vanessa oakes thesis (pdf 6mb)

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Cyclin A/CDK2 regulates

multiple G2 phase pathways

Vanessa Oakes BSc (Hons)

A thesis submitted to Queensland University of Technology

in fulfilment of the requirements for the degree of

Doctor of Philosophy

AUGUST 2012

Diamantina Institute for Cancer, Immunology and Metabolic Medicine

University of Queensland, Princess Alexandra Hospital, Woolloongabba, QLD, Australia

and

Queensland University of Technology, School of Biomedical Sciences, Faculty of Health,

Brisbane, QLD, Australia

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TABLE OF CONTENTS

Statement of originality ............................................................................. Error! Bookmark not defined.

Contribution by other colleagues .............................................................. Error! Bookmark not defined.

Acknowledgements .................................................................................................................................. vi

Abstract .................................................................................................................................................. viii

Publications ............................................................................................................................................. xii

List of Abbreviations .............................................................................................................................. xiii

List of Figures ........................................................................................................................................ xvii

List of Tables ......................................................................................................................................... xixx

Chapter 1 Summary of the literature. ................................................................................................. 1-54

1.1 The cell cycle, structure and function. ......................................................................................... 2

1.1.1 Stages of mitosis ................................................................................................................... 2

1.2 Cancer and the cell cycle .............................................................................................................. 6

1.3 Regulation of cell cycle................................................................................................................. 7

1.3.1 The cyclin/CDK complexes .................................................................................................... 8

1.3.2 Cell cycle checkpoints ......................................................................................................... 13

1.4 Exploiting the DNA damage checkpoints, a cancer therapy. ..................................................... 17

1.5 DNA damage pathways .............................................................................................................. 18

1.5.1 ATM/ATR ............................................................................................................................. 20

1.5.2 Chk1 and Chk2..................................................................................................................... 24

1.5.3 p53 ...................................................................................................................................... 27

1.5.4 Cdc25 ................................................................................................................................... 28

1.6 Other important cell cycle regulators ........................................................................................ 32

1.6.1 p38 MAPK ............................................................................................................................ 32

1.6.2 Claspin ................................................................................................................................. 35

1.6.3 Wee1 ................................................................................................................................... 37

1.6.4 Cdh1 .................................................................................................................................... 38

1.6.5 Plk1 ...................................................................................................................................... 40

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1.7 Checkpoint recovery ................................................................................................................... 42

1.8 G2/M regulation by cyclin A/CDK2 and cyclin B/CDK1 .............................................................. 44

1.9 Chk1 regulation .......................................................................................................................... 47

1.9.1 The role of Chk1 during an unperturbed cell cycle ............................................................. 50

1.9.2 Chk1 inactivation ................................................................................................................. 52

1.10 Hypothesis ................................................................................................................................ 53

1.11 Significance ............................................................................................................................... 54

1.12 Aims of the study ...................................................................................................................... 54

Chapter 2 Materials and Methods .................................................................................................... 55-82

2.1 Antibodies ................................................................................................................................... 56

2.2 Cell Biology Methods. ................................................................................................................. 58

2.2.1 Cell culture ........................................................................................................................... 58

2.2.2 Cell Synchronisation. ........................................................................................................... 59

2.2.3 Cell Treatments. .................................................................................................................. 59

2.2.4 Transfection Treatments. .................................................................................................... 60

2.2.5 Cell fixation and immunofluorescent labelling. ................................................................... 62

2.2.6 Microscopy and image acquisition ...................................................................................... 65

2.2.7 FACS Analysis ....................................................................................................................... 66

2.2.8 Cellomics .............................................................................................................................. 68

2.3 Molecular Biology Methods ....................................................................................................... 69

2.3.1 Plasmid list ........................................................................................................................... 69

2.3.2 Plasmid construct cloning........................................................................................................ 74

2.3.2.1. Generation of Cherry-Chk1 and GST-Chk1 constructs. ................................................... 74

2.3.2.1 Generation of PBD-GFP .................................................................................................... 75

2.4 Protein Biochemistry Methods. .................................................................................................. 76

2.4.1 Immunoblot Analysis ........................................................................................................... 76

2.4.2 Immunoprecipitation ........................................................................................................... 77

2.4.3 Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and immunoblot

analyses ........................................................................................................................................ 79

2.4.4 Cyclin A/CDK2 kinase assay ................................................................................................. 80

Chapter 3 Active Chk1 is present in unperturbed G2 phase ........................................................... 83-111

3.1 INTRODUCTION .......................................................................................................................... 84

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3.2 RESULTS...................................................................................................................................... 87

3.2.1 Activated Chk1 is present during an unperturbed cell cycle. ............................................. 87

3.2.2 ATR is responsible for Chk1 phosphorylation in an unperturbed cell cycle during G2 phase.

..................................................................................................................................................... 97

3.2.3 pChk1 localises to the centrosomes during mitosis. ......................................................... 101

3.3 DISCUSSION .............................................................................................................................. 104

Chapter 4 Characterising the cyclin A/CDK2 dependent G2 phase delay ..................................... 112-146

4.1 INTRODUCTION ........................................................................................................................ 113

4.2 RESULTS.................................................................................................................................... 116

4.2.1 Cyclin A depletion delays G2 phase progression and entry into mitosis .......................... 116

4.2.2 Inhibition of CDK2 with a specific inhibitor delays G2 phase progression into mitosis. ... 119

4.2.3 Cyclin A/ CDK2 regulates the timing of centrosomal cyclin B/ CDK1 activation ............... 124

4.2.4 Cyclin A / CDK2 localises to the centrosomes in late G2 phase. ....................................... 129

4.2.5 Cyclin A/ CDK2 depletion or inhibition affects the exit from G2 / M checkpoint. ............ 132

4.3 DISCUSSION .............................................................................................................................. 138

Chapter 5 Chk1 is required to implement the cyclin A/ CDK2 dependent G2 phase delay .......... 147-171

5.1 INTRODUCTION ........................................................................................................................ 148

5.2 RESULTS.................................................................................................................................... 150

5.2.1 Chk1 contributes to the cyclin A/ CDK2 dependent G2 delay. ......................................... 150

5.2.2 Inhibition of G2 phase CDK2 causes an accumulation of G2 phase cells and activated Chk1.

................................................................................................................................................... 156

5.2.3 Chk2 siRNA cannot overcome the cyclin A/CDK2 dependent G2 phase delay. ................ 158

5.2.4 ATR phosphorylates Chk1 to impose the cyclin A dependent G2 phase delay. ............... 161

5.2.5 Activated Chk1 is not a consequence of cyclin A siRNA induced DNA damage................ 162

5.3 DISCUSSION .............................................................................................................................. 165

Chapter 6 Cyclin A/CDK2 regulation of Chk1 ................................................................................ 172-207

6.1 INTRODUCTION ........................................................................................................................ 173

6.2 RESULTS.................................................................................................................................... 175

6.2.1 pChk1 and Cyclin A co-localise during G2 phase. ............................................................. 175

6.2.2 Chk1 and Cyclin A interact during G2 phase. .................................................................... 177

6.2.3 Cyclin A can phosphorylate Chk1 on Ser286 and Ser301. ................................................ 180

6.2.4 Over-expression of Chk1 delays mitotic entry. ................................................................. 181

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6.2.5 In vivo phosphorylation of Chk1 Ser286 and Ser301 is not dependent on cyclin A/CDK2.

.................................................................................................................................................... 193

6.2.6 Chk1 mutants do not change localisation. ........................................................................ 199

6.3 DISCUSSION .............................................................................................................................. 200

Chapter 7 Identification of other G2 phase pathways regulated by cyclin A/CDK2 ..................... 208-246

7.1 INTRODUCTION ........................................................................................................................ 209

7.2 RESULTS .................................................................................................................................... 212

7.2.1 Plk1 inhibition resembles cyclin A/CDK2 depletion/inhibition. ........................................ 212

7.2.2 Cyclin A siRNA causes mislocalisation of Plk1. .................................................................. 214

7.2.3 Cdh1 level is reduced with cyclin A depletion. ................................................................ 2255

7.2.4 Reduction of Cdh1 does not affect the cyclin A/CDK2 dependent G2 delay. ................. 2311

7.2.5 Claspin is required during the cyclin A/CDK2 dependent G2 phase delay to regulate pChk1

levels. ........................................................................................................................................ 2322

7.3 DISCUSSION .............................................................................................................................. 240

Chapter 8 Final Discussion ............................................................................................................. 247-263

Chapter 9 References .................................................................................................................... 264-317

Chapter 10 Appendix .................................................................................................................... .318-320

10.1 Appendix 1 ............................................................................................................................. 319

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QUT Verified Signature

QUT Verified Signature

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Acknowledgements

I am indebted to my supervisor, Associate Professor Brian Gabrielli, for his advice,

assistance and support over the last few years. I am grateful for having a great mentor that

empowered me to grow as an individual both personally and professional. Sometimes it’s

the hard times, such as writing this thesis, to realise the positive outcome of the challenges

and I am sure I will continue to reflect on the many experiences you have given me.

Thanks also to my QUT supervisor, Professor Adrian Herington. Your continued support has

allowed me to get this far and I will be forever grateful.

I would sincerely like to thank past and present Gabrielli Laboratory colleagues and friends

who supported me and contributed towards this thesis. In particular I would like to

especially thank Dr Heather Beamish who has helped greatly throughout this whole process.

I would also like to individually mention the following member of the Lab; Dr Francis

Stevens, Nichole Giles, Leonore de Boer, Puji Astuti (also for sharing her Birthday),

Stephanie Le, Sabrina Kee, Max Ranall, Brittney Harrington, and Rose Boutros. Many thanks

also to Sandrine Roy for help with the Confocal/Microscopy, as well as your life guidance.

Finally, I owe a huge thank you to my family and friends who have been there from the

beginning and have willingly shared my ups and downs of life as a student. The dedication

required to finally complete this journey is attributed to my late Grandfather, Charles

Wobcke. I will always be grateful for your constant love and support.

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Abstract

The cell cycle is a carefully choreographed series of phases that when executed successfully

will allow the complete replication of the genome and the equal division of the genome and

other cellular content into two independent daughter cells. The inability of the cell to

execute cell division successfully can result in either checkpoint activation to allow repair

and/or apoptosis and/or mutations/errors that may or may not lead to tumourgenesis.

Cyclin A/CDK2 is the primary cyclin/CDK regulating G2 phase progression of the cell cycle.

Cyclin A/CDK2 activity peaks in G2 phase and its inhibition causes a G2 phase delay that we

have termed ‘the cyclin A/CDK2 dependent G2 delay’. Understanding the key pathways that

are involved in the cyclin A/CDK2 dependent G2 delay has been the primary focus of this

study.

Characterising the cyclin A/CDK2 dependent G2 delay revealed accumulated levels of the

inactive form of the mitotic regulator, cyclin B/CDK1. Surprisingly, there was also increased

microtubule nucleation at the centrosomes, and the centrosomes stained for markers of

cyclin B/CDK1 activity. Both microtubule nucleation at the centrosomes and

phosphoprotein markers were lost with short-term treatment of CDK1/2 inhibition. Cyclin

A/CDK2 localised at the centrosomes in late G2 phase after separation of the centrosomes

but before the start of prophase. Thus G2 phase cyclin A/CDK2 controls the timing of entry

into mitosis by controlling the subsequent activation of cyclin B/CDK1, but also has an

unexpected role in coordinating the activation of cyclin B/CDK1 at the centrosome and in

the nucleus. In addition to regulating the timing of cyclin B/CDK1 activation and entry into

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mitosis in the unperturbed cell cycle, cyclin A/CDK2 also was shown to have a role in G2

phase checkpoint recovery.

Known G2 phase regulators were investigated to determine whether they had a role in

imposing the cyclin A/ CDK2 dependent G2 delay. Examination of the critical G2 checkpoint

arrest protein, Chk1, which also has a role during unperturbed G2/M phases revealed the

presence of activated Chk1 in G2 phase, in a range of cell lines. Activated Chk1 levels were

shown to accumulate in cyclin A/CDK2 depleted/inhibited cells. Further investigations

revealed that Chk1, but not Chk2, depletion could reverse the cyclin A/CDK2 dependent G2

delay. It was confirmed that the accumulative activation of Chk1 was not a consequence of

DNA damage induced by cyclin A depletion. The potential of cyclin A/CDK2 to regulate Chk1

revealed that the inhibitory phosphorylations, Ser286 and Ser301, were not directly

catalysed by cyclin A/CDK2 in G2 phase to regulate mitotic entry. It appeared that the ability

of cyclin A/CDK2 to regulate cyclin B/CDK1 activation impacted cyclin B/CDK1s

phosphorylation of Chk1 on Ser286 and Ser301, thereby contributing to the delay in G2/M

phase progression.

Chk1 inhibition/depletion partially abrogated the cyclin A/CDK2 dependent G2 delay, and

was less effective in abrogating G2 phase checkpoint suggesting that other cyclin A/CDK2

dependent mechanisms contributed to these roles of cyclin A/CDK2. In an attempt to

identify these other contributing factors another G2/M phase regulator known to be

regulated by cyclin A/CDK2, Cdh1 and its substrates Plk1 and Claspin were examined. Cdh1

levels were reduced in cyclin A/CDK2 depleted/inhibited cells although this had little effect

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on Plk1, a known Cdh1 substrate. However, the level of another substrate, Claspin, was

increased. Cdh1 depletion mimicked the effect of cyclin A depletion but to a weaker extent

and was sufficient at increasing Claspin levels similar to the increase caused by cyclin A

depletion. Co-depletion of cyclin A and Claspin blocked the accumulation of activated Chk1

normally seen with cyclin A depletion alone. However Claspin depletion alone did not

reduce the cyclin A/CDK2 dependent G2 delay but this is likely to be a result of inhibition of

S phase roles of Claspin.

Together, these data suggest that cyclin A/CDK2 regulates a number of different

mechanisms that contribute to G2/M phase progression. Here it has been demonstrated

that in normal G2/M progression and possibly to a lesser extent in G2 phase checkpoint

recovery, cyclin A/CDK2 regulates the level of Cdh1 which in turn affects at least one of its

substrates, Claspin, and consequently results in the increased level of activated Chk1

observed. However, the involvement of Cdh1 and Claspin alone does not explain the G2

phase delay observed with cyclin A/CDK2 depletion/inhibition. It is likely that other

mechanisms, possibly including cyclin A/CDK2 regulation of Wee1 and FoxM1, as reported

by others, combine with the mechanism described here to regulate normal G2/M phase

progression and G2 phase checkpoint recovery. These findings support the critical role for

cyclin A/CDK2 in regulating progression into mitosis and suggest that upstream regulators of

cyclin A/CDK2 activation will also be critical controllers of this cell cycle transition.

The pathways that work to co-ordinate cell cycle progression are very intricate and

deciphering these pathways, required for normal cell cycle progression, is key to

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understanding tumour development. By understanding cell cycle regulatory pathways it will

allow the identification of the pathway/s and their mechanism/s that become affected in

tumourgenesis. This will lead to the development of better targeted therapies, inferring

better efficacy with fewer side effects than commonly seen with the use of traditional

therapies, such as chemotherapy. Furthermore, this has the potential to positively impact

the development of personalised medicines and the customisation of healthcare.

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Publications

De Boer, L., Oakes, V., Beamish, H., Giles, N., Stevens, F., Somodevilla-Torres, M., DeSouza,

C., and Gabrielli, B (2008). "Cyclin A/cdk2 coordinates centrosomal and nuclear mitotic

events." Oncogene 27(31): 4261-4268.

Beamish, H., de Boer, L., Giles, N., Stevens, F., Oakes, V., Gabrielli B (2009). Cyclin A/cdk2

regulates adenomatous polyposis coli-dependent mitotic spindle anchoring. Journal of

Biological Chemistry, 284, 29015-23

Boutros, R., Lorenzo, C., Mondesert, O., Jauneau, A., Oakes, V., Dozier, C., Gabrielli, B.,

Ducommun, B (2011). CDC25B associates with a centrin 2-containing complex and is

involved in maintaining centrosome integrity. Biology of the Cell, 103, 55-68.

Oakes, V., Brooks, K., Edwards, B., Ranall, M., Leo, P., Pavey, S., Pinder, A., Beamish, H.,

Mukhopadhyay, P., Lambie, D and Gabrielli B (2012). “A potent Chk1 inhibitor is selectively

cytotoxic in melanomas with high levels of replicative stress.” Oncogene.

http://www.ncbi.nlm.nih.gov/pubmed/19703905




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List of Abbreviations

Ab Antibody

ACA anti-centromere antibody

Ala/A Alanine

APC/C Anaphase promoting complex/cyclosome

APS Ammonium Persulfate

ATM Ataxia telangiectasia mutated

ATP Adenosine 5'-triphosphate

ATR ataxia telangiectasia and Rad3-related protein

ATRIP ATR interacting protein

BRCA1 Breast cancer type 1 susceptibility protein

BSA Bovine Serum Albumin

Bub Budding inhibited by benomyl

BubR1 Budding inhibited by benomyl receptor 1

caff caffeine

CAK CDK activating kinase

CDK cyclin dependent kinase

CDK2i cyclin dependent kinase 2 inhibitor

Chk1 Checkpoint kinase 1

Chk1i Checkpoint kinase 1 inhibitor

Chk2 Checkpoint kinase 2

Cy3 indodicarbocyanine 3

Cy5 indodicarbocyanine 5

DAPI 4’-6-Diamidino-2-phenylindole

DDR DNA damage response

DI Diamantina Institute

DMEM Dulbecco’s Modified Eagle’s medium

DMP dimethyl pimelimidate

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DMSO dimethyl sulfoxide

DNA Deoxyribonucleic Acid

DNA-PK DNA-dependent protein kinase

DSB Double stranded break

dsDNA double stranded DNA

DTT Dithiothreitol

EDTA Ethylenediamine tetra-acetic acid

EGTA Ethylene glycol-tetra acetic acid

ERK Extracellular-signal-related kinase

FACS Fluorescence Activated Cell Sorting

FBS Foetal bovine serum

FoxM1 Forkhead protein M1

Gadd45 growth arrest and DNA damage inducible gene

G1 Gap 1

G2 Gap 2

GFP Green fluorescent protein

GST Glutathione-S transferase

h hour/s

HeLa Human cervical carcinoma cell line

H2O2 Hydrogen Peroxide

HRP Horse radish peroxidise

H3 Histone H3

HU Hydroxyurea

IgG Immunoglobulin G

IP Immunoprecipitate

IPTG Isopropyl β-D-1-thiogalactopyranoside

IR Ionising radiation

kDa kilo Dalton

LB Lysogeny broth

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M Mitosis

MAPK Mitogen-activated kinase

MeOH Methanol

MK-2 MAP kinase-activation protein kinase 2

MPM2 Mitotic Protein Marker 2

MRN Mre11, Rad50, Nbs1

MTOC Microtubule organising centre

NES Nuclear Export Signal

NLS Nuclear Localisation Signal

p38 MAPK p38 Mitogen-activated kinase

PBD Polo-box domain

PBS Phosphate buffered saline

PBST Phosphate buffered saline plus tween 20

pChk1 phosphorylated Chk1 Ser317

PCNA Proliferating cell nuclear antigen

PCR polymerase chain reaction

PI Propidium Iodide

PIKK Phosphatidylinositol 3-kinase related kinase

PFA Paraformaldehyde

Plk Polo like kinase

Plk1i Polo like kinase 1 inhibitor

PP2A Protein phosphatase 2A

PY15 CDK1 phosphorylation Tyrosine 15

QUT Queensland University of Technology

RPA Replication protein A

Rb Retinoblastoma protien

S Synthesis phase

SAC Spindle assembly checkpoint

SCF Skp, Cullin, F-box containing complex

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SDS Sodium dodecyl sulphate

SDS-PAGE Sodium dodecyl sulphate polyacrylamide gel electrophoresis

Ser/S Serine

siRNA short interfering RNA

SS Serum supreme

SSB Single stranded DNA break

ssDNA Single stranded DNA

TBE Tris-Borate-EDTA

TBS Tris buffered saline

TBST Tris buffered saline plus tween 20

TEMED N, N, N', N'-tetramethylethylenediamine

Thr/T Threonine

TKO MEFs Triple knock-out mouse embryonic fibroblasts

Tyr/Y Tyrosine

ULP Universal Lysis Buffer

UQ University of Queensland

USP ubiquitin-specific protease

UV(R) Ultraviolet (radiation)

WT Wildtype

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List of Figures

Figure 1.1 The eukaryotic cell cycle is made up of 4 separate phases. .................................................. 3

Figure 1.2 The mechanisms of the cell cycle – cellular processes and regulation of the cell cycle and

the different phases of mitosis. .............................................................................................................. 5

Figure 1.3 Cellular outcomes in response to DNA damage. ................................................................... 9

Figure 1.4 Illustration of the G2/M regulation of cyclin B/CDK1 activation. ........................................ 12

Figure 1.5 The basic structure of the DNA damage response (DDR) pathway..................................... 19

Figure 1.6 ATM-dependent response to DNA damage. ....................................................................... 23

Figure 1.7 ATR-dependent response to DNA damage. ........................................................................ 26

Figure 1.8 The ATR dependent DDR pathway is highly complex.......................................................... 33

Figure 3.1 pChk1 is present in asynchronously growing cells but strongly induced in response to DNA

damage. ................................................................................................................................................ 89

Figure 3.2 Active Chk1 is present during an unperturbed cell cycle. ................................................... 90

Figure 3.3 Chk1 siRNA effectively removes Chk1 and its typical functions. ......................................... 92

Figure 3.4 Cellomics data confirms the presence of pChk1 in an unperturbed cell cycle. .................. 95

Figure 3.5 Chk1 phosphorylation by ATR in G2 phase is required for timely mitotic entry. ................ 98

Figure 3.6 pChk1 is present at the centrosomes in G2 and Mitosis of an unperturbed cell cycle. .... 102

Figure 4.1 Cyclin A siRNA causes a G2 phase delay. ........................................................................... 117

Figure 4.2 CDK2i causes a G2 phase delay which was due to specific CDK2 inhibition. .................... 121

Figure 4.3 Cyclin A siRNA and CDK2i disconnect the coordinated activation of centrosomal and

nuclear pools of cyclin B/CDK1. .......................................................................................................... 125

Figure 4.4 Cyclin A/CDK2 is required for coordinated cyclin B/CDK1 activity. ................................... 130

Figure 4.5 Cyclin A/CDK2 localises to the centrosomes in late G2 phase. ......................................... 133

Figure 4.6 Cyclin A/CDK2 is required for exit from a G2 phase checkpoint arrest. .......................... 136

Figure 4.7 Cyclin A/CDK2 is required for exit from a G2 phase checkpoint arrest. ............................ 139

Figure 5.1 Chk1 depletion causes reversal of the cyclin A dependent G2 phase delay. .................... 151

Figure 5.2 CDK2 inhibition confirms a role for pChk1 Ser317 in the cyclin A / CDK2 dependent G2

delay. .................................................................................................................................................. 157

Figure 5.3 Chk2 does not have a role in the cyclin A dependent G2 phase delay. ............................ 159

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Figure 5.4 ATR is required to phosphorylate Chk1 to induce the cyclin A dependent G2 phase delay.

........................................................................................................................................................... 163

Figure 5.5 Cyclin A siRNA does not cause DNA damage. ................................................................... 166

Figure 6.1 Cyclin A and Chk1 co-localise at the centrosomes. ........................................................... 176

Figure 6.2 Cyclin A and Chk1 co-immunoprecipitate ......................................................................... 178

Figure 6.3 Cyclin A/CDK2 phosphorylates Chk1 in vivo ..................................................................... 182

Figure 6.4 Chk1 SS317,345DD, with S286A and S301A mutations do not delay in G2 phase ........... 183

Figure 6.5 Chk1 Ser286 and Ser301 Alanine mutants cause a delay of mitotic entry. ...................... 186

Figure 6.6 Cherry-Chk1 constructs have impaired Ser317 and Ser345 phosphorylation. ................. 190

Figure 6.7 Flag Chk1 constructs are phosphorylated on Ser317 and delay mitotic entry. ................ 192

Figure 6.8 Chk1 phosphorylation on Ser286 Ser301........................................................................ 1964

Figure 6.9 SS286,301AA mutation does not affect Chk1 localisation ................................................ 201

Figure 7.1 Plk1 inhibition causes phenotypes similar to cyclin A/CDK2 depletion/inhibition. .......... 215

Figure 7.2 Plk1 overexpression did not contribute to delayed mitotic entry caused by cyclin A siRNA

treatment. .......................................................................................................................................... 218

Figure 7.3 Cyclin A affects Plk1 localisation at the centrosomes and kinetochores in mitosis. ...... 2221

Figure 7.4 Cdh1 levels are reduced with cyclin A/CDK2 depletion/inhibition. ................................ 2243

Figure 7.5 Cdh1 overexpression does not rescue the cyclin A/CDK2 G2 phase delay and itself inhibits

mitotic entry. ................................................................................................................................... 2287

Figure 7.6 Cdh1 siRNA does not rescue the cyclin A/CDK2 G2 phase delay and delays mitotic

progression also. .............................................................................................................................. 2343

Figure 7.7 Claspin is increased with cyclin A depletion. .................................................................... 235

Figure 7.8 Claspin siRNA can reduced pChk1 levels during the cyclin A/CDK2 dependent G2 delay but

cannot rescue the delay. .................................................................................................................... 238

Figure 8.1 Model of Chk1 regulation during G2/M progression. ....................................................... 251

Figure 8.2 Claspin involvement and regulation during G2/M............................................................ 254

Figure 8.3 A simplified schematic of G2 regulation involving cyclin A/CDK2. ................................... 263

Figure 10.1 Cellomics data confirms the presence of pChk1 in an unperturbed cell cycle. .............. 320

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List of Tables

Table 1.1 Cyclin/CDKs complex are activated within specific phases of the cell cycle. ....................... 10

Table 2.1 Antibodies List. ..................................................................................................................... 57

Table 2.2 List of siRNA. ......................................................................................................................... 62

Table 2.3 List of Plasmids ..................................................................................................................... 69

Table 2.4 List of Chk1 mutagenesis primers used. ............................................................................... 70

Table 2.5 List of sequencing primers used. .......................................................................................... 74

Table 2.6 PBD-GFP cloning primers ...................................................................................................... 76

Table 2.7 Components and Volumes used for SDS-PAGE gels ............................................................. 79

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Chapter 1

Summary of the literature

Chapter 1 – Literature Review

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1.1 The cell cycle, structure and function.

An active cell division cycle consists of four stages. When quiescent, G0 cells commit to re-

enter the cell cycle they will enter the first gap phase of the cell cycle known as G1. In G1,

cell growth is initiated which involves the synthesis of organelles requiring both structural

proteins and enzymes. G1 also, prepares the cell for DNA replication in the subsequent

phase known as S phase or the synthesis phase. After duplication of the genome, cells will

grow further during G2 phase. Also during G2 phase, the cell will ensure that DNA

replication has occurred with complete fidelity and that all DNA damage is repaired prior to

mitotic entry. It is essential that G2 processes occur correctly as this is the penultimate

phase before cells enter mitosis. Mitosis is the most complex stage of the cell cycle and

involves the even segregation of the replicated chromosomes and the production of two

independent daughter cells (Alberts et al., 1994, Doree and Hunt, 2002, Tyson et al., 2002).

Mitosis itself is comprised of several stages that will be discussed in more detail in section

1.1.1. As Figure 1.1 depicts, the cell cycle may be a continuous process. Once cells complete

mitosis the daughter cells are capable of undergoing their own cellular divisions by re-

entering G1 or they may stop dividing and enter G0.

1.1.1 Stages of mitosis

There are distinct stages of mitosis; prophase, prometaphase, metaphase, anaphase,

telophase and cytokinesis (Figure 1.2). During prophase the chromosomes condense, the


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Figure 1.1 The eukaryotic cell cycle is made up of 4 separate phases.

G1 (gap 1), S (synthesis) and G2 (gap 2) phases are collectively referred to as interphase, and

completion of the cell cycle occurs with M phase (mitosis) (Alberts et al. 1994; Doree and Hunt,

2002; Tyson et al. 2002). When not dividing, cells are considered to be in G0.


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nuclear envelope breakdown (NEB) occurs and the centrosomes migrate to opposing

positions across the nucleus. The centrosome is the microtubule organising centre (MTOC)

and prior to mitosis it duplicates to facilitate the formation of a bipolar mitotic spindle

required for mitosis (See Figure 1.2). In addition the centrosomes also nucleate astral

microtubules which radiate away from the chromosomes and are required for spindle

attachment to the cell cortex, spindle orientation and positioning, and the fidelity of cell

division (Strauss et al., 2006). The duplication of the centrosomes occurs during S phase and

is regulated by cell cycle regulators including, cyclin E and CDK2 (Sterns et al., 2005).

Regulation of centrosome duplication is important as aberrant numbers of centrosomes

have been associated with increased genomic instability and cancer (Brinkley, 2001, Lingle

and Salisbury, 2000, Wang et al., 2004).

The centrosome initiated microtubules that form the spindle are captured by the

kinetochores which are associated with centromeric regions of each chromosome prior to

metaphase (prometaphase). During metaphase microtubule attachment to each

kinetochore is achieved and the tension across the kinetochore aligns the chromosomes to

the midline of the cell. Once in anaphase, sister chromatids move actively toward the

opposite centrosomes or ‘poles’ of the mitotic spindle. During telophase the chromosomes

begin to de-condense and nuclear envelopes form around the daughter chromosomes

located at opposite poles. The final stage of mitosis is cytokinesis which involves the final

separation of the daughter cells by the contracting and pinching of an actinomyosin-based

ring (Alberts et al., 1994, Nigg, 2001, Pines and Rieder, 2001) (Figure 1.2). The actinomyosin


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Figure 1.2 Mechanisms of the cell cycle - cellular processes and regulation of the cell cycle and the

different phases of mitosis.

At the top of the figure cyclin/CDKs are placed above the particular stages of the cell cycle for which

they are required. The cell schematics illustrate the structural changes that occur within particular

stages of the cell cycle mitosis. The grey object indicates the centrosomes and the lines protruding

from the centrosomes are representative of microtubules (2n = a single genome, 4n = duplicated

genome). The red arrows indicate the checkpoints of the cell cycle.


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ring is composed of several important proteins that are required to assist with the pinching

action between the two daughter cells. Each stage of mitosis can be visualised with the aid

of Figure 1.2.

1.2 Cancer and the cell cycle

The importance of the cell cycle was evident to early cell biologists and is made clear from

quotes such as that by Schleiden & Schwann in 1838: ‘Cell division is the only path to

immortality. Non dividing cells can live for as long as a hundred years, but they always

eventually die.’ (The Cell Cycle: An Introduction (1993) Murray, A & Hunt, T (W.H. Freeman

& Co., New York)). This statement perhaps overstates the case somewhat, but for growth

and homeostasis of all organisms there must be cell division. However, a hazard of cell

division is the possibility of incurring genetic or epigenetic errors. The worst outcome of

these errors is uncontrolled cell growth and division which is a feature associated with

tumour formation.

Cancer formation is associated with genes that promote or suppress tumour growth -

known as oncogenes or tumour suppressors, respectively. Oncogenes and tumour

suppressors often have one or more cell cycle function/s. Although the overall structure of

the cell cycle is well understood, the many pathways that contribute to its regulation are

less understood. The initiation and progression of cancer is a culmination of multiple

defects within the cell cycle. Defining a detailed understanding of the pathways that

contribute towards the normal regulation of cell cycle progression is essential to understand


7

how defects in these pathways contribute to cancer. This level of detail in relation to the

cell cycle is also necessary to identify potential targets, for the treatment of cancer/s, that

are characteristic of a particular defect.

The remainder of this chapter will detail the regulatory pathways critical to the research

covered in this thesis. Further in depth information will be supplied about G2/M

progression, in particular the processes and regulation required for G2/M progression in

both normal and in DNA damage situations will be discussed.

1.3 Regulation of cell cycle

The correct progression of cells through the phases of the cell cycle is controlled by the

ordered expression and activation of the cyclin dependent kinase (CDK) complexes. The

activation of cyclin/CDK complexes provides the forward impetus through the cell cycle,

whereas their inhibition can implement a ‘checkpoint’ which becomes the braking

mechanism of the cell cycle.

DNA damage activates a complex response network that incorporates cell cycle arrest, DNA

repair and possibly apoptosis if the damage cannot be repaired (Khanna and Jackson, 2001,

Niida and Nakanishi, 2006, Medema and Macurek, 2011). To repair damage to the cell,

regulatory pathways are triggered that allow the cell time for repair. This stoppage of the

cell cycle to allow repair of DNA damage is referred to as a checkpoint. In cases where

damage cannot be repaired and apoptotic signalling fails the cell will continue uncontrolled

through cell division leading to genomic mutation/s (Zhou and Elledge, 2000) (Hyka-


8

Nouspikel et al., 2012). Uncontrolled cellular division is a common characteristic of most

cancers and may contribute to the initiation and/or progression of tumourgenesis (Figure

1.3). Damaged DNA can be caused spontaneously by endogenous defects in S phase or

mitosis or indirectly by exogenous external stimuli such as exposure to ultraviolet radiation

(UV(R)) or ionising radiation (IR). The protein pathways that comprise these responses to

DNA damage are critical to ensure the production of identical progeny. Therefore the loss of

normal cell cycle controls such as checkpoints is a ‘hallmark’ of many human cancers

(Piwnica-Worms, 1999) and understanding the regulation of the cell cycle is critical for

understanding the formation of cancer.

1.3.1 The cyclin/CDK complexes

The cell cycle is regulated at the most basic level by the ordered activation of the CDK family

of Ser/Thr kinases (Morgan, 1995). As the name implies, the activity of these kinases is

dependent on their association with their regulatory subunit known as a cyclin. There are

approximately 30 cyclins (Okamoto and Beach, 1994, Rickert et al., 1996, Peng et al., 1998)

and 21 CDK genes but only 11 cyclins and 5 CDKs have known cell cycle functions (Rickert et

al., 1996). The major cyclins required for cell cycle progression are typically expressed

periodically in unique phases of the cell cycle, whereas CDKs remain at constant levels

throughout the cell cycle. Except for CDK1 which is a typical E2F-target gene and in some

circumstances E2F regulation may cause a reduction in CDK1. When complexed with its partner

CDK, the cyclins determine the complexes temporal and spatial activity as well as substrate


9

Figure 1.3 Cellular outcomes in response to DNA damage.

In all circumstances apoptosis may be initiated but is dependent on the cell, the mutations and

pathways affected. Normal cells will eventually initiate cell death once growth can no longer be

sustained.


10

specificity (Evans et al., 1983). The cyclin/CDK complexes required for cell cycle progression

are listed and shown in Table 1.1 and Figure 1.2, respectively.

Table 1.1 Cyclin/CDKs complex are activated within specific phases of the cell cycle.

CDK cyclin cell cycle phase when active

CDK4 cyclin D1, D2, D3 G1 phase

CDK6 cyclin D1, D2, D3 G1 phase

CDK2 cyclin E G1/S transition

CDK2 cyclin A S phase and G2 phase

CDK1 (cdc2) cyclin A G2/M transition

CDK1 (cdc2) cyclin B Mitosis

The predominant complexes required for G2 and M phases are cyclin A/CDK2 and cyclin

B/CDK1 respectively. CDK2 is required for two different phases of the cell cycle. Specifically,

CDK2 binds and functions with cyclin A during S and G2 phase (Table 1.1, Figure 1). The role

of cyclin A/CDK2 in normal G2 progression has been controversial, as investigations using

Cdk2 knockdown and Cdk2 knockout mice failed to reveal any defects in cell cycle

progression (Berthet et al., 2003, Ortega et al., 2003, Tetsu and McCormick, 2003). This was

later demonstrated to be caused by the functional redundancy with the highly homologous

CDK1 (Kaldis and Aleem, 2005, Malumbres and Barbacid, 2009, Satyanarayana & Kaldis,

2009). Conversely, there is evidence that under certain circumstances, i.e. embryonic

development, Cdk2 cannot compensate for Cdk1 (Satyanarayana, et al., 2008). However,

cyclin A/cdk does not appear to be the sole regulator of G2 phase as cyclin A can complex

with CDK1 in late G2/M mitosis and although its role is not well defined, it is thought to be

important for G2/M transition. Lastly, cyclin B complexed with CDK1 is the primary CDK

complex required for mitotic progression (King et al., 1994, Arellano and Moreno, 1997).


11

Inactivation of cyclin B/CDK1 is also required for mitotic exit and this is achieved by the

degradation of the cyclin B subunit which commences in anaphase with full degradation

completed by early G1 phase.

Not only are the cyclin/CDK complexes regulated by subunit interaction, their

phosphorylation/dephosphorylation adds another level of regulation ensuring tight control

of transition/s between cell cycle phases (Figure 1.4). The regulation of cyclin/CDK

complexes is controlled by reversible inhibitory phosphorylations and activating

phosphorylation. For example, CDK1 and CDK2 are inhibited by phosphorylation on Tyrosine

15 (also known as Tyr15 or PY15) and Threonine 14 (Thr14), and there is an additional

activating phosphorylation site at Thr160 on CDK2 and Thr161 on CDK1. These activating

sites are phosphorylated by CAK (CDK activating kinase) (Figure 1.4) while the inhibitory

phosphorylation sites, Tyr15 and Thr14, are catalysed by the kinases Wee1 and Myk1

(Parker and Piwnica-Worms, 1992b, Mueller et al., 1995, Liu et al., 1997, Rothblum-Oviatt et

al., 2001, Stumpff et al., 2004). The dephosphorylation of the inhibitory phosphorylation

sites, and consequent activation of cyclin/CDKs, is performed by the dual-specificity Cdc25

phosphatases. Wee1/Myk1 and Cdc25 are therefore key upstream regulators of cyclin/CDKs

that act antagonistically to regulate their activation.

The location of cyclin/CDK complexes also contributes to their regulation. Although the

cyclin/CDK complexes are predominantly activated within the nucleus, cyclin B has been

found to contain a nuclear export signal (Moore et al., 1999) which localises it to the

cytoplasm until the beginning of prophase (Porter and Donoghue, 2003). This allows for the


12

Figure 1.4 Illustration of the G2/M regulation of cyclin B/CDK1 activation.

Cyclin E/CDK2 and cyclin A/CDK2 are regulated by a similar mechanism at the G1/S and early S/G2

phase transitions, respectively. Wee1 phosphorylates Tyr15 (Y15) and Wee1-like kinase Myk1

phosphorylates Thr14 (T14). Both T14 and T15 are inhibitory phosphorylation sites, as indicated by

the red colouring. Cdc25 phosphatases are responsible for the removal of these inhibitory

phosphorylations. CAK catalyses the activating phosphorylation Thr161 (T161), shown in green.


13

initial activation of cyclin B/CDK1 to occur in the cytoplasm and controls the timing for

cyclin B/CDK1 activation in the nucleus. Therefore understanding the localised activity of

cyclin/CDK complexes will assist in understanding its role during the cell cycle. CDK activity

can be counteracted by inhibitory proteins called CDK inhibitors (CKI). There are two distinct

families of CKIs: the INK4 family and the Cip/Kip family (Sherr and Roberts, 1995). The INK4

family specifically inactivates CDK4 and CDK6 (Carnero and Hannon, 1998), while the

Cip/Kip family, is more promiscuous and can inhibit all cyclin/CDKs in vitro, but in vivo

inhibits CDK2 and in some cases CDK1 complexes only (Polyak et al., 1994, Harper et al.,

1995, Lee et al., 1995, Hengst and Reed, 1998).

1.3.2 Cell cycle checkpoints

As indicated in Figure 1.2, there are several checkpoints throughout the cell cycle. The

common goal of the checkpoint is to co-ordinate a global response to repair and maintain

genome integrity (Hartwell and Weinert, 1989, Zhou and Elledge, 2000, Harrison and Haber,

2006).

Different types of DNA damage influences the pathways that lead to a checkpoint arrest.

For example DNA damage that results in single stranded breaks (SSBs) caused by external

stimuli such as ultraviolet radiation cause the activation of ATR (Ataxia Telangiectasia and

Rad3 related) dependent pathways. Other types of DNA damage may result in double

stranded breaks (DSBs). The ATM (Ataxia Telangiectasia Mutated) protein kinase activity is

predominantly stimulated by DSBs in DNA (Chan et al., 2000), such as those caused by


14

ionising radiation (IR) (Rotman and Shiloh, 1999). p53 activity is also very critical for

checkpoint arrest to allow DNA repair and/or apoptosis if the damage cannot be repaired

(Owen-Schaub et al., 1995, Polyak et al., 1997, Gottlieb and Oren, 1998). p53 can act

dependently or independently of ATM/ATR. The checkpoints and their pathways required to

enforce the checkpoint arrest are discussed in detail in the following sections.

1.3.2.1 G1/S phase checkpoint

The G1/S checkpoint is dependent on the tumour suppressor gene, p53, which protects

cells from undergoing tumourigenic transformation (Vogelstein et al., 2000). The rapid

induction of p53 protein levels in G1 phase in turn up-regulates the CKI, p21 in response to

some forms of DNA damage (el-Deiry et al., 1993, Levine, 1997) which binds to and inhibits

G1 phase CDKs (Gu et al., 1993, Xiong et al., 1993, Harper et al., 1995). The G1 arrest

thereby prevents the replication of damaged DNA in S phase (Ko and Prives, 1996). ATM

and ATR are involved in sensing DNA damage and can activate p53 via phosphorylation

during the G1/S checkpoint (el-Deiry et al., 1993).

1.3.2.2 S phase checkpoint

The S phase checkpoint ensures the fidelity of S phase, as it has the ability to stop or at least

slow the rate of DNA synthesis (Paulovich and Hartwell, 1995, Weinert et al., 2009, Zhou

and Elledge, 2000). The checkpoint involves either ATM or ATR activity depending on the S

phase effect. Errors in S phase commonly result in exposed single stranded DNA (ssDNA),

which then becomes coated in replication protein A (RPA). RPA localised to ssDNA is


15

believed to be the ‘red flag’ that initiates the checkpoint response by binding of ATR, which

subsequently leads to the stabilisation of replication forks (Katou et al., 2003, Cortez et al.,

2004, Yoo et al., 2004, Cimprich and Cortez, 2008). In situations where ATR is defective,

SSBs can be trimmed to produce DSBs, allowing ATM signalling. Use of these pathways

during the S phase checkpoint (Boddy et al., 1998, Lindsay et al., 1998, Martinho et al.,

1998, Zeng et al., 1998, Brondello et al., 1999), acts as a further safety mechanism to ensure

a checkpoint response that guarantees cell survival (Segurado and Tercero, 2009).

1.3.2.3 G2 Phase DNA damage checkpoint

The importance of the G2 checkpoint was determined by a study using primary mouse

embryonic fibroblasts (TKO MEFs) that were manipulated to remove all three

Retinoblastoma (Rb) family members and lack the G1/S checkpoint. These TKO MEFs were

able to suppress apoptosis but did not allow unconstrained cell division, that is

characteristic of cancer cells, because they arrested in G2 phase (Foijer et al., 2005).

The G2 phase checkpoint also known as the G2 phase DNA damage checkpoint has both p53

dependent and independent mechanisms (Bache et al., 1999, Chang and Eastman, 2012).

The p53-dependent arrest involves the maintenance of a long term arrest (in terms of days),

whereas p53-independent mechanisms initiate an immediate arrest that can be stable for a

matter of hours (Bunz et al., 1998). Both mechanisms target the regulation of the cyclin

B/CDK1 complex, required for M phase progression (Table 1), however the mechanisms by

which these are imposed differ. p53-independent mechanisms involve the inhibition of


16

Cdc25 phosphatases, whereas the dissociation of cyclin B from CDK1 leading to cyclin

B/CDK1 inactivation is caused by p53-dependent induction of Gadd45 (growth arrest and

DNA damage inducible gene) (Hermeking et al., 1997, Taylor and Stark, 2001). This is one of

several ways that p53 can block mitotic entry during a G2 checkpoint.

p53-independent mechanisms involve the apical kinases, ATR/ATM, which through their

downstream targets and subsequent phosphorylation cascades leads to cyclin B/CDK1

inactivation. ATM/ATR initiates a more rapid checkpoint response to DNA damage than p53,

however if a prolonged arrest is required p53 dependent pathways will predominate. This

may be facilitated by ATM phosphorylation of the p53 inhibitor, MDM2, as well as ATM/ATR

phosphorylation of p53 itself on Ser15 to increase p53 stability (Canman et al., 1998,

Khosravi et al., 1999, Tibbetts et al., 1999, Maya et al., 2001,).

The pathways activated during the G2 checkpoint are dependent on the extent of damage.

The rapid response facilitated by ATM/ATR will be discussed in more detail, in section 1.5.

1.3.2.4 Spindle Assembly checkpoint

The mitotic checkpoint, better known as the spindle assembly checkpoint (SAC), has the

ability to stop the onset of anaphase in cells with mitotic spindle defects. For example, the

improper spindle alignment or incorrect tension of the chromosomes before division will

delay separation until the defect has been rectified (Kops et al., 2005). If a lack of tension

between the chromosomes and spindle is detected by the SAC, caused by either unattached

kinetochores or those not attached correctly, the checkpoint is activated by the inhibition of


17

the anaphase promoting complex/cyclosome (APC/C; hereafter known as APC) cofactor,

Cdc20. The inhibition of Cdc20 causes the stabilisation of key proteins, such as securins that

are normally ubiquitinated by APC associated with Cdc20 (APCCdc20) to allow sister

chromatid separation (Holloway et al., 1993, Cohen-Fix et al., 1996, Zou et al., 1999).

Bub1-3 (Budding inhibited by benomyl) and Mad1-3 (Mitotic arrest deficient), are two of

the first protein families identified to be involved in and activated by mitotic spindle

defect/s (Fang et al., 1998a, Amon, 1999). More recently, many more proteins have been

discovered to have roles during the SAC such as Aurora B. Aurora B is necessary for BubR1

(Bub Receptor 1) phosphorylation and recruitment to the kinetochores (Ditchfield et al.,

2003, Lens et al., 2003), which is subsequently required for Bub1 involvement in the SAC

(Carvalho et al., 2003, Ditchfield et al., 2003, Lampson and Kapoor, 2005, Mistry et al.,

2008). Unsurprisingly, inhibition of Aurora B overrides the SAC, affecting chromosome

segregation (Hauf et al., 2003, Santaguida et al., 2011) and also results in the inability of

Aurora B to phosphorylate histone H3 on Serine 10 (Wei et al., 1999), a common marker of

mitosis and chromosome condensation (Hendzel et al., 1997, Goto et al., 1999, Preuss et al.,

2003).

1.4 Exploiting the DNA damage checkpoints, a cancer therapy

Many conventional chemotherapeutic agents induce DNA damage and thus target cell cycle

checkpoints (Walton et al., 2010). A high proportion of tumours are deficient in the tumour

suppressor gene p53, and are thus deficient in the G1 phase checkpoint and reliant on the


18

p53-independent G2/M DNA damage checkpoint pathways (Chen et al., 2012) as a

protective response to DNA damage. Tumour reliance on the G2/M checkpoint can be used

as a selective target to distinguish between normal and cancer cells. The effectiveness of G2

phase checkpoint abrogation came from the early work of Lau and Pardee in 1982: cells

arrested in G2 phase and then treated with caffeine, which causes G2/M checkpoint

abrogation, resulted in premature and lethal mitosis (Lau and Pardee, 1982). This lethality

was caused by the inability of the cells to repair the damage before entering mitosis. Since

then numerous inhibitors that target the essential G2/M checkpoint kinases have proven to

be very effective inducers of tumour regression (Zhao et al., 2002b, Sorensen et al., 2003,

Xiao et al., 2003). There are several inhibitors for checkpoint proteins that have been used

in various phase clinical trials including, Chk1, p53, Plk1 (Mross et al., 2008, Chen et al.,

2009, Strebhardt, 2010, Carrassa and Damia, 2011, Cheok et al., 2011, Bennett et al., 2012).

The specifics of these G2/M proteins will be discussed in detail in the chapter.

1.5 DNA damage pathways

DNA lesions, either SSB or DSB, act as the initial signal to activate the appropriate DNA

damage pathway. Several layers of interlinked signalling pathways encompass DNA damage

detection and response pathway/s and the intricate co-ordination of repair processes is

becoming increasingly apparent (Thompson, 2012). These layers are known as the apical

transducers, transducers and effectors (Figure 1.5). The most common proteins within these

layers are the apical transducers ATM and ATR, with signals mediated by the transducers

Chk1 and Chk2. Chk1 and Chk2 can promote cell cycle arrest, damage repair and apoptotic


19

Figure 1.5 The basic structure of the DNA damage response (DDR) pathway.

The basic DDR pathway includes different downstream layers consisting of Apical Transducers,

Transducers and Effectors. Additional, ‘mediator proteins’ may also be involved in the DDR pathway.

The DDR pathway utilised is determined from the type of damage.


20

signalling responses enforced by regulating a number of effector kinases (Niida et al., 2010,

Kasahara et al., 2010, Smith et al., 2010), such as Cdc25 and p53. There are additional

proteins which are highly involved and required to assist ATM/ATR signalling and to arrest

cells in G2 phase such as Claspin and BRCA1 (Breast Cancer 1) (Yarden et al., 2002). There

are also independent pathways such as the p38MAPK signalling network that was recently

shown to delay G2/M progression via a non-canonical ERK (Extracellular Regulated Kinase)

pathway (Astuti et al., 2009). The common feature of these pathways is that inhibition of

the cyclin/CDK complexes causes cell cycle arrest. The mechanism for the cell cycle arrest

will be addressed in the following sections, with specific focus on the ATR dependent

pathway.

1.5.1 ATM/ATR

ATM and ATR are members of the PIKK (phosphoinositide 3-kinases) family of kinases

(Khanna and Jackson, 2001) of which there are three family members. The third member of

the PIKK family is DNA-PK (DNA-dependent protein kinase) which is also a DNA double-

strand break repair enzyme. The ATM/ATR response depends on the type of damage: ATM

responds to DSBs while ATR responds to SSBs. ATM and ATR were initially thought to act via

independent pathways, based on their different substrates (Kim et al., 1999). ATM

predominantly phosphorylates Chk2, while ATR phosphorylates Chk1. There is now

evidence that these two pathways overlap. For example, activation of ATR can occur in

response to DSBs caused by IR (Gatei et al., 2003, Jazayeri et al., 2006) and ATR-dependent

phosphorylation and activation of ATM can occur in response to SSBs caused by UV(R) or


21

replication fork stalling (Stiff et al., 2006). The overlap is supported by the phosphorylation

status of several common ATM/ATR substrates, such as p53 (Khanna et al., 1998, Tibbetts et

al., 1999) and Chk1 (Gatei et al., 2003, Ozeri-Galai et al., 2008) in response to various forms

of DNA damage. The extent and significance of ATM/ATR signalling overlap is the subject of

controversy and ongoing research.

ATM

ATM is not considered an essential gene as knockout mice are viable (Beamish et al., 1996,

Shiloh, 2001). Notably, ATM knockouts are more sensitive to DNA damage (Meyn, 1995,

Lavin and Shiloh, 1997, Shiloh, 1997, Takao et al., 1999, Worgul et al., 2002, Worgul et al.,

2005) and are defective in radiation induced G1/S, S and G2/M checkpoint control (Painter

and Young, 1980, Paules et al., 1995). ATM is present as an auto-inhibitory homodimer and

upon activation it auto-phosphorylates on Ser1981, dissociating to form active monomers

(Bakkenist and Kastan, 2003). When active, ATM preferentially binds to DNA termini (Smith

et al., 1999) so that a rapid response to DNA damage may occur (Bakkenist and Kastan,

2003).

The MRN complex (Mre11, Rad50 and Nbs1 (Nijmegen Breakage syndrome 1)), is highly

conserved and is required for ATM activation during DNA damage repair (Lee and Paull,

2004, Dupre et al., 2006). In response to DSBs the MRN complex has an initial role in

processing double strand breaks due to its DNA nuclease and binding capabilities (Petrini,

2000). The requirement for MRN activity has been demonstrated by hypomorphs

(mutations that cause reduced function of the gene product) of Mre11 and Nbs1 which


22

result in AT (ataxia-telangiectasia)-like disorders (Shiloh, 1997, Petrini, 2000, Tauchi et al.,

2002b, Costanzo et al., 2004, O'Driscoll et al., 2004). Once active, ATM phosphorylates

downstream substrates such as BRCA1, Chk2 (checkpoint kinase 2), p53, H2AX (H2A histone

family), and Nbs1 ( Gatei et al., 2000, Lee and Paull, 2004). The ATM dependent checkpoint

response is outlined in Figure 1.6.

ATR

ATR is a 303kDa (kilo Daltons) protein first identified due to its homology with ATM and

Saccharomyces pombe (sp) gene Rad3 (Cimprich et al., 1996). Unlike ATM, ATR depletion is

embryonic lethal (Brown and Baltimore, 2000), and is therefore considered essential for

cellular function under normal or stressed conditions. Expression of a kinase-inactive allele

of ATR in human fibroblasts or overexpression of a kinase inactive mutant of ATR induces

cellular sensitivity to DNA damaging agents and causes deficiencies in cell cycle checkpoints

(Cliby et al., 1998, Wright et al., 1998, Helt et al., 2005). Additionally, defective splicing of

ATR mRNA in the human autosomal recessive disorder, Seckel Syndrome (O'Driscoll et al.,

2003), results in microcephaly, growth retardation, dwarfism, mental retardation and

dysmorphic facial features, and cell cycle checkpoint defects (Tauchi et al., 2002a, Tauchi et

al., 2002b, O'Driscoll et al., 2003, Frappart et al., 2005).

ATR is activated in response to SSBs and is recruited to sites of DNA lesions. ATR interacts

with specific DNA structures (Byun et al., 2005, Cortez, 2005) by binding RPA (Itakura et al.,

2004), which then leads to ATR phosphorylation of SQ/TQ sites on its targets (Figure 1.7).

Regulation of ATR kinase activity is poorly understood, however its interaction with ATRIP


23

Figure 1.6 ATM-dependent response to DNA damage.

MRN is required for ATM activation in response to DNA damage and the subsequent activation of its

downstream targets.


24

(ATR-interacting partner) appears to regulate ATR expression and affects DNA damage

signalling. Interestingly, ATR can regulate expression of ATRIP as observed by depleting cells

of these proteins using siRNAs (Cortez et al., 2001). Coincidentally, loss of either ATR or

ATRIP also results in similar defects within the cell, notably a loss of the G2/M response to

damage (Cortez et al., 2001), and demonstrates the mutually dependent relationship

between ATR and ATRIP. ATR binding to single stranded DNA via RPA may also be

influenced by ATRIP (Zou and Elledge, 2003, Wu et al., 2005, Namiki and Zou, 2006)(Figure

1.7). In response to DNA damage, ATR-ATRIP co-localise at DNA damage foci in an RPA-

dependent manner (Itakura et al., 2004). The exact role of RPA in ATR-ssDNA binding is still

unclear, as RPA was shown to be dispensable for ATR phosphorylation of Chk1 in response

to damage (Unsal-Kacmaz and Sancar, 2004, Ball et al., 2005).

ATR, as well as its downstream target Chk1, appears to have normal G2 phase roles and

although these roles are not fully understood, it is likely to resemble the checkpoint

functions, as shown in Figure 1.7.

1.5.2 Chk1 and Chk2

Checkpoint kinases 1 and 2 were identified as respective downstream targets of ATR and

ATM (Walworth et al., 1993, Murakami and Okayama, 1995). These conserved kinases have

been identified in yeast and humans (Blasina et al., 1999, Tominaga et al., 1999). The

importance of Chk1 for cellular function is demonstrated by the death of Chk1 knockout

mice at early embryonic stages of development. These embryos have abnormal nuclei


25

within the core of the blastocyst which reflects defective cell growth and cell checkpoints

(Liu et al., 2000, Takai et al., 2000, Zachos et al., 2003).

In yeast and mammals, ATR will phosphorylate Chk1 on Ser317 and Ser345 in response to

damage (Liu et al., 2000, Lopez-Girona et al., 2001c, Zhao and Piwnica-Worms, 2001).

Phosphorylation of Chk1 on Ser317 and Ser345 are activating phosphorylations that from

low levels, present in basal conditions, (Kaneko et al., 1999, Zhao et al., 2002a, Sorensen et

al., 2003, Tapia-Alveal et al., 2009) increase rapidly in response to DNA damage (Tapia-

Alveal and O’Connell, 2011).

Once active, Chk1 is capable of targeting Cdc25 family members for phosphorylation. Chk1

phosphorylates and inhibits the Cdc25, therefore rendering the phosphatases incapable of

activating cyclin/CDK complexes (Furnari et al., 1997, Uto et al., 2004). There are several

Chk1-dependent phosphorylation sites that are critical for Cdc25 inhibition that are

catalysed not only during the G2 checkpoint but also throughout other phases of the cell

cycle. Regulation of Cdc25 will be discussed in more detail, in section 1.5.4.

Unlike Chk1, Chk2 is not essential for pre-natal development (Hirao et al., 2000, Takai et al.,

2002). Chk2 is specifically shown to be phosphorylated by ATM in response to IR on

threonine 68 (Thr68) (Matsuoka et al., 2000, Melchionna et al., 2000), implicating Chk2 in

the DSB DNA damage response pathway. Although Chk2 has a specific role in the ATM

dependent DNA damage pathway, Chk2 deficient mice revealed that its main function is in

the p53 apoptotic pathway and not the G2 DNA damage checkpoint (Hirao et al., 2002,

Takai et al., 2002). Collectively, the data suggests that the role of Chk2 during the


26

Figure 1.7 ATR-dependent response to DNA damage.

ATR binds to the single stranded binding protein, RPA, which is facilitated by ATRIP. ATR activates

Chk1 which ultimately leads to a block of mitotic cyclin/CDK activation and entry into mitosis.


27

checkpoint is less essential than that of Chk1. Further in depth discussion regarding the role

of Chk1 will be discussed later in section 1.8.

1.5.3 p53

In response to DNA damage, p53 is phosphorylated on Ser15 and Ser20 (Chehab et al.,

2000). Phosphorylation of Ser15 is mediated by ATM or ATR and promotes its role in the

apoptotic pathway (Chehab et al., 2000). Phosphorylation of Ser15 stabilises p53 by

blocking the interaction with its negative regulatory partner Mdm2 (Maya et al., 2001).

Ser20 phosphorylation is mediated by Chk2 (Chehab et al., 2000, Tian et al., 2002), and has

also been linked to p53 stabilisation (Chehab et al., 1999).

As mentioned, p53 has functions in the G1/S checkpoint where it facilitates p21 binding and

inhibition of cyclin E/CDK2 (Harper et al., 1993, el-Deiry et al., 1993). Although the G1 arrest

is just as defective in p21-/- cells as it is in p53 -/- cells, in response to IR the arrest can only

be partially maintained. This suggests there are other p53-dependent targets that can

promote G1 phase arrest (Deng et al., 1995). However, it may be the involvement of other

p53-independent pathways that promote arrest at the G1/S boundary such as; Chk2

phosphorylation and destabilisation of Cdc25A (Falck et al., 2001), degradation of cyclin D in

response to IR (Agami and Bernards, 2000), which redistributes p21 from cyclin D/CDK2 to

inhibit cyclin E/CDK2 (Sherr and Roberts, 1999, Yu et al., 2001).

The G1/S checkpoint function of p53 is similar to its role during the G2 checkpoint arrest.

Investigation of several sarcoma cell lines with different p53 mutations revealed that p53 is


28

required for the G2 checkpoint (Bache et al., 1999). The role of p53 depends on the ability

of p21 and retinoblastoma protein (RB) to implement a G2 phase arrest. This occurs by the

initial inactivation of cyclin B/CDK1 activity and is followed by a decrease in cyclin B and

CDK1 levels (Flatt et al., 2000). In addition p53 mediates a p21-dependent block of CDK2

activation which leads to delayed cyclin B/CDK1 activation. Inhibition of CDK2 may also lead

to decreased CDK2-dependent function of the transcription factor NF-Y, thereby reducing

the levels of cyclin B and CDK1 (Yun et al., 2003, Yun et al., 2006). Another transcription

factor also regulated by p53 is FoxM1 (forkhead box protein M1). p53’s repression of FoxM1

consequently disrupts the level of several important G2/M phase genes, such as cyclin B,

Cdc25B, and Aurora B (Leung et al., 2001, Wang et al., 2005). Like NY-F, both p53 and CDK2

can also regulate FoxM1. The importance of these CDK2 and p53 regulated pathways is

evident in p53-/- CDK2-/- double knockout cells that are unable to arrest in G2 phase, unlike

single knockout cells (Chung and Bunz, 2010).

1.5.4 Cdc25

Cdc25s are responsible for the removal of CDK inhibitory phosphorylations (Kristjansdottir

and Rudolph, 2004, Niida and Nakanishi, 2006). There are three human Cdc25 isoforms,

Cdc25 A, B, and C (Sadhu et al., 1990, Galaktionov and Beach, 1991, Nagata et al., 1991) of

which there are several Cdc25 splice variants (Baldin et al., 1997, Forrest et al., 1999, Bureik

et al., 2000, Wegener et al., 2000) that allow for differences in cyclin/CDK substrate

preferences (Gabrielli et al., 1997). The level of Cdc25 phosphatases at the G2/M transition

allow for the modulation of cell division in response to cell enviornment.


29

There is approximately 60% homology in the C-terminal catalytic domains of the three

isoforms and this determines the substrate specificity of the phosphatase (Gabrielli et al.,

1997, Rudolph et al., 2001). There is only 20-25% homology within the N-terminal domain

which contains several phosphorylation sites as well as nuclear import and export signals,

which regulate activity, stability and localisation. Each Cdc25 may have various functions

during the cell cycle and in response to certain cellular responses. When active, Cdc25

promotes cell cycle progression by activating cyclin/CDK complexes but when inactive it

results in cell cycle arrest due to the inability to active the cyclin/CDK target (Figure 1.4).

Localisation can be largely governed by phosphorylation events which can promote 14-3-3

binding, effecting nuclear export signal (NES) and nuclear localisation signal (NLS) (Dalal et

al., 1999, Lopez-Girona et al., 2001b, Lindqvist et al., 2004, Astuti et al., 2010, Astuti and

Gabrielli, 2011).

The most recent conclusions are that Cdc25B and Cdc25C are typically required for G2/M,

while Cdc25A is more involved in G1/S (Gabrielli et al., 1996, Morris et al., 2000, Manke et

al., 2005). However, Cdc25A, B and C all have the ability to regulate cyclin A/CDK2 and cyclin

B/CDK1 activity which both affect the entry of cells into mitosis (Gabrielli et al., 1996,

Kramer et al., 2004, Lindqvist et al., 2005) and recent evidence has demonstrated both

Cdc25B and Cdc25C are also required for S phase (Garner-Hamrick and Fisher, 1998,

Turowski et al., 2003). Once active, cyclin B/CDK1 can then promote its own activation loop

by phosphorylating any of the Cdc25s (Mailand et al., 2002, Astuti et al., 2010). This

discloses the functional redundancy between the family members as well as the intricacy of

the environmental factors that influence Cdc25 functions.


30

Overexpression studies have led to the assumption that Cdc25B functions prior to Cdc25C,

to prime the cells for mitosis whilst Cdc25C is required for the full activation cyclin B/CDK1

(Gabrielli et al., 1996, Karlsson et al., 1999, Goldstone et al., 2001). Cdc25B has the ability to

move across the nuclear membrane giving it access to cyclin A/CDK2 in the nucleus and

cyclin B/CDK1 at the centrosome (Gabrielli et al., 1996, Karlsson et al., 1999, Goldstone et

al., 2001). Cdc25C is primarily cytoplasmic in interphase, and then enters the nucleus in late

prophase (Dalal et al., 1999, Graves et al., 2001, Busch et al., 2007, Esmenjaud-Mailhat et

al., 2007, ). Cdc25C has also been reported to localise, along with active CDK1, at the

centrosome in late G2 and mitosis (Busch et al., 2007, Bonnet et al., 2008). Cdc25A, localises

in both the cytoplasm and the nucleus throughout the cell cycle and peaks within the

nucleus during G2 phase (Kallstrom et al., 2005) however mitotic regulation is presumed to

be co-ordinated by Cdc25B (Goldstone et al., 2001, Loffler et al., 2006).

As mentioned previously, Cdc25s are phosphorylated by Chk1 in response to damage,

rendering them inactive (Peng et al., 1997, Uto et al., 2004). This was first demonstrated by

Chk1 phosphorylation of Cdc25C Ser216. This phosphorylation causes the binding of 14-3-3

and blocks the nuclear importation of Cdc25C (Peng et al., 1997). Cdc25B phosphorylation

on Ser323 also leads to 14-3-3 binding and cytosolic retention (Giles et al., 2003). Cdc25A

can also be regulated by Chk1 during the DDR which blocks the activation of cyclin B/CDK1

(Falck et al., 2001, Chen et al., 2003, Sorensen et al., 2003).

In response to G2 phase checkpoint signalling, Cdc25B is phosphorylated by a number of

other checkpoint kinases. In response to UV(R), p38MAPK activates MAPKAP Kinase-2 (MK-


31

2) (Manke et al., 2005, Lemaire et al., 2006) leading to MK-2 phosphorylation of Cdc25B

Ser323, subsequent 14-3-3 binding and checkpoint arrest (Bulavin et al., 2001). Chk1 and

Chk2 are also capable of phosphorylating Cdc25B in response to UV(R) (Manke et al., 2005),

and both MK-2 and Chk1 are the two major kinases for Ser323 phosphorylation (Boutros et

al., 2007).

There is evidence that Cdc25 are activated by phosphorylation events that disrupt the

binding of 14-3-3 proteins and increase access to substrates (Wang et al., 2007, Astuti et al.,

2010). These activating phoshorylations are CDK2/ERK pathway-dependent (Margolis et al.,

2003, Wang et al., 2007), however the activating phosphorylations are largely regulated by

dephosphorylation by PP2A (protein phosphatase 2A) (Margolis et al., 2006a). Exogenous

expression of Cdc25B either mutated activating phosphorylations or not, caused abrogation

of the G2 checkpoint (Goldstone et al., 2001, Bugler et al., 2006, Astuti et al., 2010). This

may also be a consequence of the specific role of Cdc25B in checkpoint recovery (van Vugt

et al., 2004a, Karlsson-Rosenthal and Millar, 2006, Jullien et al., 2011, ) which is likely to

involve its activation of cyclin A/CDK2, which is also required for checkpoint recovery (van

Vugt et al., 2004b, Alvarez-Fernandez et al., 2010).

Evidence has demonstrated a necessity for Cdc25B during normal G2 progression (Gabrielli

et al., 1996, Karlsson et al., 1999, Bulavin et al., 2001). Cdc25B overexpression causes

activation of cyclin B/CDK1 while overexpression of a dominant negative Cdc25B results in

inhibition of cyclin A/CDK2 activation (Goldstone et al., 2001), both of which affect mitotic

progression. The regulation of Cdc25B during a normal cell cycle involves several


32

phosphorylations by Chk1 (Ser230 and Ser563) (Schmitt et al., 2006) and Aurora A (Ser353)

(Dutertre et al., 2004, Cazales et al., 2005). These phosphorylation sites control normal

mitotic entry, as mutagenesis of these sites caused defective mitotic entry (Dutertre et al.,

2004, Cazales et al., 2005). Overall, Cdc25B is the primary regulator of G2 phase progression

controlling cyclin A/CDK2 and cyclin B/CDK1 activation during normal and DDRs.

1.6 Other important cell cycle regulators

In additional to the G2 checkpoints already mentioned there are other important

components that contribute to these pathways. These can be identified in Figure 1.8 and

will be discussed in the following section.

1.6.1 p38 MAPK

p38 is a member of the Mitogen Activated Protein kinase (MAPK) family that mediates a

wide variety of cellular behaviours in response to extracellular stimuli ( Han et al., 1994, Lee

et al., 1994, Rouse et al., 1994, Raingeaud et al., 1995, Foltz et al., 1997, Freshney et al.,

1997, , Pietersma et al., 1997, Shalom-Barak et al., 1998, Shapiro et al., 1998), including

UV(R) irradiation (Rouse et al., 1994, Wang et al., 1998, Bulavin et al., 2001, Reinhardt et al.,

2007).

During a G1 arrest p38 activity is required for p21 stabilisation (Kim et al., 2002, Lafarga et

al., 2009); whereas the role of p38 during G2/M progression is less well defined and is

controversial with many reports that p38 is not functional in transformed and cancer cell


33

Figure 1.8 The ATR dependent DDR pathway is highly complex.

The Figure outlines the very basic regulatory pathways that exist between these proteins during

G2/M. Grey lines indicate regulation by degradation.


34

lines (Mikhailov et al., 2007). There is evidence that p38 can function downstream of ATM

and ATR, however in response to UV(R) damage p38 can be activated in the absence of ATM

and ATR, suggesting additional modes of p38 activation (Mikhailov et al., 2004, Reinhardt et

al., 2007). In response to UV(R) exposure, p38 can lead to the phosphorylation of Cdc25B

and Cdc25C (Bulavin et al., 2001). However, later evidence revealed that Cdc25B

phosphorylation and inhibition is catalysed by the p38 downstream kinase MK-2 (Obata et

al., 2000, Manke et al., 2005, Lemaire et al., 2006).

p38 can also enhance p53 stability by causing disassociation of p53 from its negative

regulator Mdm2 in response to UV(R) (Bulavin et al., 1999, Huang et al., 1999, Huang et al.,

2000, She et al., 2000). MK-2 is also capable of stabilising p53 by phosphorylation of sites

that can also be phosphorylated by Chk1 and Chk2 (She et al., 2002). The increase in p53

stability triggers several pathways to impose inhibition of cyclin B/CDK1 (el-Deiry et al.,

1993, Hermeking et al., 1997, Zhan et al., 1999). The p38-MK-2 pathway acts independently

of ATR and ATM (Bulavin et al., 2001, Bradham and McClay, 2006) but suggests overlap

between Chk1/2 and p38 signalling that has not been fully defined (Figure 1.8).

In addition to its checkpoint function, p38 is also present during an unperturbed cell cycle.

p38 has several mitotic roles and it has been shown to contribute toward correct spindle

assembly, spindle checkpoint function, the regulation of motor proteins required for the

correct attachment between kinetochore and microtubule ends (Lee et al., 2010). p38 also

has the ability to localise to the centrosomes from late G2 until mitosis where it can


35

phosphorylate Cdc25B (Bulavin et al., 2001, Cha et al., 2007) suggesting p38 can regulate

the timing of mitotic entry.

1.6.2 Claspin

Claspin is largely known as a checkpoint mediating protein required for the ATR dependent

activation of both Chk1 and BRCA1 and checkpoint arrest (Kumagai and Dunphy, 2000, Lin

et al., 2004, Sorensen et al., 2004, Liu et al., 2006, Wang et al., 2006, Lee et al., 2012,

Lindsey-Boltz and Sancar, 2011). Interestingly, Claspin levels peak during S and G2 phases

before being degraded during mitosis (Bennett and Clarke, 2006, Mailand et al., 2006,

Mamely et al., 2006, Peschiaroli et al., 2006) and G1 phase (Bennett and Clarke, 2006). The

cell cycle dependent fluctuations of Claspin protein level as well as the presence of low

levels of Claspin phosphorylation on Thr916, commonly observed with DNA damage, in

asynchronous untreated cells (Bennett and Clarke, 2006) suggest that Claspin has a role in

normal cell cycle progression.

Claspin phosphorylation on Thr916 and Ser945 is ATR dependent (Kumagai and Dunphy,

2003, Clarke and Clarke, 2005). Chk1 has been shown to phosphorylate Thr916 (Chini and

Chen, 2006) however others have shown that an unidentified kinase(s) distinct from ATR

and Chk1 is responsible for the direct phosphorylation of Thr916 (Bennett et al., 2008).

Phosphorylation of Claspin is required for the formation of the binding domain essential for

Chk1s interaction with Claspin and the activation of Chk1 by ATR (Clarke and Clarke, 2005,

Freire et al., 2006, Liu et al., 2006, Lindsey-Boltz et al., 2009). Claspin recruits Chk1 to ATR


36

for phosphorylation after which the Claspin-Chk1 interaction weakens (Chini and Chen,

2003, Jeong et al., 2003). In addition, Chk1 and Claspin are dependent on each other for

activation and stability, respectively (Chini et al., 2006).

Claspin degradation is required for G2 checkpoint recovery. For Claspin degradation to

occur, Plk1 phosphorylates Claspin which leads to ubiquitination by the SCFβTrCP

(Skp/Cullin/F-box containing complex) ubiquitin ligase complex (Mailand et al., 2006,

Mamely et al., 2006, Peschiaroli et al., 2006). During a G2 checkpoint arrest, Claspin

degradation is blocked by the ubiquitin specific phosphatases, USP28, allowing for

continued Chk1 activation (Peschiaroli et al., 2006, Bassermann et al., 2008). In addition,

Plk1 is degraded during the G2 checkpoint arrest thus preserving Claspin stability and

blocking mitotic entry (Smits et al., 2000). Plk1 substrates, such as Claspin, are

phosphorylated to prime them for Plk1 binding however the kinase that primes Claspin for

Plk1 binding is at present unknown. It is not known whether increased Claspin stability is

sufficient to sustain the G2 phase arrest, however its degradation is sufficient to promote

checkpoint recovery (Mailand et al., 2006, Mamely et al., 2006).

As well as a role during G2 phase, Claspin is involved in ATR-Chk1 mediated S phase

checkpoint signalling (Petermann and Caldecott, 2006) and normal S phase functions.

During an unperturbed S phase Claspin binds to chromatin, maintains integrity of

replication forks (Sar et al., 2004), and binds proteins required for replication (Lee et al.,

2003, Lee et al., 2005). In particular Claspin is required for normal replication fork

progression during an unperturbed S phase (Szyjka et al., 2005, Tourriere et al., 2005,


37

Hodgson et al., 2007, Petermann et al., 2008) which consequently gives Claspin the ability to

control the rate of proliferation. Therefore, unsurprisingly, depletion or overexpression of

Claspin causes a reduction or increase in proliferation respectively (Lin et al., 2004).

1.6.3 Wee1

Wee1 plays an important role in co-ordinating cell growth and cell division at the G2/M

transition. Wee1 is located in the nucleus and protects the cell from premature mitotic

entry (Heald et al., 1993, Liu et al., 1997). Wee1 inhibits mitosis by phosphorylating the

conserved Tyr15 on CDK1 (Russel and Nurse 1987, Featherstone and Russell, 1991,

Lundgren et al., 1991, Parker et al., 1992, Parker and Piwnica-Worms, 1992b). Wee1 activity

and stability is directly regulated by phosphorylation which produces a binding site for 14-3-

3 (Parker and Piwnica-Worms, 1992a, Honda et al., 1997b, Wang et al., 2000). In mice, 14-3-

3 binds Wee1 and increases protein stability and activity of Wee1 (Honda et al., 1997a,

Wang, 2000). Recently, Wee1 has been shown to associate and become negatively

regulated by cyclin A/CDK2 phosphorylation on Thr239. Mutation of Thr239, or the cyclin A

binding site, of Wee1 caused increased inhibitory CDK phosphorylation by Wee1 resulting in

inhibition of G2/M progression (Li et al., 2010). Furthermore, Wee1 degradation is also

mediated by phosphorylation: Wee1 is phosphorylated by CDK1 which promotes Plk1

binding required for APC ubiquitination and βTrCP E3 ligase mediated degradation

(Watanabe et al., 2004).


38

1.6.4 Cdh1

Cdh1 is an adapter protein of the ubiquitin ligase, APC, and regulates the activity and

substrate specificity of the E3 ligase which plays essential functions during late mitosis/G1.

There are two cofactors required for APC function, Cdc20 and Cdh1. Cdc20 is required for

early mitotic functions, while Cdh1 is required for late mitotic/G1 entry functions. Important

mitotic regulators degraded at the end of mitosis by Cdh1 are: cyclin A (Geley et al., 2001)

and cyclin B (King et al., 1995), Plk1 (Lindon and Pines, 2004), Aurora A (Castro et al., 2002)

and FoxM1 (Laoukili et al., 2008a, Park et al., 2008, Alvarez-Fernandez, 2010a). Cdc20 is also

a Cdh1 substrate (Prinz et al., 1998, Pfleger and Kirschner, 2000) and Cdc20 compensates

for a loss of Cdh1 by degrading accumulated Cdh1 substrates (Garcia-Higuera et al., 2008, Li

et al., 2008). However, Cdh1 is critical for the fidelity of mitosis as a loss of Cdh1 results in

mitotic defects such as an increased number of binucleated cells (Garcia-Higuera et al.,

2008, Li et al., 2008).

During S, G2 and M phases a mobility shift, on an immunoblot, indicative of

phosphorylation of Cdh1 is observed (Kramer et al., 2000) and correlates with inactivation

of Cdh1 (Kramer et al., 1998, Zachariae et al., 1998a, Jaspersen et al., 1999, Lukas et al.,

1999a, Blanco et al., 2000, , Kramer et al., 2000, Yamaguchi et al., 2000). Overexpression of

Cdh1 affects many of its substrates such as cyclin B, Plk1, Claspin and Cdc20 (Gao et al.,

2009). Therefore the regulation of Cdh1 levels and activity are important for cell cycle

progression. During normal cell cycle progression the phosphatase, Cdc14, controls Cdh1

dephosphorylation (Visintin et al., 1998, Jaspersen et al., 1999). During a G2 phase


39

checkpoint arrest, Cdc14B translocates from the nucleus to dephosphorylate and

consequently activate Cdh1 (Bassermann et al., 2008). Cdh1 activation contributes to the

G2 checkpoint arrest by ubiquitination and consequently degradation of Plk1 (Bassermann

et al., 2008). At the same time, Claspin and Wee1 stabilisation is required for the G2

checkpoint arrest. Claspin, also a Cdh1 substrate, is blocked from Cdh1 dependent

degradation during the G2 checkpoint arrest by USP28. USP28 stabilises the G2 checkpoint

arrest by stabilising Claspin dependent Chk1 activation.

APC-Cdh1 (APCCdh1) activation is regulated by several mechanisms. One is the ubiquitination

of the APC conjugated ubiquitin enzyme E2, which provides a negative feedback loop that

eventually destroys APCCdh1 activity (Rape and Kirschner, 2004, Rape et al., 2006). Second,

the accumulation of CDK activity leads to the dissociation of Cdh1 from APC (Lukas et al.,

1999a, Kramer et al., 2000). Third, phosphorylated Cdh1 is a target of SCF complex, thereby

limiting APCCdh1 activity (Benmaamar and Pagano, 2005). Finally, in G1 phase E2F activates

the transcription of Emi1, a known inhibitor of APC (Grosskortenhaus and Sprenger, 2002,

Hsu et al., 2002).

CDK dependent phosphorylation of Cdh1 occurs on multiple sites (Hall et al., 2004). Alanine

mutations of these phosphorylation sites supports increased APCCdh1 binding and activity

(Lukas et al., 1999a, Kramer et al., 2000) and may lead to a growth arrest due to the inability

of mitotic cyclins to accumulate (Hall et al., 2004). In budding yeast, CDK1 has been shown

to phosphorylate Cdh1 to provide a binding site for Plk1 which then also phosphorylates

Cdh1 to regulate its activity (Crasta et al., 2008). In mammalian cells other CDKs are most


40

likely responsible for Cdh1 phosphorylation since CDK1 along with its binding partner cyclin

B is not active during S and G2 phase, when Cdh1 phosphorylation and inactivation is

apparent. Cyclin A has been shown to phosphorylate and inactivate the APCCdh1 complex

(Lukas et al., 1999a, Sorensen et al., 2001), and it is cyclin A/CDK2 that is most likely

involved in Cdh1 regulation during S and G2. In the absence of cyclin A, premature

degradation of the APC target cyclin B can ultimately affect mitotic entry (Reber et al.,

2006). However, others have shown that disruption of cyclin A binding to Cdh1 only

affected cyclin A but not other APCCdh1 substrates such as cyclin B and securin (Sorensen et

al., 2001). In Drospholia, cyclin E/CDK2 is a negative regulator of APC (Sigrist and Lehner,

1997), but in the mammalian system cyclin E/CDK2 had little effect on APC activity (Brandeis

and Hunt, 1996, Lukas et al., 1999a).

1.6.5 Plk1

Plk1 (polo-like kinase 1) is a member of the polo-like kinase family, which consists of 5

members, Plk1-5, that all contain a conserved domain known as the polo-box domain (PBD).

Plk1 is the most studied family member and is highly expressed in a number of human

cancers. Plks are important for the fidelity of mitosis and are particularly important for

cytokinesis (Barr et al., 2004).

Plk1 is a serine/threonine kinase expressed predominantly during late G2 and mitosis, and is

activated by phosphorylation on the Thr210 residue (Lee and Erikson, 1997, Qian et al.,

1998, Jang et al., 2002a, Jang et al., 2002b, Kelm et al., 2002). Once active, Plk1


41

phosphorylates the consensus motif Glu-Asp-X-Ser/Thr-φ (X = any amino acid, φ = a

hydrophobic residue which is frequently a leucine). The PBD is critical for Plk1 activity and it

is involved in auto-inhibition of Plk1 (Lee and Erikson, 1997, Mundt et al., 1997). The PBD is

required for the subcellular localisation of Plk1 (Song et al., 2000, Lowery et al., 2004,

Lowery et al., 2005, Lowery et al., 2007) by recognising binding to a particular phospho-

epitope (Elia et al., 2003a). Although many phospho-proteins interact with the PBD, it is

unclear if all binding proteins are Plk1 substrates or if the binding is simply required to

increase the local concentration of Plk1.

The PBD is responsible for the localisation of Plk1 to the centrosomes by binding Plk1

substrates localised at the centrosomes. Like many other Plk1 substrates, Nedd1 is primed

by CDK1 phosphorylation at the centrosomes which leads to Plk1 binding (Zhang et al.,

2009a). Plk1 localisation and function at the centrosomes is important as it is involved in

centrosomal maturation (Barr et al., 2004). Its activity is required for the recruitment of γ-

tubulin to the centrosomes and is essential for microtubule nucleation (Casenghi et al.,

2003, Haren et al., 2009, Johmura et al., 2011). Depletion of Plk1 results in defects in the

mitotic spindle and incorrect cell division (Lane and Nigg, 1996, Sumara et al., 2004,

McInnes et al., 2006, Peters et al., 2006, Lenart et al., 2007, Santamaria et al., 2007, Haren

et al., 2009).

The over-expression of PBD-GFP has been shown to displace endogenous Plk1 and resulted

in a dramatic increase in the G2 phase population (Elia et al., 2003b), indicating that Plk1 is

also required for G2 phase progression. Plk1 has essential G2 phase checkpoint roles (van


42

Vugt et al., 2004a). In response to the checkpoint arrest Plk1 levels are reduced and the

protein inactivated, by Cdh1 dependent degradation ( Smits et al., 2004, Bassermann et al.,

2008, ). Inversely, recovery from a checkpoint arrest requires an increase in Plk1 levels and

activation (van Vugt et al., 2004a, van Vugt and Medema, 2004, Macurek et al., 2008). Once

active, Plk1 targets several pathways for checkpoint recovery. Plk1 phosphorylates both

Claspin and Wee1 (van Vugt et al., 2004a, Watanabe et al., 2004, Mamely et al., 2006)

which leads to SCF mediated degradation and is responsible for Cdc25C activation (Kumagai

and Dunphy, 1996, Qian et al., 1998), and Cdc25B localisation (Lobjois et al., 2009). The

mechanism by which cells increase the level of Plk1 required for checkpoint recovery is

unknown but may involve FoxM1. Plk1 is a FoxM1 regulated transcript but is also a positive

regulator of FoxM1 therefore Plk1’s checkpoint recovery role may involve a positive

feedback loop (Fu et al., 2008). Because Plk1-mediated phosphorylation of FoxM1 requires

initial priming by CDK activity, this could be a key event regulated by the low levels of CDK

activity, now known to be needed for checkpoint recovery (Duursma and Cimprich, 2010).

1.7 Checkpoint recovery

Checkpoint recovery occurs after the cell has repaired the DNA damage. Therefore,

checkpoint recovery is a crucial process that determines the fate of a cell after DNA

damage. Understanding the mechanisms governing checkpoint recovery has recently

become just as important as understanding the mechanisms of the checkpoint arrest.

Mutation of many checkpoint arrest pathways forces cells into mitosis. By silencing

checkpoint signalling such as ATM/ATR-Chk1/Chk2 signalling, recovery may be achieved.


43

Little information is known about the inactivation of ATR but possibly involves its mutually

dependent binding partner, ATRIP (Yang et al., 2004). By blocking ATM/ATR signalling

pathways it relieves the negative regulation of cell cycle controllers such as Cdc25s and

consequently G2/M progression. Some examples are; mutations of 14-3-3 binding sites on

Cdc25B caused cells to rapidly overcome G2 checkpoint arrest (Giles et al., 2003, Schmitt et

al., 2006) and mutation of Chk1 activating phosphorylation sites will overcome the

checkpoint (Walworth et al., 1993, Walworth and Bernards, 1996).

Dephosphorylation and degradation events are largely involved in checkpoint recovery.

Several phosphatases such as Wip1 and PP2A have identified roles in checkpoint recovery

to dephosphorylate such checkpoint proteins as Chk2, Chk1, γH2AX and p53 (Chowdhury et

al., 2005, Keogh et al., 2006, Shreeram et al., 2006, Lindqvist et al., 2009). Ubiquitin-

dependent proteasome-mediated degradation is also thought to play an essential role for

checkpoint recovery. The identification of SCFβTrCP as a switch between arrest and recovery

was acknowledged due to its ability to impose a checkpoint arrest by Chk1 dependent

degradation of Cdc25A, and checkpoint recovery via degradation of Wee1 and Claspin.

Interestingly, both the degradation of Wee1 and Claspin are Plk1 dependent. The trigger to

the SCFβTrCP switch is not well understood but components of the pathway such as Plk1

signalling remain less elusive.

Although it was once suspected that cyclin/CDK activity was inhibited during a checkpoint

arrest recent evidence has shown that low levels of CDKs are required during the G2

checkpoint to regulate transcription of genes required for recovery, such as Plk1, cyclin A


44

and cyclin B (Alvarez-Fernandez et al., 2010a, Alvarez-Fernandez et al., 2010b). In particular,

low levels of cyclin A/CDK are required to phosphorylate FoxM1, relieving the auto-

repressor N terminal domain of FoxM1 (Laoukili et al., 2008b) and leading to the expression

of FoxM1 dependent G2/M genes. This pathway for checkpoint recovery may also involve

Cdc25 as depletion of Cdc25B blocked cells from entering mitosis after a checkpoint arrest

(Goldstone et al., 2001).

1.8 G2/M regulation by cyclin A/CDK2 and cyclin B/CDK1

Cyclins A and B were the first cyclins identified in sea urchin eggs and were shown to be

synthesised during each cell cycle and degraded during mitosis (Evans et al., 1983, Swenson

et al., 1986, Standart et al., 1987). In human cells both cyclin A and cyclin B are essential for

G2/M progression.

There are three types of cyclin B, B1, B2 and B3. Cyclin B3 was the most recently identified,

it binds CDK2 and has important meiotic functions (Nguyen et al., 2002). Unlike cyclin B3,

cyclins B1 and B2 both bind CDK1 but localise to different sites within the cell. Cyclin B1 is

the predominant cyclin B required for mitosis, which is why cyclin B1 nullizygous mice are

not viable (Brandeis et al., 1998).

Cyclin B/CDK1 activation is highly coordinated and the initial activation of cyclin B/CDK1

occurs in the cytoplasm and at the centrosomes (De Souza et al., 2000a, Jackman et al.,

2003). This leads to the abrupt translocation and increased concentration of cyclin B/CDK1

to the nucleus approximately 5 minutes prior to NEB in HeLa cells (Pines and Hunter, 1991,


45

Bailly et al., 1992, Gallant and Nigg, 1992, Ookata et al., 1992, Hagting et al., 1999). The

increased activity of cyclin B/CDK1 is dependent on the continual nuclear translocation of

cyclin B/CDK1 which is essential for driving mitotic events. Activation of cyclin B/CDK1 in the

nucleus involves the phosphorylation of its cytoplasmic retention site and the consequent

dephosphorylation by Cdc25 family members to enhance activity (Hoffmann et al., 1993, Li

et al., 1997, Yang et al., 1998, Hagting et al., 1999, Yang et al., 2001, Bulavin et al., 2003a,

Bulavin et al., 2003b, Walsh et al., 2003, , Margolis et al., 2006b, ,). The increase of cyclin

B/CDK1 in the nucleus was thought to involve a block of nuclear export by Plk1 dependent

phosphorylation events (Toyoshima-Morimoto et al., 2002, Yuan et al., 2002, Jackman et al.,

2003). However, later evidence revealed CDK1 localisation in the nucleus can be attributed

to an increase in nuclear import (Gavet and Pines, 2010). The specifics of the nuclear import

are unclear but it may be linked to associations with unidentified, nuclear import/export

factors ( Li et al., 1997, Hagting et al., 1999).

The ability of cyclin B/CDK1 activation levels to control the timing of these mitotic events

was demonstrated by the classic study by Masui and Markert ‘cytoplasmic control of

nuclear behaviour....’, where the cytosol of a metaphase oocyte can induce metaphase in an

interphase ooctye (Masui and Markert, 1971). In addition, initial cyclin B/CDK1 activity

levels cannot promote later mitotic events that require higher cyclin B/CDK1 activity levels

(Lindqvist et al., 2007). Gavet and Pines (2010) also showed that different thresholds of

cyclin B/CDK1 activity induced different mitotic events. For example, the initial activation of

cyclin B/CDK1 at the centrosomes is linked to the conversion of an interphase centrosome

to a MTOC and regulates microtubule nucleation and mitotic spindle formation (Gabrielli et


46

al., 1996, De Souza et al., 2000a) which begins in late G2 phase. This is followed by

chromosome condensation after initial formation of the mitotic spindle (Li et al., 1997,

Hagting et al., 1999) and leads to the translocation of cyclin B/CDK1 into the nucleus and

NEB (Gavet and Pines, 2010).

The timing of cyclin B/CDK1 activation is shown to be dependent on the activity of cyclin

A/CDK2 activity (De Boer et al., 2008) and is the reason that a number of groups have

independently demonstrated that cyclin A depletion or CDK2 inhibition results in a G2 phase

delay (Fung et al., 2007, Gong et al., 2007a). There are two types of cyclin A, cyclin A1 and

A2. Studies of knockout mice revealed that cyclin A2 is required for viability whereas cyclin

A1 expression is confined to germ cells and is required only for male fertility (Liu et al.,

1998, Lele and Wolgemuth, 2004, Wolgemuth et al., 2004). However others have shown

that in fibroblasts cyclin A (A1 and A2) and cyclin E have redundant roles required for cell

proliferation but in contrast cyclin A is essential for hematopoietic and embryonic stem cell

proliferation (Kalaszczynska et al. 2009). Cyclin A associates with CDK2 (Pines and Hunter,

1990, Tsai et al., 1991, Clarke et al., 1992, Kobayashi et al., 1992, Lees and Harlow, 1993)

and localises the complex to the nucleus (Maridor et al., 1993). A small pool has also been

shown to localise to the centrosome in late G2 phase (De Boer et al., 2008). Cyclin A/CDK2

has biphasic activation, with a small pool being activated in S phase (Girard et al., 1991,

Walker and Maller, 1991, Pagano et al., 1992, Zindy et al., 1992), but the major pool being

activated in early G2 phase (Lehner and O'Farrell, 1990, Whitfield et al., 1990, Walker and

Maller, 1991, Pagano et al., 1992, Knoblich and Lehner, 1993, Furuno et al., 1999,,

Goldstone et al., 2001, Hu et al., 2001, De Boer et al., 2008).


47

The mechanism by which cyclin A/CDK2 regulates the timing of cyclin B/CDK1 activation is

unclear. The G2 phase Cdc25B-dependent activation of cyclin A/CDK2 is blocked in G2

phase checkpoint arrested cells, and the expression of a mutant form of CDK2 that does not

require Cdc25 dependent activation bypassed the G2 phase checkpoint (Goldstone et al.,

2001). Interestingly, ATM/ATR inhibitors can relinquish the block in cyclin B/CDK1

activation, but not cyclin A/CDK2 activation in response to DNA damage, suggesting that the

Cdc25B-dependent activation of cyclin A/CDK2 is independent of ATM/ATR during the

checkpoint arrest (Goldstone et al., 2001). There is also evidence that cyclin A/CDK2 can

regulate ATR signalling (Hu et al., 2001, Jazayeri et al., 2006, Myers et al., 2007). Together,

these data point to an undefined role for cyclin A/CDK2 and its immediate upstream

activator Cdc25B in the G2 phase checkpoint arrest.

Cyclin A is degraded during mitosis (Jacobs et al., 2001) but in early mitosis cyclin A/CDK2 is

required for correct spindle assembly (Beamish et al., 2009). Without cyclin A/CDK2 spindle

rotation occurs due to the absence of proper spindle attachments (Beamish et al., 2009).

Overall, cyclin A/CDK2 has important roles in both G2 and mitosis and also in response to

the checkpoint. However, the other pathways that may be involved in these cyclin A/CDK2

regulated processes are not well defined.

1.9 Chk1 regulation

The phosphorylation of Chk1 on Ser317 and Ser345 in response to DNA damage has been

identified as a major mechanism of Chk1 regulation. These sites can also be found


48

phosphorylated in unperturbed cells suggesting that they function during this time to

regulate Chk1. Mutational analysis of these sites has been used to demonstrate their roles

both in response to damage and during an unperturbed cell cycle.

Phosphorylation of Ser317 and Ser345 regulate Chk1 kinase activity, in particular during the

DDR. The basal level of these phosphorylations is increased dramatically in response to DNA

damage however mutation of Ser317 or Ser345 to either alanine (phospho-inhibitory) or

aspartate (phospho-mimicking) did not affect the basal level of Chk1 kinase activity with or

without DNA damage (Gatei et al., 2003). However, others have shown that mutation of

Ser345 affects Chk1 activity and its ability to arrest cells during the checkpoint (Capasso et

al., 2002). Only a kinase dead mutant of Chk1 (D130A) was able to abolish Chk1 activity with

or without DNA damage in response to damage (Gatei et al., 2003). Collectively these

results demonstrate that the precise role for Chk1 during the G2 checkpoint is still

undefined and that mutational analysis of Ser317 and Se345 may not be the best tool for

use in Chk1 regulatory studies.

Mutation of Ser317 to alanine caused decreased phosphorylation of Ser345 (Niida et al.,

2007, Walker et al., 2009); however mutation of Ser345 did not noticeably impact Ser317

phosphorylation suggesting that Ser317 influences phosphorylation on Ser345. Functional

analysis of Ser317 and Ser345 has also been conducted using mutational analysis of these

sites (Niida et al., 2007). The commonality that exists between the two sites is that single

inhibitory mutations result in impaired checkpoint arrests, however the literature

demonstrates the difference these sites have on Chk1 function and localisation (Niida et al.,


49

2007, Wilsker et al., 2008). Independent of Ser317 status, Chk1 phosphorylated on Ser345

was found located at the centrosome (Wilsker et al., 2008) and prevented the premature

activation of CDK1 during a normal cell cycle and a G2 phase checkpoint arrest (Kramer et

al., 2004, Loffler et al., 2006). Ser345 is located within a 14-3-3 consensus binding motif

(Capasso et al., 2002, Jiang et al., 2003). The disruption of 14-3-3 binding caused by Ser345

mutation could be the reason for the differences seen with single site Ser317 or Ser345

mutants and may also be another reason why Chk1 mutants are not useful tools for Chk1

studies.

Evidence is mounting that the C terminus of Chk1 mediates critical aspects of Chk1 function

(Palermo et al., 2008, Wang et al, 2012) . Chk1 is predominantly a nuclear protein and in the

absence of damage Chk1 phosphorylated on Ser317 and Ser345 is found associated with

chromatin (Jiang et al., 2003, Smits, 2006, Smits et al., 2006). Nuclear levels of

phosphorylated Chk1 on Ser317 and Ser345 increase in the presence of DNA damage (Jiang

et al., 2003). Phosphorylated Chk1 Ser317 and Ser345 are also evident in the cytoplasm

following DNA damage, suggesting that Chk1 phosphorylation can promote cytoplasmic

translocation from the nucleus after DNA damage. However, Alanine mutation of both

Ser317 and Ser345 promoted cytoplasmic localisation suggesting these phosphorylations

are more likely to promote nuclear localisation (Jiang et al., 2003).


50

1.9.1 The role of Chk1 during an unperturbed cell cycle

There is evidence that a basal level of Chk1 activity is present during an unperturbed cell

cycle (Capasso et al., 2002, Gatei et al., 2003, Zhang et al., 2006b, Xu et al., 2012) and

correlates with the observation that phosphorylated Chk1 is present during an unperturbed

cell cycle, which is often unmentioned by publishing authors. Along with the fact that Chk1-/-

is embroyic lethal in muce, collectively, this suggests a role for active Chk1 during an

unperturbed cell cycle.

Chk1 inhibition during an unperturbed cell cycle enhances the rate of mitotic entry (Tang et

al., 2006). Chk1 can regulate mitotic entry by two possible mechanisms. Firstly, similar to its

role in the DDR, Chk1 phosphorylates and inhibits Cdc25B during an unperturbed cell cycle;

however the phosphorylation sites differ from the sites phosphorylated in response to

damage. When cells are deficient of Chk1, Cdc25 is not inhibited by Chk1 which results in

premature activation of CDK1 (Chen et al., 2003) and cells proceed uncontrollably into

mitosis. In particular, Chk1 negatively regulates Cdc25B by phosphorylating several sites

including Ser230 and Ser563 (Schmitt et al., 2006), during G2 phase therefore governing

mitotic entry. Ser230 phosphorylation of Cdc25B occurs at the centrosomes and contributes

to the timing of mitotic entry (Schmitt et al., 2006). Chk1 phosphorylated at Ser345 can also

localise at the centrosomes (Wilsker et al., 2008) in an unperturbed cell cycle and confirms

that active Chk1 present during an unperturbed cell cycle regulates Cdc25B activation at the

centrosomes. Recent work shows that Chk1 is capable of controlling the activation of Polo-

like kinase 1 (Plk1) during an unperturbed cell cycle as well as during the DNA damage


51

response. Chk1 depletion was shown to cause an increase in Plk1 activity (Tang et al., 2006).

However enhanced Plk1 levels and activation are known to coincide with mitotic entry and

therefore could be a consequence of increased mitotic entry caused by Chk1 inhibition.

These are clear indications that Chk1 has the ability to control G2 phase progression,

however it is unknown if other mechanisms are involved.

Chk1 also has critical roles during S phase, such as maintaining replication fork integrity.

Consequently, depletion of Chk1 results in replication fork stalling, possible collapsed forks

and aberrant replication forks (Petermann et al., 2008) but at the same time can lead to the

activation of adjacent and possibly suppressed origins (Maya-Mendoza et al., 2007).

Therefore, it is unsurprising that the rate of replication forks is affected by Chk1 loss, in fact

the rate is decreased by half in Chk1 knockout cells when compared to wildtype cells

(Petermann et al., 2006).

Chk1 removal results in an inability of cells to sustain a mitotic arrest ( Zachos et al., 2007,

Zachos and Gillespie, 2007, Carrassa et al., 2009). Specifically, Chk1 has a role during mitosis

to oversee the correct segregation of the genome, thus explaining why the absence of Chk1

leads to misaligned chromosomes followed by mitotic catastrophe (Niida et al., 2005). This

may be linked to the role of Chk1 during the spindle assembly checkpoint where it is

required to control the activation of Aurora B and BubR1 by phosphorylation and

localisation (Zachos et al., 2007). Without the correct functioning of spindle checkpoint

components, such as Chk1, the fidelity of mitosis is at risk.


52

1.9.2 Chk1 inactivation

Once active, Chk1 is required until the repair of damage is complete. The inhibition of Chk1

must be timed correctly as premature inactivation results in mitotic catastrophe (Latif et al.,

2004). Decreased Ser317 and Ser345 phosphorylation correlates with checkpoint exit.

Dephosphorylation of these sites can be achieved by protein phosphatase 1 (PP1), and in

the absence of PP1 the checkpoint arrest is extended (den Elzen and O'Connell, 2004,

Kuntziger et al, 2012).

More recently a new mode of negative regulation of Chk1 has been uncovered. Chk1 was

inhibited by phosphorylation on Ser286 and Ser301 catalysed by CDK1 during mitosis. First

indications demonstrated that these phosphorylation events were required to inhibit Chk1

for mitotic progression (Shiromizu et al., 2006). Later, these phosphorylation events were

also shown to control Chk1 localisation. Mutational analysis revealed that these

phosphorylation sites are required for the cytoplasmic localisation of Chk1 from the

nucleus. Further investigation found that these inhibitory phosphorylation sites can be

phosphorylated in response to DNA damage also. During the DDR cyclin A/CDK2 is most

likely responsible for Chk1 phosphorylation on Ser286 and Ser301, as cyclin B/CDK1 is

inactivated during this time (Ikegami et al., 2008). Phosphorylation of these sites in

response to DNA damage does not correlate with the earlier suggestion that these sites

were inhibitory. The evidence that these inhibitory phosphorylations occur on the same

Chk1 molecule also phosphorylated on the activating sites implies the function of these


53

sites remains elusive. A great deal of further investigation is required to understand the

timing of these phosphorylations and their role during G2/M progression.

1.10 Hypothesis

Chk1 is a central component of the ATR/ATM dependent G2 checkpoint whose role and

regulation during G2/M progression is not yet well defined. Cyclin A/CDK2 is a major

regulator of G2 phase progression and like Chk1 has the ability to control cyclin B/CDK1

activation. The ability of Chk1 to regulate cyclin B/CDK1 activity is well known but the

mechanism/s by which cyclin A/CDK2 regulates cyclin B/CDK1 activation is less well known.

There is evidence that cyclin A/CDK2 activity is required during the G2 phase checkpoint but

its function is poorly understood. In this study the mechanism(s) by which G2 phase cyclin

A/CDK2 regulates the timing of cyclin B/CDK1 activation and entry into mitosis will be

examined, as well as the contribution of cyclin A/CDK2 to the G2 phase checkpoint. Due to

Chk1s very strong role in G2/M progression, the initial hypothesis of this study is that cyclin

A/CDK2 regulates the activity of Chk1 in normal G2 phase progression by suppressing Chk1

activity to permit cyclin B/CDK1 activation and mitotic entry. The second hypothesis is that

cyclin A/CDK2 has a similar role and utilises similar mechanisms to suppress Chk1 in

recovery from the G2 phase checkpoint arrest. Third, we will look beyond Chk1 and

determine the involvement of additional known or unknown G2/M proteins to act within

these pathways.


54

1.11 Significance

The understanding of the cell cycle and its regulation has come a long way in the last

decade. However the details of pathways involved in cell cycle regulation, particularly G2/M

phase transition and the G2 phase checkpoint, are still being determined. This study

contributes to developing and enhancing this knowledge. Checkpoints are often defective in

cancers, therefore understanding their detailed pathways and components may allow for

further development or identification of better targets for anticancer therapies.

1.12 Aims of the study

Specifically, this study will elucidate pathways involved in normal G2/M progression as well

as during a G2 checkpoint response. The regulation of G2 phase by cyclin A/CDK2 will be

particularly examined. The specific aims of this study are-

1. How does cyclin A/CDK2 regulate G2/M progression?

2. Is Chk1 active during an unperturbed cell cycle and what is its role?

3. Does cyclin A/CDK2 regulate G2 phase Chk1 to contribute to normal G2/M progression

and checkpoint recovery?

4. What other G2/M pathways may be regulated by cyclin A/CDK2 to allow mitotic entry?

55

Chapter 2

Materials and Methods

Chapter 2 – Material and Methods

56

All chemicals used were of analytical grade rand were purchased from Sigma Aldrich (Castle

Hill, NSW, Australia), Invitrogen (Mulgrave, VIC, Australia), Merck Chemicals (Kilsyth, Victoria,

Australia), Calbiochem (Darmstadt, Germany) and Univar (Taren Pt, NSW, Australia), unless

otherwise indicated. All solutions were prepared using Milli-Q purified water and autoclaved

or filter-sterilized through 0.22μm filters when appropriate. All glassware used for cell or

bacterial culture was sterilized prior to use. Restriction enzymes were purchased from New

England Bio Labs (Ipswich, MA, USA). All tissue culture materials were purchased from

Invitrogen unless otherwise indicated. Protease inhibitors (Complete Mini-EDTA-free

protease inhibitor cocktail) were obtained from Roche (Dee Why, NSW, Victoria). All PCR

primers and siRNA were synthesised and purchased from Invitrogen unless stated.

2.1 Antibodies

All antibodies were stored as per manufacturer’s instructions for short term use, at either 4°C

or -20°C. The dilution of antibodies for immunoprecipitations (IPs) refers to the final dilution

of antibody used in the total IP reaction volume, after the addition of protein A beads and

cell lysates. Dilutions used for immunoblotting (IB) and immunofluorescence (IF) are given in

Table 2.1 also.


57

Table 2.1 Antibodies List.

Dilution used per application

Antibody Name

Source IB IF IP

ACA Home made (mouse)

1:1000

Cdc25B Santa Cruz (C-20) (rabbit) 1:100

CDK1 PY15 Cell Signalling (rabbit) 1:1000

Cdh1 Abcam (mouse)

1:100

Chk1 Abcam (goat)

1:100 1:50

pChk1 S286 Gift from Dr Masaki Inagaki (mouse) 1:100

pChk1 S301 Gift from Dr Masaki Inagaki (mouse) 1:100

pChk1 S317 Bethyl Labs (rabbit)

1:50

pChk1 S317 Cell Signalling (rabbit) 1:1000

1:50

Chk1 S345 Cell Signalling (rabbit) 1:500 1:50

Chk2 Cell Signalling (rabbit) 1:500

pChk2 T68 Abcam (rabbit) 1:500

Claspin Abcam (goat) 1:5000

cyclin A Santa Cruz (H-432) (rabbit) 1:1000 1:200 1:50

cyclin B Home made (rabbit) 1:100

Flag Invitrogen (mouse) 1:1000 1:100 1:100

GFP Cell Signalling (mouse) 1:1000 1:100

pMek1 T286 Cell Signalling (rabbit) 1:500 1:100

MPM2 Abcam (mouse)

1:100


58

2.2 Cell Biology Methods.

2.2.1 Cell culture

HeLa cells (Human cervical carcinoma cell line) were grown in Dulbecco’s Modified Eagles

Medium (DMEM) supplemented with 10% Serum Supreme, 1 mM sodium pyruvate, 1 M

Hepes, 2 mM L-Glutamine. Cells were incubated at 37°C in 5% CO2.

FO2 (ATR defective; Seckel Syndrome derived cell line), NFF (Neonatal Foreskin Fibroblasts),

and WI38 (Untransformed Fibroblasts) cells were grown in DMEM supplemented with 10%

Foetal Bovine Serum and 1 mM sodium pyruvate, 1 M Hepes, 2 mM L-Glutamine.

HeLa cells stabling expressing Cherry tagged α-tubulin were grown in the same conditions as

the parental HeLa cells.

U2OS Myc-Cdh1 inducible cells were grown in DMEM supplemented with 10% Fetal Bovine

Serum, 1 mM sodium pyruvate, 1 M Hepes, 2 mM L-Glutamine and 2 µg/mL tetracycline. Cells

Myc Cell Signalling (mouse) 1:1000 1:100

PCNA DAKO (mouse) 1:10 000

Plk1 Abcam (mouse) 1:500 1:100

Plk1 pT210 Abcam (mouse) 1:500

α-tubulin Sigma Aldrich (mouse)

1:500

γ-tubulin Sigma Aldrich (mouse)

1:500

Wee1 Cell Signalling (rabbit) 1:1000


59

were incubated at 37°C in 5% CO2. For Cdh1 induction cells were washed 3 times for 10

minutes in warmed phosphate buffered saline (PBS). Cells were then incubated in the media

listed above, but without tetracycline addition.

2.2.2 Cell Synchronisation

For thymidine synchrony experiments, cells were treated with 2.5 mM thymidine

supplemented media for 16-24 h. Cells were then washed three times with pre-warmed PBS

and then once with pre-warmed DMEM before adding growth media supplemented with 24

μM thymidine and 24 μM 2’deoxycitidine. This allows a release of cells from their G1/S

arrest. After an 8 h release the majority of cells are in G2 and were then blocked again with

thymidine (2.5 mM) for 16-24 h. Cells were then washed as above, and full medium

supplemented with 24 µM thymidine and 24 μM 2’deoxycitidine was used to release cells

from the arrest. These cells were then collected at the indicated times and analysed by FACs

and immunoblotting or visualised for cell cycle progression by time lapse microscopy.

2.2.3 Cell Treatments

G2 cell cycle checkpoint arrest with Etoposide was achieved with addition of 1 μM Etoposide

(Sigma Aldrich) into complete medium and incubated for approximately 16-24 h. For mitotic

synchronisation, cells were treated with nocodazole (Sigma Aldrich) in complete medium for

16 h at a final concentration of 0.5 µg/ml. Mitotic cells lose their adherence to the dishes,

thus were collected by simply removing media and manually disrupting cell attachment, a

technique known as a “mitotic shake off”. Roscovitine (rosco) (Sigma Aldrich) was used at 50


60

μM for indicated times to inhibit both CDK1 and CDK2 activity. CDK2 inhibitor (Calbiochem),

Ro09 3033, was used at concentrations and indication times, as specified in individual

experimental figure legends. CDK1i RO-3306 (Calbiochem) was used at 9 µM, CDK2i PHA-

533533 (Courtesy of Professor Grant McArthur (Peter Mac (Victoria, Australia)) was used at 4

µM or as indicated (Deans et al., 2006, Pevarello et al., 2005a, Vassilev et al., 2006). The

proteasome inhibitor, MG132 (Calbiochem (La Jolla, CA, USA)) was used at 20 µM. Chk1

inhibitor, SB 218078 (Calbiochem), was used at 2.5 µM or as indicated in individual

experimental figure legends. Caffeine (Sigma Aldrich) was used at 5 mM. Plk1i (BI 2536)

(Selleckchem, Boston, USA) was used at concentrations indicated in individual experimental

figure legends. DMSO (Sigma Aldrich) treated samples were used as controls and cells were

treated with the same volume of DMSO as used for the active comparison.

Cells treated with UV(R) where washed three times for 10 minutes with pre-warmed PBS.

After the last wash, a minimalistic amount of PBS was applied to the monolayer of cells

adhered to the dish. Cells were then radiated with the specified amount of UV(R). Once

complete, PBS was removed and completed media was applied to the cells. Cells were

returned to the incubator until required.

2.2.4 Transfection Treatments

2.2.4.1 Plasmid transient transfection

For transient transfection of Flag-Chk1 constructs (Gift from Prof Kum Kum Khanna, Qld

Institute of Medical Research, Brisbane, Australia), Myc-Cdh1, Cherry-Cyclin A, GFP and Myc


61

plasmids, cells were plated onto 10cm2 dishes or 6-well plates and allowed to attach for 18 h.

Cells were then transfected with 1 µg/mL of the relevant DNA plasmid in either 10 cm2 or 6

well plates. Lipofectamine 2000 reagent (Life Technologies) was used according to the

manufacturer’s instructions. Transfection medium was replaced with the appropriate

medium for the cell line and incubated for a further 24 h, or until specified, at 37°C with 5%

CO2.

2.2.4.2 siRNA transfection

Table 2.2 lists the siRNA used in this study. Included are the sequences, concentrations of the

siRNA supplied by Ambion (Austin, TX, USA), Invitrogen and Dharmacon (Lafayette CO, USA).

Note that the amount of the control (N) siRNA is intentionally not listed as the concentration

used varied depending on the concentration of the targeted siRNA used in the experiment.

To transfect siRNA duplexes, Lipofectamine 2000 Reagent was used as per manufacturer’s

instructions. All samples were collected for analysis 24 h after transfection or as specified in

the experiment. However, for transfection of NFF, cells were transfected using the same

method but the transfection reagent was replaced with Darmafect 1 (Dharmacon)

transfection reagent, which provided better transfection efficiency in this cell line.


62

Table 2.2 List of siRNA.

siRNAs Target Sequence (5’ 3’) Supplier Final conc. (nM)

Control/nonsense

(Con/N) aatgatctacctgttaagagtcctgtctc Invitrogen -

Chk1 tcgtgagcgtttgttgaac Invitrogen 30

cyclin A

A1 aaggatcttcctgtaaatgatgagc Invitrogen 50

A2 taaacctaaagtgggttacatg Invitrogen 50

A3 ggaaagtcttaagccttgtctca Invitrogen 50

A4 Not Available – product # 2512 Ambion 50

A5 Not Available – product # 2514 Ambion 50

Chk2 c1 aaacgccgtcctttgaatatt Invitrogen 90

c2 aagaacctgaggagcctaccc Invitrogen 90

c3 aatgtgtgaatgacaactact Invitrogen 90

Claspin (Clasp) aaggaaagaaaggcagccaga Invitrogen 40

Cdh1

Cdh1 A ccacaggauuaacgagaau Dharmacon Smartpool 40

Cdh1 B ggaacacgcugacaggaca Dharmacon Smartpool 40

Cdh1 C gcaacgaugugucucccua Dharmacon Smartpool 40

Cdh1 D gaagaagggucuguucacg Dharmacon Smartpool 40

2.2.5 Cell fixation and immunofluorescent labelling

All cell lines were cultured to near confluence, and harvested by trypsinisation. Cells were

diluted 3- to 10-fold and cultured overnight onto poly-L-lysine- (0.1 mg/mL) or fibronectin-

(20 μg/mL) coated glass coverslips, washed twice with PBS, and fixed in either 4°C 4%

paraformaldehyde (PFA) (Electron Microscopy Sciences, Hatfield, PA, USA) for 20 minutes at

room temperature (RT), followed by permeabilisation with 0.1% saponin for 10 minutes at RT

or treated with methanol (MeOH) for 5 minutes at -20 °C. Cells were washed twice with PBS,

and blocked for 30 minutes at room temperature with 2.5% Bovine Serum albumin (BSA).


63

This was followed by primary antibody incubation at RT for 1 h, or 4C overnight diluted with

0.1% BSA in PBS. For control incubations, the primary antibody was omitted (Cellomic

experimentation (with pChk1 S317 antibody, see section 2.2.6). Cells were washed twice with

PBS and incubated simultaneously with the relevant secondary antibodies and the DNA

marker DAPI (4’-6-Diamidino-2-phenylindole) where relevant, in 0.1% BSA in PBS, for 1 h at

RT in the dark. After washing twice in PBS, coverslips were mounted onto glass slides using

Prolong Gold (Invitrogen) and stored at 4°C.

Secondary antibodies used were conjugated-Alexa 488 and Alexa 555 (Invitrogen) and were

used at a 1:250 dilution (8 μg/ml). DAPI (1 μg/mL) was included with the secondary

antibodies. Samples were imaged using either a Zeiss Apotome widefield fluorescence

microscope or a Zeiss 510 Meta confocal microscope. Images were analysed using Zeiss

AxioVision or Zen confocal software. Images were processed using Adobe Photoshop or

ImageJ freeware.

2.2.5.1 Cyclin A staining

Cells were fixed for 5 minutes with methanol at -20°C. Prior to incubation with blocking

solution (2.5% BSA/PBS) cells were incubated on ice for 5 minutes with 2% digitonin/PBS.

Cells were then blocked and stained for cyclin A (Santa Cruz (Santa Cruz, CA, USA)) as per

section 2.2.3


64

2.2.5.2 pChk1 staining

Coverslips were incubated in 3% PBA/PBS for 30 minutes on ice and permeabilised for 20

minutes on ice with 0.5% Triton X-100/PBS. Cells were then blocked and stained for pChk1

Ser345 (Cell Signaling) as per section 2.2.3.

pChk1 Ser317 immunofluorescent staining using antibody purchased from Bethyl

(Montgomery, TX, USA) was carried out after fixing cells with 4% paraformaldehyde (PFA)

(method outlined in section 2.2.5).

2.2.5.3 Plk1 staining

Coverslips were fixed for 5 minutes with 3% PFA/PBS and permeabilised for 5 minutes with

2% Triton X-100/PBS. This was then followed by 5 minutes incubation with methanol at 20°C.

Coverslips were fixed for 5 minutes with 3% PFA, 2% sucrose in PBS. Triton X-100 (0.5%) was

then added for 5 minutes at 4°C. Further permeabilisation was carried out by incubating cells

for 5 minutes at room temperature in 0.5% Triton X-100 in 20 mM HEPES pH 7.4, 3 mM

MgCl2, 50 mM sodium chloride, 300 mM sucrose, 0.02% Sodium Azide solution. This was

then immediately removed and methanol was added for 5 minutes at -20°C. Coverslips were

then washed twice with PBS and incubated in 3% BSA/PBS in a humidified chamber for 30

minutes for blocking. After blocking, primary antibody was diluted in 3% BSA/PBS plus 0.02%

Sodium Azide and incubated on cells for 1 h in a humidified chamber. Primary antibody was

removed by three 20 minute washes with PBS followed by secondary antibody incubation

along with DAPI. Secondary antibodies were diluted in 3% BSA and incubated for 30 minutes


65

in a humidified chamber. Lastly the coverslips were washed with five 20 minute washes with

PBS before mounting onto slides.

2.2.6 Microscopy and image acquisition

2.2.6.1 Immunofluorescence microscopy

Immunofluorescence microscopy was performed using a Zeiss Apotome widefield

microscope equipped with a cooled CCD camera. Images were taken using 40x, 60x or 100x

oil objectives, through appropriate filter sets for DAPI, FITC/Alexa488, and Cy3 fluorophores,

using exposure times optimised for each field photographed. ImageJ freeware or Adobe

Photoshop programmes were used in post-capture processing and analyses of digital images.

2.2.6.2 Confocal laser scanning microscopy

Confocal microscopy was performed using a Zeiss 510 Meta laser scanning spectral confocal

microscope. Images were acquired using Plan APO 63x or 100x oil objectives, and

fluorescence emissions were captured sequentially using line and frame averaging to reduce

noise through standard filters. The 488nm, 543nm and 633nm lasers were used to excite

Alexa-488, Cy3 and Cy5 fluorophores, respectively. For cell cycle-regulated expression and

co-localization studies, confocal scan settings were standardised using control cells stained

with either primary antibody in the presence of the relevant peptide, or with the secondary

antibody only, to exclude signals obtained through non-specific antibody staining.


66

2.2.6.3 Time lapse imaging

For live imaging of cells, cells were cultured in a 6 well plate format and transfected, as

specified. Plates were transferred to a live cell imaging chamber (Zeiss) and maintained at

37C with 5% CO2 in the dark until imaging was required using the Zeiss Cell observer

microscope. Transfected cells were serially imaged with a 20x objective over 15 min intervals

using transmitted light. Fluorescent filter sets were used when GFP or Cherry tagged cell lines

were used. Scoring of these samples was conducted manually and approximately 200 cells

per condition were counted.

2.2.7 FACS Analysis

All FACS samples in this study were quantified for cellular DNA content by flow cytometry,

cells were washed in PBS twice and then fixed in 70% ethanol overnight at -20°C. After

fixation cells were washed in PBS, and then resuspended in 500 μL of solution containing 25

μg/mL of propidium iodide (PI) and 25 μL of RNase A (1 mg/mL) in PBS. After staining with PI

for approximately 15 minutes, samples were filtered and analysed using the FACs Calibur (BD

Biosciences) system. 10,000 cells were acquired per sample. Cell Quest Software was used

for the acquisition and FlowJo was used for analysis.

2.2.7.1 Chk1 expressing FACS analysis

Cells co-transfected with Green Fluorescent Protein (GFP) and Flag-Chk1 constructs were

analysed by FACS to determine the DNA profile of the positively expressing GFP cells. For GFP


67

detection by FACS, cells were fixed and stained as above for DNA content with PI. Acquisition

required a 488-nm laser emitting at 635 nm to detect the positive GFP expressing cells and a

PI laser to acquire this data simultaneously. Analysis of 50,000 cells was acquired and was

conducted using FlowJo software. Only the DNA profile of the positively expressing cells was

analysed.

2.2.7.2 MPM2 FACS

To specifically detect mitotic cells, cells were fixed as above. After fixation, cells were washed

twice with PBS and then resuspended in blocking solution for 1 h at room temperature.

Blocking solution consists of 1% BSA (Sigma Aldrich) in Phosphate buffered saline plus tween

20 (PBST). Monoclonal MPM2 antibody (Abcam, 1:100 (San Francisco, CA, USA)) was added

directly to each sample, suspended thoroughly and gently agitated for 1 h at room

temperature. Cells were washed twice with PBS and then incubated with secondary

antibody, Alexa 488 goat anti mouse IgG (Invitrogen) diluted 1:250 in blocking solution, for 1

h at room temperature in the dark. All following steps had minimised exposure to light. Cells

were washed twice in PBS and resuspended in solution containing 25 μg/mL PI and 25 μL of

RNase A (1 mg/mL) in PBS. After staining with PI for 15 minutes, samples were filtered and

analysed by a FACs Calibur. For mitotic analysis 50,000 cells were scored per sample. Analysis

was conducted using FlowJo software.


68

2.2.8 Cellomics

Each cell line was cultured as indicated above and seeded into 96 well flat bottomed view

plates at 5000 cells per well for high content image analysis using the Cellomics high content

imager. Cells were then washed in PBS, fixed with ice-cold methanol for 20 minutes at -20°C

and stained with pChk1 S317 (Bethyl) as indicated above. DAPI (1 µg/mL) (Sigma Aldrich) was

used to stain DNA when incubating cells with the secondary antibody (1:250). DAPI was used

as it has superior quantitative binding to DNA, providing better discrimination of the cell

cycle phases. Cells were stored in PBS in darkened conditions at 4°C until analysis with the

Cellomics ArrayScan VTI high-content imager (from Thermo Scientific, 10x objective, XF93

filter set) using Cellomics ArrayScan software. Approximately 2000 cells were scored for each

cell line and controls stained with secondary antibody only were analysed to eliminate

background fluorescence. The DNA content/DNA intensity was used to ascertain the cell

cycle distributions. This allowed for the discrepancy between G1, S and G2/M populations.

The same conditions were also carried out on controls which were then subtracted from the

samples. Floating bar graphs display the mean including three standard deviations and the

range represents the 95 percentile of each population.


69

2.3 Molecular Biology Methods

2.3.1 Plasmid list

The following table lists all plasmids used in this thesis. Many of these plasmids were gift from various

colleagues. The methods used for the generation of the vectors required for the thesis are detailed in

the subsequent sections.

Table 2.3 List of Plasmids

PLASMID NAME VECTOR BACKBONE

Cherry α-tubulin mCherry-C1

Cherry Chk1 S286 mCherry-C1

Cherry Chk1 S301 mCherry-C1

Cherry Chk1 SS286,301AA mCherry-C1

Cherry Chk1 WT mCherry-C1

Cherry Cyclin A mCherry-C1

Flag Chk1 WT pCI-neo

Flag Chk1 S286A pCI-neo

Flag Chk1 S301A pCI-neo

Flag Chk1 SS286,301AA pCI-neo

Flag Chk1 SS317,345DD S286A pCI-neo

Flag Chk1 SS317,345DD S301A pCI-neo

Flag Chk1 SS317,345DD SS286,301AA pCI-neo

GFP pEGFP-C1

GST-Chk1 pGex 4T-1

GST-Chk1 S286A pGex 4T-1

GST-Chk1 S301 pGex 4T-1

GST-Chk1 SS286,301AA pGex 4T-1

Myc Cdh1 pX-MYC

Myc Cdh1 AAA pX-MYC

Myc-Plk1 pRcCMV

PBD-GFP pEGFP-C1


70

Cherry α-tubulin, Myc-Cdh1, Myc-Cdh1 AAA and Myc-Plk1 were gifts from Nichole Den Elzen

(Institute of Molecular Biology, University of Queensland), Claus Sorensen (Biotech Research

and Innovation Centre, University of Copenhagen) and Erich Nigg (The Centre of Molecular

Life Sciences, University of Basel) respectively.

2.3.1.1 Generation of S286A, S301A and SS286,301AA

pCI-neo Flag Chk1 constructs (Chk1 WT; Chk1 SS317,345AA; Chk1 SS317,345DD) were gifts

from Dr Kum Kum Khanna. Chk1 was cloned into this vector using EcoR1 and Sal1 restriction

sites.

Using the Stratagene site directed mutagenesis protocol both Ser286 and Ser301 were

mutated to alanine using the primers listed in Table 2.3. Primers were designed using

Intergrated DNA technologies’ web-based Oligo Analyser 3.0 primer design programs

available online.

Table 2.4 List of Chk1 mutagenesis primers used.

Primer Sequence (5' → 3')

286A Forward gag tca ctt cag gtg gtg tgt cag agg ctc cca gtg g

286A Reverse cca ctg gga gcc tct gac aca cc acct gaa gtg act c

301A Forward cca att tgg act tcg ctc cag taa aca gtg c

301A Reverse gca ctg ttt act gga gcg aag tcc aaa tgg g


71

2.3.1.2 Preparation of competent cells

2mL Lysogeny broth (LB) medium was inoculated with XL1-blue or BL21 Escherichia coli (E-coli)

bacterial strains and incubated at 37C overnight with orbital shaking at 250 rpm. A 2 mL

overnight culture was used to inoculate 200 mL LB medium in the presence of 15 μg/mL

tetracycline (XL1-blue only) and cultured for a further 4 h in 1 L culture flasks, or until the

absorbance at 600 nm read approximately 0.6. Cultures were then chilled on ice, and

centrifuged at 3,000 rpm for 15 minutes at 4C in sterile 250 mL centrifuge bottles. Pelleted

bacteria were washed twice with 200 mL sterile Milli-Q purified water. Bacteria were

transferred to 50 mL Falcon tubes and washed once with 20 mL 10% glycerol. Bacterial

pellets were then resuspended in an equal volume of 10% glycerol and either used

immediately or aliquoted into eppendorf tubes and stored at -80C until use.

2.3.1.3 Transformation of bacteria

Bacteria were transformed with plasmids or polymerase chain reaction (PCR) reactions using

the heat shock method. DNA was added to competent Escherichia Coli and incubated on ice

for 30 minutes. Samples were then heat shocked for 45 seconds at 42 °C and placed back on

ice for 2 minutes. Bacteria were then transferred into 1 mL of LB media and grown for 1 h

with gentle shaking at 37°C. Bacteria were then plated onto LB agar plates containing

appropriate antibiotics and incubated inverted overnight at 37°C.


72

2.3.1.4 Screening for positive clones

Bacterial colonies were picked into 2 mL LB medium containing appropriate antibiotics, and

shaken at 250 rpm overnight at 37°C. Bacteria were pelleted and DNA extracted as per the

Promega DNA purification kit protocol. Samples were diluted used 100 μL of RNase/DNase

free water supplied by Promega (Wisconsin, USA) and stored at -20°C. The DNA was further

subjected to sequencing to confirm positive mutants.

2.3.1.5 Sequencing

600 ng of mini-prep plasmid DNA purified using the Promega DNA miniprep kit as per

instructions, was added into a 1.5 mL eppendorf tube along with 6.4 pmol of sequencing

primer. Samples were then sent for sequencing to the Australian Genome Research Facility

(AGRF) at the University of Queensland (UQ) St Lucia campus. Sequence traces were checked

visually with Applied Biosystems sequence scanner software, to ensure that subcloned

inserts were in the correct reading frame.

The sequencing primer used for pCI-neo was the universal T7 primer (5’

TAATACGACTCACTATAGGG 3’).

2.3.1.6 Large scale plasmid DNA preparation

Following sequence verification of plasmid inserts, large quantities (100 μg-1 mg) of plasmid

DNA were prepared using the Qiagen Hi-Speed Plasmid Purification kit, according to the

manufacturer’s instructions. DNA concentration and purity were determined by absorbance


73

at 260 nm, and the ratio of absorbance at 260 nm to absorbance at 280 nm, respectively.

DNA was digested and gel electrophoresis conducted to confirm plasmid purification.

2.3.1.7 Restriction enzyme digests

Restriction digests were carried out according to manufacturer’s instructions using the

recommended buffers, except that BSA at a final concentration of 100 μg/mL was added to

all reactions. Vectors for ligation were treated with shrimp alkaline phosphatase (Promega),

according to manufacturer’s instructions prior to ligation reactions, to reduce end-to-end

vector re-ligation frequency.

2.3.1.8 Agarose gel electrophoresis

DNA of interest was electrophoresed in agarose gels prepared at appropriate concentrations

in TBE (Tris/Borate/EDTA) (pH8.0) buffer, containing 0.2 μg/mL ethidium bromide. Lambda

DNA cut with Hind III/ Eco RI or a 100 base pair DNA ladder (Invitrogen) were used as DNA

markers. Following electrophoresis, gels were photographed under UV(R) illumination with a

BioRad (Hercules, CA, USA) CCD camera using Molecular Analyst software.

2.3.1.9 Ligation

DNA ligations were performed in 10 μL reactions using 25-50 ng vector DNA and an

insert:vector ratio of approximately 5-10:1. Reactions were carried out in the presence of T4

DNA ligase (1 U/10 μL) and 100 μg/mL BSA according to manufacturer’s instructions


74

(Invitrogen). The relevant amount of vector DNA was ligated in an additional reaction as a

negative control; this was used to provide a base-line colony count for subsequent

transformations.

2.3.2 Plasmid construct cloning

2.3.2.1. Generation of Cherry-Chk1 and GST-Chk1 constructs

Chk1 was cloned from the pCI-neo vector using EcoR1 and Sal1 digestion. This produced a

fragment of 1.4 kb, which was excised from the DNA electrophoresis gel. Purification of gel

bands was conducted using the Promega gel purification kit as per instructions. mCherry

vector and pGex4T-3 were both digested with EcoR1 and Sal1.

Cherry-Chk1 ligations were screened for insertion by conduction of single and double

restriction enzyme digestions with EcoR1 and Sal1. If correct, these constructs were further

tested by cell culture transfection and immunoblotting of samples.

Both constructs were sequenced to confirm positive mutations. Sequencing was carried out

as per section 2.3.1.5 and primers used for sequencing the constructs are listed in Table 2.4.

Table 2.5 List of sequencing primers used.

Sequencing Primer Sequence (5' → 3')

mCherry (reverse) gaaatttgtgatgctattgc

pGex 4T-3 (forward) gggctggcaagccacgtttggtg


75

After positive identification, clones were subjected to large scale DNA purification as per

section 2.3.1.6. This was followed by confirmation of DNA digestion and DNA gel

electrophoresis as per sections 2.3.1.7 and 2.3.1.8 respectively.

2.3.2.2 Generation of PBD-GFP

Myc-Plk1 was used to generate PBD-GFP. The PBD was expressed from the parental plasmid

using the sequencing primers in Table 2.6. PCR products were resolved by agarose gel

electrophoresis (Section 2.3.1.8) and the amplified PBD was purified from the agarose gel

using a Promega DNA purification kit. Purified PBD was ligated into a pGEMTeasy cloning

vertor to make use of restriction sites for cloning into a fluorescenct GFP containing vector.

pGEMTeasy-PBD was digested with EcoRI. Restriction digests were resolved by agarose gel

electrophoresis and the PBD band (of approximately 800 base pairs) was purified using the

Promega DNA purification kit. The EcoRI digested PBD was then digested with SalI, which was

possible due to the removal of a SalI site from the pGEMTeasy cloning vector. PBD with EcoRI

and SalI cut ends assisted the cloning into the recipient vector in the correct orientation.

Finally, PBD digested with EcoRI and SalI was ligated into eGFP-C1 pre-digested with EcoRI

and SalI. Ligated products were screened with sequential digests using EcoRI and then SalI

enzymes. Positive clones were then transfected into cells to confirm the correct cloning of

PDB-GFP.


76

Table 2.6 PBD-GFP cloning primers

Primer name Sequence (5'-->3')

Forward Primer ctgaccattccaccaaggttttc

Reverse Primer ggcagagttccggaggatt

2.4 Protein Biochemistry Methods

2.4.1 Immunoblot Analysis

Cells were lysed by suspension in Universal Lysis Buffer (ULP) (50 mM Tris pH7.4, 150mM

NaCl, 2 mM EDTA, 2 mM EGTA, 25 mM NaF, 0.2% Triton X-100, 0.3% NP40, 25 mM β-

glycerophosphate and 5 µg/ml aprotinin, 5 µg/ml leupeptin, 0.1 mM PMSF) and incubated for

30 minutes at 4°C. Cell lysates were cleared by centrifugation at 13,200 rpm for 15 minutes.

Protein estimation by BioRad Bradford Assay allowed equalisation of all lysates, and 20 µg of

protein was resolved by 10% sodium dodecyl sulphate polyacrylamide gel electrophoresis

(SDS-PAGE). Proteins were then transferred to Hybond-C nitrocellulose membrane

(Amersham Biosciences, USA) using the semi dry transfer method. Membranes were blocked

in 5% skim milk powder solution in PBS with 0.1% Tween 20 (PBST) for 30 minutes and

incubated in the required antibody overnight at 4°C.

After incubation with the appropriate antibody, membranes were washed in PBST for 3 x 10

minutes. Membranes were then incubated with the appropriate Horseradish Peroxidase

(HRP) conjugated secondary antibody (Zymed (Mulgrave, VIC, Australia) 1:2000) for 1 h at

room temperature. Finally, membranes were washed for 10 minute intervals for 1 h with


77

PBST or TBST (Tris-buffered Saline plus 0.1% Tween 20), depending on the antibody

specificity, before detection by enhanced chemiluminescence (Perkin Elmer, Massachusetts,

USA) using the Fisher Biotech Imaging system. Quantitative image analysis was performed

using ImageJ freeware.

2.4.2 Immunoprecipitation

For co-immunoprecipitation experiments, specified amounts of protein were pre-cleared

with either protein G (Flag immunoprecipitation) or A sepharose (Chk1 and cyclin A

immunoprecipitation) beads for 1 h at 4˚C. Pre-cleared extracts were collected by

centrifugation at 2,000 rpm for 0.5 minute at room temperature (RT) then incubated with the

appropriate antibody/conjugated beads for 2 h at 4˚C. Immune complexes were collected

with protein A or G Sepharose beads for 1 h at 4˚C and washed four times with ULP. The

immunoprecipitated protein complexes were eluted with SDS sample buffer and the sample

boiled for 5 minutes.

Lysates immunoprecipitated with Cyclin A-protein A coupled beads were first pre-cleared

with protein A beads. Anti-Flag M2 affinity gel (Invitrogen) was used for Flag

immunoprecipitation and lysates were precleared with protein G. Chk1 immunoprecipitation

was carried out using protein A beads. IgG (Invitrogen) was used as a control for IP

experiments. The amount used was equivalent to the antibody being used in the experiment.


78

2.4.2.1 Antibody coupling to protein A beads

1 mL protein A beads were washed twice with PBS and then twice in conjugation buffer

(50 mM Tris pH 7.6, 150 mM NaCl, 2% BSA). The beads were then suspended in 1 mL volume

in conjugation Buffer, along with 20 μL of cyclin A (Santa Cruz H-432; 200 µg/mL) antibody or

0.4 µL of rabbit-IgG (Invitrogen; 10 mg/mL) and incubated on a rocker for 1 h at RT. The

supernatant was then removed after spinning for 5 minutes at 3,000 g. The beads were then

transferred to a 15 mL tube with 10 mL Na Borate buffer, pH 9.0 and washed twice,

centrifuging at 3,500 rpm for 4 minutes at RT. Covalent coupling of the antibody was

achieved by adding 50 mg DMP (dimethyl pimelimidate, Sigma Aldrich D8388) to 10 mL Na

Borate Buffer plus beads, ensuring a pH > 8.3 and incubating for 30 minutes at RT with

rocking.

The reaction was stopped by pelleting beads and adding 10 mL 0.2 M ethanolamine, pH 8.0.

The beads were washed once and then incubated for 2 h at RT with rocking. Ethanolamine

was removed by washing twice with PBS. The beads were further washed with 1 mL 100 mM

glycine pH 3.0, followed by a wash with 1 mL 100 mM Tris pH 8.0. Lastly, beads were washed

in PBS, ensuring pH ≈7.4 and resuspended in 1 x bed volume of PBS with an addition of Na

Azide to 0.02%.


79

2.4.3 Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and

immunoblot analyses

2.4.3.1 SDS-PAGE

Protein extracts were resolved using SDS-PAGE on 4, 7 or 10% polyacrylamide gels. The

percentage gel used would be based on the protein to be detected. Equal amounts of protein

or fractions of equivalent volume were diluted with 5x SDS-PAGE sample buffer (1 M Tris pH

6.8; 20% glycerol; 2% SDS; 20% β-mercaptoethanol; 0.01% bromophenol blue), boiled for 5

minutes, and then separated alongside a Precision Plus Protein Dual colour standard (BioRad)

on polyacrylamide minigels in running buffer (25 mM Tris; 250 mM glycine; 0.1% SDS pH 8.3)

at 130 Volts for 1½ to 2½ h. The resolving and stacking gel compositions are outlined below

in Table 2.5 (volumes are sufficient for 2 mini-gels).

Table 2.7 Components and Volumes used for SDS-PAGE gels

Resolving Gel Stacking Gel REAGENT 10% 7% 4%

1M Tris pH 8.8 (mL) 2.5 2.5 2.5 --

1M Tris pH 6.8 (mL) -- -- -- 0.5

Water (mL) 3.96 4.96 5.96 2.72

10% SDS (µL) 100 100 100 40

30% bis- acrylamide (mL) 3.3 2.33 1.33 0.68

TEMED (µL) 12 12 12 5

10% APS (µL) 100 100 100 80


80

In some cases, proteins were visualised by staining with Coomassie Brilliant Blue for 20

minutes with rocking, followed by de-staining in de-stain solution (45% methanol; 10% glacial

acetic acid) for 1 h.

2.4.3.2 Immunoblot transfer

For immunoblot analyses, proteins were separated by SDS-PAGE and electro-transferred to

nitrocellulose filters (Millipore) as follows. Transfer materials were pre-soaked in transfer

buffer (25 mM Tris; 250 mM glycine; 20% methanol; 0.005% SDS) prior to assembly in the

BioRad semidry transfer cassette. 4 pieces of filter paper were placed onto the cassette,

followed by the nitrocellulose membrane and the gel on top of the membrane. Care was

taken during each assembly step to exclude air bubbles. The gel was then overlayed with 4

more pieces of filter paper. The cassette was then closed and proteins were transferred at

either 15 Volts for 12 minutes (7 and 10% gels) or 25 minutes (4% gel). Protein loading was

analysed using Ponceau S (1% ponceau S; 1% glacial acetic acid) staining of filters.

2.4.4 Cyclin A/CDK2 kinase assay

2.4.4.1 Induction of recombinant protein expression in E.coli BL21 cells

5 mL overnight cultures of BL21 bacteria transformed with pGex4T3-Chk1 WT, pGex4T3-Chk1

S286A, pGex4T3-Chk1 S301A or pGex4T3-Chk1 SS286,301AA were used to inoculate 50 mL

cultures at 1/200 dilution. Cultures were incubated at 37C, and shaken at 250 rpm for

approximately 4 h or until the absorbance at 600 nm was approximately 0.6, indicating log-


81

phase growth. Recombinant protein expression was induced by addition of 0.5 mM IPTG

(Isopropyl β-D-1-thiogalactopyranoside) and incubating at 30C, shaking at 250 rpm for 2 h.

BL21 bacteria were pelleted and washed once with ice-cold STE (10 mM Tris pH 8, 150 mM

NaCl, 1 mM EDTA). Proteins were isolated by resuspending pellets in isotonic lysis buffer (55

mM DTT, 2 mM PMSF, 2% sarcosyl) with 1 mg/mL lysosome (1mL lysis buffer per 50mL

culture) and incubating on ice for 40 minutes. This was followed by brief sonication for 2

minutes. For the isolation of soluble extracts, lysates were centrifuged at 15,000 rpm for 30

minutes at 4C and supernatants removed. Protein product sizes were verified by

electrophoresing pre- and post-induction protein samples on 10% SDS-PAGE gels, followed

by Coomassie Brilliant Blue staining (G-250). Proteins were then bound to GST beads by

incubation of the supernatant with 400 μL of beads, along with 2.5% Triton X-100, for 2 h

rotation at 4 °C. Beads were then collected by centrifugation at 1500 rpm for 2 minutes at

4°C. To wash, beads were resuspended in PBS and collected by centrifugation at 1500 rpm

for 1 minute at 4 °C. This was repeated 3 times, followed by a high salt wash with 1 M NaCl in

PBS and another 2 washes in PBS alone. Beads were stored as a 50% slurry in PBS plus

protease cocktail inhibitor at 4°C. For prolonged storage samples were kept at either -20°C or

-80 °C

2.4.4.2 In vitro kinase assay.

Equal volumes were taken of each purified GST-Chk1 sample (300-500 ng) and washed twice

with kinase buffer (KB) (20 mM Tris pH7.4, 1 mM DTT). The supernatant was removed and

samples resuspended in 30 μL of reaction buffer (20 mM Tris pH7.4, 15 mM MgCl2, 0.2 mM

http://en.wikipedia.org/wiki/Isopropyl_%CE%B2-D-1-thiogalactopyranoside


82

ATP, 1 mM DTT and 25 mM β-glycerophosphate) containing 5 μCi *γ32P] and recombinant

cyclin A/CDK2 (Cell Signalling). The reaction was incubated for 15 minutes at 30 °C and then

stopped by the addition of 6 μL of SDS-PAGE loading buffer. Samples were then boiled at

100°C for 5 minutes before resolving by SDS-PAGE (10%).

Gels were then stained with Coomassie Brilliant Blue, as per 2.4.3.1. Gels were then sealed

and exposed to X-ray film (Fuji Film Corporation) overnight at -80°C.

83

Chapter 3

Active Chk1 is present in unperturbed G2 phase

Chapter 3 – Active Chk1 present in G2 phase

84

3.1 INTRODUCTION

The control of cell cycle progression is reliant on cyclin / CDK complexes. The progression of

cells from G2 phase into mitosis is of critical importance as mitosis is the pivotal stage where

daughter cells will segregate to form two independent functioning cells. The process of

mitosis is complex and to ensure cells are capable of completing it successfully, they are

examined for correct functionality during the G2 phase before they enter this final stage of

the cell cycle.

The cyclin/CDK complexes required during the transition from G2 into mitosis are cyclin

A/CDK2 which functions during G2 phase and cyclin B/CDK1 which is the predominant mitotic

complex. The activation of the cyclin B/CDK1 complex, and therefore subsequent mitotic

entry, is conducted by a family of phosphatases known as the Cdc25s. Cdc25s are specific

activators of cyclin/CDK complexes and catalyse the removal of inhibitory phosphorylations,

Thr14 and Tyr15. In addition to the activation of cyclin B/CDK1 initiated by Cdc25B, active G2

phase cyclin A/CDK2 is required for the subsequent activation of cyclin B/CDK1 and mitotic

entry.

If DNA damage is detected during a cell cycle, appropriate checkpoints are activated;

enabling cells to stop cycle progression until all damage is resolved. Complex and structured

signalling events take place in order for the checkpoint to be implemented, but invariably

cyclin/CDK complexes are the final target of checkpoint signalling which halts cell cycle

progression. There are a number of checkpoints throughout the cell cycle. Of specific

interest here is the G2 phase checkpoint. The type of DNA damage inflicted upon the cell


85

governs the responding G2 phase checkpoint signalling pathway utilized; the ATR-Chk1

pathway responds to single stranded DNA damage and the ATM-Chk2 pathway to double

strand DNA damage. There is evidence that these two pathways can cooperate in response

to DNA double strand breaks (DSBs), with ATR-Chk1 activated as a response to repair

intermediates involving ssDNA (Banin et al., 1998, Zhao and Piwnica-Worms, 2001).

The central effector kinase of the G2 phase DNA damage checkpoint is Checkpoint kinase 1

(Chk1). Chk1 is a multifunctional kinase which until recently was thought to only function in

response to DNA damage. The essential importance of ATR-Chk1 function is demonstrated

by the embryonic lethality of both ATR-/- and Chk1-/- mice (Liu et al., 2000, Takai et al., 2000)

compared to the viable although cancer prone ATM and Chk2 knockout mice (Barlow et al.,

1996, Elson et al., 1996, Xu et al., 1996, Takai et al., 2002). In response to DNA damage, Chk1

is phosphorylated by ATR at two activating sites in humans, Ser317 and Ser345 (Zhao and

Piwnica-Worms, 2001). Its regulation and functioning is thought to be dependent on these

phosphorylation events (Capasso et al., 2002) therefore Ser317 and Ser345 phosphorylation

have been used as indicators of Chk1 activation. Although each of these phosphorylation

sites have been linked to the control of different Chk1 functions/localizations (Wilsker et al.,

2008) they are both phosphorylated rapidly in response to damage. Phosphorylation of these

sites may be sequential as response to UV(R) damage requires Ser317 phosphorylation for

subsequent phosphorylation of Ser345 before Chk1 can activate downstream checkpoint

proteins. Despite the strong evidence that the Ser317 and Ser345 phosphorylations reflect

Chk1 activation, mutational analysis disputes this fact. The phospho inhibitory Chk1

Ser317Ala (serine mutated to alanine) and Ser317Ala/Ser345Ala double mutant is still active,


86

with kinase activity level similar to wildtype constructs, while the kinase dead mutant (lysine

38 substituted to methionine) had no kinase activity (Tapia-Alveal et al., 2009). These data

suggest that although phosphorylation sites Ser317 and Ser345 are required for activation

they are not necessarily an appropriate representation of the activity status of Chk1.

However the two remain closely linked as high levels of phosphorylated Ser317 and Ser345

correspond to the arrest of cells in G2 phase and reduction in these phosphorylations

corresponds to the resolution of the damage and re-entry into the cell cycle. The detection of

Chk1 kinase activity and the presence of Ser317 and Ser345 phosphorylations during an

unperturbed cell cycle suggests that a basal level of Chk1 remains primed during a normal

cell cycle so that the cell can rapidly respond to DNA damage (den Elzen and O'Connell, 2004,

Harvey et al., 2004, Latif et al., 2004, Niida et al., 2007, ), but the question still remains as to

the role of Chk1 during a normal cell cycle.

Although the predominant purpose of these is to implement a checkpoint response, these

phosphorylations are detected under normal conditions and this has been observed in

several publications (Harvey et al., 2004, Latif et al., 2004, Niida et al., 2007) without

investigation into the purpose of its presence. Several recent publications have proposed

important roles for Chk1 under normal conditions during G2 phase and mitosis (Schmitt et

al., 2006, Wilsker et al., 2008, Matsuyama et al., 2011). Combined, these data suggest that

the basal level of activated Chk1 present during an unperturbed cell cycle may be linked to as

yet unknown roles of Chk1 during a normal cell cycle.


87

The aim of this chapter, therefore, was to investigate the presence of Chk1

phosphorylation/activation throughout the different stages of the cell cycle in order to

determine if phosphorylated Chk1 (pChk1) levels remained constant or fluctuate throughout

the cell cycle.

3.2 RESULTS

3.2.1 Activated Chk1 is present during an unperturbed cell cycle.

To demonstrate the activation of Chk1 in the cell cycle with and without DNA damage we

used immunoblotting and FACS analysis. Chk1 phosphorylation and thereby activation was

strongly induced by G2 phase checkpoint activators Etoposide or Ultraviolet Radiation

(UV(R)) treatment, which are known to require Chk1 activity and increased Chk1

phosphorylation (Ser317 and Ser345) (Figure 3.1 A and B). Increased Ser345 phosphorylation

is only shown here in response to UV(R) treatment, however it is known to be activated in

response to DNA damage by Etoposide and, although not shown here, it is demonstrated

later in this study. The increased phosphorylation of Chk1 corresponded to a G2 checkpoint

arrest, confirmed by propidium iodide (PI) FACS (Figure 3.1 A and B, right hand panels) and

an increase of the G2 phase marker, CDK1 phosphorylated on Tyr15 (PY15) (only shown here

with Etoposide treatment). Phosphorylated Chk1 Ser317 is also clearly seen in the untreated

asynchronously growing HeLa cells in both experiments (Figure 3.1 A and B). Unlike the DNA

damage response, the presence of Chk1 phosphorylation did not appear to correspond to

activation of the G2 checkpoint arrest, as seen by FACS profiles (Figures 3.1 A and B, left hand


88

panels). These data could suggest that this low level of phosphorylated Chk1 was not

sufficient to induce an arrest but was present as part of a surveillance mechanism to rapidly

respond to any damage when it occurs. Alternatively, perhaps this phosphorylated Chk1 is

not active, with additional modifications required for either the activation or maintaining

Chk1 in an inactive state.

Although active Chk1 was phosphorylated on both Ser317 and Ser345, the phospho-Ser317

antibodies (denoted here after as pChk1) were used predominantly throughout this study as

the antibodies were more reliable than the phospho-Ser345 antibody available. As activated

Chk1 was present during an unperturbed cell cycle, as seen in Figure 3.1, the activation

status of Chk1 throughout the different stages of the cell cycle was investigated (Figure 3.2).

Samples of HeLa cells were collected after synchronization with a double thymidine

synchrony block and collected at the indicated time points and analysed by FACS and

immunoblotting for pChk1 and other cell cycle markers. FACS analysis was also performed to

determine the cell cycle phase of each sample collected. Immunoblotting using PY15, as a

marker of G2 phase, and cyclin B, which becomes elevated in mitosis, was used to further

distinguish between these two phases of the cell cycle. pChk1 (the upper band as indicated

by an arrow) (Figure 3.2) was detected at release from the thymidine arrest (0 h), increasing

in S phase and then persisting through G2 phase (6-10 h), before declining as cells began to

enter mitosis (10-12 h), as indicated by the reduction in PY15 at these later time points

(Figure 3.2). Thus, Chk1 appears to be phosphorylated in S and G2 phase cells without


89

Figure 3.1 pChk1 is present in asynchronously growing cells but strongly induced in response to

DNA damage.

A. HeLa cells were treated with 10 J/cm2 of UV(R) B (Ultraviolet radiation B) and collected 24 h

after exposure. Samples were collected for immunoblotting and FACS analysis. Cells were

immunoblotted for phospho-Chk1 Ser317 and Ser345, Chk1 and the loading control, α-

tubulin. For FACS analysis, cells were stained with propidium iodide for DNA content.

B. HeLa cells were treated with 1 µM Etoposide and collected 24 h after drug addition.

Samples were collected for immunoblotting and FACS analysis. Cells were immunoblotted

for pChk1 Ser317, Chk1 and PY15 antibody which detects CDK1 phosphorylation on tyrosine

15 and is a marker of G2 phase accumulation. α-tubulin was used as a loading control. For

FACS analysis cells were stained with propidium iodide for DNA content.


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Figure 3.2 Active Chk1 is present during an unperturbed cell cycle.

HeLa cells were synchronised using a double thymidine block release protocol (See section 2.2.2).

Cells were collected at indicated time points after blockade release (0-14 h). Samples were divided

and analysed by immunoblotting or FACS. For immunoblot analysis lysates were immunoblotted for

activated Chk1 phosphorylated on S317 (pChk1; top band as indicated by arrow), total Chk1 protein,

inactive CDK1 (PY15) and cyclin B (top band as indicated by arrow). Immunoblotting of PCNA was

used as a loading control. Asynchronous cells were used as a control. PI FACS analysis was carried out

and cell cycle populations were quantified with FlowJo software. The percentage of cell cycle phases

for each time point is represented in the graph.


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any damage present. Phospho antibodies were used to detect the activation status of other

important G2 checkpoint proteins Chk2 (pChk2 T68, data not shown) and activated p38MAPK

(pp38) and no changes were detected across the cell cycle (Figure 3.2). PCNA levels were

used as a loading control.

To confirm the presence of elevated pChk1 in G2 phase cells, and remove any potential

artifacts that may arise from a thymidine-induced synchrony, the specificity of the pChk1

siRNA was first checked. Immunoblot data was used to show the efficiency of Chk1 the siRNA

to deplete Chk1 and pChk1. The results showed an approximate 50% reduction in Chk1 levels

(Figure 3.3 A, lane 1 vs. lane 3). The effects of Chk1 siRNA on pChk1 levels and checkpoint

function was assessed by treating cells with siRNA prior to treatment with Etoposide, which

was used to induce a G2 checkpoint arrest. This clearly demonstrated the effectiveness of

the siRNA to reduce pChk1 levels and checkpoint activation (Figure 3.3 A, lanes 2 and 4.

pChk1 indicated by the arrow). The depletion of Chk1 resulted in the failure of the G2

checkpoint arrest allowing cell cycle progression and mitotic entry. This was demonstrated by

the larger presence of the G1 peak seen by FACS. An increased presence of mitotic cells was

evident by the typical mitotic rounded phenotype in the Chk1 siRNA treated sample

compared to the control (Figure 3.3 C). The reduction in PY15 levels accordingly

demonstrated that the number of G2 phase cells decreased with Chk1 siRNA and inversely

the number of mitotic cells increase as shown by cyclin B levels (Figure 3.3 A, lane 2 vs. 4)

To analyse further the presence of pChk1 in an unperturbed cell cycle, pChk1 S317 antibody

was tested for specificity using immunofluorescene. Cells treated with Chk1 siRNA were

stained with pChk1 Ser317 antibody and DAPI for DNA. The images in Figure 3.4 A


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Figure 3.3 Chk1 siRNA effectively removes Chk1 and its typical functions.

A. Exponentially growing HeLa cells were transfected with 30 nM Chk1 siRNA or a nonsense (N)

control siRNA. Cells were then either treated or not with Etoposide and collected 24 h after

drug addition. Samples were then immunoblotted for pChk1 (top band of image) and Chk1

to determine knockdown, and also markers of cell cycle progression, PY15 and cyclin B.


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PCNA was used as a loading control. The arrow indicates the pChk1 band and the lower

band is a non-specific band detected by the antibody.

B. Samples from A were fixed and stained with propidium iodide to assess DNA content by

FACS.

C. Before harvesting the cells for A and B, images of the cells were captured using a Zeiss

inverted microscope. Arrows are used to indicate mitotic cells.


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demonstrate the specificity of the antibody to be used for the analysis of the future

experiments in Figure 3.4.

With the specificity of the pChk1 antibody established, quantitative image analysis of

asynchronously grown cultures from a number of different cell lines stained with the same

pChk1 antibody and DAPI was conducted. The analysis was used to confirm the synchrony

data above and to also give a true representation of pChk1 levels during the cell cycle in the

absence of any artefacts that the thymidine synchrony may have induced. This was

performed using a Cellomics VTi ArrayScan, a high content imaging system which assessed

and quantified the immunofluorescent staining of individual cells. The cells were seeded in a

96 well plate format, fixed and immunostained for pChk1 and DAPI. The individual cell cycle

stages were determined using DNA intensity which increases from G1 to S and G2/M and was

based on DAPI staining. The fluorescent intensity of pChk1 staining was analysed for each cell

and plotted against the DNA content. Controls stained with secondary antibody alone were

used to determine the background fluorescent intensity of the cells. Quantitative image

analysis was performed on 5,000 - 10,000 cells of asynchronously growing HeLa, Neonatal

Foreskin Fibroblasts (NFF), A2058 (melanoma cell line; data not shown), HT144 (an ATM

deficient melanoma cell line), and FO2 cells (an ATR defective Seckel syndrome cell line). The

analysis displayed here is from the HeLa data (Figure 3.4 B and C), the analysis of the other

cell lines is similar to the HeLa data and can be found in Appendix 1 (Figure 10.1). The

histogram is based on the DNA content of the cells analysed for pChk1 intensity. The

histograms were used to set parameters of the cell cycle phases. The scatter plot is a

comparison between the pChk1 (dark data points) levels and the background (light data


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Figure 3.4 Cellomics data confirms the presence of pChk1 in an unperturbed cell cycle.

A. HeLa cells were seeded on to coverslips and were transfected with nonsense control siRNA

(NS) or with Chk1 siRNA (Chk1 si). 24 h post transfection cells were fixed with MeOH and

stained for pChk1 Ser317 (Bethyl Ab)(red), γ-tubulin (green) and DAPI (blue).

B. HeLa cells were seeded into a 96 well plate, fixed with MeOH and stained with or without

pChk1 Ser317, detected by secondary antibody Alexia 555, and DAPI for DNA content. Cells

stained with Alexia 555 only, were used to determine background intensities of the assay.

The dot plot displays pChk1 intensities (dark data points) and background intensities (light

data points). The histogram is based on DNA content quantified using DAPI staining of DNA

content. 2n represents G1 phase cells and 4n represents G/M cells. .

C. Quantification of the results was carried out by defining the cell cycle phases based on the


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DNA intensity histograms, as in B. The graph represents the mean plus three standard

deviations; the range represents the 95 percentile. The results were corrected for

background intensities. The Etoposide treatment was used as a control and accordingly

results in elevated pChk1 levels.


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points) levels. All cell lines showed the same trend where pChk1 levels increase with cell

cycle progression, with the highest intensity evident during G2/M (Figure 3.4 C). For HeLa cell

data, it is evident that the G2/M phase population (cells with 4 n DNA content) is

approximately one and a half fold higher than the G1 population (cells with 2n DNA content).

Etoposide treated cells were used as a positive control and accordingly showed higher levels

of pChk1. This analysis by Cellomics confirmed that pChk1 is present within unperturbed cells

and this is independent of any synchrony effects. The immunoblot and Cellomics data in

Figures 3.2 and 3.4 together demonstrate that pChk1 is readily detectable in unperturbed S

and G2 phase cells. Although the immunoblot data showed the highest level of pChk1 during

S phase (Figure 3.2) this may be a consequence of the chemical synchronization as the

Cellomics data showed the highest levels during G2/M (Figure 3.4).

3.2.2 ATR is responsible for Chk1 phosphorylation in an unperturbed cell cycle during G2

phase.

In response to damage, Chk1 is activated by ATR, but there is also evidence of overlap

between the ATM and ATR checkpoint signalling pathways (Gatei et al., 2003, Jazayeri et al.,

2006). When thymidine synchronised HeLa cells were treated with the ATM/ATR inhibitor,

caffeine, at release from the synchrony arrest, the levels of pChk1 detected in early S phase

time points (0-4 h) are relatively unaffected (Figure 3.5 A), however G2 phase pChk1 was

completely absent (compare 6 and 8 h time points of Figures 3.2 and 3.5 A; these

experiments were performed in parallel). Although caffeine addition clearly delayed S phase


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Figure 3.5 Chk1 phosphorylation by ATR in G2 phase is required for timely mitotic entry.

A. HeLa cells were treated as in Figure 3.2, however at the same time as release of cells from

thymidine block, 5 mM of caffeine was added to all thymidine treated samples. As in 3.2,


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samples were collected for immunoblotting and FACS. Lysates for immunoblotting were

immunoblotted with pChk1 S317 (pChk1), Chk1, cyclin B, PY15 and PCNA. PI FACS analysis

was carried out and cell cycle populations were quantified with FlowJo software. The

percentage of cell cycle phases for each time point is represented in the graph.

B. HeLa cells were synchronised in G2 phase and left untreated (CONT) or treated with 5 mM

caffeine (CAFF) or 2.5 µM Chk1i. Cells were then followed by time lapse microscopy,

visualised every 15 minutes and then scored for timing of entry into mitosis. Mitosis was

scored once the rounded mitotic phenotype became visible. The accumulative data is

displayed here as a percentage. The result was confirmed by conducting the experiment in

triplicate and the best representation was chosen for publishing.

C. HeLa cells were synchronised in G2 phase and left untreated (CONT) or treated with Chk1

siRNA or 2.5 µM Chk1i overnight. Cells were then followed by time lapse microscopy,

visualised every 15 minutes and then scored for timing of entry into mitosis. Mitosis was

scored once the rounded mitotic phenotype became visible. The accumulative data is

displayed here as a percentage. The result was confirmed by conducting the experiment in


D. FO2 cells were treated as in A. FO2 cells treated with 2 mM of Hydroxyurea (HU) for 24 h

were collected and used as a positive control of Chk1 activation.


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it accelerated the kinetics of entry into mitosis, with an abrupt decline in PY15 levels

occurring between 8 and 10 h after release in the presence of caffeine as opposed to 10 h in

the parallel experiment (compare Figure 3.2 with Figure 3.5 A). This acceleration was also

observed with caffeine or a Chk1i addition to thymidine synchronised G2 phase cells and

followed by live cell time lapse microscopy (Figure 3.5 B). Cells were assessed for their timing

of entry into mitosis, based on the appearance of the rounded mitotic morphology. A similar

result was observed with Chk1 siRNA treatment and Chk1i, although the effect of Chk1 siRNA

was weaker (Figure 3.5 C). Interestingly, Chk1i was consistently less efficient than caffeine at

accelerating mitotic entry which could be an effect of caffeine’s multiple targets (Bode and

Dong, 2006, Bode and Dong, 2007). This data along with additional data from the laboratory

produced using ATM and Chk2 inhibitors (data not shown) demonstrates that ATR-Chk1

signalling is present active during normal G2 phase and contributes to controlling the timing

of mitotic entry.

To confirm the presence of pChk1 as a specific ATR dependent phosphorylation event,

synchronised FO2 cells from an ATR deficient Seckel Syndrome patient (with functional ATM)

were analysed for their pChk1 levels during cell cycle progression (Figure 3.5 D). As indicated

by FACs, FO2 cells did not synchronise with thymidine as efficiently as HeLa cells which could

be due to a slower growth rate. However cyclin B and PY15 levels demonstrate the G2 and M

phase time points. pChk1 was detected in G1/S phase (0-2 h) and in the positive control

(hydroxyurea treated (HU)), however pChk1 was not detected in S and G2/M phase samples

(4-10 h) (Figure 3.5 D). Therefore Chk1 is normally activated by ATR during the S and G2

phases of the cell cycle and its activity is required to control the timing of mitotic entry.


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3.2.3 pChk1 localises to the centrosomes during mitosis.

A large congregation of proteins localise to the centrosomes during late G2 and mitosis,

including ATM and ATR (Zhang et al., 2007). During interphase, pChk1 Ser317 appears more

predominant in the nucleus, present in punctuate nuclear structures, with also weaker

dispersed staining of the cytoplasm (Figure 3.6 A). From the start of prophase when the

chromosomes begin to condense and the mitotic spindle forms, pChk1 S317 appeared to

localise to the centrosomes. This localisation is maintained until metaphase, but was not

present from telophase through to the completion of mitosis, cytokinesis (Figure 3.6 A). Co-

staining with α-tubulin, a marker of the mitotic spindle which originates from the

centrosome, confirmed the centrosomal localisation of pChk1 Ser345 (Figure 3.6 B).

Therefore, both Ser317 and Ser345 phosphorylated forms of Chk1 were present at the

centrosomes during mitosis. Chk1 siRNA, validated previously, was used to determine the

specificity of the pChk1 antibody in the immunofluorescent analysis of cells. HeLa cells

treated with or without Chk1 siRNA were immunofluorescently stained for pChk1 Ser345, α-

tubulin (used to identify the mitotic spindle and the location of the centrosomes) and DAPI to

identify the DNA. Treatment with Chk1 siRNA ablated the centrosomal staining in mitotic

cells (Figure 3.6 B). To confirm the specificity of pChk1 Ser345 staining at the centrosomes,

cells were treated with caffeine, an inhibitor of ATM/ATR. pChk1 staining was essentially lost

with caffeine treatment (Figure 3.6 C), demonstrating that this is an ATR dependent

phosphorylation event.


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Figure 3.6 pChk1 is present at the centrosomes in G2 and Mitosis of an unperturbed cell cycle.

A. Asynchronous HeLa cells were grown on poly-L-lysine coated coverslips, fixed with methanol

and then stained with pChk1 S317 antibody (pChk1) (Bethyl Ab)(red) and DAPI for DNA

(blue). Arrows indicate centrosomal localisation.

B. HeLa cells treated with or without 30 nM Chk1 siRNA were fixed with methanol and stained

for pChk1 Ser345 (pChk1) (green), α-tubulin (red) and DAPI for DNA (blue). Images were


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taken of mitotic cells to demonstrate the presence of Ser345 at the centrosomes. Arrows

indicate the loss of Ser345 localisation at the centrosomes seen with Chk1 siRNA treatment.

C. To demonstrate whether the centrosomal localisation of pChk1 is due to ATR dependent

phosphorylation, HeLa cells were treated for 10 h with or without 5 mM of caffeine and

then treated as in A. These cells were co-stained for pChk1 Ser345 (pChk1) (green) and DAPI

for DNA (blue) only.

D. Asynchronously growing HeLa cells were fixed with methanol and stained for pChk1 S317

(pChk1) (red), γ-tubulin (green) and DAPI for DNA (blue). Arrows indicate the G2 phase

separating centrosomes.


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Although pChk1 localises at the centrosomes during mitosis (Figure 3.6) the question remains

regarding the localization of the pChk1 during G2 phase. Staining with pChk1 whilst co-

staining with the centrosome marker, γ-tubulin, confirms the localisation of pChk1 S317 at

the centrosomes earlier than mitosis during late G2 phase. Late G2 phase was indicated by

the beginning of centrosome separation whilst cells displayed no chromosome condensation

(Figure 3.6 D, indicated by arrows). It also shows that pChk1 is predominantly nuclear during

G2 phase. Therefore Ser317 is able to localise at the centrosomes during both G2 and

mitosis.

3.3 DISCUSSION

The critical importance of Chk1 to the normal cell cycle is shown by the embryonic lethality

of Chk1-/- knockout mice (Takai et al., 2000), as well as the mitotic catastrophe and death

which occurs upon depletion of Chk1 (Niida et al., 2005). The presence of activated pChk1 in

unstressed cells has now been reported by a number of groups (Walworth et al., 1993,

Walworth and Bernards, 1996). It has now been shown to be present at the centrosome in

the absence of DNA damage signals, where it specifically regulates Cdc25B to control normal

mitotic progression (Capasso et al., 2002, Jackson et al., 2002, Kramer et al., 2004, Loffler et

al., 2006, Wilsker et al., 2008, Tibelius et al., 2009). The data presented in this chapter, in

combination with published work demonstrates clearly that pChk1 S317 and S345 can

localise to the centrosomes (Wilsker et al., 2008). Whilst pChk1 S317 was localised to the

centrosomes it was also present in the nucleus during an unperturbed cell cycle. Although

no specifically localized pool of pChk1 was targeted, inhibition of Chk1 in G2 phase drives


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premature entry into mitosis, demonstrating that the activated pChk1 functions during G2 to

control the timing of entry into mitosis.

ATR knockout results in embryonic lethality, but this is not the case for ATM-/- where

offspring are viable, suggesting ATM is non-essential to the unperturbed cell cycle (Beamish

et al., 1996, Brown and Baltimore, 2000, Shiloh, 2001). Here, it was found that the function

of Chk1 during G2 phase is ATR dependent. The role of ATR was determined using several

methods including the use of FO2 cells, caffeine treatment and ATR knockdown.

Unfortunately, HeLa cells treated with ATR pSuper shRNA to deplete ATR levels were not

viable. FO2 cell data did show strong evidence of ATR involvement in the S and G2 phase

Chk1 activation, and although we did not have access to the ATR complemented cells to

formally confirm ATR involvement the caffeine data confirmed the involvement of ATM/ATR

in this role. Although the data could not rule out any involvement of ATM, other data in the

laboratory has shown that a specific ATM inhibitor had little effect on the rate of mitotic

entry, and that effect was possibly to slow progression into mitosis. The lack of any

detectable level of activated Chk2 present in an unperturbed cell cycle was further evidence

for a lack of involvement of ATM in normal G2 phase progression. The levels of pChk1

strongly correlate to the extent of the G2 checkpoint induced. From the Cellomics data

(Figure 3.4 B and C) it is evident that, of the cell lines tested, HeLa cells were most strongly

arrested in G2 phase and accordingly have the highest intensity readings of pChk1 compared

to the other less affected cell lines (Figure 3.4 B and C compared to Appendix 1). The

phosphorylation of Chk1 during normal G2 phase does not induce a checkpoint arrest as seen

with pChk1s role during the DDR but inhibition or depletion of Chk1 in an unperturbed cell


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cycle, as it does in G2 checkpoint activated cells, drives premature mitotic entry, implying

that Chk1 can control the timing of mitotic entry even in the absence DNA damage. Based on

the Cellomics data, the levels of pChk1 are highest during G2/M but the data also show a

strong presence of pChk1 during S phase. In fact, the synchronization data show the highest

levels of pChk1 during S phase however this is most likely an effect of synchronization and

accounts for the discrepancy between the two sets of data. A role for pChk1 in S phase is

plausible, as mutation of Chk1 Ser317 to Alanine results in DNA replication abnormalities in

the absence of exogenous stress (Wilsker et al., 2008). Chk1 is also activated in response to

replication fork stalling. This is expected to be noticeable in tumour cell lines that have a

predisposition to DNA damage that would lead to replication stress even during an

unperturbed cell cycle (Mohindra et al., 2002, Brooks et al., 2012). In addition, Chk1 has the

ability to regulate Cdc25A degradation during the S phase checkpoint resulting in the

inhibition of CDK2 and the suppression of DNA replication by phosphorylation of serines 123,

178, 278, and 292 (Zhao et al., 2002b, Goloudina et al., 2003, Hassepass et al., 2003,

Sorensen et al., 2003). The same phosphorylations of Cdc25A by Chk1 in response to the S

phase checkpoint can also occur in an unperturbed cell cycle and therefore this may be the

same mechanism for regulation by Chk1 of normal DNA replication (Sorensen et al., 2003).

Chk1 function has been reported to be of importance during mitosis. This is based on the

observations that depletion of Chk1 by siRNA results in a metaphase arrest and activation of

the spindle assembly checkpoint due to the presence of lagging chromosomes and therefore

the inability of these chromosomes to align in a metaphase plate (Tang et al., 2006).

However, it is more likely that these mitotic effects of Chk1 inhibition are a consequence of


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premature mitotic entry caused by Chk1 inhibition. In experiments conducted by others

members of the Gabrielli laboratory on the specific consequence of Chk1 inhibition during G2

phase, it was found to have relatively little effect on the immediate mitosis, but the

continued presence of Chk1 inhibitor affected proceeding mitoses and resulted in similar

mitotic defects to those observed with inhibition of Chk1 in S phase (Brooks et al., 2012;

unpublished data from the Gabrielli Laboratory). This is similar to reports demonstrating that

either deletion of Chk1 or inhibition of Chk1 with a small molecule inhibitor in S phase

induced premature and aberrant mitosis that was dependent on Cdc25B (Niida et al., 2005,

Loffler et al., 2006). The aberrant mitotic phenotype was identical to the “paraspindle”

phenotype originally observed by this lab with Cdc25B overexpression (Gabrielli et al., 1996).

The centrosomal pool of Chk1 has been reported to contribute significantly to regulating

mitotic entry. Initial activation of cyclin B/CDK1 occurs at the centrosomes and leads to the

full activation of cyclin B/CDK1 ( De Souza et al., 2000b, Jackman et al., 2003). Premature

activation of cyclin B/CDK1 can result in premature mitotic entry which can be due to the

inability of Chk1 to regulate Cdc25B at the centrosome (Kramer et al., 2004, Loffler et al.,

2006). Enhanced mitotic entry caused by inhibition of Chk1 could be a result of lost negative

regulation of Plk1 (Tang et al., 2006), and/or regulation of Cdc25B at the centrosomes

(Schmitt et al., 2006). Because pChk1 is detected at the centrosome it is likely that Chk1 is

active during G2 phase to undertake these roles. Despite identifying the requirement of

active Chk1 to control mitotic entry, the exact phosphorylation status and the regulation of

Chk1 that is required for this control of G2/M progression is currently unknown.


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The functional studies of Ser317 and Ser345 phosphorylation have uncovered potentially

differing roles for these sites in normal G2/M regulation. Ser345 was not detected on the

centrosomes during G2, whereas we and others have found Ser317 localised at the

centrosomes during G2 phase (Wilsker et al., 2008). Mutational analysis also confirms that

phosphorylation governs the localization and, potentially, roles of Chk1. The Ser345 Alanine

mutant, Ser345Ala, is unable to dissociate from the chromatin and causes mitotic

catastrophe, whereas Ser317Ala prevents mitotic catastrophe and is able to still localize to

the cytoplasm, like wildtype Chk1 (Niida et al., 2007). Therefore, the function and localization

of Chk1 is strongly governed by site specific phosphorylation events. Although the DNA

damage response causes a rapid phosphorylation of both Ser317 and Ser345, the distinct

roles of each phosphorylation event in an unperturbed cell cycle may provide further insights

into the roles and regulation of Chk1 in normal cell cycle progression.

The surprising finding here was that pChk1 levels present in G2 phase appeared to be

relatively reduced as cells entered mitosis, although the levels of pChk1 Ser317 at the

centrosomes during mitosis remained relatively stable until anaphase. Based on Chk1’s role

during the DNA damage response, it would be expected that the presence of phosphorylated

Chk1 would result in a G2 phase arrest and an inability for cells to enter mitosis. However,

from the current data and that of others, it is clear that the presence of pChk1 does not stop

cells from entering mitosis, contrary to the checkpoint function of Chk1. Therefore, I

hypothesize that, although pChk1 Ser317 is associated with activity, during mitosis it only

indicates that the kinase is activated, but to allow mitotic entry further modifications, most

likely further phosphorylation event/s, must occur during G2 phase to inhibit the activity of


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Chk1. These hypothetical inhibitory phosphorylations may also result in changes in

localization and functioning of Chk1 away from its normal substrates and regulators. The

formulation of the hypothesis was supported by a publication that identified inhibitory

phosphorylation of Chk1 by cyclin B/CDK1 that occur in mitosis (Shiromizu et al., 2006), and

these phosphorylations affected the localization of Chk1 (Enomoto et al., 2009). The

disruption of Chk1 localisation by phosphorylation events has been shown previously with

Chk1 Ser317 and Ser345 mutants. These mutants show disruption to the 14-3-3 binding and

therefore localization within the nucleus and potential activity of the Chk1 (Jiang et al., 2003,

Dunaway et al., 2005). Alternatively, if inhibitory phosphorylations are not the answer then

another hypothesis would be a pChk1 threshold which allows for pChk1 present at low levels

to monitor cell cycle integrity without causing a G2 phase arrest.

It is common to assume Chk1 activity based on the phosphorylation of Chk1 on Ser317 and

Ser345. As already mentioned, these phosphorylation sites govern the localization and

functions of Chk1. However, the effects of mutation of Ser317 and Ser345 phosphorylation

sites would suggest a more complex picture. Analysis of the phosphorylation mutants of

these sites suggests that they are not only catalytic activation sites. Ser317 and Ser345

mutation to Alanine (Ser317Ala and Ser345Ala) cause defects to the G2 checkpoint due to

the inability of the activation caused by phosphorylation on Ser317 and Ser345 (Capasso et

al., 2002) but, confusingly, analysis of the phospho mimicking Aspartate mutants of Ser317

and Ser345 (Ser317D and Ser345D) showed they had a similar kinase activity to Alanine

mutants (Capasso et al., 2002, Gatei et al., 2003). This data suggests that the Ser345D mutant

affects the 14-3-3 binding required for Chk1 localisation and activity (Capasso et al., 2002),


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and this is also a likely explanation for the Ser317D mutant. Although these mutants have

been used to determine the importance of these phosphorylations in response to damage,

the phosphorylations themselves may not be good determinants of Chk1 activity and

function during the G2/M transition. To address this issue and to determine the true activity

of Chk1 during the G2/M transition, Chk1 kinase assays were attempted. Although a range of

conditions for the kinase assay were investigated, all methods failed to produce detectable

levels of kinase activity in vitro, even when assaying Chk1 kinase activity upon DNA damage

treatment. Therefore the assays were abandoned and the activity of Chk1 during the G2/M

transition cannot be conclusively determined. Premature mitosis caused by removal or

inhibition of Chk1 and the likelihood of inhibitory phosphorylations during mitosis suggests

that Chk1 activity is required during G2 phase and is not required during mitosis. However,

the latter is not fully established and raises some important questions.

To determine how Chk1 phosphorylation events influence mitotic entry would be difficult

based on the published data regarding Chk1 mutants, as well as the possibility of potential

inhibitory phosphorylation events. Further investigation and better tools are required to

study Chk1’s role in G2/M phase. The presence of pChk1 during an unperturbed cell cycle is

now a vastly published fact, but at the time of this study it was a novel finding that ATR-Chk1

signalling and their kinase activities were required to regulate normal mitotic entry. It was

also a novel finding that Chk1 Ser345, as well as Ser317, could also localize to the

centrosomes during mitosis. We were also able to establish that pChk1 can localize at the

centrosomes earlier during G2 phase. The centrosomal localization of Chk1 may later provide

insight into further roles of the kinase during this time. Collectively, we have added to an


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ever increasing base of evidence that investigates Chk1s normal G2/M role. Now that this

role for active Chk1 has been confirmed we believe that Chk1 could tie into another

regulatory G2/M pathway that is being currently investigated within the laboratory. This

pathway is the cyclin A/CDK2 dependent G2 delay pathway and in the next chapter its role

and some of its known mechanisms and targets will be presented.

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Chapter 4

Characterising the cyclin A/CDK2 dependent G2

phase delay

Chapter 4 – Characterising the cyclin A/CDK2 dependent G2 delay

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4.1 INTRODUCTION

Understanding the mechanisms regulating normal cell cycle control is essential to determine

how defects can contribute to tumourgenesis. The cyclin A/CDK2 complex is required

through S and G2/M phases of the cell cycle to regulate S phase progression and the timely

progression into mitosis. Cyclin A/CDK2 has an established S phase role (Hu et al., 2001), and

is the predominant cyclin/CDK complex activated during G2 phase and reaches its peak

activity in this same phase (Gu et al., 1992, Pagano et al., 1992, Rosenblatt et al., 1992)

although its contribution to G2 phase progression is not well understood. In addition, cyclin A

is not degraded by the APC/C (Anaphase promoting complex/cyclosome) proteasome

complex until prometaphase, suggesting a role for cyclin A/CDK2 during G2/M phase

progression.

The role of cyclin A/CDK2 during G2 phase of the cell cycle is controversial due to a lack of

phenotype in Cdk2 knockout mice (Berthet et al., 2003, Ortega et al., 2003) and CDK2 siRNA

treatment of cancer cells (Tetsu and McCormick, 2003). On the other hand, microinjection of

cyclin A targeted antibodies delayed tissue culture cells from entering mitosis (Clarke et al.,

1992) and induction of dominant negative CDK2 after G1 phase in synchronised U2OS cells

blocked cells in G2 phase (Hu et al., 2001). Furthermore, cells from Drosophila mutants

lacking cyclin A also arrested in G2 phase after the maternal cyclin A had been exhausted

(Knoblich and Lehner, 1993). Additionally, microinjection of active cyclin A/CDK2 caused a

more rapid mitotic entry (Furuno et al., 1999). This latter data suggest that cyclin A/CDK2

acts as a rate limiting component of G2/M progression.


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Cyclin B/CDK1, the predominant mitotic cyclin/CDK, is activated after cyclin A/CDK2 in late

G2 phase and drives mitotic entry. The initial activation of cyclin B/CDK1 occurs at the

centrosomes and within the cytoplasm and establishes positive feedback loops to further its

activation (De Souza et al., 2000a, Jackman et al., 2003). This involves the phosphorylation of

cyclin B/CDK1 on its cytoplasmic retention site leading to nuclear import and the

phosphorylation of Cdc25 family members to enhance interaction with critical phosphatases

required to dephosphorylate negative regulatory phosphorylation sites, all of which sustain

its full activation (Yang et al., 1998, Hagting et al., 1999, Yang et al., 2001, Peter et al., 2002,

Bulavin et al., 2003a, Bulavin et al., 2003b, Walsh et al., 2003, Margolis et al., 2006b). The full

activation of cyclin B/CDK1 results in chromosome condensation after initial formation of the

mitotic spindle (Li et al., 1997, Hagting et al., 1999).

Cyclin A/CDK2 has been shown as an established regulator of cyclin B stabilisation, but not

activation, through indirect inhibition of APC/C which is required for cyclin B/CDK1

destruction (Lukas et al., 1999b). There is also evidence from Xenopus studies that show

cyclin A/CDK is capable of stimulating cyclin B/CDK1 dependent mitotic events such as

microtubule nucleation at the centrosomes (Buendia et al., 1992). Collectively the data

establishes a clear, yet not fully understood role for cyclin A/CDK in the control of mitotic

machinery and mitotic entry.

Cyclin/CDK complexes also have very specific roles in checkpoint regulation. Checkpoints act

to prevent the cell from cycling while it is affected by specific stresses, halting the cell cycle

to allow repair of any damage that may have been incurred. This requires the sustained


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inhibition of cyclin/CDK complexes which is achieved by specific checkpoint signalling

pathways. The apical checkpoint kinases ATM/ATR are activated in response to specific DNA

damage and initiate checkpoint signalling upon their recruitment to the sites of the damage.

ATM and ATR activate downstream targets Chk2 and Chk1 (Checkpoint kinase 2 and 1),

respectively, and the loss of this signalling affects the cell’s ability to arrest at the checkpoint.

Chk1 has been studied extensively and its function during the G2 phase DNA damage

checkpoint is to phosphorylate and inhibit the Cdc25 family which causes sustained inhibition

of cyclin B/CDK1, therefore maintaining the checkpoint arrest and blocking mitotic entry

(Furnari et al., 1997). Chk1 has a central role during the G2 phase DNA damage checkpoint,

and loss of Chk1 or its activator ATR, results in failure of checkpoint arrest.

Although cyclin B/CDK1 is essential for mitotic entry, it is both G2 phase cyclin A/CDK2 as

well as cyclin B/CDK1 that are held inactive during a G2 phase DNA damage checkpoint to

ensure cells cannot enter mitosis (Goldstone et al., 2001). The stable expression of functional

cyclin A/CDK2 causes abrogation of the G2 phase checkpoint and CDK2 null cells were unable

to exit the checkpoint (Chung and Bunz 2010, Walker et al., 1995). Collectively, the data

implies that not only does cyclin A/CDK2 appear to be a rate limiting protein of normal G2/M

progression in non-murine cells but it may also be required in some capacity for the G2 phase

checkpoint.

Although the importance of cyclin A/CDK2 during G2 phase is evident, its precise role and the

pathways involved remain unclear. This study establishes the role of cyclin A /CDK2 in G2

phase. We have identified the requirement of cyclin A /CDK2 for both normal and checkpoint


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G2/M transition and also determined a mechanism of action for cyclin A/CDK2 in normal

G2/M transition. The mechanism identified is the requirement of cyclin A/CDK2 to correctly

coordinate cyclin B/CDK1 activity and mitotic events such as chromosome condensation and

microtubule nucleation.

4.2 RESULTS

4.2.1 Cyclin A depletion delays G2 phase progression and entry into mitosis

To examine the role of cyclin A/CDK2 in G2/M progression three different cyclin A siRNA

target sequences, each with differing efficiencies were used to deplete cyclin A (Figure 4.1 A).

After 24 h treatment with cyclin A siRNA the protein levels were reduced by up to 70% and

cell cycle profiles demonstrated that cyclin A depletion resulted in an increase in the G2/M

phase population when compared to controls (Figure 4.1 B). The G2/M phase populations of

cyclin A A1 and A3 siRNA treated cells increased by two fold, whereas the A2 siRNA which did

not deplete cyclin A as effectively (Figure 4.1 A) showed a smaller accumulation of cells in

G2/M phase. To differentiate between G2 and M phase populations a marker of G2 phase,

CDK1 phosphorylation on tyrosine 15 (PY15), was used. PY15 levels accumulate in cyclin A

siRNA treated cells compared to the controls (Figure 4.1 C), demonstrating the predominance

of the G2 phase population of cells over the mitotic population of cells. Etoposide, known to

induce a strong G2 arrest was used as a positive control and accordingly caused a strong

increase in PY15 levels (Figure 4.1 C).


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Figure 4.1 Cyclin A siRNA causes a G2 phase delay.

A. Exponentially growing HeLa cells were transfected with 50 nM of one of three siRNA

directed against cyclin A (A1, A2, or A3), a nonsense (N) siRNA, or treated with lipofectamine

alone (C). Cells were harvested after 24 h, and immunoblotted for cyclin A and CDK2. Data

contributed by Dr Leonore De Boer. Blot Quantification was conducted by Vanessa Oakes

using ImageJ software. All treatments were compared to the intensity of the CDK2 loading

control and standardised to the control (C).

B. FACS profiles of the DNA content of cells transfected with A1, A2, A3 cyclin A siRNA,

nonsense (Non) siRNA or mock transfected (Con) at 24 h, as in A. The percentage of cells in

G1, S and G2/M phases are shown in descending order in the top right corner of each

profile. Data contributed by Dr Leonore De Boer.

C. HeLa cells were treated with Etoposide (1 μM) (Etop) for 16 h to arrest the cells in the G2

phase, or transfected with either cyclin A A1 siRNA (A1), nonsense siRNA (N) or mock

transfected (C) and harvested at 24 h. Lysates were immunoblotted for PY15. Data

contributed by Dr Leonore De Boer.


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D. Mock transfected (Con) and cyclin A (A1, A2 or A3) siRNA transfected HeLa cells were

synchronised with thymidine and followed by time lapse microscopy. Cells were scored for

timing of entry into mitosis; >100 cells were counted for each treatment. This data is

representative of four independent experiments and the best representation was chosen

for publishing. Data is presented as cumulative mitotic index scored from the time (h) after

release from thymidine block. Data contributed by Dr Leonore De Boer.


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To determine the effect of cyclin A on G2 phase, HeLa cells were synchronised in early S

phase by a thymidine block and the time required to enter mitosis was assessed using live

cell time lapse microscopy. Cyclin A siRNA transfected HeLa cells showed a delay of entry into

mitosis by 3-4 h compared to the control siRNA transfected cells. This delay was observed

with all three cyclin A siRNAs (Figure 4.1 D) and confirms a G2 phase delay as a specific result

of cyclin A knockdown.

4.2.2 Inhibition of CDK2 with a specific inhibitor delays G2 phase progression into mitosis.

Cyclin A forms a complex with binding partner CDK2 in S phase and this complex reaches

maximal activity in G2 phase (Goldstone et al., 2001, De Boer et al., 2008). By inhibiting the

catalytic activity of cyclin A’s binding partner, CDK2, we can determine whether the G2 delay

seen with cyclin A siRNA was due to the specific loss of the cyclin A /CDK2 complex. The

contribution of CDK2 to the G2 phase delay was examined using a specific small molecule

inhibitor of CDK2, Ro09-3033 (CDK2i). This is a highly selective inhibitor of CDK2 (IC50 20

nM), and inhibits by binding irreversibly to the ATP binding site of CDK2 (Stead et al., 2002).

To ensure only G2 phase CDK2 was inhibited, synchronised HeLa cells were treated with two

different concentrations of CDK2i in early G2 phase (7 h post release). Mitotic entry was

assessed by live cell time lapse microscopy. When compared to the control, both CDK2i

treated samples delayed any entry into mitosis by at least 4 h (Figure 4. 2 A). 0.3 µM of CDK2i

delayed progression to a lesser extent than 0.5 µM, producing an expected dose dependent

result. This delay was similar to that caused by cyclin A knockdown (Figure 4.1 D) as would

be expected if CDK2’s binding partner was cyclin A. This experiment also demonstrated the


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role of cyclin A/CDK2 in regulating entry into mitosis was a G2 phase specific, and from here

forth this delay is termed as the cyclin A/CDK2 dependent G2 phase delay.

Published work has shown that the CDK2i used displayed 10 fold in vitro selectivity for CDK2

compared to CDK1, compared to its in vivo selectivity which may not be as strong due to the

higher dose required to inhibit CDK2 in cells (2 μM in vivo compared to its in vitro IC50 of 20

nM) (Stead et al., 2002). However, the G2 phase delay seen with the lower concentration of

CDK2i (Figure 4.2 A; 0.3µM CDK2i) may suggest otherwise. In addition, cyclin A forms an

active complex with CDK1 for a short period in late G2/beginning of mitosis, which could

conceivably be involved in regulating mitotic entry also (Mitra et al., 2006). Therefore, it was

important to determine the specificity of the CDK2i.

Recombinant Cdc25B phosphatase was used to determine the effect of CDK2i on CDK2

and/or CDK1 kinase activity. The activity of CDK2 was completely lost after CDK2i, which

could not be recovered in vitro by the addition of recombinant Cdc25B. CDK2i treatment also

blocked G2/M phase cyclin B1/CDK1 activation but this was recovered with addition of

recombinant Cdc25B, demonstrating the selectivity of the CDK2i for the cyclin A/CDK2

complex. The lack of CDK1 activity in CDK2i treated cells (without Cdc25B addition) is similar

in the control lane (Figure 4.2 B; compare lanes 5 and 1). Although this indicates that CDK2

doesn’t affect CDK1 activity, there is a small level of CDK1 activity in the control sample and

suggests that there may be some regulation of CDK1 activity by CDK2, which may contribute

to mitotic entry.


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Figure 4.2 CDK2i causes a G2 phase delay which was due to specific CDK2 inhibition.

A. Thymidine synchronised HeLa cells were treated without (Con; closed diamonds) or with 0.3

µM (open squares) and 0.5 µM CDK2i (Ro09-3033) (open triangles). CDK2i was added at 7 h

after synchrony release (early G2 phase) then followed by time lapse microscopy. The

cumulative mitotic index was scored. The result was confirmed by conducting the

experiment in triplicate and the best representation was chosen for publishing.

B. Thymidine synchronised HeLa cells were treated with 0.5 µM Ro09-3033 (CDK2i) at 6 h after

release from thymidine arrest (early G2 phase). 8 h post thymidine release/2 h after

addition of CDK2i, cells were harvested in late G2 phase for analysis. G2 phase, control cells,

was collected at the same time as CDK2i treated samples (8 h post thymidine release).

Mitotic samples (M) were collected approximately 9 h after thymidine release. Cells were

collect for FACS and activity assays. Samples for FACS analysis were collected and treated as


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detailed in Chapter 2. The DNA content of these samples is shown and demonstrates the

synchronicity of each sample. Cells were lysed and CDK2 and CDK1 immunoprecipitates

were assayed for histone kinase activity either without or with pre-incubation with

recombinant Cdc25B. Kinase data contributed by Dr Brian Gabrielli.

C. Asynchronous HeLa cells were either untreated (Con), treated with the indicated

concentrations of CDK2i (Ro09-3033), or 50 µM roscovitine (rosco) for 10 h then harvested.

The cells were analysed by MPM2 staining of the mitotic population. Quantification of the

FACS data was conducted to determine the percentage of the mitotic population in each

treatment. Total number of cells counted per condition was 10 000. Data was combined for

three independent experiments. Error bars represent the standard deviation.

D. Thymidine synchronised HeLa cells were treated without (circles) or with 1 µM (open

triangle), 2 µM (closed triangles) or 4 µM (squares) CDK2i (PHA-533533). CDK2i was added

at 7 h after synchrony release (early G2 phase) and followed by time lapse microscopy. The

cumulative mitotic index was scored. The result was confirmed by conducting the

experiment in triplicate and the best representation was chosen for publishing.


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To further demonstrate the specificity of this CDK2i Ro09-3033, its effects were compared to

the CDK 1 / 2 inhibitor, roscovitine. The CDK inhibitors were added to asynchronously

growing HeLa cell cultures for 10 h and the mitotic index was assessed by staining for the

mitotic phosphoprotein marker MPM2 and its level determined by FACS. This revealed that

Ro09-3033 treated cells are able to enter mitosis, whereas roscovitine treatment blocked

cells from entering mitosis (Figure 4.2 C). The lack of reduction of mitotic cells was

representative of the transient delay caused by CDK2i treatment (Figure 4.2 A), compared to

a complete block in mitotic entry with CDK1 inhibition, demonstrating that the delay was not

a consequence of direct inhibition of CDK1. CDK2i caused an increase in the proportion of

mitotic cells when compared to the control, however this was due to a delay of cells in

mitosis which has been previously observed with cyclin A depletion/CDK2 inhibition (Fig 4.1

D; data not shown, (Beamish et al., 2009).

During the course of the study, another inhibitor of CDK2, PHA-533533, was acquired

(Pevarello et al., 2004) (IC50 = 37 nM). The drug was tested for its ability to delay mitotic

entry like that seen with Ro09-3033 (Figure 4.2 A). The selectivity of this compound for CDK2

was previously tested and shown to have a 5 fold higher selectivity for CDK2 compared to

CDK1 (Pevarello et al., 2005b). A dose dependent delay in mitotic entry was observed, the

most effective being the highest concentration of PHA-533533 (4 µM), which delayed mitotic

progression by 2 h (Figure 4.2 D). This second CDK2i was acquired after Ro09-3033 became

unavailable for further studies, and was used throughout the second half of this study.

Collectively the data in Figure 4.2 demonstrates the G2 delay as a consequence of the G2

phase specific inhibition of cyclin A/CDK2.


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4.2.3 Cyclin A/CDK2 regulates the timing of centrosomal cyclin B/CDK1 activation

The activation of cyclin B/CDK1 in prophase is initiated at the centrosomes and within the

cytoplasm which in turn drives nuclear translocation and full activation of nuclear cyclin

B/CDK1. Activation of cyclin B/CDK1 at the centrosomes correlates with increased

microtubule nucleation required for the formation of the mitotic spindle, which occurs prior

to nuclear envelope disassembly (De Souza et al., 2000a, Jackman et al., 2003). To determine

whether the centrosomal pool of cyclin B/CDK1 was affected by cyclin A/CDK2 depletion or

inhibition, microtubule foci representing the increased microtubule nucleation at the

centrosomes were assessed in HeLa cells stably expressing a Cherry-tagged α-tubulin by live

cell time lapse microscopy. Cells treated with cyclin A siRNA formed two microtubule foci,

however these foci persisted on average approximately 40 - 50 minutes longer than in

controls (normally <30 min, Figure 4.3 A) before forming a mitotic spindle (Figure 4.3 B and

C). This result was also observed in cells treated with low dose CDK2i Ro09-3033 but to a

lesser extent than cyclin A siRNA treatment (Figure 4.3 D). Quantification of synchronized

CDK2i treated samples demonstrates a delay of microtubule foci formation by approximately

3 h (Figure 4.3 E) which corresponds to the 3-4 h delay seen in the cyclin A/CDK2 dependent

G2 delay (Figures 4.1 D and 4.2 A), indicating that although the centrosomal mitotic events

preceded the nuclear events by up to 1 h, this was still delayed when compared to the

controls. The data indicate that cyclin A/CDK2 may be required for the coordinated timing of

the centrosomal and nuclear activation of cyclin B/CDK1. Further observation of cyclin A

siRNA treated cells revealed that the formation of microtubule foci was also delayed which is

evident by the condensation of the microtubules that eventually form the mitotic spindle


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Figure 4.3 Cyclin A siRNA and CDK2i disconnect the coordinated activation of centrosomal and

nuclear pools of cyclin B/CDK1.

A. HeLa cells stably expressing cherry α-tubulin were transfected with a control, nonsense,

siRNA and synchronised with thymidine. 5 h post release for the synchrony cells were

followed by time lapse video microscopy. Cells were imaged every 10 min by live cell time

lapse microscopy (minutes are indicated in the top left corner of each imaged timeframe).

The Cherry filter set was used to produce fluorescent images of α-tubulin displayed.


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B. HeLa cells stably expressing cherry α-tubulin were transfected with cyclin A (A3) siRNA and

synchronised with thymidine. 5 h post release for the synchrony cells were followed by time

lapse microscopy. Cells were imaged every 10 min by live cell time lapse microscopy

(minutes are indicated in the top left corner of each imaged timeframe).

C. Cells shown in A and B where quantitated for the length of time taken between microtubule

foci appearance until spindle formation. Over 100 cyclin A siRNA (A1) and over 60 nonsense

(N) and mock transfected (C) cells with detectable microtubule foci were counted.

Experiment conducted by Vanessa Oakes and analysis by Dr Brian Gabrielli.

D. Thymidine synchronised cherry α-tubulin HeLa cells were treated with or without 0.3 µM

CDK2i (Ro09-3033) in G2 phase (7 h post release) and followed by time lapse microscopy.

Cells were imaged every 15 min. Cells were quantitated for the length of time starting from

the appearance of microtubule foci until spindle formation. Approximately 100-150 CDK2i

and over 80 nonsense (C) treated cells with detectable microtubule foci were counted.

Experiment conducted by Vanessa Oakes and analysed by Dr Brian Gabrielli.

E. Thymidine synchronised cherry α-tubulin HeLa cells were treated with 0.3 µM CDK2i (Ro09-

3033) in G2 phase (7 h post release) and followed by time lapse microscopy. Cells were

imaged every 15 min. Over 80 control (diamonds) and CDK2i treated (triangles) cells with

detectable microtubule (Mt) foci/nucleation were counted and scored with respect to the

timing of initial foci formation. Only cells which displayed foci for at least one frame were

counted. The result was confirmed by conducting the experiment in duplicate and the best

representation was chosen for publishing. Experiment conducted by Vanessa Oakes and

analysed by Dr Brian Gabrielli.


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(Figure 4.3 A and B). Microtubule foci, like those shown in Figure 4.3 A, also stain for γ-

tubulin indicating they are centrosome containing structures required for the formation of

the mitotic spindle (data not shown). Therefore with loss/inhibition of cyclin A/CDK2, the

activation of cyclin B/cdk1 appears to be delayed which causes the delay in time taken for

the microtubule foci to form the mitotic spindle. This delay causes the consequent delay of

mitotic nuclear events including the activation of the total pool of cyclin B/cdk1.

Examination of this mitotic phenotype by immunofluorescent staining required the use of

markers of cyclin B/cdk1 activation and mitosis. During normal mitosis phosphorylated

histone H3 Ser10 (pH3) levels increase (Hendzel et al., 1997, Goto et al., 1999, Preuss et al.,

2003), and Mitotic Protein Marker (MPM2) by immunostaining is evident. pH3 is a marker of

chromosome condensation, while MPM2 decorates both the mitotic centrosomes and

condensed chromosomes. MPM2 recognizes phospho-epitopes present in mitotic proteins.

Although the precise spectrum of epitopes that MPM2 recognizes is not fully defined, a

major MPM2 recognition sequences contains phosphorylated threonine /serine residues

followed by a proline. These specific sites are phosphorylation sites for cyclin B/cdk1

(Stukenberg et al., 1997, Westendorf et al., 1994) but are not restricted to cyclin B/cdk1

alone. Activation of other mitotic kinases such as Polo like kinase 1 (Plk1) and Mitogen

activated protein kinase (MAPK) are indicated by MPM2 staining also (Alvarez et al., 1991,

Clark-Lewis et al., 1991, Gonzalez et al., 1991, Logarinho and Sunkel, 1998, Wang et al., 2007,

Vanderheyden et al., 2009). Both of these markers were used to assess the normal

occurrence of prophase events of microtubule foci and chromosome condensation.


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During a normal prophase, these markers detect chromosome condensation and MPM2 in

addition can detect formation of microtubule foci. As seen in Figure 4.4 A control (N) a

prophase cell is stained for pH3 in the nucleus, which indicates nuclear envelop breakdown

and microtubule foci are formed as indicated by α-tubulin staining. Depletion of cyclin A

causes a noticeable portion of cells with prophase-like microtubule foci (determined by -

tubulin staining) and lack nuclear staining normally seen with pH3 staining (Figure 4.4 A;

cyclin A (A1) siRNA treatment). This corresponded to an absence of chromosome

condensation normally detectable by MPM2 or pH3 staining (Figure 4.4 A and B).

Interestingly, MPM2 staining indicated that the microtubule foci were present without any

sign of chromosome condensation in cyclin A depleted cells (indicated by arrows in Figures

4.4 B). This suggested that the two prophase events, chromosome condensation and

microtubule focus formation, that are normally closely coupled (as shown to occur at the

same time in Figure 4.4 B control (N)) became uncoordinated in cyclin A depleted cells. To

describe this phenotype better we employed the use of the term ‘preprophase’. The

preprophase cells display microtubule foci formation, without any evidence of chromosome

condensation.

MPM2 staining, which identified both chromosome condensation and microtubule foci, was

used to score the preprophase phenotype (Figure 4.4 C). As indicated by the scoring, there

was a significant accumulation of preprophase cells in cyclin A depleted samples (Figure 4.4

B, A1 treated), where only the microtubule foci were stained. Although a small portion of

preprophase cells was present in control samples, the majority have a normal prophase


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appearance where MPM2 staining of condensed DNA was evident at the same time as

microtubule foci formation (Figure 4.4 B; N).

The detection of increased microtubule nucleation and MPM2 staining at the centrosomes in

cyclin A depleted cells suggested the presence of activated cyclin B/cdk1 at the centrosomes.

To determine the activation of cyclin B/cdk1, the specific cyclin B/cdk1 mitotic

phosphorylation of Mek1 on threonine 286 (pMek1 T286), was used as a marker

(Rossomando et al., 1994). All mitotic cells stained for pMek1 T286, with specific prophase

staining of microtubule foci and condensed chromosomes. pMek1 T286 stained the

microtubule foci in both control prophase cells and cyclin A depleted preprophase cells

(Figure 4.4 D; indicated by arrows), however it was not found in the nucleus of cyclin A

depleted preprophase cells and only became present in the nucleus once chromosome

condensation had occurred (Figure 4.4 D; observe the difference between the cells in the A1

treatment image). Roscovitine, the cdk 1 and 2 inhibitor, treatment caused an almost

complete loss of pMek1 T286 MPM2 nuclear and spindle pole staining (data not shown),

confirming the use of pMek1 T286 as a marker of cyclin B/cdk1 activity. These data confirm

that the delay of cyclin B/cdk1 activation seen in these experiments is a direct result of the

cyclin A/cdk2 loss/inhibition, although the centrosomal activation of cyclin B/cdk1 appears

less affected than the nuclear mitotic activation.

4.2.4 Cyclin A / CDK2 localises to the centrosomes in late G2 phase.

These surprising effects of cyclin A /CDK2 depletion/inhibition on microtubule nucleation at

the centrosome suggested that the complex itself may localise to the centrosomes to


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Figure 4.4 Cyclin A/CDK2 is required for coordinated cyclin B/CDK1 activity.

Asynchronously growing HeLa cells were transfected with 50 nM of cyclin A siRNA A1 and A3 or a

nonsense control (N). 24 h post transfection cells were stained for;


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A. phosphoSer10 H3 (pH3), α-tubulin and DAPI for DNA. pH3 staining of cyclin A siRNA treated

cells shows nuclear staining in a metaphase cell, whereas the adjacent prophase cell (indicated

with the arrowhead) shows microtubule foci formation by α-tubulin staining but no nuclear

pH3 staining.

B. MPM2, α-tubulin and DAPI for DNA. The exposure time for MPM2 is identical for both control

and cyclin A siRNA samples. Note the staining of microtubule foci in both control (indicated by

arrowheads adjacent to brightly stained nucleus) and cyclin A depleted cells by α-tubulin and

MPM2 staining. The lack of MPM2 nuclear staining signifies no chromosome condensation has

occurred in cyclin A depleted samples.

C. Cells from B. displaying prophase-like microtubule foci were scored for detectable chromosome

condensation. Cells displaying microtubule foci in the absence of chromosome condensation

were scored as preprophase (filled columns) and those that displayed both were scored as

prophase (open columns). This data represents the average and standard deviation of three

independent experiments. *P = 0.025, **P = 0.015.

D. pMek1 T286 (marker of cyclin B / CDK1 activity), α-tubulin and DAPI for DNA. pMek1 T286

stained the microtubule foci of all mitotic cells, shown by arrows. Nuclear pMek1 T286 was

present in control cells but was lacking in cyclin A siRNA treated samples, representative of no

chromosome condensation. Exposure times for pMek1 T286 were kept constant whilst imaging.

Scale bars represent 10 µm. Arrows indicate centrosomes.


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regulate cyclin B/CDK1 activity. It has previously been suggested that cyclin A can localize to

the centrosomes (Bailly et al., 1992, Pagano et al., 1992, den Elzen and Pines, 2001), although

minimal data have been presented to support these claims, and the localization of CDK2 at

the centrosomes was unknown. A mild permeabilisation of cells prior to fixing was used to

remove cytoplasmic CDK2 and cyclin A. This allowed the visualization of centrosomal CDK2

and cyclin A and its co-localisation with the centrosomal marker, γ-tubulin (Figure 4.5 A;

indicated by arrows). The centrosomal localization of cyclin A occurs during G2 phase once

the centrosome pair begins to separate but before microtubule nucleation occurs. CDK2

staining revealed that CDK2 also localizes to the centrosomes during the same time indicated

by the condensed chromosomes. Arrows are used to mark the separating centrosomes

(Figure 4.5 B). Cyclin A depletion revealed that the centrosomal localization of CDK2 was

dependent on cyclin A (Figure 4.5 B).

4.2.5 Cyclin A/CDK2 depletion or inhibition affects the exit from G2 / M checkpoint.

Implementation of a checkpoint arrest requires the inhibition of cyclin/CDK complexes to put

a ‘brake’ on cell cycle progression. The exit of cells from a G2 checkpoint requires the

removal of this braking system to promote mitotic entry. The inhibition of cyclin B/CDK1

activation is essential for the implementation of the G2 checkpoint, and its activation is

required for mitotic entry after the release from the checkpoint arrest. The involvement of

cyclin A/CDK2 in the G2 checkpoint had not been examined in any detail previously. In light

of the data in this study where we show the control of cyclin B/CDK1 activation and normal

mitotic entry by cyclin A/CDK2, it was assumed that cyclin A/CDK2 could have a similar role in


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Figure 4.5 Cyclin A/CDK2 localises to the centrosomes in late G2 phase.

A. Asynchronous growing HeLa cells were seeded onto glass coverslips and then fixed,

detergent permeabilised and stained with DAPI for DNA (blue), cyclin A (green) and γ-

tubulin (red). Image is of the prophase cell displayed was taken using a Zeiss Apotome

widefield microscope equipped with a CCD camera. Arrows indicate the position of

centrosomal staining.

B. HeLa cells were treated without (Con) or with 50 nM cyclin A (A1) siRNA. 24 h post

transfection cells were fixed, detergent permeabilised and stained with DAPI for DNA (blue),

CDK2 (green) and γ-tubulin (red). Images show the co-localisation of CDK2 at the

centrosomes in two different control cells and the loss of CDK2 from the centrosomes


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caused by cyclin A depletion. Arrows indicate the position of centrosomal staining. Images

were taken with a Zeiss Apotome widefield microscope equipped with a CCD camera


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mitotic entry from a G2 phase checkpoint arrest. Cyclin A/CDK2 was known to be inhibited

to implement the checkpoint, and a mutant of CDK2 that did not require Cdc25-dependent

activation was able to bypass the checkpoint (Goldstone et al., 2001, Chow et al., 2003)

suggesting that the activation of cyclin A/CDK2 might impact on the G2 phase checkpoint. In

addition, Cdc25B is essential for checkpoint exit (Goldstone et al., 2001, Bugler et al., 2006)

and its major role is the activation of G2 phase cyclin A/CDK2 (Goldstone et al., 2001),

supporting the notion that cyclin A/CDK2 activation is required for exit from the checkpoint

arrest.

The DNA damaging agent Etoposide was used to activate the G2 phase DNA damage

checkpoint by activation of ATR/Chk1 and downstream signalling. Caffeine, an ATM/ATR

inhibitor, was used to block this signalling and force cells to exit the G2 phase checkpoint

arrest. Cells were visualized by live cell time lapse microscopy to determine whether exit

from the checkpoint was affected by either cyclin A depletion or CDK2 inhibition. Control

cells exited the checkpoint and entered mitosis at a much faster rate than cells treated with

either cyclin A siRNA A1 or A3 (Figure 4.6 A) or CDK2i PHA-533533 (Figure 4.6 B). Treatment

with PHA-533533 caused a greater block in checkpoint exit which was most likely a result of

more complete inhibition of CDK2 as opposed to the partial knockdown of cyclin A. This

demonstrates the specific requirement of cyclin A/CDK2 for exit from a checkpoint arrest,

commonly referred to as checkpoint recovery. With cyclin A depletion or CDK2 inhibition, a

significant portion of cells underwent apoptosis which possibly indicated that cyclin A/CDK2

was also required for the continued viability of the damaged cells (data not shown).


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Figure 4.6 Cyclin A/CDK2 is required for exit from a G2 phase checkpoint arrest.

A. Mock transfected (N) and cyclin A (A1 or A3) siRNA transfected HeLa cells were treated with

Etoposide (1 µM) overnight to arrest cells in the G2 phase checkpoint. Cells were forced to

exit the checkpoint with addition of 5 mM caffeine (at time zero). Cells were analysed for

their mitotic entry using live cell time lapse microscopy directly after caffeine addition.

>100 cells for each treatment were counted for their timing of mitotic entry. The result was

confirmed by conducting the experiment in triplicate and the best representation was

chosen for publishing.


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B. HeLa cells were treated with Etoposide overnight to arrest them in a G2 phase checkpoint.

Cells were treated, 30 min prior to caffeine (5 mM) addition, with either DMSO (Cont) or

CDK2i at a concentration of either 2 μM or 4 μM of CDK2i (PHA-533533). Cells were

analysed for their mitotic entry using live cell time lapse microscopy. >100 cells per

treatment were counted for their timing mitotic entry. A significant proportion of CDK2i

treated cells died via apoptosis which may explain why the mitotic accumulation never

exceeded 30%. The result was confirmed by conducting the experiment in triplicate and the

best representation was chosen for publishing.


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We also determined whether the Chk1i (SB 218078) had a better ability to allow exit from

the checkpoint in either cyclin A depleted cells or CDK2 inhibited cells. Compared to caffeine

treatment, Chk1i treatment more strongly inhibited the exit of cyclin A depleted cells from

the checkpoint (Figure 4.7 A). Chk1i treatment of CDK2i treated cells also showed a dramatic

decrease in the number of cells entering mitosis (Figure 4.7 B). Like the caffeine experiment

of Figure 4.6 B, many cells underwent apoptosis. Control cells in which the central kinase,

Chk1 - which is required to implement the checkpoint – has been inhibited, enter mitosis

readily. The consequence of cyclin A/CDK2 depletion/inhibition on checkpoint exit is

noticeable when compared to the control. As Chk1i inhibition could not be used to overcome

the checkpoint in cyclin A/CDK2 depleted/inhibited cells it was evident that cyclin A/CDK2

has an important role in checkpoint exit.

4.3 DISCUSSION

This study confirms the importance of cyclin A/CDK2 during G2 phase of the cell cycle. A

number of studies now confirm the role of cyclin A/CDK2 in regulating the timing of cyclin

B/CDK1 activation and entry into mitosis (Furuno et al., 1999, Hu et al., 2001, Mitra and

Enders, 2004, Fung et al., 2007, Gong et al., 2007a, ). In addition, this study has contributed a

functional role of cyclin A/CDK2 in the timing and co-ordination of the centrosomal and

nuclear activation of cyclin B/CDK1 and mitotic events.

Several publications have now determined a role for cyclin A/CDK2 during G2 phase,

however, the proposed mechanisms for its G2 phase role vary. One group reported that


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Figure 4.7 Cyclin A/CDK2 is required for exit from a G2 phase checkpoint arrest.

A. Control transfected (NS) and cyclin A (A1) siRNA transfected HeLa cells were treated with

Etoposide (1 µM) overnight to arrest cells in the G2 phase checkpoint. Cells were forced to

exit the checkpoint with either addition of 5 mM caffeine or 1 μM Chk1i (SB 218078). Cells

were analysed for their mitotic entry using live cell time lapse microscopy. >100 cells for

each treatment were counted for their timing of mitotic entry and the cumulative data is

shown here. The result was confirmed by conducting the experiment in triplicate and the

best representation was chosen for publishing.


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B. HeLa cells were treated with Etoposide overnight to arrest them in a G2 phase checkpoint.

Cells were treated either with 2 μM or 4 μM of CDK2i (PHA-533533) 30 min prior to addition

of 1 μM of Chk1i (SB 218078). Cells were analysed for their mitotic entry using live cell time

lapse microscopy. >100 cells per treatment were counted for their timing mitotic entry and

the cumulative data is shown here. A significant proportion of CDK2i treated cells died by

apoptosis which may explain why the mitotic accumulation never exceeded 20%. The result

was confirmed by conducting the experiment in triplicate and the best representation was



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cyclin A depletion by shRNA (short hairpin RNA) caused an increase of Wee1 activity along

with a decrease in Cdc25C and Plk1 activity (Fung et al., 2007). However, they reported that

only the depletion of Wee1 bypassed the cyclin A/CDK2 dependent G2 phase delay.

Consequently they found that increased Wee1 activity caused the inhibition of CDK1, and

delayed mitotic entry which reflects the results shown here of delayed cyclin B/CDK1

activation caused by cyclin A siRNA or CDK2i addition. The type of analysis carried out

however does not exclusively determine that Wee1 activity is enhanced during the specific

cyclin A/CDK2 dependent G2 delay. Therefore whether the timing of enhanced Wee1 activity

correlates to the delay in cyclin B/CDK1 activity, as seen with cyclin A depletion/CDK2

inhibition, is not known. More recently, cyclin A/CDK2 was shown to directly phosphorylate

Wee1 (Li et al., 2010). Although this phosphorylation was not shown to affect the activation

of Wee1, it does affect its localization. Normally Wee1 enters the cytoplasm upon mitotic

entry (Heald et al., 1993, Baldin and Ducommun, 1995, Katayama et al., 2005) and without

cyclin A/CDK2 phosphorylation Wee1 remains in the nucleus, therefore affecting cyclin

B/CDK1 activation and consequently mitotic entry (Li et al., 2010). This phosphorylation

event strongly suggests a role for Wee1 in the cyclin A/CDK2 dependent G2 phase delay.

Another group reported that cyclin A removal by siRNA did not affect the activation of cyclin

B/CDK1 (Gong et al., 2007b). The biochemical analysis of cyclin B/CDK1 activity assays used in

this study was most likely not sensitive enough to detect the short delay seen with cyclin A

removal. The advantage of my analysis, which uses markers of cyclin B/CDK1 activity, is that

it is single cell analysis and gives a better representation of the exact timing of cell cycle

phases and events affected by cyclin A removal.


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Activation of cyclin B/CDK1 occurs at the beginning of mitosis. Active cyclin B/CDK1 is first

detected at the centrosome during prophase before the full activation of the remaining

cytoplasmic and nuclear pools occurs. The time between the initial activation at the

centrosomes and the remainder of cyclin B/CDK1 is no more than 20-30 minutes (De Souza et

al., 2000a, Kramer et al., 2004). Depletion or inhibition of cyclin A/CDK2 dissociated this

normally tight link between centrosomal and nuclear mitotic events and cyclin B/CDK1

activation. How cyclin A/CDK2 regulates the timing of cyclin B/CDK1 activation remains

unknown. As mentioned previously, the activation of Wee1 could be a contributing factor

(Fung et al., 2007), as could the role of cyclin A which targets cyclin B into the nucleus to

promote nuclear envelop breakdown (Gong et al., 2007a). Another possible G2/M regulator

that could be involved in the cyclin A/CDK2 dependent G2 delay is Plk1. The activity of Plk1

was shown to be diminished in cyclin A depleted cells but when over-expressed,

constitutively active Plk1 did not overcome the G2 delay caused by cyclin A shRNA treatment

(Fung et al., 2007). Plk1 is a major regulator of G2/M progression regulating Wee1 stability

(Watanabe et al., 2004, Watanabe et al., 2005), Aurora A (Seki et al., 2008a, van Leuken et

al., 2009), and regulation of APC/Cdh1 (Hansen et al., 2004) and thereby the degradation of

many important mitotic proteins. Although over-expression of Plk1 did not affect the cyclin

A/CDK2 dependent G2 phase delay, it does not fully exclude the involvement of Plk1. The

activity of Plk1 is greatly dependent on its localization established by its Polo box domain

(PBD). Binding to the PBD is governed by the recognition of phospho-epitopes primed by

other kinases (Lee et al., 2008). So, even though over-expressed Plk1 could not overcome the

cyclin A/CDK2 delay, this could be due to the inability of Plk1 to localize and therefore


143

function correctly. Therefore, determining if Plk1 localisation is affected by cyclin A depletion

or CDK2 inhibition would be necessary to determine Plk1’s involvement in the cyclin A/CDK2

dependent G2 phase delay.

Cdc25B is an activator of cyclin B/CDK1 and is required for its initial activation at the

centrosomes. Aurora A phosphorylates Cdc25B at the centrosome just prior to Cdc25B

activation of cyclin B/CDK1 at the centrosome (Dutertre et al., 2004). As cyclin A siRNA and

CDK2i treatment caused a delay in cyclin B/CDK1 activation at the centrosome, this could

suggest that cyclin A/CDK2 is regulating an Aurora A/Cdc25B dependent event that is

required for the centrosomal activation of cyclin B/CDK1 (Cazales et al., 2005). Aurora A has a

prominent role during mitosis and in particular is required for correct spindle formation

(Andrews et al., 2003, Schmitt et al., 2006, Hoar et al., 2007, Sardon et al., 2008, Cowley et

al., 2009, Asteriti et al., 2011). In cyclin A/CDK2 depleted/inhibited cells the mitotic spindle

forms correctly. Therefore Aurora A is not a likely target of cyclin A/CDK2 and is not

contributing to the cyclin A/CDK2 dependent G2 phase delay. However, Aurora A dependent

regulation of Cdc25B may be responsible for overcoming the delayed activation of the

centrosomal cyclin B/CDK1 caused by cyclin A/CDK2 loss. This could further explain the time

difference between the activation of centrosomal cyclin B/CDK1 and remaining pools of

cyclin B/CDK1. The latter would not be Aurora A dependent.

The mechanism by which cyclin A/CDK2 contributes to the exit from a checkpoint arrest is

examined further in the following chapters. However, it is likely that the mechanism is

similar, if not the same, as the role of cyclin A/CDK2 during normal G2/M phase progression.


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Some recent data has shown the importance of CDK2 in the G2 checkpoint. In this report it

was shown that double knockout p53 and CDK2 cells are unable to arrest at the G2

checkpoint (Chung and Bunz, 2010) whereas p53 wildtype cells reinforce the checkpoint.

This can be achieved in many ways including p53 repression of cyclin A and cyclin B

(Desdouets et al., 1996, Innocente et al., 1999, Levesque et al., 2005, Frum et al., 2009) or

inhibition of CDK1 via p21 (Xiong et al., 1993, Harper et al., 1995, Bunz et al., 1998,

Satyanarayana et al., 2008, Lossaint et al., 2011) so that cells cannot enter mitosis. Therefore

the role of cyclin A/CDK2 during the G2 checkpoint is dependent of p53. In the model HeLa

cell system used here, p53 and Rb activities are defective due to the human papilloma virus

(HPV) but there is some evidence of minimal p53 activity in these cells which may be

sufficient enough to reinforce the checkpoint (Matlashewski et al., 1986, May et al., 1991,

Johnson et al., 2002). Other checkpoint proteins such as ATM and ATR do not seem to

reinforce the checkpoint. As treatment with caffeine demonstrates that the mechanism is

independent of ATM and ATR signalling. It is likely that the reported role for Cdc25B in the

checkpoint exit may be contributing to the required activation of cyclin A/CDK2.

Further investigation has identified several mechanisms for cyclin A/CDK2 action in the G2/M

checkpoint. One group has shown the regulation of several mitotic regulators, Cdc25A, Chk1,

cdc6 and ATRIP by cyclin A/CDK2 in response to the checkpoint activation (Chung and Bunz,

2010). Although there is conflicting data that suggests Cdc25A is the regulator of cyclin

A/CDK2 (Timofeev et al., 2009), our laboratory demonstrated that Cdc25B is the major

regulator of cyclin A/CDK2 (Gabrielli et al., 1997), and expression of a CDK2 mutant that does

not require Cdc25-depedent activation bypasses the checkpoint arrest (Goldstone et al.,


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2001), further supporting the role for cyclin A/CDK2 in checkpoint recovery. Based also on

the fact that cyclin A/CDK2 activation is required for checkpoint exit, this may indicate that

cyclin A/CDK2 could target a feedback loop that could involve indirect activation of Cdc25B.

During normal G2/M progression there is evidence that cyclin A/CDK2 can control its own

activation by the inhibition of its inhibitor Wee1 (Li et al., 2010). Therefore it seems likely

that cyclin A/CDK2 activity is required for mitotic progression from a checkpoint and that it

may stimulate its own activation and cyclin B/CDK1 activation by regulating Wee1. Another

possibility would be through its reported regulation of stimulating FoxM1 transcription

(Wierstra and Alves, 2006, Laoukili et al.,2008b, Alvarez-Fernandez et al., 2010). FoxM1 is

responsible for the transcription of several mitotic kinases such as Plk1 and Cdc25B, whose

presence is required for cells to exit a checkpoint (Costa, 2005, Alvarez-Fernandez et al.,

2010,). This provides evidence that cyclin A/CDK2 may stimulate its own positive feedback

loop by controlling the activation of FoxM1. This agrees with other data that reveals minimal

cyclin A/CDK2 activity must be retained during a G2 checkpoint, so as to allow checkpoint

recovery through activating FoxM1 and enhancing transcription of important checkpoint exit

genes (Laoukili et al., 2008b, Alvarez-Fernandez et al., 2010). Interestingly, Cdc25B

expression is also regulated by FoxM1. This implies that the activation of some cyclin A/CDK2

by Cdc25B is required for its subsequent activation of FoxM1 to allow exit from a G2

checkpoint.

Here I have demonstrated that cyclin A/CDK2 regulates the timing of cyclin B/CDK1 activation

and entry into mitosis. It also regulates the connection between the timing of the

centrosomal and nuclear mitotic events. In addition, it has a novel role in regulating exit


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from the G2 phase checkpoint arrest. Interestingly ATM/ATR or the Chk1i could not be used

to overcome the failure of cells to exit an ATR/Chk1 dependent checkpoint when

depleted/inhibited of cyclin A/CDK2. This may indicate that cyclin A/CDK2 is required to

relieve the cells from the checkpoint by somehow affecting the activity of Chk1 which is

required for the implementation of the checkpoint. In the next chapter I will further

investigate the possible mechanism by which cyclin A/CDK2 controls G2/M progression and

how this involves Chk1.

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Chapter 5

Chk1 is required to implement the cyclin A/CDK2

dependent G2 phase delay

Chapter 5 – Chk1 is required to implement the cyclin A/CDK2 dependent G2 delay

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5.1 INTRODUCTION

Cyclin A/CDK2 is a major regulator of G2 phase and its depletion delays G2 phase progression

(Fung et al., 2007). The delay is caused by a loss of coordination of key mitotic events which

is a result of delayed cyclin B/CDK1 activation. The mechanism by which cyclin A/CDK2

regulates cyclin B/CDK1 activation and subsequent progression into mitosis is unknown.

G2/M phase checkpoint mechanisms also regulate cyclin B/CDK1 and progression into

mitosis, and we have shown that cyclin A/CDK2 is required for exit from the G2 phase

checkpoint arrest, suggesting that cyclin A/CDK2 may regulate checkpoint proteins. As shown

previously in chapter 3, ATR-Chk1 has a role in the normal regulation of cyclin B/CDK1 and

G2/M transition suggesting that cyclin A/CDK2 may act by regulating this signalling pathway.

Chk1, the effector kinase of the G2 phase checkpoint, is activated by ATR on Ser317 and

Ser345 in response to checkpoint activation. These phosphorylations have also been

demonstrated to be essential for normal G2/M progression in an unperturbed cell cycle

(Kaneko et al., 1999, Zhao et al., 2002a, Zhao et al., 2002b, Sorensen et al., 2003). The

essential role of Chk1 is evident from the embryonic lethality caused by Chk1 knockout.

Although the name might imply that Chk1 and Chk2 are related they are in fact structurally

unrelated. However, they are both serine/threonine kinases activated in response to DNA

damage. Chk2 knockouts do not cause embryonic lethality and although Chk2 is stably

expressed throughout the cell cycle, Chk2 activation is not present in the absence of DNA

damage, unlike Chk1.


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Chk1 has a normal cell cycle, S phase checkpoint (Zhao et al., 2002b, Sorensen et al., 2003)

and a critical G2/M phase checkpoint function (Gatei et al., 2003, Xiao et al., 2003). siRNA

studies have demonstrated the important role of Chk1 in normally-timed mitotic entry,

where its removal causes a premature mitosis, chromosome misalignment and mitotic

spindle checkpoint activation (Tang et al., 2006, Zachos et al., 2007). Chk1 regulates Cdc25

and Wee1 activity (Peng et al., 1997, Kumagai et al., 1998, Zeng et al., 1998, Furnari et al.,

1999, Lee et al., 2001, Rothblum-Oviatt et al., 2001, Stanford and Ruderman, 2005, Schmitt

et al., 2006) and these targets are most likely responsible for the mitotic disturbances seen

with Chk1 siRNA treatment. Chk1 has specifically been shown to phosphorylate Cdc25B at

the centrosome to control the initial activation of cyclin B/CDK1 and therefore mitotic entry

(Schmitt et al., 2006). This published work, and work presented in Chapter 3, clearly

demonstrates Chk1 functions in controlling G2/M transition.

Thus, Chk1 and cyclin A/CDK2 appear to have similar roles in regulating the activation of

cyclin B/CDK1 and mitotic entry. The similarity of functions suggests that these two

important G2/M phase kinases are acting within the same pathway or adjacent pathways.

Data from the previous chapter, chapter 4, demonstrated that cyclin A/CDK2 may be

required to regulate Chk1 to release cells from a G2 Chk1-dependent checkpoint. In this

chapter I have used both Chk1 siRNA and inhibitors to determine whether Chk1 is involved in

the cyclin A/CDK2 dependent G2 delay.


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5.2 RESULTS

5.2.1 Chk1 contributes to the cyclin A/CDK2 dependent G2 delay.

CDK2 inhibition and cyclin A depletion caused a G2 phase delay in asynchronously growing

cells (Chapter 4, Figures 4.1 and 4.2). This G2 phase delay was reflected by an increase in the

G2/M population and an increase in PY15 levels and was reminiscent of the G2 phase delay

imposed in response to DNA damage, which ultimately inhibits the cdc25-dependent

activation of cyclin B/CDK1. To determine the mechanism of the G2 phase delay imposed in

response to cyclin A/CDK2 depletion/inhibition, the role and contribution of potential key G2

phase checkpoint proteins was investigated.

Immunoblot analysis of cyclin A siRNA treated cells revealed an accumulation of activated,

Ser317 phosphorylated Chk1 (pChk1) in these cells (Figure 5.1 A, lane 3 arrow indicates

pChk1). Quantification of pChk1 levels confirmed the significant increase in pChk1 levels seen

with cyclin A knockdown (Figure 5.1 A; bar graph). Cyclin A depletion caused the signature

accumulation of the G2 phase marker, PY15. Using Chk1 siRNA, as previously used in Chapter

3, revealed that the simultaneous knockdown of cyclin A and Chk1 unsurprisingly leads to a

reduction of pChk1 levels when compared to the cyclin A only depleted sample. This also

correlated to the reduction of PY15 levels, indicating a loss of the G2 phase delay (Figure 5.1

A, Lane 5). The Chk1 siRNA only treatment displayed similar levels of cyclin A and PY15 as the

controls (Figure 5.1 A; compare lanes 1 or 2 with lane 4). The level of cyclin A and Chk1

knockdown was equivalent when used either singularly or in combination and neither

resulted in destabilisation of the other protein.


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Figure 5.1 Chk1 depletion causes reversal of the cyclin A dependent G2 phase delay.

A. Asynchronous HeLa cells were either mock transfected (C) or transfected with control,

nonsense, siRNA (N), cyclin A siRNA (A1), Chk1 siRNA (Chk1) or cyclin A and Chk1 siRNA.

Cells were harvested after 24 h and analysed by immunoblotting for the activated pSer317


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Chk1 (pChk1), cyclin A (cyc A), Chk1, PY15, and PCNA as a loading control. The arrow

indicates the specific pChk1 band, whilst the asterisk indicates an unspecific band detected

by the pChk1 antibody. The lower band was not detected by the unphosphorylated Chk1

antibody and was not depleted by Chk1 siRNA indicating it as a non-specific band. The bar

graph shows the quantification of pChk1 levels in samples collected 24 h post transfection

with siRNA, cyclin A siRNA (A1 and A3) or nonsense siRNA (NS). Immunoblots were analysed

using ImageJ software. Results were formulated from 3 independent experiments. Double

asterisk indicates p value of <0.01.

B. Samples from A were analysed for their DNA content using PI FACS. The FACS data is of the

G2/M population from 3 independent experiments and was analysed by Cell Quest. Error

bars represent standard deviations.

C. HeLa cells treated with either nonsense (solid circles), cyclin A siRNA A1 (open circles), A1

and Chk1 siRNA (squares) or A1 and Chk1 inhibitor (triangles) were thymidine synchronised

and followed by time lapse microscopy 6 h after synchrony release. Cells were scored for

entry into mitosis, over 200 cells were counted in each case. This is a typical result from

three independent experiments. The result was confirmed by conducting the experiment in


D. Asynchronous HeLa cells were either mock transfected (C), transfected with 50 nM of

nonsense (N) or cyclin A siRNA (A1-A5). Etoposide treatment overnight (1 µM) was used as a

positive control. Lysates were collected 24 h post transfection and immunoblotted for

pChk1 (light exposure, top), pChk2 T68, Chk2, pp38, p38, cyclin A and PCNA as a loading

control. Arrows indicate bands of interest, whilst the asterisk indicates an unspecific band.

ImageJ was used to quantify band intensity.


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E. NFF cells were transfected with either nonsense siRNA (N) or cyclin A siRNA (A1). 24 h post

transfections cells were lysed and immunoblotted for pChk1 S317 (pChk1), cyclin A, PY15

and PCNA. Arrow indicates bands of interest, whilst the asterisk indicates an unspecific

band.


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FACS analysis of these samples correlated with the immunoblot data and revealed an

accumulation of G2/M phase cells with cyclin A siRNA treatment (Figure 5.1 B). There was

also a two fold increase in the MPM2 positive mitotic population with cyclin A depletion

(from 1.5 to 3%) (data not shown), but this small increase could not account for the large

increase in G2/M population observed with cyclin A depletion, indicating that the majority of

cells were in G2 phase (consistent with findings in Chapter 4). Co-depletion of Chk1 reduced

the G2/M phase levels to that seen in controls, confirming the reduction in G2 delay with co-

depletion observed by immunoblot analysis (Figure 5.1 A and B). Chk1 siRNA by itself did not

affect the proportion of G2/M phase cells (Figure 5.1 B). MPM2 staining demonstrated that

the M phase proportion of cells was relatively unchanged with any treatment other than

cyclin A siRNA alone treatment (data not shown). The mitotic accumulation seen with cyclin

A knockdown was shown to be a direct consequence of cyclin A/CDK2 knockdown/inhibition

which has been previously published (De Boer et al., 2008, Beamish et al., 2009).

To assess the potential involvement of Chk1 in the cyclin A dependent G2 phase delay,

synchronised HeLa cells were treated with siRNA and assessed for timing of entry into mitosis

by live cell time lapse microscopy (Figure 5.1 C). Cyclin A siRNA treated cells (open circles)

were delayed in their entry into mitosis compared to the control (C) (closed circles), as shown

previously (Chapter 4; Figure 4.1 D). When cyclin A depleted cells were treated in

combination with either Chk1 siRNA (squares) or with Chk1 inhibitor (Chk1i; SB 218078,

triangles) cells entered mitosis at a faster rate than cyclin A siRNA alone treated cells (Figure

5.1 C). Cells were treated with the Chk1i in early G2 phase (5 h after release) while Chk1

siRNA treatment caused Chk1 depletion from the beginning of the synchrony. Based on the


155

timing of Chk1 treatment, the inhibitor data shows that G2 phase Chk1 is likely to be

contributing to the cyclin A/CDK2 dependent G2 delay. Interestingly, Chk1 siRNA or inhibitor

treatment did not completely reverse the effects of cyclin A depletion, suggesting that there

could be other contributors to the cyclin A/CDK2 dependent G2 delay in addition to Chk1.

This could however be the result of incomplete Chk1 knockdown/inhibition. The results do

however, demonstrate a definite role for Chk1 in the cyclin A dependent G2 delay.

The activation status of other important checkpoint kinases, such as Chk2 and p38, that are

reported to impose a G2 phase delay in response to DNA damage were assessed in cyclin A

depleted cells. Cyclin A siRNAs A1-A3 were used as well as 2 additional cyclin A siRNA, A4 and

A5, all of which caused an efficient reduction in cyclin A levels (Figure 5.1 D). No change in

either endogenous Chk2 or p38 levels or the activated counterparts, using activation specific

phospho-antibodies p38 and pChk2 T68, was detected when comparing controls and cyclin A

depletion. However they were both elevated in the Etoposide treated positive control. The

increased pChk1 levels, and the lack of increase of other known G2 phase checkpoint

regulators indicated that only Chk1 was involved in the cyclin A/CDK2 dependent G2 phase

delay. An increase in pChk1 levels caused by cyclin A siRNA was also seen is NFF cells (Figure

5.1 E) and U20S cells (data not shown). Accordingly, further investigations focused on the

role of Chk1 during G2 phase regulation and its relationship with cyclin A/CDK2.


156

5.2.2 Inhibition of G2 phase CDK2 causes an accumulation of G2 phase cells and activated

Chk1.

To determine whether pChk1 also accumulated with direct inhibition of CDK2, cells were

treated with 0.5 μM of CDK2i (Ro09-3033) for 24 h and the level of pChk1 assessed.

Compared to an untreated control, the level of pChk1 was noticeably increased. When cells

were co-treated with CDK2i and Chk1i, the levels of pChk1 reduced to the same levels as that

seen in the control. In addition to the increase of pChk1, the levels of the G2 phase marker,

PY15, were also increased with CDK2i treatment and reduced by co-treatment with Chk1i

(Figure 5.2 A). The pChk1 and PY15 data demonstrated that CDK2i treatment, as with cyclin A

depletion, causes an accumulation of G2 phase cells that involves activated Chk1, and

indicates that normal G2 progression requires the cyclin A/CDK2 complex in some manner

regulating Chk1 activation and function.

To assess whether the G2 phase accumulation observed with CDK2i treatment was due to a

delay in G2 phase, and the degree of bypass of any delay with Chk1i co-treatment, time lapse

microscopy of synchronised HeLa cell cultures was performed. Thymidine synchronised HeLa

cells treated with CDK2i (PHA-533533) in G2 phase (6 h after synchrony release) displayed a

lengthened G2 phase by over 2 h (Figure 5.2 B). The delay was reduced by the co-treatment

of cells with Chk1i, with these cells entering mitosis at an almost identical rate as the control

cells (Figure 5.2 B). This recovery was more complete than that seen in Figure 5.1 C with

cyclin A knockdown. This data supports the conclusion that Chk1 was required to implement

the cyclin A/CDK2 dependent G2 phase delay. It does not however suggest as strongly as


157

Figure 5.2 CDK2 inhibition confirms a role for pChk1 Ser317 in the cyclin A / CDK2 dependent G2

delay.

A. HeLa cells were treated for 24 h with 0.5 µM CDK2i (Ro09-3033) then without or with Chk1i

(2.5 µM) for a further 5 h. Control cells (Con) were treated with DMSO, for the same total

time as CDK2i plus chk1i treated samples. Cells were harvested and analysed by

immunoblotting for activated Chk1 (pChk1), Chk1, PY15 and PCNA as a loading control.

B. HeLa cells were synchronised using the double thymidine method, at 6 h post the final

release when cells were entering G2 phase they were either untreated or treated with 2 µM

CDK2i (PHA-533533) . 1 h after CDK2i addition, Chk1i (2.5 µM) was added to one of the

CDK2i treated samples and then analysed by time lapse microscopy. The cumulative mitotic

index was scored. The result was confirmed by conducting the experiment in triplicate and

the best representation was chosen for publishing.


158

Figure 5.1 C that other pathways may be involved in the cyclin A/CDK2 dependent G2 phase

delay.

5.2.3 Chk2 siRNA cannot overcome the cyclin A/CDK2 dependent G2 phase delay.

As Chk1 and Chk2 appeared to have overlapping functions in terms of imposing a G2 phase

delay, it was important for this study to specifically define any potential involvement of Chk2

in the cyclin A dependent G2 phase delay. We were previously unable to detect any changes

in the level of activated Chk2, Chk2 phosphorylated on Thr68 (pChk2) in cyclin A depleted

cells (Figure 5.1 D). To investigate the possible contribution of Chk2 to the cyclin A/CDK2

dependent G2 phase delay three independent Chk2 siRNA were developed from published

sequences (Gire et al., 2004, Astuti et al., 2009, Seror et al., 2009) and tested on

asynchronously growing or Etoposide treated HeLa cells (Figure 5.3 A). Chk2 levels showed

that c3 was the most effective siRNA at reducing Chk2 levels. pChk2 levels were very low in

asynchronously growing cultures (Figure 5.3 A lanes 1-4) but there appeared to be a clear

reduction of pChk2 with c1 and c3 siRNA. Because of the low levels of asynchronous pChk2 it

was more ideal to observe pChk2 levels after Etoposide treatment. Etoposide treatment in

Chk2 siRNA treated cells showed a significant reduction in pChk2 levels (Figure 5.3 A). Of the

three Chk2 siRNA used (c1-c3), c3 was the most potent in reducing pChk2 and Chk2 levels in

both asynchronous and Etoposide treated cells.

In further experiments Chk2 siRNA c3, which appeared to be the most efficient at reducing

Chk2, was used to determine whether depleting Chk2 affected the cyclin A/CDK2 dependent

G2 delay. Synchronised HeLa cells co-depleted of cyclin A and Chk2 by siRNA were followed


159

Figure 5.3 Chk2 does not have a role in the cyclin A dependent G2 phase delay.

A. Exponentially growing HeLa cells were transfected with 90 nM of one of three siRNA

directed against Chk2 (c1, c2, or c3) or a nonsense (N) siRNA. After transfection cells were

either treated with or without 1 µM Etoposide overnight. Samples collected 24 h after siRNA


160

treatments were immunoblotted for pChk2 T68 to determine knockdown and PCNA as a

loading control.

B. HeLa cells were transfected with control, nonsense, siRNA (N) (circles), cyclin A siRNA (A3)

(triangles), nonsense and Chk2 siRNA c3 (upside down triangles), or A3 and Chk2 siRNA c3

(vertical lines). 24 h after transfection cells were synchronised with thymidine and followed

by live cell time lapse microscopy from 6 h after thymidine release. Cells were scored for

entry into mitosis; over 200 cells were counted in each case. This is a typical result from

three independent experiments. The result was confirmed by conducting the experiment in


C. After completion of the experiment in B, cells were collected and immunoblotted for siRNA

targeted proteins cyclin A and Chk2 to confirm knockdown. PCNA was used as a loading

control. Quantification of immunoblots was performed using ImageJ software. Samples

were compared to the nonsense (N) minus Chk2 siRNA treatment.


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by time lapse microscopy. Cyclin A depleted cells by A3 siRNA (dashed line) caused a typical

G2 phase delay when compared to the controls (solid line). The cyclin A siRNA A3 induced G2

delay was not affected by co-depletion of Chk2 (Figure 5.3 B). Immunoblot data of these

samples showed that cyclin A and Chk2 were successfully depleted in the relevant samples

(Figure 5.3 C). Based on this data it appears that Chk2 is not involved in the cyclin A/CDK2

dependent G2 delay.

5.2.4 ATR phosphorylates Chk1 to impose the cyclin A dependent G2 phase delay.

Since ATR appeared to be responsible for the activation of Chk1 during a normal G2 phase,

the contribution of ATR to the increased level of activated Chk1 in cyclin A depleted cells was

examined. To demonstrate ATR involvement, cyclin A depleted cells were treated with

caffeine to inhibit ATM/ATR activity. Cyclin A depletion with cyclin A A1 siRNA produced the

expected G2/M phase and PY15 accumulation, which was reduced with caffeine addition

(Figure 5.4 A). To examine the kinetics of the effects of caffeine treatment in G2 phase cells,

time lapse microscopy was conducted (Figure 5.4 B). Cyclin A depletion caused the typical

cyclin A dependent G2 phase delay (long dashed line), and this delay was reduced with the

addition of caffeine in G2 phase (6 h post release from thymidine synchrony) (short dashed

line). The caffeine treatment of cyclin A depleted cells increased the rate of mitotic entry

compared to control cells (solid line) but did not result in a complete recovery. This

correlated with previous data demonstrating that Chk1 inhibition promoted faster mitotic

entry (Chapter 3 Figure 5 B and D), indicating that ATR-Chk1 signalling regulates normal G2


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phase progression and is required to implement the cyclin A/ CDK2 dependent G2 phase

delay.

To confirm that the G2 phase delay was dependent on ATR specifically, ATR deficient FO2

cells were used to demonstrate the role of ATR. Cyclin A siRNA treatment of FO2 cells

effectively depleted cyclin A protein but was unable to induce a G2 phase delay. This was

shown by both the absence of a G2/M phase or PY15 accumulation in the cyclin A depleted

samples. Additionally, there was no accumulation of pChk1 detected in the cyclin A depleted

FO2 cells (Figure 5.4 C). This demonstrates that Chk1 is specifically phosphorylated by ATR

when cyclin A is depleted, and that this is required to implement a G2 phase delay.

5.2.5 Activated Chk1 is not a consequence of cyclin A siRNA induced DNA damage.

Because pChk1 is associated with DNA damage, it was important to determine whether the

increased pChk1 observed with cyclin A siRNA treatment was a consequence of DNA damage

caused by the siRNA. In response to DNA damage, sites of damage recruit many DNA

damage proteins required for signalling and repair of the damage. The phosphorylation of

Histone H2AX (γ- H2AX) occurs in response to DNA damage and accumulation of γ-H2AX

levels is commonly used as a marker of DNA damage (Rogakou et al., 1999, Sedelnikova et

al., 2002). Immunoblotting for γ-H2AX demonstrated the expected increase in Etoposide

treated samples. Basal levels of γ-H2AX were detected in untreated (control) and nonsense

siRNA controls (N), and there was no increase of γ-H2AX in cyclin A siRNA treated samples

(A1 and A3) (Figure 5.5 A). Cyclin A knockdown was sufficient in both A1 and A3 samples as


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Figure 5.4 ATR is required to phosphorylate Chk1 to induce the cyclin A dependent G2 phase delay.

A. HeLa cells were transfected with either nonsense (N) or cyclin A siRNA (A1) and after 8 h

treated without or with 5 mM caffeine for 16 h. Cells were harvested and analysed by FACS

or immunoblotted for the indicated proteins. Data contributed by Dr Leo De Boer.

B. HeLa cells were transfected with either nonsense (Control; solid line) or cyclin A siRNA (A1;

broken line) and synchronised with thymidine. At 6 h post release, when cells are in G2

phase, cyclin A siRNA (A1) treated cells were treated with 5 mM caffeine (A1 + caff; grey

line). Cells were then followed by live cell time lapse microscopy, visualised every 15


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minutes and then scored for timing of entry into mitosis. Approximately 200 cells were

counted per treatment. The result was confirmed by conducting the experiment in triplicate

and the best representation was chosen for publishing.

C. Asynchronously growing FO2 cells were either mock transfected (C) or transfected with

nonsense (N) or cyclin A (A1) siRNA, then harvested after 24 h. Cells were analysed by FACS

and immunoblotted as in A. PCNA was used as a loading control.


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shown by cyclin A immunoblotting (Figure 5.5 A). Cells from the same experiment were also

subjected to the Comet assay which detects DNA damage by single cell electrophoresis

where damaged/fragmented DNA damage creates tailing/smearing. DNA damage was readily

detectable in positive controls treated with hydrogen peroxide (H2O2) but no effect was seen

in controls or cyclin A siRNA treated samples (data not shown). The absence of any increase

in markers of DNA damage used, demonstrates that cyclin A siRNA does not induce any

further DNA damage above control levels. Thus increased pChk1 levels seen with cyclin A

siRNA were not a consequence of DNA damage induced by cyclin A siRNA, confirming the

involvement of pChk1 in the cyclin A/CDK2 dependent G2 delay.

5.3 DISCUSSION

It is undeniable that Chk1 is a key protein required for normal development based on the

embryonic lethality of Chk1-/- knockout mice (Takai et al., 2000). Whether this also correlates

to an essential role in the cell cycle is unknown. However, based on its important role in

several well documented cell cycle processes, this suggests that the two may be linked. On

the other hand, there is published work that provides conflicting evidence that cell cycle

progression can occur without Chk1. In DT40 cells without functional Chk1, the cells are

viable but have a slower growth rate (Zachos et al., 2003). These cells are, however, prone to

spontaneous apoptosis and so it is unknown if these cells are capable of completing a correct

cell cycle. Correct completion of the cell cycle is an essential process critical for genomic

stability and, if affected, will result in apoptosis. Therefore, this also points to Chk1 being a

critical cell cycle regulator.


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Figure 5.5 Cyclin A siRNA does not cause DNA damage.

HeLa cells were either untransfected or transfected with nonsense (N) or cyclin A siRNA (A1 and A3),

synchronised by thymidine and collected as they transited through G2 phase (6 h post release). HeLa

cells treated with 1 μM Etoposide overnight were used as a positive control. Whole cell lysates were

blotted for γH2AX, cyclin A, PY15 and α-tubulin as a loading control. Data was obtained by

collaboration with Dr Francis Stevens.


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It has been determined that not only does Chk1 have an important role in the response to

DNA damage but it also has an important role in regulating the timing of normal G2/M

transition. Both of these roles require Chk1 activity to regulate cyclin B/CDK1 activation and

thereby limit the rate of mitotic entry (Capasso et al., 2002, Gatei et al., 2003, Schmitt et al.,

2006,).

Another rate limiting complex controlling entry into mitosis is cyclin A/CDK2. Cyclin A/CDK2

has been established as a G2 phase regulator (Goldstone et al., 2001, Fung et al., 2007) that

controls the activation of cyclin B/CDK1 and, consequently, several key mitotic events,

thereby governing mitotic entry (Fung et al., 2007, Gong et al., 2007a). By depletion of cyclin

A or inhibition of CDK2, cells are delayed in G2 phase due to delayed activation of cyclin

B/CDK1. Direct regulation of cyclin B/CDK1 by cyclin A/CDK2 is unlikely and is most probably

by indirect regulation of other important G2 regulator/s that are capable of influencing cyclin

B/CDK1 activation, such as Chk1. Based on Chk1’s role during G2 phase by regulation of cyclin

B/CDK1 activation, Chk1 stands out as a likely target of cyclin A/CDK2 regulation during G2

phase.

From data presented here it is likely that this is the case. As shown, Chk1 interacts with cyclin

A/CDK2 and is thought to be regulated in some way by cyclin A/CDK2. An interaction

between cyclin A/CDK2 and Chk1 can occur at the centrosomes during G2 phase allowing

cyclin A/CDK2 to regulate Chk1. Chk1 also has other reported G2 functions at the

centrosomes and this involves the negative regulation of Cdc25B which in turn blocks cyclin

B/CDK1 activation to control mitotic entry (Schmitt et al., 2006). Therefore, it is possible that


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interaction between cyclin A/CDK2 and Chk1 at the centrosomes is required to provide relief

of the regulation of Cdc25B by Chk1 and allow timely mitotic entry.

Data from the previous chapter (Chapter 4, Figure 4.7) and this chapter imply that cyclin

A/CDK2 may inactivate Chk1 to allow mitotic entry. This potentially involves inhibitory

phosphorylation of Chk1. Known inhibitory phosphorylations on Chk1, such as Ser280 by Akt,

are required for checkpoint exit (Puc et al., 2005). More recently, two novel phosphorylation

Chk1 sites, Ser286 and Ser301 (Shiromizu et al., 2006), have been identified. These residues

were first identified as cyclin B/CDK1 target phosphorylation sites that were potentially

inhibitory with phosphorylations occurring during normal mitosis and appearing to influence

Ser317 and Ser345 phosphorylation (Ikegami et al., 2008).

If cyclin A/CDK2 can in fact negatively regulate Chk1 by phosphorylation, we hypothesised

that these inhibitory phosphorylation sites may be phosphorylated earlier in G2 phase by

cyclin A/CDK2 to permit mitotic entry. Thereafter, cyclin B/cdk1 would contribute to the

same phosphorylation events in mitosis to allow mitotic progression. Shortly after

formulating this hypothesis others published that cdk2 could phosphorylate Ser286 and

Ser301 in vitro which impacted mitotic entry (Ikegami et al., 2008).

The main activating kinase of Chk1, ATR, is necessary to phosphorylate Chk1 during G2 phase

to implement the cyclin A/cdk2 dependent G2 delay. However, the data do not rule out any

involvement of ATM. If cyclin A/cdk2 is capable of inhibiting Chk1 via phosphorylations

during the G2/M transition then this would either require the prior removal of G2 phase

Ser317 and Ser345 phosphorylations or involve Ser286 and Ser301 phosphorylation


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overriding the activating Ser317 and Ser345 phosphorylations to render Chk1 inactive.

Because the activating phosphorylations are present in G2 phase and are reduced upon

mitotic entry (Chapter 3, Figure 3.2), it is difficult to speculate on the exact status of these

phosphorylations during this transitional period. However, published work shows that Ser317

and Ser345 phosphorylations can occur at the same time as Ser286 and Ser301

phosphorylations (Ikegami et al., 2008) therefore it is likely that Ser286 and Ser301

phosphorylations occurs prior to the dephosphorylation of Chk1 on Ser317 and Ser345.

Therefore, whether Ser286 or Ser301 is required to enhance the dephosphorylation of

Ser317 or Ser345 or to simply inactive Chk1 is unknown.

The involvement of Chk1 had previously been ruled out in the cyclin A/cdk2 dependent G2

delay due to the inability to see an increase in pChk1 levels by immunoblotting (Gong and

Ferrell, 2010). Our data does not agree with this but also suggest that Chk1 is not the only

protein affected by cyclin A/cdk2 depletion/inhibition. The incomplete recovery of mitotic

entry seen with Chk1 inhibition suggested that there are other compensatory pathways that

cyclin A/cdk2 may be required to regulate in order to control mitotic entry. Another

important mitotic regulator, Wee1, has been implicated as a key component of the cyclin

A/cdk2 dependent G2 delay. This same published data ruled out the involvement of either

Cdc25 or Plk1 (Fung et al., 2007). Although the combined removal of cyclin A and Wee1 does

restore cyclin B/cdk1 activity and the number of mitotic cells, implicating the decline of the

cyclin A/cdk2 dependent G2 delay, it does not determine if this is an effect of the Wee1

removal in the cells depleted of cyclin A with siRNA. To more conclusively confirm the role of

Wee1 in the cyclin A/cdk2 dependent G2 delay further single cell analysis should be used to


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demonstrate the effects of Wee1 removal on the recovery of cyclin B/cdk1 activation, which

is delayed by cyclin A removal.

Chk1’s established roles have already identified it as a central effector of G2/M transition in

either a normal or checkpoint response transition. Chk1 has the ability to directly control the

activation of cyclin B/cdk1 through the regulation of its negative regulator, Wee1 (Lee et al.,

2001, Calonge and O'Connell, 2006), and its positive regulator, Cdc25 (Furnari et al., 1999,

Uto et al., 2004, Loffler et al., 2006, Schmitt et al., 2006). Therefore, the possible direct

inhibitory regulation of Chk1 by cyclin A/cdk2 could enhance the activation of Cdc25B and

would also cause a decrease in Wee1 activity. Once the activation of cyclin B/cdk1 is

achieved the sustained inhibition of Chk1 would promote a cyclin B/cdk1 positive feedback

loop (Pomerening et al., 2003, Lindqvist et al., 2005) where cdk1 could itself continue the

regulation of both Cdc25 (Izumi and Maller, 1993, Shen et al., 1998, Lu et al., 1999, Margolis

et al., 2006b) and the inhibition of Wee1 (Ayad et al., 2003, Watanabe et al., 2004, Kim et al.,

2005, Watanabe et al., 2005, Okamoto and Sagata, 2007).

Chk1 is not the only G2/M kinase that can regulate both Cdc25 and Wee1. There are reports

that Plk1 can phosphorylate and regulate both Cdc25B and Wee1 to promote mitotic entry

(Lobjois et al., 2009). For example, Plk1 can promote the degradation of Wee1 (Lee et al.,

2001, Rothblum-Oviatt et al., 2001, Katayama et al., 2005, Stanford and Ruderman, 2005)

and enhance the nuclear localization of Cdc25 to allow its activation of cyclin B/cdk1 in the

nucleus (Toyoshima-Morimoto et al., 2002). Furthermore, Plk1 and Chk1 seem to be able to

regulate each other - whilst Chk1 causes negative regulation of Plk1, Plk1 can likewise


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mediate the suppression of Chk1 by the indirect regulation of its activating partner Claspin

(Yoo et al., 2004). Due to Plk1’s ability to regulate G2/M transition in a similar manner to

Chk1 it is likely that it could be part of the compensatory pathway that is involved in the

cyclin A/cdk2 dependent G2 delay. Although overexpression of active Plk1 does not

overcome the cyclin A/cdk2 dependent G2 delay, it does not rule out any specific effect on

the timing of cyclin B/cdk1 activation or that of other key mitotic events (Fung et al., 2007).

In addition, the activity of Plk1 is governed by its localization and therefore determining the

effects of cyclin A/cdk2 removal/depletion on Plk1 localisation may provide further

confirmation of its non-involvement.

The data confirms that Chk1 has a definitive role in G2 phase which impacts the timed entry

into mitosis. The increase of pChk1 Ser317 seen with cyclin A depletion and cdk2 inhibition

were required to implement the G2 phase delay that was typical of cyclin A/cdk2

depletion/inhibition. The use of Chk1i and siRNA implies, further, that the activation of Chk1

is required to induce the delay and suggests that this involves Chk1 co-ordination of cyclin

B/cdk1 activation at the centrosomes to allow mitotic entry. The mode of cyclin A/cdk2’s

regulation of Chk1 is unknown but we hypothesize that the interaction between cyclin

A/cdk2 and Chk1 is required for inhibitory phosphorylation of Chk1 on Ser 286 and 301 and

accordingly for control of mitotic entry. Proving the regulation of Chk1 by cyclin A/cdk2 is the

next step to further uncovering the pathway involving cyclin A/cdk2 and Chk1 in the

regulation of G2 phase progression.

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Chapter 6

Cyclin A/CDK2 regulation of Chk1

Chapter 6 – cyclin A/CDK2 regulation of Chk1

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6.1 INTRODUCTION

The centrosome is a small membraneless organelle that is critical for cell cycle progression

(Lukasiewicz and Lingle, 2009). It has a complex structure and undergoes its own cycle of

replication, growth and division that is linked to but separate from the cell cycle (Nigg and

Stearns, 2011). The centrosome cycle regulates the duplication and maturation of a single

centrosome into mother and daughter centrosomes, so that each resultant daughter cell

from the cell division cycle is provided with a centrosome. The cycle begins during S phase

and by G2 the duplicated centrosomes migrate to opposite poles of the cell, to facilitate

formation of the mitotic spindle during mitosis and allowing each resultant daughter cell to

inherit a centrosome (Nigg and Stearns, 2011).

The spindle poles are the microtubule organising centres (MTOC) of the mitotic cell. The

spindle consists of a centrosome at either side of the cell during mitosis, which directs

microtubule polymerisation towards the condensed chromosomes. The microtubules attach

to the sister chromosomes via kinetochores and then individual chromosomes are retracted

towards opposite poles of the cell during anaphase. This allows for each daughter cell to

acquire a single copy of the genome once the cell division is complete. Once the mitotic

spindle has facilitated genome segregation during mitosis, the spindle disassembles and all

that remains is the single centrosome inherited by each of the daughter cells.

A small pool of the mitotic complex cyclin B/ CDK1 is initially activated at the centrosomes

and precedes the full activation of cyclin B/CDK1 (Jackman et al., 2003). Prior to cyclin

B/CDK1 localisation and activation at the centrosome during the G2/M transition, cyclin A/


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CDK2 localises to the centrosomes in late G2 phase when centrosome separation has

occurred, and this is required for the timely activation of cyclin B/CDK1 and mitotic entry (De

Boer et al., 2008). The mechanism by which cyclin A/CDK2 regulates this G2/M transition and

whether this involves its localisation at the centrosome is currently unknown. Chk1 is also

known to localise to the centrosome where it regulates the activation of Cdc25B which in

turn regulates cyclin B/CDK1 activation and mitotic entry (Schmitt et al., 2006). The

mechanism regulating Chk1 activation at the centrosome is currently unknown, although the

localisation of cyclin A/CDK2 and its demonstrated role in regulating centrosomal cyclin

B/CDK1 activity suggests a connection between these kinases at the centrosomes is required

to control G2/M progression.

The removal and/or inhibition of the cyclin A/CDK2 complex caused a G2 phase delay in

which Chk1 plays a critical role (Chapter 5). In this study we were also able to show that

cyclin A/CDK2 is required to facilitate the exit of cells from a Chk1 dependent G2 phase

checkpoint arrest. The use of both ATM/ATR and Chk1 inhibitors could not rescue the lack of

checkpoint exit caused by cyclin A depletion or CDK2 inhibition. This data strongly suggests

that there is a regulatory pathway during G2 phase that requires cyclin A/CDK2 regulation of

Chk1. The presence of active Chk1 during G2 phase and the ability of active Chk1 to cause a

G2 phase arrest implies the likelihood that cyclin A/CDK2 may negatively regulate Chk1 to

allow entry into mitosis. Although the mechanism by which cyclin A/CDK2 may regulate Chk1

is unknown, it is possible that this might in turn regulate Cdc25 and therefore cyclin B/CDK1

activity to control mitotic entry.


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A possible mode of Chk1 regulation by cyclin A/CDK2 is based on the ability of cyclin B/CDK1

to directly phosphorylate Chk1 Ser286 and Ser301 which was shown to inhibit Chk1 activity

(Shiromizu et al., 2006). The similar substrate specificity of cyclin B/CDK1 and cyclin A/CDK2

lead to the hypothesis that cyclin A/CDK2 catalyses Chk1 Ser286 and Ser301

phosphorylations during late G2 to promote mitotic entry. The interaction of cyclin A/CDK2

and Chk1 and this mode of regulation is investigated in this Chapter.

6.2 RESULTS

6.2.1 pChk1 and Cyclin A co-localise during G2 phase.

It has already been demonstrated that pChk1 can localise to the centrosomes during G2

phase and mitosis (Chapter 3 Figure 3.6), and cyclin A/CDK2 also localised to the

centrosomes during G2 phase (Chapter 4 Figure 4.4 and 4.5). Immunofluorescent staining of

cells for pChk1 and cyclin A demonstrated that both localise predominantly to the nucleus

but clear centrosomal staining was also evident. A cell was determined to be in late G2 phase

when the separation of the centrosomes was evident prior to nuclear envelope breakdown

and chromosome condensation. Such cells were identified and as the arrows indicate, cyclin

A/CDK2 and pChk1 co-localised to the centrosomes in late G2 phase (Figure 6.1 A, as

indicated by arrows). Staining for Chk1 confirmed Chk1 and cyclin A co-localised during G2

phase (Figure 6.1 B). Although cells were only stained for cyclin A, it was assumed that CDK2


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Figure 6.1 Cyclin A and Chk1 co-localise at the centrosomes.

A. Asynchronous HeLa cells were grown on coverslips, fixed and immunostained for cyclin A

(green), pChk1 Ser317 (pChk1) (red), and DAPI for DNA (blue). Arrow heads indicate the

centrosomes. Both images are of G2 phase cells as determined by the separation of the

centrosomes. Images were taken using a Zeiss Apotome widefield microscope equipped

with a CCD camera.

B. Cells were treated and fixed as in A), stained for Chk1 (green), Cyclin A (red) and DAPI for

DNA (blue). Arrow heads indicate the centrosomes. The image is of G2 phase cells as

determined by the separation of the centrosomes. Image was taken using a Zeiss Apotome

widefield microscope equipped with a CCD camera


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would be present at this time as CDK2 localisation at the centrosomes was previously

determined to be cyclin A dependent (Chapter 4 Figure 4.5)(De Boer et al., 2008). Although

the localisation of these proteins at the centrosomes had been reported previously, it was

important for this study to determine whether they co-localised during G2 phase. Based on

their co-localisation it was plausible to hypothesise that cyclin A/CDK2 and Chk1 may interact

during G2 phase and that this may facilitate the direct regulation of Chk1 by cyclin A/CDK2.

6.2.2 Chk1 and Cyclin A interact during G2 phase.

To confirm an interaction between cyclin A/CDK2 and Chk1, co-immunoprecipitation

experiments were conducted. Cyclin A and Chk1 are both approximately 50 kDa, which runs

very close to the IgG heavy chain when resolved by SDS-PAGE and makes detection of the

endogenous proteins problematic. To bypass this difficulty, we over-expressed a Cherry

tagged Cyclin A construct in HeLa cells. Chk1 complexes were immunoprecipitated using

polyclonal Chk1 antibody, and immunoblot analysis of co-immunoprecipitated proteins

revealed that Cherry-cyclin A specifically associates with Chk1. No Cherry-cyclin A was

detected in the control immunoprecipitation confirming the specificity of the interaction

(Figure 6.2 A). To confirm this interaction, Flag tagged Chk1 was over-expressed and cyclin A

antibody coupled to protein A beads was used to immunoprecipitate Cyclin A associated

protein complexes. Cells were either treated with or without nocodazole to arrest them in

mitosis. Nocodazole treatment was used as a negative control as cyclin A is degraded in

mitosis. Phosphorylated Mek1 T286 (pMek1 T286), a mitotic marker was used to confirm the

synchronisation. While cyclin A was reduced with nocodazole treatment, so were Chk1


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Figure 6.2 Cyclin A and Chk1 co-immunoprecipitate

A. 1 mg of protein was extracted from asynchronous HeLa cell lysates either transfected with

or without Cherry-cyclin A DNA. Immunoprecipitation using Chk1 antibody demonstrated

that Cherry-cyclin A binds to Chk1. A control sample treated with protein A sepharose beads


179

only and no Chk1 antibody (CONT) were used to demonstrate the specificity of the

interaction. 20 μg of lysates was used to observe the expression of the Cherry-cyclin A

construct and the presence of Chk1. PCNA was used as a loading control.

B. Cells transfected with or without Flag-Chk1 were treated overnight with or without

nocodazole. Nocodazole treated samples were collected by detaching the mitotic cells with

a mitotic shake off. Cyclin A antibody was coupled to protein A beads prior to

immunoprecipitation. Chk1 antibody was then used to detect endogenous and exogenous

binding of Chk1 to Cyclin A. Anti-rabbit IgG antibody coupled to protein A beads was used as

a control to confirm that no non-specific binding was occurring and that the interaction

between cyclin A and Chk1 was specific.


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levels (Figure 6.2 B). Endogenous Chk1 was stable in mitosis, but the Flag-Chk1 was unstable

in nocodazole treated cells. Endogenous Chk1 was detected in all cyclin A

immunoprecipitates and not detected in control samples immunoprecipitated with the

control anti-rabbit IgG, which confirmed the specificity of the interaction (Figure 6.2 B; lower

band). The exogenous, Flag-Chk1 was also detected in asynchronously growing cells

immunoprecipitated with cyclin A. Although this does not establish a direct interaction

between cyclin A/CDK2 and Chk1, it does demonstrate that cyclin A/CDK2 is in a complex

with Chk1.

6.2.3 Cyclin A can phosphorylate Chk1 on Ser286 and Ser301.

Two novel phosphorylation sites on Chk1, Ser286 and Ser301, were identified as inhibitory

sites phosphorylated by cyclin B/CDK1 in mitosis (Shiromizu et al., 2006). The data pointing to

cyclin A/CDK2 potentially regulating Chk1, and the interaction between these proteins

demonstrated above, suggested that cyclin A/CDK2 could phosphorylate these sites in G2

phase to inhibit Chk1 and allow timely mitotic entry. To examine whether cyclin A/CDK2

could directly phosphorylate these sites in vitro, Glutathione S-Transferase (GST) fused to full

length Chk1 wildtype (WT) or Chk1 with phospho site mutations of serines (S or Ser) 286 and

301 to alanine (A or Ala) were generated. Single site mutations, Chk1 Ser286 to Ala (S286A)

and Chk1 Ser301 to Ala (S301A), and a double mutant, Chk1 Ser286 to Ala and Ser301 to Ala

(SS286,S301AA) were compared to a wild type (WT) Chk1 control. The phosphorylation of

Chk1 was determined by assessing the ability of recombinant cyclin A/CDK2 kinase to

phosphorylate the GST-Chk1 constructs. The kinase was first tested and found to be capable


181

of phosphorylating a histone H1 control (data not shown). Cyclin A/CDK2 was capable of

phosphorylating the WT and S301A Chk1 constructs, and there also appeared to be minor

phosphorylation of the S286A mutant and even less phosphorylation of the SS286,301AA

double mutant (Figure 6.3). No phosphorylation of the GST control was expected or evident

(Figure 6.3, Lane 1). These results demonstrate that cyclin A/CDK2 is able to phosphorylate

specifically Chk1 Ser286 and to a lesser extent, possibly Ser301. The largest reduction in

phosphorylation was seen with the double, SS286,301AA, mutant and this data suggests that

the phosphorylations are perhaps an ordered event, where Ser286 is required for the

subsequent phosphorylation of Ser301.

6.2.4 Over-expression of Chk1 delays mitotic entry.

Once it had been established that cyclin A/CDK2 could phosphorylate Chk1 in vitro on Ser286

and, if it is an ordered event, Ser301 also, the Chk1 phosphorylation mutant constructs,

inserted into mammalian expression vectors, were used to determine their effect on mitotic

entry. Based on the hypothesis that phosphorylation of these sites by cyclin A/CDK2 during

G2 phase should inhibit Chk1 and allow mitotic progression, Ala mutants should delay mitotic

entry due to the lack of Chk1 phosphorylation and subsequent inhibition.

The Chk1 mutants were cloned into two mammalian expression vector systems to create

Flag-Chk1 and Cherry-Chk1 constructs. These Chk1 mutants were generated in a Chk1 Ser317

and Ser345 phospho-mimicking mutant background, where the serines were mutated to

aspartate (Asp or D) residues. This was based on the assumption that the inhibitory site


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Figure 6.3 Cyclin A/CDK2 phosphorylates Chk1 in vivo

Purchased recombinant cyclin A/CDK2 and GST-Chk1 constructs were combined and a 32P-γ-ATP in

vitro kinase assay performed. Samples were resolved by SDS-PAGE and stained with Coomassie

Brilliant Blue (G-250). Phosphorylated bands were visualised by autoradiography. A photograph of

the Coomassie stained gel (lower panel) is used to show equal loading between samples. GST-Chk1 is

75 kDa band and GST is a 25kDa band (indicated with arrows). Numbers to the left of the lower panel

indicate the molecular weight in kDa’s of the marker bands. Arrows indicate the bands as per

labelling. .


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Figure 6.4 Chk1 SS317,345DD, with S286A and S301A mutations do not delay in G2 phase

A. HeLa cells were co-transfected with GFP plasmid (2 µg) and the indicated Flag Chk1 plasmid

(2 µg) (SS317,345AA, S286A, S301A, SS286,301AA). 24 h post transfection cells were fixed

with 70 % Ethanol and stained with PI. 2D FACS was carried out to determine the DNA

content of the GFP co-transfected population. Scatter plots show the gated, GFP positive,


184

cells (red boxes, percentages of the cell population are shown within). Cell cycle profiles of

the gated population are shown on the left. Percentages of cell cycle phases, G1, S and

G2/M (of the gated population) are shown in the top right hand corner in descending order.

All samples were gated except for the ungated control (untransfected control), where a

representation of the entire cell population is shown as a control. Analysis was conducted

using FlowJo software.

B. Samples for immunoblotting were collected from the experiment conducted in A) to

confirm the expression of Flag and GFP. Samples were immunoblotted for Flag, GFP and

PCNA, used as a loading control.


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phosphorylations occurred on the already activated Chk1 to inhibit the kinase activity. Single

S286A and S301A mutants, and the double mutant, SS286A,S301AA, were made in Flag-Chk1

SS317,345DD mutant background and against the SS317,345DD construct as a control. Cells

were co-transfected with GFP to identify Chk1 transfected cells. Analysis by FACS data

revealed no difference in the G2/M populations of Flag-Chk1 expressing cells (Figure 6.4 A),

even though a similar level of overexpression of each sample was achieved (Figure 6.4 B) This

negative result may have been due to the Aspartate mutations of sites, Ser317 and Ser345.

Mutation of Ser345 had previously been reported to affect 14-3-3 binding to these sites and

therefore produce a catalytically defective Chk1 (Capasso et al., 2002, Jiang et al., 2003,

Dunaway et al., 2005). Therefore Chk1 mutations of Ser286 and Ser301 were generated in a

Chk1 Ser317 and Ser345 wild type background and the involvement of Ser286 and Ser301

was re-examined.

HeLa cells were transfected with Flag-Chk1 constructs, treated with Etoposide and then

forced into mitosis with caffeine addition, and fixed for immunofluorescent staining 6 h after

caffeine addition. We had previously used this system to demonstrate that cyclin A/CDK2

depletion/inhibition appeared to be required to inhibit Chk1 to control checkpoint exit

(Chapter 5). Flag-Chk1 expressing cells were then scored as either G2 phase or mitotic. This

experiment demonstrated that a lower proportion of wild type (WT) Flag-Chk1 expressing G2

phase cells had entered mitosis compared to either single, S286A or S301A, or double,

SS286,301AA mutants which all drove similar proportions of the over expressing cells into

mitosis (Figure 6.5 A). Cells were also scored for apoptosis which is not presented here.

Apoptosis was very noticeable in the WT samples and may indicate that the overexpression


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Figure 6.5 Chk1 Ser286 and Ser301 Alanine mutants cause a delay of mitotic entry.

A. HeLa cells were either mock transfected (CONT) or transfected with 2 μg of Flag Chk1 constructs

WT, S286A, S301A or SS286,301AA. 6 h post transfection cells were treated overnight with


187

Etoposide (1 μM). The following day 5 mM caffeine was added to each sample and incubated

for a further 6 h, after which coverslips were fixed with methanol. Cells were stained with Flag

antibody and DAPI for DNA and scored as either mitotic or G2 phase based on the morphology

of the DNA. Only the Flag staining cells were scored for the Chk1 transfected samples and the

mock control cells were counted randomly. Approximately 200 cells were counted per

experiment and the experiment was conducted in triplicate.

B. HeLa cells were co-transfected with GFP plasmid (2 μg) and the indicated Flag Chk1 plasmid (2

μg). A GFP only control (not co-transfected with Flag Chk1 plasmid) was transfected with 4µg of

plasmid. 24 h post transfection cells were collected for analysis. Cells were fixed with 70%

ethanol and stained with PI. 2D FACS was carried out to analyse cell cycle profiles of positively

transfect cells. Positively transfected cells were analysed by gating the GFP positive population.

Scatter plots show the gated, GFP positive, cells (red boxes, percentages of the cell population

are shown within). Cell cycle profiles of the gated population are shown on the left. All samples

were gated except for the ungated control (Control) which was not transfected with either GFP

or Flag-Chk1 and is a representation of the entire cell population. Analysis was conducted using

FlowJo software.

C. The experiment in B) was conducted in triplicate. All samples were analysed using FlowJo and

the percentage of cells in the representative cell cycle phases was determined. The data

presented is the mean G2/M population versus the G1 population. Error bars are representative

of standard deviations. A p value of 0.026 was obtained from a T-test conducted using the GFP

and SS286,301AA values.

D. Samples for immunoblotting were collected from the experiment conducted in B) to confirm

the expression of Flag and GFP. Samples were immunoblotted for Flag, GFP and PCNA, used as a

loading control.


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of WT Chk1 can cause cell cycle aberrations and apoptosis. This was made evident by the

large difference between the WT and control scorings. The mutants did not display a similar

presence of apoptotic cells as the WT and perhaps this reflects the inability of these cells to

enter mitosis readily. This data indicated that the inability of Chk1 to be phosphorylated on

Ser286 or Ser301 caused a delay in mitotic entry, however it opposes the ordered

phosphorylation theory suggested from Figure 6.3.

This initial observation was further investigated using FACS analysis of transfected cells to

determine whether the mutants caused an increase in the G2/M population. This required

co-transfection of Flag constructs with a GFP vector to identify the transfected cells. Two

dimensional FACS was used to analyse the DNA content of the GFP co-transfected cells

(Figure 6.5 B). As indicated in Figure 6.5 B, the double Chk1 SS286,301AA mutant produced

the greatest increase in the G2/M phase population when compared to the G1 population.

S286A caused a slightly larger increase in G2/M compared to S301A, which reflects the kinase

data and supports the theory of an ordered phosphorylation. Although this data does not

produce significant values, this could reflect the numerous pathways that are required to

regulate G2/M progression. The experiment shown in Figure 6.5 B was conducted in triplicate

and the ratio of the G2/M population compared to the G1 population is presented. Although

the WT Chk1 caused a similar result to the SS286,301AA double mutant, compared to the

GFP control the ratio of G2/M phase cells compared to the G1 population was significantly

larger (Figure 6.5 C). There was also a slight increase in both the single mutants despite the

overexpression of each sample being similar (Figure 6.5 D). Despite not being evident earlier,

this data provided new evidence that overexpression of WT Chk1 is also responsible for an


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increased G2/M population, for which the reason remains unclear. This type of analysis

detects G2/M populations and could not be used to detect specific increases in G2 phase but

provides strong evidence that overexpression of Chk1 SS286,301AA double mutant may

cause a G2 phase delay.

To further characterise the delay of these S286A and S301A mutants in G2 phase time lapse

microscopy was used. For these experiments, Cherry-Chk1 were used to identify transfected

cells. The Cherry mutants were scored for their timing of mitotic entry. It is a well noted

observation that cell transfection delays mitotic entry and this is also evident in this data

when comparing the untransfected control (CONT) to the Cherry transfected control

(Cherry). As shown in Figure 6.6 A, the Chk1 SS286,301AA double mutant did not cause a

significant delay of entry into mitosis compared to cells expressing Cherry vector alone. This

was possibly a consequence of the inability of the Cherry tagged mutant proteins to be

phosphorylated on the activating phosphorylation sites, Ser317 and Ser345. Analysis of

lysates demonstrated that both single and double mutants were devoid of activating

phosphorylations, even in the presence of DNA damage. On the other hand, Cherry Chk1 WT

(WT) could be phosphorylated on both Ser317 and Ser345 activating sites (Figure 6.6 B).

Therefore while the WT construct could be activated and it does not appear to delay mitotic

entry further than the Cherry vector control or the SS286,301AA double mutant.

As the Cherry mutants were unable to be normally phosphorylated on Ser317 and Ser345,

cells were co-transfected with GFP and Flag-Chk1 constructs and followed by time lapse

microscopy. The effect of these mutations on normal G2/M progression and checkpoint exit


190

Figure 6.6 Cherry-Chk1 constructs have impaired Ser317 and Ser345 phosphorylation.

A. HeLa cells were untransfected (CONT) or transfected with the indicated Cherry constructs

(WT or SS286,301AA). The Cherry only construct was used as a control. Cells were

synchronised with thymidine and scored for their mitotic entry using time lapse microscopy.


191

Only Cherry expressing cells (excluding the CONT), identified by the fluorescent imaging

were scored. Approximately 200 cells were counted per condition and the cumulative

mitotic index is shown. The result was confirmed by conducting the experiment in triplicate

and the best representation was chosen for publishing.

B. HeLa cells were transfected without (CONT) or with the indicated Cherry-Chk1 constructs. 6

h post transfection, cells were treated with Etoposide for 16 h. Lysates were generated from

these cells and then analysed for the levels of pChk1 Ser317, pChk1 Ser345, Chk1 and PCNA

(loading control). Endogenous Chk1 and Cherry-Chk1 are labelled and indicated by the

arrows. The asterisk indicates an unspecific band or possible truncation of the Flag-Chk1

plasmid.


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Figure 6.7 Flag Chk1 constructs are phosphorylated on Ser317 and delay mitotic entry.

A. HeLa cells were transfected without (CONT) or with the indicated Flag-Chk1 constructs. 6 h

post transfection cells were treated with Etoposide for 16 h. Cells lysates were analysed for

pChk1 Ser317, Chk1 and PCNA (loading control). The Flag constructs ran just above the

endogenous Chk1. Arrows indicate the appropriate bands.

B. HeLa cells were transfected with GFP only (CONT) or co-transfected with the indicated Flag-

Chk1 constructs. Cells were then synchronised with thymidine and scored for their mitotic

entry using time lapse microscopy. Only GFP expressing cells were scored. Approximately

200 cells were counted per condition and the cumulative mitotic indexes are shown. The

result was confirmed by conducting the experiment in triplicate and the best representation

was chosen for publishing.


193

was assessed. Normal G2/M transition was analysed by the thymidine synchronisation of

cells, while checkpoint exit was assessed by treating cells with Etoposide and forcing the exit

from the checkpoint by the addition of caffeine. The mutants were first tested for their ability

to be phosphorylated on the activating phosphorylation, Ser317. Both the wild type and

double mutant Flag-Chk1 constructs were readily phosphorylated on the activating Ser317

site (Figure 6.7 A). Time lapse microscopy revealed that during normal G2/M progression,

wild type Chk1 produced a delay in mitotic entry by less than 1 h, whereas the SS286,301AA

double mutant produced a slightly extenuated delay (Figure 6.7 B). A similar minor delay was

also observed in the G2 phase checkpoint exit system, with again the double mutant having a

similar effect as the wild type protein (data not shown). These data suggest that

phosphorylation of Ser286 and Ser301 may be required for mitotic entry, but overexpression

of wild type Chk1 also sufficiently delayed entry into mitosis.

6.2.5 In vivo phosphorylation of Chk1 Ser286 and Ser301 is not dependent on cyclin

A/CDK2.

Commercial phospho specific Ser286 and Ser301 Chk1 antibodies became available from

ABCAM during this study but were found to be inadequate. Non-commercial antibodies were

then obtained courtesy of Dr Masaki Inagaki of the Aichi Cancer Centre Research Institute,

Japan. Ser286 and Ser301 phosphorylations are known to occur during mitosis therefore the

antibodies were initially tested on cells arrested in mitosis. These phosphorylation site

specific antibodies only detected immunoprecipitated Chk1 (Figure 6.8 A, lanes 1 and 2).

While both antibodies were able to detect immunoprecipitated Chk1 phosphorylated in


194


195


196

Figure 6.8 Chk1 phosphorylation on Ser286 Ser301

A. Lysates were collected from HeLa cells treated with or without 0.5 µM nocodazole for 16 h.

These lysates were used for immunoprecipitation with Chk1 antibody or anti-goat IgG as a

control (Cont). Lysate and immunoprecipitation samples were resolved by immunoblot and

detected with pChk1 S286, pChk1 S301, and Chk1 antibodies (as indicated by arrows). The

strong bands visible in control lanes (Cont IP) are IgG bands.

B. HeLa cells transfected with nonsense (N) or cyclin A siRNA (A3) were treated with 0.5 µM

nocodazole for 16 h. Samples were immunoprecipitated with Chk1 and resolved by SDS-

PAGE. Immunoblot analysis for pChk1 S301 and Chk1 were carried out on samples that had

been immunoprecipitated for Chk1. Lysates were immunoblotted for cyclin A to check for

knockdown and PCNA to check equal sample loading. The quantification of Chk1 S301 levels

were determined by measuring the mean intensities of bands compared to the nonsense

control (N), using ImageJ software.

C. HeLa cells were either untransfected or transfected with nonsense (N) or cyclin A siRNA (A3)

and then thymidine synchronised. Untransfected synchronised cultures were treated for 2h

with 4 µM CDK2i (PHA-533533) 5 h post thymidine release. All samples were collected 7 h

post release. Nocodazole treated HeLa cells were used as controls, immunoprecipitated

with Chk1 antibody (+), or control anti-goat IgG antibody immunoprecipitation from the

same lysates (-). Immunoprecipitated complexes were immunoblotted for pChk1 S301,

pChk1 S317 and Chk1, lysates were immunoblotted for cyclin A, PY15 and PCNA. The

quantification of Chk1 S301 levels were determined by measuring the mean intensities of

bands compared to the nonsense control (N), using ImageJ software.

.


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D. HeLa cells transfected with nonsense (N) or cyclin A (A3) siRNA were treated with Etoposide

for 16 h. Untransfected HeLa cells were treated for 16 h with Etoposide, before addition of 4

µM of CDK2i (PHA-533533) for 2 h. Samples were immunoprecipitated with Chk1 antibody,

resolved by SDS-PAGE and immunoblotted for pChk1 S301, pChk1 S317 and Chk1. Lysates

were immunoblotted with cyclin A and PCNA to confirm knockdown and loading. The

quantification of Chk1 S301 levels were determined by measuring the mean intensities of

bands compared to the nonsense control (N), using ImageJ software.

E. HeLa cells transfected with scrambled (N) or cyclin A (A3) siRNA were treated with

Etoposide for 16 h. Untransfected HeLa cells were treated with 16 h with Etoposide before 2

h incubation with 4 µM CDK2i (PHA-533533) or 9 µM CDK1i (Ro 3306). All samples were

then treated for 5 h with 5 mM caffeine. Samples were immunoprecipitated with Chk1

antibody, resolved by SDS-PAGE and immunoblotted for pChk1 S301, pChk1 S317 and Chk1.

Lysates were immunoblotted with cyclin A, pMek1 T286, PY15 and PCNA to confirm

knockdown. The quantification of Chk1 S301 and S317 levels were determined by

measuring the mean intensities of bands compared to the nonsense control (N), using

ImageJ software. All samples were compared to the levels of endogenous Chk1 and then

standardised to the control (N).


198

mitosis, the Ser301 antibody was more sensitive than the Ser286 antibody. Although Ser301

was not identified as a strong phosphorylation site for cyclin A/CDK2 (Figure 6.3), there was

evidence of ordered phosphorylation and therefore, for all further analysis we used the

Ser301 antibody only.

We investigated whether the mitotic phosphorylation of Ser301 was dependent on cyclin A.

siRNA depletion greatly reduced cyclin A levels in mitotic samples, but there was little effect

on Ser301 phosphorylation in these samples (Figure 6.8 B). Because cyclin A/CDK2 controls

mitotic entry and is most active in G2 phase, the effect of cyclin A depletion on Ser301

phosphorylation was analysed in G2 phase samples. Chk1 Ser301 phosphorylation was

minimally affected by cyclin A depletion, and in some cases was slightly increased, as seen in

thymidine treated samples (Figure 6.8 C). This was not due to insufficient knockdown as

indicated by immunoblotting of the lysates. Surprisingly, CDK2i treatment caused a marked

reduction of Ser301 levels. Examination of Chk1 Ser317 phosphorylation confirmed the

increased Chk1 activation caused by either cyclin A siRNA or CDK2i treatment (Figure 6.8 C).

Immunoblot analysis of Etoposide treated samples revealed an expectedly large amount of

Chk1 pSer317 phosphorylation levels. At the same time a low level of Ser301

phosphorylation was also evident and was reduced further by either cyclin A siRNA or CDK2i

treatment (Figure 6.8 D). When the control cells (N) were released from the checkpoint

arrest with caffeine, the levels of Chk1 Ser301 increased while Ser317 was greatly reduced.

Ser301 was highest in the control (N), reduced in the cyclin A siRNA (A3) treated samples,

and absent in CDK1 and CDK2 inhibitor treated samples (Figure 6.8 E). Inversely, the level of


199

Chk1 phosphorylated on Ser317 was reduced in both the control (N) and cyclin A siRNA (A3)

treated samples but remained elevated in the CDK inhibitor treated cells (Figure 6.8 E). The

effect of the CDK2i was stronger than cyclin A knockdown, which is likely to reflect the

incomplete removal of the cyclin A/CDK2 complex by siRNA compared to degree of inhibition

by the CDK2i. This mirrors earlier findings where CDK2i more strongly affected exit from the

checkpoint arrest than did cyclin A siRNA (Chapter 4 Figure 4.6). Both CDK inhibitors are

known to block checkpoint exit and accordingly the mitotic marker pMek1 T286 was

completely absent in the CDK inhibitor treated samples, and corresponded to high levels of

the G2 phase marker, PY15. While cyclin A knockdown was sufficient, the cells were able to

exit the checkpoint as seen by elevated levels of pMek1 T286, however they were delayed

slightly as seen by a minor elevation in PY15 levels (Figure 8 E). This data supports the

suggestion that Ser301 is an inhibitory phosphorylation, as it accordingly increases in samples

that required inactive Chk1 so that they can exit a checkpoint.

6.2.6 Chk1 mutants do not change localisation.

The localisation of proteins is important for them to carry out their specific roles. To

determine whether Ser286 and Ser301 phosphorylations affect the localisation of Chk1,

immunofluorescence staining was carried out on cells transfected with S286A and S301A

mutants. HeLa cells transfected with Flag-Chk1 constructs were synchronised in G2 phase by

treatment with either thymidine or Etoposide. Cells stained with Flag antibody were analysed

for Chk1 localisation. The exposure time for all images was kept constant however the level

of expression varied from cell to cell. Despite this, the localisation of the WT and double


200

mutant appeared very similar in both G2 synchronised treatments, with predominantly

nuclear staining and some cytoplasmic staining (Figure 6.9). The single mutants also

displayed predominantly nuclear localisation with both treatments (data not shown). The

data does not correlate to published data that showed WT Chk1 to be predominantly

cytoplasmic, while the double SS286,301AA double mutant is nuclear (Enomoto et al., 2009).

This further reflects our results where WT and SS286,301AA overexpression appeared to act

similarly to delay cell cycle progression.

6.3 DISCUSSION

Data from Chapter 5 suggested cyclin A/CDK2 negatively regulates Chk1 activation. During

the time of this study novel phosphorylation sites on Chk1, Ser286 and Ser301, were

identified as cyclin B/CDK1 phosphorylation sites that inhibit Chk1 in mitosis (Enomoto et al.,

2009). It was therefore hypothesised that these sites may also be phosphorylated by cyclin

A/CDK2 in G2 phase and during exit from the G2 phase checkpoint arrest, to inhibit Chk1

thereby allowing mitotic entry. Cyclin A/CDK2 was indeed capable of associating with Chk1,

although this was not identified as a specific G2 phase interaction. However, based on the co-

localisation data cyclin A/CDK2 and Chk1 interact in G2 phase as both are detected at the

centrosomes during centrosome separation. Chk1 has been reported to negatively regulate

cdc25B at the centrosomes during G2 phase (Schmitt et al., 2006). This could explain the G2

phase delay, if cyclin A/CDK2 was not present to negatively regulate Chk1 activity, then Chk1

would continue to negatively regulate cdc25B at the centrosome and cause a delay of mitotic

entry. Based on this data, it seemed possible that cyclin A/CDK2 was imposing some mode of


201

Figure 6.9 SS286,301AA mutation does not affect Chk1 localisation

HeLa cells were transfected with the indicated Chk1 constructs. These cells were synchronised with

overnight treatment with 1 µM Etoposide (etop O/N treatment) or thymidine and collected in G2

phase (6 h post release) (G2 thy). After fixation with methanol, the cells were stained with Flag and

DAPI for DNA. Images of positively transfected cells in G2 phase were taken using the Zeiss Apotome

Fluorescent microscope with 100x oil objective.


202

regulation which involved interaction with Chk1 during G2 phase, possibly at the

centrosomes. The remainder of the study concentrated on the mode of Chk1 regulation

implemented by cyclin A/CDK2. In particular the role of Chk1 phosphorylation on, Ser286 and

Ser301 during G2 phase was investigated.

In Chapter 3, I confirmed the presence and importance of activated Chk1 during G2 phase.

Based on this data and the working hypothesis, it seemed likely that Ser286 and Ser301

phosphorylation by cyclin A/CDK2 occurred at the same time or just after activating

phosphorylations to inhibit G2 phase Chk1. It was later demonstrated that phosphorylation

of Chk1 on Ser286 and Ser301 could occur on the same molecule of Chk1 phosphorylated on

Ser317 and Ser345 (Ikegami et al., 2008). The analysis of Ser286 and Ser301 mutated to

Alanine in a Ser317 and Ser345 phospho mimicking construct presented no G2 phase delay.

This either reflects that these inhibitory phosphorylations cannot overcome the activating

phosphorylations and cannot occur at the same time on the same Chk1 molecule, unlike a

previous report (Ikegami et al., 2008), or that 14-3-3 binding of these mutants is affected. As

mentioned previously, in Chapter 3, the Ser345 phosphorylation site lies within a conserved

14-3-3 binding motif, and mutation of this site affects 14-3-3 binding and therefore

localisation and function of Chk1 (Dunaway et al., 2005). In fact the phospho mimicking Chk1

mutation disrupts 14-3-3 binding and is catalytically inactive (Jiang et al., 2003). Based on the

possible side effects of the phospho mimicking mutations, it was decided to mutate Ser286

and Ser301 to Alanine in Chk1 wildtype constructs so that Ser317 and Ser345

phosphorylation could occur naturally, and found that these constructs did cause a G2 phase

delay. Only Flag-tagged constructs and not Cherry-tagged constructs delayed entry when


203

compared to transfected controls, the identified difference being that they were able to be

phosphorylated on Chk1 Ser317. Therefore unlike the Cherry-tagged versions, these

constructs were normally activated and suggests that this phosphorylation is required to

impose a G2 phase delay. Unlike, Enomoto et al. (2009) who advocated that Ser286 and

Ser301 Alanine mutants cause a G2 phase delay of only 2 h compared to a wild type Chk1

control, the data presented here does not support this. Unexpectedly a similar delay of the

SS286,301AA double mutant compared to the Chk1 WT was seen in the present study. This

was not evident with the Cherry tagged constructs and this is possibly due to the inability of

the constructs to be phosphorylated on activating phosphorylations to the same extent as

the Flag Chk1 WT construct. Alternatively, the natural phosphorylation of inhibitory sites was

perhaps sufficient enough to allow timely mitotic progression. The discrepancy in results

between this study and others may be explained by the experimental systems used. Unlike

the analysis of transient transfections conducted as part of this study, Enomoto et al. (2009)

used inducible Chk1 cell lines. The use of inducible cell lines enables the timed expression of

Chk1 in all cells during a particular stage of the cell cycle compared to the transient Chk1

over-expression used in this study, which allowed variations in the amount of Chk1 expressed

as well as its untimely expression throughout an entire cell cycle.

Although the inducible expression of wildtype Chk1 used by Enomoto et al. et al (2009) did

not appear to induce a G2 phase delay, it has previously been reported that Chk1

overexpression causes a G2 phase delay in yeast (Walworth et al., 1993), which was linked to

enhanced phosphorylation CDK1 Tyr15 levels (O'Connell et al., 1997, Rhind et al., 1997,

Lopez-Girona et al., 1999, Raleigh and O'Connell, 2000, Lopez-Girona et al., 2001a). It is


204

unsurprising that overexpression of Chk1 could delay mitosis, as Chk1 depletion causes the

hastening of mitotic entry (Chapter 3). As reported here, Chk1 WT overexpression resulted in

a delay compared to controls. It is possible that by overwhelming the cell with a high level of

Chk1 leads to an imbalance between active and inactive Chk1, with a possible tip towards a

larger amount of active Chk1, therefore resulting in a mitotic delay. To balance out the levels

Chk1 when overexpressing Chk1 constructs, I attempted to generate siRNA resistant Chk1

constructs so that endogenous levels of Chk1 could be depleted at the same time as

overexpression. Although the use of Chk1 siRNA resistant constructs would have better

avoided an imbalance of Chk1 protein levels, unfortunately the siRNA resistant Chk1

constructs were not used in this study because no further time could be invested into the

development of these constructs.

The data presented confirms that Chk1 Ser286 and Ser301 can be phosphorylated by cyclin

A/CDK2 in vitro as reported by others (Ikegami et al., 2008). In agreement with published

data, it was also demonstrated that a larger decrease of phosphorylation occurred with the

S286A mutant than the S301A mutant. Although the in vitro data demonstrated

phosphorylation by cyclin A/CDK2, there remains no evidence for the in vivo phosphorylation

of these sites by cyclin A/CDK2. Investigation of these sites in vivo revealed that cyclin

A/CDK2 was most likely not the predominant kinase responsible for these phosphorylations.

As shown, CDK2i caused only a small reduction in G2 phase Ser301 phosphorylation, while

cyclin A siRNA did not change Ser301 phosphorylation. Although phosphorylation by cyclin

A/CDK2 cannot be completely dismissed, the CDK1i data strongly suggested that cyclin

B/CDK1 is likely to be the major kinase required for Ser286 and Ser301 phosphorylation


205

during G2 phase and mitosis. The specificity of the CDK1i inhibitor used and the

concentration at which it was used cannot be used to refute the involvement of CDK1

(Vassilev et al. 2006, Ma et al. 2009). If cyclin B/CDK1 is the predominant kinase required for

Ser286 and Ser301 phosphorylation then it is possible that a small pool of cyclin B/CDK1

becomes activated at late G2, to inhibit Chk1 by phosphorylation of Ser286 and Ser301 and

allow mitotic entry. This could also potentially feed into the already well-established cyclin

B/CDK1 self-activation loop, due to Chk1 inhibition resulting in Cdc25B, and therefore cyclin

B/CDK1, activation. However it is possible that cyclin A/CDK2 dependent activation of cyclin

B/CDK1 could contribute to the phosphorylation status of Ser286 and Ser301. Cyclin A/CDK2

could also contribute by providing the initial phosphorylation of Chk1 Ser286 and Ser301

which, soon after, is taken over by cyclin B/CDK1. The latter may be likely as Ser286 and

Ser301 can be phosphorylated in the G2 checkpoint and it is well known that cyclin B/CDK1 is

kept inactive during this time, while a small pool of cyclin A/CDK2 is required during the

checkpoint to allow checkpoint exit (Laoukili et al., 2008b, Alvarez-Fernandez et al., 2010,

Alvarez-Fernandez and Medema, 2010).

Interestingly, during the checkpoint exit, an increase in cyclin B/CDK1 is required for mitotic

entry and as shown here correlates with an increase in Ser301 phosphorylation and a

decrease in Ser317 phosphorylation. Cyclin A siRNA or CDK inhibitors, both of which are

known to affect the checkpoint exit, reduced Ser301 phosphorylation. The specificity of the

CDK1i has been shown by others (Ma et al., 2009) and infers that CDK1 is the predominate

kinase required for Ser286, Ser301 phosphorylation. However the results imply that cyclin

A/CDK2 is either directly or indirectly involved in the phosphorylation of the Ser286 and


206

Ser301 sites. The data also confirms the initial publication that these phosphorylations occur

predominantly in mitosis.

Interestingly Ikegami et al. (2008) published that Chk1 activating phosphorylations, Ser317

and Ser345, could occur at the same time as the inhibitory, Ser286 and Ser301,

phosphorylations during a G2 checkpoint. It is not uncommon for active and inhibitory

phosphorylations to occur on the same protein molecule, for example Tyr15 inhibitory

phosphorylation of CDK1 can occur at the same time as the activating phosphorylation on

Thr161. In this situation CDK1 is held inactive but once Tyr15 phosphorylation is removed it

will allow activation of CDK1. As Chk1 is required to be active during a checkpoint, this

suggests that the Chk1 inhibitory phosphorylations are present because the protein is not

fully activated yet or that Ser286 and Ser301 are not the full story required for inhibition. It

was shown here that the activating and inhibitory phosphorylations could occur at the same

time under either normal or checkpoint conditions but that during the checkpoint response

these phosphorylations occurred at markedly opposite levels to one another, further

promoting the theory that Ser286 and Ser301 are inhibitory phosphorylations required

during mitosis and for checkpoint exit. This same observation was not clear in the normal G2

samples. This may be due to the different experimental approach used for these two

experiments - thymidine synchronisation versus the DNA damaging drug Etoposide.

However, if these sites are not inhibitory sites than perhaps they indirectly impact Chk1

activity and are more strongly required to control Chk1 localisation, which is important for

Chk1 G2/M function. It has been reported that, Ser286 and Ser301 are phosphorylated to


207

permit Chk1 localisation in the cytoplasm (Enomoto et al., 2009), which most likely promotes

further cyclin B/CDK1 activation and consequently mitotic entry. On the other hand, active

Chk1 phosphorylated on Ser317 and Ser345 is predominantly nuclear (Jiang et al., 2003). We

and others have shown that the alanine mutants of Ser286 and Ser301 localise to the nucleus

(Enomoto et al., 2009), possibly resembling the active form of Chk1. Unlike the published

data, in this study WT Chk1 localised similarly to the SS286,301AA double mutant and not

evenly throughout the cell as reported by others. This could be a result of the increased

phosphorylation of Ser317 observed on wildtype Chk1 and Chk1 Ala mutants (Figure 6.7). As

both constructs, WT and SS286,301AA double mutant, localised to the nucleus (Figure 6.9)

this indicates that both are representative of the active form of Chk1, thereby functioning

accordingly and delaying mitotic entry. Therefore it is possible that the reason the

SS286,301AA double mutant does not delay mitotic entry when compared to the control is

because the WT construct is acting in a similar fashion to delay mitotic entry.

In summary, I have shown that the negative regulation of Chk1 by cyclin A/CDK2 during G2

phase is unlikely to involve phosphorylation on Ser286 and Ser301. These sites appear to be

major cyclin B/CDK1 phosphorylation sites that are catalysed during mitosis. Thus in the next

Chapter I have attempted to investigate the mechanisms by which cyclin A/CDK2 regulates

Chk1 activation.

208

Chapter 7

Identification of other G2 phase pathways

regulated by cyclin A/CDK2

Chapter 7 –Other cyclin A/CDK2 regulated pathways

209

7.1 INTRODUCTION

In the previous chapters I have presented evidence that Chk1 is a contributor to the cyclin

A/CDK2 dependent G2 phase delay, however the incomplete recovery of the cyclin A/CDK2

dependent G2 delay by Chk1 siRNA or Chk1i demonstrated that other pathways also

contribute to the cyclin A/CDK2 dependent G2 delay. One possible candidate is Plk1. Plk1 is

an essential mitotic kinase that is also required for G2/M progression. Plk1 has a role in

mitotic entry and in exit from a G2 phase checkpoint, similar to that found by us for cyclin

A/CDK2. In addition, others had demonstrated a reduction in Plk1 activity with cyclin A

depletion (Fung et al., 2007). Re-introduction of a constitutively active Plk1 construct does

not rescue the cyclin A/CDK2 dependent G2 phase delay and monopolar asters that are

diagnostic of Plk1 inhibition (van Vugt et al., 2004b, Sumara et al., 2004, Hanisch et al., 2006)

are not evident in cyclin A depleted cells, suggesting that Plk1 activity is not affected by cyclin

A/CDK2 depletion or inhibition.

Plk1 structure includes a unique conserved region known as the polo box domain (PBD) that

governs Plk1 function and localisation. The PBD recognises a phospho-specific motif on Plk1

substrates and therefore Plk1 is ‘primed’ by phosphorylation events catalysed by other G2/M

phase kinases ( Lee et al., 1999, Jang et al., 2002a, Seong et al., 2002, Elia et al., 2003b, Park

et al., 2010). Phosphorylation of these sites produces a binding site for Plk1 through its PBD

and allows for direct contact of Plk1 with its substrate. CDK1 is known to regulate Plk1

localisation by the phosphorylation of many Plk1 substrates ( Qi et al., 2006, Wong and Fang,


210

2007, Zhang et al., 2009a). As CDK1 and CDK2 have similar substrate specificities it is possible

that cyclin A/CDK2 may also prime Plk1 targets, however this has not been investigated.

Cdh1 is one of two co-activators of the E3 ubiquitin ligase APC/C (Anaphase-promoting

complex/cyclosome). Although Cdh1 is specifically required for APC/C function in late mitosis

and G1 phase (Schwab and Dreyer, 1997, Sigrist and Lehner, 1997, Visintin et al., 1997, Fang

et al., 1998b, Kramer et al., 1998, Zachariae et al., 1998b, Jaspersen et al., 1999), Cdh1

targets the destruction of many important G2/M proteins including Plk1. This function of

Cdh1 as well as others is required during the G2 phase checkpoint (Bassermann et al., 2008).

Regulation of Cdh1 occurs by cyclin/CDK inhibitory phosphorylation and Cdc14B phosphatase

that removes these phosphorylations, stimulating APC/C binding and activation. In budding

yeast the non-phosphorylated form of Cdh1 is able to bind and activate APC/C (Kramer et al.,

2000). Both cyclin A/CDK2 and cyclin B/CDK1 can regulate Cdh1 (Sorensen et al., 2001) and

cyclin A/CDK2 is reported to regulate Cdh1 in S phase (Lukas et al., 1999a). Without cyclin A’s

negative regulation of APC/C, premature degradation of APC/C targets may occur, including

cyclin A itself (Geley et al., 2001), cyclin B1 (King et al., 1995), Aurora A or B (Stewart and

Fang, 2005, van Lueken et al., 2009), Plk1 (Lindon and Pines, 2004), FoxM1 (Laoukili et al.,

2008a, Park et al., 2008) and the other APC co-activator Cdc20, (Prinz et al., 1998, Pfleger and

Kirschner, 2000), all of which will ultimately affect mitotic entry.

Cdh1 also has a role during the G2 phase checkpoint arrest, where when activated during the

checkpoint, it leads to the destabilisation of Plk1 (Bassermann et al., 2008). The reactivation

of Cdh1, required for checkpoint arrest, was shown to be a consequence of the


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dephosphorylation of the cyclin A/CDK2 phosphorylation sites on Cdh1 by Cdc14B

(Bassermann et al., 2008). Based on this data, it is important to determine if premature

activation of Cdh1 and therefore the premature degradation of its known targets could be

contributing to the cyclin A/CDK2 dependent G2 delay.

While many Cdh1 substrates are degraded during the checkpoint, one of the more important

targets of APC/CCdh1, Claspin, is not. Claspin is an adapter protein of Chk1 and is essential for

ATR phosphorylation and activation of Chk1 during a checkpoint response (Kumagai and

Dunphy, 2000, Lin et al., 2004, Sorensen et al., 2004, Liu et al., 2006, Wang et al., 2006). In

response to damage, Chk1 phosphorylation on Ser317 and Ser345 requires ATR to

phosphorylate Claspin in the Chk1 binding domain to facilitate Claspin-Chk1 binding and

further phosphorylation of Chk1 by ATR (Bennett and Clarke, 2006). Although Claspin is a

Cdh1 substrate, its stability is protected during the DNA damage response by the

deubiquitination enzyme, Usp28. This is so Claspin can maintain the checkpoint by sustaining

Chk1 activation (Bassermann et al., 2008). Claspin turnover is also controlled by SCFβTrCP, and

the deubiquitinase USP7 can block this ubiquitination-mediated degradation (Faustrup et al.,

2009). βTrCP degradation of Claspin requires Plk1 dependent phosphorylation (Peschiaroli et

al., 2006), however the mechanism for Plk1 binding to Claspin is unclear. These multiple

pathways provide a flexible means to regulate the important ATR/Chk1 pathway (Faustrup et

al., 2009). Claspin stability is variable during an unperturbed cell cycle, but its levels are

known to follow a similar pattern to pChk1, which are present from S phase and continue

until they decline with mitotic entry (Bennett and Clarke, 2006). The constant degradation

and turnover of Claspin governs its function during the checkpoint, and this mechanism may


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similarly regulate its activity during an unperturbed cell cycle. The phosphorylation of Claspin

by ATR on Ser916, which facilitates Chk1 binding and it’s phosphorylation by ATR, is present

at low levels within an asynchronous population (Bennett et al., 2008). Therefore, it is likely

that pChk1, required during an unperturbed cell cycle and hence during the cyclin A/CDK2

dependent G2 delay, requires Claspin function as it does in a checkpoint.

In this chapter I have investigated the potential contribution of Cdh1, Plk1 and Claspin during

a cyclin A/CDK2 dependent G2 delay. In detail, the specific ability of cyclin A/CDK2 to affect

Plk1 PBD functions, Cdh1 degradation and Claspin stability which impacts pChk1s role have

been examined.

7.2 RESULTS

7.2.1 Plk1 inhibition resembles cyclin A/CDK2 depletion/inhibition.

Plk1, like cyclin A/CDK2, is important for G2 phase and treatment with Plk1 inhibitor (Plk1i)

produces a phenotype similar to cyclin A/CDK2 depletion/removal. Previously, cells depleted

of cyclin A or inhibited for CDK2 were shown to produce a mitotic delay associated with off-

centred and tilted mitotic spindles (Beamish et al., 2009). Inhibition of Plk1 can lead to the

formation of monopolar asters (Sunkel and Glover, 1988, Llamazares et al., 1991, Lane and

Nigg, 1996, Qian et al., 1998, Sumara et al., 2004, van Vugt et al., 2004b, Peters et al., 2006,

McInnes et al., 2006, Lenart et al., 2007), this phenotype was observed when we treated

asynchronous cells with Plk1i and assessed after 18 h (data not shown), when Plk1i (BI 2536)

was used to treat late G2 phase cells, bipolar spindles were commonly observed. However,


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60% of these bipolar mitotic spindles were off-centred (Figure 7.1 A and E), reminiscent of

the effect caused by cyclin A depletion.

In addition to producing a similar mitotic phenotype, Plk1i treatment also delayed mitotic

entry (Figure 7.1 B) when compared to a DMSO control (crossed line). The IC50 for this Plk1

inhibitor is 0.83 nM (Steegmaier et al., 2007) and in this experiment two different

concentrations of Plk1i were tested. While 1 μM treatment with Plk1i caused a slight delay

(thin dashed line), 5 μM treatment (thick dashed line) delayed G2 phase to a similar extent as

cyclin A siRNA (A3) treatment (solid line). The concentration required to delay mitotic entry

was significantly higher than the concentrations of Plk1i, approximately 100 nM, required to

cause mitotic spindle defects (Lenart et al., 2007, Haupenthal et al., 2012). It is suspected

that the higher concentrations used here to delay mitotic entry, were required to inhibit not

only Plk1 but possibly other Plks.

The polo box domain (PBD) of Plk1 is required to localise Plk1 correctly during the cell cycle

and facilitate its function. Over expression of a PBD-GFP (PBD) construct acts as a dominant

mutant, blocking endogenous Plk1 from binding to its normal specific locations (Hanisch et

al., 2006). Using a PBD-GFP construct we were able to determine the involvement of Plk1

kinase activity and correct localisation of Plk1 in normal G2/M progression. HeLa cells

transfected with either a GFP control vector or PBD-GFP (PBD), synchronised with thymidine

block release, were analysed for mitotic entry by live cell time lapse microscopy. PBD

overexpression slowed mitotic entry significantly compared to the GFP control (Figure 7.1 C),

similar to the effect of Plk1i and cyclin A depletion (Figure 7.1 B). The expression of the


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constructs was confirmed by immunoblotting (Figure 7.1 C; immunoblot). This delay caused

by the PBD-GFP construct suggested that both the kinase activity and localisation of Plk1 are

important for normal G2/M progression. In addition, the overexpression of PBD-GFP

confirmed that results seen with Plk1i treatment was due to inhibition of Plk1 and was less

likely a consequence of the inhibition of other Plks. Cells transfected with PBD-GFP for 24 h

were collected and stained to confirm expression of the construct and with α-tubulin to

determine the position of the mitotic spindle. As can be seen in Figure 7.1 D the mitotic

spindle (red) is off centred within the cell. This is comparable to the Plk1i treated cell in

Figure 7.1 A. The quantification of this phenotype revealed that PBD expression increased the

proportion of off-centred spindles by up to 60 % of the bipolar spindles observed, and was

similar to the proportion observed with the Plk1i (Figure 7.1 E). Thus it appears that cyclin

A/CDK2 and Plk1 are acting in a common pathway to regulate proper spindle formation, and

this requires correct localisation as well as Plk1 activity.

7.2.2 Cyclin A siRNA causes mislocalisation of Plk1.

To determine the specific effect of cyclin A/CDK2 on Plk1 function and/or localisation, cyclin

A depleted cells were analysed for their Plk1 protein levels and its localisation. Whilst Plk1

levels were unchanged in cyclin A depleted G2 phase samples compared to the control

(Figure 7.2 A), the levels of Plk1 and the activating phosphorylation on Plk1 pT210 were

reduced by 60 % in cyclin A depleted cells arrested in mitosis with the microtubule

depolymerisation drug, nocodazole (Figure 7.2 B).


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Figure 7.1 Plk1 inhibition causes phenotypes similar to cyclin A/CDK2 depletion/inhibition.

A. Asynchronously growing HeLa cells were treated with 5 μM Plk1i or DMSO (CONT). 24 h post

treatment, cells were stained for α-tubulin, ACA (anti-centromere antibody), and DAPI for DNA.


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B. Thymidine synchronised HeLa cells were either transfected with cyclin A A3 siRNA (solid line),

treated with DMSO (CONT: crossed line) or with 1 µM (thin dashed line) and 5 µM Plk1i (BI

2536) (thick dashed line). The inhibitor and DMSO were added 5 h post synchrony release (early

G2 phase) then followed by time lapse microscopy. The cumulative mitotic index was scored.

The result was confirmed by conducting the experiment in triplicate and the best

representation was chosen for publishing.

C. HeLa cells were transfected with GFP or PBD-GFP (PBD) expression plasmids and synchronised

with thymidine. 5 h post thymidine release, cells were followed by time lapse microscopy. The

cumulative mitotic index of the GFP expressing cells was scored. After the completion of the

microscopy the cells were collected, lysed and immunoblotted to check for the expression of

the transfected constructs. Samples were immunoblotted with GFP antibody. The result was

confirmed by conducting the experiment in triplicate and the best representation was chosen

for publishing.

D. HeLa cells were transfected with GFP or PBD-GFP expression plasmids collect 24 h transfection

and stained for GFP, α-tubulin and DAPI for DNA. Mitotic cells were observed and imaged for off

centred spindles. Arrows indicate the centrosomal localisation of PBD-GFP. Data contributed by

Dr Brian Gabrielli.

E. Quantitation of the percentage of cells containing off centred mitotic spindles in experiments

similar to those shown in A and D. Data from three independent experiments, each counting at

least 100 mitotic cells, has been combined and error bars represent SEM. Data contributed by

Dr Brian Gabrielli.


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We confirmed that reintroduction of Plk1 could not overcome the cyclin A dependent delay

as previously reported (Figure 7.2 C; (Fung et al., 2007)). We also analysed the effect of

cyclin A depletion and simultaneous overexpression of Myc-tagged Plk1 (Myc-Plk1) on cells

exiting a G2 phase checkpoint arrest. Cell cultures were treated with Etoposide to impose a

G2 phase checkpoint arrest, then released with caffeine for 10 h, fixed and scored as either

G2 phase or mitotic cells by microscopy. Scoring demonstrated that a lower percentage of

cells were able to enter mitosis with co-transfection of cyclin A siRNA, and that over

expression of Myc-Plk1 did not reduce this delay significantly (Figure 7.2 D). Collectively,

these data demonstrate that Plk1 activity was not affected by cyclin A/cdk2

depletion/inhibition.

The ability of PBD to displace endogenous Plk1 binding and produce similar phenotypes as

cyclin A depletion indicated that perhaps the mislocalisation of Plk1 was the main

consequence of cyclin A depletion. This would explain why Plk1 overexpression could not

overcome cyclin A dependent G2 phase delay, as it was possibly due to the inability of Plk1 to

bind its critical substrates because of the loss of pre-priming of Plk1 substrates by cyclin

A/cdk2. To assess whether Plk1 substrate binding and localisation was dependent on cyclin

A/cdk2, the localisation of Plk1 was observed in cells depleted of cyclin A. Plk1 has very

distinctive mitotic localisation, specifically at the centrosomes and the kinetochores

(Golsteyn et al., 1995, Ahonen et al., 2005, Wong and Fang, 2005, Qi et al., 2006). We

examined mitotic Plk1 localisation with or without cyclin A knockdown. In nonsense siRNA

transfected controls, Plk1 localised normally to the centrosomes and kinetochores, whereas

cyclin A siRNA caused a noticeable reduction in Plk1 localisation at these sites (Figure 7.3 A


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Figure 7.2 Plk1 overexpression did not contribute to delayed mitotic entry caused by cyclin A siRNA

treatment.

A. Asynchronously growing HeLa cells were transfected with 50 nM of cyclin A siRNA A3 or a

nonsense siRNA control (N). Cells were treated with thymidine, released for 6 h, collected, lysed

and immunoblotted for Plk1, cyclin A, pMek1 T286, PY15 and PCNA.


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B. Asynchronously growing HeLa cells were transfected with 50 nM of cyclin A siRNA A3 or a

nonsense siRNA control (N). Transfected cells were treated with nocodazole (0.25 μM) for

approximately 16 h, then collected, lysed and immunoblotted for cyclin A, Plk1, Plk1 (pT210)

and PCNA.

C. Asynchronously growing HeLa cells were transfected with 50 nM of cyclin A siRNA A3 or a

nonsense siRNA control (N). Cells were also co-transfected with Myc-Plk1 or an empty Myc

vector control, and GFP to identify the transfected cells. After transfection cells were

synchronised with thymidine. 6 h post release cells were followed by time lapse microscopy.

The cumulative mitotic index of GFP expressing cells was scored. After the completion of

microscopy the cells were collected, lysed and immunoblotted for Plk1, cyclin A and PCNA to

check for the knockdown and/or expression within each sample. PCNA was used to determine

equal loading. The result was confirmed by conducting the experiment in triplicate and the best


D. Asynchronously growing HeLa cells were transfected with 50 nM of cyclin A siRNA A3 or a

nonsense siRNA control (N) and/or Myc Plk1 or the empty vector control. 6 h post transfection

cells were treated with Etoposide (1 μM) for 16 h. After Etoposide incubation, caffeine was

added to force cells into mitosis. 10 h post caffeine addition cells were fixed and stained for

Myc, α-tubulin and DAPI for DNA. Only Myc tag expressing cells were scored as either remaining

in G2 phase or progressing into mitosis. Percentage of the total cell number is presented for

each condition and approximately 200 cells were counted per condition. Results are

representative of three independent experiments.


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and B). Although specific co-staining with markers such as γ-tubulin for centrosomes and ACA for

kinetochores has not been performed it is well recognised in the literature that Plk1 localises to these

localisations and the images of Figure 7.3 A and B clearly resemble these unique localisations. The

regulation of a pool of Plk1 during mitosis by cyclin A/cdk2 suggested that cyclin A/cdk2 may

also affect Plk1 binding to specific proteins during G2 phase, although it was not possible to

readily discern specific localisation of endogenous Plk1 in G2 phase.

To determine if normal G2 phase localisation of Plk1 at the centrosomes was affected by

cyclin A/cdk2, a preliminary analysis using the PBD-GFP construct in G2 phase synchronised

cells was conducted. G2 phase cells were identified by their separating centrosomes, as

indicated by γ-tubulin staining. Immunofluorescence of PBD-GFP showed that PBD binding

was not affected by cdk2 inhibition (Figure 7.3 C). The GFP only controls showed no

localisation to the centrosomes, whereas the PBD-GFP construct was evident at the

centrosomes with or without cdk2i treatment. Therefore cyclin A/cdk2 only affects the

mitotic localisation of Plk1 which most likely leads to the similar mitotic ‘rotating spindle’

phenotype. Although this data demonstrated that G2 phase centrosome Plk1 localisation is

not affected by cyclin A/cdk2 this was a preliminary experiment and was not quantified. In

addition, the timing of sample collection and treatment with cdk2i could contribute to this

result. Therefore, further experimentation would be required to conclusively show that Plk1

centrosomal localisation during G2 phase is not affected by cdk2i.


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Figure 7.3 Cyclin A affects Plk1 localisation at the centrosomes and kinetochores in mitosis.


nonsense control (CONT). 24 h post transfection cells stained for Plk1 and DAPI for DNA.

Images were taken of cells in prophase.

B. Cells were treated as in A with additional co-staining of α-tubulin. Images were taken of

cells in metaphase. Arrows indicate centrosomes.

C. HeLa cells were transfected with GFP or PBD-GFP and synchronised with thymidine. 5 h post

thymidine release (S phase), cells were treated with either DMSO or 4 μM of CDK2i. At 8 h

post thymidine release cells were fixed with methanol and stained for γ-tubulin (red) and

DAPI for DNA (blue). Images were taken of cells in G2 phase, where evidence of separating

centrosomes was apparent. Arrows indicate the position of the centrosomes.

Exposure times for images within each experiment were kept consistent. Images were taken using a

Zeiss Apotome widefield microscope with 100x oil objective.


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Figure 7.4 Cdh1 levels are reduced with cyclin A/CDK2 depletion/inhibition.


nonsense control (N). 24 h post transfection cells were collected, lysed and immunoblotted

for cyclin A, Cdh1, and PCNA. . The bar graph shows the quantification of Cdh1 levels in

samples collected 24 h post transfection with siRNA, cyclin A siRNA (A1 and A3) or nonsense

siRNA (NS). siRNA treated samples were also compared to an untransfected control (Con).

Immunoblots were analysed using ImageJ software. Results were formulated from 5

independent experiments. The asterisks indicate p value of <0.05.

B. Asynchronously growing HeLa cells were transfected with 50 nM of cyclin A siRNA A1 and

A3, a nonsense control (N) or a lipofectamine treated only control (C). Cells were

synchronised with thymidine. 5 h post release cells were treated with or without 20 µM of

MG132 for 2h. All samples were then collected 7 h post thymidine release. All samples were

collected in G2 phase at 7 h post thymidine release. Samples were lysed and

immunoblotted for Cdh1, cyclin A, and PCNA.

C. Asynchronously growing HeLa cells were transfected with 50 nM of cyclin A siRNA A3 or a

nonsense control (N). 24 h post transfection cells were stained for Cdh1, cyclin A and DAPI

for DNA. Images on the left are the merged images.

D. HeLa cells were blocked in G2 phase with Etoposide (1 µM) for approximately 16 h. Cells

were then treated +/- CDK1i (9 µM) or CDK2i (4 µM) for the indicated time points.

Asynchronous (As) and Etoposide (et) only treated cells were used as controls. All samples

were lysed and immunoblotted for Cdh1, the G2 phase marker, PY15 and PCNA.


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7.2.3 Cdh1 level is reduced with cyclin A depletion.

Cdh1 regulates Plk1 levels, and Cdh1 activity has been reported to be regulated by cyclin

A/CDK1/2 (Zhou et al., 2003, Reber et al., 2006). To investigate the effect of cyclin A

depletion on Cdh1 levels, we examined the effect of cyclin A siRNA on Cdh1 levels. Cdh1

levels were reduced by approximately 50% in cyclin A depleted cells compared to a nonsense

transfected control (Figure 7.4 A). Quantification of Cdh1 levels confirmed the significant

decrease in Cdh1 levels seen with cyclin A knockdown (Figure 7.4 A; bar graph). Cdh1 levels

were also reduced in G2 phase synchronised cells but to a lesser extent (Figure 7.4 B; lanes 1-

4). In addition, immunofluorescence data showed that cyclin A reduction accordingly caused

a reduction in Cdh1 staining when compared to controls (Figure 7.4 C). The

immunofluorescent staining did not appear to demonstrate any Cdh1 mislocalisation caused

by cyclin A depletion. To determine whether cyclin A/CDK2 was affecting proteasome

mediated degradation of Cdh1, we treated controls and cyclin A depleted cells with the

proteasome inhibitor, MG132, for 2 h. Treatment of G2 phase cells with MG132 increased

Cdh1 levels in nonsense treated cells, demonstrating that Cdh1 is normally controlled by

proteasome mediated degradation (Figure 7.4 B). MG132 treatment also increased Cdh1

levels in the cyclin A depleted cells, and increased the levels by a similar extent to the control

samples. This was also noted by comparing the quantification of the appropriate untreated

samples to the MG132 treated sample. This revealed an average mean increase in Cdh1

levels with MG132 treatment of 0.67 with a standard deviation of 0.08. This suggests that

cyclin A/CDK2 was not affecting the stability of Cdh1 through proteasome mediated

degradation.


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To determine whether the effects of cyclin A depletion were due to inhibition of the cyclin

A/CDK2 or CDK1 complex, Etoposide treated G2 phase checkpoint arrested cells were

treated with either the CDK2 or CDK1 inhibitor (CDK2i and CDK1i), and levels of Cdh1

assessed at several time points after caffeine addition. CDK2i treatment caused a reduction

in Cdh1 levels after 5 h whereas the CDK1i had little obvious effect (Figure 7.4 D). Although

the effect of the CDK2i was not immediate, it did indicate that cyclin A/CDK2 and not CDK1

was involved in regulating the level of Cdh1. In addition, the 5 h effect of the CDK2i on Cdh1

levels suggests that it might be a transcriptional effect that cyclin A/CDK2 has on Cdh1

(Figure 7.4 D). More recent work from the Gabrielli Laboratory shows that it is not a stability

effect that cyclin A/CDK2 has on Cdh1. To determine whether the reduction in Cdh1 was

contributing to the cyclin A/CDK2 dependent G2 delay, Cdh1 was reintroduced into cells

depleted of cyclin A. U2OS cells expressing an inducible Cdh1 were transfected with

nonsense or cyclin A siRNA (A3) and then synchronised with thymidine. Upon thymidine

release, cells were treated with tetracycline to induced Cdh1 expression. Time lapse

microscopy was then used to determine the timing of mitotic entry. As previously reported,

cyclin A siRNA (A3) caused a G2 phase delay but Cdh1 induction did not positively impact the

duration of this delay (Figure 7.5 A; compare solid line with long dashed line). Likewise CDK2i

produced a G2 phase delay (Figure 7.5 A; vertical dashed line), which was also unaffected by

Cdh1 induction. Induction of Cdh1 alone produced a G2 phase delay similar to cyclin A

depletion/CDK2 inhibition. Immunoblotting of samples induced for various times showed a

steady increase of Cdh1 that reaches a maximum at 6 h (Figure 7.5 B). Compared to the 50 %

reduction of Cdh1 levels with cyclin A knockdown (Figure 7.4 A), the level of Cdh1 induction


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Figure 7.5 Cdh1 overexpression does not rescue the cyclin A/CDK2 G2 phase delay and itself inhibits

mitotic entry.

A. A U2OS cell line with a tetracycline repressed Myc-Cdh1 vector was transfected with nonsense

control (N) or cyclin A (A3) siRNA. Post transfection, cells were synchronised with thymidine.

Upon thymidine release, tetracycline was removed from the media to allow Cdh1 induction. For

CDK2i treated samples, 4 µM of inhibitor was added at 6 h post thymidine release. After CDK2i

addition cells were followed by time lapse microscopy. The cumulative mitotic index was

scored. The result was confirmed by conducting the experiment in triplicate and the best


B. U2OS cells that stably express Myc-Cdh1 were maintained in tetracycline containing media to

repress Myc-Cdh1 expression. Tetracycline was removed in induced Myc-Cdh1 expression by

washing cells three times with warmed PBS. The media was then replaced onto cells without

the addition of tetracycline. Cells were then collected at the indicated time points after

tetracycline removal. An untreated sample (uninduced) that did not express Myc-Cdh1 due to

incubation with tetracycline was used as a control. Quantification was carried out using ImageJ

software and samples were standardised to the PCNA loading control and equalised to the

uninduced control.

C. Samples from A were collected after the completion of microscopy, lysed and immunoblotted.

Samples were immunoblotted for Cdc25B, cyclin B, pChk1, cyclin A, Cdh1, PY15 and PCNA.

D. HeLa cells were co-transfected with either an empty vector control (MYC), Myc-Cdh1 WT (Cdh1

WT) or Myc-Cdh1 AAA (Cdh1 AAA), and a GFP expression vector. Cells were synchronised with

thymidine, released and followed by time lapse microscopy. Only the GFP expressing cells were

scored. The cumulative mitotic index was scored. After the microscopy was complete cells were

collected, lysed and immunoblotted for Cdh1, Plk1, cyclin A and PCNA. The result was confirmed


229

by conducting the experiment in triplicate and the best representation was chosen for

publishing.

E. U2OS cells with a tetracycline inducible Myc-Cdh1 vector were either induced for Cdh1 (-

tetracycline) or not (+ tetracycline). 24 h post induction samples were collected and fixed for

FACS with ethanol. FACS was conducted to determine the cell cycle profiles of these samples.

The dashed lines indicate the G1 and G2/M phase populations. The S phase population is in

between the lined section.


230

at 6 h was 8 fold greater than endogenous levels. Thus, the lack of rescue may simply reflect

the massive effects of Cdh1 over expression. The effects of Cdh1 overexpression on G2/M

phase regulatory proteins revealed that induction of Cdh1 decreased the levels of Cdc25B,

cyclin A and cyclin B1, and in the level of the inactive form of CDK1 (PY15), although this is a

consequence of the reduced levels of cyclin A and B (Figure 7.5 C). Cyclin A and cyclin B1 are

known targets of Cdh1 (Raff et al., 2002, Zhou et al., 2002, Zhou et al., 2003). Interestingly

there was also a very minor but noticeable reduction in pChk1 levels with Cdh1 induction.

To determine whether these results were specific for the inducible Cdh1 cell line, we

conducted transient transfection of Cdh1 in HeLa cells. We used a wild type Myc-Cdh1 (Cdh1

WT) construct and a constitutively active triple Ala Cdh1 mutant, Myc-Cdh1 AAA (Cdh1 AAA),

which is a mutation of the cyclin binding site of Cdh1 (amino acids 445-447, Arg-Val-Leu all

mutated to Ala) and cannot be inhibited by cyclin A/CDK2 (Sorensen et al., 2001). When

transiently over expressed, both constructs produced a similar mitotic delay (Figure 7.5 D) as

observed with the inducible Cdh1 (Figure 7.5 B). Cdh1 overexpression caused a decrease in

Cdh1 substrates cyclin A and Plk1, although Plk1 levels were only reduced in the WT and not

the AAA mutant as reported previously (Sorensen et al., 2001). Lastly, FACS analysis of the

inducible U2OS cell line confirmed that a large induction of Cdh1 levels, such as that seen at

24 h (Figure 7.5 B), was able to cause a large accumulation of cells in G2/M phase (Figure 7.5

E) and correlates to this microscopy data.


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7.2.4 Reduction of Cdh1 does not affect the cyclin A/CDK2 dependent G2 delay.

Cdh1 overexpression reduced the levels of several important G2/M phase regulatory

proteins, and therefore we could not rescue the effect of cyclin A/CDK2 depletion using the

high level overexpression from these expression vectors. To directly determine whether the

reduced levels of Cdh1 seen with cyclin A/CDK2 depletion did contribute to the G2 phase

delay, Cdh1 levels were depleted using specific Cdh1 siRNA. Initially we assessed the efficacy

of the Cdh1 siRNA to deplete Cdh1 levels, and its functional consequence on the level of the

APCCdh1 substrate, Plk1. Only the most effective Cdh1 siRNA (Cdh1 A siRNA) is shown in the

data present, however several Cdh1 siRNA were tested and all resulted in a similar result as

in Figure 7.6 A. The Cdh1 siRNA reduced Cdh1 levels by approximately 70 % and increased

the levels of Plk1 (Figure 7.6 A). We next determined the rate of mitotic entry of cells

depleted of cyclin A and Cdh1 using time lapse live cell microscopy. Cdh1 depletion caused a

minor delay in G2 phase compared to cyclin A depletion, and co-depletion had no effect on

the cyclin A dependent effect (Figure 7.6 B). Immunoblots show that Cdh1 siRNA depleted

Cdh1 levels to below the levels caused by cyclin A siRNA treatment (Figure 7.6 C).

Depletion of Cdh1 with or without cyclin siRNA treatment during a checkpoint exit revealed

similar results to the same treatment of asynchronous cycling cells (Figure 7.6 D and 7.6 C).

These data demonstrate that the reduction in Cdh1 levels observed with cyclin A depletion

was unlikely to contribute to the cyclin A/CDK2 dependent G2 phase delay.


232

7.2.5 Claspin is required during the cyclin A/CDK2 dependent G2 phase delay to regulate

pChk1 levels.

Claspin is a co-activator of Chk1, required for Chk1 phosphorylation by ATR during checkpoint

activation and is also a Cdh1 substrate. During a checkpoint however Claspin is stabilised by

the deubiquitination enzyme USP28. To determine the involvement of Claspin in the cyclin

A/CDK2 dependent G2 delay and its ability to impact pChk1 levels, cyclin A depleted cells

were used.

Claspin was modestly increased (20-30%) in cyclin A siRNA treated samples (Figure 7.7 A).

Quantification of Claspin levels confirmed the significant increase in Claspin levels seen with

cyclin A knockdown (Figure 7.7 A; bar graph). As a Cdh1 target, Claspin levels were increased

or decreased when Cdh1 was inducted or removed respectively (Figure 7.7 B). Cells arrested

in mitosis with nocodazole were used as a negative control, as Claspin is known to be

degraded in mitosis (Mamely et al., 2006). Claspin is required for Chk1 activation therefore it

seemed probable that the increased Claspin detected in cyclin A/CDK2 dependent G2 phase

delay was responsible for enhanced pChk1 phosphorylation (Chapter 5 Figure 5.1 A) seen in

the cyclin A/CDK2 dependent G2 phase delay. siRNA specific for Claspin was used to assess

the effects of Claspin depletion alone on G2/M phase progression and to determine whether

Claspin depletion could rescue the effect of cyclin A/CDK2 depletion. Specific siRNA

decreased Claspin levels by 50 % which resulted in a decrease in pChk1 levels by

approximately 40 % in Etoposide treated HeLa cells (Figure 7.7 C). This also led to a reduced

G2 phase checkpoint arrest, revealed by the increased G1 phase population/2n population as

seen by FACS (Figure 7.7 C; FACS histograms). This data shows that Claspin is required for


233


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Figure 7.6 Cdh1 siRNA does not rescue the cyclin A/CDK2 G2 phase delay and delays mitotic

progression also.

A. Asynchronously growing HeLa cells were transfected with 40 nM of Dharmacon smart pool

Cdh1 siRNA (Cdh1 A) or a nonsense control (N). 24 h post transfection cells were collected, lysed

and immunoblotted for Cdh1, Plk1 and PCNA. Quantification was carried out using ImageJ

software. Samples were equalised to their PCNA loading controls and Cdh1 siRNA treatment

was standardised to the control.

B. HeLa cells transfected with either nonsense (N: crosses) or cyclin A siRNA A3 (dashed line) were

either co-transfected or not with 40 nM Cdh1 siRNA (Cdh1 A). Cells were thymidine

synchronised and followed by time lapse microscopy 6 h after synchrony release. Cells were

scored for entry into mitosis, over 200 cells were counted in each case. The result was

confirmed by conducting the experiment in triplicate and the best representation was chosen

for publishing.

C. After the completion of B, samples were lysed and immunoblotted for Cdh1, cyclin A and PCNA.

D. Samples were transfected as in B and were blocked in the G2 phase checkpoint with 16 h

Etoposide (1 μM) treatment after which caffeine (5 mM) was added to drive cells into mitosis.

The mitotic entry of cells was followed immediately after caffeine addition by time lapse

microscopy. Cells were scored for entry into mitosis and over 200 cells were counted in each

case. The result was confirmed by conducting the experiment in triplicate and the best



235

Figure 7.7 Claspin is increased with cyclin A depletion.

A. Asynchronously growing HeLa cells were transfected with 50 nM of cyclin A siRNA A1 and

A3, a nonsense siRNA control (N) or a lipofectamine treated only control (C). 24 h post

transfection cells were collected, lysed and immunoblotted for Claspin and PCNA. ImageJ

was used to quantify the intensities of Claspin bands, all were normalised to PCNA bands.

The bar graph shows the quantification of Claspin levels taken from 4 independent

experiments. HeLa samples were collected 24 h post transfection with siRNA, cyclin A siRNA


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(A1 and A3) or nonsense siRNA (NS). Immunoblots were analysed using ImageJ software.

Double asterisk indicates p value of <0.01.

B. Myc-Cdh1 tetracycline inducible cells were either induced for Cdh1 (-tetracycline) or not (+

tetracycline). 24 h post induction samples were collected. HeLa cells were transfected with

nonsense siRNA or Cdh1 siRNA (Cdh1 A) or treated with 0.25 µM nocodazole (NOC) for 24 h.

After all treatments, samples were collected, lysed and immunoblotted for Claspin, Cdh1

and PCNA.

C. HeLa cells were transfected or not with Claspin siRNA. 6 h post transfection cells were

treated with 1 μM Etoposide for 16 h, after which they were collected. Samples were fixed

and analysed by PI FACS and lysed and blotted for Claspin, pChk1 and PCNA. The FACS

profiles are shown and the 2n and 4n peaks are labelled.


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Chk1 activation and checkpoint arrest. The incomplete reduction in pChk1 levels or of the G2

checkpoint arrest suggests that the Claspin levels need to be further reduced to see more of

an effective bypass of the checkpoint arrest.

To determine the role of Claspin in the cyclin A/CDK2 dependent G2 phase delay, it was

necessary to ensure that Claspin depletion was as complete as possible. To this end, the

effects of multiple rounds of Claspin siRNA treatment were assessed. Both 24 h and 48 h

treatment with Claspin siRNA efficiently depleted Claspin levels, and treatment on

consecutive days effectively reduced Claspin to below detectable levels (Figure 7.8) (Data for

24 h treatment with Claspin siRNA not shown). Immunoblotting for other G2/M phase

markers revealed that all Claspin siRNA treatments increased the level of G2 phase markers

Cdc25B, Wee1 and the inactive form of CDK1/2 (PY15). The level of pChk1 was reduced

compared to the nonsense control, which acts as a positive control for the functional

consequence of Claspin depletion.

To determine the impact of Claspin on the cyclin A/CDK2 dependent G2 delay, cells were

depleted of cyclin A and Claspin individually, or co-depleted of both cyclin A and Claspin.

These cells were then synchronised with thymidine, followed by using time lapse live cell

microscopy and scored for their entry into mitosis. After which, the cells were analysed by

immunoblotting. The siRNA effectively reduced Claspin and cyclin A levels (Figure 7.8 B).

Cyclin A depletion alone increased Claspin and pChk1 levels as shown previously (Figure 7.7 A

and Chapter 5 Figure 5.1 A, respectively) and co-depletion with Claspin siRNA effectively

reduced the pChk1 levels to a similar level as the control (Figure 7.8 B), demonstrating that


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Figure 7.8 Claspin siRNA can reduced pChk1 levels during the cyclin A/CDK2 dependent G2 delay

but cannot rescue the delay.

A. HeLa cells were transfected twice within two consecutive days with either 50 nM Claspin

siRNA or nonsense (CONT) siRNA. 48 h post the initial transfection cells were collected,

lysed and immunoblotted for Claspin, Cdc25B, Wee1, Plk1, pChk1, PY15 and PCNA.


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B. Samples from C were collected after the completion of the microscopy. These samples were

lysed and immunoblotted for Claspin, Wee1, cyclin A, pChk1 and PCNA.

C. HeLa cells transfected with either nonsense (N: circles), cyclin A siRNA A3 (dashed line) were

either co-transfected or not with 40 nM Claspin siRNA. All cells were thymidine

synchronised and followed by time lapse microscopy 5 h after synchrony release. Cells were

scored for entry into mitosis, over 200 cells were counted in each case. The result was

confirmed by conducting the experiment in triplicate and the best representation was



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the increased Claspin levels were likely to be responsible for the increased pChk1 levels seen

in cyclin A/CDK2 depleted cells. In this experiment Claspin siRNA also reduced pChk1 levels

as seen previously in Figure 7.7 C. When the effects of Claspin depletion were examined in

relation to mitotic entry, it was found that the cyclin A dependent G2 phase delay was not

overcome by co-depletion of Claspin. In fact, when Claspin siRNA was used either individually

or in combination with cyclin A siRNA, Claspin siRNA also caused a delay of entry into mitosis

(Figure 7.8 C). This finding supported the previous observation that G2 phase markers are

elevated with Claspin depletion (Figure 7.8 A). The results were not an artefact of off target

effects of the siRNA as four individual Claspin siRNAs, all which produced similar levels of

Claspin depletion, produced similar delays in entry into mitosis. This data suggested that

Claspin was responsible for the increased pChk1 levels in cyclin A depleted cells, but in

addition to its Chk1 regulatory role it may have other functions that affect the timing of

mitotic entry.

7.3 DISCUSSION

In this chapter, I have investigated the mechanism of the cyclin A/CDK2 dependent G2 phase

delay. This included investigation of Plk1, Cdh1 and Claspin, for which the level of each was

shown to be affected by cyclin A siRNA or CDK2i treatment.

The similarities between cyclin A/CDK2 depletion/inhibition and Plk1 inhibition using either a

small molecule inhibitor or overexpression of the polo box domain strongly suggest that Plk1

and cyclin A/CDK2 function in the same regulatory pathway governing both mitotic entry and

localisation of the mitotic spindle. Although the concentration of Plk1i used to implement a


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G2 phase delay was notably higher than required to produce spindle defects such as

monopolar asters. The fact that overexpression of Plk1 PBD (PBD-GFP) also results in a G2

delay suggested that the inhibition of other Plks was not greatly impacted. However, the

provider of the inhibitor, Selleckchem, has estabilished that the Plk1i BI 2536 can inhibit Plk2

and Plk3 but approximately 5 and 10 fold less, respectively. This is unsurprising as Plk2 and

Plk3 reportedly share only 39 and 36% sequence identity, respectively, to Plk1 over the PBD

residues 410–593, but not Plk4 (Chen et al., 2003). In addition, a more recent publication

showed that the PBD of Plk1-3 have overlapping localisations (Park et al. 2010). Collectively,

it appears that the high concentration of Plk1i used is likely affecting other Plk family

members. As Plk3, like Plk1, can regulate mitotic entry and cytokinesis (Ouyang et al. 1999,

Conn et al. 2000) and Plk3 has been shown to bind to Chk2 causing it to be involved in the G2

checkpoint (Bahassi el et al. 2006) means that the inhibition of Plk3 may be strongly

contributing to the results produced in this study when using high concentrations of Plk1i.

Based on the role of Plk1 and the data gathered it was hypothesised that cyclin A/CDK2 is

phosphorylating Plk1 substrates that direct Plk1 binding to its substrates through the polo

box domain of Plk1. Many Plk1 substrates are known to be phosphorylated by cyclin B/CDK1,

which facilitates Plk1 PBD binding (Yamaguchi et al., 2005, Wu and Liu, 2008, Zhang et al.,

2009b). Initial data has shown that whilst the mitotic localisation of Plk1 is affected by loss of

cyclin A, the G2 phase localisation of PBD was not affected by the loss of CDK2 activity (Figure

7.3), however further studies are required to confirm this data and the loss of Plk1 binding to

specific protein substrates. Cyclin A/CDK2 may be required for Plk1 localisation to a subset of

Plk1 substrates; however it clearly does not contribute to the Plk1 dependent separation of


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mother and daughter centrosomes as no monopolar aster phenotype was observed with

cyclin A depletion. A proteomics approach should be undertaken in order to conclusively

determine whether cyclin A/CDK2 affects the binding of Plk1 to specific substrates during G2

phase.

Although the mitotic defects of Plk1 and cyclin A depletion/inhibition may involve as yet

unidentified G2 phase specific roles for Plk1 and cyclin A/CDK2, it may also reflect a

prominent mitotic role for these proteins. The decreased levels and defective localisation of

Plk1 observed in mitotic cells depleted of cyclin A demonstrates that Plk1 is regulated either

directly or indirectly by cyclin A/CDK2 during early mitosis, while cyclin A/CDK2 still has an

active presence. Cyclin A/CDK2 has the ability to regulate Plk1 transcription by the regulation

of FoxM1. If the decreased levels of Plk1 observed in mitosis were due to the inability of

cyclin A/CDK2 to active FoxM1 leading to Plk1 transcription then the effect would have been

seen in G2 phase levels of Plk1. However, it is clear from the data that G2 phase levels of

Plk1 in cyclin A depleted cells were unaffected (Figure 7.2). Interestingly, Wee1 is also

reduced in mitosis rather than in G2 phase. Plk1 is a target of Cdh1 and may account for the

reduction of Plk1 due to regulation of Cdh1 by cyclin A/CDK2. However, Wee1 is not a target

of Cdh1 therefore the reason for reduced Wee1 levels in mitosis is unknown. Although we

have shown that cyclin A/CDK2 depletion/inhibition reduced Cdh1 levels, perhaps the

remaining Cdh1 is more active due to the fact that cyclin A/CDK2 is not present to inhibit

Cdh1 activity, therefore leading to increased reduction of Cdh1 substrates such as Wee1 and

Plk1. The degradation of Plk1 by Cdh1 (Figure 7.5 D) is likely to lead to reduced Plk1 activity,

both of which are required for mitotic exit. Why this occurs in mitosis and not G2 is unknown


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and further work is needed to confirm the activity of Cdh1 under these conditions to prove

this hypothesis. Although it is questionable, if Chk1 is able to negatively regulate Plk1 levels

(Tang et al., 2006), this pathway could also have influenced the results obtained at part of

this study.

This current study found that Cdh1 levels were reduced in cyclin A depleted cells, indicating

that this was not the pathway that cyclin A/CDK2 was affecting to cause reduced Plk1 levels.

Once active in G2 phase Plk1 has the ability to regulate its own transcription by the activation

of FoxM1. This only occurs once cyclin B/CDK1 has become active and it phosphorylates

FoxM1 allowing Plk1 to bind to FoxM1 (Fu et al., 2008). It is likely that the delayed activation

of cyclin B/CDK1 caused by cyclin A/CDK2 depletion/inhibition causes Plk1 to be unable to

bind and activate FoxM1, therefore negatively impacting on Plk1 levels. Although this

appears to be the most likely impact of cyclin A/CDK2 levels, it is possible that other

pathways not yet identified to control Plk1 stability are involved. The lack of data involving

Plk1 stability mechanisms may be an indication that this is a likely possibility.

This reduction in Cdh1 levels is unlikely to be an off target effect of the siRNA as other cyclin

A siRNA gave the same result (data not shown). The use of Etoposide demonstrated that the

effect of CDK2i on Cdh1 levels, which took approximately 5 h, was not a result of Cdh1

destabilisation caused by cyclin A/CDK2 depletion/inhibition. Although there is no real

evidence it does suggest a transcriptional effect by cyclin/CDK2 on Cdh1. In addition,

proteasome degradation of Cdh1 is not impacted by cyclin A/CDK2 as Cdh1 levels increased

in the cyclin A depleted cells as well as the controls. However, it cannot be ruled out that the


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2 h treatment with proteasome inhibitor, MG132, was not sufficient to allow the complete

recovery of Cdh1 levels. Therefore transcriptional control of Cdh1 by cyclin A/CDK2 may

contribute to the decreased levels of Cdh1, however Cdh1 has also been shown to control its

own degradation via APC/C (Listovsky et al., 2004). This was shown by the introduction of

Cdh1 resulting in the increased degradation of itself (Listovsky et al., 2004). Therefore, if it is

possible that depletion of cyclin A can lead to an increase in Cdh1 activation then this may

coincide with the over stimulation of Cdh1 activity leading to its degradation and overall

decrease in the endogenous levels of Cdh1. Based on cyclin A/CDK2s ability to regulate Plk1

levels, further studies using another impartial Cdh1 substrate is required to understand the

stability of Plk1 and the activity status of Cdh1 at this time point.

Manipulation, both the overexpression and depletion, of Cdh1 levels was ineffective at

reducing the cyclin A/CDK2 dependent G2 delay. Over expression of Cdh1 levels is capable of

causing a significant delay in mitotic entry and this correlated with its ability to regulate the

stabilisation of several crucial G2/M proteins such as cyclin B1. This study also used the

constitutively active Cdh1 AAA and Cdh1 WT constructs, both of which caused a mitotic

delay. Cdh1 AAA has previously been known to cause a G1/S delay while the WT did so to a

lesser extent (Sorensen et al., 2001) and this may correlate to the difference in Plk1 levels

seen with overexpression. WT overexpression, unlike Cdh1 AAA overexpression, caused a

reduction of Plk1 levels (one of its known targets) that is required for mitotic entry. One

possible explanation for the difference in Plk1 levels is that Plk1 localises to the cytoplasm in

S and G2 phase (Taniguchi et al., 2002), whereas Cdh1 is active within the nucleus, and our

Cdh1 AAA mutant predominantly localises to the nucleus (Zhou et al., 2003). Hence, it is


245

possible that localisation in separate cellular compartments may underlie the lack of effect

observed. Another important gene that is a reported Cdh1 substrate but was not

investigated in this study due to time constraints and lack of reagents was the transcription

factor FoxM1 (Park et al., 2008). FoxM1 is responsible for the transcription of several G2/M

proteins, such as cyclin B, cyclin A and Cdc25B (Costa, 2005), all of which we found to be

reduced with Cdh1 overexpression. Therefore, Cdh1 overexpression could cause either

increased degradation of its substrates such as cyclin B, and/or a decrease in their

transcription by FoxM1 degradation.

In response to G2 phase DNA damage, Cdh1 degradation of Claspin is blocked. Claspin is

required to promote the activation of Chk1 so that the checkpoint arrest is maintained. Our

data revealed that Claspin levels are increased and required for the activation of pChk1 in

response to cyclin A/CDK2 inhibition. Interestingly, Claspin siRNA could not rescue the

effects of cyclin A siRNA but was able to block the increase in pChk1 normally seen with

cyclin A/CDK2 depletion/inhibition. One possible explanation for this is the rate of DNA

replication, which has been shown to be greatly reduced with Claspin siRNA treatment

(Petermann et al., 2008). Others have also shown a reduced rate of proliferation with Claspin

siRNA (Lin et al., 2004) and therefore this may mask the ability of Claspin to rescue the cyclin

A/CDK2 dependent G2 delay. Further investigation is required to confirm an S phase delay

caused by Claspin depletion.

Claspin stability is governed by several deubiquitination activities. The deubiquitinase, USP7

(Faustrup et al., 2009), which contains several cyclin A binding sites, regulates SCFβTrCP


246

mediated degradation of Claspin (Mailand et al., 2006, Peschiaroli et al., 2006), whereas

USP28 can regulate the Cdh1 mediation degradation of Claspin (Zhang et al., 2006a,

Bassermann et al., 2008). In addition, Claspin degradation by SCFβTrCP involves PBD binding of

Plk1 which leads to further phosphorylation by Plk1 that promotes further degradation of

Claspin (Peschiaroli et al., 2006). As both cyclin A/CDK2 and Plk1 regulate Claspin levels,

cyclin A/CDK2 may prime Claspin to allow Plk1 PBD binding and degradation. Therefore, loss

of cyclin A/CDK2 activity would ultimately lead to the stabilisation of Claspin. Another

possibility is that the increased stability is an indirect consequence of the cyclin A/CDK2

dependent delayed activation of cyclin B/CDK1. The possibility that cyclin A/CDK2 could

control the deubiquitination enzymes, USP7 and USP28, should also be investigated.

Although the role of Claspin and the G2 phase role of Cdh1 are mostly restricted to the DNA

damage response pathways, the evidence presented here demonstrates roles for both

proteins during a normal G2 phase. In the absence of cyclin A/CDK2 activity increased Claspin

levels sustain pChk1 levels required for the cyclin A/CDK2 dependent G2 phase delay. The

regulation of Claspin stability by cyclin A/CDK may involve its regulation by Plk1, however the

data presented for Cdh1 is more inconclusive. Therefore, further investigation is required to

determine if cyclin A/CDK2 can impact Claspin stability either by 1) affecting Plk1 PBD binding

to Claspin leading to its degradation, 2) cyclin A/CDK2 dependent reduction of Cdh1 levels or

3) cyclin A/CDK2 regulation of deubiquitinases such as USP7 or USP28. Overall, it is likely that

cyclin A/CDK2 is required in multiple pathways to control the stability of Claspin which

governs pChk1 levels and G2 phase progression.

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Chapter 8

Final Discussion

Chapter 8 – Final Discussion

248

The work presented in the preceding chapters characterised a number of pathways regulated

by cyclin A/CDK2 and their involvement in the co-ordination of G2/M progression. The basis

for this study was to demonstrate the role for cyclin A/CDK2 during G2 phase progression.

We and others had demonstrated that depletion of cyclin A or inhibition of CDK2 delayed

progression through G2 phase (Fung et al., 2007, Gong et al., 2007a, De Boer et al., 2008).

Although KO MEFs demonstrate redundancies of Cdks and cyclins the somatic function of

cyclin A/CDK2 is not definitively clear from this model and the work in this thesis has

established the importance of cyclin A/CDK2 activity in the regulation of the timing of

progression from G2 phase into mitosis. Loss of cyclin A/CDK2 activity delayed the activation

of cyclin B/CDK1 and as demonstrated in Chapter 3 resulted in loss of the coordination of

centrosomal and nuclear mitotic events associated with cyclin B/CDK1 activity. Additionally,

Chapter 4 demonstrates that cyclin A/CDK2 also controls the recovery from the G2 phase

checkpoint arrest and progression into mitosis.

Chk1, a major contributor to the G2 phase checkpoint, is now known to be involved in

normal G2 phase progression (Jiang et al., 2003, Schmitt et al., 2006, Matsuyama et al.,

2011). Based on its established roles, Chk1 contribution towards the cyclin A/CDK2

dependent G2 delay was investigated. Having had established that pChk1 was present not

only during normal G2 phase progression but also with cyclin A/CDK2 depletion/inhibition,

the ability of cyclin A/CDK2 to regulate Chk1 was investigated. Previously, studies had

demonstrated that CDK1 phosphorylated Chk1 on Ser286 and Ser301 to cause inhibition in

mitosis. Therefore the hypothesis that cyclin A/CDK2 could phosphorylate Chk1 during G2

phase was formulated. Cyclin A/CDK2 phosphorylated Chk1 in in vitro kinase assays however


249

the in vivo data did not support this. Others have published that CDK2 can phosphorylate

Chk1 on Ser286 and 301 in vitro (Ikegami et al., 2008). However, in combination with other

data from this study, it appeared more likely that cyclin B/CDK1 was the main kinase required

to phosphorylate Chk1 Ser286 and Ser301 in G2 phase and mitosis. Nevertheless, the timed

activation of cyclin B/CDK1, which is cyclin A/CDK2 dependent, may account for results that

show a small effect of cyclin A/CDK2 depletion/inhibition on these phosphorylation sites.

There is also the possibility that for a brief time in G2 phase cyclin A/CDK2 is required to

directly and initially phosphorylate Ser286 and Ser301. Further investigations are required to

determine the direct involvement of cyclin A/CDK2 but regardless of the kinase/s involved

there is strong evidence to suggest that these phosphorylations contribute to Chk1

inactivation which facilitates G2/M progression and G2 phase checkpoint arrest.

The effect of Chk1 Ser286 and Ser301 phosphorylation on G2/M progression, the G2

checkpoint and regulation of Chk1 localisation was investigated as part of this study.

Phosphorylation of Ser286 and Ser301 has been reported to lead to translocation of Chk1

from the nucleus to the cytoplasm (Enomoto et al., 2009). Under normal circumstances, Chk1

is reported to be predominantly located within the nucleus (Matsuyama et al., 2011). In

response to DNA damage an increase in activating phosphorylations (Ser317 and 345)

coincides with an influx of Chk1 into the nucleus. Dephosphorylation of Ser317 and Ser345

can be catalysed by PP2A (Leung-Pineda et al., 2006, Li et al., 2007), and is thought to

facilitate translocation of Chk1 into the cytoplasm and promote Chk1 inhibition that is

required for checkpoint recovery as well as unperturbed mitotic progression. The exact

timing of these events remains unclear and the length of time Chk1 must be phosphorylated


250

on Ser286 and Ser301 to ensure dephosphorylation of Ser317 and Ser345 by PP2A, required

for mitosis, is unknown. As shown in earlier results, minimal Ser317 phosphorylation was

detected in mitosis, and published data has shown that activating phosphorylations, Ser317

and Ser345, and inhibitory phosphorylations, Ser286 and Ser301 can occur on the same Chk1

molecule during mitosis (Ikegami et al., 2008). Therefore, it is likely that both

phosphorylation events, activating and inhibitory, occur prior to mitotic entry. The theory

that phosphorylation of Ser286 and Ser301 can overcome Chk1 activation leading to Chk1

inhibition and mitotic entry, means that although the phosphorylated state of Ser317 and

Ser345 may be a surrogate of activity it may not be a marker of Chk1 activation during G2/M

phase progression. Consequently the dephosphorylation of Ser317 and Ser345 is likely to

occur at some time during mitosis. Additional work is essential to uncover the

phosphorylation status of Chk1 during specific stages of the cell cycle as the regulation of

Chk1 activity continues to be shown as a complex process (Wang et al. 2012). In fact the

contribution of Ser286 and Ser301 phosphorylation of Chk1 was shown recently to be

present and required to implement a checkpoint in response to DNA damage (Xu et al.,

2012). Although much of the work conducted in this study was used to determine the role

and regulation of Chk1 during G2/M by using overexpression vectors, it is evident that they

were not ideal for use in cell cycle investigational studies. In addition, the synchronisation

methods commonly used for cell cycle studies, such as this, never provide a solid

representation of all cells within the exact same phase of the cell cycle and although it is the

most ideal model available it cannot deliver the perfect results required. The model in Figure


251

Figure 8.1 Model of Chk1 regulation during G2/M progression.

In G2 phase ATM/ATR phosphorylates Chk1 on Ser317 and Ser345. During late G2 phase, Chk1

phosphorylation on Ser286 and Ser301 may involve either cyclin B/CDK1 phosphorylation, governed

by the cyclin A/CDK2 regulation of cyclin B/CDK1, or possibly via direct phosphorylation by cyclin

A/CDK2 itself. Once phosphorylated in late G2 phase the removal of phosphorylations Ser317 and

Ser345 is likely catalysed by PP2A in the nucleus and leads to Chk1 inactivation and translocation into

the cytoplasm. Continued phosphorylation of Ser286 and Ser301 during mitosis (after cyclin A/CDK2

degradation) is solely dependent on cyclin B/CDK1 activity. Phosphorylation of Ser286 and Ser301 by

cyclin A/CDK2 is unknown and therefore is indicated by a dashed arrow.


252

8.1 summarises the results obtained from this study and the hypothesised mode of Chk1

regulation in G2/M progression.

Claspin is a known adapter protein required for Chk1 phosphorylation. The ability of cyclin

A/CDK2 to regulate Claspin levels was a novel finding of this study. Additional work by an

Honours student that I supervised confirmed that both cyclin A depletion and CDK2

inhibition, stabilised Claspin levels (data not shown). Claspin influences the activation of Chk1

in response to DNA damage and stabilisation of Claspin has been shown to be responsible for

pChk1s involvement in response to DNA damage (Kumagai and Dunphy, 2000, Liu et al.,

2006, Wang et al., 2006, Bassermann et al., 2008, Lindsey-Boltz et al., 2009). During the cell

cycle, endogenous Claspin levels follow a similar pattern to pChk1 levels, and are highest in S

and G2 phases and drop off once cells begin to enter M phase (Mamely et al., 2006). The role

of Claspin in S phase will most likely be the reason for delayed mitotic entry caused by

Claspin siRNA (Figure 7.8 C) and hence why Claspin siRNA could not overcome cyclin A

depletion and/or CDK2 inhibition to the same extent as Chk1 siRNA treatment.

Recently, Claspin stability was shown to be involved in the release of cells from the G2

checkpoint arrest. During a G2 checkpoint arrest, Claspin is active and mediates the

activation of Chk1 whilst during recovery Claspin must be degraded by SCFβTrCP or APCCdh1

(Mailand et al., 2006, Mamely et al., 2006, Peschiaroli et al., 2006, Bassermann et al., 2008).

Addition of caffeine, and to a lesser extent Chk1 inhibitor, was unable to force G2 checkpoint

exit in cyclin A depleted cells and suggests the role of Claspin during the G2 checkpoint is not

the only pathway regulated by cyclin A/CDK2.The degradation of Claspin by the SCFβTrCP


253

ubiquitin ligase during checkpoint recovery has been shown to be Plk1 dependent (Mailand

et al., 2006, Mamely et al., 2006, Peschiaroli et al., 2006, Bassermann et al., 2008). Plk1

regulates Claspin degradation (Mailand et al., 2006, Mamely et al., 2006, Peschiaroli et al.,

2006) by generating phosphodegrons required for βTrCP binding. Plk1 targets Claspin as well

as other targets such as Wee1 for degradation during the G2/M transition (Ang and Harper,

2004, Hansen et al., 2004, Moshe et al., 2004) and therefore also facilitates checkpoint

recovery. Plk1 inhibition necessary for the G2 checkpoint is ATM/ATR dependent (van Vugt et

al., 2001, Tsvetkov and Stern, 2005) and recovery from the checkpoint requires activation of

Plk1 by Aurora A. FoxM1 is responsible for the transcription of Plk1 and other G2 phase

proteins, such as cyclin B and Cdc25B that are required for mitotic entry. FoxM1 is positively

regulated by cyclin A/CDK2 phosphorylation and PP2A antagonises cyclin A/CDK2 activation

by dephosphorylation of FoxM1 (Laoukili et al., 2008b, Alvarez-Fernandez et al., 2011). The

possibility that cyclin A/CDK2 regulates FoxM1 transcription of Plk1 during G2 phase thereby

controlling the normal increase in Plk1 levels required for mitotic entry has been

investigated. The G2 phase protein levels of Plk1, cyclin B and Cdc25B appeared unchanged

in cyclin A depleted G2 phase samples. It is impossible to comment that all G2/M genes

dependent on FoxM1 activity are affected by a loss of cyclin A/CDK2, as conclusions can only

be made for those proteins investigated but it is suggested that the effects of cyclin A/CDK2

on FoxM1 activity require further attention. Cyclin A/CDK2 regulation of FoxM1 occurs in G2

phase and based on published work is likely to be required for checkpoint recovery (Laoukili

et al., 2008b, Alvarez-Fernandez et al., 2010). Although blocking cyclin A/CDK2 activation was

shown to be required for checkpoint arrest (Goldstone et al., 2001), a small portion of cyclin


254

Figure 8.2 Claspin involvement and regulation during G2/M.

During G2 or the G2 checkpoint, Claspin is required for the ATR dependent activation of Chk1 (top

half of figure). During mitotic progression/checkpoint recovery (bottom half of figure) Plk1

phosphorylates Claspin and promotes degradation of Claspin by SCFβTrCP or APCCdh1 (dependent on the

type of stress that induces the G2 checkpoint). As indicated , cyclin A/CDK2 has the potential to

regulate Claspin levels during mitotic progression/checkpoint recovery. The mode of this regulation is

not yet clearly defined but may involve Plk1 regulation via FoxM1 or by other yet unknown

pathway/s. The grey colour specifies pathways, events, regulation of proteins that are blocked or not

functioning at that time point.


255

A/CDK2 remains active specifically to activate FoxM1, to facilitate checkpoint recovery.

Whether the expression of FoxM1 dependent genes is affected when a G2 checkpoint is

activated in cells depleted of cyclin A/CDK2 was not determined, therefore further

information regarding FoxM1 activation is required to determine the pathways involved in

G2 phase checkpoint recovery. In addition there is evidence that Plk1 has the ability to

regulate FoxM1 activity and is dependent of CDK1 priming of FoxM1 (Fu et al., 2008). The

investigation of the mechanisms of FoxM1 regulation may be more difficult to determine due

to the normal changes in Plk1 levels seen upon mitotic entry and the evident delay of mitotic

entry caused by cyclin A/CDK2 depletion/inhibition. The overall contribution of the pathway

towards Claspin stability and the level of Claspin required to control checkpoint

recovery/mitotic progression is unknown, however Figure 8.2 below encompasses the

hypothesised pathways involved.

A possible mechanism for cyclin A/CDK2 regulation of Claspin stability is via regulation of

deubiquitinases that contribute to Claspin stability. During the G2 phase checkpoint

Bassermann et al. (2008) found that Claspin stability is protected from APCCdh1 degradation

by the action of USP28 (Ubiquitin-specific-processing protease 28). In addition to USP28,

USP7 is capable of binding and regulating Claspin stability also and has been found to protect

Claspin degradation by SCFβTrCP in response to stress (Faustrup et al., 2009). Although there is

currently no data that shows cyclin/CDK regulation of USP7 or USP28, USP28 can be

regulated by phosphorylation. USP28 is phosphorylated in an ATM-dependent manner and

can stabilise Chk2 in the absence or presence of damage such as IR (Zhang et al., 2006a). This

suggests that USP28 has a basal level of activity during an unperturbed cell cycle. Recent


256

data from the Gabrielli lab has demonstrated that USP7 depletion rescues a cyclin A/CDK2

dependent reduction of Claspin levels and Chk1 activation (data not shown). This would

suggest that USP7 is involved in the regulation of Claspin and thereby pChk1 levels. To date

there are no reports of direct cyclin A/CDK2 regulation of the USP family and the Gabrielli

Laboratory have also been unable to show direct cyclin A/CDK2 interaction with USP7,

however further investigation is needed to reveal the mode of regulation.

Once Plk1 is active, how does the affinity of Plk1 for Claspin increase so as to promote

Claspin degradation? Interestingly, ATR phosphorylates Claspin in Xenopus, which facilitates

Plk1 docking to Claspin and thereby causing Chk1 inactivation and termination of the

checkpoint (Yoo et al., 2004). This has not yet been shown in a mammalian system, and the

kinase responsible remains unknown. The possibility that cyclin A/CDK2 was responsible for

priming Claspin for Plk1 binding was investigated due to the similarity between the Plk1

binding site and CDK2 phosphorylation site, but no evidence for a role of cyclin A/CDK2 in

promoting Plk1 binding to Claspin could be determined. However, the fact that cyclin A/CDK2

appeared to regulate Claspin levels suggested cyclin A/CDK2 could directly influence Claspin

levels by priming it for Plk1 dependent degradation. The localisation of Plk1 was used as an

indicator of cyclin A/CDK2 ability to prime Plk1 substrates. Although immunofluorescence

data suggested that cyclin A/CDK2 can control Plk1 localisation, it did not specifically

demonstrate Plk1 binding to particular substrates, such as Claspin. Additional

immunoprecipitation methods carried out by an Honours student at the time, Brittney

Harrington, confirmed that Plk1 and Claspin binding was not affected by cyclin A/CDK2 in G2

phase instead it appeared to be disrupted in mitosis only (data not shown). We have


257

previously shown that cyclin A/CDK2 has a role in mitosis and this was to control the

phosphorylation of APC (adenomatous polyposis coli) and proper spindle assembly (Beamish

et al., 2009). Preliminary evidence in the laboratory has shown Plk1 can bind APC, but

whether the interaction is cyclin A/CDK2 dependent is currently inconclusive. Why the

interaction between Claspin and Plk1 is more greatly impacted, by a loss of cyclin A/CDK2, in

mitosis rather that G2 phase is currently unknown. Proteomics analysis of Plk1 substrates

affected by cyclin A/CDK2 depletion/inhibition has been conducted but to date has not

revealed any Plk1 binding proteins that are specifically dependent on cyclin A/CDK2

phosphorylation.

Work from the Gabrielli group and others have shown that Cdc25B and Plk1 have critical

roles during the G2 checkpoint. Inhibition of either Cdc25B or Plk1 blocks cells exiting the G2

phase checkpoint and also delays normal mitotic progression (van Vugt et al., 2004a).

Cdc25B-dependent G2 phase activation of cyclin A/CDK2 is blocked during the checkpoint

(Goldstone et al 2001). Reports have shown that a small portion of cyclin A/CDK2 remains

active and is required for the exit of cells from the checkpoint (Goldstone et al., 2001,

Alvarez-Fernandez et al., 2010, Duursma and Cimprich, 2010), therefore this could be the

reason why Cdc25B inhibition blocks checkpoint exit. The exact role of Plk1 in normal G2/M

progression is less clear than for Cdc25B. The question that arises is whether Plk1 is involved

in the same pathway/s as cyclin A/CDK2 during the G2 checkpoint and if its prominent role is

to implement the arrest or to block recovery or both? The role of Plk1 in checkpoint

recovery is to initiate the degradation of Wee1 and Claspin. There is evidence that CDK1 may

prime Wee1 for Plk1 binding to mediate its degradation. To date, it appears that Claspin


258

stability is dependent on cyclin A/CDK2 activity, however direct priming of Claspin by cyclin

A/CDK2 to allow Plk1 binding is unknown. The fact that Plk1 overexpression cannot

overcome the effects of cyclin A depletion (Figure 7.1 and 7.2) and that the overexpression of

a PBD construct blocks cells in G2 phase, may reflect cyclin A/CDK2s ability to prime Plk1

substrates such as Wee1 and Claspin. However, others have reported that constitutively

active Plk1 can bypass the G2 checkpoint (Smits et al., 2000) and although this refutes the

suggestion that cyclin A/CDK2 is required to prime Plk1 substrates, perhaps the portion of

cyclin A/CDK2 that remains active during the checkpoint is sufficient to prime Plk1 substrates.

There is also the possibility that cyclin A/CDK2 is initially required to implement the arrest by

inhibiting Plk1 prior to regulating Plk1 localisation, explaining the observation that

constitutively active Plk1 can overcome the checkpoint. As mentioned previously, a reduction

in the activating phosphorylation of Plk1 (pT210) in G2 phase could not be accurately

assessed with the reagents available and little is known about the mechanisms that govern

Plk1 activity, except that its activation requires Aurora A activation (Macurek et al., 2008,

Seki et al., 2008b). Work done in the Gabrielli laboratory has ruled out cyclin A/CDK2s ability

to affect Aurora A activity under these circumstances, based on a lack of monospindle asters

in cyclin A depleted cells which are typical of Plk1 inhibition (Sunkel and Glover, 1988,

Llamazares et al., 1991, Lane and Nigg, 1996, Qian et al., 1998, Sumara et al., 2004, McInnes

et al., 2006, Lenart et al., 2007,) and Aurora A inhibition ( Glover et al., 1995, Roghi et al.,

1998, Berdnik and Knoblich, 2002, Hoar et al., 2007) (data not shown). However, the delay of

cyclin B/CDK1 activation at the centrosomes caused by cyclin A depletion/ CDK2 inhibition

could be a consequence of an indirect disruption to the regulation of Cdc25B by Aurora A,


259

although this was not investigated as part of this study. The reduction of Plk1 activating

phosphorylation levels in mitosis caused by cyclin A depletion suggested that cyclin A/CDK2 is

able to regulate Plk1 protein levels. Plk1 proteolysis is required for proper mitotic exit and is

initiated by APCCdc20 and is completed by APCCdh1 in late mitosis. Although this study has

provided evidence for cyclin A/CDK2 regulation of Cdh1, the data does not entirely support

the regulation of Plk1 being temporally coordinated by this mechanism of Cdh1 regulation;

further work is clearly required to determine the exact effects that cyclin A/CDK2 may have

on Cdh1 activity and therefore its ability to regulate Plk1 levels.

Wee1 catalyses the inhibitory Tyr15 phosphorylation of CDK1/2 and was recently found to be

negatively regulated by cyclin A/CDK. In particular, cyclin A/CDK phosphorylates and inhibits

Wee1, and as a consequence affects G2/M progression (For an overview see Figure 8.3). An

Alanine mutant of this phosphorylation site on Wee1 blocks CDK inhibition and therefore

inhibits G2/M progression (Li et al., 2010). Wee1 is degraded upon mitotic entry via a βTrCP-

dependent mechanism and requires CDK1 or Plk1 phosphorylation for recognition by βTrCP

(van Vugt et al., 2004a, Smith et al., 2007). The possibility that cyclin A/CDK2 phosphorylation

of Wee1 leads to Wee1 degradation was not shown by Li et al 2010. however this may have

been obscured by the overexpression setting used. The regulation of Wee1 could be an

important contributor to the cyclin A/CDK2 dependent G2 delay, where phosphorylation of

Wee1 by cyclin A/CDK2 could lead to Plk1 binding and Plk1 dependent SCFβTrCP degradation.

However, recent work from the Gabrielli lab has failed to detect any cyclin A/CDK2-

dependent Polobox domain binding within Wee1, although it may be that other regions of

Plk1 in addition to the Polobox are required for the cyclin A/CDK2-dependent binding.


260

A recurrent theme of this study was that the data provided insight into the pathways

involved in the cyclin A/CDK2 dependent G2 delay, but the data did not identify a dominant

pathway that accounted for cyclin A/CDK2 dependent G2 phase progression. This is thought

to be a result of the timing at which these events are being analysed. As a tightly regulated

and timed process the cell cycle is difficult to study and although the synchronisation

methods generate certain information about the pathways involved, it is not an exact

representation of the events occurring at a particular time point in the cell cycle. The cell line

predominantly used in this study was the HeLa cell line. These cells are used because of their

ideal cell cycle profile but unfortunately they have diminished levels of the important cell

cycle genes p53 and Rb (Hamada et al., 1996). Unlike Rb which functions during G1/S and

G1/S checkpoint, p53 does have roles during G2 phase however its roles appear to be

independent of the cyclin A/CDK2 delay as NFF cells, with functional p53, do delay in G2

phase. The cyclin A/CDK2 dependent G2 delay in NFF cells also caused an accumulation of

pChk1 (data not shown) and suggests that p53 is not involved in this cyclin A/CDK2 G2/M

regulated pathway. However, p53 has an established role during the G2 phase checkpoint

which can be impacted by a loss of cyclin A/CDK2 activity and therefore this may impact the

dependency that HeLa cells have on cyclin A/CDK2 during a G2 checkpoint arrest. In response

to IR, human cells without CDK2 rely on p53 dependent activation of the G2 checkpoint and

inversely cells without p53 rely on CDK2 dependent G2 checkpoint activation (Chung and

Bunz, 2010). The p53 dependent signalling occurs independently of ATR/ATM signalling

through p21 dependent inhibition of cyclin B/CDK1 to block mitotic entry. It is unknown if

HeLa cells have compensatory pathways used to control the G2 checkpoint in the absence of


261

CDK2 as they lack functional p53. If not, this could account for the noticeable portion of cell

death in G2 phase arrested HeLa cells treated with cyclin A/CDK2 siRNA/inhibitor and forced

to exit the checkpoint with the addition of caffeine or Chk1 inhibitor (data not shown).

All of the proteins mentioned have established roles in both an unperturbed cell cycle as well

as the G2 checkpoint. The ATR-Claspin-Chk1 pathway is critical during the G2 checkpoint and

this study has also strongly confirmed its involvement during an unperturbed cell cycle. The

role of Chk1 during an unperturbed cell cycle appears to resemble its function during the G2

checkpoint. However, there are some checkpoint pathways that act differently during normal

cell cycle progression. For example, depletion of Cdc25B and Plk1 does not affect

unperturbed mitotic entry to the same extent as mitotic entry following checkpoint recovery

suggesting a difference in pathways utilised (van Vugt et al., 2004a). Therefore it cannot be

assumed that a protein’s checkpoint function and unperturbed cell cycle function are the

same in both situations. Chk1 involvement in the cyclin A/CDK2 dependent G2 delay was

linked to the mediating protein Claspin. The regulation of Claspin in this pathway may involve

Plk1 dependent degradation by SCF. However, further investigation is needed, as cyclin

A/CDK2 appeared to only regulate Plk1 in M phase. Mitotic effects on Wee1 levels were also

noticed in cyclin A/CDK2 depleted/inhibited samples and may also involve Plk1 dependent

SCF degradation. The mitotic pathways affected by cyclin A/CDK2 were not examined as part

of this study but may be linked to the defective mitoses commonly observed in cyclin A/CDK2

depleted/inhibited samples (Beamish et al., 2009). Experimentation to determine Cdh1

involvement in the cyclin A/CDK2 dependent G2 delay was conclusive but showed only a

minor role for Cdh1. The preliminary evidence (data not shown) suggests that Cdh1 is


262

involved in the regulation of Chk1 levels required for the delay via Claspin regulation. It may

be that Cdh1 regulation of Claspin stability contributes to increased pChk1 levels seen with

cyclin A/CDK2 depletion/inhibition. Although a reduction in Cdh1 levels was observed in

cyclin A/CDK2 depleted/inhibited samples, further informative data regarding the activity

status of Cdh1 is necessary to provide further insight into its involvement. Overall, there was

no sole dominant pathway identified that was involved in the cyclin A/CDK2 dependent G2

delay. It is apparent from this study that the master regulator of G2, cyclin A/CDK2, regulates

many pathways during G2 which consequently are affected when a loss of cyclin A/CDK2

activity occurs. The model in Figure 8.3 demonstrates some of the many pathways regulated

by cyclin A/CDK2 in G2/M. Much of this model is suggestive (Figure 8.3; dashed lines) and

additional work is required to confirm the exact role of these pathways in both the

checkpoint and unperturbed G2/M progression.


263

Figure 8.3 A simplified schematic of G2 regulation involving cyclin A/CDK2.

Dashed lines indicate degradation pathways influenced by Cdh1 and Plk1 or FoxM1 transcription.

Some pathways may have more prominent roles during either an unperturbed G2/M progression or

G2 checkpoint but have not been excluded for their involvement in either of these processes. The

boxes indicate the G2/M proteins that are regulated by Cdh1 and FoxM1 that are most likely

regulated by cyclin A/CDK2.

264

Chapter 9

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Chapter 10

Appendix

Chapter 10 – Appendices

319

10.1 Appendix 1

Chapter 10 – Appendices

320

Figure 10.1 Cellomics data confirms the presence of pChk1 in an unperturbed cell cycle.

Quantification of cell lines was carried out by defining the cell cycle phases based on DNA intensity

histograms, like that seen in Figure 3.4 B. The graph represents the mean plus three standard

deviations, the range represents the 95 percentile. The results of pChk1 (Ser317) were corrected for

background intensities. Etoposide treatment was used as a control and accordingly results in elevated

pChk1 levels.

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