Characterization of G-quadruplexes in DNA repair protein gene sequences and the effects of oxidized guanine lesions on DNA base insertion and elongation

Guanine (G) is one of the four nucleo bases that make up the complex macromolecule deoxyribonucleic acid (DNA). In addition to its ability to Watson-Crick base pair with cytidine (C), guanine can base pair with itself vianon-canonical, Hoogsteen base pairing and form a secondary structure of DNA called a G-quadruplex (G4). Addressed in chapter 1, these structures, composed of four contiguous runs of at least three guanines each, are typically found in the telomeric sequence of chromosomes, but can be found in other DNA sequences and have also been implicated in transcriptional regulation of oncogenes. Using circular dichroism, thermal-melting analysis, and thioflavin T fluorescence, 8 possible G4s were identified and characterized in DNA sequences associated with base excision repair proteins (BER). The presence of G4s in these sequences can affect the transcription of these proteins. Guanine is also easily oxidized to a number of products. These products can cause base transversions and thus induce mutations that can drastically affect an organism. In chapter 2, the effect of oxidized guanine lesions was determined by examining which DNA bases each lesion inserted opposite, as well as the percent elongation past the lesion when located in a DNA strands. The two research directions intersect where BER proteins have the role of repairing mutations in DNA, such as those that result from oxidized guanine lesions.

CHARACTERIZATION OF G-QUADRUPLEXES IN DNA REPAIR PROTEIN GENE SEQUENCES AND THE EFFECTS OF OXIDIZED GUANINE LESIONS ON DNA BASE INSERTION AND ELONGATION by Joshua Visser A Senior Honors Thesis Submitted to the Faculty of The University of Utah In Partial Fulfillment of the Requirements for the Honors Degree in Bachelor of Science In Chemistry Department Approved: ______________________________ Cynthia J. Burrows, PhD Thesis Faculty Supervisor _____________________________ Cynthia J. Burrows, PhD Chair, Department of Chemistry _______________________________ Thomas G. Richmond, PhD Honors Faculty Advisor _____________________________ Sylvia D. Torti, PhD Dean, Honors College May 2016 Copyright © 2016 All Rights Reserved ABSTRACT Guanine (G) is one of the four nucleobases that make up the complex macromolecule deoxyribonucleic acid (DNA). In addition to its ability to Watson-Crick base pair with cytidine (C), guanine can base pair with itself via non-canonical, Hoogsteen base pairing and form a secondary structure of DNA called a G-quadruplex (G4). Addressed in chapter 1, these structures, composed of four contiguous runs of at least three guanines each, are typically found in the telomeric sequence of chromosomes, but can be found in other DNA sequences and have also been implicated in transcriptional regulation of oncogenes. Using circular dichroism, thermal-melting analysis, and thioflavin T fluorescence, 8 possible G4s were identified and characterized in DNA sequences associated with base excision repair proteins (BER). The presence of G4s in these sequences can affect the transcription of these proteins. Guanine is also easily oxidized to a number of products. These products can cause base transversions and thus induce mutations that can drastically affect an organism. In chapter 2, the effect of oxidized guanine lesions was determined by examining which DNA bases each lesion inserted opposite, as well as the percent elongation past the lesion when located in a DNA strands. The two research directions intersect where BER proteins have the role of repairing mutations in DNA, such as those that result from oxidized guanine lesions. ii TABLE OF CONTENTS ABSTRACT ii CHAPTER 1 1 INTRODUCTION 1 METHODS 4 RESULTS AND DISCUSSION 6 REFERENCES 11 CHAPTER 2 14 INTRODUCTION 14 METHODS 17 RESULTS AND DISCUSSION 19 REFERENCES 25 iii CHAPTER 1: Identification and characterization of G-quadruplexes in DNA repair protein gene sequences INTRODUCTION Within the primary sequence of the human genome are motifs central for directing cellular pathways. Gene promoter sequences are recognized by transcription factors for regulating transcription; chemical modification of 5’-CpG-3’ islands occurs for gene regulation (i.e., epigenetics); and certain sequence motifs can adopt alternative-secondary structures for guiding cellular processes1,2. An example of one of these secondary structures is the G-quadruplex (G4), which is composed of consecutive runs of guanine (G) nucleotides with intervening sections of a few nucleotides; and which folds around monovalent ions like potassium (K+) 3-6. This folding pattern involves Hoogsteen base pairing, hydrogen bonding using non-traditional base pair partners beyond the traditional Watson-Crick, between Gs in polymers called tetrads. Three tetrads are formed with twelve Gs with metal ions between each set of tetrads. The folding pattern, and more specifically looping, can cause the G4 to adopt a number of folding patterns that affect its optical properties and stability. It has been a mystery as to if G4s actually fold within human cells and what roles these structures have. It has been determined by immunoflouresence with a G4-binding antibody that these structures are found in human chromatin; with the maximal observation occurring during the S phase of the cell cycle7,8. Experiments to determine the role of G4s have come to the conclusion that they are critical to cellular processes and are sites of increased genome instability. Many strategies have been used to find operational G4 sequences in the genome. Prior to the sequencing of the human genome, sequences such as the c-MYC promoter were examined as possible G4s9. Once the human genome was published, more than 370,000 possible G4 sequences were identified using bioinformatics10. The bioinformatics scans searched for the classical G4 sequence motif of 5ʹ-G≥3-N1-7-G≥3-N17-G≥3-N1-7-G≥3-3ʹ. Beyond repeat sequences, such as that of telomeres, further inspection of identified G4s showed preference for promoters, 5ʹ-untranslated regions (5ʹ-UTR), and the 5ʹ end of the first intron11,12. More recently, G4-sequencing developed by the Balasubramanian laboratory using the Illumina platform has been able to identify more than 700,000 possible G4 sequences in the genome13. This platform was able to identify a large number of non-classical G4s with loops exceeding the traditional 7 nucleotides and bulges within the structure. Cell based studies to identify G4s have come back with 1000s of possible sites11,12. Studies focusing on possible regulatory G4s in the 5ʹ- and 3ʹ-UTRs in the coding and template strands of the genome have found that G4s in the coding strand can also be found in the mRNA, and some G4s in the 5ʹ-UTRs of mRNA have regulatory features14-16. The highest and most well-known occurrence of G4s has been in the 5ʹTTAGGG-3ʹ repeat sequence of the telomere of chromosomes, for which a structural purpose has been hypothesized17,18. Two roles of G4s in replication have been documented; the first of these was the observation of a G4-consensus sequence at human origins of replication19, and the second was an observed stalling of polymerase bypass by G4s that led to genome instability20. Indeed, the location of a G4 can effect transcription in two different ways; the first is if the structure is in the coding strand it speeds up transcription by making the DNA single stranded without the work of a helicase and letting the polymerase progress through faster, second is if the structure is in the template 2 strand, as stated above, it can stall the polymerase or even stop it which prevents transcription. Evolution has selected a number of G4-specific helicases (BLM, XPD, XPB) to ensure smooth replication through these knotted structures3; however mutations rendering these helicases dysfunctional lead to a number of catastrophic diseases21. For this reason, the pathogenesis of a number of diseases is attributed to expansion of repeat sequences that adopt G4 folds in DNA and RNA22. Perhaps the most important roles of G4s as transcription-regulatory elements, and the most studied, has been the presence of the structures in the promoters of oncogenes such as c-MYC23, VEGF-A24, hTERT25, KRAS26, HIF-1α27, PDGFA28, c-KIT29, and c-MYB30. Gene regulation has been documented when the G4 resides in either the coding or template strand of the promoter or 5’-UTR31. Because DNA repair is a complex cellular process that is critical to cellular viability, the use of dynamic, DNA control sequences, such as G4s, may provide finetuning mechanisms for regulation of key steps in these pathways. For this reason, six human DNA repair protein gene sequences were inspected for classical and non-classical G4s in the promoter and 5ʹ-UTR sequences of established transcripts32,33. Two of these genes were inspected in two different contexts, observing the sequence with only four of its G domains and then adding on the 5th domain. It has recently been hypothesized that the 5th run of Gs, which do not constitute the core of the G4, can act as “spare tires” that are a backup if any of the core tracks of Gs is damaged by oxidation, allowing the G4 to persist and facilitating the DNA repair process34. These eight sequences (Table 1) were synthesized and studied by three complementary methods to establish if they adopted G4 3 architectures. These results are discussed with respect to their possibility toward G4 regulation of human DNA repair genes during oxidative or inflammatory stress. METHODS DNA sequences of interest were obtained as single-stranded DNA oligonucleotides (Table 1). These oligomers were synthesized by the DNA Sequencing Core Facility at the University of Utah and purified by HPLC. After purification, the sequences were dried down and each underwent dialysis to rid the samples of purification salts. Concentrations of the samples were determined by UV-vis spectra at the lambda max wavelength of 260 nm. These samples were then used as stocks for all experiments conducted. To establish if the sequences studied folded and the topologies of the potential structures, three complementary methods were utilized. The first of these methods was recording the circular dichroism (CD) spectra for each of the sequences studied. These spectra indicate if a G4 structure is present and whether the dominant fold in solution is parallel stranded (λmax = 260 nm and λmin = 240 nm), antiparallel stranded (λmax ~ 290295 nm and λmin = 250 nm), or a mixture of parallel and antiparallel strands (λmax ~ 265 nm, λmin = 240 nm, and a strong shoulder at 290 nm)(Figure 1A) 35,36. The differences in the spectra are caused by the difference in ellipticity between folding motifs due to the chiral nature of the G4s. Two buffer systems were used, one a potassium buffer mirroring physiological conditions (120 mM KCl, 12 mM NaCl, 20 mM KPi, pH 7.4, 25 °C) and the other a sodium buffer(132 mM NaCl, 20 mM NaPi, pH 7.4, 25 °C). The second experiment determined the Tm values for each sequence, by monitoring the transitions at 295 nm37. This was done by observing the absorbance of the 4 sequence at 295 nm over a temperature range from 25 to 100 °C taking measurements at every degree. The Tm is the midpoint of the curve as it changes its curvature, and is a result of the DNA denaturing from the G4-secondary structure to single-stranded DNA (Figure 2) and vice-versa. Measurements were taken for both the increase (denaturing) in temperature and the subsequent decrease (annealing). Observing both the heating and cooling cycles allows us to see if there is a difference between the quickly forming kinetic folding pattern and the more stable thermodynamic fold that requires more energy to form. Tm values above 60 °C indicated regulation by a G4 – ability to stall polymerase bypass and impact biological processes38. In the third and final study, the emission intensity of the G4-specific fluorophore thioflavin T was measured39. This study was conducted by using the excitation wavelength of 425 nm and observing fluorescence at the emission wavelength of 490 nm. Thioflavin T in its relaxed state is not fluorescent, but when it binds a G4, the fluorescence intensity at 490 nm is significantly enhanced. Furthermore, the fluorescence does not increase in the presence of single- or double-stranded DNA. A positive control consisting of the c-MYC G4 was used along with single- and double-stranded DNA as negative controls, giving the expected fluorescence intensities. G4s were assayed under identical conditions to the controls and if the fluorescent intensity of the G4 was more than twenty-fold greater than that of thioflavin T alone, it was decided that the sequence adopted a G4. These three methods together can be used to claim the presence and identity of a G4 in the sequences studied, and this information can be used in conjunction with cell studies to explain and observe transcription for these sequences. 5 RESULTS AND DISCUSSION In the first study using CD spectrometry, all eight of the sequences studied seemed to adopt a parallel fold in the potassium buffer that models physiological conditions (Figure 1B). In the sodium buffer system, results were similar to that of potassium albeit at a lower intensity for most sequences (Figure 1C). Both conditions reinforce the hypothesis of the presence of G4s. Table 1. G4 sequences characterized from the promoters or 5’-UTR of human DNA repair genes. Gene APE APE 5th MPG NEIL1 NEIL1 5th PCNA 6th Pol η UNG Sequence (tracts of Gs involved in G4 in red) 5'-A GGG CA GGG T GGGGG T GGG T-3' 5'-CA GGG CA GGG TC GGGG T GGG TAGCCCT GGGGG TT-3' 5'-A GGGGG AGCACTC GGGGGG AT GGGGGG C-3' 5'-TTT GGGGGG A GGGGGG CGCA GGG A GGGG CTT-3' 5'-GC GGG TCCTAAGTGT GGGGG A GGGGG CGCA GGG A GGGG CG-3' 5'-CA GGG A GGG A GGG CGAC GGGGG C GGGG C GGGG CG-3' 5'-CT GGGG C GGGG AGA GGG TGTC GGG AC-3' 5'-GT GGGG TC GGGGGG A GGGGG CT GGG AA-3' 6 Role BER Protein BER Protein BER Protein BER Protein BER Protein RNA polymerase protein DNA and translesion polymerase BER Protein A 20 Elipticity (mdeg) Propeller 15 Hybrid 10 Basket 5 0 220 240 -10 2E+09 300 320 [θ] (deg*cm2*dmol-1) APE 1E+09 MPG -1E+09 270 320 Wavelength (nm) UNG APE APE 5th Domain NEIL 1 1E+09 APE 5th Domain NEIL 1 0 C 2E+09 UNG 220 280 Wavelength (nm) B [θ] (deg*cm2*dmol-1) 260 -5 NEIL1 5th Domain PCNA 6th Domain Pol η MPG 0 -1E+09 220 270 Wavelength (nm) 320 NEIL1 5th Domain PCNA 6th Domain Pol η Figure 1. CD Spectra of folding patterns and sequences. (A) Example spectra showing three folding motifs. (B) Spectra of sequences in potassium buffer. (C) Spectra of sequences in sodium buffer. Thermal melting data were used as a second source of evidence of the presence of G4s and showed results that would be expected for sequences containing this secondary structure. For all sequences studied, there was a change in the G4-specific signal intensity at 295 nm, indicating a transition from the folded G4 to single-stranded DNA (Table 2). As shown, there was not a Tm value obtained for NEIL 1 5th in the sodium buffer during cooling, despite repeated trials. It is believed that this discrepancy is due to an additional secondary structure forming. Also of interest were second melting points for APE and PCNA in the potassium buffer at 80.1 °C and greater than 90 °C, respectively, during the heating cycle. These results indicate a second stable structure, possibly another G4, forming before the DNA denatures becomes single stranded. 7 Looking at only the more applicable potassium buffer system, every sequence studied had a melting point above the requisite 60 °C during one of the cycles, indicating the presence of regulatory G4s in these sequences. The large discrepancies between heating and cooling Tm values for APE and PCNA is likely due to the fact that the noted second transitions in heating cycles is more similar to the transition recorded in the cooling cycles. 1.1 Absorbance 1.05 Heating 1 Cooling 0.95 0.9 0.85 0.8 0 20 40 60 80 100 120 Temperature (°C) Figure 2. Sample Tm curve for G4 containing sequence Table 2. Tm values for the increasing and decreasing temperature scans of sequences. Measurements have a standard deviation of 10% due to only one trial being conducted. Sequence Heating Cooling APE 5th, K 68.12 66.48 APE 5th, Na 58.16 52.51 APE, K 51.25 75.27 APE, Na 65.07 53.19 MPG, K 68.94 62.66 MPG, Na 81.07 58.97 Neil 1 5th, K 70.15 63.78 Neil 1 5th, Na 56.99 -Neil 1, K 65.68 63.85 Neil 1, Na 74.77 43.27 PCNA, K 54.14 75.59 PCNA, Na 79.19 62.28 Pol η, K 63.10 61.38 Pol η, Na 48.18 43.52 8 Using the fluorophore thioflavin T in both the potassium and sodium buffer systems, it was determined that almost all sequences contained a G4 based on the twenty fold increase in fluorescence cutoff established in the literature (Figure 3)57. However, in the sodium buffer, a false positive was obtained for MPG. MPG actually does not contain a G4, per se, but a similar secondary structure that contains only two tetrads – a result of having only three runs of Gs – and is similar to the motif of the thrombin binding aptamer, which is known to have low affinity for thioflavin T. Once again, while the sodium buffer had results that resembled that of the more pertinent potassium buffer, these results were at a lower intensity, which allowed for MPG to not pass this assay. A C B D Figure 3. Thioflavin T fluorescence spectra. (A) Raw fluorescence intensity of sequences in potassium buffer. (B) Raw fluorescence intensity of sequences in sodium buffer. (C) Fluorescence at 290 nm and G4 cutoff in potassium buffer. (D) Fluorescence at 290 nm and G4 cutoff in sodium buffer. 9 From the three techniques used, it was determined that all of the sequences studied had a G4 that could potentially actively regulate transcription. From CD studies, it was determined that all of the sequences adopt a parallel fold while Tm and fluorescence studies provided further evidence for the presence of G4s. However, it was shown that the MPG sequence did not actually form a G4, but instead a comparable structure with two tetrads that is similarly adopted by the thrombin binding aptamer. The two sequences of NEIL 1 and APE, which were studied with both four and five domains of G runs, proved to be very informative as to the effect of the “spare tire” 5th domain. The addition of the 5th domain, in both cases, did not have significant effect on the CD or Tm; however, in the fluorimetry assays, there was a significant increase in fluorescence intensity. The results from the first two assays indicate that the addition of the 5th domain did not have an effect on the G4 formed, and therefore the 5th domain was not part of the G4. Contrary to this is the increase in fluorescence intensity that indicates a more stable G4, or more likely a structure with a higher affinity for thioflavin T. All together, these studies suggest that the addition of the “spare tire” did not have an effect on the G4 because they were not part of the G4 formed. Therefore, it is concluded that there is a presence of G4s in the DNA repair protein gene sequences studied, that they adopt parallel folds, and that the addition of a 5th domain did not have an effect on the G4 formed, suggesting the first four domains are those responsible for the quadruplex. 10 REFERENCES 1. Lenhard, B.; Sandelin, A.; Carninci, P. Nat. Rev. Genet., 2012, 13, 233-245. 2. Matos, J.; West, S.C. DNA Repair (Amst), 2014, 19, 176-181. 3. Maizels, N.; Gray, L.T. PLoS Genet., 2013, 9, e1003468. 4. Bugaut, A.; Balasubramanian, S. Nucleic Acids Res., 2012, 40, 4727-4741. 5. Rhodes, D.; Lipps, H.J. Nucleic Acids Res., 2015, 43, 8627-8637. 6. Balasubramanian, S.; Hurley, L.H.; Neidle, S. Nat. Rev. Drug Discov., 2011, 10, 261275. 7. Biffi, G.; Tannahill, D.; McCafferty, J.; Balasubramanian, S. Nat. Chem., 2013, 5, 182-186. 8. Henderson, A.; Wu, Y.; Huang, Y.C.; Chavez, E.A.; Platt, J.; Johnson, F.B.; Brosh, R.M., Jr.; Sen, D.; Lansdorp, P.M. Acids Res., 2014, 42, 860-869. 9. 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Sundquist, W.I.; Klug, A. Nature, 1989, 342, 825-829. 11 19. Besnard, E.; Babled, A.; Lapasset, L.; Milhavet, O.; Parrinello, H.; Dantec, C.; Marin, J.M.; Lemaitre, J.M. Nat. Struct. Mol. Biol., 2012, 19, 837-844. 20. Lopes, J.; Piazza, A.; Bermejo, R.; Kriegsman, B.; Colosio, A.; Teulade-Fichou, M.P.; Foiani, M.; Nicolas, A. EMBO J., 2011, 30, 4033-4046. 21. Maizels, N. EMBO Rep., 2015, 16, 910-922. 22. Simone, R.; Fratta, P.; Neidle, S.; Parkinson, G.N.; Isaacs, A.M. FEBS Lett., 2015, 589, 1653-1668. 23. Brooks, T.A.; Hurley, L.H. Nat. Rev. Cancer, 2009, 9, 849-861. 24. Sun, D.; Liu, W.J.; Guo, K.; Rusche, J.J.; Ebbinghaus, S.; Gokhale, V.; Hurley, L.H. Mol. Cancer Ther., 2008, 7, 880-889. 25. Palumbo, S.L.; Ebbinghaus, S.W.; Hurley, L.H. J. Am. Chem. Soc., 2009, 131, 1087810891. 26. Cogoi, S.; Xodo, L.E. Nucleic Acids Res., 2006, 34, 2536-2549. 27. De Armond, R.; Wood, S.; Sun, D.; Hurley, L.H.; Ebbinghaus, S.W. Biochemistry, 2005, 44, 16341-16350. 28. Qin, Y.; Rezler, E.M.; Gokhale, V.; Sun, D.; Hurley, L.H. Nucleic Acids Res., 2007, 35, 7698-7713. 29. Bejugam, M.; Sewitz, S.; Shirude, P.S.; Rodriguez, R.; Shahid, R.; Balasubramanian, S. J. Am. Chem. Soc., 2007, 129, 12926-12927. 30. Miyazaki, T.; Pan, Y.; Joshi, K.; Purohit, D.; Hu, B.; Demir, H.; Mazumder, S.; Okabe, S.; Yamori, T.; Viapiano, M.; et al. Clin. Cancer Res., 2012, 18, 1268-1280. 31. Bochman, M.L.; Paeschke, K.; Zakian, V.A. Nat. Rev. Genet., 2012, 13, 770-780. 32. Wood, R.D.; Mitchell, M.; Sgouros, J.; Lindahl, T. Science, 2001, 291, 1284-1289. 33. Wood, R.D.; Mitchell, M.; Lindahl, T. Mutat. Res., 2005, 577, 275-283. 34. Fleming, A.M.; Zhou, J.; Wallace, S.S.; Burrows, C.J. (2015) ACS Cent. Sci., 2015, 1, 226-233. 35. Karsisiotis, A.I.; Hessari, N.M.; Novellino, E.; Spada, G.P.; Randazzo, A.; Webba da Silva, M. Angew. Chem., Int. Ed., 2011, 50, 10645-10648. 36. An, N.; Fleming, A.M.; Middleton, E.G.; Burrows, C.J. PNAS, 2014, 111 (40), 14325-14331. 12 37. Mergny, J.L.; Phan, A.T.; Lacroix, L. FEBS Lett, 1998, 435, 74-78. 38. Piazza, A.; Adrian, M.; Samazan, F.; Heddi, B.; Hamon, F.; Serero, A.; Lopes, J.; Teulade-Fichou, M.P.; Phan, A.T.; Nicolas, A. EMBO J., 2015, 34, 1718-1734. 39. Renaud de la Faverie, A.; Guedin, A.; Bedrat, A.; Yatsunyk, L.A.; Mergny, J.L. Nucleic Acids Res., 2014, 42, e65. 13 CHAPTER 2: Effects of oxidized guanine lesions on DNA base insertion and extension INTRODUCTION Damage of DNA bases by oxidation is of great interest due to the implication of this damage with aging, mutagenesis, and carcinogenesis1-3. The nucleotide triphosphate (dNTP) pool as well as DNA itself are subject to damage by agents both interior and exterior to the body, resulting in DNA strand breaks, abasic sites, and oxidized bases that require DNA repair proteins to repair or excise in order to maintain the integrity of the genome2. Guanine, as one of the DNA bases that can be oxidized, is of great interest due to the ease in which it undergoes oxidation. Therefore, guanine (G) and its common, first oxidation product 8-oxoguanosine (OG) have been studied; furthermore, their further oxidation products of spiroiminodihydantoin (Sp) and guanidinohydantoin (Gh), which result from nucleophilic attack of G and OG by H2O after oxidation are also of great interest (Scheme 1)4-14. Scheme 114: Oxidation pathway of G and OG to Gh and Sp It has been shown that Sp is the major product of oxidation occurring in a neutral or slightly basic environment while Gh is preferred in systems that are acidic (pH < 6) or near neutral pH in oligomers5,6,8,15. Both Sp and Gh are derived from the common 14 intermediate 5-OH-OG, but go through different mechanistic steps and rearrangements to give the respective products. The guanine oxidation products – OG, Sp, and Gh – each have a similar effect on DNA, they cause base transversions. Base transversion is a point mutation where a base switches from a purine to a pyrimidine as a result of the base pairing partners of the oxidized lesions. In the case of guanine and its oxidation products, this is a switch from guanine – a purine – to thymine (T) or cytosine (C)– pyrimidines. Whereas the base pair partner of G is C, OG base pairs with both A and C, Gh base pairs with A and G, and Sp base pairs with A and G. An example of a base transversion is the typical G:C base pair becoming OG:A upon oxidation and when the DNA is copied, this base pair becomes T:A, causing a G to T transversion. Although OG is only 5% mutagenic, causing only G to T transversions, Sp and Gh are greater than 98% mutagenic, causing G to C and G to T transversions16-18. For this reason, there is much work focused on these lesions and their incorporation into the genome. The base transversions that result from these oxidized products depend upon the relative distribution of the lesions between base pair partners. For OG, the distribution of insertion of dC and dA opposite OG in the template is 7:1 for Klenow Fragment while for Sp the distribution of dA and dG insertion opposite Sp in the template is nearly 1:1 for both diastereomers for the same polymerase19,20. Different polymerases will give different distributions of base pairs and even the concentration of polymerase and time spent reacting will affect product distribution; therefore, these distributions are subject to change based upon reaction conditions. 15 Current research is directed at understanding both the insertion of oxidized guanine nucleotides as well as extension beyond these lesions when present in a primer strand. The different ways, in which lesions are present in DNA, whether it be in the nucleotide pool – in the case of insertion – or part of DNA being actively copied – in the case of extension – can drastically change base pairing partners. Recent research has stated that despite the typical base pairing partners of A and G for Sp when the Sp is present in the template, Sp will only insert opposite C when Sp comes from the nucleotide pool as a triphosphate (dSpTP)21. The suspicious insertion of dSpTP opposite C may come from dGTP as in impurity because synthesis of dSpTP occurred from oxidation of dGTP. Therefore, this study is directed at replicating this data and then scrupulously purify the dSpTP to remove any contaminate dGTP to demonstrate in addition to testing the differences in base pairing partners for Gh when present as part of the primer and when part of the nucleotide pool. Therefore, it will first be tested which nucleotide each oxidized guanine lesion base pairs with when added to a solution of template and primer, where each template differs in the nucleotide opposite where the guanine lesion will be inserted. Second, the extension of DNA sequences containing oxidized guanine lesions will be tested by adding a dNTP mix containing A, T, C, and G to the same templates, but with primers terminating in an oxidized lesion. It is important to note that this research, in regards to primer extension, contain the oxidized guanine lesion as part of the primer as opposed to the common placement in the template. As a result, the results contained herein may differ from literature values due to the location of the lesion. As a part of the primer, 16 there is no “running start” of the polymerase before it reaches the lesion, making extension beyond a difficult task. METHODS DNA sequences of interest were obtained as single-stranded DNA oligonucleotides (Figure 1). The four template strands, primer, and primer containing OG were synthesized by the DNA Sequencing Core Facility at the University of Utah. The OG-containing primer was first cleaved from the solid support with concentrated NH4OH with 200 mM β-mercaptoethanol at 25 °C over 24 hours. This primer was subjected to oxidative conditions to give the primers containing Gh and the two Sp diastereomers. To get these higher oxidized lesions, 20 μM OG primer was reacted with 240 μM Na2IrCl6 in pH 7 ddH2O to give Gh and a pH buffer containing 20 mM NaPi to give Sp. The oligonucleotides were purified by HPLC. After purification, the sequences were dried down and each underwent dialysis to rid the samples of purification salts. The concentrations of the samples were determined by UV-vis spectra at the lambda max wavelength of 260 nm. These samples were then used as stocks for all experiments conducted. DNA primers were radiolabeled by attaching 32P to the 5’-end of the primer using T4-polynucleotide kinase and radioactive γ-32P-ATP. The dOG triphosphate (dOGTP) used in the insertion assay was obtained from Trilink BioTechnologies. The triphosphate of Sp (dSpTP) was obtained by adding 0.5 mM dGTP to a pH 7 buffer with 10 mM sodium acetate and 50 μM Rose Bengal, a singlet oxygen producer. The reaction was carried out for 30 minutes and the sample was then purified by NAP column to remove the Rose Bengal. The solution was then dried down in vacuo, rehydrated with deionized water, and purified by HPLC using a C18 17 reversed-phase column using 5 mM tetrabutylammonium sulfate, 10 mM NaPi, pH 7.0 (mobile phase A) and acetonitrile (mobile phase B) as the mobile phases. As the gradient, mobile phase B was increased from 5 to 75% over 60 minutes at a flow rate of 1.0 mL/min while monitoring the elution profile at 230 nm. The dSpTP diastereomers were collected separately and dried in vacuo. These samples were brought back up in deionized water and subjected to a second HPLC run to remove the salts. This run consisted of the mobile phases of 20 mM ammonium bicarbonate (mobile phase A) and acetonitrile (mobile phase B) with a gradient of mobile phase B being increased from 1 to 65% over 30 minutes. Samples were once again dried in vacuo and rehydrated with deionized water. The two HPLC purification runs were repeated to ensure absolute purity. To make the triphosphate of Gh (dGhTP), 0.5 mM dGTP was added to a pH 4 buffer containing 10 mM sodium acetate and 50 μM Rose Bengal. The reaction was carried out for 30 minutes, and then purified by NAP column. Sample was dried in vacuo and subjected to HPLC purification technique of dSpTP listed above, but only one cycle. The concentration of the diastereomers of dSpTP and dGhTP were determined by UV-vis at the lambda max wavelength of 240 nm using extinction coefficient values previously determined22. For insertion assays, varying concentrations of oxidized guanine lesions were added to 10 nM of radiolabeled duplex DNA (at a ratio of 1:1.25 primer to template) with Klenow Fragment exo- and incubated for 30 minutes at 37 °C in Klenow reaction buffer (New England Biolabs). The reactions were quenched with 5 μL of termination solution (95% formamide, 0.1% bromophenol, and 0.1% xylene cyanol). These samples were 18 applied to a 20% polyacrylamide gel in the presence of 7M urea, electrophoresed at 45 W for two hours, and then put on a phosphor screen for 18 hours. The screen was imaged by phophorimager autoradiography, and the band intensities quantified using ImageQuant software. Yields were based on the intensity of bands compared to the total intensity of the reaction. Relative incorporation was computed by comparing intensity of band for one template to the total intensity for all incorporation bands. For extension assays, varying concentrations of each dNTP (A, T, C, G) were added to 10 nM of radiolabeled duplex DNA and polymerase (5 units of Klenow Fragment exo- or 2 units of Dpo4) and incubated at 37 °C or 55 °C, respective of polymerase, for 30 minutes. The reaction was quenched with 5 μL of termination solution and applied to a 20% polyacrylamide gel in the presence of 7 M urea, electrophoresed at 70 W for three hours, and then put on a phosphor screen for 18 hours. The screen was imaged as above and extension percentage was based on the intensity of the extended strand relative to the total intensity of the reaction. RESULTS AND DISCUSSION The first direction of this experimentation was a focus on the insertion of the oxidized guanine lesions opposite the templates of interest using the same primer across all templates. Primer : 3’-XAG CCA TAT TGA CAC GAG AGT-5’ Template: 5ʹ-ACT TCT CCT NTC GGT ATA ACT GTG CTC TCA TAG-3ʹ Figure 1. DNA sequences used in both extension and elongation assays where N = A, T, C, G and X = OG, Gh, S-Sp, and R-Sp and is only present for extension assays. Initial studies focused on the insertion of dOGTP as a proof of concept and a test of experimental procedures. At a concentration of 300 nM dOGTP, it was found that, as 19 expected, the dOGTP inserted opposite the template strands containing A and C. Surprisingly, the distribution of products was opposite that present for the insertion of dNTPs opposite OG in the template; here there was a 7:1 ratio of insertion of A:C as opposed to the reverse (Figure 2A)Shibutani. As reaction conditions were altered to include more polymerase and a higher concentration of dOGTP, the distribution began to shift closer to a 2:1 insertion ratio in favor of A (Figure 2B, 2C). A 85% 15% C B 73% 27% A 66% 34% T C G A C A T C G Figure 1. Insertion of dOGTP opposite templates at 10 nM duplex DNA and varied concentration of dOGTP and units of Klenow fragment exo-, percentages showing relative insertion. (A) Gel of 300 nM dOGTP with one unit/μL of enzyme. (B) Gel of 300 nM dOGTP and five units/μL of enzyme. (C) Gel of 3 μM dOGTP and five units/μL of enzyme. Following this experiment, the diastereomers of dSpTP (R and S) were tested. As anticipated, the concentration of dSpTP necessary to cause insertion was higher than that of dOG or dG, due to the propeller shape of the hydantoin ring that impacts proper base stacking in DNA, and thus insertion. At concentrations of 300 nM and 3 μM dSpTP, there was no noticeable insertion for S-Sp and possible insertion at 3 μM dSpTP for R-Sp (Figure 2A). At the concentration of 30 μM – at least 100 times larger than the concentration necessary to get dOGTP insertion – there can be seen insertion for dSp (Figure 2B, 2C). 20 A C B A T C GAT C G AT C G 300 nM 3 μM T A C G A T CGA T CG Figure 2. dSpTP insertion assays gels with 10 nM duplex DNA and 5 units/μL polymerase. (A) Gel of S-Sp. (B) Gel of 30 μM R-Sp. (C) Gel of 30 μM S-Sp. For both diastereomers, the insertion was seen opposite the templates containing A or C, which is contrary to the data expected for dSp – where this data reflects insertion of dNTPs opposite dSp in the template. The ratio of insertion was approximately 2:1 in favor of C for both diastereomers of dSp. This may be due to the fact that when it is in the template, it displays one of its rings to base pair, and the other ring when it is present as a triphosphate. When in the template, the dSp is locked into a confirmation due to base stacking, but when present as the triphosphate, it has more freedom as to display the preferential ring. Succinctly, the dSp is in the syn form in one instance and the anti form in the other, giving the varied base pair partners. The other possibility is that these samples of dSpTP contain dGTP as an impurity during their synthesis; future studies will be conducted to better understand this observation. Also of interest were “satellite” bands in the gels containing high concentration of dSp. These bands were at a distance that would indicate full insertion, despite the fact that only dSpTP was present as a nucleotide. Based on the fact that dSpTP was the only nucleotide that would be inserting, it is concluded that these satellite bands are most likely left over signal from a previously imaged gel. An additional expedition into the insertion partners of Gh was to be done, but due to time constraints and the low yield of product, such gels have not yet been run. 21 Efforts were then directed toward the extension of primers that terminated in the oxidized guanine lesions. First, Klenow Fragment exo- was tested. Initial tests at 30 μM dNTPs showed no insertion opposite any of the lesions tested, so the concentration of dNTPs was increased tenfold to 300 μM. At this concentration, extension was seen to some extent for all templates that were opposite the dOG terminating primer and to a minute degree for some templates that contained the primer terminating in dGh (Figure 3). The highest percentage of extension of the dOG primer was seen off dA and dC templates, which had either the dA or dC base pairing with the dOG (Table 1). The bands for extension can be seen in for the dGh containing samples, but the percentage of extension is on the order of noise. There was no insertion seen opposite either dSp diastereomer for any of the templates. ATCGATCGA TCGATCG OG A TCGA TCGATCGATC G Gh S-Sp R-Sp Figure 3. Gel of the extension percentage off of guanine oxidation product terminating primers, 10 nM duplex DNA, 5 units/μL Klenow Fragment, 300 μM dNTP’s. Table 1. Percentage extension by Klenow Fragment exo- for various templates with primers terminating in oxidative guanine lesions. “- -“ denotes no detectable extension. OG Gh S-Sp R-Sp A 10.2 ± 2.7 0.7 -- -- T 2.9 ± 0.4 -- -- -- C 5.1 ± 1.2 1.8 -- -- G 4.1 ± 1.0 -- -- -- 22 Another polymerase lacking the exonuclease domain, Dpo4 from Sulfolobus solfataricus, a homologue of DNA Polymerase IV, was tested to determine how different polymerase deal with oxidative lesions. Trials began with 300 μM dNTPs under the impression that extension would require the same concentration of dNTPs regardless of polymerse. The gel run for Dpo4 could only be run in duplicate, so the results have a greater standard deviation with a contribution to this large distribution also coming from the low intensity of the bands in the gel (Figure 4). Using Dpo4, there was a greater percentage of extension beyond the dOG-containing primer for all templates besides the dG template (Figure 5). However, there was no indication of extension beyond dGh primer or either dSp primer. This indicates that Dpo4, the DNA translesion synthesis polymerase, perpetuates more mutations then Klenow Fragment exo- for DNA containing the common dOG oxidation product in the template, but has a lower percentage of extension, and thus less inducing of mutation, than Klenow Fragment for the hydantoins – dGh and dSp. A B A TC G A TCG ATCGA T C G OG Gh A T C GA T C G A T C G A T C G S-Sp R-Sp Figure 4. Gels showing the extension of templates using 2 units/μL of Dpo4 and 300 μM NTP’s. (A) Gel of dOG and dGh primers opposite templates. (B) Gel of dSp diastereomer terminating primers opposite templates. 23 Percent Bypass 20 15 10 5 0 Polymerase and Template Figure 5. Comparison of extension percentages of Klenow Fragment exo- versus Dpo4 for each template. In the insertion portion of this research, it was found what the insertion partners were for the three guanine oxidative lesions studied when they are present as nucleotides. It was found that there is a possible change in base pair partners or distribution of incorporation between lesions that are present as part of the template and lesions that are present as nucleotides. This can explain the purportedly odd base pairing of dSp with dA and dC as opposed to dA and dG. As stated earlier, when dSp is part of the template, it may be in one conformer (syn or anti) that displays one of its rings for base pairing while when dSp is a nucleotide it adopts the other conformer – and thus exposes a different ring for base pairing – explaining a change in base pair partner. Another explanation for the distribution of dSp incorporation products is an impurity of dG. To distinguish which explanation is more plausible, purification cycles of dSp will be run, running gels after each cycle to see if the distribution of products changes. In the elongation portion of this research, it was found that for both polymerases studied that dOG is the lesion from which it is easiest to extend beyond. The polymerase Dpo4 had a higher percentage of elongation past the dOG containing primer when it was opposite most templates, meaning that it is more likely to perpetuate the mutation caused 24 by an oxidation of a dG to dOG. Klenow Fragment exo- had a higher affinity for the hydantion dGh and was seemingly able to extend beyond dGh when it was part of the primer, whereas Dpo4 showed no extension. This means that Klenow Fragment was less selective in its processing beyond lesions, meaning it would perpetuate mutations as a result of dGh present in DNA. 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