BIOCHEMICAL CHARACTERIZATION OF APRATAXIN, THE PROTEIN DEFICIENT IN ATAXIA WITH OCULOMOTOR APRAXIA TYPE 1

This work was completed at the Queensland Institute of Medical Research by:

JANELLE LOUISE HANCOCK B.App Sci (Biochem, Hons)

and is submitted for the award of Doctor of Philosophy at the Queensland University of Technology

September 2008

i ii STATEMENT OF ORIGINALITY

I hereby declare that I am the sole author of this work and any content from other sources has been acknowledged and fully cited. The following material has not been submitted, either in part or whole, for a degree at this or any other institution. This thesis was prepared in accordance with the regulations outlined by the Queensland University of Technology, for the degree of Doctor of Philosophy. The research within this thesis was carried out under the principal supervision of Professor Martin Lavin and Dr Olivier Becherel.

……………………………………………….. Janelle Louise Hancock

iii iv ABSTRACT

Neurodegenerative disorders are heterogenous in nature and include a range of ataxias with oculomotor apraxia, which are characterised by a wide variety of neurological and ophthalmological features. This family includes recessive and dominant disorders. A subfamily of autosomal recessive cerebellar ataxias are characterised by defects in the cellular response to DNA damage. These include the well characterised disorders Ataxia-Telangiectasia (A-T) and Ataxia-Telangiectasia Like Disorder (A-TLD) as well as the recently identified diseases Spinocerebellar ataxia with axonal neuropathy Type 1 (SCAN1), Ataxia with Oculomotor Apraxia Type 2 (AOA2), as well as the subject of this thesis, Ataxia with Oculomotor Apraxia Type 1 (AOA1). AOA1 is caused by mutations in the APTX gene, which is located at chromosomal locus 9p13. This gene codes for the 342 amino acid protein Aprataxin. Mutations in APTX cause destabilization of Aprataxin, thus AOA1 is a result of Aprataxin deficiency.

Aprataxin has three functional domains, an N-terminal Forkhead Associated (FHA) phosphoprotein interaction domain, a central Histidine Triad (HIT) nucleotide hydrolase domain and a C-terminal C2H2 zinc finger. Aprataxins FHA domain has homology to FHA domain of the DNA repair protein 5’ polynucleotide kinase 3’ phosphatase (PNKP). PNKP interacts with a range of DNA repair proteins via its FHA domain and plays a critical role in processing damaged DNA termini. The presence of this domain with a nucleotide hydrolase domain and a DNA binding motif implicated that Aprataxin may be involved in DNA repair and that AOA1 may be caused by a DNA repair deficit. This was substantiated by the interaction of Aprataxin with proteins involved in the repair of both single and double strand DNA breaks (X- Ray Cross-Complementing 1, XRCC4 and Poly-ADP Ribose Polymerase-1) and the hypersensitivity of AOA1 patient cell lines to single and double strand break inducing agents.

At the commencement of this study little was known about the in vitro and in vivo properties of Aprataxin. Initially this study focused on generation of recombinant Aprataxin proteins to facilitate examination of the in vitro properties of Aprataxin. Using recombinant Aprataxin proteins I found that Aprataxin binds to double stranded DNA. Consistent with a role for Aprataxin as a DNA repair enzyme, this binding is

v not sequence specific. I also report that the HIT domain of Aprataxin hydrolyses adenosine derivatives and interestingly found that this activity is competitively inhibited by DNA. This provided initial evidence that DNA binds to the HIT domain of Aprataxin. The interaction of DNA with the nucleotide hydrolase domain of Aprataxin provided initial evidence that Aprataxin may be a DNA-processing factor.

Following these studies, Aprataxin was found to hydrolyse 5’adenylated DNA, which can be generated by unscheduled ligation at DNA breaks with non-standard termini. I found that cell extracts from AOA1 patients do not have DNA-adenylate hydrolase activity indicating that Aprataxin is the only DNA-adenylate hydrolase in mammalian cells. I further characterised this activity by examining the contribution of the zinc finger and FHA domains to DNA-adenylate hydrolysis by the HIT domain. I found that deletion of the zinc finger ablated the activity of the HIT domain against adenylated DNA, indicating that the zinc finger may be required for the formation of a stable enzyme-substrate complex. Deletion of the FHA domain stimulated DNA-adenylate hydrolysis, which indicated that the activity of the HIT domain may be regulated by the FHA domain. Given that the FHA domain is involved in protein-protein interactions I propose that the activity of Aprataxins HIT domain may be regulated by proteins which interact with its FHA domain.

We examined this possibility by measuring the DNA-adenylate hydrolase activity of extracts from cells deficient for the Aprataxin-interacting DNA repair proteins XRCC1 and PARP-1. XRCC1 deficiency did not affect Aprataxin activity but I found that Aprataxin is destabilized in the absence of PARP-1, resulting in a deficiency of DNA-adenylate hydrolase activity in PARP-1 knockout cells. This implies a critical role for PARP-1 in the stabilization of Aprataxin.

Conversely I found that PARP-1 is destabilized in the absence of Aprataxin. PARP-1 is a central player in a number of DNA repair mechanisms and this implies that not only do AOA1 cells lack Aprataxin, they may also have defects in PARP-1 dependant cellular functions. Based on this I identified a defect in a PARP-1 dependant DNA repair mechanism in AOA1 cells.

vi Additionally, I identified elevated levels of oxidized DNA in AOA1 cells, which is indicative of a defect in Base Excision Repair (BER). I attribute this to the reduced level of the BER protein Apurinic Endonuclease 1 (APE1) I identified in Aprataxin deficient cells.

This study has identified and characterised multiple DNA repair defects in AOA1 cells, indicating that Aprataxin deficiency has far-reaching cellular consequences.

Consistent with the literature, I show that Aprataxin is a nuclear protein with nucleoplasmic and nucleolar distribution. Previous studies have shown that Aprataxin interacts with the nucleolar rRNA processing factor nucleolin and that AOA1 cells appear to have a mild defect in rRNA synthesis. Given the nucleolar localization of Aprataxin I examined the protein-protein interactions of Aprataxin and found that Aprataxin interacts with a number of rRNA transcription and processing factors. Based on this and the nucleolar localization of Aprataxin I proposed that Aprataxin may have an alternative role in the nucleolus. I therefore examined the transcriptional activity of Aprataxin deficient cells using nucleotide analogue incorporation. I found that AOA1 cells do not display a defect in basal levels of RNA synthesis, however they display defective transcriptional responses to DNA damage.

In summary, this thesis demonstrates that Aprataxin is a DNA repair enzyme responsible for the repair of adenylated DNA termini and that it is required for stabilization of at least two other DNA repair proteins. Thus not only do AOA1 cells have no Aprataxin protein or activity, they have additional deficiencies in PolyADP Ribose Polymerase-1 and Apurinic Endonuclease 1 dependant DNA repair mechanisms. I additionally demonstrate DNA-damage inducible transcriptional defects in AOA1 cells, indicating that Aprataxin deficiency confers a broad range of cellular defects and highlighting the complexity of the cellular response to DNA damage and the multiple defects which result from Aprataxin deficiency. My detailed characterization of the cellular consequences of Aprataxin deficiency provides an important contribution to our understanding of interlinking DNA repair processes.

vii KEYWORDS: Ataxia with Oculomotor Apraxia Type 1 (AOA1); Early onset ataxia with hypoalbuminemia (EAOH); Autosomal recessive cerebellar ataxia (ARCA); DNA repair; Aprataxin; Base Excision Repair (BER); single strand break repair (SSBR); APTX.

viii LIST OF PUBLICATIONS

Refereed Publications:

Kijas, A. W., Harris, J. L., Harris, J. M., Lavin, M. F. (2006). Aprataxin forms a discrete branch in the HIT (histidine triad) superfamily of proteins with both DNA/RNA binding and nucleotide hydrolase activities. The Journal of Biological Chemistry. 281(20):13939-48.

Poster Presentations:

Queensland Institute of Medical Research Student Conference, 2005 Janelle L Harris, Amanda W Kijas, Martin F Lavin, The Histidine Triad domain of Aprataxin has novel DNA binding capability.

East Coast Protein Meeting, 2005 Janelle L Harris, Amanda W Kijas, Martin F Lavin, The Histidine Triad domain of Aprataxin has novel DNA binding capability.

Australian Society for Medical Research, Queensland Conference, 2007 Janelle L Harris, Amanda W Kijas, Martin F Lavin, The Histidine Triad domain of Aprataxin has novel DNA binding capability.

Ataxia Telangiectasia Workshop, 2008. Janelle L Harris, Olivier Becherel, Martin F Lavin, Biochemical Characterization of the DNA repair protein Aprataxin.

Australian Society for Medical Research, Queensland Conference, 2008. Janelle L Harris, Olivier Becherel, Martin F Lavin, Aprataxin has a unique role in DNA repair.

ix x CONFERENCES AND INVITED SEMINARS

East Coast Protein Meeting, 2007 Janelle L Harris, Martin F Lavin, Aprataxin has a Unique Role in DNA repair.

Queensland Institute for Medical Research Student Conference, 2007 Janelle L Harris, Olivier Becherel, Martin Lavin.

Queensland Institute for Medical Research Student Seminar, 2008 Janelle L Harris, Olivier Becherel, Martin Lavin, The protein Aprataxin has a novel role in DNA repair.

National Institute for Medical Research (London, UK), 2008 Janelle L Harris, Olivier Becherel, Martin Lavin, Synergistic function of Aprataxin and PARP-1 in DNA repair.

University of Sussex (Brighton, UK), 2008 Janelle L Harris, Olivier Becherel, Martin Lavin, Synergistic function of Aprataxin and PARP-1 in DNA repair.

National Institute for Aging (Baltimore MD, USA), 2008 Janelle L Harris, Olivier Becherel, Martin Lavin, Synergistic function of Aprataxin and PARP-1 in DNA repair.

Queensland Protein Group Conference, 2008 Janelle L Harris, Olivier Becherel, Martin Lavin, Synergistic function of Aprataxin and PARP-1 in DNA repair.

xi xii AWARDS, GRANTS AND PRIZES

Awards and Grants:

ASBMB student prize for Queensland Protein Group Conference seminar, to attend COMBIO 2008 ($500)

Queensland Institute for Medical Research Higher Degrees Committee Travel Grant to attend Ataxia Telangiectasia Workshop 2008 ($1,500)

Queensland University of Technology, Faculty of Science Travel Grant to attend Ataxia Telangiectasia Workshop 2008 ($2,000)

Queensland University of Technology, School of Life Sciences Travel Grant to attend Ataxia Telangiectasia Workshop 2008 ($1,750)

Queensland Cancer Fund, Travel Grant to attend Ataxia Telangiectasia Workshop 2008 ($3,000)

Queensland Institute of Medical Research, Small Equipment Grant 2006 ($2,500)

Queensland University of Technology, Blueprint Scholarship (APA equivalent, 2005- 2007)

Conference Prizes:

Queensland Protein Group Conference 2008, ASBMB Travel Scholarship for the seminar: Synergistic function of Aprataxin and PARP-1 in DNA repair.

East Coast Protein Meeting 2007, Student Oral Presentation Prize for seminar: Aprataxin has a Unique Role in DNA repair.

xiii xiv ACKNOWLEDGEMENTS

I would like to thank my supervisors Professor Martin Lavin, Dr Amanda Kijas, Dr Olivier Becherel and Associate Professor Terry Walsh. I am especially grateful to Amanda, who started me down this long road, and Olivier, for seeing me to the end of it.

I would also like to thank Dr Nuri Gueven for his ‘stimulating’ discussions and Dr Sergei Kozlov, who introduced me to most of the radioactive techniques used in this thesis. Also no words can express the gratitude the whole lab has for Aine Farrell, who looks after all of us (and our cells, and our ordering…) to keep the wheels moving. So thanks everyone, it’s been a fantastic few years.

Last (and certainly not least), I couldn’t have done this without the support of my husband Jonathan, who has had to deal with my PhD-related mood swings for nearly four years. To my best friends Carina and Crystal, our regular ‘tune-out’ nights have kept us all sane (there’s nothing three girls can’t deal with provided they have enough pizza and chocolate). I’m also grateful to my parents who taught me to question everything, especially my father Neil taught me one of the most fundamental concepts I’ve needed for this degree: if you start something, you had better finish it.

xv xvi TABLE OF CONTENTS

Statement of Originality iii Abstract v Keywords viii List of Publications ix Conferences and Invited Seminars xi Grants, Awards and Prizes xiii Acknowledgments xv Detailed Contents xvii List of Figures xxiv List of Tables and Equations xxx List of Chemicals and Suppliers xxxi List of Antibodies and Suppliers xxxiv Supplier Addresses xxxvi Commonly Used Abbreviations xxxvii

DETAILED CONTENTS CHAPTER 1 Literature Review 1

1.1 Autosomal Recessive Cerebellar Ataxias 3 1.1.1 Ataxia Telangiectasia 3 1.1.2 Ataxia Telangiectasia Like Disorder 5 1.1.3 Xeroderma Pigmentosum 6 1.1.4 Ataxia with Oculomotor Apraxia Type 2 7 1.1.5 Spinocerebellar Ataxia with Axonal Neuropathy 8 1.1.6 Ataxia with Oculomotor Apraxia Type 1 10

1.2 Structure and Function of Aprataxins domains 13 1.2.1 The FHA domain 13 1.2.2 The HIT domain 15 1.2.3 The zinc finger 23

xvii 1.3 DNA repair mechanisms 24 1.3.1 Double strand break repair pathways 26 1.3.1.1 NHEJ 27 1.3.1.2 HR 30 1.3.2 Single strand break repair pathways 31 1.3.2.1 Direct SSBR 32 1.3.2.2 Indirect SSBR (NER) 36 1.3.2.3 Indirect SSBR (MMR) 37 1.3.2.4 Indirect SSBR (BER) 38

1.4 General conclusions and aims of this thesis 41

1.5 References 42

CHAPTER 2 Generation of recombinant Aprataxin and Aprataxin-specific antibodies 63

2.1 Introduction 65 2.1.1 Expression and purification of recombinant proteins 65 2.1.2 Generation of specific antibodies 66

2.2 Materials and Methods 68 2.2.1 Expression constructs 68 2.2.2 Plasmid preparation 69 2.2.3 Preparation of Competent Bacterial Cells 69 2.2.4 Transformation of Competent Cells 69 2.2.5 Expression and purification of recombinant Aprataxin from pTYB1 70 2.2.6 Expression and purification of recombinant Aprataxin from pGEX 6.1 71 2.2.7 Purification of anti-Aprataxin specific antibodies 72 2.2.8 Estimation of protein concentration 73 xviii 2.2.8 Protein electrophoresis 73

2.3 Results 74 2.3.1 Purification of recombinant Aprataxin using pTYB1 74 2.3.2 Generation of recombinant Aprataxin using pGEX 6.1 81 2.3.3 Purification of Aprataxin-specific antibodies 86

2.4 Discussion 90

2.5 References 92

CHAPTER 3 Biochemical characterization of recombinant Aprataxin 95

3.1 Introduction 97 3.1.1 Domain structure of Aprataxin 97

3.2 Materials and Methods 99 3.2.1 Partial chymotryptic proteolysis 99 3.2.2 Electrophoretic mobility shift assay (EMSA) 99 3.2.3 Nucleotide Hydrolysis 100 3.2.4 Single strand break repair by cell extracts 102 3.2.5 Single strand break repair by recombinant ligase 104 3.2.6 Adenylation of DNA by T4 DNA ligase 104 3.2.7 Binding of Aprataxin to adenylated DNA 105 3.2.8 Hydrolysis of adenylated DNA by Aprataxin 105 3.2.9 3’ phosphatase activity assays 105

3.3 Results 107 3.3.1 Characterization of the binding activities of Aprataxin 107 3.3.2 Characterization of the Aprataxins nucleotide hydrolase activity 114

xix 3.3.3 Characterization of the interaction between DNA binding and nucleotide hydrolysis 122 3.3.4 Inhibition of single strand break repair by 3’ terminal DNA damage 127 3.3.5 Binding of Aprataxin to 5’ adenylated DNA 131 3.3.6 Hydrolysis of 5’ adenylated DNA by Aprataxin 132 3.3.7 Examination of 3’ phosphatase activity 135 of Aprataxin

3.4 Discussion 137

3.5 References 140

CHAPTER 4 Characterization of multiple DNA repair defects in AOA1 cells 143

4.1 Introduction 145 4.1.1Assembly of a ‘repairosome’ 145 4.1.2 Aprataxin localization and post-translational modifications 146 4.1.3 A role for Aprataxin in multiple DNA repair pathways 146

4.2 Materials and Methods 148 4.2.1 2D gel electrophoresis 148 4.2.2 In vitro kinase assay 148 4.2.3 DNA binding of endogenous Aprataxin 150 4.2.4 DNA-adenylate hydrolase activity of nuclear extracts 150 4.2.5 Effect of 3’ oxidation on 5’ adenylate hydrolysis by extracts 151 4.2.6 In vitro single strand break repair by cell extracts 151 4.2.7 Effect of Aprataxin on protein stability 152 xx 4.2.8 8-oxo-dG immunostaining 153 4.2.9 Nitrotyrosine immunostaining 153 4.2.10 Subcellular distribution of Aprataxin 154 4.2.11 Subcellular distribution of Aprataxin activity 155 4.2.12 DNA-adenylate hydrolase activity of PARP-1 and XRCC1 defective cell lines 155 4.2.13 PARP-1 knockdown 156 4.2.14 Impact of PARP inhibition on Aprataxin activity 157 4.2.15 Patch repair assay 158 4.2.16 Examination of Aprataxin activity in the brain 158

4.3 Results 160 4.3.1 Mutations in APTX and Aprataxin protein stability 160 4.3.2 Post-translational modification of Aprataxin 161 4.3.4 Characterization of endogenous Aprataxin- DNA binding 166 4.3.4 Characterization of endogenous Aprataxin- 5’ DNA adenylate hydrolase activity 167 4.3.5 Single strand break repair in AOA1 cell extracts 172 4.3.6 Oxidative Stress in AOA1 cells 173 4.3.7 Subcellular distribution of Aprataxin protein and activity 177 4.3.8 Regulation of Aprataxin by interacting proteins 182 4.3.9 Base excision repair in AOA1 cell extracts 188 4.3.10 Aprataxin activity in the brain 196

4.4 Discussion 198 4.4.1 Post translational modification of Aprataxin 198 4.4.2 Hydrolase activity of endogenous Aprataxin 198 4.4.3 Single strand break repair by AOA1 cell extracts 202 4.4.4 Oxidative DNA damage and base excision repair in AOA1 cells 203

xxi 4.4.5 Summary 205

4.5 References 207

CHAPTER 5 A role for Aprataxin in RNA biogenesis 215

5.1 Introduction 217 5.1.1 Structure and function of the nucleolus 217 5.1.2 Nucleolar transcription 228 5.1.3 Interaction between Aprataxin and nucleolin 219

5.2 Materials and Methods 221 5.2.1 Recruitment of proteins to dsDNA 221 5.2.2 GST pulldowns 222 5.2.3 UBF immunostaining 222 5.2.4 UBF immunoblotting 223 5.2.5 Nucleotide analogue incorporation protocols 224

4.3 Results 226 5.3.1 Aprataxin interacts with RNA transcription and processing factors 226 5.3.2 Aprataxin stabilizes UBF 235 5.3.3 Transcriptional defects in AOA1 cells 240

4.4 Discussion 263 5.4.1 Interaction of Aprataxin with transcription factors 263 5.4.2 Aprataxin dependant stabilization of UBF 264 5.4.3 Transcriptional defects in Aprataxin deficient cells 265 5.4.4 Summary 268

4.5 References 269 xxii

CHAPTER 6 General Discussion 277

6.1 Generation of reagents 279

6.2 Biochemical characterisation of Aprataxin 280 6.2.1 Aprataxin binds adenosine derivatives 280 6.2.2 Aprataxin hydrolyses adenosine derivatives 281 6.2.3 The HIT domain interacts with DNA 281 6.2.4 Aprataxin is a DNA-end processing factor 282 6.2.5 The HIT domain is regulated by the zinc finger and FHA domains 283 6.2.6 Aprataxin activity in cells is regulated by PARP-1 284

6.3 Characterisation of the defects in AOA1 cells 286 6.3.1 End processing 286 6.3.2 Base Excision Repair 288 6.3.3 Interaction of Aprataxin with transcription and RNA processing factors 291 6.3.4 Aprataxin-dependant stabilization of UBF 292 6.3.5 Aprataxin deficient cells have defective transcriptional responses to DNA damage 293

6.4 Future Directions 295

6.5 References 297

xxiii APPENDIX I

Kijas, A. W., Harris, J. L., Harris, J. M., Lavin, M. F. (2006). Aprataxin forms a discrete branch in the HIT (histidine triad) superfamily of proteins with both DNA/RNA binding and nucleotide hydrolase activities. The Journal of Biological Chemistry. 281(20):13939-48.

APPENDIX II

Janelle Harris, Burkhard Jakob, Keith W. Caldecott, Valérie Schreiber, Gisela Taucher-Scholz, Olivier J. Becherel and Martin F Lavin (2009). Multiple DNA repair defects in AOA1 reveal indirect roles for aprataxin in the DNA damage response. Human Molecular Genetics. Submitted and revision in progress.

LIST OF FIGURES CHAPTER 1 Figure 1.1: Activation of ATM by DNA damage. 5 Figure 1.2: Repair of abortive Topoisomerase I structures by Tdp1. 9 Figure 1.3: Aprataxin domain architecture and AOA1 mutations. 12 Figure 1.4: Alignment of Aprataxin and PNKP FHA domains. 14 Figure 1.5: Protein sequence alignment of HIT superfamily members. 16 Figure 1.6: Phylogenetic analysis of HIT proteins. 17 Figure 1.7: A model for the role of Aprataxin in single strand break repair. 20 Figure 1.8: DNA lesions and repair mechanisms. 26 Figure 1.9: Schematic of classical NHEJ. 28 Figure 1.10: Deficiency of XRCC1 inhibits alternative NHEJ. 29 Figure 1.11: Schematic of HR. 31 Figure 1.12: Effect of metal ions on induction of DNA breaks by hydrogen peroxide. 33 Figure 1.13: Schematic of direct SSBR. 35 Figure 1.14: Schematic of NER. 37 xxiv Figure 1.15: Schematic of short and long patch repair mechanisms. 40

CHAPTER 2 Figure 2.1: Schematic representation of the pTYB1 affinity purification system. 75 Figure 2.2: Test induction of wild-type and V263G Aprataxin fusion protein expression. 75 Figure 2.3: Cleavage efficiency of Aprataxin- Intein fusion proteins. 76 Figure 2.4. Affinity purification of wild-type and V263G recombinant Aprataxin from bacterial cell lysates. 78 Figure 2.5. Ion exchange purification of wild-type and V263G Aprataxin. 80 Figure 2.6. Quantification of recombinant Aprataxin concentrations. 81 Figure 2.7: Overview of pGEX 6.1 expression and purification system. 82 Figure 2.8: Schematic of pGEX 6.1-APTX truncation constructs. 83 Figure 2.9: Purification of recombinant Aprataxin using pGEX 6.1. 84 Figure 2.10: Purification of recombinant Aprataxin proteins. 85 Figure 2.11: Purified recombinant Aprataxin proteins. 86 Figure 2.12: Generation of crosslinked antigen-GST resin for antibody purification. 88 Figure 2.13: Purified Aprataxin antibodies. 89

CHAPTER 3 Figure 3.1: Partial Chymotryptic proteolysis of bacterial recombinant Aprataxin- (poly)nucleotide complexes. 108 Figure 3.2: Optimisation of chymotryptic proteolysis of yeast recombinant Aprataxin in the presence of DNA. 109 Figure 3.3: Comparison of Aprataxin conformational changes in the presence of various (poly)nucleotides. 110 Figure 3.4: Binding of recombinant Aprataxin proteins to dsDNA. 112 Figure 3.5: Binding of Aprataxin to different DNA structures. 114 Figure 3.6: Resolution of adenosine derivatives by strong anion exchange HPLC. 115

xxv Figure 3.7: Calibration of HPLC detector. 116

Figure 3.8: Generation of AMP from hydrolysis of AMPNH2. 120 Figure 3.9: Generation of AMP from hydrolysis of diadenosine tetraphosphate. 121 Figure 3.10. Inhibition of Aprataxins diadenosine tetraphosphate hydrolase activity by double stranded DNA. 123 Figure 3.11: Models of enzyme inhibition. 124 Figure 3.12. Impact of DNA on reaction kinetics of diadenosine tetraphosphate hydrolysis by Aprataxin. 126 Figure 3.13: Single strand break repair schematic. 128 Figure 3.14: Repair of 3’ damaged single strand breaks by cell extracts. 129 Figure 3.15: Alternative single strand break repair schematic. 130 Figure 3.16: Repair of 3’ damaged single strand breaks by T4 DNA ligase. 130 Figure 3.17: Schematic of Aprataxin cleavage of 5’adenylated DNA. 131 Figure 3.18: Aprataxin binding to 5’ adenylated DNA. 132 Figure 3.19: Hydrolysis of 5’ adenylated DNA by Aprataxin. 133 Figure 3.20: DNA-adenylate hydrolase activity of Aprataxin C and N-terminal truncation mutants. 134 Figure 3.21: Testing Aprataxin for phosphatase activity. 136 Figure 3.22: 3’ Phosphatase activity of control and AOA1 cell extracts. 136

CHAPTER 4 Figure 4.1: Schematic of the an vitro kinase assay. 149 Figure 4.2: Immunoblot of control and AOA1 lymphoblastiod cell lines. 160 Figure 4.3: Putative phosphorylation sites on Aprataxin. 162 Figure 4.4: Aprataxin is phosphorylated in vivo. 163 Figure 4.5: Tyrosine phosphorylation of Aprataxin in vivo. 164 Figure 4.6: Schematic of Aprataxin-GST fusion proteins. 165 Figure 4.7: Aprataxin is not an ATM kinase substrate. 165 Figure 4.8: Phosphorylation of Aprataxin by ATR. 166 xxvi Figure 4.9: Multiple alignment of Aprataxin sequences. 166 Figure 4.10: Endogenous Aprataxin binds DNA. 167 Figure 4.11. Schematic of Aprataxin cleavage of 5’adenylated DNA. 168 Figure 4.12: Hydrolysis of adenylated DNA by endogenous Aprataxin- titration with cell extracts. 169 Figure 4.13: Hydrolysis of adenylated DNA by endogenous Aprataxin – time-course. 170 Figure 4.14: Inhibition of DNA-adenylate hydrolysis by an Aprataxin specific antibody. 171 Figure 4.15: Schematic of double modified nick duplex. 172 Figure 4.16: Impact of adjacent 3’ modification on DNA-adenylate hydrolysis. 172 Figure 4.17: 3’ 8-oxo-dG inhibits single strand break repair. 173 Figure 4.18: Stability of APE1 in AOA1 cell lines. 174 Figure 4.19: Oxidative DNA damage in AOA1 cells. 176 Figure 4.20: Oxidative protein damage in AOA1 cells. 177 Figure 4.21: Cellular distribution of Aprataxin. 179 Figure 4.22: Cellular distribution of Aprataxin. 180 Figure 4.23: DNA-adenylate hydrolase activity of nucleoplasmic and nucleolar fractions. 181 Figure 4.24: XRCC1 deficiency does not affect Aprataxin activity. 183 Figure 4.25: PARP-1 is required for stabilization of Aprataxin. 183 Figure 4.26: Stability of Aprataxin protein after PARP-1 siRNA. 185 Figure 4.27: Lack of DNA adenylate hydrolase activity in PARP-1 knockout cells. 186 Figure 4.28: Inhibition of PARP activity by 3AB treatment. 187 Figure 4.29: PARP activity is dispensable for DNA-adenylate hydrolysis. 188 Figure 4.30: Schematic of in vitro patch repair assay. 189 Figure 4.31: Defective long patch repair in PARP-1 knockout cells. 190 Figure 4.32: Reduced long patch repair efficiency in AOA1 cells. 191 Figure 4.33: Quantification of long patch repair by control and AOA1 cells. 192

xxvii Figure 4.34: Quantification of long patch repair in control and AOA1 cells-quadruplicate experiments. 193 Figure 4.35: Patch repair by control and AOA1 cells- quantification of reaction intermediates and products. 194 Figure 4.36: Effect of recombinant Aprataxin on short patch repair. 195 Figure 4.37: Levels of PARP-1 in Aprataxin deficient and corrected cell lines. 196 Figure 4.38: Distribution of Aprataxin activity in the murine brain. 197 Figure 4.39: PARP-1 immunostaining in HeLa and MEF cells. 200 Figure 4.40. PARP-1 immunostaining in Hepa cells. 201 Figure 4.41: PARP-1 is required for appearance of XRCC1 nuclear foci at sites of oxidative DNA damage. 202 Figure 4.42: OGG1 and APE1 substrates, products and reaction kinetics. 205 Figure 4.43: Schematic representation of the indirect effects of Aprataxin deficiency. 206 CHAPTER 5 Figure 5.1: Subdomains of the nucleolus. 218 Figure 5.2: Aprataxin mediated recruitment of SFPQ to DNA. 229 Figure 5.3: Schematic of Aprataxin GST-fusion constructs. 231 Figure 5.4: Interaction of Aprataxin with RNA processing factors. 232 Figure 5.5: Domain structure of UBF proteins. 232 Figure 5.6: Polynucleotide-independent interaction between Aprataxin, UBF and hnRNP U. 233 Figure 5.7: Interaction of Aprataxin with RNA processing factors- phosphorylation dependence. 234 Figure 5.8: Protein complex assembly at the rDNA promoter. 235 Figure 5.9: UBF localization in HeLa cells. 236 Figure 5.10: Effect of Aprataxin deficiency on UBFs response to DNA damage. 237 Figure 5.11: Quantification of the effect of hydrogen peroxide on UBF staining in FD105 M20 and M21 fibroblasts. 238 Figure 5.12: Lack of Aprataxin destabilizes UBF. 239 Figure 5.13: Schematic of nucleotide analogue incorporation assay. 242 xxviii Figure 5.14. Optimization of fixation method. 243 Figure 5.15: Optimization of staining methods. 244 Figure 5.16: Optimization of incorporation conditions- BrUTP versus FUrd. 245 Figure 5.17: 5-Fluro Uridine incorporation by HeLa cells- co-localization with nucleophosmin. 246 Figure 5.18. Distribution of rRNA synthesis within the nucleolus. 247 Figure 5.19: FUrd incorporation by HeLa cells: optimization of antibody staining. 248 Figure 5.20: FUrd incorporation by HeLa cells- optimization of antibody dilution. 249 Figure 5.21: FUrd incorporation by HeLa cells- optimization of wash conditions. 250 Figure 5.22. Inhibition of RNA synthesis by Actinomycin D. 251 Figure 5.23: FUrd incorporation by HeLa cells- optimization chase time. 253 Figure 5.24: Effect of γ-irradiation on RNA synthesis in HeLa cells. 254 Figure 5.25: Quantitation of effect of γ-irradiation on RNA synthesis in HeLa cells. 255 Figure 5.26. Incorporation of 5’flurouridine by neonatal foreskin fibroblasts. 256 Figure 5.27: FUrd incorporation in FD105 M21 fibroblasts- optimization of labelling conditions. 257 Figure 5.28: Effect of Aprataxin deficiency on the transcriptional response to γ- irradiation. 258 Figure 5.29: Quantification of the effect of Aprataxin deficiency on the transcriptional response to γ-irradiation. 259 Figure 5.30: Effect of Aprataxin deficiency on the transcriptional response to oxidative stress. 260 Figure 5.31: Quantification the effect of Aprataxin deficiency on the transcriptional response to oxidative stress. 261 Figure 5.32: Distribution of nucleophosmin after DNA damage in AOA1 cells. 262 Figure 5.33: Model for UBF dimerization and DNA binding. 266

xxix LIST OF TABLES AND EQUATIONS

CHAPTER 1 Table 1.1: Frequency of symptoms in 14 AOA1 patients. 11 Table 1.2: Hydrolase activity of Aprataxin on nucleotide derivatives. 19 Equation 1.1. Reaction mechanism of DNA ligation. 21, 127 Table 1.3: Hydrolysis of 3’ substrates by Aprataxin. 22

CHAPTER 2 Table 2.1: Aprataxin pGEX 6.1 cloning primers. 68 Table 2.2: Summary of immunization and purification details for Aprataxin-specific antibodies. 87

CHAPTER 3 Table 3.1: Oligonucleotide sequences used to generate EMSA substrates. 100 Table 3.2: Oligonucleotides used to construct SSBR duplexes. 102 Table 3.3: Oligonucleotides used to generate 5’ adenylated DNA. 105 Table 3.4: Oligonucleotides used for phosphatase assays. 106

CHAPTER 4 Table 4.1: Oligonucleotides used to generate doubly modified nick structures. 151 Table 4.2: Oligonucleotides used to construct SSBR duplexes. 152 Table 4.3: PARP-1 siRNA sequences. 157 Table 4.4: Oligonucleotides used to generate patch repair substrate. 158

xxx LIST OF CHEMICALS AND SUPPLIERS

Chemical Abbreviation Supplier

2- glycerol phosphate Sigma 2-[4-(2-sulfoethyl)piperazin-1-yl]ethanesulfonic acid PIPES Sigma 2-mecaptoethanol Sigma 3-[(3-Cholamidopropyl)dimethylammonio]-1- propanesulfonate CHAPS Sigma 3-aminobenzamide 3-AB Sigma 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid HEPES Sigma 4,5,6,7-tetrabromobenzotriazole TBB Sigma 4',6-diamidino-2-phenylindole DAPI Molecular Probes 4-Morpholinepropanesulfonic acid,sodium salt MOPS Amresco 5-Bromo-4 Chloro-3-Indolyl-B-D-Galactopyranoside X-gal Sigma 5-fluorouridine FUrd Sigma Acetamide Sigma Acetic acid Merck Acetone Merck Acetonitrile Burdick & Jackson Acrylamide: bisacrylamide, 19:1 (40%) Biorad Acrylamide: bisacrylamide, 29:1 (30%) Biorad Adenosine Sigma Adenosine diphosphate ADP Sigma Adenosine monophosphate AMP Sigma Adenosine monophosphoramidate AMPNH2 Sigma Adenosine triphosphate ATP Sigma Agar BD Agarose Amresco Albumin, bovine fraction IV BSA Amresco Ammonium acetate Amresco Ammonium chloride Ajax Ammonium hydrogen carbonate Merck Ammonium persulfate APS Sigma Ammonium phosphate Sigma Ampicillin, sodium salt ICN Biorex-70 resin Biorad Boric acid Amresco Bradford reagent Biorad Bromodeoxy uridine BrdU Sigma Bromophenol blue ICN Bromo-uridine triphosphate BrUTP Sigma Butanol Fronine Calcium Chloride Sigma Chloroform Sigma Chymotrypsin Sigma Complete protease inhibitor Roche Coomassie brilliant blue R250 and G250 Sigma D-glucose Merck Diadenosine tetraphosphate AppppA Sigma Diethanolamine Sigma Dimethyl pimlimidate DMP Sigma Dimethyl Sulfoxide DMSO Amresco

xxxi Di-sodium hydrogen orthphosphate Ajax Dithiothreitol DTT Astral Dulbecco's Modified Eagle's Medium DMEM Gibco Dynabeads M-280 Streptavidin Dynal Enhanced Luminol Reagent Perkin Elmer Ethanol Ajax Ethanolamine Sigma Ethidium bromide EtBr Sigma Ethyl methanesulfonate EMS Sigma Ethylene glycol Sigma Ethylenediaminetetra-acetic acid EDTA Ajax Ficol 400 Sigma Foetal Calf Serum FCS Invitrogen Formalin Sigma Formamide Merck Fungizone Apothecon Geneticin/G418 G418 Invitrogen Scientifix/ Glutathione agarose Amersham Glycerol Ajax Glycine Ajax Hoechst 33342 Invitrogen Hybond C Amersham Hybond N+ Amersham Hydrochloric acid Ajax hydrogen peroxide Ajax Hydroxyirea HU Sigma Hygromycin Invitrogen Isoamyl alcohol BDH chemical Isopropanol Sigma Isopropylthiogalactopyranoside IPTG Sigma L-Glutamic acid, potassium salt Sigma Lipofectamine 2000 Invitrogen Lithium chloride Sigma Lysozyme, type VI MP Magnesium acetate Ajax Magnesium Chloride Ajax Magnesium Sulphate Ajax Manganese chloride Sigma Methanol Ajax Methoxymagnesium methyl carbonate MMC Sigma Methyl methanesulfonate MMS Sigma Mineral oil Sigma N,N,N', N'-tetramethydethylenediamine TEMED Sigma Newborn Calf Serum NBS Invitrogen Nitric acid Merck N-Lauroylsarcosine sodium salt Sarcosine Sigma Okadaic acid Sigma OptiMEM Gibco Orange G Merck Oxidizing Reagent Perkin Elmer PageRuler prestained protein ladder Fermentas Paraformaldehyde PFA Sigma Peptone BD

xxxii Phenol Sigma Polyoxyethylene (20) sorbitan monolaurate Tween-20 Amresco Ponceau S Sigma Potassium carbonate Sigma Potassium Chloride Sigma Potassium dihydrogen orthophosphate Sigma Potassium hydrogen orthophosphate Ajax Potassium hydroxide Sigma Potassium permanganate Ajax Protein A sepherose Amersham Protein G sepherose Amersham Puromycin Invitrogen Roswell Park Memorial Institute 1640 media RPMI-1640 Made internally Rubidium chloride Fluka Sephadex G-200 Pharmacia Sephadex G-50 Pharmacia Sepharose, fast flow Sigma Sodium acetate Merck Sodium Azide Sigma Sodium carbonate anhydrous Merck Sodium Chloride Sigma Sodium dodecyl sulphate SDS Amresco Sodium flouride Sigma Sodium hydrogen carbonate Ajax Sodium Hydroxide Merck Sodium nitrate Ajax Sodium pyrophosphate ICN Sodium sulphate Ajax Sodium sulphite Sigma Sodium vanidate Sigma Sucrose, from cane sugar Ajax Sulfuric acid Merck Triethanolamine Sigma Tris (hydroxymethyl) aminomethane hydrochloride Tris acid Sigma tris(hydroxymethyl)aminomethane Tris base Invitrogen Tri-sodium citrate Merck Trypsin, TC grade Sigma Urea, electrophoresis grade Sigma Water for irrigation Baxter X-ray film Kodak and Fujifilm Xylene cyanol FF Merck Yeast extract BD γ-P32 adenosine triphosphate (3000Ci/mmol) γP-32 ATP Amersham

xxxiii LIST OF ANTIBODIES AND SUPPLIERS

Primary antibodies Host Immunogen species Company/ Generated by Clone/Cat#

4355-MC- 8-oxo-G mouse Trevigen 100 AIF rabbit Cell Signalling 4642

Prof Grigory Dianov, University of Oxford, APE1 rabbit Oxford, UK

raised by Dr Amanda Kijas, Queensland Institute for Medical Research, Brisbane, Aprataxin rabbit Australia. Purified in chapter 2.

raised by Dr Amanda Kijas, Queensland Institute for Medical Research, Brisbane, Aprataxin sheep Australia. Purified in chapter 2. a-tubulin mouse Sigma 2-28-33 ATM mouse GeneTex 2CI

raised and purified by Dr Rick Woods (Queensland Institute for Medical Research, ATR rabbit Queensland, Australia b-actin mouse Sigma AC40 BrdU mouse Sigma B2531 BrdU rat Abcam ab6326 BrdU rat Abcam ab6326 BrdU mouse Becton Dickinson 347580 DNA-PK mouse Santa Cruz 18 2 EGFR mouse Santa Cruz sc-101

Dr Ulrich Sheer, Department of Cell and Developmental Biology, Biocenter, Fibrillarin rabbit University of Würzburg,

Dr Amila Suraweera, Queensland Institute GST rabbit for Medical Research, Brisbane, Australia. hnRNP U rabbit Abcam ab20666 Nitrotyrosine rabbit Cell Signaling 9691 nucleolin mouse MBP M019-3 nucleophosmin rabbit Cell Signaling 3542 p53 rabbit Cell Signaling 9282 PAR Mouse Bectin Dickinson 10H PARP-1 mouse Serotec MCA1522 pSer rabbit Cell Signalling 2981 pTyr mouse Cell Signalling 9411 xxxiv RNA polymerase II mouse Abcam ab5408

Dr Ingrid Grummt, Institute of Cell and Tumor Biology, German Cancer Research TAF95 rabbit Center, Heidelberg, Germany

Dr Ingrid Grummt, Institute of Cell and Tumor Biology, German Cancer Research TTF1 rabbit Center, Heidelberg, Germany UBF mouse Santa Cruz sc-13125 ubiquitin rabbit Abcam ab19247 XRCC1 sheep Bethyl A300-065A XRCC1 rabbit Serotec AHP428 XRCC4 mouse Novus 100-343

Secondary antibodies

Immunogen Host species Conjugate Company Clone/Cat#

rabbit IgG sheep HRP Chemicon AP322P mouse IgG sheep HRP Chemicon AP326P sheep IgG rabbit HRP Chemicon AP147B goat IgG rabbit HRP Chemicon AP106P

rabbit IgG donkey AlexaFluor-488 Invitrogen A21206 sheep IgG donkey AlexaFluor-594 Invitrogen A11016 sheep IgG donkey AlexaFluor-488 Invitrogen A11015 rabbit IgG donkey AlexaFluor-594 Invitrogen A21207 mouse IgG donkey AlexaFluor-594 Invitrogen A21203 mouse IgG donkey AlexaFluor-488 Invitrogen A21202 mouse IgG goat AlexaFluor-594 Invitrogen A11032 rat IgG goat AlexaFluor-488 Invitrogen A11006

Fluorophore Excitation λ (nm) Emission λ (nm) AlexaFluor-488 488 510 AlexaFluor-594 594 615 DAPI 345 455 Green Fluorescent Protein 488 510

xxxv Supplier Addresses Company Headquarters/Local distributor

Abcam Cambridge, UK Agilent Santa Clara, California, USA Ajax Taren Point, NSW, Australia American Type Culture Collection (ATCC) Manassas, Virginia, USA Amersham Biosciences Little Chalfont, Buckinghamshire, UK Amresco Solon, Ohio, USA Animal Resource Centre Canning Vale WA, Australia Apothecon Princeton, New Jersey, USA Astral Gymea NSW, Australia Baxter Deerfield, Illinois, USA BDH chemicals Carle Place, New York, USA Becton Dickinson North Ryde, NSW, Australia Bethyl Montgomery, Texas, USA Biorad Hercules, California, USA Cell Signalling Danvers, Maryland, USA Chemicon Temecula, USA Dynal Oslo, Norway Fluka now owned by Sigma-Aldrich Fronine Taren Point, NSW, Australia Fujifilm Minato, Tokyo, Japan Hewlett-Packard Palo Alto, California, USA ICN Costa Mesa, California, USA Invitrogen Carlsbad, California, USA Kodak Rochester, New York, USA MBP Gardenia, California, USA Merck Whitehouse Station, New Jersey, USA MP Solon, Ohio, USA New England Biolabs Ipswich, Maryland, USA Novus Littleton, Colorado, USA Pall Life Sciences East Hills, New York, USA Perkin Elmer Waltham, Massachusetts, USA Phenomonex Pennant Hills, Sydney, NSW, Australia Roche Indianapolis, Indianapolis, USA Santa Cruz Santa Cruz, California, USA Scientifix Clayton, Victoria, Australia Serotec Kidlington, Oxford, UK Sigma-Aldrich Castle Hill, NSW, Australia Trevigen Gaithersburg, Maryland, USA

xxxvi

COMMONLY USED ABBREVIATIONS

8-oxo-dG 8'-oxo-2'deoxy guanosine AOA1 ataxia with oculomotor apraxia type 1, synonymous with EAOH AOA2 ataxia with oculomotor apraxia type 2 APE1 apurinic endonuclease 1, synonymous with abasic endonuclease 1 ARCA autosomal recessive cerebellar ataxia A-T ataxia-telangiectasia A-TLD ataxia telangiectasia like disorder ATM ataxia telangiectasia mutated ATR ataxia telangiectasia related BER base excision repair C2ABR control lymphoblastiod cell line C3ABR control lymphoblastiod cell line Chk1 checkpoint kinase 1 Chk2 checkpoint kinase 2 CK2 caesin kinase II DBS double strand break DcpS scavenger mRNA-decapping enzyme DNA-PK DNA-dependant protein kinase DNA-PKcs DNA-dependant protein kinase catalytic subunit dRP deoxyribose phosphate DSBR double strand break repair EAOH early onset ataxia with hypoalbuminemia EMS ethyl methanesulfonate ERCC ethyl methanesulfonate repair complementing faPy 2,6-diamino-4-hydroxy-5-Nmethylformamidopyrimidine FD105 primary AOA1 patient cell line FD105 M20 immortalized AOA1 fibroblast cell line, transformed with empty vector immortalized AOA1 fibroblast cell line, transformed full length APTX FD105 M21 cDNA under constituative promoter FEN1 flap endonuclease 1- also known as endonuclease 1 FHA domain forkhead associated domain Fhit fragile HIT protein GalT galactose transferase GST glutathione-S-transferase Hint1 histidine triad nucleotide hydrolase 1 HIT domain histidine triad domain hnRNP U heterogeneous nuclear ribonucleoprotein U HR homologous recombination IR irradiation L938 AOA1 patient lymphoblastiod cell line (V263G, P206L) L939 AOA1 patient lymphoblastiod cell line (V263G,V263G) LCL lymphoblastiod cell line Lig 3 DNA ligase 3α

xxxvii LP BER long patch base excision repair MALDI- TOF Matrix Assisted Laser Desorption /Ionization- Time Of Flight MDC1 mediator of DNA damage checkpoint 1 MMC methylmercury chloride MMR mismatch repair MMS methyl methanesulfonate NEIL1 nei endonuclease VIII-like 1 NER nucleotide excision repair NFF neonatal foreskin fibrobloasts NHEJ nonhomologous end joining OGG1 8'-oxo-2'deoxy guanosine DNA glycosylase 1 PAGE polyacrylamide gel electrophoresis PAR polyADP ribose PARG polyADP ribose glycohydrolase PARP-1 polyADP ribose polymerase 1 PARP-2 polyADP ribose polymerase 2 PBS. phosphate buffered saline PCNA poliferating cell nuclear antigen PCR polymerase chain reaction PI3KK phosphatidylinositol 3-OH-kinase-related kinase PNKP 5' polynucleotide kinase 3' phosphatase Polβ DNA polymeraseβ RNAP 1 RNA polymerase I RPA replication protein A- also known as SSB RT room temperature SCA spinocerebellar ataxia SCAN1 spinocerebellat ataxia with axonal neuropathy SCID severe combined immunodeficiency SL1 selectivity factor 1 SP BER short patch base excision repair SSB single strand break SSBR single strand break repair TAE tris acetate EDTA buffer TAF95 TATA binding protein associated factor 95 TBE tris borate EDTA buffer TBS tris buffered saline Tdp1 tyrosyl-DNA phosphodiesterase TFIIH transcription factor II H Topo1 DNA topoisomerase 1 Topo2 DNA topoisomerase 2 TTF1 transcription termination factor 1 UBF1 upstream binding factor 1 UBF2 upstream binding factor 2 UCS upstream control sequence XP xeroderma pigmentosum XRCC1 X-ray cross complementing 1 XRCC4 X-ray cross complementing 4

xxxviii

xxxix

CHAPTER 1

Literature Review

1 2 1.1 AUTOSOMAL RECESSIVE CEREBELLAR ATAXIAS

Autosomal recessive cerebellar ataxias (ARCAs) are a group of rare neurological disorders which result in ataxia (a lack of muscle coordination) and a range of ophthalmological and neurological symptoms due to cerebellar dysfunction. There are many different types of ARCA, and a subset of these are caused by defective responses to DNA damage. These diseases include Ataxia Telangiectasia (A-T), Ataxia Telangiectasia Like Disorder (A-TLD), Xeroderma Pigmentosum (XP), Spinocerebellar Ataxia with Axonal Neuropathy Type 1 (SCAN1), and Ataxia with Oculomotor Apraxia Types 1 and 2 (AOA1 and AOA2). The focus of the present study is AOA1. The clinical aspects of AOA1 and related disorders will be introduced in the following sections, along with their genetic basis and our current understanding of their causal molecular and biochemical defects.

1.1.1 Ataxia Telangiectasia

1.1.1.1 Clinical presentation and genetic defect:

Ataxia Telangiectasia (A-T) is the best characterised DNA repair-defective ARCA (1,2). A-T has an early age of onset (normally between 2 and 4 years) and the initial phase of the disease is characterised by progressive cerebellar ataxia (3,4). Patients are normally wheelchair bound by the age of 10 due to worsening gait disturbances (4). Oculomotor abnormalities (difficulty controlling eye movements) are present in the majority of patients (5), and oculocutaneous telangiectasia (blood-shot eyes) appear between 2 and 8 years of age (1). The extra-neurological features of A-T include a predisposition to malignancy (especially leukaemia and lymphoma, reference 6) in addition to extreme sensitivity to many anti-cancer treatments including radiation (7,8) and mutagenic chemicals (9-11). This makes it difficult to effectively treat the malignancies frequently developed by A-T patients. A-T patients also display immunodeficiency (12). This, combined with their frequent inhalation of food and fluids (due to abnormal coordination of their breathing and swallowing cycles, personal communication with Dr Thomas Crawford, Neurologist, John Hopkins Children’s Research Hospital) leads to frequent potentially life-threatening respiratory infections (13). A-T is caused by mutation of the Ataxia-Telangiectasia

3 Mutated (ATM) gene, which is located at chromosomal locus 11q22 (14). This gene encodes the protein kinase ATM. Most A-T patients display destabilization of ATM, however some patients have intermediate to normal levels of protein with reduced activity (15). Thus A-T is caused by a deficiency of ATM kinase activity.

1.1.1.2 Function of deficient protein:

ATM belongs to a family of serine-threonine kinases which possess structural homology to yeast and mammalian phosphoinositide 3 (PI3) -kinases. This group is therefore referred to as the PI3-kinase-like-kinase (PI3KK) family (16). Cells from A- T patients display hypersensitivity to ionizing radiation (17,18) and are not arrested at the S phase checkpoint after irradiation (19), providing evidence of a role of ATM in DNA repair and cell cycle regulation. Further studies revealed that ATM is present as an inactive dimer in the nucleus and its kinase activity is activated by DNA double strand breaks (20). After induction of double strand breaks, ATM is activated by autophosphorylation on S367, S1983 and S1981 and becomes monomeric (20-22). This activation requires the MRN complex (consisting of the proteins Mre11, Rad50 and Nbs1 proteins) which senses and processes DNA damage. Activated monomeric ATM then initiates a number of cell-cycle control responses including initiation of G1/S, S and G2/M phase cell cycle blocks by phosphorylation of a number of cell- cycle regulators including p53 and Chk2 (references 20,23,24 and Figure 1.1, taken from 23).

4

Figure 1.1: Activation of ATM by DNA damage. ATM is activated by autophosphorylation which results conversion of the inactive dimer into active monomers. ATM then phosphorylates downstream targets involved in a range of regulatory processes. Proteins involved in ATM activation are shown in dark blue, in DNA repair in light blue, in apoptosis in light green, in the G1/S checkpoint in purple, in the intra-S checkpoint in yellow, in the G2/M checkpoint in dark green, in gene regulation in beige and proteins not belonging to any of these classes are shown in pink. Taken from (23).

1.1.2 Ataxia-Telangiectasia-Like Disorder

1.1.2.1 Clinical presentation and genetic defect:

The clinical presentation of Ataxia Telangiectasia-Like Disorder (A-TLD) displays similarities to the neurological symptoms of A-T. Patients display progressive cerebellar ataxia, peripheral neuropathy and oculomotor apraxia. A-TLD patients do not display telangiectasia, predisposition to malignancy or immunological deficits (25). A-TLD is caused by mutation of the MRE11A gene, at chromosomal locus

5 11q21 (26). Given the phenotypic similarities between A-T and A-TLD and the closeness of MRE11A to ATM, detailed genetic mapping was required to identify A- TLD as a separate disorder from A-T. MRE11A encodes the Mre11 protein. Attempts to generate an MRE11 knockout cells failed, indicating that Mre11 is an essential protein (27). Consistent with this, A-TLD patients express low levels of truncated or mutated Mre11 protein (26).

1.1.2.2 Function of deficient protein:

Similar to the situation with A-T, cell lines derived from A-TLD patients display hypersensitivity to ionizing radiation and cell-cycle checkpoint defects (26). Mre11 is a DNA processing factor involved in the repair of double strand breaks. Mre11 interacts with Rad50 (28) and Nbs1 (29) to form the MRN complex and deficiency of Mre11 (as in A-TLD) causes destabilization of the whole complex (26). The MRN complex is a sensor of double strand breaks and is involved in the maintenance of genomic integrity (30). MRN function is not dependant on ATM; indeed ATM activation is dependant on recruitment of the MRN complex to sites of DNA damage (Figure 1.1 and reference 31). This provides a functional explanation for the phenotypic similarities between A-T and A-TLD.

1.1.3 Xeroderma Pigmentosum

1.1.3.1 Clinical presentation and genetic defects:

The primary clinical manifestation of Xeroderma Pigmentosum (XP) is extreme photosensitivity. Patients are hypersensitive to UV irradiation and develop multiple cutaneous malignancies (32). Frequent clinical features also include skin pigmentation abnormalities, photophobia and conjunctivitis (32). XP patients display variable neurological symptoms which include varying degrees of ataxia, choreform (involuntary fidgeting) movements, spasticity and progressive mental retardation (33). XP is a syndrome with eight complementation groups and patient genotypes, designated XPA (XPA gene, 9q22.3, see (34)), XPB (ERCC3 gene, 2q21, see (35,36)), XPC (XPC gene, 3p25, see 37), XPD (ERCC2 gene, 19q13.2-q13.3, see 38), XPE

6 (DDB2 gene, 11p12-p11, see 39), XPF (ERCC4 gene, 16p13.3-p13.13, see 40), XPG (ERCC5 gene, 13q33, see 41) and XP variant (POLH gene, 6p21.1-p12, see 42).

1.1.3.2 Function of deficient proteins:

The genes mutated in XPA-G are involved in nucleotide excision repair (NER, reviewed in 43). This pathway is the principal mechanism for repair of a range of structurally unrelated DNA modifications such as UV-induced pyrimidine dimers as well and bulky chemical adducts (reviewed in 43). Cells derived from XPA-G patients are hypersensitive to UV radiation and display defects excising UV- photoproducts (43). The mechanism of NER will be detailed later in this review.

Unlike XPA-G, XP variant cells do not display NER defects (44). In addition to DNA repair pathways, cells have evolved a mechanism called trans-lesion synthesis (TLS) to tolerate nucleotide damage during replication (reviewed in 45). TLS requires low fidelity DNA polymerases which are able to synthesize nascent DNA using a damaged template. The product of the POLH gene, DNA polymerase η is able to synthesize nascent DNA using a template containing UV-induced pyrimidine dimers (45). This protein is deficient in XP variant patient cells. Thus XP variant is caused by a deficiency in the TLS pathway.

1.1.4 Ataxia with Oculomotor Apraxia Type 2

1.1.4.1 Clinical presentation and genetic cause:

Ataxia with Oculomotor Apraxia Type 2 (AOA2) patients display a similar clinical presentation to AOA1 patients. This disease is also referred to as non-Friedreich spinocerebellar ataxia type 1 (SCA1). AOA2 is a late-onset (11 to 22 years) progressive cerebellar ataxia (46,47). AOA2 patients display movement defects including gait ataxia and choreform movements as well as progressive peripheral sensory and motor neuropathy, with frequent speech (dysarthria) difficulties and oculomotor apraxia (47). AOA2 is not associated with mental impairment, or predisposition to malignant or infectious diseases. AOA2 is associated with elevated serum levels of γ-globulin, α-fetoprotein and creatin kinase and imaging studies have revealed progressive cerebellar degeneration in all patients (47). AOA2 is caused by

7 mutation of the SETX gene, located at 9q34 (46). This gene produces a primary transcript with an open reading frame of 8031 nucleotides which codes for the 2677 amino acid protein Senataxin (46). SETX mutations identified in AOA2 patients include point, frameshift and premature truncation and splicing mutations (46,48). These mutations normally lead to a deficiency of Senataxin protein in AOA2 patient cells (Dr Amila Suraweera, Queensland Institute for Medical Research, Brisbane, Australia, personal observations).

1.1.4.2 Function of defective protein:

The cellular function of Senataxin is not clearly understood. Cell lines from Senataxin-deficient AOA2 patients display hypersensitivity to single strand break inducing agents, elevated basal levels of oxidative DNA damage and elevated levels of hydrogen peroxide induced chromosome aberrations (49). They also display a defect in repair of hydrogen peroxide induced double strand breaks, indicating that Senataxin may protect cells from oxidative stress (49). Senataxin is homologous to the Saccharomyces cerevisiae DNA/RNA helicase Sen1p and recent evidence indicates a role for Senataxin in mRNA processing (Suraweera et. al; Senataxin, the homologue of yeast Sen1p, is involved in RNA metabolism; in preparation).

1.1.5 Spinocerebellar Ataxia with Axonal Neuropathy Type 1

1.1.5.1 Clinical presentation and genetic defect:

Spinocerebellar Ataxia with Axonal Neuropathy Type 1 (SCAN1) is an autosomal recessive spinocerebellar ataxia with an age of onset between 12 to 15 years of age. All patients display peripheral neuropathy, which results in the progressive loss of touch and pain sensation in the limbs (50). Patients with advanced disease develop an ataxic gait causing them to become wheelchair-dependant. Unlike many other ARCA disorders, oculomotor abnormalities have not been reported in SCAN1 patients. SCAN1 is caused by mutations of the TDP1 gene, which codes for the DNA repair protein Tyrosyl-DNA phosphodiesterase (Tdp1, reference 50).

8 1.1.5.2 Function of defective protein:

A potentially lethal DNA modification occurs when a DNA processing enzyme which acts via a covalent intermediate becomes ‘trapped’ at its site of action. This is the mode of cytotoxicity of the chemotherapeutic drug camptothecin, an inhibitor of Topoisomerase I. Camptothecin does not impair the DNA binding or backbone nicking activities of Topoisomerase I, but inhibits re-ligation, trapping Topoisomerase I to the 3’ terminus of a nick (51). Specific repair mechanisms exist which remove protein modifications from DNA. In vitro, Tdp1 repairs DNA-protein adducts which occur as a result of abortive Topoisomerase I reactions (52). This is consistent with the elevated levels of abortive Topoisomerase I complexes in SCAN1 cells and the hypersensitivity of SCAN1 patient cells to camptothecin (53). The proposed cellular function of Tdp1 in the repair of Topoisomerase I modified single strand breaks is shown in Figure 1.2.

Figure 1.2: Repair of abortive Topoisomerase I structures by Tdp1. Topoisomerase 1 creates transient single strand breaks and remains covalently attached to the 3’ break terminus. It is proposed that if resealing of this single strand break is not successful, the trapped protein-DNA complex is partially degraded by an unknown protease and Tdp1 hydrolyses the residual tyrosyl-DNA covalent bond to generate a 5’ phosphate. This can readily be repaired by DNA 3’ phosphatases (based on references 50,52).

9 1.1.7.1 Ataxia with Oculomotor Apraxia Type 1

1.1.7.1 Clinical presentation:

Ataxia with Oculomotor Apraxia Type 1 (AOA1) is an autosomal recessive spinocerebellar ataxia (54), which constitutes 5% of non-Friedreich progressive ARCA cases in a predominantly French population (55) and 21% of ARCA cases (including Friedreichs Ataxia) in a Portuguese population (56). It is synonymous with the disease Early Onset Ataxia with Hypoalbuminemia (EAOH, 57). While early studies reports that AOA1 patients develop gait ataxia between 2 and 6 years of age (54), more recent work reports a later onset (6.8 years, SD 4.8, reference 55). The gait ataxia observed in children with AOA1 has several characteristics which are typical of hereditary spinocerebellar ataxias (personal communication with Dr Thomas Crawford, Neurologist, John Hopkins Children’s Research Hospital). When standing at rest AOA1 children stand with their feet widely spaced and slowly gyrate their upper body. Their walking gait is characterised by uneven steps and may wobble side- to-side, while faster movement appears relatively normal. As the disease progresses patients develop limb dysmetria, which is the inability to accurately direct intentional movement in their limbs (54,55,58,59). This results in a progressive loss of limb control and leads to worsening of their gait and upper body control defects. As such all patients become wheelchair bound within 5 to 20 years after disease onset (55). In the early stages of AOA1 patients also display choreform movements and are unable to remain still when instructed to do so. In most instances this feature subsides as the disease progresses (55). Most AOA1 patients have absent or diminished deep tendon reflexes in either or both lower or upper limbs (54,55,58,59), indicative of peripheral motor neuropathy. Over the course of the disease children with AOA1 develop symmetric distal muscle weakness and wasting (58,59), perhaps as a result of their inability to make efficient use of their muscles. Peripheral sensory neuropathy causes some loss of sensation in patients limbs, normally in the more advanced stages of the condition (55,57,60). AOA1 patients can display a range of visual control defects including an impaired ability to perform both horizontal and vertical eye movements (54,55). A detailed opthalmological study by Le Ber et al. quantitated the visual defects in AOA1 patients and found that they take longer to change the focus of their visual attention and often ‘overshoot’ their targets (55). They also have difficulty accurately performing simultaneous movements of the eyes and head to track a

10 moving object and have prolonged saccades (jumping movements of the eyes needed to change visual focus, reference 55). Some patients also display masked facies (a reduced capacity to display facial expression) and/or dysarthria (54,55). Patients do not have a predisposition to cancer development or transmissible illnesses. Studies of the mental capacity of AOA1 patients have produced conflicting results: Aicardi et al. reported that their cohort of patients had a normal intelligence quotient range (54), while Le Ber et al. found mental impairment (either retardation or dysexecutive syndrome) in all patients examined (55). Laboratory studies have revealed motor and sensory axonal neuropathy, atrophy of the brainstem and cerebellum, hypoalbuminemia and hypercholesterolemia as pathological markers of AOA1 (55,56). Some features of AOA1 are variable and a summary of their frequency is shown in Table 1.1 (taken from Le Ber et al., reference 55).

Table 1.1: Frequency of symptoms in 14 AOA1 patients. Frequency is shown as a percentage and the number of patients examined for a particular trait is shown in brackets. Taken from Le Ber et al. (55).

11 1.1.7.2 Genetic cause:

AOA1 is caused by mutation of APTX, which is located on the long arm of chromosome 9 (9p13). Several transcripts are produced from this single gene (61); the longest and predominant transcript consists of 7 exons and 6 introns, with a coding region of 1026 nucleotides (61,62). This ‘full length’ transcript is expressed in the heart, brain, lymph nodes, liver, kidneys and spleen (61) and codes for the 342 amino acid protein Aprataxin (60,63). Disease-causing mutations include substitution, missense and premature truncation, splicing mutations as well as deletion of the whole coding region (Figure 1.3 and references 55,58-60,63-66). AOA1 causal mutations give rise to Aprataxin proteins which have reduced half-lives compared to wild-type in neuronal cells (67). The estimated population frequency of AOA1 is 1/200,000 live births in the Portuguese population (56). Based on this the allelic frequency of AOA1 causing mutations is approximately 1/450. The APTX mutations detected in AOA1 patients result in destabilization of the protein product, Aprataxin (67). To understand the biochemical deficiency which causes AOA1, the structure and function of the domains of Aprataxin will be explored.

Figure 1.3: Aprataxin domain architecture and AOA1 mutations. Adapted from Seidle et al. (66). Aprataxin is expressed from a seven exon gene, APTX, located on 9p13. A range of APTX mutations have been identified as causal for AOA1 and a selection are indicated. Such mutations include point mutations, premature termination codons and single nucleotide insertions and deletions, leading to frameshifts.

12 1.2 STRUCTURE AND FUNCTION OF APRATAXIN’S DOMAINS

Aprataxin is a 342 amino acid, 39.1kDa protein which contains three functional domains: an N-terminal Forkhead Associated (FHA) domain, a central Histidine

Triad (HIT) domain and a C-terminal C2H2 zinc finger. At the commencement of the present study the properties of Aprataxin’s domains were largely unknown. During the course of this study significant advances have been made in our understanding the function of Aprataxin’s domains. Here I will introduce separately the properties of each of Aprataxin’s functional domains, with a focus on their biochemical characteristics and links to DNA repair.

1.2.1 The FHA domain: FHA domains are phosphorylation dependant protein binding motifs, and are generally between 55 and 75 amino acids in length (68). These domains interact with serine and threonine phosphorylated proteins. FHA domains are found in all eukaryotes and in a wide range of protein types including transcription factors (69) and cell cycle control factors (70) and DNA repair proteins (71). The FHA domain of Aprataxin has homology to the FHA domain of 5’ polynucleotide kinase 3’ phosphatase (PKNP, Figure 1.4 and reference 60), a protein involved in the processing of DNA termini at single strand breaks (72). PKNP interacts, via its FHA domain, with the DNA single strand break repair scaffold protein X-Ray Cross- Complementing (XRCC)1 to form a repair complex consisting of PKNP, XRCC1, DNA ligase 3α (Lig 3α) and DNA polymerase β (Polβ, reference 73). The interaction of the PKNP FHA domain with XRCC1 is dependant on phosphorylation of XRCC1 and is abolished by inhibition of Casein Kinase II (CK2, reference 74). Based on the homology between the FHA domains of Aprataxin and PKNP, it was proposed that Aprataxin and PKNP may bind to the same region of XRCC1. It was subsequently shown that the FHA domain of Aprataxin, like PNKP, interacts with XRCC1 (75-78). This interaction is dependant on triple phosphorylation of XRCC1 on the CK2 sites pS518, pT519 and pT523 and the binding of Aprataxin and PNKP to XRCC1 is mutually exclusive (76). Deficiency of either Aprataxin or XRCC1 renders cells hypersensitive to the DNA-alkylating agent methyl methanesulfonate (MMS) providing a functional link between the two proteins (75-77).

13

Figure 1.4: Alignment of Aprataxin and PNKP FHA domains. Sequence numbers AAQ74130 and AAH02519. Generated in Clustal W using default settings (79).

Aprataxin also interacts with the double strand break repair scaffold protein XRCC4 (75). XRCC4 is involved in the repair of double strand breaks by non-homologous end joining (NHEJ, references 80,81). Analogous to the interaction between Aprataxin and XRCC1, the interaction between Aprataxin and XRCC4 has been shown to involve the FHA domain of Aprataxin and is also dependant on phosphorylation of CK2 sites on XRCC4 (75). The interaction of Aprataxin with both single and double strand break repair complexes indicates that Aprataxin may function in multiple pathways. This is substantiated by the sensitivity of AOA1 cells to agents which induce single (75-77) (low dose hydrogen peroxide and MMS) and double strand breaks (75) (γ-irradiation and high dose hydrogen peroxide), however the hypersensitivity of AOA1 cells to γ-irradiation is presently controversial. Luo et al. report mild hypersensitivity of patient fibroblasts by clonogenic survival assay (76), whereas Gueven et al. found normal sensitivity of patient-derived lymphoblastiod cell lines using trypan blue exclusion (77). To compound this confusion, I have observed delayed repair of a subset of γ-radiation induced double strand breaks in AOA1 MEFs (Becherel et. al; poster presentation at the Ataxia Telangiectasia Workshop 2008). Hopefully the availability of the APTX-/- mouse (generated in Peter McKinnon’s laboratory, St Jude Childrens Research Hospital, Memphis, USA) and human isogenic Aprataxin deficient and corrected cell lines (generated in Keith Caldecott’s laboratory, University of Sussex, Brighton, UK) will facilitate the standardization of cell-based assays, enabling the DNA repair community to develop a unified profile of the defects in AOA1.

More recently this laboratory has demonstrated an interaction between the FHA domain of Aprataxin and the nucleolar rRNA processing factor nucleolin (also

14 known as C23, reference 82). This interaction is dependant on phosphorylation of nucleolin by an unidentified kinase (82). Aprataxin is a nuclear protein with nucleolar and nucleoplasmic distribution (77,82). Given that Aprataxin does not contain a nucleolar targeting sequence, it seems likely that its localization to the nucleolus occurs indirectly via an FHA-domain interaction with a protein with a nucleolar targeting sequence like nucleolin. This was confirmed by transient depletion of nucleolin by siRNA, which blocked the accumulation of Aprataxin in nucleoli (82). The interaction of Aprataxin with nucleolin was ablated by inhibition of RNA polymerase I, indicating that it is dependant on active rRNA synthesis (82). A possible explanation for this is that nucleolin could be dephosphorylated as a result of transcriptional inhibition, which would prevent its interaction with Aprataxin. The authors also noted that AOA1 cell lines have less nucleolin than control cells and found that this is due to reduced stability of nucleolin in the absence of Aprataxin in vivo (82). Nucleolin is essential for processing of the pre- 47S rRNA (83), which ultimately generates the mature 18S, 5.8S and 28S rRNAs. Consistent with this Becherel et al. found that AOA1 cells have a mild defect in the early stages of pre-rRNA processing (82). In vitro nucleolin binds preferentially to the rDNA non-transcribed spacer (84), a region involved in initiation of rRNA synthesis. It also interacts with histone H1 to modulate chromatin structure (85). This indicates that although it does not interact with RNA polymerase I, nucleolin could be a modulator of rRNA transcription. Thus Aprataxin may have a role in rRNA synthesis and processing via stabilization of the rRNA transcription and processing factor nucleolin.

1.2.2 The HIT domain:

The HIT domain of Aprataxin has homology to members of the HIT protein superfamily, which includes the HINT, Fhit, DcpS, and GalT protein sub-families (86). This domain is characterized by the presence of the highly conserved HIT motif, HαHαHαα (where α is a hydrophobic amino acid) towards the C-terminus of the domain (shown in Figure 1.5, reference 87). Although the cellular function of most HIT domain containing proteins is unknown, these domains confer nucleotide hydrolase activity in vitro (86,88-90). Initial biochemical studies aimed at characterizing the substrate preference of Aprataxin’s HIT domain were based on

15 classification of Aprataxin as either a Hint (66) or Fhit type hydrolase (88). Sequence analysis of HIT superfamily members by this laboratory revealed that Aprataxin forms a discrete branch on the HIT superfamily tree (Figure 1.6 taken from Kijas et al., reference 91). However given that Hint1 and Fhit proteins have functional links with the maintenance of genomic stability (92-94), it seemed reasonable that Aprataxin may have a similar role on a related substrate.

Figure 1.5: Protein sequence alignment of HIT superfamily members. Protein sequences for human Aprataxin (AAQ74130), Hint (CAG33329), Fhit (ABM66093), GalT (AAB28328) and DcpS (Q96C86) proteins were obtained from the NCBI protein database and then aligned using Clustal X (79). The HIT motif is boxed.

16

Figure 1.6: Phylogenetic analysis of HIT proteins. Taken from Kijas et al., (91).

Fhit proteins are found in animals and fungi and all proteins which have been characterized biochemically possess diadenosine polyphosphate hydrolase activity in vitro and produce AMP as one of the two mononucleotide products (86). In vitro

Fhit hydrolyses its substrate (diadenosine polyphosphate, ApnA) at physiologically

relevant concentrations (Km =4.6 µM, reference 89, with estimated intracellular concentrations between 0.05 µM and 5 µM, reference 95), however the identity of the in vivo Fhit substrate has not been confirmed. Fhit was initially identified as a potential tumor suppressor by linkage analysis of a family with a history of early and severe renal carcinoma (normally a disease of the aged, references 96,97). Patients from this family were found to have a balanced translocation between the short arms of chromosomes 3 and 8, with a region on the short arm of chromosome 3 deleted. This deletion was found to disrupt expression of a 1.1kb transcript. This rather small transcript is produced from a large gene (over 1 Mb) which spans the most fragile site in the human genome (FRA3B), giving the protein product the name Fragile HIT (Fhit). Translocations and deletions in this region have subsequently been identified in a range of tumors including cancers of the gastrointestinal tract (94) and lung (93).

The HINT subfamily member Hint1 is an adenosine derivative hydrolase in vitro. This 14kDa homodimeric protein was initially described as an ADP hydrolase, albeit with an activity of 8.5/sec/M (98), 4x107 times lower than Fhit cleaving

17 AppppA (99). More recently, screening of a number of adenosine and inosine derivatives revealed that Hint1 hydrolyses adenosine monophosphoramidate

(AMPNH2) (2941176/sec/M) and adenosine monophosphate-N-ε-(N-α-Boc- lysinamide) (496000/sec/M) (90). The amide derivative structure of these molecules indicated that Hint1 may be involved in the hydrolysis of an AMP group from a protein (90). Many proteins are modified by adenylation or bind AMP covalently as part of their catalytic process (for example DNA kinases and ligases as well as protein kinases), thus Hint1 may be responsible for removal of some of these modifications. The in vivo function of Hint1 is not known however animal studies have revealed that this protein is a haplo-insufficient tumour suppressor (92).

At the commencement of this degree very little was known about the biochemical properties of Aprataxin’s domains. Given the homology of Aprataxin to Fhit and HINT family proteins and the possible roles of these proteins in the maintenance genomic stability, initial biochemical studies focused on characterization of the activity of Aprataxin on Fhit and HINT-type nucleotide substrates. Hirano et al. described the activity of Aprataxin’s HIT domain on the Fhit-type fluorescent substrate guanosine triphosphate-4,4-difluoro-4-bora-3a,4a-diaza-s-indacene (GpppBODIPY, reference 88). They determined that “WT aprataxin did not show significant activity”, however a truncated form of Aprataxin did. Aprataxin lacking the FHA domain displayed measurable GpppBODIPY hydrolase activity (although the authors do not present supporting catalytic parameters). This activity was further enhanced by the addition of the FHA domain separately (Vmax 0.00178/sec, Km 4.27 µM) which indicated possible regulation of HIT domain activity by the FHA domain. Although this study revealed an interesting relationship between the functional domains of Aprataxin, it was apparent that Fhit type substrates were not efficiently cleaved by Aprataxin (Fhit cleaves GpppBODIPY with Km 1.5 µM and Vmax 0.58/sec, reference 100).

Subsequent studies examined the activity of Aprataxin against different nucleotides. Seidle et al. (66) examined the activity of Aprataxin against several nucleotide based substrates (data reproduced in Table 1.2). It is notable that Seidle et al. found that Aprataxin has low but measurable hydrolase activity against GpppBODIPY

(Vmax 0.0004/sec, Km 13.1 µM) while Hirano et al. report no detectable activity

18 (88). The reaction conditions used are identical except for the use of a higher concentration of enzyme by Seidle et al. (12 nmole versus 40 pmol, both in 25 µL reactions) which probably accounts for their increased sensitivity. The use of very high concentrations of enzyme will result in elevated sensitivity however in this instance enzyme concentration (480 µM) exceeds the substrate concentrations used (2.5 to 250 µM). It seems likely that under these conditions the substrate is limiting and its concentration would change substantially over the reaction period. Kinetic experiments of this nature probably vastly underestimate the maximum velocity. The best substrate identified by this study, tert-butoxycarbonyl-(L)-lysine methylcoumarinamide (t-Boc-LysineAMP-MCA), is an analogue of adenylated lysine which is similar to some characterised Hint1 substrates. In this study Aprataxin demonstrated a higher turnover of this substrate than the fluorogenic molecule used by Hirano et al. (88), however the low hydrolase activities observed in both studies suggested that these substrates may not have biological relevance.

Table 1.2: Hydrolase activity of Aprataxin on nucleotide derivatives. The hydrolase activity of Aprataxin against a range of adenosine based molecules was determined by Seidle et al. (66). This table was taken from their publication.

In the same period our laboratory conducted a kinetic analysis of Aprataxin against the

adenosine derivatives AppppA and AMPNH2 (91). This study addressed the possibility that Aprataxin requires post-translational modification for activity by using recombinant Aprataxin expressed and purified from Saccharomyces cerevisiae as well as E.coli. This revealed that Aprataxin (from S.cerevisiae) hydrolyses both the Fhit

substrate AppppA (Vmax 0.0115/sec, Km 18 µM) and the Hint1 substrate AMPNH2

(Vmax 0.0195/sec, Km 837 µM). Aprataxin purified from E.coli hydrolysed AppppA with a similar maximum velocity but reduced affinity compared to the S.cerevisiae

protein (0.0171/sec, Km 27 µM). This indicated that substrate binding may be enhanced

19 by post-translational modification. I also attempted to address the findings of Hirano et al. (88) that Aprataxin’s hydrolase activity is stimulated by deletion of the FHA domain. Examination of the AppppA hydrolase activity of full length Aprataxin compared to an N-terminal truncation mutant (amino acids 118-342) revealed that deletion of the FHA domain did not have an effect on hydrolase activity (truncation

mutant Vmax 0.0176, Km 28 µM). I proposed that this may be due to the use of different substrates in these studies. I also examined the functional interaction between the HIT domain and the zinc finger of Aprataxin. I determined that double stranded DNA inhibits Aprataxin’s AppppA hydrolase activity and based on this proposed that Aprataxin binds DNA via its HIT domain in vivo and uses its hydrolase activity to repair modified DNA. Relating this with the characterised hypersensitivity of AOA1 cells to the Topoisomerase I inhibitor camptothecin (62), we hypothesized that Aprataxin has an analogous role to the 3’ processing factor Tdp1 and repairs abortive Topoisomerase I-DNA covalent structures (Figure 1.7).

Figure 1.7: A model for the role of Aprataxin in single strand break repair. Taken from Kijas et al. (91).

Subsequently Aprataxin was found to be a DNA end-processing factor, although on a 5’ substrate rather than the 3’ tyrosyl modification we had suggested. Aprataxin was reported to hydrolyse a DNA repair intermediate, 5’ adenylated DNA (101). This

20 molecule is a covalent intermediate of the ligation process (102). The mechanism of DNA ligation entails three steps, as shown in Equation 1.1. 1) Initially the ligase hydrolyses ATP to form a ligase-AMP covalent complex. 2) Following adenylation of the enzyme, the adenylate group is transferred from the ligase onto the 5’phosphate DNA terminus, generating a 5’DNA-adenylate (5’-AMP-DNA). 3) After DNA adenylation, the phosphodiester bond is restored and AMP released. A 3’hydroxyl group is absolutely required for end-joining (step 3) but is not required for the enzyme and DNA adenylation phases of the reaction mechanism (steps 1 and 2). In the absence of a 3’hydroxyl the end-joining reaction is inhibited, resulting in an accumulation of 5’DNA adenylate (101,103,104). Ahel et al. reported that Aprataxin hydrolyses this molecule, although kinetic parameters were not determined (101). Significantly they found that wild-type cell extracts are able to hydrolyse adenylated DNA, whereas AOA1 cell extracts are not. This indicated that Aprataxin is the only protein in a mammalian cell capable of repairing this DNA modification. The authors also found that 5’ adenylate modifications accumulate when DNA is treated with hydrogen peroxide before being subjected to an in vitro repair reaction. This provided a biochemical link between the newly characterised Aprataxin substrate and the established hypersensitivity of AOA1 cells to hydrogen peroxide. Following this the same laboratory reported that mutation of the zinc finger drastically reduces the DNA- adenylate hydrolase activity of Aprataxin (to approximately 1%), presumably by impairing the ability of Aprataxin to interact with this substrate (103).

Equation 1.1. Reaction mechanism of DNA ligation. Various chemical species are differentiated by colour throughout the reaction set. Based on (105).

More recently, Takahasi et al. reported that Aprataxin possesses 3’ repair activities (106). Aprataxin was reported to repair both 3’ phosphate and 3’ phosphoglycolate damaged termini (106). Reported kinetic parameters for repair of these termini by Aprataxin as well as the established 3’ phosphatase PNKP and the 3’ phosphoglycohydrolase Apurinic Endonuclease 1 (APE1) are shown in Table 1.3

21 (taken from Takahasi et al., reference 106). These figures indicate that Aprataxin repairs 3’ phosphoglycolate more efficiently than APE1 does. Comparatively, Winters et al. found that recombinant APE1 hydrolyses 3’ phosphoglycolate with a Vmax of

0.059/sec (calculated from their reported value of 0.1 fmol/min/pg of enzyme) and a Km of 50 nM (107). These parameters indicate that APE1 is capable of much more efficient phosphoglycohydrolase activity than Takahasi et al. observed for either APE1 or Aprataxin. Based on this I conclude that Aprataxin is not an efficient 3’ phosphoglycohydrolase and that the APE1 purified by Takahasi et al. may be largely inactive. Additionally Takahasi et al. reported that Aprataxin has approximately 18% the 3’ phosphate repair efficiency of PNKP (based on the Km /Vmax ratios for each enzyme). However the activity of PNKP reported by Takahasi et al. is much lower than

the activity reported by others (0.466/sec, Km 16 nM, reference 108). Based on the level of PNKP activity reported by Wiederhold et al. (108), Aprataxin is not likely to make a major contribution to the repair of 3’ phosphate modifications in a cell. The reaction kinetics presented in this publication indicate that Aprataxin hydrolyses 3’ phosphate and phosphoglycolate with even lower efficiency than early reports found for Aprataxin’s hydrolysis of small nucleotide based substrates (66,88,91). Additionally, other laboratories have been unable to detect 3’ phosphatase or 3’ phosphoglycolate hydrolase activities in recombinant Aprataxin (personal communications, Prof Steven West, Cancer Research UK, London Research Institute, UK and Prof Keith Caldecott, University of Sussex, Brighton, UK), making this aspect of Aprataxin activity controversial.

Table 1.3: Hydrolysis of 3’ substrates by Aprataxin. Taken from Takahasi et al. (106).

22 1.2.3 The Zinc Finger:

Aprataxin possesses a C-terminal zinc finger of the C2H2 type. C2H2 zinc fingers are ubiquitous domains which are found in many proteins. It is the most prevalent motif in the mammalian proteome and was first identified in transcription factor IIIA (TFIIIA) as a sequence specific DNA binding motif which is stabilized by chelation

of a zinc ion (109). Many C2H2 zinc finger containing proteins have RNA binding capacity (110-112) and this motif has also been implicated in protein-protein

interactions (113). Thus the C2H2 zinc finger appears to be a very versatile motif capable of mediating protein-DNA, protein-RNA and protein-protein interactions.

Initial characterization of the DNA and RNA binding capacity of Aprataxin was provided by this laboratory in 2006 (91). We demonstrated that Aprataxin binds to double stranded DNA and RNA with similar affinities. Transient binding to single stranded DNA was also observed. An N-terminal truncated form of Aprataxin, which lacks the FHA domain is able to bind DNA with a similar affinity as the full length protein, indicating that the FHA domain is dispensable for DNA binding. Full length Aprataxin expressed and purified from either E.coli or S.cerevisiae is able to bind double stranded DNA, indicating that post-translational modification is not required for this activity.

Following the identification of a high efficiency substrate for the Aprataxin HIT domain (adenylated DNA), Rass et al. examined the binding of Aprataxin to this modified DNA molecule (103). They found that Aprataxin is able to bind double stranded DNA which is adenylated either at an internal nick or at a blunt end. Additionally they reported that Aprataxin can stably interact with adenylated single stranded DNA, indicating that 5’ adenylate stabilizes the transient interaction observed by Kijas et al. (91). Using poly[dI٠dC] to compete with adenylated DNA for Aprataxin binding, the authors found that (catalytically inactive) Aprataxin interacts with adenylated DNA in a more stable manner than with unmodified duplex. Similar to the model suggested by Kijas et al. (91), Rass et al. proposed a model where Aprataxin scans the genome by interacting with duplex DNA in a transient manner before encountering its substrate, 5’ adenylated DNA, where it forms a stable repair complex (103).

23 At the commencement of this study the interaction of Aprataxin with DNA repair complexes, its possession of DNA binding and nucleotide hydrolysis domains and the hypersensitivity of AOA1 patient cells to genotoxic agents indicated a role for Aprataxin in DNA repair. This was substantiated by the identification of the repair intermediate 5’ adenylated DNA as a high efficiency Aprataxin substrate. Aprataxin was reported to be the only enzyme capable of hydrolysing this modification, so it follows that AOA1 cells (which have no Aprataxin) should accumulate adenylated DNA. There is strong unpublished evidence that this is the case. Aprataxin/ Tdp1 double knockout cells accumulate more breaks than Tdp1 single knockouts. Addition of recombinant Aprataxin to genomic DNA from the treated double knockout cells but not the single knockout results in release of AMP, indicating that these excess breaks are adenylated (Prof Keith Caldecott, University of Sussex, Brighton, UK, personal communication and oral presentation at Ataxia Telangiectasia Workshop 2008). This implicates Aprataxin as a DNA-adenylate hydrolase whose function is proof-reading the genome for stalled ligation structures. Ligation is a critical process in all DNA repair pathways, so these mechanisms will be introduced with a focus on proteins known to interact with Aprataxin.

1.3 DNA REPAIR MECHANISMS The genomic stability of living cells is threatened by the DNA damaging effects of endogenous free-radicals and environmental genotoxins. Endogenously generated damaging agents include reactive oxygen species such as superoxide and hydroxyl radicals, which are generated as a result of mitochondrial respiration (114). DNA double strand breaks are also generated during V(D)J recombination, which is essential for diversification of the immune system. Exogenous sources of genotoxic stress include UV and ionizing radiation. Both endogenous and exogenous sources of oxidative stress can damage DNA directly or indirectly in a replication-dependant manner by oxidation of the dNTP pool (115). DNA single and double strand breaks are also generated by normal cellular processes. Controlled generation and repair of double strand breaks is essential for normal development of the immune system, facilitating antibody diversification of the by recombination of immunoglobulin genes (116). Transient single strand breaks are generated by Topoisomerase I, which relieves topological stress on DNA created by moving replication or transcription forks (117). Topoisomerase II generates controlled double strand breaks at DNA crossover points to

24 facilitate chromosome segregation (117). As introduced previously, defects in DNA repair can have a wide range of detrimental effects including development of malignancies and neurodegeneration.

DNA repair mechanisms can be simplistically classified into two major pathways, single and double strand break repair. Regardless of the nature of the lesion DNA repair is a highly ordered, sequential process. Initially the lesion must be detected by sensor proteins which recruit repair complexes. These proteins then process the lesion to generate a break with ligatable termini, which is then sealed by a DNA ligase. Depending on the nature of the initial lesion, and to some extent the nature of the surrounding DNA (for example transcribed or not, hypermethylated or not), repair pathways can have elaborate and sometimes interlinking mechanisms. For example, damage to transcribed regions of DNA activate mechanisms which will repair these lesions preferentially (termed transcription coupled repair, TCR). Additionally, some DNA lesion types have the potential to be converted into double strand breaks by disruption of DNA replication forks and subsequent fork collapse. Such lesions include DNA single strand interruptions and nucleotide damage. When examining the repair or toxicity of such lesions, the possibility of conversion into double strand breaks via replication fork collapse should always be considered. A summary of some agents which produce DNA damage, the lesions they cause and the pathways responsible for their repair is shown in Figure 1.8.

25

Figure 1.8: DNA lesions and repair mechanisms. From de Boer et al. (43).

Previous studies established that Aprataxin deficient cells are hypersensitive to agents which induce single strand breaks (camptothecin, MMS and hydrogen peroxide, references 75-77,118). Some studies also found hypersensitivity to double strand break inducing agents (irradiation and MMC, reference 75) however not all laboratories have been unable to form a consensus on this. The observed intermediate hypersensitivity of AOA1 cells to such agents could be attributable to the abundance of single strand breaks these agents also produce (119). Regardless of this, Rass et al. found that Aprataxin efficiently repairs adenylate modifications at single and double strand breaks as well as single stranded termini (103), indicating that it has the potential to be involved in multiple DNA repair pathways. This idea is substantiated by the interaction of Aprataxin with proteins involved in both single and double strand break repair pathways (75,77,78,118). To facilitate an understanding of the potential role of Aprataxin in DNA repair, an overview of single and double strand break repair and their subpathways will be presented.

1.3.1 Double strand break repair: Double strand breaks can be generated by ionizing radiation (by direct damage to DNA or by radiolysis of water and free-radical generation, reference 119), some radiomimetic chemicals (120) or indirectly by collapse of replication forks blocked by

26 nucleotide/base damage or single strand breaks (121). The ability to accurately repair double strand breaks is a fundamental requirement for maintenance of genomic integrity. Defects in double strand break repair result in a number of genome instability disorders, some of which have been described here (sections 1.1.1 and 1.1.2). DNA double strand break repair can be divided into major two sub-pathways: Non- Homologous End Joining (NHEJ) and Homologous Recombination (HR). These pathways use different repair complexes, have different levels of repair fidelity and occur preferentially at different stages of the cell cycle.

1.3.1.1 NHEJ: NHEJ is the primary method of double strand break repair in non-proliferating or G1- phase cells (122). Classical NHEJ is initiated by the DNA-end binding heterodimer Ku70:80, which detects the break. These proteins align the broken ends (123) and together with scaffold protein XRCC4 stabilize the interaction of the PI3KK DNA- dependant Protein Kinase catalytic subunit (DNA-PKcs) with the termini (124). It is presently unclear which protein (DNA-PKcs or XRCC4) arrives at the break first. The interaction of DNA-PKcs with the Ku heterodimer and its subsequent stabilization at the break results in activation its serine/threonine protein kinase activity, with targets including XRCC4 (124,125). Double strand breaks often have overhanging or damaged termini which need to be processed by repair factors before ligation. This processing can include the removal of several nucleotides by nucleases such as Artemis and WRN (126,127), and as such NHEJ often results in microdeletions. Artemis is phosphorylated by the DNA damage inducible protein kinase ATM, and this phosphorylation is required for end-processing by Artemis. This links the signalling functions of ATM with control of DNA repair mechanisms. After processing the DNA termini are resealed by the XRCC4 interacting protein DNA ligase IV. An overview of classical NHEJ is shown in Figure 1.9.

Ligation attempts before nuclease processing is complete have the potential to cause 5’ adenylation. Aprataxin interacts with XRCC4 indicating a potential role for it in repair of abortive ligation products at sites of NHEJ (75).

27

Figure 1.9: Schematic of classical NHEJ. DNA double strand breaks are detected by the Ku heterodimer. This results in recruitment of DNA-PKcs and the XRCC4/ DNA ligase 4 complex. DNA ends are often processed by nucleases to generate blunt termini (not shown here for the sake of simplicity), which are ligated by DNA ligase 4. Summarized from (123,124). Mutation or deficiency of classical NHEJ proteins has serious consequences in mammals. Mutation of DNA-PKcs causes defective V(D)J recombination and arrested B- and T- lymphocyte development in severe combined immune-deficient (SCID) mice (128). Strangely this defect can be partially rescued by treatment of the animals with radiation or radiomimetic drugs, possibly by activation of alternative double strand break repair mechanisms (129,130). Deletion of PARP-1 (which has many roles including suppression of recombination) rescues lymphocyte development in the DNA-

28 PKcs deficient SCID mouse (131). This indicates that elevated levels of recombination due to loss of PARP-1 aid V(D)J recombination in the absence of DNA-PK activity (131,132). DNA-PKcs mutant and PARP-1 knockout SCID mice do not display a predisposition to tumour formation, however double mutant mice develop a high frequency of lymphoma (131). This indicates that DNA-PKcs and PARP-1 may function in parallel pathways.

Consistent with this idea, an alternative NHEJ pathway involving PARP-1, XRCC1, and DNA ligase 3α has been proposed (133-136). The well-characterised functions of these proteins in the repair of single strand breaks will be addressed in subsequent sections. Recombinant PARP-1, XRCC1 and DNA ligase 3α are able to repair double strand breaks with blunt or 5’ or 3’ recessed termini in vitro (133). Exclusion of any of these proteins abolished re-ligation activity indicating that the three proteins function in a concerted manner. PARP-1 and XRCC1 deficiency impairs ligation of the double strand breaks in vitro (Audebert et al., reference 133) and Figure 1.10 below, taken from Harris et al.; Synergistic function of Aprataxin and PARP-1 in DNA repair; in preparation), indicating the PARP-1/XRCC1/ DNA ligase 3α complex facilitates an alternative NHEJ mechanism. Given that this alternative NHEJ pathway requires PARP-1 and that V(D)J recombination in the DNA-PK mutant SCID mouse is corrected by PARP-1 deficiency (131), the observed correction must occur via another mechanism. As immature lymphocytes are highly proliferative this correction could occur via homology-mediated repair.

Figure 1.10: Deficiency of XRCC1 inhibits alternative NHEJ. Nuclear extracts from wild-type (AA8) and XRCC1 mutant (EM9) CHO cells were incubated with a radiolabelled 36nt duplex with a 4nt palindromic 3’ overhang for the indicated times. Reaction products were analyzed by denaturing gel electrophoresis. From Harris et al.; Synergistic function of Aprataxin and PARP-1 in DNA repair; in preparation.

29 1.3.1.2 HR: Homologous recombination is the predominant double strand break repair mechanism for cells in late S to G2 phase (122,137) and also provides a mechanism for telomere length maintenance in cells lacking telomerase (138). Activation of this pathway coincides with induction of ATM-dependant G1/S, S or G2/M cell cycle blocks until the break is resolved (139). ATM defects impair HR mediated DNA repair (140). HR defects can result in neurodegeneration or a predisposition to cancer, indicating a critical link between ATM kinase activity, cell cycle control and HR. A-TLD and Nijmegen Breakage Syndrome (NBS) are characterised by ataxia without cancer predisposition and cancer predisposition without ataxia respectively. They are caused by mutation of MRE11(A-TLD) and NBS (NBS) genes, which code for the Mre11 and Nbs1 proteins. Mre11 and Nbs1 interact with a third protein, Rad50, to form the MRN complex. This complex has a critical role in the initial phases of HR and is essential for the activation of ATM after DNA damage (141).

Recombinational repair resolves double strand breaks without introducing errors, as NHEJ can (137). Rad52 and Rad51 bind rapidly to double strand breaks, facilitating recruitment of Rad51-family proteins (including XRCC2, XRCC3, Rad51B, C and D, references 142,143). The MRN complex is subsequently recruited, resulting in resection of both termini by Mre11 and activation of ATM (141,144). After strand resection and ATM-dependant cell-cycle blockage, a region of DNA homologous to the single stranded ends is located. Subsequent assembly of the damaged strands with undamaged template is facilitated by Rad51, which has the ability to exchange a single stranded region of DNA for the same sequence in a duplex (termed strand invasion, reference 145). Nascent DNA is then synthesized using the homologous strands as templates and the non-resected strands as primers (137). After synthesis the repair structure is disassembled by resolvases and the re-sealed by DNA ligases (137). A schematic of this process is shown in Figure 1.11.

30 Figure 1.11: Schematic of HR. DNA double strand breaks are detected by Rad52, which facilitates binding of Rad51 (and Rad51 family proteins, not shown here for simplicity). 3’-5’ resection of the damaged termini is carried out by the MRN complex member Mre11. Strand invasion is facilitated by Rad51, which exchanges homologous single and double stranded DNA strands. This results in generation of a repair bubble where nascent DNA is synthesized by replicative DNA polymerases. After resolution of the repair complex, the newly synthesized complimentary regions anneal and are ligated. Summarized from (146).

1.3.2 Single strand break repair pathways:

Single strand breaks are abundant potentially deleterious lesions. Single strand breaks can arise by disintegration of the phosphodiester backbone or through processing of DNA lesions. Such indirect breaks are generated by enzymatic mechanisms responsible for repairing oxidized bases, abasic sites, mismatches, bulky DNA- adducts, and strand crosslinks. Unrepaired single strand breaks can be converted to double strand breaks in S phase by causing the stalling and subsequent collapse of replication forks (121). This is the mode of toxicity of the chemotherapeutic drug

31 camptothecin. Single strand breaks in coding regions have the potential to harm non- proliferating cells by causing the production of mutant transcripts or blocking transcription of a damaged coding region all together (147,148). Similar to the situation with the repair of double strand breaks, cells have developed an array of single strand break repair mechanisms to deal with different lesion structures. Single strand break repair can be classified into four pathways: direct single strand break repair (SSBR), Nucleotide Excision Repair (NER), MisMatch Repair (MMR) and Base Excision Repair (BER).

1.3.2.1 Direct Single Strand Break Repair

Direct single strand breaks arise by spontaneous (149) or damage induced disintegration of deoxyribose (150). Sugar damage can be the result of attack by free radicals including peroxide and superoxide, which are generated during mitochondrial respiration and by exogenous sources of stress (reviewed in 151). Oxidative DNA damage is mediated by divalent cations, particularly transition metals (Figure 1.12 and references 149,150). Such breaks often possess a single nucleotide gap with a monophosphate (70%) or phosphoglycolate (30%) 3’ terminus (151).

32

Figure 1.12: Effect of metal ions on induction of DNA breaks by hydrogen peroxide. Supercoiled plasmid DNA was incubated with hydrogen peroxide in the presence of 50 µM transition metal salts, as indicated. Single stranded breaks convert the supercoiled substrate into the open circular conformation. Double strand breaks convert it into a linear strand. Band identities are indicated. Taken from Kobayashi et al., (150).

These modified single strand gaps recruit PARP-1, a multi-functional DNA repair protein (152,153). 17 PARP proteins have been identified based on homology searching of the human genome database (154). PARP family proteins transfer ADP- ribose units to the γ-carboxyl group of specific lysine or aspartic acid residues on acceptor proteins using NAD+ as a cofactor (155-158). The resulting polymers form polyanionic linear or multiply branched polyADP ribose (PAR) chains (159). Two out of these 17 proteins (PARP-1 and PARP-2) have characterised roles in DNA repair (160-162). PARP-1 is the most abundant of the PARP proteins. Its catalytic activity is enhanced up to 500 fold by DNA breaks (163) and accounts for at least 85% of DNA damage induced PAR synthesis, with the remainder attributable to PARP-2 (164). PARP-1 itself is the predominant target for its own catalytic activity (165). Other targets for PARP-1 activity include histones (166-168), DNA topoisomerases I and II (169,170) and DNA helicases (171). PARP-1 automodification leads to an

33 accumulation of negative charge at the break site causing disassociation of PARP-1 from the break and subsequent inhibition of catalytic activity (172,173). The accumulation of negative charge at breaks protects damaged termini from unwanted recombination (50, 51) and serves as a recruitment signal for repair factors including XRCC1 (174). There is evidence that PARP-2 functions in DNA single strand break repair and may be able to partially compensate for PARP-1 deficiency (161).

Direct single strand break repair is initiated by detection of the break by PARP-1. Accumulation of PAR at the break recruits XRCC1 (174), a scaffold protein which serves as an assembly platform for discrete repair complexes containing a number of proteins including PNKP, DNA ligase 3α, DNA polymerase β, APE1 and Aprataxin (73,75,76,175). DNA ligase 3α and DNA polymerase β appear to be present in all XRCC1 complexes, and other factors are variable. Recruitment of multiple XRCC1- repair factor complexes to direct single strand breaks facilitates rapid sequential processing and repair. The 3’ phosphate termini frequently present at direct single strand breaks are repaired primarily by PNKP, restoring the 3’ hydroxyl terminus (176). APE1 also has well characterised 3’ phosphatase activity (177). The phosphatase activities of both APE1 and PNKP are stimulated by XRCC1 in vitro, indicating that XRCC1 enhances their repair activities in addition to concentrating them at break sites (175,178). Termini of the alternative structure, 3’ phosphoglycolate, are processed to generate a 3’ hydroxyl by APE1 (179) and Tdp1 (180). The resulting 3’hydroxyl 5’phosphate gap is filled by DNA polymerase β to generate a single stranded nick (181,182), which is subsequently ligated by DNA ligase 3α (Figure 1.13 and reference 183).

34

Figure 1.13: Schematic of direct SSBR. Single strand breaks are detected by PARP- 1, resulting in localized PAR synthesis. This recruits XRCC1 repair complexes which contain DNA ligase 3α and DNA polymerase β in addition to one of a number of repair factors such as Aprataxin or PNKP. This facilitates processing of damaged termini. After end processing the resulting 5’ phosphate 3’ hydroxyl single nucleotide gap is filled by DNA polymerase β producing a nick, which is ligated by DNA ligase 3α. Note that if the 5’ break terminus was unable to be processed to generate a 5’ phosphate, this pathway can link with an indirect 5’ repair mechanism (long patch BER, described in a future section).

35 Given the interaction of Aprataxin with the XRCC1 complex and the hypersensitivity of AOA1 cells to single strand break inducing agents it seems likely that Aprataxin, similar to APE1 and PNKP, functions as an accessory factor in direct SSBR. If termini are not processed before DNA ligation is attempted the 5’ break terminus may become adenylated, requiring hydrolysis by Aprataxin before repair can be completed.

1.3.2.2 Indirect SSBR (Nucleotide Excision Repair)

Indirect single strand break repair covers three distinct pathways- Nucleotide Excision Repair (NER), Base Excision Repair (BER) and Mismatch Repair (MMR). NER is responsible for the repair of a variety of lesions including UV-induced pyrimidine dimers, intra-strand crosslinks, DNA-protein crosslinks and a range of chemical adducts (184). This versatile mechanism does not rely on detection of each specific modification- rather it senses distortions of DNA structure (184). A schematic of NER is shown in Figure 1.14. The simplest model of NER involves detection of DNA distortion by the XPC or XPE complexs (consisting of XPC or XPE, Cen2, and hHR23B, reference 185) and subsequent recruitment of repair factors including the multi-protein transcription factor complex TFIIH (186). TFIIH unwinds distorted region generating a denatured bubble (187). At this point the system needs to locate the actual lesion, and in particular identify which strand is damaged. This may be performed by XPA, which has a high affinity for modified single stranded DNA (188,189). Once the damage is located the damaged strand is incised by XPG (3’ cut) and the ERCC1/XPF hetrotrimer (5’ cut), generating an approximately 30 nucleotide single stranded region (187). This is then filled in by the replicative polymerases and ligated by DNA ligase 1 or 3α (184). Bulky nucleotide damage in transcriptionally active regions is initially detected by Cockyane Syndrome proteins A and B (CSA and CSB). CSA and CSB facilitate the recruitment of NER complexes to sites of DNA damage specifically on transcribed DNA regions, enabling more efficient repair of transcribed DNA (Transcription Coupled Repair is reviewed in 190).

36

Figure 1.14: Schematic of NER. NER is initiated when the XPC complex detects distortions in chromatin structure. XPC (or XPE) recruits TFIIH to the lesion site where it unwinds the affected region to generated a denatured bubble. Once the damaged strand is identified (possibly by XPA) the damaged region is excised by XPG, XPF and ERCC1 to generate an approximately 30 nucleotide single stranded region. This is then filled in by replicative DNA polymerases and ligated by DNA ligases 1 or 3α. Summarized from information in (184-189).

1.3.2.3 Indirect SSBR (Mismatch Repair):

Substitution and insertion/deletion mismatches which occur during DNA replication are repaired my MMR (reviewed in 191). It can also function as a BER-independent mechanism to excise 8-oxo-dG, which can be wrongly incorporated due to oxidation

37 of the nucleotide pool (115). The foremost consequence of disruption of MMR is an increased abundance of genome-wide point mutations. In prokaryotes this elevated mutation rate acts to improve the survival fitness of colonies under stress (192). In humans the improved survival fitness of sub-populations of cells gives rise to tumours (193).

MMR is initated when MutSα or MutSβ complexes bind to a lesion. Studies of MMR activity in cell free systems as well as yeast an animal models indicate that MutSα detects single base insertion/deletion and mismatch mutations whereas MutSβ detects larger insertions (194). Although little is known about how the mutated strand is identified, both MutSα and MutSβ interact with the replication factor PCNA, which may assist with strand discrimination (195). The excision and subsequent repair mechanisms are presently unclear, however it has been proposed that MMR links with other pathways via the interaction of MutSα and MutSβ with multi-functional repair factors including Flap Endonuclease 1 (FEN1), DNA polymerases δ and ε and Replication Proteins A and C (RPA and RPC, reference 196).

1.3.2.4 Indirect SSBR (Base Excision Repair):

In vivo oxidation of DNA results in several modifications including 8,5’-(S)-cyclo-2’- deoxyadenosine (cyclo-dA), 2,6-diamino-4-hydroxy-5-Nmethylformamidopyrimidine (faPy) and 8-oxo-deoxyguanosine (8-oxo-dG) (reviewed in 197). Of these lesions 8- oxo-dG is the most abundant and mutagenic (197,198). BER repairs damage to DNA bases, including oxidation, ethylation and methylation (199). It also repairs abasic sites which are the generated spontaneously due to the instability of the N-glycosyl bond in DNA (149). BER is carried out as a sequential process which initially involves detection and subsequent excision of the damaged base by a DNA glycosylase (an enzyme which hydrolyses the DNA N-glycosyl bond to generate an abasic site, references 200,201). There are two classes of DNA glycosylases: those that possess intrinsic ability to cleave the abasic sites that are the product of N- glycosyl bond hydrolysis (bifunctional DNA glycosylases) and those that do not (monofunctional DNA glycosylases) (202). Bifunctional DNA glycosylases, including the 8-oxo-dG DNA glycosylase OGG1, cleave abasic sites to yield a 3’-α,β- unsaturated aldehyde and 5’ phosphate single nucleotide gap (202). In this instance,

38 repair of the 3’-α,β-unsaturated aldehyde by APE1 produces a single nucleotide gap with 3’ hydroxyl and 5’ phosphate termini. This can subsequently be repaired in the same manner as direct SSBs (175). Interestingly the glycosylase activities of bifunctional DNA glycosylases are stimulated by APE1 (203,204). Bifunctional glycosylases bind stably to abasic sites and several studies have proposed that abasic endonucleases (including APE1) can displace these enzymes from abasic sites, facilitating enzyme turnover (203-205). Abasic sites produced by monofunctional DNA glycosylases are hydrolysed by APE1 to generate a 3’ hydroxyl, 5’ deoxyribose phosphate (dRP) gap (206). 5’dRP modified gaps are repaired by polymerization of a single nucleotide from the 3’ hydroxyl terminus. The exonuclease activity of polβ can subsequently hydrolyse a ‘standard’ 5’ dRP to generate a break with ligatable termini (207). This is referred to as short patch repair (reviewed in 199,208,209). Oxidised or reduced 5’dRP termini are poor substrates for the exonuclease activity of polβ (210). In this instance the damaged 5’ strand is displaced and additional nucleotides are synthesized by DNA polymerase δ or ε, generating a ‘flap’ in a process referred to as long patch repair (reviewed in 199,208-210). This structure is a substrate for FEN1 (211) which cleaves off the flap to generate a single strand break with 5’ phosphate and 3’ hydroxyl termini, which are subsequently rejoined by DNA ligase 1 (10,29). An overview of this process and its links with direct single strand break repair is shown in Figure 1.15.

39

Figure 1.15: Schematic of short and long patch repair mechanisms. BER is initiated when a DNA glycosylase detects base damage and hydrolyses the DNA N-glycosyl bond to generate an abasic site. This is then hydrolysed by either a bifunctional DNA glycosylase to produce a 3’α,β-unsaturated aldehyde single nucleotide gap, or by an abasic lyase (generally APE1) to generate a 5’ dRP single nucleotide gap. A 3’ α,β- unsaturated aldehyde can be repaired by APE1, yielding a single nucleotide gap with 3’ hydroxyl and 5’ phosphate termini which can be repaired in the same manner as direct single strand breaks. ‘Standard’ 5’ dRP termini are repaired by the 3’-5’ exonuclease activity of DNA polymerase β. This once again generates a single strand nick with 3’ hydroxyl and 5’ phosphate termini which are ligated by DNA ligase 3α. These two subpathways are jointly referred to as short patch repair. Alternatively if the single nucleotide gap has a modified (oxidized or reduced) 5’ dRP terminus, it cannot be excised by the exonuclease activity of DNA polymerase β. In this instance replicative polymerases, bound to the PCNA trimer are recruited to synthesize several nucleotides and displace the damaged terminus. This flap is cleaved by FEN1 to generate a clean break which is ligated by DNA ligase 1. Summarized from (183,199,208-210,212-214).

40 1.4 GENERAL CONCLUSIONS AND AIMS OF THIS THESIS:

DNA repair defective cerebellar ataxias are a heterogenous family of disorders where mutation of a single gene results in disruption of one or more critical genome maintenance mechanisms. Mutations in APTX were recently identified as causative for AOA1. These mutations cause destabilization of the APTX gene product Aprataxin. Aprataxin interacts with a number of proteins involved in different DNA repair mechanisms and AOA1 cells are hypersensitive to DNA damaging agents. At the commencement of the present study little was known about the in vitro or in vivo properties and functions of Aprataxin. Thus the aim of this work was to characterise the properties of Aprataxin and the nature of the defects in Aprataxin deficient (AOA1) cells.

Specific aims included:

1. Production of recombinant Aprataxin proteins and affinity purified antibodies to facilitate cellular and biochemical analyses of Aprataxin.

2. Characterisation of the biochemical properties of recombinant Aprataxin including:

i. Substrate preference

ii. DNA binding capacity

3. Characterisation of the in vivo role(s) of Aprataxin by examination of the defects in AOA1 cells. This included:

i. Analysis of the activity of multiple DNA repair pathways in AOA1 cells

ii. Identification of a role for Aprataxin in transcription

4. Examination of the influence of other proteins on Aprataxin’s in vivo activities.

41 1.5 REFERENCES

1. Boder, E. and Sedgwick, R.P. (1958) Ataxia-telangiectasia; a familial syndrome of progressive cerebellar ataxia, oculocutaneous telangiectasia and frequent pulmonary infection. Pediatrics, 21, 526-554. 2. Lavin, M.F. and Shiloh, Y. (1997) The genetic defect in ataxia-telangiectasia. Annu Rev Immunol, 15, 177-202. 3. Farina, L., Uggetti, C., Ottolini, A., Martelli, A., Bergamaschi, R., Sibilla, L., Zappoli, F., Egitto, M.G. and Lanzi, G. (1994) Ataxia-telangiectasia: MR and CT findings. J Comput Assist Tomogr, 18, 724-727. 4. Sedgwick, R.P. and Boder, E. (1960) Progressive ataxia in childhood with particular reference to ataxia-telangiectasia. Neurology, 10, 705-715. 5. Lewis, R.F., Lederman, H.M. and Crawford, T.O. (1999) Ocular motor abnormalities in ataxia telangiectasia. Ann Neurol, 46, 287-295. 6. Taylor, A.M., Metcalfe, J.A., Thick, J. and Mak, Y.F. (1996) Leukemia and lymphoma in ataxia telangiectasia. Blood, 87, 423-438. 7. Gotoff, S.P., Amirmokri, E. and Liebner, E.J. (1967) Ataxia telangiectasia. Neoplasia, untoward response to x-irradiation, and tuberous sclerosis. Am J Dis Child, 114, 617-625. 8. Morgan, J.L., Holcomb, T.M. and Morrissey, R.W. (1968) Radiation reaction in ataxia telangiectasia. Am J Dis Child, 116, 557-558. 9. Chen, R.L., Wang, P.J., Hsu, Y.H., Chang, P.Y. and Fang, J.S. (2002) Severe lung fibrosis after chemotherapy in a child with ataxia-telangiectasia. J Pediatr Hematol Oncol, 24, 77-79. 10. Irsfeld, H., Korholz, D., Janssen, G., Wahn, V. and Schroten, H. (2000) Fatal outcome in two girls with hodgkin disease complicating ataxia-telangiectasia (Louis-Bar syndrome) despite favorable response to modified-dose chemotherapy. Med Pediatr Oncol, 34, 62-64. 11. Sandoval, C. and Swift, M. (2000) Commentary: appropriate cancer chemotherapy for ataxia-telangiectasia patients. Med Pediatr Oncol, 34, 65- 66. 12. Biggar, W.D. and Good, R.A. (1975) immunodeficiency in ataxia- telangiectasia. Birth Defects Orig Artic Ser, 11, 271-276.

42 13. Bott, L., Lebreton, J., Thumerelle, C., Cuvellier, J., Deschildre, A. and Sardet, A. (2007) Lung disease in ataxia-telangiectasia. Acta Paediatr, 96, 1021-1024. 14. Savitsky, K., Bar-Shira, A., Gilad, S., Rotman, G., Ziv, Y., Vanagaite, L., Tagle, D.A., Smith, S., Uziel, T., Sfez, S. et al. (1995) A single ataxia telangiectasia gene with a product similar to PI-3 kinase. Science, 268, 1749- 1753. 15. Lakin, N.D., Weber, P., Stankovic, T., Rottinghaus, S.T., Taylor, A.M. and Jackson, S.P. (1996) Analysis of the ATM protein in wild-type and ataxia telangiectasia cells. Oncogene, 13, 2707-2716. 16. Abraham, R.T. (2004) PI 3-kinase related kinases: 'big' players in stress- induced signaling pathways. DNA Repair (Amst), 3, 883-887. 17. Ali, S.K., Bowman, W.P. and Aur, R.J. (1981) Increased radiation sensitivity of fibroblasts in ataxia telangiectasia with acute lymphocytic leukemia. Am J Dis Child, 135, 1142-1143. 18. Littlefield, L.G., Colyer, S.P., Joiner, E.E., DuFrain, R.J., Frome, E. and Cohen, M.M. (1981) Chromosomal radiation sensitivity in ataxia telangiectasia long-term lymphoblastoid cell lines. Cytogenet Cell Genet, 31, 203-213. 19. Painter, R.B. (1981) Radioresistant DNA synthesis: an intrinsic feature of ataxia telangiectasia. Mutat Res, 84, 183-190. 20. Bakkenist, C.J. and Kastan, M.B. (2003) DNA damage activates ATM through intermolecular autophosphorylation and dimer dissociation. Nature, 421, 499- 506. 21. Kozlov, S.V., Graham, M.E., Peng, C., Chen, P., Robinson, P.J. and Lavin, M.F. (2006) Involvement of novel autophosphorylation sites in ATM activation. The EMBO Journal, 25, 3504-3514. 22. Pellegrini, M., Celeste, A., Difilippantonio, S., Guo, R., Wang, W., Feigenbaum, L. and Nussenzweig, A. (2006) Autophosphorylation at serine 1987 is dispensable for murine Atm activation in vivo. Nature, 443, 222-225. 23. Kurz, E.U. and Lees-Miller, S.P. (2004) DNA damage-induced activation of ATM and ATM-dependent signaling pathways. DNA Repair (Amst), 3, 889- 900.

43 24. Goodarzi, A.A., Block, W.D. and Lees-Miller, S.P. (2003) The role of ATM and ATR in DNA damage-induced cell cycle control. Prog Cell Cycle Res, 5, 393-411. 25. Taylor, A.M., Groom, A. and Byrd, P.J. (2004) Ataxia-telangiectasia-like disorder (ATLD)-its clinical presentation and molecular basis. DNA Repair (Amst), 3, 1219-1225. 26. Stewart, G.S., Maser, R.S., Stankovic, T., Bressan, D.A., Kaplan, M.I., Jaspers, N.G., Raams, A., Byrd, P.J., Petrini, J.H. and Taylor, A.M. (1999) The DNA double-strand break repair gene hMRE11 is mutated in individuals with an ataxia-telangiectasia-like disorder. Cell, 99, 577-587. 27. Xiao, Y. and Weaver, D.T. (1997) Conditional gene targeted deletion by Cre recombinase demonstrates the requirement for the double-strand break repair Mre11 protein in murine embryonic stem cells. Nucleic Acids Res, 25, 2985- 2991. 28. Carney, J.P., Maser, R.S., Olivares, H., Davis, E.M., Le Beau, M., Yates, J.R., 3rd, Hays, L., Morgan, W.F. and Petrini, J.H. (1998) The hMre11/hRad50 protein complex and Nijmegen breakage syndrome: linkage of double-strand break repair to the cellular DNA damage response. Cell, 93, 477-486. 29. Cox, B.S. and Parry, J.M. (1968) The isolation, genetics and survival characteristics of ultraviolet light-sensitive mutants in yeast. Mutat Res, 6, 37- 55. 30. D'Amours, D. and Jackson, S.P. (2002) The Mre11 complex: at the crossroads of dna repair and checkpoint signalling. Nat Rev Mol Cell Biol, 3, 317-327. 31. Lee, J.H. and Paull, T.T. (2004) Direct activation of the ATM protein kinase by the Mre11/Rad50/Nbs1 complex. Science, 304, 93-96. 32. van Steeg, H. and Kraemer, K.H. (1999) Xeroderma pigmentosum and the role of UV-induced DNA damage in skin cancer. Mol Med Today, 5, 86-94. 33. Chu, G. and Mayne, L. (1996) Xeroderma pigmentosum, Cockayne syndrome and trichothiodystrophy: do the genes explain the diseases? Trends Genet, 12, 187-192. 34. Robins, P., Jones, C.J., Biggerstaff, M., Lindahl, T. and Wood, R.D. (1991) Complementation of DNA repair in xeroderma pigmentosum group A cell extracts by a protein with affinity for damaged DNA. The EMBO Journal, 10, 3913-3921.

44 35. Oh, K.S., Khan, S.G., Jaspers, N.G., Raams, A., Ueda, T., Lehmann, A., Friedmann, P.S., Emmert, S., Gratchev, A., Lachlan, K. et al. (2006) Phenotypic heterogeneity in the XPB DNA helicase gene (ERCC3): xeroderma pigmentosum without and with Cockayne syndrome. Hum Mutat, 27, 1092-1103. 36. Weeda, G., van Ham, R.C., Vermeulen, W., Bootsma, D., van der Eb, A.J. and Hoeijmakers, J.H. (1990) A presumed DNA helicase encoded by ERCC-3 is involved in the human repair disorders xeroderma pigmentosum and Cockayne's syndrome. Cell, 62, 777-791. 37. Peterson, C. and Legerski, R. (1991) High-frequency transformation of human repair-deficient cell lines by an Epstein-Barr virus-based cDNA expression vector. Gene, 107, 279-284. 38. Flejter, W.L., McDaniel, L.D., Johns, D., Friedberg, E.C. and Schultz, R.A. (1992) Correction of xeroderma pigmentosum complementation group D mutant cell phenotypes by chromosome and gene transfer: involvement of the human ERCC2 DNA repair gene. Proc Natl Acad Sci U S A, 89, 261-265. 39. Keeney, S., Eker, A.P., Brody, T., Vermeulen, W., Bootsma, D., Hoeijmakers, J.H. and Linn, S. (1994) Correction of the DNA repair defect in xeroderma pigmentosum group E by injection of a DNA damage-binding protein. Proc Natl Acad Sci U S A, 91, 4053-4056. 40. Sijbers, A.M., de Laat, W.L., Ariza, R.R., Biggerstaff, M., Wei, Y.F., Moggs, J.G., Carter, K.C., Shell, B.K., Evans, E., de Jong, M.C. et al. (1996) Xeroderma pigmentosum group F caused by a defect in a structure-specific DNA repair endonuclease. Cell, 86, 811-822. 41. Lalle, P., Nouspikel, T., Constantinou, A., Thorel, F. and Clarkson, S.G. (2002) The founding members of xeroderma pigmentosum group G produce XPG protein with severely impaired endonuclease activity. J Invest Dermatol, 118, 344-351. 42. Masutani, C., Kusumoto, R., Yamada, A., Dohmae, N., Yokoi, M., Yuasa, M., Araki, M., Iwai, S., Takio, K. and Hanaoka, F. (1999) The XPV (xeroderma pigmentosum variant) gene encodes human DNA polymerase eta. Nature, 399, 700-704. 43. de Boer, J. and Hoeijmakers, J.H. (2000) Nucleotide excision repair and human syndromes. Carcinogenesis, 21, 453-460.

45 44. Lehmann, A.R., Kirk-Bell, S., Arlett, C.F., Paterson, M.C., Lohman, P.H., de Weerd-Kastelein, E.A. and Bootsma, D. (1975) Xeroderma pigmentosum cells with normal levels of excision repair have a defect in DNA synthesis after UV-irradiation. Proc Natl Acad Sci U S A, 72, 219-223. 45. Lehmann, A.R. (2005) Replication of damaged DNA by translesion synthesis in human cells. FEBS letters, 579, 873-876. 46. Moreira, M.C., Klur, S., Watanabe, M., Nemeth, A.H., Le Ber, I., Moniz, J.C., Tranchant, C., Aubourg, P., Tazir, M., Schols, L. et al. (2004) Senataxin, the ortholog of a yeast RNA helicase, is mutant in ataxia-ocular apraxia 2. Nat Genet, 36, 225-227. 47. Le Ber, I., Bouslam, N., Rivaud-Pechoux, S., Guimaraes, J., Benomar, A., Chamayou, C., Goizet, C., Moreira, M.C., Klur, S., Yahyaoui, M. et al. (2004) Frequency and phenotypic spectrum of ataxia with oculomotor apraxia 2: a clinical and genetic study in 18 patients. Brain, 127, 759-767. 48. Criscuolo, C., Chessa, L., Di Giandomenico, S., Mancini, P., Sacca, F., Grieco, G.S., Piane, M., Barbieri, F., De Michele, G., Banfi, S. et al. (2006) Ataxia with oculomotor apraxia type 2: a clinical, pathologic, and genetic study. Neurology, 66, 1207-1210. 49. Suraweera, A., Becherel, O.J., Chen, P., Rundle, N., Woods, R., Nakamura, J., Gatei, M., Criscuolo, C., Filla, A., Chessa, L. et al. (2007) Senataxin, defective in ataxia oculomotor apraxia type 2, is involved in the defense against oxidative DNA damage. J Cell Biol, 177, 969-979. 50. Takashima, H., Boerkoel, C.F., John, J., Saifi, G.M., Salih, M.A., Armstrong, D., Mao, Y., Quiocho, F.A., Roa, B.B., Nakagawa, M. et al. (2002) Mutation of TDP1, encoding a topoisomerase I-dependent DNA damage repair enzyme, in spinocerebellar ataxia with axonal neuropathy. Nat Genet, 32, 267-272. 51. Christiansen, K. and Westergaard, O. (1996) The effect of camptothecin on topoisomerase I catalysis. Ann N Y Acad Sci, 803, 50-59. 52. El-Khamisy, S.F., Saifi, G.M., Weinfeld, M., Johansson, F., Helleday, T., Lupski, J.R. and Caldecott, K.W. (2005) Defective DNA single-strand break repair in spinocerebellar ataxia with axonal neuropathy-1. Nature, 434, 108- 113. 53. Miao, Z.H., Agama, K., Sordet, O., Povirk, L., Kohn, K.W. and Pommier, Y. (2006) Hereditary ataxia SCAN1 cells are defective for the repair of

46 transcription-dependent topoisomerase I cleavage complexes. DNA Repair (Amst), 5, 1489-1494. 54. Aicardi, J., Barbosa, C., Andermann, E., Andermann, F., Morcos, R., Ghanem, Q., Fukuyama, Y., Awaya, Y. and Moe, P. (1988) Ataxia-ocular motor apraxia: a syndrome mimicking ataxia-telangiectasia. Ann Neurol, 24, 497- 502. 55. Le Ber, I., Moreira, M.C., Rivaud-Pechoux, S., Chamayou, C., Ochsner, F., Kuntzer, T., Tardieu, M., Said, G., Habert, M.O., Demarquay, G. et al. (2003) Cerebellar ataxia with oculomotor apraxia type 1: clinical and genetic studies. Brain, 126, 2761-2772. 56. Barbot, C., Coutinho, P., Chorao, R., Ferreira, C., Barros, J., Fineza, I., Dias, K., Monteiro, J., Guimaraes, A., Mendonca, P. et al. (2001) Recessive ataxia with ocular apraxia: review of 22 Portuguese patients. Arch Neurol, 58, 201- 205. 57. Shimazaki, H., Takiyama, Y., Sakoe, K., Ikeguchi, K., Niijima, K., Kaneko, J., Namekawa, M., Ogawa, T., Date, H., Tsuji, S. et al. (2002) Early-onset ataxia with ocular motor apraxia and hypoalbuminemia: the aprataxin gene mutations. Neurology, 59, 590-595. 58. Amouri, R., Moreira, M.C., Zouari, M., El Euch, G., Barhoumi, C., Kefi, M., Belal, S., Koenig, M. and Hentati, F. (2004) Aprataxin gene mutations in Tunisian families. Neurology, 63, 928-929. 59. Shahwan, A., Byrd, P.J., Taylor, A.M., Nestor, T., Ryan, S. and King, M.D. (2006) Atypical presentation of ataxia-oculomotor apraxia type 1. Dev Med Child Neurol, 48, 529-532. 60. Moreira, M.C., Barbot, C., Tachi, N., Kozuka, N., Uchida, E., Gibson, T., Mendonca, P., Costa, M., Barros, J., Yanagisawa, T. et al. (2001) The gene mutated in ataxia-ocular apraxia 1 encodes the new HIT/Zn-finger protein aprataxin. Nat Genet, 29, 189-193. 61. Hirano, M., Nishiwaki, T., Kariya, S., Furiya, Y., Kawahara, M. and Ueno, S. (2004) Novel splice variants increase molecular diversity of aprataxin, the gene responsible for early-onset ataxia with ocular motor apraxia and hypoalbuminemia. Neurosci Lett, 366, 120-125.

47 62. Mosesso, P., Piane, M., Palitti, F., Pepe, G., Penna, S. and Chessa, L. (2005) The novel human gene aprataxin is directly involved in DNA single-strand- break repair. Cell Mol Life Sci, 62, 485-491. 63. Date, H., Onodera, O., Tanaka, H., Iwabuchi, K., Uekawa, K., Igarashi, S., Koike, R., Hiroi, T., Yuasa, T., Awaya, Y. et al. (2001) Early-onset ataxia with ocular motor apraxia and hypoalbuminemia is caused by mutations in a new HIT superfamily gene. Nat Genet, 29, 184-188. 64. Habeck, M., Zuhlke, C., Bentele, K.H., Unkelbach, S., Kress, W., Burk, K., Schwinger, E. and Hellenbroich, Y. (2004) Aprataxin mutations are a rare cause of early onset ataxia in Germany. J Neurol, 251, 591-594. 65. Criscuolo, C., Mancini, P., Sacca, F., De Michele, G., Monticelli, A., Santoro, L., Scarano, V., Banfi, S. and Filla, A. (2004) Ataxia with oculomotor apraxia type 1 in Southern Italy: late onset and variable phenotype. Neurology, 63, 2173-2175. 66. Seidle, H.F., Bieganowski, P. and Brenner, C. (2005) Disease-associated mutations inactivate AMP-lysine hydrolase activity of aprataxin. J Biol Chem, 280, 20927-20931. 67. Hirano, M., Asai, H., Kiriyama, T., Furiya, Y., Iwamoto, T., Nishiwaki, T., Yamamoto, A., Mori, T. and Ueno, S. (2007) Short half-lives of ataxia- associated aprataxin proteins in neuronal cells. Neurosci Lett, 419, 184-187. 68. Hofmann, K. and Bucher, P. (1995) The FHA domain: a putative nuclear signalling domain found in protein kinases and transcription factors. Trends Biochem Sci, 20, 347-349. 69. Darieva, Z., Pic-Taylor, A., Boros, J., Spanos, A., Geymonat, M., Reece, R.J., Sedgwick, S.G., Sharrocks, A.D. and Morgan, B.A. (2003) Cell cycle- regulated transcription through the FHA domain of Fkh2p and the coactivator Ndd1p. Curr Biol, 13, 1740-1745. 70. Roche, K.C., Wiechens, N., Owen-Hughes, T. and Perkins, N.D. (2004) The FHA domain protein SNIP1 is a regulator of the cell cycle and cyclin D1 expression. Oncogene, 23, 8185-8195. 71. Li, J., Williams, B.L., Haire, L.F., Goldberg, M., Wilker, E., Durocher, D., Yaffe, M.B., Jackson, S.P. and Smerdon, S.J. (2002) Structural and functional versatility of the FHA domain in DNA-damage signaling by the tumor suppressor kinase Chk2. Mol Cell, 9, 1045-1054.

48 72. Jilani, A., Ramotar, D., Slack, C., Ong, C., Yang, X.M., Scherer, S.W. and Lasko, D.D. (1999) Molecular cloning of the human gene, PNKP, encoding a polynucleotide kinase 3'-phosphatase and evidence for its role in repair of DNA strand breaks caused by oxidative damage. J Biol Chem, 274, 24176- 24186. 73. Marintchev, A., Robertson, A., Dimitriadis, E.K., Prasad, R., Wilson, S.H. and Mullen, G.P. (2000) Domain specific interaction in the XRCC1-DNA polymerase beta complex. Nucleic Acids Res, 28, 2049-2059. 74. Loizou JI, E.-K.S., Zlatanou A, Moore DJ, Chan DW, Qin J, Sarno S, Meggio F, Pinna LA, Caldecott KW. (2004) The protein kinase CK2 facilitates repair of chromosomal DNA single-strand breaks. Cell Mol Life Sci, 117, 17-28. 75. Clements, P.M., Breslin, C., Deeks, E.D., Byrd, P.J., Ju, L., Bieganowski, P., Brenner, C., Moreira, M.C., Taylor, A.M. and Caldecott, K.W. (2004) The ataxia-oculomotor apraxia 1 gene product has a role distinct from ATM and interacts with the DNA strand break repair proteins XRCC1 and XRCC4. DNA Repair (Amst), 3, 1493-1502. 76. Luo, H., Chan, D.W., Yang, T., Rodriguez, M., Chen, B.P., Leng, M., Mu, J.J., Chen, D., Songyang, Z., Wang, Y. et al. (2004) A new XRCC1- containing complex and its role in cellular survival of methyl methanesulfonate treatment. Mol Cell Biol, 24, 8356-8365. 77. Gueven, N., Becherel, O.J., Kijas, A.W., Chen, P., Howe, O., Rudolph, J.H., Gatti, R., Date, H., Onodera, O., Taucher-Scholz, G. et al. (2004) Aprataxin, a novel protein that protects against genotoxic stress. Hum Mol Genet, 13, 1081- 1093. 78. Date, H., Igarashi, S., Sano, Y., Takahashi, T., Takahashi, T., Takano, H., Tsuji, S., Nishizawa, M. and Onodera, O. (2004) The FHA domain of aprataxin interacts with the C-terminal region of XRCC1. Biochem Biophys Res Commun, 325, 1279-1285. 79. Thompson, J.D., Higgins, D.G. and Gibson, T.J. (1994) CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res, 22, 4673-4680. 80. Koch, C.A., Agyei, R., Galicia, S., Metalnikov, P., O'Donnell, P., Starostine, A., Weinfeld, M. and Durocher, D. (2004) Xrcc4 physically links DNA end

49 processing by polynucleotide kinase to DNA ligation by DNA ligase IV. The EMBO Journal, 23, 3874-3885. 81. van Heemst, D., Brugmans, L., Verkaik, N.S. and van Gent, D.C. (2004) End- joining of blunt DNA double-strand breaks in mammalian fibroblasts is precise and requires DNA-PK and XRCC4. DNA Repair (Amst), 3, 43-50. 82. Becherel, O.J., Gueven, N., Birrell, G.W., Schreiber, V., Suraweera, A., Jakob, B., Taucher-Scholz, G. and Lavin, M.F. (2006) Nucleolar localization of aprataxin is dependent on interaction with nucleolin and on active ribosomal DNA transcription. Hum Mol Genet, 15, 2239-2249. 83. Ginisty, H., Amalric, F. and Bouvet, P. (1998) Nucleolin functions in the first step of ribosomal RNA processing. The EMBO Journal, 17, 1476-1486. 84. Olson, M.O., Rivers, Z.M., Thompson, B.A., Kao, W.Y. and Case, S.T. (1983) Interaction of nucleolar phosphoprotein C23 with cloned segments of rat ribosomal deoxyribonucleic acid. Biochemistry, 22, 3345-3351. 85. Erard, M.S., Belenguer, P., Caizergues-Ferrer, M., Pantaloni, A. and Amalric, F. (1988) A major nucleolar protein, nucleolin, induces chromatin decondensation by binding to histone H1. Eur J Biochem, 175, 525-530. 86. Brenner, C. (2002) Hint, Fhit, and GalT: function, structure, evolution, and mechanism of three branches of the histidine triad superfamily of nucleotide hydrolases and transferases. Biochemistry, 41, 9003-9014. 87. Brenner, C., Bieganowski, P., Pace, H.C. and Huebner, K. (1999) The histidine triad superfamily of nucleotide-binding proteins. J Cell Physiol, 181, 179-187. 88. Hirano, M., Furiya, Y., Kariya, S., Nishiwaki, T. and Ueno, S. (2004) Loss of function mechanism in aprataxin-related early-onset ataxia. Biochem Biophys Res Commun, 322, 380-386. 89. Barnes, L.D., Garrison, P.N., Siprashvili, Z., Guranowski, A., Robinson, A.K., Ingram, S.W., Croce, C.M., Ohta, M. and Huebner, K. (1996) Fhit, a putative tumor suppressor in humans, is a dinucleoside 5',5"'-P1,P3-triphosphate hydrolase. Biochemistry, 35, 11529-11535. 90. Krakowiak, A., Pace, H.C., Blackburn, G.M., Adams, M., Mekhalfia, A., Kaczmarek, R., Baraniak, J., Stec, W.J. and Brenner, C. (2004) Biochemical, crystallographic, and mutagenic characterization of hint, the AMP-lysine

50 hydrolase, with novel substrates and inhibitors. J Biol Chem, 279, 18711- 18716. 91. Kijas, A.W., Harris, J.L., Harris, J.M. and Lavin, M.F. (2006) Aprataxin forms a discrete branch in the HIT (histidine triad) superfamily of proteins with both DNA/RNA binding and nucleotide hydrolase activities. J Biol Chem, 281, 13939-13948. 92. Li, H., Zhang, Y., Su, T., Santella, R.M. and Weinstein, I.B. (2006) Hint1 is a haplo-insufficient tumor suppressor in mice. Oncogene, 25, 713-721. 93. Fong, K.M., Biesterveld, E.J., Virmani, A., Wistuba, I., Sekido, Y., Bader, S.A., Ahmadian, M., Ong, S.T., Rassool, F.V., Zimmerman, P.V. et al. (1997) FHIT and FRA3B 3p14.2 allele loss are common in lung cancer and preneoplastic bronchial lesions and are associated with cancer-related FHIT cDNA splicing aberrations. Cancer research, 57, 2256-2267. 94. Ohta, M., Inoue, H., Cotticelli, M.G., Kastury, K., Baffa, R., Palazzo, J., Siprashvili, Z., Mori, M., McCue, P., Druck, T. et al. (1996) The FHIT gene, spanning the chromosome 3p14.2 fragile site and renal carcinoma-associated t(3;8) breakpoint, is abnormal in digestive tract cancers. Cell, 84, 587-597. 95. Rapaport, E. and Zamecnik, P.C. (1976) Presence of diadenosine 5',5''' -P1, P4-tetraphosphate (Ap4A) in mamalian cells in levels varying widely with proliferative activity of the tissue: a possible positive "pleiotypic activator". Proc Natl Acad Sci U S A, 73, 3984-3988. 96. Glover, T.W., Coyle-Morris, J.F., Li, F.P., Brown, R.S., Berger, C.S., Gemmill, R.M. and Hecht, F. (1988) Translocation t(3;8)(p14.2;q24.1) in renal cell carcinoma affects expression of the common fragile site at 3p14(FRA3B) in lymphocytes. Cancer Genet Cytogenet, 31, 69-73. 97. Yamakawa, K., Takahashi, E., Murata, M., Okui, K., Yokoyama, S. and Nakamura, Y. (1992) Detailed mapping around the breakpoint of (3;8) translocation in familial renal cell carcinoma and FRA3B. Genomics, 14, 412- 416. 98. Lima, C.D., Klein, M.G. and Hendrickson, W.A. (1997) Structure-based analysis of catalysis and substrate definition in the HIT protein family. Science, 278, 286-290. 99. Pace, H.C., Garrison, P.N., Robinson, A.K., Barnes, L.D., Draganescu, A., Rosler, A., Blackburn, G.M., Siprashvili, Z., Croce, C.M., Huebner, K. et al.

51 (1998) Genetic, biochemical, and crystallographic characterization of Fhit- substrate complexes as the active signaling form of Fhit. Proc Natl Acad Sci U S A, 95, 5484-5489. 100. Draganescu, A., Hodawadekar, S.C., Gee, K.R. and Brenner, C. (2000) Fhit- nucleotide specificity probed with novel fluorescent and fluorogenic substrates. J Biol Chem, 275, 4555-4560. 101. Ahel, I., Rass, U., El-Khamisy, S.F., Katyal, S., Clements, P.M., McKinnon, P.J., Caldecott, K.W. and West, S.C. (2006) The neurodegenerative disease protein aprataxin resolves abortive DNA ligation intermediates. Nature. 443, 713-716. 102. Sillero, A. and Sillero, M.A. (2000) Synthesis of dinucleoside polyphosphates catalyzed by firefly luciferase and several ligases. Pharmacol Ther, 87, 91- 102. 103. Rass, U., Ahel, I. and West, S.C. (2007) Actions of aprataxin in multiple DNA repair pathways. J Biol Chem, 282, 9469-9474. 104. Ahel, I., Rass, U., El-Khamisy, S.F., Katyal, S., Clements, P.M., McKinnon, P.J., Caldecott, K.W. and West, S.C. (2006) The neurodegenerative disease protein aprataxin resolves abortive DNA ligation intermediates. Nature, 443, 713-716. 105. Lehman, I.R. (1974) DNA ligase: structure, mechanism, and function. Science, 186, 790-797. 106. Takahashi, T., Tada, M., Igarashi, S., Koyama, A., Date, H., Yokoseki, A., Shiga, A., Yoshida, Y., Tsuji, S., Nishizawa, M. et al. (2007) Aprataxin, causative gene product for EAOH/AOA1, repairs DNA single-strand breaks with damaged 3'-phosphate and 3'-phosphoglycolate ends. Nucleic Acids Res, 35, 3797-3809. 107. Winters, T.A., Henner, W.D., Russell, P.S., McCullough, A. and Jorgensen, T.J. (1994) Removal of 3'-phosphoglycolate from DNA strand-break damage in an oligonucleotide substrate by recombinant human apurinic/apyrimidinic endonuclease 1. Nucleic Acids Res, 22, 1866-1873. 108. Wiederhold, L., Leppard, J.B., Kedar, P., Karimi-Busheri, F., Rasouli-Nia, A., Weinfeld, M., Tomkinson, A.E., Izumi, T., Prasad, R., Wilson, S.H. et al. (2004) AP endonuclease-independent DNA base excision repair in human cells. Mol Cell, 15, 209-220.

52 109. Vrana, K.E., Churchill, M.E., Tullius, T.D. and Brown, D.D. (1988) Mapping functional regions of transcription factor TFIIIA. Mol Cell Biol, 8, 1684-1696. 110. Brown, R.S. (2005) Zinc finger proteins: getting a grip on RNA. Curr Opin Struct Biol, 15, 94-98. 111. Friesen, W.J. and Darby, M.K. (2001) Specific RNA binding by a single C2H2 zinc finger. J Biol Chem, 276, 1968-1973. 112. Kennedy, D., Ramsdale, T., Mattick, J. and Little, M. (1996) An RNA recognition motif in Wilms' tumour protein (WT1) revealed by structural modelling. Nat Genet, 12, 329-331. 113. McCarty, A.S., Kleiger, G., Eisenberg, D. and Smale, S.T. (2003) Selective dimerization of a C2H2 zinc finger subfamily. Mol Cell, 11, 459-470. 114. Ames, B.N., Shigenaga, M.K. and Hagen, T.M. (1993) Oxidants, antioxidants, and the degenerative diseases of aging. Proc Natl Acad Sci U S A, 90, 7915- 7922. 115. Colussi, C., Parlanti, E., Degan, P., Aquilina, G., Barnes, D., Macpherson, P., Karran, P., Crescenzi, M., Dogliotti, E. and Bignami, M. (2002) The mammalian mismatch repair pathway removes DNA 8-oxodGMP incorporated from the oxidized dNTP pool. Curr Biol, 12, 912-918. 116. Schatz, D.G., Oettinger, M.A. and Schlissel, M.S. (1992) V(D)J recombination: molecular biology and regulation. Annu Rev Immunol, 10, 359- 383. 117. Zechiedrich, E.L. and Osheroff, N. (1990) Eukaryotic topoisomerases recognize nucleic acid topology by preferentially interacting with DNA crossovers. The EMBO Journal, 9, 4555-4562. 118. Hirano, M., Yamamoto, A., Mori, T., Lan, L., Iwamoto, T.A., Aoki, M., Shimada, K., Furiya, Y., Kariya, S., Asai, H. et al. (2007) DNA single-strand break repair is impaired in aprataxin-related ataxia. Ann Neurol, 61, 162-174. 119. Ward, J.F. (1988) DNA damage produced by ionizing radiation in mammalian cells: identities, mechanisms of formation, and reparability. Prog Nucleic Acid Res Mol Biol, 35, 95-125. 120. Povirk, L.F. (1996) DNA damage and mutagenesis by radiomimetic DNA- cleaving agents: bleomycin, neocarzinostatin and other enediynes. Mutat Res, 355, 71-89.

53 121. Kuzminov, A. (2001) Single-strand interruptions in replicating chromosomes cause double-strand breaks. Proc Natl Acad Sci U S A, 98, 8241-8246. 122. Takata, M., Sasaki, M.S., Sonoda, E., Morrison, C., Hashimoto, M., Utsumi, H., Yamaguchi-Iwai, Y., Shinohara, A. and Takeda, S. (1998) Homologous recombination and non-homologous end-joining pathways of DNA double- strand break repair have overlapping roles in the maintenance of chromosomal integrity in vertebrate cells. The EMBO Journal, 17, 5497-5508. 123. Feldmann, E., Schmiemann, V., Goedecke, W., Reichenberger, S. and Pfeiffer, P. (2000) DNA double-strand break repair in cell-free extracts from Ku80-deficient cells: implications for Ku serving as an alignment factor in non-homologous DNA end joining. Nucleic Acids Res, 28, 2585-2596. 124. Leber, R., Wise, T.W., Mizuta, R. and Meek, K. (1998) The XRCC4 gene product is a target for and interacts with the DNA-dependent protein kinase. J Biol Chem, 273, 1794-1801. 125. Smith, G.C. and Jackson, S.P. (1999) The DNA-dependent protein kinase. Genes Dev, 13, 916-934. 126. Li, B. and Comai, L. (2000) Functional interaction between Ku and the werner syndrome protein in DNA end processing. J Biol Chem, 275, 28349-28352. 127. Ma, Y., Pannicke, U., Schwarz, K. and Lieber, M.R. (2002) Hairpin opening and overhang processing by an Artemis/DNA-dependent protein kinase complex in nonhomologous end joining and V(D)J recombination. Cell, 108, 781-794. 128. Blunt, T., Finnie, N.J., Taccioli, G.E., Smith, G.C., Demengeot, J., Gottlieb, T.M., Mizuta, R., Varghese, A.J., Alt, F.W., Jeggo, P.A. et al. (1995) Defective DNA-dependent protein kinase activity is linked to V(D)J recombination and DNA repair defects associated with the murine scid mutation. Cell, 80, 813-823. 129. Danska, J.S., Pflumio, F., Williams, C.J., Huner, O., Dick, J.E. and Guidos, C.J. (1994) Rescue of T cell-specific V(D)J recombination in SCID mice by DNA-damaging agents. Science, 266, 450-455. 130. Murphy, W.J., Durum, S.K., Anver, M.R., Ferris, D.K., McVicar, D.W., O'Shea, J.J., Ruscetti, S.K., Smith, M.R., Young, H.A. and Longo, D.L. (1994) Induction of T cell differentiation and lymphomagenesis in the thymus of mice

54 with severe combined immune deficiency (SCID). J Immunol, 153, 1004- 1014. 131. Morrison, C., Smith, G.C., Stingl, L., Jackson, S.P., Wagner, E.F. and Wang, Z.Q. (1997) Genetic interaction between PARP and DNA-PK in V(D)J recombination and tumorigenesis. Nat Genet, 17, 479-482. 132. Chatterjee, S., Berger, S.J. and Berger, N.A. (1999) Poly(ADP-ribose) polymerase: a guardian of the genome that facilitates DNA repair by protecting against DNA recombination. Mol Cell Biochem, 193, 23-30. 133. Audebert, S., Calsou. (2004) Involvement of Poly(ADP-ribose) Polymerase-1 and XRCC1/DNA Ligase III in an Alternative Route for DNA Double-strand Breaks Rejoining. J Biol Chem, 279, 55117-55126. 134. Audebert, S., Weinfeld , Calsou (2006) Involvement of polynucleotide kinase in a poly(ADP-ribose) polymerase-1-dependent dna double-strand breaks rejoining pathway. Journal of Molecular Biology, 356, 257-265. 135. Wang, M., Wu, W., Wu, W., Rosidi, B., Zhang, L., Wang, H. and Iliakis, G. (2006) PARP-1 and Ku compete for repair of DNA double strand breaks by distinct NHEJ pathways. Nucleic Acids Res, 34, 6170-6182. 136. Perrault, R., Wang, H., Wang, M., Rosidi, B. and Iliakis, G. (2004) Backup pathways of NHEJ are suppressed by DNA-PK. J Cell Biochem, 92, 781-794. 137. Hoeijmakers, J.H. (2001) Genome maintenance mechanisms for preventing cancer. Nature, 411, 366-374. 138. Dunham, M.A., Neumann, A.A., Fasching, C.L. and Reddel, R.R. (2000) Telomere maintenance by recombination in human cells. Nat Genet, 26, 447- 450. 139. Abraham, R.T. (2001) Cell cycle checkpoint signaling through the ATM and ATR kinases. Genes Dev, 15, 2177-2196. 140. Morrison, C., Sonoda, E., Takao, N., Shinohara, A., Yamamoto, K. and Takeda, S. (2000) The controlling role of ATM in homologous recombinational repair of DNA damage. The EMBO Journal, 19, 463-471. 141. You, Z., Chahwan, C., Bailis, J., Hunter, T. and Russell, P. (2005) ATM activation and its recruitment to damaged DNA require binding to the C terminus of Nbs1. Mol Cell Biol, 25, 5363-5379. 142. Liu, N., Lamerdin, J.E., Tebbs, R.S., Schild, D., Tucker, J.D., Shen, M.R., Brookman, K.W., Siciliano, M.J., Walter, C.A., Fan, W. et al. (1998) XRCC2

55 and XRCC3, new human Rad51-family members, promote chromosome stability and protect against DNA cross-links and other damages. Mol Cell, 1, 783-793. 143. Miyazaki, T., Bressan, D.A., Shinohara, M., Haber, J.E. and Shinohara, A. (2004) In vivo assembly and disassembly of Rad51 and Rad52 complexes during double-strand break repair. The EMBO Journal, 23, 939-949. 144. Paull, T.T. and Gellert, M. (1998) The 3' to 5' exonuclease activity of Mre 11 facilitates repair of DNA double-strand breaks. Mol Cell, 1, 969-979. 145. McIlwraith, M.J., Van Dyck, E., Masson, J.Y., Stasiak, A.Z., Stasiak, A. and West, S.C. (2000) Reconstitution of the strand invasion step of double-strand break repair using human Rad51 Rad52 and RPA proteins. J Mol Biol, 304, 151-164. 146. van den Bosch, M., Lohman, P.H. and Pastink, A. (2002) DNA double-strand break repair by homologous recombination. Biological chemistry, 383, 873- 892. 147. Zhou, W. and Doetsch, P.W. (1994) Transcription bypass or blockage at single-strand breaks on the DNA template strand: effect of different 3' and 5' flanking groups on the T7 RNA polymerase elongation complex. Biochemistry, 33, 14926-14934. 148. Sanchez, G. and Mamet-Bratley, M.D. (1994) Transcription by T7 RNA polymerase of DNA containing abasic sites. Environmental and Molecular Mutagenesis, 23, 32-36. 149. Lindahl, T. (1993) Instability and decay of the primary structure of DNA. Nature, 362, 709-715. 150. Kobayashi, S., Ueda, K. and Komano, T. (1990) The effects of metal ions on the DNA damage induced by hydrogen peroxide. Agric Biol Chem, 54, 69-76. 151. Breen, A.P. and Murphy, J.A. (1995) Reactions of oxyl radicals with DNA. Free Radical Biology & Medicine, 18, 1033-1077. 152. Woodhouse, B.C., Dianova, II, Parsons, J.L. and Dianov, G.L. (2008) Poly(ADP-ribose) polymerase-1 modulates DNA repair capacity and prevents formation of DNA double strand breaks. DNA Repair (Amst), 7, 932-940. 153. Schreiber, V., Dantzer, F., Ame, J.C. and de Murcia, G. (2006) Poly(ADP- ribose): novel functions for an old molecule. Nat Rev Mol Cell Biol, 7, 517- 528.

56 154. Ame, J.C., Spenlehauer, C. and de Murcia, G. (2004) The PARP superfamily. Bioessays, 26, 882-893. 155. Althaus, F.R. and Richter, C. (1987) ADP-ribosylation of proteins. Enzymology and biological significance. Mol Biol Biochem Biophys, 37, 1- 237. 156. Riquelme, P.T., Burzio, L.O. and Koide, S.S. (1979) ADP ribosylation of rat liver lysine-rich histone in vitro. J Biol Chem, 254, 3018-3028. 157. Burzio, L.O., Riquelme, P.T. and Koide, S.S. (1979) ADP ribosylation of rat liver nucleosomal core histones. J Biol Chem, 254, 3029-3037. 158. Suzuki, H., Quesada, P., Farina, B. and Leone, E. (1986) In vitro poly(ADP- ribosyl)ation of seminal ribonuclease. J Biol Chem, 261, 6048-6055. 159. Hayashi, K., Tanaka, M., Shimada, T., Miwa, M. and Sugimura, T. (1983) Size and shape of poly(ADP-ribose): examination by gel filtration, gel electrophoresis and electron microscopy. Biochem Biophys Res Commun, 112, 102-107. 160. Le Page, F., Schreiber, V., Dherin, C., De Murcia, G. and Boiteux, S. (2003) Poly(ADP-ribose) polymerase-1 (PARP-1) is required in murine cell lines for base excision repair of oxidative DNA damage in the absence of DNA polymerase beta. J Biol Chem, 278, 18471-18477. 161. Schreiber, V., Ame, J.C., Dolle, P., Schultz, I., Rinaldi, B., Fraulob, V., Menissier-de Murcia, J. and de Murcia, G. (2002) Poly(ADP-ribose) polymerase-2 (PARP-2) is required for efficient base excision DNA repair in association with PARP-1 and XRCC1. J Biol Chem, 277, 23028-23036. 162. Masutani, M., Nozaki, T., Nakamoto, K., Nakagama, H., Suzuki, H., Kusuoka, O., Tsutsumi, M. and Sugimura, T. (2000) The response of Parp knockout mice against DNA damaging agents. Mutat Res, 462, 159-166. 163. Benjamin, R.C. and Gill, D.M. (1980) ADP-ribosylation in mammalian cell ghosts. Dependence of poly(ADP-ribose) synthesis on strand breakage in DNA. J Biol Chem, 255, 10493-10501. 164. Shieh, W.M., Ame, J.C., Wilson, M.V., Wang, Z.Q., Koh, D.W., Jacobson, M.K. and Jacobson, E.L. (1998) Poly(ADP-ribose) polymerase null mouse cells synthesize ADP-ribose polymers. J Biol Chem, 273, 30069-30072.

57 165. Lindahl, T., Satoh, M.S., Poirier, G.G. and Klungland, A. (1995) Post- translational modification of poly(ADP-ribose) polymerase induced by DNA strand breaks. Trends Biochem Sci, 20, 405-411. 166. Boulikas, T., Bastin, B., Boulikas, P. and Dupuis, G. (1990) Increase in histone poly (ADP-ribosylation) in mitogen-activated lymphoid cells. Exp Cell Res, 187, 77-84. 167. Adamietz, P. and Rudolph, A. (1984) ADP-ribosylation of nuclear proteins in vivo. Identification of histone H2B as a major acceptor for mono- and poly(ADP-ribose) in dimethyl sulfate-treated hepatoma AH 7974 cells. J Biol Chem, 259, 6841-6846. 168. Ogata, N., Ueda, K. and Hayaishi, O. (1980) ADP-ribosylation of histone H2B. Identification of glutamic acid residue 2 as the modification site. J Biol Chem, 255, 7610-7615. 169. Ferro, A.M., Thompson, L.H. and Olivera, B.M. (1984) Poly (ADP- ribosylation) and DNA topoisomerase I in different cell lines. Adv Exp Med Biol, 179, 441-447. 170. Scovassi, A.I., Mariani, C., Negroni, M., Negri, C. and Bertazzoni, U. (1993) ADP-ribosylation of nonhistone proteins in HeLa cells: modification of DNA topoisomerase II. Exp Cell Res, 206, 177-181. 171. Adelfalk, C., Kontou, M., Hirsch-Kauffmann, M. and Schweiger, M. (2003) Physical and functional interaction of the Werner syndrome protein with poly- ADP ribosyl transferase. FEBS letters, 554, 55-58. 172. Desmarais, Y., Menard, L., Lagueux, J. and Poirier, G.G. (1991) Enzymological properties of poly(ADP-ribose)polymerase: characterization of automodification sites and NADase activity. Biochim Biophys Acta, 1078, 179-186. 173. Smulson, M., Istock, N., Ding, R. and Cherney, B. (1994) Deletion mutants of poly(ADP-ribose) polymerase support a model of cyclic association and dissociation of enzyme from DNA ends during DNA repair. Biochemistry, 33, 6186-6191. 174. El-Khamisy, S.F., Masutani, M., Suzuki, H. and Caldecott, K.W. (2003) A requirement for PARP-1 for the assembly or stability of XRCC1 nuclear foci at sites of oxidative DNA damage. Nucleic Acids Research, 31, 5526-5533.

58 175. Vidal, A.E., Boiteux, S., Hickson, I.D. and Radicella, J.P. (2001) XRCC1 coordinates the initial and late stages of DNA abasic site repair through protein-protein interactions. The EMBO Journal, 20, 6530-6539. 176. Das, A., Wiederhold, L., Leppard, J.B., Kedar, P., Prasad, R., Wang, H., Boldogh, I., Karimi-Busheri, F., Weinfeld, M., Tomkinson, A.E. et al. (2006) NEIL2-initiated, APE-independent repair of oxidized bases in DNA: Evidence for a repair complex in human cells. DNA Repair (Amst), 5, 1439-1448. 177. Dyrkheeva, N.S., Khodyreva, S.N. and Lavrik, O.I. (2007) [Multifunctional human apurinic/apyrimidinic endonuclease 1: the role of additional functions]. Mol Biol (Mosk), 41, 450-466. 178. Whitehouse, C.J., Taylor, R.M., Thistlethwaite, A., Zhang, H., Karimi- Busheri, F., Lasko, D.D., Weinfeld, M. and Caldecott, K.W. (2001) XRCC1 stimulates human polynucleotide kinase activity at damaged DNA termini and accelerates DNA single-strand break repair. Cell, 104, 107-117. 179. Parsons, J.L., Dianova, II and Dianov, G.L. (2004) APE1 is the major 3'- phosphoglycolate activity in human cell extracts. Nucleic Acids Res, 32, 3531- 3536. 180. Inamdar, K.V., Pouliot, J.J., Zhou, T., Lees-Miller, S.P., Rasouli-Nia, A. and Povirk, L.F. (2002) Conversion of phosphoglycolate to phosphate termini on 3' overhangs of DNA double strand breaks by the human tyrosyl-DNA phosphodiesterase hTdp1. J Biol Chem, 277, 27162-27168. 181. Dianov, G., Price, A. and Lindahl, T. (1992) Generation of single-nucleotide repair patches following excision of uracil residues from DNA. Mol Cell Biol, 12, 1605-1612. 182. Chang, L.M. (1975) The distributive nature of enzymatic DNA synthesis. J Mol Biol, 93, 219-235. 183. Caldecott, K.W. (2003) Protein-protein interactions during mammalian DNA single-strand break repair. Biochem Soc Trans, 31, 247-251. 184. Nouspikel, T. (2008) Nucleotide excision repair and neurological diseases. DNA Repair (Amst), 7, 1155-1167. 185. Sugasawa, K., Ng, J.M., Masutani, C., Iwai, S., van der Spek, P.J., Eker, A.P., Hanaoka, F., Bootsma, D. and Hoeijmakers, J.H. (1998) Xeroderma pigmentosum group C protein complex is the initiator of global genome nucleotide excision repair. Mol Cell, 2, 223-232.

59 186. Wood, R.D. (1999) DNA damage recognition during nucleotide excision repair in mammalian cells. Biochimie, 81, 39-44. 187. Evans, E., Moggs, J.G., Hwang, J.R., Egly, J.M. and Wood, R.D. (1997) Mechanism of open complex and dual incision formation by human nucleotide excision repair factors. The EMBO Journal, 16, 6559-6573. 188. Buschta-Hedayat, N., Buterin, T., Hess, M.T., Missura, M. and Naegeli, H. (1999) Recognition of nonhybridizing base pairs during nucleotide excision repair of DNA. Proc Natl Acad Sci U S A, 96, 6090-6095. 189. Jones, C.J. and Wood, R.D. (1993) Preferential binding of the xeroderma pigmentosum group A complementing protein to damaged DNA. Biochemistry, 32, 12096-12104. 190. Svejstrup, J.Q. (2002) Mechanisms of transcription-coupled DNA repair. Nat Rev Mol Cell Biol, 3, 21-29. 191. Kunkel, T.A. and Erie, D.A. (2005) DNA mismatch repair. Annu Rev Biochem, 74, 681-710. 192. Halas, A., Baranowska, H. and Policinska, Z. (2002) The influence of the mismatch-repair system on stationary-phase mutagenesis in the yeast Saccharomyces cerevisiae. Curr Genet, 42, 140-146. 193. Peltomaki, P. (1997) DNA mismatch repair gene mutations in human cancer. Environ Health Perspect, 105 Suppl 4, 775-780. 194. Genschel, J., Littman, S.J., Drummond, J.T. and Modrich, P. (1998) Isolation of MutSbeta from human cells and comparison of the mismatch repair specificities of MutSbeta and MutSalpha. J Biol Chem, 273, 19895-19901. 195. Modrich, P. (1997) Strand-specific mismatch repair in mammalian cells. J Biol Chem, 272, 24727-24730. 196. Kolodner, R.D. and Marsischky, G.T. (1999) Eukaryotic DNA mismatch repair. Current Opinion in Genetics & Development, 9, 89-96. 197. Croteau, D.L. and Bohr, V.A. (1997) Repair of oxidative damage to nuclear and mitochondrial DNA in mammalian cells. J Biol Chem, 272, 25409-25412. 198. Poulsen, H.E., Prieme, H. and Loft, S. (1998) Role of oxidative DNA damage in cancer initiation and promotion. Eur J Cancer Prev, 7, 9-16. 199. Caldecott, K.W. (2001) Mammalian DNA single-strand break repair: an X- ra(y)ted affair. Bioessays, 23, 447-455.

60 200. David-Cordonnier, M.H., Boiteux, S. and O'Neill, P. (2001) Excision of 8- oxoguanine within clustered damage by the yeast OGG1 protein. Nucleic Acids Res, 29, 1107-1113. 201. Parsons, J.L., Zharkov, D.O. and Dianov, G.L. (2005) NEIL1 excises 3' end proximal oxidative DNA lesions resistant to cleavage by NTH1 and OGG1. Nucleic Acids Res, 33, 4849-4856. 202. Fortini, P., Parlanti, E., Sidorkina, O.M., Laval, J. and Dogliotti, E. (1999) The type of DNA glycosylase determines the base excision repair pathway in mammalian cells. J Biol Chem, 274, 15230-15236. 203. Hill, J.W., Hazra, T.K., Izumi, T. and Mitra, S. (2001) Stimulation of human 8-oxoguanine-DNA glycosylase by AP-endonuclease: potential coordination of the initial steps in base excision repair. Nucleic Acids Res, 29, 430-438. 204. Vidal, A.E., Hickson, I.D., Boiteux, S. and Radicella, J.P. (2001) Mechanism of stimulation of the DNA glycosylase activity of hOGG1 by the major human AP endonuclease: bypass of the AP lyase activity step. Nucleic Acids Res, 29, 1285-1292. 205. Mokkapati, S.K., Wiederhold, L., Hazra, T.K. and Mitra, S. (2004) Stimulation of DNA glycosylase activity of OGG1 by NEIL1: functional collaboration between two human DNA glycosylases. Biochemistry, 43, 11596-11604. 206. Wilson, D.M., 3rd and Barsky, D. (2001) The major human abasic endonuclease: formation, consequences and repair of abasic lesions in DNA. Mutat Res, 485, 283-307. 207. Matsumoto, Y. and Kim, K. (1995) Excision of deoxyribose phosphate residues by DNA polymerase beta during DNA repair. Science, 269, 699-702. 208. Dianov, G.L. and Parsons, J.L. (2007) Co-ordination of DNA single strand break repair. DNA Repair (Amst), 6, 454-460. 209. Lindahl, T., Karran, P. and Wood, R.D. (1997) DNA excision repair pathways. Current Opinion in Genetics & Development, 7, 158-169. 210. Caldecott, K.W. (2003) XRCC1 and DNA strand break repair. DNA Repair (Amst), 2, 955-969. 211. Kim, K., Biade, S. and Matsumoto, Y. (1998) Involvement of flap endonuclease 1 in base excision DNA repair. J Biol Chem, 273, 8842-8848.

61 212. Demple, B. and Harrison, L. (1994) Repair of oxidative damage to DNA: enzymology and biology. Annu Rev Biochem, 63, 915-948. 213. Fortini, P., Pascucci, B., Parlanti, E., D'Errico, M., Simonelli, V. and Dogliotti, E. (2003) 8-Oxoguanine DNA damage: at the crossroad of alternative repair pathways. Mutat Res, 531, 127-139. 214. Wilson, D.M., 3rd. (2007) Processing of nonconventional DNA strand break ends. Environmental and Molecular Mutagenesis, 48, 772-782.

62

CHAPTER 2

Generation of recombinant Aprataxin and Aprataxin- specific antibodies

63 64 2.1 INTRODUCTION

The identification of a novel gene is often the beginning of a long road to discover the cellular function of the gene product. The functional characterization of a protein can follow several approaches. In silico technologies can be used to analyse and compare nucleotide and protein sequences for the presence of previously identified structural and functional domains or sequence similarities. They can also be used to characterize the relationship between the protein of interest and others within a protein family. This can be helpful if the domains or homologous proteins identified have previously described functions.

2.1.1 Expression and purification of recombinant proteins: Another approach involves the expression of recombinant protein, followed by purification. This is usually achieved by cloning the cDNA into an expression system that allows the generation of a fusion of the protein of interest and a polypeptide fusion partner, termed a tag. In addition to facilitating protein purification, tags are useful for manipulation of the target protein. For example, tags are useful when the protein of interest does not have an available antibody. In this instance an antibody against the tag can be used to detect the fusion protein. Tags are also commonly used to facilitate precipitation of the fusion protein. This is an invaluable technique for purification of the fusion protein from a complex mixture such as cell lysate. Tags can be used on either or both termini of the protein of interest (the use of C and N-terminal tags on the same protein to facilitate purification is termed Tandem Affinity Purification). Commonly used tags include: Glutathione-S-Transferase (GST), a polypeptide which binds to reduced glutathione immobilized on a solid matrix (1); poly histidine (hexa-His is common), which has a high affinity for immobilized divalent cations (Ni2+ is normally used, 2); Inteins, which binds to chitin (3); and FLAG peptide which can be purified by immunoprecipitation (4).

Although the use of tags simplifies and facilitates protein purification, they have a number of drawbacks. In some instances the presence of a tag may disrupt the tertiary structure of the protein of interest, and this misfolding may lead to aggregation, precipitation or degradation of the fusion protein, resulting in a poor yield. Although aggregation of proteins can often be reduced by the use of cell lines engineered to

65 express chaperone proteins (reviewed in 5) or the addition of mild detergents throughout the lysis and purification procedure, the final product, although soluble, may still be misfolded and inactive. The presence of a tag can also interfere with the function of a protein. For example some tags, particularly large ones such as GST, may prevent the target protein from binding its interaction partners. This can be particularly problematic when the target protein is small relative to the size of the tag. Additionally, fusion with GFP is known to change the sub-cellular localization of some proteins. The purification conditions required for some tags (for example elevated temperature, elevated pH, or high concentrations of reducing agents in Intein based systems, 3) may also result in inactivation of the target protein. When selecting an expression system, some factors to consider are: yield; purity; robustness of the system, and cost. FLAG and Hexa-His systems are generally expensive to use compared to Intein and GST based systems. Intein based systems are inexpensive as the affinity resin can be re-used several times, although the yields are generally not as high as GST based systems and the purification conditions are quite harsh. GST based systems normally provide good yields for proteins below 100 kDa and harsh conditions are not normally necessary, making them an ideal choice provided that the bulky tag does not interfere with function of the target protein.

Once a recombinant protein is produced, it can be tested for various activities as indicated by its homology to other proteins and domain structure. Domains can also be expressed individually to study their function independently from the rest of the protein. Point mutations (generally to alanine or glycine) can be introduced throughout the protein to determine the locations of residues critical for protein function.

2.1.2 Generation of specific antibodies: Thirdly, purified recombinant proteins may also be used for immunization of laboratory animals to generate antibodies to the proteins or domains of interest. The immunized animal may be used to generate monoclonal or polyclonal antibodies. The generation of monoclonal antibodies is a lengthy process (six months or more), and involves immunization of the animal, generation of hybridomas followed by harvesting and purification of the antibody (6). This process results in the generation of an immortal cell line and therefore can provide a virtually unlimited supply of antibody with minimal batch to batch variability; however it is very expensive. Polyclonal antibodies

66 take much less time to produce than monoclonal antibodies (the first bleed is normally two to four weeks after the initial immunization) making them often a more convenient and cheaper option (6). Depending on the animal used, the large volume of serum collected can compensate for the absence of an infinite supply of antibody-generating cells. One drawback of the use of polyclonal antibodies is variation in antibody specificity and titer between batches, however this can be overcome by pooling several bleeds. One major advantage of polyclonal antibodies over monoclonal ones is the presence of antibodies against multiple epitopes in polyclonal serum. When an animal is immunized many immune cell clones express and release a wide range of antibodies into the serum, often to epitopes at different locations along the immunogen. In contrast, monoclonal antibodies are produced by immortalization of a single antibody producing cell, which can produce only a single antibody (6). This makes the use of polyclonal antibodies advantageous for several applications, including immunoflourescence and immunoprecipitation where one or more epitopes on the target protein may be blocked by other interacting molecules. Each antibody raised will have different characteristics, dependant on the antigen it was raised against (whole protein or peptide, native or denatured, mono or polyclonal antibody). Therefore every antibody must be tested and optimized for each application such as immunoblotting, immunostaining, immunoprecipitation.

Antibodies generated by these means are commonly used to study several aspects of protein biochemistry. They may be used to enrich the protein of interest from cell or tissue lysate to study the function of the endogenous protein. They are useful tools to observe changes in abundance of the target protein under different conditions, the localization and trafficking of proteins in a cell, and the expression of the protein in different tissues.

This chapter deals with the expression of recombinant Aprataxin from two bacterial expression systems, pTYB1 (Intein based) and pGEX 6.1 (GST based), as well as the purification of Aprataxin specific antibodies from the serum of immunized animals.

67 2.2 MATERIALS AND METHODS

2.2.1 Expression constructs: pTYB1 (New England Biolabs) constructs coding for full length wild-type (NCBI accession number AAQ74130) and V263G Aprataxin cDNAs were generated during this students honours year. Briefly, Aprataxin cDNA was amplified using the primers 5’ CCTAGCTGCATGCGGGTGTGCTGGTTGGTG 3’ and 5’ TTTATAGCGGCCGCCCTGTGTCCAGTGCTTCCTGAG 3’. The PCR products were gel purified and digested with the restriction endonucleases Nhe1 and Not1 (New England Biolabs) in the recommended buffers, precipitated with 1/10 volumes of 3 M sodium acetate pH 5.2 and 1 volume of isopropanol, resuspended in 50 mM Tris pH 7.5 with 5 mM EDTA and ligated using T4 DNA ligase (New England Biolabs) with Nhe1 and Not1 digested pTYB1.

pGEX 6.1 Aprataxin expression constructs were generated by Dr Amanda Kijas (Queensland Institute for Medical Research, Brisbane, Australia). Briefly, expression constructs of full-length (Protein A), N-terminal deleted (Protein B), C-terminal deleted (Protein C), and N-and C-terminal deleted (Protein D) forms of Aprataxin were prepared using PCR of a full length Aprataxin cDNA clone using the primer sets indicated in Table 1. These amplifications introduced BamHI and XhoI sites for cloning into pGEX-6.1 (Amersham Biosciences). This cloning strategy leads to the addition of five amino acids (GPLGS) to the N terminus after cleavage with PreScission protease.

Sequence, 5'-3' Protein A Forward CGGGATCCATGATGCGGGTGTGCTGGTTG Reverse CCGCTCGAGTCACTGTGTCCAGTGCTTCCT Protein B Forward CGGGATCCTCAGGCAACAGTGATTCTATAGA Reverse CCGCTCGAGTCACTGTGTCCAGTGCTTCCT Protein C Forward CGGGATCCATGATGATGCGGGTGTGCTGGTTGG Reverse CCGCTCGAGTCAACGAAGGGGCAGCTTCA Protein D Forward CGGGATCCTCAGGCAACAGTGATTCTATAGA Reverse CCGCTCGAGTCAACGAAGGGGCAGCTTCA Table 2.1: Aprataxin pGEX 6.1 cloning primers.

68

2.2.2 Plasmid preparation: pTYB1 and pGEX 6.1 expression vectors coding for the wild-type and V263G Aprataxin proteins were prepared from transformed stock DH5α E.coli (Invitrogen) by the alkaline lysis method. Briefly, the cell pellet from 2 to 10 mL of overnight LB culture was resuspended in 50 mM Tris pH 8.0, 10 mM Na-EDTA, with 100 mg/L RNase A (New England Biolabs), and lysed in 200 mM NaOH with 1% SDS. Genomic DNA was precipitated in 3 M potassium acetate adjusted to pH 5.5 with glacial acetic acid. The supernatant was recovered, and mixed with an equal volume of 1:1 phenol:chloroform pH 8.0. The aqueous phase was recovered and the remaining phenol was extracted using one volume of chloroform. The aqueous phase was recovered, and the plasmid was precipitated using 0.1 volumes of 3 M potassium acetate and 1 volume of isopropanol. The purified plasmid was resuspended in 50 mM Tris pH 7.5 with 5 mM EDTA.

2.2.3 Preparation of Competent Bacterial Cells: All plasmid manipulations were performed in DH5α E.coli and all expressions were performed in BL-21 (DE3)pLys E.coli. Stocks for both of these cell strains were prepared by streaking a stock culture onto an LB agar plate and incubating overnight at 37°C. Single colonies were picked and inoculated into LB media and incubated overnight at 37°C. Competent cells were prepared by inoculation 20 mL of this saturated culture into 1 L of LB media. The culture was grown at 30°C, shaking at 200 rpm until the culture reached an OD600 of 0.5 (approximately 3 hours). Cells were then pelleted by centrifugation at 5,000 x g for 10 minutes in a Beckman JA-10 rotor at 4°C.

The cell pellet was then washed 3 times in ice-cold 0.1 mM CaCl2, and pelleted by centrifugation at 5,000 x g at 4°C. The cells were resuspended in 5 mL of ice-cold 0.1 mM CaCl2 with 10% glycerol and incubated on ice for 1 hour. The cells were then aliquoted and snap frozen on dry ice before being stored at -80°C.

2.2.4 Transformation of Competent Cells: Transformation of competent E.coli cells was performed by the heat shock procedure. Briefly, transformation was achieved by incubation of up to 100 ng of plasmid or 5 µL of ligation mixture with 100 µL of competent cells for 30 minutes on ice. This was followed by heat shock treatment (42° for 45 seconds) followed by a 2 minute recovery

69 on ice. 500 µL of pre-warmed LB media was then added to the cells, which were subsequently incubated at 37°C for 45 minutes under agitation at 200 rpm. Cells were then pelleted by centrifugation at 2,000 x g on a benchtop centrifuge for 1 minute, resuspended in 100 µL of LB media and spread onto LB agar plates (10 g/L NaCl, 10 g/L peptone, 5 g/L yeast extract, 7.5 g/L agar) containing 10 g/mL ampicillin. Plates were incubated at 37°C in a standard atmosphere overnight. The next day colonies were picked and grown overnight at 37°C in LB media with 100 µg/mL ampicillin (prepared the same as plates but excluding agar). Bacterial cell lines were preserved by addition of sterile glycerol (1:1 media: glycerol) and subsequent storage at -80°C.

2.2.5 Expression and purification of recombinant Aprataxin from pTYB1: Overnight cultures of the wild-type and V263G Aprataxin containing pTYB1 constructs were inoculated into fresh LB media containing 100 µg/mL ampicillin and regrown to

OD600 0.5 prior to induction for 3 hours at 30°C with 0.5 mM isopropylthioglalctopyranoside (IPTG). All following steps were performed at 4°C unless stated otherwise. Cells were washed in Column buffer (20 mM Tris pH 8.0, 1 M NaCl, 1 mM EDTA, 0.3% glycine) and resuspended in Lysis buffer (20 mM Tris pH 8.0, 1 M NaCl, 1 mM EDTA, 0.3% glycine, 0.25% Triton X-100, 0.25% NP-40 and 20 µM PMSF). Cells were frozen at -70°C, defrosted, and sonicated for 5 minutes per 500 mL of original culture on a 50% cycle (2 seconds on, 2 seconds off) at maximum intensity (Branson Sonifier model S250A). The lysate was cleared by centrifugation at 20,000 x g for 30 minutes (Beckman JA-17 rotor).

The cleared lysate was applied to a column of chitin resin (New England Biolabs). Typically, 0.5 mL of packed resin bed was used per liter of induced wild-type culture, and 0.5 mL was used per four liters of induced V263G culture. The columns were washed with 10 (wild-type) and 20 (V263G) column volumes of Column buffer containing 0.25% Triton X-100 and 0.25% NP-40. Following this, the columns were washed briefly (3 volumes) with Column buffer containing 2 M NaCl. The V263G column was subsequently washed in 3 volumes of 25 mM Tris pH 7.5, 200 mM KCl, 3

mM MgCl2, 10% glycerol, and 100 µM ATP to remove contaminating heat-shock proteins (based on information provided in (7)). The columns were equilibrated in Cleavage buffer (20 mM Tris pH 8.0, 500 mM NaCl, 1 mM EDTA, 10% glycerol, 0.3% glycine, 0.25% Triton X-100, 0.25% NP-40, 50 mM dithiothreitol (DTT), and 25

70 mM oxidized cystine), clamped and incubated for 36 hours at 16°C. The columns were eluted using Cleavage buffer without DTT and cystine.

The eluted proteins were diluted to 50 mM NaCl, 2% glycerol, 0.06% glycine, 0.05% Triton X-100, 0.05% NP-40 and 10 mM Tris, pH 8.0 and loaded onto 200 µL of Biorex-70 resin (Biorad) equilibrated in this buffer. The columns were washed in 1 mL of 10 mM Tris pH 8.0, 66 mM NaCl and eluted initially in 10 mM Tris pH 8.0 500 mM NaCl. A second elution in 10 mM Tris pH 8.0 1 M NaCl was also collected.

Protein containing Biorex-70 fractions were concentrated using 10 kDa Nanosep centrifugal devices (Pall life sciences) according to the manufacturers instructions. Concentrates were diluted 1/3 in 10 mM Tris pH 8.0, giving a final buffer constitution of 10 mM Tris pH 8.0 and approximately 300 mM NaCl and 3% glycerol. Aliquots were stored at -80ºC.

2.2.6 Expression and purification of recombinant Aprataxin from pGEX 6.1: pGEX 6.1 constructs were transformed into BL-21 E.coli (Invitrogen) and induced as for pTYB1. The cell pellet produced from IPTG-induced cells was processed and cleared lysate generated as previously. Cleared lysate was incubated with glutathione sepharose (Amersham Biosciences) for 2 hours at 4ºC. Beads were then washed in 20 volumes of Column Buffer. The beads were then washed two bed volumes of Precision Protease Buffer (20 mM Tris pH 7.5, 200 mM NaCl, 1 mM EDTA, 1 mM DTT). The beads were then incubated in 1.5 volumes of Precision Protease Buffer with 2 µg of Precision Protease (Amersham Biosciences) overnight at 4ºC. The cleaved target proteins were then eluted in Precision Protease Buffer. Protein C was stored at -80°C. Proteins A and B were subjected to further purification.

Proteins A and C were diluted 1/5 in 10 mM Tris pH 7.5, giving a final NaCl concentration of 40 mM. These proteins were passed over an equilibrated Biorex-70 strong cation exchange/ size exclusion column. The column was then washed sequentially with 10 mM Tris pH 7.5 containing increasing concentration of NaCl (between 100 and 500 mM). The elution profiles were analyzed by SDS-PAGE. Pure fractions (between 350 and 500 mM NaCl) were pooled and concentrated using 10 kDa

71 Nanosep centrifugal devices (Pall life sciences) according to the manufacturers instructions. Purified proteins were then stored at -80ºC.

2.2.7 Purification of anti-Aprataxin specific antibodies: Affinity resins for purification of the rabbit Aprataxin antibody were generated as previously described (8). Briefly, pGEX 6.1 and pGEX 5.1 and 6.1 APTX constructs were transformed into BL-21 E.coli and induced with 0.5 mM IPTG for 3 hours at 37ºC prior to lysis by sonication as described earlier. Cleared lysates were incubated with glutathione sepharose (Amersham Biosciences), which was subsequently washed in 200 mM Borate-NaOH pH 8.6. To covalently link GST and GST-Aprataxin fusion proteins to the glutathione beads, they were resuspended in 0.2 M triethanolamine pH 8.3, 20 mM dimethyl pimelimidate-HCl (DMP) and incubated for 60 minutes at room temperature with gentle agitation. The covalent cross-linking was terminated by aspiration of the supernatant and addition of 0.2 M ethanolamine pH 8.2 to the beads for 60 minutes. Beads were washed in 0.1 M glycine pH 2.5 to remove unlinked molecules prior to re-equilibration in PBS. Purity of the proteins on the resin and efficiency of crosslinking were assessed by denaturation of samples of beads in 5x SDS-PAGE loading buffer followed by protein electrophoresis.

Anti-Aprataxin antibodies were generated by Dr Amanda Kijas (Queensland Institute for Medical Research, Brisbane, Australia) by inoculation of a rabbit and several sheep with full-length Aprataxin or Aprataxin domains (generated as described in Gueven et al., 9). Immune sera were cleared overnight at 4ºC with the GST-crosslinked glutathione sepharose generated above. Cleared sera were then incubated with the GST- Aprataxin beads for 6 hours at 4ºC. Non-specific binding was removed by washing the beads in TBS Tween-20 (0.05%). Specific antibodies were removed by elution with 100 mM glycine pH 2.5 and the pH was adjusted to 7.5 by addition of 1 M Tris (pH unadjusted). Sheep antibodies were dialysed against 10 mM Tris pH 7.5, 10 mg/mL BSA, 100 mM NaCl. The rabbit antibody was not dialysed as it was already rather dilute. The rabbit antibody was concentrated using a 10kDa Nanosep centrifugal device (Pall life sciences) according to the manufacturers instructions. Aliquots were stored at - 80ºC.

72 2.2.8 Estimation of protein concentration: Protein concentrations were determined using the Biorad Bradford Protein Assay. Briefly, 200 µL of 1/5 diluted Bradford Reagent was added the wells of a 96 well plate. 2 µL of protein containing sample, or BSA standards (0.5 to 10 mg/mL) were added to each well. The reactions were mixed by pipetting, and incubated at room temperature for 5 minutes prior to reading (595 nm) on a micro plate reader (VERSAmax micro plate reader, Molecular Devices) and data analysis using SOFTmax PRO software (Version 3.1.2, Molecular Devices).

2.2.8 Protein electrophoresis Denaturing protein electrophoresis was performed according to standard laboratory protocols (10). 5x SDS-PAGE buffer is: 100 mM Tris, pH 6.8, 2% SDS, 5% ß- mercaptoethanol, 15% glycerol, with bromophenol blue to assist loading. Gels were Coomassie stained by incubation in a solution of 50% methanol, 10% glacial acetic acid and 0.2% Brilliant blue R250. Gels were destained by incubation in the above solution lacking Brilliant blue. Colloidal Coomassie staining was performed by incubation of gels overnight in a solution containing 0.08% Coomassie Brilliant Blue G250, 1.6% ortho-phosphoric acid, 8% ammonium sulphate and 20% methanol. These gels were destained in water.

73 2.3 RESULTS

2.3.1 Purification of recombinant Aprataxin using pTYB1: The APTX cDNA encodes a 342 amino acid protein with a predicted molecular weight of 39.1kDa. This cDNA was cloned into the Intein-based expression system pTYB1 during this students honours project. In this system, the target protein is fused to an Intein, in which a Chitin Binding Domain facilitates affinity purification. After binding of the fusion protein to chitin resin and washing of the column, addition of thiols such as DTT and cystine are used to induce a conformational change in the Intein, resulting in its activation and subsequent self-cleavage of the target protein from the tag. A schematic of this process is shown in Figure 2.1.

Initial induction conditions were tested by growing and inducing small scale (10 mL) cultures, lysing the cells in 1 mL of Lysis Buffer, and incubating the cleared lysate with 50 µL of chitin beads for 10 minutes at 4°C. The beads were then pelleted in a microcentrifuge, and SDS-PAGE loading buffer was added directly to the pellet. The resulting denatured proteins were then resolved by 10% SDS-PAGE and analyzed by Coomassie staining, as shown in Figure 2.2. The predicted weight of the Aprataxin- Intein fusion protein is 94 kDa (the Intein tag is 55 kDa). Figure 2.2 shows that the predominant band for both the wild-type and V263G test inductions is approximately 100 kDa, indicating that the induction of wild-type and V263G Aprataxin protein expression was successful. Figure 2.2 also shows that the V263G Aprataxin protein has not been induced to the same extent as the wild-type protein (the same volume of bead slurry eluent was loaded). This is not unexpected as mutation of Aprataxin is known to destabilize the protein (11). This may be due to one or more factors, including misfolding of the protein or toxicity of the mutated protein to the cells, which may cause precipitation or degradation of the expressed protein. As these factors are largely not controllable, the reduced levels of expression of the V263G protein compared to wild-type Aprataxin were deemed acceptable.

74 Figure 2.1: Schematic representation of the pTYB1 affinity purification system. Expression of the fusion protein is achieved by addition of IPTG to log phase bacterial culture. Cleared lysate is passed over chitin affinity resin, and non- specific binding eliminated by washing. Cleavage of the target protein from the affinity tag is induced by incubation of the resin with reducing agents such as DTT and the target protein can be subsequently eluted.

Figure 2.2: Test induction of wild-type and V263G Aprataxin fusion protein expression. Wild-type and V263G pTYB1 constructs were induced in BL-21 E.coli cells as described in section 2.2.5. Lysates were subjected to batch affinity purification using chitin resin. Enriched proteins were eluted in 5x SDS-PAGE loading buffer (without β- mercaptoethanol) and subjected to SDS- PAGE and Coomassie staining.

75 Preparation of protein from the pTYB1 system requires reducing agent-induced cleavage of the target protein from the Intein tag. I analyzed the cleavage efficiency of wild-type and V263G Aprataxin proteins by SDS-PAGE (Figure 2.3). This involves taking a sample of the bead slurry after the cleavage reaction (described in section 2.2.5) and denaturing the proteins in SDS-PAGE loading buffer without reducing agents. This is crucial as the β-mecaptoethanol present in normal loading buffer induces efficient cleavage of the target from the tag, making assessment of cleavage reaction efficiency impossible (compare mutant protein bead eluants in Figure 2.3 where beads were not reduced before electrophoresis with Figure 2.4, where they were). Under the conditions outlined in section 2.2.5, the majority of the wild-type fusion protein is cleaved to produce separate Intein and Aprataxin proteins while cleavage of the V263G protein into Intein and Aprataxin fragments was much lessefficient (Figure 2.3).

100

50

40

Figure 2.3: Cleavage efficiency of Aprataxin- Intein fusion proteins. Wild-type and V263G induced lysates (2 mL) were bound to small volumes of Chitin beads (50 µL). These beads were washed in batch mode as described before initiation of the cleavage reaction. After the designated reaction period, 100 µL of 5 x SDS-PAGE loading buffer without β-mecaptoethanol was added directly to the cleavage reactions. This facilitated elution of the proteins from the beads without causing additional thiol- induced cleavage of the fusion proteins. The eluents were then subjected to 10% SDS- PAGE and Coomassie staining.

76 After this optimization of induction and cleavage conditions, large scale inductions of wild-type and V263G proteins were performed as in Materials and Methods. This revealed binding of a 70 kDa contaminant to the V263G Aprataxin protein column (data not shown). Given the mutation of the induced protein and its subsequent poor expression, this contaminant may have been the chaperone and degradative marker Heat Shock Protein 70 (HSP 70). The column washing conditions were therefore modified to include a larger wash volume for the V263G column (20 volumes instead of 10), and the inclusion of an ATP wash, which has been reported to remove HSP 70 from its interacting proteins (7).

As can be seen in Figure 2.4, this treatment resulted in the production of large quantities of nearly homogenous wild-type protein, and very small quantities of highly contaminated V263G protein. Extensive washing and the addition of an ATP wash did not eliminate contaminating proteins from the V263G Aprataxin preparation. Thus an additional chromatographic step was performed to try and reduce the level of contamination in the V263G fractions.

77

Figure 2.4. Affinity purification of wild-type and V263G recombinant Aprataxin from bacterial cell lysates. A, wild-type protein: lanes 1 to 9- fractions collected from chitin column after cleavage of the target protein from the affinity tag. Lane 10- sample of chitin resin after elution. B, V263G protein lanes 1 to 10- fractions collected from chitin column after cleavage of the target protein from the affinity tag. Lane 11- sample of chitin resin after elution. Protein is eluted from resin using 5x PAGE buffer. Molecular weights are shown in kDa.

Wild-type and V263G Aprataxin were subsequently subjected to further purification using the dual size exclusion/cation exchange resin Biorex 70 (Biorad) as in section 2.2.5. The bed volume used here (200 µL) is not sufficient for size exclusion chromatography. Therefore this aspect of Biorex 70 resin will be disregarded in the subsequent results and discussion.

78 Proteins were bound to the Biorex resin in a low salt buffer as described. These conditions allowed the binding of Aprataxin to the resin, as shown by comparison of the protein content of the chitin column eluent (Figure 2.5a lane 2) with that of the Biorex column flow through (Figure 2.5a lane 3). This shows the presence of high levels of wild-type Aprataxin in the chitin column eluent, which are largely absent in the Biorex column flow through, indicating efficient binding. The columns were washed and then eluted in a high salt buffer. As can be seen by the extremely efficient elution of the wild-type protein (Figure 2.5a), these conditions successfully interfere with the ionic interactions between the target protein and the resin. This procedure produced 625 µg of homogenous wild-type protein (Figure 2.6). Unfortunately, the V263G protein retained high levels of contamination, even after this additional chromatographic step (Figures 2.5 and 2.6). Complete elution of the V263G protein was unable to be achieved with either the chitin affinity column (Figure 2.4b) or the subsequent ion exchange column (Figure 2.5b). The yield for the production of V263G Aprataxin was very poor (approximately 25 µg of total protein from 8 L of induced culture). It was estimated (based on visual inspection of Figure 2.6) that approximately 75% of the protein recovered from the Biorex column was contamination, and V263G Aprataxin was only 25% of the total protein in this solution. This gives a yield of V263G Aprataxin of approximately 6 µg from 8 L, an efficiency 400 times less than that of the wild-type purification.

79

Figure 2.5. Ion exchange purification of wild-type and V263G Aprataxin. A and B) Lane 1- chitin resin after cleavage of the target protein, lane 2- eluents from the chitin column. The eluents were loaded onto cation exchange columns. Lane 3- flow through from the cation exchange columns, lane 4- wash of the cation exchange columns. Lanes 5 to 9- fractions collected from elution of the cation exchange columns using 10 mM Tris pH 7.5 buffered 500 mM NaCl. Lane 10- elution from the columns using 10 mM Tris pH 7.5 buffered 1 M NaCl. Lane 11- cation exchange resin post-elution. Proteins are eluted from resin using 5x SDS-PAGE loading buffer. Molecular weights are shown in kDa.

80 WT V263G ug BSA

1 0.1 0.2 1 0.5 0.2 0.1 0.2 0.5 1 2 3

60kD 50kD

40kD

30kD

Figure 2.6. Quantification of recombinant Aprataxin concentrations. Coomassie stained 12% SDS-PAGE of recombinant Aprataxin proteins. Volumes (in µL) of wild-type and mutant Aprataxin proteins loaded are indicated above each lane.

2.3.2 Generation of recombinant Aprataxin using pGEX 6.1:

Given the failure of the pTYB1 system to readily produce homogenous point-mutated Aprataxin, the GST based system pGEX 6.1 (which had used by other members of this laboratory to produce a wide variety of mutant and truncated proteins) was used to generated C and N-terminal deletions of Aprataxin. An overview of the purification method for GST-fusion protein purification is shown in Figure 2.7. The principal difference between this system and pTYB1 is that the cleavage step requires the addition of an exogenous protease (Prescision Protease), rather than activation of the built-in protein splicing element.

81

Figure 2.7: Overview of pGEX 6.1 expression and purification system. Induction of fusion protein expression is achieved by addition of IPTG to log phase bacterial cultures. Lysate is then passed over glutathione affinity resin and nonspecific binding eliminated by washing. Cleavage and subsequent elution of the target protein from the affinity resin is achieved by incubation of the resin with Prescission Protease followed by several washes.

In order to study the functional relationship between the domains of Aprataxin a series of pGEX 6.1 domain deletion constructs were created (Dr Amanda Kijas, Queensland Institute for Medical Research, Brisbane, Australia, described in (12). These consisted of the full length protein (amino acids 1 to 342, Protein A), a C- terminal truncation (amino acids 118-342, protein B), an N-terminal truncation (amino acids 1 to 319, Protein C) and a dual truncation (amino acids 118 to 319, Protein D), as shown in Figure 2.8.

82

Figure 2.8: Schematic of pGEX 6.1-APTX truncation constructs. A series of cDNAs coding for different Aprataxin regions were cloned into pGEX 6.1 by Dr Amanda Kijas (Queensland Institute for Medical Research, Brisbane, Australia). These constructs were designed to express the full length protein (Protein A, full length, wild-type), an N-terminal truncated protein which lacks the FHA domain (Protein B), a C-terminal truncated protein lacking the zinc finger (Protein C), and a dual N- and C-terminal truncated protein which lacks both FHA and zinc finger domains (Protein D).

These Aprataxin-pGEX 6.1 vectors were transformed into BL-21 E.coli and induced in the same manner as the pTYB1 constructs (method described in section 2.2.6). After lysis and clearing, the lysates were passed over glutathione columns and washed in Column buffer. Subsequent incubation of the columns with PreScision Protease resulted in release of proteins of the appropriate size from the GST tag, as shown by SDS-PAGE of the eluents (Figure 2.9). The doubly truncated Protein D did not appear to induce well (Figure 2.9). Attempts at purification of this protein were not continued.

Based on Figure 2.9, the eluents from Aprataxin Protein A and C columns contained low level contamination across a broard range of molecular weights. Therefore these eluents were subjected to cation exchange chromatography in a manner similar to that used for the pTYB1 expressed proteins (described in section 2.2.6).

83

Figure 2.9: Purification of recombinant Aprataxin using pGEX 6.1. Coomassie stained 10% SDS-polyacrylamide gel of recombinant proteins (1 or 4 μL as indicated) eluted from glutathione resin by precision protease cleavage. BSA standards (100 to 2000 ng) are shown. Proteins A, B, C and D refer to full length, N and C- terminal truncation mutants and a dual truncation mutant as indicated in Figure 2.8. Molecular weights are shown in kDa.

Figure 2.10 shows the binding of the proteins to the Biorex 70 resin as a high concentration of protein in the affinity column eluent compared to a low concentration in the Biorex 70 column flow through. A series of washes of the column were performed, each containing a higher NaCl concentration than the last (200 to 500 mM NaCl in 50 mM increments as described in section 2.2.6). Efficient elution of the Aprataxin proteins was observed between 300 and 500 mM NaCl-containing buffers (Figure 2.10). These fractions, in which the target proteins were nearly homogenous, were pooled and concentrated. For Proteins A and C the concentration of the final products were 2.64 and 1.76 mg/mL respectively, in a volume of approximately 1 mL. Protein B, which was not concentrated, had a final concentration of 2.06 mg/mL and an approximate volume of 2.5 mL. Thus purification of Aprataxin Proteins A, B and C was achieved with yields of approximately 2.64, 5.15 and 1.76 mg respectively. Figure 2.11 shows colloidal Coomassie staining of the final products.

84

Figure 2.10: Purification of recombinant Aprataxin proteins. After affinity purification, Aprataxin proteins A and C (full length and C-terminal truncated respectiveley) were eluted from glutathione sepherose (Figure 2.9) and subsequently subjected to purification on Biorex-70 dual strong cation exchange/ size exclusion resin. Samples of GST resin and GST eluent were analyzed to determine the efficiency of cleavage from the affinity column. Efficient binding of the target protein to the cation exchange resin is evident by the difference in protein content of the GST eluent and the cation exchange (Biorex) column flow through. Aprataxin proteins were successfully eluted by washing of the columns with buffers of increasing ionic strength (eluent lanes, NaCl concentration in 10 mM Tris pH 7.5 is indicated). The extent of elution from the cation exchange resin was examined by SDS-PAGE analysis of the beads post-elution (Biorex resin lanes). Molecular weights are shown in kDa.

85

A C B

Figure 2.11: Purified recombinant Aprataxin proteins. Purified recombinant Aprataxin proteins (50 pmol each of Proteins A B and C as indicated) were analyzed for contamination by colloidal Coomassie stained 12% SDS-PAGE gel. Molecular weights are shown in kDa. Proteins A, B and C are full length, N-terminal and C-terminal truncation proteins respectivley

2.3.3 Purification of Aprataxin-specific antibodies:

In addition to recombinant proteins, antibodies against proteins or domains of interest are important tools are essential for many biochemical studies. Antisera against full length Aprataxin and specific domains were generated by Dr Amanda Kijas (Queensland Institute for Medical Research, Brisbane, Australia, using pGEX 5.1 constructs described in Gueven et al., 9). Here the generation of affinity columns and subsequent affinity purification is described. The purification of sheep anti-FHA specific antibody is shown in detail; however the process is identical for all antibodies generated here. A summary of the antigens used for immunization by Dr Kijas and the proteins used for purification by myself is shown in Table 2.2.

86

Species Antigen Purified Specific to Against Rabbit Whole protein Protein C FHA and HIT (aa 1-342)-GST (aa 1-319) domains Sheep FHA (aa 1- FHA (aa 1-118) FHA domain 118) -GST Sheep Protein B (aa Protein C (aa 1- HIT domain 118- 342) -GST 319)

Table 2.2: Summary of immunization and purification details for Aprataxin-specific antibodies. Antisera were generated against different regions of the Aprataxin protein (as indicated by “Antigen”) by inoculation of laboratory animals (species as indicated) by Dr Amanda Kijas using recombinant proteins described in Gueven et al., 9). These were purified against Aprataxin region-GST fusion proteins to generate antibodies reactive with specific Aprataxin domains.

Initially competent BL-21 E.coli were transformed with a pGEX 5.1 vector containing the sequence coding for the FHA domain of Aprataxin (amino acids 1 to 118, described in Gueven et al. 9) fused to GST, or the empty vector alone (GST only). Induction was performed in an identical manner to the pGEX 6.1 constructs (section 2.2.6). Cells were grown to mid-log phase and a sample was taken before the initiation of induction by the addition of IPTG. After 3 hours of induction a second sample of cells were taken from both cultures to verify induction of the GST and GST-FHA proteins (Figure 2.12). After induction, the cell pellet was processed as described for the purification of GST proteins already, and the lysate passed over glutathione columns and washed, also as described. An important step in the purification of antibodies is the chemical crosslinking of the target protein to its affinity resin such that it does not ‘leak’ when the antibody is eluted. To achieve covalent attachment of the fusion proteins to the glutathione resin, the washed beads were treated with the crosslinking agent DMP. The efficiency of this reaction was measured by comparing a sample of the beads pre and post crosslinking by SDS- PAGE (Figure 2.12). The presence of large amounts of protein in the “before

87 crosslinking” lanes indicates successful binding of the fusion proteins and the absence of these bands in the “after crosslinking” lanes indicated successful crosslinking of the target proteins to the glutathione resin. After equilibration of the resin in PBS-Triton X-100 (0.5%), the columns were ready to use for purification of antibodies from immune sera. A Protein C crosslinked affinity column was prepared in the same manner (data not shown).

Figure 2.12: Generation of crosslinked antigen-GST resin for antibody purification. Induction of GST and GST-FHA proteins was monitored by addition of SDS-PAGE loading buffer to uninduced and induced cell pellets. After induction the expressed proteins were bound to GST resin, which was then treated with DMP to crosslink bound proteins to the matrix. The efficiency of this crosslinking reaction was determined by comparison of the proteins eluted from a sample of each resin when boiled in SDS- PAGE buffer.

As the antisera generated by Dr Kijas were raised against fusions of Aprataxin or its domains to GST, some of the antibodies present in the serum are likely to be GST- specific. To remove these antibodies, the sera were initially cleared of GST-reactive antibodies by overnight incubation with GST-crosslinked glutathione beads. The

88 cleared sera were then allowed to bind to their respective columns for several hours at 4°C before washing with PBS- Triton X-100 as described. Specific antibodies were then eluted by incubation of the columns with 0.1 M glycine (pH 2.5). The pH of the eluents was immediately neutralized with 1 M Tris (pH unadjusted) to prevent denaturation of the antibodies. Sheep antibodies were dialysed as described in section 2.2.7 and all antibodies were concentrated before storage at -80°C. SDS-PAGE analysis of the purified antibodies is shown in Figure 2.13.

Figure 2.13: Purified Aprataxin antibodies. Coomassie stained SDS-polyacrylamide gel of 5 μL of affinity purified Aprataxin antibodies. Molecular weights are shown in kDa.

89 2.4 DISCUSSION

This chapter has dealt with the production of recombinant proteins and purification of Aprataxin-specific antibodies, tools which will be essential for characterization of the cellular function of Aprataxin.

Several factors were considered when selecting bacterial expression systems, including yield (1), purity (2), robustness of the system (3) and cost (4). Based on these criteria, the initial selection of the pTYB1 expression system was acceptable for the production of wild-type Aprataxin but not for the V263G mutant. This system performed well in the expression and purification of wild-type protein, producing approximately 625 µg of homogenous protein with minimal need for optimization of induction and purification conditions. Conversely, the performance of this system at expression and purification of the V263G mutant Aprataxin protein was poor at all stages. Mutation of a single amino acid within the target protein resulted in a drastic reduction in the levels of induction. This is consistent with the very low levels of mutated Aprataxin proteins (9) and the reduced half life of these molecules (11) in mammalian cells. Poor induction of this protein was partially compensated for by inducing larger volumes of culture. During optimization of the purification of the V263G protein, it became apparent that the protein may be partially misfolded and aggregated. This was evidenced by low enzymatic activity of the Intein protein splicing element when fused to the V263G protein (Figure 2.3). When incubated with DTT and cystine the wild-type fusion protein is readily cleaved to produce Intein (55kDa) and Aprataxin (41kDa) bands. Under the same conditions cleavage of the mutant protein is very inefficient, indicating that the activity of the Intein is reduced when it is fused with the Aprataxin point mutant. This necessitated extensive optimization of cleavage conditions, including increases in cleavage time, the testing of different thiols such as cysteine and β-mercaptoethanol as substitutes for DTT to induce cleavage and the addition of detergents to reduce protein aggregation (data not shown). This process finally defined ‘optimal’ conditions for the purification of V263G Aprataxin in pTYB1, which resulted in a production of a very low yield of V263G protein which was highly contaminated even after secondary purification.

90 The poor performance of pTYB1 for purification an Aprataxin point mutant suggested that it may not be an appropriate system to use for generation of Aprataxin truncation mutants. Therefore the GST-based system pGEX 6.1 was used as an alternative. This system proved to be far more reliable and robust than pTYB1, requiring minimal optimization and producing several milligrams of homogenous protein for three out of the four proteins initially attempted.

The second set of tools generated here, antibodies affinity purified to react with various regions of Aprataxin will be used for a wide variety of experiments discussed in future chapters of this thesis. Optimization of the conditions for the use of each of these antibodies will therefore be explained when they are initially used in each experiment.

91 2.5 REFERENCES

1. Smith, D.B. and Johnson, K.S. (1988) Single-step purification of polypeptides expressed in Escherichia coli as fusions with glutathione S-transferase. Gene, 67, 31-40. 2. Ljungquist, C., Breitholtz, A., Brink-Nilsson, H., Moks, T., Uhlen, M. and Nilsson, B. (1989) Immobilization and affinity purification of recombinant proteins using histidine peptide fusions. Eur J Biochem, 186, 563-569. 3. Watanabe, T., Ito, Y., Yamada, T., Hashimoto, M., Sekine, S. and Tanaka, H. (1994) The roles of the C-terminal domain and type III domains of chitinase A1 from Bacillus circulans WL-12 in chitin degradation. J Bacteriol, 176, 4465- 4472. 4. Hopp, T., Pricket, K., Price, V., Libby, R., March, C., Ceretti, D., Urdal, D. and Conlon, P. (1988) A short polypeptide marker sequence useful for recombinant protein identification and purification. Bio/Technology, 6, 1204-1210. 5. Guise, A.D., West, S.M. and Chaudhuri, J.B. (1996) Protein folding in vivo and renaturation of recombinant proteins from inclusion bodies. Mol Biotechnol, 6, 53-64. 6. Harlow, E. and Lane, D. (1988) "Antibodies. A Laboratory Manual. Cold Spring Harbor Laboratory Press, ISBN 0-87969-314-2. 7. Burdon, R.H. (1986) Heat shock and the heat shock proteins. Biochem J, 240, 313-324. 8. Bar-Peled, M. and Raikhel, N.V. (1996) A method for isolation and purification of specific antibodies to a protein fused to the GST. Anal Biochem, 241, 140- 142. 9. Gueven, N., Becherel, O.J., Kijas, A.W., Chen, P., Howe, O., Rudolph, J.H., Gatti, R., Date, H., Onodera, O., Taucher-Scholz, G. et al. (2004) Aprataxin, a novel protein that protects against genotoxic stress. Hum Mol Genet, 13, 1081- 1093. 10. Maniatis, T. and Sambrook, J. (1989) Molecular Cloning: A Laboratory Manual (first edistion). Cold Spring Harbor Laboratory Press, ISBN: 0-87969-309-6. 11. Hirano, M., Asai, H., Kiriyama, T., Furiya, Y., Iwamoto, T., Nishiwaki, T., Yamamoto, A., Mori, T. and Ueno, S. (2007) Short half-lives of ataxia- associated aprataxin proteins in neuronal cells. Neurosci Lett, 419, 184-187.

92 12. Kijas, A.W., Harris, J.L., Harris, J.M. and Lavin, M.F. (2006) Aprataxin forms a discrete branch in the HIT (histidine triad) superfamily of proteins with both DNA/RNA binding and nucleotide hydrolase activities. J Biol Chem, 281, 13939-13948.

93 94

CHAPTER 3 Biochemical characterization of recombinant Aprataxin

95 96 3.1 INTRODUCTION

The eukaryotic genome in under constant stress from various sources. External sources of genotoxic stress include ionizing and ultra-violet radiation and chemical agents. Endogenous sources of genotoxic stress include products of oxidative metabolism, which can generate reactive oxygen species such as superoxide. This array of genotoxic insults can induce a broad range of changes to the molecular structure of DNA. These include the formation of single or double stranded DNA breaks, oxidation of the bases within the DNA molecule and the formation of DNA- protein adducts to name a few. Failure of a cell to correctly repair DNA damage can result in apoptosis or malignant transformation. Thus cells have developed a plethora of mechanisms to repair damaged DNA and maintain genomic integrity. A family of ARCAs is characterized by DNA repair defects, caused my mutations of genes which produce DNA repair proteins. AOA1, caused by mutation of APTX, is one such disorder.

APTX mutations found in AOA1 patient cause deficiency of the protein product, Aprataxin (1,2). Cell lines from AOA1 patients are hypersensitive to a range of DNA damaging agents including hydrogen peroxide, MMS and ionizing radiation (3-5). This provided initial evidence that Aprataxin was involved in cell survival after DNA damage.

3.1.1 Domain structure of Aprataxin Aprataxin is 342 amino acid protein which contains three functional domains; an N- terminal Forkhead Associated (FHA) domain, a central Histidine triad (HIT) domain

and a C-terminal C2H2 zinc finger. The functions of these domains in general and in regards to Aprataxin have been detailed in section 1.2. Briefly, FHA domains are phosphopeptide interaction motifs (6). The FHA domain of Aprataxin interacts with the DNA repair proteins XRCC1 and XRCC4 (3-5). These proteins are involved in single and double strand break repair respectively (7,8). The sensitivity of AOA1 cells combined with this interaction data indicated that Aprataxin may be involved in single and double strand break repair. To elucidate the function of Aprataxin in DNA repair, over the last several years this study and others have characterized the function of Aprataxin’s HIT and zinc finger domains.

97

C2H2 zinc fingers are DNA binding motifs which is stabilized by chelation of a zinc ion (9). Some C2H2 zinc fingers are capable of binding to RNA (10-12) and this domain has also been implicated in protein-protein interactions (13). At the commencement of this study the in vitro function of Aprataxin zinc finger was not known. Therefore I describe here a characterization of Aprataxin’s interactions with polynucleotides through this domain.

HIT domains are characterized by the consensus sequence HαHαHαα (where α is a hydrophobic amino acid, reference 14). Although the in vivo function of HIT domains is not known, they do confer nucleotide hydrolase activity in vitro (14). HIT proteins are classified into sub-families (Hint, Fhit, GalT and DcpS) based on protein sequence relationships and subsequent differences in substrate preference. At the commencement of this study previous characterizations of Aprataxin’s enzymatic activity attempted to classify it as either a Hint or Fhit type adenosine derivative nucleotide hydrolase (15,16). The enzymatic activities reported by these studies were low compared to activities reported for Hint or Fhit on the same substrates (reviewed in section 1.2). Furthermore, estimations of Aprataxin’s activity by different laboratories against the same substrate produced variable results (15-17). Therefore this study has examined the activity of Aprataxin against both Hint and Fhit type substrates. I also examined Aprataxin’s activity against a newly identified DNA repair intermediate (18), 5’ adenylated DNA and examine the functional interactions between Aprataxin’s domains.

98

3.2 MATERIALS AND METHODS

3.2.1 Partial chymotryptic proteolysis: Bacterial or yeast recombinant Aprataxin (17 pmol) was incubated at 30°C in 20 mM HEPES pH 7.0, 50 mM NaCl, 8% sucrose, 10 ng/µL acetylated BSA in a total volume of 41 µL, in the presence (or absence) of various concentrations AMP, ATP,

AMPNH2, single and double stranded DNA as indicated for 5 to 10 minutes to allow binding. Yeast recombinant Aprataxin was supplied by Dr Amanda Kijas, and its generation is described in Kijas et al. (17). After binding, 5 ng of chymotrypsin (unless stated otherwise) was added to begin proteolysis. Reactions were terminated after the indicated times by removing a 10 µL sample of the reaction into 5uL of 5xSDS-PAGE loading dye and freezing on dry ice.

Reactions were resolved on 15% resolving 5% stacking Tris-glycine based gels (29:1 acrylamide: bisacrylamide), and transferred onto nitrocellulose membranes using standard conditions for the transfer of high molecular weight proteins (6 g/L Tris base, 3 g/L glycine, 0.36 g/L SDS, 20% methanol, 100 Vhrs).

3.2.2 Electrophoretic mobility shift assay (EMSA): The EMSA, or gel retardation assay, is a commonly used technique to examine binding of proteins to RNA and DNA. Although earlier examples of the EMSA can be found in the literature, the experiments shown here, and indeed most of those performed today, are based on the principle described by Garner et al. in 1981 (19). Single stranded synthetic DNA oligonucleotides (Table 3.1) were 5’ labeled with P-32 using γP-32 ATP and T4 Polynucleotide Kinase (PNK, Invitrogen) in the ‘exchange’ reaction as per manufacturers’ instructions. Double stranded DNA was generated by mixing the appropriate strands in equimolar amounts and heating them to 80-90°C in a dry heat block, before cooling over several hours. Labeled single stranded DNA was generated in a similar manner, with mock addition of an opposite strand. Radiolabelled DNA was diluted into unlabeled DNA of the same type to yield a final concentration of 5 pmol/µL with a specific activity of approximately 100 counts/pmol/sec.

99

homoduplex 5' ATGTGAATCAGTATGGTTCCTATCTGCTGAAGGAAAT 5' ATTTCCTTCAGCAGATAGGAACCATACTGATTCACAT mismatch 5' ATGTGAATCAGTATGGTTCCTATCTGCTGAAGGAAAT 5' ATTTCCTTCAGCAGATGGGAACCATACTGATTCACAT nick 5' ATGTGAATCAGTATGGTTCCTATCTGCTGAAGGAAAT 5' ATTTCCTTCAGCAGAT 5' GGGAACCATACTGATTCACAT gap 5' ATGTGAATCAGTATGGTTCCTATCTGCTGAAGGAAAT 5' ATTTCCTTCAGCAGAT 5' GGGAACCATACTGATTCACA Table 3.1: Oligonucleotide sequences used to generate EMSA substrates.

Recombinant Aprataxin produced as detailed in section 2.3.1, was incubated with up to 20 pmol of DNA in a 12 µL reaction in native loading buffer (20 mM HEPES pH 7.0, 50 mM NaCl, 8% sucrose, 10 ng/µL acetylated BSA). Amounts of recombinant protein are indicated on each lane. Reactions were prepared by adding protein to native loading buffer and were initiated by the addition of radiolabelled DNA. Reactions were incubated for at least 5 minutes at 4°C (unless stated otherwise) to allow the binding to reach equilibrium.

Subsequently, the equilibrated binding reactions were resolved by native polyacrylamide gel electrophoresis on a Biorad Maxi gel rig. Gels were generally between 5% and 8% 29:1 acrylamide: bisacrylamide, made and run on 0.5xTBE (1x TBE being 54 g/L Tris base, 27.5 g/L boric acid and 10 mM EDTA). 10 µL out of each 12 µL reaction was loaded onto the cool (10°C or less) pre-run gels, and resolved by electrophoresis at 20 mA for 30 to 50 minutes. The gels were dried onto 3MM Whattman paper using a Biorad gel dryer and exposed to x-ray film (Fujifilm) or a phosphor imaging screen (Amerhsam Biosciences and Molecular Dynamics). Binding was quantified based on loss of the unshifted band (i.e. the intensity of the unshifted band in each protein containing lane compared to the no protein control). This allows quantification of unstable binding which does not produce a discrete shifted band.

3.2.3 Nucleotide Hydrolysis: Stock solutions of adenosine and adenosine derivatives (ATP, ADP, AMP, AppppA, and AMPNH2) were made by preparing approximately 2 mM solutions and

100 measuring the absorbance at 259 nm in 10 mM Tris at pH 8.0. Accurate concentrations were calculated from these readings. Serial dilution of these generated stocks, which were stored in small aliquots at -70°C.

A Hypersil-120 strong anion exchange column (Phenomenex), attached to a Hewlett Packard series 1050 quaternary pump with a 200 µL injection loop, was used to resolve the components of nucleotide mixtures, and a Hewlett Packard 1040A HPLC detection system set at 259 nm was used for detection. Initial conditions recommended by the column supplier were used for resolving nucleotides were 100% 5 mM Ammonium Phosphate pH 2.8 to 750 mM Ammonium Phosphate pH 3.7 over 30 minutes. These conditions were not adequate for the resolution of AMP

from AMPNH2. The buffer composition and running conditions were optimised to

resolve AMP and AMPNH2. Optimised buffer compositions are Buffer A (2 mM ammonium phosphate at pH 2.8), and Buffer B (750 mM ammonium phosphate at pH 3.7). Running conditions for the resolution of nucleotides are 0 to 5% Buffer B over 15 minutes, followed by 5 to 100% Buffer B over the next 15 minutes at a pump speed of 1 mL per minute.

A 10 minute single step gradient from 0 to 100% Buffer B at 1 mL per minute was used for running AMP standards. Standards were generated by making several dilutions of stock AMP solutions, such that 20 µL of diluted stock contained 50 through to 19750 picomoles of AMP. 200 µL of Buffer A was added to 20 µL of each of these standard solutions. This mixture was loaded into a 200 µL injection loop, resulting in a consistent 10% loss before column injection for all reactions and standards.

Nucleotide hydrolase activity was analysed using 20 µL reactions containing 15 pmol of recombinant Aprataxin (expressed from pTYB1) in 10 mM HEPES pH 7, 30 mM NaCl and the indicated concentrations of each substrate (minimum 10 µM, maximum 1500 µM). Reactions were incubated at 37°C for 20 (AppppA or 60 minutes (AMPNH2). The effect of duplex DNA on hydrolase activity was examined by incubation of increasing concentrations of homoduplex (0 to 10 nM, generated by annealing the first two oligonucleotides in Table 3.1) with Aprataxin and 400 µM diadenosine tetraphosphate in the reaction conditions described above. Reactions were incubated for 40 minutes at 37°C. Detailed examination of the effect of DNA

101 on nucleotide hydrolysis was performed by incubation of Aprataxin (15 pmol) with increasing concentrations of diadenosine tetraphosphate (as indicated) in the presence of 10 nM duplex DNA using the same reaction conditions as previously. All reactions were terminated after the appropriate times by addition of 200 µL of Buffer A and frozen immediately. All reactions were performed in duplicate and all experimental data is shown in the appropriate figures.

The program Chemstation (Agilent Technologies) was used to identify and integrate the peaks corresponding to each compound. The areas under the curve, as calculated by Chemstation were used to generate a standard curve for picomoles of AMP versus milli Absorbance Unit seconds. The resulting equation describing the linear relationship between moles loaded and peak area was used to calculate the quantities of substrate and product in all hydrolase reactions.

3.2.4 Single strand break repair by cell extracts:

3.2.4.1 Generation of substrates:

Duplexes containing a single strand nick with a modified or unmodified 3’ terminus were generated by radiolabelling Oligo B (3’ strand) as described previously and annealing it to Ligase Bottom 36-mer and the 5’ strand (Oligo A) (Table 3.2). In this instance the radiolabel and the modified terminus are on opposite ends of the same oligonucleotide.

Ligase Bottom 36mer 5' TGTAGTCACTATCGGAATGAGGGCGACACGGATATG Oligo A 5' CCCTCATTCCGATAGTGACTACA Oligo B (unmodified) 5' CATATCCGTGTCG Oligo B (8-oxo-dG) 5' CATATCCGTGTC(8-oxo-dG) Oligo B (abasic) 5' CATATCCGTGTC(abasic site) Oligo B (phosphate) 5' CATATCCGTGTCG(phosphate) Table 3.2: Oligonucleotides used to construct SSBR duplexes.

3.2.4.2 Generation of nuclear extracts: The cell lines L938 and L939 were derived from the peripheral blood of male Japanese AOA1 patients and the control cell lines C2ABR and C3ABR were derived from the peripheral blood of healthy Caucasian males (thanks to Aine Farrel, Queensland Institute for Medical Research, Brisbane, Australia). Both of the

102 AOA1 patients have substitution mutations within the HIT domain (L938- P206L/P206L and L939- P206L/V263G).

Nuclear extracts were generated as published by Audebert et al. (20). Lymphoblastoid cells were grown in RPMI 1640 containing 6% FCS. All steps were performed at 4ºC. 2-3 x 108 cells were pelleted by centrifugation for 5 min at 1,500 x g. The cell pellet was washed twice with PBS and resuspended in 2 to 5 mL of Hypotonic Buffer (10 mM Tris pH 7.5, 10 mM KCl, 10 mM MgCl2, 1 mM DTT, 1 mM PMSF and 1 x complete inhibitor) and incubated on ice for 15 min. Cell membranes were broken by dounce homogenization (15 strokes), and the nuclei pelleted at 2,000 x g for 3 min. Nuclei were resuspended in 0.5 to 2 mL of

Extraction Buffer (10 mM Tris pH 7.5, 10 mM MgCl2, 450 mM KCl, 1 mM DTT, 1 mM PMSF and 1 x complete inhibitor) and incubated on a rotating wheel for 45 minutes. Lysate was cleared by centrifugation at 16,100 x g for 10 minutes and the supernatant was precipitated by addition of 0.313 g of ammonium sulphate per mL, neutralized with 3.13 µL NaOH per mL and incubated on a wheel for 30 minutes. Precipitated proteins were recovered by centrifugation at 16,100 x g for 20 minutes, resuspended in Dialysis Buffer (50 mM Tris pH 7.5, 1 mM EDTA, 13% glycerol, 0.1 M potassium glutamate and 1 mM DTT) and incubated on a rotating wheel for 30 minutes prior to dialysis against Dialysis Buffer. Extract was then centrifuged at 16,100 x g for 10 minutes and the supernatant stored in aliquots at -80ºC. Protein concentration was determined by Bradford assay.

3.2.4.3 Experimental conditions: 30 µg of C2ABR (control) lymphoblastiod cell extract was incubated with 2 pmol of each DNA substrate in Ligase Buffer (50 mM Tris pH 7.5, 25 µg/mL BSA, 10 mM

MgCl2, 0.5 mM DTT, 1 mM ATP) in a 10 µL volume at room temperature for the indicated times. Reactions were terminated by addition of 20 µL of formamide- EDTA denaturing gel loading buffer (50 mM EDTA in formamide, with bromophenol blue to assist loading). Complete denaturation was ensured by boiling the samples (80°C for 5 minutes) before snap-cooling.

103 3.2.4.4 Denaturing Electrophoresis: All DNA repair reactions were analysed by electrophoresis on 19% acrylamide, 1% bisacrylamide, 7 M urea, 1 x TBE sequencing gels (Biorad Sequigen gel rig, 21cm x 50 cm plates, 0.2 mm spacers and combs). Gels were run on 1 x TBE (anodic buffer) and 0.5 x TBE (cathodic buffer) at 50 W for at least 45 minutes to preheat the apparatus and remove APS from the gel. Samples were subsequently resolved at 50 W for 1 to 2 hours. Gels were transferred onto Whattman 3MM paper and dried on a Biorad gel drier (50°C for 2-3 hours and allowed to cool before removing vacuum). Where relevant, gels were used to expose a Molecular Dynamics phosphorscreen and reactions were quantified in ImageQuant 5.1. Specifically, band intensities were determined by integration and the substrate: product ratio was determined.

3.2.5 Single strand break repair by recombinant ligase: 3.2.5.1 Generation of substrates: The substrates used in this experiment were generated similarly to those in section 3.2.4.1, except that here the 3’ modification and radiolabel are on different oligonucleotides. Here Oligo A (5’ strand) was radiolabelled as described previously and annealed to Ligase Bottom 36mer and Oligo B (3’ strand). This is so that I can measure adenylation of the 5’ break terminus.

3.2.5.2 Experimental conditions: 2 pmol of each substrate was incubated with 1 unit of T4 DNA ligase (NEB) in Ligase Buffer at room temperature for the indicated times before termination of the reaction. Reactions were analysed by denaturing PAGE.

3.2.6 Adenylation of DNA by T4 DNA ligase: Adenylated DNA was generated similarly to as previously reported(18). Oligonucleotide sequences are indicated in Table 3.3. The oligonucleotide “5’ 18- mer” was radiolabelled as previously described. This was then annealed with equal amounts of the other oligonucleotides indicated in Table 3.3. 200 pmol of the resulting duplex was incubated with 50 units of T4 DNA ligase in Ligase Buffer for 24 hours at room temperature. The abortive ligation reaction was terminated by boiling for 20 minutes at 80°C. Duplex was re-formed by allowing the tube to cool

104 over several hours. In all following sections, this adenylated duplex is referred to as Duplex A. Successful adenylation was monitored by denaturing page as described previously, by comparing a sample of this reaction to a ‘no ligase’ mock reaction.

Ligase Bottom 36mer 5' TGTAGTCACTATCGGAATGAGGGCGACACGGATATG 5' 18-mer 5' ATTCCTATAGTGACTACA 3' 18-mer (dideoxy) 5' CATATCCGTGTCGCCCTC(dideoxy) Table 3.3: Oligonucleotides used to generate 5’ adenylated DNA.

3.2.7 Binding of Aprataxin to adenylated DNA: Binding of Aprataxin to Duplex A was examined as previously described (section 3.2.2), except that here the specificity of binding was also confirmed by antibody supershift. Recombinant full length Aprataxin (up to 3 pmol of protein expressed from pGEX6.1, as indicated) was incubated with 200 µg of rabbit null serum or 200 µg of affinity-purified rabbit α-Aprataxin antibody for 20 minutes at 4°C before addition of 2 pmol of Duplex A. Null serum is an appropriate control for non- specific antibody-protein interactions. Reactions were allowed to equilibrate for 5 minutes on ice before native protein electrophoresis as described in section 3.2.2.

3.2.8 Hydrolysis of adenylated DNA by recombinant Aprataxin: Where indicated, recombinant Aprataxin (200 fmol) was incubated with 200 µg of rabbit null serum or 200 µg of purified rabbit α-Aprataxin in Ligase Buffer without ATP for 20 minutes at 4°C. Hydrolase reactions were performed with the indicated amounts of Aprataxin (full length or truncated as indicated) and 2 pmol of Duplex A in 10 µL final volume at room temperature in Ligase Buffer. Reactions were terminated after the indicated times by boiling with formamide-EDTA loading buffer, and electrophoresis and reaction analysis was performed as described previously (section 3.2.3.4).

3.2.9 3’ Phosphatase activity assays: 3.2.9.1 Generation of substrates: Radiolabelled nicked duplexes with either 3’ hydroxyl or 3’ phosphate were generated using established 5’ radiolabelling and annealing protocols. Oligonucleotide sequences used for this experiment are shown in Table 3.4. Briefly,

105 Oligo B was radiolabelled and annealed to the other two strands to generate a nicked duplex with a 3’ phosphate or hydroxyl nick. Ligase Bottom 36mer 5' TGTAGTCACTATCGGAATGAGGGCGACACGGATATG Oligo A 5' CCCTCATTCCGATAGTGACTACA Oligo B (phosphate) 5' CATATCCGTGTCG(phosphate) Oligo B (unmodified) 5' CATATCCGTGTCG Table 3.4: Oligonucleotides used for phosphatase assays.

3.2.9.2 Experimental conditions: To examine the 3’ phosphatase activity of recombinant Aprataxin, the indicated quantities of full length Aprataxin (made using pGEX 6.1) were incubated with 1 pmol of 3’ phosphorylated duplex in Ligase Buffer (total volume 10 µL) at room temperature for 75 minutes. As a positive control for Aprataxin activity, Aprataxin was incubated in parallel with 1 pmol of Duplex A. Reactions were terminated and resolved as described previously. The unmodified (3’ hydroxyl) duplex was incubated and resolved in parallel with these samples as a control for successful resolution of 3’ phosphate and 3’ hydroxyl oligonucleotides.

The 3’ phosphatase activity of nuclear extract was examined in a similar fashion. Extracts were generated from control (C2ABR and C3ABR) and AOA1 (L938 and L939) lymphoblastiod cell lines as described (section 3.2.3.2). Extracts (amounts indicated) were incubated with the 1 pmol of the 3’ phosphate modified duplex in 10 µL reactions for 10 minutes at room temperature. Reactions were terminated and resolved as previously described.

106

3.3 RESULTS

3.3.1 Characterization of the binding activities of Aprataxin: Initially bacterial recombinant Aprataxin (expressed from pTYB1) was subjected to partial chymotryptic digestion in the presence or absence of nucleotide-based molecules. Partial proteolytic digestion is one technique for measurement of conformational changes in protein structure and can be used to identify movement in specific folds of the protein of interest by mapping exposed protease sites. Additionally, protein interaction with a ligand/substrate can be detected by alteration of the banding pattern when partially protease digested. At this stage identification of any of the proteolytic fragments generated is not required; observation of alterations in the banding pattern is sufficient to demonstrate an interaction between the protein of interest and its ligand.

Incubation of recombinant Aprataxin with increasing concentrations of the Hint protein family substrate AMPNH2 or AMP (the leaving group for HIT type hydrolase reactions) resulted in several alterations to the chymotryptic banding pattern of Aprataxin (Figure 3.1). These alterations were more marked at the higher concentrations of ligand, providing a crude indication of dose dependant binding. Binding of Aprataxin to dsDNA was also observed as an alteration of the chymotryptic banding pattern of the protein compared to Aprataxin with no DNA (Figure 3.1). It is interesting to note that the banding patterns of Aprataxin with AMPNH2 and AMP are similar. This is anticipated as both molecules are of a similar size and are expected bind to the same site on Aprataxin. Conversely, the banding pattern of Aprataxin incubated with dsDNA is different from those observed for AMPNH2 and AMP. This may indicate that DNA and the mononucleotides bind to different regions of the Aprataxin protein.

107 full length Aprataxin

1 2 3 4 5 6 7 8 9 10 11

Figure 3.1: Partial Chymotryptic proteolysis of bacterial recombinant Aprataxin- (poly)nucleotide complexes. 20μL reactions ran for 15 minutes. Lane 1- Aprataxin incubated with chymotrypsin in the absence of any additional chemicals. Lanes 2 to 4-

Aprataxin pre-incubated with 0.2 mM, 0.5 mM, and 0.8 mM of AMPNH2, and partially digested with chymotrypsin. Lanes 5 to 7- Aprataxin pre-incubated with 0.4 mM, 0.8 mM, and 1.6 mM AMP respectively prior to the addition of chymotrypsin. Lanes 9 to 10- Aprataxin pre-incubated with 2 mM, 4 mM, and 8 mM oligonucleotide dsDNA prior to chymotryptic digestion. Lane 10- Aprataxin incubated in parallel with the other reactions, but lacking additional nucleotides or chymotrypsin.

It was noted that recombinant Aprataxin produced in pTYB1, although homogenous on the Coomassie stained gels in Chapter 2, has some low molecular weight contamination when Western blotted with rabbit anti-Aprataxin serum (Figure 1, far right lane). This made it difficult to determine which low molecular weight bands were due to low molecular weight contamination and which were due to genuine proteolysis of the full length protein. Additional partial chymotryptic digestion experiments were therefore performed using recombinant Aprataxin expressed in yeast kindly provided by Dr Amanda Kijas (17). As the concentration of protease is critical in experiments of this nature, the quantity of chymotrypsin required to show high discrimination between protein with and without DNA bound was optimized. It was determined that 2 ng of

108 chymotrypsin with 17 pmol of Aprataxin provided an optimal amount of proteolysis to demonstrate the binding of DNA (Figure 3.2). This concentration was used in all following experiments.

1 2 3 4 5 6 7 8 9

Figure 3.2: Optimisation of chymotryptic proteolysis of yeast recombinant Aprataxin in the presence of DNA. Lane 1- Yeast recombinant Aprataxin without chymotrypsin treatment, in the absence of added nucleotides. Lanes 2 and 3- yeast recombinant Aprataxin pre-incubated with (lane 3) and without (lane 2) oligonucleotide dsDNA prior to the addition of 2 ng of chymotrypsin. Lanes 4 and 5- yeast recombinant Aprataxin pre-incubated with (lane 5) and without (lane 4) oligonucleotide dsDNA prior to the addition of 0.2 ng of chymotrypsin. Lanes 6 and 7- yeast recombinant Aprataxin pre-incubated with (lane 7) and without (lane 6) oligonucleotide dsDNA prior to the addition of 0.1 ng of chymotrypsin. Lanes 8 and 9- yeast recombinant Aprataxin pre-incubated with (lane 9) and without (lane 8) oligonucleotide dsDNA prior to the addition of 0.02 ng of chymotrypsin.

Subsequently, a range of nucleotide based molecules were tested for their ability to bind to Aprataxin (Figure 3.3). In all instances where Aprataxin was incubated with a

nucleotide based molecule (single or double stranded DNA, AMP or AMPNH2) prior to proteolysis, formation of an intense 20 kDa band was observed. This band is only weakly evident in the no substrate control (left lane), indicating that it is caused by exposure of a ‘hidden’ chymotrypsin site upon incubation of these molecules with Aprataxin. This provided initial evidence that recombinant Aprataxin was capable of binding both DNA and potential nucleotide based substrates.

109 1 2 3 4 5 6 7 Figure 3.3: Comparison of Aprataxin conformational changes in the presence of various (poly)nucleotides. Lane 1 - Aprataxin incubated with chymotrypsin in the absence of additional nucleotides. Lanes 2 and 3- Aprataxin pre-incubated with 1 mM oligo ssDNA and ds DNA prior to chymotryptic digestion. Lanes 4 to 6- Aprataxin pre- incubated with 0.5 mM AMP, 1 mM AMPNH2, and 1 mM AppppA respectively prior to the addition of chymotrypsin. Lane 7- Aprataxin incubated in the absence of chymotrypsin or additional nucleotides.

This observed binding of recombinant Aprataxin to DNA was further substantiated by EMSA, a widely used technique for studying the interaction between proteins and DNA or RNA molecules. A DNA or RNA molecule is labeled, generally with a radionuclide (however fluorescent tags are increasing in popularity) and incubated with the protein of interest. Native gel electrophoresis of the subsequent complexes can be performed using either polyacrylamide or agarose gels, depending on the expected size of the resulting complex. Nucleic acid- protein complexes have a higher mass/charge ratio than the respective unbound molecules, therefore migration of the complexes is retarded compared to the unbound nucleic acids. This resolves nucleic acids which are not protein bound from those which are. Following electrophoresis, nucleic acids can be detected by the normal autoradiographic techniques.

Figure 3.4 shows the binding of wild-type Aprataxin to a 36 base pair DNA duplex as retarded migration of the radiolabelled species. A commonly used description of the

110 affinity of a protein for DNA is the amount of protein required to bind to 50% of a set amount of DNA. Based on Figure 3.4, it was determined that 1 pmol of Aprataxin was required to bind to 50% of the 1 pmol of DNA in the reaction. Thus the Kd of Aprataxin for this duplex is 1.0. Only three the first three lanes of Figure 3.4 were used for this quantification because at higher ratios of protein: DNA multimeric complexes (two and three Aprataxin molecules/ DNA molecule) are observed, and examination of cooperative versus non-cooperative binding was not the aim of this study.

It is notable that at different concentrations of Aprataxin protein, three distinct protein- bound DNA species are observed (Figure 3.4). These most likely correspond to one, two, and three molecules of Aprataxin protein bound to the same molecule of DNA. This provides a rough estimation of the minimum size of the DNA Aprataxin can bind. As three molecules of Aprataxin can fit on a 37 bp duplex, the footprint of Aprataxin on double stranded DNA is approximately 12 base pairs or less. This is consistent with the literature at the time (17).

V263G mutant Aprataxin does not bind to duplex DNA (Figure 3.4). Increasing concentrations of this mutant recombinant protein failed to alter the electrophoretic mobility of the radiolabelled duplex used. This protein contains a single mutation in the HIT domain, and its failure to bind DNA provides initial evidence that the HIT domain may be involved in Aprataxins protein-DNA interaction. One limitation of this study was the inability to generate homogenous V263G recombinant aprataxin protein. As demonstrated in the previous chapter (Figure 2.6), this preparation is highly contaminated with what appears to be heat-shock protein. It is possible that these contaminants may interferre with any activities the mutant protein would otherwise display, however the removal of heat-shock proteins from this preparation was not achieved. Thus although this provides initial evidence that the HIT domain interacts with DNA, this must be substantiated by other means.

111

Figure 3.4: Binding of recombinant Aprataxin proteins to dsDNA. A. P-32 labeled homoduplex DNA was incubated with increasing amounts of bacterial recombinant wild-type and V263G Aprataxin, as indicated, in a total volume of 12 µL on ice prior to sample resolution and film exposure. B. The gel shown in A was used to expose a Molecular Dynamics phosphor screen and subjected to quantitative analysis to determine the proportion of free and protein-bound DNA at various wild-type Aprataxin concentrations.

112 Given that Aprataxin had been implicated in DNA repair both by the sensitivity of AOA1 patient cell lines to DNA damaging agents and by the proteins it interacts with, it was of interest to determine the relative affinity of Aprataxin to various DNA repair intermediates. A variety of DNA structures which mimic different types of DNA damage were generated. This was achieved by annealing one oligonucleotide with a complementary one to generate homoduplex, two complementary adjacent ones to generate a nick, or one oligonucleotide which is complimentary except for one base change to generate a mismatch and mock addition of an opposite strand to generate single stranded DNA (sequences are described in Table 3.1). These structures were then incubated with recombinant Aprataxin as previously and subjected to native gel electrophoresis to assess complex formation (Figure 3.5).

Binding of recombinant Aprataxin to homoduplex is observed as formation of a band of retarded electrophoretic mobility (Figure 3.5). Binding of Aprataxin to nicked, mismatched, and single stranded DNA is also demonstrated. Aprataxin appears to have a higher affinity for nicked DNA than the corresponding homoduplex (Figure 3.5). This implies that Aprataxin may have a higher affinity for DNA termini than for the central region of a strand. Aprataxin also binds to single stranded DNA, however a large amount of smearing is evident in these lanes. This indicates that the binding of Aprataxin with single stranded DNA is fairly transient, as these complexes dissociate during electrophoresis.

113

Figure 3.5: Binding of Aprataxin to different DNA structures. Various P-32 labelled DNA structures (1.0 pmol per reaction) were incubated with up to 4 pmol of recombinant bacterial recombinant Aprataxin (as described in Methods) to assess Aprataxin’s ability to bind these different structures. The types of structures are indicated above the lanes. Homoduplex indicates two complimentary 37 base primers annealed. Nick indicates a 16 and 21 base oligonucleotides annealed with a complimentary 37 base oligonucleotide. Mismatch indicates duplex formed by two 37 base primers which are complimentary except for a single internal T-G mismatch. Single strand indicates a 37 base single stranded oligonucleotide. Samples were resolved on a native TBE gel and exposed to a Molecular Dynamics phospor screen.

3.3.2 Characterization of the Aprataxin’s nucleotide hydrolase activity: The HIT domain of Aprataxin has homology to the HIT domains of both Hint and Fhit type hydrolases, therefore both Hint and Fhit substrates (AMPNH2 and AppppA) were tested as potential Aprataxin substrates. Initially, conditions were established to resolve the likely products. Strong anion exchange HPLC, which is generally appropriate for chromatographic separation of small nucleotide species, was selected for this. Optimization of chromatography protocols was performed as part of this student honors project. The optimal protocol is described in section 3.2.3. An example of resolution of adenosine based nucleotides using this protocol is shown in Figure 3.6. The detector

114 was calibrated by resolving known amounts of AMP and analyzing the resulting HPLC trace. The integrated peak areas for different amounts of AMP and the resulting standard curve are shown (Figure 3.7).

Figure 3.6: Resolution of adenosine derivatives by strong anion exchange HPLC. The indicated nucleotides were mixed prior to loading onto the HPLC column and resolved by a gradient of increasing phosphate concentration and increasing pH as described in methods.

115

Figure 3.7: Calibration of HPLC detector. AMP, in the quantities indicated, was loaded onto the HPLC column and resolved as described in methods. AMP peaks were quantified by ChemStation (raw data shown in A). Calculated peak area was plotted against picomoles loaded (shown in B) and data was analyzed by linear regression (data not shown). The equation for the fitted straight line (R2 >0.99) was used to determine the quantity of AMP present in later samples.

Analysis of the catalytic activity of Aprataxin was performed by incubation of recombinant Aprataxin with either AppppA or AMPNH2 in varying concentrations. All reactions were performed under non-substrate limiting conditions (less than 10% hydrolysis of the substrate) so that the measured reaction velocity is a good approximation for V0.

Binding of Aprataxin to the nucleotide derivative AMPNH2 has already been demonstrated by partial chymotryptic proteolysis (Figures 3.1 and 3.3). Reaction

116 velocities for a range of substrate concentrations are plotted in Figure 3.8a and were transformed to generate a double reciprocal plot (Figure 3.8b). Calculation of the linear line of best fit for the double reciprocal plot was performed using Microsoft Excel (equation shown on chart). Based on the linear nature of the hydrolysis data on the double reciprocal plot, it appears that the hydrolase activity of Aprataxin follows simple Michaelis-Menten kinetics (binds to a single molecule of substrate and releases two products, which would be typical for HIT proteins). Simple mathematical transformation of the equation describing the line of best fit of the double reciprocal plot can be used to determine the kinetic parameters for this reaction.

Any straight line can be described by the equation y= mx + c, where m is the gradient of the line and c is a constant. Substitution of the double reciprocal plot axis units and the gradient of the line (Km /Vmax) into this equation yields: (1/Velocity) = m(1/SubstrateConc) + c

It is apparent that as the concentration of substrate increases, the observed reaction velocity also increases due to increased active site occupancy on the enzyme. Theoretically, if an infinitely high substrate concentration could be achieved then the time between the release of products from an active site and the entrance of a new substrate would be zero. Therefore the limiting factor for the rate of this reaction would be the speed at which the enzyme can perform the reaction. This idea can be used in conjunction with the equation above to determine the theoretical maximum rate of an enzymatic reaction. SubstrateConc  ,

/1 SubstrateConc  0 Under these circumstances, substitution into the equation for this straight line yields: /1  cVelocity which can be rearranged to determine the velocity of reaction at infinite substrate concentration: cVelocity 1

Velocity  /1 c This is the maximum possible reaction speed for a particular enzyme, and is denoted Vmax. The equation for a double reciprocal plot can be similarly manipulated to

117 determine the affinity of an enzyme for its substrate, Km. The gradient of the line is described by:  VKmm max/ Now that Vmax is known and the gradient of the line of best fit has been determined by linear regression, this equation can easily be solved for Km:  VmKm max

All reaction parameters presented here were calculated by dividing the number of moles of AMP produced (determined by peak integration and substitution into the standard curve equation) by the number of moles of protein in the reaction (one active site/protein molecule) and the reaction period in seconds. This yields values in moles/mole/sec, which cancels out to /sec, the standard unit for expression of reaction velocities. Thus the kinetic parameters for the hydrolysis of AMPNH2 by Aprataxin are

Vmax= 0.069/sec and Km = 816 µM. Hint hydrolyses this substrate with Vmax=

0.20/sec and Km = 68 nM (0.068 µM) (21). One measure of the efficiency of an enzyme at metabolizing its substrate is the ratio of the Vmax and Km. The lower the ratio of Km

/Vmax the more efficient an enzyme is. Km /Vmax for Aprataxin hydrolyzing AMPNH2 is 11826 µM.sec, and the ratio of Hint for the same substrate is 0.34 µM.sec, indicating that Aprataxin is approximately 3,500 times less efficient at performing this reaction than Hint. Although at the time (mid 2006) this was the highest reported reaction velocity for Aprataxin on any substrate, comparison of Aprataxin’s kinetic parameters with those of Hint indicated that Aprataxin is not a Hint-type enzyme.

Based on the homology of Aprataxin to Fhit (an AppppA hydrolase) the activity of Aprataxin against AppppA was also determined. Hydrolysis of diadenosine tetraphosphate by Aprataxin resulted in formation of equimolar amounts of AMP and ATP, as analyzed by anion exchange HPLC. Quantification of AMP production over a range of substrate concentrations yielded the velocity versus concentration plot shown in Figure 3.9a. Inversion of these values generated the double reciprocal plot shown in Figure 3.9b. As before, linear regression of the double reciprocal plotted data allowed determination of the kinetic parameters of this reaction (Km = 3.87 µM and Vmax= 0.0194/sec). Similarly, hydrolysis of this substrate by Fhit results in the formation of equimolar amounts of ATP and AMP, with a similar Km but much higher velocity

118 (Vmax= 73/sec, Km = 1.9 µM) (22). The Km /Vmax ratios of these two enzymes indicate that Aprataxin is over 7,500 times less efficient at hydrolysis of AppppA than Fhit.

Based on the Km /Vmax ratio for Aprataxin on both substrates, AppppA and

AMPNH2, I determined that Aprataxin was more efficient at hydrolysis of the former. Due to the very low maximum reaction velocity for hydrolysis of this substrate, it seemed unlikely that AppppA could be Aprataxin’s physiological substrate, even though the observed Km indicated that binding may be possible at physiological concentrations (23).

119 A.

B.

Figure 3.8: Generation of AMP from hydrolysis of AMPNH2. Recombinant Aprataxin was incubated with up to 1500 µM AMPNH2 as described in methods. After the reaction period, samples were subjected to HPLC as described and quantity of AMP produced was determined by peak integration by ChemStation. A. The observed reaction velocities (picomoles of product/ picomoles of enzyme/ second) were plotted against substrate concentration. B. The observed reaction velocities and substrate concentrations were transformed to generate a double reciprocal plot. A linear line of best fit was generated by Microsoft Excel, with the equation and R2 shown.

120

A.

B.

Figure 3.9: Generation of AMP from hydrolysis of diadenosine tetraphosphate. Recombinant Aprataxin was incubated with up to 400 µM diadenosine tetraphosphate as described in methods. After the reaction period, samples were subjected to HPLC as described and quantity of AMP produced was determined by peak integration by ChemStation. A. The observed reaction velocities (picomoles of product/ picomoles of enzyme/ second) were plotted against substrate concentration. B. The observed reaction velocities and substrate concentrations were transformed to generate a double reciprocal plot. A linear line of best fit was generated by Microsoft Excel, with the equation and R2 shown.

121

3.3.3 Characterization of the interaction between DNA binding and nucleotide hydrolysis: As AppppA was determined to be the most efficient Aprataxin substrate identified at that time (section 3.3.2), this molecule was used to analyze the functional interaction between the HIT and zinc finger domains. Binding of Aprataxin to single and double stranded DNA had already been demonstrated (section 3.3.1), so I was interested to explore any interaction between the activities of the HIT domain, which is a nucleotide hydrolase, and the zinc finger, which binds polynucleotides. Initial experiments involved addition of increasing concentrations of double stranded DNA to a set of diadenosine tetraphosphate hydrolase reactions all containing the same substrate and enzyme concentration (conditions outlined in section 3.2.3). Reaction velocity was once again determined by quantification of AMP formation. A plot of the observed reaction velocities versus concentration of DNA is shown (Figure 3.10). I observed inhibition of the AppppA hydrolase activity of Aprataxin upon addition of increasing concentrations of DNA. More detailed kinetic experiments were necessary to define the nature of the relationship between DNA binding and inhibition of nucleotide hydrolase activity.

122

Figure 3.10. Inhibition of Aprataxin’s diadenosine tetraphosphate hydrolase activity by double stranded DNA. Recombinant Aprataxin was incubated with 400 µM diadenosine tetraphosphate in the presence of up to 10 nM double stranded DNA as described in Materials and Methods. Reaction constituents were resolved by HPLC as described and the amount of AMP produced analyzed by integration using ChemStation. Observed reaction velocities were plotted against concentration of DNA added. A linear trend towards lower reaction velocities at high DNA concentrations was observed.

Enzyme inhibition can be split into two basic categories: competitive and non- competitive. Competitive inhibitors bind to the active site of an enzyme and prevent binding of the substrate. Non-competitive inhibitors bind the enzyme distally from the active site but impair the ability of the enzyme to perform its reaction and release products. These two types of inhibition are shown diagrammatically in Figure 3.11.

123

Figure 3.11: Models of enzyme inhibition. In an uninhibited reaction, an enzyme binds to its substrate, forms an enzyme-substrate complex and then releases products. Binding of the enzyme to the substrate is impaired by the presence of a competitive inhibitor, preventing formation of the enzyme-substrate complex. Binding of a non-competitive inhibitor to an enzyme results in a reduced efficiency for conversion of the enzyme- substrate complex into enzyme and released products.

Under uninhibited conditions the enzyme binds its substrate to form an enzyme- substrate intermediate and then releases products. Under conditions of competitive inhibition the substrate and inhibitor bind the enzyme in a mutually exclusive manner and compete for access to the active site. It follows that at very high substrate concentrations, binding of the inhibitor will be prevented and the reaction can proceed as normal. This type of inhibition results in an increase in the apparent Km (indicating an apparent reduction in substrate affinity) and no change in the maximum reaction velocity. Under conditions of non-competitive inhibition substrate and inhibitor can bind to the enzyme at the same time, and elevation of the concentration of one molecule does not impair the enzyme binding of the other. This form of inhibition is due to the reduced (or ablated) ability of the enzyme-inhibitor complex to form reaction products. In this instance the affinity of the enzyme for the substrate is unchanged and the maximum velocity is reduced. Some inhibitors may cause both competitive and non- competitive inhibition of a reaction. This is referred to a mixed inhibition. The contribution of competitive and non-competitive inhibition to the overall reduction of

124 enzyme activity can be determined by comparing the change in apparent Km to the change in Vmax.

To determine the mode of inhibition of the hydrolase activity of Aprataxin by DNA, recombinant protein was incubated with variable concentrations of substrate as previously in the presence or absence of a low concentration of double stranded DNA (section 3.3.2). A substrate concentration versus reaction velocity plot is shown in Figure 3.12a. It is apparent from this plot that addition of DNA to these Aprataxin hydrolase reactions has both reduced the maximum velocity and increased the Km, indicating that DNA is a mixed inhibitor. Kinetic parameters for these reaction sets were once again determined from a double reciprocal plot (Figure 3.12b). Addition of 10 nM double stranded DNA resulted in a reduction of the maximum velocity to 0.0086/sec

(from 0.0194/sec) and an increase of the Km to 65.2 µM (from 3.87 µM). Inhibition constants for the competitive and non-competitive inhibition effects of DNA were determined based on this data using the standard methods (Kicomp=0.00259, Kinon-comp= 0.4273).

125

A.

B.

Figure 3.12. Impact of DNA on reaction kinetics of diadenosine tetraphosphate hydrolysis by Aprataxin. Recombinant Aprataxin was incubated with up to 1000 µM diadenosine tetraphosphate in the presence or absence of 10 nM double stranded DNA (indicated by inhibited or uninhibited) as described in methods. Reaction constituents were resolved by HPLC as described and the amount of AMP produced analyzed by integration using ChemStation. A. Observed reaction velocities were plotted against concentration of diadenosine tetraphosphate. B. These values were subsequently used to generate a double reciprocal plot. Lines of best fit for both reaction sets were generated by Microsoft Excel, with equations and R2 values shown.

126

This indicated that DNA was a mixed inhibitor of Aprataxin’s hydrolase activity. It was apparent that for DNA to be (in part) a competitive inhibitor of AppppA hydrolase activity, DNA probably binds to the active site of Aprataxin. Based on the probable binding of DNA to the HIT domain active site and the interaction of Aprataxin with DNA repair proteins, it seemed likely that Aprataxin may be a DNA modifying enzyme in vivo.

3.3.4 Inhibition of single strand break repair by 3’ terminal DNA damage: Further investigation into this possibility revealed that Aprataxin has hydrolase activity against 5’ adenylated DNA (18), an intermediate of DNA repair which can accumulates in some circumstances. 5’ adenylated DNA is generated as part of the ligation process (Equation 1.1, included here for clarity). Initially the DNA ligase forms a covalent bond with AMP (enzyme adenylation, step 1), which is then transferred to the 5’ terminus of a DNA break (DNA adenylation, step 2). After DNA adenylation, the phosphodiester bond is restored and AMP released (end-joining, step 3). A 3’hydroxyl group is absolutely required for end-joining but is not required for the enzyme and DNA adenylation phases of the reaction mechanism. If ligation is not completed this AMP molecule may remain attached to the 5’ DNA terminus, inhibiting further attempts at repair. 33

E + Appp E-AMP + PPi 1. enzyme adenylation E-AMP + 5’P-DNA E-AppDNA 2. DNA adenylation E-AppDNA + 3’OH DNA 3. end joining DNA DNA-p-DNA

Equation 1.1: Reaction mechanism of DNA ligation. Various chemical species are differentiated by colour throughout the reaction set.

Initially it was of interest to determine what circumstances can cause inhibition of ligation by cell extracts. A system to measure the impact of 3’ damage on ligation of a single strand break was devised (section 3.2.3.1 and Figure 3.13). This system involves incorporation of a variety of types of DNA damage at the 3’ terminus of a single strand break, with the 5’ terminus of the same oligonucleotide radiolabelled. Using this

127 system I show that when present at the 3’ terminus of a single strand break, three common types of base damage inhibit ligation by nuclear extracts (Figure 3.14). The presence of 8-oxo-dG, a product of oxidative DNA damage, on the 3’ terminus of a break completely inhibited single strand break ligation, as did the presence of a 3’ abasic site. Repair of a single strand break possessing a 3’ phosphate group was still possible, although only after dephosphorylation of the 3’ terminus (as seen by a half- nucleotide increase in apparent molecular weight).

Oligo A Oligo B

Figure 3.13: Single strand break repair schematic. A series of short nicked DNA duplexes were generated from a long (36 nucleotide) strand and two complimentary shorter strands. Oligo B, as indicated above, may possess either an unmodified (hydroxyl) or a modified (phosphate, 8-oxo-dG or abasic) 3’ terminus. This oligonucleotide is radiolabelled using γ P-32 ATP and T4 polynucleotide kinase as described prior to annealing with the other two oligonucleotides to generate radiolabelled duplex with a nick. Sequences are shown in Table 3.2.

128

Figure 3.14: Repair of 3’ damaged single strand breaks by cell extracts. Control (C3ABR) nuclear extracts were incubated with the unmodified or modified DNA nick structures depicted in Figure 3.13 for the indicated times, according to section 3.2.3.3. The different molecular electrophoretic mobilities of ‘reagents’ are anticipated and are due to modifications of the 3’ terminus. 3’8-oxo-dG modification (at any position in the chain) causes a marginal increase in apparent molecular weight compared to unmodified DNA. The presence of a phosphate (at either terminus) results in an aproximatley half- nucleotide decrease in apparent molecular weight, while an abasic site results in a single nucleotide decrease in apparent size. Reactions were terminated by addition of formamide-EDTA denaturing loading dye prior to resolution by denaturing gel electrophoresis (19% acrylamide 1% bis-acrylamide). The dried gel was used to expose a Molecular Dynamics phosphor screen.

The impact of modification of the 3’ terminus on DNA ligation was subsequently analyzed from the perspective of the 5’ strand. In a variation of the system used previously, the 5’ termini of the nicked duplexes (Oligo A) were radiolabelled to

129 measure ligase-dependant adenylation of the 5’ break terminus in response to 3’ terminal damage (method in section 3.2.6, schematic in Figure 3.15, result in Figure 3.16).

Oligo A Oligo B

Figure 3.15: Alternative single strand break repair schematic. The assay depicted in Figure 3.13 was modified such that the impact of ligation on the 5’ strand could be observed. The 5’ strand (instead of the 3’ strand) was radiolabelled with P-32 as described (section 3.2.6). This strand was annealed with the unlabeled ones to generate nicked duplex labeled in the indicated position. Note that repair of the 3’ strand cannot be assessed by this system.

Figure 3.16: Repair of 3’ damaged single strand breaks by T4 DNA ligase. Unmodified or modified DNA duplexes were incubated with recombinant T4 DNA ligase for the indicated times prior to reaction termination and denaturing gel electrophoresis as described (section 3.2.6). The identities of each of the observed bands are indicated. The far right hand lane is a no enzyme control used to identify non-adenylated (reagent) Oligo A.

130

We show that the presence of a phosphate group, 8-oxo-dG or an abasic site on the 3’ terminus of a nick inhibits ligation (relative to an unmodified control) and that a 3’ abasic site and 3’ 8-oxo-dG also cause adenylation of the 5’ strand (Figure 3.16). It seems likely that 3’ phosphate inhibits binding of DNA ligase to the break due to the high concentration of negative charge.

3.3.5 Binding of Aprataxin to 5’ adenylated DNA: Having shown that some types of complex DNA damage could result in the formation of adenylated DNA, the binding and hydrolase activity of Aprataxin against this structure was analyzed. To do this, adenylated DNA was generated by incubating radiolabelled nicked DNA possessing a 3’ dideoxy terminus with T4 DNA ligase in the presence of ATP (as described in section 3.2.6). The resulting adenylation of the 5’ strand can be observed as a reduction in the electrophoretic mobility of the DNA by approximately one nucleotide. This is readily resolved by denaturing gel electrophoresis and detected by subsequent autoradiography. A schematic of this DNA-based substrate and its proposed cleavage by Aprataxin is shown in Figure 3.17.

= P-32 phosphate group

5’18-mer 3’18-mer

Figure 3.17: Schematic of Aprataxin cleavage of 5’adenylated DNA. Adenylated DNA was generated by incubation of a nicked radiolabelled DNA duplex possessing a 3’ dideoxy break terminus (indicated by “dd”) with recombinant T4 DNA ligase in the presence of ATP.

The binding of Aprataxin to adenylated DNA was analyzed by EMSA as previously described (section 3.2.2). Binding was observed as a reduction in electrophoretic

131 mobility of the double stranded DNA (Figure 3.18). The specificity of this interaction was confirmed by antibody super-shift.

Figure 3.18: Aprataxin binding to 5’ adenylated DNA. Recombinant Aprataxin was incubated with affinity purified rabbit α-Aprataxin antibody or rabbit null serum as described in section 3.2.7. Duplex A was added to this and incubated on ice prior to native polyacrylamide gel electrophoresis. The antibody supershift demonstrates the specificity of the Aprataxin-Duplex A interaction.

3.3.6 Hydrolysis of 5’ adenylated DNA by Aprataxin: Having demonstrated that Aprataxin binds to adenylated DNA, I examined the whether Aprataxin hydrolyses this molecule. The hydrolysis of 5’ adenylated DNA by Aprataxin was examined by incubating varying amounts of recombinant Aprataxin with Duplex A as described in section 3.2.8 (Figure 3.19). Reaction specificity was confirmed by inhibition of hydrolysis by affinity purified rabbit α-Aprataxin. Note that some non- adenylated DNA is present in the substrate even without addition of Aprataxin. This is due to incomplete adenylation of the DNA by T4 DNA ligase, and this residual unmodified DNA was taken into account when measuring Aprataxin’s reaction kinetics on this substrate.

132 A.

B.

Figure 3.19: Hydrolysis of 5’ adenylated DNA by Aprataxin. A. The indicated quantities of recombinant Aprataxin were incubated with 2 picomoles of adenylated DNA for one minute prior to reaction termination. Adenylated and deadenylated species were resolved by denaturing gel electrophoresis as previously. Reaction specificity was by inhibition of the reaction by a rabbit α- Aprataxin purified antibody (+Ab lane) compared to an equivalent amount of null serum (+NS lane). B. The dried gel was used to expose a phosphor screen as described and the scanned image was analyzed in ImageQuant 5.1.

Based on quantitation of the autoradiograph shown in Figure 3.19a (Figure 3.19b), 25 femtomoles of recombinant Aprataxin can hydrolyse 0.7 picomoles of adenylated DNA in 60 seconds. This yields a reaction velocity of 0.47/sec, much faster than the maximum velocities I obtained for AppppA (0.0194/sec) and AMPNH2 (0.069/sec) (section 3.3.2). This, coupled with the strong binding of Aprataxin to adenylated DNA provided strong indication that an in vivo function of Aprataxin may be hydrolysis of 5’ adenylated DNA.

133 As Aprataxin lacking the FHA domain had been reported to have elevated levels of hydrolase activity on another substrate (15), the contribution of the FHA and zinc finger domains to the DNA-adenylate hydrolase activity of Aprataxin was measured. Equimolar amounts of recombinant Aprataxin Proteins A (Full Length), B (No FHA) and C (No ZnF) were incubated with adenylated DNA for the indicated times prior to reaction termination. Samples were resolved by denaturing PAGE and analyzed as described previously (Figure 3.20).

Figure 3.20: DNA-adenylate hydrolase activity of Aprataxin C and N-terminal truncation mutants. The role of the FHA and zinc fingers in the adenylate hydrolase activity of the HIT domain was explored using the truncation mutant proteins generated in Chapter 2. Equimolar amounts of Proteins A (full length) B (no FHA) and C (no ZnF) were incubated with adenylated DNA for the indicated times. Reactions were terminated, resolved and analyzed as previously.

As previously reported for an N-terminal truncation of Aprataxin, I found that deletion of the FHA domain (as in Protein C) results in an increase in activity of the HIT domain relative to the activity of full length Aprataxin (Protein A). This indicates that the FHA

134 domain may be a negative regulator of HIT domain activity. Deletion of the zinc finger (Protein B) resulted in complete loss of hydrolase activity. It is not known whether zinc finger deletion has an impact on hydrolysis of small nucleotides (for example AppppA), but it seems likely that the loss of activity observed here is due to an inability of the protein to bind this DNA-based substrate in a stable manner.

3.3.7 Examination of 3’ phosphatase activity of Aprataxin: Aprataxin has also been reported to remove 3’ phosphate from nicked DNA (24). This report suggests a role for Aprataxin as a general end-processing factor, but the reported kinetic data indicate that other enzymes (for example PNKP and APE1) are much more efficient 3’ phosphatases than Aprataxin (detailed section 1.2.2). It was therefore of interest to examine the proposed phosphatase activity of Aprataxin in more detail.

Duplex DNA containing a nick with both 5’ and 3’ phosphate termini was generated, with the 3’ strand radiolabelled (according to section 3.2.9). This duplex (1 pmol) was incubated with increasing concentrations of recombinant Aprataxin protein (between 0.1 and 4 pmol) for 75 minutes. Aprataxin incubated with 5’ adenylated DNA served as a positive control for Aprataxin hydrolase activity. Complete hydrolysis of 5’ adenylated DNA was observed even at the lowest concentration of Aprataxin. Conversely, phosphatase activity was not detected even in the presence of a molar excess of Aprataxin with a long incubation period (Figure 3.21). This indicates that the recombinant Aprataxin produced in section 2.2.6 does not have phosphatase activity. This finding is supported by others in the field (Prof Keith Caldecott, University of Sussex, Brighton, UK and Prof Steven West, Cancer Research UK, London Research Institute, UK, personal communications). Takahashi et al. additionally claimed that the 3’ phosphatase activity of Aprataxin is higher than that of the well characterised phosphatase PNKP, and that Aprataxin is the major contributor to 3’ phosphate repair in cells (24). In light of our failure to find phosphatase activity in recombinant Aprataxin, I examined the 3’ phosphatase activity of control and AOA1 cell extracts. If Aprataxin is a major contributor to the repair of 3’ phosphates in cells, AOA1 cells (which lack Aprataxin) should display a marked deficit in conversion of a 3’ phosphate to a hydroxyl. Control and AOA1 nuclear extracts display equal proficiency at hydrolysis of 3’ phosphate into a 3’ hydroxyl group (Figure 3.22). I therefore conclude that the phosphatase activity reported by Takahashi et al. is due to contamination of

135 their protein preparations and that Aprataxin does not play an evident role in the repair of 3’ phosphate modified termini {Takahashi, 2007 #231}.

Figure 3.21: Testing Aprataxin for phosphatase activity. Recombinant Aprataxin was incubated with one picomole of ‘5 adenylated nicked DNA or 3’ phosphorylated nicked DNA in the indicated quantities for 75 minutes. The phosphorylation status of the substrate duplex was confirmed by phosphorylated and non-phosphorylated molecular weight standard oligonucleotides. Hydrolysis of 5’ adenylated DNA was observed as a loss of the adenylated DNA band and an increase in intensity of the 5’ phosphate band. Protocol described in section 3.2.9.

Figure 3.22: 3’ Phosphatase activity of control and AOA1 cell extracts. Radiolabelled DNA duplex with a 3’ phosphate modified internal nick (1 pmol) was incubated with the indicated quantities of control (C2ABR and C3ABR) and AOA1 (L938 and L939) nuclear extract. Hydrolysis of the 3’ phosphate to a hydroxyl can be seen as an approximately half-nucleotide increase the in apparent molecular weight of the radiolabelled species. Protocol described in section 3.2.9.

This section has described the biochemical characterization of recombinant Aprataxin, from the initial stages of identification of interacting molecules through to enzymatic analysis of a range of substrates and examination of the contribution of each domain to hydrolase activity. These findings will now be discussed in relation to the literature.

136 3.4 DISCUSSION

This chapter has described a thorough biochemical characterization of the properties of recombinant Aprataxin, from initial binding experiments to enzymatic analysis on a variety of substrates.

At the commencement of this study, the pool of literature related to Aprataxin was limited and the biochemical studies which had been performed were not in agreement. Therefore it was initially necessary to establish the types of molecules Aprataxin can bind to. An interaction between Aprataxin and the potential substrate adenosine monophosphoramidate and product AMP was demonstrated by partial chymotryptic proteolysis. This provided evidence that Aprataxin is capable of binding these nucleotides in a stable manner. This method was not used for further characterization of Aprataxin’s binding properties due to the fine line between adequate and complete protease digestion. This interaction was then characterized in further detail by EMSA. This proved to be a far simpler and more reliable technique (although initial gels such as Figure 3.5 were a bit smeary, possibly due to inadequate cooling or pre-running).

It was important to establish the DNA structures Aprataxin binds to, so that I can begin to understand its role in normal cells and the molecular defect in AOA1. Data presented here shows that Aprataxin binds in a stable manner to double stranded DNA and transiently to single stranded DNA. Additional work by Dr Amanda Kijas has shown that the binding of Aprataxin to double stranded DNA is not sequence specific (17), which is consistent with a role for Aprataxin as a DNA repair enzyme.

An analysis of the enzymatic activity of Aprataxin against Fhit and Hint type substrates (adenosine monophosphoramidate and diadenosine tetraphosphate) revealed that Aprataxin is not efficient at hydrolysis of either of these molecules. Up until this time most studies attempted to categorize Aprataxin as either an Fhit or Hint-type enzyme. Demonstration that Aprataxin is incapable of efficient hydrolysis of these molecules (compared to the kinetic parameters of Fhit and Hint) allowed this theory to be discarded (17).

137 Even though Aprataxin displayed very low activity against diadenosine tetraphosphate and adenosine monophosphoramidate, establishment of this simple HPLC-based assay allowed us to study the interplay between nucleotide hydrolysis and DNA binding. I was subsequently able to demonstrate that DNA is a competitive inhibitor of small nucleotide hydrolysis by Aprataxin. This indicated that DNA could bind to the HIT domain active site. Combined with the sensitivity of AOA1 cells to DNA damaging agents, the interaction between the HIT domain of Aprataxin and DNA indicated that Aprataxin’s in vivo substrate could be a DNA adduct.

Following the publication of the inhibition of Aprataxin hydrolase activity by DNA (17), a novel DNA-based substrate for Aprataxin was described (18). This molecule, 5’ adenylated DNA, is formed as an intermediate during the DNA ligation process. I show that oxidative damage to the 3’ terminus of a DNA break can result in inhibition of the end joining phase of the DNA ligation reaction and generation of adenylated DNA. Using native and denaturing gel electrophoresis techniques, I confirmed that Aprataxin both binds to and hydrolyses this substrate. Even though the zinc finger is unlikely to participate directly in hydrolysis of adenylated DNA, deletion of the zinc finger ablated hydrolase activity. The most likely explanation for this is that the zinc finger is involved in orientation of the substrate or stabilization of the substrate-enzyme complex.

Additionally, I have shown that the DNA adenylate hydrolase activity of Aprataxin is enhanced by deletion of the FHA domain. This provides evidence that the hydrolase activity of Aprataxin is regulated via the FHA domain, and indicates a possible control mechanism for activation or suppression of activity. The FHA domain of Aprataxin interacts with a number of proteins including the DNA repair scaffold proteins XRCC1 and XRCC4 (outlined in section 1.2.1). Given that deletion of this region stimulates the activity of the HIT domain, in vivo the activity of Aprataxin could be modulated by interacting proteins.

It has been reported that Aprataxin possesses 3’ phosphatase activity in addition to DNA adenylate hydrolase activity (24). I was unable to replicate this finding, and note that this reported activity (Vmax= 0.00027/sec and Km = 129 µM) was very low compared to Aprataxin’s hydrolase activity on adenylated DNA (at least

138 0.47/sec). I propose that this phosphatase activity could be due to contamination of the authors protein preparation.

The biochemical properties of recombinant Aprataxin described here are consistent with a role for Aprataxin as a general DNA repair/proof-reading enzyme, with a primary function of removing the ligation-blocking modification 5’ DNA adenylate. DNA repair is a highly regulated process in which many proteins have redundant functions. Therefore to understand the role of Aprataxin within a network of DNA repair proteins, an analysis of the DNA repair capacity of Aprataxin deficient cells is necessary. This will be described in the next chapter.

139 3.5 REFERENCES

1. Date, H., Onodera, O., Tanaka, H., Iwabuchi, K., Uekawa, K., Igarashi, S., Koike, R., Hiroi, T., Yuasa, T., Awaya, Y. et al. (2001) Early-onset ataxia with ocular motor apraxia and hypoalbuminemia is caused by mutations in a new HIT superfamily gene. Nat Genet, 29, 184-188. 2. Moreira, M.C., Barbot, C., Tachi, N., Kozuka, N., Uchida, E., Gibson, T., Mendonca, P., Costa, M., Barros, J., Yanagisawa, T. et al. (2001) The gene mutated in ataxia-ocular apraxia 1 encodes the new HIT/Zn-finger protein aprataxin. Nat Genet, 29, 189-193. 3. Clements, P.M., Breslin, C., Deeks, E.D., Byrd, P.J., Ju, L., Bieganowski, P., Brenner, C., Moreira, M.C., Taylor, A.M. and Caldecott, K.W. (2004) The ataxia-oculomotor apraxia 1 gene product has a role distinct from ATM and interacts with the DNA strand break repair proteins XRCC1 and XRCC4. DNA Repair (Amst), 3, 1493-1502. 4. Gueven, N., Becherel, O.J., Kijas, A.W., Chen, P., Howe, O., Rudolph, J.H., Gatti, R., Date, H., Onodera, O., Taucher-Scholz, G. et al. (2004) Aprataxin, a novel protein that protects against genotoxic stress. Hum Mol Genet, 13, 1081- 1093. 5. Luo, H., Chan, D.W., Yang, T., Rodriguez, M., Chen, B.P., Leng, M., Mu, J.J., Chen, D., Songyang, Z., Wang, Y. et al. (2004) A new XRCC1-containing complex and its role in cellular survival of methyl methanesulfonate treatment. Mol Cell Biol, 24, 8356-8365. 6. Durocher, D. and Jackson, S.P. (2002) The FHA domain. FEBS letters, 513, 58- 66. 7. Cahill, D., Connor, B. and Carney, J.P. (2006) Mechanisms of eukaryotic DNA double strand break repair. Front Biosci, 11, 1958-1976. 8. Caldecott, K.W. (2003) XRCC1 and DNA strand break repair. DNA Repair (Amst), 2, 955-969. 9. Vrana, K.E., Churchill, M.E., Tullius, T.D. and Brown, D.D. (1988) Mapping functional regions of transcription factor TFIIIA. Mol Cell Biol, 8, 1684-1696. 10. Brown, R.S. (2005) Zinc finger proteins: getting a grip on RNA. Curr Opin Struct Biol, 15, 94-98.

140 11. Friesen, W.J. and Darby, M.K. (2001) Specific RNA binding by a single C2H2 zinc finger. J Biol Chem, 276, 1968-1973. 12. Kennedy, D., Ramsdale, T., Mattick, J. and Little, M. (1996) An RNA recognition motif in Wilms' tumour protein (WT1) revealed by structural modelling. Nat Genet, 12, 329-331. 13. McCarty, A.S., Kleiger, G., Eisenberg, D. and Smale, S.T. (2003) Selective dimerization of a C2H2 zinc finger subfamily. Mol Cell, 11, 459-470. 14. Brenner, C., Bieganowski, P., Pace, H.C. and Huebner, K. (1999) The histidine triad superfamily of nucleotide-binding proteins. J Cell Physiol, 181, 179-187. 15. Hirano, M., Furiya, Y., Kariya, S., Nishiwaki, T. and Ueno, S. (2004) Loss of function mechanism in aprataxin-related early-onset ataxia. Biochem Biophys Res Commun, 322, 380-386. 16. Seidle, H.F., Bieganowski, P. and Brenner, C. (2005) Disease-associated mutations inactivate AMP-lysine hydrolase activity of aprataxin. J Biol Chem. 280, 20927-20931. 17. Kijas, A.W., Harris, J.L., Harris, J.M. and Lavin, M.F. (2006) Aprataxin forms a discrete branch in the HIT (histidine triad) superfamily of proteins with both DNA/RNA binding and nucleotide hydrolase activities. J Biol Chem, 281, 13939-13948. 18. Ahel, I., Rass, U., El-Khamisy, S.F., Katyal, S., Clements, P.M., McKinnon, P.J., Caldecott, K.W. and West, S.C. (2006) The neurodegenerative disease protein aprataxin resolves abortive DNA ligation intermediates. Nature. 443, 713-716. 19. Garner, M.M. and Revzin, A. (1981) A gel electrophoresis method for quantifying the binding of proteins to specific DNA regions: application to components of the Escherichia coli lactose operon regulatory system. Nucleic Acids Res, 9, 3047-3060. 20. Audebert, S., Calsou. (2004) Involvement of Poly(ADP-ribose) Polymerase-1 and XRCC1/DNA Ligase III in an Alternative Route for DNA Double-strand Breaks Rejoining. J Biol Chem, 279, 55117-55126. 21. Bieganowski, P., Garrison, P.N., Hodawadekar, S.C., Faye, G., Barnes, L.D. and Brenner, C. (2002) Adenosine monophosphoramidase activity of Hint and Hnt1 supports function of Kin28, Ccl1, and Tfb3. J Biol Chem, 277, 10852- 10860.

141 22. Barnes, L.D., Garrison, P.N., Siprashvili, Z., Guranowski, A., Robinson, A.K., Ingram, S.W., Croce, C.M., Ohta, M. and Huebner, K. (1996) Fhit, a putative tumor suppressor in humans, is a dinucleoside 5',5"'-P1,P3-triphosphate hydrolase. Biochemistry, 35, 11529-11535. 23. Rapaport, E. and Zamecnik, P.C. (1976) Presence of diadenosine 5',5''' -P1, P4- tetraphosphate (Ap4A) in mamalian cells in levels varying widely with proliferative activity of the tissue: a possible positive "pleiotypic activator". Proc Natl Acad Sci U S A, 73, 3984-3988. 24. Takahashi, T., Tada, M., Igarashi, S., Koyama, A., Date, H., Yokoseki, A., Shiga, A., Yoshida, Y., Tsuji, S., Nishizawa, M. et al. (2007) Aprataxin, causative gene product for EAOH/AOA1, repairs DNA single-strand breaks with damaged 3'-phosphate and 3'-phosphoglycolate ends. Nucleic Acids Res, 35, 3797-3809.

142

CHAPTER 4 Characterization of multiple DNA repair defects in AOA1 cells

143 144 4.1 INTRODUCTION

DNA single strand break repair (SSBR) is a complex cellular process, involving many sequential steps and is highly regulated. Thousands of single strand breaks occur in each cell every day, due to both free radicals produced by mitochondrial respiration and exogenous sources of stress. These breaks are repaired by highly efficient mechanisms, however unrepaired DNA single strand breaks have the potential to be converted into highly toxic double strand breaks.

Regardless of the nature of the lesion or the exact pathway responsible for repairing it, SSBR is a sequential process (detailed in section 1.3.2). Initially the lesion is detected, either by specific proteins which scan the genome, or by proteins which detect distortions of the normal DNA conformation. The lesion, or a section of the strand containing it, is then enzymatically excised to generate a single strand break with one or more nucleotides missing. Processing is often required to restore the 5’ phosphate and 3’ hydroxyl termini prior to gap filling by a DNA polymerase and subsequent ligation.

4.1.1 Assembly of a ‘repairosome’: These sequential processes are facilitated by both constitutive and induced protein- protein interactions which assemble the repair machinery at the lesion. The presence of pre-formed complexes ensures maximum repair efficacy by ensuring that all the required proteins are present to facilitate the ordered progression of reactions. As discussed in detail in section 1.2.1, Aprataxin interacts with the core XRCC1/lig3α/ polβ single strand break repair complex in a constitutive manner (1,2). A second XRCC1-containing complex exists which contains the end-processing factor PNKP but not Aprataxin (2), indicating that Aprataxin and PNKP may not be needed for repair of the same type of breaks. This strategy ensures that direct single strand breaks can be detected, processed, and the gap filled and ligated using the same protein complex. This is more efficient recruitment of individual proteins to the break and thus reduces the time required to repair each break. Protein deficiency in such a complex could have dramatic effects. Recruitment of the complex to sites of DNA damage may be impaired. Indeed, XRCC1 is not recruited to sites of damage in the absence of PARP-1 (3). Secondly, the repair reaction sequence may be interrupted if

145 no other enzyme or pathway is able to compensate for the deficiency. This is exemplified by the elevated levels of single strand breaks in Tdp1 deficient cells (4). Finally, deficiency of one protein can cause destabilization of proteins it interacts with. One example of this is the destabilization of the whole MRN complex in Mre11 or Rad50 mutated cells (5).

Additionally, both protein-protein interactions and post-translational modifications can affect cellular processes by modulating protein localization and enzymatic activities (6,7).

4.1.2 Aprataxin localization and post-translational modifications: Previous reports have shown that Aprataxin is a nuclear protein with nucleoplasmic and nucleolar distribution (8,9), compatible with a role for Aprataxin in DNA repair. However, compartmentalized proteins may have different enzymatic properties in different sub-cellular compartments (10). Furthermore the activity of enzymes is also frequently modified by post-translational modification (for examples see 11,12). Such modifications include phosphorylation, acetylation, glycosylation and adenylation as well as attachment of protein modifiers like ubiquitin and SUMO. Although Aprataxin contains putative phosphorylation sites (including target sites for ATM and ATR, Protein Kinases A, C and G, Casein Kinases I and II, and INSR) the phosphorylation status of Aprataxin is yet to be characterised.

4.1.3 A role for Aprataxin in multiple DNA repair pathways: As introduced in section 1.3.2.2, single strand break repair is a complex process which relies on several lesion-dependant pathways. The hypersensitivity of AOA1 cells to MMS and hydrogen peroxide (which generate single strand breaks) indicate that they have a defect in their cellular response to single strand breaks (1,2,9). Aprataxin’s interaction with the direct single strand break repair and BER proteins XRCC1 and PARP-1 indicated that it may have a direct role in repair (1,2,9,13), and this was strengthened by the report of Rass et al. who identified 5’ adenylated DNA as an efficient Aprataxin substrate (14).

In the previous chapter (section 3.3.4) it was demonstrated that that adenylated DNA is formed as a by-product of abortive ligations at complex single strand breaks. Each

146 of the DNA repair pathways outlined in section 1.3 has the potential to generate an adenylated 5’ terminus if ligation is erroneously attempted before 3’ processing has occurred. If 3’ and 5’ processing is performed successfully, the subsequent ligation reaction will be successful and Aprataxin may not be required. It is proposed that Aprataxin is required at a small subset of breaks where 3’ termini have not been processed prior to the first ligation attempt (14). Such breaks could be generated directly or as intermediates of indirect single strand break repair mechanisms and thus Aprataxin’s activity may be required in multiple repair pathways.

In this chapter our analysis of effect of sub-cellular localization and protein-protein interactions on Aprataxin activity is described. The post-translational modification status of Aprataxin, in particular in response to DNA damage, was also examined. Given that Aprataxin interacts with proteins involved in both direct single strand break repair and BER, I investigated the role of Aprataxin in these processes. The role of Aprataxin in the repair of complex single strand breaks possessing an oxidized 3’ terminus was examined. Finally the relationship between Aprataxin, XRCC1 and PARP-1 in the repair of single strand breaks was determined, focusing on characterisation of the molecular defects in AOA1.

147 4.2 MATERIALS AND METHODS

4.2.1 2D gel electrophoresis: 2D electrophoresis separates protein species in two stages. Initially proteins from a complex mixture are absorbed into an acrylamide strip with an immobilized pH gradient. Proteins migrate to their isoelectric point in the immobilized gradient when a current is applied, a process called isoelectric focusing (IEF). After focusing, the strip is embedded into an SDS-polyacrylamide gel for separation of proteins based on their mass/charge ratio. After resolution of the second dimension normal protein detection methods can be used (for example total protein or modification-specific stains and Western blotting).

Nuclear extracts were generated from C3ABR and L938 as described in the previous chapter (section 3.2.3.2). Proteins from these extracts were precipitated by addition of 10 volumes of 8:1 acetone: chloroform and incubation at -20°C for 4 hours. Precipitated protein was recovered by centrifugation and resuspended in 200 µL of 7 M urea, 2 M thiourea, 4% CHAPS, 50 mM DTT, 1% pH 7-10 ampholytes (Biorad) with a trace of bromphenol blue. Proteins were passively absorbed into a pH 7-10 isoelectric focusing strip (Biorad) according to the manufacturers instructions. After absorption proteins were focused at 8000 V (maximum 50 mA per strip) for 30,000 Vhrs. Strips were then equilibrated according the manufacturers instructions and embedded in the upper portion of a 5% stacking, 12% resolving denaturing acrylamide gel for second dimension separation. Proteins were transferred onto nitrocellulose for Western blotting with rabbit-α-Aprataxin serum (affinity purified rabbit α-Aprataxin was not available at this time).

4.2.2 In vitro kinase assay: This technique relies on immunoprecipitation to enrich the kinase of interest (or the use of purified recombinant kinase) and subsequent incubation of the enriched kinase with a recombinant substrate (the potential target for kinase activity) in the presence of γ-P32 ATP. Phosphorylation of the substrate is detectable following SDS-PAGE separation and autoradiography. A schematic of the in vitro kinase assay is shown in Figure 4.1.

148

Figure 4.1: Schematic of the an vitro kinase assay. The kinase of interest is precipitated from cell extracts (after appropriate treatment) using a specific antibody and proteinA/G beads. It is subsequently incubated with the desired substrate in the presence of γ-P32 ATP. Kinase activity results in the transfer of the radiolabelled γ phosphate from ATP onto the substrate, thus labelling the substrate.

4.2.2.1 ATM kinase assay: Endogenous ATM was immunoprecipitated from irradiated (10 Gy) or non-irradiated control (C3ABR) and ATM kinase-dead (AT3ABR) cell lines as described by Lavin et al. (15). Immunoprecipitation from AT3ABR provides a control for ATM antibody specificity. Immunoprecipitates were incubated with recombinant Aprataxin-GST (generated by Dr Amanda Kijas, Queensland Institute for Medical Research, Brisbane, Australia) or p53-GST (generated by Dr Sergei Kozlov Queensland Institute for Medical Research, Brisbane, Australia, reference 15) fusion proteins in the presence of γ-P-32 ATP according to the method of Lavin et al. (15). Reactions were analysed by protein electrophoresis.

4.2.2.2 ATR kinase assay: Control (C3ABR) cells were treated with 2 µM hydroxyurea (HU) or mock treated for 2 hours. Lysates were subsequently prepared and ATR was subsequently immunoprecipitated according to the method described in Lavin et al. (15) (antibody generated by Dr Rick Woods, Queensland Institute for Medical Research, Brisbane, Australia). Immunoprecipitates were incubated with Aprataxin-GST substrates and phosphorylation analysed as for ATM. Even loading is shown using Coomassie blue staining.

149 4.2.3 DNA binding of endogenous Aprataxin: A biotinylated duplex (5’ (biotin)- TGTAGTCACTATCGGAATGAGGGCGACACGGATATG, annealed to the unmodified complementary strand) was attached to streptavidin magnetic beads (Dynal) as per manufacturers instructions. Beads were mock treated in parallel. DNA coated or uncoated beads (10 µg) were incubated with 30 µg of nuclear extract from the indicated cell lines in Ligase Buffer (total volume 50 µL) for 10 minutes on ice. Reactions were gently mixed every several minutes. DNA-binding proteins were enriched by precipitation of the beads in a magnetic rack (Invitrogen) and washed in Ligase Buffer. DNA binding proteins were eluted in 5 x SDS-PAGE loading buffer and subjected to protein electrophoresis and Western blotting.

4.2.4 DNA-adenylate hydrolase activity of nuclear extracts: Duplex A and nuclear extracts were generated as described in the previous chapter (section 3.2.6). Unless stated otherwise, 10 µL reactions were performed in Ligase Buffer at room temperature. Concentrations of extract in each reaction and reaction times are indicated in the legend for each experiment. Reactions were terminated and resolved as described previously. Reaction kinetics were analysed by quantification of a phosphor image in ImageQuant 5.1. Specifically, the intensity of both reagent and product bands was quantified by integration. ‘Percent de-adenylation’ is calculated by determining the relative amount of radiation which corresponds to product ([intensity of product band/total intensity]x 100). ‘Picomoles of substrate converted’ is obtained by multiplication of this value by the number of picomoles of Duplex A in the reaction to begin with. Logarithmic curves of best fit were generated in Microsoft Excel.

Inhibition of the hydrolase activity of cell extract was achieved by pre-incubation of extracts with the affinity purified rabbit α-Aprataxin antibody. 5 µg of nuclear extract was incubated with 200 µg of purified antibody or null serum at 4°C for 20 minutes in a total volume of 8 µL in Ligase Buffer. After antibody binding reactions were initiated by addition of 2 pmol of Duplex A (to yield a final volume of 10 µL). Reactions were terminated as previously described after the indicated times. Reactions were analysed by denaturing electrophoresis as described.

150 4.2.5 Effect of 3’ oxidation on 5’ adenylate hydrolysis by extracts: 4.2.5.1 Substrate generation: The effect of oxidation of the adjacent terminus on DNA-adenylate hydrolase activity was examined by generating DNA structures with singly (5’ adenylate) and doubly modified (5’ adenylate, 3’ 8-oxo-dG) breaks. These structures were generated using the oligonucleotides in Table 4.1. Oligo A was radiolabelled as described previously and was subsequently adenylated. This was achieved by annealing 5’ radiolabelled Oligo A to Ligase Bottom 36-mer in Ligase Buffer and incubating this structure for 24 hours at room temperature with T4 DNA ligase (50 units to 200 pmol of DNA in a 50 µL volume). The ligation reaction was terminated by heat denaturation as described previously and Oligo B subsequently annealed to generate a 5’ adenylated nick with either a 3’ hydroxyl or 3’ 8-oxo-dG terminus.

Ligase Bottom 36mer 5' TGTAGTCACTATCGGAATGAGGGCGACACGGATATG Oligo A 5' CCCTCATTCCGATAGTGACTACA Oligo B (unmodified) 5' CATATCCGTGTCG Oligo B (8-oxo-dG) 5' CATATCCGTGTC(8-oxo-dG) Table 4.1: Oligonucleotides used to generate doubly modified nick structures.

4.2.5.2 Experimental conditions: 10 µg of control nuclear extracts were incubated with 2 pmol of singly or doubly modified nicked duplex in Ligase Buffer for the indicated times. Reactions were terminated and resolved as described previously.

4.2.6 In vitro single strand break repair by cell extracts: 4.2.6.1 Substrate generation:

Duplexes containing a nick with a modified or unmodified 3’ terminus were generated by 5’ radiolabelling Oligo B as described previously and annealing it to Ligase Bottom 36-mer and the 5’ strand (Oligo A). In this instance the radiolabel and the modified terminus are on opposite ends of the same oligonucleotide (in section 4.2.5 they were on different strands). Sequences are indicated in Table 4.2. These substrates are identical to the ones generated in section 3.2.3.

151 Ligase Bottom 36mer 5' TGTAGTCACTATCGGAATGAGGGCGACACGGATATG Oligo A 5' CCCTCATTCCGATAGTGACTACA Oligo B (unmodified) 5' CATATCCGTGTCG Oligo B (8-oxo-dG) 5' CATATCCGTGTC(8-oxo-dG)

Table 4.2: Oligonucleotides used to construct SSBR duplexes.

4.2.6.2 Experimental conditions: 10 µg of control or AOA1 nuclear extract was incubated with 2 pmol of either substrate (3’ hydroxyl or 3’ 8-oxo-dG) in Ligase Buffer (total volume 10 µL) at room temperature for the indicated times. Reactions were terminated and resolved as described.

4.2.7 Effect of Aprataxin on protein stability: Corrected (FD105 M21) and uncorrected (vector only, FD105 M20) transformed AOA1 fibroblasts were kindly provided by Prof Keith Caldecott (University of Sussex, Brighton, UK). This provided us with AOA1 fibroblasts capable of indefinite expansion and enabled us to study the role of Aprataxin in isogenic cell lines. These cells were maintained in DMEM supplemented with Fungizone, penicillin, streptomycin, and 12% FCS. FD105 M20 and M21 nuclear lysates were prepared by scraping cells in Lysis Buffer B (50 mM Tris pH 7.5, 150 mM NaCl, 2 mM EGTA, 2 mM EDTA, 25 mM NaF, 25 mM β-glycerophosphate, 0.2% Triton X-100, 0.3% NP- 40, 1 x complete protease inhibitor). Equal amounts protein were subjected to electrophoresis and Western blotting. The relative abundance of proteins was determined by quantification of the intensity of the band of interest on a non-saturated exposure. This value was standardized between the two cell lines using the intensity of the loading control band. Abundance in FD105 M20 was expressed as a fraction of the abundance in FD105 M21.

152 4.2.8 8-oxo-dG immunostaining: For 8-oxo-dG detection, Normal (NFF), AOA1 (FD105) fibroblasts were grown on coverslips for 48h and processed for immunofluorescence as described in (16). Briefly, cells on coverslips were fixed with 100% pre-chilled methanol for 5 min and immersed in 100% pre-chilled acetone for 5 min. Coverslips were subsequently air- dried, treated with 0.05 N HCl for 5 min on ice and washed three times with PBS. RNA was digested by incubating the coverslips in 100 µg/ml RNase in 150 mM NaCl with 15 mM sodium citrate for 1h at 37C. After RNA digestion, coverslips were sequentially washed in PBS, 35% ethanol, 50% ethanol and 75% ethanol for 2 min each. DNA was denatured by incubating the coverslips with 0.15 N NaOH in 70% ethanol for 4 min. A series of washes were performed starting with 70% ethanol containing 4% v/v formaldehyde, then 50% ethanol, 35% ethanol and finally PBS for 2 min each. Proteins were digested with 5 µg/ml proteinase K in TE pH 7.5 for 10 min at 37C. After several PBS washes, coverslips were incubated with anti-8-oxo-dG antibody in PBT20 (1X PBS/ 1% BSA/ 0.1% Tween 20) for 1h at room temperature. Following several washes with 0.1X PBS, 8-oxoG was detected using an AlexaFluor488 secondary antibody (Invitrogen, 1/500 in PBT20). Nuclei were counterstained with DAPI and slides were mounted for immunofluorescence. Images were captured using a digital camera (Carl Zeiss, Axiocam MRm) attached to a fluorescent microscope Axioskop2 mot plus (Carl Zeiss) using Plan Apochromat 1.4 oil DIC (63x magnification). The AlexaFluor 488 excitation wavelength was 488 nm, emission wavelength is 510 nm. The DAPI excitation wavelength was 345 nm, emission wavelength of 455 nm. Zeiss software (Axiovision LE 4.3) was used to capture the individual images which were assembled using Adobe Photoshop 7.0. Fluorescence intensity was quantitated on the RAW images using the public domain software Image J version 1.34s (NIH, USA) prior their assembly in Adobe Phototoshop 7.0.

4.2.9 Nitrotyrosine immunostaining: FD105 M20 and M21 fibroblasts were trypsinized and allowed to attach to sterile glass coverslips for at least 48 hours. Sub-confluent cells were fixed in PBS containing 10% formalin and blocked in PBS containing 5% NBS and 0.05% Triton X-100 and stained with rabbit α-Nitrotyrosine (1/500 in PBS with 5% NBS) for 2

153 hours at 37ºC. Coverslips were washed in PBS before detection using an α-rabbit AlexaFluor 594 conjugated antibody (1/1000 in PBS with 5% NBS) for 2 hours at 37ºC. Cells were counterstained with DAPI and washed in PBS before mounting in Moviol. The excitation wavelength of AlexaFluor 594 was 594 nm and the emission wavelength was 615 nm. Images were captured using a digital camera (Carl Zeiss, Axiocam MRm) attached to a fluorescent microscope Axioskop2 mot plus (Carl Zeiss) using Plan Apochromat 1.4 oil DIC (63x magnification). Zeiss software (Axiovision LE 4.3) was used to capture the individual images which were assembled using Adobe Photoshop 7.0.

4.2.10 Subcellular distribution of Aprataxin: 4.2.10.1 Fluorescent microscopy: Confluent HeLa cells were transfected with the Aprataxin-GFP construct described in Gueven et al. (9) using Lipofectamine according to the manufacturers protocol. Cells were fixed in 4% paraformaldehyde (in PBS) 48 hours after transfection, blocked in PBS containing 5% NBS and 0.05% Tween-20 and stained for nucleolin (1/500 dilution overnight 4ºC) in the same buffer. Nucleolin staining was detected using an α-mouse AlexaFluor 594 conjugated IgG. Cells were analysed by microscopy as previously (section 4.2.8).

4.2.10.2 Subcellular fractionation: Nucleolar and nucleoplasmic extracts were prepared as previously described(17). Briefly, 3 x 108 cells were washed twice in PBS. Pelleted cells were resuspended in 3 mL of Buffer A (10 mM Tris pH 7.5, 10 mM NaCl and 1 mM MgCl2) and incubated for 30 min on ice. Cell membranes were broken by dounce homogenization (20 strokes) and nuclei pelleted by centrifugation (1,200 x g). The supernatant (cytoplasmic fraction) was retained. The crude nuclear pellet was resuspended in 800

µL of Buffer B (0.25 M sucrose 10 mM MgCl2), layered over 1 mL of Buffer C (0.88

M sucrose 10 mM MgCl2) and centrifuged at 1,200 x g for 10 minutes. The purified nuclear pellet was resuspended in 800 µL of Buffer D (0.34 M sucrose 10 mM

MgCl2) and lysed by two sonication bursts of 20 seconds each with a 1 minute incubation on ice between (maximum intensity on a Branson Sonifier model S250A). The homogenate was centrifuged as previously through 1 mL of Buffer C. The supernatant (nucleoplasmic fraction) was stored on ice while the nucleolar pellet was

154 further processed. The nucleolar pellet was resuspended in 100 µL of Dialysis Buffer, incubated on a rotating wheel for 30 minutes and cleared by centrifugation at 16,100 x g for 10 minutes. The supernatant (nucleolar extract) was recovered and processed in parallel with the nucleoplasmic fraction. Cytoplasmic, nucleoplasmic and nucleolar fractions were dialysed against Dialysis Buffer prior to storage in aliquots at -80ºC.

4.2.10.3 Immunoprecipitation from fractions: Aprataxin immunoprecipitations were performed from cytoplasmic, nucleoplasmic and nucleolar protein fractions. 10 μg of sheep α-FHA affinity purified antibody was incubated with 500 μg of protein G cleared extract (generated as described in section 4.2.10.2) in a 500 μL volume in Lysis Buffer B overnight at 4ºC, on a rotating wheel. Proteins were precipitated with protein G agarose for 2 hours at 4ºC on a rotating wheel. Immunoprecipitates were washed in Lysis Buffer B, eluted in 5x PAGE buffer and subjected to protein electrophoresis and Western blotting.

4.2.11 Subcellular distribution of Aprataxin activity: Nucleoplasmic and nucleolar extracts from C2ABR and C3ABR (5μg each, generated as described in section 4.2.10.2) were incubated with 2 pmol of Duplex A in a 10 μL reaction in Ligase Buffer. Reactions were terminated, resolved and analysed as described previously. Percent repair is the proportion of substrate which has been converted to product. Lines are the logarithmic curve of best fit of the mean of three independent experiments using independently generated fractions of the two cell lines.

4.2.12 DNA-adenylate hydrolase activity of PARP-1 and XRCC1 defective cell lines: 4.2.12.1 Cell lines: The dependence of Aprataxin activity on PARP-1 and XRCC1 was examined using defective cell lines. EM9 is a Chinese hamster ovary (CHO) cell line derived by random mutagenesis followed by cellular sensitivity screening{Thompson, 1982 #533}. It has no full length XRCC1 protein. AA8 is the parental cell line. PARP-1 KO is a mouse embryonic fibroblast cell line derived from a mouse with the PARP-1 gene interrupted by a transposon. This cell line has no PARP-1 protein. The cell line PARP-1 WT is derived from a wild-type littermate. PARP-1 deficiency and XRCC1

155 deficiency respectively are confirmed in these cell lines by immunoblotting. All cell lines were obtained from the American Type Culture Collection.

4.2.12.2 Experimental conditions: Nuclear extracts were prepared from 90% confluent cultures in the same manner as previous experiments examining DNA repair activity (section 3.2.3.2). The indicated amounts of each extract were incubated in Ligase Buffer with Duplex A (2 pmol) in a 10 μL volume for the indicated times. Reactions were terminated and resolved as previously.

4.2.13 PARP-1 knockdown: To confirm the requirement of PARP-1 for stabilization of Aprataxin, PARP-1 was transiently depleted using siRNA technology. siRNAs directed against PARP-1 were purchased from Invitrogen and sequences are shown in Table 4.3. HeLa cells were grown to confluence in 6-well plates. Transfection was performed according to Invitrogens recommendations. Briefly, the transfection mixture was prepared by separate incubation of 250 μL of OptiMEM with 10 μL of Lipofectamine 2000 (per reaction) and 250 μL of OptiMEM with 200 μmol of each siRNA duplex at room temperature for 5 minutes. These were then mixed and incubated for a further 20 minutes at room temperature to allow formation of transfection reagent-siRNA complexes. In the mean time, cells were washed 3 x in OptiMEM. After the 20 minute incubation, the OptiMEM was removed from the cells and the transfection mixture was added. The plate was returned to the incubator and cells were incubated in the transfection mixture for 8 hours. After this period DMEM with 12% FCS was added directly to each well. The next day this process was repeated (double transfection). 48 hours after the end of the first transfection, cells were scraped in Lysis Buffer B. Cleared lysates were subjected to protein electrophoresis and Western blotting. PARP-1 knockdown and Aprataxin abundance were quantified in the same manner as protein abundance was deter mined in the FD105 cell lines previously (section 4.2.7).

156

PARP siRNA 1 5' ACUCCUACUACAAGCUGCAGCUUCU PARP siRNA 2 5' UAUCGAUGACUUCCAAGUCAUACGC PARP siRNA 3 5' GGAGACCCAAUAGGCUUAAUCCUGU Table 4.3: PARP-1 siRNA sequences. The above sequences were synthesized, annealed to their reverse complement and purified by Invitrogen. Invitrogen product number 10620312.

4.2.14 Impact of PARP inhibition on Aprataxin activity: 4.2.14.1 PARP inhibition: C3ABR cells were incubated (or mock treated) with the PARP activity inhibitor 3- aminobenzamide (3AB, 5 mM) for 4 hours. To confirm that the efficiency of PARP inhibition, samples of cells were taken from inhibited and uninhibited flasks. These samples were then divided and treated (or mock treated) with 1 mM hydrogen peroxide in media for 10 minutes. After this period cells were washed twice in PBS and lysed immediately in 5 x PAGE loading buffer. These were then subjected to protein electrophoresis and Western blotting. DNA damage by hydrogen peroxide results in activation of PARP-1s ADP-ribose transferase activity and subsequent automodification. PARP inhibition is demonstrated by a lack of poly-ADP ribose (PAR) synthesis after DNA damage.

Nuclear extracts were generated from the mock and 3AB treated extracts in the same manner as previous in vitro DNA repair experiments (section 3.2.3.2).

4.2.14.2 Experimental conditions: Nuclear extracts from the mock and 3AB treated C3ABR cells (generated as described in section 3.2.3.2) were incubated with Duplex A for the indicated times (5 μg of extract, 2 pmol of substrate in a 10 μL volume at room temperature). Reactions were terminated after the indicated times and analysed as previously described.

157 4.2.15 Gap filling assay: 4.2.15.1 Substrate generation: The radiolabelled Oligo B (unmodified) was annealed with Ligase Bottom 36-mer and Oligo A minus 6 to generate a duplex containing a central single stranded region of 6 nucleotides. Sequences are shown in Table 4.4.

Ligase Bottom 36mer 5' TGTAGTCACTATCGGAATGAGGGCGACACGGATATG Oligo B (unmodified) 5' CATATCCGTGTCG Oligo A minus 6 5' TTCCGATAGTGACTACA Table 4.4: Oligonucleotides used to generate gap filling substrate.

4.2.15.2 Experimental conditions and data analysis: Nuclear extracts (generated as per usual for DNA repair assays, section 3.2.3.2) were incubated with the gap filling substrate (30 μg of protein, 1 pmol of substrate) in Ligase Buffer without ATP but supplemented with dTNPs (10 µM each) at room temperature for the indicated times. Recombinant Aprataxin was added as indicated. Reactions were terminated and resolved as previously. Gap filling is observed as several single nucleotide increases in molecular weight of the substrate. Reactions are quantified by integration of the intensity of each band in the 10 minute lanes using ImageQuant 5.1. The sum of these values provided the total intensity of each lane. The relative abundance of each band was determined by division of its intensity by the total lane intensity. To quantify the repair defect in AOA1 cells, the relative abundance of the fully extended product (Oligo B + 6nt) was determined for each of the four cell lines used in four independent replicates. The relative abundances for the control (C2ABR and C3ABR) and AOA1 (L938 and L939) were pooled. In each instance the control data was designated “100%” and the AOA1 pooled relative abundance was expressed as a proportion of this. This data was subjected to Students t-test in Microsoft Excel (n=4, assuming unequal variances).

4.2.16 Examination of Aprataxin activity in the brain: 4.2.16.1 Preparation of brain extracts: A 6-week female BalbC x C57 Black 6 mouse was culled by cervical dislocation (parental strains were obtained from the Animal Resource Centre). The whole brain was removed by decapitation of the animal followed by generation of two cuts in the skull from the base of the neck towards the ears which facilitated ‘peeling’ the top of

158 the skull away. The whole brain was removed in to a petri dish and dissected into cerebellum, cerebrum, and brainstem. These tissues were mechanically homogenized in Lysis Buffer B by douncing in a ground-glass homogenizer. Lysates were cleared for 1 hour at 13,000 rpm on a benchtop centrifuge. Lysates passed through Biorad G- 50 desalting columns according to manufacturers instructions to remove a layer of insoluble, floating lipids.

4.2.16.2 Experimental conditions: 30 μg of extract from each brain region was incubated in Ligase Buffer with 2 μg of purified rabbit α-Aprataxin antibody or rabbit null serum for 30 minutes at room temperature. Hydrolase reactions were performed at room temperature and were initiated by addition of 2 pmol of Duplex A. The final reaction volume was 10 μL. Reactions were terminated after the indicated times and resolved as described.

159 4.3 RESULTS

4.3.1 Mutations in APTX and Aprataxin protein stability: Ideally the molecular defect in AOA1 would be studied using patient and control neuronal cell lines, however brain biopsies from living patients are not possible due to ethical considerations. Researchers of recessive cerebellar ataxia therefore rely on immortalized lymphoblastiod cells (which can be obtained by Epstein Barr Virus treatment of cells isolated from peripheral blood) or primary fibroblasts (isolated from a skin-punch biopsy). Given the limited number of generations before senescence and relatively slow growth rate of primary fibroblasts, most DNA repair biochemical studies use lymphoblastiod cell lines. To date, APTX mutations described in AOA1 patients have been reported to lead to destabilization of the Aprataxin protein (9,18- 20). This was confirmed by immunoblotting for Aprataxin in control (C2ABR and C3ABR) and AOA1 (L938 and L939) lymphoblastiod cell lines. Aprataxin is present as an approximately 42kDa band of equal intensity in both control cell lines (Figure 4.2). Aprataxin is not detectable in either AOA1 patient cell line indicating that the point mutations in APTX result in a dramatic reduction in Aprataxin protein stability (Figure 4.2).

Figure 4.2: Immunoblot of control and AOA1 lymphoblastiod cell lines. Cell extracts from control (C2ABR and C3ABR) and AOA1 (L938 and L939) cell lines were subjected to 12% SDS-PAGE and transferred onto nitrocellulose. The membrane was subsequently probed with affinity purified rabbit α-Aprataxin antibody and sheep α- rabbit-HRP. Proteins were detected by Western blotting . Equivalent loading is shown by β-actin Immunoblot.

160 4.3.2 Post-translational modification of Aprataxin: Activities of many DNA repair processes are modulated by post-translational modifications (for examples see 21-23). Some common modifications include conjugation of the target protein with protein modifiers like Ubiquitin and SUMO (21), acetylation (23), glycosylation (24) and phosphorylation (25). Modification with Ubiquitin or SUMO (8.5 and 11.6kDa respectively) is detectable by one-dimensional PAGE as an increase in molecular weight of the protein of interest. Such an increase in the molecular weight of Aprataxin has not been reported in the literature or observed in these studies, either in untreated or DNA-damaged cells (personal observations). It is possible for Ubiquitin or SUMO modification of a protein to block antibody binding (preventing detection by Western blot) however given the relative sizes of these modifiers and Aprataxin this is unlikely. Similarly, glycosylation of proteins is detectable as an increase in molecular weight. Monoglycosylation is sometimes detectable as formation of a single higher-molecular weight band. Due to branching and the variable degree of modification of different protein molecules of the same species, protein polyglycosylation normally results in smearing on one dimensional PAGE (for an example see the PAR immunoblot in Figure 4.28). Such smearing is sometimes mistakenly attributed to DNA contamination of the protein preparation. In the absence of a modification-specific antibody, glycosylation of a protein can by treatment of the sample with a glycosylase prior to electrophoresis. Aprataxin runs as a sharp band and I have no evidence to suggest it is glycosylated. Phosphorylation can in some instances be detected as an increase in molecular weight of a protein on one dimensional PAGE, although often the addition of a single phosphate group does not produce a detectable change in the mass/charge ratio of a protein. The Aprataxin protein sequence contains putative phosphorylation sites for several protein kinases (Figure 4.3). To examine the in vivo phosphorylation status of Aprataxin, control and AOA1 (Aprataxin deficient) cell extracts were subjected to two-dimensional gel electrophoresis. Two dimensional electrophoresis of control and AOA1 extracts and subsequent α-Aprataxin immunoblotting revealed that Aprataxin is present in control cells in at least three isoforms (Figure 4.4). L938 is an Aprataxin deficient cell line and as such it was included as a control for non-specific antibody binding (at this point an affinity purified Aprataxin antibody suitable for Western blot was not available). The predominant spot present in C3ABR but not L938 corresponds to the size and pI of non-phosphorylated Aprataxin (apparent molecular

161 weight 42kDa, pI 9.3). This spot is marked ‘0’. Two clear spots of elevated pI are clearly visible (marked +1 and +2), and may correspond to single and double phosphorylation.

Figure 4.3: Putative phosphorylation sites on Aprataxin. The Aprataxin protein sequence was analysed using NetPhosK 1.0 (26), which detects consensus sequences for mammalian kinases (unfiltered, default settings).

162 non-specific

Figure 4.4: Aprataxin is phosphorylated in vivo. Control and AOA1 cell extracts (200 µg) were passively absorbed into 11cm Biorad pH 7-10 IPG strips using the manufacturers standard conditions. First and second dimension resolution was performed as described in section 4.2.1. Gels were transferred onto nitrocellulose and blotted with rabbit α-Aprataxin serum (purified antibody was not available at this time).

Phosphorylation is a common phenomenon in the DNA damage response. Phosphorylation of proteins has numerous functions including alteration of protein- protein interaction profile and enzymatic activity as well as playing a central role in cell signalling. Therefore phosphorylation has the potential to regulate the cellular functions of Aprataxin. The effect of DNA damage on the phosphorylation status of Aprataxin was examined. Control and AOA1 cells were incubated with or without hydrogen peroxide (200 µM) in their media for 10 minutes prior to preparation of total extract. I subsequently immunoprecipitated tyrosine phosphorylated proteins from these extracts using an α-pTyr antibody. Immunoprecipitates were subjected to PAGE and probed with rabbit-α-Aprataxin polyclonal serum (Figure 4.5). While this revealed that Aprataxin is tyrosine phosphorylated, the identity of the tyrosine kinase responsible remains unknown. Serine phosphorylation of Aprataxin was examined in a similar manner however the specificity of the α-pSer antibody proved problematic (data not shown).

163

Figure 4.5: Tyrosine phosphorylation of Aprataxin in vivo. Control and AOA1 cells were treated with 0.2 mM hydrogen peroxide for 10 minutes prior to the preparation of cell extracts by lysis in Lysis Buffer B. Equal quantities of lysate were incubated with protein G agarose overnight at 4°C. The resulting supernatants were incubated with 2 µg of α-pTyr antibody for 6 hours at 4°C before precipitation with protein G agarose. Interacting proteins were eluted in 5 x SDS-PAGE loading buffer prior to protein electrophoresis and Western blotting (rabbit-α-Aprataxin serum).

Possible effectors of Aprataxin’s phosphorylation include the DNA damage-inducible protein kinases ATM and ATR, which phosphorylate [S/T]Q sites. To examine wether these protein kinases phosphorylate of Aprataxin, we employed an in vitro kinase assay. Endogenous ATM was immunoprecipitated from irradiated (10 Gy) or unirradiated control (C3ABR) and ATM kinase-dead (AT3ABR) cell lines according to Lavin et al. (15). Immunoprecipitated ATM was then incubated with recombinant Aprataxin (diagram shown in Figure 4.6, described in (9) or p53 GST fusion proteins (generated by Dr Sergei Kozlov, 15). Substantial phosphorylation of the p53-GST protein was observed, indicating successful immunoprecipitation of ATM (Figure 4.7). Weak phosphorylation of the 266-342 fragment was observed, however some kinase activity is also evident in the AT3ABR immunoprecipitates, indicating that this not ATM mediated.

164

Figure 4.6: Schematic of Aprataxin-GST fusion proteins. Aprataxin domain kinase assay substrates were expressed and purified as N-terminal GST fusion proteins from constructs pGEX5.1. Described in Gueven et al. (9).

Figure 4.7: Aprataxin is not an ATM kinase substrate. Control and A-T lymphoblastiod cell lines were irradiated (10 Gy) or unirradiated and lysed one hour post-treatment. ATM was immunoprecipitated from these lysates and incubated with Aprataxin and p53 GST fusion proteins in the presence of γ-P32 ATP. Reactions were analysed by SDS-PAGE and subsequent autoradiography. Courtesy of Dr Nuri Gueven (Queensland Insititute for Medical Research, Brisbane, Australia).

Similarly, endogenous ATR was immunoprecipitated from untreated or hydroxyurea treated control cells (C3ABR). Again immunoprecipitates were incubated with Aprataxin-GST fusion protein substrates in the presence of γ-P32 ATP according to Lavin et al. (15). Very weak constitutive phosphorylation of the 1-110 fragment was observed (Figure 4.8). Strong phosphorylation of the 98-176 fragment was observed. This fragment contains a single canonical ATR phosphorylation site, serine 168. This site, and the surrounding protein sequence is conserved between human, mouse, chicken and zebrafish Aprataxin homologues (Figure 4.9). This conservation indicates that this site may be functionally significant. Phosphorylation of this site may regulate the activity of Aprataxin.

165

Figure 4.8: Phosphorylation of Aprataxin by ATR. Control cells were treated with 2 µM hydroxyurea (HU, or mock treated) for two hours. ATR was immunoprecipitated from the subsequent lysates. The immunoprecipitates were incubated with Aprataxin- GST fusion proteins in the presence of γ-P-32 ATP. Reactions were analysed by SDS- PAGE and autoradiography. Courtesy of Dr Tanya Vaughn (Queensland Insititute for Medical Research, Brisbane, Australia).

Figure 4.9: Multiple alignment of Aprataxin sequences. Aprataxin protein sequences were aligned using ClustalW, default settings. NCBI accession numbers AAQ74130, XP001477338, NP999894 and XP429199. The putative ATR phosphorylation site is indicated.

4.3.4 Characterization of endogenous Aprataxin- DNA binding: We and others have characterised the DNA binding capacity of recombinant Aprataxin (section 3.3.1 and references 14,27). However the DNA binding capability of endogenous Aprataxin has not been determined. Here I examined wether endogenous Aprataxin could bind to duplex DNA. Biotinylated duplex DNA was attached to streptavidin coated beads, which were used to precipitate DNA binding proteins from nuclear extracts. Precipitated proteins were analysed by Western

166 blotting. I found that endogenous Aprataxin interacts with duplex DNA in the APTX- wild-type cell lines (Figure 4.10).

In the previous chapter I provided a detailed characterisation of the hydrolase activity of recombinant Aprataxin. Having demonstrated that endogenous Aprataxin can also bind to DNA, I examined the hydrolytic activity of endogenous Aprataxin.

Figure 4.10: Endogenous Aprataxin binds DNA. Biotinylated double stranded DNA was attached to streptavidin magnetic beads (Dynal). DNA coated (or mock treated) beads were incubated with extracts from HeLa, control lymphoblastiod or AOA1 cell lines as described in section 4.2.3. Interacting proteins were eluted using SDS-PAGE loading buffer and analysed by electrophoresis. Proteins were transferred onto nitrocellulose and immunoblotted (rabbit-α-Aprataxin affinity purified antibody). XRCC1 was used as a positive control for DNA binding.

4.3.4 Characterization of endogenous Aprataxin- 5’ DNA adenylate hydrolase activity: In the years following the identification of APTX mutations as causal for AOA1, many biochemical studies including those described in the previous chapter, were performed to characterise Aprataxin’s substrates and binding partners in an effort to elucidate its cellular role. As reported in the previous chapter, Aprataxin displayed very poor catalytic activity on the HIT protein substrates tested initially tested (18,27,28). Substantial catalytic activity was reported against the newly identified

167 substrate, 5’ adenylated DNA (20). This particular substrate is generated during a key step in DNA replication and repair (Equation 1.1). A schematic of the 5’ adenylated DNA substrate used in these studies, Duplex A, is shown in Figure 4.11, repeated from the previous chapter for clarity. I have shown that adenylated DNA can accumulate at sites of complex DNA damage (Figure 3.16), providing a link between the biochemical activity of Aprataxin (20) and the DNA-damaging agent sensitivity of AOA1 cells (1,2,9).

= P-32 phosphate agroup 5’ 18-mer 3’ 18-mer

Figure 4.11. Schematic of Aprataxin cleavage of 5’adenylated DNA. Adenylated DNA was generated by incubation of a nicked radiolabelled DNA duplex possessing a 3’ dideoxy break terminus (indicated by “dd”) with recombinant T4 DNA ligase in the presence of ATP.

To demonstrate that Aprataxin is the only protein in a cell capable of repairing this DNA modification, extracts from control (Aprataxin containing) and AOA1 (Aprataxin deficient) cells were incubated with 5’ adenylated DNA. Hydrolysis of the adenylate modification can be observed as an increase in the electrophoretic mobility of the radiolabelled DNA. As shown in Figure 4.12, increasing concentrations of the control cell extracts (C2ABR and C3ABR) result in rapid conversion of the 5’ adenylated oligonucleotide (indicated by AMP-18mer) into the 5’ phosphate 18-mer form. AOA1 cell extract did not convert the substrate into the 5’ phosphate form providing strong evidence that Aprataxin is the only DNA 5’ adenylate hydrolase in human cells.

168 This was confirmed with a time course experiment, where the adenylated DNA substrate was incubated with control and AOA1 nuclear extract for increasing lengths of time (Figure 4.13). No hydrolysis was observed by the AOA1 nuclear extract in the time required for the control extract to hydrolyse all the substrate. This provided definitive evidence that AOA1 patient cell nuclear extracts were unable to hydrolyse 5’adenylated DNA.

Figure 4.12: Hydrolysis of adenylated DNA by endogenous Aprataxin- titration with cell extracts. A. Hydrolysis of 5’DNA adenylate by increasing concentrations of control (C2ABR and C3ABR) and AOA1 (L938 and L939) cell extracts. Reactions were initiated by addition of Duplex A (2 pmol) and allowed to proceed for 2 minutes before termination and denaturing electrophoresis. B. Graphical representation of duplicate extract titrations as in A. Lines show the logarithmic curve of best fit of the means of these duplicate sets. Method described in section 4.2.4.

169

Figure 4.13: Hydrolysis of adenylated DNA by endogenous Aprataxin – time-course. A. DNA adenylate hydrolysis over time by control (C3ABR) and AOA1 (L938) nuclear extracts. Reactions contained 2 µg of nuclear extract and were initiated by addition of Duplex A (2 pmol). Reactions were allowed to proceed for the indicated times prior to termination. B. Graphical representation of duplicate time courses. The logarithmic curve of best fit is shown. Method described in section 4.2.4.

An additional experiment was performed to rule out the possibility that another protein (different from Aprataxin) is defective in AOA1 and is responsible for the observed adenylate hydrolysis. Addition of a purified Aprataxin antibody to control cell extract impaired their ability to hydrolyse 5’ adenylated DNA (Figure 4.14). This confirmed that Aprataxin is indeed a DNA-adenylate hydrolase. Combined with the total deficiency of hydrolase activity in AOA1 cells, this provides compelling evidence that Aprataxin is the only DNA-adenylate hydrolase in human cells.

170

Figure 4.14: Inhibition of DNA-adenylate hydrolysis by an Aprataxin specific antibody. Inhibition of DNA adenylate hydrolysis by addition of Aprataxin antibody. 5 µg of control (C2ABR and C3ABR) and AOA1 (L938) extract was incubated with 200 ng of anti-Aprataxin antibody (+) or null serum (-) in a 8 µL volume for 20 minutes. Reactions were initiated by addition of 2 picomoles of Duplex A (1 pmol/µL) prior to termination after the indicated times. Method described in section 4.2.4.

We demonstrated in the previous chapter that adenylated DNA can accumulate at single strand breaks possessing a range of modified 3’ termini (Figure 3.16). To mimic an abortive ligation caused by an oxidised single strand break, 5’ adenylate and 3’ 8-oxo-dG modified oligonucleotides were annealed to a scaffold to generate a doubly modified nick (described in section 4.2.5, schematic shown in Figure 4.15). The rate of repair of the adenylate in this structure was compared to that of an adenylate adjacent to an unmodified 3’ terminus. Incubation of these two structures with control cell extracts revealed that the DNA-adenylate hydrolase activity of Aprataxin is not affected by oxidation of the adjacent terminus (Figure 4.16), suggesting that adenylates arising in wild-type cells due to abortive ligation of 3’ oxidized breaks are repaired efficiently.

171

Figure 4.15: Schematic of double modified nick duplex. Nicked DNA duplex radiolabelled and adenylated on the 5’ break terminus and possessing either an unmodified (hydroxyl) or an 8-oxo-dG 3’ terminus was generated (section 4.2.5.1) to examine the effect of 3’ terminal damage on the 5’ processing activity of Aprataxin.

Figure 4.16: Impact of adjacent 3’ modification on DNA-adenylate hydrolysis. Nicked DNA duplex (2 pmol), radiolabelled and adenylated on the 5’ break terminus and possessing either an unmodified (hydroxyl) or an 8-oxo-dG 3’ terminus (schematic Figure 4.15) was incubated with control (C2ABR and C3ABR) cell extract for the indicated time. Hydrolysis of the 5’ adenylate group over time was observed as an increase in electrophoretic mobility of the radiolabelled species on a denaturing acrylamide gel. Substrate generation and reaction conditions are described in section 4.2.5.

4.3.5 Single strand break repair in AOA1 cell extracts: To examine the effect of oxidation and Aprataxin deficiency on single strand break repair, nicked 5’ labelled duplexes possessing either unmodified or 3’ 8-oxo-dG termini were generated (identical to the substrates generated in section 3.2.3, depicted in Figure 3.13). These were incubated with control and AOA1 nuclear extracts for increasing lengths of time and formation of adenylated and ligated products was monitored (according to section 4.2.6). As shown in Figure 4.17, the presence of a 3’ 8-oxo-dG inhibits ligation in both control and AOA1 cell extract. The extent of inhibition due to the 3’ 8-oxo-dG is similar in both cell lines, indicating that the observed inhibition of single strand break repair is not Aprataxin-related.

172

Figure 4.17: 3’ 8-oxo-dG inhibits single strand break repair. Nicked duplex DNA (2 pmol) radiolabelled on the 5’ break terminus and possessing either a hydroxyl (control) or an 8-oxo-dG 3’ break terminus (for a schematic see Figure 3.13) was incubated with control (C3ABR) or AOA1 (L938) cell extract for 2 to 30 minutes. Method described in section 4.2.6.

4.3.6 Oxidative Stress in AOA1 cells: We have demonstrated that oxidation at a single strand break inhibits ligation (Figure 4.17), and using a recombinant model showed that 3’ break oxidation can lead to 5’ adenylation (Figure 3.16). Oxidative stress, which can be produced endogenously by mitochondria or exogenously by environmental agents, results in oxidation of proteins, lipids and DNA (for examples relevant to neurodegeneration see references (29-31). In vivo oxidation of DNA results in several modifications which are removed by BER, 8-oxo-dG being the most abundant (reviewed in section 1.3.2.4). Elevated levels of oxidative stress have been observed in several spinocerebellar ataxias, including A-T (32,33) and AOA2 (16). More recently elevated levels of 8-oxo-dG were observed in AOA1 patient brain specimens (34), indicating a potential defect in BER.

The primary function of APE1 appears to be incision of the phosphodiester backbone at abasic sites as part of the BER process (reviewed in section 1.3.2.4). Additional functions for APE1 have been characterised. Parsons et al. showed that APE1 is the only enzyme capable of hydrolysis of a 3’ 8-oxo-dG at a single strand break (35). It

173 also stimulates the activity of OGG1, the primary mammalian 8-oxo-dG glycosylase (36,37). The elevated level of 8-oxo-dG staining in AOA1 patient brains could be due to a deficit in APE1. To test this idea I examined the expression level of APE1 in AOA1 cells. APE1 levels were examined by immunoblotting cell extracts from corrected and uncorrected AOA1 cell lines (Figure 4.18). I found that Aprataxin deficient (FD105 M20) cells have substantially less APE1 protein than corrected cells. APE1 stimulates the activity of many bifunctional DNA-glycosylases, including OGG1 which excises 8-oxo-dG. Based on this I proposed that AOA1 cells may have a defect in excision of 8-oxo-dG.

B.

Figure 4.18: Stability of APE1 in AOA1 cell lines. A. Total extract (50 µg) from corrected (FD105 M21) or uncorrected (FD105 M20) cell lines was subjected to SDS- PAGE and transferred onto nitrocellulose. Aprataxin deficiency in FD105 M20 was confirmed by immunoblotting with the purified rabbit antibody. APE1 levels were examined using an antibody provided by Prof Grigory Dianov (University of Oxford, Oxford, UK). AIF shows equal loading. B. APE1 protein in FD105 M20 and M21 was quantified using ImageQuant 5.1 and normalized to AIF loading. The abundance of APE1 in FD105 M20 is expressed relative to FD105 M21. Method described in section 4.2.7.

174

We subsequently examined whether the observed APE1 deficiency caused a corresponding increase in 8-oxo-dG lesions in AOA1 cells. 8-oxo-dG was detected in the DNA of individual cells by immunostaining as described in section 4.2.8. I identified increased levels of 8-oxo-dG staining in AOA1 patient fibroblasts (FD105) compared to control cells (NFF) under normal growth conditions and an increase in staining of both cell lines in response to hydrogen peroxide treatment (Figure 4.19). This is consistent with the hypersensitivity of AOA1 cells to hydrogen peroxide, the reduced level of APE1 in AOA1 cells and the capacity of hydrogen peroxide to induce DNA oxidation and formation of 8-oxo-dG.

175

A. B.

Figure 4.19: Oxidative DNA damage in AOA1 cells. A. Immunofluorescence detection of 8-oxo-dG levels in normal (NFF) and AOA1 (FD105) fibroblasts either untreated or treated with 1 mM H2O2 for 30 min in aerobic conditions. After treatment, H2O2-containing media was replaced with fresh media and cells were incubated for an additional 2 h before fixation and 8-oxo-dG detection. A representative image is shown. B. Fluorescence intensity was quantitated on at least 60 individual cells for each treatment using the public domain software Image J (NIH, USA). Student’s t-test analysis of 8-oxo-dG staining in both cell lines demonstrated a significant difference in 8 oxo-dG levels between untreated NFF and

FD105 (*, p<0.001); NFF untreated and treated with H2O2 (#, p<0.001); NFF and

FD105 treated with H2O2 (**, p<0.001). No significant difference between FD105 and H2O2-treated FD105 ($, p=0.5) was observed. Method described in section 4.2.8. Method described in section 4.2.8. Experiment performed by Olivier Becherel (Queensland Institute for Medical Research, Brisbane, Australia). The scale bar indicates 20 µm.

Having determined that Aprataxin deficiency results in an elevated level of oxidative DNA damage, I examined wether Aprataxin deficient cells have elevated levels of oxidative stress generally. Free radicals are capable of damaging all biological macromolecules, including proteins and lipids as well as DNA. Elevated levels of free radicals from either endogenous or exogenous sources can cause a range of oxidative protein modifications including methionine sulfoxide, oxo-histidine, de-aminated

176 lysine, hydroxy and peroxy derivatives of valine, leucine and tyrosine, and nitro tyrosine (reviewed in 38). Tyrosine nitrosylation is a commonly used biomarker for oxidative protein damage and oxidative stress. I examined tyrosine nitrosylation in Aprataxin deficient and corrected cells by immunostaining and found that Aprataxin deficiency results in an elevated level of tyrosine nitrosylation (Figure 4.20). Combined with the elevated levels of 8-oxo-dG in AOA1 cells this indicates that Aprataxin deficiency causes an elevation in the basal level of oxidative stress.

DAPI

Figure 4.20: Oxidative protein damage in AOA1 cells. Immunofluorescent detection of oxidized proteins in AOA1 and corrected cell lines. Untreated FD105 M20 and M21 cells were fixed in 10% formalin and stained with an α-nitroTyrosine antibody. Staining was detected using an AlexaFluor594 conjugated secondary antibody. Coverslips were counterstained with DAPI and mounted in Moviol. Images were captured on a Zeiss AxioSkop using ZeissVision software. Representative images are shown. Method described in section 4.2.9. The scale bar indicates 20 µm.

4.3.7 Subcellular distribution of Aprataxin protein and activity: Aprataxin is a nuclear protein, present in both nucleoplasmic and nucleolar compartments (8,9). To confirm this the distribution of an Aprataxin-GFP fusion protein in HeLa cells was observed by fluorescent microscopy (Figure 4.21a). Diffuse nucleoplasmic and intense nucleolar fluorescence was observed, as indicated by Nucleolin and DAPI counterstaining. The distribution of Aprataxin in lymphoblastiod cells was determined by sub-cellular fractionation and immunoprecipitation (Figure 4.21b). Immunoprecipitation from an Aprataxin

177 deficient cell line (L938) was used to control for non-specific antibody binding. To compare the sub-cellular distribution of Aprataxin with its interacting proteins, nucleoplasmic and nucleolar fractions of control cells were subjected to immunoblotting (Figure 4.22). I determined that in control LCLs, Aprataxin is distributed in roughly equal concentrations between the nucleolus and the nucleoplasm (fraction integrity was confirmed by RNA polymerase II and Nucleolin distribution). This contrasts with the HeLa immunofluorescence, where Aprataxin- GFP is concentrated in the nucleolus. It seems possible that the ratio of nucleolar to nucleoplasmic Aprataxin could differ between cell types, or that over-expression of Aprataxin with a fusion partner alters the normal protein distribution. Interestingly, Aprataxin’s interacting proteins PARP-1 and XRCC1 display different distribution patterns between the nucleoplasm and the nucleolus. XRCC1 is present in both fractions in roughly equivalent concentrations, whereas PARP-1 is present almost exclusively in the nucleoplasm.

178

A. B.

Figure 4.21: Cellular distribution of Aprataxin. A.GFP-Aprataxin transiently transfected HeLa cells were fixed in 4% paraformaldehyde in PBS, stained for nucleolin and counterstained with DAPI. Images were obtained on a Zeiss AxioSkop fluorescent microscope using ZeissVision software (courtesy of Dr Olivier Becherel, construct generated by Gueven et al., 9). A representative image is shown. The scale bar indicates 20 µm. B. Cytoplasmic, nucleoplasmic and nucleolar fractions were generated from control and AOA1 lymphoblastiod cell lines. Extracts were cleared with protein G agarose overnight at 4°C. Subsequently 10 µL of sheep-α Aprataxin- FHA affinity purified antibody was added to 500 µg of extract overnight. Proteins were precipitated with proteinG agarose and eluted in SDS-PAGE sample buffer for electrophoresis and Western blotting (rabbit-α-Aprataxin serum). The sheep light chain band demonstrates that equal quantities of IP antibody were used for each sample. Cyto, NP and No denote cytoplasmic, nucleoplasmic and nucleolar extracts respectively. Method described in section 4.2.10.

179

Figure 4.22: Cellular distribution of Aprataxin. Control (C2ABR) lymphoblastiod cells were fractionated as described in methods. 50 µg each of nucleoplasmic and nucleolar protein was subjected to electrophoresis on a 6-12% gradient polyacrylamide gel and transferred onto nitrocellulose for Western blotting. NP and No denote nucleoplasmic and nucleolar extracts respectively. Method described in section 4.2.10.

Localization of a protein or relocalization to a different compartment can often modulate its activity. For example, relocalization of MDM2 from the nucleolus to the nucleoplasm enables it to interact with the tumour suppressor p53 (39). To determine whether Aprataxin possesses the same activity in both the nucleolus and the nucleoplasm I examined the 5’DNA adenylate hydrolase activity of nuclear fractions of control cells. Incubation of equal masses of nucleoplasmic and nucleolar protein extracts (generated as described in section 4.2.10.2) with adenylated DNA revealed that Aprataxin is active in both fractions (Figure 4.23). I determined that nucleolar protein fraction displays a higher specific activity than the nucleoplasmic fraction, even though Western blotting indicated that these compartments contain roughly equal relative abundances of Aprataxin (in terms of molecules of Aprataxin/µg of protein, Figure 4.22). This indicates that the elevated level of hydrolase activity in the nucleolus is not due to a higher relative abundance of Aprataxin protein.

180

Figure 4.23: DNA-adenylate hydrolase activity of nucleoplasmic and nucleolar fractions. A. DNA adenylate hydrolysis by nucleoplasmic and nucleolar extracts. Nucleoplasmic and nucleolar extract (5 µg) from C2ABR were incubated with 2 picomoles of Duplex A in 10 µL for the times indicated. N.E is a no extract control. B. Graph of replicates of A. De-adenylation activity of increasing amounts of extract was measured twice using independent preparations of C2ABR fractions and once using C3ABR fractions. Lines show the logarithmic curve of best fit of the mean of these three reaction sets. Methods described in section 4.2.10.

Possible mechanisms exist for this alternate regulation of Aprataxin activity in the nucleoplasm versus the nucleolus. These include activation or inhibition of the protein in either compartment by differential protein-protein interactions or post-translational modifications. In the previous chapter I demonstrated that HIT domain activity is regulated by the FHA domain (section 3.3.6). Thus, I propose that the difference in activity between the nucleoplasmic and nucleolar fractions may be due to different protein-protein interactions via the this motif. PARP-1 interacts with the FHA domain

181 of Aprataxin and is present in the nucleoplasm in much higher concentrations than the nucleolus (Figure 4.22). Having established in the previous chapter that deletion of the FHA domain stimulates hydrolase activity, it seems possible that protein-protein interactions through this motif have the potential to regulate the activity of the HIT domain.

4.3.8 Regulation of Aprataxin by interacting proteins: To explore the possibility of regulation of the HIT domain by Aprataxin-interacting proteins, the activity of Aprataxin was measured in a panel of cell lines defective for key proteins in DNA repair (XRCC1 and PARP-1).

The CHO cell line EM9 is deficient in full length XRCC1. This cell line was generated by ENU mutagenesis of the parental strain (AA8) to generate a cell line sensitive to X-rays (40). The mutated protein was identified by complementation of this hypersensitivity and subsequent sequencing of the XRCC1 gene in these cells. This revealed that in EM9 XRCC1 has a premature truncation codon after 220 out of 634 codons (41). I examined the effect of XRCC1 deficiency on the DNA-adenylate hydrolase activity of Aprataxin using EM9 and AA8 cell extracts. I was unable to examine the level of Aprataxin in AA8 and EM9 as our Aprataxin antibodies do not react with the hamster protein, but found that deficiency of XRCC1 does not cause a defect in Aprataxin activity (Figure 4.24).

182 AMP-18mer

18mer

Figure 4.24: XRCC1 deficiency does not affect Aprataxin activity. A. The impact of XRCC1 deficiency on Aprataxin activity was measured by incubation of control (AA8) and XRCC1 (EM9) deficient CHO cell extracts (5 µg each) with 2 picomoles of Duplex A (AMP-18mer) for the indicated times. N.E is a no extract control. B. Deficiency of XRCC1 was confirmed by immunoblotting of extracts. Loading is shown by β-actin (courtesy of Dr Olivier Becherel, Queensland Institute for Medical Research, Brisbane, Australia). Method described in section 4.2.12.

Western blotting of wild-type and PARP-1 deficient extracts revealed that loss of PARP-1 results in destabilization of the Aprataxin protein (Figure 4.25), since Aprataxin was not detectable in PARP-1 knockout cell extracts.

Figure 4.25: PARP-1 is required for stabilization of Aprataxin. Total cell extracts from wild-type and PARP-1 knockout MEFs were prepared and subjected to SDS-PAGE and Western blotting. PARP-1 and Aprataxin were detected as previously. Upstream

Binding Factor (UBF), isoforms 1 and 2 were used as a loading control.

The dependence of Aprataxin stability on PARP-1 was further confirmed by transient depletion of PARP-1 in vivo using siRNA technology. PARP-1 siRNA duplexes were

183 transfected overnight into confluent HeLa cells using Lipofectamine 2000 as described in section 4.2.13. Knockdown of PARP-1 resulted in destabilization of Aprataxin (Figure 4.26). Transfection of siRNA duplex 1 resulted in a greater than 80 percent reduction in the levels of PARP-1 and a greater than 35 percent reduction in the level of Aprataxin protein. Transfection of duplex 2 resulted in a greater than 60 percent reduction in PARP-1 protein level and a corresponding 35 percent reduction in Aprataxin level. Aprataxin protein levels were not affected by transfection with siRNA duplex 3, which failed to cause knockdown PARP-1 protein. Together with Figure 4.25, this indicates that PARP-1 is critical for the stabilization of Aprataxin. Based on this I expect that PARP-1 knockout cells would display a defect in Aprataxin activity.

184

Figure 4.26: Stability of Aprataxin protein after PARP-1 siRNA. A. Confluent HeLa cells were transfected with control or PARP-1 specific siRNA using Lipofectamine 2000 as per manufacturers instructions. Cells were harvested 48 hours after transfection and total cell extracts were analysed by SDS-PAGE and immunoblotting. PARP-1 and Aprataxin were detected as previously, and beta-actin is shown as a loading control. B. Levels of Aprataxin and PARP-1 in part A were quantified using ImageQuant 5.1.

185

The DNA-adenylate hydrolase activity of PARP-1 wild-type and knockout cells were subsequently analysed (as described in section 4.2.12). As predicted based on the apparent destabilization of Aprataxin in the absence of PARP-1, the knockout cell line displayed very low levels of DNA-adenylate hydrolase activity (Figure 4.27). Control MEF nuclear extracts hydrolysed the adenylated substrate in a rapid manner, with almost no substrate remaining at the earliest time point, 1 minute. In contrast, substrate is detectable in PARP-1 knockout cell extract beyond 5 minutes. The rapid formation of low molecular weight bands in the PARP-1 knockout lanes indicates that what initially appears to be repair of the 5’ adenylate group is most likely substrate degradation due to non-specific 3’ to 5’ nuclease activity.

Figure 4.27: Lack of DNA adenylate hydrolase activity in PARP-1 knockout cells. The impact of deficiency of PARP-1 on Aprataxin activity was measured by incubation of PARP-1+/+ and PARP-1-/- nuclear extracts (5 µg) with 2 picomoles of Duplex A (AMP-18mer) for the indicated times prior to sample denaturation. N.E is a no extract control. Methods are described in section 4.2.12.

We concluded that PARP-1 protein is critical for maintenance of normal Aprataxin levels and is therefore indirectly required for DNA-adenylate repair. PARP-1 is a DNA damage-activated ADP-ribose polymerase. The synthesis of ADP-ribose serves as a recruitment signal for many DNA repair proteins to sites of DNA breaks. To discern the importance of PARP-1’s catalytic activity with respect to DNA-adenylate

186 hydrolysis, control cells were treated with the PARP inhibitor 3AB (note that this molecule is a substrate mimetic and as such inhibits the entire PARP enzyme family). Inhibition of PARP activity was demonstrated by treatment of cells with hydrogen peroxide and PAR immunoblotting (Figure 4.28). Hydrogen peroxide treatment generates breaks which induce PAR synthesis in 3AB untreated cells. Lack of DNA- damage induced PAR synthesis in 3AB treated cells demonstrates effective PARP inhibition. PARP inhibition had no effect on the abundance of Aprataxin in the cells (Figure 4.28).

The activity of Aprataxin in 3AB treated and untreated cell extracts was subsequently determined. I found that inhibition of PARP did not result in a noticeable reduction in the activity of Aprataxin in cell extracts (Figure 4.29). From this I conclude that PARP-1 is required for Aprataxin protein stability and is therefore indirectly required for DNA-adenylate repair in vitro, but that PARP activity is not required for either stabilization or recruitment to the substrate in vitro.

Figure 4.28: Inhibition of PARP activity by 3AB treatment. Control lymphoblastoid cells were incubated with the PARP inhibitor 3-aminobenzamide (3AB) or the mock treated as described in section 4.2.14. 3AB treated and untreated cells were then mock treated or incubated with hydrogen peroxide (1 mM final concentration in media) for 10 minutes. After treatment cells were washed in PBS and lysed immediately in SDS- PAGE loading buffer. Protein equivalent to 1x106 cells was analysed. Method described in section 4.2.14.1.

187

Figure 4.29: PARP activity is dispensable for DNA-adenylate hydrolysis. The impact of PARP activity on DNA-adenylate hydrolysis by Aprataxin measured by treatment of cells with 3-aminobenzamide (3AB) (or control) prior to extract preparation. 3AB and DMSO treated extracts (5 µg each) were incubated with 2 picomoles of Duplex A (AMP-18mer) for the indicated times. Reactions were terminated with formamide loading buffer and analysed by sequencing gel electrophoresis. Method described in section 4.2.14.2.

4.3.9 Base excision repair in AOA1 cell extracts: PARP-1 constitutively interacts with base excision repair proteins including DNA polymerase β (polβ) (reviewed in 42). Many studies have found that PARP-1 deficient cells display a multitude of DNA repair defects (43-45), including a defect in the gap- filling phase of BER (46). Based on previous studies (46), I designed an oligonucleotide based substrate to assess the efficiency of PARP-1 dependant gap filling in AOA1 cell lines. This 36 nucleotide duplex contains a central 6 nucleotide single stranded region (substrate generation described in section 4.2.15, schematic shown in Figure 4.30). When supplied with dNTPs, BER proficient cell extracts extend across this single stranded region.

188

Oligo A-6

Figure 4.30: Schematic of in vitro gap filling assay. A duplex with an internal single stranded region was generated by annealing three DNA oligonucleotides as shown. One of these, denoted Oligo B, was 5’P-32 labelled. The six nucleotide single stranded region can be extended across by cell extracts in the presence of dNTPs. The addition of each nucleotide can then be examined by denaturing PAGE and autoradiography. Substrate generation is described in section 4.2.15.

Using this system I confirmed that PARP-1 deficient cells display a defect in gap filling (Figure 4.31). PARP-1 knockout cells are capable of extending a single nucleotide from the 3’ terminus of the radiolabelled strand, but are not able to efficiently extend further. This is consistent with the findings of Dantzer et al. who reported that PARP-1 knockout cells display a mild impairment of short (single nucleotide) patch repair activity and a major deficit in long (multiple nucleotide) patch repair activity (46). The authors proposed that this distinction between long and short patch repair may be due to the use of different repair complexes for single and multiple nucleotide extension reactions.

189

A.

B.

Figure 4.31: Defective gap filling in PARP-1 knockout cells. A. Wild-type (PARP-1 +/+) or deficient (PARP-1 -/-) MEF nuclear extracts were incubated with the gap filling substrate and analysed according to section 4.2.15. B. The relative abundances of each band in the 10 minute reactions were analysed as described in section 4.2.15.

Having validated that efficient long (but not short) gap filling requires PARP-1, we analysed the gap filling capacity of AOA1 cell extracts. I found that AOA1 cells

190 possess a reduced capacity to extend across a single stranded region of DNA to generate a fully extended product (Figure 4.32).

Figure 4.32: Reduced gap filling efficiency in AOA1 cells. Control (C2ABR and C3ABR) and AOA1 (L938 and L939) nuclear extracts were incubated with the short gap filling substrate (1 pmol) for the indicated times prior to sample denaturation and electrophoresis, according to section 4.2.15.

Quantification of the amount of fully extended product (Oligo B + 6) in control and AOA1 cell lines is shown in Figure 4.33. Quantification of this single experiment revealed that repair activities are similar between the control and between the AOA1 cell lines, and that both AOA1 cell lines display a repair defect. Pooled, normalized data from quadruplicate independent experiments (Figure 4.34) revealed that Aprataxin deficiency results in a statistically significant gap filling defect in generation of fully extended product (Oligo B+6 nucleotides, 54% of control activity, p = 0.001). Given that the duplexes used in these experiments are quite short (36 nucleotides), these data may represent distributive DNA synthesis by DNA polymerase β rather than PCNA-dependant repair (which would typify LP BER). However in either circumstance I have demonstrated that this activity measures PARP-1 dependant repair (Figure 4.31), thus the functional interaction between Aprataxin and PARP-1 can be assessed in this manner.

While both Aprataxin and PARP-1 deficient cells display a defect in gap filling, the nature of this defect differs between the cell types. Quantification of the intensity of each band in the 10 minute time points of PARP-1 knockout and wild-type reactions confirmed that knockout cells are able to extend a single nucleotide from the substrate

191 but cannot efficiently synthesize longer strands (Figure 4.31). Thus PARP-1 knockout cells which have a specific defect in gap filling show an accumulation of single nucleotide extended product. In contrast, quantification of control and AOA1 gap filling reactions revealed that AOA1 cells are slightly less efficient than controls at each step and do not display an accumulation of any reaction intermediates (Figure 4.35).

Figure 4.33: Quantification of gap filling by control and AOA1 cells. The phosphorimage shown in Figure 4. 34 was analysed using ImageQuant 5.1 (default settings). Shown is the intensity of the fully extended product (Oligo B+6nt) relative to the total amount of isotope in that particular lane. The 20 minute time point was used in this instance.

192

Figure 4.34: Quantification of gap filling in control and AOA1 cells- quadruplicate experiments. The 20 minute time points of four independent short gap filling experiments were quantified as shown in Figure 4. 35. For each experiment the mean abundance of fully extended product was determined for the control (C2ABR + C3ABR) and AOA1 (L938 + L939) cell lines. The control average was set to “100 percent” and the AOA1 activity expressed as a proportion of this.

193 35

30

25

20 relative abundance 15

10

5 C3ABR

0 C2ABR L938 1 2 3 L939 4 5 nucleotides extended 6 Figure 4.35: Gap filling by control and AOA1 cells- quantification of reaction intermediates and products. The relative abundance of each band in the 20 minute reactions control and AOA1 reactions were analysed in ImageQuant 5.1. Intensities for each band are plotted as relative abundance (intensity of band compared to total signal in that lane).

To determine if Aprataxin protein plays a direct role in the gap filling phase of BER, recombinant Aprataxin was added to AOA1 cell extract. If Aprataxin deficiency is directly responsible for the impaired of gap filling activity of AOA1 cells, addition of recombinant protein should enhance their activity. I found that recombinant Aprataxin had no effect on the rate of gap filling by AOA1 cell extracts, indicating that Aprataxin is not directly involved (Figure 4.36). Based on failure of recombinant Aprataxin to correct the BER defect in AOA1, this repair defect may be due to inappropriate protein modification or a deficit in stabilization or recruitment of repair complexes. Given that PARP-1 was shown to be required for stabilization of Aprataxin, it is possible that lack of Aprataxin affects the stability of its interacting proteins, including PARP-1.

194

Figure 4.36: Effect of recombinant Aprataxin on gap filling. Control (C2ABR) and AOA1 (L938) cell extracts with or without recombinant full-length Aprataxin (1 pmol) were incubated with the gap filling substrate (1 pmol) for the indicated times.

To test this hypothesis, the level of PARP-1 protein was examined in Aprataxin positive and negative isogenic cell lines, as described in section 4.2.7. Correction of AOA1 cell lines with full length APTX cDNA resulted in an increase in PARP-1 levels (Figure 4.37). The Aprataxin deficient cell line does have detectable PARP- 1. I therefore conclude that Aprataxin stabilizes PARP-1, but that Aprataxin deficiency does not lead to complete PARP-1 deficit. Given that PARP-1 plays a central role in direct single strand break repair and BER, this deficiency may be partially responsible for the hypersensitivity of AOA1 cells to agents which induce oxidative DNA damage.

195

Figure 4.37: Levels of PARP-1 in Aprataxin deficient and corrected cell lines. Total cell extracts from immortalized AOA1 fibroblasts (empty vector, FD105 M20) and corrected AOA1 fibroblasts (full length cDNA, FD105 M21) were subjected to SDS-PAGE and Western blotting for Aprataxin and PARP-1. Apoptosis Inducing Factor (AIF) is shown as a loading control.

This BER deficiency in AOA1 cells probably compounds the effects of their inability to repair 5’ adenylated DNA breaks. Long patch BER is a mechanism for removing ‘non-processible’ 5’ DNA damage (reviewed in section 1.3.2.4). The 3’ terminus of the break is used to prime DNA polymerase extension and the undamaged, intact strand is used as a template. As extension occurs the damaged 5’ terminus is displaced and is subsequently cleaved to generate a ligatable nick. AOA1 cells therefore have an absolute defect in direct (Aprataxin dependant) repair of 5’ adenylates and a reduced ability to excise 5’ DNA damage using long patch BER. They also have a reduced level of APE1 which is probably responsible for their elevated level of 8-oxo-dG modified DNA. AOA1 cells subsequently will have more 3’ 8-oxo-dG than control cells, which may cause a higher frequency of abortive ligations. The cumulative effect of these multiple DNA repair defects probably results in the gradual accumulation of 5’ adenylated DNA in the genomes of AOA1 patients. There is some evidence that high levels of DNA damage can inhibit transcription, and deficiency of vital transcripts can lead to neuronal cell death (47,48). I propose that the accumulation of DNA damage in coding regions of AOA1 patient genomes causes loss of transcripts critical to neuronal survival, resulting in progressive neurodegeneration.

4.3.10 Aprataxin activity in the brain: As AOA1 patients display neuronal degeneration specifically in the cerebellum (49), the activity of Aprataxin in three major brain regions was examined. We do not have access to human brain specimens therefore I used tissue from a wild-type mouse. This

196 mouse brain was dissected into brainstem, cerebrum and cerebellum and tissue extracts were generated by homogenization of the tissue in Lysis Buffer B. Equal amounts of protein were incubated with adenylated DNA (Duplex A) for up to 10 minutes (Figure 4.38). Analysis of the reaction products revealed that cerebellar extract has a higher DNA-adenylate hydrolase activity than brainstem or cerebrum extract. Unfortunately the Aprataxin antibodies available to us at the time were not reactive with the mouse homolog. I was therefore unable to inhibit this DNA- adenylate hydrolysis (Figure 4.38) or examine the level of Aprataxin present in these brain regions.

Figure 4.38: Distribution of Aprataxin activity in the murine brain. Murine brain regions were dissected and homogenized in Lysis Buffer B. 30 µg of extract was incubated with 2 µg of purified rabbit α-Aprataxin or rabbit null serum for 30 minutes at room temperature. Reactions were subsequently initiated by addition of Duplex A (2 pmol). Reactions were denatured after the indicated times with formamide-EDTA loading buffer and analysed by sequencing gel. Extract generation and reaction conditions are described in section 4.2.16.

197

4.4 DISCUSSION

4.4.1 Post translational modification of Aprataxin: A range of post translational modifications have the potential to modulate the activity of an enzyme, including acetylation, glycosylation, ubiquitinylation and phosphorylation. I determined that Aprataxin is phosphorylated in at least two different locations: an undetermined tyrosine and an SQ/TQ site which is likely to be the conserved residue S168. This SQ site is phosphorylated by the PI3 like protein kinase ATR but not by the homologous kinase ATM. ATR is activated single stranded regions of DNA and stalled replication forks (50), whereas ATM is activated by double strand breaks (51). AOA1 cells are hypersensitive to agents which induce direct and indirect single strand breaks but display a lesser degree of hypersensitivity to double strand break inducing agents (1,9). Given this sensitivity profile, the distinct roles of ATM and ATR in DNA damage signalling and the phosphorylation of Aprataxin by ATR, I speculate that the activity of Aprataxin may be modulated by phosphorylation by ATR. This could be further examined by analysis the DNA- adenylate hydrolase activity of ATR defective (Seckel) cells.

4.4.2 Hydrolase activity of endogenous Aprataxin: The last two years have seen substantial clarification of the enzymatic activity of Aprataxin and its role in DNA repair. Initial reports of the hydrolase activity of recombinant Aprataxin characterised its limited activity against mono or diadenosine based nucleotides (18,27,28). Following this, Aprataxin was found to hydrolyse the ligation intermediate, DNA 5’ adenylate (14,20). I demonstrated that this structure is generated as a result of attempted ligation of single strand breaks with damaged 3’ termini (Figure 3.16). Initial reports determined that Aprataxin is the only protein capable of repairing this modification (14,20). I substantiate these findings using Aprataxin deficient cell lines derived from different AOA1 patients and inhibition of endogenous Aprataxin with a function-blocking antibody.

Activity of proteins can be modulated by subcellular localization. For example the cAMP-dependant activation of PKA can be regulated by localization via its interaction with A-Kinase Anchoring Proteins, which tether the kinase to a range of

198 intracellular sites. This allows PKA to respond to discrete cAMP gradients with appropriate substrate selection (reviewed in 52). Given the localization of Aprataxin to both the nucleoplasm and nucleolus (8,9), it was important to determine whether this difference in localization corresponds to a difference in enzymatic activity. I determined that Aprataxin is active in both the nucleolus and the nucleoplasm. Interestingly I found elevated levels of DNA-adenylate hydrolase activity in the nucleolus compared to the nucleoplasm, despite the very low concentration of DNA in nucleoli. This is surprising given the proposed DNA repair function of Aprataxin. However several possibilities may explain the necessity of high Aprataxin activity in nucleoli despite their low DNA content. All are related to the very high density of transcription in the nucleolus. DNA in the nucleolus is highly transcribed and is therefore loosely packaged and thus may be more exposed to the aqueous environment than tightly packed chromatin. This elevated solvent access to the DNA molecule may result in a higher frequency of lesions (i.e. lesions/Mb) in this compartment. A higher abundance of DNA lesions would cause an elevated frequency of abortive ligations in nucleolar compared to nucleoplasmic DNA. Secondly, DNA damage in coding regions may cause transcription fork collapse resulting in double strand break formation (similar to the situation with replication fork collapse, reference 53). As the density of active transcriptional machinery in the nucleolus is very high, lesions would need to be repaired rapidly to avoid transcription fork collapse and generation of toxic double strand breaks. Finally, damage to a large enough proportion of rDNA repeats in the nucleolus could block RNA polymerase 1 and ‘starve’ a cell of vital rRNA transcripts (54,55). An elevated level of Aprataxin activity may be required to compensate for any or all of these problems. As the abundance of Aprataxin (molecules/mg of extract) is similar between nucleoplasmic and nucleolar fractions this difference in activity cannot be mediated by an unequal distribution of the enzyme (Figure 4.22). In the previous chapter I reported that the hydrolase activity of recombinant Aprataxin is stimulated by deletion of the FHA domain (Figure 3.20) and here I found that that the FHA-domain interacting protein PARP-1 plays an important role in the stabilization of Aprataxin. I also demonstrated that the DNA damage-responsive nucleoplasmic kinase ATR phosphorylates Aprataxin in vitro (Figure 4.8). I therefore propose that the activity of Aprataxin may be modulated by 1) phosphorylation and 2) protein-protein interactions via the FHA domain.

199 While apparently conflicting results on the localization of PARP-1 exist in the literature, where PARP-1 is reported to be either primarily nucleoplasmic (56) or nucleolar (57) based on immunofluorescence, I report that PARP-1 is predominantly nucleoplasmic by immunoblotting (Figure 4.22). These reported differences may be partially due to differences in the protein concentrations between the two compartments. One possibility may be that if the protein density in the nucleolus is much higher than that of the nucleoplasm immunostaining may indicate that PARP-1 is concentrated in the nucleolus, even if it is less abundant there than in the nucleoplasm (in terms of molecules/mg of total protein). This discrepancy may also be compounded by the use of different immunostaining protocols (see Figure 4.39 below, taken from Meder et al., 57). Weak fixation methods and the use of detergent extraction result in ‘washing out’ of nucleoplasmic PARP-1 protein, while nucleolar PARP-1 is retained. Additionally the sub-nuclear distribution of PARP-1 varies between different cell lines (Figure 4.39).

Figure 4.39: PARP-1 immunostaining in HeLa and MEF cells. Taken from Meder et al., (57).

Furthermore, in contrast to Meder et al. (57), some authors report that PARP-1 is almost exclusively nucleoplasmic (see Figure 4.40 below, taken from Vidakovic et al., 56). Thus the cell lines and methods of analysis used may account for these discrepancies. However it is clear using either immunofluorescence or immunoblotting of fractions that PARP-1 is a nuclear protein, consistent with its interaction with Aprataxin and its role as a DNA repair protein.

200

Figure 4.40. PARP-1 immunostaining in Hepa cells. Taken from Vidokovic et al., (56).

To discern the effect of protein interactions on Aprataxin’s function, I examined the DNA-adenylate hydrolase activity in PARP-1 and XRCC1 deficient cell lines. I first demonstrated that XRCC1 deficiency does not affect DNA-adenylate hydrolase activity. Second, I report here that PARP-1 deficiency has a negative effect on DNA adenylate repair capacity. PARP-1 knockout cells are unable to efficiently hydrolyse adenylated DNA, due to destabilization of Aprataxin protein (section 4.3.8). Aprataxin was not detectable in PARP-1 knockout cell extracts indicating that PARP- 1 is critical for stabilization of Aprataxin. Therefore I subsequently examined the dependence of adenylate hydrolase activity on PARP activity. In contrast to the requirement of PARP activity for XRCC1 recruitment, inhibition of PARP activity had no effect on DNA-adenylate hydrolysis. This indicates that although PARP-1 protein is critical for stabilization of Aprataxin, PAR synthesis is not required for stabilization or targeting Aprataxin to its substrate in vitro. On the other hand the interaction between XRCC1 and PARP-1 is dependant on PAR synthesis (46) and this interaction is required for the assembly or stability of XRCC1 protein complexes at sites of DNA damage (see Figure 4.41, taken from El-Khamisy et al., 3). Consequently, deficiency of PARP-1 has a number of genotoxic consequences including inhibition of direct single strand break repair and BER due to a failure of repair complex recruitment (3,46).

201

Figure 4.41: PARP-1 is required for appearance of XRCC1 nuclear foci at sites of oxidative DNA damage. Taken from El-Khamisy et. al (3).

4.4.3 Single strand break repair by AOA1 cell extracts: Aprataxin interacts with a number of proteins involved in the repair of direct single strand breaks, including PARP-1, XRCC1, DNA ligase 3α and DNA polymerase β (1,2). Deficiency of XRCC1 results in single strand break repair deficits, both in terms of cellular sensitivity profile (40,58,59) and in vitro repair capacity (60). The XRCC1 deficient CHO cell line EM9 exhibits hypersensitivity to ethylating and methylating agents, the topoisomerase I inhibitor camptothecin, and BrdU. EM9 also displays elevated basal and induced levels of chromosome aberrations (40,59). Reconstituted DNA repair reactions have revealed roles for XRCC1 multiple DNA repair pathways (60,61). Given that AOA1 cells are hypersensitive to agents which induce single strand breaks, such as hydrogen peroxide (1,9), I examined the in vitro single strand break repair capacity of AOA1 cell extracts. Deficiency of Aprataxin does not result in a gross in vitro single strand break repair defect, indicating that abortive ligations only occur at a small proportion of breaks (section 4.3.5).

In the previous chapter I demonstrated that oxidation (8-oxo-dG) of the 3’ terminus of a single strand break inhibits ligation and causes adenylation of the 5’ strand (section 3.3.4). I therefore examined the potential role of Aprataxin in repair of this type of complex single strand break. Repair of this damaged substrate by cell extracts was less efficient than repair of the non-oxidized control DNA, however an Aprataxin-

202 dependant repair defect was not observed at either oxidized or unmodified single strand breaks (section 4.3.5). Additionally, oxidation of the 3’ terminus did not result in adenylation of the 5’ strand by AOA1 cell extract, which I anticipated based on the ability of 3’8-oxo-dG to inhibit recombinant ligase causing 5’adenylation (section 3.3.4). This could be due to either or both of two possibilities. 1) Endogenous DNA ligase may not bind efficiently to single strand breaks with an oxidized 3’ terminus. Adenylated DNA therefore may not be produced in detectable amounts due to a reduced level of enzyme-AMP-substrate intermediate complex formation. 2) Under the conditions used, the rate of adenylate formation may be slower than the rate of non-specific 3’ to 5’ exonuclease activity (visible as laddering of the ligated products). Any adenylates formed would not be detectable due to substrate degradation. Given that the efficiency of ligation by cell extract is much lower and the non-specific nuclease activity much higher than the previous instance, this result does not conflict with the information presented in section 3.3.4. I maintain that oxidation of the 3’ terminus of a single strand break inhibits ligation and may cause 5’ adenylation, but that this is difficult to given the abundance of nucleases in cell extracts.

4.4.4 Oxidative DNA damage and base excision repair in AOA1 cells: Oxidative stress is a hallmark of many neurodegenerative disorders, including Parkinson’s and Alzheimer’s Diseases (33,62,63). A common biomarker for oxidative stress is 8-oxo-dG, a product of free-radical attack on DNA. Hydrogen peroxide treatment of cells produces 8-oxo-dG (among other lesions). Brain specimens from deceased AOA1 patients display elevated levels of 8-oxo-dG staining compared to controls (34), and AOA1 cells are hypersensitive to hydrogen peroxide (1,9). Combined this evidence suggests that AOA1 cells may have a defect in the repair of 8-oxo-dG. I found that AOA1 fibroblasts have an elevated basal level of 8-oxo-dG staining, supporting the findings of Hirano et al. (34) and providing initial evidence that AOA1 cells have a defect in their oxidative stress response.

8-oxo-dG is repaired using the BER pathway (reviewed in 42,64-66). In the initial phase of 8-oxo-dG repair by BER, 8-oxo-dG is excised to generate an abasic site by the 8-oxo-G DNA glycosylases OGG1 (67-69) or NEIL1 (69), depending on the proximity of the lesion to a single strand break. OGG1 depurinates 8-oxo-dG sites

203 present on intact DNA strands or at least three nucleotides away from a single strand break (69). NEIL1 is able to depurinate 8-oxo-dG closer to a single strand break (two or three nucleotides away) but neither enzyme can repair a site on the 3’ terminus or one nucleotide away from a single strand break (69). APE1 can hydrolyse 8-oxo-dG present at a 3’ terminus (35). NEIL1 and APE1 only repair 8-oxo-dG at or near termini, and OGG1 is the major enzyme responsible for 8-oxo-dG excision (70,71).

OGG1 is a bifunctional glycosylase (meaning that it has dual glycosylase and abasic lyase activities), however it is able to excise 8-oxo-dG faster than it can hydrolyse abasic sites, leading to an accumulation of abasic DNA (36,37). Conversely, OGG1 has a higher affinity for the product of its glycosylase reaction than the substrate (36). The 8-oxo-dG glycosylase activity of OGG1 does not follow Michaelis-Menten kinetics, due to its high affinity for abasic strands and relatively weak abasic lyase activity (36). This means that OGG1 can be ‘sequestered’ by stable binding to its product, abasic DNA. Therefore 8-oxo-dG excision in cells is rate limited by the high affinity (but relatively low Kcat) of OGG1 for the product of its glycosylase activity, abasic sites. Addition of APE1 to OGG1 glycosylase reactions stimulates depurination of 8-oxo-dG DNA by OGG1, suggesting that APE1 facilitates OGG1 turnover by competing with it for abasic sites (36). Additionally the very efficient (Kcat = 21.7x106/s) hydrolysis of abasic strands into abasic single strand breaks by APE1 reduces the concentration of abasic strands, resulting in preferential formation of OGG1-[8-oxo-dG DNA] rather than OGG1- [abasic site] complexes. A summary of this kinetic data is shown in Figure 4.42. I demonstrate here that lack of Aprataxin causes destabilization of APE1. This deficiency of APE1 likely leads to a reduced efficiency of OGG1 enzyme turnover and a subsequent decreased rate of OGG1- dependant 8-oxo-dG depurination in AOA1 cells. This may be responsible for the hypersensitivity of AOA1 cells to hydrogen peroxide and their elevated basal level of 8-oxo-dG.

204

Figure 4.42: OGG1 and APE1 substrates, products and reaction kinetics. Summarized from Hill et al., (36)

In addition to hydrolysis of abasic strands, APE1 possesses several 3’ damage processing activities. These include hydrolysis of 3’ 8-oxo-dG (35), deoxyribose phosphate (72), phosphoglycolate (73) and monophosphate groups (72,74). APE1 appears to be the primary protein responsible for the hydrolysis of 3’dRP(72), and is the only protein capable of hydrolysis of 3’ 8-oxo-dG (35). Given that AOA1 cells display elevated levels of total 8-oxo-dG, it seems likely that they have a higher incidence of 3’ terminal 8-oxo-dG than control cells. In the previous chapter I demonstrated that 3’ 8-oxo-dG inhibits ligation and results in adenylation of the 5’ terminus. I therefore propose AOA1 cells have a higher incidence of abortive ligations than control cells, due to their probable indirect 3’ processing defects.

Aprataxin also has the potential to be involved in BER via its interaction with PARP- 1. In support of Dantzer et al., I demonstrated that PARP-1 deficiency results in a defect in long but not short gap filling (46). Following this I examined the gap filling capacity of AOA1 cells and found that they display a mild gap filling defect. Examination of PARP-1 protein levels in AOA1 and APTX corrected isogenic cell lines revealed that Aprataxin stabilizes PARP-1 protein (section 4.3.8). This explains the intermediate gap filling defect displayed by AOA1 cells (less efficient than control extracts, more efficient than PARP-1 knockout extracts).

4.4.5 Summary: In this chapter I have provided a detailed characterisation of the DNA-adenylate hydrolase activity of endogenous Aprataxin. I have demonstrated that the activity of Aprataxin is elevated in the nucleolus relative to the nucleoplasm and provide evidence for mechanisms which may regulate Aprataxin’s activity between these

205 compartments. Protein-protein interactions are central for all cellular processes including DNA repair. Here I report that the interaction between Aprataxin and PARP-1 is critical for the stability of both proteins. Deficiency of Aprataxin results in a subsequent functional PARP-1 deficiency, as measured by gap filling capacity. I also found that Aprataxin deficiency results in destabilization of the BER protein APE1, which probably accounts for the elevated levels of 8-oxo- dG I observed in AOA1 patient fibroblasts. In summary I propose that Aprataxin deficiency results in at least three discrete, compounding DNA repair defects: 1) the inability to hydrolyse DNA adenylates, 2) a reduced gap filling capacity and 3) a reduced 8-oxo-dG excision capacity. These are summarized in Figure 4.43.

Figure 4.43: Schematic representation of the indirect effects of Aprataxin deficiency.

Given that BER is a mechanism capable of removing non-hydrolysable 5’ DNA damage, AOA1 cells may rely on BER to repair 5’ adenylates which they are unable to excise directly. Thus, not only are AOA1 cells unable to repair adenylates directly, they have a probably have a reduced capacity indirectly repair abortive ligations by long patch BER, leading to a gradual accumulation of adenylates in their genome.

206 4.5 REFERENCES

1. Clements, P.M., Breslin, C., Deeks, E.D., Byrd, P.J., Ju, L., Bieganowski, P., Brenner, C., Moreira, M.C., Taylor, A.M. and Caldecott, K.W. (2004) The ataxia-oculomotor apraxia 1 gene product has a role distinct from ATM and interacts with the DNA strand break repair proteins XRCC1 and XRCC4. DNA Repair (Amst), 3, 1493-1502. 2. Luo, H., Chan, D.W., Yang, T., Rodriguez, M., Chen, B.P., Leng, M., Mu, J.J., Chen, D., Songyang, Z., Wang, Y. et al. (2004) A new XRCC1- containing complex and its role in cellular survival of methyl methanesulfonate treatment. Mol Cell Biol, 24, 8356-8365. 3. El-Khamisy, S.F., Masutani, M., Suzuki, H. and Caldecott, K.W. (2003) A requirement for PARP-1 for the assembly or stability of XRCC1 nuclear foci at sites of oxidative DNA damage. Nucleic Acids Research, 31, 5526-5533. 4. El-Khamisy, S.F., Saifi, G.M., Weinfeld, M., Johansson, F., Helleday, T., Lupski, J.R. and Caldecott, K.W. (2005) Defective DNA single-strand break repair in spinocerebellar ataxia with axonal neuropathy-1. Nature, 434, 108- 113. 5. Krogh, B.O., Llorente, B., Lam, A. and Symington, L.S. (2005) Mutations in Mre11 phosphoesterase motif I that impair Saccharomyces cerevisiae Mre11- Rad50-Xrs2 complex stability in addition to nuclease activity. Genetics, 171, 1561-1570. 6. Voit, R. and Grummt, I. (2001) Phosphorylation of UBF at serine 388 is required for interaction with RNA polymerase I and activation of rDNA transcription. Proc Natl Acad Sci U S A, 98, 13631-13636. 7. Korgaonkar, C., Hagen, J., Tompkins, V., Frazier, A.A., Allamargot, C., Quelle, F.W. and Quelle, D.E. (2005) Nucleophosmin (B23) targets ARF to nucleoli and inhibits its function. Mol Cell Biol, 25, 1258-1271. 8. Becherel, O.J., Gueven, N., Birrell, G.W., Schreiber, V., Suraweera, A., Jakob, B., Taucher-Scholz, G. and Lavin, M.F. (2006) Nucleolar localization of aprataxin is dependent on interaction with nucleolin and on active ribosomal DNA transcription. Hum Mol Genet, 15, 2239-2249. 9. Gueven, N., Becherel, O.J., Kijas, A.W., Chen, P., Howe, O., Rudolph, J.H., Gatti, R., Date, H., Onodera, O., Taucher-Scholz, G. et al. (2004) Aprataxin, a

207 novel protein that protects against genotoxic stress. Hum Mol Genet, 13, 1081- 1093. 10. Grozinger, C.M. and Schreiber, S.L. (2000) Regulation of histone deacetylase 4 and 5 and transcriptional activity by 14-3-3-dependent cellular localization. Proc Natl Acad Sci U S A, 97, 7835-7840. 11. Ariza, R.R., Keyse, S.M., Moggs, J.G. and Wood, R.D. (1996) Reversible protein phosphorylation modulates nucleotide excision repair of damaged DNA by human cell extracts. Nucleic Acids Res, 24, 433-440. 12. Hu, J., Imam, S.Z., Hashiguchi, K., de Souza-Pinto, N.C. and Bohr, V.A. (2005) Phosphorylation of human oxoguanine DNA glycosylase (alpha- OGG1) modulates its function. Nucleic Acids Res, 33, 3271-3282. 13. Date, H., Igarashi, S., Sano, Y., Takahashi, T., Takahashi, T., Takano, H., Tsuji, S., Nishizawa, M. and Onodera, O. (2004) The FHA domain of aprataxin interacts with the C-terminal region of XRCC1. Biochem Biophys Res Commun, 325, 1279-1285. 14. Rass, U., Ahel, I. and West, S.C. (2007) Actions of aprataxin in multiple DNA repair pathways. J Biol Chem, 282, 9469-9474. 15. Lavin, M.F., Scott, S.P., Kozlov, S. and Gueven, N. (2004) Analyzing the regulation and function of ATM. Methods Mol Biol, 281, 163-178. 16. Suraweera, A., Becherel, O.J., Chen, P., Rundle, N., Woods, R., Nakamura, J., Gatei, M., Criscuolo, C., Filla, A., Chessa, L. et al. (2007) Senataxin, defective in ataxia oculomotor apraxia type 2, is involved in the defense against oxidative DNA damage. J Cell Biol, 177, 969-979. 17. Scherl, A., Coute, Y., Deon, C., Calle, A., Kindbeiter, K., Sanchez, J.C., Greco, A., Hochstrasser, D. and Diaz, J.J. (2002) Functional proteomic analysis of human nucleolus. Mol Biol Cell, 13, 4100-4109. 18. Seidle, H.F., Bieganowski, P. and Brenner, C. (2005) Disease-associated mutations inactivate AMP-lysine hydrolase activity of aprataxin. J Biol Chem. 19. Hirano, M., Asai, H., Kiriyama, T., Furiya, Y., Iwamoto, T., Nishiwaki, T., Yamamoto, A., Mori, T. and Ueno, S. (2007) Short half-lives of ataxia- associated aprataxin proteins in neuronal cells. Neurosci Lett, 419, 184-187. 20. Ahel, I., Rass, U., El-Khamisy, S.F., Katyal, S., Clements, P.M., McKinnon, P.J., Caldecott, K.W. and West, S.C. (2006) The neurodegenerative disease

208 protein aprataxin resolves abortive DNA ligation intermediates. Nature. 433, 713-716. 21. Hoege, C., Pfander, B., Moldovan, G.L., Pyrowolakis, G. and Jentsch, S. (2002) RAD6-dependent DNA repair is linked to modification of PCNA by ubiquitin and SUMO. Nature, 419, 135-141. 22. Thiriet, C. and Hayes, J.J. (2005) Chromatin in need of a fix: phosphorylation of H2AX connects chromatin to DNA repair. Mol Cell, 18, 617-622. 23. Sun, Y., Jiang, X., Chen, S., Fernandes, N. and Price, B.D. (2005) A role for the Tip60 histone acetyltransferase in the acetylation and activation of ATM. Proc Natl Acad Sci U S A, 102, 13182-13187. 24. Ame, J.C., Spenlehauer, C. and de Murcia, G. (2004) The PARP superfamily. Bioessays, 26, 882-893. 25. Siliciano, J.D., Canman, C.E., Taya, Y., Sakaguchi, K., Appella, E. and Kastan, M.B. (1997) DNA damage induces phosphorylation of the amino terminus of p53. Genes Dev, 11, 3471-3481. 26. Blom, N., Sicheritz-Ponten, T., Gupta, R., Gammeltoft, S. and Brunak, S. (2004) Prediction of post-translational glycosylation and phosphorylation of proteins from the amino acid sequence. Proteomics, 4, 1633-1649. 27. Kijas, A.W., Harris, J.L., Harris, J.M. and Lavin, M.F. (2006) Aprataxin forms a discrete branch in the HIT (histidine triad) superfamily of proteins with both DNA/RNA binding and nucleotide hydrolase activities. J Biol Chem, 281, 13939-13948. 28. Hirano, M., Furiya, Y., Kariya, S., Nishiwaki, T. and Ueno, S. (2004) Loss of function mechanism in aprataxin-related early-onset ataxia. Biochem Biophys Res Commun, 322, 380-386. 29. Lyras, L., Cairns, N.J., Jenner, A., Jenner, P. and Halliwell, B. (1997) An assessment of oxidative damage to proteins, lipids, and DNA in brain from patients with Alzheimer's disease. Journal of Neurochem, 68, 2061-2069. 30. Lyras, L., Perry, R.H., Perry, E.K., Ince, P.G., Jenner, A., Jenner, P. and Halliwell, B. (1998) Oxidative damage to proteins, lipids, and DNA in cortical brain regions from patients with dementia with Lewy bodies. Journal of Neurochem, 71, 302-312.

209 31. Lovell, M.A. and Markesbery, W.R. (2007) Oxidative DNA damage in mild cognitive impairment and late-stage Alzheimer's disease. Nucleic Acids Res, 35, 7497-7504. 32. Chen, P., Peng, C., Luff, J., Spring, K., Watters, D., Bottle, S., Furuya, S. and Lavin, M.F. (2003) Oxidative stress is responsible for deficient survival and dendritogenesis in purkinje neurons from ataxia-telangiectasia mutated mutant mice. J Neurosci, 23, 11453-11460. 33. Reichenbach, J., Schubert, R., Schindler, D., Muller, K., Bohles, H. and Zielen, S. (2002) Elevated oxidative stress in patients with ataxia telangiectasia. Antioxidants & Redox Signaling, 4, 465-469. 34. Hirano, M., Yamamoto, A., Mori, T., Lan, L., Iwamoto, T.A., Aoki, M., Shimada, K., Furiya, Y., Kariya, S., Asai, H. et al. (2007) DNA single-strand break repair is impaired in aprataxin-related ataxia. Ann Neurol, 61, 162-174. 35. Parsons, J.L., Dianova, II and Dianov, G.L. (2005) APE1-dependent repair of DNA single-strand breaks containing 3'-end 8-oxoguanine. Nucleic Acids Res, 33, 2204-2209. 36. Hill, J.W., Hazra, T.K., Izumi, T. and Mitra, S. (2001) Stimulation of human 8-oxoguanine-DNA glycosylase by AP-endonuclease: potential coordination of the initial steps in base excision repair. Nucleic Acids Res, 29, 430-438. 37. Vidal, A.E., Hickson, I.D., Boiteux, S. and Radicella, J.P. (2001) Mechanism of stimulation of the DNA glycosylase activity of hOGG1 by the major human AP endonuclease: bypass of the AP lyase activity step. Nucleic Acids Res, 29, 1285-1292. 38. Shacter, E. (2000) Quantification and significance of protein oxidation in biological samples. Drug Metab Rev, 32, 307-326. 39. Weber, J.D., Taylor, L.J., Roussel, M.F., Sherr, C.J. and Bar-Sagi, D. (1999) Nucleolar Arf sequesters Mdm2 and activates p53. Nat Cell Biol, 1, 20-26. 40. Thompson, L.H., Brookman, K.W., Dillehay, L.E., Carrano, A.V., Mazrimas, J.A., Mooney, C.L. and Minkler, J.L. (1982) A CHO-cell strain having hypersensitivity to mutagens, a defect in DNA strand-break repair, and an extraordinary baseline frequency of sister-chromatid exchange. Mutat Res, 95, 427-440.

210 41. Shen, M.R., Zdzienicka, M.Z., Mohrenweiser, H., Thompson, L.H. and Thelen, M.P. (1998) Mutations in hamster single-strand break repair gene XRCC1 causing defective DNA repair. Nucleic Acids Res, 26, 1032-1037. 42. Caldecott, K.W. (2003) Protein-protein interactions during mammalian DNA single-strand break repair. Biochem Soc Trans, 31, 247-251. 43. Ishizuka, S., Martin, K., Booth, C., Potten, C.S., de Murcia, G., Burkle, A. and Kirkwood, T.B. (2003) Poly(ADP-ribose) polymerase-1 is a survival factor for radiation-exposed intestinal epithelial stem cells in vivo. Nucleic Acids Res, 31, 6198-6205. 44. Masutani, M., Nozaki, T., Nakamoto, K., Nakagama, H., Suzuki, H., Kusuoka, O., Tsutsumi, M. and Sugimura, T. (2000) The response of Parp knockout mice against DNA damaging agents. Mutat Res, 462, 159-166. 45. Masutani, M., Nozaki, T., Nishiyama, E., Shimokawa, T., Tachi, Y., Suzuki, H., Nakagama, H., Wakabayashi, K. and Sugimura, T. (1999) Function of poly(ADP-ribose) polymerase in response to DNA damage: gene-disruption study in mice. Mol Cell Biochem, 193, 149-152. 46. Dantzer, F., de La Rubia, G., Menissier-De Murcia, J., Hostomsky, Z., de Murcia, G. and Schreiber, V. (2000) Base excision repair is impaired in mammalian cells lacking Poly(ADP-ribose) polymerase-1. Biochemistry, 39, 7559-7569. 47. Brooks, P.J. (2007) The case for 8,5'-cyclopurine-2'-deoxynucleosides as endogenous DNA lesions that cause neurodegeneration in xeroderma pigmentosum. Neuroscience, 145, 1407-1417. 48. Brooks, P.J., Wise, D.S., Berry, D.A., Kosmoski, J.V., Smerdon, M.J., Somers, R.L., Mackie, H., Spoonde, A.Y., Ackerman, E.J., Coleman, K. et al. (2000) The oxidative DNA lesion 8,5'-(S)-cyclo-2'-deoxyadenosine is repaired by the nucleotide excision repair pathway and blocks gene expression in mammalian cells. J Biol Chem, 275, 22355-22362. 49. Le Ber, I., Moreira, M.C., Rivaud-Pechoux, S., Chamayou, C., Ochsner, F., Kuntzer, T., Tardieu, M., Said, G., Habert, M.O., Demarquay, G. et al. (2003) Cerebellar ataxia with oculomotor apraxia type 1: clinical and genetic studies. Brain, 126, 2761-2772.

211 50. Ward, I.M., Minn, K. and Chen, J. (2004) UV-induced ataxia-telangiectasia- mutated and Rad3-related (ATR) activation requires replication stress. J Biol Chem, 279, 9677-9680. 51. Lee, J.H. and Paull, T.T. (2005) ATM activation by DNA double-strand breaks through the Mre11-Rad50-Nbs1 complex. Science, 308, 551-554. 52. Michel, J.J. and Scott, J.D. (2002) AKAP mediated signal transduction. Annu Rev Pharmacol Toxicol, 42, 235-257. 53. Liberi, G., Cotta-Ramusino, C., Lopes, M., Sogo, J., Conti, C., Bensimon, A. and Foiani, M. (2006) Methods to study replication fork collapse in budding yeast. Methods Enzymol, 409, 442-462. 54. Zhou, W. and Doetsch, P.W. (1994) Transcription bypass or blockage at single-strand breaks on the DNA template strand: effect of different 3' and 5' flanking groups on the T7 RNA polymerase elongation complex. Biochemistry, 33, 14926-14934. 55. Sanchez, G. and Mamet-Bratley, M.D. (1994) Transcription by T7 RNA polymerase of DNA containing abasic sites. Environmental and Molecular Mutagenesis, 23, 32-36. 56. Vidakovic, M., Koester, M., Goetze, S., Winkelmann, S., Klar, M., Poznanovic, G. and Bode, J. (2005) Co-localization of PARP-1 and lamin B in the nuclear architecture: a halo-fluorescence- and confocal-microscopy study. J Cell Biochem, 96, 555-568. 57. Meder, V.S., Boeglin, M., de Murcia, G. and Schreiber, V. (2005) PARP-1 and PARP-2 interact with nucleophosmin/B23 and accumulate in transcriptionally active nucleoli. J Cell Sci, 118, 211-222. 58. Barrows, L.R., Holden, J.A., Anderson, M. and D'Arpa, P. (1998) The CHO XRCC1 mutant, EM9, deficient in DNA ligase III activity, exhibits hypersensitivity to camptothecin independent of DNA replication. Mutat Res, 408, 103-110. 59. Ray, J.H., Louie, E. and German, J. (1985) Hybridization with human cells corrects the elevated SCE frequency and BrdU hypersensitivity of hamster cell line EM9. Cytogenet Cell Genet, 39, 154-157. 60. Kubota, Y., Nash, R.A., Klungland, A., Schar, P., Barnes, D.E. and Lindahl, T. (1996) Reconstitution of DNA base excision-repair with purified human

212 proteins: interaction between DNA polymerase beta and the XRCC1 protein. The EMBO Journal, 15, 6662-6670. 61. Audebert, S., Calsou. (2004) Involvement of Poly(ADP-ribose) Polymerase-1 and XRCC1/DNA Ligase III in an Alternative Route for DNA Double-strand Breaks Rejoining. J Biol Chem, 279, 55117-55126. 62. Zhang, J., Perry, G., Smith, M.A., Robertson, D., Olson, S.J., Graham, D.G. and Montine, T.J. (1999) Parkinson's disease is associated with oxidative damage to cytoplasmic DNA and RNA in substantia nigra neurons. Am J Pathol, 154, 1423-1429. 63. Markesbery, W.R. (1997) Oxidative stress hypothesis in Alzheimer's disease. Free Radical Biology & Medicine, 23, 134-147. 64. Caldecott, K.W. (2001) Mammalian DNA single-strand break repair: an X- ra(y)ted affair. Bioessays, 23, 447-455. 65. Dianov, G.L. and Parsons, J.L. (2007) Co-ordination of DNA single strand break repair. DNA Repair (Amst), 6, 454-460. 66. Lindahl, T., Karran, P. and Wood, R.D. (1997) DNA excision repair pathways. Current Opinion in Genetics & Development, 7, 158-169. 67. Dherin, C., Dizdaroglu, M., Doerflinger, H., Boiteux, S. and Radicella, J.P. (2000) Repair of oxidative DNA damage in Drosophila melanogaster: identification and characterization of dOgg1, a second DNA glycosylase activity for 8-hydroxyguanine and formamidopyrimidines. Nucleic Acids Res, 28, 4583-4592. 68. David-Cordonnier, M.H., Boiteux, S. and O'Neill, P. (2001) Excision of 8- oxoguanine within clustered damage by the yeast OGG1 protein. Nucleic Acids Res, 29, 1107-1113. 69. Parsons, J.L., Zharkov, D.O. and Dianov, G.L. (2005) NEIL1 excises 3' end proximal oxidative DNA lesions resistant to cleavage by NTH1 and OGG1. Nucleic Acids Res, 33, 4849-4856. 70. Arai, T., Kelly, V.P., Minowa, O., Noda, T. and Nishimura, S. (2002) High accumulation of oxidative DNA damage, 8-hydroxyguanine, in Mmh/Ogg1 deficient mice by chronic oxidative stress. Carcinogenesis, 23, 2005-2010. 71. Le Page, F., Klungland, A., Barnes, D.E., Sarasin, A. and Boiteux, S. (2000) Transcription coupled repair of 8-oxoguanine in murine cells: the ogg1 protein

213 is required for repair in nontranscribed sequences but not in transcribed sequences. Proc Natl Acad Sci U S A, 97, 8397-8402. 72. Demple, B. and Harrison, L. (1994) Repair of oxidative damage to DNA: enzymology and biology. Annu Rev Biochem, 63, 915-948. 73. Parsons, J.L., Dianova, II and Dianov, G.L. (2004) APE1 is the major 3'- phosphoglycolate activity in human cell extracts. Nucleic Acids Res, 32, 3531- 3536. 74. Chen, D.S., Herman, T. and Demple, B. (1991) Two distinct human DNA diesterases that hydrolyze 3'-blocking deoxyribose fragments from oxidized DNA. Nucleic Acids Res, 19, 5907-5914.

214

CHAPTER 5

A role for Aprataxin in RNA biogenesis

215 216 5.1 INTRODUCTION

In chapter 4 I reported that Aprataxin is present in the nucleolus, consistent with previous reports (1,2). Additionally I found that the specific activity of Aprataxin in the nucleolus is higher than in the nucleoplasm. This indicated that adenylate formation in the nucleolus may be very frequent or very toxic (necessitating a high level of Aprataxin activity), or that Aprataxin has an alternative role in the nucleolus.

5.1.1 Structure and function of the nucleolus:

Nucleoli are non-membrane bound structures present in eukaryotic nuclei. Nucleoli are highly transcriptionally active and are composed of many discrete transcription factories. The primary function of the nucleolus is synthesis and processing of rRNA (reviewed in 3). Each transcription factory is composed of three discrete domains, based on their morphology and function in rRNA synthesis and processing (reviewed in 4). These structures can be visualized by incorporation of BrUTP into nascent transcripts followed by immunogold labelling and electron microscopy (Figure 5.1, taken form (5). The Fibrillar Centre (FC) contains rRNA transcription and processing factors and is visible as a region of low electron density. The FC is surrounded by an electron dense region called the Dense Fibrillar Component (DFC). rRNA transcription is initiated at the border of the DFC and FC, with nascent strands synthesized into the DFC (4). Nascent rRNA transcripts then move into the GC where they are processed into pre-ribosomes (4).

217

Figure 5.5.1: Subdomains of the nucleolus. Individual sub-nucleolar components can be visualized by incorporation of BrUTP into nascent rRNA followed by immunogold staining and electron microscopy. Shown is a single BrUTP-labelled nucleolar transcription factory with individual domains identified (Fibrillar Centre-FC, Dense Fibrillar Component- DFC, and the Granular Component- GC). HeLa cells may have up to 30 such factories per nucleolus. Taken from (5).

Additional, non rRNA related roles for the nucleolus have been identified. In Saccharomyces cerevisiae immature tRNAs are present predominantly in the nucleolus, where they may be processed in parallel with rRNAs (6). The nucleolus is also involved in DNA damage signalling by controlling the shuttling of the ubiquitin ligase MDM2 which in turn regulates p53 stability (7).

5.1.2 Nucleolar transcription:

Nucleoli are clearly visible by DAPI staining as regions of low DNA content. Despite this, most of a cells transcriptional activity occurs in the nucleolus where transcription factories are arranged around rDNA tandem repeats. Approximately 180 copies of the 47 kb rDNA repeat are present on each of the five acrocentric chromosomes(4). Of these approximately 120 genes are active; these are arranged three to four genes per transcription factory (3,4). Transcription of rRNA is carried out exclusively by the RNA polymerase 1 holoenzyme (3). RNA polymerases themselves have no intrinsic ability to bind specifically to promoter sequences and instead rely on the formation of pre-initiation complexes by their respective transcription factors (8). In the case of RNA polymerases II and III, pre-initiation complexes are formed by interaction of

218 their respective transcription factors to TATA boxes upstream of the initiation site (8). RNA polymerase 1 does not require a TATA box for initiation of transcription; the rDNA core promoter contains an Upstream Control Sequence (UCS) instead (9). The UCS recruits the TATA binding complex selectivity factor 1 (SL1, also known as TIF-1B). This is a multiprotein complex consisting of TATA Binding Protein (TBP), TATA Binding Protein Associated Factor 95 or 110 (TAF 95, TAF 110), TAF 68 or 63 and TAF 48 as well as a HMG-box protein called Upstream Binding Factor (UBF) (9). Together these proteins bind the UCS forming a RNA polymerase I pre-initiation complex. The C-terminal region of UBF then recruits RNA polymerase I to the ‘primed’ promoter site (10). RNA polymerase 1 subsequently leaves the pre-initiation complex and scans the strand for its transcription initiation site, leaving the UBF and SL1 bound to the UCS ready to prime another transcription event (11). Termination of RNA polymerase 1 transcription is facilitated by Transcription Termination Factor 1 (TTF1) which binds to termination elements in the 3’-external transcribed spacer of rDNA repeats (12,13).

Transcription of rRNA is regulated by cell cycle. RNA polymerase I is active between

G1 and G2 phases (as well as G0) and is inactivated as cells enter mitosis and progress towards prophase. This inhibition is controlled by cdc2-cyclin B mediated phosphorylation of SL1 (14,15). Phosphorylation of SL1 inhibits its interaction with UBF, preventing formation of pre-initiation complexes (14). This inhibition of rRNA synthesis results in disintegration of the nucleolus and association of partially processed transcripts and the nucleolar transcription machinery into Nucleolar Organizing Regions (NORs, reference 16). After mitosis the cdc2-cyclin B mediated inhibition of the SL1/UBF interaction is relieved, allowing formation of pre-initiation complexes and RNA polymerase I activation in the NORs of daughter cells, which form new nucleoli (14,15).

5.1.3 Interaction between Aprataxin and nucleolin:

This laboratory has demonstrated an interaction between Aprataxin and the nucleolar rRNA processing factor nucleolin (1). I therefore predicted that Aprataxin may

219 interact with other proteins involved in RNA transcription and processing. In this chapter I have analysed the physical and functional interactions between Aprataxin and various proteins involved in RNA biogenesis.

220

5.2 MATERIALS AND METHODS

5.2.1 Recruitment of proteins to dsDNA:

This experiment was performed in a similar manner to the analysis of DNA binding of Aprataxin from extract in the previous chapter (section 4.2.3). Briefly, A biotinylated duplex (5’ biotin-TGTAGTCACTATCGGAATGAGGGCGACACGGATATG, annealed to the unmodified complementary strand) was attached to streptavidin magnetic beads (Dynal) as per manufacturers instructions. Beads were mock treated in parallel. DNA coated or uncoated beads (30 µg) were incubated with 100 µg of nuclear extract from the indicated cell lines (generated in the same manner as for all DNA repair assays, section 3.2.3.2) in Ligase Buffer (total volume 100 µL) for 10 minutes on ice. Reactions were gently mixed every several minutes. DNA-binding proteins were enriched by precipitation of the beads in a magnetic rack (Invitrogen) and washing in Ligase Buffer. DNA binding proteins were eluted in 5x SDS-PAGE loading buffer and subjected to protein electrophoresis and colloidal Coomassie blue staining.

Bands which were present in C3ABR but absent in L938 (indicating Aprataxin- dependant recruitment) were excised and stored in 1% acetic acid at 4°C until further processing. Bands were trypsinized and identified by mass spectrometry by Dr Geoffrey Birrel according to established protocols (17,18). Briefly, spectrometric analysis was carried out using a Microflex MALDI-TOF-PSD (Bruker Daltonics, Bremen, Germany), operated in a positive ion reflectron mode. Mass spectrometry (MS) data were acquired using 350 shots of a nitrogen laser at 355 nm with a 20 Hz repetition rate and varying intensity. MS data were calibrated via close external calibration using the peptide standards (New England Biolabs) containing Angiotensin I (MH+ 1296.69), Neurotensin (MH+ 1672.92), ACTH (1-17 clip, MH+ 2093.09), ACTH (18-39 clip, MH+ 2465.20), ACTH (7- 38 clip, MH+ 3657.93). To analyse the MS data, a Mascot search engine (Version 1.9) and the NCBI database were used. Mass tolerance was set at 150 ppm. The search took into account

221 carbomidomethylated cysteine and oxidised methionine and no other post- translational modifications were included (17,18). Thanks to Dr Amila Suraweera for this method.

5.2.2 GST pulldowns:

Aprataxin-GST fusion proteins have been previously described (1,2). Briefly, for pull- down assays, the GST, GST-FHA, GST-NL, GST-HIT and GST-ZF fusion proteins were bound to glutathione Sepharose beads (Amersham). These proteins were generated from pGEX 5.1 constructs and a schematic is shown in Figure 4.6. Total cell extracts were prepared by lysing 1 x 107 control cells (C3ABR) in 1 ml of Lysis Buffer B. 1 mg of cleared lysate was incubated with the protein coated GST beads in Lysis Buffer B in a final volume of 1 mL (using 50 µL of beads). Pull-downs were performed in batch mode for 2 hours at 4oC on a rotating wheel. Where indicated Benzonase (Roche) was included at a concentration of 50 units/mg of protein extract. Alternatively, to examine the effect of protein phosphorylation by CK2 on protein interactions, confluent HeLa cells in 6-well dishes were incubated with the CK2 inhibitor 4,5,6,7-tetrabromobenzotriazole (TBB) for 8 hours (concentrations are indicated) in DMEM with 12% FCS. After treatment cells were washed in PBS and each well was scraped in 1 mL of Lysis Buffer B. After clearing, 900 µL of extract from each well was incubated with 50 µL of protein coated GST beads. For all experiments beads were washed three times with lysis buffer B and resuspended in 20 l of 5x SDS-PAGE loading buffer. Proteins were resolved by 8-15% gradient protein electrophoresis and transferred onto nitrocellulose. Immunoblotting for TAF95, fibrillarin and TTF1 was kindly performed by Dr Valerie Schreiber (Département Intégrité du Génome, École Supérieure de Biotechnologie de Strasbourg, Strasbourg).

5.2.3 UBF immunostaining:

Subconfluent adherent cells were treated as described in figure legends and then fixed in PBS containing 10% formalin for 10 minutes at room temperature. They were then dehydrated in 100% methanol at room temperature for 10 minutes and rehydrated in

222 PBS. Cells were permeabilized and blocked in Blocking Solution (PBS containing 5% NBS and 0.05% Triton X-100) for at 1 hour at room temperature. Cells were incubated with mouse α-UBF and rabbit α-nucleophosmin antibodies (1/100 for UBF, 1/500 for nucleophosmin, in Blocking Solution for 1 hour at 37°C). Staining was detected using AlexaFluor 488-α-mouse IgG and AlexaFluor 594-α-rabbit IgG. Images were captured using a digital camera (Carl Zeiss, Axiocam MRm) attached to a fluorescent microscope Axioskop2 mot plus (Carl Zeiss) using Plan Apochromat 1.4 oil DIC (63x magnification). Zeiss software (Axiovision LE 4.3) was used to capture the individual images which were assembled using Adobe Photoshop 7.0.

UBF staining was quantified by immunofluorescence in corrected and defective AOA1 fibroblast cell lines. Staining intensity was determined by quantification of the staining of individual nuclei using the public domain software ImageJ. The freehand draw tool was used to circle each nucleus, and the measure tool used to determine the mean, minimum, and maximum intensities as well as the size of the designated area. The staining intensity of each nucleus was determined by multiplication of the area and mean intensity values. At least 40 cells from each slide were quantified and all are plotted, with the mean intensity (standardized to untreated FD105 M21) for each slide indicated by a bar. Students t-test was used to examine the level of significance of intensity differences between the slides.

5.2.4 UBF immunoblotting:

Sub-confluent FD105 M20 and M21 cells were washed and scraped in PBS and lysed in Lysis Buffer B. Equal amounts of protein were subjected to denaturing protein electrophoresis and Western blotting as indicated.

223 5.2.5 Nucleotide analogue incorporation protocols:

5.2.5.1 Passive Incorporation

Sub-confluent cells on coverslips were incubated with 2 mM FUrd in growth media (DMEM with 12% FCS) for 10 minutes at 37°C. Coverslips were washed in PBS and immediately fixed in the indicated solutions for 10 minutes. Coverslips were then washed in PBS and blocked and permeabilized for one hour at room temperature in PBS containing 5% bovine serum and 0.05% Triton X-100. Coverslips were stained as indicated before mounting in Moviol. Images were captured on a Zeiss AxioSkop as described previously.

5.2.5.2 Hypotonicity

Sub-confluent cells on coverslips were permeabilized by incubation in the hypotonic buffer KHB (30 mM KCl, 10 mM HEPES pH 7.4) for 5 minutes at 37°C. Cells were then incubated with KHB containing either BrUTP (100 µM) or FUrd (2 mM) for 15 minutes at 37°C. The nucleotide mixture was then removed and replaced with DMEM (12% FCS) for 20 minutes. Cells were subsequently fixed in PBS containing 2% paraformaldehyde (PFA) for 10 minutes at room temperature. Cells were permeabilized and blocked in PBS containing 5% bovine serum and 0.05% Triton X- 100. Immunostaining was then performed as indicated before mounting in Moviol. Images were captured on a Zeiss AxioSkop as described previously.

5.2.5.3 Lipofectamine 2000

FUrd was transfected into sub-confluent cells on coverslips using Lipofectamine 2000. The volumes indicated are for each well of a standard 6-well plate. 10 µL of transfection reagent was added to 250 µL of OptiMEM, mixed gently and incubated at room temperature for 5 minutes. This was mixed with 250 µL of OptiMEM containing 20 mM FUrd, mixed gently and incubated at room temperature for further 20 minutes. During this incubation cells on coverslips were washed once in PBS and twice in OptiMEM. The cells were incubated with the transfection mixture for 15 minutes at 37°C. The transfection mixture was removed and coverslips were washed

224 three times in PBS before being transferred to wells containing DMEM 12% FCS and incubated for the indicated times at 37°C. Cells were then fixed in 2% paraformaldehyde in PBS for 10 minutes at room temperature and washed three times in PBS. Cells were permiablized and blocked in PBS containing 5% bovine serum and 0.05% Triton X-100 for at least 60 minutes. Immunostaining was performed as indicated before mounting in Moviol. Images were captured on a Zeiss AxioSkop as described previously.

5.2.5.4 FuGene6

FUrd was transfected into sub-confluent cells on coverslips using FuGene6. The volumes indicated refer to a single well of a standard 6-well plate. 6 µL of FuGene transfection reagent was added to 250 µL of OptiMEM, mixed gently and incubated at room temperature for 5 minutes. 250 µL of OptiMEM containing 10 mM FUrd was added to this, mixed gently and incubated for a further 20 minutes at room temperature. During this incubation cells on coverslips were washed once in PBS, twice in OptiMEM and gently drained on a tissue. This transfection solution was immediately added to the drained coverslips. Coverslips were incubated with this transfection mixture for 15 minutes at 37°C. The mixture was subsequently removed and the coverslips were washed in PBS before incubation in DMEM containing 12% FCS for the indicated times. Coverslips were then washed in PBS and fixed in PBS containing 10% formalin for 10 minutes at room temperature. Coverslips were washed in PBS before dehydration in 100% methanol at room temperature for 10 minutes. Coverslips were subsequently rehydrated in PBS for 10 minutes prior to blocking in PBS containing 3% bovine serum and 2% BSA for 60 minutes at room temperature. Immunostaining was subsequently performed as indicated prior to mounting the coverslips in Moviol. Images were captured on a Zeiss AxioSkop or a DeltaVision microscope as indicated.

5.2.5.5 Quantification of FUrd incorporation:

FUrd incorporation was quantified in the same manner as UBF staining was quantified previously.

225 5.3 RESULTS

Previous chapters have focused on the biochemical characterisation of recombinant Aprataxin or examination of the DNA repair capacities of Aprataxin deficient cells. This laboratory has identified interactions between Aprataxin and several proteins which are not directly implicated in DNA repair, including the rRNA processing factor nucleolin (1). In the previous chapter and in Gueven et. al (2). I characterised the sub-cellular distribution of Aprataxin and found that Aprataxin is present in both the nucleoplasm and the nucleolus. Furthermore I found that Aprataxin in the nucleolus has a higher specific activity than in the nucleoplasm. To explain this I proposed two non-mutually exclusive possibilities. DNA in the nucleolus may be more prone to DNA damage and thus a cell may require an elevated level of Aprataxin activity to main rDNA integrity. Secondly, Aprataxin may have an alternative function in the this organelle. Given that the primary function of the nucleolus is synthesis of rRNA and processing rRNA into pre- ribosomes, I hypothesized that Aprataxin may have function in the nucleolus as a transcription/rRNA processing factor. Here I have used several approaches to examine this hypothesis, including characterisation of protein/DNA and protein- protein interactions, examination of the effect of Aprataxin on the level and localization of a transcription factor, and analysis of the transcriptional activity of Aprataxin deficient cells.

5.3.1 Aprataxin interacts with RNA transcription and processing factors:

Studies by this laboratory and others have found that Aprataxin directly interacts with DNA (19,20) and RNA (19) as well as with DNA repair proteins (1,21,22). I have demonstrated that Aprataxin deficiency results in destabilization of two additional DNA repair proteins, PARP-1 and APE1. An additional consequence of Aprataxin deficiency may be defective recruitment of proteins to DNA. This is exemplified by the inability of PARP-1 knockout cells to form XRCC1 foci at hydrogen peroxide- induced lesions (23). Recruitment of proteins to DNA can be examined using two basic approaches. 1- recruitment of proteins of interest can be examined by

226 immunostaining or fluorescent microscopy in cells after appropriate stimuli, as used by El-Khamisy et al. (23). This approach is suitable for examining the recruitment of known proteins of interest to lesions of undefined structure. 2- recruitment of any protein to a specific DNA structure can be examined in vitro by incubation of cell extracts with the biotinylated DNA structure of interest. The DNA and its interacting proteins can be precipitated with avidin or streptavidin beads and the interacting proteins examined by SDS-PAGE and subsequently identified by mass spectrometry.

To explore the role of Aprataxin in recruitment of proteins to DNA I examined differences between control and AOA1 DNA binding proteins in vitro. Streptavidin magnetic beads were coated with a biotinylated 36 nucleotide DNA duplex as outlined in section 5.2.1. Control and AOA1 nuclear extracts were incubated with the DNA coated beads. Interacting proteins were enriched by precipitation and washing of the beads. The recruited proteins were subsequently eluted with SDS-PAGE loading buffer and subjected to electrophoresis and colloidal coomassie staining (Figure 5.2). The predominant difference between the binding profiles of C3ABR and L938 extracts was the binding of an approximately 115kDa protein to DNA in C3ABR but not L938. The presence of this band amongst the DNA binding proteins of control but not AOA1 extract indicated that recruitment of this protein to DNA is Aprataxin dependant. To identify this Aprataxin-dependant DNA binding protein, the band was excised, trypsinized and subjected to MALDI-TOF mass spectrometry (thanks to Dr Geoff Birrell, Queensland Institute for Medical Research, Brisbane, Australia). This identified the DNA binding protein as Splicing Factor, Proline Glutamine rich (SFPQ). Significant mass spectrometry hits were not obtained for other bands present in C3ABR but not L938, indicating Aprataxin-dependant recruitment.

SFPQ, also known as PSF for Polypyrimidine Tract Binding-protein (PTB)-associated Splicing Factor, binds polypyrimidine tracts in mammalian mRNA introns independently of its interacting partner PTB (24). pre-mRNA splicing takes place in a multi-complex structure called a spliceosome. These complexes assemble on the pre- mRNA in a stepwise manner based on their function in the overall splicing reaction

227 and form four functional intermediates in the order E→ A →B →C (reviewed in 25,26). Complexes E and A are designated the pre-spliceosome, and B and C the spliceosome. SFPQ is present in the B complex and enriched in the C complex, whose function is excision of a lariat intron from an exon terminus so that it may be joined to the preceding exon (27). SFPQ is essential for spliceosome formation and SFPQ- depleted HeLa cell extracts are deficient in their ability to splice the α-tropomyosin pre-mRNA using an in vitro splicing assay (24). SFPQ also interacts with the 3’ pre- mRNA processing exonuclease XRN2 and is required for its recruitment to nascent pre-mRNA transcripts, where it facilitates coupling of 3’ pre-mRNA processing with degradation of excised RNA by XRN2 (28). More recently SFPQ was found to interact with the C-terminal domain of RNA polymerase II (29). The absence of SFPQ recruitment in AOA1 cell extracts indicates that this protein is either destabilized in the absence of Aprataxin or that its recruitment is Aprataxin- dependant. In either case it implies that Aprataxin may affect RNA processing. Either situation implies a role for Aprataxin in splicing and that AOA1 cells may have splicing defects.

228

SFPQ

Figure 5.2: Aprataxin mediated recruitment of SFPQ to DNA. Control (C3ABR) and AOA1 (L938) cell extracts were incubated with magnetic beads coated in a 36 nucleotide DNA duplex, which was used to analyse DNA binding proteins in control and AOA1 nuclear extract as described in section 5.2.1.

We have previously shown that Aprataxin is present in the nucleolus (Chapter 4 and references 1,2) and that it interacts with via the FHA domain with the nucleolar rRNA processing factor nucleolin (1). In Figure 5.2 I found that Aprataxin is required for recruitment of a splicing factor to DNA. This raises the possibility that AOA1 cells may lower efficiency or defective splicing compared to control cells. I subsequently looked for interactions between Aprataxin and transcription and RNA processing factors using GST pull-downs.

GST pull-downs using a panel of overlapping Aprataxin domain-GST fusion proteins (schematic Figure 5.3) demonstrated a strong interaction between the Nuclear Localization domain of Aprataxin and the rRNA transcriptional TAF95 (Figure 5.4). This interaction occurred through the Nuclear Localization Signal region of

229 Aprataxin. TAF95 is a component of the SL1 complex, which is required for initiation of rRNA transcription (11). The SL1 complex interacts with an additional trans- activating protein, UBF, on rDNA promoters to form the pre-initiation complex (9). Consistent with the findings of Becherel et al. (1), Aprataxin also interacts with UBF (Figure 5.4). This interaction occurs primarily via the FHA domain but I observed some binding of UBF to the HIT domain of Aprataxin. Binding of UBF to the nuclear localization signal region was not observed. This indicates that Aprataxin may interact with two components involved in transcription initiation, TAF95 and UBF at the same time. Two UBF proteins are produced by alternate splicing of the UBF primary transcript (30). These HMG-box proteins differ by splicing out of 37 amino acids from HMG-box 2 in the shorter isoform, UBF-2 (Figure 5.5). Both proteins interact with RNA polymerase I via their acidic C-terminal regions. UBF-1 is an RNA polymerase I transcription factor and Histone H1 transcriptional anti-repressor (10). The in vivo function of UBF-2 is less clear given that its DNA binding, trans- activation, and anti-repressor activates are at least an order of magnitude less than UBF-1 (10). A more recent study has proposed that UBF-2 is an RNA polymerase II transcription factor (31) although little additional work has been published to date. I also identified an interaction between Aprataxin and TTF1 (Figure 5.4). This nucleolar protein is required for the termination of transcription by RNA polymerase I (12,13). Similarly to the interaction with UBF, the interaction between TTF1 and Aprataxin occurs via the FHA and HIT domains of Aprataxin (Figure 5.4). Additionally I observed an interaction between Aprataxin and the rRNA processing factor fibrillarin (Figure 5.4). Fibrillarin interacted primarily with the Nuclear Localization Signal fragment (amino acids 98 to 176) and to a lesser extent with the FHA domain fragment (amino acids 1 to 110). Fibrillarin is a nucleolar protein present predominantly in the DFC which interacts with small nucleolar RNAs (U3, U8 and U13, reference 32). Deficiency of fibrillarin in a mouse system is lethal (33) but conditional depletion experiments in yeast indicate that it is involved in early phases of rRNA transcript processing (34,35). These interactions implicate Aprataxin in rRNA transcription initiation, termination and processing. These suggest a role for Aprataxin in RNA polymerase I transcription and nucleolar rRNA processing.

230 Finally, I also observed an interaction between Aprataxin and the mRNA processing factor hnRNP U (Figure 5.4). hnRNP U is present in a large ribonucleoprotein complex (36) and has been implicated as an RNA polymerase II transcriptional enhancer (37). Combined with the interaction between Aprataxin and the putative RNA polymerase II transcription factor UBF-2, this indicates that Aprataxin may have a general role in transcription. This is substantiated by the interaction of Aprataxin with the transcription and RNA processing factors RNA helicase p68, ribosomal protein L3 and the FACT complex (Dr Olivier Becherel, Queensland Institute for Medical Research, Brisbane, Australia, unpublished observations).

Figure 5.3: Schematic of Aprataxin GST-fusion constructs. These pGEX 5.1 constructs are described in Gueven et al., (2) and previously in this thesis. Proteins were expressed and attached to beads by Dr Olivier Becherel (Queensland Institute for Medical Research, Brisbane, Australia).

231

Figure 5.5.4: Interaction of Aprataxin with RNA processing factors. Aprataxin-GST pull-downs were performed as described. Precipitated proteins were eluted in 5x SDS- PAGE loading buffer. These samples were then subjected to electrophoresis and immunoblotting. GST only is a control for non-specific binding. Methods are described in section 5.2.2.

Figure 5.5: Domain structure of UBF proteins. Taken from Kuhn et al., (10). The locations of HMG-boxes 1 to 5 are indicated. UBF-2 lacks 37 amino acids from HMG-box 2. The C-terminal acidic regions (solid black boxes) are required to bind RNA polymerase II.

232 The possibility that Aprataxin interacts with UBF and hnRNP U via a DNA or RNA intermediate was addressed by performing pulldowns in the presence of a DNA/RNA exonuclease. In Figure 5.6, cell extracts were treated with Benzonase prior to incubation with Aprataxin FHA and HIT domain GST fusions proteins (described in section 5.5.2). Pretreatment with Benzonase enhanced the interaction between Aprataxin and UBF and between Aprataxin and hnRNP U. This indicates that not only are these interactions independent of DNA and RNA, they are enhanced by its degradation. A likely explanation is that UBF and hnRNP U are sequestered by binding to polynucleotides, preventing their interaction with Aprataxin.

Figure 5.6: Polynucleotide-independent interaction between Aprataxin, UBF and hnRNP U. HeLa cell extract was incubated with Aprataxin-GST fusion proteins in the presence or absence of Benzonase as described (section 5.5.2). Specifically binding proteins were eluted as previously and subjected electrophoresis and Western blotting .

FHA domains are phosphopeptide interaction motifs (38). The interaction between Aprataxin and the DNA repair scaffold protein XRCC1 has been shown to require

233 CK2 dependant phosphorylation of XRCC1 (21). UBF is an in vitro CK2 substrate and phosphorylation of its C-terminal region is essential for its trans-activation activity (39,40). I therefore examined the effect of CK2 inhibition on the interaction between Aprataxin and UBF. Confluent HeLa cells were treated with the specific CK2 inhibitor TBB for 8 hours in cell culture media prior to the preparation lysates, which were subsequently used for GST pulldown experiments. Pre-treatment with increasing concentrations of TBB inhibited the interaction between Aprataxin and UBF (Figure 5.7). I noted that the interaction between Aprataxin and hnRNP U was not inhibited by TBB treatment, indicating that the interaction between Aprataxin and hnRNP U is independent of CK2 kinase activity, similar to the situation with Aprataxin’s interaction with nucleolin (1). Inhibition of CK2 was confirmed by a reduction in the affinity of the Aprataxin FHA domain for XRCC1, as previously reported (21). The interaction of Aprataxin with multiple transcription and RNA processing factors via the FHA domain suggests that multiple Aprataxin-processing factor complexes exist within a nucleus.

Figure 5.7: Interaction of Aprataxin with RNA processing factors- phosphorylation dependence. HeLa cells were pre-treated in 6- well dishes for 8 hours with the indicated concentrations of the Casein Kinase 2 inhibitor TBB in media. Cell lysate was then incubated with the FHA-GST fusion protein as previously, and subsequently eluted, resolved by SDS-PAGE and proteins detected by immunoblotting. TBB treatment and pulldowns were performed as described in section 5.5.2.

234 5.3.2 Aprataxin stabilizes UBF:

Binding of UBF to both the rDNA promoter is essential for the initiation of rRNA transcription (10). A scheme of the assembly of protein complexes at the rDNA promoter is shown in Figure 5.8. I examined whether UBF levels or localization are abnormal in Aprataxin deficient cells. Additionally, given the role of Aprataxin in DNA damage repair, I also investigated whether DNA damage affects UBF protein level or localization. Consistent with the role of UBF as an rRNA transcription factor, UBF immunostaining detects an exclusively nucleolar protein. Becherel et al. found that Aprataxin and UBF relocalize to nucleolar caps after inhibition of RNA polymerase I activity (1). Krulak et al. reported relocalization of UBF from fibrillar centres and the dense fibrillar component to nucleolar caps in response to ionizing radiation (41). AOA1 cells display only mild hypersensitivity to ionizing radiation (21) but have a more pronounced sensitivity to hydrogen peroxide (2,21). Before examining the localization and expression of UBF in AOA1 cells, I confirmed the specificity of UBF immunostaining using HeLa. I performed mock immunostaining without a primary antibody to detect non-specific secondary antibody interactions. The no primary antibody control panel shows that at under these conditions the α- mouse AlexaFluor-488 IgG does not produce detectable levels of non-specific staining or cross-reactivity with the α-rabbit secondary (Figure 5.9).

Figure 5.8: Protein complex assembly at the rDNA promoter. Adapted from Moss (11).

235 DAPI

Figure 5.9: UBF localization in HeLa cells. HeLa cells fixed in 10% formalin for 10 minutes, dehydrated in 100% methanol for 10 minutes, and rehydrated in PBS. Cells were blocked and permeabilized in PBS containing 5% bovine serum and 0.05% Triton X-100 prior to incubation with or without mouse α-UBF antibody (1/100) for 1 hour at 37°C. Nucleophosmin staining was performed as described for the FUrd pulse experiments. Staining was detected using rabbit-α-mouse Alexafluor-488 and goat α- rabbit Alexafluor-594 IgG, and cells were counterstained with DAPI. The scale bar indicates 20 µm.

Based on the sensitivity profile of AOA1 cells, I examined the effect of hydrogen peroxide and Aprataxin protein deficiency on UBF distribution. APTX corrected (FD105 M21) and vector only (FD105 M20) AOA1 cell lines were exposed to 1 mM hydrogen peroxide (or mock treated) for 10 minutes. Cells were subsequently fixed after a 30 minute recovery. Relocalization of UBF was not observed after treatment in either cell line (Figure 5.10). This could be due to the use of different cell lines and a different DNA damaging agent from Krulak et. al (41). Comparison of UBF immunostaining of untreated corrected and uncorrected AOA1 cell lines reveals that Aprataxin deficiency results in a slightly diffuse UBF staining pattern (Figure 5.10). This could be due to weaker association of UBF-1 with rDNA promoters or an alteration in nucleolar structure or trafficking in the absence of Aprataxin.

236 DAPI

Figure 5.10: Effect of Aprataxin deficiency on UBFs response to DNA damage. The impact of Aprataxin deficiency on UBF level and localization in response to hydrogen peroxide treatment was determined by treating corrected and vector only AOA1 fibroblast cell lines (10 minutes of 1 mM H2O2 in DMEM with 12% FCS, no recovery period). Cells were then fixed, dehydrated, blocked, and stained as described in methods. The scale bar indicates 20 µm.

Following this I quantitatively examined the effect of Aprataxin deficiency and hydrogen peroxide treatment on UBF staining. UBF immunostaining of FD105 M20 and FD105 M21 cells with and without hydrogen peroxide treatment was quantified as described in section 5.2.3 (Figure 5.11). Aprataxin deficiency resulted in a significant reduction in the steady state level of UBF (20.6% reduction, p<0.05) suggesting that Aprataxin affects UBF protein stability. This is reminiscent of the reduced stability of nucleolin observed in AOA1 cells (1). Hydrogen peroxide treatment of both corrected and uncorrected AOA1 cells resulted in a significant reduction of UBF protein levels. This effect does not seem to be Aprataxin related, as a similar reduction occurs in both FD105 M20 and FD105 M21 cell lines (a 18.7% reduction in corrected cells, p<0.02 and a 21.2% reduction in uncorrected cells,

237 p<0.03) and could be due to inhibition of UBF mRNA transcription or translation, or elevated UBF degradation after DNA damage.

Figure 5.11: Quantification of the effect of hydrogen peroxide on UBF staining in FD105 M20 and M21 fibroblasts. The effect of Aprataxin deficiency on UBF steady- state protein level in response to hydrogen peroxide treatment was determined by quantification of UBF immunofluorescence in corrected and defective fibroblast cell lines. Staining intensity was quantified using ImageJ as described in materials and methods. p-values were calculated using Students t-test.

We have shown that Aprataxin deficiency results in a small but significant reduction in UBF immunostaining suggesting that Aprataxin is involved in stabilization of UBF. Given that all commercial UBF antibodies detect both UBF-1 and UBF-2 I used Western blotting to determine if Aprataxin deficiency causes differential expression of either protein. Immunoblotting of the corrected AOA1 cell line FD105 M21

238 confirmed that UBF-1 (97kDa) and UBF-2 (94kDa) are present as in comparable amounts in fibroblasts (Figure 5.12). Surprisingly, I found that in the absence of Aprataxin, full length UBF-1 and UBF-2 are barely detectable (Figure 5.12). Instead UBF Western blotting of FD105 M20 UBF detects two smaller bands at approximately 77 and 74kDa. These bands are also present, but very faintly, in the corrected cell line and probably correspond to N-terminal UBF fragments which contain the HMG-box 2 alternatively spliced region (10). Proteolysis of UBF into 77 and 74kDa HMG-box 2 containing fragments should release an approximately 20kDa C-terminal fragment. Rather I detected an α-UBF reactive band of approximately 50kDa in the Aprataxin deficient cell line. This may be the C-terminal fragment migrating abnormally due to its abundance of acidic residues. Such abnormal migration patterns are well documented in the study of acidic proteins, for example nucleolin has a molecular weight of 77kDa but actually migrates at 110kDa due to its large central acidic region. UBF also has 21 potential CK2 phosphorylation sites in its C-terminal region (42), and phosphorylation of these may alter migration of the acidic tail fragment significantly.

Figure 5.12: Lack of Aprataxin destabilizes UBF. Total extract from FD105 M20 and FD105 M21 cells was subjected to protein electrophoresis and Western blotting. The resulting nitrocellulose membrane was probed for Aprataxin and UBF. Apoptosis Inducing Factor (AIF) immunoblotting shows equal loading. Described in section 5.2.4.

239 Aprataxin deficient cells have reduced levels of UBF antibody reactive protein (Figure 5.12), with mildly abnormal localization (Figure 5.10). Furthermore in the absence of Aprataxin UBF is degraded into two fragments, which may correspond to the DNA (74 and 77kDa) and RNA polymerase I binding regions (20kDa, migrating at 50kDa) (Figure 5.12). A C-terminal deleted UBF-1 protein (ΔC552) displays no transcriptional anti-repression or trans-activation activity (10), indicating that the C- terminal region is critical for coupling UBFs DNA binding activity to other transcriptional effectors. Given the interaction of Aprataxin with RNA processing factors and its requirement for stabilization of UBF, I propose that AOA1 cells may harbour defects in rRNA synthesis and processing. This idea is supported by the observation that AOA1 cells have a defect in the early stages of pre-rRNA processing (1).

5.3.3 Transcriptional defects in AOA1 cells:

We have shown that Aprataxin interacts with proteins involved in RNA metabolism, and that Aprataxin deficient cells do not have full length UBF. I have demonstrated in the previous chapter that AOA1 cells have a range of DNA repair defects, and DNA damage in coding regions has been shown to inhibit transcription (43,44). I was therefore interested in the relationship between transcription, DNA damage and Aprataxin deficiency. To explore the role of Aprataxin in transcription I developed an in vivo nucleotide analogue incorporation protocol.

Several methods for analysis of RNA and DNA synthesis are based on the incorporation of modified nucleotides into RNA or DNA of living cells (for examples see 45,46). Nucleotide analogue incorporation assays detect only nascently synthesized strands, making them appropriate for analysis of global polynucleotide synthesis within a defined period of time. Nucleotide analogue incorporation has one major advantage over real-time PCR for analysis of changes in transcriptional activity in response to stimuli. Short-term or subtle changes in transcriptional activity may be difficult to detect using real-time PCR because of the ‘background’ of transcripts which were produced before the stimulus was applied. Comparatively, nucleotide

240 analogue incorporation measures global synthetic activity over a defined period of time. This makes them ideal for measuring inducible changes in global transcriptional profile, but their disadvantage is that they cannot measure synthesis of a single gene product, as real-time PCR can. I selected nucleotide analogue incorporation to examine the effect of Aprataxin deficiency and DNA damage on transcription for two reasons. Firstly, I was interested in measuring short-term changes in transcription, and PCR may not be able to detect subtle effects. Secondly, the majority of the RNA a cell produces is rRNA and I wanted to examine rRNA expression changes, so lack of gene-specificity is acceptable. Incorporation protocols can be broadly divided into two categories: those using radiolabelled nucleotides and those using halogenated nucleotide analogues. Radiolabelled (generally tritiated) nucleotides are ideal for experiments where high-throughput or very high sensitivity is required, as such experiments are conveniently performed in multi-well plate format and quantified by scintillation counting. On the other hand incorporation of halogenated nucleotides is detectable by immunofluorescence and as such can examine nascent synthesis in individual cells. As such, the synthetic activity of sub-populations of cells can be examined (for example, RNA synthesis can be measured with respect to cell cycle or marker expression). Additionally incorporation of halogenated nucleotides facilitates the examination of the sub-cellular distribution of RNA synthesis. Given the spatial separation of mRNA and rRNA synthesis and the interaction of Aprataxin with both RNA polymerase I and II transcription factors, I selected halogenated nucleotide incorporation and fluorescent microscopy to examine the transcriptional activity of Aprataxin deficient cells.

The experimental incorporation of halogenated nucleotides can be broken into several phases, as outlined in Figure 5.13. Initially adherent cells are incubated with the nucleotide, either in media or in a solution which will permeabilize the cell membrane. The nucleotide then enters the cell and is bound by the respective polymerases. The labelling solution is then removed and replaced with normal media so that incorporation can occur.

241 “Pulse” with analogue Remove pulse mixture.

1. 2. 3.

Figure 5.13: Schematic of nucleotide analogue incorporation assay. Nucleotide synthesis can be monitored in vivo by the incorporation of radiolabelled or halogenated nucleotides. To examine RNA synthesis by fluorescent microscopy, adherent cells are incubated with a uracil analogue, sometimes in the presence of digitonin or a transfection agent (1). This analogue enters the cells and becomes incorporated into cellular nucleotide pool (2). The analogue-containing media is then removed and replaced with fresh media for an optimized period of time to allow incorporation (3). Unincorporated molecules are then washed out prior to detection of incorporated nucleotides by immunostaining.

rRNA is synthesized only in the nucleolus and synthesis rates are elevated in rapidly growing cells. Therefore I initially examined nucleotide incorporation in HeLa cells, due to their rapid growth rate and large nucleoli. Previous authors have reported that the uridine analogue FUrd can pass through the cell membrane and is incorporated exclusively into RNA (41). Once the analogue is incorporated the cells are fixed and permeabilized to ‘wash out’ unincorporated nucleotides and subsequently stained with a BrdU antibody. To optimize this staining protocol I exposed HeLa cells to FUrd by the passive incorporation method before treatment with one of three fixatives (Figure 5.14) and analysed FUrd incorporation with mouse α-BrdU (Sigma). No specific incorporation of FUrd was observed under any of the fixation conditions, indicating that either the analogue did not enter the cells or that immunostaining failed.

242 DAPI

Figure 5.14. Optimization of fixation method. HeLa cells were treated with FUrd according to the Passive Incorporation protocol and processed as described in section 5.2.5.1. Fixation conditions tested were: 2% paraformaldehyde (PFA) in PBS, 70% ethanol in PBS and a 1:1 mixture of acetone and methanol. Coverslips were subsequently incubated with 1/500 rabbit α-nucleophosmin (NEB) and 1/100 mouse α-BrdU (Sigma) in PBS containing 5% bovine serum for 3 hours at 37°C. Coverslips were washed three times in PBS and incubated with 1/500 AlexaFluor-488 α-mouse and 1/500 AlexaFluor-594 α-rabbit IgG, washed once in PBS and counterstained with DAPI. Coverslips were washed three times in PBS and mounted in Moviol. Images were acquired on a Zeiss AxioSkop using the x63 oil immersion objective. The scale bar indicates 100 µm.

To examine the possibility of staining failure I tested several of BrdU antibodies. Shown in Figure 5.15 is immunostaining of HeLa cells treated with FUrd by the

243 passive incorporation method prior to fixation and staining with antibodies from Sigma and Becton Dickinson (using this method I have also attempted staining with two BrdU antibodies from Abcam, data not shown). No specific incorporation was observed. I concluded that either FUrd cannot pass across the mammalian cell membrane or that these antibodies can only detect bromine analogues.

DAPI

Figure 5.15: Optimization of staining methods. HeLa cells were treated with FUrd according to the Passive Incorporation protocol described in section 5.2.5.1. Coverslips were then incubated with either 1/100 mouse α-BrdU (Sigma) or 1/50 mouse α-BrdU (Becton Dickinson) in PBS containing 5% bovine serum for 3 hours at 37°C. Coverslips were washed three times in PBS and incubated with 1/500 AlexaFluor-488 α-mouse, washed once in PBS and counterstained with DAPI. Coverslips were washed three times in PBS and mounted in Moviol. Images were acquired on a Zeiss Axioskop using the x63 oil immersion objective. The scale bar indicates 100 µm.

244 We subsequently examined incorporation of both FUrd and BrUTP using an established method for membrane permeabilization, hypotonicity (section 5.2.5.2 and (47). HeLa cells were pulsed with FUrd or BrUTP prior to fixation and immunostaining (Figure 5.16). No incorporation was observed. Given that the BrdU antibody should react with bromouridine labelled RNA I concluded that membrane permeabilization using this hypotonicity method was not successful. I therefore tried two other permeabilization methods- treatment with the transfection reagents Lipofectamine 2000 (Invitrogen) and FuGene6 (Roche).

DAPI

Figure 5.16: Optimization of incorporation conditions- BrUTP versus FUrd. HeLa cells were treated with BrUTP or FUrd according to the Hypotonicity protocol described in section 5.2.5.2. Coverslips were permeabilized and blocked as described. Cells were incubated with 1/100 mouse α-BrdU (Sigma) in PBS containing 5% bovine serum for 3 hours at 37°C. Coverslips were washed three times in PBS and incubated with 1/500 AlexaFluor-488 α-mouse, washed once in PBS and counterstained with DAPI. Coverslips were washed three times in PBS and mounted in Moviol. Images were acquired on a Zeiss AxioSkop using the x63 oil immersion objective. The scale bar indicates 100 µm.

245 Successful incorporation and staining of FUrd treated HeLa cells was obtained using the Lipofectamine 2000 protocol and Sigma mouse α-BrdU antibody (Figure 5.17). Intense nucleolar incorporation and weaker nucleoplasmic incorporation was observed, based on colocalization with nucleophosmin and DAPI counterstaining respectively. This is in agreement many reports which find that rDNA is very transcriptionally active (reviewed in 3,48).

DAPI

Figure 5.17: 5-Fluro Uridine incorporation by HeLa cells- co-localization with nucleophosmin. HeLa cells were incubated for 15 minutes in a mixture of Lipofectamine 2000 transfection reagent and FUrd in OptiMEM as described in section 5.2.5.3. Coverslips were processed as described. Cells were subsequently incubated with mouse α-BrdU (1/100, Sigma) and rabbit α- nucleophosmin (1/250, NEB) at 37° for 2 hours. Staining was detected by incubation with AlexaFluor-488 α- mouse and AlexaFluor-594 α-rabbit conjugated IgG and cells were counterstained with DAPI. Coverslips were mounted in Moviol and images were acquired on a Zeiss AxioSkop fluorescent microscope using the x63 oil immersion objective. The scale bar indicates 100 µm.

rRNA is synthesized in discrete transcription factories within the nucleolus (5,16). In HeLa cells each nucleus has approximately 30 rRNA transcription factories arranged into one or more nucleoli (16). I subsequently examined the subnucleolar distribution of rRNA synthesis. Cells pulsed with FUrd according to the Lipofectamine 2000 method (section 5.2.5.3) were immunostained and examined by fluorescent microscopy using a 100x objective. The resulting images were enhanced in Adobe Photoshop 3.0 to emphasize regions of intense transcriptional activity (Figure

246 5.18). I found that FUrd incorporation into nucleolar RNA occurs in many discrete foci (Figure 5.18), in agreement with the existence of subnucleolar domains (5,16).

DAPI

Figure 5.18. Distribution of rRNA synthesis within the nucleolus. HeLa cells were pulsed with FUrd according to the Lipofectamine 2000 protocol and stained with 1/100 mouse α-BrdU (Sigma). After nucleophosmin staining, secondary detection and DAPI staining, incorporation was assessed by fluorescent microscopy on a Zeiss AxioSkop using the x100 oil immersion objective. 488 nm and 594 nm channel levels were adjusted in Adobe Photoshop 3.0 such that the weakly stained regions were rendered invisible, to highlight nucleolar transcription factories. The scale bar indicates 5 µm.

Previous immunostaining optimization experiments failed due to a lack of FUrd uptake by the cell (Figures 5.14 to 5.16). Having determined that lipofection efficiently permeabilizes mammalian cell membranes to FUrd, antibody staining procedures were re-evaluated. I compared FUrd staining in HeLa cells using two different BrdU antibodies (Figure 5.19). Both Sigma and Abcam antibodies produce diffuse non-specific staining of the whole cell and detect nucleolar incorporation of FUrd, however the Sigma antibody produces more intense specific staining (Figure 5.19). Given that the level of background for the Abcam and Sigma antibodies is the same, the Sigma one displayed a better signal-to-noise ratio and was used for subsequent experiments.

247 DAPI

Figure 5.19: FUrd incorporation by HeLa cells: optimization of antibody staining. HeLa cells were pulsed with FUrd according to the Lipofectamine 2000 method. After processing of the coverslips cells were stained with either equal dilutions (1/100) of BrdU primary antibodies (Sigma or Serotec as indicated) prior to detection with appropriate conjugated secondary antibodies as previously and DAPI counterstaining. The scale bar indicates 50 µm.

FUrd staining with the Sigma antibody was subsequently optimized by titration. FUrd pulsed and control treated cells were incubated with the antibody at 1/50 or 1/250 dilutions before detection using the same concentration of secondary antibody (Figure 5.20). A 1/50 dilution of the antibody produces very intense granular nucleolar staining and weak granular nucleoplasmic staining in FUrd treated cells. Some staining is visible in FUrd untreated cells, indicating that at this concentration the antibody interacts to a limited degree with non-halogenated nucleotides. This background is predominantly nucleolar indicating that it is due to the antibody binding non-halogenated RNA and not DNA. At a 1/250 dilution specific punctate nucleolar staining is observed however this staining is only a few times stronger than the background. Based on this titration the optimal antibody concentration for these

248 experiments is between 1/50 and 1/250. For the following experiments a 1/100 dilution was used.

DAPI

Figure 5.20: FUrd incorporation by HeLa cells- optimization of antibody dilution. HeLa cells, treated with FUrd and prepared as in Figure 5.19, were incubated with the indicated concentrations of mouse α-BrdU antibody (Sigma) in PBS containing 5% bovine serum for 2 hours at 37°C. Nucleophosmin staining, secondary antibody detection and DAPI counterstaining were performed as previously. Cells were mounted in Moviol and images acquired on a DeltaVision fluorescent microscope. The scale bar indicates 10 µm.

Following the optimization of antibody concentration, I optimized washing conditions. A number of buffer components can be adjusted to alter the strength of specific and non-specific antibody interactions including osmolarity and detergent concentration. Decreased salt concentrations provide higher stringency. I prepared FUrd treated cells on coverslips and stained them in parallel except for the washing steps after both primary and secondary antibody incubations. Coverslips were washed three times (5 minutes each wash) in 1x or 0.5x PBS as indicated after primary and secondary antibody incubations and staining examined by fluorescent microscopy

249 (Figure 5.21). Washing cells with 0.5x PBS resulted in weaker nucleolar staining compared with 1x PBS. The nucleoplasmic staining intensity appears to be similar under both conditions however under high magnification the nucleoplasmic staining of the 0.5x PBS treated slide appears slightly fuzzy and individual transcription factories are less clear than on the 1x PBS treated slide. The elevated stringency of 0.5x PBS appears to disrupt the specific antibody-nucleotide interactions and may damage the micro-structure of individual transcription factories. I have therefore used 1x PBS washes in all subsequent experiments.

DAPI

Figure 5.21: FUrd incorporation by HeLa cells- optimization of wash conditions. HeLa cells were pulsed with FUrd according to the Lipofectamine 2000 protocol. Slides were prepared and stained (Sigma mouse α-BrdU and secondary antibody as previously described) in parallel except for the washes after both primary and secondary antibody incubations. Coverslips were washed in either 1x or 0.5x PBS three times for 5 minutes per wash after both antibody incubations. Cells were subsequently counterstained with DAPI in 1x PBS and mounted. Cells were examined on a Zeiss AxioSkop (x63 oil immersion objective). The scale bar indicates 100 µm.

250 Having established specific staining conditions I confirmed that the observed nucleolar polynucleotide synthesis is due to transcription by RNA polymerase I using a specific RNA polymerase I inhibitor. Actinomycin D is a specific RNA polymerase I inhibitor when used at low concentrations (less than 5 µg/mL, references 49,50). RNA polymerase I was inhibited in HeLa cells by treatment with 2 µg/mL actinomycin D for 3 hours in cell culture media. Nascent RNA synthesis was subsequently measured by pulsing with FUrd (Lipofectamine 2000 protocol) with processing and staining as optimized (Figure 5.22). Cells not treated with actinomycin D displayed limited punctate nucleoplasmic and intense nucleolar staining. Cells treated with actinomycin D displayed nucleoplasmic staining but very weak nucleolar staining. This residual nucleolar signal may be due to incomplete RNA polymerase I inhibition or fluorescence leakage from nucleoplasmic RNA above and below the focal plane of this wide-field microscope.

DAPI

Enlargement of top

cell in + Actinomycin D

nucleolus

Figure 5.22. Inhibition of RNA synthesis by Actinomycin D. HeLa cells on coverslips were pre- treated with actinomycin D (2 µg/µL) in DMEM (12% FCS) for 3 hours at 37°C. Cells were subsequently pulsed with FUrd according to the Lipofectamine 2000 protocol outlined in section 5.2.5.3. Coverslips were fixed, permeabilized and blocked as described in section 5.2.5.3.. Incorporation was detected using the Sigma mouse α-BrdU antibody, 1/100 in PBS with 5% bovine serum for 3 hours at 37°C. The secondary antibody and DAPI counterstaining were performed as previously. Images were acquired on a Zeiss AxioSkop using the x63 oil immersion objective. The scale bar indicates 100 µm.

251

Having confirmed that the observed nucleolar RNA synthesis is dependant on RNA polymerase 1 activity, I optimized the time allowed for incorporation after FUrd pulsing. I previously observed some variability between duplicate slides when cells were treated with FUrd using the Lipofectamine protocol, indicating that the composition of Lipofectamine-FUrd complexes may not be very stable. After optimizing a new lipofection protocol for FuGene6 transfection reagent this problem has been alleviated. HeLa cells on coverslips were pulsed with FUrd according to the FuGene6 lipofection protocol (section 5.2.5.4). After pulsing cells were washed in pre-warmed growth media (DMEM with 12% FCS). They were subsequently incubated in fresh media for the indicated times prior to fixation in 10% formalin in PBS. Coverslips were stained as optimized and incorporation visualized on a DeltaVision microscope (Figure 5.23). Only weak nucleolar incorporation is visible when cells are fixed immediately after pulsing. Incorporation intensity increases up until the longest time point, 20 minutes. Longer time points were not examined as I aim to measure the acute effects of DNA damage on transcription. Longer incubations may result in higher levels of incorporation but the quantity and nature of DNA lesions may alter substantially during this time. I have therefore used a chase time of 20 minutes for the following experiments.

252 DAPI

Figure 5.23: FUrd incorporation by HeLa cells- optimization chase time. HeLa cells were pulsed FUrd according to the FuGene6 protocol outlined in section 5.2.5. After washing, cells were chased in DMEM (12% FCS) for the indicated times prior to fixation, dehydration and blocking as described. Immunostaining was performed as previously optimized (1/100 dilution of Sigma mouse α-BrdU primary antibody, 1/500 dilution of AlexaFluor-488 α-mouse) prior to DAPI staining, mounting, and imaging on a DeltaVision microscope. The scale bar indicates 10 µm.

DNA damaging agents cause formation of lesions in both coding and non-coding regions. Lesions could be formed at higher frequency in coding regions due to their open chromatin structure. Lesions in coding regions can result in generation of truncated, frameshifted or point mutated transcripts (51-53). To prevent the formation of mutated transcripts and subsequent potentially harmful proteins, mechanisms have arisen which inhibit transcription in response to DNA damage (41) and specifically repair coding regions (54). Kruhlak et al. demonstrated that cells deficient in ATM

253 kinase activity display radioresistant RNA synthesis (RRS), indicating that the ATM signalling pathway is involved in inhibition of transcription after irradiation (41).

To further validate the optimized methodology I examined the effect of γ-irradiation on RNA synthesis in HeLa cells. Cells were exposed to 10 Gy of γ-irradiation (or mock treated) and allowed to recover for 30 minutes in growth media before pulsing with FUrd (FuGene6 method). Cells were fixed after a 20 minute chase and immunostained as optimized. Consistent with the findings of Krulak et al., irradiation resulted in an inhibition of nucleolar and nucleoplasmic RNA synthesis in the cells (Figure 5.24 and reference (41). Quantification of 45 cells on each slide revealed that irradiation resulted in an average of 20% inhibition (p<0.01) of global RNA synthesis over the 65 minute period between the end of the treatment and fixation (Figure 5.25). This seems consistent with the findings of Krulak et al. who reported that nascent RNA synthesis is reduced by approximately 50 percent and 15 percent respectively 20 and 40 minutes after 5 Gy irradiation (41).

DAPI

Figure 5.24: Effect of γ-irradiation on RNA synthesis in HeLa cells. HeLa cells were treated with 10 Grays of γ-radiation (or untreated) using a MDS Nordion Gammacell irradiator. Dose rate at the time of this experiment was 1 Gy per minute. 30 minutes after the end of treatment the cells were pulsed with FUrd according to the FuGene6 protocol. After pulsing cells were chased for 20 minutes in DMEM (12% FCS) and then fixed in 10% formalin. Subsequent sample preparation and staining is as described previously. The scale bar indicates 100 µm. This trend was observed in two separate experiments.

254

Figure 5.25: Quantitation of effect of γ-irradiation on RNA synthesis in HeLa cells. The effect of irradiation on RNA synthesis was determined by quantification of immunostaining using ImageJ as described in section 5.2.5. 45 cells from each slide were quantified. The resulting data sets were compared using Students t-test in Microsoft Excel. This trend was observed in two separate experiments

Having optimized nascent RNA synthesis detection and validated the reported inhibition of transcription after γ-irradiation, I analysed the role of Aprataxin in this cellular response. At the commencement of these experiments the only adherent Aprataxin deficient cell line available to us was the non-immortalized AOA1 cell line FD105. Neonatal Foreskin Fibroblasts (NFFs) are an appropriate control cell line for FD105. I therefore examined FUrd incorporation in untreated NFFs. FUrd is not incorporated efficiently by NFFs, and individual cells display highly variable levels of incorporation (Figure 5.26). Similarly poor levels of incorporation were seen in the AOA1 cell line FD105 (not shown). This could be due to their slow growth rate compared to HeLa (they need less RNA) or differences in membrane permeability (primary cells are generally more difficult to transfect than transformed cell lines). The poor incorporation combined with slow growth rates (doubling time of 7 to 10 days for FD105) prompted us to look for an alternative system to study the role of Aprataxin in transcription.

255 DAPI

Figure 5.26. Incorporation of 5’flurouridine by neonatal foreskin fibroblasts. Sub- confluent neonatal foreskin fibroblasts were pulsed with FUrd according to the FuGene6 protocol. The chase was performed in DMEM (12% FCS) for 20 minutes. Coverslips were subsequently processed as optimised and examined using a Zeiss AxioSkop (x63 oil immersion objective). The scale bar indicates 100 µm.

We subsequently obtained hTERT immortalized AOA1 and corrected cell lines, courtesy of Professor Keith Caldecott (University of Sussex, Brighton, UK). These cell lines are ideal for biochemical experimentation as they are isogenic. I initially optimized the incorporation conditions for the corrected cell line, FD105 M21 and determined that 10 or 20 minute chase times provide good levels of incorporation (Figure 5.27).

256 DAPI

Figure 5.27: FUrd incorporation in FD105 M21 fibroblasts- optimization of labelling conditions. Corrected AOA1 fibroblasts (FD105 M21) were pulsed with FUrd and processed according to the FuGene6 protocol described in section 5.2.5.4. Chase times are indicated. Coverslips were stained with 1/100 mouse α-BrdU (Sigma) in PBS containing 5% bovine serum as described previously. Secondary antibody incubation, DAPI counterstaining and image acquisition are as described previously. The scale bar indicates 100 µm.

Having established appropriate incorporation and staining conditions for these new cell lines, I used them to investigate the role of Aprataxin in transcription inhibition after irradiation. FD105 M21 (corrected) and FD105 M20 (vector only) cell lines were mock treated or exposed to 5 Gy of γ-radiation. Nascent RNA synthesis was examined by FUrd incorporation using the FuGene6 method and subsequent immunostaining performed as optimized. Irradiation resulted in a 31.2% inhibition of RNA synthesis in the corrected cell line (Figure 5.28, quantified in Figure 5.29). The transcriptional activity of Aprataxin deficient cells is not affected by irradiation (Figure 5.28, quantified in Figure 5.29), indicating that Aprataxin causes RRS.

257 DAPI

Figure 5.28: Effect of Aprataxin deficiency on the transcriptional response to γ- irradiation. Corrected and uncorrected AOA1 fibroblasts were treated with 5 Gy γ- irradiation (or untreated), and pulsed with FUrd 30 minutes later as previously described (section 5.2.5.4). Subsequent sample processing and staining and image acquisition was performed as previously described. Scale bar indicates 100 µm.

258

Figure 5.29: Quantification of the effect of Aprataxin deficiency on the transcriptional response to γ-irradiation. The impact of Aprataxin deficiency on RNA synthesis in response to γ-irradiation was determined by quantification of FUrd immunostaining using ImageJ as described previously and analysed using the t-test in Microsoft Excel. This trend was observed in two separate experiments.

Given that AOA1 cells display limited sensitivity to γ-irradiation, I was interested to examine their transcriptional response to an agent they are more sensitive to. In Chapter 4 I identified an elevated level of oxidative DNA damage in AOA1 cells, consistent with their hypersensitivity to the oxidative stress inducing agent hydrogen peroxide. Therefore I treated FD105 M20 and FD105 M21 cell lines with hydrogen peroxide for 10 minutes (500 µM and 1 mM in media). Cells were then washed and allowed to recover in media without hydrogen peroxide for 30 minutes before initiation of the FuGene6 FUrd incorporation protocol and immunostaining as optimized previously. I found that a short period of treatment with high-dose hydrogen peroxide did not cause a significant decrease in the RNA synthesis rate of APTX-corrected AOA1 cells (Figure 5.30, quantification shown in Figure 5.31). This result demonstrates the different effects of hydrogen peroxide and ionizing radiation on transcription and highlights the complexity of the DNA damage response. In

259 contrast to irradiation, treatment of uncorrected AOA1 cells with hydrogen peroxide caused significant inhibition of nascent RNA synthesis both in the nucleolus and the nucleoplasm (Figure 5.30, quantification shown in Figure 5.31). The elevated basal level of oxidative stress present in AOA1 cells (section 4.3.6) may account for this dramatic transcriptional response. Additionally, hydrogen peroxide treatment may also induce formation of RNA polymerase-blocking lesions which AOA1 cells cannot repair and thus cause inhibition of global transcription.

DAPI

Figure 5.30: Effect of Aprataxin deficiency on the transcriptional response to oxidative stress. Corrected and vector only AOA1 fibroblasts were treated with 1 mM

H2O2 in DMEM (12% FCS) for 10 minutes. Cells were then allowed to recover in normal media for 30 minutes prior to FUrd pulsing (FuGene6 protocol), processing and imaging as previously described. Scale bar indicates 100 µm.

260

Figure 5.31: Quantification the effect of Aprataxin deficiency on the transcriptional response to oxidative stress. The impact of Aprataxin deficiency on the transcriptional response to of hydrogen peroxide treatment was determined by quantification of FUrd immunostaining (FuGene6 protocol) as described previously (section 5.2.5). This trend was observed in three separate experiments.

Some nucleolar proteins, including Aprataxin, nucleolin and UBF, have been reported to relocalize to nucleolar caps after inhibition of rRNA synthesis (1). Conflicting evidence exists within the literature on wether or not nucleophosmin relocalizes to nucleolar caps as a result of inhibition of rRNA synthesis (see reference 55 versus 56). Here I report relocalization of nucleophosmin to structures which appear to be nucleolar caps in some Aprataxin deficient cells after treatment with hydrogen peroxide (Figure 5.32). This supports the idea that RNA polymerase I is inhibited by DNA damage in Aprataxin deficient cells.

261 DAPI

Figure 5.32: Distribution of nucleophosmin after DNA damage in AOA1 cells. Nucleophosmin immunostaining of an FD105 M20 cell after hydrogen peroxide treatment (1 mM hydrogen peroxide for 10 minutes, 30 minute recovery). Cells were fixed in 10% formalin in PBS followed before permeablization, blocking and staining (1/500 rabbit α-nucleophosmin, 1/1000 AlexaFluor-594 α-rabbit IgG). Nucleophosmin appears to have re-localized to nucleolar caps. Scale bar indicates 10 µm.

262 5.4 DISCUSSION

5.4.1 Interaction of Aprataxin with transcription factors:

Transcription is an essential and highly regulated metabolic function. I have demonstrated interactions between Aprataxin and the rRNA transcription activation factors TAF95 and UBF (section 5.3.1). Two UBF isoforms, generated by alternative splicing are present in mammalian cells. The shorter isoform, UBF-2 lacks part of the second of five tandem HMG-boxes (10). Both UBF-1 and UBF-2 are able to bind to Aprataxin, indicating that HMG box 2 is not required for the interaction with Aprataxin. Given that the HMG-box is a DNA interaction domain and that the interaction between Aprataxin and UBF occurs independently of DNA, the HMG- boxes are unlikely to be directly involved in this interaction. The interaction between Aprataxin and UBF most likely occurs via the C-terminal region of UBF. This region of UBF has been shown to be essential for both trans-activation and anti-repression activities of UBF-1 (10). This region interacts with the SL1 complex and is essential for formation of the pre-initiation complex (57). Deletion of the acidic tail ablates the ability of UBF-1 to remove the transcriptional repressor Histone 1 from rDNA promoters, indicating that acidic tail interactions are essential for UBF-1 mediated transcriptional anti-repression (10). This region contains multiple phosphorylation sites and has been shown to be phosphorylated in vivo (57), indicating that it has the potential to interact with Aprataxin’s’ FHA phospho-peptide interaction domain.

TAF95 is a member of the SL1 complex, which interacts with UBF at rDNA promoters to form pre-initiation complexes (9). The interaction of SL1 and UBF at the rDNA promoter stabilizes the interaction of UBF with the promoter, enhancing rRNA transcriptional activity (58). TAF95 interacts strongly with the Nuclear Localization fragment of Aprataxin (section 5.3.1). Thus two proteins involved in the formation of pre-initiation complexes, UBF and TAF95, interact with different regions of Aprataxin, so all of these proteins may be present in a single multi-protein complex at rDNA promoters. This implies a critical role for Aprataxin in transcriptional regulation. This hypothesis is supported by the observation that the FHA domain of

263 Aprataxin also interacts with the large subunit of RNA polymerase II (Dr Olivier Becherel, Queensland Institute for Medical Research, Brisbane, Australia, unpublished observations).

We also demonstrated an interaction between Aprataxin and the rRNA processing factor fibrillarin (section 5.3.1). This interaction occurred via the nuclear localization region of Aprataxin (amino acids 98 to 176) and to a weaker extent with the FHA domain fragment (amino acids 1 to 110). Based on this I concluded that the minimal fibrillarin binding region on Aprataxin is between amino acids 98 and 110. Fibrillarin is an essential rRNA processing factor in mammals (33) and depletion of fibrillarin in yeast results impairment of all rRNA processing steps and a subsequent reduction in the number of cytoplasmic ribosomes (34,35). The interaction of Aprataxin with fibrillarin and hnRNP U, together with the previously reported interaction with nucleolin (1), implicates Aprataxin in RNA processing as well as transcription.

5.4.2 Aprataxin dependant stabilization of UBF:

In this chapter I identified a critical role for Aprataxin in stabilization of both UBF-1 and UBF-2 (section 5.3.2). Immunostaining of Aprataxin deficient cells revealed that they possess approximately 20 percent less UBF protein than corrected cells. Analysis of the molecular weight and intensity of α-UBF reactive proteins in Aprataxin deficient and corrected cells revealed that Aprataxin deficient cells only have degraded UBF protein. Thus Aprataxin stabilizes UBF and deficiency of Aprataxin results in proteolysis of UBF into HMG-box and C-terminal tail regions. This separates the two UBF functional domains and may result in disconnection between UBFs DNA binding and protein interaction functions. I noted that while Aprataxin deficient cells still display predominantly nucleolar UBF localization, this staining is more diffuse and cytoplasmic staining is notable.

264 Transcription of rRNA requires both binding of the rDNA upstream control element and the SL1 complex by UBF. Given the degradation of UBF in the absence of Aprataxin I predicted that Aprataxin deficient cells may have a transcriptional deficit.

5.4.3 Transcriptional defects in Aprataxin deficient cells:

Becherel et al. reported that Aprataxin deficient cells display a minor deficit in their levels of rRNA primary transcript (1). This was based on the analysis of steady-state levels the 47S and 45S rRNAs in AOA1 cells by real-time PCR, using primers directed against the 5’ETS. AOA1 lymphoblastiod cell lines contain 15 percent less 47S and 45S rRNA than control cells. Such a defect could be caused by either defective pre-rRNA processing or reduced levels of nascent RNA polymerase I transcription. Given that Aprataxin is required for stabilization of the RNA polymerase I transcription factor UBF-1, I hypothesized that AOA1 cells may have reduced rates of RNA polymerase I activity. I subsequently examined the rate of nascent RNA synthesis in Aprataxin deficient cells by nucleotide analogue incorporation and found that Aprataxin deficiency in fact does not cause a deficit in basal synthesis rate. This is consistient with the marginal defect in 47S and 45S rRNA levels in untreated AOA1 cells (1).

Given the critical role of UBF in RNA polymerase I transcription initiation I was surprised that Aprataxin deficient cells, which do not have full length UBF, display normal basal levels of RNA synthesis. Here I have proposed a model of how the proteolysed UBF HMG box and C-terminal acidic tail regions may retain transactivation activity. UBF forms a dimer at the rDNA promoter (59,60). The N- terminal region of each protein interacts to form a complex containing 10 HMG boxes, six of which are in contact with the promoter, with the acidic tail regions protruding from both sides of the dimer (59). As discussed previously, the acidic tail of UBF is essential for its interaction with the SL1 complex and this interaction is central to the formation of the pre-initiation complex. Given that cells containing only HMG box and acidic tail proteolysed UBF fragments are capable of rRNA synthesis, these domains must be able to interact stably and independently of the peptide

265 backbone. I propose that the proteolysed UBF fragments interact to adopt a similar conformation as the whole protein and thus remain capable of facilitating formation of pre-initiation complexes (Figure 5.33).

Figure 5.33: Model for UBF dimerization and DNA binding. In the top panel, full length UBF forms a dimer over the rDNA promoter. In the absence of Aprataxin UBF is cleaved into acidic tail and HMG box regions, as shown in the lower panel. Proteolysed UBF may be able to adopt a similar conformation on DNA to uncleaved UBF, allowing Aprataxin deficient cells to competently initiate rRNA transcription.

To investigate the functional interaction between Aprataxin, DNA damage and RNA transcription, I examined the effect of Aprataxin deficiency on transcription after DNA damage. DNA damage results in a complex set of cellular responses, including the inhibition of rRNA transcription. Irradiation of cells results in sequestration of RNA polymerase I and UBF to nucleolar caps (41), reminiscent of the staining patterns observed after treatment of cells with the RNA polymerase I inhibitor Actinomycin D (1,55,61). Inhibition of rRNA synthesis and relocalization of RNA polymerase I to nucleolar caps are not dissociated effects. The inhibition of rRNA synthesis after ionizing radiation treatment is an indirect effect of DNA damage, dependant on activity of the DNA damage signalling kinase ATM (41). DNA damage in transcribed regions has also been shown to have direct consequences on RNA synthesis. In vitro transcription studies revealed that T4 RNA polymerase can synthesize across a single nucleotide gap provided it possesses a 3’ phosphate (52). The resulting transcripts are frameshifted due to a single nucleotide deletion which

266 corresponds to the missing DNA nucleotide. Abasic sites in transcribed regions result in generation of point mutated or prematurely truncated transcripts (62). Cisplatin- induced DNA adducts inhibit RNA synthesis by sequestering UBF, which has a very high affinity for four-way junction DNA and crosslinks (63). Thus three mechanisms appear to contribute to inhibition of RNA synthesis after DNA damage, an ATM- dependant signalling pathway, direct inhibition of synthesis by lesions in coding regions, and sequestering of UBF by a subset of DNA lesions.

We therefore assessed the Aprataxin-dependant transcriptional effects of hydrogen peroxide and ionizing radiation (section 5.3.3). Irradiation of corrected AOA1 cells resulted in a significant reduction in their rate of nascent RNA synthesis (31.2% reduction, p=0.0002), whereas irradiation of Aprataxin deficient cells did not (10.2% reduction, p=0.2). This indicates that, like A-T cell lines, Aprataxin deficient cells display RRS. Clements et al. found that AOA1 cells do not have a defect in ATM activation (21), although this may need to be validated on a more extensive panel of downstream proteins. This indicates that this radioresistant RNA synthesis is probably not due to a defect in ATM activation, but could instead be due to an Aprataxin- dependant defect in some downstream part of the ATM signalling pathway. For example, while the degraded UBF protein present in Aprataxin deficient cells has normal transactivation activity, it may be a poor substrate for its regulating kinases (such as CK2). Thus Aprataxin deficient cells are able to synthesize RNA under normal conditions but after irradiation are unable to inhibit synthesis, which may result in the production of aberrant transcripts.

Conversely, treatment of corrected AOA1 cells with the oxidative stress inducing agent hydrogen peroxide did not cause a significant inhibition of nascent RNA synthesis, while Aprataxin deficient cells displayed a remarkable level of inhibition. The doses used were rather high (0.5 and 1 mM for 10 minutes) and I propose that this level of oxidative stress may have caused extensive DNA damage in transcribed regions, resulting in direct inhibition of nascent synthesis. In Chapter 4 I characterised an elevated level of oxidative stress and multiple DNA repair deficiencies in AOA1 cells. The reduced efficiency of repair of oxidative lesions may lead to extremely

267 sensitive cellular responses to oxidative stress inducing agents in Aprataxin deficient cells.

5.4.4 Summary:

In this chapter I have characterised the interaction between Aprataxin and several proteins involved in RNA metabolism. These interactions occur primarily through the FHA and nuclear localization regions of Aprataxin but some HIT domain involvement was also observed. I further characterised the interaction between Aprataxin and one of these proteins, UBF. I demonstrated that Aprataxin is necessary for stabilization of UBF and that Aprataxin deficient cells display abnormal UBF localization. Given the critical role of UBF in transcription I analysed the transcription activity of Aprataxin deficient cells and found that they do not display a defect in their basal levels of RNA synthesis. Based on this I propose that the partially degraded UBF present in AOA1 cells retains the ability dimerize and form pre- initiation complexes, and is thus able to facilitate normal rRNA transcription initiation.

We described two different transcriptional defects in Aprataxin deficient cells in response to induced DNA damage. Irradiation of control cells induces ATM- dependant inhibition of transcription. Reminiscent of ATM deficient cells, AOA1 cells display RRS, despite that they do not have an ATM activation defect (21). This implies that AOA1 cells have a signalling defect downstream of ATM. Conversely, hydrogen peroxide treatment inhibited transcription in Aprataxin deficient cells indicating that AOA1 cells have a hypersensitive transcriptional response to oxidative stress. Taken together this data adds another layer of complexity to the role of Aprataxin in the cell and suggests a role for Aprataxin in transcription and the stabilization of transcription factors.

268 5.5 REFERENCES

1. Becherel, O.J., Gueven, N., Birrell, G.W., Schreiber, V., Suraweera, A., Jakob, B., Taucher-Scholz, G. and Lavin, M.F. (2006) Nucleolar localization of aprataxin is dependent on interaction with nucleolin and on active ribosomal DNA transcription. Hum Mol Genet, 15, 2239-2249.

2. Gueven, N., Becherel, O.J., Kijas, A.W., Chen, P., Howe, O., Rudolph, J.H., Gatti, R., Date, H., Onodera, O., Taucher-Scholz, G. et al. (2004) Aprataxin, a novel protein that protects against genotoxic stress. Hum Mol Genet, 13, 1081- 1093.

3. Shaw, P.J. and Jordan, E.G. (1995) The nucleolus. Annu Rev Cell Dev Biol, 11, 93-121.

4. Dundr, M. and Misteli, T. (2001) Functional architecture in the cell nucleus. Biochem J, 356, 297-310.

5. Cheutin, T., O'Donohue, M.F., Beorchia, A., Vandelaer, M., Kaplan, H., Defever, B., Ploton, D. and Thiry, M. (2002) Three-dimensional organization of active rRNA genes within the nucleolus. J Cell Sci, 115, 3297-3307.

6. Bertrand, E., Houser-Scott, F., Kendall, A., Singer, R.H. and Engelke, D.R. (1998) Nucleolar localization of early tRNA processing. Genes Dev, 12, 2463- 2468.

7. Weber, J.D., Taylor, L.J., Roussel, M.F., Sherr, C.J. and Bar-Sagi, D. (1999) Nucleolar Arf sequesters Mdm2 and activates p53. Nat Cell Biol, 1, 20-26.

8. Dynan, W.S. and Tjian, R. (1985) Control of eukaryotic messenger RNA synthesis by sequence-specific DNA-binding proteins. Nature, 316, 774-778.

9. Learned, R.M., Learned, T.K., Haltiner, M.M. and Tjian, R.T. (1986) Human rRNA transcription is modulated by the coordinate binding of two factors to an upstream control element. Cell, 45, 847-857.

269 10. Kuhn, A., Voit, R., Stefanovsky, V., Evers, R., Bianchi, M. and Grummt, I. (1994) Functional differences between the two splice variants of the nucleolar transcription factor UBF: the second HMG box determines specificity of DNA binding and transcriptional activity. The EMBO Journal, 13, 416-424.

11. Moss, T. (2004) At the crossroads of growth control; making ribosomal RNA. Current Opinion in Genetics & Development, 14, 210-217.

12. Grummt, I., Rosenbauer, H., Niedermeyer, I., Maier, U. and Ohrlein, A. (1986) A repeated 18 bp sequence motif in the mouse rDNA spacer mediates binding of a nuclear factor and transcription termination. Cell, 45, 837-846.

13. Sander, E.E., Mason, S.W., Munz, C. and Grummt, I. (1996) The amino- terminal domain of the transcription termination factor TTF-I causes protein oligomerization and inhibition of DNA binding. Nucleic Acids Res, 24, 3677- 3684.

14. Heix, J., Vente, A., Voit, R., Budde, A., Michaelidis, T.M. and Grummt, I. (1998) Mitotic silencing of human rRNA synthesis: inactivation of the promoter selectivity factor SL1 by cdc2/cyclin B-mediated phosphorylation. The EMBO Journal, 17, 7373-7381.

15. Kuhn, A., Vente, A., Doree, M. and Grummt, I. (1998) Mitotic phosphorylation of the TBP-containing factor SL1 represses ribosomal gene transcription. J Mol Biol, 284, 1-5.

16. Hemmerich, P. and Diekmann, S. (2005) Visions of the cell nucleus, ISBN 1588830276, American Scientific Publishers.

17. Birrell, G.W., Earl, S., Masci, P.P., de Jersey, J., Wallis, T.P., Gorman, J.J. and Lavin, M.F. (2006) Molecular diversity in venom from the Australian Brown snake, Pseudonaja textilis. Mol Cell Proteomics, 5, 379-389.

18. Birrell, G.W., Earl, S.T., Wallis, T.P., Masci, P.P., de Jersey, J., Gorman, J.J. and Lavin, M.F. (2007) The diversity of bioactive proteins in Australian snake venoms. Mol Cell Proteomics, 6, 973-986.

270 19. Kijas, A.W., Harris, J.L., Harris, J.M. and Lavin, M.F. (2006) Aprataxin forms a discrete branch in the HIT (histidine triad) superfamily of proteins with both DNA/RNA binding and nucleotide hydrolase activities. J Biol Chem, 281, 13939-13948.

20. Rass, U., Ahel, I. and West, S.C. (2007) Actions of aprataxin in multiple DNA repair pathways. J Biol Chem, 282, 9469-9474.

21. Clements, P.M., Breslin, C., Deeks, E.D., Byrd, P.J., Ju, L., Bieganowski, P., Brenner, C., Moreira, M.C., Taylor, A.M. and Caldecott, K.W. (2004) The ataxia-oculomotor apraxia 1 gene product has a role distinct from ATM and interacts with the DNA strand break repair proteins XRCC1 and XRCC4. DNA Repair (Amst), 3, 1493-1502.

22. Luo, H., Chan, D.W., Yang, T., Rodriguez, M., Chen, B.P., Leng, M., Mu, J.J., Chen, D., Songyang, Z., Wang, Y. et al. (2004) A new XRCC1- containing complex and its role in cellular survival of methyl methanesulfonate treatment. Mol Cell Biol, 24, 8356-8365.

23. El-Khamisy, S.F., Masutani, M., Suzuki, H. and Caldecott, K.W. (2003) A requirement for PARP-1 for the assembly or stability of XRCC1 nuclear foci at sites of oxidative DNA damage. Nucleic Acids Research, 31, 5526-5533.

24. Patton, J.G., Porro, E.B., Galceran, J., Tempst, P. and Nadal-Ginard, B. (1993) Cloning and characterization of PSF, a novel pre-mRNA splicing factor. Genes Dev, 7, 393-406.

25. Lamond, A.I. (1993) The spliceosome. Bioessays, 15, 595-603.

26. Lamm, G.M. and Lamond, A.I. (1993) Non-snRNP protein splicing factors. Biochim Biophys Acta, 1173, 247-265.

27. Gozani, O., Patton, J.G. and Reed, R. (1994) A novel set of spliceosome- associated proteins and the essential splicing factor PSF bind stably to pre- mRNA prior to catalytic step II of the splicing reaction. The EMBO Journal, 13, 3356-3367.

271 28. Kaneko, S., Rozenblatt-Rosen, O., Meyerson, M. and Manley, J.L. (2007) The multifunctional protein p54nrb/PSF recruits the exonuclease XRN2 to facilitate pre-mRNA 3' processing and transcription termination. Genes Dev, 21, 1779-1789.

29. Emili, A., Shales, M., McCracken, S., Xie, W., Tucker, P.W., Kobayashi, R., Blencowe, B.J. and Ingles, C.J. (2002) Splicing and transcription-associated proteins PSF and p54nrb/nonO bind to the RNA polymerase II CTD. RNA, 8, 1102-1111.

30. O'Mahony, D.J. and Rothblum, L.I. (1991) Identification of two forms of the RNA polymerase I transcription factor UBF. Proc Natl Acad Sci U S A, 88, 3180-3184.

31. Grueneberg, D.A., Pablo, L., Hu, K.Q., August, P., Weng, Z. and Papkoff, J. (2003) A functional screen in human cells identifies UBF2 as an RNA polymerase II transcription factor that enhances the beta-catenin signaling pathway. Mol Cell Biol, 23, 3936-3950.

32. Tyc, K. and Steitz, J.A. (1989) U3, U8 and U13 comprise a new class of mammalian snRNPs localized in the cell nucleolus. The EMBO Journal, 8, 3113-3119.

33. Newton, K., Petfalski, E., Tollervey, D. and Caceres, J.F. (2003) Fibrillarin is essential for early development and required for accumulation of an intron- encoded small nucleolar RNA in the mouse. Mol Cell Biol, 23, 8519-8527.

34. Tollervey, D., Lehtonen, H., Jansen, R., Kern, H. and Hurt, E.C. (1993) Temperature-sensitive mutations demonstrate roles for yeast fibrillarin in pre- rRNA processing, pre-rRNA methylation, and ribosome assembly. Cell, 72, 443-457.

35. Tollervey, D., Lehtonen, H., Carmo-Fonseca, M. and Hurt, E.C. (1991) The small nucleolar RNP protein NOP1 (fibrillarin) is required for pre-rRNA processing in yeast. The EMBO Journal, 10, 573-583.

36. Dreyfuss, G., Matunis, M.J., Pinol-Roma, S. and Burd, C.G. (1993) hnRNP proteins and the biogenesis of mRNA. Annu Rev Biochem, 62, 289-321.

272 37. Kukalev, A., Nord, Y., Palmberg, C., Bergman, T. and Percipalle, P. (2005) Actin and hnRNP U cooperate for productive transcription by RNA polymerase II. Nat Struct Mol Biol, 12, 238-244.

38. Durocher, D. and Jackson, S.P. (2002) The FHA domain. FEBS letters, 513, 58-66.

39. Voit, R., Kuhn, A., Sander, E.E. and Grummt, I. (1995) Activation of mammalian ribosomal gene transcription requires phosphorylation of the nucleolar transcription factor UBF. Nucleic Acids Res, 23, 2593-2599.

40. Voit, R., Schnapp, A., Kuhn, A., Rosenbauer, H., Hirschmann, P., Stunnenberg, H.G. and Grummt, I. (1992) The nucleolar transcription factor mUBF is phosphorylated by casein kinase II in the C-terminal hyperacidic tail which is essential for transactivation. The EMBO Journal, 11, 2211-2218.

41. Kruhlak, M., Crouch, E.E., Orlov, M., Montano, C., Gorski, S.A., Nussenzweig, A., Misteli, T., Phair, R.D. and Casellas, R. (2007) The ATM repair pathway inhibits RNA polymerase I transcription in response to chromosome breaks. Nature, 447, 730-734.

42. Stefanovsky, V.Y., Pelletier, G., Hannan, R., Gagnon-Kugler, T., Rothblum, L.I. and Moss, T. (2001) An immediate response of ribosomal transcription to growth factor stimulation in mammals is mediated by ERK phosphorylation of UBF. Mol Cell, 8, 1063-1073.

43. Brooks, P.J., Wise, D.S., Berry, D.A., Kosmoski, J.V., Smerdon, M.J., Somers, R.L., Mackie, H., Spoonde, A.Y., Ackerman, E.J., Coleman, K. et al. (2000) The oxidative DNA lesion 8,5'-(S)-cyclo-2'-deoxyadenosine is repaired by the nucleotide excision repair pathway and blocks gene expression in mammalian cells. J Biol Chem, 275, 22355-22362.

44. Treiber, D.K., Zhai, X., Jantzen, H.M. and Essigmann, J.M. (1994) Cisplatin- DNA adducts are molecular decoys for the ribosomal RNA transcription factor hUBF (human upstream binding factor). Proc Natl Acad Sci U S A, 91, 5672- 5676.

273 45. Wansink, D.G., Schul, W., van der Kraan, I., van Steensel, B., van Driel, R. and de Jong, L. (1993) Fluorescent labeling of nascent RNA reveals transcription by RNA polymerase II in domains scattered throughout the nucleus. J Cell Biol, 122, 283-293.

46. Wojtowicz, J.M. and Kee, N. (2006) BrdU assay for neurogenesis in rodents. Nat Protoc, 1, 1399-1405.

47. Koberna, K., Stanek, D., Malinsky, J., Ctrnacta, V., Cermanova, S., Novotna, J., Kopsky, V. and Raska, I. (2000) In situ fluorescence visualization of bromouridine incorporated into newly transcribed nucleolar RNA. Acta Histochem, 102, 15-20.

48. Scheer, U. and Hock, R. (1999) Structure and function of the nucleolus. Curr Opin Cell Biol, 11, 385-390.

49. Baserga, R., Estensen, R.D. and Petersen, R.O. (1965) Inhibition of DNA synthesis in Ehrlich ascites cells by actinomycin D. II. The presynthetic block in the cell cycle. Proc Natl Acad Sci U S A, 54, 1141-1148.

50. Lieberman, I., Abrams, R. and Ove, P. (1963) Changes in the metabolism of ribonucleic acid preceding the synthesis of deoxyribonucleic acid in mammalian cells cultured from the animal. J Biol Chem, 238, 2141-2149.

51. Liu, J. and Doetsch, P.W. (1996) Template strand gap bypass is a general property of prokaryotic RNA polymerases: implications for elongation mechanisms. Biochemistry, 35, 14999-15008.

52. Zhou, W. and Doetsch, P.W. (1994) Transcription bypass or blockage at single-strand breaks on the DNA template strand: effect of different 3' and 5' flanking groups on the T7 RNA polymerase elongation complex. Biochemistry, 33, 14926-14934.

53. Zhou, W., Reines, D. and Doetsch, P.W. (1995) T7 RNA polymerase bypass of large gaps on the template strand reveals a critical role of the nontemplate strand in elongation. Cell, 82, 577-585.

274 54. Svejstrup, J.Q. (2002) Mechanisms of transcription-coupled DNA repair. Nat Rev Mol Cell Biol, 3, 21-29.

55. Smetana, K., Busch, R., Chan, P.K., Smetana, K., Jr. and Busch, H. (2001) Immunocytochemical localization of nucleophosmin and RH-II/Gu protein in nucleoli of HeLa cells after treatment with actinomycin D. Acta Histochem, 103, 325-333.

56. Shav-Tal, Y., Blechman, J., Darzacq, X., Montagna, C., Dye, B.T., Patton, J.G., Singer, R.H. and Zipori, D. (2005) Dynamic sorting of nuclear components into distinct nucleolar caps during transcriptional inhibition. Mol Biol Cell, 16, 2395-2413.

57. Voit, R. and Grummt, I. (2001) Phosphorylation of UBF at serine 388 is required for interaction with RNA polymerase I and activation of rDNA transcription. Proc Natl Acad Sci U S A, 98, 13631-13636.

58. Friedrich, J.K., Panov, K.I., Cabart, P., Russell, J. and Zomerdijk, J.C. (2005) TBP-TAF complex SL1 directs RNA polymerase I pre-initiation complex formation and stabilizes upstream binding factor at the rDNA promoter. J Biol Chem, 280, 29551-29558.

59. Stefanovsky, V.Y., Pelletier, G., Bazett-Jones, D.P., Crane-Robinson, C. and Moss, T. (2001) DNA looping in the RNA polymerase I enhancesome is the result of non-cooperative in-phase bending by two UBF molecules. Nucleic Acids Res, 29, 3241-3247.

60. Wolffe, A.P. (1994) Architectural transcription factors. Science, 264, 1100- 1101.

61. Puvion-Dutilleul, F., Mazan, S., Nicoloso, M., Pichard, E., Bachellerie, J.P. and Puvion, E. (1992) Alterations of nucleolar ultrastructure and ribosome biogenesis by actinomycin D. Implications for U3 snRNP function. Eur J Cell Biol, 58, 149-162.

62. Sanchez, G. and Mamet-Bratley, M.D. (1994) Transcription by T7 RNA polymerase of DNA containing abasic sites. Environmental and Molecular Mutagenesis, 23, 32-36.

275 63. Zhai, X., Beckmann, H., Jantzen, H.M. and Essigmann, J.M. (1998) Cisplatin- DNA adducts inhibit ribosomal RNA synthesis by hijacking the transcription factor human upstream binding factor. Biochemistry, 37, 16307-16315.

276

CHAPTER 6

General discussion

277 GENERAL DISCUSSION

Autosomal recessive cerebellar ataxias are a heterogeneous family of disorders which include a subset of ataxias caused by mutation of DNA damage signalling and repair proteins. These include the well-characterised disorders A-T, A-TLD, XP, and the more recently identified disorders AOA1 and 2 and SCAN1 (1-6). AOA1 is characterised by early-onset spinocerebellar ataxia, oculomotor apraxia, peripheral sensory and motor neuropathy as well as a range of other variable symptoms (5,7-9). The gene mutated in AOA1, APTX codes for the 342 amino acid protein Aprataxin (4,5) which has three predicted functional domains (FHA, HIT and zinc finger, reference (5). During the course of this study I and others have provided insight into the function of each of these domains and the protein as a whole.

6.1 GENERATION OF REAGENTS

A common approach to characterise a novel protein and determine the function of its domains involves generation of a recombinant form of the protein. As such the initial phase in the characterisation of the various domains of Aprataxin involved producing wild-type, point and truncation mutant recombinant Aprataxin proteins, using two expression systems. Wild-type, as well as C- and N-terminal truncations of Aprataxin were purified to homogeneity using a bacterial expression system (2.64, 5.15 and 1.76 mg respectively, using pGEX 6.1). Purification of a doubly truncated protein was unsuccessful due to the poor expression, binding and elution of this protein even though it appeared to be predominantly soluble. I also generated wild-type Aprataxin using another bacterial expression system, pTYB1, although the yield was substantially less than the equivalent protein expressed in pGEX (625 µg versus 2.64 mg). This study was unable to purify a HIT domain Aprataxin mutant (V263G) to homogeneity using pTYB1, due to its apparent misfolding and aggregation and its subsequent strong interaction with heat-shock proteins. The aggregation of mutant proteins is a common phenomenon in recombinant bacterial expression systems (10). After considerable optimization, enrichment of the V263G protein produced approximately 25 µg of protein containing 75% contamination. This very low yield is

278 consistent with the observation that even point mutations in Aprataxin generally render the protein unstable (11). Indeed I demonstrated in Chapter 4 that mammalian cells with this mutation do not have detectable Aprataxin (L939 is homozygous V263G).

To study Aprataxin in vivo, I also generated a set of affinity purified polyclonal Aprataxin antibodies from rabbit and sheep. The rabbit antibody was raised against the whole protein-GST fusion, while the sheep antibodies were raised against FHA- GST and HIT-ZnF-GST proteins. These antibodies were affinity purified against the FHA and/or HIT domains to yield a rabbit antibody which does not block the zinc finger and sheep antibodies against the FHA domain or the FHA and HIT domains respectively. Both the rabbit and the sheep α-FHA antibodies were useful for immunostaining and immunoflourescence, while the rabbit antibody was useful for a broad range of applications including Western blotting, antibody supershifts and inhibition of the HIT domain. Generation of these purified antibodies proved to be crucial for downstream characterisation of the in vitro and in vivo functions of Aprataxin.

6.2 BIOCHEMICAL CHARACTERISATION OF APRATAXIN

6.2.1 Aprataxin binds adenosine derivatives:

At the commencement of this study little was known about the activities of Aprataxin. Sequence analysis determined that Aprataxin contains a central HIT-type nucleotide hydrolase domain and early studies demonstrated that Aprataxin has low levels of nucleotide hydrolase activity on a range of guanosine and adenosine derivatives (12,13). The reported activities were very low compared to those displayed by other HIT proteins on similar substrates (14-17) and as such I was concerned that these reported activities could be due to low level contamination of the authors protein preparations. To confirm the specificity of these reported reactions I examined wether Aprataxin undergoes a change in conformation in the presence of nucleotide derivatives. To do this I employed partial proteolysis, which is a means to examine which protease sites are exposed to the soluble environment and accessible to the

279 chosen protease (18,19). Alterations in conformation of the protein of interest change which sites are accessible to the protease and are therefore detectable as changes in protein banding pattern by SDS-PAGE. I detected similar changes in Aprataxin proteolysis patterns in the presence of the reported substrate adenosine monophosphoramidate and the product of HIT domain activity, AMP, suggesting that adenosine monophosphoramidate and AMP induce similar conformational changes in Aprataxin. This is consistent with the proposal that Aprataxin is an adenosine- derivative hydrolase which produces AMP.

6.2.2 Aprataxin hydrolyses adenosine derivatives:

We confirmed this by examining the hydrolase activity of recombinant Aprataxin on the nucleotide analogues diadenosine tetraphosphate and adenosine monophosphoramidate. Hydrolysis of diadenosine tetraphosphate was performed with a maximum theoretical rate 0.0194/sec with a Km of 3.87 µM, and hydrolysis of adenosine monophosphate was performed faster (0.069/sec) but with a higher Km (816 µM). The kinetic parameters reported in this thesis are similar to those presented by Kijas et al. on the same substrates but using protein produced from different expression systems (20). This provided additional validation of the activity of Aprataxin reported by Kijas et al. (20). However activities obtained by different laboratories on similar or identical substrates appear highly variable. For example Seidle et al. also found that Aprataxin hydrolyses diadenosine tetraphosphate (Vmax=

0.0009/sec, Km = 39 µM), but much less efficiently than reported here and in Kijas et al. (12,20). This variability makes comparisons between different kinetic studies difficult. At this point the activity of Aprataxin’s zinc finger had not been determined and so initially I employed similar techniques (partial proteolysis and nucleotide hydrolysis) to explore the properties of this domain.

6.2.3 The HIT domain interacts with DNA:

We examined the ability of DNA to induce conformational changes in Aprataxin and found that both single and double stranded DNA induce conformational changes. The proteolysis patterns of Aprataxin with single or double stranded DNA are similar,

280 with single stranded DNA inducing a less marked conformational change than double stranded DNA. This indicates that Aprataxin can bind to both single and double stranded DNA, with a lower affinity for single stranded structures. This was confirmed by EMSA, where I observed a stable interaction between Aprataxin and double stranded DNA and a transient interaction with single stranded DNA. Interestingly I also noted that the HIT domain mutant V263G is unable to bind DNA, suggesting a crucial role for the HIT domain in the Aprataxin-DNA interaction.

Given that the HIT domain of Aprataxin hydrolyses nucleotide derivatives and appears important it in DNA binding, I proposed that Aprataxin binds to DNA through the HIT domain. This idea was explored further by examining the functional interaction between the hydrolase and DNA binding activities of Aprataxin. I found that double stranded DNA is a mixed (combined competitive and non-competitive) inhibitor of nucleotide hydrolysis, with the major component being competitive inhibition, indicating that DNA binds directly to the HIT domain of Aprataxin. This, combined with the interaction of Aprataxin with DNA repair enzymes such as XRCC1 and PARP-1 and the DNA damage sensitivity of AOA1 cells clearly implicated the HIT domain in DNA processing. Based on the hypersensitivity of AOA1 cells to the Topoisomerase I inhibitor camptothecin I proposed that Aprataxin is a repair factor which processes DNA-3’ Topoisomerase I covalent structures (20,21).

6.2.4 Aprataxin is a DNA-end processing factor:

Elucidation of the role of the HIT domain of Aprataxin in DNA binding and nucleotide hydrolysis was provided by Ahel et al. in 2006, when they identified DNA repair related substrate for Aprataxin (5’ adenylated DNA, reference 22). Although as previously proposed Aprataxin was found to be a DNA end processing enzyme, it acts on a 5’ structure rather than the 3’ modification we had suggested. 5’ adenylated DNA is generated as a reaction intermediate during DNA ligation. If a DNA ligation reaction is initiated but stalls part way through catalysis, this intermediate structure could accumulate. Given that DNA ligation is a crucial step in DNA repair, cells have

281 the potential to generate 5’ adenylated DNA in response to a variety of genotoxic chemicals. This was substantiated by Ahel et al. who demonstrated that 5’ adenylated DNA accumulates in vitro when ligation is attempted with a damaged substrate (3’ deoxy or hydrogen peroxide induced damage in their case). They found that Aprataxin hydrolyses 5’ adenylated DNA to restore the 5’ phosphate terminus, and experiments presented here substantiate these findings. Additionally, in the present study I demonstrated that 5’ adenylated DNA can accumulate during ligation of DNA single strand breaks with oxidized or abasic 3’ termini. Both endogenously generated reactive oxygen species and exogenously applied DNA damaging agents (such as camptothecin) can produce 3’ terminally damaged breaks (23,24). Our findings and those of Ahel et al. indicate that 5’ DNA adenylates can be generated after failed ligation attempts at a range of DNA structures (22). This provides a link between the hypersensitivity of Aprataxin-deficient cells to DNA damaging agents such as hydrogen peroxide (which produces 3’ modified breaks directly and indirectly) and camptothecin (which produce 3’ modified breaks directly), the inhibition of DNA ligation, the accumulation of 5’ adenylates at 3’ modified breaks, and the in vitro activity of Aprataxin.

6.2.5 The HIT domain is regulated by the zinc finger and FHA domains:

By analysing the activity of full length and truncated recombinant Aprataxin proteins against adenylated DNA, I determined the contribution of the zinc finger and FHA domains to the activity of the HIT domain. I was unable to detect hydrolase activity in a protein lacking the zinc finger. This was supported by the findings of Rass et al. who found that an Aprataxin zinc finger mutant has 1% the hydrolase activity of the wild-type protein (25). This indicates that although the zinc finger may not participate in the hydrolysis reaction per se, it may be necessary for stabilization of the enzyme- substrate complex or orientation of the substrate with respect to the active site. This is similar to the situation with PARP-1, which cannot be activated in if its DNA binding domain is deleted even though the catalytic domain is in tact. Thus the zinc finger, which binds single and double stranded DNA in a sequence non-specific manner (20,25), is critical for DNA-adenylate hydrolysis.

282 In addition, I found that the FHA domain inhibits the HIT domains DNA-adenylate hydrolase activity. Indeed, deletion of the FHA domain stimulated the DNA-adenylate hydrolase activity of Aprataxin’s HIT domain approximately two-fold. This effect is consistent with but less dramatic than the stimulation of activity by FHA domain deletion reported by Hirano et al. (13). Their study failed to detect GpppBODIPY hydrolase activity in full length protein but observed limited substrate turnover by Aprataxin lacking the FHA domain. Our finding did not support the previous report by Kijas et al. who, using protein expressed and purified in the same system as in this thesis, found that Aprataxin lacking the FHA domain had very similar kinetic parameters to the full length protein (20). The difference between the stimulation of hydrolase activity reported here and the lack stimulation reported by Kijas et al. was unexpected given that the proteins used were prepared in a similar manner from the same constructs. Thus I propose that the differences in the stimulatory effects reported by these three studies are due to the use of different substrates (GpppBODIPY and AppppA by Hirano et al. and Kijas et al. respectively, and 5’ adenylated DNA in this study, see references (13,20). As no other study has examined the effect of deletion of the FHA domain on Aprataxin’s hydrolase activity using the biologically relevant substrate, 5’ adenylated DNA, I maintain that the FHA domain has a mild inhibitory effect on the HIT domain. Furthermore I propose that the activity of the HIT domain may be regulated protein-protein interactions via the FHA domain in vivo. Aprataxin has been found to interact with a number of proteins via its FHA domain, including XRCC1, XRCC1, PARP-1, UBF, nucleolin and hnRNP U. The FHA domain of Aprataxin may adopt different conformations when interacting with these different proteins, which could in turn modulate the activity of the HIT domain.

6.2.6 Aprataxin activity in cells is regulated by PARP-1:

Aprataxin interacts with the DNA repair proteins PARP-1 and XRCC1. These interactions occur through the FHA domain of Aprataxin with the BRCT domains 1 and 2 of XRCC1 (binding to these two domains occurs independently) and the single BRCT domain of PARP-1 (26,27 and Dr Olivier Becherel, Queensland Institute for Medical Research, Brisbane, Australia, unpublished observations). Given that deletion of the FHA domain stimulates the activity of the HIT domain of recombinant

283 Aprataxin, I proposed that HIT domain activity may be regulated by PARP-1 and XRCC1 in vivo. Deficiency of XRCC1 did not affect the abundance or activity of Aprataxin in cell extracts. This is not in agreement with other studies which found that transient depletion of XRCC1 by siRNA results in destabilization of Aprataxin (26,27). The XRCC1 mutant cell line EM9 has a point mutation which causes premature translation termination of the protein, however low levels of truncated protein are present in these cells (28). This may be enough to stabilize Aprataxin. Also, EM9 cells have chronic XRCC1 deficiency, and recovery of normal Aprataxin levels over many passages probably provides a growth advantage. Thus even if XRCC1 stabilizes Aprataxin, Aprataxin levels may recover after many generations of XRCC1 deficiency.

On the other hand deficiency of PARP-1 (but not PARP activity) had a profound impact on DNA-adenylate hydrolysis. Hydrolysis of 5’ adenylated DNA was extremely slow in PARP-1 knockout cell extracts. Further investigation of the Aprataxin activity deficit in PARP-1 knockout cells revealed that they do not have detectable levels of Aprataxin. This was substantiated by transient depletion of PARP- 1 using siRNA, which caused a reduction in Aprataxin levels. This indicates that PARP-1 is necessary for stabilization of Aprataxin while XRCC1 may not be. This is not an uncommon phenomenon, for example mutation and subsequent destabilization of Mre11 or Nbs1 results in destabilization of the other proteins in the MRN complex, whereas mutation of the third complex member, Rad50 does not affect levels of Mre11 or Nbs1. This destabilization in the absence of PARP-1 but not XRCC1 also supports the idea that Aprataxin is present in a range of DNA repair complexes. This would be analogous to the situation with the FHA domain containing general repair factor PNKP, which is present in a range of repair complexes (29,30) and is consistent with the role of Aprataxin as an accessory factor in DNA ligation.

284 6.3 CHARACTERISATION OF THE DEFECTS IN AOA1 CELLS

6.3.1 End processing:

The aim of the second phase of this study was to characterise the biochemical and molecular defects caused by Aprataxin deficiency, which are expected to be causative of the neurodegenerative disorder AOA1. Initially this involved examination of the DNA-adenylate hydrolase activity of AOA1 cells. This study, together with Ahel et al. revealed that AOA1 cell extracts are deficient in DNA-adenylate hydrolase capacity, indicating that Aprataxin is the only protein in mammalian cells able to repair this modification (22). This indicated a unique, non-redundant function for Aprataxin in DNA repair.

Aprataxin is a nuclear protein with nucleoplasmic and nucleolar localization, in agreement with its zinc finger and nuclear localization sequences (31). Aprataxin’s nucleolar localization is dependant on its interaction with the rRNA processing factor nucleolin and active rRNA synthesis (32). Given the distribution of Aprataxin in both nuclear sub-compartments, I examined the DNA-adenylate hydrolase activity of nucleoplasmic and nucleolar fractions from control cells. I found that nucleolar extract has a higher specific activity than nucleoplasmic extract despite a roughly equal concentration of Aprataxin in these fractions. This indicates that this disparity in DNA-adenylate hydrolase activity is not due to differential distribution of Aprataxin protein. I noted that the distribution of Aprataxin’s interacting protein PARP-1 is uneven between these compartments, with a much higher concentration in the nucleoplasm. Based on the elevated level of hydrolase activity demonstrated by an N- terminal truncation of Aprataxin, I proposed that the activity of the HIT domain could be regulated by the FHA domain. Thus interaction with different proteins in both sub- nuclear compartments could fine-tune Aprataxin’s activity. I also found in vitro phosphorylation of Aprataxin’s HIT domain by the DNA damage signalling kinase ATR. This phosphorylation may also modulate the activity of Aprataxin directly or by inducing formation of different protein complexes.

285 Takahashi et al. recently described an additional role for Aprataxin as a 3’ processing factor (33). They reported that Aprataxin can process damaged 3’ termini (phosphate and phosphoglycolate) which are often present at direct single strand breaks (33,34). The rates of hydrolysis of 3’ phosphate and phosphoglycolate molecules by Aprataxin reported by Takahashi et al. were low compared to rates reported by other authors for the well-established processing factors PNKP and APE1 (33,35,36). Additionally, I was unable to detect 3’ phosphatase activity in recombinant Aprataxin, even after incubation of the substrate with a 5-fold molar excess of enzyme for over an hour. To examine the contribution of Aprataxin to repair of 3’ phosphate, I measured the phosphatase activities of control and AOA1 lymphoblastiod cell extracts. I found that deficiency of Aprataxin did not confer a defect in phosphatase activity, therefore Aprataxin does not make a major contribution to 3’ phosphate hydrolysis in cells. The phosphoglycohydrolase activity of Aprataxin was not examined here due to the involved process required to generate this substrate (optimization of in vitro treatment of a duplex with bleomycin, identification of the desired reaction product, purification by denaturing PAGE, desalting by chromatography, and finally annealing to form the desired duplex, described in reference (36) and communications with other researchers (Prof Keith Caldecott, University of Sussex, Brighton, UK and Prof Steven West, Cancer Research UK, London Research Institute, UK) who failed to detect activity of Aprataxin against 3’ phosphoglycolate. This indicates that Aprataxin probably does not make a major contribution to repair of these lesions in vivo. I conclude that the low level phosphatase and phosphoglycohydrolase activities reported by Takahashi et al. are due to contamination of their Aprataxin protein preparation or reagents.

In summary the primary enzymatic function of Aprataxin appears to be repair of 5’ adenylated DNA termini. We and others have been unable to replicate the findings of Takahashi et al. which implicate a role for Aprataxin in processing of 3’ termini (33).

286 6.3.2 Base Excision Repair:

Elevated levels of oxidative DNA damage are a hallmark of many neurodegenerative disorders, including some DNA repair-defective ataxias (A-T, AOA2, references 37,38). 8-oxo(d)G is one of the major products of oxidative DNA damage and is a commonly used marker of oxidative stress. Using immunostaining of brain samples from deceased AOA1 patients, Hirano et al. reported that the nuclei in AOA1 patient cerebella have elevated levels 8-oxo-(d)G (39). This staining was RNase resistant indicating that AOA1 patients have elevated levels of 8-oxo-dG (rather than 8-oxo-G, which is present in elevated levels in some Parkinsons Disease patient brains, reference 40).

To characterise the effect of Aprataxin deficiency on oxidative stress levels, we examined the abundance of oxidized DNA (8-oxo-dG) and protein (nitrosylated tyrosine) AOA1 cells. We found that Aprataxin deficiency led to elevated levels of protein and DNA oxidation, indicating that AOA1 cells have a high basal level of oxidative stress.

8-oxo-dG base-pairs with adenine preferentially over cytosine and also induces low frequency misincorporation of upstream nucleotides if nascent DNA synthesis is performed by a non-proofreading polymerase (41). Thus 8-oxo-dG can induce point and tandem mutations in proliferative cells. 8-oxo-dG can also cause transcriptional mutagenesis by facilitating synthesis C→A point mutated transcripts which may result in translation of truncated, non-functional or toxic proteins (42,43). This may also contribute to the progressive neurodegeneration in disorders which are characterised by elevated levels of oxidative DNA damage.

8-oxo-dG is repaired by BER and the primary enzyme responsible for the detection and excision of this lesion is OGG1 (44). This enzyme hydrolyses the N-glycosyl bond between the phosphodiester backbone and the damaged base, leaving an abasic site. OGG1 is a bifunctional glycosylases, meaning that it also has abasic lyase

287 activity (it can cleave abasic sites to generate a 3’-α,β-unsaturated aldehyde 5’ phosphate gap, reference (44). Its lyase activity is low compared to its glycosylase activity, but it has similar affinity for both molecules (45,46). Thus depurination by OGG1 is strongly inhibited by build-up of abasic DNA. The depurination activity of OGG1 is stimulated by the abasic lyase APE1 (45-47), which occurs via rapid hydrolysis of abasic DNA by APE1, which prevents sequesteration of OGG1 (45,46). Thus although APE1 does not participate directly in 8-oxo-dG excision, it stimulates excision by facilitating OGG1 enzyme turnover (and performs a similar function for other bifunctional DNA glycosylases, references 45,46,48). I found that deficiency of Aprataxin results in destabilization of APE1. Uncorrected AOA1 cells have approximately 50% less APE1 than corrected AOA1 cells, indicating that this protein is unstable in the absence of Aprataxin. This provides a functional explanation for the elevated level of oxidative DNA damage in AOA1 cells and indicates that AOA1 cells may have a defect in BER.

Subsequent exploration of possible Aprataxin-related BER defects demonstrated that Aprataxin deficient cells have less PARP-1 than corrected cells. PARP-1 is a multifunctional protein with diverse roles including inhibition of classical NHEJ during late S, G2 and M phases, initiation of apoptosis, is required for an alternative NHEJ pathway and recruitment of repair factors for direct SSBR and BER (49-53). The deficiency of PARP-1 in Aprataxin deficient cells indicates that Aprataxin deficiency could cause defects in cellular functions which rely on PARP-1. A defect in in vitro DNA double strand break repair was not observed in AOA1 cells (Harris et al. Synergistic function of Aprataxin and PARP-1 in DNA repair, in preparation). This is consistent with the role of PARP-1 dependant NHEJ as a backup or redundant repair mechanism (49).

Although AOA1 cells do not have a defect in PARP-1 dependant NHEJ, they do have a defect in the gap filling phase of BER. As introduced in Chapter 1, BER is a mechanism for excision of damaged nucleotides (section 1.3.2.4). XRCC1 interacts with proteins involved in both short (DNA polymerase β and DNA ligase 3α) and long patch repair (PCNA) (54-56) and PARP-1 is required for the formation of stable XRCC1 foci at sites of DNA damage (52). Thus PARP-1 is a potential recruitment

288 factor for short and long patch BER proteins. PARP-1 deficiency has a minor impact on short patch repair and a major impact gap filling in long patch repair (57). This may be due to the defective recruitment of the PCNA-DNA polymerase long patch repair complex, which is responsible for the addition of the second and subsequent nucleotides in long patch repair (58,59). The interaction profile of PARP-2 is similar to that of PARP-1 and there is evidence that PARP-2 also participates in BER (60). Using an in vitro assay I determined that AOA1 cells have approximately 50% the long gap filling capacity of control cells. This is consistent with the established role of PARP-1 in long patch repair and the stabilization of PARP-1 by Aprataxin.

In addition to repair of modified bases, BER can facilitate repair of damaged 5’ termini the cell cannot process directly. As outlined previously this is performed by extension of multiple nucleotides from the 3’ hydroxyl terminus by the PCNA-DNA polymerase complex, causing displacement of the damaged terminus. The resultant flap is cleaved by FEN1 to generate a ligatable single strand nick. I and others have shown that AOA1 cells cannot directly hydrolyse 5’ adenylated DNA (22,25) and there is unpublished evidence that this modification accumulates in AOA1 cells (Prof Keith Caldecott, University of Sussex, Brighton , UK, personal communications). As such AOA1 cells probably rely on long patch BER to indirectly resolve 5’ adenylates. This study has demonstrated that AOA1 cells possess a reduced long patch repair capacity, indicating that they are probably less capable of indirect repair of all types of 5’ DNA damage, including adenylation.

In summary, I have characterised multiple compounding DNA repair defects in AOA1 cells. AOA1 cells are unable to directly excise 5’ DNA adenylates and have a reduced level of APE1 (which would cause defects in hydrolysis of abasic sites and impair turnover of DNA glycosylases). This is supported by the elevated levels of 8- oxo-dG in AOA1 cells, which is consistent with a defect in OGG1 turnover. I found that Aprataxin deficiency also results in destabilization of PARP-1 and identified resultant defect in gap filling in AOA1 cells. Thus not only are AOA1 cells unable to directly repair adenylates, they have defects in excision of oxidative lesions and

289 indirect repair of non-hydrolysable 5’ modifications. These compounding defects are likely to contribute to the accumulation of adenylated DNA in AOA1 cells.

While DNA lesions impact on genomic integrity, they can also affect transcription in both proliferative and non-proliferative cells. Depending of the nature of the lesion, DNA damage in coding regions can result in inhibition of transcription or the production of point mutated, frameshifted or truncated transcripts (61-63). This can cause either deficiency of vital proteins or production of toxic mutated proteins. Given that 1) AOA1 cells display an elevated level of 8-oxo-dG (which can base-pair with adenine facilitating generation of mutant transcripts), 2) Aprataxin interacts with the rRNA processing factor nucleolin in a rRNA transcription-dependant manner and 3) Aprataxin is present and its activity is elevated in the nucleolus (a site of high transcriptional activity), I examined whether Aprataxin may also have a role in transcription.

6.3.3 Interaction of Aprataxin with transcription and RNA processing factors:

Previous studies have demonstrated an interaction between Aprataxin and the rRNA processing factor nucleolin (32). I identified interactions between Aprataxin and UBF isoforms 1 and 2, both of which have transactivation and transcriptional anti- repressive activities (although both activities are substantially lower in UBF-2) (64,65). Aprataxin also interacts with TAF95, a protein which interacts with UBF-1 (among other proteins) at the rDNA promoter to form the pre-initiation complex (66). TAF95 interacted strongly with the nuclear localization region of Aprataxin, while both UBF proteins interact primarily with the FHA domain and to a lesser extent with the HIT domain. This interaction map indicates that UBF and TAF95 may interact with Aprataxin simultaneously, possibly at the pre-initiation complex. In addition to its interaction with these rRNA transcription initiation factors, I identified an interaction between Aprataxin and the RNA polymerase I transcription termination factor TTF1. TTF1 interacts with the FHA and HIT domains of Aprataxin with similar affinity. TTF1 is necessary for efficient RNA polymerase I transcription termination (67,68), but has also been found to bind to rDNA upstream of the core promoter and may also be involved in activation of transcription (69). Finally, an

290 interaction between Aprataxin and the large subunit of RNA polymerase II has been identified (Dr Olivier Becherel, Queensland Institute for Medical Research, Brisbane, Australia, unpublished observations). These interactions imply that repair of DNA- adenylates may occur in a transcription-coupled manner or that Aprataxin has a role in regulation of RNA polymerase I.

In addition to the interactions with these transcription factors I identified interactions between Aprataxin and the RNA processing factors fibrillarin and hnRNP U. Fibrillarin interacted primarily with the nuclear localization region of Aprataxin, indicating that it has the potential to interact directly with Aprataxin simultaneously with FHA domain interacting proteins. Fibrillarin indispensable for the early stages of rRNA processing and deficiency of fibrillarin is lethal (70,71). hnRNP U is an mRNA processing factor (72) which interacted with the FHA domain and to a limited extent with the HIT domain. The interaction of Aprataxin with factors involved in the processing of rRNA and mRNA indicated that Aprataxin may have a general role as a transcription and RNA processing factor, or be constitutively associated with transcriptional machinery so that it may preferentially repair DNA adenylate modifications in coding regions. In support of this, AOA1 cells have been shown to have a mild transcriptional defect, having 15% less 47S and 45S pre-rRNA than control cells (32).

6.3.4 Aprataxin-dependant stabilization of UBF:

UBF is an essential RNA polymerase I transcription factor with exclusively nucleolar localization (73,74). I hypothesized that a defect in localization or stabilization of UBF could be responsible for the reduced levels of 47S/45S pre-rRNA in AOA1 cells. I examined the localization of UBF by immunostaining and noted that in Aprataxin deficient cells the staining appeared more diffuse with some nucleoplasmic staining observed. Quantification of the UBF staining intensity of individual nuclei revealed that Aprataxin deficiency results in a marginal reduction in UBF protein level. Such a deficit could explain the reduced level of 47S/45S rRNA in AOA1 cells (32). To confirm the findings of our immunofluorescence experiment, I examined

291 UBF protein levels in Aprataxin deficient and corrected cells by Western blot. Surprisingly I found that both UBF isoforms are degraded in the absence of Aprataxin. The size of the degradation products indicates that UBFs DNA binding and transactivation domains have been separated. Given the central role of UBF in RNA polymerase I transcription initiation (73,74) this raises the question of how Aprataxin deficient cells are viable and able to initiate rRNA synthesis at all. I proposed a model where the two proteolysed fragments are able to interact and adopt a similar conformation to the intact protein. Additionally if these fragments interact independently with other transcription factors, these interactions may serve to hold them in a conformation capable of activating transcription.

6.3.5 Aprataxin deficient cells have defective transcriptional responses to DNA damage:

Treatment of cells with DNA damaging agents results in a cascade of cellular responses, many of which are mediated by protein kinases (reviewed in 75,76). One such response is the inhibition of DNA synthesis in response to DNA damage (the most commonly used agent is ionizing radiation). The purpose of this response is to prevent cells continuing DNA replication with unrepaired DNA breaks (reviewed in 77,78). Failure to inhibit cell cycle progression by activation of the intra-S-phase checkpoint can have a range of deleterious effects. Single strand breaks can be converted to more toxic double strand breaks by causing collapse of replication forks (79). Unrepaired 8-oxo-dG or abasic sites in S phase result in point mutations (41). Thus activation of the intra-S-phase checkpoint is critical for maintenance of genomic stability. Radioresistant DNA synthesis (RDS) is a result of failure of the intra-S- phase checkpoint and is a hallmark of defective cell signalling after DNA damage (77,78). Activation of this mechanism is dependant on ATM but independent of p53 (77,78) and cells derived from A-T patients display RDS (80).

Similar to the well-studied situation with RDS, a recent report identified an ATM- dependant mechanism for inhibition of RNA synthesis after DNA damage (81). Cells from A-T patients continue to synthesize RNA after DNA damage, while control cells

292 inhibit transcription (81). This provides a mechanism for the cell to prevent generation of aberrant transcripts from damaged DNA coding regions. In this study I examined the effect of Aprataxin deficiency and γ-irradiation on RNA synthesis. Aprataxin deficient cells did not display significant inhibition of RNA synthesis after irradiation, whereas corrected cells displayed a 31.2% reduction. Thus Aprataxin deficiency results in RRS. This defective transcriptional response to irradiation indicates that Aprataxin deficient cells have a defect in DNA-damage signalling or downstream effectors of RNA synthesis inhibition. Clements et al. determined that AOA1 cells do not have a defect in ATM activation (26), indicating that this phenomenon is probably due a defect downstream of ATM.

Given the hypersensitivity of AOA1 cells to the oxidizing agent hydrogen peroxide, I also examined the effect of this agent and Aprataxin deficiency on transcription. Conversely to what was observed with irradiation, I found that Aprataxin deficient cells display an extremely sensitive transcriptional response to hydrogen peroxide induced DNA damage. The mean transcriptional activity of Aprataxin deficient cells decreased by 78% after treatment with 0.5 mM hydrogen peroxide (p= 0.004), whereas the transcriptional activity of corrected cells was not significantly affected (16% reduction, p= 0.305). This dramatic effect of hydrogen peroxide on transcription in Aprataxin deficient cells may be due to their elevated basal levels of oxidative stress. I demonstrated that control (corrected) cells have less oxidative DNA and protein damage than AOA1 (uncorrected) cells. Thus in the absence of Aprataxin cells may require lower doses of exogenous oxidative stress inducing cells to induce stress-response mechanisms. This is consistent with the increased number of breaks induced by the same dose of hydrogen peroxide in AOA1 compared to wild-type cells (Markus Fusser, Institute of Pharmacy, University of Mainz, Germany, personal communications), which together with this abnormal transcriptional response may explain the hypersensitivity of AOA1 cells to oxidative stress.

The hypersensitive transcriptional response of Aprataxin deficient cells to hydrogen peroxide and their RRS indicate that cells initiate different responses to γ-radiation

293 and hydrogen peroxide induced damage, and highlights the complexity of the DNA damage response.

6.4 FUTURE DIRECTIONS

It is from its catalytic activity of Aprataxin has the potential to function in many DNA repair mechanisms (as all of them require ligation as a final step) and that Aprataxin deficiency has far-reaching consequences in the cellular response to DNA damage. Now that the primary in vivo function of Aprataxin is known and some of the functional consequences of its deficiency have been identified, future work should focus on the regulation of Aprataxin’s activity and characterisation of its role in cellular responses which are defective in AOA1. Specifically:

1. Investigation of phosphorylation as a means of DNA-adenylate hydrolase activity regulation. ATR phosphorylation of S168 may regulate activity of the HIT domain directly, or indirectly by modulating protein-protein interactions or sub-nuclear distribution. Confirmation of S168 as the target of ATR phosphorylation should be confirmed using kinase assays with S168A mutated Aprataxin. The effect of ATR and DNA damage on phosphorylation of this site could be examined using phospho-specific antibodies after a range of treatments of control and ATR defective (Seckel) cells. Aprataxin activity should be measured in Seckel cells. 2. Investigation of protein-protein interactions as a means of in vivo Aprataxin- regulation. I found that the activity of recombinant Aprataxin is stimulated by deletion of the FHA domain. Given that PARP-1 deficiency results in destabilization of Aprataxin, examination of the effect of protein-protein interactions may require analysis of activity using recombinant Aprataxin with recombinant interacting proteins. 3. Examination of the contribution of SSBR protein deficiency to the DNA damage hypersensitivity of AOA1 cells. APE1 and PARP-1 are central components of SSBR and are destabilized by Aprataxin deficiency. The contribution of Aprataxin activity to SSBR cannot be determined given that

294 Aprataxin deficiency destabilizes APE1 and PARP-1. The contribution of PARP-1 and APE1 deficiency to the hypersensitivity of AOA1 cells should be investigated using AOA1 cells transfected with PARP-1 and APE1 expression constructs. 4. Investigation of the contribution of PARP-1 and APE1 deficiency to the elevated basal level of oxidative stress in AOA1 cells. PARP-1 and APE1 are involved in the cellular response to oxidative stress. The contribution of these proteins to the elevated basal level of oxidative stress in AOA1 cells can be examined in a similar manner to 3. 5. Investigation of the cause of radioresistant RNA synthesis in Aprataxin deficient cells. Radioresistant RNA synthesis in Aprataxin deficient cells indicates that they may have a defect in the ATM signalling pathway. DNA damage induced phosphorylation of a range of ATM targets should be examined in AOA1 cells. Pathways responsible for inhibition of RNA synthesis could be explored using specific protein kinase inhibitors. It would also be interesting to determine if AOA1 cells display RDS. 6. Investigation of the mechanism of inhibition of transcription in Aprataxin deficient cells by hydrogen peroxide. The hypersensitive transcriptional response of AOA1 cells to hydrogen peroxide could be due to either a very high level of DNA damage in coding regions or a defective signalling response. These mechanisms could be discriminated by determining if this defect is corrected by treatment of cells with antioxidants (increasing their capacity to cope with oxidative DNA damage) and inhibition of different stress signalling pathways.

Cerebellar degeneration, and particularly Purkinje cell loss, is a central feature of DNA repair defective ataxias. Understanding the reason for this loss is of central importance to the DNA repair/cerebellar ataxia field. Examination of the intracellular sources of oxidative stress and its effects on Purkinje cells compared to other neuronal cell populations ex vivo could provide a molecular understanding of the common neurological phenotype shared by patients with defects in such diverse cellular mechanisms.

295 6.7 REFERENCES

1. Mohamed, R. and Lavin, M.F. (1986) Ataxia-telangiectasia cell extracts confer radioresistant DNA synthesis on control cells. Exp Cell Res, 163, 337- 348.

2. Stewart, G.S., Maser, R.S., Stankovic, T., Bressan, D.A., Kaplan, M.I., Jaspers, N.G., Raams, A., Byrd, P.J., Petrini, J.H. and Taylor, A.M. (1999) The DNA double-strand break repair gene hMRE11 is mutated in individuals with an ataxia-telangiectasia-like disorder. Cell, 99, 577-587.

3. Chu, G. and Mayne, L. (1996) Xeroderma pigmentosum, Cockayne syndrome and trichothiodystrophy: do the genes explain the diseases? Trends Genet, 12, 187-192.

4. Date, H., Onodera, O., Tanaka, H., Iwabuchi, K., Uekawa, K., Igarashi, S., Koike, R., Hiroi, T., Yuasa, T., Awaya, Y. et al. (2001) Early-onset ataxia with ocular motor apraxia and hypoalbuminemia is caused by mutations in a new HIT superfamily gene. Nat Genet, 29, 184-188.

5. Moreira, M.C., Barbot, C., Tachi, N., Kozuka, N., Uchida, E., Gibson, T., Mendonca, P., Costa, M., Barros, J., Yanagisawa, T. et al. (2001) The gene mutated in ataxia-ocular apraxia 1 encodes the new HIT/Zn-finger protein aprataxin. Nat Genet, 29, 189-193.

6. Moreira, M.C., Klur, S., Watanabe, M., Nemeth, A.H., Le Ber, I., Moniz, J.C., Tranchant, C., Aubourg, P., Tazir, M., Schols, L. et al. (2004) Senataxin, the ortholog of a yeast RNA helicase, is mutant in ataxia-ocular apraxia 2. Nat Genet, 36, 225-227.

7. Le Ber, I., Moreira, M.C., Rivaud-Pechoux, S., Chamayou, C., Ochsner, F., Kuntzer, T., Tardieu, M., Said, G., Habert, M.O., Demarquay, G. et al. (2003) Cerebellar ataxia with oculomotor apraxia type 1: clinical and genetic studies. Brain, 126, 2761-2772.

296 8. Amouri, R., Moreira, M.C., Zouari, M., El Euch, G., Barhoumi, C., Kefi, M., Belal, S., Koenig, M. and Hentati, F. (2004) Aprataxin gene mutations in Tunisian families. Neurology, 63, 928-929.

9. Habeck, M., Zuhlke, C., Bentele, K.H., Unkelbach, S., Kress, W., Burk, K., Schwinger, E. and Hellenbroich, Y. (2004) Aprataxin mutations are a rare cause of early onset ataxia in Germany. J Neurol, 251, 591-594.

10. Baneyx, F. (1999) Recombinant protein expression in Escherichia coli. Curr Opin Biotechnol, 10, 411-421.

11. Hirano, M., Asai, H., Kiriyama, T., Furiya, Y., Iwamoto, T., Nishiwaki, T., Yamamoto, A., Mori, T. and Ueno, S. (2007) Short half-lives of ataxia- associated aprataxin proteins in neuronal cells. Neurosci Lett, 419, 184-187.

12. Seidle, H.F., Bieganowski, P. and Brenner, C. (2005) Disease-associated mutations inactivate AMP-lysine hydrolase activity of aprataxin. J Biol Chem. 280, 20927-20931.

13. Hirano, M., Furiya, Y., Kariya, S., Nishiwaki, T. and Ueno, S. (2004) Loss of function mechanism in aprataxin-related early-onset ataxia. Biochem Biophys Res Commun, 322, 380-386.

14. Brenner, C., Bieganowski, P., Pace, H.C. and Huebner, K. (1999) The histidine triad superfamily of nucleotide-binding proteins. J Cell Physiol, 181, 179-187.

15. Barnes, L.D., Garrison, P.N., Siprashvili, Z., Guranowski, A., Robinson, A.K., Ingram, S.W., Croce, C.M., Ohta, M. and Huebner, K. (1996) Fhit, a putative tumor suppressor in humans, is a dinucleoside 5',5"'-P1,P3-triphosphate hydrolase. Biochemistry, 35, 11529-11535.

16. Lima, C.D., Klein, M.G. and Hendrickson, W.A. (1997) Structure-based analysis of catalysis and substrate definition in the HIT protein family. Science, 278, 286-290.

17. Pace, H.C., Garrison, P.N., Robinson, A.K., Barnes, L.D., Draganescu, A., Rosler, A., Blackburn, G.M., Siprashvili, Z., Croce, C.M., Huebner, K. et al.

297 (1998) Genetic, biochemical, and crystallographic characterization of Fhit- substrate complexes as the active signaling form of Fhit. Proc Natl Acad Sci U S A, 95, 5484-5489.

18. Chu, D.M., Corbin, J.D., Grimes, K.A. and Francis, S.H. (1997) Activation by cyclic GMP binding causes an apparent conformational change in cGMP- dependent protein kinase. J Biol Chem, 272, 31922-31928.

19. Davletov, B.A. and Sudhof, T.C. (1994) Ca(2+)-dependent conformational change in synaptotagmin I. J Biol Chem, 269, 28547-28550.

20. Kijas, A.W., Harris, J.L., Harris, J.M. and Lavin, M.F. (2006) Aprataxin forms a discrete branch in the HIT (histidine triad) superfamily of proteins with both DNA/RNA binding and nucleotide hydrolase activities. J Biol Chem, 281, 13939-13948.

21. Mosesso, P., Piane, M., Palitti, F., Pepe, G., Penna, S. and Chessa, L. (2005) The novel human gene aprataxin is directly involved in DNA single-strand- break repair. Cell Mol Life Sci, 62, 485-491.

22. Ahel, I., Rass, U., El-Khamisy, S.F., Katyal, S., Clements, P.M., McKinnon, P.J., Caldecott, K.W. and West, S.C. (2006) The neurodegenerative disease protein aprataxin resolves abortive DNA ligation intermediates. Nature. 433, 713-716.

23. Christiansen, K. and Westergaard, O. (1996) The effect of camptothecin on topoisomerase I catalysis. Ann N Y Acad Sci, 803, 50-59.

24. Lindahl, T., Karran, P. and Wood, R.D. (1997) DNA excision repair pathways. Current Opinion in Genetics & Development, 7, 158-169.

25. Rass, U., Ahel, I. and West, S.C. (2007) Actions of aprataxin in multiple DNA repair pathways. J Biol Chem, 282, 9469-9474.

26. Clements, P.M., Breslin, C., Deeks, E.D., Byrd, P.J., Ju, L., Bieganowski, P., Brenner, C., Moreira, M.C., Taylor, A.M. and Caldecott, K.W. (2004) The ataxia-oculomotor apraxia 1 gene product has a role distinct from ATM and

298 interacts with the DNA strand break repair proteins XRCC1 and XRCC4. DNA Repair (Amst), 3, 1493-1502.

27. Luo, H., Chan, D.W., Yang, T., Rodriguez, M., Chen, B.P., Leng, M., Mu, J.J., Chen, D., Songyang, Z., Wang, Y. et al. (2004) A new XRCC1- containing complex and its role in cellular survival of methyl methanesulfonate treatment. Mol Cell Biol, 24, 8356-8365.

28. Shen, M.R., Zdzienicka, M.Z., Mohrenweiser, H., Thompson, L.H. and Thelen, M.P. (1998) Mutations in hamster single-strand break repair gene XRCC1 causing defective DNA repair. Nucleic Acids Res, 26, 1032-1037.

29. Koch, C.A., Agyei, R., Galicia, S., Metalnikov, P., O'Donnell, P., Starostine, A., Weinfeld, M. and Durocher, D. (2004) Xrcc4 physically links DNA end processing by polynucleotide kinase to DNA ligation by DNA ligase IV. The EMBO Journal, 23, 3874-3885.

30. Whitehouse, C.J., Taylor, R.M., Thistlethwaite, A., Zhang, H., Karimi- Busheri, F., Lasko, D.D., Weinfeld, M. and Caldecott, K.W. (2001) XRCC1 stimulates human polynucleotide kinase activity at damaged DNA termini and accelerates DNA single-strand break repair. Cell, 104, 107-117.

31. Gueven, N., Becherel, O.J., Kijas, A.W., Chen, P., Howe, O., Rudolph, J.H., Gatti, R., Date, H., Onodera, O., Taucher-Scholz, G. et al. (2004) Aprataxin, a novel protein that protects against genotoxic stress. Hum Mol Genet, 13, 1081- 1093.

32. Becherel, O.J., Gueven, N., Birrell, G.W., Schreiber, V., Suraweera, A., Jakob, B., Taucher-Scholz, G. and Lavin, M.F. (2006) Nucleolar localization of aprataxin is dependent on interaction with nucleolin and on active ribosomal DNA transcription. Hum Mol Genet, 15, 2239-2249.

33. Takahashi, T., Tada, M., Igarashi, S., Koyama, A., Date, H., Yokoseki, A., Shiga, A., Yoshida, Y., Tsuji, S., Nishizawa, M. et al. (2007) Aprataxin, causative gene product for EAOH/AOA1, repairs DNA single-strand breaks with damaged 3'-phosphate and 3'-phosphoglycolate ends. Nucleic Acids Res, 35, 3797-3809.

299 34. Breen, A.P. and Murphy, J.A. (1995) Reactions of oxyl radicals with DNA. Free Radical Biology & Medicine, 18, 1033-1077.

35. Wiederhold, L., Leppard, J.B., Kedar, P., Karimi-Busheri, F., Rasouli-Nia, A., Weinfeld, M., Tomkinson, A.E., Izumi, T., Prasad, R., Wilson, S.H. et al. (2004) AP endonuclease-independent DNA base excision repair in human cells. Mol Cell, 15, 209-220.

36. Winters, T.A., Henner, W.D., Russell, P.S., McCullough, A. and Jorgensen, T.J. (1994) Removal of 3'-phosphoglycolate from DNA strand-break damage in an oligonucleotide substrate by recombinant human apurinic/apyrimidinic endonuclease 1. Nucleic Acids Res, 22, 1866-1873.

37. Chen, P., Peng, C., Luff, J., Spring, K., Watters, D., Bottle, S., Furuya, S. and Lavin, M.F. (2003) Oxidative stress is responsible for deficient survival and dendritogenesis in purkinje neurons from ataxia-telangiectasia mutated mutant mice. J Neurosci, 23, 11453-11460.

38. Suraweera, A., Becherel, O.J., Chen, P., Rundle, N., Woods, R., Nakamura, J., Gatei, M., Criscuolo, C., Filla, A., Chessa, L. et al. (2007) Senataxin, defective in ataxia oculomotor apraxia type 2, is involved in the defense against oxidative DNA damage. J Cell Biol, 177, 969-979.

39. Hirano, M., Yamamoto, A., Mori, T., Lan, L., Iwamoto, T.A., Aoki, M., Shimada, K., Furiya, Y., Kariya, S., Asai, H. et al. (2007) DNA single-strand break repair is impaired in aprataxin-related ataxia. Ann Neurol, 61, 162-174.

40. Zhang, J., Perry, G., Smith, M.A., Robertson, D., Olson, S.J., Graham, D.G. and Montine, T.J. (1999) Parkinson's disease is associated with oxidative damage to cytoplasmic DNA and RNA in substantia nigra neurons. Am J Pathol, 154, 1423-1429.

41. Efrati, E., Tocco, G., Eritja, R., Wilson, S.H. and Goodman, M.F. (1999) "Action-at-a-distance" mutagenesis. 8-oxo-7, 8-dihydro-2'-deoxyguanosine causes base substitution errors at neighboring template sites when copied by DNA polymerase beta. J Biol Chem, 274, 15920-15926.

300 42. Chen, Y.H. and Bogenhagen, D.F. (1993) Effects of DNA lesions on transcription elongation by T7 RNA polymerase. J Biol Chem, 268, 5849- 5855.

43. Kuraoka, I., Endou, M., Yamaguchi, Y., Wada, T., Handa, H. and Tanaka, K. (2003) Effects of endogenous DNA base lesions on transcription elongation by mammalian RNA polymerase II. Implications for transcription-coupled DNA repair and transcriptional mutagenesis. J Biol Chem, 278, 7294-7299.

44. van der Kemp, P.A., Thomas, D., Barbey, R., de Oliveira, R. and Boiteux, S. (1996) Cloning and expression in Escherichia coli of the OGG1 gene of Saccharomyces cerevisiae, which codes for a DNA glycosylase that excises 7,8-dihydro-8-oxoguanine and 2,6-diamino-4-hydroxy-5-N- methylformamidopyrimidine. Proc Natl Acad Sci U S A, 93, 5197-5202.

45. Hill, J.W., Hazra, T.K., Izumi, T. and Mitra, S. (2001) Stimulation of human 8-oxoguanine-DNA glycosylase by AP-endonuclease: potential coordination of the initial steps in base excision repair. Nucleic Acids Res, 29, 430-438.

46. Vidal, A.E., Hickson, I.D., Boiteux, S. and Radicella, J.P. (2001) Mechanism of stimulation of the DNA glycosylase activity of hOGG1 by the major human AP endonuclease: bypass of the AP lyase activity step. Nucleic Acids Res, 29, 1285-1292.

47. Saitoh, T., Shinmura, K., Yamaguchi, S., Tani, M., Seki, S., Murakami, H., Nojima, Y. and Yokota, J. (2001) Enhancement of OGG1 protein AP lyase activity by increase of APEX protein. Mutat Res, 486, 31-40.

48. Marenstein, D.R., Chan, M.K., Altamirano, A., Basu, A.K., Boorstein, R.J., Cunningham, R.P. and Teebor, G.W. (2003) Substrate specificity of human endonuclease III (hNTH1). Effect of human APE1 on hNTH1 activity. J Biol Chem, 278, 9005-9012.

49. Wang, M., Wu, W., Wu, W., Rosidi, B., Zhang, L., Wang, H. and Iliakis, G. (2006) PARP-1 and Ku compete for repair of DNA double strand breaks by distinct NHEJ pathways. Nucleic Acids Res, 34, 6170-6182.

301 50. Yu, S.W., Wang, H., Poitras, M.F., Coombs, C., Bowers, W.J., Federoff, H.J., Poirier, G.G., Dawson, T.M. and Dawson, V.L. (2002) Mediation of poly(ADP-ribose) polymerase-1-dependent cell death by apoptosis-inducing factor. Science, 297, 259-263.

51. Audebert, S., Calsou. (2004) Involvement of Poly(ADP-ribose) Polymerase-1 and XRCC1/DNA Ligase III in an Alternative Route for DNA Double-strand Breaks Rejoining. J Biol Chem, 279, 55117-55126.

52. El-Khamisy, S.F., Masutani, M., Suzuki, H. and Caldecott, K.W. (2003) A requirement for PARP-1 for the assembly or stability of XRCC1 nuclear foci at sites of oxidative DNA damage. Nucleic Acids Research, 31, 5526-5533.

53. Le Page, F., Schreiber, V., Dherin, C., De Murcia, G. and Boiteux, S. (2003) Poly(ADP-ribose) polymerase-1 (PARP-1) is required in murine cell lines for base excision repair of oxidative DNA damage in the absence of DNA polymerase beta. J Biol Chem, 278, 18471-18477.

54. Marintchev, A., Robertson, A., Dimitriadis, E.K., Prasad, R., Wilson, S.H. and Mullen, G.P. (2000) Domain specific interaction in the XRCC1-DNA polymerase beta complex. Nucleic Acids Res, 28, 2049-2059.

55. Kubota, Y., Nash, R.A., Klungland, A., Schar, P., Barnes, D.E. and Lindahl, T. (1996) Reconstitution of DNA base excision-repair with purified human proteins: interaction between DNA polymerase beta and the XRCC1 protein. The EMBO Journal, 15, 6662-6670.

56. Fan, J., Otterlei, M., Wong, H.K., Tomkinson, A.E. and Wilson, D.M., 3rd. (2004) XRCC1 co-localizes and physically interacts with PCNA. Nucleic Acids Res, 32, 2193-2201.

57. Dantzer, F., de La Rubia, G., Menissier-De Murcia, J., Hostomsky, Z., de Murcia, G. and Schreiber, V. (2000) Base excision repair is impaired in mammalian cells lacking Poly(ADP-ribose) polymerase-1. Biochemistry, 39, 7559-7569.

302 58. Podlutsky, A.J., Dianova, II, Podust, V.N., Bohr, V.A. and Dianov, G.L. (2001) Human DNA polymerase beta initiates DNA synthesis during long- patch repair of reduced AP sites in DNA. The EMBO Journal, 20, 1477-1482.

59. Fortini, P., Pascucci, B., Parlanti, E., Sobol, R.W., Wilson, S.H. and Dogliotti, E. (1998) Different DNA polymerases are involved in the short- and long- patch base excision repair in mammalian cells. Biochemistry, 37, 3575-3580.

60. Schreiber, V., Ame, J.C., Dolle, P., Schultz, I., Rinaldi, B., Fraulob, V., Menissier-de Murcia, J. and de Murcia, G. (2002) Poly(ADP-ribose) polymerase-2 (PARP-2) is required for efficient base excision DNA repair in association with PARP-1 and XRCC1. J Biol Chem, 277, 23028-23036.

61. Liu, J. and Doetsch, P.W. (1996) Template strand gap bypass is a general property of prokaryotic RNA polymerases: implications for elongation mechanisms. Biochemistry, 35, 14999-15008.

62. Zhou, W. and Doetsch, P.W. (1994) Transcription bypass or blockage at single-strand breaks on the DNA template strand: effect of different 3' and 5' flanking groups on the T7 RNA polymerase elongation complex. Biochemistry, 33, 14926-14934.

63. Zhou, W., Reines, D. and Doetsch, P.W. (1995) T7 RNA polymerase bypass of large gaps on the template strand reveals a critical role of the nontemplate strand in elongation. Cell, 82, 577-585.

64. Kuhn, A., Voit, R., Stefanovsky, V., Evers, R., Bianchi, M. and Grummt, I. (1994) Functional differences between the two splice variants of the nucleolar transcription factor UBF: the second HMG box determines specificity of DNA binding and transcriptional activity. The EMBO Journal, 13, 416-424.

65. O'Mahony, D.J. and Rothblum, L.I. (1991) Identification of two forms of the RNA polymerase I transcription factor UBF. Proc Natl Acad Sci U S A, 88, 3180-3184.

66. Learned, R.M., Learned, T.K., Haltiner, M.M. and Tjian, R.T. (1986) Human rRNA transcription is modulated by the coordinate binding of two factors to an upstream control element. Cell, 45, 847-857.

303 67. Grummt, I., Rosenbauer, H., Niedermeyer, I., Maier, U. and Ohrlein, A. (1986) A repeated 18 bp sequence motif in the mouse rDNA spacer mediates binding of a nuclear factor and transcription termination. Cell, 45, 837-846.

68. Sander, E.E., Mason, S.W., Munz, C. and Grummt, I. (1996) The amino- terminal domain of the transcription termination factor TTF-I causes protein oligomerization and inhibition of DNA binding. Nucleic Acids Res, 24, 3677- 3684.

69. Langst, G., Becker, P.B. and Grummt, I. (1998) TTF-I determines the chromatin architecture of the active rDNA promoter. The EMBO Journal, 17, 3135-3145.

70. Newton, K., Petfalski, E., Tollervey, D. and Caceres, J.F. (2003) Fibrillarin is essential for early development and required for accumulation of an intron- encoded small nucleolar RNA in the mouse. Mol Cell Biol, 23, 8519-8527.

71. Tollervey, D., Lehtonen, H., Carmo-Fonseca, M. and Hurt, E.C. (1991) The small nucleolar RNP protein NOP1 (fibrillarin) is required for pre-rRNA processing in yeast. The EMBO Journal, 10, 573-583.

72. Kukalev, A., Nord, Y., Palmberg, C., Bergman, T. and Percipalle, P. (2005) Actin and hnRNP U cooperate for productive transcription by RNA polymerase II. Nat Struct Mol Biol, 12, 238-244.

73. Voit, R., Kuhn, A., Sander, E.E. and Grummt, I. (1995) Activation of mammalian ribosomal gene transcription requires phosphorylation of the nucleolar transcription factor UBF. Nucleic Acids Res, 23, 2593-2599.

74. Voit, R. and Grummt, I. (2001) Phosphorylation of UBF at serine 388 is required for interaction with RNA polymerase I and activation of rDNA transcription. Proc Natl Acad Sci U S A, 98, 13631-13636.

75. Lowndes, N.F. and Murguia, J.R. (2000) Sensing and responding to DNA damage. Current Opinion in Genetics & Development, 10, 17-25.

76. Shiloh, Y. (2001) ATM and ATR: networking cellular responses to DNA damage. Current Opinion in Genetics & Development, 11, 71-77.

304 77. Iliakis, G., Wang, Y., Guan, J. and Wang, H. (2003) DNA damage checkpoint control in cells exposed to ionizing radiation. Oncogene, 22, 5834-5847.

78. Bartek, J., Lukas, C. and Lukas, J. (2004) Checking on DNA damage in S phase. Nat Rev Mol Cell Biol, 5, 792-804.

79. Kuzminov, A. (2001) Single-strand interruptions in replicating chromosomes cause double-strand breaks. Proc Natl Acad Sci U S A, 98, 8241-8246.

80. Painter, R.B. (1981) Radioresistant DNA synthesis: an intrinsic feature of ataxia telangiectasia. Mutat Res, 84, 183-190.

81. Kruhlak, M., Crouch, E.E., Orlov, M., Montano, C., Gorski, S.A., Nussenzweig, A., Misteli, T., Phair, R.D. and Casellas, R. (2007) The ATM repair pathway inhibits RNA polymerase I transcription in response to chromosome breaks. Nature, 447, 730-734.

305 306

APPENDIX I

307

APPENDIX II