Investigation of Stathmin's Role in Neuroblastoma Drug Resistance

Investigation of stathmin’s role in

neuroblastoma drug resistance,

differentiation and metastasis

Frances Louise Byrne

Thesis submitted in fulfilment of the requirements for the degree

of Doctor of Philosophy

School of Women’s & Children’s Health

Faculty of Medicine

University of New South Wales, Sydney, Australia

2012

Table of Contents

List of Figures………………………………………………………….…vi List of Tables……………………………………………………………...ix Abbreviations………………………………………………………….…..x Publications arising from this thesis…………………………………....xv Acknowledgements……………………………………………...………xvi Abstract………………………………………………………………...xviii Chapter 1. Introduction ...... 1 1.1 Neuroblastoma ...... 3 1.1.1 Prognostic Indicators ...... 3 1.1.1.1 Age & Disease Stage ...... 3 1.1.1.2 Genetic Abnormalities ...... 4 1.1.1.3 Histology ...... 5 1.1.1.4 Gene expression ...... 6 1.1.2 Risk Classification ...... 7 1.1.3 Treatment ...... 7 1.1.4 Drug Resistance ...... 9 1.1.4.1 The MYCN Oncogene ...... 9 1.1.5 Metastatic Disease ...... 10 1.2 The Cell Cytoskeleton ...... 14 1.2.1 Microtubule Network ...... 15 1.2.1.1 Structure & Function ...... 15 1.2.2 Microtubule Dynamics & GTP hydrolysis ...... 20 1.2.3 Microtubule Assembly & Stability ...... 20 1.2.4 Tubulin Isotypes ...... 21 1.2.5 Microtubule Nucleation & Post-translational modifications (PTMs)...... 22 1.2.6 Microtubule-interacting Proteins ...... 23 1.2.6.1 Motor Proteins ...... 23 1.2.6.2 Microtubule Stabilisers ...... 24 1.2.6.3 Microtubule Destabilisers ...... 25

1.2.7 Tubulin-Binding Agents ...... 25 1.2.8 Resistance to TBAs ...... 26 1.2.9 Actin Network...... 30 1.2.9.1 Structure & Function ...... 30 1.2.10 Actin in Cell Motility ...... 31 1.2.10.1 Assembly of Actin Structures ...... 31 1.2.10.2 Disassembly of Actin Structures ...... 32 1.2.10.3 Linking Actin to Microtubules ...... 32 1.3 Stathmin ...... 33 1.3.1 Tissue Expression ...... 34 1.3.2 Structure & Function ...... 34 1.3.3 Phosphorylation ...... 40 1.3.4 Cell Cycle Regulation ...... 43 1.3.5 Stathmin-like Protein Family ...... 44 1.3.6 Cell Differentiation ...... 48 1.3.7 Neural Cell Functions ...... 49 1.3.8 Cell Migration ...... 50 1.3.9 Interphase Microtubule Network ...... 50 1.3.10 RhoGTPase Signalling Pathways ...... 51 1.3.11 Interacting Proteins ...... 52 1.4 Stathmin in Cancer ...... 53 1.4.1 Transcription Factor Deregulation ...... 54 1.4.2 Stathmin & p53 ...... 56 1.4.3 Post-transcriptional Deregulation ...... 57 1.4.4 Oncogenic Signalling Pathways ...... 58 1.4.5 Loss of Heterozygosity ...... 59 1.4.6 Altered Activity ...... 60 1.4.7 Tumour Progression ...... 61 1.4.8 Drug Resistance ...... 62 1.4.9 Metastasis...... 66 1.4.10 Stathmin in Neuroblastoma ...... 67 1.5 Thesis Aims ...... 68 Chapter 2. Materials & Methods ...... 70

2.1 Cell culture ...... 71 2.2 siRNA-mediated stathmin suppression ...... 73 2.3 shRNA-mediated stathmin suppression ...... 75 2.3.1 Upscale production of plasmid DNA...... 77 2.3.2 Plasmid transfections ...... 77 2.4 cDNA preparation & qPCR ...... 78 2.5 Protein expression analysis ...... 79 2.5.1.1 Examination of protein phosphorylation ...... 83 2.6 Cell proliferation assay ...... 83 2.7 Cell enumeration & viability ...... 84 2.8 Tubulin polymerisation assay ...... 84 2.9 Cytotoxic drugs & inhibitors ...... 85 2.10 Cell cycle analysis ...... 85 2.11 Cytotoxicity assays ...... 86 2.12 Drug-treated clonogenic assays ...... 86 2.13 Retinoic acid-induced differentiation ...... 87 2.14 Microscopy ...... 88 2.14.1 Phase contrast ...... 88 2.14.2 Immunofluorescence ...... 88 2.15 Neurite counts ...... 89 2.16 Soft agar assays ...... 89 2.17 Monolayer wound-healing assays ...... 90 2.18 Chemotaxis migration & invasion assays ...... 91 2.19 Luciferase-expressing neuroblastoma cells...... 92 2.20 In vitro luciferase assays ...... 94 2.21 Orthotopic injection of neuroblastoma cells ...... 95 2.22 In vivo & ex vivo imaging & tumour measurements ...... 96 2.23 Immunohistochemistry ...... 98 2.24 Statistical analyses ...... 99 Chapter 3. Results I ...... 101 3.1 Stathmin protein expression in neuroblastoma cell lines ...... 103 3.2 Stathmin gene silencing in neuroblastoma cells ...... 106

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3.3 Stathmin does not influence the expression of cytoskeletal-related proteins in neuroblastoma cells ...... 109 3.4 Stathmin does not influence neuroblastoma cell proliferation or viability ...... 117 3.5 Stathmin does not markedly influence neuroblastoma drug sensitivity...... 124 3.6 Discussion ...... 140 Chapter 4. Results II ...... 144 4.1 ATRA induces neurite formation in neuroblastoma cells ...... 147 4.2 ATRA reduces stathmin protein expression in neuroblastoma cells ...... 150 4.3 ATRA induces stathmin phosphorylation in neuroblastoma cells ...... 155 4.4 Stathmin regulates ATRA-induced neurite formation ...... 160 4.5 Stathmin does not influence ATRA-induced growth arrest ...... 166 4.6 Stathmin does not influence ATRA-induced alterations in differentiation marker expression ...... 169 4.7 Discussion ...... 184 Chapter 5. Results III ...... 187 5.1 Stathmin does not influence neuroblastoma anchorage-independent growth .... 189 5.2 Stathmin does not influence neuroblastoma 2D cell migration ...... 189 5.3 Stathmin mediates chemotactic-induced neuroblastoma cell migration & invasion ...... 192 5.4 Stathmin regulates neuroblastoma cell morphology ...... 198 5.5 Stathmin regulates neuroblastoma tubulin polymer levels ...... 201 5.6 Stathmin regulates cofilin phosphorylation ...... 204 5.7 Stathmin does not influence LIMK expression ...... 207 5.8 Stathmin regulates cofilin and MLC phosphorylation via ROCK ...... 207 5.9 Stathmin regulates neuroblastoma cell invasion via ROCK ...... 213 5.10 Discussion ...... 216 Chapter 6. Results IV ...... 222 6.1 Stathmin shRNA\luciferase-expressing neuroblastoma cells ...... 224 6.2 Stathmin does not influence neuroblastoma tumour engraftment or growth ..... 227 6.3 Metastatic spread in the orthotopic neuroblastoma mouse model ...... 239 6.4 Neuroblastoma lung metastasis ...... 251 6.5 Stathmin suppression reduces neuroblastoma lung metastasis ...... 256 6.6 Discussion ...... 264

Chapter 7. Conclusions & Future Directions ...... 267 Appendix…………………………………………………………...………….....275 References………………………………………………………………………..286

List of Figures

Figure 1.1The formation of a metastatic tumour...... 11

Figure 1.2 Microtubules and actin filaments...... 16

Figure 1.3 Microtubule ‘dynamic instability’ is governed by GTP hydrolysis...... 18

Figure 1.4 Tubulin-Binding Agents...... 27

Figure 1.5 The T2S complex...... 36

Figure 1.6 Stathmin mediates microtubule dynamics by sequestering tubulin and/or increasing microtubule catastrophe...... 38

Figure 1.7 Signalling pathways that regulate stathmin phosphorylation...... 41

Figure 1.8 The stathmin-like protein family...... 46

Figure 2.1 The pRS vector map...... 76

Figure 2.2 Schematic of the SFG-NESTGL triple-modality reporter construct...... 93

Figure 3.1 Stathmin protein expression in neuroblastoma cell lines...... 104

Figure 3.2 SiRNA optimisation in neuroblastoma cells...... 107

Figure 3.3 Stathmin gene expression in siRNA-transfected neuroblastoma cells...... 110

Figure 3.4 Double siRNA transfection of neuroblastoma cells...... 112

Figure 3.5 Cytoskeletal protein expression in stathmin-suppressed BE(2)-C cells...... 115

Figure 3.6 Proliferation and viability of stathmin-suppressed BE(2)-C cells...... 118

Figure 3.7 Proliferation of stathmin-suppressed SH-SY5Y cells...... 120

Figure 3.8 Cell cycle profiles of paclitaxel-treated, stathmin-suppressed BE(2)-C cells…...... 122

Figure 3.9 Short-term drug sensitivity of stathmin-suppressed BE(2)-C cells...... 126

Figure 3.10 Short-term drug sensitivity of stathmin-suppressed SH-SY5Y cells...... 128

Figure 3.11 Long-term drug sensitivity of stathmin-suppressed BE(2)-C cells...... 130

Figure 3.12 Long-term drug sensitivity of stathmin-suppressed BE(2)-C cells...... 132

Figure 3.13 Long-term drug sensitivity of stathmin-suppressed SH-SY5Y cells...... 135

Figure 3.14 Long-term drug sensitivity of stathmin-suppressed SH-SY5Y cells...... 137 vi

Figure 4.1 ATRA induces neurite formation in neuroblastoma cells...... 148

Figure 4.2 Short-term exposure to ATRA does not markedly alter stathmin protein expression in neuroblastoma cells...... 151

Figure 4.3 Long-term (7 day) exposure to ATRA reduces stathmin protein expression in neuroblastoma cells...... 153

Figure 4.4 ATRA alters the phosphorylation of stathmin’s Ser16 and Ser25 residues in neuroblastoma cells...... 156

Figure 4.5 ATRA alters the phosphorylation of stathmin’s Ser38 and Ser63 residues in neuroblastoma cells...... 158

Figure 4.6 Stathmin suppression reduces ATRA-induced neurite formation in BE(2)-C cells...... 161

Figure 4.7 Stathmin suppression reduces ATRA-induced neurite formation in SH-SY5Y cells...... 164

Figure 4.8 Stathmin suppression does not prevent ATRA-induced growth arrest in neuroblastoma cells...... 167

Figure 4.9 Short term ATRA treatment does not alter the expression of neuronal differentiation markers...... 170

Figure 4.10 Development of stathmin shRNA-expressing SH-SY5Y cells...... 173

Figure 4.11 ShRNA-mediated stathmin suppression reduces ATRA-induced neurite formation in SH-SY5Y cells...... 175

Figure 4.12 Long-term ATRA treatment reduces stathmin expression in SH-SY5Y cells...... 178

Figure 4.13 SCG10 and MAP2c expression in stathmin shRNA-suppressed SH-SY5Y cells following long-term treatment with ATRA...... 180

Figure 4.14 βIII-tubulin and NSE expression in stathmin shRNA-suppressed SH-SY5Y cells following long-term treatment with ATRA...... 182

Figure 5.1 Stathmin does not regulate neuroblastoma anchorage-independent growth.190

Figure 5.2 Stathmin does not regulate 2D neuroblastoma cell migration...... 193

Figure 5.3 Stathmin regulates chemotactic-induced neuroblastoma cell migration and invasion...... 196

Figure 5.4 Stathmin regulates neuroblastoma cell morphology...... 199

Figure 5.5 Stathmin regulates tubulin polymer levels in neuroblastoma cells...... 202

Figure 5.6 Stathmin regulates cofilin phosphorylation in neuroblastoma cells...... 205

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Figure 5.7 Stathmin does not regulate LIM kinase expression or phosphorylation in neuroblastoma cells...... 208

Figure 5.8 Stathmin suppression-induced increases in cofilin and MLC phosphorylation are reversed by treatment with the ROCK inhibitor, Y-27632...... 211

Figure 5.9 Stathmin suppression-induced reduction of cell invasion is reversed by the ROCK inhibitor, Y-27632...... 214

Figure 5.10 Schematic diagram of stathmin’s proposed regulation of neuroblastoma cell migration and invasion...... 217

Figure 6.1 Development of shRNA/luciferase-expressing neuroblastoma cells...... 225

Figure 6.2 ShRNA-mediated stathmin suppression reduces neuroblastoma cell invasion…………………………………………………………………..228

Figure 6.3 ShRNA-expressing SK-N-BE(2)/TGL cells express similar levels of luciferase...... 230

Figure 6.4 Stathmin does not influence neuroblastoma tumour burden...... 233

Figure 6.5 Stathmin does not influence neuroblastoma tumour growth...... 235

Figure 6.6 Stathmin suppression was maintained in the primary neuroblastoma tumours...... 237

Figure 6.7 Luciferase and stathmin expression in neuroblastoma primary tumours. ... 240

Figure 6.8 Bioluminescent imaging and histological analysis of the left kidney...... 242

Figure 6.9 Bioluminescent imaging and histological analysis of the right kidney...... 245

Figure 6.10 Bioluminescent imaging and histological analysis of the liver...... 247

Figure 6.11 Histological analysis of the spleen...... 249

Figure 6.12 BLI detection of potential bone and/or lymph node metastases...... 252

Figure 6.13 Bioluminescent imaging and histological analysis of a CtrlSH lung...... 254

Figure 6.14 Bioluminescent imaging and histological analysis of a Stmn Seq.3SH lung…...... 257

Figure 6.15 Neuroblastoma tumour burden in CtrlSH, Stmn Seq.2SH and Stmn Seq.3SH mouse lungs...... 259

Figure 6.16 Stathmin suppression reduces neuroblastoma tumour burden in the lungs of mice...... 262

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Appendix Figure 1. Stathmin suppression reduces cell proliferation in Calu-6 lung cancer cells……………………………………………………………………………276 Appendix Figure 2. Stathmin-like protein expression in neuroblastoma and lung cancer cells..…………………………………………………………………………………..278 Appendix Figure 3. Migration assay optimisation……………………………………280 Appendix Figure 4. EMT marker protein expression in stathmin-suppressed neuroblastoma cells…………………………………………………………………...282 Appendix Figure 5. Detection of adrenal fat pad injection spills by bioluminescent imaging………………………………………………………………………………..284

List of Tables

Table 1.1 Altering stathmin expression modifies cell sensitivity to chemotherapy & irradiation ...... 64

Table 2.1 Neuroblastoma Cell Types ...... 71

Table 2.2 Stathmin siRNA and shRNA Sequences ...... 74

Table 2.3 Antibody dilutions for western blots ...... 81

Abbreviations

ADF actin-depolymerising factor

ARF Alternative reading frame

ATCC American Type Culture Collection

ATP adenosine triphosphate

ATRA all-trans retinoic acid

BCL-2 B-cell lymphoma-2

BIM Bcl-2 interacting mediator of cell death

BLI bioluminescent imaging bp base pairs

BSA bovine serum albumin

β2M β2-microglobulin

CCNU 1-(2-chloroethyl)-3-cyclohexyl-l-nitrosourea

CDDP cisplatin cDNA complementary deoxyribonucleic acid

CIN chronophin

CML chronic myelogenous leukaemia

CNS central nervous system

Ctrl control

DAPI 4',6-diamidino-2-phenylindole

DMEM Dulbecco’s Modified Eagle Media

DMSO dimethyl sulfoxide

DNA deoxyribonucleic acid

DPBS Dulbecco’s phosphate buffered saline

DTT dithiothreitol

EB end-binding

ECM extracellular matrix

EFS event-free survival

EMT epithelial to mesenchymal transition

ERK extracellular signal-regulated kinase

FCS foetal calf serum

GAPDH glyceraldehyde 3-phosphate dehydrogenase

GDP guanosine diphosphate

GD2 disialoganglioside

GFP green fluorescent protein

GTP guanosine triphosphate

H&E haematoxylin and eosin

HCC hepatocellular carcinoma

HRP horseradish peroxidase

HSV1-tk herpes simplex virus 1- thymidine kinase

IHC immunohistochemistry

INSS International Neuroblastoma Staging System i.p. intraperitoneal kD kilodaltons

KIS kinase interacting with stathmin

KO knockout

LIMK LIM domain-containing kinase

LOH loss of heterozygosity mA milliamps

MAP microtubule-associated protein

MAPK mitogen activated protein kinase

MEFs mouse embryonic fibroblasts

MIBG meta-iodobenzyl-guanidine miRNA microRNA

MKI mitosis-karyorrhexis index

MLC myosin light chain

MMLV Moloney Murine Leukaemia Virus

MMP matrix-metalloproteinase

MQ-H2O milli-Q water mRNA messenger ribose nucleic acid

MTOC microtubule organising centre

MYCN neuroblastoma-derived v-myc myelocytomatosis viral related oncogene

NGF nerve growth factor

NSCLC non-small cell lung cancer

NSE neuron-specific enolase

PACAP pituitary adenylate cyclise-activating polypeptide

PAGE polyacrylamide gel electrophoresis

PBS phosphate buffered saline

PBS-T 0.05% Tween-20/phosphate buffered saline

PDGF platelet derived growth factor

PFA paraformaldehyde

PI propidium iodide

PI3K Phosphatidylinositol 3-kinase

PMA phorbol 12-myristate 13-acetate pRS retroviral silencing plasmid

PTM post-translational modification qPCR quantitative polymerase chain reaction

RAS renin-angiotensin system

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RhoGEFs Rho guanine nucleotide exchange factors

RIPA radio-immunoprecipitation assay

RNA ribonucleic acid

RNAi ribonucleic acid interference

ROCK Rho associated coiled-coil forming protein kinase

RPMI Roswell Park Memorial Institute

SCID Severe-combined immune-deficient

SCLC small-cell lung cancer

SDS sodium dodecyl sulphate sec second

SEM standard error of the mean

Ser serine shRNA short hairpin ribonucleic acid siRNA small interfering ribonucleic acid

SMP skim milk powder

SSH slingshot

STMN stathmin

STR short tandem repeat

TBA tubulin-binding agent

TBS Tris-buffered saline

TBS-T 0.05% Tween-20/Tris-buffered saline

TGF-β transforming growth factor beta

TMZ temozolomide

TPA tetradecanoylphorbol 13-acetate

TXL paclitaxel

VCR vincristine

VLB vinblastine

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VP-16 etoposide

VSMC vascular smooth muscle cell

WASP Wiskott-Aldrich Syndrome Protein

WAVE WASP and Verprolin homologous protein

5-FU 5-fluorouracil

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Publications arising from this thesis

1. Byrne F.L., Phillips P., Hansford L., Fletcher J., Ormandy C., McCarroll J.A.,

Kavallaris M. Stathmin suppression reduces neuroblastoma cell invasion and

metastasis. Manuscript in preparation.

2. Ng D. and Byrne F.L. (2011). Chapter 14: Stathmin and Cancer. In: Kavallaris

M. (Eds) “The Cytoskeleton and Human Disease”, Springer, NY USA. In press.

Abstracts

Byrne F.L., McCarroll J., Kavallaris M. Stathmin promotes neuroblastoma cell invasion and metastasis. ASMR MRW® XIX NSW Scientific Meeting, University of

Sydney, Sydney, NSW. Oral Presentation (awarded the University of Sydney Medal for

Best Overall Presentation).

Frances L. Byrne, Josh McCarroll, Maria Kavallaris. Stathmin mediates neuroblastoma cell migration and invasion via regulation of the actin network. American Association for Cancer Research annual meeting, Washington DC, USA 2010. Poster presentation.

Byrne F.L., McCarroll J., Kavallaris M. Stathmin contributes to the migratory and invasive phenotype of neuroblastoma. 37th Annual Meeting of the Coast Association

2009 Tow Research Awards, Sydney, NSW. Oral Presentation.

Acknowledgements

First and foremost, I would like to thank Professor Maria Kavallaris for not only allowing me to complete my PhD with her, but for her continual support and guidance throughout my project. She encouraged independent thinking and allowed me to test my own hypotheses, which for scientists is the ultimate freedom. Her motivation and dedication to science is inspirational and she has been a fantastic role model for me and many others wanting to pursue a career in this challenging field. I would also like to thank my co-supervisors Dr. Joshua McCarroll and Dr. Toby Trahair for all their informative discussions, encouragement and assistance during my PhD. In particular

Josh, who not only helped me in the lab (all those hours in the animal facility!), but was there through all the ups and downs and gave up so much of his time to talk about my project, careers and science in general. He has been a true friend and mentor to me in so many aspects of my life and I can’t imagine how I could have got through my PhD without him!

I would like to express my sincerest gratitude to Professor Michelle Haber and everyone at the Children’s Cancer Institute Australia for being so supportive and making my time at the institute a very enjoyable (and productive) experience. Past and present members of the Tumour Biology & Targeting program, including Josh, Sela, Alice, Marion,

Michael, Pei Pei, Tracy and Mel, have all been incredibly supportive and wonderful friends. I would also like to thank the Animal facility staff, Dr. Jamie Fletcher, Zillan

Neiron and Jess Koach for their training and assistance with experiments, and Amanda

Philp for organising student activities which have provided valuable insights into career opportunities for scientists. Outside of CCIA I would like to thank my collaborators Dr.

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Phoebe Phillips, Dr. Loen Hansford, Professor Chris Ormandy and Anita Ledger for all their help with my studies, and the Anthony Rothe Memorial Trust and Louiza Zervos

Memorial Trust for funding my PhD project.

And last, but definitely not least, a huge thank you to my wonderful family and friends in Sydney, Adelaide and overseas. In particular, Kyle, my parents (Nigel and Paula), brother (Rich), Anna and my fluff balls (Oskar & Sooty) who have given me so much love, support, guidance and encouragement throughout my time in Sydney and during my PhD. They have helped shaped the person and scientist I am today and I love them all dearly.

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Abstract

Neuroblastoma originates from precursor cells of the sympathetic nervous system and is one of the most deadly childhood cancers, accounting for 15% of all paediatric cancer deaths. Metastasis, drug resistance and poor tumour cell differentiation all contribute to an aggressive neuroblastoma phenotype. With a better understanding of these elements, new drug targets can be identified to improve treatment and ultimately survival rates in this aggressive disease.

Stathmin is a microtubule destabilising protein highly expressed in neuroblastoma. Until now, stathmin’s functional role in this malignancy had not been addressed. The aims of this study were to evaluate stathmin’s contribution to neuroblastoma drug resistance, differentiation and metastasis. A small interfering RNA (siRNA) approach was utilised to suppress stathmin expression and determine stathmin’s functional role in 2 independent neuroblastoma cell lines, BE(2)-C and SH-SY5Y. SiRNA-mediated stathmin suppression did not markedly influence neuroblastoma cell proliferation or sensitivity to a range of DNA-damaging and tubulin-binding agents. In contrast, stathmin played a role in the differentiation of neuroblastoma cells induced by retinoic acid, a differentiation agent used to treat high risk neuroblastoma. Short-term exposure to all-trans retinoic acid (ATRA) induced stathmin phosphorylation and neurite outgrowth in BE(2)-C and SH-SY5Y cells. Moreover, stathmin suppression significantly inhibited ATRA-induced neurite formation in both neuroblastoma cell lines. However, stathmin’s role in neuroblastoma differentiation was restricted to neurite outgrowth as stathmin suppression did not influence ATRA-induced growth

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arrest or ATRA-induced alterations in differentiation marker expression in neuroblastoma cells.

The migration of cancer cells away from the primary tumour and invasion of the surrounding extracellular matrix (ECM) are two key steps in the metastatic process.

Although stathmin did not play a role in 2D cell migration, stathmin suppression significantly inhibited the migration and invasion of BE(2)-C and SH-SY5Y cells toward chemo-attractants. To determine the mechanism by which stathmin was regulating cell migration and invasion, the cell cytoskeleton, which plays a critical role in these processes was examined in stathmin-suppressed neuroblastoma cells. Stathmin suppression increased the numbers of short and thin, actin-rich neurite-like projections, microtubule density and tubulin polymer levels in BE(2)-C cells compared to controls.

Moreover, stathmin suppression increased the phosphorylation of cofilin and myosin light chain (MLC), two key actin-regulatory proteins, compared to control BE(2)-C cells. Treatment of stathmin-suppressed BE(2)-C cells with the Rho associated coiled- coil forming protein kinase (ROCK) inhibitor, Y-27632, ablated MLC phosphorylation and returned cofilin phosphorylation and cell invasion back to that of untreated controls.

Therefore stathmin appears to regulate the phosphorylation of cofilin and MLC, and cell invasion, through ROCK. However, the increase in cofilin phosphorylation observed in stathmin-suppressed cells was only partially reversed by Y-27632 indicating that stathmin also regulates cofilin phosphorylation via an alternate pathway.

Stathmin’s effects on neuroblastoma cell migration and invasion suggested a role for stathmin in neuroblastoma metastasis. In order to examine this phenotype in vivo, stathmin shRNA/luciferase-expressing neuroblastoma cells [SK-N-BE(2)/TGL] were

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developed and characterised for use in a clinically relevant orthotopic neuroblastoma mouse model. SK-N-BE(2)/TGL cells were stably transfected with 2 individual stathmin shRNA constructs or a control shRNA construct. ShRNA-expressing SK-N-

BE(2)/TGL cells were injected into the left adrenal fat pad of SCID-Beige mice.

Tumour growth was monitored weekly using bioluminescent imaging (BLI). Although stathmin was effectively suppressed in stathmin shRNA-expressing primary tumours

(85% reduction at the protein level) compared to controls, stathmin did not influence neuroblastoma cell engraftment or tumour growth. In contrast, BLI showed that stathmin suppression significantly reduced neuroblastoma tumour burden in the lungs of mice by 71% (p<0.008) compared to controls. Luciferase immunostaining confirmed that the bioluminescent signal in lungs was due to neuroblastoma cells that had metastasised to this organ. This data highlights stathmin as a potential target to treat metastatic neuroblastoma.

In conclusion, this study has demonstrated that stathmin regulates neurite outgrowth, cell migration, invasion and metastasis in neuroblastoma; previously unreported roles for this protein in this malignancy. Moreover, stathmin’s regulation of neuroblastoma cell invasion appears to be mediated by blocking ROCK signalling which is responsible for amoeboid-like cell motility. Thus stathmin may promote the alternate mode of cell motility, mesenchymal-like cell motility, in neuroblastoma cells. Further investigations are therefore required to determine whether stathmin drives a particular type of cell motility in neuroblastoma cells and more importantly, whether targeting stathmin in neuroblastoma has the potential to reduce/prevent metastasis in this deadly childhood cancer.

Chapter 1. Introduction

Every year in Australia more than 100,000 people are diagnosed with cancer and over

40,000 will die from this disease. The most difficult to treat cancers are those that have spread throughout the body (metastatic disease). These types of cancers contain highly proliferative immature cells that are resistant to even the most advanced multi-modality treatments. As such, metastatic disease contributes to most cancer deaths (Langley and

Fidler 2011).

Cancer cells rely on their cytoskeleton for survival, growth and movement through the body, making chemotherapy drugs that target the cytoskeleton the most successful anticancer agents to date (Jordan and Wilson 1998). Despite their success, cancer cells develop resistance to these agents which hampers efforts to effectively eliminate the disease. Furthermore, cancer patients often experience severe toxicity and high morbidity from current chemotherapy regimens, re-enforcing the move towards individualised therapy and identification of new anti-cancer targets. This thesis will examine the role of a specific cytoskeletal protein, stathmin, in the highly metastatic childhood cancer neuroblastoma, to evaluate its potential as a new therapeutic target for this deadly disease.

1.1 Neuroblastoma

Neuroblastoma is the most common extra-cranial childhood tumour accounting for 6-

10% of all childhood cancers and 15% of all paediatric oncology deaths (Gutierrez,

Fischer et al. 2007; Maris, Hogarty et al. 2007). Neuroblastoma originates from precursor cells of the sympathetic nervous system that form tumours along the sympathetic chain including the abdomen (adrenal medulla), neck, pelvis, thorax and cervix (Gutierrez, Fischer et al. 2007; Maris, Hogarty et al. 2007). Neuroblastoma presents as an extremely heterogenic malignancy in regards to clinical, biological, histological and genetic features. It is predicted that germline mutations account for

20% of cases and an autosomal-dominant pattern of inheritance has been identified in some patients. However, the aetiology of neuroblastoma remains largely unknown

(Brodeur 2003).

1.1.1 Prognostic Indicators

1.1.1.1 Age & Disease Stage

Two strong prognostic indicators for neuroblastoma are patient age and disease stage, as defined by the International Neuroblastoma Staging System (INSS) (Brodeur, Pritchard et al. 1993). The median age for diagnosis is 18 months and 40% of patients are diagnosed by the age of one [reviewed in (Brodeur 2003)]. Children more than 1 year old usually present with widely disseminated disease and have a 5 year survival rate less than 50% (Gutierrez, Fischer et al. 2007). Infants are commonly diagnosed with small tumours that have often spread to the liver, skin and bone (stage 4-S) [reviewed in

(Maris, Hogarty et al. 2007)]. Although metastatic, stage 4-s tumours can spontaneously regress making them distinct from other advanced stages and have a good prognosis

(Miale and Kirpekar 1994). Approximately 40% of patients present with limited stage disease (stages 1, 2A and 2B) which is characterised by localised tumours that can be excised and that may or may not have spread to the ipsilateral lymph nodes [reviewed in

(Brodeur, Pritchard et al. 1993; Brodeur 2003; Maris, Hogarty et al. 2007)]. However,

50% of patients present with widely disseminated disease classifying them as advanced stage (stages 3, 4 and 4S) (Maris, Hogarty et al. 2007). Stage 3 tumours are those that have; i) spread across the midline, ii) are midline tumours with bilateral extension or lymph node involvement, or iii) are localised but have contralateral regional lymph node involvement (Maris, Hogarty et al. 2007). In stage 4 disease, the cancer has spread to distant organs, bones and lymph nodes and five year survival rates for these patients are less than 35% (Gutierrez, Fischer et al. 2007; Pearson, Pinkerton et al. 2008).

1.1.1.2 Genetic Abnormalities

Amplification of the neuroblastoma-derived v-myc myelocytomatosis viral related oncogene (MYCN) and other chromosomal alterations serve as strong prognostic factors in neuroblastoma. The MYCN transcription factor is amplified in 22% of patients and is associated with advanced stage disease, rapid tumour progression and poor prognosis

[reviewed in (Brodeur 2003)]. High chromosome number, or near-triploidy, is considered favourable particularly in infants (Brodeur 2003). In contrast, loss of heterozygosity (LOH) on the short arm of chromosome 1p (1p36) confers a poor prognosis and is associated with MYCN amplification and advanced stage disease

(Brodeur 2003). It has been proposed that LOH at this region is likely to affect genes involved in neural cell differentiation, cell cycle and signal transduction (Janoueix-

Lerosey, Novikov et al. 2004). Deletion of the 11q allele occurs in 43% of patients and favours a poor prognosis. This deletion is directly associated with loss of 14q, but is inversely related to 1p deletion and MYCN amplification, as reviewed in (Brodeur

2003).

More recently, activating mutations, amplification and copy number gains of the receptor tyrosine kinase gene, anaplastic lymphoma kinase (ALK) (located on chromosome 2p23) have been identified in neuroblastoma that frequently occur in combination with MYCN amplification (Azarova, Gautam et al. 2011). Activation of

ALK by ligand binding, mutations and amplification stimulate numerous oncogenic signalling pathways such as PI3K/Akt, ERK and MAPK (Azarova, Gautam et al. 2011) and dominant mutations in this gene are shown to increase MYCN expression in neuroblastoma cells and induce transformation of NIH3T3 cells (Schonherr, Ruuth et al. 2012). Thus inhibitors and antibodies against ALK have been an area of intense investigation for the potential treatment of neuroblastoma (Bresler, Wood et al. 2011; Di

Paolo, Ambrogio et al. 2011; Di Paolo, Brignole et al. 2011; Carpenter, Haglund et al.

2012).

1.1.1.3 Histology

Neuroblastoma is diagnosed by histopathological examination of tumour tissue and bone marrow aspirates or biopsies (Maris, Hogarty et al. 2007). Benign neuroblastoma tumours, termed ganglioneuromas, contain mature neuron clusters covered by a thick stroma of Schwann cells while malignant neuroblastomas are characterised by small, round and poorly differentiated neuroblasts (Brodeur 2003). These features, as well as the number of mitotic and nucleus-fragmented cells per 5000 cells [mitosis-karyorrhexis

index (MKI)], define neuroblastoma histology according to the International

Neuroblastoma Pathology Classification system (Shimada, Ambros et al. 1999).

Unfavourable tumours are those that are poorly differentiated, Schwannian stroma-poor and have a high MKI (>200 cells) (Shimada, Ambros et al. 1999).

1.1.1.4 Gene expression

The aberrant expression of three neurotrophin receptors, tyrosine kinase receptors A, B and C, are strong predictors of outcome in neuroblastoma. Unfavourable neuroblastomas express high levels of tyrosine kinase receptor B (TkB) and its ligand, brain-derived neurotrophic factor (BDNF), which induce an autocrine loop thought to promote proliferation, chemoresistance and metastasis (Maris and Matthay 1999).

Tyrosine kinase receptor A expression, with or without tyrosine kinase receptor C expression, is considered favourable as it is usually found in more differentiated tumours (Maris and Matthay 1999; Brodeur 2003). Other poor prognostic factors include elevated telomerase, B-cell lymphoma-2 (BCL2), renin-angiotensin system

(RAS), multidrug efflux pumps, nerve growth factor (NGF)-binding transmembrane receptor, p75 neurotrophin receptor (p75NTR) (Brodeur 2003) and ALK (Azarova,

Gautam et al. 2011). High levels of the cell-surface glycoprotein CD44 is associated with more differentiated and non MYCN-amplified tumours, and therefore confers a good prognosis (Brodeur 2003). High risk patients often have elevated serum ferritin, lactate dehydrogenase, disialoganglioside (GD2), chromagranin and neuron-specific enolase (NSE) which are associated with a poor outcome (Brodeur 2003; Maris,

Hogarty et al. 2007). However these features are not of prognostic value and are therefore not used in patient stratification (Brodeur 2003; Maris, Hogarty et al. 2007).

1.1.2 Risk Classification

Disease stage, age, MYCN amplification, histology and DNA index (ploidy) define neuroblastoma risk groups. All patients with stage 1, most patients with stage 2

(excluding those with MYCN amplification) and those with stage 4-S disease (non- amplified MYCN, favourable histology, DNA index > 1) are classified as low risk and account for 40% of all neuroblastoma cases (Brodeur 2003; Maris, Hogarty et al. 2007).

Non MYCN-amplified patients with stage 3 or 4 (age <1 year, favourable histology,

DNA index >1) or 4-S (age <1 year, unfavourable histology, DNA index > 1) are classified as intermediate risk and make up 25% of patients (Brodeur 2003; Maris,

Hogarty et al. 2007). Both low and intermediate risk groups have high event-free survival (EFS) rates over 80% (Maris and Matthay 1999). The remaining patients harbouring MYCN amplification or those with stage 4 disease older than 1 year are classified as high risk and have dismal prognosis with 5 year survival rates of 30%

(Matthay, Villablanca et al. 1999; Maris, Hogarty et al. 2007; Pearson, Pinkerton et al.

2008).

1.1.3 Treatment

Improved detection methods (early detection), risk stratification, surgery, pharmacological intervention (high dose chemotherapy and retinoic acid) and bone marrow or stem cell transplants have markedly increased survival rates for neuroblastoma over the past three decades (Gutierrez, Fischer et al. 2007). For low risk disease, surgery alone is sufficient or combined with low-dose chemotherapy if tumour removal is incomplete (Castleberry 1997). Removal of the primary tumour has also

proved particularly beneficial for patients with stages 2B and 3 disease (Gutierrez,

Fischer et al. 2007). However, for children with high risk disease, 10 year survival rates remain dismal (20-30%) even with high dose chemotherapy (myeloablation), successful removal of the primary tumour, local radiation and retinoid treatment (Armstrong,

Redfern et al. 2005; Pearson, Pinkerton et al. 2008). Therefore there is still an urgent need to identify new drug targets for the treatment of high risk neuroblastoma.

The most commonly used chemotherapeutics for the treatment of neuroblastoma include the tubulin-binding agent (TBA) vincristine, DNA-targeted pro-drugs

(cyclophosphamide, topotecan), DNA alkylating agents (cisplatin, carboplatin, melphalan), DNA intercalating agents (doxorubicin) and topoisomerase II inhibitors

(etoposide, teniposide) (Pearson, Pinkerton et al. 2008). Vincristine and cisplatin are some of the most effective but also the least myelotoxic of all these agents (Pearson,

Pinkerton et al. 2008). The synthetic retinoid isotretinoin (13-cis-retinoic acid) is administered to high risk patients intermittently for 6 months (160mg/m2 daily for 2 weeks/month) to induce tumour cell differentiation, an effect shown to markedly improve survival rates for these patients (Matthay, Villablanca et al. 1999; Pearson,

Pinkerton et al. 2008). For high risk patients, radioactive meta-iodobenzyl-guanidine

(MIBG) is also used for disease palliation and for monitoring tumour growth and bone metastases during therapy (Maris, Hogarty et al. 2007). New therapies in early phase clinical trials for neuroblastoma include the novel tubulin-binding agent ABT-751

(Phase II), monoclonal antibodies to GD2, fenretinide (synthetic retinoid derivative) and the small molecule Trk inhibitor CEP-701 (Maris, Hogarty et al. 2007).

1.1.4 Drug Resistance

Although most neuroblastoma patients respond well to the first round of cyto-reductive therapy, many will become resistant as treatment progresses. Drug resistance in neuroblastoma is thought to arise via multiple genetic defects or aberrations in signalling pathways. These include DNA methylation (Qiu, Mirkin et al. 2005), activation of the TrkB/BDNF pathway (Thiele, Li et al. 2009) and disruption to cell death pathways (Goldsmith and Hogarty 2005; McKee and Thiele 2006; Swarbrick,

Woods et al. 2010). Other factors that may contribute to drug resistance in neuroblastoma include alterations in cytoskeletal proteins that mediate resistance to specific cytotoxic agents, such as the TBAs (Don, Verrills et al. 2004; Po'uha, Shum et al. 2009). However, the most widely studied drug resistance mechanism in neuroblastoma is that of MYCN and its target genes.

1.1.4.1 The MYCN Oncogene

MYCN amplification remains one of the strongest prognostic indicators for neuroblastoma. The MYCN gene, located on the short arm of chromosome 2, is amplified in one fifth of all neuroblastoma tumours though the cause of amplification is unknown (Brodeur 2003). MYCN encodes the N-myc protein which when bound to max, acts as a transcriptional activator (Wenzel and Schwab 1995). Activation of MYCN target genes such as the multiple drug resistance protein 1 (MRP1), ornithine decarboxylase (ODC) and minichromosome maintenance protein promote G1 cell cycle progression and chemoresistance (Brodeur 2003). MRP1 pumps various chemotherapeutic agents including vincristine, doxorubicin and etoposide out of the cell reducing their cytotoxic effects (Haber, Bordow et al. 1999; Manohar, Bray et al. 2004;

Munoz, Henderson et al. 2007). MYCN also targets the recently discovered small non- coding RNAs, microRNAs (miRNAs) that bind to the 3ʹ untranslated region (3ʹUTR) of genes to repress protein expression. Fontana et al. identified that MYCN transactivation of the microRNA 17-5p-92 cluster confers drug resistance via repression of the cyclin- dependent kinase inhibitor p21 and Bcl-2 interacting mediator of cell death (BIM)

(Fontana, Fiori et al. 2008). Interestingly MYCN itself is regulated by miRNAs, in particular miR-34a, which is located at 1p36. LOH at this region results in reduced miR-

34a expression and up-regulation of MYCN (Wei, Song et al. 2008). Moreover, endogenous expression of miR-34a in MYCN-amplified cell lines reduced MYCN expression, DNA synthesis and induced apoptosis (Wei, Song et al. 2008). Therefore, amplification or over-expression of MYCN, and activation of its downstream targets, significantly contribute to neuroblastoma proliferation, survival and drug resistance.

Although MYCN itself has proved a difficult therapeutic target, the development of

ODC (Rounbehler, Li et al. 2009) and MRP1 inhibitors (Burkhart, Watt et al. 2009) may prove beneficial for the treatment of MYCN-amplified or over-expressed tumours.

1.1.5 Metastatic Disease

Neuroblastoma is a highly metastatic disease with more than 70% of patients harbouring widely disseminated disease at diagnosis (Ara and DeClerck 2006). Metastasis is initiated when tumour cells detach from the primary tumour, invade the surrounding tissues and enter the blood or lymphatic system (Fig. 1.1). Once in circulation, some tumour cells will lodge at a distant site and are able to grow and thrive in their new environment by developing their own blood supply (Fig. 1.1). The most common sites for metastases in neuroblastoma are the bone marrow (70% of patients), bone (56% of

Figure 1.1The formation of a metastatic tumour.

Metastasis is initiated when cancer cells detach from the primary tumour and migrate and invade the surrounding tissue. These cells then penetrate the endothelium and enter the blood or lymphatic system. Once in circulation, cancer cells can form emboli with other cancer cells or other cell types which then arrest at a distant site. Some cancer cells then invade the surrounding tissue and grow in this new environment by developing their own blood supply (vasculature). Other cells in the microenvironment secrete growth factors to help the new tumour grow. Image adapted from (Ara and

DeClerck 2006).

Endothelium

Cancer cell

Emboli

Growth factors

Vasculature

Primary Metastatic tumour Migration & Invasion tumour

patients), lymph nodes (31% of patients), liver (30% of patients), intracranial and orbital sites (18% of patients), lungs (3% of patients) and central nervous system (CNS) (> 1% of patients) (DuBois, Kalika et al. 1999; DuBois, London et al. 2008). A study of 648 stage 4 and 4S patients found that those with metastases in the bone marrow, bone, cranium, lung and CNS had lower EFS compared to those with liver and skin metastases (DuBois, Kalika et al. 1999). While the driving force underlying metastases in this cancer are unclear, it is thought that multiple factors contribute to this phenotype.

Neuroblastoma tumour cells aberrantly express cell adhesion molecules, integrins and matrix-metalloproteinases (MMPs) which influence cell-cell and cell-extracellular matrix (ECM) interactions (Meyer, van Golen et al. 2004; Ara and DeClerck 2006;

Nyalendo, Beaulieu et al. 2008). Tumour-stromal cell interactions also greatly influence the ability of neuroblastoma cells to colonise a new site. For example, bone marrow stromal cells and osteoblasts secrete various chemokines such as stromal derived factor

(SDF-1)/CXCL12 which attracts CXCR4 chemokine receptor-expressing neuroblastoma cells to the bone marrow [reviewed in (Ara and DeClerck 2006)]. The activation of the CXCR4 chemokine receptor up-regulates various growth factors and integrins which promote attachment, colonisation and proliferation of neuroblastoma cells in the bone marrow and bone [reviewed in (Ara and DeClerck 2006)].

In advanced-stage neuroblastoma, the location of metastases correlates with patient age and MYCN amplification (DuBois, Kalika et al. 1999). It is therefore thought that

MYCN may also play a key role in neuroblastoma metastasis. Interestingly, the expression of Twist, a transcription factor and promoter of metastasis in other malignancies, is over-expressed in MYCN-amplified tumours and is shown to prevent

MYCN-driven alternative reading frame (ARF)/p53-dependent apoptosis (Valsesia-

Wittmann, Magdeleine et al. 2004). More recent studies have shown that MYCN regulates the expression of miRNAs that in turn regulate the cell junction protein cadherin (Ma, Young et al. 2010). Furthermore, MYCN may also contribute to the metastatic phenotype of neuroblastoma by regulating various cytoskeletal proteins which are critical for cell migration and invasion [reviewed in (Ara and DeClerck

2006)].

1.2 The Cell Cytoskeleton

The eukaryotic cell cytoskeleton comprises three major protein filaments; microfilaments (actin), intermediate filaments such as keratin and vimentin, and microtubules, all of which provide structural support and aid multiple cell functions.

Interactions between the actin and microtubule networks in particular are critical for cell division, differentiation, morphogenesis and motility (Rodriguez, Schaefer et al. 2003).

These networks are temporally and spatially regulated by a vast array of interacting proteins that enable them to rapidly change shape and length in order to fulfil their normal functions (Rodriguez, Schaefer et al. 2003). Alterations in cytoskeletal and cytoskeletal-regulatory proteins have been identified in cancer cells that promote tumour survival, progression, drug-resistance and metastasis. Thus some of these proteins have proved extremely effective anti-cancer targets (Bhat and Setaluri 2007).

1.2.1 Microtubule Network

1.2.1.1 Structure & Function

Microtubules are a network of dynamic structures critical for diverse cell functions such as cell division, vesicular transport, polarisation and motility. Microtubules comprise of

α/β-tubulin heterodimer repeats that bind head to tail to form protofilaments (Figs. 1.2,

1.3). Each α- and β-tubulin subunit is approximately 450 amino acids with a molecular mass of 55 kilodaltons (kD) [reviewed in (Wade 2009)]. Lateral interactions between 13 protofilaments produce long, 25nm wide microtubule cylinders (Wade 2009) (Fig. 1.3).

The heterodimer repeat pattern within microtubules governs their polarity and dynamic nature of the microtubule ends. The most rapid growing microtubule end, the plus end, is capped by β-tubulin subunits while the slower growing minus end is crowned by α- tubulin subunits (Wade 2009) (Fig. 1.3).

Figure 1.2 Microtubules and actin filaments.

Black and white confocal microscopy images of filamentous actin and microtubules in neuroblastoma cells. The merged image shows actin in red, microtubules in green and nuclei in blue. Images are derived from experiments conducted in this thesis.

Actin

10 μm

Microtubules

Merge

Figure 1.3 Microtubule ‘dynamic instability’ is governed by GTP hydrolysis.

GTP-loaded tubulin heterodimers form a GTP-cap at the microtubule plus end (capped by β-tubulin) to prevent depolymerisation. Hydrolysis of GTP at the catalytic loop on α- tubulin induces conformational changes to the microtubule lattice destabilising lateral protofilament interactions and inducing depolymerisation. An abrupt change in microtubule dynamics from shortening to growth is the ‘rescue’ phase while a growth to shortening phase is termed ‘catastrophe’. Image adapted from Jordan 2004.

Stable microtubule Tubulin-bound -end + end GTP Tubulin-bound GTP or GDP

Rescue GTP Cap

RAPID TRANSITION Cap loss

Catastrophe

Tubulin-bound GDP

Depolymerising microtubule

1.2.2 Microtubule Dynamics & GTP hydrolysis

Microtubules rapidly change between phases of growth (polymerisation) or shortening

(depolymerisation), a property termed ‘dynamic instability’. An abrupt transition from microtubule shortening to growth is termed ‘rescue’ while growth to shortening is termed ‘catastrophe’ [reviewed in (Wade 2009)] (Fig. 1.3). Microtubule dynamic instability is governed by hydrolysis of guanosine triphosphate (GTP) within the microtubule lattice. GTP bound to the exchangeable site (E-site) on β-tubulin is trapped within tubulin heterodimers upon microtubule polymerisation (Wade 2009). Addition of a GTP-loaded heterodimer layer, or a GTP-cap, at the β-tubulin plus ends prevents microtubule depolymerisation while a catalytic loop on α-tubulin promotes GTP hydrolysis (Wade 2009) (Fig. 1.3). This process induces a conformational change from straight to a more energetically-favourable curved shape (Wade 2009). In this conformation, lateral protofilament interactions are destabilised resulting in microtubule depolymerisation (Wade 2009) (Fig. 1.3). Another way microtubules act dynamically is the process of ‘treadmilling’ whereby a net flow of tubulin subunits moves from the plus end to the minus end without affecting microtubule length (Nogales 2000).

1.2.3 Microtubule Assembly & Stability

During cell division specific types of microtubules, termed astral microtubules, radiate outwards from centrosomes and interdigitation of these with chromosomal microtubules, and subsequent minus end focusing, create the spindle pole to separate

chromsomes [reviewed in (Wade 2009)]. These microtubules are 4-100 times more dynamic than interphase microtubules [reviewed in (Jordan and Wilson 2004)] and function to separate duplicated chromosomes during cell division (Wade 2009).

Interphase microtubules act as tracks to carry vesicles and organelles such as mitochondria to different sub-cellular compartments [reviewed in (Wittmann and

Waterman-Storer 2001)]. The functions of interphase and mitotic microtubules are heavily reliant on their assembly and stability which are influenced by a host of factors including tubulin isotype composition, post-translational modifications (PTMs), microtubule nucleation and interactions with diverse stabilising and destabilising proteins (Nogales 2000).

1.2.4 Tubulin Isotypes

The highly conserved tubulin super-family comprises seven members (α, β, γ, δ, ε, ζ and

η- tubulin), though alterations in microtubule stability are mostly attributed to the transcription and post-translational modifications of the α- and β-tubulin isotypes

(McKean, Vaughan et al. 2001). Mammalian cells express at least six α-tubulin and eight β-tubulin isotypes each with high homology except for the last 15 amino acids at the C-terminal region (Nogales 2000; Leandro-Garcia, Leskela et al. 2010). These variations are thought to provide distinct functions for the β-tubulin isotypes (Roach,

Boucher et al. 1998) and influence interactions with various microtubule-associated proteins (MAPs) and microtubule dynamics (Nogales 2000).

Eight genes encode the β-tubulin isotypes each displaying unique patterns of tissue expression (Luduena 1998; Leandro-Garcia, Leskela et al. 2010). Class I β-tubulin

(TUBB), the most ubiquitous member of the family, is highly expressed in the thymus

(Leandro-Garcia, Leskela et al. 2010). Class IIa (TUBB2A) and IIb (TUBB2B) differ by only two amino acids and are primarily expressed in the brain (Roach, Boucher et al.

1998; Leandro-Garcia, Leskela et al. 2010). Similarly, class III β-tubulin (TUBB3) is most abundant in the brain, specifically neuronal cells, but is also expressed in sertoli cells of the testes (Roach, Boucher et al. 1998; Leandro-Garcia, Leskela et al. 2010).

Class IVa (TUBB4A) is the most abundant isotype in the brain accounting for 46% of all

β-tubulin in this tissue (Roach, Boucher et al. 1998; Leandro-Garcia, Leskela et al.

2010). Class IVb (TUBB2C) on the other hand is ubiquitous and the most prominent isotype in the heart and testes (Roach, Boucher et al. 1998; Leandro-Garcia, Leskela et al. 2010). Class V (TUBB6) and class VI (TUBB1) β-tubulin are the least abundant isotypes and are primarily found in breast/lung tissues and hematopoietic cells, respectively (Leandro-Garcia, Leskela et al. 2010).

1.2.5 Microtubule Nucleation & Post- translational modifications (PTMs)

The nucleation of microtubule polymers is critical for proper functioning of the interphase microtubule network. Microtubule nucleation predominantly occurs at microtubule organising centres (MTOCs) such as the centrosomes and is initiated by longitudinal or lateral interactions between γ-tubulin in the MTOC and α-tubulin

[reviewed in (Nogales 2000)]. The α-tubulin subunit binds to the MTOC and from there the microtubules grow outwards toward the cell periphery (Nogales 2000).

Enzymatic-induced, reversible post-translational modifications (PTMs) such as detyrosination, acetylation, and phosphorylation occur at the C-terminal regions of α- and β-tubulin and are important for microtubule functioning [reviewed in (Verhey and

Gaertig 2007)]. It is thought that PTMs act as a code read by microtubule-interacting proteins which mediates their activity and in turn leads to higher-order microtubule restructuring at specific subcellular locations (Verhey and Gaertig 2007). Tubulin PTMs are enriched in slow subunit turnover microtubules and can therefore indicate microtubule stability (Verhey and Gaertig 2007).

1.2.6 Microtubule-interacting Proteins

1.2.6.1 Motor Proteins

Motor proteins such as the kinesin and dynein families help transport cargo along microtubules to different sub-cellular locations such as cilia, axons and at the leading edge of migrating cells [reviewed in (Verhey and Gaertig 2007)]. Kinesin family members (kinesin 1-14 and orphan kinesins) contain microtubule-binding and adenosine triphosphate (ATP) binding sites within their C- or N-terminus motor domains (Wade

2009). Most kinesins form long heterotetrameric complexes that become activated by

ATP hydrolysis when in close proximity to microtubules, prompting their movement towards microtubule plus ends [reviewed in (Wade 2009)]. Dyneins on the other hand use ATP hydrolysis to ‘walk’ along microtubules towards the minus ends [reviewed in

(Wade 2009)]. It is thought that microtubule-interacting proteins and tubulin PTMs can influence motor protein-microtubule interactions (Verhey and Gaertig 2007; Wade

2009). Indeed, the activity, attachment and directional movement of motor proteins can be compromised in the presence of tau-bound microtubules (Wade 2009).

1.2.6.2 Microtubule Stabilisers

Microtubule-associated proteins (MAPs), including the large (200-300 kD) members

MAP1A-1C, MAP2a-2c, MAP3, MAP4 and the small (~55kD) member tau, promote microtubule stabilisation by binding to the C-terminal domain of tubulin [reviewed in

(Maccioni and Cambiazo 1995; Bhat and Setaluri 2007)]. More specifically, the neuron- enriched MAP2 and tau attach to the external microtubule wall acting as bridges between tubulin heterodimers [reviewed in (Wade 2009)]. MAPs can also act as intermediate proteins linking microtubules to other cytoskeletal networks. For example, unphosphorylated MAP2c binds to microtubules but can also localise and interact with actin upon phosphorylation [reviewed in (Rodriguez, Schaefer et al. 2003)]. In a similar fashion, plus end tracking proteins (+TIPs) promote microtubule stabilisation and function to mediate interactions with diverse cellular structures. Examples of +TIPs include the cytoplasmic linker proteins (CLIPs), CLIP-associating proteins (CLASPs), adenomatous polyposis coli (APC), end binding (EB)-family proteins and dynactin- associated protein, p150Glued [reviewed in (Wade 2009)]. These proteins rapidly dissociate from older microtubule structures and accumulate at new growing ends, a feature that is regulated by multiple factors including phosphorylation, motor protein transport and interactions with other +TIPs [reviewed in (Akhmanova and Hoogenraad

2005)]. One of the most interesting +TIP functions includes microtubule capture at cortical sites which initiate interactions with actin and the plasma membrane [reviewed in (Akhmanova and Hoogenraad 2005)].

1.2.6.3 Microtubule Destabilisers

Microtubule destabilising proteins such as katanin, spastin, the stathmin-like proteins and kinesin-like proteins, XKCM1 and XKIF2, play critical roles in many cell functions including cell division [reviewed in (Nogales 2000)]. Katanin severs microtubules during mitosis assisting their release from centrosomes, while XKCM1 and XKIF2 act enzymatically to bind microtubule ends thereby inducing microtubule depolymerisation, an event preceding mitotic spindle assembly (Nogales 2000). The stathmin-like proteins are small phospho-proteins able to bind tubulin heterodimers and subsequently destabilise microtubules. All members of the family are abundant in the nervous system and are important regulators of neuronal microtubule dynamics during mammalian development (Curmi, Gavet et al. 1999; Chauvin, Poulain et al. 2008). Stathmin, the founding member of the family, induces microtubule depolymerisation by promoting microtubule catastrophes and/or sequestering tubulin heterodimers (Howell, Larsson et al. 1999). Recent investigations have highlighted stathmin’s contribution to metastasis and chemotherapy resistance, particularly to tubulin-binding agents, in a number of malignancies [reviewed in (Rana, Maples et al. 2008)].

1.2.7 Tubulin-Binding Agents

Tubulin-binding agents (TBAs) are an important class of chemotherapeutic drugs that bind β-tubulin at the vinca domain (vinca alkaloids), or the colchicine and taxane

(taxanes) binding sites (Jordan 2002). At high concentrations, vinca alkaloids promote microtubule depolymerisation while at low concentrations they suppress microtubule dynamics, inducing a potent mitotic block at the metaphase/anaphase junction resulting in cell death (Jordan and Wilson 2004). These agents are extremely effective in the

treatment of haematological malignancies and neuroblastoma (Jordan and Wilson

2004). Colchicine also destabilises microtubules at low concentrations by forming a complex with β-tubulin within the microtubule lattice thereby interfering with microtubule assembly (Fig. 1.4). Colchicine is not used in cancer therapy but rather for gouty arthritis and familial Mediterranean fever (Jordan and Wilson 2004). Taxanes such as paclitaxel bind the taxane site on the inner wall of the microtubule, stabilising guanosine diphosphate (GDP)-bound tubulin within the microtubule polymer. Paclitaxel is commonly used for the treatment of solid tumours including ovarian, breast, prostate and lung cancer (Jordan and Wilson 2004) (Fig. 1.4). Other drugs that stabilise microtubules are the epothilones, including epothilone B, that also binds to the β-tubulin taxane site and are used for the treatment of drug-refractory breast cancer (Jordan and

Wilson 2004) (Fig. 1.4). Although TBAs are considered some of the most successful anti-cancer agents to date, acquired or intrinsic resistance to these agents remains a major clinical problem.

1.2.8 Resistance to TBAs

Cancer cells evade the actions of TBAs by altering the expression of cytoskeletal and cytoskeletal-related proteins (Bhat and Setaluri 2007). Aberrant expression of specific

β-tubulin isotypes have been identified in TBA-resistant cell lines and patient samples that provide a selective advantage against these agents (Burkhart, Kavallaris et al.

2001). In particular, altered expression of βIII-tubulin has been identified in Vinca alkaloid-resistant (Kavallaris, Tait et al. 2001) and desoxyepothilone B-resistant leukaemia cells (Verrills, Flemming et al. 2003), and βIII-tubulin is shown to mediate resistance to tubulin-binding and DNA-damaging agents in non-small cell lung cancer

Figure 1.4 Tubulin-Binding Agents.

Tubulin-binding agents (TBA) interfere with microtubule dynamics by binding to β- tubulin. Vinca alkaloids bind to the vinca domain inducing microtubule depolymerisation at high concentrations. Similarly, colchicine can promote depolymerisation by forming a complex with β-tubulin in the microtubule lattice. The taxanes and epothilones share the taxane-binding site on the inner microtubule wall to stabilise microtubule polymers. Image adapted from (Jordan and Wilson 2004).

Vinblastine + end Paclitaxel

Tubulin- colchicine complex

-end

Vinca alkaloids Colchicine Taxanes vincristine paclitaxel vinblastine docetaxel vindesine vinorelbine Epothilones epothilone A-F

(NSCLC) cell lines (Kavallaris, Burkhart et al. 1999; Gan, Pasquier et al. 2007). Point mutations in β-tubulin and α-tubulin have also been identified in paclitaxel-resistant ovarian (Balachandran, Welsh et al. 2003) and lung adenocarcinoma cell lines

(Martello, Verdier-Pinard et al. 2003). These mutations can affect the binding of TBAs as well as interactions with key regulatory proteins such as stathmin (Martello, Verdier-

Pinard et al. 2003).

Investigations conducted by Verrills et al. demonstrated that alterations in actin can also mediate TBA resistance. Mutations in the actin isoform, γ-actin, have been identified in

TBA-resistant leukaemia cell lines (Verrills, Po'uha et al. 2006) and γ-actin expression is down-regulated in leukaemia xenografts (Verrills, Liem et al. 2006) and patient samples from clinical relapse (Verrills, Po'uha et al. 2006). Expression of γ-actin mutants confer resistance to TBAs in mouse NIH-3T3 cells while suppressing γ-actin increases TBA resistance in a neuroblastoma cell line (Verrills, Po'uha et al. 2006). A role for the actin-regulatory protein LIM domain-containing kinase, LIM kinase 2

(LIMK2), in TBA resistance has also been described. LIMK2 is elevated in neuroblastoma cell lines selected for resistance to microtubule destabilising agents

(Po'uha, Shum et al. 2009). Furthermore, silencing LIMK2 expression in a neuroblastoma cell line increased drug sensitivity, while over-expression conferred resistance to microtubule destabilising agents (Po'uha, Shum et al. 2009). These findings strengthen the links between the actin and microtubule networks and highlights

LIMK2 as a potential target for microtubule destabilising agent-refractory neuroblastoma.

Proteins that directly regulate microtubule dynamics can also confer a drug-resistant phenotype in some malignancies. The expression of the microtubule stabilising proteins,

MAP2, MAP4 and tau, are predictive of chemotherapy response, metastatic potential and patient outcome in a number of cancers [reviewed in (Bhat and Setaluri 2007)].

MAP2 expression is associated with metastatic disease-free survival in primary melanomas and increased docetaxel-sensitivity in pancreatic ductal adenocarcinomas

(Bhat and Setaluri 2007). Moreover, exogenous MAP2 expression induces microtubule stabilisation, cell cycle arrest and growth inhibition of metastatic melanoma cells in vitro and in vivo (Bhat and Setaluri 2007). Drug-resistance can also be mediated by microtubule destabilising proteins such as stathmin [reviewed in (Rana, Maples et al.

2008)].

1.2.9 Actin Network

1.2.9.1 Structure & Function

Actin is a 42 kD protein constituting up to 25% of the total protein in eukaryotic cells.

Actin is critical for many cell functions including cell division, differentiation and migration [reviewed in (Chaponnier and Gabbiani 2004)]. The actin microfilaments are constructed from globular actin subunits that polymerise to form long filamentous actin

(F-actin) structures (Fig. 1.2). Two F-actin strands twist around one another to form helical 7nm microfilaments. In vertebrates, there are six closely related actin isoforms with varying developmental and tissue-specific expression [reviewed in (Chaponnier and Gabbiani 2004)]. Four of these are restricted to muscles (α-skeletal, α-cardiac and

α- and γ-smooth muscle actin) while β- and γ-actin are ubiquitous (Chaponnier and

Gabbiani 2004). The primary structure and amino acid composition of actin is highly

conserved, with the cytosolic β- and γ-actin isoforms differing by only four amino acids at the N-terminal (Chaponnier and Gabbiani 2004). The dynamic nature of the actin network is governed by numerous interacting proteins that can also regulate remodelling of the microtubule network. The most widely studied actin-interacting proteins are those that mediate cell movements and have been implicated in metastasis.

1.2.10 Actin in Cell Motility

1.2.10.1 Assembly of Actin Structures

Signalling from the plasma membrane induces GTP-activation of the small RhoGTPase family, including RhoA, Rac1 and Cdc42, by Rho guanine nucleotide exchange factors

(RhoGEFs) [reviewed in (Insall and Machesky 2009)]. The RhoGTPases then activate a cascade of signalling pathways leading to the formation of complex actin filament structures. The sheet-like protrusion of a lamellipod is formed when Wiskott-Aldrich

Syndrome Protein (WASP) activates the Arp2/3 complex which leads to nucleation of branched actin filaments (Insall and Machesky 2009). Actin bundles or unbranched actin filaments within filopodia are made via actin interactions with fascin or the formin family of proteins, respectively (Insall and Machesky 2009). Formins, including mDia

1-3, bind to the fast growing end of actin filaments to recruit actin monomers for new filament initiation (Insall and Machesky 2009). Formins also directly interact with, and promote the formation of stable microtubules and are important for cytokenesis, cell motility and adhesion (Insall and Machesky 2009).

1.2.10.2 Disassembly of Actin Structures

The formation of new actin filaments requires the rapid disassembly of old actin filaments into ATP-charged monomers. Key proteins involved in this process include the actin-depolymerising factor (ADF)/cofilin family and coronin. Unphosphorylated cofilin binds to ADP-bound actin to sever and de-branch actin filaments thereby assisting in the recycling of actin monomers [reviewed in (Insall and Machesky 2009)].

Cofilin’s actin-binding ability is activated by the protein phosphatases slingshot (SSH) and chronophin (CIN), and becomes inactivated by phosphorylation induced by the LIM kinases, LIMK1 and LIMK2, and testicular protein kinase (TESK) downstream of the

RhoGTPase signalling pathways (Bernard 2007). Coronin acts in concert with cofilin to aid actin disassembly by inhibiting the Arp2/3 complex (Insall and Machesky 2009).

Coronin can also recruit SSH to the leading edge of migrating cells to activate cofilin in this region (Insall and Machesky 2009). Thus, a complex cycle of activity exists between actin interacting proteins to regulate actin dynamics and structure during cell movement.

1.2.10.3 Linking Actin to Microtubules

The RhoGTPases are critical for actin restructuring and increasing evidence suggests that alterations in microtubule stability can influence RhoGTPase activity and vice versa

[reviewed in (Wittmann and Waterman-Storer 2001)]. Rac1 regulates actin remodelling during cell migration (Wittmann and Waterman-Storer 2001) and its activation promotes microtubule growth in the lamellipodia of migrating cells (Wittmann and

Waterman-Storer 2001). It is believed this latter event may be mediated through regulation of microtubule-interacting proteins. Indeed, activation of Rac1 by the brain-

enriched Dock7 or lymphocyte-specific Dock2 guanine nucleotide exchange factors

(GEFs) induces phosphorylation of the microtubule-destabilising protein stathmin

[reviewed in (Holmfeldt, Sellin et al. 2009)]. Therefore regulation of stathmin downstream of the RhoGTPases may be important for microtubule restructuring that may in turn alter RhoGTPase activity and actin remodelling.

1.3 Stathmin

Stathmin, also known as pp17 (Feuerstein and Cooper 1983), prosolin (Braverman,

Bhattacharya et al. 1986), p19 (Pasmantier, Danoff et al. 1986), p18 (Hanash, Strahler et al. 1988), metablastin (Schubart, Xu et al. 1992) and p21 (Hoelscher and Ascoli 1993) was first identified in 1983 as a 17 kilo Dalton (kD) cytosolic protein rapidly phosphorylated in response to diverse extracellular stimuli (Sobel and Tashjian 1983).

Stathmin is phosphorylated upon exposure to various growth factors (Sobel and

Tashjian 1983), hormones (Doye, Boutterin et al. 1990), differentiation-inducing peptides (Dejda, Chan et al. 2010), heat shock (Beretta, Dubois et al. 1995) and antigen receptor stimulation (Imboden, Weiss et al. 1985). Thus stathmin is considered an intracellular relay protein, hence its name which is derived from the Greek word for relay “stathmos” (Sobel, Boutterin et al. 1989). The aberrant expression of stathmin in various malignancies has led to other aliases including oncoprotein 18 (Op18) (Hailat,

Strahler et al. 1990) and leukaemia-associated phosphoprotein p18 (LAP18) (Mock,

Krall et al. 1993). Over the years a number of studies have reported contradictory findings concerning stathmin’s functional role in both normal and tumour cell biology.

Therefore we have yet to fully understand the multiple roles and perhaps undiscovered roles of stathmin in different cells and malignancies.

1.3.1 Tissue Expression

Stathmin is a phylogenetically conserved protein whose expression is tightly regulated during development (Koppel, Boutterin et al. 1990). During brain ontogenesis, stathmin expression increases at embryonic day 16, peaks at the neonatal stage and then declines at the adult stage (Doye, Soubrier et al. 1989). Stathmin is predominantly expressed in neurons within the brain (Chneiweiss, Beretta et al. 1989) and other regions of the central nervous system, including the spinal cord (Bieche, Maucuer et al. 2003). Outside of the nervous system, stathmin is abundant in haematopoietic tissues, testes and in the foetal liver while moderate expression is found in the colon, female reproductive tissues

(ovary, placenta and uterus) and trachea (Bieche, Maucuer et al. 2003). The lowest expression of stathmin is found in the adult liver (Bieche, Maucuer et al. 2003). Based on stathmin’s tissue distribution during development, early studies implicated a dual role for stathmin in “developmental” regulations, relating to cell proliferation and differentiation, and to “functional” regulations in the adult, particularly in the nervous system (Koppel, Boutterin et al. 1990).

1.3.2 Structure & Function

The stathmin gene, STMN1, covers 6.3 kilobases (kb) on chromosome 1 (band p35-

36.1) and comprises of five exons and four introns (Ferrari, Seuanez et al. 1990;

Melhem, Zhu et al. 1991). Although coding for a 149 amino acid product, the first amino acid methionine is cleaved leaving an N-acetylated alanine at the N-terminus

(Labdon, Nieves et al. 1992). Stathmin is an intrinsically disordered protein with an unstructured N-terminal domain and an α-helical C-terminal domain (Zhu, Kozarsky et

al. 1989; Maucuer, Camonis et al. 1995; Steinmetz 2007) (Fig. 1.5). Stathmin’s N- terminal domain contains multiple phosphorylation sites and is therefore considered the

‘signal-integrating domain’, while the C-terminal is thought to participate in coiled-coil protein-protein interactions (Maucuer, Camonis et al. 1995). In 1996 Belmont and

Mitchison were the first to discover stathmin’s function as an inhibitor of microtubule polymerisation (Belmont, Mitchison et al. 1996). Stathmin binds to two head-to-tail aligned α/β-tubulin heterodimers forming a T2S complex (one stathmin molecule: two tubulin heterodimers) (Steinmetz 2007) (Fig. 1.5). Critical for the formation of the T2S complex are the two tubulin-interacting sites within the C-terminal region comprising homologous 35 amino acid residues (Glu48-Val82 and Glu99-Val133) (Steinmetz

2007). In this complex, stathmin promotes microtubule catastrophes and/or sequesters tubulin heterodimers in a pH-dependent manner (Howell, Larsson et al. 1999) (Fig. 1.6).

Upon binding to tubulin, stathmin can protect tubulin from tubulin-disrupting and degrading proteins, including the tubulin folding cofactor E (TBCE) and E-like proteins, thereby regulating tubulin turnover and maintaining tubulin pools (Sellin, Holmfeldt et al. 2008).

Figure 1.5 The T2S complex.

X-ray crystal structure (3.5 Å resolution) of the ternary T2S complex formed between a fragment of the stathmin homologue RB3 (yellow and red ribbon) and two α/β-tubulin heterodimers (grey structures). The N-terminal prevents longitudinal tubulin heterodimer contacts while the C-terminal is required for tubulin sequestering. The yellow ribbon at the N-terminus represents the major helix nucleation site that initiates formation of the α-helical C-terminal domain. Image adapted from (Steinmetz 2007).

α1

N-terminal ‘capping’ domain α/β tubulin heterodimer

β1

α2 C-terminal domain α/β tubulin heterodimer β2

Figure 1.6 Stathmin mediates microtubule dynamics by sequestering tubulin and/or increasing microtubule catastrophe.

At pH 6.8 stathmin sequesters tubulin heterodimers thereby preventing microtubule polymerisation. At pH 7.5 stathmin destabilizes microtubules by promoting microtubule catastrophes from the plus end (Howell, Larsson et al. 1999). Image adapted from (Rubin and Atweh 2004).

- end + end

Microtubule

Stathmin

Tubulin Catastrophe sequestering promoting pH 6.8 pH 7.5

1.3.3 Phosphorylation

Stathmin’s tubulin-binding ability is spatially and temporally regulated by phosphorylation on three serine residues in the N-terminal (Ser16, Ser25, Ser38) and one in the C-terminal domain (Ser63) [reviewed in (Steinmetz 2007)] (Fig. 1.7). A diverse range of kinases phosphorylate stathmin during mitosis including the calcium- calmodium dependent protein kinases (CaM II/IVGr), cyclin-dependent kinase

1(CDK1)/ p34cdc2 (Beretta, Dobransky et al. 1993) aurora kinase B (AurB) (Liedtke,

Leman et al. 2002) and polo-like kinase 1 (Plx1) (Budde, Kumagai et al. 2001) (Fig.

1.7). A number of growth factor-induced kinases regulate stathmin activity during interphase including various members of the mitogen-activated protein kinase (MAPK) family (Ng, Zhao et al. 2010), cyclic adenosine monophosphate-dependent kinase

(PKA) (Cassimeris 2002), kinase interacting with stathmin (KIS) (Langenickel, Olive et al. 2008) and p21-activated kinase (Pak1) (Wittmann, Bokoch et al. 2004; Takahashi and Suzuki 2009) (Fig. 1.7). The four phosphorylation sites within stathmin constitute numerous phosphoisoforms each with varying degrees of tubulin-binding affinity

(Beretta, Dobransky et al. 1993; Steinmetz 2007). In particular, Ser16 or Ser63 phosphoisoforms have significantly reduced tubulin-binding affinity while Ser25 and

Ser38 phosphoisoforms have only marginally reduced affinity for tubulin [reviewed in

(Steinmetz 2007)]. However, phosphorylation of Ser25 and Ser38 are essential for the efficient phosphorylation of Ser16 and Ser63 indicating a sequential order of stathmin phosphorylation sites (Steinmetz 2007). This phenomenon has the most relevance during mitosis in which complete phosphorylation (inactivation) of stathmin is required for cell cycle progression (Steinmetz 2007).

Figure 1.7 Signalling pathways that regulate stathmin phosphorylation.

A diverse range of receptor, ion-channel and mitotically-activated kinases induce stathmin phosphorylation at four serine residues; three in the N-terminal domain and one in the C-terminus. These kinases include the p21-activated kinase (Pak1)

(Wittmann, Bokoch et al. 2004; Takahashi and Suzuki 2009), calcium-calmodium dependent protein kinases (CAMKIV), various members of the mitogen-activated protein kinase (MAPK) family (Ng, Zhao et al. 2010), kinase interacting with stathmin

(KIS) (Langenickel, Olive et al. 2008), cyclic adenosine monophosphate-dependent kinase (PKA) (Cassimeris 2002), aurora kinase B (AurB) (Liedtke, Leman et al. 2002), polo-like kinase 1 (Plx1) (Budde, Kumagai et al. 2001) and cyclin-dependent kinase

1(CDK1)/ p34cdc2 (Beretta, Dobransky et al. 1993). The identity of the kinase “?” phosphorylating Ser63 during mitosis is unknown. Image adapted from (Holmfeldt,

Sellin et al. 2009).

Interphase Mitosis Receptor/ion channel Cell cycle-regulated -regulated kinases kinases

Rac Pak1

AurB Ca++ CAMKIV Ser16 N-terminal domain Plx1

Ras MAPK Ser25 CDK1

KIS Ser38 C-terminal domain

cAMP PKA Ser63 ?

Interestingly, microtubule assembly itself, in both mitotic and interphase cells, induces stathmin phosphorylation at Ser16 suggesting a “positive feedback loop” exists between microtubule polymer levels and stathmin activity (Kuntziger, Gavet et al. 2001).

1.3.4 Cell Cycle Regulation

As cells enter mitosis stathmin is phosphorylated at Ser25/Ser38 then Ser16/Ser63 by mitotically-active kinases in regions close to the mitotic spindle (Brattsand, Marklund et al. 1994; Larsson, Melander et al. 1995; Niethammer, Bastiaens et al. 2004). Upon mitotic exit stathmin is reactivated (dephosphorylated) by the okadaic-acid sensitive protein phosphatases PP1, PP2A and PP2B (Gavet, Ozon et al. 1998; Mistry, Li et al.

1998; Rubin and Atweh 2004). The importance of stathmin phosphorylation- inactivation during mitosis is highlighted by studies showing the effects of impaired stathmin phosphorylation on cell cycle progression. Expression of the cyclin-dependent kinase (Ser25/38) target site-mutant in leukaemia cells induced a transient G2-M delay, followed by S phase progression without proper separation of chromosomes resulting in endoreduplication (Marklund, Osterman et al. 1994; Rubin and Atweh 2004).

Additionally, forced expression of the Ser16 phosphorylation-impaired Q18>E mutation

(Misek, Chang et al. 2002) and the phosphorylation site-deficient mutant (Op18-tetraA) destabilised spindle microtubules, arrested cells at prometaphase and increased aneugenic activity in leukaemia cells (Holmfeldt, Sellin et al. 2010). Interestingly, over- expression of exogenous stathmin, which can also increase stathmin activity, induced a potent mitotic block and chromosomal instability (micronuclei formation) in leukaemia cells (Holmfeldt, Brannstrom et al. 2006; Holmfeldt, Sellin et al. 2010). Holmfeldt et al. propose that chromosomal instability caused by excessive stathmin activity during

mitosis is a result of evasion of the spindle assembly checkpoint prior to the metaphase- anaphase transition (Holmfeldt, Sellin et al. 2010). Thus tight regulation of stathmin activity is deemed critical for mitosis. Interestingly, stathmin over-expression also interferes with meiotic cell division by destabilising non-kinetochore microtubule

“barrel” arrays, decreasing spindle length, slowing kinetochore microtubule poleward flux and increasing chromosome oscillation, as observed in Xenopus egg extracts

(Houghtaling, Yang et al. 2009).

Although inactivation of stathmin is critical for mitosis, loss of stathmin can also inhibit mitotic progression. In some non-neuronal cells, inhibition of stathmin expression induces a G2-M delay and growth arrest (Mistry, Bank et al. 2005; Singer, Ehemann et al. 2007; Wang, Dong et al. 2007; Singer, Malz et al. 2009; Mitra, Kandalam et al.

2011). These findings have led researchers to speculate a role for stathmin in the disassembly of the mitotic spindle upon mitotic exit (Iancu, Mistry et al. 2001; Rubin and Atweh 2004). However in contrast to these findings, Holmfeldt et al. have demonstrated that inhibiting stathmin expression does not affect mitotic spindle assembly nor cell proliferation in a range of leukaemia cells, which led Holmfeldt et al. to speculate that stathmin may be dispensable during mitosis in some cell types

(Holmfeldt, Brannstrom et al. 2006; Holmfeldt, Sellin et al. 2010).

1.3.5 Stathmin-like Protein Family

Stathmin is the founding member of the highly homologous and conserved, stathmin- like protein family. Family members include superior cervical ganglion-10 protein

(SCG10) (STMN2), SCG10-like protein (SCLIP) (STMN3), RB3 (STMN4) and its splice

variants RB3’ and RB3’’(Koppel, Boutterin et al. 1990; Curmi, Gavet et al. 1999;

Cassimeris 2002) (Fig. 1.8). All members are highly expressed in the mammalian nervous system with SCG10 and RB3 neural-specific while stathmin and SCLIP are ubiquitous (Sobel, Boutterin et al. 1989; Bieche, Maucuer et al. 2003). The tubulin- binding activity of the stathmin-like proteins is regulated by phosphorylation on at least one site (Ser16 on stathmin) located in their highly homologous (~70% sequence identity) “stathmin-like” domain (Cassimeris 2002) (Fig. 1.8). Unlike stathmin, which moves freely in the cytoplasm, the other stathmin-like proteins associate with vesicular and golgi membranes through palmitoylation of their N-terminal cysteines (Fig. 1.8)

(Chauvin, Poulain et al. 2008). Given their high homology and shared tubulin-binding function, some propose that the stathmin-like proteins may be functionally redundant and that loss of stathmin may lead to compensation by these proteins (Bieche, Maucuer et al. 2003; Singer, Malz et al. 2009). Indeed, RB3 expression is elevated in stathmin knockout (KO) mouse embryonic fibroblasts (MEFs) (Ringhoff and Cassimeris 2009) and SCLIP expression is up-regulated in the nervous system of aging stathmin KO mice

(Liedtke, Leman et al. 2002). Moreover, like stathmin, SCLIP is involved in regulating cell proliferation and migration in non-small cell lung cancer cells (Singer et al, 2009) while SCG10 negatively regulates neuronal cell migration (Westerlund, Zdrojewska et al. 2011). However, the stathmin-like proteins also play distinct functional roles, particularly in neuronal cells, during cell polarisation and axon formation as a result of membrane-directed sub-cellular localisation (Poulain and Sobel 2009; Higuero,

Sanchez-Ruiloba et al. 2010).

Figure 1.8 The stathmin-like protein family.

All members of the stathmin-like protein family, stathmin, SCG10, SCLIP, RB3 and its splice variants RB3’ and RB3’’, bind to tubulin, destabilise microtubules and harbour serine phosphorylation (P) sites in their stathmin-like domain (~70% amino acid identity). The four serine residues responsible for mediating stathmin’s tubulin binding affinity are Ser16, Ser25, Ser38 and Ser63. The other stathmin-like family members have larger N-terminal domains for membrane associations that dictate their sub-cellular distribution. Image adapted from (Cassimeris 2002).

Membrane-binding domain Stathmin-like domain N-terminus C-terminus 16 25 38 63 Stathmin P P P P

SCG10 P P P P

SCLIP P P P

RB3 P

RB3’ P

RB3’’ P

1.3.6 Cell Differentiation

As cells differentiate into mature cells, drastic changes in the cytoskeleton are observed that coincide with changes in stathmin activity and expression. Stathmin is rapidly phosphorylated in response to a diverse range of differentiation agents including nerve- growth factor (NGF), forskolin (Doye, Boutterin et al. 1990), tetradecanoylphorbol 13- acetate (TPA) (Chneiweiss, Cordier et al. 1992; Jones, Lord et al. 1992), phorbol 12- myristate 13-acetate (PMA) (Feuerstein and Cooper 1983; Wang, Liao et al. 1993) and pituitary adenylate cyclise-activating polypeptide (PACAP) (Dejda, Chan et al. 2010).

In some cells, stathmin expression increases during differentiation, as observed during retinoic acid-induced differentiation of embryonal carcinoma cells (Maltman, Christie et al. 2009), NGF-induced differentiation of rat pheochromocytoma PC12 cells (Di Paolo,

Pellier et al. 1996) and in all-trans retinoic acid (ATRA)- and granulocyte colony stimulating factor (G-CSF)-induced differentiation of HL60 cells into neutrophils

(Johnson, Jones et al. 1995). However in most cells, continued exposure to differentiation agents leads to a decline in stathmin expression coinciding with growth arrest (Johnson, Jones et al. 1995; Rubin, French et al. 2003; Yoshie, Kashima et al.

2008). These differentiation-induced fluctuations in phosphorylation and expression suggest that stathmin plays a functional role in cell differentiation. Indeed, a forced increase in stathmin activity, via expression of constitutively active stathmin, impaired megakaryocyte maturation and platelet production (Iancu-Rubin, Gajzer et al. 2010) while stathmin inhibition prevented dibutyryl cyclic adenosine monophosphate (db- cAMP)-induced differentiation of trophoblasts (Yoshie, Kashima et al. 2008) and NGF- induced neurite formation in PC12 cells (Di Paolo, Pellier et al. 1996). However, silencing stathmin expression did not affect PACAP-induced neurite formation in PC12

cells (Dejda, Chan et al. 2010), suggesting that stathmin’s involvement in cell differentiation may be differentiation agent- and signalling pathway-dependent.

1.3.7 Neural Cell Functions

Numerous studies have demonstrated that stathmin is important for development of the nervous system, particularly the differentiation of human embryonic stem cells into post-mitotic neurons (Delaloy, Liu et al. 2010) and for neurogenesis in the brain (Jin,

Mao et al. 2004; Giampietro, Luzzati et al. 2005). Furthermore, stathmin is thought to play a neuro-protective role via microtubule-mediated regulation of caspase 3 activity in neuronal cells (Dejda, Chan et al. 2010). Stathmin’s localisation and activity are tightly regulated in neuronal cells specifically during axon development (Watabe-Uchida, John et al. 2006), dendritic arborisation in vitro (Ohkawa, Fujitani et al. 2007) and during dendritic microtubule-restructuring and motor functioning of Purkinje cells in vivo

(Ohkawa, Fujitani et al. 2007). These studies highlight an important role for stathmin in the development and maintenance of the nervous system.

Given the proposed roles for stathmin in neuronal cells, it remains unclear why young stathmin KO mice lack a developmental phenotype (Schubart, Yu et al. 1996). Young stathmin KO mice develop and behave like wild-type mice and display no alterations in

T-cell maturation or fertility (Schubart, Yu et al. 1996). It has therefore been suggested that stathmin-like proteins such as SCG10, which is co-expressed in neurons and displays similar patterns of expression to stathmin during development of the nervous system (Grenningloh, Soehrman et al. 2004), may be sufficient to maintain neuronal cell microtubule dynamics in stathmin KO mice. Conversely, aging stathmin KO mice

develop axonopathy of the central and peripheral nervous systems (Liedtke, Leman et al. 2002), display defects in learned and innate fear processing, and innate parental behaviours and social interactions (Shumyatsky, Malleret et al. 2005; Martel, Nishi et al. 2008). Thus stathmin appears to be vital for regulating neuronal cell functions later in life. In support of this are studies showing direct correlations between loss of stathmin expression and neurofibrillary tangle formation in Alzheimer’s disease (Jin,

Masliah et al. 1996; Cheon, Fountoulakis et al. 2001) and in the brains of adult Down syndrome patients (Cheon, Fountoulakis et al. 2001).

1.3.8 Cell Migration

Stathmin is important for the migration of Drosophila germ cells (Ozon, Guichet et al.

2002), MEFs (Baldassarre, Belletti et al. 2005), human endothelial cells (Mistry, Bank et al. 2007), trophoblasts (Yoshie, Kashima et al. 2008), various cancer cells including glioma (Liang, Choi et al. 2008), NSCLC (Singer, Malz et al. 2009), hepatocellular carcinoma (HCC) (Gan, Guo et al. 2010), gastric cancer (Jeon, Han et al. 2010) and sarcoma cells (Baldassarre, Belletti et al. 2005) in vitro and rat neuronal cells in vitro and in vivo (Jin, Mao et al. 2004; Giampietro, Luzzati et al. 2005). However, the mechanism by which stathmin regulates cell migration is potentially multifaceted.

1.3.9 Interphase Microtubule Network

Wittmann et al. report that only a small fraction of stathmin is phosphorylated upon growth factor-stimulated cell migration (Wittmann, Bokoch et al. 2004) and fluorescence resonance energy transfer analyses found that stathmin’s tubulin-

sequestering activity is inactivated at the leading edge of migrating Xenopus cells

(Niethammer, Bastiaens et al. 2004). It has therefore been speculated that differential stathmin phosphorylation gradients exist within motile cells to regulate local microtubule dynamics at the cell periphery. Furthermore, stathmin assists cell movement by promoting ‘pioneer’ microtubule growth at the leading edge of motile cells (Wittmann and Waterman-Storer 2001; Niethammer, Bastiaens et al. 2004).

However, the signalling pathways that regulate stathmin activity and microtubule dynamics in motile cells appear complex.

1.3.10 RhoGTPase Signalling Pathways

Activation of the RhoGTPase signalling pathways are critical for cell migration and stathmin has been linked to these pathways in various cell types. Activation of Rac1 induces Ser16 phosphorylation in T-cells and neuronal cells (Watabe-Uchida, John et al.

2006; Tanaka, Hamano et al. 2007) and Ser25/Ser38 in breast cancer cells (Takahashi and Suzuki 2009). One kinase responsible for phosphorylating stathmin downstream of

Rac1 activation is Pak1 (Daub, Gevaert et al. 2001; Wittmann, Bokoch et al. 2004).

Interestingly, Rac1-induced phosphorylation of stathmin in breast cancer cells promoted direct interactions between stathmin and the motor protein kinesin (in a Pak1-WAVE2- kinesin complex) (Takahashi and Suzuki 2009) and the microtubule end-binding protein, EB1 (Takahashi, Tanaka et al. 2010). Moreover, inhibition of stathmin expression disrupted these interactions reducing microtubule-mediated transport of

WAVE2 to the leading edge, lamellipodia formation and cell migration in breast cancer cells (Takahashi and Suzuki 2009). Thus the regulation of stathmin activity downstream of Rac1 may be important for cell migration in various cell types.

Stathmin has also been linked to RhoA signalling through its interactions with p27kip1, a member of the Cip/Kip family of cyclin-dependent kinase inhibitors. Baldassarre et al. discovered that the C-terminal region of p27kip1 directly binds to stathmin in vitro and in vivo inhibiting stathmin’s tubulin-sequestering ability and maintaining a mesenchymal- type mode of migration in sarcoma cells (Baldassarre, Belletti et al. 2005). Furthermore, the phosphorylation of p27kip1 (at threonine 198) may dictate interactions with, and the activity of stathmin, which subsequently influences RhoA activity, microtubule lipid raft trafficking and migration in MEFs (Belletti, Pellizzari et al. 2010; Schiappacassi,

Lovisa et al. 2011). Collectively, these studies suggest p27kip1-stathmin interactions may regulate the actin network, via RhoA, and specific modes of cell migration.

1.3.11 Interacting Proteins

A number of other proteins have been identified that directly interact with and influence stathmin’s role in cell migration. Kinase-interacting with stathmin (KIS)-mediated phosphorylation of stathmin at Ser38 has been implicated in the migration of vascular smooth muscle cells (VSMCs) in vitro and in vivo (Langenickel, Olive et al. 2008).

Langenickel et al. found that KIS KO mice express high levels of stathmin in a more active state compared to wild-type mice which promotes the migration of VSMCs during wound repair (Langenickel, Olive et al. 2008). Further investigation of KIS KO

VSMCs found alterations in microtubule stability and filamentous actin distribution at the cell periphery compared to wild-type cells (Langenickel, Olive et al. 2008). Thus it appears that KIS is important for regulating stathmin activity and cytoskeletal- restructuring during VSMC migration, though this interaction has yet to be confirmed in other cell types.

Recent studies have identified a novel interaction between stathmin and the most pleiotropic member of the signal transducer and activator of transcription (STAT) family, STAT3 (Ng, Lin et al. 2006; Verma, Dourlat et al. 2009). While STAT3 predominantly exerts its function in the nucleus, new evidence reveals distinct roles for

STAT3 in the cytoplasm (Jing and Tweardy 2005). STAT3 binds to stathmin’s C- terminal domain, inhibiting microtubule depolymerisation and promotes the migration of MEFs (Ng, Lin et al. 2006) and T-cells (Verma, Dourlat et al. 2009). Interestingly,

STAT3 also binds to SCLIP, maintaining SCLIP stability and conferring a less invasive, epithelial morphology in breast cancer cells (Ng, Lim et al. 2010). Therefore, cytoplasmic STAT3 is important for regulating microtubule dynamics in motile cells by binding to and inhibiting the microtubule-depolymerising activity of multiple members of the stathmin-like protein family.

1.4 Stathmin in Cancer

Stathmin is over-expressed in numerous malignancies including leukaemia (Hanash,

Strahler et al. 1988; Brattsand, Roos et al. 1993), breast cancer (Curmi, Nogues et al.

2000; Saal, Johansson et al. 2007), gastric cancer (Jeon, Han et al. 2010), pancreatic cancer (Iacobuzio-Donahue, Maitra et al. 2002), endometrial carcinoma (Trovik, Wik et al. 2011), HCC (Yuan, Jeng et al. 2006; Singer, Ehemann et al. 2007), NSCLC (Singer,

Malz et al. 2009), prostate cancer (Friedrich, Gronberg et al. 1995), oral squamous cell carcinoma (Kouzu, Uzawa et al. 2006), cervical carcinoma (Xi, Rui et al. 2009), medulloblastoma (Kuo, Wang et al. 2009) and retinoblastoma (Mallikarjuna, Sundaram et al. 2010). In most cases, stathmin over-expression correlates with poorly differentiated, highly metastatic and drug-resistant tumours and serves as a strong

prognostic indicator of tumour grade, recurrence and survival (Saal, Johansson et al.

2007; Kuo, Wang et al. 2009; Xi, Rui et al. 2009; Hsieh, Huang et al. 2010; Jeon, Han et al. 2010; Trovik, Wik et al. 2011). Although the direct cause is unknown, it is likely that multiple defects at the transcriptional and post-transcriptional level contribute to stathmin over-expression in cancer.

1.4.1 Transcription Factor Deregulation

Sequencing of the stathmin promoter identified consensus sites for a range of proliferation-related transcription factors. The stathmin gene promoter contains fifteen

GC boxes, four E2F, two AP-2 transcription factor binding sites and a number of

CCAAT boxes (Melhem, Zhu et al. 1991). The transcription factor NF-Y is shown to predominantly bind to two CCAAT sites within the stathmin promoter (Benlhabib and

Herrera 2006). One of these sites, located 65 bases upstream of the transcriptional start site, is predicted to account for more than 60% of stathmin’s transcriptional gene activity in normal proliferating cells (Benlhabib and Herrera 2006). The E2F transcription factor may also be responsible for stathmin over-expression in some cancer cells. Indeed, Polzin et al. have shown that all four E2F binding sites account for

80% of stathmin expression in a prostate cancer cell line (Polzin, Benlhabib et al. 2004).

Moreover, it has been proposed that c-Jun, a transcriptional activator frequently over- expressed in cancer, may mediate stathmin regulation indirectly by increasing E2F activity or via direct interactions with the stathmin promoter (Kinoshita, Leaner et al.

2003). In opposition to their role as transcriptional activators, E2Fs can be converted to transcriptional repressors via binding to retinoblastoma (Rb) proteins. Studies conducted by Polager et al. demonstrate that an Rb/E2F complex, p130/E2F-4, directly binds to

the stathmin promoter repressing transcription in vivo (Polager and Ginsberg 2003). As such, disruptions to Rb/E2F binding may also contribute to stathmin over-expression in cancer. Chen et al. report that the histone-lysine N-methyltransferase gene enhancer of zeste homolog 2 (EZH2), an enzyme frequently over-expressed in aggressive malignancies, may contribute to stathmin over-expression by interfering with Rb/E2F complex formation (Chen, Lin et al. 2007). EZH2 competes with histone deacetylase

(HDAC) in binding the Rb protein p130, which may then inhibit Rb/E2F interactions and therefore Rb/E2F-mediated transcriptional repression of stathmin expression

(Harbour and Dean 2000).

More recently the role of the transforming-growth factor-beta (TGF-β) inducible early gene 1 (TIEG1) (Jiang, Chen et al. 2009) and forkhead box M1b (FoxM1b) (Park,

Gusarova et al. 2011) transcription factors have been implicated in promoting stathmin expression in cancer. The stathmin promoter contains multiple Sp1-binding sites that are targeted by Krüppel-like transcription factors, including TIEG1, which is frequently down-regulated in invasive cancers (Jiang, Chen et al. 2009). Lentiviral-induced TIEG1 expression in TGF-β-resistant pancreatic cell lines reduced stathmin expression, inhibited cell proliferation, increased apoptosis and enhanced sensitivity to the deoxycytidine analogue, gemcitabine (Jiang, Chen et al. 2009). Moreover, stathmin over-expression prevented TIEG1-induced growth arrest highlighting a direct link between TIEG1 and stathmin expression in pancreatic cancer (Jiang, Chen et al. 2009).

FoxM1b is a proliferation-related transcription factor over-expressed in numerous malignancies including breast and pancreatic cancer, glioblastoma and hepatocellular carcinoma (Carr, Park et al. 2010; Park, Gusarova et al. 2011). Recent studies have

demonstrated that FoxM1b binds directly to the stathmin promoter to stimulate expression (Carr, Park et al. 2010; Park, Gusarova et al. 2011). Moreover, over- expression of FoxM1 in human epidermal growth factor receptor 2 (HER2)-positive breast cancer cells increased stathmin expression, inhibited paclitaxel-induced tubulin polymerisation and increased resistance to Herceptin® and paclitaxel (Carr, Park et al.

2010). This study clearly demonstrated the potential impact FoxM1 has on stathmin expression and drug-resistance in breast cancer cells. However, the role FoxM1 plays in regulating stathmin expression is not restricted to breast cancer. Liver tumours derived from FoxM1b (transcript variant of FoxM1) transgenic mice that were negative for the

FoxM1 inhibitor Arf, expressed higher levels of stathmin compared to Arf-/- tumours

(Carr, Park et al. 2010). Furthermore, inhibition of stathmin expression in FoxM1b- expressing Arf-/- hepatocellular carcinoma cells inhibited lung metastasis in vivo (Park,

Gusarova et al. 2011). Thus FoxM1 may be a major contributor to stathmin over- expression, drug-resistance and metastasis in numerous malignancies.

1.4.2 Stathmin & p53

Various studies have demonstrated that activation of the tumour-repressor gene p53 represses stathmin expression in both murine and human transformed and immortalised cell lines (Ahn, Murphy et al. 1999; Johnsen, Aurelio et al. 2000). It is therefore believed that p53 mutations, frequently found in cancer, may contribute to stathmin over-expression in some malignancies (Yuan, Jeng et al. 2006). The link between p53 activation and reduced stathmin expression is reportedly mediated via an indirect mechanism as the stathmin promoter lacks the p53-binding site (TATA box) (Melhem,

Zhu et al. 1991). Transcriptional repressors such as Rb/E2F (Polager and Ginsberg

2003) and Egr1 are proposed to regulate stathmin expression downstream of p53 activation. Indeed, Fang et al. demonstrated that Egr1 directly binds to the stathmin promoter following p53 activation in lung cancer cells (Fang, Min et al. 2009). These observations have lead researchers to speculate that reducing stathmin expression may be a therapeutic strategy to restore some wild-type p53 functions in mutant p53 cells

(Alli, Yang et al. 2007). Moreover, depleting stathmin expression in a colon cancer cell line and HeLa cells, both lacking p53, induced cell death and delayed G2-M cell cycle progression (Carney and Cassimeris 2010). Conversely, over-expression of stathmin reversed p53-induced G2-M cell cycle arrest, suggesting that stathmin is a key target of p53-induced cell cycle regulation (Johnsen, Aurelio et al. 2000).

1.4.3 Post-transcriptional Deregulation

A number of studies have indicated that stathmin is deregulated at the post- transcriptional level. Singer et al. revealed that over-expression of the RNA-modifying protein, far upstream sequence element-binding protein-I (FBP-1), directly correlates with stathmin and SCLIP expression in NSCLC and that genetically altering FBP-1 levels altered stathmin and SCLIP expression in NSCLC cell lines (Singer, Malz et al.

2009). Therefore FBP-1 co-ordinates the expression of multiple microtubule- destabilising proteins in NSCLC and may be important for regulating microtubule dynamics in cancer cells over-expressing FBP-1. Other studies have shown regulation of stathmin expression by the small non-coding RNAs, microRNAs (miRNAs). Wong et al. report that loss of miRNA-223, which directly targets and represses stathmin mRNA expression, correlates with stathmin over-expression in HCC (Wong, Lung et al.

2008). More recently, Delaloy et al. propose an essential role for the brain-specific miR-

9 in targeting stathmin for the promotion of proliferation and reduction in migration of multipotent human neural progenitor cells (Delaloy, Liu et al. 2010), though whether miR-9 contributes to stathmin deregulation in cancer is unknown.

1.4.4 Oncogenic Signalling Pathways

The activation of common oncogenic signalling pathways, including phosphatidylinositol 3-kinase (PI3K) and Hedgehog (Hh) correlate with stathmin over- expression in some malignancies. Amplification or over-expression of the PI3K oncogene, PI3KCA, or loss of phosphatase and tensin homolog (PTEN), a known inhibitor of the PI3K pathway, are common lesions in PI3K-activated tumours. Recent investigations have demonstrated that stathmin over-expression strongly correlates with

PI3K activation in breast and endometrial cancers and is therefore considered a robust marker of this signature (Saal, Johansson et al. 2007; Salvesen, Carter et al. 2009;

Trovik, Wik et al. 2010). Stathmin is over-expressed in PI3KCA-amplified and overexpressed endometrial cancers as well as PTEN-negative breast and endometrial tumours (Saal, Johansson et al. 2007; Salvesen, Carter et al. 2009). Furthermore, endogenous expression of PTEN in PTEN-negative breast cancer cell lines, xenografts and glioma cells, dramatically reduced stathmin expression (Stolarov, Chang et al.

2001; Saal, Johansson et al. 2007). This phenotype was also observed upon PI3K- inhibitor treatment of PTEN-negative glioma cells in vitro and breast tumours in vivo

(Saal, Johansson et al. 2007). These studies clearly link stathmin over-expression to activation of the PI3K pathway. In another study, Yoshie et al. reported that stathmin inhibition blocked hypoxia-induced Akt activation, a key downstream signalling protein of the PI3K pathway, in endometrial and endothelial cells (Yoshie, Miyajima et al.

2009). However, further studies are required to confirm stathmin’s role in regulating the

PI3K pathway.

The Hh signalling pathway is critical during embryonic development for the regulation of cell proliferation, polarity and differentiation and is aberrantly activated in a variety of malignancies. In primary prostate tumours, stathmin expression correlates with and localises with the Hh ligand, Sonic hedgehog (Shh), and ligand receptor, Patched

(Chung, Kim et al. 2010). Treatment with the Hh signalling inhibitor, cyclopamine, decreased stathmin expression and cell proliferation while induction of the pathway, via treatment with a Shh peptide or over-expression of the glioma-associated oncogene homolog (Gli), increased stathmin expression in a prostate cancer cell line (Chung, Kim et al. 2010). Therefore, stathmin expression is linked to cell proliferation downstream of

Hh-activation in some tumours, though the direct link between Hh signalling and stathmin expression is unclear.

1.4.5 Loss of Heterozygosity

The stathmin gene is located on chromosome 1, band p35-36.1 (Ferrari, Seuanez et al.

1990), a site of frequent deletions or LOH in malignancies such as neuroblastoma, glioma and breast cancer. Although no link has been identified between high stathmin expression and LOH in breast cancer (Curmi, Nogues et al. 2000), low stathmin expression correlates with 1p LOH and recurrence-free survival in malignant gliomas

(Ngo, Peng et al. 2007). In addition, stathmin expression is reduced in some 1p LOH compared to 1p-normal neuroblastoma tumours and cell lines (Janoueix-Lerosey,

Novikov et al. 2004). However, in most cancers, stathmin over-expression and/or increased activity contribute to an aggressive cancer phenotype.

1.4.6 Altered Activity

In addition to high expression, stathmin is found in a more ‘active’ (unphosphorylated) state in some cancer cells. Stathmin is more ‘active’ in neoplastic and recurrent sarcoma tumours compared to normal tissue (Belletti, Nicoloso et al. 2008) and stathmin activity increases with MYCN copy number in primary human neuroblastoma tumours and cell lines (Hailat, Strahler et al. 1990). Furthermore, alterations in stathmin phosphorylation have been identified in in vivo-derived vincristine-resistant leukaemia xenografts

(Verrills, Liem et al. 2006) and a somatic mutation (Q18>E), identified in a human oesophageal adenocarcinoma tumour, increases stathmin activity by impairing the phosphorylation of Ser16 (Misek, Chang et al. 2002). Forced expression of this Ser16 mutation induced NIH3T3 cell transformation (Misek, Chang et al. 2002), increased aneugenic activity in leukaemia cells (Holmfeldt, Brannstrom et al. 2006) and enhanced fibrosarcoma cell migration and invasion (Belletti, Nicoloso et al. 2008). However, mutations in the stathmin gene are uncommon which suggests that other mechanisms exist to perturb stathmin activity in cancer. One proposed mechanism is via hypoxia

(oxygen deprivation); a common phenomenon in advanced tumours that promotes drug- resistance and metastasis. Recent studies report altered activity of stathmin under hypoxic conditions (1% O2) in cardiomyocytes and HeLa cells (Hu, Chu et al. 2010).

Hypoxia induced the dephosphorylation of stathmin at Ser16 and lead to destabilisation of the microtubule network (Hu, Chu et al. 2010). This implicates stathmin in mediating hypoxia-induced alterations to the cell cytoskeleton. Indeed in human endometrial

stromal cells, inhibition of stathmin expression induced microtubule stabilisation and was associated with reduced hypoxia-inducible factor (HIF) 1α and vascular epidermal growth factor (VEGF) expression (Yoshie, Miyajima et al. 2009). These studies suggest that regulation of stathmin expression and activity may be altered by the induction of hypoxia.

1.4.7 Tumour Progression

In many malignancies, stathmin over-expression strongly correlates with poorly differentiated (Friedrich, Gronberg et al. 1995; Chen, Wang et al. 2003; Jeon, Han et al.

2010) and highly proliferative tumours (Curmi, Nogues et al. 2000; Sadow, Rumilla et al. 2008; Salvesen, Carter et al. 2009; Trovik, Wik et al. 2010). When induced to differentiate, cancer cells cease to proliferate, which in many cases, coincides with a reduction in stathmin expression (Johnson, Jones et al. 1995). This suggests that high stathmin levels promote cell proliferation in some cancers. Indeed, inhibition of stathmin expression is shown to markedly reduce proliferation of a variety of non- neuronal cancer cell lines (Mistry, Bank et al. 2005; Alli, Yang et al. 2007; Singer,

Ehemann et al. 2007; Wang, Dong et al. 2007; Singer, Malz et al. 2009; Chung, Kim et al. 2010; Hsieh, Huang et al. 2010; Jeon, Han et al. 2010; Mitra, Kandalam et al. 2011).

Furthermore, inhibiting stathmin expression reduced the tumourigenicity of leukaemia cells in vitro and osteosarcoma and gastric cancer cells in vivo (Jeha, Luo et al. 1996;

Wang, Dong et al. 2007; Jeon, Han et al. 2010). Another way in which stathmin may influence cancer progression is by promoting chromosomal instability. Indeed, high stathmin expression and/or increased stathmin activity directly correlates with aneuploidy in various tumours (Holmfeldt, Brannstrom et al. 2006; Hsieh, Huang et al.

2010; Trovik, Wik et al. 2010). It has also been proposed that stathmin may be playing a role in cell survival as inhibition of stathmin expression increases apoptosis in a variety of cancer cells lines in vitro and in vivo (Alli, Yang et al. 2007; Wang, Dong et al. 2007;

Zhang, Xiong et al. 2008; Jeon, Han et al. 2010; Mitra, Kandalam et al. 2011). Although the mechanism underlying stathmin’s role in cell survival is unclear, it has been speculated that stathmin may influence microtubule-mediated translocation and regulation of apoptotic B-cell lymphoma (BCL) family members (Longuet, Serduc et al.

2004; Singer, Ehemann et al. 2007; Rana, Maples et al. 2008). Therefore a combination of high stathmin expression and activity may be contributing to cancer progression by promoting cell growth, survival and chromosomal instability.

1.4.8 Drug Resistance

Alterations in stathmin expression and activity are closely associated with drug- resistance in some malignancies. Stathmin expression is significantly reduced in gemcitabine-resistant pancreatic cancer cells (Kuramitsu, Taba et al. 2010) and is over- expressed in paclitaxel-resistant lung cancer cells (Martello, Verdier-Pinard et al. 2003), vincristine-resistant neuroblastoma cells (Don, Verrills et al. 2004), cisplatin-resistant head and neck cancer cells (Johnsson, Zeelenberg et al. 2000) and drug-resistant leukaemia patient samples (Roos, Brattsand et al. 1993). In addition, alterations in stathmin phosphoisoforms have been identified in vincristine-resistant leukaemia xenografts (Verrills, Liem et al. 2006). In a clinical setting, stathmin over-expression confers an unfavourable prognosis for ovarian cancer patients treated with paclitaxel and platinum (Su, Smith et al. 2009) and in many other malignancies is associated with paclitaxel resistance (Orr, Verdier-Pinard et al. 2003; Rana, Maples et al. 2008). This

suggests that stathmin expression may be mediating drug sensitivity and be predictive of chemotherapy response in some malignancies (Song, Choi et al. 2006). Indeed, targeting stathmin has proved effective in modifying the sensitivity of a range of cancer and endothelial cells to chemotherapeutic agents (Table 1.1). In particular, inhibition of stathmin expression enhances sensitivity to paclitaxel as observed in breast cancer (Alli,

Yang et al. 2007), leukaemia (Iancu, Mistry et al. 2000), HCC (Singer, Ehemann et al.

2007), prostate cancer (Mistry and Atweh 2006) and retinoblastoma cells in vitro

(Mitra, Kandalam et al. 2011) and osteosarcoma cells in vivo (Wang, Dong et al. 2007)

(Table 1.1). In addition, reducing stathmin expression increases sensitivity to DNA- targeted agents such as cisplatin in HCC cell lines (Singer, Ehemann et al. 2007)

(Table 1.1). However, inhibition of stathmin expression did not appear to alter the sensitivity of two ovarian cancer cell lines to paclitaxel or cisplatin (Aoki, Oda et al.

2009) (Table 1.1). Conversely, stathmin over-expression increased resistance to paclitaxel and vinblastine in breast cancer cells (Alli, Bash-Babula et al. 2002;

Balasubramani, Nakao et al. 2010) and gambogic acid and gambogenic acid in a HCC cell line (Wang, Chen et al. 2009) (Table 1.1). It therefore appears that inhibition of stathmin expression is a plausible mechanism by which to enhance the sensitivity of some cancer cells to chemotherapeutic agents. However, the exact mechanisms underlying stathmin-mediated alterations in drug sensitivity remain unclear.

It has been widely reported that stathmin over-expression in cancer cells is associated with increased resistance to paclitaxel (Orr, Verdier-Pinard et al. 2003; Rana, Maples et al. 2008). In a simplistic view, it is thought that over-expression of stathmin may enhance microtubule destabilisation, an effect that would potentially thwart the microtubule-stabilising actions of paclitaxel (Balasubramani, Nakao et al. 2010).

Table 1.1 Altering stathmin expression modifies cell sensitivity to chemotherapy & irradiation

Origin Cell Line/s Silencing Over-expression Drugs/Irradiation Response Ref. TXL ↑ (syn.) CML K562 Antisense RNA - 5-FU, doxo. ↑ (add.) (Iancu, Mistry et al. 2000) VLB, nocodazole No affect VCR, vindesine ↑ SCLC SBC-3 - cDNA (Nishio, Nakamura et al. 2001) TXL, docetaxel No affect NSCLC Calu-1, Calu-6 siRNA - Irradiation (6/8Gy) ↑ (Singer, Malz et al. 2009) TXL, VLB ↓ Breast cancer BT20 - cDNA (Alli, Bash-Babula et al. 2002) doxo., campto. No affect TXL, VLB ↑ Breast cancer BT549 siRNA - (Alli, Bash-Babula et al. 2002) doxo., CDDP, VP-16 No affect HCC Hep3B, HUH-7 siRNA - TXL, VLB, CDDP ↑ (Singer, Ehemann et al. 2007) HCC Hep2G siRNA - GBA/GBGA ↑ (Wang, Chen et al. 2009) HCC Hep2G - cDNA GBA/GBGA ↓ (Wang, Chen et al. 2009) SKOV-3, Ovarian cancer siRNA - TXL, CDDP No affect (Aoki, Oda et al. 2009) OVCAR-3 Table 1.1 continued on following page.

Table 1.1 (continued) Altering stathmin expression modifies cell sensitivity to chemotherapy & irradiation

Over- Origin Cell Line/s Silencing Drugs/Irradiation Response Ref. expression CCNU ↑ (Ngo, Peng et al. Glioma U251 shRNA - VCR, procarbazine, TMZ No affect 2007) (Ngo, Peng et al. Glioma A172 - cDNA CCNU ↓ 2007) Saos-2, (Wang, Dong et al. Osteosarcoma siRNA - TXL, docetaxel ↑ MG63 2007) TXL, VP-16 ↑ (syn.) (Mistry and Atweh Prostate cancer LNCaP Anti-stathmin ribozyme - 5-FU, doxorubicin ↑ (add.) 2006) (Mitra, Kandalam et Retinoblastoma Y79 siRNA - TXL, VCR ↑ al. 2011) (Mistry, Bank et al. Endothelial HUVEC Anti-stathmin ribozyme - TXL ↑ 2007) campto. (camptothecin), CCNU [1-(2-chloroethyl)-3-cyclohexyl-l-nitrosourea], CDDP (cisplatin), CML (chronic myelogenous leukaemia), doxo.

(doxorubicin), GBA (gambogic acid), GBGA (gambogenic acid), HCC (hepatocellular carcinoma), HUVEC (human umbilical vein endothelial cells), NSCLC (non-small cell lung cancer), SCLC (small-cell lung cancer), shRNA (short hairpin RNA), siRNA (small interfering RNA), TMZ

(temozolomide), TXL (paclitaxel), VCR (vincristine), VLB (vinblastine), VP-16 (etoposide), 5-FU (5’fluorouracil), syn. (synergistic), add.

(additive), ↑(increased sensitivity), ↓ (decreased sensitivity).

However, mounting evidence suggests that stathmin’s influence on cancer cells’ response to paclitaxel is far more complicated (Balasubramani, Nakao et al. 2010).

Early studies by Iancu et al. led to speculation that stathmin inhibition combined with anti-mitotic drug treatment may increase cell sensitivity due to disruption of distinct steps in the same (mitotic) pathway (Iancu, Mistry et al. 2000). As such, in many cells where stathmin inhibition increases sensitivity to paclitaxel, stathmin inhibition alone disrupts mitotic progression (Iancu, Mistry et al. 2000; Singer, Ehemann et al. 2007;

Wang, Dong et al. 2007; Mitra, Kandalam et al. 2011). Recent investigations have demonstrated that stathmin itself can mediate drug-binding and is a target for chemotherapeutic agents. Devred et al. report that stathmin increases the affinity of vinblastine for tubulin and vice versa in vitro, suggesting that cells with high stathmin expression may be more sensitive to this agent (Devred, Tsvetkov et al. 2008).

However, in breast cancer cells stathmin over-expression reduced sensitivity to vinblastine (Alli, Bash-Babula et al. 2002) and silencing stathmin increased HCC and breast cancer cell sensitivity vinblastine (Alli, Bash-Babula et al. 2002; Singer,

Ehemann et al. 2007) (Table 1.1). Additionally, Ngo et al. also discovered that the

DNA-alkylating agent CCNU, commonly used for the treatment of brain tumours, can covalently bind and promote carbamoylation of stathmin’s lysine residues subsequently inhibiting its microtubule depolymerizing activity (Ngo, Peng et al. 2007). Collectively, stathmin may influence cancer cell drug sensitivity by multiple mechanisms.

1.4.9 Metastasis

Stathmin over-expression correlates with highly metastatic disease in various non- neuronal cancers (Saal, Johansson et al. 2007; Belletti, Nicoloso et al. 2008; Blower,

66 Chung et al. 2008; Xi, Rui et al. 2009; Hsieh, Huang et al. 2010; Jeon, Han et al. 2010;

Kuramitsu, Taba et al. 2010; Trovik, Wik et al. 2011) and adult neuroendocrine tumours

(Sadow, Rumilla et al. 2008), and strong stathmin immunohistochemical staining in preoperative curettage samples predicts lymph node metastasis and poor survival in endometrial carcinomas (Trovik, Wik et al. 2011). Stathmin is abundant in vascular invasive HCC (Yuan, Jeng et al. 2006; Hsieh, Huang et al. 2010), endometrial carcinomas (Salvesen, Carter et al. 2009) and gastric cancers (Jeon, Han et al. 2010) and is localised at the invasive front of NSCLC (Singer, Malz et al. 2009) suggesting that stathmin plays a role in cell invasion in vivo. Interestingly, Belletti et al. demonstrated a functional role for stathmin in regulating 3 dimensional (3D) cell motility and invasion of sarcoma cells in vitro and in vivo (Belletti, Nicoloso et al. 2008). Both stathmin over- expression and increased stathmin activity, via expression of the Ser16 phosphorylation impaired Q18>E mutation, enhanced sarcoma cell invasion in vitro and metastatic potential in vivo (Belletti, Nicoloso et al. 2008). Further analyses demonstrated that this increase in stathmin activity decreased ECM contact-mediated stathmin phosphorylation, microtubule stabilisation and induced amoeboid-like invasion of sarcoma cells in vitro (Belletti, Nicoloso et al. 2008). Thus, high stathmin expression and activity may influence the mode in which some cancer cells invade and metastasize.

1.4.10 Stathmin in Neuroblastoma

Using 2 dimensional (2D) gel electrophoresis, Hailat et al. revealed that stathmin is highly expressed in some neuroblastoma tumours and that stathmin phosphorylation isoforms decreased in MYCN-amplified primary neuroblastoma tumours and cell lines compared to the non-amplified counterparts (Hailat, Strahler et al. 1990). Moreover,

67 MYCN-amplified and non-amplified cells responded differently to retinoic acid treatment in regards to stathmin phosphorylation (Hailat, Strahler et al. 1990). In adrenal tissues taken from mice orthotopically injected with neuroblastoma cells, stathmin was found to be >15 times more abundant, and its phosphorylation pattern altered, in tumour-containing adrenal glands compared to normal adrenal glands

(Campostrini, Pascali et al. 2004). Moreover, stathmin is over-expressed in malignant and metastatic phaeochromocytomas (neural crest-derived tumours of the adrenal medulla) compared to benign tumours and normal adrenal tissue (Sadow, Rumilla et al.

2008; Lin, Chen et al. 2011). It has also been reported that stathmin gene expression may be altered in neuroblastoma due to its genetic location on chromosome 1p36, a site frequently deleted in neuroblastoma strongly associated with MYCN amplification and advanced stage disease (Brodeur 2003). Janoueix-Lerosey et al. found that stathmin gene expression is reduced by 1.74 fold in 1p LOH neuroblastoma cell lines and primary tumours compared to those without 1p LOH (Janoueix-Lerosey, Novikov et al.

2004). Interestingly, Don et al. found that stathmin expression was increased in vincristine-resistant neuroblastoma cells compared to parental cells (Don, Verrills et al.

2004). Collectively, these studies indicate that altered stathmin expression and/or activity are linked to a number of unfavourable prognostic factors in neuroblastoma and other neuroendocrine tumours. However stathmin’s functional role in neuroblastoma biology remains to be determined.

1.5 Thesis Aims

In a number of epithelial cancers, stathmin is over-expressed and/or its activity altered compared to normal differentiated tissue. Suppressing stathmin expression in these

68 cancer cells can reduce proliferation, tumour growth, migration and invasion and increase drug sensitivity. All these factors have identified stathmin as a promising anticancer target and as such, a number of studies have investigated potential strategies by which to target stathmin in cancer cells.

Neuroblastoma is one of the most aggressive and deadly childhood cancers. Three key phenotypes that contribute to its aggressive nature are; i) drug resistance, ii) poor differentiation and iii) metastasis. Therefore in order to improve treatments and subsequently survival outcomes for this disease, a better understanding of these phenotypes are required.

In neuroblastoma tumours and cell lines, alterations in stathmin expression and/or activity are associated with unfavourable prognostic factors. However no study to date has addressed stathmin’s functional role in this cancer. Based on these findings, this study was designed to investigate stathmin’s contribution to drug-resistance, differentiation and metastasis in neuroblastoma. The clinical opportunity for this study is that by evaluating stathmin’s functional role in this cancer, it may identify stathmin as a potential therapeutic target for neuroblastoma. As such, any anti-stathmin therapies developed for other malignancies may also be efficacious for the treatment of neuroblastoma.

69 Chapter 2. Materials & Methods

70 2.1 Cell culture

Neuroblastoma tumours arise from multi-potential cells of the neural crest. These tumours usually contain a mixture of cell types including neuroblastic ‘N’ type, schwannian ‘S’ type (substrate-adherent) and intermediate ‘I’ type cells (Ross, Biedler et al. 2003). ‘I’ type cells express both ‘N’ and ‘S’ type marker proteins and their morphology is a mixture of these two cell types (Ross, Biedler et al. 2003). A panel of neuroblastoma cell lines, containing all these cell types, were screened for stathmin protein expression in initial experiments, as shown in Table 2.1.

Table 2.1 Neuroblastoma Cell Types

Parental Cell Line Subline/s Type Reference SKNBE(2) - I (Biedler, Roffler-Tarlov et BE(2)-C I al. 1978) SK-N-SH - I (Biedler, Roffler-Tarlov et SH-SY5Y N al. 1978) SH-EP S NBL-W - I (Foley, Cohn et al. 1991) NBL-S S NBL-WR - ? (Foley, Cohn et al. 1991) (Tumilowicz, Nichols et al. IMR-32 - I 1970)

71 The human neuroblastoma cell line SK-N-BE(2) was a generous gift from Dr. Sylvain

Baruchel (The Hospital for Sick Children, Toronto). The SK-N-BE(2) neuronal clone,

BE(2)-C, and SK-N-SH cell line and sublines, SHEP and SH-SY5Y, were obtained from Dr. June Biedler, Fordham University, New York. NBL-S, NBL-W and NBL-WR cell lines were kindly provided by Dr S. Cohn (Northwestern University, Chicago, IL).

The NBL-W and NBL-WR cells were derived from a tumour from the same patient pre- and post-relapse, respectively (Foley, Cohn et al. 1991). IMR-32 and Calu-6 (lung cancer) cell lines were obtained from the American Type Culture Collection (ATCC).

The majority of experiments conducted in this thesis utilised SK-N-BE(2), BE(2)-C and

SH-SY5Y neuroblastoma cells that were all originally derived from bone marrow biopsies of relapsed patients with advanced (stage 4) disease, their clonal derivation has been previously described (Biedler, Roffler-Tarlov et al. 1978). The SK-N-BE(2),

BE(2)-C, SH-SY5Y and Calu-6 cell lines were all validated by short tandem repeat

(STR) profiling (CellBank Australia, Westmead, New South Wales). All cells were maintained as monolayers grown in 10% foetal calf serum (FCS)/Dulbecco’s Modified

Eagle Media (DMEM) (Gibco-Invitrogen, Carlsbad, CA, USA) (neuroblastoma cells) or

10% FCS/Roswell Park Memorial Institute (RPMI) media (lung cancer cells) at 37°C in a humidified atmosphere with 5% CO2. Cells were passaged 2-3 times per week using phosphate buffered saline (PBS)/Trypsin. All cells were routinely screened and found to be free of mycoplasma.

72 2.2 siRNA-mediated stathmin suppression

RNA interference (RNAi) is an evolutionary-conserved (small RNA-mediated), post- transcriptional gene silencing mechanism found in most eukaryotes. Small interfering

RNA (siRNA) are synthetic mimics of these naturally occurring RNAs, usually 20-25 base pairs long that are used extensively in gene function studies. For human stathmin

(STMN1) suppression studies, neuroblastoma cells were transfected with STMN1 siRNA ON-TARGETplus SMARTpool reagent or ON-TARGETplus Non-targeting

Pool (control) reagent (ThermoFisher Scientific, Lafayette, CO). The ON-TARGETplus siRNA reagents are dual-modified to reduce potential off-target effects. The first modification is to the sense strand that prevents interaction with the RNA-induced silencing complex (RISC) and favours antisense strand uptake. The second modification is to the antisense strand seed region to minimise seed-related off-target effects

(Dharmacon RNAi Technologies, ThermoFisher Scientific). The AllStars Neg. control siRNA (Qiagen, Valencia, CA, USA) was also used for initial siRNA optimisations. All siRNAs were reconstituted in 1x siRNA buffer (20mmol/L KCl, 6mmol/L HEPES pH7.5, 0.2mmol/L MgCl2) (ThermoFisher Scientific) to a working concentration of

20μmol/L. The STMN1 SMARTpool reagent contains four individual siRNA sequences that target STMN1, transcript variant 2 (NM_203399) (see Table 2.2 for siRNA sequences). These four sequences also target the three other STMN1 transcript variants

(NM_005563, NM_203401 and NM_001145454) as they are homologous in these regions. The ON-TARGETplus Non-targeting Pool has been confirmed by microarray to have minimal targeting of known genes in human cells.

73 Table 2.2 Stathmin siRNA and shRNA Sequences

Target gene/Accession # siRNA Seq. siRNA Sequence Gene location

1 GAAAGACGCAAGUCCCAUGUU Exons 3-4 STMN1/ *NM_203401, NM_203399, 2 UAAAGAGAACCGAGAGGCAUU Exon 4 NM_005563, 3 GAAACGAGAGCACGAGAAAUU Exon 4 NM_001145454 4 GAAGAGAAACUGACCCACAUU Exon 4 shRNA Seq. shRNA Sequence

1 TGTCTCCTTCCATGACTGCTTTGTTT 3ʹ UTR STMN1/ *NM_203401, NM_203399, 2 TCTTACCAGATGCTACTGGACTTGAATGG 3ʹ UTR NM_005563, 3 GAATACACTGCCTGTCGCTTGTCTTCTAT 3ʹ UTR NM_001145454 4 GAGTGTGGTCAGGCGGCTCGGACTGAGCA 5ʹ UTR

Non-effective Non-applicable GCACTACCAGAGCTAACTCAGATAGTACT Non-applicable control Seq. (Sequence)

*All four stathmin siRNA and shRNA sequences target all four stathmin gene transcripts

74 Lipofectamine 2000 (Invitrogen, Carlsbad, CA, USA), a cationic liposome-based reagent, was used for siRNA delivery into cells. Cells were transfected with siRNA and

Lipofectamine 2000 (2μg/well) in suspension and seeded into 6 well plates at a density of 2x105cells/well for BE(2)-C and 2-8x105cells/well for SH-SY5Y. The optimal concentration of siRNA was 5nmol/L for BE(2)-C cells and 5 or 50nmol/L for SH-

SY5Y cells. To maximise and prolong stathmin suppression, neuroblastoma cells were double-transfected with siRNA, as previously described (Liu, Schuff-Werner et al.

2004). Briefly, cells were transfected with siRNA and Lipofectamine 2000 in suspension, seeded into 6 well plates and 24 hours later adherent cells were re- transfected with the equivalent concentrations of siRNA and Lipofectamine 2000.

Double siRNA transfections were employed for all experiments shown in Chapters 3

(from Fig. 3.4 onwards), 4 and 5.

2.3 shRNA-mediated stathmin suppression

Short hairpin RNA (shRNA) is a form of RNAi that enables prolonged suppression of target genes via continued expression of an RNA duplex from a plasmid. For sustained suppression of stathmin (STMN1), cells were transfected with HuSH™ retroviral silencing plasmids (pRS) (Origene Technologies, Inc. Rockville, MD). The pRS vector map is shown in Fig. 2.1. Four individual STMN1-specific shRNA pRS plasmids, an empty vector pRS plasmid and a HuSH™ Non-Effective (scrambled) pRS plasmid were provided (see Table 2.2 for shRNA sequences).

Figure 2.1 The pRS vector map.

The pRS vector contains a puromycin resistance gene (for mammalian cell selection) and a shRNA hairpin sequence (29 nucleotide gene-specific insert, 7 nucleotide loop and 29 nucleotide sequence in reverse compliment) followed by a TTTTTT termination sequence, under control of the U6 small nuclear RNA gene promoter (Origene

Technologies, Inc. Rockville, MD).

76 2.3.1 Upscale production of plasmid

DNA

Each shRNA construct (5μg) was reconstituted in 50μL sterile water and then further diluted to a final concentration of 1ng/μL. DH5α competent cells (Invitrogen) were thawed on ice in sterile 17x100mm polypropylene tubes. Plasmid DNA (2ng) was added to 50μL cells and then incubated on ice for 30 minutes. Cells were heated to

42°C in a water bath for 20 seconds and then returned to ice for 2 minutes. Super optimal broth with catabolite repression (SOC) media (450μL) was added to the cells prior to incubation at 37°C in a Certomat® lab shaker (D&A laboratory services,

Baulkham Hills, New South Wales, Australia) for 1 hour at 220rpm. Cells (25μL) were then streaked onto LB-agar plates containing 100μg/mL ampicillin and incubated overnight at 37°C. One colony from each plate was picked, cultured in 2.5mL of ampicillin-containing LB-Broth and incubated at 37°C for 8 hours at 200rpm. For large scale production of each plasmid, 300μL of cells were added to 150mL of ampicillin- containing LB-Broth and cultured overnight at 37°C at 200rpm. Plasmid DNA was isolated using the QIAfilter™ Plasmid Maxi Kit (Qiagen) following manufacturer’s instructions.

2.3.2 Plasmid transfections

Cells were seeded in 6 well plates at 4 x 105/well for SK-N-BE(2) and 8 x 105/well for

SH-SY5Y one day prior to transfection. Cells were transfected with 4μg of each plasmid using 10μL/well Lipofectamine 2000 (Invitrogen) as the delivery vehicle.

Twenty four hours later, media was removed and replaced with fresh 10%FCS/DMEM.

77 Cells were then selected in 1μg/mL puromycin in 10%FCS/DMEM forty eight hours post transfection and were maintained in selection media for at least 10 days. All mock

(un-transfected) cells were dead within 3-4 days. Stathmin suppression was confirmed by western blot analysis 72 hours post-transfection.

2.4 cDNA preparation & qPCR

Stathmin mRNA expression was examined using real-time quantitative polymerase chain reaction (qPCR). Cells were harvested 48-72 hours post siRNA transfection, washed with PBS and total RNA was isolated using the RNeasy Plus Mini Kit (Qiagen,

Valencia, CA) as per manufacturer’s instructions. Tumour tissue was placed in 1.5mL

RNAlater (Qiagen) for 48 hours at 4°C prior to RNA extraction using Trizol® reagent, as per manufacturer’s instructions. RNA was quantified by Nanodrop (Thermo

Scientific) and total RNA (0.25-1μg) was reverse transcribed in a total volume of 10μL with complementary DNA (cDNA) master mix. A 1x cDNA master mix contained 13 units recombinant RNasin® ribonuclease inhibitor (Promega, Madison WI), 3.3mmol/L dithiothreitol (DTT), 0.25mmol/L deoxynucleotide triphosphates (Invitrogen), 1x random primers (Promega), 200 units Moloney Murine Leukemia Virus (MMLV) reverse transcriptase (Promega) and 1x single strand buffer [50mM Tris-HCl (pH 8.3 at

25°C), 75mM KCl, 3mM MgCl2, 10mM DTT]. cDNA samples were heated at 37°C for

1 hour, diluted 1:10 in nuclease-free water and then stored at -20°C. qPCR was performed in a total volume of 25μL containing 2uL of the cDNA mix plus 12.5μL

Quantitect SYBR green (Qiagen), 2.5μL STMN1_1_SG QuantiTect Primer Assay

(contains forward and reverse primers) (Qiagen) and 8μL nuclease-free H20. Samples were placed in a 96-well reaction plate (Applied Biosystems). The plate was spun @

78 1500 x g for 2 minutes before samples were analysed on an Applied Biosystems 7500

PCR system. qPCR conditions were as follows; stage 1) 1 cycle at 50°C for 2 minutes, stage 2) 1 cycle at 95°C for 15 minutes, stage 3) 40 cycles each at 95°C for 15 seconds followed by 60°C for 1 minute, stage 4) 1 cycle each at 95°C for 15 seconds, 60°C for 1 minute, 95°C for 15 seconds. The STMN1_1_SG QuantiTect Primer Assay targets exons 3/4 of stathmin transcript variant 1 (NM_005563), a region homologous to all stathmin’s transcript variants. Stathmin mRNA expression was normalised to the housekeeping genes β2-microglobulin (β2M) or glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (Quantitect primer assays, Qiagen).

2.5 Protein expression analysis

Cells were harvested, washed twice in PBS and the cell pellet lysed in radio- immunoprecipitation assay (RIPA) buffer [150mM NaCl, 10mmol/L nonyl phenoxypolyethoxylethanol (NP)-40, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulphate (SDS), 50mmol/L Tris pH 7.5] containing 1% protease and phosphatase inhibitors (Roche Diagnostics Australia). Frozen mouse tissues were homogenised in

1mL ice-cold RIPA buffer (with protease and phosphatase inhibitors) using the OMNI

TH tissue homogeniser (Lomb Scientific Pty. Ltd., Tarren Point, New South Wales,

Australia). Cell lysates were pulse sonicated for 10 seconds using a Microson™

Ultrasonic homogenizer (Misonix Inc., Farmingdale, NY, USA) and cell debris was removed by centrifugation at 14,000 x g for 20 minutes at 4°C. Supernatant was transferred to a new eppendorf tube and total protein content was determined using a

BCA protein assay kit (Pierce). Cell lysates were diluted in an equal volume of 2x reducing sample buffer (130mM Tris pH8, 20% v/v glycerol, 4.6% w/v SDS, 0.02%

79 bromophenol blue, 129.65mmol/L DTT) and then heated to 95°C for 5 minutes. Total cellular proteins (10-30μg) were resolved on 15% SDS-polyacrylamide gel electrophoresis (PAGE) or Criterion 4-20% Tris-HCl gels (Bio-Rad Laboratories,

Hercules, CA) and electro-transferred to nitrocellulose membrane (GE Healthcare) using the Hoefer Mini Transphor unit (Hoefer Scientific Instruments) containing pre- cooled transfer buffer (0.025M Tris-Glycine, 20% methanol). Proteins were transferred at 70 milliamps (mA) overnight or 300mA for 3 hours at 4°C. Membranes were stained with 0.5% Ponceau S in 1% acetic acid to confirm protein transfer and equal loading and then blocked for 1 hour in 10% skim milk powder (SMP)/Tris-buffered saline

(TBS) or 5% bovine serum albumin (BSA)/TBS. Membranes were washed in

0.5%Tween-20/TBS and proteins detected using the antibodies listed in Table 2.3.

Primary antibodies were incubated overnight at 4ºC and were detected using horseradish peroxidase (HRP)–linked polyclonal antibodies (GE Healthcare Life

Sciences, Uppsala, Sweden). HRP-linked secondary antibodies were incubated for 90 minutes at room temperature. Membranes were developed using ECL Plus Western blotting reagent (GE Healthcare) and chemifluorescent signal was detected and imaged with a Typhoon 9410 laser scanner (GE Healthcare) or X-ray developer (Okamoto,

China).

80 Table 2.3 Antibody dilutions for western blots

Primary Supplier Dilution Secondary Dilution Diluent Total β- Sigma tubulin 1:1000 Sheep α-mouse 1:5000 TBS-T Aldrich (TUB 2.1) Total α- Sigma 0.5% SMP/ tubulin 1:3000 Sheep α-mouse 1:20,000 Aldrich TBS-T (DM1A) Class I β-tubulin Abcam 1:5000 Sheep α-mouse 1:20,000 TBS-T (SAP.4G5) Class II 0.5% SMP/ β-tubulin Covance 1:500 Sheep α-mouse 1:10,000 TBS-T (7B9) Class III β-tubulin Covance 1:1000 Sheep α-mouse 1:12,000 TBS-T (Tuj1) Class IV 0.5% SMP/ β-tubulin Abcam 1:500 Sheep α-mouse 1:2000 TBS-T (ONS.1A6) Acetylated Sigma 1:2000 Sheep α-mouse 1:25,000 TBS-T α-tubulin Aldrich Tyrosinated Sigma 1:1000 Sheep α-mouse 1:60,000 TBS-T α-tubulin Aldrich Sigma Donkey α- Total actin 1:2000 1:15,000 TBS-T Aldrich rabbit β-actin Sigma Donkey α- 1:10,000 1:50,000 TBS-T (AC-15) Aldrich rabbit Sigma Donkey α- γ-actin 1:2000 1:40,000 TBS-T Aldrich sheep MAP4 0.5% SMP/ BD 1:1000 Sheep α-mouse 1:2500 (clone 18) TBS-T Sigma 0.5% SMP/ MAP2† 1:500 Sheep α-mouse 1:500 Aldrich TBS-T Stathmin BD 1:1000 Sheep α-mouse 1:2000 TBS-T Donkey α- 0.5% SMP/ SCG10 ‡ 1:1000 1:5000 rabbit TBS-T Donkey α- 0.5% SMP/ SCLIP ‡ 1:500 1:1000 rabbit TBS-T

† Detects all MAP2 isoforms including MAP2c (~70 kilo Daltons) ‡ Generously provided by Dr. Dominic Chi Hiung Ng, University of Melbourne, Australia. Antibody clone details are written within brackets.

Table 2.3 continued on following page.

Table 2.3 (continued) Antibody dilutions for western blots

Primary Supplier Dilution Secondary Dilution Diluent 0.5% SMP/ LIMK1 • 1:500 Goat α-rat 1:500 TBS-T Donkey α- 5% BSA/ LIMK1-P* Abcam 1:1000 1:1000 rabbit TBS-T 0.5% SMP/ LIMK2 • 1:500 Goat α-rat 1:500 TBS-T Donkey α- 5% BSA/ LIMK2-P* Abcam 1:1000 1:1000 rabbit TBS-T Donkey α- 0.5% SMP/ Cofilin Abcam 1:500 1:500 rabbit TBS-T Donkey α- 0.5% SMP/ Cofilin-P* Abcam 1:500 1:500 rabbit TBS-T Sigma 0.5% SMP/ MLC 1:1000 Sheep α-mouse 1:1000 Aldrich TBS-T Donkey α- 0.5%BSA/ MLC-P* Abcam 1:1000 1:1000 rabbit TBS-T

GAPDH Abcam 1:10,000 Sheep α-mouse 1:15,000 TBS-T

1% SMP/ N-cadherin BD 1:1000 Sheep α-mouse 1:2000 TBS-T Santa Cruz 0.5% SMP/ Vimentin 1:200 Sheep α-mouse 1:5000 Biotech. TBS-T Donkey α- 0.5% SMP/ Paxillin BD 1:2000 1:4000 rabbit TBS-T Donkey α- 5%BSA/TBS- Paxillin-P* Invitrogen 1:1000 1:4000 rabbit T Stathmin Santa Cruz Donkey α- 0.5% SMP/ 1:1000 1:1000 Ser16* Biotech. rabbit TBS-T Stathmin Donkey α- 0.5% SMP/ Abnova 1:1000 1:1000 Ser25* rabbit TBS-T Stathmin Donkey α- 0.5% SMP/ Abcam 1:500 1:500 Ser38* rabbit TBS-T Stathmin Donkey α- 0.5% SMP/ Abcam 1:500 1:500 Ser63* rabbit TBS-T

*Phospho antibody • Generously provided by Dr. Ora Bernard, St. Vincent’s Institute of Medical Research, Melbourne, Australia.

82 2.5.1.1 Examination of protein phosphorylation

Forty eight hours after double siRNA-transfection, cells were serum-starved in DMEM for 2 hours before treating with 10% FCS/DMEM for the indicated time points. Cells were harvested in ice-cold PBS, centrifuged at 4500 x g and the cell pellets resuspended in RIPA buffer containing protease and phosphatase inhibitors. For experiments using the ROCK inhibitor, cells were pre-treated for 1 hour with 10-

40μmol/L Y-27632, incubated with equivalent concentrations of Y-27632-containing

DMEM for 2 hours and then treated with 10% serum/DMEM before harvesting as described above. Total cellular proteins (30μg) were resolved on Criterion 4-20% Tris-

HCl gels (Bio-Rad Laboratories, Hercules, CA) and the phosphorylation of cofilin

(serine 3), LIMK1 (threonine 508), LIMK2 (threonine 505), paxillin (tyrosine 118) and stathmin (serine’s 16, 25, 38 and 63) were analysed by western blot (see Table 2.3 for antibody information).

2.6 Cell proliferation assay

Cells were double-transfected with siRNA and 24 hours later, harvested and seeded into

96 well plates at a density of 3000 cells/well for BE(2)-C and 12,000 cells/well for SH-

SY5Y. Cellular metabolic activity (cell proliferation) was measured by adding resazurin solution [597μmol/L resazurin, 67μmol/L methylene blue, 1μmol/L potassium hexacyanoferrate (III), 1μmol/L potassium hexacyanoferrate (II) trihydrate in phosphate buffered saline] a 1:10 dilution to triplicate wells 12 hours prior to fluorometric detection [using a Victor2™ plate reader (Perkin Elmer, Waltham, USA) excitation

560nm- emission 590nm] at the indicated time points.

83 2.7 Cell enumeration & viability

Cells were double-transfected with siRNA in 6 well plates. Total cells (floating and adherent) were harvested at the indicated time points and dead cells were identified using 0.4% trypan blue stain (Invitrogen). Cell counts were obtained by counting the number of live cells/well. Percentage cell viability was determined by dividing the total number of live cells by the total number of cells (live + dead).

2.8 Tubulin polymerisation assay

Forty eight hours after double siRNA-transfection, cells were either maintained in

10%FCS/DMEM or serum-starved for 2 hours and then stimulated with 25ng/mL human recombinant platelet-derived growth factor (PDGF)-BB (Sigma Aldrich) in

10%FCS/DMEM for 1 hour. Cells were washed 2 times in warm PBS prior to incubation with 150μL/well of pre-warmed hypotonic buffer [1mmol/L MgCl2, 2 mmol/L ethylene glycol tetraacetic acid (EGTA), 0.5% NP-40, 1x protease inhibitors and 20mmol/L Tris pH6.8] for 10 minutes at 37°C. Cells were scraped from the plate and centrifuged at 14,000 x g for 10 minutes. The supernatant (soluble fraction) was transferred to a new eppendorf tube and the cell pellet was resuspended in 300μL hypotonic buffer. One hundred microliters of 4x sample buffer (45% glycerol, 9.2%

SDS, 0.3mol/L Tris pH 6.8, 0.04% bromophenol blue, 20% β-mercaptoethanol) was added to both soluble and cell pellet fractions. The cell pellet fraction containing sample buffer was then sonicated until the cell pellet dissolved. The soluble fractions were pulse sonicated for 10 seconds. All samples were then heated at 90°C for 10 minutes and 45μL of each fraction was resolved by 15% SDS-PAGE. Tubulin fractions (soluble

84 ‘S’ and polymerised ‘P’) were detected by western blot using total α-tubulin antibody.

The levels of polymerised tubulin were quantified by densitometry and the percentage of polymerised tubulin was calculated as follows [(‘P’/ ‘P’+ ‘S’) x100].

2.9 Cytotoxic drugs & inhibitors

Paclitaxel, Taxus. sp. (Calbiochem, MerckKGaA, Darmstadt, Germany), and vincristine sulfate (Sigma-Aldrich) were prepared at stock concentrations of 2mmol/L in dimethyl sulfoxide (DMSO) (Sigma Aldrich). Epothilone B (Calbiochem) and ZM447439

(Tocris, Bristol, UK) were prepared in DMSO at 100μmol/L and 10mmol/L, respectively. The final concentration of DMSO exposed to cells was no more than 0.5%

(v/v) for the duration of the experiment. Cisplatin (Pfizer, Perth, Australia), etoposide

(Pfizer) and doxorubicin hydrochloride (Mayne Pharma Pty. Ltd., Mulgrave, Australia) were provided at stock solutions of 1mg/mL (3.33mmol/L), 20mg/mL (33.98mmol/L) and 2mg/mL (3.45mmol/L), respectively. The ROCK inhibitor Y-27632 [(+)-(R)-trans-

4(1-aminoethyl)-N-(4-pyridyl)cyclohexanecarboxamide dihydrochloride] (Sigma-

Aldrich) was prepared at a concentration of 1mmol/L in sterile H20.

2.10 Cell cycle analysis

The distribution of DNA content in siRNA-transfected cells was determined by flow cytometry. Fifty six hours post transfection; cells were exposed to the indicated concentrations of paclitaxel for 18 hours. Adherent and floating cells were harvested, washed with warm PBS and stained with propidium iodide (PI) solution [(0.2% Triton

X-100 (Sigma-Aldrich), 50 μg/ml PI (Sigma-Aldrich), and 2 μg/ml DNase-free RNase

85 (Roche Diagnostics Australia), phosphate-buffered saline] for 15 min at 37°C in the dark. DNA content of each cell was measured by flow cytometry (FACSCalibur, BD

Biosciences, Franklin Lakes, NJ) and analysed using CellQuest (BD Biosciences). The

CellQuest program was used to quantitate the distribution of cells in each phase of the cell cycle: sub-G1, G1, S, and G2-M.

2.11 Cytotoxicity assays

Cells were harvested 32 hours post-siRNA transfection and seeded into 96 well plates at a density of 5000 cells/well for BE(2)-C and 12,000 cells/well for SH-SY5Y. Cells were incubated at 37°C overnight prior to drug treatment. Drugs were diluted in cell culture medium and added to each well at the indicated concentrations. Following 72 hours incubation at 37°C, cell viability was detected by addition of resazurin solution and fluorometric analysis using a Victor2™ plate reader (Perkin Elmer). Cell viability of drug treated cells is displayed as a percentage of control, untreated cells (i.e. cells with no drug). The final concentration of DMSO exposed to the cells was no more than 0.5%

(v/v) for the duration of the experiment.

2.12 Drug-treated clonogenic assays

Cells were double-transfected with siRNA and 48 hours later seeded into 6 well plates at a density of 300/well for BE(2)-C and 600/well for SH-SY5Y. Cells were incubated for 6 hours at 37°C prior to drug exposure. Drugs were diluted in cell culture medium

86 and added to each well at the indicated concentrations. After 72 hours incubation at

37°C, medium from each well was removed and replaced with 10%FCS DMEM. Media was changed every 3 days until colonies were visible. Colonies were fixed and stained with 0.5% crystal violet in methanol and individual colonies were manually counted.

Results are expressed as a surviving fraction [colony number / (number of cells seeded

× plating efficiency)]. Plating efficiency equals the colony number divided by the number of cells seeded in the drug-free medium. Dose response curves were plotted for each drug and cell line combination. Alterations in drug sensitivity were calculated as a fold change as determined by dividing the half maximal inhibitory concentration (IC50) of the stathmin siRNA-treated group by the IC50 of the control siRNA-treated group.

2.13 Retinoic acid-induced

differentiation

All-trans retinoic acid (ATRA) (Sigma-Aldrich) was prepared by reconstituting 50mg

ATRA powder in 33.3mL of 100% ethanol (5 mmol/L stock concentration). Stock solution was covered in aluminium foil and stored at -80ºC to prevent isomerisation

(Armstrong, Redfern et al. 2005). Naïve neuroblastoma cells or neuroblastoma cells double transfected with siRNA were seeded in 6 well plates and treated with either

10μmol/L ATRA or the equivalent volume of 100% ethanol (vehicle control) diluted in

10%FCS/DMEM. Tissue culture plates were covered in foil to prevent light exposure and incubated at 37ºC. Cells were imaged by phase contrast microscopy (see section

2.14.1) and then harvested for gene and protein expression analyses. For longer term

87 experiments (7 and 10 day treatments), fresh ATRA media was added to the cells every

72 hours.

2.14 Microscopy

2.14.1 Phase contrast

Cells were grown in 6 well plates and then imaged using a 10x objective on the

Axiovert S100 inverted phase-contrast microscope (Zeiss) equipped with a SPOT RT

Slider CCD scientific digital camera system driven by SPOT Basic software (Diagnostic

Instruments, Inc. Sterling Heights, MI).

2.14.2 Immunofluorescence

Cells were seeded at a density of 30,000 cells/well into sterile 4-well Nunc Lab-Tek™ chamber slides (Thermo Fisher Scientific Inc.). For experiments using the ROCK inhibitor, cells were pre-treated with 10μmol/L Y-27632 prior to incubation in serum- free media containing 10μmol/L Y-27632 for 2 hours. Cells were incubated at 37°C for

24 hours and then fixed in 100% ice-cold methanol for 8 minutes or 4% paraformaldehyde (PFM)/PBS for 12 minutes. PFM-fixed cells were permeabilised with

1% Triton X/PBS for 3 minutes and rinsed 3 times in PBS. Methanol and PFM-fixed cells were blocked in 0.45μm-filtered 10%FCS/PBS for 10 minutes at room temperature. Cells were then incubated with primary monoclonal antibodies (diluted in

5%FCS/PBS) against α-tubulin (1:400) and stathmin (1:100) for 30 minutes at room temperature. Slides were washed 3 times with 0.1%Tween/PBS for 5 minutes. Primary antibodies were detected using Alexa Fluor® 488 or 568 goat anti-mouse IgG

88 (Invitrogen) diluted 1:1000 in 5%FCS/PBS and incubated for 30 minutes at room temperature. Slides were washed as described above. To observe actin filaments, cells were stained with Alexa Fluor® 568 Phalloidin (Invitrogen) diluted 1:50 in

5%FCS/PBS and incubated for 10 minutes at room temperature. Slides were rinsed in

MQ-H20 and then mounted using VECTASHIELD® mounting medium with 4',6- diamidino-2-phenylindole (DAPI) (Vector Laboratories, Burlingame, CA). Cytoskeletal structures were then visualised and imaged using an Olympus FluoView™ FV1000 confocal microscope with a 63X 1.35 NA oil objective.

2.15 Neurite counts

Neuroblastoma cells were treated with 10μmol/L ATRA (as described in section 2.13) and changes in cell morphology were visualised by taking 5 random phase-contrast images per treatment group using a 10x objective (see section 2.14.1). A cell bearing at least one neurite twice the length of the cell body was scored as positive (Neubrand,

Thomas et al. 2010) and was counted using ImageJ software (Rasband, W.S., ImageJ,

U. S. National Institutes of Health, Bethesda, Maryland, USA, http://rsb.info.nih.gov/ij/,

1997-2009). The percentage of neurite positive cells equalled [(number of neurite positive cells/neurite positive + neurite negative cells) x100].

2.16 Soft agar assays

Forty eight hours after double-siRNA transfection, BE(2)-C (1000 cells) and SH-SY5Y

(4000 cells) were re-suspended in 0.33% agar in growth media and plated on a 0.5% solidified agar layer in triplicate dishes. After 12 days for BE(2)-C cells and 21 days for

89 SH-SY5Y cells, colonies were individually counted and photographed using the

Axiovert 200 M microscope (Zeiss, Oberkochen, Germany) coupled to an

AxioCamMR3 camera and driven by the Axio Vision software (Zeiss). The results were expressed as percentage colony formation [(number of colonies formed/total number of cells seeded) x 100%]. Colony size (perimeter) was measured manually using Axio

Vision software (Zeiss). Average colony size was calculated for each run by measuring

90 colony sizes for both control and stathmin siRNA-treated cells.

2.17 Monolayer wound-healing assays

Twenty four hours post-double siRNA transfection, cells were seeded into 60x15mm,

2mm gridded dishes (Corning Inc., Corning, NY, USA) at a density of 1.5x106 cells/well for control siRNA-treated cells and 2.5x106 cells/well for stathmin siRNA- treated cells (more stathmin siRNA-treated cells were required to ensure the same level of confluence as the control cells). Cells were left to adhere and become confluent for

24 hours before creating 5 individual scratch ‘wound’ sites in each well using 200μL tips. Cell debris was removed from wells by washing 3 times in culture medium. Phase contrast images were taken of 5 individual wound sites using a 10x objective (as described in section 2.14.1). These initial images were considered ‘time 0’ (0 hours).

The same wound sites were then imaged 8, 12, 24, 32 and 48 hours post-scratch. Using

ImageJ software, the percentage wound recovery was calculated based on the remaining surface area within each wound at each time point compared to time 0 hours. The

90 percentage wound recovery equalled 100-[(surface area of wound at x hours/surface area of wound at ‘time 0’) x 100].

2.18 Chemotaxis migration & invasion assays

Cells were double-transfected with siRNA and 24 hours later incubated in serum-free

DMEM for 2 hours. For experiments using the ROCK inhibitor, cells were pre-treated with 10μmol/L Y-27632 diluted in 10%FCS/DMEM for 1 hour prior to incubation with

10μmol/L Y-27632 diluted in serum-free media. The underside of 8μm PET 24-well migration inserts (BD Biosciences) or 8μm BD Biocoat™ BD Matrigel™ 24-well invasion chambers (BD Biosciences) were coated with 10μg/mL collagen IV (BD

Biosciences) and incubated at room temperature for 1 hour. Inserts were then pre-wet in

1mL serum-free DMEM for 30-60 minutes in companion plate wells. Cells were harvested, resuspended in 500μL serum-free DMEM and seeded into the top chamber at the following densities: BE(2)-C cells at 15,000 cells/insert for the migration assay and

25,000 cells/insert for the invasion assay, SH-SY5Y cells at 60,000 cells/insert for the migration assay and 120,000 cells/insert for the invasion assay, and SK-N-BE(2)/TGL cells at 60,000 cells/insert for the invasion assay. The lower chamber contained 10%

FCS and 25ng/mL human recombinant platelet-derived growth factor (PDGF)-BB

(Sigma-Aldrich) diluted in DMEM, to act as chemo-attractants. For experiments using the ROCK inhibitor, 10μmol/L Y-27632 was diluted in serum-free or 10%FCS/DMEM and added to the top and bottom chambers, respectively. After seeding, cells were incubated for 24 hours [for SK-N-BE(2)/TGL cells] or 48 hours [for BE(2)-C and SH-

91 SY5Y cells] at 37°C. The inserts were removed from the companion plate and fixed using 100% methanol and stained with May-Grunwald (Sigma-Aldrich) which was diluted 1:3 in MQ-H20 (0.45μm-filtered) and Giemsa (Sigma-Aldrich) which was diluted 1:25 in MQ-H20 (0.45μm-filtered). Once dry, membranes were removed with a scalpel, placed onto glass slides and covered with glass coverslips. Fifteen random images were taken of each membrane (top and bottom surfaces) with an Olympus BH-2 inverted microscope and a 20x objective. Results are expressed as a migration or invasion index [(number of cells on under surface of membrane divided by total number of cells on both surfaces of the membrane) x 100] or an invasion index (% of control) where control cells are 100% and all other treatments are calculated as a percentage of control.

2.19 Luciferase-expressing neuroblastoma cells

The SK-N-BE(2) cells provided by Dr. Sylvain Baruchel (Research Institute at The

Hospital for Sick Children, Toronto) have previously been used in a neuroblastoma xenograft (metastatic) model (Zhang, Smith et al. 2009). The SK-N-BE(2)/TGL cells were made by retroviral delivery of the SFG-NESTGL triple-modality reporter construct (Fig. 2.2) (Ponomarev, Doubrovin et al. 2004) into SK-N-BE(2) cells, as performed by Dr. Jamie Fletcher (Children’s Cancer Institute Australia, New South

Wales). This retroviral vector (SFG-Ntp backbone) harbours the Aequorea victoria green fluorescent protein (GFP), the herpes simplex virus 1 thymidine kinase (HSV1-tk) reporter gene and the Photinus pyralis (firefly) luciferase gene.

NES SD SA

LTR HSV1-tk eGFP Luc LTR

Figure 2.2 Schematic of the SFG-NESTGL triple-modality reporter construct.

The HSV-tk1 cDNA bearing a nuclear export signal (NES) from mitogen-activated protein kinase kinase (MAPKK) was fused to the N-terminus of eGFP cDNA with firefly luciferase (Luc) at the C-terminus. When transcribed and translated, these 3 functional subunits code for a single fusion protein of ~130kDa predicted molecular weight (Ponomarev, Doubrovin et al. 2004). SD- splice donor site, SA- splice acceptor site, LTR- long terminal repeat.

93 GFP-expressing SK-N-BE(2) cells were selected by fluorescence-activated cell sorting and a high GFP-expressing SK-N-BE(2)/TGL population was pooled. Luciferase assays

(see section 2.20) then confirmed that this population expressed high levels of luciferase.

2.20 In vitro luciferase assays

To evaluate luciferase expression in vitro, SK-N-BE(2)/TGL cells were seeded at

10,000 cells/well in 96 well plates. Cells were then serially diluted (1:2 dilution) across the plate to a final density of 5 cells/well. Three hours after seeding, the cells were placed inside the pre-warmed specimen chamber of the IVIS Imaging System

(Xenogen, Alameda, CA) and imaged (medium binning, F-stop of 1) using the highly sensitive, cooled CCD camera. This initial imaging was to determine the background bioluminescence emitted from the cells. Firefly D-luciferin potassium salt (Gold

Biotechnology, St. Louis, MO) was diluted to 15mg/mL in Dulbecco’s Phosphate

Buffered Saline (DPBS) (Gibco-Invitrogen), filtered (0.22μm) and then added to wells at a final concentration of 150μg/mL. The cells were then returned to the specimen chamber for 3 minutes prior to imaging, as described above. Bioluminescence was analysed using the LIVINGIMAGE (v3.2) software (Xenogen). Photon emission from the cells, as calculated by total flux (photons/sec), was measured using the region of interest (ROI) method as described in the Caliper LifeSciences manual (Caliper Life

Sciences, Inc.). Background luminescence was determined by measuring the total flux from the cells prior to the addition of the substrate. These values were then subtracted from the total flux obtained from reciprocal wells after addition of the substrate.

94 2.21 Orthotopic injection of neuroblastoma cells

Severe-combined immune-deficient (SCID)-Beige mice were chosen for this study as they are more immune-compromised than SCID mice as a result of the beige mutation

(defective natural killer cell and macrophage activity) and have successfully been used in an orthotopic neuroblastoma model (Khanna, Jaboin et al. 2002). Eight week old male mice (obtained from Professor Chris Ormandy, Garvan Institute of Medical

Research, New South Wales, Australia) were housed under specific pathogen-free conditions and fed autoclaved standard chow and water ad libitum. One day prior to surgery, surgical sites were prepared by removing fur on the left flank of the animal between the front and hind limbs. Fur was removed using Nair® depilatory cream that was applied to the area using sterile cotton swabs. The cream was removed within one minute (to prevent skin irritation) using saline solution and gauze pads. On the day of surgery, SK-N-BE(2)/TGL shRNA-expressing neuroblastoma cells were harvested using PBS/trypsin and re-suspended in culture media. Cells were then spun at 1500 x g for 5 minutes, supernatant removed and the cell pellet resuspended in warm PBS. Cell enumeration and viability were assessed using 0.4% trypan blue (Invitrogen) and experiments were continued if cells were greater than 95% viable. Mice were anesthetised using 4% isoflurane which was reduced to 2% isoflurane to sustain sedation. Pain relief [Temgesic® (buprenorphine)] was diluted in saline and administered subcutaneously at a dosage of 0.025mg/kg. Two million cells were resuspended in 15μL Cultrex® basement membrane extract (BME) (Trevigen, Inc.

Gaithersburg, MD, USA) and injected into the left adrenal fat pad, as described

95 previously (Khanna, Jaboin et al. 2002). Briefly, the surgical area was wiped with iodine and alcohol swabs and then a ~2cm vertical incision was made in the outer skin layer (located over the spleen which is visible through the skin layer). Another 1cm vertical incision was made into the peritoneum to expose the spleen, kidney and left adrenal gland. The spleen was moved gently to the side and the cell/BME mix (~20μL) was injected into the left adrenal gland fat pad using a 29ga. needle attached to a 0.5mL syringe (Terumo Medical Corporation, Elkton, MD, USA). Any mice with obvious spillage from the injection site were sacrificed immediately via CO2 asphyxiation. The peritoneum was closed with Safil 5/0 absorbable sutures (B. Braun Australia, Bella

Vista, New South Wales) and the skin layer with Reflex™ 7mm stainless steel wound clips (Fine Science Tools Inc., North Vancouver, B.C., Canada). Saline (1mL) and pain relief (Temgesic® at a dosage of 0.025mg/kg) were administered to mice immediately after surgery via subcutaneous injection. Animals were returned to clean, pre-warmed cages and monitored during recovery. Wound clips were removed from animals 9 days post-surgery. All animal experiments were approved by the Animal Ethics Committee,

University of New South Wales (ACEC#: 10/114A).

2.22 In vivo & ex vivo imaging & tumour measurements

Luciferase-expressing neuroblastoma cells were visualised in vivo via intraperitoneal

(i.p.) injection of the luciferase substrate, D-luciferin (Gold Biotechnology) at 150 mg/kg in DPBS into mice. Animals were anaesthetised using 4% isoflurane (Abbott

Laboratories, North Chicago, Illinois) and placed onto the warmed stage inside the IVIS

96 Imaging System specimen chamber where they received continuous exposure to 2% isoflurane to sustain sedation. Ten minutes after injection, mice were imaged on the ventral and left lateral sides (medium binning, F-stop 1). Mice were imaged 1 day post- surgery to check for tumour cell leakage from the injection site. Any animals with obvious leakage were sacrificed and eliminated from the study. Mice remaining in the study were imaged 9 days post-injection, every week thereafter and on the day of sacrifice. Mice were euthanised by CO2 asphyxiation when the tumour reached 1500 mm3 (by palpation) or 38 days post injection of tumour cells, whichever came first.

Tumour size was measured by weight and volume in mm3 using the formula: volume =

(length x width x depth)/2.

For ex vivo imaging, 150 mg/kg D-luciferin was injected i.p. prior to necropsy. Animals were sacrificed by C02 asphyxiation, skinned and the carcass, tumour and organs were placed in 10cm petri dishes or 6 well plates for imaging. The tumour, organs and carcasses were covered with D-luciferin (300 μg/ml in DPBS) for 30 seconds prior to imaging (medium binning, F-stop 1) using the IVIS Imaging System. To determine background bioluminescence for in vivo and ex vivo imaging, 2 naïve mice and their organs were imaged using the IVIS Imaging System, as described above.

Bioluminescent images of mice and organs were digitised and electronically displayed as a pseudocolor overlay onto a grey scale animal image on the LIVINGIMAGE (v3.2) software (Xenogen). Bioluminescence signal was then quantified from the left lateral side of the live animal and lungs post sacrifice using the ROI method. The mean bioluminescence values from naïve mice, as measured in total flux (photons/sec), were

511700 and 10478 for the whole live animal and lungs, respectively. These values were

97 subtracted from the signal obtained from similarly sized ROIs from whole animals and organs of mice injected with luciferase-expressing neuroblastoma cells.

2.23 Immunohistochemistry

Tissues were fixed in 10% formalin (Sigma, St. Louis, Missouri) or 4% paraformaldehyde (ProSciTech, Thuringowa, Queensland, Australia) for 48 hours and then washed twice in PBS and once in 70% ethanol before placing in histology cassettes

(ProSciTech). Cassettes were immersed in 70% ethanol prior to sending to the

Microscopy Unit of the University of New South Wales, Australia for processing.

Tissues were paraffin-embedded, 5μm sections cut and stained with haematoxylin and eosin (H&E) or prepared on aminoalkysilane-treated glass slides for immunohistochemistry (IHC). For IHC analyses, slides were incubated at 60°C for 30 minutes to assist tissue section attachment. Tissues were then deparaffinised with three incubations in xylene (3 minutes each) and rehydrated through graded ethanol series of

100%, 95% and 70% (5 minutes each) and two MQ-H20 washes (2 minutes each).

Antigen retrieval was performed by placing slides in 400mL citrate buffer (10mM citrate buffer, 0.05% Tween-20 pH 6) which was heated on high for 3 minutes in a microwave oven followed by 15 minutes at 104°C in a regular oven. The solution was left to cool for 1 hour at room temperature and then slides were washed twice with MQ-

H20. Endogenous peroxidase activity was blocked by placing slides in peroxidase blocking solution (1% H202, 3.3% methanol) for 10 minutes. Tissue sections were outlined with a PAP pen and then blocked with 10% goat serum/PBS for 1 hour. To detect luciferase and stathmin expression, tissue sections were incubated for 1 hour at room temperature with rabbit polyclonal antibodies against luciferase (Fitzgerald,

98 Acton, MA, USA) and stathmin (Abcam, Cambridge, UK) diluted in 1% goat serum/PBS at 1:100 and 1:250 respectively. To ensure secondary antibody specificity, one slide was incubated with rabbit IgG control antibody at a concentration equivalent to the primary antibodies. Slides were washed 3 times in 0.05% Tween-20/PBS (PBS-

T) for 5 minutes followed by 1 quick wash in PBS. Tissue sections were incubated with biotinylated secondary antibody diluted 1:200 in 1% goat serum/PBS for 10 minutes.

Slides were then washed in PBS-T as described above and incubated for 5 minutes with avidin-biotin-peroxidase complex (2 drops each of reagent A and B in 5mL PBS) from the Vectastain ABC Kit (Vector Labs, Burlingame, California, USA). Slides were washed twice in PBS and then incubated with ImmPACT™ DAB substrate (Vector

Labs) at room temperature until the desired intensity developed. Slides were rinsed in

MQ-H20, counterstained with Mayer’s haematoxylin (nuclear counterstain) for 1 minute, dehydrated and then mounted with EUKITT mounting medium (O. Kindler

GmbH and Co, Freiburg, Germany). Slides were scanned using an Aperio ScanScope

XT Slide Scanner and images were analysed using the ImageScope software (Aperio,

Vista, CA, USA).

2.24 Statistical analyses

Statistical analyses were performed using the GraphPad Prism program. For in vitro experiments, results are expressed as means of at least three independent experiments ± standard error of the mean (SEM). Unpaired, two-tailed Student’s t tests were used to determine the statistical differences between various experimental and control groups, with p<0.05 considered statistically significant. For in vivo experiments, the bioluminescent signal [measured as total flux (photons/sec)] from control and stathmin

99 shRNA-expressing groups were compared using the non-parametric Mann-Whitney U test, where p<0.05 was considered statistically significant.

100 Chapter 3. Results I

101 Evaluation of stathmin’s role in neuroblastoma cell proliferation and

drug resistance

Neuroblastoma originates from precursor cells of the sympathetic nervous system and is the most common malignancy in infants. The most aggressive neuroblastoma tumours are highly proliferative and intrinsically resistant to most chemotherapeutic agents.

Treatment for neuroblastoma often includes myeloablation (high dose) chemotherapy using various chemotherapeutic agents such as the tubulin-binding agent vincristine and various DNA-targeted agents such as cisplatin, doxorubicin and etoposide (Pearson,

Pinkerton et al. 2008). Although most children respond well to the first round of cyto- reductive therapy, many will subsequently relapse. This highlights a need to identify new drug targets to treat drug-refractory neuroblastoma.

Stathmin is a key microtubule destabilising protein over-expressed in numerous malignancies. High expression of stathmin has been closely associated with more proliferative and drug-resistant disease in a range of non-neuronal cancers, as reviewed in Chapter 1. A number of studies have shown that altering stathmin expression and activity disrupts cell cycle progression and proliferation in some cells suggesting that stathmin plays a role in regulating mitotic microtubules and cell division (Giampietro,

Luzzati et al. 2005; Mistry, Bank et al. 2005; Singer, Ehemann et al. 2007; Wang, Dong et al. 2007; Singer, Malz et al. 2009). Furthermore, numerous reports have shown that inhibiting stathmin expression can enhance cancer cell sensitivity to various chemotherapeutic agents [reviewed in (Rana, Maples et al. 2008)] (refer to Chapter 1,

Table 1.1). However, one recent report has demonstrated that inhibiting stathmin expression in ovarian cancer cell lines did not alter their sensitivity to paclitaxel and

102 cisplatin (Aoki, Oda et al. 2009). Therefore, stathmin’s role in mediating drug sensitivity may be cell type dependent.

Stathmin is highly expressed in neuroblastoma and its expression and phosphorylation is altered between MYCN-amplified and non-amplified neuroblastoma cells (Hailat,

Strahler et al. 1990). However stathmin’s functional role in this malignancy is unresolved. Based on previous studies in non-neuronal cancers, it was proposed that stathmin may enhance cell proliferation and confer a drug resistant phenotype in neuroblastoma. To address this, a small-interfering RNA (siRNA) approach was utilised to suppress stathmin expression and subsequently examine stathmin’s role in neuroblastoma cell cycle progression and proliferation in vitro. To examine stathmin’s influence on drug resistance, stathmin suppressed neuroblastoma cells were exposed to various chemotherapeutic agents and their response assessed using both short-term and long-term in vitro assays.

3.1 Stathmin protein expression in neuroblastoma cell lines

Neuroblastoma tumours arise from multi-potential cells of the neural crest. In order to investigate stathmin’s role in neuroblastoma cell proliferation and drug resistance, a panel of neuroblastoma cell lines, containing all three neuroblastoma cell types (refer to

Materials & Methods 2.1), were screened for stathmin protein expression by western blot analysis (Fig. 3.1A). In general, the ‘N’ type cells including SH-SY5Y and IMR-32

103 Figure 3.1 Stathmin protein expression in neuroblastoma cell lines.

A) A representative western blot showing stathmin protein expression in a panel of neuroblastoma cell lines. GAPDH was used as a loading control. B) Quantitation of stathmin protein expression (normalised to GAPDH) in neuroblastoma cell lines. Data represent the mean ±SEM (error bars) of at least 3 independent experiments.

104

Stathmin

GAPDH

105 cells, and ‘I’ type cells, such as the BE(2)-C cells expressed higher levels of stathmin protein than the ‘S’ type cells, such as the SHEP cells (Fig. 3.1B). Two cell lines,

BE(2)-C and SH-SY5Y, were chosen for future experiments as they expressed moderate levels of stathmin protein relative to the other cell lines. In addition, BE(2)-C cells harbour various molecular traits that may influence stathmin’s role in neuroblastoma drug resistance including MYCN amplification (Carr, Bown et al. 2007) and mutant p53

(Tweddle, Malcolm et al. 2001). BE(2)-C cells contain ~55 MYCN copies (Carr, Bown et al. 2007), are hemizygotes for the p53 locus on chromosome 17p and contain a missense mutation in exon 5 that inactivates p53 (Tweddle, Malcolm et al. 2001). In contrast, SH-SY5Y cells are wild-type for p53, are non-MYCN-amplified and express moderate levels of the N-myc protein (Chesler, Goldenberg et al. 2008).

3.2 Stathmin gene silencing in neuroblastoma cells

Small interfering RNAs (siRNAs), usually 20-25 base pairs long, silence target gene expression by exploiting a cells’ RNA interference (RNAi) machinery. SiRNA was employed in this study to examine stathmin’s functional role in neuroblastoma biology.

In both cell lines, 5nmol/L of stathmin siRNA effectively suppressed stathmin protein expression by 66% (±5.93%) for BE(2)-C cells and 66% (±7.34%) for SH-SY5Y cells relative to control#2 siRNA-treated cells (Fig. 3.2A, 3.2B). This concentration of siRNA (the lowest effective concentration) was chosen to reduce potential ‘off-target’ effects of siRNA treatment.

106 Figure 3.2 SiRNA optimisation in neuroblastoma cells.

A) A representative western blot and corresponding quantitation (lower graph) of stathmin protein expression in siRNA-transfected BE(2)-C cells 72hrs post transfection.

B) A representative western blot and corresponding quantitation (lower graph) of stathmin protein expression in siRNA-transfected SH-SY5Y cells 72hrs post transfection. In both cell lines, stathmin protein expression was normalised to GAPDH

(loading control) and calculated as a percentage of Ctrl #2 siRNA. Data represent the mean ±SEM (error bars) of 3 independent experiments.

107

A BE(2)-C

Stathmin siRNA (nM) Ctrl siRNA

Mock 1 5 25 50 100 #1 #2 Stathmin GAPDH

Stathmin siRNA (nM) Ctrl siRNA

B SH-SY5Y

Stathmin siRNA (nM) Ctrl siRNA Mock 1 5 25 50 100 #1 #2 Stathmin GAPDH

Stathmin siRNA (nM) Ctrl siRNA

108 Stathmin suppression using 5 nmol/L siRNA was then confirmed at the mRNA level.

This concentration of siRNA suppressed stathmin gene expression by 84% (±1.77%; p<0.0001) in BE(2)-C cells (Fig. 3.3A) and 62% (±6.3%; p<0.005) in SH-SY5Y cells

(Fig. 3.3B). To maximize and extend stathmin suppression, neuroblastoma cells were transfected twice with 5nmol/L siRNA (see Materials & Methods 2.2) (Fig. 3.4A).

Double siRNA-transfection of BE(2)-C cells reduced stathmin protein expression by

93% (±3.44%) at 48 hours after the second transfection, relative to controls (Fig. 3.4B).

Furthermore, double siRNA-transfection prolonged stathmin protein suppression for up to 144 hours compared to controls (Figs. 3.4B). Importantly, double siRNA-transfection of neuroblastoma cells was not toxic to the cells. Double siRNA transfections were therefore used for all further experiments.

3.3 Stathmin does not influence the expression of cytoskeletal-related proteins in neuroblastoma cells

Stathmin is the founding member of the highly homologous stathmin-like protein family. To ensure siRNA specificity, SCG10 and SCLIP expression were examined in stathmin siRNA-suppressed BE(2)-C cells (Fig. 3.4B). Results showed that stathmin siRNA was specific as it did not markedly influence the expression of either of these proteins (Fig. 3.4B).

109 Figure 3.3 Stathmin gene expression in siRNA-transfected neuroblastoma cells.

Stathmin mRNA levels in BE(2)-C (A) and SH-SY5Y cells (B) 48 hours post transfection with 5 nmol/L stathmin siRNA (STMN) or control (Ctrl) siRNA. Stathmin mRNA was normalised to β2-microglobulin (β2M). Data represent the mean ±SEM

(error bars) of 4 independent experiments. ** p<0.005, *** p<0.001.

110

A BE(2)-C

B SH-SY5Y

111 Figure 3.4 Double siRNA transfection of neuroblastoma cells.

A) Double transfection of BE(2)-C cells with stathmin siRNA (STMN) or control siRNA (Ctrl). Stathmin suppression was enhanced and maintained in STMN siRNA- treated cells up to 144 hours post-transfection, compared to Ctrl siRNA-treated cells.

Stathmin suppression did not induce compensatory expression of the stathmin-like proteins SCG10 and SCLIP. GAPDH was used as a loading control. B) Quantitation of stathmin protein expression (normalised to GAPDH) in double siRNA-transfected

BE(2)-C cells up to 144 hours post transfection. Stathmin protein expression was calculated as a percentage of Ctrl at each time point. Data represent the mean ±SEM

(error bars) of 3 independent experiments.

112

SCG10 Stathmin SCLIP GAPDH Time (hrs) 24 48 72 96 120 144

113 Stathmin is an important regulator of microtubule dynamics and has also been linked to regulation of the actin network (Daub, Gevaert et al. 2001; Wittmann, Bokoch et al.

2004; Watabe-Uchida, John et al. 2006; Tanaka, Hamano et al. 2007; Takahashi and

Suzuki 2009; Belletti, Pellizzari et al. 2010). The expression levels of microtubule and actin-related proteins were therefore examined in stathmin-suppressed BE(2)-C cells

(Fig. 3.5). Stathmin suppression did not alter the expression of the microtubule stabilising protein, microtubule-associated protein 4 (MAP4) (Fig. 3.5B), total α- and β- tubulin, the β-tubulin isotypes; βI, βII, βIII, βIV (Fig. 3.5C) or total, β- and γ-actin (Fig.

3.5D). Furthermore, the expression of the microtubule stability-related, post- translationally modified forms of α-tubulin, tyrosinated and acetylated α-tubulin, were not altered in stathmin-suppressed BE(2)-C cells (Fig. 3.5E). Similar results were seen in SH-SY5Y cells (data not shown). Therefore in normal cell culture conditions, suppressing stathmin did not alter the expression of a variety of cytoskeletal-related proteins.

114 Figure 3.5 Cytoskeletal protein expression in stathmin-suppressed BE(2)-C cells.

Representative western blots showing the expression of cytoskeletal proteins in stathmin-suppressed BE(2)-C cells 72hrs post-siRNA transfection. (A) Stathmin suppression in siRNA-transfected cells. Stathmin suppression did not alter the expression of MAP4 (B), total α- or β-tubulin or the β-tubulin isotypes (C), total actin,

β-actin or γ-actin (D) or tyrosinated or acetylated α-tubulin (E) compared to mock and control (Ctrl) siRNA-transfected cells. GAPDH was used as a loading control.

115

A B

Mock Ctrl STMN Mock Ctrl STMN GAPDH MAP4

Stathmin GAPDH

Mock Ctrl STMN Mock Ctrl STMN Total α-tubulin βI-tubulin βII-tubulin GAPDH βIII-tubulin Total β-tubulin βIV-tubulin GAPDH GAPDH

D E

Mock Ctrl STMN Ctrl STMN Tyrosinated Total actin α-tubulin β-actin GAPDH

γ-actin Acetylated α-tubulin GAPDH GAPDH

116 3.4 Stathmin does not influence neuroblastoma cell proliferation or viability

Microtubules depolymerise and re-polymerise to form the mitotic spindle during cell division. In some cell types, inhibiting stathmin expression induces a G2-M arrest which is associated with a reduction in cell proliferation and increased apoptosis

(Mistry, Bank et al. 2005; Singer, Ehemann et al. 2007; Wang, Dong et al. 2007; Singer,

Malz et al. 2009). To determine whether stathmin may be playing a role in regulating cell proliferation in neuroblastoma cells, stathmin siRNA-suppressed neuroblastoma cells were subjected to in vitro cell proliferation assays. Stathmin suppression did not markedly affect BE(2)-C cell proliferation (Fig. 3.6A-B). Only a small decrease in cell viability was observed in stathmin-suppressed BE(2)-C cells 48 hours post transfection

(87% in stathmin siRNA-treated cells compared to 91% in control cells), although this affect was not seen at other time points (Fig. 3.6C). To determine if this lack of effect on cell proliferation was cell-type specific, cell proliferation was also measured in stathmin-suppressed SH-SY5Y cells. As shown in Fig. 3.7, stathmin suppression did not alter cell proliferation in SH-SY5Y cells. In addition, stathmin suppression did not induce a G2-M delay in BE(2)-C cells (Fig. 3.8, non-treated cells) but rather a small but statistically significant decrease in G1 (61.91±0.42% in stathmin siRNA-treated cells compared to 64.3±0.69% in control cells; p<0.05) (Fig. 3.8). These results indicate that

117 Figure 3.6 Proliferation and viability of stathmin-suppressed BE(2)-C cells.

Proliferation of BE(2)-C cells transfected with stathmin (STMN) or control (Ctrl) siRNA. Cell proliferation was measured using resazurin solution (A) and by cell enumeration (B). The percentage viability of Ctrl and Stmn siRNA-transfected BE(2)-C cells (C). Data represent the mean ±SEM (error bars) of at least 3 independent experiments. Not significant (NS), **p<0.002.

118 A

119 Figure 3.7 Proliferation of stathmin-suppressed SH-SY5Y cells.

Proliferation of SH-SY5Y cells transfected with stathmin (STMN) and control (Ctrl) siRNA, as measured using resazurin solution. Data represent the mean ±SEM (error bars) of 3 independent experiments.

120 SH-SY5Y

121 Figure 3.8 Cell cycle profiles of paclitaxel-treated, stathmin-suppressed BE(2)-C cells.

SiRNA-treated BE(2)-C cells were treated with paclitaxel for 18 hours, harvested and

DNA content was analysed by flow cytometry. Representative profiles are shown. Data represent the mean ±SEM of 3 independent experiments. Significant differences between stathmin suppressed cells (STMN siRNA, bottom panel) and control cells (Ctrl siRNA, top panel) at the equivalent drug concentrations are indicated (bold, italic). * p<0.05, ** p<0.002, *** p<0.001.

122

1.12 ± 1.01 0.53 1.14 ± ± :43.8 :45.25 1 1 600 :12.65±0.06 -M:31.94 ± :11.77±0.73 -M:32.52 1 2 1 2 G S:11.69 ±0.31 S:11.69 G Sub-G S:10.61 ±0.22 S:10.61 G Sub-G G 0.2 0.94 ± ± 0.34 0.7 ± ± :48.08 :47.66 1 1 300 :18.75±0.72 -M:21.33 :17.87±0.26 -M:21.85 1 2 1 2 S:12.32 ±0.62 S:12.32 G Sub-G G Sub-G S:13.06 ±0.41 S:13.06 G G 0.94 0.97 ± ± 0.65 0.92 ± :48.56 1 :46.39 1 150 :27.08±0.21** -M:14.49 ± Paclitaxel (nmol/L) 1 2 :31.75±0.61 -M:12.53 1 2 S:10.38 ±0.52 S:10.38 G Sub-G G S:9.9 ±0.59 G Sub-G G DNA content DNA 0.14*** 0.14 ± 0.61 ± 0.21 ± :6.78 :4.81 1 1 :62.35±1.01 -M:13.2 :66.1±0.97 -M:12.87 ± 1 2 25 1 2 S:18.09 ±0.57 S:18.09 G G Sub-G S:16.57 ±0.75 S:16.57 G Sub-G G 1.12 0.42 0.21 0.47 ± ± ± ± :6.62 :6.87 1 1 :64.3±0.69 -M:12.52 :61.91±0.42* -M:13.01 1 2 1 2 Sub-G ±1.05 S:16.89 G ±0.53 S:18.49 G Sub-G G G

Untreated

Ctrl siRNA Ctrl siRNA STMN

123 stathmin does not play a major role in regulating cell proliferation in two independent neuroblastoma cell lines.

3.5 Stathmin does not markedly influence neuroblastoma drug sensitivity

Tubulin is a primary target of many anti-cancer drugs, termed the tubulin-binding agents (TBAs). TBAs include the microtubule stabilising drugs, paclitaxel and epothilone B, and the microtubule destabilising agent, vincristine (Jordan and Wilson

2004). Stathmin is an important microtubule destabilising protein that can influence cancer cell sensitivity to both TBAs and DNA-damaging agents [reviewed in (Rana,

Maples et al. 2008)]. To determine whether stathmin-suppressed neuroblastoma cells could undergo paclitaxel-induced G2-M cell cycle arrest, siRNA-transfected BE(2)-C cells were treated with this drug for 18 hours [approximate population doubling time for

BE(2)-C cells (Biedler, Roffler-Tarlov et al. 1978)] prior to cell cycle analyses (Fig.

3.8). While treatment with 300-600 nmol/L paclitaxel induced a potent mitotic block, stathmin suppression did not dramatically alter neuroblastoma cell cycle profiles in the presence of drug compared to controls (Fig. 3.8). Only a small but statistically significant increase in sub-G1 (6.78±0.14% in stathmin siRNA-treated cells compared to 4.81±0.14% in control cells; p<0.001) was observed in stathmin-suppressed cells treated with 25 nmol/L paclitaxel and a small but statistically significant decrease in G1

124 (27.08±0.21% in stathmin siRNA-treated cells compared to 31.75±0.61% in control cells; p<0.005) was observed in stathmin-suppressed cells treated with 150 nmol/L paclitaxel (Fig. 3.8). Therefore, stathmin suppression did not markedly affect the cell cycle profiles of paclitaxel-treated neuroblastoma cells compared to controls.

Short-term cytotoxicity assays were also conducted to evaluate stathmin’s influence on neuroblastoma drug sensitivity. SiRNA-mediated stathmin suppression did not alter

BE(2)-C cell sensitivity to paclitaxel (Fig. 3.9A), vincristine (Fig. 3.9B) or doxorubicin

(Fig. 3.9C), compared to control siRNA-treated cells. To ensure these results were not unique to the BE(2)-C cells, these assays were also conducted with SH-SY5Y cells (Fig.

3.10). However, similar to the BE(2)-C cells, stathmin suppression did not alter SH-

SY5Y cell sensitivity to paclitaxel (Fig. 3.10A), vincristine (Fig. 3.10B) or doxorubicin

(Fig. 3.10C).

Long-term clonogenic assays were then used to determine the replicative potential of stathmin-suppressed neuroblastoma cells following drug treatment (refer to Materials and Methods 2.12). Similar to short-term cytotoxicity assays, stathmin suppression did not significantly affect BE(2)-C sensitivity to the TBAs paclitaxel (Fig. 3.11A), vincristine (Fig. 3.11B) and epothilone B (Fig. 3.11C) or the DNA-damaging agent, etoposide (Fig. 3.12A). A small, although statistically significant, increase in sensitivity to the DNA-damaging agent cisplatin was observed in stathmin-suppressed BE(2)-C cells at one concentration (0.8μmol/L; p<0.05) (Fig. 3.12B).

125 Figure 3.9 Short-term drug sensitivity of stathmin-suppressed BE(2)-C cells.

The percentage viability of BE(2)-C cells transfected with stathmin (STMN) and control

(Ctrl) siRNA was assessed after 72 hour treatment with (A) paclitaxel, (B) vincristine or

(C) doxorubicin. Each point on the dose response curves is the mean of at least 3 independent experiments ±SEM (error bars).

126 A BE(2)-C

127 Figure 3.10 Short-term drug sensitivity of stathmin-suppressed SH-SY5Y cells.

The percentage viability of SH-SY5Y cells transfected with stathmin (STMN) and control (Ctrl) siRNA was assessed after 72 hour treatment with (A) paclitaxel, (B) vincristine or (C) doxorubicin. Each point on the dose response curves is the mean of at least 3 independent experiments ±SEM (error bars).

128 A SH-SY5Y

129 Figure 3.11 Long-term drug sensitivity of stathmin-suppressed BE(2)-C cells.

The replicative potential (expressed as a surviving fraction) of BE(2)-C cells treated with stathmin (STMN) and control (Ctrl) siRNA was assessed after 72 hour exposure to (A) paclitaxel, (B) vincristine or (C) epothilone B. Media was removed following drug treatment and fresh media replaced every 3 days until colonies were counted. Each point on the dose response curves is the mean of at least 3 independent experiments

±SEM (error bars).

130 A BE(2)-C

131 Figure 3.12 Long-term drug sensitivity of stathmin-suppressed BE(2)-C cells.

The replicative potential (expressed as a surviving fraction) of BE(2)-C cells treated with stathmin (STMN) and control (Ctrl) siRNA was assessed after 72 hour exposure to (A) etoposide, (B) cisplatin or (C) ZM447439. Media was removed following drug treatment and fresh media replaced every 3 days until colonies were counted. Each point on the dose response curves is the mean of at least 3 independent experiments ±SEM

(error bars). * p< 0.05.

132 A BE(2)-C

133 When calculating relative drug sensitivity (refer to Materials and Methods 2.12) the relative increase in sensitivity for stathmin-suppressed cells compared to control cells was only 1.15 fold (IC50 of 3.42μmol/L ± 0.46 for stathmin-suppressed cells compared to IC50 of 3.93 ± 0.44 for control cells) and not statistically significant (Fig. 3.12B).

Stathmin is phosphorylated on serine 16 by Aurora kinase B during mitosis (Gadea and

Ruderman 2006). The Aurora kinase B inhibitor ZM447439, which interferes with mitotic spindle assembly (Gadea and Ruderman 2006), was therefore used in clonogenic assays to examine whether stathmin suppression may interfere with neuroblastoma cell response to ZM447439. SiRNA-mediated stathmin suppression increased the sensitivity of BE(2)-C cells to this inhibitor, but only at the higher concentrations of 750nmol/L

(surviving fraction of 0.286±0.04 in stathmin siRNA-treated cells vs. 0.499±0.07 in control siRNA-treated cells; P<0.05) and 1000nmol/L (surviving fraction of

0.016±0.007 in stathmin siRNA-treated cells vs. 0.1±0.029 in control siRNA-treated cells; P<0.05) (Fig. 3.12C). The relative increase in sensitivity of stathmin-suppressed cells to ZM447439 compared to controls, although statistically significant was only 1.17 fold (IC50 of 0.69 ± 0.03 for stathmin-suppressed cells compared to IC50 of 0.81 ± 0.03 for control cells; p<0.05) (Fig. 3.12C). Therefore the changes observed in BE(2)-C sensitivity to both cisplatin and ZM447439 were only small and occurred at few concentrations, suggesting that these changes are unlikely to be biologically relevant.

To ensure that the effects of drug sensitivity were not specific to BE(2)-C cells, SH-

SY5Y cells were also subjected to the same clonogenic assays. However, similar to

BE(2)-C cells, stathmin suppression did not alter SH-SY5Y cell sensitivity to paclitaxel

(Fig. 3.13A), vincristine (Fig. 3.13B), epothilone B (Fig. 3.13C), etoposide (Fig.

3.14A), cisplatin (Fig. 3.14B) or ZM447439 (Fig. 3.14C).

134 Figure 3.13 Long-term drug sensitivity of stathmin-suppressed SH-SY5Y cells.

The replicative potential (expressed as a surviving fraction) of SH-SY5Y cells treated with stathmin (STMN) and control (Ctrl) siRNA was assessed after exposure to (A) paclitaxel, (B) vincristine or (C) epothilone B. Each point on the dose response curves is the mean of at least 3 independent experiments ±SEM (error bars).

135 A SH-SY5Y

136 Figure 3.14 Long-term drug sensitivity of stathmin-suppressed SH-SY5Y cells.

The replicative potential (expressed as a surviving fraction) of SH-SY5Y cells treated with stathmin (STMN) and control (Ctrl) siRNA was assessed after exposure to (A) etoposide, (B) cisplatin or (C) ZM447439. Each point on the dose response curves is the mean of at least 3 independent experiments ±SEM (error bars).

137 A SH-SY5Y

138 Collectively, these data demonstrate that stathmin does not dramatically alter the sensitivity of neuroblastoma cells to a range of chemotherapeutic agents.

139 3.6 Discussion

Stathmin is a major cytosolic phosphoprotein that destabilises microtubules and is over- expressed in a number of malignancies. In other cancers, stathmin over-expression is associated with more proliferative and drug-resistant disease. In neuroblastoma, stathmin expression and phosphorylation is altered in MYCN-amplified tumours compared to non-amplified tumours (Hailat, Strahler et al. 1990). However, the role of stathmin in neuroblastoma drug sensitivity and cell proliferation had not been addressed.

In this study, siRNA-mediated stathmin suppression did not significantly influence cell cycle progression, proliferation, viability or drug-sensitivity of neuroblastoma cells.

Similarly, it has been shown that stathmin suppression does not affect cell proliferation in a range of leukaemia cell lines (Holmfeldt, Brannstrom et al. 2006; Holmfeldt, Sellin et al. 2010) or drug sensitivity in ovarian cancer cell lines (Aoki, Oda et al. 2009). This is in contrast to other reports showing that inhibiting stathmin expression reduces cell proliferation, increase apoptosis and enhances drug sensitivity of osteosarcoma (Wang,

Dong et al. 2007), hepatocellular carcinoma (Singer, Ehemann et al. 2007) prostate cancer (Mistry, Bank et al. 2005) and breast cancer cells (Alli, Bash-Babula et al. 2002).

While it is unclear why stathmin does not appear to play a role in these functions in neuroblastoma cells, recent studies by Singer et al. have demonstrated that the highly homologous stathmin-like protein SCLIP mimics some of stathmin’s functions in lung cancer cells (Singer, Malz et al. 2009). Singer et al. found that individual suppression of stathmin and SCLIP reduced viability, proliferation, migration and invasion of lung cancer cells (Singer, Malz et al. 2009). Stathmin’s effect on lung cancer cell

140 proliferation was also confirmed in this study, as shown in Appendix Fig. 1.

Furthermore, Singer et al. found that combined suppression of stathmin and SCLIP expression further reduced cell migration compared to individual targeting of stathmin and SCLIP (Singer, Malz et al. 2009). This data suggests that these proteins are not functionally redundant in their role in cell migration (Singer, Malz et al. 2009).

However, combined suppression of stathmin and SCLIP did not lead to a further reduction in cell viability, proliferation or invasion, indicating that these proteins may be redundant at least for some functions in lung cancer cells (Singer, Malz et al. 2009).

It is therefore possible that the abundance of other stathmin-like proteins in neuronal cells may be responsible for maintaining some of stathmin’s functions in stathmin- depleted neuroblastoma cells. In particular, it has been proposed that SCG10, which is also highly expressed in neurons and shows a similar pattern of expression to stathmin during nervous system development (Grenningloh, Soehrman et al. 2004), may be able to mimic stathmin’s functions in neuronal cells. As anticipated, western blot analysis found that SCG10, a neuronal-specific protein, was highly expressed in neuroblastoma cell lines compared to the lung cancer Calu-6 cells (Appendix Fig. 2A). In contrast,

SCLIP, a ubiquitous protein, was expressed in both neuroblastoma cell lines [SH-SY5Y and BE(2)-C] and Calu-6 cells (Appendix Fig. 2B). Therefore it is possible that SCG10 and/or SCLIP may be able to compensate for some of stathmin’s functions, particularly in cell proliferation and drug response, in stathmin-depleted neuroblastoma cells.

Stathmin is transcriptionally repressed down-stream of wild-type p53 activation in both murine and human transformed and immortalised cell lines (Ahn, Murphy et al. 1999).

It has been proposed that stathmin expression may be elevated in p53 mutant cells and that suppression of stathmin could restore some wild-type p53 functions in these cells

(Alli, Yang et al. 2007). Indeed, RNAi-mediated suppression of stathmin in p53 mutant

141 cancer cells and p53-suppressed normal fibroblasts significantly reduced cell proliferation, increased apoptosis and led to a delay through G2 of the cell cycle, but had little effect on their wild-type p53 cell counterparts (Carney and Cassimeris 2010).

Experiments conducted in this chapter therefore explored the possibility that suppressing stathmin expression in mutant p53 BE(2)-C cells (Tweddle, Malcolm et al.

2001) may influence their response to drugs that activate the p53 pathway, such as the

DNA-damaging agents. However, suppressing stathmin in BE(2)-C cells did not significantly alter sensitivity to the DNA-damaging agents doxorubicin or etoposide, and only a small increase in sensitivity was observed with cisplatin at one drug concentration. Thus, no marked benefit was observed when targeting stathmin in combination with DNA-damaging agent treatment in mutant p53 neuroblastoma cells.

While it is unclear why suppressing stathmin in p53 mutant BE(2)-C cells did not enhance sensitivity to chemotherapy drugs, one possibility is that because stathmin suppression in BE(2)-C cells did not influence cell proliferation, cell cycle progression or increase apoptosis [an affect observed in other p53 mutant/p53-suppressed cells

(Carney and Cassimeris 2010)] this may have prevented these cells becoming more sensitive to chemotherapy drugs.

Studies have shown that inhibition of stathmin expression can sensitise a range of non- neuronal cancer cell lines to both DNA and microtubule-targeted agents [reviewed in

(Rana, Maples et al. 2008)]. However, the exact mechanisms underlying stathmin- mediated drug-sensitivity remain unclear. Iancu et al. propose that stathmin-mediated disruption of mitosis, via stathmin suppression, may explain the increased cell sensitivity to mitotic-disrupting agents (Iancu, Mistry et al. 2000). Indeed, this seems to be the case in hepatocellular, osteosarcoma and leukaemia cells in which stathmin suppression alone induces a G2-M arrest, reduces cell proliferation and upon treatment

142 with anti-mitotic agents enhances cell sensitivity (Iancu, Mistry et al. 2000; Singer,

Ehemann et al. 2007; Wang, Dong et al. 2007). Thus, given that stathmin suppression did not dramatically alter cell cycle progression or proliferation in either neuroblastoma cell line; this may explain why no dramatic changes in drug sensitivity were found.

In summary, although stathmin influences cell proliferation and drug resistance in other malignancies, stathmin does not appear to mediate proliferation or drug resistance in neuroblastoma. Further studies were therefore employed to determine stathmin’s role in this aggressive childhood cancer.

143 Chapter 4. Results II

144 Evaluation of stathmin’s role in retinoic acid-induced differentiation of

neuroblastoma cells

Stathmin is highly expressed in neuroblastoma and its expression and phosphorylation in response to retinoic acid differs between MYCN-amplified vs. non-amplified tumours

(Hailat, Strahler et al. 1990). In chapter 3, stathmin was shown not to influence neuroblastoma cell proliferation or drug sensitivity. Given that stathmin has been linked to cell differentiation in various cell types (as reviewed in Chapter 1, section 1.3.6), this raised the possibility that stathmin may influence neuroblastoma differentiation.

The extent of tumour differentiation serves as a strong prognostic factor in neuroblastoma. Some neuroblastoma tumours spontaneously differentiate to form benign ganglioneuromas while others contain poorly differentiated and highly proliferative neuroblasts (Brodeur 2003). Retinoids, such as 13-cis-retinoic acid, have proved particularly beneficial for inducing tumour cell differentiation in high risk neuroblastoma patients (Matthay, Villablanca et al. 1999; Pearson, Pinkerton et al.

2008). 13-cis-retinoic acid is a synthetic vitamin A derivative that becomes isomerised to the more thermodynamically stable all-trans retinoic acid (ATRA) and the unstable

9-cis retinoic acid (Armstrong, Redfern et al. 2005). ATRA functions to regulate gene expression by binding to ligand-activated nuclear retinoic acid receptors (RARs)

(Armstrong, Redfern et al. 2005) and by down regulating the MYCN transcription factor that ultimately leads to growth arrest and alterations in cell morphology (Thiele,

Reynolds et al. 1985; Armstrong, Redfern et al. 2005). More specifically, ATRA treatment of ‘N’ and some ‘I’ type neuroblastoma cells induces the formation of neurites (Thiele, Reynolds et al. 1985; Ross, Spengler et al. 1995).

145

Neurites are long and thin cylindrical processes that serve as precursor structures to axons and dendrites and are therefore indicative of cell differentiation into a mature neuron (Clagett-Dame, McNeill et al. 2006). Neurites are formed by coordinated reorganisation of the cell cytoskeleton, particularly the actin and microtubule networks.

However, the precise mechanisms underlying ATRA-induced cytoskeletal restructuring and neurite formation in neuroblastoma cells are unclear (Clagett-Dame, McNeill et al.

2006).

It has been well reported that stathmin expression and activity is altered during cell differentiation. Stathmin expression increased upon ATRA-induced differentiation of embryonal carcinoma cells into neuron-like cells (Maltman, Christie et al. 2009), decreased upon ATRA/granulocyte colony stimulating factor (G-CSF)-induced differentiation of promyeloid cells into neutrophils (Johnson, Jones et al. 1995), and stathmin expression and phosphorylation altered upon ATRA-induced differentiation of neuroblastoma cells (Hailat, Strahler et al. 1990). In PC12 cells (derived from a pheochromocytoma of the rat adrenal medulla), stathmin is required for NGF-induced neurite formation (Di Paolo, Pellier et al. 1996) but not pituitary adenylate cyclise- activating polypeptide (PACAP)-induced neurite formation (Dejda, Chan et al. 2010).

However, the role stathmin plays in neuroblastoma differentiation and neurite formation are unknown. The aims of this chapter were to examine stathmin’s role in ATRA- induced differentiation and neurite formation in neuroblastoma cells.

146 4.1 ATRA induces neurite formation in neuroblastoma cells

Retinoic acid induced-cell differentiation is characterised in part by alterations in cell morphology including neurite formation in some cell types. In particular, it has been well established that ATRA induces neurite formation in BE(2)-C and SH-SY5Y cells

[reviewed in (Clagett-Dame, McNeill et al. 2006)]. To confirm that ATRA induced neuroblastoma differentiation in BE(2)-C and SH-SY5Y cells, cells were treated with

10μmol/L ATRA, a non-toxic concentration shown to reduce proliferation and induce neurite formation in neuroblastoma cells (Thiele, Reynolds et al. 1985). Phase contrast images were taken of the cells 24-72 hours post ATRA treatment (Fig. 4.1). Short and thin processes (red arrowheads) were observed in BE(2)-C and SH-SY5Y cells after 24 hours exposure to ATRA, compared to vehicle-treated [ethanol (EtOH)] control cells

(Fig. 4.1). Longer processes (red arrows) were evident at 48 hours and became more prominent at 72 hours in both cell lines treated with ATRA (Fig. 4.1). This confirmed that ATRA was effective at inducing neurite formation in neuroblastoma cells.

147 Figure 4.1 ATRA induces neurite formation in neuroblastoma cells.

Treatment of BE(2)-C and SH-SY5Y cells with 10μmol/L ATRA induced the formation of long thin processes (neurites) compared to EtOH-treated (vehicle control) cells. Short cell extensions were present after 24 hours exposure (red arrowheads). Longer processes were evident after 48 and 72 hours exposure (red arrows).

148

BE(2)-C SH-SY5Y

EtOH

100μm

48 mol/L ATRA (hrs) ATRA μ mol/L 10

149 4.2 ATRA reduces stathmin protein expression in neuroblastoma cells

Stathmin expression alters during the differentiation of other cell types (Johnson, Jones et al. 1995; Maltman, Christie et al. 2009), though studies examining stathmin expression during neuroblastoma cell differentiation have been limited. It was therefore important to determine whether stathmin expression was altered in ATRA-treated neuroblastoma cells. Stathmin protein expression was examined 8, 24, 48 and 72 hours post ATRA treatment, the latter time points (24-72 hours) coinciding with neurite formation in 2 independent neuroblastoma cell lines, BE(2)-C and SH-SY5Y (Fig. 4.1).

As shown in Fig. 4.2, stathmin protein expression did not dramatically alter in ATRA- treated BE(2)-C cells (Fig. 4.2A) or SH-SY5Y cells (Fig. 4.2B) over this time course compared to controls. However, treatment of neuroblastoma cells for seven days with

ATRA reduced stathmin protein expression by 27.7±8.5% in BE(2)-C cells (Fig. 4.3A) and by 39.9±9.0% in SH-SY5Y cells (Fig. 4.3B) compared to controls. Therefore a reduction in stathmin protein expression only occurred after prolonged exposure to

ATRA and did not coincide with neurite outgrowth.

150 Figure 4.2 Short-term exposure to ATRA does not markedly alter stathmin protein expression in neuroblastoma cells.

Representative western blots and relative quantitation of stathmin protein expression in

BE(2)-C cells (A) and SH-SY5Y cells (B) treated with 10μmol/L ATRA for 8-72 hours.

Stathmin protein expression was normalised to GAPDH and stathmin protein expression in ATRA-treated cells (hatched bars) was calculated relative to EtOH- treated (vehicle control) cells (filled bars). Data represent the mean ± SEM of 3 independent experiments.

151

A BE(2)-C

ATRA (hr) EtOH 8244872 Stathmin

GAPDH

B SH-SY5Y ATRA (hr) EtOH 8244872 Stathmin

GAPDH

152 Figure 4.3 Long-term (7 day) exposure to ATRA reduces stathmin protein expression in neuroblastoma cells.

Representative western blots and relative quantitation of stathmin protein expression in

BE(2)-C cells (A) and SH-SY5Y cells (B) following treatment with 10μmol/L ATRA for 7 days. Stathmin protein expression was normalised to GAPDH and stathmin protein expression in ATRA-treated cells (hatched bars) was calculated relative to EtOH-treated

(vehicle control) cells (filled bars). Data represent the mean ± SEM of 3 independent experiments.

153

A BE(2)-C

EtOH ATRA Stathmin

GAPDH

B SH-SY5Y EtOH ATRA Stathmin

GAPDH

154 4.3 ATRA induces stathmin phosphorylation in neuroblastoma cells

Retinoic acid has been shown to induce stathmin phosphorylation in MYCN-amplified cells and reduce stathmin expression and phosphorylation in non-amplified neuroblastoma cells (Hailat, Strahler et al. 1990), suggesting a link between retinoic acid-induced differentiation and stathmin phosphorylation in neuroblastoma cells. To examine this phenotype, BE(2)-C (MYCN-amplified) and SH-SY5Y (non-amplified) cells were treated with 10μmol/L ATRA and the phosphorylation of stathmin’s four serine residues; Ser16, 25, 38 and 63, were examined by western blot.

Phosphorylation of Ser16 slightly increased in BE(2)-C cells following 8 and 24 hours exposure to ATRA, and then declined at 48 hours, compared to controls (Fig. 4.4A). In

SH-SY5Y cells, Ser16 phosphorylation increased by 1.5-fold at 24 hours and then declined after 48 hours exposure to ATRA, compared to controls (Fig. 4.4A).

Interestingly, this general trend, whereby phosphorylation gradually increased at 24 hours and then declined at 48 hours following ATRA treatment was also observed for

Ser25 (Fig. 4.4B), Ser38 (Fig. 4.5A) and Ser63 (Fig. 4.5B) in both cell lines. In particular, Ser63 phosphorylation increased by 1.5-fold at 8 hours, 2.2-fold at 24 hours and then declined after 48 hours exposure to ATRA in BE(2)-C cells compared to

155 Figure 4.4 ATRA alters the phosphorylation of stathmin’s Ser16 and Ser25 residues in neuroblastoma cells.

(A) Representative western blots and relative quantitation of Ser16 phosphorylation in

BE(2)-C and SH-SY5Y cells following treatment with 10μmol/L ATRA. Ser16 phosphorylation was normalised to GAPDH and Ser16 phosphorylation in ATRA- treated cells (hatched bars) was calculated relative to EtOH-treated (vehicle control) cells (filled bars). (B) Representative western blots and relative quantitation of Ser25 phosphorylation in BE(2)-C and SH-SY5Y cells following treatment with 10μmol/L

ATRA. Ser25 phosphorylation was normalised to GAPDH and Ser25 phosphorylation in ATRA-treated cells (hatched bars) was calculated relative to EtOH-treated cells

(filled bars). Data represent the mean ± SEM of 3 independent experiments.

156

A BE(2)-C SH-SY5Y ATRA (hrs) ATRA (hrs) EtOH8244872 EtOH 8244872 Ser16

GAPDH

B BE(2)-C SH-SY5Y ATRA (hrs) ATRA (hrs) EtOH8244872 EtOH 8244872 Ser25

GAPDH

157 Figure 4.5 ATRA alters the phosphorylation of stathmin’s Ser38 and Ser63 residues in neuroblastoma cells.

(A) Representative western blots and relative quantitation of Ser38 phosphorylation in

BE(2)-C and SH-SY5Y cells after treatment with 10μmol/L ATRA. Ser38 phosphorylation was normalised to GAPDH and Ser38 phosphorylation in ATRA- treated cells (hatched bars) was calculated relative to EtOH-treated (vehicle control) cells (filled bars). (B) Representative western blots and relative quantitation of Ser63 phosphorylation in BE(2)-C and SH-SY5Y cells after treatment with 10μmol/L ATRA.

Ser63 phosphorylation was normalised to GAPDH and Ser63 phosphorylation in

ATRA-treated cells (hatched bars) was calculated relative to EtOH-treated cells (filled bars). Data represent the mean ± SEM of 3 independent experiments.

158

A BE(2)-C SH-SY5Y

ATRA (hrs) ATRA (hrs) EtOH 8244872 EtOH 8244872 Ser38

GAPDH

B BE(2)-C SH-SY5Y ATRA (hrs) ATRA (hrs) EtOH 8244872 EtOH 8244872 Ser63

GAPDH

159 controls (Fig. 4.5B). Therefore ATRA induces stathmin phosphorylation in both the

MYCN-amplified BE(2)-C cells and the non-amplified SH-SY5Y cells at times coinciding with early-stage neurite formation. This data suggests that phosphorylation of stathmin, and potentially phosphorylation-inactivation of stathmin at 24 hours, may be important for ATRA-induced neurite formation in neuroblastoma cells.

4.4 Stathmin regulates ATRA- induced neurite formation

To determine whether stathmin was important for neurite formation in neuroblastoma cells, stathmin siRNA-suppressed BE(2)-C and SH-SY5Y cells were exposed to

10μmol/L ATRA for 72 hours. As shown in Fig. 4.6A, ATRA treatment induced neurite formation in BE(2)-C cells (red arrows) compared to EtOH-treated controls. However, stathmin suppression reduced the number of ATRA-induced neurites in BE(2)-C cells compared to ATRA-treated controls (Fig. 4.6A). This phenotype was quantified by counting the percentage of neurite positive cells (a cell bearing at least one neurite twice the length of the cell body) (Neubrand, Thomas et al. 2010) in each treatment group, as described in Materials & Methods section 2.15.

160 Figure 4.6 Stathmin suppression reduces ATRA-induced neurite formation in

BE(2)-C cells.

(A) Phase contrast images of control (Ctrl) and stathmin (STMN) siRNA-transfected

BE(2)-C cells treated with 10μmol/L ATRA or EtOH (vehicle control) for 72 hours.

Neurites are highlighted with red arrows. (B) The percentage neurite positive cells were counted for siRNA-treated cells following treatment with ATRA (hatched bars) or

EtOH (filled bars). Data represent the mean ± SEM of 3 independent experiments, * p<0.05, ** p<0.002.

161 A BE(2)-C EtOH ATRA Ctrl siRNA

100μm STMN siRNA

siRNA Ctrl STMN Ctrl STMN EtOH ATRA

162 ATRA significantly increased the percentage of neurite positive BE(2)-C cells

(18.62±2.47% in ATRA-treated Ctrl siRNA vs. 6.03±1.15% in EtOH-treated Ctrl siRNA cells, p<0.002) and stathmin suppression significantly reduced ATRA-induced neurite formation in BE(2)-C cells by 51% (9.06±1.83% in ATRA-treated STMN siRNA vs. 18.62±2.47% in ATRA-treated Ctrl siRNA cells, p<0.05) (Fig. 4.6B). This phenotype was also observed in SH-SY5Y cells (Fig. 4.7). ATRA treatment induced neurite formation in SH-SY5Y cells (red arrows) compared to EtOH-treated controls

(Fig. 4.7A) and stathmin suppression reduced the number of ATRA-induced neurites in

SH-SY5Y cells (Fig. 4.7A). This phenotype was confirmed by counting neurite positive cells. ATRA significantly increased the percentage of neurite positive SH-SY5Y cells

(18.22±1.20% in ATRA-treated Ctrl siRNA cells vs. 7.12±0.70% in EtOH-treated Ctrl siRNA cells, p<0.002) and stathmin suppression significantly reduced ATRA-induced neurite formation in SH-SY5Y cells by 38% (11.24± 0.84% in ATRA-treated STMN siRNA cells vs. 18.22±1.20% in ATRA-treated Ctrl siRNA cells, p<0.01) (Fig. 4.7B).

Therefore, stathmin is required for ATRA-induced neurite formation in neuroblastoma cells.

163 Figure 4.7 Stathmin suppression reduces ATRA-induced neurite formation in SH-

SY5Y cells.

(A) Phase contrast images of control (Ctrl) and stathmin (STMN) siRNA-transfected

SH-SY5Y cells treated with 10μmol/L ATRA or EtOH (vehicle control) for 72 hours.

Neurites are highlighted with red arrows. (B) The percentage neurite positive cells were counted for siRNA-treated cells following treatment with ATRA (hatched bars) or

EtOH (filled bars). Data represent the mean ± SEM of 3 independent experiments, ** p<0.01.

164 A SH-SY5Y EtOH ATRA Ctrl siRNA

100μm STMN siRNA

siRNA Ctrl STMN Ctrl STMN EtOH ATRA

165 4.5 Stathmin does not influence

ATRA-induced growth arrest

Stathmin suppression inhibits neurite formation in neuroblastoma cells indicating that stathmin was potentially influencing neuroblastoma cell differentiation. To further examine this phenotype, cell proliferation (which declines upon cell differentiation) was examined in stathmin siRNA-suppressed neuroblastoma cells following treatment with

ATRA. As shown in Fig. 4.8A, ATRA significantly reduced the proliferation of both control (4.1x105 cells in ATRA-treated Ctrl siRNA cells vs. 12.1x105 cells in EtOH- treated Ctrl siRNA cells, p<0.005) and stathmin-suppressed BE(2)-C cells (3.6x105 cells in ATRA-treated STMN siRNA cells vs. 10.4x105 cells in EtOH-treated STMN siRNA cells, p<0.0001) (Fig. 4.8A). However, cell proliferation did not differ between ATRA- treated control and ATRA-treated stathmin-suppressed BE(2)-C cells (Fig. 4.8A).

Similarly, ATRA significantly reduced cell proliferation of control (7.5x105 cells in

ATRA-treated Ctrl siRNA cells vs. 11.2x105 cells in EtOH-treated Ctrl siRNA cells, p<0.05) and stathmin-suppressed SH-SY5Y cells (6x105 cells in ATRA-treated STMN siRNA cells vs. 9.4x105 cells in ethanol-treated STMN siRNA cells, p<0.05) but there were no differences in cell numbers between ATRA-treated control and ATRA-treated stathmin-suppressed SH-SY5Y cells (Fig. 4.8B). Therefore, stathmin suppression does not prevent ATRA-induced growth inhibition in neuroblastoma cells. This indicates that stathmin’s role in neuroblastoma cell differentiation may be restricted to neurite formation.

166 Figure 4.8 Stathmin suppression does not prevent ATRA-induced growth arrest in neuroblastoma cells.

Cell counts of control (Ctrl) and stathmin (STMN) siRNA-transfected BE(2)-C (A) and

SH-SY5Y (B) cells following treatment with 10μmol/L ATRA (hatched bars) or EtOH

(filled bars) for 72 hours. Data represent the mean ± SEM of 3 independent experiments, * p<0.05, *** p<0.001.

167 A BE(2)-C

siRNA Ctrl STMN Ctrl STMN EtOH ATRA B SH-SY5Y

siRNA Ctrl STMN Ctrl STMN EtOH ATRA

168 4.6 Stathmin does not influence

ATRA-induced alterations in differentiation marker expression

Alterations in the expression of neuron-specific enolase (NSE), the stathmin-like protein

SCG10, βIII-tubulin and the low molecular weight microtubule-associated protein 2

(MAP2) isoform, MAP2c, have all been linked to neuronal cell differentiation

(Weisshaar, Doll et al. 1992; Yang, Li et al. 2008). To further clarify stathmin’s role in neuroblastoma differentiation, the expression of these proteins were examined in stathmin siRNA-suppressed neuroblastoma cells following ATRA treatment (Fig. 4.9).

However, treatment of neuroblastoma cells with ATRA for 72 hours did not alter

SCG10 expression in either control or stathmin-suppressed BE(2)-C cells (Fig. 4.9A) and only a slight decrease in expression was observed in ATRA-treated control vs.

EtOH-treated control SH-SY5Y cells (Fig. 4.9A). Furthermore, 72 hour ATRA treatment did not alter MAP2c expression in either control or stathmin-suppressed

BE(2)-C or SH-SY5Y cells (Fig. 4.9A). Only a slight increase in βIII-tubulin expression was observed in 72 hour ATRA-treated BE(2)-C cells (Fig.4.8B) while no differences in

βIII-tubulin expression were observed in ATRA-treated SH-SY5Y cells (Fig. 4.9B).

These results demonstrate that short-term ATRA treatment (72 hours) did not markedly influence neuronal differentiation marker expression in neuroblastoma cells.

169 Figure 4.9 Short term ATRA treatment does not alter the expression of neuronal differentiation markers.

(A) Representative western blots of stathmin, SCG10 and MAP2c protein expression in control (Ctrl) and stathmin (STMN) siRNA-transfected BE(2)-C and SH-SY5Y cells following 72 hour 10μmol/L ATRA treatment. GAPDH was used as a loading control.

(B) Representative western blots and quantitation of βIII-tubulin protein expression in

Ctrl and STMN siRNA-transfected BE(2)-C and SH-SY5Y cells following 72 hour

ATRA treatment. βIII-tubulin protein expression in ATRA-treated cells (hatched bars) and EtOH-treated cells (filled bars) were normalised to GAPDH. Data represent the mean ± SEM of 3 independent experiments.

170

A BE(2)-C SH-SY5Y

EtOH ATRA (72hr) EtOH ATRA (72hr) siRNA Ctrl STMN Ctrl STMN Ctrl STMN Ctrl STMN

Stathmin

GAPDH

SCG10

MAP2c

GAPDH

B BE(2)-C SH-SY5Y EtOH ATRA (72hr) EtOH ATRA (72hr) siRNA Ctrl STMN Ctrl STMN Ctrl STMN Ctrl STMN βIII-tubulin

GAPDH

171 Therefore neuroblastoma cells were treated with ATRA for a longer time period to determine whether stathmin was influencing ATRA-induced neuroblastoma differentiation.

To conduct this experiment, stable stathmin shRNA-expressing SH-SY5Y cells were developed to ensure that stathmin suppression was maintained during long-term (10 day) exposure to ATRA. The ‘N’ (neuronal) type SH-SY5Y cells were deemed more suitable for these experiments as ‘neuronal’ differentiation markers were being examined. SH-SY5Y cells were transiently transfected with four individual stathmin shRNA, a control (non-effective) shRNA and empty vector constructs. Stathmin sequences 2 and 3 (hereafter designated STMN Seq.2 and STMN Seq.3) were the most effective at reducing stathmin protein expression compared to control shRNA and empty vector-transfected cells (Fig. 4.10A). These shRNA constructs where subsequently used to make stable shRNA-expressing SH-SY5Y cells (as described in

Materials & Methods section 2.3) (Fig. 4.10B). Of note, no marked differences in the expression of the stathmin-like proteins, SCG10 and SCLIP, were observed in STMN

Seq.2 and STMN Seq.3 cells compared to control cells (Fig. 4.10B). To confirm the phenotype observed in siRNA-transfected cells, neurites were counted in 10 day ATRA- treated, shRNA-expressing SH-SY5Y cells. Treatment of control cells with 10μmol/L

ATRA for 10 days significantly increased the percentage of neurite positive cells compared EtOH-treated control cells (18.3% ATRA-treated Ctrl shRNA cells vs. 2.2% in EtOH-treated Ctrl shRNA cells, p<0.0005) (Fig. 4.11A). Furthermore, the percentage of neurite positive cells in ATRA-treated STMN Seq.3 cells was significantly reduced compared to control cells (8.3% in STMN Seq.3 cells compared to 18.3% in ATRA- treated Ctrl shRNA cells, p<0.005) (Fig. 4.10B).

172 Figure 4.10 Development of stathmin shRNA-expressing SH-SY5Y cells.

(A) Representative western blot of stathmin protein expression in mock, empty vector

(pRS), control (Ctrl) shRNA and stathmin (STMN) shRNA transiently-transfected SH-

SY5Y cells. Four individual shRNA sequences for stathmin (Seq.1, Seq.2, Seq.3 and

Seq.4.), targeting different regions of the stathmin gene, were screened. STMN shRNA

Seq.2 and Seq.3 gave the best stathmin suppression compared to mock, pRS and Ctrl shRNA (B). Representative western blots showing stathmin, SCG10 and SCLIP expression in stable stathmin shRNA (STMN Seq.2 and STMN Seq.3) and Ctrl shRNA- expressing SH-SY5Y cells.

173 A SH-SY5Y

STMN shRNA Ctrl Mock pRS shRNA Seq.1 Seq.2 Seq.3 Seq.4

Stathmin

GAPDH

B STMN STMN STMN STMN Ctrl Seq.2 Seq.3 Ctrl Seq.2 Seq.3

SCG10 SCLIP Stathmin Stathmin

GAPDH GAPDH

174 Figure 4.11 ShRNA-mediated stathmin suppression reduces ATRA-induced neurite formation in SH-SY5Y cells.

(A) Phase contrast images of control shRNA (Ctrl), stathmin sequence 2 shRNA

(STMN Seq.2) and stathmin sequence 3 shRNA (STMN Seq.3) SH-SY5Y cells treated with 10μmol/L ATRA or EtOH (vehicle control) for 10 days. Neurites are highlighted with red arrowheads. (B) The percentage neurite positive cells were counted for Ctrl and stathmin shRNA-suppressed cells following treatment with ATRA (hatched bars) or

EtOH (filled bars) cells. Data represent the mean ± SEM of 3 independent experiments,

** p<0.01, *** p<0.001, NS (not significant).

175

A EtOH ATRA (10 days) Ctrl STMN Seq.2

STMN Seq.3 100μM B

Ctrl STMN STMN Ctrl STMN STMN Seq.2 Seq.3 Seq.2 Seq.3 EtOH ATRA (10 days)

176 Although there was no significant difference between the percentages of neurite positive cells in ATRA-treated STMN Seq.2 cells (16.3%) and Ctrl shRNA-expressing cells

(18.3%), this may have been a result of greater stathmin suppression in STMN Seq.3 cells compared to STMN Seq.2 cells.

The expression of the neuronal differentiation markers in shRNA-expressing SH-SY5Y cells were then examined following 10 days treatment with ATRA (Fig. 4.12-4.14). As shown in previous experiments, stathmin protein expression was markedly reduced in

SH-SY5Y cells treated with ATRA for a long time period (10 days) (Fig. 4.12). More importantly, ATRA treatment further reduced stathmin expression in STMN Seq.2 and

STMN Seq.3 cells in a similar manner to ATRA-treated control shRNA cells (Fig.

4.12). ShRNA-mediated stathmin suppression did not alter the levels of SCG10 (Fig.

4.13A), MAP2c (Fig. 4.13B), βIII-tubulin (Fig. 4.14A) or NSE (Fig. 4.14B) in SH-

SY5Y cells following ATRA treatment as compared to control shRNA cells.

Collectively, these results show that stathmin suppression did not significantly alter neuroblastoma cell response (as determined by differentiation marker expression) to long-term ATRA treatment.

177 Figure 4.12 Long-term ATRA treatment reduces stathmin expression in SH-SY5Y cells.

Representative western blot and quantitation of stathmin protein expression in control

(Ctrl) shRNA and stathmin shRNA (STMN Seq.2 and STMN Seq.3) SH-SY5Y cells treated with EtOH (filled bars) or 10μmol/L ATRA (hatched bars) for 10 days.

Stathmin expression was normalised to GAPDH. Data represent the mean ± SEM of 3 independent experiments. **p<0.01, *** p<0.001.

178

EtOH ATRA (10 days) STMN STMN STMN STMN Ctrl Seq.2 Seq.3 Ctrl Seq.2 Seq.3 Stathmin

GAPDH

STMN STMN STMN STMN Ctrl Seq.2 Seq.3 Ctrl Seq.2 Seq.3 EtOH ATRA (10 days)

179 Figure 4.13 SCG10 and MAP2c expression in stathmin shRNA-suppressed SH-

SY5Y cells following long-term treatment with ATRA.

(A) Representative western blot and quantitation of SCG10 protein expression in control (Ctrl) and stathmin shRNA (STMN Seq.2 and STMN Seq.3) SH-SY5Y cells treated with EtOH (filled bars) or 10μmol/L ATRA (hatched bars) for 10 days. SCG10 expression was normalised to GAPDH. (B) Representative western blot and quantitation of MAP2c protein expression in Ctrl shRNA, stathmin shRNA (STMN Seq.2 and

STMN Seq.3) SH-SY5Y cells treated with EtOH (filled bars) or ATRA (hatched bars) for 10 days. MAP2c expression was normalised to GAPDH. Data represent the mean ±

SEM of 3 independent experiments. NS (not significant).

180

A EtOH ATRA(10 days) STMN STMN STMN STMN Ctrl Seq.2 Seq.3 Ctrl Seq.2 Seq.3 SCG10 GAPDH

B EtOH ATRA(10 days) STMN STMN STMN STMN Ctrl Seq.2 Seq.3 Ctrl Seq.2 Seq.3 MAP2c GAPDH

181 Figure 4.14 βIII-tubulin and NSE expression in stathmin shRNA-suppressed SH-

SY5Y cells following long-term treatment with ATRA.

(A) Representative western blot and quantitation of βIII-tubulin protein expression in control (Ctrl) shRNA and stathmin shRNA (STMN Seq.2 and STMN Seq.3) SH-SY5Y cells treated with EtOH (filled bars) or 10μmol/L ATRA (hatched bars) for 10 days.

βIII-tubulin expression was normalised to GAPDH. (B) Representative western blot and quantitation of NSE protein expression in Ctrl shRNA and stathmin shRNA (STMN

Seq.2 and STMN Seq.3) SH-SY5Y cells treated with EtOH (filled bars) or ATRA

(hatched bars) for 10 days. NSE expression was normalised to GAPDH. Data represent the mean ± SEM of 3 independent experiments. NS (not significant).

182

A EtOH ATRA(10 days) STMN STMN STMN STMN Ctrl Seq.2 Seq.3 Ctrl Seq.2 Seq.3 βIII-tubulin GAPDH

B EtOH ATRA(10 days) STMN STMN STMN STMN Ctrl Seq.2 Seq.3 Ctrl Seq.2 Seq.3 NSE GAPDH

183 4.7 Discussion

Retinoic acid is currently used to induce tumour differentiation in high risk neuroblastoma patients. However the precise mechanisms regulating retinoic acid- induced differentiation, particularly neurite outgrowth, in neuroblastoma cells are not well understood. In this chapter it has been demonstrated that stathmin is important for

ATRA-induced neurite formation in neuroblastoma cells; a function potentially mediated by stathmin’s regulation of the cell cytoskeleton.

In accordance with previous studies (Johnson, Jones et al. 1995; Rubin, French et al.

2003; Yoshie, Kashima et al. 2008), stathmin expression declines in differentiated neuroblastoma cells. However, this effect only occurs after prolonged exposure (7-10 days) and not after short-term exposure (72 hours) to ATRA, during which neurites are formed in neuroblastoma cells. The observation that ATRA altered stathmin phosphorylation during short-term exposure suggests that stathmin may be playing a role in neurite formation. Indeed, stathmin suppression significantly inhibited ATRA- induced neurite outgrowth in neuroblastoma cells, which highlights a novel role for stathmin in this process. Although the exact mechanism by which stathmin regulates neurite formation is unclear, it is likely mediated by ATRA-induced changes in stathmin phosphorylation.

Stathmin’s four serine residues are phosphorylated by multiple kinases, many of which have been implicated in cell differentiation and neurite formation (Clagett-Dame,

McNeill et al. 2006). Stathmin’s Ser16 residue is a known target of Pak1 and PKA

[reviewed in (Holmfeldt, Sellin et al. 2009)]. PKA also phosphorylates Ser63, while a

184 variety of MAPKs phosphorylate Ser25 (Holmfeldt, Sellin et al. 2009). Previous studies have shown that ATRA-induced activation of PKA, Pak1 and the MAPKs, extracellular signal-regulated kinase (ERK) and c-Jun N-terminal kinase (JNK), are important for neurite formation in neuroblastoma cells (Shea, Beermann et al. 1992; Niles 2004;

Clagett-Dame, McNeill et al. 2006). More specifically, ATRA-induced activation of

Pak1, JNK and PKA are important for neurite outgrowth in SH-SY5Y cells, while ERK activation is critical for neurite outgrowth in BE(2)-C cells [reviewed in (Clagett-Dame,

McNeill et al. 2006)]. Thus it appears that the multiple signalling pathways required for neuritogenesis in neuroblastoma cells may converge at stathmin, regulating stathmin’s activity and potential functional role in this process.

Neuritogenesis is initiated by the formation of actin-rich lamellipodia at the cell periphery [reviewed in (Poulain and Sobel 2009)]. Small microtubule bundles then extend from the soma, protrude into the lamellipodia, align in parallel and lengthen to form the neurite (Poulain and Sobel 2009). The formation of these structures requires coordinated remodelling of the actin and microtubule networks of which are mediated by the small RhoGTPases [reviewed in (Clagett-Dame, McNeill et al. 2006)]. Rac1 and

Cdc42 activation promotes neurite formation while RhoA activation blocks neurite formation and aids neurite retraction (Clagett-Dame, McNeill et al. 2006). Interestingly, numerous studies have linked stathmin to both the Rac1 and RhoA signalling pathways in various cell types (Daub, Gevaert et al. 2001; Wittmann, Bokoch et al. 2004; Watabe-

Uchida, John et al. 2006; Tanaka, Hamano et al. 2007; Takahashi and Suzuki 2009). In particular, Takahashi et al. found that EGF-induced activation of Rac1-Pak1 induced stathmin phosphorylation and microtubule-mediated transport of the actin-regulatory protein WAVE2 (Takahashi and Suzuki 2009). More importantly, inhibition of stathmin

185 expression in EGF-stimulated breast cancer cells reduced WAVE2 transport and the formation of lamellipodia (Takahashi and Suzuki 2009). Therefore given that lamellipodia are precursor structures to neurites, stathmin may be playing a role in lamellipodia formation in neuroblastoma cells. Moreover, the increases in stathmin phosphorylation that occur at all four serine residues after 24 hours exposure to ATRA indicate that phosphorylation-inactivation of stathmin may be critical for microtubule stability and neurite extension during this time. Future studies could address whether the total pool of stathmin is inactivated during this time or whether inactivation of stathmin is localised at neurites.

This chapter has demonstrated that ATRA alters stathmin expression and phosphorylation and that stathmin suppression blocked ATRA-induced neurite formation in neuroblastoma cells. However, stathmin did not influence ATRA-induced growth inhibition or changes in differentiation marker expression, indicating a restricted role for stathmin in neuritogenesis. Interestingly it has been reported that the signalling pathways important for neurite formation in SH-SY5Y cells are distinct from those that regulate cell growth arrest during differentiation (Eggert, Ikegaki et al. 2000). As such the signalling pathways implicated to regulate stathmin phosphorylation and neurite outgrowth in ATRA-treated neuroblastoma cells may be separate from those required for growth arrest. Thus it is possible that stathmin may only be required for the morphological alterations, and not growth arrest, induced by ATRA in neuroblastoma cells.

186

Chapter 5. Results III

187 Examination of stathmin’s role in anchorage-independent growth,

migration and invasion of neuroblastoma cells

The extent of metastatic spread determines neuroblastoma disease stage and risk stratification, and ultimately impacts overall survival. Patients with unresectable tumours that have spread across the midline, or tumours that have disseminated to regional lymph nodes, bone, bone marrow and other organs are classified as advanced stage (stages 3 and 4) and have a poor prognosis (Maris, Hogarty et al. 2007). In other cancers, high stathmin expression correlates with increased tumour size (Brattsand

2000; Yuan, Jeng et al. 2006) and metastatic disease (Saal, Johansson et al. 2007;

Belletti, Nicoloso et al. 2008; Kuo, Wang et al. 2009; Xi, Rui et al. 2009; Hsieh, Huang et al. 2010; Jeon, Han et al. 2010). In other neural crest-derived tumours of the adrenal medulla, stathmin is more abundant in malignant and metastatic tumours compared to the benign tumours and normal adrenal tissue (Sadow, Rumilla et al. 2008; Lin, Chen et al. 2011). Furthermore, suppressing stathmin expression reduced the tumourigenicity of osteosarcoma cells in vivo (Wang, Dong et al. 2007), cell migration and invasion of multiple cancer cells in vitro (Baldassarre, Belletti et al. 2005; Liang, Choi et al. 2008;

Singer, Malz et al. 2009; Gan, Guo et al. 2010; Jeon, Han et al. 2010) and metastatic spread of transgenic FoxM1b Arf-null hepatocellular carcinoma cells in vivo (Park,

Gusarova et al. 2011). These findings indicate that stathmin may be regulating tumour growth and metastasis in neuroblastoma. To examine these phenotypes, stathmin siRNA-suppressed neuroblastoma cells were subjected to in vitro assays to evaluate anchorage-independent growth, cell migration and invasion.

188 5.1 Stathmin does not influence neuroblastoma anchorage- independent growth

The ability of cancer cells to grow in an anchorage-independent manner is a surrogate marker of tumourigenic potential. In order to determine whether stathmin was playing a role in neuroblastoma anchorage-independent growth, stathmin siRNA-suppressed neuroblastoma cells were grown in soft agar. Stathmin suppression did not significantly reduce colony number in either BE(2)-C (Fig. 5.1A) or SH-SY5Y cells (Fig. 5.1B).

Similarly, stathmin suppression did not affect colony size in BE(2)-C (Fig. 5.1C) or SH-

SY5Y cells (Fig. 5.1D). This demonstrates that stathmin does not markedly influence neuroblastoma anchorage-independent growth.

5.2 Stathmin does not influence neuroblastoma 2D cell migration

Metastasis is a complex process involving many steps including cell migration and invasion. Cell migration and invasion can be studied in vitro using monolayer wound- healing, and transwell migration and invasion assays (Valster, Tran et al. 2005).

189 Figure 5.1 Stathmin does not regulate neuroblastoma anchorage-independent growth.

Colony formation (%) of BE(2)-C (A) and SH-SY5Y (B) cells transfected with control

(Ctrl) or stathmin (STMN) siRNA. Representative BE(2)-C colony dishes are shown below graph (A). Colony size (perimeter in μm) of BE(2)-C (C) and SH-SY5Y (D) cells transfected with Ctrl or STMN siRNA. Representative BE(2)-C colonies are shown below graph (C). Data represent the mean ±SEM (error bars) of 4 independent experiments. Not significant (NS).

190

ACBE(2)-C BE(2)-C

Ctrl STMN Ctrl STMN siRNA siRNA

500μm

B SH-SY5Y D SH-SY5Y

Ctrl STMN Ctrl STMN siRNA siRNA

191 To determine whether stathmin was playing a role in 2 dimensional (2D) neuroblastoma cell migration (migration of cells on a flat surface), stathmin siRNA-suppressed neuroblastoma cells were subjected to monolayer wound-healing assays (Fig. 5.2A). As shown in Fig. 5.2A, stathmin suppression did not alter the ability of BE(2)-C cells to move into the wound site compared to control cells. These results were confirmed by calculating the percentage wound recovery over time for both control and stathmin- suppressed BE(2)-C cells (as described in Chapter 2, section 2.17) (Fig. 5.2B).

Therefore stathmin does not markedly influence 2D neuroblastoma cell migration.

5.3 Stathmin mediates chemotactic- induced neuroblastoma cell migration & invasion

Although stathmin did not play a role in 2D migration, this did not exclude the possibility that stathmin regulated other types of motility in neuroblastoma cells. Cancer cells are exposed to gradients in growth factors and cytokines (chemo-attractants) in their surrounding extra-cellular matrix (ECM) that help stimulate cell migration, invasion and metastasis (Condeelis, Singer et al. 2005). These conditions can be mimicked in vitro using transwell chemotaxis migration and invasion assays (Valster,

Tran et al. 2005).

192 Figure 5.2 Stathmin does not regulate 2D neuroblastoma cell migration.

(A) Representative images of stathmin (STMN) and control (Ctrl) siRNA-transfected

BE(2)-C cells at time 0 (0 hr), 8, 24 and 48 hours post scratch. The black dotted lines represent the original wound site and the red lines highlight the movement of cells towards the wound. (B) The percentage wound recovery was calculated for both STMN and Ctrl siRNA-transfected cells at each time point. Data represent the mean of 3 independent experiments ±SEM (error bars).

193

A 0hr 8hr 24hr 48hr Ctrl siRNA STMN siRNA

194 To determine whether stathmin was influencing chemotactic-induced neuroblastoma cell migration and invasion, stathmin siRNA-suppressed neuroblastoma cells were subjected to transwell migration and invasion assays. Serum, PDGF and collagen IV were used as chemo-attractants in the lower chamber; conditions that were optimised in

BE(2)-C cells (Appendix Fig. 3). As shown in Fig. 5.3, stathmin suppression significantly reduced the migratory ability of BE(2)-C cells by 47% (migration index of

3.06±0.23 for stathmin siRNA-treated cells vs. 5.81±0.08 for control cells; p<0.001)

(Fig. 5.3A) and SH-SY5Y cells by 44% (migration index of 2.65±0.25 for stathmin siRNA-treated cells vs. 4.72±0.65 for control cells; p<0.05, n=3) (Fig. 5.3B). Moreover, stathmin suppression significantly inhibited the invasion of BE(2)C cells through

Matrigel™ by 62% (invasion index of 8.55±2.16 for stathmin siRNA-treated cells vs.

22.76±3.54 for control cells; p<0.05) (Fig. 5.3C) and SH-SY5Y cells by 56% (invasion index of 8.72±1.64 for stathmin siRNA-treated cells vs. 19.61±2.48 for control cells vs. cells; p<0.05) (Fig. 5.3D). Therefore stathmin is important for chemotactic-induced migration and invasion of neuroblastoma cells.

195 Figure 5.3 Stathmin regulates chemotactic-induced neuroblastoma cell migration and invasion.

Chemotactic-induced migration of stathmin (STMN) and control (Ctrl) siRNA- transfected BE(2)-C (A) and SH-SY5Y cells (B). Chemotactic-induced cell invasion of

STMN and Ctrl siRNA-transfected BE(2)-C (C) and SH-SY5Y cells (D). Data represent the mean of 3 independent experiments ±SEM (error bars). * p<0.05,*** p<0.001.

196

A BE(2)-CB SH-SY5Y

Ctrl STMN Ctrl STMN siRNA siRNA

C BE(2)-C D SH-SY5Y

Ctrl STMN Ctrl STMN siRNA siRNA

197 5.4 Stathmin regulates neuroblastoma cell morphology

The cell cytoskeleton plays a major role in cell migration and invasion, and changes in cell morphology indicate a perturbed cell cytoskeleton. Therefore in order to understand the potential mechanisms by which stathmin mediates neuroblastoma cell migration and invasion, the morphology of stathmin siRNA-suppressed neuroblastoma cells were examined. In contrast to controls, stathmin-suppressed BE(2)-C cells appeared to have more short and thin neurite-like extensions (Fig. 5.4A). The cell cytoskeleton was then visualised in serum-starved BE(2)-C cells. Stathmin suppressed cells displayed long and thin actin-rich projections (white arrows) compared to controls (Fig. 5.4B). In addition, small increases in microtubule density were observed in stathmin suppressed cells compared to controls (Fig. 5.4B, inset image). Therefore stathmin expression levels appear to influence morphology and cytoskeletal structures in neuroblastoma cells.

198 Figure 5.4 Stathmin regulates neuroblastoma cell morphology.

(A) Phase-contrast images of control (Ctrl) and stathmin (STMN) siRNA-treated BE(2)-

C cells. (B) Black and white confocal images of actin filaments and microtubules, including magnified microtubules (inset images), in Ctrl and STMN siRNA-treated

BE(2)-C cells. The merged colour image shows microtubules in green, actin filaments in red and nuclei in blue. White arrows point to long and thin actin-rich cell extensions.

199

A Ctrl siRNA STMN siRNA

100 μm

B Ctrl siRNA STMN siRNA

10 μm Actin filaments 10 μm Microtubules Merge

200 5.5 Stathmin regulates neuroblastoma tubulin polymer levels

Stathmin regulates tubulin monomer-heterodimer partitioning during interphase and total polymer levels in various cell types (Alli, Yang et al. 2007; Holmfeldt, Sellin et al.

2009; Ringhoff and Cassimeris 2009). To determine whether stathmin was influencing tubulin polymerisation in neuroblastoma cells, stathmin siRNA-suppressed BE(2)-C cells were grown in normal culture medium (basal) or serum-starved (serum-free) and then treated with culture medium containing 25ng/mL PDGF (10% serum+PDGF) for 1 hour. These conditions were based on those used in the chemotaxis assays. Cells were then harvested and the soluble (S) and polymerised (P) tubulin fractions were determined by western blot analysis (Fig. 5.5A). A trend for increased tubulin polymer levels in stathmin-suppressed cells compared to control cells was observed in all conditions (Fig. 5.5A). However, this effect was only significantly different in serum- starved (serum-free) conditions (Fig. 5.5B). Stathmin suppression significantly increased tubulin polymer levels by 29% compared to control cells under serum-starved conditions (percentage polymerised tubulin 38.04±2.07 for stathmin siRNA-treated cells vs. 27.16±2.26 for control cells; p<0.05) (Fig. 5.5B). Therefore stathmin regulates tubulin polymer levels in neuroblastoma cells, particularly under serum-starved conditions.

201 Figure 5.5 Stathmin regulates tubulin polymer levels in neuroblastoma cells.

(A) Stathmin (STMN) and control (Ctrl) siRNA-transfected BE(2)-C cells were maintained in normal cell culture medium (basal), serum-starved (serum-free) or serum- starved and then treated with growth factors (25ng/mL PDGF+10%FCS in DMEM) for

1 hour. Soluble (S) and polymerised (P) tubulin fractions were separated and α-tubulin and stathmin expression was analysed by western blot. (B) The percentage of tubulin polymer levels in STMN and Ctrl siRNA-transfected cells are shown for all conditions.

* p< 0.05, not significant (NS).

202

A 10%serum Basal Serum-free +PDGF siRNA Ctrl STMN Ctrl STMN Ctrl STMN SSPPSSPPSSPP α-tubulin

Stathmin

% polymer 27 32 27 38 28 38

siRNA Ctrl STMN Ctrl STMN Ctrl STMN Basal Serum-free 10%serum +PDGF

203 5.6 Stathmin regulates cofilin phosphorylation

The small RhoGTPases are important regulators of the cell cytoskeleton. Activation of the

Rho-Rho associated coiled-coil forming protein kinase (Rho-ROCK) signalling pathway regulates actin remodelling, through the phosphorylation of cofilin and myosin light chain

(MLC) (Torka, Thuma et al. 2006), and also microtubule dynamics (Kadir, Astin et al.

2011), both of which are important for cell migration. To further elucidate the mechanism underlying stathmin’s regulation of neuroblastoma cell migration and invasion, the expression and phosphorylation of cofilin, a major downstream effector protein of the

Rho-ROCK signalling pathway (Maekawa, Ishizaki et al. 1999), was examined in stathmin siRNA-suppressed neuroblastoma cells (Fig. 5.6). Interestingly, a marked increase in cofilin phosphorylation was observed in stathmin-suppressed BE(2)-C cells in both serum-free and serum-treated conditions compared to control cells (Fig. 5.6A, arrows). Furthermore, while cofilin phosphorylation declined in controls cells 30 minutes post serum-treatment, cofilin phosphorylation was maintained up to 60 minutes post serum-treatment in stathmin-suppressed cells (Fig. 5.6B). Total levels of cofilin were not altered. This indicates that stathmin can regulate cofilin phosphorylation, and potentially the cycling of activity following growth factor (serum) treatment, in neuroblastoma cells.

204 Figure 5.6 Stathmin regulates cofilin phosphorylation in neuroblastoma cells.

Stathmin (STMN) and control (Ctrl) siRNA-transfected BE(2)-C cells were serum- starved (serum-free) or serum-starved and then treated with 10% serum

(10%FCS/DMEM) for the indicated time points. (A) The expression of stathmin, cofilin and phospho-cofilin (cofilin-P) were determined by western blot. Increased cofilin phosphorylation is indicated (arrows). (B) The phosphorylation of cofilin was quantified for Ctrl (unfilled bars) and STMN siRNA-treated cells (black bars) at each time point. The phosphorylation of cofiln was normalised to GAPDH and calculated relative to Ctrl siRNA cells in serum-free media.

205 A 10% serum Serum-free 5 min 15 min 30 min 60 min siRNA Ctrl STMN Ctrl STMN Ctrl STMN Ctrl STMNCtrl STMN Stathmin

GAPDH

Cofilin-P

Cofilin

GAPDH

206 5.7 Stathmin does not influence

LIMK expression

Cofilin is phosphorylated by various kinases including testis-specific protein kinase 1 and

2 (TESK1 and TESK2), Nck-interacting kinase (NIK)-related kinase (NRK)/NIK-like embryo-specific kinase (NESK) (NRK/NESK) and the double LIM domain-containing kinases; LIM kinase 1 (LIMK1) and LIM kinase 2 (LIMK2) (Scott and Olson 2007).

However, due to the tissue-specific expression of the other kinases in the testes and skeletal muscle, the LIM kinases predominantly phosphorylate cofilin in other tissues

(Scott and Olson 2007). Given that cofilin phosphorylation was markedly altered in stathmin-suppressed neuroblastoma cells, the phosphorylation of LIMK1 and LIMK2 were also examined (Fig. 5.7). However, in contrast to cofilin, there were no marked alterations in LIMK1 or LIMK2 protein expression or phosphorylation (Fig. 5.7).

5.8 Stathmin regulates cofilin and

MLC phosphorylation via ROCK

Cofilin is a major actin remodelling protein and downstream effector of the Rho-ROCK signalling pathway (Maekawa, Ishizaki et al. 1999). In order to determine whether stathmin was regulating cofilin phosphorylation via the Rho-ROCK signalling pathway,

Y-27632, a compound that specifically inhibits the kinase activity of p160ROCK

(ROCKI) and ROKα/Rho-kinase (ROCKII), was employed to block ROCK signalling

207 Figure 5.7 Stathmin does not regulate LIM kinase expression or phosphorylation in neuroblastoma cells.

Stathmin (STMN) and control (Ctrl) siRNA-treated BE(2)-C cells were serum-starved

(serum-free) or serum-starved and treated with 10% serum (10%FCS/DMEM) for the indicated time points. LIMK1, phospho-LIMK1 (LIMK1-P), LIMK2, phospho-LIMK2

(LIMK2-P) and stathmin protein expression were determined by western blot.

208

10% serum Serum-free 5 min 15 min 30 min 60 min Ctrl STMN Ctrl STMN Ctrl STMN Ctrl STMN Ctrl STMN Stathmin GAPDH

LIMK1-P

LIMK1 GAPDH

LIMK2-P LIMK2 GAPDH

209 (Narumiya, Ishizaki et al. 2000). Treatment of serum-starved BE(2)-C cells with

10μmol/L Y-27632 reduced cofilin phosphorylation in both control and stathmin- suppressed BE(2)-C cells (Fig. 5.8). Notably, the levels of cofilin phosphorylation in stathmin-suppressed cells were returned to that of untreated control cells (Fig. 5.8). No further reduction in cofilin phosphorylation was observed in control or stathmin- suppressed cells treated with more than 10μmol/L Y-27632 (Fig. 5.8). This data indicates that cofilin phosphorylation is only partially influenced by ROCK and that an alternate pathway exists by which stathmin regulates cofilin phosphorylation.

In further support stathmin’s role in Rho-ROCK signalling, the phosphorylation of myosin light chain (MLC), a direct target of ROCK (Gutjahr, Rossy et al. 2005), was increased in serum-starved, stathmin-suppressed BE(2)-C cells (Fig. 5.8). Treatment of control and stathmin-suppressed cells with 10μmol/L Y-27632 ablated MLC phosphorylation (Fig. 5.8). This indicates that MLC phosphorylation is ROCK-dependent and that stathmin’s regulation of MLC phosphorylation may be solely dependent on

ROCK in neuroblastoma cells.

210 Figure 5.8 Stathmin suppression-induced increases in cofilin and MLC phosphorylation are reversed by treatment with the ROCK inhibitor, Y-27632.

Stathmin (STMN) and control (Ctrl) siRNA-transfected BE(2)-C cells were left untreated (serum-free) or were pre-treated with 10-40μmol/L of the ROCK inhibitor, Y-

27632, for 1 hour prior to incubation in serum-free alone (serum-free) or serum-free media containing the equivalent concentrations of Y-27632. The expression of stathmin, cofilin, phospho-cofilin (cofilin-P), myosin light chain (MLC) and phospho-MLC

(MLC-P) were determined by western blot. Increases in cofilin and MLC phosphorylation are indicated (arrows).

211

Y-27632

Serum-free 10μM20μM40μM Ctrl STMN Ctrl STMN CtrlSTMN Ctrl STMN Stathmin

Cofilin-P

Cofilin

α-tubulin

MLC-P

MLC

α-tubulin

212 5.9 Stathmin regulates neuroblastoma cell invasion via

ROCK

To determine whether stathmin’s role in neuroblastoma cell invasion is mediated through

ROCK, stathmin-suppressed BE(2)-C cells were treated with the ROCK inhibitor Y-

27632 and subjected to invasion assays. As shown in Fig. 5.9, treatment of stathmin- suppressed cells with the ROCK inhibitor returned the level of cell invasion back to that of untreated control cells [invasion index (% of control) is 53.37±7.3 for stathmin siRNA- treated cells vs. 93.87±9.84 for stathmin siRNA+Y-27632-treated cells; p<0.05].

Therefore the reduction in cell invasion induced by stathmin suppression is inhibited by

Y-27632. This data suggests that stathmin is regulating neuroblastoma cell invasion via

ROCK.

213 Figure 5.9 Stathmin suppression-induced reduction of cell invasion is reversed by the ROCK inhibitor, Y-27632.

Stathmin (STMN) (hatched bars) and control (Ctrl) siRNA-treated cells (filled bars) were left untreated (minus symbols) or treated with 10μmol/L Y-27632 (plus symbols) and subjected to transwell assays. Results are expressed as a percentage of untreated

Ctrl siRNA cells (% of control). Data represent the mean of 5 independent experiments

±SEM (error bars). * p< 0.05.

214 siRNA Ctrl Ctrl STMN STMN Y-27632 - + - +

215 5.10 Discussion

The Rho-ROCK signalling pathway facilitates cell migration by altering the activity of various cytoskeletal proteins, including the actin-severing protein cofilin (Maekawa,

Ishizaki et al. 1999). The cycling of cofilin between inactive (phosphorylated) and active

(dephosphorylated) states is considered a “key determinant of invasive and metastatic potential” in cancer cells (Scott, Hooper et al. 2010). Therefore stathmin’s regulation of cofilin phosphorylation in neuroblastoma cells may underpin stathmin’s role in neuroblastoma cell migration and invasion. To support this theory, treatment of stathmin- suppressed neuroblastoma cells with the ROCK inhibitor (Y-27632) returned cell invasion and cofilin phosphorylation back to that of untreated control cells, suggesting that stathmin plays a role in regulating cell invasion and cofilin phosphorylation via

ROCK. However, the increase in cofilin phosphorylation induced by stathmin suppression was not ablated by Y-27632, suggesting that stathmin’s regulation of cofilin phosphorylation is not limited this pathway (Fig. 5.10). Examination of the LIM kinases found no marked differences in LIMK1 or LIMK2 expression or phosphorylation in stathmin-suppressed neuroblastoma cells. These findings may be attributed to the other

RhoGTPase signalling pathways that regulate LIM kinase activity (Scott and Olson 2007)

(Fig. 5.10). Another possibility is that stathmin is regulating the activity of the cofilin phosphatases, slingshot (SSH) and chronophin (CIN) (Fig. 5.10) (Scott and Olson 2007).

In addition, the expression and phosphorylation of paxillin (another downstream target of the ROCK signalling pathway) were also unchanged in stathmin-suppressed BE(2)-C cells (Appendix Fig. 4A) indicating that stathmin’s regulation of the ROCK signalling pathway may not be linear and/or may be limited.

216 Figure 5.10 Schematic diagram of stathmin’s proposed regulation of neuroblastoma cell migration and invasion.

Extracellular stimuli activate the RhoGTPase signalling pathways (Rho, Cdc42, Rac) that lead to the phosphorylation of the LIM domain-containing kinases, LIM kinase 1 and LIM Kinase 2 (LIMK1/2) via Rho associated coiled-coil forming protein kinase

(ROCK), Pak 1, Pak2, Pak4 and myotonic dystrophy kinase-related Cdc42-binding kinase alpha (MRCKα) (Scott and Olson 2007). Activation of the Rho-ROCK signalling pathway induces LIMK1/2 and myosin light chain (MLC) phosphorylation. ROCK is inhibited by Y-27632. Cofilin is phosphorylated by LIMK1/2. Cofilin is dephosphorylated by chronophin (CIN) and slingshot (SSH). Based on our data, we propose that stathmin destabilises microtubules and may block ROCK signalling, which is implicated in regulating amoeboid-like cell motility (Torka, Thuma et al. 2006).

Stathmin may also be regulating the dephosphorylation of cofilin (?). Subsequently, stathmin may assist neuroblastoma cells to move in the alternate, mesenchymal-like

(elongated, proteolysis-dependent), mode by regulating both the microtubule and actin networks.

217

Extracellular stimuli: Serum-starvation/growth factors

Y-27632 Rho Cdc42/Rac Cdc42

Stathmin ROCK Pak1,2,4 MRCKα

P P LIMK1/2 MLC

P Cofilin Cofilin

? Stathmin SSH/CIN P

Microtubule destabilisation Actin reorganisation

Mesenchymal-like cell invasion

218 While the mechanisms underlying stathmin’s regulation of cofilin activity are unclear, they most likely involve the RhoGTPase signalling pathways of which stathmin is an important downstream target (Daub, Gevaert et al. 2001; Wittmann, Bokoch et al. 2004;

Takahashi and Suzuki 2009; Takahashi, Tanaka et al. 2010). In breast cancer cells, EGF- activation of Rac1-Pak1 induced stathmin phosphorylation and stimulated microtubule- mediated transport of the actin-regulatory protein WAVE2 (Takahashi and Suzuki 2009;

Takahashi, Tanaka et al. 2010). Furthermore, silencing stathmin inhibited WAVE2 transport and subsequently reduced lamellipodia formation and cell migration in breast cancer cells (Takahashi and Suzuki 2009; Takahashi, Tanaka et al. 2010). In other cell types, direct interactions between p27kip1 and stathmin regulate microtubule stability, cell morphology, motility, lipid raft trafficking and adhesion-dependent RhoA activity

(Belletti, Pellizzari et al. 2010). These studies highlight an important link between stathmin, the RhoGTPases, cytoskeletal restructuring and cell migration. In addition, recent studies have demonstrated that drug-induced alterations in microtubule stability influence cofilin expression and phosphorylation in various cell types (Balasubramani,

Nakao et al. 2010; Belletti, Pellizzari et al. 2010). Thus stathmin’s regulation of tubulin polymerisation has the potential to influence several feed-back loops that occur between microtubules and the RhoGTPases (Wittmann and Waterman-Storer 2001). As a consequence, stathmin may directly or indirectly influence the activity of numerous microtubule and actin restructuring proteins, including cofilin and MLC, in neuroblastoma cells.

Epithelial-mesenchymal transition (EMT) is a complex cellular program thought to play a major role in metastasis. During EMT, epithelial cells with apical-basal polarity, lose cell- cell contacts and switch to a mesenchymal morphology (front-back end polarity) in order

219 to invade the ECM (Yang and Weinberg 2008). In a recent study, interactions between stathmin and Siva1 (an apoptosis-related protein), and their subsequent effect on microtubule stability, have been implicated in EMT in breast cancer cells (Li, Jiang et al.

2011). Moreover, stathmin suppression increased E-cadherin expression and the phosphorylation of focal adhesion kinase, and decreased vimentin expression in breast cancer cells; phenotypes associated with reduced EMT (Li, Jiang et al. 2011). Of note, only subtle differences in neural-cadherin (N-cadherin) and vimentin expression

(Appendix Fig. 4B), and no alterations in the expression or phosphorylation of the focal adhesion adaptor protein paxillin, were observed in stathmin siRNA-suppressed neuroblastoma cells (Appendix Fig. 4A). However, these results do not exclude the possibility that stathmin is playing a role in EMT, or regulating a particular type of cell movement, in neuroblastoma cells. On the contrary, results from the ROCK inhibitor experiments suggest that stathmin is blocking ROCK signalling, which is thought to mediate amoeboid-like (rounded shape, proteolysis-independent) cell movements (Fig.

5.10) (Torka, Thuma et al. 2006). Subsequently, stathmin may promote the alternate mesenchymal (elongated shape, proteolysis-dependent)-like mode of cell motility in neuroblastoma cells (Fig. 5.10). Interestingly, in mesenchymal-like sarcoma cells, direct interactions between p27kip1 and stathmin (that inhibit microtubule destabilization) are important for maintaining cell shape and a reduced migratory phenotype (Baldassarre,

Belletti et al. 2005). However, increasing stathmin activity in these cells by overexpressing stathmin or expressing a Ser16 phosphorylation-impaired (Q18>E) mutation, promoted amoeboid-like cell invasion in vitro and higher metastatic potential in vivo

(Belletti, Nicoloso et al. 2008). Interestingly, increasing stathmin activity in sarcoma cells did not influence 2D migration (Belletti, Nicoloso et al. 2008); a finding that supports the theory that the signalling pathways and cytoskeletal restructuring that occurs in cells in

220 2D differs greatly to that of cells in chemotaxis assays (Torka, Thuma et al. 2006). This phenomenon provides potential explanation as to why stathmin did not affect 2D migration but significantly influenced chemotactic-induced migration and invasion of neuroblastoma cells. Collectively, these results suggest that stathmin’s regulation of the neuroblastoma cell cytoskeleton may determine the mode in which neuroblastoma cells migrate and invade.

In this chapter it has been demonstrated that stathmin mediates chemotactic-induced neuroblastoma cell migration and invasion and that these functions may be attributed to changes in the cell cytoskeleton downstream of the Rho-ROCK signalling pathway.

These results indicate that stathmin may be influencing neuroblastoma metastasis; a phenotype that will be examined in chapter 6.

221 Chapter 6. Results IV

222 Evaluation of stathmin’s contribution to the metastatic phenotype of

neuroblastoma

In vitro studies in Chapter 5 revealed a potential role for stathmin in neuroblastoma metastasis. To fully address whether stathmin was influencing this phenotype, an in vivo model of metastasis was required.

Xenograft models are widely used in pre-clinical studies to evaluate therapeutic efficacy. Two examples of these models are the ‘experimental’ metastasis model, in which tumour cells are intravenously injected into mice (Zhang, Smith et al. 2009), and the ‘orthotopic’ model whereby tumour cells are injected into the anatomical site of the primary tumour (Khanna, Jaboin et al. 2002). The experimental metastasis model determines the ability of tumour cells, already in circulation, to arrest and grow at different sites. However the caveat for this model is that many steps in the metastatic cascade, including tumour-stromal interactions that occur within the primary tumour organ are excluded (Langley and Fidler 2011). Thus the orthotopic model is favoured when wanting to examine the entire metastatic process. Orthotopic neuroblastoma mouse models in particular simulate many clinical features of human neuroblastoma and have proved invaluable for studying tumourigenesis and metastasis in this malignancy (Khanna, Jaboin et al. 2002; Almgren, Henriksson et al. 2004; Henriksson,

Almgren et al. 2004; Meier, Muhlethaler-Mottet et al. 2007; Nevo, Sagi-Assif et al.

2008; Fuchs, Christofferson et al. 2009).

Bioluminescent imaging (BLI) is one way in which researchers can non-invasively monitor tumour growth and metastasis in small animals. Cancer cells genetically modified to express the Photinus pyralis (firefly) luciferase gene are detected in vivo via

223 injection of the luciferase substrate (luciferin) into mice. Luciferin becomes oxidised in the presence of adenosine triphosphate and oxygen upon which a visible light is emitted

(peak emission wavelength of 560nm) and is detected externally (O'Neill, Lyons et al.

2009). BLI is a highly specific (only metabolically active cells are detected), sensitive and rapid (requires short acquisition times) imaging modality with high through-put capabilities; making this system an extremely valuable tool for pre-clinical oncology research (O'Neill, Lyons et al. 2009).

Based on the results from Chapter 5 that demonstrated that stathmin is mediating chemotactic-induced neuroblastoma cell migration and invasion, the aim of this final results chapter was to determine whether stathmin was regulating neuroblastoma metastasis. In order to investigate this, stathmin shRNA-suppressed/luciferase- expressing neuroblastoma cells were developed and utilised in a clinically-relevant, orthotopic neuroblastoma mouse model.

6.1 Stathmin shRNA\luciferase- expressing neuroblastoma cells

Luciferase-expressing SK-N-BE(2) cells [SK-N-BE(2)/TGL] (described in Chapter 2, section 2.19) were transfected with four individual stathmin shRNA constructs and a control shRNA construct (Fig. 6.1A). Stathmin shRNA sequences 2 and 3 (hereafter referred to as Stmn Seq.2SH and Stmn Seq.3SH respectively) gave the best stathmin suppression compared to the mock and control shRNA (hereafter referred to as CtrlSH) transfected cells (Fig. 6.1A). These constructs were used to make stably-transfected cells (Fig. 6.1B).

224 Figure 6.1 Development of shRNA/luciferase-expressing neuroblastoma cells.

(A) Luciferase-expressing SK-N-BE(2)/TGL cells were transiently transfected with control (Ctrl) and stathmin (STMN) shRNA constructs. Four individual stathmin shRNA constructs (Seq.1, Seq.2, Seq.3 and Seq.4), targeting different regions of the stathmin gene, were screened. STMN shRNA sequence 2 and sequence 3 gave the most effective stathmin suppression compared to mock and Ctrl shRNA-transfected cells. (B)

SK-N-BE(2)/TGL were stably transfected with Ctrl shRNA (CtrlSH), STMN sequence 2

(Stmn Seq.2SH) and STMN sequence 3 (Stmn Seq.3SH) constructs. Stathmin suppression was maintained up to 4 weeks out of selection media. GAPDH was used as a loading control.

225

A SK-N-BE(2)/TGL

STMN shRNA Ctrl Mock shRNA Seq.1 Seq.2 Seq.3 Seq.4

Stathmin

GAPDH

B Stmn Stmn Seq.2 CtrlSH SH Seq.3SH

Stathmin

GAPDH

226 Stathmin suppression was maintained in Stmn Seq.2SH and Stmn Seq.3SH cells up to 4 weeks out of selection media compared to CtrlSH cells (Fig. 6.1B). Immunofluorescence microscopy also confirmed stathmin suppression in Stmn Seq.2SH and Stmn Seq.3SH cells compared to CtrlSH cells (Fig. 6.2A). Importantly, the invasion phenotype was maintained in stathmin-suppressed SK-N-BE(2)/TGL cells (Fig. 6.2B). Stathmin suppression inhibited the invasion of SK-N-BE(2)/TGL cells through Matrigel™ by

40% for Stmn Seq.2SH (invasion index percentage of control 59.63%±3.72) and 44% for Stmn Seq. 3SH (invasion index percentage of control 56.03%±6.47) (Fig. 6.2B). In addition, the bioluminescent signal was similar for CtrlSH, Stmn Seq.2SH and Stmn

Seq.3SH cells (Fig. 6.3). Thus the shRNA-expressing SK-N-BE(2)/TGL cells were deemed suitable for use in the orthotopic neuroblastoma mouse model.

6.2 Stathmin does not influence neuroblastoma tumour engraftment or growth

The orthotopic neuroblastoma mouse model accurately simulates human neuroblastoma development. This model was therefore developed and characterised for the first time at the Children’s Cancer Institute Australia specifically to evaluate stathmin’s role in neuroblastoma metastasis. SCID-Beige mice were injected with 2 x 106 shRNA- expressing SK-N-BE(2)/TGL cells into their left adrenal fat pad (the most common site for primary neuroblastoma tumours), as described in Materials & Methods, section 2.21.

227 Figure 6.2 ShRNA-mediated stathmin suppression reduces neuroblastoma cell invasion.

(A) Confocal images showing stathmin (red) and GFP expression (green) in CtrlSH,

Stmn Seq.2SH and Stmn Seq.3SH SK-N-BE(2)/TGL cells. In addition to luciferase, these cells express GFP as the TGL construct harbours the Aequorea victoria green fluorescent protein (GFP) gene (see Fig. 2.2 in Materials & Methods). The merged image shows an overlay of stathmin and GFP expression. (B) Chemotactic-induced invasion of CtrlSH, Stmn Seq.2SH and Stmn Seq.3SH SK-N-BE(2)/TGL cells. Invasion index of Stmn Seq.2SH and Stmn Seq.3SH cells are calculated as a percentage of CtrlSH cells. Data represent the mean of 3 independent experiments ±SEM (error bars).

228

A GFPStathmin Merge Ctrl SH Stmn Seq.2 SH

10μm Stmn Seq.3

CtrlSH Stmn Stmn Seq.2SH Seq.3SH

229 Figure 6.3 ShRNA-expressing SK-N-BE(2)/TGL cells express similar levels of luciferase.

(A) A representative luminescent image of serially diluted CtrlSH, Stmn Seq.2SH and

Stmn Seq.3SH SK-N-BE(2)/TGL cells. The coloured scale bar indicates the level of bioluminescence emitted from the cells, as measured in photons/second/centimetre squared/steradian (p/sec/cm2/sr). (B) Quantification of bioluminescence emitted from serially diluted CtrlSH, Stmn Seq.2SH and Stmn Seq.3SH cells confirmed that each cell population expressed similar levels of luciferase, measured as total flux

(photons/second).

230

A Cell number

0.8

CtrlSH 0.6

Stmn Seq.2SH 0.4

Stmn Seq.3SH 0.2

x107 p/sec/cm2/sr B

231 This cell number was chosen based on previous publications using this model (Khanna,

Jaboin et al. 2002; Meier, Muhlethaler-Mottet et al. 2007; Fuchs, Christofferson et al.

2009). Any animals with obvious leakage from the injection site (observed during surgery) or 1 day post-surgery, as determined by BLI (Appendix Fig. 5), were eliminated from the study.

To determine whether stathmin was contributing to neuroblastoma tumour growth, tumour burden was measured weekly by BLI (Fig. 6.4). Representative pseudocolor luminescent images overlaid on photographic images (hereafter referred to as luminescent images) of mice from each group are shown in Fig. 6.4A. Mice were imaged on their left lateral and ventral sides in order to visualise tumour growth each week (Fig. 6.4A). Quantitation of bioluminescence (total flux in photons/second) revealed that suppressing stathmin expression did not influence tumour burden over time compared to controls (Fig. 6.4B). Similarly, representative grey scale and luminescent images of primary tumours show that stathmin does not influence neuroblastoma tumour growth (Fig. 6.5A). This phenotype was confirmed by measuring primary tumour volume (Fig. 6.5B) and weight (Fig. 6.5C). This indicates that stathmin is not important for neuroblastoma tumour growth in vivo.

Stathmin expression was then examined in the primary tumours to ensure that stathmin suppression was maintained for the duration of the experiment. As shown in Fig. 6.6A, stathmin mRNA was suppressed by 79% in Stmn Seq.2SH primary tumours (relative stathmin gene expression of 0.15±0.02 in Stmn Seq.2SH vs. 0.68±0.15 in CtrlSH, p<0.01) and 82% in Stmn Seq. 3SH primary tumours (relative stathmin gene expression of 0.13±

0.02 in Stmn Seq.3SH vs. 0.68±0.15 in CtrlSH, p<0.05) compared to controls. A western blot shows stathmin protein expression in 3 primary tumours from each group (Fig.

6.6B).

232 Figure 6.4 Stathmin does not influence neuroblastoma tumour burden.

(A) Representative luminescent images of CtrlSH, Stmn Seq.2SH and Stmn Seq.3SH mice

1 and 4 weeks after inoculation. Mice were imaged on their left lateral (L) and ventral

(V) sides. The coloured scale bar indicates the levels of bioluminescence emitted from mice, as measured in photons/second/centimetre squared/steradian (p/sec/cm2/sr). (B)

Tumour burden, as determined by measuring total flux (photons/second) from the left lateral sides, was monitored over time for CtrlSH, Stmn Seq.2SH and Stmn Seq.3SH mice.

Data represent the mean of ≥13 mice for each group ±SEM (error bars).

233

A CtrlSH Stmn Seq.2SH Stmn Seq.3SH LV LV LV 1.2

1 1.0

0.8

LV LV LV0.6

0.4 Weeks post Weeks injection 4

0.2

x109 p/sec/cm2/sr B

234 Figure 6.5 Stathmin does not influence neuroblastoma tumour growth.

(A) Representative grey scale (top images) and pseudocolor luminescent images (lower images) of CtrlSH, Stmn Seq.2SH and Stmn Seq.3SH primary tumours. The coloured scale bar indicates the levels of bioluminescence emitted from tumours, as measured in photons/second/centimetre squared/steradian (p/sec/cm2/sr). Graphs show primary tumour volumes (B) and weights (C) for each group. Data represent the mean (black bars) of at least 10 primary tumours for each group ±SEM (error bars).

235

Ctrl Stmn Seq.2 Stmn Seq.3 A SH SH SH 3.0

2.5

2.0 1cm 1.5

1.0

0.5

x109 p/sec/cm2/sr B

CtrlSH Stmn Stmn Seq.2SH Seq.3SH C

CtrlSH Stmn Stmn Seq.2SH Seq.3SH

236 Figure 6.6 Stathmin suppression was maintained in the primary neuroblastoma tumours.

(A) Stathmin mRNA levels in CtrlSH, Stmn Seq.2SH and Stmn Seq.3SH primary tumours.

Stathmin mRNA was normalised to GAPDH mRNA. Data represent the mean ±SEM

(error bars) of at least 3 primary tumours for each group. (B) Representative western blot showing stathmin expression in 3 individual CtrlSH, Stmn Seq.2SH and Stmn

Seq.3SH primary tumours. (C) Quantitation of stathmin protein expression in CtrlSH,

Stmn Seq.2SH and Stmn Seq.3SH primary tumours. Stathmin protein expression was normalised to GAPDH protein expression. Data represent the mean of 3 primary tumours for each group ±SEM (error bars). * p<0.05, ** p< 0.01.

237

CtrlSH Stmn Seq.2SH Stmn Seq.3SH B CtrlSH Stmn Seq.2SH Stmn Seq.3SH Mouse # 12333 12 12 Stathmin GAPDH

CtrlSH Stmn Seq.2SH Stmn Seq.3SH

238 Stathmin protein was suppressed by 85% in Stmn Seq.2SH primary tumours (relative stathmin protein expression of 0.33±0.14 in Stmn Seq.2SH vs. 2.19±0.54 in CtrlSH, p<0.05) and 85% in Stmn Seq. 3SH primary tumours (relative stathmin protein expression of 0.32± 0.04 in Stmn Seq.3SH vs. 2.19±0.54 in CtrlSH, p<0.05) compared to controls (Fig. 6.6C). IHC staining also confirmed stathmin suppression in a Stmn Seq.

3SH primary tumour compared to a CtrlSH primary tumour (Fig. 6.7).

6.3 Metastatic spread in the orthotopic neuroblastoma mouse model

The purpose of this study was to determine whether stathmin was influencing neuroblastoma metastasis. In order to examine this phenotype, it was imperative that metastasis could be accurately quantified. Based on previous studies in humans and mice, neuroblastoma cells spread to the lymph nodes, liver, lungs, spleen, bone/ bone marrow and kidneys as a result of direct extension from the primary tumour, locoregional or haematogenous spread (via the blood system) (Khanna, Jaboin et al.

2002; Fuchs, Christofferson et al. 2009). These sites were therefore examined for metastatic spread using BLI and IHC (as described in Chapter 2, section 2.22).

A representative luminescent image of a CtrlSH left kidney is shown in Fig. 6.8A. IHC staining for luciferase revealed luciferase-expressing neuroblastoma cells inside the left kidney (Fig. 6.8B, black arrowhead). However, the left kidney became encapsulated by the primary tumour. Although the kidney was removed from the primary tumour, it was

239 Figure 6.7 Luciferase and stathmin expression in neuroblastoma primary tumours.

Immunostaining of CtrlSH and Stmn Seq.3SH primary tumours. Tumour cells were confirmed by H&E and luciferase staining. No staining was present in the isotype control (rabbit IgG control) which confirmed the specificity of the secondary antibody

(see Materials & Methods section 2.23). Luciferase expression was similar in both

CtrlSH and Stmn Seq.3SH primary tumours. Stathmin was suppressed in the Stmn

Seq.3SH primary tumour compared to CtrlSH.

240 CtrlSH Stmn Seq.3SH H&E

100μm Isotype control Luciferase Stathmin

241 Figure 6.8 Bioluminescent imaging and histological analysis of the left kidney.

(A) A representative luminescent image of the left kidney from a CtrlSH mouse. The coloured scale bar indicates the amount of bioluminescence emitted from neuroblastoma cells, as measured in photons/second/centimetre squared/steradian (p/sec/cm2/sr). (B)

H&E and luciferase immunostaining confirmed that neuroblastoma cells were present within the left kidney (black arrowhead). No staining was present in the isotype control

(rabbit IgG control) which confirmed the specificity of the secondary antibody (see

Materials & Methods section 2.23). (C) The left kidney became encapsulated by the primary tumour during the course of the experiment. The kidney was removed from the primary tumour prior to imaging.

242

A Left kidney 3.0 2.5 2.0 1.5 1.0 0.5 x108 p/sec/cm2/sr

B H&E Isotype control Luciferase

500μm

Primary tumour

Left kidney

243 excluded from any further bioluminescence analyses due to potential interference of neuroblastoma cells remaining attached to the outside of the organ (Fig. 6.8C). BLI and

IHC confirmed that neuroblastoma tumours grew on the surface of other organs that were either in direct contact or had close proximity to the primary tumour. A representative luminescent image of a CtrlSH right kidney is shown in Fig. 6.9A. IHC analyses showed that a luciferase-expressing neuroblastoma tumour grew on the surface of the right kidney (Fig. 6.9B, black arrowhead).

Similarly, a representative luminescent image of a CtrlSH liver (Fig. 6.10A) and corresponding IHC (Fig. 6.10B, black arrowhead), and a representative luminescent image of a CtrlSH spleen (Fig. 6.11A) and corresponding IHC (Fig. 6.11B, black arrowheads), demonstrated that luciferase-expressing neuroblastoma tumours grew on the surface of these organs. However, it was unclear whether these tumours were a result of direct contact with the primary tumour, had occurred via locoregional spread or had grown from cells that may have leaked from the injection site. Subsequently, these organs were excluded from further BLI-based analyses of metastasis due to the unknown origin of these tumours. However, it is important to note that this data does not exclude the possibility that there were metastatic tumours present in these organs.

244 Figure 6.9 Bioluminescent imaging and histological analysis of the right kidney.

(A) A representative luminescent image of a right kidney from a CtrlSH mouse. The coloured scale bar indicates the amount of bioluminescence emitted from neuroblastoma cells, as measured in photons/second/centimetre squared/steradian (p/sec/cm2/sr). (B)

H&E and luciferase immunostaining confirmed that a neuroblastoma tumour grew on the outside of the right kidney (black arrowhead). No staining was present in the isotype control (rabbit IgG control) which confirmed the specificity of the secondary antibody

(see Materials & Methods section 2.23).

245

A Right kidney 3.0 2.5 2.0 1.5

1.0 0.5 x108 p/sec/cm2/sr B

H&E

500μm

Isotype control

Luciferase

246 Figure 6.10 Bioluminescent imaging and histological analysis of the liver.

(A) A representative luminescent image of a liver from a CtrlSH mouse. The coloured scale bar indicates the amount of bioluminescence emitted from neuroblastoma cells, as measured in photons/second/centimetre squared/steradian (p/sec/cm2/sr). (B) H&E and luciferase immunostaining confirmed that a neuroblastoma tumour grew on the outside of the liver (black arrowhead). No staining was present in the isotype control (rabbit

IgG control) which confirmed the specificity of the secondary antibody (see Materials &

Methods section 2.23).

247 Liver A 5.0

4.0

3.0

2.0

1.0

X107 p/sec/cm2/sr B

H&E

500μm

Isotype control

Luciferase

248 Figure 6.11 Histological analysis of the spleen.

(A) A representative luminescent image of a spleen from a CtrlSH mouse. The coloured scale bar indicates the amount of bioluminescence emitted from neuroblastoma cells, as measured in photons/second/centimetre squared/steradian (p/sec/cm2/sr). (B) H&E and luciferase immunostaining confirmed that multiple neuroblastoma tumours grew on the surface of the spleen (black arrowheads). No staining was present in the isotype control

(rabbit IgG control) which confirmed the specificity of the secondary antibody (see

Materials & Methods section 2.23).

249

Spleen A 3.0

2.5

2.0

1.5

1.0

0.5 X109 p/sec/cm2/sr B

H&E

1mm

Isotype control

Luciferase

250 The bone, bone marrow and lymph nodes are common sites for metastatic tumours in neuroblastoma. BLI was therefore utilised to determine whether neuroblastoma cells had metastasised to these regions (Fig. 6.12). Mice were culled and their carcasses skinned to reduce signal interference from the fur. The primary tumours were excised prior to imaging. As shown in Fig. 6.12, bioluminescent signal was emitted from the region in which the primary tumour was excised. Bioluminescent signals were also evident in other regions on the carcass (red arrows) (Fig. 6.12). These signals indicated that neuroblastoma cells had potentially metastasised to the bone/bone marrow and/or lymph nodes (Fig. 6.12). However, accurate quantitation of the bioluminescent signal from these regions proved difficult due to interference from residual cells from the primary tumour and the inconsistent locations of these signals between animals. Based on these findings, quantitation of bioluminescence from animal carcasses were not included in this study.

6.4 Neuroblastoma lung metastasis

In this orthotopic neuroblastoma model, the lungs are separated from the primary tumour and injection site by the diaphragm. Therefore metastases found in this organ are likely a result of haematogenous spread. Mouse lungs were examined by BLI and

IHC to identify potential neuroblastoma metastases. A representative luminescent image of a CtrlSH lung is shown in Fig. 6.13A. IHC confirmed that luciferase-expressing neuroblastoma cells were present within the lung (Fig. 6.13B, black arrowheads) and at different locations within the lung tissue (Fig. 6.13B, black arrowheads). Using the same luminescent scale as the CtrlSH lung, a luminescent image of a Stmn Seq.3SH lung

251

Figure 6.12 BLI detection of potential bone and/or lymph node metastases.

Representative luminescent image of the dorsal view of a CtrlSH mouse carcass. The culled animal was skinned and the primary tumour excised prior to imaging. Some cells from the primary tumour remained on the carcass, as highlighted by the luminescent signal from that region. Luminescent signals (red arrows) were also present on other regions on the carcass indicating potential bone or lymph node involvement. The coloured scale bar indicates the amount of bioluminescence emitted from neuroblastoma cells, as measured in photons/second/centimetre squared/steradian (p/sec/cm2/sr).

252

1.0

0.8

Primary 0.6 tumour excision 0.4

0.2

x108 p/sec/cm2/sr

253 Figure 6.13 Bioluminescent imaging and histological analysis of a CtrlSH lung.

(A) A representative luminescent image of a lung from a CtrlSH mouse. The coloured scale bar indicates the amount of bioluminescence emitted from neuroblastoma cells, as measured in photons/second/centimetre squared/steradian (p/sec/cm2/sr). (B) Lung tissue sections at 1x and 4x magnification. Luciferase immunostaining confirmed that multiple neuroblastoma tumours grew within the lung (black arrowheads).

254

1.2 1.0 A 0.8 CtrlSH 0.6 0.4 0.2 X107 p/sec/cm2/sr B H&E Isotype control Luciferase

500μm

255 (Fig. 6.14A) shows less bioluminescent signal compared to the CtrlSH lung (Fig.

6.13A). In correlation with the signal, there were fewer luciferase-expressing tumours in the Stmn Seq.3SH lung (Fig. 6.14B) compared to the CtrlSH lung (Fig. 6.13B).

Importantly, this data confirmed that the bioluminescent signal from the lungs was due to the presence of neuroblastoma cells within this organ. Furthermore, the lungs were deemed a suitable organ to accurately quantitate metastasis in this model.

6.5 Stathmin suppression reduces neuroblastoma lung metastasis

BLI was utilised to evaluate stathmin’s contribution to neuroblastoma lung metastasis.

Four representative luminescent images of CtrlSH, Stmn Seq.2SH and Stmn Seq.3SH lungs are shown in Fig. 6.15A. Initially, 13 and 17 mice were orthotopically injected with Stmn Seq.2SH cells and Stmn Seq.3SH cells, respectively. However, Stmn Seq.2SH and Stmn Seq.3SH behaved similarly in regards to stathmin suppression (Fig. 6.1B), luciferase expression (Fig. 6.3), invasion (Fig. 6.2B) and tumour growth (Fig. 6.5).

Furthermore, there was no significant difference in neuroblastoma tumour burden

6 between Stmn Seq.2SH and Stmn Seq.3SH (median total flux of 9.13x10 for Stmn

7 Seq.2SH vs. 3.69x10 for Stmn Seq.3SH, p=0.315) (Fig. 6.15B). Therefore, Stmn Seq.2SH and Stmn Seq.3SH lung data was pooled (hereafter referred to as Stmn Seq.2/3SH).

256 Figure 6.14 Bioluminescent imaging and histological analysis of a Stmn Seq.3SH lung.

(A) A representative luminescent image of a lung from a Stmn Seq.3SH mouse. The coloured scale bar (same scale used for the CtrlSH lung) indicates the amount of bioluminescence emitted from neuroblastoma cells in photons/second/centimetre squared/steradian (p/sec/cm2/sr). (B) Lung tissue sections at 1x and 4x magnification.

Luciferase immunostaining confirmed that fewer neuroblastoma tumours (black arrowheads) grew within the Stmn Seq.3SH lung compared to the CtrlSH lung (Fig. 6.13).

257

1.2 1.0 A 0.8 0.6 Stmn Seq.3SH 0.4 0.2 X107 p/sec/cm2/sr B H&E Isotype control Luciferase

500μm

258 Figure 6.15 Neuroblastoma tumour burden in CtrlSH, Stmn Seq.2SH and Stmn

Seq.3SH mouse lungs.

(A) Four individual luminescent lung images from CtrlSH, Stmn Seq.2SH and Stmn

Seq.3SH mice. The coloured scale bar indicates the level of bioluminescence emitted from the lungs in photons/second/centimetre squared/steradian (p/sec/cm2/sr). (B)

Neuroblastoma tumor burden in mouse lungs was quantified by measuring the total flux

(photons/sec) from each lung. Each dot represents one lung and the bar represents the median total flux for each group (n≥ 13 for each group). The median total flux values are displayed on the right side of the bar and the p value (>0.05, not significant) is displayed above the joining bar.

259

260 Quantitation of mouse lung bioluminescence showed significantly reduced neuroblastoma tumour burden (71% reduction) in stathmin-suppressed mice compared

7 7 to controls (median total flux of 2.17x10 for Stmn Seq.2/3SH lungs vs. 7.36x10 for

CtrlSH lungs, p<0.008) (Fig. 6.16). Thus, suppressing stathmin significantly reduced the ability of neuroblastoma cells to metastasise in vivo.

261 Figure 6.16 Stathmin suppression reduces neuroblastoma tumour burden in the lungs of mice.

Neuroblastoma tumour burden in CtrlSH and Stmn Seq.2/3SH (mice pooled from Stmn

Seq.2SH and Stmn Seq.3SH) lungs were quantified by measuring the total flux

(photons/sec) from each lung. Each dot represents one lung and the bar represents the median total flux ±SEM (error bars) for each group (n≥ 27). The median total flux value is displayed to the right of the bar,* p< 0.05.

262

CtrlSH Stmn Seq.2/3SH

263 6.6 Discussion

The highly metastatic nature of neuroblastoma makes this malignancy one of the most deadly childhood cancers. Regrettably, even the most advanced treatment regimens have little efficacy against neuroblastoma metastases, which highlights an urgent need for new therapeutic targets to treat this disease.

Stathmin is highly expressed in neuroblastoma and other neuroendocrine tumours

(Hailat, Strahler et al. 1990; Sadow, Rumilla et al. 2008), however no studies to date have determined stathmin’s functional role in neuroblastoma. In this study, a clinically- relevant orthotopic neuroblastoma mouse model was established specifically to examine stathmin’s role in neuroblastoma tumour growth and metastasis. This model mimics human neuroblastoma development as reflected by the involvement of local organs, regional lymph nodes and distant sites; the latter being associated with more aggressive and advanced-stage disease (Khanna, Jaboin et al. 2002; Almgren, Henriksson et al.

2004; Henriksson, Almgren et al. 2004; Meier, Muhlethaler-Mottet et al. 2007; Nevo,

Sagi-Assif et al. 2008; Fuchs, Christofferson et al. 2009). In some animals, BLI and

IHC detected luciferase-expressing neuroblastoma tumours on the surface of organs such as the liver, kidneys and spleen that were in close proximity to the primary tumour.

These tumours potentially arose from direct contact with the primary tumour or had occurred via locoregional spread. However, for this study it was imperative to accurately quantitate metastatic spread. Therefore these organs were excluded from further analyses due to the unknown origin of these tumours.

In neuroblastoma patients, the presence of lung metastases confer a poor prognosis

(34.5±6.8% 5-year overall survival) as they often correlate with other unfavourable

264 features such as adrenal primary tumours, liver and central nervous system metastases, elevated lactate dehydrogenase and MYCN amplification (DuBois, London et al. 2008).

In this orthotopic model, the lungs are separated from the primary tumour by the diaphragm and therefore any metastases found in this organ would be a result of systemic spread from the primary site. Using BLI and IHC analyses, luciferase- expressing neuroblastoma cells were identified in the lungs of mice and neuroblastoma tumour burden (metastasis) in this organ was quantified by BLI. Stathmin suppression significantly reduced neuroblastoma tumour burden in the lungs of mice compared to controls (71% reduction; p<0.008). This data is consistent with in vitro data demonstrating that stathmin is important for neuroblastoma cell migration and invasion

(Chapter 5). Collectively, these results suggest high expression of stathmin in the primary tumour may promote metastasis. In support of this, studies have shown that stathmin expression is increased at the invasive front of lung tumours (Singer, Malz et al. 2009) and that stathmin over-expression enhanced cell invasion in vitro and the metastatic potential of sarcoma cells in vivo (Belletti, Nicoloso et al. 2008). Therefore strategies aimed at reducing stathmin expression in the primary tumour may prevent metastatic spread from this site. It is also possible that suppressing stathmin expression may reduce the invasiveness of cancer cells that have already metastasized. However, comprehensive analyses would be required to confirm this phenotype.

In summary, this study has demonstrated that combining a luciferase-based detection method with the orthotopic neuroblastoma mouse model is a valuable tool by which to study metastasis in this malignancy. Importantly, RNAi-mediated stathmin suppression in neuroblastoma cells has revealed a novel role for stathmin in neuroblastoma metastasis. Therapeutics designed to reduce stathmin expression in neuroblastoma tumours may therefore prevent metastasis and/or reduce the invasiveness of cancer cells

265 that have already spread. These outcomes would benefit most neuroblastoma patients as the majority (more than 50%) have wide-spread disease at diagnosis [reviewed in (Ara and DeClerck 2006)]. However further investigations are required to determine whether reducing stathmin expression also influences neuroblastoma metastasis to other organs or bone, and the mechanism by which stathmin is driving metastasis in this malignancy.

266 Chapter 7. Conclusions &

Future Directions

267 Neuroblastoma is a highly metastatic and drug-refractory childhood cancer. For children with advanced-stage disease, the prognosis is poor with 5-year survival rates less than

30% (Gutierrez, Fischer et al. 2007). Therefore new drug targets are urgently required to improve survival rates in this malignancy.

Stathmin, a microtubule destabilising protein, is highly expressed in neuroblastoma but until now, stathmin’s functional role in this malignancy was unknown. Results from this thesis have demonstrated for the first time that stathmin is important for neurite formation and metastasis in neuroblastoma. In contrast to a number of other cancers, stathmin does not influence neuroblastoma cell cycle progression, proliferation, viability, tumour growth or drug-sensitivity. This suggests that some of stathmin’s functions may be cell type-specific or that the other highly homologous stathmin-like proteins, which are abundant in neuroblastoma cells, may be able to mimic some of stathmin’s functions in these cells. In lung cancer cells, both stathmin and SCLIP regulate cell viability, proliferation, migration and invasion (Singer, Malz et al. 2009).

Combined suppression of stathmin and SCLIP in lung cancer cells had an additive effect on reducing cell migration compared to individual suppression of stathmin and

SCLIP (Singer, Malz et al. 2009). In contrast, combined suppression of stathmin and

SCLIP did not further reduce cell viability, proliferation or invasion, indicating that these proteins may be redundant for some functions and not others in lung cancer cells

(Singer, Malz et al. 2009). This begs the question as to what roles the other stathmin- like proteins play in neuroblastoma and whether they contribute to the aggressive nature of this malignancy. Future studies will be required to address these questions.

268 The phosphorylation of stathmin’s serine residues regulates stathmin’s tubulin-binding affinity, thereby impacting microtubule stability and the interactions and transport of various cytoskeletal remodelling proteins (Takahashi and Suzuki 2008; Takahashi and

Suzuki 2009). Results from this thesis revealed that stathmin is phosphorylated during

ATRA-induced neurite formation in neuroblastoma cells. Furthermore, suppression of stathmin inhibited ATRA-induced neurite outgrowth in neuroblastoma cells. Taken together, these results suggest that stathmin and potentially the phosphorylation of its serine residues, may be critical for neurite formation. One way in which to examine this further would be to express stathmin phosphorylation site-mutants in neuroblastoma cells, treat these cells with retinoic acid and examine neurite formation. These experiments could potentially identify which stathmin serine residues are critical for retinoic acid-induced neurite formation in neuroblastoma cells.

Hailat et al. discovered that stathmin is in a more ‘active’ (unphosphorylated) state in

MYCN-amplified primary neuroblastoma tumours and cell lines compared to non- amplified tumours and cell lines (Hailat, Strahler et al. 1990). Therefore increased stathmin expression and activity may contribute to a more aggressive neuroblastoma phenotype. In support of this, Ser16 phosphorylation is significantly reduced in metastatic breast cancer cell lines and tumours compared to untransformed cell lines, normal and primary tumour tissues (Li, Jiang et al. 2011). A combination of increased stathmin expression and reduced Ser16 phosphorylation was found in recurrent and metastatic sarcomas compared to normal tissue and primary tumours (Belletti, Nicoloso et al. 2008). Furthermore, increasing stathmin activity (via expression of a Ser16 phosphorylation-impaired mutant or over-expression of stathmin) reduced microtubule stability and promoted extracellular matrix (ECM)-driven cell migration and invasion in

269 vitro and metastatic potential of sarcoma cells in vivo (Belletti, Nicoloso et al. 2008).

Based on these findings, and the limited available clinical studies of stathmin in neuroblastoma (Hailat, Strahler et al. 1990), it would be valuable to determine whether stathmin and/or its phosphorylation state are altered in neuroblastoma tumours collected from patients with localised vs. metastatic disease.

Stathmin is critical for neurite formation and metastasis in neuroblastoma cells; processes that require activation of multiple signalling pathways to induce dramatic remodelling of the cell cytoskeleton. The major signalling proteins involved in cytoskeletal restructuring are the RhoGTPases which are activated by extracellular stimuli including retinoic acid (Clagett-Dame, McNeill et al. 2006) and growth factors

(Wittmann and Waterman-Storer 2001). Based on previous studies and the results from this thesis, it now appears that under conditions in which signalling pathways are activated, stathmin may have a dual role in regulating both microtubules and the actin network, by direct and/or indirect mechanisms (Wittmann and Waterman-Storer 2001;

Wittmann, Bokoch et al. 2004; Takahashi and Suzuki 2009). In this thesis it was demonstrated that stathmin regulated tubulin polymer levels and the phosphorylation of the actin regulatory proteins, cofilin and myosin light chain (MLC) via ROCK, in serum-starved and growth factor-activated neuroblastoma cells. Stathmin may also regulate cofilin dephosphorylation. However, the exact mechanism by which stathmin influences ROCK signalling and cofilin activity are unknown. The most likely candidates for this would be the RhoGTPases that are upstream of ROCK and cofilin.

Alterations in microtubule stability influence RhoGTPase activity and restructuring of the cell cytoskeleton (Wittmann and Waterman-Storer 2001). Stathmin is regulated downstream of various RhoGTPase signalling pathways and therefore has the potential

270 to influence the multiple feedback loops that occur between the cell cytoskeleton and its regulatory proteins (Daub, Gevaert et al. 2001; Wittmann, Bokoch et al. 2004;

Baldassarre, Belletti et al. 2005; Watabe-Uchida, John et al. 2006; Tanaka, Hamano et al. 2007). Future studies are therefore required to determine whether stathmin is regulating the RhoGTPases in neuroblastoma cells. These studies would not only provide further clarification of stathmin’s role in neuroblastoma cell migration/invasion and metastasis, but also neurite formation, which is heavily reliant on the RhoGTPases.

Although the mechanism by which stathmin regulates neuroblastoma metastasis is unknown, results from this thesis have indicated that stathmin may regulate a specific type of cell movement in neuroblastoma cells. Stathmin potentially drives neuroblastoma cell invasion by blocking ROCK signalling, a pathway known to mediate proteolysis-independent, amoeboid-like (rounded shape) cell motility (Torka,

Thuma et al. 2006). Consequently, stathmin, via regulation of the cell cytoskeleton, may promote an alternate mode of cell motility in neuroblastoma cells.

Mesenchymal-like (elongated-shape) cell motility is dependent on proteolysis-mediated degradation of the surrounding ECM (Torka, Thuma et al. 2006). The ECM is degraded by zinc-dependent MMPs such as MMP-2 and MMP-9 that are transported along microtubules with kinesin prior to exocytosis (Schnaeker, Ossig et al. 2004 632).

Advanced-stage neuroblastoma tumours express high levels of MMP-2 and MMP-9 that promote metastasis by degrading collagens and fibronectin in the ECM [reviewed in

(Jiang, Stanke et al. 2011)]. In particular, MMP-2-mediated degradation of collagen IV is thought to play a critical role in EMT of neural crest cells (Radisky and Radisky

2010). Based on results from the chemotaxis assays that utilised collagen IV as a

271 chemo-attractant (refer to Chapter 5, Fig. 5.3), it is possible that stathmin may be regulating MMPs in neuroblastoma cells. To support this, studies have shown that stathmin activity and thus microtubule stability are influenced by cellular contact with the ECM (Belletti, Nicoloso et al. 2008) and that direct interactions between stathmin, kinesin and tubulin are important for microtubule-mediated transport of various proteins in breast cancer cells (Takahashi and Suzuki 2009). More recently it has been demonstrated that direct interactions between stathmin and Siva1 (an apoptosis-related protein) suppress EMT by inhibiting stathmin-mediated microtubule destabilisation in breast cancer cells (Li, Jiang et al. 2011). Thus a potential role for stathmin in neuroblastoma metastasis has been postulated. Stathmin promotes neuroblastoma metastasis through microtubule destabilisation which in turn regulates microtubule- mediated transport and exocytosis of MMPs, ECM degradation and mesenchymal-like invasion and metastasis. To confirm this role for stathmin, it would be critical to determine whether stathmin regulates the transport and exocytosis of MMPs in neuroblastoma cells. Interestingly, this information also has implications for stathmin’s role in neurite formation as MMPs play an important role in retinoic acid-induced neurite formation in neuroblastoma cells (Joshi, Guleria et al. 2007).

Metastasis is the major cause of death in cancer. Stathmin over-expression is associated with highly metastatic disease in other cancers and, as demonstrated in this thesis, stathmin facilitates neuroblastoma metastasis. Therefore targeting stathmin may prove a valuable strategy to combat metastasis in many cancers. While there are no known inhibitors of stathmin, a recent study has reported that a bifunctional stathmin shRNA construct, encapsulated in cationic liposomes and delivered by repeated intratumoural injections, reduced stathmin expression and subsequent growth of colorectal cancer

272 xenografts and primary melanoma and osteosarcoma xenografts in nude mice (Phadke,

Jay et al. 2011). In another study, a survivin promoter-driven stathmin siRNA vector effectively suppressed stathmin expression, induced a G2-M delay, apoptosis and inhibited cell growth in human cervical cancer and osteosarcoma cell lines but had no effect on endothelial cells (Zhang, Wang et al. 2006). Therefore stathmin-specific siRNA/shRNA constructs may be useful for tumour gene therapy. Systemic delivery of nanoparticle-siRNA complexes may also provide an avenue for stathmin-targeted therapies in human cancers (Davis, Zuckerman et al. 2010). Neuroblastoma-targeting anti-disialoganglioside (GD2) lipid nanoparticles have proved successful for the delivery of anaplastic lymphoma kinase (ALK) siRNA to neuroblastoma tumours that induced growth arrest, apoptosis and increased survival in animal models of the disease

(Di Paolo, Ambrogio et al. 2011). Future studies are therefore required to determine whether stathmin can be targeted in primary and metastatic tumours using systemically- delivered shRNA or siRNA.

Chemotherapeutics already in the clinic may also prove valuable for targeting stathmin in cancer. The DNA-alkylating agent CCNU, currently used for the treatment of brain tumours, Hodgkin's disease, non-Hodgkin's lymphoma, melanoma, lung and colon cancer, covalently binds to stathmin promoting the carbamoylation of lysine residues

(Ngo, Peng et al. 2007). This drug inhibits stathmin-mediated microtubule destabilisation and reduces the migration and invasion of glioma cells (Liang, Choi et al. 2008). Using isothermal titration calorimetry and analytical ultracentrifugation,

Devred et al. revealed that stathmin increases the affinity of the microtubule- destabilising drug vinblastine for tubulin and vice versa which suggests that cells expressing high levels of stathmin may be more sensitive to this agent (Devred,

273 Tsvetkov et al. 2008). It is therefore possible that the efficacy of these agents may in part be attributed to their regulation of stathmin-tubulin interactions and that these drugs may prove useful for treating all stathmin-expressing tumours. In addition, this thesis has demonstrated that the orthotopic neuroblastoma mouse model, coupled with a luciferase-based detection method, could be a valuable tool to assess the efficacy of any stathmin-targeted therapies in pre-clinical studies.

In conclusion, the key finding of this thesis is that stathmin mediates metastasis in the aggressive childhood cancer neuroblastoma. This study has provided the first direct evidence that stathmin regulates neuroblastoma metastasis in vivo and that suppressing stathmin in neuroblastoma may not only reduce metastatic spread, but also the invasiveness of metastatic tumours. Therapeutics aimed at reducing stathmin expression or attenuating stathmin activity in primary and/or metastatic neuroblastoma tumours may improve treatment and ultimately survival rates for this deadly disease.

274 Appendix

275 Appendix Figure 1. Stathmin suppression reduces cell proliferation in Calu-6 lung cancer cells.

(A) A representative western blot showing stathmin suppression in Calu-6 cells transfected with increasing concentrations of stathmin (STMN) siRNA compared to control (Ctrl) siRNA. GAPDH was used as a loading control. (B) Cell counts of siRNA- transfected Calu-6 cells. Cell counts of STMN siRNA-treated cells were calculated as a percentage of Ctrl siRNA-treated cells. Data represent the mean of 3 independent experiments ± SEM (error bars).

276

A CALU-6

STMN siRNA (nmol/L) Ctrl 5 10 25 50 100 Stathmin

GAPDH

B CALU-6

Ctrl 5 10 25 50 100 STMN siRNA (nmol/L)

277 Appendix Figure 2. Stathmin-like protein expression in neuroblastoma and lung cancer cells.

(A) A representative western blot showing stathmin and SCG10 expression in neuroblastoma cells [SH-SY5Y and BE(2)-C] vs. lung cancer cells (Calu-6 cells).

GAPDH was used as a loading control. (B) A representative western blot showing stathmin and SCLIP expression in neuroblastoma cells [SH-SY5Y and BE(2)-C] vs. lung cancer cells (Calu-6 cells). GAPDH was used as a loading control.

278

SCG10 Stathmin

GAPDH

SCLIP Stathmin

GAPDH

279 Appendix Figure 3. Migration assay optimisation.

BE(2)-C cells were subjected to migration assays using PDGF and collagen IV (Coll.

IV) as chemo-attractants. PDGF was diluted in culture medium to 25ng/mL and placed in either the top or bottom chamber, or both chambers. Inserts were coated with Coll. IV

(10μg/mL) on either the top or bottom, or both sides, of the insert. The migration index

(%) was highest when PDGF was placed in the lower chamber and Coll. IV coated on the underside of the insert. Data represent ≥1 experiment for each condition.

280

BE(2)-C

PDGF (top) - + - - - PDGF (bottom) - + + + + Coll. IV (top) - - - + - Coll. IV (bottom) - - - + +

281 Appendix Figure 4. EMT marker protein expression in stathmin-suppressed neuroblastoma cells.

(A) Stathmin (STMN) and control (Ctrl) siRNA-treated BE(2)-C cells were serum- starved (serum-free) or serum-starved and treated with 10% serum in DMEM for the indicated time points. Paxillin, phospho-Paxillin (Paxillin-P) and stathmin protein expression were determined by western blot. GAPDH was used as a loading control. (B)

The expression of the EMT marker proteins; neural cadherin (N-cadherin) and vimentin, were examined in STMN and Ctrl siRNA-transfected BE(2)-C and SH-SY5Y cells.

GAPDH was used as a loading control.

282

A 10% serum Serum-free 5 min 15 min 30 min 60 min siRNA Ctrl STMN Ctrl STMN Ctrl STMN Ctrl STMN Ctrl STMN Stathmin

Paxillin-P

Paxillin

GAPDH

B BE(2)-C SH-SY5Y siRNA Ctrl STMN Ctrl STMN N-cadherin

Vimentin GAPDH

Stathmin

283 Appendix Figure 5. Detection of adrenal fat pad injection spills by bioluminescent imaging.

Representative luminescent images of 3 mice injected with luciferase-expressing neuroblastoma cells into their left adrenal fat pads. Mice were imaged on their dorsal, left lateral, right lateral and ventral sides. The coloured scale bar indicates the amount of bioluminescence emitted from neuroblastoma cells, as measured in photons/second/centimetre squared/steradian (p/sec/cm2/sr). Mice 1 and 2 show no cell leakage from their injection sites (localised signal). Mouse number 3 shows cell leakage, as represented by the red arrow, on the left lateral, right lateral and ventral sides. Mouse number 3 was therefore excluded from the study.

284

Dorsal Left lateral 12 3 12 3

12 3 1 2 3

Right lateral Ventral X108 p/sec/cm2/sr

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