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HOX/PBX interaction as a therapeutic target in Acute
Myeloid Leukaemia
A thesis submitted in part requirement for the
Degree of Doctor of Philosophy
Raed Alharbi
Microbial and cellular sciences
Faculty of Health and Medical Sciences
University of Surrey
March 2015
I
Declaration of originality
This thesis and the work to which it refers are the results of my own efforts. Any ideas,
data, images or text resulting from the work of others (whether published or unpublished) are
fully identified as such within the work and attributed to their originator in the text, bibliography
or in footnotes. This thesis has not been submitted in whole or in part for any other academic
degree or professional qualification. I agree that the University has the right to submit my work
to the plagiarism detection service TurnitinUK for originality checks. Whether or not drafts
have been so-assessed, the University reserves the right to require an electronic version of the
final document (as submitted) for assessment as above.
The majority of this introduction is taken from my own review article (Alharbi,
Pettengell et al. 2012) with supplementary material added in sections 1.1, 1.5, 1.7.4 and 1.8.
II
Summary
Acute myeloid leukaemia (AML) is a disorder characterised by the accumulation of
blast cells or progenitors of one of several non-lymphoid haematopoietic cell lineages and is
classified into two types: primary and secondary. HOX genes are over-expressed in both AML
and other cancers. This over-expression is associated with an intermediate/unfavourable
cytogenetic subset of AML. Although HOX over-expression is a common feature of AML,
conventional knockout methods have failed to fully evaluate their functions due to their
functional redundancy. We have applied an alternative approach by using a synthetic peptide
called HXR9 to antagonise the interaction between HOX proteins and their cofactor PBX,
which interacts with HOX proteins in groups 1-10.
AML cell lines derived from different AML types express different subsets of HOX
genes at different levels due to the heterogeneity of AML. It is showed for the first time that
targeting the HOX-PBX interaction using HXR9 led to cell death of the tested AML cell lines.
This cell death did not appear to be through apoptosis, as there were no signs of the caspase
activation and nuclear fragmentation. Likewise, there was also no activation of key necrotic
markers such as cypD and PARP1. Instead, cell death involved, at least in part, the expression
of c-FOS and p21 in p53-independentmanner. In addition, HXR9 caused cell death in
MEK/ERK and p38 independent pathways, but the JNK pathway exerted a resistant effect in
K562 cells. It was found that inhibiting the Ca2+ downstream mediators CaM, PKC and HO-1
significantly sensitised tested AML cell lines to HXR9. Taken together, these findings indicate
a novel cell death pathway in AML cells. In vivo modelling also showed that HXR9 could delay
tumour growth in a mouse model of AML.
III
Acknowledgements
First of all, I would like to thank my first supervisor Dr. Richard Morgan for his patience
for my endless questions, great guidance throughout the last four years, and for his wonderful
help to extend my ideas and translate it to practical assays that has great impacts on both myself
and the outcome of the project. I also would like to thank Prof. Hardev Pandha for giving me
the chance to work in his group. My special thanks to Angela Boxall who trained me in my first
months in the lab. Also, I would like to thank Dr. Guy Simpson for his great help in mouse
models. I am very lucky to have such friendly colleagues and very grateful for their support and
company in the last few years.
My special thanks to my family whom I dedicate this thesis to. I am indebted to my
Mother who is an endless source of help and support throughout my life. I also would like to
thank my brothers and my little sister for their encouragement. My special thanks to my wife
Ruba and my little angels Ratille and Racille who make my life more interesting and
meaningful. I really look forward to spending all the time with you.
My sincere love is to my Dad's soul who taught me reading and writing. He was the
master teacher throughout my life. I wish he were here to see this moment that he had been
waiting for. I miss his support, encouragement and voice.
IV
SUMMARY .............................................................................................................................. II
ACKNOWLEDGEMENTS .................................................................................................. III
LIST OF FIGURES ............................................................................................................ VIII
LIST OF TABLES ................................................................................................................. IX
LIST OF ABBREVIATIONS ................................................................................................. X
CHAPTER 1 INTRODUCTION ............................................................................................ 1
1.1 Overview ................................................................................................................................................... 2
1.1.1 Genetic changes in AML ....................................................................................................................... 6
1.2 HOX Genes ................................................................................................................................................ 6
1.3 HOX Cofactors ......................................................................................................................................... 8
1.4 HOX genes in haematopoiesis .................................................................................................................. 9
1.4.1 Gain of function studies ....................................................................................................................... 10
1.4.2 Loss of function studies ....................................................................................................................... 11
1.5 Upstream regulators of HOX genes ...................................................................................................... 15
1.6 HOX downstream target genes in haematopoietic cells ...................................................................... 16
1.7 The role of HOX genes in acute leukaemia ........................................................................................... 21
1.7.1 HOX fusion proteins ............................................................................................................................ 21
1.7.2 HOX over-expression in AML ............................................................................................................. 23
1.7.3 HOX gene dysregulation in acute lymphoid leukaemia (ALL) ............................................................ 25
1.7.4 HOX genes as prognostic markers ....................................................................................................... 26
1.8 Hypothesis and aims ............................................................................................................................... 28
CHAPTER 2 MATERIALS AND METHODS .................................................................. 30
2.1 Materials ................................................................................................................................................. 31
2.1.1 Reagents ............................................................................................................................................... 31
2.1.2 Instruments........................................................................................................................................... 33
2.1.3 Cell lines .............................................................................................................................................. 33
2.1.4 HXR9 and CXR9 peptide synthesis ..................................................................................................... 33
2.1.5 Mice ..................................................................................................................................................... 34
2.2 Methods ................................................................................................................................................... 34
2.2.1 General cell culture methods ................................................................................................................ 34
2.2.1.1 Routine cell culture ..................................................................................................................... 34
2.2.1.2 Cell counting and cell density calculation using a haemocytometer ........................................... 36
2.2.1.3 Cryopreservation of cell stocks ................................................................................................... 36
2.2.1.4 Revitalisation of cryopreserved cells........................................................................................... 37
V
2.2.2 Gene expression analysis by real-time PCR (RT-PCR) ....................................................................... 37
2.2.2.1 mRNA extraction ........................................................................................................................ 37
2.2.2.2 Measuring mRNA concentration ................................................................................................. 38
2.2.2.3 Reverse transcription of mRNA into cDNA ............................................................................... 38
2.2.2.4 Complementary PCR primers design .......................................................................................... 39
2.2.2.5 RT-PCR ....................................................................................................................................... 42
2.2.2.6 RT-PCR data analysis ................................................................................................................. 43
2.2.3 Lactate dehydrogenase (LDH) assay ................................................................................................... 44
2.2.3.1 LDH assay data analysis ............................................................................................................. 45
2.2.4 Assessment of drug combination interaction by LDH ......................................................................... 46
2.2.5 Annexin V- PE assay ........................................................................................................................... 47
2.2.6 Caspase-3 activity assay ...................................................................................................................... 49
2.2.6.1 Caspase-3 activity data analysis .................................................................................................. 50
2.2.7 Cell cytospins and 4',6-diamidino-2-phenylindole (DAPI) staining .................................................... 50
2.2.8 Inhibition of caspases activity by z-VAD-FMK .................................................................................. 51
2.2.9 Cyclosporin A (CsA) protection assay ................................................................................................. 52
2.2.10 Necrostatin-1 (Nec-1) protection assay ........................................................................................... 53
2.2.11 Fructose protection assay ................................................................................................................. 53
2.2.12 The effect of ethilenediaminetetra-acetic acid (EDTA) on HXR9 cytotoxicity .............................. 54
2.2.13 The effect of HXR9 on mitogen activated protein kinase (MAPK) pathways ................................ 55
2.2.14 The effect of inhibition of NADPH oxidase (NOX) on HXR9 efficacy .......................................... 55
2.2.15 Role of μ-calpain in HXR9 cell killing ............................................................................................ 56
2.2.16 The effect of calmodulin (CaM) inhibition on HXR9 cytotoxicity ................................................. 56
2.2.17 The effect of protein kinase C (PKC) inhibition on HXR9 cytotoxicity ......................................... 57
2.2.18 The effect of the heme oxygenase-1 (HO-1) inhibition on HXR9 cytotoxicity ............................... 57
2.2.19 The effect of the p53 inhibition on HXR9 cytotoxicity ................................................................... 58
2.2.20 Western blotting (WB) for protein expression ................................................................................. 58
2.2.20.1 Preparation of cell lysate ............................................................................................................. 59
2.2.20.2 Measuring protein concentrations by pierce BCA assay ............................................................. 59
2.2.20.3 Sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE) for protein
separation …………………………………………………………………………………………………...60
2.2.20.4 Transferring proteins to polyvinylidene fluoride (PVDF) membranes ....................................... 60
2.2.20.5 Detection of proteins by antibodies ............................................................................................. 60
2.2.21 In vivo assays ................................................................................................................................... 62
2.2.21.1 Cell preparations ......................................................................................................................... 62
2.2.21.2 Systemic injection of C1498-GFP in C57BL/6 and nude mice ................................................... 62
2.2.21.2.1 Harvesting and processing PB ............................................................................................. 62
2.2.21.2.2 Harvesting and processing BM and other organs................................................................. 62
2.2.21.3 Subcutaneous (S.C.) injection of AML cells into C57BL/6 and SCID mice .............................. 63
2.3 Statistical analysis .................................................................................................................................. 63
2.3.1 Calculation of IC50 ............................................................................................................................... 64
2.3.2 The analysis of drug combination assay data ....................................................................................... 64
2.3.3 Statistical analysis of in vivo assays ..................................................................................................... 64
CHAPTER 3 IN VITRO CYTOTOXICITY OF HXR9 ..................................................... 67
3.1 Introduction ............................................................................................................................................ 68
3.1.1 Amis of chapter 3 ................................................................................................................................. 69
3.2 Results ..................................................................................................................................................... 70
3.2.1 HOX gene expression in AML cell lines .............................................................................................. 70
VI
3.2.2 HXR9 is cytotoxic on all tested AML derived cell lines ..................................................................... 73
3.2.3 DNR is cytotoxic on K562 and HL-60 ................................................................................................ 76
3.2.4 Combination effect of HXR9 and DNR ............................................................................................... 77
3.2.5 MTX induces K562 and HL-60 cell death ........................................................................................... 79
3.2.6 Combination effect of HXR9 and MTX .............................................................................................. 80
3.3 Discussion ................................................................................................................................................ 82
3.3.1 Summary of chapter ............................................................................................................................. 87
CHAPTER 4 THE MECHANISM OF HXR9 CYTOTOXICITY ................................... 88
4.1 Introduction ............................................................................................................................................ 89
4.1.1 Aims of chapter 4 ................................................................................................................................. 94
4.2 Results ..................................................................................................................................................... 96
4.2.1 HXR9 causes the up-regulation of c-FOS ............................................................................................ 96
4.2.2 HXR9 causes death of AML tested cell lines ...................................................................................... 98
4.2.3 HXR9 does not affect caspase or Bcl-2 family transcription ............................................................. 101
4.2.4 HXR9 does not activate caspase-3 in the tested AML cell lines ........................................................ 104
4.2.5 HXR9 efficacy is not affected by the general inhibition of caspases ................................................. 107
4.2.6 HXR9 does not cause nuclear fragmentation ..................................................................................... 109
4.2.7 Ca2+ chelating abrogates cell killing by HXR9 .................................................................................. 112
4.2.8 EDTA rescues cells from killing by HXR9 ....................................................................................... 114
4.2.9 Cell killing by HXR9 does not involve CypD ................................................................................... 116
4.2.10 Inhibition of RIP1 modifies the cytotoxicity of HXR9.................................................................. 118
4.2.11 HXR9 induces cell death through an ATP-independent pathway ................................................. 120
4.2.12 HXR9 induces cell death through a PARP1-independent pathway ............................................... 122
4.2.13 The effect of inhibition of MAPK pathways on HXR9 cytotoxicity ............................................. 123
4.2.13.1 HXR9 induces cell death in a MEK/ERK independent pathway .............................................. 123
4.2.13.2 HXR9-mediated cell death does not require p38 pathway signalling ....................................... 125
4.2.13.3 Inhibition of the JNK pathway sensitises K562, but not HL-60 cells to HXR9 ........................ 127
4.2.14 Blocking NOX enzymes sensitizes cells to HXR9 ........................................................................ 129
4.2.15 μ-Calpain is not required for HXR9-induced cell death ................................................................ 131
4.2.16 CaM inhibition dramatically increases the sensitivity of cells to HXR9 ....................................... 133
4.2.17 Inhibition of PKC greatly increases the sensitivity of K562 and HL-60 cells to HXR9 ............... 135
4.2.18 Simultaneous inhibition of CaM and PKC potentiates HXR9 cytotoxicity ................................... 137
4.2.19 HO-1 inhibition increases the sensitivity to HXR9 ....................................................................... 139
4.2.20 HXR9 induces p21 expression but not p53 ................................................................................... 141
4.2.21 HXR9-induces AML cell death through a p53-independent pathway ........................................... 143
4.3 Discussion .............................................................................................................................................. 144
4.3.1 Summary of chapter ........................................................................................................................... 156
CHAPTER 5 IN VIVO CYTOTOXICITY OF HXR9 ..................................................... 157
5.1 Introduction .......................................................................................................................................... 158
5.1.1 Aims of chapter 5 ............................................................................................................................... 159
5.2 Results ................................................................................................................................................... 160
5.2.1 HXR9 is cytotoxic for C1498-GFP cells ........................................................................................... 160
5.2.2 CaM blocking enhances the cytotoxicity of HXR9 ........................................................................... 161
5.2.3 Inhibition of PKC activity significantly sensitises cells to HXR9 ..................................................... 163
VII
5.2.4 Concurrent inhibition of CaM and PKC significantly enhances the efficacy of HXR9 ..................... 165
5.2.5 Efficient expression of GFP in C1498 cells ....................................................................................... 167
5.2.6 Establishment a systemic C1498-GFP model in C57BL/6 mice........................................................ 168
5.2.7 Establishment a systemic C1498-GFP model in C57BL/6 and nude mice ........................................ 170
5.2.8 Development a C1498-GFP flank model in C57BL/6 mice .............................................................. 174
5.2.9 HXR9 significantly extended the survival of C1498-GFP xenograft ................................................ 175
5.2.10 Simultaneous inhibition of CaM and PKC did not sensitise C1498-GFP flank tumour to HXR9. 177
5.2.11 HXR9 did affect the tumour growth of K562 flank model ............................................................ 179
5.3 Discussion .............................................................................................................................................. 181
5.3.1 Summary of chapter ........................................................................................................................... 187
CHAPTER 6 GENERAL DISCUSSION AND FUTURE DIRECTIONS ..................... 188
6.1 Discussion .............................................................................................................................................. 189
6.2 Conclusion ............................................................................................................................................. 203
6.3 Future directions .................................................................................................................................. 204
APPENDICES ....................................................................................................................... 206
REFERENCES ..................................................................................................................... 218
PUBLICATIONS .................................................................................................................. 247
VIII
List of Figures
Figure 1.1 A schematic diagram of haematopoiesis.. .............................................................................................. 3
Figure 1.2 A schematic structure of clustered HOX genes. ..................................................................................... 8
Figure 1.3 structures of AbdB-HOX, NUP98 and the predictive fusion protein NUP98-HOXA.. ....................... 22
Figure 2.1 A haemocytometer diagram.. ............................................................................................................... 36
Figure 2.2 LDH cytotoxicity assay.. ...................................................................................................................... 44
Figure 2.3 A flow cytometry plot shows different cell populations in the annexin V-PE assay.. ......................... 48
Figure 2.4 ATP depletion by fructose. .................................................................................................................. 53
Figure 3.1 HOX gene expression in KG-1, HEL92.1.7, KU812F, K562 and HL-60 cell lines.. ........................... 72
Figure 3.2 LDH assay for HXR9 and CXR9 cytotoxicity on AML derived cell lines.. ........................................ 75
Figure 3.3 LDH assay for DNR cytotoxicity on K562 and HL-60 cell lines......................................................... 76
Figure 3.4 LDH assay of HXR9 and DNR combination therapy for K562 and HL-60.. ....................................... 78
Figure 3.5 LDH assay for MTX cytotoxicity on K562 and HL-60 cell lines.. ...................................................... 79
Figure 3.6 LDH assay of HXR9/MTX combination therapy for K562 and HL-60.. ............................................. 81
Figure 4.1 A schematic diagram of cell death.. ..................................................................................................... 90
Figure 4.2 c-FOS expression in AML cell lines after treatment with HXR9.. ...................................................... 97
Figure 4.3 Detection of cell death in AML cell lines by flow cytometry.. .......................................................... 100
Figure 4.4 Analysis of transcriptional changes of several pro- and anti-apoptotic genes upon HXR9 treatment.
............................................................................................................................................................................. 103
Figure 4.5 Detection of caspase-3 activity in AML cell lines using Z-DEVD-R110.. ........................................ 106
Figure 4.6 General inhibition of caspase activity in HXR9 treated AML cell lines by z-VAD-FMK.. .............. 108
Figure 4.7 DAPI staining of AML cells upon HXR9 treatment.. ........................................................................ 111
Figure 4.8 Effect of EDTA on HXR9 cytotoxicity.. ............................................................................................ 113
Figure 4.9 LDH analysis of the effect of EDTA on HXR9 cytotoxicity.. ........................................................... 115
Figure 4.10 Effect of CsA on HXR9 cytotoxicity.. ............................................................................................. 117
Figure 4.11 Effect of RIP1 inhibition on HXR9 cytotoxicity.. ............................................................................ 119
Figure 4.12 The effect of ATP depletion on HXR9 cytotoxicity.. ...................................................................... 121
Figure 4.13 WB analysis of PARP1 activation after HXR9 treatment.. .............................................................. 122
Figure 4.14 The effect of inhibiting the MEK/ERK pathway on the cytotoxicity of HXR9. .............................. 124
Figure 4.15 The effect of inhibiting the p38 pathway on the cytotoxicity of HXR9.. ......................................... 126
Figure 4.16 The effect of inhibiting the JNK pathway on HXR9 cytotoxicity.. .................................................. 128
Figure 4.17 The effect of NOX inhibition on the cytotoxicity of HXR9.. ........................................................... 130
Figure 4.18 The effect of blocking μ-calpain on HXR9 cytotoxicity. ................................................................. 132
Figure 4.19 The effect of CaM inhibition on HXR9 sensitivity.. ........................................................................ 134
Figure 4.20 The effect of blocking of PKC on the cytotoxicity of HXR9 on K562 and HL-60 cells. ................ 136
Figure 4.21 The impact of concurrent inhibition of CaM and PKC on the cytotoxicity of HXR9.. .................... 138
Figure 4.22 The effect of HO-1 inhibition on the cytotoxicity of HXR9.. .......................................................... 140
Figure 4.23 RT-PCR analysis of p53 and p21 expression in response to HXR9 treatment.. ............................... 142
Figure 4.24 The effect of blocking p53 protein on HXR9 cytotoxicity. .............................................................. 143
Figure 5.1 LDH assay for the cytotoxicity of HXR9 on C1498-GFP cells. ........................................................ 160
Figure 5.2 The effect of CaM inhibition on the cytotoxicity of HXR9 for C1498-GFP.. ................................... 162
Figure 5.3 The impact of PKC inhibition on the sensitivity of C1498-GFP cells to HXR9. ............................... 164
Figure 5.4 The effect of CaM and PKC simultaneous inhibition on the efficacy of HXR9.. .............................. 166
Figure 5.5 Expression of GFP in C1498 cells.. ................................................................................................... 167
Figure 5.6 FACS analysis of PB from C57BL/6 mice 13 days after injection of C1498-GFP cells. .................. 168
Figure 5.7 FACS analysis of C1498-GFP cells shows engraftment in several organs of C57BL/6 and nude mice.
............................................................................................................................................................................. 172
Figure 5.8 HXR9 treatment effect on the growth of C1498-GFP xenograft in female C57BL/6 mice. .............. 176
Figure 5.9 Effect of HXR9 I.T. treatment on the growth of C1498-GFP flank xenograft in C57BL/6 mice.. .... 178
Figure 5.10 Impact of HXR9 treatment on the K562 flank-xenograft growth in SCID mice.............................. 180
IX
List of Tables
Table 1.1 Classification of leukaemia. .................................................................................................................... 3
Table 1.2 AML classification according WHO classification ................................................................................. 5
Table 1.3 HOX gene studies. ................................................................................................................................. 14
Table 1.4 A summary of mammalian HOX target genes....................................................................................... 20
Table 2.1 Reagents were used in this study and their suppliers. ............................................................................ 31
Table 2.2 Cell lines used in this study. .................................................................................................................. 35
Table 2.3 cDNA synthesis mix.. ............................................................................................................................ 39
Table 2.4 HOX gene primers used for PCR amplification..................................................................................... 40
Table 2.5 Pro- and anti-apoptotic and β-actin gene primers used for PCR amplification. .................................... 42
Table 2.6 RT-PCR reaction components. This table shows the components of single RT-PCR reaction.. ........... 43
Table 2.7 Different cell populations in the annexin V-PE assay. .......................................................................... 48
Table 2.8 Antibodies used in WB and working dilutions. ..................................................................................... 61
Table 2.9 Summary of the in vitro assays performed and reagents used. .............................................................. 65
Table 3.1 IC50 values of HXR9 on AML derived cell lines with SEM. ............................................................... 75
Table 3.2 The analysis of the combination effect of HXR9 and DNR by Calcusyn software.. ............................. 78
Table 3.3 The analysis of the combination effect of HXR9 and MTX by Calcusyn software. ............................. 81
Table 3.4 The IC50 values of HXR9 with cell lines derived from different cancers. ............................................. 84
Table 4.1 The mechanism of HXR9 cytotoxicity on AML cells. ........................................................................ 156
Table 5.1 A summary of injection of C1498-GFP I.V. in C57BL/6 mice. .......................................................... 169
Table 5.2 A summary of injections of C1498-GFP I.V. in C57BL/6 and nude mice. ......................................... 173
Table 5.3 A summary of injection of C1498-GFP S.C in C57BL/6 mice. .......................................................... 174
X
List of Abbreviations
7-AAD 7-Amino actinomycin D
Abd-HOXA AbdominalB-HOXA
ACPP Activatable CPP
AIF Apoptosis inducing factor
ALL Acute lymphoid leukaemia
AML Acute myeloid leukaemia
AP-1 Activator protein-1
Apaf1 Apoptosis protease-activating factor
APML Acute promyelocytic leukaemia
ATP Adenosine triphosphate
ATRA All-trans retinoic acid
BCA Bicinchonic acid
BM Bone marrow
BCA Bradford city assay
BSA Bovine serum albumin
Ca2+ Calcium ions
CaM Calmodulin
cDNA Complementary deoxyribonucleic acid
CDX Caudal-type homebox transcription factor
c-FLIP (L) Cellular FLICE (FADD-like IL-1β-converting enzyme)- inhibitory
protein (L)
CI Combination index
CML Chronic myeloid leukaemia
CPP Cell penetrating peptide
CR Complete remission
CsA Cyclosporin A
XI
CXCR4 C-X-C chemokine receptor type 4
CypD Cyclophilin D
Cyt C Cytochrome C
DAPI 4',6-Diamidino-2-phenylindole
DISC Death-inducing signalling complex
DMEM Dulbecco’s Modified Eagle’s Medium
DMSO Dimethyl sulphoxide
DNR Daunorubicin
DPI Diphenyleneiodonium chloride
ED Effective dose
EDTA Ethilenediaminetetra-acetic acid
ELISA Enzyme-linked immunosorbent assay
ERK Extracellular signal-regulated kinase
FACS Fluorescence-activated cell sorting
FBS Fetal bovine serum
FLT3 Fms-like tyrosine kinase 3
GFP Green fluorescent protein
H2O2 Hydrogen Peroxide
HO-1 Heme oxygenase-1
HOX Homeobox transcription factor
HP Haemtopoietic progenitor
HSC Haematopoietic stem cell
I.T. Intratumoural
I.V. Intravenous
IC50 The half maximal inhibitory concentration
IMDM Iscove’s Modified Dulbecco’s medium
JNK Jun N-terminal kinase
XII
LDH Lactate dehydrogenase
MAPK Mitogen activated protein kinase
MDS Myelodysplasia
MEIS Myeloid ectropic insertion site
MLL Mixed lineage, myeloid lymphoid, leukaemia
mPTP Mitochondrial permeability transition pore
MRD Minimal residual disease
mRNA Messenger ribonucleic acid
MTX Mitoxantrone
Nec-1 Necrostatin-1
NF-ҠB Nuclear factor-kappa B
NK Natural killer cells
NK-AML Normal karyotype AML
NOD-SCID Non-obese diabetic with severe combined immune-deficient
NOX NADPH oxidase
NPM1 3 Nucleophosmin 1
NUP98 Nucloporin 98
OS Overall survival
PARP Poly ADP ribose polymerase
PB Peripheral blood
PBS Phosphate buffered saline
PBX Pre B cell leukaemia
PCD Programmed cell death
PCR Polymerase chain reaction
PFT-α Pifithrin-α
PKC Protein kinase C
PPIX Protoporphyrin IX
XIII
PS Phosphatidlyserine
PVDF Polyvinylidene fluoride
R2 Linear coefficient correlation
R9 Arginine residues
RAS Rat sarcoma
RIP1 Receptor interacting protein 1
RIP3 Receptor interacting protein 3
RNA Ribonucleic acid
Ro31-8220 Methanesulfonate salt
ROS Reactive oxygen species
RPM Revolution per minute
RPMI-1640 Roswell Park Memorial Institute
RT Room temperature
RT-PCR Real-time PCR
S.C. Subcutaneous
SCID Severe combined immune-deficient
SDS-PAGE Sodium dodecyl sulphate polyacrylamide gel electrophoresis
SEM Standard error of mean
TNF Tumour necrosis factor
TRAIL Tumour necrosis-related apoptosis-inducing ligand
VEGF Vascular endothelial growth factor
W-7 N-(6-Aminohexyl)-5-chloro-1-naphthalenesulfonamide hydrochloride
WB Western blotting
XIAP X linked inhibitor of apoptosis
1
Chapter 1 Introduction
2
1.1 Overview
Haematopoiesis is defined as the formation of new blood cells. Initially,
haematopoietic stem cells (HSCs) differentiate into multipotent haematopoietic
progenitors that undergo a gradual differentiation to give rise to array of more lineage-
restricted progenitors that ultimately produce highly specialised and differentiated
mature blood cells (Figure 1.1). Mature cells can be classified into myeloid cells
including neutrophils, eosinophils, basophils, monocytes, platelets and red cells, and
lymphoid cells including B- lymphocytes, T-lymphocytes and natural killer (NK) cells
(Orkin and Zon 2008; Doulatov, Notta et al. 2012).
Haemtological malignancies are a group of heterogeneous diseases that are
initiated by leukaemic stem cells that are able to both self-renew and differentiate like
normal HSCs, although in an abnormal fashion, thereby increasing their number and
giving rise to differentiated cells that represent the majority of cells found in the tumour.
The classification of haemtological malignancies depended on the stage of the disease
(acute or chronic leukaemia) and the immunophenotype of the cells (myeloid or
lymphoid) (Table 1.1) (Warr, Pietras et al. 2011).
3
Figure 1.1 A schematic diagram of haematopoiesis. HSC differentiate to MPP that give rise to lineage-
committed progenitors which in turn are differentiated to more mature blood cells. HSC: haematopoietic
stem cell, MPP: multipotent progenitor, CLP: committed lymphoid progenitor, CMP: committed
myeloid progenitor, Pro-NK: pro-natural killer cell, MEP: megakaryocyte-erythrocyte progenitor, GMP:
granulocyte-monocyte progenitor, MP: megakaryocyte progenitor, EP: erythrocyte progenitor, GP:
granulocyte progenitor, MP: monocyte progenitor.
Table 1.1 Classification of leukaemia.
Acute Chronic
Myeloid origin Acute myeloid leukaemia Chronic myeloid leukaemia
Lymphoid origin Acute lymphoid leukaemia Chronic lymphoid leukaemia
4
Acute myeloid leukaemia (AML) is a heterogeneous group of genetically and
phenotypically aggressive disorders where the differentiation of haematopoietic
progenitors (HPs) is blocked, increasing their self-renewal ability and disturbing the
normal regulation of proliferation (Frohling, Scholl et al. 2005). In the UK, AML is
the most frequent acute leukaemia in adults, accounting for 77% of cases. The median
age at presentation is 69 years and the male: female ratio is about 5: 4 (Smith, Howell
et al. 2011). The disease is commonly classified by either the French-American-British
system, or that described by the world health organization. The former is based on
morphology and maturation stage and classifies AML into eight groups (M0-M7). The
latter is also based on morphology, but also includes immunophenotyping, genetics and
clinical manifestations, and classifies AML into four main groups: AML with recurrent
genetic abnormalities, AML with myelodysplasia (MDS)-related changes, therapy-
related AML and MDS or AML that does not fit into any of these groups (Table 1.2).
Non-random chromosomal alterations, such as balanced translocations, monosomies,
trisomies, inversion and deletions have been found in the leukaemic cells of almost 55%
of AML patients, and until recently they were considered to be the most crucial
prognostic factors for complete remission (CR), likelihood of relapse, and overall
survival (OS) (Estey and Döhner 2006; Mrózek, Marcucci et al. 2007).
About 55% of AML cases have chromosomal aberrations and about 15% have
complex karyotype, three or more cytogenetic aberrations (Betz and Hess 2010).
Cytogenetic studies is used as prognostic indicator and classify AML into three
prognostic groups: favorable, intermediate and adverse. AML cases with t(15;17),
t(8;21) and t(16;16) are associated with favorable-risk group. The intermediate-risk
group includes cases with t(9;11), along with cases exhibiting loss of the Y
chromosome or gains of whole chromosome. This prognostic group also includes AML
5
cases with normal karyotype that account almost 45% of AML cases. The adverse-risk
group includes complex karyotype, t(6;9), inv(3)/ t(3;3) (Roche, Zeng et al. 2004;
(Marcucci, Mrozek et al. 2005; Betz and Hess 2010; Buccisano, Maurillo et al. 2012).
Table 1.2 AML classification according WHO classification (WHO 2008).
Categories
AML with recurrent genetic abnormalities
AML with t(8;21)(q22;q22); RUNX1-RUNX1T1
AML with inv(16)(p13.1q22) or t(16;16)(p13.1;q22); CBFB-MYH11
APL with t(15;17)(q22;q12); PML-RARA
AML with t(9;11)(p22;q23); MLLT3-MLL
AML with t(6;9)(p23;q34); DEK-NUP214
AML with inv(3)(q21q26.2) or t(3;3)(q21;q26.2); RPN1-EVI1
AML (megakaryoblastic) with t(1;22)(p13;q13); RBM15-MKL1
Provisional entity: AML with mutated NPM1
Provisional entity: AML with mutated CEBPA
AML with myelodysplasia-related changes
Therapy-related AML
AML, not otherwise specified
AML with minimal differentiation
AML without maturation
AML with maturation
Acute myelomonocytic leukemia
Acute monoblastic/monocytic leukemia
Acute erythroid leukemia
Pure erythroid leukemia
Erythroleukemia, erythroid/myeloid
Acute megakaryoblastic leukemia
Acute basophilic leukemia
Acute panmyelosis with myelofibrosis
Down syndrome related AML
Transient abnormal myelopoiesis
Myeloid leukemia associated with Down syndrome
6
Recent advances in molecular diagnosis have resulted in both gene alterations
and the dysregulation of specific genes becoming increasingly important as prognostic
elements in AML. This has helped to clarify the numerous heterogeneities of AML
subsets, particularly AML subsets showing normal karyotype AML (NK-AML)
(Dohner and Dohner 2008) and furthered understanding of the molecular mechanisms
of leukaemogenesis.
1.1.1 Genetic changes in AML
The origin of AML is associated with two distinct genetic changes, referred to
as Class I and Class II. Class I consists of mutations that enhance proliferation signal
transduction pathways and induce the proliferation of HSCs or HPs and usually affect
kinase signaling pathways, such as FLT3, KIT, NRAS/KRAS and JAK/STAT
mutations. Class I mutations take place late and cause disease progression. Class II
mutations target haematopoietic transcription factor genes leading to a block in myeloid
differentiation and conferring the self-renewal ability of HPs. These mutations take
place early and initiate the AML disease (Dohner and Dohner 2008; Renneville,
Roumier et al. 2008; Betz and Hess 2010). One of the most affected and mutated
transcription factors are homeobox (HOX) genes.
1.2 HOX Genes
The HOX genes are a family of homeodomain-containing transcription factors
(Garcia-Fernandez 2004), initially characterized in Drosophila as master regulators of
trunk and tail development during embryogenesis (Shah and Sukumar 2010).
Duplication of the original HOX gene cluster has given rise to 39 genes in mammals,
7
separated into four clusters known as A, B, C, and D (Amores, Force et al. 1998;
Abramovich, Pineault et al. 2005; Shah and Sukumar 2010). These clusters are located
on four different chromosomes, HOXA (7p15), HOXB (17q21), HOXC (12q13), and
HOXD (2q31) (Rice and Licht 2007). Ancestors of the original gene in each of the
clusters are known as paralogs, and generally they show a high degree of sequence
similarity as well as functional redundancy (Figure 1.1) (He, Hua et al. 2011).
The arrangement of HOX genes into clusters allows for enhancer sharing which
enables a precise spatial and temporal coordination of expression during development.
The relative regulatory dominance also varies between HOX genes, giving rise to what
is often referred to as a 'HOX code' (Knittel, Kessel et al. 1995), and resulting in the
following distinctive criteria: (1) Temporal distribution. The expression of HOX genes
starts from the 3' end of the cluster and proceeds stepwise towards the 5' end. (2) Spatial
distribution. The 3’ most member of the cluster is expressed with a more anterior limit
than the next member and each subsequent member has a more posterior limit of
expression resulting in an overlapping series of expression domains. (3) Posterior
prevalence. In each individual cluster, the function of the posterior gene products is
dominant over the more anterior genes (He, Hua et al. 2011).
8
Figure 1.2 A schematic structure of clustered HOX genes. The 39 HOX genes are located on four
different chromosomes. Homology of human HOX genes HOX-C to Drosophilia HOM-C genes is
showed by colours. Blank squares show missing genes (Previously published in Alharbi, Pettengell et al.
2012).
1.3 HOX Cofactors
DNA binding site studies suggest that HOX proteins have relatively limited
selectivity and specificity, and they need cofactor interactions in order to increase both
(Phelan, Rambaldi et al. 1995; Moens and Selleri 2006; Mann, Lelli et al. 2009). The
most important HOX-cofactors are the three amino acid loop extension proteins, which
comprise the pre B cell leukaemia (PBX) and myeloid ectropic insertion site (MEIS)
families (Moens and Selleri 2006; Mann, Lelli et al. 2009). These cofactors have
crucial roles in development and haematopoiesis (Thorsteinsdottir, Kroon et al. 2001).
For example, Pbx1 null mice die during the embryonic stage as a result of severe
haematopoietic defects (DiMartino, Selleri et al. 2001) and Meis1-deficient mice fail to
generate megakaryocytes, exhibit severe hemorrhaging and likewise die during the
embryonic stage (Hisa, Spence et al. 2004). In zebrafish, Meis1 and Pbx contribute to
Drosophilia(HOM-C)
Pb
Chromosome number
Iab Dfd Scr Antp Ubx Abd A Abd B
Antennapedia Complex (Ant-C) Bithorax Complex (BX-C)
1 32 5 6 7 10 11 12 1394HOX A
1 2 3 4 5 76 9 138
Mammalian (HOX-C)
HOX B
54 136 8 9 10 11 12
93 4 8 1110 12 13
7p15
17q21
12q13
2q31
HOX C
HOX D
3` 5`
9
the production of erythropoietic cells and the inhibition of myelopoiesis (Pillay,
Forrester et al. 2010). Generally, Hox proteins 1-10 bind with Pbx1 (Shen, Chang et al.
1996), whereas Hox proteins 9-13 bind with Meis1 (Shen, Montgomery et al. 1997).
Recently, it has been reported that HOXA6 binds to MEIS1 which indicates that MEIS1
interaction is not limited to the 5' end HOXs.(Dickson, Liberante et al. 2013).
1.4 HOX genes in haematopoiesis
HOX genes are expressed in HSCs and HPs in a manner reminiscent of their
expression in early development, with lineage and differentiation stage-restricted
manners. Thus, for example, HOXB3, HOXB4, and HOXA9 are highly expressed in
uncommitted haematopoietic cells, whereas HOXB8 and HOXA10 are expressed in
myeloid committed cells. Different mammalian CD34+ cell subpopulations express at
least 22 of the 39 HOX genes (Grier, Thompson et al. 2005). Anterior '3' end' HOX
genes (HOX1-6) are highly expressed in the most primitive HSCs. Subsequently, the
anterior HOX genes are downregulated, and posterior '5' end' HOX genes are expressed
during commitment (Sauvageau, Lansdorp et al. 1994). HOX genes are highly
expressed in the most primitive HSCs and HPs, while their expression is almost absent
in CD34- cells, which are considered differentiated bone marrow (BM) cells
(Sauvageau, Lansdorp et al. 1994; Pineault, Helgason et al. 2002; Helen Wheadon
2011). The analysis of HOX gene expression in human multipotent stem cells and T-
cell progenitors showed that HOXA genes are prominently expressed during T-cell
development, in particular AbdominalB-HOXA (AbdB-HOXA) including HOXA7-
HOXA11, with only HOXB3 and HOXC3 expressed from other HOX clusters (Taghon,
Thys et al. 2003).
10
1.4.1 Gain of function studies
The function of HOX genes in normal haematopoiesis has been widely studied
using gene expression analysis and knockin or knockout studies in HSCs and early HPs
(Table 1.1). Generally the over-expression of a HOX gene leads to an expansion of stem
and progenitor cell populations together with a block on differentiation. Notable
example of this include the over-expression of murine Hoxb6, which resulted in the
expansion of HSCs and myeloid progenitors, together with the inhibition of
erythropoiesis and lymphopoiesis (Fischbach, Rozenfeld et al. 2005), and over-
expression of murine Hoxb3 that resulted in several haematological abnormalities, such
as a block of B- and T-cell differentiation as well as a delay in MP proliferation
(Sauvageau, Thorsteinsdottir et al. 1997). Over-expression of human HOXC4 resulted
in expansion of early and committed myeloid and erythroid progenitors (Daga, Podesta
et al. 2000), and knockin of human HOXA5 caused an increase in the number of myeloid
progenitors and blocked erythroid differentiation (Crooks, Fuller et al. 1999; Fuller,
McAdara et al. 1999). Likewise, over-expression of HOXA10 in human cord blood or
fetal liver CD34+ HPs resulted in a significant production of blast cells and
myelopoiesis concomitant with a complete block of erythroid differentiation and a
severe reduction in B-cell development (Buske, Feuring-Buske et al. 2001). Other HOX
genes are required for the maintenance of progenitor or stem cell status and promote
their proliferation, especially HOXA9 and HOXB4. The former is the most
preferentially expressed HOX gene in human CD34+ HSCs and early HPs and is
subsequently down-regulated during differentiation. Murine Hoxa9 over-expression
enhances HSC expansion and myeloid progenitor proliferation and, with a long latency,
leads to leukaemia (Kroon, Krosl et al. 1998; Thorsteinsdottir, Mamo et al. 2002). In
11
contrast to myeloid progenitors, Hoxa9 over-expression resulted in a partial inhibition
of pre-B cell differentiation, but did not affect T-cell development (Thorsteinsdottir,
Mamo et al. 2002). Hoxb4 is also highly expressed in HSCs and down-regulated during
differentiation (Sauvageau, Lansdorp et al. 1994; Pineault, Helgason et al. 2002). Its
over-expression in murine and human cell lines results in a remarkable expansion of
HSCs in vivo and in vitro without resulting in leukaemia or lineage disturbances
(Sauvageau, Thorsteinsdottir et al. 1995; Amsellem, Pflumio et al. 2003). Indeed, the
self-renewal ability of Hoxb4-transduced HSCs is 20-50 fold greater than untreated
cells, and can be increased still further by knocking down Pbx1 (Krosl, Beslu et al.
2003; Cellot, Krosl et al. 2007).
1.4.2 Loss of function studies
In addition to the knockin and over-expression approaches described above,
knockdown and deletion studies in murine models and cell lines have also been used to
evaluate the role of HOX genes in haematopoiesis. However, owing to the functional
redundancy of HOX genes, the results of knockdown assays are sometimes difficult to
interpret and do not always reflect the findings of studies where the gene has been over-
expressed. For example, it has been found that Hoxb4 null mice exhibit a significant
reduction in size and cellularity of haematopoietic organs, such as spleen and liver, and
a slight decrease in HSCs and HPs number without a significant disturbance of lineage
commitment (Brun, Björnsson et al. 2004; Bijl, Thompson et al. 2006). Likewise,
Hoxb3 null mice display a notable reduction in B-cell progenitors, and a reduction in
BM cellularity, but no significant reduction in B-cell numbers in the spleen (Ko, Kwan
Lam et al. 2007). HOXb3/b4 -/- mice have a greater reduction in HSCs and HPs, yet no
12
difference in haematopoietic cell commitment (Bjornsson, Larsson et al. 2003). Using
polymerase chain reaction (PCR), it has been demonstrated that fetal liver cells (c-Kit+)
of Hoxb4 null mice expressed other Hoxb genes to a significantly higher level than
control cells (Bijl, Thompson et al. 2006). Thus, Hoxb3 and Hoxb4 can regulate the
normal function of HSCs but are not individually essential for the production of major
blood lineages because of redundancy with other members of the Hox gene family.
Bijl and colleagues (2006) found that an individual loss of Hoxb4 or even the
complete loss of the Hoxb cluster (b1-b9) did not affect the ability of murine fetal liver
HSCs to self-renew. The repopulation and differentiation potential were retained,
compared with wild-type control cells. Thus, the Hoxb cluster may not be necessary for
haematopoiesis, with members of the other Hox clusters presumably having largely
duplicate roles. Analysis of Hox gene expression in fetal liver cells (c-Kit+ Hoxb1-
Hoxb9-/-), revealed genetic interactions between members of the Hoxa, Hoxb and Hoxc
clusters, whereby these cells exhibited down-regulation of all Hoxa genes, except
Hoxa13, and up-regulation of Hoxc4, Hoxc9 and Hoxc11, also suggesting functional
redundancy and complex genetic interactions between Hox genes.
An exception to the general prevalence of functional redundancy amongst the
HOX gene family is HOXA9, the most highly expressed HOX family member in HSCs.
Hoxa9-/- mice showed significant deficiencies in myeloid and lymphoid cells
concomitant with a significant defect in repopulating ability (So, Karsunky et al. 2004;
Lawrence, Christensen et al. 2005). These deficiencies include commom myeloid
progenitors, granulocyte/monocyte progenitors, common lymphoid progenitors and
lymphoid progenitors (pro- and pre-B cells, pro-T cells) (So, Karsunky et al. 2004).
There is also a corresponding reduction in spleen cellularity and size (Magnusson, Brun
13
et al. 2007). Notably, compared to the entire cluster Hoxb deficient fetal liver HSCs,
Hoxa9-/- fetal liver HSCs exhibited a more dramatic defect in repopulating ability (Bijl,
Thompson et al. 2006; Magnusson, Brun et al. 2007), and HSCs from Hoxa9/b3/b4 null
mice had the same repopulating ability as those from Hoxa9 null mice (Magnusson,
Brun et al. 2007). Gene knockdown studies have revealed that some additional HOX
genes are also essential in normal haematopoiesis. Knockdown of HOXA5 led to an
increase in erythroid progeneitors and a reduction in the number of myelomonocytic
cells (Crooks, Fuller et al. 1999; Fuller, McAdara et al. 1999), and Hoxa7 null mice
showed a reduction in megakaryocytic/erythroid progenitors as well as reticulocytosis
and thrombocytopenia (So, Karsunky et al. 2004). Knockout of Hoxb6 resulted in an
increase in early erythroid progenitors in murine BM and fetal liver cells (Kappen
2000). Likewise, Hoxc3-/- mice showed a reduction in late erythroid progenitors without
affecting the haemoglobinization size (Takeshita, Bollekens et al. 1993), and Hoxc8
deficient mice showed a significant reduction in erythroid, granulocyte and macrophage
colony formation potential (Shimamoto, Tang et al. 1999).
14
Table 1.3 HOX gene studies (Previously published in Alharbi, Pettengell et al. 2012).
HOX gene Gain of function Loss of function Species
HOXA5 ↑Myeloid progenitors and
block erythroid
differentiation (Crooks,
Fuller et al. 1999; Fuller,
McAdara et al. 1999)
↑Erythroid progenitors and
↓myelomonocytic cells (Crooks,
Fuller et al. 1999; Fuller, McAdara
et al. 1999)
Human
Hoxa7 ↓ MEP, reticulocytosis and
thrombocytopenia (So, Karsunky
et al. 2004)
Mouse
Hoxa9 ↑HSCs expansion and
myeloid progenitor
proliferation.
Block erythroid
differentiation (Kroon,
Krosl et al. 1998;
Thorsteinsdottir, Mamo et
al. 2002).
↓↓CMP, GMP, CLP, lymphoid
progenitors, repopulating ability
and spleen cellularity and size (So,
Karsunky et al. 2004; Lawrence,
Christensen et al. 2005).
Mouse
HOXA10 ↑↑ Blast cells and
myelopoiesis, ↓ B cell
differentiation and block
erythroid differentiation
(Buske, Feuring-Buske et
al. 2001).
Human
Hoxb3 Block B- and T-cell
differentiation and a delay
in myeloid progenitor
proliferation (Sauvageau,
Thorsteinsdottir et al. 1997).
↓↓ B-cell progenitors and bone
marrow cellularity (Ko, Kwan
Lam et al. 2007).
Mouse
HOXB4/
Hoxb4
↑↑ HSCs expansion
(Sauvageau,
Thorsteinsdottir et al. 1995;
Amsellem, Pflumio et al.
2003).
↓↓ Haematopoietic organs
cellularity and size, ↓ HSCs and
HPCs and ↑ Hoxb genes (Brun,
Björnsson et al. 2004; Bijl,
Thompson et al. 2006).
Human /
mouse
Hoxb3/b4 ↓↓ HSCs and HPCs (Bjornsson,
Larsson et al. 2003).
Mouse
Hoxb6 ↑HSCs expansion and
myeloid progenitors.
↓ erythropoiesis and
lymphopoiesis (Fischbach,
Rozenfeld et al. 2005).
↑ Early erythroid progenitors
(Kappen 2000).
Mouse
Hoxc3 ↓ Erythroid progenitors (Takeshita,
Bollekens et al. 1993).
Mouse
HOXC4 ↑ Early and committed
myeloid and erythroid
progenitors (Daga, Podesta
et al. 2000).
Human
Hoxc8 ↓Erythroid, granulocyte and
macrophage colony formation
potential (Shimamoto, Tang et al.
1999).
Mouse
15
1.5 Upstream regulators of HOX genes
Knockout models of HOX gene upstream regulators have helped to define their role in
normal haematopoiesis. Regulators include transcriptional activators such as mixed
lineage, myeloid lymphoid, leukaemia (MLL), and a family of caudal-type homebox
transcription factors (CDX1, CDX2, and CDX4). The existence of HOX genes in
clusters makes them particularly sensitive to changes in chromosomal organization, and
repressors of HOX transcription include genes that mediate this process, most notably
members of the polycomb group (Beuchle, Struhl et al. 2001). These regulators have
crucial roles in normal development and haematopoiesis through the regulation of HOX
genes. A number of studies demonstrated that Mll-deficient embryonic bodies and Mll-
conditional knockout mice showed a dramatic reduction in HSCs and HPs. In addition,
these embryonic bodies and the mice exhibited greatly reduced expression of a number
of Hox genes including Hoxa7, Hoxa9, Hoxa10 and other Hoxb and Hoxc genes (Ernst,
Mabon et al. 2004; Jude, Climer et al. 2007). Likewise, Cdx compound-deficient
zebrafish and murine embryonic stem cells showed dysregulation of the embryonic HPs
as well as impaired expression of Hox genes (Davidson and Zon 2006; Wang, Yabuuchi
et al. 2008). However, it has been shown that Cdxs are not essential for normal
haematopoiesis in adult mice. For example, Cdx4-deficient mice showed minimal
haematopoietic defects, though it was highly expressed in wild-type myeloid progenitor
cells (Koo, Huntly et al. 2010). In addition, human CDX2 is not expressed in normal
HSCs, or in myeloid, B-cell, or T-cell progenitors (Scholl, Bansal et al. 2007; Rawat,
Thoene et al. 2008). In addition, it has been found that HOX gene expression is also
regulated by small single-stranded RNAs (miRNAs) and the non-coding RNA
HOTAIR (Bhatlekar, Fields et al. 2014).
16
miRNAs regulate HOX expression by repressing the expression of anterior genes,
thereby supporting posterior prevalence while HOTAIR was reported as a negative
regulator of HOX expression (Yekta, Tabin et al. 2008; Nakayama, Shibazaki et al.
2013).
1.6 HOX downstream target genes in haematopoietic cells
The mechanism by which HOX proteins regulate haematopoiesis is not yet fully
understood. However, genome-wide analyses after experimentally induced changes in
HOX genes expression have identified some potential downstream targets. Amongst
these are the HOX genes themselves, some of which have been shown to cross-regulate
their neighbours, or their cofactors. HOXA9, HOXA10 and HOXB4 are the most
comprehensively studied genes in this respect because of their key roles in normal
haematopoiesis and leukaemia. It is particularly noteworthy that HOXA9 positively
regulates the transcription of other HOX genes including HOXA7 and HOXA10 and its
cofactor PBX3 and MEIS1 (Faber, Krivtsov et al. 2009). A summary of HOX
downstream target genes is presented in Table 1.2.
As described above, HOXA9 is a key regulator of haematopoiesis and behaves
as an oncogene in leukaemia. It is therefore unsurprising that it activates the
transcription of genes known to regulate cell proliferation and survival. For example,
Hoxa9 directly activates the Pim1 gene, the product of which enhances proliferation by
activating c-Myb, and also exerts an anti-apoptotic effect by phosphorylating and
inactivating the BAD protein (Leverson, Koskinen et al. 1998; Hu, Passegué et al.
2007). c-Myb has also been identified as an indirect transcriptional target of Hoxa9-
Mies1 that mediates transformation in Mll-Enl leukaemia (Hess, Bittner et al. 2006).
Other HOXA9 targets include the oncogene ID2, which is up-regulated, and BIM,
17
which encodes an apoptotic factor and is down-regulated (Nagel, Venturini et al. 2010).
HOXA9 also activates the CYBB gene, which encodes the Gp91phox (a phagocyte
respiratory burst oxidase protein), and is expressed in differentiated myeloid cells (Bei,
Lu et al. 2005). In mice, Hoxa9 has been shown to directly activate the transcription of
the flt3 gene, which is associated with an unfavourable prognosis of AML (Gwin, Frank
et al. 2010) and it also regulates its own cofactor, Meis1, through binding to Meis1
upstream regulator genes cerb1 and pknox1 (Hu, Fong et al. 2009). More proliferative
genes have been identified as downstream targets for Hoxa9 including Camk2d, Cdk6,
Erg, Etv6, Flt3, Foxp1, Gfi1, Kit, Lck, Lmo2, Myb and Sox4 (Huang, Sitwala et al.
2012). In the same study, it was shown that Hoxa9 down-regulates differentiation and
inflammation genes including Ifit1, Tlr4, Ccl3, Ccl4, Csf2rb, Ifngr1, Runx1, Cd28 and
Cd33. HOXA9 also regulates the anti-apoptotic gene Bcl-2 which may explain the cell
survival role of HOXA9 (Brumatti, Salmanidis et al. 2013). The fusion protein
nucloporin 98 (NUP98-HOXA9) has been found to stimulate the proliferation of HSCs
by activating the expression of other HOX genes; including HOXA9, HOXA7, MEIS1
and PBX3. It also up-regulates a number of leukaemogenic transcription factors
including EVI1 and MEF2C, and receptor tyrosine kinases including FLT3 and KIT
(Takeda, Goolsby et al. 2006).
It is also of note that there are both overlapping and opposing functions between
the closely related HOXA9 and HOXA10 transcription factors. For example, in a similar
manner to Hoxa9, Hoxa10 activates the expression of proliferative genes that result in
myeloid progenitor expansion such as Itgb3, Hif, Tgfβ2 and Fgf2 by direct binding to
their promoters (Bei, Lu et al. 2007; Magnusson, Brun et al. 2007; Shah, Wang et al.
2011; Shah, Bei et al. 2012). HOXA10 also activates the transcription of anti-apoptotic
18
genes such as DUSP4 which encodes mitogen-activated protein phosphatase 2.
mitogen-activated protein phosphatase 2 in turn prevents cell death by down-regulating
JNK (Wang, Lu et al. 2007). Hoxa10 decreases erythroid differentiation and
megakaryopoiesis by activating Hoxa5 and inactivating Gata-1, respectively
(Magnusson, Brun et al. 2007), and it also induces Cdx4 expression in myeloid cells
(Bei, Huang et al. 2011).
Unlike HOXA9, HOXA10 can also exert anti-proliferative effects. For
example, in cooperation with its trimeric cofactors, HOXA10 induces p21 transcription
leading to cell cycle arrest and differentiation (Bromleigh and Freedman 2000). It also
represses CYBB transcription (Eklund, Jalava et al. 2000), thereby acting in an opposing
manner to HOXA9. In a fusion form with Nup98, Hoxa10 activates more than 400
genes including the self-renewal genes Flt3, Prnp, Hlf and Jag2 (Palmqvist, Pineault et
al. 2007).
Many HOXB4 target genes have also been identified in three studies (Lee,
Zhang et al. 2010; Oshima, Endoh et al. 2011; Fan, Bonde et al. 2012). Over-expression
of Hoxb4 resulted in transcriptional up-regulation of Meis1, Dntt, Hlf, Sox4 and Runx2,
while it down-regulated the transcription of lymphoid specific genes, such as B220, and
myeloid-specific genes, such as Hmbs (Lee, Zhang et al. 2010). Some HOXB4 targets
seem to vary in a context-dependent manner, for example, it has been found to down-
regulate the transcription of c-MYC in the HL-60 cell line leading to cell differentiation
(Pan and Simpson 2001), while it activates c-Myc transcription in murine BM cells
(Satoh, Matsumura et al. 2004). As with HOXA9, Hoxb4 activates the transcription of
its neighbouring genes, Hoxb2, Hoxb3 and Hoxb5 (Satoh, Matsumura et al. 2004).
Hoxb4 also activates activator protein-1 (AP-1) complex members Fra-1 and Jun-B,
19
which leads in turn to an increase in the level of cyclin-D1 and a decrease the level of
c-Fos transcription, thereby increasing the proliferation capacity of HSCs (Krosl and
Sauvageau 2000).
20
Table 1.4 A summary of mammalian HOX target genes (Alharbi, Pettengell et al. 2012).
HOX protein Targets of transcriptional activation Targets of transcriptional repression Species
HOXA9/ Hoxa9 Pim1 (Hu, Passegué et al. 2007), ID2 (Nagel,
Venturini et al. 2010), CYBB (Bei, Lu et al.
2005), HOXA7, HOXA10, PBX3, MEIS1
(Faber, Krivtsov et al. 2009). Bcl-2
(Brumatti, Salmanidis et al. 2013), Flt3
(Gwin, Frank et al. 2010), Cerb1 and Pknox1
(Bei, Lu et al. 2005), Camk2d, Cdk6, Erg,
Etv6, Foxp1, Gfi1, Kit, Lck, Lmo2, Myb and
Sox4 (Huang, Sitwala et al. 2012).
BIM (Nagel, Venturini et al. 2010)/
Itfi1, Tlr4, Ccl3, Ccl4, Csf2rb, Ifngr1,
Runx1, Cd28, Cd33 (Huang, Sitwala et
al. 2012).
Human/
mouse
Hoxa9-Meis1 c-Myb (Hess, Bittner et al. 2006). Mouse
NUP98-HOXA9 HOXA7, HOXA9, MEIS1, PBX3, EVI1,
MEF2C, FLT3 and KIT (Takeda, Goolsby et
al. 2006).
Human
HOXA10/
Hoxa10
P21 (Bromleigh and Freedman 2000),
DUSP4 (Wang, Lu et al. 2007), Itgb3 (Bei,
Lu et al. 2007), Hlf (Magnusson, Brun et al.
2007), Tgfβ2 (Shah, Wang et al. 2011), Fgf2
(Shah, Bei et al. 2012), Dusp4 (Wang, Lu et
al. 2007), Hoxa5 (Magnusson, Brun et al.
2007), Cdx4 (Bei, Huang et al. 2011).
CYBB (Eklund, Jalava et al. 2000)/
Gata1 (Magnusson, Brun et al. 2007)
Human/
mouse
Nup-Hoxa10 Flt3, Prnp, Hlf and Jag2 (Palmqvist, Pineault
et al. 2007).
Mouse
HOXB4/ Hoxb4 MEIS1, DNTT, HLF, SOX4, RUNX2 (Lee,
Zhang et al. 2010), c-MYC (Pan and Simpson
2001), Hoxb2, Hoxb3 (Satoh, Matsumura et
al. 2004), Fra-1, JunB (Krosl and Sauvageau
2000).
B220 and HMBS (Lee, Zhang et al.
2010), c-Myc (Satoh, Matsumura et al.
2004).
Human/
mouse
21
1.7 The role of HOX genes in acute leukaemia
Numerous studies have now shown that HOX genes can promote the
development of AML by forming chimeric fusions with other genes, but more recent
work has also shown that their miss-expression, in particular their over-expression, is
also important in the formation of malignancy.
1.7.1 HOX fusion proteins
One of the most frequent fusion partners for HOX genes is nucloporin (NUP98),
a member of the nuclear pore family (Figure 1.2). It is localized in the nuclear
membrane and functions as a selective transporter for ribonucleic acid (RNA) and
proteins between the nucleus and cytoplasm. NUP98-HOX fusion proteins have been
reported in AML and other leukaemias. In AML, NUP98-HOXA9 is associated with a
t(7;11)(p15;p15) translocation (Borrow, Shearman et al. 1996; Nakamura, Largaespada
et al. 1996). There are eight other HOX genes that can be fused with NUP98, including
HOXA11 and HOXA13 (Fujino, Suzuki et al. 2002; Suzuki, Ito et al. 2002), HOXD11
and HOXD13 (Raza-Egilmez, Jani-Sait et al. 1998; Taketani, Taki et al. 2002), and
HOXC13 (Taketani, Taki et al. 2002). Thus only the 5’ most members of each HOX
complex have been documented to be fused with NUP98 in AML. However, Hoxb3
has been shown to be a potential leukaemogenic partner with Nup98 (Pineault,
Abramovich et al. 2004), suggesting that the ability to be a fusion NUP98 partner is not
limited to the 5’ most HOX genes. Generally, NUP98-HOX fusions induce cell
proliferation and function as transcriptional activators and NUP98 fusions with HOX
proteins are more oncogenic than fusions with other partners (Saw, Curtis et al. 2013).
22
Nup98-Hox fusion proteins result in AML with a long latency, around 11-12
months (Kroon, Thorsteinsdottir et al. 2001). However, this latency can be reduced to
two months by co-over-expression of the Hox cofactor Meis1 (Pineault, Buske et al.
2003) and the receptor tyrosine kinase Flt3 (Palmqvist, Argiropoulos et al. 2006). For
example, concurrent translocation of Nup98-Hoxd13 and Flt3-ITD developed AML in
3 months (Greenblatt, Li et al. 2012). FLT3 has an essential role in the regulation of
early HPs growth, and causes increased and uncontrolled self-renewal of these cells
through a FLT3 ligand-independent pathway (Tosic, Stojiljkovic et al. 2009).
Figure 1.3 structures of AbdB-HOX, NUP98 and the predictive fusion protein NUP98-HOXA. A) A
general structure of AbdB-HOX (9-13) proteins that have been reported to fuse with NUP98. B) Structure
of normal NUP98 protein. C) Structure of predictive NUP98-HOX fusion protein. This fusion eliminates
MD and RBD from AbdB and NUP98, respectively. The arrows represent the breakpoints. MD: MEIS
domain, PM: Pbx motif, H: hexapeptide, HD: homedomain, FG: phenylalanine-glycine, GLEBS: gle2p-
binding-like motif, RBD: RNA binding domain (Previously published in Alharbi, Pettengell et al. 2012).
PMMD COOHNH2
HOX
GLEBS RBDFGNH2 COOH
NUP98
HDPMGLEBS COOHNH2
NUP98-HOX
A)
B)
C)
FG
FG FG
HDH
H
23
1.7.2 HOX over-expression in AML
HOX genes may also be indirectly involved in AML through chromosomal
rearrangements that involve their upstream regulators, such as MLL. MLL fusion
proteins constitute about 10% of therapy related AML and 3% of de novo AML (Slany
2009). There are more than 64 translocation partner genes that fuse with the MLL N-
terminus (Meyer, Kowarz et al. 2009). Normally, Mll positively regulates the
transcription of Hox genes by maintaining their expression through direct binding to a
proximal promoter region (Milne, Briggs et al. 2002). MLL fusion proteins activate
HOX gene transcription more efficiently than MLL alone (Liu, Cheng et al. 2009; Slany
2009), especially the 5’ end members of the HOXA cluster, together with their co-
activator MEIS1. As a consequence, myeloid differentiation is blocked and proliferation
is enhanced (Marschalek 2011). Consistent with this proposed mechanism, it has been
reported that MLL-AF9, like NUP98-HOXA9, leads to a block in erythroid/myeloid
maturation and to erythroid hyperplasia (Abdul-Nabi, Yassin et al. 2010), and the Mll-
Enl fusion protein requires Hoxa7 and Hoxa9 for efficient immortalization of HPs
(Ayton and Cleary 2003). Conversely, a number of studies demonstrated that the
expression of HOXA genes is not crucial for MLL leukaemogenesis, yet their
expression affects disease phenotype. For instance, Hoxa7 and Hoxa9 influence AML
latency and phenotype; yet they are not essential to initiate Mll-Gas7-mediated
leukaemogenesis (So, Karsunky et al. 2004). Furthermore, suppression of HOXA9
expression in cells with a chimeric MLL-AF9 gene reduces the survival of leukaemic
cells and changes the disease phenotype, but it does not affect AML initiation (Faber,
Krivtsov et al. 2009).
24
The dysregulation of another regulator of HOX genes, the CDX gene family, has
also been shown to drive the development of AML. CDX2 is expressed in the majority
of AML cases (90%), but not in normal adult haematopoiesis, and Cdx2-elevated
expression leads to AML with only a short latency period (Scholl, Bansal et al. 2007).
In contrast, the closely related CDX4 gene is expressed in 25% of AML cases, and is
expressed in normal adult haematopoiesis, and Cdx4 over-expression in murine BM
results in AML but only with a long latency period (Bansal, Scholl et al. 2006). This
latency can be accelerated in mice through cooperation of Meis1 which results in the
over-expression of a number of Hox genes including Hoxa6, Hoxa7, Hoxa9, Hoxb4,
Hoxb8 and Hoxc6 (Bansal, Scholl et al. 2006). Cdx2 expression alone is sufficient to
drive the up-regulation of a related set of HOX genes (Rawat, Thoene et al. 2008),
demonstrating the importance of the Cdx family in the dysregulation of Hox genes
during AML.
The dysregulation of HOX gene expression is also associated with the
nucleophosmin 1 (NPM1) mutation. NPM1 is a chaperone protein that shuttles between
the nucleus and cytoplasm, although its predominant localization is in the nucleus (Rau
and Brown 2009). NPM1 has a crucial role in several biological processes, such as
ribosome biogenesis, genomic stability and cell cycle progression. In adult AML,
NPM1 mutation is the most common genetic aberration, reported in about 35% of all
adult AML and approximately in 45-55% of NK-AML (Falini, Mecucci et al. 2005). In
pediatric AML, NPM1 mutations are significantly less common, occurring in 8-10% of
cases, and in about a quarter of normal karyotype cases (Brown, McIntyre et al. 2007;
Hollink, Zwaan et al. 2009). The relocation of NPM1 into the cytoplasm (NPMc+)
occurs only in AML (Falini, Bolli et al. 2009). This relocation causes up-regulation of
25
a number of HOX genes, some of which are similar to those seen in AML initiated by
a MLL chimeric gene fusion, while some are distinct. Thus for example, HOXA4,
HOXA6, HOXA7, HOXA9, HOXB9 and MEIS1 are over-expressed in both contexts,
while HOXB2, HOXB3, HOXB5, HOXB6 and HOXD4 are up-regulated in NPMc+
AML only (Mullighan, Kennedy et al. 2007). It has been reported that activation of a
humanized Npm1 allele led to over-expression of Hoxa5, Hoxa7, Hoxa9 and Hoxa10,
induction of HSC self-renewal and the expansion of myelopoiesis (Vassiliou, Cooper
et al. 2011). The exact mechanism of the association between the NPM1 mutation and
the up-regulation of HOX genes is still unclear. A possible explanation is that NPM1
directly disturbs the expression of HOX genes, or alternatively, that NPM1 mutations
arrest the differentiation of early HPs in which HOX expression is up-regulated (Rau
and Brown 2009).
1.7.3 HOX gene dysregulation in acute lymphoid leukaemia (ALL)
The dysregulation of HOX genes has also been reported in ALL including both
B- and T-progenitor ALL (B-ALL and T-ALL), especially when MLL translocations
are involved. For example, human HOXA9, HOXA10 and HOXC6, and their cofactor
MEIS1 were up-regulated in both MLL-ENL T-ALL and MLL B-ALL (Ferrando,
Armstrong et al. 2003). Human HOXA9 and MEIS1, and murine Hoxa5, Hoxa9 and
Meis1 were likewise up-regulated in MLL-AF4 B-ALL (Rozovskaia, Feinstein et al.
2001; Krivtsov, Feng et al. 2008). A common signature of MLL rearrangements in
AML and ALL is the over-expression of HOXA genes including HOXA3, HOXA5,
HOXA7, HOXA9 and HOXA10 (Zangrando, Dell'Orto et al. 2009). In addition, HOX
genes can be involved in other translocations such as CALM-AF10 T-ALL, where
human HOXA5, HOXA9, HOXA10 and MEIS1 are over-expressed (Dik, Brahim et al.
26
2005). HOXA proteins can also form chimeric fusion proteins with T-cell receptor in
T-ALL, which results in the global over-expression of HOXA genes (Soulier, Clappier
et al. 2005; Speleman, Cauwelier et al. 2005). Surprisingly, the aberrant expression of
CDX2 in ALL, an upstream regulator of HOX genes, is not correlated with HOX gene
dysregulation (Thoene, Rawat et al. 2009).
1.7.4 HOX genes as prognostic markers
HOX gene expression has become an important prognostic factor in AML.
Over-expression of HOX genes is associated with an intermediate/unfavorable
cytogenetic subset of AML. For example, among 6817 genes that have been
investigated in AML patients, the single gene correlated with the worst outcome and
relapse of disease as well as short survival was HOXA9 (Golub, Slonim et al. 1999).
Likewise, high expression of HOXA9 associates with low CR rate (Li, Li et al. 2013).
Correspondingly, low HOXA9 expression was found to correlate with the best outcome
and response to therapy (Andreeff, Ruvolo et al. 2008). It is also noteworthy that low
expression of HOXA4 and MEIS1 are also favorable predictors for AML patient
outcome (Zangenberg, Grubach et al. 2009). In addition, a signature of up-regulation
of HOXA6, HOXA9 and PBX3 with a low expression of MEIS1 is an independent poor
prognostic marker in NK-AML, and simultaneous knockdown of HOXA6 and HOXA9
caused cell death and AML cell lines became more sensitive to cytarabine (Dickson,
Liberante et al. 2013). Furthermore, high expression of HOXA7, HOXA9 and HOXA11
along with the cofactor PBX3 is an independent prognosis marker of adverse OS in
abnormal karyotype AML (Li, Huang et al. 2012). The global levels of HOX expression
also seem to reflect the outcome of disease, possibly reflecting the functional
redundancy exhibited by many of the HOX genes. Thus the highest levels of HOX genes
27
are seen in FLT3 mutation cases, which have unfavorable outcomes, and generally the
HOX genes are expressed only at a very low level in favorable subsets of AML.
Although recent work has established the importance of HOX genes in the
development of AML, it is still not clear exactly what the functions of these genes are
beyond a general inhibition of differentiation and an increase in cell proliferation. This
lack of precise mechanistic knowledge for individual HOX genes may be owing to the
functional redundancy showed by many members of this family, making the
knockdown of single HOX genes generally difficult to interpret, and it may also help to
explain the contrast in gene knockin and knockout results. In myeloma and some solid
malignancies this has been addressed by targeting the HOX proteins as a group by
antagonizing their interactions with the PBX cofactor using the peptide inhibitor HXR9
(Morgan, Pirard et al. 2007; Shears, Plowright et al. 2008; Plowright, Harrington et al.
2009; Daniels, Neacato et al. 2010; Morgan, Plowright et al. 2010).
PBX binds to PBX motif in HOX paralogus group 1-10. HOX-PBX dimers have
more binding affinity and specificity for target DNA sequences than the HOX monomer
alone. Of note, HOX-PBX dimers are strictly involved the interaction of PBX to the
conserved hexapeptide sequence WYPWMK that is found N terminal to the
homeodomain of HOX proteins from paralogous groups 1-10 for cooperative DNA
binding by PBX and HOX partners (Chang, Shen et al. 1995; Shen, Rozenfeld et al.
1997; Medina-Martinez and Ramirez-Solis 2003). The peptide inhibitor HXR9 contains
a hexapeptide sequence mimics the HOX protein hexapeptide sequences that binds to
PBX and thereby interfering the DNA binding of HOX-PBX. HXR9 is cytotoxic to
cells, predominantly through the induction of apoptosis, which in turn depends upon a
rapid increase in expression of c-FOS (Morgan, Pirard et al. 2007; Shears, Plowright et
28
al. 2008; Espinosa, Shinohara et al. 2009; Plowright, Harrington et al. 2009; Daniels,
Neacato et al. 2010). A similar approach may be useful in AML for addressing
redundant functions of HOX genes.
1.8 Hypothesis and aims
The hypothesis of this study was that AML would also be sensitive to killing by
HXR9, and indeed it may be that the HOX genes could represent a useful therapeutic
target, especially in AMLs that show high levels of HOX expression and a
correspondingly poor prognosis. In order to assess this hypothesis, three main
objectives were set up.
The first objective of this study was to investigate the cytotoxicity of HXR9 on
AML cell lines derived from different AML subtypes and if so to investigate the
combination therapy of HXR9 with AML first line chemotherapy. The strategies for
conducting this were:
To measure the expression of the whole panel of 39 HOX genes in five
AML cell lines.
To investigate the cytotoxicity of HXR9 on the five cell lines either
alone or in combination with AML first line chemotherapy.
The second objective was to investigate the mechanism of HXR9 cytotoxicity
on the AML-derived cell lines. The strategies for conducting this were:
To determine whether HXR9 induced cell death through apoptosis or
necrosis.
29
To assess the involvement of some signaling pathways such as MAPK
and pro-apoptotic proteins such as p53.
The third objective was to investigate the efficacy of HXR9 against AML
disease in vivo. The strategies for conducting this were:
To establish an AML model.
To assess the efficacy of HXR9 either alone or with co-treatment with
some non-toxic inhibitors.
30
Chapter 2 Materials and Methods
31
2.1 Materials
2.1.1 Reagents
Table 2.1 Reagents were used in this study and their suppliers.
Reagent Supplier
SYBR® green jumpstatTM Taq ready mixTM Sigma (Dorset, UK)
Penicillin-streptomycin
μI-1640
DMEM
Trypan blue
HBSS
Histopaque®-1077 Hybri-Max
β-mercaptoethanol
Daunorubicin
Mitoxantrone
DAPI stain
z-VAD-FMK
Cyclosporin A
Necrostatin-1
Fructose
EDTA
U0126 monoethanolate
SP600125
SB 203580
DPI
Calpain inhibitor I
W-7
Ro31-8220 methanesulfonate and pifithrin-α
32
Reagent Supplier
Protoporphyrin IX Sigma (Dorset, UK)
Carestream® Kodak BioMax® light film
Fixer and replenisher
Developer and replenisher
Cloned AMV first-strand synthesis kit invitrogen, Thermo scientific (Paisley, UK)
Glutamine
FBS
IMDM medium
Enz Chek® Caspase-3 Assay Kit#2
NuPAGE® LDS sample buffer
BenchmarkTM prestained protein ladder
NuPAGE novex 12% Bis Tris
DMSO
Pierce® RIPA buffer
Protease inhibitor cocktail
NuPAGE® sample reducing agent
NuPAGE® MES SDS running buffer
InvitrolonTM PVDF filter paper sandwich
Novex cryoTubeTM vials
SuperSignal® West Pico Chemiluminescent substrate
Annexin V-PE apoptosis detection kit BD pharmingen (Cambridge, UK)
Cytotoxicity detection kit Roche (Mannheim, Germany)
RNeasy mini kit and RBC lysis solution Qiagen (West Sussex, UK)
Gene Primers Eurgentec (Seraing, Belgium)
Mycoplasma detection kit lonza (Slough, UK)
33
2.1.2 Instruments
Instruments used in this study were: Beckman coulter a plate reader, Miltenyibiotec a
fluorescence-activated cell sorting (FACS) machine, Lebtech a Nanodrop spectrophotometer
ND-1000, and Stratagene Mx 3005P, SHADON cytospin 4, a fluorescent microscope,
eppendorf centrifuge 5415R.
2.1.3 Cell lines
Human and mouse derived AML cell lines were used in this study. Human derived cell
lines used were primary or secondary. Primary AML cell lines derived from patients with de
novo mutations including KG-1, a cell line derived from an erythroleukaemia (Koeffler and
Golde 1978), HEL 92.1.7, an erythroleukaemia cell line (Martin and Papayannopoulou 1982)
and HL-60, a cell line obtained from acute promyelocytic leukaemia (APML) (Collins, Ruscetti
et al. 1978). Secondary AML cell lines are derived from patients that developed AML from
MDS, chronic myeloid leukaemia (CML) or following previous treatment with chemotherapy.
These include K562, a CML cell in blast crisis (Lozzio and Lozzio 1975) and KU812F, also a
CML cell in blast crisis (Kishi 1985). The mouse derived cell line G1498-GFP was a kind gift
from Dr. Justin Kline (Department of Medicine Section of Haematology/Oncology, Chicago,
USA).
2.1.4 HXR9 and CXR9 peptide synthesis
The synthesis of these peptides has been described previously (Morgan, Pirard et al.
2007). Briefly, HXR9 is a short peptide of 18 amino acids, comprising a hexapeptide sequence
and nine C-terminal arginine residues (R9) that enables cell penetration. The hexapeptide
mimics the hexapeptide sequence of HOX proteins of paralogs 1-10, this sequence functions as
a competitive inhibitor of HOX/PBX. CXR9 is a control peptide that does not have a functional
hexapeptide portion but has the R9. The stability of the peptides was increased using D-isomers
34
NH2- and COOH- terminals. The half-life of both peptides is about 12 hours at the serum
(Morgan, Pirard et al. 2007). A conventional column-based technique was used to synthesise
HXR9 and CXR9 and were purified to at least 80% (Biosynthesis Inc., Lewisville, TX, USA).
The peptide sequences are as follows:
HXR9: WYPWMKKHHRRRRRRRRR (2700.06 Da)
CXR9: WYPAMKKHHRRRRRRRRR (2604.14 Da)
2.1.5 Mice
Female C57BL/6, nude and severe combined immune-deficient (SCID) mice,
approximately 6 to 8 weeks age, were purchased from Jackson laboratory, Kent, UK. These
mice were maintained in pathogen-free cages and fed with irradiated food and acidified water.
All experiments were performed under the compliance of Home Office and institutional
instructions.
2.2 Methods
2.2.1 General cell culture methods
2.2.1.1 Routine cell culture
All cell culture techniques were performed in a class II microbiological safety cabinet
and all equipment and reagent bottles were sterilised using 70% ethanol solution before use.
K562, KU812F and HEL92.1.7 cells were grown in a culture medium consisting of Roswell
Park Memorial Institute (RPMI-1640) medium complemented with 10% fetal bovine serum
(FBS), 5% L-glutamine and 5% antibiotics (1000 units of penicillin/ml and 10 mg/ml
streptomycin, P/S). KG-1 and HL-60 cells were cultured in Iscove’s Modified Dulbecco’s
medium (IMDM) supplemented with 20% FBS and 5% antibiotics. C1498-GFP cells were
maintained as a suspension culture in Dulbecco’s Modified Eagle’s Medium (DMEM)
35
supplemented with 10% FBS, 5% L-glutamine and 5% antibiotics. All cell lines were cultured
in T-75 sterile cell culture flasks at 37°C in a humidified environment containing 5% CO2 and
95% air. Cells were sub-cultured routinely three times a week (Table 2.2). Mycoplasma
contamination test was carried out at monthly intervals using MycoAlert™ Mycoplasma
Detection Kit.
Table 2.2 Cell lines used in this study. This table shows sources and diseases that cell lines were derived from.
All cell lines were cultured at 37°C, 5% CO2 and split 3 times a week.
Cell line Source Disease Culure media References
KG-1 Human Erythroleukaemia IMDM, 20%FBS, P/S (Koeffler and
Golde 1978)
HEL92.1.7 Human Erythroleukaemia RPMI, 10%FBS, P/S
and L-glutmaine
(Martin and
Papayannopoulou
1982)
HL-60 Human APML IMDM, 20%FBS, P/S (Collins, Ruscetti
et al. 1978)
KU812F Human CML in blast
crisis
RPMI, 10%FBS, P/S
and L-glutmaine
(Kishi 1985)
K562 Human CML in blast
crisis
RPMI, 10%FBS, P/S
and L-glutmaine
(Lozzio and
Lozzio 1975)
C1498-GFP Murine AML DMEM, 10%FBS,
P/S and L-glutmaine
(Zhang, Gajewski
et al. 2009)
36
2.2.1.2 Cell counting and cell density calculation using a haemocytometer
A 10μl aliquot of cell suspension was mixed with 90μl of trypan blue stain and 10μl of
the mixture was loaded onto the grid of a Neubauer haemocytometer. Only healthy and round
cells that were located in the four corner squares of the Neubauer haemocytometer were counted
and averaged in order to determine the cell density (Figure 2.1).
Figure 2.1 A haemocytometer diagram. It indicates the four corner squares that were used for counting cell
number; only cells that were healthy and round cells that only were counted. The number of cells in one set of 16
squares is equal to the number of cells x104/ml.
2.2.1.3 Cryopreservation of cell stocks
After calculating the cell density, a volume containing 5x106 cells was transferred into a
25ml universal tube and centrifuged at 1500rpm for 3 minutes. Then, supernatant was removed
and cell pellets were re-suspended in 1ml storage medium, which was full growth medium
supplemented with 5% (v/v) dimethyl sulphoxide (DMSO). The cell suspension was then
transferred into 1.8ml cryovials. Cells were stored at -80ºC for overnight before being
transferred to liquid nitrogen for long term storage.
37
2.2.1.4 Revitalisation of cryopreserved cells
Cryopreserved cells were thawed rapidly by incubating in a 37ºC water bath for 2
minutes. Then, cells were transferred to 10ml of pre-warmed full growth medium and
centrifuged at 1500rpm for 3 minutes. The supernatant was discarded in order to remove DMSO
traces and cell pellets were re-suspended in 10ml full growth medium in a T-25 sterile cell
culture flask and cultured at 37°C in a humidified environment containing 5% CO2. After cell
growth had resumed, cells were transferred to a T-75 flask and used for experiments.
2.2.2 Gene expression analysis by real-time PCR (RT-PCR)
Gene expression analysis using SYBR greenTM was carried out in order to measure the
changes in expression of genes of interest. The RT-PCR method is characterised by its
sensitivity and specificity for the detection of the level of messenger RNA (mRNA) expression
(Marone, Mozzetti et al. 2001). This technique was performed in three stages. Firstly, mRNA
was extracted from cells of interest. Secondly, mRNA was reverse transcribed to
complementary deoxyribonucleic acid (cDNA). Thirdly, the resultant cDNA was employed as
a PCR amplification template using specific primers for the genes of the interest. During PCR
amplification, the products of the genes of the interest were measured in real-time by assessing
the fluorescence produced by SYBR green, when it binds to double-stranded DNA. The
intensity of fluorescence reflects the amount of the target gene in the sample. The relative
expression of the target genes was assessed by comparing their expression to the expression of
the housekeeping gene β-actin.
2.2.2.1 mRNA extraction
This procedure was applied in order to extract and purify mRNA that will be used in
synthesis of cDNA. mRNA was extracted using the RNeasy® Plus Mini Kit according to the
manufacturer's instructions. Cells were suspended in 5% FBS media at a concentration of 5×105
38
cell/ml, seeded in 6-well plates, and treated in triplicate with the half maximal inhibitory
concentrations (IC50) of HXR9, unless otherwise specified, with the equivalent concentrations
of CXR9, or with media only. After treatment for two hours, cells were transferred into
eppendorfs. Then eppendorfs were spun at 1000rpm for five minutes and supernatants were
removed. After that, cells were lysed using RLT plus lysis buffer and β-mercaptoethanol, which
inhibits RNase. Next, cell lysates were vortexed in order to lyse more cells and yield the
maximum amount of RNA. Then, cell lysates were passed through a genomic DNA eliminator
spin column to remove the genomic DNA. Ethanol was added to cell lysates to allow RNA
binding to the RNeasy® silica gel membrane spin column. Then, cell lysates were washed using
the provided buffer RW1 to remove biomolecules that were non-specifically bound to the
membrane, such as carbohydrates, proteins, fatty acids, the small RNAs, ribosomal RNA and
transferase RNA. After that, the silica membrane was washed twice by RPE buffer to remove
ethanol. Finally, mRNA was eluted by using 35μl RNase-free water. mRNA samples were
stored at -80ºC until needed.
2.2.2.2 Measuring mRNA concentration
mRNA concentration was quantified using the Nanodrop® spectrophotometer ND-
1000, which is based on the Beer Lambert law to measure RNA concentration of 1μl of a
sample in ng/μl by measuring the ultra violet light absorbance at 260nm. RNA sample purity
was assessed using the 260/280 ratio, which should be more than two.
2.2.2.3 Reverse transcription of mRNA into cDNA
mRNA was reverse transcribed to cDNA using the Cloned AMV First-Strand cDNA
Synthesis Kit, which initiates the reverse transcription to cDNA of all mRNA molecules present
in a sample using the random primers method. mRNA samples were thawed on ice and 15μl
containing 100ng RNA was prepared in 1.5ml eppendorfs using DEPC-treated water. An
adequate volume of cDNA synthesis mix was prepared according to the manufacturer's
39
instructions and 5μl was added to each RNA sample (Table 2.3). The mixture was incubated at
50°C for 30 minutes to synthesise cDNA followed by heating at 85°C for 5 minutes to terminate
the reaction. Assuming cDNA synthesis efficiency was 100%, this produced almost 100ng of
cDNA in 20μl at a concentration of 5ng/μl, which was further diluted to 2ng/μl by adding 30μl
of DEPC-treated H2O and stored at -20°C.
Table 2.3 cDNA synthesis mix. This table shows the components of the cDNA synthesis mix and volumes used
to prepare a 5μl of the solution.
cDNA synthesis mix components Volume per reaction (μl)
5× Buffer 2
10mM dNTP Mix 1
0.1M DTT 0.5
Oligo (dT)20 (50μM) 0.5
RNaseOUTTM (40U/μl) 0.5
Cloned AMV RT (15U/μl) 0.5
Total volume 5
2.2.2.4 Complementary PCR primers design
Sequences of complementary primers for PCR amplification were designed using
(http://simgene.com/Primer3). Tables 2.4 and 2.5 show the complementary primer sequences
for the whole set of HOX genes and pro- and anti-apoptotic genes, respectively, that were
assessed in this study. All used primers were stored at -20ºC and thawed on ice before use.
40
Table 2.4 HOX gene primers used for PCR amplification.
Gene Forward Sequence 5’-3’ Reverse Sequence 5’-3’ Amplicon Length
(bp)
HOXA1 CTGGCCCTGGCTACGTATAA TCCAACTTTCCCTGTTTTGG 153
HOXA2 TTCAGCAAAATGCCCTCTCT TAGGCCAGCTCCACAGTTCT 176
HOXA3 ACCTGTGATAGTGGGCTTGG ATACAGCCATTCCAGCAACC 227
HOXA4 CCCTGGATGAAGAAGATCCA AATTGGAGGATCGCATCTTG 271
HOXA5 CCGGAGAATGAAGTGGAAAA ACGAGAACAGGGCTTCTTCA 193
HOXA6 AAAGCACTCCATGACGAAGG TCCTTCTCCAGCTCCAGTGT 158
HOXA7 TGGTGTAAATCTGGGGGTGT TCTGATAAAGGGGGCTGTTG 285
HOXA9 AATAACCCAGCAGCCAACTG ATTTTCATCCTGCGGTTCTG 203
HOXA10 ACACTGGAGCTGGAGAAGGA GATCCGGTTTTCTCGATTCA 159
HOXA11 CGCTGCCCCTATACCAAGTA GTCAAGGGCAAAATCTGCAT 279
HOXA13 GGATATCAGCCACGACGAAT ATTATCTGGGCAAAGCAACG 176
HOXB1 TTCAGCAGAACTCCGGCTAT CCTCCGTCTCCTTCTGATTG 157
HOXB2 CTCCCAAAATCGCTCCATTA GAAAGGAGGAGGAGGAGGAA 259
HOXB3 TATGGCCTCAACCACCTTTC AAGCCTGGGTACCACCTTCT 299
HOXB4 TCTTGGAGCTGGAGAAGGAA GTTGGGCAACTTGTGGTCTT 155
HOXB5 AAGGCCTGGTCTGGGAGTAT GCATCCACTCGCTCACTACA 189
HOXB6 ATTTCCTTCTGGCCCTCACT GGAAGGTGGAGTTCACGAAA 184
HOXB7 CAGCCTCAAGTTCGGTTTTC CGGAGAGGTTCTGCTCAAAG 249
HOXB8 GTAGGCTTCAGCTGGGACTG GGGAGCCTTTGCTTAAATCC 265
HOXB9 TAATCAAAGACCCGGCTACG CTACGGTCCCTGGTGAGGTA 198
41
Gene
Forward Sequence 5’-3’
Reverse Sequence 5’-3’
Amplicon Length
(bp)
HOXB13 CTTGGATGGAGCCAAGGATA CCGCCTCCAAAGTAACCATA 234
HOXC4 CGCTCGAGGACAGCCTATAC GCTCTGGGAGTGGTCTTCAG 276
HOXC5 CAGTTACACGCGCTACCAGA AGAGAGGAAAGGCGAAAAGG 268
HOXC6 AAGAGGAAAAGCGGGAAGAG GGTCCACGTTTGACTCCCTA 190
HOXC8 CTCAGGCTACCAGCAGAACC TTGGCGGAGGATTTACAGTC 150
HOXC9 AGACGCTGGAACTGGAGAAG AGGCTGGGTAGGGTTTAGGA 190
HOXC10 CGCCTGGAGATTAGCAAGAC GGTCCCTTGGAAGGAGAGTC 289
HOXC11 CGGAACAGCTACTCCTCCTG CAGGACGCTGTTCTTGTTGA 186
HOXC12 CAAGCCCTATTCGAAGTTGC GCTTGCTCCCTCAACAGAAG 180
HOXC13 GTGGAAATCCAAGGAGGACA TTGTTGAGGGACCCACTCTC 170
HOXD1 TTCAGCACCAAGCAACTGAC TAGTGGGGGTTGTTCCAGAG 232
HOXD3 CAGCCTCCTGGTCTGAACTC ATCCAGGGGAAGATCTGCTT 176
HOXD4 TCAAATGTGCCATAGCAAGC TCCATAGGGCCCTCCTACTT 173
HOXD8 TCAAATGTTTCCGTGGATGA GCTCTTGGGCTTCCTTTTTC 290
HOXD9 TCCCCCATGTTTCTGAAAAG GGGCTCCTCTAAGCCTCACT 236
HOXD10 GCTCCTTCACCACCAACATT AAATATCCAGGGACGGGAAC 154
HOXD11 GGGGCTACGCTCCCTACTAC GCTGCCTCGTAGAACTGGTC 253
HOXD12 CGCTTCCCCCTATCTCCTAC CTTCGGGCGCATAGAACTTA 201
HOXD13 GGGGATGTGGCTCTAAATCA AACCTGGACCACATCAGGAG 265
42
Table 2.5 Pro- and anti-apoptotic and β-actin gene primers used for PCR amplification.
Gene Forward Sequence 5’-3’
Reverse Sequence 5’-3’
Amplicon Length
(bp)
Bad CGGAGGATGAGTGACGAGTT GATGTGGAGCGAAGGTCACT 180
Bak1 TTTTCCGCAGCTACGTTTTT GGTGGCAATCTTGGTGAAGT 248
Bax TTTGCTTCAGGGTTTCATCC CAGTTGAAGTTGCCGTCAGE 246
Bcl2 GAGGATTGTGGCCTTCTTTG ACAGTTCCACAAAGGCATCC 170
Bid CTGCAGGCCTACCCTAGAGA ACTGTCCGTTCAGTCCATCC 195
XIAP GGGGTTCAGTTTCAAGGACA CGCCTTAGCTGCTCTTCAGT 182
Apaf1 TTCTGATGCTTCGCAAACAC CTGGCAAATCTGCCTTCTTC 237
Caspase-3 TTTTTCAGAGGGGATCGTTG CGGCCTCCACTGGTATTTTA 151
Caspase-6 ATCCTCACCGGGAAACTGTG AATTGCACTTGGGTCTTTGC 161
Caspase-7 AGTGACAGGTATGGGCGTTC CGGCATTTGTATGGTCCTCT 164
Caspase-9 CTAGTTTGCCCACACCCAGT GCATTAGCGACCCTAAGCAG 172
PARP1 GCTCCTGAACAATGCAGACA CATTGTGTGTGGTTGCATGA 233
c-FOS CCAACCTGCTGAAGGAGAAG GCTGCTGATGCTCTTGACAG 232
p21 GACACCACTGGAGGGTGACT
CAGGTCCACATGGTCTTCCT 171
p53 GTGGAAGGAAATTTGCGTG CCAGTGTGATGATGGTGAGG
183
β-actin ATGTACCCTGGCATTGCCGAC GACTCGTCATACTCCTGCTTG 227
2.2.2.5 RT-PCR
Each gene of interest was tested in triplicate. The housekeeping gene β-actin was used
as an endogenous control. An adequate volume of PCR mastermix was prepared in an
eppendorf as detailed in table 2.6. This mastermix contains cDNA, SYBR® Green JumpStartTM
Taq ReadyMixTM, ROX reference dye and DEPC-treated H2O. 20μl of the PCR mastermix was
added into three wells of a Mx 3000P® 96-well plate for each gene. 5μl of 1μM of each primer
pair was added into the designated wells. Plates were then sealed with optical caps and
centrifuged at 1500rpm for 1 minute to bring all reagents to the well bottoms and remove air
43
bubbles. The Plates were then run on a Stratagene Mx3005P thermal cycler. The thermal
profiles employed were: heating once for 10 minutes at 95°C, followed by 30 seconds at 95°C,
1 minute at 60°C, 30 seconds at 72°C for 40 cycles. After completing the thermal profiles, the
detected fluorescence of each reaction was converted to cycle threshold using MxPro software.
The mean of the cycle threshold values of the triplicate repeats for each gene was calculated
and used for measuring gene expression, as detailed below.
Table 2.6 RT-PCR reaction components. This table shows the components of single RT-PCR reaction. All
components were stored at -20°C and thawed on ice before use.
PCR reaction components Volume per reaction (μl)
cDNA (2ng/μl) 1
DEPC-treated H2O 6.25
SYBR® Green JumpStartTM Taq ReadyMixTM 12.5
ROX reference dye 0.25
Primers-forward and reverse mixture 5
Total volume 25
2.2.2.6 RT-PCR data analysis
A previously described relative quantification method was used to analyse RT-PCR data
(Livak and Schmittgen 2001). Two steps were performed in this analysis. Firstly, the expression
of genes of interest was normalised to the expression of the housekeeping gene β-actin.
Secondly, the normalised data of genes of interest of HXR9 or CXR9 treated samples were
compared to that of untreated control samples. Microsoft Excel was used to perform all
calculations. Experiments were repeated three times. Results were expressed as the mean of the
relative gene expression of the three experiments with error bars to show the standard error of
mean (SEM).
44
2.2.3 Lactate dehydrogenase (LDH) assay
LDH is present in the cytoplasm of all cells. It is released into culture medium if the cell
membrane is permeablized. Therefore, the amount of this enzyme in the culture medium
indicates to the integrity of cell membrane and hence provides an estimation of cytotoxicity
(Haslam, Wyatt et al. 2000). LDH activity is assessed in two steps. Firstly, LDH oxidises lactate
to pyruvate by reducing NAD+ to NADH/H+. Secondly, diaphorase, a catalyst, transfers 2H
from NADH/H+ to a tetrazolium salt (INT) which results in the formation of the red salt
formazan. Hence, the amount of formazan formed directly correlates to the amount of LDH in
the culture medium and thereby to the number of damaged cells (Figure 2.2).
Figure 2.2 LDH cytotoxicity assay. LDH catalyses the reduction of NAD⁺ to NADH⁺/H⁺ through the conversion
of lactate to pyruvate. Then, diaphorase reduces the yellow tetrazolium salt INT to the red formazan salt in the
presence of NADH⁺/H⁺.
The LDH assay was used to measure the cytotoxicity of HXR9, daunorubicin (DNR)
and mitoxantrone (MTX). In HXR9 cytotoxicity determination, cells were treated with HXR9
or its negative control CXR9 for 2 hours. In DNR or MTX cytotoxicity determination, cells
were treated with either drug for 24 hours as per the following:
Cells were suspended in 5% FBS of the appropriate culture medium at a concentration of 5×105
cell/ml and seeded in 96-well flat-bottom plates at a concentration of 5×104 cell/well. Then cells
were treated with 100μl of a serial dilution of the tested drug, which have been already diluted
45
in 5% FBS of the appropriate culture medium. Each plate included untreated cells (negative
control), 2% triton-treated cells (positive control), and 5% FBS medium only (background
control). Each condition was repeated six times. The plates were incubated in a humidified
incubator at 37ºC, 5% CO2 for either 2 or 24 hours according to the tested drug. After treatment,
the plates were spun at 1000rpm for 5 minutes and 100μl of supernatant was decanted from
each well into a fresh, 96-well flat-bottom plate. An equal amount of LDH reagent was prepared
according the manufacturer's instructions and added to the supernatants for two minutes, and
then LDH enzymatic activity was estimated by reading the absorbance at 492nm, using a plate
reader.
2.2.3.1 LDH assay data analysis
In order to assess the percentage of surviving cells, the cytotoxicity percentage was
calculated first. The average absorbance values of the six repeats of each condition were
calculated. Then, the average absorbance value of the background control was subtracted from
each of the other conditions average absorbance values. After that, the cytotoxicity percentage
was calculated according to the manufacturer's instructions as per the following:
Cytotoxicity% = treated value−negative control
positive control−negative control × 100
Then, the proportion of surviving cells was determined as the following:
%surviving cells = 100 - cytotoxicity
The %surviving cells was plotted on X-Y scatter diagrams using GraphPad Prism
software (California, USA) in order to create cytotoxicity curves for each cell line and drug.
Therefore, the surviving fractions of treated cells can be compared to those of untreated cells.
Additionally, the cytotoxicity results of the different cell lines can be compared.
46
Each experiment was repeated at least three times (biological repeats) and each valid
experimental curve should have a linear correlation coefficient (R2) ≥ 0.95. The cytotoxicity
curves were generated using the mean %surviving cells at different drug concentrations.
Additionally, the SEM of each single dose was determined and included as an error bar on the
cytotoxicity curves.
IC50 was then calculated by CalcuSyn software (Biosoft, Cambridge, UK) as described
in statistical analysis.
2.2.4 Assessment of drug combination interaction by LDH
This assay was performed in vitro to evaluate the interaction levels (synergism, additive,
or antagonism) between HXR9 and the chemotherapeutic drugs DNR or MTX.
Cells were suspended in 5% FBS medium and seeded in 96-well flat-bottom plates at a
concentration of 5×104 cell/well. Then, cells were treated with 100μl of five increasing
concentrations (IC12.5, IC25, IC50, IC75, IC100) of each drug alone or in combination with HXR9.
For single treatments, cells were treated with HXR9 or with a chemotherapeutic drug, DNR or
MTX, for 2 or 24 hours, respectively. For combination treatment, cells were treated with either
DNR or MTX for 24 hours and HXR9 was added for the last 2 hours of treatment. Each
treatment was repeated in triplicate. Each plate included negative, positive and background
controls. After treatment, plates were spun for 5 minutes at 1000rpm, 100μl of supernatant was
decanted into new 96-well flat-bottom plates and 100μl LDH reagent was added. The
cytotoxicity was then determined as described for the LDH assay data analysis section and the
evaluation of the drug interaction between HXR9 and the chemotherapeutic drugs was
performed as described for the analysis of drug combination assay data section. The
combination assay data were plotted on column graphs using GraphPad Prism software.
47
2.2.5 Annexin V- PE assay
The annexin V-PE assay was used to quantify the percentage of the apoptotic cells and
evaluate the mechanism of cell death. This assay is based on the cell membrane permeability
of the apoptotic cells. During early stages of apoptosis, the membrane bound phospholipid
phosphatidlyserine (PS) translocates from the inner surface of the cellular plasma membrane to
the external surface, where it can be detected by annexin V protein that has a high phospholipid-
binding affinity. Annexin V is conjugated to PE (a fluorochrome) and this compound is used as
a probe for cell analysis by flow cytometry (Vermes, Haanen et al. 1995). Therefore,
quantification of exposed PS using PE labelled annexin V by flow cytometry is considered a
sensitive measurement of apoptosis (Koopman, Reutelingsperger et al. 1994). In addition, 7-
amino actinomycin D (7-AAD), which is a membrane impermeable fluorescent DNA binding
stain that selectively interacts with guanine and cytosine bases in double-stranded DNA, was
used to distinguish between viable and non-viable cells (Philpott, Turner et al. 1996). Viable
cells stain negative with both reagents. Early apoptotic cells stain positive for annexin V-PE
only. Late apoptotic, necrotic, or dead cells stain positive for both stains, as shown in table 2.7.
The late apoptotic/ necrotic cells will be reflected to in this study as dead cells, because this
assay cannot definitely differentiate between these two conditions (Figure 2.3).
Cells were suspended in 5% FBS of the appropriate culture medium at a concentration
of 1×106 cell/ml and seeded in 96-well flat-bottom plates at a concentration of 1×105 cell/well.
Then cells were treated either with 100μl of a serial HXR9 dilution or with a high CXR9
concentration in triplicate. Each plate included untreated cells (negative control). The plates
were incubated in a humidified incubator at 37ºC, 5% CO2 for two hours. After two-hour
treatment, the plates were spun at 1000rpm for five minutes and supernatants were removed.
Cell pellets were washed twice with cold phosphate buffered saline (PBS) and then cells re-
suspended in 20μL of 1X binding buffer (0.1M Heps/NaOH (pH 7.4), 1.4M NaCl, 25mM
48
CaCl2). Cell suspensions were transferred to culture tubes. 5μL of both annexin V-PE and 7-
AAD were added and then cell suspensions were incubated at the room temperature (RT) for
15 minutes in the dark. Then, 200μL of 1X binding buffer were added to tubes and analyzed by
a flow cytometry.
The percentage of the different cell populations, viable, early apoptotic and dead cells,
were plotted by GraphPad Prism software (California, USA) on column graphs to allow the
comparison of treated and untreated negative control cells. Each experiment was repeated at
least three times independently.
Table 2.7 Different cell populations in the annexin V-PE assay.
Annexin V 7-AAD Cell Stage
Negative Negative Viable
Positive Negative Early apoptosis
Positive Positive Late apoptosis or necrosis (dead
cells)
Figure 2.3 A flow cytometry plot shows different cell populations in the annexin V-PE assay. In the left bottom
corner are the viable cells that are negative for annexin V and 7-AAD. In the right bottom corner are the early
apoptotic cells that are stained with annexin V only. In the top part are dead cells that are stained with both annexin
V and 7-AAD.
49
2.2.6 Caspase-3 activity assay
Activation of caspases is a key event in apoptosis. In particular, caspases-3/7 are
considered as executioner caspases that recognise and cleave numerous proteins containing the
DEVD sequence such as poly ADP ribose polymerase (PARP), resulting in cell apoptosis. The
caspase-3 assay is based on the ability of active caspase-3 to cleave a synthetic DEVD sequence.
DEVD is covalently linked to R110 in a compound called Z-DEVD-R110, which suppresses
the fluorescence R110. The caspase cleavage of DEVD results in R110 fluorescence, which is
measured at excitation wavelength 492nm and emission wavelength 520nm. Ac-DEVD-CHO
is a caspase-3/7 inhibitor that can be used to verify the resultant fluorescence is due to caspase-
3 activity. A reference standard can be used to quantify the R110 produced in the reaction.
Cells were suspended in the appropriate culture medium plus 5% FBS at a concentration
of 2×106 cell/ml and seeded in eppendorfs at a concentration of 1×106 cell/reaction. Cells were
treated, in triplicate, with 500μl of different HXR9 concentrations or with 500μl of an
equivalent CXR9 concentration to the highest HXR9 concentration. Each experiment included
untreated cells (negative control). The plates were then incubated in a humidified incubator at
37ºC, 5% CO2 for two hours. After the treatment for two hours, eppendorfs were spun at
1000rpm for 5 minutes and supernatants were removed. Cell pellets were then washed with
PBS and re-suspended in 100μl of 1X cell lysis buffer and incubated on ice for 30 minutes. The
lysed cells were then spun at 5000rpm for 5 minutes and 50μl of the supernatant was transferred
to a new 96-well flat bottom black plate. 50μl of the lysis buffer was added into separate wells
as non-enzyme controls in order to measure the substrate background fluorescence. As an
additional control, 1μl of 1mM Ac-DEVD-CHO inhibitor was added to selected samples that
were incubated at RT for 10 minutes in the dark. During this time, the remaining samples
(without inhibitors) were kept on ice. 50μl of 500μM Z-DEVD-R110 substrate was then added
50
to all samples. The supernatants were incubated with the substrate for 30 minutes in the dark.
Serial concentrations of the reference standard were added to the plates in order to quantify
R110 fluorescence produced in the reaction. Finally the caspase-3 activity was measured by
reading the plates at 492nm by the plate reader.
2.2.6.1 Caspase-3 activity data analysis
For each individual experiment, the average fluorescent value for the different
conditions was calculated, and the lysis buffer reading (background) was subtracted from the
other average values. Then, the resultant values were plotted on column graphs using GraphPad
Prism software to compare caspase-3 activity and calculate the statistical significance between
untreated control cells and cells treated either with HXR9 or CXR9.
In order to validate each experiment, the reference standard data were plotted separately
on X-Y scatter graphs and each valid reference standard curve should have R2 ≥ 0.95.
Each experiment was performed at least in triplicate. The caspase-3 activity data were
generated by the mean of the fluorescent values of the different cell conditions. The SEM of
each single dose was determined and included on the graphs.
2.2.7 Cell cytospins and 4',6-diamidino-2-phenylindole (DAPI) staining
Cytospin is a method used to concentrate cells. After cell cytospinning, cell morphology
can be evaluated by staining. DAPI is a cell-permeable blue fluorescent stain that binds
specifically to the minor groove adenine-thymine regions of double-stranded DNA. The
binding of DAPI with DNA intensifies the fluorescence about 20 fold and this can be imaged
using a fluorescence microscope or flow cytometry. DAPI stain is used to assess apoptosis by
visualising the nuclear morphology changes such as chromosome condensation and
fragmentation (Kapuscinski 1995).
51
Cells were suspended in 5% FBS culture medium at a concentration of 1×106 cell/ml
and seeded in 24-well plates at a concentration of 5×105 cell/well. Then cells were treated either
with 500μl of HXR9 the IC50 or double the IC50, in duplicate. Each plate included untreated
negative control cells. The plates were incubated in a humidified incubator at 37ºC, 5% CO2
for two hours, after which cells were transferred into eppendorfs. Harvested cells were washed
twice with PBS, and fixed in 4% formaldehyde/PBS solution.
Cytospinning was then performed. Microscope-slides, filter cards and cytofunnels were
assembled using cytospin holders and then the entire apparatus was placed in a cyto-centrifuge.
Then, 300μl of 4% formaldehyde/PBS fixed cells was added to the cytofunnels and spun at
500rpm for 5 minutes. Next, the apparatus was dis-assembled. Slides were stained with 1 drop
of 1:1000 DAPI/PBS for 5 minutes in the dark. After staining, excess DAPI/PBS was removed
and 1 drop of the mounting media 40% glycerol/PBS added. Then, slides were covered with
slip covers and sealed using nail vanish. Finally, images were captured with 20x lenses by a
fluorescent microscope.
2.2.8 Inhibition of caspases activity by z-VAD-FMK
Z-VAD-FMK is a cell permeable general caspase inhibitor. It can prevent the induction
of apoptosis through irreversible binding to caspase protease catalytic sites (Marcelli,
Cunningham et al. 1999). Pre- and co-treatment of cells with z-VAD-FMK assay was used to
investigate the role of caspases in cell death. Z-VAD-FMK was diluted in DMSO at 43mM and
stored at -20ºC.
Some wells were pre-treated with 50μM z-VAD-FMK. Cells were suspended in 5%
FBS culture media at a concentration of 1×106 cell/ml and seeded in 96-well flat-bottom plates
at a concentration of 1×105 cell/well. The plates were then pre-incubated in a humidified
incubator at 37ºC, 5% CO₂ for one hour. Meanwhile, HXR9 and CXR9 were prepared in 5%
52
FBS media either with or without 50μM z-VAD-FMK. After one hour pre-incubation, cells
were treated in triplicate with 100μl of HXR9 IC50, double the IC50, or with a CXR9
concentrations equivalent to double the IC50. Additionally, each plate included untreated
negative control cells and DNR treated cells as positive control cells. The plates were then
incubated in a humidified incubator at 37ºC, 5% CO2 for two hours. After treatment for two
hours, the plates were spun at 1000rpm for five minutes and supernatants were discarded. Cell
pellets were washed twice with cold PBS and then stained with annexinV-PE as described for
the annexinV-PE assay section. Data were plotted as described for the annexinV-PE assay.
Statistical significance was assessed by comparing the data of untreated cells to those of pre-
and co-treated with 50μM z-VAD-FMK.
2.2.9 Cyclosporin A (CsA) protection assay
CsA attenuates mitochondrial permeability transition pore (mPTP) formation by
binding to cyclophilin D (cypD), which is required for mPTP formation in necrotic
mitochondrial cell death (Li, Johnson et al. 2004; Basso, Fante et al. 2005; Dube, Selwood et
al. 2012). CsA was diluted in DMSO to a stock concentration of 20mM, aliquoted and stored
at -20ºC.
In this assay cells were pre- and co-treated with or without 5μM CsA and hydrogen
peroxide (H2O2) treatment was used as a positive control. Cells were suspended in 5% FBS
culture medium at a concentration of 1×106 cell/ml and seeded in 96-well flat-bottom plates at
a concentration of 1×105 cell/well. The plates were then pre-incubated in a humidified incubator
at 37ºC, 5% CO2 for one hour. Meanwhile, HXR9 and H2O2 were prepared in 5% FBS medium
either with or without 5μM CsA. After one hour pre-incubation, cells were treated in triplicate
with 100μl of HXR9 at the IC50, double or triple the IC50 or with 20mM H2O2. Additionally,
each plate included untreated negative control cells. The plates were then incubated in a
53
humidified incubator at 37ºC, 5% CO2 for two hours. After treatment for two hours, the plates
were spun at 1000rpm for 5 minutes and supernatants were discarded. Cell pellets were washed
twice with cold PBS and then stained with annexinV-PE as described for the annexinV-PE
assay. Data analysis was performed by flow cytometry.
The percentage of viable cells was plotted using GraphPad Prism software on column
graphs. Each experiment was repeated at least three times independently.
2.2.10 Necrostatin-1 (Nec-1) protection assay
Nec-1 is a selective inhibitor of receptor interacting protein 1 (RIP1) (Degterev, Hitomi
et al. 2008). RIP1 inhibition prevents a form of programmed necrosis called necroptosis
(Degterev, Huang et al. 2005). Nec-1 was diluted in DMSO to a stock concentration 40mM and
stored at -20ºC. In this experiment, some cells were pre- and co-treated with 50μM Nec-1. The
experiment and data analysis was performed as described for the CsA protection assay.
2.2.11 Fructose protection assay
Fructose is a glycolytic substrate that at high concentrations significantly sequesters
phosphate and accordingly intracellular adenosine triphosphate (ATP) is depleted (Figure 2.4).
This selective depletion of ATP protects cells from apoptosis, but leaves enough levels (15%)
of ATP to rescue cells from spontaneous necrosis (Nieminen, Saylor et al. 1994).
Figure 2.4 ATP depletion by fructose.
54
Cells were pre- and co-treated with a high concentration of fructose (40mM). Fructose
was freshly dissolved in 5% FBS culture medium. Cells were suspended in 5% FBS culture
medium supplemented with or without 40mM fructose and seeded in 96-well flat-bottom plates
at a concentration of 5×104 cell/well. The plates were then incubated at 37ºC, 5% CO2 for one
hour. Serial HXR9 concentrations were prepared in 5% FBS media either with or without
40mM fructose. After an hour pre-incubation, cells were treated in triplicate with 100μl of the
different HXR9 concentrations or 2% triton as a positive control. Additionally, each plate
contained cells incubated with 5% FBS medium as a low control and background wells
contained 5% FBS medium only. After treatment for two hours, the plates were spun at
1000rpm for 5 minutes and 100μl of each supernatant was decanted into fresh, 96-well flat-
bottom plates. Finally, LDH reagents were added as described in the LDH assay section.
Similarly, data analysis was performed as for the LDH assay. The statistical significance of the
effect of fructose on the cytotoxicity of HXR9 was performed by comparing the survival
fractions of fructose treated and untreated cells.
2.2.12 The effect of ethilenediaminetetra-acetic acid (EDTA) on HXR9 cytotoxicity
EDTA is a calcium ions (Ca2+) chelator that was used to investigate the role of
extracellular Ca2+ in the cytotoxicity of HXR9. The role of Ca2+ was assessed by two different
assays, LDH and annexin V, as described for the fructose protection and CsA protection assays,
respectively. In both assays, cells were pre- and co-treated with or without 20mM EDTA
dissolved in 5% FBS medium. The statistical significance of the effect of EDTA on the
cytotoxicity of HXR9 assessed by comparing the survival fraction of EDTA treated cells to
untreated cells.
55
2.2.13 The effect of HXR9 on mitogen activated protein kinase (MAPK) pathways
MAPKs are a family of protein kinases regulating many essential cellular pathways
including cell survival, proliferation and apoptosis. The most extensive studied MAPK
pathways are extracellular signal-regulated kinase (ERK), Jun N-terminal kinase (JNK) and
p38 (Dhillon, Hagan et al. 2007). The effect of HXR9 on those pathways was studied by
blocking each pathway using a specific inhibitor. The inhibitors used were U0126
monoethanolate, SP600125 and SB203580 for ERK, JNK and p38, respectively. The inhibitor-
stocks were dissolved in DMSO at concentrations 1mM, 25mM and 5mM of U0126
monoethanolate, SP600125 and SB203580, respectively. U0126 monoethanolate and
SP600125 were stored at 4ºC while SB203580 stored at -20ºC until needed.
In all of the assays that investigate the effect of HXR9 on ERK, JNK and p38 pathways,
cells were pre- and co-treated with or without the inhibitors. The working concentrations used
were 20μM, 60μM and 25μM for U0126 monoethanolate, SP600125 and SB203580,
respectively. Experiments were performed as described for the fructose protection assay;
however, data were plotted in column diagrams. The statistical significance of the effect of
blocking MAPK pathways on the cytotoxicity of HXR9 was determined by comparing the
survival fraction of the different-inhibitors-treated and untreated cells.
2.2.14 The effect of inhibition of NADPH oxidase (NOX) on HXR9 efficacy
NOX is a generator of intracellular reactive oxygen species (ROS) (Lambeth 2004). The
effects of NOX on the efficiency of HXR9 were assessed using an inhibitor called
diphenyleneiodonium chloride (DPI). DPI is widely used to block the effect of NOX in spite of
its non-specific binding (Park, Song et al. 2007). DPI stock was dissolved in DMSO at 15mM
and stored at -20ºC until used.
56
Cells were pre- and co-treated with or without 60μM DPI. The assessment of DPI effect
on the cytotxicity of HXR9 was performed as described for the fructose protection assay and
data were plotted in column diagrams. The statistical significance was assessed by comparing
the survival fraction of DPI treated and untreated cells.
2.2.15 Role of μ-calpain in HXR9 cell killing
Calpains are Ca2+-dependent cysteine proteases that commonly activated in Ca2+
involved cell death mechanisms (Liu, Van Vleet et al. 2004). Among them, μ-calpain is one of
the most extensively studied in apoptosis/necrosis processes. Calpain inhibitor I (N-Acetyl-
Leu-Leu-Norleu-al) was used to inhibit the Ca2+/μ-calpain pathway. Calpain inhibitor I was
dissolved in DMSO at a concentration 20mM and stored at -20ºC until used.
Cells were pre- and co-treated with or without 60μM calpain inhibitor I. The assessment
of the role of μ-calpain in the cytotxicity of HXR9 was performed as described for the fructose
protection assay and data were plotted in column diagrams. The statistical significance was
assessed by comparing the survival fractions of calpain inhibitor I treated cells to untreated
cells.
2.2.16 The effect of calmodulin (CaM) inhibition on HXR9 cytotoxicity
CaM is one of the main intracellular regulators of Ca2+ (Yuan, Jing et al. 2011). W-7
(N-(6-Aminohexyl)-5-chloro-1-naphthalenesulfonamide hydrochloride) was used to inhibit
CaM and assess its role in HXR9 cytotoxicity. W-7 was dissolved in DMSO at a concentration
50mM and stored at -20ºC.
57
Cells were pre- and co-treated with or without 50μM W-7. The assessment of the effect
of the inhibition of CaM on the cytotoxicity of HXR9 was performed as described for the
fructose protection assay and data were plotted in column diagrams. The statistical significance
was determined by comparing the survival fractions of W-7 treated cells to untreated cells.
2.2.17 The effect of protein kinase C (PKC) inhibition on HXR9 cytotoxicity
PKC is one of the kinases that are affected by intracellular Ca2+ concentrations. Ro31-
8220 (methanesulfonate salt) was used to inhibit and assess the role of PKC in HXR9
cytotoxicity. Ro31-8220 was dissolved in H2O at a stock concentration 20mM and stored at
4ºC. Cells were pre- and co-treated with or without 30μM Ro31-8220. The assessment of the
effect of the inhibition of PKC on the cytotxicity of HXR9 was done as performed for the
fructose protection assay and data were plotted in column diagrams. The statistical significance
was assessed by comparing the survival fractions of Ro31-8220 treated cells to untreated cells.
2.2.18 The effect of the heme oxygenase-1 (HO-1) inhibition on HXR9 cytotoxicity
HO-1 is an anti-apoptotic protein (Rushworth and MacEwan 2008). Protoporphyrin IX
(PPIX) was used to inhibit HO-1 protein and assess its role in the cytotoxicity of HXR9. PPIX
was dissolved in DMSO at a stock concentration 18mM and stored at -20ºC. Cells were pre-
and co-treated with or without 50μM of PPIX. The effect of HO-1 inhibition on HXR9
cytotoxicity was determined as described for the fructose protection assay and data were plotted
in column diagrams. The statistical significance was performed by comparing the survival
fraction of PPIX treated cells and untreated cells.
58
2.2.19 The effect of the p53 inhibition on HXR9 cytotoxicity
Pifithrin-α (PFT-α) is an inhibitor of p53-dependent apoptosis and was used to block
and assess the role of p53 in the cytotoxicity of HXR9. PFT-α was dissolved in DMSO at a
stock concentration 20mM and stored at -20ºC. Cells were pre- and co-treated with or without
40μM of PFT-α. The role of p53 in HXR9 cytotoxicity was assessed as described for the
fructose protection assay and data were plotted in column diagrams. The statistical significance
was calculated by comparing the survival fraction of PFT-α treated cells and untreated cells.
2.2.20 Western blotting (WB) for protein expression
WB was conducted to assess the difference in protein expression between untreated
cells and HXR9 treated cells. This assay requires treatment of cell lines with HXR9, extraction
of cytosolic proteins, proteins separation based on their sizes and blotting on a membrane.
Finally, the proteins of interest are detected by labelled antibodies. In order to compare the
expression of protein in different samples, the same amount of protein was loaded in each well
and the same dilution of antibodies was used.
Cells were suspended in 5% FBS medium at a concentration of 1.5×106 cell/ml and
seeded in 24-well plates. Then cells were treated with 1ml of 15μM or 30μM HXR9. Each plate
included untreated cells as a negative control. Each condition was repeated three times. The
plates were incubated in a humidified incubator at 37ºC, 5% CO₂ for two hours. After treatment
for two hours, the three repeats of each condition were gathered in one 25ml universal tube and
centrifuged for 5 minutes at 1500rpm at 4ºC. Supernatants were decanted and cell pellets were
washed twice with 5ml ice cold PBS. Cell pellets were collected in eppendorfs and lysed as
described for the preparation of cell lysate.
59
2.2.20.1 Preparation of cell lysate
Cell pellets containing 4.5×106 cells were suspended in 300μl RIPA solution with 10μl
of inhibitor cocktail (protease and phosphatase inhibitor) and 10μl of 1mM EDTA. Cell
suspensions were incubated on ice for 15 minutes and agitated at least twice. Cell membranes
were sheared by passing cells through 21 gauge needles attached to 1ml syringes for at least 4-
5 times. Cell suspensions were centrifuged at 13000rpm for 5 minutes at 4ºC and supernatants
were transferred to new eppendorfs. The protein concentration in cell lystaes was determined
using a bicinchonic acid (BCA) assay.
2.2.20.2 Measuring protein concentrations by pierce Bradford city assay (BCA)
This assay was used to measure protein concentrations in cell lysates. It is based on
reduction of copper Cu2+ to cuprous ion Cu+ by proteins. Cu+ then chelates with two molecules
of BCA to form a purple-coloured product that can be detected by a 96-well plate reader at
562nm. The intensity of the formed colour increases linearly with increasing protein
concentrations allowing protein concentrations to be calculated using a standard curve made of
bovine serum albumin (BSA). This assay was performed as follows:
25μl of samples were added in duplicate in 96-well flat-bottom plates. Additionally, each plate
contained a BSA series dilution added in duplicate. To each sample and standard well, 200μl
of BCA working reagent, a combination of reagents A and B in a ratio 50:1, was added. The
plates were gently agitated on a plate shaker and incubated for 30 minutes at 37ºC. After the
incubation, the plates were left at RT for 5 minutes in the dark before reading the absorbance
on a plate reader. The absorbance means for the duplicate repeats of each sample and standard
were calculated. The standard curve was then generated by the absorbance means of BSA series
dilutions and their corresponding concentrations using Microsoft Excel. Finally, the protein
concentrations of the samples were calculated using the standard curve linear regression
equation.
60
2.2.20.3 Sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE) for
protein separation
Cellular proteins were separated based on size using SDS-PAGE. First, 20μl samples
were prepared (13μl cell lysate, 5μl sample buffer and 2μl reducing agent) and heated at 70ºC
for 10 minutes. Second, a Novex® NuPAGE 4-12% Bis-Tris Gel, which is used to separate
proteins, was placed in an XCell SureLockTM Mini-Cell electrophoresis system that contained
800ml NuPAGE® MES SDS running buffer, which prepared by adding 20 parts of water to the
buffer. Third, 10μl of Novex® sharp pre-stained marker was loaded in the first well of every
gel. Forth, 20μl of samples were loaded in the remaining wells. Finally, protein separation was
performed by running gels at 200 volt for one hour.
2.2.20.4 Transferring proteins to polyvinylidene fluoride (PVDF) membranes
After electrophoretic separation, proteins were transferred to PVDF membranes. Firstly,
PVDF membranes were soaked in methanol for 2 minutes while filter papers were soaked in
water. Membranes were then placed in Invitrogen Electrophoresis System and covered with the
gel and then the filter papers and on top of that transfer sponges were placed. Finally, proteins
were transferred by running at 30V for 7 minutes.
2.2.20.5 Detection of proteins by antibodies
After transferring proteins, PVDF membranes were blocked in 5% blocking buffer (5%
(w/v) dried non-fat milk powder, 0.1% (v/v) Tween-20) with gentle rocking for 2 hours at RT.
Then, the membranes were incubated with primary antibodies with gentle rocking overnight at
4ºC. After overnight incubation, the membranes were washed 4 times with 0.1% Tween PBS
solution for 10 minutes at RT with gentle rocking to remove unbound antibodies. Then, the
membranes were incubated with secondary antibodies for 2 hours at RT with gentle rocking;
the antibodies used in WB are listed in table 2.8. After 2 hours incubation, membranes were
washed 3 times, as above. Then, membranes were exposed to superSignal® West Pico
61
Chemiluminescent substrate according to the manufacturer's recommendation. Next,
membranes were exposed to carestream® Kodak BioMax® light film for 3 minutes to visualise
the chemiluminescence. After that, films were developed and then fixed using developer and
fixer, respectively.
Table 2.8 Antibodies used in WB and working dilutions.
Antibody Type Isotype Molecular
weight (KDa)
Working
concentration
Supplier
Primary anti-
PARP1
Monoclonal Mouse IgM 116 1/4000 Abcam
Primary anti-β-
actin
Polyclonal Rabbit IgG 42 1/10000
Secondary for
anti-PARP
Polyclonal Goat IgG 1/4000
Secondary for
anti-β-actin
Polyclonal Goat IgG 1/2000
62
2.2.21 In vivo assays
2.2.21.1 Cell preparations
Cells used for in vivo experiments were C1498-GFP and K562 cell lines. Cells were
spun for 3 minutes and re-suspended and washed twice in HANKS medium. C1498-GFP and
K562 cells were then suspended in serum-free HANKS medium at density 1x107cell/ml and
5x107 cell/ml, respectively.
2.2.21.2 Systemic injection of C1498-GFP in C57BL/6 and nude mice
C57BL/6 and nude mice were warmed by a heat lamp in order to dilate the vain. C1498-
GFP cells were then injected into the tail vain (1x106 cell/mouse) using 23 gauge needles. Mice
were then culled randomly during the experiment duration using CO2 to harvest peripheral
blood (PB) and organs including BM, liver, spleen and kidney to assess the engraftment of
AML.
2.2.21.2.1 Harvesting and processing PB
PB was harvested through cardiac puncture using heparinised needles. Blood were then
suspended in 10ml PBS and washed twice. Red blood cells were lysed by harvesting cells in
RBC lysis buffer for 10 minutes. Cells were then spun and supernatant was removed. Cells
were then re-suspended in 20μL of 1X binding buffer. AML engraftment in PB was assessed
by the presence of green fluorescent protein (GFP+) cells using flow cytometry.
2.2.21.2.2 Harvesting and processing BM and other organs
BM was harvested from femurs by flushing with HANKS medium; cells were spun and
re-suspended in HANKS medium. Cells were obtained from spleen, liver and kidney by
mincing and filtration through a cell strainer (70μM). AML engraftment was then assessed as
described for PB.
63
2.2.21.3 Subcutaneous (S.C.) injection of AML cells into C57BL/6 and SCID mice
C57BL/6 and SCID mice were anesthetised using isoflurane. C1498-GFP (1x106
cell/mouse) and K562 (5x106 cell/mouse) cells were then S.C. injected in the flank of C57BL/6
and SCID mice, respectively. Flank tumour sizes were measured three times a week using
calliper measurement. Tumour sizes were calculated using the formula: (length x width x
width)/2. When flank tumour became palpable, the mice were divided into two groups with one
introduced intratumourly (I.T.) with 50μl of the indicated doses of HXR9 or HXR9 + (500μM
W-7 + 300μM Ro31-8220) in the result section and the other group (control group) were
introduced with I.T. with 50μl of PBS. Mice were sacrificed when tumour size ≥ 1500mm3 or
when mice became sick.
2.3 Statistical analysis
Calcusyn software was used to determine the IC50 of the different drugs for the different
cell lines and for assessing whether the interaction between HXR9 with DNR or MTX was
synergistic, additive or antagonistic, as described below.
Student's t-test was applied for data analysis of all assays using GraphPad Prism
software. Results are expressed as the mean of three separate experiments with error bars to
show the SEM. Statistical significance was determined by comparing the different conditions
in each assay with each other and considered significant when p < 0.05 (*), highly significant
when p < 0.01 (*), or very highly significant when p < 0.001 (***) or p < 0.0001 (****).
64
2.3.1 Calculation of IC50
IC50 of HXR9, DNR and MTX for different cell lines were calculated by the median-
effect equation of Chou (Chou 1974) using the Clcusyn software. The equation is:
fa/fu = (D/Dm)m
Where D is the dose of the drug; Dm is the dose need to produce a 50% effect (IC50); fa is the
fraction affected by the dose; fu is the fraction unaffected by the dose, fu = 1-fa; m is the exponent
signifying the shape (sigmoidicity) of the dose effect curve.
2.3.2 The analysis of drug combination assay data
The combination effects of HXR9 with either DNR or MTX were calculated by a
combination index (CI) equation introduced by Chou-Talalay (Chou 1991), using Calcusyn
Software. The equation is
CI = (D)1
(Dx)1+
(D)2
(Dx)2 +
(D)1(D)2
(Dx)1(Dx)2
Where CI > 1, CI = 1, CI < 1 indicate antagonism, an additive effect and synergism,
respectively. X is the inhibition percentage so for example, 50%, (Dx)1 and (Dx)2 are the
concentrations of drug 1 and drug 2 alone, respectively, that cause 50% of cell death and can
be calculated by the median-effect equation; (D)1 and (D)2 are the concentrations of the drugs
in combination that also cause 50% of cell death.
2.3.3 Statistical analysis of in vivo assays
Data were expressed as the average tumour sizes for each group at the represented day
with SEM as error bars. The statistical significance in tumour sizes was determined using two-
way anova analysis.
65
Table 2.9 Summary of the in vitro assays performed and reagents used.
Assay Investigation reagent solvent Stock
concentration
Storing Working
concentration
End point
measurement
LDH HXR9, DNR
and MTX
cytotoxicity
LDH
detection
kit
-20ºC LDH activity
Combination
cytotoxicity
HXR9
interaction
with DNR or
MTX
LDH
detection
kit
-20ºC LDH activity
Annexin V Cell death
mechanism
Annexin V
and 7-
AAD
4ºC PS exposure
Caspase-3 Caspase-3
role
z-DEVD-
R110 and
Ac-
DEVD-
CHO
-20ºC LDH activity
DAPI
staining
Nuclear
fragmentation
DAPI
stain
PBS -20ºC 1:1000 Nuclear
fragmentation
General
caspase
activity
General
caspase
activity
z-VAD-
FMK
DMSO 43mM -20ºC 50μM PS exposure
CsA
protection
CypD role CsA DMSO 20mM -20ºC 50μM PS exposure
Nec-1
protection
RIP1 role Nec-1 DMSO 40mM -20ºC 50μM PS exposure
Fructose
protection
ATP role Fructose medium RT 40mM LDH activity
EDTA effect Extracellular
Ca2+
EDTA medium RT 20mM LDH activity
and PS
exposure
ERK
inhibition
EERK role U0126 DMSO 1mM 4ºC 20μM LDH activity
JNK
inhibition
JNK role SP600125 DMSO 25mM 4ºC 60μM LDH activity
p38
inhibition
p38 role SB203580 DMSO 5mM -20ºC 25μM LDH activity
66
Assay Investigation reagent solvent Stock
concentration
Storing Working
conentration
End point
measurement
NOX
inhibition
NOX role DPI DMSO 15mM -20ºC 60μM LDH activity
μ-calpain μ-calpain
role
Calpain
inhibitor I
DMSO 20mM -20ºC 60μM LDH activity
CaM
inhibition
CaM role W-7 DMSO 50mM -20ºC 50μM LDH activity
PKC
inhibition
PKC role Ro31-
8220
H2O 20mM 4ºC 30μM LDH activity
Simultaneous
CaM and
PKC
inhibition
Simultaneous
CaM and
PKC
inhibition
W-7 and
Ro31-
8220
50μM W-7
and 30μM
Ro31-8220
LDH activity
p53
inhibition
p53 role PFT-α DMSO 20mM -20ºC 40μM LDH activity
HO-1
inhibition
HO-1 role PPIX DMSO 18mM -20ºC 50μM LDH activity
WB PARP1
activation
Anti-
PARP1
antibody
Blocking
buffer
-20ºC 1/4000 Protein level
67
Chapter 3 In vitro cytotoxicity of HXR9
68
3.1 Introduction
The dysregulation of HOX genes is a common to many types of haematological
malignancies, including AML (Ferrando, Armstrong et al. 2003), and solid
malignancies including renal (Shears, Plowright et al. 2008), ovarian (Cheng, Liu et al.
2005; Morgan, Plowright et al. 2010), lung (Abe, Hamada et al. 2006; Plowright,
Harrington et al. 2009) and other cancers. Furthermore, over-expression of HOXA9 in
AML is a poor prognostic factor (Golub, Slonim et al. 1999), whilst down-regulation
of HOX genes is a favourable prognostic factor (Andreeff, Ruvolo et al. 2008;
Zangenberg, Grubach et al. 2009). HOX genes have an oncogenic function in
leukaemias (Eklund 2007) and their over-expression in melanoma maintain cell
survival, in part through preventing c-FOS transcription (Morgan, Pirard et al. 2007).
The small cell-permeable peptide HXR9 was used to disrupt the interaction
between HOX proteins and their partner transcription factor, PBX. This resulted in cell
death after a two-hour treatment in several cancer cell lines including melanoma, renal
and ovarian cells (Morgan, Pirard et al. 2007; Shears, Plowright et al. 2008; Morgan,
Plowright et al. 2010). Therefore, it was investigated whether AML cell lines were also
sensitive to HXR9.
I also studied the effect of combining HXR9 with commonly used
chemotherapies in AML treatment, MTX and DNR (Piccaluga, Visani et al. 2002;
Minotti, Menna et al. 2004; Fernandez 2010; Larson, Campbell et al. 2012).
69
3.1.1 Amis of chapter 3
To assess the expression of HOX genes in AML and the cytotoxicity of HXR9
on several AML cell lines, either alone or in combination with commonly used AML
chemotherapies.
To achieve these aims, the following experiments were performed:
1. The expression of the whole panel of 39 HOX genes was measured in five AML
cell lines (KG-1, HEL92.1.7, HL-60, KU812F and K562) by RT-PCR.
2. The cytotoxicity of HXR9 on the five AML cell lines was assessed by LDH
assay.
3. The cytotoxicity of common AML chemotherapy drugs including DNR and
MTX on two AML cell lines was assessed by LDH assay.
4. The interaction between HXR9 and DNR or MTX was conducted by LDH
assay.
70
3.2 Results
3.2.1 HOX gene expression in AML cell lines
Since HXR9 disrupts the interaction of HOX (1-10)-PBX, the expression of
these HOX genes was assessed by RT-PCR. The expression of HOX genes was studied
in the five AML derived cell lines HL-60, KG-1, HEL92.1.7, KU812F and K562. This
revealed a number of differences in HOX gene expression, with HOXA and HOXB
cluster genes (Figure 3.1A and B) more highly expressed than those of HOXC and
HOXD clusters (Figure 3.1C and D). It is also noteworthy that many HOXA cluster
were highly expressed in KG-1 cell line including HOXA5, HOXA6, HOXA9 and
HOXA10. As in KG-1, HOXA9 and HOXA10 were highly expressed in the KU812F
cell line, while HOXB9 was the most highly expressed gene in K562 and HEL92.1.7.
Intriguingly, HL-60 cells show very little HOX expression.
71
A)
B)
C)
72
D)
Figure 3.1 HOX gene expression in KG-1, HEL92.1.7, KU812F, K562 and HL-60 cell lines. A) The most
expressed HOXA genes in the KG-1 cell line were HOXA5, HOXA6, HOXA9 and HOXA10, while HOXA9 and
HOXA10 were most highly expressed in KU812F. B) HOXB9 was the most expressed HOX gene in HEL92.1.7
and K562. C) HOXC and D) HOXD genes showed very little expression in tested AML derived cell lines. The
expression of HOX genes were represented as the logarithm of the ratio of the expression of HOX genes to the
housekeeping gene β-actin (×10 000). Graphs express the mean of three independent experiments and error bars
show the SEM.
73
3.2.2 HXR9 is cytotoxic on all tested AML derived cell lines
To determine whether disrupting the interaction between HOX proteins and PBX
triggers cell death, the AML derived cell lines including KG-1, HEL 92.1.7 and HL-60, which
are derived from primary AML patients, and KU812F and K562, which are derived from
secondary AML patients, were exposed to titrations of HXR9 and its negative control CXR9
for two hours. The cytotoxic effect of HXR9 and CXR9 was then measured by LDH assay.
HXR9 had a dramatic cytotoxic effect on all tested AML cell lines, while CXR9 that lacks a
functional domain had no cytotoxicity (Figure 3.2). The IC50 values were measured by Calcusyn
software and were 4.5μM, 6.1μM, 19.9μM, 9.1μM and 10.4μM for KG-1, HEL 92.1.7, HL-60,
KU812F and K562, respectively (Table 3.1).
74
A)
B)
C)
75
D)
E)
Figure 3.2 LDH assay for HXR9 and CXR9 cytotoxicity on AML derived cell lines. The cell lines were treated
with titrations of HXR9, red curve, and CXR9, blue line, for two hours, and then the LDH enzyme activity was
measured in supernatants, which indirectly reflects cell survival. IC50s of HXR9 were calculated using Calcusyn
software. Graphs show the mean of three independent repeats that had R2 ≥ 0.95 and error bars show SEM. A)
KG-1, B) HEL 92.1.7, C) HL-60, D) KU812F and E) K562 cell lines.
Table 3.1 IC50 values of HXR9 on AML derived cell lines with SEM.
Cell line Source IC50 of HXR9 (μM) SEM
KG-1 Primary AML
patients
4.5 ± 0.353
HEL 92.1.7 6.1 ±0.720
HL-60 19.9 ±0.368
KU812F Secondary AML
patients
9.1 ±1.886
K562 10.4 ±0.281
The IC50 values represent the mean IC50 of three independent experiments that were calculated by Calcusyn
software.
76
3.2.3 DNR is cytotoxic on K562 and HL-60
In attempt to investigate the combination effect of HXR9 and chemotherapeutic drugs
on AML-derived cell lines, two of the conventional chemotherapies were selected including
DNR and MTX. It was reported that CD34+ AML cells, KG-1 and HEL92.1.7, were resistant
for both drugs (Bailly, Muller et al. 1995; Bailly, Skladanowski et al. 1997). Therefore, HL-60
and K562 that are represented primary and secondary AML, respectively, were selected to
conduct the combination therapy effect.
To determine the cytotoxicity of DNR on the AML cell lines K562 and HL-60, the cell
lines were treated with a titration of DNR for 24 hours and the cytotoxicity was assessed by
LDH assay. Both cell lines were sensitive to DNR at approximately the same concentration
after 24-hour treatment with an IC50 of 17.7μM and 17.4μM for K562 and HL-60, respectively
(Figure 3.3).
Figure 3.3 LDH assay for DNR cytotoxicity on K562 and HL-60 cell lines. The cell lines were treated with a
titration of DNR (range 2.5μM - 30μM) for 24 hours. The cytotoxicity of DNR was estimated by measuring LDH
enzyme activity in cell free supernatants, which indirectly reflects cell survival. The IC50 of DNR calculated using
Calcusyn software that was 17.7μM for K562 and 17.4μM for HL-60. The Graph shows the mean of three
independent repeats that had R2 ≥ 0.95 and error bars show the SEM. The red and blue curves represent K562 and
HL-60, respectively.
77
3.2.4 Combination effect of HXR9 and DNR
To determine the effect of a combination between HXR9 and DNR, the cell lines K562
and HL-60 were treated concurrently with HXR9 and DNR. The cell lines were exposed to each
drug alone or to the combination of the two drugs (Figure 3.4). In the single drug exposure, the
cell lines were exposed to HXR9 for two hours or to DNR for 24 hours. In the combination
therapy, the cell lines were exposed to DNR for 24 hours and in the last two hours HXR9 was
added. The combination effect of HXR9 and DNR was then estimated for significance using
Calcusyn software by calculating CI values, as described in the analysis of drug combination
assay data. The combination therapy had an antagonistic effect at the effective dose 50 (ED50)
on both cell lines, K562 (ED50 1.34 ± 0.75) and HL-60 (ED50 1.32 ± 0.016), while at ED90, the
combination therapy was synergistic on K562 (ED90 0.72 ± 0.12) and nearly additive on HL-
60 (ED90 0.98 ± 0.07) (Table 3.2).
78
A) B)
.
Table 3.2 The analysis of the combination effect of HXR9 and DNR by Calcusyn software. HXR9 and DNR
combination effect on K562 (A) and HL-60 (B) cells.
A) HXR9 and DNR combination effect on K562.
B) HXR9 and DNR combination effect on HL-60.
ED CI Combination effect
ED50 1.34 ± 0.75 - slight antagonism
ED75 0.98 ± 0.91 ± Nearly additive
ED90 0.72 ± 0.12 ++ Moderate synergism
ED CI Combination effect
ED50 1.32 ± 0.016 -- Moderate antagonism
ED75 1.129 ± 0.045 -- Moderate antagonism
ED90 0.98 ± 0.07 ± Nearly additive
Figure 3.4 LDH assay of HXR9 and DNR combination therapy for K562 and HL-60. Drug
concentrations used were 0.25, 0.5, 1, 2, 3 times the calculated IC50 of each drug in a constant ratio,
HXR9 IC50 was 21μM and 20μM, DNR IC50 was 17.7μM and 17.4μM for K562 and HL-60,
respectively. Cells were treated with HXR9, blue bars, or DNR, red bars, for 2 hours or 24 hours,
respectively. In addition, cells were treated with DNR for 24 hours and in the last two hours HXR9 was
added, green bars, in the combination therapy. The graphs show the mean of three independent repeats
and error bars show the SEM. A) is K562 and B) is HL-60 combination therapy graphs.
79
3.2.5 MTX induces K562 and HL-60 cell death
To assess the cytotoxicity of MTX on K562 and HL-60, cells were exposed to increasing
concentrations of MTX (range 2.5μM - 40μM) for 24 hours. The induction of cell death was
then determined by LDH assay. MTX induced cell death in both cell lines with an IC50 20.6μM
and 12.8μM for K562 and HL-60, respectively (Figure 3.5).
Figure 3.5 LDH assay for MTX cytotoxicity on K562 and HL-60 cell lines. The cell lines were treated with
different concentrations of MTX (range 2.5μM - 40μM) for 24 hours. Cell death was assessed by measuring LDH
enzyme activity in cell free supernatants that indirectly reflects cell survival. The IC50 of MTX was calculated
using Calcusyn software and was 20.6μM for K562 and 12.8μM for HL-60. The Graph shows the mean of three
independent repeats that had R2 ≥ 0.95 and error bars show the SEM. The red and blue curves represent K562 and
HL-60, respectively.
80
3.2.6 Combination effect of HXR9 and MTX
To assess the effect of HXR9 and MTX combination therapy, K562 and HL-60 cells
were incubated with each agent alone or concurrently (Figure 3.6). Cells were treated for two
hours and 24 hours with HXR9 and MTX, respectively. In the co-administration therapy, cells
were treated for 24 hours with MTX and in the last two hours HXR9 was administrated. The
effect of the HXR9/MTX combination was then analysed by the median effect method, as
described in the analysis of drug combination assay data. Using this method, HXR9/MTX
appeared an antagonistic interaction at all drug concentrations used on both cell lines with CI
values more than 1 (Table 3.3).
81
A) B)
Figure 3.6 LDH assay of HXR9/MTX combination therapy for K562 and HL-60. Drug concentrations used were
0.25, 0.5, 1, 2, 3 times the calculated IC50 of each drug in a constant ratio, the HXR9 IC50 was 21μM and 20μM,
and the MTX IC50 was 20.6μM and 12.8μM for K562 and HL-60, respectively. Cells were treated with HXR9,
blue bars, or MTX, red bars, for 2 hours or 24 hours, respectively. In addition, cells were treated with DNR for 24
hours and in the last two hours HXR9 was added, green bars, in the combination therapy. The Graphs show the
mean of three independent repeats and error bars show SEM. A) is K562 and B) is HL-60 combination therapy
graphs.
Table 3.3 The analysis of the combination effect of HXR9 and MTX by Calcusyn software.
A) HXR9 and MTX combination effect on K562.
B) HXR9 and MTX combination effect on HL-60.
ED CI Combination effect
ED50 1.47 ± 0.019 - - - Antagonism
ED75 1.62 ± 0.34 - - - Antagonism
ED90 1.53 ± 0.175 - - - Antagonism
ED CI Combination effect
ED50 2.06 ± 0.50 - - - Antagonism
ED75 1.75 ± 0.28 - - - Antagonism
ED90 1.63 ± 0.19 - - - Antagonism
82
3.3 Discussion
The expression of all 39 HOX genes was evaluated in this study in five AML derived
cell lines including KG-1 and HEL92.1.7 from erythroleukaemic patients (Koeffler and Golde
1978; Martin and Papayannopoulou 1982), HL-60 from APML (Collins, Ruscetti et al. 1978),
and KU812F and K562 derived from CML patients in blast crisis (Lozzio and Lozzio 1975;
Kishi 1985). Our findings of HOX expression concur with previous studies. Whilst the myeloid
origin cell line KU812F highly expressed HOXA9 and HOXA10 genes, the erythroid cell line
HEL92.1.7 highly expressed HOXB9 gene consistent with previous findings (Giampaolo,
Sterpetti et al. 1994; Kawagoe, Humphries et al. 1999; Pineault, Helgason et al. 2002). Although
K562 originated from a CML patient in blast crisis, this cell highly expressed HOXB9 gene
which supports previous studies suggesting that K562 is in fact a human erythroleukaemia cell
line (Andersson, Jokinen et al. 1979; Andersson, Nilsson et al. 1979). Interestingly, KG-1
highly expressed HOXA genes including HOXA5, HOXA6, HOXA9 and HOXA10 even though
it is an erythroid line. It was reported that KG-1 highly expressed HOXA9 and HOXA10 genes
due to the presence of a trisomy 8 mutation (Kok, Brown et al. 2010). However, the associating
mechanism between up-regulation of HOXA genes and trisomy 8 is unclear. Intriguingly, the
APML cell line HL-60 hardly expressed HOX genes, but this is in line with previous studies in
which APML was found to be characterised by global down-regulation of HOX genes
(Thompson, Quinn et al. 2003). Notably, the most expressed subset of HOX genes were the
posterior (6-13) HOX genes. One potential explanation for their up-regulation in AML cell lines
is their ability to interrupt haematopoietic differentiation (Pineault, Abramovich et al. 2004).
Overall, the expression of HOX genes in the tested AML lines was consistent with the pre-
established notion that HOX genes are expressed in a lineage-restricted manner, whereby HOXA
and HOXB genes are expressed in myeloid and erythroid cells, respectively (Giampaolo,
Sterpetti et al. 1994; Kawagoe, Humphries et al. 1999; Pineault, Helgason et al. 2002).
83
Here it is showed that the high expression of HOX genes can be used as a therapeutic
target in AML as targeting the HOX-PBX interaction with HXR9 caused cell death in the tested
AML derived cell lines. The IC50 of HXR9 for the treated AML derived cell lines ranges from
4.5μM to 20μM after a two-hour treatment. The sensitivity of the tested cell lines to HXR9
directly correlated to the level of HOX genes expression, for example, KG-1 cells that highly
expressed HOXA5, HOXA6, HOXA9 and HOXA10 were the most sensitive to HXR9 with an
IC50 4.5μM after a two-hour treatment while HL-60 cells, which show a global down-regulation
of HOX genes were the least sensitive to HXR9, with an IC50 19.9μM after a two-hour treatment.
Furthermore, there was no difference in terms of HXR9 sensitivity between the HEL92.1.7 cells
derived from a primary AML patient and KU812F and K562, derived from CML patients in
blast crisis (secondary AML) which might be because these cells express the most highly
expressed HOX gene at the same level. In general, the tested AML cell lines were more sensitive
to HXR9 than other cell lines derived from solid malignancies including melanoma, renal,
ovarian and lung cancers (Table 3.4) (Shears, Plowright et al. 2008; Plowright, Harrington et
al. 2009; Morgan, Plowright et al. 2010). The difference in the sensitivity between AML cells
and solid cancers might be because survival of AML cells is totally depending upon the HOX-
PBX interaction, or due to the abundant expression of HOX genes in AML cells compared to
the solid cancers.
84
Table 3.4 The IC50 values of HXR9 with cell lines derived from different cancers.
Cell line Cell line source IC50 Treatment period Reference
KG-1, HEL92.1.7
and HL60
Primary AML 4.5μM , 6.1μM and
19.9μM
2 hours
This study
KU812F and K562 Secondary AML 9.1μM and 10.4μM
769-p and CaKi-2 Renal 40μM and 44μM (Shears, Plowright et al.
2008)
B16F10 Melanoma 20μM (Morgan, Pirard et al.
2007)
SK-OV3 Ovarian 70μM (Morgan, Plowright et
al. 2010)
A549 and H23 Lung 32.5μM and 69μM 24 hours (Plowright, Harrington
et al. 2009)
This table shows the difference in the IC50 values between AML cell lines investigated in this study and cell lines
derived from solid cancers including renal, melanoma, ovarian and lung cancers established in previous studies.
All cells were treated with HXR9 for two hours except lung cancer cells, which were treated for 24 hours.
Generally, AML cell lines were the most sensitive to HXR9.
The drug-interaction effect of HXR9 and DNR was performed using the Chou and
Talaly method of calculating the median effect dose (Chou 1991). Results showed that the
combination effect was antagonistic at ED50 in both of the tested AML cell lines K562 and HL-
60, but it was either synergistic or additive at ED90 in K562 or HL-60, respectively. DNR causes
apoptosis by blocking topoisomerase II, intercalating with DNA, and producing ROS, and leads
to nuclear factor-kappa B (NF-ҠB)-DNA binding as a cellular response to stress (Lanzi,
Gambetta et al. 1991; Boland, Foster et al. 1997; Boland, Fitzgerald et al. 2000; Laurent and
Jaffrézou 2001; Campbell, O'Shea et al. 2006; Karl, Pritschow et al. 2009). Interestingly, NF-
ҠB has an anti-apoptotic role in DNR treatment, as reducing the activity of NF-ҠB and reducing
the degradation of the NF-ҠB inhibitor IҠB using bortezomib increases the sensitivity of K562
to DNR (Wang, Wang et al. 2012). In addition, DNR causes up-regulation of PKC that reduces
DNR cytotoxicity by enhancing NF-ҠB-DNA binding and reducing ROS production (Lanzi,
85
Gambetta et al. 1991; Laurent and Jaffrézou 2001; Bezombes, de Thonel et al. 2002). However,
Inhibition of PKC sensitized the AML cell line U937 to DNR (Mas, Hernandez et al. 2003). In
addition, it has been found that ROS production as a result of DNR treatment slows the kinetics
of NF-ҠB activation and DNA binding (Campbell, O'Shea et al. 2006), which might explain the
difference in effect of combining low or high doses of HXR9 and DNR. In the low dose DNR
and HXR9 combination therapy, the level of ROS production might be insufficient to delay
NF-ҠB-DNA binding long enough for HXR9 to cause the up-regulation of c-FOS that mediates
HXR9 toxicity and is also a negative regulator of NF-ҠB (Morgan, Pirard et al. 2007; Plowright,
Harrington et al. 2009; Takada, Ray et al. 2010). However, at high concentrations of both drugs,
DNR might produce more ROS that would delay the activation of NF-ҠB, allowing HXR9 to
cause an up-regulation of c-FOS that would inhibit NF-ҠB activity and ultimately lead to either
a synergistic effect in K562 or at least additive effect in HL-60. These data suggest that HXR9
is synergistic or at least additive with DNR, a drug commonly used for AML treatment (Figure
3.7).
Figure 3.7 The proposed interaction mechanism between DNR and HXR9. DNR blocks topoisomerase II that
leads to activation of the surviving pathway PKC/NF-κB, which in turn reduces necrosis by HR9 through reduction
of c-FOS activity, and the apoptotic pathway through ROS signalling that delays the activity of NF-κB.
86
Unlike DNR, MTX has an antagonistic effect on HXR9. This antagonism is owing to
the conflict in the mechanism of both drugs. MTX causes apoptosis by inhibiting topoisomerase
II, which involves the production of a transient DNA double-strand break, thereby damaging
DNA that in turn activates NF-ҡB-DNA binding as a consequence of cellular stress response
(Boland, Foster et al. 1997; Boland, Fitzgerald et al. 2000; Neuhaus, Kieseier et al. 2004;
Campbell, O'Shea et al. 2006; Karl, Pritschow et al. 2009). Unlike DNR, ROS production is not
involved in MTX mediated apoptosis (Campbell, O'Shea et al. 2006). In addition, MTX leads
to activation of PKC, which is a positive upstream regulator of NF-ҡB and it is has been
reported that pre-treating cells with a PKC activator enhanced the cytotoxicity of MTX (Bhalla,
Ibrado et al. 1993; Estévez, Vieytes et al. 1996; Lin, O'Mahony et al. 2000; Mut, Amos et al.
2010; Lutzny, Kocher et al. 2013). However, PKC appeared to cause resistance to HXR9 as it
was found in this study. While c-FOS, c-JUN and EGR-1 over-expression caused relapse in
AML patients treated with MTX, c-FOS over-expression is considered as a key event in
apoptosis induced by HXR9; EGR-1 and c-JUN are also up-regulated by HXR9 (Bailly,
Skladanowski et al. 1997; Potapova, Basu et al. 2001; Staber, Linkesch et al. 2004; Morgan,
Pirard et al. 2007; Plowright, Harrington et al. 2009). Whilst disruption of NF-ҡB by short
interfering RNA caused a dramatic reduction in apoptosis upon MTX treatment, c-FOS was
reported as a negative regulator of NF-ҡB mediated stress response and c-fos-/- mice showed an
elevation in NF-ҡB activity (Ray, Kuwahara et al. 2006; Karl, Pritschow et al. 2009; Takada,
Ray et al. 2010). Likewise, MTX treatment leads to a significant reduction of c-Myc, while c-
Myc was found to be a downstream effector of c-FOS-induced apoptosis (Bhalla, Ibrado et al.
1993; Kalra and Kumar 2004). In addition, the expression of c-FOS was high in MTX-resistant
AML cell lines (Bailly, Skladanowski et al. 1997) and c-FOS positively regulates ABCG2, also
known as breast cancer resistance protein (BCRP), which mediates MTX-resistance in the
87
breast cancer cell line MCF-7 (Bailey-Dell, Hassel et al. 2001; Schwabedissen, Grube et al.
2006). Taken together, these data suggest that HXR9 and MTX exerted a reciprocal inhibition.
3.3.1 Summary of chapter
In summary, this chapter studies show that:
AML cell lines derived from different AML types express different subsets of HOX
genes at different levels due to the heterogeneity of AML.
The HOX-PBX interaction is crucial for the tested AML cell lines survival and targeting
this interaction using HXR9 leads to cell death.
HXR9/DNR combination therapy is synergistic or at least additive.
HXR9/MTX combination therapy is antagonistic due to the conflict in their mechanisms
of action.
88
Chapter 4 The mechanism of HXR9 cytotoxicity
89
4.1 Introduction
Better understanding of the molecular aberrations in cancers resulted in the development
of several new potential therapeutic agents such as HXR9 that target HOX-PBX interaction.
However, molecular agents combined with conventional chemotherapies have given rise to
more curative therapies. In order to provide a rational basis for the selection of a synergistic
partner for HXR9, the mechanism by which HXR9 induces cell death needs to be identified.
The specificity of HXR9 for the HOX-PBX interaction was previously demonstrated in
melanoma and lung cancer cells (Morgan, Pirard et al. 2007; Plowright, Harrington et al. 2009),
as were downstream transcriptional changes. Notably, the increased expression of c-FOS was
responsible, at least in part, for the HXR9 mediated cell death (Morgan, Pirard et al. 2007;
Shears, Plowright et al. 2008; Plowright, Harrington et al. 2009; Morgan, Plowright et al. 2010;
Morgan, Boxall et al. 2012; Morgan, Boxall et al. 2014), subsequently, the expression of c-FOS
was assessed in this study. In addition, the up-regulation of other genes including ATF1, ATF3,
NRUA3, JUN, DUSP1 was also observed (Morgan, Pirard et al. 2007; Shears, Plowright et al.
2008; Morgan, Boxall et al. 2012). Previous studies showed that the general inhibition of
caspases did not completely prevent HXR9-triggered cell death, which suggests that there may
be additional mediators of cell death (Shears, Plowright et al. 2008; Morgan, Boxall et al. 2012;
Morgan, Boxall et al. 2014). However, the nature of these potential mediators of cell death by
HXR9 remains unknown. Therefore, the possible involvement of necrotic pathways in HXR9
induced cell death was investigated. In addition, the molecular mechanism of HXR9 action is
yet to be identified, subsequently, the potential role of MAPK pathways including MEK/ERK,
p38 and JNK were assessed.
90
Cell death can be classified into accidental cell death and programmed cell death (PCD).
Accidental cell death results from extreme environmental disturbances including severe
hypoxia, and extreme temperature changes. Necrosis was described as an unregulated and
accidental cell death. However, it has also been suggested that necrosis might result from
regulated enzyme-catalysed biochemical reactions (Fink and Cookson 2005). Therefore,
necrosis can be categorised as PCD when it results from proteolytic reactions and called
necrotic-like PCD or regulated necrosis (Galluzzi, Kepp et al. 2014). Regulated necrosis is
classified into mitochondrial necrosis, necroptosis and parthanatos (Figure 4.1). This type of
cell death is characterised by the absence of hallmarks of apoptosis such as caspase activation,
Bcl-2 family member involvement, cyt C release, chromatin condensation, nuclear
fragmentation and PARP1 cleavage (Saraste and Pulkki 2000; Ziegler and Groscurth 2004;
Miao and Degterev 2009).
Figure 4.1 A schematic diagram of cell death. It can be classified into programmed and accidental cell death. The
former is further sub-classified into apoptosis and regulated necrosis. Apoptosis is sub-divided into extrinsic and
intrinsic pathways, both of which are caspase-dependent. Regulated necrosis, which is caspase-independent, is
subdivided into mitochondrial necrosis, necroptosis and parthanatos.
91
Cells undergo apoptosis via two main pathways, extrinsic and intrinsic pathway, both
of which eventually result in caspase activation (Figure 4.1). The extrinsic pathway is initiated
by the binding of ligands to cell surface death receptors such as those of the tumour necrosis
factor (TNF) receptor family that in turn leads to activation of initiator pro-caspases such as
pro-caspase-8 and/or pro-caspase-10 (Jin and El-Deiry 2005). Active caspase-8 and-10 cleave
the executioner pro-caspases-3/7 (Huerta, Goulet et al. 2007), which in turn cleave many
substrates including PARP1 and cytoskeletal proteins, and ultimately cause cell death by
apoptosis (Vermeulen, Van Bockstaele et al. 2005). The intrinsic pathway, also called the
mitochondrial pathway, involves the release of pro-apoptotic proteins that are trapped in
mitochondria such as cyt C and apoptosis inducing factor (AIF) (Martin 2010). Mitochondrial
membrane integrity is controlled by Bcl-2 family members (Galluzzi, Kepp et al. 2014), which
include anti-apoptotic proteins such as Bcl-2 and Bcl-xL, and pro-apoptotic proteins such as
Bad, Bid, Bax and Bak1 (Jin and El-Deiry 2005). Under normal conditions, the anti-apoptotic
members of Bcl-2 and Bcl-xL heterodimerize with the pro-apoptotic members Bax and Bak1
and consequently preventing their apoptotic function (Martin 2010). During apoptosis, caspase-
8 cleaves Bid that in turn with dephosphorylated Bad promotes apoptosis by binding to the anti-
apoptotic Bcl-2 and Bcl-xL proteins (Li, Zhu et al. 1998; Ghibelli and Diederich 2010; Martin
2010). As a result, Bax and Bak1 are released and form Bax/Bak channels in the outer
mitochondrial membrane causing mitochondrial permeabilization (Martin 2010). Cyt C is then
released into the cytosol and, in the presence of ATP, binds to Apaf-1 and pro-caspase-9
forming the pro-apoptotic complex apoptosome that leads to auto-cleavage and activation of
caspase-9 and caspase-3 and -7 (Shi 2002). Then, active caspases 3, 7 and 9 proteolytically
cleave XIAP which in healthy viable cells binds and inhibits the three caspases (Vermeulen,
Van Bockstaele et al. 2005). Therefore, both extrinsic and intrinsic apoptotic pathways was
investigated to assess the involvement of apoptosis in HXR9 cell death. In addition, due its
92
known role in both apoptotic pathways, p53 and its downstream effector p21 involvement in
HXR9 caused cell death were investigated.
Ca2+ is one of the main mediators of regulated necrosis and studying its role in necrosis
is common. Subsequently, assessing the involvement of extracellular Ca2+ in HXR9 induced
cell death was addressed in this study. In addition, the role of Ca2+-downstream effectors
including NOX, µ-calpain, CaM, PKC was assessed.
CypD is a member of the cyclophilin family located in the mitochondrial matrix (Elrod
and Molkentin 2013). It is considered as the most important regulator of mPTP because it is the
only protein known to be required for mitochondrial permeability transition-dependent necrosis
(Alam, Baetz et al. 2014).
Necroptosis is a type of regulated necrosis that is mediated by the interaction between
RIP1 and RIP3 to form a necrosome (Marshall and Baines 2014). RIP1 can be either a
component of a pro-survival complex inducing the expression of NF-ĸB, an apoptotic complex
enhancing caspase-dependent apoptosis, or a necrosome complex initiating necroptosis
(Vandenabeele, Grootjans et al. 2013). However, RIP1 kinase activity and necrosome formation
are only required in necroptosis (Cho, Challa et al. 2009; He, Wang et al. 2009; Zhang, Shao et
al. 2009). The molecular downstream mechanism of the necrosome that eventually results in
necroptotic cell death is yet to be identified, and it remains unclear whether mitochondria are
involved (Cho, Challa et al. 2009; Zhang, Shao et al. 2009; Tait, Oberst et al. 2013). In addition,
mixed lineage kinase domain-like protein and phosphoglycerate mutase family member 5 were
identified as RIP3 downstream mediators of necroptosis (Sun, Wang et al. 2012; Wang, Jiang
et al. 2012).
Parthanatos is a form of regulated necrosis that is PARP1-dependent and caspase-
independent. This form of necrosis is characterised by hyper-activation of PARP1 that leads to
loss of mPTP and subsequently nuclear translocation of AIF from mitochondria, loss of ATP
93
and ultimately cell necrosis (Andrabi, Dawson et al. 2008). PARP1 is a DNA repair protein that
is activated in response to DNA damage by alkylating agents and Ca2+ signalling pathways
(Galluzzi, Kepp et al. 2014).
94
4.1.1 Aims of chapter 4
To investigate the mechanism of HXR9 action.
To achieve this aim, the following studies were performed:
1. Assess whether HXR9 causes up-regulation of c-FOS in AML cell lines as it did in all
previously tested cancers.
2. To determine whether HXR9 induced apoptosis in AML cells the following assays were
used:
a. Annexin V/7-AAD and DAPI staining assays to assess the morphological
changes.
b. RT-PCR to measure the expression of some members of the Bcl-2 family,
initiator and executioner caspases, apoptosis protease-activating factor 1
(Apaf1), X linked inhibitor of apoptosis (XIAP) and PARP1.
c. Inhibition of either caspase-3 by Z-DEVD-R110 and general inhibition of
caspases by z-VAD-FMK to analyse the role of caspases in the cytotoxicity of
HXR9.
3. To assess whether necrosis is involved in the cytotoxicity of HXR9, the following
studies were performed:
a. Determining the effect of chelating extracellular Ca2+ by EDTA on the efficacy
of HXR9 by LDH and annexin V/7-AAD assays.
b. Assessing the role of CypD in the HXR9 cytotoxicity by inhibiting its role using
CsA.
c. Assessing the role of RIP1 in the HXR9 cytotoxicity by inhibiting its role using
Nec-1.
d. Assessing the effect of depleting ATP by fructose on the effectiveness of HXR9.
95
e. Determining whether HXR9 causes activation of PARP1 protein.
4. To determine the role of MAPK pathways and their downstream targets, the following
assays were analysed:
a. Members of MAPK pathway MEK/ERK, p38 and JNK.
b. NOX proteins.
c. μ-calpain, CaM, PKC and HO-1 proteins.
5. To assess whether p53 is involved in HXR9-induced cell death, the following assays
were done:
a. Measuring the expression of p53 and its downstream target p21 mRNA.
b. Inhibiting p53 protein.
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4.2 Results
4.2.1 HXR9 causes the up-regulation of c-FOS
c-FOS up-regulation was found to be a mediator of apoptosis triggered by HXR9 in
melanoma, lung, ovarian, breast and prostate derived cell lines (Morgan, Pirard et al. 2007;
Plowright, Harrington et al. 2009; Morgan, Plowright et al. 2010; Morgan, Boxall et al. 2012;
Morgan, Boxall et al. 2014). To determine whether HXR9 also causes up-regulation of c-FOS
in AML derived cell lines, the expression of c-FOS was measured in untreated, CXR9 or HXR9
treated cells by RT-PCR. Cells were treated for two hours with the IC50s of HXR9 or equivalent
concentrations of the negative control peptide CXR9, which were 4.5μM, 6μM, 20μM, 9μM
and 10.5μM for KG-1, HEL92.1.7, HL-60, KU812F and K562, respectively. A two-hour
treatment with HXR9 resulted in a 7.5, 16.7, 3.3, 50, and 7.5 fold increase in c-FOS expression
in KG-1, HEL92.1.7, HL-60, KU812F and K562, respectively, compared to untreated cells.
However, there was no significant change in c-FOS expression between untreated cells and
CXR9 treated cells (Figure 4.1).
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Figure 4.2 c-FOS expression in AML cell lines after treatment with HXR9. AML cell lines were treated with the
following concentrations of HXR9 or CXR9: 4.5μM, 6μM, 20μM, 9μM and 10.5μM for KG-1, HEL92.1.7, HL-
60, KU812F and K562, respectively, or were untreated (control), for two hours. After a two-hour treatment, mRNA
was extracted from cells and c-FOS expression was measured by RT-PCR. Results are presented as a ratio between
c-FOS and β-actin (x10 000). A two-hour treatment with HXR9 caused an elevation of c-FOS transcript levels
comparing to untreated cells. Graphs show the mean of three independent repeats and error bars show the SEM.
*p < 0.05, **p < 0.01 and ***p < 0.001 with respect to untreated cells.
98
4.2.2 HXR9 causes death of AML tested cell lines
In order to determine the mode of cell death following HXR9 treatment, an annexin V-
PE assay was used. Cell lines were treated with the IC50, double and triple the IC50 of HXR9 or
of CXR9 equivalent to the highest concentration of HXR9 for two hours and then stained with
annexin V and 7-AAD. Cells were classified either as viable, early apoptotic or dead cells
according to 7-AAD penetration and annexin V binding (Table. 2.7). After a two-hour
treatment, there was no significant change in untreated cells and CXR9 treated cells, whilst
cells treated with the IC50 of HXR9 rapidly entered apoptosis, and many were dead (Figure 4.2).
For example, in untreated KU812F cells, 95% were viable cells, 2% and 3% were in early
apoptosis or dead, respectively, while 55% were viable, 20% were early apoptotic and 30%
were dead after 9μM HXR9 treatment (Figure 4.2D). Generally, cell treatment with either
HXR9 concentration killed the majority of cells. For example, treating HL-60 cells with 20μM
HXR9 killed 55% of cells, while 27% and 18% were viable or in an early apoptotic state,
respectively (Figure 4.2C).
99
A)
B)
C)
100
D)
E)
Figure 4.3 Detection of cell death in AML cell lines by flow cytometry. Cells were treated with the IC50, 2xIC50,
or 3xIC50 of HXR9, and with a concentration of CXR9 equivalent to the highest concentration of HXR9 for two
hours. Cells were then stained with annexin V and 7-AAD for 15 minutes in the dark and analysed by flow
cytometry. In all the tested cell lines, the number of early apoptotic and dead cells increased with increased HXR9
concentrations. Results are expressed as the mean of three separate experiments and error bars show the SEM. *p
< 0.05, **p < 0.01, ***p < 0.001 and ****p < 0.0001 with respect to untreated cells. A) KG-1, B) HEL92.1.7, C)
HL-60, D) KU812F and E) K562 cells.
101
4.2.3 HXR9 does not affect caspase or Bcl-2 family transcription
RT-PCR was used to analyse the transcriptional changes of several pro- and anti-
apoptotic genes upon disruption of the HOX/PBX dimer by HXR9. This analysis revealed that
HXR9 did not cause transcriptional changes in the initiator caspase-9 and its partner Apaf1 that
together form an apoptosome complex with cytochrome C (cyt C) thereby starting the intrinsic
apoptosis pathway (Adrain and Martin 2001). In addition, executioner caspases-3, -6 and -7,
which are key mediators of apoptosis (Huerta, Goulet et al. 2007), were not up-regulated after
HXR9 treatment. Likewise, none of the pro-apoptotic members of the Bcl-2 family including
Bad, Bax, Bak1 and Bid, nor the anti-apoptotic member Bcl-2, which mediate mitochondrial
cell death (Green and Kroemer 2004), showed transcriptional changes after HXR9 treatment.
In addition, neither the transcription of PARP1 nor XIAP was affected by HXR9 treatment
(Figure 4.3).
102
A)
B)
C)
103
D)
E)
Figure 4.4 Analysis of transcriptional changes of several pro- and anti-apoptotic genes upon HXR9 treatment.
Cells were treated with the IC50 of HXR9, CXR9 or untreated (control) for two hours. Then, mRNA was harvested
from the AML derived cell lines KG-1, HEL92.1.7, HL-60, KU812F and K562 and gene expression was analysed
using RT-PCR. There was no significant change in any of the analysed genes. Results are presented as a ratio with
β-actin (x10 000). Statistical analysis was performed using student's t-test by comparing the relative expression of
a gene of interest in HXR9 or CXR9 treated cells to its counterpart in untreated cells. Graphs show the mean of
three independent experiments and error bars show the SEM. Results are shown for A) KG-1, B) HEL92.1.7, C)
HL-60, D) KU812F and E) K562 cells.
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4.2.4 HXR9 does not activate caspase-3 in the tested AML cell lines
In order to study the mechanism of cell death, caspase-3 activity was measured after
treating cells with increasing concentrations of HXR9 or with a CXR9 concentration that was
equivalent to the highest concentration of HXR9. A fluorescently labelled caspase-3 substrate
and Ac-DEVD-CHO inhibitor were also added. As an additional control, R110 standard curves
were generated to validate individual experiments (Appendix 1). There were no significant
changes between untreated and HXR9 treated cells with respect to caspase-3 activity in any of
the AML cell lines (Figure 4.4).
105
A)
B)
C)
106
D)
E)
Figure 4.5 Detection of caspase-3 activity in AML cell lines using Z-DEVD-R110. Cells were treated with IC50,
2xIC50, 3xIC50 of HXR9, with an equivalent CXR9 to the highest concentration of HXR9 or untreated (control) for
two hours. Then, cells were harvested and lysed. Next, Ac-DEVD-CHO, a caspase inhibitor, was added to some
cells as a control. The Z-DEVD-R110 substrate was then added to cells and the fluorescence was read at 492nm.
Results were then normalised to lysis buffer fluorescence reading, background levels. There was no statistical
difference in caspase-3 activity between both HXR9- or CXR9-treated cells and untreated control cells. Graphs
show the mean of three independent experiments and error bars show the SEM. A) A graph represents caspase-3
activity of KG-1, B) HEL92.1.7, C) HL-60, D) KU812F and E) K562 cells.
KU812F
Untreated 27M CXR9 9M HXR9 18M HXR9 27M HXR9100
101
102
103
104
105
106
107
HXR9
HXR9+Ac-DEVD-CHO
Treatment
Flu
ore
scence
107
4.2.5 HXR9 efficacy is not affected by the general inhibition of caspases
In order to further study the role of caspases in HXR9-induced cell death, K562 and
HL-60 cells were pre-treated with or without the general inhibitor of caspase activity z-VAD-
FMK. DNR that has been shown to activate caspases was used as a positive control (Liu, Kelsey
et al. 2002; Kim, Park et al. 2003). Results revealed that 50μM z-VAD-FMK pre-treatment
dramatically increased the percentage of viable cells in DNR treated cells, p < 0.0001. However,
there was no statistical difference in terms of the percentage of viable cells between z-VAD-
FMK non- or pre-treated cells. For example, the percentage of viable cells was 62% and 58%
for z-VAD-FMK non- or pre-treated K562 cells, respectively (Figure 4.5). Likewise, pre-
treatment with z-VAD-FMK did not cause significant changes in the early apoptotic and dead
cells comparing with z-VAD-FMK non-treated cells (Appendix 2). This suggests that HXR9
kills cells in a caspase-independent pathway.
108
A)
B)
Figure 4.6 General inhibition of caspase activity in HXR9 treated AML cell lines by z-VAD-FMK. Cells were
pre-treated either with or without 50μM z-VAD-FMK for one hour, and then treated with the IC50, or 2xIC50 of
HXR9 for two hours or with 17.5μM DNR for 24 hours, which was used as a positive control. Next, cells were
harvested and stained with annexin V and 7-AAD for 15 minutes in the dark and analysed by flow cytometry. Data
showed that there was no statistical difference in terms of sensitivity to HXR9 between pre-treated cells with or
without 50μM z-VAD-FMK. Graphs show the mean of three independent experiments and error bars show the
SEM. ****p < 0.0001 with respect to z-VAD-FMK untreated cells. A) K562 and B) HL-60 cells.
109
4.2.6 HXR9 does not cause nuclear fragmentation
In order to assess changes in nuclear morphology upon HXR9 treatment, AML derived
cell lines were stained with the fluorescent stain DAPI and analysed by a fluorescence
microscope. Changes in the nuclear morphology such as chromatin condensation and nuclear
fragmentation are markers of late stage apoptosis (Collins, Schandl et al. 1997; Ziegler and
Groscurth 2004; Huerta, Goulet et al. 2007). It was found that treatment of cells with the IC50s
of HXR9 did not result in changes to the nuclear morphology when comparing to untreated
cells in all investigated cell lines (Figure 4.6). This suggests that HXR9-induces cell death in a
nuclear fragmentation-independent pathway.
110
Untreated IC50 HXR9
KG-1
HEL92.1.7
HL-60
111
Untreated IC50 HXR9
Figure 4.7 DAPI staining of AML cells upon HXR9 treatment. Cells were treated with the IC50s of HXR9 for 2
hours. Cells were then harvested, fixed in 4% formaldehyde, and cytospan. Fixed cells were stained with DAPI
for 5 minutes at the dark. Finally, stained cells were analysed by a fluorescent microscope at 20x lenses.
KU812F
K562
112
4.2.7 Ca2+ chelating abrogates cell killing by HXR9
As HXR9 seems to cause cell death in a caspase-independent manner, a number of
alterative pathways were investigated. Ca2+ is known to mediate caspase-independent cell death
(Dong, Li et al. 2005; Festjens, Vanden Berghe et al. 2006). Therefore, to analyse the role of
Ca2+ in HXR9-induced cell death, the extracellular Ca2+ chelator EDTA was used. Interestingly,
chelating extracellular Ca2+ rapidly attenuated the cytotoxicity of HXR9 and rescued cells from
cell death. For example, the percentage of viable cells increased from 50% to 80%, ****p <
0.0001, upon co-treatment with 20mM EDTA in 20μM HXR9 treated K562 cells (Figure 4.7A).
Likewise, in 20μM HXR9 treated HL-60 cells, the percentage of viable cells increased threefold
in 20mM EDTA co-treated cells compared to untreated cells (Figure 4.7B). In addition, co-
treatment with EDTA dramatically decreased the percentage of early apoptotic and dead cells
in both cell lines (Appendix 3).
113
A)
B)
Figure 4.8 Effect of EDTA on HXR9 cytotoxicity. Cells were pre-treated with or without 20mM EDTA for one
hour, and then treated with 10μM, 20μM and 40μM HXR9, prepared in media either with or without 20mM EDTA,
for two hours. Next, cells were harvested and stained with annexin V and 7-AAD for 15 minutes in the dark and
analysed by flow cytometry. Results showed that Pre- and co-treatment of cells with 20mM EDTA significantly
increased the % viable cells and thereby inhibited the cytotoxicity of HXR9. Graphs show the mean of three
independent experiments of viable cells, non-stained cells with either stain, and error bars show the SEM. ***p <
0.001 and ****p < 0.0001 with respect to cells not exposed to 20mM EDTA. A) K562 and B) HL-60 cells.
114
4.2.8 EDTA rescues cells from killing by HXR9
In order to confirm the previous results that showed chelating extracellular Ca2+ reduced
the cytotoxicity of HXR9, an LDH assay was performed to analyse the effect of EDTA pre-
and co-treatment. The results of the LDH assay were consistent with the results obtained using
the annexin V assay. Chelating extracellular Ca2+ by EDTA markedly reduced the levels of
LDH enzyme detected in K562 and HL-60 cells. In addition, HXR9 did not have an IC50 up to
the highest concentration used, 50μM (Figure 4.8). These results showed that EDTA prevented
cell killing by HXR9.
115
A)
B)
Figure 4.9 LDH analysis of the effect of EDTA on HXR9 cytotoxicity. Cells were pre-treated with or without
20mM EDTA for one hour, and then with varying concentrations of HXR9 for two hours prepared in media either
with or without 20mM EDTA, red and blue curves, respectively. LDH activity was then measured in the
supernatants, which indirectly reflects the fraction of surviving cells. Pre- and co-treatment of cells with 20mM
EDTA significantly attenuated the cytotoxicity of HXR9. Graphs show the mean of three independent experiments
and error bars show the SEM. *p < 0.05, **p < 0.01, ***p < 0.001 and ****p < 0.0001 with respect to cells not
treated with 20mM EDTA. A) K562 and B) HL-60 cells.
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4.2.9 Cell killing by HXR9 does not involve CypD
To assess whether HXR9 causes mitochondrial necrotic cell death through activation of
CypD that in turn leads to formation of mPTP, cells were treated with CsA that binds to CypD
and inhibits mitochondrial cell death (Li, Johnson et al. 2004; Basso, Fante et al. 2005; Dube,
Selwood et al. 2012). H2O2, which triggers mPTP formation and mitochondrial necrotic cell
death, was used as a positive control for this assay (Baines, Kaiser et al. 2005; Nakagawa,
Shimizu et al. 2005; Schinzel, Takeuchi et al. 2005; Jobe, Wilson et al. 2008). Cell death was
measured using the annexin V assay. Treatment with 5μM CsA significantly increased the
percentage of viable cells exposed to H2O2 from 44.9% ± 1.7 to 69.2% ± 1.34, ****p < 0.0001,
and from 0.92% ± 0.3 to 34.2% ± 2.4, ****p < 0.0001, of H2O2-treated K562 and HL-60,
respectively. However, CsA at 5μM could not rescue K562 cells from death caused by HXR9
treatment, and interestingly, concurrent CsA treatment with HXR9 significantly decreased the
survival of HXR9 treated HL-60 cells (Figure 4.9, appendix 4). Taken together, it can be
concluded that CypD activation is not required for HXR9-induced K562 and HL-60 cell death.
117
A)
B)
Figure 4.10 Effect of CsA on HXR9 cytotoxicity. Cells were pre-treated with or without 5μM CsA for one hour,
and then with the IC50, 2xIC50, 3xIC50 of HXR9, 20mM H2O2 (positive control), or were left untreated (negative
control) for two hours with or without 5μM CsA. The cells were then harvested and stained with annexin V and
7-AAD for 15 minutes in the dark and analysed by flow cytometry. Results showed no statistical difference in the
proportion of viable cells between K562 cells (A) incubated with or without 5μM CsA, while incubation with 5μM
CsA led to a significant decrease in proportion of viable HL-60 cells (B). Graphs show the mean of three
independent experiments and error bars show the SEM. *p < 0.05, ***p < 0.001 and ****p < 0.0001 with respect
to 5μM CsA untreated cells.
118
4.2.10 Inhibition of RIP1 modifies the cytotoxicity of HXR9
Activation of RIP1 is a key event in necroptosis (Holler, Zaru et al. 2000). To assess
whether HXR9 causes necroptosis through activation of RIP1, the effect of RIP1 inhibition by
its specific inhibitor Nec-1 on HXR9 cytotoxicity was evaluated using the annexin V assay.
Nec-1 has been reported to inhibit H2O2-induced cell death; therefore H2O2 was used as a
positive control for this assay (Hanus, Zhang et al. 2013). Treatment with 50mM Nec-1 rescued
cells from 20mM H2O2 mediated cell killing. Nec-1 pre- and co-incubation with 20mM H2O2-
treated K562 cells significantly increased the percentage of viable cells from 34.5% ± 3.4 to
67.1% ± 2.3 and decreased the percentage of early apoptotic and dead cells from 41.3% ± 2.5
to 27.2% ± 1.4 and from 24.6% ± 3.6 to 7.3% ± 0.25, respectively, compared to Nec-1untreated
cells (Figure 4.10A). In HXR9-treated K562 cells, there was no significant impact of inhibition
of RIP1 on the cytotoxicity of increasing concentrations of HXR9 (Figure 4.10A). However,
Nec-1 at 50μM significantly changed the cytotoxicity of all concentrations of HXR9 on HL-60.
For example, at 40µM HXR9 treatment, Nec-1 significantly increased the percentage of viable
cells from 3.7% ± 1.21 to 9.2% ± 1.73 and dramatically delayed cell death by increasing the
percentage of early apoptotic cells from 16.3% ± 1.2 to 46.4% ± 2.4 and decreasing the
percentage of dead cells from 80% ± 1.1 to 44.3% ± 2.4, compared to Nec-1 untreated cells
(Figure 4.10B). Therefore, it can be concluded that the inhibition RIP1 has an impact on the
cytotoxicity of HXR9 and this effect depends on the cell type.
119
A)
B)
Figure 4.11 Effect of RIP1 inhibition on HXR9 cytotoxicity. Cells were pre-treated with or without 50μM Nec-1
for one hour, and then with increasing concentrations of HXR9, with an equivalent CXR9 concentration to the
highest concentration of HXR9, 20mM H2O2 (positive control), or were untreated (negative control), in media
supplemented either with or without 50μM Nec-1 for two hours. Cells were then harvested and stained with
annexin V and 7-AAD for 15 minutes in the dark and analysed by flow cytometry. Treatment with 50μM Nec-1
did not affect the sensitivity of K562 cells (A) to HXR9, while for HL60 (B) it significantly increased the
percentage of viable and early apoptotic cells and decreased the percentage of dead cells. Graphs show the mean
of three independent experiments and error bars show the SEM. *p < 0.05, **p < 0.01, ***p < 0.001 and ****p <
0.0001 with respect to 50μM Nec-1 untreated cells.
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4.2.11 HXR9 induces cell death through an ATP-independent pathway
High levels of fructose have been found to cause ATP depletion (Nieminen, Saylor et
al. 1994; Latta, Künstle et al. 2000). ATP is necessary for PARP1-dependent necrosis and
caspase-dependent apoptosis (Los, Mozoluk et al. 2002; Latta, Künstle et al. 2007). To analyse
whether ATP has a role in HXR9-induced AML cell death, the impact of ATP depletion by a
high concentration of fructose on the cytotoxicity of HXR9 was measured by the LDH assay
using an enzyme-linked immunosorbent assay (ELISA). ATP depletion did not reduce the
cytotoxicity of HXR9 on K562 and HL-60 (Figure 4.11). These results were consistent with a
caspase-independent and PARP1-independent pathway.
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A)
B)
Figure 4.12 The effect of ATP depletion on HXR9 cytotoxicity. Cells were pre-incubated with or without
40mM fructose for one hour, and then with a range of HXR9 concentrations (2.5μM – 50μM) for two hours
prepared in media either with or without 40mM fructose, red and blue curves, respectively. HXR9 cytotoxicity
was then measured by assessing LDH enzyme activity in cell-free supernatants. ATP depletion did not inhibit
HXR9-induced death in either (A) K562 or (B) HL-60 cells. Graphs show the mean of three independent
experiments and error bars show the SEM.
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4.2.12 HXR9 induces cell death through a PARP1-independent pathway
The activation of PARP-1 results in ATP depletion, which is a key event in necrotic cell
death (Bouchard, Rouleau et al. 2003; Kim, Zhang et al. 2005). To assess the involvement of
PARP1 in HXR9 induction of AML cell death, the expression of PARP1 protein was estimated
by WB assay. By comparing the band density of treated cells to their untreated counterparts, it
can be concluded that HXR9 did not cause PARP1 activation (Figure 4.12). These findings
concur with those of the ATP depletion assay which suggests that HXR9 treatment results in
cell death independently of ATP depletion.
Figure 4.13 WB analysis of PARP1 activation after HXR9 treatment. HL-60 and K562 cells were treated with
increasing concentrations of HXR9 for two hours and then PARP1 protein expression was examined by WB. β-
actin protein was used as a loading control. There was no difference in band densities between treated and treated
cells after a two hour-treatment with HXR9.
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4.2.13 The effect of inhibition of MAPK pathways on HXR9 cytotoxicity
The results presented earlier in this chapter showed that chelating extracellular Ca2+ by
EDTA prevented HXR9 cytotoxicity. Potential downstream effectors of Ca2+-mediating
necrosis are MAPK pathway members and therefore the effect of HXR9 treatment on three key
components of this pathway, MEK/ERK, p38 and JNK was evaluated (Dhillon, Hagan et al.
2007).
4.2.13.1 HXR9 induces cell death in a MEK/ERK independent pathway
Ca2+ enhances the activation of the MEK/ERK pathway (Egea, Espinet et al. 1999;
Huang, Maher et al. 1999; Lallemend, Lefebvre et al. 2003). To test whether the MEK/ERK
pathway has a role in HXR9 cytotoxicity, this pathway was blocked by a specific inhibitor,
U0126, and the cytotoxicity of HXR9 was examined using the LDH assay. H2O2 that causes
cell death through activation of the MEK/ERK pathway was used as a positive control (Chung,
Jeong et al. 2002; Ray, Huang et al. 2012). Treatment with 20μM U0126 significantly inhibited
the cytotoxicity of H2O2 (Appendix 5). However, U0126 could not attenuate the cytotoxicity of
HXR9 on K562 or HL-60 cells (Figure 4.13). These findings suggest that MEK/ERK pathway
is not required for HXR9-mediated cell death of these cells.
124
A)
B)
Figure 4.14 The effect of inhibiting the MEK/ERK pathway on the cytotoxicity of HXR9. Cells were pre-treated
with or without 20μM of the MEK/ERK inhibitor U0126 for one hour, and then with increasing concentrations of
HXR9 prepared in media with or without 20μM U0126, red and blue curves, respectively. The proportion of
surviving cells was then measured using the LDH assay. Pre- and co-treatment of cells with 20μM U0126 did not
prevent the cytotoxicity of HXR9 in (A) K562 or (B) HL60 cells. Graphs show the mean of three independent
experiments and error bars show the SEM.
125
4.2.13.2 HXR9-mediated cell death does not require p38 pathway signalling
Ca2+ positively regulates the p38 pathway (Huang, Maher et al. 1999; Hsu, Huang et al.
2007). To evaluate the role of the p38 pathway in the HXR9-mediated cell killing, its specific
inhibitor SB302580 was used and the cytotoxicity of HXR9 measured by LDH assay. The p38
pathway is involved in H2O2 cytotoxicity (Pinkus, Weiner et al. 1996; Tobiume, Matsuzawa et
al. 2001; Yoon, Kim et al. 2001; Carvalho, Evelson et al. 2004), and therefore H2O2 was used
as a positive control for this assay. Blocking p38 remarkably inhibited the cytotoxicity of H2O2
(Appendix 6), but not HXR9 on the AML cell lines (Figure 4.14).
126
A)
B)
Figure 4.15 The effect of inhibiting the p38 pathway on the cytotoxicity of HXR9. Cells were pre-treated with or
without 25μM of the p38 inhibitor SB302580 for one hour, and then with a titration of HXR9 prepared in media
with or without 25μM SB302580, red and blue curves, respectively. The cytotoxicity of HXR9 was analysed by
measuring LDH enzyme activity in cell-free supernatants. Pre- and co-treatment of cells with 25μM SB302580
did not affect the cytotoxicity of HXR9 in either (A) K562 or (B) HL-60 cells. Statistical significance was tested
using the student's t-test. Graphs show the mean of three independent experiments and error bars show the SEM.
127
4.2.13.3 Inhibition of the JNK pathway sensitises K562, but not HL-60 cells to HXR9
Like the MEK/ERK and p38 pathways, the JNK pathway is activated by Ca2+ signalling
(Huang, Maher et al. 1999; Son, Lee et al. 2010; Jan, Su et al. 2013). To test the involvement
of JNK pathway for HXR9-induced cell death, it was inhibited using the specific inhibitor
SP600125. Due its ability to induce cell death through activation of the JNK pathway, H2O2
was used as a positive control for this assay (Pinkus, Weiner et al. 1996; Tobiume, Matsuzawa
et al. 2001; Yoon, Kim et al. 2001; Carvalho, Evelson et al. 2004; Kadowaki, Nishitoh et al.
2005). SP600125 at 60μM efficiently abrogated the cytotoxicity of H2O2 (Appendix 7).
Interestingly, inhibiting the JNK pathway resulted in inconsistent results in terms of HXR9
cytotoxicity, significantly sensitizing K562 cells, but not HL-60 cells to killing by HXR9.
Blocking the JNK pathway sharply decreased the percentage of surviving K562 cells from 56%
± 1.7 to 35.4 ± 2.5, ***p < 0.001, after 20μM HXR9 treatment (Figure 4.15).
128
A)
B)
Figure 4.16 The effect of inhibiting the JNK pathway on HXR9 cytotoxicity. Cells were pre-incubated in media
supplemented with or without 60μM of the JNK inhibitor SP600125 for one hour, and then treated with increasing
concentrations of HXR9 prepared in media supplemented with or without 60μM SP600125, red and blue curves,
respectively. The percentage of surviving cells was measured using the LDH assay. Graphs show the mean of three
independent experiments and error bars show the SEM. *p < 0.05, **p < 0.01 and ***p < 0.001 with respect to
60μM SP600125 untreated cells.
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4.2.14 Blocking NOX enzymes sensitizes cells to HXR9
Previous studies have reported that members of NOX enzyme family such as NOX1
and NOX5 are regulated by Ca2+ (Bánfi, Molnár et al. 2001; Bánfi, Tirone et al. 2004; Brueckl,
Kaestle et al. 2006; Valencia and Kochevar 2007). To test whether NOXs have a role in HXR9
mediated cell killing, they were inhibited using DPI. Inhibition of NOX activity by DPI
remarkably sensitised both K562 and HL-60 cells to HXR9. For example, after 20μM HXR9
treatment, the proportion of surviving K562 and HL-60 cells decreased from 57.9% ± 3.54 to
32.6% ± 0.21, *p < 0.05, and from 55.2% ± 1.9 to 26.6 % ± 2.7, *p < 0.05, respectively,
compared to DPI untreated cells (Figure 4.16). These results show that NOX enzymes exert an
anti-necrotic effect in HXR9 treatment.
130
A)
B)
Figure 4.17 The effect of NOX inhibition on the cytotoxicity of HXR9. Cells were pre-treated with or without
60μM of the NOX inhibitor, DPI, for one hour, and then with a range of HXR9 concentrations (2.5μM – 50μM)
prepared in media with or without 60μM DPI, red and blue curves, respectively. Next, the cytotoxicity of HXR9
was assessed using the LDH assay. NOX inhibition significantly sensitised both (A) K562 and (B) HL-60 cells to
HXR9. Graphs show the mean of three independent experiments and error bars show the SEM. *p < 0.05 and **p
< 0.01 with respect to 60μM DPI untreated cells.
131
4.2.15 μ-Calpain is not required for HXR9-induced cell death
μ-Calpain or calpain type I is a downstream effector of Ca2+ (Mathiasen, Sergeev et al.
2002). To analyse whether HXR9 caused AML cell death through the activation of μ-Calpain,
its activity was blocked by the specific inhibitor calpain inhibitor I and assessed using the LDH
assay. Due to its ability to activate μ-Calpain, H2O2 was used as a positive control for this assay
(McClung, Judge et al. 2009; Pallepati and Averill-Bates 2011). Calpain inhibitor I significantly
inhibited the cytotoxicity of H2O2 (Appendix 8). However, calpain inhibitor I at 60μM could
not abrogate the cytotoxicity of HXR9 on tested AML cell lines, K562 and HL-60 (Figure 4.17).
132
A)
B)
Figure 4.18 The effect of blocking μ-calpain on HXR9 cytotoxicity. Cells were pre-treated with or without 60μM
calpain inhibitor I for one hour, and then for two hours with increasing concentrations of HXR9 prepared in media
with or without 60μM calpain inhibitor I, red and blue curves, respectively. Next, the cytotoxicity of HXR9 was
assessed in cell-free supernatants using the LDH. μ-calpain inhibition did not affect HXR9 cytotoxicity in either
(A) K562 or (B) HL-60 cells. Graphs show the mean of three independent experiments and error bars show the
SEM.
133
4.2.16 CaM inhibition dramatically increases the sensitivity of cells to HXR9
CaM is one of the main Ca2+ concentration sensor (Balshaw, Xu et al. 2001; Hoeflich
and Ikura 2002; Yuan, Jing et al. 2011). To assess the involvement of CaM in the cytotoxicity
of HXR9, its role was blocked by the specific inhibitor W-7. Interestingly, treating cells with
50μM W-7 alone did not affect cells, yet co-treatment of cells with HXR9 significantly reduced
cell survival. Co-treatment of K562 and HL-60 cells with 20μM HXR9 and 50μM W-7 sharply
decreased the percentage of surviving cells from 57% ± 1.6 to 15% ± 1.8, ****p < 0.0001, and
from 61% ± 2.4 to 26% ± 2.1, ***p < 0.001, respectively (Figure 4.18). It also reduced the IC50
of HXR9 on K562 cells by threefold, from 21μM to 7μM, and on HL-60 cells by twofold, from
24μM to 13μM (Figure 4.18C).
134
A)
B)
C)
IC50
Cell line/condition HXR9 HXR9 + 50μM W-7
K562 21.4μM ± 3.1 7μM ± 2.9
HL-60 24μM ± 1.7 13μM ± 1.2
Figure 4.19 The effect of CaM inhibition on HXR9 sensitivity. Cells were pre-treated with or without 50μM
W-7 for one hour, and then for two hours with increasing concentrations of HXR9 prepared in media with
or without 50μM W-7, red and blue curves, respectively. The LDH assay was used to measure cytotoxicity.
W-7 dramatically increased the sensitivity to HXR9 of both (A) K562 cells and (B) HL-60 cells. Graphs
show the mean of three independent experiments and error bars show the SEM. C) A table shows the
difference in HXR9 IC50 between 50μM W-7-treated and untreated cells. *p < 0.05, **p < 0.01, ***p < 0.001
and ****p < 0.0001 with respect to 50μM W-7 untreated cells.
135
4.2.17 Inhibition of PKC greatly increases the sensitivity of K562 and HL-60 cells to
HXR9
PKC is a family of kinases that are regulated by intracellular Ca2+ (Zhu, Fang et al. 1999;
Reyland 2007). To analyse the impact of blocking PKC on HXR9-induced AML cell death, the
specific inhibitor Ro31-8220 was used and the cytotoxicity of HXR9 was assessed with the
LDH assay. 30μM Ro31-8220 was not cytotoxic for AML cells, yet it significantly increased
HXR9-mediated cell killing. In K562 cells, Ro31-8220 remarkably sensitized cells to a very
low concentration of HXR9, 1.25μM, *p < 0.05 (Figure 4.19A). In addition, it caused a huge
reduction in the IC50 of HXR9, by 4.3 fold, from 21.4μM ± 1.5 in K562-HXR9 only treated
cells to 5μM ± 2.9 (Figure 4.19C). Similarly, inhibition of PKC in HL-60 increased cell killing
by HXR9 and decreased the IC50 by 2.7 fold from 29.3μM ± 1.9 to 10.7μM ± 0.8 (Figure 4.19B
and C). These results showed that PKC activity protects against HXR9 cytotoxicity.
136
A)
B)
C)
Figure 4.20 The effect of blocking of PKC on the cytotoxicity of HXR9 on K562 and HL-60 cells. Cells were pre-
treated with or without 30μM Ro31-8220 for one hour, and then for two hours with different concentrations of
HXR9 in media supplemented with or without 30μM Ro31-8220, red and blue curves, respectively. The LDH
assay was then used to measure the cytotoxicity of HXR9 in cell-free media. Ro31-8220 co-treatment significantly
reduced cell survival. Graphs show the mean of three independent experiments for (A) K562 cells and (B) HL-60
cells and error bars show the SEM. *p < 0.05, **p < 0.01, ***p < 0.001 and ****p < 0.0001 with respect to 30μM
Ro31-8220 untreated cells. C) Differences in the IC50 values between HXR9-only treated cells and HXR9 + 30μM
Ro31-8220 treated cells.
IC50
Cell line/condition HXR9 HXR9 + 30μM Ro31-8220
K562 21.4μM ± 1.5 5μM ± 2.9
HL-60 29.3μM ± 1.9 10.7μM ± 0.8
137
4.2.18 Simultaneous inhibition of CaM and PKC potentiates HXR9 cytotoxicity
To test whether simultaneous inhibition of CaM and PKC produced a further
sensitisation to HXR9, K562 and HL-60 were exposed to the inhibitors of both proteins and
treated with a titration of HXR9 and the cytotoxicity was measured using the LDH assay. The
concurrent regimen significantly increased the sensitivity to HXR9. For example, at 10μM
HXR9 treatment, the concurrent inhibition of both proteins dramatically decreased the
percentage of surviving cells by 55%, ****p < 0.0001, and 57%, ****p < 0.0001, for K562 and
HL-60, respectively, comparing with HXR9-only treated cells (Figure 4.20A and B). In
addition, the IC50 of HXR9 was reduced by 4.3 and 3.1 fold for K562 and HL-60, respectively
(Figure 4.20C).
138
A)
B)
C)
IC50
Cell line/condition HXR9 HXR9 + (50μM W-7 30μM Ro31-8220)
K562 21.1μM ± 4.5 4.9μM ± 3.5
HL-60 14.8μM ± 3.9 4.8μM ± 2.7
Figure 4.21 The impact of concurrent inhibition of CaM and PKC on the cytotoxicity of HXR9. Cells were pre-
incubated with or without 50μM W-7 and 30μM Ro31-8220 for one hour, and then for two hours with a range of
HXR9 concentrations (0.625μM – 50μM) prepared in media with or without the inhibitors, red and blue curves,
respectively. The cytotoxicity of HXR9 was assessed using the LDH assay. The concurrent inhibition of CaM and
PKC significantly increased the HXR9-mediated cell death. Graphs show the mean of three independent
experiments with (A) K562 cells and (B) HL-60 cells, and error bars show the SEM. **p < 0.01, ***p < 0.001 and
****p < 0.0001 with respect to W-7 and Ro31-8220 untreated cells. C) The difference in the IC50s between HXR9-
only treated cells and HXR9 + (50μM W-7 + 30μM Ro31-8220) treated cells.
139
4.2.19 HO-1 inhibition increases the sensitivity to HXR9
HO-1 acts as an anti-apoptotic protein in AML (Rushworth and MacEwan 2008). In
addition, HO-1 is a downstream effector of NOXs and PKC (Tsoyi, Jang et al. 2011; Schroder,
Zhang et al. 2012; Lee, Yang et al. 2012; Nguyen, Kim et al. 2013). To assess whether the
inhibition of HO-1 could increase the sensitivity to HXR9, HO-1 activity was blocked by its
specific inhibitor PPIX and the sensitivity of AML cells to HXR9 assessed. HO-1 inhibition
resulted in a significant reduction in the survival of both cell lines (Figure 4.21). PPIX decreased
the IC50 of HXR9 by 2.2 fold and 1.4 fold for K562 and HL-60, respectively (Figure 4.21C).
Therefore, HO-1 activity inhibits HXR9-mediated cell killing.
140
A)
B)
C)
IC50
Cell line/condition HXR9 HXR9 + 50μM PPIX
K562 23.7μM ± 3.4 10.9μM ± 4.2
HL-60 22.5μM ± 5.1 16μM ± 1.4
Figure 4.22 The effect of HO-1 inhibition on the cytotoxicity of HXR9. Cells were pre-treated with or without
50μM PPIX for one hour and then for two hours with HXR9 in media with or without 50μM PPIX, red and blue
curves, respectively. Cell death was assessed using the LDH assay. Graphs show the mean of three independent
experiments for (A) K562 and (B) HL-60 cells, and error bars show the SEM. *p < 0.05, **p < 0.01, ***p < 0.001
and ****p < 0.0001 with respect to 50μM PPIX untreated cells. C) Differences in the IC50s between HXR9-only
treated cells and HXR9 + 50μM PPIX treated cells.
141
4.2.20 HXR9 induces p21 expression but not p53
To analyse whether HXR9 causes up-regulation of p53 and its downstream effector p21,
the expression of both genes was analysed by RT-PCR. Results showed that HXR9 did not
affect the expression of p53 in K562 or HL-60 cells, but it did lead to a significant increase in
the expression of p21. For example, the expression of p21 in K562 cells increased by 2.5, **p
< 0.01, and 4.5, ****p < 0.0001, fold after treatment with 15μM and 30μM of HXR9,
respectively (Figure 4.22B).
142
A)
B)
Figure 4.23 RT-PCR analysis of p53 and p21 expression in response to HXR9 treatment. RNA extracted from
cells after a two-hour treatment with 15μM and 30μM HXR9, or 30μM CXR9. The expression of p53 and p21
genes are given as ratio to the expression of the housekeeping gene β-actin (×10 000). Graphs show the mean of
three independent experiments and error bars show the SEM. **p < 0.01, ***p < 0.001 and ****p < 0.0001 with
respect to untreated cells.
143
4.2.21 HXR9-induces AML cell death through a p53-independent pathway
To further investigate the role of p53 in HXR9-mediated cell killing, the activity of p53
protein was blocked by PFT-α, and the cytotoxicity of HXR9 was measured by LDH assay.
Blocking p53 protein did not affect the sensitivity of AML cells to HXR9 (Figure 4.23).
A)
B)
Figure 4.24 The effect of blocking p53 protein on HXR9 cytotoxicity. Cells were pre-treated with or without
40μM PFT-α for one hour, and then for two hours with a titration of HXR9 prepared in media supplemented with
or without 40μM PFT-α, red and blue curves, respectively. Graphs show the mean of three independent
experiments for (A) K562 and (B) HL-60 cells, and error bars show the SEM.
144
4.3 Discussion
HXR9 has previously been shown to induce cell death in a range of solid malignancies,
and here it is demonstrated that HXR9 also induces AML cell death. The efficacy of HXR9
could be improved by better understanding its mechanism of action, allowing synergistic
therapies to be identified.
Antagonism of the HOX/PBX interaction resulted in a remarkable up-regulation in the
expression of c-FOS. The induction of c-FOS expression by HXR9 was previously documented
to be responsible, at least partially, for triggering cell death in human melanoma, renal, lung,
ovarian, breast and prostate cancer cell lines (Morgan, Pirard et al. 2007; Shears, Plowright et
al. 2008; Plowright, Harrington et al. 2009; Morgan, Plowright et al. 2010; Morgan, Boxall et
al. 2012; Morgan, Boxall et al. 2014), and knocking down c-FOS expression partially rescued
melanoma and prostate cancer cell lines from HXR9-induced apoptosis (Morgan, Pirard et al.
2007; Morgan, Boxall et al. 2014). c-FOS is a component of the AP-1 transcription activating
complex, and upon dimerisation with its binding partner JUN, regulates cell proliferation and
cell cycle progression (Durchdewald, Angel et al. 2009). It has been shown to be a proto-
oncogene in a variety of human tumours including those of the bone, brain and skin (Gamberi,
Benassi et al. 1998; Silvestre, Gil et al. 2010; Guinea-Viniegra, Zenz et al. 2012). However, a
reduction in c-FOS expression in epithelial ovarian carcinoma was found to be an independent
marker of shorter progression-free and OS (Mahner, Baasch et al. 2008). In addition, c-FOS
over-expression has a pro-apoptotic function in vitro, and caused a delay in tumour growth and
metastasis of the ovarian cancer OvCa cells in vivo by dysregulating adhesion proteins
(Oliveira-Ferrer, Rossler et al. 2014). It was also found to be involved in the induction of the
human ovarian carcinoma cell line A2780 in response to fenrtinide treatment (Appierto, Villani
et al. 2003). Moreover, c-FOS exerted a pro-apoptotic function by repressing the anti-apoptotic
gene cellular FLICE (FADD-like IL-1β-converting enzyme)-inhibitory protein (L) (c-FLIP (L))
145
which resulted in sensitising prostate cells to tumour necrosis-related apoptosis-inducing ligand
(TRAIL) (Zhang, Zhang et al. 2007). It also induced human hepatoma Huh7 apoptosis by
mediating c-MYC expression (Kalra and Kumar 2004). c-Fos over-expression resulted in cell
cycle arrest by up-regulating the expression of the nuclear protein CHOP and the cell cycle
inhibitors p16INK4A and p57KIP2; suppressing rat sarcoma (Ras)-mediated malignancy that
ultimately led to apoptosis of murine epithelial hepatocytes (Mikula, Gotzmann et al. 2003). Its
prolonged expression also suppressed cell cycle entry of murine BM cells (Okada, Fukuda et
al. 1999). Furthermore, c-Fos enhanced hepatocyte apoptosis by directly up-regulating the
major isoform of bim gene BimEL (Kitamura, Ogawa et al. 2003).
Externalisation of PS was seen in all tested AML cell lines after HXR9 treatment. This
finding was consistent with previous studies in different cancer cell lines upon HXR9 treatment
(Morgan, Pirard et al. 2007; Shears, Plowright et al. 2008; Plowright, Harrington et al. 2009;
Daniels, Neacato et al. 2010; Morgan, Plowright et al. 2010; Morgan, Boxall et al. 2012;
Morgan, Boxall et al. 2014). In viable cells, PS is localised on the inner leaflet of the cell surface
and its transfer to the outer leaflet is thought to be an engulfment signal by macrophages (Fink
and Cookson 2005). It is well-known that the exposure of PS to the outer leaflet of the cell
surface is a distinctive feature for apoptosis and results from the activation of caspase-3
(Mandal, Moitra et al. 2002; Bonomini, Dottori et al. 2004). Therefore, the binding-ability of
annexin V to PS was used specifically to detect and quantify apoptotic cells. However, the outer
leaflet exposure of PS before the damage of the plasma membrane was reported in thymocytes
at the prelethal stage of necrosis, oncosis (Waring, Lambert et al. 1999). In addition, oncotic
L1210 cells, a mouse leukaemia cell line, showed annexin V+ /PI- phenotype after serum amine
oxidase exposure (Bonneau and Poulin 2000), and exposure of Jurkat cells to different oncotic
stimuli led to externalisation of PS, annexin V+ /7-ADD-, to the outer leaflet before the loss of
cell membrane integrity (Lecoeur, Prevost et al. 2001; Lecoeur, de Oliveira-Pinto et al. 2002).
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Likewise, caspase-8 deficient and Bcl-2 over-expressing Jurkat cells exhibited PS exposure
during the oncotic stage (Krysko, de Ridder et al. 2004). H2O2 also was reported to cause PS
externalisation in a cypD-dependent pathway, i.e. mitochondrial necrosis (Jobe, Wilson et al.
2008). Therefore, externalisation of PS does not actually differentiate between apoptosis and
necrosis (Fink and Cookson 2005; Verheij 2008; Blankenberg and Strauss 2012).
The hallmark of apoptosis is caspase activation, and this was not found to be necessary
in HXR9-induced cell death. These data are consistent with the likely mechanism of HXR9-
induced cell death in the renal cancer cell lines Caki-2 and 769-P, in malignant B cell lines and
some prostate cell lines (Shears, Plowright et al. 2008; Daniels, Neacato et al. 2010; Morgan,
Boxall et al. 2014). However, the general inhibition of caspases decreased the cytotoxicity of
HXR9 on the tested renal, ovarian, breast and prostate cancer cell lines, although this decrease
did not completely abrogate the effect of HXR9 (Shears, Plowright et al. 2008; Morgan,
Plowright et al. 2010; Morgan, Boxall et al. 2014). This may be due to the presence of other
mediators of HXR9-induced cell death. The results of this study support this. For example,
HXR9 treatment of AML cells did not change the expression of PARP1 at the RNA or protein
level. Nuclear fragmentation of apoptotic cells results from the cleaving of the cytoskeletal
protein lamin by caspase-3 or -6 (Kagawa, Go et al. 2001; Ruchaud, Korfali et al. 2002), which
might explain the absence of nuclear fragmentation in HXR9 treated AML cell lines. In
addition, the expression of the anti-apoptotic genes Bcl-2 and XIAP that were known as targets
of active caspases (Jin and El-Deiry 2005; Vermeulen, Van Bockstaele et al. 2005) were not
affected by HXR9 treatment.
The findings of this study suggest that mitochondrial apoptosis is not required for
HXR9-mediated cell death. For example, an unchanged ratio between the anti-apoptotic Bcl-2
and the pro-apoptotic Bax and Bak1 may reflect the intactness of mitochondrial membranes,
since the presence of excess pro-apoptotic members of the Bcl-2 family enhances mitochondrial
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permeabilization (Martin 2010). Furthermore, the mitochondrial release of cyt C would
ultimately lead to caspase activation. In addition, the cleavage and activation of Bid that
mediates mitochondrial apoptosis depends on the activation of caspase-8, which was not
required for HXR9 cytotoxicity. Likewise, HXR9 seemed to induce cell death in a Bax-
independent process, since Bax-dependent apoptosis involves either caspase-9 or p53 activity,
neither of which was needed for HXR9 mediated-cell killing (Janssen, Pohlmann et al. 2007;
Gogada, Prabhu et al. 2011; Meng, Zhang et al. 2012).
The absence of apoptotic-hallmarks activation, including caspase activation, Bcl-2
family involvement, chromatin condensation and nuclear fragmentation suggest that HXR9
induces AML cell death through a necrotic pathway.
EDTA is a general cation-chelating agent, despite being widely used as an extracellular
Ca2+ chelator (Dong, Li et al. 2005; Allen, Gemel et al. 2011; de Araújo Leite and Marques-
Santos 2012). For example, EDTA can form complexes with cations such as Na+, Mg+, Fe2+,
Cu2+, Zn2+ that can be easily removed from the extracellular environment (Flora and Pachauri
2010). Interestingly, the cell penetrating peptide (CPP) of HXR9 is R9 that is a highly cationic
sequence (Mitchell, Steinman et al. 2000; Sarko, Beijer et al. 2010; Regberg, Srimanee et al.
2012; Bechara and Sagan 2013). In addition, the binding of annexin V to PS is a Ca2+ dependent
reaction (Vermes, Haanen et al. 1995). Therefore, the abrogation of HXR9 effect by EDTA
might be due to the chelating of HXR9 and the absence of Ca2+ in annexin V/7-AAD assay.
CypD knockout studies have demonstrated its crucial role in mitochondrial necrosis by
Ca2+ overload and oxidative stress, but is not required for apoptosis (Li, Johnson et al. 2004;
Baines, Kaiser et al. 2005; Nakagawa, Shimizu et al. 2005; Schinzel, Takeuchi et al. 2005). In
addition, CypD is involved in Ca2+-independent, RIP3- and p53-dependent mPTP necrosis
(Lerch, Halangk et al. 2013; Tian, Xu et al. 2013; Pei, Shang et al. 2014). Accordingly, HXR9
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might mediate necrosis of AML in an mPTP-independent pathway, or at least, in a CypD-
independent mechanism.
In this study it was shown that HXR9 led to cell death through a necroptotic pathway,
at least in part, through RIP1 activity in HL-60 cells, whilst K562 cell death was RIP1-
independent. It is possible though that the effect of blocking RIP1 HXR9-induced HL-60 cell
death could reflect a sub-effective dose of the RIP1 inhibitor, as Nec-1 showed some toxic
effects at higher doses. The latter explanation is consistent with the fact that Nec-1 partially
protected HL-60 cells from HXR9. Alternatively, other mediators in addition to RIP1 may be
required in HXR9-induced cell death. It is noteworthy that a RIP1-independent cell death
pathway does not rule out a potential role for necroptosis since it was reported that viral
infection resulted in RIP3-dependent but RIP1-independent necroptosis through TNF mediated
necroptosis (Zhang, Shao et al. 2009; Upton, Kaiser et al. 2010). Therefore, the role of RIP3 in
HXR9-induced K562 cell death requires further investigation.
The presence of ATP is crucial for caspase-dependent apoptosis and PARP1 hyper-
activation. In caspase-dependent apoptosis, ATP is involved in the formation of the apoptosome
complex that mediates apoptotic signalling by cleaving and activating executioner caspases
(Jochen, Richter et al. 2002). It was reported that ATP depletion by fructose in primary rat and
mouse hepatocytes abrogated cyt C release and activation of executioner caspases, and DNA
fragmentation by TNF-mediated apoptosis, and that these events were restored after ATP
repletion (Latta, Kunstle et al. 2000; Jochen, Richter et al. 2002). In addition, the presence of
ATP is essential for chromatin condensation and nuclear fragmentation (Tsujimoto 1997). ATP
depletion by fructose prevented caspase activation and subsequently shifted cell death toward
necrosis in the leukaemic Jurkat and K562 cells (Leist, Single et al. 1999; Verrax, Dejeans et
al. 2011). Likewise, PARP1 involves ATP in the transfer of poly (ADP-ribose) from NAD+
(Devalaraja-Narashimha and Padanilam 2009; Ethier, Tardif et al. 2012). Taking together, the
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depletion of ATP elicits the role of caspases and PARP1 hyper-activation in the mechanism of
HXR9-induced cell death of AML cells. It is well-known that fructose at high concentrations
traps intracellular ATP and maintains a sufficient cytoplasmic concentration to prevent cell
necrosis (Latta, Künstle et al. 2000; Latta, Künstle et al. 2007; Speicher, Köhler et al. 2012).
The results of this study revealed that ATP depletion by a high concentration of fructose did
not prevent HXR9-induced necrotic death of the tested AML cells, which also supports a
caspase- and PARP1- independent mechanism for HXR9-induced cell death.
Despite its well-known crucial role in TNF-induced necrosis, PARP1 was dispensable
in TNF-mediating necroptosis (Xu, Huang et al. 2006). Accordingly, it can be concluded that
TNF activates necroptosis and necrosis in distinct pathways, although PARP1 was also shown
to be a downstream effector of the necrosome in TRAIL-induced necroptosis (Jouan-Lanhouet,
Arshad et al. 2012). In this study, it was reported that HXR9-induced necrosis in a PARP1-
independent mechanism. While the JNK pathway is involved in PARP1-induced necrosis,
HXR9-induces necrosis of AML cells in JNK-independent pathway (Degterev, Huang et al.
2005; Xu, Huang et al. 2006; Jog, Dinnall et al. 2009; Ethier, Tardif et al. 2012; Sosna, Voigt
et al. 2014). Since PARP1 is a crucial player in mPTP, the results of PARP1 activation in this
study support a mitochondria-independent pathway for HXR9-induced necrosis (Abramov and
Duchen 2008).
The MAPK family of protein kinases consists of four members; ERK, p38, JNK and the
big mitogen activated protein kinase. MAPK proteins regulate different cellular pathways
including cell survival, proliferation and cell death (Chen and Sommer 2009). In this study
inhibition of MEK/ERK and p38 did not affect HXR9-induced cell necrosis. JNK inhibition
also did not affect the efficacy of HXR9 in HL-60, although it increased the cytotoxicity of
HXR9 for K562. This effect of JNK inhibition might be due to the expression of the BCR-ABL
oncoprotein in K562 cells that is reported to induce leukaemogenesis through activation of JNK
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signalling pathway (Hess, Pihan et al. 2002; Hantschel, Warsch et al. 2012; Chen, Gallipoli et
al. 2013). In addition, the JNK pathway is reported to be an inducer of clonal evolution of
Fanconi anaemia to AML (Li, Sejas et al. 2007) and sustained activation of the JNK pathway
with AKT/FOXO signalling maintains AML cells in an undifferentiated state (Sykes, Lane et
al. 2011). In FL15.12 cells, a haemopoietic pro-B cell line, IL-3 promoted cell survival by
phosphorylating BAD through JNK signalling (Yu, Minemoto et al. 2004). More recently, it
was reported that co-inhibition of the TNF-JNK pathway increased the sensitivity of primary
human AML cells in vitro to NF-ĸB inhibitors (Volk, Li et al. 2014), and that the JNK pathway
promoted proliferation and metastasis in several cancers including those of the liver and skin
(Chen, Nomura et al. 2001; Han, Moon et al. 2009; Bettermann, Vucur et al. 2010; Ke, Harris
et al. 2010; Ebelt, Cantrell et al. 2013).
NOXs are a family of proteins that are considered to be a primary resource of ROS in
living cells and regulated by intracellular Ca2+ signalling (Yu, Kim et al. 2006; Morgan, Kim et
al. 2008; El Jamali, Valente et al. 2010; Hole, Darley et al. 2011; Jiang, Zhang et al. 2011;
Pandey, Gratton et al. 2011) and mediate necrotic cell death (Ouyang, Shi et al. 2012). DPI is
a widely used NOX inhibitor, though it is also a broad inhibitor of other flavoenzymes including
nitric oxide synthase and xanthine oxidase (Li, Ragheb et al. 2003; Park, Song et al. 2007). In
this study DPI increased the efficacy of HXR9 on the tested AML cells, suggesting that NOXs
has survival roles upon HXR9 treatment. In line with these findings, it was found that DPI
mediated the apoptosis of the pancreatic cancer cell line PANC-1 by blocking the survival
pathway NOX4/ROS/AKT (Mochizuki, Furuta et al. 2006). DPI was also shown to mediate
p53-dependent apoptosis (Kim, Song et al. 2007; Park, Song et al. 2007). In addition, NOXs
were found to be regulated by PKC and CaM kinases that were identified in this study as
mediators of cell resistance to HXR9 (Inoguchi, Sonta et al. 2003; Cai, Torreggiani et al. 2010;
Pandey, Gratton et al. 2011).
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Calpains are a family of 14 cysteine proteases that are regulated by Ca2+. The most
extensively studied members of this family are μ-calpain or calpain I that is present in the
cytosol and is stimulated by micro-molar Ca2+ concentrations, and m-calpain or calpain II that
is present in the cell membrane and is activated by mille-molar concentration of Ca2+ (Kar,
Chakraborti et al. 2009; Szydlowska and Tymianski 2010). In this study, the potential role of
μ-calpain was studied because it was reported a mediator of Ca2+-dependent necrosis
(Mathiasen, Sergeev et al. 2002; Zhivotovsky and Orrenius 2011; Ouyang, Shi et al. 2012).
However, HXR9-induced necrosis of AML cells in a μ-calpain-independent manner. This
might be explained by the caspase-independent pathway of HXR9-mediated cell death, as μ-
calpain was shown to cleave and activate pro-caspases-3/9 in human lung adenocarcinoma cells
(Liu, Xing et al. 2009). This finding is also consistent with the suggestion that HXR9-induced
cell death does not require mitochondria, since μ-calpain was found to cause cleavage and
activation of the pro-apoptotic members of Bcl-2 family Bax and Bid (Chen, He et al. 2001;
Mandic, Viktorsson et al. 2002; Toyota, Yanase et al. 2003; Sanchez-Gomez, Alberdi et al.
2011). It has also been shown that μ-calpain is a downstream mediator of PARP1 in the release
of AIF that ultimately leads to mitochondria-dependent necrosis (Cao, Xing et al. 2007; Liu,
Xing et al. 2009; Vosler, Sun et al. 2009; Storr, Carragher et al. 2011).
CaM is a small 17KDa intracellular protein that is considered to be the main regulator
and Ca2+-binding protein in all eukaryotic cells (Wang, Li et al. 2010; Berchtold and Villalobo
2014). In this study, the CaM antagonist W-7 enhanced HXR9-induced necrosis. CaM exerts
surviving effects by positively regulating pro-survival and proliferative proteins, and inhibition
of CaM alone or in combination therapy promotes cell death in many cancer models. CaM
inhibition sensitised human lung cancer H1299 cells to TRAIL-induced apoptosis by enhancing
death-inducing signalling complex (DISC)-triggered apoptosis and/or preventing the
phosphorylation of the survival protein AKT, leading to a reduction in expression of the anti-
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apoptotic protein c-FLIP (Hwang, Min et al. 2009). CaM also regulates the PI3K survival
pathway (Lasbury, Durant et al. 2009), and inhibition of CaM attenuated the ERK1/2-dependent
survival pathway and up-regulated the Bax- and caspase-3-dependent pathway (Bartolomé, de
las Cuevas et al. 2007). CaM can also maintain NF-ĸB activity by phosphorylating and
degrading its binding inhibitor IĸB (Shumway, Berchtold et al. 2002; Tano and Vazquez 2011).
In vitro and in vivo studies of cholangiocarcinoma showed that the inhibition of CaM prevented
the phosphorylation of AKT and c-FLIP recruitment to DISC, thereby increasing the
recruitment and activation of caspase-2, caspase-8 and caspase-9. This in turn leads to cleavage
and activation of caspase-3, releasing cyt C and triggering apoptosis (Ahn, Pan et al. 2003;
Pawar, Ma et al. 2009; Jing, Yuan et al. 2011; Jing, Yuan et al. 2012).
PKCs are a family of protein kinases that regulate several cellular functions including
cell cycle progression, cellular proliferation, differentiation, survival and apoptosis. The PKC
family is composed of 12 members and is classified into four classes according to structural
and regulatory characteristics including the conventional/classical PKCs, the novel PCKs, the
atypical PKCs and PKC-related kinases. Of these, only the conventional PKCs are Ca2+-
dependent (Bosco, Melloni et al. 2011; Shen, Kim et al. 2012; Parker, Justilien et al. 2014). Of
the 12 PKCs, 7 are anti-apoptotic proteins including PKCα, PKCβ, PKCι, PKCγ, PKCζ, PKCε
(Reyland 2007). In this study, the general inhibitor of PKC Ro31-8220 increased the sensitivity
of the tested AML cell lines to HXR9. This supports the findings that NOX inhibition also
sensitized cells to HXR9, as PKCα and PKCβ are positive regulators for NOXs (Shiraki, Oyama
et al. 2012; Paneni, Beckman et al. 2013). Up-regulation of PKCα and PKCι were documented
in breast cancer stem cells, human glioma (Desai, Pillai et al. 2011; Tam, Lu et al. 2013). In
addition, PKC was found to increase the stability of the anti-apoptotic protein c-FLIP (Kaunisto,
Kochin et al. 2009). PKCι was documented to cause cell cycle progression by phosphorylating
CdK7 and CdK2 (Win and Acevedo-Duncan 2009; Pillai, Desai et al. 2011), and in the cancer
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prostate cell line LNCaP, PKC promoted cell survival through the
PI3K/PKD1/PKC/ERK1/2/NF-ҡB signalling pathway (Chen, Giridhary et al. 2011). It was also
reported that PKC enhanced cell survival and invasion by inducing the expression of the growth
factor bone morphogenic protein-4, components of the Ras signalling pathway, and matrix
metalloproteinase 2 and 9 (Hwang, Yun et al. 2011; Shimizu, Kayamori et al. 2012; Lonic,
Powell et al. 2013). During TRAIL-mediated apoptosis, PKC activation was reported to prevent
DISC formation and consequently inhibited the apoptosis of HeLa cells (Harper, Hughes et al.
2003). In ALL, PKC inhibition sensitised cells to Ca2+-mobilizers (Zhu, Fang et al. 1999).
Furthermore, inhibition of PKC resulted in a dephosphorylation and activation of Bad
concurrently with a down-regulation of Bcl-2 and ultimately to mitochondrial dysfunction and
caspase-dependent apoptosis (Win and Acevedo-Duncan 2009; Desai, Pillai et al. 2011;
Whang, Jo et al. 2013).
The concurrent inhibition of CaM and PKCs alongside with HXR9 treatment resulted
in AML cell death at very low IC50 in compared to HXR9-alone. This might result from the
inhibition of both cell survival pathways such as PI3K/AKT, and anti-apoptotic proteins such
as c-FLIP and anti-apoptotic members of Bcl-2 family.
HO-1 protein is an important cytoprotective enzyme that has anti-apoptotic properties
and promotes cancer cell survival, angiogenesis and drug-resistance (Jozkowicz, Was et al.
2007; Lee, Chun et al. 2011; Park, Jung et al. 2011; Lee, Yang et al. 2012). It is constitutively
expressed in primary human AML samples, cell lines and multiple myeloma cell lines (Barrera,
Rushworth et al. 2012; Herrmann, Kneidinger et al. 2012). Inhibiting HO-1 increased the
sensitivity of AML cells to HXR9. This is consistent with a number of other findings. For
example, c-FOS is suppressed by HO-1 (Sun 2008), and PKC promotes cell survival through
two different pathways that both involve HO-1, for example, through PKC/p38/Nrf2/HO-1 or
PKC/ROS/PI3K/AKT/Nrf2/HO-1 (Tsoyi, Jang et al. 2011; Lee, Yang et al. 2012; Nguyen, Kim
154
et al. 2013). In addition, NOX4 was reported to induce HO-1 expression through the ROS/Nrf2
pathway (Schroder, Zhang et al. 2012). HO-1 was also found to reduce the expression of p21
that was found to play a pro-necrotic role following HXR9 treatment in this study (Was,
Sokolowska et al. 2011). In addition, HO-1 prevents TNF-induced apoptosis in both c-FLIP
and NF-ҡB knockdown AML cells (Rushworth and MacEwan 2008; Rushworth, Zaitseva et
al. 2010). HO-1 also protected AML cells from the front-line chemotherapeutic drugs DNR and
cytarabine and its inhibition enhanced the efficacy of both drugs (Heasman, Zaitseva et al.
2011).
p53 is one of the most extensively studied cell death proteins, and stimulates c-FOS
transcription in cells undergoing p53-dependent apoptosis (Elkeles, Juven-Gershon et al. 1999).
The results of this study showed that HXR9 did not change the expression of p53 at the RNA
or at the protein level. These results were consistent with the mechanism of HXR9 killing of
breast cancer cells that is p53-independent (Morgan, Boxall et al. 2012). In addition, p53 protein
directly interacts with the anti-apoptotic members of the Bcl-2 family, Bcl-2 and Bcl-XL,
resulting in the release of the pro-apoptotic members Bak and Bax, leading to caspase-
dependent apoptosis (Mihara, Erster et al. 2003; Vaseva and Moll 2009; Chakraborty,
Mazumdar et al. 2014; Chi 2014). p53 also up-regulates the transcription of, and physically
interacts with Bak, Bax and Bad resulting in mitochondrial apoptosis (Chipuk, Maurer et al.
2003; Chipuk, Kuwana et al. 2004; Leu, Dumont et al. 2004; Jiang, Du et al. 2006; Pietsch, Leu
et al. 2007; Wenzel, Wunderlich et al. 2012; Chi 2014; Lee, Lee et al. 2014; Saha, Bhattacharjee
et al. 2014). However, none of the Bcl-2 family members or Apaf1, a mediator of p53-dependent
apoptosis, changed in expression during HXR9-mediated AML cell necrosis. Furthermore,
calpain, an inducer of p53 apoptosis, was not required for HXR9-induced AML cell death (Del
Bello, Moretti et al. 2007; Bukowska, Lendeckel et al. 2012; Woo, Xue et al. 2012). More
recently, the role of p53 in cypD- and PARP1-dependent programmed necrosis was identified
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(Vaseva, Marchenko et al. 2012; Montero, Dutta et al. 2013; Pei, Shang et al. 2014). However,
neither cypD nor PARP1 were involved in HXR9-induced AML necrosis. Taken together, these
findings suggest that HXR9-indcues AML necrosis in a p53-independent manner.
p21 is a member of the family of cyclin kinase inhibitors that include p27 and p57
(Kwon, Jovanovic et al. 2003; Masgras, Carrera et al. 2012). It was originally recognised as a
main regulator of the progression of the cell cycle (Harper, Adami et al. 1993). However, p21
also regulates a number of other cellular functions, including apoptosis, cell proliferation and
autophagy (Asada, Yamada et al. 1999; Dong, Li et al. 2005; Luo, Zou et al. 2011). Although
it was thought that the regulation of p21 was totally dependent on p53, p21 can in fact be
regulated by several transcription factors in a p53-independent manner (Liu and Huang 2006;
Lafarga, Cuadrado et al. 2009; Shin, Kim et al. 2011; Han, Kim et al. 2012; Masgras, Carrera
et al. 2012). For example, the early growth response-1 protein can induce the transcription of
p21 independently of p53 (Choi, Kim et al. 2008). The early growth response-1 gene is induced
in lung cancer cells after HXR9 treatment (Plowright, Harrington et al. 2009). Interestingly,
p21 has been shown to be involved in p53-independent necrotic cell death (Ussat, Werner et al.
2002; Kwon, Jovanovic et al. 2003). The results of this study show that the induction of p21 in
response to HXR9 was p53-independent.
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4.3.1 Summary of chapter
In the summary, HXR9-mediated cell death in AML cells was characterised by the
absence of the well-known features of apoptosis including caspase activation, the up-regulation
of Bcl-2 family-members, chromatin condensation and nuclear fragmentation. The mechanism
was also p53-independent. However, the mechanism of cell death involved, at least in part, c-
FOS and p21 over-expression. In addition, RIP1 was partially involved in the cell death of HL-
60, but not K562 cells, and the JNK pathway exerted pro-survival signalling in the HXR9-
mediated killing of K562 cells. Likewise, CaM, PKC and HO-1 exerted resistance to HXR9-
mediated cell death (Table 4.1).
Table 4.1 The mechanism of HXR9 cytotoxicity on AML cells.
Causes up-regulation of Involves Independent of Induced by inhibition of
c-FOS
p21
RIP1 (in HL-60) Caspases
Bcl-2-family members
Nuclear fragmentation
CypD
ATP
PARP-1 activation
MEK/ERK pathway
p38 pathway
µ-calpain
p53
JNK pathway (in K562)
NOX
CaM
PKC
HO-1
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Chapter 5 In vivo cytotoxicity of HXR9
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5.1 Introduction
In vitro cytotoxicity of HXR9 on the tested AML cell lines was proven. In addition, this
study data revealed that the concurrent inhibition of CaM and PKC dramatically increased the
sensitivity of the tested AML cell lines to HXR9. Likewise, in vivo efficacy of HXR9 to
eradicate solid tumours of different origins including melanoma, lung, ovarian, breast and
prostate was demonstrated (Morgan, Pirard et al. 2007; Plowright, Harrington et al. 2009;
Morgan, Plowright et al. 2010; Morgan, Boxall et al. 2012; Morgan, Boxall et al. 2014). In the
previous studies, HXR9 was administrated via different routes including intravenously (I.V.),
I.T. or intraperitoneally. In addition, HXR9 did not show adverse side effects on PB and BM
after 10 days of daily I.V. administration at 15mg/kg/day (Morgan, Pirard et al. 2007).
However, the in vivo efficacy of HXR9 on AML cell lines and the side effect of I.V.
administration of high doses of HXR9 have not been studied, yet.
To determine the cytotoxicity of high doses of HXR9 on normal PB, a mouse strain with
the whole blood cells was used, C57BL/6. Due to the intact immunity of C57BL/6 mice, the
mouse AML cell line C1498 that is originated from C57BL/6 mouse strain, a syngeneic tumour
model, labelled with GFP (C1498-GFP) was used to evade the immune system and engraft in
mice. In addition, SCID mice were used to engraft the human AML cell line K562.
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5.1.1 Aims of chapter 5
To assess the efficacy of HXR9 against AML disease in vivo either alone or in
combination with the simultaneous inhibition of CaM and PKC, and to assess the adverse effect
of high doses of HXR9 on normal PB.
The main objectives were:
1. To determine the cytotoxicity of HXR9 either alone or with the simultaneous
inhibition of CaM and PKC on the C1498-GFP cell line in vitro.
2. To establish an AML mouse model by engrafting C1498-GFP or K562 cells in either
C57BL/6 or SCID mice, respectively.
3. To treat these models with high doses of HXR9.
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5.2 Results
5.2.1 HXR9 is cytotoxic for C1498-GFP cells
To test whether HXR9 induces the cell death of the murine AML cell line C1498-GFP,
C1498-GFP cells were treated with a series of HXR9 concentrations for two hours, and
cytotoxicity was determined by LDH assay using ELISA. HXR9 induced C1498-GFP cell death
(Figure 5.1) with an IC50 15.8μM ± 2.6 that determined using Calcusyn software.
Figure 5.1 LDH assay for the cytotoxicity of HXR9 on C1498-GFP cells. Cells were exposed to a titration of
HXR9 for two hours and the cytotoxicity of HXR9 was assessed by measuring LDH enzyme activity in cell-free
supernatants. The graph shows the mean of three independent repeats that had R2 ≥ 0.95 and error bars show the
SEM.
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5.2.2 CaM blocking enhances the cytotoxicity of HXR9
To assess whether the inhibition of CaM produces the same effect on the cytotoxicity
of HXR9 as its inhibition in human AML cell lines, CaM was blocked in the murine AML cell
line C1498-GFP cells using the W-7 inhibitor. As with human AML cells, blocking CaM
increased the sensitivity of C1498-GFP cells to HXR9. For example, co-treatment of C1498-
GFP cells with 10μM HXR9 and 50μM W-7 significantly decreased the percentage of surviving
cells from 72% ± 6.2 to 37% ± 3.3, **p < 0.01, compared to HXR9-only treated cells (Figure
5.2). In addition, co-treatment caused a dramatic decrease in the IC50 of HXR9 by two folds,
from 14.4μM ± 1.2 to 7.5μM ± 2.3 (Figure 5.2).
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IC50
Cell line/ condition HXR9 HXR9+50μM W-7
C1498-GFP 14.4μM ± 1.2 7.5μM ± 2.3
Figure 5.2 The effect of CaM inhibition on the cytotoxicity of HXR9 for C1498-GFP. Cells were pre-incubated
with or without 50μM W-7 for one hour and then with HXR9 for two hours, with or without 50μM W-7, red and
blue curves, respectively. The cytotoxicity of HXR9 was then measured by LDH assay in cell-free media. Co-
treatment with 50μM W-7 significantly increased the cytotoxicity of HXR9 on C1498-GFP cells. The graph shows
the mean of three independent experiments and error bars show the SEM. *p < 0.05, **p < 0.01 and ***p < 0.001
with respect to 50μM W-7 untreated cells. The table shows the difference in the IC50 between HXR9-only treated
cells and HXR9 + 50μM W-7 treated cells.
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5.2.3 Inhibition of PKC activity significantly sensitises cells to HXR9
To assess the effect of inhibition of PKC activity on the sensitivity of C1498-GFP cells
to HXR9, PKC activity was blocked using Ro31-8220 and the sensitivity to HXR9 was
measured by LDH. Ro31-8220 at 30μM dramatically enhanced cell killing by HXR9. For
example, co-treatment of cells with 30μM Ro31-8220 resulted in a huge reduction, p < 0.0001,
in the percentage of surviving cells from 89% ± 3.5 to 58% ± 3.3, compared to cells treated
with 5μM HXR9 alone (Figure 5.3). Likewise, co-treatment with Ro31-8220 decreased the IC50
of HXR9 by twofold from 14.2μM ± 1.9 to 6.6μM ± 2.1 compared to HXR9-only treated cells
(Figure 5.3).
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IC50
Cell line/ condition HXR9 HXR9+30μM Ro31-8220
C1498-GFP 14.2μM ± 1.9 6.6μM ± 2.1
Figure 5.3 The impact of PKC inhibition on the sensitivity of C1498-GFP cells to HXR9. Cells were pre-treated
with or without 30μM Ro31-8220 for one hour and then with HXR9 for two hours, with or without 30μM Ro31-
8220, red and blue curves, respectively. The sensitivity of C1498-GFP cells to HXR9 was then assessed by LDH
assay using ELISA. Results show that blocking PKC dramatically increases the sensitivity of C1498-GFP cells to
HXR9. Graphs show the mean of three independent experiments and error bars show the SEM. *p < 0.05 and
****p < 0.0001 with respect to 30μM Ro31-8220 untreated cells. The table shows the difference in the IC50
between HXR9-only treated cells and HXR9 + 30μM Ro31-8220 treated cells.
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5.2.4 Concurrent inhibition of CaM and PKC significantly enhances the efficacy of
HXR9
To assess the impact of the concurrent inhibition of CaM and PKC on the sensitivity of
C1498-GFP cells to HXR9, both kinases were concurrently blocked and cells were treated with
increasing concentrations of HXR9 and the cytotoxicity of HXR9 was analysed using the LDH
assay. The simultaneous inhibition of both proteins remarkably sensitised C1498-GFP cells to
HXR9. For example, at 5μM HXR9 treatment, the inhibition of CaM and PKC hugely reduced
the percentage of surviving cells from 83% ± 3.9 to 30.4% ± 4.4 (p < 0.0001), compared to
HXR9-only treated cells (Figure 5.4). Likewise, the concurrent blocking of CaM and PKC
remarkably decreased the IC50 of HXR9 by approximately tenfold, from 14.2μM ± 1.3 to 1.5μM
± 1.4 (Figure 5.4).
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IC50
Cell line/ condition HXR9 HXR9+(30μM Ro31-8220+50μM
W-7)
C1498-GFP 14.2μM ± 1.3 1.5μM ± 1.4
Figure 5.4 The effect of CaM and PKC simultaneous inhibition on the efficacy of HXR9. Cells were pre-treated
with or without the combination of the inhibitors, 50μM W-7 and 30μM Ro31-8220, for one hour and then with
HXR9 for two hours, with or without both inhibitors (red and blue curves, respectively). The sensitivity of C1498-
GFP cells to HXR9 was then estimated by measuring LDH enzyme activity in cell-free media using ELISA. The
simultaneous blocking of CaM and PKC remarkably enhanced the efficacy of HXR9. The graph shows the mean
of three independent experiments and error bars show the SEM. *p < 0.05, **p < 0.01 and ****p < 0.0001 with
respect to W-7 and Ro31-8220 untreated cells. The table shows the difference in the IC50 between HXR9-only
treated cells and HXR9 + (50μM W-7 + 30μM Ro31-8220) treated cells.
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5.2.5 Efficient expression of GFP in C1498 cells
The murine AML cell line C1498 was previously transduced to express GFP in order to
monitor tumourgenicity (Zhang, Gajewski et al. 2009). The expression of GFP was confirmed
by fluorescence microscopy, and flow cytometry, which revealed that approximately 95% of
C1498 cells express GFP (Figure 5.5).
A) B)
Figure 5.5 Expression of GFP in C1498 cells. A) Detection of GFP expression, green fluorescence, by fluorescent
microscope (x10). B) Analysis of GFP expression in C1498 by flow cytometry. The percentage of C1498 was
calculated as the percentage of cells that express GFP.
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5.2.6 Establishment a systemic C1498-GFP model in C57BL/6 mice
To establish a murine AML model to assess the cytotoxicity of HXR9 in vivo, C57BL/6
mice (n=10) were inoculated I.V. with 1x106 C1498-GFP cells on day 0 (Table 5.1). The
leukaemia burden was monitored in PB by harvesting blood from random mice on days 3, 7,
13, 17 and 20 (Appendix 9), and estimated by calculating the percentage of GFP+ cells within
the peripheral white blood cells (Figure 5.6). GFP+ cells were found to have engrafted in 2 of
10. This engraftment rate was too low to study the effect of HXR9 in vivo.
A) B)
Figure 5.6 FACS analysis of PB from C57BL/6 mice 13 days after injection of C1498-GFP cells. C1498-GFP
cells were injected in C57BL/6 mice on day 0 and PB was collected from two mice. Peripheral blood mononuclear
cells were analysed by FACS. A) A plot shows no engraftment of C1498-GFP cells. B) A plot shows engraftment
of C1498-GFP cells.
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Table 5.1 A summary of injection of C1498-GFP I.V. in C57BL/6 mice.
Mouse strain C57BL/6
Number of mice 10
Cell line C1498-GFP
Cell inoculation number 1x106 cell/mouse
Inoculation site I.V.
Harvested organ PB
Experiment length (days) 20
Engraftment rate 2 out of 10
(1 in day 13 and 1 in day 17)
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5.2.7 Establishment a systemic C1498-GFP model in C57BL/6 and nude mice
As the first xenograft model did not work, two further mouse strains, C57BL/6 (n=10)
and nude mice (n=5), were inoculated with 1x106 C1498-GFP cells I.V. on day 0 (Table 5.2).
In this experiment, the leukaemia burden was monitored by calculating the percentage of GFP+
cells within the whole population of white blood cells in many sites including PB, BM, liver,
kidney and spleen on days 8, 15, 22, 25, 28 and 35 (Figure 5.7 and appendix 10). Results showed
that the first engraftment site was the liver in both strains. However, cells were engrafted in 3
and 2 mice in C57BL/6 and nude strains, respectively, and two C57BL/6 mice died in the day
22 after injection. The low engraftment rate, the late stage engraftment of cells in PB and the
death of two mice suggested that this xenograft model was not appropriate for analysing the
efficacy of HXR9 in vivo.
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A) C57BL/6 mice
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B) Nude mice
Figure 5.7 FACS analysis of C1498-GFP cells shows engraftment in several organs of C57BL/6 and nude mice.
C1498-GFP cells were inoculated I.V. in both strains on day 0. Several sites including PB, BM, liver, kidney and
spleen were harvested and white blood cells were purified as described in methods. Then, the engraftment of
C1498-GFP cells was analysed by FACS. A) A plot shows C1498-GFP cells engraftment in organs from a
C57BL/6 mouse, B) a nude mouse.
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Table 5.2 A summary of injections of C1498-GFP I.V. in C57BL/6 and nude mice.
Mouse strain C57BL/6 Nude
Number of mice 10 5
Cell line C1498-GFP
Cell inoculation number 1x106 cell/mouse
Inoculation site I.V.
Harvested organs PB, BM, liver, kidney and spleen
1st engraftment site Liver
Experiment length (days) 28 35
Engraftment rate 3 out of 10
(2 and 1 in days 22 and 28,
respectively)
2 out of 5
(1 in day 22 and 1 in day 28)
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5.2.8 Development a C1498-GFP flank model in C57BL/6 mice
As the I.V. inoculation of C1498-GFP cells was not appropriate to establish it as an
AML model to study the efficacy of HXR9 in vivo, the flank xenograft attempt was done to
assess its efficacy to be as a model. C57BL/6 mice (n=5) were injected S.C. with 1x106 C1498-
GFP cells on day 0 (Table 5.3). The leukaemic tumours started to be measured on day 7 and all
mice had tumours by day 20. Therefore, the flank xenograft was used to conduct in vivo
experiments.
Table 5.3 A summary of injection of C1498-GFP S.C in C57BL/6 mice.
Mouse strain C57BL/6
Number of mice 5
Cell line C1498-GFP
Cell inoculation number 1x106 cell/mouse
Inoculation site S.C.
Experiment length (days) 20
Engraftment rate 100%
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5.2.9 HXR9 significantly extended the survival of C1498-GFP xenograft
Due to its successful engraftment, the flank xenograft of C1498-GFP cells in C57BL/6
mice was established in order to assess the effect of HXR9 in vivo. C1498-GFP cells were
implanted S.C. in the flank, 1x106 cell/mouse. When tumours became palpable, the mice were
divided into two cohorts of 9 mice each with one cohort treated with 50mg/kg HXR9 three
times/week I.T. and the other with PBS as a control. Comparison of tumour growth delay
between the different cohorts at each time point showed that there was no statistical difference
(p = 0.23) in tumour sizes although the mean tumour volume on day 12 was less than half that
of the control group (Figure 5.8A). OS data showed that HXR9-treated mice survived
significant longer than their counterpart (p = 0.036), 22% compared to 0% for HXR9 and PBS
treated mice, respectively (Figure 5.8B).
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A)
B)
Figure 5.8 HXR9 treatment effect on the growth of C1498-GFP xenograft in female C57BL/6 mice. Mice were
injected with 1x106 C1498-GFP cells S.C. in the flank. When xenografts became palpable, day 0, mice were treated
I.T. with 50mg/kg HXR9 (n=9) or PBS (n=9) three times/week. Mice were culled when tumour sizes reached ≥
1500 mm3. A) Mean changes in xenograft volumes following HXR9 or PBS treatment and error bars show SEM.
B) The corresponding survival curve for mice treated with HXR9 or PBS shown in A.
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5.2.10 Simultaneous inhibition of CaM and PKC did not sensitise C1498-GFP flank
tumour to HXR9
To assess whether the concurrent inhibition of CaM and PKC in vivo could enhance the
efficacy of HXR9 as it did in vitro, the same drug combination was used in the C1498-GFP
flank tumour. Flank tumours were initiated in C57BL/6 mice by S.C. administration of C1498-
GFP cells, 1x106cell/mouse. When tumours became measureable, mice were divided into four
groups. Two control groups were treated with either PBS or CaM and PKC inhibitors, 500μM
W-7 and 300μM Ro31-8220, respectively; the other groups were either treated with 50mg/kg
HXR9 alone or 50mg/kg HXR9 plus the inhibitors simultaneously for three times/week.
Tumour growth data demonstrated that there were no statistical variances among the different
groups at any time point. Although the mean tumour volume in HXR9 and the inhibitor treated
mice on days 21, 23 and 25 showed an entire absence of tumour, this data represented the
tumour volume of one mouse out of 10 mice (Figure 5.9A). Consequently, the OS of the treated
groups with HXR9 alone or HXR9 and the inhibitors were not statistically significant different
compared to the OS of the control groups (Figure 5.9B).
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A)
B)
Figure 5.9 Effect of HXR9 I.T. treatment on the growth of C1498-GFP flank xenograft in C57BL/6 mice. C1498-
GFP cells (1x106) were inoculated S.C. in the mice flanks. When tumours became palpable, mice were split into
four groups, two control groups were administrated PBS (n=9) and CaM and PKC inhibitors (n=10), 500μM W-7
and 300μM Ro31-8220, respectively; and the other two groups were administrated either 50mg/kg HXR9 alone
(n=10) or 50mg/kg HXR9 + (500μM W-7 and 300μM Ro31-8220) (n=10) I.T. for three times/week. When tumours
reached ≥ 1500 mm3, mice were excised. A) Changes in tumour volumes resulting from treatment with PBS,
500μM W-7 and 300μM Ro31-8220, 50mg/kg HXR9 or 50mg/kg HXR9 + (500μM W-7 and 300μM Ro31-8220)
and error bars show SEM. B) Corresponding survival curve for mouse groups shown in A.
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5.2.11 HXR9 did affect the tumour growth of K562 flank model
As an additional in vivo model, human AML cell line K562 was used in the SCID mice.
K562 cells were inoculated S.C. in the flank of SCID mice at 1x106 cell/mouse. When tumours
became measurable, the mice were split into two groups of 6 mice each. One group was injected
I.T. with 100mg/kg HXR9 three times/week, except on the 10 and 12 mice were treated with
300mg/kg and 200mg/kg HXR9, respectively. The control group was injected with PBS.
Tumour growth delay and median survival data revealed that there was no statistical difference
between the two groups (Figure 5.10). Of note, 300mg/kg and 200mg/kg HXR9 led to the death
of two mice each.
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A)
B)
Figure 5.10 Impact of HXR9 treatment on the K562 flank-xenograft growth in SCID mice. The xenograft was
initiated by S.C. injection of 1x106 K562 cells in the flank. When tumours became measureable, day 0, mice were
treated I.T. with HXR9 (n=6) or PBS (n=6) three times/week. HXR9-treated mice were treated with 100mg/kg
overall the experiment except on day 7 and 10 where they were treated with 300mg/kg and 200mg/kg respectively.
Mice were euthanized when tumour volumes reached ≥ 1500 mm3. A) Mean values of tumour volumes of HXR9-
treated or PBS-treated mice and error bars show SEM. B) OS of HXR9- and PBS-treated mice.
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5.3 Discussion
Leukaemic mouse models were established four decades ago after the development of
SCID mice. This model is characterised by the lack of active T and B lymphocytes (Bosma and
Carroll 1991). Subsequently, the non-obese diabetic with severe combined immunodeficiency
(NOD-SCID) model was developed; the mice lack functional T and B lymphocytes and have
fewer NKs (Shultz, Schweitzer et al. 1995). Another model is nude mice, which are
characterised by the absence of functional T lymphocytes and a partial absence of B
lymphocytes. There are also other leukaemic mouse models available including NOD-SCID-
β2 null and NOD-SCID-IL-2Rγ null. The leukaemic xenotransplantations in mice can be
established by different administrations including I.V., intrafemoral and S.C. inoculations. The
intrafemoral injection of leukaemic cells into the BM is the most common and effective
xenotransplantation administration. These preclinical models are used to assess the anti-cancer
efficacy of drugs.
One of the aims of this chapter was to measure the cytotoxicity of HXR9 and its
cytotoxic efficacy on AML cells in vivo. Therefore, C57BL/6 mouse strain that has a full
functional immune system and the murine AML cell line C1498 labelled with GFP were used
for this purpose. The in vitro cytotoxicity assay showed the sensitivity of C1498-GFP to HXR9
and this sensitivity was further induced by either inhibiting CaM or PKC activity alone or
concurrently, which indicates that C1498-GFP cells respond to HXR9 as human AML cells.
The systemic injection of C1498-GFP in C57BL/6 mice resulted in only a low engraftment rate.
This low engraftment in PB might be the result of C1498-GFP cell lysis by NKs and CD8+
cytotoxic lymphocytes (Boyer, Orchard et al. 1995). In addition, the level of engraftment
measured in this study is typically seen in AML xenotransplantation experiments (Sanchez,
Perry et al. 2009).
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As a result, the systemic xenotransplantation was redone in two mouse strains including
C57BL/6 and nude mice, monitoring engraftment in the PB, BM, liver, kidney and spleen. This
experiment showed engraftment of C1498-GFP cells in all harvested organs with low
engraftment in PB and BM. Extramedullary infiltration, the presence and proliferation of
leukaemic cells in sites beyond the medullary spaces of BM including liver, kidney and spleen,
is a common symptom in AML and associated with poor prognosis and poor OS (Ravindranath,
Steuber et al. 1991; Creutzig, Harbott et al. 1995; Rubnitz, Raimondi et al. 2002; Chang,
Brandwein et al. 2004). Extramedullary infiltration was also reported in mouse
xenotransplantations (Krivtsov, Twomey et al. 2006; Sanchez, Perry et al. 2009). The causes of
extramedullary infiltration are not fully understood. However, the interaction between the
chemokine stromal cell-derived factor-1 alpha produced from human and murine liver, kidney
and spleen, and its receptor C-X-C chemokine receptor type 4 (CXCR4), is highly expressed
by AML and other leukaemic cells is considered as a crucial step in AML cell proliferation and
trafficking, AML disease progression and extramedullary infiltration (Mohle, Schittenhelm et
al. 2000; Kollet, Spiegel et al. 2001; Kollet, Shivtiel et al. 2003; Kato, Niwa et al. 2011). In
addition, targeting CXCR4 prevented the engraftment of AML cells in the BM and spleen of
transplanted mice (Tavor, Petit et al. 2004; Zhang, Patel et al. 2012). Obviously, the overall
engraftment rate was relatively low in both strains which may due to some immune activity in
both strains or due to technical issues such as failure to inject cells in the tail vain.
The I.V. injection of C1498-GFP cells in C57BL/6 and nude mice was clinically
relevant to AML disease. However, this systemic injection has many limitations. For example,
the low engraftment rate which could be overcome by pre-treating mice with a combination
regimen of total body irradiation and a pharmacological immunosuppressive agents. It is well-
documented that total body irradiation of recipient mice sustained engraftment by decreasing
the homing ability to the marrow microenvironment, decreasing the repopulating ability and
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causing cell death of the resident stem and progenitor cells and accordingly allowing injected
cells to engraft (Kahl, Mielcarek et al. 2004). In addition, the combination regimen inhibited
graft rejection by host T-cells which increased the engraftment rate (Mielcarek, Torok-Storb et
al. 2011; Shan and Ma 2013). Furthermore, the high engraftment rate of C1498-GFP cells in
extramedullary organs and the low engraftment rate in PB was a limitation due to the lack of
non-invasive monitoring system of the progression of AML which could be overcome by using
bioluminescence imaging. This technique can rapidly and noninvasively detect disease
progression from early stages by measuring light emission from cells labelled with the
bioluminescent reporter genes.
Due to the low systemic engraftment rate; the lack of radiation and bioluminescence
imaging facilities, the S.C. engraftment model was used in this study. This model is not a
realistic representation of AML, but it is an acceptable model for the preclinical evaluation of
drugs (Vinante, Rigo et al. 1999; He, Liu et al. 2003; Siegler, Kalberer et al. 2005; Motiwala,
Majumder et al. 2009; Zhang, Zhang et al. 2012; Chen, Yang et al. 2013; Park, Mishra et al.
2013; Sukhai, Prabha et al. 2013; Li, Li et al. 2014; Yang, Yu et al. 2014). The flank model in
C57BL/6 mice showed 100% engraftment rate that indicated to the usefulness of this model to
assess the efficacy of HXR9. At the first experiment, the concentration of HXR9 used was
50mg/kg, which was the half of the maximum concentration that previously used (Morgan,
Plowright et al. 2010; Morgan, Boxall et al. 2012; Morgan, Boxall et al. 2014). The I.T injection
of HXR9 showed that HXR9 was effective in delaying the growth of C1498-GFP cell tumour.
At the second experiment, the concurrent inhibition of CaM and PKC was introduced with the
same concentration of HXR9, 50mg/kg. Similar to the first experiment, using concurrent
inhibition of CaM and PKC did not sensitize tumours to 50mg/kg HXR9. This result was
consistent with in vitro results that showed the concurrent inhibition of CaM and PKC was not
cytotoxic itself, but increased the sensitivity to HXR9.
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The human AML cell line K562 and SCID mice were used as a second in vivo model.
At 100mg/kg, HXR9 was not effective in reducing tumour sizes of treated mice. In addition, at
300mg/kg and 200mg/kg, HXR9 caused the death of two mice at each concentration. This effect
might be owing to the systemic leakage of HXR9 through tumour microvessels which might
result in HXR9 mediated toxicity. The acute toxicity of HXR9 might result from the cationic
R9 which is well-known for its nonspecific interactions and ability to penetrate most organs
(Vives, Schmidt et al. 2008; Sarko, Beijer et al. 2010).
The ethics regulations might underestimate the in vivo cytotoxicity of HXR9 on AML
cells. According to the ethics mice were culled when tumours reached 1500 mm3, this action
might underestimate the difference in tumour sizes between different mouse groups, treated and
untreated mice. This is one of the limitations of using the flank model for AML
xenotransplantation and could be overcome by using a systemic in vivo model.
The main challenge of treating cancer is drug resistance. To understand the mechanisms
of chemoresistance, the structure of solid tumours should be known. Solid tumours are
heterogenous environment compose of cancer and stromal cells. The stromal cells includes
fibroblasts, immune, endothelial and mesothelial cells, adipocytes and extracellular matrix
(Castells, Thibault et al. 2012). The resistance to therapy is complicated and caused by several
factors. Some of resistance factors are cell intrinsic ''biochemical'' including genetic mutations
that affect drug uptake, over-expression of antiapoptotic and/or survival genes (Wilson,
Longley et al. 2006; Bleau, Hambardzumyan et al. 2009). Other factors are cell extrinsic
''physiological'' that resulted from the stromal microenvironment (Williams, den Besten et al.
2007; Eckstein, Servan et al. 2009). For example, tumours characterised by their extracellular
acidity and their alkaline or neutral intracellular environments (Tredan, Galmarini et al. 2007).
The extracellular pH influences the cellular uptake and thereby the efficacy of chemotherapies.
For example, increasing the extracellular acidity by I.V. glucose administration, through an
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unknown mechanism, promoted the uptake and the efficacy of weak acid drugs such as
chlorambucil, while it decreased the tumour growth delay of the small-cell lung carcinoma cell
line 54A by the weak base doxorubicin (Gerweck, Vijayappa et al. 2006). In contrast, in vitro
and systemic in vivo alkalinisation of extracellular environment by sodium biocarbonate
increased the uptake and the efficiency of weak base drugs including anthracyclines, taxanes
and alkylating agents on the breast cancer cell line MCF-7, whereas it reduced the cytotoxicity
of weak acid drugs including cyclophosphamide, 5-fluorouracil and chlorambucil (Mahoney,
Raghunand et al. 2003; Raghunand, Mahoney et al. 2003). It is noteworthy that HXR9 is
composed of the functional peptide and R9 which is one of the cell-penetrating peptides that
characterised by its cationic feature (Jin, Zhang et al. 2013). Therefore, the tumour extracellular
acidity might affect the uptake and the in vivo efficacy of HXR9 on the tested cell lines
including C1498-GFP and K562 cells. Tumour microenvironments are also characterised by
the presence of some hypoxia regions (Tredan, Galmarini et al. 2007; De Bock, Mazzone et al.
2011). Hypoxia was documented to cause in vivo drug resistance by activating the expression
angiogenic, anti-apoptotic and survival genes through triggering the over-expression of hypoxia
inducible factor-1 (Pouyssegur, Dayan et al. 2006). hypoxia inducible factor-1 in turn induces
the expression of NF-ĸB that activates the survival pathway by enhancing PI3K/Akt (Eltzschig
and Carmeliet 2011). NF-ĸB also mediates the expression of the inflammatory molecules such
INF-γ and TNF-α that induces the expression of the angiogenic factor vascular endothelial
growth factor (VEGF) from endothelial cells and cancer cells (Eltzschig and Carmeliet 2011).
VEGF is a mediator of the over-expression of the anti-apoptotic proteins including Bcl-2 and
XIAP (Nör, Christensen et al. 1999; Lee, Shanafelt et al. 2005). However, the inhibition of
VEGF expression in a S.C. model of K562 led to a decrease in the tumour microvessels and
caused cell apoptosis (He, Liu et al. 2003). Hypoxia was also reported to cause up-regulation
of P-glycoprotein, which is responsible of drug efflux (Comerford, Wallace et al. 2002).
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Tumours cause infiltration and accumulation of immune cells such as macrophages in tumour
microenvironments (De Palma and Lewis 2011). This subset of macrophages called tumour
associated macrophages and functions as a tumour growth inducer by increasing tumour
vascularisation and angiogenesis through over-expressing VEGF (Stockmann, Doedens et al.
2008). In addition, those macrophages facilitate tumour growth by providing
immunosuppressive microenvironment through expressing anti-inflammatory molecules such
as interleukin 10, transforming growth factor β (Hao, Lu et al. 2012). The concurrent treatment
regimen of tumour associated macrophages and chemotherapy caused a significant reduction
in tumour sizes (DeNardo, Brennan et al. 2011). Adhesion of stromal cells to cancer cells also
promoted the survival of cancer cells (Correia and Bissell 2012). Resistance to new therapies
is multi-factorial and tumour microenvironment has an integral part in it.
The discrepancy between in vitro and in vivo response of both tested mouse and human
AML cell lines C1498-GFP and K562, respectively, indicated that tumour microenvironment
might played a crucial role in the in vivo chemoresistance to HXR9. However, this potential
explanation does not exclude the acquired chemoresistance that resulted from cell intrinsic
mutations. To understand and improve the in vivo efficacy of HXR9, more molecular assays
and the physiology of tumour microenvironment are needed to be investigated. Of note, the
S.C. injection does not realistically reflect the nature of AML disease. Therefore, there is a need
to develop a reliable preclinical in vivo model for the assessment of the efficacy of HXR9.
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5.3.1 Summary of chapter
In summary, in vitro results showed that HXR9 has a significant anti-tumour activity on
C1498-GFP cells that was sharply promoted by either inhibition of CaM/PKC or both of which.
However, S.C. injection of C1498-GFP mouse model results demonstrated that this cell line
was chemo-resistant to either HXR9-alone treatment or combination treatment of HXR9 and
concurrent inhibition of CaM and PKC. The chemoresistance to HXR9 was also seen in K562
S.C. implemented into SCID mice. In addition, HXR9 resulted in death of some mice at high
concentrations. Finally, the attempt to develop a systemic mouse model for C1498-GFP cells
in C57BL/6 and nude mice resulted in low engraftment rate.
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Chapter 6 General Discussion and Future Directions
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6.1 Discussion
AML is considered to be the most frequent acute leukaemia to affect adult patients.
Conventionally, AML is classified cytogenetically according to the cell karyotypes into three
prognostic groups: favourable prognostic group including t(8;21), inv(16) and t(15;17), an
adverse prognostic group including complex karyotypes, trisomy 8 and CML-blast crisis, and
an intermediate prognostic group including most other abnormalities and NK-AML (Roche,
Zeng et al. 2004). Of note, about 45% of AML cases exhibit normal karyotypes (Marcucci,
Mrozek et al. 2005). The treatment regimen for all prognostic groups has not been changed
substantially over the last 4 decades, with the exception of APML. The current standard
induction therapy is a combination of DNR for 3 days with cytarabine for 7 days in a so-called
''3+7 regimen''. This regimen usually achieves 70%-75% CR in young (< 60 years), and 50%
CR in old patients. A post-induction, consolidation regimen consisting of high cytarabine doses
or alternatively allogeneic transplantation improves the outcome of AML patients. However,
the outcome of this regimen is heterogonous and the majority of patients will relapse and die
even after achieving clinical CR (Burnett 2012; Patel and Levine 2012).
The significant heterogeneity in the outcome of AML patients might be due to several
reasons. First, the current CR criteria are broad and totally dependent on microscopic counting
of the number of blast cells in BM to determine if the make-up ≤ 5% of the cell population, and
has not changed substantially since 1956 (Hourigan and Karp 2013). This limitation could be
overcome by including the detection of AML-specific molecular stable targets using PCR or
flow cytometry in CR criteria. The detection of AML cells after chemotherapy at levels under
the sensitivity level of the microscopic detection is known as minimal residual disease (MRD)
(Ossenkoppele and Schuurhuis 2013). Second, the low specificity of the current treatment
regimen also plays a role in the heterogeneity of response. To overcome this limitation, AML
patients should be treated with more cell-specific drugs. Third, there is heterogeneity of AML
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cells with respect to treatment sensitivity; for example, favourable prognostic groups respond
better than poor prognostic groups to the currently used chemotherapy.
Currently, the evaluation of relapse risk of AML (except for APML) patients in initial
clinical CR is based on the average clinical results of large historical populations who had the
same pre-treatment chromosomal and molecular abnormalities at the presentation time
(Mrozek, Marcucci et al. 2012; O'Donnell, Abboud et al. 2012). This assessment method lacks
the evaluation of the induction treatment efficiency by patient-individualised-evaluation and
can be improved by including MRD detection to guide decisions to the most-suitable post-
induction treatment. MRD is currently applicable in CML to detect the BCR-ABL fusion that
is present in almost all CML cases (Cross, White et al. 2012; O'Hare, Zabriskie et al. 2012). In
AML, MRD is now only used in APML to detect the PML-RARA fusion (Grimwade,
Jovanovic et al. 2009; Chendamarai, Balasubramanian et al. 2012). However, MRD is not
involved in the rest of AML cases; therefore, finding genes that are over-expressed in AML
cells is still an open area for more studies. For example, the most common mutated genes in
AML, including NPM1 and FLT3, might be potential detectable by MRD (Kristensen, Møller
et al. 2011; Buccisano, Maurillo et al. 2012). The detection of molecular targets offers a higher
sensitive than the current microscopic detection and presents a more-personalised approach to
diagnosis. It was reported that detection of the NPM1mut MRD preceded haematological relapse
by a median time of 8 weeks (Schnittger, Kern et al. 2009). Analogously, therapeutic
intervention after the detection of molecular relapse by MRD improved clinical outcomes
(Rubnitz, Inaba et al. 2010; Inaba, Coustan-Smith et al. 2012). It is noteworthy that HOX gene
over-expression is a common event in many AML subtypes and is strongly associated with a
poor prognosis, as previously mentioned in the introduction, and thus they are potential targets
in MRD.
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The heterogeneity of AML is due to the presence of various cytogenetic abnormalities
in 55% of AML cases including numerical aberrations, structural aberrations and balanced
translocations (Buccisano, Maurillo et al. 2012). In addition, molecular studies revealed a
diversity of molecular markers that further sub-classified the majority of NK-AML patients
(Marcucci, Mrozek et al. 2005). The discovery of molecular markers in NK-AML leads to the
hypothesis that leukaemogenesis results from the interaction of many mutations. These lead to
either an increase in cell proliferation (class I mutations) or a block of cell differentiation (class
II mutations), ultimately leading to AML (Dohner and Dohner 2008; Renneville, Roumier et
al. 2008; Betz and Hess 2010).
The burgeoning molecular understanding of AML could be valuable in many ways,
including the further sub-classification of the disease and the development of more personal
prognostic predictions that could form the cornerstone for future therapeutic regimens.
Currently, all-trans retinoic acid (ATRA) is the only widely used molecularly targeted therapy
in AML, for the treatment of APML characterised by the presence of t(15;17)/PML-RARA.
Understanding the molecular mechanism, signalling pathways and cooperating mutations will
pave the way for new potential molecular drugs that hopefully will result in more personalised
treatment.
The diversity of cytogenetic and molecular aberrations is one of the main obstacles to
the development of targeted therapies for AML, and the definition of the common genetic
abnormalities in patients with different kinds of mutations may lead the way to subtype specific
therapies. MLL fusion proteins were reported in 10% of therapy-related AML and 3% of de
novo AML (Slany 2009). Core binding factor translocations including CBFB/MYH11 and
RUNX1/RUNX1T1 fusions constitute 6% and 10% of all AML cases, respectively (Haferlach
2008). NUP98-HOX fusion proteins were also reported in AML (Fujino, Suzuki et al. 2002;
Suzuki, Ito et al. 2002; Taketani, Taki et al. 2002). Advances in molecular techniques also
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defined genetic aberrant expressions in AML. For example, the dysregulation of CDX2 and
CDX4 occur in 90% and 25% of AML cases, respectively (Bansal, Scholl et al. 2006; Scholl,
Bansal et al. 2007). Wilms' tumour gene 1 over-expression was seen in 80%-90% of AML cases
(Ostergaard, Olesen et al. 2004; Weisser, Kern et al. 2005), and the NPM1 mutation is present
in 35% of all AML cases and approximately 55% of NK-AML (Falini, Mecucci et al. 2005;
Thiede, Koch et al. 2006; Schlenk, Döhner et al. 2008). FLT3 mutations are present in 25% of
all AMLs and in 70% of NK-AMLs (Thiede, Steudel et al. 2002; Yanada, Matsuo et al. 2005).
The NRAS mutation was reported in 11% of AMLs (Bacher, Haferlach et al. 2006). These
aberrations can be found alone or in combination. For example, FLT3 is often coincident with
NPM1 or MLL mutations (Döhner, Schlenk et al. 2005; Olesen, Nyvold et al. 2005).
The above-mentioned combinations of FLT3 with NPM1 or MLL indicates that genetic
and cytogenetic aberrations cannot be assessed independently, and may in fact reflect the early
mechanism of leukaemogenesis and narrow down the potential genetic and signalling pathways
that are suitable for therapeutic approaches. An in depth analysis of AML subtypes suggest that
HOX genes can be one of the common denominators of different cytogenetic and molecular
aberrations. HOX gene over-expression was reported in AML with chromosomal rearrangement
including MLL fusion proteins and MYST3-CREBBP translocation (Ayton and Cleary 2003;
Camos, Esteve et al. 2006; Slany 2009). Mll fusion was found to function in AML by
collaborating with Ras to drive the over-expression of Hoxa9 and activate Raf (Ono, Kumagai
et al. 2009). Hox proteins can also form fusion proteins with Nup98 that mediate AML
formation in combination with flt3 (Borrow, Shearman et al. 1996; Nakamura, Largaespada et
al. 1996; Raza-Egilmez, Jani-Sait et al. 1998; Kroon, Thorsteinsdottir et al. 2001; Fujino,
Suzuki et al. 2002; Suzuki, Ito et al. 2002; Taketani, Taki et al. 2002; Pineault, Abramovich et
al. 2004; Palmqvist, Argiropoulos et al. 2006). In addition, NUP98-HOXA9 protein was
significantly associated with KRAS and Wilms' tumour gene 1 mutations (Chou, Chen et al.
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2009). Hox genes are regulated by the Cdx family in AML (Bansal, Scholl et al. 2006; Rawat,
Thoene et al. 2008),and, likewise, gene profiling studies showed a strong association between
FLT3 mutations and HOXA7, HOXA9 and HOXA10 expression (Roche, Zeng et al. 2004;
Eklund 2011). Over-expression of several HOX genes was also seen in the presence of NPM1
mutations (Mullighan, Kennedy et al. 2007; Vassiliou, Cooper et al. 2011). These studies
suggested that HOX genes are novel targets for molecular targeted therapies in various AML
subtypes.
Targeting a group of HOX proteins by disrupting their interactions with their cofactor
PBX was previously achieved using the small cell-permeable peptide HXR9. This approach
illustrated the importance of HOX genes in the tumorigenesis and survival of the cancer cells
derived from several solid cancers (Morgan, Pirard et al. 2007; Shears, Plowright et al. 2008;
Plowright, Harrington et al. 2009; Morgan, Plowright et al. 2010; Morgan, Boxall et al. 2012;
Morgan, Boxall et al. 2014). Therefore, this study was carried out based on the hypothesis that
AML cells would be sensitive to killing by HXR9 and indeed HOX genes might be a useful
therapeutic target in a subset of AML patients that over-express HOX genes.
In order to test this hypothesis I measured HOX gene expression in AML cell lines
derived from different AML subtypes and assessed their susceptibility in vitro to HXR9
treatment either alone or in combination with first line AML chemotherapies (chapter 3). I also
performed a comprehensive study of the mechanism of cell killing by HXR9, (chapter 4) and
assessed the in vivo efficacy of HXR9 to kill AML cells either alone or in combination with
non-toxic reagents (chapter 5).
The AML cell lines used in this study represented various AML sybtypes and prognoses.
These included AML patients with de novo mutations (primary AML), including KG-1,
HEL92.1.7 and HL-60. In addition, cell lines were derived from CML patients in blast crises
(secondary AML), including K562 and KU812F. KG-1 is an erythroleukaemia cell line
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characterised by the expression of CD34 and the presence of a complex karyotype, and is
consequently associated with a poor prognosis (Koeffler and Golde 1978; Ahmed, Laverick et
al. 1999; Mrózek, Tanner et al. 2003; Mrozek 2008). HEL92.1.7 is also an erythroleukaemia
cell line carrying the Janus kinase 2 mutation that is associated with a poor prognosis: 80% of
patients relapse (Martin and Papayannopoulou 1982; Illmer, Schaich et al. 2007; Fiskus,
Verstovsek et al. 2011). HL-60 is an APML cell line associated with a favourable prognosis
and characterised by the presence of t(15;17)/(PML-RARA) (Collins, Ruscetti et al. 1978;
Dores, Devesa et al. 2012). Both K562 and KU812F are characterised by the presence of the
Philadelphia (Ph) chromosome t(9;22)/(BCR-ABL) and derived from CML patients in blast
crisis, which is indicative of a poor prognosis (Lozzio and Lozzio 1975; Kishi 1985; Roche,
Zeng et al. 2004; Rucker, Sander et al. 2006; Weisberg, Wright et al. 2008).
The expression of HOX genes in the tested AML lines was consistent with the
established idea that HOX genes are expressed in a lineage-restricted manner, whereby HOXA
and HOXB genes are expressed in myeloid (KU812F) and erythroid (HEL92.1.7 and K562)
cells, respectively. The exception to this was the erythroid cell line KG-1, which expressed
HOXA genes that were linked to the presence of trisomy 8. In addition, the APML cell line HL-
60 was characterised by a global down-regulation of HOX genes. It is noteworthy that the
CD34+ cell line KG-1, which is characterised by a poor response to conventional
chemotherapies, strongly expresses a set of HOX genes compared with the CD34- cell line HL-
60 that represents AML with a favourable prognosis (Ahmed, Laverick et al. 1999; Marone,
Scambia et al. 2002; Dores, Devesa et al. 2012). This difference in the prognosis is associated,
at least in part, to the expression of HOX genes since the high and low expression of HOX genes
corresponds to a poor and favourable prognosis, respectively (Golub, Slonim et al. 1999;
Andreeff, Ruvolo et al. 2008; Zangenberg, Grubach et al. 2009). In addition, the comparison
between KG-1 and HL-60 in terms of HOX expression is consistent with the previous
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observation that HOX genes are highly expressed in CD34+ cells but show virtually no
expression in CD34- cells (Sauvageau, Lansdorp et al. 1994; Pineault, Helgason et al. 2002).
The limitation of assessing the expression of HOX genes was the lack of normal CD34+ cells,
which could be addressed in future studies.
The functional redundancy of HOX genes obscures our understanding of their
importance in the survival and proliferation of cancer cells, even though their over-expression
has been reported in different kinds of cancers. In this study I showed for the first time that the
high expression of HOX genes could form the basis of individualised medicine in AML as
antagonising the interaction between a set of HOX proteins and their cofactor PBX by the small
cell-permeable peptide HXR9 resulted in cell death. I also found that the susceptibility of the
tested AML cell lines to HXR9 was correlated with the expression level of HOX genes. The
most susceptible cell line was KG-1 that highly expresses a set of HOX genes, while HL-60
showed very low HOX gene expression had the lowest susceptibility to HXR9. Interestingly,
the expression of CD34 is considered to be a marker of a poor prognosis, an adverse clinical
course and a low CR rate of AML (Repp, Schaekel et al. 2003). In addition, CD34+ AML cells
were reported to be resistant to the current chemotherapies, yet CD34+ might predict high
sensitivity to HXR9 since HOX genes are highly expressed in CD34+ cells (Bailly, Muller et al.
1995; Bailly, Skladanowski et al. 1997). Likewise, gene expression profiling of CD34+ cells in
CML patients in chronic phase reveals a higher expression of HOXA9, which may suggest that
HXR9 could be used in combination with imatinib, the first line treatment for CML, which
targets the BCR-ABL fusion protein (Diaz-Blanco, Bruns et al. 2007; Palandri, Castagnetti et
al. 2008; Hehlmann 2012). Consistent with this suggestion, the cell lines that express BCR-
ABL fusion, KU812F and K562, were sensitive to HXR9. In addition, it was reported that Bcr-
Abl cooperated with Nup98-Hox proteins to progress to the ultimate phase of CML that is blast
crisis (Mayotte, Roy et al. 2002; Ito, Kwon et al. 2010; Di Giacomo, Pierini et al. 2014).
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Analogously, the sensitivity of Ph+ AML cells to HXR9 might suggest that HXR9 could be
used in a combination with imatinib to kill ALL cells that express both BCR-ABL fusion and
HOX genes (Redaelli, Piazza et al. 2009). Since HL-60 is sensitive to HXR9, a rational
approach may be to combine HXR9 with the current first line treatment for APML, ATRA
(Tallman and Altman 2009).
Combination therapy is a common approach to treating drug-resistant cancer, and
depends on a synergistic interaction between drugs that presumably have complementary
modes of action. Multiple DNA-damaging agents were reported to activate NF-ҡB, such as
anthracyclines, alkalyting agents and topoisomerase inhibitors (Boland, Fitzgerald et al. 2000;
Laurent and Jaffrézou 2001; Campbell, O'Shea et al. 2006). However, NF-ҡB is negatively
regulated by c-FOS, which is the main pro-apoptotic downstream target of HXR9 (Ray,
Kuwahara et al. 2006; Takada, Ray et al. 2010). In addition, DNA-damage is known to stimulate
PKC activity that was found to have an antagonistic, anti-apoptotic effect on HXR9 in this study
(Johnson, Lu et al. 2002; Bluwstein, Kumar et al. 2013). The findings of this study therefore
suggest that DNA-damaging agents that activate NF-ҡB and PKC might not be suitable
combination partners to HXR9. The molecular mechanism of HXR9 is yet to be fully
characterised. The specificity of HXR9 to target the HOX-PBX interaction and modify the
transcription of some genes were previously assessed, and the mode of cell death of the tested
solid cancers showed that HXR9 led to cell death by either apoptosis or necrosis (Morgan,
Pirard et al. 2007; Shears, Plowright et al. 2008; Plowright, Harrington et al. 2009; Morgan,
Plowright et al. 2010; Morgan, Boxall et al. 2012; Morgan, Boxall et al. 2014). However, the
findings of this study indicate that there is difference, possibly unique mechanism of cell death
by HXR9 in AML.
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The study of cell death was carried out by assessing the changes in various cellular
compartments after HXR9 treatment. For example, nuclear morphology was assessed by DAPI
staining. The limitation of this assay was the lack of a positive control for nuclear
fragmentation. The cytosolic changes were studied by analysing the changes in different
members including the pro-apoptotic member of the Bcl-2 family, XIAP, Apaf1 and caspase
activation. In addition, the mitochondrial role was investigated by measuring the expression of
the pro-apoptotic and anti-apoptotic genes of the Bcl-2 family including Bad, Bax, Bak1 and
Bcl-2. All the above-mentioned studies revealed that HXR9-induced cell death through an
apoptosis-independent pathway, involving programmed necrosis.
Ca2+ and ROS are two main regulators of necrosis (Festjens, Vanden Berghe et al.
2006). The role of extracellular Ca2+ was assessed using EDTA. However, EDTA is a general-
cation inhibitor that might chelate HXR9 since R9 is a cation sequence. EGTA is another cation
chelator used in a preliminary experiment, data not shown. EGTA increased the sensitivity to
HXR9, although this might be the effect of its alkaline solvent NaOH that might increase the
pH of the extracellular environment and thereby strengthen the interaction between the
polycation HXR9 and the acidic cell surface (Raghunand, Mahoney et al. 2003). To the best of
my knowledge, there is no specific cell impermeable Ca2+ chelator. In addition, intracellular
Ca2+ concentration was not measured in this study, although Ca2+ downstream mediators of
necrosis were.
Programmed necrosis is classified into various types such as mitochondrial necrosis,
parthanatos and necroptosis (Andrabi, Dawson et al. 2008; Elrod and Molkentin 2013; Marshall
and Baines 2014). The assessment of CypD and PARP1 activation suggested that HXR9-
induced necrosis in mitochondria- and parthanatos-independent pathways, respectively. Of
note, the limitation of PARP1 activation assay was the lack of a positive control. However,
RIP1 is involved, at least in part, in HL-60 necroptosis and the absence of any RIP1 inhibition
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effect on K562 does not exclude a necroptotic pathway since necroptosis can be RIP1-
independent, but RIP3-dependent. Therefore, a RIP1 inhibition assay might suggest the need to
block the role of RIP1 and RIP3 by short interfering RNA or a pharmacological inhibitor to
analyse whether HXR9-induced necrosis in the only necroptosis-dependent pathway or whether
there is another pathway involved.
ATP depletion was achieved by incubating cells with a high concentration of fructose,
which is known to deplete intracellular ATP. The limitation of this assay was the lack of a
positive control. In addition, it might be better to measure intracellular ATP using more specific
kits in the future.
Mitochondria do not seem necessary for HXR9-induced death in the tested AML cells.
Mitochondria participate in apoptosis and some forms of necrosis. In apoptosis, it is involved
in caspase- and Bax-dependent pathways, yet neither pathway was essential in HXR9-induced
AML cell death (Martin 2010; Gogada, Prabhu et al. 2011). In necrosis, mitochondria are
indispensable for CypD- and PARP1-dependent necrosis, but neither CypD nor PARP1
activation was involved in HXR9-induced AML cell necrosis (Andrabi, Dawson et al. 2008;
Elrod and Molkentin 2013) .
HXR9 caused cell death in c-FOS-dependent pathway, at least in part, in previously
tested cell lines that derived from different kinds of solid cancers (Morgan, Pirard et al. 2007;
Shears, Plowright et al. 2008; Plowright, Harrington et al. 2009; Morgan, Plowright et al. 2010;
Morgan, Boxall et al. 2012; Morgan, Boxall et al. 2014). Likewise, c-FOS up-regulation was
seen in the tested AML cell lines. In an attempt to understand more about the molecular
mechanism that is involved in HXR9 cell killing, MAPK pathways were studied using
pharmacological inhibitors to block their potential role in HXR9 treatment. It was found that
HXR9 induced necrosis of the tested AML cells in a MAPK-independent pathway. As Ca2+ is
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a main inducer of necrosis, the role of some of the Ca2+ downstream effectors that might
regulate HXR9-induced necrosis was also analysed. For example, μ-Calpain was not involved
in HXR9-induced necrosis. Interestingly, the other analysed Ca2+ downstream mediators
including NOXs, CaM, PKC and HO-1 had an anti-necrotic effect, as their inhibition
accelerated the necrosis of the tested AML cells. In addition, it was found that c-FOS induced
p21 expression in p53-independent pathway.
The anti-necrotic effect of NOXs, CaM, PKC and HO-1 after HXR9 treatment for AML
cells might direct the selection of combination drugs that would be used in the future with
HXR9. For example, cisplatin was reported to result in cell apoptosis through pathways that
involve the activation of PKC, NOXs and JNK (Persaud, Hoang et al. 2005; Kim, Lee et al.
2010; Itoh, Terazawa et al. 2011; Pabla, Dong et al. 2011; Pereira, Igea et al. 2013). In addition,
c-FOS over-expression was highly associated with cisplatin resistance (Muscella, Urso et al.
2009). Likewise, the alkylating agents treosulfan and etoposide were reported to induce
apoptosis in a PKC-dependent manner (Schmidmaier, Oellerich et al. 2004; Day, Wu et al.
2009; Zhao, Duan et al. 2009). Therefore, cisplatin, treosulfan and etoposide might have
antagonistic effects on HXR9 treatment. However, drugs that inhibit NOXs, CaM, PKC and
HO-1 might produce synergistic effect upon combination with HXR9.
ROS seems to exert an anti-necrotic effect in HXR9 treated AML cells. The primary
sources of ROS are mitochondria, NOXs, and some enzymes and growth factors (Festjens,
Vanden Berghe et al. 2006; Morgan, Kim et al. 2008; Jiang, Zhang et al. 2011). The results of
this study suggested that mitochondria are not required for HXR9 cytotoxicity. However,
inhibition of NOXs enhanced the efficacy of HXR9. In addition, the results of this study showed
that inhibition of PKC, which is an upstream regulator for HO-1 and JNK pathway and
downstream effector for ROS also augmented the effect of HXR9 (Liu, Nishitoh et al. 2000;
Kadowaki, Nishitoh et al. 2005; Tsoyi, Jang et al. 2011). Therefore, HXR9 might increase the
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expression of anti-oxidant transcription factors and it will be informative to investigate their
expression and measure ROS levels after HXR9 treatment.
The in vitro results were promising and were followed up by in vivo models to assess
the efficacy of HXR9 in inducing the necrosis of AML cells. In addition, one of the aims of this
study was to assess the cytotoxicity of HXR9 on normal PB; subsequently, the mouse AML
cell line C1498-GFP and the C57BL/6 mouse strain that has a full immune system were used.
However, using both this strain and a nude mouse strain for I.V. injection of cell lines and
HXR9 had several limitations. There was a low engraftment rate of AML cells, which could be
a result of multiple factors. These include the presence of either a full or partial immune system
in C57BL/6 and nude mice, respectively. This limitation could be overcome by eradicating the
immune system by pre-treating mice with immunosuppressive agents and total body irradiation.
Alternatively, the most common mouse strains for in vivo AML models that are characterised
by the lack of the immune system are NOD/SCID or NOD/SCID gamma mice that have a high
rate of engraftment could be used (Shan and Ma 2013). The low engraftment rate could also be
explained by the low number of injected cells or a failure to inject cells in tail vein. Another
limitation is the presence of extramedullary infiltration that could be monitored using a
bioluminescence imaging system.
The S.C. model of AML cells is an acceptable model for preclinical evaluations of new
agents, but it does not recapitulate the BM microenvironment. Recent studies demonstrated
multiple differences among different stromal cells originating from different organs. The
differences include phenotypes, proliferation, the expression levels of SDF-1, VEGF and
TGFβ1 (Dmitrieva, Minullina et al. 2012; De Luca, Verardi et al. 2013). As stromal cells play
an integral part in chemoresistance, this difference in tumour microenvironments should be
considered in terms of assessing the in vivo efficacy of HXR9 to induce AML necrosis.
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Despite the heterogeneity of the S.C. and BM tumour microenvironments, the in vivo
chemoresistance to HXR9 could be a result of cell intrinsic or extrinsic factors. Therefore, the
potential causes of chemoresistance should be studied in order to improve the in vivo efficacy
of HXR9. Chemoresistance studies could be performed using a short-duration, ex vivo/co-
culture model, and could allow the role of stromal cells in HXR9 resistance to be determined.
Possible mechanisms of resistance could include an interaction with AML cells, and or the
expression of specific cytokines, growth factors and matrix proteins (Garrido, Appelbaum et al.
2001; Konopleva, Konoplev et al. 2002; van Gosliga, Schepers et al. 2007; Schuringa and
Schepers 2009; Civini, Jin et al. 2013; Lopez, Garcia et al. 2014).
The specificity of HXR9 to target HOX-PBX interaction was previously studied
(Morgan, Pirard et al. 2007; Plowright, Harrington et al. 2009). However, one of the main
limitations of CPPs, which is the R9 sequence in HXR9 peptide, its non-specific uptake. In
other words, CPPs are able to penetrate all tissues and organs, regardless of the administration
route. This drawback can be overcome by binding targeting peptides with an R9 sequence that
will produce a selective delivery system. For example, CXCR4 that is over-expressed in many
cancers including AML can be targeted by a ligand that is bound to CPPs in order to inhibit the
non-specific interaction of CPPs. This approach was done by attaching CXCR4 ligand to TAT,
a CPP, and the anticancer peptide p53-activating peptide that led to increase the specificity of
the delivery system and the efficacy of the anticancer peptide comparing to unguided CPP
(Snyder, Saenz et al. 2005). Analogously, the specificity can be established by creating
activatable CPP (ACPP) that constitute of a polycationic CPP linked to a neutralising polyanion
motif via a cleavable linker. This structure blocks the electrostatic interaction of the ACPP with
the negative charge cell surface by decreasing the charge of ACPP to approximately zero and
ultimately blocks uptake into cells. Proteolysis of the cleavable linker by a specific protease
releases the polycationic ACPP domain and enables it to enter cells. Many cancer associated
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proteases are known, which are highly expressed in cancer tissues, but are absent or expressed
at low concentrations in normal tissues such as metalloproteases (Vives, Schmidt et al. 2008;
Regberg, Srimanee et al. 2012). For example, binding doxorubicin to ACPP that was sensitive
to metalloproteases 2/9 effectively decreased the toxicity of doxorubicin and inhibited the
proliferation of HT-1080 cells (Shi, Gao et al. 2012).
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6.2 Conclusion
In vitro results showed that HOX-PBX dimer might be a potential therapeutic target in
a subset of AML that over-express HOX genes. This may potentially help development
of personalised medicine for AML disease. The comprehensive analysis of the
mechanism of HXR9 cytotoxicity may guide in the selection of the potential combination
drugs. For example, agents that cause NOX, CaM, PKC activation should be avoided and
vise versa. A proper in vivo systemic model should be used to assess the in vivo efficacy
of HXR9 cytotoxicity, along with conducting more in vitro assays to investigate the
mechanism of HXR9 in order to choose potential synergistic agents.
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6.3 Future directions
Combination therapy is becoming increasingly important in cancer treatment. The
findings of this study reveal that HXR9 is able to trigger the necrosis of K562 and KU812F
cells that express the BCR-ABL fusion protein; and HL-60 that expresses PML-RARA. It may
therefore be valuable to assess combination therapy with HXR9 and first line treatment drugs
such as imatinib and ATRA that target BCR-ABL and PML-RARA fusion proteins,
respectively.
Resistance to cell killing by HXR9 may involve anti-necrotic proteins, and thus agents
that target such proteins are potential synergistic agents with HXR9. For example, tamoxifen
and trifluoperazine, CaM antagonists, might be used in a combination with HXR9 (Ahn, Pan et
al. 2003; Wang, Li et al. 2010). Likewise, enzastaurin that inhibits PKC activity and increased
p21 levels in multiple myeloma cells could have potential synergistic effect with HXR9 (Raab,
Breitkreutz et al. 2009). The HO-1 inhibitors, pegylated zinc-protoporphyrin and styrene-
maleic-acid-copolymer-micelle-encapsulated particles, which showed apoptotic effects in
AML cell lines are also potential synergistic drugs with HXR9 (Herrmann, Kneidinger et al.
2012). Arsenic trioxide exerts its cytotoxicity by inhibiting the JNK pathway thereby increasing
the activity of c-FOS and p21 proteins; it will also be valuable to assess its combinatory effect
with HXR9 (Huang, Liu et al. 2006; Liu and Huang 2008).
Silencing c-FLIP in combination with HXR9 treatment seems an interesting
combination to perform since c-FLIP is negatively regulated by c-FOS (Zhang, Zhang et al.
2007). In addition, c-FLIP is a downstream effector for CaM and PKC (Hwang, Min et al. 2009;
Kaunisto, Kochin et al. 2009). It is also able to inhibit apoptosis and necrosis by suppressing
procaspase-8 and RIP1, respectively (Day, Huang et al. 2008; Wang, Du et al. 2008; He and
He 2013).
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The mechanism of cell killing by HXR9 requires further investigation. To understand
how RIP1 activity can cause c-FOS over-expression after HXR9 treatment, at least in HL-60
cells, the upstream regulators of c-FOS and downstream effectors of RIP1 such as apoptosis
signal-regulating kinase 1 might be involved in this mechanism (Liu, Nishitoh et al. 2000;
Vanlangenakker, Vanden Berghe et al. 2012). Such investigations may also include some c-
FOS upstream regulators, including FOS related kinase and ribosomal S6 kinase 2 (Zhang,
Zhang et al. 2007). In addition, p21 downstream effectors might be assessed. It is noteworthy
to assess the role of RIP3 in HXR9 cytotoxicity since necroptosis was involved, at least in HL-
60 cells.
A co-culture model to assess the efficacy of HXR9 in inducing AML necrosis needs to
be conducted. Such a model can mimic the BM microenvironment in in vivo models. AML cells
could be co-cultured either in direct or indirect contact with stromal cells such as human MS-5
cells to evaluate the influence of stromal cells and the secretome in the efficacy of HXR9.
A systemic in vivo model is needed to be established to assess the efficacy of HXR9 on
AML disease. In this study, the nude mouse strain was used, but the growth rate was low which
might due to the immune activity. In addition, C57BL/6 and SCID flank models do not
represent AML disease. Therefore, it is noteworthy to establish a systemic in vivo model in
other mouse mouse strains including NOG or NOD, the most common AML models, which are
characterised by the lack of T and B lymphocytes, impaired NK cells and antigen-presenting
cells (Shan and Ma 2013).
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Appendices
207
A)
B)
C)
208
D)
E)
Appendix 1. R110 standard curves for the caspase-3 assay. R110 standard curves were used as additional controls
for caspases-3 activity to show that free R110 reagent was fluorescent and its fluorescence correlated with its
concentration. A range of concentrations (5μM -25μM) of R110 were prepared by diluting 5mM R110 into 1×
reaction buffer. Then, R110 fluorescence was measured at 492nm. To validate individual experiments, R2 should
be ≥ 0.95. Graphs show the mean of three independent experiments and error bars show the SEM.
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A)
B)
Appendix 2. General inhibition of caspase activity in HXR9 treated AML cell lines by z-VAD-FMK. Cells were
pre-treated either with or without 50 μM z-VAD-FMK for one hour. Then, cells were treated with the IC50 and 2×
IC50 of HXR9 for two hours, or with 17.5 μM of DNR for 24 hours, which was used as a positive control. Next,
cells were harvested and stained with annexin V and 7-AAD for 15 minutes in the dark and analysed by flow
cytometry. There was no statistical difference in terms of sensitivity to HXR9 between pre-treated cells with or
without 50μM z-VAD-FMK. Graphs show the mean of three independent experiments and error bars show the
SEM. *p < 0.05, **** p < 0.0001 with respect to z-VAD-FMK untreated cells. A) Inhibition of caspase activity
by z-VAD-FMK in K562 and B) HL-60 cells.
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A)
B)
Appendix 3. Effect of EDTA on HXR9 cytotoxicity. Cells were pre-treated with or without 20mM EDTA for one
hour. Then, cells were treated with 10μM, 20μM, 40μM HXR9 that prepared in media either with or without
20mM EDTA or with an equivalent CXR9 concentration to the highest concentration of HXR9 for two hours. The
cells were then harvested and stained with annexin V and 7-AAD for 15 minutes in the dark and analysed by flow
cytometry. Pre- and co-treatment of cells with 20mM EDTA significantly inhibited the cytotoxicity of HXR9
compared to EDTA untreated counterparts. Graphs show the mean of three independent experiments and error
bars show the SEM. *p < 0.05, **p < 0.01, ***p < 0.001 and ****p < 0.0001 with respect to 20mM EDTA treated
cells. A) K562 cells and B) HL-60 cells.
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A)
B)
Appendix 4. The Effect of CsA on HXR9 cytotoxicity. Cells were pre-treated with or without 5μM CsA for one
hour. Then cells were treated with 1×, 2×, or 3× IC50 of HXR9, with an equivalent CXR9 concentration to the
highest concentration of HXR9, with 20mM H2O2 as a positive control, or were untreated (negative control). Next,
cells were harvested and stained with annexin V and 7-AAD for 15 minutes in the dark and analysed by flow
cytometry. Results show 5μM CsA leads to a significant decrease in % viable cells and an increase in % early
apoptotic and dead cells of HXR9 treated cells comparing to their counterparts. Graphs show the mean of three
independent experiments and error bars show the SEM. *p < 0.05, **p < 0.01, ***p < 0.001 and ****p < 0.0001
with respect to 5μM CsA untreated cells. A) K562 and B) HL-60 cells.
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A)
B)
Appendix 5. The effect of blocking the MEK/ERK pathway in H2O2 cytotoxicity. Cells were pre-treated with or
without 20μM of the MEK/ERK inhibitor U0126 for one hour. Next, cells were treated with 20mM H2O2 for two
hours and the cytotoxicity was measured by LDH assay. U0126 significantly inhibited the cytotoxicity of H2O2.
The graphs show the mean of three independent experiments and the error bars show the SEM. **p < 0.01 and
****p < 0.0001 with respect to 20μM U0126 untreated cells. A) K562 and B) HL-60 cells.
213
A)
B)
Appendix 6. The effect of inhibiting the p38 pathway on H2O2 cytotoxicity. Cells were pre-treated with or without
25 μM SB302580 for one hour, and were then treated with 20mM H2O2 for two hours. Cytotoxicity was measured
using the LDH assay. Blocking the p38 pathway significantly reduced the cytotoxicity of H2O2. The graph shows
the mean of three independent experiments and the error bars show the SEM. **p < 0.01 and ***p < 0.001 with
respect to 25μM SB302580 untreated cells. A) K562 and B) HL-60 cells.
214
Appendix 7. The impact of JNK pathway inhibition in the cytotoxicity of H2O2 in HL-60 cells. Cells were pre-
incubated either with or without 60μM SP600125 for one hour, and then incubated with 20mM H2O2 in media
supplemented with or no supplemented with 60μM SP600125 (red and blue bars, respectively). Next, the
cytotoxicity was assessed using the LDH assay. 60μM SP600125 dramatically decreased the cytotoxicity of H2O2.
The graph shows the mean of three independent experiments and the error bars show the SEM. ***p < 0.001 with
respect to 60μM SP600125 untreated cells.
0
25
50
75
100HL-60
20mM
% S
urv
ival cells
H2O2
H2O2+60M SP600125
***
215
A)
B)
Appendix 8. The effect of blocking μ-calpain on the cytotoxicity of H2O2. Cells were pre-incubated with or
without 60μM calpain inhibitor I for one hour, and then incubated with 20mM H2O2 prepared in media with or
without calpain inhibitor I (red and blue bars, respectively). The cytotoxicity of H2O2 was then measured using the
LDH assay. Blocking μ-calpain significantly inhibited the cytotoxicity of H2O2. Graphs show the mean of three
independent experiments and error bars show the SEM. **p < 0.01 with respect to calpain inhibitor I untreated
cells. A) K562 and B) HL-60 cells.
216
Appendix 9. Establishment of a systemic C1498-GFP model in C57BL/6 mice, only PB was harvested.
Harvesting day Number of mice harvested Engraftment rate
3 2 0
7 2 0
13 2 1
17 2 1
20 2 0
217
Appendix 10.
A) Establishment a systemic C1498-GFP model in C57BL/6, PB, BM, kidney, liver and spleen were harvested.
B) Establishment a systemic C1498-GFP model in C57BL/6, PB, BM, kidney, liver and spleen were harvested.
Harvesting day Number of mice harvested Engraftment rate
8 2 0
15 2 1
22 2 1
25 2 0
28 2 1
Harvesting day Number of mice harvested Engraftment rate
8 1 0
15 1 0
22 1 1
28 1 0
35 1 1
218
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