the functional characterisation of human rmnd5 …...5.2.2 rmnd5 proteins colocalise with nkx3.1 in...

The Functional Characterisation of Human RMND5 Proteins in Normal

Physiology and Prostate Cancer

Alison Louw

Bsc (Hons)

Student Number 10476359

School of Pathology and Laboratory Medicine

This thesis is submitted in fulfilment of the requirements for the award of Doctor of Philosophy at the University of Western Australia

DECLARATION FOR THESES CONTAINING PUBLISHED WORK AND/OR WORK PREPARED FOR PUBLICATION The examination of the thesis is an examination of the work of the student. The work must have been substantially conducted by the student during enrolment in the degree. Where the thesis includes work to which others have contributed, the thesis must include a statement that makes the student’s contribution clear to the examiners. This may be in the form of a description of the precise contribution of the student to the work presented for examination and/or a statement of the percentage of the work that was done by the student. In addition, in the case of co-authored publications included in the thesis, each author must give their signed permission for the work to be included. If signatures from all the authors cannot be obtained, the statement detailing the student’s contribution to the work must be signed by the coordinating supervisor. Please sign one of the statements below.

1. This thesis does not contain work that I have published, nor work under review for publication. Student Signature .........................................................................................................................................................

2. This thesis contains only sole-authored work, some of which has been published and/or prepared for publication under sole authorship. The bibliographical details of the work and where it appears in the thesis are outlined below. Student Signature .........................................................................................................................................................

3. This thesis contains published work and/or work prepared for publication, some of which has been co-authored. The bibliographical details of the work and where it appears in the thesis are outlined below. The student must attach to this declaration a statement for each publication that clarifies the contribution of the student to the work. This may be in the form of a description of the precise contributions of the student to the published work and/or a statement of percent contribution by the student. This statement must be signed by all authors. If signatures from all the authors cannot be obtained, the statement detailing the student’s contribution to the published work must be signed by the coordinating supervisor.

Student Signature …………………………………………………………………………………………. Coordinating Supervisor Signature. ..……………………………………………………………………

Contents

Declaration i Acknowledgements ii Awards iii Publications iv List of Figures vi List of Tables ix Abbreviations x Abstract xvi 1.0 General Introduction 1 1.1 Prostate Cancer 1

1.1.1 Prostate Cancer Incidence and Mortality 1

1.1.2 Prostate Cancer Risk Factors 2

1.1.3 Prostate Cancer Diagnosis and Treatment 2

1.1.4 Castration Resistant Prostate Cancer 4

1.1.5 Molecular Alterations in Prostate Cancer 5

1.2 NKX3.1 7

1.2.1 NKX3.1 Protein Binding Partners 9

1.2.2 NKX3.1 Target Genes 11

1.2.3 Regulation of NKX3.1 Gene Expression 13

1.3 RMND5 Proteins 16

1.4 Ubiquitin and Ubiquitin-like Proteins 16

1.4.1 Ubiquitin Cascade 17

1.5 Ubiquitin Activating Enzymes (E1) 19

1.5.1 Discovery of E1 Enzymes 19

1.5.2 Structure and Function of E1 Enzymes 20

1.6 Ubiquitin Conjugating Enzymes (E2) 22

1.6.1 The Structure of E2 Enzymes 22

1.6.2 Classification of E2 Conjugating Enzymes 25

1.6.3 E2 Conjugating Enzyme Interactions with E1 and E3 Enzymes 26

1.6.4 Roles of E2 Enzymes in Ubiquitin Chain Formation 27

1.6.4.1 Chain Initiating and Chain Elongating E2 Enzymes 27

1.6.4.2 E2 Enzyme Processivity 28

1.3.4.3 E2 Enzyme Ubiquitin Chain Assembly and Linkage Selection 29

1.7 E3 Ubiquitin Ligases 30

1.7.1 HECT Domain E3 Ubiquitin Ligases 31

1.7.1.1 Structure and Mechanisms of Ubiquitin Transfer by HECT

Domains 32

1.7.1.2 Regulation of HECT E3 Ubiquitin Ligases 33

1.7.2 RING Domain E3 Ubiquitin Ligases 35

1.7.2.1 RING Domain Structure 36

1.7.2.2 Mechanism of Ubiquitin Transfer by RING Domain E3s 38

1.7.2.3 Regulation of RING E3 Ubiquitin Ligases 39

1.7.2.4 Single Subunit RING E3 Ubiquitin Ligases 39

1.7.2.5 Multisubunit RING E3 Ubiquitin Ligases 40

1.8 Outcomes of Ubiquitination 42

1.8.1 Ubiquitin Chain Topology Determines Ubiquitinated Protein Fate 42

1.8.2 Ubiquitin Binding Domains Determine Ubiquitinated Protein

Outcome 45

1.9 E3 Ubiquitin Ligases and Cancer 46

1.9.1 E3 Ubiquitin Ligases and the Cell Cycle 46

1.9.2 E3 Ubiquitin Ligases and DNA Damage 47

1.9.2.1 Tumour Suppressor p53 47

1.9.2.2 BRAC1/BARD1 49

1.9.3 E3 Ubiquitin Ligases and Signal Transduction 49

1.10 Statement of Aims 51

2.0 Materials 53

2.1 Reagents 53

2.1.1 Cell Culture 53

2.1.2 Primers 53

2.1.3 Reverse Transcription - Polymerase Chain Reaction (PCR) 54

2.1.4 Plasmids 54

2.1.5 Cloning 55

2.1.6 GST Fusion Protein Production 55

2.1.7 Immunoprecipitation 55

2.1.8 Western Blotting 56

2.1.9 Fluorescence Microscopy 56

2.1.10 General Laboratory Reagents 56

2.2 Commercial Kits 57

2.3 Equipment 59

2.4 Computer Software 60

3.0 Methods 61

3.1 Cell Culture 61

3.1.1 Routine Maintenance of Mammalian Cell Lines 61

3.1.2 Cryopreservation and Thawing of Mammalian Cells 61

3.1.3 Preparation of Cells for Fluorescence Microscopy 62

3.1.4 Transfection of Mammalian Cells 62

3.1.5 Treatment of Mammalian Cells 63

3.2 RNA Extraction and DNase Treatment 64

3.2.1 RNA Extraction 64

3.2.2 DNase Treatment of RNA 65

3.3 Reverse Transcription 65

3.4 Polymerase Chain Reaction (PCR) 65

3.4.1 PCR 65

3.4.2 “A” Tailing of PCR Products 67

3.4.3 Site Directed Mutagenesis 67

3.4.3.1 Mutagenesis PCR 67

3.5 Spectrophotometric Quantitation of RNA/DNA 68

3.6 Agarose Gel Electrophoresis 68

3.7 DNA Purification 68

3.7.1 Purification of DNA 68

3.7.2 Gel Purification of DNA 69

3.8 Cloning of PCR Products 69

3.8.1 Plasmids 69

3.8.2 Restriction Enzyme Digestion of Plasmids 70

3.8.3 Shrimp Alkaline Phosphatase Digestion 70

3.8.4 Ligation Reactions 70

3.8.5 Preparation of Competent Bacterial Cells 75

3.8.5.1 Preparation of Competent Escherichia coli DH5α 75

3.8.5.2 Preparation of Competent Escherichia coli BL21 75

3.8.6 Transformation of Bacterial Cells 75

3.8.7 Preparation of Bacterial/Glycerol Stocks 76

3.9 Small Scale Plasmid Purification 76

3.10 Large Scale Plasmid Purification 77

3.11 GST Fusion Protein Production and Purification 77

3.11.1 Small Scale Production of GST Fusion Proteins 77

3.11.2 Large Scale GST Fusion Protein Production 79

3.11.3 Total Protein Extraction from E. coli BL21 Cells 80

3.12 DNA Sequencing 80

3.13 Immunoprecipitation 81

3.14 Ubiquitin Assays 82

3.14.1 In Vitro Ubiquitin Assay 82

3.14.2 In Vivo Ubiquitin Assay 83

3.15 Western Blotting 84

3.15.1 Preparation of Whole Cell Lysates 84

3.15.2 Polyacrylamide Gel Electrophoresis 84

3.15.3 Western Transfer 85

3.15.4 Immunoblotting (Western Blotting) 85

3.15.5 Coomassie Blue Staining 86

3.16 Microscopic Imaging of Cells 86

3.16.1 Preparation of Slides for Fluorescence Microscopy 86

3.16.2 Preparation of Slides for Immunofluorescence Microscopy 87

3.16.3 Fluorescence Microscopy 87

3.17 Mass Spectrometry 88

4.0 Characterisation of RMND5 E3 Ubiquitin Ligase Activity 90

4.1 Introduction 90

4.1.1 Yeast RMD5/Gid2 90

4.1.2 Human RMND5 Proteins 94

4.1.2.1 RMND5A 94

4.1.2.2 RMND5B 94

4.1.3 Protein Domains 94

4.1.3.1 Lissencephaly 1 Homology Motif (LisH) 94

4.1.3.2 C-Terminal to LisH (CTLH) Domain 98

4.1.3.3 CT11-RanBPM (CRA) Domain 99

4.1.3.4 Really Interesting New Gene (RING) Domain 99

4.1.4 RMND5 Proteins and Cancer 99

4.2 Results 101

4.2.1 Bioinformatics Analyses of RMND5 Protein Architecture 101

4.2.2 Cloning of Full Length RMND5A into pGEX-2TK and Expression of

GST-RMND5 Proteins for In Vitro Ubiquitination Assays 101

4.2.2.1 Cloning of RMND5A into pGEX-2TK 101

4.2.2.2 Small Scale Production of GST, GST-RMND5A and

GST-RMND5B 106

4.2.2.3 Large Scale Production of GST-RMND5A 106

4.2.3 Cloning of RMND5 RING Domains for In Vitro Ubiquitination Assays 107

4.2.3.1 Cloning of RMND5 Proteins into pGEX-2TK 107

4.2.3.2 pGEX-RING Domain Protein Expression 112

4.2.4 In Vitro Auto-Ubiquitination Assays 113

4.2.4.1 Optimisation of In Vitro Auto-Ubiquitination Assays using the

GST-CBL RING Domain 113

4.2.4.2 In Vitro Auto-Ubiquitination Assays Using the GST-RMND5A

and GST-RMND5B RING Domains 116

4.2.4.3 Screening of E2 Conjugating Enzymes in In Vitro Ubiquitination

Assays 116

4.2.4.4 Control In Vitro Auto-Ubiquitination Assays 119

4.2.5 In Vivo Ubiquitination Assays 119

4.2.6 Investigation of the E3 Ubiquitin Ligase Activity of the

RMND5A and RMND5B RING Domains using RMND5A (C356S) and

RMND5B (C358S) RING Domain Mutants 121

4.2.6.1 Introduction of C356S into the RMND5A RING Domain 121

4.2.6.2 Introduction of C358S into the RMND5B RING Domain 127

4.2.6.3 Cloning of the RMND5A (C356S) and RMND5B (C358S) RING

Domains into pGEX-2TK 131

4.2.6.4 Expression and Intracellular Localisation of RMND5A

(C356S) and RMND5B (C358S) 134

4.2.6.5 In Vivo Ubiquitination Activity of RMND5A (C356S) and

RMND5B (C358S) 137

4.2.6.6 In Vitro Auto-Ubiquitination Activity of RMND5A

(C356S) and RMND5B (C358S) RING Domains 139

4.2.7 Examination of the E3 Ubiquitin Ligase Activity of the

RMND5A and RMND5B RING Domains by the Introduction

of Dual Mutations in the RMND5 RING Domain 143

4.2.7.1 Mutation of the RMND5A (C358A/H360A) RING Domain 145

4.2.7.2 Cloning of the RMND5A (C356A/H358A) RING Domain into

pGEX-2TK 150

4.2.7.3 Mutation of the RMND5B (C358A/H360A) RING Domain 152

4.2.7.4 Cloning of the RMND5B (C358A/H360A) RING Domain into

pGEX-2TK 153


(C356A/H358A) and RMND5B (C358A/H360A) 157

4.2.7.6 In Vivo Ubiquitination Activity of RMND5A

(C356A/H358) and RMND5B (C358A/H360A) Mutant Proteins 157

4.2.7.7 In Vitro Auto-Ubiquitination Activity of RMND5A

(C356A/H358A) and RMND5B (C358A/H360A) RING Domains 163

4.3 Discussion 170

5.0 RMND5 Proteins Ubiquitinate NKX3.1 188

5.1 Introduction 188

5.2 Results 193

5.2.1 RMND5 Proteins Interact with NKX3.1 in LNCaP Prostate Cancer

Cells 193

5.2.1.1 RMND5A Interacts with NKX3.1 193

5.2.1.2 RMND5B Interacts with NKX3.1 193

5.2.2 RMND5 Proteins Colocalise with NKX3.1 in LNCaP Cells 195

5.2.3 Regulation of NKX3.1 Expression in Prostate Cancer Cells 196

5.2.4 RMND5 Protein Effects on NKX3.1 Protein Expression 203

5.2.4.1 RMND5A and RMND5B Reduce NKX3.1 Protein Levels 203

5.2.4.2 RMND5A (C356S and C356A/H358A) and RMND5B

(C358S and C358A/H360A) Reduce NKX3.1 Protein Levels 206

5.2.5 NKX3.1 is Ubiquitinated in LNCaP Cells 208

5.2.5.1 RMND5 Proteins Ubiquitinate NKX3.1 208

5.2.5.2 RMND5A (C356A/C358A) and RMND5B (C358A/H360A)

Ubiquitinate NKX3.1 210

5.3 Discussion 213

6.0 Characterisation of RMND5 Protein Binding Partners 223

6.1 Introduction 223

6.1.1 Characterisation of the CTLH Complex Components 223

6.1.2 Protein Domain Architecture of the CTLH Complex Members 224

6.1.3 The Yeast Vid30 Complex 227

6.1.4 CTLH Complex Components 229

6.1.4.1 Muskelin 230

6.1.4.2 ARMC8α 231

6.1.4.3 RanBPM 232

6.1.4.4 EMP 233

6.2 Results 236

6.2.1 Transcripts Encoding the CTLH Complex Components are

Expressed in Prostate Cancer Cells 236

6.2.2 Cloning of RanBPM 239

6.2.2.1 Cloning of Full Length RanBPM (90kDa) 239

6.2.2.2 Cloning of the RanBPM 55kDa Isoform into pmCherry-C1 243

6.2.3 Interaction between RanBPM and RMND5A/RMND5B 245

6.2.3.1 RMND5A Interaction with RanBPM (55kDa) 245

6.2.3.2 RMND5B Interaction with RanBPM (55kDa) 247

6.2.3.3 RanBPM (55kDa) Interaction with RMND5 proteins 247

6.2.4 Colocalisation of RanBPM with RMND5A and RMND5B 249

6.2.5 Interaction Between RMND5A and RMND5B 251

6.2.5.1 Cloning of RMND5B into pmCherry-C1 251

6.2.5.2 RMND5A and RMND5B Interact in LNCaP Cells 257

6.2.5.3 RMND5A and RMND5B Colocalise in LNCaP Cells 257

6.2.6 Mass Spectrometric Identification of RMND5 Binding Partners 259

6.2.6.1 Identification of RMND5A Binding Proteins 259

6.2.6.2 Investigation of a Putative RMND5A Binding Partner 265

6.2.8.3 Identification of RMND5B Binding Proteins 268

6.3 Discussion 273

7.0 General Discussion 285

7.1 Discussion 285

7.2 Future Directions 293

8.0 References 302 Appendix I: Buffers and Solutions 350 Appendix II: Primer Sequences 366 Appendix III: Sequencing 369 Appendix IV: Mass Spectrometry Mascot Data 381

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Declaration I certify that this thesis does not incorporate, without my acknowledgement, any

material previously submitted for a degree or diploma from any university and that to

the best of my knowledge and belief, does not contain any material previously

published or written by another person except where due reference is made in text.

Candidate: Alison Louw ___________________ Date: ________ Supervisors: Dr Jacqueline Bentel ___________________ Date: ________ Winthrop Professor Jennet Harvey ___________________ Date: ________

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Acknowledgements There are so many people who I would like to thank and without whom this thesis would not have eventuated if not for their support, guidance and distraction! Firstly, I would like to sincerely thank my supervisors Jacky and Jennet. Jacky, I have learned so much from you and I would like to thank you for your mentorship, guidance and advice over the past few years. To Jennet, thank you for your positive advice and much needed perspective when required! To my good friend and lab mate Jasmine, I had a great time shopping, eating great food, drinking cocktails, laughing (and crying) with you! You always provided sound advice and reminded me that I was not alone in this and I hope we will remain good friends for a long time to come. To Lisa, I will remember all of our conversations and the laughs in the bug lab (my refuge!), breakfasts/lunches and your much loved cupcakes. Diana and Dian, thanks for taking the time to listen and offer advice (with Diana’s always starting, “You know Alison, you should just…”) and support – I know I can always count on you! Marc, you were always available to answer any questions and offer much needed advice and perspective – both work and running related! Thank you. To the current lab members, Jamie, Vivian and Abbie, and past members, Darren, Ebony, Agata, Cheryl, Chris, Lily and Ivy thanks for the conversations, including but not limited to experimental advice and entertaining stories about experimental fails (aka “optimisation”) reminding me that I am not the only one to make stupid mistakes (usually essential to the experiment’s success)! I have enjoyed getting to know every one of you and I have learned so much from everyone. I will cherish the good times, which were many – what ever happened to Friday morning coffee (or hot/iced chocolate)? A ritual that definitely needs to be resurrected I think! To my wonderful family, my Moms, Janet, Johan, Amber, Chelsey and my grandparents, I would never have made it this far without your love and never ending support through good and bad. Thank you for listening to my practice talks – even though you probably didn’t know what I was talking about always pointing to smears and bands all the time! – if it was important to me you were always there to help (and agree with me about the all-important colour schemes). Last but not least, the sixth family member my Chippey, thanks for the welcome feathery distraction and chats – even though I know most of the time you were only interested in my food or colourful shoes! I will be forever grateful to pump classes and running training for the chance to vent my frustrations about science in general and certain experiments in particular (which were seemingly endless), I have retained my sanity! To ASOS etc and the city of Perth, I appreciate the evening and lunch time retail therapy that was badly needed to gain some much needed clarity! Thank you too to the Head of School and Graduate Research Coordinator of the School of Pathology and Laboratory medicine and the members of the Research Centre next door, especially Kay for putting up with me always reporting broken equipment (I promise I didn’t break all that stuff!). Again to everyone, I say thank you and I hope to keep in touch in the future - Alison

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Awards During my studies I have been the recipient of a Prescott Postgraduate Scholarship from

the University of Western Australia.

I was the recipient of the Australia and New Zealand Society for Cell and

Developmental Biology Poster Prize for my poster presentation entitled, “ RMND5

Proteins Target the Prostatic Tumour Suppressor, NKX3.1 for Proteasomal

Degradation” at the Combined Biological Sciences Meeting in Perth, Australia , August

2012

I received the Murdoch University Prize for my oral presentation entitled, “RMND5 E3

Ubiquitin Ligases Ubiquitinate the Prostatic Tumour Suppressor, NKX3.1 Targeting it

for Proteasomal Degradation” at the Australian Society for Medical Research (ASMR)

Medical Research Week WA Student Symposium in Perth, Australia, June 2012

I was the recipient a Protein Synthesis, Targeting and Quality Control Best Thematic

Poster Award for my poster presentation entitled, “RMND5 Proteins Function as E3

Ubiquitin Ligases in Prostate Cancer Cells” at the American Society for Biochemistry

for Biochemistry and Molecular Biology in San Diego, USA, April 2012

I received the Murdoch University Prize for my oral presentation entitled, “RMND5

Proteins Function as E3 Ubiquitin Ligases in Prostate Cancer Cells” at the Australian

Society for Medical Research (ASMR) Medical Research Week WA Student

Symposium in Perth, Australia, June 2011

I was the recipient of the Scientific Encouragement Award for my oral presentation

entitled, “The Functional Characterisation of Human RMND5 Proteins in Prostate

Cancer Cells” at the Royal Perth Hospital Young Investigators Day in Perth, Australia,

August 2010

I received the Australian Society for Biochemistry and Molecular Biology Poster Prize

for a poster presentation entitled, “Functional Characterisation of Human RMND5

Proteins in Prostate Cancer Cells” at the Combined Biological Sciences Meeting

(CBSM) in Perth, Australia, August 2010

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Publications Conference Abstracts/Presentations Louw, A. , Thomas, M., Harvey, J., and Jacqueline Bentel (2012) “RMND5 Proteins

are Novel E3 Ubiquitin Ligases that Target the Prostatic Tumour Suppressor,

NKX3.1” ComBio 2012, Adelaide, Australia September 2012

Louw, A. Harvey, J., and Jacqueline Bentel (2012) “RMND5 Proteins Target the

Prostatic Tumour Suppressor, NKX3.1 for Proteasomal Degradation”

Combined Biological Sciences Meeting, Perth, Australia, August 2012

Louw, A, Harvey, J., and Jacqueline Bentel (2012) “RMND5 E3 Ubiquitin Ligases

Ubiquitinate the Prostatic Tumour Suppressor, NKX3.1 Targeting it for

Proteasomal Degradation”, Australian Society for Medical Research WA

Student Symposium, Perth, Australia, June 2012.

Louw, A, Harvey, J., and Jacqueline Bentel (2012) “RMND5 Proteins Function as E3

Ubiquitin Ligases in Prostate Cancer Cells”, American Society for

Biochemistry and Molecular Biology (ASBMB) Annual Meeting, San Diego,

CA, April 2012.


Ubiquitin Ligases in Prostate Cancer Cells”, Royal Perth Hospital Young

Investigators Day, Perth, Australia, November 2011

Louw, A, Harvey, J., and Jacqueline Bentel (2011) “Interaction of RMND5 Proteins

with the Prostatic Tumour Suppressor, NKX3.1”, Combined Biological

Sciences Meeting, Perth, Australia, August 2011


Ubiquitin Ligases in Prostate Cancer Cells”, Australian Society for Medical

Research WA Student Symposium, Perth, Australia, June 2011.

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Louw, A, Harvey, J., and Jacqueline Bentel (2011) “Human RMND5 Proteins Function

as E3 Ubiquitin Ligases in Prostate Cancer Cells”, Ubiquitin Satellite Meeting,

Melbourne, Australia, February 2011.

Louw, A, Harvey, J., and Jacqueline Bentel (2010) “The Functional Characterisation of

Human RMND5 Proteins in Prostate Cancer Cells”, 12th IUBMB OZBIO

Conference, Melbourne, Australia, October 2010.


Human RMND5 Proteins in Prostate Cancer Cells”, Royal Perth Hospital

Young Investigators Day, Perth, Australia, August 2010


Human RMND5 Proteins in Prostate Cancer Cells”, Combined Biological

Sciences Meeting, Perth, Australia, August 2010

Louw, A, Harvey, J., and Jacqueline Bentel (2010) “Functional Characterisation of

Human RMND5 Proteins in Prostate Cancer Cells”, Australian Society for

Medical Research WA Student Symposium, Perth, Australia, June 2010.

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List of Figures

1.0 General Introduction 1

Figure 1.1: Prostate cancer incidence 1

Figure 1.2: Chromosomal losses associated with human prostate cancer

initiation and progression 7

Figure 1.3: The ubiquitination cascade 18

Figure 1.4: Domain structure and sequence conservation of UBE1 and UBA6 21

Figure 1.5: Mechanism of ubiquitin activation by E1 ubiquitin conjugating

Enzymes 21

Figure 1.6: Three dimensional structure of E2 enzyme ubiquitin

conjugating domains 23

Figure 1.7: E2 ubiquitin chain linkage selection model 30

Figure 1.8: HECT domain structure 33

Figure 1.9: Regulation of HECT domain E3 catalytic activity 34

Figure 1.10: Mechanism of ubiquitin transfer by HECT and RING domain

containing E3 ubiquitin ligases 35

Figure 1.11: RING domain structure 36

Figure 1.12: RING and U-box domain amino acid residues involved in E2

enzyme interaction 37

Figure 1.13: Multisubunit E3 ubiquitin ligases 40

Figure 1.14: APC and SCF multisubunit E3 ubiquitin ligases 41

Figure 1.15: The type of ubiquitination determines substrate protein fate 43

Figure 1.16: p53 regulation in response to genotoxic stress 48

Figure 1.17: Roles of CBL in the regulation of receptor tyrosine kinase

Signalling 51

3.0 Methods 61

Figure 3.1: Map of the pGEM®-T Easy cloning vector 71

Figure 3.2: Map of the pEGFP-C2 expression vector 72

Figure 3.3: Map of the pmCherry C1 expression vector 73

Figure 3.4: Map of the pGEX-2TK bacterial expression vector 74

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Figure 4.1: Proposed mechanism of action of the Vid30 complex 92

Figure 4.2: Protein domain architecture of RMND5 proteins 102

Figure 4.3: Cloning of full length RMND5A into pGEX-2TK 104

Figure 4.4: Expression and purification of full length GST-RMND5A

and GST-RMND5B 108

Figure 4.5: Cloning of the RING domains of RMND5A, RMND5B and CBL

into pGEX-2TK 110

Figure 4.6: Expression and purification of the GST-RING domains of

RMND5A, RMND5B and CBL 114

Figure 4.7: Optimisation of in vitro ubiquitination assays 117

Figure 4.8: RMND5 RING domains mediate ubiquitin transfer with

specific E2 conjugating enzymes 118

Figure 4.9: Control reactions for in vitro ubiquitination assays 120

Figure 4.10: In vivo ubiquitination activity of RMND5 proteins 122

Figure 4.11: Site directed mutagenesis of the RING domains of

RMND5A and RMND5B 123

Figure 4.12: Preparation of pEGFP-RMND5A (C356S) 125

Figure 4.13: Preparation of pEGFP-RMND5B (C358S) 129

Figure 4.14: Cloning the RMND5A (C356S) and RMND5B (C358S) RING

domains into pGEX-2TK 132

Figure 4.15: Expression and cellular localisation of GFP-RMND5A (C356S)

and GFP-RMND5B (C358S) proteins 135

Figure 4.16: In vivo ubiquitination activity of RMND5A (C356S) and

RMND5B (C358S) 138

Figure 4.17: Expression and purification of GST-RMND5A (C356S) RING

and GST-RMND5B (C358S) RING domains 140

Figure 4.18: In vitro ubiquitination activity of GST-RMND5A (C356S)

and GST-RMND5B (C358S) RING domains 144

Figure 4.19: Site directed mutagenesis of the RING domains of RMND5A

and RMND5B to produce RMND5A (C356A/H358A) and

RMND5B (C358A/H360A) 146

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Figure 4.20: Generation of RMND5A (C356A/H358A) by site directed

mutagenesis and cloning of pEGFP-RMND5A (C356A/H358A) 148

Figure 4.21: Cloning of the RMND5A (C356A/H358A) RING domain into

pGEX-2TK 151

Figure 4.22: Generation of RMND5B (C358A/H360A) by site-directed

mutagenesis and cloning into pEGFP-C2 154

Figure 4.23: Cloning of sequences encoding the RMND5B (C358A/H360A)

RING domain into pGEX-2TK 156

Figure 4.24: Expression and cellular localisation of GFP-RMND5A

(C356A/H358A) and GFP-RMND5B (C358A/H360A) 158

Figure 4.25: In vivo ubiquitination activity of GFP-RMND5A (C356A/H358A)

and GFP-RMND5B (C358A/H360A) 161

Figure 4.26: Expression and purification of GST-RMND5A (C356A/H358A)

RING and GST-RMND5B (C358A/H360A) RING 164

Figure 4.27: Optimisation of in vitro ubiquitination assays for

GST-RMND5A (C356A/H358A) RING and GST-RMND5B

(C358A/H360A) RING 168

Figure 4.28: In vitro ubiquitination activity of GST-RMND5A (C356A/H358A)

and GST-RMND5B (C358A/H360A) 169

5.0 RMND5 Proteins Ubiquitinate NKX3.1 188

Figure 5.1: Post-translational modification of NKX3.1 190

Figure 5.2: NKX3.1 interacts with RMND5A and RMND5B in prostate cancer

Cells 193

Figure 5.3: RMND5 proteins colocalise with NKX3.1 in LNCaP cells 197

Figure 5.4: Determination of NKX3.1 half-life 199

Figure 5.5: Degradation of NKX3.1 by the proteasome 200

Figure 5.6: Lysosomal processing of NKX3.1 202

Figure 5.7: Overexpression of RMND5A or RMND5B reduces NKX3.1 levels 204

Figure 5.8: Proteasome inhibition restores NKX3.1 protein levels following

RMND5 overexpression 205

Figure 5.9: Overexpression of wild-type and mutant RMND5 proteins reduces

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NKX3.1 levels 207

Figure 5.10: Ubiquitination of NKX3.1 in vivo by RMND5A and RMND5B 209

Figure 5.11: In vivo ubiquitination of NKX3.1 following overexpression of

wild-type and mutant RMND5 proteins 212

6.0 The CTLH Complex 223

Figure 6.1: Protein domain architecture of the CTLH complex components 225

Figure 6.2: Predicted Vid30 and CTLH complex topology 229

Figure 6.3: Optimisation of RMND5A, RMND5B, RanBPM, muskelin,

Twa1, EMP, ARMC8α and C17orf39 PCR conditions 237

Figure 6.4: The CTLH complex components are expressed in prostate and

breast cancer cells 240

Figure 6.5: Cloning of the RanBPM coding region 241

Figure 6.6: Cloning of RanBPM (55kDa) into the pGEM®-T Easy cloning

Vector 244

Figure 6.7: Preparation of the pmCherry-RanBPM (55kDa) expression

plasmid 246

Figure 6.8: RMND5A and RMND5B interact with RanBPM in LNCaP cells 248

Figure 6.9: Optimisation of immunoprecipitation reactions using

Protein G Sepharose 250

Figure 6.10: Cherry-RanBPM (55kDa) colocalises with GFP-RMND5A and

GFP- RMND5B 252

Figure 6.11: Cloning of RMND5B into pGEM®-T Easy 253

Figure 6.12: Cloning of RMND5B into pmCherry-C1 255

Figure 6.13: RMND5A and RMND5B interact and colocalise in LNCaP cells 258

Figure 6.14: Immunoprecipitation of GFP-RMND5A and its binding partners 261

Figure 6.15: Identification of putative GFP-RMND5A binding partners 267 Figure 6.16: Immunoprecipitation of GFP-RMND5B and its binding partners 271

x

List of Tables

1.0 General Introduction 1

Table 1.1 – Human ubiquitin conjugating enzymes 24

3.0 Methods 61

Table 3.1 – Cell seeding density and reagents for transfection of mammalian

cells 63

Table 3.2 – LNCaP cell treatments 64

Table 3.3 – Addition of DNA polymerases to PCRs 66

Table 3.4 – PCR conditions 66

Table 3.5 – Site directed mutagenesis PCR conditions 67

Table 3.6 – Beads and buffers utilised for immunoprecipitation reactions 82

Table 3.7 – In vitro auto-ubiquitination assay components 83

Table 3.8 – In vivo ubiquitination assay plasmid combinations 84

Table 3.9 – Primary and secondary antibodies and their respective dilutions 86

Table 3.10 – Excitation and emission wavelengths of fluorescent labels 88


Table 4.1 – Human LisH domain containing proteins 95

Table 4.2 – Optimisation of in vitro ubiquitination assay enzyme

Concentrations 166

6.0 Characterisation of RMND5 Protein Binding Partners 223

Table 6.1 – The human CTLH complex components and their yeast orthologues 228

Table 6.2 – Candidate RMND5A binding partners identified by mass

spectrometry 263

Table 6.3 – Function of RMND5A binding proteins identified by mass

Spectrometry 266

Table 6.4 – Putative RMND5B binding partners identified by mass

Spectrometry 270

xi

Abbreviations

°C Degrees Celcius aa Amino Acids AJCC American Joint Committee on Cancer ALLN N-Acetyl-Leu-Leu-Nle-CHO APC/C Anaphase Promoting Complex/Cyclosome APS Ammonium Persulphate AR Androgen Receptor ARF ADP Ribosylation Factor ARF-BP1 ARF-Binding Protein 1 (alias MULE/HUWE1) Arg Arginine ARMC8α/β Armadillo Repeat Containing α/β AS Antisense ATP Adenosine Triphosphate BARD1 BRCA1 Associated RING Domain 1 BCL-2 B Cell Leukemia BDNF Brain-Derived Neurotrophic Factor BIRC6 Baculoviral Inhibitor of Apoptosis Repeat Containing

Protein 6 BLAST Basic Local Sequence Alignment bp Base Pairs BRCA1 Breast Cancer Susceptibility Gene 1 BRCC BRCA1-BRCA2 Containing Complex BRET Bioluminescence Resonance Energy Transfer BRUCE Baculovirus Inhibitor of Apoptosis Repeat Containing

Ubiquitin Conjugating Enzyme (alias BIRC6/Apollon) BSA Bovine Serum Albumin β-TrCP β-Transducin Repeat Containing Protein C2 Ca2+ Binding Motif Ca2Cl Calcium Chloride CBL Casitas B-Lineage Lymphoma CCD Catalytic Cysteine Domain Cdc20 Cell Division Cycle Protein 20 Cdh1 Cadherin 1 CDHC Cytoplasmic Dynein Heavy Chain Cdk Cyclin Dependent Kinase cDNA Complementary DNA CDS Coding Sequence CHIP Carboxyl Terminus of Hsc70 Interacting Protein CK2 Casein Kinase 2 c-MDH Cytoplasmic Malate Dehydrogenase CO2 Carbon Dioxide CRA CT11-RanBPM CRL4 Cullin4A-RING E3 Ubiquitin Ligase CSTF1 Cleavage Stimulation Factor 1 C-terminal Carboxy-Terminal CTLH C-Terminal to LisH Cul Cullen Protein Cys Cysteine DAPK Death Associated Protein Kinase DCAF-1 DDB1 and Cul4 associated factor 1

xii

ddH2O Deionised Water DEPC Diethylpolycarbonate DHT 5α-dihydrotestosterone DMSO Dimethylsulphoxide DNA Deoxyribonucleic Acid dNTP Deoxynucleoside Triphosphate DRE Digital Rectal Examination DSB Double Stranded Breaks DUB Deubiquitinating Enzyme E1 Ubiquitin Activating Enzyme E2 Ubiquitin Conjugating Enzyme E3 Ubiquitin Ligase E4 Conjugation Factor/Ubiquitin Chain Elongation Factor EBS ETS1 Binding Site EDTA Ethylenediaminetetraacetic Acid EEAP Early Endosome Associated Protein eGFP Enhanced Green Fluorescent Protein EGFR Epidermal Growth Factor Receptor ElaC2 ElaC Homologue 2 EMP Erythroblast Macrophage Protein (alias MAEA) ERAD Endoplasmic Reticulum Associated Degradation ERG Ets Related Gene ERK Extracellular Signal Regulated Kinase ESE3 Epithelium Specific Ets Transcription Factor 3 ETS Ethrythroblastosis Virus E26 ETS1 Erythroblastosis Virus E26 Oncogene Homolog 1 EZH2 Enhancer of Zeste Homologue 2 FANCD2 Fanconi Anaemia Group D2 FAT10 F Adjacent Transcript 10/ Human Leukocyte Antigen F

Associated FBPase Fructose 1,6 Bisphosphatase FDA US Food and Drug Administration FCS Fœtal Calf Serum FGFR1 Fibroblast Growth Factor Receptor 1 FMRP Fragile X Mental Retardation Protein FOP FGFR1 Oncogene Partner FRET Fluorescence Resonance Energy Transfer G1/2 Growth Phase1/ 2 (Cell Cycle) G2BR Ube2G2 Binding Region GABAAR Gamma-aminobutyric Acid A Receptor GAT domain GGA and Tom1 Domain Gid Glucose Induced Degradation GST Glutathione S Transferase GWAS Genome Wide Association Studies H1 Helix 1 H2O2 Hydrogen Peroxide HACE1 HECT and Ankyrin Repeat Containing E3 Ubiquitin

Ligase 1 HAUSP Herpes-virus Ubiquitin Specific Protease HDAC Histone Deacetylase HDAC1 Histone Deacetylase 1 HECT Homologous to E6-AP Carboxyl Terminus

xiii

HEK Human Embryonic Kidney Cells HeLa Henrietta Lacks Ovarian Carcinoma Cells HERC HECT and RCC1 Domain Containing Protein HGF Hepatocyte Growth Factor His Histidine HPN Hisitidine-Proline-Asparagine HPV Human Papilloma Virus HRP Horse Radish Peroxidase HRS Hepatocyte Growth Factor-Regulated Tyrosine Kinase

Substrate HSMpp8 Matrix Metalloprotease 8 HUWE1 HECT, UBE, WWE Domain Containing Protein 1 (alias

MULE/ARF-BP1) Hxt7p Hexose Transporter Protein 7 IGF1 Insulin-like Growth Factor 1 IGF1R IGF1 Receptor IGFBP3 IGF1 Binding Protein 3 IgG Immunoglobulin G IKK IKappaβ Kinase IP Immunoprecipitation ITCH Ubiquitin Protein Ligase Itchy Homologue JFK Just One F-box and Kelch Domain Containing Protein JNK c-Jun N-terminal Kinase kbp Kilobase Pairs kDa Kilo Daltons L4/7 Loop 4/7 LAMP1/2 Lysosome Associated Membrane Protein 1/2 LB Luria Bertani LHR Linker Helix Region LHRH Luteinising Hormone Releasing Hormone LIS1 Lissencephaly Protein 1 LisH Lissencephaly 1 Homology Domain LOH Loss of Heterozygosity LUBAC Linear Ubiquitin Chain Assembly Complex Lys Lysine MAEA Macrophage Erythroblast Attacher MAPK Mitogen Activated Protein Kinase MCS Multiple Cloning Site MDH2 Malate Dehydrogenase 2 MDM2 Murine Double Minute Homologue 2 MDMX Murine Double Minute Homologue X MEKK1 Mitogen Activated Portien/ERK Kinase Kinase 1 MG132 Carbobenzoxy-Leu-Leu-Leucinal MgCl2 Magnesium Chloride mL Millilitres mM Millimolar MMS2 Methyl Methanesulphonate Sensitive Gene Product 2 MOPS 3(N-Morpholino)propanesulphonic Acid mRNA Messenger Ribonucleic Acid MS Mass Spectrometry MSR1 Macrophage Scavenging Receptor 1 MULE Mcl-1 Ubiquitin Ligase E3 (alias HUWE1/ARF-BP1)

xiv

MYC Myelocytomatosis Viral Oncogene Homologue N Amino N4BP Nedd4-Binding Protein NaOH Sodium Hydroxide NCBI National Centre for Biotechnology Information N-CoR Nuclear Receptor Corepressor NEDD4/8 Neural Precursor Cell-Expressed, Developmentally Down

Regulated 4/8 NEMO NF-κB Essential Modulator NETN NP-40, EDTA, Tris, NaCl Buffer NF-κB Nuclear Factor Kappa Light Chain Enhancer of Activated

B Cells NHS Normal Horse Serum NP-40 Nonidet P-40 NPAT Nuclear Protein of the Ataxia Telangiectasia Mutated

Locus NSLC Non-Small Cell Lung Carcinoma NTD N-Terminal Domain OD Optical Density OFD1 Oral Facial Density ORF Open Reading Frame p75NTR p75 Neurotrophin Receptor PBGD Porphobilinogen Deaminase PBS Phosphate Buffered Saline PBST Phosphate Buffered Saline Tween-20 PCAN Prostate Cancer Gene 1 PCNA Proliferating Cell Nuclear Antigen PCR Polymerase Chain Reaction PEPCK Posphoenolpyruvate Carboxykinase PHD Plant Homeodomain PIN Prostatic Intraepithelial Neoplasia PIPES Piperazinediethanesulphonic Acid PKC Protein Kinase C pmol Pecomole PMSF Polymethylsulphonyl Fluoride POTEE Prostate Ovary Testis Expressed Protein E PPxY Proline-Proline-any amino acid- Tyrosine PRR Proline Rich Repeat PS Penicillin/Streptomcyin PSA Prostate Specific Antigen PTEN Phosphate and Tensin Homologue Deleted on

Chromosome Ten PTM Post-Translational Modification PUF60 Poly (U) Binding Splicing Factor RA Retinoic Acid RanBP9/10 Ran Binding Protein 9/10 RanBPM M Ran Binding Protein in the Microtubule Organising

Centre (RanBP9) RAR Retinoic acid Receptor RCC1 Regulator of Chromosome Condensation 1 RGG box Arginine (R), Glycine (G) Rich Region RHOT Ras homolog family member T2

xv

RING Really Interesting New Gene RIPA Radioimmune Precipitation Buffer RLD RCC1-like Domains RMD5 Required for Meiotic Nuclear Division 5 (Saccharomyces

cerevisiae) RMND5A Required for Meiotic Nuclear Division 5 homologue A RMND5B Required for Meiotic Nuclear Division 5 homologue B RNA Pol II RNA Polymerase II RNA Ribonucleic Acid rpm Revolutions Per Minute RPMI Roswell Park Memorial Medium RTK Receptor Tyrosine Kinase RT-PCR Reverse Transcription-Polymerase Chain Reaction S phase Synthesis Phase (DNA Replication Phase - Cell Cycle) S Sense SAP Shrimp Alkaline Phosphatase SCF Skip-Cullin-F-box SDS Sodium Dodecyl Sulphate SH2 SRC Homology 2 Siah Seven In Absentia Homologue Sic1 S-phase cyclin/cyclin-dependent kinase inhibitor Sif2p Sir4p Interacting Protein 2 SILAC Stable Isotope Labelling with Amino Acids in Culture siRNA Small Interfering RNA SKIP Ski Interacting Protein 1 SKP S-Phase Kinase-Associated Protein SMAC Second Mitochondria-Derived Activator of Caspases SMART Simple Modular Architecture Research Tool SMGA Smooth Muscle Gamma Actin SMRT Silencing Mediator for Retinoic and Thyroid Receptor SMURF SMAD Ubiquitination Regulatory Factor SNP Single Nucleotide Polymorphism SNW1 SNW Domain Containing Protein 1 (alias SKIP) SOCS Suppressor of Cytokine Signalling SPRY Spla and Ryanodine Receptor SRB Suppressor of RNA Polymerase B SRC Rous Sarcoma Viral Oncogene Homoloue SRE Serum Response Element SRF Serum Response Factor SUMO Small Ubiquitin Like Modifier TAD Transcriptional Activation Domains TAF5/6 Transcription initiation factor TFIID subunit 5/6 T-ALL T Cell Acute Lymphoblastic Leukemias Taq Thermus Aquaticus TBL1 X/Y Transducin β Like Protein 1 X/Y TBL1 Transducin β-like Protien 1 TBLR1 Transducin β-like Protein 1 Related Protein TBS Tris Buffered Saline TBST Tris Buffered Saline Tween-20 TCA Tricarboxylic Acid TCOF1 Treacher Collins-Franceschetti Syndrome 1 TCR T Cell Receptor

xvi

TE Tris/EDTA TKBD Tyrosine Kinase Binding Domain TNF Tumour Necrosis Factor TNM Tumour, Node and Metastasis TOPO I Topoisomerase I TOPORS Topoisomerase Binding RING Finger Protein TOR Target of Rapamycin TPR Tetrapeptide Repeat TRAF TNF Receptor Associated Factor TRITC Tetramethylisothiocyanate TSG Tumour Suppressor Gene TSP-1 Thrombospondin 1 Twa1 Two-Hybrid Associated Protein 1 U Units Uae/UBA Ubiquitin Activating Enzyme Ub Ubiquitin UBA Ubiquitin Associated Domain UBC Ubiquitin Conjugating Domain/Enzyme UBD Ubiquitin Binding Domain UBE Ubiquitin-Like Modifier Activator Enzyme Ubl Ubiquitin-Like Protein/Domain UBM Ubiquitin Binding Motif UBZ Ubiquitin Binding Zinc Finger UEV Ubiquitin Enzyme Variants UFD2 Ubiquitin Fusion Degradation UIM Ubiquitin Interacting Motif UPS Ubiquitin Proteasome System USP Ubiquitin Specific Protease UTR Untranslated Region UV Ultraviolet V Volts VEGF-C Vascular Endothelial Growth Factor C VEGF-C Vascular Endothelial Growth Factor C VHL Von Hippel Landau VHL-CBC VHL-cullin 2-elongin B-elongin C Vid Vacuolar Import and Degradation WD40 Tryptophan-Aspartic Acid (W-D) dipeptide WDR26 WD repeat containing protein 26 WW Tryptophan-Tryptophan WWE Tryptophan-Tryptophan-Glutamic Acid WWP1 WW Domain Containing Protein 1 Y/FPPxxP Tyrosine/Phenylalanine-Proline-Proline-any amino acid-

any amino acid-Proline

xvii

Abstract

RMND5A and RMND5B are highly homologous uncharacterised proteins named after

their yeast orthologue, Required for Meiotic Nuclear Division 5 (RMD5), a RING

domain-containing E3 ubiquitin ligase. RMND5B was originally identified in our

laboratory to interact with the prostatic tumour suppressor, NKX3.1, expression of

which is reduced or undetectable in up to 80% of metastatic prostate tumours.

Bioinformatics analyses of the cDNA and translated sequences of RMND5A and

RMND5B identified four protein-protein interaction domains, a Lissencephaly 1

homology (LisH), a C-terminal to LisH (CTLH), a CT11-RanBPM (CRA) and a Really

Interesting New Gene (RING) domain. Alignment of the RING domains of RMND5

proteins with that of yeast RMD5 identified that all eight amino acid residues essential

for RING domain folding and therefore activity were identical between the proteins,

suggesting that RMND5A and RMND5B function as E3 ubiquitin ligases. In vitro

ubiquitination assays carried out using the RING domains of RMND5A and RMND5B

and a panel of 11 E2 conjugating enzymes identified that RMND5A interacted with the

E2 enzymes UbcH2, UbcH5b and UbcH5c, whilst RMND5B associated with UbcH5b

and UbcH5c to mediate ubiquitin transfer to substrate lysine residues. Consistent with

this finding, full length RMND5 proteins were associated with ubiquitinated proteins in

vivo in LNCaP prostate cancer cells and this effect was augmented by proteasome

inhibition. Site-directed mutagenesis reduced the in vitro autoubiquitination activity of

the RMND5A (C356A/H358A) and RMND5B (C358A/H360A) RING domain

mutants, while in vivo, interaction of RMND5B (C358A/H360A) with ubiquitinated

proteins was decreased.

In prostate cancer cells, both RMND5A and RMND5B were found to

coimmunoprecipitate with NKX3.1 and overexpression of either RMND5A or

RMND5B enhanced NKX3.1 ubiquitination, resulting in a dose-dependent decline in

NKX3.1 protein levels that was reversed upon proteasome inhibition. These findings

indicated that RMND5 proteins interact with NKX3.1, promoting its ubiquitination and

proteasome-mediated degradation. RMND5A has been reported to form part of a large,

multi-protein complex, the CTLH complex, human orthologue of the yeast Vid30 E3

ubiquitin ligase complex which contains RMD5. All CTLH complex members were

shown to be expressed in prostate and breast cancer cell lines and RMND5A and

RMND5B were identified to interact and colocalise with RanBPM, a proposed core

xviii

component of the CTLH complex. These findings suggested that the CTLH complex is

able to form in prostate cancer cells potentially containing either or both RMND5

proteins. Mass spectrometry, used to identify proteins that interacted by

coimmunoprecipitation with RMND5A or RMND5B, resulted in the isolation of a

number of candidate binding partners including mitochondrial proteins such as ATPase

synthase subunits α and β, and nuclear proteins such as SNW1 and XRCC6. These

results suggest the involvement of RMND5 proteins in the regulation of other cellular

and metabolic processes, which may be investigated in future studies. This thesis has

therefore determined that both RMND5A and RMND5B function as E3 ubiquitin

ligases in prostate cancer cells and are able to target the prostatic tumour suppressor,

NKX3.1 for ubiquitination and proteasome-dependent degradation. As the RMND5

chromosomal loci are frequently disrupted in cancer, further characterisation of the

biological activity of RMND5A and RMND5B will elucidate their roles in normal

physiological processes and the contribution of their deregulated function to cancer

formation and progression.

Chapter 1 General Introduction

Chapter 1: General Introduction


1

1.0 General Introduction

The expression of factors involved in the development and maintenance of organs

including the prostate is meticulously regulated at all stages, from transcription to

translation, ensuring the appropriate temporal and spatial expression of each gene. Once

produced, the cell maintains tight control over protein function through a range of post-

translational modifications and protein-protein interactions, ensuring the precise activity

of each protein and its degradation when damaged or unnecessary. Deviation of any of

these regulatory pathways is therefore associated with a number of abnormalities

including aberrant gene expression and improper protein folding, activity or turnover,

potentially leading to the development of pathological states including cancer.

1.1 Prostate Cancer

1.1.1 Prostate Cancer Incidence and Mortality

Prostate cancer is the second most common cause of cancer and the sixth leading cause

of cancer related mortality of men worldwide, however in developed countries, prostate

cancer is typically the most commonly diagnosed cancer of men and the third leading

cause of cancer related mortality (Figure 1.1) (Ferlay et al., 2009). The higher incidence

of prostate cancer in western countries such as Australia, USA, UK, New Zealand and

Europe is well documented, and although the prostate cancer mortality rates in these

countries are declining, the mortality rates in less developed countries in Africa, Asia

and Eastern Europe are rising (Center et al., 2012).

Figure 1.1: Prostate cancer incidence. The incidence of prostate cancer is particularly high in developed countries including Australia/New Zealand and is lowest in Asia and Africa (Ferlay et al., 2010).


2

It is proposed that differences in prostate cancer incidence and mortality between

developed and non-developed countries are due to genetic and lifestyle factors as well

as differences in disease diagnosis and treatment availability (Center et al., 2012).

1.1.2 Prostate Cancer Risk Factors

A number of risk factors for prostate cancer development have been proposed including

diet, occupation and infection, however the best defined risk factors are age, family

history and ethnicity (Schottenfeld and Fraumeni, 2006; Brawley, 2012).

Approximately 30% of men over the age of 50 and 75% of men over the age of 80 years

exhibit evidence of prostate cancer at autopsy, while the median age at diagnosis of

clinical prostate cancer between 2001 – 2010 was 67 years of age (Brawley, 2012;

Eylert and Persad, 2012). Prostate cancer incidence rates have increased since the mid-

1980s and in 2005, the incidence rates of prostate cancer were 3.64 and 7.23 times

higher among males aged 50-59 and <50, respectively compared to those in 1986

(Brawley, 2012). The younger age at diagnosis is likely to be due in part to improved

screening and detection methods including measurement of circulating prostate specific

antigen (PSA) levels (Section 1.1.3), more sensitive imaging techniques and greater

public awareness of the disease (Damber and Aus, 2008). Although familial prostate

cancer syndromes are relatively rare, accounting for <10% of cases (Section 1.1.5), men

with an affected first degree relative have an estimated 2-3 fold increased risk for

development of the disease themselves, which is likely to result from both genetic and

lifestyle factors, including diet (Powell, 2011). African American men have a 1.4 times

higher risk of being diagnosed with prostate cancer than European American men and

are at a 2-3 times higher risk of dying from the disease, and this has been attributed to a

combination of dietary, genetic and socioeconomic factors (Chornokur et al., 2011;

Powell, 2011).

1.1.3 Prostate Cancer Diagnosis and Treatment

Screening and detection of prostate cancer can involve digital rectal examination (DRE)

and measurement of serum PSA levels, with many men asymptomatic during the early

stages of disease. PSA is present in both normal and malignant prostatic epithelial cells,

and although increased serum PSA levels may indicate the presence of prostate cancer,

other conditions such as prostatitis and benign prostatic hyperplasia may also be

associated with elevated serum levels of PSA (Porter and Brawer, 1993; Hochreiter,


3

2008). Abnormalities detected using DRE and PSA testing may be further investigated

by transurethral ultrasound guided needle biopsy to sample the prostate tissue for

histopathological analysis and diagnosis of prostate cancer (Greene et al., 2009; Bailey

and Brewster, 2011).

Treatment of prostate cancer depends upon the stage of the tumour, which can be

categorised using the American Joint Committee on Cancer (AJCC) Tumour, Node and

Metastasis (TNM) classification system (Edge et al., 2010). The Gleason score obtained

from histological examination of the tumour tissue is recognised as the preferred

grading system (Gleason and Mellinger, 1974; Greene and Sobin, 2002). Along with

this information, the patient’s age and general health are also important determinants for

assessing potential treatments. Watchful waiting or active surveillance is recommended

for those patients with low grade, small volume tumours and this is due to the side

effects associated with surgery or radiation therapy which include urinary incontinence

and erectile dysfunction (Singer et al., 2012). For organ confined prostate cancer, active

surveillance may also be employed. However in fitter younger men, radical

prostatectomy can be performed to surgically remove the prostate and this treatment is

reported in some studies to be more effective in the longer term for these patients

compared to surveillance (Bill-Axelson et al., 2005; Heidenreich et al., 2008; Wiltz et

al., 2009; Hugosson et al., 2011; Holmberg et al., 2012).

Androgen ablation and anti-androgen therapies may be used in the clinical management

of patients with high-risk localised disease (frequently in combination with

radiotherapy), for patients with elevated PSA levels after local treatment and to treat

patients with metastatic disease (Sharifi et al., 2010). The approach of androgen

depletion is based on the knowledge that prostate tumours are dependent on androgens,

which mediate their effects via the androgen receptor (AR), for their growth and

survival. Androgen deprivation treatment usually involves either surgical

(orchidectomy) or chemical castration using oestrogens (rarely), anti-androgens or

luteinising hormone releasing hormone (LHRH) agonists or antagonists to halt the

production of testosterone by the testes (Harris et al., 2009; Sharifi et al., 2010). LHRH

agonists and antagonists prevent testicular androgen production by antagonising the

release of pituitary gonadotropins, however LHRH agonists are associated with an

initial increase in gonadotropin and therefore testosterone production, leading to acute

growth of the tumour (“tumour flare”) until circulating testosterone levels fall (Harris et


4

al., 2009). To prevent tumour flare associated with initiation of LHRH agonist

treatment, non-steroidal AR antagonists such as bicalutamide, flutamide and more

recently enzalutamide (MDV3100) are used to block the actions of androgens

(Perlmutter and Lepor, 2007; Connolly et al., 2012; Scher et al., 2012). The

combination of LHRH agonists with anti-androgen treatments, particularly bicalutamide

is known as maximum/combined androgen blockade and is widely used for the

treatment of advanced prostate cancer (Connolly et al., 2012). In addition, treatments

such as ketoconazole which reduce adrenal androgen synthesis may be used in

combination with chemical and surgical castration to further decrease circulating

androgen levels (Perlmutter and Lepor, 2007). Treatments which prevent the synthesis

of androgens such as the CYP17A inhibitor, abiraterone acetate may be used in the

management of prostate cancer, however treatment resistance in the form of upregulated

CYP17A or mutated AR has already been noted (Cai et al., 2011; de Bono et al., 2011).

Inhibitors of 5α-reductase such as finasteride may also be used to block the production

of the potent AR activator 5α-dihdrotestosterone (DHT) from testosterone, reducing

prostate cancer growth (Nacusi and Tindall, 2011).

1.1.4 Castration Resistant Prostate Cancer

Hormonal therapies for prostate cancer are effective in the short term with remissions

typically lasting 2-3 years. However, most patients develop resistance to these

treatments, termed castration-resistant prostate cancer, and due to the lack of effective

therapies, patient survival from the time of progression is ~16-18 months (Pienta and

Bradley, 2006). Although prostate tumours become castration resistant they still make

use of the AR signalling pathway and a number of mechanisms by which prostate

cancer cells survive the androgen deprived environment have been proposed. These

include androgen hypersensitivity, in which prostate tumours circumvent the depletion

of androgens by developing the ability to respond to very low levels of circulating

androgens (Gregory et al., 2001b). Prostate cancer cells achieve this by increasing

expression of the AR, which may be due to gene amplification, by increased AR

sensitivity to androgens due to AR mutations, or in association with the intratumoral

production of androgens (Visakorpi et al., 1995a; Gregory et al., 2001b; Linja et al.,

2001; Chen et al., 2004; Locke et al., 2008). AR mutations, which have been reported to

occur in ~10% of prostate tumours following androgen deprivation therapy, may also

allow the receptor to bind nonandrogenic or nonsteroidal ligands or undergo ligand

independent activation, as evidenced by the identification of AR splice variants lacking


5

the ligand binding domain which are constitutively active (Veldscholte et al., 1992;

Zhao et al., 2000; Ueda et al., 2002; Taplin et al., 2003; Marques et al., 2005; Bluemn

and Nelson, 2012). Cross-talk between the AR and other pathways that are able to

activate the AR, for example HER2 and MAPK has been observed (Craft et al., 1999;

Ueda et al., 2002). In a proportion of castrate-resistant prostate cancers, the balance

between AR coactivators and repressors is altered and in particular, increased

coactivator levels have been found in these cells (Gregory et al., 2001a; Ngan et al.,

2003). Another mechanism by which castration resistance can arise is by the use of

alternative non-androgen mediated mechanisms, for example upregulation of

antiapoptotic factors including Bcl-2 has been reported in advanced castration resistant

prostate tumours (McDonnell et al., 1992; July et al., 2002; Pootrakul et al., 2006).

Chemotherapeutic agents such as docetaxel and cabazitaxel, immunotherapies such as

sipuleucel-T and CYP17 inhibitors including abiraterone acetate may be used for the

treatment of castrate-resistant prostate cancer, increasing survival by 2-5 months

(Rehman and Rosenberg, 2012; Gerritsen, 2012). Bone metastases are common in men

with castrate-resistant prostate cancer and treatments include radiotherapy to reduce

bone pain, bisphosphonates such as zoledronic acid and denosumab an antibody

treatment targeting RANK ligand (Shore et al., 2012).

1.1.5 Molecular Alterations in Prostate Cancer

In addition to AR signalling, aberrant expression of many other proteins, tumour

suppressor gene or oncogene products and genetic alterations are common in prostate

cancer and as in many cancers, these involve hereditary and sporadic mutations which

accumulate over time, driving prostate tumour initiation and progression or determining

treatment responses (Visakorpi, 2003). A number of susceptibility loci for prostate

cancer development have been identified by genome wide linkage studies in families

with a high prostate cancer incidence, and these include chromosome 1q24-25

(RNASEL), 17p (ElaC homologue 2 (ElaC2)), 20q13 and chromosome Xq (Smith et al.,

1996; Xu et al., 1998; Berry et al., 2000; Tavtigian et al., 2001; Wang et al., 2002b;

Adler et al., 2003; Wiklund et al., 2004). Germline mutations in the AR, BRCA1,

BRCA2, CYP1B1 and MSR1 genes have also been associated with prostate cancer

susceptibility (Dong, 2006). The identification of genes conferring a predisposition to

prostate cancer development has been challenging in part due to the late age of disease

onset, making it difficult to screen two or more generations for susceptibility loci and

therefore to discern which mutations are associated with disease development (Rubin


6

and De Marzo, 2004, Xu et al., 2005). In addition, the likelihood that mutations or

variants of multiple genes predispose to prostate cancer development rather than one or

two further complicates the identification of susceptibility loci (Rubin and De Marzo,

2004; Xu et al., 2005).

Sporadic chromosomal abnormalities or mutations in specific genes are commonly

present in prostate cancers, with the most common chromosomal abnormalities

including loss of heterozygosity at 8p, 10q, 13q and 17p and chromosomal gains at 7,

8q, 18q and Xq (Cher et al., 1994; Joos et al., 1995; Visakorpi et al., 1995b; Nupponen

et al., 1998; Dong, 2006). Genes within these loci associated with prostate cancer

initiation and progression include the homeobox gene NKX3.1 (8p21.2), the AKT

signalling regulator PTEN (10q), the Retinoblastoma gene Rb (13q), p53 (17p) and the

MYC proto-oncogene (8q24) (Abate-Shen and Shen, 2000; Visakorpi, 2003). More

recently identified common chromosomal aberrations in prostate cancer involve ETS

gene fusions, particularly the TMPRSS2-ERG gene fusion, which is present in

approximately 50% of localised prostate cancers and is proposed to result in the

overexpression of oncogenic transcription factors including ERG (Mosquera et al.,

2009; Tomlins et al., 2009). Consistent with a multistep theory of cancer development,

the loss of specific chromosomal regions is proposed to be associated with particular

stages of prostate cancer formation and disease progression (Figure 1.2) (Abate-Shen

and Shen, 2000). For example, the preneoplastic lesion, prostatic intraepithelial

neoplasia (PIN) is associated with the loss of chromosomal regions such as 8p

(NKX3.1), followed by tumour initiation and progression which are associated with the

loss of 10q (PTEN), 13q (Rb) and TMPRSS2 gene rearrangement, and invasion,

metastasis and progression to castration resistance mediated in part by the loss of 17p

(p53) (Figure 1.2) (Abate-Shen and Shen, 2000).

The advent of newer sequencing technologies including next generation sequencing

techniques has allowed the identification of specific single nucleotide polymorphisms

(SNP) or base substitutions prevalent in prostate tumours. For example, whole exome

sequencing has identified repeated somatic mutations in SPOP, FOXA1 and MED12,

with base substitutions in the cullin based E3 ubiquitin ligase SPOP present in 6-13% of

prostate tumours (Barbieri et al., 2012). In addition to chromosomal abnormalities and

gene mutations, interruption in caretaker gene expression such as the pi class of

Glutathione-S Transferases (GST) via promoter hypermethylation is present in a high


7

percentage of prostate tumours (He et al., 1997; Millar et al., 1999). Accumulating data

documenting the spectrum of molecular abnormalities present in prostate cancers may

identify prostate cancer subtypes and so increase understanding of the formation and

indolence or progression of individual prostate tumours. Depending on the major

genetic changes present, targeted therapies could be developed for treatment of specific

prostate cancer subtypes.

1.2 NKX3.1

NKX3.1 is a homeodomain transcription factor, expression of which is localised to the

prostatic epithelium and to a lesser extent, the testis (Prescott et al., 1998; Meeks and

Schaeffer, 2011). In mice, Nkx3.1 expression is the earliest marker of prostate

development and is required for the formation and maintenance of the prostate gland,

with disruption of Nkx3.1 expression during embryogenesis resulting in defects in

ductal morphogenesis and the abnormal production of secretory proteins (Sciavolino et

al., 1997; Bhatia-Gaur et al., 1999; Tanaka et al., 2000; Matusik et al., 2008). Nkx3.1

expression is maintained in adulthood and adult heterozygous (Nkx3.1+/-) and

homozygous (Nkx3.1-/-) mutant mice exhibit prostatic epithelial hyperplasia and

dysplasia which increases in severity with age to resemble the preneoplastic lesion, PIN

(Bhatia-Gaur et al., 1999; Abdulkadir et al., 2002; Kim et al., 2002a; Abdulkadir, 2005;

Abate-Shen et al., 2008). An interesting finding is that PIN lesions in heterozygous

Nkx3.1 mutant mice express undetectable levels of Nkx3.1 protein, suggesting that

Figure 1.2: Chromosomal losses associated with human prostate cancer initiation and progression. Each stage in prostate cancer formation and progression is associated with the disruption of specific chromosomal regions containing tumour suppressor genes and the accumulation of gene mutations or dysregulation of gene expression (Abate-Shen and Shen, 2000).


8

prostate carcinogenesis is facilitated by loss of Nk3.1 expression (Abdulkadir et al.,

2002). This is supported in cell culture studies where Nkx3.1 overexpression inhibits

cell proliferation, indicating a role for Nkx3.1 in tumour suppression (Kim et al.,

2002a). Although loss of Nkx3.1 expression alone is not sufficient for prostate tumour

formation, Nkx3.1 cooperates with the loss of expression of other tumour suppressor

gene products such as p27 and Pten to promote the development of invasive prostate

adenocarcinomas and tumour metastasis (Kim et al., 2002b; Abate-Shen et al., 2003;

Gary et al., 2004).

In humans, the NKX3.1 gene is located at the chromosomal locus, 8p21.2, a region that

undergoes loss of heterozygosity (LOH) in approximately 80% of advanced prostate

tumours, with NKX3.1 protein levels reduced or undetectable in up to 80% of

metastatic tumours (Emmert-Buck et al., 1995; Vocke et al., 1996; He et al., 1997;

Bowen et al., 2000; Aslan et al., 2006; Barnabas et al., 2011). LOH at 8p21.2 has also

been reported in 20—80% of high grade PIN lesions, indicating that NKX3.1 loss is an

early event in prostate carcinogenesis (Emmert-Buck et al., 1995; Haggman et al.,

1997; Bowen et al., 2000; Asatiani et al., 2005; Aslan et al., 2006; Bethel et al., 2006).

Despite the frequent loss of NKX3.1 expression, epigenetic changes such as classical

hypermethylation of the NKX3.1 promoter or mutations in the NKX3.1 coding region

that would account for the lack of NKX3.1 expression are not commonly reported and

do not correlate with NKX3.1 levels in prostate tumours (Section 1.2.3) (Voeller et al.,

1997; Xu et al., 2000; Ornstein et al., 2001; Asatiani et al., 2005). Additionally, NKX3.1

mRNA levels may not be reduced in prostate tumours, whilst a discordance between

NKX3.1 mRNA and protein levels has been observed in human prostate cancer,

suggesting that translational or post-translational control of NKX3.1 expression may be

altered in prostate tumour cells (Xu et al., 2000; Ornstein et al., 2001; Bethel et al.,

2006). In the prostate tumours of Nkx3.1+/-/Pten+/- mice, Nkx3.1 mRNA expression is

maintained, however Nkx3.1 is mislocalised to the cytoplasm in a subset of prostate

tumour cells, indicating its inability to perform transcriptional regulatory functions

(Kim et al., 2002b; Mimeault and Batra, 2011). Thus, in addition to NKX3.1 LOH in

prostate tumours, altered NKX3.1 gene transcription, potentially involving epigenetic

alterations, and translational or posttranslational dysregulation may contribute to the

reduced or lack of function of NKX3.1 in prostate cancer cells.


9

Recent studies have shown that NKX3.1 is involved in the DNA damage response by

regulating the activation of the kinases ATM and ATR and thereby their

phosphorylation of histone 2AX (termed γH2AX), an indicator of DNA damage

(Bowen and Gelmann, 2010; Erbaykent-Tepedelen et al., 2011). Upon DNA damage,

NKX3.1 also colocalised with ATM, ATR and γH2AX at DNA damage foci and in the

longer term, NKX3.1 expression reduced the accumulation of γH2AX foci in DNA

damage induced cells, indicative of lower levels of DNA damage (Bowen and Gelmann,

2010). Erbaykent-Tepedelen et al. (2011) determined that the more efficient response

of cells expressing high levels of NKX3.1 to DNA damage is likely to be due to higher

basal levels of phosphorylation of proteins involved in the DNA damage response such

as ATM, CHK2 and H2AX, indicating a constitutively active DNA damage response in

these cells (Erbaykent-Tepedelen et al., 2011). Given the involvement of NKX3.1 in the

DNA damage response, loss of NKX3.1 may result in a defective response to DNA

damage, thereby contributing to prostate tumour initiation and progression.

1.2.1 NKX3.1 Binding Partners

A number of NKX3.1 binding partners have been identified and of interest is the finding

that the majority of these are also transcription factors with which NKX3.1 interacts to

regulate gene expression (Section 1.2.2). In addition, NKX3.1 is able to associate with

other factors that influence gene expression such as histone deacetylase 1 (HDAC1) and

topoisomerase I.

The first NKX3.1 binding partner identified was serum response factor (SRF), a

transcription factor that plays important roles in embryogenesis and in the adult is

required for the growth and differentiation of skeletal muscle (Carson et al., 2000; Li et

al., 2005). The binding of NKX3.1 and SRF has been shown to require the NKX3.1

tinman motif (amino acids 29-35), the acidic domain (amino acids 88-96) and the SRF

interacting motif (amino acids 99-105) located in the amino–terminal region as well as

the carboxy-terminal amino acids 216-234 of NKX3.1, which interact with the SRF

MADS box (Ju et al., 2006; Zhang et al., 2008b). The MADS box of SRF is a DNA

binding and dimerisation motif and has been documented to facilitate interactions of

SRF with the transcription factor, TEL1 (Gupta et al., 2001), as well as the

homeodomain transcription factor, Nkx2.5 (Chen and Schwartz, 1996). The interaction

between SRF and Nkx2.5 requires the homeodomain of Nkx2.5, which is traditionally

characterised as a DNA binding domain, and although the homeodomain had previously


10

been reported to mediate NKX3.1 interaction with SRF, more recent studies have

shown that it is not necessary for this interaction (Chen and Schwartz, 1996; Carson et

al., 2000; Ju et al., 2006; Zhang et al., 2008b). The amino- and carboxy-terminal

regions of NKX3.1 facilitate its interaction with other proteins (Ju et al., 2006; Zhang et

al., 2008b; Chen et al., 2002), however fine mapping of the specific protein domains

involved has only been performed in the above-mentioned studies.

Other NKX3.1-interacting transcription factors include PDEF, the SP transcription

factors and MYC. Prostate derived ETS factor (PDEF) is a recently isolated member of

the ETS transcription factor family and its expression is confined to epithelial cells of a

number of tissues, including the prostate and breast, where it is purported to function as

a tumour suppressor and in some studies as an oncogene (Oettgen et al., 2000; Steffan

and Koul, 2011; Sood et al., 2012). The interaction between NKX3.1 and PDEF was

shown to be dependent on the NKX3.1 homeodomain and a tyrosine rich sequence

carboxy-terminal to the homeodomain, which interacts with the PDEF ETS domain and

linker region (Chen et al., 2002a; Chen and Bieberich, 2005). Nkx3.1 and SP family

members form complexes both in vitro and in vivo that require the homeodomain and

amino-terminal of Nkx3.1 and the DNA-binding domain of SP proteins (Simmons and

Horowitz, 2006). Interestingly, Nkx3.1 DNA binding is not required for its interaction

with SP family members (Simmons and Horowitz, 2006). Recently, direct interaction of

NKX3.1 with both AR and FOXA1 was reported, with the novel transcriptional

complex forming in an androgen dependent manner (Tan et al., 2012). Nkx3.1 has also

been identified to interact with the oncoprotein Myc, with the process requiring the Myc

II box but occurring in a DNA independent manner (Anderson et al., 2012).

In addition to transcription factors, NKX3.1 has been shown to interact with HDACI, an

enzyme that plays a role in transcriptional repression by deacetylating histones and

thereby aiding in chromatin condensation (Doetzlhofer et al., 1999). As such, NKX3.1

recruits HDAC1 to NKX3.1 response elements to downregulate target gene expression

(Zhang et al., 2008a). HDAC1 is also able to deacetylate (acetylated) proteins, for

example p53, resulting in p53 ubiquitination and degradation (Ito et al., 2002) and it has

been proposed that NKX3.1 interaction with HDAC1 recruits it away from MDM2-p53

complexes, thereby stabilising p53 protein levels and activity (Li et al., 2006). NKX3.1

interaction with the DNA resolving enzyme topoisomerase I, which requires the

homeodomain, enhances topoisomerase activity on DNA, and this effect is supported in


11

vivo by the observation that prostate tissue from Nkx3.1-/- and Nkx3.1+/- mice exhibits

reduced topoisomerase I activity (Bowen et al., 2007). Given the cellular functions of

topoisomerase I, this interaction, and therefore NKX3.1, have the potential to play a role

in the regulation of transcription, DNA replication or DNA repair.

1.2.2 NKX3.1 Target Genes

Using its homeodomain, NKX3.1 is able to bind to its ‘TAAGTA’ consensus sequence

located in the regulatory regions of target genes, and in cooperation with cofactors,

modulates gene expression. NKX3.1 functions predominantly as a transcriptional

repressor, with analysis of gene expression profiles in the prostates of conditional

Nkx3.1-/- mice identifying that the majority (~80%) of differentially expressed genes

were overexpressed in PIN lesions compared to the percentage of genes exhibiting

reduced expression (~20%) (Song et al., 2009). Although NKX3.1 has typically been

documented to act as a transcriptional repressor, it upregulates the expression of another

prostate specific protein, prostate cancer gene 1 product (PCAN1/GDEP) via binding to

two NKX3.1 binding sites (NBS) located in the PCAN1 promoter (Olsson et al., 2001;

Liu et al., 2008). In mice, Nkx3.1 interaction with Srf activates transcription from the

smooth muscle gamma-actin (SMGA) promoter in a mechanism that is proposed to

involve Nkx3.1 binding to an Nkx3.1 binding site in the SMGA promoter and

recruitment of Srf to nearby Srf binding elements, thereby increasing transcription

(Carson et al., 2000).

NKX3.1 interacts with steroid hormone receptors in the hormonal regulation of gene

expression, competing with the oestrogen receptor for binding sites in the promoter

regions of a subset of oestrogen responsive genes, and thereby acting as a

transcriptional repressor of oestrogen responsive genes (Holmes et al., 2008). In

addition, NKX3.1 represses expression of the AR via binding to an NKX3.1 responsive

element in the AR promoter, in vitro results that are supported by in vivo observations

that prostatic tissue in Nkx3.1-/- mice exhibits higher levels of AR expression (Lei et al.,

2006). However, other studies have reported that AR expression is correlated with

NKX3.1 protein levels in human prostate tumours and that knockdown of NKX3.1

reduces AR expression (Xu et al., 2000; Tan et al., 2012). It has also recently been

shown that NKX3.1 and the AR co-operate to regulate the expression of a number of

androgen-responsive genes and that AR and NKX3.1 are able to bind both the AR and

NKX3.1 promoter regions, implicating the transcription factors in auto-regulation and


12

coregulation (Tan et al., 2012). Tan et al. (2012) determined that in the presence of

androgens, NKX3.1 binds to an intragenic region 79kb downstream of the AR

transcriptional start site where it is suggested to enhance AR gene expression (Tan et al.,

2012). Similarly, the oncoprotein Myc and Nkx3.1 have been determined to bind to

regulatory sites within the promoters of many common target genes, thereby suggesting

that these proteins coregulate gene expression, although Nkx3.1 was reported to

antagonise the transcriptional activity of Myc on a subset of coregulated target genes

including Hk2 (Anderson et al., 2012). Nkx3.1 loss cooperates with Myc

overexpression in the promotion of prostate carcinogenesis, highlighting the importance

of interactions between these transcriptional regulators (Anderson et al., 2012).

In genome-wide screens performed to identify genes differentially expressed in Nkx3.1

null prostates during various stages of prostate carcinogenesis it was observed that the

differentially expressed gene signatures were similar in Nkx3.1-/- and Pten-/- mouse

prostate tissues (Song et al., 2009). Further, these gene signatures were similar to those

of prostate tumours in mice with prostate restricted expression of constitutively active

Akt (Song et al., 2009). The differential expression of a common subset of genes in

these mouse models was dependent on the loss of Nkx3.1, which occurs early during

prostate carcinogenesis in Pten-/- mice (Song et al., 2009). NKX3.1 has also been

implicated in the regulation of PI3K/Akt signalling by antagonising the expression of

ligands responsible for activating the signalling pathway (Sarker et al., 2009). In its

negative regulation of expression of one of these ligands, vascular endothelial growth

factor C (VEGF-C), NKX3.1 binds to an NKX3.1 response element located at -997 of

the VEGF-C promoter and recruits HDAC1, with this effect abrogated upon siRNA

mediated knockdown of NKX3.1 expression or the use of HDAC1 inhibitors (Zhang et

al., 2008a). By binding to its cognate receptor, VEGFR3, VEGF-C enhances

lymphangiogenesis, and increased levels of VEGF-C are correlated with lymph node

metastasis in prostate cancer (Zhang et al., 2008a). Therefore, given its negative

regulation of VEGF-C expression, NKX3.1 loss may aid in lymphangiogenesis, and this

is supported by the finding that deletion of 8p21.1-21.2 is associated with the

development of lymph node metastases (Oba et al., 2001).

An alternative mechanism by which NKX3.1 negatively regulates PI3K/Akt signalling

is by its downregulation of the expression and function of IGF1, a ligand for the IGF1

receptor (IGF1-R) (Muhlbradt et al., 2009; Zhang et al., 2012). In the PC-3 prostate


13

cancer cell line, ectopic expression of NKX3.1 was found to reduce IGF1 expression at

both the mRNA and protein level, implicating NKX3.1 in the transcriptional regulation

of IGF1 expression (Zhang et al., 2012). In addition, by upregulating the expression of

IGF1 binding protein 3 (IGFBP3) which antagonises IGF1 activity, NKX3.1 acts to

reduce IGF1 signalling and suppress IGF1 induced cell proliferation (Muhlbradt et al.,

2009). NKX3.1 overexpression was also found to reduce the phosphorylation of

downstream IGF1 signalling factors such as IGF1-R and AKT, whilst in the presence of

an IGF1 variant which is unable to bind IGFBP3, NKX3.1 mediated inhibition of IGF1

signalling was reduced (Muhlbradt et al., 2009; Zhang et al., 2012). Overall, these

studies have demonstrated that by modifying IGF1 signalling, NKX3.1 is able to

downregulate cell growth, thus providing a potential mechanism by which NKX3.1 loss

leads to deregulated cell proliferation in prostate cancer.

NKX3.1 interaction with transcription factors can also modulate their transcriptional

activity, thereby indirectly regulating gene expression. For example, NKX3.1 binding to

PDEF and SP family members is able to repress their transcriptional activation function,

thereby negatively regulating expression of their target genes, including PSA (Chen et

al., 2002a; Simmons and Horowitz, 2006). Although numerous putative SP recognition

elements are located in the human PSA promoter, only those at the distal end were

necessary for NKX3.1 mediated suppression of promoter activity (Simmons and

Horowitz, 2006). In addition to its role as an intracellular transcription factor, NKX3.1

has been investigated as a paracrine transcription factor, and was recently shown to be

secreted from prostate epithelial cells, regulating gene expression in nearby cells (Zhou

et al., 2012). Supporting this finding, the effect was abolished in the NKX3.1 (T164A)

mutant, a germline NKX3.1 mutation reported to cosegregate with hereditary prostate

cancer, which is not able to be secreted (Zhou et al., 2012). Although the NKX3.1

(T164A) mutant has been documented to disturb the NKX3.1 homeodomain and

therefore reduce DNA binding, in a human prostate epithelial cell line the NKX3.1

(T164A) mutant was able to repress transcription from the PSA promoter (Zheng et al.,

2006; Zhou et al., 2012). This indicated that the NKX3.1 (T164A) mutant was still able

to function as a transcription factor, perhaps by its interaction with other transcriptional

regulators or cofactors and therefore that the mutation affected expression of a specific

subset of NKX3.1 target genes (Zhou et al., 2012).


14

1.2.3 Regulation of NKX3.1 Gene Expression

NKX3.1 expression is regulated by a number of factors including androgens (AR),

retinoids (RAR/RXR) and the ETS transcription factors, ETS1, ERG and ESE3. These

transcription factors include NKX3.1 target genes which NKX3.1 regulates in a

feedback loop. The androgen regulated expression of NKX3.1 is well characterised in

vitro and in vivo (He et al., 1997; Prescott et al., 1998), with the original report of the

cloning of Nkx3.1 identifying that following castration of mice, Nkx3.1 mRNA levels

rapidly declined but were restored by the administration of androgens (Bieberich et al.,

1996). Similarly in human androgen-responsive prostate cancer cell lines, NKX3.1

expression is markedly upregulated upon treatment with androgens and decreases

following androgen withdrawal (He et al., 1997; Prescott et al., 1998). The effects of

androgens on NKX3.1 expression are mediated at least in part by an androgen response

element located in the NKX3.1 5’ promoter (-3013) which negatively regulates NKX3.1

expression and two androgen responsive elements in the NKX3.1 3’UTR which enhance

NKX3.1 expression (Yoon and Wong, 2006; Thomas et al., 2010). The AR is recruited

in a ligand dependent manner to the NKX3.1 promoter (+2 and +39 from the

transcriptional start site), with siRNA-mediated knockdown of AR expression reducing

NKX3.1 mRNA and protein levels (Tan et al., 2012). Thus NKX3.1 and AR coregulate

each other’s expression by binding to consensus sequences located in their own

promoter regions, as well as common target genes (Tan et al., 2012).

The tumour suppressor p53 reduces NKX3.1 expression by preventing androgen-

induced NKX3.1 promoter transactivation, and this effect is opposed by AR

overexpression (Jiang et al., 2006a). p53 was hypothesised to repress NKX3.1

expression by preventing AR binding to an (uncharacterised) androgen response

element located in the NKX3.1 promoter (Jiang et al., 2006a). This is in contrast to the

role of NKX3.1 in stabilising p53 protein levels (Lei et al., 2006). The relationship

between p53 and NKX3.1 is proposed to be linked via PTEN which regulates both p53

stabilisation and transcriptional activity and is suggested to regulate NKX3.1 as Nkx3.1

is down regulated in Pten null prostate tissues (Wang et al., 2003b). Therefore, several

transcription factors with which NKX3.1 cooperates in the regulation of gene

expression and whose expression NKX3.1 regulates also modulate NKX3.1 expression

in a feedback loop. This has also been suggested for Myc, where PIN lesions

overexpressing Myc exhibit low levels of Nkx3.1 expression in comparison to normal


15

epithelial tissue, indicating a role for Myc in the regulation of Nkx3.1 expression (Iwata

et al., 2010).

NKX3.1 expression is induced by retinoids (9-cis retinoic acid (9-cis RA) and all-trans

retinoic acid (tRA)) which are present at high levels in the prostate (Jiang et al., 2006b;

Thomas et al., 2006). The effects of 9-cis RA are reported to be mediated via two RA

response elements located in the NKX3.1 promoter, while the tRA responsive regions in

the NKX3.1 promoter have not been defined (Jiang et al., 2006b; Thomas et al., 2006).

NKX3.1 expression is regulated both negatively and positively by ETS transcription

family members. ETS1 overexpression induces an increase in NKX3.1 mRNA and

protein levels in prostate cancer cells, and these effects are mediated in part by an ETS1

binding site (EBS) located in the NKX3.1 5’ promoter (Preece et al., 2011). The ETS

transcription factors ESE3 and ERG mediate opposing effects on NKX3.1 expression,

with ERG reducing NKX3.1 expression directly by binding to ERG binding sites in the

NKX3.1 promoter and indirectly by inducing the polycomb group protein, EZH2, which

mediates H3K27 methylation of the NKX3.1 promoter (Kunderfranco et al., 2010).

Conversely, ESE3 is a positive regulator of NKX3.1 expression, binding to the same

response element in the NKX3.1 promoter and activating transcription as well as binding

to the EZH2 promoter, reducing its expression and therefore downregulating NKX3.1

promoter methylation (Kunderfranco et al., 2010). ERG displaces ESE3 binding on the

NKX3.1 and EZH2 gene promoters, indicating that overexpression of ERG has a

negative effect on NKX3.1 expression and therefore promotes cell proliferation

(Kunderfranco et al., 2010). This finding may have implications for prostate cancers

with TMPRSS2-ERG gene fusions that result in ERG overexpression, thus providing a

mechanism for the reduction in NKX3.1 expression in cancers where NKX3.1 is not

disrupted (Clark and Cooper, 2009). Interestingly, transgenic mice overexpressing

either ERG or a TMPRSS2-ERG fusion protein do not develop prostate cancer but

develop PIN. However, simultaneous loss of Pten in these mice results in the

development of invasive carcinoma (Carver et al., 2009; King et al., 2009). As

mentioned previously, mice exhibiting overactive PI3K/Akt signalling display low or

undetectable Nkx3.1 protein levels, suggesting that loss of Nkx3.1 expression caused by

increased PI3K/Akt signalling or ERG overexpression may aid in part the initiation or

progression of prostate cancers in these mice.

Overexpression of the transcription factor SP-1 was found to enhance NKX3.1

expression at both the mRNA and protein level, an effect mediated by SP-1 binding to


16

two SP-1 recognition elements (+29 to +43, and -60 to -46 from the transcription start

site), which are located in the NKX3.1 promoter (Yu et al., 2009). A SNP in the NKX3.1

5’UTR associated with prostate cancer susceptibility has been determined to encode a

SP-1 binding site, with the susceptible G allele associated with lower levels of NKX3.1

expression in patients carrying this variant (Akamatsu et al., 2010). Thus, patients with

this polymorphism may be predisposed to the development of prostate cancer. NKX3.1

LOH and dysregulation of NKX3.1 regulators provides one mechanism by which

NKX3.1 expression is reduced or lost in prostate tumours, however studies have

reported that NKX3.1 mRNA levels are high in prostate tumours and that there is a

discordance between NKX3.1 mRNA and protein levels (Xu et al., 2000; Ornstein et al.,

2001; Bethel et al., 2006; Kim et al., 2002b). Additionally, Nkx3.1 is reported to be

mislocalised to the cytoplasm in a proportion of prostate tumours, suggesting that in

these cases, aberrant NKX3.1 function is not due to transcriptional abnormalities but

may be due to dysfunctional translation or post-translational control of NKX3.1.

1.3 RMND5 Proteins

In order to identify novel NKX3.1 binding partners, yeast two hybrid assays were

carried out in our laboratory, which identified the FLJ22318 gene product as a potential

NKX3.1 interacting protein. This protein was later renamed RMND5B (NM_022762.3)

and together with RMND5A (NM_022780.3) it forms the RMND5 protein family

(Section 4.1.2). RMND5 proteins share 70% amino acid homology and are highly

conserved between diverse mammalian species, suggesting that they play similar

cellular roles. Named after their yeast orthologue, RMD5, a RING domain E3 ubiquitin

ligase, both RMND5 proteins also possess carboxy-terminal RING domains, suggesting

that they too possess this enzymatic activity (Section 4.1.1, 4.2.1) (Santt et al., 2008).

The aims of this thesis included characterisation of the biological activity of RMND5

proteins and the functional consequences of the interaction between NKX3.1 and

RMND5B, which had been confirmed previously (Dawson, 2006), and potentially

RMND5A. These studies were based on the investigation of RMND5 proteins as E3

ubiquitin ligases.

1.4 Ubiquitin and Ubiquitin-like Proteins Post-translational modifications (PTMs) add a layer of complexity and diversity to the

proteome and currently more than 200 types of PTMs have been described, most of


17

which are carried out by enzymes (Walsh, 2005). These protein modifications include

the covalent linkage of small molecules such as phosphoryl or glycosyl groups or the

attachment of whole proteins, such as ubiquitin to amino acid side chains. Their

presence may alter the function, stability and cellular localisation of the modified

protein (Walsh, 2005).

The best characterised example of a protein PTM is the small 76 amino acid protein

ubiquitin which was first identified in the 1970s (Ciechanover et al., 1980; Hershko et

al., 1980; Hershko et al., 1981). Ubiquitin, which adopts a characteristic β-grasp or

ubiquitin-like fold, is highly conserved in eukaryotes but is absent from bacteria and

archaea (Vijay-Kumar et al., 1987a; Vijay-Kumar et al., 1987b; Hochstrasser, 2009).

Following the characterisation of ubiquitin as a PTM, a family of small related proteins,

termed ubiquitin-like proteins (Ubl) were identified, all of which are activated and

conjugated to their specific substrates via a similar mechanism, with each containing a

ubiquitin-like fold, even when their sequences are not conserved with that of ubiquitin

(Kerscher et al., 2006). The Ubls, which include small ubiquitin related modifier

(SUMO), neural precursor cell expressed, developmentally down-regulated 8 (NEDD8)

and human leukocyte antigen F adjacent transcript 10 (FAT10), all contain a carboxy-

terminal glycine residue which is the site of conjugation of ubiquitin or the Ubl to its

target site (Kerscher et al., 2006; Hanzelmann et al., 2012). Ubiquitin and most Ubls

must undergo processing to expose the carboxy-terminal glycine residue as they are

produced as inactive precursors, and deubiquitinating enzymes (DUBs) or Ubl-specific

proteases (USPs) cleave ubiquitin or Ubls, respectively before they can be attached to

substrate proteins (Pickart and Rose, 1985; Monia et al., 1989; Kamitani et al., 1997;

Layfield et al., 1999).

1.4.1 Ubiquitin Cascade

In order for the carboxy-terminal glycine residue of ubiquitin and Ubls to be attached

covalently to a target lysine residue, it must undergo activation via an enzymatic

cascade consisting of three steps (Figure 1.3). In the first step, a ubiquitin activating

enzyme (E1) activates free ubiquitin in an ATP dependent manner, forming an E1-

ubiquitin thioester linkage with the carboxy-terminal glycine residue of ubiquitin

(Ciechanover et al., 1981; Haas et al., 1982). The ubiquitin is then passed from the

active site cysteine of the E1 enzyme to that of the next enzyme in the cascade, the E2

or ubiquitin conjugating enzyme (Hershko et al., 1983). The activated ubiquitin


18

molecule is then attached to the ε-amino group of a lysine residue in the substrate or

previously attached ubiquitin, or in some cases the ubiquitin is linked to the α-amino

group of the amino-terminal residue of ubiquitin or cysteine side-chain of the substrate

protein with the aid of a ubiquitin ligase, the E3 enzyme, which is responsible for

substrate protein recognition and binding (Hershko et al., 1983; Ciechanover and

Schwartz, 1989; Ciechanover and Ben-Saadon, 2004; Cadwell and Coscoy, 2005).

Two main types of E3 ubiquitin ligases have been identified, the homologous to E6-AP

carboxy-terminus (HECT) proteins and the really interesting new gene (RING) proteins,

which are classified according to the domain that is required for interaction with the E2

conjugating enzyme and thus ubiquitin transfer (Deshaies and Joazeiro, 2009). In

addition to this three-step reaction, complexity is added in the form of E4 and DUB

enzymes. E3 enzymes that elongate polyubiquitin chains in cooperation with other E3

enzymes are known as E4 enzymes (Koegl et al., 1999; Hoppe, 2005), and DUB

enzymes remove ubiquitin by hydrolysing the ubiquitin-substrate bond thereby

recycling ubiquitin chains (Reyes-Turcu et al., 2009). The ubiquitin enzymatic cascade

is hierarchical, there are two E1 enzymes, approximately 40 E2 enzymes and over 600

Figure 1.3: The ubiquitination cascade. The ubiquitin conjugating enzyme, E1 activates free ubiquitin in an ATP dependent manner, forming a ubiquitin thioester with the carboxy-terminal glycine residue of ubiquitin. The E1 then associates with the E2 ubiquitin conjugating enzyme resulting in the transfer of the activated ubiquitin molecule to the active site cysteine residue of the E2. The E2 enzyme next binds to an E3 ubiquitin ligase, which recognises and binds the substrate, resulting in the transfer of ubiquitin to the target lysine residue of the substrate, with the carboxy-terminal glycine of ubiquitin linked via an isopeptide bond to the lysine residue of the substrate protein (Adapted from (Fang and Weissman, 2004)).


19

ubiquitin protein ligases, and therefore it is the E3 enzymes that confer the most

specificity to the pathway (Deshaies and Joazeiro, 2009; Schulman and Harper, 2009).

1.5 Ubiquitin Activating Enzymes (E1)

1.5.1 Discovery of E1 Enzymes

In order for ubiquitin to become attached to its substrate protein or another ubiquitin

molecule, the carboxy-terminal glycine residue must be activated and this initial

activation step is carried out by a ubiquitin activating enzyme, E1 (UBE1) which was

originally described in the early 1980s (Ciechanover et al., 1982; Haas and Rose, 1982).

UBE1 is present in the cell as two isoforms, E1a (117kDa) and E1b (110kDa) that are

produced from alternative translation start sites, with E1b lacking the amino-terminal 40

amino acid residues (Cook and Chock, 1992; Shang et al., 2001). E1a and E1b are

ubiquitously expressed in equal amounts, their levels are steady during all stages of the

cell cycle, both proteins have a half-life of ~20 hours and both appear to possess the

same E2 enzyme charging ability (Handley et al., 1991; Stephen et al., 1996; Shang et

al., 2001). E1a contains a nuclear localisation signal and is itself subject to post-

translational modification by serine phosphorylation, which causes its nuclear

translocation during the G2 phase of the cell cycle (Cook and Chock, 1995). As E1b

remains unphosphorylated and localised in the cytoplasm, E1a is the predominant

isoform in the nucleus, where it is involved in cell cycle progression, with E1 mutant

cell lines undergoing cell cycle arrest at S/G2 and G2 (Finley et al., 1984; Cook and

Chock, 1995).

Plants, marsupials and mice contain more than one Ube1 or Ube1-like gene, and in mice

and marsupials the second Ube1-like gene encodes Ube1y, a protein that shares 90%

amino acid homology to UbeE1x and is a testis-specific enzyme, the gene for which is

encoded on the Y chromosome (Kay et al., 1991; McGrath et al., 1991). In 2007, three

groups reported the existence of a second mammalian E1 enzyme, UBA6/E1-

L2/UBE1L2, encoded by a gene on human chromosome 4 and sharing 40% amino acid

homology with UBE1 (Chiu et al., 2007; Jin et al., 2007; Pelzer et al., 2007). This

enzyme, now termed UBA6 is found in vertebrates, but not in worms, plants or yeast

(Pelzer et al., 2007). It is able to link covalently with ubiquitin both in vitro and in vivo,

and transfers activated ubiquitin onto a subset of E2 enzymes, including UbcH5b

(Section 1.6) (Jin et al., 2007; Pelzer et al., 2007). Interestingly, Chiu et al (2007)


20

reported that UBA6 can activate the ubiquitin-like protein, FAT10 whose cognate E1

enzyme had not previously been identified, and that RNAi against UBA6 reduced the

formation of FAT10 conjugates in cells (Chiu et al., 2007). FAT10, also known as

diubiquitin, contains two ubiquitin-like domains and its expression is induced by

tumour necrosis factor α (TNFα) and interferon γ (IFNγ), however the cognate E2 and

E3 enzymes for FAT10 are yet to be reported as UBA6 was not able to transfer FAT10

to any E2 enzymes tested (Chiu et al., 2007).

UBA6, like UBE1, is widely expressed in tissues and cell lines, with UBE1 exhibiting

an approximately ten-fold higher level of expression, although UBA6 exhibits higher

expression levels in the testis, suggesting a role for this E1 enzyme in testis

development and function (Pelzer et al., 2007). Jin et al. (2007) reported an E2 enzyme

specific for UBA6 interaction and ubiquitin transfer, USE1, which is coexpressed with

UBA6 in most human tissues, implicating UBA6 in the activation of a distinct

ubiquitination pathway with its own E2 and perhaps E3 enzymes. Uba6 homozygous

knockout mice are embryonically lethal, indicating UBA6 involvement in development,

however, since Fat10 knockout mice show no abnormalities in development, this

lethality is likely not due to defects in the FAT10 pathway (Chiu et al., 2007).

Therefore, although UBE1 is the most abundant of the E1 enzymes, suggesting that it is

responsible for mediating the activation of most ubiquitin dependent pathways, the

newly discovered UBA6 may be responsible for other as yet uncharacterised ubiquitin

and FAT10 pathways in embryonic and/or adult tissues. In addition, UBA6 may be able

to contribute its activity to a proportion of the same regulatory pathways as UBE1, as

both enzymes are able to transfer activated ubiquitin to a subset of the same E2

conjugating enzymes (Jin et al., 2007).

1.5.2 Structure and Function of E1 Enzymes

All E1 enzymes contain a similar three domain architecture, an adenylation domain

made up of two ThiF-homology motifs, a catalytic cysteine domain (CCD), and a

carboxy-terminal ubiquitin fold domain or β-grasp fold which is responsible for E2

binding (Figure 1.4) (Lake et al., 2001; Walden et al., 2003; Huang et al., 2005a). Each

domain plays a role in ubiquitin activation and the transfer of ubiquitin to cognate E2

conjugating enzymes.


21

The mechanism of catalysis of E1 enzymes, for example UBE1, has been well studied

(Haas and Rose, 1982; Haas et al., 1982). Using their adenylation domain, E1 enzymes

activate ubiquitin by initially binding Mg-ATP and catalysing the formation of a

ubiquitin adenylate with the carboxy-terminal glycine residue of ubiquitin (Figure 1.5)

(Ciechanover et al., 1982; Haas et al., 1982).

Figure 1.5: Mechanism of ubiquitin activation by E1 ubiquitin conjugating enzymes. Ubiquitin is activated in an ATP dependent manner resulting in the formation of a ubiquitin-adenylate, Ub(A) and the release of inorganic phosphate. The E1 active site cysteine residue then attacks the ubiquitin-adenylate, resulting in the formation of a ubiquitin thioester, S~Ub(T) leaving the E1 enzyme able to load a second ubiquitin molecule as a ubiquitin-adenylate Ub(A) before the transfer of the ubiquitin thioester to the ubiquitin conjugating enzyme, E2 (Schulman and Harper, 2009).

Figure 1.4: Domain structure and sequence conservation of UBE1 and UBA6. The E1 enzymes UBE1 and UBA6 contain an adenylation domain, consisting of two ThiF motifs which between UBE and UBA6 share 43% and 57% amino acid homology. The catalytic cysteine domain (CCD) and ubiquitin fold domain of UBE1 and UBA6 share 41%, 32% amino acid homology, respectively (Jin et al., 2007).


22

This ubiquitin-adenylate is subject to attack by the E1 active site cysteine residue in the

CCD domain, resulting in the formation of a high energy E1-ubiquitin linkage, with the

cysteine residue of the E1 attached to the carboxy-terminal glycine of ubiquitin (Figure

1.5) (Haas and Rose, 1982; Haas et al., 1982). The E1 enzyme is then able to load a

second ubiquitin molecule by the formation of another ubiquitin-adenylate on its

adenylation domain (Figure 1.5), with one ubiquitin covalently bound at the active

cysteine site and the other non-covalently bound at the adenylation active site (Haas et

al., 1982). A thioester transfer reaction subsequently occurs whereby the ubiquitin-

thioester on the E1 is transferred to the active cysteine residue of its cognate E2 enzyme

(Figure 1.5) (Haas et al., 1982).

1.6 Ubiquitin Conjugating Enzymes (E2)

1.6.1 The Structure of E2 Enzymes

The human genome encodes at least 37 ubiquitin conjugating enzymes (E2) (Table 1.1),

however for many of these, the particular E1 enzymes with which they are able to

associate in order to become charged with ubiquitin or Ubl, and the E3 enzymes with

which they interact to transfer activated ubiquitin/Ubl molecules to the substrate remain

poorly characterised (Michelle et al., 2009). All E2 conjugating enzymes contain a

conserved ~150 – 200 amino acid ubiquitin conjugating domain (UBC) (Hofmann and

Falquet, 2001; Burroughs et al., 2008). Structurally, the UBC is composed of four α-

helices, a four stranded anti-parallel β-sheet and a 310 helix located near the active site,

which contains a catalytically active cysteine residue located in a shallow groove (Lin et

al., 2002, Ozkan et al., 2005, Eddins et al., 2006). This is surrounded by conserved

amino acid residues involved in E2~Ub thioester formation and ubiquitination of target

lysine residues (Lin et al., 2002; Ozkan et al., 2005; Eddins et al., 2006).

Within the UBC, the conserved active site catalytic cysteine residue is responsible for

the formation of the ubiquitin thioester (Wu et al., 2003b), while a tripeptide histidine-

proline-asparagine (HPN) motif is located 7-10 amino acid residues to the amino-

terminal of the active cysteine residue (Wenzel et al., 2011). Within this motif, the

histidine residue is believed to be involved in maintaining the structure of the active

site, and the asparagine residue stabilises the oxyanion intermediate formed when the

target lysine residue of the substrate attacks the E2~Ub thioester, resulting in the

formation of an isopeptide linkage between the Ub molecule and target lysine residue


23

(Figure 1.6) (Wu et al., 2003b; Cottee et al., 2006). Mutation of the asparagine residue

does not impede E1 and E3 interactions with the E2 enzyme or ubiquitin transfer

between the enzymes, suggesting that it is only important for ubiquitin transfer to target

lysine residues (Wu et al., 2003b). Another motif present in the UBC of many E2

enzymes, the Y/FPPxxP*

Martinez-Noel et al., 2001

motif, is found 7 to 11 amino acids amino-terminal to the

HPN motif (Figure 1.6). Structural studies have shown that in the folded protein, the

carboxy-terminal proline residues are located close to a highly conserved tryptophan

residue, positioned carboxy-terminal to both the HPN motif and the catalytic cysteine

residues and this interaction may stabilise the L7 loop and aid in the positioning of the

L4 and L7 loops (Figure 1.6) ( ; Michelle et al., 2009).

* Y = tyrosine, F = phenylalanine, P = proline, x = any amino acid

Figure 1.6: Three dimensional structure of E2 enzyme ubiquitin conjugating domains. The generalised E2 enzyme structure consists of loops labelled L1 to L8, four β-sheets (S1 to S4), four α-helices (H1 to H4), and the 310 helix (h). Conserved amino acid residues are the HPN motif, the catalytic cysteine (C) residue, a conserved tryptophan residue (W) and a PxxPP motif (Michelle et al., 2009).


24

Table 1.1 – Human ubiquitin conjugating enzymes

Gene Name

Alternative Name(s)

Class Cognate Ub/Ubl and Chain Specificity (in vitro/in vivo)

Biological Role

UBE2A HR6A, RAD6A I Ub (mono, K11, K48,) DNA repair

UBE2B HR6B, RAD6B I Ub (K11, K48, K63) DNA repair

UBE2C UBCX, UbcH10 II Ub (K11) DNA repair

UBE2D1 UbcH5A, E2-17K1 I Ub (mono, K11, K48) Cell cycle regulation

UBE2D2 UbcH5B, UBC4/5, E2-17K2

I Ub (mono, K11, K48, K63) p53, immune

UBE2D3 UbcH5C, E2-17K3 I Ub (mono, K11, K48) DNA repair, apoptosis

UBE2D4 HBUCE1 I Ub (K11, K48) Unknown

UBE2E1 UbcH6 II Ub (K48) Unknown

UBE2E2 UbcH8 II Ub (K11, K48, K63) Unknown

UBE2E3 UBCE4, UbcH9, UbcM2, E2-23K

II Ub (K11, K48, K63) Growth regulation

UBE2F NCE2 II NEDD8 SCF regulation

UBE2G1 Ubc7 I Ub (K48, K63) ER

UBE2G2 Ubc7 I Ub (K48) ER

UBE2H UbcH2, E2-20K III Ub (K11, K48) Unknown

UBE2I Ubc9 I SUMO Cell cycle, chromosome segregation

UBE2J1 NCUBE1 III Ub (K11) ER

UBE2J2 NCUBE2 III Ub (K48) ER

UBE2K Ubc1, HIP-2, E2-25K

III Ub (K48) Protein quality control

UBE2L3 UbcH7, E2-F1, L-UBC, UBCE7

I Unknown Cell Cycle, transcription

UBE2L6 UbcH8, RIG-B I Ub, ISG15 IFN signalling

UBE2M Ubc12 II NEDD8 SCF regulation

UBE2N Ubc13, BLU I Ub (K63) NK-κB signalling, DNA repair

UBE2NL UEV Unknown

UBE2O E2-230K IV Unknown Haematopoiesis

UBE2Q1 NICE-5, Ube2Q II Unknown Unknown

UBE2Q2 II Unknown Mitosis


25

UBE2R1 Cdc34, Ubc3, E2-32K

III Ub (K48) Cell cycle regulation

UBE2R2 Cdc34B, Ubc3B III Ub (K48) Cell cycle regulation

UBE2S E2-EPF5, E2-24K III Ub (K11) Cell cycle regulation

UBE2T HSPC150 III Ub (mono, K11, K48, K27, K63))

DNA repair

UBE2U III Unknown Unknown

UBE2V1 UEV-1 II UEV, Ub (K63) NK-κB signalling, DNA repair

UBE2V2 MMS2, EDPF-1, UEV2

I UEV, Ub (K63) NK-κB signalling, DNA repair

UBE2V3 UEVLD, UEV-3 Unknown Unknown

UBE2W I Ub (MONO, K11) DNA repair

UBE2Z Use1, HOYS7 IV Acts with UBA6 Apoptosis

AKTIP FTS Unknown Apoptosis, vesicular transport

BIRC6 Bruce, Apollon IV Unknown Apoptosis, cytokinesis

1.6.2 Classification of E2 Conjugating Enzymes

E2s can be classified into four groups depending on their domain architecture. Class I

E2 enzymes contain only the UBC, the minimal region required for E2 activity and the

UbcH5 E2 conjugating enzymes are included in this class (Hofmann and Falquet, 2001;

Winn et al., 2004). Class II E2 enzymes are those which contain the UBC and an

amino-terminal domain, while those containing the UBC with a carboxy-terminal

extension are classified as Class III E2s (Hofmann and Falquet, 2001; Winn et al., 2004;

van Wijk and Timmers, 2010). The amino- and carboxy-terminal extensions of E2

enzymes are important for their activity and regulation of ubiquitination (Kolman et al.,

1992; Silver et al., 1992; Summers et al., 2008). Class IV E2 enzymes possess both

amino- and carboxy-terminal extensions and include two large E2 enzymes, E2-230K

(UBE2O) and apollon (BIRC6/BRUCE). E2-230K is composed of 1292 amino acids, it

plays a role in reticulocyte maturation and haematopoiesis, and is able to ubiquitinate its

substrates directly without the need for an E3 enzyme (Klemperer et al., 1989; Berleth

and Pickart, 1996). The second large E2 enzyme in this class, apollon (BIRC6/BRUCE)

is a 528kDa protein which plays an anti-apoptotic role in the cell by mediating the

ubiquitination and subsequent degradation of SMAC, a proapoptotic protein (Hao et al.,


26

2004; Sekine et al., 2005). Apollon is also able to ubiquitinate its target proteins directly

without the association of an E3 ubiquitin ligase (Hao et al., 2004).

Alternatively, Michelle et al., (2009) have proposed a different E2 classification system

based on phylogenetic data (Michelle et al., 2009). By this system, E2 enzymes may be

subdivided into 17 subfamilies which do not resemble the abovementioned

classification system, and although the newer proposed classification method takes into

account the functional domains and structure of the E2 enzymes, the 17 families mostly

contain only one or two E2 enzymes, making it more complex compared to the initial

method.

1.6.3 E2 Conjugating Enzyme Interactions with E1 and E3 Enzymes

The function of E2 enzymes in the ubiquitin pathway is to interact with both E1 and E3

enzymes in order to facilitate the transfer of activated ubiquitin to target lysine residues.

Initially the E2 enzyme must interact with a ubiquitin E1 activating enzyme that is

loaded with two ubiquitin molecules, and transfer the ubiquitin to its own cysteine

residue (He and Rape, 2009). The E1 enzyme undergoes conformational alterations

upon ubiquitin binding, which make available negatively charged sites within the UBC

domain of the E1 that can be recognised by lysine residues within the E2 α-helix 1

(H1), with these residues are only present in E2 enzymes that bind ubiquitin (Lee and

Schindelin, 2008). As such, the high affinity of the E2 for the E1 enzyme occurs only

when the E1 is charged with two ubiquitin molecules, thus ensuring specificity of the

E1-E2 interaction and ubiquitin transfer (Huang et al., 2007; Lee and Schindelin, 2008).

The mechanisms by which individual E2 enzymes are able to distinguish between the

two human E1 enzymes, UBE1 and UBA6 are not well understood (Jin et al., 2007).

Following the formation of the E2~Ub thioester, the E2 enzyme must associate with the

E3 ubiquitin ligase to allow substrate ubiquitination. The E1 and E3 interaction sites of

the E2 enzyme are overlapping, requiring the amino-terminal H1 and loops L4 and L7

and as such, the E2 enzyme must disengage from the E1 enzyme before it is able to

interact with the E3 enzyme to enable ubiquitin transfer. Furthermore, the E2 cannot be

recharged with ubiquitin whilst bound to the E3 (Figure 1.6) (Eletr et al., 2005). E2

enzymes can interact with either of the two classes of E3 ubiquitin ligases, HECT and

RING domain containing proteins, with these domains of the E3s interacting with

similar regions of the E2 UBC domain (L4 and L7) (Section 1.6.1). The L4 hydrophobic


27

phenylalanine amino acid residues appear to be more important for E2-HECT domain

interactions compared to E2-RING interactions, whilst hydrophobic residues in L7 are

required for both HECT and RING interactions with the E2 (Nuber and Scheffner,

1999; Christensen et al., 2007).

The ability of an E2 enzyme to interact with several E3 ubiquitin ligases relates to

multiple E3 interacting residues in the UBC. For example, UBE2N uses the amino acids

arginine 6 and lysine 10 in its recognition of the E3 TRAF6, but residues arginine 7 and

lysine 10 when interacting with CHIP (Zhang et al., 2005; Yin et al., 2009). E2

enzymes are additionally able to recognise and associate with residues outside of the

RING or HECT domains, for example the E2 enzyme UBE2G2 binds the E3 gp78 at its

RING domain and Ube2G2 Binding Region (G2BR) motif, enhancing the interaction

between this E2 and E3 enzyme pair (Chen et al., 2006a; Li et al., 2009). Although the

binding of the E2 and E3 are weak, E2-E3 binding facilitates the release of the ubiquitin

molecule from the E2 active site. This is thought to be due to a conformational change

that takes place upon E3 binding, where the asparagine residue in the HPN motif is

positioned close to the E2 active site, thus stabilising the oxyanion intermediate formed

between the acceptor lysine residue and ubiquitin carboxy-terminal glycine, described

above (Wu et al., 2003b; Ozkan et al., 2005). The low E2 affinity for the E3 is also

hypothesised to favour ubiquitin chain formation because the E2 must disengage from

the E3 before it can become recharged with ubiquitin (Eletr et al., 2005; Ye and Rape,

2009).

1.6.4 Roles of E2 Enzymes in Ubiquitin Chain Formation

1.6.4.1 Chain Initiating and Chain Elongating E2 Enzymes

Ubiquitination of a substrate is initiated when a ubiquitin molecule is attached to the

target lysine residue of the substrate and this monoubiquitination can be built upon by

the attachment of multiple ubiquitin molecules to produce a polyubiquitin chain. E2

enzymes show a preference for ubiquitin chain initiation or elongation (Windheim et

al., 2008), with the E2 enzymes UBE2W and UBE2E2 involved in ubiquitin chain

initiation when they associate with the E3 heterodimer breast cancer type 1

susceptibility protein (BRCA1)- associated RING domain 1 (BARD1), performing

monoubiquitination of the target lysine residue of the substrate (Christensen et al.,

2007). In contrast, the E2s Ubc13-Mms2 and UBE2K are involved in ubiquitin chain


28

elongation, forming lysine 48 or lysine 63 linked ubiquitin chains upon association with

this E3 complex (Christensen et al., 2007). Some E2 enzymes which are involved in

chain initiation are not specific for the lysine residues in the target which become

ubiquitinated. For example, UbcH5 (UBE2D) family members are able to ubiquitinate

various substrates on multiple lysine residues to initiate chain formation (Kirkpatrick et

al., 2006; Windheim et al., 2008). However, the E2 UBE2T preferentially

monoubiquitinates FANCD2 on specific lysine residues (Alpi et al., 2008). Many chain

elongation E2 enzymes are only able to recognise ubiquitinated substrates, and interact

with the ubiquitin bound to the substrate to attach more ubiquitin molecules, thereby

creating specific chain topologies. UBE2S specifically forms lysine 11 linked

polyubiquitin chains, whereas UBE2K shows a preference for lysine 48 linked ubiquitin

chains, however neither of these E2 enzymes are able to initiate chain formation

(Haldeman et al., 1997; Hofmann and Pickart, 1999; Windheim et al., 2008). As such,

when associated with the E3, TRAF6, UbcH5 (UBE2D) initiates ubiquitin chain

formation and the E2 UBE2N-UBE2VI attaches additional ubiquitin molecules through

lysine 63, activating NF-κB (Petroski et al., 2007). A few E2 enzymes such as yeast

Cdc34 are able to initiate the formation and elongation of ubiquitin chains, and Cdc34

interacts with the E3 SCF to ubiquitinate S-phase cyclin/cyclin-dependent kinase

inhibitor (Sic1), forming lysine 48 linked polyubiquitin chains and targeting Sic1 for

degradation (Verma et al., 1997). Other E2 enzymes are able to initiate chain formation

and catalyse the formation of short ubiquitin chains before a second, chain elongation

E2 extends the short chains (Ye and Rape, 2009).

1.6.4.2 E2 Enzyme Processivity

Ubiquitin chain length and therefore the ultimate fate of the ubiquitinated protein are

dependent on the processivity of ubiquitin chain formation, which is defined as “the

number of ubiquitin molecules transferred to a growing chain during a single round of

substrate association with an E3” (Ye and Rape, 2009). Factors influencing the

processivity of ubiquitin chain formation are the affinity of the E3 for the substrate and

the rate at which the E2 catalyses ubiquitin transfer (Ye and Rape, 2009). For a protein

to be targeted for degradation, a lysine 11 and 48 linked polyubiquitin chain of at least

four ubiquitin molecules is required, and some E2 enzymes have developed novel

mechanisms to increase their processivity. The human anaphase promoting complex

(APC) binds the substrate securin, resulting in its rapid polyubiquitination through

lysine 11, a process catalysed by the E2s UBE2C and UBE2S that results in its


29

degradation (Rape and Kirschner, 2004). UBE2C recognises a specific region in securin

and ubiquitin known as a TEK box, thereby allowing the E2 to rapidly catalyse

ubiquitin chain initiation by ubiquitinating securin. Elongation occurs by attaching

multiple ubiquitin molecules through UBE2C recognition of the TEK box in both

proteins which, along with the chain elongating E2 UBE2S, increases processivity (Jin

et al., 2008; Williamson et al., 2009). In order to increase processivity, some E2

enzymes, for example UBE2G2, pre-assemble polyubiquitin chains and the chains are

then transferred en bloc to the target lysine residue (Li et al., 2007b; Ravid and

Hochstrasser, 2007). Others form E2-Ub complexes, bound both covalently to the

catalytic cysteine residue of the active site and non-covalently to the backside or β-sheet

of the E2 . This results in the concentration of active E2 enzymes, loaded with ubiquitin,

in close proximity to the E3 enzyme, thereby increasing processivity as occurs (for

example) when the BRCA1-BARD1 complex is autoubiquitinated using UbcH5

(UBE2D) family members (Brzovic et al., 2006).

1.6.4.3 E2 Enzyme Ubiquitin Chain Assembly and Linkage Selection

The ultimate fate of the ubiquitinated protein is dependent on the types of ubiquitin

linkages formed and it is the E2 enzyme which plays a major role in determining not

only the lysine residue on the substrate that is ubiquitinated (Section 1.7) but also the

type of polyubiquitin chains that are formed (Ye and Rape, 2009). Some E2 enzymes

are specific for the types of polyubiquitin chains that they synthesise, with UBE2S

synthesising lysine 11 linked polyubiquitin chains whilst UBE2K and UBE2RI catalyse

the formation of lysine 48 linked ubiquitin chains. A number of E2 enzymes are

additionally able to synthesise ubiquitin chains of these topologies even in the absence

an E3 enzyme (Chen and Pickart, 1990; Haas et al., 1991; Ye and Rape, 2009). In order

to achieve this linkage specificity, the E2 enzyme positions the lysine residue of the

acceptor ubiquitin so that it is exposed to the active site of the E2 which contains the

activated donor ubiquitin, thereby ensuring that specific lysine linkages are formed

between ubiquitin molecules (Figure 1.7) (Petroski and Deshaies, 2005; Eddins et al.,

2006).

Not all E2s are able to confer linkage specificity and members of the UbcH5 (UBE2D)

E2 family which only contain the core UBC domain, do not form specific ubiquitin

chains but rather are able to catalyse the formation of ubiquitin chains of all linkages,


30

leading to the hypothesis that in this case linkage specificity may be determined by the

E3 enzyme (Brzovic et al., 2006, Ye and Rape, 2009).

Recently, it has been reported that although the E2 enzyme plays a major role in

determining which lysine residues of ubiquitin are linked upon polyubiquitin chain

formation, the E3 enzymes are important in the determination of the type of

ubiquitination (mono- or poly-ubiquitination), ubiquitin linkage type and the identity

and specificity of target lysine residue of the substrate by directing the E2 enzyme

(David et al., 2011). E2 enzymes are therefore an integral part of the ubiquitin

proteasome system and do not merely function to transfer ubiquitin. They are important

for the determination of the length and topology of ubiquitin chains and the

determination of the target lysine residue of the substrate protein thereby determining

the fate of the ubiquitinated substrate.

1.7 E3 Ubiquitin Ligases E3 ubiquitin ligases are involved in the final step of the ubiquitin cascade and are

responsible for bringing together the substrate and activated ubiquitin molecule by

recognising and binding the substrate protein as well as the E2 enzyme loaded with the

Figure 1.7: E2 ubiquitin chain linkage selection model. The E2 enzyme bound to the donor ubiquitin is able to orient the acceptor ubiquitin exposing the correct lysine residue to the active site (Ye and Rape, 2009).


31

activated ubiquitin. As mentioned previously (Section 1.4.1), E3 enzymes can be

classified according to the domain required for interaction with the ubiquitin-charged

E2 enzyme and ubiquitin transfer. HECT and RING domains are the two main groups

of E3 enzymes, however other domains similar to the RING domain are also involved in

ubiquitination, including U-box and plant homeodomain (PHD) motifs. Another protein

that is able to ubiquitinate proteins without the presence of a recognisable HECT or

RING domain is the deubiquitinating enzyme UCH-L1, which is able to

monoubiquitinate α-synuclein by reversing its deubiquitinating action, thereby

bypassing the need for E1 and E2 enzymes (Liu et al., 2002).

1.7.1 HECT Domain E3 Ubiquitin Ligases

The HECT domain was initially identified in the human papilloma virus E6-associated

protein (E6-AP), which was shown to ubiquitinate p53, targeting it for proteasomal

degradation (Huibregtse et al., 1994; Scheffner et al., 1994). The domain functions by

interacting with the E2 enzyme, which is followed by transfer of the activated ubiquitin

to the HECT domain, resulting in the formation of a HECT~Ub intermediate. The

activated ubiquitin is then transferred to the target lysine residue of the substrate

protein, which is bound to amino-terminal domains of the HECT domain protein

(Scheffner et al., 1995). The HECT domain has been identified in 28 human proteins

and these HECT domain proteins can be divided into three subfamilies, the Nedd4

family consisting of 9 family members, the HERC family (6 members) and finally other

HECT domain containing proteins comprising 13 members (Huibregtse et al., 1995;

Harvey et al., 1999; Rotin and Kumar, 2009).

In addition to the HECT domain, all Nedd4 family members contain an amino-terminal

C2 domain and between two and four WW domains, which regulate substrate binding

and cellular localisation (Ingham et al., 2004). C2 domains are able to bind

phospholipids and proteins and recruit proteins to membranes, thereby regulating

protein localisation (Plant et al., 2000; Dunn et al., 2004). WW domains contain two

conserved tryptophan residues and recognise proline rich motifs including the PPxY†

Sudol et al., 1995

motif, which is recognised by most Nedd4 WW motifs ( ; Staub and

Rotin, 1996). Nedd4 family members include NEDD4, SMURF1, SMURF2 and ITCH

(Rotin and Kumar, 2009). HERC family members consist of proteins ranging from † P = proline, Y = tyrosine, x = any amino acid


32

100kDa to 500kDa in size dependent on the number of regulator of chromosome

condensation 1 (RCC1)-like domains (RLDs) that they contain. Although most HERC

family members contain only the RLD and HECT domains, some large HERC proteins

such as HERC1 also contain other protein-protein interaction domains such as Spla and

Ryanodine receptor (SPRY) domains and WD40 repeats (Cruz et al., 2001; Garcia-

Gonzalo et al., 2005; Edwin et al., 2010). Other HECT domain containing proteins

contain several different domains located in their carboxy terminus, including HUWE1

(MULE/ARF-BP1) which contains a WWE (Tryptophan-Tryptophan-Glutamine) and a

ubiquitin associated domain (UBA), and HACE1 which contains ankyrin repeats

(Anglesio et al., 2004).

1.7.1.1 Structure and Mechanisms of Ubiquitin Transfer by HECT Domains

The HECT domain, which is always located towards the carboxy terminus of the

protein, is composed of ~350 amino acid residues and can be divided into amino (N)

and carboxy (C) terminal lobes which are joined by a linker region (Huang et al., 1999).

The N terminal lobe associates with the E2 conjugating enzyme whilst the C terminal

lobe contains an active site cysteine residue to which the active ubiquitin molecule is

transferred from the E2 conjugating enzyme active site in a transthiolation reaction

(Huang et al., 1999). The last 60 amino acid residues of the C terminal lobe also appear

to direct ubiquitin chain linkage specificity (Kim and Huibregtse, 2009). Structural

studies of HECT domain E3s bound to E2 enzymes have shed light on the mechanism

of action of these proteins. Crystal structures of the HECT E3s E6-AP and WWP1 in

complex with UbcH7 revealed distances of 41Å or ~16Å, respectively separating the

HECT domain C lobe cysteine of the E3 and the UbcH7 active site cysteine suggesting

that, as this distance is too far for transthiolation to occur, the flexible linker region must

permit conformational changes to reduce the gap to within 6Å (Figure 1.8) (Huang et

al., 1999;(Verdecia et al., 2003).

The type of polyubiquitin chain that is formed upon ubiquitination of a substrate

molecule is important as this determines the ultimate fate of the ubiquitinated protein.

As HECT domain E3 ubiquitin ligases form a thioester intermediate with the activated

ubiquitin molecule prior to the transfer of ubiquitin to the substrate, it is the HECT E3

that determines the ubiquitin chain linkage attached to the substrate, and this is in

contrast to the mechanism of action of RING domain E3 enzymes (Section 1.7.2, 1.8)

(Wang and Pickart, 2005). The mechanism by which HECT E3s synthesise ubiquitin


33

chains differs, for example, the HECT E3, E6-AP is able to build Lys48 linked ubiquitin

chains on its active site cysteine in the HECT domain in vitro, whereas the HECT E3,

KIAA10 synthesises Lys29 and Lys48 linked free ubiquitin chains (Wang and Pickart,

2005).

1.7.1.2 Regulation of HECT E3 Ubiquitin Ligases

HECT E3 ligases are able to regulate themselves by autoubiquitination as well as being

regulated by other interacting proteins which either enhance or inhibit the HECT E3

activity by releasing autoinhibition, aiding in E2-E3 interaction or blocking E3 substrate

binding. For example, ITCH is regulated by autoinhibition mediated by the interaction

of its WW domain containing a proline rich region (PRR) and its HECT domain

(Gallagher et al., 2006). ITCH becomes activated upon phosphorylation of the PRR

motif by JNK1, which interacts with the HECT domain of ITCH on three residues

(Ser199, Thr222, Ser232) thereby disrupting the intramolecular interactions between the

ITCH domains (Figure 1.9) (Gallagher et al., 2006; Bruce et al., 2008). ITCH activity is

also regulated by binding of N4BP1 to the ITCH WW2 domain, which inhibits the

binding of ITCH to target proteins such as p73α, JUN and p63, thus inhibiting their

ubiquitination (Oberst et al., 2007). SMURF2 is also regulated by auto-inhibition,

Figure 1.8: HECT domain structure. (A) Crystal structure of the HECT domain of E6-AP and UbcH7 showing the N terminal lobe (red) and C terminal lobe (blue) structure of the HECT domain in complex with UbcH7 (yellow) and a 41Å distance between the catalytic cysteine residues (blue squares). (B) Structure of WWP1 bound to UbcH7 showing a distance of only 16Å between the catalytic cysteine residues due to the orientation of the C lobe of WWP1 (Huang et al., 1999, Verdecia et al., 2003, adapted from Rotin and Kumar, 2009)


34

Figure 1.9: Regulation of HECT domain E3 catalytic activity. (A) SMAD7 activates SMURF2 by using its N-terminal domain (NTD) to interact with both SMURF2 and the E2 UbcH7, bringing them into close proximity. This interaction increases the affinity of SMURF2 for UbcH7. (B) Auto-inhibition of ITCH, mediated by the interaction of its HECT and WW-PRR domains, is relieved by JNK phosphorylation of the PRR domain (Adapted from Kee and Huibregtse, 2007).

thereby protecting it from auto-ubiquitination and degradation (Figure 1.9). In this

process, the C2 and HECT domains of SMURF2 associate to inactivate the enzyme,

with this mechanism of auto-regulation common among HECT E3 ligases including

Nedd4L, NEDD4 and WWP2 (Wiesner et al., 2007; Bruce et al., 2008; Wang et al.,

2010). Interestingly, the auto-inhibition of NEDD4 is released by increasing

intracellular calcium levels, which disrupts the binding of its C2 and HECT domains

(Wang et al., 2010).

The E3 ubiquitin ligase activity of the HECT domain protein SMURF2 is regulated by

the adaptor protein SMAD7, which is required as the conformation of SMURF2 and the

E2 UbcH7 in complex is suboptimal, with the catalytic cysteine residues separated by

50Å (Ogunjimi et al., 2005). SMAD7 uses its amino-terminal to bind both SMURF2

and UbcH7, thereby acting as an adaptor and bringing the two enzymes into close


35

proximity to allow ubiquitin transfer (Figure 1.9) (Ogunjimi et al., 2005). The low

affinity of SMURF2 for UbcH7 is due to the presence of two hydrophilic amino acids

(His547, Tyr581) within the E2 binding pocket and replacement of the two hydrophilic

amino acids with hydrophobic residues critical for mediating E6-AP-UbcH7 interaction

results in the constitutive activation of SMURF2 (Figure 1.9) (Ogunjimi et al., 2005;

Kee and Huibregtse, 2007).

1.7.2 RING Domain E3 Ubiquitin Ligases

The second major family of E3 ubiquitin ligases possesses a RING domain or RING-

like domain such as the U-box or PHD domain. The RING domain was first described

in 1993 however the function of the domain was not characterised until 1999

(Freemont, 1993; Lorick et al., 1999) and in the intervening years the RING domain

was hypothesised to function as a DNA binding motif (Freemont, 1993; Lovering et al.,

1993; Lorick et al., 1999). RING domains have been identified in over three hundred

proteins and are therefore the most common type of E3 ubiquitin ligase with five times

more members than the HECT family (Fang et al., 2003). RING domain proteins differ

from HECT ligases in the manner in which the charged ubiquitin molecule is transferred

to the target lysine residue of the substrate molecule (Figure 1.10).

Figure 1.10: Mechanism of ubiquitin transfer by HECT and RING domain containing E3 ubiquitin ligases. (a) HECT domains accept the ubiquitin from the ubiquitin conjugating enzyme (E2), forming a ubiquitin thioester before transfer of the activated ubiquitin molecule to the target lysine residue of the substrate. (b) RING domain proteins bind the E2 enzyme and facilitate the direct transfer of the activated ubiquitin molecule to the target lysine residue of the substrate (Adapted from Rotin and Kumar, 2004).


36

RING domain proteins associate with both the E2 enzyme and the substrate, and

essentially act as a scaffold bringing the E2 and substrate into close proximity and

thereby allowing the transfer of the ubiquitin molecule (Deshaies and Joazeiro, 2009).

In contrast, following interaction with the E2 enzyme, HECT domain E3s form a

HECT~Ub intermediate before the ubiquitin molecule is transferred to the target lysine

residue of the substrate or acceptor ubiquitin (Figure 1.10) (Rotin and Kumar, 2009).

1.7.2.1 RING Domain Structure

The RING domain consists of a sequence of distinctively placed cysteine (C) and

histidine (H) residues which chelate two zinc (Zn2+) ions, thereby forming a cross brace

structure which promotes correct conformation of the domain (Figure 1.11) (Lorick et

al., 1999).

The first, second, fifth and sixth C/H amino acids coordinate the first Zn2+, while the

third, fourth, seventh and eighth C/H residues bind the second Zn2+ ion (Lorick et al.,

1999). The two main RING domain variants present are the C3HC4 (RING-HC) and

C3H2C3 (RING-H2) types, which vary as to the number and placement of their

Figure 1.11: RING domain structure. (A) The canonical RING-HC sequence consists of seven Cysteine (C) residues and one Histidine (His) residue where X is any amino acid and the number refers to the number of amino acids in the linker regions. (B) The cysteine (C) and histidine (H) amino acid residues coordinate two Zinc (Zn2+) ions, forming a cross brace structure (Adapted from Deshaies and Joazeiro, 2009).


37

histidine and cysteine amino acids. The RING domain, which is catalytically inactive, is

able to associate with the E2 enzyme, acting as a scaffold to facilitate ubiquitin transfer.

Variations in the canonical RING domain sequence have been identified in functional

E3 ubiquitin ligases, for example RBQ1 contains an aspartate at position eight and

RBX1 contains an asparagine residue at position 4 (Zheng et al., 2002; Deshaies and

Joazeiro, 2009). Other conserved amino acid residues in the RING domain that are

important for E2 enzyme binding have been identified with most positioned close to the

zinc coordinating residues and corresponding to the characterised points of contact

between the RING domain and E2 (Figure 1.12) (Deshaies and Joazeiro, 2009).

Many of the three hundred RING domain containing proteins have not yet been

characterised, and although it is believed that the majority of these proteins possess E3

ubiquitin ligase activity, a number of RING domain containing proteins are known to be

non-functional as E3s on their own. For example, the RING domain containing proteins

BARD1 and MDMX do not possess intrinsic E3 activity, but interact with other RING

domain containing proteins, BRCA1 and MDM2, respectively through their RING

domains, enhancing the E3 activity of their interacting E3 (Hashizume et al., 2001;

Linares et al., 2003). In addition to their ability to heterodimerise, RING domain

Figure 1.12: RING and U-box domain amino acid residues involved in E2 enzyme interaction. Amino acid residues important for mediating RING/HECT domain-E2 interactions (red) are distinct from the Zinc coordination residues (blue) of the RING domain proteins c-Cbl and cIAP2. The corresponding residues are marked in the U-box ubiquitin ligase CHIP (Deshaies and Joazeiro, 2009).


38

proteins are able to form homodimers, as has been reported for the E3 ubiquitin ligases

Siah and c-Cbl (Polekhina et al., 2002; Kozlov et al., 2007). These E3 ubiquitin ligase

dimers can be formed by interaction of the RING domains, as is the case for the

MDM2-MDMX heterodimer (Poyurovsky et al., 2007), however amino acid residues

carboxy-terminal to the RING domain are also involved in dimerisation, and are

involved for example in cIAP2 homodimerisation (Linke et al., 2008; Mace et al.,

2008). The ubiquitin ligase c-Cbl homodimersises using its UBA domain, which is

located distal to the RING domain and is also required for c-Cbl/Cbl-b

heterodimerisation (Kozlov et al., 2007).

1.7.2.2 Mechanisms of Ubiquitin Transfer by RING Domain E3s

Following E2-E3 binding, the RING domain is thought to aid in ubiquitin transfer by

acting as a scaffold bringing the E2 and substrate into close proximity and it is also

hypothesised that upon E2-E3 binding, conformational change is effected in the E2

enzyme, thereby augmenting ubiquitin transfer from the E2 active site to the substrate

lysine (Seol et al., 1999; Deshaies and Joazeiro, 2009). Although generally it is the E2

enzyme that determines substrate lysine specificity and ubiquitin chain linkage

topology, the RING E3 ubiquitin ligase does play a role in this determination. Recent in

vitro studies have ascertained that without an E3 enzyme, E2 enzymes were not

selective of the substrate lysine residue or the type of ubiquitin linkage formed in

studies using multiple well characterised E2-E3 pairs and characterised E3 substrates

including E2G2-gp78 and the gp78 substrate HERP (David et al., 2011). The role of the

E3 enzyme in determining which lysine residue of the substrate was ubiquitinated and

the type of ubiquitin chain linkages formed was hypothesised to be due to the

positioning of the E2 and substrate upon binding by the E3 enzyme (David et al., 2011).

It is important to note that the ubiquitin chain linkage topologies formed by particular

E2-E3 pairs are constrained to the particular chain topologies preferred by the E2

conjugating enzyme, therefore the E3 does not seem to direct new chain topologies to

be formed, but merely guides the E2 enzyme (David et al., 2011). This result highlights

the importance of the E2-E3 complex in directing ubiquitination and specificity.

Additionally, one E3 enzyme is typically able to interact with a number of E2 enzymes,

for example BRCA1 has been reported to interact with ten E2 enzymes (Christensen et

al., 2007). Therefore depending on the type of E2 enzyme with which the E3 interacts,

the ubiquitin chain topology and substrate lysine residues that are ubiquitinated may


39

change, resulting in different outcomes of ubiquitination of a single substrate.

Furthermore, substrates such as p53 may be coregulated by a number of E3 enzymes

depending upon the cellular environment, resulting in different outcomes for the

substrate depending upon the E3 enzyme responsible for its ubiquitination (Brooks and

Gu, 2006).

1.7.2.3 Regulation of RING E3 Ubiquitin Ligases

The expression and activity of RING E3 ubiquitin ligases are regulated by a variety of

PTMs including ubiquitination, sumoylation, neddylation and phosphorylation which

can be attached to the substrate, E2 or E3 to regulate their activity or recognition by the

E3 enzyme. RING domain containing proteins are able to regulate their own activity by

autoubiquitination, for example, the E3 ubiquitin ligase TRAF6 undergoes site-specific

lysine 63 autoubiquitination in response to interleukin 1 treatment, resulting in the

activation of Iκβ kinase (IKK), whilst MDM2 autoubiquitinates its own lysine residues

in response to DNA damage, resulting in its proteasome-dependent degradation, thus

leaving its target p53 to participate in the DNA damage response (Stommel and Wahl,

2004; Lamothe et al., 2007). MDM2 autoubiquitination is regulated by the

deubiquitinating enzyme, HAUSP which deubiquitinates MDM2 and p53, thereby

additionally regulating cellular p53 protein levels (Li et al., 2002; Cummins et al., 2004;

Cummins and Vogelstein, 2004). As mentioned above, RING domain proteins may be

regulated by other PTMs such as phosphorylation. Many E3 ubiquitin ligases are only

able to recognise their substrates once they are phosphorylated, as is the case for c-CBL,

which binds and ubiquitinates phosphorylated RTKs, propagating both the RTK signal

as well as receptor endocytosis and degradation by the lysosome (Waterman et al.,

1999; Garcia-Guzman et al., 2000). E3 ubiquitin ligases can also be regulated by the

attachment of Ubls, for example the Cullin subunit of Skip-Cullin-F-box E3 complexes

is neddylated, thereby activating the complex by enhancing the recruitment of the E2

enzyme Ubc4 (Kawakami et al., 2001).

1.7.2.4 Single Subunit RING E3 Ubiquitin Ligases

As the name suggests, single subunit E3 ubiquitin ligases are able to ubiquitinate target

proteins themselves, and this is due to the presence of a substrate recognition

component and E2 binding module on the same protein. For example, MDM2 contains

an amino-terminal p53 binding domain and a carboxy-terminal RING domain and is


40

therefore able to recruit p53 and the E2 enzymes, UbcH5 and E2-25K to ubiquitinate

p53, leading to its proteasome dependent degradation (Fang et al., 2000; Honda and

Yasuda, 2000; Saville et al., 2004).

1.7.2.5 Multisubunit RING E3 Ubiquitin Ligases

Multisubunit E3 ubiquitin ligases are complexes consisting of multiple proteins each

playing a distinct role within the complex. These include the E3 ubiquitin ligase

responsible for E2 recruitment and binding, the substrate recognition element and

adaptors. Three E3 ubiquitin ligase complexes that have been well studied are the APC

and SCF complexes, which regulate each other in a cell cycle dependent manner, and

the structurally related von Hippel Lindau-Cul2/elongin B/elongin C (VHL-CBC)

complex (Figure 1.13).

The APC is a large E3 ubiquitin ligase complex that consists of eleven core subunits

which associate in a cell cycle dependent manner with two different activators and

substrate adaptors, Cdc20 and Cdh1 (Thornton et al., 2006; McLean et al., 2011). The

APC has over 100 substrates, many of which are recognised and recruited to the

complex by Cdc20 and Cdh1 owing to the presence of short motifs such as the

Figure 1.13: Multisubunit E3 ubiquitin ligases. Well characterised multisubunit E3 ubiquitin ligases include SCF (left panel), VCB-CUL2 (middle) and APC (right panel) (Weissman, 2001)


41

destruction box (RxxL(x)nN/D/E‡) and KEN(x)nP§

Barford, 2011

box in the substrates (Figure 1.13)

( ; Meyer and Rape, 2011).

The APC has been proposed to consist of two subcomplexes, joined by a linker protein

APC1 (Thornton et al., 2006). The catalytic core of the APC consists of the cullin

protein, APC2 and the RING protein APC11, and together these two proteins are able to

bind the E2 activating enzyme and in vitro can ubiquitinate proteins but with little

substrate specificity (Figure 1.14A) (Tang et al., 2001). Also forming part of the core is

APC10, another substrate recognition component of the complex (Kurasawa and

Todokoro, 1999). The second subunit of the complex contains the tetrapeptide repeat

(TPR) containing proteins APC3, APC6, APC6 and APC8, which recruit Cdc20 and

Cdh1 to the complex (Figure 1.14A) (Vodermaier et al., 2003).

Another E3 ubiquitin ligase complex that is involved in cell cycle regulation is the SCF

E3 complex, which causes the degradation of cyclins, cyclin dependent kinase inhibitors

and transcription factors (Marti et al., 1999; Sutterluty et al., 1999; Nakayama et al.,

2004). SCF complex E3s consist of an invariable core containing the RING E3 Rbx1,

‡ R = arginine, L = leucine, N = asparagine, D = aspartic acid, E = glutamic acid x = any amino acid

§ K = lysine, E = glutamic acid, N = asparagine, P = proline

Figure 1.14: APC and SCF multisubunit E3 ubiquitin ligases. (A) The APC consists of 11 components including the RING domain protein Apc11 which binds the E2 conjugating enzyme and the activation components Cdc20 or Cdh1. (B) The SCF family of E3 ubiquitin ligases which contain the Rbx1 RING domain protein, Cul1 and the adaptor Skp1 that recruits F-box proteins, the substrate recognition components of the complex (Adapted from (Willems et al., 2004).


42

the cullin protein Cul1 and the adaptor Skp1, with the cullin protein interacting with

linker proteins in order to recruit substrates (Figure 1.13, 1.14B) (Willems et al., 2004).

The variable unit in the complex is the substrate recognition component, the F-box

protein, and over 70 F-box proteins have been identified in humans with many able to

recognise multiple substrates. Three of the best characterised SCF complex F-box

proteins are Skp2, Fbw7 and βTrCP (Skowyra et al., 1997; Yu et al., 1998). SCF E3

complexes ubiquitinate a multitude of proteins, usually phosphorylated proteins,

targeting them for degradation (Willems et al., 2004). For example SCFFbw, SCFβTrCP

and SCFSkp2 complexes recognise phosphorylated cyclin E, β-catenin and p27, with F-

box proteins of these complexes interacting directly through the phosphoresidues

(Orlicky et al., 2003; Wu et al., 2003a; Hao et al., 2005; Hao et al., 2007).

1.8 Outcomes of Ubiquitination

1.8.1 Ubiquitin Chain Topology Determines Ubiquitinated Protein Fate

Although the attachment of ubiquitin to a substrate is usually associated with its

proteasomal degradation, this is not the only function of ubiquitination, which can lead

to a variety of different outcomes for the substrate, depending on the type of ubiquitin

linkages (Figure 1.15) (Ikeda et al., 2010). Ubiquitin itself contains seven lysine

residues (Lys6, Lys11, Lys27, Lys29, Lys33, Lys48, and Lys63) all of which can form

chain linkages, whereby the carboxy-terminal glycine residue of the donor ubiquitin is

attached to the ε amino group of a specific lysine residue of the acceptor ubiquitin

molecule (Ikeda and Dikic, 2008). Additionally ubiquitin molecules can be linked in a

linear fashion (Tokunaga et al., 2009; Walczak et al., 2012). Ubiquitin chain formation

through lysine 48 and 63 of ubiquitin are referred to as canonical ubiquitination whereas

polyubiquitination through the remaining lysine residues and linear ubiquitination are

often referred to as atypical or non-canonical ubiquitination as much less is understood

about these types of ubiquitin linkages (Wickliffe et al., 2011).

Attachment of a single ubiquitin molecule at one or multiple sites on the substrate is

known as mono- or multi-monoubiquitination, and this signal does not target proteins

for degradation but rather is associated with a change in protein-protein interactions,

subcellular localisation or modified function (Figure 1.15) (Kerscher et al., 2006;

Deribe et al., 2010). For example, the transcription factor Met4 is polyubiquitinated by

the SCFMET30 complex E3 leading to its inability to associate with its cofactor Cbf1,


43

resulting in its transcriptional inactivation (Kaiser et al., 2000). The E3 ubiquitin ligase

MDM2 polyubiquitinates p53, targeting it for proteasome dependent degradation,

however under low MDM2 conditions, MDM2 monoubiquitinates p53, causing its

cytoplasmic accumulation (Li et al., 2003).

Polyubiquitination of a substrate through lysine 11 and 48 targets the protein for

degradation by the 26S proteasome (Chau et al., 1989; Kirkpatrick et al., 2006; Jin et

al., 2008). Mutation of lysine 48 of ubiquitin in yeast is lethal, with protein degradation

and therefore cell cycle progression disturbed, highlighting the importance of this

ubiquitin residue in proteasomal degradation (Chau et al., 1989; Xu et al., 2009).

Lysine 11 polyubiquitin chain linkages are involved in endoplasmic reticulum

associated degradation (ERAD), as evidenced by ER stress resulting in the

accumulation of lysine 11 linked ubiquitin chains (Adhikari and Chen, 2009; Xu et al.,

2009). The lysine 11 mutants lead to an increase in the unfolded protein response,

which is activated upon the accumulation of unfolded proteins in the ER lumen

(Adhikari and Chen, 2009; Xu et al., 2009).

Figure 1.15: The type of ubiquitination determines substrate protein fate. A single ubiquitin can be attached to a substrate protein (mono or multi-monoubiquitination) or multiple ubiquitin molecules can be attached via specific lysine residues, forming a polyubiquitin chain, with the topology of the ubiquitin chain determining the outcome of the ubiquitinated protein (Ye and Rape, 2009).


44

Ubiquitin linkages through lysine 63 are typically not associated with protein

degradation and mutation of yeast ubiquitin lysine 63 leads to abnormalities in DNA

repair (Spence et al., 1995). Ubiquitination through this lysine residue plays a role in

receptor endocytosis, DNA damage repair, protein trafficking and signalling (Spence et

al., 1995; Deng et al., 2000; Zhong et al., 2005; Sorrentino et al., 2008; Yang et al.,

2009; Boname et al., 2010). Lysine 63 ubiquitination is important for the activation of

IKK by TRAF6 in response to proinflammatory cytokines, resulting in the activation of

the transcription factor NF-κβ which regulates diverse processes such as inflammation,

apoptosis and immunity (Section 1.7.2.3) (Chen et al., 1996; Deng et al., 2000). The

most recent type of ubiquitin linkage to be discovered, linear ubiquitination, also plays a

role in the activation of the NF-κβ signalling pathway. Upon genoxotic stress or TNFα

exposure, the linear ubiquitination chain assembly complex, LUBAC ubiquitinates

NEMO, a regulator subunit of the IKK complex, resulting in the activation of the NF-κβ

pathway (Kirisako et al., 2006; Tokunaga et al., 2009; Niu et al., 2011).

Few studies have addressed the fate of proteins modified by the remaining ubiquitin

linkages (Figure 1.15). Lysine 6 ubiquitination has been associated with the prevention

of ubiquitin-mediated protein degradation in a study using biotinylated lysine 6 and

lysine 6 mutants in vitro and in mammalian cells respectively, in which lysine 6 linked

(biotinylated or K6W mutants) were resistant to proteasomal degradation (Shang et al.,

2005). Additionally, in response to DNA damage, BRCA1 has been associated with the

attachment of polyubiquitin linkages through lysine 6 to proteins in foci containing

BRCA1/BARD1 heterodimers (Morris and Solomon, 2004). The transcription factor

Jun, which together with c-Fos forms the transcription factor AP1, may be ubiquitinated

through lysine 27, resulting in its lysosomal localisation and degradation (Ikeda and

Kerppola, 2008). The attachment of lysine 29 polyubiquitin chains to proteins is

associated with both lysosomal and proteasomal protein degradation (Lindsten et al.,

2002; Chastagner et al., 2006). The E3 ubiquitin ligases Cbl-b and Itch induce T cell

receptor-ζ (TCR-ζ) lysine 33 polyubiquitination, modifying its phosphorylation and

protein interactions without targeting the TCR for degradation either by the proteasome

or lysosome (Huang et al., 2010). Aytpical or branched ubiquitin chains, which consist

of ubiquitin molecules linked through various ubiquitin lysines as well as mixed

ubiquitin and Ubl chain linkages have been described, however the cellular roles of

these linkages have not yet been elucidated (Ikeda and Dikic, 2008; Trempe, 2011).


45

1.8.2 Ubiquitin Binding Domains Determine Ubiquitinated Protein

Outcome

The reason for the variation in outcomes of ubiquitinated proteins is due to the presence

of specialised motifs known as ubiquitin binding domains (UBDs) in many cellular

proteins, which interact with ubiquitinated proteins through these motifs, thereby

interpreting the ubiquitin signal (Hurley et al., 2006). There are currently over 20 types

of UBDs found in over 150 proteins which utilise the ubiquitin β-sheet for their

interaction with ubiquitinated proteins (Dikic et al., 2009). UBDs differ in the type of

ubiquitin linkages that they recognise and therefore have different structures which can

be categorised according to the types of folds that they form, including α-helical, zinc

finger, ubiquitin conjugating domains and plekstrin homology folds (Dikic et al., 2009).

UBDs are found in a variety of proteins including those associated with the proteasome,

E2 conjugating enzymes, ERAD, DNA repair, endocytosis, multivesicular body

biogenesis, kinase regulation and cell signalling (Young et al., 1998; Hofmann and

Falquet, 2001; VanDemark et al., 2001; Fisher et al., 2003; Swanson et al., 2003; Prag

et al., 2005; Stamenova et al., 2007; Fu et al., 2009).

UBDs that recognise monoubiquitin include ubiquitin interacting motifs (UIMs),

ubiquitin binding zinc finger (UBZ) and ubiquitin-associated domains (UBA). For

example, all Y-family translesion synthesis family polymerases, which function in

translesion synthesis upon DNA damage, contain UBM and UBZ domains which

recognise monoubiquitinated PCNA (Lehmann et al., 2007). PCNA monoubiquitination

following DNA damage recruits Y-family polymerases to sites of DNA damage,

allowing the switch from DNA replication polymerases to translesion synthesis

polymerases (Bienko et al., 2005).

Proteins targeted to the proteasome are recognised by ubiquitin receptors, which are

associated with the proteasome, either stably or transiently. These receptors bind the

polyubiquitin chain through their UBDs and bind the proteasome through their

ubiquitin-like domain (Ubl), exhibiting a high affinity for lysine 48 linked ubiquitin

chains (Schauber et al., 1998; Verma et al., 2004). Proteins containing domains that

recognise ubiquitin play an integral part in the ubiquitin system by determining the

consequences of protein ubiquitination. As such the existence of diverse UBDs is

essential for the correct interpretation of the ubiquitin signal and the regulation of


46

ubiquitinated proteins, leading to their degradation or their function in novel cellular

roles.

1.9 E3 Ubiquitin Ligases and Cancer

Given the involvement of the ubiquitin system in many different cellular processes such

as DNA repair, the cell cycle, proteasomal and lysosomal degradation, cell signalling,

protein trafficking and ERAD, it is not surprising that abnormalities in proteins that

form part of the ubiquitin system are implicated in pathological states including cancer.

The substrate recognition components of the ubiquitin system including E3 ubiquitin

ligases and DUBs are often dysregulated in cancer and are therefore potential targets for

drug development. It is not surprising when their expression is disrupted in cancer, that

several E3 ubiquitin ligases are considered oncogenes or tumour suppressor genes

depending upon the function of their cellular substrates (Lipkowitz and Weissman,

2011). A few E3 ubiquitin ligases may play either role in malignancy due to the diverse

roles of a single substrate or the roles of multiple substrates (Lipkowitz and Weissman,

2011).

1.9.1 E3 Ubiquitin Ligases and the Cell Cycle

Mutation or abnormal expression of members of the APC and SCF E3 ubiquitin ligase

complexes which regulate the cell cycle have been documented in cancers (Section

1.7.2.5). Several members of the APC are mutated in colon cancer cells and

overexpression of an APC8/CDC23 mutant in colon epithelial cells leads to abnormal

levels of cyclin B1 and dysregulated cell cycle progression (Wang et al., 2003a).

Knockout of the Cdc20 substrate adaptor of the APC in mouse embryos results in cell

cycle inhibition at mitosis when embryos are at the two-cell stage (Li et al., 2007a).

Mice lacking Cdh1 (Cdh1-/-), the second APC substrate adaptor, exhibit genomic

instability and do not survive past embryonic day 12.5, while aged Cdh1+/- heterozygous

mice (25 months) develop epithelial neoplasias in a number of tissues, suggesting that

Cdh1 functions as a tumour suppressor gene (Li et al., 2007a; Garcia-Higuera et al.,

2008).

High levels of SKP2, the substrate recognition element in SCFSKP2 E3 ubiquitin ligase

complexes have been detected in a number of cancers (Hershko et al., 2001; Latres et

al., 2001). High SKP2 levels are predicted to be oncogenic due its control of the G1/S


47

checkpoint by ubiquitinating and targeting for degradation the CDK inhibitor p27

(Skaar and Pagano, 2009). In high grade lymphomas, breast and colorectal tumours,

high SKP2 levels inversely correlate with p27 levels suggesting that a direct effect of

SKP2 on p27 leads to a failure in G1/S checkpoint control and aberrant cellular

proliferation (Hershko et al., 2001; Latres et al., 2001; Traub et al., 2006). The

oncogenic potential of SKP2 is supported by several observations in Skp2 mouse

models, with Skp2-/- mice expressing increased p27 levels and exhibiting reduced cell

growth (Lin et al., 2010).

1.9.2 E3 Ubiquitin Ligases and DNA Damage

1.9.2.1 Tumour Suppressor p53

The transcription factor p53 regulates cellular responses to DNA damage by inducing

cell cycle arrest or apoptosis (Figure 1.16) (Livingstone et al., 1992; Shaw et al., 1992;

O'Connor et al., 1993). p53 is negatively regulated by the E3 ubiquitin ligases MDM2

and MDMX, with MDMX enhancing MDM2 E3 ubiquitin ligase activity toward p53

(Honda et al., 1997; Stad et al., 2001). MDM2 is the primary regulator of p53, with

MDM2 ubiquitinating p53 and itself, targeting both proteins for proteasomal

degradation (Haupt et al., 1997; Honda et al., 1997; Honda and Yasuda, 2000). The

MDM2 gene is disrupted in a number of cancers, with its amplification documented in

osteosarcomas, oesophageal carcinomas and breast cancers (Oliner et al., 1992; Quesnel

et al., 1994; Momand et al., 1998). Mdm2-/- knockout mice exhibit embryonic lethality

due to p53 dysregulation, whilst crossing Mdm2-/- mice with p53-/- mice rescues this

phenotype (Jones et al., 1995; Montes de Oca Luna et al., 1995). Mice expressing low

levels of Mdm2 display reduced tumour formation due to higher levels of p53, and in

Mdm2 null mice, expression of inducible p53 results in growth arrest (Mendrysa et al.,

2006; Ringshausen et al., 2006).

These studies highlight the importance of the balance between MDM2 and p53 which is

supported by the finding that approximately 33% of human sarcomas that display wild-

type p53 exhibit MDM2 amplification, thereby suggesting that MDM2 overactivation

represents one mechanism by which cells escape growth regulation (Leach et al., 1993;

Marine and Lozano, 2010). Several compounds have been designed to reduce MDM2

activity or MDM2-p53 binding, including the small molecule inhibitor Nutlin-3a.

Nutlin-3a disrupts MDM2-p53 interaction, causing cell cycle arrest or apoptosis in


48

tumour cells, with the apoptotic response strongest in cells overexpressing MDM2

(Tovar et al., 2006). MDM2 activity is regulated by other proteins such as the tumour

suppressor, ARF and posttranslational modifications such as neddylation,

phosphorylation and acetylation of p53, MDM2 and MDMX (Pise-Masison et al., 1998;

Pomerantz et al., 1998; Luo et al., 2004; Xirodimas et al., 2004; Wade et al., 2010).

Additionally, p53 is regulated by several other E3 ubiquitin ligases including CHIP,

PIRH2 and TOPORS (Leng et al., 2003; Rajendra et al., 2004; Esser et al., 2005).

Therefore, although dysregulated expression of MDM2 is present in a percentage of

human tumours that express wild-type p53, potentially representing a therapeutic target

in these tumours, abnormal p53 levels or function in tumours exhibiting wild-type p53

may be due to the altered function of other E3 ubiquitin ligases and proteins regulating

p53 activity.

Figure 1.16: p53 regulation in response to genotoxic stress. Upon genotoxic stress, p53 and MDM2 undergo post-translational modification, disrupting their association and preventing MDM2 mediated p53 ubiquitination. Additionally, ARF and ribosomal proteins interact with MDM2 and inhibit its ability to bind p53, leading to an increase in p53 activity and the induction of proteins involved in cell cycle arrest or apoptosis (Lipkowitz and Weissman, 2011).


49

1.9.2.2 BRAC1/BARD1

The E3 ubiquitin ligase BRCA1 is frequently mutated in familial breast and ovarian

cancers and plays an important role in the sensing of DNA damage and in DNA damage

repair (Welcsh and King, 2001). Sporadic breast tumours can also exhibit a reduction or

loss of BRCA1 expression, with BRCA1 levels and tumour grade being negatively

correlated (Taylor et al., 1998; Wilson et al., 1999). BRCA1 functions as a heterodimer

with BARD1 in several complexes which are involved in DNA damage detection, cell

cycle checkpoint regulation and recruitment of DNA damage repair proteins (Cantor et

al., 2001; Yarden et al., 2002; Wang et al., 2007). For example, BRCA1 interacts with

serine 406 phosphorylated abraxas and receptor associated protein 80 (RAP80), which

contains a UIM (Wang et al., 2007). Following DNA damage, BRCA1/BARD1-

abraxas-RAD80 complexes are localised to foci of DNA damage (Wang and Elledge,

2007, Wang et al., 2007). This is due to interaction of the UIM of RAP80 with proteins

such as histones that become ubiquitinated as a consequence of DNA damage repair

signalling pathways involving the E3 ubiquitin ligases RNF8 and RNF168 (Wang and

Elledge, 2007; Wang et al., 2007; Yan et al., 2007). Although the precise role of

BRCA1/BARD1 heterodimers in DNA repair once localised to DNA double strand

breaks (DSB) is unknown, it is hypothesised that BRCA1/BARD1 ubiquitinates

proteins at these sites. This is evidenced by the requirement of BRCA1/BARD1 for the

accumulation of ubiquitinated proteins at sites of DNA damage (Mallery et al., 2002;

Morris and Solomon, 2004; Polanowska et al., 2006). Members of the BRCA1/BARD

complex also form part of the BCRA1/BARD1-abraxas-RAD80BRCC36-BRCC45

complex required for regulation of the G2/M checkpoint, which ensures that

chromosomal segregation has occurred before entry into mitosis (Kim et al., 2007;

Sobhian et al., 2007).

1.9.3 E3 Ubiquitin Ligases and Signal Transduction

The CBL proteins, Cbl (also known as c-Cbl), Cbl-b and Cbl-c are a family of E3

ubiquitin ligases that also function as adaptor proteins, thereby acting as both positive

regulators propagating RTK downstream signalling or as negative regulators by

ubiquitinating RTKs, resulting in the internalisation and trafficking of the receptor for

recycling or degradation by the lysosome (Figure 1.17) (Levkowitz et al., 1998;

Ettenberg et al., 1999; Joazeiro et al., 1999; Kales et al., 2010). Mutations in RTKs

causing constitutive activation of the receptor in the absence of ligand or amplification


50

of RTK genes are common in cancers (Blume-Jensen and Hunter, 2001).

Approximately 5% of human myeloid neoplasms contain mutations that inactivate the

E3 ubiquitin ligase function of CBL implicating a role for CBL as a tumour suppressor

gene (Kales et al., 2010). Mutations in Cbl that inactivate its E3 ubiquitin ligase activity

typically occur within the RING domain and linker region (Lipkowitz and Weissman,

2011). The linker region which is located immediately amino-terminal to the RING

domain is phosphorylated, resulting in a conformational change in CBL and the

activation of the RING domain (Kassenbrock and Anderson, 2004). CBL mutants are

proposed to act via two mechanisms. Firstly, they no longer ubiquitinate RTKs which

are therefore not internalised and degraded or recycled. Secondly, the RING domain

inactive mutants are hypothesised to function in a dominant negative manner by binding

to the activated RTK and thereby blocking the binding of other CBL proteins. The

observation that Cbl and Cbl-b double knockout mice develop early onset

myeloproliferative disease provides support for the tumour suppressor role of CBL

(Naramura et al., 2010).

CBL also exhibits oncogenic activity under specific circumstances as evidenced by the

observation that CBL RING domain mutants which have therefore lost their E3

ubiquitin ligase activity do not induce cellular transformation and that mutations in both

RING domain and linker region are required for transformation (Thien et al., 2001).

This is supported by the finding that a deletion encompassing parts of the linker region

and RING domain of Cbl results in the activation of the epidermal growth factor

receptor (EGFR) and Flt3 in the absence of ligand, and in the presence of ligand

enhances EGFR activity (Thien and Langdon, 1997; Sargin et al., 2007; Sanada et al.,

2009). Furthermore, the oncogene v-Cbl (amino acids 1-355 of CBL), the transforming

gene from the Cas NS-1 murine retrovirus, which lacks the Cbl RING domain, induces

the development of murine lymphomas and leukaemias (Langdon et al., 1989). Studies

in NIH 3T3 cells have demonstrated that a CBL mutant containing the first 357 amino

acids of the CBL protein (similar to v-Cbl), functions by associating with RTKs and

activating RTK mediated signalling (Bonita et al., 1997). The transforming properties

of Cbl mutations are therefore hypothesised to result from the loss of the tumour

suppressor functions of Cbl and unmasking of its oncogenic activity facilitated by

amino-terminal regions which mediate RTK signalling (Kales et al., 2010).


51

By targeting proteins for ubiquitin-dependent degradation, E3 ubiquitin ligases control

the levels of oncogene and tumour suppressor gene products. Signal transduction

regulators, DNA repair related proteins and cell cycle regulators are all regulated by E3

ubiquitin ligases, with aberrant expression of the E3s hypothesised to result in the

enhanced degradation of tumour suppressors and accumulation of oncogene products

(Kitagawa et al., 2009). Depending on the ubiquitination substrate of the E3 ubiquitin

ligase and its function, the E3 itself can act as an oncogene or tumour suppressor. From

these examples it is evident that disruption of E3 ubiquitin ligases plays a role in the

initiation and/or progression of multiple cancers, highlighting the diverse roles of these

proteins in the regulation of cellular functions.

1.10 Statement of Aims

The homeodomain containing transcription factor NKX3.1 regulates prostatic

development and is expressed in the luminal epithelial cells of the adult prostate (He et

al., 1997; Bhatia-Gaur et al., 1999; Bowen et al., 2000; Chen et al., 2002a). NKX3.1

expression is reduced or undetectable in up to 80% of prostate tumours and it is

proposed to function as a prostate-specific tumour suppressor (Bowen et al., 2000;

Asatiani et al., 2005). Evidence of discordance between NKX3.1 mRNA and protein

levels and its aberrant cytoplasmic mislocalisation have suggested that translational or

post-translational dysregulation may contribute to a loss of NKX3.1 function in prostate

tumours (Kim et al., 2002b; Bethel et al., 2006; Bethel and Bieberich, 2007). Few

Figure 1.17: Roles of CBL in the regulation of receptor tyrosine kinase signalling. CBL ubiquitinates RTKs resulting in their internalisation and lysosomal degradation and thereby functioning as a tumour suppressor. CBL also functions as an oncogene by acting as an adaptor and activating downstream RTK signalling pathways (Kales et al., 2010).


52

NKX3.1 interacting proteins that regulate NKX3.1 post-translational modification have

been identified, therefore in order to identify NKX3.1 binding partners that potentially

modify its activity or expression, a yeast two hybrid analysis was performed in our

laboratory which determined the FLJ22318 gene product, later renamed RMND5B to

be an NKX3.1 interacting protein (Dawson, 2006). RMND5B and its homologue

RMND5A are named after their yeast orthologue, RMD5 a RING domain containing E3

ubiquitin ligase (Santt et al., 2008) and as RMND5A and RMND5B each contain

putative RING domains, it is feasible that they too function as E3 ubiquitin ligases. To

determine the biological activity of human RMND5A and RMND5B, the aims of this

thesis were:

1. To characterise the E3 ubiquitin ligase activity of RMND5 proteins in prostate cancer

cells.

2. To determine whether RMND5A and/or RMND5B ubiquitinate NKX3.1 and to

characterise the outcome of NKX3.1 ubiquitination by RMND5 proteins.

3. To identify additional RMND5A and RMND5B interacting proteins.

Chapter 2 Materials

Chapter 2: Materials

Chapter 2 Materials

53

2.1 Reagents 2.1.1 Cell Culture Item Supplier Ammonium Chloride Sigma Aldrich, USA Charcoal Stripped Serum Sigma Aldrich, USA Chloroquine Sigma Aldrich, USA Cycloheximide AG Scientific, USA Dihydrotestosterone Sigma Aldrich, USA DU145 Prostate Cancer Cell Line American Type Culture Collection, USA Foetal Calf Serum Trace Scientific Ltd, Australia Lactacystin Sigma Aldrich, USA LNCaP Prostate Cancer Cell Line American Type Culture Collection, USA Metafectine PRO® Reagent Biontex Laboratories, GmbH MG132 AG Scientific, USA Penicillin/Streptomycin Gibco® Life Technologies, Australia RPMI 1640 with L-Glutamine Thermo Electron Corporation, Australia Sodium Hydrogen Carbonate Merck Pty Ltd., Germany Trypsin/EDTA (0.25%) Gibco® Life Technologies, Australia 2.1.2 Primers Primer sequences are located in Appendix II Item Supplier ARMC81103-S Geneworks, Australia ARMC81668-AS Geneworks, Australia β Actin146-S Geneworks, Australia β Actin464-AS Geneworks, Australia C17orf39298-S Geneworks, Australia C17orf39814-AS Geneworks, Australia CBLRING1141-S Geneworks, Australia CBLRING1257-AS Geneworks, Australia EMP1083-AS Geneworks, Australia EMP580-S Geneworks, Australia Muskelin1675-S Geneworks, Australia Muskelin2120-AS Geneworks, Australia pEGFP1266-S Geneworks, Australia M13-S Geneworks, Australia M13-AS Geneworks, Australia pGEX-AS Geneworks, Australia pGEX-S Geneworks, Australia RanBPM1-S Geneworks, Australia RanBPM1029-S Geneworks, Australia RanBPM1259-AS Geneworks, Australia RanBPM1550-S Geneworks, Australia RanBPM2190-AS Geneworks, Australia RanBPM687-S Geneworks, Australia RMND5A(C356A)1045-S Geneworks, Australia RMND5A(C356A)1083-AS Geneworks, Australia RMND5A(C356A/H358A)1045-S Geneworks, Australia RMND5A(C356A/H358A)1083-AS Geneworks, Australia RMND5A(C356S)1045-S Geneworks, Australia RMND5A(C356S)1083-AS Geneworks, Australia

Chapter 2 Materials

54

RMND5A1-S Geneworks, Australia RMND5A1176-AS Geneworks, Australia RMND5A490-AS Geneworks, Australia RMND5A603-S Geneworks, Australia RMND5ABamHI1-S Geneworks, Australia RMND5ARING1006-S Geneworks, Australia RMND5ARING1131-AS Geneworks, Australia RMND5B(C358A/H360A)1083-AS Geneworks, Australia RMND5B(C358A/H360S)1054-S Geneworks, Australia RMND5B(C358S)1051-S Geneworks, Australia RMND5B(C358S)1057-S Geneworks, Australia RMND5B(C358S)1080-AS Geneworks, Australia RMND5B(C358S)1089-AS Geneworks, Australia RMND5B790-S Geneworks, Australia RMND5BBamHI 1-S Geneworks, Australia RMND5BBamHI 1182-AS Geneworks, Australia RMND5BRING1012-S Geneworks, Australia RMND5BRING1137-AS Geneworks, Australia RMND5BSalI 1-S Geneworks, Australia RMND5BSalI 1182-AS Geneworks, Australia RMND5BTOPO 1182-AS Geneworks, Australia Twa1335-S Geneworks, Australia Twa1681-AS Geneworks, Australia 2.1.3 Reverse Transcription - Polymerase Chain Reaction (PCR) Item Supplier AmpliTaq Gold® 360 DNA Polymerase Applied Biosystems, Australia AmpliTaq Gold® Mastermix Applied Biosystems, Australia DMSO Thermo Fischer Scientific, Finland DTT (0.1M) Invitrogen, Australia dNTP 100mM (dATP,dGTP,dCTP,dTTP) Promega, Australia 5x First Strand Buffer Invitrogen, Australia MgCl2 (50mM) Invitrogen Technologies, Australia MgCl2 (25mM) Applied Biosystems, Australia Oligo(dT) Primer 0.5µg/µL Promega, Australia 10x PCR Buffer (No MgCl2) Invitrogen, Australia 5x Phusion® HF Buffer Thermo Fischer Scientific, Finland Phusion® High-Fidelity DNA Polymerase Thermo Fischer Scientific, Finland 5x Phusion® High GC Buffer Thermo Fischer Scientific, Finland Platinum Taq Polymerase 5U/µL Invitrogen, Australia Sterile ddH2O Baxter Healthcare Pty Ltd, Australia Superscript II Reverse Transcriptase® Invitrogen, Australia 2.1.4 Plasmids Item Supplier pcDNA3.1-V5 Invitrogen, Australia pCMV-HA-Ubiquitin Gift from Professor Wallace Langdon,

University of Western Australia pGEX-2T-CBL Gift from Professor Wallace Langdon,

University of Western Australia pEGFP-C2 Clontech, USA

Chapter 2 Materials

55

pGEM®-T Easy Cloning Vector Promega, Australia pGEX-2TK GE Healthcare Life Sciences, Australia pmCherry C1 Gift from Dr Archa Fox, West Australian

Institute for Medical Research 2.1.5 Cloning Item Supplier Agar Bacteriological Amresco®, USA Ampicillin Amresco®, USA BamHI-HF Restriction Enzyme New England Biolabs, USA 10x Buffer 4 Promega, Australia 10x Buffer L Invitrogen, Australia DpnI Restriction Enzyme New England Biolabs, USA EcoR1-HF Restriction Enzyme New England Biolabs, USA Escherichia coli BL21 GE Healthcare Life Sciences, Australia HindIII Restriction Enzyme New England Biolabs, USA Escherichia coli DH5α Invitrogen Technologies, Australia Kanamycin CSL Limited, Australia KpnI Restriction Enzyme Invitrogen, Australia NdeI Restriction Enzyme Invitrogen, Australia 10x React Buffer 6 Invitrogen, Australia RNaseA Roche Diagnostics, Germany Rosetta Escherichia coli BL21 Gift from Dr Evan Ingley, West Australian

Institute for Medical Research SalI-HF Restriction Enzyme New England Biolabs, USA SAP Dephosphatase Buffer Roche Diagnostics, Germany Shrimp Alkaline Phosphatase (SAP) Roche Diagnostics, Germany T4 DNA Ligase New England BioLabs, UK 10x T4 DNA Ligase Reaction Buffer New England BioLabs, UK Tryptone Amresco®, USA Yeast Extract Becton Dickinson, Australia 2.1.6 GST Fusion Protein Production Glutathione S Transferase Agarose Beads GE Healthcare Life Sciences, Australia

Isoproyl β-D-1-thiogalactopyranoside(IPTG) Astral Scientific, Australia Lysozyme (10mg/mL) Amresco®, USA

Dithiothreitol (DTT) Sigma Aldrich, USA L-Glutathione, reduced Sigma-Aldrich, USA Coomassie Brilliant Blue G Sigma Aldrich, USA 2.1.7 Immunoprecipitation Item Supplier Anti-GFP Microbeads Miltenyi Biotec, Germany Complete Protease Inhibitor Tablets Roche Diagnostics, Germany Protein A Microbeads Miltenyi Biotec, Germany Protein A Sepharose Beads GE Healthcare, Australia Protein G Microbeads Miltenyi Biotec, Germany Protein G Sepharose Beads GE Healthcare, Australia Sodium Deoxycholate Sigma Aldrich, USA Sodium Orthovanadate Sigma Aldrich, USA

Chapter 2 Materials

56

2.1.8 Western Blotting Item Supplier 40% Acrylamide Amresco®, USA Ammonium Persulphate BioRad, Australia BSA Fraction V High-Grade Fatty Acid Free Roche Diagnostics, Germany ColorPlus Prestained Protein Marker New England Biolabs, USA Donkey Anti-Goat HRP Conjugate Santa Cruz Biotechnology Inc., USA ECLTMWestern Blotting Detection Reagent GE Healthcare, Australia Goat Anti-Actin IgG Santa Cruz Biotechnology Inc., USA Goat Anti-NKX3.1 IgG Santa Cruz Biotechnology Inc., USA Goat Anti-Rat IgG Jackson ImmunoResearch Europe, UK HybondTMC Extra Nitrocellulose Membrane GE Healthcare, Australia 2-Mercaptoethanol BDH Chemicals, Australia Mouse Anti-GFP IgG Clontech, USA Mouse Anti-HA HRP Conjugate Cell Signaling Technology, USA Mouse Anti-RFP IgG Clontech, USA Mouse Anti-V5 IgG Invitrogen, Australia 4-12% Precast Bis-Tris Acrylamide Gels Invitrogen, Australia 12% Precast Polyacrylamide Gels BioRad, Australia Rat Anti-Cherry IgG ChromoTek, Germany Sheep Anti-Mouse HRP Conjugate Chemicon, Australia Skim Milk Powder Bonlac Foods Inc, Australia Tetramethylethylenediamine (TEMED) BioRad, Australia Tween-20 Sigma Aldrich, USA 2.1.9 Fluorescence Microscopy Item Supplier Anti-goat Alexa Fluor® 546 Molecular Probes, USA Chlorobutanol Sigma Aldrich, USA 40% Formaldehyde BDH Chemical, Australia Hœchst 33258 Dye Sigma Aldrich, USA Normal Horse Serum GIBCO® , Australia Immersion Oil for Fluorescence Microscopy Leitz Wetzlar, Germany Phalloidin TRITC 77418 Sigma Aldrich, USA Polyvinyl Alcohol Sigma Aldrich, USA Sodium Azide BDH Chemicals, Australia 2.1.10 General Laboratory Reagents Item Supplier Ammonium Sulphate Sigma Aldrich, USA Agarose ITM Amresco®, USA Big DyeTM Terminator Sequencing Buffer Applied Biosystems, USA Big DyeTM Terminator Applied Biosystems, USA Bromophenol Blue Sigma Aldrich, USA Calcium Chloride Ajax Chemicals, Australia Chloroform Ajax Chemicals, Australia Diethylpyrocarbonate (DEPC) Sigma Aldrich, USA Disodium Hydrogen Orthophosphate BDH Chemicals, Australia Dimethyl Sulfoxide (DMSO) Thermo Fischer Scientific, Finland Ethanol Rowe Scientific, Australia Ethidium Bromide ICN Biochemicals, USA

Chapter 2 Materials

57

Ethylenediaminotetraacetic Acid (EDTA) BDH Chemicals, Australia Glacial Acetic Acid BDH Chemicals, Australia Glycerol Sigma Aldrich, USA Glycine BioRad, Australia Hydrochloric Acid BDH Chemicals, Australia Isopropanol BDH Biochemicals, England Magnesium Chloride BDH Chemicals, Australia Methanol Biolab, Australia Igepal CA630 (NP-40) Sigma Aldrich, USA Orthophosphoric Acid Ajax Finechem, Australia Piperazinediethanesulfonic acid (PIPES) Amresco®, USA Phenylmethanesulfonylfluoride (PMSF) Sigma Aldrich, USA Potassium Acetate BDH Chemicals, Australia Potassium Chloride BDH Chemicals, Australia Potassium Dihydrogen Orthophosphate BDH Chemicals, Australia RQ1 RNase Free DNase Promega, Australia RQ1 Stop Solution Promega, Australia Sodium Acetate BDH Chemicals, Australia Sodium Chloride Rowe Scientific, Australia Sodium Dodecyl Sulphate BioRad, Australia Sodium Fluoride Sigmal Aldrich, USA Sodium Hydroxide Sigma Aldrich, USA Sodium Phosphate BDH Chemicals, Australia Sodium Vanadate Sigmal Aldrich, USA Sucrose Sigma Aldrich, USA Tris Amresco®, USA Triton X-100 Sigma Aldrich, USA Zinc Chloride Sigma Aldrich, USA 2.2 Commercial Kits Item Supplier PureLinkTM HiPure Plasmid Midiprep Life Technologies, Australia Kit (Catalog Number K210004) Equilibration Buffer, EQ1 Resuspension Buffer, R3 Lysis Buffer, L7 Precipitation Buffer, N3 Wash Buffer, W8 Elution Buffer, E4 TE Buffer QIAquick Gel Purification Kit Qiagen, Australia (Catalog Number 28106) QIAquick Spin Columns Buffer QG Buffer PB Buffer EB Collection Tubes (2mL) Loading Dye

Chapter 2 Materials

58

QIAquick PCR Purification Kit Qiagen, Australia (Catalog Number 28706) QIAquick Spin Columns Buffer PBI Buffer PB Buffer EB Collection Tubes (2mL) Loading Dye Quant-iTTM dsDNA BR Assay Kit Life Technologies, Australia (Catalog Number Q32853) Quant-iTTM dsDNA BR Reagent (Component A) Quant-iTTM dsDNA BR Buffer (Component B) Quant-iTTM dsDNA BR Standard #1 (Component C) Quant-iTTM dsDNA BR Standard #2 (Component D) Ubiquitinylation Kit Enzo Life Sciences, USA (Catalog Number BML-UW9920-001) 20X Ubiquitin Activating Enzyme Solution (E1) 10X Ubiquitin Conjugating Enzyme Solutions (E2) - UbcH1 - UbcH2 - UbcH3 - UbcH5a - UbcH5b - UbcH5c - UbcH6 - UbcH7 - UbcH8 - Ubc13/Mms2 - UbcH10 20x Biotinylated Ubiquitin Solution (Bt-Ub) 20x Mg-ATP Solution 2x Non-reducing Gel Loading Buffer 10x Ubiquitinylation Buffer UltraspecTM RNA Isolation System Fischer Biotech, Australia (Biotecx Laboratories, Catalog NumberBL-10200) UltraspecTM RNA µMACSTM GFP Purification Kit Miltenyi Biotech, Germany (Catalog Number 130-091-125) anti-GFP microbeads Lysis Buffer Wash Buffer 1 Wash Buffer 2 Elution Buffer Vectastain Elite ABC Streptavidin-HRP Kit Vector Laboratories, USA (Catalog Number PK-6100) Solution A

Chapter 2 Materials

59

Solution B 2.3 Equipment Item Supplier 0.2µM Filters Pall Corporation, USA 23G Needles Becton Dickinson, USA AGFA CP-1000 Film Developer AGFA-Gevaert NV, Belgium Avanti J-25I Centrifuge Beckman Coulter Inc., Australia Avanti JA-25-5 Rotor Beckman Coulter Inc., Australia C1000 Thermal Cycler Bio-rad Laboratories, USA CellStar® Tissue Culture 10cm Dishes Greiner Labortechnik, Germany DNA Sub CellTM Electrophoresis Tank Bio-rad Laboratories, USA Dry Block Heater Thermoline, Australia Eppendorf 0.5, 1.5 mL Microcentrifuge Eppendorf, Germany Tubes 5mL, 10mL, 50mL Tubes Sarstedt, Germany Eppendorf 5415R Centrifuge Eppendorf, Germany Eppendorf 5804R Centrifuge Eppendorf, Germany Glass Coverslips Menzel-Glaser, Germany Haemocytometer Hawksley, UK Heidolph MR 100 Magnetic Stirrer John Morris Scientific Pty Ltd, Australia Hoefer Mini VE Electrophoresis Unit Hoefer Inc., USA L420S Precision Balance Sartorius, Germany Laminar Flow Hood Email-Westinghouse, Australia Microscope Coverslips Menzel-Glaser®, Germany Mini Sub DNA CellTM Electrophoresis Tank Bio-rad Laboratories, USA NanoDrop® ND-1000 Spectrophotometer Biolab Australia, Australia Nikon Eclipse TS100 Microscope Nikon, Japan Nikon Eclipse TiE Inverted Microscope Nikon, Japan Nunc Cryotubes Inter Med, Denmark Olympus IX71 Inverted Microscope Olympus, USA Orbital Mixer Incubator Ratek Instruments Pty.Ltd., Australia Parafilm American National CanTM, USA PHM 83 AutoCal pH Meter Radiometer, Copenhagen pH Cube pH Meter TPS, Australia Pipetman® Automatic pipettes Gilson, USA Pipette Tips Sarstedt, USA PowerPacTM 300 and 3000 Bio-rad Laboratories, USA PTC-100TM Programmable Thermal Cycler MJ Research Inc, Australia Protean® 3 Cell Acrylamide Gel Apparatus Bio-rad Laboratories, USA Red Rotor Orbital Shaker Hoefer Scientific, Australia Rotator 360º Ratek Instruments Pty.Ltd., Australia Sanyo Incubator Sanyo Electric Co., Japan Sharp Microwave Sharp, Australia Syringes (1mL) Becton Dickinson, USA Tissue Culture Flasks (T75, T25) Sarstedt, USA Tissue Culture Plates Becton Dickinson, USA Transfer Apparatus Bio-rad Laboratories, USA UV Sterile Cabinet Starkeys, Australia UV Transilluminator Hoefer Scientific, Australia Whatman Paper 3mm Whatman International, Ausltralia Zx3 Vortex VELP® Scientifica, Italy

Chapter 2 Materials

60

X-ray film (CL-XPosure) Thermo Fischer Scientific, Finland 2.4 Computer Software Item Supplier Adobe Illustrator Adobe Systems Inc., USA Adobe Photoshop CS Adobe Systems Inc., USA EndNote X1 ISI ResearchSoft, USA Image Pro Plus (Autoquant) Media Cybernetics, USA Microsoft Office® Microsoft® Corporation, USA Nanodrop 1000 Software Thermo Scientific, USA NIS Elements Nikon, Japan Quantity One® Imaging and Quantitation Bio-rad Laboratories, USA Software

Chapter 3 Methods

Chapter 3: Methods

Chapter 3 Methods

61

3.1 Cell Culture

3.1.1 Routine Maintenance of Mammalian Cell Lines

The human prostate cancer cell lines, DU145 and LNCaP obtained from the American

Tissue Culture Collection (ATCC) were used in these studies. Cells were maintained in

75cm2 culture flasks in ~10mL RPMI 1640 medium71*

supplemented with 10% (v/v)

foetal calf serum (FCS), 100U/mL penicillin and streptomycin (RPMI/PS/10%FCS74) in

humidified incubators at 37 ºC and 5% CO2. Medium was replaced every 2-3 days and

cells were passaged every 5-7 days as required.

To passage cells, RPMI/PS/10%FCS74 was aspirated, cells were rinsed with ~2mL

PBS56, then 1.5mL trypsin/EDTA was added to each flask and the flasks were incubated

at 37°C for ~2 minutes to dislodge the cells. RPMI/PS/10%FCS74 medium was added to

inactivate the trypsin and the cells were aliquoted into fresh 75cm2 flasks or used as

required (Section 3.1.3, 3.1.4, 3.1.5). For routine maintenance of cells, DU145 cultures

were passaged at a 1:10 dilution while LNCaP cells were passaged at a 1:5 dilution.

3.1.2 Cryopreservation and Thawing of Mammalian Cells

Cell lines growing in 75cm2 flasks were trypsinised when ~90% confluent (Section

3.1.1) and RPMI/PS/10%FCS74 was added to inactivate the trypsin. Cell suspensions

were centrifuged at 1000rpm for 5 minutes at room temperature, the supernatant

removed and the cells resuspended in 1mL per 75cm2 flask

RPMI/PS/10%FCS/10%DMSO75. Cell suspensions were aliquoted into pre-cooled 2mL

cryotubes, frozen at -80ºC in insulated containers then transferred to liquid nitrogen

storage.

Mammalian cells stored in liquid nitrogen were rapidly thawed in a waterbath at 37ºC

and the thawed cell suspensions pipetted into 75cm2 flasks containing pre-warmed

RPMI/PS/10%FCS74. Flasks were incubated overnight at 37ºC and 5% CO2, the

following morning the medium was replaced with fresh RPMI/PS/10%FCS74 and the

cells were cultured as usual (Section 3.1.1).

* Buffers and Solutions referenced by superscript numbers are described in Appendix I.

Chapter 3 Methods

62

3.1.3 Preparation of Cells for Fluorescence Microscopy

For microscopic imaging, cells were grown on glass coverslips in 6 well plates. Prior to

commencement of the experiments, coverslips were wiped with 70% ethanol25, placed

one per well into 6-well tissue culture plates then exposed to UV light in a laminar flow

hood for 45 minutes to sterilise. To wet the coverslips, ~200µL RPMI/PS/10%FCS74

was placed on the centre of each coverslip and culture plates were incubated at 37°C

and 5% CO2 for 1-2 hours prior to seeding of the cells. Cells were trypsinised, counted

using a haemocytometer, medium was removed from the coverslips and the appropriate

volume of cell suspension was added to each coverslip (Sections 3.1.1, 3.1.5) (Table

3.1). Cells were incubated overnight and the following day were transfected with the

appropriate plasmid DNA (Section 3.1.4), then incubated a further 48 hours before

processing for microscopy (Section 3.16). To preserve fluorescence, coverslips were

processed in minimal light.

3.1.4 Transfection of Mammalian Cells

Transient transfection of the cell lines was carried out using MetafectineTM PRO

(Biontex). For these experiments, cells were trypsinised (Section 3.1.1), counted using a

haemocytometer, the appropriate number of cells per well was added to each culture

plate in RPMI/PS/10%FCS74 (Table 3.1) and the plates were incubated overnight. The

following day, medium was replaced with the appropriate volume of

RPMI/PS/10%FCS74 (Table 3.1), plasmid for each well was made up to the appropriate

volume with RPMI 1640 medium71, MetafectineTM PRO reagent was made up to the

same volume with RPMI 1640 medium71 (Table 3.1), and the two solutions were

combined and incubated for 20 minutes to allow liposomes to form around the DNA.

The solution was added dropwise to each well or coverslip and cultures were incubated

at 37ºC, 5% CO2 for 48 hours. For cells growing on coverslips, medium on the

coverslips was replaced 6 hours post-transfection with fresh RPMI/PS/10%FCS74 and

the coverslips were incubated for a further 42 hours.

Chapter 3 Methods

63

Table 3.1 – Cell seeding density and reagents for transfection of mammalian cells

12 well plate

6 well plate Coverslips in 6 well plate

10 cm petri dish

Total volume 1.1mL 2.2mL 2.2mL 11.4mL RPMI/PS/10%FCS74 1mL 2mL 2mL 10mL MetafectineTM PRO 3µL 6µL 6µL 42µL

Amount of DNA transfected

2µg 4µg 4µg 10-30µg

Total transfection volume

2x 50µL 2x100µL 2x100µL 2x700μL

Number of DU145 cells seeded

1.5 x105

cells/well 4x105

cells/well 0.75x105

cells/coverslip 2x106

cells/well Number of LNCaP cells

seeded 2 x105

cells/well 5x105

cells/well 2x105

cells/coverslip 4x106

cells/well

3.1.5 Treatment of Mammalian Cells

For experiments involving the depletion or addition of androgens to cultures, LNCaP

cells were trypsinised (Section 3.1.1), aliquoted into the required culture dishes (Table

3.1) in RPMI/PS/10%FCS74 and incubated overnight at 37ºC and 5% CO2. The

following morning the medium was replaced with RPMI/PS/5% charcoal treated FCS

(CSS)73. For experiments examining the effect of androgens on NKX3.1 protein levels,

the cells were cultured in RPMI/PS/5%CSS73 for 24 hours (androgen depletion) before

the addition of 10-8M Dihydrotestosterone (DHT)18. Cells were cultured for 8-48 hours

at 37ºC and 5% CO2 prior to cell lysis. For experiments examining the effects of

androgen depletion on NKX3.1 protein levels, the cells were cultured in

RPMI/PS/5%CSS73 at 37ºC and 5% CO2 for 8-24 hours prior to cells lysis.

For treatment with proteasome or lysosome inhibitors, cells were trypsinised (Section

3.1.1) and the appropriate numbers of cells were seeded into 12 or 6 well plates (Table

3.1) in RPMI/PS/10%FCS74 then incubated at 37°C and 5% CO2 overnight. Where cells

were treated for 24-48 hours with proteasome or lysosome inhibitors, the medium was

replaced with RPMI/PS/10%FCS74 containing the inhibitor and the cells were cultured

for up to 48 hours prior to harvest in Whole Cell Lysis Buffer112 (Table 3.2) (Section

3.15.1). Cells treated for shorter period of time were grown to ~80% confluency prior to

2-8 hours of treatment with the inhibitor and harvest in Whole Cell Lysis Buffer112

(Table 3.2) (Section 3.15.1). Cells treated with inhibitors and 10-8M DHT18 were

prepared as described above, with culture of cells in RPMI/PS/5%CSS73 for 24 hours

prior to the addition of DHT and/or inhibitors. Additionally, where cells were depleted

Chapter 3 Methods

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of androgens as described above by culture in RPMI/PS/5%CSS73, inhibitors were

added at the time the medium was changed for 8-24 hours prior to cell lysis.

Table 3.2 –LNCaP cell treatments

Treatment Final Concentration Hours of Treatment

NH4Cl3 10mM – 20mM 8 - 24

Chloroquine 25µM - 100µM 8-48

Lactacystin37 10µM 2-8

MG13250 10µM 3-8

DHT18 10-8 M 24-48

Cycloheximide13 10µg/mL or 20µg/mL 0.25-4

For treatment of cultures with cycloheximide13, LNCaP cells were grown in 6 well

plates in RPMI/PS/10%FCS74 until ~80% confluent, either 10µg/mL or 20µg/mL

Cycloheximide13 was added to the medium and the cells were cultured for a further 15-

240 min prior to harvesting in Whole Cell Lysis Buffer112.

3.2 RNA Extraction and DNase Treatment

3.2.1 RNA Extraction

For RNA extraction, cells cultured in 75cm2 flasks were trypsinised (Section 3.1.1),

passaged as required and the remaining cell suspensions were transferred into sterile

10mL tubes, centrifuged at 5000rpm for 5 minutes at room temperature, the

supernatants removed and the cell pellets stored at -80ºC until use. To extract RNA, cell

pellets were lysed in 1mL of Ultraspec® RNA and pipetted to disperse the solution.

Lysates were transferred to fresh 1.5mL microcentrifuge tubes, incubated on ice for 5

minutes at 4ºC, 0.2 volumes of chloroform was added to each tube and the tubes were

shaken vigorously for 15 seconds then immediately placed on ice for 5 minutes. Tubes

were centrifuged at 12000rpm for 15 minutes and the upper aqueous layer containing

the RNA was transferred to a fresh 1.5mL microcentrifuge tube, an equal volume of

isopropanol was added to each tube and the tubes were shaken to mix, then incubated

on ice for 10 minutes. Tubes were centrifuged at 12000rpm and 4ºC for 10 minutes to

precipitate the RNA, the supernatants were removed, the pellets were washed twice with

1mL 75% ethanol25 per 1mL Ultraspec® RNA followed by centrifugation at 7500rpm

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65

for 5 minutes at 4ºC. After the final centrifugation, supernatants were removed and

pellets were air dried then dissolved in 10-100µL DEPC-treated ddH2O15 and stored at

-80ºC.

3.2.2 DNase Treatment of RNA

For DNase treatment of RNA (Section 3.2.1), 10μL RNA, 20μL RQ1 RNase Free

DNase, 4μL 10x Reaction buffer and ddH2O to 40μL were added to a 0.5mL tube and

the solution was incubated at 37ºC for 30 minutes. Tubes were immediately placed on

ice, 6μL RQ1 Stop buffer was added then the tubes were heated at 95ºC to inactivate the

DNase. For ethanol precipitation of RNA, 6.6μL 3M Sodium Acetate pH4.679 (1/10th

sample volume) and 156μL 100% ethanol (2.5-3 times sample volume) were added to

each tube, the tubes were vortexed, placed on dry ice for one hour then centrifuged at

12000rpm for 30 minutes and 4ºC. Supernatants were removed, the RNA pellets washed

with 100μL 75% ethanol25 and the tubes again centrifuged at 12000rpm and 4ºC for 5

minutes. The supernatants were discarded, tubes were heated at 60ºC for ~5 minutes to

dry the pellets and the RNA was dissolved in ~10μL DEPC-treated ddH2O15. RNA

concentrations were measured using a NanoDrop® ND1000 UV/Vis spectrophotometer

and RNA solutions were stored at -80°C (Section 3.5).

3.3 Reverse Transcription To reverse transcribe RNA into cDNA, 1µg total RNA (Sections 3.2.1, 3.2.2) was added

to a 0.5mL microcentrifuge tube along with 1µL (0.5μg) Oligo(dT), 1µL 10mM dNTP23

and ddH2O to a final volume of 12µL, the tube was incubated at 65ºC for 5 minutes to

denature the RNA, then immediately placed on ice for 2-5 minutes. Following

incubation, 4µL 5X First Strand Buffer and 2µL 0.1M DTT were added to each tube

and the tubes were incubated at 42°C for 2 minutes. One µL (200U) Superscript II

Reverse Transcriptase® was added and the tubes were incubated at 42°C for 50 minutes,

then heated at 70°C for 15 minutes to degrade the reverse transcriptase enzyme. cDNA

was stored at -20ºC.

3.4 Polymerase Chain Reaction (PCR)

3.4.1 PCR

PCRs were performed in BioRad C1000TM Thermal Cyclers. Each reaction contained

1µL (~50ng) cDNA (Section 3.3) or 15ng plasmid DNA (Section 3.10), PCR buffer

Chapter 3 Methods

66

(containing dNTPs), MgCl2, 15 pmol sense and antisense primers (Appendix II), DNA

Polymerase and ddH2O to 25μL, with the PCR buffer used dependent upon the type of

DNA Polymerase utilised (Table 3.3). Reactions were carried out for 30-35 cycles, with

the annealing temperature and extension times optimised for each primer pair (Table

3.4, Appendix II) and amplified PCR products were stored at -20°C or electrophoresed

in 1-2% agarose gels2 (Section 3.6).

Table 3.3 – Addition of DNA polymerases to PCRs

Platinum® Taq DNA Polymerase

Phusion®

High-Fidelity DNA Polymerase

Combination Platinum® Taq DNA Polymerase + Phusion

High-Fidelity DNA Polymerase

AmpliTaq® Gold 360 DNA Polymerase

Amount DNA Polymerase per Reaction (Units)

0.5 0.4 0.5/0.2 0.625

PCR Buffer 5x PCR Buffer54

5x Phusion HF Buffer (1.5 mM MgCl2)

5x Phusion HF Buffer (1.5 mM MgCl2)

12.5µL AmpliTaq Gold Mastermix

Table 3.4 – Primer pair PCR conditions

Primersa MgCl2 (mM)

DNA Denaturation/ Time

Annealing Temperature (°C)/Time

Extension Time (72°C)

Number of Cycles

RMND5B BamHI 2 95°C/1 min 55°C /1 min 100 sec 35 Twa1 1.5 95°C/1 min 60°C / 1 min 1 min 35 EMP 1.5 95°C/1 min 57°C /1 min 1 min 35 ARMC8 2 95°C/1 min 55°C /1 min 1 min 35 Muskelin 1.5 95°C/1 min 55°C /1 min 1 min 35 C17orf39 1.5 95°C/1 min 55°C /1 min 1 min 35 RMND5A700S/RMND5A AS

1.5 95°C/1 min 55°C /1 min 1 min 35

RMND5B789-S/RMND5BTOP0-AS

1.5 95°C/1 min 55°C /1 min 1 min 35

RMND5A RING 2 95°C/1 min 55°C /1 min 1 min 35 RMND5B RING 2 95°C/1 min 55°C /1 min 1 min 35 CBL RING 2 95°C/1 min 55°C /1 min 1 min 35 RanBPM687-S/RanBPM2190-AS

1.5 98°C/10 sec 60°C /30 sec 90 sec 35

RMND5A BamHI1-S / RMND5A1176-AS

1.5 98°C/10 sec 64°C /30sec 90 sec 35

a = Appendix II

Chapter 3 Methods

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3.4.2 “A” Tailing of PCR Products

For TA cloning of PCR amplified products where Phusion® High Fidelity Polymerase

was used, ‘A’ tailing of the PCR products was carried out using Platinum® Taq DNA

Polymerase. For each reaction, 7µL purified PCR product, 1µL 10X Taq PCR Buffer,

0.5µL 10mM dATP14, 5U Platinum® Taq DNA Polymerase and 0.75mM MgCl2 were

combined and the reaction was incubated at 70ºC for 30 minutes. ‘A’-tailed products (1-

2µL) were used in ligation reactions (Section 3.8.4).

3.4.3 Site Directed Mutagenesis

Site directed mutagenesis was performed, based on the Stratagene QuikChange®

Protocol. Overlapping forward and reverse mutagenesis primers of 25-45 bases with 3’

overhangs were designed, with estimated melting temperatures of between 60ºC and

70ºC and the mutation located in the middle of both sequences (Appendix II).

3.4.3.1 Mutagenesis PCR

Mutagenesis PCRs were performed in a BioRad C1000TM Thermal Cycler using 15ng

template plasmid DNA (Sections 3.9, 3.10), 0.5U Phusion® High Fidelity Polymerase

polymerase, 4µL 5x Phusion® HF Buffer, 200µM dNTPs23, 0.5µM forward and reverse

mutagenesis primers (Appendix II), 3% DMSO and ddH2O to 20µL (Table 3.5). To

digest methylated wild-type plasmid DNA, 0.5 µL (10U) DpnI was added to each

reaction and the tubes were incubated for 60 minutes at 37ºC, then heated at 80ºC for 20

minutes to inactivate the DpnI and the PCR products transformed into competent E. coli

DH5α cells (Section 3.8.6).

Table 3.5 – Site directed mutagenesis PCR conditions

Primers (15 pmol/μL)

RMND5A (C356S)

FWD/ RVSE

RMND5A (C356A/H358A)

FWD/RVSE pair 1 and 2

RMND5B (C358S)

FWD/ RVSE

RMND5B (C358A/H360A)

FWD/RVSE

MgCl2 (mM) 1.5mM 1.5mM – 2.5mM 1.5mM 1.5mM – 3mM DNA

denaturation time (98ºC)

30s 40s 30s 40s

Annealing Temperature (°C)

65ºC - 72ºC 64°C-70°C 58ºC - 68ºC 65°C

Annealing Time (seconds)

30s 50s 30s 50s

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Extension Time (72ºC)

3 min 4min 3 min 4min

Number of Cycles 26 26 26 26

3.5 Spectrophotometric Quantitation of RNA/DNA A NanoDrop® ND-1000 spectrophotometer was used to determine the concentration of

both RNA and DNA while a Qubit fluorometer with Quant-iTTMdsDNA BR Assay Kit

was used to determine the concentration of double-stranded (plasmid) DNA. For

measurements using the Nanodrop® ND-1000, the spectrophotometer was blanked with

the appropriate solution (DEPC-treated ddH2O15 or TE buffer) and the optical density

(OD) of the RNA/DNA solutions at 260nm and 280nm was determined. An OD of 1.0

represented 40µg/mL RNA or 50µg/mL DNA, while OD260/OD280 ratios of between 1.8

and 2.0 indicated relatively pure RNA/DNA solutions. For DNA measurements using

the Qubit fluorometer, a Quant-iTTM working solution was prepared by the addition of

1μL Quant-iTTM Reagent to 199μL Quanti-iTTM Buffer. For the standards, 190μL

Quanti-iTTM working solution was aliquoted into two tubes into which 10μL standard

solutions S1 and S2 were added, respectively. Between 1-20μL samples were added to

180-199μL Quant-iTTM working solution to obtain a final volume of 200μL in all tubes.

Tubes were vortexed for 3 seconds and allowed to stand for >2 minutes before reading

the samples using the QubitTM fluorometer. DNA concentrations in μg/mL determined

by the fluorometer were adjusted according to the dilutions used for each sample.

3.6 Agarose Gel Electrophoresis DNA was electrophoresed in 1% or 2% (w/v) agarose gels2 in 1X TAE buffer100. The

appropriate volume of 6X DNA loading dye22 was added to each sample, with 5-15μL

sample loaded into each well and a lane containing 5μL (250ng) 1Kb PlusTM DNA

ladder61 included in each gel. Gels were electrophoresed at 100V for 20-40 minutes and

visualised under UV transillumination using a Gel Doc 2000, with images analysed

using BioRad Quantity One® software.

3.7 DNA Purification

3.7.1 Purification of DNA

PCR products and plasmids were purified using a QIAquick® PCR purification kit and

according to the manufacturer’s instructions. Briefly, 5 volumes of PBI buffer was

Chapter 3 Methods

69

added to each sample, the solution was mixed then added to a QIAquick® column,

which was centrifuged at 13000rpm for 1 minute at room temperature and the flow

through discarded. PE buffer (750µL) was added to each column, columns were

centrifuged at room temperature for 1 minute at 13000rpm at room temperature, the

flow through discarded and the column centrifuged for a further 1 minute at 13000rpm

and room temperature to dry the column matrix. Columns were placed into fresh 1.5mL

microcentrifuge tubes and the DNA eluted by the addition of 50μL EB buffer to the

centre of the QIAquick® column membrane. Columns were incubated at room

temperature for 1 minute then centrifuged for 1 minute at room temperature and

13000rpm. Purified DNA was stored at -20ºC.

3.7.2 Gel Purification of DNA

To gel purify DNA using a Qiagen QIAquick® Gel Purification Kit, samples

electrophoresed in agarose gels (Section 3.6), were visualised under UV

transillumination, the appropriate bands were excised from the gel using sterile scalpel

blades and placed into preweighed 1.5mL microcentrifuge tubes. Tubes were

reweighed, the weight of the agarose calculated and 100μL Buffer QG per 100mg gel

slice was added to each tube. Tubes were incubated at 50ºC for 10 minutes or until the

agarose had melted, 1 gel volume of isopropanol was added to the tube and the solution

mixed by pipetting. The solution was then transferred to a QIAquick® spin column and

centrifuged at 13000rpm for 1 minute at room temperature. The flow through was

discarded, 750μL Buffer PE was added to each column, which was centrifuged at room

temperature for 1 minute at 13000rpm and the flow through discarded. Columns were

centrifuged for a further 1 minute at 13000rpm and room temperature to dry the column

matrix and the columns were transferred to fresh 1.5mL microcentrifuge tubes. To elute

the DNA, 20-50μL Buffer EB was added to each column, columns were incubated for 1

minute at room temperature then centrifuged at 13000rpm for 1 minute. Purified DNA

was stored at -20ºC.

3.8 Cloning of PCR Products

3.8.1 Plasmids

The pGEM®-T Easy cloning vector (Promega) was used for cloning of PCR-amplified

fragments using the A overhangs generated by Platinum® Taq DNA polymerase (Figure

3.1, Section 3.4). The pEGFP-C2 and pmCherry-C1 mammalian expression vectors

Chapter 3 Methods

70

(Clontech) were used to enable the expression of EGFP- and Cherry-tagged proteins in

mammalian cells (Figures 3.2, 3.3). The pGEX-2TK bacterial expression vector (GE

Healthcare) was used to allow the expression of GST-fusion proteins in bacterial cells

(Figure 3.4).

3.8.2 Restriction Enzyme Digestion of Plasmids

For 15μL restriction enzyme digests, purified plasmid (Section 3.7.1, 3.9, 3.10), 1μL

(5U) appropriate restriction enzyme and 1.5μL 10X buffer (appropriate for restriction

enzyme) were added to 0.5mL microcentrifuge tubes and the volume made up to 15μL

using ddH2O. Tubes were incubated for 3 hours at the optimum temperature for each

restriction enzyme in BioRad PTC-100TM Programmable Thermal Cyclers and the

digested plasmids were stored at -20°C.

3.8.3 Shrimp Alkaline Phosphatase Digestion

In order to prevent recircularisation of restriction enzyme digested plasmids (Section

3.8.2), the 5’ phosphate groups were removed using shrimp alkaline phosphatase (SAP).

For 40µL digests, digested plasmid (Section 3.8.2), 4μL 10X SAP dephosphatase buffer

and 1.5μL (1.5U) SAP were made up to 40µL with ddH2O, digests were incubated at

37ºC for 30 minutes, then at 65ºC for 15 minutes to inactivate the SAP. Plasmids were

purified using a QIAquick® Purification kit (Section 3.7.1) and stored at -20ºC or

electrophoresed in agarose gels (Section 3.6).

3.8.4 Ligation Reactions

For 10μL ligation reactions, 50ng linear vector (restriction enzyme digested, SAP

treated (Section 3.8.2, 3.8.3)), 1μL 10X T4 DNA Ligase Reaction Buffer, the

appropriate amount of insert and 1μL T4 DNA Ligase were made up to 10μL with

ddH2O. Ligation reactions were incubated at 16°C overnight in BioRad PTC-100TM

Programmable Thermal Cyclers and stored at -20°C. The amount of insert required for

1:1 molar ratios of insert:vector was calculated using the following equation:

x(ng)insert = 50ng vector x y(kb)insert z(kb)vector

Chapter 3 Methods

71

Figure 3.1: Map of the pGEM®-T Easy cloning vector showing the multiple cloning site (MCS) and the insert position (*). Ligation of inserts into the MCS results in the disruption of the lacZ gene thus enabling the selection of transformed bacterial colonies containing plasmids with insert s (white colonies) and those containing recirularised vector (blue colonies) on LB agar/Ampicillin39 plates containing X-gal (Adapted from www.promega.com/vectors).

*

Chapter 3 Methods

72

Figure 3.2: Map of the pEGFP-C2 expression vector showing the multiple cloning site (MCS) and the EcoRI restriction digest site where RMND5A or RMND5B was inserted (*). The enhanced green fluorescent protein (EGFP) coding region is located upstream of the MCS, therefore in-frame ligation of RMND5A into the plasmid results in the expression of RMND5A with an amino-terminal EGFP tag (EGFP-RMND5A) following transfection of the plasmid into mammalian cells (adapted from www.clontech.com/images/pt/dis_vectors).

*

Chapter 3 Methods

73

Figure 3.3: Map of the pmCherry C1 expression vector showing the multiple cloning site (MCS) and the EcoRI/BamHI restriction enzyme digest sites where RMND5B and RanBPM were inserted. The Cherry fluorescent tag was inserted into a pEGFP-C1 expression vector with the EGFP tag removed. The Cherry fluorescent tag is amino-terminal to the MCS, therefore in-frame ligation of RMND5B and RanBPM into the MCS resulted in the expression of amino-terminally Cherry-tagged RMND5B and RanBPM (adapted from www.staff.ncl.ac.uk/p.dean/pEGFP_C1_map.pdf)

http://www.staff.ncl.ac.uk/p.dean/pEGFP_C1_map.pdf�

Chapter 3 Methods

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Figure 3.4: Map of the pGEX-2TK bacterial expression vector showing the multiple cloning site (MCS) and the BamHI/EcoRI restriction enzyme insert position. The GST tag is upstream from the MCS thus allowing the expression of amino-terminal GST tagged fusion proteins in bacterial systems (adapted from http://www.gelifesciences.com/aptrix/upp00919.nsf/Content).

http://www.gelifesciences.com/aptrix/upp00919.nsf/Content�

Chapter 3 Methods

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3.8.5 Preparation of Competent Bacterial Cells

3.8.5.1 Preparation of Competent Escherichia coli DH5α

To prepare competent bacterial cells, an aliquot of E. coli DH5α glycerol stocks was

streaked onto a Luria-Bertani (LB) Agar38 plate and incubated inverted at 37ºC. The

following day, one colony from the overnight plate was inoculated into 10mL LB

Broth43 and incubated overnight at 37ºC with shaking at 225rpm. The next morning,

1mL of the culture was inoculated into 200mL LB Broth43 and the culture incubated at

37ºC with shaking at 225rpm for ~3 hours until the OD600 was between 0.4-0.5. The

culture was divided into four 50mL centrifuge tubes and centrifuged at 4000rpm for 10

minutes at 4ºC to sediment the bacterial cells, the supernatant was discarded and the

bacterial pellets were resuspended in 5mL ice cold Glycerol/PIPES33 buffer until

homogeneous. The cell suspensions were pooled, incubated on ice for 30 minutes and

the bacterial cells repelleted by centrifugation at 4000rpm for 10 minutes at 4ºC. The

supernatant was discarded and the bacterial pellet was resuspended in 2mL

Glycerol/PIPES33 buffer until homogeneous, divided into 100μL aliquots in sterile

1.5mL tubes and stored at -80ºC.

3.8.5.2 Preparation of Competent Escherichia coli BL21

To prepare competent E. coli BL21 bacterial cells, BL21 cells from glycerol stocks

stored at -80ºC were streaked onto an LB Agar38 plate and incubated inverted overnight

at 37ºC. The following day, a single colony from the plate was inoculated into 5mL LB

Broth43 and incubated overnight at 37ºC and 225rpm. The next morning, 0.5mL of

overnight culture was transferred to 7.5mL fresh LB Broth43 and incubated at 37ºC,

225rpm for 1-1.5 hours until the OD600 was 0.4-0.5. At this time, the bacterial cells were

collected by centrifugation at 2500rpm for 10 minutes at 4ºC, the supernatant decanted

and the pellet resuspended in 0.5mL 50mM CaCl27. Another 2mL 50mM CaCl2

7 was

added to the tube and the cell suspension was mixed then stored on ice for 30 minutes.

The bacterial cells were again collected by centrifugation at 2500rpm for 10 minutes at

4ºC, resuspended in 0.5mL 50mM CaCl27 and 50μL aliquots were used immediately in

transformation reactions (Section 3.8.6).

3.8.6 Transformation of Bacterial Cells

Aliquots of 100 µL competent E. coli DH5α cells (Section 3.8.5.1) stored at -80°C, or

50μL freshly prepared competent BL21 cells (Section 3.8.5.2) were thawed on wet ice,

Chapter 3 Methods

76

2-10µL ligation reaction (Section 3.8.4) was added to each tube, the solution was stirred

to mix then incubated on ice for 30 minutes, heated at 42°C for 90 seconds then placed

on ice for 2 minutes. Eight-hundred µL LB broth43 was added to each tube and the tubes

were incubated at 37°C with shaking at 225rpm for 1 hour. Aliquots of 50µL, 100µL

and 150µL of the transformed cells were spread onto LB agar/ampicillin39 or LB

agar/kanamycin41 plates, or for blue/white colony selection, LB/ampicillin/IPTG/X-

gal40 or LB/kanamycin/IPTG/X-gal42 plates and the plates were incubated inverted at

37°C overnight then stored at 4°C.

3.8.7 Preparation of Bacterial/Glycerol Stocks

Five mL bacterial cultures in LB broth/ampicillin44 or LB broth/kanamycin46 were

prepared by overnight incubation at 37°C and 225rpm. The following day, the cultures

were centrifuged at 5000rpm for 8 minutes at 4°C and the supernatants discarded. A

1mL aliquot of prechilled LB broth/10% glycerol45 was added to each pellet, the cells

were evenly suspended, transferred to 2mL cryotubes and stored at -80°C.

3.9 Small Scale Plasmid Purification Bacterial cultures were inoculated into 5mL LB broth43 containing the appropriate

antibiotic and incubated overnight at 37°C and 225rpm. The following day, 1.5mL from

each culture was transferred to a 1.5mL microcentrifuge tube, which was centrifuged at

14000rpm for 30 seconds at room temperature. Supernatants were removed from the

tubes, bacterial pellets were resuspended in 100µL ice cold Solution I92 by vortexing,

then cells were lysed by the addition of 200µL freshly prepared Solution II93 to each

tube. Tubes were immediately inverted 6 times, 150µL ice cold Solution III94 was added

to each tube, and the tubes were gently vortexed three times, incubated on ice for 5

minutes and then centrifuged for 5 minutes and 14000rpm at room temperature.

Supernatants were transferred into fresh 1.5mL microcentrifuge tubes, 2 volumes of

100% ethanol (~900µL) was added, tubes were briefly vortexed, incubated for 2

minutes at room temperature then centrifuged at 12000rpm for 5 minutes at room

temperature. Supernatants were decanted, 600µL 70% ethanol25 was added to each tube,

the tubes were inverted to mix, centrifuged at room temperature for 5 minutes at

12000rpm, the supernatants drained and the tubes inverted to air dry for 20 minutes.

Plasmid pellets were resuspended in ~30µL ddH2O, 1µL RNase A (10mg/mL)70 was

Chapter 3 Methods

77

added to each tube, tubes were incubated at 37°C for 1 hour and purified plasmids were

stored at -20°C.

3.10 Large Scale Plasmid Purification Large scale plasmid preparation was carried out using an Invitrogen PureLinkTM HiPure

Plasmid Midiprep Kit. For this procedure, 5mL bacterial cultures were incubated for 6

hours at 37ºC and 225rpm in LB Broth43 containing the appropriate antibiotic, then

inoculated into 100mL LB broth43 containing the appropriate antibiotic and incubated

overnight at 37°C and 225rpm. The following day the culture was centrifuged at

4000rpm for 10 minutes at room temperature. Supernatants were removed, the pellets

were collected in 4mL Resuspension Buffer containing RNase A, the mixture was

pipetted until homogeneous and transferred to a sterile 15mL tube. Four mL Lysis

Buffer was added, the mixture was inverted to mix, incubated at room temperature for 5

minutes, then 4mL Precipitation Buffer was added and the tube was inverted to mix.

Tubes were centrifuged at 12000rpm for 20 minutes at 4°C and supernatants were

transferred to columns which had been pre-equilibrated by the addition of 10mL

Equilibration Buffer that had been allowed to flow through by gravity flow.

Supernatants were passed through each column by gravity flow, columns were washed

twice with 10mL Wash Buffer and the flow throughs discarded. A sterile 15mL tube

was placed under each column and the DNA was eluted from the column by adding

5mL Elution Buffer and allowing the buffer to pass through the column by gravity flow.

To precipitate the DNA, 3.5mL isopropanol was added to each elution tube, the solution

was inverted to mix and the tubes were centrifuged at 15000rpm for 30 minutes at 4°C.

Pellets were washed with 3mL 70% ethanol25, centrifuged at 15000rpm for 20 minutes

at 4°C, the ethanol removed and the pellets were air dried for 10 minutes then

resuspended in 100µL TE buffer. Purified plasmid DNA was stored at -20°C.

3.11 GST Fusion Protein Production and Purification

3.11.1 Small Scale Production of GST Fusion Proteins

For small scale production of GST fusion proteins, BL21 cells that had been

transformed with pGEX-2TK containing the insert of interest were spread onto LB

Agar/Ampicillin42 plates and incubated overnight at 37ºC (Section 3.8.6). The following

morning, one colony was inoculated into 5mL LB Broth/Ampicillin44 and incubated at

37ºC with shaking at 225rpm for ~2-3 hours until the OD600 was ~0.5. For cultures

Chapter 3 Methods

78

where GST-RING domain proteins were induced, 250µM ZnCl2 was added to the

medium to facilitate RING domain folding. Cultures were then divided in two and 25μL

100mM IPTG35 was added to one culture in order to induce GST fusion protein

production. The cultures were incubated for an additional 2.5 hours at 30ºC and 225rpm

and the OD600 recorded. One mL of each culture was transferred to a sterile 1.5mL tube

and the cells were centrifuged at 14000rpm for 1 minute at 4°C, the supernatants

discarded and the cell pellets resuspended in the required volume of Whole Cell Lysis

Buffer112 according to the formula:

Lysates were drawn through a 23G needle 20-30 times to decrease their viscocity and

stored at -20°C.

For small scale GST Fusion protein production where both the soluble and insoluble

protein fractions were to be collected, LB Agar/Ampicillin42 plates were streaked with

frozen glycerol stocks of BL21 transformed with the pGEX-2TK expression vector

containing the insert of interest (Section 3.8.6, 3.8.7). The plates were incubated

overnight at 37ºC and the following day one colony was inoculated into 5mL LB

Broth/Ampicillin44 and incubated overnight at 37ºC and 225rpm. The next morning,

0.5mL culture was transferred into a fresh 50mL tube containing 10mL LB

Broth/Ampicillin44 and incubated at 37ºC and 225rpm for ~1.5 hours until the OD600

was ~0.5. A 1mL aliquot was taken (uninduced control sample) and 95µL 100mM

IPTG35 was added to the remaining ~9.5mL culture, which was incubated for 3 hours at

30ºC and 225rpm to induce GST fusion protein production. The uninduced cells were

centrifuged at 14000rpm for 1 minute, the pellet resuspended in 200μL Whole Cell

Lysis Buffer112 and the lysate drawn through a 23G needle to decrease the viscosity.

GST fusion proteins were prepared by centrifuging the IPTG-treated cultures at

4000rpm for 5 minutes, discarding the supernatants and resuspending the pellets in

10mL Sonication Buffer95. One hundred μL 10mg/mL Lysozyme48 was added to the

resuspended cells and the tubes were inverted to mix, then incubated on ice for 30

minutes. One mL 10% Triton X-100111 was then added to each tube, the tubes were

inverted to mix and the solution sonicated on ice at 3 x 30sec (Setting 7, 25 watt output)

to lyse the bacterial cells. The solution was centrifuged at 14000rpm for 15 minutes and

the supernatant collected as the soluble fraction whilst the sedimented material

(insoluble fraction) was resuspended in 200μL Whole Cell Lysis Buffer112, as described

Volume (µL) = 1000 x OD600 8

Chapter 3 Methods

79

above. Ten μL of each sample was electrophoresed in 12% separating SDS

polyacrylamide gels78,96 (Section 3.15.2), which were stained with Coomassie blue11

(Section 3.15.5) and destained with Coomassie blue destaining solution10 to visualise

the proteins.

3.11.2 Large Scale GST Fusion Protein Production

To produce larger quantities of GST fusion proteins, BL21 bacterial cells transformed

with pGEX-2TK expression vectors containing the inserts of interest were inoculated

into 5mL LB Broth/Ampicillin44 and incubated overnight at 37ºC and 225rpm. The

following day, the 5mL culture was inoculated into 100mL LB Broth/Ampicillin44 and

incubated at 37ºC and 225rpm for ~2 hours until the OD600 was 0.6. For cultures where

GST-RING domain proteins were induced, 250µM ZnCl2 was added to the medium to

facilitate RING domain folding. GST fusion protein production was induced by the

addition of 1mL 100mM IPTG35 and the culture was incubated at 30ºC and 225rpm for

3 hours. Following incubation, the cultures were centrifuged at 3000rpm for 15 minutes

at 4ºC, the supernatants discarded and the cell pellets resuspended in 20mL Sonication

Buffer95. 200μL 10mg/mL Lysozyme48 was added to each suspension, the tubes were

inverted to mix then incubated on ice for 15 minutes prior to the addition of 144μL

200mM PMSF60, 50μL 0.5M EDTA28, 20μL 1M DTT20, 200μL 1M MgCl249 and 2mL

10% Triton X-100111 to each tube. Tubes were mixed by inversion and the cell

suspensions were lysed by sonication at 3 x 30 seconds (setting 7, 25 watt output).

Lysates were centrifuged at 27000rpm for 15 minutes at 4ºC and to isolate GST fusion

proteins, the supernatants were transferred to fresh 50mL tubes and incubated with

300μL 50% Glutathione agarose beads30 for 15 minutes at 4ºC with rotation. The beads

were collected by centrifugation at 3000rpm/4ºC for 2 minutes, the supernatants

removed and the beads resuspended in 10mL NETN Buffer53 then incubated for 10

minutes with rotation at 4ºC. The beads were again collected by centrifugation for 2

minutes at 3000rpm and 4ºC and washed in NETN Buffer53 as described above. To

elute the GST fusion proteins, the beads were pelleted by centrifugation at 3000rpm and

4ºC for 2 minutes, the supernatants removed and the beads resuspended in 300μL GST

Elution Buffer32 and incubated for 10 minutes at 4ºC with rotation. Beads were

collected by centrifugation at 3000rpm for 2 minutes at 4ºC and the supernatants

transferred to sterile 0.5mL tubes and stored at -80ºC. To confirm expression of the

GST fusion protein, an aliquot was electrophoresed in a 12% SDS polyacrylamide

gel78, 96 (Section 3.15.2) along with BSA standards (1μg - 10μg) and the gel stained with

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Coomassie blue11 and destained with Coomassie blue destaining solution10 (Section

3.15.5).

3.11.3 Total Protein Extraction from E. coli BL21 Cells

To obtain total protein from E. coli BL21 cells, BL21 cells were inoculated from

glycerol stocks into 5mL LB Broth43 and incubated overnight at 225rpm. Two mL

overnight culture was inoculated into 10mL LB Broth43 and the culture was incubated

for 4 hours at 37°C/225rpm. Bacterial cells were harvested by centrifugation, the

supernatant discarded and the cells were resuspended in 5mL Sonication Buffer95 with

the addition of 5µL 10mg/mL Lysozyme48. Cells were lysed by sonication at 3 x 30

seconds on ice (setting 7, 25 watt output), the debris sedimented by centrifugation for 5

minutes at 5000rpm and the supernatant collected and stored at -80°C in 0.5mL

aliquots.

3.12 DNA Sequencing Sequencing reactions were carried out using the dideoxy chain termination method

with Big DyeTM Terminator and Big DyeTM Terminator sequencing buffer (Sanger et

al., 1977). Sequencing reactions were prepared in 0.5mL microcentrifuge tubes and

contained 8μL 2.5x Big DyeTM Terminator sequencing buffer, 3pmol primer (Appendix

II), 250-500ng plasmid DNA, 0.5μL Big DyeTM terminator (v3.1) and ddH2O to a final

volume of 20µL. The sequencing reactions were carried out in PTC-100TM BioRad

Programmable Thermal Cyclers with 25 cycles of DNA denaturation at 96ºC for 15

seconds, primer annealing at 50ºC for 10 seconds, and primer extension at 60ºC for 4

minutes. Reactions were precipitated by adding 2μL 3M sodium acetate81 (pH4.6) and

50μL 95% ethanol25 to each tube, the tubes were vortexed then incubated on ice for 10

minutes. Tubes were centrifuged at 12000rpm for 30 minutes at 4°C, supernatants were

removed, pellets were rinsed with 70% ethanol25, centrifuged at 12000rpm for 5

minutes at 4°C and the supernatants removed. Tubes were air dried for 10 minutes at

room temperature, stored at -20ºC and reactions were sequenced by the The Lotterywest

State Biomedical Facility at Royal Perth Hospital using an ABI Prism 3730 capillary

sequencer. Sequencing chromatograms were viewed using Chromas Lite version 2.23

and analysed using Basic Local Alignment Search Tool (BLAST)

(http://blast.ncbi.nlm.nih.gov/Blast).

http://blast.ncbi.nlm.nih.gov/Blast)�

Chapter 3 Methods

81

3.13 Immunoprecipitation Immunoprecipitation reactions were carried out using Protein A or Protein G Sepharose

beads using an in-house laboratory protocol and buffers, or alternatively

immunoprecipitations using Protein A/G or anti-GFP microbeads performed using a

µMACSTM GFP purification kit and supplied buffers. Immunoprecipitation reactions

were carried out at 4°C using cells growing in 10cm petri dishes that had been

transfected with expression plasmids and cultured for 48 hours following transfection

(Section 3.1.4). To harvest cells, culture plates were placed on ice, the medium was

aspirated and cells were rinsed 3 times with 1mL ice cold PBS56. The PBS56 was

aspirated, 1mL ice cold RIPA buffer69 or Lysis Buffer47 (anti-GFP microbeads) was

added to each plate, cells were scraped into the buffer then transferred to pre-cooled

1.5mL microcentrifuge tubes on ice. The suspensions were pipetted to disperse, tubes

were rotated for 30 minutes at 4°C to lyse the cells, centrifuged at 10000rpm for 10

minutes to collect the cell debris and the ~1mL supernatant placed in a fresh pre-cooled

1.5mL microcentrifuge tube. A 50µL aliquot of the lysate (total input) was transferred

to a 0.5mL microcentrifuge tube and stored at -20°C. Where microbeads were utilised

for immunoprecipitation reactions, 2-4μg antibody (Table 3.6) and 100μL Protein A/G

microbeads (Miltenyi Biotec) were added to the remaining ~950μL lysate and the tubes

were incubated for 30 minutes on ice. For immunoprecipitation of GFP-tagged proteins,

50μL anti-GFP microbeads (Miltenyi Biotec) was added to the ~950μL cell lysate and

the tubes incubated on ice for 30 minutes.

To prepare μ columns (Miltenyi Biotec), 200μL RIPA Buffer69 or Lysis Buffer47 was

applied to each column and allowed to drain by gravity flow. Cell lysates were added to

the columns and allowed to drain through by gravity flow, the columns were washed 5x

with 200µL RIPA Buffer69 or 4x with 200µL Wash Buffer 1 and once with 100µL

Wash buffer 2 (anti-GFP microbeads). To elute bound proteins, 20μL pre-heated 1X

SDS-PAGE gel loading buffer77 or 20μL Elution buffer (anti-GFP microbeads) was

added to each column, the columns were incubated for 5 minutes at room temperature, a

further 50μL buffer was added to the column and the eluate was collected and analysed

by SDS-PAGE and western blotting (Section 3.15).

Where Protein A64 or Protein G66 Sepharose beads were used, 300µL Sepharose bead

slurry per sample was prepared by washing the beads in 1.5mL PBS56 five times and

centrifugating the slurry at 4000rpm for 2 minutes at 4ºC between washes. The beads

Chapter 3 Methods

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were then resuspended in 150µL PBS56 to create a 50% slurry and stored in three 100µL

aliquots at 4°C. Cell lysates were prepared as above, then pre-cleared by the addition of

100µL 50% Protein A65/G67 slurry and incubation of the lysates for 1 hour at 4°C with

rotation. The beads were collected by centrifugation at 4000rpm for 2 minutes at 4ºC

and each supernatant divided equally into two fresh 1.5mL tubes, one containing 5µg

antibody (Table 3.6), and the other containing no antibody as a control (mock

immunoprecipitation). For the untransfected control, untransfected cell lysates were

prepared as above and incubated with 5μg antibody (Table 3.6). Tubes were incubated

overnight at 4°C with rotation and the next morning, 100µL 50% Protein A65 or Protein

G67 slurry was added to each tube, the tubes were incubated for 2 hours with rotation at

4°C to allow immunocomplex binding to Protein A or G and the beads collected by

centrifugation at 4000rpm for 2 minutes at 4°C. Supernatants were discarded, the beads

were washed with 1mL ice cold PBS56, then collected by centrifugation at 4000rpm and

4°C for 2 minutes and the supernatants discarded. This washing procedure was carried

out a further 4 times. After the last wash was removed, the proteins were eluted by

heating the beads in 50µL 2X SDS PAGE gel loading buffer77 at 95°C for 5 minutes.

The beads were collected by centrifugation at 4000rpm for 2 minutes at 4°C and the

supernatants were collected and analysed by SDS-PAGE and western blotting (Section

3.15).

Table 3.6 – Beads and buffers utilised for immunoprecipitation reactions

Buffer Utilised/ Microbeads Utilised

Protein A or G Microbeads

Anti-Tag Microbeads

Protein A or G Sepharose

Cell Lysis Buffer RIPA Buffer69 Lysis Buffer47 RIPA Buffer69

Column/Bead Preparation Buffer

RIPA Buffer69 Lysis Buffer47 PBS56

Wash Buffer RIPA Buffer69 4x Wash Buffer 1*

1x Wash Buffer 2* RIPA Buffer69

Column Elution Buffer 1X SDS PAGE gel loading buffer77

Elution Buffer* 2X SDS PAGE gel loading buffer77

* = µMACSTM GFP purification kit

3.14 Ubiquitin Assays

3.14.1 In Vitro Ubiquitin Assay

In vitro ubiquitin assays were carried out using a Ubiquitinylation kit (Enzo Life

Sciences). Each reaction required the addition of E1, E2, E3 enzymes, ATP, ubiquitin

Chapter 3 Methods

83

and ubiquitinylation buffer in a 0.5mL tube (Table 3.7) and the reactions were carried

out at 37ºC for 60 minutes. To stop the reactions, 50µL 2x SDS-PAGE loading buffer79

was added to each tube, samples were heated at 95ºC for 8 minutes and the reaction

products separated in 12% polyacrylamide gels78, 96 or in 4-12% gradient

polyacrylamide gels. Biotinylated-ubiquitin was detected by western blotting using a

Vectastain ABC Elite Streptavidin-HRP Kit97 (Section 3.15).

Table 3.7 – In vitro auto-ubiquitination assay components

Auto-ubiquitination Assay (μL)

Minus ATP Control (μL)

Minus E3 enzyme Control (μL)

ddH2O To 50μL To 50μL To 50μL 10X Ubiquitinylation Buffer

5 5 5

100U/mL IPP68 10 10 10 50mM DTT21 1 1 1 0.1M Mg-ATP 2.5 - 2.5 20x E1 5 5 5 10x E2 2.5 2.5 2.5 E3 To 4µM To 4µM - 20x Bt-Ubiquitin 2.5 2.5 2.5

3.14.2 In Vivo Ubiquitin Assay

In vivo ubiquitination assays were performed by transfecting LNCaP cells growing in

10cm petri dishes with the appropriate plasmid combinations (Section 3.1.4, Table

3.8). Transfected cells were incubated at 37°C with 5% CO2 for 42-45 hours at which

time proteasome inhibitors (10µM MG13250 or 10µM Lactacystin37) were added to

allow the accumulation of ubiquitinated proteins, then the cultures were incubated for a

further 3-6 hours (Section 3.1.4). Cells were lysed 48 hours post-transfection and the

protein of interest, either GFP-RMND5 proteins or NKX3.1-V5, was

immunoprecipitated (Section 3.13) using anti-GFP microbeads or Protein A microbeads

with anti-V5 antibody, and the eluate analysed by 4-12% gradient SDS-PAGE and

western blotting (Section 3.15).

Table adapted from Ubiquitinylation Product Data Sheet UW9920 (Enzo Life

Sciences)

Chapter 3 Methods

84

Table 3.8 – In vivo ubiquitination assay plasmid combinations

Immunoprecipitated Protein

Transfected Plasmids (total=30µg/10cm plate)

Protein Tag Immunoprecipitated

RMND5A pEGFP-RMND5A, pCMV-HA-Ubiquitin

GFP

RMND5B pEGFP-RMND5B, pCMV-HA-Ubiquitin

GFP

NKX3.1 pcDNA3.1-NKX3.1-V5, pCMV-HA-Ubiquitin, pEGFP-RMND5A or pEGFP-RMND5B

V5

3.15 Western Blotting

3.15.1 Preparation of Whole Cell Lysates

To prepare whole cell lysates, medium was removed from cells growing in 6 or 12 well

plates (Section 3.1.4, 3.1.5), wells were rinsed with PBS56 and 100-250µL Whole Cell

Lysis Buffer112 was added to each well. Cells were scraped into the buffer using a

spatula and the lysates were transferred to 1.5mL microcentrifuge tubes. Lysates were

drawn through 23G needles into 1mL syringes until no longer viscous and stored at -

20ºC.

3.15.2 Polyacrylamide Gel Electrophoresis

Polyacrylamide gel electrophoresis was performed using a BioRad Protean® 3 Cell or

XCell SureLockTM Mini Cell Electrophoresis system. To prepare polyacrylamide gels,

plates were assembled according to the manufacturer’s instructions, 12% SDS-PAGE

separating gels78 were prepared and pipetted between the glass plates to ~0.5cm below

the level of the wells. The solution was overlayed with ddH2O and gels were

polymerised for 45 minutes at room temperature. The ddH2O was removed and the gel

space rinsed and then filled with 4% stacking gel96 solution. Combs were inserted

between the plates and the stacking gels were polymerised for 45 minutes at room

temperature.

Protein samples were prepared in 0.5mL microcentrifuge tubes and contained 10-20µL

sample (Sections 3.13, 3.14, 3.15.1) and 1-2µL 10X SDS PAGE Loading Buffer77.

Samples were denatured at 95°C for 8 minutes then allowed to cool to room

temperature. The electrophoresis apparatus was assembled according to the

manufacturer’s instructions, filled with 1X Running Buffer76 and samples were added to

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85

the wells in a fumehood. Where NUPAGE® Bis-Tris 4-12% gradient gels were used, the

XCell SureLockTM Mini Cell electrophoresis system was assembled according to the

manufacturer’s instructions and filled with 1X MOPS buffer51. Each gel included a well

containing 5μL ColorPlus Prestained Protein Marker. Gels were electrophoresed for ~45

minutes at 200V.

3.15.3 Western Transfer

Following electrophoresis, the apparatus was disassembled and gels were transferred to

containers of ice cold Transfer Buffer105. Transfer cassettes were assembled from

“black” to “white” and contained two sponges, two pieces of Whatman filter paper, the

gel, a piece of HybondTM-C Extra nitrocellulose membrane cut to the size of the gel,

two pieces of Whatman filter paper and two sponges. Each component was wetted with

ice cold Transfer Buffer105 before being added to the cassette and air bubbles were

removed by rolling a tube over the surface as each component was added to the cassette.

Cassettes were closed and inserted into a Mini-PROTEAN III cell transfer apparatus

that contained an ice block and magnetic stirrer. Tanks were filled with Transfer

Buffer105, placed on a magnetic stirrer and proteins were transferred overnight at 30V

with gentle stirring. Following overnight transfer, the transfer apparatus was

disassembled and filters were either stored at 4ºC between Whatman filter paper and

wrapped in aluminium foil or used immediately for immunoblotting (western blotting)

(Section 3.15.4).

3.15.4 Immunoblotting (Western Blotting)

Immunoblotting was performed at room temperature with horizontal rotation unless

otherwise indicated. Prior to immunoblotting, nitrocellulose membranes (Section

3.15.3) were cut to size and incubated for 90 minutes in TBS/3% Blotto103 blocking

solution. Filters were then sequentially incubated with primary antibody diluted in

TBST/1% Blotto103 for 90 minutes (Table 3.9), TBST102 for 3 x 10 minute washes,

secondary antibody diluted in TBST/1% Blotto103 for 90 minutes and washed 3 x 10

minutes with TBST102 (Table 3.9). For membranes blotted for biotinylated-ubiquitin,

the membrane was blocked for 1 hour in TBST/1% BSA104, washed 3x 10 minutes with

TBST102, incubated for 1 hour with streptavidin-HRP99 solution diluted in

TBST/1%BSA104 followed by 6 x 10 minute washes with TBST102. Filters were drained

and incubated with Enhanced Chemiluminescence (ECLTM) Western Blotting Detection

Reagent24 for 1 minute, the filters again drained, wrapped in plastic wrap and exposed to

Chapter 3 Methods

86

X-ray film for 10 seconds – 30 minutes as required. Exposed filters were developed in

an AGFA developer, scanned using an HP Photosmart 2710 scanner and analysed using

BioRad Quantity One® software. Using densitometry of the protein bands (BioRad

Quantity One®) proteins of interest were normalised to the housekeeping gene β-actin.

Table 3.9 – Primary and secondary antibodies and their respective dilutions

Primary Antibody Secondary Antibody Antibody Dilution Antibody Dilution

Goat anti-actin IgG 1:2000 Donkey anti-goat HRP conjugate

1:2000

Goat anti-NKX3.1 IgG 1:1000 Donkey anti-goat HRP conjugate

1:2000

Mouse anti-V5 IgG 1:2000 Sheep anti-mouse HRP conjugate

1:2000

Mouse anti-GFP IgG 1:2000 Sheep anti-mouse HRP conjugate

1:2000

Mouse anti-androgen receptor IgG

1:2000 Sheep anti-mouse HRP conjugate

1:2000

Rat anti-RFP IgG* 1:2000 Anti-rat HRP Conjugate

1:5000

Mouse anti-RFP IgG* 1:2000 Sheep anti-mouse HRP conjugate

1:2000

Mouse anti-HA (HRP labelled) 1:1000 - - Streptavidin-HRP97 33.3:1000 - -

* Red fluorescent antibodies show immunogenicity towards Cherry fluorescent tag which is a DsRed derivative

3.15.5 Coomassie Blue Staining

Following polyacrylamide gel electrophoresis (Section 3.15.2), gels were incubated in

Coomassie Blue staining solution11 for 2-16 hours with gentle rotation then destained

using Coomassie Blue Destaining10 solution for 40-60 minutes. Stained gels were dried

at room temperature between cellophane wrap and stored at room temperature. For mass

spectrometric analysis, 12% acrylamide gels were stained with a colloidal Coomassie

Blue solution12 and destained using 1% Acetic Acid1, both for 16-24 hours (Section

3.17).

3.16 Microscopic Imaging of Cells

3.16.1 Preparation of Slides for Fluorescence Microscopy

To prepare cells growing on coverslips in 6 well plates, 0.5μL Hœchst 33258 dye

(10mg/mL)34 was added to each well at 46 hours after transfection (Section 3.1.3, 3.1.4)

Chapter 3 Methods

87

and the cells were incubated for 2 hours at 37º/5% CO2 to allow uptake of the dye.

Medium was removed from the wells and cells were washed 3 x 5 minutes with PBS56,

the PBS56 was aspirated and cells were fixed in 4% formaldehyde29 for 15 minutes at

room temperature. The cells were again washed 3 x 5 minutes with PBS56, the PBS56

was aspirated and the cells were permeabilised by the addition of 200µL 0.1% Triton-X

100111 for 5 minutes at room temperature. Cells were washed for 3 x 5 minutes with

PBS56, the PBS56 was removed then 50µL TRITC-phalloidin dye55 was added to each

coverslip and the coverslips incubated at room temperature for 40 minutes. Phalloidin is

a compound extracted from Amanita phalloides that binds filamentous actin allowing

cytoplasmic staining of cells. Coverslips were washed for 3 x 5 minutes with PBS56 then

mounted onto glass microscope slides using 10µL mounting medium52 and stored in the

dark at 4ºC.

3.16.2 Preparation of Slides for Immunofluorescence Microscopy

For immunofluorescence microscopy, coverslips were initially processed as described

in Section 3.16.1, and following permeabilisation with 0.1% Triton-X100110, coverslips

were washed for 3 x 5 minutes with PBS56, the PBS56 aspirated, 400µL Confocal

Blocking Buffer9 was added to each coverslip and the coverslips incubated for 30

minutes at room temperature. The coverslips were washed 3 x 5 minutes with PBS56 and

incubated with primary antibody diluted 1:2000 in PBS/1% BSA57 overnight at 4ºC,

protected from light. The following morning the coverslips were washed 3 x 5 minutes

with PBS56, 100µL AlexaFluor®546 secondary antibody diluted 1:400 in PBS/1%

BSA57 was added to the coverslips and the coverslips were incubated at room

temperature for 1 hour. Coverslips were washed 5 x 5 minutes with PBS56, the PBS56

aspirated, 50µL phalloidin dye55 was added to each coverslip and the coverslips were

incubated at room temperature for 40 minutes. Coverslips were washed 5 x 5 minutes

with PBS56, mounted onto microscope slides using 10µL mounting medium52 then

stored at 4ºC protected from light.

3.16.3 Fluorescence Microscopy

An Olympus IX71 Inverted Microscope or a Nikon Ti-E Inverted Fluorescence

Microscope were used to visualise the prepared slides (Section 3.16.1, 3.16.2). The

Olympus IX71 microscope uses 6V30W Halogen illumination, whilst the Nikon Ti-E

microscopy uses TI-DS Diascopic illumination pillar 30W, and both microscopes use a

Chapter 3 Methods

88

range of filters to excite fluorophores using specific excitation wavelengths (Table

3.10). UV excitation wavelengths (10-400nm) were used to visualise blue fluorophores

such as Hœchst 3325834, blue excitation wavelengths (400-500nm) were used to

visualise green fluorophores such as EGFP, and green excitation wavelengths (500-

580nm) were used for the visualisation of red fluorophores such as

Tetramethylisothiocyanate (TRITC) phalloidin55 (Table 3.10). Photographs were taken

of the images using Image-Pro® Plus or Nikon Elements software and the images were

overlayed using Adobe Photoshop (Adobe Systems, Inc., San Jose, Calif.) and Confocal

Assistant 4.02 (Todd Clark Brelje) software then saved in .TIFF format.

Table 3.10 – Excitation and emission wavelengths of fluorescent labels

Excitation Wavelength (nm)

Emission Wavelength (nm)

Enhanced Green Fluorescent Protein

488nm 507nm

Hœchst 3325834 360nm-365nm 465nm Phalloidin (TRITC)55 540nm-545nm 570nm-573nm Alexa Fluor 546 (goat anti-mouse antibody)

546nm 570nm

Cherry 587nm 610nm

3.17 Mass Spectrometry To prepare proteins for mass spectrometric analysis, LNCaP cells growing in 4 x 10cm

dishes were transfected with plasmids encoding either GFP-RMND5A or GFP-

RMND5B (Section 3.1.4), the cells lysed and GFP immunoprecipitation of the lysates

performed using anti-GFP microbeads (Section 3.13). Mock immunoprecipitation

reactions using lysates from 3 x 10 cm2 dishes of untransfected cells and anti-GFP

microbeads was also performed as a control (Section 3.13). The immunoprecipitation

products were separated in 12% SDS-PAGE polyacrylamide gels 78, 96 (Section 3.15.2),

with a small aliquot of each sample analysed by GFP western blotting to confirm

efficient immunoprecipitation (Section 3.15.3, 3.15.4). The electrophoresed samples

were stained with Coomassie blue 12 (Section 3.15.5) and the protein bands of interest

were excised, with each added to a 1.5mL tube. 50µL acetonitrile was added to each

tube and the bands were dried by vacuum centrifugation for 1 hour at room temperature

then processed and analysed by the Australian Proteome Analysis Facility, Sydney.

Chapter 3 Methods

89

At APAF, bands were cut into small pieces, destained, dried and trypsin digested in

ammonium bicarbonate (pH8.0) overnight. The peptides were made up to 40µL in ESI

buffer and preconcentrated by injecting the sample onto a Michrome peptide Captrap

then desalted with 0.1% formic acid and 2% acetonitrile at a flow rate of 8µL/min.

Peptides were separated using an Exigent TEMPO nanoflow liquid chromatography

system using an SGE ProteCol C18, 300A, 3µm, 150µm x 10cm analytical column and

eluted from the column using a linear solvent gradient (steps from H2O:CH3CN (100:0,

+0.1% formic acid) to H2O:Ch3CN (10:90, +0.1% formic acid) over 80 minutes at a

flow rate of 500nL/min. The liquid chromatography products were analysed using a Q

Star Elite Mass Spectrometer (AB Sciex), and positive ion nanoflow electrospray, with

the mass spectrometer operated in an information dependent acquisition mode (IDA)

using a TOF mass analyser. The TOF MS survey scan was acquired with the three

largest multiply charged ions (counts >25) in each survey scan subjected to MS/MS

analysis and MS/MS spectra were accumulated for 2s (m/z 100-1600). The resulting

data were submitted to the search program Mascot (Matrix Science Ltd, London, UK)

and the peaklists were searched using the SwissProt database against Homo sapiens.

High scores were confirmed or qualified by operator inspection as a match. A decoy

database was also searched, providing a false positive identification percentage.

Chapter 4 Characterisation of RMND5 E3 Ubiquitin Ligase Activity

Chapter 4: Characterisation of RMND5 E3 Ubiquitin Ligase Activity


90

4.1 Introduction

Human RMND5 proteins are named after their yeast orthologue, RMD5, and although

the cellular roles of the human orthologues are not well characterised, the functions of

RMD5 are being progressively determined in S.cerevisiae and as such may shed light

on the functions of human RMND5A and RMND5B.

4.1.1 Yeast RMD5/Gid2

Yeast cells growing on non-fermentable carbon sources synthesise glucose via

gluconeogenesis, a pathway that involves the enzyme fructose-1,6-bisphosphatase

(FBPase), which under these conditions has a life-life of approximately 20 hours

(Funayama et al., 1980). Upon exposure to glucose, FBPase gene expression is

repressed and the existing protein undergoes rapid degradation with its half-life reduced

to ~20 minutes in a process known as catabolite inactivation (Gancedo, 1971; Schork et

al., 1995). The route by which FBPase is degraded is dependent on the amount of time

the cells have been starved of glucose prior to its addition to the medium. If cells have

been glucose starved for a short amount of time (<24 hours), FBPase is

polyubiquitinated and degraded by the ubiquitin-proteasome system, while for cells

growing on non-fermentable medium such as acetate for a longer time period (>24

hours), FBPase undergoes vacuolar import and degradation (Schork et al., 1994; Chiang

and Chiang, 1998; Hammerle et al., 1998).

Proteins involved in the degradation of FBPase by the ubiquitin-proteasome system are

known as glucose induced degradation of FBPase (Gid) proteins. Gid1-3, the first of the

Gid proteins identified to be involved in the degradation of FBPase was characterised

following the isolation of three mutants defective in glucose induced degradation

(Hammerle et al., 1998). These mutants were also defective in their ability to degrade

N-end rule proteins which are short lived proteins degraded by the ubiquitin-proteasome

system and recognised due to the presence of defined amino-terminal amino acid

residues (Varshavsky, 1997; Hammerle et al., 1998). Gid2 was also identified in a

screen of a single-gene deletion mutant databank for Saccaromyces cerevisiae genes

required for sporulation and meiosis. Enyenihi and Saunders identified 12 genes that

were not essential for meiotic entry but that were required for meiotic nuclear division

(RMD 1-12), including RMD5/Gid2, although its role in this process has not yet been

elucidated (Enyenihi and Saunders, 2003).


91

Gid2 is not present as a monomeric protein in the cell but is associated with a large,

600kDa protein complex, the remaining members of which, Gid4-9, were identified in a

mutational screen (Regelmann et al., 2003). The components of the complex have also

been identified in yeast systematic interaction studies (Ho et al., 2002; Krogan et al.,

2006; Pitre et al., 2006). The complex, named the Vid30/Gid complex, consists of

Gid1/Vid30, RMD5/Gid2, Gid4/Vid24, Gid5/Vid28, Gid7, Gid8 and Gid9/Fyv10, and

includes proteins which function in both of the FBPase degradation pathways

(Regelmann et al., 2003). The Gid proteins that degrade FBPase by the proteasome and

vacuolar import and degradation (Vid) proteins, discussed below, are responsible for

FBPase breakdown by the vacuole (Regelmann et al., 2003). The Vid30 complex is an

E3 ubiquitin ligase complex, with RMD5/Gid2 contributing its enzymatic activity to the

complex, a function mediated by its carboxy-terminal RING domain (Figure 4.1) (Santt

et al., 2008). In RMD5/Gid2 mutant cells, and in in vitro and in vivo ubiquitination

assays utilising an RMD5 RING domain mutant, FBPase is no longer ubiquitinated,

further supporting the role of this protein and the Vid30 complex in targeting FBPase

for degradation by the ubiquitin-proteasome system (Regelmann et al., 2003). Recently,

a second protein within the complex, Gid9/Fyv10, which contains a degenerate RING

domain, has been shown to associate with RMD5, contributing to the E3 ubiquitin

ligase activity of the complex (Braun et al., 2011).

Vid30, another member of the complex appears to be the substrate recognition

component as it was shown to interact with FBPase in immunoprecipitation reactions

and, along with Vid28, Vid30 is proposed to function as a core component of the Vid30

complex (Pitre et al., 2006; Santt et al., 2008). An additional associated member,

Gid4/Vid24 is not present in cells growing on ethanol but its expression is rapidly

induced upon growth of cells on glucose, however as Gid4/Vid24 mRNA levels are

similar under both conditions, Gid4/Vid24 regulation appears to occur at the

translational level (Pitre et al., 2006; Santt et al., 2008). Gid4/Vid24 is hypothesised to

activate the Vid30 complex by altering its conformation following glucose shift,

resulting in the ubiquitination and degradation of FBPase, which is dependent on RMD5

(Santt et al., 2008). Following FBPase degradation, Gid4/Vid24 is itself ubiquitinated

by RMD5 resulting in its proteasomal degradation in a regulatory feedback mechanism

(Figure 4.1) (Braun et al. 2011). The ubiquitin conjugating enzyme, Gid3/Ubc8 is

associated with the Vid30 complex and is involved in the ubiquitination of FBPase and

Gid4/Vid24.


92

Another protein associated with the complex, Gid6/Ubp14 is a deubiquitinating enzyme

that cleaves ubiquitin chains on FBPase, thereby reducing the competition for the

proteasome between polyubiquitin chains attached to the substrate and the substrate

itself (Amerik et al., 1997; Regelmann et al., 2003). Supporting this, cells lacking

Gid6/Ubp14 exhibit a three-fold reduction in FBPase degradation (Regelmann et al.,

2003). In addition to its regulation of FBPase, the Vid30 complex, including

RMD5/Gid2 and Gid9/Fyv10, are implicated in the regulation of other enzymes

involved in the irreversible steps of glucose synthesis, including the gluconeogenic

enzyme phosphoenolpyruvate decarboxykinase (PEPCK) and the TCA cycle enzyme

cytoplasmic malate dehydrogenase (c-MDH) (Figure 4.1) (Santt et al., 2008; Braun et

al., 2011). Interestingly, all three enzymes contain amino terminal proline residues, and

RMD5/Gid2 mutants are associated with defective degradation of N end rule pathway

proteins, while mutation of the amino terminal proline residue of FBPase renders it

incapable of degradation by the proteasome (Hammerle et al., 1998). These findings

suggest that the Vid30 complex plays a broader role in the regulation of responses of the

yeast cells to changing nutrient conditions by recognising substrates with amino

terminal proline residues.

Three members of the Vid30 complex, Gid1/Vid30, Gid5/Vid28 and RMD5/Gid2 have

also been implicated in the degradation of the high affinity hexose transporter 7 (Hxt7)

in response to rapamycin or nitrogen-starvation (Snowdon et al., 2008). Gid1/Vid30 and

Figure 4.1: Proposed mechanism of action of the Vid30 complex. Upon exposure to glucose, the Vid30 complex is activated by Gid4/Vid24, which enables RMD5/Gid2 and Gid9/Fyv10 to ubiquitinate FBPase, PEPCK and c-MDH, thereby targeting them for degradation by the proteasome (Braun et al., 2011).


93

Gid5/Vid28 play overlapping roles in Hxt7 internalisation and degradation as only

Vid30/Vid28 double mutants display substantial Hxt7 stabilisation (Snowdon et al.,

2008). Although RMD5/Gid2 mutant cells also exhibited delayed internalisation and

degradation of Hxt7, the transporter was eventually internalised and degraded in all

conditions implicating the involvement of other proteins in this process (Snowdon et al.,

2008). Vid30 has been shown to play a role in nitrogen catabolite repression by shifting

the cells towards glutamate production, particularly under low ammonia conditions (van

der Merwe et al., 2001). Therefore, Gid1/Vid30, Gid5/Vid28 and RMD5/Gid2 and the

Vid30 complex were hypothesised to function closely with the target of rapamycin

(TOR) pathway (Snowdon et al., 2008). A human orthologue of the Vid30 complex has

been reported and this complex, named the CTLH complex will be discussed in detail in

Chapter 6 (Kobayashi et al., 2007).

As described above, FBPase can also be degraded by the vacuole. FBPase is recognised

by Vid proteins and packaged into Vid vesicles, which fuse with the vacuole, resulting

in its degradation (Hoffman and Chiang, 1996; Chiang and Chiang, 1998). One protein

involved in this process is Gid4/Vid24, which is synthesised in the presence of glucose,

is associated with Vid vesicles and is required for the transport of FBPase in Vid

vesicles to the vacuole. Consistent with this function, Gid4/Vid24 mutants show an

accumulation of FBPase in Vid vesicles which do not move to the vacuole (Chiang and

Chiang, 1998). Coatomer proteins such as Sec28 which play roles in the endocytic

trafficking of proteins in yeast and mammalian cells, form part of Vid vesicles and

associate with Gid4/Vid24, leading to the hypothesis that Vid vesicles merge with the

endocytic pathway in the transport of FBPase to the vacuole for degradation (Brown et

al., 2008). Recently it has been shown that the Vid pathway does indeed merge with the

endocytic pathway at actin patches at the plasma membrane where early endocytosis

takes place, thereby utilising the endocytic pathway to deliver Vid vesicles and their

cargo including FBPase and MDH2 to the vacuole for degradation (Brown et al., 2010).

When glucose is added to starved cells, FBPase, MDH2 and the Vid vesicle associated

proteins Gid4/Vid24 and Sec28 are present at actin patches where Gid1/Vid30 attached

to Vid vesicles associates with the actin patches, thereby merging the two pathways

(Brown et al., 2010; Alibhoy et al., 2012).


94

4.1.2 Human RMND5 Proteins

Human RMND5A (Gene ID 64795) and RMND5B (Gene ID 64777) are the

uncharacterised human orthologues of yeast RMD5. The RMND5 proteins share 70%

amino acid homology and are highly conserved between mammalian species, indicating

that they perform both similar and important cellular roles.

4.1.2.1 RMND5A

RMND5A (p44CTLH/FLJ13910), which is located at the chromosomal locus 2p11.2,

spans approximately 54.7 kilobase pairs (kbp) and contains an open reading frame of 9

exons, producing an mRNA of 6201bp. Translation results in a 391 amino acid protein

that contains four protein-protein interaction domains, a Lissencephaly 1 homology

motif (LisH), a C-terminal to LisH motif (CTLH), a CT11-RanBPM (CRA) motif and a

Really Interesting New Gene (RING) domain. The RMND5A gene is conserved in dog,

cow, chimpanzee, mouse, rat, fruit fly, mosquito, Arabidopsis plants, nematode, mould

and rice (Homologene 5668).

4.1.2.2 RMND5B

RMND5B (FLJ22318), which has provisional protein coding status, is located at 5q35.3

and spans ~17.4kbp with an open reading frame of 11 exons. The transcribed mRNA is

1825bp and encodes a protein of 393 amino acid residues which, like RMND5A,

consists of four protein-protein interaction domains, a LisH, CTLH, CRA and RING

domains. RMND5B orthologues are found in dog, cow, chimpanzee, mouse, rat,

zebrafish and Arabidopsis plants (Homologene 100777).

4.1.3 Protein Domains

4.1.3.1 Lissencephaly 1 Homology Motif (LisH)

There are 2382 LisH domains present in 2364 proteins in the SMART non-redundant

database, with 1.09% (26) of these found in human proteins including RanBPM,

RanBP10 and OFD (SMART SM00667) (Schultz et al., 1998). In all 26 of the human

proteins, the LisH domain is found at the N-terminal except for one protein, DDB1 and

CUL4 Associated Factor 1 (DCAF1) in which the domain is found midway through the

protein (Table 4.1). Six of these 26 LisH-containing proteins also contain a CTLH and a

CRA domain, with a further two containing a LisH and CTLH domain. In addition to a

LisH domain seven proteins contain WD40 repeats. A further seven contain no other


95

identified protein domains, but may contain coiled coil repeats (Table 4.1) (Schultz et

al., 1998).

Table 4.1 – Human LisH domain containing proteins

Human LisH Domain Containing Proteins

Full Name Other protein Domains or Regions

Cellular Role

OFD1 Oral-facial-digital syndrome type 1

Coiled coil regions

Neural cell migration

NOLC1 Nucleolar and coiled-body phosphoprotein

SRP40 Transcription

SSBP2 Single-stranded DNA-binding protein 2

SSDP Regulator of haematopoietic growth/differentiation


SSDP Regulation of development


SSDP Unknown

DCAF1 DDB1 and Cul4 associated factor 1

Coiled coil region

SCF E3 ubiquitin ligase copmplex member

RMND5A Required for Meiotic Nuclear Division 5A

CTLH, CRA, RING

Unknown

RMND5B Required for Meiotic Nuclear Division 5B

CTLH, CRA, RING

Unknown

FR1OP FGFR1 oncogene partner - Erythroid proliferation/differentiation

MKLN1 Muskelin 1 CTLH, Discoidin-like domain, Kelch Repeat

Mediates cell spreading by interacting with TSP-1

TBL1X Transducin β Like Protein 1 X (F-Box like protein)

WD40, F Box Transcription

TBL1Y Transducin β Like Protein 1 Y

WD40 Transcription

ARMC9 Armadillo repeat containing protein 9 isoform 2

Coiled coil region

Downregulates α-catenin

LIS1 Lissencephaly 1/ Platelet-activating factor acetylhydrolase IB subunit alpha

WD40, coiled coil region

Neural cell migration

CP063/RHOT2 C16orf63/Ras homologue family member T2

- Rho GTPase

NPAT Nuclear protein of the ataxia telangiectasia mutated locus

- Transcription/histone gene expression

TWA1/CT011 C20orf11/Twa1 CTLH, CRA Unknown MAEA Macrophage erythroblast

attacher/Erythroblast macrophage protein

CTLH, CRA, putative RING

Erythroblast maturation


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A8MX09 Putative uncharacterised protein WDR47

CTLH, WD40, coiled coil

Unknown

RanBP9/RanBPM Ran binding protein 9 SPRY, CTLH, CRA

Adaptor protein/cell signalling

RanBP10 Ran binding protein 10 SPRY, CTLH, CRA

Guanine nucleotide exchange factor

TBL1XR1 Transducin β-like 1 X-linked receptor 1

WD40 Transcription

SMU1 Suppressor of mec-8 and unc-52 homologue

CTLH, WD40 RNA splicing

Treacle Treacle Treacle domains

Nuclear trafficking phosphoprotein

TAF5 Transcription initiation factor TFIID subunit 5

TFIID, WD40 Transcription

KIAA1468 LisH domain and HEAT repeat-containing protein KIAA1468

Coiled coil regions

Unknown

The LisH domain was originally identified in the Lissencephaly 1 (LIS1) protein to

mediate LIS1 dimerisation and oligomerisation, along with the LIS1 coiled-coil domain

(Tai et al., 2002; Kim et al., 2004). By associating with Cytoplasmic Dynein Heavy

Chain (CDHC), which plays an integral role in the microtubule based transport of

organelles and cytoskeletal components, LIS1 modulates CDHC activity during neural

cell migration, axon growth and retrograde transport and is thereby proposed to regulate

microtubule dynamics (Sasaki et al., 2000; Smith et al., 2000). Mutations in the LIS1

protein, including those in the LisH domain, lead to Miller-Dieker lissencephaly, a

disease caused by defective neural cell migration, leading to mental retardation,

epilepsy and premature death (Cardoso et al., 2000; Emes and Ponting, 2001).

Similarly, mice that are heterozygous for a truncated Lis1 gene in which residues

encoding the LisH domain and a coiled coil region (1-63) are missing, display abnormal

cortex morphology hypothesised to be due to defective neuronal migration (Cahana et

al., 2001).

The LisH domain is also present in other proteins which are involved in neural cell

migration, including Oral-facial-digital syndrome type 1 (OFD1), Transducing β like 1

(TBL1) and Treacle, abnormalities in which give rise to neurological or craniofacial

disorders, suggesting that the LisH domain plays a similar role in each of these proteins

(Emes and Ponting, 2001; Kim et al., 2004). OFD1 is an X-linked dominant disorder

caused by mutations in the CXorf5 gene. The condition is lethal in males, while affected

females exhibit facial and oral malformations, and malformation of their digits,


97

suggesting the involvement of neural cell migration (Towfighi et al., 1985; Gerlitz et

al., 2005). Treacher Collins Syndrome arises from mutations in the Treacher Collins-

Franceschetti syndrome 1 (TCOF1) gene which encodes Treacle and results in the

abnormal migration of cells of the neural crest, leading to craniofacial abnormalities and

hearing impairment (Towfighi et al., 1985; Emes and Ponting, 2001; Marszalek et al.,

2002). TBL1, an F-box containing protein that functions as a substrate recognition

component in SCF E3 ubiquitin ligase complexes, has been implicated in the

pathogenesis of ocular albinism (Dimitrova et al., 2010). Mutations of key amino acids

in the LisH domains of LIS1, OFD1 and TBL1 reduce their half-life and lead to their

abnormal cellular localisation suggesting that the LisH domain is involved in the

regulation of these aspects of protein function (Gerlitz et al., 2005). Recently, the LisH

domain of muskelin has been shown to direct the nuclear localisation of this protein and

in a muskelin LisH domain–vinculin chimera, mutation of the LisH domain resulted in

the cytoplasmic mislocalisation of the vinculin chimera (Valiyaveettil et al., 2008).

Thus, the LisH domain appears to be an important determinant of intracellular

localisation.

The LisH domains of TBL1 and LIS1 are also essential for their dimerisation and

resolution of the crystal structure of the fibroblast growth factor receptor 1 (FGFR1)

oncogene partner (FOP), including its LisH domain identified that the LisH domain is

important for FOP dimerisation and centrosomal localisation (Gerlitz et al., 2005;

Mikolajka et al., 2006). Oligomerisation of the TBL1 and TBLR1 proteins is reliant on

the LisH domain. TBL1 and its receptor TBLR1 are associated with nuclear receptor

corepressor (N-CoR) and silencing mediator for retinoid and thyroid receptors (SMRT)

in large protein complexes which are associated with HDAC3. TBL1-TBLR1 stabilise

the structure of the corepressor complexes by forming interactions with HDAC3 and

histones H2B and H4, thereby aiding in chromatin-substrate recognition (Li et al., 2000;

Zhang et al., 2002; Yoon et al., 2003). Deletion of the TBL1 and TBLR1 LisH domains

results in their failure to interact with histone H4 and the inability of the N-CoR

complex to associate with chromatin, thereby inhibiting the function of the complex to

act as a transcriptional repressor (Choi et al., 2008). Similarly, in the yeast orthologue

of TBL1, Sif2, the LisH domain mediates protein tetramerisation and Sif2 interaction

with the Set3 complex, which shows homology to the human N-CoR and SMRT

corepressor complexes (Cerna and Wilson, 2005). The LisH domain of DCAF1 is

required for the formation of Cullin4A-RING E3 Ubiquitin Ligase (CRL4)


98

supramolecular complexes (Ahn et al., 2011). Thus, in multiple proteins, the LisH

domain is required for the formation of large protein complexes, including those

involved in the ubiquitin-proteasome system such as TBL1 and DCAF1.

Although its function is incompletely characterised, based on the functions of LisH

domain containing proteins, the LisH motif is proposed to be involved in determining

protein half-life, facilitating the formation of protein complexes and regulating

microtubule function by mediating protein dimerisation or by binding microtubules or

CDHC directly (Emes and Ponting, 2001). In addition, the LisH domain has been

proposed to be involved in cell migration, nucleokinesis and chromosome segregation

(Emes and Ponting, 2001).

4.1.3.2 C-Terminal to LisH (CTLH) Domain

The CTLH domain is an ill-conserved 58 amino acid residue motif with 5

characteristically arranged leucine residues (residues 12, 21, 31, 42, 50) which form a

“U” shaped domain structure (Zeng et al., 2006). According to the SMART database,

there are 708 CTLH domains in 679 proteins, however only nine of these (1.03%) are

found in humans (SMART SM0068) (Schultz et al., 1998). Eight of these proteins also

contain an N-terminal LisH domain and many also contain C-terminal WD40 repeats

(Table 4.1). A single protein, WD repeat containing protein 26 (WDR26), contains only

a CTLH domain and WD40 repeats and is therefore similar in domain structure to other

LisH and CTLH domain containing proteins. WDR26, originally described in 2004, is a

G-beta-like protein which is involved in the regulation of MAPK signalling pathways

and mediates MEKK1 repression of the transcriptional activity of the ETS transcription

factor ELK-1 (Zhu et al., 2004). WDR26 is also implicated in mediating the

transcriptional activity of other proteins. For example, WDR26 opposes H2O2 induced

cell death by down regulating AP-1 transcriptional activity (Zhao et al., 2009; Feng et

al., 2012). These findings therefore implicate the CTLH domain, in part, in

transcriptional regulation. Interestingly, the LisH and CTLH domains of yeast Vid30 are

important for its function in merging the Vid and endocytic pathways for FBPase and

MDH2 degradation as deletion of these domains results in the impairment of FBPase

association with actin patches (Section 4.1.1) (Alibhoy et al., 2012).


99

4.1.3.3 CT11-RanBPM (CRA) Domain

The CRA domain was initially identified in the C-terminus of RanBPM, and according

to the SMART database there are six human proteins, including RanBPM, which

contain this domain, all of which also contain the LisH and CTLH domains (Table 4.1)

(Schultz et al., 1998; Menon et al., 2004). RanBPM uses its CRA domain to interact

with the C-terminus of the Fragile X Mental Retardation Protein (FMRP), a region of

FMRP that contains an RGG box with which it interacts with RNA. For this reason it

was proposed that by using its CRA domain to interact with FMRP, RanBPM is able to

modulate the RNA binding-properties of FMRP (Menon et al., 2004). Additional

functions of the CRA domain of either RanBPM or of other CRA domain containing

proteins (Table 4.1) have not been reported.

4.1.3.4 Really Interesting New Gene (RING) Domain

The RING domain, typically found in E3 ubiquitin ligases contains the characteristic

consensus sequence, CX2X(9-39)CX(1-3)HX(2-3)C/HX2CX(4-48)CX2C (Section

1.7.2). In addition, RING variants such as RBQ1 and RBX1 (Section 1.7.2.1) are able to

function as E3 ubiquitin ligases with aspartate and asparagine residues in place of

conserved cysteine or histidine residues (Deshaies and Joazeiro, 2009). Yeast RMD5 is

another example of a functional E3 ubiquitin ligase that does not possess all eight

cysteine/histidine amino acids and analysis of the RING domains of RMND5A and

RMND5B indicated that they are similar in amino acid composition to yeast RMD5

(Section 4.2.1) (Santt et al., 2008).

4.1.4 RMND5 Proteins and Cancer

Disruption of RMND5A and RMND5B has been detected in a number of cancers. Using

differential display, quantitative RT-PCR and RNA in situ hybridisation, Li et al. (2008)

identified that RMND5A is overexpressed in ovarian cancer (Li et al., 2008). The

RMND5A chromosomal locus, 2p11.2 is amplified in pilocytic astrocytomas and

atypical lobular hyperplasia of the breast, whilst this region undergoes loss in mantle

cell lymphoma cell lines (Camps et al., 2006; Mastracci et al., 2006; Belirgen et al.,

2012). The chromosomal locus of RMND5B, 5q35.3 has been found to be deleted in a

number of cancers including neuroblastoma and non-small cell lung carcinomas

(NSLC) (Mendes-da-Silva et al., 2000; Mosse et al., 2005). Additionally, amplification

of this chromosomal locus has been identified in patients with asbestos related lung


100

tumours, clear cell renal carcinoma and uterine leiomyosarcoma (Nymark et al., 2006;

Chen et al., 2009; Raish et al., 2012). The 5q35.3 chromosomal locus undergoes loss of

heterozygosity (LOH) in 82% of breast tumours that also display mutations of the breast

cancer 1 (BRCA1) gene and in 44% of BRCA2 mutated breast tumours (Johannsdottir et

al., 2006). The region is also located within an uncharacterised prostate cancer

heritability locus identified in a genome-wide linkage analysis of 1233 prostate cancer

families, however the genes with positive linkage within the 5q35 locus have not been

further investigated (Xu et al., 2005; Christensen et al., 2010). As such, although the

chromosomal regions continuing the RMND5A and RMND5B genes are disrupted in a

number of cancers, characterisation of specific genes in the two loci, 5q35.3 and 2p11.2

which are responsible for tumour initiation and/or progression has not been reported.


101

4.2 Results

4.2.1 Bioinformatics Analyses of RMND5 Protein Architecture

In order to identify potential cellular functions of RMND5 proteins, the protein domain

architecture of RMND5A and RMND5B was predicted using online protein domain

prediction tools including the National Centre for Biotechnology Information (NCBI)

(http://www.ncbi.nlm.nih.gov/gene/64795), SBASE (http://www.icgeb.trieste.it/sbase)

and Simple Modular Architecture Research Tool (SMART) (http://smart.embl-

heidelberg.de/) databases (Vlahovicek et al., 2005, Schultz et al., 1998). These

databases identified four protein-protein interaction domains present in each of

RMND5A and RMND5B. RMND5A contained a Lissencephaly 1 homology (LisH)

(amino acids 114-146), a C-Terminal to LisH (CTLH) (amino acids 153-210), a CT-11

RanBPM (CRA) (amino acids 209-302) and a Really Interesting New Gene (RING)

(amino acids 336-377) domain, while RMND5B contained a LisH (amino acids 116-

148), a CTLH (amino acids 155-212), a CRA (amino acids 211-305) and a RING

(amino acids 338-379) domain (Figure 4.2A). Additionally, SBASE database identified

loosely conserved putative protein domains in the amino-terminal regions of RMND5A

and RMND5B, with RMND5A containing a ribulose phosphate 3-epimerase like

domain (amino acids 11-40) and a GAT-like domain (amino acids 42-94), and

RMND5B containing a putative myosin tail like domain (amino acids 26-91) (not

shown) (Vlahovicek et al., 2005). The functions of LisH, CTLH and CRA domain have

not been well characterised, however RING domains are well characterised motifs

commonly found in E3 ubiquitin ligases, suggesting that RMND5 proteins are able to

function as E3 ubiquitin ligases. Alignment of the RING domains of RMND5A,

RMND5B and yeast RMD5 identified that all eight zinc coordinating residues required

for RING domain folding and function are identical between the three proteins,

supporting the hypothesis that RMND5 proteins possess this enzymatic activity (Figure

4.2B).

4.2.2 Cloning of Full Length RMND5A into pGEX-2TK and Expression of

GST-RMND5 Proteins for In Vitro Ubiquitination Assays

4.2.2.1 Cloning of RMND5A into pGEX-2TK

To determine whether RMND5 proteins possess E3 ubiquitin ligase activity, full length

RMND5A was cloned into the pGEX-2TK expression vector to allow expression of

GST-RMND5A in bacteria. The pGEX-RMND5B construct was already available in

http://www.ncbi.nlm.nih.gov/gene/64795�

http://www.icgeb.trieste.it/sbase�

http://smart.embl-heidelberg.de/�

http://smart.embl-heidelberg.de/�

Chapter 4 Characterising RMND5 E3 Ubiquitin Ligase Activity

102

Figure 4.2: Protein domain architecture of RMND5 proteins. (A) Bioinformatics analyses identified four protein-protein interaction domains in RMND5 proteins with (i) RMND5A containing a Lissencephaly 1 homology (LisH) (aa 114-146), a C-Terminal to LisH (CTLH) (aa 153-210), a CT-11 RanBPM (CRA) (aa 209-302) and finally a Really Interesting New Gene (RING) (aa 336-377) domain. (ii) RMND5B was identified to contain a LisH (aa 116-148), a CTLH (155-212aa), a CRA (aa 211-305) and a RING (aa 338-379) domain. The protein domains of RMND5A and RMND5B exhibit a high degree of amino acid homology. (B) Alignment of the RING domains of RMND5A, RMND5B and yeast RMD5 identified that all zinc co-ordinating residues required for RING domain folding and function (red) were identical between the three proteins.

A (i)

(ii)

B

Amino acid position

1

1

80%

391

Lissencephaly 1 Homology Domain C-Terminal to LisH Domain CT-11 RanBPM Domain Really Interesting New Gene Domain

114 146 153 210 209 302 336 377

393

116 148 155 212 211 305 338 379

RMND5A

RMND5B

76% 60% 68% 74% Amino acid homology 84%

CPILRQQTTDNNPPMKLVCGHIISRDALNKMFNGS--KLKCPYC

CPILRQQTSDSNPPIKLICGHVISRDALNKLINGG--KLKCPYC

CPVLKEETTTENPPYSLACHHIISKKALDRLSKNGTITFKCPYC

RMND5A

RMND5B

RMD5 361 404

336 377

338 379


103

the laboratory (Dawson, 2006). To prepare RMND5A insert for ligation, the RMND5A

coding region was amplified by PCR using pEGFP-RMND5A as a template, the

RMND5ABamHI1-S and RMND5A1176-AS primers and a range of annealing

temperatures from 64°C - 68°C for 35 cycles (Sections 3.4.1, 3.8, Appendix II). The

PCR products were electrophoresed in a 1% agarose gel, identifying a product at the

expected size of ~1.2kb in PCRs using all annealing temperatures tested (Section 3.4,

Figure 4.3A). Five PCRs were performed using the optimised annealing temperature of

64°C, the PCR products were combined, “A” tails added and a 5µL aliquot of the

purified products electrophoresed in a 1% agarose gel, verifying a single band at the

expected size of ~1.2kb (Sections 3.4.1, 3.4.2, Figure 4.3B).

To obtain pGEMT-RMND5A, 100ng (2µL) of the RMND5A insert was ligated into

50ng (1µL) pGEM®T-Easy by TA cloning and the ligation products were transformed

into competent E. coli DH5α (Sections 3.8.4, 3.8.5.1, 3.8.6). Transformed bacteria were

spread on LB Agar/Ampicillin plates containing X-gal for blue/white colony selection

and after overnight incubation, 4 white colonies were picked, inoculated into LB

Broth/Ampicillin and incubated overnight (Sections 3.8.6, 3.9). The following day,

plasmids were isolated from the bacteria, RNase treated then digested with EcoRI to

identify clones containing an RMND5A insert (Sections 3.8.2, 3.9). The products were

electrophoresed in a 1% agarose gel which identified the correctly sized ~1.2kb insert in

all pGEMT-RMND5A clones 1-8 (Section 3.6, Figure 4.3C). To confirm the correct

sequence of RMND5A inserts, pGEMT-RMND5A clones 1 to 4 were purified, 5µL of

each was electrophoresed in a 1% agarose gel to estimate plasmid DNA concentration

and 2µL of pGEMT-RMND5A clones 1 to 4 was sequenced using M13-S and M13-AS

primers (Sections 3.6, 3.7.1, 3.12, Appendix II, not shown). Analysis of chromatograms

using BLASTTM verified the correct RMND5A sequence in all pGEMT-RMND5A

clones (Section 3.12, not shown).

To prepare RMND5A insert for ligation into pGEX-2TK, the RMND5A insert from

pGEMT-RMND5A clone 1 was released by EcoRI digestion, the products were

electrophoresed in a 1% agarose gel and the ~1.2kb RMND5A insert gel purified

(Sections 3.6, 3.8.2). Five µL of the purified RMND5A insert was electrophoresed in a

1% agarose gel, with 50ng (5µL) ligated into 50ng (2µL) pGEX-2TK that had been

EcoRI digested, SAP treated and purified (Sections 3.8.2, 3.8.3, Figure 4.3D, E). The

ligation products were transformed into E. coli DH5α and selected by growth on


104

Lane 1: MW Marker Lane 2: 64°C Lane 3: 65°C Lane 4: 66°C Lane 5: 67°C Lane 6: 68°C Lane 7: Negative Control

Lane 1: MW Marker Lane 2: Purified RMND5A Lane 3: Negative Control

A

~1.2kb

~1.2kb

Lane 1: MW Marker Lane 2, 4, 6, 8, 10, 12, 14, 16: Undigested pGEMT-RMND5A clones 1, 2, 3, 4, 5, 6, 7, 8 Lane 3, 5, 7, 9, 11, 13, 15, 17: EcoRI digested pGEMT- RMND5A clones 1, 2, 3, 4, 5, 6, 7, 8

C

D

~1.2kb ~4.2kb

~5kb

B

Lane 1: MW Marker Lane 2: pGEX-2TK Lane 3: BamHI/EcoRI/SAP digested pGEX-2TK


105

Figure 4.3: Cloning of full length RMND5A into pGEX-2TK. (A) pEGFP-RMND5A (template) and an annealing temperature gradient (64°C to 68°C) were used to amplify the RMND5A coding region. The products were electrophoresed in a 1% agarose gel, identifying the ~1.2kb insert in all reactions. (B) Following optimisation, the RMND5A coding region was amplified in quadruplicate reactions, the reactions were combined and purified. 5µL product was electrophoresed in a 1% agarose gel from which the DNA concentration was estimated to be ~50ng/µL. (C) pGEMT-RMND5A clones 1 to 8 were EcoRI digested to release the ~1.2kb RMND5A insert and the products were electrophoresed in a 1% agarose gel identifying inserts in all clones. (D) 5µL BamHI/EcoRI digested and purified pGEX-2TK plasmid was electrophoresed in a 1% agarose gel, from which the concentration of DNA was estimated to be 25ng/µL. (E) pGEMT-RMND5A was EcoRI digested, releasing the ~1.2kb RMND5A insert and 5µL gel purified product was electrophoresed in a 1% agarose gel. The DNA concentration was estimated from the gel to be ~10ng/µL. (F) pGEX-RMND5A clones 1-6 were EcoRI digested and the products were electrophoresed in a 1% agarose gel, identifying the RMND5A insert in all clones.

~1.2kb ~5kb

~1.2kb

E

F

Lane 1: MW Marker Lane 2: Purified RMND5A

Lanes 1, 14: MW Marker Lane 2, 4, 6, 8, 10, 12: Undigested pGEX-RMND5A clones 1, 2, 3, 4, 5, 6 Lane 3, 5, 7, 9, 11, 13: EcoRI digested pGEX- RMND5A clones 1, 2, 3, 4, 5, 6


106

LB Agar/Ampicillin plates (Section 3.8.6). Six colonies of pGEX-RMND5A were

inoculated into LB Broth/Ampicillin and small scale plasmid isolation was performed

followed by RNase treatment then EcoRI digestion to release the insert (Sections 3.8.2,

3.9). The reaction products were electrophoresed in a 1% agarose gel, identifying that

all six clones contained a product of the expected insert size of ~1.2kb (Section 3.6,

Figure 4.3F). pGEX-RMND5A clones 1 to 4 were purified and 5µL aliquots were

electrophoresed in a 1% agarose gel (Sections 3.6, 3.7.1, not shown). Based on the gel

image, 2µL each of pGEX-RMND5A clones 1 to 3 were sequenced using the pGEX-S

and pGEX-AS primers, with no mutations identified in pGEX-RMND5A clone 2 using

BLASTTM analysis (Section 3.12, Appendix II, Appendix III).

4.2.2.2 Small Scale Production of GST, GST-RMND5A and GST-RMND5B

pGEX-RMND5A clone 2 was transformed into E. coli BL21 cells and a glycerol stock

was prepared (Section 3.8.7, 4.2.2.1). To verify GST-fusion protein expression in

bacteria, pGEX-RMND5A and pGEX-RMND5B transformed E. coli BL21 cells were

selected on LB Agar/Ampicillin plates along with E. coli BL21 cells transformed with

pGEX-2TK (without insert) as a control. Expression of the GST-fusion proteins was

induced by the addition of 1mM IPTG (final concentration) and lysates of the cells were

electrophoresed in 12% polyacrylamide gels then stained with Coomassie blue to

visualise cellular proteins (Sections 3.11.1, 3.15.5, 3.15.1, 3.15.2). The presence in

lysates from IPTG-induced cultures of bands of ~25kDa (GST) and ~60kDa (GST-

RMND5A, GST-RMND5B) indicated that the transformed bacteria expressed the GST

fusion proteins, although the apparent molecular size of the GST-RMND5A and GST-

RMND5B was smaller than the expected molecular size of ~69kDa (Figure 4.4A).

4.2.2.3 Large Scale Production of GST-RMND5A

Following confirmation of bacterial production of GST-RMND5A (Section 4.2.2.2),

large scale purification of GST-RMND5A (Section 3.11.2) was performed and the

purified GST-RMND5A protein, along with aliquots collected from lysates of the

uninduced and IPTG induced cells, and the insoluble and unbound fractions were

electrophoresed in a 12% polyacrylamide gel, which was stained with Coomassie blue

(Sections 3.15.2, 3.15.5, Figure 4.5B). Although a prominent ~60kDa band was evident

in lysates from IPTG-induced cells, a ~60kDa band of purified GST-RMND5A was

only barely visible (Figure 4.4B). The prominent ~60kDa band was not evident in the


107

unbound fraction, which would usually indicate that the majority of the GST-RMND5A

had been bound to the glutathione sepharose beads, however, electrophoresis of the

insoluble fraction, which included a prominent ~60kDa band, showed that the majority

of the GST-RMND5A fusion protein was present in insoluble aggregates (Figure 4.4B).

To optimise the extraction of soluble GST-RMND5A, culture and purification

conditions were adjusted including IPTG induction of the cultures at room temperature,

28°, 30°C and 37°C, and from 30 minutes to 4 hours to capture the induced protein prior

to aggregation (not shown). In order to extract GST-RMND5A from the insoluble

aggregates, the bacterial lysates were sonicated for 4-6 x 30 second pulses on ice, and

up to seven elutions of the GST-RMND5A fusion proteins from the glutathione

sepharose was carried out at both room temperature or 4°C, however, similar low yields

of purified GST-RMND5A were obtained (Figure 4.4B). Finally, the pGEX-RMND5A

plasmid was transformed into Rosetta BL21 cells, which are codon-optimised bacterial

cells, and the production of GST-RMND5A was performed to determine whether this

strain also packaged the GST-RMND5A fusion protein into insoluble aggregates,

however, again the GST-RMND5A protein was present in the inclusion bodies with

very little in the soluble fraction (not shown). Thus, full length GST-RMND5A and

GST-RMND5B could not be purified from bacterial cells and an alternative approach

was taken whereby the RING domains of RMND5A and RMND5B would be cloned

into pGEX-2TK to generate GST-RMND5A RING and GST-RMND5B RING for use

in in vitro ubiquitination assays.

4.2.3 Cloning of RMND5 RING Domains for In Vitro Ubiquitination Assays

4.2.3.1 Cloning of RMND5 RING Domains into pGEX-2TK

To investigate the E3 ubiquitin ligase activity of RMND5A and RMND5B using the

RING domains of each of the proteins, the RMND5A (126bp) and RMND5B (126bp)

RING domains were PCR amplified from pEGFP-RMND5A and pEGFP-RMND5B,

respectively in PCRs utilising 1-2mM MgCl2 and a 55°C annealing temperature for 35

cycles (Section 3.4.1). The 117bp RING domain of CBL, a well-characterised E3

ubiquitin ligase was similarly PCR amplified from pCMV-CBL and electrophoresis of

each of the RING domains resulted in identification of bands of the expected sizes of

117bp (CBL) or 126bp (RMND5A, RMND5B) (Section 3.6, Figure 4.5A). The optimum

PCR conditions of 1.5mM MgCl2 and a 55°C annealing temperature were utilised to

amplify in quadruplicate the RING domains of each of RMND5A, RMND5B and CBL,


108

Figure 4.4: Expression and purification of full length GST-RMND5A and GST-RMND5B. (A) Expression of full length GST-RMND5A and GST-RMND5B was verified by electrophoresis and Coomassie blue staining of lysates from IPTG induced bacterial cells transformed with pGEX-RMND5A and pGEX-RMND5B. Protein bands at ~60kDa corresponding in size to GST-RMND5A and GST-RMND5A were identified in lysates from the IPTG-induced cultures. (B) GST-RMND5A was purified from IPTG induced pGEX-RMND5A transformed bacterial cells and the products electrophoresed in a 12% polyacrylamide gel which was stained with Coomassie blue. A ~60kDa band corresponding in size to GST-RMND5A was identified in lysates and purified products from IPTG induced cultures.

1 2 4 5 7

Lane 1: MW Marker Lane 2: GST: no IPTG Lane 3: GST: IPTG Lane 4: GST-RMND5A: no IPTG Lane 5: GST-RMND5A: IPTG Lane 6: GST-RMND5B: no IPTG Lane 7: GST-RMND5B: IPTG

~25kDa

~60kDa

1 2 3 4 5 6 7

Lane 1: MW Marker Lane 2: GST-RMND5A: no IPTG Lane 3: GST-RMND5A: IPTG, whole lysate Lane 4: GST-RMND5A: IPTG, soluble fraction Lane 5: GST-RMND5A: IPTG, unbound fraction Lane 6: GST-RMND5A: IPTG, purified fraction Lane 7: GST-RMND5A: IPTG, insoluble fraction

~60kDa

A

B

6 3


109

the four reactions for each RING domain were combined, purified and 5µL aliquots

were electrophoresed in a 2% agarose gel to verify product amplification (Sections

3.4.1, 3.6, Figure 4.5B).

The purified RING domains of RMND5A, RMND5B and CBL were ligated into the

pGEM®T-Easy cloning vector by ligating 30ng (1µL) of each PCR product into 50ng

(1µL) pGEM®T-Easy by TA cloning (Section 3.8.4). The ligation products were

transformed into competent E. coli DH5α and the transformed bacteria spread onto LB

Agar/Ampicillin plates containing X-gal to identify plasmids containing inserts using

blue/white colony selection (Section 3.8.6). Four white colonies for each of pGEMT-

RMND5A RING, pGEMT-RMND5B RING and pGEMT-CBL RING were picked,

inoculated into LB Broth/Ampicillin and incubated overnight. Plasmids were isolated

by small scale plasmid isolation, RNase treated, digested with BamHI and EcoRI and

the products were electrophoresed in 2% agarose gels (Section 3.6, 3.8.2, 3.9). Faint

insert bands of the appropriate size were identified in all of the plasmids and glycerol

stocks were prepared for each clone (Section 3.8.7, not shown).

To confirm that the correct sequences of the RMND5A, RMND5B and CBL RING

domains were present, plasmids from two clones of each of pGEMT-RMND5A RING,

pGEMT-RMND5B RING and pGEMT-CBL RING were purified, 2µL of each purified

plasmid was electrophoresed in a 1% agarose gel and based on the gels, 2µL of each

plasmid was sequenced using M13-S and M13-AS primers (Section 3.6, 3.7.1, 3.12, not

shown). Using BLASTTM, the correct RING domain sequences in pGEMT-RMND5A

clone 1, pGEMT-RMND5B clone 1 and pGEMT-CBL clone 1 were verified (Section

3.12, Appendix II, not shown).

To prepare the RING domains of RMND5A, RMND5B and CBL for subcloning into

pGEX-2TK, the RING domains of pGEMT-RMND5A RING clone 1, pGEMT-

RMND5B RING clone 1 and pGEMT-CBL RING clone 1 were released by digestion

with BamHI and EcoRI, the products were electrophoresed in 2% agarose gels, the

RING domain inserts were gel purified and 5µL of each electrophoresed in a 2%

agarose gel (Section 3.6, 3.7.1, 3.8.2, Figure 4.5C). Based on the gel, 7µL (30ng) of

each RING domain insert was ligated into 2µL (20ng) pGEX-2TK which had been

digested with BamHI and EcoRI and SAP treated (Section 3.8.2, 3.8.3, 3.8.4, Figure

4.5D). The ligation products were transformed into E. coli DH5α, selected by growth on


110

C Lane 1: MW Marker Lane 2: Purified RING RMND5A Lane 3: Purified RING RMND5B Lane 4: Purified RING CBL

~120bp

~120bp

Lane 1: MW Marker Lane 2: RMND5A RING (1mM MgCl2) Lane 3: RMND5A RING (1.5mM MgCl2) Lane 4: RMND5A RING (2mM MgCl2) Lane 5: RMND5B RING (1mM MgCl2) Lane 6: RMND5B RING (1.5mM MgCl2) Lane 7: RMND5B RING (2mM MgCl2) Lane 8: Negative Control Lane 9: MW Marker Lane 10: CBL RING (1mM MgCl2) Lane 11: CBL RING (1.5mM MgCl2) Lane 12: CBL RING (2mM MgCl2) Lane 13: Negative Control

A

~120bp

B Lane 1: MW Marker Lane 2: Purified RMND5A RING Lane 3: Purified RMND5B RING Lane 4: Purified CBL RING Lane 5: Negative Control


111

Figure 4.5: Cloning of sequences encoding the RING domains of RMND5A, RMND5B and CBL into pGEX-2TK. (A) PCR conditions for the amplification of the RING domains of RMND5A, RMND5B and CBL were optimised using LNCaP cDNA, an annealing temperature of 55°C and 1mM – 2mM MgCl2. (B) Following PCR optimisation, each RING domain was amplified in quadruplicate reactions, the reactions for each RING domain were combined and purified and 5µL of each purified RING domain was electrophoresed in a 2% agarose gel, from which the DNA concentrations were estimated to be ~30ng/µL. (C) pGEMT-RMND5A RING, pGEMT-RMND5B RING and pGEMT-CBL RING were digested with BamHI and EcoRI to release the RING inserts, the inserts were gel purified and 5µL of each purified insert was electrophoresed in a 2% agarose gel from which the concentration of each was estimated to be ~5ng/µL. (D) 5µL purified BamHI/EcoRI digested and SAP treated pGEX-2TK plasmid was electrophoresed in a 1% agarose gel from which the DNA concentration was estimated to be ~10ng/µL. (E) pGEX-RMND5A RING, pGEX-RMND5B RING and pGEX-CBL RING were purified and 5µL each purified product was electrophoresed in a 1% agarose gel.

Figure 4.9: BamHI and EcoRI digests and purification of pGEX-RING domains of RMND5A, RMND5B and CBL. (B) pGEX-RMND5A RING, pGEX-RMND5B RING and pGEX-CBL RING (i) clone 3 and (ii) clone 4 were purified and 5µL of each plasmid was electrophoresed in 1% agarose gels.

Lane 1: MW Marker Lane 2: Undigested pGEX-2TK Lane 3: Purified BamHI/EcoRI SAP digested pGEX-2TK

D

~5kb

Lane 1: MW Marker Lane 2: Purified pGEX-RING RMND5A clone 4 Lane 3: Purified pGEX-RING RMND5B clone 4 Lane 4: Purified pGEX-RING CBL clone 4

E

~5kb


112

LB Agar/Ampicillin plates and 4 colonies of bacteria transformed with each of pGEX-

RMND5A RING, pGEX-RMND5B RING and pGEX-CBL RING were inoculated into

LB Broth/Ampicillin for small scale plasmid isolation (Section 3.8.6, 3.9). Purified

plasmids were digested with RNase and following BamHI and EcoRI digestion to

release the inserts, plasmids were electrophoresed in 2% agarose gels, however no

inserts were visualised in the gels, potentially due to poor visibility of the small (117bp

and 126bp) sizes of the RING domain inserts (not shown). pGEX-RMND5A RING

clone 4, pGEX-RMND5B RING clone 4 and pGEX-CBL RING clone 4 were purified,

5µL aliquots of each were electrophoresed in a 1% agarose gel and 2µL of each plasmid

was sequenced using the pGEX-S and pGEX-AS primers (Section 3.6, 3.7.1, 3.12,

Appendix II, Figure 4.5E). Sequencing results indicated the presence of RING domain

inserts containing no mutations in all plasmids (Appendix III).

4.2.3.2 pGEX-RING Domain Protein Expression

Since each RING domain chelates two zinc ions, the medium in which bacterial

production of the GST fusion proteins was induced was supplemented with ZnCl2, as

was the buffer in which the eluted GST-fusion proteins was stored. This addition was

included in order to aid in the correct conformation and therefore functioning of the

RING domains. A range of ZnCl2 concentrations from 50µM - 250µM was tested to

determine whether the growth of E. coli BL21 cells was inhibited by the addition of

ZnCl2. For these experiments, the production of the GST-RMND5A RING domain

fusion protein was induced by the addition of 1mM IPTG to 5mL cultures, with 15µL

aliquots of the cell lysates electrophoresed in 12% polyacrylamide gels. Coomassie blue

staining of the gels identified that the amounts of bacterial proteins and the expression

of the GST-RMND5A RING domain were similar under all ZnCl2 concentrations

tested, indicating that the growth of E. coli BL21 cells and the production of GST-

fusion proteins was not affected by the addition of ZnCl2 to the culture medium (Section

3.11.1, 3.11.2). As such, 250µM ZnCl2 was added to the LB broth and 25µM ZnCl2 was

added to the elution buffer for subsequent experiments (Figure 4.6A).

For small scale production of GST-RING domains, the pGEX-RMND5A RING clone

4, pGEX-RMND5B RING clone 4 and pGEX-CBL RING clone 4 plasmids (Section

4.2.3.1) were transformed into E. coli BL21 cells and glycerol stocks were generated

(Section 3.8.7). To verify GST-fusion protein expression in bacteria, E. coli BL21 cells

transformed with pGEX-RMND5A RING clone 4, pGEX-RMND5B RING clone 4 and


113

pGEX-CBL RING clone 4 were streaked onto LB Agar/Ampicillin plates, individual

colonies were inoculated into LB broth/Ampicillin and expression of GST or the GST-

fusion proteins was induced by the addition of 1mM IPTG for 2.5 hours. Cells were

lysed, 15µL aliquots were electrophoresed in 12% polyacrylamide gels and the gels

were stained with Coomassie blue to visualise cellular proteins (Sections 3.11.1, 3.15.2,

3.15.5, Figure 4.6B). A prominent band of ~25kDa was evident in lysates of IPTG

treated cells transformed with pGEX-2TK (GST), while in those extracts of cells

transformed with pGEX-RMND5A RING, pGEX-RMND5B RING and pGEX-CBL

RING, prominent protein bands of the expected size of ~28kDa were present (Figure

4.6B). In lysates of cells transformed with pGEX-CBL RING, a strong band running at

a slightly higher than expected molecular weight of ~30kDa was evident (Figure 4.6B).

These findings indicated that following IPTG induction, each of the GST fusion

proteins was strongly expressed. For in vitro ubiquitination assays, GST and each of the

GST-RING domains were purified from 100mL bacterial cultures (Section 3.11.2,

4.2.4) and the purified proteins were electrophoresed in 12% polyacrylamide gels along

with BSA standards (Section 3.15.2). Gels were stained with Coomassie blue, indicating

successful production of GST and each of the GST-RING domain fusion proteins

(Section 3.15.5, Figure 4.6C, D).

4.2.4 In Vitro Auto-Ubiquitination Assays

4.2.4.1 Optimisation of In Vitro Auto-ubiquitination Assays using the GST-

CBL RING Domain

To optimise experimental conditions, in vitro ubiquitination assays were performed

using 4µM (final concentration) of the GST-RING domain of CBL and the E2

conjugating enzyme UbcH5b, which has been shown to mediate ubiquitin transfer with

CBL in vitro (Huang et al., 2009). These 50µL reactions were set up using an in vitro

Ubiquitinylation kit (ENZO Biosciences), reactions were terminated by the addition of

50µL 2x non-reducing loading dye and the products were electrophoresed in 12%

polyacrylamide gels (Section 3.14.1). Western blotting for biotinylated ubiquitin

resulted in the identification of a prominent band at >100kDa, as expected for a positive

result, however multiple additional bands at lower molecular weights were also present,

representing proteins bound to ubiquitin via isopeptide and thioester bonds (Figure 4.7).

In order to minimise the thioester-linked ubiquitinated protein bands, subsequent in

vitro ubiquitination assays were terminated by the addition of 2x reducing loading dye


114

A

B

1 2 3 4 5 6 7 8 9 10 Lane 1: MW Marker

Lane 2: 1µg BSA

Lane 3: 2.5µg BSA

Lane 4: 5µg BSA

Lane 5: 7.5µg BSA

Lane 6: 10µg BSA

Lane 7: 1µL GST-RMND5A RING

Lane 8: 3µL GST-RMND5A RING

Lane 9: 3µL GST-RMND5B RING

Lane 10: 1µL GST-RMND5B RING

~28kDa

C

Lane 1: MW Marker

Lane 2: GST: no IPTG

Lane 3: GST: IPTG

Lane 4: GST-RMND5A RING: no IPTG

Lane 5: GST-RMND5A RING: IPTG

Lane 6: GST-RMND5B RING: no IPTG

Lane 7: GST-RMND5B RING: IPTG

Lane 8: GST-CBL RING: no IPTG

Lane 9: GST-CBL RING: IPTG

~28kDa

1 2 3 4 5 6 7 8 9

~28kDa

1 2 3 4 5 6 7

Lane 1: MW Marker

Lane 2: GST-RMND5A RING: no IPTG

Lane 3: GST-RMND5A RING: IPTG

Lane 4: GST-RMND5A RING: IPTG and 50µM ZnCl2


Lane 6: GST-RMND5A RING: IPTG 200µM ZnCl2



115

Figure 4.6: Expression and purification of the GST-RING domains of

RMND5A, RMND5B and CBL. (A) The production of the GST-RMND5A RING

domain (~28kDa) was assessed by IPTG induction of pGEX-RMND5A RING

transformed E. coli BL21 cells that had been cultured in LB broth containing 50µM-

250µM ZnCl2. 15µL aliquots of each bacterial lysate was electrophoresed in a 12%

polyacrylamide gel and the gel was stained with Coomassie blue. (B) The

production of GST-RMND5A RING, GST-RMND5B RING and GST-CBL RING

domains was induced by the addition of IPTG to E. coli BL21 cultures transformed

with pGEX-RMND5A RING, pGEX-RMND5B RING and pGEX-CBL RING,

respectively. 15µL aliquots of each bacterial lysate was electrophoresed in 12%

polyacrylamide gels which were stained with Coomassie blue to determine GST

fusion protein production. The GST-RMND5A RING, GST-RMND5B RING and

GST-CBL RING domains were purified from large scale IPTG induced bacterial

cultures and the purified products electrophoresed in 12% polyacrylamide gels then

stained with Coomassie blue. The concentration of each GST-RING domain was

estimated in comparison to BSA standards, with the concentration of (C) GST-

RMND5A RING domain and GST-RMND5B RING domain estimated to be

~0.7µg/µL and ~0.9µg/µL, respectively, whilst (D) the concentration of GST-CBL

and GST were estimated to be ~1.9µg/µL and ~2.1µg/µL, respectively.

Lane 1: MW Marker

Lane 2: 1µg BSA

Lane 3: 2.5µg BSA

Lane 4: 5µg BSA

Lane 5: 7.5µg BSA

Lane 6: 10µg BSA

Lane 7: 1µL GST

Lane 8: 3µL GST

Lane 9: 1µL GST-CBL RING

Lane 10: 3µL GST-CBL RING

1 2 3 4 5 6 7 8 9 10

~25kDa ~28kDa

D


116

and the samples were boiled at 95°C for 5 minutes (Section 3.14.1). This resulted in the

detection of fewer bands compared to experiments where non-reducing loading dye was

used and the samples were not boiled (Figure 4.7).

4.2.4.2 In Vitro Auto-ubiquitination Assays Using the GST-RMND5A and

GST-RMND5B RING Domains

In vitro ubiquitination assays were performed using 4µM (final concentration) each of

the GST-RMND5A RING and GST-RMND5B RING domains along with the GST-

CBL RING domain as a positive control (Section 4.2.3.2). A negative control reaction

that lacked ATP was also included as the E1 enzyme requires ATP for the activation of

ubiquitin. Again the reactions were terminated using 2x reducing loading dye and the

products boiled at 95°C for 5 minutes prior to electrophoresis in 12% polyacrylamide

gels and western blotting for biotinylated ubiquitin (Section 3.14.1, 3.15). Under these

conditions, the negative control reaction resulted in a single band at ~10kDa

corresponding to free ubiquitin, as expected, whilst reactions where the GST-RING

domains of RMND5A and RMND5B were utilised resulted in a prominent band at

>100kDa, similar to that of the CBL RING domain (Figure 4.7). Lower molecular

weight bands were also present, corresponding to ubiquitinated RING domains and

ubiquitin isopeptide and thioester linked to the E2 conjugating enzyme UbcH5b. Initial

experiments therefore indicated that these experimental conditions would be suitable for

evaluation of RMND5A and RMND5B auto-ubiquitination activity.

4.2.4.3 Screening of E2 Conjugating Enzymes in In Vitro Ubiquitination

Assays

In order to determine the E2 enzymes with which RMND5A and RMND5B could

interact to mediate the transfer of ubiquitin, a panel of eleven E2 conjugating enzymes

including UbcH5b was utilised in in vitro ubiquitination assays (Section 3.14.1). The E2

enzymes UbcH1, UbcH2, UbcH5a, UbcH5b, UbcH5c, UbcH6, UbcH7, UbcH8,

UbcH10 and UbcH13 were tested with the GST-RING domains of RMND5A and

RMND5B in these experiments. Control reactions included UbcH5b and GST-CBL

RING domain (positive control), a reaction lacking ATP (negative control) and a

reaction lacking a GST-RING domain (negative control). Following electrophoresis of

reaction products in 12% polyacrylamide gels and western blotting for ubiquitin, high

molecular weight bands were present when the GST-RMND5A RING domain was


117

Figure 4.7: Optimisation of in vitro ubiquitination assays. The GST-CBL RING domain and UbcH5b were used to optimise in vitro ubiquitination assays. Reactions were initially terminated by the addition of 2x non-reducing loading dye (left panel) and 10µL of each reaction product was electrophoresed in a 12% polyacrylamide gel then analysed by western blotting for biotinylated ubiquitin. To reduce the presence of thioester-ubiquitinated proteins, subsequent reactions were terminated by the addition of 2x reducing loading dye and heating at 95°C for 5 minutes (middle panel). Using this method, in vitro ubiquitination assays containing the GST-CBL, GST-RMND5A or GST-RMND5B RING domains resulted in high molecular weight protein bands corresponding to polyubiquitinated proteins (right panel). Experiment was performed twice.

Chapter 4

Characterisation of RM

ND5 E3 U

biquitin Ligase Activity

Figure 4.8: RMND5 RING domains mediate ubiquitin transfer with specific E2 conjugating enzymes. In vitro ubiquitination assays were carried out using the GST-RMND5A or GST-RMND5B RING domains along with a panel of 11 E2 conjugating enzymes and 10µL each reaction product was electrophoresed in 4-12% gradient gels. Control reactions were also performed lacking ATP, E3 enzyme (negative controls) or containing the functional E3 ubiquitin ligase CBL (positive control). Following western blotting for biotinylated ubiquitin, high molecular weight bands corresponding to polyubiquitinated proteins were visible in assays containing the GST-RMND5A RING domain with UbcH2, UbcH5b and UbcH5c and the GST-RMND5B RING domain with UbcH5b and UbcH5c. Experiment was performed once.

118

RMND5A RING domain RMND5B RING domain

Ub- RING(>100kDa)

UbProteins

(~15-50kDa)

Free Ubiquitin (~10kDa)

Controls


119

utilised in in vitro reactions with the E2 enzymes UbcH2, UbcH5b and UbcH5c, whilst

similar high molecular weight bands were present when the GST-RMND5B RING

domain was utilised in in vitro ubiquitination assays with UbcH5b and UbcH5c (Section

3.14.1, 3.15, Figure 4.8). These results indicated that RMND5A and RMND5B both

exhibit E3 ubiquitin ligase activity in vitro and that RMND5A interacts with UbcH2,

UbcH5b and UbcH5c, while RMND5B interacts with UbcH5b and UbcH5c to mediate

ubiquitin transfer.

4.2.4.4 Control In Vitro Auto-Ubiquitination Assays

In order to determine whether the results obtained in in vitro ubiquitination assays

(Section 4.2.4.2, 4.2.4.3) were specific to the inclusion of GST-RMND5A RING or

GST-RMND5B RING domains, the assays were repeated using the E2 enzyme UbcH5b

and each time omitting a reaction ingredient (Section 3.14.1). In addition, GST alone

was used in a control reaction. Where GST alone was utilised, lower molecular weight

bands were present, corresponding to free ubiquitin, polyubiquitin or ubiquitinated E2

enzyme (Figure 4.9). In negative control reactions where ubiquitin, ATP, E1 or E2 were

omitted, only free ubiquitin or polyubiquitin bands were present, whilst reactions where

GST-RMND5A RING or GST-RMND5B RING domains were omitted resulted in the

same bands as well as additional products corresponding to ubiquitinated E2 enzyme

(Figure 4.9). In reactions where GST-RMND5A RING and GST-RMND5B RING

domains were added, bands corresponding to free ubiquitin, polyubiquitin and

ubiquitinated E2 enzymes at lower molecular weights were evident, in addition to a

band of ~38kDa that was not present in other reactions, which corresponded in size to

the monoubiquitinated RING domains (Figure 4.9). These results indicated that the

positive results in in vitro ubiquitination assays (Section 4.2.4.2, 4.2.4.3) were specific

for the GST-RMND5A RING and GST-RMND5B RING domains and that RMND5

proteins were able to auto-monoubiquitinate their RING domains.

4.2.5 In Vivo Ubiquitination Assays

In order to determine whether full length RMND5 proteins possess E3 ubiquitin ligase

activity in mammalian cells, in vivo ubiquitination assays were performed by

transfecting LNCaP cells with plasmids encoding GFP-RMND5A or GFP-RMND5B

and HA-Ubiquitin, and following 6 hours of proteasome inhibition with 10µM

lactacystin the cells were lysed at 48 hours post-transfection (Section 3.1.4, 3.1.5, 3.10).


120

-

RMND5BRING-Ub~38kDa

Free Ub(~10kDa)

RMND5ARING-Ub~38kDa

Free Ub(~10kDa)

ATPUbiquitin

E1UbcH5bGST-RINGGST

++++

+-

++++-

-+

+++-

-

++

++-

-

+++

+-

-

++++

-

++++

-+

WB: Bt-Ubiquitin

Figure 4.9: Control reactions for in vitro ubiquitination assays. In vitro

ubiquitination assays were performed using UbcH5b and a full set of

controls for each of RMND5A and RMND5B. Controls included the

omission of one ingredient per reaction and inclusion of GST instead of

GST-RMND5A RING domain or GST-RMND5B RING domain. The

presence of a band at ~38kDa corresponding in size to the

monoubiquitinated RING domain was only present in reactions containing

all assay ingredients and either GST-RMND5A or GST-RMND5B. The

experiment was performed twice and representative results are shown.


121

GFP immunoprecipitation was used to isolate GFP-tagged RMND5A and RMND5B

proteins, which were electrophoresed in 4-12% gradient polyacrylamide gels to allow

separation of high molecular weight ubiquitinated proteins. Increased expression of high

molecular weight protein bands were detected by HA western blotting (for HA-

ubiquitin) in lysates of cells that overexpressed GFP-RMND5A or GFP-RMND5B,

indicating that RMND5A and RMND5B were associated with ubiquitinated proteins in

LNCaP cells (Section 3.13, 3.15, Figure 4.10). These results were enhanced when

cultures were treated with the proteasome inhibitor, lactacystin which allowed

accumulation of ubiquitinated proteins, producing intense smears of high molecular

weight proteins of >80kDa (Figure 4.10). Western blotting of the total cell lysates of

these samples identified the presence of GFP, GFP-RMND5A and GFP-RMND5B and

the similar expression of HA-ubiquitin in all samples, indicating that the results were

due to differences in levels of GFP-RMND5A/GFP-RMND5B associated ubiquitinated

proteins (not shown). These experiments provided evidence that RMND5 proteins are

associated with ubiquitinated proteins in LNCaP cells, which is consistent with the in

vitro ubiquitination assay results.

4.2.6 Investigation of the E3 Ubiquitin Ligase Activity of the RMND5A and

RMND5B RING Domains using RMND5A (C356S) and RMND5B (C358S)

RING Domain Mutants

In order to demonstrate that the RING domains of RMND5 proteins were responsible

for their E3 ubiquitin ligase activity in in vitro and in vivo ubiquitination assays, a single

amino acid change was introduced into the RING domains of RMND5A (C356S) and

RMND5B (C358S) using site-directed mutagenesis (Section 3.4.3, Figure 4.11). The

corresponding mutation in the yeast RMD5 (C379S) was previously shown to inhibit its

ability to ubiquitinate its substrate protein, fructose-1, 6-bisphosphatase (Santt et al.,

2008). Once the full length pEGFP-RMND5A (C356S) and pEGFP-RMND5B (C358S)

mutants were generated, the RMND5A (C356S) and RMND5B (C358S) RING domains

were cloned into the pGEX-2TK expression vector to allow the expression of GST-

RMND5A (C356S) RING and GST-RMND5B (C358S) RING.

4.2.6.1. Introduction of C356S into RMND5A RING Domain

Site-directed mutagenesis was performed using a PCR based method and initial

optimisation of PCR conditions was carried out in 26 cycle reactions using a gradient of


122

Figure 4.10: In vivo ubiquitination activity of RMND5 proteins. LNCaP cells were transfected with plasmids encoding HA-Ubiquitin, GFP-RMND5A or GFP-RMND5B or empty vector and cultured for 48 hours post-transfection, with 10µM Lactacystin added for the final 6 hours of culture. Following GFP immunoprecipitation of cell lysates, HA western blotting was performed on the immunoprecipitated proteins, identifying increased association of GFP-RMND5A and GFP-RMND5B with ubiquitinated proteins that was enhanced by proteasome inhibition. The experiment was performed twice and representative results are shown.


123

Figure 4.11: Site directed mutagenesis of the RING domains of RMND5A and RMND5B. Alignment of the RING domains of RMND5A, RMND5B and yeast RMD5 showing all eight amino acid residues required for RING domain activity (red/orange). These include the conserved cysteine residues (bold, orange) to be mutated to serine (green) in order to reduce or inactivate the RING domain activity of RMND5A and RMND5B. Numbers indicate amino acid number.




RMND5A (C356S)

RMND5B (C358S)

RMD5 (C379S)

S

336 377

338 379

361 404


124

annealing temperatures ranging from 65°C-72°C, 1.5mM MgCl2 and the pEGFP-

RMND5A plasmid as the DNA template (Section 3.4.3). Electrophoresis of 10µL of the

amplified products identified a single band at the expected size of ~5.9Kb, which was

present at all annealing temperatures tested but was more prominent at lower annealing

temperatures (Figure 4.12A). The reaction products from PCRs performed with

annealing temperatures of 65°C and 65.5°C were digested with DpnI to degrade the

parental methylated plasmid DNA and the reaction products were transformed into E.

coli DH5α and selected by plating on LB Agar/Kanamycin (Section 3.4.3.1, 3.8.6). Six

of the resulting colonies were picked, cultured in LB Broth/Kanamycin and plasmids

isolated by small scale plasmid purification (Section 3.9). Plasmids were treated with

RNase then digested with EcoRI to release the RMND5A insert and 10µL of the reaction

products were electrophoresed in a 1% agarose gel (Section 3.6, 3.8.2, Figure 4.12B).

Bands corresponding to the ~1.2kb RMND5A (C356S) insert were identified in all

clones and the pEGFP-RMND5A (C356S) clones 1 to 4 were purified, 5µL of each

purified clone was electrophoresed in a 1% agarose gel to verify the presence of the

~5.9kb insert and based on the gel, 2µL of each clone was sequenced using the

RMND5A1176-AS primer (Section 3.6, 3.7.1, 3.12, Appendix II, not shown).

BLASTTM analysis of the sequencing chromatograms verified the presence of the

C356S mutation in clones 1 and 4, whilst clones 2 and 3 each had a region of inserted

DNA, therefore, the entire coding region of the pEGFP-RMND5A (C356S) clones 1

and 4 was sequenced using the RMND5A603-S, RMND5A1-S and RMND5A490-AS

primers (Section 3.12, Appendix II). Sequencing analysis detected no mutations aside

from the G1061C base substitution that encoded the C356S amino acid change (not

shown).

To ensure that no further mutations were present in the pEGFP-C2 vector that contained

the RMND5A (C356S) RING domain mutation, the RMND5A coding region was

excised from the pEGFP-C2 plasmid by digestion with EcoRI, gel purified and the

purified insert electrophoresed in a 1% agarose gel, verifying the presence of the ~1.2kb

product (Sections 3.6, 3.7.2, 3.8.2, Figure 4.12C). Fresh pEGFP-C2 vector was digested

with EcoRI, SAP treated, purified and a 2µL aliquot electrophoresed in a 1% agarose

gel (Section 3.8.2, 3.8.3, 3.6, Figure 4.12D). Sixty ng (4µL) RMND5A (C356S) was

ligated with fifty ng (0.5µL) pEGFP-C2 and the products were transformed into

competent E. coli DH5α (Sections 3.8.2, 3.8.4). Transformed bacteria were grown on

LB Agar/Kanamycin plates, 4 colonies were inoculated into LB Broth/Kanamycin


125

~1.2kb

~4.7kb

Lane 1: MW Marker Lane 2: Undigested pEGFP-RMND5A (C356S) Clone 1 Lane 3-6: EcoRI digested pEGFP-RMND5A (C356S) Clone 1-4

E

A Lane 1: MW Marker Lane 2: 72°C Lane 3: 71.6°C Lane 4: 70.9°C Lane 5: 69.5°C Lane 6: 67.8°C Lane 7: 66.4°C Lane 8: 65.5°C Lane 9: 65°C Lane 10: Negative Control

~5.9kb

Lane 1: MW Marker Lane 2: Undigested pEGFP-RMND5A (C356S) Clone 1 Lane 3-8: EcoRI digested pEGFP-RMND5A (C356S) Clones 1-6

B

~1.2kb

~4.7kb

Lane 1: MW Marker Lane 2: RMND5A (C356S) Clone 1 Lane 3: RMND5A (C356S) Clone 4

~1.2kb

Lane 1: MW Marker Lane 2: Undigested pEGFP-C2 Lane 3: EcoRI digested pEGFP-C2

~4.7kb

C D


126

Figure 4.12: Preparation of pEGFP-RMND5A (C356S). (A) Wild type pEGFP-RMND5A and an annealing temperature gradient (65°C to 72°C) were used to optimise site directed mutagenesis PCR conditions. 10µL of each DpnI digested PCR product was electrophoresed in a 1% agarose gel, visualising the mutagenesis products at ~5.9kb in all reactions. (B) Plasmids isolated from pEGFP-RMND5A (C356S) clones 1 to 6 were digested with EcoRI to liberate the ~1.2kb insert then electrophoresed in a 1% agarose gel confirming the presence of insert in all clones. (C) pEGFP-RMND5A (C356S) clones 1 and 4 were digested with EcoRI to release the ~1.2kb RMND5A (C356S) coding region which was gel purified. 5µL of each purified product was electrophoresed in a 1% agarose gel from which the DNA concentration was estimated to be ~15ng/µL. (D) 2µL of purified EcoRI/SAP digested pEGFP-C2 plasmid was electrophoresed in a 1% agarose gel from which the concentration of DNA was estimated to be ~100ng/µL. (E) Plasmids isolated from pEGFP-RMND5A (C356S) clones 1 to 4 were digested with EcoRI to liberate the ~1.2kb insert, then electrophoresed in a 1% agarose gel, confirming that all clones contained an insert. (F) To determine the insert orientation, pEGFP-RMND5A (C356S) mutant clones 1 to 4 were digested with SacI, then electrophoresed in a 1% agarose gel. The presence of a band at ~300bp in the digested plasmids of clones 1-4 indicated that the RMND5A (C356S) insert was in the sense orientation in all plasmids.

~300bp

~5.6kb

Lane 1: MW Marker Lane 2: SacI digested wild type pEGFP-RMND5A Lane 3: Undigested pEGFP-RMND5A (C356S) Clone 1 Lane 4-7: SacI digested pEGFP-RMND5A (C356S) Clone 1-4

F


127

cultures for small scale plasmid purification and the purified plasmids RNase treated

then digested with EcoRI to release the insert (Section 3.6, 3.8.2, 3.9). Following

electrophoresis in a 1% agarose gel, a band at the expected size of ~1.2kb was present in

all four clones, and to determine insert orientation, the plasmids were digested with SacI

then electrophoresed in a 1% agarose gel (Sections 3.6, 3.8.2, Figure 4.12E). The

presence of a band at ~300bp indicated that all clones were inserted in the pEGFP-C2

vector in the sense orientation (Figure 4.12F). Each of the pEGFP-RMND5A (C356S)

clones was purified and 5µL product electrophoresed in a 1% agarose gel to estimate

the DNA concentration (Sections 3.6, 3.7.1, not shown). Based on the gel, 2µL of each

of clones 2 and 3 was utilised in sequencing reactions using the RMND5A1-S,

RMND5A1176-AS, RMND5A603-S and RMND5A490-AS primers, the products of

which were analysed using BLASTTM (Section 3.12, Appendix II). From this analysis,

pEGFP-RMND5A (C356S) clone 2 was verified as mutation free, the sense orientation

of the insert was confirmed and glycerol stocks of this clone were prepared (Section

3.8.7, 3.12, Appendix III).

4.2.6.2 Introduction of C358S into the RMND5B RING Domain

Initial site-directed mutagenesis PCRs using RMND5B (C358S) Primer set 1 (Appendix

II) to incorporate the RMND5B (C358S) RING domain mutation did not amplify

products and optimisation of PCR conditions including the use of a range of annealing

temperatures from 50°C - 72°C, 1.5mM and 2mM MgCl2, high fidelity and high GC

content buffers and increased initial denaturation, annealing and extension times were

not successful (not shown). The mutagenesis PCR primers were redesigned and

shortened (RMND5B (C358S) Primer set 2) and the PCR conditions re-optimised using

a range of annealing temperatures from 65°C - 72°C, which resulted in amplification of

similar levels of a single product at the expected size of ~5.9kb at all annealing

temperatures (Section 3.4.3, 3.6, Figure 4.13A). The negative control contained a faint

smear of DNA which was attributed to primer self-annealing (Figure 4.13A). PCR

products from reactions amplified with an annealing temperature of 65°C and 65.5°C

were digested with DpnI to degrade the wild-type parental DNA, the products were

transformed into competent E. coli DH5α cells and selected by plating onto LB

Agar/Kanamycin (Sections 3.4.3, 3.8.6). Four of the resulting colonies were picked,

incubated overnight in LB Broth/Kanamycin and plasmids isolated by small scale

plasmid purification then treated with RNase and SalI digested to liberate the insert

(Section 3.8.2, 3.9). Electrophoresis of 10µL of each product identified a band at the


128

expected size of ~1.2kb and each of the plasmids was purified then electrophoresed in

preparation for sequencing reactions (Section 3.6, 3.7.1, Figure 4.13B). Based on the

gel, 3µL of each of clones 1 to 4 was sequenced using the RMND5B790-S primer, and

the sequencing chromatograms analysed using BLASTTM, verifying the presence of the

RMND5B C358S mutation in each of the four clones (not shown). To confirm the entire

RMND5B sequence, pEGFP-RMND5B (C358S) clones 1 and 2 were sequenced using

the RMND5BSalI1-S, RMND5BSalI1182-AS and pEGFP1266-S primers, verifying the

absence of mutations in the remainder of the RMND5B coding region (Appendix II, not

shown).

To subclone RMND5B (C358S) into pEGFP-C2, pEGFP-RMND5B (C358S) clone 1

was digested with SalI to release the RMND5B (C358S) insert, the insert was gel

purified then 5µL was electrophoresed in a 1% agarose gel (Section 3.6, 3.7.2, 3.8.2,

Figure 4.13C). pEGFP-C2 was prepared by digestion with SalI, SAP treatment,

purification, then 2µL product was electrophoresed in a 1% agarose gel (Section 3.6,

3.8.2, 3.8.3, Figure 4.13C). The ligation reaction contained 50ng (5µL) RMND5B

(C358S) insert and 50ng pEGFP-C2, the ligation products were transformed into

competent E. coli DH5α and transformed bacteria were selected by growth overnight on

LB Agar/Kanamycin (Section 3.8.4, 3.8.6). Ten colonies were picked, cultured in LB

Broth/Kanamycin for small scale plasmid purification, the resulting plasmids treated

with RNase, digested with SalI to liberate the insert and 10µL of each product was

electrophoresed in a 1% agarose gel (Section 3.6, 3.8.2, 3.9, Figure 4.13D). A band at

the expected size of ~1.2kb was present in clones 5, 6 and 10, and to determine the

insert orientation, these were digested with PstI, with clone 6 the only plasmid

producing a single band at ~315bp, indicating that this clone contained the insert in the

sense orientation (Section 3.6, 3.8.2, Figure 4.13E). pEGFP-RMND5B (C358S) clone 6

was purified, sequenced using the RMND5BSalI1-S, RMND5BSalI1182-AS,

RMND5B790-S and pEGFP1266-S primers and the sequences verified to be mutation

free (apart from the G1067C base change resulting in the C358S mutation) and in the

sense orientation (Section 3.12, Appendix II, Appendix III). A glycerol stock was

prepared of this mutant (Section 3.8.7).


129

Lane 1: MW Marker Lane 2: 72°C Lane 3: 71.6°C Lane 4: 70.9°C Lane 5: 69.5°C Lane 6: 67.8°C Lane 7: 66.4°C Lane 8: 65.5°C Lane 9: 65°C Lane 10: Negative Control

~5.9kb

Lane 1: MW Marker Lane 2: Undigested pEGFP-RMND5B (C358S) Clone 1 Lane 3-6: SalI digested pEGFP-RMND5B (C358S) Clone 1-4

~1.2kb ~4.7kb

B

~1.2kb ~4.7kb

Lane 1: MW Marker Lane 2: RMND5B (C358S) Clone 1 Lane 3: Undigested pEGFP-C2 Lane 4: SalI digested pEGFP-C2

C

A

~1.2kb ~4.7kb

Lane 1: MW Marker Lane 2: Undigested pEGFP-RMND5B (C358S) Clone 1 Lane 3 - 10: SalI digested pEGFP-RMND5B (C358S) Clone 1 - 10

D


130

Figure 4.13: Preparation of pEGFP-RMND5B (C358S). (A) Wild type pEGFP-RMND5B and an annealing temperature gradient ranging from 65°C to 72°C were used to optimise RMND5B site-directed mutagenesis PCR conditions. 10µL of each DpnI digested PCR product was electrophoresed in a 1% agarose gel, and the mutagenesis products visualised at ~5.9kb. (B) Plasmids isolated from pEGFP-RMND5B (C358S) clones 1 to 4 were digested with SalI to liberate the ~1.2kb insert then electrophoresed in a 1% agarose gel, confirming the presence of insert in all clones. (C) pEGFP-RMND5B (C358S) mutant clone 1 was SalI digested to liberate the RMND5B (C358S) coding region and 5µL of the gel purified insert was electrophoresed in a 1% agarose gel from which the concentration was estimated to be 10ng/µL. 2µL of purified SalI/SAP digested pEGFP-C2 plasmid was electrophoresed in a 1% agarose gel from which the DNA concentration was estimated to be 300ng/µL. (D) Plasmids isolated from pEGFP-RMND5B (C358S) clones 1 to 10 were digested with SalI then 5µL of each digested product was electrophoresed in a 1% agarose gel. Clones 5, 6 and 10 contained the RMND5B (C358S) insert at ~1.2kb. (E) To determine insert orientation, pEGFP-RMND5B (C358S) clones 4, 5, 6, 10 were digested with PstI then electrophoresed in a 1% agarose gel. The presence of two bands at ~1kb and ~300bp indicated that the RMND5B (C358S) inserts in clones 5 and 6 were in the antisense orientation whilst a single band at ~300bp indicated that the RMND5B (C358S) insert was present in the sense orientation.

~1 kb

Lane 1: MW Marker Lane 2: Undigested pEGFP-RMND5B (C358S) Clone 4 Lane 3: PstI digested pEGFP-RMND5B (C358S) Clone 4 Lane 4: PstI digested pEGFP-RMND5B (C358S) Clone 5 Lane 5: PstI digested pEGFP-RMND5B (C358S) Clone 10 Lane 6: MW Marker Lane 7: Undigested pEGFP-RMND5B (C358S) Clone 6 Lane 8: PstI digested pEGFP-RMND5B (C358S) Clone 6

~5.6kb

~300bp

E ~5.6kb ~300bp


131

4.2.6.3 Cloning of the RMND5A (C356S) and RMND5B (C358S) RING

Domains into pGEX-2TK

Following development of RMND5A (C356S) (Section 4.2.6.1) and RMND5B (C358S)

(Section 4.2.6.2), the mutant RING domains were PCR amplified from these plasmids

using the previously optimised PCR conditions, the amplified products purified,

electrophoresed in a 2% agarose gel, then 60ng (2µL) of each PCR product was ligated

into 50ng (1µL) pGEM®T-Easy cloning vector (Section 3.4, 3.6, 3.7.1, Figure 4.14A).

The ligation products were transformed into competent E. coli DH5α cells and selected

by growth of the transformed bacteria on LB Agar/Ampicillin plates with blue/white

colony selection, then 4 colonies each of the pGEMT-RMND5 RING domain mutants

were picked and incubated in LB Broth/Ampicillin for small scale plasmid purification

(Section 3.8.6). Purified plasmids were treated with RNase, digested with BamHI and

EcoRI to release the insert, and the digestion products electrophoresed in a 2% agarose

gel (Section 3.6, 3.8.2). A band of the expected molecular weight of ~126bp was

present in all digested plasmids of each of pGEMT-RMND5A (C356S) RING and

pGEMT-RMND5B (C358S) RING, therefore clones 1 and 2 were purified then

electrophoresed in a 1% agarose gel in preparation for sequencing (Section 3.6, 3.7.1,

Figure 4.14B). Based on the gel, 4µL pGEMT-RMND5A (C356S) RING clones 1 and 2

and 2µL pGEMT-RMND5B (C358S) RING clones 1 and 2 were sequenced using the

M13-S and M13-AS primers, with BLASTTM analysis of the sequencing products

verifying the presence of the RMND5A (C356S) and RMND5B (C358S) RING domain

mutations and no additional base changes (Section 3.12, Appendix II, not shown).

To prepare the RMND5A (C356S) RING and RMND5B (C358S) RING domains for

ligation, pGEMT-RMND5A (C356S) RING clone 1 and pGEMT-RMND5B (C358S)

RING clone 1 were each digested with BamHI and EcoRI to release the ~126bp inserts,

the products gel purified and 5µL of each electrophoresed in a 2% agarose gel in

preparation for ligation reactions (Sections 3.6, 3.7.2, 3.8.2, Figure 4.14C). The pGEX-

2TK vector was prepared by small scale plasmid purification, RNase treatment, BamHI

and EcoRI digestion and SAP treatment, prior to purification and electrophoresis in a

1% agarose gel (Sections 3.6, 3.7.1, 3.8.2, 3.9, Figure 4.14D). Based on the gels, 40ng

(8µL) RMND5A (C356S) RING and 60ng (4µL) RMND5B (C358S) RING were ligated

with 50ng (2.5µL) pGEX-2TK, the resulting products were transformed into competent

E. coli DH5α cells and selected by plating cultures on LB Agar/Ampicillin (Sections


132

Lane 1: MW Marker Lane 2: RMND5A (C356A) RING Lane 3: RMND5B (C358S) RING Lane 4: Negative Control

Lane 1: MW Marker Lane 2: Undigested pGEMT-RMND5A (C356S) RING clone 1 Lane 3-6: EcoRI digested RMND5A (C356S) RING clone 1-4 Lane 7-10: EcoRI digested RMND5B (C358S) RING clone 1-4 Lane 11: MW Marker

~126bp

~126bp

~3kb

A

B

Lane 1: MW Marker Lane 2: RMND5A (C356S) RING Lane 3: RMND5B (C358S) RING Lane 4: Negative Control

~126bp

~5kb

Lane 1: MW Marker Lane 2: Undigested pGEX-2TK Lane 3: BamHI/EcoRI SAP digested pGEX-2TK

C

D


133

Figure 4.14: Cloning of sequences encoding the RMND5A (C356S) and RMND5B (C358S) RING domains into pGEX-2TK. (A) The RING domains of pEGFP-RMND5A (C356S) and pEGFP-RMND5B (C358S) were PCR amplified and 5µL of each purified product was electrophoresed in a 2% agarose gel from which the DNA concentration of each was estimated to be 30ng/µL. (B) Plasmids isolated from pGEMT-RMND5A (C356S) RING and pGEMT-RMND5B (C358S) RING clones 1 to 4 were BamHI and EcoRI digested and the products were electrophoresed in a 2% agarose gel confirming that each clone contained an insert. (C) pGEMT-RMND5A (C356S) RING and pGEMT-RMND5B (C358S) RING were BamHI/EcoRI digested, gel purified and 5µL product was electrophoresed in a 2% agarose gel from which the concentration of RMND5A (C356S) and RMND5B (C358S) were estimated to be ~5ng/µL and 15ng/µL, respectively. (D) 5µL of purified BamHI/EcoRI digested pGEX-2TK plasmid was electrophoresed in a 1% agarose gel and the plasmid concentration estimated from the gel to be ~20ng/µL. (E) pGEX-RMNDA (C356S) RING and pGEX-RMND5B (C358S) RING clones 1 and 2 were purified and 5μL product electrophoresed in a 1% agarose gel.

Lane 1: MW Marker Lane 2: Purified pGEX-RMND5A (C356S) RING clone 1 Lane 3: Purified pGEX-RMND5A (C356S) RING clone 2 Lane 4: Purified pGEX-RMND5B (C358S) RING clone 1 Lane 5: Purified pGEX-RMND5B (C358S) RING clone 2

~5kb

E


134

3.8.4, 3.8.6). Four colonies of each of the pGEX-RMND5 RING domain mutants were

inoculated into LB Broth/Ampicillin for small scale plasmid purification, plasmids were

treated with RNase, digested with BamHI and EcoRI to liberate the RING domain

inserts and 10µL of each product was electrophoresed in a 2% agarose gel (Sections 3.6,

3.8.2, 3.9). A faint ~126bp band was evident in all pGEX-RMND5A (C356S) RING

clones 1-4 and pGEX-RMND5B (C358S) RING clones 1-4 (not shown), and pGEX-

RMND5A (C356S) RING and pGEX-RMND5B (C358S) RING clones 1 and 2 were

purified and electrophoresed in a 1% agarose gel in preparation for sequencing

(Sections 3.6, 3.7.1, Figures 4.14E). Based on the gel, 4µL of each of pGEX-RMND5A

(C356S) RING clones 1 and 2 and 7µL pGEX-RMND5B (C358S) RING clones 1 and 2

were sequenced using the pGEX-S and pGEX-AS primers, with BLASTTM analysis

identifying that pGEX-RMND5A (C356S) RING clone 2 and pGEX-RMND5B

(C358S) RING clone 2 each contained no additional mutations (Section 3.12, Appendix

II, not shown). These plasmids were transformed into E. coli BL21 cells and glycerol

stocks were prepared (Section 3.8.6, 3.8.7).

4.2.6.4 Expression and Intracellular Localisation of RMND5A (C356S) and

RMND5B (C358S)

In preparation for transfection into mammalian cells, large scale plasmid preparations of

pEGFP-RMND5A (C356S) and pEGFP-RMND5B (C358S) were performed and the

plasmid DNA concentrations determined to be 1.93µg/µL for pEGFP-RMND5A

(C356S) and 2.2µg/µL for pEGFP-RMND5B (C358S) (Section 3.10, 4.2.6.1, 4.2.6.2).

To verify the expression of GFP-RMND5A (C356S) and GFP-RMND5B (C358S),

LNCaP cells growing in 6 well plates were transfected with 4µg plasmids encoding

either wild-type or mutant GFP tagged-RMND5A or RMND5B (Section 3.1.4).

Cultures were harvested 48 hours following transfection and GFP western blotting

identified the presence of a ~70kDa band in the lysates of cells transfected with each of

the constructs (Section 3.15, Figure 4.15A). These findings indicated that mutation of

the RING domains of RMND5A and RMND5B did not inhibit the expression or levels

of expression of the proteins.

The intracellular localisation of the GFP-RMND5A (C356S) and GFP-RMND5B

(C358S) was investigated by transfection of LNCaP cells growing on coverslips with

2µg plasmids encoding GFP-tagged wild-type or mutant RMND5 proteins, with the


135

A

B GFP Nucleus Cytoplasm Overlay

GFP-RMND5A

Wild Type

C356S

GFP-RMND5B

Wild Type

C358S

Untransfected Negative Control


136

Figure 4.15: Expression and cellular localisation of GFP-RMND5A (C356S) and GFP-RMND5B (C358S) proteins. LNCaP cells were transfected with plasmids encoding either GPF-RMND5A, GFP-RMND5B, GFP-RMND5A (C356S) or GFP-RMND5B (C358S) and at 48 hours post-transfection (A) the cells were harvested for GFP western blotting, which identified bands corresponding to GFP-RMND5A, GFP-RMND5A (C356S), GFP-RMND5B and GFP-RMND5B (C358S) at ~70kDa. (B) Alternatively, the cells were prepared for fluorescence microscopy and stained with Hœchst 33258 and tetramethylisothiocyanate (TRITC)-Phalloidin to image the nucleus and cytoplasm, respectively. GFP-RMND5A, GFP-RMND5B, GFP-RMND5A (C356S) and GFP-RMND5B (C358S) displayed diffuse nuclear and cytoplasmic localisation, with GFP-RMND5B and GFP-RMND5B (C358S) also exhibiting a punctate cytoplasmic distribution (Magnification x1000,). Experiments were performed twice and representative results are shown. (WT= wild-type).


137

cells fixed at 48 hours post transfection and viewed by fluorescence microscopy

(Section 3.1.3, 3.1.4, 3.16.1). Both GFP-RMND5A and GFP-RMND5A (C356S)

proteins exhibited a diffuse appearance and were localised in the nucleus and

cytoplasm, with occasional cells transfected with wild-type RMND5A exhibiting

punctate cytoplasmic staining (Figure 4.15B). The GFP-RMND5B and GFP-RMND5B

(C358S) proteins similarly displayed a diffuse appearance and both nuclear and

cytoplasmic localisation, with most cells also exhibiting a punctate cytoplasmic

distribution (Figure 4.15B). These findings indicated that the RING domain mutations

did not markedly alter the cellular intracellular localisation of RMND5 proteins in

comparison to the wild-type RMND5A and RMND5B.

4.2.6.5 In Vivo Ubiquitination Activity of RMND5A (C356S) and RMND5B

(C358S)

The activity of RMND5A (C356S) and RMND5B (C358S) was evaluated in in vivo

ubiquitination assays in comparison to the activity of wild-type RMND5 proteins. For

these experiments, LNCaP cells were cotransfected with 15µg of plasmids encoding

HA-Ubiquitin and GFP-RMND5A, GFP-RMND5A (C356S), GFP-RMND5B or GFP-

RMND5B (C358S), cultured for 42 hours, then treated for the final 6 hours with the

proteasome inhibitor, MG132 to allow the accumulation of ubiquitinated proteins within

the cells (Section 3.1.4, 3.1.5, 3.14.2). The cells were harvested at 48 hours post-

transfection, GFP-tagged RMND5 proteins were immunoprecipitated, and the

immunoprecipitated proteins analysed by HA western blotting (for HA-Ubiquitin)

(Section 3.13, 3.15, Figures 4.18, 4.16). All GFP immunoprecipitated products

contained high molecular weight HA (ubiquitin) bands, corresponding to ubiquitinated

proteins which were more prominent in immunoprecipitates from the MG132 treated

cells (Figure 4.16). Both of the (wild-type) GFP-RMND5A and GFP-RMND5B

immunoprecipitates were associated with increased levels of high molecular weight

ubiquitinated proteins in comparison to cells overexpressing GFP alone, and the levels

of ubiquitinated proteins were further enhanced when cells were treated with MG132

(Figure 4.16). However, GFP-RMND5A, GFP-RMND5A (C356S), GFP-RMND5B and

GFP-RMND5B (C358S) immunoprecipitated proteins were each associated with

similar amounts of high molecular weight proteins corresponding to ubiquitinated

proteins, indicating that the RMND5A (C356S) and RMND5B (C358S) RING domain

mutations did not markedly reduce the in vivo ubiquitination activity of the GFP-


138

Figure 4.16: In vivo ubiquitination activity of RMND5A (C356S) and RMND5B

(C358S). LNCaP cells were cotransfected with plasmids encoding HA-ubiquitin and

GFP-RMND5A, GFP-RMND5B, GFP-RMND5A (C356S), GFP-RMND5B

(C358S) or GFP (empty vector). Cells were cultured for 48 hours, with 10μM

MG132 added for the final 6 hours of culture. Following GFP immunoprecipitation

of the cell lysates, HA western blotting was performed on the immunoprecipitated

proteins, identifying low levels of ubiquitinated proteins associated with GFP, (A)

GFP-RMND5A and GFP-RMND5A (C356S), or (B) GFP-RMND5B and GFP-

RMND5B (C358S). The levels of ubiquitinated proteins were increased following

MG132 treatment of cultures and in comparison to GFP-expressing cells, the levels

of ubiquitinated proteins were evaluated in cells that overexpressed wild-type or

mutant RMND5A and RMND5B. The experiment was performed twice and

representative blots are shown (WT= wild-type).

A

B

WT (C356S)

- - - + + +

Empty Vector

10µM MG132

GFP (~70kDa)

HA (Ubiquitin)

Ubn-Proteins

(~80kDa – 175kDa)

GFP-RMND5A

- - - + + +

Empty Vector WT (C358S)

10µM MG132

HA-Ubiquitin

GFP (~70kDa)

Ubn-Proteins (~80

– 175kDa)

GFP-RMND5B


139

RMND5 proteins. Alternatively, the E3 ubiquitin ligase activity of endogenous protein

may be masking the reduced activity of the mutant proteins. Western blotting of the

total cellular inputs for HA-Ubiquitin detected similar expression in all lysates of cells

transfected with pCMV-HA-Ubiquitin, and similarly GFP western blotting of the total

cellular inputs showed that GFP, and wild-type or mutant GFP-RMND5A and GFP-

RMND5B were each expressed (not shown), indicating that the above result was not

due to markedly altered expression from the transfected plasmids.

4.2.6.6 In Vitro Auto-Ubiquitination Activity of RMND5A (C356S) and

RMND5B (C358S) RING Domains

To verify that the GST-RMND5A (C356S) and GST-RMND5B (C358S) RING

domains (Section 4.2.6.3) could be expressed in bacterial cells, each plasmid was

transformed into competent E. coli BL21 cells, glycerol stocks were prepared and cells

from each stock were streaked onto LB Agar/Ampicillin plates (Section 3.8.7).

Individual colonies were inoculated into LB Broth/Ampicillin, expression of the GST-

fusion proteins was induced by the addition of 1mM IPTG, then aliquots of the cells

were lysed and electrophoresed in 12% acrylamide gels, which were stained with

Coomassie blue (Sections 3.11.1, 3.15.2, 3.15.5). The remaining aliquots of bacterial

cells were sonicated, the soluble and insoluble fractions collected and 15µL aliquots

electrophoresed in 12% polyacrylamide gels, which were similarly stained with

Coomassie blue to determine the localisation of the mutant GST-RING domains in the

bacterial cells (Section 3.11.1, 3.15, Figure 4.17A). Only cellular proteins were present

in lysates from untreated cells, whilst a prominent band at ~28kDa was visible in

extracts of cells where expression of GST-RMND5A (C356S) RING or GST-RMND5B

(C358S) RING domains had been induced by IPTG treatment. A prominent band at

~28kDa was present in both the soluble and insoluble fractions, indicating that the GST-

RMND5A (C356S) RING and GST-RMND5B (C358S) RING proteins were packaged

into insoluble inclusion bodies but were also present in the soluble fraction in sufficient

quantities to be purified for in vitro auto-ubiquitination assays. To obtain purified GST-

RMND5A (C356S) RING and GST-RMND5B (C358S) RING domains for in vitro

ubiquitination assays, the proteins were extracted from 100mL bacterial cultures and

electrophoresed along with BSA standards in 12% polyacrylamide gels, which were

stained with Coomassie blue (Section 3.11.2, 3.15.2, 3.15.5, Figure 4.17B, C). In vitro

ubiquitination assays were also performed using bacterial proteins as the substrate, and


140

Lane 1: MW Marker

Lane 2: GST-RMND5A (C356S) RING: no IPTG

Lane 3: GST-RMND5A (C356S) RING: IPTG

Lane 4: GST-RMND5A (C356S) RING: IPTG, soluble fraction

Lane 5: GST-RMND5A (C356S) RING: IPTG, insoluble fraction

Lane 6: GST-RMND5B (C358S) RING: IPTG

Lane 7: GST-RMND5B (C358S) RING: IPTG, soluble fraction

Lane 8: GST-RMND5B (C358S) RING: IPTG, insoluble fraction

1 2 3 4 5 6 7 8

~28kDa

1 2 3 4 5 6 7 8

Lane 1: MW Marker

Lane 2: 1µg BSA

Lane 3: 2.5µg BSA

Lane 4: 5µg BSA

Lane 5: 7.5µg BSA

Lane 6: 10µg BSA

Lane 7: 1µL GST-RMND5B (C358S) RING

Lane 8: 3µL GST-RMND5B (C358S) RING

~28kDa

1 2 3 4 5 6 7 8 Lane 1: MW Marker

Lane 2: 1µg BSA

Lane 3: 2.5µg BSA

Lane 4: 5µg BSA

Lane 5: 7.5µg BSA

Lane 6: 10µg BSA

Lane 7: 1µL GST-RMND5A (C356S) RING

Lane 8: 3µL GST-RMND5A (C356S) RING

~28kDa

B

C

A


141

Figure 4.17: Expression and purification of GST-RMND5A (C356S) RING

and GST-RMND5B (C358S) RING domains. E. coli BL21 cells transformed

with pGEX-RMND5A (C356S) RING or pGEX-RMND5B (C358S) RING were

induced with IPTG to produce GST-RMND5A (C356S) RING or GST-RMND5B

(C358S) RING domains (~28kDa). (A) The bacterial cells were separated into

soluble and insoluble fractions and a 15µL aliquot of each sample was

electrophoresed in a 12% polyacrylamide gel followed by Coomassie blue

staining to visualise the compartmentalisation of the induced GST fusion proteins.

(B) GST-RMND5A (C356S) RING and (C) GST-RMND5B (C358S) RING

domains were purified from the bacterial cells, an aliquot electrophoresed in

polyacrylamide gels and stained with Coomassie blue, from which the

concentration of the protein was estimated in comparison to BSA standards. The

concentrations of the GST-RMND5A (C356S) RING and the GST-RMND5B

(C358S) RING domain was estimated to be ~2.3µg/µL and ~2.8µg/µL,

respectively. (D) Proteins were extracted from E. coli BL21 cells and 1µL, 5µL

and 8µL aliquots were electrophoresed in a 12% polyacrylamide gel to visualise

the bacterial proteins.

1 2 3 4

Lane 1: MW Marker

Lane 2: 1µL extracted bacterial proteins



D


142

to generate proteins for these assays, total proteins were isolated from E. coli BL21

bacterial cells (as bacteria do not ubiquitinate proteins) (Section 3.11.3). Electrophoresis

and Coomassie blue staining of the bacterial proteins indicated that a range of low-high

molecular weight proteins were present, which was suitable for these analyses (Sections

3.15.2, 3.15.5, Figure 4.17D).

In vitro ubiquitination assays using the E2 enzyme UbcH5b were carried out to compare

the activity of wild-type GST-RMND5A and GST-RMND5B RING domains to that of

the GST-RMND5A (C356S) and GST-RMND5B (C358S) RING domains. In vitro

ubiquitination reactions were carried out at 37°C for 1 hour and the reaction products

were electrophoresed in 12% polyacrylamide gels followed by western blotting for

biotinylated ubiquitin (Section 3.14.1, 3.15). The negative control reaction lacking ATP

yielded a single protein band at ~10kDa corresponding to free ubiquitin, whilst the

negative control reactions lacking GST-RING domains (E3) yielded protein bands

corresponding to ubiquitinated E2 enzyme, as observed previously (Section 4.2.4.3).

Biotinylated-ubiquitin western blotting of reactions containing wild-type GST-RMND5

RING domains yielded prominent protein bands at >100kDa corresponding to

polyubiquitinated proteins, and the appearance or intensity of the protein bands in

reactions containing wild-type GST-RMND5A RING and GST-RMND5A (C356S)

RING domain proteins was similar (Figure 4.18). These results indicated that the

RMND5A (C356S) RING domain mutation did not reduce RMND5A RING domain E3

ubiquitin ligase activity. Biotinylated ubiquitin western blotting of in vitro

ubiquitination assays using the GST-RMND5B (C358S) RING domain identified a

small reduction in the intensity of prominent high molecular weight protein bands at

>100kDa, corresponding to polyubiquitinated proteins in comparison to reactions

containing GST-RMND5B RING domain, indicating a reduction in the E3 ubiquitin

ligase activity of the RMND5B (C358S) RING domain (Figure 4.18). In vitro

ubiquitination assays using the GST-RMND5A RING, GST-RMND5A (C356S) RING,

GST-RMND5B RING or GST-RMND5B (C358S) RING domains were repeated with

the addition of 2µL extracted bacterial proteins as substrates (Section 3.14.1, 3.15). In

these assays, the presence of protein bands ranging from 7kDa to >100kDa in western

blots for biotinylated ubiquitin, indicated that all RING domains were able to

ubiquitinate bacterial proteins (Figure 4.18). Evidence of a small reduction in protein

banding present in the reactions using GST-RMND5B (C358S) RING domain


143

compared to reactions containing the GST-RMND5B RING domain supported initial

findings that mutation of the RMND5B RING domain had reduced its ubiquitination

activity. However, the appearance and intensity of protein bands corresponding to

ubiquitinated proteins were similar in reactions containing GST-RMND5A RING or

GST-RMND5A (C356S) RING domains indicating that the mutation had not altered the

ubiquitination activity of the RMND5A RING domain (Figure 4.18).

To determine whether the GST-RMND5A (C356S) RING domain exhibited differences

in the rate at which it was able to mediate ubiquitin transfer compared to the wild-type

GSTRMND5A RING domain, in vitro ubiquitination assays were repeated at a reaction

temperature of 30°C (rather than 37°C), and with aliquots of each reaction taken at 10,

30, 60 and 90 minutes (Section 3.14.1, Figure 4.18). Biotinylated ubiquitin western

blotting of reaction products from the in vitro ubiquitination assays identified high

molecular weight protein bands corresponding to polyubiquitinated proteins at >100kDa

of a similar intensity in reactions containing either GST-RMND5A RING or GST-

RMND5A (C356S) under all conditions tested, indicating no alterations in the

ubiquitination activity of the mutant RMND5A RING domain. The only notable

difference in in vitro ubiquitination activity of the GST-RMND5A (C356S) RING

domain was evident in the appearance of a band at ~38kDa corresponding in size to the

monoubiquitinated GST-RING domain (Figure 4.18). This band appeared more intense

in western blots of products from reactions performed using the GST-RMND5A

(C356S) RING domain, suggesting that the mutant RMND5A RING domain favoured

automonoubiquitination in comparison to the wild-type RMND5A RING domain under

these conditions. The intensity of this band was similar for both GST-RMND5A RING

and GST-RMND5A (C356S) RING domains when reactions were carried out at 37°C

(Figure 4.18).

4.2.7 Examination of the E3 Ubiquitin Ligase Activity of the RMND5A and

RMND5B RING Domains by the Introduction of Dual Mutations in the

RMND5 RING Domains

As introduction of a single amino acid change into the RING domains of RMND5A and

RMND5B did not cause a marked reduction in E3 ubiquitin ligase activity, substitution

of two amino acids in each of the RING domains was introduced by site-directed

mutagenesis, generating RMND5A (C356A/H358A) and RMND5B (C358A/H360A)


144

WT WTC356S C358S WT WTC356S C358S- - - - + + + + Bacterial Protein Substrate

Ubn Proteins (~8 - >100kDa)

10 10 30 60 90 30 60 90 minutes

Ubn Proteins (~8kDa ->100kDa)

RING-Ub (~38kDa)

Figure 4.18: In vitro ubiquitination activity of GST-RMND5A (C356S) and GST-RMND5B (C358S) RING domains. (A) GST-RMND5A RING, GST-RMND5B RING, GST-RMND5A (C356S) RING or GST-RMND5B (C358S) RING were used in in vitro ubiquitination assays with UbcH5b. Reactions were incubated at 37°C for 60 minutes and 10μL of each product was electrophoresed in 4-12% gradient gels. Western blotting for biotinylated ubiquitin identified high molecular weight bands in reactions containing both wild-type and mutant GST-RMND5 RING domains (left panel). In reactions containing bacterial proteins (right panel) and either wild-type GST-RMND5A RING or mutant GST-RMND5A (C356S) RING, similar levels of ubiquitinated proteins spanning the blot were evident. In vitro ubiquitination assays containing GST-RMND5B (C358S) RING exhibited a slight reduction in banding corresponding to ubiquitinated proteins compared to reactions using wild-type GST-RMND5B RING. (B) Wild-type GST-RMND5A or GST-RMND5A (C356A) RING domains were used in in vitro ubiquitination assays with UbcH5b and the reactions were incubated at 30°C for 10 - 90 minutes. Ten μL of each product was electrophoresed in 4-12% gradient gels followed by western blotting for biotinylated ubiquitin, identifying high molecular weight protein bands of a similar intensity spanning the blot in reactions containing both wild-type GST-RMND5A and GST-RMND5A (C356S) RING domains. Experiments were performed twice and representative blots are shown (WT= wild-type).

A

B B


145

(Section 3.4.3, Figure 4.19). The two amino acids were selected for mutation as they are

the third and fourth zinc coordinating residues in the RING domain and mutation of the

homologous residues in other E3 ubiquitin ligases has been reported to reduce their

activity (Zhang et al., 2009). In addition, in RMND5 proteins these two residues are the

canonical RING domain zinc coordinating residues (cysteine and histidine) as opposed

to the first two and fifth and sixth zinc coordinating residues which are not canonical

zinc binding amino acids (Figure 4.19) (Zhang et al., 2009).

4.2.7.1 Mutation of the RMND5A (C356A/H358A) RING Domain

PCR primers were designed to introduce three base changes into the RMND5A RING

domain using RMND5A (C356S) as a template, however although multiple PCR

conditions were utilised to optimise the reaction including a range of annealing

temperatures between 50°C - 72°C, 1.5 – 3mM MgCl2 and high fidelity and GC buffers,

no PCR products were amplified (Section 3.4.3) (not shown). The mutagenesis PCR

primers were therefore redesigned to introduce the RMND5A (C356A/H358A)

mutations in two consecutive PCRs with the C356A mutation introduced via a single

base change in the first round of PCR (RMND5A Primer Set 1) and once confirmed, the

two additional base changes required to introduce the H358A mutation were introduced

using the pEGFP-RMND5A (C356A) plasmid DNA as a template (RMND5A Primer

Set 2). Initial PCRs to introduce the RMND5A (C356A) mutation were carried out using

a gradient of annealing temperatures between 58°C - 66°C for 26 cycles, which

produced a faint band at the expected size of ~5.9kb at all annealing temperatures tested

(Section 3.4.3, 3.6, Figure 4.20A). Amplified products from PCRs performed at

annealing temperatures of 64°C and 66°C were treated with DpnI to digest the parental

methylated plasmid DNA and the reaction products were transformed into E. coli DH5α

then selected by plating on LB Agar/Kanamycin (Sections 3.8.4, 3.8.6). Four of the

colonies were inoculated into LB Broth/Kanamycin for small scale plasmid purification,

the plasmids RNase treated, purified and 5µL electrophoresed in a 1% agarose gel

which resulted in the identification of ~5.9kb bands corresponding in size to pEGFP-

RMND5A (C356A) (Section 3.6, 3.9, not shown). Two µL of each of pEGFP-

RMND5A (C356A) clones 1-4 were sequenced using the RMND5A1176-AS primer,

and BLASTTM analysis of the sequences verified the presence of the C356A mutation in

all clones (Section 3.12, Appendix II, not shown).


146

Figure 4.19: Site directed mutagenesis of the RING domains of RMND5A and RMND5B to produce RMND5A (C356A/H358A) RING and RMND5B (C358A/H360A) RING. Alignment of the RING domains of RMND5A, RMND5B and yeast RMD5 showing all eight amino acid residues required for RING domain folding and function (red/orange) including the conserved cysteine residues (bold, orange) to be mutated to alanine (blue) to reduce or inactivate the RING domain activity of RMND5A and RMND5B. Numbers indicate amino acid number.




RMND5A (C356A/H358A)

RMND5B (C358A/H360A)

RMD5 (C379S)

A A

336 377

338 379

361 404


147

The pEGFP-RMND5A (C356A) plasmid was then used in site-directed mutagenesis

reactions to incorporate the remaining two base changes in the RMND5A RING domain.

These PCRs were carried out using a gradient of annealing temperatures between 64°C

and 72°C for 26 cycles, and electrophoresis of the amplified product identified a faint

band at the expected size of ~5.9kb with more intense bands present in reactions

performed with the lower annealing temperatures of 64°C and 66.4°C (Section 3.4.3,

3.6, Figure 4.20B). The PCR products from reactions with the annealing temperatures

of 64°C and 66.4°C were digested with DpnI then transformed into competent E. coli

DH5α and selected on LB/Kanamycin plates (Section 3.4.3, 3.8.6). Four colonies were

picked and cultured in LB Broth/Kanamycin for small scale plasmid purification, the

plasmids were treated with RNase, purified and 5µL of each of pEGFP-RMND5A

(C356A/H358A) clones 1-4 were electrophoresed in a 1% agarose gel (Section 3.6, 3.9,

Figure 4.20C). Based on the gel, 2µL of each was used in sequencing reactions with the

RMND5A1176-AS primer, and BLASTTM analysis of the chromatograms verified the

presence of the RMND5A (C356A/H358A) mutations in all clones (Section 3.12,

Appendix II, not shown). The entire coding region of pEGFP-RMND5A

(C356A/H358A) clones 1 and 2 was sequenced using the RMND5A603-S, RMND5A1-

S and RMND5A490-AS primers, verifying that no additional mutations had been

incorporated in the site-directed mutagenesis procedure (Appendix II, not shown).

To ensure that no further mutations were present in the pEGFP-C2 vector in which the

RMND5A (C356A/H358A) RING mutations were induced, the RMND5A coding

region was excised from the pEGFP-RMND5A (C356A/H358A) plasmid by digestion

with EcoRI, gel purified and the products electrophoresed in a 1% agarose gel (Sections

3.6, 3.7.2, 3.8.2, Figure 4.20D). Fresh pEGFP-C2 vector was prepared by digestion with

EcoRI, SAP treatment, purification then electrophoresis of the purified products in a 1%

agarose gel (Section 3.6, 3.8.2, 3.8.3, Figure 4.20E). Based on these gels, 60ng (2µL)

RMND5A (C356A/H358A) insert was ligated into 50ng (0.5µL) pEGFP-C2 and the

products were transformed into competent E. coli DH5α cells (Sections 3.8.4, 3.8.6).

Transformed bacteria were selected by growth on LB Agar/Kanamycin plates, three

colonies were picked, cultured in LB Broth/Kanamycin for small scale plasmid

purification, and the plasmids were RNase treated and digested with EcoRI to release

the RMND5A (C356A/H358A) insert (Sections 3.8.2, 3.9). Electrophoresis of the

digested plasmids identified a band at the expected size of ~1.2kb in pEGFP-RMND5A

(C356A/H358A) clones 1 and 2, these clones were purified then 5µL of each was


148

Lane 1: MW Marker Lane 2: Undigested pEGFP-C2 Lane 3: EcoRI and SAP digested pEGFP-C2

~4.7kb

Lane 1: MW Marker Lane 2-6: pEGFP-RMND5A (C356A/H358A) clone 1-4

Lane 1: MW Marker Lane 2: RMND5A (C356A/H358A)

~1.2kb

D E

Lane 1: MW Marker Lane 2: 66ºC Lane 3: 64ºC Lane 4: 60ºC Lane 5: 58ºC Lane 6: Negative Control

~5.9kb

A

B ~5.9kb

~5.9kb

C

Lane 1: MW Marker Lane 2: 70°C Lane 3: 69.5°C Lane 4: 68.4°C Lane 5: 66.4°C Lane 6: 64°C Lane 7: Negative Control


149

Figure 4.20: Generation of RMND5A (C356A/H358A) by site-directed mutagenesis and cloning of pEGFP-RMND5A (C356A/H358A). (A) pEGFP-RMND5A (C356S) and an annealing temperature gradient ranging from 58°C to 66°C were used to optimise RMND5A (C356A) site-directed mutagenesis PCR conditions. PCR products were digested with DpnI and 10µL of each digested product was electrophoresed in a 1% agarose gel, identifying the ~5.9kb mutagenesis products. (B) An annealing temperature gradient from 64°C to 70°C and the pEGFP-RMND5A (C356A) template were used to optimise RMND5A (C356A/H358A) site directed mutagenesis, and 10µL of each of the DpnI digested PCR products were electrophoresed in a 1% agarose gel. The mutagenesis products were identified at ~5.9kb. (C) 5µL of each of purified pEGFP-RMND5A (C356A/H358A) clones 1 to 4 was electrophoresed in a 1% agarose gel. (D) pEGFP-RMND5A (C356A/H358A) was EcoRI digested to release the RMND5A (C356A/H358A) coding region which was gel purified. 5µL purified product was electrophoresed in a 1% agarose gel from which the concentration of the ~1.2kb insert was estimated to be 30ng/µL. (E) 2µL of purified EcoRI/SAP digested pEGFP-C2 plasmid was electrophoresed in a 1% agarose gel from which the concentration of DNA was estimated at ~100ng/µL. (F) Plasmids isolated from pEGFP-RMND5A (C356A/H358A) clones 1 to 3 were EcoRI digested to liberate the ~1.2kb insert and the products were electrophoresed in a 1% agarose gel. Clones 1 and 2 contained the RMND5A (C356A/H358A) insert. (G) pEGFP-RMND5A (C356A/H358A) clones 1 and 2 were purified and 5µL of each purified plasmid was electrophoresed in a 1% agarose gel.

Lane 1: MW Marker Lane 2: Undigested pEGFP-RMND5A (C356A/H358A) Lane 3-5: EcoRI digested pEGFP-RMND5A (C356A/H358A) clones 1-3

~1.2kb

~4.7kb

Lane 1: MW Marker Lane 2: Purified pEGFP-RMND5A (C356A/H358A) clone 1 Lane 3-5: Purified pEGFP-RMND5A (C356A/H358A) clones 2

F

G

~5.9kb


150

electrophoresed in a 1% agarose gel in preparation for sequencing reactions (Sections

3.6, 3.7.1, Figure 4.20E, F). Based on the gel, 1µL of each clone was used for

sequencing reactions with the RMND5A1-S, RMND5A1176-AS, RMND5A603-S and

RMND5A490-AS primers, BLASTTM analysis of which indicated that pEGFP-

RMND5A (C356A/H358A) clone 2 contained no additional mutations (aside from the

T1060G/G1061C/C1066G/A1067C base changes corresponding to the C356A/H358A

amino acids changes) and was inserted in the sense orientation (Appendix II, Appendix

III).

4.2.7.2 Cloning of the RMND5A (C356A/H358A) RING Domain into pGEX-

2TK

Following production of RMND5A (C356A/H358A) (Section 4.2.7.1), the mutant

RMND5A RING domain was PCR amplified in quadruplicate using

RMND5ARING1006-S and RMND5ARING1131-AS primers and the previously

optimised PCR conditions, the products were purified and 5µL was electrophoresed in a

2% agarose gel (Section 3.4.3, 3.6, 4.2.3.1, Figure 4.21A, Appendix II). Based on the

gel, 100ng (2µL) of the RMND5A (C356A/H358A) RING domain PCR product was

ligated into 50ng (1µL) pGEM®T-Easy cloning vector and the products transformed

into competent E. coli DH5α cells and selected by growth on LB Agar/Ampicillin plates

(Section 3.8.4). Four colonies were inoculated into LB Broth/Ampicillin for small scale

plasmid purification, the purified plasmids were RNase treated, digested with BamHI

and EcoRI to release the insert, and 10µL of each product was electrophoresed in a 2%

agarose gel (Section 3.6, 3.8.2, 3.9, Figure 4.21B). A band of the expected molecular

weight of ~126bp was present in pGEMT-RMND5A (C356A/H358A) RING clones 1,

3 and 4, clones 1 and 3 were purified and 5µL of each was electrophoresed in a 1%

agarose gel in preparation for sequencing (Section 3.7.1, not shown). Based on the gel,

3µL of each of clones 1 and 2 were sequenced using M13-S and M13-AS primers, and

the presence of the C356A and H358A mutations in pGEMT-RMND5A

(C356A/H358A) clone 1 were verified using BLASTTM analysis (Section 3.12).

To prepare the RMND5A (C356A/H358A) RING domain for ligation into pGEX-2TK

to generate pGEX-RMND5A (C356A/H358A) RING, the pGEMT-RMND5A

(C356A/H358A) RING clone 1 plasmid was digested with BamHI and EcoRI to release

the ~126bp insert, the product gel purified and 5µL of the purified product

Chapter 4 Characterisation of RMND5 E3 ubiquitin ligase activity

151

Figure 4.21: Cloning of sequences encoding the RMND5A (C356A/H358A) RING

domain into pGEX-2TK. (A) The RING domain sequence of pEGFP-RMND5A

(C356A/H358A) was PCR amplified and 5µL purified PCR product was

electrophoresed in a 2% agarose gel from which the DNA concentration was

estimated to be 50ng/µL. (B) Plasmids isolated from pGEMT-RMND5A

(C356A/H358A) RING clones 1 to 4 were BamHI/EcoRI digested to release the

~126bp insert and 10µL of each product was electrophoresed in a 2% agarose gel. (C)

pGEMT-RMND5A (C356A/H358A) RING was digested with BamHI and EcoRI, the

insert was gel purified and 5µL purified product was electrophoresed in a 2% agarose

gel, from which the concentration was estimated to be ~5ng/µL. (D) 5µL of purified

BamHI/EcoRI and SAP digested pGEX-2TK plasmid was electrophoresed in a 1%

agarose gel from which the concentration was estimated to be ~10ng/µL.

B

Lane 1: MW Marker

Lane 2: RMND5A (C356A/H358A) RING

~126bp

Lane 1: RMND5A (C356A/H358A)

RING

Lane 2: Negative Control

Lane 3: MW Marker

~126bp

~3kb

Lane 1: MW Marker

Lane 2: Undigested pGEMT-RMND5A

(C356A/H358A) RING

Lane 3-6: BamHI/EcoRI digested

pGEMT-RMND5A (C356A/H358A)

RING clones 1-4

A

~126bp

p

C

D Lane 1: MW Marker

Lane 2: Undigested pGEX-2TK

Lane 3: BamHI/EcoRI/SAP treated pGEX-2TK ~1.2kb


152

electrophoresed in a 2% agarose gel (Section 3.6, 3.8.2, Figure 4.21C). The pGEX-2TK

vector was prepared by small scale plasmid purification, the plasmid RNase treated,

BamHI and EcoRI digested and SAP treated, then purified and 5µL of the product was

electrophoresed in a 1% agarose gel (Section 3.6, 3.8.2, 3.8.3, 3.9, Figure 4.21D). Based

on the gel, 40ng (8µL) RMND5A (C356A/H358A) RING domain was ligated into 50ng

(5µL) pGEX-2TK and the products were transformed into competent E. coli DH5α cells

then selected by plating on LB Agar/Ampicillin (Section 3.8.4, 3.8.6). Four colonies of

pGEX-RMND5A (C356A/H358A) RING were picked and cultured in LB

Broth/Ampicillin for small scale purification. The purified plasmids were RNase treated

then digested with BamHI and EcoRI to liberate the ~126bp RMND5A (C356A/H358A)

RING domain and 10µL of each product was electrophoresed in a 2% agarose gel

(Section 3.6, 3.8.2, 3.9, not shown). Although no insert bands were visible on the gel

(potentially due to their small size of ~126bp), pGEX-RMND5A (C356A/H358A)

RING clones 1 and 3 were purified and 3µL of each of clone 1 and 3 was sequenced

using the pGEX-S and pGEX-AS primers. BLASTTM analysis of the sequencing

products indicated that pGEX-RMND5A (C356A/H358A) RING clone 3 contained no

additional mutations, apart from those encoding the C356A/H358A amino acid changes

(Section 3.7.1, 3.12, Appendix II, not shown). This plasmid was then transformed into

competent E. coli BL21 cells and a glycerol stock was prepared (Section 3.8.7).

4.2.7.3 Mutation of the RMND5B (C358A/H360A) RING Domain

To introduce the C358A and H360A mutations into the RMND5B RING domain, PCR

primers were designed to introduce the three base changes required, with the pEGFP-

RMND5B (C358S) plasmid to be used as the DNA template. Initial PCRs were carried

out using an annealing temperature of 65°C and a range of MgCl2 concentrations from

1.5 – 3mM MgCl2 for 26 cycles, producing a faint band at the expected size of ~5.9kb

in reactions containing 2mM, 2.5mM and 3mM MgCl2 when the PCR products were

electrophoresed in a 1% agarose gel (Sections 3.4.3, Figure 4.22A). The products from

reactions using 2mM and 3mM MgCl2 were treated with DpnI to digest the parental

methylated plasmid DNA, purified, then the digest products were transformed into E.

coli DH5α and selected by plating on LB Agar/Kanamycin (Section 3.8.6). Four

resulting colonies were picked, cultured in LB Broth/Kanamycin for small scale

plasmid purification, the plasmids treated with RNase and purified, with electrophoresis

of the products in a 1% agarose gel identifying bands corresponding to the ~5.9kb

pEGFP-RMND5B (C358A/H360A) (Section 3.6, 3.9, not shown). Based on the gel,


153

1µL of each of pEGFP-RMND5B (C358A/H360A) clones 1-4 were sequenced

reactions using the RMND5B790-S primer, which verified the presence of the

C358A/H360A mutations in all clones (Section 3.12). The entire coding region of

pEGFP-RMND5B (C358A/H360A) clones 1 and 2 was sequenced using the

RMND5BSalI1-S, RMND5BSalI1182-AS and pEGFP1266-S primers, and BLASTTM

analysis of the chromatograms confirmed that both inserts contained no additional

mutations (Appendix II, not shown).

To subclone the RMND5B (C358A/H360A) insert into pEGFP-C2, the RMND5B

coding region was excised from the plasmids by digestion with SalI, gel purified and

electrophoresed in a 1% agarose gel (Sections 3.6, 3.7.2, 3.8.2, Figure 4.22B). The

pEGFP-C2 vector was prepared by SalI digestion, SAP treatment, purification then

electrophoresis of the product in a 1% agarose gel (Section 3.7.1, 3.8.2, 3.8.3, Figure

4.22C). Based on the gels, 35ng (7µL) RMND5B (C358A/H360A) insert was ligated

into 50ng (0.5µL) pEGFP-C2 and the products were transformed into competent E. coli

DH5α cells (Section 3.8.4). Three colonies were inoculated into LB Broth/Kanamycin

for small scale plasmid purification, the plasmids RNase treated and digested with SalI

to release the insert then electrophoresed in a 1% agarose gel (Section 3.6, 3.8.2, 3.9,

Figure 4.22D). A band at the expected size of ~1.2kb was present in clone 1, and this

plasmid was purified and 5µL product electrophoresed in a 1% agarose gel (Sections

3.6, 3.7.1, Figure 4.22E). Based on the gel, 1µL of clone 1 was sequenced using the

RMND5BSalI1-S, RMND5BSalI1182-AS and pEGFP1266-S primers and BLASTTM

analysis verified that pEGFP-RMND5B (C358A/H360A) clone 1 contained no

additional mutations (aside from the T1066G/G1067/C1072G/A1073C base changes

corresponding to the C358A/H360A amino acid changes) and was in sense orientation,

therefore glycerol stocks were prepared of this clone (Section 3.8.7, 3.12, Appendix II,

Appendix III).

4.2.7.4 Cloning of the RMND5B (C358A/H360A) RING Domain into pGEX-

2TK

Following the production of pEGFP-RMND5B (C358A/H360A) (Section 4.2.7.3), the

RMND5B (C358A/H360A) RING domain was PCR amplified in quadruplicate using

the RMND5BRING1012-S and RMND5BRING1137-AS primers, the reactions were


154

Figure 4.22: Generation of RMND5B (C358A/H360A) by site-directed mutagenesis. (A) pEGFP-RMND5B (C358S) template, an annealing temperature of 65°C and 1.5mM-3mM MgCl2 were used to optimise RMND5B (C358A/H360A) site-directed mutagenesis PCR conditions. PCR products were DpnI digested and 10µL of each product was electrophoresed in a 1% gel to visualise the ~5.9kb mutagenesis products. (B) pEGFP-RMND5B (C358A/H360A) was SalI digested, releasing the RMND5B (C358A/H360A) insert. 5µL gel purified insert was electrophoresed in a 1% agarose gel from which the concentration was estimated to be ~5ng/µL. (C) 2µL purified SalI and SAP digested pEGFP-C2 was electrophoresed in a 1% agarose gel, from which the DNA concentration was estimated to be ~100ng/µL. (D) pEGFP-RMND5B (C358A/H360A) clones 1 to 3 were digested with SalI to liberate the ~1.2kb insert then electrophoresed in a 1% agarose gel. Clone 2 contained an insert. (E) 5µL purified pEGFP-RMND5B (C358A/H360A) clone 2 was electrophoresed in a 1% agarose gel.

Lane 1: MW Marker Lane 2: 1.5mM MgCl2 Lane 3: 2mM MgCl2 Lane 4: 2.5mM MgCl2 Lane 5: 3mM MgCl2 Lane 6: Negative Control

~5.9kb

A

Lane 1: MW Marker Lane 2: Purified pEGFP-RMND5B (C358A/H360A) clone 1 ~5.9kb

E

Lane 1: MW Marker Lane 2: Undigested pEGFP-C2 Lane 3: SalI digested pEGFP-C2

~4.7kb

C

Lane 1: MW Marker Lane 2: RMND5B (C358A/H360A)

~1.2kb

B

D

~1.2kb

~4.7kb

Lane 1: MW Marker Lane 2: Undigested pEGFP-RMND5B (C358A/H360A) Lane 3-5: SalI digested pEGFP-RMND5B (C358A/H360A) clones 1-3


155

combined and purified and 5µL of the purified product was electrophoresed in a 2%

agarose gel (Section 3.4, 3.6, 3.7.1, Figure 4.23A). Based on the gels, 60ng (4µL)

RMND5B (C358A/H360A) was ligated with 50ng (1µL) pGEM®T-Easy and the

products were transformed into competent E. coli DH5α then selected by growth on LB

Agar/Ampicillin plates (Section 3.8.4, 3.8.6). Four colonies of pGEMT-RMND5B

(C358A/H360A) RING were inoculated into LB Broth/Ampicillin and incubated

overnight prior to small scale plasmid purification, the plasmids were RNase treated,

BamHI and EcoRI digested to release the insert and electrophoresed in a 2% agarose gel

(Section 3.6, 3.8.2, Figure 4.23B). A band of ~126bp was present in all clones, pGEMT-

RMND5B (C358A/H360A) RING clones 1-4 were purified and 5µL of each product

was electrophoresed in a 1% agarose gel (Section 3.6, 3.7.1, not shown). Based on the

gel, 3µL of each of pGEMT-RMND5B (C358A/H360A) RING clones 1 and 2 were

sequenced using the M13-S and M13-AS primers (Section 3.12, Appendix II).

BLASTTM analysis verified that RMND5B (C358A/H360A) RING clone 1 contained no

additional mutations (not shown).

To subclone the RMND5B (C358A/H360A) RING into pGEX-2TK, pGEMT-RMND5B

(C358A/H360A) RING clone 1 was digested with BamHI and EcoRI, the ~126bp insert

gel purified and 5µL of the product electrophoresed in a 2% agarose gel (Section 3.8.2,

Figure 4.23C). The pGEX-2TK vector was prepared by small scale plasmid

purification, RNase treatment, BamHI and EcoRI digestion and SAP treatment, then

5µL of the purified product was electrophoresed in a 1% agarose gel (Section 3.6, 3.8.2,

3.8.3, 3.9, Figure 4.23D). Based on the gels, 40ng (8µL) RMND5B (C358A/H360A)

RING and 50ng (5µL) pGEX-2TK were ligated and the products transformed into

competent E. coli DH5α then selected by plating on LB Agar/Ampicillin (Section 3.8.4,

3.6). Four colonies of pGEX-RMND5B C358A/H360A RING were picked and cultured

overnight in LB Broth/Ampicillin for small scale purification. Plasmids were RNase

treated, digested with BamHI and EcoRI to liberate the ~126bp RING domain and 10µL

of each product was electrophoresed in a 2% agarose gel (Section 3.6, 3.8.2). Although

no insert bands were visible due to the small size of the products (not shown), pGEX-

RMND5B (C358A/H360A) RING clones 1 and 2 were purified and 2µL of each clone

was sequenced using the pGEX-S and pGEX-AS primers, BLASTTM analysis of which

confirmed that pGEX-RMND5B (C358A/H360A) RING clone 1 was mutation free

apart from the base substitutions encoding the C358A/H360A amino acid changes


156

Figure 4.23: Cloning of sequences encoding the RMND5B (C358A/H360A) RING

domain into pGEX-2TK. (A) 5µL purified PCR amplified RMND5B

(C358A/H360A) was electrophoresed in a 2% agarose gel and the DNA concentration

was estimated to be ~15ng/µL. (B) pGEMT-RMND5B (C358A/H360A) RING

clones 1 to 4 were BamHI and EcoRI digested to release the ~126bp insert and the

products were electrophoresed in a 2% agarose gel, identifying inserts in all clones.

(C) 5µL gel purified BamHI and EcoRI digested pGEMT-RMND5B

(C358A/H360A) RING was electrophoresed in a 2% agarose gel from which the

DNA concentration was estimated to be ~5ng/µL. (D) BamHI/EcoRI and SAP

digested pGEX-2TK plasmid was purified and 5µL purified product was

electrophoresed in a 1% agarose gel from which the concentration was estimated to

be ~10ng/µL.

Lane 1: MW Marker

Lane 2: RMND5B RING (C358A/H360A)

Lane 3: Negative Control

~126bp

Lane 1: MW Marker

Lane 2: Undigested pGEMT-RMND5B (C358A/H360A) RING

Lane 3-6: BamHI/EcoRI digested pGEMT-RMND5B (C358A/H360A)

RING clones 1-4

~126bp

~3kb

A

B

~5kb

Lane 1: MW Marker

Lane 2: Undigested pGEX-2TK

Lane 3: BamHI/EcoRI SAP

digested pGEX-2TK

D

Lane 1: MW Marker

Lane 2: RMND5B

(C358A/H360A) RING

C

~126bp


157

(Section 3.12, Appendix II). This plasmid was then transformed into E. coli BL21 cells

and a glycerol stock was prepared (Section 3.8.7).


(C356A/H358A) and RMND5B (C358A/H360A)

To examine the expression and intracellular localisation of the mutant RMND5

proteins, large scale purification of the pEGFP-RMND5A (C356A/H358A) and

pEGFP-RMND5B (C358A/H360A) plasmids was performed and the concentration of

the purified plasmid was determined to be 1.2µg/µL for pEGFP-RMND5A

(C356A/H358A) and 1.3µg/µL for pEGFP-RMND5B (C358A/H360A) (Section 3.10,

4.2.7.1, 4.2.7.3). Expression of pEGFP-RMND5A (C356A/H358A) and pEGFP-

RMND5B (C358A/H360A) was evaluated by western blotting following transient

transfection of LNCaP cells with 4µg plasmids encoding each of wild-type or mutant

GFP-RMND5 proteins (Section 3.1.4, 3.15, Figure 4.24A). GFP western blotting

identified a prominent band at ~70kDa in the lysates of all transfected cells, indicating

expression of each of GFP-RMND5A, GFP-RMND5B, GFP-RMND5A

(C356A/H358A) and GFP-RMND5B (C358A/H360A) (Figure 2.24A).

To determine the cellular localisation of GFP-RMND5A (C356A/H358A) and GFP-

RMND5B (C358A/H360A), LNCaP cells growing on coverslips were transfected with

2µg plasmid encoding either wild-type or mutant GFP-RMND5 proteins (Section 3.1.3,

3.1.4). The cells were fixed 48 hours post transfection and viewed by fluorescence

microscopy, which identified that GFP-RMND5A and GFP-RMND5A

(C356A/H358A) proteins were diffusely distributed in the nucleus and cytoplasm, with

some cells exhibiting a punctate cytoplasmic distribution (Section 3.16, Figure 4.24B).

The GFP-RMND5B and GFP-RMND5B (C358A/H360A) proteins also appeared to be

diffusely distributed in the nucleus and cytoplasm, with most cells also containing

punctate cytoplasmic granules with GFP fluorescence (Figure 4.24B). These results

indicated that the RMND5 RING domain mutations did not alter the cellular

localisation of the RMND5 proteins in comparison to that of the wild-type protein.


158

A

B GFP Nucleus Cytoplasm Overlay

GFP-RMND5A

Wild Type

C356A/ H358A

GFP-RMND5B

Wild Type

C358A/ H360A

Untransfected Negative Control


159

Figure 4.24: Expression and cellular localisation of GFP-RMND5A (C356A/H358A) and GFP-RMND5B (C358A/H360A). LNCaP cells were transfected with plasmids encoding GFP-RMND5A, GFP-RMND5B, GFP-RMND5A (C356A/H358A) or GFP-RMND5B (C358A/H360A). At 48 hours post-transfection, (A) cells were harvested for GFP western blotting, which identified ~70kDa bands corresponding to GFP-RMND5A, GFP-RMND5A (C356A/H358A), GFP-RMND5B or GFP-RMND5B (C358A/H360A). (B) Alternatively, the cells were prepared for fluorescence microscopy and stained with Hœchst 33258 and tetramethylisothiocyanate (TRITC)-Phalloidin to visualise the nucleus and cytoplasm, respectively. GFP-RMND5A, GFP-RMND5B, GFP-RMND5A (C356A/H358A) and GFP-RMND5B (C358A/H360A) displayed diffuse nuclear and cytoplasmic localisation, with GFP-RMND5B and GFP-RMND5B (C358A/H360A) also exhibiting a punctate distribution in the cytoplasm (Magnification x1000). Experiments were preformed twice and representative results are shown. (WT= wild type).


160

4.2.7.6 In Vivo Ubiquitination Activity of RMND5A (C356A/H358A) and

RMND5B (C358A/C360A)

The activity of RMND5A (C356A/H358A) and RMND5B (C358A/H360A) was

assessed using in vivo ubiquitination assays performed following cotransfection of

LNCaP cells plasmids encoding HA-ubiquitin and either GFP-RMND5A, GFP-

RMND5A (C356A/H358A), GFP-RMND5B or GFP-RMND5B (C358A/H360A). To

promote the accumulation of ubiquitinated proteins, cells were incubated with 10µM

MG132 for the final 2 hours prior to harvest at 48 hours following transfection (Section

3.1.4, 3.1.5, 3.14.2). The shorter time period of proteasome inhibition was used to

facilitate identification of differences in the rate at which the wild-type and mutant

RMND5 proteins were able to associate with ubiquitinated proteins. GFP-tagged

proteins were immunoprecipitated from the cell lysates and electrophoresed in 4-12%

gradient polyacrylamide gels, with HA western blotting of GFP-RMND5A, GFP-

RMND5A (C356A/H358A) and empty vector immunoprecipitates identifying protein

bands corresponding to polyubiquitinated proteins at a range of molecular weights

(~80kDa – 175kDa) (Section 3.15, Figures 4.25A). The levels of ubiquitinated proteins

were markedly increased in the immunoprecipitates of GFP-RMND5A and GFP-

RMND5A (C356A/H358A) expressing cells that had been MG132 treated (Figure

4.25A). GFP-RMND5A and GFP-RMND5A (C356A/H358A) were associated with

similar amounts of ubiquitinated proteins, indicating that either the RMND5A

(C356A/H358A) mutation did not reduce the E3 ubiquitin ligase activity of RMND5A

(C356A/H358A) or that endogenous active proteins were masking alterations in the

activity of the mutant protein.

Similar in vivo ubiquitination assays were performed using GFP-RMND5B and GFP-

RMND5B (C358A/H360A), with HA (ubiquitin) western blotting identifying high

molecular weight proteins (~80kDa – 175kDa) corresponding to ubiquitinated and

polyubiquitinated proteins associated with both wild-type and mutant GFP-RMND5B

(Figure 4.25B). Increased levels of ubiquitinated proteins were detected in

immunoprecipitates from cells overexpressing wild-type GFP-RMND5B that had been

treated with MG132 (Figure 4.25B). In comparison, markedly reduced amounts of

ubiquitinated proteins were detected in association with GFP-RMND5B

(C358A/H360A), suggesting that the RING domain mutations had reduced the E3

ubiquitin ligase activity of RMND5B (Figure 4.25B). Western blotting of the total


161

A

B

WT (C356S)

- - - + + +

Empty Vector

10µM MG132

GFP (~70kDa)

HA (Ubiquitin)

Ubn-Proteins (~80kDa – 175kDa)

GFP-RMND5A

- - - + + +

Empty Vector WT (C358S)

10µM MG132 HA-Ubiquitin

GFP (~70kDa)

Ubn-Proteins (~80 – 175kDa)

GFP-RMND5B


162

Figure 4.25: In vivo ubiquitination activity of GFP-RMND5A (C356A/H358A) and GFP-RMND5B (C358A/H360A). LNCaP cells were cotransfected with plasmids encoding HA-ubiquitin and GFP-RMND5A, GFP-RMND5A (C356A/H358A) GFP-RMND5B, GFP-RMND5B (C358A/H360A) or GFP (empty vector). Cells were cultured for 48 hours with 10μM MG132 added for the final 2 hours of culture. Following GFP immunoprecipitation of the cell lysates, HA western blotting was performed on the immunoprecipitated proteins and identified enhanced accumulation of ubiquitinated proteins in lysates from MG132 treated cultures. (A) GFP-RMND5A and GFP-RMND5A (C356A/H358A) were associated with similar levels of ubiquitinated proteins, while (B) in comparison to GFP-RMND5B, GFP-RMND5B (C358A/C360A) was associated with reduced levels of ubiquitinated proteins. Experiments were performed twice and representative results are shown. (WT = wild type)


163

cellular input samples for HA (Ubiquitin), and GFP (wild-type/mutant

RMND5A/RMND5B) detected similar levels in each sample, suggesting that the results

were not due to differences in the transfection efficiency or expression of the fusion

proteins (not shown).

4.2.7.7 In Vitro Auto-Ubiquitination Activity of RMND5A (C356A/H358A)

and RMND5B (C358A/H360A) RING Domains

To verify that the GST-RMND5A (C356A/H358A) RING and GST-RMND5B

(C358A/H360A) RING domains were expressed in E. coli BL21 cells, the pGEX-

RMND5A (C356A/H358A) RING and pGEX-RMND5B (C358A/H360A) RING

plasmids were each transformed into competent E. coli BL21 cells and glycerol stocks

were prepared (Sections 3.8.7, 4.2.7.2, 4.2.7.4). Transformed E. coli BL21 cells were

selected on LB Agar/Ampicillin plates and individual colonies were picked and

inoculated into LB Broth/Ampicillin (Section 3.11.1). Expression of the GST-fusion

proteins was induced by the addition of 1mM IPTG for 2.5 hours, cells were harvested

and 15µL aliquots of lysates from each culture was electrophoresed in 12%

polyacrylamide gels and stained with Coomassie blue (Sections 3.11.1, 3.15.2, 3.15.5,

Figure 4.26A). A prominent band at ~28kDa was present in lysates from the IPTG-

induced cells, with GST-RMND5A (C356A/H358A) RING and GST-RMND5B

(C358A/H360A) RING domains in both the soluble and insoluble fractions, indicating

that RMND5 RING domain mutants were packaged into inclusion bodies, however

sufficient levels of the proteins were present in the soluble fractions for purification and

use in in vitro ubiquitination assays. To obtain purified GST-RMND5A

(C356A/H358A) RING and GST-RMND5B (C358A/H360A) RING domain proteins

for in vitro ubiquitination assays, proteins were induced by the addition of 1mM IPTG

and extracted from 100mL bacterial cultures (Section 3.11.2). Purified GST-RMND5

RING proteins were electrophoresed in 12% polyacrylamide gels with BSA standards

and Coomassie blue stained to estimate the concentration of the purified GST-fusion

proteins (Section 3.25.2, 3.15.5, Figure 4.26B, C).

Prior to the assessment of the GST-RMND5A (C356A/H358A) RING and GST-

RMND5B (C358A/H360A) RING domain activity in comparison to that of the wild-

type GST-RMND5A RING and GST-RMND5B RING domains, in vitro ubiquitination

assays were re-optimised as the enzyme concentrations used in the Ubiquitinylation kit


164

B

C 1 2 3 4 5 6 7 8 Lane 1: MW Marker Lane 2: 1µg BSA Lane 3: 2.5µg BSA Lane 4: 5µg BSA Lane 5: 7.5µg BSA Lane 6: 10µg BSA Lane 7: 1µL GST-RMND5B RING Lane 8: 3µL GST-RMND5B RING Lane 9: 1µL GST- RMND5B (C358A/H360A) RING Lane 10:1µL GST- RMND5B (C358A/H360A) RING

~28kDa

9 10

1 2 3 4 5 6 7 8

Lane 1: MW Marker Lane 2: 1µg BSA Lane 3: 2.5µg BSA Lane 4: 5µg BSA Lane 5: 7.5µg BSA Lane 6: 10µg BSA Lane 7: 1µL GST-RMND5A RING Lane 8: 3µL GST-RMND5A RING Lane 9: 1µL GST-RMND5A (C356A/H358A) RING Lane 10: 3µL GST-RMND5A (C356A/H358A) RING

~28kDa

9 10

1 2 3 4 5 6 7

Lane 1: MW Marker Lane 2: GST-RMND5A (C356A/H358A) RING: no IPTG Lane 3: GST-RMND5A (C356A/H358A) RING: IPTG Lane 4: GST-RMND5A (C356A/H358A) RING: IPTG, insoluble fraction Lane 5: GST-RMND5A (C356A/H358A) RING: IPTG, soluble fraction Lane 6: GST-RMND5B (C358A/H360A) RING: no IPTG Lane 7: GST-RMND5B (C358A/H360A) RING: IPTG Lane 8: GST-RMND5B (C358A/H360A) RING: IPTG, insoluble fraction Lane 9: GST-RMND5B (C358A/H360A) RING: IPTG, soluble fraction

~28kDa

8 9 A


165

Figure 4.26: Expression and purification of GST-RMND5A (C356A/H358A) RING and GST-RMND5B (C358A/H360A) RING. (A) E. coli BL21 cells transformed with pGEX-RMND5A (C356A/H358A) RING or pGEX-RMND5B (C358A/H360A) RING were induced with IPTG and the presence of the GST-RMND5A (C356A/H358A) RING or GST-RMND5B (C358A/H360A) RING domains in the soluble and insoluble fractions prepared from the cultures was investigated by electrophoresis of the fractions in 12% polyacrylamide gels, which were stained with Coomassie blue. Following purification, the RING domain proteins were electrophoresed in 12% polyacrylamide gels, and the gels stained with Coomassie blue. In comparison to BSA standards, the concentration of (B) GST-RMND5A RING and GST-RMND5A (C356A/H358A) RING was determined to be ~2.6µg/µL and ~1.26µg/µL, respectively. (C) The concentration of GST-RMND5B RING and GST-RMND5B (C358A/H360A) RING was estimated to be ~4.3µg/µL and ~0.7µg/µL, respectively.


166

(ENZO Life Sciences) were high in comparison to those used in the literature (David et

al., 2011, Yang et al., 2000). Although this was suitable for initial in vitro

ubiquitination assays, in order to determine differences in the activity of wild-type and

mutant RMND5A and RMND5B RING domains, the concentrations of each enzyme

were re-evaluated (Table 4.2). In vitro ubiquitination assays were also carried out for 30

minutes at 37°C (compared to 60 minutes), and the reactions terminated with the

addition of 50µL 2x reducing loading dye and heating for 5 minutes at 95°C (Section

3.14.1).

Table 4.2 – Optimisation of in vitro ubiquitination assay enzyme concentrations

Enzyme Concentration

Utilised in Initial

Screen

Optimised

Concentration 1

Optimised

Concentration 2

E1 Activating Enzyme 100nM 25nM 50nM

E2 Conjugating

Enzyme, UbcH5b

2.5µM 0.25µM 0.5µM

GST-RING domain 4µM 1µM 2.5µM

Ten µL of each reaction product was electrophoresed in 4-12% gradient polyacrylamide

gels followed by western blotting for biotinylated ubiquitin. All reactions yielded

multiple protein bands, corresponding to ubiquitinated proteins, with most of these of a

similar appearance and intensity in reactions containing GST-RMND5A, GST-

RMND5A (C356A/H358A), GST-RMND5B and GST-RMND5B (C358A/H360A)

RING domain proteins. However, a band at ~38kDa, corresponding to

monoubiquitinated GST-RMND5 RING domains which was present at both enzyme

concentrations tested (Table 4.2), appeared to be reduced in in vitro ubiquitination

reactions containing the GST-RMND5A (C356A/H358A) RING and GST-RMND5B

(C358A/H360A) RING domains compared to those reactions containing wild-type

GST-RMND5A and GST-RMND5B RING domains (Section 3.14.1, Figure 4.27).

Additionally, high molecular weight protein bands at >100kDa corresponding to

polyubiquitinated proteins were of a greater intensity in reactions containing the wild-

type GST-RMND5B RING domain compared to the GST-RMND5B (C358A/H360A)

RING domain.


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Experiments were repeated using Condition 1 (Table 4.2), along with negative control

reactions lacking ATP or the GST-RMND5 RING domain, and 10µL of each product

was electrophoresed in 4-12% gradient gels (Section 3.14.1, 3.15). Western blotting for

biotinylated ubiquitin and GST were used to detect ubiquitinated proteins and GST-

fusion proteins, respectively (Section 3.15, Figure 4.28). Reactions lacking ATP yielded

a single band corresponding to free ubiquitin, whilst reactions lacking the GST-RMND5

RING yielded bands corresponding to ubiquitinated E2 enzyme only (Figure 4.28).

Again, multiple biotinylated ubiquitin bands of similar intensity were present in

reactions containing GST-RMND5A RING, GST-RMND5B RING, GST-RMND5A

(C356A/H358A) RING and GST-RMND5B (C358A/H360A) RING domains.

However, in reactions containing the GST-RMND5A (C356A/H358A) RING and GST-

RMND5B (C358A/H360A) RING domain proteins, the protein band at ~38kDa

corresponding to monoubiquitinated GST-RING domain proteins was reduced

compared to reactions containing wild-type GST-RMND5A RING and GST-RMND5B

RING domain proteins (Figure 4.28). GST western blotting identified expression of the

GST-RING domains, while high molecular weight bands corresponding to

polyubiquitinated proteins (>100kDa) were not visible in this set of in vitro

ubiquitination reaction products at the short x-ray film exposure times used to visualise

differences in the appearance and intensity of monoubiquitinated GST-RMND5 RING

domain protein bands (Figure 4.28). These results are consistent with the in vivo

ubiquitination assay results where mutation of the RMND5B RING domain resulted in

a larger reduction in ubiquitination activity compared to analogous mutations of the

RMND5A RING domain (Section 4.2.7.6) and are consistent with the RMND5A

(C356A/H358A) and RMND5B (C358A/H360A) mutations disrupting RMND5 RING

domain activity.


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Condition 1 Condition 2

Monoubiquitinated RMND5A RING domain

MonoubiquitinatedRMND5B RING domain

WT WT

C356A/H358A

A

B

WT WT

C358A/H360A

C358A/H360A

C356A/H358A

GST-RMND5A RING domain

GST-RMND5B RING domain

Figure 4.27: Optimisation of in vitro ubiquitination assays for GST-RMND5A

(C356A/H358A) RING and GST-RMND5B (C358A/H360A) RING. The

concentrations of E1, E2 and GST-RING domains used in in vitro ubiquitination

assays were optimised, with condition 1 containing 25nM E1, 0.25µM E2 and

1µM GST-RING, whilst condition 2 contained 50nM E1, 0.5µM E2 and 2.5µM

GST-RING. The in vitro ubiquitination assays were carried using either (A) GST-

RMND5A RING or GST-RMND5A (C356A/H358A) RING or (B) GST-

RMND5B RING or GST-RMND5B (C358A/H360A) RING. Western blotting for

biotinylated ubiquitin identified protein bands corresponding to ubiquitinated

proteins in reactions containing both wild-type and mutant GST-RMND5 RING

domains, including a band corresponding to monoubiquitinated GST-RMND5

RING domains in each reaction. (WT= wild-type).


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Figure 4.28: In vitro ubiquitination activity of GST-RMND5A (C356A/H358A) and GST-RMND5B (C358A/H360A). In vitro ubiquitination assays using GST-RMND5A RING, GST-RMND5B RING, GST-RMND5A (C356A/H358A) RING or GST-RMND5B (C358A/H360A) RING were performed and 10µL of each product was electrophoresed in 4-12% gradient gels. GST western blotting identified the presence of GST-RING domain proteins in all reactions except the negative control lacking E3 enzyme. Western blotting for biotinylated ubiquitin yielded multiple bands including a ~38kDa band corresponding in size to monoubiquitinated GST-RMND5 RING domains that was reduced in reactions containing GST-RMND5A (C356A/H358A) RING and GST-RMND5B (C358A/H360A) RING compared to reactions containing GST-RMND5A or GST-RMND5B, respectively. The experiment was performed twice and representative results are shown. (WT= wild-type).


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4.3 Discussion

The homologues RMND5A and RMND5B are highly conserved between diverse

species ranging from mammals to Arabidopsis. In this thesis, analysis of RMND5

protein domain composition and amino acid sequence identified that each contains 4

protein-protein interaction domains, a LisH, CTLH, CRA and RING domain and that

the amino acid variation between RMND5A and RMND5B is not confined to a

particular protein domain but rather is scattered throughout the protein sequences,

including areas of the proteins that do not contain identifiable protein domains.

Bioinformatics analyses indicated that the general protein domain architecture of

RMND5 proteins was similar to that of other LisH domain containing proteins, which

also commonly possess a CTLH and CRA domain, and that the protein, EMP has the

same protein domain architecture as human RMND5A and RMND5B (Schultz et al.,

1998). The six human proteins that possess a LisH, CTLH and CRA domain either form

part of the human CTLH complex or are orthologues of the known CTLH complex

members, suggesting that the presence of this particular protein domain architecture

allows the proteins to function in complementary cellular pathways or roles (Santt et al.,

2008).

The existence of orthologues of the CTLH complex members such as RMND5B and

RanBP10 (orthologue of RanBPM) provides evidence that these proteins may be able to

replace or join their paralogue in the CTLH complex, perhaps altering the function or

substrates of the complex. Alternatively, the function of the orthologues may have

diverged, as has been documented for other paralogous genes (Sahdev et al., 2008;

Singh and Hannenhalli, 2008). The LisH and CTLH domains are co-expressed in

RMND5A and RMND5B in addition to many as yet uncharacterised proteins,

suggesting that the domain pair functions together in its interaction with cellular

proteins or in alternative activities. However, although the LisH, CTLH and CRA

domains are proposed protein-protein interaction domains, their functions are not well

characterised, with the majority of information inferred from the activities of proteins

that contain these domains. For example, involvement of LisH domain containing

proteins in microtubule based protein transport was hypothesised from the function of

LIS1, the protein in which the LisH domain was originally identified (Emes and

Ponting, 2001).


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The C-termini of RMND5A and RMND5B that contain the RING domain are the most

highly conserved regions of the two proteins, indicating that the RING domain performs

similar functions in both proteins. This hypothesis is consistent with the findings of this

study that the RING domains of both RMND5 proteins possess E3 ubiquitin ligase

activity. Bioinformatics analyses of the RING domains of RMND5A and RMND5B

identified that although all eight residues predicted to be required for zinc coordination

are not the canonical cysteine and histidine residues, they are identical to those present

in yeast RMD5, a functional E3 ubiquitin ligase (Santt et al., 2008). Furthermore,

comparison of the RING domains of RMND5A and RMND5B to RING domain

consensus sequences identified that additional residues in the RING domains of

RMND5 proteins were conserved, with these residues present in other well

characterised E3 ubiquitin ligases such as MDM2 and plant E3 ubiquitin ligases (Stone

et al., 2005). Arabidopsis E3 ubiquitin ligases are well studied, with nine RING domain

variants characterised that differ in their zinc coordinating and intervening residues,

providing support for the existence of additional RING domain variants still to be

identified in human proteins (Stone et al., 2005).

The 34 amino acid residues located immediately amino-terminal to the RING domain

share 84% amino acid conservation between RMND5A and RMND5B. Since this

region is adjacent to the RING domain it may be important for the E3 ubiquitin ligase

activity of RMND5 proteins, as identified in CBL where the linker region, which is

located N-terminal to the RING domain, is important for interaction with the E2

enzyme UbcH7 (Zheng et al., 2000). As such, this region of RMND5A and RMND5B

may perform similar functions, potentially contributing to E2 enzyme binding, and the

high degree of homology indicates that both RMND5 proteins may be able to interact

with similar E2 enzymes via this region. Alternatively, the domain may be required for

the regulation of RING domain activity or structure. The amino-terminal of RMND5

proteins is not as well conserved as the carboxy-terminal, with the LisH and CTLH

region sharing between 60-68% amino acid identity and the carboxy-terminal located

CRA domain sharing 74% amino acid homology. Gene duplication, through which

RMND5A and RMND5B may have arisen, has been suggested to allow one copy of the

gene to maintain the normal gene function, whilst the other gene is able to undergo

divergence and acquire new functions, thereby allowing the evolution of new

morphology (Mazet and Shimeld, 2002). Thus, RMND5 proteins may have arisen from

the same gene and over time they have acquired different cellular substrates or binding


172

partners to allow divergent roles. This, together with the lower amino acid homology of

the LisH and CTLH domains, suggests that these domains may have diverged in their

protein recognition or interaction roles, indicating that RMND5 proteins may utilise

these domains to interact with different proteins. The interacting proteins may be

substrates of RMND5 E3 ubiquitin ligase activity, however, the protein domain

architecture of RMND5 proteins with the RING domain located carboxy-terminally and

the 3 additional protein-protein interaction domains located towards the amino-terminus

suggests that the amino-terminal domains may be able to mediate additional cellular

roles. This has been suggested for other RING domain containing proteins with a

similar multi-domain architecture, including BARD1. The RING domain-containing

protein BARD1, which functions with and enhances the activity of the functional E3

ubiquitin ligase and tumour suppressor BRCA1, contains an amino-terminal RING

domain and two additional protein-protein interaction domains towards the C-terminus

of the protein, it required for cell viability and has tumour suppressor properties of its

own (Irminger-Finger and Jefford, 2006). In addition to its activity with BRCA1,

BARD1 interacts with the mRNA polyadenylation cleavage factor CSTF1 and represses

its cellular polyadenylation activity, a mechanism by which BARD1 is proposed to

regulate cell proliferation (Kleiman and Manley, 1999; Irminger-Finger and Jefford,

2006).

The amino-termini of RMND5 proteins, which do not contain identifiable protein

domains or localisation signals exhibit a high degree of amino acid identity (76%),

which suggests that this stretch of 113 amino acids is important in RMND5 protein

function. According to the SBASE protein domain prediction database, RMND5

proteins also contain putative protein domains in their amino-terminal region which are

loosely conserved. RMND5A contains a ribulose phosphate 3-epimerase-like domain

(amino acids 11-40) and a GAT-like domain (amino acids 42-94), whilst RMND5B

contains a myosin tail like domain (amino acids 26-91) (Vlahovicek et al., 2005).

Although these domains are only weakly conserved in RMND5A and RMND5B, their

functions are interesting. Ribulose phosphate 3-epimerase is activated by Zn2+ binding,

similar to the RING domain, and functions in the pentose phosphate pathway (Akana et

al., 2006). The domain is also present in many other enzymes that use phosphorylated

proteins as substrates (Akana et al., 2006). The presence of this putative domain is

intriguing as many E3 ubiquitin ligases contain phosphorecognition motifs to allow

recognition and binding to their substrates for ubiquitination. For example, CBL


173

recognises phosphorylated receptor tyrosine kinases via its SH2 domain, which it then

ubiquitinates (Joazeiro et al., 1999). Therefore it would be interesting to determine

whether the ribulose phosphate 3-epimerase-like domain, although perhaps non-

functional with regards to epimerase activity, retains phosphate binding ability. The

GGA and Tom1 (GAT) domain is a ubiquitin binding domain that is present in GGA

clathrin coat adaptors, which function in the trans-Golgi network by sorting

monoubiquitinated cargo (Scott et al., 2004). While GAT domains are also found in

proteins involved in the sorting of ubiquitinated proteins in multi-vesicular bodies, it is

presently unknown whether this putative domain facilitates binding of ubiquitinated

proteins with RMND5A (Scott et al., 2004; Hurley et al., 2006). As will be discussed in

Chapter 6, a member of the CTLH complex, ARMC8α interacts with the endosomal

sorting protein Hrs which binds ubiquitinated proteins, thus the CLTH complex with

RMND5A as a member may ubiquitinate proteins and/or transport ubiquitinated

proteins to the endosomal system (Tomaru et al., 2010).

Finally, RMND5B contains a putative myosin tail like domain, the function of which

varies between myosin family members, either interacting with other myosin tails or

binding cargo proteins functioning in the intracellular trafficking of organelles (Sellers,

2000). Myosin motors traffic organelles along actin “tracks” whilst dyneins and

kinesins are involved in intracellular transport using microtubules, which consist of α-

tubulin and β-tubulin (Nelson et al., 2005). The presence of this putative domain in

RMND5B is interesting as the LisH domain is also implicated in organelle trafficking

by binding cytoplasmic dynein heavy chain and regulating microtubule function (Emes

and Ponting, 2001). Additionally, members of the yeast Vid30 complex, the human

orthologues of which contain LisH and CTLH domains, are involved in the vacuole-

based transport of FBPase and also interact with actin patches, thereby merging the

vacuole with the endocytic system (Brown et al., 2010; Alibhoy et al., 2012). The

presence of the LisH, CTLH, putative ribulose phosphate 3-epimerase-like, GAT-like

and myosin tail-like domains, which may recognise and transport ubiquitinated proteins

to the endosome/lysosome for degradation, implicate RMND5 proteins and members of

both the yeast Vid30 and human CTLH complexes in the intracellular transport of

cargo. As RMND5 proteins are E3 ubiquitin ligases, they may play dual roles, either

ubiquitinating proteins for degradation or recognising ubiquitinated proteins and

sorting/transporting them within the cell. In this study, proteasome inhibitors were used

to determine the accumulation of ubiquitinated proteins upon overexpression of


174

RMND5 proteins in vivo, however given the aforementioned information, future

experiments may more extensively investigate the use of lysosome inhibitors to

determine whether RMND5 proteins are involved in the lysosomal degradation of

proteins.

The presence in both RMND5A and RMND5B of the LisH and RING domains, which

function as dimerisation motifs, indicates that RMND5 proteins may use these domains

to form homodimers or heterodimers with each other, as demonstrated in this thesis

(Chapter 6), or with other similar proteins such as EMP (Kim et al., 2004). RING

domains are commonly used as dimerisation motifs, often resulting in their enhanced

activity, especially where RING domain heterdimersation is involved (Brzovic et al.,

2001; Linke et al., 2008). For example, dimerisation of BRCA1/BARD1 enhances the

activity of BRCA1 as does the association of MDM2 with MDMX, while RING domain

homodimerisation is also essential for the functioning of some E3 ubiquitin ligases

including the E3 enzyme RNF4 (Brzovic et al., 2001; Liew et al., 2010; Pant et al.,

2011). Thus it is feasible that RMND5 proteins interact with other RING domain

containing proteins to enhance their own E3 ubiquitin ligase activity or that of an

interacting RING domain containing protein. This hypothesis is consistent with findings

in this thesis that although mutant RMND5A (C356A/H358A) and RMND5B

(C358A/H360A) exhibited reduced RING domain automonoubiquitination in vitro, in

in vivo ubiquitination assays no reduction in the activity of the mutant RMND5A was

detected.

RING domain E3 ubiquitin ligases may function as single subunit or multi-subunit

(complex) proteins. Whether RMND5 proteins are able to function in either or both

modalities is yet to be determined. As mentioned previously, RMND5A has been shown

form to part of the large multi-protein CTLH complex and as such could impart its

activity to the complex (Kobayashi et al., 2007). However, both RMND5 proteins

contain LisH, CTLH and CRA domains in addition to the RING domain, and therefore

the additional protein-protein interaction domains could act in a substrate recognition

capacity, allowing RMND5A and/or RMND5B to function as E3 ubiquitin ligases

independently of the CTLH complex. The tissue distribution or endogenous levels of

RMND5A and RMND5B proteins are unknown, although both genes are expressed in

the majority of tissue types (NCBI Unigene EST Profiles

ftp://ftp.ncbi.nih.gov/repository/UniGene/Homo_sapiens/Hs75277, Hs27222). Further


175

investigation of the tissue distribution of these proteins will help to elucidate the cellular

roles of RMND5A and RMND5B and to identify whether they function together or

have divergent targets in different tissues. The LisH domain has been documented to aid

in the formation of other protein complexes, including E3 ubiquitin ligase complexes

such as the Cullin4A-RING DCAF1 complex (Cerna and Wilson, 2005; Choi et al.,

2008; Ahn et al., 2011). As other LisH, CTLH and CRA domain containing proteins

form part of the CTLH complex, the LisH domain may similarly be responsible for

complex formation (the CTLH complex will be discussed in detail in Chapter 6).

RMND5A and RMND5B were found to be localised in both the nucleus and cytoplasm,

however in approximately 60% of transfected cells, RMND5B exhibited a punctate

cytoplasmic distribution. A similar intracellular distribution was also displayed by

RMND5A in a lower proportion of GFP-RMND5A overexpressing cells. Due to the

lack of suitable commercially available antibodies against either RMND5A or

RMND5B, the intracellular distribution of endogenous RMND5A or RMND5B proteins

is unknown at this stage and as such it is not clear whether their punctate appearance is

due to a normal cellular function or results from their overexpression following

transfection. The punctate appearance of RMND5A and RMND5B has now been

identified in multiple cell lines, including both breast and prostate cancer cell lines (not

shown) and for RMND5B, following overexpression with multiple N and C terminal

protein tags, suggesting that the localisation is specific to RMND5 proteins (Dawson,

2006). The cellular localisation of RMND5A has been reported previously as diffusely

nuclear and cytoplasmic in HEK293 cells, which is in agreement with the findings in

this study, including those cells that displayed a punctuate cytoplasmic distribution of

RMND5A (Kobayashi et al., 2007).

The well characterised E3 ubiquitin ligase, Siah1 is also reported to exhibit a punctate

cytoplasmic distribution, due to its association with mitochondria, however cells

expressing Siah1 RING deletion mutants no longer displayed this staining, implicating

the RING domain structure or function in its punctate distribution (Hu and Fearon,

1999). As such, RMND5 proteins could be localising to these punctate speckles as part

of a normal cellular activity, which may be due to their RING domain and/or E3

ubiquitin ligase activity or unrelated cellular role, and which may be investigated in

future studies using RING domain deletion mutants of RMND5A and RMND5B. To

determine the specific localisation of RMND5 proteins, organelle stains may also be


176

used, for example early endosomal markers (EEA1, Rab4, Rab5 antibodies), late

endosome markers (Rab7, Rab9 antibodies) lysosomal markers (LAMP1, LAMP2

antibodies, Lyso Tracker® (Life Technologies)) or mitochondrial markers (Mito

Tracker® (Life Technologies)). Localisation of RMND5 proteins in the lysosome for

example may suggest either that they play a particular role associated with the

lysosome, or that they are packaged and degraded in the lysosome when overexpressed,

therefore additional functional studies will be required to characterise the relationship

between RMND5 protein intracellular localisation, activity and processing. RMND5

proteins are degraded at least in part by the proteasome as they show accumulation upon

proteasome inhibition (Chapter 6), and although mutation of the RING domains did not

markedly alter RMND5 protein localisation, their localisation may be determined by

their interaction with other proteins through either the RING domain or via one of their

other protein-protein interaction domains, the LisH, CTLH or CRA domains. Therefore,

single or multiple domain deletion mutants of RMND5A or RMND5B may enable the

determination of protein domain(s) that direct their intracellular localisation, including

their punctate distribution.

To characterise the E3 ubiquitin ligase activity of RMND5 proteins in this study, full

length RMND5A was cloned into the pGEX-2TK expression vector to allow the

expression of full length GST-tagged RMND5 proteins for use in in vitro ubiquitination

assays. However, the expression and purification of both GST-RMND5A and GST-

RMND5B fusion proteins was hindered by the packaging of the proteins into inclusion

bodies in E. coli BL21 bacterial cells. The misfolding of expressed proteins in bacteria

into insoluble aggregates, which are biologically inactive, has been widely documented

(Francis and Page, 2010). Proteins may be misfolded due to their requirement for longer

folding times or the need for chaperones to facilitate correct folding and, due to the

inability of bacteria to accomplish numerous eukaryotic post-translational modifications

such as phosphorylation, larger proteins and specific protein domains may be difficult to

obtain as soluble proteins (Villaverde and Carrio, 2003; Esposito and Chatterjee, 2006;

Francis and Page, 2010). The production of insoluble proteins may also be due to weak

promoter sequences, inefficient initiation of translation and the presence of rare codons

(codon bias) which may be rectified by the modification of promoter and translation

initiation sequences and the use of codon optimised bacterial cells, respectively (Sahdev

et al., 2008; Malhotra, 2009). In these experiments, western blotting of whole cell

lysates of the transformed bacteria indicated that GST-RMND5 fusion proteins were


177

produced in large quantities in the bacteria, suggesting that promoter sequences and

translation initiation were not causing the production of insoluble aggregates.

The E. coli BL21 strain utilised for the present studies was codon optimised and the

amount of full-length (insoluble) protein produced by the bacteria indicated that the low

yields of purified protein were not likely to be due to codon bias. Furthermore, a second

codon optimised strain, Rosetta BL21 cells was also tested, producing similar results.

The use of BL21 cells for routine production of fusion proteins has been suggested due

to their codon optimisation and the absence of expression of specific intracellular

proteases, thereby promoting stabilisation of the fusion proteins (Sahdev et al., 2008).

The GST protein tag, which is 211 amino acids in length, is also considered desirable as

it acts as a chaperone to aid in protein folding and can increase the solubility of the

fusion protein (Malhotra, 2009). In addition, the GST-fusion proteins can be

immobilised on glutathione agarose beads which have a high affinity for GST ensuring

low non-specific binding and therefore reducing contamination of the final purified

product with other proteins (Malhotra, 2009; Harper and Speicher, 2011). As an

alternative to the GST tag, smaller tags including a His-tag (6-10 histidine residues) or

Strep-II tag (8 amino acid residues WSHPQFEK) or an amino-terminal fusion with a

highly translated native protein such as maltose binding protein (396 amino acids) or

thioredoxin (109 amino acids) could have been tested to maximise solubility of the

fusion protein (Yasukawa et al., 1995; Malhotra, 2009).

Modification of expression conditions may also be used to increase solubilisation of

proteins, including lowering of the induction temperature, increasing aeration and the

co-expression of chaperones, however many proteins will remain insoluble (Esposito

and Chatterjee, 2006; Lorick et al., 2006). At lower temperatures, the expression of

chaperones is induced, the activity of proteases is reduced, and protein interactions

aiding in inclusion body formation and toxic phenotypes of fusion proteins are

suppressed (Sahdev et al., 2008). In this study, expression conditions were modified by

increasing aeration, reducing induction temperature and optimising GST-fusion protein

induction time, however altering these conditions did not increase solubilisation of

GST-RMND5A. The medium in which the bacterial cells were grown was also

optimised to obtain soluble protein by the addition of ZnCl2 to aid correct folding of the

RING domain. This modification of growth medium conditions has been suggested to

enhance protein solubilisation for the production of RING domain containing and other


178

proteins (Lorick et al., 2006; Sahdev et al., 2008). A further option was to purify the

inclusion bodies containing the misfolded proteins, disaggregate the proteins by

denaturation using buffers containing urea and then following dialysis, refold the

proteins. However this is a long process which is not considered optimal as the protein

yield is often low and the production of correctly folded proteins must be stringently

optimised and is not guaranteed, therefore potentially yielding non-functional proteins

or proteins with variably reduced activity (Middelberg, 2002; Rabhi-Essafi et al., 2007;

Sahdev et al., 2008). As such, this option for the production of GST-RMND5A was not

pursued.

Cell free systems may also be used for the production of proteins for biochemical assays

and can be advantageous because the in vitro system is directed to the production of a

single protein in comparison to in vivo systems that require the synthesis of normal

cellular proteins as well as the protein of interest, which may be cytotoxic to the cells

(Iskakova et al., 2006). Such methods involve the production of fusion proteins from

mRNA or DNA templates using coupled or linked in vitro transcription and translation.

These systems combine the use of a prokaryotic phage RNA polymerase and promoter

(e.g. T7, T3) with eukaryotic or prokaryotic extracts from human cells, wheat germ or

rabbit reticulocytes (for example) to provide the transcription and translation machinery

and are supplemented with energy regenerating solutions, amino acids and accessory

proteins (Stueber et al., 1984; Mikami et al., 2008). Newer methods for the production

of larger proteins such as the 200kDa Dicer, which possess biological activity have been

developed (Mikami et al., 2008). The use of human cell line extracts and advances in

cell free protein production systems are proposed to facilitate proper protein folding and

post-translational modification of the fusion proteins by the addition of reagents and

chaperones. Although these methods are an improvement on prokaryotic cell free

systems, which have been found to produce proteins with low activity, the production of

proteins that require post-translational modifications remains a challenge as it is

difficult to obtain homogeneously modified protein products in cell free systems

(Katzen et al., 2005; Iskakova et al., 2006). While these methods often involve the use

of radioactively labelled amino acids they can be modified to produce unlabelled tagged

proteins, with in vitro transcribed/translated proteins reported to be successfully used in

in vitro ubiquitination assays (Lorick et al., 2006; Jin et al., 2009).


179

An alternative to the production of proteins in bacterial cells would be their production

in mammalian, yeast or insect cells such as Trichoplusia or Spodoptera. For these

methods, insect cells are infected with the baculovirus expression vector system, and

this method has been used extensively for the production of complex proteins, including

membrane bound proteins, as well as in the pharmaceutical industry for the manufacture

of animal and human vaccines such as Cervarix (Kost et al., 2005; Cox, 2012).

Establishment of the method is time-consuming as the insect cells must be cultured and

infected with the baculovirus at particular stages of cell growth to achieve maximal

infection and protein production, the type of insect cells must be selected and the ability

of the cells to produce the protein of choice must be verified (Reuveny et al., 1993). The

use of insect cells is advantageous as they are often able to produce large quantities of

heterologous protein, and the cells can be cultured at room temperature, do not require

CO2 and in most cases can be grown in serum free media (Kost et al., 2005). Although

insect cells are eukaryotic, they are not able to perform all types of mammalian post-

translational modifications such as glycosylation, however insect cells engineered to

produce glycosylated proteins by the incorporation of mammalian genes encoding N-

glycan activity have now made it possible to produce glycosylated proteins in insect

cells (Hollister et al., 2002; Kost et al., 2005). Baculovirus expression systems

containing mammalian cell-active expression cassettes (BacMam), can also be utilised

to infect mammalian cells for the expression of recombinant proteins, and human

osteosarcoma and hepatic cells have been reported to produce high levels of gene

expression, however not all cell lines are able to be transduced or show effective

expression of the gene/protein of interest (Gao et al., 2002; Song et al., 2003; Kost et

al., 2005). There are therefore a number of alternative means for the production of large

quantities of recombinant proteins, which are misfolded such as RMND5 proteins or

that are quickly degraded in bacterial cells. However, due to time constraints an

alternative means to examine the activity of RMND5 proteins was to clone smaller,

soluble peptides containing the active domain of interest, the RING domain, into the

pGEX-2TK expression vector.

GST-RING domain fusion proteins were therefore used in in vitro ubiquitination assays

for this thesis in order to test the E3 ubiquitin ligase activity of RMND5 proteins. In

vitro ubiquitination assays are routinely employed to determine the E3 ubiquitin ligase

activity of putative ubiquitin ligases, and although where possible full length proteins

are used, shorter peptides encompassing the active domain, RING/HECT/U-box or


180

PHD domain, may also be used (Furukawa et al., 2005). The use of the GST moiety can

be advantageous when testing autoubiquitination activity as the GST tag dimerises,

which can aid in the detection of E3 activity as many RING domains also dimerise

(Lorick et al., 2006). However, in vitro auto-ubiquitination assays are prone to artefacts

due to non-specific protein aggregation formed during incubation and there are a

number of methods to minimise these, which were used where possible in this study. A

positive control in the form of CBL was utilised, and reactions lacking ATP and GST-

RING domains (negative controls) were included as part of the evaluation of the panel

of 11 E2 enzymes. Although the positive results expected from the in vitro assay

included a single high molecular weight band representing polyubiquitinated proteins

and lower molecular weight bands corresponding to mono- or multi-ubiquitinated

proteins, multiple additional bands corresponding to thioester linked proteins were also

visualised. Thus controls were used to investigate the identity of the additional bands

evident in ubiquitin western blots of the reaction products, specifically whether their

presence was related to the addition of GST-RMND5 RING domains and therefore

represented their ubiquitination products. Moreover, a full set of control reactions was

performed with each reaction omitting a reaction ingredient, following selection of the

E2 enzyme UbcH5b for further study. In this way the in vitro ubiquitination assays

consistently identified specific ubiquitination patterns (including bands corresponding

to auto-monoubiquitination) when the GST-RMND5 RING domains were included in

the reactions. These bands were not present when GST-CBL was used as a positive

control or when reaction ingredients were omitted, suggesting that the results were

specific to the presence of the RMND5 RING domains.

The in vitro ubiquitination assays performed using the RMND5 RING domains that

were carried out for this thesis provided a starting point for the analysis of RMND5

protein E3 ubiquitin ligase activity in vitro, and formed the preliminary information

required for further assessment of RMND5 protein E3 ubiquitin ligase activity.

However, a number of other assays may be performed in the future to further assess

RMND5 E3 ubiquitin ligase activity in vitro. Longer isoforms of GST-RMND5A or

GST-RMND5B that include additional domains required for substrate recognition, that

facilitate protein folding or that increase the number of lysine residues available for

ubiquitination (assisting in the visualisation of (larger) ubiquitinated proteins) may be

used in future studies (Zhang et al., 2009). To improve visualisation of ubiquitinated

proteins, in vitro ubiquitination assays may be performed with the GST fusion proteins


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attached to GST beads, allowing the removal (by washing) from the beads of non-

specific reaction products and E1 or E2 enzymes with thioester linked ubiquitin

(Furukawa et al., 2005; Lorick et al., 2006). Alternatively, RMND5 proteins may be

overexpressed by transfection of mammalian cells, immunoprecipitated and the purified

proteins used in in vitro assays, which also overcomes the problem of insoluble full

length protein production in bacteria (Furukawa et al., 2002; Burger et al., 2005; Lorick

et al., 2006). An advantage to this method is that the protein has been produced in

mammalian cells and as such would have undergone the correct post-translational

modifications, and in addition, if the E3 ubiquitin ligase requires binding partners or

cofactors, these may be coimmunoprecipitated, strengthening the in vitro assays (Lorick

et al., 2006).

Most E3 ubiquitin ligases are able to interact with more than one E2 conjugating

enzyme, with at least one of the UbcH5 family of E2 enzymes interacting with most E3

ligases. However, the identity of E2 enzymes that interact with an E3 cannot be

predicted and need to be determined in functional studies such as the in vitro

ubiquitination assays used in this thesis, which utilised a panel of 11 E2 enzymes. For a

variety of reasons, in vitro ubiquitination assays may not identify all possible E2

enzymes that interact with RMND5 proteins to mediate ubiquitin transfer. For example,

the RING domain alone was used in these assays and as such areas outside of the

domain required for E2 interaction would not be present, including regions immediately

adjacent to the RING domain that are known to be important for E2-E3 interactions

(Zheng et al., 2000). In addition, the interaction between the E2 and E3 may not be

sufficient to result in ubiquitin transfer in vitro, for example CBL can interact with

UbcH5b and UbcH7, but in vitro only the interaction with UbcH5b results in ubiquitin

transfer, although CBL uses UbcH7 in vivo (Huang et al., 2009).

In this study, in vitro ubiquitination assays were performed using a single set of

conditions with only 11 E2 enzymes evaluated and as such, RMND5 proteins may be

able to interact in vivo with other E2 enzymes that were or were not present in the panel.

In humans, ~40 E2 enzymes have been characterised, many of which are ubiquitously

expressed and exhibit both cytoplasmic and nuclear localisation, similar to both

RMND5A and RMND5B (van Wijk and Timmers, 2010). Of the E2 enzymes in the

panel that did not interact with RMND5 proteins in in vitro ubiquitination assays, it is

possible with optimal experimental conditions, specific cellular environments or the


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presence of cofactors or complex components including the RING domains of other

proteins that they are be able to interact with RMND5 proteins to mediate ubiquitin

transfer. Furthermore, as some E2 enzymes are chain initiating enzymes, the use of only

the RMND5 RING domains and single E2 enzymes may have failed to detect

interactions between RMND5 proteins and important chain elongating E2 enzymes that

require mono- or poly-ubiquitinated substrates for their activity (Windheim et al., 2008;

Ye and Rape, 2009). For example, TRAF6 utilises UbcH5 to initiate chain formation by

monoubiquitating the substrate, and this is followed by its interaction with the chain

elongating E2 enzyme UBE2N-UBE2V1 to polyubiquitinate the substrate NF-κB. As

the E2 enzymes UBE2N-UBEV1 and UBE2S cannot perform ubiquitin chain initiation,

their substrate activity is dependent on substrate chain initiation by other E2 enzymes

(Christensen et al., 2007; Petroski et al., 2007; Windheim et al., 2008).

Finally, the in vitro assays used in this project assessed the auto-ubiquitination activity

of RMND5 RING domains and therefore did not allow evaluation of potential

interactions of full-length RMND5 proteins with different E2 enzymes depending on the

substrate and cellular environment. To identify E2 enzymes that interact with RMND5

proteins in vivo, future studies may use immunoprecipitation assays from whole cell

lysates to determine E2-E3 interaction pairs, which may be confirmed by GST pull-

down assays and colocalisation microscopy (Lorick et al., 2006). Additionally, the type

of ubiquitin linkage attached to the substrate following the E3 interaction with specific

E2 enzymes can be assessed in either in vitro or in vivo assays with the use of linkage

specific antibodies or ubiquitin mutants (Lorick et al., 2006).

The use of GST-RMND5A and GST-RMND5B RING domains in in vitro assays

showed that RMND5 proteins were able to interact with a number of E2 enzymes to

mediate ubiquitin transfer and that both RMND5A and RMND5B were able to interact

with UbcH5b to mediate their auto-monoubiquitination. The finding that RMND5A and

RMND5B interacted with members of the UbcH5 family of E3 ubiquitin ligases was

not unusual as almost all E3 ubiquitin ligases are able to interact with at least one of the

family members, including the E3 ubiquitin ligase AO7, which only interacts with

members of this E2 enzyme family (Hakli et al., 2004; Lorick et al., 2006). The UbcH5

E2 conjugating enzyme family contains only the ubiquitin conjugating domain and as

such, members of this family are classified as class I E2 enzymes. UbcH5 members are

able to interact with a multitude of E3 ubiquitin ligases to mediate the transfer of


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ubiquitin to many different substrates, and in most cases the enzymes are not specific

for the target lysine residues on the substrate that are ubiquitinated (Kirkpatrick et al.,

2006; Ye and Rape, 2009). UbcH5 enzymes are associated with monoubiquitination and

are therefore known as chain initiating E2 enzymes, responsible for the initial

monoubiquitination of the substrate but subsequently replaced on the E3 interacting site

by a chain elongating E2 enzyme, which is specific for the type of ubiquitin chain

linkages that are formed (Ye and Rape, 2009). However, members of the UbcH5 family

are also able to form ubiquitin chains through lysine 11, 48 or 63 in vitro and are not

specific for the ubiquitin chain linkages that they form (Windheim et al., 2008; Boname

et al., 2010; Dynek et al., 2010). It is therefore proposed that UbcH5 enzymes are able

to perform polyubiquitination under some conditions but usually perform

monoubiquitination.

In a global yeast two hybrid screen for E2-E3 interaction pairs, RMND5B was

identified to interact with UbcH5 family members but not with UbcH2, consistent with

the results of the present study (van Wijk et al., 2009). Given the function of UbcH5

enzymes, RMND5A and RMND5B may utilise these enzymes for chain initiation and

use other E2 enzymes to extend these chains in elongation reactions. In contrast to

RMND5B, under the assay conditions used for this study, RMND5A was able to

interact with UbcH2 to mediate ubiquitin transfer, suggesting that RMND5 proteins

target different substrates or direct alternative outcomes of their ubiquitinated substrates

due to their interaction with different E2 enzymes. Interestingly the yeast orthologue of

UbcH2, Ubc8 (Gid 3) is reported to be the E2 enzyme associated with the Vid30

complex, and as RMND5A is a proposed member of the CTLH complex, human

orthologue of the Vid30 complex, it is feasible that the CTLH complex similarly utilises

UbcH2 (Kaiser et al., 1994; Santt et al., 2008). UbcH2, a class 3 E2 conjugating

enzyme, is associated with the formation of lysine 11 and 48 linked ubiquitin chains

which target substrate proteins for degradation by the proteasome, indicating that if

RMND5A is able to utilise this E2 enzyme in vivo, its ubiquitinated protein substrates

may be similarly targeted for proteasomal degradation (Santt et al., 2008).

In order to confirm E3 ubiquitin ligase activity of RMND5 proteins, site directed

mutagenesis of the RING domains was performed. As described previously (Section

1.7.2), the RING domain contains eight conserved cysteine or histidine residues that are

required to chelate two zinc ions which hold the RING domain in a cross brace structure


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for the correct functioning of the domain (Deshaies and Joazeiro, 2009). In order to

show that the RING domain is responsible for the E3 ubiquitin ligase activity of a

protein, the RING domain is mutated to reduce or abolish E2 enzyme binding, with

these mutants used in in vitro and in vivo ubiquitination assays. Conventionally, one or

two of the eight zinc coordinating residues are mutated to disrupt domain function,

however not all mutations will disrupt the RING structure, as was shown in this study.

For these experiments, a conserved amino acid (C379S), mutation of which in yeast

RMD5 rendered it unable to ubiquitinate its substrate fructose-1,6-bisphosphatase, was

mutated in the RMND5 proteins, however this mutation did not grossly affect RMND5

protein E3 ubiquitin ligase activity. Despite the lack of major effects on RING domain

function, the mutation may affect the ability of the RING domains to ubiquitinate

specific substrates as was reported for yeast RMD5 (Santt et al., 2008). In agreement

with this, the RMND5A (C356S) mutant appeared to preferentially mediate auto-

monoubiquitination compared to wild-type RMND5A, consistent with a difference in a

specific activity of the mutant. Disruption of a single amino acid would leave 7 other

zinc coordinating residues and potentially other conserved residues in the RING domain

that maintain protein folding. Results of the experiments indicated that the single

mutation permitted E2 enzyme binding but may have distorted the positioning of the

ubiquitin molecules and their attachment to each other upon elongation of the

monoubiquitin chain, thereby rendering the mutant RMND5A (C356S) RING domain

with a preference for auto-monoubiquitination.

As the C356S mutation was not sufficient to disrupt E2 enzyme binding of the

RMND5A RING domain, and a similar result was obtained for RMND5B (C358S), two

amino acid residues were mutated in both RMND5A and RMND5B. For these studies,

the 2 amino acids, C356A/H358A (RMND5A) and C358A/H360A (RMND5B) were

chosen as each binds a separate zinc residue in both RING domains. Therefore

theoretically, mutation of both residues in the RING domains could reduce or negate the

ability to bind two zinc residues, disrupting the structure and E3 ubiquitin ligase activity

of the domain, as has been observed following mutation of the equivalent amino acids

in other RING domain containing proteins in the literature (Zhang et al., 2009). The

residues mutated in RMND5A and RMND5B were also canonical cysteine and histidine

RING domain residues, although not all 8 coordinating residues present in the RING

domains of RMND5 proteins are these conserved amino acids (Section 4.2.1).

Alternatively, if the site of E2 interaction was known, the coordinating amino acids of


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the RING domain important for E2 interaction could be mutated for investigation of E2-

E3 interaction as this strategy would be more likely to inhibit E2-E3 interaction and

disrupt substrate ubiquitination. In this case, the amino acid residues required for

UbcH5B interaction were unknown, however, bioinformatics analysis and comparison

of the common E2 interaction residues in RING domains in different E3 ubiquitin

ligases has subsequently identified that RMND5 proteins both contain conserved E2

interaction residues within their RING domains (not shown) which could be mutated in

further studies to abolish RING domain activity (Deshaies and Joazeiro, 2009). For

example, mutation of a conserved tryptophan residue in the RING domains of CBL

(W408A) and TOPORS (W131A) renders these E3 ubiquitin ligases inactive, and this

residue is hypothesised to be critical for the physical interaction of the E3 with its

cognate E2 (Joazeiro et al., 1999; Rajendra et al., 2004).

In in vitro ubiquitination assays, the RMND5 RING domains were able to auto-

monoubiquitinate, although the physiological relevance of this observation, if any,

remains to be determined. Due to their enzymatic activity, E3 ubiquitin ligases are able

to ubiquitinate their own lysine residues either as a by-product of their activity or as a

means of auto-regulation. For example, MDM2 ubiquitinates itself and additionally

ubiquitinates p53, targeting both proteins for degradation (Fang et al., 2000). Whether

RMND5 proteins are able to auto-ubiquitinate their RING domains in vivo, the type of

ubiquitin chains formed (as the E2 enzyme UbcH5b used in in vitro ubiquitination

assays prefers monoubiquitination) and the outcome of ubiquitination will need to be

addressed in future studies.

Mutation of the RING domain can result in the altered cellular localisation of the E3

ubiquitin ligase, however mutant RMND5A (C356A/H358A) and RMND5B

(C358A/H360A) exhibited a similar nuclear and cytoplasmic localisation compared to

the wild-type protein under the culture conditions used in this study. In contrast,

mutation of the first RING domain in Parkin was reported to result in its mislocalisation

and packaging into aggresomes in which misfolded proteins are localised, but did not

cause the abnormal localisation of wild-type Parkin (Cookson et al., 2003). Similarly,

deletion of the RING domain of Siah1 also resulted in its altered cellular localisation

from a punctuate cytoplasmic distribution to a diffuse cytoplasmic localisation (of the

mutant but not endogenous wild-type Siah1) (Hu and Fearon, 1999). These findings

suggest that the mutant RMND5 proteins are still able to maintain interactions with


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endogenous RMND5 proteins or other cellular binding partners that determine their

cellular distribution, which may or may not be involved in their E3 ubiquitin ligase

activity (e.g. CTLH complex members).

In in vitro ubiquitination assays performed to assess RMND5A (C356A/H368A) mutant

activity, a small reduction in activity was observed, however no differences in the

activity of the wild-type and mutant RMND5A proteins were detected in the in vivo

ubiquitination assays used in this thesis project. Mutation of the RING domains of E3

ubiquitin ligases can result in the stabilisation of their substrates as well as their own

stabilisation as they are no longer able to regulate their own activity by

autoubiquitination or that of their substrate (Hu and Fearon, 1999; Fang et al., 2000).

Therefore, if RMND5 proteins form homo- or hetero-dimers in vivo, mutant RMND5

proteins may interact with wild-type functional RMND5A or RMND5B or other

proteins and increase their stability, resulting in no loss of E3 ubiquitin ligase activity in

vivo, as was observed for the RMND5A (C356S) and (C356A/C358A) mutants.

Although the RMND5A (C356A/H358A) mutant did not exhibit a general reduction in

E3 ubiquitin ligase activity in vivo, its ability to ubiquitinate specific substrates may be

disrupted, as was described previously for yeast RMD5 (Santt et al., 2008).

Additionally, whilst the RMND5A (C356A/H358A) mutation may have disrupted the

RING domain and its interaction with UbcH5b in vitro, the mutation may not have

affected the mutant RMND5A binding to other E2 enzymes in vivo, with several E2

enzymes such as Rad18 and gp78 reported to interact with E2 enzymes outside of the

RING domain (Bailly et al., 1997; Das et al., 2009; Li et al., 2009). Another

explanation for the apparently unaltered association of the RMND5A (C356A/H358A)

mutant with ubiquitinated proteins in vivo is that disruption of the RMND5A RING

domain resulted in the misfolding of RMND5A and its recognition and ubiquitination

by proteins functioning in quality control. Therefore, RMND5A (C356A/H358A) may

seem to be maintaining its E3 ubiquitin ligase activity in vivo whilst the (unidentified)

ubiquitinated proteins detected in the assay may represent increased ubiquitination of

the misfolded RMND5A (C356A/H358A). Alternative methods to abolish the E3

ubiquitin ligase activity of RING domain containing proteins are available, for example,

RING domain deletion mutants are commonly used in the literature for in vivo or in

vitro studies, or for in vitro ubiquitination assays, zinc chelators such as EDTA or

TPEN may be used to inhibit the RING domain activity (Lorick et al., 2006; Gao et al.,

2009).


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The activity of RMND5B (C358A/H360A) was markedly reduced in in vitro and in vivo

assays, suggesting that mutation of the two amino acids residues resulted in substantial

disruption of RING domain function. Alternatively, in in vivo ubiquitin assays the

structure of RMND5B as a whole may have been altered by mutation of the RING

domain, thereby disrupting other RMND5B protein-protein interactions in addition to

that of the RMND5B RING domain interaction with E2 enzymes. E3 ubiquitin ligase

RING domain mutants have been reported to function in a dominant negative manner,

for example, a RING domain deleted Siah1 mutant without E3 ubiquitin ligase activity

was shown to accumulate in cells due to its inability to auto-ubiquitinate and target

itself for degradation (Hu and Fearon, 1999). Furthermore, the mutant Siah1 protein was

still able to associate with other Siah and Sina proteins and a Siah substrate DCC.

Mutant Siah1 was proposed to prevent substrate ubiquitination and degradation by the

proteasome due to its sequestration of the substrate away from wild-type Siah1, thereby

providing evidence that the mutant Siah1 functions in a dominant negative manner (Hu

and Fearon, 1999). However, mutation of the RING domain may result in the enhanced

degradation of the misfolded protein and in this study it was noted that it was difficult to

immunoprecipitate equal amounts of mutant RMND5 proteins compared to wild-type

proteins which may be due to an increased rate of degradation of the misfolded mutants

(or alternatively a reduction in the production of these mutants).

The results from this chapter have shown that RMND5A and RMND5B are

multidomain proteins with a similar protein domain architecture including a RING

domain, typically present in E3 ubiquitin ligases. These studies have provided evidence

that both RMND5 proteins function as E3 ubiquitin ligases in vitro and in vivo in

LNCaP cells. As RMND5B was originally identified to interact with NKX3.1, the

ability of RMND5A and RMND5B to bind and ubiquitinate NKX3.1 was investigated.

Chapter 5 RMND5 Proteins Ubiquitinate NKX3.1

Chapter 5: RMND5 Proteins Ubiquitinate

NKX3.1


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5.1 Introduction

Expression of the prostatic tumour suppressor NKX3.1 is reduced in many high grade

prostate cancers and is undetectable in up to 80% of metastatic prostate tumours

(Section 1.2). Loss of heterozygosity (LOH) at the 8p21.2 chromosomal locus that

includes the NKX3.1 gene is common, however inactivating mutations in the coding

region of the remaining allele are not yet reported (Voeller et al., 1997; Xu et al., 2000;

Ornstein et al., 2001). NKX3.1 gene promoter hypermethylation is also not widely

detected in prostate tumours and studies have reported increased NKX3.1 mRNA levels

and discordance between NKX3.1 mRNA and protein levels in prostate tumours,

suggesting that the NKX3.1 gene is transcribed (Voeller et al., 1997; Ornstein et al.,

2001; Asatiani et al., 2005; Lind et al., 2005; Bethel et al., 2006; Bethel and Bieberich,

2007). As such, the mechanism(s) by which NKX3.1 protein levels are reduced or

undetectable in prostate cancer cells are not well understood but provide evidence for

altered regulation of NKX3.1 at the translational or post-translational level (Bethel et

al., 2006). In addition, it has been documented that NKX3.1 is aberrantly localised in

the cytoplasm in a proportion of prostate tumours, indicating its inability to perform

transcriptional regulatory roles and suggesting that deregulation of proteins involved in

the post-translational regulation of NKX3.1 may contribute to prostate carcinogenesis

(Kim et al., 2002b).

The transcriptional regulation of NKX3.1 by the androgen receptor (AR), retinoic acid

receptor (RAR), ETS1 and ERG/ESE has been characterised, however post-

transcriptional and post-translational regulation of NKX3.1 in the normal prostate and

in prostate tumour cells are less well understood (Section 1.2.3) (He et al., 1997;

Prescott et al., 1998; Kunderfranco et al., 2010; Thomas et al., 2010; Preece et al.,

2011). Phosphorylation and ubiquitination are closely related post-translational

modifications, with many E3 ubiquitin ligases and substrates requiring phosphorylation

prior to activation or ubiquitination, respectively (Yamamoto et al., 2005; Lin et al.,

2006; Suizu et al., 2009). The relationship between post-translational modification and

protein processing is evident in the regulation of NKX3.1, which has been documented

to undergo phosphorylation on a number of serine and threonine residues that affect

protein ubiquitination and stability. NKX3.1 levels are regulated by the protein kinase

casein kinase 2 (CK2), with the CK2α’ catalytic subunit of CK2 specifically

phosphorylating NKX3.1 on threonine residues 89 and 93 (Thr89, Thr93) resulting in


189

NKX3.1 protein stabilisation (Figure 5.1) (Li et al., 2006). Reduction of CK2 activity

by siRNA or specific inhibitors decreases NKX3.1 protein levels, which are restored

following proteasome inhibition, thereby demonstrating that CK2-mediated

phosphorylation prevents NKX3.1 degradation by the proteasome (Li et al., 2006). The

mechanisms by which Thr89 and Thr93 phosphorylation in the NKX3.1 acidic domain

affect NKX3.1 stability have been investigated by nuclear magnetic resonance imaging,

identifying that the homeodomain associates with the acidic domain and upon DNA

binding, this association is relieved (Ju et al., 2006; Ju et al., 2009). Phosphorylation of

the acidic domain enhances its interaction with the homeodomain and increased stability

of the interaction is predicted to prevent ubiquitination of NKX3.1, which is

hypothesised to occur in the NKX3.1 homeodomain (Ju et al., 2009). These events

inhibit ubiquitin-mediated degradation of NKX3.1 and therefore NKX3.1 levels are

stabilised.

Conversely, the inflammatory cytokines, tumour necrosis factor α (TNFα) and

interleukin 1β accelerate the degradation of NKX3.1 by phosphorylating NKX3.1 on

serine 196 (Ser196) in the carboxy-terminal, promoting ubiquitination (Figure 5.1)

(Markowski et al., 2008). NKX3.1 is not believed to be ubiquitinated at its carboxy

terminus near Ser196, although NKX3.1 has been proposed to be ubiquitinated on

lysine residues within its homeodomain, which is located close to the carboxy terminus

(Markowski et al., 2008; Ju et al., 2009). The lysine residues of NKX3.1 that are

ubiquitinated have not been investigated, and in addition, the kinase and ubiquitin ligase

responsible for cytokine-mediated degradation of NKX3.1 have not been identified

(Markowski et al., 2008). The finding that inflammatory cytokines are involved in

NKX3.1 degradation is interesting as Nkx3.1 expression is markedly reduced in

bacterial prostatitis and inflammation has been implicated in prostate carcinogenesis,

although the drivers and mediators of these events have not been elucidated (De Marzo

et al., 2007; Khalili et al., 2010). Phosphorylation of NKX3.1 can also occur on the

closely located serine residues Ser185 and Ser195 (Figure 5.1) (Markowski et al.,

2008). Phosphorylation of NKX3.1 at Ser185 promotes its ubiquitination and

degradation under steady state conditions, with Ser185Ala mutants exhibiting increased

stability, while Ser195 phosphorylation enhances both cytokine mediated Ser196 and

steady state Ser185 phosphorylation, leading to NKX3.1 degradation (Markowski et

al., 2008). NKX3.1 phosphorylation on serine 48 (Ser48) was originally investigated

due its proximity to arginine 52 (R52), which is substituted with cysteine (R52C) due to


190

the C154T polymorphism in ~11% of healthy men (Gelmann et al., 2002). In prostate

cancer patients, the presence of this polymorphism (R52C) is associated with a ~1.8

fold increased risk of stage C or D disease (or Gleason Score of 7 or greater) (Gelmann

et al., 2002). R52 is important for directing Ser48 phosphorylation by protein kinase C

(PKC) and mutation of either of these residues (R52C or S48A) results in reduced

NKX3.1 phosphorylation. Consistent with previous studies reporting that NKX3.1

phosphorylation facilitates NKX3.1 self-association thereby reducing NKX3.1 DNA

binding (through Ser89 and Ser93), phosphorylation at this site affects NKX3.1 DNA

binding (Gelmann et al., 2002). Both R52C and S48A mutants exhibit enhanced binding

compared to the wild-type NKX3.1 protein on an NKX3.1 consensus DNA sequence

(Gelmann et al., 2002), therefore suggesting that NKX3.1 phosphorylation on multiple

residues may aid in its self-association. Although the R52C polymorphism alters

NKX3.1 phosphorylation and DNA binding it does not affect NKX3.1 coactivation of

SRF transcriptional activity, however the transcriptional activity of other NKX3.1

cofactors may be affected. Supporting this, a more recent study has found that men who

were either heterozygous or homozygous for the C form of the R52C polymorphism

were at 1.6x risk for enlarged prostate size with a high proportion also developing

benign prostatic hyperplasia (Rodriguez Ortner et al., 2006).

Figure 5.1: Post-translational modification of NKX3.1. NKX3.1 is reported to be phosphorylated on serine 48 (Ser48) (Gelmann et al., 2002), threonine residues 89 and 93 (Thr89, Thr 93) (Li et al., 2006), and serine residues 185, 195 and 196 (Ser185, Ser195, Ser196) (Markowski et al., 2008) and to undergo ubiquitination on lysine residues within the homeodomain (Ju et al., 2009).


191

Although its ubiquitination has been incompletely characterised, NKX3.1 is

ubiquitinated by the E3 ubiquitin ligase TOPORS both in vitro and in vivo, resulting in

its proteasome dependent degradation in vivo (Guan et al., 2008). Supporting these

findings, siRNA mediated knockdown of TOPORS resulted in NKX3.1 protein

accumulation and its prolonged half-life (Guan et al., 2008). Using NKX3.1 deletion

proteins, it was determined that the homeodomain was required for the interaction

between NKX3.1 and TOPORS, and although the amino-terminus of NKX3.1 was

sufficient for binding to TOPORS, the interaction was weaker than that of proteins

containing either the homeodomain and the amino-terminus, or the homeodomain and

the carboxy-terminus (Guan et al., 2008). Consistent with previous reports that the

homeodomain was a possible site of NKX3.1 ubiquitination, the study also

demonstrated that the homeodomain alone was ubiquitinated by TOPORS, although to a

lesser degree than other proteins containing the homeodomain and additional protein

sequences. These results supported a hypothesis that the homeodomain is likely to be

the site of ubiquitination, with other residues outside of the homeodomain increasing the

efficiency of NKX3.1 interaction with TOPORS (Guan et al., 2008). NKX3.1 contains

fourteen lysine residues which are potential ubiquitination sites with nine of these

located in the homeodomain, although the specific lysine residues in the homeodomain

that are ubiquitinated by TOPORS or by other E3 ubiquitin ligases, and the

ubiquitination of other lysine residues in the NKX3.1 protein remain uncharacterised.

Interestingly, both TOPORS and NKX3.1 interact with the DNA Helicase

Topoisomerase I (TOPO I). TOPORS was originally identified as a TOPO I binding

partner and acts as a SUMO-1 E3 ligase that regulates TOPO I and chromatin binding

protein activity (Hammer et al., 2007; Pungaliya et al., 2007). TOPORS was

hypothesised to interact with TOPO I, recruiting it to RNA polymerase II transcriptional

complexes, whilst NKX3.1 was found to enhance TOPO I DNA helicase activity

(Haluska et al., 1999; Bowen et al., 2007). Additionally, upon exposure to DNA

damaging agents, NKX3.1 and TOPO I comigrate within the nucleus (Bowen et al.,

2007). These studies implicate both NKX3.1 and TOPORS in the regulation of TOPO I

activity, and therefore suggest their ability to regulate DNA replication, gene expression

and DNA repair.

RMND5B was originally identified in our laboratory as an NKX3.1 binding partner in a

yeast two-hybrid screen. In order to determine the possible outcome(s) of the interaction

between RMND5B and NKX3.1, the function of RMND5B and its homologue


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RMND5A was investigated in this thesis. Both RMND5 proteins were determined to

function as E3 ubiquitin ligases (Chapter 4), leading to the hypothesis that NKX3.1 may

be a substrate of RMND5 mediated ubiquitination. Therefore, the ubiquitination of

NKX3.1 by RMND5A and RMND5B and the outcome of NKX3.1 ubiquitination was

investigated in prostate cancer cells.


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5.2 Results

5.2.1 RMND5 Proteins Interact with NKX3.1 in LNCaP Prostate Cancer Cells

5.2.1.1 RMND5A Interacts with NKX3.1

RMND5B was initially identified in our laboratory due to its interaction with the

prostatic tumour suppressor NKX3.1 (Dawson, 2006), however the interaction between

RMND5A and NKX3.1 had not been determined. To investigate NKX3.1 interaction

with RMND5A, LNCaP cells growing in 10cm petri dishes were cotransfected with

plasmids encoding GFP-RMND5A and NKX3.1-V5 (Section 3.1.4). At 48 hours post-

transfection, the cells were lysed, an aliquot (50µL) of the cell lysate was taken, and the

remaining lysate was subjected to either GFP or V5 immunoprecipitation (Section

3.13). The immunoprecipitation reaction products were electrophoresed in 12%

polyacrylamide gels and western blotting was performed for GFP or V5 (Section 3.15).

For GFP-RMND5A immunoprecipitations, GFP-RMND5A was present at the expected

size of ~70kDa in the immunoprecipitate, but not in the input lysate indicating that the

expression levels of GFP-RMND5A were below the sensitivity of detection of the

antibody in this dilute fraction. GFP-RMND5A was not detected in the mock

immunoprecipitation control, indicating successful immunoprecipitation (Figure 5.2A).

Western blotting of the lysates for NKX3.1-V5 identified a ~35kDa protein band

corresponding to NKX3.1-V5 in both the input lysate and immunoprecipitate, with no

bands in the mock immunoprecipitated control, indicating that RMND5A and NKX3.1

interact in LNCaP cells. When NKX3.1-V5 was immunoprecipitated, western blotting

for NKX3.1-V5 identified a band of ~35kDa corresponding to NKX3.1-V5 in the input

lysate and immunoprecipitate, whilst western blotting for GFP-RMND5A resulted in a

band of ~70kDa in the input lysate and immunoprecipitated fractions (Figure 5.2A). No

bands were detected in the untransfected or mock immunoprecipitated controls and

together these results indicated that RMND5A and NKX3.1 interact in LNCaP cells.

5.2.1.2 RMND5B Interacts with NKX3.1

To confirm the interaction between RMND5B and NKX3.1, LNCaP cells growing in

10cm petri dishes were cotransfected with plasmids encoding GFP-RMND5B and

NKX3.1-V5, lysed at 48 hours post-transfection, an aliquot of the cell lysate taken, and

the remaining lysate immunoprecipitated using GFP antibodies (Sections 3.1.4, 3.13).

Immunoprecipitated proteins were electrophoresed in 12% polyacrylamide gels and

western blotting was performed for GFP-RMND5B and NKX3.1-V5 (Section 3.15). A


194

Figure 5.2: NKX3.1 interacts with RMND5A and RMND5B in prostate cancer cells. (A) LNCaP cells were cotransfected with plasmids encoding GFP-RMND5A and NKX3.1-V5. At 48 hours post-transfection, (A) (i) GFP-RMND5A was immunoprecipitated from the cells using GFP antibodies or (ii) NKX3.1-V5 was immunoprecipitated using V5 antibodies. Western blotting for GFP and V5 was performed, identifying co-immunoprecipitation of GFP-RMND5A and NKX3.1-V5. (B) LNCaP cells were cotransfected with plasmids encoding GFP-RMND5B and NKX3.1-V5 and 48 hours post-transfection, GFP-RMND5B was immunoprecipitated using anti-GFP antibodies. Western blotting for GFP-RMND5B and NKX3.1-V5 identified both proteins in the immunoprecipitated samples. Each experiment was performed twice and representative results are shown.


195

band corresponding to GFP-RMND5B was present in the immunoprecipitate,

identifying successful GFP-RMND5B immunoprecipitation (Figure 5.2B). GFP-

RMND5B bands were not detected in the input lysate, indicating that GFP-RMND5B

levels were below the sensitivity of the antibody in this (dilute) fraction, and similarly

no bands were detected in the untransfected and mock immunoprecipitated control

reactions (Figure 5.2B). Western blotting identified a ~35kDa band corresponding in

size to NKX3.1-V5 in the immunoprecipitated and input lysate samples (Figure 5.2B).

These results confirmed the interaction between RMND5B and NKX3.1 and with

results described in Section 5.2.1.1 indicated that both RMND5A and RMND5B are

able to interact with NKX3.1 in LNCaP cells.

5.2.2 RMND5 Proteins Colocalise with NKX3.1 in LNCaP Cells

The localisation and colocalisation of NKX3.1 and RMND5 proteins in LNCaP cells

were investigated using fluorescence microscopy. For these studies, LNCaP cells

growing on coverslips were cotransfected with plasmids encoding NKX3.1-V5 and

either GFP-RMND5A or GFP-RMND5B, then cultured for 48 hours post-transfection

prior to V5 immunostaining of the cells (Section 3.1.3, 3.1.4, 3.16). When coexpressed

with NKX3.1-V5, both GFP-RMND5A and GFP-RMND5B exhibited a predominantly

nuclear localisation, with some diffuse cytoplasmic staining (Figure 5.3). In contrast, in

the absence of NKX3.1-V5 overexpression, GFP-RMND5A and GFP-RMND5B

exhibited a diffuse nuclear and cytoplasmic distribution (Section 4.2.6.4, 4.2.7.5), whilst

in cells expressing NKX3.1-V5 alone, NKX3.1 displayed a predominantly nuclear

localisation (Figure 5.3). In cells coexpressing NKX3.1-V5 and GFP-RMND5A or

GFP-RMND5B, NKX3.1-V5 exhibited a mainly cytoplasmic localisation which also

contrasted previous reports and findings in our laboratory, including those observed in

this study (in the absence of RMND5 protein overexpression) where NKX3.1 and

NKX3.1-V5 were predominantly nuclear (Asatiani et al., 2005, Dawson, 2006). In these

experiments, colocalisation of NKX3.1 with GFP-RMND5A or GFP-RMND5B was

evident predominantly in the cytoplasm of LNCaP cells (Figure 5.3). To investigate

whether the reduced nuclear localisation of NKX3.1-V5 in RMND5A/RMND5B

overexpressing cells resulted from its increased ubiquitin-mediated degradation by the

proteasome, these experiments were repeated using LNCaP cells treated with 10µM

MG132, a proteasome inhibitor (Section 5.2.3) for the final 3 hours of culture prior to

preparation of the cells for viewing by microscopy at 48 hours post-transfection

(Section 3.1.3, 3.1.4, 3.15). Under these conditions, NKX3.1-V5 exhibited a similar


196

cytoplasmic localisation in cells co-expressing GFP-RMND5A/GFP-RMND5B, which

was predominantly nuclear (not shown). Under these culture conditions, GFP-

RMND5A/GFP-RMND5B and NKX3.1-V5 colocalisation occurred mainly in the

cytoplasm of LNCaP cells (not shown). The lack of MG132 effects on NKX3.1

localisation may be due in part to the short MG132 treatment period, however, longer

MG132 treatment of LNCaP cells growing on coverslips resulted in cytotoxic effects

due to proteasome inhibition (Section 5.2.3). Fluorescence microscopy to determine the

localisation and colocalisation of GFP-RMND5A (C356A/H358A) or GFP-RMND5B

(C358A/H360A) and NKX3.1-V5 determined that similar to wild-type GFP-RMND5

proteins, GFP-RMND5A (C356A/H358A) and GFP-RMND5B (C358A/H360A)

exhibited diffuse nuclear and cytoplasmic distribution, with GFP-RMND5B

(C358A/H360A) also exhibiting a punctate cytoplasmic appearance when expressed

alone (Section 3.1.3, 3.1.4, 3.15, 4.2.7.5). Coexpression of GFP-RMND5A

(C356A/H358A) or GFP-RMND5B (C358A/H360A) with NKX3.1-V5 identified that

NKX3.1-V5 exhibited a mainly cytoplasmic distribution whilst GFP-RMND5A

(C356A/H358A) and GFP-RMND5B (C358A/H360A) displayed a predominantly

nuclear cellular localisation with some cytoplasmic staining (not shown). GFP-

RMND5A (C356A/H358A) and GFP-RMND5B (C358A/H360A) colocalised with

NKX3.1-V5 predominantly in the cytoplasm of LNCaP cells (not shown), in a similar

manner to the colocalisation of wild-type GFP-RMND5A or GFP-RMND5B with

NKX3.1-V5. Furthermore, in cells coexpressing GFP-RMND5A (C356A/H358A) or

GFP-RMND5B (C358A/H360A) and NKX3.1-V5, 3 hours of treatment with 10µM

MG132 did not markedly alter the intracellular distribution or colocalisation of the

exogenously expressed proteins (not shown). These experiments may be further

optimised in future studies to allow for more extensive investigation of the effects of

RMND5 protein overexpression on the intracellular localisation of NKX3.1, including

the analysis of the fluorescence intensity of NKX3.1-V5 following proteasome

inhibition which may indicate its accumulation in the cell.

5.2.3 Regulation of NKX3.1 Expression in Prostate Cancer Cells

In order to determine the effects of RMND5 proteins on NKX3.1 expression, NKX3.1

protein half-life was initially investigated by determination of NKX3.1 levels following

treatment of cultures with 10µg/mL cycloheximide for 0 – 240 minutes (Section 3.1.5).


197

Figure 5.3: RMND5 proteins colocalise with NKX3.1 in LNCaP cells. To investigate the localisation and colocalisation of RMND5 proteins and NKX3.1, LNCaP cells growing on coverslips were transfected with plasmids encoding (A) NKX3.1-V5, (B) GFP-RMND5A and NKX3.1-V5, (C) GFP-RMND5B and NKX3.1-V5 or (D) untransfected. At 48 hours post-transfection, cells were fixed, permeabilised and stained for NKX3.1-V5 using an anti-V5 primary antibody and secondary anti-goat AlexaFluor® 546 antibody. Coverslips were viewed by fluorescence microscopy, identifying that GFP-RMND5A and GFP-RMND5B displayed a predominantly nuclear localisation with some diffuse cytoplasmic staining and that NKX3.1-V5 exhibited a predominantly nuclear localisation when expressed alone whilst when coexpressed with either GFP-RMND5A or GFP-RMND5B in the cell, NKX3.1-V5 displayed a predominantly cytoplasmic cellular localisation. Colocalisation of NKX3.1-V5 with GFP-RMND5A or GFP-RMND5B was predominantly detected in the cytoplasm of LNCaP cells (Magnification x1000). The experiment was performed three times and representative results are shown.

Overlay

NKX3.1-V5 GFP-RMND5A Hœchst 33258 Overlay

NKX3.1-V5 GFP-RMND5B Hœchst 33258 Overlay

B

C

NKX3.1-V5 488 channel Hœchst 33258 Overlay A

Hœchst 33258 No 1° Ab D Overlay


198

Under these culture conditions (RPMI/PS/10% FCS), the half-life of NKX3.1 was

estimated to be ~15.8 minutes, which is consistent with a previous report where the

NKX3.1 protein half-life was determined to be ~25 minutes (Section 3.1.5, 3.15, Figure

5.4) (Thomas et al., 2006).

Regulation of NKX3.1 levels by the proteasome has been reported previously (Guan et

al., 2006, Li et al., 2006), and to establish experimental conditions for these studies,

degradation of NKX3.1 by the proteasome and lysosome were examined. To

demonstrate NKX3.1 degradation by the proteasome, NKX3.1 levels were examined

following treatment of LNCaP cells with the proteasome inhibitors lactacystin and

MG132. For these experiments, 10µM lactacystin was added to cultures growing in

RPMI/PS/10%FCS for 8 hours prior to lysis, followed by NKX3.1 and β-actin western

blotting (Section 3.1.5, 3.15). Under these culture conditions, β-actin levels remained

relatively constant during the initial 6 hours of lactacystin treatment while NKX3.1

protein levels were not markedly altered until 6 hours of exposure to lactacystin (Figure

5.5). By 8 hours, both β-actin and NKX3.1 levels were decreasing and, due to the light

microscopic appearance of the cultures (not shown), it was likely that these results were

in part due to the cytotoxicity of the lactacystin.

A second proteasome inhibitor, MG132 was also utilised to investigate NKX3.1 protein

degradation by the proteasome and for these experiments LNCaP cells growing in 6

well plates in RPMI/PS/10%FCS were treated with 10µM MG132 for either 8 or 24

hours prior to western blotting for NKX3.1 and β-actin (Section 3.1.5, 3.15). In initial

experiments, NKX3.1 protein levels were similar in treated and untreated cultures (data

not shown), therefore since NKX3.1 expression is androgen regulated (He et al., 1997;

Prescott et al., 1998), NKX3.1 protein levels were monitored during androgen treatment

of LNCaP cells, which were grown in RPMI/PS/5%CSS for 24 hours to deplete the

cultures of androgens, then treated with 10-8M 5α-dihydrotestosterone (DHT) in

conjunction with 10µM MG132 for 8 or 24 hours (Section 3.1.5, 3.15). NKX3.1 protein

levels were low in cultures depleted of androgens (RPMI/PS/5%CSS) and in the

presence of androgens, NKX3.1 levels were markedly increased (not shown). As such,

accumulation of NKX3.1 protein levels due to proteasome inhibition were concealed by

the strong induction of NKX3.1 expression by DHT (not shown). Therefore, NKX3.1

protein levels were examined during androgen depletion of LNCaP cells cultured in

RPMI/PS/5%CSS and 10µM MG132 for 8 or 24 hours (Section 3.1.5, 3.15). Under


199

β-actin

NKX3.1

15 30 60 120 240

Cycloheximide

00.20.40.60.8

11.2

0 15 30 60 120 240

Norm

alise

dN

KX

3.1

Pro

tein

Lev

els

Cycloheximide Treatment (minutes)

(minutes)

A

B

0

Figure 5.4: Determination of NKX3.1 half-life. The half-life of NKX3.1 in LNCaP cells was estimated by treatment of cultures with 10µg/mL cycloheximide for 0-240 minutes to inhibit new protein production. (A) Cells were lysed at the indicated time points and the lysates electrophoresed in 12% polyacrylamide gels for NKX3.1 and β-actin western blotting. (B) Quantitation of normalised NKX3.1 protein levels indicated that the half-life of NKX3.1 was ~15.8 minutes under these culture conditions. The experiment was performed twice and representative results are shown.


200

(hours)NKX3.1

β-actin

~30kDa

~44kDa

2 4 6 8

0

0.5

1

1.5

Vehicle 2 4 6 8

Nor

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ised

NK

X3.

1 P

rote

in

Leve

ls

10µM Lactacystin (hours)

Lactacystin

β-actin

NKX3.1

8 24 8 24

Control MG132

0

1

2

3

4

5

Vehicle 10µM MG132

Nor

mal

ised

N

KX

3.1

Pro

tein

Le

vels

8 24

~30kDa

~44kDa

(hours)

hours

A

B

0

0

Control

Figure 5.5: Degradation of NKX3.1 by the proteasome. LNCaP cells were treated with 10µM of the proteasome inhibitors, lactacystin or MG132. Following treatment, cells were harvested, lysates were electrophoresed in 12% polyacrylamide gels and western blotting was performed for endogenous NKX3.1 and β-actin. (A) When cells were cultured in RPMI/PS/10% FCS, NKX3.1 protein levels were not altered during 2-4 hours of lactacystin treatment, but were reduced to ~40% of controls by 8 hours. (B) When LNCaP cells were cultured in RPMI/PS/5%CSS for 8 hours, NKX3.1 protein levels were initially low due to androgen deprivation, but were increased following 8 hours of culture in RPMI/PS/5%CSS with MG132 treatment. By 24 hours of culture with MG132, NKX3.1 levels were decreased, potentially due to nonspecific toxic effects of MG132. Experiments were performed twice and representative results are shown.


201

these culture conditions, NKX3.1 remained at low levels in vehicle treated cells (as

expected due to androgen depletion) however, following 8 hours of MG132 treatment,

NKX3.1 protein levels were markedly increased (Figure 5.5). This result is therefore

consistent with findings in the literature that NKX3.1 undergoes proteasomal

degradation. By 24 hours of MG132 treatment, both NKX3.1 and β-actin levels were

reduced and, similar to cultures treated with lactacystin for longer time periods, there

was evidence of cytotoxicity owing to long term proteasome inhibition. For these

reasons, LNCaP cells were treated with lactacystin and MG132 for only short periods of

time.

To assess whether NKX3.1 may also undergo degradation by the lysosome, NKX3.1

protein levels were monitored following treatment of cells with two lysosome

inhibitors, NH4Cl or choloroquine. For these experiments, LNCaP cells growing in 6

well plates were treated with 10mM or 20mM NH4Cl for either 6 or 24 hours prior to

western blotting for NKX3.1 and β-actin (Section 3.1.5, 3.15). Under these culture

conditions, NKX3.1 levels were reduced to similar levels at 6 and 24 hours of treatment

with 10mM or 20mM NH4Cl, results that were in part due to nonspecific cytotoxic

effects of NH4Cl on LNCaP cells (Figure 5.6). Preliminary testing of chloroquine

indicated an optimum concentration of 25µM, with 50µM or 100µM chloroquine

treatments resulting in marked cytotoxicity and large reductions in NKX3.1 protein

levels (Section 3.1.5, 3.15). For these experiments, NKX3.1 protein levels in LNCaP

cells treated with 25µM chloroquine for up to 48 hours were found to increase ~1.5 fold

following 6 hours of chloroquine treatment (Figure 5.6). The increased NKX3.1 levels

were maintained until 48 hours of treatment, providing evidence that NKX3.1

degradation may also be regulated in part by the lysosome (Figure 5.6). NKX3.1 protein

levels were similarly monitored in cultures that had been androgen depleted in

RPMI/PS/5%CSS for 24 hours prior to the addition of 10-8M DHT and 10mM or 20mM

NH4Cl, or 25μM chloroquine for 6 or 24 hours (Section 3.1.5, 3.15). However similar to

proteasome inhibition, changes in NKX3.1 protein levels due to lysosome inhibition

were not evident due to the rapid accumulation of NKX3.1 protein after DHT treatment

(not shown). NKX3.1 levels were then monitored during androgen withdrawal by

culture of LNCaP cells in RPMI/PS/5%CSS with lysosome inhibition using 10mM or

20mM NH4Cl, or 25μM chloroquine for 6 or 24 hours (Section 3.1.5, 3.15). In these

experiments, low levels of NKX3.1 were evident in vehicle treated cells (as expected


202

(Hours)

Control10mM NH4Cl

20mM NH4Cl

6 6 624 24 24

NKX3.1

β-actin

~30kDa

~44kDa

β-actin

4 6 8 24 482

NKX3.1~30kDa

~44kDa

Chloroquine

00.40.81.21.6

2

Nor

mal

ised

NK

X3.

1 P

rote

in L

evel

s

25µM Chloroquine (hours)

0

0.5

1

1.5

Control 10mM NH4Cl20mM NH4Cl

Nor

mal

ised

N

KX

3.1

Pro

tein

Le

vels

6 hours 24 hours

(hours)

A

B

10mM NH4Cl

20mM NH4Cl

0

Figure 5.6: Lysosomal processing of NKX3.1. To determine whether NKX3.1 was degraded by the lysosome, endogenous NKX3.1 levels were monitored following treatment of LNCaP cells with (A) 10mM or 20mM NH4Cl for 6 or 24 hours, or (B) 25µM chloroquine for 2-48 hours. Following treatment, cells were harvested and lysates electrophoresed in 12% polyacrylamide gels then analysed by western blotting for NKX3.1 and the housekeeping protein β-actin. NKX3.1 levels were reduced at 6 and 24 hours of NH4Cl treatment, whilst cells treated with chloroquine showed a ~1.5-fold increase in NKX3.1 protein levels at 6 hours which was maintained until 48 hours of treatment. Experiments were performed twice and representative results are shown.


203

due to androgen depletion) and upon lysosome inhibition NKX3.1 protein levels

remained at low levels compared to controls at both time points, providing no additional

evidence for lysosomal degradation of NKX3.1 (not shown). The contribution of the

lysosome in the regulation of NKX3.1 levels was not investigated further for this thesis

and subsequent experiments examined the role of RMND5 proteins in proteasome-

mediated NKX3.1 degradation. However, lysosomal degradation of NKX3.1 may be

investigated in future studies to determine whether it plays a direct or indirect role in the

control of NKX3.1 expression.

5.2.4 RMND5 Protein Effects on NKX3.1 Protein Expression

5.2.4.1 RMND5A and RMND5B Reduce NKX3.1 Protein Levels

To determine whether NKX3.1 was a substrate of RMND5 protein ubiquitination,

NKX3.1 protein levels in LNCaP cells were monitored following overexpression of

either RMND5A or RMND5B. For these studies, LNCaP cells growing in 6 well plates

were transfected with increasing concentrations (0-4µg) of plasmids encoding GFP-

RMND5A or GFP-RMND5B, cultures were lysed at 48 hours post-transfection and

analysed by western blotting for GFP and endogenous NKX3.1 (Section 3.1.4, 3.15).

GFP western blotting confirmed increasing expression of GFP-RMND5A or GFP-

RMND5B in cultures transfected with increasing amounts of the expression plasmids

(Figure 5.7). While western blotting for NKX3.1 identified a band at the expected size

of ~30kDa in lysates from GFP-RMND5A and GFP-RMND5B overexpressing cells,

NKX3.1 levels were reduced as the concentration of GFP-RMND5A or GFP-RMND5B

increased. These results indicated that overexpression of either RMND5A or RMND5B

downregulated NKX3.1 levels (Figure 5.7).

To investigate whether the reduction in NXK3.1 levels following RMND5

overexpression was due to its proteasomal degradation, LNCaP cells transfected with

plasmids encoding GFP-RMND5A or GFP-RMND5B were treated with 10µM MG132

for the final 6 hours prior to harvesting of cells at 48 hours post-transfection and

western blotting for GFP and endogenous NKX3.1 (Section 3.1.4, 3.1.5, 3.15). NKX3.1

levels were reduced following either GFP-RMND5A or GFP-RMND5B

overexpression, with these effects partially reversed when GFP-RMND5A or GFP-

RMND5B overexpressing cells were treated with MG132 (Figure 5.8). GFP-RMND5A

and GFP-RMND5B were also increased in MG132 treated cultures, indicating that they


204

Figure 5.7: Overexpression of RMND5A or RMND5B reduces NKX3.1 levels. LNCaP prostate cancer cells were transfected with increasing concentrations of plasmids encoding either (A) GFP-RMND5A or (B) GFP-RMND5B and at 48 hours post-transfection, cells were harvested for western blotting. GFP western blotting confirmed increasing expression of GFP-RMND5 proteins, whilst NKX3.1 protein levels were reduced as GFP-RMND5A and GFP-RMND5B expression increased.


205

- + - + - +Control

pEGFP-RMND5A

pEGFP-RMND5B

10µM MG132

NKX3.1

GFP

β-actin

0

0.5

1

1.5

2

2.5

Vehicle RMND5A RMND5B

Norm

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dNK

X3.1

Pr

otei

n Le

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Ethanol MG132

~30kDa

~44kDa

~74kDa

NKX3.1

GFP

β-actin

- + - + - +Control

pEGFP-RMND5A

pEGFP-RMND5B

10µM MG132~30kDa

~44kDa

~74kDa

0

0.4

0.8

1.2

1.6

Vehicle RMND5A RMND5B

Norm

alise

dNK

X3.1

Pr

otei

n Le

vels Ethanol MG132

A

B

Control

Control

Vehicle

Vehicle

Figure 5.8: Proteasome inhibition restores NKX3.1 protein levels following RMND5 overexpression. LNCaP cells were transfected with plasmids encoding GFP-RMND5A or GFP-RMND5B and at 42 hours post-transfection the medium was replaced with (A) RPMI/PS/10%FCS or (B) RPMI/PS/5%CSS and the culture treated with 10µM MG132 as indicated. Cells were harvested at 48 hours post-transfection and analysed by western blotting for NKX3.1, GFP-RMND5A or GFP-RMND5B. Under both conditions, NKX3.1 protein levels were reduced following RMND5 overexpression and NKX3.1 protein levels were increased following MG132 treatment, as were the protein levels of GFP-RMND5A and GFP-RMND5B. Experiments were performed twice and representative results are shown.


206

too are regulated by proteasomal degradation (Figure 5.8). As NKX3.1 expression is

regulated by androgens, the above experiment was repeated and the cells were grown in

medium depleted of androgens (RPMI/PS/5%CSS) for the final 6 hours of culture,

during which time they were treated with MG132 (Section 3.1.4, 3.1.5, 3.15). Western

blotting of the lysates for NKX3.1 determined that upon androgen depletion, NKX3.1

protein levels were low, consistent with previous experiments (Section 5.2.3, Figure

5.8). Under these conditions, the reductions in NKX3.1 protein levels following GFP-

RMND5A or GFP-RMND5B overexpression were diminished or not evident, however

the accumulation of NKX3.1 in MG132-treated GFP-RMND5A and GFP-RMND5B

overexpressing or control cultures was enhanced (Figure 5.8). Together, these results

provided evidence that RMND5 proteins were involved in the targeting of NKX3.1 for

degradation by the proteasome, and that RMND5 proteins are themselves degraded by

the proteasome.

5.2.4.2 RMND5A (C356S and C356A/H358A) and RMND5B (C358S and

C358A/H360A) Reduce NKX3.1 Protein Levels

To determine the effects of mutant RMND5A (C356S and C356A/H358A) and

RMND5B (C358S and C358A/H360A) on NKX3.1 expression, NKX3.1 protein levels

were monitored following transfection of LNCaP cells with 4µg pEGFP-RMND5A,

pEGFP-RMND5A (C356S), pEGFP-RMND5A (C356A/H358A), pEGFP-RMND5B,

pEGFP-RMND5B (C358S) or pEGFP-RMND5B (C358A/H360A). Cells were lysed at

48 hours post-transfection and GFP-RMND5 protein and endogenous NKX3.1 levels

were determined by western blotting (Section 3.1.5, 3.15). In these experiments, GFP

western blotting resulted in the identification of ~70kDa bands in lysates from all

cultures transfected with GFP-RMND5 proteins, confirming expression of both wild-

type and mutant GFP-RMND5A and GFP-RMND5B (Figure 5.9). NKX3.1 was

expressed in all cultures, with NKX3.1 protein levels reduced following wild-type

RMND5 protein overexpression, consistent with previous findings (Figure 5.9).

Surprisingly, the reduction in NKX3.1 protein levels was greater in lysates expressing

mutant GFP-RMND5A (C356S or C356A/H358A) or GFP-RMND5B (C358S or

C358A/H360A). Although the western blotting technique is only semi-quantitative and

these results were not evaluated statistically, NKX3.1 levels in cultures overexpressing

mutant RMND5 proteins were consistently lower than NKX3.1 levels in cultures

overexpressing wild-type RMND5 proteins (and control cultures). Therefore, while


207

NKX3.1

GFP

β-actin

pEGFP-RMND5A

pEGFP-RMND5B

WT WT

~30kDa

~70kDa

~44kDa

0

0.2

0.4

0.6

0.8

1

1.2

Untransfected RMND5A RMND5B

Nor

mal

ised

NKX

3.1

Prot

ein

Leve

ls WT

(C356S)/(C358S)

(C356A/H358A)/(C358A/H360A)

Figure 5.9: Overexpression of wild-type and mutant RMND5 proteins reduces NKX3.1 levels. LNCaP cells were transfected with plasmids encoding GFP-RMND5A, GFP-RMND5B, GFP-RMND5A (C356S), GFP-RMND5B (C358S), GFP-RMND5A (C356A/H358A) or GFP-RMND5B (C358A/H360A) and at 48 hours post-transfection cells were harvested and analysed by western blotting for NKX3.1, GFP and β-actin. Endogenous NKX3.1 protein levels were reduced by >30% following overexpression of wild-type or mutant GFP-RMND5 proteins. Experiment was performed twice and representative results are shown.


208

overexpression of RMND5 proteins reduced NKX3.1 protein levels, mutant RMND5

proteins resulted in greater decreases in NKX3.1 protein levels.

5.2.5 NKX3.1 is Ubiquitinated in LNCaP Cells

5.2.5.1 RMND5 Proteins Ubiquitinate NKX3.1

To determine whether RMND5A and RMND5B ubiquitinate NKX3.1, targeting it for

degradation by the proteasome, in vivo ubiquitination assays were carried out. For these

experiments, LNCaP cells were cotransfected with plasmids encoding NKX3.1-V5,

HA-ubiquitin and either GFP-RMND5A or GFP-RMND5B, and 48 hours post-

transfection, cells were lysed, input lysate samples were collected and NKX3.1-V5 was

immunoprecipitated with anti-V5 antibodies (Section 3.1.4, 3.13). Immunoprecipitation

products were electrophoresed in 4-12% gradient polyacrylamide gels and analysed by

HA (ubiquitin), V5 (NKX3.1) and GFP (RMND5A/RMND5B) western blotting

(Section 3.15). Western blotting of the input lysate samples for NKX3.1-V5 produced

bands at the expected size of ~35kDa in all lysates, and GFP western blotting identified

GFP-RMND5A and GFP-RMND5B in cultures transfected with the respective plasmids

(with the GFP-RMND5B protein band exhibiting low intensity due to dilution in the

total cell lysate) (Figure 5.10 (i)). In V5-immunoprecipitated samples, the presence of

~35kDa bands following V5 western blotting indicated successful immunoprecipitation

of V5-tagged NKX3.1. In the absence of GFP-RMND5A or GFP-RMND5B

overexpression, HA-ubiquitin western blotting detected multiple bands ranging in size

from ~35kDa – 175kDa, consistent with the presence of ubiquitinated NKX3.1 in the

V5 immunoprecipitation samples, however few bands corresponding to ubiquitinated

NKX3.1-V5 were detected when GFP-RMND5A or GFP-RMND5A was overexpressed

(Figure 5.10 (i)). To verify that this was not due to failure of transfections with the HA-

Ubiquitin expression plasmid, HA western blotting was performed on the input lysates

from these experiments, demonstrating that all transfections were successful (Section

3.15, Figure 5.10 (i)). These results suggested that although NKX3.1 is ubiquitinated in

vivo, overexpression of RMND5 proteins resulted in their rapid degradation, markedly

reducing detection of ubiquitinated NKX3.1 in the cultures.

To investigate this hypothesis, the experiments were repeated and cells were treated

with 10µM MG132 for the final 3 hours prior to harvest to allow the accumulation of

ubiquitinated proteins (Section 3.1.4, 3.1.5, 3.13). In these experiments, V5-


209

Figure 5.10: Ubiquitination of NKX3.1 in vivo by RMND5A and RMND5B. (i) LNCaP cells were cotransfected with plasmids encoding NKX3.1-V5, HA-Ubiquitin and GFP-RMND5A or GFP-RMND5B, and at 48 hours post-transfection NKX3.1-V5 was immunoprecipitated using anti-V5 antibodies. Western blotting for HA-ubiquitin demonstrated that NKX3.1 was ubiquitinated in vivo and that there was a reduction in NKX3.1-associated ubiquitination following RMND5A or RMND5B overexpression that was potentially due to its rapid degradation in the proteasome. (ii) The experiment was repeated with cells treated with the proteasome inhibitor 10µM MG132 for the final 3 hours of culture prior to immunoprecipitation of NKX3.1-V5 with anti-V5 antibodies at 48 hours post-transfection. Western blotting for HA-ubiquitin indicated an increase in NKX3.1-associated ubiquitination following RMND5A and RMND5B overexpression. Experiments were performed twice and representative results are shown.

Input lysate (2%)

+ + + - + + + +

- - - -

- - +

+

- - - -

IP

HA (Ubiquitin, ~35 – 175kDa)

V5 (NKX3.1, ~35kDa)

NKX3.1-V5

GFP-RMND5A

GFP-RMND5B

HA-Ubiquitin

10µM MG132 + + + +

+ + + - + + + +

- - - -

- - +

+

HA (Ubiquitin, 20 – 200kDa)

V5 (NKX3.1, ~35kDa)

GFP (RMND5A/RMND5B, ~70kDa)

(i) (ii)


210

immunoprecipitated and total cellular input samples were electrophoresed in 4-12%

gradient polyacrylamide gels and western blotting was performed for HA (ubiquitin),

V5 (NKX3.1) and GFP (RMND5A/RMND5B) (Section 3.13, 3.15). GFP western

blotting of the total cellular inputs identified GFP-RMND5A and GFP-RMND5B

expression in lysates of cells transfected with the appropriate plasmids and V5 western

blotting of all total cellular inputs and V5 immunoprecipitation samples indicated

successful transfection and immunoprecipitation of the ~35kDa NKX3.1-V5 protein,

respectively (Figure 5.10 (ii)). Following proteasome inhibition by MG132,

accumulation of high molecular weight HA-ubiquitin corresponding to ubiquitinated

and polyubiquitinated proteins was evident in the total cellular input fractions and HA-

ubiquitinated NKX3.1 (or NKX3.1-associated proteins) were evident in V5

immunoprecipitated samples (Figure 5.10 (ii)). In contrast to experiments performed in

the absence of MG132 treatment (Figure 5.10 (i)), the levels of HA-ubiquitinated

NKX3.1 (or NKX3.1-associated proteins) in GFP-RMND5A and GFP-RMND5B

overexpressing cells were markedly increased in comparison to cells that did not

overexpress RMND5 proteins (Figure 5.10 (ii)). These results indicated that both

RMND5A and RMND5B promoted the ubiquitination of NKX3.1, leading to its

proteasome-mediated degradation.

5.2.5.2 RMND5A (C356A/C358A) and RMND5B (C358A/H360A)

Ubiquitinate NKX3.1

Previous experiments had identified that overexpression of mutant RMND5A

(C356A/H358A) or RMND5B (C358A/H360A) was associated with reduced NKX3.1

protein levels (Section 5.2.4.2). To examine whether RMND5A (C356A/H358A) and

RMND5B (C358A/H360A) were able to ubiquitinate NKX3.1, NKX3.1 ubiquitination

was assessed following transfection of cells with plasmids encoding NKX3.1-V5, HA-

Ubiquitin and GFP-RMND5A, GFP-RMND5B, GFP-RMND5A (C356A/H358A) or

GFP-RMND5B (C358A/H360A) (Section 3.1.4). Cells were treated with 10µM MG132

for 3 hours prior to harvesting, total cellular input samples were taken and NKX3.1-V5

was immunoprecipitated with anti-V5 antibodies (Section 3.1.4, 3.1.5, 3.13).

Immunoprecipitates and total cellular input samples were electrophoresed in 4-12%

gradient polyacrylamide gels and analysed by western blotting for V5 (NKX3.1), HA

(Ubiquitin) and GFP (RMND5 proteins) (Section 3.15). Western blotting of the total

cellular input samples demonstrated that HA-ubiquitin, NKX3.1-V5 and (wild-type and


211

mutant) GFP-RMND5A proteins were expressed at the expected molecular weights in

the lysates from transfected cells, however (wild- type and mutant) GFP-RMND5B was

barely visible in the (dilute) input lysates samples (Figure 5.11). Western blotting of

lysates following V5 immunoprecipitation showed that NKX3.1-V5 was successfully

immunoprecipitated and HA-ubiquitin western blotting of the immunoprecipitated

samples identified smears of ubiquitinated proteins ranging in size from ~35kDa -

~175kDa (Figure 5.11). Ubiquitinated NKX3.1 (or NKX3.1 associated proteins) were

evident in all lysates, however NKX3.1 ubiquitination was enhanced in lysates of

cultures that overexpressed GFP-RMND5A, GFP-RMND5B or mutant GFP-RMND5B

(C358A/H360A), whilst a reduction in NKX3.1 ubiquitination following

overexpression of GFP-RMND5A (C356A/H358A) was observed (Figure 5.11). These

experiments demonstrated that mutation of key amino acid residues in RMND5A

(C356A/H368A) reduced its ability to ubiquitinate NKX3.1-V5, however similar

mutations in RMND5B (C358A/H360A) did not markedly alter its ability to

ubiquitinate of NKX3.1-V5.


212

Figure 5.11: In vivo ubiquitination of NKX3.1 following overexpression of wild-type and mutant RMND5 proteins. LNCaP cells were cotransfected with plasmids encoding NKX3.1-V5, HA-Ubiquitin and GFP-RMND5A, GFP-RMND5B, GFP-RMND5A (C356A/H358A) or GFP-RMND5B (C358A/H360A). At 48 hours post-transfection, NKX3.1-V5 was immunoprecipitated using anti-V5 antibodies. Western blotting for HA-ubiquitin showed that NKX3.1 was ubiquitinated in vivo and that there was an increase in NKX3.1-associated ubiquitination following RMND5A and RMND5B overexpression. HA western blotting also demonstrated that NKX3.1 ubiquitination was reduced following overexpression of GFP-RMND5A (C356A/H358A) compared to GFP-RMND5A. In contrast, levels of NKX3.1 ubiquitination following GFP-RMND5B (C358A/H360A) or GFP-RMND5B overexpression were similar. The experiment was performed twice and representative results are shown.

GFP (RMND5A/RMND5B, ~70kDa)

NKX3.1-V5

GFP-RMND5A

GFP-RMND5B

HA-Ubiquitin

MG132

+ + + - + + + +

-

-

- -

- -

+

+

GFP-RMND5A (C356A/H358A)

GFP-RMND5B (C358A/H360A)

+

+

- - - - - - -

- - - - - - -

+ +

+ +

+ + + + + +

HA (Ubiquitin, 35kDa – 175kDa)

V5 (NKX3.1, ~35kDa)

HA (Ubiquitin, ~20 – 200kDa)

V5 (NKX3.1, ~35kDa)

IP

Input Lysate (2%)


213

5.3 Discussion Expression of the prostatic tumour suppressor, NKX3.1 is reduced or undetectable in up

to 80% of prostate tumours, and although loss of heterozygosity at the NKX3.1 locus is

common, mutation or epigenetic inactivation of the remaining allele, which could

account for the loss of NKX3.1 expression is believed to be a rare occurrence (Voeller

et al., 1997; Bowen et al., 2000; Asatiani et al., 2005). Discordance between NKX3.1

mRNA and protein levels in prostate tumours and prostate cancer cell lines has led to

the hypothesis that aberrant post-translational modification of NKX3.1 may contribute

to the low protein levels detected in prostate tumour cells, however mechanisms

involved in this process have not been described (Xu et al., 2000; Bethel et al., 2006).

RMND5B was originally identified in our laboratory to interact with NKX3.1 in a yeast

two hybrid screen and this binding was initially confirmed using GST-pulldown and

coimmunoprecipitation assays (Dawson, 2006). In this thesis, RMND5B and its

homologue RMND5A were identified to function as E3 ubiquitin ligases (Chapter 4)

and therefore characterisation of RMND5 protein function was extended to investigate

whether NKX3.1 was a ubiquitination target of either or both RMND5 proteins.

Initially, interaction between RMND5B and NKX3.1 was confirmed and the interaction

between RMND5A and NKX3.1 was investigated using coimmunoprecipitation assays,

establishing that both RMND5 proteins interacted with NKX3.1 in LNCaP cells. In this

thesis, only immunoprecipitation assays were used to identify NKX3.1 interaction with

the RMND5 proteins although protein-protein interactions are usually confirmed using

a variety of methods. These most commonly include yeast two-hybrid assays, which

enable the identification of novel interactions as well as validation of protein-protein or

protein domain interactions, GST-pulldown, Far western blotting,

coimmunoprecipitation and bioluminscence resonance energy transfer (BRET) or

fluorescence resonance energy transfer (FRET) assays (Fuks et al., 2003; Ciruela et al.,

2010). GST-pulldown assays are useful in situations where expression of high levels of

a protein of interest is limiting, for example where the protein product inhibits

mammalian cell growth or reduces cell viability. However, as proteins are produced in

bacteria, posttranslational modifications such as phosphorylation, which may be

required for specific protein-protein interactions may not occur, limiting the usefulness

of the method (Sahdev et al., 2008). Although yeast-two hybrid assays are performed in

eukaryotic cells, cell type specific factors including cofactors required for protein-


214

protein interactions may not be present, therefore some interactions may not be detected

(Luo et al., 1997). Other limitations can include the generation of false positive binding

as interacting proteins in yeast may not be localised in the same cellular compartments

in mammalian cells (Luo et al., 1997). Far western blotting is similar to western blotting

but instead of using antibodies for the detection of a particular protein (bait), another

protein (prey) is utilised, thereby detecting specific protein-protein interactions (Wu et

al., 2007b). FRET and BRET rely on the transfer of energy from a

fluorescent/luminescent donor to a fluorescence acceptor to detect protein-protein

interactions where proteins are within 1-10nm of each other (Wu and Brand, 1994; Xu

et al., 1999; Ciruela et al., 2010). An advantage of BRET/FRET is that the techniques

can be utilised to determine protein-protein interactions in real time in living cells,

including interactions that are transient or unstable and which therefore would be

difficult to detect using coimmunoprecipitation or GST pulldown assays (Pfleger and

Eidne, 2006; Kocan et al., 2010). While the present studies have provided good

evidence for RMND5 protein interaction with NKX3.1 in prostate cancer cells, future

studies that characterise the protein-protein interaction domains of either protein and the

physiological conditions that promote or inhibit these interactions may use one or more

additional methods to investigate or confirm results.

Following identification of the interaction between RMND5A/RMND5B and NKX3.1,

fluorescence microscopy was used to determine their localisation/colocalisation and

therefore the potential location of their interaction in prostate cancer cells. The results

obtained were interesting as the localisation of both NKX3.1 and the RMND5 proteins

was altered upon co-expression of either RMND5 protein with NKX3.1. As discussed

previously, RMND5A and RMND5B generally display a diffuse nuclear and

cytoplasmic distribution (Section 4.2.6.4, 4.2.7.5) with a proportion of cells exhibiting a

punctate cytoplasmic localisation. NKX3.1 is an androgen regulated transcription factor

and as such it localises predominantly to the nucleus in the presence of androgens

(Bowen et al., 2007; Guan et al., 2008). Overexpression of NKX3.1 with either

RMND5A or RMND5B resulted in the cytoplasmic localisation of NKX3.1 and the

predominantly nuclear localisation of RMND5 proteins, although some colocalisation in

the cytoplasm was observed. NKX3.1 protein levels are reduced in prostate tumours,

however NKX3.1 cytoplasmic mislocalisation has also been hypothesised as a possible

contributor to loss of NKX3.1 function, a finding that is consistent with the results of

this study where overexpression of RMND5 proteins resulted in the cytoplasmic


215

mislocalisation of NKX3.1 (Kim et al., 2002b). The mechanism by which RMND5

proteins are localised to the nucleus and NKX3.1 is localised to the cytoplasm under

these culture conditions is unknown. It is feasible that nuclear RMND5A and RMND5B

promote the ubiquitination of NKX3.1 and its relocalisation to the cytoplasm for

degradation or an as yet unidentified cytoplasmic role. Alternatively, ubiquitinated

NKX3.1 may be degraded in the nucleus. As the functions of RMND5 proteins are

incompletely characterised, it is also possible that their effects on the intracellular

localisation of NKX3.1 do not involve their E3 ubiquitin ligase activity.

Mechanisms regulating the intracellular distribution of RMND5 proteins in cells

transiently overexpressing RMND5A or RMND5B with or without NKX3.1

overexpression are also unknown at this time, however it is conceivable that this

involves post-translational modification. RMND5 protein auto-ubiquitination which

was observed in this study, may result in their altered intracellular localisation. In

addition, the increased RMND5 protein levels detected following proteasome inhibition

indicate that RMND5 protein levels are regulated by proteasomal degradation, which

may be related to their auto-ubiquitination or to ubiquitination by other E3 ubiquitin

ligases. E3 ubiquitin ligase auto-regulation has been widely documented with MDM2

and TRAC-1 two examples of ubiquitin ligases that regulate their own protein levels by

auto-ubiquitination, thereby resulting in their own proteasome dependent degradation

(Fang et al., 2000; Giannini et al., 2008). The findings of this study that RMND5

proteins are able to mono- and poly-auto-ubiquitinate their RING domains suggests that

RMND5A and RMND5B may regulate their own activity, where for example

monoubiquitination may result in altered cellular function or localisation and

polyubiquitination could lead to proteasome-dependent degradation. The type of

ubiquitin linkages, which direct the fate of RMND5 proteins may be ascertained in

future studies with the use of linkage specific antibodies or ubiquitin mutants (Wu-Baer

et al., 2003; Newton et al., 2008).

RMND5 protein effects on NKX3.1 may be related to their level of expression

following transient transfection of LNCaP cells. Such findings have been observed for

MDM2, which ubiquitinates p53 and depending upon the levels of MDM2, the type of

ubiquitination and the outcome of p53 ubiquitination differ (Boyd et al., 2000, Geyer et

al., 2000, Li et al., 2003). When MDM2 protein levels are high, p53 is ubiquitinated

and targeted for nuclear degradation, however, lower levels of MDM2 result in p53


216

monoubiquitination and its nuclear export and cytoplasmic localisation where it is

unable to function as a transcription factor (Boyd et al., 2000; Geyer et al., 2000; Li et

al., 2003). Since the levels and cellular localisation of endogenous RMND5 proteins are

unknown, it is not possible at present to determine the contribution of RMND5A and/or

RMND5B to steady-state or androgen-induced NKX3.1 levels and localisation.

However, these may be investigated in future studies following development of

antibodies to the endogenous proteins (Section 7.2). Similarly, the dose-dependent

effects of RMND5 proteins on NKX3.1 levels and localisation may also be examined

using inducible methods of RMND5 protein overexpression. Interestingly, muskelin and

RanBPM, members of the CTLH complex (along with RMND5A) are hypothesised to

function in nucleocytoplasmic shuttling, and it is therefore feasible that changes in the

intracellular localisation of NKX3.1 following RMND5 overexpression are mediated in

part by the CTLH complex (Section 6.1.1, 6.1.4.3) (Kobayashi et al., 2007;

Valiyaveettil et al., 2008). Following RMND5 overexpression, a dose-dependent

reduction in NKX3.1 protein levels was observed, although changes in NKX3.1 mRNA

were not investigated but could be determined in future studies to assess whether

RMND5A and RMND5B regulate NKX3.1 solely at the protein level. As RMND5B

has been demonstrated to exert transcriptional repressor effects on an NKX3.1

responsive element, one of which is present in the NKX3.1 promoter, the nuclear

localisation of RMND5 proteins may reflect alternative cellular functions of RMND5A

and RMND5B (Dawson, 2006).

In order to examine RMND5 protein regulation of NKX3.1 levels, proteasomal and

lysosomal degradation of NKX3.1 were initially investigated. Culture of cells with the

proteasome inhibitor, lactacystin initially increased NKX3.1 protein levels, supporting

its processing by the proteasome, with progressive reduction in protein levels over eight

hours of treatment likely due to in part to nonspecific toxic effects of lactacystin or the

expected damaging effects of longterm disruption of the normal regulation of protein

processing. The concentration of 10µM lactacystin to inhibit the proteasome in LNCaP

cells has been commonly used in the literature, however duration of exposure to

lactacystin is generally limited due to the accumulation of nonspecific effects (Huang et

al., 2005b; Shirley et al., 2005; Chen et al., 2011a). Cells treated with a second

proteasome inhibitor MG132 did not show an accumulation of NKX3.1 under normal

culture conditions (RPMI medium supplemented with FCS) nor did cells cultured in

medium depleted of steroid hormones and supplemented with physiological levels of


217

the androgen metabolite DHT (RPMI supplemented with charcoal stripped FCS and

DHT), highlighting the importance of performing studies under experimental conditions

where proteasomal processing of the target protein is likely to be enhanced. Since

NKX3.1 is an androgen regulated gene, NKX3.1 protein levels were monitored

immediately following androgen depletion (RPMI medium supplemented with charcoal

stripped FCS) when NKX3.1 levels are known to decline rapidly (He et al., 1997;

Prescott et al., 1998). Again, the concentration of 10µM MG132 used was obtained

from previous studies (Huang et al., 2005b; Chen et al., 2011a) and under these culture

conditions, the rapid reduction of NKX3.1 protein levels was reversed following

MG132 treatment. These experiments therefore demonstrated that NKX3.1 was

degraded by the proteasome and that androgen deprivation of LNCaP cells promoted

proteasome-mediated degradation of NKX3.1. Proteasome inhibitors such as MG132

and ALLN, which are peptide aldehydes, predominantly inhibit the chymotrypsin-like

activity of the proteasome and generally do not affect cell viability or growth for 10-20

hours of treatment (Rock et al., 1994). However, peptide aldehydes can also inhibit

some lysosomal cysteine proteases and calpains, thereby affecting lysosomal activity

(Lee and Goldberg, 1998). Lactacystin is an irreversible proteasome inhibitor that is a

more complete inhibitor of proteasome activity than the peptide aldehydes due to its

ability to inhibit both the chymotrypsin and trypsin-like activities of the proteasome,

however it also inhibits cathepsin A, another lysosomal protease (Craiu et al., 1997).

Validation of the mechanism of protein degradation using more than one proteasome

inhibitor is important and in this study both lactacystin and MG132 were tested, with

results both comparable between the two treatments (Section 4.2.5, 4.2.6.5, 4.2.7.6) and

consistent with previous studies that identified regulation of NKX3.1 levels by

proteasomal degradation (Li et al., 2006; Guan et al., 2008).

Lysosomal degradation of NKX3.1 was examined in cells treated with the lysosome

inhibitors ammonium chloride and chloroquine, although both proteasome inhibitors

tested are also able to inhibit lysosomal activity to some extent. While studies using

ammonium chloride found no evidence of lysosomal processing of NKX3.1, in

experiments using chloroquine, the increased NKX3.1 protein levels suggested that

under specific culture conditions, NKX3.1 may be degraded by a lysosomal mechanism.

In contrast to results from the experiments testing proteasome inhibitors, there was no

evidence that lysosomal degradation mediated the rapid decrease in NKX3.1 levels

following removal of androgens from the culture medium. Both chloroquine and


218

ammonium chloride inhibit lysosomal acidification and thus to confirm that NKX3.1 is

able to undergo degradation by the lysosome, a lysosomal inhibitor that functions by

inhibiting lysosomal proteases such as leupeptin or E-64 may be utilised (Lee and

Goldberg, 1998). Findings in this thesis of degradation of NKX3.1 by the proteasome

are consistent with the canonical pathway in which up to 90% of short-lived cellular

proteins undergo proteasomal degradation (Hicke, 1999). In contrast, those proteins that

undergo lysosomal degradation are generally membrane bound proteins (Lee and

Goldberg, 1998; Hicke, 1999), although a number of proteins have been reported, for

example α-synuclein and connexin43, whose degradation is mediated by both protein

degradation pathways (Tofaris et al., 2001; Qin et al., 2003; Webb et al., 2003). Both

the proteasomal and lysosomal pathways of protein degradation are regulated by

ubiquitination and cross-talk between the two pathways of protein degradation has been

reported (Qiao and Zhang, 2009; Ciechanover, 2012). Inhibition of the proteasome

frequently leads to an increase in lysosomal proteases and therefore function, however

lysosomal inhibition most commonly results in a reduction in proteasomal function but

an increase in heat shock proteins, suggesting that lysosome inhibition enhances

autophagy that is mediated by chaperones (Rideout et al., 2004; Pandey et al., 2007;

Qiao and Zhang, 2009). Therefore, similar to other cellular proteins, it is feasible that

depending upon the cellular environment and the presence of NKX3.1 regulators,

NKX3.1 may be degraded by either the proteasome or lysosome, although proteasomal

degradation is likely to predominate.

Overexpression of RMND5A or RMND5B resulted in reduced NKX3.1 protein levels

that were partially restored upon proteasome inhibition, indicating that RMND5

proteins regulated cellular NKX3.1 levels and that this was in part mediated by

promotion of NKX3.1 ubiquitination and proteasomal degradation. This mechanism

was further supported by in vivo ubiquitination assays which determined that upon

RMND5 overexpression, the levels of ubiquitinated NKX3.1 were markedly enhanced.

NKX3.1 ubiquitination and proteasomal degradation may be mediated by a direct

interaction of RMND5A or RMND5B with NKX3.1 or may occur via an intermediate

RMND5 protein regulated pathway. For example, Runx2 is reported to be ubiquitinated

by the E3 ubiquitin ligase Smurf1, however deletion of the PY domain of Runx2 with

which Smurf1 interacts resulted in only a partial reduction of Runx2 degradation (Shen

et al., 2006). In this study it was determined that Smad6 interacted with both Smurf1

and Runx2 and enhanced Smurf1 induced Runx2 proteasomal degradation, providing a


219

mechanism by which Smurf1 induced Runx2 proteasomal degradation both directly and

indirectly (Shen et al., 2006). The protein-protein interaction domains of RMND5

proteins and NKX3.1 have not been mapped, however their elucidation would enable

mutation studies to be performed to differentiate between direct or indirect mechanisms.

Although endogenous RMND5A or RMND5B protein levels in LNCaP cells are

unknown due to the lack of suitable commercially available antibodies, further

confirmation of the involvement of RMND5 proteins in the normal regulation of

NKX3.1 may be achieved with the use of siRNA for knockdown of RMND5 protein

levels. Increased NKX3.1 levels following RMND5 knockdown, which has been

performed to confirm the effect of other E3 ubiquitin ligases on their substrates would

indicate an important role of RMND5 proteins in the regulation of NKX3.1 levels that

may have significant implications in normal physiology and in pathological conditions

including prostate cancer (Guan et al., 2008; Tatham et al., 2008). Whether the outcome

of RMND5 mediated ubiquitination of NKX3.1 solely results in the proteasomal

degradation of NKX3.1 remains to be determined. The type and therefore outcome of

NKX3.1 ubiquitination may be investigated by western blotting using the linkage

specific antibodies to characterise the specific ubiquitin linkages associated with

NKX3.1 during RMND5-mediated ubiquitination (Newton et al., 2008). Additionally,

in vitro ubiquitination assays could also be performed to verify NKX3.1 ubiquitination

by RMND5 proteins. This would require the immunoprecipitation of both

RMND5A/RMND5B and NKX3.1 from mammalian cells as neither full length GST

fusion protein is able to be purified in sufficient quantities from bacterial cells. An

advantage to this assay is that the type of NKX3.1-associated ubiquitin linkages

promoted by RMND5 proteins can be more specifically determined incorporating use of

ubiquitin mutants or linkage specific antibodies (Wu-Baer et al., 2003; Newton et al.,

2008).

Overexpression of mutant RMND5 proteins in LNCaP cells resulted in similar

reductions in NKX3.1 protein levels compared to that associated with overexpression of

wild-type RMND5 proteins. These unexpected results may reflect residual activity of

the mutant RMND5 proteins which was evident in in vitro ubiquitination assays using

RMND5A (C356A/H358A) and RMND5B (C358A/360A) RING domains (Section

4.2.7.7). Alternatively, the reduced E3 ubiquitin ligase activity of RMND5 proteins may

be compensated for in vivo by RMND5 binding partners that possess E3 ubiquitin ligase

activity or that augment the activity of RMND5 proteins in mammalian cells. For


220

example RMND5A forms part of the CTLH complex which contains a second putative

RING domain protein, EMP that may contribute E3 ubiquitin ligase activity to the

complex, thereby maintaining all or part of its E3 ubiquitin ligase activity following

mutation of RMND5A (Santt et al., 2008). E3 ubiquitin ligases may function as homo-

or heterodimers (Hashizume et al., 2001; Linke et al., 2008; Johnson et al., 2012) and

therefore the reduced enzymatic activity of the mutant RMND5 proteins may be

compensated for by the E3 ubiquitin ligase activity of its interacting partner, including

wild-type endogenous RMND5 proteins. The reduction in NKX3.1 protein levels upon

RMND5 mutant protein overexpression may also indicate that RMND5 proteins are

able to reduce NKX3.1 protein levels, in part, by a mechanism unrelated to their E3

ubiquitin ligase activity and that RING domain inactivation may enhance this activity.

Finally, the in vivo function of RMND5 proteins in relation to NKX3.1 ubiquitination

may be dependent on specific environmental conditions such as androgen withdrawal or

activity of signalling pathways, and therefore changes in NKX3.1 levels following

overexpression of mutant RMND5 proteins were not evident using the experimental

parameters employed for these studies.

The similar effects of mutant RMND5A and RMND5B on NKX3.1 levels in LNCaP

cells were expected to be reflected by similar levels of NKX3.1 ubiquitination in cells

overexpressing each of RMND5A (C356A/H358A) and RMND5B (C358A/H360A).

However, under the experimental conditions used, overexpression of RMND5A

(C356A/H358A) was associated with reduced levels of ubiquitinated NKX3.1, while

overexpression of RMND5B (C358A/H360A) was associated with similar levels of

NKX3.1 ubiquitination compared to LNCaP cells overexpressing wild-type RMND5B.

These findings for RMND5A (C356A/C358A) are consistent with previous studies

where mutation of E3 ubiquitin ligases typically results in a reduction in substrate

ubiquitination, including the finding by Santt et al. (2008) that mutant RMD5 (C379S)

no longer ubiquitinates its substrate FBPase (Qiu et al., 2000). Thus the experiments

performed for this thesis provided evidence that NKX3.1 is a ubiquitination target of

RMND5A and to confirm that RMND5A is able to ubiquitinate NKX3.1, in vitro

ubiquitination assays could be carried out in future studies using full length RMND5A

and RMND5A (C356A/H358A). Ubiquitination of NKX3.1 by RMND5A and reduced

levels of NKX3.1 ubiquitination by RMND5A (C356A/H358A) would verify that the

specific RING domain mutations introduced reduced activity toward this substrate as

has been demonstrated in the literature for further E3 ligases (Ryu et al., 2011).


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Contrary to results of in vitro ubiquitination assays and to its association with

ubiquitinated proteins in vivo (Section 4.2.7.6, 4.2.7.7), overexpression of the RMND5B

(C358A/H360A) mutant did not alter NKX3.1 ubiquitination compared to wild-type

RMND5B in LNCaP cells. While the reasons for this cannot be ascertained without

further studies, it is feasible that this may be due to the maintenance of residual E3

ubiquitin ligase activity of the RMND5B (C358A/H360A) mutant which was evident in

in vitro ubiquitination assays and supported by the reduction in NKX3.1 protein levels

following RMND5B (C358A/H360A) overexpression. Mutant RMND5B may interact

with another E3 ubiquitin ligase that is able to ubiquitinate NKX3.1, allowing

accumulation of ubiquitinated NKX3.1 to similar levels that accompany overexpression

of wild-type RMND5B. In these studies, the levels of ubiquitinated NKX3.1 were

determined 48 hours after transfection of cells to overexpress RMND5B

(C358A/H360A), and it is also feasible that alterations in NKX3.1 ubiquitination are

more clearly evident at earlier time points. Although RMND5A and RMND5B are

highly homologous proteins that are likely to have overlapping substrates, a proportion

of their targets are expected to be unique. Similarly, the mutations introduced into each

of RMND5A and RMND5B may have differentially affected their affinity for NKX3.1

and the efficiency of their ability to ubiquitinate NKX3.1. Each of these aspects of

RMND5 protein activity may be evaluated in future studies using in vitro and in vivo

binding and ubiquitination assays.

The regulation of NKX3.1 at the protein level and the mechanisms by which NKX3.1

protein levels are reduced in prostate tumours are not well understood and it is likely

that a number of proteins including protein kinases and E3 ubiquitin ligases regulate the

post-translational modification and degradation of NKX3.1. The E3 ubiquitin ligase,

TOPORS plays a role in NKX3.1 ubiquitination and proteasomal degradation (Guan et

al., 2008), and results of this thesis indicate that both RMND5 proteins and TOPORS

may regulate NKX3.1 levels in the prostate, potentially under different environmental

conditions. Many cellular proteins including p53 are ubiquitinated by more than one E3

ubiquitin ligase resulting in their degradation by the proteasome, and deregulation of

individual E3 ubiquitin ligases may produce minimal or profound changes in the

regulation of cellular levels of the target protein due to contributions of the other E3

ubiquitin ligases (Scheffner et al., 1993; Kubbutat et al., 1997; Esser et al., 2005).

Although it is unknown whether the RMND5A and RMND5B gene loci are themselves

disrupted in prostate cancer, the RMND5B gene is located at chromosome 5q35, a


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prostate cancer heritability locus, and the chromosomal regions of both RMND5A and

RMND5B are disrupted in a number of cancer types, with RMND5A overexpressed in

ovarian cancer (Xu et al., 2005; Li et al., 2008; Christensen et al., 2010). In the absence

of well-characterised RMND5 antibodies, the effects of RMND5 loss of expression or

overexpression on NKX3.1 expression in human prostate tumours remain to be

determined. Previous studies have shown that the TOPORS chromosomal locus

undergoes LOH in 21.8%-50% of prostate tumours, however as TOPORS levels or

function have not been characterised in human prostate tumours, the relationship

between TOPORS LOH and reduced NKX3.1 levels are unknown (Perinchery et al.,

1999). It is also likely that the reduced NKX3.1 levels seen in prostate tumours results

from the dysregulation of other proteins that mediate NKX3.1 post-translational

modifications and stability, including CK2, which phosphorylates NKX3.1, increasing

its half-life (Li et al., 2006). The chromosomal loci encoding the CK2α and α’ catalytic

subunits are deleted in a proportion of prostate tumours, with further studies required to

determine correlations between alterations in CK2 expression or activity and NKX3.1

levels (Best et al., 2005; Jin et al., 2011).

Results from these studies have therefore identified that both RMND5A and RMND5B

interact with NKX3.1 and that NKX3.1 is a ubiquitination target of RMND5 proteins in

prostate cancer cells. Although aberrant RMND5 expression has not been investigated

in prostate tumours, the findings that RMND5A is overexpressed in ovarian tumours and

that the RMND5A and RMND5B loci are amplified in a number of cancers suggests that

RMND5 overexpression may contribute, in part, to the reduced NKX3.1 levels present

in prostate tumours. As NKX3.1 expression is largely restricted to the prostate in the

adult and according to the NCBI Gene Expression Omnibus Database, RMND5 proteins

are ubiquitously expressed, it is likely that RMND5 proteins have additional cellular

substrates and binding partners. These binding partners may be substrates of

RMND5A/RMND5B induced ubiquitination or may be involved in additional activities

of RMND5 proteins that are potentially mediated by the LisH, CTLH and CRA

domains found in both proteins. To further characterise RMND5 protein function,

additional RMND5 protein binding partners were therefore investigated, in both the

context of the previously reported CTLH complex and in the identification of novel

interactors.

Chapter 6 Characterisation of RMND5 Binding Partners

Chapter 6: Characterisation of RMND5

Protein Binding Partners

Chapter 6 Characterisation of RMND5 Protein Binding Partners

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6.1 Introduction The CTLH complex is the proposed human orthologue of the yeast E3 ubiquitin ligase

Vid30 complex (Santt et al., 2008). Although the biological activity of the RMND5A-

containing CTLH complex has not yet been characterised, the function of RMND5A as

an E3 ubiquitin ligase and the protein domain architecture of other CTLH complex

components suggests that this complex may also function as an E3 ubiquitin ligase

complex.

6.1.1 Characterisation of the CTLH Complex Components

RanBPM was originally identified to form part of a large 670kDa complex, with the

GTPase Ran coimmunoprecipitating with RanBPM as part of this complex (Nishitani et

al., 2001). Yeast two-hybrid studies characterising the members of the RanBPM

associated complex identified the muskelin, Twa1 and matrix metalloprotease 8

(HSMpp8) gene products as RanBPM binding partners, however only muskelin and

Twa1 were confirmed as complex members by subsequent co-immunoprecipitation and

gel filtration analyses (Umeda et al., 2003). Although Twa1 was described as a

predominantly nuclear protein, co-expression of muskelin with either Twa1 or RanBPM

resulted in the diffuse cytoplasmic and nuclear redistribution of Twa1 and RanBPM,

indicating the potential in vivo interaction between these proteins (Umeda et al., 2003).

In an effort to identify additional complex components, Kobayashi et al. (2007)

performed linear sucrose gradient centrifugation and immunoprecipitation of

endogenous RanBPM from HEK293 cells and identified the co-immunoprecipitating

proteins muskelin, p48EMLP (EMP/MAEA), p44CTLH (RMND5A) and the armadillo

repeat containing proteins ARMC8α and ARMC8β by mass spectrometry (Kobayashi et

al., 2007). Due to the finding that five of the CTLH complex components contained

similar protein domain architectures, including LisH and CTLH domains, the complex

was named the CTLH complex (Figure 6.1) (Kobayashi et al., 2007).

Three of the CTLH complex components were novel proteins including ARMC8α and

ARMCβ, alternatively spliced products of the same gene, with ARMC8α encoding the

larger isoform which was identified to co-immunoprecipitate with the other complex

components and with p44CTLH/RMND5A (Kobayashi et al., 2007). Although Twa1

was identified in the original screen, it was not detected as coimmunoprecipitating with

RanBPM and the associated CTLH complex, but was shown to interact with each


224

complex component in coimmunoprecipitation assays (Kobayashi et al., 2007). Using

these assays, it was determined that all complex components interacted with RanBPM,

Twa1, muskelin and ARMC8α and that these members were also able to self-associate

(Section 4.1.3.1) (Kobayashi et al., 2007). It was further demonstrated in

coimmunoprecipitation assays that RMND5A and p44EMLP interacted, that the

RMND5A amino-terminal and carboxy-terminal regions were necessary for this

interaction and that the CTLH domain was required for the interaction between

RMND5A and ARMC8α (Kobayashi et al., 2007). Fluorescence microscopy indicated

that endogenous RMND5A, ARMC8α, Twa1 and RanBPM displayed diffuse nuclear

and cytoplasmic localisation in HEK293 cells, p44EMLP exhibited a nuclear

distribution whilst muskelin was mainly cytoplasmic (Kobayashi et al., 2007). Although

individually not all components were expressed in the same cellular compartment, when

co-expressed, the cellular distribution of CTLH complex members was altered and the

proteins colocalised either in the nucleus or the cytoplasm or both (Kobayashi et al.,

2007).

6.1.2 Protein Domain Architecture of the CTLH Complex Members

Each of the CTLH complex components contains a similar protein domain architecture,

notably a LisH, CTLH and CRA domain, and it has been proposed that the proteins

utilise one or more of these domains to associate with each other (Figure 6.1). For

example, it has been demonstrated that RMND5A uses its CTLH domain and part of the

CRA domain to interact with ARMC8α (Kobayashi et al., 2007). As described

previously (Section 4.1.3.1), the LisH domain is a dimerisation motif and therefore

could be used by CTLH complex members to associate with each other. Alternatively,

the domain may be utilised for complex multimerisation in a similar manner to that of

the LisH domain of DCAF4, which forms Cullin4A RING E3 ubiquitin ligase

supramolecular complexes via its LisH domain (Ahn et al., 2011). The functions of the

CTLH and CRA domains are not well characterised, however they are also proposed

protein-protein interaction domains (Section 4.1.3.2, 4.1.3.3). Several CTLH complex

members contain additional protein domains including RMND5A which contains a

RING domain, RanBPM, which contains a SPRY domain, muskelin which contains a

discoidin-like and a Kelch repeat domain and ARMC8α/β, which possess Armadillo

repeat domains (Figure 6.1).


225

.

The SplA and ryanodine receptor (SPRY) domain is a protein-protein interaction

domain originally identified in the Dictyostelium discoideum tyrosine kinase spore lysis

A (SplA) and the mammalian Ryanodine receptor (Ponting et al., 1997). The domain is

found in a number of proteins including the tripartite motif (TRIM) RING domain

containing E3 ubiquitin ligases and is responsible for substrate binding of TRIM21

(Stacey et al., 2012). Additionally, suppressor of cytokine signalling (SOCS) box

proteins, e.g. SPSB2, which contain SPRY and/or WD40 repeats, function as substrate

recognition components in Cullin-Rbx1 E3 ubiquitin ligase complexes, with the SPRY

domain responsible for substrate recognition and binding (Kuang et al., 2010; Linossi

and Nicholson, 2012). The SPRY domains of SPSB1 to 4, are also able to bind the

intracellular domain of the human growth factor receptor MET (Wang et al., 2005).

Interestingly, the CTLH complex member RanBPM and its homologue RanBP10 also

use their SPRY domains to interact with MET, implicating the SPRY domain in similar

functions in these proteins and suggesting that this domain of RanBPM/RanBPM10

may function as a substrate recognition element in the CLTH complex (Wang et al.,

2002a; Wang et al., 2004).

Figure 6.1: Protein domain architecture of the CTLH complex components. The CTLH complex components possess a similar domain structure that includes Lissencephaly 1 homology (LisH), C-terminal to LisH (CTLH) and CT-11 RanBPM (CRA) domains as well as unique domains such as SplA and Ryanodine receptor (SPRY), discoidin-like domains, Kelch repeats and Armadillo repeat domains (Kobayshi et al., 2007).

(RMND5A)


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The discoidin domain, which plays a role in membrane anchoring, is found in a variety

of extracellular and membrane bound proteins, with many being involved in cell

adhesion and migration, in particular during development (Baumgartner et al., 1998;

Arakawa et al., 2007). Kelch repeats form a β-propeller structure which is also formed

by the WD40 repeat domain and due to their similar structure, both domains possess

similar functions (Hudson and Cooley, 2008). WD40 repeats are known to bind

phosphoresidues on the substrate, however, this recognition activity of Kelch repeats is

uncharacterised (Pickart, 2001). Kelch and WD40 repeats are often present in F-box

proteins which are the substrate recognition components of SCF (SKP1, Cullin1-Rbx1,

F-box protein) E3 ubiquitin ligase complexes (Sun et al., 2007). The amino-terminal F-

box motif binds the SKP1 component of the complex whilst the carboxy-terminal

contains other protein-protein interaction motifs including the Kelch repeat or WD40

domain which bind the substrates of these complexes. For example, the F-box and

Kelch repeat containing protein Just one F-box and Kelch domain containing protein

(JFK) forms part of an SCF complex binding p53 through its Kelch domain and

resulting in its ubiquitination and targeting it for proteasomal degradation (Sun et al.,

2007; Sun et al., 2009). The Kelch repeat containing protein KLHL20 forms part of the

KLHL20-Cul3-Rbx E3 ubiquitin ligase complex and acts as the substrate recognition

component, using its Kelch repeat to interact with death associated protein kinase

(DAPK), resulting in the polyubiquitination of DAPK and its proteasome-mediated

degradation (Lee et al., 2010). This emerging function of Kelch repeat and SPRY

domains as substrate recognition domains for proteins forming part of E3 ubiquitin

ligase complexes is intriguing as little is known about the cellular functions of these

domains. The presence of the SPRY and Kelch repeat domains in RanBPM and

muskelin, respectively suggests that they may play roles as substrate recognition

components in the CTLH complex. Indeed, the yeast orthologue of RanBPM, Vid30

recognises and binds FBPase, a substrate of the Vid30 complex (Santt et al., 2008).

Armadillo repeats are found in a range of proteins involved in diverse cellular functions

including signalling, cytoskeleton formation/regulation and protein folding (Tewari et

al., 2010). The Armadillo repeat domain is also present in proteins involved in protein

degradation, such as the Arm-HECT E3 ubiquitin ligases, the F-box proteins Aardvark

and Arabidillo, the U-box ubiquitin ligase UFD2, and ARMC8α, which as will be

discussed shortly is involved in the degradation of α-catenin (Section 6.1.4.2) (Suzuki et

al., 2008; Tewari et al., 2010). Thus the presence of the LisH, SPRY, Kelch repeat and


227

Armadillo repeat domains within CTLH complex members provides some evidence that

the CTLH complex functions as an E3 ubiquitin ligase complex, similar to the yeast

Vid30 complex.

.

6.1.3 The Yeast Vid30 Complex

The yeast orthologue of the human CTLH complex is a well characterised E3 ubiquitin

ligase complex that ubiquitinates and targets for degradation a subset of gluconeogenic

enzymes including fructose 1,6 bisphosphatase (Section 4.1.1). The function of the

yeast Vid30 complex and the specific roles of each of the complex components may

indicate the function of the members of the human CTLH complex (Table 6.1). For

example, RMND5A, and potentially p44EMLP, yeast orthologues of which impart the

Vid30 complex with E3 ubiquitin ligase activity, may contribute this enzymatic activity

to the CTLH complex. Similarly, RanBPM, the human orthologue of Vid30 which

functions both as a core component and in substrate recognition of the complex, may

perform similar roles in the CLTH complex. Recently, Menssen et al. (2012) predicted

the architecture of the yeast Vid30 complex, and substitution of the Vid30 complex

members with the human CTLH complex components suggests the possible topology of

the CTLH complex as the human and yeast orthologues share the same protein domain

architecture (Figure 6.2) (Santt et al., 2008; Menssen et al., 2012).

The proposed CTLH complex architecture is consistent with the findings of Kobayashi

et al. (2007) in that each of the complex components are able to interact with each other

and RMND5A is able to interact with both ARMC8α and EMLP, with these interactions

potentially able to occur simultaneously as different protein domains are required

(Figure 6.2) (Kobayashi et al., 2007). Suzuki et al. (2008) determined that ARMC8α

and β are not essential for the formation of the CTLH complex, suggesting that they

function as adaptors (Suzuki et al., 2008). However, the yeast orthologue of ARMC8α,

Gid5 is a proposed core component of the Vid30 complex which binds the proposed E3

ubiquitin ligase components, Gid2/RMD5 (RMND5A) and Gid9 (p44EMLP) and links

the complex with Gid4, the activator of the Vid30 complex (Section 4.1.1) (Menssen et

al., 2012).


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Table 6.1 – The human CTLH complex components and their yeast orthologues

The human orthologue of Gid4, C17orf39 has not been identified as a member of the

CTLH complex, suggesting either that the CTLH complex has been investigated in an

inactive form and requires ARMC8α to bind C17orf39 for activation of the complex,

which may be the reason for the finding that ARMC8α was not necessary for CTLH

complex formation (Suzuki et al., 2008). Or, alternatively the CTLH complex does not

require activation in mammalian cells. However, as Gid4 is only transiently associated

with the Vid30 complex, C17orf39 may not be identified as a cofactor for the CTLH

complex unless investigated under the specific set of environmental conditions required

for activation of the CTLH complex, which have yet to be determined.

CTLH Complex Component

Protein Domains

Protein Function/Cell Type Identified or Investigated

Yeast Orthologue

Yeast Orthologue Function in Vid30 Complex

RMND5A LisH, CTLH, CRA, RING

Proposed E3 ubiquitin ligase/HEK cells

Gid2/RMD5 E3 ubiquitin ligase

RanBPM SPRY, LisH, CTLH, CRA

Scaffolding protein; immune and neural cell interactions/many cell types

Gid1/Vid30 Proposed core component, substrate recognition component

Muskelin LisH, CTLH, Kelch Repeat

Mediator of cell spreading and morphology; neural cell and cardiomyocyte interactions/many cell types

Gid7/Moh2 Unknown

EMP LisH, CTLH, CRA

Mediates maturation of erythroblasts/HEK cells

Gid9/Fyv10 E3 ubiquitin ligase

ARMC8α/β Armadillo repeats/β-catenin-like repeats

Associates with HRS and is involved in α-catenin degradation/HEK cells

Gid5/Vid28 Proposed core component

Twa1 LisH, CTLH, CRA

Unknown/HEK cells Gid8/Dcr1 Unknown

C17orf39 Protein kinase-like domain

Unknown; Not identified as a CTLH complex component

Gid4/Vid24 Activator of Vid30 complex


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The human orthologues of two proteins associated with the yeast Vid30 complex, the

E2 conjugating enzyme Ubc8 and the deubiquitinating enzyme Gid6/Ubp14, whose

human orthologues are UbcH2 and isopeptidase T, have not yet been determined to

interact with members of the human CTLH complex. However, in this thesis, RMND5A

was found to associate with UbcH2 in in vitro ubiquitination assays, and additional

studies may confirm UbcH2 interaction with the CTLH complex (Section 4.2.4.3).

These findings provide support further investigation of human CTLH complex function

as an E3 ubiquitin ligase complex that promotes protein degradation by the proteasome

and/or lysosome, as is the case for the yeast Vid30 complex.

6.1.4 CTLH Complex Components

The functions of most of the CTLH complex components have not been well

characterised, however reports in the literature have provided some indication of their

biological activities and thus their potential roles in the CTLH complex.

C17 orf39

ARMC8α

RMND5A EMLP

Twa1

RanBPM Muskelin

Yeast Vid30 Complex Human CTLH Complex

A B

Figure 6.2: Predicted Vid30 and CTLH complex topology. (A) Menssen et al. 2012 predicted the architecture of the yeast Vid30 complex by performing coimmunoprecipitation studies using full length and deletion constructs of the individual Vid30 complex members (B) Replacement of the Vid30 complex components with their proposed human orthologues in the predicted Vid30 complex structure indicates the possible structure of the CTLH complex. (Adapted from Menssen et al., 2012).


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6.1.4.1 Muskelin

Muskelin is a cytoplasmic and membrane associated protein that was originally

demonstrated to mediate the intracellular responses of cells growing on thrombospondin

by integrating cell spreading, adhesion and cytoskeletal organisation by as yet

uncharacterised mechanisms (Adams et al., 1998). Apart from its LisH and CTLH

domains, muskelin contains a carboxy-terminal Kelch repeat domain which forms a β-

propeller structure that functions as an actin binding motif in many proteins, and

although muskelin does not directly bind actin or tubulin, it is associated with the actin

cytoskeleton and modifies cytoskeletal organisation (Adams et al., 1998).

The muskelin amino-terminal contains a discoidin-like domain which, along with the

carboxy-terminal Kelch repeat domains, mediates self-association in a head to tail

orientation (Prag et al., 2004). PKC phosphorylates muskelin on two residues, Ser324

and Thr515, and mutation of these residues interferes with the ability of muskelin to

self-associate (Prag et al., 2007). The functional outcomes of this self-association are

incompletely characterised although other Kelch repeat containing proteins, including

Kelch, Actinfilin and Mayven undergo oligomerisation (Robinson and Cooley, 1997;

Soltysik-Espanola et al., 1999; Chen et al., 2002b). Multimerisation is required for the

mediation of actin filament crosslinking by Drosophila Kelch, the first identified Kelch

containing protein (Robinson and Cooley, 1997), whilst muskelin self-association has

recently been proposed to play a role in directing its cellular localisation (Valiyaveettil

et al., 2008). Although muskelin is predominantly localised in the cytoplasm, it is also

detectable in the nucleus, with studies in C2C12 skeletal myoblasts, COS-7 epithelial

cells and SW1222 carcinoma cells indicating that the LisH domain directs nuclear

localisation while the carboxy-terminal 35 amino acids localise muskelin to the

cytoplasm, even in the presence of the LisH domain (Valiyaveettil et al., 2008).

Muskelin self-association leads to autoinhibition, which is hypothesised to be relieved

by phosphorylation of Thr723, which releases the self-association and promotes nuclear

translocation (Valiyaveettil et al., 2008). The specific cellular environment required for

phosphorylation or dephosphorylation have not yet been determined.

Few muskelin binding partners have been reported, however those identified to date

support the involvement of muskelin in the endocytosis and intracellular transport of

cargo proteins as well as in cytoskeletal organisation. Muskelin interacts with the

Gamma-aminobutyric Acid A Receptor (GABAAR) α1 and aids in the transport of this


231

receptor to the lysosome for degradation. Muskelin and GABAAR α1 form a complex

with myosin VI and dynein, thereby facilitating the F-actin and microtubule based

internalisation and intracellular transport of GABAAR α1 to the lysosome for

degradation (Heisler et al., 2011). Delayed internalisation and intracellular transport of

GABAA R α1 are evident in muskelin knockout mice, which also exhibit diluted coat

colour, supporting a role for muskelin in intracellular transport in melanocytes (Heisler

et al., 2011). Muskelin associates with p39, a cyclin dependent kinase 5 (CDK5)

activator involved in the adhesion and migration of epithelial cells of the lens and

cornea (Ledee et al., 2005). p39 relocates muskelin to the cell periphery and is

hypothesised to link muskelin to the actin cytoskeleton by associating with α-actinin,

thereby facilitating its function in cytoskeletal organisation (Dhavan et al., 2002; Ledee

et al., 2005). Muskelin has also been identified as a prostaglandin EP3α receptor

binding partner that inhibits EP3α internalisation and promotes the association of this G

protein coupled receptor with Gi, enhancing Gi activity (Hasegawa et al., 2000).

6.1.4.2 ARMC8α

Although both ARMC8α and ARMC8β were identified as members of the CTLH

complex, ARMC8α is the longer isoform and is the isoform whose function has been

further investigated (Kobayashi et al., 2007; Suzuki et al., 2008; Tomaru et al., 2010).

ARMC8α is among several proteins including importin-α and β-catenin that contain

armadillo repeats, which are involved in intracellular transport and cell adhesion

(Franke et al., 1989; McCrea et al., 1991; Kobayashi et al., 2007). α-catenin colocalises

and interacts with ARMC8α and β-catenin at the cell membrane using its N-terminal

(amino acids 82-148). Overexpression of ARMC8α results in the rapid degradation of

α-catenin by the proteasome, whereas ARMC8α knockdown results in a prolonged α-

catenin half-life and reduced degradation (Suzuki et al., 2008). Together with β-catenin,

α-catenin forms part of the cadherin-catenin complex and is important in linking these

complexes to the actin cytoskeleton and directing actin assembly (Maiden and Hardin,

2011). ARMC8α also interacts with the endosomal protein, hepatocyte growth factor-

regulated tyrosine kinase substrate (HRS), which is involved in membrane protein

trafficking by recognising monoubiquitinated receptors such as the epidermal growth

factor receptor (EGFR), directing the ubiquitinated receptors for sorting through the

endosome and lysosomal degradation (Raiborg et al., 2003; Tomaru et al., 2010). The

interaction of HRS with ubiquitinated proteins through its ubiquitin interacting motif is

enhanced by ARMC8α and identification of the ubiquitinated proteins and the


232

mechanism by which ARMC8α is able to aid in their association with HRS will assist in

the understanding of ARMC8α function (Tomaru et al., 2010).

6.1.4.3 RanBPM

RanBPM has been reported to interact with at least 43 proteins involved in diverse

processes, identifying it to function as a protein stabiliser, transcriptional regulator, cell

cycle and neuronal function regulator and as a scaffolding/adaptor protein (Murrin and

Talbot, 2007; Suresh et al., 2012). RanBPM interacts with the cytoplasmic domain of

many membrane bound receptors, including MET, p75NTR, Axl/Sky, TrkA, TrkB and

with proteins associated with the intracellular domains of receptors including TRAF6

(Bai et al., 2003; Wang et al., 2004; Hafizi et al., 2005; Yuan et al., 2006; Yin et al.,

2010). Additionally, RanBPM interacts with other membrane bound proteins, including

the membrane transporter Dectin-1, the ecto-nuclease CD39 and the calcium CaV3.1

channel (Wu et al., 2006; Xie et al., 2006; Kim et al., 2009; Wang et al., 2012). The

interaction of RanBPM with specific receptors or associated proteins has been shown to

either enhance or repress signalling from that receptor. For example, RanBPM binding

to TRAF6 inhibits TRAF6 ubiquitination and its downstream signalling, leading to a

reduction in TRAF6 related NF-κB signalling (Wang et al., 2012). Alternatively,

RanBPM interaction with TrkB, a receptor tyrosine kinase for brain-derived

neurotrophic factor (BDNF), enhances BDNF-induced activation of the MAPK and Akt

signalling pathways and promotes neuronal survival (Yin et al., 2010).

RanBPM is also hypothesised to play a role in receptor endocytosis (Bai et al., 2003), it

reduces agonist-induced endocytosis of the mu opioid receptor without affecting

signalling from the receptor (Talbot et al., 2009), and by interacting with Plexin A,

RanBPM has been proposed to connect Plexin 3A receptors to retrograde transport

(Togashi et al., 2006). As well as binding to membrane bound receptors and proteins,

RanBPM interacts with steroid hormone receptors such as the AR, thyroid hormone

receptor and glucocorticoid receptor in which role it functions as a ligand dependent

coactivator. For example, RanBPM increases by 300% the transcriptional activity of

thyroid receptor alpha 1 on the thyrotropin releasing hormone and thyroid stimulating

hormone promoters (Rao et al., 2002; Poirier et al., 2006). RanBPM interacts with

numerous other transcription factors such as TAF4, which is involved in neural

development and the p53 associated nuclear transcription factor, p73α (Brunkhorst et

al., 2005; Kramer et al., 2005).


233

RanBPM may also play a role in the regulation of cytoskeletal organisation due to its

interaction with numerous proteins that mediate cell morphology, spreading and

migration, with RanBPM interaction with these proteins most frequently enhancing

these activities. For example, RanBPM has been characterised to interact with muskelin,

and knockdown of either of these proteins results in “protrusive cell morphologies with

enlarged perimeters” and altered F-actin distribution (Valiyaveettil et al., 2008). By

interacting with the lethal giant larvae (Mgl-1) tumour suppressor, RanBPM enhances

Mgl-1 associated cell migration and colony formation, and by interacting with MET, the

receptor tyrosine kinase for hepatocyte growth factor (HGF), RanBPM augments HGF-

MET signalling and enhances cell migration (Wang et al., 2004; Suresh et al., 2010). In

neural cells, RanBPM reduces cell proliferation, migration and neurite outgrowth by

interacting with and modulating the activity of CD39, the neural cell adhesion molecule

L1 and β1-integrin (Denti et al., 2004; Cheng et al., 2005; Wu et al., 2006). Together

with muskelin, RanBPM has been implicated in nucleocytoplasmic shuttling, as

supported by the original isolation of RanBPM as a Ran binding protein and by the roles

of RanBPM in escorting acetylcholinesterase and porphobilinogen deaminase (PBGD)

from the cytoplasm into the nucleus (Greenbaum et al., 2003; Gong et al., 2009).

The diverse activities of RanBPM have led to the hypothesis that it functions as an

adaptor or scaffolding protein, which is consistent with its proposed role in the CTLH

complex. The nucleocytoplasmic and cytoskeletal functions of RanBPM following its

interaction with muskelin suggest complementary or coordinated functions of these two

proteins, potentially including their roles as members of the CTLH complex.

6.1.4.4 Erythroblast Macrophage Protein (EMP)

Erythroblast macrophage protein (EMP/MAEA) was originally identified as a ~30kDa

protein mediating the attachment of erythroblasts to macrophages in erythroblast islands

(Hanspal and Hanspal, 1994). A number of lines of evidence have demonstrated that

EMP is required for the direct association of macrophages and erythroblasts in these

islands. Firstly, EMP possesses a short amino-terminal extracellular domain, and HeLa

cells expressing an amino-terminal deletion mutant are unable to associate with

erythroblasts (Hanspal et al., 1998). Additionally, Emp null mice display

haematopoietic defects, including the inability of macrophages to form erythroblastic

islands (Soni et al., 2006; Soni et al., 2008). In immature erythroblasts, EMP is


234

associated with the nuclear matrix whilst in mature erythroblasts, EMP is localised at

the cell membrane. Immature erythroblasts are unable to associate with macrophages,

suggesting that EMP must be attached to the cell membrane to directly mediate cell

attachment and that the protein may have additional functions in the nucleus (Soni et

al., 2007). The cell membrane association of EMP in HeLa cells indicates that EMP can

exhibit an extracellular amino-terminus in cell types other than macrophages, with this

domain potentially functioning in cell-cell or cell-matrix adhesion. Since the carboxy-

terminal intracellular domain of EMP contains multiple tyrosine residues, it has been

hypothesised that these residues may become phosphorylated, allowing association of

phospho-EMP with signalling kinases and therefore with intracellular signalling

pathways (Hanspal et al., 1998). Although the nuclear role of EMP has yet to be

investigated, in order to assess the cellular roles of EMP in non-haematopoietic cells,

Bala et al. (2006) determined the localisation of EMP in HEK cells. In these cells,

recombinant EMP was localised to either the nuclear matrix or cell membrane and

depending upon the stage of the cell cycle, EMP was localised to distinct nuclear

regions. During interphase, EMP colocalised with nuclear actin and SC35, a

spliceosome assembly factor whilst during mitosis, EMP localised to mitotic spindles

(Bala et al., 2006).

Another mechanism by which EMP is involved in erythroblast-macrophage maturation

is via its association with and regulation of the distribution of F-actin in erythroblasts

and macrophages, contributing to the enucleation of erythroblasts and the formation of

macrophage filopodia (Soni et al., 2006). These results are supported by the finding that

EMP null embyros exhibit little cytoplasmic F-actin, defects in erythropoiesis including

incomplete terminal differentiation of erythroblasts which display nuclei, and

macrophages that do not display cytoplasmic protrusions (Soni et al., 2006; Soni et al.,

2008). EMP additionally possesses anti-apoptotic functions and EMP deletion in

erythroblasts results in apoptosis (Hanspal et al., 1998). This activity is consistent with

that of its yeast orthologue, Fyv10/Gid9 which also plays an anti-apoptotic role

(Hanspal et al., 1998; Khoury et al., 2008). However, the mechanism by which EMP

inhibits apoptosis is yet to be established.

Members of the CTLH complex and their associated protein domains are therefore

implicated in broadly similar cellular pathways including cytoskeletal organisation,

protein trafficking and protein degradation. The widespread expression of many of the


235

CTLH complex members suggests that the complex may form in diverse cell types,

although the existence of the CTLH complex in cells other than HEK293 cells has not

been investigated. Additionally, the inclusion of orthologues of the complex members

RanBPM (RanBP10) and RMND5A (RMND5B) in the CTLH complex has not yet

been described. In addition to their proposed activities as part of the CTLH complex,

the multidomain CTLH complex members and their orthologues may also function in

additional roles. In this thesis, further characterisation of RMND5 protein function was

initiated by investigation of the expression of CTLH complex members in prostate

cancer cells and identification of RMND5A and RMND5B binding proteins.


236

6.2 Results

6.2.1 Transcripts encoding the CTLH Complex Components are Expressed

in Prostate Cancer Cells

In order to determine whether it was possible for the human CLTH complex to form in

prostate cancer cells, RT-PCR was used to identify whether the complex components

were expressed in LNCaP cells. Between 300bp-600bp of the coding region of each of

the CTLH complex components was PCR amplified using LNCaP cDNA template and

specific primer pairs for each CTLH complex member (Sections, 3.2, 3.3, Appendix II).

For RMND5A, RMND5B, RanBPM and muskelin, PCRs were carried out using 1.5mM

MgCl2 and an annealing temperature of 55°C for 35 cycles and the products

electrophoresed in a 2% agarose gel, identifying bands at the expected size for each

cDNA, RMND5A (~574bp), RMND5B (~393bp), RanBPM (~641bp) and muskelin

(~446bp) (Sections 3.4, 3.6, Figure 6.3A). Twa1 and EMP PCR amplification was

optimised using a gradient of annealing temperatures between 53°C and 60°C with

1.5mM MgCl2, and the PCR products were electrophoresed in a 2% agarose gel from

which single bands corresponding in size to Twa1 (~347bp) and EMP (~504bp) were

identified when higher annealing temperatures were used (Sections 3.4, 3.6, Figure

6.3B). ARMC8α was PCR amplified using a range of 1-2mM MgCl2 and an annealing

temperature of 55°C for 35 cycles, with analysis of the PCR products by electrophoresis

in a 2% agarose gel verifying the presence of the ~566bp ARMC8α fragment at all

MgCl2 concentrations tested (Sections 3.4, 3.6, Figure 6.3C). PCRs optimising the

amplification of C17orf39 were carried out using a range of annealing temperatures

from 53°C-60°C and 1-2mM MgCl2 concentrations for 35 cycles (Sections 3.4, 3.6).

Electrophoresis of the products identified a band corresponding in size to C17orf39

(~517bp) when an annealing temperature of 55°C and 2mM MgCl2 were used (Sections

3.4, 3.6, Figure 6.3D, E).

To confirm expression of each CTLH complex member in LNCaP cells and to ascertain

their expression in the DU145 prostate cancer and MCF-7 breast cancer cell lines, the

optimised PCR conditions for each complex component were used to amplify each

member using LNCaP, DU145 and MCF-7 cDNA, which was obtained by extraction of

the RNA from each cell line and reverse transcription of the RNA using oligo-dT

primers (Sections 3.2, 3.3). The PCR products were electrophoresed in 2% agarose gels,

verifying the presence of each complex component at the expected molecular sizes in


237

*

*

C Lane 1: MW Marker Lane 2: 1mM MgCl2 Lane 3: 1.5 mM MgCl2 Lane 4: 2mM MgCl2 Lane 5: Negative Control

Lane 1: MW Marker Lane 2: 53˚C Lane 3: 54˚C Lane 4: 55˚C Lane 5: 56˚C Lane 6: 57˚C Lane 7: Negative control

B

Lane 8: MW Marker Lane 9: 56˚C Lane 10: 57˚C Lane 11: 58.2˚C Lane 12: 59.2˚C Lane 13: 60˚C Lane 14: Negative Control

(i)

(ii)

Lane 1: MW Marker Lane 2: RMND5A (574bp) Lane 3: RMND5B (393bp) Lane 4: RanBPM (641bp) Lane 5: Muskelin (446bp) Lane 6: Negative Control

A


238

Figure 6.3: Optimisation of RMND5A, RMND5B, RanBPM, muskelin, Twa1, EMP, ARMC8α and C17orf39 PCR conditions. (A) PCRs for RMND5A, RMND5B, RanBPM and muskelin using LNCaP cDNA, 1.5mM MgCl2 and an annealing temperature of 55°C resulted in amplification of products of the expected size. (B) Gradient PCRs using 1.5mM MgCl2 and LNCaP cDNA were performed to optimise the amplification of (i) Twa1 (347bp) and (ii) EMP (504bp). Lower annealing temperatures resulted in the amplification of multiple bands, while higher temperatures yielded a single band of the expected molecular size (*) for both Twa1 and EMP. (C) PCRs to amplify ARCM8α (566bp) were carried out using an annealing temperature of 55°C, LNCaP cDNA and 1mM, 1.5mM or 2mM MgCl2, with all three conditions yielding products of the expected molecular size. (D) Gradient PCRs with annealing temperatures ranging from 53°C to 60°C were performed to amplify C17orf39 (517bp). (E) A prominent band corresponding to C17orf39 at ~500bp was amplified at an annealing temperature of 56°C, which was utilised in further PCRs using 1mM, 1.5mM and 2mM MgCl2. Ten µL of each reaction was electrophoresed in 2% agarose gels to visualise the products.

Lane 1: MW Marker Lane 2: 53˚C Lane 3: 54.2˚C Lane 4: 55˚C Lane 5: 56˚C Lane 6: 57.3˚C Lane 7: 58˚C Lane 8: 59.5˚C Lane 9: 60˚C Lane 10: Negative Control

D

E Lane 1: MW Marker Lane 2: 56˚C (1mM MgCl2) Lane 3: 56˚C (1.5 mM MgCl2) Lane 4: 56˚C (2mM MgCl2) Lane 5: Negative Control


239

each cell line (Section 3.4, 3.6, Figure 6.4). The amplified PCR products were not

sequenced at this stage to confirm that the appropriate cDNAs had been amplified,

however this could be performed in future studies to continue investigation of CTLH

complex members.

6.2.2 Cloning of RanBPM

The yeast orthologue of RanBPM, Gid1/Vid30, is a proposed core component of the

yeast Vid30 complex, suggesting that human RanBPM may fulfil a similar role in the

CTLH complex (Pitre et al., 2006). This is supported by the finding that RanBPM

interacts with each complex member in coimmunoprecipitation assays in Cos-7 cells

(Kobayashi et al., 2007). To investigate its similar function in prostate cancer cells, the

interaction between RMND5A and RanBPM was verified in the LNCaP cell line, and to

examine the potential for RMND5B to join or replace RMND5A in the CTLH complex,

interaction between RanBPM and RMND5B was also investigated. To perform these

assays, the RanBPM coding region was cloned into the pmCherry-C1 expression vector

to allow the expression of Cherry-RanBPM. Two isoforms of RanBPM have been

identified, a full length 90kDa isoform, and a smaller 55kDa isoform which has a

truncated amino-terminus but does not lack the currently identified protein domains

(Nakamura et al., 1998; Nishitani et al., 2001; Kobayashi et al., 2007) (Section

6.1.4.1). The PCR primers used to amplify RanBPM (Section 6.2.1) would not

distinguish between the 90kDa and 55kDa isoforms.

6.2.2.1 Cloning of Full Length RanBPM (90kDa)

To clone the 90kDa RanBPM isoform, PCR primers were designed to amplify the entire

RanBPM coding region using the RanBPM1-S and RanBPM2190-AS primers from

LNCaP, DU145 and MCF-7 cDNA, however, no products were amplified even after

extensive optimisation with a range of annealing temperatures, MgCl2 concentrations,

high fidelity and high GC content buffers, DMSO, the use of different RNA extraction

methods, reverse transcriptase enzymes and Taq polymerases (Section 3.4, 3.6, not

shown, and Figure 6.5). The second approach was to amplify the RanBPM coding

region in two fragments, and using an endogenous XbaI site, the two fragments were to

be ligated into the pGEM®T-Easy cloning vector (Figure 6.5). However, again, no

bands were present upon agarose gel electrophoresis of the PCR products (Section 3.4,

3.6, not shown). Therefore, amplification of the smaller 55kDa isoform of RanBPM


240

Figure 6.4: The CTLH complex components are expressed in prostate and breast cancer cells. RNA was extracted from the (A) LNCaP, (B) DU145 (prostate cancer) and (C) MCF-7 (breast cancer) cell lines, reverse transcribed and the resulting cDNA utilised in PCRs for each of the CTLH complex members. Ten µL of each reaction was electrophoresed in 2% agarose gels, identifying that all CTLH complex members were expressed in each cell line.

LNCaP

DU145

MCF-7

A

B

C

Lane 1: MW marker Lane 2: RMND5A (573bp) Lane 3: RMND5B (393bp) Lane 4: RanBPM (618bp) Lane 5: Twa1 (347bp) Lane 6: EMP (504bp) Lane 7: ARMC8α (566bp) Lane 8: Muskelin (446bp) Lane 9: C17orf39 (517bp) Lane 10: Negative Control


241

XbaI 1 2190 1190

RanBPM1-S

RanBPM 2190-AS

1 2190

1

2190

1259

1029

RanBPM1-S

RanBPM2190-AS

RanBPM 1029-S primer

RanBPM1259-AS

A

B

C

1 2190

SalI

SalI

SalI

HindIII

HindIII

HindIII

687 2190 HindIII SalI

RanBPM2190-AS

RanBPM687-S

D


242

Figure 6.5: Cloning of the RanBPM coding region. (A) Full length RanBPM is encoded by a 2190bp coding region that (B) could be amplified as a single fragment using the RanBPM1-S and RanBPM2190-AS primers (Appendix II). As the full length isoform could not be amplified in this manner, (C) the second method utilised the presence of an endogenous XbaI restriction site at position 1190bp to aid in the cloning of RanBPM as two fragments using the RanBPM1-S and RanBPM1259-AS primers to amplify the amino-terminal 1259bp and the RanBPM1029-S and RanBPM2190-AS primers to amplify the carboxy-terminal 1162bp of the RanBPM coding region which could both be ligated into the pGEM®T Easy cloning vector using TA cloning. The carboxy-terminal fragment could then be excised from the pGEM®T Easy cloning vector by restriction enzyme digestion with XbaI and SalI and the pGEM®T Easy cloning vector containing the amino-terminal fragment digested with XbaI to and SalI (at the 3’ end of the multiple cloning site of pGEM®T Easy) to allow the ligation of the second fragment into the cloning vector containing the first 1259bp fragment and thereby the production of full length RanBPM (2190bp). (D) If this could not be achieved, the smaller 55kDa isofom of RanBPM (55kDa) which arises from an alternative start site in the RanBPM coding region (687bp) could be PCR amplified using the RanBPM687-S and RanBPM2190-AS primers. The protein sequence encoded by the first 687bp of RanBPM does not contain identifiable protein domains or localisation signals.


243

was pursued as it was not clear whether the failure to amplify full-length RanBPM was

due to technical difficulties, whether the full-length isoform was not expressed or was

expressed at low levels in LNCaP cells, or whether the full-length RanBPM mRNA was

not reverse transcribed efficiently under the conditions used. Thus, as the 55kDa

isoform of RanBPM has been used in previous studies characterising RanBPM function,

and the isoform retains all identified RanBPM protein domains, this approach was

considered acceptable for these studies (Figure 6.5) (Nakamura et al., 1998).

6.2.2.2 Cloning of the RanBPM 55kDa Isoform into pmCherry-C1

The RanBPM (55kDa) coding region was PCR amplified using the RanBPM687-S and

RanBPM2190-AS primers, a range of annealing temperatures from 52°C-60°C and

LNCaP cDNA (Section 3.2, 3.3, 3.4, Appendix II). The PCR products were

electrophoresed in a 1% agarose gel, identifying a product at the expected size of

~1.5kb at the annealing temperatures of 60°C, 58.6°C and 54.8°C (Figure 6.6A). The

PCR was repeated in quadruplicate at the optimum annealing temperature of 60°C, the

products combined, “A” tails added, the product purified and 5µL electrophoresed in a

1% agarose gel to confirm amplification of the ~1500bp product (Section 3.4, 3.6, 3.7.1,

Figure 6.6B).

To obtain pGEMT-RanBPM (55kDa), 40ng (4µL) purified RanBPM (55kDa) was

ligated into 50ng (1µL) pGEM®T-Easy and the ligation products were transformed into

competent E. coli DH5α cells (Sections 3.8.4, 3.8.5.1, 3.8.6). Colonies were selected by

growth on LB Agar/Ampicillin plates with blue/white colony selection and 6 white

colonies were picked and cultured in LB Broth/Ampicillin overnight (Section 3.8.6,

3.9). Plasmids were isolated by small scale plasmid purification, RNase treated and

EcoRI digested to release the RanBPM (55kDa) insert, and the products electrophoresed

in a 1% agarose gel (Section 3.6, 3.8.2, 3.9, Figure 6.6C). Inserts of the expected size

were identified in all clones, pGEMT-RanBPM (55kDa) clones 1 to 6 were purified and

2µL of each was electrophoresed in a 1% agarose gel (not shown). Based on the gel,

2µL of each of clones 1 to 3 was used in sequencing reactions using the M13-S, M13-

AS, RanBPM1029-S and RanBPM1259-AS primers (Appendix II, Section 3.12).

BLASTTM analysis of the sequencing chromatograms identified that pGEMT-RanBPM

(55kDa) clone 2 was mutation free (not shown).


244

Figure 6.6: Cloning of RanBPM (55kDa) into the pGEM®-T Easy cloning vector. (A) To optimise amplification of the RanBPM (55kDa) isoform (~1500bp), PCRs containing 1.5mM MgCl2 and LNCaP cDNA were performed using annealing temperatures from 52°C to 60°C. (B) Using the optimum annealing temperature (60°C), four 25µL PCRs were performed, the amplified products pooled and purified. A 5µL aliquot of the purified product was electrophoresed in a 1% agarose gel from which the DNA concentration was estimated to be ~10ng/µL. (C) Purified RanBPM (55kDa) was ligated into the pGEM®-T Easy cloning vector, the ligation reaction transformed into DH5α cells and plasmid DNA extracted from 6 colonies. Plasmids isolated from clones 1 to 6 were digested with EcoRI and electrophoresed in a 1% agarose gel. Inserts were identified in all clones.

~3000bp

C

Lane 1: MW marker Lane 2, 4, 6, 8, 10, 12: Undigested cut pGEMT-RanBPM (55kDa) clones 1-6 Lane 3, 5, 7, 9, 11, 13: EcoRI digested pGEMT-RanBPM (55kDa) clones 1-6 Lane 14: MW marker

~540bp ~970bp

Lane 1: MW marker Lane 2: 60˚C Lane 3: 58.6˚C Lane 4: 57.8˚C Lane 5: 56.5˚C Lane 6: 54.8˚C Lane 7: 53.4˚C Lane 8: 52.5˚C Lane 9: 52˚C Lane 10: Negative Control

Lane 1: MW marker Lane 2: Purified RanBPM

(55kDa) Lane 3: Negative Control

A B ~1500bp ~1500bp

1 2 3 4 5 6 7 8 9 10


245

To subclone RanBPM (55kDa) into pmCherry-C1, pGEMT-RanBPM (55kDa) clone 2

was digested with HindIII and SalI to release the insert, the insert was gel purified and

5µL of the purified product was electrophoresed in a 1% agarose gel (Section 3.6, 3.7.2,

3.8.2, Figure 6.7A). pmCherry-C1 was prepared by digestion with HindIII and SalI,

SAP treatment and 2µL product was electrophoresed in a 1% agarose gel (Section 3.6,

3.7.2, 3.8.2, Figure 6.7B). The ligation reaction containing 30ng (6µL) RanBPM

(55kDa) insert and 30ng (2µL) pmCherry-C1, was transformed into competent E. coli

DH5α cells and the transformed bacteria were selected by growth on LB

Agar/Kanamycin (Sections 3.8.4, 3.8.6). Plasmids extracted from 7 colonies were NdeI

digested and the purified products were electrophoresed in a 1% agarose gel (Section

3.6, 3.8.2, 3.9, Figure 6.7C), with products of the expected size identified in pmCherry-

RanBPM (55kDa) clones 1, 2, 3, 4 and 6 (Figure 6.7C). Clones 1-3 were purified and

sequenced using the RanBPM1-S, RanBPM1029-S, RanBPM2190-AS and

RanBPM1259-AS primers from which pmCherry-RanBPM (55kDa) clone 2 was

verified as mutation free using BLASTTM analysis (Section 3.12, Appendix III).

Large scale preparation of pmCherry-RanBPM (55kDa) clone 2 was performed to

obtain purified plasmid for transfection, the concentration of which was determined

using spectrophotometry to be 1.5µg/µL (Section 3.6, 3.10, Figure 6.7D). To confirm

expression from Cherry-RanBPM (55kDa) clone 2, LNCaP cells were transfected with

4µg pmCherry-RanBPM (55kDa), the cells were harvested at 48 hours post-transfection

and 10µL lysate was electrophoresed in a 12% polyacrylamide gel for western blotting

(Section 3.1.4, 3.15). Cherry western blotting identified a band at the expected

molecular size of ~85kDa in lysates from transfected cells, indicating successful

expression of the pmCherry-RanBPM (55kDa) construct (Figure 6.7E).

6.2.3 Interaction between RanBPM and RMND5A/RMND5B

6.2.3.1 RMND5A Interaction with RanBPM (55kDa)

To investigate the interaction of RMND5A with RanBPM, LNCaP cells growing in

10cm petri dishes were cotransfected with plasmids encoding GFP-RMND5A and

Cherry-RanBPM (55kDa), at 48 hours post-transfection the cells were lysed, an aliquot

was taken (total cellular input) and the lysate was immunoprecipitated with GFP

antibodies (Section 3.1.4, 3.13). The GFP immunocomplexes were electrophoresed in

12% polyacrylamide gels and GFP and Cherry western blotting was performed


246

Cherryβ-actin

~85kDa

~45kDa

Figure 6.7: Preparation of the pmCherry-RanBPM (55kDa) expression plasmid. (A) To prepare pmCherry-RanBPM (55kDa), pGEMT-RanBPM (55kDa) clone 2 was digested with HindIII and SalI to release the ~1500bp insert, which was gel purified. 5µL of the purified insert was electrophoresed in a 1% agarose gel from which the insert concentration was estimated to be ~5ng/µL. (B) To prepare pmCherry-C1, the plasmid was digested with HindIII and SalI, purified, SAP treated, re-purified and 2µL of the purified product was electrophoresed in a 1% agarose gel from which the concentration was estimated to be ~15ng/µL. (C) pmCherry-RanBPM (55kDa) was digested with NdeI, and the digested products electrophoresed in a 1% agarose gel. Clones 1, 2, 3, 4 and 6 contained an insert of the expected size (~1.16kb). (D) Large scale purification of the pmCherry-RanBPM (55kDa) plasmid was performed, and the purified plasmid was digested with NdeI and electrophoresed in a 1% agarose gel. (E) Expression of RanBPM (55kDa) from the pmCherry-RanBPM (55kDa) plasmid was determined by transfection of LNCaP cells with 4µg pmCherry-RanBPM (55kDa) and Cherry western blotting of lysates from cells harvested 48 hours post-transfection. A Cherry fusion protein of the expected size of ~85kDa was identified in transfected cultures.

E

Lane 1: MW Marker Lane 2: Undigested pmCherry-RanBPM (55kDa) Lane 3: NdeI digested pmCherry-RanBPM

(55kDa)

D

~1.16kb

~5kb

Lane 1: MW Marker Lane 2: 5µL gel purified

RanBPM (55kDa)

~1.5kb

A B

Lane 1: MW Marker Lane 2: 2µL Undigested pmCherry-C1 Lane 3: 2µL HindIII/SalI, SAP digested

pmCherry C1

~4.7kb

C

Lane 1: MW marker Lane 2, 4, 6, 8, 10, 12, 14: Undigested pmCherry-RanBPM55 clones 1-7 Lane 3, 5, 7, 9, 11, 13, 15: NdeI digested pmCherry-RanBPM55 clones 1-7 Lane 16: MW marker

~1.16kb

~5kb


247

(Section 3.15). A band corresponding in size to GFP-RMND5A at ~70kDa was present

in both the total cellular input and immunoprecipitated sample indicating successful

immunoprecipitation of GFP-RMND5A (Figure 6.8A). No bands were present in the

untransfected and mock immunoprecipitation samples. Western blotting for Cherry-

RanBPM (55kDa) identified a band at ~85kDa corresponding in size to Cherry-

RanBPM (55kDa) in the total cellular input and immunoprecipitated samples, indicating

successful co-immunoprecipitation of Cherry-RanBPM (55kDa) with GFP-RMND5A.

No bands were present in the untransfected and mock immunoprecipitation samples

(Figure 6.8A). These results confirmed an interaction between RMND5A and RanBPM

(55kDa).

6.2.3.2 RMND5B Interaction with RanBPM (55kDa)

To determine whether RMND5B and RanBPM interact in LNCaP cells, cells growing

in 10cm petri dishes were cotransfected with pEGFP-RMND5B and pmCherry-

RanBPM (55kDa) and at 48 hours post-transfection the cells were lysed, and the lysates

immunoprecipitated with GFP antibodies (Section 3.1.4, 3.13). Samples were

electrophoresed in 12% polyacrylamide gels and GFP western blotting verified the

presence of a ~70kDa band corresponding in size to GFP-RMND5B in the

immunoprecipitate, indicating successful GFP immunoprecipitation (Section 6.8B). No

GFP immunoreactive bands were present in the total cellular input indicating that GFP-

RMND5B was present at low levels in this (dilute) fraction (Figure 6.8B). Similarly,

GFP-RMND5B was not detected in the untransfected and mock immunoprecipitated

samples, while Cherry western blotting identified an ~85kDa band corresponding in size

to Cherry-RanBPM (55kDa) in both the total cellular input and immunoprecipitated

samples, with no bands visible in the untransfected and mock immunoprecipitated

controls (Figure 6.8B). These results indicated that RMND5B was also able to associate

with RanBPM (55kDa).

6.2.3.3 RanBPM (55kDa) Interaction with RMND5 proteins

To perform reciprocal coimmunoprecipitation studies to identify coimmunoprecipitation

of RMND5 proteins with Cherry-RanBPM, Cherry-RanBPM immunoprecipitation

using Protein G beads was optimised as the antibody subtype (rat IgG2a) is not

efficiently bound by Protein A (Section 3.15, 3.16, not shown). LNCaP cells growing in

10cm dishes were cotransfected with plasmids encoding GFP-RMND5A and Cherry-


248

WB: Cherry (RanBPM)

Untransfected

WB: GFP (RMND5B)

IP: GFP (RMND5B)Cotransfected

WB: GFP (RMND5A)

WB: Cherry (RanBPM)

IP: GFP (RMND5A)CotransfectedUntransfectedA

B

~70kDa

~85kDa

~70kDa

~85kDa

Figure 6.8: RMND5A and RMND5B interact with RanBPM in LNCaP cells. LNCaP cells were cotransfected with plasmids encoding Cherry-RanBPM (55kDa) and either GFP-RMND5A or GFP-RMND5B, the cells were harvested 48 hours post-transfection, proteins were immunoprecipitated from cell lysates using anti-GFP antibodies and Protein A beads and immunoprecipitated/coimmunoprecipitated proteins were detected by GFP and Cherry western blotting. (A) In cells cotransfected with GFP-RMND5A and Cherry-RanBPM (55kDa), western blotting detected both proteins in the total cell lysate and in immunoprecipitated samples. (B) In cells cotransfected with GFP-RMND5B and Cherry-RanBPM (55kDa), both proteins were detected in the immunoprecipitated sample and Cherry-RanBPM (55kDa) was detected in the total cell lysate. Levels of GFP-RMND5B were usually too low to be detected by GFP western blotting in total cell lysates (total input). Immunoprecipitated proteins were not detected in lysates of untransfected cells or in mock immunoprecipitation reactions (Mock). Experiments were performed twice and representative blots are shown.


249

RanBPM (55kDa) and 48 hours post-transfection, the cells were lysed and an aliquot

(total cellular input) taken (Section 3.1.4, 3.13). The remaining lysate was subjected to

immunoprecipitation using either GFP antibodies (mouse IgG1) or Cherry antibodies

(rat IgG2a), the immunoprecipitates were electrophoresed in 12% polyacrylamide gels

prior to GFP and Cherry western blotting (Section 3.13, Figure 6.9A, B). Although

both antibody subtypes, mouse IgG1 and rat IgG2a, should be recognised and bound by

Protein G, western blotting for both GFP-RMND5A or Cherry-RanBPM (55kDa)

identified only a small amount of either GFP-RMND5A or Cherry-RanBPM (55kDa) in

the immunoprecipitated samples, even though both proteins were readily detected in the

total cellular input samples (Figure 6.9A, B). These findings indicated that

immunoprecipitation using Protein G beads was inadequate. To further optimise the

methods, the experiment was repeated using Protein G beads from Miltenyi Biotec,

cells were lysed at 48 hours post-transfection and after an aliquot was taken (total

cellular input) the remaining lysate was immunoprecipitated with a Cherry antibody (rat

IgG2a) and Protein G beads (Section 3.1.4, 3.13). The Cherry immunocomplexes were

electrophoresed in 12% polyacrylamide gels and then analysed by Cherry and GFP

western blotting (Section 3.15, Figure 6.9C). Cherry western blotting identified a band

corresponding to Cherry-RanBPM (55kDa) at ~85kDa in the immunoprecipitated

samples, indicating successful immunoprecipitation. No bands were present in the total

cellular input samples, indicating that Cherry-RanBPM (55kDa) levels were below the

level of detection of the antibody. Similarly no bands were present in the untransfected

and unbound control fractions. GFP western blotting identified bands corresponding to

GFP-RMND5A and GFP-RMND5B in the immunoprecipitated samples at ~70kDa

(Figure 6.9C). Again, no bands corresponding to either GFP-RMND5A or GFP-

RMND5B were present in the total cellular input samples, indicating that these proteins

were present at low levels in this (dilute) fraction. Similarly, no bands were detected in

the untransfected and unbound control samples. These results confirmed the interaction

between RanBPM and RMND5A/RMND5B in LNCaP cells. Therefore, the studies

identified and confirmed RanBPM interaction with both RMND5A and RMND5B in

LNCaP cells.

6.2.4 Colocalisation of RanBPM with RMND5A and RMND5B

To investigate the localisation and colocalisation of RanBPM with RMND5A and

RMND5B using fluorescence microscopy, LNCaP cells growing on coverslips were


250

IP: Cherry (RanBPM)WB: Cherry (RanBPM)

Immunoprecipitating antibody: Cherry (rat IgG2a)

IP: GFP (RMND5A)WB: GFP (RMND5A)

Immunoprecipitating antibody: GFP (mouse IgG1) A

B

~70kDa

~85kDa

GFP-RMND5A GFP-RMND5B

WB: GFP

WB: Cherry (RanBPM)

IP: Cherry (RanBPM)

~70kDa

~85kDa

C

Figure 6.9: Optimisation of immunoprecipitation reactions using Protein G Sepharose. To optimise immunoprecipitation reactions using Protein G sepharose, LNCaP cells were cotransfected with GFP-RMND5A and Cherry-RanBPM (55kDa), harvested at 48 hours following transfection and immunoprecipitation reactions performed on the cell lysates using either (A) anti-GFP (mouse IgG1) antibody or (B) anti-Cherry (rat IgG2a), with western blotting using the same antibody performed to evaluate the immunoprecipitation reaction. Both of the immunoprecipitating antibodies are reported to be recognised by Protein G. The immunoprecipitated protein of interest was detected in both the total cell lysate and to a lesser extent in the immunoprecipitated samples. (C) Miltenyi Biotec Protein G sepharose was used to immunoprecipitate Cherry-RanBPM with an anti-Cherry antibody (rat IgG2a) and Cherry and GFP western blotting identified Cherry-RanBPM (55kDa) and either GFP-RMND5A or GFP-RMND5B in the immunoprecipitated samples (IP). Experiments were performed twice and representative blots are shown.


251

cotransfected with plasmids encoding Cherry-RanBPM (55kDa) and either GFP-

RMND5A or GFP-RMND5B and at 48 hours post-transfection the cells were prepared

for microscopy (Section 3.1.3, 3.1.4, 3.16). Under these culture conditions, Cherry-

RanBPM (55kDa) displayed a diffuse nuclear and cytoplasmic cellular localisation

when coexpressed with GFP-RMND5A, which also exhibited a diffuse localisation in

the cytoplasm and nucleus (Figure 6.10). These findings are consistent with a previous

report investigating the intracellular localisation of RanBPM and RMND5A in HEK293

cells (Kobayashi et al., 2007). When coexpressed with GFP-RMND5B, both Cherry-

RanBPM (55kDa) and GFP-RMND5B exhibited a diffuse cellular localisation in the

nucleus and cytoplasm, however consistent with a previous study, GFP-RMND5B also

displayed punctate cytoplasmic staining in a proportion of cells (Dawson, 2006) (Figure

6.10). In these cells, Cherry-RanBPM (55kDa) colocalised with GFP-RMND5B in a

proportion of these speckles (Figure 6.10). The results identified that RMND5A and

RMND5B colocalise with RanBPM (55kDa) in LNCaP cells.

6.2.5 Interaction Between RMND5A and RMND5B

To determine whether RMND5 proteins interacted with each other, the

coimmunoprecipitation and colocalisation of RMND5A and RMND5B was investigated

in LNCaP cells. For these studies, RMND5B was initially cloned into the pmCherry

expression vector to allow the expression of a Cherry-RMND5B fusion protein.

6.2.5.1 Cloning of RMND5B into pmCherry-C1

To generate pmCherry-RMND5B, the RMND5B coding region was PCR amplified

using pEGFP-RMND5B as a template, the RMND5BBamHI1-S and RMND5BBamHI-

AS primers, a 55°C annealing temperature and 1-2mM MgCl2 for 35 cycles (Section

3.4). The PCR products were electrophoresed in a 1% agarose gel, identifying a ~1.2kb

product when 1.5mM MgCl2 and 2mM MgCl2 were used (Section 3.6, Figure 6.11A).

The optimised PCR conditions with 2mM MgCl2 were used to amplify RMND5B in

quadruplicate, the products were combined, “A” tails added, and 5µL of the purified

products electrophoresed in a 1% agarose gel (Section 3.4, 3.7.1, Figure 6.11B). To

obtain pGEMT-RMND5B, 80ng (2µL) RMND5B was ligated with 50ng (1µL)

pGEM®T-Easy and the ligation products were transformed into competent E. coli DH5α

(Section 3.8.4, 3.8.6). Transformants were selected by growth on LB Agar/Ampicillin

using blue/white colony selection, plasmids were purified from six white colonies,


252

Figure 6.10: Cherry-RanBPM (55kDa) colocalises with GFP-RMND5A and GFP-RMND5B. LNCaP cells were cotransfected with plasmids encoding Cherry-RanBPM (55kDa) and either GFP-RMND5A or GFP-RMND5B and the cells fixed and prepared for microscopy 48 hours post-transfection. The three fusion proteins exhibited a diffuse nuclear and cytoplasmic localisation in LNCaP cells with accumulations of GFP-RMND5B also having a punctate appearance. Cherry-RanBPM (55kDa) colocalised with (A) GFP-RMND5A and (B) GFP-RMND5B in both the nucleus and cytoplasm of LNCaP cells (C) Untransfected cells were imaged under the same conditions as the transfected cells (Magnification x1000). The experiment was performed twice and representative results are shown.

Cherry-RanBPM GFP-RMND5B Hoechst33258 Overlay

Cherry-RanBPM GFP-RMND5A Hoechst 33258 Overlay A

B

-ve control (546nm) Hœchst 33258 Overlay C -ve control (488nm)

Chapter 6 The CTLH Complex

253

Figure 6.11: Cloning of RMND5B into pGEM®-T Easy. (A) RMND5B was amplified from the pEGFP-RMND5B expression vector using 1-2mM MgCl2 and 55°C annealing temperature for 35 cycles. PCR products (10µL) were electrophoresed in a 1% agarose gel. (B) RMND5B was amplified using the optimised PCR conditions, the PCR products purified and 5µL electrophoresed in a 1% agarose gel from which the concentration was estimated to be 40ng/µL. (C) Plasmids isolated from pGEMT-RMND5B clones 1 to 6 were digested with BamHI to release the ~1.2kb insert and the products electrophoresed in a 1% agarose gel. Inserts were identified in all clones.

Lane 1: MW Marker Lane 2: 1mM MgCl2 Lane 3: 1.5mM MgCl2 Lane 4: 2mM MgCl2 Lane 5: Negative Control

Lane 1: MW Marker Lane 2: RMND5B Lane 3: Negative Control

~1.2kb

~1.2kb

A

Lane 1: MW Marker Lane 2, 4, 6, 8, 10, 12: Undigested pGEMT-RMND5B clones 1-6 Lane 3, 5, 7, 9, 11, 13: BamHI digested pGEMT-RMND5B clones 1-6

~1.2kb

~3kb

C


254

RNase treated then BamHI digested to release the ~1.2kb insert and the products

electrophoresed in a 1% agarose gel (Section 3.6, 3.8.2, 3.9, Figure 6.11C). The ~1.2kb

RMND5B insert was identified in all 6 clones, and to confirm the correct insert

sequences, pGEMT-RMND5B clones 1-3 were purified then sequenced using M13-S

and M13-AS primers (Appendix II, Section 3.12), with BLASTTM analysis of the

chromatograms identifying pGEMT-RMND5B clone 1 to be mutation free (not shown).

To subclone RMND5B into the pmCherry-C1 expression vector, the RMND5B insert

from pGEMT-RMND5B clone 1 was released by BamHI digestion, the insert was gel

purified and 5µL purified plasmid was electrophoresed in a 1% agarose gel (Section 3.6,

3.7.2, 3.8.2, Figure 6.12A). pmCherry-C1 was prepared by BamHI digestion and SAP

treatment, and 5µL purified plasmid was electrophoresed in a 1% agarose gel (Section

3.6, 3.8.2, 3.8.3, Figure 6.12B). Sixty ng (3µL) RMND5B insert was ligated into 50ng

(1µL) pmCherry-C1 and the ligation products were transformed into competent E. coli

DH5α then selected by growth on LB Agar/Kanamycin (Section 3.8.4, 3.8.6). Plasmids

purified from 16 colonies were digested with RNase then BamHI and an aliquot of each

plasmid was electrophoresed in a 1% agarose gel, from which ~1.2kb (RMND5B)

inserts were identified in pmCherry-RMND5B clones 2, 7 and 11 (Section 3.6, 3.8.2,

3.9, Figure 6.12C). These clones were purified, digested with KpnI to determine insert

orientation and the products were electrophoresed in a 2% agarose gel (Section 3.6,

3.7.1, 3.8.2, Figure 6.12D). The presence of a ~1kb band upon digestion of pmCherry-

RMND5B clone 7 indicated that the RMND5B insert was in the sense orientation

(Figure 6.12D). pmCherry-RMND5B clone 7 was sequenced using the

RMND5BBamHI1-S, RMND5BBamHI1182-AS and RMND5B790-S primers and

BLAST™ analysis of the chromatograms identified that this clone was mutation free

(Section 3.12, Appendix II, Appendix III).

To prepare pmCherry-RMND5B for transfection, a large scale plasmid preparation of

pmCherry-RanBPM was performed and an aliquot of the purified plasmid

electrophoresed in a 1% agarose gel (Section 3.6, 3.10, Figure 6.12E). The plasmid was

also BamHI digested to verify the presence of the ~1.2kb RMND5B insert and the

concentration of the plasmid was determined to be 1.6µg/µL (Section 3.6, 3.8.2, 3.10,

Figure 6.12E). To investigate expression of Cherry-RMND5B, LNCaP cells were

transfected with 4µg pmCherry-RanBPM and at 48 hours post-transfection the cells


255

14

~5.9kb ~5.2kb

~1kb

~280bp

Lane 1: MW Marker Lane 2, 4, 6: Undigested pmCherry-RMND5B Clones 2, 3, 11 Lane 3, 5, 7: KpnI digested pmCherry-RMND5B Clones 2, 3, 11

D

~1.2kb

Lane 1: MW Marker Lane 2: RMND5B (clone 1)

~5kb

Lane 1: MW Marker Lane 2: Undigested pmCherry C1 Lane 3: BamHI/SAP digested pmCherry C1

A

C

~5kb ~1.2kb

~5kb

~1.2kb

Lanes 1, 18: MW Marker Lanes 2, 4, 6, 8, 10, 12, 14, 16, 19, 21, 23, 25, 27, 29, 31: Undigested pmCherry-RMND5B clones 1-16 Lanes 3, 5, 7, 9, 11, 13, 15,17, 20, 22, 24, 26, 28, 30, 32: BamHI digested pmCherry-RMND5B clones 1-16

1 2 3 4 5 6 7 8 9 10 11 12 13 15 16 17

18 19 20 21 23 24 25 26 27 28 29 30 31 32 22

B


256

Cherry

β-actin

~75kDa

~45kDa

F

Figure 6.12: Cloning of RMND5B into pmCherry-C1. (A) pGEMT-RMND5B clone 1 was digested with BamHI to release the insert, the ~1.2kb insert was gel purified and 5µL was electrophoresed in a 1% agarose gel from which the concentration was estimated to be ~20ng/µL. (B) pmCherry-C1 was prepared by digesting the plasmid with BamHI and SAP, followed by purification and electrophoresis of 5µL prepared plasmid in a 1% agarose gel. The plasmid concentration was estimated to be ~50ng/µL. (C) pmCherry clones 1-16 were digested with BamHI which identified that clones 2, 7 and 11 contained the ~1.2kb RMND5B insert. (D) To determine the orientation of the RMND5B insert, pGEMT-RMND5B clones 2, 7 and 11 were digested with KpnI and electrophoresed in a 1% agarose gel. Digestion of clones 2 and 11 produced bands at approximately 5.9kb and 280bp (antisense orientation), while KpnI digestion of clone 7 resulted in a ~1kb band (sense orientation). (E) Large scale plasmid preparation of pmCherry-RMND5B was performed and BamHI digestion released the ~1.2kb insert. (F) Expression of Cherry-RMND5B was confirmed by transfection of LNCaP cells with pmCherry-RMND5B and anti-Cherry western blotting performed using lysates prepared at 48 hours post-transfection.

~4.7kb

~1.2kb

Lane 1: MW Marker Lane 2: Undigested pmCherry-RMND5B Lane 3: BamHI digested pmCherry-RMND5B

E


257

were harvested, an aliquot electrophoresed in a 12% polyacrylamide gel and Cherry

western blotting was performed (Section 3.1.4, 3.15). Western blotting identified a

~75kDa band corresponding to Cherry-RMND5B in lysates from transfected cultures

only, confirming expression of the Cherry-RMND5B fusion protein (Figure 6.12F).

6.2.5.2 RMND5A and RMND5B Interact in LNCaP Cells

To examine whether RMND5 proteins interact in LNCaP cells, cells growing in 10cm

petri dishes were cotransfected with plasmids encoding GFP-RMND5A and Cherry-

RMND5B (Section 3.1.4). At 48 hours post-transfection, the cells were lysed, an aliquot

taken (total cellular input) and the lysate immunoprecipitated with GFP antibodies

(Section 3.13). GFP immunocomplexes were electrophoresed in 12% polyacrylamide

gels and GFP and Cherry western blotting was performed (Section 3.15). GFP western

blotting identified a ~70kDa band corresponding in size to GFP-RMND5A in the total

cellular input and in immunoprecipitated samples, indicating successful GFP

immunoprecipitation (Figure 6.13A). No bands were present in lysates from

untransfected cells or in mock immunoprecipitated samples. Cherry western blotting

identified a band corresponding to Cherry-RMND5B in the total cellular input and the

immunoprecipitated samples, indicating that RMND5A interacts with RMND5B

(Figure 6.13A). Again, no bands were detected in lysates from untransfected cells and

in mock immunoprecipitated samples.

6.2.5.3 RMND5A and RMND5B Colocalise in LNCaP Cells

To assess whether RMND5 proteins colocalise, LNCaP cells growing on coverslips

were cotransfected with plasmids encoding GFP-RMND5A and Cherry-RMND5B and

at 48 hours post-transfection the cells were prepared for fluorescence microscopy

(Section 3.1.3, 3.1.4, 3.16). Both RMND5 proteins exhibited a diffuse nuclear and

cytoplasmic distribution when coexpressed, which was consistent with previous results

in cells overexpressing either RMND5A or RMND5B alone (Section 4.2.6.4, 6.13B). In

a proportion of cells coexpressing GFP-RMND5A and Cherry-RanBPM, both proteins

also exhibited a punctate cytoplasmic distribution. This was more pronounced compared

to cells overexpressing only one of the RMND5 proteins although it was not possible to

quantitate these results due to a lack of suitable equipment (Figure 6.13B). The

colocalisation of RMND5 proteins in the nucleus and cytoplasm and in punctate

cytoplasmic speckles was indicated by yellow staining when the GFP-RMND5A and


258

WB: Cherry (RMND5B)

WB: GFP (RMND5A)

IP: GFP (RMND5A)

~75kDa

~70kDa

Figure 6.13: RMND5A and RMND5B interact and colocalise in LNCaP cells. (A) To assess whether RMND5A and RMND5B interact, LNCaP cells were transiently transfected with plasmids encoding GFP-RMND5A and Cherry-RMND5B. At 48 hours post-transfection, the cells were lysed and GFP-RMND5A was immunoprecipitated. GFP and Cherry western blotting identified that GFP-RMND5A and Cherry-RMND5B were both present in the total cell lysate and immunoprecipitated samples. (B) RMND5 protein colocalisation was determined by cotransfection of LNCaP prostate cancer cells with plasmids encoding GFP-RMND5A and Cherry-RMND5B and preparation of cells for fluorescence microscopy at 48 hours post-transfection. GFP-RMND5A and GFP-RMND5B displayed colocalisation in the nucleus and cytoplasm and in punctate cytoplasmic speckles (Magnification x1000). Experiments were performed twice and representative results are shown.

A

B GFP-RMND5A Cherry-RMND5B Hœchst 33258 Overlay

GFP-RMND5A Cherry-RMND5B Overlay Hœchst 33258

-ve control (488nm) Hœchst 33258 Overlay -ve control (546nm)


259

Cherry-RMND5B images were overlayed. These findings therefore demonstrated that

RMND5A and RMND5B colocalise in LNCaP cells.

6.2.6 Mass Spectrometric Identification of RMND5 Binding Partners

Both RMND5A and RMND5B were found in this thesis to possess E3 ubiquitin ligase

activity and to ubiquitinate the prostatic tumour suppressor NKX3.1, targeting it for

proteasomal degradation. As NKX3.1 expression is localised to the prostate and

RMND5 proteins are widely expressed, they are potentially able to ubiquitinate proteins

in the other tissue types in which they are expressed, and in all tissues would be able to

ubiquitinate multiple proteins. Furthermore, RMND5 proteins contain multiple protein-

protein interaction domains, suggesting that they possess cellular functions distinct from

their E3 ubiquitin ligase activity. The identification of RMND5 binding partners may

elucidate additional cellular roles of RMND5 proteins, for example by determining the

outcome(s) of the interaction. To commence this investigation, which may be continued

in future studies, RMND5 binding partners were identified using mass spectrometry.

For these studies 1D nano liquid chromatography electrospray ionisation-MS/MS was

used to determine proteins co-immunoprecipitating with either GFP-RMND5A or GFP-

RMND5B in LNCaP cells.

6.2.6.1 Identification of RMND5A Binding Proteins

To determine RMND5A binding partners, LNCaP cells growing in 4 x 10cm petri

dishes were transfected with pEGFP-RMND5A and at 48 hours post-transfection, the

cells were lysed, an aliquot was taken and the lysates were combined and

immunoprecipitated using GFP antibodies bound to Protein A beads (Section 3.1.4,

3.13). For the control, 4 petri dishes of untransfected cells were treated in the same

manner (Section 3.13). Samples were electrophoresed in 12% polyacrylamide gels and a

5µL aliquot was separately analysed by GFP western blotting, which identified a

~70kDa band corresponding in size to GFP-RMND5A in the total input and

immunoprecipitated sample, confirming successful GFP immunoprecipitation (Section

3.15, Figure 6.14A). A band corresponding to GFP-RMND5A was also present in the

unbound fraction, indicating that GFP-RMND5A immunoprecipitation was incomplete.

No bands were present in the mock immunoprecipitated samples. The remaining

immunoprecipitate, which was also electrophoresed in a 12% polyacrylamide gel, was

stained with Coomassie blue to visualise immunoprecipitated proteins (Section 3.15).


260

Eight bands present in the immunoprecipitate which were not present in the mock

immunoprecipitated control were excised from the gel, dried and sent to the Australian

Proteome Analysis Facility (APAF) for mass spectrometric analysis (Section 3.17,

Figure 6.14B). At APAF, all bands were cut into smaller pieces, destained, dried, then

trypsin digested from which the peptides were extracted using acetonitrile/formic acid

then concentrated ready for analysis by 1D nanoLC ESI-MS/MS (Section 3.17). Band 1

was analysed separately, and a small portion of each of bands 1-8 were combined and

analysed by 1D nanoLC ESI-MS/MS, resulting in the identification of multiple proteins

from the peptide fragments by analysis of the raw data using MASCOT, a program that

uses mass spectrometry data to identify proteins from primary sequence databases. To

screen the results, all peptide matches to a particular protein were individually analysed,

with those peptides having an individual ion score of >30 from the mass spectrometry

data indicating identity or extensive homology (p<0.05) to the matched protein.

Additionally, all proteins represented by two or more unique peptides with an individual

ion score >30 were considered to be positively identified in the screen.

The ~70kDa band 1, which corresponded in size to GFP-RMND5A, was found to

contain RMND5A (score = 369) when the raw mass spectrometry data was analysed

using MASCOT (Appendix IV, Figure 6.14C) (Perkins et al., 1999). RMND5A was the

seventh most abundant protein identified in this gel band and upon analysis, the peptide

coverage of RMND5A was 33% with 9 unique peptides (individual ion score >30)

(Appendix IV, Figure 6.14C, D). RMND5A was also identified in the combined

analysis of bands 1-8 (score = 113), with 4 unique peptides (individual ion score >30)

which were also present in band 1. The remaining list of identified proteins (from band

1 and bands 1-8) was then filtered against two contaminants databases, the Max Planck

Institute contaminants database (www.mpg.de/en) and the common repository of

adventitious proteins (cRAP) database (www.thegpm.org/crap/index.html), which

resulted in the removal of keratins and other proteins that are considered to be external

contaminating proteins and resulting in the identification of multiple putative RMND5A

binding partners (Table 6.2).


261

1 2 3 4 5

4 3

1 7

6 5 2 8

A

B

Lane 1: Total input (mock immunoprecipitated control) Lane 2: Total input (immunoprecipitated sample) Lane 3: Molecular weight marker Lane 4: Mock immunoprecipitation control Lane 5: Immunoprecipitated sample C


262

Figure 6.14: Immunoprecipitation of GFP-RMND5A and its binding partners. To identify RMND5A binding partners, LNCaP cells were transfected with plasmid encoding GFP-RMND5A and at 48 hours post-transfection, GFP immunoprecipitation was performed followed by electrophoresis of the immunoprecipitated proteins in 12% polyacrylamide gels. (A) GFP western blotting identified a ~70kDa band corresponding to GFP-RMND5A in both the total input and immunoprecipitated samples. (B) Coomassie blue staining identified multiple bands in the immunoprecipitated sample that were not present in the mock immunoprecipitated control. These bands (bands 1-8) were excised and analysed by 1D nano LC ESI/MS/MS to identify GFP-RMND5A and its binding partners (C) RMND5A was the seventh most abundant protein in band 1 with a score of 369 and 9 unique peptides corresponding to RMND5A identified. (D) Analysis of the RMND5A matches identified peptides distributed over the length of the protein and covering 33% of the protein sequence (red). Numbers show amino acid residues.

D

MDQCVTVERELEKVLHKFSGYGQLCERGLEELIDYTGGLKHEILQSHGQDAELSGTLSLVLTQCCKRIK

DTVQKLASDHKDIHSSVSRVGKAIDKNFDSDISSVGIDGCWQADSQRLLNEVMVEHFFRQGMLDVAEEL

CQESGLSVDPSQKEPFVELNRILEALKVRVLRPALEWAVSNREMLIAQNSSLEFKLHRLYFISLLMGGT

TNQREALQYAKNFQPFALNHQKDIQVLMGSLVYLRQGIENSPYVHLLDANQWADICDIFTRDACALLGL

SVESPLSVSFSAGCVALPALINIKAVIEQRQCTGVWNQKDELPIEVDLGKKCWYHSIFACPILRQQTTD

NNPPMKLVCGHIISRDALNKMFNGSKLKCPYCPMEQSPGDAKQIFF

1

391


263

Table 6.2 – Candidate RMND5A binding partners identified by mass spectrometry

Protein Abbreviation/Name No. of Unique Peptide Hits (band 1/all bands) *

Required for Meiotic Nuclear Division 5A RMND5A 10 and 4

Cytoskeletal

Tubulin TBA1A 5

TBA4A 10

TBB5 10

Actin ACTB 12

Lamin Lamin B1 1

Myosin Unconventional myosin VI 2

Myosin regulatory light chain 10

1

Microtubule Microtubule-actin cross linking factor 1 isoform 1/2/3/5

1

Merlin Merlin 3

Flotillin 2 FLOT2 1

Mitochondrial

ATP synthase subunit (Ox Phos) ATPα 4

ATPβ 8

Isocitrate dehydrogenase (NADP) ICD 1

Phosphoenolpyruvate carboxykinase PEPCK/ PCKGM 1

Dihydrolipoyllysine-residue acetyltransferase component of pyruvate dehydrogenase complex

ODP2 0

Stress 70 protein (mitochondrial HSP) GRP75 9 and 1

RNA Processing

Heterogeneous nuclear ribonuclearprotein

HNRNPM 11

HNRNPQ 1

HNRNPC 8

HNRNPF 2

HNRNPH1 4

Probable ATP-dependent RNA-helicase DDX41 9 and 4


264

DDX5 3

DHX35 3

ATP-dependent RNA helicase DDX39A

Polyadenylate binding protein PABP1 8 and 2

PABP5 1

KH domain-containing RNA binding, signal transduction associated protein 1

KHDR1 1

THO complex subunit 3 THOR3 0

SNW domain containing protein 1 SNW1 6

Eukaryotic initiation factor IF4A3 9

IF4A1 1

Elongation factor 1α EF1A1 1

RNA binding Raly-like protein RALYL 1

Splicing factor SF3B3 2

SPF45 1

Poly(U)-binding splicing factor PUF60 18 and 11

Crooked neck-like protein 1 CRNL1 1

Poly (rC) binding protein 1 PCBP1 0

Heat Shock Proteins HSP71 9 and 4

HS71L 0

Heat shock cognate 71kDa HSPC7C 24 and 15

78kDa glucose regulated protein GRP78 9

Miscellaneous

X-ray repair cross complementing protein 6

XRCC6 4

Histone H3 1

Ubiquitin-40S ribosomal protein S27a RS27A 2 and 1

Exportin 2 XPO2 1

Interleukin enhancer binding factor 2 2

Inositol hexakisphosphate and diphosphoinositol-pentakisphosphate kinase 2

VIP2 1

Uncharacterised protein C14orf166B C14orf166B 0

POTE ankyrin domain family member E POTEE 1

Ubiquitin-40S ribosomal protein S27a RS27A 2

* Peptide individual ion score >30


265

The remaining proteins were filtered for proteins that bind sepharose and magnetic

beads, including cytoskeletal proteins, DEAD box proteins, hnRNPs, and eukaryotic

translation and initiation factors and those proteins that bind the GFP tag alone (Trinkle-

Mulcahy et al., 2008). The remaining proteins were then classified according to their

cellular localisation and/or role (Table 6.3), resulting in the identification of six

potential RMND5A binding partners, two of which are mitochondrial proteins, one of

which is a cytoplasmic protein and the remainder of which are nuclear proteins (Table

6.3). None of the identified proteins were CTLH complex components.

6.2.6.2 Investigation of a Putative RMND5A Binding Partner

Further investigation of the putative RMND5A binding partners identified by mass

spectrometry would involve confirmation of their interaction with RMND5A using

coimmunoprecipitation assays and specific antibodies to individual proteins. As the

number of significant peptide hits identified in the mass spectrometry screen for PUF60

were high, the peptide coverage of PUF60 from peptides identified in the mass

spectrometry screen was 44% and a PUF60 antibody was commercially available, it was

selected for further analysis (Figure 6.15A). Following purchase of the antibody, PUF60

western blotting was performed using LNCaP whole cell lysates and the optimum

primary antibody concentration was determined to be 1:1000 (Section 3.15, not shown).

To confirm the mass spectrometry results, LNCaP cells were transfected with plasmid

encoding GFP-RMND5A and at 48 hours post-transfection, the cells were lysed, an

aliquot taken (total input) and the remaining lysate was immunoprecipitated using GFP

antibodies (Section 3.1.4, 3.13). Samples were electrophoresed in 12% polyacrylamide

gels and analysed by GFP and PUF60 western blotting (Section 3.15). GFP western

blotting identified a ~70kDa band corresponding in size to GFP-RMND5A in the total

cellular input and immunoprecipitated fractions, indicating successful GFP-RMND5A

immunoprecipitation (Figure 6.15B). No GFP-immunoreactive bands were detected in

the mock immunoprecipitated control or the unbound fractions. Western blotting for

PUF60 identified a ~60kDa band in the total cellular input, the mock

immunoprecipitated control and immunoprecipitated sample fractions (Figure 6.15B).

These results indicated that PUF60 bound strongly to the bead matrix and was not a

specific GFP-RMND5A binding partner. During the time frame of this thesis, no

additional candidate RMND5A interactors have been investigated.


266

Table 6.3 – Function of RMND5A binding proteins identified by mass spectrometry

Protein No. of

Unique Peptide Hits*

Protein Coverage

Function Cellular Localisation

RMND5A 10 and 4 33% Member of the CTLH complex Nuclear/ cytoplasmic

Poly(U)-splicing factor PUF60

18 and 11

44% A splicing factor involved in splicing pre-mRNA that recognises 3’ splice sites. Together with another splicing factor U2AF65 enhances splicing. PUF60 can replace U2AF65 in vitro.

Nuclear

ATP synthase subunits α and β, mitochondrial

4 and 8 12.3% and 22%

Both form part of the catalytic core (F1) of the mitochondrial ATP synthase which synthesise ATP during oxidative phosphorylation.

Mitochondria

SNW domain containing protein 1 SNW1

6 17.4% SNW1/Ski interacting protein (SKIP), interacts with the Ski oncoprotein. SNW1 is a transcriptional coactivator or repressor. It binds the ligand binding domain of vitamin D and retinoic acid receptors, augmenting vitamin D, retinoic acid, oestrogen and glucocorticoid gene expression. SNW1 also functions as a splicing factor by interacting with U2AF65, recruiting it to p21 pre-mRNA. SNW1 controls BMP signalling in embryogenesis.

Mainly nuclear some light cytoplasmic

X-ray repair cross-complementing protein 6 XRCC6

4 13.6% XRCC6/ Ku70 plays a role in telomere maintenance by functioning as an ATP dependent DNA dependent helicase (together with Ku80 XRCC6 forms the Ku autoantigen nuclear complex), which is involved in DSB repair and antibody VDJ recombination, both by the NHEJ DNA repair pathway.

Nuclear

Ubiquitin-40S ribosomal protein S27a

2 21.1% Fusion protein between ubiquitin and ribosomal protein S27a. Post-translationally modified to produce ubiquitin and S27a, a component of the 40S ribosome.

Cytoplasmic

*Peptide individual ion score >30


267

Figure 6.15: Identification of putative GFP-RMND5A binding partners. PUF60 was the fourth most abundant protein present in band 1 (score=483) and the most abundant protein present when all bands in the immunoprecipitated sample were combined (bands 1-8, score=1245) for analysis. (A) Peptides matching PUF60 were located throughout the protein and covered 44% of the PUF60 amino acid sequence. (B) To assess whether PUF60 was a GFP-RMND5A binding partner, LNCaP cells were transfected with plasmid encoding GFP-RMND5A and harvested at 48 hours post-transfection for GFP immunoprecipitation. Electrophoresis of the immunoprecipitated proteins in 12% polyacrylamide gels and GFP western blotting identified a ~70kDa band corresponding in size to GFP-RMND5A in both the total input and immunoprecipitated sample. PUF60 western blotting identified a ~60kDa band in the total input, immunoprecipitated and mock immunoprecipitated control fractions indicating that PUF60 nonspecifically binds to the Protein A beads. Numbers indicate amino acid residues.

A PUF60

B

MATATIALQVNGQQGGGSEPAAAAAVVAAGDKWKPPQGTDSIKMENGQSTAAKLGLPPLTPEQQEALQKA

KKYAMEQSIKSVLVKQTIAHQQQQLTNLQMAAVTMGFGDPLSPLQSMAAQRQRALAIMCRVYVGSIYYEL

GEDTIRQAFAPFGPIKSIDMSWDSVTMKHKGFAFVEYEVPEAAQLALEQMNSVMLGGRNIKVGRPSNIGQ

AQPIIDQLAEEARAFNRIYVASVHQDLSDDDIKSVFEAFGKIKSCTLARDPTTGKHKGYGFIEYEKAQSS

QDAVSSMNLFDLGGQYLRVGKAVTPPMPLLTPATPGGLPPAAAVAAAAATAKITAQEAVAGAAVLGTLGT

PGLVSPALTLAQPLGTLPQAVMAAQAPGVITGVTPARPPIPVTIPSVGVVNPILASPPTLGLLEPKKEKE

EEELFPESERPEMLSEQEHMSISGSSARHMVMQKLLRKQESTVMVLRNMVDPKDIDDDLEGEVTEECGKF

GAVNRVIIYQEKQGEEEDAEIIVKIFVEFSIASETHKAIQALNGRWFAGRKVVAEVYDQERFDNSDLSA

1

559


268

6.2.6.3 Identification of RMND5B Binding Proteins

To identify RMND5B binding partners, LNCaP cells growing in 4 x 10cm petri dishes

were transfected with plasmid encoding GFP-RMND5B and at 48 hours post-

transfection, the cells were lysed, a total cellular input sample was taken and the

remaining lysate was immunoprecipitated using GFP antibodies (Section 3.1.4, 3.13).

For the mock immunoprecipitation controls, the same procedure was performed using

untransfected LNCaP cells (Section 3.13). A 5µL aliquot of each sample was

electrophoresed in a 12% polyacrylamide gel and analysed by GFP western blotting,

which identified a ~70kDa band corresponding in size to GFP-RMND5B in the total

input and immunoprecipitated samples, indicating successful immunoprecipitation

(Figure 6.16A). No bands were present in the mock immunoprecipitation control sample

or the unbound fraction. The remainder of the immunoprecipitated samples were

electrophoresed in a precast 12% polyacrylamide gel (to minimise the presence of

contaminating proteins) and stained with Coomassie blue to visualise the proteins

(Section 3.15, Figure 6.16B). The mock immunoprecipitated control contained a

prominent band at ~70kDa as well as a number of minor bands and the lane was divided

into 4 bands (Figure 6.16B). The immunoprecipitated sample, which was

electrophoresed in two lanes due to its higher volume (~60µL), was separated into 6

bands. Band 1 comprised a section of the gel at ~70kDa from both lanes and the

remainder of lane 3 was separated into 5 bands (bands 2-5), each of which was dried

and sent to APAF for mass spectrometric identification of the protein bands (Section

3.17, Figure 6.16B).

For mass spectrometry analysis at APAF, the mock immunoprecipitated control bands

(Mock bands 1-4, Figure 6.16B) and immunoprecipitated sample bands (IP bands 1-6,

Figure 6.16B) were cut into smaller fragments, destained, dried, trypsin digested and the

peptides extracted with acetonitrile/formic acid and then concentrated (Section 3.17).

Five µL of the mock immunoprecipitated bands (M1-4) were combined and 5µL of the

immunoprecipitated sample bands (IP 1-6) were combined and from each combined

sample, a 5µL aliquot was analysed separately by 1D nanoLC ESI-MS/MS. The raw

data from the mass spectrometry screen was analysed by MASCOT, resulting in the

identification of multiple peptides corresponding to numerous proteins (Appendix IV)

(Perkins et al., 1999). The peptide matches to a particular protein were analysed

individually. An ion score of >30 for a peptide from the mass spectrometry data

indicated identity or extensive homology (p<0.05) to the matched protein. Where two or


269

more unique peptides were identified with an ion score >30 for a particular protein, it

was considered a positive identification of that protein.

The mock immunoprecipitated control sample contained proteins known to be external

contaminants as well as proteins that have been reported to bind non-specifically to the

magnetic bead matrix used in the immunoprecipitation reaction (Trinkle-Mulcahy et al.,

2008). PUF60 was identified among these proteins (score = 1105) as were other splicing

factors, multiple keratins, heat shock proteins belonging to the hsp70 family, hnRNPs,

the cytoskeletal protein actin, eukaryotic transcription initiation and elongation factors

and DEAD box proteins (Appendix IV). Similarly, the immunoprecipitated sample also

yielded proteins that have been determined to be external contaminants and to bind the

bead matrix utilised. In the initial analysis, all of the proteins identified in the mass

spectrometric screens to present in both the immunoprecipitated sample and the mock

immunoprecipitated control were removed. The list of remaining that

immunoprecipitated with GFP-RMND5B was filtered against two contaminants

databases, the Max Planck Institute contaminants database (www.mpg.de/en) and the

common repository of adventitious proteins (cRAP) database

(www.thegpm.org/crap/index.html). The remaining proteins in the GFP-RMND5B

immunoprecipitated sample were classified according to their cellular localisation

and/or role (Table 6.4).

Of the proteins identified in the immunoprecipitated but not mock immunoprecipitated

samples, albumin and Elongation factor 1 alpha 1 have been identified as binding to

sepharose bead proteomes and therefore are considered unlikely to be specific binding

partners of GFP-RMND5B at this stage (Trinkle-Mulcahy et al., 2008). RMND5B

(score = 70) was identified in this screen from two unique peptides with individual ion

scores of >30, which gave a peptide coverage of 4.6% for RMND5B (Figure 6.16C, D).

One protein identified in this screen that obtained two unique peptide matches

(individual ion score >30), was ubiquitin-40s ribosomal protein S27a (score = 97) which

yielded a peptide coverage of 21.1% and also immunoprecipitated with GFP-RMND5A

(Figure 6.16E, F, Table 6.3). Other proteins identified obtained unique peptide matches

of 1, which is considered not to represent a positive identification for a particular

protein, however these results will be useful in the analysis of future mass spectrometric

investigations of RMND5B interacting partners. Although many proteins were removed

from the list of potential RMND5B (and RMND5A) interacting proteins due to their


270

inclusion in lists of known contaminants in this type of assay, both RMND5A and

RMND5B are multidomain proteins that may function in nucleocytoplasmic shuttling or

in other roles involving the cytoskeleton, and therefore specific interaction of RMND5

proteins with a subset of these proteins may be examined in future studies.

Table 6.4 – Putative RMND5B binding partners identified by mass spectrometry

Protein Number of Unique Peptide Hits*

Function Cellular Localisation

RMND5B 2 Unknown Nuclear/ cytoplasmic

Ubiquitin-40s ribosomal protein S27a

2 Ubiquitin-S27A fusion protein post-translationally processed to yield ubiquitin and the ribosomal 40S component S27A.

Cytoplasmic

POTE ankryin domain family member E

1 POTE proteins are expressed in prostate, testis, ovary and placenta. Ten homologues exist with a proposed signalling function.

Plasma membrane

Tubulin beta-4-A chain

1 A beta tubulin isoform, tubulins are a component of microtubules which comprise part of the cytoskeletal structure.

Cytoplasmic

Neprilysin 1 Membrane metallo-endopeptidase. Cytoplasmic

ATP synthase subunit beta

1 Forms part of a mitochondrial ATP synthase. Mitochondrial

Unconventional myosin-VI

1 Unconventional myosin VI is an ATPase that uses actin for the intracellular transport of vesicles and is involved in cell migration.

Nuclear/ cytoplasmic

Elongation factor 1-alpha 1

1 Protein biosynthesis/With PARP1 and TXK forms part of a transcription factor complex that regulates IFN-gamma expression.

Nuclear/ cytoplasmic

Serum albumin 3 Plasma carrier for steroid hormones, heme and fatty acids/Essential for the maintenance of osmotic pressure.

Plasma

* Peptide individual ion score >30


271

A

1 2 3 4 5

M1

M2

M3

M4

IP2 IP3 IP1 IP4

IP5

IP6

B

Lane 1: Molecular weight marker Lane 2: Mock immunoprecipitated control Lane 3: Molecular weight marker Lane 4: Immunoprecipitated sample 1 Lane 5: Immunoprecipitated sample 2

C


272

MEQCACVERELDKVLQKFLTYGQHCERSLEELLHYVGQLRAELASAALQGTPLSATLSLVMS

QCCRKIKDTVQKLASDHKDIHSSVSRVGKAIDRNFDSEICGVVSDAVWDAREQQQQILQMAI

VEHLYQQGMLSVAEELCQESTLNVDLDFKQPFLELNRILEALHEQDLGPALEWAVSHRQRLL

ELNSSLEFKLHRLHFIRLLAGGPAKQLEALSYARHFQPFARLHQREIQVMMGSLVYLRLGLE

KSPYCHLLDSSHWAEICETFTRDACSLLGLSVESPLSVSFASGCVALPVLMNIKAVIEQRQC

TGVWNHKDELPIEIELGMKCWYHSVFACPILRQQTSDSNPPIKLICGHVISRDALNKLINGG

KLKCPYCPMEQNPADGKRIIF

Figure 6.16: Immunoprecipitation of GFP-RMND5B and its binding partners. To identify RMND5B interacting proteins, LNCaP cells were transfected with pEGFP-RMND5B and at 48 hours post-transfection, GFP immunoprecipitation was performed followed by electrophoresis of the immunoprecipitated proteins in a precast 12% polyacrylamide gel. (A) GFP western blotting identified a ~70kDa band corresponding to GFP-RMND5B in both the total input (barely visible in image) and immunoprecipitated sample. (B) Coomassie blue staining identified multiple bands in the immunoprecipitated and mock immunoprecipitated sample. The mock immunoprecipitated control sample was divided into 4 bands (M1-4), and the immunoprecipitated sample, which was electrophoresed in two lanes, was separated into eleven bands, six (IP1-6) of which were analysed by mass spectrometry. (C) Two unique peptides were matched to RMND5B (score=70) using MASCOT. (D) Analysis of the peptide matches identified a protein coverage of 4.6%. (E) Ubiquitin-40S ribosomal protein S27a (score=97) obtained two unique peptide matches in the mass spectrometry screen for GFP-RMND5B binding partners. (F) The peptide matches for ubiquitin-40S ribosomal S27a produced a protein coverage of 21.1%. Numbers indicate amino acid residues.

D RMND5B

F Ubiquitin-40S ribosomal protein S27a

MQIFVKTLTGKTITLEVEPSDTIENVKAKIQDKEGIPPDQQRLIFAGKQLEDGRTLSDYNIQ

KESTLHLVLRLRGGAKKRKKKSYTTPKKNKHKRKKVKLAVLKYYKVDENGKISRLRRECPSD

ECGAGVFMASHFDRHYCGKCCLTYCFNKPEDK

E

1

393

1

156


273

6.3 Discussion

In this chapter, expression of the CTLH complex members in prostate cancer cells was

evaluated using RT-PCR. The method was used to rapidly and qualitatively identify the

expression of the CTLH complex components, however since only mRNA levels were

examined, protein levels of the complex members in the prostate and breast cancer cells

lines used are unknown at this stage. Western blotting was not able to be used to

determine expression of each of the CTLH complex members for this thesis as suitable

antibodies against each complex component are not commercially available. However,

to continue investigation of the CTLH complex members in future studies, real time

PCR (RT-qPCR) may be used to quantitate mRNA levels (VanGuilder et al., 2008) and

the results can be augmented with western blotting as suitable antibodies are generated.

The finding that each complex component is expressed in two prostate and one breast

cancer cell line suggests that it is possible for the CTLH complex to form in these cell

types as has been demonstrated in HEK293 cells (Kobayashi et al., 2007). It is also

possible that the complex exists in many cell types as a large ~670kDa RanBPM

associated complex has been detected in a number of cell lines including HeLa cells

(Nishitani et al., 2001; Ideguchi et al., 2002; Umeda et al., 2003). Members of the

CTLH complex are also widely expressed with EMP, Twa1, muskelin and RanBPM

expression documented in a variety of human and mouse tissues and RanBPM

exhibiting highest expression in the prostate, ovaries and testes (Adams et al., 1998;

Rao et al., 2002; Umeda et al., 2003; Bala et al., 2006).

Interaction of RMND5A and RMND5B with RanBPM was investigated in this study as

a measure of whether the CTLH complex could form in prostate cancer cells using

either RMND5A or potentially RMND5B. These experiments were performed due to

the finding that the yeast orthologue of RanBPM, Vid30 functions as a core component

of the Vid30 complex, and, based on its multidomain structure RanBPM may fulfil a

similar role in the human CTLH complex (Pitre et al., 2006). The formation of the

CTLH complex was identified in initial reports using sucrose gradient sedimentation

and coimmunoprecipitation assays due to the association of the CTLH complex

components with RanBPM, further supporting the hypothesis that RanBPM is a core

component of the complex (Kobayashi et al., 2007). Similarly, RanBPM has been

hypothesised to function as a scaffolding protein, again supporting its role as an integral

member of the CTLH complex (Murrin and Talbot, 2007). Glycerol gradient

sedimentation has been used to demonstrate that yeast RMD5 forms part of a large


274

~600kDa protein complex, whilst in human HEK293 cells RMND5A was identified as

a CTLH complex member, with both of these findings supporting the inclusion of

RMND5 proteins in the CTLH complex (Regelmann et al., 2003; Kobayashi et al.,

2007). Both RMND5 proteins were confirmed to interact with RanBPM in prostate

cancer cells, providing additional evidence that the CTLH complex may form in this

cell type and further that RMND5B may join or replace RMND5A in the CTLH

complex. Thus, although RMND5 proteins were only identified in this thesis to interact

with a single core member of the CTLH complex, the results suggest that the CTLH

complex can form in prostate cancer cells and provide a starting point for additional

assays confirming the existence of the complex in these cells.

Additional methods for the detection of the CTLH complex in prostate cancer cells

include co-immunoprecipitation assays with each complex member, and although as

mentioned previously antibodies against endogenous members of the complex are not

commercially available, these studies may be performed following cloning and

transfection of the individual complex components into prostate cancer or other cell

types. Results may be confirmed by sucrose gradient centrifugation of the high

molecular weight CTLH complex from prostate cancer cells with mass spectrometry

used to identify complex components (Kobayashi et al., 2007). In the present study,

novel binding partners of RMND5 proteins were isolated by immunoprecipitation of

RMND5A/RMND5B and their interacting partners, with the coimmunoprecipitated

proteins identified using mass spectrometry. Although this is a commonly used method

for the identification of novel proteins, members of the CTLH complex were not were

not detected, which may have resulted from weak or transient interaction between the

complex components or the low levels of expression of complex members in this cell

type (Vasilescu et al., 2004). It is also feasible that the complex forms under specific

cellular conditions that remain to be identified, as has been shown for the yeast Vid30

complex, which is only activated under changing nutrient environments (Santt et al.,

2008).

The interaction of RMND5 proteins with RanBPM was used as an indicator of the

potential formation of the CTLH complex, however for these investigations, the larger,

90kDa isoform of RanBPM could not be isolated from three cell lines, although

multiple optimisation methods were employed. Thus, the smaller, 55kDa isoform of

RanBPM was cloned and used in immunoprecipitation and colocalisation assays with


275

RMND5 proteins. The 55kDa RanBPM isoform was the initial isoform reported, and in

the literature most studies that use the RanBPM (90kDa) isoform have obtained the

expression construct encoding the isoform from the group that originally described its

isolation (Nishitani et al., 2001). It is therefore possible that other groups have

experienced similar difficulty in amplifying the full length transcript, and indeed,

Nishitani et al. (2001) derived the original construct by amplifying the 5’ region of the

RanBPM (90kDa) isoform and ligating it with the RanBPM (55kDa) isoform,

supporting this hypothesis. PCR difficulties may be due to a stretch of repeating

nucleotides at the 5’ end of the transcript which encodes a string of proline and

glutamine residues in the amino-terminal region of the RanBPM (90kDa) protein

(Nishitani et al., 2001). The RanBPM (55kDa) isoform does not lack identifiable

protein domains and in this study the exogenous Cherry-tagged protein was found to

exhibit the same cellular distribution as that reported for the full length protein

(Kobayashi et al., 2007). Furthermore, RanBPM was able to interact and colocalise with

both RMND5 proteins in prostate cancer cells, which is consistent with previously

reported findings that RMND5A and RanBPM interact and colocalise in HEK293 cells

(Kobayashi et al., 2007). Interestingly, when coexpressed with either RMND5A or

RMND5B, the intracellular localisation of RanBPM was altered and it colocalised with

RMND5 proteins in punctate cytoplasmic speckles. Another CTLH complex

component, muskelin has been reported to exhibit a similar cytoplasmic punctate

appearance that is not associated with aggresomes and is dependent on its discoidin-like

domain and Kelch repeat domain (Prag et al., 2004). Although the localisation of the

three complex members in similar structures is interesting and supports formation of the

CTLH complex, the functional relevance of these cytoplasmic speckles is unknown but

may be investigated in future studies using antibodies or stains for intracellular

organelles such as endosomes or mitochondria.

The function of the CTLH complex has not yet been identified, and individually the

complex members are involved in diverse cellular processes, however there are several

lines of evidence supporting a role for the CTLH complex in protein degradation.

Firstly, as mentioned previously, the yeast Vid30 complex functions as an E3 ubiquitin

ligase complex, and this study has shown that both RMND5 proteins possess E3

ubiquitin ligase activity, which mirrors the function of their yeast orthologue RMD5

that provides the Vid30 complex with its enzymatic activity (Santt et al., 2008).

ARMC8α is reported to be associated with the proteasomal degradation of α-catenin and


276

enhances the interaction of the endosomal sorting protein HRS with ubiquitinated

proteins, potentially bringing ubiquitinated proteins in close proximity to HRS (Suzuki

et al., 2008; Tomaru et al., 2010), while RanBPM is also associated with the

deubiquitinating enzyme USP11 (Ideguchi et al., 2002). The protein domain

architecture of members of the CTLH complex also supports an E3 ubiquitin ligase

function, with RanBPM and muskelin containing SPRY and Kelch repeat domains,

respectively, both of which function as substrate recognition motifs in proteins forming

part of other E3 ubiquitin ligase complexes (Sun et al., 2009; Kuang et al., 2010; Lee et

al., 2010). Finally, the armadillo repeat domain is also present in proteins that are

associated with protein degradation, either those that possess intrinsic E3 ubiquitin

ligase activity or those that form part of E3 ubiquitin ligase complexes such as F-box

proteins (Tewari et al., 2010). In future studies, E3 ubiquitin ligase activity of the

CTLH complex could be investigated using in vitro ubiquitination assays by

reconstituting the complex either by coimmunoprecipitation of the entire complex or by

production of each of the components in bacterial cells or by other methods including in

vitro transcription/translation (as discussed in Section 4.3) (Skowyra et al., 1999; Chen

et al., 2006b; Ahn et al., 2011). In yeast, RMD5 and Gid9 function together to provide

the Vid30 complex with its E3 ubiquitin ligase activity, and collaboration of the human

orthologues of these factors, RMND5A and EMP may also be specifically investigated

in in vitro ubiquitination assays used to evaluate CTLH complex function (Braun et al.,

2011).

Another common feature of CTLH complex members is their role in cytoskeletal

organisation and cytoskeleton-based intracellular transport. The cytoskeleton consists of

microtubules, intermediate filaments and microfilaments (actin filaments) (Alberts et

al., 2004). Molecular motor proteins use these cytoskeletal elements for intracellular

transport, for example myosin molecules move cargo along actin filament “tracks”

whilst dyneins are microtubule associated proteins that among other functions carry

endosomal vesicles to the lysosome for degradation (Alberts et al., 2004; Nelson et al.,

2005). Actin is also essential for endocytosis and in particular, F-actin is important for

the transport of proteins such as caveolin from the plasma membrane to early

endosomes (Samaj et al., 2004). Members of the yeast Vid30 complex are not only

involved in the proteasomal degradation of FBPase, but under specific conditions,

FBPase is trafficked by members of the Vid30 complex to the endosomal system and

the yeast version of the lysosome, the vacuole for degradation (Brown et al., 2010;


277

Alibhoy et al., 2012). Specifically, Vid30 associates with Vid vesicles that contain

FBPase and with actin patches, thereby merging the Vid vesicles with the endocytic

pathway (Brown et al., 2010; Alibhoy et al., 2012). Muskelin, EMP, ARMC8α and

RanBPM are all involved in the intracellular transport of cargo or cytoskeletal

organisation, with muskelin mediating the intracellular cytoskeletal response to

thrombospondin and involved in both membrane bound receptor endocytosis and

endocytic transport (Adams et al., 1998; Ledee et al., 2005; Heisler et al., 2011). In a

similar role, muskelin connects F-actin and microtubule based transport of the

GABAAR α1 receptor by interacting with both myosin VI and dynein, thereby bridging

the two cytoskeletal based transport systems (Heisler et al., 2011), and together,

muskelin and RanBPM play a role in nucleocytoplasmic transport (Valiyaveettil et al.,

2008). EMP is associated with F-actin and due to this association, is vital for

enucleation and filopodia formation in erythroblasts and macrophages, respectively

(Soni et al., 2006). The LisH and Kelch repeat domains present in CTLH complex

members are also associated with intracellular transport of cargo, with the LisH domain

functioning as a dynein binding domain and the Kelch repeat an actin binding motif

(Adams et al., 2000; Emes and Ponting, 2001). In the initial screen for RMND5 binding

partners by mass spectrometry, multiple cytoskeletal proteins including actins, tubulins,

myosins, microtubule associated proteins, merlin, flotillin and lamins were identified,

suggesting that RMND5 proteins are associated with cytoskeletal organisation or

transport. However, as cytoskeletal proteins have been documented to non-specifically

bind the sepharose and magnetic bead matrix used, it will be particularly important to

confirm the interaction of individual cytoskeletal proteins with RMND5A or RMND5B

using additional assays (Trinkle-Mulcahy et al., 2008).

The CTLH complex components, RanBPM and RMND5A each have cellular

homologues, suggesting that RanBP10 and RMND5B may replace their paralogue in

the complex, thereby altering its function(s) or targets. Each of the homologous pairs of

proteins exhibits a high degree of amino acid identity, however RanBPM and RanBP10

have been documented to possess different functions while RMND5A and RMND5B

have not been characterised. For example, although both RanBPM and RanBP10 can

interact with the intracellular domain of the human growth factor receptor MET, only

the interaction of MET with RanBPM results in the activation of the Ras/ERK

signalling pathway (Wang et al., 2002a; Wang et al., 2004). RanBP10 has also been

reported as a tubulin binding protein that localises to microtubules in megakaryocytes


278

thereby playing a role in platelet shape and degranulation (Schulze et al., 2008; Kunert

et al., 2009). This function is consistent with that of other CTLH complex members

which are involved in microtubule dynamics and therefore cytoskeletal organisation.

Formation of the CTLH complex with either or both paralogues may extend the

function or substrates of the complex due to the proposed divergence in function of the

homologues and may be investigated in future studies.

Due to the presence of the LisH and RING dimerisation motifs in RMND5 proteins,

their association with each other was investigated. Many RING domain proteins use this

domain to form homodimers or heterodimers, which can enhance their enzymatic

activity (Section 4.2.1, 4.3), with the LisH domain additionally associated with the

formation of large protein complexes (Ahn et al., 2011). Coimmunoprecipitation and

colocalisation assays demonstrated that RMND5 proteins were able to interact and

colocalise in the nucleus and cytoplasm of prostate cancer cells. The finding suggests

that RMND5 proteins could function as heterodimers, however as RMND5 proteins

share a high degree of amino acid homology it may be that they are able to interact with

each other due to their similarity but that this interaction is not functionally relevant and

that they preferentially form homodimers in vivo. Whether RMND5 proteins

simultaneously form part of the CTLH complex and whether they function as single

subunit or multi-subunit E3 ubiquitin ligases or both remains to be addressed. Several

E3 ubiquitin ligases are reported to function in both modalities including Siah1 which

forms part of an SCF E3 ubiquitin ligase complex that ubiquitinates β-catenin or

alternatively functions as a single subunit that ubiquitinates a number of targets

including synphilin-1 (Liani et al., 2004; Dimitrova et al., 2010). Furthermore, it will be

interesting to determine whether RMND5A and EMP are able to function as E3

ubiquitin ligase heterodimers given the finding that yeast RMD5 and Gid9 function

together in the Vid30 complex (Santt et al., 2008; Braun et al., 2011). Typical E3

ubiquitin ligase complexes contain a single RING or HECT domain containing protein

with associated interchangeable complex members including adaptors and substrate

recognition subunits. For example, SCF and VHL E3 ubiquitin ligase complexes usually

contain the RING domain protein Rbx1, whilst the cullin and adaptor proteins may

change (Willems et al., 2004). Since it is the RING domain that interacts with E2

enzymes, substitution of the RING domain containing protein within an E3 complex

may alter the type of ubiquitination of the substrate proteins.


279

The cellular localisation of RMND5 proteins is interesting considering the identification

of multiple mitochondrial proteins as potential RMND5A and RMND5B binding

partners, including the α and β subunits of the mitochondrial ATP synthase involved in

oxidative phosphorylation, the TCA cycle enzyme isocitrate dehydrogenase and the

gluconeogenic enzyme phosphoenolpyruvate carboxykinase (PEPCK). Although all

protein-protein interactions identified using mass spectrometry need to be confirmed by

further studies, the Vid30 complex ubiquitinates and targets for degradation the

gluconeogenic enzyme FBPase (Santt et al., 2008), and therefore it is feasible that

RMND5 proteins and potentially the CTLH complex, regulate levels of mitochondrial

enzymes by ubiquitination.

The mass spectrometric techniques employed in this study to identify putative RMND5

binding partners were based on the premise that by immunoprecipitating the protein of

interest, the endogenous binding partners of that protein may be coimmunoprecipitated

and therefore identified. Due to technical limitations of this method, it will important in

future studies to confirm protein-protein interactions of individual molecules using

additional methods such as co-immunoprecipitation, GST-pulldown and additionally

colocalisation (Guenther et al., 2000; Liang et al., 2008; Paul et al., 2011). In this study,

different controls were used including a mock reaction where untransfected cells were

immunoprecipitated with the same anti-GFP microbeads used to immunoprecipitate

GFP-RMND5 proteins. In this way, proteins that bound to the microbead matrix could

be identified. A preclearing step prior to immunoprecipitation with the micro beads

alone (unbound with GFP antibody) to remove proteins non-specifically bound to the

beads, was not included in these experiments and is not recommended by the GFP

microbead manufacturers for immunoprecipitation reactions (Vostal and Shulman,

1993; Falsone et al., 2008). Nonetheless this could be added in future experiments. In

the immunoprecipitation using GFP-RMND5A, the bands present in the mock control

were used to guide excision of bands for mass spectrometric analysis, with two

contaminants databases used to eliminate commonly contaminating proteins, including

external contaminants and proteins used in mass spectrometry quantitation (Keller et al.,

2008; Bell et al., 2009). A publication identifying the sepharose and magnetic bead

proteome was also used to eliminate proteins binding to the microbead matrix and GFP

tag (Trinkle-Mulcahy et al., 2008).


280

Mass spectrometric analysis of both immunoprecipitation assays identified many

common contaminating proteins including keratins and non-specific bead proteome

binding proteins, for example heat shock proteins (Trinkle-Mulcahy et al., 2008). These

proteins were also present in the mock immunoprecipitation control used for RMND5B

immunoprecipitation and thus were eliminated as external contaminants or non-specific

binding proteins. Although precautions were taken when preparing all reagents for

immunoprecipitation and mass spectrometry with ultrapure water, precast acrylamide

gels, nitrile gloves, a face mask and hair clips utilised, keratins were the predominant

proteins identified (Mann et al., 2001). As keratins are external contaminants they are

present on the outside of the gel, on the glass plates and may be added upon gel

handling and as such are exposed to trypsin during protein digestion and ionise

efficiently, resulting in their identification by mass spectrometry (Pandey et al., 2000;

Keller et al., 2008). In the future, mass spectrometry grade reagents and acrylamide gels

and tubes may be utilised to reduce the presence of external contaminants. The GFP tag

has been determined to bind multiple cellular proteins, and although a published

database was used to exclude GFP binding proteins, an additional control which may be

used in future experiments is the expression of the GFP tag alone (empty vector) in

mammalian cells followed by its immunoprecipitation with anti-GFP microbeads

(Trinkle-Mulcahy et al., 2008; Paul et al., 2011).

As a result of using contaminants databases, many proteins were eliminated from the

original mass spectrometry screen, some of which may have been real RMND5 binding

partners. Alternative approaches that may be employed to diminish interference by

non-specific binding proteins involve the use of quantitative-mass spectrometry and

isotope labelling such as stable isotope labelling with amino acids in culture (SILAC),

which includes a negative control to account for proteins non-specifically binding the

affinity matrix or protein tag (Ong et al., 2002). In these studies, the negative control

culture is labelled with light 12C medium and the experimental sample is labelled with

heavy 13C medium (Trinkle-Mulcahy et al., 2008). Following cell lysis the samples are

mixed, resulting in both negative control and experimental samples being

immunoprecipitated simultaneously and analysed by mass spectrometry in the same run,

reducing variability between experiments (Trinkle-Mulcahy et al., 2008). This mass

spectrometry based technique is now widely used to identify novel protein-protein

interactions and its use has been expanded to study post-translational modifications and


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to identify cell proteomes under various conditions (Dammer et al., 2012; Deeb et al.,

2012; Udeshi et al., 2012; Yin et al., 2012).

Proteins with significant unique peptide matches of 2 or more were considered a match

for the presence of that protein in this study and therefore among the list of putative

RMND5 binding partners there were a range of proteins with significant hits from 2 to

18 (Higdon and Kolker, 2007). These results will be further analysed in future studies as

the top peptide matches do not necessarily reflect the strength of the protein-protein

interaction and any of the proteins may be genuine binding partners. For the methods

used, differences in binding affinity, the proportion of each protein that interacts with

the protein of interest in the cell at any given time, the abundance of the protein, the

efficiency of trypsin digestion and peptide ionisation may all account for differences in

the number of significant peptide matches identified (Nesvizhskii and Aebersold, 2005;

Higdon and Kolker, 2007; Trinkle-Mulcahy et al., 2008).

Another important element to consider in protein identification by mass spectrometry is

the peptide coverage of the identified proteins as a more complete protein coverage

lends more weight to the presence of that protein in the immunoprecipitate. Although

the entire protein may be present as trypsin digested peptides in the sample, usually only

between 20-30% are identified by mass spectrometry and there are multiple proposed

reasons for this. Some trypsin digested peptides do not ionise well, or due to unknown

mechanisms, the intensity of the peptide may be suppressed in a mixture of peptides,

long stretches of arginine and lysine lacking peptides may fall outside of the measured

m/z interval or the protein may be present at a low abundance (Nesvizhskii and

Aebersold, 2005; Hjerno and Hojrup, 2006). Therefore, whilst the presence of only a

portion of the protein represented by peptide matches does not rule out the presence of

that protein due to the above-mentioned reasons, it is also possible for example that a

degradation product of the protein has non-specifically bound to the protein of interest

or the affinity matrix. When RMND5A was immunoprecipitated, 33% protein coverage

was obtained, and the peptide matches were found throughout the RMND5A protein

sequence, with 9 unique peptide matches identified when protein band 1 was analysed

alone and 4 unique significant peptide matches when all protein bands were combined

and analysed together. In the literature, protein sequence coverage of 14% representing

eight unique peptides has been used as a positive protein identification (Block et al.,


282

2011). For RMND5B, 2 unique peptides obtained a protein coverage of 4.6% which is

low, however 2 unique peptides is considered a positive protein identification.

The top ranked putative RMND5A binding partner identified was PUF60, which had 18

significant peptide matches accounting for 46% of the PUF60 protein sequence. PUF60

was identified when the band containing only RMND5A was analysed and also when

all 8 protein bands of interest were combined and analysed, and importantly, different

peptides were identified when only band 1 was analysed compared to when all of the

protein bands were combined and analysed. As PUF60 (~60kDa) corresponded in size

to the region excised from the gel (~60-70kDa), the overall findings provided good

support of the further investigation of PUF60 as an RMND5A binding partner. While

other splicing factors were identified and eliminated in this screen due to their reported

ability to bind the affinity matrix (Trinkle-Mulcahy et al., 2008), PUF60 was not

eliminated as it did not appear in the published databases and the number of peptide

matches to PUF60 were high compared to other splicing factors identified (Trinkle-

Mulcahy et al., 2008). Subsequent mass spectrometry analysis of GFP-RMND5B

binding partners including a mock immunoprecipitated control confirmed PUF60 as

binding to the magnetic bead matrix. This experiment therefore highlights the

importance of analysis of the mock immunoprecipitation control by mass spectrometry,

as visual analysis of bands present in the immunoprecipitated sample that are not

present in the mock control is not sufficient.

Proteins identified as putative RMND5A and RMND5B binding partners, with at least 2

unique peptides andbetween 12.3% - 33% protein sequence coverage included

mitochondrial, cytoplasmic protein and nuclear proteins, a splicing factor, a

transcription factor and a protein involved in DNA repair. The transcription factor

SNW1, identified as a putative RMND5A binding partner has been demonstrated to

interact with the 3’ splicing factor U2AF65, in particular in relation to p21 pre-mRNA

splicing, and is therefore hypothesised to couple transcription and RNA splicing (Zhang

et al., 2003; Chen et al., 2011b). PUF60, which was also identified in this screen, is

homologous to and interacts with U2AF65, and PUF60 can substitute for U2AF65 in

vitro (Hastings et al., 2007; Corsini et al., 2009). Therefore, whether SNW1 specifically

interacts with RMND5A or whether SNW1 was coimmunoprecipitated with PUF60,

remains to be determined. However, SNW1 was not identified in the mock

immunoprecipitated control or GFP-RMND5B immunoprecipitated sample mass


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spectrometry results, therefore an interaction between RMND5A and SNW1 will be

ascertained by future coimmunoprecipitation reactions.

The peptide hits and sequence coverage identified in the GFP-RMND5B mass

spectrometry screen were low compared to that of the peptide matches obtained when

GFP-RMND5A and its binding partners were analysed by mass spectrometry, which

resulted in the identification of RMND5B and ubiquitin-40s ribosomal protein S27A as

the only two proteins represented by two unique peptides (individual ion score >30).

This may have been due to division of the GFP-RMND5B immunoprecipitation sample

between two lanes for electrophoresis, with only one of the lanes analysed (thereby

diluting the amount of sample used). Additionally, in the GFP-RMND5A mass

spectrometry analysis, individual bands were excised in comparison to the GFP-

RMND5B mass spectrometry screen where large sections of the gel were excised for

analysis. Thus, to concentrate the proteins present, the removal of smaller gel bands

from the gel which may be analysed individually may yield results with higher

scores/peptide matches to each protein. In addition, identification of low abundance

proteins present in each band may be possible, which are not identified when large

sections of gels are combined and analysed.

Although several of the proteins identified in the GFP-RMND5A and GFP-RMND5B

mass spectrometry screens had only one unique peptide (individual ion score >30)

where two or more are generally considered to be a positive identification for a protein,

some of these proteins were identified in both GFP-RMND5A and GFP-RMND5B

mass spectrometry screens. These proteins were ubiquitin-40s ribosomal protein S27A,

mitochondrial ATP synthase subunit β, POTE ankyrin domain family member E,

tubulin β-4A and unconventional myosin-VI. As the proteins were not present in the

mock immunoprecipitation control or in the contaminants databases, the proteins are

potential RMND5 protein binding partners, which may be investigated in future studies

using additional analyses including coimmunoprecipitation assays and colocalisation

microscopy (Ciruela et al., 2010). RMND5A contains a putative GAT-like domain, a

ubiquitin-binding domain (Section 4.2.1, 4.3), therefore if RMND5A/RMND5B is

confirmed to interact with ubiquitin-40S ribosomal protein S27A it will be interesting to

identify the protein domain responsible for this interaction. Additionally, as ubiquitin-

40S ribosomal protein S27A is a fusion protein of ubiquitin and ribosomal protein S27A

prior to its processing, it will be important to determine results of the outcome of this


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interaction (Wong et al., 1993). Thus, at the close of this project, RMND5A and

RMND5B have been demonstrated to bind NKX3.1 and RanBPM, with mass

spectrometric analysis of RMND5A and RMND5B interacting proteins identifying

several candidate binding partners. These may be further investigated in conjunction

with additional mass spectrometric screens to delineate other RMND5 protein

interactors including CTLH complex members.

Chapter 7 General Discussion

Chapter 7: General Discussion


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7.1 General Discussion – The Role of E3 Ubiquitin Ligases in

Transcriptional Regulation Gene expression is a tightly controlled process and as such transcription, the initial step

in this process, is regulated by a number of mechanisms, in particular post-translational

modifications including acetylation, phosphorylation and ubiquitination. Ubiquitin and

the proteasome participate in the regulation of transcription by modifying the function

of histones, RNA Polymerase II (RNA Pol II) and transcription factors (Davie and

Murphy, 1990; Li et al., 1993; Fuchs et al., 1997; Mitsui and Sharp, 1999; Starita et al.,

2005). Histone ubiquitination alters chromatin structure, affecting the accessibility of

the transcriptional machinery to sites of transcription and also acts as a signal to recruit

regulators of transcription (Fierz et al., 2011; Hammond-Martel et al., 2012). By

regulating RNA Pol II and the transcription factor machinery directly, ubiquitination

affects their protein levels and activity via proteolytic and non-proteolytic mechanisms

(Mitsui and Sharp, 1999; Akiyama et al., 2005; Yan et al., 2009; Dao et al., 2012).

Together with other post-translational modifications, this multi-levelled approach to

transcriptional regulation exerts fine control over gene expression, ensuring that genes

will only be expressed in response to the correct cellular signals.

Since transcription factors, which possess activator and/or repressor functions, play a

significant role in determining which genes are expressed at any given time, they form

an important point of transcriptional regulation. Perhaps the best known and most

comprehensively characterised outcome of transcription factor ubiquitination by E3

ubiquitin ligases is the proteasome dependent degradation of the transcription factor. In

the present study, the transcription factor NKX3.1 was determined to be ubiquitinated

by the E3 ubiquitin ligases RMND5A and RMND5B, thus targeting NKX3.1 for

degradation by the proteasome. Along with the ability of polyubiquitination to promote

the degradation of transcription factors, there is an established role of ubiquitination in

modulating the activity of the transcription factor prior to its degradation. The potency

of transcriptional activation domains (TADs), present in activators and coactivators, has

been shown to inversely correlate with their half-life, particularly in the case of acidic

TADs (Molinari et al., 1999). Molinari and colleagues demonstrated that TADs from

various transcription factors fused to the GAL4 DNA binding domain were able to

induce transcription and that their half-lives were dependent on the strength with which

these chimeric transcription factors activated transcription (Molinari et al., 1999). This


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rapid protein turnover was also dependent on the presence of the GAL4 DNA-binding

domain, suggesting that the transcription factor must be bound to DNA and activated to

trigger ubiquitination (Molinari et al., 1999). A further example of this mechanism was

demonstrated in yeast where yeast transcription factors containing the VP16 TAD are

ubiquitinated by the SCFMET30 E3 ubiquitin ligase complex, however in cells lacking

Met30, the LED-VP16 transcription factor does not undergo ubiquitination and

subsequent degradation and loses its transactivation ability (Salghetti et al., 2001).

These findings suggest that in addition to its role in transactivation, the TAD acts as a

destabilisation domain and that ubiquitination of this domain modulates LED-VP16

degradation and transactivation activity. Therefore, as the activation and destruction of

the transcription factor are linked though the TAD, when the transcription factor has

fulfilled its transcriptional activity and is no longer needed, its degradation ensures that

it can no longer affect transcription. NKX3.1, which has a short half-life of ~25 minutes

contains an acidic domain and is able to activate the transcription of genes such as

PCAN1, although it is better known as a transcriptional repressor (Olsson et al., 2001,

Liu et al., 2000, Zhang et al., 2008a, Chen et al., 2002, Thomas et al., 2006). Further

investigation of the role NKX3.1 ubiquitination in the regulation of both its degradation

and its transcriptional activity, will identify the involvement of RMND5 proteins in this

process.

Transcription factor degradation can also be regulated by multiple E3s in response to

different cellular pathways, thereby allowing fine-tuned regulation of transcription

factor expression. Similarly, NKX3.1 protein levels are likely to be regulated by

multiple E3 ubiquitin ligases including TOPORS, which has been established to

ubiquitinate NKX3.1 resulting in its proteasome dependent degradation (Guan et al.,

2008). The present study has demonstrated that two additional E3 ubiquitin ligases,

RMND5A and RMND5B also regulate NKX3.1 protein levels by ubiquitination,

although their contribution in the normal maintenance of NKX3.1 levels is yet to be

determined. As NKX3.1 is involved in multiple cellular functions including

transcription and the DNA damage response, it is feasible that each E3 ubiquitin ligase

ubiquitinates NKX3.1 in response to different environmental conditions and in relation

to its different biological activities (Gelmann et al., 2003; Guan et al., 2008; Erbaykent-

Tepedelen et al., 2011). While individual transcription factors may be regulated by

multiple E3 ubiquitin ligases, single E3 ubiquitin ligases may target multiple

transcription factors, playing a central role in transcriptional regulation. For example,


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TOPORS is able to ubiquitinate both NKX3.1 and p53, and therefore deregulation of

TOPORS expression in prostate tumours may affect a number of transcriptional

pathways (Rajendra et al., 2004; Guan et al., 2008). In addition to the findings in this

thesis that RMND5 proteins ubiquitinate NKX3.1 and target it for proteasomal

degradation, overexpression of either RMND5A or RMND5B were also found to result

in a dose-dependent reduction in AR levels (not shown). The transcription factor SNW1

was additionally identified as a candidate RMND5A binding partner, and when

overexpressed with NKX3.1, RMND5 proteins were predominantly located in the

nucleus of prostate cancer cells, implicating both RMND5 proteins in the regulation of

several transcription factors.

The regulation of transcription factors by E3 ubiquitin ligases is not solely associated

with protein degradation, and both monoubiquitination and polyubiquitination of

transcription factors has been linked to alterations in other aspects of their cellular

activity or function. While proteolytic degradation is the ultimate means of halting the

activity of a transcription factor, the cellular localisation of transcription factors is also

an important factor in the regulation of their activity. Ubiquitination of a number of

transcription factors has been reported to alter their intracellular localisation and in the

present study, overexpression of RMND5 proteins resulted in reduced levels of nuclear

NKX3.1 and its predominant cytoplasmic localisation. A similar mechanism occurs

following ubiquitination of p53 by MDM2, with high levels of MDM2 resulting in p53

polyubiquitination and degradation, however when MDM2 levels are low, MDM2

monoubiquitinates p53 resulting in its nuclear export where it cannot directly regulate

transcription (Section 5.3) (Boyd et al., 2000; Geyer et al., 2000; Li et al., 2003). In the

cytoplasm, p53 plays roles in apoptosis and autophagy, indicating that nuclear export

does not always abrogate the activity of the transcription factor protein, and it is

possible that NKX3.1 similarly performs alternative functions when relocated to the

cytoplasm following overexpression of RMND5 proteins (Mihara et al., 2003; Tasdemir

et al., 2008). Conversely, the proteolytic cleavage of transcription factors triggered by

ubiquitination can promote their nuclear localisation to fulfil their transcriptional

regulatory role and well known examples of this include the activation of the nuclear

factor kappa B1 (NF-κB1) subunits p52 and p50 (Lin et al., 1998, Orian et al., 1999,

Orian et al., 2000). This occurs following cleavage of their precursors p105 and p100

via ubiquitination by the SCFβ-TRCP E3 ubiquitin ligase complex, leading to the


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degradation of all but the amino-terminal transcriptionally active p52 and p50 domains

(Lin et al., 1998; Orian et al., 1999; Orian et al., 2000).

Monoubiquitination of transcription factors is another non-proteolytic mechanism of

transcription factor modulation, but although well-established, the mechanism by which

monoubiquitination regulates transcription factor function remains incompletely

understood. An unusual example of this mechanism is the ubiquitination of SRC3, a

transcriptional coactivator. The phosphorylation-dependent monoubiquitination of

SRC3 activates its transcriptional regulatory activity, however, this initial ubiquitination

is extended and progressively results in the formation of a polyubiquitin chain, which

triggers the proteasomal degradation of SRC3, thereby allowing a specific interval

during which SRC3 can function as transcriptional coactivator (Wu et al., 2007a). In

addition to its effects on transcription factor activity, non-proteolytic ubiquitination can

also affect cofactor binding. The activity of the Met4 transcription factor is regulated in

this manner, and ubiquitination of Met4 by the SCFMET30 E3 ubiquitin ligase does not

alter its promoter binding capacity but interferes with its ability to interact with the Cbf1

transcription factor, thereby resulting in the failure to form an active Met4

transcriptional complex (Kaiser et al., 2000).

In addition to their ability to regulate transcription factors by ubiquitination, it is evident

that a subset of E3 ubiquitin ligases are able to function directly in transcription

themselves by forming part of the transcriptional machinery. One of the well-

characterised E3 ubiquitin ligases with this dual function in transcription is BRCA1,

with the BRCA1/BARD1 E3 ubiquitin ligase heterodimer proteins forming part of the

RNA Pol II holoenzyme (Chiba and Parvin, 2002). This complex consists of RNA Pol

II and general transcription factor members as well as suppressor of RNA Pol B (SRB),

which are all required for the initiation of transcription (Koleske and Young, 1995;

Scully et al., 1997; Chiba and Parvin, 2002). BRCA1 associates with the RNA Pol II

holoenzyme complex by interacting with RNA Helicase, and with its amino-terminal

which contains the RING domain, BRCA1 is hypothesised to associate with the

complex via BARD1 (Anderson et al., 1998; Chiba and Parvin, 2002). In response to

DNA damage, the BRCA1/BARD1 heterodimer ubiquitinates RNA Pol II, targeting it

for proteasomal degradation (Kleiman et al., 2005; Starita et al., 2005). However, in

unstressed cells BRCA1 functions as a co-activator of a subset of genes including p53

by interacting with enhancers, thereby linking the enhancers to the RNA Pol II


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holoenzyme to promote transcription (Ouchi et al., 2000; MacLachlan et al., 2002).

Other examples of E3 ubiquitin ligases that participate in transcription are the yeast E3

ubiquitin ligase Rsp5 and its human orthologue hPRF1 as well as the HECT E3

ubiquitin ligase E6-AP which both function as coactivators for steroid hormone

receptors (Imhof and McDonnell, 1996; Nawaz et al., 1999). In a previous study in the

laboratory, RMND5B was identified to possess transcriptional repressor activity,

implying that RMND5B and perhaps RMND5A play additional as yet uncharacterised

roles in transcriptional regulation (Dawson, 2006). Although RMND5 proteins do not

contain defined DNA binding domains, the presence of multiple protein-protein

interaction domains implies that they are able to interact with a number of proteins

including transcription factors, or form part of transcription factor complexes, thereby

exerting transcriptional activator or repressor activities in a similar manner to other

LisH domain containing proteins (Section 4.1.3.1) (Li et al., 2000; Zhang et al., 2002;

Yoon et al., 2003, Choi et al., 2008). This role for RMND5 proteins is further supported

by the presence of putative steroid hormone receptor binding domains, nuclear receptor

boxes (NRBs) present in both proteins and the finding in this study that both proteins

accumulate in the nucleus following overexpression with the transcription factor,

NKX3.1. Additionally, the DNA damage repair factor, XRR5C and the transcription

factor SNW1 were identified in this study as potential RMND5A binding partners.

The ability of BRCA1 and BARD1 to participate in a variety of cellular processes

including DNA repair, cell cycle control and apoptosis is due to their multi-domain

architecture (Jin et al., 1997; Westermark et al., 2003; Fabbro et al., 2004; Schuchner et

al., 2005). Multi-functional E3 ubiquitin ligases possess a similar protein domain

architecture, with the RING domain located at the far amino- or carboxy-terminal and

additional protein domains located in the remaining length of the protein, suggesting

that in many cases the RING domain activity is separate from that of the residual

protein domains. However, the remaining protein domains may also function as

substrate recognition elements in single subunit E3 ubiquitin ligases or as association

domains for those E3 ubiquitin ligases forming part of multi-subunit E3 ubiquitin ligase

complexes. Although BARD1 interacts with BRCA1 through its RING domain, thereby

enhancing BRCA1 E3 ubiquitin ligase activity, BARD1 contains carboxy-terminal

ankyrin repeats and two BRCT domains which mediate BRCA1 independent cellular

roles in cell cycle regulation, p53 stabilisation and apoptosis (Wu et al., 1996; Xia et al.,

2003). By interacting with the DNA-PK subunit Ku-70 and p53, BARD1 induces p53


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phosphorylation and stabilisation, and in the cytoplasm, BARD1 associates with the

mitochondria, triggering apoptosis by stimulating Bax oligomerisation (Feki et al.,

2005; Tembe and Henderson, 2007). In comparison, the proteins investigated in the

present study, RMND5A and RMND5B possess an analogous protein domain

architecture, a carboxy-terminal RING domain and three additional protein-protein

interaction domains, the LisH, CTLH and CRA domains located amino-terminally to

the RING domain, suggesting that they too are multi-functional proteins. This is further

supported by the findings in this study that both proteins exhibit diffuse nuclear and

cytoplasmic localisation, with accumulation in cytoplasmic vesicles in a proportion of

cells that potentially signifies multiple functions of RMND5 proteins in distinct cellular

compartments. This hypothesis is supported by the identification of mitochondrial

proteins as putative RMND5A and RMND5B binding partners.

The ubiquitination of transcription factors, which has the ability to regulate transcription

factor activity and function, can be carried out either by single subunit or multi-subunit

E3 ubiquitin ligases. Single subunit E3 ubiquitin ligases typically contain a

HECT/RING/U-box domain and substrate recognition domains and tend to be multi-

functional proteins. One such example is the U-box E3 ubiquitin ligase CHIP which

functions as a co-chaperone and as an E3 ubiquitin ligase, interacting with the

chaperones Hsc70-Hsp70 and Hsp90 through its amino terminal tetricopeptide repeat,

whilst retaining its ubiquitin ligase activity mediated by its carboxy-terminal U-box

domain (Ballinger et al., 1999; Jiang et al., 2001). By interacting with molecular

chaperones, CHIP tips the balance of the protein folding machinery towards protein

ubiquitination and degradation, thereby connecting these two integral systems

responsible for the regulation of protein integrity. In its role as a transcriptional

regulator, CHIP ubiquitinates p53 and c-Myc resulting in their proteasomal degradation

(Esser et al., 2005; Naito et al., 2010; Paul et al., 2012). An interesting aspect of single

subunit E3s is their ability to form homodimers or heterodimers, most often resulting in

enhancement of their enzymatic activity. This is perhaps best typified by the

MDM2/MDMX heterodimer which targets p53 for proteasomal degradation, although

individually neither MDM2 nor MDMX are able to polyubiquitinate p53 in vitro,

demonstrating the importance of both proteins in the activity of the heterodimer (Wang

et al., 2011). In the absence of heterodimerisation, MDM2 and MDMX perform

distinct cellular roles that are independent of each other. MDM2 is able to ubiquitinate a

number of proteins including the transcription factor p73 and is a coactivator of the


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ApoCIII promoter, while MDMX contributes to p53 induced apoptosis by interacting

with BCL-2 in the mitochondria, facilitating the interaction between p53 and BCL2 and

thereby inducing cytochrome C release and apoptosis (Mancini et al., 2009; Kubo et al.,

2010; Yang et al., 2012). Similarly, in this study, the structurally related RMND5

proteins were demonstrated to interact and colocalise, implicating the proteins in

homodimer or heterodimer formation, conceivably with respect to the coregulation of

specific cellular targets as is the case for MDM2 and MDMX. As each RMND5 protein

was identified to possess E3 ubiquitin ligase activity in vitro it is also possible that they

have individual cellular targets and due to their multi-domain architecture that they

mediate alternative cellular functions similar to that of other heterodimeric E3 ubiquitin

ligases such as BRCA1/BARD1.

Proteins possessing E3 ubiquitin ligase activity, particularly those containing RING

domains are also capable of forming part of large, multi-protein E3 ubiquitin ligase

complexes, thus broadening the list of potential substrates of the E3 by extending the

range of available substrate recognition domains. E3 ubiquitin ligase complexes

involved in the regulation of transcription factors tend to be of the SCF (Skp-Cullin-F-

box) family, which all contain the RBX1 RING domain protein, cullin scaffolding

protein and a Skp 1 linker protein which binds a number of substrate recognition

components, including F-box proteins (Willems et al., 2004, Sun et al., 2007). In this

manner, the invariant core E3 complex is able to ubiquitinate and thereby regulate a

range of target proteins depending on the F-box protein with which it is associated.

Upon formation of the SCF complex with the F-box protein Fbw7, the KLF5

transcription factor is rapidly degraded whilst replacement of Fbw7 for β-TrCP results

in the ubiquitination and degradation of the transcription factor β-catenin (Latres et al.,

1999; Zhao et al., 2010). As a proposed E3 ubiquitin ligase complex, the CTLH

complex investigated in the present study is comprised of proteins of similar protein

domain architecture. This complex includes RMND5A, an E3 ubiquitin ligase, the

ARMC8α putative adaptor protein and RanBPM and muskelin, potential substrate

recognition components that contain SPRY and Kelch repeat domains present in F-box

and SOCS box proteins which function as substrate recognition components in other E3

ubiquitin ligase complexes (Kobayashi et al., 2007; Kuang et al., 2010; Lee et al.,

2010). Although the function of the CTLH complex as a multiprotein E3 ubiquitin

ligase complex is supported by the characterised function of several of its components

and the reported activity of its yeast orthologue as an E3 ubiquitin ligase complex (Santt


292

et al., 2008), the transcriptional regulatory activity of the CTLH complex is more

speculative. However, RanBPM is a binding partner and transcriptional activator of

steroid hormone receptors and multiple transcription factors, while RMND5A contains

putative NRBs and regulates the transcription factor NKX3.1 (Rao et al., 2002;

Brunkhorst et al., 2005; Dawson, 2006; Poirier et al., 2006). Therefore, the

transcriptional regulatory function of RMND5 proteins and the CTLH complex may be

further investigated in future studies.

Due to their important roles as regulators of gene transcription, it is unsurprising that the

dysregulation of multiple E3 ubiquitin ligases is associated with a variety of cancers.

Furthermore, as many E3 enzymes, not limited to those with transcriptional regulatory

roles, are involved in multiple cellular processes including apoptosis, cell cycle

regulation and genome stability, their disruption not only affects their transcriptional

regulatory roles but their broader cellular functions, thereby associating the abnormal

functions of E3 ubiquitin ligase with all stages of malignancy. The number of E3

ubiquitin ligases implicated in disease is constantly increasing and includes well-

characterised proteins such as BRCA1, MDM2, SCFFbw7 and WWP1, which have been

classified as oncogenes (e.g. MDM2) or tumour suppressor genes (e.g. BRCA1),

depending upon the cancer-associated abnormality involved (Oliner et al., 1992;

Hashizume et al., 2001; Chen et al., 2007; O'Neil et al., 2007). This classification is not

invariant but depending upon the cellular role(s) of the substrate(s), the E3 enzyme may

function in either capacity. In many cases, due to their pervasive role in cellular

pathways, it is the E3 ubiquitin ligase that balances opposing oncogenic and tumour

suppressor functions of abnormal or aberrantly regulated genes, and due to their defined

substrate specificity, it is not surprising that multiple E3s are being investigated as

possible therapeutic targets (Chen et al., 2008; Edelmann et al., 2011; Buckley et al.,

2012; McCormack et al., 2012). As E3 ubiquitin ligases and regulators of the

transcription factor NKX3.1 as well as other unidentified cellular targets, RMND5

proteins are themselves dysregulated in a number of cancers. Particularly of interest is

the RMND5B chromosomal locus, 5q35.5 which is located in a region associated with

prostate cancer heritability (Xu et al., 2005; Christensen et al., 2010). Additionally, the

RMND5A and RMND5B gene loci are disrupted in a number of cancers including

ovarian carcinoma, mantle cell lymphoma, pilocytic astrocytoma, non-small lung cell

carcinoma, neuroblastoma and breast carcinoma implying that RMND5 proteins

perform important cellular functions in a number of different cell types (Section 4.1.4)


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(Mendes-da-Silva et al., 2000; Mosse et al., 2005; Camps et al., 2006; Johannsdottir et

al., 2006; Li et al., 2008; Belirgen et al., 2012). The identification of these cellular roles

and additional target proteins of RMND5 ubiquitination will aid in the understanding of

the consequences of RMND5 disruption in cancers and if appropriate, may add to the

many E3 ubiquitin ligases targeted for cancer therapeutics.

7.2 Future Directions The investigations in this thesis, that RMND5 proteins function as E3 ubiquitin ligases

and ubiquitinate the transcription factor, NKX3.1 leading to its proteasomal

degradation, have yielded results which may be expanded in future studies

characterising the cellular functions of RMND5 proteins, particularly with regard to

their ubiquitination targets, including NKX3.1.

The cellular functions of proteins are in part mediated by their intracellular localisation,

expression levels and binding partners, elucidation of which requires the availability of

specific antibodies against the protein of interest. Although for the short term,

exogenous tagged protein may be used in in vivo or in vitro functional assays, and

indeed in some case may be more suitable than use of the endogenous protein, it is

ultimately essential that the endogenous protein should be investigated. In this

laboratory, previous attempts to generate monoclonal or polyclonal antibodies against

RMND5B have not been successful (not shown), and while two commercial RMND5A

antibodies are currently available, neither is validated and where western blotting is

reported, the molecular size of the protein band does not correspond to that of

RMND5A. The RMND5A and RMND5B amino acid sequences are highly similar in

mouse, rat, rabbit and other species commonly used to generate antibodies, potentially

reducing the antigenicity of the proteins, however it will be important for future studies

to develop antibodies against RMND5A and RMND5B in order to investigate the

cellular localisation and protein levels of endogenous RMND5 proteins both in

mammalian cell lines and in normal and malignant tissues. These antibodies may also

be used to assess whether endogenous RMND5 proteins interact in mammalian cells

and to provide evidence supporting their heterodimer formation or their inclusion in the

CTLH complex.


294

In this study, transient overexpression of RMND5 proteins was used to determine their

effects on substrate proteins as the endogenous levels of either RMND5A or RMND5B

were not known (although the expression of both genes was verified using RT-PCR).

Following determination of endogenous RMND5A and RMND5B levels, future in vitro

studies to determine alternative functions or additional substrates of RMND5 proteins

can include the use of siRNA knockdown of either or both RMND5A and RMND5B.

Due to the high degree of similarity between RMND5A and RMND5B, it is likely that

they share cellular targets or that one may compensate for loss of the other, therefore the

function of both RMND5 proteins should be investigated together. Knockdown of E3

ubiquitin ligase expression by siRNA is commonly used to examine E3 function and

should result in increased expression of substrate proteins where ubiquitination by the

E3 ubiquitin ligase (e.g. RMND5A or RMND5B) results in proteasomal degradation

(Zhong et al., 2005; Nishitani et al., 2006). These experiments will therefore be critical

for confirmation of the identity of endogenous RMND5 substrate proteins such as

NKX3.1.

Investigation of the in vitro function of both RMND5A and RMND5B RING domains

confirmed that, like their yeast orthologue, both proteins possess E3 ubiquitin ligase

activity (Santt et al., 2008). Future studies expanding on these findings would confirm

the activity of full length RMND5 proteins in in vitro ubiquitination assays by

producing RMND5A/RMND5B using alternative protein tags in bacteria or in vitro

transcription/translation methods as discussed previously (Section 4.3). The presence of

all protein domains including substrate recognition domains following full length

protein production, would allow the inclusion of putative substrate proteins in in vitro

ubiquitination assays. In the presence of particular substrates or in auto-ubiquitination

assays, it will be useful if both RMND5 proteins are assayed simultaneously to assess

whether they are able to function as heterodimers and whether this enhances their

ubiquitination activity (Hashizume et al., 2001). The soluble full length RMND5

proteins may also be used to obtain crystal structures of both proteins which could be

used to assess whether RMND5 proteins form homodimers or heterodimers, and the

mechanism by which this activity may enhance their individual activity or allow them to

ubiquitinate distinct proteins. Crystal structures may also aid in the characterisation of

RMND5 protein interactions with other CTLH complex members and delineate

RMND5 protein interactions with different E2 conjugating enzymes, a finding of this

study.


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Both RMND5 proteins were determined to interact with a number of E2 conjugating

enzymes in in vitro ubiquitination assays performed for this thesis. As some E2

enzymes are chain initiating enzymes while others appear to function predominantly as

chain elongating E2 enzymes, the E2 enzymes that require prior monoubiquitination of

the substrate would not be identified using these methods (Windheim et al., 2008; Ye

and Rape, 2009; Williamson et al., 2011). Thus, it will be important to determine more

comprehensively which E2 conjugating enzymes are able to interact with RMND5

proteins in vivo, as the range of E2 enzymes may differ between the two proteins, in

vivo results may differ compared to in vitro findings, and interactions may depend on

specific culture (environmental) conditions. Identification of the E2 enzymes with

which RMND5 proteins interact to mediate ubiquitin transfer will be important as it is

the E2 enzyme that determines the ubiquitin chain topology attached to the substrate

and therefore the ultimate fate of the substrate, although the E3 ubiquitin ligase also

plays an important role in this process (Windheim et al., 2008; David et al., 2011).

Once the E2 conjugating enzymes have been identified, the outcome of substrate

ubiquitination by RMND5 proteins can be predicted. These predictions can then be

assessed biochemically using linkage specific antibodies or ubiquitin mutants to

determine the ubiquitin linkages of specific RMND5 substrate proteins, for example

NKX3.1 (Wu-Baer et al., 2003; Newton et al., 2008; Matsumoto et al., 2010). As

NKX3.1 is targeted for degradation following RMND5 overexpression, this implies that

together with the relevant E2 enzyme, RMND5 proteins are involved in the formation

of lysine 11, lysine 29 or lysine 48 polyubiquitin chains, therefore the type of ubiquitin

chains attached to NKX3.1 under these conditions can be verified experimentally (Chau

et al., 1989; Gregori et al., 1990; Johnson et al., 1995; Matsumoto et al., 2010).

Similarly, following the identification of additional substrates of RMND5

ubiquitination, the types of ubiquitin chains attached by the E2/E3 enzyme pair can be

determined as at this stage it is unknown whether RMND5 proteins always target their

substrate proteins for proteasomal degradation. Due to the inclusion of RMND5A and

possibly RMND5B in the human CTLH complex, and the association of members of

the complex with endosomal protein degradation (where monoubiquitination serves as

an internalisation trigger for membrane bound proteins and receptors), it will be

interesting to determine the outcomes of ubiquitination of substrates of RMND5

proteins and the CTLH complex. These include whether the outcomes of the substrate

proteins differ depending on the function of RMND5A or RMND5B as single subunit


296

E3 ubiquitin ligases or as part of the CTLH complex (Haglund et al., 2003a; Haglund et

al., 2003b). The studies will be assisted in part by use of the above mentioned linkage

specific antibodies (see also Sections 4.3, 5.3).

In the present study, the prostatic tumour suppressor, NKX3.1 was investigated as a

substrate of RMND5 mediated ubiquitination. In the future, the RMND5 mediated

degradation of NKX3.1 should be further examined including the specific

environmental or cellular signals under which RMND5 proteins mediate NKX3.1

ubiquitination, in particular as another E3 ubiquitin ligase, TOPORS is also involved in

NKX3.1 degradation by the proteasome (Guan et al., 2008). NKX3.1 expression is

androgen (AR) regulated and although this is in part mediated by androgen regulation of

NKX3.1 gene transcription (Prescott et al., 1998), it is apparent that NKX3.1 protein is

also androgen regulated as androgen withdrawal induces proteasome mediated

degradation of NKX3.1. The effects of androgen addition or depletion on NKX3.1

ubiquitination could be investigated in future studies, including the identification of the

E3 ubiquitin ligases involved. Since NKX3.1 is regulated by phosphorylation and

phosphorylation by CK2 is reported to protect NKX3.1 from ubiquitin-mediated

degradation, it is likely that there is an interplay between these two types of post-

translational modifications, which could be further elucidated by identification of the E3

ubiquitin ligase(s) whose activity is modified by CK2 phosphorylation of NKX3.1 (Li et

al., 2006; Markowski et al., 2008). Under conditions where NKX3.1 ubiquitination is

induced, it may be determined whether phosphorylation of particular NKX3.1 residues

is a signal for its ubiquitination as it is for other proteins (Punga et al., 2006; Scaglioni

et al., 2008). Furthermore, if RMND5 proteins either alone or as part of the CTLH

complex are able to mediate phospho-NKX3.1 ubiquitination, the means by which they

are able to recognise phosphorylated NKX3.1 will require investigation as neither

protein contains a canonical phospho-recognition motif. Of additional interest would be

the identification of specific lysine residues of NKX3.1 which are ubiquitinated by

RMND5 proteins and/or TOPORS, with ubiquitination of lysine residues within the

homeodomain hypothesised to target NKX3.1 for proteasomal degradation (Ju et al.,

2009).

NKX3.1 is a DNA binding transcription factor, and it is feasible that RMND5 proteins

ubiquitinate DNA-bound NKX3.1, implying that it is active NKX3.1 which is

ubiquitinated and hence targeted for degradation once it is “spent”, thereby allowing its


297

removal from the DNA and replacement with a fresh transcription factor. Another

aspect of the interaction between RMND5A/RMND5B and NKX3.1 that would

increase understanding of its biological consequences would be to determine the

domains of each protein required for this interaction. Using yeast two-hybrid assays, it

has been determined in this laboratory that the interaction between RMND5B and

NKX3.1 may be mediated in part by the CTLH and CRA domains of RMND5B,

although the result requires confirmation, and RMND5A-NKX3.1 interaction domains

have not been reported (Lau, 2008). The CTLH domain and part of the CRA domain are

also required for the interaction of RMND5A with ARMC8α (Kobayashi et al., 2007),

suggesting that if RMND5 proteins use these domains to form part of the CTLH

complex, they cannot simultaneously interact with NKX3.1. As such, RMND5 protein

interaction with NKX3.1 may not occur in conjunction with the CTLH complex but

may be mediated by RMND5 proteins functioning as single subunit E3 ubiquitin

ligases.

In this study, mass spectrometry was used to identify putative RMND5 binding partners

and the candidate binding partners isolated could be further investigated and confirmed

using additional functional studies such as co-immunoprecipitation assays, GST-

pulldown assays and colocalisation microscopy before additional experiments can be

performed assessing the functional outcomes of the interactions. Characterisation of the

interacting proteins will facilitate identification of additional functions of RMND5

proteins potentially mediated by the RING domain or by the additional protein domains

present in the proteins. Alternatively, individual RMND5 protein interacting factors

may function to regulate RMND5 activity. Although the proteins identified in these co-

immunoprecipitation experiments should contain RMND5 binding partners, the proteins

are not expected to be substrates of RMND5 ubiquitination as the interaction between

an E3 ubiquitin ligase and its substrate is usually transient and may be weak, especially

if the E3 ubiquitin ligase is a multimeric complex (Burande et al., 2009; Andrews et al.,

2010). In the future, experiments involving mass spectrometric identification of

RMND5 protein interacting factors may be performed using either wild-type

RMND5A/RMND5B or mutant RMND5A/RMND5B, including RING domain

mutants. Mutation of the RING domain may prolong the interaction between the E3

ubiquitin ligase and its substrate as the substrate can no longer be ubiquitinated by the

E3 enzyme (Hu and Fearon, 1999). Comparison of the proteins binding to wild-type or

mutant RMND5A/RMND5B may therefore identify proteins which are more likely to


298

be substrate proteins (Hu and Fearon, 1999). Alternatively, for large scale screening of

potential RMND5 substrate proteins, protein microarrays may be used employing a

microarray based system and in vitro ubiquitination assays which are each carried out

using the E3 ubiquitin ligase of interest and different potential substrate proteins (Gupta

et al., 2007; Persaud et al., 2009; Andrews et al., 2010). In this way, multiple

potentially relevant substrates of RMND5 protein ubiquitination could be identified

which may then be confirmed individually using in vitro and in vivo assays.

Although RMND5A and RMND5B are widely expressed, factors that regulate their

expression have not been investigated and these studies may be commenced using

transcription factor binding bioinformatics tools to predict potential transcription factor

binding sites in the promoter or untranslated regions (UTR) of the RMND5A and

RMND5B genes. Following the identification of potential transcription factor binding

sites, chromatin immunoprecipitation (ChIP), DNase I footprinting and electrophoretic

mobility shift assays (EMSA) can be performed to specifically detect protein-DNA

interactions, and luciferase assays may be used to assess transcriptional activation or

repressor functions of the transcription factors on the putative binding elements in the

regulatory regions of the RMND5A and RMND5B genes (Liu et al., 2010; Kerschner

and Harris, 2012). Finally, site directed mutagenesis of putative transcription factor

binding sites may be included in EMSA and luciferase assays to confirm the function of

the elements and associated transcription factors in the regulation of RMND5A and/or

RMND5B expression.

Previous studies in the laboratory identified that RMND5B exerted a transcriptional

repressor activity on an NKX3.1 consensus binding element (Dawson, 2006) and to

further characterise the transcriptional regulatory role of RMND5B and the potential

transcriptional function of RMND5A, luciferase assays may be used to determine

whether RMND5A is also able to exert transcriptional repressor activity upon an

NKX3.1 responsive element, with results confirmed using ChIP assays and EMSAs. If

RMND5 proteins are shown from mass spectrometry screens to bind transcription

factors or mediators of transcription factor complexes, this may be further investigated

using immunoprecipitation and ChIP assays, which will assist in characterising the

mechanisms of transcriptional regulatory activity, as RMND5 proteins do not contain

identifiable DNA binding domains (Shang et al., 2002; Perissi et al., 2004). In

conjunction with these studies, microarray technology may be used to identify potential


299

RMND5 target genes following overexpression or knockdown of the expression of each

protein (Yang et al., 2008; Gorte et al., 2011).

Both RMND5A and RMND5B were found to exhibit a diffuse nuclear and cytoplasmic

localisation, however a punctate cytoplasmic distribution was also evident in a

proportion of cells. The characterisation of these punctate bodies and whether they are

of functional significance may be ascertained in future studies (Section 4.3) in particular

to follow up results of mass spectrometric identification of RMND5A and RMND5B

binding partners which included mitochondrial proteins. This finding is interesting

given that gluconeogenic and TCA cycle enzymes have been identified as substrates of

the yeast Vid30 complex (Santt et al., 2008). In relation to the broad intracellular

distribution of RMND5 proteins, it will be important to determine where the CTLH

complex forms, whether it functions as an E3 ubiquitin ligase complex, and whether the

CTLH and Vid30 complexes retain overlapping substrates (Kobayashi et al., 2007;

Santt et al., 2008). To assess its function, the CTLH complex may be reconstituted in

vitro, as has been successfully performed for other E3 ubiquitin ligase complexes

(Skowyra et al., 1999; Chen et al., 2006b). These studies may then be used to determine

whether either or both RMND5A and RMND5B can form part of the CTLH complex,

and whether the inclusion of RMND5A and/or RMND5B in the CTLH complex alters

the substrates or type of ubiquitination of the substrates. In yeast, Gid9 functions with

RMD5 to provide the Vid30 complex with its E3 ubiquitin ligase activity and a similar

function of human EMP, orthologue of Gid9, may be investigated in relation to the

CTLH complex in future studies (Braun et al., 2011). The functional consequences of

an additional E3 ubiquitin ligase in the CTLH complex can also be determined,

including augmentation of the E3 activity of RMND5A/RMND5B and alternative types

of ubiquitination of substrate proteins caused by interaction with additional E2

conjugating enzymes. If the CTLH complex can be reconstituted in vitro it is possible

that all or part of the complex could be crystallised, providing information related to the

roles of each complex member that would complement mapping of the protein-protein

interaction domains to delineate CTLH complex topology (Menssen et al., 2012).

Accumulation of this information may also resolve the substrate recognition

components of the CTLH complex, which are currently hypothesised to be RanBPM

and muskelin due to the presence of their SPRY and Kelch repeat domains (Kuang et

al., 2010; Lee et al., 2010), as well as substrates of the CTLH complex.


300

Since the expression, tissue distribution and biological activities of RMND5 proteins

are incompletely characterised, the generation of RMND5A and RMND5B transgenic

or knockout mice may aid in the elucidation of their normal physiological roles. Based

on the similar activities and high amino acid sequence homology of RMND5A and

RMND5B, it is feasible that RMND5A and/or RMND5B knockout mice may not

exhibit a distinctive phenotype as loss of expression of one RMND5 protein may be

compensated by the function of the other, therefore knockout of both RMND5A and

RMND5B may need to be generated. Conversely, if RMND5 knockout or transgenic

mice display embryonic lethality or die shortly after birth, conditional knockout or

transgenic mice may be an option for the investigation of RMND5 protein functions.

These animals may be observed for changes in development, growth, fertility and

behaviour and their tissues may be analysed using, for example, immunohistochemistry

and gene expression analysis to determine the effects of the abnormal expression of

RMND5 proteins (Crawley, 2007).

The ultimate aim of this research would be to determine the expression and function of

each RMND5 protein in normal human tissues as well as in cancer or other pathological

states. For these studies, mRNA expression, protein levels and the tissue distribution of

each of RMND5A and RMND5B may be investigated if appropriate tissues are

available. As RMND5 proteins are highly homologous they may have similar or

overlapping targets, however their tissue distribution may differ resulting in distinct

expression profiles of RMND5 regulated proteins which include a spectrum of

commonly regulated proteins amongst proteins whose expression is specifically

regulated by RMND5A or RMND5B. Tissue microarrays would also allow the

comparison of RMND5 protein expression in normal versus tumour tissue samples for

different cancers types, with protein analysis and real-time RT-PCR of these tissues

potentially identifying concordant changes in the expression of specific RMND5

substrates in cancer types where RMND5 proteins function as important determinants of

gene or protein expression. Analysis of the chromosomal loci of RMND5A and

RMND5B, 2p11.2 and 5q35.3, respectively using FISH or other technologies would

enable determination of abnormalities involving the RMND5 gene loci (e.g. gene copy

number), and sequencing of each gene would enable the assessment of mutations in the

RMND5 coding regions in malignant tissues that may affect their protein function and

therefore the regulation of their substrates (Bartlett, 2004).


301

7.3 Concluding Remarks This study has demonstrated that human RMND5 proteins function as E3 ubiquitin

ligases in prostate cancer cells, with RMND5A interacting with UbcH2, UbcH5b and

UbcH5c, and RMND5B interacting with UbcH5b and UbcH5c to mediate ubiquitin

transfer. Both RMND5A and RMND5B are able to ubiquitinate the prostate specific

tumour suppressor, NKX3.1 resulting in its proteasome-dependent degradation. It is

possible that RMND5A and RMND5B form part of the multiprotein CTLH complex,

and that the additional LisH, CTLH and CRA domains of each protein are able to

mediate alternative, as yet uncharacterised functions. As both RMND5 chromosomal

loci are disrupted in a number of cancers, the identification of RMND5 substrates of

ubiquitination and their additional cellular functions may increase knowledge of their

contributions to normal physiology and to cancer initiation or progression.

Chapter 8 References

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Appendix I Buffers and Solutions

Appendix I: Buffers and Solutions


350

Buffers and Solutions 1. 2% Acetic Acid Acetic Acid 10mL ddH2O 490mL Acetic acid and ddH2O were combined and the solution was stored at room temperature. 2. 1% and 2 % Agarose Gel 1%

Agarose Gel

2% Agarose

Gel Agarose Powder

3g 6g

1X TAE100 300mL 300mL Ethidium Bromide26(10mg/mL)

8μL 8μL

Agarose powder was added to TAE100, the solution was microwaved until the agarose had dissolved and ethidium bromide26 was added. Agarose was stored at room temperature and liquefied by heating in a microwave prior to use. 3. 1M Ammonium Chloride (NH4Cl) Ammonium Chloride 5.36g ddH2O 100mL Ammonium chloride was dissolved in ddH2O and the solution was stored at room temperature.

4. 10% Ammonium Persulphate (APS) Ammonium Persulphate 0.1g ddH2O 1mL Ammonium persulphate was dissolved in ddH2O and the solution was stored at 4°C. 5. Ampicillin (100 mg/mL) Ampicillin Powder 100mg ddH2O 1mL Ampicillin powder was dissolved in ddH2O and the solution was stored at -20°C. 6. 50mM Calcium Chloride (CaCl2) 1M Calcium Chloride7 5mL ddH2O 100mL The reagents were mixed and the solution sterilised using a 0.2μM filter and stored at room temperature. 7. 1M Calcium Chloride (CaCl2) Calcium Chloride 27g ddH2O 100mL Calcium chloride was dissolved in ddH2O and the solution then sterilised using a 0.2μM filter and stored at room temperature. 8. 50mM Chloroquine Chloroquine 257mg ddH2O 10mL Chloroquine was dissolved in ddH2O and the solution stored shielded from light at room temperature.


351

9. Confocal Blocking Buffer Horse Serum 5mL BSA Fraction V 500mg Sodium Azide 10mg PBS56 45mL Reagents were combined and the solution was stored at 4˚C shielded from light. 10. Coomassie Blue Destaining Solution Methanol 450mL Glacial Acetic Acid 100mL ddH2O 450mL Reagents were combined and the solution stored at room temperature. 11. Coomassie Blue Staining Solution Coomassie Brilliant Blue G 1g Methanol 450mL Glacial Acetic Acid 100mL ddH2O 450mL Coomassie Brilliant Blue G was dissolved in the methanol and ddH2O mixture. The glacial acetic acid was added in a fumehood and the solution was stored at room temperature.

12. Coomassie Blue Staining Solution for Mass Spectrometry Ammonium Sulphate 100g Coomassie Brilliant Blue G 1.2g Orthophosphoric Acid 100mL Methanol 200mL ddH2O 700mL Coomassie Brilliant Blue G was dissolved in 200mL methanol. In a separate bottle, ammonium sulphate was dissolved in 600mL ddH2O, orthophosphoric acid was added and the solution was made up to 800mL using ddH2O. The Coomassie Brilliant Blue G solution was then added and the solution was stored at room temperature.

13. 20mg/mL Cycloheximide Cycloheximide 20mg DMSO 1mL Cycloheximide was dissolved in DMSO and the solution stored shielded from light at -20°C in 100µL aliquots.

14. 10mM dATP 100mM dATP 10µL ddH2O 90µL Reagents were combined and the solution stored at -20°C.

15. DEPC Treated ddH2O Diethylpyrocarbonate (DEPC) 1mL ddH2O 1L DEPC was added to ddH2O in a fumehood, the solution was shaken, left overnight to allow the DEPC to evaporate, autoclaved and stored at room temperature.


352

16. 10-2M Dihydrotestosterone 5α-Dihydrotestosterone 2.9024mg Absolute Ethanol 1mL Reagents were combined and the solution stored at -20°C shielded from light. 17. 10-4M Dihydrotestosterone 10-2M DHT16 10µL Absolute Ethanol 990µL Reagents were combined and the solution stored at -20°C shielded from light. 18. 10-5M Dihydrotestosterone 10-4M DHT17 10µL Absolute Ethanol 90µL Reagents were combined and the solution stored at -20°C, shielded from light. The solution was diluted 1:1000 in culture medium to obtain a concentration of 10-8M. 19. 1M Disodium Hydrogen Orthophosphate (Na2HPO4) Disodium Hydrogen Orthophosphate 35.48g ddH2O 250mL Disodium Hydrogen Orthophosphate was dissolved in ddH2O by heating in a waterbath at 30°C. Once dissolved the solution was stored at room temperature.

20. 1M Dithiothreitol (DTT) Dithiothreitol 1.25g ddH2O 8mL DTT was dissolved in ddH2O, the solution was divided into 1mL aliquots and stored at -20°C. 21. 50mM Dithiothreitol (DTT) in 20mM Tris pH7.5 1M DTT20 0.5mL 1M Tris108 pH7.5 0.2mL ddH2O 9.3mL Reagents were combined and the solution used immediately. 22. 6X DNA Loading Dye Bromophenol Blue 10mg Sucrose 7g 0.1M EDTA27 pH8.0 2mL ddH2O 8mL Reagents were combined and the solution was made up to 10mL with ddH2O then stored at room temperature. 23. 10mM and 25mM dNTP (PCR) 10mM

dNTP 25mM dNTP

100mM dATP

10µL 25µL

100mM dTTP

10µL 25µL

100mM dGTP

10µL 25µL

100mM dCTP

10µL 25µL

ddH2O 60µL - Reagents were combined and the solution was stored at -20ºC.


353

24. ECL PlusTM Detection Reagent Equal volumes of Solution A and Solution B (GE Healthcare) were combined at room temperature immediately prior to use. 25. 70% , 75% and 95% (v/v) Ethanol 70%

Ethanol 75%

Ethanol 95%

Ethanol Absolute Ethanol

70mL 75mL 95mL

ddH2O 30mL 25mL 5mL Reagents were combined and the solution was stored at room temperature. 26. Ethidium Bromide (10mg/mL) Ethidium Bromide 10mg ddH2O 1mL Ethidium bromide was dissolved in ddH2O and the solution was stored shielded from light at room temperature. 27. 0.1M Ethylenediaminetetra-acetic acid (EDTA) pH8.0 0.5M EDTA26 pH8.0 100mL ddH2O 400mL 0.5M EDTA29 was diluted in ~400mL ddH2O, the pH adjusted with sodium hydroxide, and the solution made up to 500mL, autoclaved and stored at room temperature.

28. 0.5M Ethylenediaminetetra-acetic acid (EDTA) pH 8.0 EDTA 186.12g Sodium Hydroxide ~20g EDTA was dissolved in 500mL ddH2O, the pH adjusted to 8.0 with sodium hydroxide pellets and the volume adjusted to 1L with ddH2O. The solution was mixed and left overnight, autoclaved and stored at room temperature. 29. 4% Formaldehyde 40% Formaldehyde 5 mL PBS56 45mL Reagents were combined and the solution was stored at room temperature shielded from light. 30. 50% Glutathione Sepharose Stock Glutathione Sepharose 4B 1 bottle PBS56 pH7.3 500mL PBS pH7.3 /0.02% Sodium Azide58

~20mL Glutathione Sepharose 4B beads were transferred into a 50mL tube and allowed to settle at 4°C. The supernatant was discarded and the beads resuspended in 50mL cold PBS56 pH7.3 then stored at 4°C to allow to settle again. This procedure was carried out five times. After the last wash, the beads were allowed to settle at 4°C, the supernatant removed and the beads resuspended in an equal volume PBS pH7.3/0.02% sodium azide58 then stored at 4°C.


354

31. 50% Glutathione Sepharose 50% Glutathione Sepharose Stock30

300μL PBS56 pH 7.3 ~30mL For each experimental sample, 300μL 50% Glutathione sepharose stock30 bead slurry was centrifuged for 1 minute at 3000rpm 4°C, the supernatant removed and the beads resuspended in 5mL cold PBS56. This wash was repeated four times then the 150μL beads were resuspended in 150μL PBS56 to produce a 50% Glutathione sepharose slurry which was used immediately. 32. GST Elution Buffer Reduced Glutathione 0.062g 1M Tris108 pH8 1mL 50mM ZnCl2

115 25µL ddH2O 9mL Reagents were combined and the solution used immediately. 33. Glycerol/PIPES Buffer 1M CaCl2

7 12mL 0.5M PIPES pH7.059 4mL Glycerol 30mL ddH2O 138mL Reagents were combined and the solution sterilised using a 0.2μM filter then stored at 4°C. 34. 10mg/mL Hœchst 33258 Hœchst 33258 25mg DMSO 2.5mL Hœchst 33258 was dissolved in DMSO, the solution divided into 50μL aliquots and stored at -20°C.

35. 100mM Isopropyl β-D-1-thiogalactopyranoside (IPTG) IPTG 1.2g ddH2O 50mL IPTG was dissolved in ~40mL ddH2O and the final volume adjusted to 50mL, the solution was sterilised using a 0.2μM filter, divided into 1mL aliquots and stored at -20°C. 36. Kanamycin (100 mg/mL) Kanamycin Powder 100mg ddH2O 1mL Kanamycin powder was dissolved in ddH2O and the solution was stored at -20°C. 37. 10mM Lactacystin Lactacystin 2mg ddH2O 531μL Lactacystin was dissolved in ddH2O, the solution was divided into 50μL aliquots and stored at -20°C. The solution was diluted 1:1000 to obtain a working concentration of 10µM in culture medium. 38. LB Agar LB Broth43 500mL Agar Bacteriologica 7.5g Agar was dissolved in the LB broth43, the solution was autoclaved then stored at room temperature. Prior to use, agar was liquefied in a microwave.


355

39. LB Agar/Ampicillin LB agar38 was liquefied in a microwave, 15mL per petri dish was poured into a sterile 50mL tube, the solution was cooled to ~60°C, 1µL per mL of 100mg/mL ampicillin5 was added (final concentration 100µg/mL ampicillin), the solution was inverted to mix, poured into petri dishes and allowed to set at room temperature. 40. LB Agar/Ampicillin/IPTG/X-gal LB Agar/Ampicillin plates39 were spread with 100μL 100mM IPTG35 and 20μL 50mg/mL X-Gal113 then allowed to dry for 30 minutes at 37°C before use. 41. LB Agar/Kanamycin LB agar38 was liquefied in a microwave, 15mL per petri dish was poured into a sterile 50mL tube, the solution was cooled to ~60°C, 1µL per mL of 100mg/mL kanamycin36 was added (final concentration 100µg/mL kanamycin), the solution was inverted to mix, poured into petri dishes and allowed to set at room temperature. 42. LB Agar/Kanamycin/IPTG/X-gal LB Agar/Kanamycin plates41 were spread with 100μL 100mM IPTG35 and 20μL 50mg/mL X-Gal113 then allowed to dry for 30 minutes at 37°C before use.

43. Luria-Bertani (LB) Broth Tryptone 10g Yeast Extract 5g Sodium Chloride 10g Reagents were dissolved in ~800mL ddH2O, the pH was adjusted to 7.0, the solution was made up to 1L with ddH2O, autoclaved and stored at room temperature. 44. LB Broth/Ampicillin For a final concentration of 100µg/mL ampicillin, 1µL 100mg/mL ampicillin5 was added per mL LB Broth43 immediately prior to use. 45. LB Broth/10% Glycerol LB Broth43 9mL Glycerol 1mL Glycerol was mixed with LB broth43 and the solution was stored at -20°C. 46. LB Broth/Kanamycin For a final concentration of 100µg/mL kanamycin, 1µL of 100mg/mL kanamycin36 was added per 1mL of LB Broth43. 47. Lysis Buffer (Miltenyi) for Immunoprecipitation Lysis Buffer 4850µL 200mM PMSF 60 25µL 40x Protease Inhibitor Cocktail63

125µL Reagents were combined and used immediately at 4°C.


356

48. Lysozyme (10mg/mL) Lysozyme 100mg 1M Tris108 pH8.0 2mL ddH2O 8mL Reagents were combined, the solution was sterilised using a 0.2µM filter then stored at -20ºC in 1mL aliquots. 49. 1M Magnesium Chloride (MgCl2) MgCl2 203g ddH2O 1L MgCl2 was dissolved in ddH2O, the solution was autoclaved then stored at room temperature. 50. 10mM MG132 MG132 4.756mg Ethanol 1mL MG132 was dissolved in ethanol and the solution stored at -20°C in 100µL aliquots. The solution was diluted 1:1000 in culture medium to obtain a concentration of 10µM. 51. 20x MOPS Buffer (Gradient Polyacrylamide Gels) MOPS 104.6g Tris Base 60.6g SDS 10g EDTA 3g ddH2O 500mL Reagents were dissolved in 400mL ddH2O, the volume made up to 500mL and the solution stored at 4°C. 20X MOPS buffer was diluted to 1X with ddH2O and used as required.

52. Mounting Medium Tris-PO4 buffer110 5mL ddH2O 75mL Polyvinylalcohol 20g Glycerol 30mL Chlorobutanol 100mg 1% Phenol Red 2-3 drops Tris-PO4 buffer and ddH2O were combined in an Erhlemeyer flask, PVA and 2-3 drops Phenol red were added and the flask placed in a 60ºC waterbath and shaken intermittently to dissolve. Glycerol was slowly added, then chlorobutanol. The pH was adjusted to 8.2 with Tris-PO4 buffer and the mounting medium stored at 4º. 53. NETN Buffer NP40 2.5mL 4M NaCl81 12.5mL 1M Tris108 pH8.0 10mL 0.5M EDTA27 pH8.0 1mL Reagents were combined and the solution stored at 4°C. For the production of GST-RING domains, 0.5M EDTA27 pH8.0 was omitted from the NETN buffer. 54. 5X PCR Buffer Taq 10X PCR Buffer 5mL (without MgCl2) 25mM dNTP23 200µL ddH2O 4.8mL Reagents were combined and the solution stored at -20°C in 1mL aliquots.


357

55. 300μM Phalloidin Stock TRITC-Phalloidin 0.1mg DMSO 255μL Phalloidin was dissolved in DMSO and stored in 20μL aliquots at -20°C. 56. Phosphate Buffered Saline (PBS) pH7.3, pH7.4 Sodium Chloride 8g Potassium Chloride 0.2g Disodium Hydrogen Orthophosphate 1.44g Potassium Dihydrogen Orthophosphate 0.44g Reagents were dissolved in ~800mL ddH2O, the pH was adjusted using 10M HCl, the solution was made up to 1L using ddH2O, autoclaved, and stored at room temperature. 57. PBS/1% BSA BSA Fraction V 100mg PBS56 pH7.4 10mL Reagents were combined and the solution was stored at 4˚C. 58. Phosphate Buffered Saline (PBS) pH7.3/0.02% Sodium Azide PBS56 pH7.3 19.96mL 10% Sodium Azide80 40μL Reagents were combined and the solution stored at 4°C.

59. 0.5M PIPES PIPES 15.15g ddH2O 100mL PIPES was dissolved in ~80mL ddH2O, the pH adjusted to 7.0 using 1M NaOH and the volume made up to 100mL with ddH2O. The solution was sterilised by passing through a 0.2μM filter and stored at room temperature. 60. 200mM PMSF PMSF 348.4mg Isopropanol 10mL Reagents were combined in a fumehood and the solution was stored at room temperature. 61. 1Kb PlusTM Molecular Weight Marker (DNA Ladder) 1Kb Plus Molecular Weight Marker 250µL 6X DNA Loading Dye22 833.3µL The reagents were combined, the volume adjusted to 5mL using Storage Buffer98 and the solution was stored in 1mL aliquots at -20º. 62. 5M Potassium Acetate Potassium Acetate 24.5g ddH2O 50mL Potassium acetate was dissolved in ~40mL ddH2O, the solution was made up to 50mL with ddH2O, autoclaved then stored at room temperature.


358

63. 40X Protease Inhibitor Cocktail Protease Inhibitor Cocktail Tablets 4 ddH2O 1mL Tablets were dissolved in ddH2O, the solution was divided into aliquots and stored at 4°C. 64. 50% Protein A Sepharose Stock Protein A Sepharose 1 bottle 50mM Tris107 pH 7.0 500mL 50mM Tris pH7.0 /0.02% Sodium Azide109 ~20mL

Protein A sepharose beads were transferred into a 50mL tube and allowed to settle at 4°C. The supernatant was discarded and the beads resuspended in 50mL cold 50mM Tris107 pH7.0 and stored at 4°C to allow the beads to settle again. This procedure was carried out five times. After the last wash, the beads were allowed to settle at 4°C, the supernatant removed, the beads resuspended in an equal volume of 50mM Tris pH7.0/0.02% sodium azide109 and stored at 4°C.

65. 50% Protein A Sepharose 50% Protein A Sepharose Stock64

300μL 50mM Tris107 pH7.0 ~10mL For each experimental sample, 300μL 50% Protein A sepharose stock64 bead slurry was centrifuged for 1 minute at 3000rpm/4°C, the supernatant removed and the beads resuspended in 1mL cold 50mM Tris107 pH7.0. The wash was repeated four times then the 150μL beads were resuspended in 150μL PBS56 pH7.4 to produce 300μL of a 50% Protein A sepharose slurry, which was divided into 100μL aliquots, stored at 4°C and used within 2 days. 66. 50% Protein G Sepharose Stock Protein G Sepharose 4 Fast Flow 1 bottle 20mM Sodium Phosphate90 pH 7.0 500mL 20mM Sodium Phosphate pH7.0 /0.02% Sodium Azide91 ~20mL

Protein G sepharose beads were transferred into a 50mL tube and allowed to settle at 4°C. The supernatant was discarded and the beads resuspended in 50mL cold 20mM sodium phosphate90 pH7.0 and stored at 4°C to allow the beads to settle again. This procedure was carried out five times. After the last wash the beads were allowed to settle at 4°C, the supernatant removed and the beads resuspended in an equal volume of 20mM sodium phosphate pH7.0/0.02% sodium azide91 and stored at 4°C.


359

67. 50% Protein G Sepharose 50% Protein G Sepharose Stock66

300μL 50mM Tris107 pH7.0 ~10mL For each experimental sample, 300μL 50% Protein G sepharose stock66 bead slurry was centrifuged for 1 minute at 3000rpm/4°C, the supernatant removed and the beads resuspended in 1mL cold 50mM Tris107 pH7.0. The wash was repeated four times then the 150μL beads were resuspended in 150μL PBS56 pH7.4 to produce 300μL of a 50% Protein G sepharose slurry, which was divided into 100μL aliquots, stored at 4°C and used within 2 days. 68. 100U/mL Pyrophosphatase Pyrophosphatase 96.8 units 20mM Tris107 pH7.5 968μL Pyrophosphatase was dissolved in 20mM Tris107 pH7.5, the solution divided into 100μL aliquots and stored at -20°C.

69. RIPA Buffer 1% Sodium Deoxycholate87 625µL 1M Tris108 pH7.4 125µL NP40 25µL 200mM Sodium Orthovanadate88 12.5µL 0.5M EDTA28 pH8.0 5µL 0.5M Sodium Fluoride85 5µL 200mM PMSF60 12.5µL 40X Protease Inhibitor Cocktail63 50µL 4M NaCl81 93.8μL ddH2O 1.6 mL Reagents were combined, with PMSF60 and 40x protease inhibitor cocktail63 added last, and the solution was used immediately. 70. RNase A (10mg/mL) RNase A 100mg 1M Tris108 pH 7.4 100µL 4M Sodium Chloride81 37.5µl Reagents were combined, the volume was adjusted to 10mL using ddH2O, heated to 100ºC for 15 minutes and allowed to cool to room temperature. The solution was then stored in 1mL aliquots at -20ºC. 71. RPMI 1640 Media RPMI 1640 1 sachet Sodium Hydrogen Carbonate 2g ddH2O 1L RPMI was dissolved in ddH2O, sodium hydrogen carbonate was added and the solution was filtered using a 0.2µm filter into 1L sterile bottles, then stored at 4°C.


360

72. RPMI 1640/PS RPMI 164071 990mL PS 10mL Reagents were combined and stored at 4°C. 73. RPMI 1640/PS/5%CSS Charcoal Stripped Serum 10mL RPMI1640/PS72 190mL Reagents were combined and the solution stored at 4°C. 74. RPMI 1640/PS/10% FCS RPMI 1640/PS72 450mL Foetal Calf Serum 50mL Reagents were combined and the solution was stored at 4°C. 75. RPMI 1640/PS/10% FCS/10% DMSO RPMI 1640/PS/10% FCS74 9mL DMSO 1mL

DMSO was dissolved in RPMI 1640/ PS/10%FCS74 and the solution was used immediately. 76. 10X Running Buffer Tris 30g Glycine 144g SDS 10g ddH2O 1L Reagents were dissolved in ddH2O, the volume adjusted to 1L and the solution was stored at room temperature. 10X Running Buffer was diluted to 1X with ddH2O and used as required.

77. 2X and 10X SDS-PAGE Loading Buffer 2X 10X Glycerol 1mL 12.5mL 1M Tris108 pH6.8

2.5mL 2.5mL

20% SDS84 2mL 2.5mL 2-Mercaptoethanol

200µL 2.5mL

Bromophenol Blue

0.01g 0.25g

ddH2O To 10mL

To 25mL

Reagents were combined, the volume adjusted with ddH2O and the solution was stored at -20ºC in 1mL aliquots. 78. 12% Separating Gel (Western Blotting) 1 M Tris108 pH8.8 1.775mL 40% Acrylamide 3mL (37:5:1) 20% SDS84 37.5µL 10% APS4 37.5µL TEMED 3.75µL ddH2O 2.515mL Reagents were combined, with 10% APS and TEMED added last, the solution was inverted to mix and used immediately. 79. 3M Sodium Acetate (pH4.6) Sodium Acetate 49.2g ddH2O 200mL Sodium acetate was dissolved in 160mL ddH2O, the pH adjusted to 4.6, the volume adjusted to 200mL and the solution autoclaved and stored at room temperature.


361

80. 10% Sodium Azide (NaN3) Sodium Azide 0.1g ddH2O 1mL Sodium azide was dissolved in ddH2O and the solution stored at 4°C. 81. 4M Sodium Chloride (NaCl) Sodium Chloride 117.2g ddH2O 500mL Sodium chloride was dissolved in ddH2O, the solution was autoclaved and stored at room temperature. 82. 1% Sodium Deoxycholate Sodium Deoxycholate 1g ddH2O 100mL Sodium deoxycholate was dissolved in ddH2O, the solution was autoclaved and stored at -20°C shielded from light. 83. 1M Sodium Dihydrogen Orthophosphate (NaHPO42H2O) Sodium Dihydrogen Orthophosphate 39g ddH2O 250mL Reagents were combined and the solution was stored at room temperature. 84. 20% Sodium Dodecyl Sulphate (SDS) Sodium Dodecyl Sulphate 10g ddH20 50mL SDS was dissolved in ddH2O and the solution was stored at room temperature.

85. 0.5M Sodium Fluoride (NaF) Sodium Fluoride 2.1g ddH2O 100mL Sodium fluoride was dissolved in ddH2O and the solution was stored at room temperature. 86. Sodium Hydrogen Phosphate (Na2HPO4) Na2HPO4 156.2g ddH2O 1L Na2HPO4 was dissolved in ~800mL ddH2O, the solution was made up to 1L and stored at room temperature. 87. 10M Sodium Hydroxide (NaOH) Sodium Hydroxide 20g ddH2O 50mL Sodium hydroxide was dissolved in ddH2O and the solution was stored at room temperature. 88. 200mM Sodium Orthovanadate Sodium Orthovanadate 3.7g ddH2O 100mL Sodium orthovanadate was dissolved in ~80mL ddH2O, the pH adjusted to 10.0, the solution heated at ~95°C until colourless and the pH readjusted to 10.0. The procedure was repeated until the solution was colourless at pH 10.0, then the solution was made up to 100mL with ddH2O, divided into 1mL aliquots and stored at -20°C.


362

89. 0.1M Sodium Phosphate pH7 1M Na2HPO4

86 57.7mL 1M NaHPO42H2O83 42.3mL ddH2O 900mL Reagents were dissolved in ~800mL ddH2O and the pH adjusted to 7. The solution was made up to 1L, autoclaved and stored at room temperature. 90. 20mM Sodium Phosphate pH7 Sodium Phosphate 100mL ddH2O 400mL The reagents were combined, the solution autoclaved then stored at room temperature. 91. 20mM Sodium Phosphate pH7/0.02% Sodium Azide 20mM Sodium Phosphate90 19.96mL 10% Sodium Azide80 40μL Reagents were combined and the solution stored at 4°C. 92. Solution I (Plasmid Preparation) Glucose 0.9g 1M Tris108 pH 8.8 2.5mL 0.5M EDTA28 pH8.0 2.0mL ddH2O 95.5mL Reagents were combined and the solution was autoclaved then stored at 4°C.

93. Solution II (Plasmid Preparation) 20% SDS84 250µL 10M Sodium Hydroxide87 100µL ddH2O 4.65mL Reagents were combined and the solution was stored at room temperature and used on the day of preparation. 94. Solution III (Plasmid Preparation) 5M Potassium Acetate62 60mL Glacial Acetic Acid 11.5mL ddH2O 28.5mL The reagents were combined, the pH adjusted to 8.0 and the solution was autoclaved and stored at 4°C. 95. Sonication Buffer 1M Tris108 pH8.0 25mL 4M NaCl81 6.25mL 0.5M EDTA28 pH 8 1mL ddH2O 467.5mL Reagents were combined and the solution stored at 4°C. 96. 4% Stacking Gel (Western Blotting) 1 M Tris108 pH6.8 1.25mL 40% Acrylamide (37:5:1) 1.3mL 20% SDS84 50µL 10% APS4 50µL TEMED 10µL ddH20 7.4mL Reagents were combined, with 10% APS and TEMED added last, the solution was inverted to mix and used immediately.


363

97. Streptavidin-HRP Solution TBS101 5mL Solution A (Vectastain) 2 drops Solution B (Vectastain) 2 drops Solution A and Solution B were added to the TBS101, the solution mixed and allowed to stand for 30 minutes. A working solution was made by diluting 100µL Streptavidin-HRP solution per 3mL buffer in 1% TBST/BSA104 and used on the day of preparation. 98. Storage Buffer (for 1Kb Plus DNA Ladder) 1M Tris108 pH 7.4 50µL 0.5M EDTA28 pH 8.0 10µL 4M Sodium Chloride81 62.5µL Reagents were combined, the volume adjusted to 5mL with ddH2O and the solution was stored at room temperature. 99. 20% Sucrose Sucrose 10g ddH20 50mL Sucrose was dissolved in ~25mL ddH2O, the solution was made up to 50mL with ddH2O and stored at room temperature.

100. 50X Tris Acetate EDTA (TAE) Tris Base 242g Glacial Acetic Acid 57.1mL 0.5M EDTA28 pH8.0 100mL Tris base was dissolved in ddH2O, the glacial acetic acid and EDTA were added, the volume made up to 1L with ddH2O and the solution autoclaved then stored at room temperature. 50X TAE was diluted to 1X with ddH2O as required and stored at room temperature. 101. Tris Buffered Saline (TBS) 1M Tris108 pH 7.4 50mL 4M Sodium Chloride81 37.5mL Tris and sodium chloride were combined, the volume adjusted to 1L with ddH2O and the solution was stored at room temperature. 102. TBS Tween (TBST) TBS101 500mL Tween-20 1mL The reagents were combined, mixed by inversion and the solution was stored at room temperature. 103. TBST/1%, 3% and 5% Blotto

Skim milk powder was dissolved in TBST102. The solution was made on the day of use and stored at room temperature.

1% Blotto

2% Blotto

5% Blotto

Skim Milk Powder

0.3g 0.9g 1.5g

TBST102 30mL 30mL 30mL


364

104. TBST/1% BSA Bovine Serum Albumin 0.1g TBST102 10mL BSA was dissolved in TBST102 and the solution used immediately. 105. Transfer Buffer (Western Blotting) Glycine 14.4g Tris 3.03g Methanol 200mL ddH2O 800mL Glycine and Tris were dissolved in ~500mL ddH2O, the volume made up to 800mL, 200mL methanol was added and the solution was stored at -20°C for >1 hour prior to use. Solution was used on the day of preparation. 106. 0.5M Tris pH7.0 Tris Base 60.55g ddH2O 1L Tris was dissolved in ~800mL ddH2O, the pH was adjusted using 10M HCl to 7.0 and the solution was made up to 1L with ddH2O. The solution was autoclaved and stored at room temperature. 107. 20mM and 50mM Tris pH7.0 20mM 50mM

0.5M Tris106 pH7.0

40mL 100mL

ddH2O 1L 1L 0.5M Tris was diluted to ~800mL with ddH2O, the pH adjusted to 7.0 using 10M HCl, the solution made up to 1L, autoclaved and stored at room temperature.

108. 1M Tris (pH6.8, 7.5, 7.5, 8.0, 8.8) Tris Base 121.1g ddH2O 1L Tris was dissolved in ~800mL ddH2O, the pH was adjusted using 10M HCl and the solution was made up to 1L with ddH2O. The solution was autoclaved and stored at room temperature. 109. 50mM Tris pH7/0.02% Sodium Azide 50mM Tris pH7.0107 19.96mL 10% Sodium Azide80 40μL Reagents were combined and the solution stored at 4°C. 110. Tris-PO4 pH9.0 Tris Base 121.9g ddH2O 1L Tris was dissolved in ~800mL ddH2O, the pH adjusted to 9 with 1M NaH2PO4

86 and the volume made up to 1L with ddH2O. The solution was autoclaved and stored at room temperature. 111. 0.1%/10% (v/v) Triton X-100 0.1% Triton

X-100 10% Triton

X-100 Triton X-100

50μL 5mL

PBS56 pH7.4

49.95mL 45mL

Reagents were combined and the solution was stored at room temperature shielded from light.


365

112. Whole Cell Lysis Buffer 20% Sucrose99 2.5mL 20% SDS84 0.5mL 1M Tris108 pH6.8 0.25mL 2-Mercaptoethanol 0.25mL ddH2O 1.5mL Reagents were combined, the solution inverted to mix and stored in a fumehood protected from light for up to 4 weeks. 113. 50mg/mL X-gal X-gal 0.5g Dimethylformamide 10mL X-gal was dissolved in dimethylformamide, the solution aliquoted and stored at -20°C. 114. 1M Zinc Chloride (ZnCl2) ZnCl2 6.815g ddH2O 50mL Reagents were combined and the ZnCl2 dissolved by heating. The solution was stored at room temperature. 115. 50mM ZnCl2 1M ZnCl2

114 5mL ddH2O 95mL Reagents were combined and the solution stored at room temperature.

Appendix II Primer Sequences

Appendix II: Primer Sequences


366

Sequencing Primers pEGFP1266-S 5’ CAT GGT CCT GCT GGA GTT CGT G 3’ M13-S 5’ GTT TTC CCA GTC ACG AC 3’ M13-AS 5’ CAG GAA ACA GCT ATG AC 3’ pGEX-S 5’ GGG CTGGCA AGC CAC GTT TGG TG 3’ pGEX-AS 5’ CCG GGA GCT GCA TGT GTC AGA GG 3’ RMND5BSalI1-S 5’ ATG GGA TCC ATG GAG CAG TGT 3’ RMND5BSalI1182-AS 5’ATG GGA TCC TCA GAA TAT GAT GCG 3’ RMND5B790-S 5’ ATC TGT GAG ACC TTT ACC CGG 3’ RMND5A603-S 5’ CTT GTT AAT GGG TGG AAC CA 3’ RMND5A490-AS 5’CTC CAC CAT CAC TTC ATT GA 3’ RanBPM1259-AS 5’ CCC ATT CTT CCT GCT AAT AC 3’ RanBPM687-S 5’ AAG CTT TAA TGG GAA TTG GTC TTT CTG 3’ Cloning Primers RMND5BBamHI1-S 5’ATG GGA TCC ATG GAG CAG TGT 3’ RMND5BBamHI1182-AS 5’ ATG GGA TCC TCA GAA TAT GAT GCG 3' RMND5A1-S 5’ GAA TTC ATG GAT CAG TGC GTG ACG 3’ RMND5ABamHI1-S 5’ GGA TCC ATG GAT CAG TGC GTG ACG 3’ RMND5A1176-AS 5’ GAA TTC TCA GAA AAA TAT CTG TTT GGC 3’ RanBPM1-S 5’ AAG CTT TAA TGT CCG GGC AGC CGC 3’ RanBPM1029-S 5’ GGA GTG GAG AAC CAA AAT CC 3’ RanBPM2190-AS 5’ GTC GAC CTA ATG TAG GTA GTC TTC CAC 3’


367

RMND5ARING1006-S 5’ GGA TCC TGC CCC ATT CTT CGT CAG 3’ RMND5ARING1131-AS 5’ GAA TTC CAC AGT AGG GAC ATT TTA ATT 3’ RMND5BRING1012-S 5’ GGA TCC TGC CCC ATC CTC CGC CAG 3’ RMND5BRING1137-AS 5’ GAA TTC CAC AGT AGG GAC ACT TCA GC 3’ CBLRING1141-S 5’ GGA TCC TGT AAA ATA TGT GCT GAA AAT GA 3’ CBLRING1257-AS 5’ GAA TTC CGC AGA AAG GAC AGC CCT G 3’ Gene Expression Primers βActin146-S 5’ AGA AGG ATT CCT ATG TGG GCG ACG A 3’ βActin464-AS 5’ CGA GTC CAT CAC GAT GCC AGT GGT A 3’ Twa1335-S 5’ TGA TCC GCC AGC GGG AGA CA 3’ Twa1681-AS 5’ GGG CTC CTC AAT CAC ACC CTT GC 3’ EMP580-S 5’ AGC TGC CTG GAG TTC AGC CTC A 3’ EMP1083-AS 5’ GAC GTA GCC GTT GGG CAG CA 3’ ARMC81103-S 5’ AGG TGC GGT TAG CTG CCG TC 3’ ARMC81668-AS 5’ TGT CCC ATC CGC TAT GTT GGC T 3’ Muskelin1675-S 5’ TGT CCA AGG TTT GCC CAT CAG C 3’ Muskelin2120-AS 5’ ACC AGG TTG CCT TTA GGA GGA GT 3’ C17orf39298-S 5’ TCC GCG GCC TCA CTC ATC CC 3’ C17orf39814-AS 5’ AGC CCT CTA TGG AGG CTG CTG A 3’ RanBPM1550-S 5’ TAATATCAAATAAAGCACATCAAT CAT 3’ RanBPM2190-AS 5’ GTC GAC CTA ATG TAG GTA GTC TTC CAC 3’


368

RMND5B790-S 5’ ATC TGT GAG ACC TTT ACC CGG 3’ RMND5BTOPO1182-AS 5’ GAA TAT GAT GCG TTT CCC ATC TGC 3’ RMND5A603-S 5’ CTT GTT AAT GGG TGG AAC CA 3’ RMND5A1176-AS 5’ GAA TTC TCA GAA AAA TAT CTG TTT GGC 3’ Site Directed Mutagenesis Primers RMND5A(C356S)1045-S 5’ CCC ATG AAA TTG GTC TCTG GTCA TAT TAT ATC AAG AGA TGC C 3’ RMND5A(C356S)1083-AS 5’ ATC TCT TGA TAT AAT ATG ACC AGA GAC CAA TTT CAT GGG TGG 3’ Primer Set 1 RMND5A(C356A)1045-S 5’ CCC ATG AAA TTG GTC GCT GGT CAT ATT ATA TCA AGA 3’ RMND5A(C356A)1083-AS 5’ ATC TCT TGA TAT AAT ATG ACC AGC GAC CAA TTT CAT GGG TGG 3’ Primer Set 2 RMND5A(C356A/H358A)1045-S 5’ CCC ATG AAA TTG GTC GCT GGT GCT ATT ATA TCA AGA GAT GCC 3’ RMND5A(C356A/H358A)1083-AS 5’ ATC TCT TGA TAT AAT AGC ACC AGC GAC CAA TTT CAT GGG TGG 3’ Primer Set 1 RMND5B(C358S)1051-S 5’ CCC ATC AAG CTC ATC TCT GGC CAT GTT ATC TCC CGA GAT GCA 3’ RMND5B(C358S)1089-AS 5’ ATC TCG GGA GAT AAC ATG GCC AGA GAT GAG CTT GAT GGG AGG 3’ Primer Set 2 RMND5B(C358S)1057-S 5’ AAG CTC ATC TCT GGC CAT GTT ATC TCC 3’ RMND5B(C358S)1080-AS 5’ GAT AAC ATG GCC AGA GAT GAG CTT GAT GGG 3’ RMND5B(C358A/H360S)1054-S 5’ATC AAG CTC ATC GCT GGC GCT GTT ATC TCC CGA 3’ RMND5B(C358A/H360A)1083-AS 5’ GGA GAT AAC AGC GCC AGC GAT GAG CTT GAT GGG AGG 3’

Appendix III Sequencing

Appendix III: Sequencing

Appendix III Sequences

369

pGEX-RMND5A – Representative Sequence >gi|15082505|gb|BC012165.1| Homo sapiens required for meiotic nuclear division 5 homolog A (S. cerevisiae), mRNA (cDNA clone MGC:20406 IMAGE:4636136), complete cds Length=3239 GENE ID: 64795 RMND5A | required for meiotic nuclear division 5 homolog A (S. cerevisiae) [Homo sapiens] (10 or fewer PubMed links) Score = 1581 bits (1752), Expect = 0.0 Identities = 975/1023 (96%), Gaps = 23/1023 (2%) Strand=Plus/Plus Query 99 CATGGATCAGTGCGTGACGGTGGAGCGCGAGCTGGAGAAGGTGCTGCACAAGTTCTCAGG 158 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 98 CATGGATCAGTGCGTGACGGTGGAGCGCGAGCTGGAGAAGGTGCTGCACAAGTTCTCAGG 157 Query 159 CTACGGGCAGCTGTGCGAGCGCGGCCTGGAGGAGCTCATCGACTACACCGGCGGCCTCAA 218 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 158 CTACGGGCAGCTGTGCGAGCGCGGCCTGGAGGAGCTCATCGACTACACCGGCGGCCTCAA 217 Query 219 GCACGAGATCCTGCAGAGCCACGGCCAAGATGCTGAATTATCAGGGACACTTTCACTTGT 278 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 218 GCACGAGATCCTGCAGAGCCACGGCCAAGATGCTGAATTATCAGGGACACTTTCACTTGT 277 Query 279 TTTGACACAGTGCTGTAAAAGAATAAAGGATACTGTTCAAAAATTGGCCTCCGACCACAA 338 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 278 TTTGACACAGTGCTGTAAAAGAATAAAGGATACTGTTCAAAAATTGGCCTCCGACCACAA 337 Query 339 AGACATCCACAGCAGTGTTTCTCGGGTTGGAAAAGCCATTGATAAGAATTTTGATTCTGA 398 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 338 AGACATCCACAGCAGTGTTTCTCGGGTTGGAAAAGCCATTGATAAGAATTTTGATTCTGA 397 Query 399 CATTAGCAGTGTGGGAATAGATGGCTGCTGGCAGGCAGACAGCCAAAGGCTTCTCAATGA 458 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 398 CATTAGCAGTGTGGGAATAGATGGCTGCTGGCAGGCAGACAGCCAAAGGCTTCTCAATGA 457 Query 459 AGTGATGGTGGAGCACTTCTTTCGACAAGGAATGCTGGATGTGGCTGAGGAGCTCTGTCA 518 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 458 AGTGATGGTGGAGCACTTCTTTCGACAAGGAATGCTGGATGTGGCTGAGGAGCTCTGTCA 517


370


371

pGEX-RMND5A wild type RING domain

RefSeq -----------------------------------------TGCCCCATTCTTCGTCAGC RING5A CTCCCGGCCGCCATGGCGGCCGCGGGAATTCGATTGGATCCTGCCCCATTCTTCGTCAGC ******************* RefSeq AAACAACAGATAACAATCCACCCATGAAATTGGTCTGTGGTCATATTATATCAAGAGATG RING5A AAACAACAGATAACAATCCACCCATGAAATTGGTCTGTGGTCATATTATATCAAGAGATG ************************************************************ RefSeq CCCTGAATAAAATGTTTAATGGTAGCAAATTAAAATGTCCCTACTGT------------- RING5A CCCTGAATAAAATGTTTAATGGTAGCAAATTAAAATGTCCCTACTGTGGAATTCAATCAC *********************************************** RefSeq = RMND5A RING domain reference sequence

RING5A = pGEX-RMND5A RING domain (Wild-type)


372

pGEX-RMND5B wild type RING domain

RefSeq ---------------------------------------TGCCCCATCCTCCGCCAGCAG RING5B CCCGGCCGCCATGGCGGCCGCGGGAATTCGATTGGATCCTGCCCCATCCTCCGCCAGCAG ********************* RefSeq ACGTCAGATTCCAACCCTCCCATCAAGCTCATCTGTGGCCATGTTATCTCCCGAGATGCA RING5B ACGTCAGATTCCAACCCTCCCATCAAGCTCATCTGTGGCCATGTTATCTCCCGAGATGCA ************************************************************ RefSeq CTCAATAAGCTCATTAATGGAGGAAAGCTGAAGTGTCCCTACTGT--------------- RING5B CTCAATAAGCTCATTAATGGAGGAAAGCTGAAGTGTCCCTACTGTGGAATTCAATCACTA *********************************************

RefSeq = RMND5B RING domain reference sequence

RING5B = pGEX-RMND5B RING domain (Wild-type)


373

pGEX-CBL RING domain

RefSeq --------------------------------------TGTAAAATATGTGCTGAAAATG CBL CCGGCCGCCATGGCGGCCGCGGGAATTCGATTGGATCCTGTAAAATATGTGCTGAAAATG ********************** RefSeq ATAAGGATGTAAAGATTGAGCCCTGTGGACACCTCATGTGCACATCCTGTCTTACATCCT CBL ATAAGGATGTAAAGATTGAGCCCTGTGGACACCTCATGTGCACATCCTGTCTTACATCCT ************************************************************ RefSeq GGCAGGAATCAGAAGGTCAGGGCTGTCCTTTCTGC------------------------- CBL GGCAGGAATCAGAAGGTCAGGGCTGTCCTTTCTGCGGAATTCAATCACTAGTGAATTCGC ***********************************

RefSeq = CBL RING domain reference sequence

CBL = pGEX-CBL RING domain


374

pEGFP-RMND5A (C356S) RING domain

RING5A TGCCCCATTCTTCGTCAGCAAACAACAGATAACAATCCACCCATGAAATTGGT ||||||||||||||||||||||||||||||||||||||||||||||||||||| RefSeq TGCCCCATTCTTCGTCAGCAAACAACAGATAACAATCCACCCATGAAATTGGT RING5A CTCTGGTCATATTATATCAAGAGATGCCCTGAATAAAATGTTTAATGGTAGCAAATTAAA || ||||||||||||||||||||||||||||||||||||||||||||||||||||||||| RefSeq CTGTGGTCATATTATATCAAGAGATGCCCTGAATAAAATGTTTAATGGTAGCAAATTAAA RING5A ATGTCCCTACTGT ||||||||||||| RefSeq ATGTCCCTACTGT

RefSeq = RMND5A RING domain reference sequence

RING5A = pEGFP-RMND5A RING (C356S) mutant

G1061C (C356S) (Base change marked in red)

Single base change at position 425 C (*) on chromatogram

*


375

pEGFP-RMND5B (C358S) RING domain

RING5B TGCCCCATCCTCCGCCAGCAGACGTCAGATTCCAACCCTCCCATCAAGCTCATCTCTGGCC ||||||||||||||||||||||||||||||||||||||||||||||||||||||| ||||| RefSeq TGCCCCATCCTCCGCCAGCAGACGTCAGATTCCAACCCTCCCATCAAGCTCATCTGTGGCC RING5B ATGTTATCTCCCGAGATGCACTCAATAAGCTCATTAATGGAGGAAAGCTGAAGTGTCCCT |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| RefSeq ATGTTATCTCCCGAGATGCACTCAATAAGCTCATTAATGGAGGAAAGCTGAAGTGTCCCT RING5B ACTGT ||||| RefSeq ACTGT


RING5B = pEGFP-RMND5B RING (C358S) mutant

G1067C (C358S) (Base change marked in red)

Single base change at position 251 C (*) on chromatogram

*


376

pEGFP-RMND5A (C356A/H358A) RING domain

RING5A TGCCCCATTCTTCGTCAGCAAACAACAGATAACAATCCACCCATGAAATTGGTC |||||||||||||||||||||||||||||||||||||||||||||||||||||| RefSeq TGCCCCATTCTTCGTCAGCAAACAACAGATAACAATCCACCCATGAAATTGGTC RING5A GCTGGTGCTATTATATCAAGAGATGCCCTGAATAAAATGTTTAATGGTAGCAAATTAA |||| |||||||||||||||||||||||||||||||||||||||||||||||||| RefSeq TGTGGTCATATTATATCAAGAGATGCCCTGAATAAAATGTTTAATGGTAGCAAATTAA RING5A AATGTCCCTACTGT |||||||||||||| RefSeq AATGTCCCTACTGT

RefSeq = RMND5A RING domain reference sequence

RING5A = pEGFP-RMND5A RING (C356A/H358A) mutant

T1060G/G1061C, C1066G/A1067C (C356A/H358A) (Base changes marked in red)

Base changes at positions 434 G, 435 C (*) and 440 G, 441 C (#) on chromatogram

* #


377

pEGFP-RMND5B (C358A/H360A) RING domain

RING5B TGCCCCATCCTCCGCCAGCAGACGTCAGATTCCAACCCTCCCATCAAGCTCATCGCTGGC |||||||||||||||||||||||||||||||||||||||||||||||||||||| |||| RefSeq TGCCCCATCCTCCGCCAGCAGACGTCAGATTCCAACCCTCCCATCAAGCTCATCTGTGGC RING5B GCTGTTATCTCCCGAGATGCACTCAATAAGCTCATTAATGGAGGAAAGCTGAAGTGTCCC |||||||||||||||||||||||||||||||||||||||||||||||||||||||||| RefSeq CATGTTATCTCCCGAGATGCACTCAATAAGCTCATTAATGGAGGAAAGCTGAAGTGTCCC RING5B TACTGT |||||| RefSeq TACTGT


RING5B = pEGFP-RMND5B RING (C358A/H360A) mutant

T1066G/G1067C, C1072G/A1073C (C358A/H360A) (Base changes marked in red)

Base changes at positions 161 G, 162 C (*) and 167 G, 168 C (#) on chromatogram

* #


378

pmCherry-RanBPM (55kDa) - Representative Sequence >gi|39812377|ref|NM_005493.2| Homo sapiens RAN binding protein 9 (RANBP9), mRNA Length=3132 GENE ID: 10048 RANBP9 | RAN binding protein 9 [Homo sapiens] (Over 10 PubMed links) Score = 1842 bits (2042), Expect = 0.0 Identities = 1088/1126 (96%), Gaps = 10/1126 (0%) Strand=Plus/Plus Query 11 AATAGACTACCAGGTTGGGATAAGCATTCATATGGTTACCATGGGGATGATGGACATTCG 70 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 783 AATAGACTACCAGGTTGGGATAAGCATTCATATGGTTACCATGGGGATGATGGACATTCG 842 Query 71 TTTTGTTCTTCTGGAACTGGACAACCTTATGGACCAACTTTCACTACTGGTGATGTCATT 130 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 843 TTTTGTTCTTCTGGAACTGGACAACCTTATGGACCAACTTTCACTACTGGTGATGTCATT 902 Query 131 GGCTGTTGTGTTAATCTTATCAACAATACCTGCTTTTACACCAAGAATGGACATAGTTTA 190 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 903 GGCTGTTGTGTTAATCTTATCAACAATACCTGCTTTTACACCAAGAATGGACATAGTTTA 962 Query 191 GGTATTGCTTTCACTGACCTACCGCCAAATTTGTATCCTACTGTGGGGCTTCAAACACCA 250 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 963 GGTATTGCTTTCACTGACCTACCGCCAAATTTGTATCCTACTGTGGGGCTTCAAACACCA 1022 Query 251 GGAGAAGTGGTCGATGCCAATTTTGGGCAACATCCTTTCGTGTTTGATATAGAAGACTAT 310 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 1023 GGAGAAGTGGTCGATGCCAATTTTGGGCAACATCCTTTCGTGTTTGATATAGAAGACTAT 1082 Query 311 ATGCGGGAGTGGAGAACCAAAATCCAGGCACAGATAGATCGATTTCCTATCGGAGATCGA 370 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 1083 ATGCGGGAGTGGAGAACCAAAATCCAGGCACAGATAGATCGATTTCCTATCGGAGATCGA 1142


379


380

pmCherry-RMND5B - Representative Sequence >gi|95044658|gb|DQ494789.1| Homo sapiens RMND5B mRNA, complete cds Length=1825 GENE ID: 64777 RMND5B | required for meiotic nuclear division 5 homolog B (S. cerevisiae) [Homo sapiens] (10 or fewer PubMed links) Score = 609 bits (674), Expect = 2e-170 Identities = 337/337 (100%), Gaps = 0/337 (0%) Strand=Plus/Plus Query 22 CCCCCTTAGCGTCAGCTTTGCCTCTGGCTGTGTGGCGCTGCCTGTGTTGATGAACATCAA 81 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 1026 CCCCCTTAGCGTCAGCTTTGCCTCTGGCTGTGTGGCGCTGCCTGTGTTGATGAACATCAA 1085 Query 82 GGCTGTGATTGAGCAGCGGCAGTGCACTGGGGTCTGGAATCACAAGGACGAGTTACCGAT 141 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 1086 GGCTGTGATTGAGCAGCGGCAGTGCACTGGGGTCTGGAATCACAAGGACGAGTTACCGAT 1145 Query 142 TGAGATTGAACTAGGCATGAAGTGCTGGTACCACTCCGTGTTCGCTTGCCCCATCCTCCG 201 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 1146 TGAGATTGAACTAGGCATGAAGTGCTGGTACCACTCCGTGTTCGCTTGCCCCATCCTCCG 1205 Query 202 CCAGCAGACGTCAGATTCCAACCCTCCCATCAAGCTCATCTGTGGCCATGTTATCTCCCG 261 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 1206 CCAGCAGACGTCAGATTCCAACCCTCCCATCAAGCTCATCTGTGGCCATGTTATCTCCCG 1265 Query 262 AGATGCACTCAATAAGCTCATTAATGGAGGAAAGCTGAAGTGTCCCTACTGTCCCATGGA 321 |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| Sbjct 1266 AGATGCACTCAATAAGCTCATTAATGGAGGAAAGCTGAAGTGTCCCTACTGTCCCATGGA 1325 Query 322 GCAGAACCCGGCAGATGGGAAACGCATCATATTCTGA 358 ||||||||||||||||||||||||||||||||||||| Sbjct 1326 GCAGAACCCGGCAGATGGGAAACGCATCATATTCTGA 1362


381

Appendix IV Mass Spectrometry

Appendix IV: Mass Spectrometry

User : Matt F

Email :

Search title : Submitted from PROJ13881 with Decoy database by Mascot Daemon on APAF-WS-08

MS data file : \\Apaf-hpv-file\User_Shared\MFitzhenry\2012 Reports\PROJ13881_JackyBentel(RPerthH)\Band1.mgf

Database : SwissProt 2012x (536029 sequences; 190235160 residues)

Taxonomy : Homo sapiens (human) (20319 sequences)

Timestamp : 8 Jun 2012 at 04:52:21 GMT

Protein hits : HSP7C_HUMAN Heat shock cognate 71 kDa protein OS=Homo sapiens GN=HSPA8 PE=1 SV=1

HSP71_HUMAN Heat shock 70 kDa protein 1A/1B OS=Homo sapiens GN=HSPA1A PE=1 SV=5

HNRPM_HUMAN Heterogeneous nuclear ribonucleoprotein M OS=Homo sapiens GN=HNRNPM PE=1 SV=3

PUF60_HUMAN Poly(U)-binding-splicing factor PUF60 OS=Homo sapiens GN=PUF60 PE=1 SV=1

GRP75_HUMAN Stress-70 protein, mitochondrial OS=Homo sapiens GN=HSPA9 PE=1 SV=2

K2C1_HUMAN

Keratin, type II cytoskeletal 1 OS=Homo sapiens GN=KRT1 PE=1 SV=6

RMD5A_HUMAN Protein RMD5 homolog A OS=Homo sapiens GN=RMND5A PE=1 SV=1

K1C9_HUMAN

Keratin, type I cytoskeletal 9 OS=Homo sapiens GN=KRT9 PE=1 SV=3

K1C10_HUMAN Keratin, type I cytoskeletal 10 OS=Homo sapiens GN=KRT10 PE=1 SV=6

SNW1_HUMAN

SNW domain-containing protein 1 OS=Homo sapiens GN=SNW1 PE=1 SV=1

DDX41_HUMAN Probable ATP-dependent RNA helicase DDX41 OS=Homo sapiens GN=DDX41 PE=1 SV=2

XRCC6_HUMAN X-ray repair cross-complementing protein 6 OS=Homo sapiens GN=XRCC6 PE=1 SV=2

K22E_HUMAN

Keratin, type II cytoskeletal 2 epidermal OS=Homo sapiens GN=KRT2 PE=1 SV=2

K2C6B_HUMAN Keratin, type II cytoskeletal 6B OS=Homo sapiens GN=KRT6B PE=1 SV=5

DDX5_HUMAN

Probable ATP-dependent RNA helicase DDX5 OS=Homo sapiens GN=DDX5 PE=1 SV=1

K2C5_HUMAN


MERL_HUMAN

Merlin OS=Homo sapiens GN=NF2 PE=1 SV=1

RS27A_HUMAN Ubiquitin-40S ribosomal protein S27a OS=Homo sapiens GN=RPS27A PE=1 SV=2

PABP1_HUMAN Polyadenylate-binding protein 1 OS=Homo sapiens GN=PABPC1 PE=1 SV=2

ALBU_HUMAN

Serum albumin OS=Homo sapiens GN=ALB PE=1 SV=2

LMNB1_HUMAN Lamin-B1 OS=Homo sapiens GN=LMNB1 PE=1 SV=2

DHX35_HUMAN Probable ATP-dependent RNA helicase DHX35 OS=Homo sapiens GN=DHX35 PE=1 SV=2

KHDR1_HUMAN KH domain-containing, RNA-binding, signal transduction-associated protein 1 OS=Homo

sapiens GN=KHDRBS1 PE=1 SV=1

PCKGM_HUMAN Phosphoenolpyruvate carboxykinase [GTP], mitochondrial OS=Homo sapiens GN=PCK2 PE=1 SV=3

HNRPQ_HUMAN Heterogeneous nuclear ribonucleoprotein Q OS=Homo sapiens GN=SYNCRIP PE=1 SV=2

MYL10_HUMAN Myosin regulatory light chain 10 OS=Homo sapiens GN=MYL10 PE=2 SV=2

ODP2_HUMAN

Dihydrolipoyllysine-residue acetyltransferase component of pyruvate dehydrogenase complex, mitochondrial OS=Homo

CN16B_HUMAN Uncharacterized protein C14orf166B OS=Homo sapiens GN=C14orf166B PE=2 SV=2

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MS data file

Database

Taxonomy

Timestamp

Protein hits

: Matt F

: : Submitted from PROJ13881 with Decoy database by Mascot Daemon on APAF-WS-08

: \\Apaf-hpv-file\User_Shared\MFitzhenry\2012 Reports\PROJ13881_JackyBentel(RPerthH)\Band-all.mgf

: SwissProt 2012x (536029 sequences; 190235160 residues)

: Homo sapiens (human) (20319 sequences)

: 8 Jun 2012 at 04:50:59 GMT

:PUF60_HUMAN Poly(U)-binding-splicing factor PUF60 OS=Homo sapiens GN=PUF60 PE=1 SV=1

K2C1_HUMAN


K1C9_HUMAN


ACTB_HUMAN

Actin, cytoplasmic 1 OS=Homo sapiens GN=ACTB PE=1 SV=1

HSP7C_HUMAN Heat shock cognate 71 kDa protein OS=Homo sapiens GN=HSPA8 PE=1 SV=1

TBB5_HUMAN

Tubulin beta chain OS=Homo sapiens GN=TUBB PE=1 SV=2


TBB4B_HUMAN Tubulin beta-4B chain OS=Homo sapiens GN=TUBB4B PE=1 SV=1

GRP78_HUMAN 78 kDa glucose-regulated protein OS=Homo sapiens GN=HSPA5 PE=1 SV=2

HNRPC_HUMAN Heterogeneous nuclear ribonucleoproteins C1/C2 OS=Homo sapiens GN=HNRNPC PE=1 SV=4


K22E_HUMAN


IF4A3_HUMAN Eukaryotic initiation factor 4A-III OS=Homo sapiens GN=EIF4A3 PE=1 SV=4

ATPB_HUMAN

ATP synthase subunit beta, mitochondrial OS=Homo sapiens GN=ATP5B PE=1 SV=3





HS71L_HUMAN Heat shock 70 kDa protein 1-like OS=Homo sapiens GN=HSPA1L PE=1 SV=2

POTEE_HUMAN POTE ankyrin domain family member E OS=Homo sapiens GN=POTEE PE=1 SV=3


K2C6C_HUMAN Keratin, type II cytoskeletal 6C OS=Homo sapiens GN=KRT6C PE=1 SV=3

HNRH1_HUMAN Heterogeneous nuclear ribonucleoprotein H OS=Homo sapiens GN=HNRNPH1 PE=1 SV=4

K2C5_HUMAN


TBA1A_HUMAN Tubulin alpha-1A chain OS=Homo sapiens GN=TUBA1A PE=1 SV=1


K2C8_HUMAN


SF3B3_HUMAN Splicing factor 3B subunit 3 OS=Homo sapiens GN=SF3B3 PE=1 SV=4


IF4A1_HUMAN Eukaryotic initiation factor 4A-I OS=Homo sapiens GN=EIF4A1 PE=1 SV=1

MYO6_HUMAN

Unconventional myosin-VI OS=Homo sapiens GN=MYO6 PE=1 SV=4

RMD5A_HUMAN Protein RMD5 homolog A OS=Homo sapiens GN=RMND5A PE=1 SV=1


TBA4A_HUMAN Tubulin alpha-4A chain OS=Homo sapiens GN=TUBA4A PE=1 SV=1


ATPA_HUMAN

ATP synthase subunit alpha, mitochondrial OS=Homo sapiens GN=ATP5A1 PE=1 SV=1

FLOT2_HUMAN Flotillin-2 OS=Homo sapiens GN=FLOT2 PE=1 SV=2

HNRPF_HUMAN Heterogeneous nuclear ribonucleoprotein F OS=Homo sapiens GN=HNRNPF PE=1 SV=3

ILF2_HUMAN

Interleukin enhancer-binding factor 2 OS=Homo sapiens GN=ILF2 PE=1 SV=2

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DX39A_HUMAN ATP-dependent RNA helicase DDX39A OS=Homo sapiens GN=DDX39A PE=1 SV=2

IDHP_HUMAN

Isocitrate dehydrogenase [NADP], mitochondrial OS=Homo sapiens GN=IDH2 PE=1 SV=2

EF1A1_HUMAN Elongation factor 1-alpha 1 OS=Homo sapiens GN=EEF1A1 PE=1 SV=1


GRP75_HUMAN Stress-70 protein, mitochondrial OS=Homo sapiens GN=HSPA9 PE=1 SV=2

H3C_HUMAN

Histone H3.3C OS=Homo sapiens GN=H3F3C PE=1 SV=3

RALYL_HUMAN RNA-binding Raly-like protein OS=Homo sapiens GN=RALYL PE=2 SV=2

SPF45_HUMAN Splicing factor 45 OS=Homo sapiens GN=RBM17 PE=1 SV=1

CRNL1_HUMAN Crooked neck-like protein 1 OS=Homo sapiens GN=CRNKL1 PE=1 SV=4

XPO2_HUMAN

Exportin-2 OS=Homo sapiens GN=CSE1L PE=1 SV=3

MACF1_HUMAN Microtubule-actin cross-linking factor 1, isoforms 1/2/3/5 OS=Homo sapiens GN=MACF1 PE=1 SV=4

HORN_HUMAN

Hornerin OS=Homo sapiens GN=HRNR PE=1 SV=2

VIP2_HUMAN

Inositol hexakisphosphate and diphosphoinositol-pentakisphosphate kinase 2 OS=Homo sapiens GN=PPIP5K2 PE=1 SV=3

THOC3_HUMAN THO complex subunit 3 OS=Homo sapiens GN=THOC3 PE=1 SV=1

ANXA5_HUMAN Annexin A5 OS=Homo sapiens GN=ANXA5 PE=1 SV=2

PCBP1_HUMAN Poly(rC)-binding protein 1 OS=Homo sapiens GN=PCBP1 PE=1 SV=2

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User : Matt F

Email :

Search title : Submitted from JackyBentel by Mascot Daemon on APAF-WS-08

MS data file : \\Apaf-hpv-file\User_Shared\MFitzhenry\2012 Reports\PROJ14099_Bentel\IP_ALL.mgf



Timestamp : 25 Jul 2012 at 22:29:54 GMT

Protein hits : PUF60_HUMAN Poly(U)-binding-splicing factor PUF60 OS=Homo sapiens GN=PUF60 PE=1 SV=1


K22E_HUMAN


K2C1_HUMAN


K1C9_HUMAN



ACTB_HUMAN




POTEE_HUMAN POTE ankyrin domain family member E OS=Homo sapiens GN=POTEE PE=1 SV=3




ALBU_HUMAN

Serum albumin OS=Homo sapiens GN=ALB PE=1 SV=2

K2C8_HUMAN


PGAM5_HUMAN Serine/threonine-protein phosphatase PGAM5, mitochondrial OS=Homo sapiens GN=PGAM5 PE=1 SV=2




SF3A3_HUMAN Splicing factor 3A subunit 3 OS=Homo sapiens GN=SF3A3 PE=1 SV=1


HNRPK_HUMAN Heterogeneous nuclear ribonucleoprotein K OS=Homo sapiens GN=HNRNPK PE=1 SV=1

RMD5B_HUMAN Protein RMD5 homolog B OS=Homo sapiens GN=RMND5B PE=2 SV=1

PTBP1_HUMAN Polypyrimidine tract-binding protein 1 OS=Homo sapiens GN=PTBP1 PE=1 SV=1

U5S1_HUMAN

116 kDa U5 small nuclear ribonucleoprotein component OS=Homo sapiens GN=EFTUD2 PE=1 SV=1

TBB4A_HUMAN Tubulin beta-4A chain OS=Homo sapiens GN=TUBB4A PE=1 SV=2

EF1A1_HUMAN Elongation factor 1-alpha 1 OS=Homo sapiens GN=EEF1A1 PE=1 SV=1

TAF6_HUMAN

Transcription initiation factor TFIID subunit 6 OS=Homo sapiens GN=TAF6 PE=1 SV=1


PRP19_HUMAN Pre-mRNA-processing factor 19 OS=Homo sapiens GN=PRPF19 PE=1 SV=1

K1H1_HUMAN

Keratin, type I cuticular Ha1 OS=Homo sapiens GN=KRT31 PE=2 SV=3

HSPB1_HUMAN Heat shock protein beta-1 OS=Homo sapiens GN=HSPB1 PE=1 SV=2

NEP_HUMAN

Neprilysin OS=Homo sapiens GN=MME PE=1 SV=2

ATPB_HUMAN

ATP synthase subunit beta, mitochondrial OS=Homo sapiens GN=ATP5B PE=1 SV=3

MYO6_HUMAN

Unconventional myosin-VI OS=Homo sapiens GN=MYO6 PE=1 SV=4

RN128_HUMAN E3 ubiquitin-protein ligase RNF128 OS=Homo sapiens GN=RNF128 PE=1 SV=1

PLCA_HUMAN

1-acyl-sn-glycerol-3-phosphate acyltransferase alpha OS=Homo sapiens GN=AGPAT1 PE=2 SV=2

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Email :

Search title : Submitted from JackyBentel by Mascot Daemon on APAF-WS-08

MS data file : \\Apaf-hpv-file\User_Shared\MFitzhenry\2012 Reports\PROJ14099_Bentel\M_ALL.mgf



Timestamp : 25 Jul 2012 at 22:27:48 GMT

Protein hits : PUF60_HUMAN Poly(U)-binding-splicing factor PUF60 OS=Homo sapiens GN=PUF60 PE=1 SV=1


K2C1_HUMAN



K2C6A_HUMAN Keratin, type II cytoskeletal 6A OS=Homo sapiens GN=KRT6A PE=1 SV=3




K22E_HUMAN





IF4A3_HUMAN Eukaryotic initiation factor 4A-III OS=Homo sapiens GN=EIF4A3 PE=1 SV=4

K1C9_HUMAN




HNRPF_HUMAN Heterogeneous nuclear ribonucleoprotein F OS=Homo sapiens GN=HNRNPF PE=1 SV=3

PGAM5_HUMAN Serine/threonine-protein phosphatase PGAM5, mitochondrial OS=Homo sapiens GN=PGAM5 PE=1 SV=2


U5S1_HUMAN

116 kDa U5 small nuclear ribonucleoprotein component OS=Homo sapiens GN=EFTUD2 PE=1 SV=1

ACTB_HUMAN


ILF2_HUMAN

Interleukin enhancer-binding factor 2 OS=Homo sapiens GN=ILF2 PE=1 SV=2

DHX35_HUMAN Probable ATP-dependent RNA helicase DHX35 OS=Homo sapiens GN=DHX35 PE=1 SV=2

RALY_HUMAN

RNA-binding protein Raly OS=Homo sapiens GN=RALY PE=1 SV=1

KRT84_HUMAN Keratin, type II cuticular Hb4 OS=Homo sapiens GN=KRT84 PE=1 SV=2

PRP19_HUMAN Pre-mRNA-processing factor 19 OS=Homo sapiens GN=PRPF19 PE=1 SV=1


TAF6_HUMAN

Transcription initiation factor TFIID subunit 6 OS=Homo sapiens GN=TAF6 PE=1 SV=1

RBMX_HUMAN

RNA-binding motif protein, X chromosome OS=Homo sapiens GN=RBMX PE=1 SV=3

SF3A3_HUMAN Splicing factor 3A subunit 3 OS=Homo sapiens GN=SF3A3 PE=1 SV=1

CDC5L_HUMAN Cell division cycle 5-like protein OS=Homo sapiens GN=CDC5L PE=1 SV=2

PRP8_HUMAN

Pre-mRNA-processing-splicing factor 8 OS=Homo sapiens GN=PRPF8 PE=1 SV=2

FL2D_HUMAN

Pre-mRNA-splicing regulator WTAP OS=Homo sapiens GN=WTAP PE=1 SV=2

HNRPR_HUMAN Heterogeneous nuclear ribonucleoprotein R OS=Homo sapiens GN=HNRNPR PE=1 SV=1



RSMB_HUMAN

Small nuclear ribonucleoprotein-associated proteins B and B' OS=Homo sapiens GN=SNRPB PE=1 SV=2

THOC1_HUMAN THO complex subunit 1 OS=Homo sapiens GN=THOC1 PE=1 SV=1


SNW1_HUMAN

SNW domain-containing protein 1 OS=Homo sapiens GN=SNW1 PE=1 SV=1

PTBP2_HUMAN Polypyrimidine tract-binding protein 2 OS=Homo sapiens GN=PTBP2 PE=1 SV=1

U520_HUMAN

U5 small nuclear ribonucleoprotein 200 kDa helicase OS=Homo sapiens GN=SNRNP200 PE=1 SV=2

HNRPK_HUMAN Heterogeneous nuclear ribonucleoprotein K OS=Homo sapiens GN=HNRNPK PE=1 SV=1


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CCD25_HUMAN Coiled-coil domain-containing protein 25 OS=Homo sapiens GN=CCDC25 PE=1 SV=2

VAPA_HUMAN

Vesicle-associated membrane protein-associated protein A OS=Homo sapiens GN=VAPA PE=1 SV=3

DX39B_HUMAN Spliceosome RNA helicase DDX39B OS=Homo sapiens GN=DDX39B PE=1 SV=1

CC150_HUMAN Coiled-coil domain-containing protein 150 OS=Homo sapiens GN=CCDC150 PE=2 SV=2

RU2A_HUMAN

U2 small nuclear ribonucleoprotein A' OS=Homo sapiens GN=SNRPA1 PE=1 SV=2

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1. PUF60_HUMAN Mass: 59838 Score: 1105 Matches: 47(30) Sequences: 19(17) emPAI: 2.34

Poly(U)-binding-splicing factor PUF60 OS=Homo sapiens GN=PUF60 PE=1 SV=1

Check to include this hit in error tolerant search or archive report

Query

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12 8

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the functional characterisation of human rmnd5 …...5.2.2 rmnd5 proteins colocalise with nkx3.1 in...

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