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A Novel Mechanism for Nicotinic Acetylcholine Receptor
Antagonist-Induced Cancer Cell Cytotoxicity
By Mitch Clarke
Master of Science Thesis
University of Otago, Dunedin
New Zealand
10 February 2016
ii
Abstract
Nicotinic acetylcholine receptors (nAChRs) are common ligand-activated neurotransmitter
receptors located throughout the body. The up-regulation of nAChRs subunits has been observed in
a number of cancer types; while current literature has identified various nAChR pathways that
contribute to the development of cancer. The present study identified a novel mechanism for the
observed cytotoxicity of the highly potent nAChR antagonist, pinnatoxin (PnTX), on human
epithelial carcinoma cell lines. Cell cycle and intracellular calcium studies suggest that PnTX exerts
its cytotoxic effects by inducing changes in the cell cycle via antagonism of the highly calcium
permeable α7-, and to a certain extent, the α4β2-nAChR receptors. This leads to an induction of cell
apoptosis via calcium-independent pathways. The use of nAChRs in combination with other known
cytotoxic agents, such as cyclosporin A, or apoptotic inducing chemotherapy agents, may provide a
sensitising tool that can reduce the dose required to elicit a cytotoxic response on developing
tumours.
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Acknowledgements
I would first like to thank Dr Sarah Baird for her time, guidance and support over my
extended time in her lab. As my Masters and previously Honours supervisor, she has contributed a
great deal of knowledge and encouragement throughout my time in Dunedin. Her mentorship and
independent thinking have been invaluable while keeping me motivated throughout the many
repeated experiments. Her patience and willingness to let her students investigate any idea, no
matter how lateral, will continue to be her greatest strength as an academic mentor. It has been a
pleasure working alongside Sarah and I wish her all the best for the future.
I would like to thank the fellow students of the Baird Lab, Ben, Sean, Chloe and Irene, for
their contribution to this project. They have all provided a great deal of knowledge and willingness
to learn that have kept me motivated during my research. Their readiness to lend a hand or provide
helpful suggestions in times of experimental agony has made this experience a lot more enjoyable. I
can only hope I have contributed as much to their academic progress as they have to mine.
To the members of the various other labs within the department, in particular Floriane and
Sebastien of the Rosengren Lab, thank you for your time and effort. The materials, technical
knowledge and guidance they both provided were invaluable. Furthermore, their willingness to
dedicate time at the expense of their own research was truly humbling and a reflection of what
incredible individuals they are. To Shane of the Kerr Lab, thank you for your constant supply of
experimental materials, knowledge surrounding the concepts of this project and sense-of-humour in
times of insanity. I am grateful to you all.
Finally, to the University of Otago, and more importantly the Pharmacology and Toxicology
Department, I would like to express my gratitude for allowing me to use the facilities during my
time in Dunedin. In particular, I would like to thank all the staff and students of the Pharmacology
and Toxicology for being such genuine people, while never falling short of providing a laugh at the
most stressful of times.
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Table of Contents
A Novel Mechanism for Nicotinic Acetylcholine Receptor Antagonist-Induced Cancer Cell
Cytotoxicity ........................................................................................................................................... i
Abstract ................................................................................................................................................ ii
Acknowledgements ............................................................................................................................. iii
Table of Contents ................................................................................................................................ iv
List of Tables .................................................................................................................................... viii
List of Figures ..................................................................................................................................... ix
List of Abbreviations .......................................................................................................................... xi
1 Introduction .................................................................................................................................. 1
1.1 Nicotinic Acetylcholine Receptors ........................................................................................ 1
1.1.1 Structure, Diversity and Localisation ............................................................................. 1
1.1.2 Neuronal nAChR Function ............................................................................................ 3
1.1.3 Non-Neuronal nAChR Function .................................................................................... 5
1.1.4 nAChRs in Disease ........................................................................................................ 7
1.2 nAChRs in Cancer Development .......................................................................................... 8
1.2.1 nAChR Expression in Cancer ........................................................................................ 9
1.2.2 nAChRs in Cancer Development and Proliferation ..................................................... 12
1.2.3 Role of nAChRs in Cancer Angiogenesis and Metastasis ........................................... 14
1.2.4 Anti-Apoptotic Effects of nAChRs in Cancer Survival ............................................... 16
1.2.5 nAChRs and Cancer Cell Death................................................................................... 18
1.3 nAChR-Modulators and nAChR-Based Therapies ............................................................. 19
v
1.3.1 nAChR-Based Therapies for Common Diseases ......................................................... 20
1.3.2 nAChR-Modulators for Cancer Treatment .................................................................. 22
1.3.3 Marine Toxins .............................................................................................................. 25
1.4 Aims and Hypothesis........................................................................................................... 26
2 Methods ...................................................................................................................................... 29
2.1 Cell Culture ......................................................................................................................... 29
2.1.1 Materials....................................................................................................................... 29
2.1.2 Cell Lines and Cell Culture .......................................................................................... 29
2.1.3 MTT Assay .................................................................................................................. 30
2.1.4 Statistical Analysis ....................................................................................................... 31
2.2 Cell Viability Assay ............................................................................................................ 31
2.2.1 Materials....................................................................................................................... 31
2.2.2 Cell Viability Assay ..................................................................................................... 31
2.3 Western Blot ........................................................................................................................ 32
2.3.1 Materials....................................................................................................................... 32
2.3.2 Antibodies .................................................................................................................... 33
2.3.3 Protein Extraction ........................................................................................................ 33
2.3.4 Western Blotting .......................................................................................................... 34
2.4 Immunocytochemistry ......................................................................................................... 35
2.4.1 Materials....................................................................................................................... 35
2.4.2 Sample .......................................................................................................................... 35
2.4.3 Fluorescence Microscopy ............................................................................................ 35
vi
2.5 Flow Cytometry ................................................................................................................... 36
2.5.1 Materials....................................................................................................................... 36
2.5.2 Cell Treatment and Staining ........................................................................................ 36
2.5.3 Fluorescence-Activated Cell Sorting (FACS).............................................................. 37
2.6 Intracellular Calcium ........................................................................................................... 38
2.6.1 Materials....................................................................................................................... 38
2.6.2 Fura-2 AM Fluorescence Measurement ....................................................................... 38
2.7 Cell Death Inhibitors ........................................................................................................... 39
2.7.1 Materials....................................................................................................................... 39
2.7.2 Cell Culture and Inhibitors ........................................................................................... 39
2.7.3 Isobologram ................................................................................................................. 39
2.8 Combination Therapy .......................................................................................................... 40
2.8.1 Materials....................................................................................................................... 40
2.8.2 Chemotherapy Combination Therapy .......................................................................... 40
3 Results ........................................................................................................................................ 41
3.1 Effects of nAChR Antagonists on Cell Viability ................................................................ 41
3.1.1 Cell Viability Assay ..................................................................................................... 41
3.1.2 Western Blot - Basal nAChR Levels ........................................................................... 47
3.1.3 Immunocytochemistry ................................................................................................. 48
3.1.4 Discussion .................................................................................................................... 52
3.2 Investigation of nAChR Antagonist Induced Cell Death .................................................... 57
3.2.1 Western Blot – α7 nAChR Levels following Treatment ............................................. 57
vii
3.2.2 Flow Cytometry - Cell Cycle Analysis ........................................................................ 60
3.2.3 Flow Cytometry - TMRE ............................................................................................. 66
3.2.4 Flow Cytometry - Annexin V/PI .................................................................................. 68
3.2.5 Discussion .................................................................................................................... 75
3.3 Inhibition of nAChR Antagonist Induced Cell Death ......................................................... 78
3.3.1 Cell Death Inhibitors .................................................................................................... 79
3.3.2 Intracellular Calcium .................................................................................................... 83
3.3.3 Discussion .................................................................................................................... 87
3.4 Combination Treatment ....................................................................................................... 91
3.4.1 CsA + MECA ............................................................................................................... 91
3.4.2 Chemotherapy Agents + nAChR Antagonists ............................................................. 98
3.4.3 Discussion .................................................................................................................. 104
4 Discussion ................................................................................................................................ 107
4.1 Signalling through nAChRs is linked to Cancer Development ......................................... 107
4.1.1 nAChR Agonist Induced Cancer Progression ............................................................ 107
4.1.2 nAChR Antagonist Driven Cytotoxicity .................................................................... 107
4.2 The nAChR Antagonist PnTX Induces Cytotoxicity by Altering Intracellular Calcium
Levels ........................................................................................................................................... 108
4.3 Synergistic Cytotoxic Effects of Selective nAChR Antagonists with Common
Chemotherapeutic Agents ............................................................................................................ 109
4.4 Future Directions ............................................................................................................... 109
4.5 Final Conclusion................................................................................................................ 111
5 References ................................................................................................................................ 112
viii
List of Tables
Table 1: Examples of Selective α7 nAChR-Modulators Investigated ............................................... 22
Table 2: Details of the Carcinoma Cell Lines .................................................................................... 30
Table 3: Cell Viability Assay Treatment Profile ............................................................................... 32
Table 4: Cell Death Inhibitor Treatment Profile ................................................................................ 39
ix
List of Figures
Figure 1: Protein ribbon structure of a nicotinic acetylcholine. ........................................................... 2
Figure 2: The regulation of intracellular pathways .............................................................................. 9
Figure 3: The determination of MDA-MB-231, HT29 ...................................................................... 42
Figure 4: The evaluation of MDA-MB-231, HT29 ........................................................................... 46
Figure 5: Western blot analysis showing the basal level ................................................................... 48
Figure 6: Immunocytochemistry images of non-permeabilised ........................................................ 50
Figure 7: Immunocytochemistry images of non-permeabilised ........................................................ 51
Figure 8: Western blot analysis investigating the effects .................................................................. 59
Figure 9: Cell cycle analysis of antagonist treated MDA-MB-231 ................................................... 61
Figure 10: Cell cycle analysis of treated MDA-MB-231 ................................................................... 63
Figure 11: Cell cycle analysis of treated HT29 .................................................................................. 64
Figure 12: Cell cycle analysis of treated PC3 .................................................................................... 65
Figure 13: Evaluation of the mitochondrial transmembrane ............................................................. 67
Figure 14: Flow cytometric detection of apoptotic and necrotic cells ............................................... 70
Figure 15: Flow cytometric detection of apoptotic and necrotic cells. .............................................. 72
Figure 16: Flow cytometric detection of apoptotic and necrotic cells ............................................... 74
Figure 17: Examination of the effect of the cell death inhibitor Cyclosporin A ............................... 80
Figure 18: Examination of the effect of the cell death inhibitor ALLN ............................................ 81
Figure 19: Examination of the effect of the cell death inhibitor BAPTA .......................................... 82
Figure 20: Examination of the effect of the cell death inhibitor zVAD ............................................ 83
Figure 21: Ca2+ accumulation through Fura-2 AM detection ............................................................ 86
Figure 22: The cytotoxic effect of combined MECA and CsA treatment ......................................... 92
Figure 23: Isobologram example displaying the line of additivity .................................................... 94
Figure 24: Isobologram demonstrating the relationship between MECA ......................................... 95
x
Figure 25: Flow cytometric detection of apoptotic and necrotic cells ............................................... 97
Figure 26: The cytotoxic effect of DOXO, DTX and ADI treatment ................................................ 99
Figure 27: The cytotoxic effect of DOXO, DTX and ADI treatment .............................................. 100
Figure 28: Determination of cell viability following treatment with DOXO .................................. 101
Figure 29: Evaluation of cell viability following treatment with DOXO ........................................ 103
xi
List of Abbreviations
[Ca2+] Intracellular Calcium Concentration
5-HT Serotonin or 5-hydroxytryptamine
ACh Acetylcholine
AChE Acetylcholinesterase
ADI Adiphenine
AEBSF 4-(2-Aminoethyl)benzenesulfonyl fluoride hydrochloride
Akt Protein Kinase B
ALLN N-Acetyl-Leu-Leu-Nle-al
AM Acetoxymethyl Ester
ANOVA Analysis of Variance
API Activating Protein 1
APS Alkypyridinium Salt
APS Ammonium Persulfate
BAPTA 1,2-bis(o-Aminophenoxy)Ethane-N,N,N’,N’-Tetraacetic Acid
Bcl B-cell Lymphoma Protein
BSA Bovine Serum Albumin
CA1 Cornu Ammonis 1
Ca2+ Calcium Ion
CaCl2 Calcium Chloride
cAMP Cyclic Adenosine Monophosphate
CD4+ Cluster of Differentiation 4-Positive
CD8+ Cluster of Differentiation 8-Positive
cdc42 Cell Division Control Protein 42
CDDP Cisplatin
xii
CDK4 Cyclin-Dependent Kinase 4
CDX Candoxin
ChAT Choline Acetyltransferase
CHT1 Choline Transporter 1
CNS Central Nervous System
CO2 Carbon Dioxide
COX Cyclooxygenase
CREB cAMP Response Element Binding Protein
CsA Cyclosporin A
DAPI 4,6-Diamidino-2-Phenylindole
DeN N-Nitrosodiethylamine or Diethylnitrosamine
DMEM Dulbecco’s Modified Eagle Medium
DMSO Dimethyl Sulfoxide
DNA Deoxyribonucleic Acid
DNAse Deoxyribonuclease
DOXO Doxorubicin
DTT Dithiothreitol
DTX Docetaxel
E2 Estradiol
ECL Electrochemiluminescence
EDTA Ethylenediaminetetraacetic Acid
EGCG (–)-Epigallocatechin-3-Gallate
EGF Epidermal Growth Factor
EGFR Epidermal Growth Factor Receptor
EMT Epithelial-Mesenchymal Transition
ER Estrogen Receptor
xiii
ERK Extracellular Signal-Regulated Kinases
FACS Fluorescence-Activated Cell Sorting
FAK Focal Adhesion Kinase
FBS Foetal Bovine Serum
FGF Fibroblast Growth Factor
FITC Fluorescein Isothiocynate
GABA Gamma-Aminobutyric Acid
GYM Gymnodimine
H2O Water
HBSS Hanks Balanced Salt Solution
HEPES 4-(2-Hydroxyethyl)-1-Piperazineethanesulfonic Acid
HMEC-L Human Microvascular Endothelial Cells – Lung
IAP Inhibitor of Apoptosis Protein
IC50 Half Maximal Inhibitory Concentration
ICC Immunocytochemistry
IgG Immunoglobulin G
KCl Potassium Chloride
KH2PO4 Potassium Di-Hydrogen Orthophosphate
LD10 Lethal Dose to 10% of Individuals
LPS Lipopolysaccharide
mAChR Muscarinic Acetylcholine Receptor
MAPK Mitogen-Activated Protein Kinases
MCF Michigan Cancer Foundation
MECA Mecamylamine
MEK Mitogen-Activated Protein Kinase
MeSH 2-Mercaptoethanol
xiv
MgSO4 Magnesium Sulphate
MLA Methyllycaconitine
MOPS 3-(N-Morpholino) Propanesulfonic Acid
mTOR Mammalian Target of Rapamycin
MTT 3-(4,5-Dimethylthiazol-2-yl)-2,5-Diphenyltetrazolium Bromide
Na+ Sodium Ion
Na2HPO4 Di-Sodium Hydrogen Orthophosphate
nAChR Nicotinic Acetylcholine Receptor
NaCl Sodium Chloride
NAD Nicotinamide Adenine Diculeotide
NaHCO3 Sodium Bicarbonate
NDNI N-n-Decylnicotinium Iodide
NF-κB Nuclear Factor-Kappa B
NIC Nicotine
NMDA N-methyl-D-aspartate
NNK Nicotine-Derived Nitrosamine Ketone
NNN N-Nitrosonornicotine
NOD/SCID Non-Obese Diabetic/Severe Combined Immunodeficiency
NOS Nitric Oxide Synthase
NSCLC Non-Small Cell Lung Carcinoma
PBS Phosphate Buffered Saline
PEG-PLA Poly(Ethylene Glycol)-Poly(D,L-Lactide)
PI Propidium Iodide
PI3K Phosphatidylinositol 3-Kinase
PKC Protein Kinase C
PnTX Pinnatoxin
xv
Raf Rapidly Accelerated Fibrosarcoma Protein
Ras Rat Sarcoma Protein
RNA Ribonucleic Acid
RPMI Roswell Park Memorial Institute
SCLC Small Cell Lung Carcinoma
SDS Sodium Dodecyl Sulfate
SEM Standard Error of the Mean
SLURP Secreted Ly-6/uPAR-Related Protein
SMAC Second Mitochondria-Derived Activator of Caspase
TEMED Tetramethylethylenediamine
TMRE Tetramethylrhodamine, Ethyl Ester, Perchlorate
TRAIL TNF-Related Apoptosis-Inducing Ligand
uPAR Urokinase Plasminogen Activator Receptor
VAChT Vesicular Acetylcholine Transporter
VEGF Vascular Endothelial Growth Factor
XIAP X-linked Inhibitor of Apoptosis Protein
zVAD-FMK N-Benzyloxycarbonyl-Val-Ala-Asp-Fluoromethyl Ketone
α-BTX α-Bungarotoxin
α-CbT α-Cobratoxin
1
1 Introduction
1.1 Nicotinic Acetylcholine Receptors
Nicotinic acetylcholine receptors (nAChRs) are common ligand-activated neurotransmitter
receptors located throughout the body. nAChRs respond to the binding of the endogenous
neurotransmitter, acetylcholine (ACh), causing fast ionotropic cationic transmission through the cell
membrane (Albuquerque et al., 2009). The receptors are expressed on both neuronal and non-
neuronal cells, resulting in a diverse range of functions. Consequently, the receptors are linked to
many human disease states, making them one of the most intensely researched areas of molecular
biology and therapeutics.
1.1.1 Structure, Diversity and Localisation
The acetylcholine receptor family is comprised of two major sub-groups of receptors;
nAChRs and muscarinic acetylcholine receptors (mAChRs). Unlike mAChRs that relay
intracellular messages though G-protein-coupled transmission, nAChRs gate the flow of various
ions through the cell membrane to induce a cellular response (Albuquerque et al., 2009). The ability
of nAChRs to regulate ion transmission stems from their pentagonal structure of five subunits,
arranged around a central pore. Each subtype spans the length of the cell membrane, with both the
N- and C-terminal of the subtype protein located extracellularly. The nAChR is comprised of an
arrangement of different α, β, γ, δ and ε subtypes that dictate the flow of ions through the receptor
pore (Kalamida et al., 2007).
As with all members of the acetylcholine receptor family, receptor function is regulated by the
interaction between ACh and the ligand-binding site. Binding of ACh to the receptor occurs at the
interface between two subunits, with at least one subunit residing from the α-subunit family, on the
extracellular domain of the receptor, as represented in Figure 1. ACh binding causes a
conformational shift in the pentagonal structure of the receptor as a result of changes to the van der
2
Waal interactions between the receptor subunits. Opening of the transmembrane pore allows the
movement of positively charged ions, including sodium and calcium (Ca2+) into the cell, and
potassium out of the cell, to elicit the cellular response (Miyazawa et al., 2003).
Figure 1: Protein ribbon structure of a nicotinic acetylcholine receptor, as viewed from the synaptic cleft (left) and
parallel with the cell membrane (right). The diagram displays the pentagonal structure of a common heteromeric
(α)2βγδ-nAChR, configured around a central pore with sections of each subunit located within the intracellular (I) and
extracellular (E) space. The αTrp149 ligand binding region of the α-subunit has been displayed in yellow. Images
adopted from (Unwin, 2005).
The localisation of the receptor also plays a role in the expression of specific nAChR
subunits. The sub-group of nAChRs is broadly split into two types, muscle-type and neuronal-type
nAChRs. Muscle-type nAChRs are located at the neuromuscular junction of muscle terminals,
where they play an important role in muscle contraction. Muscle-type nAChRs are comprised
strictly of a (α1)2β1εδ receptor configuration in the adult muscle (embryonic muscle contains the γ-
subunit in place of the ε-subunit, with the transition of γ- to ε-subunit expression occurring during
development) (Tzartos et al., 1998). Neuronal-type nAChRs demonstrate a higher degree of
diversity in terms of subunit expression. Receptor pentamers can be comprised of homomeric
3
configurations of α-subunits (e.g. (α7)5), or heteromeric configurations consisting of both α- and β-
subunits (e.g. (α4)2(β2)3). As a result, the function, ionic permeability and responsiveness to various
exogenous compounds of the neuronal-type nAChRs can differ greatly between receptor
configurations (Sargent, 1993).
Traditionally, the expression of nAChRs consisting of homo- and heteromeric configurations
of α- and β-subunits was thought to be localised to within the nervous system. However, recent
research has identified the expression of non-neuronal nAChRs on various cells outside of the
nervous system. Various nicotinic α- and β-subunits are expressed on non-neuronal tissues,
including endothelial cells, epithelial cells, mesenchymal cells and a variety of immune cells
(reviewed in (Arias et al., 2009). The function of the different neuronal and non-neuronal nAChRs
is discussed in more detail in the following sections.
1.1.2 Neuronal nAChR Function
Neuronal nAChRs are widely spread throughout the nervous system, including certain areas
of the brain and within the peripheral ganglia. The nAChRs are comprised of combinations of nine
α (α2-α10) and three β (β2-β4) subunits, forming operational ion channels at the pre-terminal, pre-
synaptic and post-synaptic sites of neuronal cells (Dajas-Bailador et al., 2004). Pre-synaptically,
nAChRs influence neurotransmitter release through membrane depolarisation, or by altering
intracellular Ca2+ concentration due to the high permeability of some nAChR subunit configurations
to Ca2+. Post-synaptically, nAChRs contribute to the fast excitatory transmission of signals within
the central nervous system (CNS), enhancing post-synaptic excitation. However, the post-synaptic
effects of neuronal nAChRs are thought to be less relevant to their overall function in the CNS
(Kalamida et al., 2007).
Based on the expression of particular subunits on the pre-synaptic terminal, nAChRs in the
CNS can regulate the release of particular neurotransmitters. For example, dopamine release has
been found to be regulated by the expression of α4β2 and α6β2 subtypes in the mesolimbic pathway
of the CNS (Gotti et al., 2010); while, α7 subtype expression is linked to glutamatergic
4
neurotransmission in various regions of the brain (Cui et al., 2013). The ability of nAChRs to
regulate neurotransmission is due to their ability to cause an increase in intracellular Ca2+ levels at
the pre-synaptic terminal. Non-α7 subtype receptor configurations, such as the α4β2, have relatively
low permeability to Ca2+ when activated. However, these receptors are functionally coupled to
voltage-gated calcium channels, resulting in an influx of Ca2+ into the pre-synaptic terminal
following ACh binding (Garduño et al., 2012). Alternatively, nAChRs, such as the homomeric α7
receptor, can induce glutamine release without the involvement of coupled ion channels, due to its
high permeability to Ca2+ through the central pore of the receptor once activated (Garduño et al.,
2012).
The ability of nAChR to regulate intracellular Ca2+ levels within the nervous system plays
another important role in the regulation of neuronal gene expression. The influx of Ca2+ into the cell
following nAChR activation of Ca2+ permeable subunits results in the downstream activation of
transcription factors (Nakayama et al., 2006). One example is the established role of nAChRs in
memory and plasticity events. The activation of nAChRs, especially those containing the highly
Ca2+ permeable α7 subunits, regulates the ERK/MAPK intracellular signalling cascade. The
prolonged activation of this signalling cascade results in the phosphorylation of various gene
transcription factors, such as the cAMP response element binding protein (CREB), Sp1 and c-myc.
The increase in gene transcription results in the expression of several proteins involved in memory
formation and long-term plasticity within the nervous system (Nishimoto et al., 2011). Furthermore,
the activation of nAChRs via exogenous agonists such as nicotine plays an important role in
addiction through the activation of similar intracellular signalling cascades (Changeux, 2010).
Finally, neuronal nAChR activation has been shown to be linked to neuroprotection within the
CNS, while contributing to protection against neuronal toxicity. nAChRs containing the α7 subtype
have been linked to the intracellular activation of the phosphatidylinositol 3-kinase (PI3K) cascade.
The neuronal activation of the PI3K pathway results in downstream phosphorylation of Akt, and
subsequently Bcl-2 and Bcl-xl proteins, a subset of anti-apoptotic proteins that can prevent cell
5
death during neuronal toxicity (Kawamata et al., 2012). The interaction of neuronal nAChRs
interaction with N-methyl-D-aspartate (NMDA) glutamate receptors has also been shown to protect
against Ca2+ induced neurotoxicity. NMDA receptor stimulation induces an influx in Ca2+ during
ischaemic brain injury, resulting in the over-activation of intracellular nitric oxide synthase and
subsequent glutamate-related cell death (Akaike et al., 1994). However, activation of nAChRs
during glutamate-induced neurotoxicity has demonstrated potential neuroprotective effects of the
nAChRs. Pre-treatment with nicotine, a potent nAChR agonist, prior to glutamate exposure
significantly increased cell viability in cultures of rat cortical neurons, compared to glutamate
exposure alone (Kaneko et al., 1997). The neuroprotective effects were shown to be related to the
inhibition of NOS through the activation of α4β2 and α7 nAChRs (Takada et al., 2003).
1.1.3 Non-Neuronal nAChR Function
ACh is synthesised by a large range of non-neuronal cells, with functions that differ greatly
from the neuronal neurotransmitter role of the cholinergic system. The role of ACh in non-neuronal
cells plays an intermediate role between the cells and the surrounding environment, facilitating cell
proliferation, differentiation and migration, amongst other physiological processes. Blockage of
these receptors leads to cellular dysfunction and death, highlighting their importance in normal
cellular processes of non-neuronal cells (Wessler et al., 2008). The expression of specific nAChR
subtypes on non-neuronal cells varies according to the cell phenotype and the intra- and
intercellular environment. Analogous to the presence of nAChRs in the nervous system, both
homomeric and heteromeric combinations of α- and β-nAChR subunits are expressed amongst non-
neuronal cells (Arias et al., 2009).
The major function of nAChRs in non-neuronal cells is the regulation of intercellular signal
transduction through auto- and paracrine secretions of ACh (Chen et al., 2002). One example of
non-neuronal nAChR signal transduction is the role of these receptors in epidermal cells, where the
concentration of ACh is found at levels significantly higher than in other non-neuronal regions of
the body (Grando et al., 2006). Real-time polymerase chain reaction and immunostaining analysis
6
of keratinocytes within the epidermis has eluded to the presence of α3, α5, α7, α9, α10, β2 and β4
subunits; in addition to the expression of the functional enzymes required for the synthesis and
breakdown of ACh (Kurzen et al., 2004; Nguyen et al., 2000). Autocrine and paracrine secretions
of ACh activate keratinocyte α7-nAChRs, utilising the influx of Ca2+ into the cell as a second
messenger. The downstream effect of the intracellular influx of Ca2+ results in the regulation of
cellular differentiation, such as cytoskeleton reorganisation and expression of migratory integrins.
Interestingly, the removal of the α7 signalling pathway in keratinocytes not only prevented
differentiation of these cells, but also decreased the expression of pro-apoptotic signalling
molecules, suggesting a secondary role of nAChR activation in keratinocyte apoptosis (Arredondo
et al., 2002).
Much like the role of nAChRs in epidermal signalling transduction, non-neuronal nAChRs
are critical in the formulation of new blood vessels. Non-neuronal nAChRs have been shown to
contribute to angiogenesis through the expression of homomeric α7 nAChRs and several
heteromeric nAChRs, including α3, α5, β2 and β4 subunits, on endothelial cells (Arias et al., 2009).
Stimulation of nAChRs of endothelial cells with the non-selective nAChR agonist, nicotine, induces
endothelial tube formation in vitro (Heeschen et al., 2002). Investigation into the activation of
endothelial cell nAChRs established that α7 homomeric receptors were the primary receptor
involved following activation with a α7-selective agonist, dimethoxybenzylidene (Li et al., 2006);
while tube formation was significantly reduced in the presence of the α7-selective antagonist,
methyllycaconitine (Beckel et al., 2006). Furthermore, the non-selective blockade of nAChRs with
α-bungarotoxin (α-BTX) reduces endothelial cell migration by significantly inhibiting fibroblast
growth factor (FGF) and vascular endothelial growth factor (VEGF), important factors for
regulating the adhesion between endothelial cells (Ng et al., 2007).
Research has also shown the ability of ACh to modify the immune response through the
expression of non-neuronal nAChRs on various immune cells via similar auto- and paracrine
functions. The presence of ACh during in vitro immune models has demonstrated the role of ACh
7
in the induction of CD4+ T-cell maturation and in the differentiation of CD8+ T-cells into cytolytic
T-cells (Basso et al., 2008; Zimring et al., 2005). Similarly, several nAChR subtypes have been
identified on B lymphocytes, with both α4α5β2 and α7α5β4 receptor configurations identified.
Murine knockouts of the α4, α7 or β2 subtypes showed that the nAChR subtypes are critically
involved in supporting B lymphocyte survival and cellular growth (Skok et al., 2007). Conversely,
knockdown of the α7 subunit in a murine model has been shown to increase the expression of IgG1
antibodies in response to an antigen, compared to wild-type mice, suggesting a possible immune
suppression role of nAChRs (Kawashima et al., 2007).
1.1.4 nAChRs in Disease
Due to the vast array of neuronal and non-neuronal processes that are governed by nAChRs, a
number of pathological conditions can arise when substantial alterations of the neuronal and non-
neuronal cholinergic system occur. Neuronal nAChR’s are involved in the fine control of complex
behaviours within the CNS, consequently, alterations to their function can contribute to the
development of several behavioural diseases (Gotti et al., 2006). In contrast, the role of the
cholinergic system of non-neuronal cells in disease is not well known.
Alzheimer’s disease, the most common cause of dementia, is characterised by a distinct loss
of cholinergic function within parts of the CNS. Recent research has alluded to the reduced numbers
of nAChRs in Alzheimer’s patients, with the α7 subunit being of greatest importance. Treatment
with nAChR agonists was shown to protect and delay the loss of neurons (Parri et al., 2011).
Similarly, changes in neuronal nAChR function have been associated with schizophrenia disease.
Studies investigating the post-mortem brains of schizophrenia patients have observed a decrease in
cholinergic receptor expression involving the α7 and α4β2 subunits in the hippocampal and striatal
regions of the brain (Durany et al., 2000; Freedman et al., 2000). Changes in α7 and α4β2 subunit
expression within the CNS have also been associated with Parkinson’s disease (Meyer et al., 2009),
autism (Martin-Ruiz et al., 2004) and attention-deficit hyperactivity disorder (Wilens et al., 2007),
with potential selective agonists offering potential treatment benefits in these disease states.
8
Alterations in the non-neuronal cholinergic system, such as small changes in the expression
pattern of selected subtypes, result in similar cellular stress outside of the nervous system (Wessler
et al., 2008). Studies have indicated a possible role of nAChRs in dermatitis, with an increase in
acetylcholine of 14-fold in superficial and 3-fold in the underlying portion of patients with atopic
dermatitis. In the same patients, a significant increase in choline-acetyltransferase, the enzyme
responsible for the production of ACh, was seen in the various skin and immune cells surrounding
the affected site (Wessler et al., 2003). In contrast, ACh content is substantially reduced in the
blood cells and bronchi of cystic fibrosis patients (Wessler et al., 2007). The cholinergic
dysfunction is thought to contribute to the characteristic alteration in water and ion balance during
cystic fibrosis; while agonists of the α7-nAChR have been shown to produce anti-inflammatory
effects in the epithelium of patients with cystic fibrosis (Greene et al., 2010). Finally, the
overexpression of nAChR on cancer cells stimulates tumour growth when activated in various
cancer types, with the role of nAChR in cancer development discussed in more detail in the
following section (Paleari et al., 2008).
1.2 nAChRs in Cancer Development
Cancer is defined as the uncontrolled growth of abnormal cells within the body. The process
by which healthy cells develop into malignant cancerous cells is complex, with a number of
endogenous and exogenous factors contributing to cancer development. However, recent research
has identified a range of common hallmarks acquired by cells during carcinogenesis, which allow
the abnormal cells to proliferate and spread throughout the body. In the most simplistic form, these
hallmarks include the ability of the cancer to sustain proliferation signalling, evade growth
suppressors, resist cell death, enable replicative immortality, induce angiogenesis, and activate
invasion and metastasis (Hanahan et al., 2011). The following section details the role of neuronal
and non-neuronal nAChRs in cancer development through facilitating some of these key functional
changes, with the key intracellular pathways summarised in Figure 2.
9
Figure 2: The regulation of intracellular pathways and pro-carcinogenic effects following nAChR activation.
Endogenous and exogenous ligands bind to the specific nAChRs, causing an influx of Ca2+and other cations through
the nAChRs and voltage-gated Ca2+channels. The large Ca2+influx triggers the activation of various second
messengers, resulting in angiogenesis, cell proliferation and the inhibition of apoptosis. Figure adopted from (Schuller,
2009).
1.2.1 nAChR Expression in Cancer
Similarly to the expression of nAChRs on normal healthy cells, multiple nAChR subtypes are
expressed on both neuronal and non-neuronal cancer cells. The expression pattern of these subtypes
may vary greatly between normal and cancerous cells, where variations in nAChR expression levels
facilitate cancer development and proliferation. Extensive research has been conducted on the
nAChR patterns of various lung cancer types, due to the link of nicotine exposure and lung cancer
development of smokers (Egleton et al., 2008). Comparison of the expression patterns of nAChR
10
subtypes between normal and cancerous lung cell types have revealed a significant upregulation of
the β4 subunit in non-small cell lung carcinoma (NSCLC) cells. Conversely, a decrease in α4 levels
was shown across the same comparison (Lam et al., 2007). Interestingly, an increase in the
expression of nAChRs has also been observed in lung cancer cells of non-smokers. A comparison
between NSCLCs in smokers and non-smokers showed variations in the expression patterns of
some nAChR subunits, with a higher expression of α6β3 receptors on NSCLCs of non-smokers,
compared to those from smokers (Lam et al., 2007). One hypothesis is that other exogenous
compounds act on nAChRs in a similar fashion to nicotine. N-nitrosodiethylamine (DeN), a
compound commonly found in food, contains a similar structure to ACh, with a high affinity for
both homo- and heteromeric nAChRs (Schuller, 2007). Compounds such as DeN may lead to the
non-tobacco-regulated modulations of nAChRs and development of lung cancer in non-smokers
(Niu et al., 2014).
The up-regulation of nAChRs subunits has also been observed in a number of cancer types,
with the majority of research focused on the α7 and α4β2 subtypes due to their abundance across all
cell types (Le Novere et al., 1995). Significant levels of either receptor subtype have been identified
on human mesothelioma (Trombino et al., 2004), colon (Wong et al., 2007), lung (Lam et al.,
2007), oesophageal (Arredondo et al., 2007), breast and pancreatic cells (Dasgupta et al., 2009).
Nicotine affinity studies have shown the ability of nicotine to bind with a higher affinity to α4β2
receptors, than to α7 nAChRs. The higher affinity of nicotine to α4β2 receptors causes a long-term
desensitisation of the receptor following chronic exposure, while having no effect on the sensitivity
of the α7 subtype (Kawai et al., 2001). Consequently, the biological function of the α7 nAChRs are
increased following nicotine exposure, resulting in an increase in cancer cell proliferation due to the
pro-carcinogenic effects of α7 nAChRs. Alternatively, the α4β2 receptor is mainly involved in
inhibitory cellular functions, with desensitisation of the receptor contributing to an environment
ideal for cancer development and proliferation (Schuller, 2009).
11
Interest surrounding the overexpression of another nAChR subtype, the α9-subunit, has grown
recently with the subunit being linked to the development of breast cancer. Microdissected breast
tumours of 50 human breast cancer samples have shown an average 7.84-fold increase in α9-
nAChR mRNA expression in 67.3% of the samples tested. In vivo studies inducing overexpression
of the α9-nAChR of MCF-10A cell xenografts in nude mice have revealed a substantial increase in
tumour growth following treatment with nicotine (mean 84% increase in tumour size) (Lee et al.,
2010). The up-regulation of the α9-nAChR expression in breast cancer has been linked to the
activation of the estrogen receptor (ER). Prolonged exposure to estrogen and nicotine are known
risk factors for the formation of breast cancer (Daniell, 1980). Recent studies have shown the ability
of nicotine to act in a similar fashion to estrogen, binding to the ER and inducing gene transcription.
The activated ER, following nicotine or estrogen exposure, can specifically bind the activating
protein 1 (API)-binding site of the α9-nAChR gene, inducing transcription and overexpression of
the α9-nAChR. The results indicate that the ER-induced α9-nAChR overexpression may play a
central role in the association between nicotine or hormonal exposure and human breast cancer
formation (Lee et al., 2011).
The expression of nAChRs in cancers of the CNS is less characterised than the expression
on non-neuronal cancer types. The expression of nAChRs on normal healthy neuronal cells is well
known, with multiple subtypes expressed, playing an important part in many physiological
processes (see Neuronal nAChR Function) (Kalamida et al., 2007). One common form of CNS
cancer is glioblastoma multiforme, an aggressive type of malignant glial cancer originating from
healthy glial cells that comprise the supportive tissue within the brain (Parsons et al., 2008).
Isolation of various glioblastoma cell types, including U87, LN18 and U373 cells, have allowed for
effective in vitro studies into the characteristics of this aggressive tumour. However, due to the
difficulty of delivering anti-cancer drugs through the blood brain barrier and the widespread role of
nAChRs in many other normal CNS functions and pathological diseases, only limited investigations
into the role of nAChRs in the development of glioblastoma have been conducted. A study
12
undertaken by (Zhan et al., 2011), has exploited the expression of nAChRs on U87 cells to facilitate
the delivery of the chemotherapy agent paclitaxel to glioblastoma cells. Poly(ethylene glycol)-
poly(D,L-lactide) (PEG-PLA) micelles encapsulating paclitaxel were coupled with candoxin
(CDX), a selective α7-nAChR antagonist, and injected intravenously into nude mice bearing
intracranial U87 glioblastoma. The paclitaxel-encapsulated PEG-PLA micelles significantly
prolonged average survival time by 7 days when coupled with CDX, compared to PEG-PLA-
Paclitaxel treatment alone. The study demonstrated the possible mechanism for exploiting
differential expression of nAChRs on the surface of glioblastoma cancer cells (Zhan et al., 2011).
1.2.2 nAChRs in Cancer Development and Proliferation
One of the most fundamental characteristics of cancer cells to develop and grow is their
ability to sustain rapid proliferation. Normal tissue cells regulate the production and release of
various growth factors that ensure controlled progression through the cell cycle, limiting tissue
growth. However, cancer cells lose the ability to regulate these growth factors, resulting in
sustained and uncontrolled proliferative signalling (Hanahan et al., 2011). nAChRs have been
shown to aid in the proliferation of cancer cells through changes in receptor expression and
downstream signalling pathways of various receptor subtypes that contribute to the loss of
homeostatic regulation of cell growth.
Investigation into the proliferative signalling of nAChRs in cancer cells using potent nAChR
agonists, such as nicotine-derived nitrosamines, has revealed several pathways important for tumour
growth. Selective binding of nitrosamines to α7-nAChRs of pulmonary neuroendocrine cells and
small cell lung carcinoma (SCLC) cells activates multiple intracellular signalling messengers,
including protein kinase C (PKC), Raf1, the mitogen-activated kinases ERK1 and ERK2, and
several transcription factors (FOS, JUN and MYC) (Jull et al., 2001). The transcription factors
FOS, JUN and MYC induce transcription of specific promoter elements located on proliferative
genes, such as c-erbB-1 (Volm et al., 1998). The proliferative effects following nAChR activation
were significantly reduced through the use of a selective α7-nAChR inhibitor, Ca2+ channel
13
blockers or pharmacological inhibition of the ERK1 and ERK2 kinases, further validating the
central role of nAChRs in cancer cell proliferation (Sheppard et al., 2000; Xu et al., 2004).
Similar proliferative pathways related to nAChR-activation have been identified in other
cancer models. Studies using NSCLC and other lung epithelial carcinomas have shown the
nitrosamine-induced activation of the PI3K-Akt pathway, resulting in the phosphorylation of the
proliferation specific p70S6-kinase messenger. The effects on cancer cell proliferation were
comparable between selective agonists of nAChRs containing α3, α4 or α7 subunits (West et al.,
2003). Furthermore, the activation of nAChRs in breast cancer cell lines, MCF10A and MCF7 cells,
enhances the activity of PKC-α and cell division control protein 42 (cdc42) involved in cellular
proliferation and migration (Guo et al., 2008). While studies have suggested that the nAChR-
proliferative effects on breast cancer cells are dependent on the expression of the α9 subunit (Chen
et al., 2011). Interestingly, the desensitisation of α4β2 receptors during prolonged exposure to
nicotine or estrogen, as previously discussed, contributes to the proliferative state of a cancer cell.
Studies have shown the ability of the α4β2-nAChR to regulate the release of gamma-aminobutyric
acid (GABA), an important neurotransmitter with tumour suppressing functions. A reduction in
GABA release has been demonstrated in nitrosamine-stimulated pulmonary adenocarcinoma cells,
suggesting a link between α4β2 desensitising and reduced GABA expression (Schuller et al., 2008).
Furthermore, reduced GABA levels have been observed in the brains of smokers (Epperson et al.,
2005); while a reduction in the RNA levels of the α4 subunit gene have been shown in pulmonary
adenocarcinoma cells (Lam et al., 2007).
nAChR signalling can influence cancer cell proliferation through synergistic functions with
various growth factors. Stimulation of nAChRs of bovine adrenal pheochromatocytes induces the
production and release of FGF-2, a mitogenic neurotrophic protein that controls the development
and plasticity of various neuronal cell types. Induction of the FGF-2 growth hormone is controlled
through the cAMP/PKC signalling pathway, a common cascade linked to nAChRs and cancer cell
growth; while selective inhibition of the α7-nAChRs significantly reduces FGF-2 expression
14
(Brown et al., 2012; Moffett et al., 1998). The activation of nAChRs by nicotine has also been
shown to increase the expression of epidermal growth factor receptor (EGFR), which when
activated by extracellular growth factors, signals cancer cell proliferation. Nicotine treatment of
MDA-MB-231 breast cancer cells was shown to stimulate EGFR expression, via the ERK1/2
signalling pathway, and promote breast cancer growth (Nishioka et al., 2011). Likewise, nicotine
induced glioma cell growth and colony formation in U87 and GBM12 cell lines through the
increased phosphorylation of EGFR, AKT and ERK. The proliferative effects were reduced
following the inhibition of the EGFR, PI3K and MEK pathways, suggesting the importance of
nAChRs in cancer cell sensitisation to extracellular growth signals (Khalil et al., 2013).
1.2.3 Role of nAChRs in Cancer Angiogenesis and Metastasis
Analogous to normal tissues, tumours require constant access to nutrients and oxygen in order
to proliferate. The physiological process of angiogenesis in which new vasculature is formed
provides the required circulation for a tumour to grow. During controlled processes, such as wound
healing, the angiogenic response is transiently activated; however, during tumour progression, the
process of angiogenesis is poorly regulated. A phenomenon called the ‘angiogenic switch’ occurs
where prolonged and sustained signalling for the development of new vasculature within the tumour
microenvironment results in continuous vessel formation (Hanahan et al., 1996; Hanahan et al.,
2011). Similar to the regulation of apoptosis, angiogenesis is thought to be regulated by the
counterbalance of pro- and anti-angiogenic factors. Several angiogenic inducers have been
identified in the process of both healthy and tumour-associated angiogenesis. VEGF, a key player in
the initiation of new blood vessel formation, is up-regulated in response to hypoxia and oncogene
signalling. In addition, the pro-angiogenic factor, FGF, has been linked to sustaining tumour
angiogenesis during prolonged expression (Baeriswyl et al., 2009; Ferrara, 2009; Hanahan et al.,
2011).
The role of nAChRs in both normal physiological angiogenesis and tumour-associated
angiogenesis is well established. Studies inducing nAChR activation through nicotine
15
administration in both the Lewis lung cancer model and colon cancer xenografts in mice have
demonstrated an increase in tumour vascularity, resulting in accelerated tumour growth (Heeschen
et al., 2001; Natori et al., 2003). Mecamylamine has been shown to partially inhibit the nicotine
induced angiogenic response and return capillary density to control levels, suggesting the
importance of the α7-subunit (Zhu et al., 2003). The pro-angiogenic effects mediated by the α7-
nAChR are linked to the induction of VEGF, involving various nAChR-linked signalling pathways.
nAChR activation significantly increases VEGF expression by >2-fold in NSCLC cells (A549 and
H157) in the presence of nicotine. The pharmacological blocking of various nAChR-mediated
signalling pathways, including Ca2+/calmodulin, c-Src, PKC, PI3K/Akt, MAPK/ERK and mTOR,
significantly reduced the nicotine induced up-regulation of the pro-angiogenic factor (Zhang et al.,
2007). Comparable effects have also been observed in gastric cancers by up-regulating both COX2
and VEGF-receptors in response to nAChR stimulation (Shin et al., 2005).
Once established, malignant tumour cells begin a process of local invasion and formation of
distant colonies, termed metastasis. Cancer cells of the primary tumour follow a sequence of key
events that characterise the process of metastasis. First, the cells invade the surrounding tissue, and
enter the microvasculature of the lymph and blood systems (intravasation). The cancer cells then
migrate from the bloodstream into distant tissues (extravasation), where the cells must adapt to the
new environment in order to form new colonies and proliferate (Chaffer et al., 2011). An integral
process in tumour metastasis is the ability of cancer cells to undergo the epithelial-mesenchymal
transition (EMT). EMT is a mechanism by which cancer cells lose their endothelial characteristics
and develop a more migratory mesenchymal phenotype, stimulating invasion and migration. The
initiation of EMT involves the action of several transcription factors that regulate genes involved in
the migratory process, allowing cancer cells to down-regulate adhesive junctions with neighbouring
cells and intravate the local vasculature (Grando, 2014).
Signalling through α9-containing nAChRs is critical in the migration of several cancer cell
types. The expression of the α9 subunit in breast cancer cell lines, MCF-7 and MDA-MB-231,
16
increases cell migration in response to nicotine; while the effects are blocked through the selective
inhibition of the α9 subunit both in vitro and in vivo (Hung et al., 2011; Lee et al., 2010). The
stimulation of α9-containing nAChRs results in the phosphorylation of several adhesion and
intercellular junction proteins, such as FAK, paxillin and β-catenin, key steps in the EMT
(Chernyavsky et al., 2007). Furthermore, the proinvasive effects of nAChRs have been linked to
α7-subunit expression on NSCLCs, breast cancer and pancreatic cancer cell lines. Nicotine
stimulation of the receptor facilitated cellular invasion across all cell lines, and most importantly
induced changes in gene expression consistent with the EMT. The reduction in epithelial markers,
such as E-cadherin, and the increased expression of mesenchymal proteins, such as vimentin and
fibronectin, in response to nAChR stimulation, further validate the role of nAChRs in the EMT and
tumour metastasis (Dasgupta et al., 2009).
1.2.4 Anti-Apoptotic Effects of nAChRs in Cancer Survival
Apoptosis serves as a natural barrier against the development of cancerous cells. Some of the
notable signals that induce apoptosis include uncontrolled cellular signalling and DNA damage,
both of which are essential precursors to the development of cancer. The intracellular activation of
apoptosis is controlled by the balance of pro- and anti-apoptotic regulator messengers, that when
altered towards a pro-apoptotic state, induce a cascade of downstream effectors that result in
programmed cell death. One family of anti-apoptotic molecules, the Bcl-2 family, are of particular
relevance to cancer. Bcl-2 acts as an inhibitor of apoptosis, by binding to and suppressing the
effects of the pro-apoptotic proteins Bax and Bak. When activated, the Bax and Bak proteins disrupt
the outer mitochondrial membrane, causing the release of pro-apoptotic factors, such as cytochrome
c, that induce the cascade of proteolytic caspases, resulting in apoptosis. Overexpression of Bcl-2 in
cancer cells inhibits the downstream effector molecules of apoptosis, thus allowing the cancer cells
to continue to proliferate (Adams et al., 2007; Hanahan et al., 2011).
Recent studies into the effects of nicotine on human MCF10A and MDA-MB-231 breast
cancer cells have shown an association between nAChR activation and Bcl-2 expression. Treatment
17
of breast cancer cells with 0.5 μM nicotine for 48 hours produced a significant 2-fold increase in
Bcl-2 expression, coinciding with a 2-fold increase in cell colony formation. Interestingly, the
increase in expression of Bcl-2 was inhibited by the co-administration of KP372-1, a selective Akt
inhibitor. The inhibition of Bcl-2 up-regulation in the presence of nicotine was again reflected in the
number of cancer cell colonies, with the number of colonies returning to baseline in the presence of
KP372-1. The results demonstrated that persistent activation of nAChRs induces an up-regulation in
Bcl-2, enhancing cancer cell survival through the induction of anti-apoptotic pathways, with the
effects regulated by the intracellular messaging of the Akt pathway (Nishioka et al., 2011).
Additional studies have further added to the role of nAChR in inducing Bcl-2 expression, with
nicotine treatment significantly prolonging cell survival and protecting breast cancer cells against
doxorubicin induced cell death through mechanisms involving Bcl-2 phosphorylation (Mai et al.,
2003).
Similar increases in Bcl-2 expression of various other cancer cell types have been observed
following nAChR activation. Human colorectal cancer cell lines treated with 1 μM nicotine induced
a significant increase in phosphorylated Akt and Bcl-2. Further investigation into the specific
nAChR subtypes involved in this process found that co-treatment with α-Bungarotoxin, a selective
α7-nAChR antagonist, inhibited both cell growth and the anti-apoptotic pathways (Cucina et al.,
2012). Similarly, the anti-apoptotic activity of human bronchial epithelial cells is thought to be
mediated via the expression of α3-, α4- and α7-containing nAChRs, resulting in the downstream
activation of the Akt pathway (West et al., 2003). Exposure of lung epithelial and cancer cells to
nicotine and nicotine-derived nitrosamines promotes cell survival through the increased expression
of Bcl-2, complemented by an increased resistance to cisplatin-induced apoptosis. Comparable
effects were observed between the healthy lung epithelial cells and the cancer cells, further
consolidating the idea that nAChRs play a key role in the development of cancer through the
induced expression of anti-apoptotic factors (Nishioka et al., 2010).
18
1.2.5 nAChRs and Cancer Cell Death
The anti-apoptotic effects of nAChRs in cancer survival are well defined in current literature.
nAChR stimulation during cancer development induces the Akt signalling pathway, increasing the
phosphorylation of the known anti-apoptotic messenger, Bcl-2. These anti-apoptotic effects of
nAChRs expressed on various cancer cell types have been discussed. However, the relationship
between nAChR blockade via antagonists and the apoptotic activity of cancer cells is less well
defined by the current literature.
Recent studies have demonstrated that various lung cancer cell lines express all the functional
components of the ACh autocrine pathway, including acetylcholinesterase (AChE), choline
acetyltransferase (ChAT), vesicular acetylcholine transporter (VAChT) and choline transporter 1
(CHT1). The complete functional loop of ACh signalling allows cancer cells to promote
proliferation through the autocrine activation of the nAChRs (Lau et al., 2013; Song et al., 2008).
Lau and colleagues investigated the possible pro-apoptotic effects of inhibiting the ACh autocrine
pathway in bronchioalveolar carcinoma cells. Initially, A549 and H358 cells treated with 100 nM
nicotine demonstrated a significant increase in cancer cell ACh secretion, through the up-regulation
of VAChT and ChAT, acting specifically through the α7-, α3β2 and β3-containing nAChR
subunits. The increase in AChR-related enzymatic activity and cell signalling coincided with an
increase in bronchioalveolar carcinoma cell proliferation. The carcinoma cells were then co-treated
with 50 μM vesamicol, a VAChT antagonist, to investigate whether the nicotine-ACh induced
growth of human bronchioalveolar carcinoma cells could be alleviated. Vesamicol caused
significant apoptosis in A549 and H358 cells, with a 2.5 to 3-fold increase in apoptosis relative to
nicotine-only treated carcinoma cells. The pro-apoptotic effects of vesamicol were linked to a
significant decrease in phosphorylated Akt. The findings can be explained by decreasing nAChR
activation swaying the balance of apoptotic mediators towards a pro-apoptotic state, thus inducing
cancer cell death (Lau et al., 2013).
19
The homomeric α7 nAChR is of particular interest given its abundant expression and well-
established role in the activation of signalling transduction pathways with anti-apoptotic and
proliferative effects. APS8, a polymeric alkypyridinium salt, was investigated by (Zovko et al.,
2013) for its ability to induce NSCLC cell apoptosis through selective antagonistic binding to α7
nAChRs. APS8 showed a concentration dependent decrease in lung cancer cell viability (IC50 =
~370 nM), while having little effect on normal fibroblast cells. The decrease in cancer cell viability
was correlated with a prominent induction of apoptotic morphology, with ~40% of cells found to be
apoptotic following exposure to 500 nM of APS8 for 48 hours. Radioligand displacement studies
demonstrated that the pro-apoptotic effects were mediated via antagonising the α7 nAChRs through
displacing the specific binding of 125I-α-BTX to the receptors. The effects of APS8 on a number of
apoptotic proteins was also investigated using a Human Apoptosis Antibody Array. APS8 induced
the up-regulation of several pro-apoptotic proteins, including bad, bax, cytochrome C, SMAC,
TRAIL-R1, Fas, and various caspases, while down-regulating anti-apoptotic proteins, such as Bcl-2
and Bcl-W. The ratio between the levels of bax and Bcl-2 proteins determines whether apoptosis
will be induced, with an increase in the bax/Bcl-2 ratio causing a switch towards mitochondrial
membrane permeabilisation and activation of apoptosis. Therefore, the change in bax and Bcl-2
expression following α7 nAChR antagonism may facilitate the observed APS8 induced cancer cell
death (Zovko et al., 2013). Given the effects of agonists and antagonists of nAChR in controlling
cellular growth, they are being explored as potential therapeutics for a number of diseases.
1.3 nAChR-Modulators and nAChR-Based Therapies
An improved understanding of various neurological and inflammatory diseases has implicated
the potential role of nAChRs in these conditions. Furthermore, the differences in the expression
pattern of the various nAChR subtypes within a disease state allows for selective pharmacological
targeting of these receptors. The following section explores some of the nAChR-modulators being
developed to target a number of nAChR-associated diseases.
20
1.3.1 nAChR-Based Therapies for Common Diseases
Agonists of α7-nAChRs have been identified as pharmacologically relevant compounds for
the treatment of several neuronal or inflammatory associated diseases. Examples of nAChR-
modulators currently under investigation for use in various disease states, as reviewed by (Pohanka,
2012), are presented in Table 1. The association between α7 nAChRs and the accumulation of
Amyloid β peptide within the CNS of Alzheimer’s patients has been well characterised. The
development of agonists to target the nAChR subtype has shown promise in the treatment of
Alzheimer’s disease by inducing neuroprotection and slowing disease progression (Lombardo et al.,
2014). A potent α7 nAChR agonist, SEN1233, is of current interest due to its ability to cross the
blood brain barrier when administered orally, thus alleviating some of the common problems
associated with current CNS-associated treatments. ABT-107, another agonist of α7 nAChR has
shown promise in the treatment of Alzheimer’s disease through protecting rat cortical cultures from
glutamate induced toxicity (Pohanka, 2012).
The use of α7 nAChR modulators in the treatment of schizophrenia is another avenue of
current research. Reduced levels of α7-subtype expression have been observed in the post-mortem
brains of schizophrenic patients (Martin-Ruiz et al., 2003). GTS-21, a partial α7 nAChR agonist, is
one of the most commonly explored treatments for schizophrenia. Clinical studies of GTS-21 have
reported improvements in neurocognitive function and gating of P50 auditory evoked potentials in
schizophrenic patients (Olincy et al., 2006). Promising results have also been observed in
schizophrenic models with another full agonist of the α7 nAChRs, TC-5619; however, these
compounds currently have limited clinical ability (Lieberman et al., 2013). Nevertheless, studies
demonstrating an improvement of cognition in schizophrenic patients following nicotine treatment
reinforce the possibility of developing selective nAChR treatments (Young et al., 2013).
The development of compounds that target the anti-inflammatory pathways of the cholinergic
system is another emerging area of therapeutic drug research. Several diseases associated with an
overstimulation of the immune system have led to the theory that pharmacological targeting of the
21
cholinergic anti-inflammatory pathways may alleviate disease symptoms and progression (Pohanka,
2012). The GTS-21 compound, currently being investigated in the treatment of schizophrenia, also
has implications in preventing inflammatory injury through reducing TNF-α and the production of
other pro-inflammatory cytokines (Rosas-Ballina et al., 2009). Vc1.1, an α9-nAChR modulator, has
demonstrated clear analgesic effects in a variety of animal pain models with encouraging pre-
clinical results (Livett et al., 2006). Another compound, CNI-1493, is a potential pro-inflammatory
cytokine inhibitor via its selective binding to the α7 nAChR. CNI-1493 has been investigated
clinically for use as a treatment in Crohn’s disease, with chronic dosing having positive effects on
disease progression (Dotan et al., 2010). Selective α7 nAChR agonists may also have implications
in other clinical conditions of the digestive tract, such as in the treatment of postoperative ileus. The
α7 agonist, AR-R17779, potently prevents postoperative ileus in mice, restoring the propulsive
function of the entire gastrointestinal tract following abdominal surgery (The et al., 2007). Although
considerable promise with α7 nAChR compounds has been demonstrated in the laboratory, there
are currently no clinically approved drugs that target the cholinergic system of neurological and
inflammatory pathways.
22
Table 1: Examples of Selective α7 nAChR-Modulators Investigated for the Treatment of Various
Disease States
Name Structure Use
ABT-107 5-(6-[(3R)-1- azabicyclo[2.2.2]oct-3-
yloxy]pyridazin-3-yl)-1H-indole
Tested for the treatment of Alzheimer’s
disease and cognitive deficits associated
with schizophrenia, not commercially
available.
SEN12333 5-morpholin-4-yl-pentanoic acid (4-pyridin-3-
yl-phenyl)-amide
TC-5619
N-[(2S,3S)-2-(pyridin-3- ylmethyl)-1-
azabicyclo[2.2.2]oct- 3-yl]-1-benzofuran-2-
carboxamide
Vc1.1 Peptide structure similar to α-conotoxin Potential analgesic effect for the
treatment of neuropathic pain.
CNI-1493
N,N′-bis[3,5-bis[N-
(diaminomethylideneamino)-
Cmethylcarbonimidoyl]phenyl] decanediamide
tetrahydrochloride
Investigated under clinical trials (as
Semapimod) for its ability to inhibit
inflammation and NO production.
GTS-21
3-[(3E)-3-[(2,4-
dimethoxyphenyl)methylidene]- 5,6-dihydro-
4H-pyridin-2-yl] pyridine
Under clinical testing for the treatment
of Alzheimer’s, cognitive deficits
associated with schizophrenia and anti-
inflammatory properties.
AR-R17779 (2S)- 2′H-spiro[4-azabicyclo[2.2.2]octane-2,5′-
[1,3]oxazolidin]-2’-one
Preclinical testing to prevent
postoperative ileus following abdominal
surgery.
1.3.2 nAChR-Modulators for Cancer Treatment
The connection of nAChRs with cancer development is well established with various nAChR
subtypes contributing to tumour proliferation, survival and metastasis. Antagonism of the nAChRs
has the potential to inhibit these effects and induce cancer cell apoptosis, as discussed in Section
1.2.5 nAChRs and Cancer Cell Death. Many experimental compounds, including those investigated
for the use in various disease states, may also have important implications in cancer treatment. The
following section explores the novel compounds used to selectively target the nAChR pathway to
regulate cancer development.
Overexpression of the nAChR in human NSCLC tissues has led to a number of studies
attempting to modulate the nAChRs to limit tumour progression. MG624, a synthetic α7-nAChR
antagonist has displayed both anti-proliferative and anti-angiogenic properties in SCLC. Human
microvascular endothelial cells derived from the lung (HMEC-L) co-treated with both MG624 and
23
nicotine supressed the nicotine-associated proliferation of the cancer cells, consolidating the
nAChR-action of MG624. In vivo studies administering MG624 in a SCLC tumour xenografted
murine model demonstrated potent anti-angiogenic effects, supressing angiogenesis through
reducing both EGR-1 and FGF2 promoters (Brown et al., 2012; Thornhill et al., 2013). Similar
effects have also been observed when co-administering the α7-nAChR antagonist, α-BTX, with
cisplatin (alkylating-like chemotherapy drug). The cytotoxic effectiveness of α-BTX alone and in
combination with cisplatin was investigated in A549 lung adenocancer and SK-MES-1 lung
squamous cells. The combination of 0.1 μM cisplatin with low levels of α-BTX (1.0 ρM)
significantly increased the expression of the apoptotic factors Bax, and Caspase-3, while decreasing
Bcl-2 expression, compared to individual treatment (Balkan et al., 2012).
Naturally occurring compounds have been implicated as possible anticancer treatments due to
their interaction with the overexpressed α9-nAChRs in breast cancer. The green tea-polyphenol (–)-
epigallocatechin-3-gallate (EGCG) has been explored as a nAChR-blocker of nicotine or estradiol
(E2) induced breast cancer proliferation. MCF-7 human breast cancer cells pre-treated with EGCG
completely abolished the nicotine and E2 induced α9-nAChR activity. Co-treatment with EGCG
also inhibits the binding activity of 3H-nicotine to the α9-nAChRs of breast cancer cells. EGCG
(1 μM) was shown to competitively bind the α9-nAChR and induce receptor down-regulation,
reducing the nAChR proliferative effects of nicotine and E2 (Tu et al., 2011). Continual studies
have compared additional green and black tea extracts to the inhibitory effects of EGCG α9-
nAChRs. Human breast cancer cells with different ER status, MDA-MB-231 (ER–) and MCF-7
(ER+) were pre-treated with the extracts, having prominent effects on the inhibition of nicotine-
induced α9-nAChR protein expression. Extract treatment also inhibited colony formation of MCF-7
cells, with some extracts demonstrating a higher degree of α9-nAChR regulation than that observed
by EGCG treatment (Ho et al., 2013). Similar results have also been observed with studies treating
MDA-MB-231 cells with polyphenolic compounds, luteolin and quercetin, by significantly down-
24
regulating α9-nAChR expression (Shih et al., 2009). The results demonstrate the potential of diet-
based extracts as possible natural modulators of the α9-nAChRs to limit cancer progression.
In addition to the naturally occurring polyphenols, various endogenous compounds have
demonstrated the potential to regulate nAChR activity. The endogenous secreted mammalian Ly-
6/urokinase plasminogen activator receptor-related protein-1 and -2 (SLURP-1 and SLURP-2),
responsible for signal transduction, immune activation and cell adhesion, has shown α7-nAChR
positive allosteric functions with possible anti-carcinogenic properties (Pettersson et al., 2009).
HT29 human colon carcinoma cells incubated with 1 μM of SLURP-1 and SLURP-2 significantly
reduced cell numbers by ~54 and 63%, respectively, compared to controls. Neither necrotic nor
apoptotic cell death was observed following fluorescent microscopy, with the researchers
concluding that the anti-proliferative effects of SLURP-1 and SLURP-2 were manifested through
the interactions with the α7-nAChR (Lyukmanova et al., 2014). The endogenous neurotransmitter,
serotonin (5-hydroxytryptamine, 5-HT), also displays possible anti-carcinogenic activity with links
to nAChR function. Treatment of HT29 and LS1034 carcinoma cells with the selective serotonin
reuptake inhibitor, sertraline, induced caspase-3 activation and Bcl-2 inhibition, causing a dose-
dependent decrease in cancer cell viability. In vivo, sertraline significantly inhibited tumour growth
in CD1 nude mice xenografted with HT29 cells (Gil-Ad et al., 2008). While the effects of serotonin
reuptake inhibitors have been shown to eliminate the effects of nicotine and nicotine-derived
nitrosamines, suggesting a link between increased serotonin levels and nAChR inhibition (Schuller,
2009).
The use of nAChR modulators as possible cancer sensitisers to current chemotherapy drugs
has been investigated. The nAChR antagonist adiphenine (ADI) was investigated in combination
with the chemotherapy drugs, doxorubicin and docetaxel. A vast range of cancer cell lines were
explored, with cell lines grouped according to their sensitivity (based on the IC50 value from the
chemotherapy treatment) to discriminate between drug-sensitive and -resistant cell lines. 250 μM
ADI treatment of resistant breast carcinoma cell lines did not affect cell proliferation; however,
25
when combined with 0.1 μM doxorubicin + 100 μM docetaxel treatment, caused a significant ~40%
reduction in cell viability, compared to doxorubicin + docetaxel treatment alone. The sensitising
effects were observed at 4-fold lower doses of ADI, even when combined with half of the
doxorubicin + docetaxel dose. Similar effects were also observed with the nAChR antagonists
proadifen and NDNI. The researchers claimed that the chemo-sensitising effects of the nAChR
antagonists were operating through the α9 subunit, having the potential to reduce the amount of
toxic chemotherapy agents required to induce tumour cell death in vivo (Hallett et al., 2012).
Neurotoxins, commonly used to distinguish between nAChR subunit combinations through
their selectivity to distinct receptor configurations, demonstrate potential cancer modulating effects.
Studies have suggested that cancer cell proliferation can be arrested through the use of snake
neurotoxins or snail conotoxins due to their selective antagonistic effects at the α7 nAChRs (Wu et
al., 2011). One particular toxin of interest, α-cobratoxin (α-CbT), has produced anticancer effects in
various cancer cell models, both in vitro and in vivo. The treatment of mesothelioma MPP89 cells
showed a notable induction of apoptosis following treatment with α-CbT, significantly inducing
caspase-3 cleavage and down-regulating several anti-apoptotic proteins (survivin, XIAP, IAP1,
IAP2 and Bcl-xl). In vivo, orthotopically transplanted NOD/SCID mice treated with a 1/1000 LD10
dose of α-CbT (0.12 ng/kg) significantly reduced tumour growth without any signs of significant
toxicity to healthy cells (Catassi et al., 2008). Comparable effects have also been observed in A549
NSCLC orthotopically grafted nude mice and MCF-7 breast tumour-bearing nu/nu mice following
treatment with α-CbT (CATASSI et al., 2006; Mei et al., 2015).
1.3.3 Marine Toxins
The potential anticancer effects of various marine-derived agents has been long known, with
several compounds approved for the use in treating various human cancers, including cytarabine,
eribulin and monomethylauristatin E (Newman et al., 2014). However, the discovery of marine
toxins that have regulatory effects on the nAChR-system has been a new avenue for marine-derived
anticancer therapeutic research. One of the first marine toxins investigated for their potential
26
anticancer effects were the polymeric alkylpyridinium salts (poly-APS), isolated from the marine
sponge Reniera sarai, for their AChE inhibitory activity. Fluorescence-activated cell sorting
analysis of NSCLC cells treated with poly-APS showed a selective cytotoxicity of the marine salts
towards cancer cells, compared to normal lung fibroblasts. Non-toxic doses of poly-APS also
reduced NSCLC cell-to-cell adhesion in vitro, while having no significant organ toxicity in healthy
C57BL/6N mice. The study indicated the possibility of regulating cancer cell proliferation through
the cholinesterase inhibitory effects of marine toxins (Paleari et al., 2006).
Since the discovery of poly-APS, several toxins have been extracted from marine shellfish
that act directly on nAChRs, possessing both potent and selective properties. Extracts related to the
cyclic imine group of toxins, such as gymnodimine (GYM) and spirolides, potently antagonise both
homomeric and heteromeric nAChRs (Bourne et al., 2010). Pinnatoxin (PnTX), a newly discovered
group of cyclic imines, has demonstrated high antagonistic affinity towards α7-nAChR. Recordings
of gamma oscillations in response to tetanic stimulation of hippocampal CA1 slices of male
Sprague-Dawley rats showed a reduction in gamma pulse shape discrimination and spike count in
the presence of PnTX E and F, similar to that seen with the selective α7 nAChR antagonist
methyllycaconitine (Hellyer et al., 2011). Furthermore, studies suggest that cyclic amines
specifically target the α7-subtype, with relatively less potent effects at the heteromeric subtypes,
such as α4β2-nAChRs (Hellyer et al., 2011; Kharrat et al., 2008). The potential nAChR-
antagonistic effects of pinnatoxins suggest their possible use in elucidating the contribution of
nAChRs to cancer development and as therapeutics for regulating cancer cell proliferation.
1.4 Aims and Hypothesis
Current literature has identified the various nAChR pathways that contribute to the
development of cancer. The majority of the published studies focus on the effects of nAChR
agonists, such as nicotine, on enhancing the carcinogenic properties of various lung, breast and
27
colon cancer cell lines. However, investigation into the role of nAChRs in the development of
glioblastoma is limited.
Recent studies investigating the use of nAChR antagonists for regulating cancer development
have highlighted the possible advantages of targeting these receptors directly. The use of these
antagonists has often been investigated in the presence of nicotine, where the main effects of the
investigational compounds are to negate the effects of nicotine on progressing cancer development.
However, many carcinoma cell types, such as mesothelioma and colon cancer types, display an
increase in nAChR expression independent of exogenous influence from compounds such as
nicotine (Egleton et al., 2008). These carcinoma cell types also contain the complete functional loop
of ACh signalling that allows cancer cells to promote proliferation through the autocrine activation
of nAChRs (Lau et al., 2013; Song et al., 2008). The present study looked to target these
endogenous functions of nAChRs using a range of pilot studies to better understand the mechanism
behind nAChR antagonist-induced cancer cell death, where clinical application of the compounds is
not an immediate focus of the various investigations performed.
In this research, the effects of both established and novel nAChR antagonists in regulating
carcinogenesis have been explored directly, and in combination with common chemotherapy
agents, utilising a variety of different methods. GYM and PnTX are considered novel compounds
due to the lack of published literature relating to their effects on carcinogenesis. Details regarding
the various compounds, selectivity towards nAChRs and the concentration ranges investigated have
been provided in Table 3 (Section 2.2.2. Cell Viability Assay). Specifically the aims of this research
were:
Aim I: To examine the cytotoxic effects of nAChR modulators on a variety of carcinoma cell
types, and to determine the key nAChR subtypes involved in mediating these
responses.
28
Aim II: To investigate the mechanism(s) of the newly identified role of nAChR modulators in
inducing carcinoma cell death.
Aim III: To determine whether inhibiting common apoptotic pathways would reduce the
cytotoxic effects of nAChR modulators.
Aim IV: To explore the synergistic capacity of the most effective nAChR modulators
identified in Aims I-III to sensitise cancer cells to common chemotherapy agents.
The results of this research suggest a possible mechanism for nAChR antagonist induced cancer cell
cytotoxicity, as highlighted in the following sections. The implications of which are discussed in
context with the current literature.
29
2 Methods
2.1 Cell Culture
2.1.1 Materials
Dulbecco’s Modified Eagle Medium (DMEM), foetal bovine serum (FBS), 0.5% Trypsin
EDTA, and Penicillin/Streptomycin were purchased from Gibco, Life Technologies (California,
USA). Phosphate buffered saline (PBS) was purchased from Invitrogen, Life Technologies
(California, USA). Haemocytometer, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide
(MTT) and Roswell Park Memorial Institute (RPMI) 1640 Medium was purchased from Sigma-
Aldrich (St Louis, MO, USA). Dimethyl sulfoxide (DMSO) was purchased from Scharlau
(Sentmenat, Barcelona, Spain). GraphPad Prism 5 was purchased from GraphPad Software (La
Jolla, CA, USA).
2.1.2 Cell Lines and Cell Culture
A total of eight human carcinoma cell lines were used in this research, as described in Table
2. MDA-MB-231, SKBR3, HT29 and PC3 cell lines were a gift from Associate Professor R.
Rosengren of the Pharmacology Department, University of Otago (Dunedin, NZ). U87, U373 and
LN18 cell lines were a gift from Dr. K. Greish of the Pharmacology Department, University of
Otago (Dunedin, NZ). A549 cells were a gift from Dr. G. Giles of the Pharmacology Department,
University of Otago (Dunedin, NZ).
30
Table 2: Details of the Carcinoma Cell Lines
Cell Line Disease State Cell Type Tissue
MDA-MB-231 Breast adenocarcinoma Epithelial
Mammary gland/breast,
derived from metastatic
site (pleural effusion)
HT29 Colorectal adenocarcinoma Epithelial Colon
PC3 Grade IV prostate
adenocarcinoma Epithelial
Prostate, derived from
metastatic site (bone)
SKBR3 Breast adenocarcinoma Epithelial
Mammary gland/breast,
derived from metastatic
site (pleural effusion)
A549 Lung carcinoma Epithelial Lung
U87 Grade IV astrocytoma Epithelial Brain
LN18 Grade IV astrocytoma Epithelial Brain/cerebrum, right
temporal lobe
U373 Grade IV astrocytoma Epithelial Brain
MDA-MB-231, HT29, PC3, A549 and SKBR3 cells were cultured in DMEM with 10% heat
inactivated FBS, 100 units/mL penicillin and 100 µg/mL streptomycin. U87, U373 and LN18 cells
were cultured in RPMI medium with 10% heat inactivated FBS. Cells were incubated at 37°C in
5% CO2 humidified conditions. Cell lines grew to ~80% confluence in 75 cm2 culture flasks before
passaging. The sub-culturing process involved discarding the media, washing with 2 mL trypsin
solution (1x trypsin, 0.5% EDTA, PBS), and incubated in 2 mL trypsin solution until cells no
longer adhered. 5 mL fresh media was added to the trypsin-cell solution, transferred to a 15 mL
falcon tube and centrifuged at 1200 rpm for 3 minutes at 4°C. The supernatant was discarded, with
the cell pellet re-suspended in 5 mL fresh medium. Cells were counted visually using a
haemocytometer under 20x magnification. The number of cells were counted in all four quadrants,
averaged and multiplied by 104, to give the cell number per mL of media. Cells were then seeded
according to experiment requirements.
2.1.3 MTT Assay
The MTT colourmetric assay was employed to determine cell viability following treatment
(refer below). The assay utilises the reduction of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl
tetrazolium bromide) (MTT) by NAD-dependant enzymatic activity, to form formazin crystals, as
31
an indicator of metabolically active viable cells (Mosmann, 1983). The MTT solution was prepared
at 5 mg/mL in PBS and combined with cells at a final concentration of 0.5 mg/mL. The MTT-media
solution was incubated for ~3 hours in the humidified incubator, solution removed, and replaced
with an equal volume of DMSO (to dissolve the formazin crystals, forming a purple solution). The
reduced MTT solution was analysed spectrophotometrically at 560 nm and processed using
microplate manager 5.2.1 software to quantify the amount of viable cells.
2.1.4 Statistical Analysis
For all cellular viability MTT assay results, GraphPad Prism software was utilised to average
and normalise the absorbance values as a percentage of the control mean ± the standard error of the
mean (SEM) following technical replicates of the method. Significance was determined by
analysing the difference of the means, using either a one-way or a two-way analysis of variance
(ANOVA) with a Bonferroni post-hoc test, where the p-value between the compared groups was
less than 0.05 (p<0.05).
2.2 Cell Viability Assay
2.2.1 Materials
Acetylcholine chloride, (–)-nicotine hydrogen tartrate salt, methyllycaconitine citrate,
serotonin creatine sulphate complex, adiphenine hydrochloride and isopropanol were purchased
from Sigma-Aldrich (St Louis, MO, USA). Mecamylamine hydrochloride was purchased from
Abcam (Cambridge, UK). Pinnatoxin G and Gymnodimine were gifted from Assoc. Prof. S. Kerr of
the Pharmacology Department, University of Otago (Dunedin, NZ), and Dr Paul McNabb and Dr
Andrew Selwood of the Cawthron Insititute (Nelson, NZ).
2.2.2 Cell Viability Assay
MDA-MB-231, HT29, PC3, U87, U373 and LN18 cell lines were seeded in 96-well plates at
a density of 3,000 cells/100µL/well in their respective media, 24 hours prior to treatment. Cells
were treated with acetylcholine (ACh), nicotine (NIC), mecamylamine (MECA), pinnatoxin G
32
(PnTX), gymnodimine (GYM), methyllycaconitine (MLA), adiphenine (ADI) or serotonin (5-HT)
at final well concentration ranging from 0.1 µM to 1000 µM. The vehicle control contained the
highest concentration of the solvent for the corresponding treatment. Treatment profiles are
displayed in Table 3. Treated cells were analysed by MTT assay at 24, 48 and 72 hours post
treatment. The absorbance values for the treated samples were normalised against the untreated
group for each corresponding time point prior to statistical analysis.
Table 3: Cell Viability Assay Treatment Profile
Treatment nAChR Selectivity Solvent
Maximum
Volume of
Vehicle
Control (µL)
Minimum
Final Well
Concentration
(µM)
Maximum
Final Well
Concentration
(µM)
Acetylcholine Broad-Spectrum
Agonist H2O 10 0.1 1000
Nicotine Broad-Spectrum
Agonist H2O 10 0.1 1000
Mecamylamine Broad-spectrum
Antagonist H2O 10 0.1 1000
Pinnatoxin G α7, α4β2 Antagonist Isopropranolol 0.193 0.1 5
Gymnodimine α7, α4β2, α3β2
Antagonist H2O 10 0.1 50
Methyllycaconitine α7 Antagonist H2O 10 0.1 100
Adiphenine
Antagonist at α3 or
α4 Containing
Heteromeric
nAChRs
H2O 10 0.1 1000
Serotonin
Secondary Effects
on Decreasing
nAChR Expression
H2O 10 0.1 1000
2.3 Western Blot
2.3.1 Materials
Albumin from bovine serum (BSA), 3-(N-morpholino) propanesulfonic acid (MOPS),
Tween® 20, sodium dodecyl sulphate (SDS), Trizma®-base, Trizma®-hydrochloride, Triton® X-100,
33
2-mercaptoethanol (MeSH) and tetramethylethylenediamine (TEMED) were purchased from
Sigma-Aldrich (St Louis, MO, USA). 30% acrylamide/bis solution 19:1 and ammonium persulfate
(APS), were purchased from Bio-Rad Laboratories Ltd (Hercules, CA, USA).
Ethylenediaminetetraacetic acid as purchased from BDH/VWR International Ltd (Radnor, PA,
USA). PageRuler™ Prestained Protein Ladder, CL-XPosure Film, Pierce® Protein Inhibitor Tablets,
Pierce® BCA Protein Assay Kit, and dithiothreitol (DTT) were purchased from Thermo-Fisher
Scientific (Waltham, MA. USA). NuPage LPS Sample Buffer and iBlot® Gel Nitrocellulose transfer
Stacks were purchased from Novex, Life Technologies (California, USA). Amersham ECL Prime
Western Blotting Detection Reagents were purchased from GE Healthcare (Little Chalfont, UK).
2.3.2 Antibodies
Rabbit monoclonal anti-nicotinic acetylcholine receptor alpha 4 antibody, rabbit polyclonal
anti-nicotinic acetylcholine receptor alpha 7, rabbit polyclonal anti-CHRNA9 antibody (alpha 9),
goat polyclonal secondary antibody to mouse IgG, and goat polyclonal secondary antibody to rabbit
IgG were purchased from Abcam, Inc (Cambridge, UK). Mouse monoclonal anti-β-actin antibody
was purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). The primary anti-α4-
nAChR, anti-α7-nAChR, and anti-β-actin antibodies were used at 1:25,000 dilution; while primary
anti-α9-nAChR was used at 1:12,500. The secondary antibodies were incubated at 1:10,000
dilution. Antibodies were diluted in 10 mL PBS containing 1% w/v BSA and 0.1% v/v Tween.
2.3.3 Protein Extraction
MDA-MB-231, HT29, PC3, U87, U373 and LN18 cell lines were seeded in 6-well plates at a
density of 200,000 cells/mL/well in their respective media. For baseline levels of nAChR
expression, cells were grown until ~80% confluence before harvesting. Treated samples were
allowed to adhere for 24 hours prior to treatment, and then harvested 72 hours post treatment, with
appropriate untreated controls. MDA-MB-231 cells used in the time-course study were treated after
24 hour adherence with 10 µM nicotine, and harvested at 30 min, 60 min, 2 h, 6 h, and 24 h post-
treatment, with corresponding untreated controls. Cell samples were harvested by removing media,
34
washing with PBS and incubating in 200 µL lysis buffer (NaCl 150 mM, Tris Base 50 mM, SDS
0.05%, Triton X100 1%, AEBSF, aprotinin, bestatin, E64, leupeptin, pepstatin A, pH 8.0) on ice
for 30 min. Cell lysate samples were collected, sonicated, and homogenised by centrifugation
(18,000 rpm, 8 min, 4°C). The protein concentration of the lysate supernatant was measured using
the Pierce® BCA Protein Assay Kit, combined with loading buffer, reducing buffer (DTT 500 mM)
and heat denatured at 70°C for 10 min.
2.3.4 Western Blotting
Loading of 5 µg protein per 20 µL per well and protein reference ladder (7 µL per well) was
resolved by 8% sodium dodecyl sulphate-polyacrylamide gel electrophoresis in MOPS buffer
(MOPS 50 mM, Tris Base, 50 mM, SDS 1%, EDTA 1 mM, pH 7.7) at 150 V for 65 min. Resolved
proteins were electroblotted onto a nitrocellulose membrane (20 V for 1 min, 23 V for 4 min, 25V
for 2 min) and blocked (5% nonfat milk solution in PBS-Tween 0.1%) for 100 min at room
temperature. The membranes were then incubated in the anti-nAChR primary antibody overnight at
4°C with gentle agitation, washed in PBS-Tween 0.1% (6 x 5 min), and incubated in the appropriate
secondary antibody at room temperature for 60 min. Membranes were washed as described above,
visualised with enhanced chemiluminescence reagent (used at a 1:1 concentration, as per the
manufacturer’s instructions) and immediately exposed to X-ray film for developing. The
membranes were then stripped (Tris HCl 62.5 mM, SDS 2%, MeSH 0.1 M, pH 6.8) for 30 min at
50°C, washed, blocked and re-probed with anti-β-actin primary antibody (for normalisation of
loading). Developed films were scanned using an imaging densitometer and densitometry data for
the individual Western blot bands determined by Quantity One software. GraphPad Prism software
was used to determine the mean density of each band.
35
2.4 Immunocytochemistry
2.4.1 Materials
Paraformaldehyde and Hoechst BisBenzimide H33342 trihydrochloride were purchased from
Sigma-Aldrich (St Louis, MO, USA). Alexa Fluor® 488 Goat Anti-Rabbit IgG (H+L) Antibody and
ProLong® Gold antifade reagent were purchased from Molecular Probes, Life Technologies
(California, USA).
2.4.2 Sample
MDA-MB-231 cells were seeded at 10,000 cells/500µL/well onto sterile micro-circular
coverslips in 24-well plates for 72 hours. Cells were fixed with 4% paraformaldehyde- PBS pH 7.4
for 15 min at room temperature. The paraformaldehyde fixed samples were treated with either
0.25% Triton X-100-PBS (for permeabilisation of cell membranes) or PBS (non-permeabilised) for
10 min. Following this step, samples were blocked with 1% BSA in PBS-Tween for 30 min, then
incubated with primary antibody (nAChR-α7 or nAChR-α9 at 1:100 in PBS-Tween) overnight at
4oC. The probed samples were incubated with fluorescent secondary antibody (488 Goat Anti-
Rabbit at 1:500 in PBS-Tween) for 1 hour under constant oscillation at room temperature in the
dark. The coverslips were briefly incubated with 1 µg/mL Hoescht stain for 1 min under dark
conditions, dried, and transferred to microscope slides, bound by ProLong Gold gel for
preservation. The prepared microscope slides were dried and stored in the dark until visualisation.
2.4.3 Fluorescence Microscopy
Cells were visualised using an upright Nikon Eclipse Ni-E Motorised microscope with
ultraviolet or blue illumination from a mercury fibre illuminator lamp through a 100x Nikon
florescence oil immersion objective (numerical aperture 1.3). Images were captured using NIS-
Elements Microscope Imaging Software through a 4,6-diamidino-2-phenylindole (DAPI) or
Fluorescein isothiocynate (FITC) filter. For image adjustments, the captured images were merged
and examined using ImageJ software.
36
2.5 Flow Cytometry
2.5.1 Materials
Propidium iodide (PI), citric acid and 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid
(HEPES) were purchased from Sigma-Aldrich (St Louis, MO, USA). Tetramethylrhodamine, ethyl
ester, perchlorate (TMRE) and Annexin V (Alexa Fluor® 488 conjugate) were purchased from
Molecular Probes, Life Technologies (California, USA). Di-sodium hydrogen orthophosphate
(Na2HPO4) and calcium chloride (CaCl2) was purchased from BDH/VWR International Ltd
(Radnor, PA, USA). Ribonuclease A was purchased from Invitrogen, Life Technologies (California,
USA). Ethanol and sodium chloride (NaCl) were purchased from Scharlau (Sentmenat, Barcelona,
Spain). Cisplatin (CDDP) was purchased from Ebewe Pharma (Unterach, Austria).
2.5.2 Cell Treatment and Staining
MDA-MB-231, HT29, PC3, and LN18 cell lines were seeded in 6-well plates at a density of
100,000 cells/mL/well in their respective media. Samples were allowed to adhere for 24 hours prior
to treatment with either MECA 1000 µM, PnTX 5 µM, ADI 1000 µM, 5-HT 1000 µM or CDDP
20 µM, with appropriate untreated controls. Cell samples were harvested using trypsin at 72 hours
post-treatment, and probed using various fluorescent stains, including PI, TMRE, or a combination
of Annexin V and PI.
Propidium Iodide
Cells were washed in cold PBS and spun at 2000 rpm for 3 min. The samples were fixed by
discarding the supernatant and mixing the cell plate with 500 µL per tube of 4°C ethanol 70%, then
stored at 4°C for 30 minutes. The fixed cell solution was washed twice in phosphate-citrate buffer
(Na2HPO4 0.192 M, Citric Acid 0.004 M), spinning at 2000 rpm for 3 min between each wash. The
cell supernatant was removed then treated with 50 µL per tube of 100 µg/mL Ribonuclease A,
followed by 200 µL per tube of 50 µg/mL propidium iodide, and kept on ice until analysis.
TMRE
37
The treated cell samples were re-suspended in 500 µL per tube of appropriate media
following trypsin harvesting. 2 µL per tube of 10 µM TMRE was added to the samples under dark
conditions, and incubated at 37°C protected from light for 10 minutes. Cell samples were washed in
PBS, spun at 2000 rpm for 3 min, and re-suspended in 300 µL per tube of media. Stained samples
were kept on ice, protected from light until analysis.
Annexin V/PI
The cell samples were washed twice in cold PBS, with spinning at 2000 rpm for 3 min, and
re-suspended in 100 µL per tube Annexin V binding buffer (HEPES 0.1 M, NaCl 1.4 M, CaCl2 25
mM, pH 7.4). 2 µL per tube Annexin V conjugate and 2 µL per tube of 50 µg/mL PI were added to
the cell solution and incubated at room temperature for 15 min protected from light. 400 µL per
tube of additional Annexin V binding buffer was combined to each sample and stored on ice until
analysis.
2.5.3 Fluorescence-Activated Cell Sorting (FACS)
Stained samples were loaded into FACS tubes and run through the FACSCalibur™. Data was
acquired using CellQuestPro software with an acquisition criteria of 10,000 events per sample, and
analysed using Flowing Software 2.5.0. The PI cell samples were measured using the yellow 585/42
nm (FL2-H) detector and analysed to generate cell cycle phase distribution data. The data was gated
to ensure only single cells were analysed, with the cell cycle phases (sub-G1, Go/G1, S and G2/M)
determined manually based on the methods described in the literature (Wlodkowic et al., 2011).
TMRE samples were measured via the same yellow 585/42 nm detector and histograms produced to
quantify the change in TMRE peaks between control and treated samples. Double-stained, Annexin
V and PI samples were measured using the green 530/30 nm (FL1-H) and yellow 585/42 nm (FL2-
H) detectors, graphed accordingly on a dot-plot with quadrants for double stain and individual
stains measured. Values were entered into GraphPad Prism software and analysed using a one-way
ANOVA with a Bonferroni post-hoc test, where the significance was determined by p<0.05.
38
2.6 Intracellular Calcium
2.6.1 Materials
Fura-2 acetoxymethyl ester (AM) was purchased from Abcam, Inc (Cambridge, UK).
Potassium chloride (KCl) was purchased from Scharlau (Sentmenat, Barcelona, Spain). Glucose,
and sodium bicarbonate (NaHCO3) were purchased from Sigma-Aldrich (St Louis, MO, USA).
Potassium di-hydrogen orthophosphate (KH2PO4) was purchased from BDH/VWR International
Ltd (Radnor, PA, USA). Magnesium sulphate (MgSO4) was purchased from Merck Millipore
(Darmstadt, Germany).
2.6.2 Fura-2 AM Fluorescence Measurement
MDA-MB-231, HT29 and PC3 cells were seeded at 40,000 cells/100µL/well in 96-well black
plates for fluorescence. Cells were allowed to adhere overnight before being loaded with 100 µL
per well of 4 µM of the Ca2+ indicator Fura-2 AM (from 1 mM stock in DMSO) in Hanks Balanced
Salt Solution (HBSS; NaCl 0.137 M, KCl 5.4 mM, Na2HPO4 0.25 mM, glucose 0.01%, KH2PO4
0.44 mM, CaCl2 1.3 mM, MgSO4 1.0 mM, NaHCO3 4.2 mM) for 60 min in a humidified incubator
protected from light. The cells were then washed in HBSS and incubated in fresh HBSS 100 µL per
well for 30 min at room temperature protected from light to allow Fura-2 AM cleavage. Cells
transfected with Fura-2 were treated with Ach 10 µM, MECA 1000 µM, PnTX 5 µM, GYM 10
µM, ADI 1000 µM, MLA 100 µM or 5-HT 1000 µM at time = 0 min. Fura-2 fluorescence was then
measured every minute for the first 20 min, 1 h, 2 h, 4 h, and 6 h post-treatment. Softpro Max
Software 4.8 was used to determine Fura-2 fluorescence using a fluorescent plate reader at an
excitation of 340 nm (λ1) and 380 nm (λ2), with an emission of 510 nm, at each time point. The
fluorescence intensity ratio (R = Fλ1/Fλ2) of Fura-2 fluorescence detected at 510 nm for each time
point was then calculated, indicating the ratio of Fura-2 in a completely Ca2+-saturated state against
Fura-2 in a Ca2+-free state. GraphPad Prism software was used to construct time-course graphs of
the fluorescence intensity ratio to display the difference between treatment groups over time.
39
2.7 Cell Death Inhibitors
2.7.1 Materials
1,2-bis(o-aminophenoxy)ethane-N,N,N’,N’-tetraacetic acid – acetoxymethyl ester (BAPTA-
AM), N-Acetyl-Leu-Leu-Nle-al (ALLN) and N-benzyloxycarbonyl-Val-Ala-Asp-fluoromethyl
ketone (zVAD-FMK) were purchased from Abcam, Inc (Cambridge, UK). Cyclosporin A (CsA)
was purchased from Sigma-Aldrich (St Louis, MO, USA).
2.7.2 Cell Culture and Inhibitors
MDA-MB-231, HT29, PC3, and LN18 cell lines were seeded in 96-well plates at a density of
3,000 cells/100µL/well in their respective media. Cells were allowed to adhere for 24 hours, then
treated with specific concentrations of various cell death inhibitors, as shown in Table 4, alone and
in combination with varying concentrations of PnTX, MECA, MLA, ADI or 5-HT. Treated cell
viability was measured via MTT assay at 48 and 72 hours post-treatment.
Table 4: Cell Death Inhibitor Treatment Profile
Cell Line
Final Well Concentration (µM)
CsA ALLN BAPTA-AM zVAD
MDA-MB-231 10 5 1 30
HT29 1 1 1 20
PC3 1 5 10 30
LN18 1 1 0.5 10
2.7.3 Isobologram
Following treatment of the various cancer cell types with cell death inhibitors, as described in
Section 2.7.2, an isobologram was constructed to investigate the relationship between CsA and
MECA in reducing cell viability. MDA-MB-231 cells were seeded in 96-well plates at a density of
3,000 cells/100µL/well in DMEM. The cells were allowed to adhere for 24 hours before treatment.
Cells were treated with CsA at a range of 0.1 to 40 µM, MECA at a range of 10 to 1500 µM, and
combinations of the two (CsA 0.1, 1.0 or 5.0 µM; MECA 100 or 500 µM). The treated MDA-MB-
40
231 cells were then analysed by MTT assay 72 hours post treatment. IC50 values for CsA and
MECA were calculated for the individual compounds and in combination to determine the overall
cytotoxic relationship between the two compounds. An isobologram was constructed according to
the methods described by (Tallarida, 2006) from the IC50 values calculated using OriginPro
software (OriginLab Corporation, MA, USA).
2.8 Combination Therapy
2.8.1 Materials
Doxorubicin (DOXO) and Docetaxel (DTX) were purchased from Cayman Chemical Company
(Michigan, USA).
2.8.2 Chemotherapy Combination Therapy
The methods for chemotherapy combination treatment of various cancer cell lines were
adopted from (Hallett et al., 2012). MDA-MB-231, HT29, PC3, A549 and SKBR3 cells were
seeded in 96-well plates at a density of 20,000 cells/100µL/well in their respective media. Cells
were allowed to adhere for 4 hours, followed by compound addition of various concentrations of
chemotherapy agents and nAChR antagonists, with appropriate vehicle controls. Chemotherapy
final well concentrations included doxorubicin at 0.1-1.0 µM and docetaxel at 100 µM. After 24
and 48 hours post-treatment, cell viability was measured via the MTT assay.
41
3 Results
3.1 Effects of nAChR Antagonists on Cell Viability
The link between nAChR activation and cancer development is well established. Several
nAChR subtypes have been shown to contribute to tumour proliferation, survival and metastasis
(refer to Section 1.2 nAChRs in Cancer Development). To test the hypothesis that the more selective
modulators of the nAChRs would display the most potent cytotoxic effects, a colourimetric cell
viability assay was used to measure the effects of exposure of a range of carcinoma cell types (refer
Table 2) to both established and novel nAChR modulators (Aim I).
3.1.1 Cell Viability Assay
Initial experiments were undertaken using the cell viability assay to establish an effective
modulating concentration for each test compound. Contrary to expectations, the nAChR agonists
acetylcholine (ACh) and nicotine (NIC) did not exhibit a consistent change in cell viability across
the range of concentrations studied (0.1 µM to 1000 µM), with the exception of HT29 cells (data
not shown). ACh and NIC concentrations of 10 µM were therefore chosen based on the observed
effects on HT29 cells, an established history of increased cancer cell viability and a potential
modification of nAChR expression observed at this concentration in published studies (Al-Wadei et
al., 2012; Dasgupta et al., 2009). Following treatment with ACh 10 µM for 72 hours, a significant
increase (P < 0.001) in HT29 viability was detected, corresponding to a 28.50% increase in cell
viability compared to untreated control cells (Figure 3A). A similar increase in cell viability was
seen following treatment of HT29 cells with NIC 10 µM (19.91% increase in cell viability) (Figure
3B). No significant increase in cell viability was seen in any other cell line when treated with either
ACh or NIC at 10 µM. Furthermore, visual analysis of MDA-MB-231, HT29 and PC3 cell lines,
provided for reference only, at 72 hours post treatment showed no changes in cell morphology or
confluency, compared to control cells (refer to microscopic images in Figure 3A and Figure 3B).
42
A
0 24 48 720
20
40
60
80
100
120
140
MDA-MB-231
HT29
PC3
U87
LN18
U373
Time (Hours)
24 48 72
MDA-MB-231 ns ns ns
HT29 ns ns ***
PC3 ns ns ns
U87 ns ns ns
LN18 ns *** *
U373 ns ns ns
ns = not significant, * = p<0.05, ** = p<0.01, *** = p<0.001
Time (Hours)
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
B
0 24 48 720
20
40
60
80
100
120
MDA-MB-231
HT29
PC3
U87
LN18
U373
Time (Hours)
24 48 72
MDA-MB-231 ns ns ns
HT29 ns ns **
PC3 ns ns ns
U87 ns * ***
LN18 * ns *
U373 ns ** ***
ns = not significant, * = p<0.05, ** = p<0.01, *** = p<0.001
Time (Hours)
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
Figure 3: The determination of MDA-MB-231, HT29, PC3, U87, LN18 and U373 cell viability following treatment with
ACh 10 µM (A) or NIC 10 µM (B). Cells were treated following a 24 hour seeding period, with cell viability tested at 24
hour intervals over a 72 hour study timeframe, with viability established using the MTT assay. Bright-field microscopic
images of MDA-MB-231, HT29 and PC3 treated cells (primary image) versus control cells (inset image) at 72 hours
are shown (20x magnification). Values are displayed as the mean percent of the viability of control (untreated) cells,
with error bars representing SEM (n = 2). A two-way ANOVA with a Bonferroni post-hoc test was used to determine
significance (* P < 0.05, ** P < 0.01, *** P < 0.001).
MDA-MB-231
HT29
PC3
MDA-MB-231
HT29
PC3
ACh 10 μM
NIC 10 μM
43
An array of nAChR antagonists were investigated for their effects on cancer cell viability. The
selectivity of the antagonists towards specific nAChR subtypes ranged from relatively non-selective
to highly selective, as summarised in Table 3. Again, concentration dependence studies were
conducted on each compound to determine which concentration had the greatest effect on cell
viability (data not shown). Contrasting effects on cell viability, compared to ACh and NIC, were
seen for the majority of compounds tested. Mecamylamine (MECA) 1000 µM, pinnatoxin G
(PnTX) 5 µM, serotonin (5-HT) 1000 µM and adiphenine (ADI) 1000 µM caused a significant
decrease in cell viability after 72 hours of treatment for all cell lines tested (P < 0.01 for all
treatments and cell lines). MECA 1000 µM (Figure 4A) and 5-HT 1000 µM (Figure 4D) produced
the largest range of viability results; causing a 21.53% (U87) to 83.28% (HT29) and 16.66% (U373)
to 72.74% (PC3) decrease in cell viability, respectively. ADI 1000 µM (Figure 4E) treatment
caused at least 50% decrease in cell viability by the 48 and 72 hour time points in all cell lines
tested, with MDA-MB-231s demonstrating the greatest susceptibility to ADI treatment (97.62%
decrease in cell viability at 72 hours compared to control). PnTX 5 µM (Figure 4B) was the most
potent of the antagonists studied, causing between 28.11% (U87) to 68.32% (HT29) decrease in cell
viability at a concentration 200-fold lower than all other nAChR antagonists tested. Gymnodimine
(GYM) 10 µM (Figure 4C) and methyllycaconitine (MLA) 100 µM (Figure 4F) produced the least
significant change in cell viability of the compounds tested.
Bright-field microscopic images of MDA-MB-231, HT29 and PC3 cell lines, as a
representation of all the cell lines tested, at 72 hours post treatment showed obvious visual changes
in cell morphology and confluency, consistent with the viability results. MECA, 5-HT and ADI
treatment showed a widespread decrease in cell viability, compared to control (untreated) cells. A
similar change in cell viability was seen following PnTX treatment, with a significant depletion in
viable cells compared to control cells. Microscopic images following GYM and MLA treatment
were consistent with the viability assay results, with little observable changes to cell morphology or
confluency.
44
A
0 24 48 720
10
20
30
40
50
60
70
80
90
100
110
MDA-MB-231
HT29
PC3
U87
LN18
U373
Time (Hours)
24 48 72
MDA-MB-231 *** *** ***
HT29 *** *** ***
PC3 *** *** ***
U87 ns * **
LN18 *** *** ***
U373 * *** ***
ns = not significant, * = p<0.05, ** = p<0.01, *** = p<0.001
Time (Hours)
No.
of
Via
ble
Cel
ls C
om
pare
d t
o C
on
trol
B
0 24 48 720
10
20
30
40
50
60
70
80
90
100
110
MDA-MB-231
HT29
PC3
U87
LN18
U373
Time (Hours)
24 48 72
MDA-MB-231 ns ** ***
HT29 *** *** ***
PC3 ns *** ***
U87 *** *** ***
LN18 *** *** ***
U373 ** *** ***
ns = not significant, * = p<0.05, ** = p<0.01, *** = p<0.001
Time (Hours)
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
MDA-MB-231
HT29
PC3
MDA-MB-231
HT29
PC3
MECA 1000 μM
PnTX 5 μM
45
C
0 24 48 720
10
20
30
40
50
60
70
80
90
100
110
120
MDA-MB-231
HT29
PC3
U87
LN18
U373
Time (Hours)
24 48 72
MDA-MB-231 ns * **
HT29 ns ns ns
PC3 ns ns ns
U87 ns ns ns
LN18 ns ns ns
U373 ns ns ns
ns = not significant, * = p<0.05, ** = p<0.01, *** = p<0.001
Time (Hours)
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
D
0 24 48 720
10
20
30
40
50
60
70
80
90
100
110
120
MDA-MB-231
HT29
PC3
U87
LN18
U373
Time (Hours)
24 48 72
MDA-MB-231 *** *** ***
HT29 ns *** ***
PC3 ns *** ***
U87 *** *** ***
LN18 *** ns ***
U373 ** *** ***
ns = not significant, * = p<0.05, ** = p<0.01, *** = p<0.001
Time (Hours)
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
MDA-MB-231
HT29
PC3
MDA-MB-231
HT29
PC3
GYM 10 μM
5-HT 1000 μM
46
E
0 24 48 720
10
20
30
40
50
60
70
80
90
100
110
MDA-MB-231
HT29
PC3
U87
LN18
U373
Time (Hours)
24 48 72
MDA-MB-231 *** *** ***
HT29 *** *** ***
PC3 *** *** ***
U87 ns *** ***
LN18 *** *** ***
U373 *** *** ***
ns = not significant, * = p<0.05, ** = p<0.01, *** = p<0.001
Time (Hours)
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
F
0 24 48 720
20
40
60
80
100
120
140
MDA-MB-231
HT29
PC3
U87
LN18
U373
Time (Hours)
24 48 72
MDA-MB-231 ns ns ns
HT29 ns ns ns
PC3 ns ns **
U87 ns ** ns
LN18 ns ns ns
U373 ns ns *
ns = not significant, * = p<0.05, ** = p<0.01, *** = p<0.001
Time (Hours)
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
Figure 4: The evaluation of MDA-MB-231, HT29, PC3, U87, LN18 and U373 cell viability following treatment with
MECA 1000 µM (A), PnTX 5 µM (B), GYM 10 µM (C), 5-HT 1000 µM (D), ADI 1000 µM (E) or MLA 100 µM (F).
Cells were treated following a 24 hour seeding period, with cell viability tested using the MTT assay at 24 hour
intervals over a 72 hour study timeframe. Bright-field microscopic images of MDA-MB-231, HT29 and PC3 treated
cells (primary image) versus control cells (supplementary image) at 72 hours are shown (20x magnification). Values
are displayed as the mean percent of the control viability, with error bars representing SEM (n = 2). A two-way
ANOVA with a Bonferroni post-hoc test was used to determine significance (* P < 0.05, ** P < 0.01, *** P < 0.001).
MDA-MB-231
HT29
PC3
MDA-MB-231
HT29
PC3
ADI 1000 μM
MLA 100 μM
47
3.1.2 Western Blot - Basal nAChR Levels
To investigate the possible correlation between nAChR subtype expression and susceptibility
to nAChR antagonism-induced changes in cell viability, the basal levels of nAChR subtypes in the
cell lines tested were determined using Western blot analysis. The relative densities of the bands on
the Western blots were quantified using laser densitometry in comparison to β-actin.
As described in the literature, the relative abundance of the different nAChR subtypes within
a cell varies depending on cell type. HT29 cells demonstrated the greatest level of nAChR
expression across all cell lines, as shown in Figure 5. HT29 cell expression of the α4, α7 and α9
nAChR subtypes were shown to be at least 142.6%, 7.43% and 86.80%, higher than the expression
observed in all other cell lines for the respective subtypes. PC3 cells showed a similar level of α7
nAChR expression as the HT29 cells; while LN-18 possessed the greatest total nAChR expression
among the glioblastoma cell lines.
48
MD
A-M
B-2
31
HT29
PC3
U87
LN
18
U37
3
0.00
0.25
0.50
0.75
1.00
1.25
1.50
4 nAChR
7 nAChR
9 nAChR
Den
sity
Rati
o
Figure 5: Western blot analysis showing the basal level of α4, α7 and α9 nAChR expression in MDA-MB-231, HT29,
PC3, U87, LN18 and U373 cancer cell lines. β-actin protein was used as a loading control to ensure equal loading of
proteins. Each well was adjusted according to the corresponding β-actin control. The columns in the graph represent
the density ratio for each nAChR subtype.
3.1.3 Immunocytochemistry
To further demonstrate the presence and localisation of the nAChR subtypes on cancer cells,
immunocytochemistry was performed on untreated MDA-MB-231 cells. The immunocytochemistry
images for non-permeabilised and permeabilised cell samples stained for the nAChR α7 and α9
subtypes are presented in Figure 6 and Figure 7, respectively. Staining with the α7 nAChR subtype
antibody in non-permeabilised cell samples showed widespread expression (Figure 6A). Subtype
expression was clearly displayed on the surface of the cancer cells, with much of the α7 labelling
49
following the numerous cell processes. However, significant unknown staining of the α7 antibody
was seen in the extracellular environment. A high cluster of staining was also observed around the
nucleus of the carcinoma cells, predominately in the permeabilised MDA-MB-231 samples (Figure
6B).
A diverse pattern of receptor subtype expression was observed for MDA-MB-231 cells
treated with the α9 antibody. Non-permeabilised cell samples showed specific staining of receptor
subtypes that mimic the flow of the cellular cytoskeleton (Figure 7A). A similar trend was seen in
the permeabilised cell samples (Figure 7B). Furthermore, the α9 antibody of permeabilised samples
stained the region of mitotic spindles within the nucleus of replicating cells. Slight extracellular and
high-density staining was seen with the α9 antibody; however, not as predominately as compared to
that seen for the α7 antibody.
50
A
B
Figure 6: Immunocytochemistry images of non-permeabilised (A) and permeabilised (B) MDA-MB-231 cells showing
α7 nAChR subtype expression. Cells were treated with the primary α7 nAChR antibody with a fluorescent Alexa Fluor®
488 secondary antibody (green) and the Hoechst nuclear stain (blue). The treated cells were visualised using an upright
Nikon Eclipse Ni-E Motorised microscope with a 100x objective lense. DAPI and FITC images were overlayed using
Image J software, with the corresponding bright-field microscopy images shown.
10 μm
10 μm
51
A
B
Figure 7: Immunocytochemistry images of non-permeabilised (A) and permeabilised (B) MDA-MB-231 cells showing
α9 nAChR subtype expression. Cells were treated with the primary α9 nAChR antibody with a fluorescent Alexa Fluor®
488 secondary antibody (green) and the Hoechst nuclear stain (blue). The treated cells were visualised using an upright
Nikon Eclipse Ni-E Motorised microscope with a 100x objective lense. DAPI and FITC images were overlayed using
Image J software, with the corresponding bright-field microscopy images shown.
10 μm
10 μm
52
3.1.4 Discussion
Currently, there is only limited information in the literature describing the effects of nAChR
antagonists on cancer cell development. The present studies examined the effects of both
established and novel nAChR modulators on cell viability of a variety of carcinoma cell types using
a cell viability assay. In addition, the study aimed to determine the key nAChR subtypes involved in
mediating these responses. It was predicted that the selective nAChR antagonists, such as the
marine toxins, would display the most potent effects on viability of the carcinoma cell types; while
the α7 and α9 receptor subtypes were anticipated to be the most prominently expressed of the
nAChR subtypes.
Both ACh and NIC were used as positive controls to investigate whether the characteristic
increase in cell viability and expression of nAChRs could be replicated due to their strong agonistic
effects at nAChRs. Viability assay results demonstrated an ~20-30% increase in cell viability of
HT29 cells over 72 hours following treatment with 10 μM ACh or NIC. However, the change in
viability following treatment with ACh or NIC was not reflected in the remaining carcinoma cell
types investigated. Previous studies have demonstrated the ability of NIC to increase cancer cell
proliferation, with the most significant effects seen in lung cancer cell lines (not tested in the
present study) (Dasgupta et al., 2009; Tsurutani et al., 2005). The current literature has shown that
changes in lung cancer cell proliferation and cellular activity could be observed within 5 hours of
NIC treatment. However, it is possible that the extensive nAChR expression on lung cancer cells
differs from the expression patterns of the carcinoma cell types tested in this research. This theory is
supported by the Western Blot results, where lower levels of the key nAChR subtypes linked to cell
proliferation, α7 and α9, were shown compared to the highly responsive HT29 cells. The 10 μM
concentration of ACh and NIC was continued due to the possibility of the compounds having an
underlying effect on nAChR expression or changes in intracellular calcium concentration ([Ca2+]),
based on previous literature findings.
53
Similar effects on HT29 cell proliferation have been previously shown with NIC promoting
cell proliferation, with a dose-dependent increase in DNA synthesis and adrenaline production
(Wong et al., 2007). The study suggested a link between the expression of the α7-nAChR subtype
and NIC-induced proliferation, with selective α7-nAChR blockage reversing the stimulatory effects
on cell proliferation. The present study is consistent with these findings, in that a significant
increase in HT29 cell proliferation was observed following nAChR stimulation that correlated with
a higher level of α7 expression in HT29 cells, compared to all other cell lines tested.
Significant decreases in cell viability were observed following MECA, PnTX, 5-HT and
ADI treatment. PnTX was the most potent of the compounds tested, causing as much as a ~70%
decrease in cell viability at a concentration 200-fold lower than that of the other nAChR
antagonists, with the most prominent effects seen in HT29 cells. PnTX A and G have been
previously shown to potently antagonise human α7 and α4β2 nAChRs (Araoz et al., 2011). Human
nAChR-expressing Xenopus oocytes showed significant blocking of α7 nAChR with nM
concentrations of PnTX A that was not reversible after 10-15 minutes washout. While PnTX A also
blocked the human α4β2 nAChR at higher concentrations, indicating that the toxin may be more
selective for the α7 subtype. PnTX G shows similar effects in oocyte-expressing nAChRs, but at
significantly higher concentrations than PnTX A (Araoz et al., 2011). The present results suggest
that PnTX is inducing carcinoma cell cytotoxicity through similar mechanisms, with both α7 and α4
nAChR subtypes being widely expressed amongst the cell lines tested. In addition, the
responsiveness of the cell lines was also correlated to the expression of the receptor subtypes, with
the highly expressive HT29 cells being the most sensitive to PnTX treatment. Conversely, the U87
cells had low nAChR expression and were the least sensitive to PnTX treatment.
The results of the non-selective nAChR antagonist, MECA, showed similar levels of
viability decrease compared to PnTX, but at a 200-fold higher concentration. It is reported in the
literature that MECA inhibits various nAChRs, including α4β2, α3β2, α3β4 and α7, with the least
potent effects observed at α7 nAChRs (Papke et al., 2001). Due to this relatively low selectivity for
54
the α7 nAChR, the present study suggests that higher concentrations of MECA are required to
induce α7-related cytotoxicity, compared to the selective α7-antagonist, PnTX. A potent decrease
in cell viability was also observed following ADI treatment of the various carcinoma cell types,
similar to that seen for MECA. ADI acts via blocking heteromeric nAChR containing α3 or α4
subunits (eg α3β4, α4β2, α4β4) suggesting the cytotoxic effects may be mediated through the α4
expression on carcinoma cell types (Gentry et al., 2001). The comparative effects of MECA and
ADI, compared to PnTX, demonstrate the importance of both the α7 and α4 nAChR subtypes in
inducing cell cytotoxicity. However, the high concentrations of both MECA and ADI required to
induce a decrease in carcinoma cell viability question the mechanism behind the observed results.
Both MECA and ADI exhibit IC50 values <15 μM at nAChRs. The use of concentrations well
above the IC50 for both compounds in the present study may reduce the selectivity of these
compounds at the specific nAChRs, potentially inducing changes in cell viability through alternate
mechanisms. Additional studies using siRNA knockdown of critical nAChRs would be required to
confirm the mechanism behind the changes in carcinoma cell viability as a result of MECA and
ADI treatment.
Significant effects on cell viability were also observed following treatment with 5-HT, with
a significant decrease in cell viability across all cell lines tested at 72 hours post treatment. The
effects were consistent with those previously observed with 5-HT reuptake inhibitors (Gil-Ad et al.,
2008; Schuller, 2009). It is possible that these effects of 5-HT are mediated through the expression
of chimeric nACh–5-HT receptors that are expressed within the nervous system. Over-activation of
5-HT receptors has been shown to induce a significant downregulation of 5-HT receptors on the cell
surface of neurons (Gray et al., 2001). The treatment of 5-HT in the present study may induce
similar effects on chimeric nACh–5-HT receptors, reducing the expression of functional nACh–5-
HT receptors on the cell surface through receptor internalisation. The decrease in the nAChR
component may cause a fatal decrease in [Ca2+] levels, relating to the observed cancer cell
cytotoxicity at high doses of 5-HT. This theory would explain why previous literature has shown
55
the ability of 5-HT reuptake inhibitors to eliminate the effects of nicotine and nicotine-derived
nitrosamines.
Interestingly, both MLA and GYM treatment had little effect on cancer cell viability with
only MDA-MB-231 and U373 cells showing a significant decrease in cell viability at 72 hours.
MLA exhibits a very high selectivity towards homomeric α7 nAChR, with the ability to bind to one
of five identical binding sites present on the α7 nAChR with an IC50 in the picomolar range (Palma
et al., 1996). MLA is thought to share the same binding sites to the α7 AChRs as the endogenous
agonist ACh, with the binding of a single MLA molecule to the receptor sufficient to prevent its
channel opening. However, unlike many antagonistic toxins that irreversibly bind the nAChR,
recovery of nAChR function is observed following MLA treatment (Palma et al., 1996). These
findings suggest that MLA may only have a significant effect on cellular function in the presence of
an agonist, such as ACh, compared to treatment with the agonist alone due to the competitive
binding nature of the compound. This idea was further investigated in Chapter 3 where [Ca2+] levels
of MDA-MB-231 cells were measured following MLA treatment. In addition, the reversible
binding of MLA may explain the lack of cytotoxicity observed following high concentrations of
MLA treatment, with 72 hours providing sufficient time for the cancer cells to recover and
proliferate.
The treatment of cancer cells with GYM was predicted to induce similar effects on cell
viability as PnTX due to their similarity in chemical structure and nAChR selectivity. GYM has
been previously shown to selectively block homomeric α7 nAChRs expressed in Xenopus oocytes
and heteropentameric α3β2 and α4β2 nAChRs during competition-binding assays (Kharrat et al.,
2008). Despite GYM displaying similar IC50 values of 0.33 and 15.5 nM at human α7 and α4β2
nAChRs, respectively, compared to PnTX, the effects on in vitro cell viability have been opposing
(Shin et al., 2005). Treatment of Neuro2a cells incubated with 10 μM GYM for 24 hours showed no
significant change in cell viability by MTT assay, while GYM at concentrations 10-fold higher have
also been relatively non-toxic on primary cultures of mouse cortical neurons (Alonso et al., 2011;
56
Dragunow et al., 2005). The result of the present study show similar effects on cell viability, with
no significant changes in cell viability, with the exception of MDA-MB-231 cells, following
treatment with 10 μM GYM for 72 hours. The difference in effects between GYM and PnTX may
be due to the reversible binding nature of GYM, showing full recovery within 30 minutes of
washout in hemidiaphragm preparation, compared to the slow and incomplete recovery of tissue
following PnTX treatment (Hellyer, 2014; Hellyer et al., 2011).
The effects of the various compounds of cancer cell viability have been demonstrated. As
noted in Section 2.1.3 (MTT Assay), the assay method measures viable cell activity through the
enzymatic reduction of MTT. However, the method is unable to distinguish between a decrease in
cell activity due to cytotoxicity or an inhibition of cell proliferation. Therefore, the data from the
cell viability assay cannot clarify whether nAChR antagonism is having a cytotoxic effect on the
cells, or preventing the cells from replicating. Microscopic imaging of the treated cells indicate that
some compounds, such as PnTX, may be preventing cell proliferation due to a lack of cell death
observed. However, additional tests, such as flow cytometry analysis of the treated cells discussed
in the following sections, was required to further investigate the effects of these compounds on cell
viability.
To further investigate the presence and localisation of the α7 and α9 nAChR subunits on
cancer cells, immunocytochemistry (ICC) was performed on untreated MDA-MB-231 breast
carcinoma cells. The α9 nAChR is an integral subunit in the development of breast cancer due to its
upregulation following activation of the estrogen receptor, having implications in cancer growth
and metastasis (Lee et al., 2011). Furthermore, the importance of the α7 nAChR subunit has been
linked to several aspects of cancer development. ICC images from non-permeabilised cells treated
with the α7 antibody displayed significant staining on the surface of the cells, including the
numerous cell processes extending from the cell body. Significant levels of expression were also
observed with the α9 antibody, with a pattern that more closely resembles the cell cytoskeleton,
rather than small clusters of receptors on the cell surface. According to current knowledge, this is
57
the first attempt to identify the expression of the α9 nAChR using an ICC method in human breast
cancer cells, making a comparison of the expression pattern with current literature difficult. Overall,
the ICC images exhibit the widespread expression of the key nAChR subtypes linked to
carcinogenesis on the cell surface of human cancer cells, with expression levels that are consistent
with those seen during the Western blot analysis for MDA-MB-231 cells. However, a lack of
reproducibility of the experiments due to difficulty with the method and optimisation of the
antibodies means further experiments would be required to confirm these conclusions.
The significant amount of non-specific binding of both antibodies within the extracellular
region and nucleus of the cancer cells questions the specificity of the antibodies to selectively detect
the α7 and α9 nAChR subunits. It is possible that extracellular staining may be abridged by changes
to the blocking and washing steps, reducing the non-specific expression and binding of the antibody
to cell fragments. Alternatively, the observed expression of the antibodies within the nucleus may
be due to known nuclear binding characteristic of the ER in breast carcinomas and link to α9
nAChR expression (Lee et al., 2011).
Having demonstrated an effect with various nAChR modulators on carcinoma cell viability,
the next step was to explore possible mechanisms of action.
3.2 Investigation of nAChR Antagonist Induced Cell Death
In the previous chapter it was demonstrated that selective nAChR antagonists induce cell
death in carcinoma cell lines. To investigate the mechanism(s) of nAChR modulator-induced
cytotoxicity (Aim II), Western blotting and flow cytometry were used to further characterise the
treated carcinoma cells.
3.2.1 Western Blot – α7 nAChR Levels following Treatment
Western blot analysis was used to investigate the effects of the various nAChR modulators on
α7 nAChR subtype expression (Figure 8). The α7 nAChR receptor was the main focus of the
Western blot study due to its widespread expression and involvement in cancer development.
58
Treatment with the nAChR agonists, ACh 10 µM and NIC 10 µM, showed little effect on α7
expression compared to untreated cells. Of the four cell lines tested, only slight increases in α7
expression were seen in MDA-MB-231. MDA-MB-231 treatment with various nAChR antagonists,
including MECA, PnTX, GYM, MLA and ADI, produced a similar increase in α7 expression
ranging from 1.32 to 1.47-fold compared to untreated cells. No consistent increases in α7
expression were seen across all other cell lines or treatments. LN-18 cells were the least responsive
to treatment with nAChR modulators, with α7 expression decreasing following treatment with all
compounds, with the exception of MECA 1000 µM.
59
Untr
eate
d M
AC
h 10
M
NIC
10
M
MEC
A 1
000
M
PnTX
5
M
GY
M 1
0 M
5-H
T 1
000
M
MLA
100
M
AD
I 25
0
0.0
0.5
1.0
1.5
2.0MDA-MB-231
HT29
PC3
LN18
a7 n
AC
hR
Den
sity
Rati
o
(Fold
Ch
an
ge)
Figure 8: Western blot analysis investigating the effects of various nAChR modulating compounds on the expression
level of the α7 nAChR subtype in MDA-MB-231, HT29, PC3 and LN18 cancer cell lines. Cells were either left untreated
or treated with ACh 10 µM, NIC 10 µM, MECA 1000 µM, PnTX 5 µM, GYM 10 µM, 5-HT 1000 µM, MLA 100 µM and
ADI 250 µM for 72 hours prior to harvesting and analysis. β-actin protein was used as a loading control to normalise
loading of proteins. The columns in the graph represent the fold change in density ratio between the treatment group
and the untreated control (n = 1).
60
3.2.2 Flow Cytometry - Cell Cycle Analysis
Cell cycle analysis was conducted using the propidium iodide (PI) DNA stain to determine
the proportion of cells in each stage of the cell cycle. The percentage of cells in the sub-G1 stage of
the cell cycle was measured using Fluorescent Activated Cell Sorting (FACS) following treatment
with the various nAChR modulators, as displayed in Figure 9. An increase in sub-G1 percentage
represents an increase in fractional DNA content, characteristic of apoptotic cells. CDDP was
chosen as it is known to induce cell apoptosis through mitochondrial depolarisation,
phosphatidylserine translocation and caspase activation (Cregan et al., 2013). Treatment with ACh
10 µM, NIC 10 µM and MLA 100 µM had no effect on the percentage of cells in sub-G1. A
significant (P < 0.001) increase in DNA fragmentation was seen for all other treatments in all cell
lines compared to untreated controls, with the exception of GYM 10 µM and ADI 250 µM
treatment of MDA-MB-231 cells (no significant change in DNA fragmentation).
MECA 1000 µM and 5-HT 1000 µM produced consistent increases in the percentage of cells
in sub-G1 compared to untreated cells, at similar levels to the positive control (CDDP 20 µM)
(Figure 9). The significant change in cells in sub-G1 was consistent with the cell viability results for
each treatment, where significant decreases in cell death were seen over the same concentrations
(Figure 4A and Figure 4D). A significant ~2-fold increase in the sub-G1 cells was also seen
following PnTX 5 µM treatment of all cell lines (Figure 9). However, despite similar viability
losses between the treatments (Figure 4B), PnTX sub-G1 levels were substantially lower than those
seen following MECA (>10-fold) and 5-HT (>12-fold) treatment. Treatment with GYM 10 µM
produced a significant increase in apoptotic HT29 (Figure 9B) and PC3 (Figure 9C) cells, with
levels similar to PnTX treatment, which was not reflective of cell viability results (Figure 4C).
61
A
Untr
eate
d M
AC
h 10
M
NIC
10
M
MEC
A 1
000
M
PnTX
5
M
GY
M 1
0 M
5-H
T 1
000
M
AD
I 25
0 M
MLA
100
M
CD
DP 2
0
0
1
2
3
4
5
20
30
40
50
***
***
***
***
Cell
s in
Su
b-G
1 (
%)
B
Untr
eate
d M
AC
h 10
M
NIC
10
M
MEC
A 1
000
M
PnTX
5
M
GY
M 1
0 M
5-H
T 1
000
M
AD
I 25
0 M
MLA
100
M
CD
DP 2
0
0
5
10
15
2030
40
50
60
70
80 ***
***
***
***
***
***
Cell
s in
Su
b-G
1 (
%)
C
Untr
eate
d M
AC
h 10
M
NIC
10
M
MEC
A 1
000
M
PnTX
5
M
GY
M 1
0 M
5-H
T 1
000
M
AD
I 25
0 M
MLA
100
M
CD
DP 2
0
0
2
4
6
8
1030
40
50
60
***
***
***
***
***
***
Cell
s in
Su
b-G
1 (
%)
Figure 9: Cell cycle analysis of antagonist treated MDA-MB-231 (A), HT29 (B) and PC3 (C) cell lines. Cells were
treated with ACh 10 µM, NIC 10 µM, MECA 1000 µM, PnTX 5 µM, GYM 10 μM, 5-HT 1000 µM, ADI 250 µM, and
MLA 100 µM for 72 hours prior to ethanol-fixation, propidium iodide staining and fluorescence-activated cell sorting
(FACS). Additionally, untreated cells were used as a negative control, while CDDP 20 µM treatment was used as a
positive control. The mean percentage of cells in the sub-G1 stage of the cell cycle has been calculated for each sample.
Values are displayed as the mean percent of cells in sub-G1, with error bars representing SEM (n = 5). A one-way
ANOVA with a Dunnett’s post-hoc test was used to determine significance compared to the untreated control (*** P <
0.001).
In addition to the Sub-G1 apoptotic group, the percentage of cells within each stage of the
cell cycle, including, G0/G1, S and G2/M, were determined to examine whether nAChR treatment
caused mitotic inhibition during any stage of cell division. The various stages of the cell cycle were
elucidated from the interactions of DNA with the fluorescent PI stain. Cells preparing for cell
division will contain increasing amounts of DNA; therefore, displaying a comparative increase in PI
fluorescence (Wlodkowic et al., 2011).
62
No consistent change in the percentage of cells within each cell cycle region was seen
following GYM 10 μM and MLA 100 μM treatment of MDA-MB-231 (Figure 10), HT29 (Figure
11) and PC3 (Figure 12) cells. MECA 1000 μM treatment produced a significant decrease in the
percentage of cells in G0/G1, S and G2/M of the cell cycle, compared to untreated cells. The
observed decrease correlated with a large increase in sub-G1 cells following MECA treatment.
Similar effects were observed following 5-HT treatment of MDA-MB-231, HT29 and PC3 cells.
Changes to the number of cells within each stage of the cell cycle were variable following
PnTX 5 μm and ADI 250 μm treatment. PnTX treatment caused a significant increase (P < 0.001) in
MDA-MB-231 (Figure 10B) and PC3 (Figure 12B) cells in the G0/G1 region of the cell cycle,
suggesting PnTX may induce cell growth inhibition through G0/G1 arrest. This effect was not
observed in HT29 cells following PnTX treatment. Interestingly, PnTX treatment of HT29 cells did
produce a significant increase in cells within the S and G2/M regions, compared to untreated cells
(P < 0.001) (Figure 11B). Comparable results were seen following ADI 250 μm treatment of MDA-
MB-231 (Figure 10E), HT29 (Figure 11E) and PC3 (Figure 12E) cells.
63
A Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100Untreated
MECA 1000 M
***
***
***
***
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
B Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100Untreated
PnTX 5 M
***
***
***
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
C Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100Untreated
GYM 10 M
***
***
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
D Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100Untreated
5-HT 1000 M
***
***
***
***
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
E
Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100 Untreated
ADI 250 M
***
***
***
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
F Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100Unteated
MLA 100 M
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
Figure 10: Cell cycle analysis of treated MDA-MB-231 cells measured via flow cytometry. Cells were treated with
MECA 1000 µM (A), PnTX 5 µM (B), GYM 10 μM (C), 5-HT 1000 µM (D), ADI 250 µM (E), and MLA 100 µM (F) for
72 hours prior to ethanol-fixation, propidium iodide staining and fluorescence-activated cell sorting. Untreated cells
were used as a negative control. The mean percentage of cells in each stage of the cell cycle has been calculated for
each sample. The error bars represent SEM (n = 5). A two-way ANOVA with a Bonferroni post-hoc test was used to
determine significance compared to the untreated control (*** P < 0.001).
64
A Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100Untreated
MECA 1000 M******
***
***
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
B Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100Untreated
PnTX 5 M
***
***
***
***
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
C Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100Untreated
GYM 10 M
***
**
***
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
D Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100Untreated
5-HT 1000 M
***
***
***
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
E
Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100 Untreated
ADI 250 M
***
******
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
F Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100Unteated
MLA 100 M
***
***
***
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
Figure 11: Cell cycle analysis of treated HT29 cells measured via flow cytometry. Cells were treated with MECA 1000
µM (A), PnTX 5 µM (B), GYM 10 μM (C), 5-HT 1000 µM (D), ADI 250 µM (E), and MLA 100 µM (F) for 72 hours
prior to ethanol-fixation, propidium iodide staining and fluorescence-activated cell sorting. Untreated cells were used
as a negative control. The mean percentage of cells in each stage of the cell cycle has been calculated for each sample.
The error bars represent SEM (n = 5). A two-way ANOVA with a Bonferroni post-hoc test was used to determine
significance compared to the untreated control (*** P < 0.001).
65
A Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100Untreated
MECA 1000 M
***
*
***
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
B Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100Untreated
PnTX 5 M
***
***
***
***
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
C Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100Untreated
GYM 10 M
***
***
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
D Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100Untreated
5-HT 1000 M
***
***
**
***
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
E
Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100 Untreated
ADI 250 M
***
**
***
***
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
F Sub-G1
G0/
G1 S
G2/
M
0
20
40
60
80
100Unteated
MLA 100 M
Pro
po
rtio
n o
f cell
s in
ea
ch
ph
ase
(%
)
Figure 12: Cell cycle analysis of treated PC3 cells measured via flow cytometry. Cells were treated with MECA 1000
µM (A), PnTX 5 µM (B), GYM 10 μM (C), 5-HT 1000 µM (D), ADI 250 µM (E), and MLA 100 µM (F) for 72 hours
prior to ethanol-fixation, propidium iodide staining and fluorescence-activated cell sorting. Untreated cells were used
as a negative control. The mean percentage of cells in each stage of the cell cycle has been calculated for each sample.
The error bars represent SEM (n = 5). A two-way ANOVA with a Bonferroni post-hoc test was used to determine
significance compared to the untreated control (* P < 0.05, ** P < 0.01, *** P < 0.001).
66
3.2.3 Flow Cytometry - TMRE
To further investigate early apoptosis of the various cancer cell lines using flow cytometry,
the TMRE stain was used on cell samples treated with the most cytotoxic nAChR modulating
compounds (MECA, PnTX, 5-HT and ADI). TMRE is a positively-charged dye that accumulates in
active mitochondria due to the negatively charged nature of the mitochondrial membrane potential.
Therefore, a left shift in the FL2-H fluorescence peak compared to untreated cells indicates an
inability of the mitochondria to sequester the TMRE due to depolarisation of the mitochondrial
membrane potential, as observed during cell apoptosis..
The TMRE histograms for the individual treatments and cell types have been presented in
Figure 13A, with percentage of cells with low TMRE intensity graphed (Figure 13B). High levels
of TMRE staining were shown in the untreated cell samples, with consistent shifts in FL2-H peaks
following treatment with the CDDP 20 µM positive control (indicative of a loss in mitochondrial
membrane permeability during early apoptosis). Results following ADI 1000 µM for 72 hours
showed a complete shift in FL2-H, with diminutive staining of active mitochondria. MECA 1000
µM and 5-HT 1000 µM treatment showed shifts in FL2-H peak comparable to the CDDP positive
control. PnTX 5 µM induced only a minor shift in FL2-H while still showing considerable TMRE
staining of active mitochondria. The results seen following PnTX 5 µM treatment were
considerably less pronounced compared to all other treatments (Figure 13B), producing only a small
but significant change from the untreated control, despite the similar effects on cell viability loss
(Figure 4B).
67
A Untreated MECA 1000 µM PnTX 5 µM 5-HT 1000 µM ADI 1000 µM CDDP 20 µM
MDA-
MB-
231
HT29
PC3
TMRE
B
Untr
eate
d M
MEC
A 1
000
M
PnTX
5
M
5-H
T 1
000
M
AD
I 10
00
M
CD
DP 2
0
0
10
20
30
40
50
60
70
80
90
100MDA-MB-231
HT29
PC3
All Combinations = ***
Cell
s w
ith
Low
TM
RE
In
ten
sity
(%
)
Figure 13: Evaluation of the mitochondrial transmembrane potential of MDA-MB-231, HT29 and PC3 cell lines
measured via flow cytometry to identify the early stages of cell apoptosis. Cells were treated with MECA 1000 µM,
PnTX 5 µM, 5-HT 1000 µM or ADI 1000 µM for 72 hours prior to TMRE staining and fluorescence-activated cell
sorting. Additionally, untreated cells were used as a negative control, while CDDP 20 µM treatment was used as a
positive control. Dot-plots represent FL2-H fluorescence versus cell number, where a shift in peak from left to right
signifies a collapse in mitochondrial transmembrane potential (A). The relative percentage of cells with low TMRE
intensity was graphed (B) with the error bars representing the mean ± SEM (n = 3). A one-way ANOVA with a
Bonferroni post hoc was used to determine significance between all treatments for each cell line (*** P < 0.001).
68
3.2.4 Flow Cytometry - Annexin V/PI
The Annexin V/PI double staining was assessed by flow cytometry to distinguish between
early apoptotic and necrotic cell death following the treatment of cancer cells with the same nAChR
modulating compounds used in TMRE studies. Quadrants were determined based on the Annexin V
(FL1-H) and PI (FL2-H) staining of untreated cells, and kept constant across all treatments. Cells
within the left-bottom quadrant represent viable cells. A shift of cells into the right-bottom quadrant
represents phosphatidylserine translocation to the cell surface through an increase in Annexin V
staining, indicative of early apoptosis. As cells enter late apoptosis, PI may penetrate the
compromised cell wall and stain the DNA, as indicated by the presence of cells within the right-top
quadrant. Similarity, secondary necrotic cells demonstrate a similar increase in both PI and Annexin
V staining as the cell wall deteriorates during necrosis; while an increase in PI staining with no
change in Annexin V staining has not been clearly characterised in the literature (Wlodkowic et al.,
2011).
The results of Annexin V/PI double staining of treated MDA-MB-231 cells have been
presented in Figure 14. Low basal levels of apoptotic and necrotic cells were seen in the untreated
control cells (6.55%). Considerable increases in Annexin+/PI- (early apoptosis) and Annexin+/PI+
(late apoptosis) stained cells were seen across all treatments, including the CDDP positive control.
ADI 1000 µM and MECA 1000 µM produced the greatest increase in apoptotic cells, with 80.49%
and 71.53% of cells, respectively, within the positively stained Annexin V region. Lower levels of
Annexin+/PI- and Annexin+/PI+ stained cells were seen following PnTX 5 µM (41.85%) and 5-HT
1000 µM (31.13%) treatment, relative to ADI and MECA treatment. Nevertheless, the results
obtained following PnTX and 5-HT treatment were still well above the 7.57% seen within the
untreated sample. The same substantial increases seen in Annexin+/PI- and Annexin+/PI+ stained
cells for all treatments compared to untreated cells was not seen within the Annexin-/PI+ quadrant.
A maximum 3-fold increase in Annexin-/PI+ cells following MECA 1000 µM treatment was
observed compared to untreated cells.
69
The percentage of MDA-MB-231 cells within the Annexin+/PI- bottom right quadrant has
been graphed in Figure 14G. All treatments, including the CDDP positive control, caused a
significant increase in Annexin+/PI- cells compared to control (P < 0.001). The significant increase
in Annexin+/PI- cells following treatment with MECA, PnTX, 5-HT and ADI indicates an increase
in the presence of early apoptotic cells.
70
Untreated MECA 1000 µM PnTX 5 µM P
rop
idiu
m I
od
ide
A B C
Annexin V
5-HT 1000 µM ADI 1000 µM CDDP 20 µM
Pro
pid
ium
Io
did
e
D E F
Annexin V
G
Untr
eate
d M
MEC
A 1
000
M
PnTX
5
M
5-H
T 1
000
M
AD
I 10
00
M
CD
DP 2
0
0
5
10
15
2040
45
50
*** ***
***
***
***
Earl
y A
pop
toti
c C
ells
(%
)
Figure 14: Flow cytometric detection of apoptotic and necrotic cells by fluorescence-activated cell sorting of propidium
iodide and Annexin V labelled MDA-MB-231 cells. Cells were treated with MECA 1000 µM (B), PnTX 5 µM (C), 5-HT
1000 µM (D) or ADI 1000 µM (E) for 72 hours prior to staining and analysis. Untreated cells (A) were used as a
negative control, while CDDP 20 µM (F) treatment was used as a positive control. Dot-plots represent individual
stained cells. The left-bottom quadrant represents viable cells (Annexin-/PI-). The right-bottom quadrant represents
early apoptotic cells (Annexin+/PI-). Late apoptotic or secondary necrotic cells are represented by the right-top
quadrant (Annexin+/PI+). Early apoptotic (Annexin+/PI-) cells were graphed (G) with the error bars representing the
mean ± SEM (n = 3). A one-way ANOVA with a Dunnett’s post-hoc test was used to determine significance compared
to the untreated control (*** P < 0.001).
5.91%
62.96%
20.19%
10.94%
4.77%
14.75%
36.08%
44.41%
6.97%
40.05%
44.82%
8.15%
2.95%
89.48%
6.55%
1.02%
9.92%
18.55%
58.51%
13.02%
5.16%
53.00%
28.03%
13.82%
71
Comparable results were seen within the HT29 treated cell samples, as shown in Figure 15,
with considerable increases in apoptotic and necrotic cells observed following the various
treatments, compared to the low basal levels of the control. ADI 1000 µM and MECA 1000 µM
treatment again produced the greatest increase in apoptotic cells, with 97.59% and 62.96% of cells
within the Annexin+ region for the respective treatments. Lower levels of apoptotic cells were seen
following PnTX 5 µM and 5-HT 1000 µM treatment. PnTX treatment produced a slight increase in
Annexin+ cells (24.96%) compared to the 7.81% for untreated cells; while 5-HT was comparable to
PnTX treatment with 28.74% of cells within the Annexin+ region. Significant increases in Annexin-
/PI+ stained cells were seen following MECA 1000 µM, 5-HT 1000 µM and CDDP 20 µM
treatment, with values of 22.57%, 19.93% and 14.26%, respectively, compared to untreated cells
(3.90%). Such large increases in Annexin-/PI+ stained cells was unique to HT29 cells, and not seen
in the other cell lines tested.
Significant increases in early apoptotic Annexin+/PI- cells were seen following treatment with
MECA (P < 0.001), PnTX (P < 0.001) and ADI (P < 0.05), compared to untreated cells (Figure
15G). No significant change in Annexin+/PI- cells was observed following 5-HT treatment,
compared to control. However, a large increase in secondary necrotic/late apoptotic cells following
5-HT treatment was observed, compared to untreated cells, despite the lack of early apoptotic cells.
72
Untreated MECA 1000 µM PnTX 5 µM P
rop
idiu
m I
od
ide
A B C
Annexin V
5-HT 1000 µM ADI 1000 µM CDDP 20 µM
Pro
pid
ium
Io
did
e
D E F
Annexin V
G
Untr
eate
d M
MEC
A 1
000
M
PnTX
5
M
5-H
T 1
000
M
AD
I 10
00
M
CD
DP 2
0
0
5
10
15
20
*** ******
*
Earl
y A
pop
toti
c C
ell
s (%
)
Figure 15: Flow cytometric detection of apoptotic and necrotic cells by fluorescence-activated cell sorting of propidium
iodide and Annexin V labelled HT29 cells. Cells were treated with MECA 1000 µM (B), PnTX 5 µM (C), 5-HT 1000
µM (D) or ADI 1000 µM (E) for 72 hours prior to staining and analysis. Untreated cells (A) were used as a negative
control, while CDDP 20 µM (F) treatment was used as a positive control. Dot-plots represent individual stained cells.
The left-bottom quadrant represents viable cells (Annexin-/PI-). The right-bottom quadrant represents early apoptotic
cells (Annexin+/PI-). Late apoptotic or secondary necrotic cells are represented by the right-top quadrant
(Annexin+/PI+). Early apoptotic (Annexin+/PI-) cells were graphed (G) with the error bars representing the mean ±
SEM (n = 3). A one-way ANOVA with a Dunnett’s post-hoc test was used to determine significance compared to the
untreated control (* P < 0.05, *** P < 0.001).
19.93%
51.32%
24.33%
4.41%
1.92%
0.49%
89.16%
8.43%
14.26%
51.56%
24.76%
9.41%
3.90%
88.28%
4.24%
3.57%
22.57%
14.47%
53.11%
9.85%
7.72%
67.32%
15.02%
9.94%
73
The cell death profiles of PC3 cells displayed a high level of apoptotic cells within the
untreated control, with 54.34% of cells within the Annexin+ region due to excessive confluency of
the cells at 72 hours (Figure 16). Increases in apoptotic cells were still however seen following
treatment with the various compounds. ADI 1000 µM and 5-HT 1000 µM produced the greatest cell
death, with respective Annexin+ values of 98.37% and 92.62 %. PnTX 5 µM treatment displayed
apoptotic results similar to the CDDP 20 µM control, with 85.57% of cells within the Annexin+
region. MECA 1000 µM treatment produced only a minor increase in Annexin+ apoptotic cells, with
a value of 62.83%, compared to untreated cells. Similar levels of Annexin-/PI+ stained cells were
observed across all treatments, contrary to results seen for the other cell lines tested.
Treatment with MECA and PnTX were the only two treatments to produce significant
increases in early apoptotic cells, compared to untreated cells (Figure 16G). However, further
analysis of the dot-plots show all treatments produced a consistent reduction in viable cells, with
large populations of cells within the secondary necrotic/late apoptotic region.
74
Untreated MECA 1000 µM PnTX 5 µM P
rop
idiu
m I
od
ide
A B C
Annexin V
5-HT 1000 µM ADI 1000 µM CDDP 20 µM
Pro
pid
ium
Io
did
e
D E F
Annexin V
G
Untr
eate
d M
MEC
A 1
000
M
PnTX
5
M
5-H
T 1
000
M
AD
I 10
00
M
CD
DP 2
0
0
5
10
15
20
***
*
Earl
y A
pop
toti
c C
ell
s (%
)
Figure 16: Flow cytometric detection of apoptotic and necrotic cells by fluorescence-activated cell sorting of propidium
iodide and Annexin V labelled PC3 cells. Cells were treated with MECA 1000 µM (B), PnTX 5 µM (C), 5-HT 1000 µM
(D) or ADI 1000 µM (E) for 72 hours prior to staining and analysis. Untreated cells (A) were used as a negative
control, while CDDP 20 µM (F) treatment was used as a positive control. Dot-plots represent individual stained cells.
The left-bottom quadrant represents viable cells (Annexin-/PI-). The right-bottom quadrant represents early apoptotic
cells (Annexin+/PI-). Late apoptotic or secondary necrotic cells are represented by the right-top quadrant
(Annexin+/PI+). Early apoptotic (Annexin+/PI-) cells were graphed (G) with the error bars representing the mean ±
SEM (n = 3). A one-way ANOVA with a Dunnett’s post-hoc test was used to determine significance compared to the
untreated control (* P < 0.05, *** P < 0.001).
3.41%
3.98%
88.62%
4.00%
1.23%
0.40%
96.40%
1.97%
5.04%
4.68%
86.51%
3.77%
8.13%
37.53%
48.60%
5.74%
5.04%
32.13%
51.57%
11.26%
3.50%
10.93%
71.56%
14.01%
75
3.2.5 Discussion
Western blot analyses and flow cytometry studies were used to examine the role of the most
potent nAChR modulators in inducing carcinoma cell death. It was initially hypothesised that the
selective nAChR antagonists would display the greatest cytotoxic effects on the various carcinoma
cell lines. The cell viability results previously discussed in Section 3.1 Effects of nAChR
Antagonists on Cell Viability demonstrated that the selective α7/α4β2 nAChR antagonist, PnTX,
displayed the most potent cytotoxic effects, while other selective antagonists, such as GYM and
MLA, were less toxic. The results of the cell viability studies were predicted to translate to the flow
cytometry results. In other words, the compounds with the greatest cytotoxic effects were expected
to have a significant impact on nAChR subtype expression, while inducing the greatest level of cell
apoptosis.
The Western blot analysis focused on the α7 nAChR subtype because of its widespread
expression, as observed in the previous studies. MDA-MB-231 cells showed a small increase in α7
expression following treatment with the various nAChR modulators, with the exception of 5-HT
treatment, while varying results were observed for all other cell lines. The literature reports that the
biological functions of the α7 nAChRs are increased following NIC exposure (Schuller, 2009).
Plummer and colleagues demonstrated the ability of the α7 nAChR to be significantly upregulated
in NCL-H69 human lung carcinoma cells following treatment with 100 ρM NNK. Changes in the
expression of the α7 nAChR subtype could be seen as early as 60 minutes following treatment;
however, the increase in expression was not observed at 24 hours post treatment (Plummer 3rd et
al., 2005). The present results following ACh or NIC treatment demonstrate little change in α7
expression across the cell types tested. A lack of a consistent change in α7 expression may be
attributed to the 72 hour treatment time, with changes in expression potentially occuring earlier in
the treatment period, as suggested by the literature. Similarly, no consistent changes in α7
expression were observed following treatment with the various nAChR antagonists. It was predicted
that the nAChR antagonists would produce a reduction in the α7 nAChR expression; however, the
76
observed results showed inconsistent changes between cell lines with no clear trends in α7
expression following treatment.
Tests to investigate the mechanism of cell death for the most cytotoxic nAChR modulators
were conducted on the MDA-MB-231, HT29 and PC3 cell lines. Flow cytometry cell cycle analysis
of treated cells following PI staining showed a significant increase in fractional DNA content
following MECA, PnTX, GYM (excluding MDA-MB-231 cells), 5-HT and ADI treatment,
compared to untreated cells. Interestingly, PnTX treatment induced small, but significant increases
in the percentage of cells in the sub-G1 fraction, indicating lower levels of apoptosis compared to
the other treatments investigated. An analysis of the complete cell cycle of MDA-MB-231 and PC3
cells indicated that PnTX treatment delayed the progression of cells from the G0/G1 phase into the
S phase. Similar results were observed following TMRE staining, with all treatments investigated
inducing significant mitochondrial membrane permeability, characteristic of early apoptosis.
Annexin V/PI double stain demonstrated significant increases in the presence of early apoptotic
cells and late apoptotic/necrotic cells following a 72 hour treatment period. PnTX showed a
consistently high level of early apoptotic cells compared to the other treatments tested, however
displayed lower levels of late apoptotic/necrotic cells, indicating a delayed onset of cell apoptosis
compared to the other nAChR antagonists.
The PnTX induced cell-cycle arrest in the G0/G1 phase may result from changes in [Ca2+]
through antagonism of α7 and α4β2 nAChRs. Transient rises in [Ca2+] are integral in the process of
cell division. Rises in [Ca2+], both from intracellular pools and the extracellular environment, occur
during the early stages of the cell cycle G1 phase. Although cell division requires a vast array of
physiological responses, Ca2+ acts as a ubiquitous second messenger, initiating many of the
processes involved during the cell cycle (Kahl et al., 2003). Current literature has demonstrated that
Ca2+ channel blockers, such as amlodipine, induce G1 phase accumulation by alleviating Ca2+ entry
into the cell. Amlodipine (30 μM) treatment of A431 human epidermoid carcinoma cells
demonstrated an ~20% increase in cells within the G0-G1 phase of the cell cycle after 48 hours of
77
treatment, compared to control cells. The reduction in [Ca2+] significantly decreased the expression
of cyclin D1 and CDK4, specific proteins involved in cell progression through the G1 phase
(Yoshida et al., 2007). The present results suggest a possible link between PnTX treatment and
reductions in [Ca2+], resulting in G0/G1 arrest and an inhibition of cell proliferation. Changes in
[Ca2+] would explain why relatively lower levels of sub-G1 cells and TMRE staining were observed
following PnTX treatment, despite possessing similar reductions in cell viability compared to other
treatments investigated. To support this theory, studies looking at the changes in [Ca2+] following
PnTX treatment were undertaken, as discussed in the next chapter.
The majority of treatments investigated induced significant levels of apoptosis in the treated
cells following 72 hours of treatment, demonstrating increases in fragmented DNA, mitochondrial
membrane permeability and phosphatidylserine translocation, indicative of early apoptosis. The
present results are consistent with those previously seen for nAChR antagonists. APS8, a non-
competitive and selective α7 nAChR antagonist, has been investigated due to its ability to inhibit
NSCLC cells growth and activate apoptotic pathways. APS8 treatment of A549 and SKMES-1 cells
caused significant mitochondrial depolarisation, leading to the release and activation of several
apoptotic factors involved in the intrinsic pathway (Zovko et al., 2013). Increased expression of
cytochrome C and SMAC, which are released following mitochondrial permeabilisation, were
observed following APS8 exposure, inducing the activation of caspase-9, a common inducer of the
intrinsic pathway (Zovko et al., 2013). PnTX, the most selective and cytotoxic α7 nAChR
antagonists investigated, induced similar mitochondrial permeabilisation in MDA-MB-231, HT29
and PC3 carcinoma cell lines. In addition, significant levels of Annexin+/PI-, indicative of early
apoptosis, suggest PnTX may induce apoptosis in a similar manner to that seen following APS8
treatment.
The cell death profile of PnTX differed from the other nAChR antagonists investigated,
despite showing similar effects in the earlier cell viability studies. PnTX treated cells displayed a
lower proportion of cells in the sub-G1 phase, lower TMRE staining and higher levels of early
78
apoptotic cells, during the flow cytometric studies. The changes in the cell cycle of the investigated
cells suggest a possible mechanism by which PnTX treatment could reduce cancer cell proliferation.
Alternatively, the shift in TMRE staining suggests possible induction of the intrinsic apoptotic
pathway, contributing to the early apoptotic effects of PnTX. The following chapter looked at
investigating the inhibition of various downstream apoptotic messengers to further deduce the
mechanism behind the cytotoxic effects of PnTX.
3.3 Inhibition of nAChR Antagonist Induced Cell Death
In the previous studies it was shown that nAChR treatment induces potent apoptotic effects in
carcinoma cell lines. To investigate this phenomenon further, apoptotic inhibitors were used to
determine whether inhibiting common apoptotic pathways would reduce the cytotoxic effects of the
nAChR modulators (Aim III).
The apoptotic pathway, as detailed in Section 1.2.4 Anti-Apoptotic Effects of nAChRs in
Cancer Survival, is regulated by several key intermediates. Following activation of the extrinsic
death receptors, or the intrinsic response to cellular stress, the apoptotic pathway is initiated. One
key pathway is the activation of the Ca2+-dependent proteases, calpains, in response to excessive
intracellular Ca2+ levels causing cleavage of the Na+/Ca2+ mitochondrial exchanger leading to
mitochondrial Ca2+ accumulation. This induces changes in the inner mitochondrial membrane
potential, which stimulates the opening of the mitochondrial permeability transition pore (Elmore,
2007; Smith et al., 2012). Cytochrome C and small mitochondria-derived activator of caspases
proteins are released through the pore, activating several downstream mediators, such as cysteine-
dependent aspartic proteases (also known as caspases). The downstream activation of caspase-3
stimulates the final stages of apoptosis, inducing cytoskeletal reorganisation and disintegration of
the cell into apoptotic bodies (Elmore, 2007; Smith et al., 2012). Several of these integral steps
were targeted by the following studies to better understand the cytotoxic effects of nAChR
modulators on carcinoma cells.
79
3.3.1 Cell Death Inhibitors
To investigate the possible apoptotic pathways affected by the treatment with the most potent
nAChR modulator tested, PnTX, various cell death inhibitors were used. MDA-MB-231, HT29,
PC3 and LN18 cells were treated with cell death inhibitors alone and in combination with PnTX 5
µM to examine any possible decrease in cell death caused by the PnTX treatment.
Cyclosporin A (CsA), a potent inhibitor of the opening of the mitochondrial permeability
transition pore during apoptosis through binding to the cyclophilin D protein, was the first inhibitor
investigated (Basso et al., 2008) (Figure 17). Other inhibitors tested included ALLN (Figure 18),
BAPTA (Figure 19) and zVAD (Figure 20). ALLN reversibly blocks the Ca2+-dependent neutral
cysteine protease calpain I, commonly involved in various cellular processes, including
proliferation, apoptosis and differentiation (Chen et al., 2002). BAPTA, a known Ca2+ chelator,
inhibits apoptosis through decreasing [Ca2+] levels critical in the activation of apoptotic caspases
(Zhao et al., 2005). Furthermore, zVAD inhibits the proteolytic activity of apoptotic enzymes
through irreversibly binding to the catalytic site of caspase proteases (Gamen et al., 2000).
The combination treatment of CsA + PnTX 5 µM did no improve the viability of any of the
four cell lines tested, compared to PnTX 5 µM treatment alone (Figure 17). Conversely, CsA 10
µM + PnTX 5 µM treatment produced a significant (P < 0.01) 38% decrease in cell viability of
MDA-MB-231, compared to PnTX 5 µM treatment alone (Figure 17A).
Similar results were seen across other cell death inhibitors tested with decreases in cell
viability seen for MDA-MB-231 cells following ALLN 5 µM + PnTX 5 µM (P < 0.001), BAPTA 1
µM + PnTX 5 µM (not significant) and zVAD 30 µM + PnTX 5 µM (P < 0.01), compared to PnTX
treatment alone. No significant changes in MDA-MB-231 cell viability were seen following CsA,
BAPTA or zVAD treatment alone, compared to untreated cells, with only ALLN treatment alone
showing a significant decrease in cell viability (P < 0.05).
80
A U
ntrea
ted
DM
SO M
PnTX
5
M
CsA
10
CsA
+ P
nTX
0
20
40
60
80
100
120
**
No.
of
Via
ble
Cel
ls C
om
pare
d t
o C
on
trol
BU
ntrea
ted
DM
SO M
PnTX
5
M
CsA
1
CsA
+ P
nTX
0
20
40
60
80
100
120
No.
of
Via
ble
Cel
ls C
om
pare
d t
o C
on
trol
C U
ntrea
ted
DM
SO M
PnTX
5
M
CsA
1
CsA
+ P
nTX
0
20
40
60
80
100
120
No.
of
Via
ble
Cel
ls C
om
pare
d t
o C
on
trol
D U
ntrea
ted
DM
SO M
PnTX
5
M
CsA
1
CsA
+ P
nTX
0
20
40
60
80
100
120
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
Figure 17: Examination of the effect of the cell death inhibitor Cyclosporin A (CsA) on PnTX induced cytotoxicity in
MDA-MB-231 (A), HT29 (B), PC3 (C) or LN18 (D) cell lines, following 72 hours of treatment. Cells were treated with
either PnTX 5 µM, CsA 1 μM or 10 µM, or a combination of the two following a 24 hour seeding period. Untreated
cells were used as a negative control, while DMSO was included as a vehicle control. Cell viability was established via
the MTT assay. Values are displayed as the mean viability (as a percentage) of the viability of control cells, with error
bars representing SEM (n = 3). A one-way ANOVA with a Bonferroni post-hoc test was used to determine significance
between treatment groups. ** represents P < 0.01 between the PnTX and the CsA + PnTX treatment groups.
81
A
Untr
eate
d
DM
SO M
PnTX
5
M
ALLN
5
ALLN
+ P
nTX
0
20
40
60
80
100
120
***
No.
of
Via
ble
Cel
ls C
om
pare
d t
o C
on
trol
B
Untr
eate
d
DM
SO M
PnTX
5
M
ALLN
1
ALLN
+ P
nTX
0
20
40
60
80
100
120
No.
of
Via
ble
Cel
ls C
om
pare
d t
o C
on
trol
C
Untr
eate
d
DM
SO M
PnTX
5
M
ALLN
5
ALLN
+ P
nTX
0
20
40
60
80
100
120
*
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
D
Untr
eate
d
DM
SO M
PnTX
5
M
ALLN
1
ALLN
+ P
nTX
0
20
40
60
80
100
120
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
Figure 18: Examination of the effect of the cell death inhibitor ALLN on PnTX induced cytotoxicity in MDA-MB-231
(A), HT29 (B), PC3 (C) or LN18 (D) cell lines, following 72 hours of treatment. Cells were treated with either PnTX 5
µM, ALLN 1 μM or 5 µM, or a combination of the two following a 24 hour seeding period. Untreated cells were used as
a negative control, while DMSO was included as a vehicle control. Cell viability was then established via the MTT
assay. Values are displayed as the mean viability (as a percentage) of the viability of control cells, with error bars
representing SEM (n = 3). A one-way ANOVA with a Bonferroni post-hoc test was used to determine significance
between treatment groups. * represents P < 0.05 while *** represents P < 0.001 between the PnTX and the ALLN +
PnTX treatment groups.
82
A
Untr
eate
d
DM
SO M
PnTX
5
M
BA
PTA
1
BA
PTA
+ P
nTX
0
20
40
60
80
100
120
No.
of
Via
ble
Cel
ls C
om
pare
d t
o C
on
trol
B
Untr
eate
d
DM
SO M
PnTX
5
M
BA
PTA
1
BA
PTA
+ P
nTX
0
20
40
60
80
100
120
No.
of
Via
ble
Cel
ls C
om
pare
d t
o C
on
trol
C
Untr
eate
d
DM
SO M
PnTX
5
M
BA
PTA
10
BA
PTA
+ P
nTX
0
20
40
60
80
100
120
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
D
Untr
eate
d
DM
SO M
PnTX
5
M
BA
PTA
0.5
BA
PTA
+ P
nTX
0
20
40
60
80
100
120
140
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
Figure 19: Examination of the effect of the cell death inhibitor BAPTA on PnTX induced cytotoxicity in MDA-MB-231
(A), HT29 (B), PC3 (C) or LN18 (D) cell lines, following 72 hours of treatment. Cells were treated with either PnTX 5
µM, BAPTA 1 μM or 5 µM, or a combination of the two following a 24 hour seeding period. Untreated cells were used
as a negative control, while DMSO was included as a vehicle control. Cell viability was then established via the MTT
assay. Values are displayed as the mean viability (as a percentage) of the viability of control cells, with error bars
representing SEM (n = 3). A one-way ANOVA with a Bonferroni post-hoc test was used to determine significance
between treatment groups. No significant difference was seen between the PnTX and the BAPTA + PnTX treatment
groups.
83
A
Untr
eate
d
DM
SO M
CD
DP 1
0 M
PnTX
5
M
zVA
D 3
0
zVA
D +
CD
DP
zVA
D +
PnT
X
0
20
40
60
80
100
120
**
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
B
Untr
eate
d
DM
SO M
CD
DP 1
0 M
PnTX
5
M
zVA
D 2
0
zVA
D +
CD
DP
zVA
D +
PnT
X
0
20
40
60
80
100
120
No.
of
Via
ble
Cel
ls C
om
pare
d t
o C
on
trol
C
Untr
eate
d
DM
SO M
CD
DP 1
0 M
PnTX
5
M
zVA
D 3
0
zVA
D +
CD
DP
zVA
D +
PnT
X
0
20
40
60
80
100
120
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
D
Untr
eate
d
DM
SO M
CD
DP 1
0 M
PnTX
5
M
zVA
D 1
0
zVA
D +
CD
DP
zVA
D +
PnT
X
0
20
40
60
80
100
120
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
Figure 20: Examination of the effect of the cell death inhibitor zVAD on PnTX induced cytotoxicity in MDA-MB-231
(A), HT29 (B), PC3 (C) or LN18 (D) cell lines, following 72 hours of treatment. Cells were treated with either PnTX 5
µM, zVAD 10, 20 or 30 µM, or a combination of the two following a 24 hour seeding period. Untreated cells were used
as a negative control, DMSO was included as a vehicle control, while CDDP and CDDP + zVAD was used as a
positive control. Cell viability was then established via the MTT assay. Values are displayed as the mean viability (as a
percentage) of the viability of control cells, with error bars representing SEM (n = 3). A one-way ANOVA with a
Bonferroni post-hoc test was used to determine significance between treatment groups. ** represents P < 0.01 between
the PnTX and the zVAD + PnTX treatment groups.
3.3.2 Intracellular Calcium
Intracellular calcium ([Ca2+]) is involved in various cell signalling pathways, including the
cell apoptotic pathway. Based on the early results linking nAChR modulation and Ca2+ influx
within a cell (refer to the discussion in Section 3.2 Investigation of nAChR Antagonist Induced Cell
Death), the [Ca2+] levels of MDA-MB-231, HT29 and PC3 cells were measured following
84
treatment with the various nAChR modulators. The MDA-MB-231, HT29 and PC3 cell lines were
chosen based on their range of nAChR expression observed during Western blot studies (Figure 5).
Cells were incubated with 4 µM of the intracellular fluorescent Ca2+ indicator, Fura-2 AM, prior to
treatment. The [Ca2+] levels were then measured over 360 minutes post treatment, with the
fluorescence intensity ratio (340 nm/380 nm) displayed in Figure 21.
Characteristic results were seen following treatment with the known nAChR agonist, ACh 10
μM, in all cells lines tested. An instant increase in [Ca2+] was seen after 1-2 minutes of ACh
treatment, compared to untreated control cells. The increase in [Ca2+] was sustained over the first 15
minutes of recording, eventually returning to baseline after ~60 minutes in both HT29 and PC3 cell
lines. MDA-MB-231 [Ca2+] levels were less defined following ACh treatment. An initial increase in
[Ca2+] was seen, however results varied over the 360 minutes of recording with [Ca2+] levels
remaining above baseline.
No obvious change in [Ca2+] levels was seen in HT29 and PC3 cell lines following GYM 10
μM and MLA 100 μM treatment (Figure 21). The level of [Ca2+] for both treatments remained close
to baseline for the 300 minutes of recordings. Slight variations in [Ca2+] levels of MDA-MB-231
cells were seen following GYM and MLA treatment. However, [Ca2+] changes were inconsistent
and not characteristic of the trends seen in the other cell lines tested. The lack of [Ca2+] variation
following GYM and MLA treatment is consistent with the small changes in cell viability shown for
both treatments in Figure 4C and Figure 4F.
Despite the observed decrease in cell viability following treatment with the nAChR
antagonists MECA and ADI (Figure 4A and Figure 4E), treatment with each compound produced
an increase in [Ca2+] levels. MECA 1000 μM treatment of HT29 cells produced a noticeable
increase in [Ca2+], compared to untreated cells. The increase in [Ca2+] was delayed compared to the
response seen following ACh 10 μM treatment, and returned to baseline after ~60 minutes. A
similar, however more variable effect was seen in MDA-MB-231 cells, while the increase in [Ca2+]
levels following MECA treatment could not be repeated in PC3 cells. An obvious increase in [Ca2+]
85
levels was also observed following treatment of MDA-MB-231, HT29 and PC3 cells with ADI
1000 μM. The increase in [Ca2+] was similar across all three cell lines, lasting for ~15 minutes
following treatment with ADI.
Treatment with PnTX 5 μM or 5-HT 1000 μM produced the only consistent decreases in
[Ca2+] levels among the compounds tested. 5-HT 1000 μM treatment in HT29 and PC3 cells
produced a noticeable decrease in [Ca2+] levels, which returned to baseline ~120 minutes following
treatment. A greater and more sustained decrease in [Ca2+] levels was observed following PnTX 5
μM treatment across all cell lines tested. [Ca2+] levels for cell lines treated with PnTX did not return
to baseline levels, remaining consistently below the untreated cell group.
MDA-MB-231 HT29 PC3
ACh 10 µM
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
UntreatedACh 10 M
Time (Min)
340/3
80 R
ati
o
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
2.6
60 120 180 240 300 360
ACh 10 M
Untreated
Time (Min)
340/3
80 R
ati
o
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
ACh 10 M
Untreated
Time (Min)
340/3
80 R
ati
o
MECA 1000 µM
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
UntreatedMECA 1000 M
Time (Min)
340/3
80 R
ati
o
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
UntreatedMECA 1000 M
Time (Min)
340/3
80 R
ati
o
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
UntreatedMECA 1000 M
Time (Min)
340/3
80 R
ati
o
PnTX 5 µM
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
UntreatedPnTX 5 M
Time (Min)
340/3
80 R
ati
o
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
UntreatedPnTX 5 M
Time (Min)
340/3
80 R
ati
o
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
UntreatedPnTX 5 M
Time (Min)
340/3
80 R
ati
o
86
MDA-MB-231 HT29 PC3
GYM 10 µM
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
UntreatedGYM 10 M
Time (Min)
340/3
80 R
ati
o
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
UntreatedGYM 10 M
Time (Min)
340/3
80 R
ati
o
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
UntreatedGYM 10 M
Time (Min)
340/3
80 R
ati
o
5-HT 1000 µM
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
Untreated5-HT 1000 M
Time (Min)
340/3
80 R
ati
o
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
Untreated5-HT 1000 M
Time (Min)
340/3
80 R
ati
o
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
Untreated5-HT 1000 M
Time (Min)
340/3
80 R
ati
o
ADI 1000 µM
0 5 10 15
1.0
1.5
2.0
2.5
3.0
3.5
4.0
4.5
60 120 180 240 300 360
UntreatedADI 1000 M
Time (Min)
340/3
80 R
ati
o
0 5 10 15
1.0
1.5
2.0
2.5
3.0
3.5
4.0
60 120 180 240 300 360
UntreatedADI 1000 M
Time (Min)
340/3
80 R
ati
o
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
UntreatedADI 1000 M
Time (Min)
340/3
80 R
ati
o
MLA 100 µM
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
UntreatedMLA 100 M
Time (Min)
340/3
80 R
ati
o
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
UntreatedMLA 100 M
Time (Min)
340/3
80 R
ati
o
0 5 10 15
0.8
1.0
1.2
1.4
1.6
1.8
2.0
2.2
2.4
60 120 180 240 300 360
UntreatedMLA 100 M
Time (Min)
340/3
80 R
ati
o
Figure 21: Ca2+ accumulation through Fura-2 AM detection within MDA-MB-231, HT29 and PC3 cells following ACh
10 µM, MECA 1000 µM, PnTX 5 µM, GYM 10 µM, 5-HT 1000 µM, ADI 1000 µM or MLA 100 µM treatment.
Untreated cells were used as the negative control Cells were allowed to adhere overnight before being treated with 100
µL/well of 4 µM of the Ca2+ indicator Fura-2 AM. Cells were then treated with one of the corresponding nAChR
modulators. Fura-2 fluorescence was measured using a fluorescent plate reader at an excitation of 340 nm (λ1) and
380 nm (λ2), with an emission of 510 nm, over 300 minutes. The fluorescence intensity ratio (R = Fλ1/Fλ2) was
calculated for each time point and graphed with error bars representing SEM (n = 3).
87
3.3.3 Discussion
The effects of various nAChR antagonists on cancer cell viability and their capacity to induce
cell apoptosis has been discussed in the previous results sections. The current section further
investigated the mechanism of nAChR-induced cancer cell death. PnTX was chosen as the lead
compound based on its high affinity to antagonise nAChRs and interesting cytotoxic profile
compared to the other antagonists investigated. Various inhibitors of common apoptotic effectors
were employed in an attempt to reduce the cytotoxic effects of PnTX and further deduce the
mechanism of cell death. The results showed no significant inhibition of cancer cell apoptosis
following co-treatment of PnTX with CsA, ALLN, BAPTA and zVAD. Interestingly, the majority
of apoptotic inhibitors induced a further decrease in cell viability when combine with PnTX, while
having little effect on cell viability alone.
The flow cytometry results have suggested an increase in mitochondrial permeabilisation
following PnTX treatment (refer to Section 3.2.3 Flow Cytometry - TMRE). Changes in
mitcochondrial permeabilisation occur during activation of the intrinsic apoptotic pathway,
resulting in release of cytochrome C and a consequent downstream activation of the caspases. CsA
acts via inhibiting the cyclophilin D protein, preventing opening of the mitochondrial permeability
transition pore during apoptosis. It was hypothesised that co-treatment with CsA might alleviate
PnTX induced cytotoxicity through inhibition of the intrinsic apoptotic pathway. Interestingly
however, CsA treatment with PnTX induced a significant decrease in cell viability, compared to
PnTX treatment alone in MDA-MB-231 cells, while no change was observed in all other cell lines
tested. CsA possesses a secondary action through inhibiting phosphatase 2B activity, which may
explain why a further decrease in cell viability was observed (Marszalec et al., 2005).
Supplementary studies to investigate the effects of combination treatment with CsA and the broad-
spectrum nAChR inhibitor, MECA, in MDA-MB-231 cells were investigated in the following
section (Combination T).
88
Similarly, combination treatment of the caspase inhibitor zVAD and PnTX induced a decrease
in cell viability compared to PnTX treatment alone. Caspases exhibit proteolytic activity, which
when activated play an almost irreversible role in apoptosis. The activation of effector caspases
(caspase-3, -6 and -7) facilitates the execution pathway of apoptotic cell death, resulting in
breakdown of nuclear material and the formation of apoptotic bodies (Elmore, 2007). The present
results are inconsistent with those seen previously following inhibition of the nAChRs. APS8, a
known antagonist of α7 nAChRs, induced significant activation of caspase-9 and caspase-3
following treatment of A549 and SKMES-1 cancer cells (Zovko et al., 2013). Interestingly, NIC has
also been shown to inhibit the activation of caspase-3 following the treatment of Tca8113 oral
cancer cells with the known caspase inducer, cisplatin, suggesting a link between nAChR regulation
and caspase activation (Xu et al., 2007). A lack of restoration of cell viability following caspase
inhibition in the presence of PnTX suggests PnTX-induced cancer cell cytotoxicity acts via a
caspase-independent mechanism. The mechanisms mediating caspase-independent apoptosis are not
well characterised; however, several key cellular events are capable of occurring in the absence of
caspase activity. Phosphatidylserine externalisation (as demonstrated during flow cytometry studies
with Annexin V in Section 3.2.4) can occur independently of caspase activation through changes in
intracellular Ca2+ activity, altering the activity of the key mediators of phosphatidylserine
externalisation (Cummings et al., 2004). Furthermore, nuclear fragmentation (evident from flow
cytometry studies with PI in Section 3.2.2) as a result of the activation of DNAses may be
controlled through non-caspase-related pathways involving Ca2+-dependent DNAse proteins
(Nitahara et al., 1998). These findings again suggest a role of intracellular Ca2+ alterations in PnTX
cancer cytotoxicity.
The detrimental effects of PnTX on MDA-MB-231 and PC3 cell viability increased following
treatment with the Ca2+ activated calpain inhibitor, ALLN, while having no significant effect on
HT29 and LN18 viability compared to PnTX treatment alone. Calpains play a role in the execution
process of apoptosis following activation of the enzymes due to an increase in intracellular Ca2+.
89
Once activated, calpains degrade the membrane, cytoplasmic and nuclear structures during cell
apoptosis (Momeni, 2011). Changes in mitochondrial permeability and release from intracellular
stores during apoptosis induce an influx in the intracellular concentration of Ca2+, activating such
proteins as calpains. The present results suggest that the cytotoxic effects of PnTX may result from
decreases in intracellular Ca2+ due to the antagonistic effects at α7 and α4β2 nAChRs. This theory is
further consolidated through the observed effects of BAPTA on PnTX cytotoxicity. The Ca2+
chelator, BAPTA, has been utilised in apoptotic studies as a possible inhibitor by reducing the
intracellular influx in Ca2+ (Zhao et al., 2005). The present results demonstrated that, in some cases,
BAPTA potentiated the cytotoxic effects of PnTX on cancer cells.
Due to the results obtained in the apoptosis inhibitory studies, it was hypothesised that the
cytotoxic effects of the various antagonists, specifically PnTX, were due to changes in [Ca2+] influx
into the cell, inducing carcinoma cell apoptosis. PnTX treatment demonstrated an obvious and
sustained decrease in [Ca2+] levels of various carcinoma cells over the 360 minutes of testing
(Figure 21). Similar effects were not observed following treatment with the other antagonists, with
only 5-HT treatment producing a transient decrease in [Ca2+].
PnTX has been identified as a potent antagonist of α7 nAChRs. Recent literature analysing
the functional capacity of PnTX A and G revealed high affinity binding and potent antagonism for
the α7 subtype during binding and voltage-clamp electrophysiology studies (Bourne et al., 2015).
Activation of the α7 nAChRs causes large influxes of intracellular Ca2+, capable of inducing various
intracellular signalling pathways, without the additional coupling to voltage-gated calcium channels
(Garduño et al., 2012). This characteristic influx in [Ca2+] is responsible for many of the pro-
carcinogenic effects of nAChRs. The effects of PnTX on [Ca2+] were first identified during studies
conducted on P388 leukaemia cancer cells (Geiger et al., 2013); however, limited discussions of the
effects on PnTX on [Ca2+] have since been reported. The present results suggest that PnTX’s ability
to antagonise the highly Ca2+ permeable α7 nAChRs may be responsible for the observed decrease
in [Ca2+] levels. These changes in [Ca2+] levels following PnTX treatment could potentially explain
90
the differences in flow cytometric and apoptosis inhibition studies compared to the other treatments
tested. However, further studies are required to confirm the relationship between PnTX and the
decrease in [Ca2+] via α7 nAChR antagonism.
The potential effects of Ca2+ in context of the PnTX induced cell-cycle arrest in the G0/G1
phase have been discussed in the previous chapter. Cells have been shown to be most susceptible to
depletions in Ca2+ during the G1 phase of the cell cycle. Ca2+ is important for the expression of the
cell cycle genes, FOS, JUN and MYC, which when depleted, reduce cell cycle progression through
the G1 phase (Kahl et al., 2003). The cytotoxic effects of PnTX may also result from the decreases
in intracellular Ca2+. A decrease in Ca2+ influx from the extracellular environment can cause the cell
to release intracellular Ca2+ reserves from the endoplasmic reticulum. In cases such as in the
presence of PnTX, stress through the chronic depletion of Ca2+ from the endoplasmic reticulum can
result in cell apoptosis via the intrinsic apoptotic pathway (Delom et al., 2007). The cytotoxic
effects of PnTX through endoplasmic reticulum stress-induced apoptosis correlates with the
increases in fragmented DNA and mitochondrial membrane permeability observed during the flow
cytometry studies.
In contrast to PnTX, MECA and ADI treatment induced a significant transient increase in
intracellular Ca2+ in the three carcinoma cell lines tested. Increases in intracellular Ca2+ can also
initiate cell apoptosis. A significant influx in intracellular Ca2+ can activate cell death effectors,
such as calpains, which stimulate the cleavage of downstream apoptotic effectors. Initiation of this
pathway ultimately results in cell apoptosis through mitochondrial permeabilisation and cytochrome
C release (Roderick et al., 2008). On the basis of this information, the various apoptotic inhibitors
investigated with PnTX would theoretically have a significant effect on MECA and API induced
cell death. However, the increase in [Ca2+] due to MECA and ADI treatment was unexpected due to
the similar antagonistic effects of both compounds on nAChRs. In addition, a similar increase in
[Ca2+] was shown following treatment with ACh, while ACh has no cytotoxic effects on carcinoma
91
cells. This suggests that the cytotoxic effects of both MECA and ADI may operate through a Ca2+-
independent pathway, contrary to that of PnTX.
3.4 Combination Treatment
The final studies were conducted to explore the synergistic capacity of the most effective
nAChR modulators identified in Aims I-III to sensitise cancer cells to common chemotherapy
agents. It was hypothesised that the nAChR antagonists would enhance the cytotoxic nature of
chemotherapy agents, at concentrations that were relatively non-cytotoxic when administered alone.
Cell viability assay, flow cytometry and an isobologram analysis were utilised to identify the most
effective cytotoxic combination of nAChR modulators and chemotherapy agents.
3.4.1 CsA + MECA
Due to the consistent decrease in cell viability observed following treatment with the cell
death inhibitors in combination with PnTX in MDA-MB-231 cells, MECA (due to the limited
availability of PnTX) and CsA were further investigated across a range of toxic and non-toxic
concentrations to determine the combination relationship between the two compounds.
The effect of MECA 100 µM and 500 µM combination treatment with CsA 0.1 µM, 1.0 µM
and 5.0 µM on MDA-MB-231 cell viability was studied following 72 hours of exposure to the
treatment (Figure 22). No significant combinational effect was seen in cells treated with CsA 0.1
µM (Figure 22A). However, significant decreases in cell viability were seen following treatment
with CsA 1.0 µM and CsA 5.0 µM. CsA 1.0 µM + MECA 500 µM combination treatment produced
a significant decrease in cell viability (P < 0.001), compared to the single treatment with MECA
500 µM or CsA 1.0 µM (Figure 22B) alone. A similar effect was not seen with CsA 1.0 µM +
MECA 100 µM; while a strong combinational effect was seen with CsA 5.0 µM treatment (Figure
22C). CsA 5.0 µM + MECA 100 µM and CsA 5.0 µM + MECA 500 µM produced significant
decreases in cell viability, compared to the individual treatments (P < 0.001). The CsA 5.0 µM +
MECA 500 µM treatment demonstrated the greatest combinational effect, resulting in a 91.2%
92
decrease in cell viability, compared to a 23.2% and 8.7% decrease for the respective CsA 5.0 µM
and MECA 500 µM treatments alone.
A
Untr
eate
d M
MEC
A 1
00
M
MEC
A 5
00
M
CsA
0.1
M
M +
MEC
A 1
00
CsA
0.1
M
M +
MEC
A 5
00
CsA
0.1
0
20
40
60
80
100
120
******
No
. o
f V
iab
le C
ell
s C
om
pa
red
to
Co
ntr
ol
B
Untr
eate
d M
MEC
A 1
00
M
MEC
A 5
00
M
CsA
1.0
M
M +
MEC
A 1
00
CsA
1.0
M
M +
MEC
A 5
00
CsA
1.0
0
20
40
60
80
100
120
***
***
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
C
Untr
eate
d M
MEC
A 1
00
M
MEC
A 5
00
M
CsA
5.0
M
M +
MEC
A 1
00
CsA
5.0
M
M +
MEC
A 5
00
CsA
5.0
0
20
40
60
80
100
120
* ***
******
***
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
Figure 22: The cytotoxic effect of combined MECA and CsA treatment in MDA-MB-231 cells. MDA-MB-231 cells were
treated with MECA 100 µM or 500 µM and CsA 0.1 μM (A), 1.0 µM (B) or 5.0 µM (C), or a combination of the two for
72 hours following a 24 hour seeding period. Untreated cells were used as a negative control. Cell viability was then
established via the MTT assay. Values are displayed as the mean percent of the control viability, with error bars
representing SEM (n = 3). A one-way ANOVA with a Bonferroni post-hoc test was used to determine significance
between the different treatment groups (* P < 0.05, *** P < 0.001).
93
Given the substantial decrease in MDA-MB-231 viability following the combination
treatment with MECA and CsA, an isobologram was constructed to establish the relationship
between the two compounds. First, the IC50 values for each individual MECA and CsA treatment
were calculated and plotted on a dotplot. The line of additivity was then added, connecting the two
IC50 values. The line of additivity represents the theoretical effect of the two treatments, given each
treatment acts independently of the other. Next, the IC50 value for each treatment was calculated in
the presence of a known and constant concentration of the other treatment (e.g. IC50 of MECA in the
presence of CsA 5.0 μM). The various IC50 values are then plotted on the isobologram to determine
the relationship between the two treatments. In the event that the interaction between the two
treatments is more effective than the sum of the single effects (the line of additivity), then it can be
concluded that the treatments act synergistically. Alternatively, should the effect of the two
treatments be less effective than the sum of the individual effects, than an antagonistic relationship
is concluded. A sample diagram highlighting the line of additivity and the two possible
relationships between two treatments has been provided in Figure 23.
94
Figure 23: Isobologram example displaying the line of additivity, connecting the IC50 values for two individual
treatments. The line of additivity represents the theoretical effect of the two treatments, given each treatment acts
independently of the other. The line of synergy represents an interaction between two treatments that is more effective
than the sum of the individual effects. Conversely, the line of antagonism characterises an interaction between the two
treatments that is less effective than the sum of the individual effects for each treatment.
An isobologram demonstrating the relationship between MECA and CsA treatment of MDA-
MB-231 cells, with respect to cell death, is displayed in Figure 24. The IC50 values for the single
treatment of MECA and CsA were determined to be 673.6 μM and 28.90 μM respectively. In
addition, the IC50 values for MECA and CsA were determined when in the presence of a constant
concentration of the alternate treatment. The IC50 value for CsA decreased to 11.40 μM in the
presence of MECA 100 μM, and further decreased to 0.9482 μM in the presence of MECA 500 μM.
Similar reductions in the IC50 value for MECA were seen in the presence of CsA. The IC50 value
dropped to 364.2 μM for MECA in the presence of CsA 1.0 μM, while further decreasing to 149.0
μM when in the presence of CsA 5.0 μM. The decrease in IC50 values for MECA and CsA have
been displayed visually in Figure 24. The data points for the fixed ratio treatment combinations fall
95
well below the line of additivity, representing a pattern of synergistic effect, similar to that shown in
the example diagram in Figure 23.
Figure 24: Isobologram demonstrating the relationship between MECA and CsA treatment of MDA-MB-231 cells. Cells
were treated with a range of MECA (10 μM – 1500 μM) and CsA (0.1 μM – 40 μM) for 72 hours, following a 24 hour
seeding period, to determine the IC50 for each treatment. The line of additivity has been added between the MECA IC50
value (673.6 μM) and CsA IC50 value (28.90 μM). IC50 values for MECA and CsA, in the presence of constant
concentrations of the other treatment, are shown to determine the relationship between the two compounds, with
respect to cell death.
The Annexin V/PI double staining was assessed by flow cytometry to distinguish between
early apoptotic and necrotic cell death following the individual and combination treatment of MDA-
MB-231 cells. Cells were either left untreated, or treated with CsA 5 μM, MECA 500 μM, or the
combination of the two. The dotplots showing viable, apoptotic and necrotic cells are presented in
Figure 25, with the shift in Annexin V staining presented histographically. The results demonstrate
an increase in Annexin+/PI- apoptotic cells following the various treatments. Small levels of
apoptotic and necrotic cells were seen in the untreated control cells. A significant increase in early
apoptotic Annexin+/PI- cells was seen in all treatment groups (P < 0.001). CsA and MECA
treatment caused moderate increases in Annexin+/PI- cells, with MECA producing a greater
increase in apoptotic cells (10.50% and 20.34%, respectively). However, combination of the two
96
treatments caused a shift in early apoptotic cells that was significantly higher than either individual
treatment (44.44%; P < 0.001). Furthermore, the cytotoxic effects of the combination treatment
caused a dramatic reduction in viable Annexin-/PI- cells to 19.44%, compared to the individual
treatments (CsA 70.52%, MECA 57.16%) and the untreated control (19.44%).
97
Untreated CsA 5 µM P
rop
idiu
m I
od
ide
A B
Annexin V
MECA 500 µM CsA + MECA
Pro
pid
ium
Io
did
e
C D
Annexin V
E
Untr
eate
d M
CsA
5
M
MEC
A 5
00
CsA
+ M
EC
A
0
10
20
30
40
50
All Combinations = ***
Earl
y A
pop
toti
c C
ells
(%
)
Figure 25: Flow cytometric detection of apoptotic and necrotic cells by fluorescence-activated cell sorting of propidium
iodide and Annexin V labelled MDA-MB-231 cells. Cells were treated with CsA 5.0 μM (B), MECA 500 μM (C) or CsA
5.0 μM + MECA 500 μM (D) for 72 hours prior to staining and analysis. Untreated cells (A) were used as a negative
control. Dot-plots represent viable and apoptotic/necrotic cells. The bottom-left quadrant represents viable cells
(Annexin-/PI-). The bottom-right quadrant represents early apoptotic cells (Annexin+/PI-). Late apoptotic or secondary
necrotic cells are represented by the top-right quadrant (Annexin+/PI+). Early apoptotic (Annexin+/PI-) cells were
graphed (E) with the error bars representing the mean ± SEM (n = 3). A one-way ANOVA with a Bonferroni post-hoc
test was used to determine significance between all combinations (*** P < 0.001).
88.05%
7.39%
1.98%
2.58%
70.52%
14.76%
10.50%
4.22%
57.16%
15.06%
20.34%
7.44%
19.44%
30.92%
44.44%
5.20%
98
3.4.2 Chemotherapy Agents + nAChR Antagonists
To further investigate the combinational effect of nAChR antagonists with common
chemotherapy agents in susceptible cell lines, the methods of (Hallett et al., 2012) were adopted.
A549 and SKBR3 cells were first treated with doxorubicin (DOXO) and docetaxel (DTX), and in
combination with ADI, to observe the corresponding reduction in cell viability. DOXO 1.0 μM,
DTX 100 μM and the combination of the two chemotherapy agents in A549 cells caused a moderate
reduction in cell viability, ranging from ~25-40% reduction in viable cells (Figure 26). Combination
of the treatments with a relatively weak cytotoxic concentration of ADI 200 μM caused little
change in the viability of the cell lines, with only DOXO + ADI causing a significant reduction
from the chemotherapy treatment alone (Figure 26A; P < 0.05).
Similar results were observed following individual and combination treatment with SKBR3
cells (Figure 27). A similar decrease in cell viability was seen following DOXO, DTX and DOXO
+ DTX treatment, resulting in a ~25-60% reduction in viable cells. No significant decrease in cell
viability was shown when combined with ADI 200 μM, compared to chemotherapy treatment
alone, for all combinations tested, despite the consistent ~20% reduction in cell viability caused by
ADI treatment alone.
99
A
Untr
eate
d M
DO
XO
1.0
M
AD
I 20
0
DO
XO
+ A
DI
0
20
40
60
80
100
120
*N
o.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
B
Untr
eate
dM
DTX
100
D
MSO M
AD
I 20
0
DTX
+ A
DI
0
20
40
60
80
100
120
*
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
C
Untr
eate
d M
M +
DTX
100
DO
XO
0.1
M
AD
I 20
0
DO
XO
/DTX
+ A
DI
0
20
40
60
80
100
120
No.
of
Via
ble
Cel
ls C
om
pare
d t
o C
on
trol
Figure 26: The cytotoxic effect of DOXO, DTX and ADI treatment in A549 cells. A549 cells were treated with DOXO
1.0 μM (A), DTX 100 μM (B) and DOXO + DTX (C), alone and in combination with ADI 200 μM for 48 hours
following a 4 hour seeding period. Untreated cells were used as a negative control, while a DMSO vehicle control was
used for DTX treatment. Cell viability was established via the MTT assay. Values are displayed as the mean percent of
the control viability, with error bars representing SEM (n = 3). A one-way ANOVA with a Bonferroni post-hoc test was
used to determine significance between the treatment groups (* P < 0.05).
100
A
Untr
eate
d M
DO
XO
1.0
M
AD
I 20
0
DO
XO
+ A
DI
0
20
40
60
80
100
120N
o.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
B
Untr
eate
dM
DTX
100
D
MSO M
AD
I 20
0
DTX
+ A
DI
0
20
40
60
80
100
120
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
C
Untr
eate
d M
M +
DTX
100
DO
XO
0.1
M
AD
I 20
0
DO
XO
/DTX
+ A
DI
0
20
40
60
80
100
120
No.
of
Via
ble
Cel
ls C
om
pare
d t
o C
on
trol
Figure 27: The cytotoxic effect of DOXO, DTX and ADI treatment in SKBR3 cells. SKBR3 cells were treated with
DOXO 1.0 μM (A), DTX 100 μM (B) and DOXO + DTX (C), alone and in combination with ADI 200 μM for 48 hours
following a 4 hour seeding period. Untreated cells were used as a negative control, while a DMSO vehicle control was
used for DTX treatment. Cell viability was established via the MTT assay. Values are displayed as the mean percent of
the control viability, with error bars representing SEM (n = 3). A one-way ANOVA with a Bonferroni post-hoc test was
used to determine significance between treatment groups.
Further analysis of the combinational effects of various nAChR antagonists with the
combined cytotoxic treatment revealed similar results to those seen for ADI treatment. Relatively
mild cytotoxic concentrations of nAChR antagonists, including MECA 500 μM, PnTX 1 μM and
MLA 100 μM, were investigated in combination with DOXO 0.1 μM + DTX 100 μM treatment. No
significant decrease in cell viability was observed for A549 (Figure 28A) or SKBR3 (Figure 28B)
101
when treated with nAChR antagonists and DOXO/DTX, compared to DOXO/DTX treatment alone
(P >0.05).
A
Untr
eate
d M
M +
DTX
100
DO
XO
0.1
M
MEC
A 5
00
M
PnTX
1
M
MLA
100
DO
XO
/DTX
+ M
EC
A
DO
XO
/DTX
+ P
nTX
DO
XO
/DTX
+ M
LA
0
20
40
60
80
100
120
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
B
Untr
eate
d M
M +
DTX
100
DO
XO
0.1
M
MEC
A 5
00
M
PnTX
1
M
MLA
100
DO
XO
/DTX
+ M
EC
A
DO
XO
/DTX
+ P
nTX
DO
XO
/DTX
+ M
LA
0
20
40
60
80
100
120
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
Figure 28: Determination of cell viability following treatment with DOXO, DTX and various nAChR antagonists in
A549 (A) and SKBR3 (B) cell lines. Cells were treated with DOXO 0.1 μM + DTX 100 μM, MECA 500 μM, PnTX 1μM,
MLA 100 μM or in various combinations for 48 hours, following a 4 hour seeding period. Untreated cells were used as
a negative control. Cell viability was established via the MTT assay. Values are displayed as the mean percent of the
control viability, with error bars representing SEM (n = 3). A one-way ANOVA with a Bonferroni post-hoc was used to
determine significance between the treatment groups.
102
Finally, the combination of DOXO 0.1 μM with nAChR antagonists was evaluated to
investigate any significant synergistic effects during cytotoxic combination treatment of cancer
cells. The viability of MDA-MB-231, HT29 and PC3 cells was measured following treatment with
DOXO 0.1 μM in combination with ADI 250 μM, MECA 100 μM, MLA 100 μM and 5-HT 100
μM. A significant decrease in cell viability was observed following DOXO + ADI treatment in
MDA-MB-231 cells, compared to DOXO treatment alone (P < 0.01; Figure 29A). A similar
significant decrease in DOXO + ADI and DOXO + MECA was seen in HT29 cells, compared to
DOXO treatment alone (P < 0.001; Figure 29B). However, no significant decrease in cell viability
was observed for all other treatment combinations in MDA-MB-231 and HT29 cells, and for all
combination treatments in PC3 cell lines (P > 0.05; Figure 29C).
103
A
Untr
eate
d M
DO
XO
0.1
M
AD
I 25
0 M
MEC
A 1
00
M
MLA
100
M
5-H
T 1
00
DO
XO
+ A
DI
DO
XO
+ M
EC
A
DO
XO
+ M
LA
DO
XO
+ 5
-HT
0
20
40
60
80
100
120**
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
B
Untr
eate
d M
DO
XO
0.1
M
AD
I 25
0 M
MEC
A 1
00
M
MLA
100
M
5-H
T 1
00
DO
XO
+ A
DI
DO
XO
+ M
EC
A
DO
XO
+ M
LA
DO
XO
+ 5
-HT
0
20
40
60
80
100
120***
***
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
C
Untr
eate
d M
DO
XO
0.1
M
AD
I 25
0 M
MEC
A 1
00
M
MLA
100
M
5-H
T 1
00
DO
XO
+ A
DI
DO
XO
+ M
EC
A
DO
XO
+ M
LA
DO
XO
+ 5
-HT
0
20
40
60
80
100
120
No.
of
Via
ble
Cell
s C
om
pare
d t
o C
on
trol
Figure 29: Evaluation of cell viability following treatment with DOXO and various nAChR antagonists in MDA-MB-
231 (A), HT29 (B) and PC3 (C) cell lines. Cells were treated with DOXO 0.1 μM, ADI 250μM, MECA 500 μM, MLA
100 μM, 5-HT 100 μM or in various combinations for 48 hours, following a 4 hour seeding period. Untreated cells
were used as a negative control. Cell viability was established via the MTT assay. Values are displayed as the mean
percent of the control viability, with error bars representing SEM (n = 3). A one-way ANOVA with a Bonferroni post-
hoc was used to determine significance between the treatment groups.
104
3.4.3 Discussion
The final studies investigated the possible synergistic effects of the most cytotoxic nAChR
modulators in combination with various chemotherapy agents. It was hypothesised that the
combination of nAChR antagonists would enhance the cytotoxic nature of the chemotherapy agents,
at concentrations that were relatively non-cytotoxic when administered alone. MECA co-treatment
with CsA was first investigated due to the synergistic effects observed during the apoptotic
inhibition studies with PnTX and CsA in various carcinoma cell lines (see Section 3.3.1 Cell Death
Inhibitors). MECA was chosen for continuation of the combination studies due to the unavailability
of PnTX. Treatment with a combination of MECA and CsA produced significant increases in
carcinoma cell death, compared to the individual treatments. The relationship between the two
compounds was determined as synergistic, with a significant reduction in IC50 value for CsA when
in the presence of relatively non-cytotoxic concentrations of MECA. From the isobologram, it
seems CsA has the greater effect on the IC50 of MECA.
The isobologram results demonstrated a synergistic effect between MECA and CsA. The
swayed nature of the IC50 points suggests that the synergistic effects are more evident under
different combinations of the two compounds. The presence of CsA has a greater effect on the IC50
of MECA, potentiating the cytotoxic effects of MECA presumably through enhancing the
downstream effects of nAChR antagonism. CsA acts via inhibiting the cyclophilin D protein,
preventing the opening of the mitochondrial permeability transition pore during apoptosis.
However, the results from studies with both MECA and PnTX suggest a secondary action of CsA,
whereby CsA acts as a phosphatase 2B inhibitor, with the ability to affect the desensitisation status
of nAChRs. Marszalec and colleagues previously investigated the desensitisation of nAChR during
modulation of kinase activation and phosphatase inhibition (Marszalec et al., 2005). The study
showed that nAChRs are present as two interconverting states of desensitisation, a rapidly
recovering D1 state, and a slower recovering D2 state. nAChRs important in cancer development,
such as the α4β2 subtype, are often driven towards the slowly recovering D2 state following
105
continuous exposure to receptor modulators. The blocking action of CsA reduces the overall
desensitisation of nAChRs by facilitating a shift toward the faster recovering D1 receptor state
through the inhibition of nAChR dephosphorylation (Marszalec et al., 2005). The phosphatase 2B
inhibitory effects in the present combination study would effectively enhance the activity of the
potent nAChR antagonists, MECA and PnTX, resulting in the observed synergistic cytotoxic
effects. Consequently, CsA could potentially be used as a sensitising agent to enhance the effects of
nAChRs antagonists in cancer cell death.
Studies involving the co-treatment of carcinoma cell lines with nAChR modulators and
chemotherapy agents, doxorubicin and docetaxel, were conducted to further explore the possible
synergistic effects of the combination treatments. The present studies were based on those
previously performed by Hallet et al, where co-treatment of ADI with the chosen chemotherapy
agents produced a significant synergistic effect in a range of ‘sensitive’ carcinoma cell lines. The
study showed a significant synergistic effect was observed at concentrations of the two treatments
that would not elicit a cytotoxic response when administered individually. The present study used a
range of carcinoma cell lines, including the A549 and SKBR3 cell lines used in the Hallet et al
study, with similar concentrations of the treatments in an attempt to replicate and further the studies
previously reported. Contrasting results were observed during the present study, with co-treatment
of ADI and other nAChR modulators with the chemotherapy agents producing no clear synergistic
cytotoxic effects. Differences in the method used to determine cell viability between the present
studies (MTT assay), and the literature (AlamarBlue reduction method) may have contributed to the
conflicting results. The AlamarBlue viability assay is based on the reduction of the non-fluorescent
reagent resazurin, to the fluorescent compound resarufin, by viable cells. However, studies have
shown the reagent can interact and convert to the fluorescent compound through interactions with
various cell media types, independent of the presence of living cells. Incubation periods up to 4
hours produced ~15% increase in relative fluorescence when incubated in cell media without the
presence of cells (Munshi et al., 2014). This effect may have exacerbated the difference in treatment
106
groups in the results by Hallet et al during the 24 hour incubation of AlamarBlue to determine cell
viability; while little differences were observed in the present study.
The concept of combining selective nAChR antagonists with chemotherapeutic agents is one
that has been alluded to throughout the current literature. Studies, such as that conducted with the
selective α7 antagonist, APS8, have demonstrated an ability to sensitise cells to apoptosis. NIC
treatment of SKMES-1 cells has been shown to increase the expression of a family of pro-survival
proteins, the inhibitors of apoptosis proteins (IAPs). The endogenous proteins bind and inactivate
caspases, effectively inhibiting cancer cell apoptosis. APS8 treatment was shown to be an effective
antagonist IAP activity. The results suggest that APS8 treatment was capable of preventing NIC-
induced resistance to apoptosis, by interfering with IAP activity, allowing the downstream
activation of caspases and ultimately cell apoptosis to occur (Zovko et al., 2013). Theoretically,
such effects of α7-nAChR antagonism would sensitise cancer cells to conventional chemotherapy
agents that attempt to induce apoptotic pathways. Doxorubicin and docetaxel both act via cytotoxic
mechanisms that do not directly target cancer cell apoptotic pathways. The present studies showed
no significant sensitisation to both chemotherapeutic agents when combined with various nAChR
antagonists. Combination of such antagonists with chemotherapeutic compounds that target specific
apoptotic pathways may better exploit the sensitising effects of nAChR antagonists.
107
4 Discussion
4.1 Signalling through nAChRs is linked to Cancer Development
4.1.1 nAChR Agonist Induced Cancer Progression
In 2009, Schuller published a pivotal review highlighting the potential causal link between
smoking and nAChR driven tumour growth and metastasis. Nicotine and the highly potent nicotine-
derived nitrosamines are now recognised as potent agonists of nAChRs (Schuller, 2009). These
agonists promote carcinogenesis by stimulating angiogenesis and inhibiting apoptosis through
binding to the α7 subunit of the nAChR (Jull et al., 2001). Blocking these effects through the use of
selective antagonists offers new opportunities for the development of mechanisms that reduce
cancer development and progression.
4.1.2 nAChR Antagonist Driven Cytotoxicity
The recently identified family of shellfish toxins, PnTXs, demonstrate high antagonistic
affinity towards α7-nAChRs (Hellyer et al., 2011). In this research, PnTX-driven nAChR
antagonism demonstrated the most profound effect on cancer cell viability of all the antagonists
tested. The cytotoxic effects were more distinguishable in the adenocarcinoma cells lines,
highlighting expression of nAChRs outside of the nervous system.
Flow cytometry studies showed that treatment of the MDA-MB-231, HT29 and PC3
carcinoma cell lines with the nAChR antagonists MECA, ADI and 5-HT (through secondary
antagonistic effects) induces DNA fragmentation and opening of the mitochondrial transition pore,
indicative of early apoptosis. Interestingly, flow cytometry and apoptotic inhibition studies
suggested an alternative mechanism for PnTX induced cytotoxicity compared to the other nAChR
antagonists investigated. PnTX treatment induced a small, but significant increase in apoptotic cells
within the sub-G1 fraction of the cell cycle; while only displaying a slight increase in the
mitochondrial membrane permeability of treated cells. The flow cytometry profile of PnTX was
significantly different to that of the MECA, ADI and 5-HT treatment, despite displaying
108
comparable cytotoxic effects during viability studies. Although a proposed mechanism for PnTX-
induced cell death of carcinoma cells has not been previously documented in the literature, the
present study suggests a cytotoxic mechanism associated with the ability of the α7-nAChR to
regulate intracellular calcium levels.
4.2 The nAChR Antagonist PnTX Induces Cytotoxicity by Altering
Intracellular Calcium Levels
The present study suggests that the cytotoxic effects of PnTX are mediated through the
observed antagonism of the highly permeable α7-, and to a lesser extent the α4β2-nAChRs,
inducing cell apoptosis through calcium-independent pathways.
Cell cycle analysis showed a potential G0/G1 phase arrest following PnTX treatment.
Transient rises in [Ca2+] are integral to the process of cell division (Kahl et al., 2003), suggesting
that changes in [Ca2+] through antagonism of α7 and α4β2 may play a role during PnTX treatment.
To investigate the mechanism of PnTX cytotoxicity further, inhibitors of common apoptotic
pathways were used in an attempt to reduce the cytotoxic effects of PnTX. PnTX-induced cell
cytotoxicity was not affected by the inhibition of the various intrinsic apoptotic processes,
suggesting an alternative mechanism for PnTXs cytotoxic effects. Intracellular calcium studies
revealed that following PnTX treatment there is a prolonged decrease in [Ca2+], suggesting a
possible cytotoxic action by inducing cell apoptosis through Ca2+-independent pathways. Although
the limited supply of PnTX available at the time of this research meant the experiments could not be
repeated, the promising initial findings warrant further investigation.
109
4.3 Synergistic Cytotoxic Effects of Selective nAChR Antagonists with
Common Chemotherapeutic Agents
A second promising outcome of this research was the demonstration that selective nAChR
antagonists enhance the cytotoxicity of common chemotherapeutic agents. The effect of nAChR
antagonist treatment was assessed in combination with CsA and common chemotherapeutic agents
to explore the synergistic capacity of the compounds. The combination of nAChR antagonists was
expected to enhance the cytotoxic nature of the chemotherapeutic agents through sensitisation of the
cancer cells (Hallett et al., 2012). Contrary to expectations, combination of the nAChR antagonists
with doxorubicin and docetaxel displayed no synergistic effects, inconsistent with previous
literature. Alternatively, MECA treatment in combination with CsA produced a significant increase
in carcinoma cell death and early apoptotic cells, compared to the individual treatments. The
relationship between the two compounds was determined as synergistic based on the isobologram
results. The synergistic cytotoxic effects are attributed to the secondary action of CsA through
phosphatase 2B inhibition, effectively enhancing the activity of the potent nAChR antagonists. CsA
could potentially be used as a sensitising agent to enhance the effects of nAChRs antagonists in
cancer cell death. Although similar results were demonstrated when CsA was further investigated
with MECA, PnTX would have been the preferred investigational treatment.
4.4 Future Directions
The present study has established the relative cytotoxic nature of several nAChR modulators.
To further investigate the importance of nAChR in cancer development and whether nAChR
overexpression of nAChR can be used to selectively exploit cancer cells, studies on healthy
epithelial cells should be conducted. Previous studies, such as that conducted by Zovko and
colleagues on the apoptotic effects of APS8, have suggested that the antagonistic effects of selective
α7-nAChRs do not affect the proliferation of normal healthy cells. A possible theory for why
selective α7-nAChR antagonists, such as PnTX, may have a differential effect between normal and
110
cancer cells, could be due to the number of binding sites present on the different cell types. Current
literature has repeatedly demonstrated the upregulation of the α7-nAChR in various cancer cell
types; while the present study has hinted at a significant relationship between greater levels of α7-
nAChR expression and an increased response to nAChR modulators. Studies looking at the effect of
PnTX on the viability and apoptotic induction of healthy epithelial cells would assist in determining
the possibility of exploiting nAChR overexpression in cancer treatment.
Further investigation into the cytotoxic mechanism of PnTX on the various carcinoma cell
types would play an important role in deducing the effects of nAChR in cancer cell death.
Observations from the present study have determined that decreases in intracellular calcium
following PnTX treatment are the likely cause of the cytotoxic effects. Knockout studies of various
nAChR subtypes, including the highly permeable α7 subunit, would further consolidate the present
findings. However, the complete mechanism and second messengers involved downstream of the
changes in intracellular calcium have not been determined. As discussed previously, Ca2+ is
important for the expression of the cell cycle genes, FOS, JUN and MYC. Investigation into the
changes in FOS, JUN and MYC expression during PnTX treatment would assist in determining
whether the decrease in intracellular calcium is having a significant effect on the cancer cell cycle.
Additionally, studies using PnTX to replicate the combination effect seen between MECA and CsA
treatment should be conducted, to investigate whether similar synergistic effects are observed.
Combination treatment of PnTX and CsA conducted on nAChR knockout cells would supplement
the current theory that CsA has a significant effect on the sensitivity of nAChRs to PnTX treatment.
Although the findings of this research highlight the potential application of nAChR
modulators in controlling cancer cell proliferation, further development is essential before they
could be used in a clinical setting. Naturally occurring antagonists often display potent inhibitory
actions on neuromuscular nAChRs, making them highly toxic when administered in vivo (Zovko et
al., 2013). The antagonistic behaviour at the neuromuscular junction often occurs at concentrations
below those required to induce cytotoxic effects on cancer cells, limiting the potential clinical use of
111
such nAChR antagonists. PnTX falls into this category with potent effects on neuromuscular
nAChRs at a concentration of ~500 nM (Hellyer et al., 2011). The potential clinical use of nAChR
antagonists in cancer treatment would require alteration to the chemical structures that limit their
effects at the neuromuscular junction, while sustaining the potent effect on non-neuronal nAChRs.
Alternatively, a better understanding of how nAChR antagonists induce cancer cell cytotoxicity
may allow targeting of downstream effector molecules that eliminate the need for selective
targeting of nAChRs.
4.5 Final Conclusion
This research has identified a novel mechanism for the observed cytotoxicity of the highly
potent nAChR antagonist, PnTX, on human epithelial carcinoma cell lines. Cell cycle and
intracellular calcium studies suggest that PnTX exerts its cytotoxic effects by inducing changes in
the cell cycle via antagonism of the highly calcium permeable α7-, and to a certain extent, the α4β2-
nAChR receptors. This leads to an induction of cell apoptosis via calcium-independent pathways.
The use of nAChRs in combination with other known cytotoxic agents, such as CsA or apoptotic
inducing chemotherapy agents, may provide a sensitising tool that can reduce the dose required to
elicit a cytotoxic response.
112
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