Genetic Stability in the Hyperthermophilic Archaeon Sulfolobus Acidocaldarius

Home , Sulfolobus, Sulfolobus solfataricus

Genetic stability in the hyperthermophilic archaeon

Sulfolobus acidocaldarius

A thesis submitted to the Graduate School

of the University of Cincinnati

in partial fulfillment of the

requirements for the degree of

Master of Science

Department of Biological Sciences

McMicken College of Arts and Sciences

by Xinyu Cong

2015

B.A. Huazhong Agricultural University

June 2012

Dennis Grogan, Ph.D., Committee chair

Brian Kinkle, Ph.D.

Stephanie Rollmann, Ph.D

Abstract

Hyperthermophilic archaea (HA) grow optimally at high temperatures that can easily destroy

DNA structure Not much is known about how these organisms maintain genetic fidelity and genome stability. The aim of this thesis research was to investigate the strategies adopted by

Hyperthermophilic archaea Sulfolobus acidocaldarius to maintain genetic fidelity and genome stability. Three studies were conducted: (i) genetic effects of disrupting Sulfolobus acidocaldarius B-family polymerase pol 2 or 3 gene, (ii) the specificity and consequence of DNA lesion bypass in vivo among wild type and DNA polymerases mutant strains, (iii) the molecular requirements for endogenous mutations of highly expressed “lacS” reporter gene in a shuttle plasmid pJlacS. An important component of maintaining genetic fidelity, DNA polymerase, was investigated in Sulfolobus acidocaldarius. Two B-family DNA polymerases mutants were used individually to test the functions of these two polymerases in sensitivity to DNA damaging factors and in effects on spontaneous mutation in a selectable gene pyrE. Disruption of either

B-family DNA polymerase did not lead to increased sensitivity to chemicals or UV light; however, compared to wild type strains, both B-family DNA polymerases mutants showed altered spontaneous mutation spectra; for example, they both had increased level of transition mutations. The DNA lesion bypass events were determined by transforming a short gapped plasmid pDM8, which has single chemically defined DNA lesion located opposite the gap. This gapped plasmid requires replication filling the gap and bypassing the lesion in order to replicate inside cells and clones that have bypassed the DNA lesion were recovered. The gap region was sequenced to identify consequence of DNA lesion bypass. The results showed that DNA lesion bypass introduced different events such as base pair changes and short deletions. Finally, to

i investigate the mechanism of endogenous mutation of a highly expressed gene, one hypothesis was tested, that the formation of R-loop during transcription leaving non-transcribed DNA single-stranded. To examine this hypothesis, a lacS gene which is normally expressed on a plasmid pClacS was made into large gapped. This gap covers the lacS gene to simulate the single-stranded structure during transcription. The lacS mutants were calculated and analyzed for the mutation types. Results showed that lacS mutation rate of gapped pClacS were significantly higher than that of intact pClacS but the mutation spectrum was different from that of pJlacS.

iii

Acknowledgements

I thank Dr. Dennis Grogan (chair) for being my mentor. He is so generous and always ready for helping me with any troubles. I also would like to thank my other committee members Dr. Brian Kinkle, and Dr. Stephanie Rollmann. Each member has shown great support and kindness towards my career pursuit.

I also would like to thank current and former members of the Grogan lab, Joseph Fackler, Dominic Mao, and Cynthia Sakofsky. They have provided great help in my experiments.

I would like thank Wieman Wendel Benedict funding for supporting many laboratory related purchases.

Finally, I would like thank my family and friends. They have provided great encouragement and support for my pursuit of master degree.

Chapter 1

Introduction to the study of hyperthermophilic archaea (HA) 1

1.1 The unique molecular properties of archaea 1 1.2 Sulfolobus acidocaldarius as a model organism 5 1.3 Overview of DNA polymerases in HA and other organisms 5 1.4 Overview of Transcription associated mutagenesis(TAM) 7

Chapter 2

Genetic effects of disrupting Sulfolobus acidocaldarius B-family polymerase Pol 2 or 3 11

2.1 Introduction 11

2.2 Method and Materials 13

2.2.1 Construction of the polymerases mutants 13

2.2.2 Assay of spontaneous mutation, sensitivity to DNA damaging chemicals and UV light. 14

2.3 Results 16

2.3.1 Construction of the polymerases mutants 16

2.3.2 Analysis of sensitivity to DNA damaging chemicals and UV light 17

2.3.3 Assay of spontaneous mutation 19

2.4 Discussion 23

Chapter 3

The specificity and consequence of DNA lesion bypass in vivo among wild type and DNA polymerases mutant strains 24

3.1 Introduction 24

3.2 Method and Materials 26

3.2.1 Construction of small gapped pDM8 with single chemically defined DNA lesions 26

3.2.2 Electroporation and genetic analysis 27

3.3 Results 27

3.4 Discussion 30

Chapter 4

The molecular requirements for endogenous mutations of “lacS” reporter gene in a shuttle plasmid pJlacS 34

4.1 Introduction 34

4.2 Method and Materials 36

4.2.1 Construction of large gapped pClacS 36

4.2.2 Electroporation and screening lacS mutants with X-gal 36

4.3 Results 38

4.4 Discussion 40

Chapter 5

Conclusions 43

Reference 46

Chapter 1 Introduction to the study of hyperthermophilic archaea (HA)

1.1 The unique molecular properties of archaea

Based on the sequence of 16S rRNA, archaea were first identified as the third domain of life by Carl Woese and colleagues in the 1970’s (Woese & Fox, 1977). According to Woese, evolution of cells is the central problem of evolutionary biology and evolution of microbes will be the core of the new evolutionary biology (Koonin, 2014). Along with bacteria and eukaryotes, they make up the three domains of the phylogenic tree which is also called ribosomal tree of life (Woese et al, 1990). To compare the genomes of bacteria and archaea, followed up by molecular biological and evolutionary experiments can reveal novel evolutionary phenomena; for example, pervasive horizontal gene transfer (HGT), to a large extent mediated by viruses and plasmids, that shapes the genomes of archaea and bacteria, requires a radical revision of the tree of life concept (Koonin & Wolf, 2012).

Archaea share many similarities with bacteria in cellular structure. For instance, both are unicellular, have no membrane-bound nucleus and contain circular chromosomes. When it comes to information-processing systems, such as transcription and translation, however, archaea are more like eukaryotes. The RNA polymerase and ribosomes in archaea are more closely related to those in eukaryotes, like the eukaryote RNA polymerase II. Archaea RNA polymerase also requires TATA-box binding protein (TBP) and transcription factor B (TFB) for transcription recognition (Allers & Mevarech, 2005).

Archaea, especially the hyperthermophilic archaea (HA), thrive in extreme environments such as geothermal regions (80 ℃ or higher). Sulfolobus, a genus of HA, grow optimally at high temperature and acidic conditions (pH 2.5-3.5) where they have to cope with the threats from

1 hyperthermal environments, and the temperature that can destabilize DNA basic chemical structure (Lindahl.

1993). Despite the pressure from environment, Sulfolobus acidocaldarius has been observed to have similar spontaneous mutation rate to Escherichia coli (Grogan, 2001).

Therefore, S. acidocaldarius must have mechanisms to maintain the integrity and stability of their genome DNA under such harsh environments.

It is widely accepted that cellular organisms require high fidelity of genome replication in order to achieve biological success (Fijalkowska et al, 2012; Shin et al, 2014). This needs high fidelity DNA polymerases to replicate and DNA repair pathways to remove replication errors, base damage or deal with unrepaired lesions. These DNA repair pathways are broadly conserved from bacteria to humans (Table 1). However, under conditions where organism fitness is not yet maximized for a particular environment, competitive adaptation may be facilitated by enhanced mutagenesis. (Loh et al, 2010), indicating that microorganisms may benefit from low genetic fidelity.

HA have to cope with the threats from hyperthermal environments. However, the lack of certain significant repair proteins makes HA quite different from the other two domains organisms (DiRuggiero et al, 1999). Mismatch repair (MMR) is a highly conserved pathway in both eukaryotes and prokaryotes, which repairs mismatch generated during DNA replication.

Usually the mismatch is detected and a part of the newly synthesized DNA strand is removed,

2 leaving a gapped DNA. DNA polymerases fill the gap and therefore repair the mismatch (Iyer et al, 2006).The genome of HA does not encode homologues of MutSL proteins of E. coli, which are used for post-replication MMR in all other systems, from bacteria to eukaryotes. However, it is complicated in mesophilic archaea, some have MutSL homologues, such as methanomicrobia and halobacteria, but some do not, like methanobacteriales and thermococcales (Banasik & Sachadyn, 2014)

Nucleotide excision repair (NER) is an important repair pathway that removes various

DNA damage, especially those causing helix-distortion and it can, for example, repair bulky lesions caused by UV light. The damage is recognized first and a short single strand DNA that contains the lesion is removed, and DNA polymerase uses the undamaged strand as template to fill the gap (Melis et al, 2013). In contrast, another repair system, base excision repair (BER) can only repair non-helix-distorting base lesions. DNA glycosylases find the lesion and remove the damaged base, leaving an abasic site (AP). The abasic site is cleaved by AP endonuclease and DNA polymerase fills the gap (Wallace, 2014). The homologues of bacterial UvrAB or eukaryotic XPA, XPC proteins, which are DNA damage-recognition proteins required for initiation of nucleotide excision repair (NER), are also absent in HA genomes (Sachadyn 2010;

White & Grogan 2008). However, the absence of a homologue does not necessarily mean the absence of the repair activity; in spite of the lack of damage-recognition homologues in NER, which is a major repair pathway for UV-induced lesions, some HA has shown the ability to repair UV lesions in their genome efficiently (Dorazi et al, 2007; Romano et al, 2007). The fact that HA can maintain their genome stability under stressful environments but lack

3 corresponding proteins that are essential in DNA repair pathways in other organisms makes HA unique.

In contrast, divergence of HA from other model organisms is also reflected in proteins they have. HA genome encodes a DNA reverse gyrase that can positively supercoil DNA , which is not seen in mesophilic bacteria or eukaryotes (Forterre, 2002) and HA as prokaryotic organisms, have the homologs of eukaryotic CMG complex, which function as a key component of DNA replication machinery in HA (Makarova et al, 2012). HA also seems to need active recombination functions to live. It was documented that HA cells cannot grow when they are deficient of recombinase RadA, Rad50, Mre1 and a helicase HerA which is only found in archaea

(Fujikane et al, 2010; Zheng et al, 2012); however, the homologous proteins in bacteria, yeast and even mesophilic archaea can be deleted (Miller & Kokjohn, 1990; Heyer, 1994; Woods &

Dyall-Smith, 1997;).

Even though certain repair pathways may be missing or highly diverged among HA, it seems like that HA have strongly effective systems to maintain genomic DNA integrity. So far, based on the available evidence, thermophiles depend on various strategies to maintain their genome integrity. Some factors that determine the melting temperature of DNA in vitro may be indications of the strategies adopted by thermophiles to prevent strand separation in vivo. First of all, there are some extrinsic factors responsible for this, such as ions and polyamines. It was found various novel polyamines in thermophilic archaea, which are more effective at increasing

Tm of DNA than inorganic cations (Terui et al, 2005). It was also shown that thermophilic archaea have high levels of small, basic proteins that have high affinity for double-strand DNA

(Robb et al, 2008). However, it should be noted that the biological roles of these proteins are not necessarily confined to stabilization of double-strand DNA for example; polyamines were proved not only to stabilize DNA against thermal pressure, but also confer additional compaction to DNA with histones (Higashibata et al, 2000). Second, as to intrinsic factors, it was previously assumed that thermophiles have a high level of G+C content. Some thermophilic bacteria do have high G+C contents, but most do not and there is no proved correlation between optimal growth temperature and G+C contents among bacteria and archaea (Galtier et al, 1997; Hurst and Merchant, 2001). Therefore, the G+C content is not likely to be the determinant of the thermophiles’ optimal growth temperature. Last but not the least, positive supercoiling is believed to be able to stabilize double strand DNA. It was found that, as to all bacteria and archaea which grow optimally above 75 ℃, their genome encodes a reverse DNA gyrase (Rgy) and this gyrase is absent from all other prokaryotic genomes (Forterre, 2002).

However, the deletion of reverse DNA gyrase was found to be non-lethal in one thermophile, but the deletion decreased its range of growth temperature (Atomi et al, 2004), indicating that the reverse DNA gyrase does have a role in maintaining proper cellular function.

1.2 Sulfolobus acidocaldarius as a model organism

There are some advantages to choose Sulfolobus acidocaldarius as a study species. First of all, in laboratory conditions, it can be easily cultured with doubling time of 4 hours at about

80 ℃, and grows aerobically (Grogan, 1989). The genomic sequencing of many Sulfolobus spp., including Sulfolobus acidocaldarius, has been accomplished recently, providing great help to molecular genetic experiments (Chen et al, 2005). Moreover, Sulfolobus acidocaldarius as a model organism can be transformed with foreign DNA by electroporation (Kurosawa and

Grogan, 2005). More and more widely used genetic tools in bacteria are now available in

Sulfolobus, for example, some E.coli and Sulfolobus shuttle vectors have been constructed and played important roles in experiments (Berkner et al, 2010; Liang et al 2013) and these vectors can be equipped with useful genes, such as β-D-glycosidase reporter gene lacS and orotate phosphoribosyltransferase gene pyrE, which will support auxotroph transformation recipient cells to grow on selective plates.

1.3 Overview of DNA polymerases in HA and other organisms

It has been documented that organisms from all three domains have more than one

DNA polymerase, and it is a significant issue to understand why organisms have so many different kinds of DNA polymerases and what their functional differences are. Based on the primary sequence homologies and crystal structure analyses, DNA polymerases are divided into six families (A, B, C, D, X, and Y) (Rothwell & Waksman, 2005). Family A polymerases are replicative and repair enzymes found in eukaryotic mitochondrion and bacteria. Family B polymerases are the major replicative polymerases of eukaryotes but not replicative polymerases in prokaryotes. Family C polymerases are the major bacteria chromosomal replicative enzymes. The majority of X family are nucleotidyl transferase proteins. So far, only six X-family DNA polymerases have been identified in eukaryotes and they function as replicative and repair polymerases. DNA polymerases from Y family are identified to be able to recognize and bypass different classes of lesions on DNA (Rothwell & Waksman, 2005). The D- family polymerases are only found in Euryarchaeota and in roles of replicating DNA (Cann et al,

1998). E. coli, for example, has five DNA polymerases, Pol I (Family A), functions in processing of

Okazaki fragments and excision repair with 3'-5' and 5'-3' exonuclease activity; Pol II (Family B)

6 are responsible for DNA replication, translesion-DNA synthesis (TLS) and excision repair with 3'-

5' exonuclease activity; Pol III (Family C) has roles in DNA replication and excision repair with 3'-

5' exonuclease activity ; Pol IV and Pol V are Family Y polymerases and carry out TLS in vivo

(Iwona et al, 2012). Yeast has more polymerases than E. coli does, for instance, Family-A Pol γ is mitochondrial DNA replication polymerase; Pol α, Pol ε, Pol δ and Pol ζ are Family-B polymerases and they have functions in chromosomal DNA replication and repair; while, Pol ζ is also involved in Error-prone TLS; Pol β and Pol σ are Family-X polymerases. Pol β is required for double-strand break repair and base excision repair; however, Pol σ is needed for sister chromatid cohesion; Pol η and Rev1 are Family-Y polymerases. Pol η is responsible for error- free translesion DNA synthesis and Rev1 is required for synthesis opposite an abasic site

(Hubscher et al, 2002; Kawasaki & Sugino, 2001).

In Sulfolobus, there are only polymerases from B (more than one) and Y families. Some

B-family polymerases have been suggested to replicate genome DNA and some are suggested to be involved in non-replicative roles, for instance, translesion DNA synthesis (TLS). (Hubscher et al, 2002).It is widely accepted that Polymerase B1, which has high fidelity and strong affinity to DNA (Zhang et al, 2009), replicates the Sulfolobus genome, The Y-family polymerase is reported to be error-prone polymerase that bypasses a variety of DNA lesions (Boudsocq et al,

2001). But it is unclear about the other two B family DNA polymerases’ natural roles. Since the organizations of prokaryotes are highly streamlined, they generally do not have unnecessary structures or enzymes. It is therefore reasonable to think that each DNA polymerase in HA plays significant roles.

1.4 Overview of Transcription associated mutagenesis (TAM)

Higher fidelity of genome replication does not necessarily mean stronger fitness. High level of fidelity in replication or transcription may negatively affect genomes’ adaption potential, while decreasing replication or transcription’s velocity and increasing its costs, which will have an impact on fitness; for example RNA viruses fast replication sacrifices replication fidelity but is critical for survival (Furió et al, 2007; de Visser & Rozen, 2005). In addition, under certain circumstances, moderately enhanced mutagenesis can facilitate adaptation (Loh et al, 2010).

Stress-induced adaptive mutagenesis is proposed as a new evolutionary viewpoint and gene transcription under stress condition increases the instability of genome in response to DNA damage, resulting in transcription-associated DNA mutagenesis (TAM) (Zhu & Li, 2014).

Transcription was first indicated to be involved in mutagenesis in the 1970s. It was documented that in E. coli, an exogenous mutagen was more effective at inducing mutations under highly transcribed condition (Herman & Dworking, 1971). In the 1990s, however, there was a debate over Cairnsian mutation or adaptive mutation that mutations are beneficial and specific to stress, instead of being random (Foster, 1993). Transcription is considered to be one of the stresses that cause adaptive mutation (Galhardo et al, 2007). Also in eukaryotes, specifically in budding yeast, transcription was connected to mutagenesis. It was demonstrated that the rate of spontaneous reversion increased when the mutant gene was highly transcribed

(Datta & Jinks-Robertson, 1995). This phenomenon is defined as transcription associated mutagenesis (TAM), which locally and permanently alters the DNA template sequence (Kim &

Jinks-Robertson, 2012). Now, TAM has been observed in most of the model organisms, such as

8 in bacteria E. coli, archaea Sulfolobus and in eukaryotes, yeast (Wright et al, 1999; Sakofsky &

Grogan, 2013).

Recent efforts have been focused on explaining the mechanisms behind TAM and transcription level was thought to be a key factor in TAM. It was demonstrated that in yeast the mutation rate is directly proportional to transcription level (Kim et al, 2007). Recently, studies showed that there are multiple mechanisms that contribute to TAM, such as collisions with the replication machinery, formation of co-transcriptional R-loops, engagement of Top1 activity, alterations in DNA base (Kim & Jinks-Robertson, 2012; Figure 1).

A :

B :

Chemical modification Strand breaks

RNA polymerase

Figure 1 A: Replisome and RNA polymerase are in head-on orientation. B: R-loop formation, Factors can affect the non-transcribed strand.

But it is still not known how specifically a given mechanism makes contribution to a particular TAM event. One possible mechanism is the structure of R-loop, which is formed during transcription that the nascent RNA is annealed back to the transcribed strand, creating a stable RNA-DNA hybrid and leaving the non-transcribed strand (NTS) exposed as a single- stranded DNA (Reaban et al., 1994) and this single-stranded DNA can be affected by mutagenic factors such as chemical modification, strand breaks, etc. This was observed in E. coli that absence of TOPO 1 facilitated the formation of R-loop, which caused serious problems to cells growth (Drolet et al, 1995). Moreover, it was also documented in yeast that the formation of

DNA: RNA hybrids, namely R-loop, lead to impaired transcription elongation (Huertas &

Aguilera, 2003). In addition, with respect to higher eukaryotes, metazoan cells, the depletion of

ASF/SF2 protein promoted R-loop formation, which resulted in a hyper-mutation phenotype (Li

& Manley, 2005).

It sounds logical that cells which grow optimally at high temperature require more effective DNA repair strategies than those at low temperature (Robb et al, 2008). Thus, compared with other model organisms, it seems like HA either have functional alternatives that can replace conventional DNA repair pathways, or maintain ancestral DNA repair systems that were abandoned during evolution by other model organisms. In that case, analysis of HA may uncover novel but effective strategies that maintain DNA integrity. More importantly, study of

HA may provide significant inspirations to understanding molecular diversity of DNA repair systems across biology. Even though, until now people have known a lot about HA, such as the whole genome sequence, the biochemical features and crystal structures of various proteins, it still remains unclear how the DNA replication and repair function in vivo, how the DNA damage

10 tolerance system, which have been studied well in other model organisms, functions in HA, and how to explain the DNA repair systems fail to function in HA with respect to highly expressed genes.

To gain a better understanding of how HA maintain the integrity of their genomes, this thesis investigated the biological roles of two non-essential B-family DNA polymerases in spontaneous mutagenesis and responses to DNA damage (Chapters 2 and 3). To investigate the mechanism behind TAM in Sulfolobus, the mutagenic effect of single-stranded DNA was measured (Chapter 4).

Chapter 2 Genetic effects of disrupting Sulfolobus acidocaldarius B-family polymerase pol 2 or 3

2.1 Introduction

It has been documented that all cellular organisms have more than one DNA polymerases, and it is a significant issue to understand why organisms have so many different kinds of DNA polymerases and what their functional differences are. To answer this question, people have to investigate not only these polymerases catalytic and biochemical properties, but also their natural functions in vivo. For instance, in archaea Sulfolobus acidocaldarius there are three B-family DNA polymerases and one Y-family polymerase; however, two non-essential B- family DNA polymerases natural roles remain mysterious in spite that people have obtained a lot their catalytic and biochemical properties as discussed below.

There are studies about the corresponding polymerases from a related species

Sulfolobus solfataricus (three B-family polymerases: Dpo1, Dpo 2 and 3, one Y-family polymerase: Dpo4) in vitro. Catalytic and biochemical tests shows that, compared to Dpo 1 and

Dpo4, Dpo2 and Dpo3 have low polymerase and 3’ to 5’ exonuclease activities (only Dpo4 has no exonuclease activity) and weak DNA binding. Although Dpo1 cannot bypass DNA lesions,

Dpo2 and Dpo3 can bypass hypoxanthine, 8-oxoguanine, and either uracil or cis-syn cyclobutane thymine (Choi et al, 2011). In addition, during nucleotide incorporation opposite undamaged DNA templates, Dpo3 had about 60-fold higher misinsertion frequencies than Dpo1

(Dpo2 was not able to be determined accurately) (Choi et al, 2011; Bauer et al, 2012). In that case, based on the biochemical test and enzyme assay in vitro, it is reasonable to think that Pol

2 and Pol 3 are additional TLS DNA polymerases of S.acidocaldarius, with properties of

12 bypassing DNA lesions related to those of Y-family polymerase, and disruption of either of this enzyme will hinder the cell’s bypass of certain DNA damage, which will increase its sensitivity to those DNA damage. This prediction also indicates that Dpo1 or Pol 1 is the only replicative DNA polymerase. Having only one replicative DNA polymerase is typical in bacteria; for example, in

E.coli Pol III is the only replicative enzyme. As to other DNA polymerases’ functions in E.coli, Pol

I is used to remove RNA primers from Okazaki fragments during lagging strand synthesis, Pol II can restart stalled replication forks when there are DNA damages on the genome, and Pol IV, V are TLS polymerases (Rangarajan et al, 1999; Rattray & Strathern, 2003). Therefore, the molecular functions of Pol 2 and Pol 3 in S.acidocaldarius may be similar to those non- replicative enzymes in E.coli.

With respect to the functions of B-family DNA polymerases, the only B-family DNA polymerase in E.coli, Pol II, is believed to have high fidelity, 3’ to 5’ exonuclease activity and a function of replication restart (When the replication is stalled at DNA damages like UV-induced damage, it needs to be restarted; Rangarajan et al, 1999; Iwona et al, 2012). Nevertheless, it was also reported that Pol II has error-prone TLS ability at certain DNA lesions in vitro, like bulky adducts (Wang & Yang, 2009), indicating that Pol II in E.coli may be involved in both chromosomal replication and TLS. In addition, eukaryotic B-family polymerases Pol α, Pol δ and

Pol ε are documented to be high-fidelity enzymes; Pol δ and Pol ε have 3’ to 5’ exonuclease activity and a function of chromosomal replication. However, Pol α has a primase instead of exonuclease activity and synthesize primer (Hubscher, 2002). However, Pol ζ in eukaryotes is reported to be error-prone TLS polymerase, which has low fidelity (Nelson et al, 1996). It was documented that in yeast, B-family DNA polymerase Pol ζ defective cells have reduction in

13 spontaneous mutation (Huang et al, 2002). In addition, as to human cells, disruption of Pol ζ does not affect cell cycle or viability, but decreases UV-induced mutagenesis (Zan, 2001) and overexpression of Pol ζ in yeast leads to an increase in UV-induced mutagenesis (Rajpal, 2000).

It suggests that some spontaneous mutations may actually require the presence of Pol ζ or other DNA polymerases.

Nevertheless, there is no published study probing the properties of Sulfolobus accessory

Pol 2 and Pol 3 (i.e. Dpo2 and Dpo3) in vivo, and researchers have emphasized the need for knockouts of these genes and study of the mutants (Bauer et al, 2012). Study 1 using species

Sulfolobus acidocaldarius may be the first such analysis of Pol 2 and Pol 3 functional mechanisms in vivo.

2.2 Method and Materials

2.2.1 Construction of the polymerases mutants

The host cell used is S. acidocaldarius auxotroph MR 31 with inactivated Sac pyrE (the pyrE gene of S. acidocaldarius) gene. Two pairs of oligos (pol 2: GTTTTGTAGAACTTCTTTAGCGA

GATCACACCAATTATTCCGATATGAGAGAGGTTTATC & TGTCTTAATCTCACAAAGAGATAAAGAAAC

TAAGTAAGCCCTCCTCTCCTATAACCAATA; pol 3: GTGTTTCAGAGAGGTGATAGTAGAGGCTACA

AAGAGGGTCAGTGTCTTAATCTCACAAAG & ATTGGATATGGGGTTCCGTCTTTAGCGTTAA

ATAGCATTACGATATGAGAGAGGTTTATC) with flanks of targeted polymerases DNA sequence were used to amplify Sso pyrE (the pyrE gene of S. solfataricus) followed by the PCR program

DG 48 (initial denaturation at 95 ℃ for 2 minutes, followed by 30 cycles of 95 ℃ for 22 seconds,

48 ℃ for 22 seconds, 72 ℃ for 1 minutes 33 seconds and a final extension of 72 ℃ for 3

14 minutes). PCR products were purified on MILLIPORE Centrifugal Filter Units (Billerica, MA) to eliminate ions. 5 microliters of the PCR products were used for electroporation and plates were incubated at 70 ℃ for 6 days.

2.2.2 Assay of spontaneous mutation, sensitivity to DNA damaging chemicals and UV light.

Minimum inhibitory concentration (MIC) test.

The chemicals chosen are MNNG (N-methyl N’-nitro N-nitrosoguanidine) , Butadiene diepoxide, Cisplatin (cisplatinum(II) diammine dichloride), Mechlorethamine(2-chloro-N-(2- chloroethyl)-N-methyl-ethanamine), Metronidazole (2-(2-methyl-5-nitro-1H-imidazol-1-yl) ethanol), Hydrogen peroxide . Each chemical was dissolved in water or ethanol and made into serial 1:2 dilutions in liquid growth medium and dispensed on a 96-well plate first and equal inoculum of polymerase mutants and WT cells with same cells density were added to the same plate. The 96-well plate was incubated at 70 ℃ for 2-3 days. The MIC of each strain to a certain chemical is the lowest concentration where cells cannot grow.

UV radiation assay.

Both mutant and WT cultures were grown in liquid medium to exponential phase, harvested by centrifuge, and resuspended in UV-transparent dilution buffer to about 1×108 cells/ml. Suspensions were spread on a glass plate lid as a thin (∼2 mm) layer and irradiated under a germicidal UV lamp and aliquots were withdrawn at regular time intervals, diluted and plated in a dark room before incubation at 70℃ (in the dark) to determine viable counts.

Spontaneous mutation test on S. solfataricus pyrE gene (Sso pyrE).

I tested the spontaneous mutation rate for Sso pyrE of wild type, pol 2 mutant and pol 3 mutant strains by fluctuation assays. About 25 independent cultures (0.2 ml) for each strain were grown at 70℃ for 3-4 days to densities of about 108 cells/ ml. The entire culture was then applied to selective plate containing both 5-Fluoroorotic acid (5-FOA) and uracil. The plates were incubated at 70 ℃ for 6 days. All FOA-resistant colonies were counted and one of them was picked randomly for each culture. Picked colonies were restreaked on plates containing only uracil for later analysis. The identity of the spontaneous pyrE mutations was determined by

PCR amplification with program DG 48 and sequencing from each isolated colonies on restreaked plates. The primers used were JRset1.fwd (ACGCCCTTAAATAAGGTTAG) and JRset1. rev (GGGACATTGAAAGAACTAGA).

Spontaneous mutation test on S. acidocaldarius pyrE gene (Sac pyrE).

The Sac pyrE gene was restored in Sso pyrE & pol 2 (or pol 3) double mutant cells by transforming them with the intact Sac pyrE gene cloned in plasmid pLk3a (Sakofsky et al, 2011).

The pyrE gene on the pLK3a replaced the old one and restored Sac pyrE function.

Transformants were plated on plates without uracil and cells with successfully restored Sac pyrE can form colonies on plates. Sac pyrE + cells were used for spontaneous mutation test and the test follows the same procedures described above. The primers used for amplifying Sac pyrE are pol 2 : saci0074regionF (AAG AGA GGA AGT GGT ATT GGC), saci0074regionR (AAC AAG AGG CTC

AAC AGG C); pol 3: saci1256regionF (GAA CCT TTC TCA GCC CTG T), saci1256regionR (CGT CTC

CCA TCT CCT CAA T).

2.3 Results

2.3.1 Construction of the polymerases mutants

The host cell used is S. acidocaldarius auxotroph with inactivated Sac pyrE (the pyrE gene of S. acidocaldarius) gene. The pyrE gene, encoding orotate phosphoribosyltransferase, is indispensable in the synthetic process of UMP and cells with a disrupted pyrE gene can only grow in the media containing uracil. Two pairs of oligos with flanks of targeted polymerases

DNA sequence were used to amplify Sso pyrE (the pyrE gene of S. solfataricus), and these amplified products were electroporated into host cells to accomplish homologous recombination with polymerase genes respectively, which switched the middle parts of polymerase genes with the smaller Sso pyrE gene. Successful disruption resulted in decreased size of polymerase genes as shown in Figure 1.

pol 2 or 3 gene Disrupted pol 2 or 3 with pyrE inserted pyrE cassette

Figure 1 Independent disruption of two polymerase genes. Gray line represents the polymerase genes and black line represents the pyrE gene and the pyrE cassette combined with polymerase genes sequence at the flanks. After homologous recombination, the pyrE cassette replaced a larger part of the polymerase genes, which decreases the distance between the primers.

So far the two polymerase mutated strains have been successfully constructed and confirmed by PCR as shown in Figure 2. Recovery of these mutants demonstrates that both Pol2 and Pol3 are not essential to the survival of Sulfolobus cells.

14.1 14.1 4.8 14.1 2.3

1.4 14. 10.7 A B C 14. 1

14. Figure 2: 1 Confirm ation of polymerase disruption. A: PCR amplification of the polymerase gene region. Lane 1, 2 show amplification14. of pol 2 in disruptant and WT cells, respectively. Lane 3, 4 show amplification of pol 3 in disruptant and WT cells, respectively. Successful disruption decreased the size of polymerases genes, which is shown in A. B: Amplification of pyrE gene1 from pol 2, 3 disruptants as shown in lane 1, 2. C: PCR amplification of polymerase disruptant but with combined primers from A, B. Lane 1, 2 show the combination of pol 2 forward primer with pyrE reverse primer and pol 2 reverse with pyrE forward. The same goes with lane 3, 4 of pol 3 disruptant. MW: molecular weight marker (bacteriophage λ DNA digested with BstEII).

2.3.2 Analysis of sensitivity to DNA damaging chemicals and UV light

MIC assays were performed as a preliminary assay of DNA lesions survival capacity. If inactivation of pol 2 or pol 3 gene causes deficiencies of DNA damage repair or tolerance, this may increase the sensitivity to various DNA damaging chemicals which can be revealed by comparing polymerase mutants and WT cells. The chosen chemicals produce diverse DNA lesions such as, cross-linked, covalently modified, oxidized bases, and each chemical represents one particular type of those DNA lesions. These compounds have been used to distinguish DNA polymerases mutants and WT strains in E.coli and Sulfolobus (Kim et al, 2001; Wong et al, 2010).

Polymerases disruptants only showed sensitivity to cisplatin which causes intra- and inter-strand crosslinks (Table 2). These results suggested that Pol2 and Pol3 may have some

contribution to tolerance of strand crosslinks damage; however, it is not necessary to say Pol2

and Pol3 are not involved in tolerance of other DNA lesions. Because it is possible that TLS is

carried out by multiple polymerases in vivo and the inactivation of one DNA polymerase can be

compensated by others; also it is possible that the tests were not extensive enough to find the

DNA damages that Pol 2 and Pol 3 bypass such as deamination.

Table 2: Sensitivity to DNA-damaging chemicals. DG185 is the wild type strain and Δpol 2 and Δpol 3 are the polymerases disruptants. MIC values were determined from 3 replicates with each chemical. They only showed different sensitivity to cisplatin which causes intra- and inter-strand crosslinks. Values are minimum inhibitory concentrations (ug/ml).

Some translesion synthesis (TLS) polymerases can bypass DNA damage brought by UV

light. In order to test whether Pol 2 and Pol 3 have functions in this way, mutants were

evaluated for UV survival under different exposure time: 0 second, 6 seconds, 10 seconds and

20 seconds as shown in Figure 3. Results showed no significant difference of sensitivity to UV

light between WT cells and polymerase mutants despite UV doses that kill over 99.99% of the

cells.

UV exposure time (second)

0 5 10 15 20 0 -1 -2 DG185 -3 Δpol 2 -4 Δpol 3 -5

Log (Survival fraction) (Survival Log -6

Figure 3: Sensitivity to UV light. There was no significant difference in sensitivity to UV between the DNA polymerase mutants and wild-type cells.

2.3.3 Assay of spontaneous mutation

Spontaneous SSo pyrE mutations

The pyrE gene is indispensable in the synthetic process of UMP, and the auxotroph cannot grow without uracil. However, when 5-FOA is added to the medium, the PyrE processes

5-FOA into 5-fluorouridine-5’-phosphate, a lethal toxin (Yamagishi et al, 1996).Spontaneous mutation rate of Sso pyrE gene was tested on both mutant and WT cells. In order to do this, mutants and WT cells were spread on plates that contain both uracil and 5-FOA. In that case, only those mutants that lose pyrE function can grow on the plate. Mutants on the plate were counted and revived in liquid media. The mutated pyrE gene was amplified and sent for sequencing to determine what specific mutations occur at the pyrE loci.

The spontaneous mutation test of Sso pyrE gene was compared with that of Dbh disruptants and wild type strains (Table 3) (Sakofsky et al, 2012). All the mutations observed from Δpol2 are base pair substitutions (BPS); however, the mutations of Δpol3 can be divided into 2 classes, BPS (62%) and frameshift (38%). What is interesting is, for the mutations found in

Δdbh cells, they were all single mutation events; however, there were multiple mutations in

Δpol2 and Δpol3 single mutant strains. Even though it was not confirmed by statistical tests because we don't have enough mutants to construct full mutations spectra, the preliminary results suggested that inactivation of Pol 2 or Pol 3 had different effects on spontaneous mutagenesis, indicating their distinct roles in vivo. Compared to dbh+ (namely, WT cells), Pol2 seems to avoid BPS and generate frameshift as well as indels mutations; while, Pol3 may have roles in avoiding BPS and generating indels.

Table 3: Spontaneous mutation rate test on SSo pyrE. dbh+ is the wild type strain, Δdbh, Δpol2 and Δpol3 are the polymerase disruptants. PyrE spontaneous mutation rate was measured and mutations are categorized into different classes. Total mutations means the mutation events numbers. BPS: base pair substitution; Indels: insertions and deletions

Spontaneous Sac pyrE mutations

Although mutations of Sso pyrE gene could be analyzed without any additional genetic manipulation, this gene is located differently in the polymerases defective cells and it is possible that the position inconsistency could have an effect on the spontaneous mutation of

Sso pyrE gene. Therefore a more detailed analysis should be performed with the Sac pyrE gene.

After statistics test, the spontaneous mutation rate of Sac pyrE was not significantly different among DG185 and two DNA polymerase mutants (Table 4). Also there was no significant difference in the three categories of mutations among wild type and DNA polymerase mutants. However, when considering only BPS or not BPS category, there was difference between WT vs pol 2 mutants (p value: 0.16), and WT vs pol 3 mutants (p value:

0.08).The same goes with frameshift or not frameshift category (p value: 0.23, 0.09). Even though the values did not yield significant difference, they were close to 0.05. Therefore, results still indicated that disruption of pol 2 or pol 3 changed both BPS and frameshift mutations spectra compared to WT. This was also indicated in the preliminary date obtained from analysis of Sso pyrE mutations (Talbe 3). Both polymerase mutants had increased transition mutations; however, in pol 2 mutants, there were 8 G: CA: T and 7 A: TG: C transition events. On the other hand, there were 13 G: CA: T and 4 A: TG: C transition events in pol 3 mutants and χ2 test showed that they were different (Table 5). In addition, comparison of transitions between

DG 185 and pol3 mutant also showed difference, which indicated that Pol 3 may emphasize on inhibiting G: CA: T transition; while Pol 2 may inhibit G: CA: T and A: TG: C transitions equally. Moreover, comparisons of transversion mutations between DG185 and pol 2-, or DG

185 and pol 3-, or pol 2- and pol 3- showed no difference at all.

CS2 Δdbh (on Sac PyrE) DG 185 (on Sac PyrE) Δpol 2(on Sac PyrE) Δpol 3(on Sac PyrE) Total mutations 111 100% 102 100.0% 109 100.0% 112 100.0% BPS 48 43% 18 17.6% 28 25.7% 31 27.7% Transitions 17 15% 5 4.9% 15 13.8% 17 15.2% Transversions 31 28% 13 12.7% 13 11.9% 14 12.5% Frameshift 53 48% 67 65.7% 63 57.8% 61 54.5% +1 bp in runs 25 23% 27 26.5% 27 24.8% 30 26.8% - 1 bp in runs 18 16% 32 31.4% 33 30.3% 21 18.8% +1 bp not in runs 8 7% 5 4.9% 3 2.8% 7 6.3% -1 bp not in runs 1 1% 2 2.0% 0 0.0% 2 1.8% +2 bp 0 0 1 1.0% 0 0.0% 1 0.9% Indels 10 9% 17 16.7% 18 16.5% 20 17.9% -3 bp 0 0 0 0.0% 0 0.0% 0 0.0% Duplications >6 bp 9 8% 5 4.9% 7 6.4% 6 5.4% Deletions >3 bp 1 1% 9 8.8% 11 10.1% 14 12.5% "+3 or +6" 0 0 0 0.0% 0 0.0% 0 0.0% Duplications <6 bp 3 2.9% 0 0.0% 0 0.0%

Table 4: Spontaneous Sac pyrE mutations. DG 185 is the wild-type strain; Δpol2 and Δpol3 are the polymerase disruptants; CS2 Δdbh is the Dbh- strain and pyrE spontaneous mutation of this strain was done by Cynthia Sakofsky (Sakofsky et al, 2012). The rate of spontaneous mutation of pyrE was measured, and mutations are categorized into different classes. The “total mutations” means the number of mutation events and corresponding percentage. BPS: base pair substitution; Indels: insertions and deletions

DG 185 vs Δpol 2 DG 185 vs Δpol 3 Δpol 2 vs Δpol 3 Three types mutations: BPS, Frameshift, Indels 0.35 0.17 0.88 BPS, not BPS 0.16 0.08 0.7 Frameshift, not frameshift 0.23 0.09 0.61 Transitions, transversions (BPS) 0.08 0.07 0.92 A:T to G:C, G:C to A:T (Transitions) 0.61 0.12 0.17 A<->G, C<-> T (Transitions) 0.80 0.03* <0.0001* A-T, A-C, G-T, G-C (Transversions) 0.48 0.53 0.65

Table 5: Statistical analyses of mutation spectra. Estimated p-values from χ2 test. Three types mutation categories included BPS, frameshift and indels mutations. A<->G, C<-> T (Transitions) includes transition between A,G and transition between C,T. Transversions categories include A-T, A-C, G-T and G-C mutations. * indicates significant.

2.4

Discussion

The properties of B-family DNA polymerases from a closely related species Sulfolobus solfataricus have been well documented in vitro. These biochemical tests included polymerase activity, exonuclease activity, DNA binding power, and lesions bypass (Choi et al, 2011; Bauer et al, 2012). However, properties of these polymerases in their natural context have never been documented. This study, therefore, provided the first phenotypic analysis of pol 2 and pol 3 mutants of S. acidocaldarius.

It is possible to disrupt two B-family DNA polymerases in S. acidocaldarius, which means that they are not essential for Sulfolobus cell. In terms of sensitivity to DNA damaging chemicals, however, DNA polymerase disruptants only showed possible sensitivity to cisplatin. Disruption of pol 2 or pol 3 has no significant effect on UV light survival or spontaneous mutation rate of pyrE gene. This does not mean that Pol 2 and Pol 3 are not involved in tolerance of other DNA lesions, because it is possible that TLS is carried out by multiple polymerases in vivo and the inactivation of one DNA polymerase can be compensated by other DNA polymerases or DNA repair pathways. In E. coli, DNA lesions bypass is carried out by multiple DNA polymerases; Pol I,

Pol II, Pol IV, and Pol V have been proved to have roles in TLS (Fuchs & Fujii, 2013).

However, deleting pol 2 or pol 3 changed the spontaneous mutation spectrum, indicating their natural roles in replication accuracy. Both Δpol 2 and Δpol 3 exhibited lower frequencies of frameshift mutations, but these lower frequencies were offset by higher frequencies of BPS. Furthermore, both Δpol 2 and Δpol 3 mutants had increased transition mutations. These results indicate that S. acidocaldarius B-family DNA polymerases Pol 2 and Pol

3 may have at least two roles in vivo: one that generates frameshift mutations, and one that inhibits BPSs, more specifically the transition mutations.

The observation of Pol 2 (or Pol 3)-suppressed mutations suggests that some DNA lesions in S. acidocaldarius genomes are bypassed accurately by Pol 2 or Pol 3 but erroneously by other DNA polymerases. The transition mutations observed in the pyrE gene accounted for nearly all of the mutations inhibited by Pol 2 or Pol 3. Specifically, Pol 3 seems to suppress G:C to A:T transition; while Pol 2 seems to have less roles in suppressing both G:C to A:T and A:T to

G:C transitions. Mechanisms that generate these mutations have significant implications for the biological roles of the Pol 2 and Pol 3 in the natural context.

Both two types of transition mutations can arise from deamination. Deamination of cytosine to uracil and of adenine to hypoxanthine can result in transition of G:C to A:T and A:T to G:C, respectively. The deamination of cytosine in nucleotides can be measured at elevated temperatures (Lindahl & Nyberg, 1974) and considering the living condition of S. acidocaldarius, it is possible that deamination happens in vivo. Since the deamination of cytosine is the major source of G:C to A:T transition and It was documented that eukaryotic Pol ι can bypass uracil and its derivatives accurately in vitro (Friedberg, 2006; Vaisman & Woodgate, 2001). Therefore, it is possible that Pol 3 have functions in bypassing uracil (not the hypoxanthine) accurately in vivo. On the other hand, Pol 2 can bypass both uracil and hypoxanthine accurately in vivo.

However, these results are partially contrary to a recent study about Sulfolobus B-family

DNA polymerase function in vitro. According to a recent study, both Dpo2 and Dpo3 were able to bypass hypoxanthine; while only Dpo2 (and not Dpo3) was able to bypass uracil (Choi et al,

2011). These results combined with the studies I did indicate that Pol 2 can bypass hypoxanthine and uracil accurately; however Pol 3 can bypass uracil instead of hypoxanthine accurately in vivo.

In summary, the analysis of B-family polymerase mutants’ phenotype indicates that both Pol 2 and Pol 3 have roles in avoiding transition mutations (specifically Pol 2 avoids G:C to

A:T as well as A:T to G:C and Pol 3 avoids G:C to A:T) that otherwise accumulate in S. acidocaldarius genomes. This remains partially consistent with biochemical studies about

Sulfolobus B-family polymerases’ functions in vitro (Choi et al, 2011). However, data also shows that both Pol 2 and Pol 3 generate frameshift mutations, which was not seen in vitro.

Chapter 3 The specificity and consequence of DNA lesion bypass in vivo among wild type and DNA polymerases mutant strains

3.1 Introduction

Translesion DNA synthesis is regarded as a DNA damage tolerance system, which is a strategy of cells coping with damaged DNA. The Sulfolobus Y-family DNA polymerases crystallize well, and offer many advantages for structural and biochemical analysis. There are over 100 published studies about crystal structure and catalytic properties of Sulfolobus Y-family polymerase Dpo4 and Dbh in vitro. However, the mechanism of TLS, especially with respect to the natural roles of Y-family polymerases, is not well understood. There are only two studies investigating them in vivo (Wong et al, 2010; Sakofsky et al, 2012) and both studies showed that there were relatively subtle defects in Dpo4/Dbh null mutants, indicating that multiple TLS polymerases may work together or there are other polymerases that are able to bypass diverse

DNA lesions. This situation is consistent with study of TLS in other organisms (Wagner et al.

2002; Godoy et al, 2007; Livneh et al, 2010).

There are some DNA lesions, if not repaired, can cause mutations in vivo; for example, deamination of cytosine and adenine can result in transition mutations (Friedberg et al, 2006); oxidation of guanine to 7,8-dihydro-8-oxoguanine or 8-oxoguanine (8-oxoG) results in transversion mutations following replication (Cheng et al, 1992); misalignment of the primer and the template can cause frameshift mutation, either 1 bp addition or deletion (Drake, 1991).

TLS system helps organisms survive unrepaired or unrepairable damage to DNA (Rattray &

Strathern, 2003). TLS as a DNA damage tolerance mechanism is a way that a DNA polymerase inserts nucleotides opposite DNA lesions that are either noninstructional or misinstructional.

Sometimes TLS bypass a particular DNA lesion can be accurate however, in other cases it can be highly mutagenic (Friedberg et al, 2005). It has been recognized that, all the Y-family polymerases that have been found so far are specialized for TLS. These are found both in bacteria such as in E. coli DNA Pol IV & V and in eukaryotes Saccharomyces cerevisiae DNA Pol η, and human DNA Pol η (Rattray & Strathern, 2003). As to archaea, it was shown that, based on the crystallized and biochemical analysis, S. solfataricus Y-family polymerase Dpo4 as well as two B-family polymerases Dpo2 and Dpo3 have DNA lesions bypass functions in vitro (Wong et al, 2010; Choi et al, 2011). Abasic (AP) site is a location in DNA that has neither a purine nor a pyrimidine base. Another common DNA lesion is 8-oxoG, which is guanine with saturated imidazole ring caused by oxygen free radicals. AP sites, if not repaired, can cause replication fork stalling and translesion DNA synthesis; however, 8-oxoG can cause mispairing with adenine in DNA. In E.coli, for example, Pol II as well as Pol IV and Pol V can bypass AP sites and 8-oxoG

(Fuchs & Fujii, 2015). The bypass of DNA lesions, however, is more complicated in eukaryotes.

For example, in yeast, the AP site can be tolerated by error free post-replication repair (PRR) and recombination processes, or translesion DNA synthesis carried out by Rev1, Pol δ and Pol η

(Boiteux & Guillet, 2004). 8-oxoG can be bypassed by yeast Pol δ and Pol η; however, Pol η is more accurate than δ (McCulloch et al, 2009). According to an in vitro enzyme assay, all four

Sulfolobus DNA polymerases fail to bypass AP site; however, two B-family DNA polymerases

Dpo 2, Dpo 3 and Y-family DNA polymerase Dpo 4 can effectively bypass 8-oxoG (Choi et al,

2011).

Nevertheless, as to in vivo test, knockout of Y-family polymerase Dbh in S. acidocaldarius indicates that there was no significant difference in sensitivity toward a diverse

28 array of DNA-damaging chemicals (Sakofsky et al, 2012), which indicates that these lethal DNA lesions induced by various chemicals are not actually bypassed by Dbh in vivo, or other DNA polymerases or DNA repair pathways compensate significantly effectively for the absence of

Dbh in vivo.

Thus, studying the properties of TLS in wild-type and DNA polymerases mutant

Sulfolobus cells is necessary to uncover what Y-family or B-family DNA polymerases contribute to TLS in their native contexts and whether certain DNA lesions lead to consistent mutations in

WT and TLS-deficient cells.

3.2 Method and Materials

3.2.1 Construction of small gapped pDM8 with single chemically defined DNA lesions

The methylated plasmid pDM8 was extracted from E. coli strain ER2566 [p6b, pC ins 4] with large scale plasmid extraction kit. The plasmid was first nicked at the bottom strand at 37℃ overnight (44 microliters of plasmid, 5 microliters reaction buffer, 1 microliter Nb.BbvCI), followed by heating at 75 ℃ for two minutes, releasing the bottom strand. Reaction was purified into 35 microliters with Biolne Sureclean (Tauton, MA). 10 microliters of oligo with a

DNA lesion (abasic site and 8-oxo G) was added together with 4 microliters ligase buffer and 1 microliter of DNA ligase at room temperature overnight for ligation of bottom strand. Then the top strand was nicked with 1 microliter of nicking enzyme Nt.BbvCI at 37℃ for 2 hours and heated at 75℃ for 2 minutes to release the top strand, which results in a short gap opposite a

DNA lesion as shown in Figure 1 B and Figure 2. This reaction was followed by adding 1 microliter of NruI at 37℃ for 2 hours for eliminating background plasmid which is not

29 gapped. The final reaction was purified with Sureclean into 30 microliters in ddH2O and kept frozen at -20℃.

3.2.2 Electroporation and genetic analysis

This short-gap plasmid was electroporated (3 microliters /electroporation) into

Sulfolobus wild-type cells MR 31 and the transformants were plated and incubated at 70 ℃ for

6 days. Transformants were picked into 2 ml liquid cultures and incubated at 70 ℃ for 2-4days until the culture was turbid. After incubation, genomic DNA was extracted from each culture and amplified with primers pDMMMRA1f (CAG GAA ATG GCG TAG GTT), pDMMMRA1r (TCG

CTT GAC TTG ACC AGA) following PCR program DG 48. PCR amplicons were digested by StuI for screening and the digestion recipe for each reaction is 3 microliter of PCR amplicon, 6 microliters of ddH2O, 1 microliter of NEB Cutsmart (Ipswich, MA), 0.5 microliter of StuI. StuI resistant amplicons were sent for sequencing with primer pDMMMRA1r. The results were used to construct spectra of mutations from the specific DNA lesion.

3.3 Results

Abasic site

The in vivo assay of translesion DNA synthesis is based on electroporation of gapped plasmid. One single DNA lesion was placed into a specially designed E.coli –Sulfolobus shuttle plasmid pDM8 by chemical synthesis of an oligonucleotide and a series of nicking, annealing, and ligation steps as in Figure 1. This shuttle plasmid has selective genes pyrE and bla, which support Sulfolobus auxotroph to grow in medium without uracil and E.coli resistant to ampicillin respectively (Berkner et al, 2007). In addition, there are several restriction enzyme sites near where the DNA lesion was introduced, such as StuI, nicking enzyme BbvCI. What should be noticed is that the top and bottom sequences shown in Figure 2 are actually on a plasmid. Thus first replication of this gapped plasmid in vivo should lose the StuI cut site, which can be used as a screen of successful preparations as shown in Figure 2. Also after ligation of the oligo, it was introduced a mismatch which is 2 bases before the lesion, changing a “G” into a “T” and this destroyed StuI enzyme cut site as well (Figure 2) . Only if TLS successfully bypasses DNA lesion can this plasmid replicate and generate a transformant. Thus, each transformant shows an individual by-pass event in vivo.

FIGURE 1: Short-gap plasmid assay. TLS must bypass DNA lesion (x) in single-stranded region in order for the plasmid to replicate and generate a transformant.

A total of 94 transformations were performed, 101 colonies were screened and 9 samples were confirmed to have lesion bypasses and sequenced; however no background reversion was observed. The low transformation efficiency of gapped pDM8 with AP lesion

(average of 1 transformant per electroporation) is consistent with transformation results using abasic-site oligonucleotide (Sakofsky and Grogan, unpublished). DNA lesion by-passed transformants showed several TLS induced mutations opposite the abasic site as shown in

Figure 2, including 5 short deletions and 4 out of them have the same size which are also at the same location of the gap previously made on the plasmid before transformation and one has a

shorter deletion and different location; an insertion of G, which is an transversion compared to

original base; and an insertion of T opposite the abasic site, which is the same as original base.

FIGURE 2: Results of short-gap plasmid assay. The top gray sequence is the region where gap is. The bottom strand is where the abasic site is introduced, indicated as only an underline. The oligo used to introduce AP site has a mismatch two bases before lesion, which changes original “G” into “T” at the bottom strand and this is also a mark of true replications bypass AP site. Sequencing results showed different bypass events.

8-oxo G

Preliminary data of transformants (wild type strain) with 8-oxoG gapped pDM8 was also

obtained. A total of 93 colonies were yielded from 12 individual transformations with an

average of 8 colonies per electroporation, which is much higher than that of AP pDM8

transformation. Confirmed 48 bypass events were categorized into 4 different groups: 1) an

insertion of C opposite the 8-oxoG followed by a deletion, or the 8-oxoG was skipped 2) an

insertion of C, 3) a precise short deletion same in AP bypass, 4) an insertion of T opposite the 8-

oxoG, followed by an extra insertion (Figure 3).

FIGURE 3: Preliminary results of 8-oxoG short-gap plasmid assay. The confirmed mutations are into 4 different categories with numbers of occurrence shown at the right side.

3.4 Discussion

The extremely low transformation efficiency of gapped plasmid with AP site lesion, which is obvious when compared with 8-oxoG transformation, could be due to the fact that enzymes recognized the AP and destroyed it. Both eukaryotic AP endonucleases and archaeal, yeast glycosylases are shown to be responsible for removing AP in vivo, leaving 3' hydroxyl and

5' deoxyribosephosphate termini, and the cleavage forms a single strand break on DNA

(Berquist et al, 2008; Hardeland et al, 2003; Sartori et al, 2001). In the experiment here, the break on AP site makes gapped plasmid linear, and this linear plasmid loses functions. This explanation can be tested by incubating the gapped pDM8 with Sulfolobus cell’s lysis extraction.

Gapped pDM8 is expected to be cut into linear form. On the other hand, another possibility is that if AP is not removed, it will prevent replication of the plasmid, which causes low

34 transformation efficiency. An in vitro biochemical test on four Sulfolobus DNA polymerases showed that none of them can bypass AP lesion efficiently (Choi et al, 2011).

With respect to the precise deletions on the oligo inserting region, it could be due to the fact that after the first nicking and releasing step, the top stand can form a hairpin structure in three possible ways as shown in Figure 3 in vitro.

FIGURE 3: Three hairpin structures. Three possible hairpin structures are formed by the top single-stranded DNA.

This hairpin structure can bring the two bottom ends close. After the ligation step, the

bottom strand is ligated instead of ligating AP site oligo; while the top strand is looped out as

shown in Figure 4.

Hairpin structure

Nicking sites Nicking sites

Top strand

Bottom strand

Ligated place

FIGURE 4: Loop structure on oligo insert region. The hairpin structure on the top stand loops out and bottom strand is ligated back together.

After the second nicking on the top strand and followed up with heating, the hairpin

structure would be released, leaving an almost intact plasmid with only a cut on the top as

shown in Figure 5.

Cut Top strand

Bottom strand

FIGURE 5: Structure of the plasmid after second nicking. Nicking and heating release the top hairpin structure and result in an almost intact plasmid.

Transformation of this special structured plasmid created in vitro could lead to the precise deletion results seen in Figure 2. It would be possible to test this explanation by skipping the addition of the synthesized oligo and following the rest steps of preparing the gapped plasmid. If this hypothesis is true, short deletions at the nicking region should be the dominant results. Another possibility that the damaged oligo on the plasmid causes the precise deletions can be tested by switching the AP site oligo with another oligo without any damage and following the rest steps of preparing the gapped plasmid.

The analysis of bypassing 8-oxoG in vivo shows that the insertion of C opposite 8-oxoG was the dominant results. This is different from the observation that many DNA polymerases misincorporate A opposite 8-oxoG (Shibutani et al, 1991; Pavlov et al, 1994). It is possible that one or more DNA polymerases in S. acidocaldarius can bypass 8-oxoG accurately. A recent study about Saccharomyces cerevisiae 8-oxoG bypass in vivo showed similar results. Translesion synthesis by Pol η replicates 8-oxoG with an accuracy (insertion of a C opposite the 8-oxoG) of approximately 94%, and Pol η mutants had dramatic lower accuracy of 8-oxoG bypass

(Rodriguez et al, 2013). Based on an unpublished research study (Sakofsky and Grogan, unpublished), S. acidocaldarius Y-family DNA polymerase dbh mutants had lower accuracy and efficiency of bypassing 8-oxoG in vivo when compared to wild type cells. This result indicates that Dbh may have functions in incorporating C opposite 8-oxoG. Also the experiment was conducted via transforming an oligo that has 8-oxoG at a synonymous base next to a selective marker; however compared to gapped plasmid assay, this experiment was not able detect frameshift and insertion or deletion (indel) mutations. On the other hand, in vivo test on 8- oxoG bypass in E. coli yielded different results that most bypass events over 8-oxoG were G-to-

T base substitution (Shikazono et al, 2013). This inconsistence may indicate that archaea are more similar to eukaryotes in translesion DNA synthesis.

In summary, the gapped plasmid assay of TLS provides a useful analysis of TLS in

Sulfolobus, especially compared to oligo transformation assay which can only measure the incorporation opposite 8-oxoG but not frameshift or indel mutations that arise from bypassing

8-oxoG. Also whether Pol 2 and Pol 3 have functions in bypassing 8-oxoG accurately is not known, even though gapped plasmid assay and in vitro experiments indicate that there may be more than one DNA polymerase that bypass 8-oxoG accurately and both Pol 2 and Pol 3 replicate through 8-oxoG efficiently in vitro as undamaged G (Choi et al, 2011). Therefore, to conduct 8-oxoG gapped plasmid and oligo assays in pol 2 and pol 3 mutants appears to be a necessary complement to the understanding of Pol 2 and Pol 3 natural roles as well as DNA lesion bypass in Sulfolobus.

Chapter 4 The molecular requirements for endogenous mutations of “lacS” reporter gene in a shuttle plasmid pJlacS

4.1 Introduction

Transcription-associated mutagenesis (TAM) is related to a high level of expression in vivo and is believed to be a major force for adaptation (Zhu & Li, 2014). Unresolved questions include how the DNA repair systems fail to function in response to highly expressed genes and how the mutations form. TAM has been studied in E.coli and yeast for decades (Herman &

Dworking, 1971; Datta & Jinks-Robertson, 1995), but was only observed in archaea recently, by analysis of the reporter gene lacS on a plasmid pJlacS in S. acidocaldarius (Sakofsky & Grogan,

2013).

The research was based on a Sulfolobus β-D-glycosidase gene (lacS), placed in pRN1 that originates in S. islandicus strain REN1H1, which is a shuttle vector for Sulfolobus species (Zillig et al, 1994) and has a strong promoter for lacS gene. Sulfolobus lacS gene can be used as a reporter for its β-D-glycosidase function and mutation of it can be scored with chromogenic indicators 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside, namely “X-gal”. High rate mutation of this gene was observed in the plasmid pJlacS, which has a strong promoter for lacS gene (Berkner et al, 2007). After correction of plasmid copy number and the growth advantage of LacS- cells, it was still observed that the lacS mutation rate is about 800 times higher than in the S. acidocaldarius chromosomal pyrE and pyrF genes (Sakofsky & Grogan, 2013). Seventy two pJlacS mutants were sequenced, and the results indicated 3 major categories of mutations, base-pair substitutions (BPS), large deletions, and slipped-strand events (Sakofsky & Grogan,

2013). It is believed that this high rate of mutation of lacS in pJlacS is relevant to the

39 documented ‘transcription-associated mutagenesis’ (TAM) in eukaryotic cells and bacteria

(Wright et al, 1999; Datta & Jinks-Robertson, 1995; Pybus et al, 2010).

The unusually frequent spontaneous mutation observed in pJlacS was the first recorded mutagenesis in HA which are caused by endogenous or native factors (Sakofsky & Grogan,

2013). This endogenous process has its unusual properties that the magnitude is extraordinarily higher than that of genomic DNA, indicating that mutation-avoidance systems have been impaired or overwhelmed in replicating this plasmid. The endogenous mutations observed in pJlacS have the features of TAM (transcription-associated mutagenesis) in that it increases with the transcription level. It is logical to think that most TAM observed in bacteria and eukaryotes involves the formation of single-stranded DNA, which can form an unusual structure called R- loop that have potentials to be mutagenic (Beletski & Bhagwat, 1996; Kim & Jinks-Robertson,

2012). It is hypothesized that the structure of single-stranded DNA leads to TAM observed in pJlacS.

Understanding the molecular basis of this endogenous mutagenesis will provide new insight into mutagenic processes that can impair HA genome stability and the corresponding mechanisms that can be used to cope with these threats. This chapter presents the first examination of the role of single-stranded DNA in the endogenous mutations in HA.

Understanding the molecular mechanisms of this mutagenesis will be a hint of solving the puzzle that why the HA can maintain genome integrity under stressful environments but at the same time, they fail to avoid spontaneous mutation of a plasmid-borne gene. Also, since mutations are strongly related to evolution in that under certain circumstance, selective growth

40 advantages can be conferred by enhanced mutagenesis (Loh et al, 2010) , uncovering the molecular requirements for this endogenous mutagenesis is contributive to answering evolutionary questions of HA.

4.2 Method and Materials

4.2.1 Construction of large gapped pClacS

The nicking reaction included 90 microliters of pClacS, 10 microliters of NEB Cutsmart buffers, 2 microliters Nt. BvCI and stayed at 37℃ overnight. Reaction was then diluted by 100 microliters of ddH2O as nicked pClacS working stock. ExoIII working stock was made by 0.5 microliter of ExoIII, 45 microliters of ddH2O, and 5 microliters of NEB Cutsmart buffer. The digesting reaction consisted of 10 microliters of nicked pClacS, various amounts of ExoIII (4 to 8 microliters) and stayed at 37 ℃ for 20 minutes, followed by heat inactivation at 80 ℃ for 30 minutes. A three microliter aliquot was picked from each ExoIII reaction and mixed with 6 microliters of ddH2O, 1 microliter of NEB Cutsmart buffer and 0.5 microliter of StyI & 0.5 XhoI for monitoring. This reaction was at 37 ℃ for 1 hour and the entire reaction was loaded on an agarose gel as shown in Figure 2. The preparations used for electroporation were 10 microliters of nicked pClacS, 6 microliters of ExoIII and stayed at 37 ℃ for 20 minutes. Half a microliter of

StyI was added to the preparation and stayed at 37℃ for 1 hour, followed by purification with

BIOLINE Sureclean into 10 microliters of ddH2O.

4.2.2 Electroporation and screening lacS mutants with X-gal and enzyme assay

Each electroporation consumed 5 microliters of these gapped pClacS preparations, plated and incubated at 70 ℃ for 6 days. Colonies were screened for lacS mutants by blue/white screening using 5-bromo-4-chloro-3-indolyl-β-D-galactoside (X-gal) solution (200

μg/ml X-gal, 0.1 M KCL, 0.1 M β-alanine, 10% ethanol v/v, 0.2% GELZAN) sprayed on plates and incubated at 55 ℃ for 6-10 hours. White and lighter in blue colonies were considered potential

Lac- and they were restreaked on plates and incubated at 70 ℃ for 6 days.

The potential Lac- mutants were confirmed by quantitative measurements of β-D glycosidase (lacS) activity (Berkner et al, 2007). Isolated mutant colonies were picked up and grown in 2ml XT culture at 70 ℃ for 2 days. The density of the cells was measured at an absorbance of 600 nm and about 106 cells were added to enzyme assay reagent (1 M KCL, sodium citrate pH5, 10 % N-lauroyl sarcosine, 0.05 M para-nitrophenyl-β-D-galactopyransoside) and incubated at 70 ℃ for 30-60 minutes depending on the density of the cell. The reaction was then added with 0.3 M Na2CO3 to present color. Absorbance readings at 420 nm were recorded and the activity of β-D glycosidase was determined by the following formula: OD420/OD600 (of corresponding cell suspension), where OD is the absorbance readings at a wavelength of 420 and 600 nm.

Strains having an activity less than 30% of that of LacS+ cells were considered LacS- mutants and confirmed by PCR followed by sequencing of the lacS gene. The PCR program used is initial denaturation at 95 ℃ for 2 minutes, followed by 30 cycles of 95 ℃ for 22 seconds, 48 ℃ for 22 seconds, 72 ℃ for 3 minutes and a final extension of 72 ℃ for 7 minutes. The primers used to amplify lacS were PClacS3712f (GATATCTGATAGTTGGAGAAATGC), pClacS3712reverse

(CTGTCTCTTATACACATCTGGATC) and sequenced with an additional middle primer LacSmidf

(GATGTGACAGAGGTTGAGATAAA).

4.3 Results

The experiments tested whether single-stranded regions created in a Sulfolobus plasmid produces mutations like those observed in TAM. To test this, plasmid pClacS was first nicked once on the bottom strand at the downstream, followed by Exo III 3’ to 5’ digestion which digests only one strand. This process was monitored by other restriction enzymes StyI and Xho I as shown in Figure 1. The Sty I cut site is at the beginning of lacS gene and the Xho I cut site is about 2.0 kb upstream from the Sty I cut site. Nicked pClacS can be cut into two fragments by

Sty I and Xho I; nevertheless, if the exonuclease digestion reaches to the beginning of lacS, which destroys the Sty I site, the treatment of both Sty I and Xho I can only make this plasmid into a linear form.

Control experiments showed that StyI treatment could identify the length of time of Exo

III digestion needed to remove one strand from the entire lacS gene. Nicked pClacS was treated by different amounts of Exo III as shown in Figure 2. These results provided guidelines for the treatment conditions for preparing plasmid with single-stranded lacS. However, all the preparations were treated by Sty I before being electroporated as a final purification step.

Unsuccessful or incomplete gap formation leaves the StyI site intact, and this treatment cut those into linear form of the plasmid, eliminating interference of intact lacS gene.

StyI site

FIGURE 1: Preparing large-gap plasmid. The plasmid was nicked at the downstream of lacS gene first, followed by digestion of only one strand and monitored by Sty I and Xho I (as the arrows show), which results in a single-stranded lacS gene.

XhoI site

MW 1 2 3 4 5 6 7 8

FIGURE 2: Monitoring the Exo III digestion on pClacS. Lane 1 is the original pClacS, and lane 2 is pClacS after nicking. Lane 3 shows nicked pClacS cut by Sty I and Xho I, that 2.0 kb fragment cut out from pClacS is marked by an arrow. Lanes 4 to 8 are pClacS cut by Sty I and Xho I after treatment by various amounts of Exo III (4 to 8 microliters). The 2.0 kb fragment is marked by an arrow, but is faint compared with that of lane 3. Based on the amount of DNA shown on the gel, it is measured by ImageJ that there was only about 5% of the pClacS left in lane 6 that can be cut by both Sty I and Xho I, which means that there were only 5% of the preparation in lane 6 that were unsuccessful. MW: molecular weight marker (bacteriophage λ DNA digested with BstEII).

In order to identify the plasmids that contained a mutation in lacS, X-gal was used to

screen the colonies of pClacS transformants on the plate. LacS- colonies were colorless, while

LacS+ colonies can be stained bluish. The non-staining mutant colonies were revived in liquid

44 media and the whole lacS gene was amplified and sent for sequencing. The spontaneous mutation rate and spectra of lacS were compared with that observed in pJlacS.

So far, 1057 colonies that generated from transformation of large gapped pClacS were screened and 22 sequenced. There were only 5 strains that showed mutations (Table 1), and the resulting mutation rate is about 0.47%. This result can be compared to the transformation of intact pClacS, which yielded 2 mutant colonies out of 6104, corresponding to a mutation rate of about 0.032%. Therefore, the increased mutation rate (by a factor of about 15) seems due to the formation of gapped plasmid.

large gapped pClacS intact pClacS induced pClacS Total mutations 5 2 35 Fraction of total mutations BPS 0.40 0.00 0.37 Transitions 0.20 0.00 0.34 Transversions 0.20 0.00 0.03 Frameshift 0.60 0.00 0.28 +1 bp 0.40 0.00 0.17 - 1 bp 0.20 0.00 0.11 +2 bp 0.00 0.00 0.00 -2 bp 0.00 0.00 0.00 Indels 0.00 1.00 0.34 small deletions (<=5 bp ) 0.00 1.00 0.31 large deletions >5 bp 0.00 0.00 0.03

Table 1: Mutation spectrum of lacS on pClacS under different conditions. Transformation of induced pClacS was conducted by Cynthia Sakofsky (Sakofsky & Grogan, 2013).

4.4 Discussion

Due to the low recovery of mutants, there were not enough for a full analysis of mutation spectrum. A χ2 test shows that there is no significant difference of the major mutation categories (BPS, Frameshift, and Indels) between the gapped pClacS and induced pClacS. However, the observed mutations may differ from those induced pClacS, in some respects, based on the following comparison. For example, there is no indel mutation observed in gapped pClacS, but these make up about one-third of the lacS mutations seen in this same plasmid under TAM conditions (Table 1). Moreover, the mutations found in the lacS gene after strand removal have a different spatial pattern than those induced in the same plasmid under

TAM conditions. The former occur at 705, 768, 848, 1294, 1430 bp in the coding sequence, whereas lacS mutations that form under TAM conditions nearly all occur in bp 938-952 and

212-217 of lacS (Sakofsky & Grogan, 2013). Finally, neither of the BPS mutations from single- stranded plasmid generated stop codons, but all of the BPS mutations generated stop codons in pClacS under TAM conditions (Sakofsky & Grogan, 2013). The results therefore show that the mutations observed in highly expressed plasmid-borne lacS gene in Sulfolobus can’t be simply explained as the formation of single-stranded DNA.

It has been documented that replication and transcription conflicts or collisions can cause genome instability (Rudolph et al, 2007; Pomerantz & O’Donnell, 2010). There are two possibilities of the collisions: the transcription and replication are either co-directional or head- on. Evidence showed that transcription of all the rRNA operons in E.coli are in the same direction with DNA replication and this pattern has been observed in over 80 bacterial species

(Ellwood & Nomura, 1982; Guy & Roten, 2004). In addition, reversing the orientation of rRNA

46 genes which results in transcription and replication head-on in Bacillus subtilis causes negative effects on fitness (Srivatsan et al, 2010). These indicate that the head-on collisions are more problematic. Therefore the alternative hypothesis will be the collisions of replication and transcriptions.

There are some other experiments that need to be done in future to clarify these questions in the Sulfolobus system. For example, the lacS gene on pClacS should be reversed and re-tested under TAM conditions. To accomplish this, the lacS gene together with its promoter need to be cut out by HaeII and NaeI first and followed by ligation back to pClacS again. This can yield lacS gene in inversion position on pClacS. To select successful inverted lacS gene on pClacS, people can transform the modified plasmid into E. coli, extract plasmid from transformants and use lacS downstream sequence as forward primer, lacS promoter region as reverse primer to amplify lacS gene. Only inverted lacS gene can be amplified with these primers. Therefore, people can extract pClacS with inverted lacS and induce it in S. acidocaldarius. In spite of the old orientations of replication and transcription of lacS on plasmid, the reverse will make the new orientations either co-directional or head-on and new hypotheses will be tested based on whether there are significant changes of mutation rate and spectra of lacS gene in induced pClacS.

In other organisms there are several enzymes that can prevent the accumulation of R- loops. These include topoisomerase 1, RNase H class, and RecG helicase of E.coli (Tuduri et al,

2009; Cerritelli et al, 2009; Rudolph et al, 2010). The relative genes in Sulfolobus acidocaldarius are found, such as topoisomerase 1 (Saci_1371), ribonuclease HII (Saci_0958 rnhB). The

47 possibility of deletions of these genes’ homologues in Sulfolobus will be tested first. Then people will construct single or multiple genes disruptant cells. The plasmid pClacS will be transformed into these cells. At the background of promoted accumulation of R-loop, normally expressed lacS gene should have more frequent endogenous mutations, and the spectra of mutations will also be compared with pJlacS expressed in WT cells. In addition, pJlacS will be transferred into cells that have overexpression of those enzymes, and this should inhibit the unusually frequent mutations of lacS.

Chapter 5 Conclusions

Archaea, a group of fascinating organisms, are usually found in the extreme environments on earth. They are set apart from bacteria and eukaryotes, encompassing unique molecular characteristics. Sulfolobus spp. thriving in extreme environments (usually 80℃, pH2-

3), have to maintain cellular processes functioning normally and cope with the condition that can increase DNA damages. In order to have a broader understanding of archaea, my thesis mainly focuses on the various aspects of molecular functions involved in maintaining genetic fidelity and genome stability in Sulfolobus acidocaldarius.

As mentioned in the introduction (Chapter 1), some significant questions raised by

Sulfolobus spp include how the DNA replication and repair of Sulfolobus acidocaldarius function in vivo, how the DNA damage tolerance system, which have been studied well in other model organisms, functions in HA, and how certain DNA repair systems may fail to function in HA with respect to highly expressed genes. After conducting the research in this thesis, at least parts of these questions can be answered.

It seems that both Pol 2 and Pol 3 have functions in accurate replication and bypass DNA lesions. An indispensable component of genetic fidelity is the DNA polymerases. Genetic analysis showed that DNA polymerases pol 2, pol 3 and dbh in Sulfolobus acidocaldarius can be deleted individually, and the deletion of each DNA polymerase did not cause obvious growth defect or sensitivity to DNA damaging agents or overall spontaneous mutation rate at a well- characterized marker gene (Sakofsky et al, 2012). Analysis of spontaneous mutations from the marker gene showed that both pol 2 and pol 3 mutants had increased BPS mutations, while decreased frameshift mutations. These results indicated that pol 2 and pol 3 may have

49 functions that affect DNA accuracy in avoiding BPS mutations and generating frameshift mutations meanwhile. Even though the pol 2 and pol 3 mutants have slight difference in mutation spectrum from wild type strains, the data still suggested that the inactivation of one

DNA polymerase could be compensated by others and Sulfolobus may have complicated pathways to maintain their DNA integrity. To have a better understanding of these non- essential DNA polymerases’ (Pol 2, Pol 3 and Dbh) natural functions, the next step in the investigation will be to construct and investigate multiple polymerases mutants.

The only confirmed genetic tool available to accomplish this is the pyrE gene.

Theoretically one can construct a double DNA polymerases mutant by transforming a Sso pyrE cassette into a single polymerase mutant to disrupt a second DNA polymerase; however, there is a very good chance that this new Sso pyrE cassette will integrate with the old Sso pyrE gene that was used to construct the single polymerase mutant instead of integrating with the target second DNA polymerase. Therefore, the need of developing new genetic tools in Sulfolobus is widely recognized. According to recent studies, there are some selectable markers developed in

Sulfolobus spp; for example, argD gene is required for agmatine prototroph in S. islandicus; herA gene in S. islandicus functions for simvastatin resistance as well (Zhang et al, 2013; Zheng et al, 2012). Therefore, it should be possible to construct those corresponding genes knockout mutants in S. acidocaldarius pol 2 or pol 3 disruptant first and then use same gene as selectable marker to disrupt the second polymerase.

With regard to damage tolerance, AP sites seem to be removed frequently in vivo; however, 8-oxoG can be present and bypassed efficiently as well as accurately. I tested the

50 capacity of S. acidocaldarius to bypass abasic site lesion; however, electroporation yielded only a few transformants, perhaps because abasic site is cleaved by endonuclease in vivo, and results in extremely low transformation efficiency. This observation, however, shows that

Sulfolobus may have strict screening systems to remove abasic sites. To investigate how

Sulfolobus deal with unrepaired DNA damages, it is reasonable to focus on other DNA lesions or multiple enzyme deficient cells that lose abilities of repairing abasic sites. To transform a gapped pDM8 plasmid with 8-oxoguanine (8-oxoG) DNA lesion into wild type strain is a good replacement for the abasic site experiment since the 8-oxoG oligo transformation was proved to be more efficient than that of abasic site (Sakofsky & Grogan, unpublished) and I have already obtained preliminary date of 8-oxoG gapped pDM8 bypass events. The analysis of 8- xoxG bypass in WT strain showed that the insertion of “C” opposite 8-oxoG were the dominant results, which indicated that S. acidocaldarius can bypass 8-oxoG accurately in vivo and this was consistent with 8-oxoG oligo transformation experiment (Sakofsky & Grogan, unpublished) as well as in yeast (Rodriguez et al, 2013) but not in E. coli (Shikazono et al, 2013). These results altogether implies that Sulfolobus may resemble eukaryotes in DNA lesion bypass process. Also as a contribution to the broader understanding of Sulfolobus TLS, it will be helpful if some people use the single polymerase mutant constructed already or the multiple polymerases mutant obtained in future as recipient cells for the gapped plasmid transformation experiments.

Finally, as to the mechanism of highly expressed gene generating frequent spontaneous mutations, it appears that the single-stranded DNA structure is not the primary cause of DNA repair systems failure to function. Transcription associated mutagenesis (TAM) was observed on a plasmid-borne lacS (pJlacS) gene in S. acidocaldarius and I tested one hypothetical

51 mechanism which is the unusual single-stranded DNA structure leads to frequent spontaneous mutations. Compared to the positive control (intact pClacS), the construction of large gapped- pClacS did result in higher spontaneous mutation rate; however, it was not possible to construct complete mutation spectrum because of low transformation efficiency. Based on the mutations I have so far, they may not resemble those previously observed on pJlacS in terms of mutation categories for three reasons: i) there was no insertion and deletion mutations observed from gapped pClacS, , but these make up about one-third of the lacS mutations seen in this same plasmid under TAM conditions, ii) the mutations found in the lacS gene after strand removal have a different spatial pattern than those induced in the same plasmid under TAM conditions, iii) neither of the BPS mutations from single-stranded plasmid generated stop codons, but all of the BPS mutations generated stop codons in pClacS under TAM conditions .

Thus the single-stranded DNA structure appears not to be the reason that causes the TAM observed on pJlacS and there are necessities to exam other explanations. Therefore, the mechanism of transcription-associated mutagenesis in Sulfolobus remains mysterious and it will be reasonable to think that not only one mechanism functions behind this.

It is therefore important to test more hypotheses and try to interconnect these tests.

One possible experiment is to test the topoisomerase 1, and in S. acidocaldarius topoisomerase1 is located at the chromosomes (Saci_1371). People can design PCR primers with flanks of topoisomerase 1 to amplify pyrE gene and transform this cassette to disrupt topoisomerase 1. Confirmed topoisomerase 1 mutants can be used as recipients for transformation of pJlacS and pClacS. Mutation rate and spectra of lacS gene will be compared

52 with those observed in wild type strain. This will demonstrate whether some specific enzymes play significant roles in TAM.

Reference

1. Allers T, Mevarech M: Archaeal genetics - the third way. Nat Rev Genet 2005, 6(1):58-73. 2. Ambur OH, Davidsen T, Frye SA, Balasingham SV, Lagesen K, Rognes T, Tonjum T: Genome dynamics in major bacterial pathogens. FEMS Microbiol Rev 2009, 33(3):453-470. 3. Atomi H, Matsumi R, Imanaka T: Reverse Gyrase Is Not a Prerequisite for Hyperthermophilic Life. J Bacteriology 2004, 186(14):4829-4833. 4. Banasik M, Sachadyn P: Conserved motifs of MutL proteins. Mutat Res 2014, 769:69-79. 5. Bauer RJ, Begley MT, Trakselis MA: Kinetics and fidelity of polymerization by DNA polymerase III from Sulfolobus solfataricus. Biochemistry 2012, 51(9):1996-2007. 6. Beletskii A, Bhagwat AS: Transcription-induced mutations: increase in C to T mutations in the nontranscribed strand during transcription in Escherichia coli. Proc Natl Acad Sci U S A 1996, 93(24):13919-13924. 7. Berkner S, Grogan D, Albers SV, Lipps G: Small multicopy, non-integrative shuttle vectors based on the plasmid pRN1 for Sulfolobus acidocaldarius and Sulfolobus solfataricus, model organisms of the (cren-)archaea. Nucleic Acids Res 2007, 35(12):e88. 8. Berkner S, Lipps G: Characterization of the transcriptional activity of the cryptic plasmid pRN1 from Sulfolobus islandicus REN1H1 and regulation of its replication operon. J Bacteriol 2007, 189(5):1711-1721. 9. Berkner S, Wlodkowski A, Albers SV, Lipps G: Inducible and constitutive promoters for genetic systems in Sulfolobus acidocaldarius. Extremophiles 2010, 14(3):249-259. 10. Berquist BR, McNeill DR, Wilson DM: Characterization of abasic endonuclease activity of human Ape1 on alternative substrates, as well as effects of ATP and sequence context on AP site incision. J Mol Biol 2008, 379(1):17-27. 11. Boiteux S, Guillet M: Abasic sites in DNA: repair and biological consequences in Saccharomyces cerevisiae. DNA Repair (Amst) 2004, 3(1):1-12. 12. Boudsocq F, Iwai S, Hanaoka F, Woodgate R: Sulfolobus solfataricus P2 DNA polymerase IV (Dpo4): an archaeal DinB-like DNA polymerase with lesion-bypass properties akin to eukaryotic poleta. Nucleic Acids Res 2001, 29(22):4607-4616. 13. Cann IK, Komori K, Toh H, Kanai S, Ishino Y: A heterodimeric DNA polymerase: evidence that members of Euryarchaeota possess a distinct DNA polymerase. Proc Natl Acad Sci U S A 1998, 95(24):14250-14255. 14. Cariello NF, Piegorsch WW, Adams WT, Skopek TR: Computer program for the analysis of mutational spectra: application to p53 mutations. Carcinogenesis 1994, 15(10):2281-2285. 15. Cerritelli SM, Crouch RJ: Ribonuclease H: the enzymes in eukaryotes. FEBS J 2009, 276(6):1494- 1505. 16. Chen LM, Brugger K, Skovgaard M, Redder P, She QX, Torarinsson E, Greve B, Awayez M, Zibat A, Klenk HP et al: The genome of Sulfolobus acidocaldarius, a model organism of the Crenarchaeota. J Bacteriology 2005, 187(14):4992-4999. 17. Cheng KC, Cahill DS, Kasai H, Nishimura S, Loeb LA: 8-Hydroxyguanine, an abundant form of oxidative DNA damage, causes G----T and A----C substitutions. J Biol Chem 1992, 267(1):166- 172. 18. Choi JY, Eoff RL, Pence MG, Wang J, Martin MV, Kim EJ, Folkmann LM, Guengerich FP: Roles of the four DNA polymerases of the crenarchaeon Sulfolobus solfataricus and accessory proteins in DNA replication. J Biol Chem 2011, 286(36):31180-31193. 19. Datta A, Jinks-Robertson S: Association of increased spontaneous mutation rates with high levels of transcription in yeast. Science 1995, 268(5217):1616-1619.

20. de Koning B, Blombach F, Brouns SJ, van der Oost J: Fidelity in archaeal information processing. Archaea 2010, 2010. 21. de Visser JA, Rozen DE: Limits to adaptation in asexual populations. J Evol Biol 2005, 18(4):779- 788. 22. Deng L, Zhu H, Chen Z, Liang YX, She Q: Unmarked gene deletion and host-vector system for the hyperthermophilic crenarchaeon Sulfolobus islandicus. Extremophiles 2009, 13(4):735-746. 23. DiRuggiero J, Brown JR, Bogert AP, Robb FT: DNA repair systems in archaea: mementos from the last universal common ancestor? J Mol Evol 1999, 49(4):474-484. 24. Dorazi R, Gotz D, Munro S, Bernander R, White MF: Equal rates of repair of DNA photoproducts in transcribed and non-transcribed strands in Sulfolobus solfataricus. Mol Microbiol 2007, 63(2):521-529. 25. Drake JW: Spontaneous mutation. Annu Rev Genet 1991, 25:125-146. 26. Drolet M, Phoenix P, Menzel R, Masse E, Liu LF, Crouch RJ: Overexpression of RNase H partially complements the growth defect of an Escherichia coli delta topA mutant: R-loop formation is a major problem in the absence of DNA topoisomerase I. Proc Natl Acad Sci U S A 1995, 92(8):3526-3530. 27. Ellwood M, Nomura M: Chromosomal locations of the genes for rRNA in Escherichia coli K-12. J Bacteriol 1982, 149(2):458-468. 28. Fijalkowska IJ, Schaaper RM, Jonczyk P: DNA replication fidelity in Escherichia coli: a multi-DNA polymerase affair. FEMS Microbiol Rev 2012, 36(6):1105-1121. 29. Fijalkowska IJ, Schaaper RM, Jonczyk P: DNA replication fidelity in Escherichia coli: a multi-DNA polymerase affair. Fems Microbiology Reviews 2012, 36(6):1105-1121. 30. Forterre P: A hot story from comparative genomics: reverse gyrase is the only hyperthermophile-specific protein. Trends Genet 2002, 18(5):236-237. 31. Foster PL: Adaptive mutation: the uses of adversity. Annu Rev Microbiol 1993, 47:467-504. 32. Frederico LA, Kunkel TA, Shaw BR: A sensitive genetic assay for the detection of cytosine deamination: determination of rate constants and the activation energy. Biochemistry 1990, 29(10):2532-2537. 33. Friedberg EC: DNA repair and mutagenesis, 2nd edn. Washington, D.C.: ASM Press; 2006. 34. Fuchs RP, Fujii S: Translesion DNA synthesis and mutagenesis in prokaryotes. Cold Spring Harb Perspect Biol 2013, 5(12):a012682. 35. Fujikane R, Ishino S, Ishino Y, Forterre P: Genetic analysis of DNA repair in the hyperthermophilic archaeon, Thermococcus kodakaraensis. Genes Genet Syst 2010, 85(4):243- 257. 36. Furio V, Moya A, Sanjuan R: The cost of replication fidelity in human immunodeficiency virus type 1. Proc Biol Sci 2007, 274(1607):225-230. 37. Galhardo RS, Hastings PJ, Rosenberg SM: Mutation as a stress response and the regulation of evolvability. Crit Rev Biochem Mol 2007, 42(5):399-435. 38. Galtier N, Lobry JR: Relationships between genomic G+C content, RNA secondary structures, and optimal growth temperature in prokaryotes. J Mol Evol 1997, 44(6):632-636. 39. Godoy VG, Jarosz DF, Simon SM, Abyzov A, Ilyin V, Walker GC: UmuD and RecA directly modulate the mutagenic potential of the Y family DNA polymerase DinB. Mol Cell 2007, 28(6):1058-1070. 40. Grogan DW: Phenotypic Characterization of the Archaebacterial Genus Sulfolobus - Comparison of 5 Wild-Type Strains. J Bacteriology 1989, 171(12):6710-6719. 41. Grogan DW, Carver GT, Drake JW: Genetic fidelity under harsh conditions: analysis of spontaneous mutation in the thermoacidophilic archaeon Sulfolobus acidocaldarius. Proc Natl Acad Sci U S A 2001, 98(14):7928-7933.

42. Gruz P, Shimizu M, Pisani FM, De Felice M, Kanke Y, Nohmi T: Processing of DNA lesions by archaeal DNA polymerases from Sulfolobus solfataricus. Nucleic Acids Res 2003, 31(14):4024- 4030. 43. Guy L, Roten CA: Genometric analyses of the organization of circular chromosomes: a universal pressure determines the direction of ribosomal RNA genes transcription relative to chromosome replication. Gene 2004, 340(1):45-52. 44. Hardeland U, Bentele M, Jiricny J, Schar P: The versatile thymine DNA-glycosylase: a comparative characterization of the human, Drosophila and fission yeast orthologs. Nucleic Acids Res 2003, 31(9):2261-2271. 45. Hendel A, Ziv O, Gueranger Q, Geacintov N, Livneh Z: Reduced efficiency and increased mutagenicity of translesion DNA synthesis across a TT cyclobutane pyrimidine dimer, but not a TT 6-4 photoproduct, in human cells lacking DNA polymerase eta. DNA Repair (Amst) 2008, 7(10):1636-1646. 46. Herman RK, Dworkin NB: Effect of gene induction on the rate of mutagenesis by ICR-191 in Escherichia coli. J Bacteriol 1971, 106(2):543-550. 47. Heyer WD: The search for the right partner: homologous pairing and DNA strand exchange proteins in eukaryotes. Experientia 1994, 50(3):223-233. 48. Higashibata H, Fujiwara S, Ezaki S, Takagi M, Fukui K, Imanaka T: Effect of polyamines on histone-induced DNA compaction of hyperthermophilic archaea. J Biosci Bioeng 2000, 89(1):103-106. 49. Hopfner KP, Karcher A, Craig L, Woo TT, Carney JP, Tainer JA: Structural biochemistry and interaction architecture of the DNA double-strand break repair Mre11 nuclease and Rad50- ATPase. Cell 2001, 105(4):473-485. 50. Huang ME, Rio AG, Galibert MD, Galibert F: Pol32, a subunit of Saccharomyces cerevisiae DNA polymerase delta, suppresses genomic deletions and is involved in the mutagenic bypass pathway. Genetics 2002, 160(4):1409-1422. 51. Hubscher U, Maga G, Spadari S: Eukaryotic DNA polymerases. Annu Rev Biochem 2002, 71:133- 163. 52. Hurst LD, Merchant AR: High guanine-cytosine content is not an adaptation to high temperature: a comparative analysis amongst prokaryotes. Proc Biol Sci 2001, 268(1466):493- 497. 53. Iyer RR, Pluciennik A, Burdett V, Modrich PL: DNA mismatch repair: functions and mechanisms. Chem Rev 2006, 106(2):302-323. 54. Jacobs KL, Grogan DW: Rates of spontaneous mutation in an archaeon from geothermal environments. J Bacteriol 1997, 179(10):3298-3303. 55. Kawasaki Y, Sugino A: Yeast replicative DNA polymerases and their role at the replication fork. Mol Cells 2001, 12(3):277-285. 56. Kelman Z, White MF: Archaeal DNA replication and repair. Curr Opin Microbiol 2005, 8(6):669- 676. 57. Kim N, Abdulovic AL, Gealy R, Lippert MJ, Jinks-Robertson S: Transcription-associated mutagenesis in yeast is directly proportional to the level of gene expression and influenced by the direction of DNA replication. DNA Repair 2007, 6(9):1285-1296. 58. Kim SR, Matsui K, Yamada M, Gruz P, Nohmi T: Roles of chromosomal and episomal dinB genes encoding DNA pol IV in targeted and untargeted mutagenesis in Escherichia coli. Mol Genet Genomics 2001, 266(2):207-215. 59. Koonin EV: Carl Woese's vision of cellular evolution and the domains of life. RNA Biol 2014, 11(3):197-204.

60. Koonin EV, Wolf YI: Evolution of microbes and viruses: a paradigm shift in evolutionary biology? Front Cell Infect Microbiol 2012, 2:119. 61. Kurosawa N, Grogan DW: Homologous recombination of exogenous DNA with the Sulfolobus acidocaldarius genome: Properties and uses. Fems Microbiol Lett 2005, 253(1):141-149. 62. Li X, Guo L, Deng L, Feng D, Ren Y, Chu Y, She Q, Huang L: Deletion of the topoisomerase III gene in the hyperthermophilic archaeon Sulfolobus islandicus results in slow growth and defects in cell cycle control. J Genet Genomics 2011, 38(6):253-259. 63. Li X, Manley JL: Inactivation of the SR protein splicing factor ASF/SF2 results in genomic instability. Cell 2005, 122(3):365-378. 64. Liang PJ, Han WY, Huang QH, Li YZ, Ni JF, She QX, Shen YL: Knockouts of RecA-Like Proteins RadC1 and RadC2 Have Distinct Responses to DNA Damage Agents in Sulfolobus islandicus. Journal of Genetics and Genomics 2013, 40(10):533-542. 65. Lindahl T: Instability and decay of the primary structure of DNA. Nature 1993, 362(6422):709- 715. 66. Lindahl T, Nyberg B: Heat-induced deamination of cytosine residues in deoxyribonucleic acid. Biochemistry 1974, 13(16):3405-3410. 67. Livneh Z, Ziv O, Shachar S: Multiple two-polymerase mechanisms in mammalian translesion DNA synthesis. Cell Cycle 2010, 9(4):729-735. 68. Loh E, Salk JJ, Loeb LA: Optimization of DNA polymerase mutation rates during bacterial evolution. Proc Natl Acad Sci U S A 2010, 107(3):1154-1159. 69. Makarova KS, Koonin EV, Kelman Z: The CMG (CDC45/RecJ, MCM, GINS) complex is a conserved component of the DNA replication system in all archaea and eukaryotes. Biol Direct 2012, 7. 70. Makarova KS, Krupovic M, Koonin EV: Evolution of replicative DNA polymerases in archaea and their contributions to the eukaryotic replication machinery. Front Microbiol 2014, 5. 71. Mao D, Grogan D: Genomic evidence of rapid, global-scale gene flow in a Sulfolobus species. ISME J 2012, 6(8):1613-1616. 72. McCulloch SD, Kokoska RJ, Garg P, Burgers PM, Kunkel TA: The efficiency and fidelity of 8-oxoguanine bypass by DNA polymerases delta and eta. Nucleic Acids Res 2009, 37(9):2830-2840. 73. Melis JP, van Steeg H, Luijten M: Oxidative DNA damage and nucleotide excision repair. Antioxid Redox Signal 2013, 18(18):2409-2419. 74. Miller RV, Kokjohn TA: General microbiology of recA: environmental and evolutionary significance. Annu Rev Microbiol 1990, 44:365-394. 75. Minnick DT, Pavlov YI, Kunkel TA: The fidelity of the human leading and lagging strand DNA replication apparatus with 8-oxodeoxyguanosine triphosphate. Nucleic Acids Res 1994, 22(25):5658-5664. 76. Nelson JR, Lawrence CW, Hinkle DC: Thymine-thymine dimer bypass by yeast DNA polymerase zeta. Science 1996, 272(5268):1646-1649. 77. Newman M, Murray-Rust J, Lally J, Rudolf J, Fadden A, Knowles PP, White MF, McDonald NQ: Structure of an XPF endonuclease with and without DNA suggests a model for substrate recognition. Embo Journal 2005, 24(5):895-905. 78. Pomerantz RT, O'Donnell M: What happens when replication and transcription complexes collide? Cell Cycle 2010, 9(13):2537-2543. 79. Rajpal DK, Wu XH, Wang ZG: Alteration of ultraviolet-induced mutagenesis in yeast through molecular modulation of the REV3 and REV7 gene expression. Mutat Res-DNA Repair 2000, 461(2):133-143.

80. Rangarajan S, Woodgate R, Goodman MF: A phenotype for enigmatic DNA polymerase II: a pivotal role for pol II in replication restart in UV-irradiated Escherichia coli. Proc Natl Acad Sci U S A 1999, 96(16):9224-9229. 81. Rattray AJ, Strathern JN: Error-prone DNA polymerases: When making a mistake is the only way to get ahead. Annu Rev Genet 2003, 37:31-66. 82. Reaban ME, Lebowitz J, Griffin JA: Transcription induces the formation of a stable RNA.DNA hybrid in the immunoglobulin alpha switch region. J Biol Chem 1994, 269(34):21850-21857. 83. Resende BC, Rebelato AB, D'Afonseca V, Santos AR, Stutzman T, Azevedo VA, Santos LL, Miyoshi A, Lopes DO: DNA repair in Corynebacterium model. Gene 2011, 482(1-2):1-7. 84. Rodriguez GP, Song JB, Crouse GF: In vivo bypass of 8-oxodG. PLoS Genet 2013, 9(8):e1003682. 85. Rothwell PJ, Waksman G: Structure and mechanism of DNA polymerases. Adv Protein Chem 2005, 71:401-+. 86. Rudolph CJ, Dhillon P, Moore M, Lloyd RG: Avoiding and resolving conflicts between DNA replication and transcription. DNA Repair 2007, 6(7):981-993. 87. Rudolph CJ, Upton AL, Briggs GS, Lloyd RG: Is RecG a general guardian of the bacterial genome? DNA Repair (Amst) 2010, 9(3):210-223. 88. Sakofsky CJ, Foster PL, Grogan DW: Roles of the Y-family DNA polymerase Dbh in accurate replication of the Sulfolobus genome at high temperature. DNA Repair (Amst) 2012, 11(4):391- 400. 89. Sakofsky CJ, Grogan DW: Endogenous mutagenesis in recombinant sulfolobus plasmids. J Bacteriol 2013, 195(12):2776-2785. 90. Sakofsky CJ, Runck LA, Grogan DW: Sulfolobus Mutants, Generated via PCR Products, Which Lack Putative Enzymes of UV Photoproduct Repair. Archaea-an International Microbiological Journal 2011. 91. Sartori AA, Schar P, Fitz-Gibbon S, Miller JH, Jiricny J: Biochemical characterization of uracil processing activities in the hyperthermophilic archaeon Pyrobaculum aerophilum. J Biol Chem 2001, 276(32):29979-29986. 92. Shikazono N, Akamatsu K, Takahashi M, Noguchi M, Urushibara A, O'Neill P, Yokoya A: Significance of DNA polymerase I in in vivo processing of clustered DNA damage. Mutat Res 2013, 749(1-2):9-15. 93. Shin DS, Pellegrini L, Daniels DS, Yelent B, Craig L, Bates D, Yu DS, Shivji MK, Hitomi C, Arvai AS et al: Full-length archaeal Rad51 structure and mutants: mechanisms for RAD51 assembly and control by BRCA2. EMBO J 2003, 22(17):4566-4576. 94. Shin DS, Pratt AJ, Tainer JA: Archaeal Genome Guardians Give Insights into Eukaryotic DNA Replication and Damage Response Proteins. Archaea-an International Microbiological Journal 2014. 95. Srivatsan A, Tehranchi A, MacAlpine DM, Wang JD: Co-orientation of replication and transcription preserves genome integrity. PLoS Genet 2010, 6(1):e1000810. 96. Terui Y, Ohnuma M, Hiraga K, Kawashima E, Oshima T: Stabilization of nucleic acids by unusual polyamines produced by an extreme thermophile, Thermus thermophilus. Biochem J 2005, 388(Pt 2):427-433. 97. Tuduri S, Crabbe L, Conti C, Tourriere H, Holtgreve-Grez H, Jauch A, Pantesco V, De Vos J, Thomas A, Theillet C et al: Topoisomerase I suppresses genomic instability by preventing interference between replication and transcription. Nat Cell Biol 2009, 11(11):1315-U1125. 98. Vaisman A, Woodgate R: Unique misinsertion specificity of poliota may decrease the mutagenic potential of deaminated cytosines. EMBO J 2001, 20(22):6520-6529.

99. Wagner J, Etienne H, Janel-Bintz R, Fuchs RP: Genetics of mutagenesis in E. coli: various combinations of translesion polymerases (Pol II, IV and V) deal with lesion/sequence context diversity. DNA Repair (Amst) 2002, 1(2):159-167. 100. Wallace SS: Base excision repair: a critical player in many games. DNA Repair (Amst) 2014, 19:14-26. 101. Wang F, Yang W: Structural Insight into Translesion Synthesis by DNA Pol II. Cell 2009, 139(7):1279-1289. 102. Woese CR, Fox GE: Phylogenetic structure of the prokaryotic domain: the primary kingdoms. Proc Natl Acad Sci U S A 1977, 74(11):5088-5090. 103. Woese CR, Kandler O, Wheelis ML: Towards a natural system of organisms: proposal for the domains Archaea, Bacteria, and Eucarya. Proc Natl Acad Sci U S A 1990, 87(12):4576-4579. 104. Wong JH, Brown JA, Suo Z, Blum P, Nohmi T, Ling H: Structural insight into dynamic bypass of the major cisplatin-DNA adduct by Y-family polymerase Dpo4. EMBO J 2010, 29(12):2059-2069. 105. Woods WG, Dyall-Smith ML: Construction and analysis of a recombination-deficient (radA) mutant of Haloferax volcanii. Mol Microbiol 1997, 23(4):791-797. 106. Wright BE, Longacre A, Reimers JM: Hypermutation in derepressed operons of Escherichia coli K12. Proc Natl Acad Sci U S A 1999, 96(9):5089-5094. 107. Yamagishi A, Tanimoto T, Suzuki T, Oshima T: Pyrimidine biosynthesis genes (pyrE and pyrF) of an extreme thermophile, Thermus thermophilus. Appl Environ Microbiol 1996, 62(6):2191-2194. 108. Zan H, Komori A, Li Z, Cerutti A, Schaffer A, Flajnik MF, Diaz M, Casali P: The translesion DNA polymerase zeta plays a major role in Ig and bcl-6 somatic hypermutation. Immunity 2001, 14(5):643-653. 109. Zhang C, Cooper TE, Krause DJ, Whitaker RJ: Augmenting the genetic toolbox for Sulfolobus islandicus with a stringent positive selectable marker for agmatine prototrophy. Appl Environ Microbiol 2013, 79(18):5539-5549. 110. Zhang L, Brown JA, Newmister SA, Suo Z: Polymerization fidelity of a replicative DNA polymerase from the hyperthermophilic archaeon Sulfolobus solfataricus P2. Biochemistry 2009, 48(31):7492-7501. 111. Zheng T, Huang Q, Zhang C, Ni J, She Q, Shen Y: Development of a simvastatin selection marker for a hyperthermophilic acidophile, Sulfolobus islandicus. Appl Environ Microbiol 2012, 78(2):568-574. 112. Zhu L, Li Q: [Stress-induced cellular adaptive mutagenesis]. Yi Chuan 2014, 36(4):327-335.