BIOCHEMICAL CHARACTERIZATION OF APRATAXIN, THE … · 2013-07-02 · STATEMENT OF ORIGINALITY ... (A-TLD) as well as the recently identified diseases Spinocerebellar ataxia with axonal

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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

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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

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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

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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.

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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.

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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.

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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



Australian Society for Medical Research, Queensland Conference, 2007



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.

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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



National Institute for Aging (Baltimore MD, USA), 2008



Queensland Protein Group Conference, 2008



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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.

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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.

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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

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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


Aprataxin from pTYB1 70


Aprataxin from pGEX 6.1 71

2.2.7 Purification of anti-Aprataxin specific antibodies 72

2.2.8 Estimation of protein concentration 73

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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


3.2.1 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

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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


4.2.1 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

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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

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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


5.2.1 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

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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

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APPENDIX I

Kijas, A. W., Harris, J. L., Harris, J. M., Lavin, M. F. (2006). Aprataxin forms a


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

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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

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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

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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

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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

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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

xxx

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

xxxi

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

xxxii

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

Glutathione agarose Scientifix/

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

xxxiii

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

xxxiv

LIST OF ANTIBODIES AND SUPPLIERS

Primary antibodies

Immunogen Host species Company/ Generated by Clone/Cat#

8-oxo-G mouse Trevigen 4355-MC-100

AIF rabbit Cell Signalling 4642

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

Aprataxin rabbit

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

Aprataxin sheep

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

a-tubulin mouse Sigma 2-28-33 ATM mouse GeneTex 2CI

ATR rabbit

raised and purified by Dr Rick Woods (Queensland Institute for Medical Research, 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

Fibrillarin rabbit

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

GST rabbit Dr Amila Suraweera, Queensland Institute 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

xxxv

RNA polymerase II mouse Abcam ab5408

TAF95 rabbit

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

TTF1 rabbit

Dr Ingrid Grummt, Institute of Cell and Tumor Biology, German Cancer Research 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

xxxvi

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, AustraliaRoche 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

xxxvii

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

FD105 M21 immortalized AOA1 fibroblast cell line, transformed full length APTX 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

xxxviii

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

xxxix

1

CHAPTER 1

Literature Review

2

3

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

4

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).

5

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


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

6

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

7

(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

8

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


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).

9

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).

10

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

11

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).

12

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.

13

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).

14

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

15

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

16

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.

17

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

18

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

19

(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

20

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

21

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

22

(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).

23

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).

24

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

25

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.

26

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

27

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


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).

28

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-

29

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.

30

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.

31

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

32

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).

33

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

34

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).

35

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).

36

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).

37

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

38

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,

39

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


40

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).

41

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.

42

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63

CHAPTER 2

Generation of recombinant Aprataxin and Aprataxin-

specific antibodies

64

65

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

66

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

67

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.

68

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.

69

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

70

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

71

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

72

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.

73

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.

74

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.

75

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.

76

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).

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.

100

50

40

77

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.

78

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.

79

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.

80

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.

81

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.

1 0.1 0.2

WT V263G

1 0.5 0.2 0.1 0.2 0.5 1 2 3

ug BSA

50kD

40kD

30kD

60kD

82

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.

83

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).

84

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.

85

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.

86

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.

A C B

87

Species Antigen Purified

Against

Specific to

Rabbit Whole protein

(aa 1-342)-GST

Protein C

(aa 1-319)

FHA and HIT

domains

Sheep FHA (aa 1-

118) -GST

FHA (aa 1-118) FHA domain

Sheep Protein B (aa

118- 342) -GST

Protein C (aa 1-

319)

HIT domain

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

88

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

89

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.

90

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.

91

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.

92

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.




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.




93

12. Kijas, A.W., Harris, J.L., Harris, J.M. and Lavin, M.F. (2006) Aprataxin forms a



13939-13948.

94

95

CHAPTER 3

Biochemical characterization of recombinant

Aprataxin

96

97

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.

98

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.

99


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.

100

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

101

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

102

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

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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.

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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:


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.


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

105

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:


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,

106

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.


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.

107

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.

108

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

1 2 3 4 5 6 7 8 9 10 11

full length Aprataxin

109

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.

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


prior to the addition of 0.2 ng of chymotrypsin. Lanes 6 and 7- yeast recombinant


prior to the addition of 0.1 ng of chymotrypsin. Lanes 8 and 9- yeast recombinant


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.

1 2 3 4 5 6 7 8 9

110

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

1 2 3 4 5 6 7

111

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.

112

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.

113

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.

114

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

115

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.

116

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

117

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.

0/1

,

oncSubstrateC

oncSubstrateC

Under these circumstances, substitution into the equation for this straight line yields:

cVelocity /1

which can be rearranged to determine the velocity of reaction at infinite substrate

concentration:

cVelocity

cVelocity

/1

1

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

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determine the affinity of an enzyme for its substrate, Km. The gradient of the line is

described by:

max/VKmm

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:

maxVmKm

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

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(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).

120

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.

A.

B.

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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.

A.

B.

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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.

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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.

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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

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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).

126

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.

A.

B.

127

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

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

E + Appp

E-AMP + 5’P-DNA

E-AppDNA + 3’OH DNA DNA

E-AMP + PPi

E-AppDNA

DNA-p-DNA

1. enzyme adenylation

2. DNA adenylation

3. end joining

128

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).

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.

Oligo A Oligo B

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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

130

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).

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.

Oligo A Oligo B

131

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.

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

= P-32 phosphate group

5’18-mer 3’18-mer

132

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.

133

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.

A.

B.

134

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

135

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

136

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.

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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).

138

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

139

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.

140

3.5 REFERENCES


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.








interacts with the DNA strand break repair proteins XRCC1 and XRCC4. DNA

Repair (Amst), 3, 1493-1502.




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.

141

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.



Res Commun, 322, 380-386.


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



13939-13948.




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.




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.

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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".






35, 3797-3809.

143

CHAPTER 4

Characterization of multiple DNA repair defects in

AOA1 cells

144

145

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

146

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

147

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.

148


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.

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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.

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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.

151

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.


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:


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.

152

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.


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.

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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 37C. 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 37C. 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

154

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

155

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

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deficiency respectively are confirmed in these cell lines by immunoblotting. All cell

lines were obtained from the American Type Culture Collection.


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).

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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).


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.

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4.2.15 Gap filling assay:


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

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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.


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.

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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.

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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

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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).

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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).

non-specific

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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.

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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.

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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

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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

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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).

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.

= P-32 phosphate agroup

5’ 18-mer 3’ 18-mer

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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.

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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.

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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.

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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.

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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

174

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.

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.

B.

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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.

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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

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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.

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

DAPI

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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.

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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.

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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.

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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

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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).

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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

AMP-18mer

18mer

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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.

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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.

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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

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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.

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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.

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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.

Oligo A-6

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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

A.

B.

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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

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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.

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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.

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12

34

56

L939

L938

C2ABR

C3ABR

0

5

10

15

20

25

30

35

relative abundance

nucleotides extended

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.

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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.

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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

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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.

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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

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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.

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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.

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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).

202

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-

203

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

204

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.

205

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

206

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.

207

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215

CHAPTER 5

A role for Aprataxin in RNA biogenesis

216

217

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).

218

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

219

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

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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.

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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

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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

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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.

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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

225

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.

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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

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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

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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.

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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

SFPQ

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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.

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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).

232

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.

233

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

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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.

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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).

236

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.

DAPI

237

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,

DAPI

238

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

239

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.

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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

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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.

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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.

“Pulse” with analogue

1. 2. 3.

Remove pulse mixture.

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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

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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.

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.

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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).

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.

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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).

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

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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).

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.

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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

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experiments is between 1/50 and 1/250. For the following experiments a 1/100

dilution was used.

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

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(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.

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.

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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.

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.

nucleolus

Enlargement of top

cell in + Actinomycin D

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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.

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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

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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).

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.

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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.

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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).

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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.

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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.

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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

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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.

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.

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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.

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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.

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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

264

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.

265

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

266

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

267

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

268

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.

269

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277

CHAPTER 6

General discussion

278

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

279

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

280

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,

281

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

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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.

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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

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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.

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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.

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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).

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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

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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

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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

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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

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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

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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

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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

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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

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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.

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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

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307

APPENDIX I

halla

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APPENDIX II

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