd Fidelity Sugar template (s-1) (µM-1s-1) selectivityb,c base UTP dA (D-1) 0.053 ± 0.001 114 ± 8 4.6 × 10-4 3.1 × 10-4 3,000 rCTP dG (D-6) 0.020 ± 0.001 52 ± 8 3.8 × 10-4 2.1 × 10-4 4,000 rATP dT (D-7) 0.0045 ± 140 ± 20 3.2 × 10-5 1.9 × 10-5 50,000 0.0002 rGTP dC (D-8) 0.00311 ± 36 ± 3 8.6 × 10-5 1.6 × 10-5 14,000 0.00007 a Calculated as (kp/Kd)matched rNTP/[(kp/Kd)correct dNTP + (kp/Kd)matched rNTP]. b Kinetic parameters of (kp/Kd)correct dNTP are from reference [117]. c Calculated as (kp/Kd)correct dNTP/(kp/Kd)matched rNTP.
92
Table 4.2 Kinetic parameters of mismatched rNTP incorporation into single- nucleotide gapped D-1 DNA catalyzed by Pol λ at 37 °C. a,b rNTP kp Kd kp/Kd Fidelity Sugar (s-1) (µM) (µM-1s-1) Selectivityb,c dTTPb 3.9 ± 0.2 2.6 ± 0.4 1.5 UTP 0.026 ± 0.002 110 ± 20 2.4 × 10-4 1.6 × 10-4 6,000 rCTP 0.0036 ± 0.0006 309 ± 132 1.2 × 10-5 8.0 × 10-6 125,000 rATP No observed incorporation rGTP 0.0061 ± 0.0007 313 ± 86 1.9 × 10-5 1.3 × 10-5 79,000 a Calculated as (kp/Kd)rNTP/[(kp/Kd)dTTP + (kp/Kd)rNTP]. bKinetic parameters for dTTP are from reference [117]. c Calculated as (kp/Kd)dTTP/(kp/Kd)rNTP.
93
Table 4.3 Kinetic parameters of dTTP or UTP incorporation into single-nucleotide gapped D-1 DNA catalyzed by Pol λ at 37 °C.
Pol λ Nucleotide kp Kd kp/Kd Sugar mutants (s-1) (µM) (µM-1s-1) selectivitya Wild type dTTPb 3.9 ± 0.2 2.6 ± 0.4 1.5 UTP 0.053 ± 0.001 114 ± 8 4.6 × 10-4 3,000 Y505G dTTP 0.85 ± 0.02 0.81 ± 0.09 1.0 UTP 0.067 ± 0.005 47 ± 11 1.4 × 10-3 700 Y505A dTTP 1.30 ± 0.03 1.0 ± 0.1 1.3 UTP 0.0251 ± 0.0006 7.0 ± 0.7 1.4 × 10-3 360 F506A dTTP 7.1 ± 0.2 2.8 ± 0.4 2.5 UTP 0.183 ± 0.007 218 ± 28 8.4 × 10-4 3,000 a Calculated as (kp/Kd)dTTP/(kp/Kd)UTP. bKinetic parameters were obtained from reference [117].
94
Table 4.4 Kinetic parameters of rCTP analogs incorporated into single-nucleotide gapped D-6 DNA catalyzed by Pol λ at 37 °C. rCTP analog kp Kd kp/Kd Sugar (s-1) (µM) (µM-1s-1) selectivitya dCTPb 1.57 ± 0.04 0.9 ± 0.1 1.7 rCTP 0.020 ± 0.001 52 ± 8 3.8 × 10-4 4,000 araCTP 1.28 ± 0.04 1.8 ± 0.2 0.71 2.4 2′-F-CTP 0.83 ± 0.02 3.4 ± 0.3 0.24 6.3 GemCTP 0.30 ± 0.02 4 ± 1 0.075 23 -5 2′-NH2-CTP 0.0084 ± 0.0003 206 ± 22 4.1 × 10 37,000 -6 2′-OCH3-CTP (1.6 ± 0.1) × 10 1,000,000 a Calculated as (kp/Kd)dCTP/(kp/Kd)CTP analog. bKinetic parameters were obtained from reference [117].
95 4.6 Figures
A
B
Figure 4.1 Sequence alignment of human X-family DNA polymerases and the active site of Pol λ. (A) Sequence alignment of the α-helix M in the thumb domain of each human X- family DNA polymerase. Residue numbers correspond to positions with respect to the N- terminal methionines. The black background indicates which residues are conserved among the family members and the arrow head denotes the potential “steric gate” residue. (B) A close-up view of potential steric interactions with the C2′ position of an incoming ddTTP are depicted in the active site of Pol λ (PDB 1XSN). Distances are listed in the table for the Y505 side chain and peptide backbone residues Y505-G508 (purple) near the C2′ position.
96 A
B C
30 0.05
25 0.04
20
) 0.03 -1
15 (s obs
k 0.02
Product (nM) Product 10
0.01 5
0 0 0 100200300400500600 0 200 400 600 800 1000 Time (s) UTP (μM)
Figure 4.2 Concentration dependence on the pre-steady state rate constant of matched ribonucleotide incorporation. (A) Each D-DNA substrate is composed of a 5′- radiolabeled 21-mer, a 5′-phosphorylated 19-mer, and a 41-mer template where ‘X’ represents A for D-1, C for D-8, G for D-6, and T for D-7. (B) A pre-incubated solution of Pol λ (120 nM) and 5′-[32P]-labeled D-1 (30 nM) was rapidly mixed with increasing concentrations of UTP•Mg2+ (5 μM, z; 10 μM, {; 25 μM, ; 50 μM, ; 100 μM, S; 200 μM, U; 400 μM, ; and 800 μM, ) for various time intervals. The solid lines are the best fits to a single-exponential equation (Equation 1 in Section 4.2) which determined the observed rate constants, kobs. (C) The kobs values were plotted as a function of UTP concentration. The data (z) were then fit to a hyperbolic equation -1 (Equation 2 in Section 4.2), yielding a kp of 0.053 ± 0.001 s and a Kd of 114 ± 8 μM.
97 2000 )
-1 1000
dmol 0 2
-1000
-2000
-3000
-4000
-5000 Molar Ellipticity (deg cm -6000 200 210 220 230 240 Wavelength (nm)
Figure 4.3 CD spectra of wild-type Pol λ and its mutants. The CD spectra were collected at 37 °C and overlaid with wild-type Pol λ and the variants Y505A (green), F506A (blue), and Y505d-T507d (red). The CD signal indicated that each protein was folded. Please note, the CD spectra for Y505G were not examined due to its low purification yield. However, based upon the similar enzymatic activities (Table 4.3), the secondary structure of Y505G was expected to mirror the spectra obtained for Y505A.
98
Figure 4.4 Chemical structures of CTP analogs. The chemical structures are shown for all CTP analogs used in this work. Chemical variations at the 2′-position are highlighted in red and abbreviations are defined in the text.
99
Figure 4.5 Model of a ribonucleotide bound at the active site of Pol λ. Using the ternary complex Pol λ•gapped DNA•ddTTP (PDB 1XSN), the 2′- and 3′-hydroxyl groups were modeled onto the ribose of the ddTTP substrate. The hydroxyl groups were extracted from the ATP substrate in complex with a reverse transcriptase (PDB 2IAJ). The model presented here shows a steric clash between the 2′-hydroxyl of a dTTP analog (rTTP) and the backbone carbonyl group of Y505.
100 Chapter 5: Identification of Critical Residues for the Tight Binding of Both Correct and Incorrect Nucleotides to Human DNA Polymerase λ
5.1 Introduction
DNA polymerases, which are organized into the A, B, C, D, X, or Y families [82, 83], have been shown to share a minimal kinetic mechanism for DNA polymerization [54]. This minimal mechanism indicates that DNA polymerases discriminate between a correct or incorrect nucleotide (dNTP) substrate predominantly during two steps: dNTP binding affinity to the enzyme•DNA complex and the rate of dNTP incorporation. Interestingly, DNA polymerases exhibit a wide range of binding affinity values for correct and incorrect dNTPs, thereby suggesting that these enzymes employ different discriminatory mechanisms. Using transient state kinetic methods, the equilibrium dissociation constants
(Kd) of an incoming nucleotide have been determined for replicative [88-92, 146], repair [114, 116, 117], and lesion bypass DNA polymerases [68, 105, 115, 186-190]. For example, the discrimination factor for nucleotide binding, defined as the ratio of the equilibrium dissociation constants for incorrect and correct nucleotides
(Kd,incorrect/Kd,correct), is 250 on average for an exonuclease-deficient mutant of the replicative human mitochondrial DNA polymerase γ [90, 91], 110 on average for the rat DNA repair polymerase β (rPol β) [114], and 4 for the human DNA lesion bypass polymerase η [186].
Human DNA polymerase λ (Pol λ), an X-family member which shares 34% sequence identity with Pol β [74], has been postulated to play a role in base excision repair (BER), non-homologous end joining, and V(D)J recombination while the participation of Pol β in BER has been confirmed [78, 172, 174-177, 191]. Intriguingly, Pol λ exhibits tight
101 nucleotide binding affinity for both correct and incorrect dNTPs in which the discrimination factor is 3 on average for nucleotide incorporation into a single-nucleotide gap DNA substrate [117]. Despite the varying discrimination factors (110 versus 3) between these two similar DNA repair enzymes, the base substitution fidelity of rPol β and human Pol λ is within the same range: 10-4 to 10-5 [114, 117].
Based on the crystal structure of Pol β, the tight binding of correct nucleotides by Pol β in the presence of single-nucleotide gap DNA is predicted to be partly due to the contribution of its 5′-deoxyribose-5-phosphate lyase (dRPase) domain which interacts strongly with the downstream primer and the polymerase domain [119]. This reasoning does not apply to Pol λ because all nucleotides have similarly high affinity to Pol λ, even with a non-gapped DNA substrate [192]. Additionally, the N-terminal BRCT and Proline-rich domains of Pol λ are not responsible because the C-terminal Pol β-like domain binds to all nucleotides as tightly as Pol λ [117, 182]. To better understand the kinetic basis for the tight binding of all nucleotides, we have created site-specific mutants of Pol λ that are designed to target key interactions between the polymerase and its substrates (Figure 5.1). Furthermore, we investigated the kinetic contributions from the intrinsic properties of a nucleotide, such as hydrogen bonding, base stacking, and steric bulkiness, using three non-natural nucleotide analogs (Figure 5.2): 1-naphthalene 5′- triphosphate (1-dNaTP), pyrene 5′-triphosphate (dPTP), and 5-nitroindole 5′-triphosphate (5-dNITP).
5.2 Materials and Methods
Materials 32 These chemicals were purchased from the following companies: [γ- P]ATP, MP Biomedicals; deoxyribonucleotides-5′-triphosphates (dNTP), GE Healthcare; 5- nitroindole 5′-triphosphate, TriLink Biotechnologies; Biospin columns, Bio-Rad Laboratories; OptiKinase™, USB Corporation; Quikchange XL site-directed mutagenesis
102 kit, Stratagene; synthetic oligodeoxyribonucleotides, Integrated DNA Technologies. The mutagenesis, expression, and purification steps of full-length wild-type Pol λ (1-575) and the full-length mutants were performed as described previously [117, 193]. CD spectroscopic studies were performed for the Pol λ mutants as described previously [193]. Pyrene 5′-triphosphate was synthesized as described previously [194]. 1- naphthalene 5′-triphosphate was synthesized according to the literature [195] except that the epimerization followed a different procedure [196] as did the formation of the triphosphate [197].
DNA binding assay DNA The equilibrium dissociation constant (Kd ) of the Pol λ•DNA binary complex was measured using a fluorescence titration assay. Increasing concentrations of Pol λ (10 - 2000 nM) were titrated into a fixed concentration of F-DNA (100 nM) in buffer L (50 mM Tris-Cl pH 8.4 at 37 °C, 5 mM MgCl2, 100 mM NaCl, 0.1 mM EDTA, 5 mM DTT, and 10% glycerol). The F-DNA substrate (Table 5.1) was excited at a wavelength of 312 nm with emission and excitation slit widths of 5 nm. The emission spectra were collected at 1 nm intervals from 320 to 500 nm using a Fluoromax-4 (Jobin Jvon Horiba). Emission background from the buffer and intrinsic protein fluorescence were subtracted from each spectrum. A modified form of the quadratic equation (Equation 1) DNA DNA 2 1/2 [F] = Fmax + [(Fmin – Fmax)/(2D0)] {(Kd + E0 + D0) – [(Kd + E0 + D0) – 4E0D0] } (1) was applied to a plot of the fluorescence intensity (F) measured at 370 nm versus enzyme concentration. Fmax and Fmin represent the maximum and minimum fluorescence intensity, respectively.
Measurement of the kp and Kd for single-nucleotide incorporation Kinetic assays were completed using optimized buffer L (50 mM Tris-Cl pH 8.4 at 37 °C,
5 mM MgCl2, 100 mM NaCl, 0.1 mM EDTA, 5 mM DTT, 10% glycerol, and 0.1 mg/ml BSA) as previously described [182]. The purification, 5′-[32P] radiolabeling, and annealing of single-nucleotide gapped D-DNA substrates (Table 5.1) was performed as described previously [117, 182]. All kinetic experiments described herein were 103 performed at 37 °C and the reported concentrations are final after mixing all the components. A pre-incubated solution containing wild-type Pol λ (120 nM) or a Pol λ mutant (300 nM) and a single-nucleotide gap DNA substrate (30 nM) was mixed with increasing concentrations of nucleotide (0.25-1500 μM) in buffer L at 37 °C. Aliquots of the reaction mixtures were quenched at various times using 0.37 M EDTA. A rapid chemical-quench flow apparatus (KinTek) was utilized for fast nucleotide incorporations. Reaction products were resolved using sequencing gel electrophoresis (17% acrylamide, 8 M urea) and quantitated with a Typhoon TRIO (GE Healthcare) and ImageQuant software (Molecular Dynamics). The time course of product formation at each nucleotide concentration was fit to a single-exponential equation (Equation 2)
[Product] = A[1 – exp(–kobst)] (2) using a nonlinear regression program KaleidaGraph (Synergy Software) to yield an observed rate constant of nucleotide incorporation (kobs). The kobs values were then plotted as a function of nucleotide concentration and fit using the hyperbolic equation (Equation 3)
kobs = kp[dNTP]/{[dNTP] + Kd} (3)
which resolved the kp and Kd values for nucleotide incorporation catalyzed by wild-type Pol λ or a mutant.
5.3 Results
The kinetic basis of Pol λ’s tight nucleotide binding was examined by creating several single-point mutants which disrupt specific interactions between each of the amino acid residues and the incoming dNTP or the template base. Based on an X-ray crystal structure of human Pol λ’s C-terminal Pol β-like domain in complex with gap DNA and a correct incoming ddTTP [55], the following amino acid residues were examined: R386, R420, K422, Y505, F506, A510, R514, and R517 (Figure 5.1). These amino acids are in close proximity to the triphosphate moiety (R386, R420, and K422), sugar moiety (Y505 and F506), or base of an incoming dNTP (A510) and the DNA template (R514, and
104 R517) (Figure 5.1). Most of these residues are conserved in both Pol λ and Pol β. However, K422, A510, and R514 in Pol λ are A185, D276, and K280 in Pol β, respectively. Following the successful completion of site-directed mutagenesis and purification of the Pol λ mutants, we examined whether the structure was altered due to the mutation(s). Therefore, we employed circular dichroism (CD) spectroscopy (Section 5.2). These data showed that all of the tested mutants had secondary structure similar to wild-type (WT) Pol λ which suggested that the mutations did not induce major structural changes (Figure 5.3). Pol λ R386E, R420A, and Y505G were not examined using CD spectroscopy due to their low purification yields.
Measurement of DNA binding affinity Next, we determined whether the mutants were able to form a binary complex (Pol λ•DNA) by using a fluorescence titration assay (Section 5.2) to measure the equilibrium DNA dissociation constant (Kd ). The F-DNA substrate contains 2-aminopurine, an analog of adenine, at the template base position (Table 5.1). Increasing concentrations of Pol λ were titrated into F-DNA, and then a plot of the corrected 2-aminopurine intensity and DNA concentration of Pol λ were fit to Equation 1 in Section 5.2 which resolved a Kd of 110 ± 20 nM (Figure 5.4 and Table 5.2). Similar binding constants (70 – 300 nM) were measured for the Pol λ mutants, indicating that these enzymes have a folded structure that can recognize and bind a single-nucleotide gap DNA substrate similar to WT Pol λ.
Measurement of nucleotide binding and incorporation rate catalyzed by Pol λ mutants The single-point Pol λ mutants were kinetically characterized by measuring the maximum rate of nucleotide incorporation (kp) and the equilibrium dissociation constant
(Kd) of an incoming nucleotide under single-turnover conditions. Single-turnover conditions prevent any complications from the steady-state phase that may originate from the rate of DNA dissociation being comparable to the rate of nucleotide incorporation for Pol λ [63, 117, 182]. Recent studies with T7 DNA polymerase, an A-family enzyme, reveal that nucleotide binding to a polymerase complex with DNA induces several conformational changes preceding the incorporation step which may mask the measured
105 Kd value under single-turnover reaction conditions [112]. Since similar conformational changes have not been detected with Pol λ during nucleotide binding and incorporation [55, 198] and the kinetic mechanisms of Pol λ with gap DNA and T7 DNA polymerase
with non-gap DNA may be different [54, 56], the Kd values measured in this paper likely reflect the true nucleotide binding affinity. To perform the single-turnover kinetic assay (Section 5.2), a pre-incubated solution of Pol λ A510E and 5′-[32P]-labeled D-1 DNA (Table 5.1) was reacted with increasing concentrations of mismatched dGTP (1-160 µM). After quantitating DNA product formation at each nucleotide concentration (Figure 5.5), -1 the pre-steady state kinetic parameters were determined: a kp of 0.0102 ± 0.0003 s and a
Kd of 15 ± 2 μM for Pol λ A510E. The pre-steady state kinetic parameters for all Pol λ mutants incorporating incorrect dGTP (Table 5.3) or correct dTTP (Table 5.4) into single-nucleotide gap D-1 DNA (Table 5.1) were determined using this single-turnover
kinetic methodology. The kp and/or Kd values were then used to calculate the substrate
specificity constant (kp/Kd), Kd ratio, and fidelity as defined in Tables 5.3 and 5.4. In this study, fidelity describes the probability of Pol λ making a base substitution error for the dGTP:dA mismatch. Our previous work determined the kinetic parameters for WT Pol λ [117].
Pol λ mutants targeting interactions near the triphosphate moiety of an incoming dNTP Pol λ single-point mutants of residues R386, R420, and K422 demonstrated a kinetically
significant (i.e. >2-fold relative to WT) weaker binding affinity (1/Kd) for both correct dTTP and incorrect dGTP incorporation opposite a template base dA (Tables 5.3 and
5.4). The Kd value increased dramatically (250- and 270-fold) for Pol λ R420A, a residue situated near the β-phosphate of an incoming dNTP (Figure 5.1). In contrast, the neutrally-charged R386A mutant, which is located near the γ-phosphate of an incoming dNTP, had a more mild effect, as the binding was weakened by a modest 6- and 16-fold for a correct and incorrect dNTP, respectively. K422 is another positively-charged residue in the vicinity of the triphosphate moiety, although, the alanine substitution mutant revealed a mild, 3-fold weaker binding affinity for both dTTP and dGTP. Interestingly, the fidelity of Pol λ R420A dropped by almost two orders of magnitude
106 (Table 5.3 and Figure 5.6). This effect was mostly due to a large decrease in the rate for dTTP incorporation (330-fold) while the rate for dGTP misincorporation (~4-fold) remained similar to WT Pol λ.
Pol λ mutants with altered interactions near the ribose of an incoming dNTP For a misincorporation, the binding affinity of dGTP was weakened by 19-, 9-, and 6-fold for Y505G, Y505A, and F506A, respectively (Table 5.3). As a result, these mutants displayed a consistent increase in enzyme fidelity (4- to 13-fold) among the mutant enzymes examined in this work (Table 5.3 and Figure 5.6).
Pol λ mutants which disrupt interactions with bases of an incoming dNTP or DNA template Replacing A510 with a glutamate resulted in modest 3- to 5-fold perturbations for the
dTTP and dGTP Kd values (Tables 5.3 and 5.4). Additionally, the R517A mutant exhibited 2- to 3-fold tighter binding affinities for both dTTP and dGTP relative to WT
Pol λ. However, the dTTP and dGTP Kd values for R514A were weakened by 2- and 25- fold, respectively. The fidelity of A510E dropped by approximately 17-fold, and this was caused by a 26-fold rate increase for a misincorporation (Table 5.3). In contrast, the fidelity of R517A dropped by three orders of magnitude which was due to a 250-fold rate decrease for a correct incorporation.
Kinetic characterization of double-point Pol λ mutants Our above results suggested that the tight dNTP binding by Pol λ is a function of multiple amino acid residues. To examine if these residues behave individually or cooperatively, we created two double mutants: R386A/A510E and R386A/R514A. These three residues
were selected because the Kd ratios for correct and incorrect dNTPs were modest so that,
if binding was weaker for the double mutant, we could accurately measure the Kd. After determining the kp and Kd values under single-turnover conditions, the correct and
incorrect Kd ratios for the double mutants ranged from 27- to 100-fold relative to WT Pol λ (Tables 5.3 and 5.4). Interestingly, the sum of the binding affinities for the single-
107 substitution mutants did not equal the binding affinities for the double-substitution mutants. Thus, these dNTP binding data indicated that nucleotide binding was synergistic or cooperative for these sites tested. Similar to A510E, the fidelity for R386A/A510E dropped by 26-fold. This loss of fidelity was mostly due to a 17-fold increase in the rate of dGTP misincorporation. In contrast, the fidelity for R386A/R514A was similar to WT Pol λ but slightly higher than the single-point mutant R386A and lower for R514A (Table 5.3 and Figure 5.6), suggesting that the effects of these two point mutations were either offset or compensated by repositioning other active site residues.
Effects of hydrogen bonding, base stacking, and steric interactions during DNA synthesis Nucleotide binding and selection may be influenced by the properties of the substrates, such as Watson-Crick hydrogen bonding, base stacking, steric interactions, and hydrogen bonds between the enzyme and DNA minor groove. To discern the role of these factors, we have determined the pre-steady state kinetic parameters (Table 5.5) for the incorporation of three non-natural nucleotide analogs (Figure 5.2) into D-7 DNA, a single-nucleotide gap DNA substrate with dT as the template (Table 5.1). This allows us to compare the kinetic parameters for the analogs with dATP, the nucleotide with the strongest base-stacking energy among the four natural bases [199]. The three non-natural nucleotide analogs (dPTP, 5-dNITP, and 1-dNaTP) lack the ability to form hydrogen bonds with a template base, possess stronger base stacking energy, and increased steric bulkiness compared to a natural nucleotide [199]. Nonetheless, WT Pol λ inserted dPTP, 5-dNITP, and 1-dNaTP, although, the incorporation efficiency was reduced significantly, mostly due to a slower rate of incorporation rather than nucleotide binding. Compared to dATP, the nucleotide binding was weakened by 24-, 12-, and 2- fold for dPTP, 1-dNaTP, and 5-dNITP, respectively. Overall, these results suggested that (i) Watson-Crick hydrogen bonding between the incoming nucleotide and template base is not essential for nucleotides with strong base-stacking energy but the hydrogen bonds likely enhance incorporation efficiency, (ii) the ground-state binding affinity of a nucleotide depends moderately on the formation of Watson-Crick hydrogen bonds, and
108 (iii) the active site of WT Pol λ can accommodate an oversized base, although, the bulky
pyrene moiety likely leads to a higher Kd.
5.4 Discussion
Tight nucleotide binding depends partially on the properties of nascent base pair We have previously demonstrated that all dNTPs, correct and incorrect, have tight and peculiarly similar binding affinities to Pol λ and single-nucleotide gap DNA [117]. The biological relevance of tight nucleotide binding affinity likely facilitates efficient DNA repair synthesis by Pol λ, such as non-homologous end joining, when cellular dNTP pools are low outside of S phase [182, 200]. The tight binding of all dNTPs is rare because most kinetically characterized DNA polymerases generally discriminate against incorrect nucleotides via weak binding and slow incorporation [201]. For example, incorrect dNTPs have 35- to 340-fold weaker binding affinity than correct dNTPs to the binary complex of Pol β and single-nucleotide gap DNA [114]. To identify the major factors contributing to Pol λ’s tight nucleotide binding, we examined the physical properties of nascent base pairs including hydrogen bonding, base stacking, and base size. Previously, the importance of those factors for other DNA polymerases has been probed using non-natural nucleobases (e.g. difluorotoluene and pyrene) substituted in an incoming dNTP or a DNA template [202]. The A- and Y-family DNA polymerases incorporate non-hydrogen bonding nucleotide analogs, albeit less efficiently than a correct, natural dNTP based on pre-steady state kinetic results [203-205]. These findings suggested that Watson-Crick hydrogen bonds formed between a dNTP and a template base are not essential for nucleotide incorporation by the A- and Y-family DNA polymerases [64, 65, 202]. Our kinetic data in Table 5.5 for the three nucleotide analogs (5-dNITP, dPTP, and 1-dNaTP) suggested that human Pol λ followed the same trend. Moreover, the moderately weaker binding affinities of dPTP, 5-dNITP, and 1-dNaTP suggested that Watson-Crick hydrogen bonds between canonical base pairs were not important for the tight binding of a nucleotide to Pol λ•DNA, but formation of these
109 hydrogen bonds greatly accelerated the catalytic rate of Pol λ. Although base stacking contributed to the 7-fold tighter binding of 5-dNITP over 1-dNaTP, it is not the major factor to dictate nucleotide binding because correct and incorrect dNTPs exhibit tight binding affinities regardless of the base identity, e.g. purine versus pyrimidine [117].
Active site residues coordinate unprecedentedly tight nucleotide binding. The X-ray crystal structure of truncated human Pol λ [55] was used to rationally select residues to mutate and to determine the individual roles of eight active site residues (R386, R420, K422, Y505, F506, A510, R514, and R517) which interact with an incoming nucleotide or the template base (Figure 5.1). The common structural moieties between correct and incorrect nucleotides are triphosphate, ribose, and nucleobase. The Pol λ residues interact with specific structural moieties of a dNTP, therefore, the kinetic significance of binding
each dNTP moiety can be dissected by averaging the Kd values for both correct and incorrect dNTPs using the mutants (Tables 5.3 and 5.4) that interact with the triphosphate (R386A, R420A, and K422A), ribose (Y505A and F506A), or nucleobase (A510E). The impact on dNTP binding generated the following ranking for each of the common structural moieties: triphosphate (Kd,average = 265 μM) >> nitrogenous base (Kd,average = 18
μM) > ribose (Kd,average = 13 μM). Stabilizing the triphosphate group of an incoming nucleotide by R386, R420, and K422 was critical for tight nucleotide binding. For example, dramatic changes in nucleotide binding were observed when R386A or R386E disrupted the salt bridge between the guanidino group of R386 and the γ-phosphate of an incoming nucleotide [55, 198]. Similarly, the electrostatic interactions between R420 and the β-phosphate of an incoming nucleotide maintain a low Kd. In comparison, the alanine mutations of Y505 and F506, which interact with the ribose of an incoming nucleotide [55], enhanced the ability of Pol λ to discriminate between correct and incorrect dNTPs
by increasing the Kd value of an incorrect nucleotide by 6- to 9-fold (Tables 3 and 4). A510 of Pol λ is one of the few active site residues not conserved in Pol β. Previously, we [182] and others [200, 206] have proposed that Pol λ’s tight dNTP binding affinity may be due to a neutral residue A510 which is D276 in Pol β. Interestingly, the A510E substitution weakened the dNTP binding affinity of Pol λ by 5- to 8-fold (Tables 5.3 and
110 5.4) while mutating D276 to a neutral residue (e.g. valine and glycine) increases the correct nucleotide binding affinity of Pol β by 4- to 9-fold and incorrect nucleotide binding affinity by 3-fold [201, 207]. Thus, a bulky, negatively-charged residue weakens the nucleotide binding affinity for both Pol λ A510E and Pol β. R514, which stacks against the template base dA, is another residue that enhanced the binding discrimination factor, for R514A weakened the binding of correct and incorrect nucleotides by 2- and 25-fold, respectively (Tables 5.3 and 5.4).
Several of the active site residues (R386, R420, K422, Y505, F506, A510, and R514) in Pol λ interact with a nascent base pair to cooperatively stabilize nucleotide binding in the ground state. Since the Kd values for the double-point mutants (R386A/A510E and
R386A/R514A) were greater than the sum of the Kd values for the single-point mutants by 2- to 5-fold (Tables 5.3 and 5.4), these residues contribute to nucleotide binding in a synergistic manner. However, certain residues, e.g. R420, play a more important role in nucleotide binding than others. It is possible that other active site residues, which we did not examine here, may also participate in nucleotide binding. Importantly, the tight nucleotide binding affinity of Pol λ was not manifested in a single amino acid residue, indicating the mechanism of how Pol λ and Pol β discriminate at the ground-state nucleotide binding level is more complex. The different conformational dynamics of these enzymes preceding catalysis may play a role. For example, Pol β undergoes a conformational change during the binary (Pol β•DNA) to ternary (Pol β•DNA•dNTP) complex transition where as Pol λ only repositions selected active site residues [55, 58, 119, 198, 208]. Furthermore, the conformational change observed for correct and incorrect nucleotide incorporations is different for Pol β [119, 209]. Thus, additional studies are needed to better understand the relationship between substrate recognition and polymerase conformational dynamics during the catalytic cycle.
Fidelity of Pol λ is maintained by minor groove interactions Among ten single-substitution Pol λ mutants, this work identified R517 as an important residue in maintaining polymerase fidelity. R517, which is R283 in Pol β, is involved in
111 minor groove interactions with the DNA template (Figure 5.1) [55]. The R517A mutation did not significantly alter nucleotide binding affinity but dramatically decreased the kp value of correct dTTP by 250-fold, leading to a 1,360-fold loss of polymerase fidelity (Tables 5.3 and 5.4). The Pol β R283A mutant followed a similar kinetic trend as Pol λ R517A, whereby the rate of a correct dTTP incorporation dropped by 310-fold for an overall 260-fold loss of fidelity [210]. Crystal structures have been solved for human Pol β R283A and truncated Pol λ R517A; both of these studies conclude that these arginine residues are essential for properly positioning the template strand through hydrogen bonding and/or van der Waal’s interactions [211, 212]. Thus, these interactions provide Pol β and Pol λ with greater polymerization efficiency for correct incorporations [117, 210]. In the absence of a large conformational change [55, 198], Pol λ relies, in part, on non-specific minor groove interactions to detect the proper nascent base pair geometry in a manner similar to that observed for A- and B-family DNA polymerases [121, 122, 155, 156].
112 5.5 Tables
Table 5.1 DNA substrates. F-DNAa 5’-CGCAGCCGTCCAACCAACTCAG GTCGATCCAATGCCGTCC-3’ 3’-GCGTCGGCAGGTTGGTTGAGTCFCAGCTAGGTTACGGCAGG-5’ D-DNAb 5’-CGCAGCCGTCCAACCAACTCA CGTCGATCCAATGCCGTCC-3’ 3’-GCGTCGGCAGGTTGGTTGAGTXGCAGCTAGGTTACGGCAGG-5’ aThe ‘F’ in the 41mer template represents 2-aminopurine. The downstream 18-mer is 5′-phosphorylated. bThe ‘X’ in the 41mer template represents dA for D-1 DNA and dT for D-7 DNA. The downstream 19-mer is 5′-phosphorylated.
113
Table 5.2 Equilibrium dissociation constants of the Pol λ•DNA complex at 37 °C. DNA Enzyme Kd (nM) WT 110 ± 20 R386A 90 ± 20 R386E 300 ± 100 R420A 200 ± 100 K422A 150 ± 40 Y505G 150 ± 70 Y505A 150 ± 50 F506A 140 ± 40 A510E 120 ± 60 R514A 70 ± 10 R517A 80 ± 30 R386A/A510E 120 ± 40 R386A/R514A 80 ± 30
114
Table 5.3 Kinetic parameters for incorrect nucleotide incorporation (dGTP) into single- nucleotide gapped DNA (D-1) catalyzed by human Pol λ at 37 °C. Pol λ k K k /K K p d p d d Fidelityb mutant (s-1) (μM) (μM-1s-1) ratioa WTc (4.0 ± 0.2) × 10-4 3.2 ± 0.5 1.3 × 10-4 - 8.3 × 10-5 R386A (1.55 ± 0.06) × 10-3 50 ± 10 3.1 × 10-5 16 3.6 × 10-4 R386E No observed incorporation R420A (1.1 ± 0.2) × 10-4 800 ± 200 1.4 × 10-7 250 8.1 × 10-3 K422A (6.3 ± 0.3) × 10-4 8 ± 1 7.9 × 10-5 3 3.4 × 10-4 Y505G (1.23 ± 0.07) × 10-3 60 ± 10 2.1 × 10-5 19 2.0 × 10-5 Y505A (7.5 ± 0.2) × 10-4 30 ± 3 2.5 × 10-5 9 1.9 × 10-5 F506A (2.9 ± 0.3) × 10-4 19 ± 6 1.5 × 10-5 6 6.0 × 10-6 A510E (1.02 ± 0.03) × 10-2 15 ± 2 6.8 × 10-4 5 1.4 × 10-3 R514A (8.2 ± 0.8) × 10-4 80 ± 20 1.0 × 10-5 25 1.1 × 10-5 R517A (1.5 ± 0.1) × 10-3 1.0 ± 0.3 1.5 × 10-3 0.3 1.1 × 10-1 R386A/A510E (6.9 ± 0.6) × 10-3 170 ± 40 4.1 × 10-5 53 2.3 × 10-3 R386A/R514A (7 ± 1) × 10-4 320 ± 90 2.2 × 10-6 100 6.4 × 10-5 a Calculated as (Kd)Mutant/(Kd)WT. b Calculated as (kp/Kd)Incorrect/[(kp/Kd)Correct + (kp/Kd)Incorrect]. cThe kinetic parameters for WT Pol λ are from reference [117].
115 Table 5.4 Kinetic parameters for correct nucleotide incorporation (dTTP) into single-nucleotide gapped DNA (D-1) catalyzed by human Pol λ at 37 °C. Pol λ kp Kd kp/Kd Kd mutant (s-1) (μM) (μM-1s-1) ratioa WTb 3.9 ± 0.2 2.6 ± 0.4 1.5 - R386Ac 1.3 ± 0.1 15 ± 5 8.7 × 10-2 6 R386Ec 0.005 ± 0.001 1000 ± 410 5.0 × 10-6 380 R420A 0.0119 ± 0.0005 710 ± 60 1.7 × 10-5 270 K422A 1.62 ± 0.08 7 ± 1 2.3 × 10-1 3 Y505Gd 0.85 ± 0.02 0.81 ± 0.09 1.0 0.3 Y505Ad 1.30 ± 0.03 1.0 ± 0.1 1.3 0.4 F506Ad 7.1 ± 0.2 2.8 ± 0.4 2.5 1 A510E 10.2 ± 0.8 21 ± 4 4.9 × 10-1 8 R514A 4.0 ± 0.1 4.1 ± 0.7 9.8 × 10-1 2 R517A 0.0153 ± 0.0004 1.3 ± 0.1 1.2 × 10-2 0.5 R386A/A510E 3.0 ± 0.2 170 ± 20 1.8 × 10-2 65 R386A/R514A 2.4 ± 0.1 70 ± 10 3.4 × 10-2 27 a Calculated as (Kd)Mutant/(Kd)WT. bThe kinetic parameters for WT Pol λ are from reference [117]. cThe kinetic parameters for Pol λ R386A and R386E are from reference [198]. dThe kinetic parameters for Pol λ Y505G, Y505A, and F506A are from reference [193].
116
Table 5.5 Kinetic parameters for non-natural nucleotide analog incorporation into single-nucleotide gapped D-7 DNA catalyzed by WT human Pol λ at 37 °C. k K k /K K dNTP p d p d d (s-1)a (μM) (μM-1s-1) ratioa dATPb 1.5 ± 0.18 0.9 ± 0.3 1.7 dGTPb (1.0 ± 0.2) × 10-2 8.4 ± 0.6 1.4 × 10-3 8 dTTPb (2 ± 1) × 10-4 7 ± 4 1.4 × 10-3 8 dCTPb (1.0 ± 0.2) × 10-2 7 ± 4 1.4 × 10-3 8 dPTP (3.3 ± 0.2) × 10-4 22 ± 2 1.5 × 10-5 24 1-dNaTP (3.6 ± 0.3) × 10-4 11 ± 3 3.3 × 10-5 12 5-dNITP (1.31 ± 0.04) × 10-2 1.5 ± 0.2 8.7 × 10-3 2 a Calculated as (Kd)Analog/(Kd)dATP. bThe kinetic parameters for natural dNTPs are from reference [117].
117 5.6 Figures
Figure 5.1 Active site of truncated Pol λ. Residues selected for mutagenesis and kinetic characterization are shown as sticks: R386 (brown), R420 (yellow), K422 (orange), F506 (dark blue), Y505 (green), A510 (cyan), R514 (purple), and R517 (red) (PDB 1XSN). The incoming ddTTP and DNA template base dA are shown as gray sticks with the important atoms in color. The remainder of the protein and DNA substrate is a solid, light gray ribbon.
118 A B
C
Figure 5.2 Chemical structure of nucleotide analogs. (A) 1-dNaTP, (B) dPTP, and (C) 5- dNITP.
119 A
2000 )
-1 1000
0 dmol 2
-1000
-2000
-3000
-4000
-5000 Molar Ellipticity (deg cm -6000 200 210 220 230 240
Wavelength (nm)
B C 2000 2000 ) )
-1 1000 -1 1000
0 0 dmol dmol 2 2
-1000 -1000
-2000 -2000
-3000 -3000
-4000 -4000
-5000 -5000 Molar Ellipticity (deg cm Molar Ellipticity (deg cm -6000 -6000 200 210 220 230 240 200 210 220 230 240 Wavelength (nm) Wavelength (nm)
Continued
120 Figure 5.3: Continued
D E
2000 2000 ) )
-1 1000 -1 1000
0 0 dmol dmol 2 2
-1000 -1000
-2000 -2000
-3000 -3000
-4000 -4000
-5000 -5000 Molar Ellipticity (degcm Molar Ellipticity (degcm -6000 -6000 200 210 220 230 240 200 210 220 230 240 Wavelength (nm) Wavelength (nm)
Figure 5.3 Circular dichroism spectra for WT and mutant Pol λ. The CD spectra were collected and overlaid for WT (black), R386A (brown), K422A (orange), Y505A (green), F506A (blue), A510E (cyan), R514A (purple), R517A (red), R386A/A510E (pink), and R386A/R514A (gray). The CD spectra indicated that all the proteins were folded. Individual spectra overlaid with wild-type Pol λ are shown for (B) R386A and K422A; (C) Y505A and F506A; (D) A510E, R514A, and R517A; and (E) R386A/A510E and R386A/R514A. The CD spectra for WT, Y505A, and F506A Pol λ are from reference [193].
121 A B
1.2 x 105 1 x 105
5 1 x 10 4 8 x 10
8 x 104 6 x 104
6 x 104 4 x 104 4 x 104 Intensity (c.p.s.) Intensity (c.p.s.)
2 x 104 2 x 104
0 0 0 200 400 600 800 1000 1200 1400 1600 0 200 400 600 800 1000 1200 Pol λ (nM) R386E Pol λ (nM)
Figure 5.4 Equilibrium dissociation constant for the dissociation of the binary complex Pol λ•DNA. A plot of 2-aminopurine intensity versus the concentration of (A) WT Pol λ DNA or (B) R386E Pol λ was fit to equation 1 (Section 5.2) which resolved a Kd = 110 ± 20 DNA nM and Kd = 300 ± 100 nM for WT and the R386E mutant, respectively.
122 A 30
25
20
15
10 Product (nM) Product
5
0 0 500 1000 1500 2000 2500 Time (s)
B 0.01
0.008
) 0.006 -1 (s
obs 0.004 k
0.002
0 0 50 100 150 200 dGTP (μM)
Figure 5.5 Concentration dependence on the pre-steady state rate constant of nucleotide misincorporation. (A) A pre-incubated solution of Pol λ A510E (300 nM) and 5′-[32P]- labeled D-1 DNA (30 nM) was rapidly mixed with increasing concentrations of dGTP•Mg2+ (1 μM, z; 2 μM, {; 5 μM, ; 10 μM, ; 20 μM, S; 40 μM, U; 80 μM, ; 160 µM, ) for various time intervals. The solid lines are the best fits to a single- exponential (Equation 2 in Section 5.2) which determined the observed rate constants, kobs. (B) The kobs values were plotted as a function of dGTP concentrations. The data (z) were then fit to a hyperbolic equation (Equation 3 in Section 5.2), yielding a kp of 0.0102 -1 ± 0.0003 s and a Kd of 15 ± 2 μM.
123
Figure 5.6 Fidelity of mutant enzymes versus WT. The base substitution fidelity calculated in Table 5.3 is plotted for each enzyme. The dashed line coincides with the fidelity of WT Pol λ. Thus, mutants with a calculated fidelity value above the dashed line exhibit lower polymerase fidelity than WT Pol λ.
124 Chapter 6: Efficiency and Fidelity of Human DNA Polymerases λ and β during Gap-Filling DNA Synthesis
6.1 Introduction
Base excision repair (BER) is the major pathway to repair single-base lesions in mammalian cells [213]. BER can proceed through the subpathways of short-patch (SP) or long-patch (LP) which result in the replacement and resynthesis of 1 or 2-10 nucleotides, respectively. For both of these subpathways, the damaged DNA base is recognized and removed by a DNA glycosylase, the DNA backbone is cleaved by apurinic/apyrimidinic endonuclease 1, nucleotide(s) is(are) incorporated by a DNA polymerase, and eventually the nicked DNA is sealed by a DNA ligase. However, these subpathways have several distinct variations: (i) one branch of LP-BER depends on the proliferating cell nuclear antigen (PCNA) [214-217], (ii) DNA polymerases β [76, 77, 214, 216, 218-225], δ [217- 219, 226, 227], ε [217, 226], λ [75, 78, 172, 173, 228], ι [229-231], and θ [12, 15] are implicated in nuclear gap-filling and/or strand-displacement DNA synthesis, (iii) the 5′- deoxyribose-5-phosphate (dRP) moiety is removed by the dRP lyase activity of a DNA polymerase in SP-BER [222, 232], and (iv) the displaced DNA strand containing the dRP group is cleaved by flap endonuclease 1 in LP-BER [219, 226]. In regards to the DNA synthesis step of BER, it remains unclear which polymerase is specifically involved in repairing each type of damaged DNA base and the preferred BER route. Clearly, each DNA polymerase functioning in BER must be able to coordinate interactions with multiple DNA repair proteins and to catalyze DNA synthesis using DNA substrates that vary in structure (e.g. gap size or DNA flap).
125 Pol β, a X-family DNA polymerase, is known to play a central role in SP-BER in vivo [76, 77] and is a strong candidate for functioning in LP-BER, likely independent of PCNA [225]. Pol λ, a X-family member that shares 34% sequence identity with Pol β [74], is postulated to complement or to support the in vivo function of Pol β in BER. This proposal is based on several observations: Pol λ possesses two key enzymatic activities (dRPase and gap-filling polymerase) that are required for SP-BER [172], Pol λ can repair uracil in a DNA duplex when substituted for Pol β in an in vitro BER reconstitution system [172], Pol λ is active in mediating BER when Pol β is absent or neutralized in cell extract from mouse embryonic fibroblasts [78], and Pol λ is involved in repairing oxidative DNA damage [173, 228]. Unlike Pol β, Pol λ has two non-enzymatic domains at its N-terminus: a breast cancer susceptibility gene 1 C-terminal (BRCT) domain and a proline-rich domain (Figure 6.1). Interestingly, the proline-rich domain has been shown to suppress polymerase activity [233], to limit strand displacement synthesis [174], and to increase polymerase fidelity on a single-nucleotide gapped DNA substrate [117]. In addition, DNA structure affects the polymerization efficiency of Pol λ [192] and Pol β [114, 201, 234]. Both enzymes prefer a single-nucleotide gapped DNA substrate with a 5′-phosphate on the downstream strand over a non-gapped primer-template DNA substrate. Based on the ternary crystal structures of Pol β [119] and a truncated form of Pol λ [55] in complex with gapped DNA and an incoming nucleotide, this substrate preference is likely due to the favorable interactions between the 5′-phosphate of the downstream strand and the dRPase domain. During BER, the DNA gap size may vary from 1-10 nucleotides with 2-4 being the most common in LP-BER [214, 219, 226, 227]. An increase in gap size would likely disrupt protein interactions with the downstream strand. Therefore, the incorporation efficiency and fidelity of a gap-filling DNA polymerase may depend on gap size. To better understand the kinetic relationship between gap size and polymerase function in BER, we have employed single-turnover kinetic assays to determine the polymerization efficiency and fidelity of Pol λ and Pol β incorporating dNTPs into DNA substrates with various gap sizes. In addition, we discerned the roles of the BRCT and proline-rich domains on gap-filling DNA synthesis
126 catalyzed by Pol λ as well as the effects of the DNA template sequence on its nucleotide selection.
6.2 Materials and Methods
Materials Biochemicals and reagents were purchased from the following companies: [γ-32P]ATP, MP Biomedicals; deoxyribonucleotide-5′-triphosphates, GE Healthcare; Bio-Spin 6 columns, Bio-Rad Laboratories; OptiKinase™, USB Corporation; synthetic oligodeoxyribonucleotides 21-mer, 5′-phosphorylated 19-mers, and 41- to 50-mers, Integrated DNA Technologies.
Mutagenesis, expression and purification of DNA polymerases Deletions, expression, and purification of wild-type human Pol λ (1-575), dPol λ (132- 575), tPol λ (245-575), and human Pol β were described previously [117, 182, 235].
Single-nucleotide gapped DNA substrates Commercially synthesized oligomers in Table 6.1 were purified using denaturing polyacrylamide gel electrophoresis. The 21-mer primer was radiolabeled with [γ-32P]ATP and OptiKinase™ and the unreacted [γ-32P]ATP was removed using a Bio-Spin 6 column; both steps were completed according to each of the manufacturers’s protocol. The gapped DNA substrates were prepared by mixing the 5′-[32P]-radiolabeled 21-mer, the appropriate non-radiolabeled downstream strand 19-mer, and the appropriate template (41- to 50-mer) at a 1:1.25:1.15 molar ratio, respectively. The primer-template DNA substrate was prepared by mixing 5′-[32P]-radiolabeled 21-mer and the 41-mer template at a 1:1.15 molar ratio. Then, the annealing mixture was denatured at 95 °C for 6 minutes and slowly cooled to room temperature over several hours.
127 Measurement of the kp and Kd for single-nucleotide incorporation assay Kinetic assays were completed using optimized buffer L (50 mM Tris-Cl, pH 8.4 at 37
°C, 5 mM MgCl2, 100 mM NaCl, 0.1 mM EDTA, 5 mM DTT, 10% glycerol, and 0.1 mg/ml BSA) for Pol λ [182] and buffer B (50 mM Tris-Cl, pH 7.8 at 37 °C, 5 mM
MgCl2, 50 mM NaCl, 0.1 mM EDTA, 5 mM DTT, 10% glycerol, and 0.1 mg/ml BSA) for Pol β. All kinetic experiments described herein were performed at 37 °C and the reported concentrations were final after mixing all the components. A pre-incubated solution containing Pol λ (120 nM) or Pol β (300 nM) and a DNA substrate (30 nM) was mixed with increasing concentrations (0.2-1500 μM) of nucleotide in the appropriate buffer at 37 °C. Aliquots of the reaction mixtures were quenched at various times using 0.37 M EDTA. A rapid chemical-quench flow apparatus (KinTek) was utilized for fast nucleotide incorporations. Reaction products were resolved using sequencing gel electrophoresis (17% acrylamide, 8 M urea) and quantitated with a Typhoon TRIO (GE Healthcare). The time course of product formation at each nucleotide concentration was fit to a single-exponential equation (Equation 1)
[Product] = A[1 – exp(–kobst)] (1) using a nonlinear regression program, KaleidaGraph (Synergy Software), to yield an observed rate constant of nucleotide incorporation (kobs). The kobs values were then plotted as a function of nucleotide concentration and a hyperbolic equation (Equation 2)
kobs = kp[dNTP]/{[dNTP] + Kd} (2)
was applied to resolve the kp and apparent Kd values for nucleotide incorporation catalyzed by Pol λ or Pol β.
6.3 Results
Effect of gap size on the kinetics of Pol λ
The incorporation efficiency (kp/Kd) of Pol λ inserting a dNTP opposite template dC with various DNA substrates (Table 6.1) was measured under single-turnover conditions (Section 6.2). As a representative example, Pol λ catalyzed the incorporation of dGTP into a 21/41-mer DNA substrate, and a hyperbolic dependence on the observed rate 128 constant was observed with increasing concentrations of dGTP, thereby yielding a -1 maximum rate of nucleotide incorporation (kp) equal to 0.0353 ± 0.0008 s and an
apparent equilibrium dissociation constant (Kd) of the Pol λ•DNA•dGTP complex equal to 0.82 ± 0.08 μM (Figure 6.2 and Table 6.2). Similar assays were performed for correct and incorrect nucleotide incorporations into gapped and non-gapped DNA, and the kinetic parameters, incorporation efficiency, efficiency ratio relative to a gap size of 1, and base substitution fidelity are provided in Table 6.2. Compared to DNA with a gap size of 1, the absence of a downstream strand decreased the incorporation efficiency of a correct dGTP by 33-fold for Pol λ. This result suggested that there is a threshold on the DNA gap size for maintaining efficient catalysis. Therefore, to identify the maximum gap width for efficient gap-filling synthesis by Pol λ, the gap size was expanded by inserting nucleotides into the DNA template so that the lengths of the 21-mer primer and 19-mer strand were held constant, the template base was dC, and the first downstream base was dG (Table 6.1). By increasing the gap size at 1-nucleotide intervals, the maximum gap width was determined to be 9 nucleotides for a correct dNTP to be inserted relatively efficient by Pol λ, since the incorporation efficiency for a gap size of 10 was reduced by 38-fold, a level comparable to the non-gapped DNA substrate (Table 6.2 and Figure 6.3 part (A)). This kinetic effect between 1- and 10-nucleotide gapped DNA was due to a
100-fold drop in kp, since the Kd remained within 3-fold (Table 6.2).
In regards to a misincorporation, the efficiency of dCTP was increased by 7- to 41-fold for gap sizes 2 to 8 when compared to a gap size of 1, thereby leading to a similar drop in polymerase fidelity (Table 6.2 and Figures 6.3 part (A) and 6.4 part (A)). However, a different dependence on gap size was observed for incorrect dATP and dTTP. A kinetically significant increase in incorporation efficiency (5- to 12-fold) and decrease in -4 -3 fidelity (10 to 10 ) for Pol λ appeared with gap sizes of approximately 4 to 7, a narrower gap range than dCTP. The lower fidelity is due to a ~40-fold increase in the rate of incorrect dNTP incorporation which leads to a higher incorporation efficiency. Pol λ did not incorporate dATP and dTTP for gap sizes greater than 8 nucleotides. Overall, these results suggested that the gap-filling activity of Pol λ would be efficient for gap
129 widths of 9 nucleotides or less, although, Pol λ may be a poor candidate for LP-BER due to the higher probability of a misincorporation for gap sizes greater than 1 nucleotide. Therefore, we decided to examine the gap-filling activity of Pol β which has stronger evidence to support a biological role in LP-BER [225].
Effect of gap size on the kinetics of Pol β The kinetic parameters were measured for Pol β inserting nucleotides into a recessed primer-template DNA substrate or gapped DNA with the selected sizes of 1, 2, 5, 7, and 10 (Tables 6.1 and 6.3). The kinetic behavior of Pol β synthesizing DNA on the aforementioned substrates was different from Pol λ. For a correct incorporation, the catalytic efficiency of Pol β differed by a mere 3-fold for non-gapped versus single- nucleotide gapped DNA (Table 6.3 and Figure 6.3 part (B)). Surprisingly, a dramatic 160-fold reduction in the polymerase efficiency was determined with a gap size of 10 while a more modest 6-fold decrease occurred with gap sizes of 5 and 7. In contrast to
Pol λ, the lower incorporation efficiency for Pol β was due to a 90-fold increase in Kd
whereas the kp was within two-fold for gap sizes of 1- and 10-nucleotides (Table 6.3). Like a correct dGTP incorporation, Pol β misincorporated dCTP, dATP, and dTTP with a lower catalytic efficiency as the gap size increased to 10 and then improved for non- gapped DNA. This finding suggested a “rebound effect”, whereby Pol β is capable of more efficient nucleotide incorporation when the gap size is greater than 10 nucleotides. For polymerase fidelity, a moderate 3- and 6-fold drop was observed for dCTP when the gap size was 5 and 7, respectively (Table 6.3 and Figure 6.4 part (B)). In contrast to Pol λ, the fidelity of Pol β incorporating dATP and dTTP increased up to 20-fold for gap widths greater than 5 nucleotides. Thus, these kinetic data suggested that Pol β would be less efficient but more accurate than Pol λ in LP-BER. Since different structure-function relationships were established for two similar polymerases, it was possible that the non- enzymatic BRCT and proline-rich domains may be influencing the catalytic activity of Pol λ.
130 Effect of non-enzymatic domains on Pol λ’s gap-filling activity To discern the impact of the BRCT and proline-rich domains on the catalytic activity of Pol λ, we created two truncated Pol λ mutants: dPol λ which is missing the BRCT domain and tPol λ which is missing the BRCT and proline-rich domains so that it is “Pol β-like” (Figure 6.1). Single-nucleotide incorporation assays were performed for dPol λ (Table 6.4) and tPol λ (Table 6.5) with the following set of DNA substrates: primer-template and gapped DNA with sizes of 1, 2, 5, 7, and 10 (Table 6.1). Like Pol λ, correct dGTP incorporation catalyzed by both Pol λ mutants remained similar for gap sizes 1, 2, 5, and 7 but decreased with a gap size of 10 or when the downstream strand was absent (Figure 6.3 parts (C) and (D)). Interestingly, among the three Pol λ enzymes, there was a gradient for the reduced catalytic efficiency when comparing 1-nucleotide gapped DNA with non- gapped: Pol λ (33) > dPol λ (28) > tPol λ (6) (Tables 6.2, 6.4, and 6.5). Here, this gradient was closely related to the 76-, 28-, and 6-fold slower rate of dGTP incorporation for Pol λ, dPol λ, and tPol λ, respectively. Furthermore, both truncated Pol λ mutants exhibited increased incorporation efficiency (11- and 34-fold) of dCTP when the gap size was 2, thereby leading to a concomitant drop in polymerase fidelity (Figure 6.4 parts (C) and (D)). However, beyond a gap size of 2, dPol λ continued to show improved catalytic efficiency when the gap sizes were 5 and 7 while tPol λ did not. For the dATP and dTTP misincorporations, the catalytic efficiency of both enzymes gradually decreased as the gap size increased. In general, the fidelity of both mutants for dATP increased up to 10- fold while the fidelity of dTTP remained relatively unchanged when the gap width was expanded. Together, the Pol λ results indicated that the N-terminal BRCT and proline- rich domains of Pol λ were responsible for (i) decreasing the incorporation efficiency of a correct dNTP when the gap size exceeds 9 nucleotides, (ii) enhancing misincorporations when the gap size is greater than 1 nucleotide, and (iii) downregulating polymerase fidelity when the gap size is greater than 1 nucleotide.
Effects of DNA template sequence on nucleotide preference In general, dCTP was the preferred misincorporation for the four enzymes examined in this work. Analysis of the DNA template sequence suggested that the favorable dCTP
131 incorporation was being instructed by the identity of the first downstream template base: dG (Table 6.1). For all three of the Pol λ constructs, the catalytic efficiency of dCTP misincorporation was consistently strong at a gap size of 2 (Figure 6.3), therefore, the 42- mer DNA template sequence was changed from 3′-CGG-5′ to 3′-CGA-5′ and 3′-CAG-5′ (Table 6.1). The kinetic parameters were measured for the four polymerases using the modified DNA templates (Tables 6.6-6.9). Compared to the 42-merCGG template, wild- type Pol λ and the truncated mutants displayed a reduced preference for dCTP and a greater preference for dTTP when dA was the first downstream base (Figure 6.5 parts (A- C)). Thus, the fidelity of Pol λ improved for dCTP but dropped for dTTP (Figure 6.6 parts (A-C)). Also notable is that two consecutive downstream dG bases did not strongly influence nucleotide specificities, since similar kinetic results were obtained for 42- merCGG and 42-merCGA. In regards to Pol β, the sequence-dependent effect was most pronounced at a gap size of 7 (Figures 6.3 part (B) and 6.4 part (B)) so the 47-mer template was changed from 3′-CGA-5′ to 3′-CAT-5′. Like Pol λ with the 42-mers, a similar kinetic trend was observed for Pol β: dTTP misincorporation was preferred for the 47-merCAT template while dCTP was preferred for 47-merCGA (Figures 6.5 part (D) and 6.6 part (D)). Thus, the favored misincorporation during gap-filling DNA synthesis is promoted by the base identity at the first downstream position when the gap size is 2 or greater. However, the magnitude of the misincorporation efficiency for the preferred dNTP likely depends on gap size, since Pol λ inserted dTTP with a greater catalytic efficiency into 47-merCAT than 42-merCAG (Table 6.6).
6.4 Discussion
Currently, at least six mammalian DNA polymerases (Pols β, δ, ε, λ, ι, and θ) are implicated in BER. Unfortunately, confirming their exact cellular roles in BER remains uncertain due to the functional redundancy of the polymerases, therefore, in vitro biochemical characterization has been instructive in understanding how these enzymes function in BER. This work investigated the enzymology of Pol λ and Pol β during gap-
132 filling DNA synthesis. DNA substrates with various gap sizes were used to model DNA in SP-BER (1-nucleotide gap) and LP-BER (2- to 10-nucleotide gap). In accordance with previous studies on Pol λ and Pol β, the presence of a 5′-phosphorylated downstream strand is critical for maintaining the highest catalytic efficiency during correct nucleotide incorporation (Tables 6.2 and 6.3) [114, 192, 201, 234]. Examining the kinetics of nucleotide incorporation as a function of gap width at 1-nucleotide intervals for Pol λ and at key gap lengths for Pol β, dPol λ, and tPol λ revealed several important findings: (i) the gap threshold for Pol λ was 9 for a correct dNTP and as low as 7 for an incorrect dNTP, (ii) the fidelity of Pol λ dropped for gap sizes of 2 through 10 nucleotides, (iii) the BRCT and proline-rich domains downregulate the fidelity of Pol λ with the multi-nucleotide gapped DNA substrates, (iv) the preferred incorrect dNTP was influenced by the first downstream template base, and (v) for Pol β, both correct and incorrect nucleotide incorporation efficiencies dropped with an increase in gap size while its average fidelity was nearly unaffected.
Kinetic barriers for efficient nucleotide incorporation The incorporation efficiency of a correct nucleotide catalyzed by Pol λ and Pol β vary as a function of gap size (Figure 6.3 parts (A) and (B)). Catalysis remained relatively efficient when the gap size is small, presumably due to strong interactions between the dRPase domain and the 5′-phosphorylated downstream strand based upon crystallographic evidence [55, 119]. Pol λ can remain engaged with both the 3′-hydroxyl of the upstream primer strand and 5′-phosphate of the downstream strand when the gap width is extended to 2 [236]. Interestingly, this task is achieved by a template scrunching mechanism, whereby Pol λ accommodates the first downstream template base in an extrahelical position and forms a “scrunching” binding pocket that consists of L277, H511, and R514 [236]. Since Pol λ maintained efficient catalysis of dGTP until a gap size of 10, these results supported a model in which Pol λ binds both the 3′-hydroxyl and 5′-phosphate moieties so that a DNA loop emerges between the thumb subdomain and dRPase domain when the gap is 9 nucleotides or less (Complex I in Figure 6.7). In this model, we speculate that the interactions between the “scrunching” binding pocket and
133 the first downstream template base stabilize the conformation of the nascent base pair at the polymerase active site which facilitates efficient catalysis (Complex II in Figure 6.7). Once the intervening singlestranded DNA template surpasses 9 nucleotides, there may be an energetic penalty to accommodate such a large DNA loop, thereby preventing the polymerase domain from properly binding the 3′-hydroxyl and the nascent base pair while the dRPase domain is engaged with the 5′-phosphate. Structural studies indicate that the dRPase domain of Pol λ governs the positioning of the polymerase domain on the DNA substrate [237], therefore, sub-optimal positioning of the primer 3′-terminus at Pol λ’s active site may lead to the reduced catalytic efficiency. In addition, efficient gap- filling polymerization by Pol λ was also regulated by the non-enzymatic N-terminal domains. For a correct incorporation, the proline-rich domain was predominantly responsible for decreasing incorporation efficiency with increasing gap size (Figure 6.3). The crystal structure of full-length Pol λ has not been solved; therefore, it is difficult to determine how these domains alter polymerase function. Based on the dGTP incorporation efficiency differences among Pol λ, dPol λ, and tPol λ (Figure 6.3 parts (A), (C), and (D)), one can postulate that there is steric interference between the protruding template loop and the N-terminal BRCT and proline-rich domains of Pol λ.
Pol λ and Pol β exhibited distinct kinetic trends for the various DNA substrates which reiterated how these two X-family homologs possess different enzymatic properties [117, 206]. Pol β showed the lowest level of catalytic efficiency with a gap size of 10 but surprisingly was able to recover when the downstream strand was absent (Table 6.3 and Figure 6.3 part (B)). Previously, it has been observed that long gap-filling DNA synthesis performed by Pol β results in product accumulation at a position that corresponds to a gap size of 10 nucleotides [238]. Based on our single-turnover kinetic studies, product accumulation is likely due to the lower incorporation efficiency of Pol β. Unfortunately, there is no crystallographic evidence to show that Pol β is able to scrunch a DNA template like tPol λ. In Pol β, the analogous “scrunching” pocket residues are N37, I277, and K280, therefore, nuances likely exist in the long-patch gap-filling mechanisms of Pol
134 β and may not follow the model for tPol λ in Figure 6.7. Nonetheless, Pol β may still form a binary complex similar to Complex I (Figure 6.7) for gapped DNA. Removing the downstream strand eliminated the steric constraints of a looped single-stranded DNA template and reduced its inhibitory effect on polymerization efficiency, thereby indicating that Pol β could achieve a more catalytically-competent ternary complex. However, such a catalytic recovery was not significant for Pol λ and its truncated mutants. Although the reason for lack of recovery is unclear, one possibility is that the absence of the interactions between the downstream primer and the dRPase domain significantly affected the conformation of the nascent base pair at Pol λ’s active site.
Pol λ was error-prone when gap size was increased The base substitution fidelity of Pol λ dropped up to 40-fold when the gap size was increased from 1 to 10 nucleotides (Table 6.2). Moreover, the fidelity of Pol λ was regulated by its non-enzymatic N-terminal domains to varying degrees for each incorrect dNTP (Figure 6.4). For 1-nucleotide gapped DNA, the proline-rich domain of Pol λ was previously shown to increase polymerase fidelity up to 100-fold [117]. In contrast, dCTP misincorporation was not affected by the N-terminal domains at a gap size of 2, as all three Pol λ constructs showed a similar drop in fidelity (Figure 6.4 parts (A), (C), and (D)). Unlike Pol λ and dPol λ, tPol λ did not sustain a lower fidelity for dCTP misinsertion into gap sizes 5 and 7, thereby indicating that the proline-rich domain was responsible for downregulating the fidelity of Pol λ during long-patch gap-filling DNA synthesis. Interestingly, both the BRCT and proline-rich domains contributed to a decrease in polymerase fidelity for dATP and dTTP misincorporations into DNA gaps with widths exceeding 2 nucleotides. Thus, deleting Pol λ’s N-terminal domains resulted in an enzyme that was kinetically more Pol β-like, i.e. the fidelity remained more constant or increased as the gap size was expanded. The relationship among polymerase fidelity, gap size, and the N-terminal domains of Pol λ can be summarized as follows: the proline-rich domain upregulates Pol λ’s fidelity during short gap-filling DNA synthesis (1-nucleotide gap) while the BRCT and proline-rich domains downregulate polymerase fidelity during long gap-filling DNA synthesis.
135
The incorrect nucleotide with the highest probability of being inserted by Pol λ was dCTP, and this preference was shown to be sequence dependent due to the presence of a 5′-dG adjacent to the template base dC (Figure 6.3 part (A) and Figure 6.5 part (A)). This result is indicative of a strand misalignment which leads to the template base dC being looped out so the adjacent 5′-dG template base can base pair with dCTP. Such a dNTP- stabilized strand misalignment is supported by the fact that the preferred misincorporation was correspondingly changed from dCTP to dTTP when the first downstream template base 5′-dG was changed to 5′-dA (Table 6.6) and has been observed in the crystal structures of tPol λ [212, 239]. By repositioning the first downstream template base dG, the second downstream template base may occupy the “scrunching” binding pocket to further stabilize the conformation of a misaligned dCTP:dG intermediate at the active site of Pol λ (Complex IV in Figure 6.7). This pathway would lead to the generation of frameshift deletions (Complex V in Figure 6.7), a characteristic of Pol λ. Due to multiple steric constraints in the DNA template, the second downstream template base may not be oriented properly in the “scrunching” binding pocket (Complex IV in Figure 6.7). In addition, the conformation of Complex IV is less optimal for catalysis than Complex II (Figure 6.7) because the incorporation efficiency of correct dGTP is approximately two orders of magnitude greater than incorrect dCTP (Figure 6.3). Assimilating these kinetic and structural findings indicates that the incoming dNTP and DNA template sequence may dictate whether Pol λ forms a slipped DNA intermediate (Complex II versus IV in Figure 6.7). Our results suggested that upon dCTP binding Pol λ prefers the strand misalignment route to enable dCTP:dG base pair formation, since dCTP incorporation remained enhanced for gap sizes 2 through 8 (Figure 6.3 part (A)). Similar effects were observed for dPol λ and tPol λ, thereby indicating the BRCT and proline-rich domains did not significantly influence template misalignment (Figure 6.3 parts (C) and (D)). Like Pol λ, both dPol λ and tPol λ misincorporated dCTP with a greater catalytic efficiency for 2-nucleotide gapped DNA than 1-nucleotide gapped DNA. For a 1-nucleotide gap, the potential to loop out the downstream template dG was likely attenuated by the annealed downstream strand.
136 Interestingly, Figure 6.3 part (B) shows that dCTP misincorporation was not preferred by Pol β with most tested gap sizes. This is consistent with published results which demonstrate that Pol λ [240] has a stronger propensity to create -1 deletions than Pol β [241].
Potential roles of Pol β and Pol λ in BER Both Pol β and Pol λ exhibit similar efficiency and accuracy for filling single-nucleotide gapped DNA (Tables 6.2 and 6.3). This suggests that Pol λ, like Pol β, would be competent in SP-BER in vivo. Increasing gap size led to unexpected enzyme-specific properties of Pol λ and Pol β with a single-stranded DNA break. Based on polymerase fidelity (Tables 6.2 and 6.3), Pol β would be better than Pol λ for accurately filling larger than 1-nucleotide gaps in LP-BER. However, Pol β is not ideal for this role either since its catalytic efficiency significantly decreased with increasing gap size (Table 6.3 and Figure 6.3 part (B)). Notably, gap sizes greater than 5 nucleotides may not be physiologically relevant because most gap sizes are reported to be 2-4 nucleotides [226, 227]. In addition, the gap-filling role of Pol β and Pol λ in LP-BER may be confined to 1- nucleotide gaps generated by the alternating activities of flap endonuclease 1 and Pol β if the repair pathway proceeds via a “Hit and Run” mechanism as proposed by Wilson and coworkers [242]. Although less likely, LP-BER may also follow a tandem process in which the first insertion at the repair site is performed by Pol β or Pol λ followed by strand-displacement DNA synthesis by a replicative polymerase such as Pol δ or ε [221, 224]. In addition, the mutational spectrum in BER is predominantly single-nucleotide deletions in cell extracts [243]. Therefore, Pol λ may function in LP-BER because it has a strong propensity to create -1 deletions. Taken together, we cannot exclude potential roles of Pol β and Pol λ in LP-BER.
6.5 Future Directions
LP-BER can proceed through a PCNA-dependent or PCNA-independent pathway. It is not known whether Pol λ would participate exlusively in one pathway or both. However, 137 Pol λ has been shown to interact with PCNA and the site of interaction has been mapped to residues that interact with the DNA substrate [244]. The proximity of PCNA to the active site and DNA binding cleft may prevent template slippage by Pol λ. Thus, it would be interesting to determine if the presence of PCNA would improve the fidelity of Pol λ filling a gap size of 2- to 7-nucleotides. In addition, the presence of PCNA may alter Pol λ-mediated DNA loop formation and stability (Figure 6.7).
138 6.6 Tables
Table 6.1 DNA substrates*.
21-19/ 5’-CGCAGCCGTCCAACCAACTCA pCGTCGATCCAATGCCGTCC-3’ 41-mer 3’-GCGTCGGCAGGTTGGTTGAGTCGCAGCTAGGTTACGGCAGG-5’
21-19/ 5’-CGCAGCCGTCCAACCAACTCA pCGTCGATCCAATGCCGTCC-3’ 42-mer 3’-GCGTCGGCAGGTTGGTTGAGTCGGCAGCTAGGTTACGGCAGG-5’
21-19/ 5’-CGCAGCCGTCCAACCAACTCA pCGTCGATCCAATGCCGTCC-3’ 43-mer 3’-GCGTCGGCAGGTTGGTTGAGTCGAGCAGCTAGGTTACGGCAGG-5’
21-19/ 5’-CGCAGCCGTCCAACCAACTCA pCGTCGATCCAATGCCGTCC-3’ 44-mer 3’-GCGTCGGCAGGTTGGTTGAGTCGAGGCAGCTAGGTTACGGCAGG-5’
21-19/ 5’-CGCAGCCGTCCAACCAACTCA pCGTCGATCCAATGCCGTCC-3’ 45-mer 3’-GCGTCGGCAGGTTGGTTGAGTCGAGTGCAGCTAGGTTACGGCAGG-5’
21-19/ 5’-CGCAGCCGTCCAACCAACTCA pCGTCGATCCAATGCCGTCC-3’ 46-mer 3’-GCGTCGGCAGGTTGGTTGAGTCGAGTGGCAGCTAGGTTACGGCAGG-5’
21-19/ 5’-CGCAGCCGTCCAACCAACTCA pCGTCGATCCAATGCCGTCC-3’ 47-mer 3’-GCGTCGGCAGGTTGGTTGAGTCGAGTGCGCAGCTAGGTTACGGCAGG-5’
21-19/ 5’-CGCAGCCGTCCAACCAACTCA pCGTCGATCCAATGCCGTCC-3’ 48-mer 3’-GCGTCGGCAGGTTGGTTGAGTCGAGTGCGGCAGCTAGGTTACGGCAGG-5’
21-19/ 5’-CGCAGCCGTCCAACCAACTCA pCGTCGATCCAATGCCGTCC-3’ 49-mer 3’-GCGTCGGCAGGTTGGTTGAGTCGAGTGCGAGCAGCTAGGTTACGGCAGG-5’
21-19/ 5’-CGCAGCCGTCCAACCAACTCA pCGTCGATCCAATGCCGTCC-3’ 50-mer 3’-GCGTCGGCAGGTTGGTTGAGTCGAGTGCGACGCAGCTAGGTTACGGCAGG-5’
21-19T/42- 5’-CGCAGCCGTCCAACCAACTCA pTGTCGATCCAATGCCGTCC-3’ merCGA 3’-GCGTCGGCAGGTTGGTTGAGTCGACAGCTAGGTTACGGCAGG-5’
21-19/42- 5’-CGCAGCCGTCCAACCAACTCA pCGTCGATCCAATGCCGTCC-3’ merCAG 3’-GCGTCGGCAGGTTGGTTGAGTCAGCAGCTAGGTTACGGCAGG-5’
21-19/47- 5’-CGCAGCCGTCCAACCAACTCA pCGTCGATCCAATGCCGTCC-3’ merCAT 3’-GCGTCGGCAGGTTGGTTGAGTCATGTGCGCAGCTAGGTTACGGCAGG-5’ *Nucleotides located in the gap are in bold, and those that were inserted to expand the gap size
are underlined. p denotes the 5′-end is phosphorylated.
139
Table 6.2 Kinetic parameters for nucleotide incorporation into gapped or recessed DNA catalyzed by Pol λ at 37 °C. k K k /K Efficiency dNTP p d p d Fidelityb (s-1) (μM) (μM-1s-1) ratioa 21-19/41-mer (1-nucleotide gap) dGTP 2.7 ± 0.1 1.9 ± 0.2 1.4 - dCTP 0.00145 ± 0.00005 1.0 ± 0.1 1.5 × 10-3 - 1.0 × 10-3 dATP 0.00047 ± 0.00002 0.9 ± 0.1 5.2 × 10-4 - 3.7 × 10-4 dTTP 0.00135 ± 0.00009 2.9 ± 0.6 4.7 × 10-4 - 3.3 × 10-4 21-19/42-mer (2-nucleotide gap) dGTP 1.51 ± 1.2 1 1.77 ± 0.02 0.06 dCTP 0.69 ± 2.3 × 10-2 16 ↑ 2.0 × 10-2 0.0161 ± 0.0004 0.08 dATP 0.00037 ± 0.00002 1.2 ± 0.2 3.1 × 10-4 2 ↑ 2.6 × 10-4 dTTP 0.00070 ± 0.00003 3.8 ± 0.5 1.8 × 10-4 3 ↑ 1.6 × 10-4 21-19/43-mer (3-nucleotide gap) dGTP 1.99 ± 0.06 1.5 ± 0.2 1.3 1 dCTP 0.79 ± 1.0 × 10-2 7 ↑ 7.5 × 10-3 0.0079 ± 0.0002 0.08 dATP 0.00044 ± 0.00003 0.6 ± 0.2 7.3 × 10-4 1 5.5 × 10-4 dTTP 0.00095 ± 0.00008 2.8 ± 0.8 3.4 × 10-4 1 2.6 × 10-4 21-19/44-mer (4-nucleotide gap) dGTP 1.64 ± 0.03 1.6 ± 0.1 1.0 1 dCTP 0.0247 ± 0.0006 0.8 ± 0.1 3.1 × 10-2 21 ↑ 2.9 × 10-2 dATP 0.48 ± 2.6 × 10-3 5 ↑ 2.6 × 10-3 0.00126 ± 0.00004 0.07 dTTP 0.00129 ± 0.00007 2.8 ± 0.5 4.6 × 10-4 1 4.5 × 10-4 21-19/45-mer (5-nucleotide gap) dGTP 2.15 ± 0.06 1.6 ± 0.2 1.3 1 dCTP 0.053 ± 0.001 0.9 ± 0.1 5.9 × 10-2 41 ↑ 4.2 × 10-2 dATP 0.54 ± 5.7 × 10-3 11 ↑ 4.3 × 10-3 0.0031 ± 0.0001 0.07 dTTP 0.0118 ± 0.0006 2.7 ± 0.4 4.4 × 10-3 9 ↑ 3.2 × 10-3 21-19/46-mer (6-nucleotide gap) dGTP 1.53 ± 0.06 0.7 ± 0.1 2.2 2 ↑ dCTP 0.79 ± 5.7 × 10-2 39 ↑ 2.5 × 10-2 0.0450 ± 0.0008 0.07 dATP 0.46 ± 3.2 × 10-3 6 ↑ 1.5 × 10-3 0.00149 ± 0.00002 0.03 dTTP 0.0115 ± 0.0003 2.0 ± 0.2 5.8 × 10-3 12 ↑ 2.6 × 10-3
Continued
140 Table 6.2: Continued
21-19/47-mer (7-nucleotide gap) dGTP 1.86 ± 0.04 1.2 ± 0.1 1.6 1 dCTP 0.049 ± 0.001 0.9 ± 0.1 5.4 × 10-2 38 ↑ 3.4 × 10-2 dATP 0.24 ± 2.8 × 10-3 5 ↑ 1.8 × 10-3 0.00066 ± 0.00001 0.02 dTTP 0.0051 ± 0.0002 1.1 ± 0.2 4.6 × 10-3 10 ↑ 3.0 × 10-3 21-19/48-mer (8-nucleotide gap) dGTP 1.01 ± 0.03 1.6 ± 0.2 0.6 2 ↓ dCTP 0.47 ± 1.7 × 10-2 12 ↑ 2.7 × 10-2 0.0082 ± 0.0002 0.06 dATP 0.00023 ± 0.00001 0.8 ± 0.2 2.9 × 10-4 2 ↓ 4.6 × 10-4 dTTP 0.00072 ± 0.00002 5.1 ± 0.4 1.4 × 10-4 3 ↓ 2.2 × 10-4 21-19/49-mer (9-nucleotide gap) dGTP 0.83 ± 0.03 3.0 ± 0.3 0.3 5 ↓ dCTP 0.38 ± 5.9 × 10-3 4 ↓ 2.1 × 10-2 0.00226 ± 0.00005 0.05 dATP No incorporation dTTP No incorporation 21-19/50-mer (10-nucleotide gap) dGTP 0.026 ± 0.001 0.7 ± 0.1 3.7 × 10-2 38 ↓ dCTP 0.00164 ± 0.00007 1.1 ± 0.2 1.5 × 10-3 1 3.9 × 10-2 dATP No incorporation dTTP No incorporation 21/41-mer (no gap) dGTP 0.82 ± 4.3 × 10-2 33 ↓ 0.0353 ± 0.0008 0.08 dCTP 0.00044 ± 0.00003 3.0 ± 0.7 1.5 × 10-4 10 ↓ 3.4 × 10-3 dATP No incorporation dTTP No incorporation a An upward-pointing arrow (↑) indicates the ratio was calculated as (kp/Kd)≥2-nucleotide gap/(kp/Kd)1-nucleotide gap; a downward-pointing arrow (↓) indicates the calculation used a reciprocal of the equation as follows: (kp/Kd)1-nucleotide gap/(kp/Kd)≥2-nucleotide gap. b Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
141
Table 6.3 Kinetic parameters for nucleotide incorporation into gapped or recessed DNA catalyzed by Pol β at 37 °C. k K k /K Efficiency dNTP p d p d Fidelityb (s-1) (μM) (μM-1s-1) ratioa 21-19/41mer (1-nucleotide gap) dGTP 18.8 ± 0.4 8.7 ± 0.4 2.2 - dCTP 0.059 ± 0.002 140 ± 20 4.2 × 10-4 - 1.9 × 10-4 dATP 0.32 ± 0.02 280 ± 60 1.1 × 10-3 - 5.3 × 10-4 dTTP 0.27 ± 0.01 330 ± 40 8.2 × 10-4 - 3.7 × 10-4 21-19/42mer (2-nucleotide gap) dGTP 39 ± 1 12 ± 2 3.3 1 dCTP 0.0153 ± 0.0002 34 ± 3 4.5 × 10-4 1 1.4 × 10-4 dATP 0.212 ± 0.009 200 ± 20 1.1 × 10-3 1 3.3 × 10-4 dTTP 0.173 ± 0.009 340 ± 50 5.1 × 10-4 2 ↓ 1.6 × 10-4 21-19/45mer (5-nucleotide gap) dGTP 36 ± 2 100 ± 10 3.6 × 10-1 6 ↓ dCTP 0.065 ± 0.002 450 ± 30 1.4 × 10-4 3 ↓ 4.0 × 10-4 dATP 0.0188 ± 0.0006 540 ± 40 3.5 × 10-5 33 ↓ 9.7 × 10-5 dTTP 0.0117 ± 0.0008 900 ± 100 1.3 × 10-5 63 ↓ 3.6 × 10-5 21-19/47mer (7-nucleotide gap) dGTP 37 ± 5 100 ± 30 3.7 × 10-1 6 ↓ dCTP 0.203 ± 0.006 500 ± 40 4.1 × 10-4 1 1.1 × 10-3 dATP 0.013 ± 0.002 800 ± 300 1.6 × 10-5 70 ↓ 4.4 × 10-5 dTTP 0.0096 ± 0.0005 1400 ± 100 6.9 × 10-6 120 ↓ 1.9 × 10-5 21-19/50mer (10-nucleotide gap) dGTP 10.5 ± 0.6 780 ± 90 1.3 × 10-2 160 ↓ dCTP 0.0023 ± 0.0001 580 ± 60 4.0 × 10-6 110 ↓ 2.9 × 10-4 dATP 0.00166 ± 0.00009 440 ± 60 3.8 × 10-6 300 ↓ 2.8 × 10-4 dTTP 0.00090 ± 0.00006 620 ± 90 1.5 × 10-6 560 ↓ 1.1 × 10-4 21/41mer (no gap) dGTP 31.4 ± 0.6 48 ± 2 6.5 × 10-1 3 ↓ dCTP 0.024 ± 0.001 230 ± 30 1.0 × 10-4 4 ↓ 1.6 × 10-4 dATP 0.111 ± 0.007 650 ± 90 1.7 × 10-4 7 ↓ 2.6 × 10-4 dTTP 0.094 ± 0.005 1100 ± 100 8.5 × 10-5 10 ↓ 1.3 × 10-4 a A downward-pointing arrow (↓) indicates the ratio was calculated as (kp/Kd)1-nucleotide gap/(kp/Kd)≥2-nucleotide gap. b Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
142
Table 6.4 Kinetic parameters for nucleotide incorporation into gapped or recessed DNA catalyzed by dPol λ at 37 °C. k K k /K Efficiency dNTP p d p d Fidelityb (s-1) (μM) (μM-1s-1) ratioa 21-19/41mer (1-nucleotide gap) dGTP 3.1 ± 0.1 1.7 ± 0.2 1.8 - dCTP 0.00135 ± 0.00007 1.9 ± 0.4 7.1 × 10-4 - 3.9 × 10-4 dATP 0.00066 ± 0.00007 1.8 ± 0.7 3.7 × 10-4 - 2.0 × 10-4 dTTP 0.00130 ± 0.00009 7 ± 1 1.9 × 10-4 - 1.0 × 10-4 21-19/42mer (2-nucleotide gap) dGTP 1.24 ± 2.3 1 2.80 ± 0.05 0.09 dCTP 0.85 ± 2.4 × 10-2 34 ↑ 1.1 × 10-2 0.0208 ± 0.0005 0.09 dATP 0.00031 ± 0.00002 3.0 ± 0.6 1.0 × 10-4 4 ↓ 4.6 × 10-5 dTTP 0.00070 ± 0.00004 4.6 ± 0.7 1.5 × 10-4 1 6.7 × 10-5 21-19/45mer (5-nucleotide gap) dGTP 1.57 ± 2.4 1 3.83 ± 0.06 0.09 dCTP 0.0060 ± 0.0003 1.5 ± 0.2 4.0 × 10-3 6 ↑ 1.6 × 10-3 dATP 0.00042 ± 0.00003 3.3 ± 0.6 1.3 × 10-4 3 ↓ 5.2 × 10-5 dTTP 0.00154 ± 0.00009 7 ± 1 2.2 × 10-4 1 9.0 × 10-5 21-19/47mer (7-nucleotide gap) dGTP 1.28 ± 2.0 1 2.62 ± 0.05 0.09 dCTP 0.77 ± 9.4 × 10-3 13 ↑ 4.5 × 10-3 0.0072 ± 0.0002 0.07 dATP 0.000102 ± 0.000008 2.1 ± 0.5 4.9 × 10-5 8 ↓ 2.4 × 10-5 dTTP 0.0006 ± 0.0001 6 ± 3 1.0 × 10-4 2 ↓ 4.9 × 10-5 21-19/50mer (10-nucleotide gap) dGTP 0.27 ± 0.01 2.4 ± 0.3 1.1 × 10-1 16 ↓ dCTP 0.00019 ± 0.00001 5 ± 1 3.8 × 10-5 19 ↓ 3.4 × 10-4 dATP No incorporation dTTP No incorporation 21/41mer (no gap) dGTP 0.109 ± 0.007 1.7 ± 0.3 6.4 × 10-2 28 ↓ dCTP 0.00030 ± 0.00002 4.2 ± 0.7 7.1 × 10-5 10 ↓ 1.1 × 10-3 dATP No incorporation dTTP No incorporation a An upward-pointing arrow (↑) indicates the ratio was calculated as (kp/Kd)≥2-nucleotide gap/(kp/Kd)1-nucleotide gap; a downward-pointing arrow (↓) indicates the calculation used a reciprocal of the equation as follows: (kp/Kd)1-nucleotide gap/(kp/Kd)≥2-nucleotide gap. b Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
143
Table 6.5 Kinetic parameters for nucleotide incorporation into gapped or recessed DNA catalyzed by tPol λ at 37 °C. k K k /K Efficiency dNTP p d p d Fidelityb (s-1) (μM) (μM-1s-1) ratioa 21-19/41mer (1-nucleotide gap)c dGTP 4.1 ± 0.2 1.9 ± 0.4 2.2 - dCTP 0.0098 ± 0.0002 1.5 ± 0.2 6.5 × 10-3 - 3.0 × 10-3 dATP 0.0046 ± 0.0001 1.4 ± 0.3 3.3 × 10-3 - 1.5 × 10-3 dTTP 0.0065 ± 0.0001 4.7 ± 0.5 1.4 × 10-3 - 6.4 × 10-4 21-19/42mer (2-nucleotide gap) dGTP 3.7 ± 0.2 2.3 ± 0.3 1.6 1 dCTP 1.12 ± 7.2 × 10-2 11 ↑ 4.3 × 10-2 0.081 ± 0.001 0.07 dATP 0.0019 ± 0.0002 2.4 ± 0.6 7.9 × 10-4 4 ↓ 4.9 × 10-4 dTTP 0.0030 ± 0.0009 6 ± 3 5.0 × 10-4 3 ↓ 3.1 × 10-4 21-19/45mer (5-nucleotide gap) dGTP 5.1 ± 0.2 3.3 ± 0.4 1.5 1 dCTP 0.0123 ± 0.0003 1.4 ± 0.1 8.8 × 10-3 1 5.7 × 10-3 dATP 0.0011 ± 0.0002 2 ± 1 5.5 × 10-4 6 ↓ 3.6 × 10-4 dTTP 0.006 ± 0.002 9 ± 4 6.7 × 10-4 2 ↓ 4.3 × 10-4 21-19/47mer (7-nucleotide gap) dGTP 3.78 ± 0.08 2.5 ± 0.2 1.5 1 dCTP 0.028 ± 0.002 3.5 ± 0.8 8.0 × 10-3 1 5.3 × 10-3 dATP 0.00035 ± 0.00002 1.6 ± 0.4 2.2 × 10-4 15 ↓ 1.4 × 10-4 dTTP 0.0027 ± 0.0002 11 ± 2 2.5 × 10-4 6 ↓ 1.6 × 10-4 21-19/50mer (10-nucleotide gap) dGTP 1.43 ± 0.05 5.1 ± 0.4 2.8 × 10-1 8 ↓ dCTP 0.00067 ± 0.00006 5 ± 1 1.3 × 10-4 49 ↓ 4.8 × 10-4 dATP 0.000350 ± 9.7 × 10-5 34 ↓ 3.5 × 10-4 0.000009 3.6 ± 0.3 dTTP No incorporation 21/41mer (no gap) dGTP 0.68 ± 0.02 2.0 ± 0.2 3.4 × 10-1 6 ↓ dCTP 0.0007 ± 0.0001 5 ± 2 1.4 × 10-4 47 ↓ 4.1 × 10-4 dATP No incorporation dTTP No incorporation a An upward-pointing arrow (↑) indicates the ratio was calculated as (kp/Kd)≥2-nucleotide gap/(kp/Kd)1-nucleotide gap; a downward-pointing arrow (↓) indicates the calculation used a reciprocal of the equation as follows: (kp/Kd)1-nucleotide gap/(kp/Kd)≥2-nucleotide gap. b Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect]. cKinetic parameters are from reference [182].
144
Table 6.6 Kinetic parameters for nucleotide incorporation into gapped DNA catalyzed by Pol λ at 37 °C. k K k /K Efficiency dNTP p d p d Fidelityb (s-1) (μM) (μM-1s-1) ratioa 21-19/41mer (1-nucleotide gap) dGTP 2.7 ± 0.1 1.9 ± 0.2 1.4 - dCTP 0.00145 ± 0.00005 1.0 ± 0.1 1.5 × 10-3 - 1.0 × 10-3 dATP 0.00047 ± 0.00002 0.9 ± 0.1 5.2 × 10-4 - 3.7 × 10-4 dTTP 0.00135 ± 0.00009 2.9 ± 0.6 4.7 × 10-4 - 3.3 × 10-4 21-19/42mer (2-nucleotide gap) dGTP 1.77 ± 0.02 1.51 ± 0.06 1.2 1 dCTP 0.0161 ± 0.0004 0.69 ± 0.08 2.3 × 10-2 16 ↑ 2.0 × 10-2 dATP 0.00037 ± 0.00002 1.2 ± 0.2 3.1 × 10-4 2 ↑ 2.6 × 10-4 dTTP 0.00070 ± 0.00003 3.8 ± 0.5 1.8 × 10-4 3 ↑ 1.6 × 10-4 21-19T/42merCGA (2-nucleotide gap) dGTP 2.4 ± 0.1 2.4 ± 0.4 1 1 dCTP 0.0305 ± 0.0007 0.52 ± 0.06 5.9 × 10-2 40 ↑ 5.5 × 10-2 dATP 0.00069 ± 0.00003 0.6 ± 0.1 1.2 × 10-3 2 ↑ 1.1 × 10-3 dTTP 0.00075 ± 0.00004 3.3 ± 0.7 2.3 × 10-4 2 ↓ 2.3 × 10-4 21-19/42merCAG (2-nucleotide gap) dGTP 2.9 ± 0.1 1.7 ± 0.3 1.7 1 dCTP 0.00076 ± 0.00007 1.7 ± 0.5 4.5 × 10-4 3 ↓ 2.6 × 10-4 dATP 0.00049 ± 0.00002 0.9 ± 0.2 5.4 × 10-4 1 3.2 × 10-4 dTTP 0.0025 ± 0.0002 4.4 ± 0.8 5.7 × 10-4 1 3.3 × 10-4 21-19/47mer (7-nucleotide gap) dGTP 1.86 ± 0.04 1.2 ± 0.1 1.6 1 dCTP 0.049 ± 0.001 0.9 ± 0.1 5.4 × 10-2 38 ↑ 3.4 × 10-2 dATP 0.00066 ± 0.00001 0.24 ± 0.02 2.8 × 10-3 5 ↑ 1.8 × 10-3 dTTP 0.0051 ± 0.0002 1.1 ± 0.2 4.6 × 10-3 10 ↑ 3.0 × 10-3 21-19/47merCAT (7-nucleotide gap) dGTP 3.3 ± 0.2 3.1 ± 0.6 1.1 1 dCTP 0.0087 ± 0.0001 0.58 ± 0.03 1.5 × 10-2 10 ↑ 1.4 × 10-2 dATP 0.0045 ± 0.0001 1.2 ± 0.1 3.8 × 10-3 7 ↑ 3.5 × 10-3 dTTP 0.146 ± 0.002 2.5 ± 0.1 5.8 × 10-2 125 ↑ 5.2 × 10-2 a An upward-pointing arrow (↑) indicates the ratio was calculated as (kp/Kd)≥2-nucleotide gap/(kp/Kd)1-nucleotide gap; a downward-pointing arrow (↓) indicates the calculation used a reciprocal of the equation as follows: (kp/Kd)1-nucleotide gap/(kp/Kd)≥2-nucleotide gap. b Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
145
Table 6.7 Kinetic parameters for nucleotide incorporation into gapped DNA catalyzed by dPol λ at 37 °C. k K k /K Efficiency dNTP p d p d Fidelityb (s-1) (μM) (μM-1s-1) ratioa 21-19/41mer (1-nucleotide gap) dGTP 3.1 ± 0.1 1.7 ± 0.2 1.8 - dCTP 0.00135 ± 0.00007 1.9 ± 0.4 7.1 × 10-4 - 3.9 × 10-4 dATP 0.00066 ± 0.00007 1.8 ± 0.7 3.7 × 10-4 - 2.0 × 10-4 dTTP 0.00130 ± 0.00009 7 ± 1 1.9 × 10-4 - 1.0 × 10-4 21-19/42mer (2-nucleotide gap) dGTP 2.80 ± 0.05 1.24 ± 0.09 2.3 1 dCTP 0.0208 ± 0.0005 0.85 ± 0.09 2.4 × 10-2 34 ↑ 1.1 × 10-2 dATP 0.00031 ± 0.00002 3.0 ± 0.6 1.0 × 10-4 4 ↓ 4.6 × 10-5 dTTP 0.00070 ± 0.00004 4.6 ± 0.7 1.5 × 10-4 1 6.7 × 10-5 21-19T/42merCGA (2-nucleotide gap) dGTP 2.57 ± 0.08 0.9 ± 0.1 2.9 1 dCTP 0.0200 ± 0.0006 0.80 ± 0.07 2.5 × 10-2 35 ↑ 8.7 × 10-3 dATP 0.00027 ± 0.00002 1.3 ± 0.4 2.1 × 10-4 2 ↓ 7.3 × 10-5 dTTP 0.00024 ± 0.00002 3.3 ± 0.8 7.3 × 10-5 3 ↓ 2.5 × 10-5 21-19/42merCAG (2-nucleotide gap) dGTP 3.9 ± 0.2 1.6 ± 0.3 2.4 1 dCTP 0.00116 ± 0.00008 2.3 ± 0.5 5.0 × 10-4 1 2.1 × 10-4 dATP 0.00030 ± 0.00003 2.5 ± 0.8 1.2 × 10-4 3 ↓ 4.9 × 10-5 dTTP 0.0027 ± 0.0002 4.0 ± 0.9 6.8 × 10-4 4 ↑ 2.8 × 10-4 a An upward-pointing arrow (↑) indicates the ratio was calculated as (kp/Kd)≥2-nucleotide gap/(kp/Kd)1-nucleotide gap; a downward-pointing arrow (↓) indicates the calculation used a reciprocal of the equation as follows: (kp/Kd)1-nucleotide gap/(kp/Kd)≥2-nucleotide gap. b Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
146
Table 6.8 Kinetic parameters for nucleotide incorporation into gapped DNA catalyzed by tPol λ at 37 °C. k K k /K Efficiency dNTP p d p d Fidelityb (s-1) (μM) (μM-1s-1) ratioa 21-19/41mer (1-nucleotide gap)c dGTP 4.1 ± 0.2 1.9 ± 0.4 2.2 - dCTP 0.0098 ± 0.0002 1.5 ± 0.2 6.5 × 10-3 - 3.0 × 10-3 dATP 0.0046 ± 0.0001 1.4 ± 0.3 3.3 × 10-3 - 1.5 × 10-3 dTTP 0.0065 ± 0.0001 4.7 ± 0.5 1.4 × 10-3 - 6.4 × 10-4 21-19T/42mer (2-nucleotide gap) dGTP 3.7 ± 0.2 2.3 ± 0.3 1.6 1 dCTP 0.081 ± 0.001 1.12 ± 0.07 7.2 × 10-2 11 ↑ 4.3 × 10-2 dATP 0.0019 ± 0.0002 2.4 ± 0.6 7.9 × 10-4 4 ↓ 4.9 × 10-4 dTTP 0.0030 ± 0.0009 6 ± 3 5.0 × 10-4 3 ↓ 3.1 × 10-4 21-19/42merCGA (2-nucleotide gap) dGTP 4.8 ± 0.1 2.1 ± 0.2 2.3 1 dCTP 0.141 ± 0.003 0.78 ± 0.06 1.8 × 10-1 28 ↑ 7.3 × 10-2 dATP 0.0014 ± 0.0001 1.3 ± 0.3 1.1 × 10-3 3 ↓ 4.7 × 10-4 dTTP 0.0028 ± 0.0002 5 ± 2 5.6 × 10-4 2 ↓ 2.4 × 10-4 21-19/42merCAG (2-nucleotide gap) dGTP 4.6 ± 0.2 2.6 ± 0.4 1.8 1 dCTP 0.0064 ± 0.0003 1.3 ± 0.2 4.9 × 10-3 1 2.8 × 10-3 dATP 0.002 ± 0.0001 1.3 ± 0.3 1.5 × 10-3 2 ↓ 8.7 × 10-4 dTTP 0.016 ± 0.001 4.3 ± 0.9 3.7 × 10-3 3 ↑ 2.1 × 10-3 a An upward-pointing arrow (↑) indicates the ratio was calculated as (kp/Kd)≥2-nucleotide gap/(kp/Kd)1-nucleotide gap; a downward-pointing arrow (↓) indicates the calculation used a reciprocal of the equation as follows: (kp/Kd)1-nucleotide gap/(kp/Kd)≥2-nucleotide gap. b Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect]. cKinetic parameters are from Reference [182].
147
Table 6.9 Kinetic parameters for nucleotide incorporation into gapped DNA catalyzed by Pol β at 37 °C. k K k /K Efficiency dNTP p d p d Fidelityb (s-1) (μM) (μM-1s-1) ratioa 21-19/41mer (1-nucleotide gap) dGTP 18.8 ± 0.4 8.7 ± 0.4 2.2 - dCTP 0.059 ± 0.002 140 ± 20 4.2 × 10-4 - 1.9 × 10-4 dATP 0.32 ± 0.02 280 ± 60 1.1 × 10-3 - 5.3 × 10-4 dTTP 0.27 ± 0.01 330 ± 40 8.2 × 10-4 - 3.7 × 10-4 21-19/42mer (2-nucleotide gap) dGTP 39 ± 1 12 ± 2 3.3 1 dCTP 0.0153 ± 0.0002 34 ± 3 4.5 × 10-4 1 1.4 × 10-4 dATP 0.212 ± 0.009 200 ± 20 1.1 × 10-3 1 3.3 × 10-4 dTTP 0.173 ± 0.009 340 ± 50 5.1 × 10-4 2 ↓ 1.6 × 10-4 21-19T/42merCGA (2-nucleotide gap) dGTP 41 ± 4 10 ± 3 4.1 2 ↑ dCTP 0.041 ± 0.006 300 ± 100 1.4 × 10-4 3 ↓ 3.3 × 10-5 dATP 0.094 ± 0.005 190 ± 30 4.9 × 10-4 2 ↓ 1.2 × 10-4 dTTP 0.26 ± 0.07 1500 ± 600 1.7 × 10-4 5 ↓ 4.2 × 10-5 21-19/42merCAG (2-nucleotide gap) dGTP 44 ± 1 10 ± 1 4.4 2 ↑ dCTP 0.072 ± 0.004 340 ± 60 2.1 × 10-4 2 ↓ 4.8 × 10-5 dATP 0.180 ± 0.008 140 ± 20 1.3 × 10-3 1 2.9 × 10-4 dTTP 0.0103 ± 0.0003 17 ± 2 6.1 × 10-4 1 1.4 × 10-4 21-19/47mer (7-nucleotide gap) dGTP 37 ± 5 100 ± 30 3.7 × 10-1 6 ↓ dCTP 0.203 ± 0.006 500 ± 40 4.1 × 10-4 1 1.1 × 10-3 dATP 0.013 ± 0.002 800 ± 300 1.6 × 10-5 70 ↓ 4.4 × 10-5 dTTP 0.0096 ± 0.0005 1400 ± 100 6.9 × 10-6 120 ↓ 1.9 × 10-5 21-19/47merCAT (7-nucleotide gap) dGTP 32 ± 4 230 ± 60 1.4 × 10-1 16 ↓ dCTP 0.0116 ± 0.0009 1200 ± 200 9.7 × 10-6 44 ↓ 6.9 × 10-5 dATP 0.04 ± 0.01 1300 ± 700 3.1 × 10-5 37 ↓ 2.2 × 10-4 dTTP 0.0113 ± 0.0003 850 ± 40 1.3 × 10-5 62 ↓ 9.6 × 10-5 a An upward-pointing arrow (↑) indicates the ratio was calculated as (kp/Kd)≥2-nucleotide gap/(kp/Kd)1-nucleotide gap; a downward-pointing arrow (↓) indicates the calculation used a reciprocal of the equation as follows: (kp/Kd)1-nucleotide gap/(kp/Kd)≥2-nucleotide gap. b Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
148
6.7 Figures
Figure 6.1 Domains of Pol λ, dPol λ, tPol λ, and Pol β. Each domain is labeled in the rectangular box and the residue numbers are noted above. N-terminal residues 1-35 of Pol λ contain a nuclear localization sequence.
149 A
30
25
20
15
10 Product (nM) Product
5
0 0 100 200 300 400 500 600 Time (s)
B
0.04
0.035
0.03
) 0.025 -1
(s 0.02 obs
k 0.015
0.01
0.005
0 0 5 10 15 20 25 30 dGTP (μM)
Figure 6.2 Single-turnover kinetic parameters of nucleotide incorporation. (A) A pre- incubated solution of Pol λ (120 nM) and 5′-[32P]-labeled 21/41-mer (30 nM) was rapidly mixed with increasing concentrations of dGTP•Mg2+ (0.2 μM, z; 0.5 μM, {; 1 μM, ; 2 μM, ; 5 μM, S; 10 μM, U; 25 μM, ) for various time intervals. The solid lines are the best fits to a single-exponential equation which determined the observed rate constants, kobs. (B) The kobs values were plotted as a function of dGTP concentration. The -1 data (z) were then fit to a hyperbolic equation, yielding a kp of 0.0353 ± 0.0008 s and a Kd of 0.82 ± 0.08 μM.
150 A B
101 101 ) -1 ) s 0 -1 -1 10 0 s 10 M -1 μ M μ 10-1 10-1 10-2
-2 10 10-3
10-4 10-3 -5
Incorporation Efficiency ( 10
-4 Incorporation Efficiency ( 10 10-6 1 2 3 4 5 6 7 8 9 10 P/T 12345678910 P/T Gap Size Gap Size (Number of Nucleotides) (Number of Nucleotides)
C D 1 101 10 ) ) -1 -1 s s 0 0 -1 -1 10 10 M M μ μ 10-1 10-1
10-2 10-2
10-3 10-3
10-4 10-4 Incorporation Efficiency ( Incorporation Efficiency ( 10-5 10-5 1 2 3 4 5 6 7 8 9 10 P/T 1 2 3 4 5 6 7 8 9 10 P/T Gap Size Gap Size (Number of Nucleotides) (Number of Nucleotides)
Figure 6.3 Effect of gap size on polymerization efficiency. Polymerization efficiency is plotted as a function of gap size for (A) Pol λ, (B) Pol β, (C) dPol λ, and (D) tPol λ. The incoming nucleotide is represented as follows: for dGTP, z for dCTP, for dATP, and S for dTTP. P/T represents data for the recessed 21/41-mer DNA substrate.
151 A B
10-1 10-2
10-2 10-3 Fidelity Fidelity 10-3 10-4
10-4 10-5 12345678910 P/T 1 2 3 4 5 6 7 8 9 10 P/T Gap Size Gap Size (Number of Nucleotides) (Number of Nucleotides)
C D
10-1 10-1
10-2 10-2
10-3 Fidelity Fidelity 10-3 10-4
10-5 10-4 1 2 3 4 5 6 7 8 9 10 P/T 1 2 3 4 5 6 7 8 9 10 P/T Gap Size Gap Size (Number of Nucleotides) (Number of Nucleotides)
Figure 6.4 Effect of gap size on polymerase fidelity. Polymerase fidelity is plotted as a function of gap size for (A) Pol λ, (B) Pol β, (C) dPol λ, and (D) tPol λ. The incoming nucleotide is represented as follows: for dGTP, z for dCTP, for dATP, and S for dTTP. P/T represents data for the recessed 21/41-mer DNA substrate.
152 A B 1 10 101 ) 41merCGC ) 41merCGC -1 42merCGG -1 s s 0 42merCGG
-1 0 -1 10 10 42merCGA M 42merCAG M 42merCGA μ μ 47merCGA -1 42merCAG 47merCAT 10 47merCGA 10-1 10-2 10-2 10-3
10-3 10-4 Incorporation Efficiency ( Incorporation Efficiency ( -4 10 10-5 dGTP dCTP dATP dTTP dGTP dCTP dATP dTTP Incoming Nucleotide Incoming Nucleotide
C D 101 101 ) 41merCGC ) 41merCGC -1 42merCGG -1 0
s 42merCGG s 10
-1 0 10 42merCGA -1 42merCGA M 42merCAG M 42merCAG μ μ -1 47merCGA 10 47merCGA -1 47merCAT 10 -2 10
-3 10-2 10
10-4 10-3 10-5 Incorporation Efficiency ( Incorporation Efficiency ( 10-4 10-6 dGTP dCTP dATP dTTP dGTP dCTP dATP dTTP Incoming Nucleotide Incoming Nucleotide
Figure 6.5 Effect of DNA sequence on polymerization efficiency. Incorporation efficiency is plotted for each of the incoming nucleotides for (A) Pol λ, (B) dPol λ, (C) tPol λ, and (D) Pol β. Nucleotide incorporation into the different DNA substrates is represented in the legend as follows: solid black bars for 21-19/41merCGC, solid scarlet bars for 21-19/42merCGG, scarlet grid bars for 21-19T/42merCGA, scarlet criss-cross bars for 21-19/42merCAG, solid blue bars for 21-19/47merCGA, and blue criss-cross bars for 21-19/47merCAT.
153 A B
-1 10-1 10 41merCGC 41merCGC 42merCGG 42merCGG 42merCGA 42merCGA 42merCAG 10-2 42merCAG -2 47merCGA 47merCGA 10 47merCAT
10-3 Fidelity Fidelity 10-3 10-4
-5 10-4 10 dCTP dATP dTTP dCTP dATP dTTP Incoming Nucleotide Incoming Nucleotide
C D
-1 10 10-2 41merCGC 41merCGC 42merCGG 42merCGG 42merCGA 42merCGA 42merCAG 42merCAG -2 47merCGA 10 -3 47merCGA 10 47merCAT Fidelity -3 Fidelity 10 10-4
10-4 10-5 dCTP dATP dTTP dCTP dATP dTTP Incoming Nucleotide Incoming Nucleotide
Figure 6.6 Effect of DNA sequence on polymerization fidelity. The base substitution fidelity is plotted for each of the incoming nucleotides for (A) Pol λ, (B) dPol λ, (C) tPol λ, and (D) Pol β. The fidelity was calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect]. Nucleotide incorporation into the different DNA substrates is represented in the legend as follows: solid black bars for 21-19/41merCGC, solid scarlet bars for 21- 19/42merCGG, scarlet grid bars for 21-19T/42merCGA, scarlet criss-cross bars for 21- 19/42merCAG, solid blue bars for 21-19/47merCGA, and blue criss-cross bars for 21- 19/47merCAT.
154
Figure 6.7 Model for long gap-filling DNA synthesis catalyzed by tPol λ. Complex I shows the polymerase domain bound to the 3′-hydroxyl while the dRPase domain is bound to the 5′-phosphate (yellow dot) so that the single-stranded DNA template is looped out between the polymerase and dRPase domains. In the presence of a correct dNTP (Complex II), the first downstream template base is in the template “scrunching” binding pocket. Successive incorporations allow the downstream nucleotide that is immediately 5′ of the template base to reside in the scrunching pocket during each catalytic cycle (Complex III). In the presence of an incorrect dNTP that is complementary to the first downstream template base (Complex IV), the template base is looped out and the second downstream template base partially occupies the “scrunching” binding pocket. This complex would generate a frameshift deletion (Complex V).
155 Chapter 7: Single-Turnover Kinetic Analysis of the Mutagenic Potential of 8-Oxo-7,8-dihydro-2′-deoxyguanosine during Gap-Filling Synthesis Catalyzed by Human DNA Polymerase λ and β
7.1 Introduction
Oxidative DNA damage commonly generates 8-oxo-7,8-dihydro-2′-deoxyguanosine (8- oxodG) lesions in genomic DNA and less often in the deoxyribonucleotide pools, although both of these may contribute to the spontaneous mutations implicated in cancer and aging [245-247]. These potentially mutagenic 8-oxodG species exist in either an anti or syn conformation depending on their base pairing partner: an anti conformation coordinates Watson-Crick base pairing with dC while a syn conformation facilitates Hoogsteen base pairing with dA. According to NMR and X-ray crystal structures, the electronegative oxygen at the C8 position of 8-oxodG induces subtle, local alterations in the conformation of duplex DNA that are confined to the modified site [248-250], however, Fourier transform-infrared spectroscopy with multivariate statistics suggests that base interactions and the phosphodeoxyribose backbone structure near the lesion are perturbed in single-stranded DNA, the form present at DNA replication forks [251]. Consequently, the summation of these physical, chemical, and structural properties affects the proper mechanistic functioning of a DNA polymerase when it encounters 8- oxodG template lesions.
As observed in several binary (E•DNA) and ternary (E•DNA•dNTP) X-ray crystal structures, each polymerase tailors its active site to accommodate the additional carbonyl moiety of the 8-oxodG lesion responsible for the dual coding feature [252-258]. Numerous kinetic and biochemical studies reveal that efficient incorporation of dCTP or
156 dATP opposite 8-oxodG is highly polymerase-dependent [259]. For example, preferential insertion of dCTP over dATP has been reported for the following DNA polymerases: Escherichia coli DNA polymerases I and II (exo-), T7 DNA polymerase (exo-), RB69 DNA polymerase, Xenopus laevis DNA polymerase γ, Saccharomyces cerevisiae DNA polymerase η, Sulfolobus solfataricus Dpo4, human DNA polymerase β (Polβ), bovine DNA polymerase δ in the presence of human proliferating cell nuclear antigen (PCNA), and human DNA polymerase ι [23, 253, 256, 258, 260-267]. In contrast, dATP is selected over dCTP to form a nascent basepair with 8-oxodG by HIV-1 reverse transcriptase (HIV-1 RT), calf thymus DNA polymerases α and δ, Bacillus stearothermophilus DNA polymerase I fragment (BF), and human DNA polymerase κ [254, 259, 262, 268]. The erroneous incorporation of 8-oxodG 5′-triphosphate (8-oxodGTP) has also been examined albeit much less extensively [263, 269].
In this present study, we employed single-turnover kinetic assays to determine which nucleotide is preferentially incorporated opposite 8-oxodG catalyzed by full-length human DNA polymerase λ (Pol λ), a recently discovered X-family member [74, 200, 270, 271]. Although its physiological role is not yet known, many lines of biochemical and in vitro evidence suggest that Pol λ, like its X-family homolog Pol β, may play a role in base excision repair (BER) [74, 78, 172, 173, 200, 270-272]. Recent cellular data support a protective function for Pol λ against oxidative DNA damage [173, 191]. Pol λ and Pol β share 34% sequence identity and 54% sequence homology [74, 270]. The C-terminal region of Pol λ, also referred to as truncated DNA polymerase λ (tPol λ), contains 5′- deoxyribose-5-phosphate lyase (dRPase) and DNA polymerase domains similar to Pol β (Figure 7.1) [270]. However, the N-terminal region of Pol λ consists of a nuclear localization signal (NLS) motif, a breast cancer susceptibility gene 1 C terminus (BRCT) domain, and a Proline-rich domain (Figure 7.1) which collectively distinguish it from Pol β [74, 270]. These distinctive N-terminal domains likely influence the enzymatic activities of Pol λ, for our recent studies reveal that the Proline-rich domain enhances the polymerase fidelity up to 100-fold [117]. Thus, Pol λ may not conform to the trend set by Pol β and other polymerases which favor dCTP:8-oxodG over dATP:8-oxodG. Until
157 now, Pol β is the only X-family DNA polymerase characterized in the presence of the potentially mutagenic 8-oxodG lesion. We also tested whether or not this favored base pairing is symmetrical by determining the preferred DNA template base for the incorporation of 8-oxodGTP. For a thorough kinetic investigation, we measured the pre- steady state kinetic parameters for all possible dNTP:8-oxodG and 8-oxodGTP:dN incorporations catalyzed by both Pol β and Pol λ, the only two DNA polymerases knowingly or likely involved in short-patch BER.
7.2 Materials and Methods
Materials 32 These chemicals were purchased from the following companies: [γ- P]ATP and dNTPs, GE Healthcare; Biospin columns, Bio-Rad Laboratories; 8-oxodGTP, TriLink BioTechnologies; synthetic oligodeoxyribonucleotide 40-mer containing an 8-oxodG, Midland Certified Reagent Company; synthetic oligodeoxyribonucleotides 21-mer, 18- mer, 19-mer, and 41-mer, Integrated DNA Technologies.
Expression and purification of Pol λ and Pol β The preparation of Pol λ was previously described [117]. For human Pol β, TAP56 cells containing pWL11 were used to overexpress Pol β with a scheme similar to that which is described in Patterson et al. [273, 274]. The enzyme was purified from E. coli lysate through a P11 phosphocellulose column (Whatman), a Heparin Sepharose column (GE Healthcare), a Mono S column (GE Healthcare), and a DEAE-Sepharose column (GE Healthcare). Pure Pol β was then concentrated and flash frozen in buffer (50 mM Tris-Cl, pH 7.5 at 9 °C, 50% glycerol, 2 mM DTT, 100 mM NaCl). The concentration of purified protein was determined by UV spectrophotometry using a calculated extinction coefficient of 2.1 x 104 (M-1 cm-1) at 280 nm.
158 Single-nucleotide gapped DNA substrates The synthesized oligomers in Figure 7.2 were purified and the 21mer primer radiolabeled with [γ-32P]ATP as previously described [117]. The single-nucleotide gapped DNA substrates were prepared by mixing the 5′-[32P]-radiolabeled 21-mer, the non- radiolabeled downstream strand, and the corresponding template at a 1:1.25:1.15 molar ratio, respectively. Then, the annealing mixture was denatured at 95 °C for 8 minutes and slowly cooled to room temperature over several hours.
Reaction buffers For kinetic studies involving Pol λ, buffer L (50 mM Tris-Cl, pH 8.4 at 37 °C, 5 mM
MgCl2, 100 mM NaCl, 0.1 mM EDTA, 5 mM DTT, 10% glycerol, and 0.1 mg/ml BSA) was used as previously described [182]. For kinetic studies involving Pol β, buffer B (50
mM Tris-Cl, pH 7.8 at 37 °C, 5 mM MgCl2, 50 mM NaCl, 0.1 mM EDTA, 5 mM DTT, 10% glycerol, and 0.1 mg/ml BSA) was used. All kinetic experiments described in this paper were performed at 37 °C and the reported concentrations are final after mixing all the components.
Measurement of the kp and Kd for nucleotide incorporation A pre-incubated solution containing DNA polymerase in molar excess of the single- nucleotide gapped DNA substrate was mixed with increasing concentrations (0.25-1000 μM) of dNTP•Mg2+ in the appropriate buffer at 37 °C. Reactions catalyzed by Pol λ and Pol β contained a 4-fold [117] and 10-fold [210] molar ratio excess of enzyme:DNA, respectively. Aliquots of the reaction mixtures were quenched at various times using 0.37 M EDTA. A rapid-chemical quench flow apparatus (KinTek) was utilized for fast nucleotide incorporations with time intervals ranging from milliseconds to several minutes. Reaction products were resolved using sequencing gel electrophoresis (17% acrylamide, 8 M urea) and quantitated with a PhosphorImager 445 SI (Molecular Dynamics). The time course of product formation at each concentration of dNTP•Mg2+ was fit to a single-exponential equation (Equation 1)
[Product] = A[1 – exp(–kobst)] (1)
159 using a nonlinear regression program KaleidaGraph (Synergy Software) to yield an observed rate constant of nucleotide incorporation (kobs). The kobs values were then plotted against the corresponding concentrations of dNTP•Mg2+ and fitted using the hyperbolic equation (Equation 2)
kobs = kp[dNTP]/{[dNTP] + Kd} (2) 2+ which resolved the Kd and kp values for the dNTP•Mg incorporation.
Incorporation and ligation of 8-oxodGMP A pre-incubated solution of purified human Pol β (185 nM) in molar excess of the single- nucleotide gapped D-1 or D-8 (23 nM) was mixed with 8-oxodGTP•Mg2+ (100 μM) in buffer B at 37 °C. Five minutes after initiating the polymerization reaction, 10 nM human ∆235 DNA Ligase I (generous gift from the laboratory of Dr. Tom Ellenberger) and 1 mM ATP (MBI Fermentas) were added directly to the polymerization reaction mixture. Aliquots were quenched at various times (5, 30, and 60 minutes) by adding them to 0.37 M EDTA and immediate heat denaturation for 2 minutes at 95 °C. The DNA products were resolved by sequencing gel electrophoresis and visualized using autoradiography.
7.3 Results
Our previous pre-steady state kinetic studies confirm the application of single-turnover techniques towards elucidating the kinetic mechanism of Pol λ [117, 182, 192]. Thus, the experiments in this paper were performed with human Pol λ in molar excess over DNA to allow the direct observation of nucleotide incorporation in a single pass of the reactants through the enzymatic pathway without complications resulting from the steady-state formation of products [63].
Single-turnover kinetic analysis of 8-oxodGTP incorporation Nucleotide pools are susceptible to oxidative DNA damage induced by reactive oxygen species, therefore, it is possible that a DNA polymerase may encounter and subsequently
160 incorporate 8-oxodGTP in vivo, particularly under conditions of severe oxidative stress and/or a disfunctional 8-oxodGTPase [275]. Our results reinforce the slow incorporation of 8-oxodGTP into undamaged single-nucleotide gapped D-DNA (Figure 7.2) by human Pol β (Figure 7.3) and Pol λ [263, 276]. To determine which undamaged D-DNA template Pol λ would preferentially incorporate 8-oxodGTP, we employed pre-steady state kinetic techniques to measure the substrate specificity (kp/Kd) of this oxidized nucleotide under single-turnover conditions.
To initiate the reaction, a pre-incubated solution of 30 nM 5′-[32P]-labeled D-1 (Figure 7.2) and 120 nM Pol λ was reacted with increasing concentrations of 8-oxodGTP in buffer L (Section 7.2) at 37 °C. The extended DNA product and unreacted primer (21- mer) were separated by sequencing gel electrophoresis and quantitated with a PhosphorImager. A plot of product formation versus reaction time intervals was fit to
equation 1 (Section 7.2) which yielded a kobs value for each concentration of 8-oxodGTP (Figure 7.4 part (A)). These single-turnover rates were plotted against the 8-oxodGTP concentrations, and after fitting the plot to equation 2 (Section 7.2), a kp of 0.049 ± 0.002 -1 s for the maximum 8-oxodGTP incorporation rate constant and a Kd of 9 ± 1 µM for 8- oxodGTP binding were obtained (Figure 7.4 part (B)). Finally, the substrate specificity
(kp/Kd) of 8-oxodGTP incorporation opposite template base dA (D-1) was calculated to be 5.6 × 10-3 µM-1s-1 (Table 7.1). Similarly, the kinetic parameters for 8-oxodGTP incorporation into dG (D-6), dT (D-7), and dC (D-8) (Figure 7.2) were measured and
listed in Table 7.1. Interestingly, both the calculated kp/Kd values and efficiency ratios indicated that 8-oxodGTP incorporation occurred most efficiently against template dA followed by dC, dG, and dT. The calculated probability of incorporating 8-oxodGTP opposite dC (2.8%) was dramatically lower than opposite dA (96.6%) (Table 7.1).
Steady-state kinetic analysis determined the incorporation efficiency (kcat/Km) for dCTP and dATP opposite 8-oxodG and the incorporation of 8-oxodGTP against dA and dC catalyzed by human Pol β [263], but the steady-state kinetic parameters (kcat and Km) are complex functions of many microscopic rate-constants, especially the DNA dissociation
161 rate constant [63]. Therefore, these cannot fully evaluate the kinetic effect of 8-oxodG as an incoming nucleotide nor as a template base. To improve the mechanistic understanding of 8-oxodGTP incorporation by Pol β, we utilized pre-steady state kinetic methodology to measure the kinetic parameters (Table 7.2) of 8-oxodGTP with all four template bases in parallel single-turnover experiments (data not shown). The substrate specificity, nucleotide incorporation efficiency ratio, and probability values all suggested that Pol β, like Pol λ, preferentially incorporated 8-oxodGTP opposite template base dA (Table 7.2).
Ligation of 8-oxodG in DNA primer To complete the BER pathway, a ligation step catalyzed by either DNA ligase III/XRCC1 or DNA ligase I is required for phosphodiester bond formation. It has been previously shown that human DNA ligases I, IIIα, or IIIβ can seal the nick when an 8-oxodG template lesion is paired with either dA or dC [277]. Although DNA polymerases may incorporate 8-oxodGTP, it has not been determined if this structurally-altered nucleotide base can be subsequently ligated to the downstream DNA strand by a DNA ligase. Perhaps inefficient ligation may safeguard the integrity of genomic DNA. This possibility was qualitatively examined in vitro by allowing Pol β, the predominant BER polymerase, to fill the gap opposite template dA or dC with 8-oxodGTP. Then, recombinant ∆235 human DNA ligase I [278] and ATP were added to the reaction mixture. Full-length reaction products (41-mer) were obtained for both of the DNA substrates with 8- oxodGMP paired with template dA or dC at various reaction times, although, the reaction appeared to be less than 100% efficient for both of the DNA substrates (Figure 7.3). Nonetheless, human DNA ligase I was catalytically active on these mutagenic DNA substrates, suggesting that the ligation step cannot prevent integration of this oxidatively- damaged substrate into DNA. Notably, Pol β performed two sequential 8-oxodGTP incorporations into D-8 within 5 min. Perhaps Pol β preferentially extends from an 8- oxodGMP:dC base pair. Meanwhile, the resulting 23-mer product could not be ligated to the downstream 19-mer strand as observed by the unchanged intensity level of the 23- mer throughout the 60 min incubation (Figure 7.3).
162 Pre-steady state kinetic analysis of single-nucleotide incorporation opposite 8-oxodG Since Pol λ and Pol β exhibited similar template base selectivity when incorporating 8- oxodGTP (Tables 7.1 and 7.2), it is plausible that these structurally-homologous, X- family polymerases may possess similar nucleotide preference when bypassing an 8- oxodG template lesion in O-6 DNA (Figure 7.2). To evaluate this hypothesis, we determined the pre-steady state kinetic parameters (Tables 7.3 and 7.4) in a manner similar to those described in Figure 7.4 for all four nucleotide incorporations into O-6 (Figure 7.2) as individually catalyzed by Pol λ and Pol β (data not shown). In Table 7.3,
the Kd for each of the four nucleotides, both correct and incorrect, to the Pol λ•O-6 binary -1 complex remained comparable (1.2-9 μM). However, the kp values (0.00071-0.91 s ) differed by approximately three orders of magnitude, leading to a large range of substrate specificities (9.0 × 10-5-0.68 μM-1s-1). The calculated efficiency ratios and probabilities indicated that Pol λ predominantly selected dCTP and dATP when bypassing 8-oxodG, and more interestingly, these two nucleotides were incorporated with almost equal efficiency and probability (Table 7.3).
Pol β incorporated each of the four nucleotides against 8-oxodG with a relatively small -1 range of Kd values (11-660 μM) and a relatively large range of kp values (0.043-7.5 s ), thus resulting in a proportionately large range of substrate specificities (6.5 × 10-5-0.58 μM-1s-1) (Table 7.4). The calculated nucleotide incorporation efficiency ratios and probabilities in Table 7.4 indicated that Pol β, unlike Pol λ, preferentially incorporated dCTP opposite 8-oxodG, although efficient incorporation of dATP was also possible. The ~2:1 selection of dCTP:dATP opposite a template 8-oxodG was consistent with the published steady-state kinetic analysis [263] and X-ray crystal structure studies of Pol β [252].
163 7.4 Discussion
Pre-steady state kinetic methodologies were applied to determine which template base human Pol λ would most efficiently incorporate 8-oxodGTP and which nucleotide would preferentially be incorporated opposite 8-oxodG. With regard to in vivo relevance, we conducted analogous assays using human Pol β, the X-family homolog of Pol λ. Our pre-
steady state kinetic parameters (kp and Kd) and their corresponding values with undamaged DNA and dNTPs (Tables 7.1-7.4) kinetically illustrated how the commonly oxidized-nucleotide base dG adversely affects DNA synthesis catalyzed by two gap- filling DNA polymerases in vitro and possibly in vivo.
Effect of 8-oxodGTP on the kinetics of gap-filling synthesis catalyzed by Pol λ and Pol β We have previously determined the pre-steady state kinetic parameters (Tables 7.1 and 7.3) for Pol λ incorporating all normal dNTPs into D-DNA (Figure 7.2) [117]. In Table
7.1, the Kd range (5-23 μM) for the binding of 8-oxodGTP to the Pol λ•DNA binary complex remained comparable to the binding of undamaged dGTP (2.1-7 μM) [117]. The kp for the incorporation of 8-oxodGTP versus dGTP opposite template dC was dramatically reduced (2.5 → 0.0008 s-1) by 3,000-fold while the maximum rate of incorporation opposite template dA increased (0.00040 → 0.049 s-1) by 100-fold (Table
7.1). Hence, variations in the kp produced the most significant changes in the substrate specificity. Most notably, a comparison of the substrate specificities for 8- oxodGTP:dC and dGTP:dC revealed a 7,500-fold difference for these Watson-Crick base pairing partners in the anti conformation. Incorporation of 8-oxodGTP was most efficient opposite template dA as indicative of the 97% insertion probability (Table 7.1).
Relative to the pre-steady state kinetic results of rat DNA polymerase β with undamaged single-nucleotide gapped DNA [114], the weaker ground-state binding affinities of 8- oxodGTP (70-570 μM) to the human Pol β•DNA binary complex for all the DNA template bases resembled the mismatches for dGTP (360-1600 μM) as listed in Table 7.2. Kinetic comparisons between human and rat Pol β are suitable because they share 95%
164 sequence identity and possess almost identical polymerase efficiency [274]. Like Pol λ, a
similar trend for the kp results emerged: there is an approximate 20-fold increase and decrease for templates dA and dC, respectively. Comparing the substrate specificities for 8-oxodGTP:dA and dGTP:dA revealed a 410-fold difference, therefore, the likelihood of Pol β creating an A:T → C:G transversion is enhanced with 8-oxodGTP. Additionally, the binding of 8-oxodGTP likely occurs in the energetically favored syn conformation as supported by this kinetic study and earlier studies examining Pol β, T7 DNA polymerase, Dbh, Dpo4, and human pol η [263, 269, 279]. However, structural studies of an E•DNA•8-oxodGTP ternary complex are necessary to verify this kinetics-based hypothesis.
Oxidized dGTP pools are substrates for DNA polymerases if an 8-oxodGTPase (e.g. E. coli MutT or MTH1) fails to hydrolyze it to 8-oxodGMP. Although 8-oxodGTPases are quite efficient enzymes, it is possible that intracellular 8-oxodGTP levels may increase under conditions of severe oxidative stress and/or a disfunctional 8-oxodGTPase [275]. The selective incorporation of 8-oxodGTP opposite template dA and dC catalyzed by Pol λ and Pol β (Tables 7.1 and 7.2) followed by the effective ligation of incorporated 8- oxodGMP to a downstream strand catalyzed by human DNA ligase I (Figure 7.3) may lead to A:T → C:G and G:C → T:A transversions, respectively, if 8-oxodGMP is not repaired in vivo.
Effect of template lesion 8-oxodG on the kinetics of gap-filling synthesis catalyzed by Pol λ and Pol β When comparing the kinetic data obtained for Pol λ with damaged (O-6) and undamaged (D-6) DNA [117], the substrate specificity for dCTP decreased by ~2-fold (Table 7.3). Interestingly, the 8-oxodG lesion selectively enhanced the misincorporation efficiency of dATP by 20,600-fold (Table 7.3). This efficiency change was due to a dramatic increase (8,100-fold) in the maximum incorporation rate constant, although, the ground-state binding affinity (1/Kd) also increased by ~2-fold. Consequently, the probability of inserting dCTP dropped from essentially 100% with D-6 to approximately 50% with O-6 (Table 7.3). This result supports 8-oxodG as a true dual-coding base which equally 165 instructs the incorporation of both dCTP and dATP at the active site of Pol λ. Furthermore, the 2- to 3-fold lower substrate specificities of dCTP and dATP with O-6 compared to matched dCTP with undamaged D-6 strongly suggest that bypassing the 8-oxodG lesion, although mutagenic, will not significantly hinder short-patch BER catalyzed by human Pol λ.
For polymerization by human Pol β, the oxidation of dG to 8-oxodG led to a dramatic
increase (400-fold) in the kp of mismatched dATP incorporation as well as a 10-fold increase and a 5-fold decrease in the apparent ground-state binding affinity of dATP and dCTP, respectively, when compared with undamaged single-nucleotide gapped DNA
(Table 7.4) [114]. Coupling the contributions from both the kp and Kd enhanced the substrate specificity 4,000-fold for mismatched dATP incorporation opposite 8-oxodG compared to dG. Despite the remarkable kinetic effect, the probability of inserting dCTP was only reduced from essentially 100% with D-6 to approximately 70% with O-6 which is in contrast to the Pol λ results (Tables 7.3 and 7.4). Nonetheless, the incorporation of dATP was only 2-fold less efficient than dCTP into O-6 and as such the mutagenic potential of G:C → A:T transversions persists.
In vivo, a specific DNA glycosylase (e.g. E. coli MutY or human hMYH) is known to selectively remove the adenine base from an 8-oxodG:dA mismatch, leading to the formation of DNA gaps [280], like the one in O-6 (Figure 7.2). Either Pol β or Pol λ may be recruited to fill these gaps opposite 8-oxodG, for both of them exhibit a protective effect against DNA oxidizing agents in vivo [173, 191, 223]. If recruited, this will be problematic, especially to the short-patch BER pathway, because (i) Pol β is known to
function in short-patch BER [76, 77] while Pol λ is likely a secondary polymerase in this DNA repair pathway [78]; (ii) both polymerases are error prone when synthesizing opposite a single 8-oxodG gap, especially Pol λ which incorporated both dATP and dCTP with almost equal probability and only slightly lower efficiency than a correct dCTP into normal DNA (Table 7.3); and (iii) the single-nucleotide gap-filling efficiency of Pol λ was more efficient than Pol β due to a mere 2-fold versus 10-fold decrease in the
166 substrate specificity of matched dCTP against 8-oxodG versus dG (Tables 7.3 and 7.4). Furthermore, the ligation step of the BER pathway compounds the mutagenic potential as a previous study shows human DNA ligases I, IIIα, or IIIβ can efficiently seal the nick when an 8-oxodG template lesion is paired with either dA or dC [277].
Three classes of polymerases when bypassing 8-oxodG A collection of DNA polymerases from the A-, B-, X-, Y-, and RT-families have been evaluated previously for their nucleotide selection when polymerizing over 8-oxodG. Both dCTP and dATP are much more efficient substrates than dGTP and dTTP for almost all of these polymerases. Based on their 8-oxodG coding potential with dCTP and dATP, we propose to divide polymerases into three classes: (i) dCTP:dATP ratio > 10: Dpo4 [256, 281], yeast DNA polymerase η [264, 267], RB69 DNA polymerase [253]; (ii)
dCTP:dATP ratio 1-10: E. coli DNA polymerase I (exo-) [259, 261], bovine DNA polymerase δ in the presence of human PCNA [266], human polymerase ι [265], human
Pol β [263], T7 DNA polymerase (exo-) [257, 262], E. coli DNA polymerase II (exo-) [261] and human Pol λ; (iii) dCTP:dATP < 1: yeast DNA polymerase ζ [23], calf thymus
DNA polymerases α [259] and δ [259], BF [254], and HIV-1 RT [262] (Table 7.5). These defined classes correspond to the mode of 8-oxodG-lesion bypass. For example, class i exhibits predominantly error-free bypass while classes ii and iii, the majority of DNA polymerases that have been kinetically characterized, are mutagenic in the presence of an 8-oxodG template. Lastly, we found no direct correlation between polymerase families and the above classes in which the extension of 8-oxodG bypassed products was not considered.
Asymmetric active sites of both Pol λ and Pol β The discrepancy for Pol λ to exhibit a strong preference for template dA when incorporating 8-oxodGTP compared to the equivalent preference for dATP and dCTP when opposite template base 8-oxodG indicated that the polymerase active site of Pol λ is asymmetrical. This similar yet contradictory base pairing scheme for Pol β coincided
with Miller et al. [263] and reiterated that the active site of human Pol β, like Pol λ, is
167 also asymmetrical. These findings were also supported by the fact that the formation of 8- oxodGTP:dA (or dC) was much less efficient than dATP (or dCTP):8-oxodG based on
their kp/Kd values (Tables 7.1-7.4). The substrate specificities, predominantly affected by a change in the kp, for even the most favored incorporation of 8-oxodGTP against dA were 100- and 200-fold lower for Pol λ and Pol β relative to their substrate specificities for dGTP opposite dC (Tables 7.1 and 7.2) [117, 263]. Hence, 8-oxodGTP was not as kinetically competent as a normal and correct dNTP substrate for these two polymerases as well as others [263, 269], and this may be one way to attenuate the mutagenicity of 8- oxodGTP in vivo.
Structural basis for the mutagenic potential of 8-oxodG Based on our above analysis, the 8-oxodG template lesion exhibited dual coding potential during gap-filling synthesis catalyzed by both Pol λ and Pol β. Several structural factors influencing the syn-anti equilibrium govern whether dCTP or dATP will be incorporated more efficiently against 8-oxodG in the polymerase active site as revealed by the X-ray crystal structure studies of the binary and ternary structures for five different DNA polymerases [252-257, 281]. First, steric clashes between the phosphodeoxyribose backbone and 8-oxodG, particularly in the anti conformation, are minimized by three DNA template-distorting modes: (i) sharp kinks in the active site of T7 DNA polymerase [255], BF [254], and bacteriophage RB69 DNA polymerase [253]; (ii) 180° rotation of
the 5′ phosphate in the active sites of Pol β [252] and Dpo4 [281]; and (iii) repositioning
active site residues, e.g. K280 of Pol β [252] and Q797 of BF [254]. Second, the dA:8- oxodG mismatch evades intrinsic proofreading mechanisms of T7 DNA polymerase
[255] and BF [254] because the minor groove resembles a correct dA:dT base pair. Third, hydrogen bonding between the C8 oxygen and amino acid side chain residues, e.g. R332 of Dpo4 [256, 281], may stabilize the anti conformation to favor dCTP incorporation [255-257, 281]. Based upon these structural observations, Pol λ may utilize some unique combination to non-discriminatively incorporate both dATP and dCTP against 8-oxodG.
168 Because tPol λ shares homology with Pol β (Figure 7.1) [74, 270] and their ternary structures with gapped DNA and an incoming nucleotide superimpose well with a root- mean-square deviation of 1.4 Å for 113 C-α atoms [55, 119, 237], it is reasonable that the 5′-phosphodeoxyribose backbone of the templating 8-oxodG may flip at the active site of Pol λ as observed with Pol β [252]. When the undamaged DNA and an incoming ddTTP
from the ternary structure of human tPol λ [55] were swapped with the gapped DNA containing 8-oxodG and dCTP substrates from the Pol β ternary structure [252], the dCTP:8-oxodG base pair was well accommodated within the active site of tPol λ (Figure 7.5 part (A)). However, our single-turnover kinetic analysis has demonstrated that Pol λ, unlike Pol β, incorporated both dATP and dCTP against 8-oxodG with equal efficiency. Krahn et al. proposes that the anti-syn equilibrium of 8-oxodG may be shifted towards the anti conformation due to the flexibility of the DNA template backbone in the active site of Pol β [252]. If this is true, certain structural factors in Pol λ have to counteract such DNA conformational perturbations targeting the template by Pol β in order to stabilize the nascent base pair dATP:8-oxodG at the active site of Pol λ. Sequence alignment analysis [200, 240] and X-ray crystal structures [55, 119] indicate that the residues surrounding the nascent base pair in human Pol β (K27, R40, A185, D276, and K280) are analogous to S268, A280, K422, A510, and R514 in human Pol λ, respectively. To accommodate 8-oxodG in the anti conformation and form a Watson- Crick base pair with an incoming dCTP at the active site of Pol β, the ternary crystal structure of Pol β [252] shows that only K280 significantly repositions itself and forms hydrogen bonding with N37. Interestingly, the ternary structure of tPol λ, undamaged
DNA, and ddTTP [55] reveals a spacious environment for the repositioning of R514 in Pol λ, the counterpart of K280 in Pol β, to accommodate 8-oxodG in the anti conformation. However, this reorientation will not be stable in Pol λ because the corresponding residue of N37 in Pol β is L277 in Pol λ, which precludes hydrogen bonding with R514 through side chain interactions. Thus, 8-oxodG in the anti conformation at the active site of Pol λ may not be as stable as it is at the active site of Pol β, leading to a slight shift of the anti-syn equilibrium of 8-oxodG towards the syn- conformation.
169 To further examine the above hypothesis, we modeled base pair dATP:8-oxodG directly from the ternary structure of the K536A mutant of T7 DNA polymerase [257] into the ternary structure of tPolλ [55] (Figure 7.5 part (B)). The model showed no evidence of steric hindrance from the tPol λ active site, but rotations at the N9-C1′ and C4′-C5′ bonds of 8-oxodG were necessary to overlay the 5′-phosphodeoxyribose backbone of 8-oxodG with the counterpart of template base dA of the undamaged DNA substrate. Interestingly, R514 was sufficiently close to form a hydrogen bond with the C6 oxygen, which may be enough to compensate for the energy cost of the aforementioned rotations and to stabilize the syn conformation of 8-oxodG. Also notable, similar hydrogen-bonding between residue R332 and the C8 oxygen has been found to stabilize the anti conformation of 8- oxodG at the active site of Dpo4 which favors dCTP incorporation against 8-oxodG [256, 281]. Thus, the balanced contribution from these structural factors likely coordinated a true equilibrium between the anti and syn conformations of 8-oxodG in the active site of Pol λ. However, further structural and biochemical studies are required to verify these predicted structure-function relationships in Pol λ, since the polymerase active site played a major role in kinetically modulating the mutagenic potential of 8-oxodG and 8- oxodGTP.
In conclusion, this single-turnover kinetic analysis established the dual coding potential of 8-oxodG as a template base at the active sites of both Pol λ and Pol β in which lesion bypass occurred with relatively high efficiency but low accuracy. These two enzymes also favored the incorporation of 8-oxodGTP against dA. Surprisingly, our results suggested 8-oxodG, if not repaired, will cause mutagenic transversions during polymerization and ligation in the BER process.
7.5 Future Directions
Soon after the completion of this study, Maga et al. showed the bypass of 8-oxodG on a primer-template DNA substrate to be relatively error-free when Pol λ is in the presence of human proliferating cell nuclear antigen (PCNA) and replication protein A (RP-A) [282]. 170 However, this study did not determine the kinetic basis of the enhanced polymerase fidelity of Pol λ. In addition, the work herein only examined 8-oxodG bypass for short- patch base excision repair, although, the process may proceed through a long-patch base excision repair pathway. Therefore, using the knowledge gained from Chapter 6, transient state kinetic techniques could be applied to establish a comprehensive mechanism of Pol λ, in the presence of PCNA and/or RP-A, bypassing site-specific 8-oxodG lesions in the templates of gapped DNA substrates with various gap widths.
171 7.6 Tables
Table 7.1 Kinetic parameters of nucleotide incorporation into single-nucleotide gapped DNA catalyzed by human Pol λ at 37 °C. DNA K k kp/Kd Efficiency template d p Probabilityb (μM) (s-1) -1 -1 ratioa base (μM s ) 8-oxodGTP dC 5 ± 7 0.0008 ± 0.0003 1.6 × 10-4 1 2.8% dA 9 ± 1 0.049 ± 0.002 5.6 × 10-3 35 96.6% dG 15 ± 12 0.0005 ± 0.0002 3.6 × 10-5 0.23 0.6% dT 23 ± 8 (7.5 ± 0.7) × 10-6 3.3 × 10-7 2.1 × 10-3 ~0% dGTPc dC 2.1 ± 0.3 2.5 ± 0.1 1.2 1 99.9% dA 3.2 ± 0.5 0.00040 ± 0.00002 1.3 × 10-4 1.1 × 10-4 ~0% dG 4 ± 1 0.00020 ± 0.00001 5.0 × 10-5 4.2 × 10-5 ~0% dT 7 ± 4 0.010 ± 0.002 1.4 × 10-3 1.2 × 10-3 0.1% a Calculated as (kp/Kd)dN/(kp/Kd)dC. b Calculated as {(kp/Kd)/[∑(kp/Kd)]} × 100. c The kp and Kd values for dGTP are from reference [117].
172
Table 7.2 Kinetic parameters of nucleotide incorporation into single-nucleotide gapped DNA catalyzed by human Pol β at 37 °C. DNA K k kp/Kd Efficiency template d p Probabilityb (μM) (s-1) -1 -1 ratioa base (μM s ) 8-oxodGTP dC 230 ± 40 0.90 ± 0.06 3.9 × 10-3 1 8.7% dA 70 ± 10 2.8 ± 0.2 4.1 × 10-2 10.5 91.3% dG 400 ± 40 0.00139 ± 0.00006 3.5 × 10-6 9.0 × 10-4 ~0% dT 570 ± 80 0.0014 ± 0.0001 2.5 × 10-6 6.4 × 10-4 ~0% dGTPc dC 3.6 ± 0.8 18.4 ± 0.8 5.1 1 99.97% dA 1600 ± 300 0.16 ± 0.05 1.0 × 10-4 2.0 × 10-5 ~0% dG 620 ± 150 0.96 ± 0.01 1.5 × 10-3 2.9 × 10-4 0.03% dT 360 ± 40 0.040 ± 0.002 1.1 × 10-4 2.2 × 10-5 ~0% a Calculated as (kp/Kd)dN/(kp/Kd)dC. b Calculated as {(kp/Kd)/[∑(kp/Kd)]} × 100. c The kp and Kd values for dGTP are from reference [114] which examines rat Pol β.
173 Table 7.3 Kinetic parameters of nucleotide incorporation into single-nucleotide gapped DNA catalyzed by human Pol λ at 37 °C.
K k kp/Kd Efficiency dNTP d p Probabilityb (μM) (s-1) (μM-1s-1) ratioa Template 8-oxodG (O-6) dCTP 1.3 ± 0.3 0.91 ± 0.04 0.70 1 50.7% dATP 1.2 ± 0.3 0.81 ± 0.05 0.68 0.97 49.3% dGTP 4 ± 1 0.00071 ± 0.00006 1.8 × 10-4 2.6 × 10-4 ~0% dTTP 9 ± 2 0.00081 ± 0.00007 9.0 × 10-5 1.3 × 10-4 ~0% Template dG (D-6)c dCTP 0.9 ± 0.1 1.57 ± 0.04 1.8 1 99.98% dATP 3 ± 1 0.00010 ± 0.00004 3.3 × 10-5 1.8 × 10-5 ~0% dGTP 4 ± 1 0.00020 ± 0.00001 5.0 × 10-5 2.8 × 10-5 ~0% dTTP 2.5 ± 0.3 0.00070 ± 0.00001 2.8 × 10-4 1.6 × 10-4 0.02% a Calculated as (kp/Kd)dNTP/(kp/Kd)dCTP. b Calculated as {(kp/Kd)/[∑(kp/Kd)]} × 100. c The kp and Kd values for template dG (D-6) are from reference [117].
174
Table 7.4 Kinetic parameters of nucleotide incorporation into single-nucleotide gapped DNA catalyzed by human Pol β at 37 °C.
K k kp/Kd Efficiency dNTP d p Probabilityb (μM) (s-1) (μM-1s-1) ratioa Template 8-oxodG (O-6) dCTP 11 ± 4 6.4 ± 0.7 0.58 1 67.4% dATP 27 ± 6 7.5 ± 0.7 0.28 0.48 32.6% dGTP 660 ± 70 0.043 ± 0.002 6.5 x 10-5 1.1 x 10-4 ~0% dTTP 400 ± 100 0.046 ± 0.005 1.2 x 10-4 2.1 x 10-4 ~0% Template dGc dCTP 1.9 ± 0.2 12.5 ± 0.5 6.6 1 ~100% dATP 270 ± 40 0.019 ± 0.001 7.0 x 10-5 1.1 x 10-5 ~0% dGTP 360 ± 40 0.040 ± 0.002 1.1 x 10-4 1.7 x 10-5 ~0% dTTP 650 ± 300 0.37 ± 0.09 5.7 x 10-4 8.6 x 10-4 ~0% a Calculated as (kp/Kd)dNTP/(kp/Kd)dCTP . b Calculated as {(kp/Kd)/[∑(kp/Kd)]} × 100. c The kp and Kd values for template dG are from reference [114] which examines rat Pol β.
175
Table 7.5 Class assignments for DNA polymerases based upon dCTP:dATP incorporation ratios opposite template base 8-oxodG.
dCTP:dATP DNA Polymerase dNTP incorporation Kinetic method Reference(s)
Class polymerase family preference ratio
Dpo4 Y dCTP 91, 73 steady-state [256, 281] steady-state and Yeast Pol η Y dCTP 68, 19 [264, 267] pre-steady state
Class i RB69 DNA B dCTP 20 steady state [253] polymerase E. coli Pol Ia A dCTP 7.4, 4.0 steady state [259, 261] Bovine Pol δb B dCTP 3.2 steady-state [266] Pol ι Y dCTP 2.1 steady-state [265] steady-state and [263]; Pol β X dCTP 2.1, 1.8 pre-steady state this chapter T7 DNA a A dCTP 1.8 steady-state [262]
Class ii polymerase E. coli DNA dCTP or B 1 steady-state [261] polymerase IIa dATP dCTP or Human Pol λ X 1 pre-steady state this chapter dATP Yeast Pol ζ B dATP 0.71 steady-state [23] Bovine Pol δ B dATP 0.44 steady-state [266] Bovine Pol α B dATP 0.14 steady-state [259]
Class iii BF A dATP 0.11 steady-state [254] HIV-1 RT RT dATP 0.075 steady-state [262] aexonuclease deficient mutant bassayed in complex with human PCNA
176 7.7 Figures
Figure 7.1 Schematic representations of human Pol λ and Pol β. Each domain, with amino acid residue numbers indicated above, is shown as a rectangle. The N-terminal 35 residues of Pol λ containing a nuclear localization signal motif are represented as a line.
177
5’-CGCAGCCGTCCAACCAACTCA CGTCGATCCAATGCCGTCC-3’ D-DNA 3’-GCGTCGGCAGGTTGGTTGAGTXGCAGCTAGGTTACGGCAGG-5’ 5’-CGCAGCCGTCCAACCAACTCA GTCGATCCAATGCCGTCC-3’ O-6 3’-GCGTCGGCAGGTTGGTTGAGTYCAGCTAGGTTACGGCAGG-5’
Figure 7.2 Single-nucleotide gapped-DNA substrates 21-19/41-mer (D-DNA) and 21- 18/40-mer (O-6). The same 5′-[32P]-radiolabeled 21-mer and a 5′-phosphorylated downstream strand (18- or 19-mer) were annealed to the different templates (bottom strand, 40- or 41-mer) to create single-nucleotide gapped DNA substrates. In D-DNA, X represents A, C, G, or T for D-1, D-8, D-6, and D-7, respectively. In O-6, Y denotes 8- oxodG.
178
Figure 7.3 Autoradiographed gel image depicting the ligation of incorporated 8-oxodG to a downstream primer of single-nucleotide gapped DNA. A pre-incubated solution of 23 nM of 5′-[32P]-labeled D-1 or D-8 and 185 nM Pol β was reacted first with 100 µM 8- oxodGTP•Mg2+, then with 10 nM ∆235 human DNA ligase I and 1 mM ATP for indicated time intervals before being quenched and analyzed by sequencing gel electrophoresis. The DNA sizes are marked in the left margin.
179 A 30
25
20
15
10 Product (nM)
5
0 0 200 400 600 800 1000 1200
Reaction Time (s)
B 0.04
0.035
0.03
0.025
(1/s) 0.02 obs
k 0.015
0.01
0.005
0 0 5 10 15 20 25 30 35
8-oxodGTP (μM)
Figure 7.4 Concentration dependence on the pre-steady state rate constant of modified nucleotide incorporation. (A) A pre-incubated solution of Pol λ (120 nM) and 5′-[32P]- labeled D-1 (30 nM) was rapidly mixed with increasing concentrations of 8- oxodGTP•Mg2+ (0.5 μM, z; 1 μM, {; 2 μM, ; 4 μM, ; 8 μM, S; 16 μM, U; 32 μM, ) for various time intervals. The solid lines are the best fits to Equation 1 (Section 7.2) which determines the observed rate constants, kobs. (B) The kobs values were plotted as a function of 8-oxodGTP concentration. The data (z) were then fit to Equation 2 (Section -1 7.2), yielding a kp of 0.049 ± 0.002 s and a Kd of 9 ± 1 μM.
180 A
B
Figure 7.5 Model of 8-oxodG in the polymerase active site of human Pol λ. (A) DNA containing anti 8-oxodG and dCTP substrates (red) in the ternary structure of human Pol β (PDB entry 1MQ3) [252] replaced their counterparts in the ternary structure of the C- terminal Pol β-like domain of human Pol λ (PDB entry 1XSP, blue) [55]; (B) Model of syn 8-oxodG:dATP (red) from the ternary structure of the K536A mutant of T7 DNA polymerase (PDB entry 1ZYQ) [257] into the ternary structure (PDB entry 1XSP, blue) of human tPol λ fragment, undamaged DNA, and an incoming ddTTP [55]. The software Swiss PDB v.3.7 was used for the modeling.
181 Chapter 8: Pre-Steady State Kinetic Analysis of Truncated and Full-Length Saccharomyces cerevisiae DNA Polymerase Eta
8.1 Introduction
DNA polymerases are organized into seven families: A, B, C, D, X, Y, and reverse transcriptase [7, 82, 83]. Among these families, DNA polymerases are involved in DNA replication, DNA repair, DNA lesion bypass, antibody generation, and sister chromatid cohesion [5]. Despite these diverse roles, DNA polymerases catalyze the nucleotidyl transfer reaction using a two divalent metal ion mechanism [283] with at least one positively charged residue [198] that functions as a general acid [284] at their active site, follow a similar minimal kinetic pathway [54], and share a similar structural architecture consisting of the fingers, palm, and thumb domains [41, 54]. Surprisingly, the polymerization fidelity of eukaryotic DNA polymerases spans a wide range: one error per 0 -9 one to one billion nucleotide incorporations (10 to 10 ) [66].
The Y-family DNA polymerases are known for catalyzing nucleotide incorporation with low fidelity and poor processivity. These enzymes are specialized for translesion DNA synthesis which involves nucleotide incorporation opposite and downstream of a damaged DNA site. Lesion bypass can be either error-free or error-prone depending on the DNA polymerase and DNA lesion combination. To accommodate a distorted DNA substrate, Y-family DNA polymerases utilize several features: a solvent accessible [101] and conformationally flexible active site [103], smaller fingers and thumb domains [101], an additional domain known as the little finger [101], the little finger and polymerase core domains move in opposite directions during a catalytic cycle [62], and a lack of 3′ → 5′ exonuclease activity [6]. Unfortunately, these features, which facilitate
182 lesion bypass, may also contribute to the low fidelity of a Y-family DNA polymerase during replication of a damaged or undamaged DNA template. Thus, it is important to understand the mechanism and fidelity of Y-family DNA polymerases.
Saccharomyces cerevisiae DNA polymerase η (yPol η), a Y-family DNA polymerase, is critical for the error-free bypass of UV-induced DNA damage such as a cis-syn thymine- thymine dimer [33, 285-288]. To date, Pol η remains the only Y-family DNA polymerase with a confirmed biological function [36]. yPol η is organized into a polymerase domain, ubiquitin-binding zinc finger (UBZ) domain, and proliferating cell nuclear antigen (PCNA)-interacting peptide (PIP) motif (Figure 8.1). X-ray crystal structures of yPol η’s catalytic core have been solved alone [289] as well as in complex with a cisplatin-DNA adduct and an incoming nucleotide [290]. Due to a lack of structures for full-length yPol η, it is unclear if the C-terminal residues 514-632 interact with DNA and contribute to the polymerase function of yPol η. Using pre-steady state kinetic techniques, we have measured the base-substitution fidelity of full-length and truncated yPol η (Figure 8.1) catalyzing nucleotide incorporation into undamaged DNA. In addition, we have determined the DNA binding affinity of both full-length and truncated yPol η. Our results show that the C-terminus of yPol η has a minor effect on the DNA binding affinity and the base substitution fidelity of this lesion bypass DNA polymerase.
8.2 Materials and Methods
Materials Materials were purchased from the following companies: [γ-32P]ATP, MP Biomedicals (Solon, OH); Biospin columns, Bio-Rad Laboratories (Herclues, CA); dNTPs, GE Healthcare (Piscataway, NJ); oligodeoxynucleotides, Integrated DNA Technologies, Inc. (Coralville, IA); OptiKinase, USB (Cleveland, OH).
183 Preparation of substrates and enzymes The synthetic oligodeoxyribonucleotides listed in Table 8.1 were purified as described previously [111]. The primer strand 21-mer or blunt-end 16-mer was 5′-radiolabeled with [γ-32P]ATP and OptiKinase. Then, the 21-mer was annealed to the appropriate 41mer template (Table 8.1) and the palindromic blunt-end substrates were annealed as described previously [111]. The catalytic core of yPol η (1-513) containing an N-terminal
MGSSH6SSGLVPRGSH tag was purified as described previously [291]. The full-length yPol η (1-632) was expressed and purified from yeast [292]. Pyrene 5′-triphosphate (dPTP) was synthesized as described previously [194].
Pre-steady state kinetic assays All experiments were performed in reaction buffer A which contained 40 mM Tris-HCl
pH 7.5 at 23 °C, 5 mM MgCl2, 1 mM DTT, 10 μg/mL BSA, and 10% glycerol. A rapid chemical-quench flow apparatus (KinTek, PA, USA) was used for fast reactions. For burst assays, a pre-incubated solution of yPol η (320 nM) and 5′-[32P]-labeled D-1 DNA 2+ (480 nM) was mixed with dTTP•Mg (100 μM). To measure the dissociation rate of the yPolη•DNA binary complex, a pre-incubated solution of yPol η (50 nM) and 5′-[32P]- labeled D-1 DNA (100 nM) was mixed with a molar excess of unlabeled D-1 DNA (2.5 μM) for various time intervals prior to initiating the polymerization reaction with 2+ dTTP•Mg (150 and 400 μM for truncated and full-length yPol η, respectively) for 15 s. For single-turnover kinetic assays, a pre-incubated solution of yPol η (150 nM) and 5′- 32 2+ [ P]-labeled DNA (30 nM) was mixed with an incoming dNTP•Mg (0.4-800 μM). Reactions were quenched at the designated time by adding 0.37 M EDTA. Reaction products were analyzed by sequencing gel electrophoresis (17% acrylamide, 8M urea, 1× TBE running buffer), visualized using a Typhoon TRIO (GE Healthcare), and quantitated with ImageQuant software (Molecular Dynamics).
DNA binding assays DNA The equilibrium dissociation constant (Kd ) of the yPol η•DNA binary complex was determined using two techniques. First, an electrophoretic mobility shift assay (EMSA)
184 was employed by adding increasing concentrations of yPol η (10 - 450 nM) into a fixed concentration of 5′-[32P]-labeled D-1 DNA (10 nM) in buffer A. The solution established equilibrium during a 20 min incubation period. Then, the binary complex was separated from unbound DNA using a 4.5% native polyacrylamide gel and running buffer as previously described except the final concentration of Tris was adjusted to 40 mM [194]. Second, a fluorescence titration assay was used. Increasing concentrations of yPol η (2- 300 nM) were titrated into a fixed concentration of F-8 DNA (25 nM) in buffer A (devoid of BSA). The F-8 DNA substrate (Table 8.1) was excited at a wavelength of 312 nm with emission and excitation slit widths of 5 nm. The emission spectra were collected at 1 nm intervals from 320-500 nm using a Fluoromax-4 (Jobin Jvon Horiba). Emission background from the buffer and intrinsic protein fluorescence were subtracted from each spectrum.
Data Analysis For the pre-steady state burst assay, the product concentration was graphed as a function of time (t) and the data were fit to the burst equation (Equation 1)
[Product] = A[1 – exp(–k1t) + k2t] (1) using the non-linear regression program, KaleidaGraph (Synergy Software). A represents
the fraction of active enzyme, k1 represents the observed burst rate constant, and k2 represents the observed steady-state rate constant. Data for the EMSA were graphed by plotting the concentration of the binary complex as a function of enzyme concentration
(E0) and fitting it to a quadratic equation (Equation 2). DNA DNA 2 1/2 [E•DNA] = 0.5(Kd + E0 + D0) – 0.5[(Kd + E0 + D0) – 4E0D0] (2)
D0 is the DNA concentration. For the fluorescence titration experiments, a modified quadratic equation (Equation 3) DNA DNA 2 1/2 [F] = Fmax + [(Fmin – Fmax)/(2D0)]{( Kd + E0 + D0) – [(Kd + E0 + D0) – 4E0D0] } (3) was applied to a plot of the fluorescence intensity (F) measured at 370 nm versus enzyme concentration. Fmax and Fmin represent the maximum and minimum fluorescence intensity,
185 respectively. For the rate of DNA dissociation from the binary complex, a single- exponential equation (Equation 4)
[Product] = A[exp(–kofft)] + C (4) was applied to a plot of product concentration versus time. A represents the reaction
amplitude, koff is the observed rate constant of DNA dissociation, and C is the concentration of the radiolabeled DNA product in the presence of a DNA trap for unlimited time. For the single-turnover kinetic assays, a plot of product concentration versus time was fit to a single-exponential equation (Equation 5)
[Product] = A[1 – exp (–kobst)] (5)
to extract the observed rate constant of nucleotide incorporation (kobs). To measure the
maximum rate constant of incorporation (kp) and the apparent equilibrium dissociation
constant (Kd) of an incoming nucleotide, the extracted kobs values were plotted as a function of nucleotide concentration and fit to a hyperbolic equation (Equation 6).
[kobs] = kp[dNTP]/(Kd + [dNTP]) (6) The free energy change (ΔΔG) for a correct and incorrect nucleotide substrate dissociating from the E•DNA•dNTP complex was calculated according to equation 7.
ΔΔG = RTln[(Kd)incorrect /(Kd)correct] (7) Here, R is the universal gas constant and T is the reaction temperature in Kelvin.
8.3 Results and Discussion
Truncated and full-length yPol η display biphasic kinetics Previously, transient state kinetic techniques have been used to characterize full-length yPol η at 30 °C [28]. Therefore, we first performed a burst assay (Section 8.2) to ensure that our purified proteins, truncated and full-length yPol η (Figure 8.1), behaved in a similar manner at 23 °C. Compared to wild-type yPol η, the truncated construct contains only the polymerase domain (Figure 8.1). A pre-incubated solution of yPol η (320 nM) 32 2+ and 5′-[ P]-labeled 21/41mer D-1 DNA (480 nM) was mixed with dTTP•Mg (100 μM) and quenched with EDTA at various times. Product concentration was plotted as a
186 function of time and was fit to equation 1, since there were two distinct kinetic phases: a rapid, exponential phase and a slow, linear phase (data not shown). These burst results were similar to those previously published [115]. Biphasic kinetics of nucleotide incorporation indicated that the first turnover rate was the rate of nucleotide incorporation occurring at the enzyme’s active site while subsequent turnovers (i.e. linear phase) were likely limited by the DNA product release step as demonstrated by full-length yPol η at 30 °C [115] and other DNA polymerases [60, 110, 111].
The C-terminal 119 residues slightly enhance DNA binding affinity of yPol η DNA The equilibrium dissociation constant for the binary complex of yPol η•DNA (Kd ) was measured to determine if the C-terminus of yPol η affects DNA binding affinity DNA (Scheme 8.1). First, the Kd was estimated using the EMSA (Section 8.2). For example, varying concentrations of full-length yPol η (10-450 nM) were incubated with a fixed concentration of 5′-[32P]-labeled D-1 DNA (10 nM) before separating the binary complex from the unbound DNA on a native gel (Figure 8.2 part (A)). Then, a quadratic equation (Equation 2) was applied to a plot of the binary complex concentration versus yPol η DNA concentration which resolved a Kd of 16 ± 1 nM (Figure 8.2 part (B) and Table 8.2). DNA Under similar reaction conditions, the Kd of truncated yPol η was estimated to be 34 ± DNA 3 nM; a binding affinity (1/Kd ) value that is 2-fold weaker than that of full-length yPol η (Table 8.2).
DNA DNA To corroborate these estimated Kd values, we measured the true Kd for the yPol η•DNA complex using a fluorescence titration assay. An analog of dA, 2-aminopurine, was embedded into the 41mer template of F-8 DNA which is identical to 21/41mer D-8 DNA except 2-aminopurine flanks the 5′ end of the templating dC base (Table 8.1). The F-8 DNA substrate (25 nM) was excited at 312 nm, and the emission spectrum was collected from 320 to 500 nm. After serial additions of full-length or truncated yPol η in independent titrations, a decrease in the fluorescence intensity of F-8 was observed. These changes in fluorescence intensity at 370 nm were plotted as a function of the yPol DNA η concentration and were fit to equation 3 to extract a Kd equal to 7 ± 4 nM for full-
187 length yPol η (Figure 8.2 part (C)) and 13 ± 5 nM for truncated yPol η (Table 8.2). These DNA Kd measurements were tighter than those determined using EMSA, since the fluorescence titration assay allows yPol η to associate and dissociate during data collection. In contrast, EMSA does not maintain a constant equilibrium because dissociated yPol η cannot re-associate with DNA during electrophoresis separation. Nonetheless, there was a confirmed ~2-fold difference in the DNA binding affinity between full-length and the catalytic core of yPol η which indicates the C-terminal 119 amino acid residues of yPol η slightly enhance the binding of the enzyme to DNA.
Next, we directly measured the rate of DNA dissociation from the yPol η•DNA complex (Section 8.2). A pre-incubated solution of yPol η (50 nM) and 5′-radiolabeled D-1 DNA (100 nM) was combined with a 50-fold molar excess of unlabeled D-1 DNA for various time intervals before dTTP was added for 15 s to allow ample extension of the labeled D- 1 DNA that remained in complex with yPol η. A plot of product concentration versus the incubation time with the unlabeled DNA trap (data not shown) was fit to equation 4 -1 -1 which yielded DNA dissociation rates (koff) of 0.008 ± 0.001 s and 0.0041 ± 0.0008 s for truncated and full-length yPol η, respectively (Table 8.2 and Scheme 8.1). Interestingly, the rate of DNA dissociation from full-length yPol η is 2-fold slower than from truncated yPol η, which indicated that the C-terminus of yPol η may slightly contribute to this polymerase’s DNA binding affinity.
DNA Based on the measured Kd from Figure 8.2 part (C) and koff values, the apparent DNA second-order association rate constant (kon = koff/Kd ) of the binary complex yPol -1 -1 η•DNA was calculated to be 0.62 and 0.59 μM s for truncated and full-length yPol η, respectively (Table 8.2). These similar kon values indicate that the slightly stronger DNA binding affinity of full-length yPol η is mainly due to a slightly slower rate of DNA dissociation (koff). Taken together, the data in Table 8.2 suggest that the C-terminal 119 amino acid residues of yPol η slightly hinder the dissociation of DNA from the binary complex yPol η•DNA. This hindrance is through either direct physical interactions
188 between the C-terminus of yPol η and DNA, modulation of the conformation of the polymerase domain by the C-terminus of yPol η, or both.
Base substitution fidelity of truncated yPol η Since a pre-steady state burst was observed for truncated yPol η, we continued to
investigate the nucleotide incorporation efficiency (kp/Kd) by measuring the maximum
rate of nucleotide incorporation (kp) and the apparent equilibrium dissociation constant
(Kd) of an incoming nucleotide under single-turnover conditions [112]. By performing
these experiments with yPol η in molar excess over DNA, the conversion of D-DNAn to
D-DNAn+1 (Scheme 8.1) was directly observed in a single pass through the enzymatic pathway [63]. A pre-incubated solution of truncated yPol η (150 nM) and 5′-[32P]-labeled 2+ D-7 DNA (30 nM) was mixed with varying concentrations of dATP•Mg (0.4 - 80 μM) and quenched with EDTA at various times (Section 8.2). A plot of product concentration versus time was fit to equation 5 to extract the observed rate constant
(kobs) for dATP incorporation (Figure 8.3 part (A)). Then, the kobs values were plotted as a function of dATP concentration and fit to a hyperbolic equation (Equation 6) which -1 resolved a kp of 6.9 ± 0.4 s and an apparent Kd of 17 ± 3 μM (Figure 8.3 part (B)). The pre-steady state kinetic parameters for the remaining 15 possible dNTP:dN base pair combinations were determined under single-turnover conditions and were used to
calculate the substrate specificity constant (kp/Kd), discrimination factor ((kp/Kd)correct/(
kp/Kd)incorrect), and fidelity ((kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect]) of truncated yPol η (Table 8.3). Overall, the base substitution fidelity of truncated yPol η was in the range of -2 -4 10 to 10 which translates into 1 misincorporation per 100 to 10,000 nucleotide incorporations (Table 8.3). Depending on the mispair, truncated yPol η catalyzed a misincorporation with 30- to 2,700-fold (640-fold on average) lower efficiency than the corresponding correct base pair. To better understand the mechanistic basis of truncated yPol η’s fidelity, the equation for polymerase fidelity can be simplified as follows:
Fidelity = (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect] ≈ (kp/Kd)incorrect/(kp/Kd)correct = -1 -1 -1 [(kp)incorrect/(kp)correct] [(Kd)correct/(Kd)incorrect] = (rate difference) (binding affinity -1 difference) (8)
189 Thus, fidelity is inversely proportional to the rate difference and apparent binding affinity difference between correct and incorrect nucleotide incorporation. In general, the mechanistic basis of yPol η’s discrimination was due to a 3- to 68-fold (18-fold on average) weaker apparent binding affinity (1/Kd) and 5- to 220-fold (50-fold on average) slower rate constant of incorporation for a mismatched dNTP.
Kinetic significance of base stacking contributing to the binding affinity of an incoming nucleotide Although all four correct dNTPs were bound with similarly high affinity (Table 8.3), mismatched purine deoxyribonucleotides have 2- to 6-fold lower apparent Kd values than mismatched pyrimidine deoxyribonucleotides. Because 5′-protruding purines have been found to have stronger stacking interactions with a terminal DNA base pair than 5′- protruding pyrimidines [199], the difference in apparent Kd values suggest that base- stacking interactions between an incorrect dNTP and the terminal primer/template base pair dA:dT (Table 8.1) play a role on the binding of dNTP by truncated yPol η. Interestingly, we have previously demonstrated that the preferred nucleotide for template-independent nucleotide incorporation catalyzed by Dpo4, another Y-family DNA polymerase, is dATP mainly due to its strong intrahelical base-stacking ability [194]. To further evaluate the role of base stacking, we first examined if truncated yPol η can catalyze template-independent nucleotide incorporation of dATP or dPTP (Figure 8.4) onto four palindromic, blunt-end DNA substrates (BE1, BE2, BE3, and BE4 in Table 8.1). The base of dPTP, a dNTP analog, has four conjugated benzene rings but possesses no hydrogen bonding abilities. The DNA substrates possess all four possible terminal base pairs and each molecule of them can be bound by a single polymerase molecule. Our radioactive experiments showed that truncated yPol η was able to incorporate dATP and dPTP (data not shown). Then, we individually measured the kinetic parameters for dATP and dPTP incorporation under single-turnover reaction conditions (Table 8.4). Interestingly, the apparent Kd values of dATP were 3- to 5- fold smaller with a purine than with a pyrimidine on the primer’s 3′-base, indicating that base stacking is also important for the binding of dATP to the binary complex of yPolη•blunt-
190 end DNA. This base-stacking effect is more dramatic for dPTP incorporation onto blunt-
end DNA because the apparent Kd values of dPTP are 10- to 80-fold tighter than dATP incorporation onto the same blunt-end DNA substrate (Table 8.4). Thus, the binding free energy difference between dATP and dPTP is 1.4 to 2.6 kcal/mol. Previously, we have obtained a comparable binding free energy difference of 2.3 kcal/mol for similar blunt- end dATP and dPTP incorporation at 37 °C catalyzed by Dpo4 [194]. Although neither dATP nor dPTP form any hydrogen bonds with a template base when bound by yPol η•blunt-end DNA, the bases of these two nucleotides should have different base-stacking interactions with a terminal base pair of a blunt-end DNA substrate considering that a dangling pyrene base (1.7 kcal/mol) has previously been found to possess a higher base- stacking free energy than a dangling adenosine (1.0 kcal/mole) [199]. However, the base-stacking free energy difference (0.7 kcal/mole) between pyrene and adenosine is smaller than the aforementioned binding free energy difference (1.4-2.6 kcal/mol) between dPTP and dATP. Thus, other sources likely contribute to the tighter binding of dPTP over dATP. One possible source is favorable van der Waals interactions between pyrene and active site residues of truncated yPol η. In addition, the base-stacking effect and van der Waals interactions may stabilize the ternary complex of yPol η•blunt end
DNA•nucleotide and facilitate catalysis, leading to much higher kp values with dPTP than
with dATP (Table 8.4). Due to the differences in kp and apparent Kd, the substrate specificity values of dPTP are 100- to 1,000-fold higher than those of dATP with blunt- end DNA (Table 8.4) and 10- to 100-fold higher than mismatched dATP with regular DNA (Table 8.3).
Base substitution fidelity of full-length yPol η The base substitution fidelities of full-length and truncated yPol η may differ because the C-terminal, non-enzymatic regions may alter the polymerization fidelity. For example, the proline-rich domain of human DNA polymerase λ has been shown to upregulate the polymerase fidelity up to 100-fold [117]. To determine if the C-terminus of yPol η influences polymerization fidelity, we measured the pre-steady state kinetic parameters for dNTP incorporation into D-1 DNA (template dA) catalyzed by full-length yPol η
191 (Table 8.5). The fidelity was calculated to be in the range of (1.4 to 2.6) × 10-3 for full- length yPol η (Table 8.5). Relative to the fidelity of truncated yPolη with D-1 (Table 8.3), full-length yPol η has a 3-fold higher fidelity. Therefore, the C-terminus of yPol η slightly affects the base substitution fidelity. Moreover, truncated yPol η discriminated between a correct and incorrect dNTP by ~30-fold on average based on the kp difference while the discrimination for full-length yPol η was ~170-fold on average for incorporation into D-1 DNA (Tables 8.3 and 8.5). The incorporation rate constant for -1 correct dTTP was ~4 s for both yPol η enzymes, but the misincorporation rate was 3- to 23-fold faster for truncated yPol η. This rate enhancement for truncated yPol η is partially offset by a greater discrimination at the apparent ground-state binding level so that the fidelity of truncated yPol η was only 3-folder lower than that of full-length yPol η.
Effect of the non-enzymatic C-terminus of yPol η on its polymerase activity Our above studies demonstrated that the C-terminus of yPol η enhances this enzyme’s DNA binding affinity and base substitution fidelity by 2- and 3-fold, respectively. These results suggest that the non-enzymatic, C-terminal region of yPol η (Figure 8.1) has a mild impact on the N-terminal polymerase domain and its activity. This conclusion is inconsistent with previous studies which have qualitatively demonstrated that mutations or deletions in the UBZ domain or PIP motif do not affect polymerase activity [293-295]. However, these reported qualitative assays are not sufficiently sensitive to detect the small perturbation on polymerase activity as described in this chapter. The presence of the C-terminal 119 residues of yPol η may either interact with DNA, slightly alter the conformation of the polymerase domain, or both (see above discussion), thereby enhancing its DNA binding affinity and polymerase fidelity.
Kinetic comparison among Y-family DNA polymerases The fidelity of several Y-family DNA polymerases synthesizing undamaged DNA has been determined by employing steady-state [100, 104, 296-304], pre-steady state [60, 68, 105, 115, 186, 189, 305], or M13-based mutation assays [104, 297, 299, 300, 306, 307]. 0 -4 From these studies, the fidelity ranges from 10 to 10 . Under steady-state reaction
192 conditions, the base substitution fidelity of yPol η and human Pol η have been measured -2 -4 -2 -3 to be in the range of 10 to 10 and 10 to 10 , respectively [296, 298], which is similar to our pre-steady state kinetic results. Consistently, Pol η displays the highest substrate specificity for the dCTP:dG base pair under both steady-state and pre-steady state reaction conditions (Table 8.3 and unpublished data in Chapters 11 and 12) [296, 298]. This may seem surprising, since Pol η participates in the efficient bypass of UV-induced DNA damage such as a cis-syn thyminethymine dimer (i.e. a dATP:dT base pair) [33, 35, 36, 285-288, 308]. However, Pol η has also been shown to be efficient at bypassing guanine-specific damage such as 8-oxo-7,8-dihydro-dG [264, 267], 1,2- cisdiammineplatinum(II)-d(GpG) intrastrand cross-links [309-312], and various N2-dG lesions [313, 314].
Among the four eukaryotic Y-family DNA polymerases [i.e. Pol η, DNA polymerase κ, DNA polymerase ι (Pol ι), and Rev1], Rev1 exhibits low fidelity on undamaged DNA due to its strong preference for inserting dCTP [189, 302] while Pol ι has an unusual preference for dGTP:dT mispairs over dATP:dT due to Hoogsteen base pair formation [68, 69]. Interestingly, the lowest fidelity base pair for truncated yPol η was dGTP:dT (Table 8.3). This observation likely results from the formation of a wobble base pair. The two hydrogen bonds established in the wobble base pair may enhance the catalytic efficiency of yPol η since hydrogen bonding is important for the efficiency and accuracy of yPol η [315]. Also noteworthy, the truncated versions of eukaryotic Y-family DNA polymerases have been used for many biochemical studies in literature. Based on our quantitative kinetic analysis of yPol η, these results suggested the non-enzymatic regions of Y-family DNA polymerases do not alter the polymerase activity significantly.
Fidelity comparison among various DNA polymerase families As a Y-family DNA polymerase, yPol η displays low fidelity on undamaged DNA (Tables 8.3 and 8.5) [296]. In contrast, replicative DNA polymerases in the A- and B- families have a polymerization fidelity that is 1-3 orders of magnitude greater than the Y- family DNA polymerases (Table 8.6). DNA polymerases with higher fidelity are more
193 proficient at using the ground-state binding affinity to discriminate between a correct and incorrect dNTP. The Y-family DNA polymerases provide little to no discrimination
based on the Kd difference while replicative DNA polymerases discriminate up to almost three orders of magnitude. This lack of selection in the ground state by the Y-family DNA polymerases may be due to the relatively loose and solvent-accessible active site which has minimal contacts with the nascent base pair [46, 101, 289]. Moreover, nucleotide selection by the Y-family DNA polymerases in the ground state may be mainly governed by Watson-Crick base pairing, since the calculated ΔΔG values (0.95 - 1.7 kcal/mol) are similar to the free energy differences between correct and incorrect base pairs (0.3 - 1.0 kcal/mol at 37 °C) at the primer terminus based on DNA melting studies (Table 8.6) [124]. However, with ΔΔG values ≥3.0 kcal/mol, the replicative DNA polymerases harness the additional 2.0 kcal/mol of energy from other sources such as a tight active site or close contacts with the nascent base pair. One common fidelity checkpoint among DNA polymerases is the varying rate differences between a matched and mismatched base pair. These large differences may correspond to different rate- limiting steps (e.g. protein conformational change, or phosphodiester bond formation) during nucleotide incorporation [46, 54, 60]. For yPol η, kinetic data suggests correct and incorrect dNTPs are limited by a conformational step preceding chemistry, although, additional studies are needed to confirm these results [115].
This work presents the mechanistic basis of the base substitution fidelity of yPol η on undamaged DNA, which examined all possible dNTP:dN base pair combinations for the first time. yPol η discriminates against incorrect nucleotides at both the ground-state nucleotide binding and incorporation steps. Furthermore, base stacking contributes to tighter binding for a misincorporation. Finally, the 119 residues at the C-terminus have a mild impact on the kinetic mechanism of yPol η.
194 8.4 Tables
Table 8.1 Sequences of DNA substratesa D-1 5’-CGCAGCCGTCCAACCAACTCA-3’ 3’-GCGTCGGCAGGTTGGTTGAGTAGCAGCTAGGTTACGGCAGG-5’ D-6 5’-CGCAGCCGTCCAACCAACTCA-3’ 3’-GCGTCGGCAGGTTGGTTGAGTGGCAGCTAGGTTACGGCAGG-5’ D-7 5’-CGCAGCCGTCCAACCAACTCA-3’ 3’-GCGTCGGCAGGTTGGTTGAGTTGCAGCTAGGTTACGGCAGG-5’ D-8 5’-CGCAGCCGTCCAACCAACTCA-3’ 3’-GCGTCGGCAGGTTGGTTGAGTCGCAGCTAGGTTACGGCAGG-5’ F-8 5’-CGCAGCCGTCCAACCAACTCA-3’ 3’-GCGTCGGCAGGTTGGTTGAGTCXCAGCTAGGTTACGGCAGG-5’ BE1 5’-ATGAGTTGCAACTCAT-3’ 3’-TACTCAACGTTGAGTA-5’ BE2 5’-TTGAGTTGCAACTCAA-3’ 3’-AACTCAACGTTGAGTT-5’ BE3 5’-CTGAGTTGCAACTCAG-3’ 3’-GACTCAACGTTGAGTC-5’ BE4 5’-GTGAGTTGCAACTCAC-3’ 3’-CACTCAACGTTGAGTG-5’ aThe template base highlighted in bold is unique to each strand and X denotes 2- aminopurine.
195
Table 8.2 Rate and equilibrium dissociation constants for the binary complex yPol η•DNA at 23 °C. Kinetic parameter Truncated yPol η Full-length yPol η -1 -1 a kon (μM s ) 0.62 0.59 -1 koff (s ) 0.008 ± 0.001 0.0041 ± 0.0008 DNA b Kd (nM) 34 ± 3 16 ± 1 DNA c Kd (nM) 13 ± 5 7 ± 4 a DNA DNA Calculated as koff/Kd . The Kd value was measured from a fluorescence titration assay. bEstimated using EMSA. cMeasured using a fluorescence titration assay.
196
Table 8.3 Kinetic parameters of nucleotide incorporation into D-DNA catalyzed by truncated yPol η at 23 °C. k K k /K Discrimination dNTP p d p d Fidelityb (s-1) (μM) (μM-1s-1) factora Template dA (D-1) dTTP 3.9 ± 0.2 15 ± 2 2.6 × 10-1 dATP 0.089 ± 0.005 80 ± 20 1.1 × 10-3 230 4.3 × 10-3 dCTP 0.43 ± 0.06 210 ± 60 2.0 × 10-3 130 7.8 × 10-3 dGTP 0.15 ± 0.01 80 ± 20 1.9 × 10-3 140 7.2 × 10-3 Template dG (D-6) dCTP 15.6 ± 0.3 11.2 ± 0.8 1.4 dATP 0.071 ± 0.002 138 ± 9 5.1 × 10-4 2700 3.7 × 10-4 dGTP 0.116 ± 0.006 80 ± 10 1.5 × 10-3 960 1.0 × 10-3 dTTP 0.92 ± 0.07 330 ± 40 2.8 × 10-3 500 2.0 × 10-3 Template dT (D-7) dATP 6.9 ± 0.4 17 ± 3 4.1 × 10-1 dCTP 1.00 ± 0.04 210 ± 20 4.8 × 10-3 85 1.2 × 10-2 dGTP 0.55 ± 0.01 46 ± 3 1.2 × 10-2 30 2.9 × 10-2 dTTP 0.62 ± 0.02 280 ± 20 2.2 × 10-3 180 5.4 × 10-3 Template dC (D-8) dGTP 6.3 ± 0.1 6.8 ± 0.4 9.3 × 10-1 dATP 0.087 ± 0.003 90 ± 10 9.7 × 10-4 960 1.0 × 10-3 dCTP 0.127 ± 0.007 200 ± 30 6.4 × 10-4 1500 6.9 × 10-4 dTTP 1.39 ± 0.06 460 ± 40 3.0 × 10-3 310 3.3 × 10-3 a Calculated as (kp/Kd)correct/(kp/Kd)incorrect. b Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
197
Table 8.4 Kinetic parameters for nucleotide incorporation onto blunt-end DNA catalyzed by truncated yeast Pol η at 23 °C. DNA kp Kd kp/Kd Efficiency (Terminal dNTP -1 -1 -1 ratioa base pair) (s ) (μM) (μM s ) BE1 (dT:dA) dATP 0.026 ± 0.002 1200 ± 200 2.2 × 10-5 - dPTP 1.27 ± 0.08 60 ± 10 2.1 × 10-2 980 BE2 (dA:dT) dATP 0.036 ± 0.002 220 ± 30 1.6 × 10-4 - dPTP 0.68 ± 0.03 23 ± 3 3.0 × 10-2 180 BE3 (dG:dC) dATP 0.0087 ± 0.0003 360 ± 30 2.4 × 10-5 - dPTP 0.22 ± 0.01 9 ± 2 2.4 × 10-2 1000 BE4 (dC:dG) dATP 0.032 ± 0.001 930 ± 70 3.4 × 10-5 - dPTP 0.74 ± 0.03 12 ± 2 6.2 × 10-2 1800 a Calculated as (kp/Kd)dPTP/(kp/Kd)dATP.
198
Table 8.5 Kinetic parameters of nucleotide incorporation into D-1 DNA catalyzed by full-length yPol η at 23 °C. k K k /K Discrimination dNTP p d p d Fidelityb (s-1) (μM) (μM-1s-1) factora Template dA (D-1) dTTP 4.2 ± 0.5 40 ± 10 1.1 × 10-1 dATP 0.0235 ± 0.0003 156 ± 7 1.5 × 10-4 700 1.4 × 10-3 dCTP 0.019 ± 0.001 70 ± 10 2.7 × 10-4 390 2.6 × 10-3 dGTP 0.043 ± 0.003 170 ± 40 2.5 × 10-4 420 2.4 × 10-3 a Calculated as (kp/Kd)correct/(kp/Kd)incorrect. b Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
199
Table 8.6 Comparison of base substitution fidelity for various DNA polymerases. Polymerase K k ΔΔGo Polymerase Fidelitya d p family differenceb differenceb (kcal/mol)c Truncated Y 3.7 × 10-4 to 2.9 × 10-2 3 to 68 5 to 220 1.6 yPol ηd Dpo4e Y 1.5 × 10-4 to 3.2 × 10-3 1 to 18 240 to 1700 0.95 rPol βf X 1.1 × 10-5 to 5.9 × 10-4 35 to 342 28 to 708 3.0 PolB1 exo-g B 3.5 × 10-6 to 1.2 × 10-4 109 to 918 4 to 589 3.7 hPol γh A 4.6 × 10-7 to 2.9 × 10-4 42 to 900 39 to 12000 3.4 a b Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect]. Calculated as defined in equation 8 (Section 8.3). cCalculated using equation 7 (Section 8.2). dAt 23 °C (this work). eAt 37 °C [105]. fAt 37 °C [114]. gAt 37 °C, excluding the fidelity contribution from the 3′ → 5′ exonuclease activity [146]. hAt 37 °C, excluding the fidelity contribution from the 3′ → 5′ exonuclease activity [91].
200 8.5 Figures
Figure 8.1 Schematic illustration of yPol η. The polymerase domain of yPol η is at the N-terminus while a ubiquitin-binding zinc finger (UBZ) domain and PCNA-interacting peptide (PIP) motif is at the C-terminus. Residue numbers are denoted above each region. For this study, the truncated construct contains only the polymerase domain.
201 A
B C
10 6 x 104
5.5 x 104 8
5 x 104 6 4.5 x 104
4 4 4 x 10
4
Intensity (cps) 3.5 x 10 2 Binary Complex (nM) Complex Binary 3 x 104
0 2.5 x 104 0 100 200 300 400 500 0 50 100 150 200 250 300 350 yPol η (nM) yPol η (nM)
Figure 8.2 Equilibrium dissociation constant for full-length yPol η. (A) Gel image showing binary complex formation at various concentrations of full-length yPol η (10 - 450 nM) in the presence of 5′-[32P]-labeled D-1 DNA (10 nM). (B) The concentration of the binary complex was plotted as a function of full-length yPol η concentration and fit to DNA equation 2 (Section 8.2) to yield a Kd = 16 ± 1 nM. (C) For the fluorescence titration assay, a plot of fluorescence intensity versus full-length yPol η concentration was fit to DNA equation 3 which resolved a Kd = 7 ± 4 nM.
202 A 30
25
20
15
10 Product (nM) Product
5
0 02468 Time (s)
B 7
6
5 )
-1 4 (s
obs 3 k 2
1
0 0 20406080100 dATP (μM)
Figure 8.3 Concentration dependence on the pre-steady state rate constant of nucleotide incorporation catalyzed by truncated yPol η. (A) A pre-incubated solution of truncated yPol η (150 nM) and 5′-[32P]-labeled D-7 DNA (30 nM) was mixed with dATP•Mg2+ (0.4 μM, z; 0.8 μM, {; 2 μM, ; 4 μM, ; 8 μM, S; 16 μM, U; 40 μM, ; 80 µM, ) and quenched with EDTA at various time intervals. The solid lines are the best fits to a single-exponential equation which determined the observed rate constant, kobs. (B) The kobs values were plotted as a function of dATP concentration. The data (z) were then fit -1 to a hyperbolic equation, yielding a kp of 6.9 ± 0.4 s and a Kd of 17 ± 3 μM.
203
Figure 8.4 Chemical structure of a non-natural nucleotide analog, dPTP.
204 8.6 Scheme
Scheme 8.1
205 Chapter 9: Kinetic Basis of Nucleotide Selection Employed by a Protein Template- Dependent DNA Polymerase
9.1 Introduction
The human genome encodes at least 16 DNA polymerases (Pol) that are involved in replicating and maintaining the integrity of genomic DNA. Human DNA polymerases are classified into four families: A, B, X, and Y. Y-family DNA polymerases are involved in DNA damage tolerance pathways, whereby a Y-family enzyme rescues stalled DNA replication at sites of DNA damage. Humans have four known Y-family members: Pol η, Pol ι, Pol κ, and Rev1. Rev1 is found in the genome of all eukaryotes [5] and is capable of functioning in both catalytic and structural roles. Composed of 1,251 amino acids [70], human Rev1 (hRev1) is organized into a central catalytic domain that is flanked by an N- terminal BRCT domain and a C-terminus with two ubiquitin-binding motifs and a domain for polymerase interactions [67]. As a scaffold protein, Rev1 interacts with proliferating cell nuclear antigen (PCNA) [316-319], ubiquitinated proteins [317, 318], and DNA polymerases η, κ, ι, and ζ [320-327]. These findings support a model, whereby Rev1 is involved in polymerase switching at sites of DNA damage [79, 328, 329]. In regards to enzymatic activity, hRev1 preferentially inserts dCTP opposite a templating base dG [70, 304, 330-332], however, unlike other human DNA polymerases, this incorporation event proceeds in a protein template-directed manner rather than a DNA template-dependent manner with Watson-Crick base pairing [72]. Instead, the incoming dCTP hydrogen bonds with R357, and the extrahelical template base dG is accommodated in a hydrophobic pocket while L358 rests in the conventional location of a templating base (Figure 9.1) [72].
206 Rev1 and Pol ζ are responsible for the majority of spontaneous and DNA damage- induced mutagenic events in yeast; early studies reveal similar findings in mammalian cell lines [333-335]. In human tissues, the rev1 gene is ubiquitously expressed, but the highest level of expression is in human testis and ovary based on RT-PCR results [70, 320, 330]. Furthermore, hRev1 has been observed at replication foci during both G1 and S phases following UV-irradiation [336]. However, it has also been reported that the protein levels of hRev1 are unaffected by UV irradiation or cell cycle progression [337]. In addition to a role in translesion synthesis, Rev1 has been implicated in somatic hypermutation, and current data suggests the catalytic domain participates in the generation of C to G transversions [338, 339]. To better understand the enzymatic function of hRev1, we have performed pre-steady state kinetic analysis on a truncated version of hRev1. Our studies established a kinetic basis for nucleotide selection by hRev1.
9.2 Materials and Methods
Materials These chemicals were purchased from the following companies: [γ-32P]ATP, MP Biomedicals; deoxyribonucleotide 5′-triphosphates, GE Healthcare; ribonucleotide 5′- triphosphates, MBI Fermentas; 2′-aracytidine-5′-triphosphate (araCTP), 2′-deoxy-2′,2′- difluorocytidine-5′-triphosphate (GemCTP), 2′-fluoro-2′-deoxycytidine-5′-triphosphate
(2′-FCTP), 2′-O-methylcytidine-5′-triphosphate (2′-OCH3-CTP), and 5-nitroindole 5′- triphosphate (dNITP), TriLink Biotechnologies; Bio-Spin 6 columns, Bio-Rad Laboratories; OptiKinase™, USB Corporation; synthetic oligodeoxyribonucleotides 21- mer, 5′-phosphorylated 19-mer, and 41-mers, Integrated DNA Technologies. Pyrene 5′- triphosphate (dPTP) was a generous gift from Dr. John-Stephen Taylor (Washington University at St. Louis).
207 Expression and purification of hRev1 The expression plasmid pBAD-REV1S, a generous gift from K. Kamiya at Hiroshima University, encoded a truncated version of human Rev1 (341-829) [340]. The expression and purification of truncated human Rev1 was performed as previously described [330].
DNA substrates Commercially synthesized oligomers in Table 9.1 were purified using polyacrylamide gel electrophoresis [105, 182]. The 21-mer primer was radiolabeled with [γ-32P]ATP and OptiKinase™ according to the manufacturer’s protocol, and the unreacted [γ-32P]ATP was subsequently removed via a Bio-Spin 6 column. The primer-template DNA substrates [105] and single-nucleotide gap DNA substrate [182] were annealed as described previously.
Measurement of the kp and Kd for single-nucleotide incorporation Kinetic assays were completed using buffer R (50 mM HEPES, pH 7.5 at 37 °C, 5 mM
MgCl2, 50 mM NaCl, 0.1 mM EDTA, 5 mM DTT, 10% glycerol, and 0.1 mg/ml BSA). All kinetic experiments described herein were performed at 37 °C, and the reported concentrations were final after mixing all of the components. A pre-incubated solution containing hRev1 (120 nM) and 5′-[32P]-radiolabeled DNA substrate (30 nM) was mixed with increasing concentrations (0.02-800 μM) of nucleotide in buffer R at 37 °C. Aliquots of the reaction mixtures were quenched at various times using 0.37 M EDTA. A rapid chemical-quench flow apparatus (KinTek) was utilized for fast nucleotide incorporations. Reaction products were resolved using sequencing gel electrophoresis (17% acrylamide, 8 M urea) and quantitated with a Typhoon TRIO (GE Healthcare). The time course of product formation at each nucleotide concentration was fit to a single- exponential equation (Equation 1)
[Product] = A[1 – exp (–kobst)] (1) using a nonlinear regression program, KaleidaGraph (Synergy Software), to yield an observed rate constant of nucleotide incorporation (kobs). The kobs values were then plotted
208 as a function of nucleotide concentration and fit using the hyperbolic equation (Equation 2)
[kobs] = kp[dNTP]/(Kd + [dNTP]) (2) which resolved the kp and Kd values for nucleotide incorporation catalyzed by hRev1.
9.3 Results Kinetic basis of dNTP selection Transient state kinetic methods were employed to measure the substrate specificity and polymerase fidelity of a truncated form of hRev1. A pre-incubated solution of hRev1 (120 nM) and 5′-[32P]-labeled D-G DNA (30 nM) was mixed with increasing 2+ concentrations of dCTP•Mg (Section 9.2). These single-turnover conditions in which hRev1 is in molar excess over DNA permits the direct observation of the DNA substrate being converted to the extended DNA product in a single pass through the enzymatic pathway [63]. The extended DNA product was quantitated, plotted (Figure 9.2), and fit to the appropriate equations (Equations 1 or 2) that resolved a maximum rate
of nucleotide incorporation (kp) of 22.4 ± 0.9 s-1 and an equilibrium dissociation constant
(Kd) of 2.2 ± 0.3 μM (Table 9.2). Notably, Tsai and Johnson report that nucleotide binding to T7 DNA polymerase, an A-family enzyme, induces several conformational changes preceding the incorporation step, thereby arguing that the measured Kd value under single-turnover reaction conditions is not a true equilibrium dissociation constant [112]. Since there is no published evidence to support the existence of such
conformational changes for the protein template-directed hRev1, we assume the Kd values measured in this chapter reflect the true nucleotide binding affinity (1/Kd). To examine how efficient hRev1 incorporates dCTP opposite other templating bases, we performed similar single-turnover assays using DNA substrates with dA (D-A), dC (D-C), and dT
(D-T) as the template base (Table 9.2). The substrate specificity constants (kp/Kd), efficiency ratio, and fidelity were calculated. The ground-state binding affinity dropped 4- to 55-fold while the rate for dCTP incorporation was reduced by 7- to 12-fold when the templating base was not dG. Overall, the catalytic efficiency was up to 360-fold
209 greater when dCTP was inserted into D-G. The preferential order of dCTP incorporation opposite the four template bases was dG >> dA > dT ≈ dC.
Next, we measured the catalytic efficiency of nucleotide incorporation for the three remaining Watson-Crick base pair combinations under single-turnover conditions and the kinetic data are listed in Table 9.2. Compared to dCTP:dG, the catalytic efficiency of hRev1 dropped 4,900-, 12,000- and 42,000-fold for dTTP:dA, dATP:dT, and dGTP:dC, respectively. Despite a change in the identity of an incoming dNTP, the template preference remained the same based on the substrate specificity constant as observed with dCTP. The binding affinity remained high for dATP but was ~14- and 20-fold weaker for dGTP and dTTP. Furthermore, the rate of dCTP incorporation into D-A, D-T, and D-C DNA was up to 820-fold faster than the canonical dNTP, therefore, the strong dCTP preference by hRev1 with templating bases dA, dT, and dC leads to an extremely low fidelity of ~1 (Table 9.2). Please note, enzyme fidelity is calculated using the
standard kinetic equation, (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect]. When fidelity approaches a value of 1, this indicates that a misincorporation is favored over the canonical Watson-Crick base pair and that a correct incorporation is not likely to occur. Therefore, to better understand the frequency of a correct incorporation catalyzed by
hRev1, the following equation was used: (kp/Kd)correct/(kp/Kd)dCTP:dN. Here, the frequency -2 -2 -3 of a correct incorporation is calculated to be 1.1 × 10 , 1.6 × 10 , and 8.6 × 10 for dTTP:dA, dATP:dT, and dGTP:dC, respectively. These values translate into approximately one correct incorporation (dTTP, dATP, or dGTP) per 100 dCTP misincorporations.
Since hRev1 displayed greater catalytic efficiency when dG is the template base, we determined the substrate specificity constant for the incorporation of the other dNTPs into D-G DNA (Table 9.2). The efficiency to form base pairs dATP:dG, dGTP:dG, and dTTP:dG was 1-, 290-, and 20-fold greater than dATP:dT, dGTP:dC, and dTTP:dA, respectively. Surprisingly, relative to the other template bases, the rate of nucleotide incorporation was up to 860-fold faster when the substrate had dG positioned as the
210 template base. Meanwhile, the Kd value was at least 10-fold higher for non-dCTP addition into D-G DNA. The fidelity of hRev1 inserting dNTPs opposite dG ranged from -3 -5 10 to 10 . It has been shown that hRev1 may participate in cellular processes that involve gapped DNA [341]. Determining the pre-steady state kinetic parameters for dCTP incorporation into a single-nucleotide gapped DNA substrate (D-G Gap) revealed that hRev1 is 7-fold more efficient with the primer-template D-G DNA substrate (Table 9.2). This modest effect can be attributed to a 2-fold slower rate and a 4-fold weaker binding affinity for dCTP incorporation.
Importance of hydrogen bonding and base stacking Crystallographic studies have shown that hRev1 utilizes a protein template-directed mechanism to instruct dCTP incorporation through hydrogen bonding between cytosine and residue R357 of hRev1 (Figure 9.1) [72]. To evaluate the roles of hydrogen bonding, base stacking, and base size during DNA synthesis, we have measured the catalytic efficiency of hRev1 incorporating two non-natural nucleotide analogs into D-G DNA (Figure 9.3 part (A) and Table 9.3). Both dPTP and dNITP lack the ability to form strong hydrogen bonds, possess greater base stacking energy, and are physically larger than dCTP [199]. hRev1 can incorporate both analogs, although, the incorporation efficiency drops by 3,500- and 11,000-fold for dNITP and dPTP, respectively. Both analogs are incorporated with significantly reduced rates (at least 490-fold) and modestly weakened binding affinities (at least 7-fold). These data suggested that hydrogen bonding is not essential for catalysis, but it does enhance the rate and binding affinity for dCTP incorporation.
Kinetic basis of ribonucleotide selection The concentration of cellular dNTP pools fluctuate during the cell cycle, and the levels are 10- to 200-fold less than the ribonucleotide (rNTP) pools which remain relatively high and constant [178, 179]. Since hRev1 has been shown to be present outside of S phase [337], we have evaluated the sugar selectivity of hRev1 by measuring the
211 substrate specificity constant for various CTP analogs (Figure 9.3 part (B) and Table 9.4). hRev1 discriminates between dCTP and rCTP by 280-fold, and this is mostly due to a 230-fold rate decrease. To better understand how size and orientation affect the degree of sugar selectivity, we have used araCTP (an anti-cancer drug that is a steric isomer of rCTP with the 2′-OH pointed above the ribose ring), 2′-F-CTP (the 2′-F group is smaller than the 2′-OH), GemCTP (an anticancer drug with two fluorines at the 2′ position), and
2′-OCH3-CTP (the 2′-OCH3 group is larger than the 2′-OH). Orientation and reduced size of the 2′ group are important factors because the efficiency of hRev1 incorporating araCTP and 2′-F-CTP was similar to dCTP. In contrast, the increased volume of the 2′- methoxy group enhanced the magnitude of discrimination to 6,700. Surprisingly, most of the sugar selection was kp driven for hRev1. The one exception is for GemCTP where the
Kd value increased by 13-fold.
9.4 Discussion
Comparison of base substitution fidelity As a dCTP transferase, Rev1 is a DNA polymerase with extremely low fidelity due to the preference to form dCTP:dN base pairs over canonical Watson-Crick base pairs dTTP:dA, dATP:dT, and dGTP:dC. Using pre-steady state kinetic methods, we have 0 -5 established a base substitution fidelity of 10 to 10 for truncated hRev1 synthesizing undamaged DNA (Table 9.2). This fidelity range is similar to other human Y-family 0 -4 DNA polymerases [66] and a fidelity range of 10 to 10 that was estimated for full- length hRev1 under semi-steady-state kinetic conditions by Zhang et al. [304]. In their studies, Zhang et al. used too much full-length hRev1 (14 fmol) in the reactions with 50 fmol of DNA and various dNTPs at 30 °C [304], possibly due to the lack of quantifiable reaction products during non-dCTP incorporations. Thus, their semi-steady-state kinetic parameters cannot be used to kinetically describe nucleotide incorporation catalyzed by hRev1. In this chapter, we employed pre-steady state kinetic methods to investigate the kinetic basis for nucleotide selection and enzyme fidelity for hRev1. Our kinetic data
212 revealed that hRev1 discriminates at both the nucleotide binding (Kd) and incorporation
(kp) steps. Overall, hRev1 prefers dCTP:dG with a 20-fold tighter binding affinity and 14- fold faster rate of incorporation (on average) with undamaged DNA relative to the other tested dNTP:dN base pair combinations (Table 9.2).
Pre-steady state kinetic analyses have been completed with a truncated form of yeast Rev1 (yRev1, 1-746) [189]. In stark contrast, yRev1 selects incoming nucleotides mostly at the nucleotide binding step (Kd). The catalytic efficiency for dCTP:dG is 660-fold greater for the human enzyme, and this effect is governed by a ~1,900-fold faster rate of dCTP incorporation catalyzed by hRev1 at 37 °C (22.4 s-1) versus yRev1 at 22 °C (0.012 s-1), although, hRev1 (2.2 μM) binds dCTP with a 3-fold weaker affinity than yRev1 (0.78 μM) [189]. Interestingly, significant kinetic differences have been observed for human and yeast Pol η at varying reaction temperatures, too [186]. Thus, it is important to exercise caution when extending conclusions about DNA polymerase homologs derived from different organisms [326, 342].
Effect of DNA substrate on the catalytic efficiency of hRev1 Translesion DNA synthesis has been proposed to proceed through a polymerase- switching or gap-filling model [40]. Also, Rev1 has been shown to be important during UV-induced post replicative gap-filling processes that likely occur outside of S phase [40, 341]. Although the incorporation efficiency dropped by ~7-fold from non-gapped to gapped DNA, hRev1 is capable of accommodating a single-nucleotide gap DNA substrate despite lacking the signature helix-hairpin-helix (HhH) motif that Pol β and Pol λ, two X-family DNA polymerases specialized for gap-filling DNA synthesis, use to -1 -1 bind the downstream strand. Moreover, the gap-filling efficiency of 1.4 μM s for hRev1 is close or similar to the values measured for rat Pol β (6.6 μM-1s-1) and human Pol λ (1.8 μM-1s-1) (Table 9.2) [114, 117]. More studies are needed to evaluate whether hRev1 plays a role in gap-filling DNA synthesis in vivo.
213 Kinetic basis for nucleotide selection Watson-Crick hydrogen bond formation between the template base and incoming dNTP has been shown to play an important role in nucleotide selection by many DNA polymerases including T7 DNA polymerase [202]. However, hRev1 does not use this DNA template-dependent mechanism to select incoming dNTPs. Instead, it uses the protein template-directed mechanism while the templating base dG is evicted from the active site by L358 so that it fits into a hydrophobic pocket surrounded by F525, K770, and H774 (Figure 9.1). To probe whether hydrogen bonds between cytosine and R357 are essential for catalysis by hRev1, we examined if hRev1 could incorporate dNITP and dPTP which are unable to form hydrogen bonds. Although efficiency was reduced dramatically (Table 9.3), these non-natural nucleotide analogs were incorporated into DNA by hRev1. These results suggested that hydrogen bonds formed between the incoming dNTP and R357 are important, but not absolutely essential for efficient nucleotide incorporation catalyzed by hRev1 and that an oversized nucleobase with strong base stacking energy can be accommodated. To better understand the role of hydrogen bonds, additional studies need to be performed using isosteric, non-hydrogen bonding dCTP analogs.
Previously, Howell et al. [189] proposed possible interactions (i.e. hydrogen bonds and base conformations) for the four dNTP:Arg combinations based on the X-ray crystal structures of yRev1•DNA•dCTP [71] and E. coli MutM DNA glycosylase•DNA [280]. Interestingly, the number of hydrogen bonds correlates with the substrate specificity of dNTP incorporation into DNA with dG as the template for both yRev1 and hRev1: dCTP (2 hydrogen bonds) > dGTP (2 hydrogen bonds if dGTP adopts a syn conformation) ≈ dTTP (1 hydrogen bond) > dATP (0 hydrogen bonds) [189, 302]. However, the identity of the template base also contributes to catalytic efficiency since dCTP misincorporation is less efficient for hRev1 (Table 9.2). Thus, optimal catalytic activity (kp/Kd) of hRev1 depends on both substrates: an incoming dCTP and the template base dG.
214 Kinetic basis for ribonucleotide exclusion Most DNA polymerases prevent ribonucleotide incorporation via a steric clash between the 2′-OH group of an incoming rNTP and a protein backbone segment [193] or bulky side chain residue of the enzyme [162-164, 166, 167]. This mechanism usually yields sugar selectivity values greater than 1,000-fold [162, 164, 166, 167, 169, 193]. hRev1 discriminates between dCTP and rCTP by 280-fold, a value that is relatively low compared to other DNA polymerases (Table 9.4). Like other DNA polymerases, hRev1 possesses a putative steric gate residue F428 but its benzene ring almost parallels and stacks to the ribose ring (Figure 9.1) [72]. Thus, it is unclear how hRev1 discriminates against rNTPs. In general, the kinetic basis for rNTP discrimination by most DNA polymerases is via weakened binding and slowed incorporation of rNTPs. Using CTP analogs, we showed that the mechanism of ribonucleotide selection employed by hRev1 is influenced by both the size and orientation of the 2′ group (Table 9.4). With varying
sizes of the 2′ substituent, the Kd values for 2′-F-CTP, rCTP, araCTP, and 2′-OCH3-CTP were not affected significantly. This is probably due to the favorable hydrogen bonding interactions between residue R357 of hREV1 and the cytosine base which compensated for the steric effect of the 2′ substituent. However, the binding of GemCTP to hRev1•D- G DNA was perturbed the most with a 13-fold lower affinity than dCTP. The geminal difluoro group of GemCTP has more electronegativity than the deoxyribose of dCTP, and an embedded GemCMP residue in duplex DNA adopts a C3′-endo pucker [343]. These may affect how GemCTP was positioned in the active site and how it interacted with R357 and F428 of hRev1, leading to the lower affinity. A similar conclusion has been drawn for the human mitochondrial DNA polymerase γ incorporating GemCTP [344]. In comparison, Table 9.4 shows that the kp variation is much larger than the Kd range for the CTP analogs. If the ribose 2′substituent either has a small size (e.g. 2′-F in both 2′-F-CTP and GemCTP) or is orientated above the ribose ring (e.g. 2′-OH in araCTP), it has a small
impact on the kp value. Contrary to these trends, the kp values for rCTP and 2′-OCH3-CTP are 200- to 2,000-fold lower than that of dCTP. Together, these results suggested that, inconsistent with the general kinetic trends observed with other DNA polymerases (see above), the steric clash of the 2′-OH of the incoming rCTP with F428 of hRev1 mostly
215 impacts the incorporation step (kp) rather than the ground-state binding step (Kd). In addition to the major contribution of the templating base dG to the high dCTP incorporation efficiency (see above discussion), our kinetic data further dissected the contribution of each chemical moiety toward the high efficiency of dCTP incorporation
catalyzed by hRev1: the ribose 2′-H of dCTP significantly contributes to the fast kp while the cytosine of dCTP contributes to the low Kd for dCTP binding. We are currently elucidating the kinetic mechanism of dCTP incorporation in order to mechanistically
understand how these chemical moieties of dCTP influence its kp and Kd.
9.5 Future Directions
Ideally, an isosteric dCTP analog should be used to better understand the role of hydrogen bonding between R357 and an incoming dNTP. In addition, it would be interesting to replace R357 with different amino acid residues (e.g. Lys, Asn, Gln, Asp, Glu, Leu, or Ala). Arg is unable to form hydrogen bonds with the Watson-Crick face of dATP, dGTP, and dTTP [345]. However, Asn or Gln can form hydrogen bonds with the Watson-Crick face of the four dNTPs [345]. Thus, it would be interesting to determine if mutating a single amino acid side chain could alter the nucleotide specificity of Rev1. Lastly, it would be useful to know the lesion bypass capacity of Rev1, especially DNA damage that targets guanine (e.g. 8-oxo-7,8-dihydro-dG).
216 9.6 Tables
Table 9.1 Sequences of the D-DNA substratesa D-G 5’-CGCAGCCGTCCAACCAACTCA-3’ 3’-GCGTCGGCAGGTTGGTTGAGTGTCAGCTAGGTTACGGCAGG-5’ D-A 5’-CGCAGCCGTCCAACCAACTCA-3’ 3’-GCGTCGGCAGGTTGGTTGAGTATCAGCTAGGTTACGGCAGG-5’ D-T 5’-CGCAGCCGTCCAACCAACTCA-3’ 3’-GCGTCGGCAGGTTGGTTGAGTTTCAGCTAGGTTACGGCAGG-5’ D-C 5’-CGCAGCCGTCCAACCAACTCA-3’ 3’-GCGTCGGCAGGTTGGTTGAGTCTCAGCTAGGTTACGGCAGG-5’ D-G Gap 5’-CGCAGCCGTCCAACCAACTCA AGTCGATCCAATGCCGTCC-3’ 3’-GCGTCGGCAGGTTGGTTGAGTGTCAGCTAGGTTACGGCAGG-5’ aEach DNA substrate is composed of a 5′-radiolabeled 21-mer and a 41-mer template which has the unique template bases in bold. D-G Gap has a 5′-phosphorylated 19-mer.
217
Table 9.2 Kinetic parameters for nucleotide incorporation into D-DNA catalyzed by hRev1 at 37 °C. k K k /K Efficiency Fidelityb dNTP p d p d (s-1) (μM) (μM-1s-1) ratioa Template dG (D-G) dCTP 22.4 ± 0.9 2.2 ± 0.3 10 dATP 0.050 ± 0.004 70 ± 20 7.1 × 10-4 1.4 × 104 7.0 × 10-5 dGTP 6.3 ± 0.3 90 ± 10 7.0 × 10-2 1.5 × 102 6.8 × 10-3 dTTP 0.88 ± 0.06 22 ± 7 4.0 × 10-2 2.5 × 102 3.9 × 10-3 Template dA (D-A) dTTP 0.092 ± 0.007 44 ± 6 2.1 × 10-3 4.9 × 103 dCTP 1.87 ± 0.05 9.5 ± 0.8 2.0 × 10-1 5.2 × 101 9.9 × 10-1 Template dT (D-T) dATP 0.00235 ± 0.00008 2.7 ± 0.4 8.7 × 10-4 1.2 × 104 dCTP 1.93 ± 0.05 35 ± 2 5.5 × 10-2 1.8 × 102 9.8 × 10-1 Template dC (D-C) dGTP 0.0073 ± 0.0010 30 ± 10 2.4 × 10-4 4.2 × 104 dCTP 3.4 ± 0.2 120 ± 10 2.8 × 10-2 3.6 × 102 9.9 × 10-1 Template dG (D-G Gap) dCTP 11 ± 1 8 ± 2 1.4 7.4 a Calculated as (kp/Kd)dCTP:D-G/(kp/Kd)dNTP:dN. b Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
218
Table 9.3 Kinetic parameters for non-natural nucleotide analog incorporation into D-G DNA catalyzed by hRev1 at 37 °C. k K k /K Efficiency dNTP p d p d (s-1) (μM) (μM-1s-1) ratioa dCTP 22.4 ± 0.9 2.2 ± 0.3 10 dATP 0.050 ± 0.004 70 ± 20 7.1 × 10-4 1.4 × 104 dNITP 0.0457 ± 0.0006 15.8 ± 0.6 2.9 × 10-3 3.5 × 103 dPTP 0.0228 ± 0.0008 25 ± 3 9.1 × 10-4 1.1 × 104 a Calculated as (kp/Kd)dCTP/(kp/Kd)dNTP.
219
Table 9.4 Kinetic parameters for CTP analog incorporation into D-G DNA catalyzed by hRev1 at 37 °C.
NTP kp Kd kp/Kd Sugar (s-1) (μM) (μM-1s-1) selectivitya dCTP 22.4 ± 0.9 2.2 ± 0.3 10 rCTP 0.098 ± 0.002 2.7 ± 0.2 3.6 × 10-2 280 araCTP 6.3 ± 0.5 4 ± 1 1.6 6 2′-F-CTP 19.2 ± 0.5 3.5 ± 0.4 5.5 2 GemCTP 6.8 ± 0.4 29 ± 6 2.3 × 10-1 43 -3 2′-OCH3-CTP 0.0122 ± 0.0006 8 ± 1 1.5 × 10 6,700 a Calculated as (kp/Kd)dCTP/(kp/Kd)analog.
220 9.7 Figures
Figure 9.1 Active site of hRev1. Important active site residues that interact with an incoming dCTP or the templating base dG are shown (PDB 3GQC). The dashed lines represent hydrogen bonds, and the four magnesium ions are shown as gray spheres.
221 A 30
25
20
15
10 Product (nM)
5
0 00.511.522.53 Time (s)
B 25
20
) 15 -1 (s
obs 10 k
5
0 0 5 10 15 20 25 30
dCTP (μM)
Figure 9.2 Concentration dependence on the pre-steady state rate constant of deoxycytidyl transferase catalyzed by hRev1. (A) A pre-incubated solution of hRev1 (120 nM) and 5′-[32P]-labeled D-6T (30 nM) was rapidly mixed with increasing concentrations of dCTP •Mg2+ (0.2 μM, z; 0.5 μM, {; 1 μM, ; 2 μM, ; 5 μM, S; 10 μM, U; and 25 μM, ) for various time intervals. The solid lines are the best fits to a single-exponential equation which determined the observed rate constant, kobs. (B) The kobs values were plotted as a function of dCTP concentration. The data (z) were then fit -1 to a hyperbolic equation, yielding a kp of 22.4 ± 0.9 s and a Kd of 2.2 ± 0.3 μM.
222
Figure 9.3 Chemical structures of nucleotide analogs. (A) non-natural nucleotide analogs and (B) CTP analogs used in this work.
223 Chapter 10: Mechanism of Double-Base Lesion Bypass Catalyzed by a Y-family DNA Polymerase
10.1 Introduction
cis-Diamminedichloroplatinum(II) (cisplatin, DDP) is a potent anticancer drug that is effective for the treatment of testicular, ovarian, head, neck, and nonsmall cell lung cancers. During the biotransformation of cisplatin in vitro and in vivo, cisplatin reacts with the N7 position of purines to generate several possible adducts: cis-
[Pt(NH3)2{d(GpG)-N7(1),-N7-(2)}] intrastrand cross-links (cisplatin-d(GpG)), cis-
[Pt(NH3)2{d(ApG)-N7(1),-N7-(2)}] intrastrand cross-links, cis-[Pt(NH3)2{d(GpNpG)- N7(1),-N7-(3)}] intrastrand cross-links, 1,3-interstrand cross-links with guanines and monofunctional cross-links with guanine [346-349]. These cisplatin–DNA adducts can severely inhibit DNA replication, thereby highlighting this cytotoxic effect as an important mode of drug action. Upon encountering a cisplatin–DNA adduct in vitro, replicative DNA polymerases, e.g. eukaryotic DNA polymerases α [350, 351], δ [351] and ε [352], stall while most DNA repair and lesion bypass polymerases can traverse the lesion. Y-family DNA polymerases, characterized by low fidelity and processivity, are notorious for bypassing DNA lesions. Of the four Y-family members in humans, DNA polymerase η (Pol η) bypasses cisplatin–DNA adducts in a relatively efficient and error- free manner in vitro [309-312], and cell-based assays provide additional evidence that Pol η is involved in the bypass of this adduct in vivo [353-355]. DNA polymerases ι (Pol ι) [356] and κ (Pol κ) [357, 358] are impeded by cisplatin–DNA adducts, however, it remains to be explored whether bypass is possible via the two-polymerase model of lesion bypass [20, 359]. Recent studies provide evidence that Rev1, both a deoxycytidyl transferase and structural factor during replication, modulates cisplatin mutagenicity
224 [360, 361], but the catalytic activity appears to be dispensable [316]. Other relevant eukaryotic DNA polymerases implicated in the bypass of cisplatin–DNA adducts are DNA polymerases γ [362], ζ [362-364], β [311, 312, 351, 365-367] and μ [368]. Finally, a variety of cellular processes have been proposed to promote cancer cell survival, which contributes to clinical drug resistance associated with cisplatin. Some of these in vivo processes are the following: reduced drug uptake, enhanced drug inactivation, increased DNA repair, disabled apoptotic signaling machinery and translesion DNA synthesis (TLS) [see reviews in references [369-373] for a comprehensive overview of postulated mechanisms]. Thus, to circumvent the hurdles of drug resistance and to design improved anticancer drugs, it is imperative to understand the effect of cisplatin as it pertains to lesion bypass at the molecular level.
For this study, we investigated the mechanistic basis of lesion bypass catalyzed by Sulfolobus solfataricus P2 DNA polymerase IV (Dpo4) for a single, site-specifically placed cisplatin-d(GpG) adduct using pre-steady state kinetics. Utilizing this methodology, we have previously established (i) the minimal kinetic mechanism and fidelity of Dpo4 incorporating a single nucleotide into undamaged DNA [105, 111] and (ii) the mechanistic basis of Dpo4 bypassing an abasic site, which is a prototype single- base lesion [374]. To provide a deeper mechanistic understanding of how Y-family DNA polymerases catalyze the bypass of other DNA lesions, we used the cisplatin-d(GpG) adduct as a model double-base lesion and Dpo4 as a model Y-family DNA polymerase. Among the aforementioned cisplatin–DNA adducts, cisplatin-d(GpG) is the predominant species (~65%) and has been correlated with clinical efficacy of the drug [347, 375, 376]. Although Dpo4 shares more sequence similarity with Pol κ as a DinB homolog, one study suggests that the lesion bypass properties of Dpo4 are more similar to Pol η [100]. Consequently, establishing the kinetic mechanism of Dpo4 bypassing the cisplatin- d(GpG) adduct may (i) establish the mechanistic basis of double-base lesion bypass, (ii) illuminate the molecular basis of drug resistance due to cisplatin bypass, (iii) assess the mutagenic potential of inducing secondary malignancies and (iv) enable scientists to design more effective anticancer drugs.
225 10.2 Materials and Methods
Reaction buffers Optimized reaction buffer D contains the following: 50mM HEPES (pH 7.5 at 37 °C),
5mM MgCl2, 50mM NaCl, 5mM DTT, 10% glycerol, 0.1mM EDTA and 0.1 mg/ml bovine serum albumin (BSA) [105]. For the gel mobility shift assay, reaction buffer E
contains the following: 50mM Tris–Cl (pH 7.5 at 23 °C), 5mM MgCl2, 50mM NaCl, 5mM DTT, 10% glycerol and 0.1 mg/ml BSA. All reported concentrations were the final concentrations upon mixing. All reactions, unless noted, were performed at 37 °C.
DNA substrates The cisplatinated-DNA oligonucleotide (Table 10.1) was modified, ligated and purified previously [377]. All DNA primers (Table 10.1) were prepared previously [377] except for the 21-mer purchased from Integrated DNA Technologies, Inc. (Coralville, IA, USA) and subsequently gel purified. The DNA primers were 5′-radiolabeled with [γ-32P]ATP (GE Healthcare, Picataway, NJ, USA) and Optikinase (USB) [105]. To anneal, the [γ- 32P]-radiolabeled DNA primer and unlabeled template were combined at a 1:1.15 molar ratio, respectively, heated to 85 °C for 6 min and cooled slowly to room temperature.
Running start assay Using a rapid-chemical quench flow apparatus (KinTek), a solution of pre-incubated Dpo4 (100 nM) and 5′-[32P] DNA (100 nM) was mixed with an equal volume (15 μl) of all four dNTPs (200 μmM each) in buffer D and quenched with 0.37M EDTA at times ranging from ms to min. The incorporation pattern was resolved via denaturing polyacrylamide gel electrophoresis (PAGE).
Electrophoretic mobility shift assay (EMSA) DNA To determine the Kd for the Dpo4•DNA binary complex, Dpo4 (15–600 nM) was added to a solution containing 5′-[32P]-labeled DNA (100 nM) in buffer E and allowed to equilibrate for 15 min at 23 °C. Then, the binary complex was separated from unbound
226 DNA using a 4.5% native polyacrylamide gel with a running buffer as described previously [374] and quantitated using a PhosphorImager 445 SI (Molecular Dynamics, Sunnyvale, CA, USA). A plot of binary complex formation versus the concentration of Dpo4 was fit to Equation 1 DNA DNA 2 1/2 [E•DNA] = 0.5(Kd + E0 + D0) – 0.5[(Kd + E0 + D0) – 4E0D0] (1)
where E0 and D0 represent the active enzyme and DNA concentrations, respectively.
Determination of the kp and Kd of an incoming nucleotide A pre-equilibrated solution of Dpo4 (120 nM) and 5′-[32P]-labeled DNA (30 nM) in buffer D was combined with increasing concentrations of a single dNTP (25–1600 μM) in buffer D. Aliquots of the reaction mixtures were quenched at various times using 0.37 M EDTA. For fast reaction times, a rapid chemical-quench flow apparatus was used. Reaction products were resolved using sequencing gel electrophoresis and quantitated with a PhosphorImager 445 SI. The time course of product formation at each nucleotide concentration was fit to a single-exponential equation (Equation 2)
[Product] = A[1 – exp (–kobst)] (2) using a nonlinear regression program, KaleidaGraph (Synergy Software, Essex Junction, VT, USA). A represents the reaction amplitude. The extracted values for the observed 2+ rate constant (kobs) of nucleotide incorporation were plotted as a function of dNTP•Mg concentration and fit to a hyperbolic equation (Equation 3)
kobs = kp[dNTP]/(Kd + [dNTP]) (3) which resolved the kp and Kd values for nucleotide incorporation. The substrate
specificity constant (kp/Kd), efficiency ratio [(kp/Kd)control/(kp/Kd)damaged], and fidelity
{(kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect]} were then calculated.
Biphasic kinetic assays A pre-equilibrated solution of Dpo4 (120 nM) and 5′-[32P]-labeled DNA (30 nM) in buffer D was mixed with a DNA trap [5 μM, 21-mer/41CTL D-1 [105]] and dNTP (1.0 mM) in buffer D supplemented with Mg2+, which may be chelated by the DNA trap as previously described [374]. A rapid chemical-quench flow apparatus was used to quench
227 aliquots of the reaction mixtures at various times using 0.37 M EDTA. Reaction products were resolved and quantitated as described earlier. The concentration of product formation was graphed as a function of time and fit to a double-exponential equation (Equation 4)
[Product] = E0A1[1 – exp (–k1t)] + E0A2[1 – exp (–k2t)] (4)
using KaleidaGraph. E0 represents the total enzyme concentration, A1 and A2 represent
the first and second phase reaction amplitudes, respectively, and k1 and k2 represent the first and second phase rate constants, respectively.
10.3 Results
Bypass of a cisplatin-d(GpG) adduct We have previously constructed a 44-mer DNA template (44DDP) containing a site- specific cisplatin-d(GpG) adduct (Table 10.1) [377]. To generate a nucleotide incorporation profile using this adduct-containing DNA substrate, we first performed a ‘running start’ assay (Section 10.2) to visualize the extension of the 5′-[32P]-labeled 19- mer/44DDP (Table 10.1) catalyzed by Dpo4 in the presence of all four dNTPs (Figure 10.1 part (B)). Our results showed that Dpo4 bypassed the single cisplatin-d(GpG) adduct, as full-length product (44-mer) was detected at 60 s (Figure 10.1 part (B)). Although Dpo4 synthesized full-length product, the significant accumulation of intermediate products revealed two distinct pause sites (23- and 24-mers) corresponding to nucleotide incorporations opposite the cisplatin-d(GpG) adduct. In comparison, Dpo4 did not pause at these positions in the control experiment with an undamaged DNA substrate (19-mer/44CTL, Table 10.1) in which full-length product was generated within 10 s (Figure 10.1 part (A)). The purity of 44DDP was examined to ensure that the bypass products in Figure 10.1 part (B) were not due to a subpopulation of unmodified or deplatinated 44-mer (Figure 10.2). Also notable, the observation of 45- and 46-mer products suggested Dpo4 catalyzed template-independent nucleotide incorporations (Figure 10.1) [194]. However, this result warrants further investigation due to the
228 possibility of nucleotide deletions, nucleotide additions or other complex transactions occurring in the vicinity of the lesion as previously demonstrated [311, 365, 378-382].
Binding of Dpo4 to cisplatin-modified DNA Various structural studies reveal a significant bend in the DNA, ~40–80°, which may preclude binary complex formation [383-385]. Since the two strong pause sites occur opposite the cisplatin-d(GpG) lesion, it is possible that a weaker binding affinity of Dpo4 to DNA may be a contributing factor. Therefore, we determined the equilibrium DNA dissociation constant (Kd ) of several binary complexes using the EMSA (Section DNA 10.2). EMSA has been proven to be a reliable assay for estimating the Kd value of the Dpo4•DNA binary complex [194, 374]. By varying the primer length, a series of DNA substrates was designed to mimic the progression of DNA synthesis catalyzed by Dpo4 as it approached, encountered and bypassed the damaged site. For example, to measure DNA the Kd value at the second pause site (Figure 10.1 part (B)), we titrated increasing concentrations of Dpo4 into a fixed concentration of 5′-[32P]-labeled 24-mer/44DDP (Table 10.1). After equilibration, the binary complex Dpo4•24-mer/44DDP and free DNA were separated via native PAGE (Figure 10.3 part (A)). Next, the concentration of binary complex formation was plotted versus Dpo4 concentration (Figure 10.3 part (B)) DNA and fit to a quadratic equation (Equation 1 in 10.2), which yielded a Kd of 26 ± 3 nM DNA (Table 10.2). Similarly, the Kd for the Dpo4•24-mer/44CTL (Table 10.1) binary DNA DNA complex was measured to be 8 ± 1 nM, and the affinity ratio [(Kd )damaged/(Kd )control] was calculated to be 3.3-fold (Table 10.2). Analysis of the first strong pause site (23- mer/44DDP) and the first downstream, nonpause site (25-mer/44DDP) revealed a similar DNA effect on the Kd value (Table 10.2). Dpo4 binding upstream of the cisplatin lesion was unaffected, yet the binding affinity downstream was not restored until the second downstream site (Table 10.2). These modest, ~3-fold lower affinity values indicated that the overall binding affinity of Dpo4 to a cisplatin-d(GpG) adduct was not significantly affected and was localized to the vicinity of the lesion. Furthermore, these results suggested that the association and dissociation rates of DNA from the Dpo4•DNA complex at or near the double-base lesion were not significantly altered, and the pausing
229 of Dpo4 observed in Figure 10.1 was not due to the faster dissociation of the Dpo4•DNA complex. This conclusion was supported by the biphasic kinetics of nucleotide incorporations at the pause sites in the presence of a DNA trap (see Biphasic kinetics section below). Thus, we hypothesized that the structurally distorted DNA significantly decreased the incorporation efficiency of an incoming nucleotide at the aforementioned pause sites.
Efficiency of nucleotide incorporation opposite a cisplatin-d(GpG) adduct Transient state kinetics performed under single-turnover conditions measures the
maximum incorporation rate (kp) and the equilibrium dissociation constant (Kd) of nucleotide incorporation which are the two defining kinetic parameters of catalytic
efficiency or substrate specificity constant (kp/Kd). To resolve the kp and Kd at the second pause site, a solution containing Dpo4 (120 nM) was pre-incubated with 5′-[32P]-labeled 24-mer/44DDP (30 nM) before initiating the reaction with increasing concentrations of dCTP for various times (Section 10.2). A primary plot of product concentration versus
time conformed to a single-exponential fit (Equation 2 in 10.2) so that the kobs values were extracted for each dCTP concentration (Figure 10.4 part (A)). Then, a secondary
plot of the kobs values versus dCTP concentration was fit to Equation 3 which yielded a kp -1 of 0.043 ± 0.004 s and a Kd of 588 ± 145 μM (Figure 10.4 part (B) and Table 10.3). The calculated substrate specificity constant of 7.3 × 10-5 μM-1s-1 was ~860-fold lower than dCTP incorporation into undamaged 24-mer/44CTL (Tables 10.3 and 10.4). Any bias due to a DNA sequence-dependent effect was attenuated by determining the kinetic parameters for all correct incorporations into 44CTL, and these results indicated that the incorporation efficiency varied over a 30-fold range (Table 10.4). Furthermore, the substrate specificity constant for dCTP incorporation into 23-mer/44DDP was reduced by 72-fold relative to the control DNA substrate 23-mer/44CTL (Tables 10.3 and 10.4).
Please note, these are apparent kp and Kd values at the strong pause sites due to the detection of biphasic kinetics of nucleotide incorporation at these sites. Misincorporation at the two strong pause sites opposite the cisplatin cross-linked guanines followed similar trends in which (i) the order of preferred dNTP incorporation was dATP > dTTP > dGTP,
230 -6 -7 -1 -1 (ii) the kp/Kd values were in the range of 10 to 10 μM s and (iii) the fidelity was in the low range of 10-4 to 10-3 at the 3′-dG site and 10-3 to 10-2 at the 5′-dG site (Table 10.3). Collectively, these kinetic data supported our hypothesis that a decreased nucleotide incorporation efficiency explained the significant product accumulation induced by the cisplatin–DNA adduct (Figure 10.1 part (B) and Table 10.3).
Efficiency of nucleotide incorporation proximal to the cisplatin-d(GpG) adduct In parallel single-turnover experiments, the substrate specificity constants for events both upstream and downstream from the lesion were kinetically examined using a series of DNA primers with the same cisplatinated or control DNA templates (Table 10.1). The efficiency ratio, defined as the substrate specificity constant for the control DNA divided by the substrate specificity constant for damaged DNA, provides a foundation for our kinetic comparisons. All upstream events were essentially normal (efficiency ratios of 1.1 and 1.5), while the downstream events revealed an intriguing trend (Figure 10.5 part (A)). Interestingly, the incorporation efficiency did not gradually decrease as a linear function before resuming to normal. Instead, the most significant downstream perturbation occurred at the -2 and -3 positions relative to the cisplatinated 5′-dG (Table 10.3 and Figure 10.5 part (A)). At these ‘weak’ pause sites, the efficiency ratio for correct dNTP incorporation into 26-mer/44DDP and 27-mer/44DDP was decreased by 36- and 17-fold, respectively, but there was not a dramatic effect on fidelity (Table 10.3 and Figure 10.5 part (B)). In addition, these ‘weak’ pause sites were observable in the running start assay as a minor accumulation of intermediate products (26- and 27-mers) (Figure 10.1 part (B)). Other notable sites in which Dpo4 appeared to pause during dNTP incorporation include 25-mer/44DDP, 29-mer/44DDP and 30-mer/44DDP (Figure 10.5 part (A)). Slowing the rate of maximum nucleotide incorporation lead to a waning of the substrate specificity constant in an unprecedented cyclic pattern during the downstream extension steps. These observations reinforced the kinetic impact of a severely distorted DNA molecule within or near the active site of a DNA polymerase [383-385]. Overall, the catalytic efficiency of Dpo4 synthesizing on a cisplatin-modified DNA substrate returned to normal after seven downstream incorporations (Table 10.3 and Figure 10.5 part (A)).
231 Biphasic kinetics Biphasic kinetic analysis is useful for extracting the rate constants and amplitudes for reactions in which distinct phases may be hidden during dNTP incorporation due to the inability to isolate different DNA polymerase binding modes [374, 377]. The amplitude of the first phase represents the fraction of E•DNA bound in a productive mode, whereas the amplitude of the second phase represents the nonproductively bound fraction that is slowly converted into a productive mode (Scheme 10.1) [374, 377]. Thus, examining biphasic kinetics advances our mechanistic understanding of the events occurring at the active site of a Y-family DNA polymerase upon encountering a cisplatin–DNA adduct. To perform this assay, a pre-incubated solution of Dpo4 (120 nM) and 5′-[32P]-labeled 23-mer/44DDP (30 nM) was mixed with a solution of 21-mer/41CTL D-1 DNA trap (5 μM) and dCTP•Mg2+ (1.0 mM) for various times before being quenched with 0.37 M EDTA (Section 10.2). The purpose of the unlabeled D-1 trap is to sequester any Dpo4 molecules that dissociate from the damaged DNA substrate so that product formation is solely due to a single-binding event. The effectiveness of the D-1 DNA trap was tested and confirmed to be sufficient (Figure 10.6). A plot of product concentration as a function of time (Figure 10.7) was fit to Equation 4 in 10.2, which yielded the following -1 kinetic parameters: A1 = 2.1 ± 0.1nM (7%), k1 = 10 ± 2 s , A2 = 19 ± 1 nM (63%), k2 = 0.0009 ± 0.0001 s-1 (Table 10.5). Indeed, biphasic kinetics was observed at the cisplatin cross-linked 3′-dG, whereby nucleotide incorporation initially proceeded with a fast phase of low amplitude followed by a slow phase of large amplitude. It is conceivable that the fast phase was due to unmodified or deplatinated 44-mer; however, this is unlikely because (i) 44DDP has been purified to apparent homogeneity (Figure 10.2) and (ii) platinum adducts are stable under harsh temperature and pH conditions [386]. Interestingly, a different biphasic trend emerged at the second strong pause site as evident -1 by these kinetic values: A1 = 1.11 ± 0.01nM (3.7%), k1 = 4.0 ± 0.7 s , A2 = 3.2 ± 0.4 nM -1 (11%), k2 = 0.00016 ± 0.00003 s (Table 10.5 and Figure 10.7). These results indicated that a minute portion of Dpo4 rapidly incorporated dCTP into the Dpo4•24-mer/44DDP complex, while most of Dpo4 either dissociated or failed to achieve a catalytically competent complex. Thus, these biphasic kinetic assays provided a basis for explaining
232 the more significant reduction in catalytic efficiency (72- versus 860-fold) for the two strong pause sites. As proof of principle, nucleotide incorporation into a non-pause site, 21-mer/44DDP, exhibited a single fast phase with a rate constant of 5.7 ± 0.6 s-1 in the presence of an unlabeled DNA trap (data not shown).
10.4 Discussion
Kinetic basis of cisplatin resistance mediated by a Y-family DNA polymerase The clinical effectiveness of cisplatin chemotherapy is limited by drug resistance in which the process of TLS is a likely factor [361, 363, 364, 387, 388]. The newly discovered Y-family DNA polymerases are suspected to participate in the bypass of platinated-DNA adducts [350-352]. Of the four human Y-family polymerases, Pol η is the likely candidate responsible for bypassing platinated-DNA adducts in vivo [354, 355]. Interestingly, results from P. Blum’s laboratory showed that a Dpo4-knockout cell line of S. solfataricus is more sensitive to cisplatin treatment than the wild-type strain (personal communication). This suggested that Dpo4, the lone Y-family DNA polymerase in S. solfataricus, participates in the resistance of cisplatin in vivo. To explore the kinetic basis of cisplatin resistance caused by the Y-family enzyme and to better understand double- base lesion bypass, we investigated the bypass of a site-specifically placed cisplatin- d(GpG) adduct catalyzed by Dpo4. Consistent with the in vivo observation, Figure 10.1 part (B) showed that Dpo4 was able to incorporate nucleotides opposite and extend from the cisplatin-d(GpG) adduct. However, the observation of intermediate product accumulation ignited further interest into elucidating the mechanistic basis of Dpo4 pausing during the insertion step as well as the lack of significant pausing during the downstream extension steps.
X-ray and NMR structural studies reveal numerous distortions when cisplatin forms a covalent linkage with two neighboring guanines in duplex DNA [383-385]. Notable structural consequences in accommodating an intrastrand cross-link include the
233 following: (i) DNA is significantly bent (~40–80°) over several base pairs, (ii) the minor groove is widened and flattened, (iii) there is a 26–49° dihedral angle between coordinated d(GpG) bases, (iv) the platinum atom is displaced from the guanine plane by ~1 Å , (v) A- and B-forms of DNA are observed, (vi) the d(GpG) bases are propeller twisted yet maintain Watson–Crick hydrogen bonding with the complementary strand and (vii) the amine group of cisplatin is hydrogen bonded to the phosphate oxygen of the DNA backbone [384, 385]. Despite the distorted DNA structure, the binding affinity of the binary complex was only modestly weakened (~3-fold) at the lesion site and one base downstream of the lesion (Table 10.2). Instead, DNA structural perturbations had a more profound effect on nucleotide incorporation efficiency and fidelity.
All incorporation events upstream of the lesion were catalyzed by Dpo4 with an efficiency comparable to undamaged DNA (Table 10.3). Upon encountering the cisplatin-coordinated 3′-dG, Dpo4 correctly inserted dCTP with a 72-fold decrease in efficiency relative to an undamaged dG base (Tables 10.3 and 10.4). The kinetic parameters suggested that weak dCTP binding (11-fold) accounted more for the reduction of catalytic efficiency than did the rate of dCTP incorporation (7-fold). However, the substrate specificity constant of dCTP incorporation at the cisplatin-coordinated 5′-dG was decreased by 860-fold due to a decrease in the rate of incorporation by 105-fold (4.5 s-1 → 0.043 s-1) (Tables 10.3 and 10.4). The weak apparent ground-state binding affinity
(1/Kd) of dCTP at both strong pause sites may indicate poor base stacking. Using pyrene nucleoside 5′-triphosphate, we have shown previously that base stacking governs the binding affinity of a nucleotide opposite an abasic site [374], undamaged DNA of a recessed primer/template substrate [194], and undamaged DNA of a blunt-end substrate [194]. Due to the covalent linkage of cis-diammineplatinum(II) with the adjacent guanine residues, the orientation of dCTP, the templating base, and the 3′-OH may preclude normal base stacking interactions, thereby weakening the binding affinity of an incoming nucleotide.
234 Interestingly, as we completed our kinetic studies reported here, an X-ray crystallographic study of yeast Pol η (yPol η) in the presence of a cisplatin-d(GpG) intrastrand cross-link and dCTP or dATP was published [290]. This study provided a structural basis for the two aforementioned slow translesion steps: the incoming nucleotide’s distance from the 3′-OH of the modeled primer terminus is unfavorably long (5 Å and 7.5 Å for the first and second elongation steps, respectively) [290]. However, the inefficient bypass catalyzed by Dpo4 may originate from a different combination of subtle structural factors. Further kinetic analysis uncovered two distinct kinetic phases occurring at the active site of Dpo4 in the presence of a DNA trap. Bypassing the cisplatin cross-linked 3′-dG showed the rapid incorporation (10 s-1) of dCTP for a small percentage (7%) of the Dpo4•23-mer/44DDP complexes and then a larger percentage (63%) gradually turned over (0.0009 s-1) to product (Table 10.5 and Figure 10.7). These P results suggested the existence of two distinct binary complexes: E•Dn (productive) and N E•Dn (nonproductive) (Scheme 10.1). Initially, 7% of the Dpo4-bound species achieved a productive binary complex that was competent for catalysis, while 63% existed as a nonproductive binary complex that slowly converted into a productive state without dissociating (Table 10.5 and Figure 10.7). For the remaining 30%, which did not generate product, these Dpo4-bound species were either arrested in a ‘dead-end’ state or simply dissociated and diluted by the non-radiolabeled DNA trap. Structural evidence of yPol η in a pre-elongation and elongation ternary complex at the cisplatin-coordinated 3′-dG position supports the existence of the nonproductive and productive modes, respectively, described by the kinetic data herein [290]. Unfortunately, the biphasic kinetic assay cannot distinguish between the presence of binary and ternary complexes in a productive or nonproductive state due to the rapid binding of an incoming nucleotide. Thus, in light of the ternary species observed in the X-ray crystal structures of yPol η, the kinetic scheme derived from biphasic kinetic analysis (Scheme 10.1) has been expanded to include the possible existence of productive and nonproductive ternary complexes as shown in Scheme 10.2 [290]. Lastly, it is possible that there is a heterogeneous mixture of various nonproductive binding modes as binary and/or ternary complexes.
235 The biphasic kinetic trace during the bypass of the cisplatin-coordinated 5′-dG revealed a different trend than the cisplatin-coordinated 3′-dG as catalyzed by Dpo4 (Table 10.5 and Figure 10.7). Although dCTP was still rapidly incorporated (4 s-1) by a small percentage (3.7%) of productively bound Dpo4 species, a greater percentage (~96%) was either present as a nonproductive binary complex (11%) or dissociated in favor of binding to the nonradiolabeled DNA trap (85%) (Table 10.5 and Figure 10.7). HIV type 1 reverse transcriptase (HIV-1 RT), an enzyme with low fidelity and processivity, exhibits a similar biphasic kinetic trend as Dpo4 in which the total amplitude for the first (~25%) cross- linked guanine is greater than the second (~2%) [377]. In contrast, the total amplitudes for an exonuclease-deficient mutant of T7 DNA polymerase (T7 exo-) were 2.6%, 3.0%, and 4.6% for correct nucleotide incorporation into 23-mer/44DDP, 24-mer/44DDP, and 25-mer/44DDP, respectively [377]. Together, these biphasic kinetic data illustrate the spectrum of cisplatin-d(GpG) bypass in which the rigid active site of a replicative polymerase (e.g. T7 exo-) is blocked, while flexible active sites are more suitable for TLS (e.g. Dpo4 and HIV-1 RT). Despite possessing flexible active sites, not all human Y- family polymerases can independently bypass a cisplatin–DNA adduct in vitro [309-312, 356-358]. Thus, active site flexibility does not insure lesion bypass. Presumably, the flexible active site of Pol η can readily accommodate productive binary and/or ternary complex formation, while the active sites of Pol ι and Pol κ fail to achieve proper alignment for nucleotidyl transfer. Interestingly, the same mechanism shown in Scheme 10.1 occurs during the bypass of an abasic site, a model single-base lesion, catalyzed by Dpo4 [374]. Thus, Schemes 10.1 and 10.2 represent a universal kinetic mechanism for the bypass of single-base and double-base DNA lesions catalyzed by a variety of translesion DNA polymerases including the Y-family enzymes and HIV-1 RT.
From the overall reaction rate measured under single-turnover conditions, the individual contribution of the productive and nonproductive binary complexes can be extracted, since product formation under single-turnover conditions is comprised of all productive and nonproductive binding states. For example, opposite the cisplatin cross-linked 3′-dG, contributions from the fast phase of nucleotide incorporation [(10 s-1) × (7% reaction
236 amplitude)] and the slow phase [(0.0009 s-1) × (63% reaction amplitude)] are predicted to yield an overall product formation rate of 0.70 s-1. Since the biphasic kinetic assays in Figure 10.7 were performed at sub-saturating concentrations of dCTP (1.0 mM), the observed rate at the first strong pause site was calculated to be 0.70 s-1 based on the
apparent kp and Kd values for dCTP (Table 10.3) substituted into Equation 3. This value of 0.70 s-1 matched 0.70 s-1 estimated from the biphasic kinetic parameters. Thus, the observed rate constant extracted from single-turnover kinetic analyses was dominated by
the rate constant of the fast phase, k1. A similar connection between single-turnover and biphasic kinetic assays was obtained for abasic lesion bypass [374]. Due to the complication of at least two phases occurring within a single turnover, an apparent rate of nucleotide incorporation and an apparent equilibrium dissociation constant of an incoming nucleotide was obtained at the strong pause sites (Table 10.3). An advantage of the biphasic kinetic assay is that the fast and slow phases can be discerned in the presence of a DNA trap. Thus, the combination of single-turnover experiments and the DNA trap assay is a powerful approach to elucidate the detailed kinetic mechanisms of lesion bypass.
To complete the TLS process, Dpo4 was able to extend the primer beyond the lesion, although, there were downstream events in which Dpo4 appeared to pause slightly. These ‘weak’ pause sites appeared when incorporating dNTPs 1, 2, 3, 5 and 6 positions downstream from the cisplatin-coordinated 5′-dG (Figures 10.1 part (B) and 10.5 part (A)). Differential domain interactions of Dpo4 with the damaged DNA substrate may account for this unusual cyclic pattern. For instance, the little finger, thumb and/or linker region may be orienting the cisplatinated DNA into the most stable conformation, but the templating base within the active site interior may not be properly aligned for nucleotidyl transfer. As the DNA helix turns and the distance between the lesion and the Dpo4 active site increases, this downstream effect eventually dissipated after seven nucleotide incorporations (Table 10.3 and Figure 10.5 part (A)). Steady-state kinetic data show that human Pol η efficiently inserts dCTP opposite cisplatin-d(GpG) adducts, but the extension step is impaired [309, 312]. In contrast, human Pol ι and Pol κ are inhibited
237 by a cisplatin-d(GpG) adduct [356-358]. Dpo4 exhibited a different pattern during TLS in which the efficiency of dCTP insertion was decreased, particularly at the cisplatin- coordinated 5′-dG lesion and the subsequent extension step was only modestly reduced (Table 10.3). Notably, Dpo4 seemingly behaved as a hybrid of human Pol ι, Pol κ and Pol η: Dpo4 catalyzed synthesis was problematic at the lesion similar to Pol ι and Pol κ yet paused during the extension period like Pol η [309, 312, 356-358].
Mutagenic potential of a cisplatin-d(GpG) adduct Several groups have investigated the mutational spectra of cisplatin in eukaryotic cell- based assays as well as in vivo [378-382]. At cisplatin-d(GpG) cross-links, some highlights of the reported mutation spectra include: (i) single-base substitutions, insertions and deletions, (ii) G:C→A:T transitions and G:C→T:C transversions and (iii) most of the mutagenic events were located opposite the lesion [380-382]. These results suggest that the overall TLS process by human DNA polymerases is error-prone in the presence of cisplatin-damaged DNA, although, individual DNA polymerases (e.g. Pol η) may replicate in an error-free manner. For the model Y-family polymerase, the likelihood of misincorporation increased, as the fidelity of Dpo4 decreased up to two orders of magnitude when compared to the fidelity studies of normal DNA (Figure 10.5 part (B)) [105]. For both cisplatin cross-linked guanines, dATP was promiscuously inserted (Table 10.3). This preferential misinsertion of dATP over dTTP and dGTP may be due to a hydrogen bond between the C6 amino group of dATP and the C6 carbonyl oxygen of cisplatin-dG. This observation is depicted in the cisplatin-coordinated 5′-dG elongation structure of yPol η with dATP [290]. In regards to substrate specificity, Dpo4 selectively incorporated dCTP over dATP by 163- and 13-fold against the cisplatin-coordinated 3′- dG and 5′-dG, respectively (Table 10.3). Fidelity during the extension steps was relatively normal except for the ‘weak’ pause site at 26-mer/44DDP, whereby dATP was preferentially incorporated over the other incorrect dNTPs opposite template dC (Table 10.3 and Figure 10.5). Typically with an undamaged DNA substrate, Dpo4 discriminates between dGTP and dATP opposite template dC by 2100-fold [105], however, a discrimination factor of only 170-fold was achieved with 26-mer/44DDP (Table 10.3).
238 Interestingly, the 5′ template base is dT which may explain the preferential selection of dATP. In general, the promiscuity of Dpo4 appeared to be confined to the site of the lesion, as downstream events were essentially error-free (Table 10.3 and Figure 10.5 part (B)). More importantly, this result suggested that human Y-family DNA polymerases may generate mutations during cisplatin bypass, thereby promoting the risk of secondary malignancies observed in cisplatin-treated cancer patients.
The misinsertion frequency of Dpo4 on a cisplatin-d(GpG) substrate quantitatively suggested that the overall bypass process was potentially error-prone. Although the fidelity of a DNA polymerase may be less stringent at the cisplatin-d(GpG) lesion, such as what we have observed with Dpo4, other repair pathways may compensate by preferentially recognizing misincorporations opposite the cisplatin-d(GpG) cross-link. For example, the mismatch repair complex preferentially distinguishes a mismatch depending on the base pair opposite the lesion (dT > dG > dA) as well as the type of cisplatin–DNA adduct (cisplatin-d(GpG) > cisplatin-d(ApG) > cisplatin-d(GpNpG) [389, 390]). Thus, in the presence of ciplatin, the overall integrity of genomic DNA is a complex function of several different cellular pathways.
Kinetic comparison of double-base lesion bypass Using pre-steady state kinetics, the mechanism of correct nucleotide incorporation opposite a cis-syn thymine–thymine (TT) dimer catalyzed by yPol η is kinetically efficient, which is in contrast to the bypass mechanism of cisplatin-d(GpG) catalyzed by Dpo4 presented here [35]. Relative to undamaged DNA, the substrate specificity for correct dNTP insertion opposite cisplatin-d(GpG) is decreased 72- to 860-fold (i.e. inefficient) for Dpo4 versus 1.2- to 6-fold (i.e. efficient) for the TT lesion and yPol η (Table 10.3) [35]. However, binary complex formation was not significantly weakened for either enzyme despite the lesion-induced structural perturbations of the DNA helix (Table 10.2) [35, 383-385, 391]. Examination of the crystal structures containing the DNA lesion in complex with a polymerase and dNTP suggest hydrogen bonding is a
239 critical factor during TLS [101, 290]. Further kinetic and structural evidence is a requisite in order to fully understand the differences in the bypass efficiency and accuracy of a TT dimer and a cisplatin-d(GpG) adduct. Since yPol η is dramatically more efficient at bypassing a TT lesion, the productively bound ternary complex in Scheme 10.1 is likely dominant over the nonproductively bound ternary complex during TLS.
In summary, this study elucidated the kinetic mechanism of Dpo4 bypassing an important double-base lesion: a cisplatin-d(GpG) intrastrand cross-link. Although Dpo4 was capable of bypassing this double-base lesion at 37 °C, the TLS process was kinetically characterized as relatively inefficient and error-prone, especially opposite the cisplatin- coordinated 5′-dG. Using the knowledge gleaned from this study, improved platinum- based anticancer drugs may be rationally designed.
10.5 Future Directions
Dpo4 was used as a model Y-family DNA polymerase. However, in the past two years, it has become increasingly apparent that each DNA polymerase may employ a unique mechanism to bypass DNA lesions. Therefore, establishing kinetic mechanisms of cisplatin bypass for the human Y-family DNA enzymes (Pol η, Pol ι, Pol κ, and Rev1) is necessary. Another route of investigation is to examine the bypass of other cisplatin- DNA adducts. To complement the kinetic studies, it is important to determine the exact sequence of the lesion bypass product which can be determined using a short oligonucleotide sequencing assay [392]. This DNA sequence information is useful for discerning which human Y-family DNA polymerase is responsible for bypassing cisplatin-DNA adducts because the sequences obtained in vitro can be compared to those found in a mouse model [382].
240 10.6 Tables
Table 10.1 DNA sequences of primers and templates. Primers 19-mer 5’-GTCCCTGTTCGGGCGCCAG-3’ 21-mer 5’-GTCCCTGTTCGGGCGCCAGGA-3’ 22-mer 5’-GTCCCTGTTCGGGCGCCAGGAG-3’ 23-mer 5’-GTCCCTGTTCGGGCGCCAGGAGA-3’ 24-mer 5’-GTCCCTGTTCGGGCGCCAGGAGAC-3’ 25-mer 5’-GTCCCTGTTCGGGCGCCAGGAGACC-3’ 26-mer 5’-GTCCCTGTTCGGGCGCCAGGAGACCA-3’ 27-mer 5’-GTCCCTGTTCGGGCGCCAGGAGACCAG-3’ 28-mer 5’-GTCCCTGTTCGGGCGCCAGGAGACCAGA-3’ 29-mer 5’-GTCCCTGTTCGGGCGCCAGGAGACCAGAG-3’ 30-mer 5’-GTCCCTGTTCGGGCGCCAGGAGACCAGAGG-3’ 31-mer 5’-GTCCCTGTTCGGGCGCCAGGAGACCAGAGGC-3’ 32-mer 5’-GTCCCTGTTCGGGCGCCAGGAGACCAGAGGCT-3’ Templates 44DDPa 3’-CAGGGACAAGCCCGCGGTCCTCTGGTCTCCGATCAGAGCACTAG-5’ 44CTL 3’-CAGGGACAAGCCCGCGGTCCTCTGGTCTCCGATCAGAGCACTAG-5’ aThe cisplatin-modified guanines are in bold.
241
Table 10.2 Binding affinity of Dpo4 to control and damaged DNA substrates at 23 °C. DNA DNA Damaged Kd Control Kd Affinity substrate (nM) substrate (nM) ratioa 21-mer/44DDP 8 ± 1 21-mer/44CTL 18 ± 3 0.4-fold 22-mer/44DDP 21 ± 4 22-mer/44CTL 16 ± 3 1.3-fold 23-mer/44DDPb 56 ± 9 23-mer/44CTL 17 ± 2 3.3-fold 24-mer/44DDPb 26 ± 3 24-mer/44CTL 8 ± 1 3.3-fold 25-mer/44DDP 17 ± 4 25-mer/44CTL 5 ± 1 3.4-fold 26-mer/44DDP 12 ± 2 26-mer/44CTL 11 ± 2 1.1-fold 27-mer/44DDP 17 ± 3 27-mer/44CTL 13 ± 4 1.3-fold a DNA DNA Calculated as (Kd )damaged/(Kd )control. bDNA substrates at strong pause sites.
242
Table 10.3 Kinetic parameters of nucleotide incorporation into cisplatin-modified DNA. DNA a a kp Kd kp/Kd Efficiency c substrate dNTP -1 -1 -1 b Fidelity (P/T) (s ) (μM) (μM s ) ratio dGTP 6.0 ± 0.2 (2.0 ± 0.2) × 102 3.0 × 10-2 1.5 - 21-mer/ dATP 0.026 ± 0.002 (1.0 ± 0.1) × 103 2.7 × 10-5 0.97 9.1 × 10-4 44DDP dCTP 0.039 ± 0.007 (6 ± 3) × 102 6.7 × 10-5 2.6 2.3 × 10-3 dTTP 0.011 ± 0.001 (1.3 ± 0.2) × 103 8.6 × 10-6 1.4 2.9 × 10-4 dATP 3.8 ± 0.2 (3.9 ± 0.7) × 102 9.8 × 10-3 1.1 - 22-mer/ dCTP 0.0041 ± 0.0004 (1.4 ± 0.2) × 103 2.9 × 10-6 29 2.9 × 10-4 44DDP dGTP 0.0081 ± .0009 (1.1 ± 0.2) × 103 7.1 × 10-6 10 7.2 × 10-4 dTTP 0.0021 ± 0.0002 (7 ± 2) × 102 3.2 × 10-6 5.5 3.3 × 10-4 dCTP 1.6 ± 0.2 (1.3 ± 0.2) × 103 1.3 × 10-3 72 - 23-mer/ dATP 0.0056 ± 0.0005 (7 ± 1) × 102 8.0 × 10-6 3.4 6.3 × 10-3 44DDP dGTP 0.00050 ± 0.00008 (6 ± 2) × 102 9.1 × 10-7 67 7.2 × 10-4 dTTP 0.0017 ± 0.0003 (6 ± 2) × 102 2.8 × 10-6 21 2.2 × 10-3 dCTP 0.043 ± 0.004 (6 ± 1) × 102 7.3 × 10-5 860 - 24-mer/ dATP 0.00265 ± 0.00009 (4.6 ± 0.4) × 102 5.7 × 10-6 4.7 7.3 × 10-2 44DDP dGTP 0.000171 ± 0.000008 (8.0 ± 0.8) × 102 2.1 × 10-7 290 2.9 × 10-3 dTTP 0.00075 ± 0.00003 (6.4 ± 0.7) × 102 1.2 × 10-6 51 1.6 × 10-2 dATP 9.6 ± 0.4 (5.8 ± 0.6) × 102 1.6 × 10-2 6.1 - 25-mer/ dCTP 0.031 ± 0.004 (1.6 ± 0.3) × 103 1.9 × 10-5 4.4 1.2 × 10-3 44DDP dGTP 0.0115 ± 0.0009 (1.1 ± 0.2) × 103 1.0 × 10-5 6.8 6.3 × 10-4 dTTP 0.025 ± 0.003 (1.3 ± 0.2) × 103 1.9 × 10-5 0.92 1.2 × 10-3 dGTP 1.9 ± 0.1 (4.0 ± 0.7) × 102 4.7 × 10-3 36 - 26-mer/ dATP 0.04 ± 0.01 (1.5 ± 0.8) × 103 2.7 × 10-5 0.98 5.6 × 10-3 44DDP dCTP 0.007 ± 0.002 (1.4 ± 0.5) × 103 5.0 × 10-6 35 1.1 × 10-3 dTTP 0.0032 ± 0.0006 (1.1 ± 0.4) × 103 2.8 × 10-6 4.3 6.0 × 10-4 27-mer/ dATP 0.20 ± 0.02 (2.7 ± 0.7) × 102 7.5 × 10-4 17 - 44DDP dCTP 0.0016 ± 0.0003 (8 ± 3) × 102 2.1 × 10-3 40 2.8 × 10-3 28-mer/ dGTP 4.4 ± 0.3 (5.0 ± 0.9) × 102 8.9 × 10-3 2.8 - 44DDP dATP 0.0057 ± 0.0006 (1.5 ± 0.3) × 103 3.9 × 10-6 6.7 4.3 × 10-4 29-mer/ dGTP 0.72 ± 0.05 (5.0 ± 0.9) × 102 1.4 × 10-3 4.0 - 44DDP dATP 0.0016 ± 0.0001 (6 ± 1) × 102 2.8 × 10-6 9.4 1.9 × 10-3 30-mer/ dCTP 3.3 ± 0.2 (2.7 ± 0.4) × 102 1.2 × 10-2 7.1 - 44DDP dGTP 0.0022 ± .0002 (9 ± 2) × 102 2.5 × 10-6 25 2.0 × 10-4 31-mer/ dTTP 5.5 ± 0.3 (1.5 ± 0.3) × 102 3.7 × 10-2 2.1 - 44DDP dCTP 0.019 ± 0.003 (1.0 ± 0.3) × 103 1.8 × 10-5 0.69 4.9 × 10-4 32-mer/ dATP 18.9 ± 0.8 (1.3 ± 0.2) × 102 1.5 × 10-1 0.53 - 44DDP dCTP 0.032 ± 0.002 (8 ± 1) × 102 3.9 × 10-5 2.2 2.6 × 10-4
Continued 243 Table 10.3: Continued
a The kp and Kd obtained with 23-mer/44DDP and 24-mer/44DDP are apparent kinetic parameters due to biphasic kinetics of nucleotide incorporations at these pause sites. b Calculated as (kp/Kd)control/(kp/Kd)damaged in which the (kp/Kd)control is derived from Table 10.4 or reference [105] for correct and incorrect incorporations, respectively. c Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
244
Table 10.4 Kinetic parameters of correct nucleotide incorporation into the 44CTL DNA template.
DNA substrate kp Kd kp/Kd dNTP (P/T) (s-1) (μM) (μM-1s-1) 21-mer/44CTL dGTP 9.3 ± 0.2 (2.1 ± 0.2) × 102 4.5 × 10-2 22-mer/44CTL dATP 4.1 ± 0.1 (3.8 ± 0.4) × 102 1.1 × 10-2 23-mer/44CTL dCTP 10.8 ± 0.4 (1.2 ± 0.1) × 102 9.3 × 10-2 24-mer/44CTL dCTP 4.5 ± 0.2 (7 ± 2) × 101 6.3 × 10-2 25-mer/44CTL dATP 15.4 ± 0.8 (1.6 ± 0.3) × 102 9.7 × 10-2 26-mer/44CTL dGTP 9.3 ± 0.2 (5.5 ± 0.4) × 101 1.7 × 10-1 27-mer/44CTL dATP 5.8 ± 0.2 (4.0 ± 0.5) × 102 1.3 × 10-2 28-mer/44CTL dGTP 15.0 ± 0.8 (6.1 ± 0.8) × 102 2.5 × 10-2 29-mer/44CTL dGTP 5.2 ± 0.5 (9 ± 2) × 102 5.6 × 10-3 30-mer/44CTL dCTP 14.5 ± 0.9 (1.7 ± 0.3) × 102 8.5 × 10-2 31-mer/44CTL dTTP 15.7 ± 0.9 (2.0 ± 0.4) × 102 7.7 × 10-2 32-mer/44CTL dATP 21 ± 1 (2.6 ± 0.4) × 102 8.0 × 10-2
245
Table 10.5 Biphasic kinetic parameters of dCTP incorporation into cisplatin-modified DNA. DNA k A k A substrate 1 1 2 2 -1 (nM) -1 (nM) (P/T) (s ) (s ) 23-mer/44DDP 10 ± 2 2.1 ± 0.1 (7%) 0.0009 ± 0.0001 19 ± 1 (63%) 24-mer/44DDP 4.0 ± 0.7 1.11 ± 0.01 (3.7%) 0.00016 ± 0.00003 3.2 ± 0.4 (11%)
246 10.7 Figures
Figure 10.1 Running start nucleotide incorporation assay. A pre-equilibrated solution of Dpo4 (100 nM) and 5′-[32P]-labeled DNA (100 nM) were mixed with all four dNTPs (200 µM each) for various reaction times before terminating with EDTA. Lengths of the resolved products and the site of the cisplatin adduct (*) are designated along the right margin. (A) reaction with 19-mer/44CTL substrate; (B) reaction with 19-mer/44DDP substrate.
247
Figure 10.2 Purity of template 44DDP. To examine the purity of DNA templates 44CTL and 44DDP (Table 10.1), the oligonucleotides were 5′-[32P]-labeled, analyzed using PAGE (17% acrylamide, 8 M urea, 1× TBE), and visualized via autoradiography. The cisplatin modification induces a shift of approximately one nucleotide. A single band was detected for both DNA templates, thereby confirming the apparent homogeneity of each oligonucleotide.
248 A
B
100
80
60
40 24-mer/44CP (nM) z 20 Dpo4 0 0 100 200 300 400 500 600 Dpo4 (nM)
Figure 10.3 Measurement of Dpo4 binding to 24-mer/44DDP. (A) Reactions containing 5′-[32P]-labeled 24-mer/44DDP (100 nM) were incubated with increasing concentrations of Dpo4 (25-425 nM). The binary complex was separated from the unbound DNA substrate via native polyacrylamide gel electrophoresis. (B) A plot of binary complex formation (Dpo4•24-mer/44DDP) versus Dpo4 concentration was fit to a quadratic DNA equation (Equation 1) which yielded a Kd of 26 ± 3 nM.
249 A
30
25
20
15
10 Product (nM)
5
0 0 100 200 300 400 500 600 700 800
Time (s)
B 0.035
0.03
0.025 )
-1 0.02 (s
0.015 obs k 0.01
0.005
0 0 500 1000 1500 2000 dCTP (μM)
Figure 10.4 Concentration dependence on the pre-steady state rate constant of dCTP incorporation. A pre-incubated solution of Dpo4 (120 nM) and 5′-[32P]-labeled 24- mer/44DDP (30 nM) was rapidly mixed with increasing concentrations of dCTP•Mg2+ (100 μM, U; 200 μM, S; 400 μM, ; 700 μM, ; 1300 μM, {; 1600 μM, z) for various time intervals. (A) The solid lines represent the best fits to a single-exponential equation (Equation 2) which determined the observed rate constants, kobs. (B) The kobs values were plotted as a function of dCTP concentration and fit to a hyperbolic equation -1 (Equation 3) whereby a kp of 0.043 ± 0.004 s and a Kd of 588 ± 145 μM were obtained.
250 A
B
Figure 10.5 Quantitative effect of the cisplatin-DNA adduct on nucleotide incorporation and fidelity. (A) A plot of the efficiency ratio (extracted from Table 10.3) for each correct incorporation into DNA substrates with varying primer lengths is shown. (B) The fidelity of all misincorporations into undamaged [bottom most row, from reference [105]] and damaged (upper rows) DNA by Dpo4 are shown. The two dashed vertical lines represent the outermost limits of “normal” misincorporations by Dpo4. Each shape designates a specific dNTP: dATP, z; dCTP, S; dGTP, ; dTTP, .
251
Figure 10.6 Effectiveness of the D-1 DNA trap. Dpo4 (120 nM) was added to a solution of 5′-[32P]-labeled 23-mer/44DDP (30 nM) and 21-mer/41CTL D-1 DNA (5 µM). The reaction was initiated with dCTP (1.0 mM) and terminated with 0.37 M EDTA after various reaction times. Products were resolved using PAGE (17% acrylamide, 8 M urea, 1× TBE). The autoradiographed gel image revealed minimal product formation (24-mer) after 30 min. Thus, a molar ratio of 167:1 for the D-1 DNA trap to the radiolabeled damaged DNA substrate was effective at sequestering Dpo4 and was used in all biphasic kinetic assays.
252 20
15
10
Product (nM) 5
0 0 500 1000 1500 2000 Time (s)
3
2.5 Inset
2
1.5
1 Product (nM) Product
0.5
0 0 5 10 15
Time (s)
Figure 10.7 Biphasic kinetics observed in the presence of a DNA trap when Dpo4 incorporated dCTP opposite the cisplatin lesion. A pre-incubated solution of Dpo4 (120 nM) and 5′-[32P]-labeled DNA (30 nM) was rapidly mixed with 1.2 mM dCTP•Mg2+ in the presence of a DNA trap (5 µM) for various time intervals. Applying a double- exponential equation (Equation 4) to a plot of product concentration versus time resolved the fast and slow phase amplitudes and rate constants for 23-mer/44DDP () and 24- mer/44DDP (z). For 23-mer/44DDP, the following kinetic parameters were extracted: -1 -1 A1 = 2.1 ± 0.1 nM (7%), k1 = 10 ± 2 s , A2 = 19 ± 1 nM (63%), k2 = 0.0009 ± 0.0001 s . For 24-mer/44DDP, the following kinetic parameters were resolved: A1 = 1.11 ± 0.01 nM -1 -1 (3.7%), k1 = 4.0 ± 0.7 s , A2 = 3.2 ± 0.4 nM (11%), k2 = 0.00016 ± 0.00003 s . The inset shows product formation within 15 s which corresponds to the fast phase.
253 10.8 Schemes
Scheme 10.1
254
Scheme 10.2
255 Chapter 11: Pre-Steady State Kinetic Analysis of the Incorporation of Anti-HIV Nucleotide Analogs Catalyzed by Human X- and Y-family DNA Polymerases
11.1 Introduction
More than 30 million people worldwide are infected with the human immunodeficiency virus (HIV) which is the causative agent of acquired immunodeficiency syndrome (AIDS). To manage the life-threatening effects of HIV replication, nucleoside reverse transcriptase inhibitors (NRTIs) have been a mainstay in effective combination antiretroviral therapy. NRTIs undergo phosphorylation by host cell kinases so that they are converted into their active di- or triphosphate (DP or TP) forms which can serve as nucleotide substrates for HIV reverse transcriptase (RT). Incorporation of the NRTIs into the viral genome by HIV RT terminates the replication process due to the lack of a 3′- hydroxyl in these drugs. However, this mechanism of drug action does not exclusively target the viral enzyme, HIV RT. Host DNA polymerases (pols), organized into the A, B, X, or Y families, are susceptible to drug inhibition because these enzymes catalyze a similar nucleotidyl transfer reaction as HIV RT. If human DNA polymerases incorporate the drug analog into human DNA, then this event will inhibit DNA replication and possibly lead to cell death and drug toxicity. For example, a correlation has been established between the kinetics of nucleotide analog incorporation catalyzed by human DNA polymerase γ (Pol γ), an A-family member, and the observed clinical toxicity that presents as mitochondrial dysfunction [393-396]. However, this molecular basis of toxicity does not account for all of the unwanted side effects [397]. For example, zidovudine has been associated with bone marrow toxicity [398, 399] while tenofovir [400-403] can cause kidney toxicity likely via Pol γ-independent mechanisms [397, 404]. Another possible mechanism is analog incorporation into nuclear DNA [398, 405, 406],
256 since NRTIs induce genomic instability by increasing mutations, structural chromosomal aberrations, abnormal chromatin structure, sister chromatid exchanges, and shorter telomeres [397, 407, 408]. Candidates responsible for NRTI insertion into nuclear DNA include three replicative, B-family DNA polymerases (Pols α, δ, and ε) and the important base excision repair enzyme, Pol β (a X-family member). Although human replicative DNA polymerases perform the majority of nuclear DNA synthesis, most of these pols have stringent nucleotide selection mechanisms, therefore, they usually exhibit weak inhibition with antiretroviral nucleotide analogs that may not be sufficient to result in cytotoxicity [409, 410].
In the past 15 years, ten novel human DNA polymerases such as Pol λ [74, 270], Pol η [36], Pol ι [411], Pol κ [412], and Rev1 [413] have been identified. Thus, the discovery of these new human enzymes was after the approval of some NRTIs by the United States Food and Drug Administration (FDA). Pol λ is an X-family member while Pol η, Pol ι, Pol κ, and Rev1 belong to the Y-family of DNA polymerases [6]. The Y-family DNA polymerases are proposed to function in DNA lesion bypass, somatic hypermutation, base excision repair, nucleotide excision repair, and recombination pathways [39, 40, 80]. For example, human Pol η is responsible for the error-free bypass of cis-syn pyrimidine- pyrimidine dimers in vivo [36]. In vitro, Pol ι has been shown to incorporate incorrect dGTP opposite a templating base thymine more efficiently than canonical dATP, Pol κ can efficiently elongate mispaired primer termini, and Rev1 functions as a dCTP transferase by using hydrogen bonds with the side chain of Arg357, rather than a template base, to direct dCTP incorporation [414]. Pol λ, which shares 34% sequence identity with Pol β [74], has putative roles in base excision repair, non-homologous end joining, and V(D)J recombination [30]. Incorporation of a chain terminator during these biological processes would result in DNA replication inhibition and single- or double- strand DNA breaks which can lead to apoptosis, immunosuppression, and genetic diseases [397]. These specialized human X- and Y-family DNA polymerases exhibit low 0 -5 fidelity (10 to 10 ) on undamaged DNA [66], lack 3′ → 5′ exonuclease activity, and possess more flexible active sites which can accommodate altered DNA and nucleotide
257 structures [67]. Overall, the X- and Y-family DNA polymerases are up to 100,000 times more error prone than Pol γ synthesizing undamaged DNA [66, 90, 91].
Due to the relatively recent discovery of these novel DNA polymerases, little is known whether these non-canonical human pols can incorporate the active di- or triphosphate forms of the NRTIs. In vitro kinetic studies are a good starting point because the human genome encodes 16 DNA pols with overlapping functions; all of these nucleotidyl transferases may potentially be unwanted targets. Currently, cell-based assays and animal models cannot unambiguously identify which DNA polymerases are responsible for inducing cellular toxicity. Except for Pols β and η, the physiological roles and regulation of these specialized X- and Y-family pols have not been confirmed under normal conditions. Moreover, the expression patterns and activity of these X- and Y-family enzymes may be altered in virus-infected and NRTI-treated cells. For example, the expression levels of these non-canonical DNA polymerases in stressed human cells are known to be raised significantly. Thus, the goal of this study was to understand the kinetic basis of incorporation for four widely-prescribed anti-HIV nucleoside analogs [emtricitabine (L-FTC), lamivudine (L-3TC), tenofovir (PMPA), and zidovudine (AZT)] in their active forms by Pol β, Pol λ, Pol η, Pol ι, Pol κ, and Rev1 using pre-steady state kinetic methods (Figure 11.1). This study established a structure-function relationship which may be useful in designing more effective and less toxic NRTIs.
11.2 Materials and Methods
Materials These chemicals were purchased from the following companies: [γ-32P]ATP, MP Biomedicals; deoxyribonucleotide 5′-triphosphates, GE Healthcare; Bio-Spin 6 columns, Bio-Rad Laboratories; OptiKinase™, USB Corporation; DNA oligomers in Table 11.1, Integrated DNA Technologies; 3′-azido-3′-deoxythymidine 5′-triphosphate (AZT-TP), TriLink BioTechnologies. Diphosphate of 9-[2-(phosphonomethoxy)propyl]adenine
258 (PMPA-DP), L-2′,3′-dideoxy-5-fluoro-3′-thiacytidine 5′-triphosphate (L-FTC-TP), and L- 2′,3′-dideoxy-3′-thiacytidine 5′-triphosphate (L-3TC-TP) were kind gifts from Gilead Sciences, Inc.
Preparation of human DNA polymerases and DNA substrates The plasmids, expression, and purification of human DNA polymerases β [235, 274], λ [117], η [392], truncated ι (1-420) [392], truncated κ (9-518) [392], and truncated Rev1 (341-829) [330, 340] were described previously. Commercially synthesized oligomers in Table 11.1 were purified using polyacrylamide gel electrophoresis [105, 182](41,42). The 21-mer primer was radiolabeled with [γ-32P]ATP and OptiKinase™ according to the manufacturer’s protocol, and the unreacted [γ-32P]ATP was subsequently removed via a Bio-Spin 6 column. The 21-41mer primer-template DNA substrates [105] and 21-19- 41mer single-nucleotide gap DNA substrates [182] were annealed as described previously.
Measurement of the kp and Kd for single-nucleotide incorporation Kinetic assays were completed using buffer B (50 mM Tris-HCl pH 7.8 at 37 °C, 5 mM
MgCl2, 50 mM NaCl, 0.1 mM EDTA, 5 mM DTT, 10% glycerol, and 0.1 mg/ml of bovine serum albumin (BSA)) for Pol β, buffer L (50 mM Tris-HCl pH 8.4 at 37 °C, 5
mM MgCl2, 100 mM NaCl, 0.1 mM EDTA, 5 mM DTT, 10% glycerol, and 0.1 mg/ml of
BSA) for Pol λ [182], and buffer Y (50 mM HEPES pH 7.5 at 37 °C, 5 mM MgCl2, 50 mM NaCl, 0.1 mM EDTA, 5 mM DTT, 10% glycerol, and 0.1 mg/ml of BSA) for Pols η, ι, κ, and Rev1. Pols η, ι, κ, and Rev1 were assayed with the 21-41mer DNA substrates, and Pols β and λ were assayed with the 21-19-41mer single-nucleotide gap DNA substrates. All kinetic experiments described herein were performed at 37 °C, and the reported concentrations were final after mixing all the components. Please note, AZT-TP was not pre-incubated with DTT to avoid reduction of the azido group [415]. A pre- incubated solution of the DNA polymerase (120 or 300 nM) and 5′-[32P]-radiolabeled DNA substrate (30 nM) was mixed with increasing concentrations (0.025-2,000 μM) of nucleotide or nucleotide analog in the appropriate buffer at 37 °C. The polymerase was in
259 molar excess over DNA, whereby the enzyme to DNA ratio was 4:1 for Pols λ, η, ι, and Rev1 and 10:1 for Pols β and κ. Aliquots of the reaction mixtures were quenched at various times using 0.37 M EDTA. A rapid chemical-quench flow apparatus (KinTek) was utilized for fast nucleotide incorporations. Reaction products were resolved using sequencing gel electrophoresis (17% acrylamide, 8 M urea) and quantitated with a Typhoon TRIO (GE Healthcare). The time course of product formation at each nucleotide concentration was fit to a single-exponential equation (Equation 1)
[Product] = A[1 – exp (–kobst)] (1) using a nonlinear regression program, KaleidaGraph (Synergy Software), to yield an observed rate constant of nucleotide incorporation (kobs). The kobs values were then plotted as a function of nucleotide concentration and fit using the hyperbolic equation (Equation 2)
kobs = kp[dNTP]/(Kd + [dNTP]) (2)
which resolved the maximum rate of nucleotide incorporation (kp) and equilibrium
dissociation constant (Kd) for nucleotide incorporation catalyzed by each enzyme.
11.3 Results
Measurement of selection factors Single-turnover kinetic assays were used to determine the kinetic basis of how Pols β, λ, η, ι, κ, and Rev1 discriminate between a correct natural nucleotide versus a nucleotide analog based on the equilibrium dissociation constant (Kd) and the maximum rate of
incorporation (kp) for an incoming nucleotide. To directly observe the conversion of the DNA substrate into the extended DNA product during a single pass through the enzymatic pathway, a pre-incubated solution of Pol η and 5′-[32P]-labeled 21-41merGT DNA (Table 11.1) was mixed with increasing concentrations of L-3TC-TP (0.2-25 μM) -1 (Section 11.2) [63]. The data were analyzed to resolve a kp of 0.0296 ± 0.0009 s and a Kd of 3.3 ± 0.3 μM (Figure 11.2). Similar single-nucleotide incorporation assays were performed for each enzyme incorporating dATP, PMPA-DP, dCTP, L-3TC-TP, L-FTC-
260 TP, dTTP, or AZT-TP opposite the complementary template base. Please note, a single- nucleotide gap DNA substrate was used in the assays for the gap-filling Pols β and λ, since the catalytic efficiency of these enzymes is enhanced by a 5′-phosphorylated downstream strand [114, 192] and the gap DNA substrate models a more physiologically- relevant DNA substrate. Meanwhile, Pols η, ι, κ, and Rev1 were assayed with a primer-template DNA substrate. The kinetic parameters are listed in Tables 11.2-11.4
where the incorporation efficiency (kp/Kd) and selection factors are defined and calculated. All polymerases prefer the natural dNTP more than the nucleotide analog.
Selection factors for L-cytosine-based analogs Lamivudine (L-3TC) is a first generation NRTI that was approved in 1995, and it has two unique features: L-steroechemistry and an oxathiolane ring (Figure 11.1). Despite lacking the natural D-stereochemistry, L-3TC-TP was incorporated by the six pols examined in this work with selection factors ranging from 40 to 3,300 (Table 11.2). In general, the mechanism of L-3TC-TP incorporation was similar for most of the pols: a slower rate of incorporation (~6,100-fold on average) but tighter nucleotide binding affinity (1/Kd, ~6- fold on average) than dCTP. In contrast, a slightly weaker ground-state binding affinity for L-3TC-TP to the Pol ι•DNA complex was measured. Emtricitabine (L-FTC), a second generation NRTI approved in 2003, consists of L-3TC with a fluorine atom added to the C5 position (Figure 11.1). This subtle modification resulted in selection factors (120 to 7,900) for L-FTC-TP that were equal to or greater than those determined for L-3TC-TP (Table 11.2). The greatest effect was observed with Pols β and ι, whereby the selection factor for L-FTC-TP increased 9- and 4-fold, respectively, relative to L-3TC-TP. This
kinetic effect was due to larger Kd values. In general, the C5 fluoro group of L-FTC-TP
did not affect the rate of incorporation, since the kp values for L-FTC-TP and L-3TC-TP remained similar. Taken together, these results showed that the stereospecificity of an incoming dNTP is important for Pols β, λ, η, ι, κ, and Rev1 during the nucleotide incorporation step.
261 Selection factors for an acyclic nucleotide analog Next, we examined tenofovir (PMPA), one of the second generation NRTIs, which possesses several novel structural characteristics: an acyclic moiety and a phosphonate group that is ~1 Å closer to the base than the phosphate of dAMP (Figure 11.1) [416]. The kinetic parameters for Pol ι and Rev1 inserting PMPA-DP could not be determined, since product formation was barely detectable after 3 h at a relatively high PMPA-DP concentration of 500 μM (data not shown). However, the selection factors of Pols β, λ, η, and κ were calculated to be 40, 65, 1,800, and 110,000, respectively (Table 11.3). For
Pols β and λ, the Kd was weakened by 5-fold on average and the kp was reduced by 11- fold on average compared to dATP. Notably, Pols β and λ have different kinetic parameters which support our previous discovery that these two homologs of the X- family possess different enzymatic properties [114, 117]. In contrast, Pol η maintained a tight enzyme•DNA•PMPA-DP complex, but the rate of PMPA-DP incorporation opposite template dT dropped by 2,600-fold. Pol κ discriminated PMPA-DP from dATP
by decreasing the rate by 250-fold and increasing the Kd value by 430-fold. Thus, the Y- family enzymes use the ribose moiety as a mechanism of discrimination more than the X- family enzymes. Moreover, most of the PMPA-DP selection factors were higher than those determined for L-3TC-TP and L-FTCTP.
Selection factors for AZT-TP The kinetic parameters were measured and the selection factors were calculated for the 5′-triphosphate form of zidovudine, the first NRTI approved by the FDA, being incorporated by the six human pols (Table 11.4). AZT has a 3′-azido group that would be larger in size than the 3′-hydroxyl (Figure 11.1). The selection factors for AZT-TP ranged from 110 to 2,600, and the mechanism of incorporation was altered mostly at the level of incorporation (270-fold reduction on average) and modestly at the ground-state binding step (11-fold weaker on average) compared to dTTP. Although Rev1 exhibits the lowest discrimination factor for AZT-TP, the incorporation efficiency is the lowest among the -6 -1 -1 pols at 5.9 × 10 μM s .
262 Different mechanisms observed for different NRTIs As noted above, different mechanisms of analog incorporation were determined for the four NRTIs, indicating the altered chemical structures differentially affect each polymerase activity as the preferential order of incorporation varies among the DNA polymerases. Furthermore, it is important to note that the order based on selection factors does not coincide with the order based on substrate specificity constants except for Pol η and Pol κ (i.e. L-FTC-TP ≈ L-3TC-TP > AZTTP > PMPA-DP for Pol η and AZT-TP > L-3TC-TP > L-FTC-TP > PMPA-DP for Pol κ). Analyzing these orders reveals the following general trends: (i) L-3TC-TP or AZT-TP are usually the most preferred NRTIs, (ii) the fluoro group on L-FTC-TP leads to a higher level of discrimination compared to L-3TC-TP, and (iii) the Y-family enzymes prefer PMPA-DP the least.
11.4 Discussion
Kinetic basis of NRTI selection and inhibition among viral and human DNA polymerases Using transient state kinetic techniques, this work determined the incorporation efficiency values for the active forms of four FDA-approved NRTIs catalyzed by six non- canonical human DNA polymerases: β, λ, η, κ, ι, and Rev1. Most of the NRTIs were inserted into DNA by these DNA polymerases, suggesting that incorporation into nuclear DNA and interference of genomic replication and repair are possible in vivo. The four NRTIs examined herein are associated with a low potential of producing mitochondrial dysfunction [417, 418], thereby suggesting toxicity associated with these NRTIs is induced by other mechanisms [397]. The selection factors for an exonuclease-deficient mutant of human mitochondrial DNA polymerase γ, an A-family member, have been measured or estimated using pre-steady state kinetic techniques [393, 394, 396, 419]. Based on the selection factors, Pol γ exhibited a greater level of discrimination than most of the X- and Y-family DNA polymerases: up to 70-, 2,400-, 270-, and 90,000-fold for L- 3TC-TP, L-FTC-TP, PMPA-DP, and AZT-TP, respectively (Tables 11.2-11.4). This kinetic finding strengthens the possible role of human X- and Y-family DNA
263 polymerases in the etiology of NRTI toxicity. The kinetics of Pol γ-catalyzed AZT-TP incorporation is complicated by the slow release of the pyrophosphate product, therefore, the lack of a hyperbolic-concentration dependence on the rate of incorporation prevented -7 the measurement of single-turnover kinetic parameters, but a selection factor of 1 × 10 was calculated [419]. The unique mechanism of AZT-TP selection by Pol γ does not apply to the non-canonical polymerases examined in this work, since their observed rate constants were dependent on the concentration of AZT-TP. For the unnatural L-cytosine analogs, Pol γ exhibits a weaker ground-state binding affinity which is in contrast to Pols β, λ, η, κ, and Rev1. Perhaps the more spacious active site of low fidelity enzymes provides greater accessibility for the L-stereoisomer to enter and the oxathiolane ring to
bind favorably, leading to a tighter Kd value. Similar to the Y-family enzymes, Pol γ depends on the presence of a ribose moiety for efficient nucleotide incorporation, since
PMPA-DP reduced the kp and weakened the Kd. Similar kinetic results were obtained for PMPA-DP incorporation catalyzed by an exonuclease-deficient mutant of T7 DNA polymerase, a model A-family enzyme [416]. Unlike Pol γ, the X- and Y-family enzymes lack 3′ → 5′ exonuclease activity, therefore, the incorporated NRTIs can only be removed by pyrophosphorolysis which requires high cellular pyrophosphate levels due to the weak binding affinity of pyrophosphate, meaning NRTIs will not be excised easily from DNA.
The efficacy of NRTIs depends partially on the selection factor of HIV-1 RT relative to the host DNA polymerases. In general, the selection factors for HIV-1 RT during DNA- dependent DNA synthesis are lower by at least 1-, 8-, 7-, and 22-fold for L-3TC-TP, L- FTC-TP, PMPA-DP, and AZT-TP, respectively, than those measured for human X- and Y-family pols (Tables 11.2-11.4). Improving the efficacy of NRTIs may be a challenge because the kinetic basis of analog incorporation for HIV-1 RT is sometimes similar to the X- and Y-family DNA polymerases. For example, HIV-1 RT incorporates the L- oxathiolane cytosine analogs with a ~4-fold tighter binding affinity and ~100-fold reduced rate of incorporation, a general trend that is similar to the X- and Y-family polymerases but different than Pol γ (Table 11.2) [393, 396, 420]. In contrast, HIV-1 RT prefers L-FTC-TP over L-3TC-TP whereas Pols β, λ, κ, ι, and Rev1 prefer L-3TC-TP.
264 These results may explain why emtricitabine is a more effective NRTI than lamivudine
[421]. The smaller and more flexible acyclic moiety of PMPA-DP only affects the Kd during catalysis by HIV-1 RT unlike the human enzymes. A crystal structure of the HIV- 1 RT•DNA•PMPA-DP ternary complex revealed that PMPA-DP forms a non-canonical Watson-Crick base pair [422]. Thus, this sub-optimal base pairing scheme for PMPA- DP:dT may contribute to the lower catalytic efficiency for the human enzymes, since hydrogen bonding has been shown to be important for incorporation catalyzed by Pol γ [205], Pol λ [423], yeast Pol η [315], Pol κ [424], and Rev1 [190]. Overall, PMPA-DP appears to be a superior drug analog because it requires only two phosphorylation events to become activated, it is a better substrate for HIV-1 RT than the seven human enzymes examined thus far (Table 11.3) [394, 416], and it has a favorable resistance profile characterized by activity against most NRTI-resistant viruses and a low propensity for resistance development [425, 426].
Potential cellular implications of NRTIs incorporated into human genomic DNA Although the X- and Y-family DNA polymerases perform fewer incorporation events than Pol γ, most of them show a lesser degree of discrimination for the nucleotide analogs versus natural dNTPs compared to Pol γ (Tables 11.2-11.4). The potential incorporation of nucleotide analogs in vivo depends on intracellular concentrations of natural nucleotides relative to the nucleotide analogs. The ratio of dCTP:L-3TC-TP is approximately 10:1 in non-infected phytohemagglutinin (PHA)-activated peripheral blood mononuclear cells (PBMC) [427]. Gao et al. have measured the dTTP:AZT-TP ratio to be 21:1 and 1.6:1 for resting and PHA-activated PBMC, respectively [428]. Unlike the aforementioned works, most studies do not measure the levels of both the natural dNTPs and the nucleotide analogs. Therefore, the dATP:PMPA-DP and dCTP:L- FTC-TP ratios are predicted by us to be 10:1 and 1:1, respectively, using the values for natural dATP or dCTP (Table 2 in reference [428]) and the average values for PMPA-DP [429, 430] and L-FTC-TP [431, 432], all of which were derived from PBMC. These predicted or measured dNTP:dNTP analog ratios suggest that the relative intracellular concentrations can approach 1:1, therefore, the selection factors (Tables 2-4) can be
265 interpreted as the insertion frequency. For example, the selection factors of AZT-TP range from 110 to 2,600 (Table 4). Thus, the six non-canonical human DNA polymerases are predicted to incorporate one AZT-TP molecule for every 110 to 2,600 dTTP incorporations when the cellular concentrations of AZT-TP and dTTP are equal.
Pols β [433], λ [200, 270], η [434], κ [412, 435], ι [436], and Rev1 [70, 330] are expressed at the mRNA level in various human tissues, including the brain, heart, lung, liver, pancreas, kidney, spleen, peripheral blood leukocyte, ovary, testis, placenta, and skeletal muscle. These DNA polymerases participate in critical cellular pathways that maintain genomic integrity and enhance cell survival, such as base excision repair, non- homologous end joining, and translesion DNA synthesis [30, 39, 40, 80, 437]. In addition, Pols λ, η, ι, and Rev1 have been implicated in antibody generation processes, including V(D)J recombination and somatic hypermutation [80, 437]. Some of these pathways involve single-strand and double-strand DNA breaks, therefore, incorporation of a chain terminator would inhibit the subsequent DNA ligation step due to the lack of a 3′-hydroxyl. Such DNA breaks may lead to genomic instability [407, 408, 438] and eventually trigger apoptosis in non-infected cells that could possibly lead to unwanted side effects [397]. The side effects associated with NRTIs include the following: nausea, anorexia, headache, fatigue, dizziness, diarrhea, myopathy, cardiomyopathy, liver failure, hepatic steatosis, pancreatitis, lactic acidosis, lipodystrophy, peripheral neuropathy, and bone marrow toxicity [438, 439]. Specifically, tenofovir and zidovudine are known for possibly inducing kidney toxicity [400-403] and bone marrow suppression [398, 399, 439], respectively, while lamivudine and emtricitabine have few adverse effects [439, 440]. The X- and Y-family DNA polymerases are expressed in the tissues afflicted with toxicity so it is plausible that NRTI incorporation by these enzymes may play a role. Furthermore, NRTI treatment has been associated with genotoxicity, and these effects are well documented for zidovudine [408, 438] in which one postulated route of toxicity involves AZT-TP incorporation into nuclear DNA [398, 406]. Since these selected X- and Y-family DNA polymerases can use AZT-TP as a substrate (Table 11.4), these enzymes are potential candidates for causing this observed genotoxicity, for AZT causes
266 cells to accumulate in S phase [441] and induces apoptosis via double-strand DNA breaks [442]. Besides NRTI incorporation by host enzymes, other processes can affect the efficacy and toxicity of the analog, such as drug uptake, transport, catabolism, and metabolism. For example, NRTIs affect nucleotide concentrations which would reduce polymerase selectivity and fidelity [408, 438]. This interplay of pathways is likely important for a comprehensive understanding of NRTI toxicity.
Concluding remarks Although our in vitro kinetic findings do not provide direct evidence that X- and Y- family DNA polymerases are a causative factor of in vivo toxicity, our data did establish the following important points: (i) most antiviral nucleotide analogs are substrates for human X- and Y-family DNA polymerases, (ii) these non-canonical pols are less selective than Pol γ, (iii) the basis of nucleotide analog selection is sometimes similar for the X- and Y-family pols and HIV-1 RT, and (iv) NRTI toxicity may involve nucleotide analog incorporation by DNA repair and lesion bypass DNA polymerases.
11.5 Future Directions
This study investigated nucleotide analog incorporation opposite undamaged DNA. However, a more physiologically-relevant DNA substrate for lesion bypass DNA polymerases is damaged DNA. Three common forms of DNA damage are 8-oxo-7,8- dihydro-dG, pyrimidine-pyrimidine dimers, and abasic sites. Therefore, it would be insightful to understand the selection factors for the drugs being incorporated opposite these and other forms of DNA damage.
267 11.6 Tables
Table 11.1 Sequences of oligonucleotidesa. 21mer 5’-CGCAGCCGTCCAACCAACTCA-3’ 19merC 5’-CGTCGATCCAATGCCGTCC-3’ 19merA 5’-AGTCGATCCAATGCCGTCC-3’ 41merGT 3’-GCGTCGGCAGGTTGGTTGAGTGTCAGCTAGGTTACGGCAGG-5’ 41merTG 3’-GCGTCGGCAGGTTGGTTGAGTTGCAGCTAGGTTACGGCAGG-5’ 41merAG 3’-GCGTCGGCAGGTTGGTTGAGTAGCAGCTAGGTTACGGCAGG-5’ aThe 21mer strand was 5′-radiolabeled. For single-nucleotide gap DNA substrates, the downstream 19mer strand was 5′-phosphorylated. The identity of important base positions is highlighted in bold.
268
Table 11.2 Kinetic parameters for nucleotide incorporation opposite template base dG at 37 °C. DNA k K k /K Selection dNTP p d p d polymerase (s-1) (μM) (μM-1s-1) factora Pol β dCTP 5.02 ± 0.07 0.71 ± 0.04 7.1 L-3TC-TP 0.00390 ± 0.18 ± 0.02 2.2 × 10-2 325 0.00010 L-FTC-TP 0.027 ± 0.001 11 ± 2 2.5 × 10-3 2,900 Pol λ dCTPb 1.57 ± 0.04 0.9 ± 0.1 1.7 L-3TC-TP 0.00402 ± 0.106 ± 3.8 × 10-2 46 0.00008 0.010 L-FTC-TP 0.0049 ± 0.0001 0.36 ± 0.03 1.4 × 10-2 120 Pol η dCTP 49 ± 2 25 ± 4 2.0 L-3TC-TP 0.0296 ± 0.0009 3.3 ± 0.3 9.0 × 10-3 220 L-FTC-TP 0.0244 ± 0.0004 2.7 ± 0.1 9.0 × 10-3 220 Pol κ dCTP 11.8 ± 0.6 32 ± 5 3.7 × 10-1 L-3TC-TP 0.00045 ± 4.0 ± 0.5 1.1 × 10-4 3,300 0.00001 L-FTC-TP 0.000225 ± 4.8 ± 0.2 4.7 × 10-5 7,900 0.000003 Pol ι dCTP 0.075 ± 0.002 50 ± 5 1.5 × 10-3 L-3TC-TP 0.00356 ± 96 ± 8 3.7 × 10-5 40 0.00009 L-FTC-TP 0.0020 ± 0.0001 230 ± 20 8.7 × 10-6 170 Rev1 dCTPc 22.4 ± 0.9 2.2 ± 0.3 10 L-3TC-TP 0.0236 ± 0.0006 1.04 ± 0.10 2.3 × 10-2 450 L-FTC-TP 0.0222 ± 0.0007 2.7 ± 0.3 8.2 × 10-3 1,200 Pol γ dCTPd 44 ± 2 1.1 ± 0.1 40 L-3TC-TPd 0.125 ± 0.005 9.2 ± 0.9 1.4 × 10-2 2,900 L-FTC-TPe 0.0086 ± 0.0015 62.9 ± 8.4 1.4 × 10-4 290,000 HIV-1 RT dCTPf 2.9 ± 0.2 56 ± 10 5.2 × 10-2 L-3TC-TPg 0.019 ± 0.001 15 ± 3 1.3 × 10-3 41 L-FTC-TPe 0.039 ± 0.003 12 ± 3 3.3 × 10-3 16 a Calculated as (kp/Kd)dCTP/(kp/Kd)Analog. bKinetic parameters are from reference [117]. cKinetic parameters are from reference [190]. dKinetic parameters are from reference [393]. eKinetic parameters are from reference [396]. fKinetic parameters are from reference [443]. gKinetic parameters are from reference [420].
269
Table 11.3 Kinetic parameters for nucleotide incorporation opposite template base dT at 37 °C. DNA k K k /K Selection dNTP p d p d polymerase (s-1) (μM) (μM-1s-1) factora Pol β dATP 32 ± 1 9.2 ± 1.0 3.5 PMPA-DP 4.7 ± 0.5 50 ± 10 9.4 × 10-2 40 Pol λ dATPb 1.5 ± 0.1 0.9 ± 0.3 1.7 PMPA-DP 0.095 ± 0.008 3.7 ± 0.9 2.6 × 10-2 65 Pol η dATP 35 ± 3 130 ± 26 2.7 × 10-1 PMPA-DP 0.0134 ± 0.0007 90 ± 10 1.5 × 10-4 1,800 Pol κ dATP 2.49 ± 0.08 7.0 ± 1.0 3.6 × 10-1 PMPA-DP 0.010 ± 0.005 3000 ± 3.3 × 10-6 110,000 2000 Pol ι dATP 0.015 ± 0.001 260 ± 40 5.8 × 10-5 PMPA-DP Could not measure high Rev1 dATP 0.00152 ± 0.00005 2.0 ± 0.3 7.6 × 10-4 PMPA-DP Could not measure high Pol γ dATPc 45 ± 1 0.8 ± 0.1 56 PMPA-DPd 0.21 ± 0.01 40.3 ± 5.7 5.2 × 10-3 10,800 T7 exo- dATPe 156 ± 8 8 ± 2 19.5 PMPA-DPe 0.096 ± 0.009 268 ± 39 3.6 × 10-4 54,400 HIV-1 RT dATPe 41.3 ± 0.6 8.1 ± 0.9 5.1 PMPA-DPe 49 ± 5 58 ± 11 8.4 × 10-1 6 a Calculated as (kp/Kd)dATP/(kp/Kd)PMPA-DP. bKinetic parameters are from reference [117]. cKinetic parameters are from reference [90]. dKinetic parameters are from reference [394]. eKinetic parameters are from reference [416] and T7 exo- was assayed at 20 °C.
270
Table 11.4 Kinetic parameters for nucleotide incorporation opposite template base dA at 37 °C. DNA k K k /K Selection dNTP p d p d polymerase (s-1) (μM) (μM-1s-1) factora Pol β dTTP 12.3 ± 0.5 3.5 ± 0.5 3.5 AZT-TP 0.0199 ± 0.0007 15 ± 2 1.3 × 10-3 2,600 Pol λ dTTPb 3.9 ± 0.2 2.6 ± 0.4 1.5 AZT-TP 0.0104 ± 0.0004 6 ± 1 1.7 × 10-3 865 Pol η dTTP 35 ± 1 41 ± 5 8.5 × 10-1 AZT-TP 0.31 ± 0.05 478 ± 155 6.5 × 10-4 1,300 Pol κ dTTP 4.35 ± 0.04 11.0 ± 0.5 4.0 × 10-1 AZT-TP 0.0144 ± 0.0005 80 ± 9 1.8 × 10-4 2,200 Pol ι dTTP 0.75 ± 0.02 13 ± 1 5.8 × 10-2 AZT-TP 0.0037 ± 0.0003 90 ± 20 4.1 × 10-5 1,400 Rev1 dTTP 0.0038 ± 0.0003 6 ± 1 6.3 × 10-4 AZT-TP 0.00119 ± 202 ± 26 5.9 × 10-6 110 0.00005 Pol γ dTTPc 25 ± 2 0.6 ± 0.16 42 AZT-TPd 10,000,000 HIV-1 RT dTTPe 16.7 19 8.8 × 10-1 AZT-TPe 0.7 2 3.5 × 10-1 2.5 a Calculated as (kp/Kd)dTTP/(kp/Kd)AZT-TP. bKinetic parameters are from reference [117]. cKinetic parameters are from reference [90]. dKinetic data are from reference [419]. eKinetic parameters are from reference [444].
271 11.7 Figures
Figure 11.1 Chemical structures of NRTIs investigated in this study and their natural counterpart.
272 A
B C
30 0.03
25 0.025
20 0.02 ) -1 15
(s 0.015 obs 10 k
Product (nM) Product 0.01
5 0.005
0 0 0 200 400 600 800 1000 0 5 10 15 20 25 30 Time (s) L-3TC-TP (μM)
Figure 11.2 Concentration dependence on the pre-steady state rate constant of L-3TC-TP incorporation catalyzed by Pol η. A pre-incubated solution of Pol η (120 nM) and 5′- [32P]-labeled 21-41merGT DNA (30 nM, Table 1) was rapidly mixed with increasing concentrations of L-3TC-TP•Mg2+ (0.2 μM, z; 0.5 μM, {; 1 μM, ; 2 μM, ; 5 μM, S; 10 μM, U; and 25 μM, ) for various time intervals. (A) A representative gel image is shown for Pol η inserting L-3TC-TP at 25 μM. The lengths of the DNA primer are indicated in the right margin. (B) The concentration of DNA product was plotted as a function of time. The solid lines are the best fits to a single-exponential equation which determined the observed rate constant, kobs. (C) The kobs values were plotted as a function of L-3TC-TP concentration. The data (z) were then fit to a hyperbolic equation, yielding -1 a kp of 0.0296 ± 0.0009 s and a Kd of 3.3 ± 0.3 μM.
273 Chapter 12: Kinetic Investigation of the Incorporation of Anti-HBV Nucleotide Analogs Catalyzed by Non-Canonical Human DNA Polymerases
12.1 Introduction
With more than two billion people infected worldwide, hepatitis B virus (HBV) remains an important global health concern. Chronic HBV infection, which affects more than 350 million people, is a major cause of hepatocellular carcinoma and liver cirrhosis, two life- threatening disease states of the liver. Thus, HBV treatment is important to prevent or to slow the progression of these severe liver complications. Currently, seven antiviral agents are approved by the United States Food and Drug Administration (FDA) for treatment of HBV: two immune modulators (interferon alfa and pegylated interferon alfa) and five nucleoside analogs [adefovir (PMEA), tenofovir (PMPA), lamivudine (L-3TC), telbivudine (L-TBV), and entecavir (ETV) (Figure 12.1)]. Following cellular uptake, these analogs undergo either two (PMEA and PMPA) or three (L-3TC, L-TBV, and ETV) phosphorylation events to be activated to their di- (PMEA-DP and PMPA-DP) or triphosphate (L-3TC-TP, L-TBV-TP, and ETV-TP) forms, respectively. These activated nucleotide analogs target the HBV DNA polymerase (Pol) which has enzymatic activity for a unique protein-priming event, RNA-dependent and DNA-dependent DNA synthesis, and degradation of RNA in a RNA/DNA duplex (i.e. RNase H). Depending on the analog, these drugs may function as competitive inhibitors against natural dNTP substrates and/or as obligate or masked chain terminators that inhibit the priming and/or polymerization activities of the HBV Pol [445]. Unfortunately, the usage of anti-HBV nucleoside analogs is limited by drug resistance and adverse side effects [446, 447]. It has been postulated that cellular DNA polymerases, such as human DNA polymerase γ (Pol γ), may be potential drug targets and the cause of observed clinical toxicity, since
274 nucleoside analogs approved for human immunodeficiency virus type 1 (HIV-1) are associated with mitochondrial toxicity [395, 448]. However, mitochondrial toxicity induced by nucleotide analog incorporation catalyzed by Pol γ does not account for all of the unwanted side effects [397]. The human genome encodes at least 15 other Pols, members of the A-, B-, X- or Y-family, that may be potential candidates for generating cellular toxicity via analog incorporation into nuclear DNA [397, 398, 405, 406].
Using pre-steady state kinetic techniques, we determined the incorporation efficiency of five anti-HBV nucleotide analogs (Figure 12.1) with six non-canonical human DNA polymerases: Pols β, λ, η, ι, κ, and Rev1. Human beings possess five X-family DNA polymerases and the two selected X- family members (Pols β and λ) in this paper participate in cellular processes such as base excision repair, nonhomologous end-joining repair, and antibody generation [30]. In human beings, there are four Y-family DNA polymerases (Pols η, ι, κ, and Rev1) that catalyze translesion DNA synthesis and may play a part in somatic hypermutation [80]. Therefore, inhibition of these selected X- and Y-family pols could lead to unwanted toxicity including apoptosis, genetic instability, and immunodeficiency. Our kinetic data showed that most of the analogs are substrates for the non-canonical pols and that the kinetic basis of incorporation varies for each analog. These results suggest that human X- and Y-family enzymes are capable of inserting nucleotide analogs in vivo and establish structure-function relationships that are important for future anti-HBV drug design.
12.2 Materials and Methods
Materials 32 These chemicals were purchased from the following companies: [γ- P]ATP, MP Biomedicals; ATP, GE Healthcare; Bio-Spin 6 columns, Bio-Rad Laboratories; deoxyribonucleotide 5′-triphosphates, GE Healthcare; β-L-2′-deoxythymidine 5′- triphosphate (L-TBV-TP), TriLink Biotechnologies; OptiKinase™, USB Corporation;
275 synthetic oligodeoxyribonucleotides 21-mer, 5′-phosphorylated 19-mer, and 41-mers, Integrated DNA Technologies. Diphosphate of 9-[2-(phosphonomethoxy)ethyl]adenine (PMEA-DP) and 9-[2-(phosphonomethoxy)propyl]adenine (PMPA-DP), β-L-2′,3′- dideoxy-3′-thiacytidine 5′-triphosphate (L-3TC-TP), and entecavir 5′-triphosphate (ETV- TP) were kind gifts from Gilead Sciences, Inc.
Preparation of human DNA polymerases and DNA substrates The plasmids, expression, and purification of human DNA polymerases β [235, 274], λ [117], η [392], truncated ι (1-420) [392], truncated κ (9-518) [392], and truncated Rev1 (341-829) [330, 340] were described previously. Purified human Δ235 DNA ligase I was a kind gift of Dr. Tom Ellenberger [278]. Commercially synthesized oligonucleotides in Table 12.1 were purified using polyacrylamide gel electrophoresis [105, 182]. The 21- mer primer was radiolabeled with [γ-32P]ATP and OptiKinase™ according to the manufacturer’s protocol, and the unreacted [γ-32P]ATP was subsequently removed via a Bio-Spin 6 column. The 21-41mer primer-template DNA substrates [105] and 21-19- 41mer single-nucleotide gapped DNA substrates [182] were annealed as described previously.
Single-turnover kinetic assays to measure the kp and Kd Kinetic assays were completed using buffer B (50 mM Tris-HCl pH 7.8 at 37 °C, 5 mM
MgCl2, 50 mM NaCl, 0.1 mM EDTA, 5 mM DTT, 10% glycerol, and 0.1 mg/ml of bovine serum albumin (BSA)) for Pol β, buffer L (50 mM Tris-HCl pH 8.4 at 37 °C, 5
mM MgCl2, 100 mM NaCl, 0.1 mM EDTA, 5 mM DTT, 10% glycerol, and 0.1 mg/ml of
BSA) for Pol λ [117], and buffer Y (50 mM HEPES pH 7.5 at 37 °C, 5 mM MgCl2, 50 mM NaCl, 0.1 mM EDTA, 5 mM DTT, 10% glycerol, and 0.1 mg/ml of BSA) for Pols η, ι, κ, and Rev1. The primer-template DNA substrates were used in assays with Pols η, ι, κ, and Rev1 while the gap-filling Pols β and λ were provided 21-19-41mer gap DNA. All kinetic experiments described herein were performed at 37 °C, and the reported concentrations were final after mixing all the components. A pre-incubated solution of the Pol (120 or 300 nM) and 5′-[32P]-radiolabeled DNA substrate (30 nM) was mixed
276 with increasing concentrations (0.01-900 μM) of a single nucleotide or nucleotide analog in the appropriate buffer at 37 °C. The Pol was in molar excess over DNA, whereby the enzyme to DNA ratio was 4:1 for Pols λ, η, ι, and Rev1 and 10:1 for Pols β and κ. Aliquots of the reaction mixtures were quenched at various times using 0.37 M EDTA. A rapid chemical-quench flow apparatus (KinTek) was utilized for fast nucleotide incorporations. Reaction products were resolved using sequencing gel electrophoresis (17% acrylamide, 8 M urea) and quantitated with a Typhoon TRIO (GE Healthcare). The time course of product formation at each nucleotide concentration was fit to a single- exponential equation (Equation 1)
[Product] = A[1 – exp(–kobst)] (1) using a nonlinear regression program, KaleidaGraph (Synergy Software), to yield an observed rate constant of nucleotide incorporation (kobs). The kobs values were then plotted as a function of nucleotide concentration and fit using the hyperbolic equation (Equation 2)
kobs = kp[dNTP]/{[dNTP] + Kd} (2)
which resolved the maximum rate of nucleotide incorporation (kp) and equilibrium
dissociation constant (Kd) for nucleotide incorporation catalyzed by each enzyme.
Extension assay for non-chain terminators A pre-incubated solution of Pol (240 or 600 nM) and 5′-[32P]-radiolabeled 21/41mer DNA (60 nM) in the appropriate buffer was mixed with either ETV-TP•Mg2+ (1-50 μM) or L-TBV-TP•Mg2+ (100-500 μM) to allow sufficient extension (at least 8 half-lives) of the primer before adding the four natural dNTPs (200 μM) for various reaction times. The enzyme to DNA ratio was 4:1 for Pols λ, η, ι, and Rev1 and 10:1 for Pols β and κ. Aliquots of the reaction mixtures were quenched using 0.37 M EDTA, and the reaction products were resolved using sequencing gel electrophoresis (20% acrylamide, 8 M urea).
277 DNA ligation assay for non-chain terminators A pre-incubated solution of Pol β (120 nM) and 5′-[32P]-radiolabeled 21-19-41mer DNA (30 nM) in buffer B was mixed in independent reactions with dGTP•Mg2+ (1 μM), dTTP•Mg2+ (1 μM), ETV-TP•Mg2+ (1 μM) or L-TBV-TP•Mg2+ (500 μM) to allow sufficient extension (at least 8 half-lives; 30 s for dGTP and dTTP, 5 min for ETV-TP, and 120 min for L-TBV-TP) of the primer before adding human Δ235 DNA ligase I (30 nM) and 1 mM ATP to the polymerization mixture. Aliquots of the reaction mixtures were quenched at various times by adding them to 0.37 M EDTA and immediate heat denaturation for 2 min at 95 °C. The DNA products were resolved using sequencing gel electrophoresis (20% acrylamide, 8 M urea).
12.3 Results
Determination of selection factors The mechanism of anti-HBV nucleotide analog incorporation catalyzed by Pols β, λ, η, ι, κ, and Rev1 was determined by utilizing single-turnover kinetic methodology. When the enzyme is in molar excess over DNA, the conversion of the DNA substrate into product is observed directly in a single pass through the enzymatic pathway so that the nucleotide concentration dependence on the observed rate constant (kobs) can resolve the equilibrium
dissociation constant (Kd) and maximum rate of incorporation (kp) for an incoming nucleotide [63]. As a representative example, a pre-incubated solution of Pol λ and 5′- [32P]-radiolabeled 21-19C-41merTG (Table 12.1) was mixed with increasing concentrations of PMEA-DP (Section 12.2, Figure 12.2 part (A)). After plotting and fitting the data into Equations 1 and 2 (Figure 12.2 parts (B) (C)), the single-turnover -1 kinetic parameters were resolved: a kp of 0.175 ± 0.004 s and a Kd of 4.5 ± 0.3 μM. Using similar single-turnover kinetic assays, the kinetic parameters were measured for each enzyme incorporating a natural nucleotide or one of the analogs to calculate the
substrate specificity constants (kp/Kd) and selection factors ((kp/Kd)dNTP/(kp/Kd)analog) in Tables 12.2-12.5.
278 Incorporation of acyclic adenine analogs Tenofovir (PMPA) and adefovir (PMEA) both have an acyclic moiety and a phosphonate group, but PMPA has an additional methyl group (Figure 12.1). In general, the selection factors were lower for PMEA-DP (44 to 250) than PMPA-DP (40 to >110,000) (Table 12.2). The X-family DNA polymerases exhibited the least amount of discrimination for
both analogs, whereby the kp value dropped by an average of 14-fold and the Kd value increased by an average of 5-fold. In contrast, the additional methyl group in PMPA-DP led to a greater degree of discrimination for the Y-family DNA polymerases, although, different mechanisms of discrimination were observed for the four enzymes. For Pol η, the rate of PMPA-DP and PMEA-DP incorporation dropped by an average of 420-fold while the binding was slightly tighter than dATP. For Pols κ, ι, and Rev1, the rate of PMEA-DP incorporation decreased by 60-, 30-, and 4-fold, and the ground-state binding
affinity (1/Kd) was 3-fold weaker, unchanged, and 60-fold weaker, respectively. The catalytic efficiency of these three enzymes dropped dramatically for PMPA-DP insertion, and the kinetic parameters could only be measured for Pol κ as noted previously (Chapter 11) that PMPA-DP incorporation was too inefficient to be observed with Pol ι and Rev1.
Incorporation of nucleotides with L-stereochemistry In the preceding chapter, the selection factors were determined for Pols β, λ, η, ι, κ, and
Rev1 inserting L-3TC-TP, and the following general mechanism emerged: the kp drops significantly (6,100-fold on average) while the Kd decreases (6-fold on average) compared to dCTP (Table 12.3). Telbivudine (L-TBV) is also an L-nucleoside analog approved for HBV, however, it has thymidine as the base and the ribose remains unmodified (Figure 12.1). No observable L-TBV-TP incorporation was detected for Rev1, likely because it is a deoxycytidyl transferase and prefers dCTP regardless of the template base [190]. For the remaining DNA polymerases, relatively large selection factors were determined (Table 12.4). Unlike L-3TC-TP, the binding of L-TBV-TP to the Pol•DNA complex was weakened by 10-fold on average but the rate of incorporation remained slow compared to dTTP. These kinetic results suggested that the oxathiolane ring of L-3TC-TP is important for the tight ground-state binding affinity of L-3TC-TP
279 and that the polymerase active sites of these Y-family polymerases could accommodate L-dNTPs.
Incorporation of entecavir 5′-triphosphate Entecavir (ETV) is a deoxyguanosine analog with a cyclopentyl sugar ring (Figure 12.1). Interestingly, Pols β, λ, η, ι, κ, and Rev1 incorporated ETV-TP opposite dC with much lower selection factors (Table 12.5) compared to the other anti-HBV analogs (Tables 12.2-12.4). Except for Rev1, the lack of discrimination between ETV-TP and dGTP was due to the binding affinity of ETV-TP being approximately 20-fold tighter on average. However, the rate of ETV-TP incorporation was on average 120-fold slower than dGTP.
Qualitative analysis of non-chain terminators being embedded into DNA ETV and L-TBV are not obligate chain terminators like PMEA, PMPA, and L-3TC (Figure 12.1), since these two analogs possess the required 3′-hydroxyl group for downstream enzymatic steps such as primer extension or DNA ligation if incorporated into a gapped DNA substrate. Therefore, it is possible that these analogs could become embedded into the human genome. To examine the primer extension possibility, we pre- incubated a solution of the enzyme and the appropriate 5′-[32P]-radiolabeled 21-41mer DNA substrate before initiating the reaction with either ETV-TP or L-TBV-TP (Section 12.2). After allowing sufficient time for the analog to be incorporated, the four natural dNTPs were added to the reaction mixture. Representative gel images are shown in Figures 12.3 and 12.4 for an X-family (Pol β) and a Y-family (Pol η) DNA polymerase, respectively. Both Pol β and η efficiently catalyzed full-length product (41mer) for the control and ETV-TP as early as 10 s after the addition of the natural dNTPs. In contrast, minimal extension of an L-TBV-MP terminated primer was observed after 1 h, and the small amount of 41mer may be partially due to the extension of the unreacted 21mer substrate rather than the 22mer terminated with the analog. Similar extension activities of both analogs were detected for Pols λ, ι, and κ (data not shown). For Rev1, the main product after ETV-TP incorporation was 23mer. Subsequent incorporations likely did not occur because Rev1 can only efficiently catalyze the transfer of dCTP when dG is the
280 template [190]. Nonetheless, ETV-TP may be a masked chain terminator for Rev1. Together, these preliminary data suggested L-TBV-TP was a chain terminator while ETV-TP was not for Pols β, λ, η, ι, κ, and Rev1.
Next, we determined whether single-nucleotide gap DNA substrates having primers terminated with ETV-MP or L-TBV-MP are substrates for human DNA ligase I. A pre- incubated solution of Pol β and the appropriate 5′-[32P]-radiolabeled 21-19/41mer DNA were reacted with either ETV-TP or L-TBV-TP before initiating the ligation reaction with human Δ235 DNA ligase I and ATP (Section 12.2). As shown in Figure 12.5, the ligated DNA product (41mer) was detected for ETV-TP and the control reactions but not L-TBV-TP. However, the ligation of ETV-MP was less efficient than the dGMP control, for the first visible appearance of 41mer was at 10 s and 300 s for dGMP and ETV-MP, respectively. Based on the extension and ligation assays, these results suggested that ETV-TP can be incorporated and embedded into the genome via primer extension or subsequent ligation while L-TBV-TP would act as a chain terminator that may lead to single-strand DNA breaks. To eliminate ambiguity in interpreting the likelihood of extension and ligation, we are currently preparing primers with L-TBV-MP or ETV-MP at the 3′-terminus and will directly examine these reactions.
12.4 Discussion
Overall trends of anti-HBV drugs by non-canonical human DNA polymerases Ranking the selection factors of the five anti-HBV analogs for each enzyme (Tables 12.2- 12.5) generated the following profiles: the order is identical for Pols λ, η, and Rev1 (ETV-TP < PMEA-DP < L-3TC-TP < PMPA-DP < L-TBV-TP) and Pols κ and ι (ETV- TP < PMEA-DP < L-3TC-TP < L-TBV-TP < PMPA-DP) while Pol β is unique (ETV-TP < PMPA-DP < PMEA-DP < L-3TC-TP < L-TBV-TP). Consistently, the lowest level of discrimination occurred with ETV-TP whereas PMPA-DP and/or L-TBV-TP had the highest selection factors for the six human pols examined in this work. The different
281 chemical modifications resulted in unique mechanisms of incorporation. Some modifications (i.e. L-stereochemistry and acyclic ribose) were unfavorable for incorporation while others (i.e. oxathiolane and methylene moieties in the sugar ring) enhanced the incorporation efficiency by forming a more stabile ternary complex through tighter binding.
Potential relationship between clinical toxicity and drug incorporation by host DNA pols Nucleoside analogs have been successful at managing viral infections such as HIV and HBV, although, drug usage is limited by drug resistance and unwanted side effects [446]. Most of the anti-HBV nucleoside analogs are relatively well tolerated and are associated with mild side effects that include headache, fatigue, dizziness, nausea, diarrhea, and flu- like symptoms [446, 449-451]. Unfortunately, a striking percentage of HBV patients report moderate to severe side effects: 22% on telbivudine [450], 15% on entecavir [452], and 18% on lamivudine [452]. Specifically, telbivudine has been associated with myopathy [453] and myalgia [454]; tenofovir [455] and adefovir [446, 451] have been associated with nephrotoxicity [447]; lamivudine has been associated with myopathy [456, 457], peripheral neuropathy [458], and Fanconi syndrome [456] on occasion [447]; and entecavir has been associated with myopathy [447]. Most of these adverse events are thought to be derived from mitochondrial toxicity due to their resemblance of mitochondrial diseases [454, 459, 460], although, some assays indicate the anti-HBV nucleoside analogs induce relatively low mitochondrial toxicity [418, 453, 461-463] and/or are weak inhibitors of Pol γ [394, 396, 463, 464]. However, problems induced by the incorporation of nucleotide analogs into nuclear DNA can also lead to mitochondrial dysfunction [397], therefore, the other 15 known human DNA polymerases are candidates for incorporating the drug analogs and potentially causing unwanted side effects. Since human replicative polymerases (e.g. Pols α, δ, and ε) employ stringent mechanisms of nucleotide selection, these enzymes are unlikely to incorporate PMEA- DP [465], PMPA-DP [410], ETV-TP [445], L-TBV-TP [453], or L-3TC-TP [466], indicating these drugs are relatively weak inhibitors. Furthermore, the selection factors measured for Pol γ [90, 394] and T7 DNA polymerase [416], two replicative A-family
282 enzymes, incorporating PMPA-DP is 10,800 and 54,400, respectively, which is significantly larger than the values determined for Pol β (40), Pol λ (65), and Pol η (1,800) (Table 12.2). Similarly, Pol γ discriminates between dCTP and L-3TC-TP by 2,900-fold which is larger than most of the selection factors calculated for the non- canonical human pols (46 to 3,300) in Table 12.3. Thus, based on the relatively low selection factors measured herein (Tables 12.2-12.5), human X- and Y-family DNA polymerases are likely to incorporate anti-HBV nucleotide analogs in vivo during DNA damage repair and lesion bypass processes, thereby inducing a cascade of cellular events associated with nucleoside analog toxicity: genomic instability → apoptosis/cell death → side effects [397, 438]. Another important consideration is that Pols β [433], λ [200, 270], η [434], ι [436], κ [412, 435], and Rev1 [70, 330] are expressed at the mRNA level in various human tissues, including those afflicted with the abovementioned side effects such as the kidney and muscle.
Kinetic results support entecavir as a potential carcinogen According to the US prescribing information sheet for entecavir, solid tumors were detected in rodents that were exposed to high doses of entecavir [467]. Entecavir possesses a 3′-hydroxyl group, therefore, the carcinogenic activity may arise when this drug is embedded into genomic DNA. It has been reported that entecavir functions as a masked chain terminator for HBV pol and cellular Pols α, β, γ, δ, and ε [445]. However, our efficient incorporation (Table 12.5), extension (Figures 12.3 and 12.4), and ligation (Figure 12.5) results suggested that there is a high potential for entecavir to become embedded into DNA. Importantly, entecavir is not a masked chain terminator for Pols β, λ, η, ι, and κ, therefore, these polymerases can likely rescue a stalled replication fork due to entecavir incorporation. Together, these processes are likely to contribute to a putative mechanism of carcinogenicity, especially if the embedded drug induces higher error rates during subsequent rounds of replication. Langley et al. [445] did not provide information on their assays with Pol β, therefore, we are unsure of the discrepancy in our incorporation and extension results. Also, ETV-TP has been shown to be a substrate for HIV-1 reverse transcriptase (RT), although, the enzyme discriminates between ETV-TP
283 and dGTP by 220-fold [468], a selection factor that is greater than those measured for the human pols (2 to 32 in Table 12.5). These kinetic results suggested that entecavir would not be a good drug candidate for treating HIV-1 infected patients.
Telbivudine is unlikely to be embedded in DNA Unlike entecavir, telbivudine exhibited an extremely low potential to become embedded in the human genome, since the selection factors were large (8,100 to >126,000) and an L-TBV-MP-terminated primer was poorly extended (Figures 12.3 and 12.4) and was not a substrate for human DNA ligase I (Figure 12.5). The unnatural L-stereochemistry of telbivudine is likely problematic for extension and ligation because of the unfavorable alignment of the 3′-hydroxyl group for catalysis. Thus, L-stereochemistry is less problematic for the incorporation step than subsequent enzymatic steps. For Pols α, β, and ε, the incorporation and extension of L-dCTP or L-dTTP (or L-TBV-TP) at low concentrations was not observed [469] which is likely due to the weak binding affinity and slow rate of incorporation for unmodified L-nucleotides as demonstrated here for L- TBV-TP insertion by six non-canonical human pols (Table 12.4). Interestingly, the oxathiolane ring of L-3TC and L-FTC improves the incorporation efficiency for L-dCTP analogs (Table 12.3). Also noteworthy, human DNA ligase I can tolerate a primer terminated with a modified chemical structure such as a ribonucleotide [278], 8-oxo-7,8- dihydro-2′-deoxyguanosine [235], and 2′-deoxy-2′,2′-difluorocytidine-5′-triphosphate [470].
Concluding remarks Unfortunately, similar transient state kinetic approaches have not been used to determine the selection factors for the HBV polymerase incorporating nucleotide analogs, so it is difficult to discern how selective the anti-HBV drugs are for the viral enzyme versus the human DNA polymerases. Nonetheless, these findings highlight the importance of the in vivo investigation to confirm whether the non-canonical human DNA polymerases contribute to drug toxicity. In addition, this work established structure-function relationships that will be important in designing nucleoside analogs to overcome the
284 limitations of clinical toxicity and drug resistance associated with current FDA-approved drugs.
12.5 Future Directions
As mentioned in the previous chapter (Section 11.5), the selection factors for nucleotide analog incorporation opposite sites of DNA damage may be more relevant for assessing potential drug toxicity. More importantly, the bypass of entecavir needs to be investigated. Our preliminary data suggests the bypass is inefficient and error-prone for the six non-canonical DNA polymerases. The preferred nucleotide opposite entecavir, a dG analog, is dATP, although, the incorporation is inefficient. To better understand the molecular basis of entecavir inducing a carcinogenic effect in rodents, we are in the process of using the methodology described in Chapter 10 (Section 10.2) to establish a comprehensive mechanism of entecavir bypass by Pols β, λ, η, ι, κ, and Rev1.
285 12.6 Tables
Table 12.1 Sequences of oligonucleotidesa. 21mer 5’-CGCAGCCGTCCAACCAACTCA-3’ 19merC 5’-CGTCGATCCAATGCCGTCC-3’ 19merA 5’-AGTCGATCCAATGCCGTCC-3’ 41merGT 3’-GCGTCGGCAGGTTGGTTGAGTGTCAGCTAGGTTACGGCAGG-5’ 41merTG 3’-GCGTCGGCAGGTTGGTTGAGTTGCAGCTAGGTTACGGCAGG-5’ 41merCG 3’-GCGTCGGCAGGTTGGTTGAGTCGCAGCTAGGTTACGGCAGG-5’ 41merAG 3’-GCGTCGGCAGGTTGGTTGAGTAGCAGCTAGGTTACGGCAGG-5’ aThe 21mer strand was 5′-radiolabeled. For single-nucleotide gap DNA substrates, the downstream 19mer strand was 5′-phosphorylated. The identity of important base changes is highlighted in bold.
286
Table 12.2 Kinetic parameters for nucleotide incorporation opposite template dT at 37 °C.
DNA kp Kd kp/Kd Selection dNTP polymerase (s-1) (μM) (μM-1s-1) factora Pol β dATPb 32 ± 1 9.2 ± 1.0 3.5 PMPA-DPb 4.7 ± 0.5 50 ± 10 9.4 × 10-2 40 PMEA-DP 1.31 ± 0.04 65 ± 6 2.0 × 10-2 170 Pol λ dATPc 1.5 ± 0.1 0.9 ± 0.3 1.7 PMPA-DPb 0.095 ± 0.008 3.7 ± 0.9 0.026 65 PMEA-DP 0.175 ± 0.004 4.5 ± 0.3 0.039 44 Pol η dATPb 35 ± 3 130 ± 26 0.27 PMPA-DPb 0.0134 ± 0.0007 90 ± 10 1.5 × 10-4 1,800 PMEA-DP 0.069 ± 0.008 55 ± 18 1.3 × 10-3 208 Pol κ dATPb 2.49 ± 0.08 7.0 ± 1.0 0.36 PMPA-DPb 0.010 ± 0.005 3000 ± 2000 3.3 × 10-6 110,000 PMEA-DP 0.039 ± 0.001 23 ± 3 1.7 × 10-3 210 Pol ι dATPb 0.015 ± 0.001 260 ± 40 5.8 × 10-5 PMPA-DPb Could not measure high PMEA-DP 0.00047 ± 0.00004 180 ± 40 2.6 × 10-6 48 Rev1 dATPb 0.00152 ± 0.00005 2.0 ± 0.3 7.6 × 10-4 PMPA-DPb Could not measure high PMEA-DP 0.00037 ± 0.00002 121 ± 23 3.1 × 10-6 250 Pol γ dATPd 45 ± 1 0.8 ± 0.1 56 PMPA-DPe 0.21 ± 0.01 40.3 ± 5.7 5.2 × 10-3 10,800 T7 exo- dATPf 156 ± 8 8 ± 2 19.5 PMPA-DPf 0.096 ± 0.009 268 ± 39 3.6 × 10-4 54,400 a Calculated as (kp/Kd)dATP/(kp/Kd)Analog. bKinetic parameters are from preceding chapter (Table 11.3). cKinetic parameters are from reference [117]. dKinetic parameters are from reference [90]. eKinetic parameters are from reference [394]. fKinetic parameters are from reference [416] and T7 exo- was assayed at 20 °C.
287
Table 12.3 Kinetic parameters for nucleotide incorporation opposite template dG at 37 °C.
DNA kp Kd kp/Kd Selection dNTP polymerase (s-1) (μM) (μM-1s-1) factora Pol β dCTPb 5.02 ± 0.07 0.71 ± 0.04 7.1 L-3TC-TPb 0.00390 ± 0.00010 0.18 ± 0.02 2.2 × 10-2 325 Pol λ dCTPc 1.57 ± 0.04 0.9 ± 0.1 1.7 L-3TC-TPb 0.00402 ± 0.00008 0.106 ± 0.010 0.038 46 Pol η dCTPb 49 ± 2 25 ± 4 2.0 L-3TC-TPb 0.0296 ± 0.0009 3.3 ± 0.3 9.0 × 10-3 220 Pol κ dCTPb 11.8 ± 0.6 32 ± 5 0.37 L-3TC-TPb 0.00045 ± 0.00001 4.0 ± 0.5 1.1 × 10-4 3,300 Pol ι dCTPb 0.075 ± 0.002 50 ± 5 1.5 × 10-3 L-3TC-TPb 0.00356 ± 0.00009 96 ± 8 3.7 × 10-5 40 Rev1 dCTPd 22.4 ± 0.9 2.2 ± 0.3 10 L-3TC-TPb 0.0236 ± 0.0006 1.04 ± 0.10 2.3 × 10-2 450 Pol γ dCTPe 44 ± 2 1.1 ± 0.1 40 L-3TC-TPe 0.125 ± 0.005 9.2 ± 0.9 1.4 × 10-2 2,900 a Calculated as (kp/Kd)dCTP/(kp/Kd)Analog. bKinetic parameters are from the preceding chapter (Table 11.2). cKinetic parameters are from reference [117]. dKinetic parameters are from reference [190]. eKinetic parameters are from reference [393].
288
Table 12.4 Kinetic parameters for nucleotide incorporation opposite template dA at 37 °C.
DNA kp Kd kp/Kd Selection dNTP polymerase (s-1) (μM) (μM-1s-1) factora Pol β dTTPc 12.3 ± 0.5 3.5 ± 0.5 3.5 L-TBV-TP 0.00055 ± 0.00001 11 ± 1 5.0 × 10-5 70,000 Pol λ dTTPb 3.9 ± 0.2 2.6 ± 0.4 1.5 L-TBV-TP 0.00072 ± 0.00002 53 ± 6 1.4 × 10-5 11,000 Pol η dTTPc 35 ± 1 41 ± 5 0.85 L-TBV-TP 0.0028 ± 0.0003 410 ± 100 6.8 × 10-6 126,000 Pol κ dTTPc 4.35 ± 0.04 11.0 ± 0.5 0.40 L-TBV-TP 0.00065 ± 0.00006 91 ± 15 7.1 × 10-6 55,000 Pol ι dTTPc 0.75 ± 0.02 13 ± 1 5.8 × 10-2 L-TBV-TP 0.00071 ± 0.00006 100 ± 30 7.1 × 10-6 8,100 Rev1 dTTPd 0.0038 ± 0.0003 6 ± 1 6.3 × 10-4 L-TBV-TP No observed incorporation High a Calculated as (kp/Kd)dTTP/(kp/Kd)L-TBV-TP. bKinetic parameters are from reference [117]. cKinetic parameters are from preceding chapter (Table 11.4). dKinetic parameters are from reference [190].
289
Table 12.5 Kinetic parameters for nucleotide incorporation opposite template dC at 37 °C.
DNA kp Kd kp/Kd Selection dNTP polymerase (s-1) (μM) (μM-1s-1) factora Pol β dGTP 18.8 ± 0.4 8.7 ± 0.4 2.2 ETV-TP 0.054 ± 0.002 0.26 ± 0.04 0.21 10 Pol λ dGTPb 2.5 ± 0.1 2.1 ± 0.3 1.2 ETV-TP 0.034 ± 0.002 0.09 ± 0.02 0.38 3 Pol η dGTP 38 ± 2 80 ± 12 0.48 ETV-TP 0.24 ± 0.01 2.4 ± 0.4 0.1 5 Pol κ dGTP 4.4 ± 0.1 10.6 ± 0.9 0.42 ETV-TP 0.370 ± 0.004 4.2 ± 0.1 8.8 × 10-2 5 Pol ι dGTP 0.095 ± 0.002 141 ± 8 6.7 × 10-4 ETV-TP 0.0097 ± 0.0004 33 ± 5 2.9 × 10-4 2 Rev1 dGTP 0.0038 ± 0.0002 3.9 ± 0.8 9.7 × 10-4 ETV-TP 0.0017 ± 0.0001 56 ± 10 3.0 × 10-5 32 HIV-1 RT dGTPc 18.3 ± 1.30 1.76 ± 0.51 10.4 ETV-TPc 0.107 ± 0.007 2.23 ± 0.67 4.8 × 10-2 220 a Calculated as (kp/Kd)dGTP/(kp/Kd)ETV-TP. bKinetic parameters are from reference [117]. cKinetic parameters are from reference [468].
290 12.7 Figures
Figure 12.1 Chemical structures of anti-HBV nucleoside analogs and their natural counterpart.
291 A
B C
30 0.16
0.14 25 0.12 20
) 0.1 -1
15 (s 0.08 obs
k 0.06 10 Product (nM) 0.04 5 0.02
0 0 0 50 100 150 200 250 300 350 400 0 5 10 15 20 25 30 Time (s) PMEA-DP (μM)
Figure 12.2 Concentration dependence on the pre-steady state rate constant of PMEA-DP incorporation catalyzed by Pol λ. A pre-incubated solution of Pol λ (120 nM) and 21-19C- 41merTG DNA (30 nM) was rapidly mixed with increasing concentrations of PMEA- DP•Mg2+ (0.2 μM, z; 0.5 μM, {; 1 μM, ; 2 μM, ; 5 μM, S; 10 μM, U; and 25 μM, ) for various time intervals. (A) A representative gel image for PMEA-DP incorporation at 1 μM is shown. The length of the DNA primer is indicated in the right margin. (B) The concentration of the DNA product is plotted as a function of time. The solid lines are the best fits to a single-exponential equation which determined the observed rate constant, kobs. (C) The kobs values were plotted as a function of PMEA-DP concentration. The data -1 (z) were then fit to a hyperbolic equation, yielding a kp of 0.175 ± 0.004 s and a Kd of 4.5 ± 0.3 μM.
292
Figure 12.3 Extension of ETV-MP and L-TBV-MP catalyzed by Pol β. A pre-incubated solution of Pol β (600 nM) and the appropriate 5′-[32P]-radiolabeled 21/41mer DNA (60 nM) was rapidly mixed with (A) four natural dNTPs (200 μM each), (B) ETV-TP (1 μM), or (C) L-TBV-TP (500 μM) to allow sufficient extension (5 min for ETV-TP and 120 min for L-TBV-TP) of the 21mer to a 22mer that is terminated with the analog. Then, the four natural dNTPs (200 μM each) were added to the reaction mixtures containing ETV-TP or L-TBV-TP for various times. The length of important oligonucleotides is marked in the right margin, and the reaction times with (+) and without (-) the four natural dNTPs are listed below each lane.
293
Figure 12.4 Extension of ETV-MP and L-TBV-MP catalyzed by Pol η. A pre-incubated solution of Pol η (240 nM) and the appropriate 5′-[32P]-radiolabeled 21/41mer DNA (60 nM) was rapidly mixed with (A) four natural dNTPs (200 μM each), (B) ETV-TP (2 μM), or (C) L-TBV-TP (500 μM) to allow sufficient extension (1 min for ETV-TP and 60 min for L-TBV-TP) of the 21mer to a 22mer that is terminated with the analog. Then, the four natural dNTPs (200 μM each) were added to the reaction mixtures containing ETV-TP or L-TBV-TP for various times. The length of important oligonucleotides is marked in the margins, and the reaction times with (+) and without (-) the four dNTPs are listed below each lane.
294
Figure 12.5 Ligation of ETV-MP and L-TBV-MP catalyzed by human DNA ligase I. A pre-incubated solution of Pol β (120 nM) and the appropriate 5′-[32P]-radiolabeled 21- 19/41mer DNA substrate (30 nM) was rapidly mixed with (A) dGTP (1 μM), (B) ETV- TP (1 μM), (C) L-TBV-TP (500 μM), or (d) dTTP (1 μM) to allow sufficient extension (30 s for dGTP and dTTP, 5 min for ETV-TP, and 120 min for L-TBV-TP) of the 21mer to a 22mer that is terminated in the added dNTP or analog. Then, the ligation reaction was initiated by adding human Δ235 DNA ligase I (30 nM) and ATP (1 mM) for various times. The length of important oligonucleotides is marked in the right margin, and the ligation times are listed below each lane.
295 Chapter 13: Additional Kinetic Studies on the Mechanism of Nucleotide Incorporation Catalyzed by Human DNA Polymerases λ and β
13.1 Introduction
Among the various DNA polymerases organized into the A-, B-, C-, D-, X-, or Y-family [7, 82, 83], these enzymes have been shown to share a minimal kinetic mechanism for DNA polymerization and a common structural organization of the fingers, palm, and thumb subdomains [41, 54]. In short, a DNA polymerase binds to the DNA substrate and incoming nucleotide (dNTP) to catalyze phosphodiester bond formation via a two metal- ion mechanism which is associative-like [41, 50, 51]. More recently, the minimal kinetic mechanism has been expanded with the addition of Steps 4 and 6 which represent local active site rearrangements (Scheme 13.1). Extensive kinetic, structural, and computational characterization of these enzymes has sparked a debate about the rate- limiting step of nucleotide incorporation catalyzed by DNA polymerases [54, 56, 57]. Possible rate-limiting steps include a pre-chemistry conformational change (Step 3), local active site rearrangements (Step 4), or the chemical reaction (Step 5) (Scheme 13.1).
We investigated the rate-limiting step of nucleotide incorporation catalyzed by two human X-family DNA polymerases: DNA polymerase λ (Pol λ) and DNA polymerase β (Pol β). These enzymes share 34% sequence similarity, possess 5′-deoxyribose-5- phosphate lyase (dRPase) and DNA polymerase domains, lack 3′ → 5′ exonuclease activity, and exhibit a base substitution fidelity of 10-4 to 10-5 [114, 117, 172, 270, 471]. The primary cellular role of Pol β is gap-filling DNA synthesis during base excision repair (BER) [76, 77, 225] while Pol λ has been postulated to function in BER, non- homologous end joining, and V(D)J recombination [78, 172-177, 191, 228, 472]. The
296 participation of Pol λ in multiple gap-filling processes is partially due to its N-terminal region (i.e. nuclear localization signal, BRCT domain, and proline-rich linker to its C- terminal dRPase and polymerase domains), since the BRCT domain is important for protein-protein interactions during recruitment of DNA repair proteins [473].
Despite possessing a similar primary sequence and basic enzymatic activities, Pol λ and Pol β exhibit various nuances in their mechanisms of nucleotide incorporation. For example, Pol λ binds tightly to both correct and incorrect dNTPs whereas Pol β binds tightly to correct dNTPs but weakly to incorrect dNTPs (Chapter 5) [114, 117]. Also, Pol β and Pol λ undergo different conformational changes during catalysis. Binary (E•DNA) and ternary (E•DNA•ddNTP) crystal structures have been solved for human Pol β or Pol λ in complex with single-nucleotide gapped DNA. Overlapping these structural snapshots reveals a dramatic open-to-closed conformational change for Pol β, however, only four active site residues reposition upon ternary complex formation for a truncated form of Pol λ [55, 119]. In addition, no significant conformational changes were detected for full- length Pol λ using a mass spectrometry-based protein footprinting approach [198]. For rat Pol β, stopped-flow kinetic studies have demonstrated that the pre-chemistry step is faster than the rate of nucleotidyl transfer, thereby suggesting another step is rate-limiting in the kinetic pathway (Scheme 13.1) [58]. To help resolve the “rate-limiting step” debate, the mechanism of nucleotide incorporation catalyzed Pol β or Pol λ was examined using three kinetic probes: (i) alpha-thio elemental effect, (ii) pulse-chase and pulse-quench experiments, and (iii) Arrhenius activation energy barrier.
13.2 Materials and Methods
Materials 32 These chemicals were purchased from the following companies: [γ- P]ATP and [α- 32 P]dATP, MP Biomedicals (Solon, OH); deoxyribonucleotides-5′-triphosphates (dNTP), GE Healthcare (Piscataway, NJ); Biospin columns, Bio-Rad Laboratories (Hercules,
297 CA); OptiKinase™, USB Corporation (Cleveland, OH); Sp-dATPαS and Sp-dTTPαS, Biolog-Life Science Institute (Bremen, Germany); synthetic oligodeoxyribonucleotides 21-mer, 5′-phosphorylated 19-mer, and 41-mers, Integrated DNA Technologies (Coralville, IA).
Pre-steady state kinetic assays Wild-type human DNA polymerase β, wild-type human Pol λ (1-575), dPol λ (131-575), and tPol λ (245-575) were expressed in E. coli and purified as described previously [117, 182, 235, 273, 274]. The single-nucleotide gapped DNA substrates shown in Table 13.1 were prepared as described previously [182]. All experiments using a form of Pol λ were performed in optimized reaction buffer L which contained 50 mM Tris-HCl (pH 8.4 at
17, 21, 25, 29, 33, or 37 °C), 5 mM MgCl2, 100 mM NaCl, 5 mM DTT, 10% glycerol, and 0.1 mg/ml BSA [182]. All experiments using Pol β were performed at 37 °C in reaction buffer B which contained 50 mM Tris-HCl (pH 7.8 at 17, 21, 25, 29, 33, or 37
°C), 5 mM MgCl2, 50 mM NaCl, 5 mM DTT, 10% glycerol, and 0.1 mg/ml BSA. Fast reactions were carried out at 37 °C using a rapid chemical-quench flow apparatus (KinTek, PA). All reported concentrations are the final reaction values.
Measurement of the phosphorothioate elemental effect A pre-incubated solution of Pol λ (300 nM) and 5′-radiolabeled 21-19/41mer D-T (30 nM) was rapidly mixed with either dATP (100 µM) or Sp-dATPαS (100 µM, >95% purity) in buffer L at 37 °C and terminated with 0.37 M EDTA. For incorporation of an incorrect nucleotide, the reaction was initiated with either dTTP (100 µM) or Sp-dTTPαS (100 µM, >95% purity). The Sp isomers were used, opposed to the Rp isomers, due to the stereoselectivity observed previously [147, 148].
Pulse-chase and pulse-quench experiments A pre-incubated solution containing Pol λ or Pol β (1 μM) and unlabeled D-T DNA substrate (100 nM) was loaded into one sample loop and rapidly mixed with the 32 appropriate buffer containing [α- P]dATP (10 μM) from a second sample loop for 298 reaction times ranging from 28 ms to 6 s at 37 °C. In the pulse-quench experiment, reactions were immediately quenched with 1 M HCl. In the pulse-chase experiment, reactions were immediately chased with 1.0 mM unlabeled dATP for 30 s, followed by quenching with 1 M HCl. In both cases, quenched reactions were treated with chloroform and neutralized with 1 M NaOH.
Measurement of the kinetic parameters (kp and Kd) for single-nucleotide incorporation A pre-incubated solution containing Pol λ or Pol β (300 nM) and a single-nucleotide gapped D-T DNA substrate (30 nM) was mixed with increasing concentrations of nucleotide (0.2-2000 μM) in the appropriate buffer and reaction temperature. Aliquots of the reaction mixtures were quenched at various times using 0.37 M EDTA.
Product analysis Reaction products were analyzed by sequencing gel electrophoresis (17% acrylamide, 8 M urea, 1× TBE running buffer) and quantitated using a Typhoon TRIO (GE Healthcare) and ImageQuant software (Molecular Dynamics). For the pulse-chase and pulse-quench assays, DNA products were resolved using a 20% highly crosslinked polyacrylamide gel matrix as described previously [149].
Data analysis Data were fit by nonlinear regression using KaleidaGraph (Synergy Software). Data from the single-turnover assays performed at a single dNTP concentration were fit to a single- exponential equation (Equation 1)
[Product] = A[1 – exp(–kobst)] (1)
where A is the reaction amplitude and kobs is the observed single-turnover rate stant. For
measurements of the maxiumum rate of nucleotide incorporation (kp) and equilibrium
dissociation constant (Kd) of an incoming dNTP, the kobs values were then plotted as a function of nucleotide concentration and fit using a hyperbolic equation (Equation 2)
kobs = kp[dNTP]/{[dNTP] + Kd} (2)
299 which resolved the kp and Kd values for nucleotide incorporation. Using kp measured at specific temperatures, the activation energy was determined using the Arrhenius equation (Equation 3)
[kp] = Ar[exp(–Ea/RT)] (3) where kp is the maximum rate of nucleotide incorporation (or kobs at a saturating
concentration of dNTP was substituted for kp), Ar is the proportionality constant, Ea is the activation energy, R is the universal gas constant, and T is the reaction temperature in Kelvin.
13.3 Results, Discussion, and Future Studies
Kinetic probe #1: the alpha-thio elemental effect To begin probing the rate-limiting step of nucleotide incorporation catalyzed by Pol λ, we examined the effect of an α-substituted phosphorothioate dNTP analog on the observed rate constant. Under single-turnover conditions, Pol λ (300 nM) and D-T DNA (30 nM, Figure 13.1) was mixed with 100 μM dATP or Sp-dATPαS in independent reaction mixtures. Fitting the curves in Figure 13.1 part (A) to a single-exponential equation -1 -1 (Equation 1 in Section 13.2) yielded kobs values of 2.6 ± 0.1 s and 0.58 ± 0.04 s for dATP and Sp-dATPαS, respectively, for a calculated elemental effect, defined as the
ratio of kobs,dNTP/kobs,Sp-dNTPαS, of 4.5. This value is highly similar to what has been measured for rat Pol β (4.3-5) with the same dT:dATP base pair [148, 474], thereby suggesting a common rate-limiting step for correct dNTP incorporation catalyzed by Pol λ and Pol β.
Next, we performed an analogous experiment as described above to investigate the
elemental effect of a misincorporation by using 100 μM dTTP or Sp-dTTPαS. The kobs values were 0.0031 ± 0.0001 s-1 and 0.00017 ± 0.00001 s-1 for dTTP and Sp-dTTPαS, respectively, which equates to an elemental effect of 18 (Figure 13.1 part (B)). For rat Pol β, an elemental effect of 9 was obtained for the dT:dGTP base pair [474]. Based on the
300 work of Herschlag et al., an elemental effect of 4 to 11 has been considered evidence of a rate-limiting chemistry step [50]. Thus, these elemental effects of 4.3 to 18 suggest the chemistry step is at least partially rate-limiting for both correct and incorrect nucleotide incorporations catalyzed by Pol λ and Pol β. However, the alpha-thio effect has been recognized as an unreliable mechanistic probe for determining whether Step 5 is rate- limiting (Chapter 3) [54], therefore, we continued our investigation of the rate-limiting step by performing a more convincing assay: pulse-chase and pulse-quench.
Kinetic probe #2: the pulse-chase and pulse-quench experiment The pulse-chase and pulse-quench assay can provide evidence for the existence of a conformational change (Steps 3 and/or Step 4) preceding the chemistry step (Step 5). The presence of a conformationally-distinct nucleotide-bound complex (e.g. E′•DNA•dNTP) is distinguished from the ground-state ternary complex (E•DNA•dNTP) by the presence of additional product formation in the pulse-chase reaction compared to the pulse-quench reaction. This assay was performed by mixing a pre-incubated solution of Pol λ or Pol β 32 (1 μM) and unlabeled D-T DNA (100 nM) with 10 μM [α- P]dATP (Section 13.2). For the pulse-quench reaction, the reaction was immediately quenched with 1 M HCl, denatured with chloroform, and neutralized with 1 M NaOH. For the pulse-chase reaction, the reaction was chased with an excess of unlabeled dATP (1 mM) for 30 s prior to the aforementioned quenching, denaturing, and neutralization steps. A single- exponential equation (Equation 1 in Section 13.2) was applied to the plots of product concentration versus time for Pol λ (Figure 13.2 part (A)) and Pol β (Figure 13.2 part (B)). For Pol λ, the reaction amplitudes of the pulse-chase and pulse-quench reactions were 36 ± 1 nM and 11 ± 1 nM, respectively, for an amplitude difference of 25 nM [475]. For Pol β, an amplitude difference of 12 nM was calculated from the pulse-chase and pulse-quench amplitudes of 62 ± 3 nM and 50 ± 2 nM, respectively. These marked differences between the pulse-chase and pulse-quench amplitudes indicated that (i) the polymerization pathways of both Pol λ and Pol β proceed with a ternary intermediate that is conformationally-distinct from the ground state ternary complex and (ii) a
301 conformational change (Steps 3 and/or Step 4) before the chemistry step (Step 5) is slower than the nucleotidyl transfer reaction (Scheme 13.1).
The pulse-chase and pulse-quench results suggested Steps 3 and/or 4 are limiting the rate of a correct nucleotide incorporation. However, based on the stopped-flow findings using rat Pol β and a DNA substrate with a site-specific 2-aminopurine base, Step 3 can be excluded as a putative rate-limiting step because the open-to-closed conformational change is approximately 10-fold faster than the rate of nucleotide incorporation measured using radioactive rapid chemical-quench flow technique [58]. Employing a similar strategy for Pol λ generated the same conclusion: a pre-chemistry conformational change does not limit correct nucleotide incorporation (Cuiling Xu and Suo, unpublished data). However, more subtle motions, such as the repositioning of select active site residues, are likely undetectable using stopped-flow fluorescence techniques, therefore, Step 4 remains a candidate for being the rate-limiting event during nucleotide incorporation.
Kinetic probe #3: the Arrhenius activation energy barrier To provide additional evidence about the rate-limiting step of nucleotide incorporation, we measured the Arrhenius activation energy barrier (Ea) for Pol λ and Pol β. The Ea is the minimum amount of energy required to convert reactants into products during
catalysis. The Ea was measured for Pol λ and Pol β incorporating correct (dATP) and incorrect (dCTP, dGTP, and dTTP) nucleotides opposite template dT. To determine the
Ea, we measured the maximum rate of nucleotide incorporation (kp) and the equilibrium
dissociation constant of an incoming nucleotide (Kd) under single-turnover conditions at 17, 21, 25, 29, 33, and 37 °C. To perform this assay (Section 13.2), a pre-incubated solution of Pol λ (300 nM) and 5′-[32P]-labeled D-T DNA (30 nM, Table 13.1) was reacted with increasing concentrations of dATP (0.2-10 µM) in optimized reaction buffer L at 25 °C. Reactions were quenched using EDTA, substrate and products were resolved using polyacrylamide gel electrophoresis, and quantitated using ImageQuant software. A plot of product formation versus time was fit to a single-exponential equation to determine the observed rate constant (kobs) (Figure 13.3 part (A)). The extracted kobs
302 values were plotted versus the corresponding concentration of dATP (Figure 13.3 part (B)). These data were fit to a hyperbolic equation which resolved the pre-steady state -1 kinetic parameters: a kp of 1.13 ± 0.04 s and a Kd of 1.0 ± 0.1 μM. The pre-steady state kinetic parameters for Pol λ incorporating dATP into single-nucleotide gapped D-T DNA
at 17, 21, 25, 29, 33, and 37 °C are presented in Table 13.2. Next, the natural log of the kp was plotted as a function of temperature (K-1) to generate the Arrhenius plot in Figure
13.4 part (A) and to resolve an Ea of 19 ± 2 kcal/mol using a linear equation (Equation 3 in Section 13.2). Similar single-turnover kinetic methodology was used to measure the maximum rate of nucleotide incorporation for correct and incorrect nucleotide incorporations catalyzed by Pol λ and Pol β at different reaction temperatures (Tables
13.2 and 13.3). Then, those kp values were subsequently used to determine the Ea (Table 13.4).
The Ea determined for Pol λ and Pol β had different values and trends for both correct and
incorrect nucleotide incorporations into single-nucleotide gap DNA. The Ea for Pol β was about 6 kcal/mol greater than the Ea for Pol λ during correct dATP insertion. When compared to a correct incorporation, the Ea values for a misincorporation increased 8-11
kcal/mol for Pol β but decreased 1-6 kcal/mol for Pol λ. The Ea values for correct dATP and incorrect dCTP were strikingly similar for Pol λ whereas the other misincorporations were 4-6 kcal/mol less than dATP. Pol λ has a strong propensity to generate -1 deletions [240] via strand misalignment [212, 239]. Since the first downstream template base is dG (Table 13.1), it is possible that dCTP may be forming a correct base pair through a
slipped DNA intermediate. Therefore, we estimated the Ea for the other three correct
dNTPs by measuring the kobs at a saturating dNTP concentration of 50 μM, which
approximates the kp, at six different temperatures (Table 13.5). Interestingly, the Ea depended on the identity of the Watson-Crick base pair: ~19-20 kcal/mol for dA:dT and
~23-24 kcal/mol for dC:dG. Thus, the Ea for an aligned dCTP:dG base pair was ~5 kcal/mol greater than a misaligned dCTP:dG (downstream) or aligned dCTP:dT base pair which is more similar to the dTTP:dT and dGTP:dT mismatches than the correct dATP:dT base pair (Tables 13.4 and 13.5).
303 Effect of Pol λ’s N-terminal domains on Ea Previously, it has been shown that the non-enzymatic N-terminal domains, especially the proline-rich domain, of Pol λ affect the catalytic functions of the C-terminal domains by suppressing polymerase activity [233], restricting strand displacement DNA synthesis [174], and altering polymerase fidelity on gap DNA of various gap widths [117] (Chapter
6). Since Pol λ and Pol β have different Ea values, we wondered whether the N-terminal
domains were responsible. To determine if the N-terminal domains influence the Ea, we estimated the Ea values for tPol λ and dPol λ (Figure 13.5) by measuring the kobs at a saturating dNTP concentration of 50 μM (data not shown) at six different temperatures
(Table 13.4). The estimated Ea values were 29 ± 1 and 14 ± 2 for tPol λ and dPol λ,
respectively. tPol λ catalyzed a correct dATP incorporation with an Ea more like Pol β whereas dPol λ had a smaller Ea like Pol λ. Thus, these preliminary results suggested that the proline-rich region contributes to Pol λ’s lower Ea. More interestingly, this finding also suggested that long-range protein motions alter the activation energy barrier. Crystal structures have been solved for only a truncated form of Pol λ, therefore, it is unknown how the structure of the N-terminal domains affects polymerase activity. Thus, additional kinetic and structural studies will need to be performed to better understand this finding. Nonetheless, the preliminary findings suggest the mechanism of correct and incorrect nucleotide incorporation is different for Pol λ and Pol β.
Comparison of Ea values for polymerase families B, X, and Y
Similar assays were performed to determine the Ea values for correct dTTP:dA and incorrect dNTP:dA base pairs catalyzed by Sulfolobus solfataricus DNA polymerase B1 (PolB1 exo-, Chapter 3), a B-family member, and DNA polymerase IV (Dpo4), a Y-
family member [60]. Compared to X-family Pols λ and β, the Ea values are different,
although, some of the trends are similar. For example, PolB1 exo- has Ea values of 38 and
55 kcal/mol for correct dTTP:dA and incorrect dATP:dA, respectively. A higher Ea for a misincorporation than the correct incorporation is a trend similar to Pol β. However,
Dpo4 has Ea values of 32.9 and 24.2 kcal/mol for correct dTTP:dA and incorrect
dGTP:dA, respectively. A higher Ea for a correct incorporation than an incorrect
304 incorporation is a trend similar to Pol λ. Crystallographic studies have shown that Pol λ and Dpo4 do not undergo a major open-to-closed conformational change from binary to ternary complex formation [46, 55]. In contrast, Pol β and replicative polymerases like PolB1 do experience a dramatic movement of the fingers subdomain [45, 119]. Furthermore, the few available crystal structures of mispairs in a polymerase active site indicate the ternary complex exists in a partially closed intermediate state [209]. Thus, conformational changes may contribute to the different Ea values and trends among the polymerase families.
Comparison of experimental and theoretical Ea values Computational studies have simulated uncatalyzed phosphodiester formation in aqueous solution based on an associative mechanism and determined the rate-limiting microscopic
step of the chemical reaction to be cleavage of the phosphate bond which has an Ea of
33.7 kcal/mol [152]. According to Pauling’s transition state theory, this Ea of ~34 kcal/mol should be lower when the chemical reaction occurs in the polymerase active site [153]. Indeed, applying similar computational methods to a dGTP:dC base pair resolved
from a ternary crystal structure of T7 DNA polymerase generated an Ea of 12.3 kcal/mol which represents cleavage of the phosphate bond during the chemistry step [152]. Computer simulations on Pol β indicated that the rate-limiting microscopic step of the nucleotidyl transfer reaction is the initial deprotonation of the 3′-hydroxyl of the primer strand. This step, which involves the proton being transferred to a water molecule before migrating to Asp, has Ea of at least 17 and 21 kcal/mol for dCTP:dG and dATP:dG base pairs, respectively [154]. Using a truncated form of Pol λ with magnesium as the active site metal ion, the potential energy barrier of a correct incorporation is estimated to be 17.6 kcal/mol, whereby the rate-limiting event is proton transfer from the primer 3′- hydroxyl to Asp [476]. For Dpo4, the free energy has been determined to be 13 kcal/mol for a dCTP:dG base pair, and the rate-limiting microscopic step was deprotonation of the 3′-hydroxyl [477]. Unfortunately, the studies on Dpo4 and truncated Pol λ did not determine an energy barrier for a mispair.
305 In general, the theoretical values for the chemistry step are less than the experimental Ea values for a correct nucleotide incorporation catalyzed by tPol λ, Pol β, Dpo4, and PolB1 exo- (Table 13.4). [Please note, the Ea for PolB1 (38 kcal/mol) is being compared with the computed Ea of T7 DNA polymerase (12.3 kcal/mol), a replicative enzyme. Also, the
Ea of tPol λ (29 kcal/mol), rather than Pol λ (18 kcal/mol), is being compared with the
computed Ea (17.6 kcal/mol) since the simulations use a truncated form of Pol λ.] Together, the experimental and theoretical values suggest the chemistry step is not rate limiting for a correct incorporation. Local active site rearrangements (Step 4) are likely rate-limiting as they are critical for substrate alignment. Once aligned, the faster chemistry step will occur. In regards to a misincorporation, there is only a theoretical value for Pol β which is 21 kcal/mol [154]. As illustrated by a correct base pair, the experimental values (33-36 kcal/mol) are larger than the theoretical value (21 kcal/mol), thereby suggesting Step 4 is rate-limiting. However, additional studies are needed to
verify this preliminary finding. It is surprising how relatively large the experimental Ea values are compared to the theoretical values for most of the polymerases. Some of the Ea values (e.g. 38 kcal/mol for PolB1 exo-) are larger than the value predicted for uncatalyzed phosphodiester bond formation in solution (33.7 kcal/mol). One possible explanation may be the structure of the transition state, for the pentacovalent intermediate may proceed via a completely associative, semi-associative, or completely dissociative mechanism. Computer simulations use a completely associative mechanism, although, some structural and kinetic studies have indicated the mechanism may be partially associative [50, 478, 479].
Please note, most of these molecular dynamic studies are completed using quantum mechanical/molecular mechanical methods. Currently, only a small subset of atoms (~60- 90) remain unconstrained during the computer simulations. Meanwhile, the remainder of the atoms, especially those at long range, are in a fixed position. Moreover, the starting reactants of a crystal structure may require the 3′-hydroxyl group of the primer strand or other hydrogen atoms to be added. Such modeling often requires the crystal-structure atoms to be repositioned. Altogether, these alterations likely alter the final energy
306 barriers. Based on the different experimental Ea values obtained for Pol λ and tPol λ (Table 13.4), these results suggest it is important for the computer simulations to consider the atoms of the entire enzyme, not only the atoms directly involved in the chemical reaction. Unfortunately, this is not feasible using current computer technology and chemical theory.
Future studies Additional kinetic, structural, and computational experiments need to be performed in order to have a comprehensive understanding of the rate-limiting step in the first turnover
of correct and incorrect nucleotide incorporation. To complete the Ea measurements, the kp values need to be determined at various reaction temperatures for tPol λ incorporating correct and incorrect nucleotides into D-T DNA. As recently reported for Dpo4 [62], the
Ea of conformational changes induced by nucleotide binding in Pol β, Pol λ, and tPol λ would be useful to compare to those measured herein (Table 13.4) and those generated from computer simulations. Crystal structures provide the structural basis for polymerase fidelity by showing polymerase-substrate and substrate-substrate interactions. In addition, crystal structures establish a molecular framework for the computational studies to
predict the Ea of the chemistry step. Therefore, ternary complexes need to be solved for PolB1 exo-•DNA•correct/incorrect dNTP and full-length Pol λ•gapped DNA•correct/incorrect dNTP. Crystal structures are available for Dpo4•DNA•incorrect dGTP and various truncated forms of Pol λ with gapped DNA and misaligned intermediates with correct and incorrect incoming dNTPs [239, 303], therefore, it may be possible to begin computational studies using the aforementioned crystal structures.
Enzyme fidelity as a function of temperature
Using the kinetic parameters obtained for determining the Ea, fidelity can be calculated (Tables 13.2 and 13.3) and plotted as a function of temperature for Pol λ and Pol β (Figure 13.6). In general, the fidelity remained relatively constant for each misincorporation examined over a 20 °C change in reaction temperature (i.e. 17-37 °C). However, both Pol λ and Pol β exhibited a modest 5-fold increase in fidelity as the
307 temperature decreased. A similar trend has been observed over a temperature range of 19 to 50 °C for PolB1 exo- (Figure 2.4) and a temperature range of 26 to 56 °C for Dpo4 [60].
308 13.4 Tables
Table 13.1 Sequences of the D-DNA substrates.a D-T 5’-CGCAGCCGTCCAACCAACTCA CGTCGATCCAATGCCGTCC-3’ 3’-GCGTCGGCAGGTTGGTTGAGTTGCAGCTAGGTTACGGCAGG-5’ D-A 5’-CGCAGCCGTCCAACCAACTCA CGTCGATCCAATGCCGTCC-3’ 3’-GCGTCGGCAGGTTGGTTGAGTAGCAGCTAGGTTACGGCAGG-5’ D-G 5’-CGCAGCCGTCCAACCAACTCA CGTCGATCCAATGCCGTCC-3’ 3’-GCGTCGGCAGGTTGGTTGAGTGGCAGCTAGGTTACGGCAGG-5’ D-C 5’-CGCAGCCGTCCAACCAACTCA CGTCGATCCAATGCCGTCC-3’ 3’-GCGTCGGCAGGTTGGTTGAGTCGCAGCTAGGTTACGGCAGG-5’ aEach DNA substrate is composed of a 5′-radiolabeled 21-mer, a 5′-phosphorylated 19-mer, and a 41-mer template which has the unique template bases in bold.
309
Table 13.2 Kinetic parameters for nucleotide incorporation into D-T DNA catalyzed by Pol λ at varying reaction temperatures. k K k /K dNTP p d p d Fidelitya (s-1) (μM) (μM-1s-1) 17 oC dATP 0.34 ± 0.01 0.53 ± 0.08 0.64 dCTP 0.00399 ± 0.00006 0.83 ± 0.08 4.8 × 10-3 7.4 × 10-3 dGTP 0.0106 ± 0.0006 5 ± 1 2.1 × 10-3 3.3 × 10-3 dTTP 0.00063 ± 0.00002 6.2 ± 0.5 1.0 × 10-4 1.6 × 10-4 21 oC dATP 0.61 ± 0.01 0.58 ± 0.05 1.1 dCTP 0.0065 ± 0.0002 0.74 ± 0.08 8.8 × 10-3 8.3 × 10-3 dGTP 0.0151 ± 0.0004 2.9 ± 0.2 5.2 × 10-3 4.9 × 10-3 dTTP 0.00088 ± 0.00002 2.9 ± 0.3 3.0 × 10-4 2.9 × 10-4 25 oC dATP 1.13 ± 0.04 1.0 ± 0.1 1.1 dCTP 0.0112 ± 0.0002 0.65 ± 0.05 1.7 × 10-2 1.5 × 10-2 dGTP 0.0248 ± 0.0007 1.9 ± 0.2 1.3 × 10-2 1.1 × 10-2 dTTP 0.00151 ± 0.00004 2.5 ± 0.3 6.0 × 10-4 5.3 × 10-4 29 oC dATP 1.65 ± 0.03 0.72 ± 0.06 2.3 dCTP 0.0179 ± 0.0002 0.43 ± 0.05 4.2 × 10-2 1.8 × 10-2 dGTP 0.035 ± 0.003 2.4 ± 1.0 1.5 × 10-2 6.3 × 10-3 dTTP 0.00220 ± 0.00004 1.7 ± 0.2 1.3 × 10-3 5.6 × 10-4 33 oC dATP 2.38 ± 0.09 1.1 ± 0.2 2.2 dCTP 0.0234 ± 0.0006 0.6 ± 0.1 3.9 × 10-2 1.8 × 10-2 dGTP 0.048 ± 0.001 1.4 ± 0.2 3.4 × 10-2 1.6 × 10-2 dTTP 0.00233 ± 0.00006 1.2 ± 0.2 1.9 × 10-3 9.0 × 10-4 37 oC dATP 2.68 ± 0.08 1.0 ± 0.1 2.7 dCTP 0.028 ± 0.001 0.7 ± 0.2 4.0 × 10-2 1.5 × 10-2 dGTP 0.053 ± 0.002 1.3 ± 0.2 4.1 × 10-2 1.5 × 10-2 dTTP 0.0026 ± 0.0002 1.6 ± 0.4 1.6 × 10-3 6.1 × 10-4 a Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
310
Table 13.3 Kinetic parameters for nucleotide incorporation into D-T DNA catalyzed by Pol β at varying reaction temperatures. k K k /K dNTP p d p d Fidelitya (s-1) (μM) (μM-1s-1) 17 oC dATP 2.08 ± 0.07 6.3 ± 0.8 0.33 dCTP 0.0207 ± 0.0007 220 ± 30 9.4 × 10-5 2.9 × 10-4 dGTP 0.018 ± 0.001 270 ± 60 6.7 × 10-5 2.0 × 10-4 dTTP 0.0026 ± 0.0001 650 ± 40 4.0 × 10-6 1.2 × 10-5 21 oC dATP 3.43 ± 0.07 5.3 ± 0.5 0.65 dCTP 0.0575 ± 0.0008 200 ± 10 2.9 × 10-4 4.4 × 10-4 dGTP 0.064 ± 0.006 330 ± 100 1.9 × 10-4 3.0 × 10-4 dTTP 0.011 ± 0.001 1100 ± 200 1.0 × 10-5 1.6 × 10-5 25 oC dATP 7.5 ± 0.3 3.6 ± 0.5 2.1 dCTP 0.18 ± 0.01 240 ± 50 7.5 × 10-4 3.6 × 10-4 dGTP 0.146 ± 0.008 320 ± 50 4.6 × 10-4 2.2 × 10-4 dTTP 0.0132 ± 0.0004 370 ± 30 3.6 × 10-5 1.7 × 10-5 29 oC dATP 11.7 ± 0.1 4.7 ± 0.2 2.5 dCTP 0.25 ± 0.02 370 ± 60 6.8 × 10-4 2.7 × 10-4 dGTP 0.308 ± 0.009 270 ± 20 1.1 × 10-3 4.6 × 10-4 dTTP 0.043 ± 0.005 500 ± 100 8.6 × 10-5 3.5 × 10-5 33 oC dATP 20.8 ± 0.4 4.6 ± 0.3 4.5 dCTP 0.61 ± 0.05 390 ± 80 1.6 × 10-3 3.5 × 10-4 dGTP 0.47 ± 0.02 220 ± 40 2.1 × 10-3 4.7 × 10-4 dTTP 0.071 ± 0.004 480 ± 70 1.5 × 10-4 3.3 × 10-5 37 oC dATP 32 ± 1 9 ± 1 3.6 dCTP 0.77 ± 0.05 250 ± 40 3.1 × 10-3 8.7 × 10-4 dGTP 1.29 ± 0.08 340 ± 50 3.8 × 10-3 1.1 × 10-3 dTTP 0.13 ± 0.03 500 ± 300 2.6 × 10-4 7.3 × 10-5 a Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect].
311
Table 13.4 Arrhenius activation energy barriers (kcal/mol) for nucleotide incorporation opposite template dT. Pol λ Pol β tPol λa dPol λa dATP 19 ± 2 24.9 ± 0.9 29 ± 1 14 ± 2 dCTP 18 ± 1 33 ± 3 ND ND dGTP 15 ± 1 36 ± 2 ND ND dTTP 13 ± 2 34 ± 3 ND ND a based on kobs values measured at a saturating dNTP concentration of 50 μM for six different temperatures (17, 21, 25, 29, 33, and 37 °C).
312
Table 13.5 Arrhenius activation energy barriers (kcal/mol) for correct nucleotide incorporations catalyzed by Pol λ.
Base pair Ea (kcal/mol) dATP:dT 19 ± 2 dTTP:dAa 20 ± 1 dCTP:dGa 23.3 ± 0.7 dGTP:dCa 24 ± 1 a based on kobs values measured at a saturating dNTP concentration of 50 μM for six different temperatures (17, 21, 25, 29, 33, and 37 °C).
313 13.5 Figures
A B
30 30
25 25
20 20
15 15
10 10 Product (nM) Product Product (nM) Product
5 5
0 0 0 5 10 15 20 25 30 0 3000 6000 9000 1.2x104 1.5x104 Time (s) Time (s)
Figure 13.1 Alpha-thiol elemental effect. Pol λ (300 nM) and 5′-[32P]-labeled D-T DNA (30 nM) was rapidly mixed with (A) 100 µM dATP (z) or Sp-dATPαS ({) in parallel time courses. The data were fit to a single-exponential equation (Equation 1 in Section -1 -1 13.2) to extract kobs values of 2.6 ± 0.1 s and 0.58 ± 0.04 s for dATP and Sp-dATPαS, respectively, for a calculated elemental effect of 4.5. (B) The same Pol λ•D-T solution was mixed with 100 µM dTTP (z) or Sp-dTTPαS ({) in parallel time courses. The data were fit to a single-exponential equation (Equation 1 in Section 13.2) to extract kobs values of 0.0031 ± 0.0001 s-1 and 0.00017 ± 0.00001 s-1 for dTTP and Sp-dTTPαS, respectively, for a calculated elemental effect of 18.
314 A B
50 70
60 40 50 30 40
20 30
Product (nM) Product (nM) Product 20 10 10
0 0 01234567 00.511.522.53 Time (s) Time (s)
Figure 13.2 Pulse-chase and pulse-quench experiments. A pre-incubated solution of (A) Pol λ or (B) Pol β (1 μM) and unlabeled D-T DNA (100 nM) was rapidly mixed with 10 µM [α-32P]dATP. The pulse-quench ({) reactions were immediately quenched with HCl (1 M) while the pulse-chase (z) reactions were chased with 1.0 mM of non-radioactive dATP for 30 s prior to acid quenching. Fitting the data to equation 1 (Section 13.2) yielded respective pulse-chase and pulse-quench reaction amplitudes of 36 ± 1 nM and 11 ± 1 nM for Pol λ and 62 ± 3 nM and 50 ± 2 nM for Pol β. Plot (A) was obtained from reference [475].
315 A B
30 1.2
25 1
20 0.8 ) -1
15 (s 0.6 obs k 10 0.4 Product (nM) Product
5 0.2
0 0 0 5 10 15 20 0246810 Time (s) dATP (μM)
Figure 13.3 Concentration dependence on the pre-steady state rate constant of nucleotide incorporation. (A) A pre-incubated solution of Pol λ (300 nM) and 5′-[32P]-labeled D-T DNA (30 nM) was rapidly mixed with increasing concentrations of dATP•Mg2+ (0.2 μM, z; 0.5 μM, {; 1 μM, ; 2 μM, ; 5 μM, S; 10 μM, U) for various time intervals at 25 °C. The solid lines are the best fits to a single-exponential equation (Equation 1 in Section 13.2) which determined the observed rate constants, kobs. (B) The kobs values were plotted as a function of dATP concentration. The data (z) were then fit to a hyperbolic -1 equation (Equation 2 in Section 13.2), yielding a kp of 1.13 ± 0.04 s and a Kd of 1.0 ± 0.1 μM.
316 A B 2 4
0 2 ) )
-1 -2 -1 0 ) (s ) (s p p k k -4 -2 ln( ln(
-6 -4
-8 -6 3.2 3.25 3.3 3.35 3.4 3.45 3.2 3.25 3.3 3.35 3.4 3.45 -1 -1 1000/T (K ) 1000/T (K )
Figure 13.4 Arrhenius activation energy barriers. The kp values for nucleotide incorporation [dATP (z), dCTP ({), dGTP (), and dTTP ( )] into 5′-[32P]-labeled D- T DNA were plotted as a function of reaction temperature and subsequently fit to equation 3 (Section 13.2). The activation energy barriers extracted from these plots are presented in Table 13.4 for (A) Pol λ and (B) Pol β.
317
Figure 13.5 Domains of Pol λ, dPol λ, tPol λ, and Pol β. Each domain is labeled in the rectangular box and the residue numbers are noted above. N-terminal residues 1-35 of Pol λ contain a nuclear localization sequence.
318 A B 10-1 10-2
10-2 10-3 Fidelity Fidelity 10-3 10-4
10-4 10-5 15 20 25 30 35 40 15 20 25 30 35 40 o o Temperature ( C) Temperature ( C)
Figure 13.6 Fidelity as a function of temperature. The base substitution fidelity of each misincorporation [dCTP ({), dGTP (), and dTTP ( )] opposite template dT is shown for (A) Pol λ and (B) Pol β. The fidelity values were extracted from Tables 13.2 and 13.3 for Pol λ and Pol β, respectively.
319 13.6 Scheme
Scheme 13.1
320
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