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Biochemical basis of SOS-induced mutagenesis: UmuD'(2)C is an error-prone DNA polymerase, Escherichia coli Pol V

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Biochemical basis of SOS-induced mutagenesis: UmuD'(2)C is an error-prone DNA polymerase, Escherichia coli Pol V

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Content INFORMATION TO USERS This manuscript has been reproduced from the microfilm master. UM I films the text directly from the original or copy submitted. Thus, some thesis and dissertation copies are in typewriter face, while others may be from any type of computer printer. The quality of this reproduction is dependent upon the quality of the copy submitted. Broken or indistinct print, colored or poor quality illustrations and photographs, print bleedthrough, substandard margins, and improper alignment can adversely affect reproduction. In the unlikely event that the author did not send UMI a complete manuscript and there are missing pages, these will be noted. Also, if unauthorized copyright material had to be removed, a note will indicate the deletion. Oversize materials (e.g., maps, drawings, charts) are reproduced by sectioning the original, beginning at the upper left-hand comer and continuing from left to right in equal sections with small overlaps. Photographs included in the original manuscript have been reproduced xerographically in this copy. Higher quality 6" x 9” black and white photographic prints are available for any photographs or illustrations appearing in this copy for an additional charge. Contact UMI directly to order. ProQuest Information and Learning 300 North Zeeb Road, Ann Arbor, Ml 48106-1346 USA 800-521-0600 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. BIOCHEMICAL BASIS OF SOS-INDUCED MUTAGENESIS: UMUD’2 C IS AN ERROR-PRONE DNA POLYMERASE, E.COLI POL V by MENGJIA TANG A Dissertation Presented to the FACULTY OF THE GRADUATE SCHOOL UNIVERSITY OF SOUTHERN CALIFORNIA In Partial Fulfillment of the Requirements for the Degree DOCTOR OF PHILOSOPHY (MOLECULAR BIOLOGY) August 2000 Copyright 2000 Mengjia Tang Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. UMI Number: 3018134 UMI UMI Microform 3018134 Copyright 2001 by Bell & Howell Information and Learning Company. All rights reserved. This microform edition is protected against unauthorized copying under Title 17, United States Code. Bell & Howell Information and Learning Company 300 North Zeeb Road P.O. Box 1346 Ann Arbor, Ml 48106-1346 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. UNIVERSITY OF SOUTHERN CALIFORNIA THE GRADUATE SCHOOL UNIVERSITY PARK LOS ANGELES, CALIFORNIA 90007 This dissertation, written by ' ylBNt'SlA. TAM under the direction of he.tr. Dissertation Committee, and approved by all its members, has been presented to and accepted by The Graduate School, in partial fulfillm ent of re quirements fo r the degree of DOCTOR OF PHILOSOPHY Dean of Graduate Studies Date ...2909. DISSERTATION COMMITTEE Chairperson Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. To my parents Shi-Ji Tang and Shi-Fan Li Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Acknowledgements This dissertation would never be finished without the participation and encouragement of many individuals. I would like to thank my advisor, Or. Myron Goodman, for his enthusiastic support and creative guidance throughout my graduate studies. I am grateful to Dr. Roger Woodgate (NIH) for his collaboration and scientific inspiration. I also like to thank Dr. Rahul Warrior for bringing me into the exciting MolBio program and for his constant support to my scientific career. I am also grateful to Dr. John Petruska for his insightful discussions and to Dr. Joseph Landolph for his time and advice on my dissertation. I am deeply grateful to Xuan Shen for his help in purifying the mutant UmuD’2C104 complex and wildtype UmuD’2 C complex from pol lllfs strain, and for some fidelity data on pol III and pol IV. I would also like to thank Dr. Phuong Pham, who purified pol IV and investigated its ability to bypass lesions, and participated in the fidelity measurement on pol III and pol IV. I would like to thank my past and present colleagues in Myron’s group - Alice, Brigette, Eli, Jeff, Mike, Phoung, Savi, Xiluo, Xuan, and Zhi-Hao - for their friendship and scientific interactions. Lastly, I would like to thank my sister and my parents for their love and encouragement, and my husband, Xiang, for his love, dedication and everything he has done for me. iii Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Table of Contents Dedication ii Acknowledgements iii List of Tables vii List of Figures viii Abstract x Chapter 1. Introduction 1 1. Escherichia coli DNA polymerases 3 1.1 DNA polymerase I 4 1.2 DNA polymerase II 5 1.3 DNA polymerase III 6 1.3.1 The functional components/subassemblies of the pol III holoenzyme 7 (a). The polymerase core 7 (b). The p sliding clamp and the y clamp loader complex 8 (c). The x subunit 9 1.3.2 pol III HE in replication 9 1.3.3 Mechanisms for maintaining high fidelity replication 10 2. Major DNA repair systems in E. coli 1 1 2.1 Methyl-directed mismatch repair 1 1 2.2 UvrABC-dependent nucleotide excision repair 12 2.3 Recombinational repair 13 3. The SOS response 15 3.1 The SOS regulatory network 16 3.2 SOS mutagenesis 18 3.2.1. Multiple roles of RecA 19 3.2.2. The UmuC and UmuD proteins 20 (a). The umuDC genes are under SOS control 20 (b). Posttranslational regulation of Umu activity 21 (c). umu mutants 23 (d). Homologs of the Umu proteins 24 3.2.3. Models for UmuD’aC-mediated translesion synthesis 25 Chapter 2. Materials and Methods 34 1. Materials 34 1.1 Bacterial strains and plasmids 34 1.2 Nucleotides 34 1.3 Enzymes and antibodies 35 iv Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 2. Methods 36 2.1 Primer/template construction 36 2.2 Purification of soluble UmuD’aC complex 37 2.2.1 Purification of UmuD’2 C from a wild type strain 37 2.2.2 Partial purification of UmuD’aC complex from a ApolB dnaE1026 strain 38 2.2.3 Purification of the mutant UmuD’aC104 complex from a wild type strain 39 2.3 Replicative bypass assay 40 2.4 Nucleotide incorporation kinetics 41 2.5 Kinetic analysis of nucleotide incorporation efficiency and polymerase fidelity 42 Chapter 3. Function of UmuD'2 C in SOS Mutagenesis: Discovery of the Error-Prone E. Coli Polymerase V 49 1. Introduction 49 2. Results 51 2.1 Purification of UmuD’aC complex 51 2.2 Reconstitution of in vitro SOS translesion synthesis 52 2.3 UmuD'aC catalyzes RecA*-dependent lesion bypass in the absence of exogeneous pol III core 53 2.4 UmuD’aC has intrinsic DNA polymerase activity 55 2.5 Other proteins required for UmuD’aC catalyzed translesion synthesis 57 2.6 UmuD'aC competes with pol III for binding primer-3'OH ends. 60 2.7 UmuD'aC catalyzed base misincorporation and mismatch extension at aberrant and normal template 62 3. Discussion 64 3.1 Reconstitution of a UmuD'aC-RecA-dependent SOS lesion bypass system in vitro. 65 3.2 UmuD'2 C is an error-prone DNA polymerase, E. coli pol V. 67 3.3 Pol III and pol V compete for primer-3'-OH ends. 68 3.4 Requirement of RecA*, SSB, p and y- complex for pol V- catalyzed lesion bypass 69 (a). Role of RecA protein - an insight to the targeting mechanism. 69 (b). SSB protein is essential for pol V’s activity 70 (c). What is the role of p sliding clamp in translesion synthesis? 73 3.5 UmuD'aC is error-prone at both damaged and normal template sites 74 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Chapter 4. Roles of E. coli DNA Polymerases IV and V in Lesion Targeted and Untargeted SOS Mutagenesis 87 1. Introduction 87 1.1 The UmuC/DinB/Rad30 superfamily 87 1.2 Common DNA lesions 89 2. Result 90 2.1 Comparison of translesion synthesis by pol V Mut, pol IV and pol III HE 90 2.2 Incorporation specificities opposite lesion sites 91 2.3 Incorporation fidelities on normai template sites 92 3. Discussion 94 Chapter 5. Conclusions and Perspectives 110 1. Summary of this dissertation 111 2. A few further thoughts 113 (a). Two sides of a coin - temporal actions of the inducible polymerases after UV irradiation 113 (b). Survival vs. evolution - purposeful conservation of the UmuC superfamily 115 Appendices 1. RecA activation in the in vitro lesion bypass assay 118 2. ATPase assay for the in vitro TLS system 120 3. N-ethylmaleimide inhibits pol V activity 122 4. Pol V does not incorporate ddNTP 123 5. A poly(A) polymerase activity in the UmuD’2 C preparation? 125 References 128 vi Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. List of Tables Table 1 -1. E. coli DNA polymerase III holoenzyme subunits and 27 subassemblies Table 1 -2. Properties of some important alleles of recA 28 Table 2-1. Bacterial strains and plasmids 45 Table 2-2. Template/Primer sequences for TLS assay 46 Table 2-3. Primer sequences for fidelity measurement 48 Table 4-1. Pol V Mut insertion specificity at DNA template lesions: 108 comparing in vitro and in vivo data Table 4-2. Fidelity of pol V Mut, pol IV, and pol III a HE by steady state 109 kinetics vii Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. List of Figures Figure 1-1. Damage tolerance strategies in E. coli 29 Figure 1-2. Proposed action of pol III HE at a replication fork 30 Figure 1-3. The SOS regulatory network 31 Figure 1-4. Regulation of UmuD’2 C activity in SOS mutagenesis 32 Figure 1-5. The two-step model of SOS translesion synthesis 33 Figure 3-1. Purification of UmuD’2 C complex from a wild-type strain 76 Figure 3-2. Reconstituting in vitro SOS lesion bypass system 77 Figure 3-3. UmuD’2 C (E coli pol V) has polymerase activity 78 Figure 3-4. Protein cofactor requirements for UmuD’2 C (E coli pol V)- catalyzed lesion bypass 79 Figure 3-5. Effect of RecA on incorporation of correct dAMP at an undamaged template by pol V 80 Figure 3-6. TLS in the presence of increasing concentration of ATP or ATPyS 8 1 Figure 3-7. Role of p, y complex in TLS in the presence of ATP or ATPyS 82 Figure 3-8. Effect of SSB on pol V-catalyzed TLS 83 Figure 3-9. Inhibition of pol V-catalyzed translesion synthesis by pol III 84 Figure 3-10. Pol V is error-prone on both damaged and normal template sites 85 Figure 3-11. A model for pol V-catalyzed translesion synthesis 86 Figure 4-1. The UmuC/Rad30/DinB/Rev1 DNA polymerase superfamily 99 Figure 4-2. Structures of three common lesions 100 Figure 4-3. Comparison of translesion synthesis catalyzed by pol III HE, pol IV and pol V Mut 101 viii Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Figure 4-4. Incorporation of A compared with G opposite a TT (6-4) photoproduct 103 Figure 4-5. Incorporation of A compared with G opposite a TT cis-syn photodimer 105 Figure 4-6. Incorporation of T or C opposite template lesion sites 106 Figure 4-7. Incorporation of A compared with G opposite an abasic site 107 Figure A-1. Activation of RecA in the in vitro lesion bypass assay 119 Figure A-2. ATPase assay for the in vitro TLS system 121 Figure A-3. NEM inhibits pol V activity 122 Figure A-4. Pol V is unable to utilize ddNTPs as substrates 124 Figure A-5. Primer extension in the absence of template and dNTPs 126 Figure A-6. A terminal transferase activity in the UmuD’2 C preparation 127 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Abstract SOS mutagenesis requiring the UmuD’C proteins occurs as part of the cells’ global response to DNA damage. The biochemical mechanism of SOS mutagenesis has remained a mystery for the past two decades due to the difficulties in obtaining biologically active UmuC protein, which is insoluble in aqueous solution when overproduced. Using purified UmuD’2 C complex in its native form, we have reconstituted an SOS lesion bypass system in vitro. We discovered that UmuD’aC is the fifth DNA polymerase in E. coli. Efficient lesion bypass requires the function of pol V mutasome (Mut), consisting of pol V, activated RecA protein (RecA*), single-stranded binding protein (SSB), processivity factor | 3 sliding clamp, and y clamp loading complex. Effect of RecA on pol V causes dramatic increase in DNA synthesis efficiency. Pol V is nonprocessive, and competes with replicative pol III at lesion sites for free primer-3’-OH end. Yet it can interact with pol III in the vicinity of lesion sites, allowing pol III to take over after lesion bypass occurs. We have examined the efficiency and specificity of pol V in bypassing common template lesions: pyrimidine-pyrimidone (6-4) photoproduct, cis-syn cyclobutane dimer, and abasic site. Pol V bypasses all three lesions within 30 seconds. In comparison, there is no detectable bypass by either pol III or the newly discovered pol IV on this time scale. A mutagenic “signature” for pol V is its incorporation of G opposite the 3’-T of a TT (6-4) photoproduct in agreement x Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. with mutational spectra. In contrast, pol III and pol IV incorporate A almost exclusively, albeit with much reduced efficiency. We have also measured the nucleotide incorporation fidelity of pol V at undamaged DNA template sites. Pol V exhibits low fidelity with misincorporation frequencies ranging from - 10'3 to 1C4; pol IV being 5- to 10-fold more accurate. Both pol V and pol IV exhibit enhanced ability to extend mismatched primer ends, the latter via a misalignment mechanism. While a definitive role of pol IV in untargeted mutagenesis remains to be clarified, our results suggest that pol V is responsible for both targeted and untargeted SOS mutagenesis in vivo. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Chapter 1 Introduction The survival of every organism requires high accuracy in maintaining its genome integrity. To achieve this, faithful duplication of the genome information is essential. In Escherichia coli, the sequential application of three mechanisms that will be discussed later in the chapter, base selection, proofreading editing, and postreplicative mismatch repair, yields precisely duplicated DNA, with error frequencies typically in the very low range of 10'9 to 10* 1 ° per base replicated (58, 238). In addition, cells also possess a variety of DNA repair systems that remove and repair modified or damaged bases from DNA. One such pathway involves direct reversal of damaged bases, using highly specialized enzymes such as photoreactivating enzymes in bacteria and an 0 6 -methylguanine specific methyltransferase in E. coli and mammals (241, 275). Another pathway, the base excision repair (BER) pathway, removes altered bases by DNA glycosyiases followed by apurinic/ apyrimidinic (AP) endonuclease and polymerase gap filling (146). Two additional repair systems, nucleotide excision repair and reccmbinational repair, will be discussed in detail in the U DNA repair” section in this dissertation. These repair systems protect genome from the potentially harmful effects of damages caused by exogenous and endogenous sources. i Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Despite the combined actions of highly accurate replication and repair pathways, situations can arise in which DNA lesions have escaped the repair processes. Based on their effects on DNA replication, lesions fall into two categories: (i) error-promoting lesions that can increase mispairing of bases (e.g. 0 6 -methyl guanine and 2-aminopurine), and (ii) replication-blocking lesions that introduce substantial distortion into DNA (e.g. pyrimidine dimers) and effectively impede replication fork advancement (58). In eukaryotic cells, the introduction of replication-blocking lesions into DNA causes cell cycle arrest or apoptosis. In E. coli and other eubacteria, however, replication-blocking lesions on DNA induce the SOS response, which includes phenomena such as enhanced capacity for DNA repair by means of excision repair and recombinational repair, cell cycle inhibition, and elevated mutation frequencies (76). Associated with the SOS response is a recovery of DNA synthesis. The renewed DNA replication is achieved through a group of mechanistically different processes, collectively referred to as damage tolerance strategies (9) (Figure 1-1). The first strategy involves reinitiation of replication downstream of the lesion (known as “replication-restarf), creating a gap in the newly synthesized daughter strand that is later repaired by recombinational repair (Fig. 1-1 A). In the second strategy, the polymerase can transiently copy the newly synthesized daughter strand from the sister chromatid, which has the same polarity as the damaged parental strand, and then switch back to the original template downstream of the lesion (Fig. 1-1B). In these two strategies, Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. the replication machinery avoids directly copying the damaged template. Hence they are known as damage avoidance (DA) strategies (76,118). The third strategy, however, occurs when the replication machinery directly incorporates a nucleotide opposite a noncoding or miscoding template lesion site and subsequently elongates the terminus (Fig. 1-1C). This process, called translesion synthesis (TLS), is potentially mutagenic because the polymerase may insert an incorrect base. In most organisms, most mutagenesis resulting from DNA damages caused by UV radiation, ionizing radiation and various chemicals appears to be due to TLS (280). In E. coli, TLS requires the function of UmuDC and RecA proteins, and results in a phenomenon called SOS mutagenesis, which is manifested by a 100-fold increase in mutation frequency after exposure to DNA damaging agents (76). This dissertation focuses on studying the biochemical mechanism of SOS mutagenesis, in particular, the role of UmuD’C proteins in translesion synthesis. This chapter is organized as follows: the first section is a brief introduction of the E. coli polymerases, followed by description of several major DNA repair systems, and finally, the SOS regulatory network is discussed, with an emphasis on SOS mutagenesis. 1. Escherichia coli DNA polymerases To date, five DNA polymerases (pol) have been identified in E. coli. The first three well-characterized polymerases ~ pol I, II, and III - will be discussed 3 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. in this section, focusing on the replicative polymerase, pol III holoenzyme. Pol IV (encoded by the dinB gene), discovered last year, will be discussed later in Chapter 4. The discovery and characterization of pol V is the focus of this work, and will be discussed throughout this dissertation. 1.1 DNA polymerase I Discovered by A. Komberg in 1956 (122), DNA polymerase I (pol I) was the first polymerase identified in E. coli. Encoded by the polA gene (108), pol I contains a single polypeptide (109 kDa) that can be separated by proteolytic cleavage into two active fragments: a large C-terminal fragment, called the Klenow fragment, and a small N-terminal fragment. The major activities of pol I can be summarized as: 5’->3’ polymerization activity, 3’-*5’ proofreading exonuclease activity, and a unique 5’-*-3’ exonuclease activity, with the first two activities carried on the Klenow fragment, and the latter on the small fragment (123). Pol I functions both in replication and repair [reviewed in ref. (123)]. With its 5’->3’ polymerase activity, pol I joins Okazaki fragments on the lagging strand during DNA replication, and participates in the gap filling step of the UvrABC excision repair, which will be discussed later in the chapter. Pol I can also initiates DNA replication in certain plasmids with a ColE1 or pMB1 origin. The proofreading 3'-*-5’ exonuclease activity recognizes and removes a mismatched primer terminus while the polymerase moves along the DNA template, a role crucial for the fidelity of polymerization reactions. The 5’->3’ Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. exonuclease activity is unique not only in its direction, but also in its requirement for base paired region. It is involved in removing the RNA primers from a RNA-DNA hybrid in Okazaki fragments during lagging strand synthesis. 1.2 DNA polymerase II DNA polymerase II was discovered in 1970 from mutant strains lacking pol I activity (114,124). Encoded by the polB gene (183) [also known as dinA (17)], pol II is a single polypeptide (89.9 kDa) having both 5’-> -3 ’ polymerase activity and 3’-*5’ exonuclease activity (125, 248). Mutant strains defective in pol II activity have been isolated (37, 94), and mutants of pol II deficient in either polymerase activity (214) or exonuclease activity (36) have been purified and wall characterized in vitro. Based on its amino acid sequence similarity to the eukaryotic DNA polymerase o c , pol II has been classified as a group B (“a-like”) polymerase (243). In 1988, pol II was rediscovered as a damage-inducible polymerase that is capable of copying and bypassing an abasic lesion in vitro (17, 18). It is induced 7-fold after DNA damage occurs, implying that it may be involved in some DNA repair processes (16). The biological function of pol II, however, remained ill characterized due to the lack of apparent cellular phenotypes in pol II deletion strains. The in vivo role of DNA polymerase II is, however, no longer a mystery. It was suggested that pol II is involved in episomal replication in nondividing cells 5 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. (69) and repair of DNA interstrand cross-links (10). Rangarajan ef a/in our laboratory were able to show that, when competing with an antimutator allele of pol III, dnaE915, pol II can function in both chromosomal and episomal DNA replication in vivo (220). The breakthrough in studying the function of pol II came from the observation that a ApolB AumuDC strain is markedly more UV sensitive than the strains devoid of either gene, suggesting that pol II plays a role in DNA repair. It was further revealed that, in the ApolB strain, there is a 50- min delay in recovery of DNA synthesis after UV-irradiation, confirming that pol II indeed participates in the replication restart process as part of damage avoidance strategies (221). 1.3 DNA polymerase III The DNA polymerase III holoenzyme (Pol III HE) is the replicative enzyme in E. coli whose main function is to replicate the E.coli chromosome [reviewed in ref. (110) and (166)], although it functions in repair and mutagenesis as well (59). Pol III HE shares special features with replicases of eukaryotes, viruses, prokaryotes and their phages, which distinguishes holoenzyme from single-subunit polymerases such as Pol I and pol II. Among these features are multi-subunit structure, the requirement for ATP to clamp tightly to DNA, the rapid speed of DNA synthesis, and a remarkably high processivity such that the enzyme remains bound to DNA for thousands of polymerization events (110). Table 1-1 lists the 10 subunits of pol III HE and 6 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. their subassemblies. This section discusses three aspects of the pol III holoenzyme: (i) the functional components of the holoenzyme: the core, the p sliding clamp, and the y clamp loader complex, (ii) the asymmetric structure of the holoenzyme, and (iii) the fidelity mechanisms of replication. 1.3.1 Functional components/subassemblies of the pol III holoenzyme (a) The polymerase core The polymerase core consists of three tightly associated subunits: (i) the polymerase subunit a , (ii) the proofreading exonuclease e , and (iii) the 6 subunit, whose function is not clear. The a subunit is encoded by dnaE gene, with a molecular mass of 130 kDa. It synthesizes DNA at a rate of 8 nucleotides per second. The e subunit, which is a 27.5 kDa polypeptide encoded by dnaQ/MutD gene, forms a tight complex with a subunit, resulting in increases in both polymerase activity and exonuclease activity (110). The function of 0 subunit has yet to be identified. It binds £ but not a, suggesting a linear a-£-0 arrangement in core (258). The polymerase core is present in approximately 20 to 40 copies per cell and less than half of them are assembled into the holoenzyme (155). In the absence of its accessory proteins, core is a weak polymerase, synthesizing DNA at a rate of 20 nucleotides per second, and is processive for about 1 1 nucleotides (65). However, it becomes the fastest polymerase in the presence 7 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. of its accessory proteins, and its processivity increase to more than 5000 nucleotides (discussed below). (b). The P sliding clamp and the y clamp loader complex The p sliding clamp is required by the holoenzyme to achieve high processivity (197). The sliding clamp, made of a homodimer of the p subunit, has a doughnut-shaped structure that can encircle and slide along DNA (see Fig. 1-2) (121). It binds pol III core through the interaction with the a subunit, thus tethering it to DNA (133, 259). As a consequence, the processivity increases dramatically, reaching about 5,500 nucleotides (65, 66,134). However, the p sliding clamp does not assemble onto DNA by itself. The y complex, consisting of five different subunits in the stiochiometry Y 2 Si8’iXiVi (156), is the molecular matchmaker that hydrolyzes ATP to load p clamps onto DNA (110). ATP hydrolysis provides the energy required to complete the assembly reaction that involves loading p onto DNA followed by dissociation of y complex (11, 272). The DNA-dependent ATPase activity is associated with y subunit and can be stimulated by p subunit and primed template. Ason et al (2) recently suggested that a DNA-triggered switch between active and inactive states of y complex provides a two-tiered mechanism enabling y complex to recognize primed template sites and load p, while preventing y complex from competing with DNA polymerase III. The mechanism by which y complex loads p onto DNA is as follows: One dimer of p is chaperoned to DNA by the y complex in the absence of core to 8 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. form the “preinitiation complex” (198). This step depends on binding of ATP to the y complex. In the second step, the core assembles with the preinitiation complex to form the “initiation complex” (198) by direct protein-protein interaction between the p and the a subunit (259), forming a processive replication complex. (c). The x subunit The y subunit is the translational frameshift product of the dnaX gene, whose full translation product is the x subunit (68). The x dimer binds two molecules of core, thus dimerizing two cores at replication fork, one on the leading strand, the other on the lagging strand, x also interacts with Y-complex. The resulting structure, pol III*, consists of 2 cores bound to one x dimer and one y complex. 1.3.2 Pol III HE in replication During DNA replication, synthesis of the leading strand and synthesis of the lagging strand are quite different (123). The leading strand polymerase need only remain clamped to DNA continuously, but the lagging strand is synthesized discontinuously as a series of short Okazaki fragments. Accordingly, the pol III holoenzyme adapts an asymmetric structure in which the accessory proteins are distributed asymmetrically (100,167), achieved by the asymmetric structure of Y-complex through the interaction between the Ysubunit and the x dimer (156). The discontinuity of lagging strand synthesis also 9 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. requires the lagging strand polymerase be repeatedly clamped and unclamped from DNA to cycle from one fragment to the next (246). Kinetic studies have demonstrated that the half-life for loading is on the order of 60 ms, rapid enough to support in vivo Okazaki fragment synthesis occurring about every 1 to 2 seconds (15). Figure 1-2 depicts the asymmetric structure of pol III holoenzyme and its action during DNA replication. 1.3.3 Mechanisms for maintaining high fidelity replication For DNA polymerases, precise replication is accomplished by three steps (58): (i) Base selection. Polymerases can distinguish and incorporate a correct deoxynucleoside triphosphate (dNTP) substrate 104 to 105 times more efficiently than an incorrect nucleotide. Such discrimination against wrong nucleotides is exerted by the polymerase active sites that have evolved mechanisms to select for the precise geometry of the Watson-Crick base pairs within DNA double helix (58,59,84). (ii) Proofreading editing. Mismatched primer termini tend to melted out at the 3’ end, allowing their excision by the 3’->5’ exonuclease activity associated with some DNA polymerases. This editing mechanism contributes to about 100-fold increase in replication fidelity, (iii) Reduced mismatch extension. Mismatched 3' termini cannot be further extended efficiently. Although polymerases appear to have similar binding affinity to matched and mismatched primer termini, extension of a mismatched primer terminus is usually kinetically blocked (52,169). This delay in addition of 10 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. next nucleotides also provides longer time for the exonuclease proofreading activity to excise a mismatched nucleotide. 2. Major DNA Repair Systems in E. coli 2.1 Methyl-directed mismatch repair The methyl-directed mismatch repair pathway is capable of repairing base mismatches, small insertions and deletions that arise from polymerase errors during DNA replication (174-176). In all eight types of base-base mismatches, A-A, A-C, G-G and G-T are repaired efficiently, A-G, T-C and T-T are repaired with moderate efficiency, whereas C-C is poorly repaired (174). The key genes involved in this pathway are: mutH, mutL, and mutS, whose mutations exhibit mutator phenotypes that cause a 1,000-fold increase in spontaneous mutation frequency (238). In addition, several other proteins are required in the reconstituted in vitro mismatch repair systems: helicase II, single-stranded binding protein (SSB), DNA polymerase III, DNA ligase, and one of the exonucleases (Exol, ExoVII, or RecJ) (135). The strand specificity of mismatch repair is signaled by the methylation state of GATC sequences, where the newly synthesized strand is transiently unmethylated (213). The repair process is initiated by binding of MutS homodimer to mispaired sites on the heteroduplex, which in turn recruits MutH and MutL. The MutS-MutL-MutH complex then begins to search for a nearby hemimethylated GATC site in a bidirectional, ATP-dependent manner to u Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. generate a looped structure, called a-loop. Upon reaching a GATC site, MutH, whose latent endonuclease activity is stimulated by the interaction with MutS and MutL, makes a nick at the unmethylated site on the newly synthesized strand. Helicase II then comes along and unwinds the nicked strand toward the mismatched site. Since the nick can occur at either side of the mismatch, different exonucleases are required for the subsequent digestion of unwound ssDNA: if the nick is located 3’ to the mismatch, exonuclease I degrades DNA in 3’-»5’ direction; otherwise, RecJ or exo VII hydrolyzes DNA in 5’->3’ direction. The resultant gap that can be several thousands base pair in length is filled in by DNA pol III and ligated by DNA ligase (174). Eukaryotic homologs of MutS and MutL have been identified in many organisms that share the same functional properties, but are hetero- rather than homodimeric [reviewed in ref. (176) (99) and (120)]. Interestingly, no MutH homolog has been identified, suggesting that a methyl-directed strand recognition and nicking mechanism may not exist in eukaryotes. 2.2 UvrABC-dependent nucleotide excision repair Nucleotide excision repair (NER) is an error-free repair process that removes the region containing a UV dimer, bulky adduct, or interstrand cross link [reviewed in ref. (232) (234) and (274)]. In E. coli, this repair pathway requires the function of 6 genes: uvrA, uvrB, uvrC, uvrD, polA and lig, the first two of which are under the control of the SOS response (233,235). Like those Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. mutators defective in mismatch repair, NER-deficient cells are highly mutable and very sensitive to UV irradation. The molecular mechanism of excision repair has been best elucidated in E. coli. In the presence of ATP, a UvrA dimer binds to single strand or damaged DNA and provides a binding site for UvrB. However, this initial binding of UvrA2 B complex to DNA is non-specific and unstable, allowing the UvrA2 B complex to travel along DNA as a helicase until it encounters a lesion at which point UvrA is released (204, 205). The resultant stable UvrB-DNA complex serves as the binding site for UvrC. UvrC has endonuclease activity (295) that makes two incisions: one is 7-8 bases 5' to the lesion and the other 3-4 bases 3’ to the lesion (231). After the region between the two cuts is removed in the presence of UvrD helicase, the 12-13 base gap is filled in by DNA polymerase I and ligated by DNA ligase (40,130, 206). Eukaryotic NER differs from that of E. coli in two major aspects. First, although eukaryotes share a common biochemical mechanism with E. coli, many more proteins are needed to carry out each step in eukaryotes compared to bacterial NER [reviewed in ref. (7, 53,146, 210, 290)]. Second, there is little homology between bacterial and eukaryotic NER proteins (60). 2.3 Recombinational repair Two recombinational repair pathways have been identified in E. coli, both dependent on the recA gene: the RecF pathway and the RecBC pathway. Both pathways are induced by the SOS response (207) and contribute equally to 13 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. repair of damaged DNA during normal growth (43, 46, 47,119,132). The RecF pathway repairs daughter strand gaps produced by the replication-restart process, while the RecBC pathway repairs double-strand breaks and double strand ends generated by disintegration of stalled replication folk (132). Another form of double strand lesions, psoralen intrastrand cross-links, can be repaired by both pathways (245). The three common phases of the two repair reactions are (i) presynapsis, during which the damaged DNA is prepared for homology search, followed closely by synapsis, during which homologous pairing and strand exchange with the intact sister duplex occur; (ii) DNA replication restart; and (iii) postsynapsis, during which the recombination intermediates are resolved (132). The postulated events in the RecF pathway are as follows (132): in preparation for synapsis, a RecOR complex descends on the SSB-coated daughter strand gap, perhaps guided by a RecFR complex. The presence of RecO allows RecA polymerization on the SSB-complexed ssDNA. During synapsis, the RecA filament finds an intact duplex homologous to the single strand gap and pairs them. Paring the damaged and the intact DNA molecules allows filling in of the gap by a DNA polymerase. In the postsynapsis phase, RuvABC resolvasome or RecG helicase removes Holliday junctions and the associated RecA filament, completing faithful repair of the daughter strand gap. The RecBC pathway differs from the RecF pathway in that, in the presynapsis phase, a single strand region has to be generated in preparation for strand pairing (46). This is done by ExoV (RecBCD), which degrades the 14 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. double strand end until a property oriented Chi site converts RecBCD degradase into RecBCD* recombinase. After Chi, RecBCD* continues to degrade DNA but only the 5’-end strand, generating a 3’ single-stranded overhang to which RecA binds. The RecA protein promotes homologous pairing and strand exchange, forming a D-loop with a single three-strand junction. The junction may be resolved by RecG helicase followed by PriA mediated replication restart. Alternatively, the 3’ end of an invading strand can be extended by pol I, followed by PriA mediated replication restart, converting it into a four-strand junction that is resolved by RuvABC. 3. The SOS Response The idea that E. coli possesses a regulatory network in response to DNA damaging agents was developed by Radman (217) and Witkin (286). As Radman stated in the written proceedings of the 1973 conference on mutagenesis: The principal idea is that E. coli possesses a DNA repair system which can be repressed under normal physiological conditions but which can be induced by a variety of DNA lesions. Because of its “response" to DNA-damage agents we call this hypothetical repair “SOS repair”. The “danger" signal which induces SOS repair is probably just a temporary blockage of the normal DNA replication and possibly just the presence of DNA lesions in the cell. In E. coli, the SOS response controls more than 40 genes, known as SOS genes, whose functions enhance the survival after cells experience 15 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. DNA damage. The SOS phenomena include enhanced DNA repair events, inhibition of cell division, prophage induction and mutagenesis (76). 3.1 The SOS regulatory network The regulation of the SOS response in E. coli is achieved through the coordinated control of two proteins: LexA protein as the repressor, and RecA protein as the activator (Fig. 1-3). Under normal (uninduced) conditions, LexA protein binds to operator sequences known as SOS boxes, allowing only basal level expression of the SOS genes (20-22,149). When DNA damage happens or when replication is inhibited, an inducing signal for SOS response is generated. Both in vivo and in vitro data suggest that the inducing signal is single-stranded regions of DNA that are generated when the cells attempt to replicate a damaged template or when normal replication is interrupted (230, 237). RecA protein then binds to these single-stranded regions in the presence of a nucleoside triphosphate to form a nucleoprotein filament (49, 50), reversibly converting to its activated form, referred to as RecA* (147). RecA* possesses a coprotease activity that promotes the proteolytic autocleavage of LexA at the position of Ala^-Glyas (95,148). As the intracellular level of active LexA repressor drops, the originally repressed SOS genes are now expressed at increased levels, and SOS phenomena mediated by these genes can be observed (induced state). As the cells recover from the damage by means of DNA repair, the levels of inducing signal and RecA* coprotease decrease, 16 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. allowing accumulation of the LexA repressor. The cells eventually return to the uninduced state. The mechanism of SOS regulation was revealed largely by isolation of lexA and recA mutants. Two classes of lexA mutants have been identified. The lexA (Ind ) mutants prevent SOS induction and are dominant to lexA* (88,185). These mutants often have missense mutations that alter the Ala^-Glyas cleavage site, resulting in resistance to RecA* mediated autocleavage (143). The lexA (Def) mutants, on the other hand, fail to produce functional LexA repressor and thus constitutively express the SOS response (184). They are recessive to lexA+ , and can arise from a variety of mutations such as missense, amber, deletions and Tn5-generated insertions (76). Similarly, various types of recA mutants have been isolated and will be discussed in section 3.2.1. A key property of the SOS regulatory network is that various SOS genes, including the lexA and recA genes themselves, are differentially regulated. Those that are repressed weakly are induced early and to a lesser extent. Examples of these include the uvrAB and lexA genes, with the induction level ranging from 4 to 6-fold. Other genes that are regulated tightly are induced late and to a much greater degree. The mostly tightly repressed SOS gene, the sulA gene, whose product prevents cell division and causes formation of long cell filament, is induced more than 100-fold. Such a “graded response” (117) allows cells with minimal damage to induce error-free repair processes, such as nucleotide excision repair and recombinational repair, before inducing other repair pathways that’s are potentially error-prone. Indeed, more extensive 17 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. cellular DNA damage is needed for expression of the umuDC genes required for error-prone translesion synthesis of damaged DNA (discussed in the next section). 3.2 SOS mutagenesis Both excision repair and recombinational repair pathways are error-free pathways, and account for repair of about 85% of the lesions resulting from DNA damaging agents. Nonetheless, the induction of the SOS response is accompanied by a 100-fold increase in mutation frequency, a phenomenon referred to as “SOS error-prone repair” or “SOS mutagenesis” (217, 286). As discussed earlier in the chapter, SOS mutagenesis results from error-prone translesion synthesis, where most of the mutations appear to be targeted to the sites of DNA damage. Weigle first reported SOS-induced error-prone repair through the observation that UV irradiation of E. coli prior to infection with phage increases the survival and mutation frequency of UV irradiated phage (284). The phenomena, known as Weigle reactivation and Weigle mutagenesis, respectively, are induced in a recA+ -/exA+ -dependent fashion by UV irradiation (54). The genes responsible for Weigle mutagenesis are recA, umuC and umuD, all of which are inducible and regulated as part of the SOS response. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 3.2.1 Multiple roles of RecA RecA is a 37.8 kDa protein (165) that participates in a number of processes that both maintain and diversify the genetic material of the bacterial cell, including homologous recombination and repair, replication restart after UV-irradiation, chromosomal segregation, SOS induction and mutagenesis (42, 128). In SOS mutagenesis, activated RecA protein is required for cleavage of the LexA repressor to turn on the umu genes, and for cleavage of the UmuD protein to activate the mutagenic pathway (see next section). In addition, RecA has a direct role in mutagenesis, revealed by the observation that constitutive expression of UmuD’C proteins in a ArecA strain did not lead to mutagenesis above spontaneous background levels (192). The function of RecA in this process might be the targeting UmuD’C proteins to lesions in DNA, a notion supported by the observations that RecA physically interacts with UmuD and that a mutant RecA defective in this interaction in vitro fails to exhibit SOS mutagenesis in vivo (74, 250). Several classes of recA mutants have been isolated (Table 1-2) (127, 128,278). recA (Cptc) alleles show enhanced activities relative to the wild-type. Many of these mutants, such as recA441 and recA730, were isolated on the basis of their ability to induce the SOS response or prophage constitutively in the absence of DNA damage (288). recA (Cpt-) alleles are partially defective mutants, an example being RecA430, which has a lower binding affinity for 19 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. ssDNA and is partially defective in recombination, SOS response and prophage induction (154,170). recA (Def) alleles, including recA1, recA 13 and recA56, are completely inactive in vivo and indistinguishable from ArecA. The fourth class of recA mutants display differential alterations, some of which are constitutive for coprotease activity but are recombination defective [e.g. recA1201 (268)], while others show differential repressor and UmuD cleavage specificities [e.g. recA1734 (57)]. Lastly, the RecA1730 mutant is defective in protein filament formation (57) and fails to interact with UmuD (74). It is recombination deficient unless overexpressed (57) but remains defective in SOS mutagenesis, suggesting that the filament formation defect, but not the improper targeting of the UmuD’C complex, can be overcome at high protein concentrations (3). 3.2.2 The UmuD and UmuC proteins The umuDC aenes are under SOS control. - In addition to RecA, the UmuC and UmuD proteins are required for most chemical and UV-induced mutagenesis. Mutations in either gene result in a nonmutable phenotype by a variety of agents (109, 252, 253). The umuDC genes are expressed from one operon, umu, located near 25 min on the E. coli genetic map, with a one-base overlap (61,244). In fact, umu is the only operon that must be induced to achieve the cell’s full mutagenic potential, since the basal level of RecA, about Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 7,200 molecules in uninduced cells, appears to be adequate for SOS mutagenesis (251). The umu genes are among the last genes induced during the SOS response, whose expression levels increase 20 minutes after SOS induction. The cellular levels of UmuD protein are about 180 molecules before and 2400 molecules after SOS induction. The cellular level of UmuC in the absence of SOS induction is undetectable, and about 200 molecules after induction (292). Production of UmuD’, a posttranslational processed form of UmuD (discussed next), switches repair from homologous recombination to SOS mutagenesis (249), possibly by inhibiting the formation of joint molecules in RecA-mediated recombination (223, 263, 276). Posttranslational regulation of Umu activity. -The umuC and umuD genes encode proteins of 46 kDa and 16 kDa, respectively (244). Intact UmuD protein, however, is inactive in SOS mutagenesis. The mutagenic active form, known as UmuD’ (14 kDa), has to be generated by posttranslational proteolytic autocleavage of UmuD, occuring at the position of Cys2 4 -Gly2 5 (33,208). The cleavage is mediated by activated RecA*, in a manner similar to, but much less efficient than, the cleavage of LexA. Both homodimers and heterodimers of UmuD and UmuD’ are formed under physiological conditions, but only UmuD’2 C is mutagenically active when complexed with UmuC (6,293). The co-existence of three UmuD dimer species in vivo implicates a third mechanism to regulate SOS mutagenesis in which intact UmuD acts as an inhibitor of SOS 21 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. mutagenesis by forming the inactive homodimer UmuD2 or heterodimer UmuDD’ (6,107). Support for this notion is that the heterodimer UmuDD’ is more stable than the two homodimer species (6) and can complex with UmuC (73, 294), thereby precluding the formation of the active UmuD^C complex. UmuD may also help shut off the error-prone translesion synthesis once the damaged DNA has been repaired. Working together, the three regulatory mechanisms ensure that error-prone translesion synthesis is activated late in SOS induction, possibly after other error-free repair systems fail (Fig. 1-4). Both UmuC and UmuD proteins are labile, with in vivo half-life of 6-7 minutes (72), and are susceptible to Lon protease degradation (73). The UmuD’ homodimer, however, is much more stable than the other two dimeric species. UmuD’ is subject to CIpXP protease degradation only when complexed with UmuD protein (73). An in vitro experiment demonstrated that UmuC is dramatically stabilized when complexed with UmuD’ (72). Before forming a complex with UmuD’, which does not occur until about 45 minutes after SOS induction (250), UmuC is stabilized by the GroEL and GroES proteins, which are molecular chaperones that play roles in preventing denaturation and aggregation of other proteins and assisting protein folding (297). Such a notion is supported by the observations that groEl and groES mutants have reduced Umu-dependent mutability to UV radiation (56) and that modest overexpression of UmuD’ and UmuC can suppress this deficiency (55). Moreover, UmuC protein coimmunoprecipitates with GroEL, suggesting an interaction between the two proteins (55). 22 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. umu mutants - In general, umuC and umuD mutants are largely nonmutable in the presence of a variety of mutagenic agents including UV radiation, 4-NQO, MMS, and neocarcinostatin, but they can still be mutated by chemical agents such as MNNG or EMS, which produce directly mispairing lesions such as 0 6 -alkylguanine (76). The umu mutants are modestly sensitive to UV killing but are much less sensitive than uvr, recA (Def), or lexA (Ind ) mutants. Several types of UmuD mutations have been isolated. One type produces altered proteins that cannot be cleaved to generate active UmuD’, such as umuD44 (6). The second type is defective in protein dimerization, while the third is defective in its ability to complex with UmuC protein. Likewise, UmuC mutants also differ in their abilities to interact with UmuD’. umuC36, umuC25 and umuC104 are all nonmutagenic, but unlike the first two mutants, UmuC104 is capable of interacting with UmuD’ (107). Overexpression of wild-type umuDC in a lexA (Def) strain results in cold- sensitivity, due to a rapid and reversible inhibition of DNA synthesis at nonpermissive temperature (30°C) (159,203). A umuC mutant, umuC125, was isolated based on its ability to suppress the conditional cold-lethality. umuC125 cells are proficient in UV radiation mutagenesis (158), suggesting that the cold- sensitivity results from an activity distinct from that in mutagenesis. It is now confirmed that UmuDC, but not UmuD’C, functions as a damage checkpoint protein by inhibiting polymerase replication after exposure to DNA damaging agents (186, 202). 23 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Homoloas of the Umu proteins - Several UmuDC homologs have been identified in other bacteria, such as umuDCsr in Salmomella typhimurium (115, 194), and on naturally occurring plasmids, including mucAB (209), impAB (151, 257), rumAB (129) and samAB (193,194). Like E. coli umuDC, these genes are also organized into operons and are damage-inducible. The pKM101plamsid carried mucA and mucB genes are homologous to umuD and umuC, respectively (182, 208, 209). Expression of mucAB suppresses the nonmutable phenotype of the umuDC mutant (281), and results in an even higher mutation frequency (12, 279). This enhanced mutability stems from the MucA protein, which is cleaved more efficiently than UmuD to generate the mutagenic active form (MucA’) (92). Interestingly, despite their high amino acid sequence similarities, UmuDC, MucAB and RumAB proteins generate different mutation spectra when participating in translesion synthesis (140, 262). Oda et al. reported that UmuD’C mRNAs injected into Xenopus oocytes can alleviate replication arrest of single-stranded M13 DNA containing cyclobutane pyrimidine dimers (196), suggesting eukaryotic cells contain lesion bypass systems related to prokaryotes. As expected, distant homologs of UmuC are now identified in eukaryotes (163, 229). These homologs share only limited conserved motifs that, not realized until last year, are important for their conserved functions, which will be discussed in chapter 4. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 3.2.3 Models for UmuD’aC-mediated SOS translesion synthesis Resulting from translesion synthesis, SOS mutagenesis clearly requires the function of a DNA polymerase. Both pol I (5) and pol II (97,111,126) are not required for the process. The participation of pol III, on the other hand, is implicated from a set of genetic studies involving temperature-sensitive pol III mutants (25, 26, 32, 89). B. Bridges and R. Woodgate observed that, although umuC, umuD and /exA(lnd') strains are largely nonmutable by UV radiation, they can be mutated to a significant degree if, following exposure to UV radiation, the cells are incubated for up to 4 hours and then exposed to photoreactivating light (27, 28). They hence proposed a two-step model for SOS mutagenesis (28, 29) (Fig. 1- 5). In the first step, pol III inserts a nucleotide opposite the lesion, but continued chain elongation is blocked. Although they proposed that this step doesn’t depend on the functions of UmuD^C and RecA, certain recA alleles can depress misincorporation (24). The second step involves UmuD’C- and RecA- dependent elongation of the primer terminus, the molecular mechanism of which was obscure. A number of models have been proposed for the role of UmuD’C in translesion synthesis, including suppression of the proofreading activity, increase in processivity of the polymerase, or relaxation of its requirement for a proper Watson-Crick DNA structure for continued replication (76). However, it remained open to speculation due to lack of evidence, or sometimes the existence of conflicting results. 25 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. The major difficulty in elucidating the translesion synthesis mechanism stemmed from the inability to obtain active UmuC protein, which is insoluble in aqueous solution. In previous studies, the UmuC protein was purified in a denatured form followed by refolding it in the presence of equimolar amounts of S9 protein or heat shock proteins (211,294). In vitro replicative assay using these proteins resulted in very limited, and often spurious, lesion bypass (218). Recently, a soluble UmuC protein has been purified in our lab, in the form of UmuD’2 C complex (31). It binds cooperatively to long stretches of ssDNA, but not dsDNA. It also interacts with RecA*-coated ssDNA (31) and effectively blocks recombinational strand exchange in vitro (223). The purpose of this dissertation is to study the biochemical basis of SOS translesion synthesis, by reconstituting an in vitro lesion bypass system using purified proteins. This dissertation presents evidence leading to the exciting discovery that UmuD’2 C itself is a bona fide DNA polymerase, designated as E. coli pol V. Pol V is error-prone on both normal and damaged templates, catalyzing efficient bypass of common UV lesions that would otherwise severely inhibit replication by pol III holoenzyme. In disagreement with the two-step model, UmuD’2 C functions at both misincorporation and bypass steps in the process of translesion synthesis, but may interact with pol III in the vicinity of the lesion sites, allowing normal replication to resume downstream of the lesion. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Reproduced with permission o f th e copyright owner. Further reproduction prohibited without permission. Table 1-1. E. coli DNA polymerase III holoenzyme subunits and subassemblies Subunit Gene Mass (kDa) Function Subassembly a dnaE 129.9 DNA polymerase “ e DnaQ, 27.5 Proofreading 3’-+5’ exonuclease core e holE 8.6 Stimulates e exonuciease pol III’ T dnaX 71.1 Dimerizes core, DNA-dependent ATPase pol III* Y DnaX 47.5 Binds ATP 5 holA 38.7 Binds p 5’ HolB 36.9 Cofactor for y ATPase and stimulates clamp y complex X holC 16.6 Binds SSB ¥ holD 15.2 Bridge between % and y - P dnaN 40.6 Sliding clamp on DNA Reproduced from ref. (110) to Table 1-2. Properties of some important alleles of recA Allele Phenotype Biochemical Alteration recA441 Spontaneous SOS induction at 42°C in the absence of DNA damage Coprotease more easily activated recA730 Constitutive expression of SOS genes, Enhanced Recombination Coprotease constitutively active, enhanced recombinase activity recA430 Partially defective in recombination, SOS response and prophage induction Lower binding affinity for ssDNA, and lower rate of association with ssDNA recA1, A recA Recombination-deficient, SOS-deficient, Extremely sensitive to DNA damage Defective RecA recA1201 Constitutive expression of SOS genes, Deficient in recombination ? recA1734 Proficient in SOS induction, Deficient in SOS mutagenesis Coprotease proficient in LexA cleavage, but defective in UmuD cleavage recA1730 Proficient in recombination when over expressed, Deficient in SOS mutagenesis Defective in protein filament formation and interaction with UmuD Adapted from (127, 128). Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. y Replication block y y y X X D am age Avoidance B Translesion synthesis f X- m R ecom binational ! Polym erase template repair switch Error-free V M utagenic Figure 1-1. Damage tolerance strategies in E. coli. [Reproduced from ref. (9)] 29 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Figure 1-2. Proposed action of poi III HE at a replication fork. The holoenzyme structure is placed at a replication fork with one core polymerase on each strand. The y complex is asymmetrically disposed relative to the two cores such that it points toward the lagging strand to load g clamps on primers repeatedly to initiate processive extension of Okazaki fragments. A. As lagging- strand polymerase extends an Okazaki fragment, the y complex assembles a p clamp onto an RNA primer. B. Upon completing an Okazaki fragment, the core disengages its ( 3 clamp, creating a vacancy for the new p clamp. C. The new p clamp falls into place with the lagging-strand core polymerase to start the next Okazaki fragment. [Reproduced from ref. (110)] 30 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. UNINDUCED STATE f To other genes controlled By LexA repressor LexA repressor Protein recA gene/ Inducible gene Operator Messenger RNA Repressor accumulates I Drop in RecA coprotease level Drop in lcwel of signal DNA repaired DNA Damage Inducing signal RecA* coprotease activated LexA repressor cleaved TT V W W lA iV W 1 ^ °°g ® ° Activated RecA* " S . coprotease Cleaved LeexA repressor f r repressor; lexA gene INDUCED STATE recA gene Induced gene Figure 1-3. The SOS regulatory network. [Modified from ref. (76)]. 3 1 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. LexA 1 I umuD umuC ------------------ y . RecA* -r~ □ □ umuD 1 ------------------- T----- umuC --------1 ------------------- ► UmuD RecA* UmuD’ j ■ ■ ■ ► C -Z l GroEL GroES X z x Q Q Q « — - ■ ■ -*- QJQQ ------------ ► Proteolysis x i 111 1 i Cell Cycle Control SOS Mutagenesis Figure 1-4. Regulation of UmuD’2 C activity in SOS mutagenesis. [Modified from ref. (76)]. 3 2 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Pol III HE P-clamp 7 -complex 3’ core 5’ 3 ’ RecA* UmuD’,C 3 ’ 5 ’ 3 ’ . 5 ’ Figure 1-5. The two-step model of SOS translesion synthesis. [Modified from ref. (76)]. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Chapter 2 Materials and Methods This chapter describes the materials and methods employed throughout this work. It is organized into two sections: the general materials such as enzymes and dNTP substrates are listed in the Materials section; the second section describes methods including construction of DNA primer/templates, purification of the UmuD’2 C complex, and in vitro translesion synthesis assays. 1. Materials 1.1 Bacterial strains and plasmids Table 2-1 lists all the plasmids and bacterial strains used to overproduce Umu proteins in this study. Details about strain and plasmid construction (Dr. Roger Woodgate, National Institutes of Health) are described in ref. (31) and (267). All strains, including “wild-type” strain, are devoid of the umuDC operon to avoid interference from the chromosomal UmuD protein. In this study, the term “wild-type strain” refers to the strains whose genes encoding for the E. coli DNA polymerases I, II, III and IV are normal. 1.2 Nucleotides Ultrapure ATP, deoxynucieoside triphosphates (dNTP), nucleoside triphosphates (NTP) and dideoxynucleoside triphosphates (ddNTP) were 34 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. purchased from Amersham Pharmacia. Radioactive [y -3 2 P] ATP (4000 Ci/mmol; 1 Ci = 37 GBq) and [a -3 2 P] ATP (3000 Ci/mmol; 1 Ci = 37 GBq) were from ICN. Nonhydrolyzable ATP-5’-[yS]thiotriphosphate (ATP7S) was purchased from Boehringer Mannheim. 1.3 Enzymes and antibodies T4 DNA ligase was purchased from Promega. EcoRI restriction enzyme was from New England Biolabs. T4 polynucleotide kinase, pol I Klenow fragment, E. coli single-stranded binding protein (SSB) and RecA protein were from Amersham Pharmacia. Purification of pol II was described in ref. (35). Purified pol III and its accessory proteins (188) were provided by Dr. Mike O’Donnell at Rockefeller University. Pol IV, fused to maltose binding protein (MBP), was purified as described (266). Purified UmuD, mutant UmuDK79A, mutant RecA1730 (74), and LexA proteins were provided by Dr. Roger Woodgate (NIH). Anti-RecA, UmuC and UmuD antibodies were raised in rabbits and affinity purified (Roger Woodgate, NIH). Pol I antibody was a generous gift from Lawrence Loeb (University of Washington, Seattle), and pol III a subunit antibody was from Dr. Mike O’Donnell. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 2. Methods 2.1 Primer/template construction All oligonucleotides were synthesized on an Applied Biosystems model 392 DNA/RNA synthesizer and gel purified. Biotinylated-CPG columns were purchased from Glen Research. The abasic (1,4-anhydro-2-deoxy-D-ribitol) phosphoramimide was synthesized as described (62) by Dr. Ramon Eritja at EMBO, and the TT (6-4) pyrimidine-pyrimidone photoproduct and the TT c/s- syn cyclobutane photodimer were synthesized by Dr. John-Stephen Taylor at Washington University (98). Circular single-stranded M13mp7 DNA was prepared using PEG/CTAB method. The templates used in the replicative bypass assay were 7.2-kb linear single-stranded DNA, constructed by ligating linearized M13mp7 molecules with lesion-containing 60-mers in three steps: (i) Single-stranded M13mp7 molecule was linearized with EcoRI (1 pg DNA /1 Unit enzyme), which cuts the DNA at the hairpin-structured multiple cloning site, at 37°C for 1 hr, followed by inactivation of the enzyme at 65°C for 20 minutes, (ii) The linearized M13mp7 DNA was then ligated with the lesion-containing 60-mers in the presence of a linker (16°C, overnight), using T4 DNA ligase. To make the abasic lesion- containing template (M13GM202X2 ), the ratio of M13mp7/30-mer linker/60-mer was 1/2/5. In the case of making the TT cis-syn photodimer (M13GMcs) or TT (6-4) photoproduct (M13GM64) templates, where the lesion-containing oligonucleotides were in limited supply, the lesion containing 12-mers were first 36 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. ligated with a 48-mer, and the resultant 60-mers were then ligated with the linearized M13mp7 in the ratio of M13mp7/linker/60-mer being 1/1.2/1.5. (iii) To remove the linkers from the ligated products, the ligation mixtures were incubated (75°C, 20 min) with excess amount (about 5 to 10-fold) of anti-sense linkers that have the complementary sequences of the linkers, then slowly cooled to room temperature. The ligated products were purified either by running through a gel filtration column (A-0.5 agarose resin, Bio-Rad) with 40 mM NaCI in TE buffer, or in the case of biotinylated anti-sense linkers, by using streptavidin-coated magnetic beads (Pierce) according to manufacture’s suggestion. The efficiencies of ligation reaction were greater than 95%, as judged by the percentage of full length primer extension products on ligated DNA. For the template carrying an abasic site (M13GM202X2 ), the abasic lesion is located 50 bases from the 5' end, and for the templates with a TT cis- syn photodimer (M13GMcs) or TT (6-4) photoproduct (M13GM64), the lesion sites are 53 and 54 bases from the 5’ end. The primers were 5'-end 3 2 P-labeled 30-mers. Table 2-2 lists the sequences of all the primers and their positions on individual template. 2.2 Purification of UmuD’2 C complex 2.2.1 Purification of UmuD’aC complex from a wild-type E. coli strain Purification was carried out as reported (31) with the following modifications: after cell lysis and poiyetheleneimine (1.1% wt/vol, pH 7.6) 37 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. precipitation, proteins were extracted by stirring the pellet in R-buffer (20 mM Tris-HCI, pH 7.5/0.1 mM EDTA/1 mM DTT/20% glycerol) containing 1 M NaCI. Ammonium sulfate was added to reach 50% of saturation, followed by centrifugation of the suspension. The pellet was dissolved in R-buffer containing 1 M NaCI, dialyzed against R-buffer with 1 M NaCI, followed by a second ammonium sulfate precipitation step in which UmuDzC complex precipitated at 30% saturation. The spun pellet was again dissolved in R-buffer with 1 M NaCI and dialyzed against R-buffer containing 50 mM NaCI before chromatography using DEAE followed by phosphocellulose as described (31). Phosphocellulose fractions containing UmuDaC were concentrated and applied onto Superdex 75 column (Amersham Pharmacia) and eluted in R-buffer containing 1 M NaCI. Fractions containing UmuD’2 C proteins were detected by Western blotting using antibodies against UmuC and UmuD/D’ in the early, bulk precipitation steps, and visualized by staining with Coomassie blue R-250 for the later, chromatography steps. 2.2.2 Partial purification of UmuD’2 C complex from a dnaE1026 ApolB strain. The earlier purification steps were carried out as described above (see section 3.2.1). In addition, the cell debris obtained after cell lysis was reextracted with R buffer with 1 M NaCI and precipitated with polyethyleneimine (1.1% wt/vol). The supernatant containing UmuD'2 C was precipitated with 50% saturation of ammonium sulfate, and the pellet was combined with that from cell 38 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. lysate, dissolved and dialyzed in R-buffer with 1 M NaCI. Proteins from the second ammonium sulfate precipitation, as described in the previous section, were centrifuged, resuspended, and dialyzed in R buffer containing 1 M NaCI. The dialyzed proteins were loaded onto a Superdex 200 size exclusion column (16/60, Amersham Pharmacia) and run in R buffer with 1 M NaCI, but containing no glycerol. Fractions (1.1 mi) were collected, and glycerol was added to individual fractions to reach a final concentration of 20%. Fractions containing UmuD’2 C proteins were visualized by staining with Coomassie blue R-250, resolved on a 12% SDS/PAGE gel. Fractions containing pol III temperature sensitive (ts) were detected by Western blotting by using antiserum directed against the a subunit. 2.2.3 Purification of the mutant UmuD’2C104 complex from a wild-type strain The mutant UmuD’2C104 (D101N) protein was purified in a similar manner as described above (see section 3.2.2) except that gel filtration was carried out by using Superdex 75 (26/60, Amersham Pharmacia). The fractions that contained abundant UmuD’2 C104 protein complex but had barely detectable pol III were pooled and further purified by phosphocellulose column chromatography (265). Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 2.3 Replicative bypass assay. The reaction mixture (10 pi) contains 20 mM Tris-HCI (pH 7.5), 8 mM MgCI2, 5 mM DTT, 0.1 mM EDTA, 25 mM sodium glutamate, 40 pg/ml BSA, and 4% (vol/vol) glycerol. ATP is at 1 mM, unless otherwise specified. Running-start reactions were performed as follows: 2 nM primed DNA substrate was incubated for 3 min at 37°C with 40 nM ( 3 protein dimer, 10 nM y complex, 1 pM RecA, and 100 pM each of dATP and dCTP. Replication was initiated by addition of UmuD^C and SSB (300 nM as tetramer), and dGTP and dTTP (100 pM each). Other polymerase, either pol III core, pol III a subunit, or pol II, when present at concentrations between 0 and 20 nM, was added together with UmuD’aC. Reactions were carried out at 37°C for 10 min, then quenched by adding 20 pi EDTA (20 mM) in 95% formamide. The product DNA was heat-denatured run on a 12% polyacrylamide denaturing gel, dried and exposed to a phosphor screen (Molecular Dynamics). Replication products were quantitated by using a Phosphorlmager (Molecular Dynamics). Standing-start reactions were carried out in a similar manner as running-start reactions, except that all four dNTPs were added after preincubation with RecA, SSB, (3 , and y complex. The replicative products were resolved on a 16% polyacrylamide gel. 40 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 2.4 Nucleotide incorporation kinetics A gel kinetic analysis was used to determine the incorporation efficiency and polymerase fidelity (51). All reaction mixtures (10 pi) contained 20 mM Tris (pH 7.5), 8 mM MgCI2 , 5 mM DTT, 0.1 mM EDTA, 25 mM sodium glutamate, 40 pg/ml BSA and 4% (v/v) glycerol. For incorporation opposite lesion sites, the reactions were carried out in 1 mM ATP, except for that at the 5’-T site of TT cis-syn cyclobutane dimer where reduced ATP concentration (100 pM) was used to avoid misincorporation of ribo-AMP. Primer/template DNA (2 nM) was incubated at 37°C for 3 min at varying dNTP substrate concentrations, 40 nM of p (as dimer), 10 nM y complex, and RecA (1 pM, for pol V, but not pol III and IV). Running-start reactions were initiated by adding 10 pM running start dNTP, 300 nM SSB (as tetramer), and either pol III a-subunit (1 to 5 nM), pol IV (20 nM) or pol V (25 to 50 nM). The running start nucleotide was absent in standing-start reactions. Reactions were run at 37°C for 5 min and quenched by adding 20 pi of 20 mM EDTA in 95% formamide, and the products were separated on a 16% polyacrylamide denaturing gel. Polymerase fidelities were measured using standing-start reactions on undamaged linear M13mp7 DNA templates annealed to 3 2 P-labeled 30-mer primers. The reactions were similar to those for the lesion sites, except that the template /primer concentration was 0.5 nM, and ATP was at 100 pM. When measuring incorporation of the right nucleotides, the reactions were done for 30 41 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. sec for pol V (10 nM) and pol IV (1 nM), and 4 min for pol III a (0.5 nM). When measuring incorporation of the wrong nucleotides, the reactions were done for 2 min for pol V (20 nM) and pol IV (2 nM), and 8 min for pol III a (2 nM). The following template sequences were used: 3’...GTTGCCG...; 3’...CGTAGCC...; 3’TCGTAGC...; 3’TATCAAC, where the base in boldface is the template target site at which the nucleotide misincorporation frequency, fin c , was measured. 2.5 Kinetic analysis of nucleotide incorporation efficiency and polymerase fidelity A steady-state gel kinetic analysis was used to determine the rate of abasic translesion synthesis as a function of dNTP concentration (51). Elongated 3 2 P-labeled primer extension products were separated by using 16% 8M urea-polyacrylamide gels. Integrated polyacrylamide gel band intensities were measured with a Phosphorlmager by using ImageQuant software (Molecular Dynamics). For running-start reaction, the relative rates of lesion bypass, vre t, were determined by measuring where Ixs designates the integrated gel band intensities of primers extended opposite the lesion site, T, and beyond, i.e., translesion synthesis, and IT ., is 42 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. the integrated gel band intensity of primers extended by addition of a single running-start nucleotide opposite the template site before the lesion site (51). For standing-start reaction, the relative rates of incorporation opposite lesion or normal target sites, \/e l, were determined by I i vr e i = ---------------= r for the right nucleotides, (Ip+0.5xIT" )xTime(min) and vrtl = ----------------- x ^E -n z y .me^R for the wrong nucleotides, (Ip +0.5x1^ )xTime(min) [Enzyme ]w where IT£ is the integrated gel band intensities of primers extended opposite and beyond the target site, I p is the integrated gel band intensity of unextended primers, and [Enzyme]R and [Enzyme]w refer to the enzyme concentration for right and wrong nucleotide incorporation, respectively, at the same target site. A plot of the relative velocity, vrB l, as a function of dNTP substrate concentration results in a rectangular hyperbola whose slope in the initial linear region is the apparent Vm a x /Km . Apparent Km and relative Vm a x values were obtained from a nonlinear least squares fit to the rectangular hyperbola, or from double reciprocal Lineweaver-Burk plots. In reactions where misincorporation opposite the target site was relatively inefficient, the plots showed little or no curvature, and apparent V m a x/K m values were obtained by a least squares fit of 43 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. the data to a straight line. The misincorporation efficiency, fin e , which is the inverse of the fidelity, is given by f in c = fidelity'1 = -V ,n i“ /Km)'v ( V ^ / K J , where the subscripts Wand R refer to incorporation of the wrong and right nucleotide, respectively. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Table 2-1. Bacterial strains and plasmids. Strains____________ Relevant Genotypes____________Selective Markers RW82 recAf lexA* AumuDC595::cat CamR RW86 recAf lexA51 (Def) AumuDC595::cat Tcn CamR RW88 recA443 lexA51(Def) AumuDC595::cat Tcr CamR DV08 ApolB::Q AumuDC596::ermG T Str0 SpcR RW384 ApolA 1 ApolB::Q AumuDC595::cat lon146::Tn10 Str0 SpcRCamR RW510 polAl ApolBwCl AumuDC: .ermGT Str° SpcR ermGT RW588 ApolBv.Cl. dnaE1026ts AumuDC::ermGT Str0 SpcR ermGT RW610 polA 12ts ApolB::Q dnaE1026ts AumuDC595::cat Str0 SpcRTetR CamR RW626 ApolBv.Q dnaE486 AdinBr.kan AumuDC595::cat Str0 SpcRTetR KanR CamR Plasmids pOS1 UmuD’C overexpression plasmid under ptac control pEC47 UmuC overexpression plasmid under pT7 control pRW252 UmuD’C overexpression plasmid under pT7 control pRW392 Mutant UmuD’CI 04 overexpression plasmid under ptac control 45 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Table 2-2. Template/Primer sequences for TLS assay. Template M13GM202X2 (M13mp7 derivative containing a synthetic abasic lesion X, the italic portion of the sequence is from a synthetic 60-mer, the rest is from M13mp7.) 5 ' - TGTATTTTCT ACGTTTGCTA ACATACTTCG TAATAAGGAG TCTTAATCTX GCCAGTTCTT AATTCACTGG CCGTCGTTTT ACAACGTCGT GACTGGGAAA ACCCTGGCGT TACCCAACTT AATCGCCTTG .... - 3 ' Primers annealed to M13GM202X2: 1 . P1 mp7: (30-mer whose 3’-end is 46 nt upstream from the abasic site, annealed to M13 sequence ) 5 ' - GCGATTAAGT TGGGTAACGC CAGGGTTTTC - 3 ' 2. PRmp7: (30-mer whose 3’-end is 2 nt upstream from the abasic site, annealed to both 60-mer and M13 sequence ) 5 ' - TAAAACGACG GCCAGTGAAT TAAGAACTGG - 3 ' 3 . PSmp7: (30-mer whose 3’-end is 1 nt upstream from the abasic site, annealed to both 60-mer and M13 sequence ) 5 ' - AAAACGACGG CCAGTGAATT AAGAACTGGC - 3 ' Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Table 2-2 (cont.) Template M13GMcs or M13GM64 (M13mp7 derivative containing pyrimidine dimers TT, the italic portion of the sequence is from the synthetic 60-mer, the rest is from M13mp7.) 5 ' - TGTATTTTCT ACGTTTGCTA ACATACTTCG TAATAAGGAG TCTTAATCGA GTATTATGAG AATTCACTGG CCGTCGTTTT ACAACGTCGT GACTGGGAAA ACCCTGGCGT TACCCAACTT AATCGCCTTG .... - 3 ' Primers annealed to M13GMcs or M13GM64: 4. P 1 mp7: (as above, 43 nt upstream from the 3’-T ) 5 . PRGM: (30-mer whose 3’-end is 2 nt upstream from the 3’-T , annealed to both 60-mer and M13 sequence ) 5 ' - CGTTGTAAAA CGACGGCCAG TGATTCTCCA - 3 ' 6. PSGM: (30-mer whose 3’-end is 1 nt upstream from the 3’-T, annealed to both 60-mer and M13 sequence ) 5 ' - GTTGTAAAAC GACGGCCAGT GATTCTCCAT - 3 ' 7. PmAGM: (30-mer whose 3’-end covers the 3’-T with an A, annealed to both 60-mer and M13 sequence ) 5 ' - TTGTAAAACG ACGGCCAGTG ATTCTCCATA - 3 ' 8. PmGGM: (30-mer whose 3’-end covers the 3’-T with an G, annealed to both 60-mer and M13 sequence ) 5 ' - TTGTAAAACG ACGGCCAGTG ATTCTCCATG - 3 ' Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Table 2*3. Primer sequences for fidelity measurement. 1. XS3 (Position on M13mp7 DNA: 1338-1367 Target G. Template sequence 3’ -CTTGCCG- 5’) 5'- CGAAAGACAG CATCGGAACG AGGGTAGCAA -3' 2. Ast (Position on M13mp7 DNA: 1356-1385 Target A. Template sequence 3' -CGTAGCC- 5’) 5'- GATCGTCACC CTCAGCAGCG AAAGACAGCA -3' 3. Tst (Position on M13mp7 DNA: 1357-1386 Target T. Template sequence 3' -TCGTAGC- 5') 5'- GGATCGTCAC CCTCAGCAGC GAAAGACAGC -3’ 4. Cst_AA (Position on M13mp7 DNA: 1477-1506 Target C. Template sequence 3' -TATCAAC- 5') 5’- GTGAATTTCT TAAACAGCTT GATACCGATA -3’ Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Chapter 3 Function of UmuD’2 C in SOS Mutagenesis: Discovery of Error-Prone E. coli Polymerase V 1. Introduction Escherichia coli normally replicates its DNA accurately, but the fidelity of replication decreases dramatically after cells are exposed to a variety of DNA- damaging agents that induce the SOS response (76, 278, 286). Given a choice, E. coli will evoke a damage avoidance pathway that in all likelihood involves polymerase strand switching to a nondamaged DNA template (58,118). Nevertheless, situations arise where the DNA replication machinery encounters a lesion, and the only recourse is the direct replication of the damaged template. Genetic characterization of the SOS-induced error-prone translesion DNA synthesis pathway shows that it depends on the UmuD’2 C complex (61, 109, 244), activated RecA protein (RecA*) (57, 261), and DNA polymerase III (89). A prevailing model for translesion DNA synthesis, based on genetic experiments, suggests that it can be separated into two steps: nucleotide misincorporation directly opposite the lesion, formerly believed to require pol III and RecA (29, 242), and lesion bypass, formerly believed to involve pol III and UmuD’C proteins (27,189, 242). 49 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. A major roadblock in the path toward understanding the phenomenon of Umu-dependent translesion DNA synthesis had been the inability to purify biologically active UmuC protein. By using a denatured-renatured form of UmuC, H. Echols and colleagues (294) were able to purify UmuC and demonstrate that together with UmuD’ and RecA*, DNA pol III was able to facilitate limited translesion DNA synthesis of a synthetic abasic site (218). Their results, while extremely encouraging, were not unequivocal owing to the variable properties of the denatured-renatured UmuC protein. Recently, our lab has succeeded in purifying the native UmuD^C complex directly, in soluble form (31). It binds cooperatively to single-stranded DNA, having similar affinities for damaged and undamaged DNA (31), and effectively blocks recombinational strand exchange in vitro (223). In this dissertation, we successfully reconstituted an in vitro lesion bypass assay by using the soluble UmuD^C complex (265). Surprisingly, we discovered that, contrary to the two-step model in which UmuD’aC proteins were speculated to be fidelity-lowering accessory factors of pol III, UmuD’2 C itself is a bona fide DNA polymerase. As the fifth identified DNA polymerase, we designated the UmuD’2 C complex as E. coli pol V (267). Pol V by itself has very weak polymerase activity, and the optimal activity of pol V requires the presence of RecA, the p sliding clamp, the y clamp-loading complex, and E. coli single-stranded binding protein (SSB). Pol V is highly error-prone at both damaged and undamaged DNA template sites, catalyzing efficient bypass of 50 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. abasic lesions that would otherwise severely inhibit replication by pol III holoenzyme complex (HE). 2. Results Until our recent success in purifying a soluble, active UmuD’aC protein complex (31), studies on the biochemical basis of SOS-induced mutagenesis had been hampered by the absence of a reconstituted in vitro lesion bypass assay using purified proteins. The goal of this work was to study the role of UmuDaC complex in translesion synthesis by reconstituting such an in vitro system. 2.1 Purification of UmuD^C complex The soluble UmuD’2 C or the mutant UmuD’2C104 complex was purified by modifying the protocol developed by Irina Bruck (31), as described in Chapter 2. A purification gel for the wild-type UmuD’2 C complex from the wild- type strain RW82 is shown in figure 3-1. The final preparation is estimated to be at least 95% pure, based on the absence of contaminating bands on silver- or Coomassie-stained polyacrylamide gels. Unless otherwise specified, this UmuD’2 C complex preparation was used throughout the study in reconstitution of lesion bypass and characterization of its enzymatic properties. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 2.2 Reconstitution of in vitro SOS translesion synthesis. A linearized M13 DNA template containing a single. site-directed abasic lesion was copied by using different combinations of pol III core, pol III accessory proteins ( J S , 7 complex), RecA, SSB, and UmuD’2 C (Fig. 3-2 Left). The primer-template (Fig. 3-2 Upper) was designed to permit loading of the | 3 processivity clamp by the y complex (15) and to allow binding of UmuD’a C , RecA, and SSB (31). Pol III core alone catalyzes relatively weak, nonprocessive DNA synthesis (Fig. 3-2, lane 1). In the presence of ( 3 , y complex and SSB, DNA synthesis by pol III core becomes much more processive, terminating one base before reaching the abasic site, X-1 (lane 2). Addition of RecA, either to pol III core (lane 3) or to pol III core, ( 3 , y complex, and SSB (lane 4) stimulates each reaction, but primer elongation still terminates at the X-1 position, one base before the lesion. Addition of UmuDaC, however, enables significant bypass of the lesion, with continued synthesis to the end of the template (lane 5). The presence of an intense pause band at X-1 suggests that incorporation opposite the lesion is still rate limiting, although it is possible that this band could arise by excision of a nucleotide incorporated opposite the abasic site by the proofreading exonuclease of pol III core. The polymerization reaction leading up to the lesion and beyond is essentially nonprocessive in the presence of UmuDaC (lane 5), although pol III and |3 , y complex are both present along with ATP. A summation of integrated band intensities beyond the lesion (summed 52 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. from site X + 1 to the end of the template) relative to the bands extended up to and including the lesion (summed from site 1 to X) shows that the amount of bypass observed after a 10 min incubation is 20%; i.e., the amount of lesion bypass corresponds to 20% of the total synthesis. ATP is needed in the lesion bypass reaction for loading ( 3 on DNA by y complex (201) and also for binding RecA to DNA, thus converting it to RecA* (227). ATP is hydrolyzed throughout the reaction, resulting in a reduction to 70% of its initial level after a 10-min reaction (see Appendix-1, Fig. A-2). To verify that the initial input concentration of ATP (1 mM) is sufficient to sustain the conversion of RecA RecA*, we added LexA protein to the reaction at 10 min and observed RecA*-mediated cleavage of LexA (see Appendix-1, Fig. A- 1). 2.3 UmuD’2 C catalyzes RecA*-dependent lesion bypass in the absence of exogeneous pol III core Unexpectedly, lesion bypass was observed to occur in a "control" reaction containing UmuD'2 C, RecA, ( 3 , y complex, and SSB, in the absence of exogeneous pol III core (Fig. 3-2, lane 9). Lesion bypass does not occur in the absence of either RecA (lane 7) or ( 3 , y complex plus SSB (lane 8). However, one interesting difference in the gel band patterns is the appearance of two adjacent bands just before the lesion (at X -1) and directly opposite the abasic site (X) (lanes 7 and 8). Thus, a band corresponding to stable incorporation 53 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. opposite the lesion persists in the absence of added pol III core (in contrast to the presence of pol III core, compare with lanes 2-4). It is possible that the weak polymerase activity in the UmuD’2 C preparation (Fig. 3-2, lane 6) that is stimulated in the presence of RecA (lane 8) is due to trace amount of contaminating polymerases, although we estimate the UmuD’2 C preparation is greater than 95% pure (see section 2.1). The presence of pol I contamination can be ruled out because primer extension is unaltered when polymerization occurs in the presence of a potent neutralizing pol I antibody (data not shown). We have verified that the UmuD’2 C complex migrates on a gel filtration column with an expected molecular mass of 70 kOa, indicating that it is unlikely to be directly bound to a known polymerase, although it is possible that pol II, whose molecular mass is 89.9 kDa (17), is present as a contaminating polymerase. However, UmuD’aC complex purified from strains devoid of either polB or both polA and polB genes showed the same lesion bypass activity (data not shown), suggesting that UmuD'2 C itself contains an intrinsic, low fidelity, DNA polymerase activity. An E. coli strain carrying the recA 1730 mutation (S117F) is proficient for most of RecA's activities but is specifically defective for Umu-dependent mutagenesis (3, 57, 250), probably because RecA1730 is unable to target the Umu proteins to lesion-containing DNA (74, 250). Accordingly, bypass of the abasic lesion no longer occurs when wild-type RecA protein is replaced with purified RecA1730 (Fig. 3-2, lane 10), although weak DNA synthesis continues to take place. Instead, the primer is extended to one base prior to the lesion, 54 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. with a much smaller amount of incorporation occurring directly opposite the lesion. These data are in contrast to the lesion bypass promoted by wild-type RecA protein (lanes 5 and 9), but are consistent with the activity of RecA1730 in vivo (3, 57), and in vitro (74). 2.4 UmuD’2 C has intrinsic DNA polymerase activity To determine whether UmuD’2 C complex has an intrinsic DNA polymerase activity separate from pol II and pol III, we purified UmuD’2 C from an E. coli mutant strain containing a deletion of pol II (Apo/S) (63) and a temperature-sensitive allele of DNA pol III (danE1026) (283). Cell lysates were enriched for soluble UmuD'2 C complex by precipitation with polyethyleneimine and then with ammonium sulfate (31, 265). The UmuD'2 C-enriched fraction was run on a high-resolution Superdex 200 column, and individual fractions were assayed for the presence of pol III a subunit by Western blots (Fig. 3-3A upper gel) and for UmuC by using a Coomassie-stained gel (Fig 3-3A. lower gel). Western blots with polyclonal antibodies directed against UmuC and UmuD' showed that the fractions containing the UmuD'2 C complex correspond precisely to the locations determined with the Coomassie-stained gel (data not shown). Three Superdex fractions were chosen to measure polymerase and abasic lesion bypass activities: fraction (fx) 50 contains predominantly pol lilts core with a small amount of UmuD'2 C; fx 56 contains overlapping pol lilts + 55 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. UmuO’aC; fx 64 contains UmuD’2 C having no detectable pol lilts present (Fig. 3- 3B). The lesion bypass assay was performed on the same abasic lesion containing template (see section 2.2 & 2.3), with a 5’-3 2 P-labeled 30-mer primer (PRmp7) located two bases upstream from the abasic lesion (Fig. 3-3B). Primer extension was carried out with each of the three fractions in the presence of RecA, p, y complex, SSB, 4 dNTP substrates, and ATP. A single running-start nucleotide (dCMP) is incorporated opposite template G to reach the abasic lesion X. A primer band extended directly opposite X corresponds to nucleotide incorporation directly opposite the lesion. Longer primer extension bands reflect synthesis proceeding past the lesion site, i.e., lesion bypass. At the permissive temperature (37°C), fx 50 (containing pol lilts) incorporates a running-start C opposite G along with a small amount of incorporation opposite X and 1 nt beyond, but all synthesis is essentially absent at the nonpermissive temperature of 47°C (Fig. 3-3B). The extremely weak lesion bypass band observed at 37°C can be attributed to the presence of a small amount of UmuD'2 C (Fig. 3-3B). In contrast, fx 64, which contains no detectable pol lilts, carries out running-start incorporation and lesion bypass at both permissive and nonpermissive temperatures (Fig. 3-3B). We conclude, therefore, that UmuD'2 C has its own polymerase activity that can incorporate a running-start C opposite G in a template-directed reaction and can also catalyze nucleotide incorporation and continued extension at an abasic template lesion. 56 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Polymerization carried out by fx 56 (UmuD’2 C + pol lilts) revealed an interesting result. The robust incorporation and lesion bypass observed at 37°C is consistent with our previous observation that UmuD'aC catalyzes incorporation directly opposite X and then extends several bases beyond the lesion, at which point UmuD'aC dissociates and is replaced by pol III HE. However, unexpectedly, this synergistic effect on downstream synthesis also occurs at the nonpermissive temperature. As seen in Fig. 3-3B, the amount of total DNA synthesis is much greater for fx 56 (UmuD'aC + pol lilts) compared with essentially no DNA synthesis for fx 50 (pol lilts) and moderate synthesis for fx 64 (UmuD'aC). It appears that the temperature-sensitive mutant pol III is stabilized in the presence of high concentrations of UmuD'aC, suggesting that the two polymerases might interact in the vicinity of the lesion. Perhaps once lesion bypass has occurred, normal DNA replication can resume by replacing a distributive Umu polymerase with a processive pol III HE. 2.5 Other proteins required for UmuD’2 C catalyzed translesion synthesis We investigated requirements for other protein cofactors by permuting RecA, p, y complex, and SSB in the lesion bypass reactions (Fig. 3-4). We observed that UmuD’aC alone clearly catalyzes incorporation of the running- start C but cannot measurably catalyze further extension opposite the lesion (lane 1). Inclusion of either p, y complex (lane 2), SSB (lane 3), or RecA (lane 5) stimulates incorporation of C, but there is essentially no incorporation opposite 57 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. X. Weak UmuD’2 C-catalyzed incorporation opposite X occurs in the presence of P , y complex + either SSB (lane 4) or RecA (lane 6), but lesion bypass is not observed. However, a strong lesion bypass signal was observed once RecA was included along with these other three components (lane 8). Indeed, RecA appears to be the most important stimulatory factor required for translesion synthesis by UmuD’2 C (discussed later in the section). Primer extension is reduced significantly when dCTP is omitted from the reaction (Fig. 3-4, lane 9). The small amount of synthesis occurring in the absence of dCTP can be attributed to UmuD’2 C-catalyzed misincorporation found to occur at undamaged template sites (see section 2.10). Note the presence of a termination band appearing before the second template G (lane 9) that disappears when all four dNTPs are present (lane 8). These data demonstrate that UmuD’2 C is behaving as a DNA polymerase, not a terminal transferase. A mutant UmuD’2 C complex containing UmuC104 (D101N) cannot copy past the abasic lesion (lane 11), thereby confirming that the error-prone polymerase activity is intrinsic to UmuD’2 C and cannot be explained by contamination with some other polymerase, e.g., the recently discovered pol IV [DinB (277), see also Chapter 4]. In support, a UmuD’2 C complex purified from a ApolB AdinB strain exhibits the same lesion bypass polymerase activity (X. Shen & M. F. Goodman, unpublished data). The effect of RecA on UmuD’2 C catalyzed translesion synthesis was investigated by using standing-start kinetic experiments where the efficiency of 58 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. dAMP incorporation opposite a target T site was compared in the presence or absence of RecA (Figure 3-5). All reactions were carried out in the presence of p, y complex, SSB, and ATP. Addition of RecA to the reaction causes a dramatic increase, ~15,000-fold, in incorporation efficiency, as measured by the V m a x/K m value. The increased efficiency is reflected in a marked reduction in Km for dATP; the steady-state Km value is reduced from 1.2 mM to 0.08 pM. This effect is greatly reduced when wild-type RecA is replaced by the mutant RecA1730, with only a 10-fold reduction (data not show). Thus, in agreement with a model proposed by R. Devoret and colleagues (250), it appears that RecA protein serves to target UmuDzC to the single-stranded DNA regions generated by replication complexes stalled at DNA template lesions. Either ATP or the nonhydrolyzable ATP analog adenosine 5'-[vS] thiotriphosphate (ATPyS) is required for translesion synthesis. Interestingly, synthesis in the presence of ATPyS becomes much more processive (Fig. 3-6, compare the ATP panels with ATPyS panels), suggesting that RecA filament disassembly, occurring in the presence of ATP but not ATPyS (171), "extinguishes” pol V activity. Furthermore, when ATPyS replaces ATP in the in vitro reaction, p, y complex can be dispensed with (Figure 3-7, compare lanes 7,8 with lanes 15,16). This result appears to resolve a perplexing difference in requirements for pol V-catalyzed translesion synthesis when comparing our data with recent results of Z. Livneh and colleagues, using a UmuC-MBP fusion protein (225), 59 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. which did not require p, y complex. In our experiments there was 1 RecA per 5 to 15 nt compared to a much higher 5 RecA/nt in Livneh’s experiment, probably sufficient to maintain an intact RecA filament. Thus, RecA’s role may be to activate UmuD’2 C while targeting it to a DNA damage site, while p, y complex could help tether UmuD’aC at the 3’-primer end during the ATP-driven dynamic RecA assembly-disassembly process. The role of SSB was also studied by titrating SSB and measuring the effect on UmuD’aC-catalyzed lesion bypass. All reactions were carried out with non-hydrolyzable ATPyS and in the absence of p, y complex (Figure 3-8). When RecA is absent, UmuD’aC itself catalyzes very weak synthesis. The amount of synthesis greatly increases as the concentration of SSB increases, but primer elongation terminates one base before the lesion (Figure 3-8, left panel). When RecA is present in the reaction, however, both synthesis and lesion bypass are strongly stimulated by addition of SSB. This absolute requirement for RecA and SSB by translesion synthesis suggests that the two proteins might interact with each other and/or with UmuD’aC at the lesion site. When ATP was used in place of ATPyS, the same result was observed except that p and y complex are again required in order to bypass the lesion (data not shown). 2.6 UmuD’2 C competes with pol III for binding primer-3’OH-ends. Kinetic experiments were performed to determine how the presence of pol III a influences UmuD'aC-catalyzed lesion bypass (Fig. 3-9). A running-start reaction, in which a single nucleotide (C) is incorporated before reaching an 60 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. abasic lesion, was used to measure incorporation directly opposite and downstream from the lesion as a function of dATP substrate concentration. RecA, p sliding clamp, y clamp loading complex, and SSB were present in all reactions. In the absence of pol III a, UmuD’2 C incorporates A opposite X, and further extension to the next template site occurs by incorporation of an A A mispair (Fig. 3-9A). In contrast, a HE incorporates a much lower amount of A opposite X, with no observable bypass. The key result is that the rate of UmuD'2 C-catalyzed lesion bypass is reduced significantly as the concentration of the pol III a HE is increased (Fig. 3-9B). It is important to emphasize that we measure lesion bypass synthesis by the amount of incorporation opposite the lesion and extension past the lesion, normalized to the amount of primer extended by incorporation of running-start C to reach the lesion site. Although incorporation of running-start C just before the lesion is strongly stimulated as the concentration of pol III is increased relative to UmuDaC, the efficiency of translesion synthesis is reduced substantially. In other words, the presence of pol III a HE at the primer-3'-end prevents UmuDaC from binding at the lesion site. A Lineweaver-Burk double reciprocal plot of the data is used to illustrate this important point (Fig. 3-9C). We observed that the apparent Km values for incorporation of A opposite X and opposite the downstream A are relatively insensitive to the concentration of pol III a holoenzyme. The apparent Km values 61 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. are roughly the same compared with UmuD’2 C alone (= 12 pM), and are approximately 40-fold lower than pol III a HE (= 500 pM). Similar results were observed when pol III HE was used in place of pol III a HE (267). This apparent noncompetitive inhibition of UmuD’2 C by pol III, with respect to dNTP incorporation at X, implies that UmuD’2 C and pol III are acting independently and competing for the same primer-3’-ends. Thus, whenever UmuD’2 C succeeds in binding to a primer-end and catalyzes bypass synthesis in the presence of pol III, it does so with a characteristically low Km value. Conversely, large reductions in Vm a x occur because there are fewer bound UmuD’2 C-primer- template DNA complexes due to competition by pol III. 2.7 UmuD’2 C catalyzed base misincorporation and mismatch extension at aberrant and normal template sites Contrary to the prevailing two-step model, in which UmuD’2 C was proposed to act as a fidelity-reducing factor for pol III, we have discovered that UmuD’2 C has intrinsic polymerase activity. Experiments were therefore set up to demonstrate the marked low fidelity of UmuD’2 C, in comparison to pol II and pol III, when incorporating at both aberrant and normal template sites, by using just one dNTP substrate (Fig. 3-10). A standing-start protocol was used to measure incorporation at an abasic lesion, X (Fig. 3-10A) or at a normal template T site (Fig. 3-1 OB). All reactions contained RecA, p, y complex, SSB, and ATP. The levels of pol III core (1 nM) and pol II (0.2 nM) were chosen to 62 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. give similar (or greater) amounts of synthesis than UmuD^C (200 nM), on an undamaged DNA template with the four dNTPs present (Fig. 3-1 OB, compare lanes 4 for the three panels). There is essentially no stable incorporation catalyzed by pol III opposite X by using either a single dNTP or in the presence of all four dNTPs (a faint band is seen in lanes A and 4, representing a small incorporation of dAMP). For pol II, faint bands are observed for incorporation of dGMP, dAMP, and dTMP, but not dCMP. Pol II catalyzes a small amount of full-length product in the presence of all four dNTPs, consistent with previous demonstration that pol II can copy past abasic lesions with a substantially higher efficiency than pol III (18). Synthesis carried out with UmuD’2 C in the presence of only dGTP or dATP substrates results in the incorporation of five consecutive G’s or A’s, whereas synthesis with either dTTP or dCTP results in the incorporation of either three T’s or one C (Fig. 3-10A). A much larger amount of synthesis occurs in the presence of four dNTPs (Fig. 3-10A, lane 4). The weak primer extension band observed in the absence of dNTPs (Fig. 3-10A, lane 0) is caused by incorporation of ribo AMP. The first incorporation occurs opposite the abasic lesion. Synthesis taking place at template sites downstream from the lesion corresponds mainly to the incorporation of mismatched nucleotides, followed by extension of mismatched termini and by additional misincorporations. These results stand in marked contrast to the relatively weak primer extension catalyzed by pol III and pol II, and clearly demonstrate the 63 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. remarkable effect of UmuD^C in relaxing the specificity of nucleotide incorporation at aberrant and normal template sites. Given the paucity of incorporation by pol III and pol II on the abasic template, it is important to show that these polymerases carry out normal DNA synthesis on a normal template, in which X is replaced by T (Fig. 3-1 OB). As expected, the predominant reaction for pol III and pol II is the incorporation of dAMP opposite T when dATP is the only substrate present. A small amount dTMP misincorporation is also occurring, possibly by a transient misalignment mechanism involving the downstream template A (14). In the case of UmuD’2 C, either four G’s, four A’s, three T’s, two C’s, or two ribo A’s are incorporated (Fig. 3-1 OB), indicating that the mutagenic Umu complex catalyzes misincorporations and mismatch extensions on normal DNA templates as well as those containing DNA damage. 3. Discussion Despite our ever-increasing understanding of the SOS response and its many repair pathways, little was known about the biochemical mechanisms of SOS mutagenesis (216, 286). Most of our knowledge is based on genetic data which indicate that UmuC, UmuD’, RecA, and pol III holoenzyme are involved in SOS mutagenesis (278), perhaps as a "mutasome" complex (58). The basic principle underlying translesion DNA synthesis is that a replication fork stalls when encountering a DNA damage site, and that SOS-induced UmuC, UmuD’, 64 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. and RecA* proteins interact with pol III to shepherd it past a template lesion, resulting primarily in a base substitution mutation at the lesion site (139). The principal difficulty in investigating the biochemical basis of SOS lesion bypass stemmed from problems in purifying biologically active UmuC protein, which is inherently insoluble in aqueous solution when overproduced (294). H. Echols and coworkers (218) were the first to observe UmuC-UmuD’- dependent lesion bypass by purifying UmuC that was purified in a denatured state in the presence of 8 M urea and subsequently renatured by dialyzing out the urea (211, 294). There were several major difficulties working with renatured UmuC, including small yields of renatured soluble protein, poor signal-to-noise in the bypass reaction, variability in the conditions for bypass: some preparations showing bypass with activated wild-type RecA protein whereas others did not (211). Owing to uncertainties in the biological activity of refolded UmuC protein and to circumvent the problems relating to insoluble UmuC protein, we succeeded in purifying sizable quantities of soluble UmuC tightly complexed to UmuD’ (31, 265). 3.1 Reconstitution of a UmuD’2 C-RecA-dependent SOS lesion bypass system in vitro. Lesion bypass occurs as a two-step reaction with a nucleotide initially incorporated opposite the lesion, followed by extension from a distorted primer terminus. Both steps involve aberrant synthetic reactions catalyzed at greatly reduced efficiencies (about 104 - to 105 -fold) compared with normal synthetic 65 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. reactions, as measured in vitro using Drosophila Pol a (219) and HIV reverse transcriptase (34). In our in vitro system, when pol III HE was used to copy a site-directed abasic template lesion, incorporation opposite and extension beyond the lesion were barely detectable. However, both reactions were stimulated strongly when UmuDaC, RecA, and SSB were included in the reaction. Bypass of an abasic lesion depends on the presence of UmuD’2 C, RecA*, P sliding clamp, y clamp loading complex, and SSB (Fig. 3-2). When either UmuD’2 C or RecA* is excluded from the reaction (Fig. 3-2, lanes 5 and 9), or when RecA1730, a missense mutant refractory to Umu-dependent mutagenesis, is present in the reaction (Fig. 3-2, lane 10), synthesis terminates at one base prior to reaching the abasic lesion, indicating that both bypass steps require the presence of UmuD’2 C and RecA*. Remarkably, however, omitting pol III core had essentially no effect on lesion bypass (Figure 3-2, left panel). These data led us to suggest that UmuD’2 C is a low fidelity DNA polymerase that incorporates a nucleotide opposite a damaged template site and then synthesizes past the lesion (265), although it remained a formal but unlikely possibility that there was a trace contaminant polymerases present whose properties were altered when interacting with UmuD’2 C. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 3.2 UmuD’2 C is an error-prone DNA polymerase, E. coli pol V. Several lines of evidence demonstrated that UmuD'2 C has intrinsic polymerase activity distinct from pol III or pol II. We purified an intact native UmuD’2C complex from cells lacking pol II and containing a temperature- sensitive mutant pol III. UmuD’2 C and pol lilts were separated by Superdex 200 gel filtration, and three column fractions were used to investigate bypass of a site-directed abasic lesion (Fig. 3-3). One fraction (fx 50) contained the pol lilts peak along with a small amount of UmuD’2 C, another (fx 64) contained UmuD’2 C peak with no detectable pol lilts, while the third fraction (fx 56) contained both UmuD’2 C and a small, yet significant, amount of pol lilts. The pol lilts (fx 50), as expected, synthesizes on normal template at 37°C, but its activity is reduced substantially at 47°C. In contrast, UmuD'2 C (fx 64) catalyzes lesion bypass at both permissive and nonpermissive temperatures (Fig. 3-3). Thus, UmuD^C's polymerase activity cannot be explained by the presence of pol III. Contamination with pol II is ruled out because a ApolB strain was used to overexpress UmuD'2 C, and pol I is ruled out because the assays were performed in the presence of large excess of neutralizing antibody directed against pol I. Contamination with the newly discovered pol IV (277) is also excluded because UmuD’2 C expressed from a A polB AdinB strain exhibits the same lesion bypass polymerase activity (data not shown). 67 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. In addition, the mutant UmuD’aC104, which is deficient in SOS mutagenesis in vivo (116, 252), fails to catalyze lesion bypass in vitro (Fig. 3-3, lane 11), proving that the lesion bypass polymerase activity resides in UmuDaC, and can not be attributed to any contaminant polymerases. As the fifth DNA polymerase discovered, we designated UmuD’2 C as E. coli pol V (267). Reuven et al, who took a different approach and purified UmuC as a fusion protein with maltose binding protein (MBP), reported that lesion bypass activity depended on the presence of pol III HE, UmuD’, RecA and SSB (226). After a more careful re-examination, they later observed, in agreement with our earlier work, that their UmuC-MBP protein also has a weak polymerase activity that is stimulated by UmuD’, RecA and SSB (225). 3.3 Pol III and pol V compete for primer-3’-OH ends. Kinetic data show that UmuDaC-catalyzed lesion bypass activity is reduced as the concentration of pol III is increased, a result that cannot be attributed to pol III proofreading (Fig. 3-9). The fact that catalysis by pol III at a lesion site is negligible compared with UmuD'aC enables a straightforward interpretation of the kinetic data. The observation that Vm a x is reduced by addition of pol III while the apparent Km for lesion bypass is essentially the same as for UmuD'aC alone indicates that both enzymes are able to bind primer-3'-ends, providing strong independent evidence that UmuD'aC is a low- fidelity DNA polymerase. 68 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 3.4 Requirement for RecA*, SSB, ( 3 and y complex during pol V-catalyzed lesion bypass UmuD^C-catalyzed lesion bypass requires the presence of activated RecA (RecA*) (Fig. 3-2; & Fig. 3-4), in agreement with genetic data (57,192, 261). However, we found that, in addition to RecA*, lesion bypass also requires ( 3 , y complex and SSB. Indeed, incorporation of a running-start nucleotide at an undamaged template site is stimulated by the presence of each of these accessory proteins (Fig. 3-4), but only when all these proteins are present does lesion bypass occur. We therefore use the term “pol V mutasome (Mut)” to designate this functional multi-protein complex in the vicinity of DNA template lesion sites, in accordance with a previous suggestion made by H. Echols (59). The role of RecA protein - an insight to the targeting mechanism. RecA plays a more direct role in SOS mutagenesis beyond its involvement in cleavage of LexA and UmuD proteins (57, 74, 261). Omission of wild-type RecA or addition of mutant RecA1730 in the lesion bypass reaction resulted in no detectable bypass. More strikingly, kinetic studies revealed that addition of RecA to pol V causes a remarkable, -15,000 fold, reduction in the apparent Km value for dATP incorporation opposite a target T site, but having no detectable effect on the Vm a x (Fig. 3-5). It’s possible that this reduction in apparent Km value actually reflects increased affinity of pol V for DNA templates. Thus, the 69 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. third essential role of RecA in SOS mutagenesis is not only interacting with pol V and targeting it to the DNA template (3, 74), but dramatically increasing the affinity of pol V for dNTP and/or DNA substrates, thereby improving the efficiency of dNTP incorporation. One perplexing question arises from the observations that in the presence of ATP, addition of RecA stimulates pol V’s polymerase activity, whereas in the presence of nonhydrolyzable ATPyS, addition of RecA inhibits pol V’s activity (Fig. 3-7, compare lanes 5, 6 with lanes 13,14). Such inhibition, however, no longer exists when SSB is presence in the reaction (Fig. 3-7, lane 15). One plausible explanation is that although pol V can’t synthesize on a RecA filament, the presence of RecA at the site of polymerization is important for pol V activity. Thus a dissociating RecA filament occurring in the presence of ATP, or a RecA filament that can be gradually replaced by SSB even in the presence of nonhydrolyzable ATPyS, provides the optimal DNA template substrate for pol V. Exactly how RecA interacts with pol V on the template remains to be resolved. SSB protein is essential for pol V’s activity. Addition of SSB greatly increases pol V's polymerase activity. Such a stimulation may be indirect because SSB eliminates DNA secondary structure (134,187), keeps ( 3 clamp from sliding off linear DNA (14), and helps RecA achieve its activated state (142,227). However, the titration experiment (Fig. 3-8) argued that SSB play a Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. more direct role in pol V-catalyzed transiesion synthesis. SSB is known to interact with and stimulate pol III HE (70), pol II (177,178) and other proteins involved in DNA replication and repair pathways (136,142,152). More importantly, SSB was shown to interact with MucB, a homolog of UmuC, in a yeast two-hybrid assay (236). One can therefore speculate that SSB directly interacts with UmuC, thus promoting DNA synthesis and lesion bypass. It’ll be of great importance to test whether SSB interacts physically with pol V and if the interaction depends on the same functional domain that interacts with pol III. One important question remains to be answered, that is, how does SSB function on a RecA-bound DNA template in transiesion synthesis? Although some data suggested that SSB and RecA bind simultaneously to M13 ssDNA (93, 271) and interact while both are bound on DNA (222), it’s still an open question whether or not they interact physically. Our observation that both proteins must be present in order to bypass lesions argues that there exists a highly ordered protein-DNA structure that positions pol V for its optimal activity. In vivo, single-stranded DNA is promptly coated by SSB. Under SOS conditions where there’s higher concentration of ATP and Mg2 + (1 3 2 ) , RecA is able to replace SSB, binding to DNA unidirectionally with a 5’-»3’ polarity (222). One can therefore envision a boundary between SSB and RecA, possibly in the vicinity of lesion sites, to which pol V is recruited. Transiesion synthesis can then be driven by subsequent replacement of RecA by SSB, taking place while both proteins are in contact with pol V. Such a dynamic boundary between RecA and SSB accounts for the dual requirements for SSB and RecA, and is in 71 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. accordance with the observations that (i) pol V preferentially binds to the tip of a RecA filament (71) and (ii) in the in vitro system, addition of SSB before RecA filament formation inhibits lesion bypass (data not shown). Clearly, more experiments are required to dissect the mechanism. Two particular SSB mutants, namely SSB-113, which is normal for DNA binding but displays temperature sensitive DNA replication due to deficiency in interacting with pol III HE (41,173), and SSB-3, which is normal in DNA replication but defective in RecA mediated recombir.ationa! repair (173, 240), may serve to identify or uncouple the roles of RecA and SSB in pol V-catalyzed transiesion synthesis. Another remaining question is what factor(s), if any, target pol V to the lesion site? Using UV irradiated dsDNA, H. Echols and colleagues had reported that RecA bound preferentially to damaged DNA (153, 228). Such preferential binding, however, may actually reflect RecA’s ability to recognize a single stranded feature of DNA caused by damaged nucleotides. Another candidate is SSB protein, whose mammalian counterpart, RPA, binds UV-damaged ssDNA with higher affinity than undamaged DNA (137). In support of this idea, preliminary data from our lab showed that, similar to RecA’s effect on pol V, SSB also causes a 6-fold increase in dNTP incorporation efficiency on damaged DNA template (P. Pham & M. F. Goodman, unpublished data). The biological significance of such a difference remains to be elucidated. Alternatively, contrary to the targeting hypotheses, it’s possible that there is no specific damage recognition factor. Rather, lesions are recognized as single stranded DNA regions by pol V mutasomal proteins. Under this scenario, pol V 72 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. would have similar properties on both damaged and normal DNA templates and, consequently, generate both targeted and untargeted mutagenesis (see next chapter). What is the role of p sliding clamp in transiesion synthesis? In addition to pol III, pol II is also able to utilize p, y complex to achieve high processivity (19). By analogy, a requirement for p, y complex by pol V-catalyzed lesion bypass might be to increase the residence time of the mutasome at the lesion site. Indeed, when present alone in the assay, pol V is extremely weak and strictly distributive, adding only one nucleotide before falling off the DNA template. In the presence of p, y complex, SSB and RecA, however, the processivity of pol V Mut increases to 6 - 8 nucleotides (266). The same result has also been observed with pol IV. Furthermore, similar to RecA's effect on pol V, p and y complex increase the efficiency of pol IV by -3 ,000-fold (266). It thus appears that the P sliding clamp serves as a global processivity factor for all DNA polymerases, with pol I being an exemption. How does p improve the processivity of pol V? Two observations, that transiesion synthesis is inhibited by overproduction of the p subunit (264) and that overexpression of UmuC is poisonous for strains with defective DNA pol III (195), suggested that UmuD’2 C comes into direct contact with the p sliding clamp. This direct interaction, which has now been demonstrated in vitro (260), 73 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. implies that the p clamp may help tether pol V at the 3’-primer end during the ATP-driven dynamic RecA assembly-disassembly process. 3.5 UmuD’2 C is error-prone at both damaged and normal template sites The most noteworthy property of pol V is reflected in its ability to catalyze nucleotide misincorporation and mismatch extension at aberrant and normal template sites. These properties are illustrated vividly in an experiment in which just a single dNTP is present for incorporation at an abasic site (Fig. 3-1OA) or at a normal T (replacing the abasic site), in the same sequence context (Fig. 3- 10B). Pol V stimulates incorporation of each dNTP substrate opposite the lesion (including incorporation of riboA), and causes multiple misincorporations downstream from the lesion, whereas pol III core and pol II catalyze negligible incorporation either at the abasic site or beyond (Fig. 3-9A). in a similar vein, pol V stimulates mis-incorporations opposite a normal template T site and beyond, whereas pol III core and pol II predominantly incorporate only A opposite T (Fig. 3-1 OB). These data illustrate how pol V can be involved in both SOS-targeted and untargeted mutagenesis. In the case of mutagenesis targeted to the site of a lesion, UmuD'2 C catalyzes misincorporation opposite the lesion and subsequent extension past the lesion using an aberrant primer terminus. In a recent study, it was demonstrated that SOS-dependent spontaneous mutator activity reflects the processing of replication errors containing normal bases, 74 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. rather than errors opposite cryptic lesions (67). Indeed, extension of natural base mispairs occurs with efficiencies of about 10'4 to 10'5 compared with extension of correctly matched base pairs (168, 212), and thus presents a strong kinetic block to further elongation. Therefore, pol V may function at any kineticaliy unfavorable primer terminus, not just those at a DNA lesion, with the caveat that both RecA* and SSB are required to bypass the lesion. In summary, we have demonstrated that UmuD’2 C is the fifth DNA polymerase in E. coli whose function is essential for SOS-induced transiesion synthesis. How can these data be reconciled with genetics experiments that have shown a clear requirement for pol III in SOS mutagenesis (24, 89)? Our experiments show that pol V is distributive, having a processivity of 6 to 8 nt under the most optimal conditions, and cannot duplicate the 4.6-Mb E. coli chromosome, a task normally performed by pol III HE. Pol III HE must, therefore, take over from pol V once bypass has occurred. If this interpretation is valid, then one can visualize how pol V might displace pol III core blocked at template lesion, interact with (J , RecA*, and SSB to copy past the lesion, and subsequently be released from the DNA to allow reentry of pol III several bases downstream (Fig. 3-11). Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Figure 3-1. Purification of UmuD’2 C complex from a wild-type strain. Fractions from each purification step were separated on a 12% SDS- polyacrylamide gel and stained with Coomassie Brilliant Blue R-250. Lane M, molecular weight markers, whose molecular weight are given to the left of the gel (in kDa); lane 1, crude lysate; lane 2, polyethyleneimine extract; lane 3, first round of ammonium sulfate precipitation (50% cut); lane 4, second round of ammonium sulfate precipitation (30% cut); lane 5, DEAE chromotography pool; lane 6, phosphocellulose chromatography pool; lane 7, Superdex 75 concentrated pool. Lanes 1 to 5 contain 30 pg protein, and lanes 6 and 7 contain 5 pg protein. 76 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Abasic Site Template 3( 7KB (M13) I 30 nt I 46 nt 50 nt s’ Primer ^ pol III core (1 nM) UmuD’2C (200 nM) UmuD’jC - + + + RecA _ + + - + 1730 SSB, P + y _ + _ + + _ + _ + + <-E nd • m m ^-X-1 P 1 2 3 4 5 6 7 8 9 10 Figure 3-2. Reconstituting in vitro SOS lesion bypass system. Standard polymerization reactions, using a running-start protocol, were carried out in the presence or absence of exogenous pol III core by using combinations of UmuD’2 C, RecA, p , y complex, and SSB. Four dNTPs (100 pM) and ATP (1 mM) were present in all reactions. A 3 2 P-labeled primer was annealed to a DNA template containing an abasic lesion, X (top of the figure), and the replication products were separated in 10% denaturing PAGE gels and visualized by phosphorimaging. Locations of the unextended primer band, abasic (site X), upstream site adjacent to the lesion (X-1), and the end of template are indicated on the right. Lane P contains the primer in the absence of proteins. Additions to the replication reaction mixtures are shown at the top of the gel. 77 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. A Fraction # pol III a ts UmuC 48 50 52 54 56 58 60 62 64 66 68 70 72 74 t42 ^ 'Vr B T A A T C A X G I ^3 2 p Fraction # P 50 56 64 50 56 64 Temperature 37 °C 47 °C Figure 3-3. UmuD’2 C (E. coli polV) has DNA polymerase activity. A. Separation of pol lilts from the UmuD’2 C complex by using Superdex 200 gel filtration from a ApolB dnaE1026\s strain. (Upper) Superdex 200 fractions (30 pi) containing pol III a subunit [designated as a (Left)] were resolved on an 8% SDS/PAGE gel and visualized by chemiluminescent immunodetection by using antiserum directed against a subunit. (Lower) Superdex 200 fractions (20 pi) containing UmuC protein [designated as C (Left)] were visualized on a 12% SDS/PAGE gel stained with Coomassie blue R-250. The presence of UmuC and UmuD proteins was verified by using UmuC and UmuD1 antisera (data not shown). B. Sephadex 200 fractions fx (50) containing predominantly pol lilts; fx (56) containing pol lilts + UmuD'2 C and fx (64) containing UmuD'aC having no detectable pol lilts were assayed for polymerase activity by extension of a 3 2 P- labeled 30-mer primer annealed to a linear M13 DNA template, at permissive (37°C) and nonpermissive (47°C) temperatures. A single running-start base, C, is incorporated opposite G to reach the abasic lesion, X. The left-hand lane contains the primer (P) in the absence of proteins. Each reaction mixture contains 1.5 pi of the indicated Superdex 200 fraction. All reactions were carried out in the presence of pol I antibody, RecA, SSB, p, y complex, 4 dNTPs, and ATP. 78 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Lane P 1 2 3 4 5 6 7 8 9 10 11 UmuD’2C + + + + + + + + + + 104 P + Y + - + - + - + + + + SSB - + + - - + + + + + RecA - - - + + + + + + + Figure 3-4. Protein cofactor requirements for UmuD’2 C (E. coli pol V)- catalyzed lesion bypass. Each reaction in lanes 1-9 contains 1.5 pi of Superdex 200 fx 64 containing UmuD'2 C (having no detectable pol lilts), pol I antibody, and various combinations of RecA, ( 3 , y complex, and SSB, as indicated (lanes 1-9). Running-start reactions were carried out at 37°C, with all four dNTPs present in lanes 1 to 8, but with dCTP omitted in lane 9. Reactions were run in the presence of 5% polyethylene glycol. A portion of the template sequence is shown at the right of lane 9, where X represents an abasic site. The left-hand lane contains the primer (P) in the absence of proteins. Standing- start reactions, in which the first incorporated nucleotide occurs opposite X, were run for wild-type UmuD‘2 C (lane 10) and for the mutant UmuD’2 C104 (D101N) (lane 11), each at a concentration of 200 nM. A portion of the template, shown at the right of lane 11, has the same sequence as the running- start template, but uses a primer terminating one base before the lesion. The asterisk (*) designates a P-label at the 5'-primer terminus. 79 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. pol V pol V + RecA Primer —* j»Tn/, u\ O M W i M ! ! O O O - 1 W M Cn d A T P (|i.M ) S o o § 8 b - bi 0 , 0 a S O ' Fold- Stimulation KmftlM) 1200 0.08 15,000 vm a x (rnin -1 ) 0.7 0.7 1 V m ax/K n. (nM'1 min*1 ) 0.6 x 10- 3 8.8 15,000 Figure 3-5. Effect of RecA on incorporation of correct dAMP at an undamaged template by pol V. Reactions were carried out with undamaged circular ssDNA M13mp7 annealed to a 30-mer 3 2 P-labeled primer. Synthesis by pol V was performed in the presence of p, y complex, SSB and ATP. Apparent Km and Vm a x values were obtained by plotting nucleotide incorporation rate versus dNTP concentration and fitting the data to a saturation curve (rectangular hyperbola) using nonlinear least squares (see Chapter 2 for detail) [courtesy of Phuong Pham (266)]. 80 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 3 2 p" 3 2 p End> Figure 3-6. TLS in the presence of increasing concentration of ATP or ATPyS. All reactions were carried out at 37°C for 10 min in the presence of 2 nM of DNA substrate, pol V (50 nM), RecA (1 pM), SSB (300 nM as tetramer), p (40 nMO and y complex (10 nM). Two primers were used that differ in their relative positions to the abasic lesion as depicted above the gel. The positions of the unextended primer P, the upstream site prior to the lesion (X-1), and the end of template are indicated to the side. 8 1 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Lane 1 2345 6 7 8 9 1 0 1 1 1213 1 4 is 1 6 P / Y - + - + - + - + - + - + -+ - + SSB - + - + - + - + RecA + - + ATP ATPyS Figure 3-7. Role of p, y complex in TLS in the presence of ATP or ATPyS. All lesion bypass reactions were carried out at 37°C for 10 min in the presence of 2 nM DNA substrate, pol V (100 nM), 4 dNTPs (100 pM each) and 1 mM of either ATP or ATPyS. If present, RecA protein was at 1 pM, SSB was at 300 nM as tetramer, and p and y comlex were at 40 nM and 10 nM, respectively. The DNA substrate is a 3 2 P-labeled primer annealed to a 7.2 kb linear single stranded DNA with an abasic lesion as depicted in Fig. 3-2. The unextended primer P, the upstream site prior to the lesion (X-1), and the end of template are indicated to the left. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. SSB RecA Figure 3-8. Effect of SSB on pol V*catalyzed TLS. All reactions were carried out with 2 nM DNA substrate, 100 nM pol V in the presence of 1 mM ATP7S. When present, RecA was at 0.5 pM. The SSB concentrations were at 0, 50, 100, 200,400 and 800 nM. The DNA substrate is a 3 2 P-labeled primer annealed to a 7.2 kb linear single stranded DNA with an abasic lesion as depicted in Fig. 3-2. The unextended primer P, the upstream site prior to the lesion (X-1), and the end of template are indicated to the left. 83 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Reproduced with permission o f th e copyright owner. Further reproduction prohibited without permission. Primer Template dCTP dATP i l / ACCGXACTAATTCTG dATP Enzyme: pol III a (25 nM) PolV + a (25 nM) P0 IV + a (5 nM) Pol V (200nM) B U m u 0 6 > 0 4 0 2 0 2 D 0 400 6 0 0 800 c 5 2 D Uni -at 0 at 0 3 0 4 as 0 2 (M lF tyiN t WAIF] Figure 3-9. Inhibition of pol V-catalyzed transiesion synthesis by pol III. A. Incorporation of dAMP opposite abasic lesion, X, by either pol V, pol III a, or a combination of both proteins was measured by varying dATP concentration, with one base running-start C (20 pM dCTP) inserted opposite template G before reaching X. RecA, SSB, P and y complex were present in the reactions. The dATP concentrations used were 0, 2,10, 50, 200, 500, and 1,000 pM. B. Michaelis-Menten saturation plots of the relative transiesion synthesis rate (v) vs. dATP concentration. C. Lineweaver-Burk double reciprocal plots suggest that pol III behave as a noncompetitive inhibitor of pol V at lesion site. Reproduced with permission o f th e copyright owner. Further reproduction prohibited without permission. dNTP dNTP A Primer 5 ' C B Primer 5 ' — C Template 3 '— GXACTAATTC — 5- Template 3’ — GTACTAATTC — 5- < - End - H < - End a o < x P 4 G A T C 0 4 G A T C 0 4GA TC 0 pol III core (1 nM ) pol II (0.2 nM ) Pol V (200 nM ) c < I- o < P 4 G A T C 0 4 GATC 0 4 GA TC 0 dNTP pol III core (1 nM) pol II (0.2 nM ) Pol V (200n M ) Figure 3-10. Pol V is error-prone on both damaged and normal template sites. Standard standing-start polymerization reactions were carried out by using pol III core, pol II, or pol V in the presence of p, ycomplex, SSB and RecA. A. Reactions carried out by using a DNA template containing an abasic lesion, X. B. Reactions carried out by using a natural DNA template in which X is replaced by T. The lanes labeled as G, A, T, and C denote reactions carried out with a single dNTP substrate, dGTP, dATP, dTTP, and dCTP, respectively. The lanes labeled as 4 and 0 denote reactions carried out in the presence and absence of four dNTPs, respectively. Lane P contains the 3 2 P-labeled primer in the absence of proteins. The abasic lesion containing and natural DNA templates are shown above each gel. Pol III HE P-clamp y^omplex 3’ 5’ core RecA* f t Pol V (UmuD’jC) SSB Pol V Mut 3’ ■ X 5’ 3’. X- A“ • • 5’ Figure 3-11. A model for pol V-catalyzed translesion synthesis, (see text for detail) 86 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Chapter 4 Roles of E. coli DNA Polymerases IV and V in Lesion Targeted and Untargeted SOS Mutagenesis 1. Indroduction 1.1 The UmuC/DinB/Rad30 superfamily Thanks to recent advances in genome projects (1,13,172, 269), it is now clear that UmuC belongs to a large superfamily of proteins that can be broadly subclassified into the UmuC-, DinB-, Rev1-, and Rad30-subfamilies [for recent reviews see ref. (23, 75, 87,101, 291)]. All members of the superfamily share 5 conserved motifs, l-V (Fig. 4-1), and possess the ability to catalyze phosphodiester bond formation. The UmuC subfamily is only found in prokaryotes. Members of the subfamily differ from those of other subfamilies in that their biological function requires UmuD-like proteins (see Chapter 1), and, in the case of pol V, other accessory proteins such as RecA and SSB (225,267). The DinB subfamily is the most conserved, discovered from bacteria to yeast to human (79,199,277). The E. coli DinB [pol IV (277), previously cloned as DinP (200)] is required for untargeted mutagenesis in X phage (30) and, when overexpressed, causes 1,000-fold increase in spontaneous mutation frequency on F episome, most of which are -1 frameshifts (113). Likewise, 87 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. transient expression of mouse DinB1 cDNA in cultured mouse cells results in a nearly 10-fold increase in the incidence of point mutations, among which about 30% were frameshift mutations (199). The Rad30 subfamily is identified only in eukaryotes (164). The Saccharomyces cerevisiae RAD30, encoding Pol r|, is involved in error-free bypass of UV-induced damages, and its mutations render increased UV sensitivity and altered mutability (105,163,229). Pol r| efficiently bypasses a cis-syn TT cyclobutane dimer in an error-free manner, putting two As opposite the lesion sites (106). The human counterpart of yeast RAD30, hRad30A, showed similar activity as pol ti (44, 45,103, 160,161). Patients having xeroderma pigmentosum variant type (XPV) are defective in pol r| activity and predisposed to UV-induced skin cancers (103,161). A second RAD30 homolog in human, hRad30B, has been cloned (164) and shown to be an error-prone DNA polymerase, Pol i (270). Members of the Rev1 subfamily are also only found in eukaryotes. The S. cerevisiae Rev1 protein is not a polymerase. Rather, it has a template- specific deoxycytidyl transferase activity that preferentially inserts a dCMP opposite an abasic site (190). Members of the Rev1 subfamily have been identified in S. pombe, C. elegans and D. melanogaster. Recently, the human Rev1 protein has been shown to possess the same dCMP transferase activity (8,144). 88 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 1.2 Common DNA lesions Three common DNA lesions are pyrimidine-pyrimidone (6-4) photoproducts [(6-4)PD] and cyclobutane photodimers (CPD) occurring at adjacent pyrimidines caused by UV radiation, and abasic (apurinic/apyrimidinic, AP) sites arising from the spontaneous loss of a DNA base, or when glycosylases excise damaged bases or uracil from DNA (145). Structural studies have revealed that, while (6-4) PDs cause substantial alterations in the DNA double-helix, naturally occurring CPDs and AP sites produce little distortions in the B-DNA structure, with the main conformational change occurring at the lesion sites [reviewed in ref. (76) and (91)]. Nonetheless, all three lesions present strong blocks to in vivo (4,141, 239) and in vitro (131, 179) replication [reviewed in ref. (86) and (150)]. These lesions are potentially lethal; yet, when bypassed in vivo and in vitro, they vary in their abilities to generate mutations. CPDs are mostly correctly replicated, the extent and accuracy of bypass depending on the polymerase used and the sequence context. AP sites and (6-4) photoproducts, on the other hand, are largely mutagenic. When AP sites are bypassed, A is preferentially inserted followed by G or T. This “A rule" (256), however, does not hold in S. cerevisiae, where C is the preferred insertion (82). Likewise, a TT (6-4) photoproduct is highly mutagenic in E. coli (141) and mammalian cells (77), but much less so in yeast (81). 89 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 2. Results We studied translesion synthesis efficiencies and base incorporation specificities of the two newly discovered E. coli DNA polymerases, pol V and pol IV, and compared them with the replicative enzyme, pol III. Three types of lesions were used: a TT (6-4) photoproduct, a TT cis-syn photodimer, and an abasic site. The templates were M13mp7 derived single-stranded DNA carrying site-specific lesions 50 to 54 nt away from the 5'-end (see Chapter 2, Materials and Methods). 2.1 Comparison of translesion synthesis by pol V Mut, pol IV and pol III HE. The forms of the enzymes used to compare translesion synthesis (TLS) efficiency were: pol V Mut (or mutasome) consisting of pol V (UmuD’2 C), RecA*, ( 3 , y complex, and SSB; pol IV (DinB) together with p, y complex and SSB, and pol III HE (holoenzyme of pol III core, p, y complex + SSB). As seen in Fig. 4-3, pol V Mut catalyzes efficient bypass of all three lesions within 30 s, with estimated lesion bypass rates of ~ 30, 20, 80%/min for the TT (6-4) photoproduct, TT photodimer, and abasic moiety, respectively. No detectable bypass is observed in 30 seconds for pol III HE or pol IV, the latter showing incorporation opposite each of the lesions but with no further extension. At later time points, > 8 min, faint bypass product bands appear for pol III HE and pol IV, with bypass rates of 1.2 and 0.7 %/min, respectively for 90 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. TT (6-4) photoproduct; 1 and 0.5 %/min forTT dimer, and 0.4 and 0.1 %/min for the abasic moiety. In contrast, extensions from a normal template position (T site, Fig. 4-3, left gel) are - 88, 98, and 68% at 30 s for pol V Mut, pol III HE, and pol IV, respectively. Enzyme levels were chosen to allow similar primer utilization on undamaged DNA templates; roughly 50% of each primer being extended at 16 min (Fig. 4-3, left gel). 2.2 Incorporation specificities opposite lesion sites. A gel kinetic assay (51) was used to measure nucleotide incorporation specificities for the three polymerases opposite both positions of a TT (6-4) photoproduct. A 3 2 P-labeled primer is extended by incorporating a single “running-start” nucleotide to reach the 3’-T of the (6-4) photoproduct (Fig. 4-4A); and standing start reactions were used for 5’-T (Fig. 4-4B). Both pol V Mut and pol IV have no measurable 3’-exonuclease proofreading activity, and are therefore compared with pol III a HE, the proofreading-deficient form of the pol III HE. We found that, for pol V Mut, the Vm a x /Km value for insertion of Q opposite the 3’-T site is 6-fold higher that for insertion of A (Fig. 4-4A). In contrast, pol III a HE and pol IV favor incorporation of A over G by 27- and 3.3- fold, respectively (Fig. 4-4A). At the 5’-T, pol V Mut incorporates A by about 8- or 13-fold over G, when either A*T or G*T primer ends are extended, respectively (Fig. 4-4B). Incorporation of G is not detectable at the 5’-T site with either pol III a HE or pol IV (data not shown). Our data for pol V Mut agree Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. qualitatively and quantitatively with in vivo mutational data showing increased 3’ T -> C transition mutations with a much smaller increase in 5’-T mutations (141, 247) (Table 4-1). The absence of detectable amounts of incorporation of either C or T at either 3’- or 5’-T site by pol V is consistent with the absence of transversions in vivo (see Fig. 4-6A). Similar kinetic analyses were performed for the other two lesions. In the case of a TT cis-syn photodimer, all three enzymes incorporate A in favor of G at both 3’- and 5’-T sites, with pol V Mut being ~ 100-fold more efficient than pol IV and pol III HE, as indicated by the individual Vm a x /Km values (see Fig. 4-5A & B for pol V and III; pol IV, P. Pham and M. F. Goodman, unpublished data). Incorporation of T or C is not detectable for all three polymerases (see Fig. 4-6B for pol V; pol III, data not showm; and pol IV, P. Pham and M. F. Goodman, unpublished data). Similarly, in accordance with the “A-rule” (256), A is favored for incorporation opposite an abasic site, with pol V being 1000-fold more efficient than pol III (Fig. 4-7). Table 4-1 summarizes the pol V incorporation specificity data from kinetic measurements, and compares it with the mutational spectra reported by two other research groups (138,139,141, 247). Thus, our in vitro kinetic data agree remarkably well with in vivo mutational data for all tested lesion sites. 2.3 Incorporation fidelities on normal template sites A standing-start gel kinetic assay (51) was used to determine nucleotide incorporation fidelity for three polymerases - pol III a HE, pol IV and pol V - on 92 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. undamaged DNA template G, A, T and C sites. Time course experiments were performed prior to the kinetic measurements to ensure that primer utilization remained below 20% to satisfy single-completed hit conditions (51) (data not shown). The misincorporation efficiency, fin e , is measured as the ratio of Vm a x /Km values comparing wrong to right nucleotides (see Chapter 2, Materials and Methods). The results are represented in Table 2. For pol V Mut, we found misincorporation efficiencies ranging between 1 to 5 x 10'3 for G*T, T«G, G*G, A*G and T«T mispairs; 3 to 7 x 10"4 for G*A, C*A, C*T, T*C, and A»C mispairs; 7 x 10'5 for A*A mispairs; and < 10'5 for C*C mispairs. These data are comparable with those obtained by another group from a gap-filling experiment (157). For pol IV, the error frequencies ranged from - 2 x 10'3 for G*G to 3 x 10'5 for C«C mispairs. However, the G«G mispair is unlikely to be caused by direct misincorporation of dGMP opposite G, but rather by dNTP-stabilized misalignment (14) whereby incorporation of G occurs opposite a template C base located immediately downstream from G. This result is consistent with pol IV’s propensity for catalyzing -1 frameshift errors (277). For comparison with pols IV and V Mut, the proofreading-defective pol III a HE makes base substitution errors in the 10"4 to 10‘6 range, consistent with previous measurements (14). in conclusion, pol V Mut and pol IV tend to make more errors, about 100- to 10-fold higher, respectively, than the proofreading deficient pol III a HE, when acting on normal template DNA. 93 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. 3. Discussion Persisting lesions in DNA greatly impede DNA replication and ultimately induce the SOS response. The latter is accompanied by an increased occurrence of mutations, most of which occur at damaged template sites and are hence referred to as “SOS targeted mutagenesis”. In addition, mutations also occur in the apparent absence of DNA damage, termed “untargeted mutations” (78, 285, 289). Both targeted and untargeted SOS mutagenesis require the function of UmuD'C proteins (pol V) (64, 67). However, the recent discovery of polymerase IV (277), which has been linked to untargeted mutagenesis in X phage and F’ plasmid (30,113), raises an interesting question: what are the biological roles of the two inducible DNA polymerases following SOS induction? We addressed the question by analyzing the efficiencies and specificities of the two enzymes in copying three common lesions as well as normal DNA template sites, and compared them with that of the replicative pol III. As expected, pol V Mut efficiently bypasses all three lesions, with bypass rates of - 30, 20, 80%/min for TT (6-4) photoproduct, TT cis-syn photodimer, and abasic site, respectively. In contrast, pol IV and pol III are essentially blocked, although they can insert nucleotides opposite the lesion sites very inefficiently (Fig. 4-3). When we measured the specificity of nucleotide incorporation opposite the 3’-T of TT (6-4) photoproduct, we found that pol V favors incorporation of G over A by 6-fold, whereas both pol IV and pol III favor 94 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. incorporation of A (Fig. 4-4). This result is consistent with the in vivo observation that mutations occur predominantly at the 3’-T site of a TT (6-4) photoproduct, with 5’-T mutations occurring much less frequently (76). Furthermore, the pol V incorporation profile opposite all the lesion sites correlates closely with in vivo mutation spectra (Table 4-1). We therefore conclude that pol V Mut, but not pol IV or pol III, is responsible for SOS mutations targeted to DNA damage sites, based on its high translesion synthesis efficiency and recapitulation of in vivo mutational specificity. We also measured the fidelity of pol V Mut, pol IV, and pol III a HE at undamaged DNA template sites. Our results indicate that, in contrast to proofreading-defective pol III a HE, which copies DNA with error rate of 1C4 to 10*6 , both pol V and pol IV misincorporate dNMPs with a frequency of 10'3 to 1C4, with pol IV exhibiting about 5- to 10-fold higher fidelity than pol V Mut (Table 4-2). It is likely that both enzymes function in untargeted mutagenesis, as implicated by their low fidelities. Vet, the role of pol IV in SOS mutagenesis needs to be clarified. Ren et al. recently reported that overexpression of DinB confers a striking mutator effect detected as spontaneous forward mutation to rifampicin resistance (224). Both pol V and pol IV exhibit enhanced ability to extend mismatched primer ends, the latter via a template misalignment mechanism (265, 267,277). Whether they operate on different primer/template structures, or they perform redundant functions on undamaged DNA in vivo, remains to be determined. A systematic characterization of the extension 95 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. efficiency from various primer/template termini may give better clues to their biological functions on undamaged DNA templates. Several pathways may result in untargeted mutagenesis in vivo. Mutations may arise when a polymerase act on ssDNA regions that arise during SOS-induction. Such regions may also form when replication is interrupted at secondary structures in DNA, or during the process of recombination or transposition. A good candidate working under these circumstances would be pol V, since SSB and/or RecA are likely involved in these processes and to recruit pol V. Alternatively, mutations are generated when a mispaired primer terminus is extended at a stalled replication fork. Stalling of pol III HE can be caused by mismatched or misaligned primer-ends that cannot be proofread, a suggestion supported by in vitro properties of the pol III HE (14). Perhaps pol IV and pol V are required to alleviate stalled replication forks on undamaged DNA. In summary, error-prone pol IV and pol V may have complementary yet overlapping roles in E. coli, with pol V Mut catalyzing TLS while pol IV and pol V extending aberrant primer-3’-ends. As pol IV and pol V Mut are both poorly processive and dissociate after relieving a blocked replication fork, pol III HE can then replace the errant polymerases to complete chromosomal replication. To date, all the characterized UmuC/DinB/Rad30 members exhibit low fidelities on undamaged DNA templates, with UmuC and DinB in the range of 10'3 and I ff 4, respectively [this work, (157) and (104)] and Rad30 in the range of 10‘2 to 10*3 (102, 270,282). It is not surprising that these enzymes lack the Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. three features that lead to highly accurate DNA duplication as exemplified in replicative polymerases: strong discrimination between correct and incorrect base pairs during polymerization, 3’-*5’ proofreading, and low efficiency in extending mismatched primer termini (see Chapter 1, section 1-3). In addition, all the enzymes are very distributive, synthesizing one to a few base pairs before failing off the template. Such properties suggest that they only function under special circumstances where the replicative polymerases are highly disfavored, for instance, at stalled replication forks at lesion sites. The enzymes, however, differ in their ability to bypass DNA lesions. Both E. coli DinB and human DINB1 are unable to bypass TT (6-4) photoproducts, TT cis-syn photodimers, and abasic sites [this study and (104)]. On the other hand, members of Rad30 and UmuC subfamilies are capable of bypassing various lesions. Yeast pol t) is able to incorporate C, and less efficiently A, opposite 8-oxoguanine, and further extend the 8-oxoG*C or 8-oxoG*A termini (296). It can also incorporate opposite an AAF bulky lesion or an abasic site, but fails to extend further (296). In addition, human pol r\ catalyzes translesion synthesis past bulky oxaliplatin and cisplatin GpG adducts more efficiently and less accurately than that seen for pol £, pol [3 , and pol y (273). Both yeast and human Pol r| efficiently bypasses TT cis-syn photodimers in an error-free manner (106,161). Interestingly, E. coli pol V is the only polymerase reported to date that bypasses a TT (6-4) photoproduct. As discussed in Chapter 1, despite their structural diversities, all polymerases (especially those involved in replication) share similar 97 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. mechanisms to achieve high replication accuracy (58, 254,255). One of the critical steps is the demand for the precise geometry of Watson-Crick base pairs by polymerase active sites. The low fidelity of the newly discovered polymerases in the UmuC/DinB/Rad30/Rev1 superfamily suggests that, compared with other replicative enzymes, these polymerases have more flexible active sites that can accommodate incorrect base pairs. The ability to synthesize opposite and extend from template lesions therefore stems from an increased tolerance of the active sites to distorted DNA structures. It seems reasonable to speculate that the active site of Rad30 is more flexible than that of DinB. An interesting case is pol V, which by itself cannot bypass lesions but does so in the presence of RecA, SSB, p and y complex. Do these cofactors bring about any conformational changes in the enzyme? Future incorporation studies using nucleoside analogs, such as difluorotoluene nucleoside triphosphate (dFTP)(180,181) and pyrene nucleoside triphosphate (dPTP) (162), may shed light on the structural properties of these error-prone polymerases. Furthermore, studies are under way to determine crystal structures for several of these enzymes which should provide invaluable insight into their underlying”fidelityn mechanisms. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. motif concenaua n i n i v v I Y « A H | | S *O E 11 H hH il I H hH i I Pol V Ee UmuC Pol i hRAD30B Pol 11 Sc Rad30 HRAD30A Sp SPBC16A3.11 hDINBI Ce F22B7.6 Pol IV Ec DinB Sc Ravi Sp SPB1347.01c Ce ZK675.2 C,HC CjHC , x y z i io ■ " O a c i w H o @ m o BRCT (870) (598) (351) (985) (935) (1027) Figure 4-1. The UmuC/DinB/Rad30/Rev1 DNA polymerase superfamily. Highly conserved motifs l-V containing putative catalytic residues and helix- hairpin-helix DNA binding domains are denoted by Roman numbers. UmuC and hRad30B have unique C-terminal sequences. The DinB-specific sequences are indicated by x, y, and z. Zinc binding motifs are indicated as C2 H2 or C2 Hc. Numbers in parentheses represent protein length in amino acids [Modified from ref. (101)]. 99 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Reproduced with permission o f th e copyright owner. Further reproduction prohibited without permission. o 3’ H-N H-N i Figure 4-2. Structures of three template lesions. A. Synthetic abasic lesion (tetrahydrofuran moiety). B. TT (6-4) pyrimidine-pyrimidone photoproduct. C. TT cis-syn cyclobutane photodimer. 8 Reproduced w ith permission o f th e copyright owner. Further reproduction prohibited without permission. End-> Normal Template 3 2^ TATGAG------- pol V Mut pol III HE pol IV iiil •»S*B (6-4) Photoproduct 3 2 p — n ___ ATTATG------ pol V Mut pol III HE pol IV cis-syn Cyclobutane Dimer 32 p ^ --- A ATTATG------- pol V Mut pol III HE pol IV iitu ■ Abasic Lesion 3 2 p GXACTA------ pol V Mut pol III HE pol IV » s 4 " r? n * r - is s if P B fl ■ ■1 '! ■ S J 1 * (Min) 0.5 32 0.5 32 0.5 32 0.5 32 0.5 32 0.5 32 0.5 32 0.5 32 0.5 32 0.5 32 0.5 32 0.5 32 Figure 4-3. Comparison of translesion synthesis by pol III HE, pol IV and pol V Mut. Normal and translesion synthesis by pol III core (1 nM), pol IV (25 nM) and pol V (25 nM) were carried out in reactions containing 2 nM primer-templates (shown at top), 1 mM ATP, 300 nM SSB, 40 nM p-subunit and 10 nM y-complex. RecA protein (1 pM) was present in pol V reactions to give mutasome, pol V Mut. Reactions were initiated by adding the polymerase and four dNTP’s (100 pM each) - see Methods. P indicates the location of the non-extended primer. The locations of a TT (6-4) photoproduct, TT cis-syn cyclobutane dimer and abasic sites are indicated by the square bracket, diagonal bracket and symbol X, respectively. The lesion bypass rate is calculated as the fraction of primers extended beyond the lesion per min divided by the total amount of primer extended (see text). V ) o < g •a < H a. H Z ■ O s id cd U Q . .§ S 1: 31 0 0 m cd id CN ID r» o x CN X © 0 0 o cd i d —I vO cd r - cd 0 0 CD CD CD CD X CN CD c * i o X C " 0 0 t" CN X cd rf n o x CN r-‘ © vO S .= c • — t U J > > c o at I 102 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Reproduced w ith permission o f th e copyright owner. Further reproduction prohibited without permission. 4-4 B. 32p dNTP ^ —ATG I t -TA’ JJATGAG' 3 2 p dNTP ATA l / TATTATGAG' u dATP dGTP dGTP dATP dNTP (jiM ) pol V Mul dATP dGTP dATP pol V Mut dGTP Km (nM) 1.8 16.0 2.0 9.0 VnuuOnin'1 ) 7.3 5.0 9.1 5.8 Vm „/K r o (fiM '1 min'1 ) 4.0 0.3 4.4 0.6 Ratio 1 3 1 8.0 1 Figure 4-4. Incorporation of A compared with G opposite a TT (6-4) photoproduct. A. The incorporation kinetics of either A or G were measured opposite the 3’-T site of a TT (6-4) photoproduct using pol V Mut, pol IV or pol III aHE. To reach the target site, a running-start T (10 pM dTTP) is incorporated opposite A as shown in sketch at top. The concentrations of dNTP = dATP or dGTP for incorporation opposite the target site, were varied. B. The incorporation kinetics of either A or G were measured opposite the 5’-T of a TT (6-4) photoproduct using pol V Mut to extend a primer terminating with either G (left gel) or A (right gel) situated opposite the 3’-T of the TT (6-4) photoproduct. The kinetic analysis, summarized in Chapter 2, is described in detail in Ref.(51). Relative efficiencies of incorporation of dATP vs. dGTP (ratios shown at the bottom of the tables) are fold-differences between V m ax/K m for - incorporation of dATP compared to dGTP. Reproduced w ith permission o f th e copyright owner. Further reproduction prohibited without permission. 4-5A. 3 2 P - - P dNTP (pM) dTTP Primer Template dNTP TATTATGAG 5’ S 8 8 8 J B 01 ° 8 8 8 8 ^ ^ dATP dGTP n l U - ‘ A - ‘ M A M - ‘ A O I O O Q O O O I Q O O O © ° dATP dGTP pol V Mut dATP dGTP pol III a HE dATP dGTP Km (pM) 33.8 ±1.7 36.2 ± 8.2 24.0 ND V m a x 4.1 ± 0.3 0.09 ± 0.005 0.04 ND Vmax/Km ruM n 1.2 x 10'1 2.5x10 3 1.7x10 3 Ratio 48 1 8 2000 Reproduced with permission o f th e copyright owner. Further reproduction prohibited without permission. 4-5B. 32 p 5’ 3’ dNTP ATA % TATTATGAG 5’ 32 p 5’ 3’ dNTP ATG £ TATTATGAG V 5' T A 3 2 p ' dNTP O N O l -> M O l -1 M < B ° 8 8 8 8 8 dATP (nM) Km(nM) V m a * (min1 ) Vm ai/Kjj, (fiM 1 min1 ) Ratio O M o i r j M t n — JO Ol O O C J I O u i dATP 0.2 ± 0.04 14.0 ± 2.0 87.5 20 dGTP (HM ) dGTP 2.0 ± 0.4 8.6 ± 6.8 4.6 1 o i o t n ^ i o u t ^ i o tn o o cn o o “ o o o g dATP (nM) dATP 0.3 ± 0.3 8.3 ± 4.8 60.8 o io oi rt ro c n — . U , f° O i o o i dGTP (HM ) dGTP ND ND S Figure 4-5. incorporation of A compared with G opposite a TT cis-syn photodimer. A, The incorporation kinetics of either A or G were measured opposite the 3’-T site of a TT cis-syn photodimer using pol V Mut or pol III aHE. B, The incorporation kinetics of either A or G were measured opposite the 5’-T of a TT cis-syn photodimer using pol V Mut to extend a primer terminating with either G (left gel) or A (right gel) situated opposite the 3’-T of the TT cis-syn photodimer. The kinetic analysis, summarized in Chapter 2, is described in detail in Ref.(51). Relative efficiencies of incorporation of dATP vs. dGTP (ratios shown at the bottom of the tables) are fold-differences between Vm ax/Km for incorporation of dATP compared to dGTP. S Reproduced with permission o f th e copyright owner. Further reproduction prohibited without permission. J - T S ™ 3------ TA'JJATG AG------- 5 - dNTP (pM) dTTP dCTP pol V Mut B. dj T^ d N T P 3 TATTATGAG 5 ' 1 - v ■ T A dNTP (pM) dTTP dCTP pol V Mut J2p dNTP 5’ ATG i f 3 ' TA'JJATGAG 5’ I T A dTTP dCTP pol V Mut 3 2 P dNTP 5’ ATG i f 3 TATJATGAG------- 5' A dTTP dCTP pol V Mut Figure 4-6. Incorporation of T or C opposite template lesion sites. A, Incorporation of either T or C opposite the 3’- or 5’-T of a (6-4) photoproduct by pol V. B, Incorporation of either T or C opposite the 3’- or 5’-T of a cis-syn - photodimer by pol V. The template sequences are indicated on top of each gel. Reproduced w ith permission o f th e copyright owner. Further reproduction prohibited without permission. Primer Template 3’ “ df 'F P dNTP CGXACTAG 5’ j & > 4 m m ft* # n m ** m m m *m m m m dNTP (pM) o “ 1 0 o i ro ^ o o £ w o g g 8 8 - - > dATP ro oi - » • ro o o ro cn o o o o o k, o o o o - t ro < n - t ro 4* O D 8 8 8 ro cn o cn dGTP dATP to O) to -U CD S wo8 8 8 8 dGTP pol V Mut dATP dGTP pol III a HE dATP dGTP Km; V m a x V m a x /K m (mM-M Ratio 18.9 ± 1.0 23.9 ± 2.4 5 47 ± 79 781 8.5 ± 0.6 6.1 ± 0 .9 0.3 ± 0.07 0.04 0.45 0.25 5.5 x 10 4 5 .4 x 10 s 1.8 10 Figure 4-7. Incorporation of A compared with G opposite an abasic site. The kinetic analysis, summarized in Chapter 2, is described in detail in Ref.(51). Relative efficiencies of incorporation of dATP vs. dGTP (ratios shown at the bottom of the tables) are fold-differences between Vm a x/K ,n for incorporation of dATP compared to dGTP. O 5 Reproduced with permission o f th e copyright owner. Further reproduction prohibited without permission. Table 4-1. Pol V Mut insertion specificity at DNA template lesions: comparing in vitro and in vivo data TT (6-4) Photoproduct TT cis-syn Photodimer Abasic Lesion Opposite 3’ T Opposite 5’ T Opposite 3’ T Opposite 5’ T In vitro in vivo kinetics 1* 2f In vitro in vivo kinetics 1* 2t in vitro kinetics in vivo 1* 2f In vitro kinetics in vivo 1* 2f In vitro kinetics in vivo § A 14.3 1 1 27 92.3 94 98 98 94 98 95.2 99 98 63.6 54-80 G 85.7 87.5 69 7.7 2 2 2 1 4.8 <1 1 36.4 15-20 The data represent the percentage of either A or G incorporated opposite each template lesion site. * From ref. (141) t From ref. (247) $ From ref. (138) § From ref. (139) Reproduced with permission o f th e copyright owner. Further reproduction prohibited without permission. Table 4-2. Fidelity of pol V Mut, pol IV, and pol III a HE by steady state kinetics pol V Mut pol IV pol III a HE TP*Template V m a x / K f ln (pM ’min1 ) f. * • m e V m a j/K fn (pM ’min1 ) f * • m e V m a > /k m (pM ’min'1 ) f * • m e dGTP*G 0.23 2.7 x 10 3 0.047 1.7 x 10 3 0.00073 2.5 x 10 dATP-G 0.11 1.3 x 10 3 0.018 6.7 x 10 4 0.0029 1.0 x 10 dTTP-G 0.41 4.8 x 10 3 0.023 8.5 x 10 4 0.014 4.8 x 10 dCTP*G 86 1 27 1 29 1 dGTP»A 0.033 3.3 x 10 4 0.0031 1.5 x 10 ^ 0.0038 3.2 x 10 dATP*A 0.0070 7.0 x 10 5 0.0011 5.2x10 5 0.0011 9.2 x 10 dTTP*A 100 1 21 1 120 1 dCTP-A 0.055 5.5x10 4 0.0020 9.5x10 5 0.019 1.6 x 10 dGTP*T 0.91 2.4 x 10 3 0.0027 3.6 x 10 4 0.0035 2.5 x 10 dATP*T 380 1 7.6 1 140 1 dTTP.T 1.4 3.7x10 3 0.00067 8.8x10 5 0.017 1.2x 10 dCTP«T 0.31 8.1 x 10 4 0.0017 2 .2x 10 4 0.0069 4.9 x 10 dGTP«C 360 1 12 1 56 1 dATP-C 0.19 5.3 x 10 4 0.0016 1.3 x 10 4 0.0040 7.1 x 10 dTTP.C 0.26 7.2x10 4 0.0017 1.4 x 10 4 0.00029 5.2 x 10 dCTP-C nd t <1.0x10 5 0.00043 3.6 x 10 5 0.00016 2.9 x 10 8 * The nucleotide misincorporation ratio, fm c = (Vfn a x/KI T 1 )w/(VlT )a x /K m )R , where W and R refer to incorporation of either a wrong or right nucleotide. The sensitivity of the assay is dependent on polymerase activity, i.e., (Vm ax/Km )R , and can detect a larger misincorporation range for pol III a HE (~10'6 ) compared to pol V Mut (< 10'5 ). The S.E. values for fin e are ±30%. t nd refers to “not detected”. (Fidelity data for pol IV at target A site and for pol III a HE at target C site, courtesy of Xuan Shen; Fidelity data for pol IV at target G site and for pol III a HE at target G site, courtesy of Phuong Pham.) Chapter 5 Conclusions and Perspectives DNA damage arising from endogenous or exogenous sources presents a great challenge for the survival of all organisms. In response, cells possess a variety of repair mechanisms to restore altered genetic information. Situations nonetheless can arise where some lesions may escape repair processes and, consequently, DNA replication is blocked when a DNA polymerase encounters template lesions. Additional measures have to be taken to avoid such deleterious effects. One such strategy, translesion synthesis, enables direct incorporation of a nucleotide opposite a template lesion and subsequent chain elongation. Because of the noncoding or miscoding nature of the lesions, this type of DNA synthesis is error-prone and leads to mutations. Genetic studies revealed that, in E. coli, error-prone translesion DNA synthesis depends on the UmuD’2 C complex, activated RecA protein (RecA*) and the replicative polymerase III. The molecular mechanism of the process, however, has remained a mystery for more than two decades due to the difficulty in obtaining soluble, active UmuC protein. Four years ago, our lab successfully purified a native UmuD’2 C complex that binds cooperatively to ssDNA and inhibits RecA-mediated recombination in vitro (31). Using this active UmuD’2 C complex, we set up an in vitro lesion bypass system in an effort to elucidate the biochemical mechanism of the UmuD’2 C-mediated SOS no Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. mutagenesis. What we discovered is remarkable, namely that UmuD’2 C is a bona fide DNA polymerase that not only efficiently bypasses common template lesions but also functions on normal templates in an error-prone manner. 1. Summary of this dissertation Several lines of evidence were presented in Chapter 3 that led to the conclusion that UmuD’2 C is a bona fide DNA polymerase, E. coii pol V: (i) Purified UmuD’2 C complex catalyzes efficient lesion bypass in the absence of exogenous polymerase. Such activity can’t be attributed to contamination by any other polymerases because the UmuD’2 C complex purified from a ApolB dnaE1026 strain has the same activity at nonpermissive temperaturea where the temperature-sensitive pol III is inactivated, and contamination of pol I is ruled out using neutralizing pol I antibody, (ii) A mutant UmuC protein, UmuC104, fails to catalyze translesion synthesis although it can complex with UmuD’. (iii) When both are included in the in vitro assay, UmuD’aC competes with pol III for free primer-3’-ends, indicating that instead of acting as a cofactor of pol III, as suggested by the two-step model, UmuD’2 C functions independently as a polymerase. We observed that translesion synthesis depends on the function of a multiple protein complex, pol V mutasome. Pol V by itself is a very weak polymerase whose activity is stimulated upon the addition of each of the following factors: RecA, SSB, and p, y complex, but lesion bypass only occurs when all the proteins are present. Omitting RecA or addition of a mutant RecA, in Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. RecA1730, abolishes bypass ability in the in vitro assay. Both in vivo and in intro studies have shown that RecA interacts with UmuD’aC. This observation led to the suggestion that RecA participate directly in translesion synthesis by targeting UmuD’zC to the lesion site (31,74,250). Using kinetic measurements, we further observed that RecA dramatically increases pol V’s affinity for dNTP and/or DNA substrates, as indicated by ~15,000-fold decrease in Km values. Therefore the proposed third role of RecA in SOS mutagenesis is not only bringing pol V to the template target site, but markedly improving the enzyme's efficiency for nucleotide incorporation by increasing the affinity of pol V for primer/template DNA. The role of SSB in the process, however, is less clear. It’s possible that SSB interacts directly with UmuD^C during transiesion synthesis, analogous to its interaction with pol III holoenzyme during replication. Whether SSB serves as a lesion recognition factor, or if it provides specificity by interacting with RecA, remains to be established. We examined the efficiency of pol V in bypassing three common UV lesions: pyrimidine-pyrimidone (6-4) photoproducts, cis-syn cyclobutane dimers, and abasic sites. Pol V bypasses all three lesions within 30 seconds, with bypass rates of ~ 30, 20, 80%/min for the TT (6-4) photoproduct, TT photodimer, and abasic moiety, respectively. In contrast, pol III and pol IV are both effectively blocked by the three lesions, with faint bypass bands only seen after 8 minutes. Incorporation specificity studies further revealed that, when copying the 3’-T of a TT (6-4) photoproduct, pol V favors incorporation of G over A by 6-fold, whereas pol III and pol IV incorporate A almost exclusively. Such a 1 1 2 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. "signature” T-»C transition agrees with in vivo mutational spectra, suggesting that pol V is responsible for SOS mutations targeted to DNA damage sites. We also measured the nucleotide incorporation fidelity of pol V when copying undamaged DNA template sites, and compared it with those of pol III and pol IV. We found that pol V is error-prone on normal DNA as well, with misincorporation frequencies ranging from - 10'3 to 10‘4. Pol IV is less error- prone than pol V, with error rates varying from - 10'4 to 10'5 . The proofreading- defective pol III a HE, on the contrary, makes base substitution errors in the 10'4 to 10'6 range. Pol V and pol IV also exhibit an enhanced ability to extend mismatched primer ends. As an increase in untargeted mutations is likely to result from the combined effects of a relatively high error rate followed by efficient mismatch extension, our data suggest that both pol V and pol IV are capable of making mistakes on normal templates. In light of their biological functions in generating untargeted mutations, genetic evidence has demonstrated that pol V is required for untargeted SOS mutagenesis. However, a clear role of pol IV in untargeted mutagenesis remains to be determined. 2. A few further thoughts Two sides of a coin - Temporal actions of the inducible polymerases after UV irradiation. It is intriguing that E. coli possesses three DNA polymerases - pol II, IV, and V - that are inducible by DNA damage. Although the functions of these enzymes are just coming to light, it is not surprising that all three polymerases are, in one way or another, involved in rescuing cells from Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. the deleterious effects of stalled replication forks (48). The three enzymes, however, are differentially induced after damage occurs. Both pol II and pol IV are induced early, about 4 to 5 minutes after SOS induction. Our lab recently reported that pol II plays a pivotal role in “induced replisome reactivation” or “replication restart” (221). In wild-type cells, following a transient arrest after DNA damage, DNA synthesis resumes at a normal rate within 5 minutes (112, 287). However, the resumption of DNA synthesis was delayed by -50 minutes in ApolB cells, and the delay was prolonged to -100 minutes in a AumuDC ApolB double mutant. The double mutant also showed decreased survival to UV irradiation, compared with either of the single mutants (221). These data suggested that pol II and pol V function in different pathways that complement each other. Furthermore, the two pathways are likely to be sequential, as the 50-minute recovery in DNA synthesis coincides with the appearance of UmuD’aC complexes in vivo (250, 251). In support of the hypothesis, no delay was observed in a ApolB recA730 strain, which constitutively expresses pol V activity (S. Rangarajan & M. F. Goodman, unpublished data). It thus appears that cells possess two distinct temporally-spaced pathways, with pol II involved in rapid, error-free replication restart beginning almost immediately after UV irradiation, and pol V catalyzing error-prone translesion synthesis that coming into play almost an hour later (83). Much less is known about the role of pol IV in DNA repair or damage tolerance mechanisms. Pol IV is induced 15-fold by DNA damage (30). When overexpressed, pol IV causes increased untargeted mutagenesis, 11 4 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. predominantly transversions and -1 frameshift mutations, on F plasmid (113). It’s likely that pol IV is activated early in SOS response, but exactly when and how it functions remains to be addressed. With the biochemical properties of pol IV being characterized, and more genetic experiments on the way, a well- defined cellular function of pol IV can be expected within the near future. Survival vs. evolution ~ purposeful conservation of the UmuC superfamilv. There has been a debate as to what the purpose of SOS mutagenesis is. M. Radman and H. Echols proposed the idea of inducible evolution (76). Such a notion seems less satisfactory since most of the induced SOS mutations are actually deleterious to cells and therefore lack advantage for environmental selection. The fact that the umu operon is tightly controlled at the transcriptional level and that posttranslational cleavage of UmuO is much slower than LexA (293) suggests that pol V-catalyzed error-prone translesion synthesis is the last resort for the cell to survive. In agreement with this idea, there appears to be a competition between error-free recombinational repair and error-prone translesion synthesis, with UmuD’aC interacting with the growing end of RecA filament and blocking homologous pairing during RecA- mediated recombination, thereby switching repair from homologous recombination to SOS mutagenesis (223, 250, 251). The observation that intact UmuDC inhibits resumption of DNA replication after cells experience DNA damage, and hence provides more time for error-free excision repair (186, 202), further strengthens the idea that the umuDC gene products provide cells with 115 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. an additional means for survival in adverse environments, and as a consequence, confer an evolutionary advantage (85). On the other hand, evolutionary advantages conferred by the action of error-prone DNA polymerases become more plausible in higher organisms. One particular case is mammalian somatic hypermutation where single base changes are introduced, with frequencies as high as 10'3 per base pair, into the rearranged variable (V) regions of antigen activated B cells, most probably as a result of DNA replication errors [for recent review see ref. (90)]. One candidate responsible for somatic hypermutability is human hRad30B (87, 270), an extremely error-prone polymerase (pol i) with base substitution error rate of ~10‘2 . More strikingly, pol i tends to make errors at template T sites, where misincorporation of G is favored 4:1 over incorporation of the correct nucleotide A (270)! This mutase activity (215) would explain the observed strand-bias that mutations from T occur with a much lower frequency than from the complementary A. In vivo studies are in progress to verify the involvement of pol t in the process. With the recent discovery of an error-prone DNA polymerases superfamily, more questions arise: why do cells possess so many errant polymerases? Is there any functional relevance of these enzymes? In S. cerevisiae, pol r\ is required for error-free bypass of cis-syn cyclobutane dimers, whereas pol £, consisting of Rev3 and Rev7, possibly working in concert with Rev1, is involved in error-prone translesion synthesis (191). In humans, the number of these errant polymerases doubles, with Pols r\ (161), i (270), C , (80), 116 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. and 6 [hOinB, as named in (104)] identified, with possibly more to be discovered [for review see (96)]. It’ll be interesting to determine whether different polymerases are selected for specific types of template lesions (or aberrant structures) and how this selectivity is achieved between error-free and error- prone synthesis. Additionally, it will be important to address how these polymerases are regulated so that normal replication is not interfered with and, once these polymerases come into play, how they operate in conjunction with the replicative machinery to allow continuation of normal DNA replication. With the rapid advances in the study of translesion synthesis, definitive answers to these questions will undoubtedly emerge in the near future. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Appendix 1 RecA activation in the in vitro lesion bypass assay Binding of RecA to single-stranded DNA reversibly converts it to the activated form, RecA*, that is important for both SOS induction and SOS mutagenesis (76). To verify that RecA is activated in our in vitro translesion synthesis system, we monitored the ability of RecA* to facilitate cleavage of LexA at the beginning and end of the replicative bypass assay (Fig. A-1). In each case, LexA was cleaved, indicating that RecA is activated throughout the in vitro lesion bypass system. LexA cleavage experiment- 10 pi reactions were carried out as described in Chapter 2, section 2.3, except that the time point at which SSB was added differed, either at time 0 followed by 4 min preincubation with RecA and ( 3 , y complex, or at 4 min when replicative bypass reaction was initiated by pol III and dNTP. Final concentrations were 2 nM P/T, 1 pM RecA, 40 nM p sliding clamp, 10 nM y clamp loading complex, 10 nM pol III core, 20 pM LexA, 100 pM each dNTP, and 1 mM ATP. Following addition of LexA, reaction mixtures were incubated for another 30 min, and terminated with 20 pi of 20 mM EDTA in 95% formamide. The reaction mixtures were separated on a 15% SDS-polyacrylamide gel, and visualized by staining with Coomassie Blue R- 250. 118 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Set 1 Set 2 Set 3 Set 4 SSBconc. (nM) ° g g ° g g ° g g ° g g C o o o o M o o o 5 pol III u RecA LexA SSB Cleaved LexA — ► ^ (Carboxyl fragment) ' IP «*. m m Figure A-1. Activation of RecA in the in vitro lesion bypass assay. Reactions were carried out as described in Chapter 2, section 2.3 except the time point at which SSB was added differed(described as follows). Final concentrations were 2 nM P/T, 1 pM RecA, 40 nM |3 sliding clamp, 10 nM y clamp loading complex, 10 nM pol III core, 20 pM LexA, 100 pM each dNTP, and 1 mM ATP. SSB concentrations were varying as indicated on top of the gel. Lane M contains molecular marker. Lane C is the control reaction where RecA protein was omitted. The four sets of reactions differ in the time points when SSB and LexA were added. Set 1 and 3, SSB was preincubated with RecA, ( 3 and y complex for 4 min before initiation of reaction with pol III and dNTP; Set 2 and 4, SSB was added at the time of reaction initiation, together with pol III and dNTP. Set 1 and 2, LexA was added at the time of reaction initiation; set 3 and 4, LexA was added 10 min after initiation of reaction. 1 1 9 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Appendix 2 ATPase assay for the in vitro TLS system ATP is required in the lesion bypass assay for loading p onto DNA by y complex (201) and also for binding RecA to DNA, converting it to RecA* (227). To verify that the initial input concentration of ATP is sufficient for the reaction, the level of ATP is examined during the course of replicative bypass assay. ATPase assay- Reactions were carried out as described in Chapter 2, section 2.3, except that [a-3 2 P]ATP was used in place of ATP. Final concentrations were 2 nM p/t DNA, 1 mM RecA, 40 nM p sliding clamp, 10 nM y clamp loading complex, 300 nM SSB, and 1 mM [a-^PJATP in 40 pi mixture. A 5-pl aliquot was removed at 6, 9,12,16, 26, and 36 min and quenched with 5-pl of 10% SDS, 40 mM EDTA mixture. Labeled ATP was separated from labeled ADP by spotting 2-pl aliquots on silica gel sheets (Kodak chromagram) and developing with an 11:7:2 NH4 OH:isopropanol:water solution. The radiolabeled reactants and products were detected and quantitated on a Phosphorlmager (Molecular Dynamics). Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. ATP hydrolysis in the in vitro lesion bypass assay 40 35 30 25 20 15 10 5 0 0 5 10 15 20 25 30 35 40 Time (min) Figure A-2. ATPase assay for the in vitro TLS system. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Appendix 3 N-ethylmaleimide inhibits pol V activity N-ethylmaleimide (NEM) is known to inhibit pol III but not pol I or pol II (123). We therefore investigated effect of NEM on pol V activity. Reactions were carried out using running-start replicative bypass assay as described in Chapter 2, section 2.3. NEM, at varying concentrations, was added at time 0 with RecA, ( 3 , and y complex followed by 3 min preincubation. Polymerases and dNTPs were then added to initiate the reaction. As can be seen in Fig. A-3, pol V is inhibited by even 2 mM NEM, as does pol III. Concentration of each polymerase in the reaction: pol II and pol III werelO nM, and pol V was 200 nM. P + y, SSB P - + - + - + - + + - + + - + +:- + + - + + - + + RecA - - - + - + - +; - + - + - + NEM (mM) 0 2 5 0 2 5 0 2 5 Polymerase Pol II Pol III core Pol V Figure A-3. NEM inhibits pol V activity. 122 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Appendix 4 Pol V does not incorporate ddNTP We investigated the ability of pol V to incorporate dideoxynucleoside triphosphate (ddNTP), and compared it with those of pol II and pol III. All reactions were carried out with |3 , y complex, SSB and RecA, in the presence of a ddNTP and the 3 remaining dNTPs. As demonstrated in figure A-4, pol III is able to incorporate ddNTP resulting in strong termination of chain elongation at the position of base-pairing nucleotides on the template strand (“target” sites), similar to those observed in sequencing reactions using T7 polymerase. Pol II and pol V, on the other hand, fail to exhibit a “sequencing” pattern, with products extended beyond the first “target” site. A closer examination of the synthesis pattern revealed an interesting difference between pol II and pol V. Although both polymerases are able to extend beyond “target” sites, major stop bands are observed at the “target” sites for pol II, whereas for pol V, major stop bands are observed one nucleotide before “target” sites. Since primer extension products beyond “target” sites are generated by extension from misincorporated primer terminus at target sites, the result implies that pol V is unable to utilize ddNTP but, in accordance with pol V's error-prone property, it is capable of inserting a wrong dNMP opposite and extending from the "target” sites. As for pol II, it appears unable to utilize ddNTP substrates. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. ddNTP: P G A T C Pol III a HE G A T C G A T C Pol II Pol V Mut Figure A-4. Pol V is unable to utilize ddNTPs as substrates. All reactions were carried out at 37°C for 10 min with |3 , y complex, SSB and RecA, in the presence of a ddNTP and the 3 remaining dNTPs. The inclusion of a ddNTP is denoted by one of the following: G, A, T, and C under each lane. Lane P contains the 3 2 P-labeled primer in the absence of proteins. Part of the template sequence is shown at right of the gel. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Appendix 5 A poly(A) polymerase activity in the UmuD’2 C preparation? Poly(A) polymerase (PAP) catalyzes template-independent sequential addition of AMP to the 3'-terminal hydroxl groups of RNAs. Two poly(A) polymerases have been identified in E. cloi (38, 39). By using an OligoT2 3 - 2 8 (Pharmacia) as primer, we observed a terminal transferase activity in our UmuD’2 C preparation that resembles PAP activity. It catalyzes template- independent addition of AMP or GMP, but fail to use dNTPs as substrates (data not shown). Interestingly, unlike the PAPs in E. coli whose activities are dependent on the presence of an RNA primer, such activity was observed with DNA primers. More experiments are needed to address (i) if the activity is truly a poly(A) polymerase; and (ii) whether the activity is due to an contaminant or intrinsic to UmuD’aC. Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. Set 1 Set 2 OligoT2 3 .2 8 primers ATP: Yes Yes Template: Yes No Figure A-5. Primer extension in the absence of template and dNTPs. All reactions were carried out in the presence of 200 nM UmuD^C, 1 pM RecA, 300 nM SSB, 40 nM p, 10 nM y complex and 1 mM ATP. The OligoT2 3 - 2 8 primer, either annealed to a PolyA template (Pharmacia, average length 400 bases) (set 1), or present alone without any template (set 2), was at 2 nM. The lanes labeled as G, A, T, and C denote reactions carried out with a single dNTP substrate, dGTP, dATP, dTTP, and dCTP, respectively. The lanes labeled as 4 and 0 denote reactions earned out in the presence and absence of four dNTPs, respectively. Lane P contains the 3 2 P-labeled primer in the absence of proteins. 1 2 6 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. OligoT2 3 .2 8 primers SfffSTs 'T 'W W NTP: 4 G A U C 0 4 G A U C 0 4 G A UCO 4 G A U C O RecA, SSB, p/y ATP Yes Yes Yes No No Yes No No Figure A-6. A terminal transferase activity in the UmuD’2 C preparation. All reactions were carried out with 200 nM UmuD’2 C and 2 nM OligoT2 3 .2 8 primer in the absence of any templates. RecA, SSB, p and y complex, if present, were at 1 pM, 300 nM, 40 nM and 10 nM, respectively. ATP, when present, was at 1 mM. The lanes labeled as G, A, U, and C denote reactions carried out with a single NTP substrate, GTP, ATP, UTP, and CTP, respectively. The lanes labeled as 4 and 0 denote reactions carried out in the presence and absence of four NTPs, respectively. Lane P contains the “ P-labeled primers in the absence of proteins. 1 2 7 Reproduced with permission of the copyright owner. Further reproduction prohibited without permission. References 1. Adams, M. D., S. E. Celniker, R. A. Holt, C. A. Evans, J. D. Gocayne, P. G. Amanatides, S. E. Scherer, P. W. Li, et al. (2000). The genome sequence of Drosophila melanogaster. Science. 287:2185-2195. 2. Ason, B., Bertram, JG., Hingorani, MM., Beechem JM, O’Donnell, M., Goodman, MF., Bloom, LB. (2000). A Model for Escherichia coli DNA Polymerase III Holoenzyme Assembly at Primer/Template Ends: DNA triggers a change in binding specificity of the y complex clamp loader. J Biol Chem. 275(4):3006-3015. 3. Bailone, A., S. Sommer, J. Knezevic, M. Dutreix, and R. Devoret. (1991). A RecA protein mutant deficient in its interaction with the UmuDC complex. Biochemie. 73:479-484. 4. Banerjee, S. K., R. B. Christensen, C. W. Lawrence, and J. E. LeClerc. (1988) . Frequency and spectrum of mutations produced by a single cis- syn thymine-thymine cyclobutane dimer in a single-stranded vector. Proc. Natl. Acad. Sci. USA. 85:8141-8145. 5. Bates, H., S. K. Randall, C. Rayssiguier, B. A. Bridges, M. F. Goodman, and M. Radman. (1989). 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Creator Tang, Mengjia (author)

Core Title Biochemical basis of SOS-induced mutagenesis: UmuD'(2)C is an error-prone DNA polymerase, Escherichia coli Pol V

School Graduate School

Degree Doctor of Philosophy

Degree Program Molecular Biology

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag biology, molecular,chemistry, biochemistry,OAI-PMH Harvest

Language English

Contributor Digitized by ProQuest (provenance)

Advisor Goodman, Myron (committee chair), Landolph, Joseph (committee member), Warrior, Rahul (committee member)

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c16-87892

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Legacy Identifier 3018134.pdf

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