Thesis Template

An investigation into the effect of DNA structural polymorphism and single-stranded DNA binding proteins on repair of disease-associated slipped-DNA repeats

Jennifer Jing Luo

A thesis submitted in conformity with the requirements for the degree of Master of Science Molecular Genetics University of Toronto

An investigation into the effect of DNA structural polymorphism and single-stranded DNA binding proteins on repair of disease- associated slipped-DNA repeats

Jennifer Jing Luo

Master of Science

Molecular Genetics University of Toronto

2015 Abstract

Gene-specific repeat expansions are the cause of a growing list of neurological diseases, including myotonic dystrophy type 1 and Huntington's disease. The formation of slipped-DNA structures in the expanded repeat sequences is thought to drive repeat instability and pathogenesis by impairing normal DNA metabolic processes. Here I show that slipped-DNAs with nicks located within the repeat tract displayed increased structural heterogeneity relative to slipped-DNAs with nicks located in the flanking sequence. Nick-in-repeat slipped-DNAs were repaired better than nick-in-flank slipped-DNAs, likely due to increased amounts of single- stranded DNA at the nicked repeat ends allowing for better repair factor binding. Single-stranded

DNA binding proteins RPA and aRPA seem to play an important part in tissue-specific instability as both complexes are overexpressed in the brains of HD patients. Neither RPA nor aRPA was required for slipped-DNA repair, although they both enhanced slipped-DNA repair efficiency.

Acknowledgments

I would like to thank my supervisor Dr. Christopher Pearson for his guidance, enthusiasm and encouragement over the years. I have learned much about science and life in your lab and I will always be grateful for the opportunities that you have given me. I would like to thank my committee members, Dr. David Bazett-Jones and Dr. Irene Andrulis, for the much-appreciated advice and feedback that you have always given me. I would like to thank our collaborators, especially Dr. Marc Wold, who has provided valuable resources and expertise for my project.

A big thank you for all the members of the Pearson lab, both past and present, for the help and support you have given me. I am so grateful that I had such wonderful colleagues to depend on, to turn to for help, and most importantly I am grateful for the laughs. I would like to thank all of my GGB and Mol Gen friends who all contributed to make my graduate school experience a fun and productive one. Thank you to my friends and family who gave me love, support and encouragement.

Thank you to my parents, Hui Luo and Aishe Sun. I am so grateful for everything that you have given me and all the sacrifices you have made for me. I hope to make you proud in everything I do.

Thank you to Petro, the best partner-in-life I could ask for. Thank you for your love and support. Thank you for feeding me when I forget to eat. Thank you for making me sleep when work becomes all-consuming. Thank you for the walks and the heated debates. Most importantly, thank you for making me want to be better in everything I do.

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Table of Contents

Acknowledgments ...... iii

List of Tables ...... vii

List of Figures ...... viii

List of Abbreviations ...... ix

1 Introduction ...... 1

1.1 Disease-causing trinucleotide repeats ...... 1

1.2 Pathogenic mechanisms in trinucleotide repeat diseases ...... 3

1.2.1 Gain of toxic protein function: Huntington’s disease ...... 3

1.2.2 Gain of toxic RNA function: myotonic dystrophy type 1 ...... 5

1.3 DNA structures and repeat instability ...... 6

1.3.1 CTG/CAG slipped-DNA ...... 7

1.4 Mechanisms of repeat instability ...... 9

1.4.1 DNA repair and repeat instability ...... 11

1.4.2 DNA replication and repeat instability ...... 14

1.5 Replication protein A: eukaryotic singled-stranded DNA binding protein ...... 16

1.5.1 RPA in DNA replication ...... 16

1.5.2 RPA in nucleotide excision repair ...... 17

1.5.3 RPA in base excision repair ...... 17

1.5.4 RPA in mismatch repair ...... 18

1.5.5 An alternative form of RPA: aRPA ...... 18

1.6 Thesis Goals ...... 19

2 Effect of DNA structural hyper-polymorphism and single-stranded DNA binding proteins on repair of disease-associated slipped-DNA repeats ...... 22

2.1 Introduction ...... 22

2.2 Materials and Methods ...... 25

2.2.1 Slipped-intermediate DNA (SI-DNA) substrate preparation ...... 25

2.2.2 Human cell extract preparation ...... 26

2.2.3 SV40 DNA replication ...... 27

2.2.4 In vitro DNA repair ...... 27

2.2.5 Western blotting ...... 28

2.2.6 DNA binding assays ...... 29

2.3 Results ...... 29

2.3.1 Slipped-DNAs with increased single-stranded potential ...... 29

2.3.2 RPA4 expression is elevated above RPA2 in HD patient brains ...... 38

2.3.3 RPA and aRPA can enhance slipped-DNA repair ...... 40

2.3.4 Repair is nick-directed ...... 45

2.3.5 Yeast RPA and bacterial SSB cannot substitute for human RPA in enhancing repair ...... 48

2.3.6 Inhibition of slipped-DNA repair by high concentrations of aRPA ...... 50

2.3.7 RPA is limiting for the repair of nick-in-repeat slipped-DNAs, this effect is independent of MutSβ ...... 53

2.3.8 FEN1 is not required for slipped-DNA repair ...... 56

2.3.9 RPA and aRPA bind to and melt slipped-DNAs differently ...... 59

2.4 Conclusions and Discussion ...... 63

2.4.1 Nick-in-repeat slip-outs have greater structural heterogeneity than nick-in- flank slip-outs ...... 63

2.4.2 Nick-in-repeat slip-outs are better repaired ...... 63

2.4.3 RPA and aRPA can enhance slipped-DNA repair ...... 65

2.4.4 Inhibition of slipped-DNA repair by high concentrations of aRPA ...... 67

2.4.5 Repair of large slipped-DNAs is independent of MMR and BER ...... 68

2.4.6 MMR does not require RPA ...... 69

2.4.7 RPA and aRPA bind to and denature slipped-DNAs differently ...... 69

2.4.8 aRPA function ...... 70

3 Summary and future directions ...... 72

3.1 Thesis summary ...... 72

3.2 Future Directions ...... 73

3.2.1 What are the effects of sequence on repair of TNR slip-outs? ...... 73

3.2.2 What causes tissue-specific repeat instability? ...... 74

3.2.3 How are long slip-outs repaired? ...... 75

3.2.4 What are potential therapies? ...... 76

3.3 Concluding remarks ...... 77

References ...... 79

List of Tables

Table 1-1. Common repeat-associated disease...... 2

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List of Figures

Figure 1-1. Formation of slipped-DNA structures...... 8

Figure 1-2. Eukaryotic mismatch repair pathway...... 10

Figure 1-3. Slipped-DNA repair outcomes...... 12

Figure 1-4. Features of slipped-DNAs affect their repair outcome...... 15

Figure 2-1. Nick-in-repeat slipped-DNAs have increased single-stranded potential...... 31

Figure 2-2. Nick-in-repeat slip-outs have greater structural heterogeneity than nick-in-flank slip- outs...... 33

Figure 2-3. Repair of slipped-DNAs depends on cell extract concentration and nick location. ... 35

Figure 2-4. RPA4 expression is elevated above RPA2 in HD patient brains...... 37

Figure 2-5. SV40 DNA replication requires RPA, but not aRPA...... 39

Figure 2-6. RPA is not required for repair, but RPA and aRPA can both enhance repair...... 41

Figure 2-7. Repair of G-T mismatches and small slip-outs do not require RPA...... 43

Figure 2-8. Repair is nick-directed...... 46

Figure 2-9. Yeast RPA and bacterial SSB cannot substitute for human RPA in enhancing repair...... 49

Figure 2-10. aRPA inhibits RPA at higher concentrations...... 51

Figure 2-11. Repair of small slip-outs, but not large slip-outs, is sensitive to MutSβ concentration...... 55

Figure 2-12. FEN1 is not required for slipped-DNA repair...... 57

Figure 2-13. RPA and aRPA bind to and melt slipped-DNA differently...... 60

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List of Abbreviations

ALS amyotrophic lateral sclerosis aRPA alternative replication protein A BER base excision repair bp base pair CNS central nervous system DM1 myotonic dystrophy type 1 DMPK dystrophica myotonica protein kinase gene DNA deoxyribonucleic acid dNTP deoxynucleoside triphosphate E. coli Escherichia coli EMSA electrophoretic mobility shift assay FMR1 fragile X mental retardation gene 1 FRAXA fragile X syndrome A FRDA Friedreich’s ataxia HD Huntington’s disease IDL insertion-deletion loop LLP large loop repair MBNL1 muscleblind protein MMR mismatch repair MSH2/3/6 MutS homolog 2/3/6 NER nucleotide excision repair nt nucleotide OGG1 oxoguanineglycosylase 1 protein pol polymerase RNA ribonucleic acid rNTP ribonucleoside triphosphate RPA replication protein A SCA spinocerebellar ataxia S-DNA slipped-DNA (homoduplex) SI-DNA slipped intermediate-DNA (heteroduplex) ssDNA single-stranded DNA SV40 simian virus 40 TC-NER transcription-coupled nucleotide excision repair TNR trinucleotide repeat UTR untranlated region

ix 1

1 Introduction 1.1 Disease-causing trinucleotide repeats

Gene-specific repeat expansions are the cause of an ever-growing number of disorders. At the present time there are more than 40 known neurological, neurodegenerative and neuromuscular inherited disorders that are associated with changes in DNA repeat tract length. Changes in the number of repeat units in a tract within a particular gene can alter its expression and the function of the RNA and/or protein it encodes. Disease-causing repetitive sequences include trinucleotides, tetranucleotides, pentanucleotides, minisatellites and megasatellites. Of these repetitive sequences, trinucleotide repeats (TNRs) are the most common unstable disease- associated DNA repeats and the focus of my thesis.

TNRs contribute to a wide range of disorders including myotonic dystrophy type 1 (DM1) and Huntington’s disease (HD). Each of these diseases is clinically distinct and involves expansion of a TNR within a transcribed gene in the human genome (See Table 1-1 for examples of the various repeat unit lengths and sequences that are associated with different diseases). TNR mutations can occur at various locations within a gene, from the promoter region (i.e. the (CAG)n repeat of spinocerebellar ataxia type 12); the 5’ untranslated region (UTR) (i.e. the (CGG)n of fragile X type A); exons (i.e. the (CAG)n of HD and SCA7); introns (i.e. the (GAA)n of Freidreich’s ataxia); and the 3’ UTR (i.e. the (CTG)n repeats of DM1). In most cases, the unaffected population harbors a short repeat (generally less than 35) that remains stable throughout each individual’s life and across generations. Expansions beyond the disease-specific threshold length (typically 35 repeat units) result in repeat tract length instability (expansions and contractions of repeat units within the tract) across a patient’s lifetime at different rates in different tissues (somatic instability) and across generations through instability in gametes (germ line instability) (Pearson, Nichol Edamura, and Cleary 2005, López Castel, Cleary, and Pearson 2010). The repeat instability in each disease is locus-specific, with no generalized genomic instability. The cause of tandem repeat tract expansions into disease-associated lengths is currently unknown despite extensive investigation.

Table 1-1

Normal Disease Gene Premutation Disease Gene Repeat sequence tract tract location tract length* length* length*

Myotonic dystrophy DMPK (CTG)•(CAG) 3’ UTR 5-37 34-90 90-6500 type 1 (DM1)

Amyotrophic lateral sclerosis-frontotemporal C9orf72 (GGGGCC)•(GGCCCC) Intron 2-22 ND 23-4400 dementia (ALS-FTD)

Fragile X syndrome type FMR1 (CGG)•(CCG) 5’ UTR 6-52 59-230 230-2000 A (FRAXA)

Huntington’s disease Coding HTT (CAG)•(CTG) 10-34 29-35 >35 (HD) exon

Spinal and bulbar Coding muscular atrophy AR (CAG)•(CTG) 9-36 ND 40-55 exon (SBMA)

Spinocerebellar ataxia Coding Ataxin1 (CAG)•(CTG) 6-39 ND 39-81 type 1 (SCA1) exon

Friedreich’s ataxia FXN (GAA)•(TTC) Intron 6-32 40-200 >200 (FRDA)

Table 1-1. Common repeat-associated disease.

“ND” denotes not determined due to insufficient data or inconsistent reporting. “*” indicates that repeat lengths may overlap between normal, permutation and disease lengths without necessarily causing disease. Modified from (Pearson, Nichol Edamura, and Cleary 2005).

In general, TNR diseases display genetic anticipation, which is the clinical observation that the disease has an earlier age-of-onset in subsequent generations, usually accompanied by symptoms that are more severe than the previous affected generation. TNR expansions have been shown to be unstable within an individual, with various tissues showing different repeat tract lengths (López Castel et al. 2011, Taylor et al. 1999, Tanaka et al. 1996, Benitez et al. 1995). The mechanisms of repeat instability are not fully clear, although there is evidence that certain DNA metabolic processes are involved. These processes include DNA replication, transcription, repair, as well as alternative DNA structure formation (see “Mechanisms of Instability” below).

1.2 Pathogenic mechanisms in trinucleotide repeat diseases

Trinucleotide repeat diseases develop through expansions of a gene-specific repeat tract resulting in four recognized mechanisms. Expansions may result in 1) loss of transcription resulting in reduced protein as in fragile X type A (FRAXA) and Friedreich’s ataxia, 2) gain of toxic protein function as in Huntington’s disease and various spinocerebellar ataxias, as well as 3) gain of toxic RNA function (protein sequestration) as in myotonic dystrophy types 1 and 2, fragile X tremor ataxia syndrome and amyotrophic lateral sclerosis and frontotemporal dementia (Pearson, Nichol Edamura, and Cleary 2005, López Castel, Cleary, and Pearson 2010, van Blitterswijk, DeJesus-Hernandez, and Rademakers 2012, Nelson, Orr, and Warren 2013). More recently 4) non-canonical RAN (repeat associated non-ATG) translation was identified and several repeat expansions have been linked to this process though roles in pathogenesis remain unclear (Zu et al. 2011, Mori et al. 2013). The gain of toxic protein function and gain of toxic RNA function mechanisms are described in detail below.

1.2.1 Gain of toxic protein function: Huntington’s disease

The hereditary nature of chorea was noted in the 19th century by several doctors, but George Huntington’s vivid description led to the designation of the disorder as Huntington’s disease (Huntington 2003). In 1993 The Huntington’s Disease Collaborative Research Group discovered that this disorder is caused by a (CAG) repeat expansion in the first exon of the HTT gene located on the short arm of chromosome 4 (1993). HD is an autosomal dominant neurodegenerative disorder which usually presents in adult life with involuntary movements, cognitive decline, psychiatric problems and progression to death 15-20 years from time of onset.

Although individual with HD typically become symptomatic in midlife, symptoms can present as early as age 1 and as late as age 80 or more (Myers 2004).

Like DM1, HD has a range of repeat sizes associated with the unaffected and affected populations. Unaffected individuals can have up to 35 CAG repeats, which are intergenerationally and somatically stable. Premutation lengths are between 29 and 35 repeats. CAG repeat expansions that are longer than 35 repeats result in disease. Interestingly, a disproportionate number of HD cases with onset before the age of 21 (juvenile onset HD) had inherited the HD gene from affected fathers (Kremer et al. 1995, Ranen et al. 1995, Trottier, Biancalana, and Mandel 1994). CAG repeats that exceed 28 show instability during replication, which grows with increasing size of the repeat (Djoussé et al. 2004, MacDonald et al. 1999). Instability is also greater in spermatogenesis than oogenesis, in that larger expansions of CAG repeats during replication occur almost exclusively in males (Kremer et al. 1995, Ranen et al. 1995, Trottier, Biancalana, and Mandel 1994). These findings account for the occurrence of anticipation, in which the age of onset of HD becomes earlier in successive generations, and the likelihood of paternal inheritance in children with juvenile onset HD. Similarly, new on-set cases of HD with a negative family history typically arise because of expansion of an allele in the permutation or normal range (28-35 CAG repeats), most usually on the paternal side (Walker 2007).

The cause of HD disease pathology is mostly attributed to the toxic gain of function protein entity that is produced by the expanded polyglutamine tract (encoded by CAG repeats) within the HTT gene encoding the multifunctional huntingtin protein. The mutant huntingtin protein bearing the expanded polyglutamine tract is improperly folded leading to ribonuclear and cytoplasmic aggregation with the ability to bind other proteins (Davies et al. 1997). Although extensively studied, the link between mutant huntingtin aggregation and pathogenesis is not clear, with the central unanswered question: are mutant huntingtin protein aggregates protective or toxic to cells? Many studies have found polyglutamine aggregation to be associated with neurodegeneration (Davies et al. 1997, DiFiglia et al. 1997, Becher et al. 1998, Ordway et al. 1997). However, in other studies there was no or negative correlation. For example, in a study using a knock-in mouse model of HD, protein aggregates arose only after symptoms began (Menalled, Sison, and Dragatsis… 2003). Another study using transfected rat neurons found that cells that have aggregated proteins survived longer than those without and led to decreased levels

5 of mutant huntingtin elsewhere in the neuron (Arrasate et al. 2004). Yet another study found that a compound that enhances aggregate formation might actually lessen neuronal pathological findings (Bodner, Outeiro, and Altmann… 2006). Towards differentiating between these two possible scenarios, there is growing interest in elucidating all the normal functions of the non- mutant huntingtin protein to aid in identifying the pathogenic roles of the mutant, toxic protein.

1.2.2 Gain of toxic RNA function: myotonic dystrophy type 1

DM1 was first described by the German physician Hans Steinert in 1909, thus it is sometimes referred to as Steinert’s disease. It is an autosomal dominant neuromuscular disease and it is the most common form of muscular dystrophy affecting adults. In 1992 Brook et al. discovered that the disease is caused by a (CTG) expansion in the 3’ UTR of the myotonic dystrophy protein kinase gene (DMPK) on the long arm of chromosome 19 (Brook et al. 1992). Although DM1 is generally considered a disease of muscle, with myotonia, progressive weakness and wasting, the disease affects multiple systems in the body. Additional symptoms include cardiac conduction defects, hypersomnia, cataracts, apathy, learning difficulties, and male infertility (Brook et al. 1992).

DM1 has a range of repeat sizes associated with the unaffected and affected populations, and patients exhibit symptoms at varying ages, generally dependent upon repeat length. Unaffected individuals can have up to 35 CTG repeats, which are intergenerationally and somatically stable. Premutation lengths are up to 90 or 100 repeats and unstable. Full disease-length repeats can be up to 6000 repeats (López Castel, Cleary, and Pearson 2010). Although repeat contractions are reported to have an occurrence rate of between 4.2 and 6.4% upon transmission (Musova et al. 2009), the majority of unstable alleles are expansions. The somatic instability of CTG repeats has been documented in DM1 patients, with repeat length differences being in the thousands between more affected and less affected tissues (López Castel et al. 2011). At the present time the mechanism by which this somatic instability occurs is not fully understood, despite intensive investigation.

The cause of DM1 disease pathology is mostly attributable to a toxic gain of function of the expanded (CUG)n RNA, which sequesters other proteins from their normal functions and creates foci of protein aggregates within the nucleus. One of the sequestered proteins is muscleblind (MBNL1), a protein that normally acts as a splicing factor (Miller et al. 2000, Jiang et al. 2004).

The sequestration of MBNL1 is directly connected to many of the symptoms of DM1. For example, MBNL1 is known to affect the alternative splicing of cardiac troponin t (cTNT), which is then misregulated when MBNL1 is sequestered in DM1, contributing to the cardiac symptoms seen in the disease (Philips, Timchenko, and Cooper 1998). Knockout of MBNL1 in a mouse model for DM1 recapitulated key muscle and eye phenotypes, supporting the hypothesis that sequestration of MBNL1 proteins by toxic RNA contributes to DM1 pathogenesis (Kanadia et al. 2003).

1.3 DNA structures and repeat instability

The key determinant of unstable repeat disease that falls upstream of all pathogenic features is the initial expansion of the repeat tract in the DNA (Pearson, Nichol Edamura, and Cleary 2005, López Castel, Cleary, and Pearson 2010). Once expanded, the repeat continues to expand throughout the life of the individual and at different rates in different tissues leading to worsening of disease severity. The mechanism that causes DNA expansion is not known. Repeat expansions and ongoing repeat instability are hypothesized to occur through the formation of non-Watson-Crick alternative DNA structures (Pearson, Nichol Edamura, and Cleary 2005, López Castel, Cleary, and Pearson 2010). There are three general classes of DNA structures associated with disease-causing repeat tracts: 1) slipped-DNA, 2) G-quadruplex DNA, and 3) triplex DNA. My study will focus on the formation and repair of slipped-DNAs, which are hairpins (or slip-outs) that form from out-of-register misalignment of repeats. Slipped-DNA structures have been shown to occur in the expanded (CGG)•(CCG) repeats in FRAXA and expanded (CTG)•(CAG) repeats in DM1 (Pearson and Sinden 1996). G-quadruplex DNA can form at G-rich TNRs such as the (CGG)•(CCG) repeats in FRAXA (Fry and Loeb 1994). G- quadruplex structures involve non-canonical hydrogen base pairing between four guanine residues either in DNA or RNA in a planar configuration to form G-quartets. The G-quartets stack upon each other to form G-qudruplexes either within a molecule or between multiple molecules. G-quadruplex structures are known to form at other G-rich sequences such as telomere repeats, promoters, immunoglobulin class switch repeats (Lipps and Rhodes 2009, Wu and Brosh 2010) and the (GGGGCC)•(GGCCCC) hexanuclotide repeat associated with amyotrophic lateral sclerosis (Reddy et al. 2013). Triplex DNA (or triple helical DNA) involves 3 DNA strands of an alternating polypurine, polypyrimidine sequence tract, where one single strand invades and pairs with a Watson-Crick duplex through Hoogsteen hydrogen bonding.

Triplex DNA structures have been shown in the expanded (GAA)•(TTC) repeats associated with Friedreich’s ataxia (FRDA) (Sakamoto et al. 1999).

1.3.1 CTG/CAG slipped-DNA

Based on the observation that repeat expansions associated with disease can be relatively large and heterogeneous, where patients may have between tens and thousands of repeats, it was hypothesized that unusual DNA structures are being formed in the repetitive DNA tracts through denaturation and renaturation of the two DNA strands during DNA metabolism that causes replication and repair errors leading to instability (Pearson and Sinden 1996). Due to the highly repetitive nature of the complementary repeat strands, perfect reannealing may be difficult thus leading to misalignment and the formation of abnormal structures. This hypothesis was tested by modeling the reannealing of the complementary DNA strands at a repeat sequence in vitro using plasmids bearing expanded trinucleotide repeat tracts. Denaturation and renaturation of this repeat-containing DNA as would occur during any DNA metabolic process results in the formation of stable alternative structures termed slipped-DNAs (S-DNA) with a melting temperature higher than physiological temperature in mammalian cells (Pearson and Sinden 1996, Gacy et al. 1995, Petruska, Arnheim, and Goodman 1996). Analyses of (CTG)•(CAG) repeat tracts using a combination of gel electrophoresis, single-strand specific nucleases (such as mung bean nuclease and T7 endonuclease), antibody probing and electron microscopy revealed intrastrand hairpin formation in the CTG strand and coiled structures in the CAG strand of the S- DNA. The extent of slipped-DNA positively correlates with repeat tract length as well as disease severity (Pearson, Eichler, et al. 1998, Pearson and Sinden 1996, Pearson et al. 2002, Tam et al. 2003). Further supporting the hypothesis that alternative DNA structures that form within repeat tracts contribute to repeat instability, work from our lab has shown that S-DNAs are found at the expanded allele of the DM1 locus in patient tissues and the level of S-DNA detected in different tissues correlates with repeat instability (Axford et al. 2013).

Slipped-DNAs formed by out-of-register misalignment of the repeats have long been thought to be transient mutagenic intermediates that arise during DNA metabolic processes such as DNA replication, repair, transcription or recombination (Figure 1-1). Characterization and understanding of the formation of slipped-DNA structures, as well as which factors might be involved in their processing, are important for understanding the mechanisms of repeat-

Figure 1-1

Figure 1-1. Formation of slipped-DNA structures.

Slip-outs can form at repeat tracts during various DNA metabolic processes including DNA replication, repair, transcription, and recombination. Depending upon the location of the slip-out, the out-of-register pairing can lead to either contraction or expansion mutations. Modified from (Axford et al. 2013).

9 associated diseases. Likewise, disease therapy might be aimed at compounds that specifically recognize these structures.

1.4 Mechanisms of repeat instability

The mechanisms that cause repeat expansions are currently unknown. The formation of aberrant DNA structures by repeat sequences is hypothesized to drive repeat tract instability. Any DNA metabolic process that causes the two strands of the duplex repeat tract to become separated and then re-anneal may lead to alternative DNA structure formation within the expanded repeat (Pearson, Nichol Edamura, and Cleary 2005, López Castel, Cleary, and Pearson 2010, Pearson and Sinden 1996). At sites of DNA replication, DNA slippage can occur in the template strand leading to contraction intermediates when the slipped-out region is not copied. This is more likely to occur on the lagging strand template, as long stretches of this strand remain single- stranded until Okazaki fragments are synthesized. Slip-outs can also form on the nascent strand, leading to expansion intermediates when parts of the template are copied more than once. This may be more likely during Okazaki fragment synthesis due to strand displacement and self- association. If these intermediates are not repaired, subsequent rounds of replication will produce either contractions or expansions. Instability can also occur in non-replication DNA during DNA repair. If a slip-out forms on the continuous or damaged/nicked strand, this can lead to contractions or expansions, respectively. During transcription of one strand, the opposite strand is temporarily left single-stranded, which could allow for structure formation. This could be exacerbated if two transcription complexes collide since many of the disease-associated repeats are bidirectionally transcribed, or if a transcription complex collides with a replication form. During recombination, slip-outs may form due to unequal crossing over or else during recombination-mediated repair. Repeat instability occurs at different times and levels in different tissues (both proliferative and non-proliferative), thus it is likely that a combination of these DNA metabolic processes, not just one mechanism alone, that leads to repeat instability. Each of the major DNA processes, DNA replication, transcription, repair and recombination, have been studied for their effect on repeat instability (Figure 1-1). My study will focus on the effect of DNA repair on repeat instability. The effects of DNA repair and replication on repeat instability are described in detail below.

Figure 1-2

Figure 1-2. Eukaryotic mismatch repair pathway.

Effective MMR involves structure-specific detection of the mismatched DNA or insertion/deletion loop, excision along the error-containing strand, error-free polymerization across the excision gap, followed by nick ligation. Modified from (Jiricny 2006).

1.4.1 DNA repair and repeat instability

DNA repair was suggested to be involved in gene-specific repeat instability due to the extensive levels of instability in the central nervous system (CNS) for various diseases. A contribution of DNA replication to CNS repeat instability is possible, but may be limited to early development when brain cells are being produced or to instability in non-neural (glial) cell populations. Since mismatch repair (MMR) is involved in maintaining genome stability at other microsatellites, and MMR is known to repair some random sequence insertion/deletion loops which could perhaps be recognized in a similar fashion as repeat slip-outs, much research has focused on how MMR affects TNR instability.

1.4.1.1 Mismatch repair

The function of the mismatch repair (MMR) system is to protect against genome-wide mutations and instability by repairing base-base mismatches and small insertion/deletion loops (IDLs). Effective MMR involves structure-specific detection of the mispaired DNA or insertion/deletion loop, excision along the error-containing strand, error-free polymerization across the excision gap, followed by nick ligation (Figure 1-2) (Jiricny 2006). In eukaryotes, there are two protein complexes involved in mismatch recognition: MutSα (MSH2+MSH6), the major mismatch recognition complex, and MutSβ (MSH2+MSH3) (Clark et al. 2000, Kleczkowska et al. 2001, Masih, Kunnev, and Melendy 2008). MutSα recognizes base-base mismatches and IDLs of one or two extra helical nucleotides while MutSβ recognizes longer IDLs, up to 13 extra nucleotides (Acharya et al. 1996, Genschel et al. 1998). Subsequent to mismatch recognition, MutSα or MutSβ recruits a heterodimer of MLH1 and PMS2 (MutLα). PMS2 has endonuclease activity and can generate single strand breaks near the mismatch, perhaps creating an entry site for ExoI (Kadyrov et al. 2006). The MutS/MutL complex translocates along the DNA in an ATP- dependent manner (Blackwell et al. 2001) until it encounters a strand break (a nick) where ExoI starts to degrade the error-containing strand (Genschel, Bazemore, and Modrich 2002). Once the mismatch has been removed, DNA polymerse δ fills in the gap (Longley, Pierce, and Modrich 1997). The complete MMR process has been reconstituted using purified recombinant proteins (Constantin et al. 2005, Zhang et al. 2005). In vitro studies use a nick to direct repair, stimulating a free end such as would be found at the ends of the leading or lagging strands of replication.

Figure 1-3

Figure 1-3. Slipped-DNA repair outcomes.

Long CTG/CAG slip-outs display three different repair outcomes: 1) correct repair, 2) escaped repair, and 3) error-prone repair. Correct repair involves full excision of the slip-out and polymerization across the gap to leave the correct number of repeats on each strand. Escaped repair involves sealing of the nick, leading to an expansion or a contraction mutation, depending on whether the slip-out is on the nicked strand or the continuous strand. Error-prone repair only creates expansion mutations, and involves incomplete removal of the slip-out, leading to a variety of different possible expansion lengths. Modified from (Panigrahi et al. 2005).

1.4.1.2 Repair of slipped-TNRs

To study the mechanism of TNR slip-out repair, in vitro assays have been developed in which circular slipped-DNAs are processed by cell extracts. The circular repair substrates are plasmids which have two features which lead to repair: a nick in the backbone to direct repair, and an excess of repeats on one strand versus the other which form a slipped-DNA structure. Unlike random heteroduplex repair, loop-directed repair has not been observed for TNR slip-outs, and thus repair will not occur without the presence of a nick (Panigrahi et al. 2005). The nick directs repair by acting as a strand discrimination signal for repair proteins – a nick mimics the unligated ends of a leading or lagging strand of replication, and thus distinguishes between error- containing nascent strand and the template strand. The second feature of the circular repair substrates, the slip-out, can be made to model expansion intermediates or contraction intermediates. An expansion occurs when the nascent/discontinuous strand has excess repeats when compared with the template strand, and thus an expansion intermediate repair substrate has a nick on the slip-out strand. A contraction occurs when the template strand has slipped-out and several repeats are not copied into the nascent strand, and thus a contraction intermediate repair substrate would have a nick on the strand opposite the slip-out. The following section discusses how slipped-TNRs are repaired.

Studies focusing on the processing of long (20-25) DNA slip-outs have found three distinct repair outcomes: 1) correct repair, 2) escaped repair, 3) and error-prone repair (Figure 1-3) (Panigrahi et al. 2005). Correct repair restores the parental repeat size; the process involves removal of the slip-out (or the region across from the slip-out for a contraction substrate), polymerization across the gap, and then nick ligation leading to perfectly paired DNA. Escaped repair involves sealing of the nick, with no change to the number of repeats on either strand, and thus retention of the full slip-out. In error-prone repair (which can only occur in expansion substrates), some but not all of the excess repeats are excised, leaving a variety of heteroduplexed expansion sizes. The relative levels of each type of repair may differ between tissues due to differences in protein expression. Currently the proteins required for long TNR slip-out repair are not known. Similar to long IDLs, repair outcomes for long TNR slip-outs are not dependent upon MMR and NER proteins (Hou et al. 2009); however, XPG (but not XPF) may slightly enhance repair by cleaving 5’ of the slip-out (Hou et al. 2011). WNR (but not BLM or ReqQ1) can also slightly improve repair through its helicase activity (Chan et al. 2012).

Repair of short TNR slip-outs is dependent upon MutSβ, and repair of intermediate lengths (5-10 excess repeats) is moderately improved by the presence of MutSβ (Zhang et al. 2012). The proportions of the various repair outcomes differ depending upon characteristics of the repair substrate.

Several features of slipped-DNA repair substrates can affect their repair outcome. CAG slip-outs are repaired better than CAT slip-outs, nicks located 5’ of the slip-out direct repair better than nicks 3’ to the slip-out, and slip-outs in the continuous strand are repaired better than those in the nicked strand (Figure 1-4). Additionally, the slip-out size as well as the number of slip-outs also affects repair efficiency. Short slip-outs are repaired with higher efficiency than long slip outs (1>3>20) and isolated slip-outs are repaired while multiple slip-outs (clustered slip-outs) reduce repair efficiency. The repair of short slip-outs and clustered slip-outs is dependent upon MMR proteins (mainly MutSβ) while the repair of long slip-outs is MMR-independent.

1.4.2 DNA replication and repeat instability

Initial studies on the causes of instability focused upon DNA replication due to early observations linking cell proliferation to TNR instability in DM1 (Pearson, Nichol Edamura, and Cleary 2005, López Castel, Cleary, and Pearson 2010). Replication through expanded (CGG)•(CCG) and (CTG)•(CAG) repeat tracts in bacterial cells was shown to cause replication fork pausing within the repeat in a length-dependent manner, where longer repeats induced greater stalling (Samadashwily, Raca, and Mirkin 1997). Replisome stalling was subsequently demonstrated in yeast and mammalian cells at expanded (CTG)•(CAG) templates, where repeat length, orientation of the repeat tract (CAG versus CTG repeat in the leading and lagging strands) as well as the distance of the repeat tract from the origin of replication all influenced instability (levels of instability as well as contractions versus expansions) (Cleary et al. 2002).

Primary DM1 fetal cells were shown to undergo ongoing expansions of the (CTG)216 repeat tract from the disease-causing allele but not the (CTG)12 non-expanded allele (Yang et al. 2003). Drug-induced perturbation of replication fork progression in these primary DM1 patient cells resulted in increased instability of the expanded repeat tract but not the normal repeat tract (Yang et al. 2003). These findings from multiple model systems all support a role for DNA replication in repeat instability. However, studies focused upon DNA replication alone do not address the high levels of instability observed in non-proliferative cells such as neurons of patients with

Figure 1-4

Figure 1-4. Features of slipped-DNAs affect their repair outcome.

Top to bottom shows decreasing repair efficiency. Contraction intermediates > expansion intermediates; 5’ nicks > 3’ nicks; CAG slip-outs > CTG slip-outs.

DM1. Therefore, several studies have been aimed at elucidating the roles of DNA repair and recombination in repeat expansions (Pearson, Nichol Edamura, and Cleary 2005, López Castel, Cleary, and Pearson 2010). Clearly, various mechanisms arising in proliferating and non- proliferating cells contribute to repeat instability.

1.5 Replication protein A: eukaryotic singled-stranded DNA binding protein

Replication protein A (RPA) is a eukaryotic single-stranded DNA-binding complex; it is essential for multiple processes in cellular DNA metabolism, including replication, multiple repair pathways, recombination and telomere maintenance. The canonical RPA is composed of three subunits (RPA1, RPA2, and RPA3) and has an extremely high affinity for single-stranded DNA (ssDNA) with an occluded binding site size of 30 nucleotides (Wold 1996). The DNA binding domains (DBDs) within RPA1 are the primary contacts with ssDNA in the initial multistep binding process; they binding to ssDNA with high affinity and 5’ to 3’ polarity (de Laat et al. 1998, Kolpashchikov et al. 2001, Iftode and Borowiec 2000). RPA’s affinity for double-stranded DNA (dsDNA) is very weak, with nearly a thousand-fold difference in affinity compared with ssDNA (Kim, Snyder, and Wold 1992, Blackwell, Borowiec, and Mastrangelo 1996). In fact, the dsDNA binding that is observed in vitro, is likely the result of RPA denaturation of duplex DNA followed by binding to ssDNA (Lao, Lee, and Wold 1999, Treuner, Ramsperger, and Knippers 1996). In this binding, the DBDs are believed to disrupt the hydrogen bonds between complementary DNA molecules. In addition to ssDNA binding, RPA interacts with multiple protein partners involved in DNA metabolism through interactions with the N- terminus of RPA1; this interaction is generally regulated by the phosphorylation of the N- terminal domain of RPA2.

1.5.1 RPA in DNA replication

RPA was first identified and purified as a protein factor required for both initiation and elongation phases of SV40 DNA replication (Wold and Kelly 1988, Wobbe et al. 1987). Much of our understanding of RPA in DNA replication comes from the SV40 viral replication system. In this system, the viral genome is replicated using the large T antigen helicase along with proteins supplied by the host cell. RPA is recruited to the origin of replication by T antigen and assists in the origin unwinding (Wold and Kelly 1988, Wobbe et al. 1987, Iftode and Borowiec 1997). The

17 high ssDNA affinity and the ability to denature duplex DNA are the properties of RPA that seem important in the initial replication phases. It is generally believed that RPA functions similarly in eukaryotic DNA replication. In the elongation phase of DNA replication, RPA is believed to play a role in stimulating DNA polymerase δ and ε (Waga and Stillman 1994).

1.5.2 RPA in nucleotide excision repair

RPA has been shown to be required for nucleotide excision repair (NER) (Mu et al. 1995, Aboussekhra et al. 1995), which is the main mechanism in humans for removing helix-distorting lesions induced by agents such UV light. Initially, RPA was believed to play some role in DNA damage recognition when studies demonstrated a binding preference to duplex damaged DNA compared with undamaged DNA (Burns et al. 1996, Patrick and Turchi 1998). However, studies looking at kinetics and protein assembly during NER suggest that RPA is recruited at a later stage in the pathway (Riedl, Hanaoka, and Egly 2003). The DNA binding polarity of RPA, the decreased affinity of RPA for damaged ssDNA and the ability to interact with the endonucleases XPF-ERCC1 and XPG support a role in protein positioning at the damaged DNA site (Matsunaga et al. 1996). The binding polarity of RPA and decreased affinity for damaged ssDNA would position the RPA1 subunit to the 5’ side of the undamaged DNA strand opposite the lesion and protect the undamaged DNA strand from nuclease cleavage (Evans et al. 1997). RPA1 interacts with XPG and would help position this endonuclease to the 3’ side of the lesion on the damaged DNA strand (He et al. 1995). Following cleavage by the endonuclease, RPA would be in position to stimulate the gap filling reaction performed by polymerase δ or ε (Shivji et al. 1995).

1.5.3 RPA in base excision repair

Base excision repair (BER) is the mechanism that removes small, non-helix-distorting base lesions from the genome; the pathway is initiated by DNA glycosylases, which recognize and remove specific damaged or inappropriate bases. The base lesion can either be removed by single-nucleotide BER (SN-BER) or long-patch BER (LP-BER); the two pathways differ in the size of the repair patch and the enzymes utilized to restore the lesion site. RPA has been shown to interact with DNA glycosylases in the base BER pathway, including UNG2 and hMYH (Mer et al. 2000, Nagelhus et al. 1997, Parker et al. 2001). In addition, studies in yeast containing a mutant RFA1 gene (RPA1 homolog) demonstrated sensitivity to methyl methane sulfonate,

18 which is an agent that produces DNA damage that is repaired by the BER pathway (Umezu et al. 1998). These data strongly suggest that RPA is required for BER. The role of RPA in long-patch BER where the damaged base and subsequent 2-8 bases are removed has been extensively studied (Matsumoto, Kim, and Bogenhagen 1994, Krokan et al. 2000). RPA stimulates long- patch BER by enhancing primer extension and unwinding the 5’ end of the downstream strand (DeMott, Zigman, and Bambara 1998). In addition, RPA has been shown to stimulate DNA ligase I in the final step of BER (Ranalli, DeMott, and Bambara 2002).

1.5.4 RPA in mismatch repair

The mismatch repair (MMR) system is discussed in detail above (see section 1.4.1.1 and Figure 1-3) The first evidence that RPA was required for MMR was from in vitro experiments neutralizing RPA activity with specific antibodies (Lin et al. 1998). Interestingly, mutant RPA with a point mutation in the RPA1 zinc finger domain did not support MMR or DNA replication, but fully supported NER (Lin et al. 1998). This suggests that different protein-protein interactions occur throughout the RPA structure that can influence and regulate various DNA metabolic pathways. Further biochemical studies identified specific functions of RPA in the MMR pathway. One function of RPA is to bind ssDNA and protect the template strand from nuclease degradation (Ramilo et al. 2002). RPA also functions to stimulate the excision process of EXO1 when a mismatched base is present (Genschel and Modrich 2003, 2009). Following removal of the mismatched base, RPA helps regulate and terminate EXO1 excision. This was initially believed to be dependent on MutLα (Zhang et al. 2005), but more recently, experiments with extracts devoid of MutLα have shown that RPA functions independently to terminate MutLα-activated EXO1 excision (Genschel and Modrich 2009).

1.5.5 An alternative form of RPA: aRPA

RPA is a highly conserved complex as all eukaryotes contain homologs of each of the RPA subunits (Wold 1996). The canonical RPA is composed of three subunits called RPA1, RPA2, and RPA3. In addition to the three canonical subunits of RPA, the human genome contains an additional subunit called RPA4, which was originally identified in a screen for proteins that interact with RPA1 (Keshav, Chen, and Dutta 1995). RPA4 homologs with complete coding sequences are only found in primates and horse(Haring, Humphreys, and Wold 2010). RPA4 shares 63% homology with RPA2, have similar domain organization and can substitute for

RPA2 in complex formation (Keshav, Chen, and Dutta 1995). The resulting alternative RPA (aRPA) is a stable heterotrimeric complex similar in size and stability to the canonical RPA (Keshav, Chen, and Dutta 1995), and aRPA interacts with ssDNA in a manner indistinguishable from canonical RPA (Mason et al. 2009). Protein and mRNA expression studies have shown that the RPA4 gene is expressed in normal human tissues but its expression is reduced in cancerous tissues and is very low in stable human cell lines (Keshav, Chen, and Dutta 1995, Kemp et al. 2010). Surprisingly, aRPA is unable to support in vitro SV40 DNA replication or chromosomal DNA replication in human cells and acts in a dominant-negative fashion to inhibit DNA replication in the presence of canonical RPA (Mason et al. 2009, Haring, Humphreys, and Wold 2010). There is evidence that aRPA can partially support certain types of DNA repair in the absence RPA (Kemp et al. 2010, Zhang et al. 2005, Ramilo et al. 2002). The role of aRPA and its relationship with RPA is not known, and it is not known if aRPA has functions outside of those performed by RPA.

It has been demonstrated that RPA4 expression in human cells does not allow the cell to replicate its genome nor proceed through the cell cycle. In addition, RPA4 expression seems to occur in predominantly quiescent cells and not in cell lines, which are by definition proliferative. Thus a possible model for the relationship between canonical RPA and aRPA is that in cells that need to perform genome maintenance, but not genome duplication (i.e. nonproliferating cells), aRPA might be able to substitute for canonical RPA. More research is needed to investigate whether aRPA can support basal processes normally performed by RPA to shed light on the function of aRPA and its relationship with RPA.

1.6 Thesis Goals

TNR diseases are caused by the inheritance of an unstable, gene-specific TNR tract, which can continue to expand during a person’s lifetime, leading to disease progression and greater repeat lengths in the offspring. The mechanism of repeat expansion is not clear, although it is believed to involve strand slippage during DNA synthesis mediated by the formation of an alternative hairpin structure by the repeat tract. Understanding how these hairpin structures may be aberrantly processed to expansions is vital to understanding how to prevent repeat tract expansions and how to induce contractions in the expanded repeat tract. Knowledge of the

20 alternative DNA structures assumed by the disease-associated repeat sequences may also aid in elucidating the normal biological roles of repeat sequences in the human genome.

Previous studies on slipped-DNA repair have used slipped-DNA substrates that have a nick in the flanking region proximal to the repeat tract (nick-in-flank substrates). While these models allow for studies of the effects of a hairpin structure, they are missing an important feature of TNR slip-outs: a nick that most likely forms within the repeat tract. A nick within a repeat tract can affect slip-out structure by allowing more flexibility and introduce more single-stranded DNA on both strands of the slip-out. Since altering DNA structures can affect repair outcomes, I sought to determine whether moving the nick location from the flanking region to within the repeat tract changes the structure of the slip-out and thus can have an effect on repair of TNR slip-outs.

There are many proteins involved in the repair of hairpin structures, and their roles may differ depending upon the size and conformation of the hairpin. An important group of proteins involved in repair are single-stranded DNA binding proteins, and the predominant ssDNA binding protein is replication protein A (RPA), which bind and protect ssDNA generated by excision as well as promoting excision and DNA re-synthesis (Li 2008, Oakley and Patrick 2010). Nick-in-repeat substrates have the potential for more single-stranded DNA on both strands of the slip-out, thus single-stranded binding proteins such as RPA might play a greater role in the repair of these slip-outs. I sought to determine the effects of RPA and its related complex aRPA on the repair of slip-outs, and how slip-out structure affects the function of these proteins.

Chapter 2

Effect of DNA structural hyper-polymorphism and single-stranded DNA binding proteins on repair of disease-associated slipped-DNA repeats

This work involved collaborations between the labs of Christopher Pearson, Marc Wold, and Albert La Spada. The Wold lab prepared HeLa cell extracts, RPA-deficient HeLa cell extracts, purified human RPA, aRPA as well as the antibodies against RPA and aRPA used in Western blots. The Wold Lab performed the helix destabilization assay seen in Figure 2-13B. The La Spada Lab performed transcript analysis of RPA2 and RPA4 in human tissue samples (Figure 2- 4B). From the Pearson lab, Dr. Gagan Panigrahi and Shirley Guan made slipped-intermediate DNA substrates and prepared cell extracts. Jodie Simard prepared the FEN1-deficient cell extract and performed Western blot analysis (Figure 2-12A). I prepared LoVo cell extracts, made slipped-intermediate DNA substrates, performed repair assays, replication assays, electrophoretic mobility shift assays and Western blots. I was involved in experimental planning, the interpretation of results, and the preparation of this manuscript.

2 Effect of DNA structural hyper-polymorphism and single-stranded DNA binding proteins on repair of disease-associated slipped-DNA repeats 2.1 Introduction

Over 40 neurological, neurodegenerative and neuromuscular diseases are caused by expansions of trinucleotide repeats (TNRs) (Cleary and Pearson 2003, López Castel, Cleary, and Pearson 2010, Mirkin 2007). At least 15 of these diseases are the result of (CTG)•(CAG) repeat tract expansions, including Huntington disease (HD) and myotonic dystrophy type 1 (DM1). A striking feature of (CTG)•(CAG) repeat diseases is the non-random pattern of repeat length mosaicism between different central nervous system regions. Both human patients and mouse models show that the striatum contains the longest and largest range of CAG repeats, while the cerebellum contains CAG repeats that are much smaller in size and range (Ishiguro et al. 2001, Pearson, Nichol Edamura, and Cleary 2005, Maciel et al. 1997, Kennedy et al. 2003, Takano et al. 1996). How these expansions occur, particularly in post-mitotic tissues such as neurons, is the subject of ongoing research.

One model suggests that TNR sequences can form slipped-DNA structures via strand slippage in the newly synthesized or nicked strand during DNA replication and repair, leading to TNR expansion (Mirkin 2007, Pearson, Nichol Edamura, and Cleary 2005). This model is consistent with the observation that CTG and CAG repeats form slipped-DNAs with a melting temperature higher than physiological temperature in mammalian cells (Pearson and Sinden 1996, Gacy et al. 1995, Petruska, Arnheim, and Goodman 1996). This model is further supported by the finding that slipped-DNAs are present at the expanded (CTG)•(CAG) repeat tract of the DM1 locus in patient tissues and the level of slipped-DNA detected in different tissues correlates with repeat instability (Axford et al. 2013). Therefore (CTG)•(CAG) repeat slipped-DNAs persist in vivo once they form and require a repair mechanism to remove the structure in order to prevent TNR expansion.

To understand the processing of slipped-DNAs, it is critical to appreciate their structural features. Slipped-DNAs form by out-of-register mis-pairings between complementary repeat

23 strands at sites of replication, repair, DNA damage, or recombination. In vitro slipped- intermediate DNAs (SI-DNAs) are heteroduplexes of (CTG)x•(CAG)y where x ≠ y. When x>y or x

Previous studies have shown that human cells are capable of repairing slipped-DNAs that form at (CTG)•(CAG) repeats in a nick-dependent manner. Repair outcome was highly sensitive to the structural features of the slipped-DNA, depending on the slip-out sequence (CTG or CAG), on whether the slip-out was in the nicked or continuous strand, and on whether the nick was located upstream or downstream of the slip-out (Panigrahi et al. 2005). Small slipped-DNAs (slip-out of 1-3 repeats) are repaired via the mismatch repair (MMR) mechanism while larger slipped-DNAs are repaired by a mechanism that is independent of MMR and nucleotide excision repair (NER) proteins (Littman, Fang, and Modrich 1999a, Genschel et al. 1998, Umar, Boyer, and Kunkel 1994, Hou et al. 2009, Panigrahi et al. 2005, Panigrahi et al. 2010, López Castel, Tomkinson, and Pearson 2009, Tian et al. 2009). How this MMR-independent system repairs large slipped-DNAs and the protein factors that participate are unknown. Small slipped-DNAs are repaired efficiently and accurately by MMR but the repair of larger slip-outs is less efficient and can generate error- prone repair products that lead to further repeat tract expansions (Panigrahi et al. 2005, Panigrahi et al. 2010). Therefore it is of interest to understand the mechanism of large slipped-DNA repair and the protein factors that are involved.

Previous studies investigating large slipped-DNA repair have used slipped-DNA substrates that have a nick in the flanking region proximal to the repeat tract (nick-in-flank substrates). While these models allow for studies of the effects of a hairpin structure, they are missing an important feature of TNR slip-outs: a nick that most likely forms within the repeat tract. A nick within a repeat tract can affect slip-out structure by allowing more flexibility and introduce more single- stranded DNA on both strands of the slip-out, which can affect the binding of repair proteins

(See Figure 2-1). Here I examine whether there are difference in repair efficiency and repair factor requirement between nick-in-flank substrates and nick-in-repeat substrates.

An important group of proteins involved in DNA repair are single-stranded DNA (ssDNA)- binding proteins that prevent ssDNA from premature reannealing, protect ssDNA from nuclease digestion, and remove secondary structures from ssDNA to allow other enzymes to bind. The major ssDNA binding protein in human cells is replication protein A (RPA), which bind and protect ssDNA generated by excision as well as promoting excision and DNA re-synthesis (Li 2008, Oakley and Patrick 2010). In addition to DNA repair, RPA has been shown to be an essential protein in many other processes that allow the cell to proliferate and maintain genomic integrity, including DNA replication, recombination and telomere maintenance (Oakley and Patrick 2010, Wold 1996). The human RPA complex is composed of three subunits: RPA1 (70kDa), RPA2 (32kDa), and RPA3 (14kDa). RPA is a highly conserved complex as all eukaryotes contain homologs of each of the RPA subunits (Wold 1996). In addition to the three canonical subunits of RPA, the human genome contains an additional subunit called RPA4 (30kDa). Unlike other RPA subunits, which have homologs in all eukaryotes, RPA4 homologs with complete coding sequences are only found in primates and horse (Haring, Humphreys, and Wold 2010). RPA4 shares 63% homology with RPA2, have similar domain organization and can substitute for RPA2 in complex formation (Keshav, Chen, and Dutta 1995, Mason et al. 2009). The resulting alternative RPA (aRPA) is a stable heterotrimeric complex similar in size, stability and biochemical properties to the canonical RPA (Mason et al. 2009) (See Figure 2-4A). Binding assays show that aRPA binds to single-stranded DNA in a manner that is indistinguishable from RPA, with high affinity and low cooperativity (Mason et al. 2009).

Previous studies using mouse and human tissues have shown that repeat instability may be related to levels of protein expression in different tissues (Mason et al. 2014, Tomé, Manley, et al. 2013, Tomé, Simard, et al. 2013, Goula et al. 2009, Goula et al. 2012). Both HD transgenic mice and human HD patients show higher expression levels of certain DNA repair genes, including RPA, in the cerebellum (least repeat instability) than the striatum (highest repeat instability) (Mason et al. 2014, Goula et al. 2009). Here we compare the expression of RPA2 and RPA4 in the cerebellum and striatum between HD patients and human controls to assess whether different ratios of RPA:aRPA can contribute to the repeat instability observed in different tissues of the CNS.

Protein and mRNA expression studies have shown that the RPA4 gene is expressed in normal human tissues but its expression is reduced in cancerous tissues and is very low in stable human cell lines (Keshav, Chen, and Dutta 1995, Kemp et al. 2010). Surprisingly, aRPA is unable to support DNA replication in human cells and acts in a dominant-negative fashion to inhibit DNA replication in the presence of canonical RPA (Haring, Humphreys, and Wold 2010, Mason et al. 2009). There is evidence that aRPA can partially support certain types of DNA repair in the absence of RPA (Ramilo et al. 2002, Kemp et al. 2010, Zhang et al. 2005). The role of aRPA and its relationship with RPA remains conjectural and it is not known if aRPA has functions outside of those performed by RPA. Here I assess differences between RPA and aRPA in helix destabilization, slipped-DNA binding and their roles in the repair of slipped-DNAs. Notably, we present evidence that the relative expression levels of RPA and aRPA may contribute to somatic instability in (CTG)•(CAG) repeat expansion diseases.

2.2 Materials and Methods

2.2.1 Slipped-intermediate DNA (SI-DNA) substrate preparation

Heteroduplex SI-DNA substrates were prepared using 1.5 kb circular plasmids containing pure (CTG)n•(CAG)n repeats (n=30, 47, 48, or 50) cloned from the human flanking DM1 sequences as previously described (Panigrahi et al. 2005, Pearson et al. 2002). SI-DNA Substrates were prepared by hybridizing linearized double-stranded DNAs to complementary circular single- stranded DNAs containing different numbers of repeats thus forcing a slip-out in the excess repeats. Heteroduplexing was performed by using an optimal molar ratio of single-stranded to double-stranded DNA and denatured in a solution of 0.5 M NaCl (yielding pH 13) and 500 mM NaOH at room temperature for 5 minutes. Samples were neutralized with 50-fold volume of 50 mM Tris-HCl (pH 8), 5 mM EDTA, and 0.2755 M NaCl, resulting in a solution having 0.01 M NaOH and 0.03 M NaCl. These conditions promote full renaturation. Samples were immediately placed at 68°C for 3 hours for renaturation, followed by purification by ethanol precipitation. The denaturation/renaturation products were then electrophoretically resolved on 1% agarose gel and gel-purified (heteroduplexes with slip-outs migrate slower than homoduplexes and hence can be separated). Each substrate contained a nick whose location (site and strand) was determined by the choice of restriction enzyme used to linearize the double-stranded plasmid DNA and choice of single-stranded circular DNA (CTG or CAG). Nick-in-flank substrates (1, 2 and 3)

26 used double-stranded plasmids that were linearized with EcoRI, which created a nick 56 nucleotides upstream of the (CTG/CAG) repeat tract. Nick-in-repeat substrates (4, 5 and 6) used double-stranded plasmids that were linearized with BsmI, which created a nick within the first repeat of the CTG/CAG repeat tract.

2.2.2 Human cell extract preparation

Cell extracts were prepared from the repair-proficient HeLa S3 cells and repair-deficient LoVo cells (hMSH2-null, undetectable levels of hMSH3 and hMSH6 proteins). HeLa S3 cell line was purchased from National Cell Culture Center, National Center for Research Resources, National Institute of Health; LoVo cell line was purchased from the American Type Culture Collection. Whole cell extract was prepared as described previously (Roberts and Kunkel 1988, Thomas, Roberts, and Kunkel 1991). Briefly, cells were grown at 37°C in Eagle Minimum Essential Medium supplemented with 10% fetal bovine serum. Cells were harvested in mid-log phase at a density of 5 x 105 cells per ml by centrifugation at 1,000 x g for 5 min. The cell pellet was rapidly washed in 500 ml of ice-cold hypotonic buffer (20 mM HEPES [N-2- hydroxyethylpiperazine-N'-2 ethanesulfonic acid; pH 7.5], 5 mM KCl, 1.5 mM MgCl2, 1 mM dithiothreitol [DTT]). The washed cell pellet was suspended in hypotonic buffer at a density of 6 x 107 to 7 x 107cells per ml and allowed to swell on ice for 15 min. The cells were then lysed with three to five strokes of a tightly-fitting pestle in a Dounce Homogenizer. After a 30 min to 60 min incubation on ice, the lysate was centrifuged at 10,000 x g for 10 min at 0°C. The clarified lysate was frozen in liquid nitrogen drop-wise and the frozen extract beads were stored at -80°C.

RPA-deficient HeLa cell extracts (RPA-/-) were gifts of Dr. Marc Wold (University of Iowa; Iowa City, Iowa) and they were prepared by siRNA knockdown as described (Haring, Humphreys, and Wold 2010, Li and Kelly 1984). Only cell extract preparations that were functional in SV40 in vitro replication were used. The Fen-Rex cell line used to make FEN1- deficient cell extract was a gift of Dr. Torgeir Holen (University of Oslo; Oslo, Norway). The cells were cultured as previously described (Moe, Sorbo, and Holen 2008). Briefly, the Fen-Rex cell line was stably transfected with a pENTR/H1/TO-Fen1-sh plasmid and maintained in Dulbecco’s Modified Eagle Medium, 10% fetal bovine serum and 1% glutamine. Down- regulation of FEN1 by RNAi was induced in the presence of 1 µg/mL tetracycline (FEN1-/-);

27 control cell line was grown in the absence of tetracycline (FEN1+/+). Whole cell extracts were prepared as described above.

2.2.3 SV40 DNA replication

The SV40 in vitro replication assay was performed as previously described (Roberts and Kunkel 1988, Li and Kelly 1984). DNA templates (150 ng) were replicated in reactions containing the following final concentrations: 100 µM each dATP, dGTP, dTTP, and dCTP; 200 µM each GTP, UTP, and CTP; 4 mM ATP; 40 mM creatine phosphate (Roche); 100 µg/ml creatine kinase (Roche); 1 µg SV40 T-antigen (Chimerx); 70 µg cell extracts and 300 ng of the indicated protein. For direct analysis of the replication products, 0.033 µM 32P-α-dCTP (3000 Ci/mmol, PerkinElmer Life Sciences) was included in each 50 µl reaction. Reactions were incubated for 4 hours at 37°C and terminated with 50 µl of stop solution (2 µg/µl proteinase K, 2% SDS, and 50 mM EDTA, pH 8) with further incubation for 30 min at 37°C. Carrier tRNA (15 µg) was added, and protein was extracted twice with phenol/chloroform and chloroform. Replication products were precipitated with ethanol and resuspended in water. Replication efficiency was determined by linearizing radioactive replication products with BamHI, and treated with DpnI, which digests fully methylated DNA. Equal quantities of reaction products digested with BamHI only or BamHI + DpnI were resolved by electrophoresis on a 15 cm 1% agarose gel. The gel was run for 16 hours at 4 V/cm in TBE buffer, dried, and exposed to Kodak film.

2.2.4 In vitro DNA repair

Slipped-strand repair reactions (50 µl) were conducted as described (Panigrahi et al. 2005). Briefly, 20 ng of SI-DNA substrate was incubated in 15 mM sodium phosphate, 40 mM creatine phosphate (Roche), 100 µg/ml creatine kinase (Roche), 4 mM ATP (Roche), 200 µM each CTP, GTP and UTP (Roche), 16.5 nM each [α-32P]dATP, [α-32P]dGTP, [α-32P]dCTP and [α-32P]dTTP (3000Ci/mmol, PerkinElmer Life Sciences), and 25 µg cell extract at 37°C for 1 hour. To assess repair efficiency, nonradioactive dNTPs were used (33 µM). The repair reaction was terminated with 50 µl of stop solution (2µg/µl proteinase K, 2% SDS, and 50mM EDTA, pH 8) with further incubation for 1 hour at 37°C. Proteins were extracted twice with phenol/chloroform and chloroform. Repair products were further purified using MinElute spin columns (Qiagen) and eluted with water. The repeat-containing fragment was liberated by digestion with EcoRI and HindIII and analyzed on a 4% native polyacrylamide gel and 6% denaturing sequencing gel.

Repair efficiencies were determined on a molar level by Southern blot probing of reaction products that had been generated in the presence of nonradioactive dNTPs (3 3µM). After release of the repeat-containing fragment (EcoRI/HindIII) and resolution on 4% native polyacrylamide gel, products were electrotransferred to nylon membrane and hybridized to a radiolabeled EcoRI/HindIII fragment containing 17 repeats. Membranes were then exposed Fujifilm imaging plates and digitally scanned using Typhoon FLA-9500 (GE Healthcare Life Sciences). Repair efficiency was determined using ImageQuant 5.1 (Molecular Dynamic); it was calculated as the proportion of radiointensity of the repeat-containing product relative to all repeat-containing fragments, as done for base-base MMR and heteroduplex repair (Thomas, Roberts, and Kunkel 1991, Littman, Fang, and Modrich 1999b).

2.2.5 Western blotting

Human cell extracts (50 µg of protein) were separated on a Mini-PROTEAN® TGX Stain- FreeTM 4-20% gradient gel (Bio-Rad) and transferred to PVDF membrane and probed overnight at 4°C. Membranes were probed with the 71-9A antibody (recognizes RPA2 subunit) to detect RPA and Anti-RPA4 antibody to detect aRPA. Immunoblot for RPA detection was incubated in HRP-conjugated sheep anti-mouse secondary antibody (GE Healthcare, 1:5000) and immunoblot for aRPA detection was incubated in HRP-conjugated donkey anti-sheep secondary antibody (Sigma, 1:5000). Chemilumiescent signals were generated using Amersham ECL Plus Western Blotting Detection Reagent (GE Healthcare). Images were captured on BioRad ChemiDocTM MP. The homogeneity of loading was verified by activating and visualizing the stain-free gel using BioRad ChemiDocTM MP. Primary antibodies were gifts of Dr. Marc Wold (University of Iowa, Iowa City, Iowa).

Western blotting for FEN1 was done by Jodie Simard in the Pearson lab. Human cell extracts (40 µg of protein) were separated on a 10% SDS-PAGE followed by immunoblotting and probed with FEN1 antibody (clone B4, Santa Cruz; 1:40,000) and actin antibody (AB-5, BD biosciences; 1:5000). Chemilumiescent signals were generated using Amersham ECL Plus Western Blotting Detection Reagent (GE Healthcare) then exposed to Kodak film.

2.2.6 DNA binding assays

Indicated amounts of RPA and aRPA were incubated with 20 ng of DNA substrates in reaction buffer containing 30 mM HEPES, 100 mM NaCl, 5 mM MgCl2, 0.5% inositol, and 1 mM dithiothreitol in a final volume of 20 µl. Reaction mixtures were incubated at 25°C for 30 min followed by restriction digestion by EcoRI and HindIII to release the repeat-containing fragment. Samples were then separated on a 4% native polyacrylamide gel in Tris-borate EDTA running buffer. Products were electrotransferred to membrane and hybridized to a radiolabeled EcoRI/HindIII fragment containing 17 repeats. Membranes were subsequently exposed to Kodak film.

2.3 Results

2.3.1 Slipped-DNAs with increased single-stranded potential

Repair of slipped-DNAs, like base-base mismatches, is nick-directed; the nick defines the strand that will be repaired using the continuous strand as the template for repair. Nick-in-flank Slipped-Intermediate DNA (SI-DNA) substrates have been well-characterized, both structurally (Pearson et al. 2002, Pearson and Sinden 1996, Tam et al. 2003, Pearson, Eichler, et al. 1998, Pearson, Wang, et al. 1998, Pearson et al. 1997) and for repair outcome (Panigrahi et al. 2005, Panigrahi et al. 2010, López Castel, Tomkinson, and Pearson 2009, Panigrahi et al. 2012). These substrates have the nick approximately 60 base pairs away from the repeat tract (EcoRI site, Figure 2-1A, B). The nick location ensure complementary annealing of the flanking sequence on either side of the repeat tract, forcing a slip-out to form within the center of the repeat tract on the strand that has an excess number of repeats (Pearson et al. 2002). This essentially anchors the slip-out in place. This anchoring does not occur when the nick is located in the repeat tract, such as when DNA slippage occurs during replication or repair. When a nick is in the repeat tract, the free ends have increased structural heterogeneity due to the absence of the anchoring non- repetitive flanking sequence (Figure 2-1C). The structure of a heteroduplex is known to dramatically affect its repair outcome; therefore, I assessed slipped-DNA repair of both nick-in- repeat and nick-in-flank substrates. Nick-in-repeat substrates were prepared by taking advantage of the unique BsmI recognition site that is naturally just upstream of the CTG/CAG tract in the cloned DM1 region, leading to cleavage within the first CTG unit of the CTG tract, and the last CAG unit of the CAG tract (Figure 2-1B, C). Nick-in-repeat substrates are molecularly identical

30 to nick-in-flank substrates, but differ only in the nick location and potentially in secondary structure. Nick-in-flank substrates have limited structural freedom (Pearson and Sinden 1996, Pearson et al. 2002) while the nick-in-repeat substrates are expected to have high structural variability, with increased potential for single-stranded DNA formation on both strands (Figure 2-1C). Such structural heterogeneity might have dramatic effects upon repair outcome, as well as the requirement for specific repair factors such as the ssDNA binding proteins RPA and aRPA.

The high structural variability of nick-in-repeat substrates was evident following substrate preparation by denaturation and renaturation of a DNA duplex linearized with BsmI and a single- stranded circular plasmid, both containing either 30 or 50 repeats. The starting materials for substrate preparation and the end products were run on agarose gels to assess degree of heteroduplex formation. Whereas the preparation of nick-in-flank substrates, with the DNA free ends in the flanking region, yielded only one heteroduplex product (Figure 2-2A), the preparation of nick-in-repeat substrates, with the DNA free ends located within the repeat region, yielded multiple heteroduplex products (Figure 2-2B and C). All the identified heteroduplex bands were purified from agarose gel and run on native polyacrylamide gels along with their repair products after processing by HeLa cell extract. Repair reactions for the 3 sub-species of substrate 5 showed that sub-species 1 and 3 underwent error-prone repair, as is expected for expansion substrates, while sub-species 2 showed no discernible starting heteroduplex substrate (Figure 2-2D). Sub-species 1 was the most concentrated following purification and thus 5-1 was used in all subsequent experiments. Repair reactions for the 7 sub-species of substrate 6 showed that sub-species 1, 5 and 7 underwent error-prone repair, while the rest of the sub-species showed no discernible starting heteroduplex or were too dilute (Figure 2-2E). All sub-species showed a contaminating species at the 30 duplex position. Sub-species 5 was the most concentrated following purification and thus 6-5 was used in all subsequent experiments. Agarose gels of heteroduplex formation products show that nick-in-repeat substrates have much higher structural variability than nick-in-flank substrates due to the lack of anchoring by flanking sequences. This increased structural variability might dramatic effects upon repair outcome, as well as the requirement for specific repair factors.

Figure 2-1

Figure 2-1. Nick-in-repeat slipped-DNAs have increased single-stranded potential.

(A) Schematic representation of circular hybrids with slipped CAG or CTG repeats and a unique nick at EcoRI (E) or BsmI (B). The CAG and CTG repeat tracts are colored blue and red, respectively. As indicated, the nicks are located 5’ and 3’ of the slip-out. Substrate numbers identify the repeat lengths, sequences, strand polarity and nick location (arrow). For example, substrate 1 is composed of a (CTG)30 strand, nicked 5’ of the slip-out at the EcoRI site, hybridized to a continuous (CAG)50 strand. The top panel shows nick-in-flank substrates, where the nick created by EcoRI is located 56bp away from the repeat tract. The bottom panel shows nick-in-repeat substrates, where the nicked created by BsmI is located within the first repeat of the repeat tract. (B) Schematic representation of the EcoRI/HindIII fragment harboring the CAG/CTG repeats, the BsmI cleavage site, human DM1 flanking sequence (thick black lines) and vector sequences (thin black lines). (C) Nick-in-flank substrates have limited structural freedom due to anchoring by the flanking sequences. Nick-in-repeat substrates have a nick within the first repeat of the CTG/CAG repeat tract, allowing the repeat tract on the nicked strand to slip and slide along the continuous strand. Thus nick-in-repeat substrates have high structural variability and increased single-stranded DNA formation in both strands.

Figure 2-2

Figure 2-2. Nick-in-repeat slip-outs have greater structural heterogeneity than nick-in-flank slip- outs.

Heteroduplex substrates were prepared by hybridizing single-stranded circular plasmid with linearized duplex DNA. (A) Duplex linearized by EcoRI gave rise to nick-in-flank SI-DNA 2 (arrowhead) Image adapted from (Panigrahi et al. 2005). (B, C) Duplex linearized by BsmI gave rise to multiple species of nick-in-repeat SI-DNA 5 and SI-DNA 6 (denoted by arrowheads). (D) Southern blot analysis of in vitro repair for the 3 sub-species of SI-DNA 5. Lanes 1, 3 and 5 show SI-DNA 5 sub-species without processing by cell extract. Lanes 2, 4 and 6 show SI-DNA sub-species processed by 25 µg of HeLa cell extract. The SI-DNA substrates and repaired products are indicated by arrows. (E) Southern blot analysis of in vitro repair for the 7 sub- species of SI-DNA 6. Lanes 1, 4, 7, 10, 13, 16 and 18 show SI-DNA 6 sub-species without processing by cell extract. Lanes 2, 5, 8, 11, 14 and 19 show substrates processed by 25 µg HeLa cell extract. Lanes 3, 6, 9, 12, 15, 17 and 20 show substrates processed by 120 µg of HeLa cell extract. The SI-DNA substrates and repaired products are indicated by arrows.

Towards assessing the structure and repair efficiency of the nick-in-repeat slip-outs versus that of nick-in-flank slip-outs, we resolved unprocessed substrates (Figure 2-3, schematics shown on top) and substrates processed by HeLa cell extracts on high-resolution native polyacrylamide gels, which can sensitively detect structural variations in slipped-DNAs (Panigrahi et al. 2005, Panigrahi et al. 2010, Pearson et al. 2002, Tam et al. 2003). Interestingly, even though the nick- in-repeat substrates (4 and 5) were molecularly the same as the nick-in-flank substrates (1 and 2), they migrated to different points on the Southern blot native gel. The nick-in-repeat substrates migrated slightly faster than the nick-in-flank slipped-DNAs – suggesting that they had different secondary structures (Figure 2-3, lanes 1, 4, 7 and 10, black arrowheads). One interpretation of this aberrant electrophoretic migration is increased flexibility, as has been observed for DNAs containing nicks or extended gaps, molecules, including DNA-junctions (Mills, Cooper, and Hagerman 1994, Guo and Tullius 2003, Pöhler, Duckett, and Lilley 1994). The location of the nick within the repeat, which has been documented to be hyper-flexible (Chastain et al. 1995, Chastain and Sinden 1998) might lead to the faster electrophoretic migration relative to the same DNAs with nicks in their flanks. We previously demonstrated that substrates with CTG slip-outs migrate electrophoretically faster than substrates with CAG slip-outs because the excess CAG repeats extruded as a single-stranded random coil, whereas the excess CTG repeats extruded as an intrastrand hairpin (Panigrahi et al. 2005, Panigrahi et al. 2010). This pattern was observed again both for nick-in-flank substrates (Figure 2-3, compare lanes 1 and 4). Substrates 4 and 5 both have CAG20 slip-outs but substrate 5 migrates slower than substrate 4 (Figure 2-3, compare lanes 7 and 10), which suggested the nick on the slip-out strand caused a bulkier slip-out.

In vitro processing by HeLa extract yielded a set of DNA products with varying electrophoretic mobility, some of which was the expected repair product, as well as some of the starting material (Figure 2-3). The expected repair products are nick-directed where the nicked strand is repaired using the continuous strand as template for repair. All substrates showed an increase in the expected correct repair product with increased amounts of HeLa extract (Figure 2-3). The nick- in-flank substrates SI-DNA 1 and 2 had considerable amounts of unrepaired substrate following HeLa extract processing (Figure 2-3, lanes 2, 4, 5 and 6). In contrast, the nick-in-repeat substrates were efficiently repaired, leaving no unrepaired substrate (Figure 2-3, lanes 8, 9, 10 and 11). These results indicate that the nick-in-repeat substrates were more efficiently repaired correctly than nick-in-flank substrates. For substrate 1, 2 and 4, processing by HeLa cell extract

Figure 2-3

Figure 2-3. Repair of slipped-DNAs depends on cell extract concentration and nick location.

Southern blot analysis of in vitro repair for a nick-in-flank 20-repeat SI-DNA (1, 2) and two nick-in-repeat 20-repeat SI-DNAs (4, 5). Repair reactions were carried out as described in Materials and Methods. Substrates were digested with EcoRI/HindIII, resolved on 4% polyacrylamide gel, electrotransferred onto nylon membrane and probed with a radiolabeled EcoRI/HindIII fragment containing 17 repeats. Lanes 1, 4, 7 and 10 show SI-DNA substrates without processing by cell extract. Lanes 2, 5, 8 and 11 show processing by 25 µg of HeLa cell extract. Lanes 3, 6, 9 and 12 show SI-DNA substrates processed by 120 µg of HeLa cell extract. The starting SI-DNA substrates are indicated by black arrowheads. The slow-migrating species that migrate above the starting substrate after processing by cell extracts are indicated by white arrowheads. Repaired products of (CTG)50•(CAG)50 and (CTG)30•(CAG)30 are indicated by arrows. Reaction products of substrate 5 are shown with a schematic of the interpreted repair products, with newly synthesized regions indicated by a tract of dots. Autoradiograph is from a single gel, identical exposure time, with intervening lanes excised for presentation.

36 gave rise to the correct repair product of 50 repeat duplex, while the correct repair product of substrate 5 was a 30 repeat duplex.

Curiously, following exposure to the cell extract the nick-in-repeat substrate reactions yielded a product or series of products that co-migrated with the starting nick-in-flank substrates (Figure 2- 3, white arrowheads). This phenomenon has never been observed for nick-in-flank substrates. These bands indicate that processing by HeLa cell extract either gave rise to repeat expansions for a small number of substrates, or induced structural conformations that migrate slower than the starting substrates. We assessed these slower migrating bands by resolving repair by radio- incorporation products on denaturing polyacrylamide gels, which denatures DNA and resolves DNA based on size alone (See Section 2.3.4 below).

For substrate 5, processing by HeLa cell extract also gave rise to a series of ladder bands (Figure 2-3, substrate 5). We believe this represents error-prone repair, previously observed as incomplete excision of the slip-out followed by gap-filling and ligation (Panigrahi et al. 2005). This process leads to DNA expansions, where some of the excess repeats in the starting SI-DNA are retained (Panigrahi et al. 2005). The retention of between 1-, 2-, 3-…. up to 20 repeats gives rise to the ladder of bands observed for substrate 4 (Figure 2-3, see schematic to the right of substrate 5). In a previous study of nick-in-flank substrates, we demonstrated that error-prone repair occurs predominantly for substrates with a nick on the slip-out strand (expansion substrates) and only to a limited degree for substrates with a nick on the strand opposite the slip- out (contraction substrates), which could explain the expansion bias of repeat instability observed in affected families (Panigrahi et al. 2005). Here we show that error-prone repair is also observed in expansion substrates for the nick-in-repeat substrates, but not in contraction substrates.

Figure 2-4

Figure 2-4. RPA4 expression is elevated above RPA2 in HD patient brains.

(A) Schematic representation of the protein subunits that make up RPA and aRPA complexes. (B) Real-time RT-PCR analysis of RPA2 and RPA4 expression levels in the striatum and cerebellum of unaffected human adults (n = 3; mean ± s.e.m., three independent experiments; **p = 8.6×10-7 – 6.6×10-6 for RPA2 and p = 2.8×10-7 – 9.5×10-5 for RPA4; t-test). Β-actin served as the normalization control. RT-PCR analysis was done by La Spada lab (UC San Diego).

2.3.2 RPA4 expression is elevated above RPA2 in HD patient brains

Previous studies of mouse and human tissues suggested that repeat instability may in some cases be related to levels of repair protein expression (Mason et al. 2014, Tomé, Manley, et al. 2013, Tomé, Simard, et al. 2013, Goula et al. 2009, Goula et al. 2012). Both HD transgenic mice and human HD patients show higher levels of factors, including RPA, in the cerebellum than the striatum, tissues that show the lowest and highest levels of somatic CAG repeat expansions (Goula et al. 2009, Mason et al. 2014). Human cells contain two RPA complexes: a canonical RPA complex that is composed of RPA1, RPA3, and RPA2 and an alternative RPA complex (aRPA) that is composed of RPA1, RPA3, and a homolog of RPA2, RPA4 (Figure 2-4A). RPA and aRPA have similar biochemical and DNA-binding properties; however, aRPA does not function in DNA replication but does act in certain repair processes (Mason et al. 2009, Haring, Humphreys, and Wold 2010). Our collaborators from the La Spada Lab (UC San Diego) analyzed gene expression levels for RPA2 and RPA4 in HD patient and control human brain regions by qPCR. Strikingly high levels of RPA2 and RPA4 were present in HD patient brains, elevated about 6- to 8-fold, respectively, compared to control brains (Figure 3B, p = 8.6×10-7 – 6.6×10-6 for RPA2 and p = 2.8×10-7 – 9.5×10-5 for RPA4, two-tailed T-test). RPA4 was more elevated than RPA2 in HD patient brains, and RPA4 levels are higher in the striatum (highest repeat instability) than cerebellum (lowest repeat instability) (Figure 2-4B). The difference in RPA4 expression between HD patient tissues and control tissues suggests that RPA4 may be involved in CAG repeat instability.

RPA and aRPA bind to ssDNA with strong affinity (Mason et al. 2009), and may be involved in processing slipped-DNAs. Towards determining the roles of RPA and aRPA in the in vitro processing of CTG/CAG slip-outs, our collaborators from the Wold lab (U of Iowa) prepared HeLa extracts depleted of RPA2 (Haring, Humphreys, and Wold 2010). My Western blot analysis confirmed RPA knockdown in extracts of anti-RPA2 siRNA treated HeLa cells but not in mock-treated HeLa cells (Figure 2-5A). The aRPA complex could not be detected in either cell extract (Figure 2-5A), which is in agreement with previous findings that RPA4 is expressed at very low levels in established cell lines (Keshav, Chen, and Dutta 1995, Kemp et al. 2010).

Figure 2-5

Figure 2-5. SV40 DNA replication requires RPA, but not aRPA.

(A) Western blots of Hela cell extract and RPA-depleted HeLa cell extract (RPA2 siRNA). Antibody that recognizes RPA2 (32 kDa) was used to detect RPA; antibody that recognizes RPA4 (30 kDa) was used to detect aRPA. (B) The pKN16 template, which contains the SV40 origin of replication but no repeat tract, was replicated in vitro by the indicated cell extracts (70 µg of protein per reaction) and 300 ng of RPA or aRPA where indicated. Purified replication products were linearized with BamHI, and an equal amount of this material was digested with both BamHI and DpnI. The digestion products were electrophoresed on a 1% agarose gel to resolve the completely replicated and the incompletely replicated material. An audioradiograph of the dried gel is shown. The DpnI-resistant material shown in lanes 2, 4 and 8 represents the products of at least one complete round of replication. A plus sign in the table indicates the addition of the indicated component.

The functionality of HeLa and RPA-deficient cell extracts and purified RPA and aRPA proteins was assessed using the SV40 in vitro replication assay. A replication template containing SV40 origin of replication was replicated in vitro with HeLa cell extract, RPA-deficient extract, RPA- deficient extract complemented with purified RPA and RPA-deficient extract complemented with purified aRPA. Reaction products were linearized (BamHI) and then digested with DpnI, which digests fully-methylated DNA. DNA resistant to DpnI digestion represents products of at least one complete round of replication. As shown in Figure 2-5B, HeLa cell extract facilitates in vitro replication and RPA-deficient HeLa extract does not. To learn if replication is dependent upon RPA or aRPA, replication assays were performed using RPA-deficient cell extract supplemented with exogenous recombinant human RPA or aRPA proteins. Addition of RPA restored replication to levels better than those seen with HeLa cell extract while aRPA was not able to restore replication at all (Figure 2-5B). These results were in agreement with previously published studies (Mason et al. 2009, Haring, Humphreys, and Wold 2010) and confirmed the functionality of cell extracts and purified proteins.

2.3.3 RPA and aRPA can enhance slipped-DNA repair

To determine the roles of RPA and aRPA in the repair of slip-outs, we performed in vitro repair assays using various nick-in-flank and nick-in-repeat slipped-DNA substrates. RPA was not required for the correct repair of either nick-in-flank or nick-in-repeat substrates (Figure 2-6, compare lanes 1 and 2 in each panel). For nick-in-repeat substrates, processing by HeLa and RPA-deficient extracts also gave rise to products that electrophoretically migrated slower than their starting SI-DNAs, but coincident with the starting nick-in-flank SI-DNA1 (Figure 2-6B and C, lanes 2 and 3). For substrate 5, processing by either cell extract gave rise to a series of ladder bands indicative of error-prone repair. Taken together, this suggested that RPA is not essential for the repair of large slip-outs, regardless of the slip-out structure.

Although RPA was not required to repair slipped-DNAs, adding RPA or aRPA to RPA-deficient cell extract enhanced correct repair of substrates with 20-repeat slip-outs. Repair enhancement is particularly pronounced for SI-DNA1; addition of RPA or aRPA more than doubled the repair efficiency of non-depleted HeLa extract (Figure 2-6A). For SI-DNA4 and SI-DNA5, addition of RPA or aRPA removed the slow-migrating species above the starting SI-DNA species and gave rise to almost complete repair of the SI-DNA (Figure 2-6B and C). Interestingly, addition of

Figure 2-6

Figure 2-6. RPA is not required for repair, but RPA and aRPA can both enhance repair.

Southern blot analysis of in vitro repair for a nick-in-flank 20-repeat SI-DNA (1) and two nick- in-repeat 20-repeat SI-DNAs (4, 5). Repair reactions were carried out as described in Materials and Methods. Substrates were digested with EcoRI/HindIII, resolved on 4% polyacrylamide gel, electrotransferred onto nylon membrane and probed with a radiolabeled EcoRI/HindIII fragment containing 17 repeats. Lanes 1 of each panel shows SI-DNA substrates without processing by cell extract. The subsequent lanes show each substrate processed by 25 μg of HeLa cell extract (lane 2), 25 μg RPA-deficient extract HeLa cell extract alone (RPA2 siRNA, lane 3), or supplemented with 600 ng RPA (lane 4), or aRPA (lane 5). The starting SI-DNA substrates are indicated by black arrowheads. The slow-migrating species that migrate above the starting

42 substrate after processing by cell extracts are indicated by white arrowheads. Repaired products of (CTG)50•(CAG)50 and (CTG)30•(CAG)30 are indicated by arrows. Repair efficiencies were calculated for three to six replicates and adjusted for starting background. Graph shows repaired material (gray bars) and processed material that migrates above the starting SI-DNA (white bars). (A) Repair of SI-DNA 1, with a (CAG)20 slip-out and a 5’ nick in the flanking region on the strand opposite the slip-out. (B) Repair of SI-DNA 4, with a (CAG)20 slip-out and a 5’ nick in the repeat tract on the strand opposite the slip-out. (C) Repair of SI-DNA 5, with a (CAG)20 slip- out and a 5’ nick in the repeat tract on the slip-out strand.

Figure 2-7

Figure 2-7. Repair of G-T mismatches and small slip-outs do not require RPA.

(A) Schematic of a G-T mismatch repair assay assessed by Southern blot. Following the repair reaction, substrates are linearized with XmnI and assessed for nick-directed repair by HindIII digestion. Products are then run on a 0.7% agarose gel to separate the repaired and unrepaired products, transferred to nitrocellulose membrane and probed with a radiolabeled plasmid. (B) Southern blot analysis of starting unprocessed G-T mismatch substrate, and their processed products, after digestion (XmnI/HindIII), resolution on 0.7% agarose gel, electrotransfer and probing. Lane 1 shows starting unprocessed G-T mismatch substrate. Lane 2 shows substrate

44 processed by 25 μg of HeLa cell extract. Lane 3 shows substrate processed by 25 μg of RPA- deficient HeLa cell extract (RPA2 siRNA). Lane 4 shows RPA-/- extract supplemented with 600 ng RPA. Lane 5 shows RPA-/- extract supplemented with 600 ng aRPA. (C) Repair of SI-DNA

3, with a (CTG)1 slip-out and a 5’ nick in the flanking region on the slip-out strand. Repair reactions were carried out as described in Materials and Methods. Repair efficiencies were calculated for three replicates and adjusted for starting background.

RPA or aRPA not only removed the slow-migrating ladder bands above the starting SI-DNA5, but the lower ladder bands were removed as well (Figure2-6C). These results suggest that RPA is not required for the repair of slip-outs, but supplementing RPA or aRPA can enhance repair for larger slip-outs (20-repeat slip-outs) and remove the slow-migrating species for nick-in-repeat slip-outs.

Repair of DNA base-base mismatches short single-repeat slip-outs requires the mismatch repair proteins MutS. Similarly, repair of base-base mismatches requires the MMR protein complex MutS (Modrich 2006). Experiments using purified proteins in a reconstituted repair system suggested that G-T mismatch repair requires the single-stranded DNA binding abilities of RPA (Ramilo et al. 2002, Zhang et al. 2005). My in vitro repair assays using the RPA-depleted HeLa cell extract showed that G-T mismatches and single repeat slip- outs (SI-DNA2) underwent repair in the absence of RPA (Figure 2-7B and C). Repair efficiencies for the two substrates were only mildly enhanced by the addition of RPA or aRPA (Figure 2-7B and C). These results suggest that RPA is not absolutely required for either base- base or short slip-out repair.

2.3.4 Repair is nick-directed

Since the nick-in-repeat substrates appear to have different secondary structures than the nick-in- flank substrates and the structure appears to change after exposure to cell extract (Figure 2-3, Figure 2-6, and Figure 2-8A, white arrowheads), we questioned whether these were repair products or structural changes. The slow-migrating bands observed for nick-in-repeat substrates indicate that processing by cell extract either gave rise to repeat expansions for a small number of substrates, or induced structural conformations that migrate slower than the starting substrates. These slow-migrating bands were removed by the addition of RPA and aRPA. This suggests that the bands are likely structural conformations and not repeat expansions. To test this possibility, repair reactions were performed as above with non-radio labeled nucleotides, and in parallel with radio-labeled nucleotides ([α-32P]-dNTPs), and products were assessed by both Southern blotting and radio-incorporation. Southern blotting detects the repeat-containing fragment whether or not it has been repaired; radio-incorporation detects DNA fragments that has incorporated [α-32P]- dNTPs by DNA synthesis (unprocessed starting substrate cannot be visualized). The radio-

Figure 2-8

Figure 2-8. Repair is nick-directed.

(A) Southern blot analysis of starting unprocessed SI-DNA substrates 1, 4 and 5, and their processed products, after digestion (EcoRI/HindIII), resolution on 4% native acrylamide, electrotransfer and probing. Lanes 1, 5 and 9 show starting unprocessed substrates 1, 4 and 5, respectively. Lanes 2, 6 and 10 show substrates processed by HeLa cell extract. Lanes 3, 7 and

11 show substrates processed by RPA-deficient (RPA2 siRNA) HeLa cell extract. Lanes 4, 8 and 12 show substrates processed by RPA-deficient extract supplemented with 600 ng RPA. Substrate schematics are shown at the top. A plus sign in the table indicates the addition of the indicated component. The starting SI-DNA substrates are indicated by black arrowheads. The slow-migrating species that migrate above the starting substrate after processing by cell extracts are indicated by white arrowheads. Repaired products of (CTG)50•(CAG)50 and

(CTG)30•(CAG)30 are indicated by arrows. (B and C) SI-DNA substrates 1, 4 and 5 were incubated with cell extracts with all four radio- [α-32P]dNTPs; the products were digested (EcoRI/HindIII) and resolved on 4% native acrylamide (B) or 6% denaturing sequencing gel (C). Lanes 1, 4 and 7 show substrates processed by HeLa cell extract. Lanes 2, 5 and 8 show substrates processed by RPA-deficient HeLa cell extract. Lanes 3, 6 and 9 show substrates processed by RPA-deficient extract supplemented with 600 ng RPA. Duplex markers for 50 repeats and 30 repeats are indicated in lane M.

48 incorporated repair products were resolved on a denaturing gel as well as a native gel in order to resolve the CTG from the CAG strand and to determine if there were changes in repeat tract size.

DNA synthesis (as indicated by [α-32P]-dNTP incorporation) occurred predominantly in the nicked strand which means that the continuous strand was used as a template for all substrates (Figure 2-8C). The ladder bands seen for the repair of substrate 5 via Southern (Figure 2-8A) were labeled with [α-32P]-dNTP incorporation (although the bands are faint) (Figure 2-8B, C). This is consistent with the ladder being the result of error-prone repair products. In contrast, there was no [α-32P]-dNTP incorporation observed in slow-migrating forms for substrates 4 and 5 (Figure 2-8B and C). Based on the results of this experiment, we concluded that the slow- migrating bands seen by Southern (Figure 2-8A, white arrowheads) were not DNA synthesis products but rather structural alterations to a small subset of unrepaired substrates. The slow- migrating bands occurred in the absence of RPA, thus the structural alterations were not caused by RPA binding but could be the result of altered conformations caused by other proteins (single-strand binding or other). The addition of RPA to the RPA-deficient cell extract removed the slow-migrating bands (Figure 2-3, Figure 2-6, and Figure 2-8A), indicating that RPA either prevented the formation of this subset of molecules or promoted their repair. This conclusion is a tentative one as this experiment was done only once; this experiment will be repeated by other members of the lab to ensure reproducibility.

2.3.5 Yeast RPA and bacterial SSB cannot substitute for human RPA in enhancing repair

RPA is a highly conserved complex, as all eukaryote have homologs of each of the three RPA subunits (Wold 1996). To assess whether scRPA or E. coli single-stranded DNA binding protein (bSSB) can substitute for hRPA in enhancing repair of SI-DNA substrates, we performed in vitro repair assays using RPA-deficient cell extract supplemented with scRPA or bSSB. As we have seen in previous experiments, hRPA was not required for repair as RPA-deficient cell extract repaired as well as RPA-proficient HeLa cell extract. However, adding hRPA to RPA-deficient extract enhanced repair efficiency. For SI-DNA 4, addition of hRPA enhanced repair efficiency to nearly 100% and removed the slow-migrating DNA structures formed by cellular processing. For SI-DNA 5, addition of hRPA enhanced repair and notably decreased the amount of slow- migrating DNA structure (Figure 2-9, compare lane 4 to lane 3). These effects were not observed

Figure 2-9

Figure 2-9. Yeast RPA and bacterial SSB cannot substitute for human RPA in enhancing repair.

Southern blot analysis of starting unprocessed SI-DNA substrates 4 and 5, and their processed products, after digestion (EcoRI/HindIII), resolution on 4% native acrylamide, electrotransfer and probing. A plus sign in the table indicates the addition of the indicated component. The starting SI-DNA substrates are indicated by black arrowheads. The slow-migrating species that migrate above the starting substrate after processing by cell extracts are indicated by white arrowheads. Repaired products of (CTG)50•(CAG)50 and (CTG)30•(CAG)30 are indicated by arrows. Lane 1 shows starting unprocessed SI-DNA substrate. Lane 2 shows substrate processed by HeLa cell extract. Lane 3 shows substrate processed by RPA-deficient cell extract (RPA2 siRNA). Lanes 4 to 6 show RPA-deficient cell extract supplemented with 600 ng of the indicated protein.

50 when scRPA or bSSB were added to RPA-deficient extract (Figure 2-9, compare lanes 5 and 6 to lane 3). The addition of hRPA to the RPA-deficient cell extract removed the slow-migrating unrepaired SI-DNA isomers, indicating that RPA either prevented the formation of this subset of molecules or promoted their repair. In contrast, neither scRPA nor bSSB could block the formation of this SI-DNA form, nor could they enhance their repair. These results show that scRPA and bSSB are not able to substitute for hRPA in enhancing repair of slipped-DNA. This suggests that species-specific RPA-protein interactions may be necessary for repair.

2.3.6 Inhibition of slipped-DNA repair by high concentrations of aRPA

HD patient brains have RPA4 levels that are 6-8 fold higher than levels of RPA2 (Figure 2-4B) suggesting that aRPA levels may be higher in HD brains than canonical RPA. It is known that aRPA has a dominant negative effect on the DNA replication activity of canonical RPA in DNA replication (Haring, Humphreys, and Wold 2010, Mason et al. 2009). We assessed the effect of high levels of aRPA on the repair of slipped-DNA to see if a dominant negative effect can be observed for repair. When aRPA was added to RPA-deficient cell extract in the presence of equal amounts of RPA, repair efficiency remained unchanged from adding RPA or aRPA alone (Figure 2-10A, compare lanes 4 and 5). However, when greater ratios of aRPA were added to RPA-deficient cell extract in the presence of RPA, repair was increasingly inhibited until repair is almost completely inhibited at a ratio of 9:1 (Figure 2-10B, lane 7). Addition of the highest concentration of aRPA in the complete absence of RPA inhibited repair completely (Figure 2- 10B, lane 8). These results demonstrated that while low concentrations of aRPA enhance repair as does RPA, high concentrations of aRPA inhibit repair. In addition, high concentrations of aRPA have a dominant negative effect on the function of canonical RPA in repair. This difference in activity between aRPA and RPA could either be caused by differences in interactions with the SI-DNA substrates or other repair proteins or both.

It is noteworthy that the production of the slower-migrating unrepaired SI-DNA isomers was evident in the absence of RPA or aRPA (Figure 2-10A and B, white arrowheads) and that the addition of RPA eliminated its detection coincident with enhanced correct repair (Figure 2-10A and B, lane 4). Inclusion of aRPA at all concentrations ablated the detection of the slow- migrating SI-DNA species. With increasing aRPA concentrations there was an increase in the

Figure 2-10

Figure 2-10. aRPA inhibits RPA at higher concentrations.

52 arrows. (A) Lane 1 shows starting unprocessed SI-DNA substrate. Lane 2 shows substrate processed by HeLa cell extract. Lane 3 shows substrate processed by RPA-deficient HeLa cell extract (RPA2 siRNA). Lane 4 shows substrate processed by RPA-deficient extract supplemented by 600 ng RPA. Lane 5 shows substrate processed by RPA-deficient extract supplemented by 600 ng RPA as well as 600 ng aRPA (1:1). (B) Lanes 1-4 show the same conditions as in (A). Lane 5 shows substrate processed by RPA-deficient extract supplemented by 600 ng RPA and 1.2 µg aRPA (1:2). Lane 6 shows substrate processed by RPA-deficient extract supplemented by 600 ng RPA and 3.6 µg aRPA (1:6). Lane 7 shows substrate processed by RPA-deficient extract supplemented by 600 ng RPA and 6 µg aRPA (1:10). Lane 8 shows substrate processed by RPA-deficient extract supplemented by 6 µg aRPA only.

53 amount of starting nick-in-repeat SI-DNA, coincident with a reduction of correct repair products. Considering that the nick-in-flank SI-DNAs are directly repaired without being structurally modified prior to repair, this observation supports the idea that the nick-in-repeat SI-DNAs are structurally morphed into forms that are similar to the nick-in-flank slipped-DNAs. Together these results suggest that the starting nick-in-repeat SI-DNAs (black arrowheads) transitioned to the slower-migrating SI-DNA species (white arrowheads) prior to its being repaired (arrows) and that high levels of aRPA inhibited the ability of RPA to facilitate this transition. Thus, it seems like the nick-in-repeat structures must first be converted to structures similar to the nick-in-flank structures prior to the repair and that this conversion does not require any repair synthesis (Figure 2-8). This conversion process can be enhanced by either RPA or aRPA (Figure 2-6).

2.3.7 RPA is limiting for the repair of nick-in-repeat slipped-DNAs, this effect is independent of MutSβ

To determine whether RPA or aRPA is limiting in the repair of slipped-DNAs in the absence of MMR proteins, we processed SI-DNA substrates with hMSH2-null (LoVo) extract, which are devoid of MMR proteins (hMSH2, hMSH3, and hMSH6) (Chang et al. 2000, Panigrahi et al. 2010). Substrates were processed with LoVo cell extract alone, or in combination with RPA, aRPA and MutSβ. Analysis of repair products by Southern blotting compared with starting material permitted quantitative assessment of repair efficiency at a molar level. The results of this analysis are summarized in Figure 2-11.

For nick-in-repeat slipped-DNAs, addition of RPA or aRPA to MMR-deficient LoVo extract enhanced repair efficiency from LoVo extract alone (Figure 2-11C and D, compare lanes 3 and 4 with lane 2). Furthermore, the slow migrating material above the starting substrate (white arrowhead) was eliminated with addition of RPA and aRPA to LoVo extract. The same pattern of repair enhancement by RPA and aRPA was also be observed when LoVo extract is complemented with MutSβ (Figure 2-11C and D, compare lanes 6 and 7 with lane 5). Thus RPA seems to be limiting for the repair of nick-in-repeat slipped-DNAs, as addition of RPA or aRPA enhanced the repair of these substrates. This effect was observed in MMR-deficient LoVo cell extract and when LoVo was complemented with MutSβ, thus this repair enhancement effect by RPA and aRPA is independent of MutSβ.

Figure 2-11

Figure 2-11. RPA is limiting for repair of nick-in-repeat slipped-DNAs.

Southern blot analysis of in vitro repair using SI-DNA substrates, processed by 25 μg of MMR- proficient cell extract (HeLa, lane 1), MMR-deficient cell extract (LoVo, lane 2), LoVo extract supplemented with 600 ng RPA (lane 3), or 600 ng aRPA (lane 4), LoVo extract supplemented with 600 ng MutSβ (lane 5), LoVo extract supplemented with 600 ng MutSβ and 600 ng RPA (lane 6), and LoVo exract supplemented with 600 ng MutSβ and 600 ng aRPA (lane 7). Repair efficiencies were calculated for three to six replicates and adjusted for starting background. Starting SI-DNAs and repaired products are indicated. The slow-migrating species that migrate above the starting substrate after processing by cell extracts are indicated by white arrowheads. Graph shows repaired material (gray bars) and processed material that migrates above the starting SI-DNA (white bars). Asterisk denotes a LoVo-specific product. (A) Repair of SI-DNA

1, with a (CAG)20 slip-out and a 5’ nick in the flanking region on the strand opposite the slip-out.

(B) Repair of SI-DNA 3, with a (CTG)1 slip-out and a 5’ nick in the flanking region on the slip- out strand. (C) Repair of SI-DNA 4, with a (CAG)20 slip-out and a 5’ nick in the repeat tract on the strand opposite the slip-out. (D) Repair of SI-DNA 5, with a (CAG)20 slip-out and a 5’ nick in the repeat tract on the slip-out strand.

For nick-in-flank substrates, addition of RPA or aRPA to LoVo extract did not affect repair efficiency (Figure 2-11A and B, compare lanes 3 and 4 with lane 2). In contrast, addition of MutSβ to LoVo extract more than doubled the repair efficiency for SI-DNA 1 and improved the repair efficiency of SI-DNA 3 to better than HeLa levels. Supplementing RPA or aRPA to LoVo extract in addition to MutSβ improved repair efficiency still further for SI-DNA 1 (Figure 2- 11A), but not for SI-DNA 3 (Figure 2-11B), possibly because the long slip-out in SI-DNA 1 offers more protein-ssDNA interaction opportunities for RPA or aRPA. For nick-in-flank substrates, RPA and aRPA did not enhance repair in the absence of MutSβ, thus it seems the repair enhancement effect by RPA and aRPA is dependent on MutSβ for nick-in-flank silpped- DNAs.

LoVo extracts processed long slip-outs of 20 repeats (SI-DNA 1, 4 and 5) as well as the MMR- proficient HeLa extract, confirming that MutSα and MutSβ are not required for the repair of large slip-outs, regardless of nick location (Figure 2-11A, C, D, compare lanes 1 and 2). In contrast, SI-DNA 3, with a slip-out of 1 repeat, was poorly repaired by LoVo extract compared to the efficient processing by HeLa extract (Figure 2-11B, compare lanes 1 and 2). This is in agreement with previous work that showed the repair of long slip-outs is independent of MMR proteins while repair of short slip-outs relies upon MMR proteins (Panigrahi et al. 2010, Panigrahi et al. 2005).

These results confirm previous observations that the repair of short slip-outs requires MMR proteins while large slip-outs are repaired independently of MMR regardless of nick location. These results also show that RPA and aRPA can enhance repair efficiency for nick-in-repeat slip-outs independently of MutSβ. RPA and aRPA can also enhance repair efficiency for nick-in- flank slip-outs, but this effect is dependent upon MutSβ.

2.3.8 FEN1 is not required for slipped-DNA repair

Flap endonuclease 1 (FEN1) is a structure-specific endonuclease that functions both in DNA replication during Okazaki fragment processing and in long-patch base excision repair (LP-BER) by removing single-stranded DNA flaps before strand ligation (Harrington and Lieber 1994). FEN1 is a critical enzyme for LP-BER in both cell extracts and reconstituted systems (Frosina et al. 1996, Klungland and Lindahl 1997, Balakrishnan and Bambara 2013). A previous study using purified yeast proteins in a reconstituted system showed that the yeast homolog of FEN1, Rad27,

Figure 2-12

Figure 2-12. FEN1 is not required for slipped-DNA repair.

(A) Western blots of HeLa cell extract and FEN1-deficient cell extract (FEN1 RNAi). Actin was used as a loading control. Western blot was performed by Jodie Simard. (B) Southern blot analysis of starting unprocessed SI-DNA substrates 4 and 5, and their processed products, after digestion (EcoRI/HindIII), resolution on 4% native acrylamide, electrotransfer and probing. A plus sign in the table indicates the addition of the indicated component. The starting SI-DNA substrates are indicated by black arrowheads. The slow-migrating species that migrate above the

58 starting substrate after processing by cell extracts are indicated by white arrowheads. Repaired products of (CTG)50•(CAG)50 and (CTG)30•(CAG)30 are indicated by arrows. Lane 1 shows starting unprocessed SI-DNA substrate. Lane 2 shows substrate processed by HeLa cell extract. Lane 3 shows substrate processed by FEN1-deficient cell extract (FEN1 RNAi). Lanes 4 shows FEN1-deficient cell extract supplemented with 600 ng of RPA.

59 was required for the repair of large nick-in-flank slip-outs of random sequence (Sommer et al. 2008).

In order to assess the role of FEN1 in the repair of nick-in-repeat slip-outs, we used a previously established human Fen-Rex cell line which allowed reversible FEN1 knock-down (Fen-Rex cell line was a gift from Dr. Torgeir Holen, University of Oslo; Oslo, Norway). FEN1 knock-down in the Fen-Rex cell line was mediated by a stably-transfected, tetracycline-inducible siRNA against FEN1. The knock-down of FEN1 in the Fen-Rex (+tet) cell line was previously found to be 10- fold compared to the Fen-Rex control (-tet) (Moe, Sorbo, and Holen 2008). Our Western blot analysis confirmed FEN1 knockdown with non-detectable levels of FEN1 in the FEN1-/- cell extract compared to the FEN1+/+ control cell line (Figure 2-12A). Repair assays showed that FEN1 was not required in the repair of nick-in-repeat slip-outs, as the substrates were repaired just as well in the absence of FEN1 (Figure 2-12B, compare lanes 2 and 3). In addition, the repair-enhancing effects of RPA are not dependent upon FEN1, as these effects can still be seen in the absence of FEN1; the addition of RPA removed the slow-migrating DNA structures formed by cellular processing (Figure 2-12B, lane 4). Thus unlike the repair of slip-outs of random sequence, the repair of nick-in-repeat CTG/CAG slip-outs does not require FEN1.

2.3.9 RPA and aRPA bind to and melt slipped-DNAs differently

Previous studies have shown that RPA and aRPA have similar binding properties to short single- stranded oligonucleotides (Mason et al. 2009). However, partially duplexed DNA and slipped- CTG/CAG repeats, in particular, may require different DNA binding kinetics to be processed properly. To assess the binding affinity of purified RPA and aRPA to CTG/CAG slip-outs, we performed DNA binding assays using our disease-length SI-DNAs. Both RPA and aRPA formed protein-DNA complexes with the SI-DNA substrates but not the control fully duplexed CTG/CAG DNA (Figure 2-13A). Both proteins have a binding site size of 20-30 nucleotides (Mason et al. 2009); this indicates that the substrates contain at least 20-30 nucleotides of ssDNA. Binding of RPA gave rise to two distinct protein-DNA bands for higher concentrations of RPA, suggesting that at higher concentrations, two molecules of RPA bound to the substrate. Binding of aRPA gave rise to only one protein-DNA band (Figure 2-13A). Additionally, the protein-DNA complex formed between aRPA and substrate 5 migrated faster than the complex formed between RPA and substrate 5. These results indicate differential binding of RPA and

Figure 2-13

Figure 2-13. RPA and aRPA bind to and melt slipped-DNA differently.

(A) DNA binding assays were carried out as described in Materials and Methods. Audoradiograms of representative DNA binding assays of RPA and aRPA using SI-DNA 4, SI- DNA 5 and a linear DNA substrate containing 50 CTG/CAG repeats are shown. Non- radiolabelled DNA substrates (20 ng) were incubated with 600 ng (lanes 2, 5), 1200 ng (lanes 3, 6), or 2400 ng (lanes 4, 7) of RPA or aRPA, as indicated, digested to release the repeat- containing fragment (EcoRI/HindIII), resolved on 4% native acrylamide, electrotransferred and probed. The positions of free DNA are indicated by black arrowheads; the positions of shifted protein-DNA bound complexes are indicated by white arrowheads. (B) DNA helix

61 destabilization assays were carried out as described in Materials and Methods. Lane 1 shows radiolabeled bubble substrate alone. Lanes 2 and 5 show radiolabeled bubble substrate in the presence of 6.7 nM of the indicated protein. Lanes 3 and 6 show substrate in the presence of 31 nM of the indicated protein. Lanes 4 and 7 show substrate in the presence of 66.7 nM of the indicated protein. Lane 8 shows radiolabeled bubble substrate that has been boiled. DNA helix destabilization assays done by Wold lab (U of Iowa).

62 aRPA to slipped-CTG/CAG repeats. They also suggest that RPA has higher levels of helix destabilization activities than aRPA, which can generate longer stretches of ssDNA for RPA to bind to, as well as affect the final structure of the protein-DNA complex.

We next assessed the helix destabilization abilities of RPA and aRPA. These assays utilized a partially single-stranded substrate with a 20-nucleotide bubble and monitored melting of the DNA in the presence of the different forms of RPA. We found that RPA unwound the bubble substrate more efficiently than aRPA (Figure 2-13B). Taken together, this data suggests that RPA is better able to destabilize hairpins and slip-outs than aRPA and thus can generate longer stretches of ssDNA for RPA to bind to, as well as affect the final structure of the protein-DNA complex.

2.4 Conclusions and Discussion

2.4.1 Nick-in-repeat slip-outs have greater structural heterogeneity than nick-in-flank slip-outs

Circular slipped-DNA substrates are prepared by hybridizing a single-stranded circular plasmid with a linearized duplex DNA. The nick-in-flank substrates were prepared using duplex DNA that was linearized at an EcoRI or HindIII site that is 60 base pairs proximal to the repeat tract, creating a nick in the region flanking the repeat tract (Figure 2-1B). The hybridization reaction for nick-in-flank substrates yielded one heteroduplex product that migrates above the linearized duplex DNA on agarose gel (Figure 2-2A) (Panigrahi et al. 2005). Nick-in-repeat substrate have never been used in previously published studies and I did not know whether the hybridization reaction would yield one product or multiple products. I prepared the nick-in-repeat substrates by using duplex DNA that was linearized at a BsmI cleavage site that is located within the first CTG unit of the CTG tract, creating a nick within the repeat tract (Figure 2-1C). We hypothesized that the nick-in-repeat substrates would have greater structural freedom and increased potential for single-stranded DNA formation on both strands due to the ability of the DNA free ends to slip and slide within the repeat tract. Indeed the hybridization reaction for nick-in-repeat substrates yielded multiple heteroduplex products that migrate above the linearized duplex DNA on agarose gel (Figure 2-2B, C). Repair reactions using these heteroduplex products purified from agarose gel produced the correct repair products (Figure 2-2D, E). Taken together, I concluded that the multiple heteroduplex products seen on agarose gel following hybridization are different structural variations of the same substrate. This confirmed our hypothesis that nick-in-repeat substrates have much higher structural variability than nick-in-flank substrates due to the lack of anchoring by flanking sequences.

2.4.2 Nick-in-repeat slip-outs are better repaired

Slipped-DNA structures can form at repeats in proliferating or non-proliferating cells and serve as mutagenic intermediates for expansions or deletions. The substrates used in this study mimic slipped-DNA intermediates that can form at replication forks, nicked or damaged DNA, and recombination sites. Previous studies using nick-in-flank substrates have shown that repair mechanism is dependent upon slip-out size. Small slip-outs (1-3 excess repeats) repair well via the mismatch repair (MMR) pathway, and large slip-outs (20 excess repeats) repair poorly and

64 independent of MMR and NER proteins (Panigrahi et al. 2005, Panigrahi et al. 2010, Hou et al. 2009, Tian et al. 2009, López Castel, Tomkinson, and Pearson 2009, Littman, Fang, and Modrich 1999a), suggesting that large slip-outs are repaired by another repair mechanism. How this mechanism repairs large slip-outs is not well-understood, although various models have been suggested, including MMR-like error-prone repair (Panigrahi et al. 2005) and incision-based repair (Hou et al. 2009). In this study, I investigated the repair of 20 repeat slip-outs that have a nick within the repeat tract (nick-in-repeat substrates), which allow greater structural freedom than nick-in-flank substrates and better mimics the nicks that can form during replication or repair of repeat tracts (Figure 2-1). I showed that despite having a large slip-out (20 excess repeats), nick-in-repeat substrates repaired accurately and with higher efficiency than their nick- in-flank counterparts (Figure 2-3). This suggests the greater structural freedom of the nick-in- repeat slip-outs allow better repair, possibly through stronger interactions with repair proteins.

A previous study reported that the repair of nick-in-flank CTG/CAG slip-outs can either be correct repair or error-prone repair, which results from incomplete excision of the slip-outs followed by gap-filling and ligation. Notably, error-prone repair is always associated with substrates with a slip-out in the nicked strand (the newly synthesize strand) (Panigrahi et al. 2005). This pattern was observed again for the nick-in-repeat substrates, both by Southern blot and [α-32P]-dNTP incorporation (Figure 2-8). This supports the hypothesis that the error-prone repair of CTG/CAG slip-outs on the nicked strand (the newly synthesized strand) provides a path through which repeat tract expansions (and not deletions) may arise. Error-prone repair removes some, but not all of the excess repeats, leaving a series of heteroduplexes that can lead to repeat expansions.

Interestingly, the repair of nick-in-repeat substrates generates bulky DNA structures within the repeat tract that migrate slower than the starting substrates electrophoretically. These slow- migrating products produced by cell extract processing correspond with the starting material for nick-in-flank substrates. The slow-migrating products are not seen in the repair reactions of nick- in-flank substrates. This indicates that a different set of DNA-binding proteins are involved in the repair of nick-in-repeat substrates, and they can disrupt the conformation of the slip-out structure such that it migrates differently. Taken together, these observations suggest that the repair of large TNR slip-outs is a complex process that may involve different enzymes depending on the structural features and sequence of the slip-out.

2.4.3 RPA and aRPA can enhance slipped-DNA repair

Given that nick-in-repeat substrates have greater potential for single-stranded DNA (ssDNA) formation, we theorized that RPA, the major ssDNA binding protein in humans, would play an important role in the repair of these substrates. RPA’s sister complex aRPA is also a ssDNA binding protein whose affinity for ssDNA is indistinguishable from that of RPA, but the role of aRPA in cellular mechanisms and its relationship with RPA is not clearly understood at the present time. We theorized that aRPA can also affect slipped-DNA repair due to its ssDNA binding ability.

Ever since the detection of RPA4 in human tissues and the discovery that it can complex with RPA1 and RPA3 to form the aRPA complex, much research has gone its elucidating its role in cellular mechanisms. That aRPA seems to be able to substitute RPA in certain types of repair but not in replication (Haring, Humphreys, and Wold 2010, Kemp et al. 2010, Mason et al. 2009), coupled with the finding that RPA4 expression is reduced in cancerous tissues and is very low in stable human cell lines (Haring, Humphreys, and Wold 2010, Keshav, Chen, and Dutta 1995) suggests that aRPA plays an active role in non-proliferating cells but not in proliferating cells. This has led to the hypothesis that RPA4 harbors anti-proliferative properties and has the potential to prevent harmful DNA replication (i.e. in cancer or viral infections). The results presented in this study corroborates the view that aRPA functions in genome maintenance but not replication. The aRPA complex supported the repair of small and large slipped-DNAs and base-base mismatches in the absence of RPA and there was little difference between repair efficiencies by RPA or aRPA (Figure 2-6, 2-7, 2-8). Thus we can conclude that in addition to nucleotide excision repair and repair by homologous recombination, aRPA is also functional in mismatch repair and slipped-DNA repair.

Neither RPA nor aRPA were required for the repair of large slip-outs, small slip-outs or base- base mismatches (Figure 2-6, 2-7, 2-8), but they both enhanced repair efficiency for large slip- outs (Figure 2-6). For nick-in-repeat substrates, addition of RPA or aRPA to the repair action not only increased the amount of correct repair products, it also eliminated the DNA structures that migrate slower than the starting substrates. Thus it seems that the cellular concentrations of RPA and aRPA can affect slipped-DNA repair efficiency and in turn modulate repeat instability. We tentatively conclude that the repair process for nick-in-repeat slipped-DNAs involves first

66 converting the slipped-DNAs to structures similar to the nick-in-flank slip-DNA structures prior to nick-directed excision and re-synthesis. Addition of RPA or aRPA enhances repair by either increasing the speed of repair for the intermediate structures, or preventing the intermediates from forming in the first place and go directly to the repair step.

As demonstrated by Figure 2-9, the repair enhancing effect of RPA is species-specific. RPA activities are conserved to a certain extent as in vitro SV40 DNA replication can be efficiently supported when human RPA (hRPA) is replaced by bovine (Bos Taurus) RPA (Nasheuer et al. 1992) and fruit fly RPA (dmRPA) (Kamakaka et al. 1994). However, even though human RPA and yeast RPA (scRPA) are somewhat homologous and can substitute for each other in certain reactions (Brill and Stillman 1989), the yeast protein cannot replace human RPA in SV40 replication (Melendy and Stillman 1993), and the essential scRPA genes cannot be replaced by the corresponding human coding sequences (Heyer et al. 1990, Brill and Stillman 1989). The ssDNA-binding properties of hRPA, scRPA and bSSB are similar, they all bind to ssDNA with high affinity (Brill and Stillman 1989). Despite this similarity, scRPA and bSSB are unable to substitute for hRPA in SV40 replication or substitute for hRPA in enhancing repair of SI-DNA, likely for the same reasons. Previous studies have found that although hRPA and scRPA both bind ssDNA with high affinity and low cooperativity, they have different binding parameters including binding site size and nucleotide preference (Sibenaller, Sorensen, and Wold 1998). The binding site of scRPA is 10-20nt larger than that of hRPA (Sibenaller, Sorensen, and Wold 1998), thus it is possible that scRPA was not able to enhance repair efficiency because the slipped-DNA site was too small for the complex to bind. The binding site size for bSSB is similar to that of hRPA, which suggests that bSSB was not able to enhance efficiency due to its lack of specific protein-protein interactions within the repair pathway.

In SV40 DNA replication, any number of SSBs can replace RPA during the initial ssDNA binding and T antigen stimulation step to unwind an origin of replication (Kenny, Lee, and Hurwitz 1989, Virshup and Kelly 1989). However, specific protein-protein interactions between RPA and DNA primase-α or T antigen are necessary for the assembly of the initiation complex (Melendy and Stillman 1993, Weinberg et al. 1990). Here I have shown that yeast RPA and bacterial SSB, both of which bind ssDNA with high affinity, are not able to substitute RPA to enhance the repair of large slip-outs (Figure 2-9). This indicates that the ability of RPA to enhance repair requires specific protein-protein interactions in addition to ssDNA-binding. Based

67 on these previous observations and the results shown in this study, we propose a model where another human ssDNA binding protein (SSB), for example the recently identified hSSB1 and 2 (Richard et al. 2008), can substitute for RPA to protect the ssDNA template from degradation or structure-formation in the absence of RPA and thus allow repair of the slip-out. However, the substitute SSB is not able to interact with and stimulate the excision proteins and polymerases that RPA interacts with and thus is not able to enhance repair. RPA has been shown to interact with and stimulate many proteins that are important in DNA repair, including EXO1 (Genschel and Modrich 2003, 2009), XPG (He et al. 1995), XPA (Stigger, Drissi, and Lee 1998) and DNA Ligase I (Ranalli, DeMott, and Bambara 2002). More investigation is needed to elucidate which protein-protein interactions enhance the repair of large slip-outs.

2.4.4 Inhibition of slipped-DNA repair by high concentrations of aRPA

Previous studies have shown evidence that repeat instability may be related to levels of protein expression in different tissues (Mason et al. 2014, Tomé, Manley, et al. 2013, Tomé, Simard, et al. 2013, Goula et al. 2009, Goula et al. 2012). In HD patients and transgenic mice, expression levels of certain DNA repair genes (including RPA) are higher in the striatum (highest repeat instability) than cerebellum (least repeat instability) (Goula et al. 2009, Mason et al. 2014). Additionally, performing repair assays using protein stoichiometry found in HD patients and mice, or with protein extracts prepared from HD mouse brains, showed reduced repair efficiency under striatal conditions, likely because of lower levels of repair proteins (Goula et al. 2012).

The data presented here show that RPA2 and RPA4 expression levels are different in HD patient brains compared to control human brain regions (Figure 2-4). Both RPA2 and RPA4 expression levels were elevated in HD patient brains, but RPA4 expression was elevated 8-fold whereas RPA2 expression was elevated 2-fold. RPA4 expression levels were higher in the striatum (highest repeat instability) than cerebellum (least repeat instability). Based on this data and previous observations that different levels of protein expression may affect repeat instability, we theorized that different concentrations of RPA and aRPA would affect slipped-DNA repair. Figure 2-10 shows that as in DNA replication, the presence of aRPA inhibited RPA activity in enhancing repair. However, the inhibition of repair requires much higher ratios of aRPA:RPA than in replication (Mason et al. 2009) and this ratio is similar to that found in HD patients (Figure 2-4). Taken together, these results suggest that stoichiometry of aRPA:RPA expressed in

68 different tissues affects repair of slipped-DNA and might explain the repeat instability between tissues in TNR diseases.

2.4.5 Repair of large slipped-DNAs is independent of MMR and BER

Slipped-DNAs can form at repeats in proliferating or non-proliferating cells and serve as mutagenic intermediates for expansions or deletions. The substrates used in this study mimic slipped-DNA intermediates that can form at replication forks, nicked or damaged DNA or recombination sites. Previous studies using nick-in-flank substrates have shown that repair mechanism is dependent upon slip-out size. Small slip-outs (1-3 excess repeats) repair well via the mismatch repair (MMR) pathway, and large slip-outs (20 excess repeats) repair poorly and independent of MMR and NER proteins (Hou et al. 2009, Tian et al. 2009, López Castel, Tomkinson, and Pearson 2009, Panigrahi et al. 2010, Littman, Fang, and Modrich 1999a, Panigrahi et al. 2005), suggesting that large slip-outs are repaired by another repair mechanism. How this mechanism repairs large slip-outs is not well understood, although various models have been suggested, including MMR-like error-prone repair (Panigrahi et al. 2005) and incision- based repair (Hou et al. 2009).

Mismatch repair protects against genome-wide mutations and instability by repairing base-base mismatches and small insertion/deletion loops (IDLs). In eukaryotes, there are two protein complexes involved in mismatch recognition: MutSα (MSH2+MSH6), the major mismatch recognition complex, and MutSβ (MSH2+MSH3). MMR is capable of repairing nick-in-flank slip-outs of up to 5 repeats, while larger slip-outs are repair via a poorly understood pathway that is independent of MMR (Hou et al. 2009, Tian et al. 2009, López Castel, Tomkinson, and Pearson 2009, Panigrahi et al. 2010, Littman, Fang, and Modrich 1999a, Panigrahi et al. 2005). Little is known about the repair pathway of nick-in-repeat slip-outs. The data presented here support the hypothesis that large slip-outs are repaired via a novel repair pathway that is independent of known repair pathways. These substrates repaired just as well in the absence of MMR proteins MutSα and MutSβ and the base excision repair (BER) protein FEN1 (Figure 2- 11, 2-12), indicating these large slip-outs are being repaired by another pathway using different protein factors. This MMR- and BER-independent repair mechanism seems to favor the greater structural freedom and single-stranded formation allowed by the nick location within the repeat

69 tract, as these substrates are repaired with greater efficiency than nick-in-flank substrates (Figure 2-3).

2.4.6 MMR does not require RPA

Previous studies using fractionated cell extracts, reconstituted systems or inactivating antibodies have reported that RPA is a required factor in MMR (Lin et al. 1998, Ramilo et al. 2002, Genschel and Modrich 2003). However, RPA requirement was not observed in this effect. SI- DNA 2 and a G-T mismatch substrate, both MMR-dependent substrates, were repaired by RPA- deficient extract as well as RPA-proficient extracts (Figure 2-7), indicating that RPA is not necessary for these substrates to undergo MMR. We propose that another single-stranded DNA binding protein (SSB) can substitute for RPA’s role of protecting ssDNA from degradation or structure formation during MMR (discussed in detail in section 2.4.3). This discrepancy is likely due to differences in the assay systems used in this study and previous studies. In this study I used HeLa cell extract, and its matched, RPA-deficient extract by siRNA knockdown so that the only difference between the cell extracts is the presence or absence of RPA. Previous studies used purified proteins in reconstituted systems or fractionated cell extracts.

2.4.7 RPA and aRPA bind to and denature slipped-DNAs differently

In RPA, two domains of RPA1 (DBD A and B) are both necessary and sufficient for high affinity DNA binding, and RPA2 contributes little to the overall affinity of the complex for ssDNA (Walther et al. 1999, Sibenaller, Sorensen, and Wold 1998). Therefore, aRPA, which contains RPA1, RPA4 and RPA3, was expected to have similar binding properties as RPA. Previous studies have shown that RPA and aRPA indeed have similar binding properties and modes (Mason et al. 2009). Biochemical and binding studies have shown that aRPA is a stable heterotrimeric complex similar in size and stability to the canonical RPA, and aRPA interacts with ssDNA in a manner indistinguishable from canonical RPA (Kemp et al. 2010, Keshav, Chen, and Dutta 1995). Given the similarities between the two complexes it was expected that both would bind slipped-DNAs in the same way. However, the binding assay in this study showed that RPA and aRPA bind to slipped-DNAs differently (Figure 2-13A). Two distinct protein-DNA bands were observed for RPA whereas only one was observed for aRPA. This indicates that two molecules of RPA can bind to the nick-in-repeat substrates while only one molecule of aRPA can bind. In addition, the protein-DNA complex formed by RPA and aRPA

70 migrate differently for SI-DNA 5, which indicates that the protein-DNA complexes formed by the two different proteins gave rise to different structures.

RPA and aRPA are the same size and structure, and the ssDNA-binding properties of RPA and aRPA have been shown to be indistinguishable, with similar affinity, cooperativity, and binding site requirement (Keshav, Chen, and Dutta 1995, Kemp et al. 2010). Thus we hypothesized that RPA and aRPA have different helix destabilization properties wherein RPA can better destabilize the DNA double helix to provide more ssDNA to bind to. This would allow more RPA complexes to bind to the substrates and alter the substrate structure in a way that differs from aRPA. Indeed the helix destabilization assay showed that RPA was able to unwind a bubble substrate better than aRPA (Figure 2-13B). Thus RPA can better destabilize the DNA double helix to provide more ssDNA to bind to, which allows more RPA complexes to bind to slipped- DNAs and alter the DNA structure

2.4.8 aRPA function

The results presented in this study corroborates the view that aRPA functions in genome maintenance but not replication. The aRPA complex supported the repair of SI-DNA substrates in the absence of RPA and there was little difference between repair efficiencies by RPA or aRPA (Figure 2-6, 2-7). As in DNA replication, the presence of aRPA inhibited RPA activity in repair (Mason et al. 2009). However, the inhibition of repair requires much higher ratios of aRPA:RPA than in replication (Figure 2-10). Interestingly, mRNA studies showed that RPA4 expression is much higher than RPA2 expression in the brains of HD patients, and RPA4 expression levels are higher in the striatum (highest repeat instability) than cerebellum (least repeat instability). Control brains showed no difference between RPA4 expression and RPA2 expression and between tissues (Figure 2-4). Previous studies have shown in HD transgenic mice that expression levels of certain DNA repair genes are higher in the cerebellum (least repeat instability) than the striatum (highest repeat instability) (Goula et al. 2009, Mason et al. 2014). Additionally, performing repair assays (BER) using the protein stoichiometry mimicking the levels present in HD mouse striatum or cerebellum, or with protein extracts prepared from HD mouse striatum or cerebellum showed reduced repair efficiency under striatal conditions, likely because of the lower levels of BER proteins (Goula et al. 2012). Taken together, these results

71 suggest that stoichiometry of aRPA:RPA expressed in different tissues might explain the vast repeat instability difference that can exist between tissues in TNR diseases.

3 Summary and future directions 3.1 Thesis summary

This thesis addresses the role of DNA repair in disease-associated trinucleotide repeat instability, specifically focusing on CTG/CAG repeats. TNR instability is the cause of several human diseases, including myotonic dystrophy type 1, Huntington’s disease, several spinocerebellar ataxias, and fragile X syndrome. The levels of instability differ between tissues in a manner dependent upon the affected locus. It is important to understand how repeat expansions occur, as increased instability is believed to coincide with increased disease severity and progression. The mechanism behind instability is thought to involve the formation or aberrant processing of slipped-DNA structures which can form during DNA metabolic processes including DNA replication and repair. While certain DNA repair proteins are implicated in causing ongoing instability, the method by which this occurs is still not well understand and is an area of active investigation. My experiments addressed how slipped-DNA structures formed by CTG/CAG repeats are processed (occasionally in a mutagenic fashion) and the proteins that are involved in slipped-DNA repair.

I assessed how characteristics of repeat slip-outs including slip-out size, slip-out sequence and nick location affect their repair, and also determined whether these repair outcomes were dependent upon the presence of certain repair factors. In agreement with previous studies, I showed that shorter slip-outs repair much better than longer slip-outs, and short slip-outs are repaired via mismatch repair while longer slip-outs are repaired independently of MMR. I showed that nick location has a striking effect on repair efficiency, as long slip-outs that have a nick within the repeat tract repair much better than their counterparts with a nick in the flanking region. I confirmed that slip-outs on the continuous strand are repaired correctly while slip-outs on the nicked strand undergo error-prone repair that generates a series of heteroduplexes that can lead to expansion mutations. That error-prone repair is restricted to expansion SI-DNA substrates may explain the repeat expansion bias that occurs in repeat diseases. Like the nick-in-flank repeats with long slip-outs, the nick-in-repeat substrates with long slip-outs are also repaired independently of MMR proteins and the crucial BER protein FEN1. Thus it is likely that while short TNR slip-outs are repaired via the mismatch repair pathway, long slip-outs are repaired by an alternative repair pathway involving different proteins.

I assessed the role of RPA and aRPA in the repair of slip-outs and found that RPA and aRPA are not required for MMR nor for the alternative repair pathway that repairs long slip-outs. It is likely that another single-stranded DNA binding protein can substitute for RPA’s role of protecting ssDNA. Although RPA is not required for repair, it can enhance repair efficiency, indicating that RPA protein-protein interactions stimulate other proteins involved in the repair pathway. Additionally, I showed that like RPA, aRPA is also able to enhance repair, which supports the hypothesis that aRPA has RPA-like roles in genome maintenance but not genome replication. Our collaborators from the La Spada Lab (UC San Diego) assessed mRNA expression levels of RPA2 (RPA complex) and RPA4 (aRPA complex) and found both to be over-expressed in HD patient brains compared to human brains, although the level of overexpression differed between different regions of the brain. I assessed the effects of aRPA:RPA stoichiometry on repeat instability and found that higher concentrations of aRPA (as might be found in HD brains) inhibits repair and eliminates repair enhancement by RPA. This suggests that the stoichiometry of key repair proteins such as the RPA complexes in different tissues can explain the vast differences in tissue instability that is observed in some repeat diseases. Lastly, the differential DNA binding of the two RPA complexes to the SI-DNA substrates used in this study suggests that there are be subtle differences between the DNA binding properties of the two complexes that require further investigation.

3.2 Future Directions

3.2.1 What are the effects of sequence on repair of TNR slip-outs?

Most studies that investigate the role of repair proteins in repeat instability have centered upon CTG/CAG repeats; however, the role of these proteins in instability at other TNRs (CGG/CCG, GAA/TTC) may differ, if they play a role at all. In a yeast model of Fragile X (CGG) expansions, knocking out msh2, msh3 and msh6 did not affect repeat instability (White, Borts, and Hirst 1999). In a human Friedreich’s ataxia (FRDA) cell line model and in FRDA induced pluripotent stem cells, knock down of MSH2 led to a decrease in GAA repeat instability (Ku et al. 2010, Halabi et al. 2012). This effect was also seen in FRDA mice, where Msh2 knockout led to somatic stability of the repeat (similar to DM1 and HD mice) (Bourn et al. 2012). Recently it was found that many cases of sporadic amyotrophic lateral sclerosis (ALS) are linked to a hexanucleotide expansion (GGGGCC) (DeJesus-Hernandez et al. 2011). There is currently no

74 knowledge about how MMR proteins interact with this repeat sequence. Different repeat sequences can form different structures, and this could affect either their recognition by repair proteins, or the conditions under which the structures may form. While CTG/CAG repeats form hairpins and unpaired loops, GAA repeats are believed to form a triplex structure, and G-rich sequences such as CGG repeats and GGGGCC repeats, can form quadruplexes (Reddy et al. 2013). In the case of DM2 and SCA10, only one of the two strands is capable of forming structures which may affect replication-mediated instability (Dere et al. 2004, Handa et al. 2005). It would be informative to repeat the in vitro repair and replication assays described in this thesis to investigate the repair of other slipped repeat sequences, or replication through these repeat tracts both with and without MMR proteins present, in order to learn further about the mechanism behind their expansions.

3.2.2 What causes tissue-specific repeat instability?

Disease-causing repeat tracts can continue to expand throughout a patient’s lifetime with longer repeat tracts being more prone to further expansions; however, even with identical starting numbers of repeats, instability differs between the various TNR loci and between tissues within a patient. One of the key questions in the field of repeat diseases is what causes tissue-specific instability. For example, most CTG/CAG expansion diseases show instability in the striatum but not in the cerebellum (López Castel, Cleary, and Pearson 2010, López Castel et al. 2011). This is a very complex question due to the numerous factors that differ between tissues (including the multitude of cell types and different gene expression patterns), and also due to the various ways in which instability can occur. Ongoing instability occurs in neurons, indicating that replication is not required for instability; however, in primary fibroblast cell lines of DM1 patients it has been shown that instability does not occur without replication (Yang et al. 2003).

The specific factors which lead to a repeat tract being unstable in one tissue but stable in another are not currently known. Studies using a transgenic HD mouse model have shown that increased repeat instability in certain tissues correlates with decreased expression of genes involved in replication and repair (Goula et al. 2009, Goula et al. 2012). It would be illuminating to investigate if this correlation between protein expression and repeat instability can be found in human patients. Western blots using protein extracts derived from different tissues of patients as well as control individuals can either confirm or deny the hypothesis that expression levels of

75 certain replication and repair genes governs repeat instability. It would be ideal to confirm this correlation in multiple repeat diseases to rule out disease-specific biases, as different diseases have different patterns of tissue-specific repeat instability. The results from my experiments suggest that higher concentrations of aRPA inhibit repair and can lead to increased repeat instability. Preliminary mRNA studies show that aRPA expression levels are increased in the brains of HD patients compared with control brains, while RPA expression levels remain the same. This suggests that protein stoichiometry of the RPA complexes contributes to tissue- specific repeat instability in repeat diseases. Further protein-based investigation involving more tissues and more repeat diseases is needed to determine the role of RPA complexes in repeat instability. It would be interesting to assess whether different protein stoichiometry in different tissues translates to different repair capacities. This can be tested by performing repair experiments with tissue-specific cell extracts. For long repeat slip-outs, the relative levels of correct/error-prone repair could differ. Perhaps unstable tissues have low expression of vital genome maintenance genes, which leads to decreased repair capacity and more error-prone repair, which leads to more expansion events and more slip-out structures.

3.2.3 How are long slip-outs repaired?

My results and other studies have shown that while short TNR slip-outs are repaired via the mismatch repair pathway, repair of long TNR slip-outs are independent of MMR and BER (Panigrahi et al. 2010, Hou et al. 2009). In addition, long slip-outs with a nick on the slip-out strand undergoes error-prone repair, which is not observed for any other substrate. This suggests that long TNR slip-outs are repaired via an unknown pathway and the repair mechanism is different depending on nick location. Which begs the question, how are long slip-outs repaired and which proteins are involved? To answer this question, we can investigate which proteins bind to long slipped-out repeats and whether different proteins bind depending on nick location. Biotin-labelled DNA substrates with a long repeat slip-out on either strand could be incubated with cell extracts, and then streptavidin-coated beads could be used to separate the proteins which bind to the slip-out. Substrates with fully duplexed CTG/CAG repeats could be included as a competitor to prevent enriching proteins that simply bind to DNA. Mass spectrometry could be used to identify the proteins that bind to the different substrates, which can then be compared to find protein that may be contributing to instability. These candidates could be tested for their role in instability via the in vitro replication/repair assays, or by siRNA knockdown (or

76 overexpression) in cell lines that display repeat instability. This method can also be used to study protein interaction differences between tissues to understand which proteins are involved in processing large slip-outs and how they differ between tissues.

3.2.4 What are potential therapies?

At the present time, there are no treatments which attack the root cause of TNR diseases – the repeat expansion mutation; current therapies target disease symptoms as they arise. For the repeat diseases where the pathogenic mechanism is believed to be a toxic gain of function at the RNA level, such as DM1 and several forms of SCA, the search for treatments has focused on targeting the disease at the RNA level with antisense oligonucletoides (ASOs) or small molecules. The intention is to either cause the degradation of the mutant RNA, or decrease its interaction with the proteins that they bind to and sequester (MBNL1 in the case of DM1) (Foff and Mahadevan 2011, Gao and Cooper 2013). Several studies have effectively employed ASOs for targeting the toxic RNA with therapeutic benefit in human cell and mouse models of myotonic dystrophy (Wheeler et al. 2009, Mulders et al. 2009). Technology is constantly developing on this front to improve ASO stability within cells and efforts are continually being made to improve delivery and as a result several therapies are entering clinical trials (Gao and Cooper 2013). In the case of myotonic dystrophy, the major therapeutic target tissue is muscle which is considerably easier to administer ASOs to, compared to neurons which are also affected in myotonic dystrophy but are the primary affected cell type in other neurodegenerative diseases such as fragile X tremor ataxia syndrome, amyotrophic lateral sclerosis and frontotemporal dementia (Nelson, Orr, and Warren 2013). The major limitation to ASO treatment is currently administration across the blood brain barrier (Gao and Cooper 2013). Efforts are currently underway to facilitate neuronal targeting and crossing of the blood-brain barrier with ASOs but other groups have focused upon screening small drug molecules to identify compounds with high therapeutic potential and targeting efficacy (Guan and Disney 2012).

Therapies with targets downstream of the DNA mutation (such as targeting the MBNL1-RNA interaction) would be less effective for treating TNR diseases than stopping or reversing the ongoing DNA expansions. Identifying factors (most likely proteins or protein complexes) that are required for expansions or factors that cause repeat tract contractions provides targets for therapeutic intervention. For example, MutSβ has been found to be necessary for ongoing

77 expansions in mice (van den Broek et al. 2002, Foiry et al. 2006), thus targeting this MMR protein could decrease instability and perhaps be effective for several different CTG/CAG repeat diseases. Recently the structure of MutSβ bound to DNA was solved (Gupta, Gellert, and Yang 2012), and as such, directed drug discovery could be used to find small molecules which can inhibit this binding. The mechanism of mismatch detection differs between MutSβ and MutSα, so despite the redundancy in their substrates, it should be possible to disrupt the actions of one without affecting the other. Initial trials of efficacy could employ the in vitro repair assay described in this thesis using short slip-out substrates. Good candidates would inhibit repair of a short slip-out substrate, which requires MutSβ for repair, but not the repair of a mismatch substrate, which requires MutSα for repair (Panigrahi et al. 2010); the goal is to inhibit MutSβ activity but not MutSα. The loss of MSH3 does not lead to tumorigenesis despite increased instability at certain microsatellites (Campregher et al. 2012); additionally, Msh3-/- DM1 mice displayed CTG/CAG contractions instead of expansions (van den Broek et al. 2002, Foiry et al. 2006). Thus targeting this protein has the potential to cure disease if perturbations to this protein can cause repeats to contract into the normal range.

DM1 families typically display CTG repeat expansions of varying magnitudes but also show contractions and stable transmissions (Salehi et al. 2007, Ashizawa et al. 1994). A recent study reported that in the French-Canadian DM1 population, intergenerational contractions occur in about 7.4% of transmissions (Puymirat et al. 2009). The source of variations in instability patterns is unknown; understanding how these intergenerational contractions occur can offer new targets in the search for therapeutics. One possible explanation for the variations in instability patterns is genetic variation. A SNP analysis could be done to identify polymorphic variants that are unique to the families with intergenerational contractions. Such variants may serve as targets for therapeutic treatment either through enhancing or ablating their attributes. Additionally, such gene variants may serve as predictors of instability, age-of-onset and disease severity.

3.3 Concluding remarks

The slipped-DNA structures assumed by disease-associated CTG/CAG repeats contribute to repeat instability and affect multiple downstream processes to cause disease. The formation of these structures and the repeat instability that is associated seem to be affected by protein stoichiometry in different tissues. Understanding how these structures are repaired and which

78 proteins are involved in the repair pathway will allow therapeutic treatments to attack these diseases at their root – prevent the disease-associated repeat tracts from expanding or induce contractions. The mechanism of instability is proving to be quite complex; however, advances in our understanding of these mechanisms could allow for highly specific and effective treatments for this growing list of repeat diseases.

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