nanotechnology-based strategies to enhance chemo- and ... · iv acknowledgments i am truly grateful...
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Nanotechnology-Based Strategies To Enhance Chemo- And
Radiation Therapy In Breast Cancer
by
Preethy Prasad
A thesis submitted in conformity with the requirements
for the degree of Doctor of Philosophy
Pharmaceutical Sciences University of Toronto
© Copyright by Preethy Prasad 2014
ii
Nanotechnology-Based Strategies to Enhance Chemo- and
Radiation Therapy in Breast Cancer
Preethy Prasad
Doctor of Philosophy
Graduate Department of Pharmaceutical Sciences
University of Toronto
2014
Abstract
A major cause of cancer treatment failure is multidrug resistance (MDR) and radioresistance to
standard therapies. Overexpression of ATP-binding cassette (ABC) transport proteins by cancer
cells, which actively transport anti-cancer agents (e.g. doxorubicin, Dox) out of the cells against
concentration gradients, is a major barrier to effective chemotherapy. Low levels of oxygen in tumors
are responsible for radioresistance contributing to the failure of radiation therapy (RT) of solid
tumors. This thesis concerns the development and evaluation of three nanoparticle delivery systems
for overcoming tumor resistance to chemo- and radiotherapy. System 1: polymer lipid hybrid
nanoparticles (PLN), co-loaded with a synergistic combination of anticancer agents Dox and
mitomycin C (MMC) (DMsPLN), were found to overcome multiple membrane efflux pumps
mediated MDR in vitro. Systemic administration of DMsPLN significantly enhanced therapeutic
efficacy in orthotopic tumor models of Dox-sensitive and resistant human breast cancer cells, with
low systemic toxicity compared to a clinically used liposomal formulation of Dox. System 2: cyclic
Arg-Gly-Asp (RGD), a ligand that binds with αvβ3 intergin receptors preferentially expressed in
angiogenic tumor blood vessels and certain cancer cells, was conjugated to DMsPLN. The Integrin-
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targeted RGD-DMsPLN resulted in a significant reduction in lung metastases of human breast cancer
cells without producing drug-associated systemic toxicity as observed in mice treated with free Dox-
MMC solutions. System 3: Manganese dioxide nanoparticles (MnO2 NPs) were developed and the
reactivity of MnO2 towards peroxides was utilized to regulate the tumor microenvironment in a
murine breast tumor. Intratumoral administration of MnO2 NPs simultaneously increased tumor
oxygenation by 45%, and tumor pH from pH 6.7 to pH 7.2 by reacting with endogenous H2O2
produced within the tumor. Combination treatment of the tumors with NPs and ionizing radiation
significantly inhibited breast tumor growth, increased DNA double strand breaks and cancer cell
death as compared to RT alone. The design and application of these three novel nanotechnology
platforms, in pharmaceutically acceptable NP formulations, provide promising therapeutic strategies
for enhanced chemo- and radiation therapy of breast cancer.
iv
Acknowledgments
I am truly grateful to all the people that have supported me throughout my Ph.D.
I would like to express my gratitude to my supervisor, Dr. X.Y.Wu, for offering me the
opportunity to train in her laboratory. The duration of my training in her laboratory has been one
of great intellectual creativity for me. Thank you Dr. Wu for your continuous guidance, and
support and also allowing me the freedom to explore my research interests. I also want to thank
you for your support and understanding of my participating in various extra-curricular activities
during the course of my graduate studies. Last but not least, thank you for all the opportunities
you have provided me to develop as a competent researcher and professional.
I would like to acknowledge my advisory committee members, Dr. Michael Rauth, Dr. Rob
Bristow and Dr. Peter O’Brien. Your valuable suggestions and feedback during my annual
meetings were most essential in shaping the direction of my research. Most notably, I would like
to extend my appreciation to Dr. Rauth, who always offered to meet with me to discuss the
project, reviewing all my manuscripts and being a great mentor.
Thank you to Dr. Ralph DaCosta and Azusa Maeda for contributing to this thesis. Dr. DaCosta
provided much guidance and graciously allowed me to use his facilities and equipment.
I am grateful for having wonderful lab mates throughout the years. Thank you to Dr. Claudia
Gordijo, Dr. Ping Cai, Michael Chu, Dr. Azhar Abbasi, Mary Shen, Jason Li, Jamie-Lugtu, Ji
Chen and Gary Chen whom I have had many helpful and enjoyable discussions and
conversations. I am grateful to Adam Shuhendler whose project I continued and I thank him for
being so helpful and patient while I learned the basics of nanoparticle drug delivery research.
Ping, thanks a lot for all your assistance with the in vivo work. You were so patient and always
available. I would also like to acknowledge the contributions of two of my summer students:
Wendy Xiong and Angela Ip. Both of you had been a great help.
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I would especially like to extend my thanks to Dr. Claudia Gordijo who has played an
instrumental role in my Ph.D. training. She has been a wonderful colleague and I thank her for
giving me the support, encouragement, enthusiasm and most of all her friendship over the past 5
years. I am definitely going to miss our morning coffee sessions.
I am grateful to the Canadian Breast Cancer Foundation, Natural Sciences and Engineering
Research Council of Canada, Canadian Institutes of Health Research, Ontario Graduate
Scholarship Program, University of Toronto and Leslie Dan Faculty of Pharmacy for
scholarships and research funding.
I owe the deepest gratitude to my parents, who have never stopped to encourage me to achieve
my full potential and pursue my goals. To my parents, Mummy and Papa, it is only because of
everything you have sacrificed and your unconditional love that I have been able to accomplish
this chapter of my life. I would also like to thank my sister, Prachy Mohan for her continued
support and guidance in many different things in life. Even though she is younger, I have learnt
many things from her.
Finally, I thank my best friend and husband, Nikhil, for his unbounded love. You have been a
pillar of strength, comforting me in my worries and supporting my aspirations. Without you,
completing the Ph.D. would not have been possible.
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Dedicated to my parents, Poonam and Bhuwan Prasad and my husband,
Nikhil Khunteta
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Table of Contents
Acknowledgments.......................................................................................................................... iv
List of Tables ................................................................................................................................ xv
List of Figures .............................................................................................................................. xvi
List of Abbreviations ................................................................................................................... xix
Chapter 1 Introduction .................................................................................................................... 1
1. Breast Cancer ....................................................................................................................... 1
1.1 Incidence and etiology ..................................................................................................... 1
1.2 Classification .................................................................................................................... 2
1.3 Development of breast cancer .......................................................................................... 3
1.4 Tumor microenvironment ................................................................................................ 5
1.4.1 Cells of the tumor microenvironment ........................................................................... 5
1.4.2 Hypoxia in solid tumours ............................................................................................. 7
1.4.3 Angiogenesis ................................................................................................................ 8
1.4.4 Integrins ...................................................................................................................... 10
1.4.5 Metastasis ................................................................................................................... 12
1.5 Breast cancer therapy ..................................................................................................... 14
1.5.1 Local therapy .............................................................................................................. 15
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1.5.2 Systemic therapy......................................................................................................... 16
2. Barriers to cancer therapy .................................................................................................. 21
2.1 Therapeutic resistance .................................................................................................... 21
2.1.1 Cellular and molecular causes of drug resistance ....................................................... 22
2.1.2 Mechanisms of resistance that relate to tumor microenvironment ............................. 25
2.2 Treatment side effects .................................................................................................... 27
3. Nanoparticulate systems in cancer therapy ........................................................................ 29
3.1 Optimal characteristics of nanoparticle delivery system................................................ 30
3.2 Targeting tumor with nanoparticles ............................................................................... 32
3.2.1 Passive targeting ..................................................................................................... 32
3.2.2 Active targeting ....................................................................................................... 33
3.2.3 Solid lipid and polymer-lipid hybrid nanoparticles ................................................ 35
4. Goal for this work .............................................................................................................. 36
5. Organization of thesis ........................................................................................................ 37
Chapter 2 A novel nanoparticle formulation overcomes multiple types of membrane efflux
pumps in human breast cancer cells.............................................................................................. 40
1 Abstract .................................................................................................................................. 41
2 Introduction ............................................................................................................................ 42
3 Materials and methods ........................................................................................................... 46
ix
3.1 Chemicals and reagents .................................................................................................. 46
3.2 Formulation and characterization of PLN ...................................................................... 47
3.3 Measurement of drug loading and encapsulation efficiency of PLN ............................. 48
3.4 Cell maintenance ............................................................................................................ 48
3.5 Clonogenic assay ............................................................................................................ 48
3.6 Median Effect Analysis .................................................................................................. 49
3.7 Fluorescence microscopy of cellular PLN uptake.......................................................... 50
3.8 Statistical analysis .......................................................................................................... 51
4 Results .................................................................................................................................... 52
4.1 Properties of PLN ........................................................................................................... 52
4.2 Dose-response of MCF human breast cancer cells treated with Dox and MMC ........... 53
4.3 Synergistic effect of Dox and MMC in MCF7 human breast cancer cells .................... 57
4.4 PLN formulations are more effective than free drugs against MCF7 cancer cells ........ 59
4.5 Cellular uptake and intracellular localization of PLN .................................................... 62
5 Discussion .............................................................................................................................. 63
6 Conclusion .............................................................................................................................. 65
7 Acknowledgements ................................................................................................................ 66
Chapter 3 Doxorubicin and mitomycin C co-loaded polymer-lipid hybrid nanoparticles inhibit
growth of sensitive and multidrug resistant human mammary tumor xenografts ........................ 67
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1 Abstract .................................................................................................................................. 68
2 Introduction ............................................................................................................................ 69
3 Materials and methods ........................................................................................................... 72
3.1 Chemicals and reagents .................................................................................................. 72
3.2 Preparation and characterization of stealth polymer lipid hybrid Nanoparticles ........... 73
3.3 Cell culture ..................................................................................................................... 74
3.4 Orthotopic Model Development and Treatments ........................................................... 74
3.5 Evaluation of therapeutic efficacy.................................................................................. 75
3.6 Determination of median survival time and percentage increase in life span ................ 76
3.7 Evaluation of safety and normal tissue toxicity ............................................................. 77
3.8 CD31 expression and assessment of microvessel density of tumors (MVD) ................ 77
3.9 Statistical analysis and graphing .................................................................................... 78
4 Results .................................................................................................................................... 78
4.1 Determination of PLD dose for the treatment ................................................................ 78
4.2 Anti-tumor efficacy of DMsPLN in sensitive and MDR tumor models ........................ 79
4.3 Survival of tumor bearing mice following treatment ..................................................... 84
4.4 Systemic toxicity of DMsPLN ....................................................................................... 85
4.5 In vivo anti-tumor mechanism of DMsPLN .................................................................. 88
5 Discussion .............................................................................................................................. 90
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6 Acknowledgements ................................................................................................................ 95
Chapter 4 Integrin-targeted polymer-lipid nanoparticles encapsulating doxorubicin and
mitomycin C enhance treatment of lung metastases of human triple negative breast cancer in a
SCID mouse model ....................................................................................................................... 96
1 Abstract .................................................................................................................................. 97
2 Introduction ............................................................................................................................ 98
3 Materials and methods ......................................................................................................... 100
3.1 Materials ....................................................................................................................... 100
3.2 Synthesis and characterization of myrj56-cRGDfK targeting constructs .................... 101
3.3 Preparation and characterization of polymer lipid nanoparticles ................................. 102
3.4 Cell culture ................................................................................................................... 103
3.5 Metastasis model development .................................................................................... 104
3.6 Biodistribution study .................................................................................................... 104
3.7 In vivo treatments ......................................................................................................... 105
3.8 Evaluation of liver and cardiotoxicity .......................................................................... 106
3.9 Statistical analysis ........................................................................................................ 106
4 Results .................................................................................................................................. 106
4.1 In vivo biodisitribution of nanoparticles in tumor bearing mice .................................. 106
4.2 Maximum tolerable dose assessment ........................................................................... 109
4.3 Inhibition of tumor metastasis in vivo.......................................................................... 110
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4.4 Evaluation of liver toxicity ........................................................................................... 114
4.5 Evaluation of caridiotoxicity ........................................................................................ 116
5 Discussion ............................................................................................................................ 118
6 Conclusion ............................................................................................................................ 122
7 Acknowledgements .............................................................................................................. 122
Chapter 5 Multifunctional albumin based MnO2 nanoparticles modulate solid tumor
microenvironment by attenuating hypoxia, acidosis, VEGF and enhance radiation response ... 123
1 Abstract ................................................................................................................................ 124
2 Introduction .......................................................................................................................... 125
3 Methods ................................................................................................................................ 127
3.1 Nanoparticle synthesis.................................................................................................. 127
3.2 Cell lines, tumor models and treatments ...................................................................... 128
3.3 Quenching of H2O2 by nanoparticles ........................................................................... 128
3.4 In vitro oxygen and pH measurements ......................................................................... 129
3.5 Cellular uptake of NPs ................................................................................................. 129
3.6 Tumor retention of NPs ................................................................................................ 130
3.7 Tumor pH measurements ............................................................................................. 130
3.8 Tumor oxygenation measurements .............................................................................. 130
3.9 Immunohistochemistry detection of tumor hypoxia .................................................... 131
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3.10 In vivo radiation treatment............................................................................................ 131
3.11 Statistical analysis ........................................................................................................ 133
4 Results and discussion .......................................................................................................... 133
4.1 Preparation of A-MnO2 NPs ........................................................................................ 133
4.2 Multifunctionality of A-MnO2 NPs in culture medium ............................................... 135
4.3 Uptake of A-MnO2 NPs by breast cancer cells ............................................................ 137
4.4 Oxygen generation in the presence of hypoxic cancer cells ........................................ 139
4.5 Effect of A-MnO2 NPs on tumor oxygenation ............................................................ 140
4.6 Effect of A-MnO2 NPs on tumor pH ............................................................................ 142
4.7 Prolonged regulation of tumor hypoxia, HIF-1α and VEGF is related to extended tumor
retention of A-MnO2 NPs ........................................................................................................ 143
4.8 A-MnO2 NPs enhanced anti-tumor effect of radiation................................................. 146
5 Conclusions .......................................................................................................................... 149
6 Acknowledgements .............................................................................................................. 149
7 Supporting information ........................................................................................................ 150
7.1 pKa Calibration forSNARF in tissue like phantoms. ................................................... 150
7.2 Consumption of A-MnO2 Nanoparticles by H2O2: ...................................................... 154
7.3 Nanoparticle structure and MnO2 quantification: ........................................................ 155
Chapter 6 Overall conclusions, major contributions and Future Perspectives ........................... 157
xiv
1 Overall Conclusions ............................................................................................................. 157
2 Major contributions of this thesis ......................................................................................... 161
3 Future Directions .................................................................................................................. 162
3.1 Delivery of Dox-MMC using nanoparticles in other metastatic models of breast cancer
162
3.2 Determine the distribution of Dox and MMC in vivo at both macroscopic and
microscopic level ..................................................................................................................... 163
3.3 Optimizing the time of MnO2 NP administration prior to irradiation .......................... 165
3.4 Application of MnO2 nanoparticles for enhancement of Chemotherapy Therapy ....... 166
Appendix ..................................................................................................................................... 168
Appendix 1: To elucidate the mechanism of cellular uptake and intracellular transport of fatty
acid-based nanoparticles in cells ............................................................................................. 168
References ................................................................................................................................... 176
xv
List of Tables
Table 2.1: Composition of various PLN formulations
Table 2.2: Particle size, zeta potential, polydispersity index, drug loading efficiency and loading
levels of the drug-loaded PLN formulations
Table 2.3: Dose-effect relationship parameters for Dox and MMC in MCF7 WT, MCF7 MX
(BCRP+), and MCF7 VP (MRP1+) human breast cancer cell lines
Table 2.4: Cytotoxicity of Dox solution and dual drug (Dox and MMC) loaded PLN (DM-PLN)
against wild type (MCF7), BCRP+ (MCF7 MX) and MRP1+ (MCF7 VP) human breast cancer
cells.
Table 3.1: Effect of DMsPLN and PLD treatment on the tumor growth delay (TGD), median
survival time (MST) and increase in life span (ILS%) of tumor bearing mice.
Table 3.2: Immunohistochemical evaluation of the vascularisation of orthotopically implanted
MDA-MB 435/LCC6 breast cancer cells after treatment
Table 4.1: Determining MTD by measuring the number of mice showing sever signs of acute
toxicity, 7 days following treatment.
Table 4.2: Metastatic burden on mice as measured by total flux (p/s), 28 days following
treatment.
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List of Figures
Figure 2.1: A typical particle size and size distribution plot of PLN
Figure 2.2: Surviving fraction, measured by a clonogenic assay, of (A) MCF7 WT, (B) MCF7
VP (MRP1+) and (C) MCF7 MX (BCRP+) cells
Figure 2.3: Percent kill of cell ability to expand clonogenically vs. drug dose exposure for 1 hr
to Dox or MMC alone or in combination
Figure 2.4: Combination Index analysis of the interaction of Dox and MMC in (A) MCF7 WT,
(B) MCF7 VP and (C) MCF7 MX cells following treatment for 1 hour.
Figure 2.5 Comparison of anti-cancer efficacy of single agent Dox or MMC free in solutions or
in PLN with dual agent PLN formulation in (A) MCF7 WT, (B) MCF7 VP (MRP1+) and (C)
MCF7 MX (BCRP+) cells
Figure 2.6: Intracellular localization of fluorescent PLN in breast cancer cell lines MCF7 WT,
MCF7VP and MCF7MX.
Figure 3.1: Percent change in body weight of MDA-MB 435/LCC6/WT tumor bearing mice
treated with (A) saline, (B) 50 mg/m2 PLD and (C) 50 mg/m
2 DMsPLN
Figure 3.2: Individual tumor growth curves over 60 days for mice for each treatment group
Figure 3.3: Average tumor volume of each treatment group vs. time for mice bearing (A) MDA-
MB 435/LCC6/WT and (B) MDA-MB 435/LCC6/MDR1 tumors.
Figure 3.4: Normalized average tumor volume at specific time points (days) for mice treated
with saline, 25 mg/m2
PLD, 25 mg/m2
DMsPLN or 4 × 25 mg/m2
DMsPLN in (A) MDA-MB
435/LCC6/WT and (B) MDA-MB 435/LCC6/MDR1 tumor models.
xvii
Figure 3.5: Kaplan-Meier survival curves for (A) MDA-MB 435/LCC6/WT and (B) MDA-MB
435/LCC6/MDR1 tumor bearing mice treated with saline (green), 25 mg/m2
PLD (red), 25
mg/m2
DMsPLN (blue) and 4 × 25 mg/m2
DMsPLN (brown).
Figure 3.6: Percent change in body weight as a function of time in mice treated with different
formulations
Figure 3.7: Blood enzymes were used to assess toxicity. Serial blood collection and analysis of
plasma enzyme levels
Figure 3.8: Antiangiogenic effect following treatment. Hematoxylin-eosin staining and
immunohistochemical staining with CD31 in tumor sections
Fig. 4.1: Nanoparticle distribution in mice bearing MDA MB 231-luc-D3H2LN metastatic
breast cancer.
Fig. 4.2: Determining tumor metastasis burden over time
Fig. 4.3: High dose of PLN significantly inhibits tumor burden
Fig. 4.4: Free drug shows hepatotoxicity not seen in DMsPLN treatment groups
Fig. 4.5: Free drug shows cardiotoxicity not seen in DMsPLN treatment groups
Figure 5.1: Characterization of A-MnO2 NPs
Figure 5.2: In vitro reactivity of A-MnO2 NPs towards hydrogen peroxide
Figure 5.3: Cellular uptake, cellular oxygen generation and cytotoxicity of A-MnO2 NPs
Figure 5.4: Effect of A-MnO2 NPs on tumor oxygenation
Figure 5.5: Effect of A-MnO2 NPs on tumor pH
Figure 5.6: Tumor retention of A-MnO2 NPs and effect on tumor hypoxia, HIF-1α and VEGF
xviii
Figure 5.7: Effect on tumor growth after treatment with radiation and A-MnO2 NPs.
Figure S2: Calculation of pKavalue for SNARF in biological phantoms.
Figure S3: In the graph we show the consumption of the MnO2 NPs (90 µM) by various
endogenous concentrations of H2O2 (up to 1 mM).
Figure S4: TEM image of nanoparticle
Figure 6.1: Transmission electron micrograph of cellular uptake and intracellular localization of
SLN in MDA 435/LCC6/MDR cells with 1h incubation at 37ºC.
Figure 6.2: TEM imaging of MDA 435/LCC6/MDR cells with 1h incubation with SLN at 37ºC
depicting nuclear localization.
Figure 6.3: Confocal imaging of cellular uptake and intracellular distribution of SLN various
cell lines at 37ºC.
Figure 6.4: Confocal imaging of cellular uptake and intracellular distribution of SLN various
cell lines at 4ºC.
Figure 6.5: Cellular association of SLN analyzed using fluorescent spectrometer in various cell
lines.
Figure 6.6: SLN uptake analyzed using fluorescent spectrometer in the presence of different
inhibitors of specific endocytic process.
Figure 6.7: Intracellular distribution of SLN and lipid droplets in MDA-MB 231 cells.
xix
List of Abbreviations
αMEM: Alpha minimal essential medium
(αvβ3): alpha v beta 3
ABC: ATP-binding cassette
ALA: δ-aminolevulinic acid
ANOVA: Analysis of variance
Bcl-2: B-cell lymphoma 2
BCS: Breast conserving surgery
BCRP: Breast cancer resistance protein
BSA: Bovine serum albumin
CAM: Cell–cell adhesion molecules
CAIX: Carbonic anhydrase IX
CDCl3: deuterated chloroform
c-Myc: Myelocytomatosis viral oncogene
cRGDfK: cyclo-Arginine-Gycine-Aspartate-D Phenylalanine-Lysine
CT: Chemotherapy
cTnT: Cardiac troponin T
Dox: Doxorubicin
xx
DMsPLN: Dox- Mitomycin C polymer lipid nanoparticle
DSB: Double strand break
DSWC: Dorsal window chamber
ECM: Extracellular matrix
EMT: Epithelial-to-mesenchymal transition
EPR: Enhanced permeability and retention
FABP: Fatty acid binding protein
FDA: Food and drug administration
FBS: Fetal bovine serum
GLUT-1: Glucose transporter 1
GST: Glutathione-S-transferase
H2O2: Hydrogen peroxide
HER2: Human epidermal growth factor receptor -2
HIF1: Hypoxia-inducible factor–1
H&E: Haematoxylin and eosin
HPESO: Hydrolyzed polymers of epoxidized soybean oil
ICG: Indocyanine green
ICP: Inductively coupled plasma
IFP: Interstitial fluid pressure
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i.t.: Intratumoral
i.v.: Intravenous
mAbs: Monoclonal antibodies
MDR: multidrug resistance
MDSC: Myeloid-derived suppressor cells
MMC: Mitomycin C
MMP: Matrix metaloproteases
MnO2: Manganese dioxide
MPS: Mononuclear phagocytic system
MRP-1: Multidrug-resistance associated protein
MSFI: Multispectral fluorescence imaging
NF-κB: Nuclear factor-κB
NIR: Near infrared
NPs: Nanoparticles
NSABP: National Surgical Adjuvant Breast and Bowel Project
PA: Photoacoustic
PAH: Poly(allylamine hydrochloride)
PDGFB: Platelet-derived growth factor B
PDT: Photodynamic Therapy
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PEG: Polyethyleneglycol
PLD: PEGylated liposomal doxorubicin
PLN: Polymer lipid nanoparticle
P-gp: P-glycoprotein
p-NPC: p-nitrophenylchloroformate
PPE: Palmar plantar erythrodysenthesia
PS: Photosensitizer
ROI: Region of interest
ROS: Reactive oxygen species
RT: Radiation therapy
SEM: Standard error of mean
SLN: Solid lipid nanoparticles
sO2: Vascular saturated oxygen
TAM: Tumor-associated macrophages
TCA: Tricarboxylic acid
TEM: Transmission electron microscopy
TGF-beta: Tumor growth factor beta (TGF-beta)
TME: Tumor microenvironment
TNBC: Triple negative breast cancer
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TUNEL: TerminaldeoxynucleotidyltransferasedUTP nick end labelling
VEGF: Vascular endothelial growth factor
WT: Wildtype
1
Chapter 1
Introduction
1. Breast Cancer
1.1 Incidence and etiology
Cancer is a life threatening illness and is a major cause of death around the world. In 2012, the
World Health Organization reported a total of 14.1 million new cancer cases and 8.2 million
cancer deaths worldwide [1]. Based on 2013 incidence and mortality rate, the Canadian Cancer
Society estimates that about 2 in 5 Canadians will develop cancer in their lifetime and 1 in 4 will
die of the disease [2].
Among women, breast cancer is one of the most common manifestations of the disease, falling
second only to lung cancer in overall mortality with an estimated 1.67 million new cases
diagnosed worldwide in 2012 [1]. Despite advancement in both early diagnosis and treatment, it
still remains a major health concern for Canadian women with about 23,800 new cases of breast
cancer and 5,000 deaths from it in 2013[2]. It is estimated that approximately 65 Canadian
women will be diagnosed with breast cancer every day. Due to the high prevalence and mortality
rate of breast cancer, it represents a significant health care burden requiring extensive research in
developing diagnostic and treatment options for the disease,
The exact cause of breast cancer is not completely understood. However, the development of the
disease has been attributed to multiple risk factors, including increasing age, family history, diet,
exposure to female reproductive hormones, and environmental factors [3]. The risk of
2
developing breast cancer is strongly related to age, with the highest incidence rates in women
over 50 (150 cases/100,000) compared to <5 cases/100,000 population in females before age 30
[4]. Family history does not play a large role, compared to other factors such as age in
development of breast cancer, even though women have an increased risk of developing breast
cancer if they have a first degree relative with a history of breast cancer [5]. The majority of
women diagnosed do not have a family member with the disease. Hormonal factors that increase
exposure to estrogen, including early age menstrual cycle, late menopause and late age
pregnancy are also risk factors for developing breast cancer [5]. Several environmental factors
such as exposure to radiation are also involved in development of breast cancer.
1.2 Classification
Breast cancer can be broadly categorized by the extent of tumor growth into in situ carcinoma
and invasive (infiltrating) carcinoma [6]. Breast carcinoma in situ is the early stage of breast
cancer where the cancer is confined to the breast and has not infiltrated the surrounding tissues.
Breast carcinoma in situ is further sub-classified as either ductal or lobular [7]. Ductal carcinoma
is the most common type of breast cancer, found in the cells of the ducts. Lobular carcinomas are
in situ results in the presence of abnormal cells in the milk-producing glands of the breasts and
are usually non-invasive. Invasive breast cancer invades to other tissues of the breast or to other
parts of the body. Invasive carcinomas are also a heterogeneous group of tumors differentiated
into histological subtypes: infiltrating ductal, invasive lobular, ductal/lobular, mucinous
(colloid), tubular, medullary and papillary carcinomas. Of these, invasive ductal carcinoma is the
most common and accounts for more than 75% of breast cancer cases [8]. Molecular markers are
3
also used to further classify breast cancer as histologically similar tumors are phenotypically
different. These subtypes based on gene expression profile segregate breast cancer into four
types (i) luminal, (ii) basal, (iii) human epidermal growth factor receptor -2 (HER2) positive and
(iv) normal type [9, 10]. The current clinical classification system relies heavily on the
histological aspects of breast cancer; however advances in technology are allowing further tumor
characterization to improve treatment choice and prognosis.
1.3 Development of breast cancer
Initiation of a breast tumour is a direct result of aberrant genetic events within a single cell which
occur spontaneously and randomly during cell division resulting in uncontrolled cell
proliferation [11, 12]. Continual replication results in the formation of a colony of abnormal cells
which is comprised of multicellular components with intricate interactions with each other and
the surrounding tissue. Hananhan and Weinberg identified six essential alterations to cell
function and described these various defects in the regulatory circuits which drive the
progressive alteration of normal cells into cells with a malignant phenotype [13, 14].
One alteration is the ability of cancer cells to proliferate continuously, independent of their
microenvironment [13, 14]. In normal cells mitogenic growth signals are transmitted into the cell
by transmembrane receptors that bind signalling molecules such as diffusible growth factors,
extracellular matrix components, and cell-to-cell to adhesion/ interaction molecules to allow cell
proliferation. However, tumor cells generate their own growth signals, thereby reducing their
dependence on stimulation from their normal tissue microenvironment [13, 14]. The second
essential alteration of cancer cells is their ability to acquire resistance to anti- proliferative
4
signals, often through loss of function mutations to anti-mitogenic signal transduction effectors
[15-17]. Thirdly, cancer cells are able to alter the signaling system that activates apoptosis,
programmed cell death designed to remove unwanted or abnormal cells [13, 14]. Resistance to
apoptosis can be acquired through loss of function mutations of pro-apoptotic factors such as p53
[18, 19] or through upregulation of anti-apoptotic genes such B-cell lymphoma 2 (Bcl-2) or
myelocytomatosis viral oncogene (c-Myc) [20-22]. The fourth alteration to cell behaviour is the
ability of cancer cells to replicate without limits [13, 14]. Normal cells are programmed to
reproduce only a finite number of times mediated by telomere shortening through successive
cycles of replication [23, 24]. This loss of terminal telomeres leads to end to-end chromosomal
fusion and karyotypic disarray eventually resulting in massive cell death [23, 24]. However,
malignant cells including breast cancer cells up regulate telomerase, an enzyme which maintains
the length of telomeres, allowing unlimited replication of cancer cells [25, 26]. The fifth essential
alteration for cancer cells is the ability to induce angiogenesis; growth of new blood vessels [13,
14]. Increased blood flow to the tumor site is necessary to deliver nutrients and oxygen for rapid
tumor growth. This increase in blood supply is achieved by activating the angiogenic switch
through increased expression of vascular endothelial growth factor (VEGF) [27]. Finally, the
sixth alteration of cellular physiology essential to the malignant phenotype is the ability of cancer
cells to invade surrounding stroma and metastasize to distant sites [13, 14]. Metastasis is the
result of a multistep process requiring local tissue invasion, intravasation, embolization and
transit, extravasation, and finally colonization of cancer cells to a distant tissue. Several classes
of proteins are involved in this process which includes cell–cell adhesion molecules (CAMs),
immunoglobulin and calcium-dependent cadherin families which mediate cell-to-cell interactions
5
and integrins which attach cells to extracellular matrix substrates [28-30]. Metastasis of cancer,
including breast cancer is a major clinical challenge and is currently considered a major
challenge to tumor curability.
1.4 Tumor microenvironment
1.4.1 Cells of the tumor microenvironment
The microenvironment of developing tumor tissue does not just consist of a homogenous group
of cancer cells but is a complex tissue consisting of proliferating tumor cells, the tumor stroma,
blood vessels, infiltrating inflammatory cells and a variety of associated tissue cells [31, 32].
Dynamic interactions between all these components support tumor growth and invasion, protect
the tumor from host immunity, promote therapeutic resistance, and provide niches for dormant
metastases to thrive [31, 32]. Although tumors are monoclonal in origin, they eventually become
heterogeneous due to random mutation in certain cells and selective pressure of the TME that
cause genotypic and phenotypic divergence as the tumor grows [33]. Research has shown that
cancer stem cells which share many characteristics with normal stem cells, including self-
renewal and differentiation, play a major role in tumor heterogeneity [34, 35]. The ability of
cancer stem cells to seed new tumor growth and their resistance to chemotherapy makes it
difficult to eradicate tumors [34, 35].
Apart from the malignant cells, the TME contains endothelial cells, immune inflammatory cells
and cancer associated fibroblasts [31, 36]. Endothelial cells are actively recruited by fibroblasts
in the tumor stroma to form tumor neovasculature [37, 38]. These new blood vessels supply the
tumor with oxygen and nutrients essential for tumors to grow beyond 1-2mm and also allow
6
tumor cells to enter the circulation, enabling the metastasis of cancer cells to multiple organs [24,
38]. Pericytes, also known as perivascular stromal cells, are an integral component of the tumor
vasculature that provides structural support to blood vessels [39, 40]. Paracrine signaling via
platelet-derived growth factor B (PDGFB), expressed by endothelial cells, and its receptor
PDGFR-β, expressed by pericytes, plays a central role in blood vessel maturation, essential for
tumor growth [41]. Tumor-infiltrating immune cells including myeloid-derived suppressor cells
(MDSC), tumor-associated macrophages (TAM), and cytotoxic lymphocytes are critical
determinants of tumor growth. Many studies have shown that increased densities of MDSC and
TAM promote tumor progression via multiple immunosuppressive mechanisms [42, 43]. TAMS
have also been shown to be major contributors to tumor angiogenesis [44, 45]. In contrast, the
presence of cytotoxic B and T lymphocytes detect and eliminate cancer cells and are also
associated with good prognosis in numerous cancers [46, 47]. Other relevant components of the
TME are cancer associated fibroblasts (CAF) and myofibroblasts, which constitute the most
abundant mesenchymal cells found within most carcinomas including breast cancer [48, 49].
They secrete various growth factors and cytokines such as tumor growth factor beta (TGF-beta)
and hepatocyte growth factor promoting the growth and survival of malignant cells [50, 51].
CAFs also play a major role in cell invasion by promoting epithelial-to-mesenchymal transition
of tumor cells, secretion of pro-invasive factors and production of matrix metaloproteases
(MMPs) which play a major role in cancer metastasis [52]. CAFs also synthesize extracellular
matrix components such as collagen, fibronectin and laminin to expediate the process of tumor
growth and invasion [53]. Furthermore, CAFs also enhance angiogenesis by secreting factors
that activate endothelial cells and pericytes [52, 54]
7
Tumor development is a complex multistep process in which a plethora of tumor and stromal
cells play important roles. The cross-talk between different components of the tumor influences
the growth, survival, invasiveness, and metastatic ability of neoplastic epithelial cells. The
components and molecules implicated in this cross-talk are attractive targets in anticancer
therapeutic intervention.
1.4.2 Hypoxia in solid tumours
Hypoxia occurs as a consequence of a disrupted balance between supply and consumption of
oxygen, owing to large tumor size and vascular abnormalities [55]. As a result, the tumor
contains interspersed regions of well oxygenated (pO2 >2.5 mmHg) and poorly oxygenated (pO2
≤ 2.5 mmHg) areas, heterogeneously distributed throughout the tumor mass [56]. One third of
breast tumors have hypoxic regions with O2 concentrations less than 0.3%, compared to normal
tissue concentrations of approximately 9% [57]. Tumor hypoxia occurs as the result of two
independent, but non-exclusive mechanisms. The first mechanism is due to the abnormal tumor
vasculature which causes functional deficits in tumor perfusion, resulting in intermittent blood
flow to cancer cell leading to acute hypoxia or perfusion limited hypoxia [56]. This intermittent
blood flow can be due to temporary occlusions of blood vessels, which expose tumour cells to
repeated cycles of hypoxia and reoxygenation (termed “cycling hypoxia”) [58, 59]. The second
mechanism is due to the irregular distribution of blood vessels within tumor which results in
chronic or diffusion limited hypoxia. Rapid tumor proliferation outgrows the vasculature,
8
resulting in the inhibition of growth of cancer cells at distances greater than the 200 µm from the
blood vessel, due to the consumption of oxygen by the intervening cells [60, 61].
Hypoxic cancer cells develop an efficient adaptive metabolic response to ensure their survival
and proliferation in low oxygen regions that would induce normal cell death [62, 63]. Hypoxic
cells can undergo a shift from aerobic oxidative phosphorylation to anaerobic glycolysis for the
production of energy, also known as the Warburg effect [62, 63]. Glycolysis produces pyruvate,
which is then converted into lactate instead of being oxidized via the tricarboxylic acid (TCA)
cycle and oxidative phosphorylation [64, 65]. Since glycolysis is considerably less efficient than
oxidative phosphorylation in producing energy, the hypoxic cell increases the rate of glucose
uptake and glycolysis to meet its energy demands [64, 65]. This metabolic shift is driven by the
hypoxia-inducible factor–1 (HIF1) [64, 66]. HIF-1 leads to a cascade of expression of numerous
genes, proteins , and enzymes including those involved in glycolysis (glucose transporter 1
[GLUT-1]), angiogenesis (VEGF), and low pH (carbonic anhydrase IX [CAIX]) to facilitate
malignant tumor growth and survival [63, 67].
1.4.3 Angiogenesis
One of the hallmarks of cancer is the acquired capability of tumors to induce angiogenesis, the
process of new vessel formation from the endothelium of existing vasculature. For the growth of
tumor beyond 2 mm3 and for the metastatic spread of cancer tissue, growth of the vascular
network is important [68]. Formation of new vascular network supplies the tumor with nutrients,
oxygen and immune cells and also removes waste products, to support the continuous growth of
the tumor [69]. Formation of new vessels is a highly regulated process so that angiogenesis is in
9
“on” or “off” state depending on the conditions governed by the net balance between pro and
anti-angiogenic molecules [70, 71]. However during tumor growth and progression, numerous
factors cause disruption in the balance between pro- and anti- angiogenic molecules triggering
the angiogenic switch from the “off” state to “on” state. These factors include production of
growth factors by tumor cells, changes in the tumor microenvironment, the recruitment of
progenitor endothelial cells from bone marrow, and the down-regulation of natural inhibitors of
angiogenesis. Microenvironmental alterations such as hypoxia (low oxygen levels), low glucose
levels, low pH, and inducers of ROS stimulate angiogenic signals.
Upon angiogenic stimulus such as hypoxia, VEGF is produced and secreted into the surrounding
tissue. VEGF binds to its receptor in endothelial cells, inducing various signaling pathways
which promote migration, survival and proliferation of the endothelial cells [37, 68, 72, 73] .
Endothelial cells activated by VEGF produce matrix metaloproteinases, which degrade vascular
basement membrane, allowing migration of activated endothelial cells into the interstitium [74].
The endothelial cells divide as they migrate into the surrounding tissues, organize into hollow
tubes that evolve gradually into a mature network of blood vessels with the help of adhesion
factors, such as integrins [75]. The newly formed blood vessels are further stabilized by
Angiotensin-1, -2, and their receptor Tie-2 [76-78]. VEGF also acts as anti-apoptotic factor for
the newly formed blood vessels, as they induce expression of anti-apoptotic molecules, such as
Bcl-2, promoting endothelial cell survival [73, 79].
Tumor blood vessels are very different from those of normal adult tissue [60, 80]. Tumor
angiogenesis leads to the formation of irregular, dilated, highly branched, tortuous and
disorganized micro-vessels with compressed lumen, inconsistent diameter, highly branched
10
structures and lack of differentiation [81, 82]. Tumor vessels become hyper-permeable, described
as ‘leaky’, due to loss of adherence between endothelial junctions as well as a discontinuous
basement membrane [60, 82]. The structural aberrations described are unique to tumor
neovasculature allowing extravasation of therapeutic macromolecules and small colloidal
particles [83, 84]. These newly formed tumor blood vessels are also biochemically unique,
expressing endothelial cell surface receptor proteins [85]. These specific and distinct receptors,
unique to tumor vasculature, can be targeted with synthetic ligands possessing high specificity
and affinity, presenting an opportunity for tumor-specific therapy [86, 87].
1.4.4 Integrins
Integrins are part of the cell adhesion receptor family that regulates a diverse array of cellular
functions crucial to the initiation, progression and metastasis of solid tumors [88]. Structurally,
integrins are transmembrane receptor proteins composed of heterodimeric complexes of
noncovalently linked alpha and beta chains [88-90]. There are at least 18α and 8β subunits,
capable of forming 24 distinct heterodimers that account for the structural and functional
diversity of the integrin family [88-90]. The extracellular domain of integrins binds to the
extracellular matrix (ECM), and the intracellular domain binds to cellular cytoskeletal elements
such as actin filaments; facilitating cell adhesion, invasion and proliferation [89, 91, 92] . The
binding of integrins with ECM ligands also induces a variety of intracellular signals for major
processes such as transcriptional control, cell death, proliferation, and cell migration [93].
Furthermore, integrins have been implicated in all steps of tumor metastasis, including
detachment of tumor cells from the primary site, invasion of ECM, intravasation into the blood
11
stream, extravasation into distant target organs, and finally formation of metastatic lesions [90,
94, 95].
Integrins meditate cell adhesion to the ECM by providing a dynamic physical linkage between
the ECM and the actin cytoskeleton. Binding of integrins with ECM ligands triggers integrin
clustering, disassembly and reorganization of actin filaments and the formation of focal adhesion
complexes [89, 91, 92]. Integrins recruit cytoskeletal proteins α-actinin, talin, and skelemin, to
facilitate this integrin-actin linkage [89, 91, 92]. Integrin ligand binding also activates
intracellular signaling pathways including focal adhesion kinase, integrin-linked kinase, and Src
kinases which are required for cell migration and proliferation [91, 96, 97].
Integrins exists in either the ligated or the unligated state, in which they regulate tumour cell
survival and malignancy [88]. Ligated integrins enhances cell survival through increased
expression of Bcl-2, activation of the PI3K–AKT pathway or nuclear factor-κB (NF-κB)
signaling, and/or p53 inactivation [88, 91]. However unligated integrins on adherent cells can
recruit and activate caspase 8, resulting in apoptotic cell death [98].
Integrins not only binds to ECM but also are a major contributor to the malignant transformation
of tumor cells by initiating signaling that induce cell spreading, migration, survival, proliferation,
and differentiation [88, 91]. Expression of integrins, such as αVβ3, have been correlated with
metastatic progression in various cancers including melanoma, breast carcinoma, prostate and
pancreatic and lung cancer [88]. Integrin αvβ3 binds a wide range of ECM molecules with an
Arg-Gly-Asp (RGD) triple-peptide motif, including fibronectin, vitronectin, and proteolysed
forms of collagen and laminin [88, 91]. Furthermore, αVβ3 integrin on the tumor cells binds to
12
platelet via fibrinogen, facilitating tumor cell arrest in the vasculature and metastasis to various
tissues, including bone marrow and lungs [94, 99]. Integrins also regulate the activity of matrix-
degrading proteases, such as matrix metalloprotease 2 (MMP2) and urokinase-type plasminogen
activator (uPA), thereby facilitating tumor cell invasion by degradation of ECM [88, 94].
Integrins further contribute to tumor progression and metastasis and play a major role in tumor
angiogenesis. Proliferating tumor endothelial cells overexpress αvβ3, a key molecule for
capillary formation, however it is absent on quiescent endothelial cells and normal tissues [100,
101].
The expression of integrins in cells of various cancers and their involvement in tumor
progression has made them appealing therapeutic targets. Preclinical studies showed that integrin
antagonists inhibit tumor growth by affecting both tumor cells and cells of the tumor
endothelium [102]. Integrin antagonists, including monoclonoal antibodies MEDI-523 and
MEDI-522, among the first integrin antagonists developed, showed considerable anti-angiogenic
effect in preclinical models [103]. However, they did not show significant efficacy in phase II
clinical trials. Currently, the αvβ3 and αvβ5 inhibitor cilengitideis being pursued in a Phase III
trial in patients with glioblastoma , following encouraging activity found in Phase II clinical
trials [104, 105]..
1.4.5 Metastasis
Metastasis is a complex process and occurs in about 90% of breast cancer patients [106].
Metastasis is the stage where the tumor cells spread from a primary site to distant organs such as
the bone, lungs, regional lymph nodes, liver and brain, to form secondary tumors [107, 108].
13
Management of metastatic breast cancer is a major problem in the clinic and is currently
considered incurable.
Metastasis is a multistep process known as the invasion-metastasis cascade [14]. In order to
metastasize; cancer cells must detach and extravasate from the primary tumor, invade through
surrounding tissues and basement membrane, enter into and survive in the blood circulation,
arrest at a distant organ by adhesion to a specific endothelium receptor, extravasate across the
endothelium, migrate through extracellular matrix and finally proliferate in the target organ
[109, 110].
The potential of a tumor cell to metastasize depends on its interactions with local factors and the
microenvironment that promotes tumor-cell growth, survival, angiogenesis, invasion and
metastasis, as explained by the "seed and soil” hypothesis [109].
Invasion, the first critical step in the metastatic process, requires the cancer cells to lose their
epithelial phenotype, decrease their cell-cell attachments and break down the extracellular matrix
[108, 110]. Cancer cells can undergo an epithelial-to-mesenchymal transition (EMT) by loss of
E-cadherin which disrupts adhesion junctions between neighboring cells and thereby supports
detachment of malignant cells from the epithelial cell layer [111, 112]. Loss of E-cadherin, as
well as the gain of N-cadherin leads to the rearrangement of the cytoskeleton by mediating Rho-
induced stress fibers and the formation of lamellopodia and filopodia by Rac1 and Cdc42
activation, respectively, resulting in enhanced motility of EMT-transformed cells [113, 114].
Following an EMT-like alteration of breast cancer cells, proteases secreted by tumor-associated
macrophages and fibroblasts result in the degradation of the ECM, enabling cancer cells to
penetrate tissue boundaries [115, 116]. These proteases include matrix MMPs, cathepsin B, and
14
plasminogen activators which not only degrade ECM but also induce cleavage of matrix proteins
to generate binding sites for integrins and cell-substrate adhesion molecules which are required
to anchor cells during tissue invasion [117, 118]. EMT-like cancer cells move to the vasculature
and intravasate through the loose endothelial junctions of the tumor blood vessels [13, 119].
The bloodstream can be a highly unfavourable environment for tumor cells owing to physical
forces, the presence of immune cells, and anoikis [120]. Tumor cells bind to coagulation factors,
including the tissue factors fibrinogen, fibrin and thrombin, to create an embolus and facilitate
arrest in capillary beds of distant tissues [121]. Following extravasation into the distant tissue,
the metastatic cancer cells recruit macrophages and fibroblasts to assist with the invasion of the
new host organ [115, 122]. The seeded cancer cells further undergo a mesenchymal-epithelial
transition in order to successfully colonize the new host tissue through proliferation,
angiogenesis, and secondary breast tumor characteristic development [107, 110].
1.5 Breast cancer therapy
Breast cancer is a heterogeneous disease, with clinically distinct biological subtypes, each with
different treatment options and response to therapy. Treatment is planned based on tumor
staging (tumor size, regional lymph node metastasis and distant metastasis), biological
characteristics (receptor status) as well as the patient’s age, health, and personal preference.
Most treatment regiments require a combination of surgery, radiation therapy, chemotherapy,
and hormone therapy for successful management of breast cancer.
15
1.5.1 Local therapy
Local intervention such as surgery or radiotherapy is used as the primary treatment for early
stage breast cancer. Women can either undergo mastectomy, the complete removal of the entire
breast or have lumpectomy/breast conserving surgery (BCS) where the cancerous area and a
small amount of surrounding normal tissue is removed [123]. Breast cancer surgery has changed
dramatically over the past decade as there is a steady decline in mastectomy and more females
are opting for BCS [124, 125]. For earlier stage breast cancer mastectomy is nearly curative
(98%) unlike BCS which carries a risk of local recurrence ranging from 6-19% [126-128].
Postoperative RT following BCS has become the standard of care for patients with early-stage
breast cancer to remove any residual cancer cells after surgery. Over 50% of all cancer patients
receive RT during the course of their illness, equating to over 500,000 patients worldwide each
year [129]. RT is known to substantially reduce the risk of loco-regional recurrence and improve
breast cancer mortality [130]. Several clinical trials have shown the benefit of RT following BCS
results in equivalent survival rates as compared to radical mastectomy [130-132]. The National
Surgical Adjuvant Breast and Bowel Project (NSABP) B-24 trial found a reduction in incidence
of local recurrence to 14% from 39% for women who received RT after BCS versus women who
underwent BCS alone [127]. Even though RT is critically important in the locoregional
management of early breast cancer, it has a few limitations as well. Tumors are radiated locally
therefore, RT is limited in its utility against systemic disease when cancer spreads to other parts
of the body [127, 130, 133]. Radiation also poses risk of injury to critical organs that may be in
16
the radiation field, such as the heart, lungs, and brachial plexus since it not only affects the tumor
but also normal tissues in the field [134, 135].
The efficacy of RT can also be limited by the tumor microenvironment characterized by low
oxygen concentration (hypoxia) and acidic pH (acidosis) [136, 137]. The effect of RT is
enhanced by molecular oxygen which potentiates radiation damage to DNA that results in cell
death [138, 139]. However, studies have demonstrated that nearly 40% of breast cancers exhibit
tumor regions with oxygen concentrations below that required for half maximal radiosensitivity
enhancement, hence reducing the effectiveness of radiation therapy [140]. Hypoxia leads to
activation HIF-1 which in turn induces the expression of various genes involved in cell
death/survival pathways such as VEGF, leading to a further increase in tumor radioresistance
[65, 141, 142]. Hypoxia and HIF-1a overexpression have been shown to correlate closely with
poor prognosis in breast cancer [67]. Acidic microenvironment also activates several genes
responsible for increased DNA damage repair and induction of an aggressive cell phenotype
leading to increased radioresistance [143]. The effect of tumor microenvironment on
radioresistance is discussed in more details in section 2.1.2.
1.5.2 Systemic therapy
Despite advances in early detection and increasing use of surgery and RT in the management of
primary tumors, approximately 30% of all patients with early-stage breast cancer develop
recurrence, metastatic in most cases [144]. Systemic treatment using cytotoxic, hormonal or
immunotherapeutic agents are used either in adjuvant or neoadjuvant settings in management of
both local and systemic forms of breast cancer [145]. In an adjuvant setting, anti-cancer
17
chemotherapeutic agents are administered systemically to patients following surgical resection of
primary tumor and axillary lymph nodes, preventing disease recurrence by eliminating cancer
cells at the treatment margins and eradicating micrometastases [146, 147]. Multiple studies have
demonstrated a 23% or greater improvement in disease free survival following adjuvant therapy
in early stage breast cancer patients [148]. Treatment can also be administered prior to local
intervention in a neoadjuvant role to reduce tumor size for BCS and decrease probability of
metastasis [149, 150]. Since both neoadjuvant and adjuvant therapies have shown similar
survival rates, the particular treatment regimen for breast cancer is chosen according to
individual tumor characteristics of the patient [123, 149].
Chemotherapy is considered the first treatment option to reduce tumor burden especially in
patients with hormone resistant/refractory metastatic cancer. Chemotherapy with anthracyclines
or taxanes is widely used for the treatment of breast cancer and is considered to be the most
effective treatment especially for triple negative metastatic breast cancer [144, 147, 150]. The
chemotherapeutic agents can be given individually, in sequence or in combination. Effective
anticancer therapy for many tumors requires a combination of multiple drugs that have different
mechanisms of action, different resistance mechanisms, and different dose-limiting toxicities
[151]. The chemotherapeutic agents for breast cancer used in this thesis will be discussed below,
followed by the utility of combination chemotherapy.
1.5.2.1 Doxorubicin
Doxorubicin (Dox), an anthracycline antibiotic is one of the most widely used chemotherapeutic
agents towards breast cancer. Dox was first extracted from Streptomyces peucetius in the 1970s
18
and has since been used for the treatment of several cancers including breast, soft tissue
sarcomas, pediatric cancers, and lymphomas [152-154]. Dox usage as a first line chemotherapy
agent, most often in combination with other anticancer agents, has demonstrated benefits in
terms of response rate, time to disease progression, and overall survival [155]. In a meta-analysis
of adjuvant chemotherapy, the anthracycline-containing regimens were more effective in
preventing recurrence and increasing survival in breast cancer patients compared to patients
given non-anthracycline containing regimens [156-158]. Dox was indicated as the preferred
single agent as well within the combination regimens for treatment of recurrent or metastatic
breast cancer resulting in disease decreased mortality rates [147, 159].
Dox exerts its cytotoxic action by multiple mechanisms including DNA intercalation to induce
DNA damage; disruption of topoisomerase II mediated DNA repair and generation of free
radicals which cause further damage to DNA, cellular membranes and proteins [154, 160]. Dox
is highly effective in oxygenated regions of the tumor, where it is oxidized to an unstable
semiquinone metabolite, which when converted back to doxorubicin releases ROS. The
generation of ROS leads to DNA and cell membrane damage, lipid peroxidation and oxidative
stress resulting in cell death [160-162]. Dox has also been reported to intercalate mitochondrial
DNA and affect the cell membrane directly by binding plasma proteins resulting in formation of
ROS [163]. Dox interferes with cellular processes by activating both intrinsic and extrinsic
apoptotic pathways, allowing it to be efficacious against various cancers.
Despite the vast utility of Dox in clinical oncology, its usage is limited by serious adverse drug
reactions including myelosuppression and congestive heart failure, arising from its unselective
cytotoxicity towards both cancerous and normal cells [164-166]. Thus, its clinical use is limited
19
to a cumulative maximum dose of 450mg/m2 in humans preventing aggressive chemotherapy.
Studies have shown that approximately 10% of patients treated with Dox or its derivatives will
develop heart complications up to 10 years after treatment [167] . The exact mechanism of Dox
induced cardiotoxicity is still unclear. It is believed that the toxicity of Dox includes disturbance
of calcium homeostasis, formation of iron-Dox complexes that generate free radicals,
mitochondrial dysfunction, and damage to cell membranes [168-171]. Due to its excellent anti-
cancer efficacy, a significant amount of research is going on to further increase the clinical
usefulness of Dox by developing new agents to reduce Dox induced toxicity [172].
1.5.2.2 Mitomycin C
Mitomycin C (MMC) is one of the most active single agents against several types of cancer
types; however currently, it is only used as a second-line adjunctive agent due to its severe
toxicity [173-175]. MMC, a naturally occurring antibiotic was isolated originally from the
microorganism Streptomyces caespitosus in 1958 and became commercially available in 1974
[176]. MMC is an alkylating agent with three active groups: quinone, urethane and an aziridine
group which requires activation by reduction of the quinone. Under anaerobic conditions
reduction of the quinone initiates a cascade of reactions, opening the aziridine ring. This highly
unstable and reactive quinone can react with DNA to form monoadducts or crosslinks, where
both the urethane and aziridine moieties cross- link DNA causing lethal damage [176-178].
Under aerobic conditions, the quinone is reduced to semi-quinone via cytochrome P-450 leading
to the generation of reactive oxygen species (ROS) which can cause toxicity to cancer cells [179,
20
180]. Crosslinks can also be generated by two-electron reduction of the quinone by DT-
diaphorase to a hydroquinone resulting in cell damage [181, 182]. The preferential bioreductive
activation of MMC in hypoxic conditions makes it an ideal candidate for a selective toxin
towards hypoxic malignant cells; however administration of MMC is limited by its severe
myelosuppression toxicity [173, 176].
1.5.2.3 Drug combination therapy
Combinations of multiple anti-cancer drugs that have different mechanisms of action and
different dose-limiting toxicities are often administered for effective treatment of tumors [151].
In metastatic settings, combination chemotherapy regimens are frequently favoured over single
agents in an attempt to achieve superior tumor response rates. Ideally, a combination therapy
should meet the following criteria: 1) each drug should have single-agent activity with no cross-
resistance, 2) synergy between the two components should be determined in preclinical research
and 3) the two drugs should have non-overlapping safety profiles [183, 184]. However, all three
criteria are rarely met and many combinatorial regimens are determined empirically.
Anthracycline containing regimens have become a standard therapy in the adjuvant setting as
they are found significantly more effective in the reduction of recurrence and death over non-
anthracycline containing regimens [147]. However, many of these regimens carry a risk of
cardiotoxicity [185]. Induction of cardiotoxicity by Dox has also been observed in anthracycline-
taxane combinations without significant enhancement of treatment efficacy [186, 187]. Additive
toxicity frequently limits the development of successful combinatorial drug regimens for cancer
[151]. Consequently, many combination therapies have failed to significantly improve outcomes
21
in clinical trials compared with sequential administration of single agents [183, 188]. Therefore,
it has been proposed that an effective combination treatment regimen requires justification of the
combination with preclinical demonstration of the synergy and, not just lack of cross resistance
[183, 189].
Benefits of combination therapy have been demonstrated when combination regimens are
determined based on pre-clinical data of drug synergy with defined dosing schedules and dose
ratios [183, 190]. Anti-cancer synergy has been observed pre-clinically with concurrent
administration of Dox and MMC both in vitro and in vivo [191-193]. Mechanistic studies
suggested that synergy between Dox and MMC was due to the interaction of drug-DNA adducts
and cross-link-activated DNA repair machinery with a covalent topoisomerase IIa–DOX–DNA
complex resulting in a supra-additive level of DNA double strand breaks [191, 193]. The
observed anti-cancer synergy between the two drugs was further exploited in this thesis
(Chapters 2, 3 & 4) to reduce the tumor of breast cancer carrying animals while overcoming
barriers to chemotherapy.
2. Barriers to cancer therapy
2.1 Therapeutic resistance
Resistance to therapy — chemotherapy or radiotherapy, remains a major problem that has
hindered the effective treatment of patients with breast cancer. The mechanisms that underlie
clinical resistance are complex and multifactorial; occurring at the molecular, cellular, and
physiological level of the tumor, allowing cancer cells a number of ways to survive cancer
therapy. Resistance to therapy may be caused by alterations in the intracellular machinery of
22
cancers cells themselves or associated with the anatomical and physiological properties of the
tumor, resulting in decreased sensitivity to anti-cancer agents or radiation [194, 195]. These
mechanisms are described in more detail below.
2.1.1 Cellular and molecular causes of drug resistance
Many cellular and molecular mechanisms have been identified that contribute to development of
resistance to a multitude of anticancer agents, a phenomena termed multidrug resistance (MDR).
MDR is a major obstacle in effective chemotherapy of cancer. MDR can be intrinsic, present at
the inception of tumorigenesis, or acquired after the initial treatment with anticancer drugs [145,
195]. Often, more than one mechanism, either simultaneously or sequentially, is responsible for
the MDR phenotype. These mechanisms include over expression of drug efflux transporters,
increase drug metabolism and up-regulation of target enzymes.
Upregulation of efflux transporters: The most frequently occurring causes of MDR include the
up-regulation of membrane bound ATP-binding cassette (ABC) transporters including P-
glycoprotein (P-gp), multidrug-resistance associated protein (MRP-1) and breast cancer
resistance protein (BCRP) [196, 197]. These transport proteins serve as energy dependent drug
efflux pumps exporting anticancer drugs such as Dox, from the cell membrane/cytoplasm to
outside of cell against a concentration gradient, thus lowering the effective drug concentrations
within the cells [198]. P-gp, a 170kDa membrane glycoprotein, coded by multidrug resistant
type 1 (mdr1) gene in humans, has been regarded as the most significant cause of MDR
phenotype since its discovery in the 1970s [199]. In general, P-gp substrates are usually organic
molecules that are uncharged or weakly basic and hydrophobic in nature [200]. P-gp mediates
23
cellular drug resistance to diverse antitumor drugs such as anthracyclines (doxorubicin,
daunorubicin, mitoxantrone), vinca alkoids (vincristine, vinblastine), epipodophyllotoxins
(etoposide), and taxanes (taxol or paclitaxol). In general, P-gp substrates are organic molecules
that are uncharged or weakly basic and hydrophobic in nature [197, 200]. In addition to P-gp,
there are other drug transporters that may confer MDR phenotype to cancer cells. MRP-1, a
larger protein than P-gp, has been found to be ubiquitously expressed throughout the body [201].
Unlike P-gp, which preferentially transports neutral or mildly cationic substrates, the substrates
of MRP are generally glutathione conjugated drugs. Like P-gp, MRP-1 has been found to efflux
a variety of chemotherapeutic agents including anthracyclines, anthrecediones, vinca alkaloids
and methotrexate [202-204]. BCRP, also known as mitoxantrone resistance protein, was
originally cloned from a highly doxorubicin resistant human breast cancer cell line (MCF-
7/AdrVp) [205]. In terms of substrate specificity, BCRP seems to confer resistance to a narrower
range of drugs than P-gp and MRP-1 that includes anthracyclines, methotrexate and
camptothecins but does not include vinca alkaloids epipodophyllotoxins, paclitaxel or cisplatin
[206]. More than one of the aforementioned membrane-associated drug transporters may be
present in the same cancer cell and render the cell even more resistant to chemotherapy [205,
207, 208].
Activation of detoxification systems: Cytotoxic activity of anti-cancer agents can also be
decreased by metabolic biotransformation via glutathione-S-transferase (GST), an enzyme that
conjugates drugs with polar molecules to facilitate their excretion out of the cell [209]. It has
been shown that GST acts to sequester both MMC and DOX, preventing their toxic interaction
with the cell [210]. GST has been shown to be overexpressed in various resistant cancer cell
24
lines such as MCF-7/Adr [211-213]. The overexpressed GST modifies the drug into an end
product with reduced activity and facilitates the removal of the drug-glutathione conjugates from
the cell via MRP1.
Increased DNA repair mechanism: Most anti-cancer agents exert their cytotoxic actions through
DNA damage. However, many cancer cells have increased DNA repair mechanisms through up-
regulation of DNA repair proteins, making them resistant towards the drug [145]. Cancer cells
can become more efficient in repairing DNA damage caused by MMC and Dox promoting the
restitution of DNA structure and hence preventing cancer cell death [214]. In addition, cancer
cells can become resistant by shutting down the signaling pathways responsible for relaying the
information regarding genotoxicity to effectors of cell death resulting in continued survival and
proliferation [145].
Resistance to apoptosis: Inhibition of apoptotic pathways may also contribute to drug resistance.
Apoptosis is the process of programmed cell death characterized by cell shrinkage, membrane
blebbing, chromatin condensation and nuclear fragmentation that can be caused by DNA damage
by certain anti-cancer agents such as Dox [215]. Chemotherapy induced apoptosis may occur via
the intrinsic pathway mediated by the Bcl-2 family of proteins which consists of more than 30
anti and pro-apoptotic molecules [216]. Over-expression of Bcl-2, an anti-apoptotic factor, may
prevent apoptosis due to Dox treatment of cancer cells [217]. Over-activation of proliferation
pathways such as MAPK/ERK, PI3K/Akt can also induce resistance to apoptosis in cancer cells
[215]. Therefore, by avoiding occurrence of apoptosis, cancer cells become less sensitive to
chemotherapeutic agents.
25
From this section it is evident that resistance to chemotherapy is complex and varied. Resistance
in breast cancer can be mediated by any combination of the mechanisms mentioned above,
representing a major obstacle to treatment. All cancer patients develop resistance to therapies,
however they may have very different resistance mechanisms at work [173].
2.1.2 Mechanisms of resistance that relate to tumor
microenvironment
In addition to cellular mechanisms of resistance, there are factors related to the tumor
microenvironment that can lead to decreased efficacy of both chemotherapy and RT. The tumor
microenvironment properties that may influence the sensitivity of tumors to anti-cancer drugs
include: the requirement for drugs to penetrate tumor tissue from blood vessels and distribute
widely enough to reach target cancer cells, despite heterogeneous vascular density and blood
flow, increased interstitial fluid pressure, presence of low oxygenated hypoxic environment, low
extracellular pH and the regulation of the expression of various genes [194, 218]. The strong
interplay between various microenvironmental factors can result in strong drug resistance
phenotypes.
Tumor vasculature and blood flow: The abnormal tumor vasculature influences distribution and
delivery of drugs within the tumor [219]. Blood vessels in tumors are often dilated, disorganized,
irregular and tortuous with discontinued or absent basement membrane [82, 220]. Fenestrations
within the walls of tumor vessels make them leaky and highly permeable. As a consequence of a
disorganized vascular network and the absence of functional lymphatics, there is increased
26
interstitial fluid pressure (IFP) within tumor which impairs drug penetration into the tumor [221,
222]. These factors affect therapeutic response and lead to drug and radiation resistance
Hypoxia: The hypoxic microenvironment within the tumor can play a major role in treatment
failure. Studies have demonstrated that nearly 40% of breast cancers exhibit tumor regions with
oxygen concentrations below that required for half maximal radiosensitivity, reducing the
effectiveness of radiation therapy [140]. The efficacy of RT critically depends on the relative
level of oxygen in the tumor at the time of irradiation as oxygen enhances the formation of DNA
double strand breaks caused by free radicals generated during RT. Under well oxygenated
condition, oxygen can combine with radiation produced DNA damage to “fix” this damage and
make its repair more difficult or impossible [58]. Various studies have demonstrated that the
colony forming ability of hypoxic cells is two-to-three times more resistant to a single dose of
ionizing radiation than cells in normal levels of oxygen [223]. In addition, tumor hypoxia can be
associated with resistance to some chemotherapeutics such as bleomycin and neocarzinostatin
[224].
Low extracellular pH: Another consequence of a poorly formed and irregular vascular system is
the accumulation of breakdown products of metabolism, resulting in an increasingly acidic
microenvironment [225, 226]. Insufficient oxygen supply forces cancer cells to undergo glucose
metabolism through the glycolytic pathway instead of respiration, thereby resulting in the
formation of lactic acid. It has also been demonstrated that hypoxia activates carbonic anhydrase,
which converts CO2 and H2O molecules to carbonic acid. Both of these mechanisms culminates
in the accumulation of acidic metabolic products in the extracellular space (i.e., H+ and lactate),
rendering a mildly acidic interstitial pH (pHe < 6.9) [55, 227]. Tumor acidosis may render
27
cancer cells chemoresistant as many cancer drugs, such as Dox , are mildly basic (pKa > 7.5) and
their protonation in the extracellular space of tumors may decrease the ability of the drug to
permeate through the cell membrane (ion trapping phenomenon) [225]. In vivo studies have
shown that the uptake and efficacy of several clinically used cancer drugs are reduced by the
acidic pHe of solid tumors [228]. Also, the physiologic changes cancer cells undergo in response
to low pHe can also contribute to chemoresistance, including reduced apoptotic potential, genetic
alteration (p53 mutations) and elevated activity of Pgp [228].
Regulation of gene expression: Hypoxia and low extracellular pH within the tumor causes stress
induced alteration of gene expression which can further contribute to treatment failure in cancer
[229]. Hypoxia leads to chronic over activation of hypoxia-inducible-factor-1 (HIF-1) which
plays a pivotal role in adaptive responses to hypoxia by modulating various cellular functions
[230]. Upon activation, HIF-1 binds to the hypoxic response element, thereby promoting
transcription of various genes including, genes involved in angiogenesis(VEGF), glycolysis
(GLUT-1) and low pH (CAIX) to facilitate malignant tumor growth and survival [231]. It has
also been shown that acidic pH can also induce VEGF expression distinct from the HIF mediated
pathway [232]. Hypoxia and acidosis are hallmarks of the metabolic environment of solid
tumors, and together they make the tumor more resistant to therapy.
2.2 Treatment side effects
The use of radiation therapy and chemotherapy has contributed to significant improvements in
survival among patients with early stage breast cancer. However, dose-limiting acute toxicities
and long-term adverse effects are the major drawbacks of conventional therapy. Radiation to the
28
breast causes injury to the skin, chest wall, lung and heart [233]. Pericarditis, i.e. inflammation to
the pericardium, is a typical acute manifestation of high dose radiation injury, while chronic
pericardial disease, coronary artery disease, cardiomyopathy, and valvular disease can manifest
themselves years after the initial treatment [234, 235]. Meta-analysis of randomized control trials
has demonstrated that patients who received RT had a higher risk of vascular mortality than
those who did not [130]. The risk of cardiac damage correlates with radiation dose-volume and
fractionation, hence preventing administration of higher doses of radiation to control tumors
[236, 237].
Anti-cancer drugs are administered systemically in order to control both local and metastatic
breast cancer. These anti-cancer drugs are chosen/ designed to kill proliferating cells without any
specificity for cancerous cells per se. As a result, these drugs such as Dox and MMC also cause
toxicity to proliferating cells in the gastrointestinal tract, hair follicles and bone marrow resulting
in common short term side effects such as vomiting, nausea and alopecia as well as therapy
limiting side effect of myelosuppression [173, 238]. In addition, Dox carries a significant risk for
dose dependent longer term cardiac toxicity that is attributed to its major phase I metabolite
doxorubicinol [239, 240]. The incidence of congestive heart failure reaches 5% for Dox treated
patients with cumulative doses of 400mg/m2[241]. Doxorubicinol induces cardiomyocyte death
resulting in congestive heart failure by interfering with the sarcoplasmic reticulum of the heart.
In an effort to overcome this cardiotoxicity, co-administration of cardioprotective agents along
with Dox or the use of other anthracycline derivatives with lower demonstrated cardiotoxicity
have been attempted [242-245]. However, these approaches have often resulted in reduced anti-
cancer efficacy, preventing effective outcomes.
29
3. Nanoparticulate systems in cancer therapy
The effectiveness of conventional cancer therapy is often limited by dose-restricting systemic
toxicity, tumor hypoxia, and the MDR phenotypes of cancer cells, as previously discussed. Over
the past two decades, significant progress has been achieved in the field of nanotechnology to
overcome these problems and offers promising and effective alternatives for cancer treatment
[246, 247].
Due to the small size of anti-cancer drugs, they are rapidly cleared from the bloodstream, thus
reducing their effective concentration within the tumor [248]. Nanoparticle drug delivery
systems allow the manipulation of the biodistribution properties of anticancer drugs by
modifying the physiochemical properties (such as hydrophobicity) of the drug delivery system
[246, 248]. This can result in the prolonged bloodstream circulation time of the drug, enabling an
adequate amount of the drug to reach the target tumor site. The encapsulated drug is also
protected from harsh environments of the body, drug metabolizing enzymes and extensive
binding to serum proteins while in the blood circulation, resulting in enhanced efficacy [249].
Additionally, the properties of nanoparticles can be tuned to improve delivery of
insoluble/exposure sensitive drugs and to specifically control the site and rate of drug cargo
release [246, 248].
Nanoparticle drug delivery system can exploit the tumor architecture to selectively target their
payloads to cancer cells, either by passive or active targeting [249]. Nanoparticles with an
optimum size (50 nm - 300 nm) are able to take advantage of the leaky tumor vasculature and its
poor lymphatic drainage, to preferentially accumulate in tumor tissue, a phenomenon known as
30
enhanced permeability and retention (EPR) effect [83, 84]. The enhanced accumulation of drug
encapsulated nanoparticles at the tumor site, and, hence, lower drug concentrations in healthy
tissues, results in increased anticancer efficacy and decreased systemic side effects. Once the
nanoparticle delivery system reaches the tumor, they are able to enter tumor cells and overcome
the MDR efflux pump mechanism [246, 249], currently one of the major causes of treatment
failure in clinic. Therefore, the rational design of nanoparticle systems has the potential to
enhance chemotherapy by delivering enhanced levels of drug to the tumor site for effective
periods of time.
3.1 Optimal characteristics of nanoparticle delivery system
Nanoparticles are composed of three basic components: a core, surface and a payload [250]. The
payload can be an anti-cancer agent which is contained within the core of the nanoparticle. Both
the core and surface must be rationally designed to deliver high payload to the tumor while
minimizing toxicity to other healthy tissues [250, 251]. One of the most important considerations
for nanoparticle design is the safety of the delivery system [250, 251]. The nanoparticle core
materials must be biodegradable, safe for repeated administration and should not produce any
toxic byproducts [250, 251]. In addition, the core of the nanoparticle should have high drug
loading capacity with maximum drug release from the particle at the tumor site [250].
The surface characteristics of the nanoparticle must ensure the stability of the formulation prior
to injection (i.e. prevent aggregation), and the in vivo biocompatibility of the nanoparticle
following administration [249, 250, 252]. Anionic surface charge is preferred over cationic
charge as cationic nanoparticles induce significantly more hemolysis than anionic nanoparticles
31
[249]. Therefore, for clinical applications, neutral or negatively charged nanoparticles are
preferred. Nanoparticles with prolonged circulation time are desired to increase accumulation at
the tumor site [252, 253]. This can be achieved by modifying the surface of nanoparticles with
the addition of a hydrophilic polymer [253]. Poly(ethyleneglycol) (PEG) is the most widely used
hydrophilic polymer and is able to confer ‘stealth’ properties to nanoparticles [253, 254].
PEGylation of the nanoparticle surface prevents opsonin binding by mononuclear phagocytic
system (MPS), improves the colloidal stability, increases circulation time resulting in increased
nanoparticle accumulation at the tumor site [254].
The size of the nanoparticle also needs to be optimized for systemic administration to prevent
embolus formation, which can occur when the particles are too large, or rapid clearance of
particles from the circulation when they are less than 10 nm [249, 252]. The size range of 50 nm
to 100 nm has been found to be optimal in promoting the passive targeting of nanoparticles to
tumor tissue by the EPR effect [249, 251]. However, the acceptable nanoparticle size is also very
material dependent and can vary from polymeric to inorganic to lipid-based nanoparticle
formulations [255].
The formulation of nanoparticle system should be simplistic in order to allow scale up
production for clinical application. Careful selection of core and surface materials are required
for the development of safe and biocompatible nanoparticles with prolonged circulation, high
accumulation in tumor and good drug loading and drug release characteristics [250].
32
3.2 Targeting tumor with nanoparticles
Conventional chemotherapy, i.e. the administration of anti- cancer agents systemically is
subject to a potential non-specific mechanism of action, a lack of selectivity for tumor cells, and
an undesired killing of normal cells [248, 249]. Additionally, due to low bioaccessibility of these
drugs to tumor tissue larger doses are required, leading to increased toxicity to normal cells and
an increased incidence of multi-drug resistance [246, 249]. Nanoparticle drug delivery systems
carrying the anti-cancer agents exploit the abnormal pathophysiology of tumor tissues to
selectively deliver their payloads to cancer cells either by passive or active targeting [249]. In the
passive mode, nanoparticles take advantage of the unique tumor pathophysiology [84, 256] and
in the active mode; nanoparticles selectively bind to receptors expressed on tumor tissue [249,
257].
3.2.1 Passive targeting
Passive targeting is the most frequently used mechanism to selectively deliver nanoparticles
loaded with anti-cancer agents to the tumor site. Passive targeting exploits the anatomical and
physiopathological characteristics of tumor vasculature, that give rise to the EPR effect [83, 84,
249]. The growing tumor mass triggers the development of new blood vessels to supply
proliferating cancer cells with oxygen and nutrients [60, 69]. This process, known as
angiogenesis, promotes the rapid development of new, irregular blood vessels that present a
discontinuous epithelium and lack the basal membrane of normal vascular structures [60, 69, 83,
84]. These characteristics result in leaky blood vessels which are structurally unique to tumor
vasculature. Accumulation of nanoparticles within the tumor is further enhanced due to poor
33
lymphatic drainage in tumors [83, 84]. The leaky blood vessels and compromised lymphatic
drainage together result in the EPR effect, which constitutes an important mechanism by which
nanoparticles can selectively accumulate in the tumor interstitium [83, 84]. Nanoparticles need
to be between 10nm - 500nm in size (depending on nanoparticle material) with anionic or neutral
surface charge to take advantage of the EPR effect [83, 84]. The surface of the nanoparticle must
also resist opsonization and MPS uptake to ensure long circulation of the nanoparticles, as EPR
effect-mediated passive tumor uptake is optimized with particle circulation times of at least 6
hours [83, 84].
3.2.2 Active targeting
Active targeting of nanoparticles involves attaching a specific ligand(s) to the surface of
nanoparticles that can recognize and bind to complementary molecules, or receptors, found on
the surface of tumor cells. Representative ligands include antibodies, antibody fragments,
proteins, aptamers, peptides, or small molecules [258, 259]. Targeting ligands are often selected
because of their high specificity and high affinities towards overexpressed receptors on cancer
cells or the cells present in the tumor microenvironment [257, 259, 260]. In order to benefit from
this increased affinity; actively targeted nanoparticles need to be in the proximity of their target
[260, 261]. Currently, actively targeted nanoparticles are used as a complementary strategy to
EPR to further augment the efficiency of cancer nanomedicines.
Designing actively- targeted nanoparticle drug carriers is a complex process as one needs to
consider the nanoparticle architecture, the ligand conjugation chemistry and the types of ligand
available all of which can contribute to the efficacy of the system. Furthermore, physicochemical
34
properties like the choice of the targeting ligand [262], the ligand density [262], and the size of
the nanoparticles [263] can also affect the efficacy of the active targeting strategy in vivo.
Practical aspects of particle design must also be taken into account, such as the production cost,
scalability and stability of the ligand.
Various monoclonal antibodies (mAbs) have been approved by the United States Federal Drug
Administration (FDA) including cetuximab, rituximab, trastuzumab and bevacizumab for
successful tissue based targeting [264, 265]. However, incorporation of mAbs to nanoparticles
have been challenging and currently unsuccessful since mAbs are complex and large (~150 kDa)
molecules and require complex conjugation techniques to be effective [259, 264]. Antibodies are
also recognized by the immune cells and MPS resulting in faster clearance from the circulation
[264, 266]. Furthermore, the increased sensitivity of mAbs to temperature, enzyme and organic
solvents causes technical challenges in the preparation of nanoparticles incorporating them. [264,
266]. Recently, peptides have been used for active targeting of nanoparticles to cancer cells.
Their small size, high stability and ease of synthesis for large scale production with excellent
quality control, make peptides attractive candidates for active targeting. RGD peptide which
strongly and specifically binds αvβ3 integrin has been extensively investigated in targeting
nanoparticles for disrupting tumor angiogensis [267]. αvβ3 integrin is highly expressed on breast
cancer cells, making it an appealing target [88, 99].
Despite numerous preclinical publications, only one actively targeted nanoparticle formulation,
Abraxane is currently used clinically [268-270]. The major shortcomings associated with active
targeting include particle recognition by the immune system, the identification of ligands specific
to cancer cells and the technical complexity of the formulations all of which limit the clinical
35
applicability of actively targeted nanoparticles [249, 260]. However, continuing research in
development of novel ligands may overcome these limitations and significantly improve the
utility of nanoparticles.
3.2.3 Solid lipid and polymer-lipid hybrid nanoparticles
Solid lipid nanoparticles (SLN) and polymer-lipid nanoparticles (PLN), both involving lipid
emulsions, are an alternative class of drug carrier systems [271, 272]. They remain solid at body
temperature and are characterized by high drug loading capacity, controlled drug release,
improved drug stability and excellent biocompatibility [271]. The core of the nanoparticles is
made of biodegradable lipids [273] and the surface of the nanoparticles has a net negative charge
shielded by a PEG corona [273, 274]. The PEG layer prevents uptake by activated MPS and
activated macrophages resulting in a longer circulation time [273, 275]. The lipid core of SLN
allows high levels of drug loading of hydrophobic drugs, but only low levels of hydrophilic
drugs, such as Dox [275]. In order to achieve high level loading of hydrophilic drugs, an anionic,
biocompatible polymer was introduced in the core of the nanoparticle resulting in a novel
formulation called PLN [276]. The anionic polymer is capable of complexation to cationic drug
species to form a neutral drug-polymer conjugate, easily loaded into the lipid core [276-278].
The drug-polymer ionic complex can be disrupted by physiological levels of divalent cations like
calcium, to enhance the release of complexed chemotherapeutic drugs. PLN allow the
simultaneous loading of hydrophilic and hydrophobic drugs with high loading capacity and good
release kinetics [276-278]. The small size of PLN allows for passive tumor uptake by
extravasation at the tumor mass through the vascular fenestrations, and retention in the tumor
36
mass due to lack of lymphatic drainage [84, 249]. The PEGylated surface of PLN allows taking
advantage of active targeting by linking short peptides, such as cyclo-Arginine-Gycine-
Aspartate-D Phenylalanine-Lysine (cRGDfK) which binds to αvβ3 integrin receptors
overexpressed by breast cancer cells [102].
SLN and PLN formulations have shown potential for application to cancer chemotherapy in
preclinical investigations [193, 277, 278]. It was demonstrated that Dox and Mitomycin C
(MMC) loaded PLN (DMsPLN) had improved cytotoxicity compared to the free drug solutions
in MDR breast cancer cell lines overexpressing P-gp transporter [193]. The systemic
administration of DMsPN have shown superior anti-cancer efficacy and tolerability of therapy in
a murine EMT6 mammary carcinoma model compared to equal concentrations of the free drugs
[279]. Furthermore, SLN have been conjugated with cRGD and an optimized concentration of
cRGD was determined to increase tumor specific drug delivery [280]
4. Goal for this work
The overall goal for this project was to improve breast cancer chemo- and radiation therapy using
nanoparticulate delivery system. This has been achieved by developing and assessing 3 systems.
The following hypotheses were investigated to support the goal:
1) Dox-MMC co- loaded PLN (DMsPLN) will overcome multiple membrane transporters
(MRP and BCRP) mediated MDR in breast tumor cells, resulting in enhanced cancer cell
kill compared to the free drug.
2) DMsPLN will preferentially accumulate in solid tumors and result in enhanced efficacy and
more tolerable chemotherapy in both sensitive and resistant breast tumor model.
37
3) RGD conjugated PLN will accumulate in lung metastasis of integrin αvβ3-overexpressing
breast cancer cells and will be more effective at arresting the growth of lung metastasis of
human breast cancer
4) MD nanoparticles will produce oxygen to attenuate tumor hypoxia and acidosis in vivo and
enhance radiation therapy.
5. Organization of thesis
The remainder of the thesis is organized into six chapters with the following contents:
In Chapter 2, a demonstration is made that the synergism seen between Dox and MMC in human
MDA-MB 435 breast cancer cells, a P-gp overexpressing cell line can also be extended to other
human MDR breast cancer cells (MCF 7 MX and MCF7 VP) that overexpress MRP1 or BCRP. It is
also demonstrated that PLN have the ability to overcome multiple membrane efflux pumps that
confer MDR phenotype to cancer cells and enhance the efficacy of Dox, MMC and dual agent loaded
PLN in both wild type and MDR human breast cancer cell lines in vitro. Treatment of MDR cells
with PLN encapsulating anticancer agents result in significantly enhanced cell kill compared with
free Dox or MMC solutions at equivalent doses. These finding have been published:
P. Prasad, J. Cheng, A. Shuhendler, A.M. Rauth, and X.Y. Wu, A novel nanoparticle formulation
overcoming multiple multidrug efflux pumps in human breast cancer. Drug Delivery and
Translational Research, 2012, 2 (2) 95-105.
Chapter 3 demonstrates the in vivo efficacy and safety of Dox–MMC co-loaded stealth polymer lipid
hybrid nanoparticles (DMsPLN). The efficacy and systemic toxicity of DMsPLN are evaluated
38
against clinically used PEGylated liposomal doxorubicin (PLD) in mice bearing MDA- MB
435/LCC6 wildtype (WT) or multidrug resistant (MDR) orthotopic tumors. Tumor size as a function
of time is used to determine the efficacy of therapy and changes in mouse body weight and damage-
associated blood enzymes were measured to assess the toxicity of the treatment. The potential of
DMsPLN to inhibit angiogenesis in solid tumors is also investigated as free Dox was reported to
exhibit anti-angiogenic effects. DMsPLN are superior to PLD in both animal models in terms of both
anti-cancer efficacy and tolerability of therapy and also demonstrated anti-angiogenic effect. These
finding have been published:
P. Prasad, A. Shuhendler, P. Cai, A.M. Rauth, and X.Y. Wu. Doxorubicin and mitomycin C co-
loaded polymer-lipid hybrid nanoparticles inhibit growth of sensitive and multidrug resistant human
mammary tumor xenografts. Cancer Letters, 2013, 334(2):263-73.
Chapter 4 presents the biodistribution, in vivo efficacy and safety of DMsPLN and RGD
conjugated DMsPLN in a murine lung metastatic model of human breast cancer. A metastatic
breast tumor model is successfully developed using the MDA-MB 231-luc-D3H2LN which
allowed for non-destructive monitoring of tumor growth and metastases using bioluminescence
imaging. Whole animal imaging demonstrates the localization of the fluorescent nanoparticles in
the metastatic breast cancer site. The efficacy and systemic toxicity of nanoparticles are evaluated
against free Dox-MMC solutions. Integrin-targeted RGD-DMsPLN resulted in a significant
reduction in lung metastases. Notably, DMsPLN treated mice also demonstrate complete absence
39
of cardiac toxicity which was prevalent in the group treated with free Dox-MMC solutions. The
findings from this work will be submitted to Journal of Controlled Release.
In Chapter 5, a highly reactive, biocompatible and colloidally stable albumin-based manganese
dioxide nanoparticle (A-MnO2 NP) system for the modulation of the in vivo tumor
microenvironment is developed. It is demonstrated that A-MnO2 NPs can simultaneously attenuate
tumor hypoxia and acidosis in vivo in a murine model of breast cancer by reacting with excessive
levels of endogenous H2O2 produced by cancer cells while down regulating expression of
hypoxia-inducible factor-1 alpha (HIF-1α) and vascular endothelial growth factor (VEGF) in solid
tumors. The oxygen generating property of A-MnO2 NP is further utilized to enhance radiation
treatment in mice bearing solid tumor. It is demonstrated that combination treatment of the tumors
with NPs and ionizing radiation significantly inhibits breast tumor growth, increases DNA double
strand breaks and cancer cell death as compared to radiation therapy alone.
The results from this study have been published:
P.Prasad, C.R. Gordijo, A.Z. Abbasi, A. Maeda, A. Ip, A. M. Rauth, R.S DaCosta, and X.Y. Wu,
Multifunctional Albumin–MnO2 Nanoparticles Modulate Solid Tumor Microenvironment by
Attenuating Hypoxia, Acidosis, Vascular Endothelial Growth Factor and Enhance Radiation
Response, ACS Nano, 2014, 8(4), 3202-3212.
In Chapter 6, a final summary and analysis of the work accomplished is presented. Possible
directions for future work are also discussed.
40
Chapter 2 A novel nanoparticle formulation overcomes
multiple types of membrane efflux pumps in human breast
cancer cells
Preethy Prasad1, Ji Cheng1, Adam Shuhendler1, Andrew M. Rauth2, Xiao Yu Wu1
1Department of Pharmaceutical Sciences, Leslie Dan Faculty of Pharmacy, University of
Toronto, 144 College Street, Toronto, Ontario, Canada, M5S 3M2
2Division of Experimental Therapeutics, Ontario Cancer Institute, 610 University Ave,
Toronto, Ontario, Canada M5G 2M9
This work has been published in Drug Delivery and Translational Research, 2012, 2 (2) 95-105
All work in this manuscript was performed by P.Prasad with assistance from the co-authors.
Permission to reproduce the publication in this thesis was received from the publisher (see
attached Reproduction Permission).
41
1 Abstract
Multidrug resistance (MDR) in cancer cells can involve overexpression of different types of
membrane drug efflux pumps and other drug resistance mechanisms. Hence, inhibition of one
resistance mechanism may not be therapeutically effective. Previously we demonstrated a new
polymer lipid hybrid nanoparticle (PLN) system was able to circumvent drug resistance of P-
glycoprotein (P-gp) over-expressing breast cancer cells. The objectives of the present study were
two-fold: 1) To evaluate the ability of the PLN system to overcome two other membrane efflux
pumps - multidrug resistance protein 1 (MRP1+) and breast cancer resistance protein (BCRP+)
overexpressed on human breast cancer cell lines MCF7 VP (MRP1+) and MCF7 MX (BCRP+).
2) To evaluate possible synergistic effects of doxorubicin (Dox)-mitomycin C (MMC) in these
cell lines. These objectives were accomplished by measuring in vitro cellular uptake,
intracellular trafficking, and cytotoxicity (using a clonogenic assay and median effect analysis),
of Dox, MMC or Dox-MMC co-loaded PLN. Treatment of MDR cells with PLN encapsulating
single anticancer agents significantly enhanced cell kill compared to free Dox or MMC solutions.
Dox-MMC co-loaded PLN were 20-30 folds more effective in killing MDR cells than free drugs.
Co-encapsulated Dox-MMC was more effective in killing MDR cells than single agent-
encapsulated PLN. Microscopic images showed perinuclear localization of fluorescently-labelled
PLN in all cell lines. These results are consistent with our previous results for P-gp
overexpressing breast cancer cells suggesting the PLN system can overcome multiple types of
membrane efflux pumps increasing the cytotoxicity of Dox-MMC at significantly lower doses
than free drugs.
42
Key words: Multidrug resistance, Breast Cancer, PLN-nanoparticles, Doxorubicin-
mitomycin C
2 Introduction
A major clinical obstacle in cancer therapy is the resistance to a multitude of anticancer
agents, a phenomena termed multidrug resistance (MDR) [194, 197, 272, 281, 282]. MDR can be
intrinsic, present at the inception of tumorigenesis, or acquired after the initial treatment with
anticancer drugs [194, 195]. The development of drug resistance in tumor cells is believed to be
a cause of treatment failure despite aggressive chemotherapy. The mechanisms that underlie
clinical drug resistance are complex and multifactorial, allowing the cancer cells to survive
chemotherapy via many escape routes [283]. Often, more than one mechanism, either
simultaneously or sequentially, is responsible for the MDR phenotype. Alterations in the
intracellular machinery of cancer cells are commonly implicated in the development of MDR,
including over-expression of membrane spanning adenosine tri-phosphate (ATP)-dependent drug
efflux pumps, enhanced drug inactivation, enhanced DNA damage repair mechanisms, reaction
with increased levels of glutathione, and evasion of apoptosis [195, 283].
One of the most frequently occurring causes of drug resistance is the up-regulation of
membrane bound ATP-binding cassette (ABC) efflux transporters in cancer cells. These
transporter proteins mediate resistance to a broad range of structurally diverse anti-cancer agents
by actively pumping cytotoxic drugs outside the cell against a concentration gradient, thus
lowering effective drug concentrations within the cell [202, 281, 284]. Several proteins including
P-glycoprotein (P-gp), multidrug-resistance associated protein (MRP1) and breast cancer
43
resistance protein (BCRP) have been identified in cancer cells [196, 208, 285-287]. P-gp
mediates cellular drug resistance to diverse antitumor drugs such as anthracyclines (doxorubicin,
daunorubicin, mitoxantrone), vinca alkoids (vincristine, vinblastine), epipodophyllotoxins
(etoposide), and taxanes (taxol or paclitaxol). In general, P-gp substrates are organic molecules
that are uncharged or weakly basic and hydrophobic in nature [200, 288]. MRP1, a larger protein
than P-gp, has been found to be ubiquitously expressed throughout the body [201, 202]. Unlike
P-gp, which preferentially transports neutral or mildly cationic substrates, MRP generally
transports neutral or anionic hydrophobic compounds and glutathione conjugated drugs. Like P-
gp, MRP1 has been found to efflux a variety of chemotherapeutic agents including
anthracyclines, anthrecediones, vinca alkoids and methotrexate [201-204, 288]. BCRP, also
known as mitoxantrone resistance protein (MXR), was originally cloned from a highly
doxorubicin resistant human breast cancer cell line (MCF-7/AdrVp) and is a homodimer of two
half transporters [205]. In terms of substrate specificity, BCRP seems to confer resistance to a
narrower range of drugs than P-gp and MRP1 that includes anthracyclines, methotrexate and
camptothecins but does not include vinca alkoids, epipodophyllotoxins, paclitaxel and cisplatin
[206]. It has been shown that some stem cells and tumor cells, present in a hypoxic environment,
may exhibit protection from anti-cancer agents due to overexpression of BCRP induced by
hypoxia [289]. The exact cross-resistance profile for each transporter is distinct, yet overlapping.
A number of studies have been conducted to relate the expression of ABC transporters to the
response of breast cancer to chemotherapy with mixed results. Some reported no significant
influence of P-gp or MRP1 expression on survival of breast cancer patients receiving adjuvant
therapy [290, 291]. In contrast, one study demonstrated a link between expression levels of MDR
44
transporters with progression of disease and response to treatment in primary invasive breast
cancer [292, 293]. These contradictory findings in different studies may be due to the complexity
of MDR mechanisms, which make it difficult to establish a link between levels of ABC
transporter and response to chemotherapy [294]. Also, more than one membrane drug transporter
may be present in the same cancer cell rendering the cell even more resistant to chemotherapy
[196, 208, 287].
Various strategies have been developed for overcoming drug resistance. One of the most
prominent strategies is the development and use of P-gp inhibitors for the circumvention of
MDR in cancer patients. The first generation P-gp inhibitors (e.g. cyclosporine A, verapamil)
have undergone phase III clinical trials for various types of cancer [295-298] with only a few
demonstrating statistically significant positive outcomes in overall survival in patients with
breast cancer [297] and non small cell lung cancer [298]. However, these inhibitors have not
reached routine clinical use due to unacceptable toxicity and the negative results obtained from
the majority of the phase III clinical trials. Second generation P-gp inhibitors, such as the
cyclosporin derivative PSC-833 (Valspodar™), were developed with lower inherent toxicities,
but their use has been limited due to unpredictable pharmacokinetic interactions and
development of secondary toxicities [295, 296]. Clinical trials with third generation modulators
(e.g. tariquidar, zosuquidar and laniquidar) are still ongoing; however some trials have been
terminated due to increased toxicity [214, 215, 287, 299-301]. Several antisense oligonucleotides
and small interfering RNA (siRNA) have been used to inhibit P-gp in vitro [302-304]. Though
these studies have shown promising results, inhibition of one transporter may be insufficient to
reverse chemoresistance because a single anticancer agent can be a substrate of multiple efflux
45
transporters. For example, doxorubicin is a substrate of P-gp/MDR1, BCRP and MRP1 [272,
294]. Moreover, MDR events independent of drug efflux may not be circumvented by using
these specific transporter inhibitors.
Another strategy is to use nanoparticulate drug carriers (e.g. liposomes, nanoemulsions
and solid lipid nanoparticles (SLN) to deliver anticancer drugs directly to tumor cells [193, 277,
278, 305-310]. The nanocarriers can circumvent the efflux pumps allowing the delivery of drugs
at high concentrations to the cancer cells. Our group has previously developed a polymer lipid
nanoparticle (PLN) system and demonstrated that PLN formulations improved cytotoxicity of
anticancer agents compared to the free drug solutions in MDR human breast cancer cell lines
[193, 277, 278, 310]. Dox loaded PLN resulted in approximately an eight-fold increase in cell
kill of P-gp overexpressing human breast cancer cells [278]. In addition, Dox and MMC dual-
agent loaded PLN (DM-PLN) were effective in killing MDR breast cancer cells at 20-30 fold
lower doses than the free drugs [193]. The enhancement in anticancer activity was attributable to
the more efficient delivery of the drugs to the site of drug action (i.e., the nuclei of the cells)
where Dox and MMC interacted synergistically, causing more DNA double strand breaks, and
were inaccessible to the effect of the drug efflux pumps [191]. Thus, the co-encapsulation of Dox
and MMC in the same nanoparticle not only overcame P-gp-mediated MDR but also could
reduce dose-limited side effects of conventional chemotherapy.
The objective of this work was to determine if the synergism between Dox and MMC and
the efficacy of DM-PLN seen in P-gp over-expressing breast cancer cells can also be obtained in
breast cancer cells expressing other types of efflux pumps, i.e., MRP1 (MCF7 VP) and BCRP
(MCF7 MX). The parental MCF7 breast cancer cell line was derived from pleural effusion from
46
a metastatic breast cancer patient [311] and its derivative cell lines MCF7 VP and MCF7 MX
were selected in etoposide and mitoxantrone, respectively [312, 313]. The efficacy of Dox,
MMC and dual agent loaded PLN was assessed in these MDR human breast cancer cell lines
using a clonogenic assay and compared with wild type MCF7 cells. Possible synergistic effects
of the Dox -MMC combination were evaluated by the median effect analysis. Intracellular
transport of the nanoparticles was examined by fluorescence microscopy.
3 Materials and methods
3.1 Chemicals and reagents
Myristic acid, poly(ethylene glycol)-100-stearate (PEG100SA), poly(ethylene glycol)-40-stearate
(PEG40SA) and all other chemicals, unless otherwise mentioned, were purchased from Sigma-
Aldrich Canada (Oakville, ON, Canada). Mitomycin C and doxorubicin were purchased from
Polymed Therapeutics (Houston, TX, USA). Hydrolyzed polymers of epoxidized soybean oil
(HPESO) was a gift from Drs. Z. Liu and S. Erhan (Food and Drug Administration, Washington,
DC, USA). Pluronic F68 (PF68) (non ionic block copolymer) was a kind gift from BASF Corp.
(Florham Park, NJ, USA). All cell culture plastic ware was purchased from Sarstedt (Montreal,
QC, Canada). Cell culture medium, Dulbecco’s Modified Eagle Medium (DMEM) and
phosphate buffered saline (PBS) were obtained from Tissue Culture Media Facility, Ontario
Cancer Institute (Toronto, ON, Canada). Fetal bovine serum (FBS) and trypsin were purchased
from Invitrogen, Inc. (Burlington, ON, Canada).
47
3.2 Formulation and characterization of PLN
Drug loaded PLNs were prepared as previously described with a few modification [193, 277,
278, 309, 310]. Briefly, a mixture of 50mg of myristic acid, 4mg of PEG100SA and 8mg of
PEG40SA was melted in a 15ml conical tube at 65°C. Drugs (Dox, MMC or Dox and MMC
together), HPESO, PF68, and water were added in different amounts depending on the
formulation required, according to Table 2.1. The solution was stirred for 20 minutes and then
ultrasonicated using a Hielscher UP100H probe ultrasonicator (Hielscher USA, Inc. Ringwood
NJ, USA) at 80% peak amplitude and 5 mm probe depth in solution for 5 minutes. The entire
emulsion was immediately transferred into 5 mL of distilled deionized water and stirred on ice.
Particle size and zeta potential were measured by dynamic light scattering and electrophoretic
mobility, respectively, using a NICOMPTM
380ZLS (PSSNICOMP, Santa Barbara, CA, USA)
apparatus. The morphology of PLN was examined using transmission electron microscopy
(TEM) (Hitachi Canada, Ltd., Mississauga, ON, Canada) following negative staining with
phosphotungstic acid.
Table 2.1. Composition of various PLN formulations
Formulation Drugs added
HPESO (µl of 50
g/l stock)
PF68 (µl of 100
g/l stock) Water (µl)
Blank n/a n/a 50 388
Dox 400 µl of 12.5g/l
Dox
50 50 0
MMC 4 mg of dry
MMC
n/a 50 388
Dual Agent 400 µl of 12.5 g/l
Dox + 4 mg of
dry MMC
50 50 0
48
3.3 Measurement of drug loading and encapsulation efficiency of PLN
A filtration method, as previously described, was used to determine drug loading of PLN [309].
Immediately after formulation, the PLN suspension was centrifuged through a 0.1 µm filter and
the drug in the filtrate was assayed spectrophotometrically at 540 nm for Dox and 364 nm for
MMC. Drug loading (%wt drug/wt lipid) and encapsulation efficiency (%wt loaded drug/wt total
drug) were then calculated.
3.4 Cell maintenance
Human breast cancer cell line MCF7 WT, its etoposide (VP-16)-selected derivative MCF7 VP
(MRP1+) and mitoxantrone-selected derivative MCF7 MX (BCRP+) were kindly provided by
Dr. Stuart A. Berger (University Health Network, Toronto, ON, Canada) and Dr. Erasmus
Schneider (Wadsworth Center, Albany, NY, USA) respectively. MCF7 WT and MCF7 VP were
grown in DMEM-high glucose (4.5 g/L) medium and supplemented with 10% FBS. MCF7 MX
were grown in DMEM–low glucose (1.0g/L) medium supplemented with 10% FBS. Cells were
grown as monolayers in plastic flasks at 37°C in a humidified incubator with 5% CO2. Cell
doubling times were typically 24h. Cells were trypsinized and subcultured at 50-fold dilution
once they were confluent. Every three months, new cultures were initiated from frozen stocks of
cells.
3.5 Clonogenic assay
Clonogenic assays were performed to evaluate the effectiveness of the tested treatment on cancer
cell proliferation. In this assay, 1× 105 cells were plated in a 6 cm Petri dish with 5ml of growth
medium and incubated for 24 hours at 37°C in a 5% CO2 atmosphere. These cells were treated
49
for 1hour with one of the following treatments: (1) Dox, (2) MMC, (3) Dox and MMC together,
(4) Dox-PLN, (5) MMC-PLN and (6) DM-PLN. Cells were then washed with PBS and detached
by typsinization. The cells were diluted and replated at 100 or 1000 cells per six cm dish in 5ml
of growth medium for 10 days at 37°C. Growth medium was removed and the cells were fixed
and stained with a 0.5% solution of methylene blue in 70% ethanol. The number of colonies
formed was counted and percent plating efficiencies (PE) were calculated (number of colonies
formed × 100%/ number of cells plated) for each treatment. The results were reported as
normalized PE, which were determined by dividing the PE of treated cells by the PE of the
control (untreated cells) [313]. For every concentration point of each treatment, 6 samples were
prepared and the experiment was repeated at least three times with cells from different passages.
The control plating efficiencies were found to be 54 ±13 for MCF 7WT, 72 ±16 for MCF 7 VP
and 68 ±13 for MCF 7 MX cells (Identify means +/- standard error of the mean (SEM).
3.6 Median Effect Analysis
Median effect analysis was employed to analyze the results from the clonogenic assay [314,
315]. The cells were treated with the drugs alone or in combination at constant molar ratios
(MMC: Dox molar ratio of 2:1) for five levels of drug dose. This ratio was chosen based on
previous studies demonstrating that Dox was twice as effective as MMC per mole at killing
tumor cells [193]. The median effect plot of log [(fa)-1
-1]-1
vs. log [D] was generated for the
three treatment groups (1) Dox alone, (2) MMC alone and (3) Dox and MMC together. Here, D
is the drug concentration and fa is the fraction of cells affected (unable to form colonies). Dox
concentration is used for the x-axis of the drug combination plot. The plot allows a
50
determination to be made of the slope (m), a measure of the sigmoidicity of the dose effect
relationship, and the x-intercept (Dm), the median effect dose.
Mathematical linearization and definition of efficacy parameters then allowed the
generation of a combination index as previously described [315]. The dose of the individual
drugs and both drugs together that affect a given percent (x%) of the plated colonies, Dx1, Dx2,
Dx1,2 respectively was calculated from Eq. 1 of reference [315].
m
a
amx
f
fDD
1
1
Based on the above equation, the combination index (CI) for quantification of synergism or
antagonism for the two drugs was determined.
A CI < 1, =1 and > 1 indicates synergism, additive effect and antagonism, respectively. The
dose-reduction index (DRI) provides a measure of how much dose of each drug is reduced in
synergistic combination at a given effect level compared with the doses of the individual drug.
Based on the above CI equation, (DRI)1= (Dx)1/D1 and (DRI)2= (Dx)2/D2
3.7 Fluorescence microscopy of cellular PLN uptake
For studies of cellular uptake and intracellular localization, fluorescent PLN were formulated by
encapsulating a fluorescent dye, nile-red, rather than utilizing the fluorescence of Dox to avoid
interference of released Dox with the detection of the nanocarriers. Briefly, 50mg of myristic
21
21
2
2
1
1
xxxx DD
DD
D
D
D
DCI
51
acid, 4mg of PEG100SA and 8mg of PEG40SA was melted. 25µL of nile-red was added to the
solution and stirred until it melted. Fifty µL of 100 g/L of PF68 and 388 µL of water were added
to the solution. The solution was stirred for 20 minutes and then ultrasonicated using a Hielscher
UP100H probe ultrasonicator (Hielscher USA, Inc., Ringwood NJ, USA) at 80% peak amplitude
and 5 mm probe depth in solution for 5 minutes. The entire emulsion was immediately
transferred into 5 mL of distilled deionized water.
MCF7 WT, MCF7 VP and MCF7 MX cells were seeded overnight in a 6cm petri dish with 5ml
of growth medium at 37°C in a 5% CO2 atmosphere. 100µL of fluorescent PLN was added to the
dish for 1hour. Cells were washed with warmed growth medium to remove free PLN. Following
washing, nuclei were stained with 0.5µg/ml Hoescht 33342 for 10 min at 37°C and imaged using
Zeiss LSM510 deconvolution fluorescence microscope (Carl Zeiss Canada, Ltd., Toronto, ON,
Canada). The images were acquired using 4’,6-diamidino-2-phenylindole (DAPI) filter to
visualize DAPI and red fluorescence protein (RFP) filter to visualize nile-red loaded PLN and
analyzed using AxioVision software(Carl Zeiss Canada, Ltd., Toronto, ON, Canada).
3.8 Statistical analysis
Data are presented as the mean ± standard error of mean (SEM) for results obtained from three
independent trials unless otherwise indicated. Statistical significance between two groups was
tested with Student’s t-test in MS Excel.
52
4 Results
4.1 Properties of PLN
The particle size (average diameter) zeta potential, polydispersity index, encapsulation
efficiencies, and drug loading of the PLN were measured and are presented in Table 2.2. A
typical particle size distribution plot determined by the dynamic light scattering method and
transmission electron microscopic (TEM) photograph are shown in Fig. 2.1. The PLN with
single or dual agents are similar in particle size, surface charge with average diameters around
160nm and zeta potential from -18.0 to -20.4mV. The TEM micrographs show a similar
morphology (spherical shape) and particle size for all PLN formulations. The polydispersity
index ranged from 0.36-0.40 for single and dual agent PLN. As shown in Table 2.2, the
encapsulation efficiency can be reached as high as >90% for the water-soluble drug, Dox. The
high encapsulation efficiency of Dox stems from the complexation of cationic Dox HCl with the
anionic HPESO polymer, which enhances partition of the drug into the lipid phase and thus
results in high drug loading efficiency [193, 277, 278, 309, 310].
Figure 2.1. A typical particle size and size distribution plot of PLN determined by dynamic light
scattering method (left) and TEM photograph of PLN (right). The particles have average
diameters of about 160 nm (Table 3) and spherical shape.
100nm
53
Table 2.2. Particle size, zeta potential, polydispersity index, drug loading efficiency and loading
levels of the drug-loaded PLN formulations
PLN
formulation
Average
diameter
(nm)
Zeta
potential
(mV)
Polydispersity
Index
Encapsulation
efficiency (%)
Drug
loading
(w/w %)
5mg Dox 160 (4) -20.4
(0.3)
0.36 (0.04) 90.1 (3.6) 9.0 (0.3)
4mg MMC 158 (4) -18.0
(0.2)
0.40 (0.03) 73.6 (4.1) 7.2 (0.6)
Drug
combination
( 5mg Dox
+ 4 mg MMC)
162 (6) -18.7
(0.7)
0.38 (0.03) 92.5 (2.4)
37.8 (4.6)
9.2 (0.9)
3.0 (0.4)
Particle diameter refers to the number-weighted diameter of readings averaged over 5min.
Polydispersity Index was calculated by dividing the standard deviation of the Gaussian
distribution by the mean diameter. Encapsulation efficiency is the % of drug added initially that
was incorporated into the PLN. Drug loading is the % of drug comprising the total PLN mass.
All values are the mean (standard error of mean) of three independent trials.
4.2 Dose-response of MCF human breast cancer cells treated with Dox and MMC
In an effort to evaluate the toxicity of Dox and MMC towards breast cancer cells, the
cells were exposed to these anti-cancer agents for 1 hour and their ability to form colonies was
evaluated. Figs. 2.2A-C compare the cytotoxicity of Dox and MMC towards MCF7 WT, MCF7
VP and MCF7 MX cells respectively. Dox and MMC exhibit a similar degree of cytotoxicity
against MCF7 WT breast cancer cells at equimolar concentrations up to 5µM. However, lower
cytotoxicity (high survival fraction) was observed in MCF7 VP and MCF7 MX cells especially
when treated with Dox ascribed to the fact that Dox is a substrate of MRP1 and BCRP efflux
pumps [281, 294]. At the same doses, the survival fraction of MCF7 MX cells treated with MMC
is greater than MCF VP cells, suggesting that this BCRP+ cell line is more resistant to MMC
54
treatment than the MRP1+ cells, although MMC is reported to be a non-substrate of BCRP [281,
294].
Figure 2.2. Surviving fraction, measured by a clonogenic assay, of (A) MCF7 WT, (B) MCF7
VP (MRP1+) and (C) MCF7 MX (BCRP+) cells after exposure to increasing concentrations of
Dox or MMC for 1 hour. Error bars represent S.E.M. In some cases the error bars are smaller
than data points.
The dose-response relationship of studied cell lines was further analyzed by the median
effect analysis (Fig. 2.3). From the plots of log[(fa)-1
-1]-1
vs. log[D] (Figs. 2.3 D-F), the median
effect doses (Dm) of Dox required for 50% cell kill were determined by taking the anti-log of the
x-intercept and found to be 0.81µM for MCF7 WT cells, 5.1µM for MC7 VP, and 6.5µM for
MCF7 MX cells respectively, which increased up to 8-fold comparing WT to MX cells (Table
2.3). In contrast, the Dm of MMC only slightly increased from 0.84 for MCF7 WT to 1.2 µM for
MCF7 VP cells (Table 2.3).
55
Figure 2.3. Percent kill of cell ability to expand clonogenically vs. drug dose exposure for 1 hr
to Dox or MMC alone or in combination (dose-effect curves) for (A) MCF7 WT, (B) MCF7 VP
and (C) MCF7 MX cells. Median effect plots for the interaction of MMC and Dox in (D) MCF7
WT, (E) MCF7 VP and (F) MCF7 MX cells following 1 hr of drug exposure. Cells were treated
with Dox (circles) or MMC (squares) or Dox and MMC (Dox-MMC) in a 2:1 molar ratio
(triangles). The doses for the Dox-MMC combination treatment are the Dox doses. Error bars
represent S.E.M. In some cases the error bars are smaller than data points.
A
A
D
A
B
A
C F
A
E
56
Table 2.3. Dose-effect relationship parameters for Dox and MMC in MCF7 WT, MCF7 MX
(BCRP+), and MCF7 VP (MRP1+) human breast cancer cell lines
Cell Type Dox MMC Dox+MMC DRI
Dm (µM) m r Dm (µM) m r Dm
(µM)
m r Dox MMC
MCF 7 WT 0.81 2.3 0.95 0.84 2.8 0.94 0.40 3.0 0.90 2.05 2.1
MCF 7 MX 6.5 1.1 0.97 1.03 1.6 0.91 0.68 1.8 0.94 9.5 2.4
MCF 7 VP 5.1 2.3 0.98 1.2 3.0 0.97 0.91 3.9 0.91 5.6 1.3
Potency, shape (sigmoidicity) and conformity of dose effect curve (linear correlation coefficient)
are represented by Dm, m and r respectively, where Dm is antilog of x-intercept in µM, m is the
slope of the median-effect plot and r is the linear correlation coefficient of the median effect plot.
Dose reduction indices for Dox-MMC combination (DRI) were determined by (DRI)Dox =
(Dm)Dox /(Dm)Dox-MMC and (DRI)MMC = (Dm)MMC /(Dm)Dox-MMC (see material and methods).
The slope m in the log[(fa)-1
-1]-1
vs. log[D] plot is a measure of the shape of dose-effect
curve, with m=1, >1 and <1 indicating hyperbolic, sigmoidal and negative sigmoidal curve,
respectively. The m values for all treatment groups were determined from the slopes of log[(fa)-1
-1]-1
vs. log[D] plots in Fig. 2.3D-F and are shown in Table 2.3. The m values for all cell lines are
greater than unity indicating a sigmoidal dose-effect relationship for all these cell lines. Since a
larger m value indicates a more sensitive dose response, the lower m values for MCF7 MX cells
treated with Dox or MMC suggest that this BCRP+ cell line is less sensitive to small changes in
drug concentrations than the MCF7 WT and MCF7 VP cells.
57
4.3 Synergistic effect of Dox and MMC in MCF7 human breast cancer cells
Simultaneous administration of 2:1 (MMC:Dox) molar ratio significantly enhanced the
cell kill over individual drug treatments in all three cell lines (Figs. 2.3A-C). It was observed that
over 90% of the cells were killed at low doses of the combination treatment for all three cell
lines, which was not achieved with single agent treatment. For example, treatment of MCF7 WT
cells with 2 µM of Dox and MMC together resulted in near complete cell death (99.09 ± 0.03%,
P<0.001) as compared to Dox and MMC treatment alone. Various parameters were obtained
from the median effect plot Fig. 2.3D-F, where fa is the fraction of cells unable to form colonies
and D is the concentration of drug (Dox only in Dox plus MMC plots). The linear correlation
coefficient (R2) indicates the conformity of the data to the median effect plot of the mass- action
law. As seen in Fig. 2.3D-F and Table 3 all treatments have R2 >0.90, indicating the validity of
the analysis.
The dose reduction index (DRI) for the Dox-MMC combination treatment relative to the single
agent treatment were calculated by dividing Dm of single agent by Dm of dual agents. As shown
in Table 2.3, the DRI for MCF7 WT cells is similar with a dose reduction about 2-fold, which is
comparable to previous findings by Shuhendler et al. in MDA MB 435/LCC6/WT breast cancer
cells [193]. However, for the MDR cells the DRI for Dox treatment is much larger with DRI =
5.6 in MCF7 MX (BCRP+) cells and 9.5 in MCF7 VP (MRP1+) cells. In contrast, the DRI for
MMC ranges from 1.3 to 2.4. The results indicate that the stronger the drug resistance of the
cells, the greater is the dose reduction.
58
The combination index allows the quantitative determination of drug interactions, where
CI < 1, = 1 and >1 indicate synergism, additive effect and antagonistic effect, respectively. With
the knowledge of m and Dm values for each drug and the combination at a constant molar ratio,
the combination index for a series of values of fa (Figs. 2.3A-C) were calculated and are plotted
in Figs. 2.4A-C. The CI values are less than unity over the entire range of fa values for all three
cell lines, suggesting a strong synergistic interaction of MMC and Dox at a 2:1 molar ratio.
Figure 2.4. Combination Index analysis of the interaction of Dox and MMC in (A) MCF7 WT,
(B) MCF7 VP and (C) MCF7 MX cells following treatment for 1 hour. All the curves drop
below unity indicating Dox and MMC exhibit a synergistic effect against all studied cell lines.
C
A B
59
As occurs in synergistic interactions, the DRI for all studied cell lines is >1 (Table 2.3).
Also shown in Table 3 the m value increases for the combination treatment in all cell lines. The
combination treatment produced steeper dose-effect curves than single treatments indicating that
small changes in doses will produce greater effects, i.e. increased growth inhibition.
4.4 PLN formulations are more effective than free drugs against MCF7 cancer cells
The efficacy of PLN loaded with Dox, MMC or the Dox-MMC combination was
assessed against wild type and resistant MCF 7 breast cancer cell lines using the clonogenic
assay. PLN were prepared containing a low (0.3µM) and a high (1.2µM) drug level based on
previous work [193]. The dose of dual agent PLN applied was calculated based on Dox loading
efficiency. The doses (0.3µM and 1.2µM) applied in PLN were much lower compared to the
concentration of free drug used (Figs. 2.2A-C). Encapsulation of chemotherapeutic agents
resulted in significant increases in cell kill compared to the unencapsulated drugs at equivalent
doses (Fig. 2.5). For example, 75 ± 11% of MCF7 WT cells were killed when 0.3µM of Dox was
delivered via PLN while only 30 ±6.5% of cells was killed when Dox was used as free agent.
Similar results were also observed with MMC – 87 ±1% cell kill for PLN versus 34±3% cell kill
when MMC was given as free agent at 0.3µM. The effect of PLN formulation was highly
significant in the resistant cell lines. The surviving fraction of MCF7 VP cells decreased from
0.96±0.13 for free Dox to 0.51±0.04 for Dox-PLN. Encapsulation of MMC showed similar
results where the surviving fraction decreased to 0.48±0.01 from 0.69±0.14 when delivered as
free agent at low concentration. MCF7 MX cells also showed a significant decrease (p<0.001) in
survival when Dox and MMC were delivered via PLN (Fig. 2.5C).
60
A dose dependent increase in cell kill was observed for all drug loaded PLN formulations
in all three cell lines. The wild type cell line showed a higher cell kill for any given dose (0.3µM
and 1.2µM) compared to either of the resistant cell lines. Co-encapsulation of both Dox and
MMC in a single PLN generated a significantly higher cell kill (p< 0.001) in both sensitive and
resistant cell line (Table 2.4). High doses of Dox-MMC encapsulated in PLN resulted in cell kills
of 99.2 ± 0.4%, 94.7 ± 1%, 98.2 ± 0.6% for MCF7 WT, MCF7 VP and MCF7 MX cells
respectively (Figs. 2.5A-C). The cytotoxicity enhancement ratio (CER) was calculated and
presented in Table 4. For the resistant cell lines (MCF7 VP and MCF7 MX), Dox and MMC
PLN resulted in more than 20 times cell kill (CERDox > 20) compared to the free Dox solution at
0.3µM. A high dose of 1.2 µM resulted in over 17 times cell kill in MCF7 VP cell line and 9
times in MCF7 MX cell line when compared to the free Dox solution (Table 4). However, only
up to a 3 fold higher cell kill (CERMMC) was observed using Dox-MMC PLN as compared to the
free MMC solution at 0.3µM (Table 2.4).
61
Figure 2.5. Comparison of anti-cancer efficacy of single agent Dox or MMC free in solutions or
in PLN with dual agent PLN formulation in (A) MCF7 WT, (B) MCF7 VP (MRP1+) and (C)
MCF7 MX (BCRP+) cells. Cells were treated with various formulations at the equivalent doses
for 1 hour and assessed using a clonogenic assay. The data represent the mean ± SEM of three
independent trials. *Statistically significant decrease in cell survival relative to free agent
treatment (P < 0.05)
*
* *
* *
*
* * * * *
*
* * *
*
*
*
C
B
A
62
Table 2.4. Cytotoxicity of Dox solution and dual drug (Dox and MMC) loaded PLN (DM-PLN)
against wild type (MCF7), BCRP+ (MCF7 MX) and MRP1+ (MCF7 VP) human breast cancer
cells.
Cell
Type
Free Dox
(% Kill)
Free MMC
(% Kill)
DM-PLN
(%Kill)
Cytotoxicity
enhancement
ratio (CERDox)b
Cytotoxicity
enhancement
ratio
(CERMMC)c
0.3µM 1.2µM 0.3µM
1.2µM Low
a High
a Low High Low High
MCF7
WT
30.5
±6.7
52.1
±3.2
33.6
±3.2
64.7
±1.6
97.7
±0.8
99.2
±0.4
3.2 1.9 2.9 1.5
MCF7
MX
4.20
±0.01
10.9
±0.2
29.7
±0.005
59.6
±0.4
89.3
±3.3
98.2
±0.6
21.2
9.0
3.0
1.6
MCF7
VP
3.40
±0.12
5.40
±0.002
30.6
±0.2
50.3
±0.02
76.6
±2.0
94.7
±1.0
22.4
17.5
2.5
1.9
a Low dose: 0.3µM Dox, and High dose: 1.2µM Dox in the dual agent loaded PLN (DM-PLN) at
a 2:1 Dox;MMC molar ratio.
b CERDox = %Kill (DM-PLN) / % Kill (Free Dox)
c CERMMC = %Kill (DM-PLN) / % Kill (Free MMC)
4.5 Cellular uptake and intracellular localization of PLN
Fluorescent PLN were formulated by encapsulating nile-red within PLN. Following 1
hour of incubation with nile-red loaded PLN; cellular uptake of PLN was examined using
fluorescence microscopy (Fig. 2.6). Strong fluorescent signals of PLN appeared inside the cell
63
and near the perinuclear region in both wild type and resistant cell lines. Shuhendler et al. have
demonstrated the uptake of PLN via an endocytosis mechanism, allowing it to overcome
membrane efflux pumps and deliver Dox and MMC to the perinuclear region [193].
Figure 2.6. Intracellular localization of fluorescent PLN in breast cancer cell lines MCF7 WT,
MCF7VP and MCF7MX. Cells were incubated with nile red-loaded PLNs (red) for 1 hour and
visualized with a RFP filter (PLNs). Nuclei were stained with Hoescht 3342 (blue) and
visualized with a DAPI filter (Nucleus) and PLN/Nucleus images overlayed (Merge). Images
were acquired with a 20× objective lens.
5 Discussion
Drug combination therapy has been investigated to increase the effectiveness of drugs
while decreasing systemic toxicity by dose reduction. In this study, the median effect analysis
64
and combination index equation of Chou and Talalay [315, 316] were applied to quantify the
potency of Dox and MMC, their combination and the shapes of dose-effect curves, as well as
their synergism at different concentrations. The usefulness of this approach has been
demonstrated in analysis of combinations of other anticancer agents [317, 318]. A synergistic
growth inhibition was found in all three cell lines (MCF7 WT, MCF7 VP and MCF7 MX)
following treatment with a 2:1 molar combination of Dox and MMC. Synergistic interaction
between Dox and MMC was also observed previously in murine breast cancer EMT6/AR1.0
cells and P-gp/MDR1 overexpressing MDA435/LCC6/MDR1 [191, 193]. Together all results
indicate that the Dox-MMC synergistic effect is not cell line specific. This synergistic interaction
resulted in a dose reduction for a given degree of effect which could lead to a reduction in
toxicity to normal tissue if applied in vivo.
The PLN formulation enhanced the efficacy of Dox, MMC and their combination against
all studied cell lines. The PLN loaded with anticancer agents were highly efficient in overcoming
MRP and BCRP efflux pumps with up to 22-fold cytotoxicity enhancement ratio (Table 2.4) as
compared to the free drugs at a low dose of 0.3M. The co-encapsulation of the cancer drugs
inside PLN, allowed Dox and MMC to be delivered simultaneously to the perinuclear region of
the cells. Simultaneous delivery of Dox and MMC is important as it is postulated that they
produce synergism by induction of DNA double strand breaks via covalent topoisomerase IIα-
Dox-DNA complex [191, 193]. Even at 1.2µM, a much lower dose than the free Dox, the
combination treatment of DM-PLN reached 99% cell kill. This low dose could significantly
reduce systemic toxicity and yet achieve higher tumor cell kill. PLN uptake by endocytosis
protected the drugs from access of the membrane transporters, i.e., MRP1 and BCRP in this
65
work and P-gp in our previous work [193] hence preventing efflux of Dox and MMC. The
nanoparticles were internalized into the cells, not just adhering to the cell membrane surface,
allowing the cargo to be delivered near its intended site of action (i.e., DNA). This explains why
the PLN formulation of Dox-MMC combination is more efficacious than the free drug
combination.
Despite the development of more potent and more specific transporter inhibitors [294-
296], clinical trials of these agents have led to unsatisfactory outcomes as many of them are also
substrates for other enzymes and transporter systems, resulting in unpredictable
pharmacokinetics. In addition these MDR reversing agents are unable to inhibit other ABC
transporters co-existing in a solid tumor, or being upregulated due to inhibition of another
transporter or exposure to a different chemotherapeutic agent. This study has shown that the
same PLN carrier system is able to overcome two membrane efflux transporters MRP1 and
BCRP and our previous work has demonstrated its capability of circumventing P-gp efflux pump
[193, 277, 278]. Since P-gp, MRP1 and BCRP are all implicated in drug resistance of cancer
cells to a variety of chemotherapeutic agents [272, 294], the results of this study and previous
studies suggest that use of nanoparticulate drug combination formulations may be a more
beneficial therapeutic approach for overcoming multiple membrane efflux pumps in MDR
cancer cells than inhibiting a single type of efflux pump.
6 Conclusion
In conclusion, the results of this study suggest a beneficial therapeutic strategy for
overcoming multiple membrane efflux pumps which are one of the most frequently occurring
66
causes of drug resistance in cancer therapy. The co-encapsulation of Dox and MMC in PLN
showed a higher cytotoxic effect due to the synergy of the two agents and the advantages of the
nanoparticle system. As overcoming MDR is clinically important, these results suggest a
promising therapeutic strategy to improve chemotherapeutic efficacy and decrease systemic
toxicity.
7 Acknowledgements
This work was funded by the Canadian Institutes of Health Research and Canadian Breast
Cancer Research Alliance. The University of Toronto Fellowship to P.P. and Scholarship from
the National Science and Engineering Research Council of Canada and the Ben Cohen Fund to
AJS are also acknowledged.
67
Chapter 3 Doxorubicin and mitomycin C co-loaded polymer-lipid
hybrid nanoparticles inhibit growth of sensitive and multidrug
resistant human mammary tumor xenografts
Preethy Prasad1, Ping Cai1, Adam Shuhendler1, Andrew M. Rauth2, Xiao Yu Wu1
1Department of Pharmaceutical Sciences, Leslie Dan Faculty of Pharmacy, University of Toronto, 144 College Street, Toronto, Ontario, Canada, M5S 3M2
2Division of Experimental Therapeutics, Ontario Cancer Institute, 610 University Ave, Toronto, Ontario, Canada M5G 2M9
This work has been published in Cancer Letters, 2013, 334(2):263-73.
All work in this manuscript was performed by P.Prasad with assistance from the co-authors,
except for the paraffin embedding, sectioning, and staining of the histological samples
68
1 Abstract
Multidrug resistance (MDR) and drug toxicity are two major factors responsible for the failure of
cancer chemotherapy. Herein the efficacy and safety of combination therapy using doxorubicin
(Dox, D)–mitomycin C (MMC, M) co-loaded stealth polymer-lipid hybrid nanoparticles
(DMsPLN) were evaluated in sensitive and MDR human mammary tumor xenografts. DMsPLN
demonstrated enhanced efficacy compared to liposomal Dox (PLD) with up to a 3-fold increase
in animal life span, a 10-20% tumor cure rate, undetectable normal tissue toxicity and decreased
tumor angiogenesis. These results suggest DMsPLN have potential as an effective treatment of
breast cancer.
Keywords
Doxorubicin, mitomycin C, nanoparticles, efficacy, toxicity, anti-angiogenesis
69
2 Introduction
Breast cancer remains one of the leading causes of cancer death in women with
approximately 1.38 million new diagnoses and 458,400 deaths in 2008 worldwide [319]. Despite
advances in treatment and early diagnosis, about 20-30% of all treated patients eventually
undergo relapse, of which most cases are metastatic [145, 320]. For the successful management
of breast cancer, chemotherapy is often employed to complement surgery and radiation therapy,
particularly when the cancer cells have spread or are suspected of spreading from the primary
tumor site to other parts of the body [238].
Doxorubicin (Dox) is one of the most effective chemotherapeutic anthracycline agents. It
is widely employed alone or often in combination with other agents for adjuvant breast cancer
chemotherapy [238]. Dox is highly effective in oxygenated regions of the tumor, exerting its
cytotoxic effects through DNA intercalation, topoisomerase II inhibition, prevention of DNA and
RNA synthesis, and generation of reactive oxygen species [321-323]. Nevertheless, its
application is associated with severe adverse effects, including myelosuppression, cardiotoxicity
and palmar plantar erythrodysenthesia (PPE), which lead to a very narrow therapeutic window
[238, 324, 325]. Moreover, its anticancer efficacy is limited by elements of the tumor
microenvironment, such as hypoxia, acidity, and defect vasculature and lymphatic vessels [218,
326], as well as multidrug resistance (MDR) of cancer cells [218, 272, 282].
MDR can be inherent or acquired following chemotherapy. It is complex and
multifactorial affording cancer cells many escape routes from chemotherapy. The most frequent
causes of MDR include the up-regulation of membrane bound ATP-binding cassette (ABC)
70
efflux transporters such as P-glycoprotein (P-gp), multidrug-resistance associated protein
(MRP1) and breast cancer resistance protein (BCRP), which have been identified in cancer cells.
These transporters increase the ability of cancer cells to actively transport anti-cancer agents,
such as Dox, out of the cells against concentration gradients, causing a reduction in drug
cytotoxicity [196, 218, 272, 282, 285].
Nanoparticle formulations have been shown to overcome multiple membrane efflux
transporter-mediated MDR by entering the cells via endocytosis and releasing therapeutic agents
inside the cells [193, 272, 277, 278, 310]. A polymer-lipid hybrid nanoparticle (PLN) system
developed in our laboratory is able to load hydrophobic and hydrophilic drugs with high
efficiency and good release kinetics [193, 277, 310]. Hydrolyzed polymers of epoxidized
soybean oil (HPESO), derived from a naturally occurring renewable source of soybean oil was
used to develop the PLN due to its amphiphilic properties brought by the long fatty chains, ether
bonds and carboxylic groups [327] and absence of cytotoxicity [278]. The PLNs with co-loaded
Dox and GG918 (a P-gp inhibitor) or Dox and mitomycin C (MMC) exhibited much greater
cytotoxicity than the free drugs against MDR breast cancer cells that overexpress P-gp, MRP1 or
BCRP [193, 277, 278, 328]. We have shown that the Dox-MMC combination can generate a
synergistic effect on both sensitive and MDR breast cancer cells in vitro [191] and that Dox (D)
and MMC (M) co-loaded stealth PLNs (DMsPLN) can further enhance their synergy at
significantly reduced dose [280, 328]. The transport of Dox-MMC to the perinuclear region of
the cancer cells by the PLNs and Dox-MMC enhanced DNA double strand breaks are believed to
contribute to the in vitro synergistic cytotoxicity [191, 280]. The efficacy of Dox-MMC
combination has also been demonstrated in a murine mammary mouse breast tumor model
71
resulting in a 185% delay in tumor growth when delivered by microspheres intratumorally [192].
Part of the enhanced in vivo therapeutic efficacy could be due to the higher toxicity of MMC in
the hypoxic environment of solid tumors [329]. Because of its severe toxicity, in particular its
myelosuppression effect, MMC has not been used widely as a first line chemotherapeutic agent
[330, 331]. Nevertheless, the interests in MMC combination with other anticancer drugs for the
treatments of unresectable solid tumors have been renewed and many clinical trials have been
pursued [332]. The unique bioreductive mechanism of MMC activation and the in vitro synergy
of Dox-MMC combination in PLNs warrant further investigation of DMsPLN for the treatment
of MDR tumors.
Solid tumors are known to possess “leaky” tumor neovasculature and malfunctioning
tumor lymphatics [218, 333], which enables accumulation of nanoparticulate therapeutics in
tumor tissue by passive targeting via the enhanced permeability and retention (EPR) effect [334].
This effect has been utilized to deliver a variety of anticancer drugs to tumor by nanocarriers
such as liposomes, soluble polymers and polymer micelles [251, 335-337]. Some of the
nanoparticle formulations have been approved for clinical use, e.g. the stealth doxorubicin
liposomal (PLD) formulation known as Doxil® or Caleyx®. The PLD formulation has shown
reduced cardiomyopathy and myelosuppression compared with free Dox in the treatment of
various cancers. This has been attributed to polyethylene glycolylation (PEGylation) that alters
the pharmacokinetic profile of Dox resulting in a higher drug concentration in the tumor and
decreased volume of distribution [338-341]. However, the enhancement in the therapeutic
efficacy is insignificant and PPE occurs in at least 45% of patients treated with PLD [342-344],
suggesting that further development of nanocarrier systems is necessary.
72
Given the superior in vitro efficacy of DMsPLN in overcoming MDR in cancer cells and
excellent systemic circulation, tumor accumulation and reduced liver uptake in vivo, as
demonstrated by whole body and microscopic imaging of mice bearing orthotropic human breast
tumors [345, 346], the therapeutic efficacy and safety of DMsPLN in sensitive and MDR human
breast tumor xenografts was evaluated in this work. Tumor size as a function of time was used to
determine the efficacy of therapy and changes in mouse body weight and damage-associated
blood enzymes were measured to assess the toxicity of the treatment. The potential of DMsPLN
to inhibit angiogenesis in solid tumors was also investigated as free Dox was reported to exhibit
anti-angiogenic effects [347]. The United States Federal Drug Agency (FDA) approved PLD
formulation (Caleyx®) was used as a comparator in this work as it has been employed clinically
[338-344].
3 Materials and methods
3.1 Chemicals and reagents
Myristic acid, poly(ethylene glycol)-100-stearate (PEG100SA), poly(ethylene glycol)-40-
stearate (PEG40SA) and all other chemicals, unless otherwise mentioned, were purchased from
Sigma-Aldrich Canada (Oakville, ON, Canada). Mitomycin C and doxorubicin were purchased
from Polymed Therapeutics (Houston, TX, USA). PEGylated liposomal doxorubicin (PLD)
(Caelyx©) was purchased from the Pharmacy at the Princess Margaret Hospital (Toronto,
Ontario, Canada). HPESO was a gift from Drs. Z. Liu and S. Erhan (Food and Drug
Administration, Washington, DC, USA). Pluronic F68 (PF68) (nonionic block copolymer) was a
kind gift from BASF Corp. (Florham Park, NJ, USA). All cell culture plastic ware was
purchased from Sarstedt (Montreal, QC, Canada). Cell culture medium, Dulbecco’s Modified
73
Eagle Medium (DMEM), and phosphate buffered saline (PBS) were obtained from Tissue
Culture Media Facility, Ontario Cancer Institute (Toronto, ON, Canada). Fetal bovine serum
(FBS) and trypsin were purchased from Invitrogen, Inc. (Burlington, ON, Canada).
3.2 Preparation and characterization of stealth polymer lipid hybrid Nanoparticles
This nanoparticle system has been completely characterized and described in our previous
work [193, 328]. Briefly, a mixture of 50 mg of myristic acid, 4 mg of PEG100SA and 8 mg of
PEG40SA was melted in a 15 ml conical tube at 65°C. Once the fatty acid was melted, 4 mg of
MMC powder was added. This was followed by the addition of 50 μL of a 50 g/L solution of
HPESO and 400 μL of a 12.5 mg/mL solution of Dox. In addition 50 μL of a 100 g/L solution of
PF68 was also added. The solution was stirred for 20 minutes and then ultrasonicated using a
Hielscher UP100H probe ultrasonicator (Hielscher USA, Inc. Ringwood NJ, USA) at 80% peak
amplitude and 5 mm probe depth in the solution for 5 minutes. The entire emulsion was
immediately transferred into 5 mL of sterile 0.9% NaCl and stirred on ice. Particle size and zeta
potential were measured by dynamic light scattering and electrophoretic mobility, respectively,
using a NICOMPTM
380ZLS (PSSNICOMP, Santa Barbara, CA, USA) apparatus. Immediately
after formulation, the PLN suspension was centrifuged through a 0.1µm filter and the drug in the
filtrate was assayed spectrophotometrically at 540 nm for Dox and 364 nm for MMC. Drug
loading (%wt drug/wt lipid) and encapsulation efficiency (%wt loaded drug/wt total drug) were
then calculated. The remaining PLN were incubated with Sephadex SP C-25 anionic dextran
microspheres at 4oC overnight to remove any unencapsulated drug. The suspension was
74
centrifuged to remove the Sephadex SP C-25 microspheres. DMsPLN were made fresh before
each injection.
3.3 Cell culture
The human breast cancer cell line, MDA-MB 435/LCC6/WT and the P-gp over expressing
MDA-MB 435/LCC6/MDR1 were kindly provided by Dr. Robert Clarke (Georgetown
University, Washington, DC, U.S.A.). Both MDA-MB 435/LCC6 cell lines were incubated at
37°C in a humidified incubator with 5% CO2 and maintained in pH 7.2 alpha modified minimal
essential medium (Ontario Cancer Institute Media Laboratory, Toronto, Ontario, Canada)
supplemented with 10% fetal bovine serum. Cell doubling times were typically 24h. Cells were
trypsinized and subcultured at 50-fold dilution once they were confluent. MDA-MB 435 cells
were tested for their suitability for use in animals and were found to be pathogen-free (Research
Animal Diagnostics Laboratory, Columbia, MO, USA)
3.4 Orthotopic Model Development and Treatments
All experiments and procedures used in the animal studies were approved by the Animal
Care Committee at the Ontario Cancer Institute (protocol # 1844). Eight-ten weeks old female
nu/nu mice were purchased from Taconic Farm Inc (Hudson, NY, USA). Solid tumors of MDA-
MB 435/LCC6/ WT and MDR breast cancer cells were grown orthotopically in mouse mammary
fat pads. In brief, the cells from frozen storage were cultured and passed at least 3 times,
followed by injection of 1 million cells in growth medium into the inguinal mammary fat pad of
each mouse. Animals had free access to food (Irradiated Tecklad LM485, Harland Tecklad,
Indianapolis, IN, USA) and sterile water. Mice were kept in Allentown ventilated microisolator
75
cages (Allentown Inc. Allentown, NJ, USA) with each cage bearing 5 mice. An initial test was
conducted using a clinically used dose (50 mg/m2) [338] to determine a tolerable Dox dose for a
complete therapeutic study. For this test, tumor-bearing mice were randomly allocated to three
different treatment groups (five mice per group): 1) control (saline), 2) PLD, 3) DMsPLN and
received an equivalent Dox dose of 50 mg/m2 via tail vein injection. For the therapeutic study,
tumor bearing mice were randomly allocated to four different treatment groups: 1) control
(saline), 2) PLD, 3) DMsPLN, 4) DMsPLN 4× (given 4 times every 4 days). Each treatment was
given equivalent to a free Dox dose of 25 mg/m2 administered by tail vein injection with a
maximum volume of any one injection of 200 μl. For the DMsPLN treatment group, additional
MMC dose of 8 mg/m2 was administered together with Dox in the formulation. Each group
contained 4-5 mice and the experiments were repeated twice. The results from the two
experiments were combined. Treatment was initiated when the tumors reached a size of 50 mm3.
Once tumors reached 300 mm3 in size or once the mice exhibited signs of discomfort (i.e. weight
loss, lack of grooming, signs of self mutilation, resistance to ambulation), mice were euthanized
by cervical dislocation under 1% isofluorane anaesthesia.
3.5 Evaluation of therapeutic efficacy
The tumor size was measured as a function of time twice weekly with vernier calipers in
two dimensions. Tumor volumes were calculated by the formula V= [(length) × (width)2
]/2,
where length is the longest diameter and width is the shortest diameter perpendicular to length
[348]. At the end of experiment, the animals were sacrificed and the tumor masses were excised
and stored in 10% buffered formalin for histological examination. Average time of tumor growth
was calculated by determining the average time required for tumor to reach the endpoint, defined
76
as the survival time. Tumor growth delay (TGD) was calculated from the mean survival time of
each group according to TGD (%) = (Tcontrol - TRx)/Tcontrol x100% [277], where the subscripts
control and Rx indicate the control and treatment group, respectively. The normalized tumor size
(V̂ ) and percentage tumor growth inhibition (TGI %) at specific days were computed as follows:
0/ˆtt VVV
%100ˆ/)ˆˆ(% controlttRxcontrolt VVVTGI
Where 0tV is the average tumor volume of a treatment group at the time when the initial
treatment was given; tV is the average tumor volume of the same group at a specific day other
than the day of initial treatment; controltV̂ and
RxtV̂ are respectively normalized tumor volume of
control (saline) group and a therapeutic (i.e., PLD, DMsPLN or DMsPLN 4×) group at time t.
3.6 Determination of median survival time and percentage increase in life span
Animal survival time was monitored and the median survival time (MST) and percentage
increase in life span (ILS %) were calculated for the mice treated with PLD, DMsPLN and
DMsPLN 4× using the following formula:
MST = (day of first death + day of last death)/2
%100)1/(% controlRx MSTMSTILS
77
3.7 Evaluation of safety and normal tissue toxicity
For safety and toxicity evaluation of the saline and nanoparticulate formulations, the body
weight of each mouse was measured every other day following treatment and was related to the
value on the first day as percent change in body weight. Body weight loss greater than 20% was
regarded as a sign of extensive systemic toxicity and the mice were sacrificed for humane
reasons rather than allowing them to die of cancer or drug toxicity. In addition, blood samples
were withdrawn via the saphenous vein every 7 days for up to five weeks after treatment. Blood
was centrifuged at 1,400 g for 20 minutes at 4oC to isolate plasma, which was immediately flash
frozen in liquid nitrogen until processing. Commercially available kits and their corresponding
methods were used to assay plasma for lactate dehydrogenase (LDH) (Cayman Chemical Co.,
Ann Arbor, MI, USA), alanine transaminase (ALT) (Cayman Chemical Co.), and creatine kinase
(CK) (BioAssay Systems, Hayward, CA, USA).
3.8 CD31 expression and assessment of microvessel density of tumors (MVD)
Tumor specimens to be evaluated for MVD were harvested and fixed in 10% buffered
formalin for 24 h before being transferred to 70% ethanol. The specimens were subsequently
paraffin-embedded and sectioned. Tumor microvessels were visualized using
immunohistochemical detection of CD31 using a 1:50 dilution of anti-CD31 antibody (abcam,
CA) and visualized using biotinylated goat anti-rabbit antibody (1:500 dilution) (Vector labs,
CA). All of the immunostained sections were counterstained using hematoxylin. To calculate
microvessel number and MVD, four different areas of the tumor were randomly selected from
78
three different animals in the same treatment group. MVD was determined as the percent total
area within a tumor section that stained positively for CD31.
3.9 Statistical analysis and graphing
GraphPad Prism® Software was used for all graphs and statistical analysis. The statistical
significance of differences between experimental and control groups was determined using t-test.
P < 0.05 was considered significant, and significant differences are shown by asterisks in the
figures. Pairwise comparisons between survival times of each treatment group between each
trial, as well as each treatment group within each cell line were performed using the Breslow
Survival Test.
4 Results
4.1 Determination of PLD dose for the treatment
The acute toxicity of PLD and DMsPLN at an equivalent Dox dose of 50 mg/m2 was
tested in nude mice bearing MDA-MB 435/LCC6/WT tumors to determine the dose of PLD and
DMsPLN that can be administered in nude mice. This dose is employed in the clinic in the
treatment of human breast cancer [DOXIL® Monograph]. Mouse body weight was measured
every 2 days to determine the systemic toxicity of the treatment. Mice in the control (saline, Fig.
3.1A) and DMsPLN (Fig. 3.1C) groups did not show any significant weight loss. However loss
of 20% of initial body weight was observed in nude mice treated with PLD (Fig. 3.1B), which is
ruled as a toxic endpoint. Figure 1E presents typical pictures of representative mice at time of
sacrifice. Since the toxicity of PLD at this dose level was too severe, for further studies PLD and
DMsPLN were administered at an equivalent Dox dose of 25 mg/m2.
79
Figure 3.1: Percent change in body weight of MDA-MB 435/LCC6/WT tumor bearing mice
treated with (A) saline, (B) 50 mg/m2 PLD and (C) 50 mg/m
2 DMsPLN. Treatment was injected
intravenously and total body weight was generated by serial weighing over length of time after
treatment. Each curve represents one animal. (D) Percent change in body weight represented by
mean ± SEM. Termination of curves indicates sacrifice of animal due to tumor size limitation.
(E) Image of a representative mouse from mice treated with saline, PLD or DMsPLN at time of
sacrifice.
4.2 Anti-tumor efficacy of DMsPLN in sensitive and MDR tumor models
Tumors established from MDA-MB435/LCC6 cells were grown to 50 mm3 size and then
groups containing 5 mice each, were treated with one of the following 4 treatments: (1) saline,
(2) PLD (25 mg/m2), (3) DMsPLN (25 mg/m
2), or (4) DMsPLN 4× (given a dose of 25 mg/m
2, 4
80
times every 4 days). Because of the aggressive growth, the MDR tumors reached the 50 mm3
size earlier than the WT tumors (Figs. 3.2 and 3.3). Thus the mice-bearing the MDR tumors were
treated at 14 days post inoculation, while those with the WT tumors were treated at 20 days.
Tumor volume was determined at predetermined times. The results from two separate
experiments have been combined. The tumor volumes of individual mice are plotted as a
function of time up to 60 days in Figure 3.2. The numbers in the brackets indicate the number of
long term survivors that were tumor-free for more than 120 days. As it can be seen in Figs. 3.2A
and 3.2E, tumor volumes in saline treated mice increase rapidly in particular in the MDR1
tumor-bearing mice (Fig. 3.2E), with an average time of tumor growth of 38.8 and 25.4 days for
WT and MDR tumors, respectively (Table 3.1). The PLD treatment shows modest inhibitory
effect on tumor growth (Figs. 3.2B and 3.2F) with 23%TGD in WT tumor model and 30% in the
MDR tumor model (Table 3.1). The treatment with a single dose of DMsPLN (25 mg/m2)
resulted in a significant delay in the growth of tumor (Figs. 3.2C and 3.2G) with 108% and
120%TGD in WT and MDR tumors, respectively (Table 3.1). When this dose of DMsPLN was
given in four rounds, once every 4 days (DMsPLN 4×), even greater efficacy was obtained with
151%TGD (Fig. 3.2D, Table 3.1). The efficacy of DMsPLN 4× treatment in the WT and MDR
tumor model was similar to that of single DMsPLN treatment (Figs. 3.2D and 3.2H, Table 3.1).
The DMsPLN and DMsPLN 4× treatments were significantly more effective than the PLD
treatment with at least one mouse in each group showing complete disappearance of tumor.
81
Figure 3.2: Individual tumor growth curves over 60 days for mice bearing MDA-MB
435/LCC6/WT (A, B, C and D) and MDA-MB 435/LCC6/MDR1 (E, F, G and H) tumors. Mice
were treated with saline (A and E), 25 mg/m2 PLD (B and F), 25 mg/m
2 DMsPLN (C and G) or 4
× 25 mg/m2
DMsPLN. Each curve represents one animal. Data are the combination of two
separate experiments. In DMsPLN treated groups, 5 mice were still alive and tumor-free after
120 days, which are not presented in the plot. The exact number of long term survivors in each
DMsPLN treatment group is indicated in the brackets.
82
Figure 3.3: Average tumor volume of each treatment group vs. time for mice bearing (A) MDA-
MB 435/LCC6/WT and (B) MDA-MB 435/LCC6/MDR1 tumors. Tumor volume is represented
by the average of tumor volume of all mice in the treatment group ± SEM. Note that the number
of mice in each group decreased in later time points which are depicted in Figure 2, and that in
DMsPLN treated groups, 5 mice were still alive and tumor-free after 120 days, which are not
presented in the plot.
Table 3.1: Effect of DMsPLN and PLD treatment on the tumor growth delay (TGD), median
survival time (MST) and increase in life span (ILS%) of tumor bearing mice.
Average Time of
Tumor Growth
Average Tumor
Growth Delay (%)
Median Survival
Time (MST) Days
Increase in Life
Span (ILS) %
WT MDR1 WT MDR1 WT MDR1 WT MDR1
Saline 38.8
(2.1)*
25.4
(0.82)
- - 40 25 - -
PLD 47.9
(2.5)
33.0
(2.2)
23 30 50 36 26 44
DMsPLN 80.8
(18.8)
55.8
(14.8)
108 120 129 104 225 316
DMsPLN
4×
97.3
(19.5)
56.7
(14.6)
151 123 123 100 210 300
*The values in the brackets are standard error of means (SEM).
As a measure of the effect of treatment on rate of tumor growth, the data in Fig. 3.2 was
replotted as average tumor size, normalized by the average tumor size when the treatment
commenced, for three representative times. For the WT tumors, 23, 26 and 29 days post
tumor inoculation were selected. For the MDR tumors, the times of 20, 24 and 26 days were
A B
83
selected, because they grew faster with 50% of the control mice having reached their end
point at 28 days. The tumor growth inhibition (TGI%) data for PLD, DMsPLN and DMsPLN
4× treatment on the three representative days and their average values are listed beneath Fig.
3.4. The representative data indicate that the TGI of the three treatment groups is in an order:
DMsPLN 4× > DMsPLN > PLD in the WT tumor model, and DMsPLN 4× DMsPLN >
PLD in the MDR model. The results indicate that DMsPLN treatment was superior to PLD
treatment in the present experiments with 2 – 3 times enhanced inhibitory effect of PLD.
*average of the three day TGI values.
Figure 3.4: Normalized average tumor volume at specific time points (days) for mice treated
with saline, 25 mg/m2
PLD, 25 mg/m2
DMsPLN or 4 × 25 mg/m2
DMsPLN in (A) MDA-MB
435/LCC6/WT and (B) MDA-MB 435/LCC6/MDR1 tumor models. The percent tumor growth
inhibition of PLD, DMsPLN and DMsPLN 4× treatment relative to the saline at these time
points are listed in the table. Error bars represent SEM.
Type of tumor Tumor Growth Inhibition (TGI %)
PLD DMsPLN DMsPLN 4×
WT tumor
(day 20, 24,
26)
4, 21, 37 /
20.7*
53, 56, 63 /
57.3*
57, 63, 77 /
65.7*
MDR1 tumor
(day 23, 26,
29)
46, 32, 26
/ 34.7*
74, 81, 74 /
76.3*
71, 71, 80 /
74.0*
B A
84
4.3 Survival of tumor bearing mice following treatment
The effect of DMsPLN on the survival of tumor-bearing mice is shown in Table 3.1 and
Fig. 3.5. Patterns of enhanced survival seen in Figs. 3.2 and 3.3 were recapitulated in Kaplan
Meier survival analysis for WT (Fig. 3.5A) and MDR1 (Fig. 3.5B) tumor-bearing mice.
DMsPLN treatment significantly increased the median survival time of the tumor-bearing mice
compared to the PLD group (Table 3.1). MST of the mice receiving saline was only 40 days in
the sensitive tumor model and 25 days in the resistant tumor model. However MST increased to
129 days and 104 days respectively for groups treated with DMsPLN and DMsPLN 4×. The
mice bearing resistant tumors also showed significant increases in survival time when treated
with DMsPLN and DMsPLN 4× with a median survival of 123 and 100 days respectively. The
increase in life span of WT tumor-bearing mice treated with DMsPLN and DMsPLN 4× ranged
up to 225%, whereas it was only 26% with PLD. Mice bearing the resistant tumor showed an
ILS% of 300 - 316% for DMsPLN, while it was only 44% for PLD treated group. At least one
mouse showed complete tumor disappearance and lived tumor-free for more than 120 days,
which is considered de facto a cure (15% life span, e.g. ~3 months) [349], in DMsPLN single
and 4× groups for both WT and MDR tumors (Fig. 3.2).
85
A B
Figure 3.5: Kaplan-Meier survival curves for (A) MDA-MB 435/LCC6/WT and (B) MDA-MB
435/LCC6/MDR1 tumor bearing mice treated with saline (green), 25 mg/m2
PLD (red), 25
mg/m2
DMsPLN (blue) and 4 × 25 mg/m2
DMsPLN (brown). (In black & white reproduction the
key top to bottom corresponds to curves left to right). In DMsPLN treated tumor-bearing mice, 5
mice were still alive and tumor-free after 120 days, which are not presented in the plot.
4.4 Systemic toxicity of DMsPLN
Safety of the drug nanocarrier administered as PLD and in DMsPLN formulations was
evaluated by measuring the changes in body weight and toxicity-associated blood enzyme levels
as a function of time after treatment. These parameters are generally used as safety indicators in
cancer chemotherapy. The body weight of each group was monitored throughout the course of
the treatment (Fig. 3.6) and loss of 20% of total initial body weight was ruled as a toxic endpoint
requiring euthanasia of the animal. All treated groups underwent an initial decrease in body
weight; however there was moderate recovery from this decline within a few days post
treatment. There was no difference in weight change patterns as a function of time among the
animals receiving saline, PLD, DMsPLN and DMsPLN 4× treatment.
MDA-MB 435/LCC6/WT MDA-MB 435/LCC6/MDR1
86
Figure 3.6: Percent change in body weight as a function of time in mice bearing orthotopic
MDA-MB 435/LCC6/WT (A-D) and MDA-MB 435/LCC6/MDR1 (F-I). Mice were treated with
(A and F) saline, (B and G) PLD, (C and H) DMsPLN or (D and I) DMsPLN 4×. Each curve
represents one animal. Termination of a line represents the time after treatment the animal’s
tumor reached the study size limit and the animal was sacrificed. The number of long term
survivors in each DMsPLN treatment group is indicated in the brackets. Average percent body
weight change as a function of time in mice bearing orthotopic (E) MDA-MB 435/LCC6/WT
and (J) MDA-MB 435/LCC6/MDR1 tumor. Mice were treated with (●) saline, (□) PLD, (▲)
DMsPLN or (x) DMsPLN 4×. The data are presented as average percent change in body weight
± SEM. Data from two separate experiments were combined. The intravenous Dox dose of 25
mg/m2 was administered in all of the formulations.
87
Blood samples were also collected once every 7 days for 5 weeks following treatment or until
animals were euthanized and were analyzed for levels of LDH, ALT and CK. As shown in Fig.
3.7, there was no significant difference in any of the three enzyme levels between the treatment
groups versus the control group though the standard deviations (SD) were large within each
group. The trends for all blood enzyme levels suggested no toxicity in any treatment group.
Figure 3.7: Blood enzymes were used to assess toxicity. Serial blood collection and analysis of
plasma enzyme levels were performed for MDA-MB 435/LCC/WT (A, C and E) and MDA-MB
435/LCC6/MDR1 (B, D, F) tumor bearing mice. LDH (A and B), ALT (C and D) and CK (E and
F) were assayed in all plasma formulations. Each data point represents mean ± SD with n= 5. (In
black & white reproduction, early termination curves correspond to Saline and PLD).
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4.5 In vivo anti-tumor mechanism of DMsPLN
In an attempt to understand some of the details of the mechanism of action of DMsPLN in
breast cancer, tumors were removed at the time of sacrifice and CD31 expressing endothelial
cells were identified using immunohistochemistry (IHC) techniques in harvested tumor tissues.
Typical haematoxylin and eosin (H & E) and CD31 antibody stained tissue samples are
displayed in Figs. 3.8A-3.8D. All tumors were assessed at approximately the same tumor size at
the end point of animal sacrifice though the time after commencement of treatment that tumors
were harvested varied from group to group and animal to animal. Vessel numbers per 0.11 mm2
area of tissue and MVD were determined randomly by selecting four different tumor areas in
three different animals from each treatment group. Animals treated with PLD, DMsPLN and
DMsPLN 4× showed decreases in vessel number (Table 3.2) and MVD (Fig. 3.8E) compared to
the saline treated group in both sensitive and resistant tumor models. The average vessel number
decreased from 12.5 in the saline group to 3.9 in both DMsPLN and DMsPLN 4× treated groups,
in the WT tumor model (Table 3.2). The average vessel number in the mice bearing MDR tumor
decreases from 12.1 in the saline group to 4.7 in DMsPLN and to 4.3 DMsPLN 4× treated
groups, respectively (Table 3.2). Animals treated with DMsPLN and DMsPLN 4× showed
statistically significant differences in MVD compared to the PLD treated animals in both the WT
and MDR tumor models (Fig. 3.8E). The MVD in mice treated with PLD was reduced to about
60%, while in the DMsPLN groups was reduced to about 30% of MVD of the control group.
Both vessel number and vessel density in the tumor tissue after treatment with DMsPLN were
significantly lower than those after treatment with PLD.
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Table 3.2: Immunohistochemical evaluation of the vascularisation of orthotopically implanted
MDA-MB 435/LCC6 breast cancer cells after treatment. All tumors were assessed when they
reached the tumor volume end point and mice were sacrificed. Average vessel number was
assessed from four different areas of the tumor (0.11 mm2) randomly selected from three
different animals in the same treatment group using anti CD31 staining.
Treatment Average vessel number in an area of 0.11mm2
MDA-MB 435/LCC6/WT MDA-MB 435/LCC6/MDR1
Saline 12.5± 1.9 12.1± 2.0
PLD 8.1± 0.8 7.6± 0.4
DMsPLN 3.9± 0.3 4.7± 0.2
DMsPLN 4× 3.9± 0.5 4.3± 0.7
All values are the mean ± SEM
Figure 3.8: Antiangiogenic effect following treatment. Hematoxylin-eosin staining and
immunohistochemical staining with CD31 in tumor sections of (A) saline, (B) PLD, (C) DMsPLN
and (D) DMsPLN 4× treated mice implanted with MDA-MB 435/LCC6/WT orthotopic tumor.
Scale bar in A, B, C, D corresponds to 200 µm. (E) Comparison of normalized microvessel density
determined from four different areas (0.11 mm2) of the tumor were randomly selected from three
different animals in each treatment group. *statistically significant decrease in CD31 staining as
compared to saline treated group (P<0.05). γ statistically significant decrease in CD31 staining as
compared to PLD treated group (P<0.05). Error bars represent SEM. In some cases the error bars
are smaller than data points.
* *
*
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5 Discussion
The present study was conducted to evaluate in vivo toxicity profile and therapeutic
efficacy of DMsPLN in the treatment of primary human breast tumor xenografts. Orthotopic
tumors from MDA-MB 435/LCC6/WT and MDA-MB 435/LCC6/MDR1 cell lines were
implanted in the mammary fat pad, which is more resemble to the microenvironment of human
breast tumor and thus may provide more clinically relevant information than subcutaneous
tumors [350, 351]. Although the WT xenograft may acquire MDR phenotype after treatment
with the Dox formulations, the P-gp MDR1 gene expression in the MDA-MB 435/LCC6/MDR1
xenograft is unlikely to decrease in vivo as the MDR1 gene was stably transduced [352].
Therefore, it serves as a good model of MDR tumor together with its WT counterpart. Herein the
efficacy and tolerability of DMsPLN was compared with clinically available PLD formulation
instead of free Dox solution as PLD has previously been compared to free Dox solution in
clinical trials [338, 340].
The development of MDR due to overexpression of P-gp that reduces intracellular drug
accumulation is a major obstacle to Dox effectiveness [218, 272, 285]. High dose chemotherapy
is thus applied to increase drug accumulation in cancer cells; however it is often accompanied by
higher dose-related normal tissue toxicities, e.g. cardiotoxicity [325, 353]. The PLD, PEGylated
liposomes with encapsulated Dox have an improved pharmacokinetic profile and reduced
cardiotoxicity, while exhibiting similar efficacy, compared to conventional Dox treatment [338-
340] .
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The present results show that DMsPLN treatment resulted in significantly higher efficacy
than clinically used PLD against the MDA-MB 435/LCC6 WT and MDR human mammary
tumor xenografts (Figs. 3.2- 3.5, Table 3.1) while exhibiting similar toxicity (Figs. 3.6 and 3.7).
The mice bearing human breast tumors showed a 108 - 151% increase in tumor growth delay,
210 – 316% increase in life span, and a 10-20% de facto cure after treatment with single or
DMsPLN 4 (Figs. 3.2, 3.3 and 3.5, Table 3.1). During the early days of treatment, the DMsPLN
formulation resulted in 57 – 76% tumor growth inhibition, as compared to 21 – 35% for the PLD
treatment (Fig. 3.4). The therapeutic efficacy of DMsPLN demonstrated in the present work, to
the best of our knowledge, is much higher than previously reported results for other
nanoparticulate formulations of Dox alone or in combination with other agents studied in the
same MDA-MB 435 tumor models. Various studies have been conducted to improve anti-cancer
agent efficacy against MDA-MB 435/LCC6/MDR1 tumors in vivo, but most demonstrated only
a modest or no enhancement of efficacy over saline or free drug controls [354-356]. Liposomal
Dox combined with a P-gp inhibitor and B cell lymphoma 2 (Bcl-2) antisense RNA induced
better tumor suppression than single agent or dual agent treatment, but resulted in up to only a
~60% tumor growth delay compared to control [355]. A polymer micellar formulation of Dox
with integrin-targeted CDCRGDCFC (RGD4C) increased the life span of MDA-MB
435/LCC6/MDR1 tumor model to 46.6 days, 29% longer than the control [357]. In comparison,
treatment with DMsPLN resulted in a MST of over 100 days in the MDR tumor model as
compared to 25 days for the control, which is 300% longer than the control.
The superior anti-tumor efficacy of the DMsPLN group is likely attributable to both the
passive targeting of the PLN to the tumor tissue [345, 346] as well as the efficient cellular uptake
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and perinuclear trafficking of PLN and the synergistic action of Dox and MMC in cancer cells
[193, 328]. Encapsulation of Dox-MMC in the same nanoparticle carrier allows the delivery of
this synergistic drug combination to the tumor site at a predetermined drug ratio, which cannot
be done with free drug cocktails in vivo. The nanocarriers are able to bypass P-gp efflux pumps
and deliver Dox and MMC simultaneously to the site of drug action, i.e., DNA in the nuclei,
resulting in increased DNA double strand breaks thus overcoming several cellular mechanisms
of MDR [191, 193, 328].
Angiogenesis, the development of new blood vessels, also plays a key role in breast
cancer development and metastasis and has been shown to be correlated with stage, grade and
prognosis of patients [358-361]. Microvessel density (MVD) has been shown to be highest in
invasive breast cancer and associated with increased VEGF expression [361, 362]. Several
chemotherapeutic agents (e.g. Dox), used routinely in breast cancer treatment, have been
reported to exhibit anti-angiogenic effects [347, 363]. An inhibitory effect of Dox on hypoxia
inducible factor (HIF1) transcription was found to be responsible for the decrease in vascular
endothelial growth factor (VEGF) and anti-angiogenic effect [347]. In the present study,
treatment with DMsPLN or PLD resulted in significant reductions in tumor blood vessel density
in both sensitive and resistant human orthotopic breast tumor models (Fig. 3.8 and Table 3.2).
However, the DMsPLN inhibited tumor vasculature more profoundly than PLD with both the
number of vessels and normalized microvessel density being about a half of those in the PLD
group.
Many experimental and preclinical studies have demonstrated that frequent
administration of anticancer agents in low doses over an extended period of time, namely
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metronomic chemotherapy, can inhibit tumor angiogenesis [364-366]. In the present work, a
single treatment with DMsPLN brought about an anti-angiogenic effect similar to the DMsPLN
4× treatment. This result implies that the first injection of DMsPLN probably has generated a
depot of Dox and MMC. Previously, we have demonstrated the retention of PLN, up to 7 days
after i.v. administration, in the tumor [345]. Therefore, we speculate that longer tumor retention
of DMsPLN, combined with sustained release of Dox-MMC, may provide anti-vascular effects,
similar to that achieved using low metronomic doses [359, 363] but with a single administration.
This additional anti-angiogenic benefit of DMsPLN treatment may complement the therapeutic
efficacy from direct cytotoxic effects of Dox and MMC on the tumor cells.
DMsPLN treatment was well tolerated in the mice even at a single dose of 50 mg/m2 and
a total dose of 100 mg/m2 from four treatments over three weeks. Unlike PLD, DMsPLN did not
result in body weight loss when given at the clinically applicable Dox dose of 50 mg/m2 (Fig.
3.1). The weight loss due to PLD treatment has previously been observed [367]. This difference
in toxicity between PLD and DMsPLN could be due to the difference in their biodistribution
profiles as PLD has large hepatic retention [339] whereas the PLN formulation has been shown
to have reduced hepatic uptake [345]. Doxorubicinol (DoxOL), a major metabolite of Dox has
been implicated in cardiotoxicity and is predominantly produced by the liver and the heart [239].
DMsPLN may limit DoxOL production by evading liver and heart resulting in decreased
toxicity. Due to the observed weight loss with the 50 mg/m2 PLD treatment, the rest of the study
was conducted at 25 mg/m2 Dox equivalent dose. DMsPLN and PLD treatment showed
insignificant changes in weight (Fig. 3.6) when administered at 25 mg/m2 Dox equivalent dose.
LDH is often elevated in cancer due to tissue damage, making it a common marker for disease
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and toxicity [368]. Over the course of treatment, there was no significant difference in LDH
between the saline group and the groups treated with chemotherapy (Fig. 3.7A and B). An
increase in ALT is often associated with liver damage [369]. Herein no significant difference
was observed between the treated and the saline group in both tumor models (Fig. 3.7C and D).
Serum CK levels, a well characterized marker for cardiac cell damage [370] also did not show
any significant difference between the groups (Fig. 3.7E and F). Collectively, the data of body
weight and the blood enzyme analysis suggest a favourable toxicity profile of DMsPLN when
administered as single or 4× dose, which is similar to that of single PLD dose at 25 mg/m2 Dox.
The DMsPLN treatments showed superior anti-cancer efficacy against the sensitive and
the MDR breast cancer tumors. However, the DMsPLN 4× treatment did not result in significant
improvement in the therapeutic outcomes as compared to a single DMsPLN treatment though a
four-fold increase in total dose was administered to the animals (Figs. 3.2 – 3.5). This could be
due to the anti-angiogenic effect of DMsPLN (Fig. 3.8, Table 3.2). The reduction in MVD may
reduce further delivery of DMsPLN to the tumor after the initial dose in the DMsPLN 4× treated
animals. Another possibility may be the development of anti-PEG antibody, which could reduce
tumor uptake of the nanoparticles decorated with PEG chains [371-373]. Future studies are
needed to elucidate the probable mechanisms and to evaluate the effect of dose and treatment
schedule on the efficacy, toxicity and anti-angiogenic effects of DMsPLN.
In summary, the present study demonstrates that DMsPLN significantly inhibit tumor
growth in both sensitive and MDR orthotopic mammary tumor models, attributable to the
synergistic effects of Dox-MMC, tumor passive targeting and possibly anti-angiogenic effects.
Treatment with DMsPLN does not result in acute or systemic toxicity suggesting it is well-
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tolerated by the experimental animals. Therefore, DMsPLN could be considered as an alternative
to PLD in the treatment of breast cancer.
6 Acknowledgements
The authors sincerely acknowledge the Canadian Breast Cancer Foundation – Ontario Region for
funding this project, Ontario Graduate Scholarship to PP, Dr. Z. Liu (National Center for
Agricultural Utilization Research, US Department of Agriculture) for providing HPESO sample,
and Jean Flanagan for technical assistance with the animal model and blood collection
procedures.
Conflict of Interest
The authors have no conflicts of interests.
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Chapter 4 Integrin-targeted polymer-lipid nanoparticles
encapsulating doxorubicin and mitomycin C enhance treatment of
lung metastases of human triple negative breast cancer in a SCID
mouse model
Preethy Prasad1, Ping Cai1, Dan Shan1, Hibret A. Adissu2, Andrew M. Rauth3, Xiao Yu
Wu1
1Department of Pharmaceutical Sciences, Leslie Dan Faculty of Pharmacy, University of Toronto, 144 College Street, Toronto, Ontario, Canada, M5S 3M2 2Toronto Centre for Phenogenomics, 25 Orde St. 3rd fl., Toronto, Ontario, Canada M5T 3H7 3Department of Medical Biophysics, University of Toronto, 610 University Ave, Toronto, Ontario, Canada M5G 2M9
This manuscript will be submitted to Journal Controlled Release
All work in this manuscript was performed by P.Prasad with assistance from the co-authors,
except for the paraffin embedding, sectioning, and staining of the histological samples.
97
1 Abstract
Triple negative breast cancer (TNBC), a subtype of breast cancer, accounts for about 15% of all
human breast cancers and is known for its aggressive characteristics leading to metastases. The
integrin receptor ανβ3 has been shown to play a critical role in tumor angiogenesis and
metastases and be highly expressed on angiogenic endothelium in malignant tissues and TNBC
MDA-MB 231 cells. Our laboratory thus developed ανβ3 integrin-targeted nanoparticles with
surface conjugated Arg-Gly-Asp (RGD) which exhibited inhibitory effects on cancer cell
adhesion and invasion. The present work aimed to prepare RGD-conjugated stealth polymer-
lipid hybrid nanoparticles encapsulating doxorubicin (Dox) mitomycin C (MMC) (RGD-
DMsPLN) and evaluate their in vivo efficacy and safety in a murine lung metastatic model of
human breast cancer. Lung metastasis of breast tumor was established using human MDA-MB
231-luc-D3H2LN, a luciferase-transfected cell line that was derived from a spontaneous lymph
node metastasis. The bio-distribution and tumor accumulation of the nanoparticles were
examined by whole animal optical imaging using near infrared fluorescence-labeled
nanoparticles. The efficacy and systemic toxicity of nanoparticles were evaluated against free
Dox-MMC solutions. Whole animal imaging demonstrated the localization of the nanoparticles
in the lung metastasis site of breast cancer. Integrin-targeted RGD-DMsPLN resulted in a
significant reduction in lung metastases without producing drug-associated systemic toxicity as
observed in the group treated with free Dox-MMC solutions. The results from a murine model of
aggressive metastatic human breast cancer suggest that RGD-DMsPLN may provide a clinically
relevant, improved intervention of TNBC.
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Key words: Lung metastases, Triple negative breast cancer, doxorubicin and mitomycin C
coencapsulation, polymer-lipid hybrid nanoparticles, integrin targeting, efficacy, toxicity
2 Introduction
Despite advances in diagnosis and treatment, breast cancer remains the second most frequent
cause of cancer death in women (behind lung cancer) [145]. Triple negative breast cancer
(TNBC), a subtype of breast cancer characterized by lack of receptors (estrogen/progesterone,
human epidermal growth factor -2 (HER-2)) accounts for about 15% of breast cancer and is
known for its aggressive characteristics leading to metastases [374, 375]. The metastatic spread
of breast cancer from the primary tumor site to a secondary site, such as the lung, is the major
cause of death in patients [376], in which integrin receptors have been found to play an important
role [5-8]. They are overexpressed in both tumor cells and angiogenic endothelial cells and
promote tumor progression and metastases [94, 377-379]. In breast cancer patients, a strong
correlation has been found between the percent of alpha v beta 3 (αvβ3)-positive vessels within
the tumor and disease progression [377]. These findings suggest an opportunity to target αvβ3
integrin for improving treatment of TNBC.
Treatment of TNBC is a major challenge due to its aggressiveness, poor prognosis, lack of
therapeutic target due to the absence of receptor proteins, and rapid development of resistance to
chemotherapeutic agents [376]. Lack of HER-2 prevents treatment of TNBC with standardized
therapies for breast cancer such as trastuzumab and lapatinib [380]. Chemotherapy remains the
only possible therapeutic option in the adjuvant or metastatic setting of TNBC [381].
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Anthracycline drug doxorubicin (Dox) is currently employed to treat patients with metastatic
cancer with overall response rates between 35-50% in patients with TNBC [382]. A higher
response rate is observed in TNBC treated with anthracycline-based or anthracycline/taxane–
based chemotherapy than other combination regimens; however, relapse is frequently observed
resulting in shorter survival times [383, 384]. Improved overall survival is achieved by dose
intensification of conventional chemotherapeutic agents including Dox [385]. However,
cumulative cardiotoxicity is a major limitation to the increased dose intensity of doxorubicin and
can lead to potentially fatal congestive heart failure [324]. Therefore, new strategies are needed
for targeting and treating tumor metastases which are currently considered to be incurable.
Nanoparticles as drug carriers have achieved clinical success in improving the tolerability of Dox
chemotherapy [367, 386], however new dose-limiting effects have been noted in the clinic in the
form of palmar plantar erythrodysethesia [342, 343]. Previously, we have developed a
biocompatible stealth polymer lipid nanoparticle (PLN) system carrying Dox and mitomycin C
(MMC) with good drug release kinetics [193, 310]. The combination of Dox and MMC resulted
in a synergistic cell kill in both sensitive and multidrug resistant (MDR) breast cancer cells
attributable to Dox-MMC enhanced DNA double strand breaks [191, 193, 387]. The synergistic
cell kill of Dox and MMC was further enhanced by co-encapsulation in the PLN (DMsPLN) and
delivering the synergistic combination to the perinuclear region of cells [191, 387]. The efficacy
of Dox-MMC has further been demonstrated in a murine mammary mouse model when
delivered by microspheres intratumorally [192] and in an orthotopic human mammary mouse
model when delivered using DMsPLNs intravenously [388]. Treatment with DMsPLNs
demonstrated enhanced efficacy compared to liposomal Dox (PLD) with up to a 3-fold increase
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in animal life span, compared to untreated animals, and undetectable normal tissue
toxicity[388]. To enhance the targetability and tumor retention of the nanoparticles, we have
further functionalized the surface of PLNs with cyclic Arg-Gly-Asp (cRGD) to interact with αvβ3
integrin receptors overexpressed on breast cancer cells [377, 389]. Our earlier study on cRGD-
conjugated solid lipid nanoparticles (RGD-SLNs) has demonstrated that, even without a drug,
the RGD-SLNs could inhibit the adhesion and invasion of αvβ3 integrin-overexpressing TNBC
MDA-MB 231 cells in vitro [26].
Herein, we have further investigated the potential of DMsPLN and RGD conjugated DMsPLN
(RGD-DMsPLN) to treat lung metastases of breast cancer. An optimal concentration of one
percent mole of RGD on the surface of DMsPLN was used, which was previously identified to
have maximum tumor uptake and retention [389]. An experimental lung metastasis model of
human breast cancer was established in mice using a highly invasive human MDA-MB 231-luc-
D3H2LN cell line. The effect on metastatic burden in the lungs of treatment with DMsPLN and
actively targeted RGD-DMsPLN was compared with free Dox-MMC solutions was evaluated
using bioluminescence imaging. In addition, histopathology was performed on liver and heart
tissues and serum cardiac troponin (cTnT) was measured to assess treatment induced systemic
toxicity.
3 Materials and methods
3.1 Materials
Myristic acid, poly(ethylene glycol)-100-stearate (PEG100SA), poly(ethylene glycol)-40-stearate
(PEG40SA), and all other chemicals, unless otherwise mentioned, were purchased from Sigma-
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Aldrich Canada (Oakville, ON, Canada). Mitomycin C and doxorubicin HCl were purchased
from Polymed Therapeutics (Houston, TX, USA). Hydrolyzed polymers of epoxidized soybean
oil (HPESO) were a gift from Drs. Z. Liu and S. Erhan (Food and Drug Administration,
Washington, DC, USA). Pluronic F68 (PF68) (non-ionic block copolymer) was a kind gift from
BASF Corp. (Florham Park, NJ, USA). Cyclo(-RGDfK) was purchased from AnaSpec, Inc.
(Fremont, CA, USA) and used without modification. All cell culture plastic ware was purchased
from Sarstedt (Montreal, QC, Canada). Cell culture medium, Alpha Modified Eagle Medium
(αMEM), and phosphate buffered saline (PBS) were obtained from Tissue Culture Media
Facility, Ontario Cancer Institute (Toronto, ON, Canada). Fetal bovine serum (FBS) and trypsin
were purchased from Invitrogen, Inc. (Burlington, ON, Canada). Human breast cancer MDA-MB
231-luc-D3H2LN cells and D-luceferin were purchased from Caliper. Female SCID mice were
purchased from Ontario Cancer Institute (Toronto, ON, Canada). All studies in mice were
performed in accordance with the guidelines and regulations of the Animal Care Committee at
the University Health Network.
3.2 Synthesis and characterization of myrj56-cRGDfK targeting constructs
The targeting construct was prepared by first activating Myrj59 (PEG100SA) with p-
nitrophenylchloroformate (p-NPC) as previously described [280].The structure of Myrj56-NPC
was confirmed by 1H- and 13C-NMR in deuterated chloroform (CDCl3) by standard pulse
sequences on a Varian Mercury 300 MHz NMR (Agilent Technologies, Inc., Santa Clara, CA,
USA). For the conjugation of cRGDfk to Myrj59-NPC, 5 mg of cRGDfk (Peptides International,
Inc., Louisville, KY, USA) was dissolved in 2 ml of 0.1 M sodium bicarbonate (pH 8.3) at room
102
temperature. 20mg of Myrj59-NPC was added to the solution and was stirred overnight at room
temperature. Unreacted p-NPC was removed by dialysis against distilled, deionized (DDI) water
using Slide-A-Lyzer® 3500 Da molecular weight cut off mini dialysis unit (Thermo Scientific,
Rockford, IL, USA) for 48 hours with changing the dialysate every 12 hours. The Myrj59-
cRGDfK targeting construct was isolated by freeze drying and the structure was confirmed by
1H-NMR in D2O by standard pulse sequences on a Varian Mercury 300 MHz NMR (Agilent
Technologies, Inc., Santa Clara, CA, USA).
3.3 Preparation and characterization of polymer lipid nanoparticles
The nanoparticles were prepared and characterized as previously described [193, 387]. Briefly,
50 mg of myristic acid, 8 mg of Myrj52 (PEG40SA) and 4mg Myrj59 (PEG100SA) were melted
at 65°C. Under stirring, 4 mg of MMC was added to the molten fatty acid, followed by the
simultaneous addition of 400 μL of a 12.5 mg/mL solution of Dox in distilled, deionized water
and 50 μL of a 50 g/L solution of HPESO in distilled, deionized water. In addition, 50 μL of 100
mg/mL PF68 was added and the emulsion was stirred at 65°C for 20 min followed by
ultrasonification for 5 min (80% peak amplitude and 5 mm probe depth in the solution) using a
Hielscher UP100H probe ultrasonicator (Hielscher USA, Inc. Ringwood NJ, USA). For the
preparation of RGD-DMsPLNs, 10 μL of 4.4 mg/mL Myrj59-cRGDfk was also added at this
step resulting in 1% RGD peptide concentration on the nanoparticle surface [389]. The entire
emulsion was immediately transferred into 5 mL of 5% dextrose solution. Particle size and zeta
potential were measured by dynamic light scattering and electrophoretic mobility, respectively,
using a NICOMPTM 380ZLS (PSSNICOMP, Santa Barbara, CA, USA) apparatus. Immediately
103
after formulation, the DMsPLN suspension was centrifuged through a 0.1µm filter and the drug
in the filtrate was assayed spectrophotometrically at 540 nm for Dox and 364 nm for MMC.
Drug loading (%wt drug/wt lipid) and encapsulation efficiency (%wt loaded drug/wt total drug)
were then calculated in the filtrate. The remaining DMsPLNs were incubated with Sephadex SP
C-25 anionic dextran microspheres at 4°C overnight to remove any unencapsulated drug. The
microspheres were then pelleted by centrifugation and the DMsPLNs were decanted into a
separate vial for injection. DMsPLNs were made fresh before each injection.
For the in vivo fluorescence investigation, the near infra-red fluorescent probe indocyanine green
(ICG) was loaded into the nanoparticles. 50 mg of myristic acid, 8 mg of Myrj52 (PEG40SA)
and 4 mg of Myrj59 (PEG100SA) were melted at 75°C. 25µL of 0.1M ICG dye in ethanol was
added and stirred for 10 minutes. 10 μL of 4.4 mg/mL Myrj59-cRGDfk, 50 μL of 100 mg/mL
PF68 and 378 μL of distilled water were added, stirred for 20 min followed by ultrasonification
for 5 min (80% peak amplitude and 5 mm probe depth in the solution). The entire emulsion was
transferred to 5% dextrose solution and stirred on ice for 5 minutes.
3.4 Cell culture
Highly metastatic triple negative breast cancer MDA-MB 231-luc-D3H2LN cells were obtained
from Caliper and grown in pH 7.2 alpha modified minimal essential medium (Ontario Cancer
Institute Media Laboratory) supplemented with 10% FBS (fetal bovine serum, Invitrogen) in a
37°C humidified incubator with 5% CO2. Cell doubling times were typically 24h. Cells were
trypsinized and subcultured at 50-fold dilution once they were confluent. Cells were tested for
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their suitability for use in animals and were found to be pathogen-free (Research Animal
Diagnostics Laboratory, Columbia, MO, USA).
3.5 Metastasis model development
Female, SCID mice (6 weeks old) were injected with 0.5 million MDA-MB 231-luc-D3H2LN
cells in 100 µL of growth medium via the tail vein to establish tumor metastasis in the lungs.
Animals had free access to food (Irradiated Tecklad LM485, Harland Tecklad, Indianapolis, IN,
USA), sterile water and were kept in Allentown ventilated microisolator cages (Allentown Inc.
Allentown, NJ, USA) with each cage bearing 5 mice. After 1 week, D-luciferin solution
(150mg/kg) was injected intraperitonealy into the mice and bioluminescent images were
obtained 10 min post injection using an IVIS Spectrum (Caliper Life Sciences, Inc. Hopkinton,
MA, USA) whole animal imager.
3.6 Biodistribution study
The biodistribution studies were performed one week after tumor inoculation in female SCID
mice. 200 µL of PLN or RGD-PLN (1% RGD) loaded with ICG was injected into the lateral tail
vein of SCID mice (n=3/group). Biodistribution of the nanoparticles was recorded at various
time points after injection with an excitation and emission wavelengths of 710 nm and 820 nm,
respectively, using the IVIS Xenogen whole animal fluorescence imaging system. In a separate
experiment, mice were euthanized by CO2 asphyxiation after 2 hours, 4 hours and 8 hours, and
the liver, spleen, kidneys, gut, heart, and lungs were excised and immediately imaged with the
whole animal fluorescent imager.
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3.7 In vivo treatments
One week after tumor inoculation treatments were initiated in the SCID mice. An initial test was
conducted to determine the tolerable free drug (Dox-MMC) and DMsPLN dose. Tumor free
mice, (5/group) were randomly allocated to different treatment groups: 1) Free Dox-MMC (3
mg/kg), 2) Free Dox-MMC (6 mg/kg), 3) Free Dox-MMC (10 mg/kg), 4) Free Dox-MMC (15
mg/kg), 5) DMsPLN (3mg/kg), 6) DMsPLN (6mg/kg), 7) DMsPLN (6 mg/kg), 8) DMsPLN (15
mg/kg). The treatment was based on the Dox dose and was given intravenously via the tail vein.
The body weight of each mouse was measured every other day following treatment and was
related to the first day as percent change in body weight.
For the therapeutic study, tumor bearing mice (5/group) were randomly allocated to different
treatment groups: 1) control (5% dextrose), 2) free Dox-MMC (3 mg/kg), 3) DMsPLNs (3
mg/kg), 4) RGD-DMsPLNs (3smg/kg), 5) DMsPLN (15 mg/kg), RGD-DMsPLNs (15 mg/kg).
Each treatment was given equivalent to a free Dox dose via tail vein injection. Tumor growth
was monitored weekly using bioluminescent imaging. Mice were given D-luciferin (150 mg/kg)
substrate by intraperitoneal injection and bioluminescent imaging was initiated 10 min after
injection with 1 min exposure time. The signal intensity of lung metastasis was quantified as the
sum of all detected photon counts within the region of interest (ROI). At Day 28, mice were
euthanized and lungs were excised for histology.
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3.8 Evaluation of liver and cardiotoxicity
Tumor metastasis bearing mice received i.v administration of saline, free drug (3 mg/kg), free
drug (15 mg/kg), DMsPLNs (3mg/kg) and DMsPLNs (15 mg/kg) via tail vein intravenous (i.v.)
injection (n=5/group). 7 days post treatment, mice were euthanized, liver and hearts were
excised, formalin fixed for histological section and stained with haematoxylin and eosin (H&E)
stain. Prepared slides were analyzed by a pathologist at Toronto Centre for Phenogenomics
(Toronto, ON, Canada).
In a separate set of experiments, blood was collected into heparin sulphate-coated capillary
tubes (Microvette CB 300 LH, Sarstedt Inc., Montreal, QC, Canada) 7 days post treatment from
metastatic tumor bearing mice. Blood was centrifuged at 1.4 g for 20 min at 4oC to isolate
plasma, which was immediately flash frozen on liquid nitrogen until processing. Plasma was
assayed for cardiac troponin T (cTnT) levels using mouse cardiac troponin T ELISA kit from
Kamiya Biomedical Company (Seattle, WA, USA).
3.9 Statistical analysis
Results are presented as mean ± standard error of mean (SEM). Statistical comparison was made
using one-way ANOVA. P values <0.05 were considered significant.
4 Results
4.1 In vivo biodisitribution of nanoparticles in tumor bearing mice
The biodistribution of nanoparticles was assessed in female SCID mice with established
metastatic foci in the lungs, one week after tumor inoculation. MDA MB 231-luc-D3H2LN
metastatic breast cancer cell line was given intravenously to establish metastatic foci and was
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visualized using bioluminescence imaging. To visualize the nanoparticles, a near infrared (NIR)
dye ICG was encapsulated within the nanoparticle core. Fig. 4.1A shows the time dependent
distribution profile of PLN and RGD-PLN given intravenously and monitored non-invasively at
different time points using Xenogen whole body animal imager with an excitation of 740 nm and
emission 820 nm. Significant near infrared fluorescence from the nanoparticles was observed in
the tumor metastasized lungs within 15 minutes post injection (Fig. 4.1B). The NIR intensity of
the whole body decreased overtime and was barely observable at 24 h post injection for PLN and
36 h post injection for RGD-PLN. Both PLN and RGD-PLN accumulated in the lung bearing
metastatic MDA MB 231-luc-D3H2LN breast tumor (Fig. 4.1B). However a much higher
accumulation and retention of RGD-PLN was observed compared to the non- targeted PLN in
the lung (Fig. 4.1B). RGD-PLN was retained in the lung for up to 36h, unlike the PLN which
was barely observable at 36h.
In a separate set of experiments, mice were sacrificed and major organs such as liver, spleen,
kidney, gut, heart and lungs were isolated to evaluate tissue distribution of nanoparticles. As
shown in Fig. 4.1C, a strong NIR signal was observed in lung bearing metastatic tumor while
other tissues showed negligible NIR fluorescence signal, except for liver and gut. At 2 hours,
PLN showed higher accumulation in the liver and the gut compared to the targeted RGD-PLN.
At 4 hours, an increased NIR signal of both PLN and RGD-PLN was detected in lungs.
However, RGD-PLN showed longer retention in the lungs compared to the PLN. No substantial
accumulation of RGD-PLN and PLN was observed in the heart and kidneys.
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Fig. 4.1: Nanoparticle distribution in mice bearing MDA MB 231-luc-D3H2LN metastatic
breast cancer. ICG loaded PLN or RGD PLN were injected intravenously in the tail vein 7 days
post tumor inoculation. Tumor inoculation in the lung was evaluated using bioluminescent
imaging (mice in far left) A) Whole body biodistrubition was imaged over time using IVIS
Xenogen animal imager with Ex: 745nm and Em: 820. B) Zoomed in image of the accumulation
of nanoparticles in the lung. The red box in A depicts where the mice was zoomed in figure B.
(C) Organs were excised for ex vivo imaging at 2 h, 4 h and 8 h after injection.
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4.2 Maximum tolerable dose assessment
Maximum tolerable dose (MTD) study was conducted for further toxicity assessment in SCID
mice without tumor. 5 mice per group were injected i.v with either free drug (free Dox-MMC) or
DMsPLN at various doses (3 mg/kg, 6 mg/kg, 10 mg/kg and 15 mg/kg equivalent Dox dose),
and were evaluated for 1 week after treatment injection (Table 4.1). The MTD was determined
based on the dose at which all animals survived without any significant presence of acute
toxicity. Most of the mice treated with free Dox-MMC (6-15 mg/kg) showed weight loss and
lack of grooming. All 5 of the mice treated with free Dox-MMC at 6 mg/kg, 10 mg/kg and 15
mg/kg exhibited clinical signs of severe toxicity such as significant weight loss of over 20%
from the initial weight, hunched back and ruffled fur coats, thereby reaching clinical end point
(Table 4.1). Therefore, MTD of free Dox-MMC in SCID mice was about 3 mg/kg Dox dose
which is consistent with previous results [390]. Treatment with DMsPLN with all doses (3
mg/kg – 15 mg/kg Dox dose) showed no apparent signs of toxicity in terms of weight loss and
fur ruffling. All 5 mice remained alive for 7 days in all DMsPLN treated groups (Table 4.1).
Table 4.1: Determining MTD by measuring the number of mice showing sever signs of acute
toxicity, 7 days following treatment.
Treatment # of mice with severe signs of acute toxicity (ie.
Body weight and fur ruffling)
Free Dox- MMC (3mg/kg Dox dose) 0
Free Dox- MMC (6mg/kg Dox dose) 5
Free Dox- MMC (10mg/kg Dox dose) 5
Free Dox- MMC (15mg/kg Dox dose) 5
DMsPLN (3mg/kg Dox dose) 0
DMsPLN (6mg/kg Dox dose) 0
DMsPLN (10mg/kg Dox dose) 0
DMsPLN (15mg/kg Dox dose) 0
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4.3 Inhibition of tumor metastasis in vivo
Having seen evidence of RGD tumor targeting of nanoparticle in vivo [280, 389], we evaluated
the suppression of metastasis by nanoparticles loaded with Dox-MMC. Highly metastatic breast
cancer MDA-MB 231-luc-D3H2LN cells were implanted into SCID mice by i.v. injection
through the tail vein and tumor progression was monitored by bioluminescence imaging, a
sensitive and non-invasive method. This technique allows us to visualize and quantify tumor
burden without the need of animal sacrifice at each point of analysis. In general, the relative level
of bioluminescence signal correlates with metastatic burden [391].
Treatments with 1) Saline, 2) Free drug (Dox-MMC), 3) DMsPLN and 4) RGD-DMsPLN were
injected i.v. 1 week after tumor inoculation and bioluminescent images were acquired every
week (Fig. 4.2 and 4.4). In Fig. 4.2, a Dox dose of 3mg/kg body weight was used in all
treatments groups. Fig. 4.2B shows decreased bioluminescent signal in mice treated with RGD-
DMsPLN at the end of Day 28. Quantitative analysis of bioluminescence signal showed a
significant inhibition of tumor metastasis by RGD-DMsPLN (3 mg/kg) treatment compared to
the saline treated mice (p< 0.05) (Fig. 4.2A). However, treatment with free Dox-MMC and
DMsPLN (3 mg/kg) did not show significant differences from the saline treated group.
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Fig. 4.2: RGD-PLN reduced the metastasis burden in vivo. (A) Quantification of tumor
metastasis burden in mice over time by bioluminescence imaging. Mice (n=5/group) were treated
with Saline, Free Drug (Dox-MMC), DMsPLN and RGD- DMsPLN, 7 days after iv. tumor
inoculation. Bioluminescent images were acquired 10 mins after injection of D-luciferin
(150mg/kg) intraperitonealy using IVIS Xenogen whole animal imager. Data represents mean ±
SEM (n=6) (B) Representative in vivo bioluminescent images of mice on Day 28 after tumor
inoculation. * indicates statistically significant decrease in RGD-DMsPLN group compared to
the saline control (p<0.05). Please note that the treatment is based on Dox dose.
*
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Since the therapeutic efficacy of the treatments was not significantly different from each other,
we decided to investigate the effect of treating tumor metastasis by injecting a higher Dox dose
of 15 mg/kg body weight. Mice treated with free Dox-MMC at 15 mg/kg Dox dose showed
severe toxicity resulting in death by 7 days post treatment. However, no signs of toxicity were
observed when free drug was encapsulated within the nanoparticle (DMsPLN) and injected at a
15 mg/kg Dox dose (Table 4.1). Due to the early death the free drug (free Dox-MMC) was given
at a maximum tolerable Dox dose of 3 mg/kg and was compared to DMsPLN given at 15 mg/kg
Dox dose.
Bioluminescence imaging clearly showed that both DMsPLN (15 mg/kg) and RGD-DMsPLN
(15 mg/kg) markedly suppressed tumor metastasis (p<0.001), as evidenced by the lowest
luciferace activity in tumor metastasis foci (Fig. 4.3A and Table 4.2). In contrast much higher
bioluminescence signals were observed in both saline and free drug (3 mg/kg) treated group (Fig.
3B). RGD-DMsPLN (15 mg/kg) exhibited the highest therapeutic effect and also showed
significant improvement in efficacy compared to the DMsPLN (15 mg/kg) treated group (p<
0.001). Free Dox-MMC (3 mg/kg) was not significantly different from the saline control
(p>0.05) in reducing tumor burden. At the end of Day 28, mice were killed, lungs were removed
and weighed. As shown in Fig. 4.3C, metastatic nodules were significantly reduced in the lungs
evidenced by much lower lung weights (p< 0.001) from the mice treated with DMsPLN (15
mg/kg) and RGD-DMsPLN (15 mg/kg), whereas free Dox-MMC (3 mg/kg) had little therapeutic
effect. H&E staining also showed the presence of very few metastatic nodules (dark purple
regions) in lungs of mice treated with DMsPLN (15 mg/kg) and RGD-DMsPLN (15 mg/kg) with
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most of the lungs free of tumor (Fig. 4.3D). In contrast, metastasis nodules occupied most of the
lung tissue from the saline and free Dox-MMC (3 mg/kg) treated groups (Fig. 4.3D).
Fig. 4.3: High dose of PLN significantly inhibits tumor burden. (A) Quantification of tumor
burden measured using bioluminescent imaging once a week. Treatments (Saline, Free Drug
[3mg/kg Dox-MMC] , DMsPLN[15mg/kg] , RGD-DMsPLN [15 mg/kg] ) were given
intravenously via the tail one week after tumor inoculation. Data represents mean ± SEM (n=6)
Table 4.2: Metastatic burden on mice as measured by total flux (p/s), 28 days following
treatment. Data represents mean ± SEM.
Treatment Total Flux (p/s)
Saline 3.14 x 108 ± 1.70 x 10
8
Free Dox- MMC (3mg/kg Dox dose) 2.47 x 108 ± 0.88 x 10
8
DMsPLN (15mg/kg Dox dose) 6.08 x 106 ± 1.80 x 10
6
RGD-DMsPLN (15mg/kg Dox dose) 2.33 x 106 ± 0.45 x 10
6
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4.4 Evaluation of liver toxicity
Microscopic changes were evaluated in the liver tissue, one week post treatment (Fig. 4.4).
Tumor bearing mice were treated with 1) Saline, 2) Free drug (Dox-MMC, 3 mg/kg), 3) Free
drug (Dox-MMC, 15 mg/kg), 4) DMsPLN (3 mg/kg) and 5) DMsPLN (15 mg/kg). One week
post treatment, mice (n= 5) were sacrificed and histopathological evaluation of the liver tissue
was performed. Results indicated that both the free drug groups (3 mg/kg and 15 mg/kg)
exhibited the most severe pathological changes compared with the other groups (Fig. 4.4A).
Livers from saline, DMsPLN (3 mg/kg) and DMsPLN (15 mg/kg) appeared normal with no
signs of toxicity. Mice treated with the lower free drug dose (3 mg/kg Dox) showed mild
microvesicular lipid accumulation within hepatocytes in midzonal and centrilobular regions (Fig.
4.4A, white arrows). Much greater liver toxicity was observed in mice treated with the free drug
at 15 mg/kg Dox resulting in microvesicular lipidosis affecting 60-70% of the hepatocytes (Fig.
4.4A, white arrows). These mice also exhibited moderate number of single cell necrosis evident
by nuclear fragmentation and cytoplasmic hypereosinophilia (Fig. 4.4B, white arrows). In
addition, binucleated hepatocytes and karyomegaly (enlarged nuclei) were frequent in the liver
of mice treated with free drug (15 mg/kg Dox dose) (Fig. 4.4B, black arrows). The
morphological diagnosis of the entire treated group is listed in Fig. 4.4C.
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Fig. 4.4: Free drug shows hepatotoxicity not seen in DMsPLN treatment groups: Livers were
removed from tumor bearing mice receiving Saline, Free Drug (3 mg/kg), Free Drug (15 mg/kg),
DMsPLN (3 mg/kg) or DMsPLN (15 mg/kg), 7 days after treatment. Liver tissue was stained
with H&E and histopathology was performed. (A) H&E stained liver sections (40×
magnification) showed hepatic microvesicular lipidosis (indicated by arrows) in free drug (Dox-
MMC) treated group both at 3 mg/kg and 15 mg/kg Dox dose. (B) H&E liver sections (100×
magnification) from mice treated with free drug (Dox-MMC) at 15 mg/kg showed liver necrosis
(white arrows) and karyomegaly (black arrows). (C) Image analysis of H&E describing the
distribution and severity of toxicity. Please note that the treatment is based on Dox dose.
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4.5 Evaluation of caridiotoxicity
Histological assessments of the hearts of tumor bearing mice in different treatment groups
(saline, free drug [3 mg/kg], free drug [15 mg/kg], DMsPLN [3 mg/kg] and DMsPLN [15
mg/kg]) were assessed 7 days after treatment to evaluate the Dox induced cardiotoxicity. At the
level of whole heart in cross-sectional view, all the mice treated with free drug at 15 mg/kg Dox
dose had blood clotting within the right ventricle (RV) chamber due to incomplete ejection of
blood (Fig. 4.5A). This observation was not present in other treatment groups. Focally extensive
mineralization within the left ventricle (LV) was also observed in the 15 mg/kg free drug treated
group with extensive necrosis of the RV (Fig. 4.5B, white arrow). Treatment with free drug at 3
mg/kg Dox dose resulted in mild myocardial necrosis (Fig. 4.5B, white arrow) and myofiber loss
within the RV subepicardial zones. However, such pronounced vacuolar degeneration and
necrosis was not prevalent in other groups including saline, DMsPLN (3 mg/kg) and DMsPLN
(15 mg/kg).
Cardiac troponin levels (cTnT) were also measured to confirm the cardiotoxicity seen in some
treatment groups. cTnT is a sensitive and specific biomarker for detection of myocardial
infarction [392]. Seven days post treatment, cTnT levels were greatly increased in the free drug
(3 mg/kg and 15 mg/kg) treatment groups compared to the saline control and DMsPLN groups
(Fig. 4.5C). Free drug at 15 mg/kg Dox dose had the highest level of cTnT levels indicative of
cardiac injury. No significant difference was observed in the saline treated and the DMsPLN
treated groups.
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Fig. 4.5: Free drug shows cardiotoxicity not seen in DMsPLN treatment groups: Hearts from
tumor bearing mice receiving Saline, Free Drug (3 mg/kg), Free Drug (15 mg/kg), DMsPLN (3
mg/kg) and DMsPLN (15 mg/kg) were removed 7 days post treatment, flash frozen, formalin
fixed, paraffin embedded and stained with H & E. Hearts. (A) Whole organ heart morphology at
1.2× magnification. Mice treated with Free Drug (15 mg/kg) had blood clots within the right
ventricular chamber. (B) Longitudinal section of heart analyzed at 40× magnification indicated
occasional nuclear fragments (white arrows) in the Free Drug (15 mg/kg) treated group resulting
in epicardial necrosis. (C) Assessment of Cardiac troponin levels in serum samples measured 7
days post treatment. All the mice were inoculated with tumor a week before treatment initiation.
Error bars represent S.E.M. In some cases the error bars are smaller than data points.
*Significantly different from both Free Drug (3 mg/kg) and Free Drug (15 mg/kg) groups.
Please note that the treatment is based on Dox dose.
* * *
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5 Discussion
Chemotherapy is the only treatment option available for TNBC since currently available targeted
endocrine and HER2-directed therapies are ineffective [375, 393]. Chemotherapy with Dox is
considered to be the most effective towards TNBC [382]; however its clinical utility is limited
due to low therapeutic index and serious adverse drug reactions [324]. Non-targeted long
circulating nanoparticulate systems such as Doxil have been extensively used in the clinic to
improve Dox tolerability while maintaining similar efficacy levels as free Dox [242].
Administration of Doxil has significantly reduced the risk of cardiotoxicity and
myelosuppression [394]; however new side effects in the form of palmar plantar
erythrodysethesia (PPE) occur in at least 45% of patients [242]. Recent studies have utilized
specific-targeted delivery of nanoparticles using surface functionalization to further enhance
efficacy and minimize toxicity [395-397].
Functionalizing nanoparticles with RGD peptides targeting αvβ3 integrin receptors overexpressed
on tumor neovasculature [398] is a potential approach for targeted delivery. αvβ3 integrin is also
expressed by invasive breast cancer where it promotes recruitment of blood vessels and invasive
function of sprouting endothelial cells [94, 398]. Therefore targeting αvβ3 integrin is currently
being explored by various researchers to improve diagnosis via imaging [399-401] or to deliver
anti-cancer agents to solid tumors [395-397]. However, the success of functionalizing
nanopaticles and delivering them in vivo has been limited. Previously, we have conjugated
cRGD to nanoparticle resulting in increased binding with the angiogenic vessels in the tumor and
prolonged tumor retention [280]. The concentration of RGD was further optimized to 1% total
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surface cRGD concentration to exhibit maximal tumor uptake and retention in vivo while
inhibiting adhesion and invasion characteristics of metastatic breast cancer cells in vitro [389].
In the present study, we demonstrated the efficacy of integrin αvβ3 targeted, RGD- nanoparticle
co-loaded with Dox-MMC in an experimental metastatic human breast cancer SCID mouse
model. At a low dose (3 mg/kg), both the nanoparticle formulations (DMsPLN and RGD-
DMsPLN) showed modest effects in inhibiting tumor growth. Previously we have shown that
free Dox-MMC combination exerts synergistic cytotoxic action against breast cancer cells in
vitro [191, 193, 387, 402]. However, in the present study treatment with the free Dox-MMC
combination did not show any advantage over the saline control treated mice (Fig. 4.2 and 4.3),
attributable to the huge difference in their pharmacokinetics. The pharmacokinetic profiles of
Dox and MMC are completely different with elimination half-lives (t1/2) being 7~20 hours for
Dox and 7~90 minutes for MMC [403, 404]. Therefore, it is necessary to encapsulate both Dox-
MMC within the same nanoparticle to ensure simultaneous delivery of both drugs to the same
cancer cells to realize their synergism in vivo.
Encapsulating the anti-cancer drugs Dox and MMC within the same nanoparticle would have
allowed simultaneous release of both drugs at the site of action. Biodistribution studies in the
present work showed co-localization of nanoparticles within the tumor in the lung (Fig. 4.1),
suggesting this advantage of co-delivery of Dox and MMC by DMsPLN and RGD-DMsPLN.
The nanoparticle formulation also allowed mitigation of the system toxicity usually observed
with Dox administration [324]. Owing to the extreme acute toxicity observed at higher doses
(Table 4.1), we were only able to evaluate the efficacy of free Dox-MMC at 3 mg/kg (toxicity-
limited dose). Both nanoparticle systems (DMsPLN and RGD-DMsPLN) demonstrated a
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significant higher efficacy in controlling the metastatic growth of tumor at 15 mg/kg dose (Fig.
4.3 and Table 4.2) without inducing noticeable systemic toxicity. The enhanced efficacy with the
DMsPLN treatment is consistent with our previous observations in an orthotopic human breast
cancer tumor model [388]. Previously we have demonstrated a significantly higher tumor growth
delay with DMsPLN treatment compared to Doxil, in immunocompromised mice bearing human
MDA MB-435 breast tumors and immunocompetent mice [388, 405].
Dox-MMC delivery to the tumor and tumor vasculature using RGD conjugated DMsPLN (RGD-
DMsPLN) resulted in 106-fold and 2.6 fold improvement in controlling lung metastasis burden
compared to free Dox-MMC and DMsPLN treatment, respectively, as detected by the
bioluminescence signals (Fig. 3A). This enhanced efficacy observed with RGD-DMsPLN
treatment could be attributable to several factors. Biodstribution studies showed increased
accumulation and retention of RGD targeted nanoparticles compared to the untargeted particles
in the lung bearing metastatic MDA-MB 231-luc-D3H2LN breast tumor (Fig. 4.1). In addition,
enhanced accumulation of RGD conjugated nanoparticles in tumor neovasculature was observed
compared to the untargeted nanoparticle using intra-vital fluorescence microscopy [280]. Since
drug release from the nanoparticles is slow and sustained [193, 277], the enhanced retention of
RGD nanoparticles in tumor tissue could enhance anti-cancer efficacy. The ability of RGD-
DMsPLN to specifically target and be retained by tumor vasculature, can result in release of
cytotoxic Dox-MMC to disrupt tumor vascular endothelium. Destabilizing tumor vasculature
will prevent delivery of nutrients and oxygen important for continuous tumor growth [238],
hence reducing tumor metastasis.
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Integrin αvβ3 is highly expressed in metastatic breast cancer, and has been shown to be involved
in cross talk with growth promoting factors involved in cell survival, migration and proliferation
[377-379]. Therefore, the enhanced efficacy with the targeted nanoparticles could also be due to
the binding of RGD to αvβ3 integrin receptors expressed on breast cancer cells. Binding of RGD
peptides with αvβ3 integrin has been shown to inhibit cancer cell migration and adhesion,
preventing cell metastasis [406, 407]. Though Phase II clinical trials have been conducted using
MEDI-523 and MEDI-522, antibodies against αvβ3 integrin, to inhibit integrin reception [408-
410], these studies have been put on hold as they were unable to demonstrate significant
efficacy. In addition, administration of unmodified peptides results in their rapid hydrolysis and
elimination, making them unfavourable for clinical usage as anti-metastatic therapeutics [411-
413]. The use of RGD-conjugated nanoparticle systems may be a more effective therapeutic
option. The enhanced efficacy of DMsPLN towards metastatic tumors is a novel finding and
shows for the first time that conjugation of RGD to DMsPLN further enhances the therapeutic
effect of the nanoparticle towards metastatic cancer.
Dose dependent toxicity of anti-cancer agents limits the cumulative drug dose received by the
patient, which in turn can limit the therapeutic efficacy of the drug. Encapsulation of Dox-MMC
inside the nanoparticle significantly reduced manifestations of drug toxicity in tested tissue (heart
and liver). It is reported that the involvement of reactive oxygen species produced by both Dox
and MMC can cause hepatocyte damage [414-417]. As reported, the majority of Doxorubicinol
(DOXol) production, the major phase I metabolite of Dox, has been implicated in Dox associated
cardiotoxicity [239, 240]. Therefore, the lower toxicity of DMsPLN, in comparison with free
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Dox, may be ascribed to reduced tissue accumulation, lower drug concentrations in normal
organs, including the liver and the heart, and reduced DOXol production.
6 Conclusion
The present study demonstrates that DMsPLN significantly inhibit lung metastases in a SCID
mouse model of highly aggressive metastatic MDA-MB 231-luc-D3H2LN breast tumor. The
conjugation of RGD to DMsPLN for the purpose of targeting the tumor endothelium and cancer
cells further enhances the efficacy of this drug delivery vehicle. Treatment with DMsPLN, at the
levels tested, also does not result in acute or systemic toxicity suggesting it is well-tolerated by
the experimental animals. Therefore, the use of RGD-DMsPLN containing synergistically acting
Dox and MMC combination could be a promising therapeutic strategy for metastatic TNBC due
to their fewer adverse side effects and more effectiveness compared to the currently employed
chemotherapeutic regimens.
7 Acknowledgements
The authors gratefully thank the Canadian Breast Cancer Foundation-Ontario Region for
supporting this work, the Canadian Institutes of Health Research and the National Science and
Engineering Research Council of Canada (NSERC) for the Equipment Grant. Ontario Graduate
Scholarship and Pfizer scholarship to P. Prasad, NSERC Graduate Scholarship to D. Shan, the
University of Toronto Fellowship, University of Toronto Nanotechnology Network Award and
Anna and Alex Beverly Fellowship to D. Shan are also gratefully acknowledged.
The authors declare that they have no competing financial interests.
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Chapter 5 Multifunctional albumin based MnO2 nanoparticles
modulate solid tumor microenvironment by attenuating hypoxia,
acidosis, VEGF and enhance radiation response
Preethy Prasad1ǂ, Claudia R Gordijo1ǂ, Azhar Z Abbasi1, Azusa Maeda2, Angela Ip1, Andrew M Rauth3, Ralph S DaCosta2 and Xiao Yu Wu1
1Department of Pharmaceutical Sciences, Leslie L. Dan Faculty of Pharmacy, University of Toronto, Toronto, Ontario, Canada, M5S 3M2. 2Ontario Cancer Institute, The Campbell Family Institute for Cancer Research, Princess Margaret Cancer Center, 610 University Avenue, Toronto, Ontario Canada, M5G 2M9. 3Department of Medical Biophysics, University of Toronto, ,610 University Avenue, Toronto, Ontario, Canada, M5G 2M9
ǂThese authors contributed equally to this work.
This work has been published in ACS Nano, 2014, 8(4), 3202-3212.
Reprinted with permission from {P.Prasad, C. Gordijp, A. Abbasi, A. Maeda, A. Ip, A.M.Rauth, R.S.
DaCosta and X.Y.Wu, Multifunctional Albumin–MnO2 Nanoparticles Modulate Solid Tumor
Microenvironment by Attenuating Hypoxia, Acidosis, Vascular Endothelial Growth Factor and Enhance
Radiation Response, ACS Nano20148 (4), 3202-3212}. Copyright {2014} American Chemical
Society
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1 Abstract Insufficient oxygenation (hypoxia), acidic pH (acidosis), and elevated levels of reactive oxygen
species (ROS), such as H2O2, are characteristic abnormalities of the tumor microenvironment
(TME). These abnormalities promote tumor aggressiveness, metastasis and resistance to
therapies. To date, there is no treatment available for comprehensive modulation of the TME.
Approaches so far have been limited to regulating hypoxia, acidosis or ROS individually,
without accounting for their interdependent effects on tumor progression and response to
treatments. Hence we have engineered new multifunctional and colloidally stable bioinorganic
nanoparticles composed of polyelectrolyte-albumin complex and MnO2 nanoparticles (A-MnO2
NPs) and utilized the reactivity of MnO2 towards peroxides for regulation of the TME with
simultaneous oxygen generation and pH increase. In vitro studies showed that these NPs can
generate oxygen by reacting with H2O2 produced by cancer cells under hypoxic conditions. A-
MnO2 NPs simultaneously increased tumor oxygenation by 45% while increasing tumor pH from
pH 6.7 to pH 7.2by reacting with endogenous H2O2 produced within the tumor in a murine breast
tumor model. Intratumoral treatment with NPs also led to the downregulation of two major
regulators in tumor progression and aggressiveness, i.e., hypoxia-inducible factor-1 alpha (HIF-
1α) and vascular endothelial growth factor (VEGF) in the tumor. Combination treatment of the
tumors with NPs and ionizing radiation significantly inhibited breast tumor growth, increased
DNA double strand breaks and cancer cell death as compared to radiation therapy (RT) alone.
These results suggest great potential of A-MnO2 NPs for modulation of the TME and
enhancement of radiation response in the treatment of cancer.
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Keywords: Multifunctionalnanoparticles, manganesedioxide, modulating tumor
microenvironment, hypoxia, acidosis, HIF-1, VEGF, radiation response, breastcancer.
2 Introduction
In solid tumors hypoxia (low oxygenation) often occurs as a consequence of a disrupted
balance between the supply and consumption of O2, owing in part to tumor growth and vascular
abnormalities, the latter also affecting O2 transport insufficiencies [55]. Hypoxia, a characteristic
of the tumor microenvironment (TME), has been shown to contribute to the resistance to
radiation therapy (RT) and to promote clinically aggressive phenotypes in cancer [139, 418].
Studies have demonstrated that nearly 40% of breast cancers exhibit tumor regions with oxygen
concentrations below that required for half maximal radiosensitivity, reducing the effectiveness
of radiation therapy [140].
Hypoxia also leads to chronic over activation of hypoxia-inducible-factor-1 (HIF-1) which
plays a pivotal role in adaptive responses to hypoxia by modulating various cellular functions
like proliferation, apoptosis, angiogenesis, pH balance and anaerobic glycolysis [229, 230]. Upon
activation, HIF-1 binds to the hypoxic responsive element, thereby promoting transcription of
various genes including VEGF (vascular endothelial growth factor) and genes encoding for
glucose transporters [419]. The expression of VEGF further induces angiogenesis and plays a
key role in promoting malignant tumor growth [420, 421]. HIF-1-also mediates the switch from
aerobic to anaerobic metabolismin hypoxic tumors for energy preservation by activating glucose
transporters and glycolytic enzymes leading to an increase in levels of lactic acid and acidosis
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(lower extracellular pH, pHe< 6.9) [64, 422]. In addition, hypoxia and high proliferation of
cancer cells produce excess amounts of reactive oxygen species (ROS), e.g. hydrogen peroxide
(H2O2) [423]. Together, hypoxia, acidosis and ROS promote mutagenesis, metastasis of cancer
cells, angiogenesis and resistance to therapies, contributing to treatment failure.
To date, various strategies have been proposed to modify the TME, aimed at the (systemic)
improvement of tumor oxygenation to surmount hypoxia-associated radioresistance. These
strategies include hyperbaric oxygen therapy [424], artificial blood substitutes [425], and drugs
which preferentially kill or sensitize hypoxic cells to radiation [426]. Unfortunately, the utility of
such methods in clinical settings is limited due to safety concerns, reagent stability and/or
inconsistent clinical response. Therefore, there is a continued and urgent need for new strategies
to improve tumor oxygenation in vivo to enhance the radiation response in solid tumors.
Here, we have taken advantage of the high reactivity and specificity of manganese dioxide
nanoparticles (MnO2 NPs) towards H2O2 for the simultaneous and sustained production of O2
and regulation of pH [427, 428] to modulate the TME. Unlike other strategies to increase tumor
oxygenation, mostly by the delivery of molecular oxygen by nanoparticles with limited O2
loading capacity [139], MnO2 NPs are able to generate O2 in situ for a prolonged time by
reacting with undesirable and abundantly available tumor metabolites (H2O2 and H+). Another
advantage of MnO2 NPs is their dual functions as both catalyst and reactant. In the latter case,
they are decomposed to harmless, water-soluble Mn2+
ions [428], avoiding the in vivo
accumulation of the metal oxide commonly observed for other metal-based nanoparticle (NP)
systems [429]. Compared with other metal nanoparticles extensively explored for biological
applications, MnO2NPs have been limited to use in biochemical sensors [428, 430] and bioassays
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[431]. To our knowledge, their reactivity towards tumor H2O2 has not been studied for in vivo
ROS reduction, O2 production or for the regulation of pH in biological systems. Therefore, for
the first time, the development and in vivo characterization of MnO2 NPs are reported for
simultaneous modulation of hypoxia and acidosis of the TME, and for enhancement of ionizing
radiation-induced tumor cell cytotoxicity in a murine breast tumor animal model.
3 Methods
3.1 Nanoparticle synthesis
MnO2 NPs were prepared by directly mixing the aqueous solutions of KMnO4 and
poly(allylamine hydrochloride) (PAH, 15kDa).Briefly, 18 mL of KMnO4 solution (3.5mg mL-1
)
was mixed with 2 mL of PAH solution (37.4 mg mL-1
), the mixture was left for 15 min at room
temperature until all permanganate was converted to MnO2. NP formation was confirmed by
recording UV-Vis absorption spectrum. NPs were washed three times with doubly distilled
(DDI) water using ultracentrifugation (50k rpm for 1hr). This step led to small (~15 nm) MnO2
NPs stabilized with PAH. At the final step, BSA was added to the MnO2 NP solution at a
BSA/NP ratio 2.5 % (wt/wt), and NaCl was added to make the solution normal saline (0.9%
NaCl). This step led to the formation of A-MnO2 NPs (~50 nm), with several MnO2 NPs
entrapped in a PHA/BSA complex due to strong electrostatic interaction between the protein and
the polymer. A typical preparation led to a ≈ 0.7% A-MnO2 NPs solution, corresponding to
≈1.1mM MnO2 as determined by inductively coupled plasma (ICP) analysis. A-MnO2NPs were
further diluted with cell medium or sterile saline for in vitro and in vivo studies, respectively.
Protein labelling kits AnaTag™ HiLyte Fluor™ 594 (Texas Red) and AnaTag™ 750
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(AnaspecInc, USA) were used to label albumin for the preparation of red fluorescent and near-
infrared NPs, respectively.
3.2 Cell lines, tumor models and treatments
In vitro: A murine EMT6 breast cancer cell line was utilized and cultured following standard cell
culture procedures [432]. For all in vitro experiments, cells in αMEM medium (105 cells per mL)
were treated with A-MnO2 NPs for 1h. Cell viability was measured using a standard MTT
protocol [432]. In vivo: All procedures strictly complied with the ethical and legal requirements
under Ontario’s Animals for Research Act and the Federal Canadian Council on Animal Care
guidelines for the care and use of laboratory animals and were approved by the University
Animal Care Committee of the University of Toronto. Solid tumors of EMT6 breast cancer cells
(106) were grown orthotopicly in Balb/c mice and animals were randomly allocated for all
treatments (n=3/group). For in vivo experiments, tumors were injected with 50 µL of A-MnO2
NPs solution in saline (0.2mM MnO2), which made the MnO2 concentration ≈ 45 µM in a ≈200
mm3 tumor. Controls were injected with equivalent volume of sterile saline.
3.3 Quenching of H2O2 by nanoparticles
For the quenching experiments, A-MnO2 NPs (90 µM) in cell medium containing 10% FBS at
370C and H2O2 (1 mM) was added to initiate the reaction. The residual concentration of H2O2
was determined over time using a PeroXOquant assay kit (Pierce, USA), at 37°C. Cell medium
with 10% FBS was used as a vehicle control.
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3.4 In vitro oxygen and pH measurements
O2 generated by A-MnO2NPs and pH changes were measured in a semi-sealed chamber coupled
with a MI-730 micro-oxygen electrode and a MI-407 pH + MI 402 reference microelectrodes
(Microelectrodes Inc, USA), at 37°C. A-MnO2 NPs were dispersed in αMEM cell medium
containing 10% FBSto give various MnO2 concentrations (10 – 90 µM). The system was made
hypoxic by bubbling with N2. Endogenous level of H2O2 (250 µM) was injected into the
chamber to initiate O2 generation. For experiments with hypoxic cells: Murine breast cancer
EMT6 cells (105 cells per mL) were suspended and stirred in αMEM medium in glass vials
plugged with rubber stoppers and pierced with two hypodermic needles for gassing. The cell
suspension was made hypoxic by introducing a mixture of 95% N2 and 5% CO2 for 20 min at
37°C. A-MnO2 NPs (45 µM) were then injected and the oxygen levels monitored over time. For
all experiments: pH or O2 were monitored every 60 s using an Oakton pH 1100 (Thermo Fisher
Scientific Inc, USA) coupled with O2-ADPT Oxygen Adapter (Microelectrodes Inc, USA) for O2
measurements. All electrodes were calibrated according to manufacturer’s instructions. αMEM
medium with 10% FBS, with or without cells was used as control.
3.5 Cellular uptake of NPs
Murine EMT6 breast tumor cells (105 cells) were incubated for 1h with A-MnO2 NPs (45 µM) at
37°C before microscopic analysis. Cell uptake of NPs by transmission electron microscopy
(TEM) was performed using a H7000 TEM microscope (Hitachi, Japan), following standard
methods for sample preparation [432]. An EVOS fluorescence microscope (AMG, USA) was
used to image live cells following incubation with red fluorescent dye labelled A-MnO2 NPs.
Cell nuclei were stained blue with HOESCHT 33342 (Invitrogen Molecular Probes, USA).
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3.6 Tumor retention of NPs
A Xenogen IVIS Spectrum Imaging System (Caliper Life Sciences Inc., USA) was used to
image tumor bearing animals over time following i.t. treatment with near-infrared labelled A-
MnO2 NPs. At each time point, a bright field image was acquired and fluorescence-labelled A-
MnO2 NPs were imaged at 754 nm excitation and 778 nm emission. Image fluorescence was
quantified by equalizing the fluorescence intensity range across all images.
3.7 Tumor pH measurements
A pH-sensitive fluorophore SNARF-4F (Life technologies S23920, NY USA) was used for ex
vivo tumor pH imaging following an established protocol [433]. Tumor bearing mice were
injected i.t. with NPs in saline followed by i.v. injection of the dye (1 nmol of SNARF-4F in 200
µL of sterile saline). Animals were sacrificed 20 min following injections, tumor tissue was
immediately harvested, cut in half and imaged with Xenogen IVIS Spectrum (Caliper Life
Sciences Inc., USA). For control experiment sterile saline was injected i.t. followed by
intravenous (i.v.) injection of SNRAF, tumors were imaged ex vivo using the same conditions.
All the necessary calibration curves of dye were performed following published protocols [434,
435] and biological tissue-like phantoms were prepared following standard procedures [433] (see
Supporting Information for details). Tumor pH was also measured using a MI-407 pH + MI 402
reference microelectrodes following a standard protocol [436] (Microelectrodes Inc, USA).
3.8 Tumor oxygenation measurements
A Vevo LAZR Photoacoustic Imaging System (VisualSonics Inc., Canada) with a 21 MHz
centre frequency transducer (LZ-550, VisualSonics Inc., Canada) was used to measure vascular
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oxygen saturation (sO2) in situ over time before and after i.t. treatment with A-MnO2 NPs.
Ultrasound was utilized to guide NP injection in order to administer treatments to the tumor.
Animals were maintained below 7% oxygen atmosphere during the experiment and sO2
measurements were assessed using standard multispectral photoacoustic imaging in the tumors in
vivo using two excitation wavelengths (750 nm and 850 nm) for deoxygenated and oxygenated
hemoglobin, respectively.
3.9 Immunohistochemistry detection of tumor hypoxia
The hypoxia marker pimonidazole hydrochloride (HypoxyprobeTM
-1 plus kit, HypoxyprobeInc,
USA) was used for ex vivo tissue staining of hypoxia following the protocol provided with the
kit. Rabbit polyclonoal HIF-1α antibody (dilution 1:100, Novus Biologicals, Catalog number:
NB100-134) and Rabbit anti-VEGF (dilution 1: 100, Thermo Scientific, Catalog number: ab-
222-P) were used for the staining of HIF-1α and VEGF, respectively. Briefly, tumor bearing
mice (n=3/group) were treated i.t. with A-MnO2 NPs or saline (control). After pre-determined
times animals were sacrificed and tumor tissues were harvested and fixed with 10% neutral
buffered formalin solution (Sigma Aldrich, USA) for histological analysis. Tumor tissue
preparation and analysis were performed by the CMHD Pathology Core laboratory at Mount
Sinai Hospital, Toronto. Slides were scanned with a NanoZoomer 2.0 RS whole slide scanner
(Hamamatsu, Japan) and images were analysed with Visiopharm 4.4.4.0 software.
3.10 In vivo radiation treatment
Solid tumor of EMT6 murine breast cancer cells were grown orthotopically in Balb/c mice.
Mice were divided into 4 groups (n=3/group): 1) Saline, 2) Saline + RT, 3) A-MnO2 NP, 4) A-
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MnO2 NP + RT. Treatmentswere initiated when the tumors reached an approximate volume of
100 mm3. The mice were restrained in a specially designed acrylic box, and the tumors were
irradiated locally with 10Gy, 30mins after i.t administration of saline and A-MnO2 NP. The
tumor size was measured as a function of time with vernier calipers in two dimensions and tumor
volumes were calculated by the formula V= [(length) × (width)2
]/2. At the end of experiment, the
animals were sacrificed and the tumor masses were excised and weighed. Tumor tissue was also
formalin fixed and stained with terminal deoxynucleotidyltransferasedUTP nick end labelling
(TUNEL) and haematoxylin andeosin (H&E) to determine percent apoptosis and necrosis.
In another set of experiments, mice were sacrificed 24 hours after radiation treatment. Tumor
tissue was excised, formalin fixed, sectioned and gamma H2AX measured to evaluateDNA
DSBs. Slides were scanned with a NanoZoomer 2.0 RS whole slide scanner (Hamamatsu, Japan)
and images were analysed with Visiopharm 4.4.4.0 software.
The enhancement of DNA DSBs due to the combination of RT + A-MnO2 NP was also evaluated
in a dorsal skin-fold window chamber (DSWC) EMT6 mouse model [437]. Treatments (Saline
and A-MnO2 NP) were injected i.t. and only half of the chamber was irradiated at 10Gy.
Irradiating only half of the chamber allowed us to determine the effect of treatment alone in the
same mice. 24 hours post irradiation tumor tissue was excised and stained with gamma H2AX
staining to evaluate DNA DSBs. Slides were scanned and images were analysed with
Visiopharm 4.4.4.0 software.
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3.11 Statistical analysis
Data are presented as mean ± standard error of the mean for results obtained from three
independent trials unless otherwise indicated. Student’s t-test or analysis of variance (ANOVA)
followed by Tukey t-test (OriginPro8©
) were utilized to determine statistical significance
between two or more groups, respectively. p-values< 0.05 were considered statistically
significant.
4 Results and discussion
4.1 Preparation of A-MnO2 NPs
For the synthesis of NPs, we employed a one-step method to reduce manganese
permanganate (KMnO4) to MnO2 NPs with cationic polyelectrolyte poly(allylamine
hydrochloride) (PAH). This synthesis procedure is rapid, reproducible and gives stable MnO2
colloidal dispersions with an average NP size distribution of 15 nm (Fig. 5.1a-b). In the present
synthesis method, we were able to decrease by 50% the amount of PAH normally used in
polyelectrolyte-based NPs synthesis [438] , as shown in the ultraviolet-visible (UV-Vis)
spectrum of samples prepared with various polyelectrolyte ratios (Fig. 5.1c). The decrease in the
amount of PAH utilized in the NP formulation is very important for in vivo applications, since
cationic polyelectrolytes can show pronounced concentration-dependent cytotoxicity. The
polyelectrolyte used here served not only as a reducing reagent to reduce KMnO4 to MnO2, but
also as a protective layer to stabilize as-formed NPs due to electrostatic repulsion (zeta potential
+30 mV, Fig. 5.1d).
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Figure 5.1.Characterization of A-MnO2 NPs:(a) Diagram & TEM images of MnO2 and A-MnO2
NPs. Precursor MnO2 NPs (~15 nm) are stabilized by positively charged PAH. In A-MnO2 (~50
nm) several MnO2 particles are entrapped in a PHA/BSA complex due to strong electrostatic
interaction between the protein and the polymer.(b) Size distribution of NPs. (c)UV-Vis
absorption spectra of KMnO4 solution and MnO2 NPs prepared at various molar ratios between
PAH andMnO2. After the reaction with PAH at ratios 2:1 and 3:1, the KMnO4 peaks (315, 525
and 545 nm) disappeared, and a new broad peak around 300 nm appeared for these samples, an
indicator of the formation of MnO2 nanoparticles. The new peak around 300 nm is attributed to
the surface plasmon band of colloidal manganese dioxide.Error! Reference source not
found.(d) Effect of coating of MnO2 NPs with BSA on zeta potential for various BSA/NPs
ratios. By adding BSA to a MnO2 NP aqueous suspension, the zeta potential of the NPs
decreased from +30 mV to -25 mV. (e)Picture of polyelectrolyte MnO2 NPs (left) and A-MnO2
NPs (right) (1mM) in various aqueous media: DDI water, normal saline (0.9% NaCl) and αMEM
cell medium containing 10% fetal bovine serum (FBS). MnO2NPs undergo aggregation in saline
or cell culture medium, while A-MnO2 NPs are stable in these media. The red color observed in
the vials comes from the pH indicator in the αMEM medium.
135
To formulate MnO2 nanoparticles for biomedical in vivo applications we took into
account the issues of nanoparticle stability, size control and toxicity. For biological applications,
NPs must be stable in cell culture medium or normal saline required for in vitro and in vivo
studies, respectively. Polyelectrolyte-coated MnO2 nanoparticles are too small and positively
charged, which can cause high instability in cell medium or saline and result in toxicity. To solve
these problems, we have conjugated small polyelectrolyte-coated MnO2 nanoparticles with
bovine serum albumin (BSA) and obtained particles of suitable size, charge, colloidal stability
and biocompatibility for in vitro and in vivo applications, while maintaining the MnO2 reactivity
towards H2O2 for the production of oxygen and increase in pH.BSA can form stable non-
covalent complexes with cationic polyelectrolytes [439], leading to lower NP toxicity [440]. The
MnO2 NP-albumin conjugates (A-MnO2) prepared were approximately 50 nm in size (Fig. 5.1b),
negatively charged (-25 mV) (Fig. 5.1d) and stable in alpha minimal essential medium (αMEM)
cell medium and saline (Fig 5.1e), making them suitable for in vivo applications. The albumin
coating also provided the NPs with different surface charge and chemistry allowing us to further
functionalize the NP surface with protein-reactive fluorescent dyes such as Texas RedTM
(excite
596/emit 617 nm) and amine-reactive near infrared dye (excite 754/emit 778 nm). These
fluorescence-labelled NPs were utilized in our subsequent in vitro and in vivo studies.
4.2 Multifunctionality of A-MnO2 NPs in culture medium
We first investigated the multifunctionality of the A-MnO2 NPs to generate O2 and to
increase the medium pH in vitro upon reaction with H2O2 at endogenous levels. The reaction
between MnO2 and H2O2 is a complex reaction leading to the decomposition of H2O2 and the
production O2 as summarized in Fig. 5.2a. Besides the production of O2, the reaction causes an
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increase in the local pH by the consumption of H+ ions and the production of an intermediate
Mn-oxo-hydroxide (MnOOH) [428]. This phenomenon can be particularly useful for the
regulation of local pH in cancer cells and tumour tissue. Hence we studied if A-MnO2 NPs
would generate measurable amounts of oxygen and increase pH at low concentrations of H2O2
found in the human body (i.e., 100 µM and up to 1 mM) [441]. We found that at a very low
concentration (~45 µM of MnO2), the NPs were able to completely quench 1 mM H2O2 in cell
medium within 40 minutes (Fig. 5.2b). We further investigated the O2 generating properties of
the NPs using an in-house-made hypoxia-maintaining chamber coupled with both a
commercially available oxygen probe and a pH microelectrode. Significant amounts of O2 was
produced (Fig. 5.2c) accompanied by an increase in the pH of physiological buffer
(phosphate/saline buffer) by one pH unit from pH 6.8 to pH 7.8 (Fig. 5.2d) by the reaction of
45µM of MnO2with 250 µM H2O2. In an attempt to simulate in vivo conditions where H2O2 is
continuously generated by tumor cells, we measured the O2 production by the NPs during the
continuous addition of exogenous H2O2 (250 µM) into the chamber every 30 min. We observed
that a single dose of the NPs (90 µM MnO2) continuously generated O2 for at least 6 cycles of 30
min each (Fig. 5.2e). These results demonstrated that H2O2 and protons can diffuse rapidly
across the polyelectrolyte-albumin complex, access the reactive sites of the MnO2 cores, produce
O2 and increase pH in a sustained manner under hypoxic conditions.
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Figure 5.2. In vitro reactivity of A-MnO2 NPs towards hydrogen peroxide:(a) Scheme showing
the reactivity of MnO2 towards H2O2 for the production of O2 and removal of protons. (b)
Quenching of endogenous level H2O2 (1 mM) by A-MnO2 NPs (45 µM). (c) Oxygen generation
at various A-MnO2 NP contents (numbers indicate MnO2 in µM). (d) Simultaneous O2
generation and pH increase vs. time by the A-MnO2 NPs. (e) O2 generation by addition of H2O2
to an A-MnO2 NP suspension. All experiments were performed (n = 3) in cell culture medium
containing 10% FBS at 37°C. Error bars are standard error of the mean.
4.3 Uptake of A-MnO2 NPs by breast cancer cells
It is known that the aberrant metabolism of cancer cells leads to significantly elevated
cellular concentrations of H2O2 [423]. We hypothesized that if the NPs could be taken up by
cancer cells, they could react quickly with endogenous H2O2 produced by cancer cells under
hypoxic stress, thus producing O2 in situ. To test this hypothesis, we first examined the cellular
uptake of the NPs by incubating EMT6 murine breast cancer cells with fluorescence-labelled A-
138
MnO2 NPs, and observed significant cellular uptake of NPs within 60 min of incubation (Fig.
5.3a). This finding was confirmed by transmission electron microscopy (TEM). TEM images
(Fig. 5.3b) showed EMT6 cells in vitro underwent membrane invagination and engulfment of the
NPs and the NPs taken up by the cell were distributed within the cell cytoplasm and vesicles
after 1 h incubation.
Figure 5.3. Cellular uptake, cellular oxygen generation and cytotoxicity of A-MnO2 NPs:(a)
Fluorescence images of cellular uptake of A-MnO2 NPs at 37°C by murine EMT6 breast cancer
cells following 1h incubation with NPs. (b) TEM images of cellular uptake of A-MnO2 NPs. (c)
O2 generation by A-MnO2 NPs incubated with hypoxic cancer cells (n = 3). Suspended cells are
made hypoxic and upon addition of A-MnO2 to the culture oxygen is generated by the reactivity
of NPs towards H2O2 released by hypoxic cancer cells. (d) Viability of murine EMT6 cancer
cells (105 cells/ mL) exposed to various concentrations of A-MnO2 NPs for 48 h. Percent of cell
viability was determined with MTT assay. (n=3) Error bars represent standard errors of the
mean.
139
This cellular uptake of A-MnO2 NPs may be attributable to the NPs interaction with the
albumin-binding protein SPARC (Secreted Protein, Acidic and Rich in Cysteine) on the cancer
cells. It has been reported that albumin receptor gp60 at endothelial cells in angiogenic tumor
vasculature and the albumin-binding protein SPARC overexpressed in a majority of tumors
including breast cancer are responsible for tumor targeting and cancer cell uptake of albumin-
bound taxanes [442-445] . Since A-MnO2 NPs are completely saturated by BSA (Fig. 5.1d), they
may be recognized by SPARC thus facilitating cellular uptake of the NPs. Nevertheless,
confirmation of this mechanism is outside the scope of the present work and will be conducted in
future experiments.
4.4 Oxygen generation in the presence of hypoxic cancer cells
We found that the NPs incubated with hypoxic breast cancer cells could react quickly with
endogenous H2O2 produced by the cells under hypoxic stress, thus producing O2in situ (Fig.
5.3c). Significant amounts of O2 (~6-fold increase of O2 levels in the medium) were detected
within 2 min by reacting with H2O2 released by the cancer cells (Fig. 5.3c). These results
indicate that the endogenous levels of H2O2 released by hypoxic cancer cells in vitro is sufficient
to react with the NPs and generate measurable O2 without addition of exogenous H2O2 to the
culture medium. Moreover, at the concentration used for in vitro O2 generation (45 µM MnO2),
A-MnO2 NPs showed relatively low cytotoxicity to EMT6 cancer cells (~80% cell viability)
(Fig. 5.3d). Based on these data, we hypothesize that elevated levels of H2O2 in solid tumors
could serve as a reactant for O2 production in vivo.
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4.5 Effect of A-MnO2 NPs on tumor oxygenation
The effect of A-MnO2 NPs on oxygen saturation within orthotopic murine EMT6 cell breast
tumors was assessed with a small animal photoacoustic (PA) imaging system following
intratumoral (i.t.) injection of 50 µL of A-MnO2 NP suspension in saline. PA imaging measures
vascular saturated O2 (sO2) by the differential optical absorption of oxygenated and
deoxygenated hemoglobin at different wavelengths, which is directly correlated with changes in
O2 concentration in the blood [446]. To ensure similar localization of the NPs in each tumor we
used ultrasound image-guidance to inject the NPs into the tumor in vivo. Since the blood vessels
maintain a saturated level of oxygen under normoxic conditions, the mice were maintained under
7% O2 during the experiments to visualize the enhancement of oxygen production by the A-
MnO2 NPs. We measured vascular sO2 before and after i.t. injections of A-MnO2 NPs
suspension or saline only (control) and found that sO2 increased promptly by approximately 45%
as compared with control tumors (Fig. 5.4 a-c). It is important to point out that PA imaging of
sO2 depends on the presence of blood flow, which is lacking in the necrotic and avascular tumor
core [333]. Thus the PA images revealed sO2 was mainly generated within the tumor periphery
(Fig. 5.4a). However, this does not imply that the NPs are unable to produce O2 in the hypoxic
region close to the tumor core, as the O2 generating capacity of the NPs is limited only by the
presence of H2O2. Interestingly, the nearly immediate detection of O2 at the tumor periphery
suggests the rapid distribution of the NPs within the tumor mass from the injection site (i.e.,
tumor center) perhaps due to the interstitial pressure gradient with a higher pressure in the tumor
core than within the peripheral region [333].
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Figure 5.4. Effect of A-MnO2 NPs on tumor oxygenation:(a) Representative2D photoacoustic
images of EMT6 solid tumors showing parametric map of estimated oxygen saturation (sO2) pre-
and post-i.t. injection of saline (control) or A-MnO2 NPs. (b) Average total sO2 in the tumor
over time. (c) Comparison of average tumor sO2 levels before and after treatments (n =3). Error
bars represent standard errors of the mean. *statistically significant increase (*p = 1.8E-5) in sO2
as compared to saline (control) treated group.
In this study, we used i.t. delivery of A-MnO2 NPs for the assessment of the effect of the NPs
on tumor oxygenation in vivo. The reason for using i.t. treatment is two-fold. Firstly, the local
delivery method is better than systemic delivery (e.g. intravenous injection) in terms of
uniformity of NP dose delivered to each tumor owing to a broad variation from tumor to tumor
in morphology and NP penetration. Secondly, local intratumoral delivery of therapeutics has
been emerging as an effective treatment of many types of localized operable and inoperable solid
tumors (e. g. breast, colorectal, lung, prostate, skin, head and neck and brain tumors) due to its
advantages over systemic methods, including dramatically higher local drug concentration, better
therapeutic outcomes and minimal systemic toxic side-effects [447, 448].
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4.6 Effect of A-MnO2 NPs on tumor pH
To measure intratumoral pH, we employed complementary fluorescence imaging [433] and
microneedle [449] methods. We applied an established ex vivo tissue protocol for mapping tumor
tissue pH which utilizes multispectral fluorescence imaging (MSFI) in conjunction with a pH-
sensitive fluorescent dye (SNARF-4F) injected in vivo prior to animal sacrifice [433]. In our
study, EMT6 tumor bearing animals were first injected i.t. with A-MnO2 NPs, followed by
intravenous injection of the SNARF-4F dye to stain for tumor pH. MSFI images of the
intratumoral facets of dye-perfused tumors were then acquired ex vivo in tissue sections and
correlated to local pH from the calibration curves obtained earlier with biological tissue-like
phantoms (see Supplementary Information). The tumor pH was also accessed ex vivo with a pH
microneedle probe [449] immediately after the MSFI procedure. We found that intratumoral
injection of A-MnO2 in orthotopic solid tumors led to higher intratumoral pH (Fig. 5.5). After a
single i.t. injection of A-MnO2 NPs, the tumor pH increased, after only 20 min, from 6.2 to 6.7
(as determined by MSFI) and from 6.7 to 7.3 (as determined by microneedle probe) (Fig. 5.5).
The difference in pH values obtained with the two different methods can be attributed to
interferences such as tissue autofluorescence and/or dye bleaching for MSFI images.
Nonetheless, both methods revealed tumor pH values consistent with pH ranges reported in
literature (e.g. pH 6.3 – 6.9, depending on the tumor model, cell line and measurement method)
[436]. The ex vivo tumor tissue pH measurements (Fig. 5.5b) revealed that A-MnO2 NPs can
quickly decrease tumor acidosis (i.e., within 30 min), most probably by quenching excessive
protons produced by cancer cell glycolysis [422].
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Figure 5.5. Effect of A-MnO2 NPs on tumor pH: (a) Ex vivo pH imaging of solid tumors i.t.
treated with A-MnO2 NPs or saline (control). Tumor pH was determined with by multispectral
fluorescence imaging (MSFI) using a pH-sensitive fluorescent dye (SNARF-4F). The bottom
insert shows the pH scale obtained with biological phantoms. (b) Tumor pH after treatment with
A-MnO2 NPs or saline (control) (n =3). Tumor pH was determined ex vivo both by MSFI (black
bars) and with a microneedle pH probe (gray bars). Error bars represent standard errors of the
mean. *statistically significant increase (*p = 0.004, **p = 0.007) in pH as compared to saline
(control) treated group.
4.7 Prolonged regulation of tumor hypoxia, HIF-1α and VEGF is related to extended tumor retention of A-MnO2 NPs
The extended retention of A-MnO2 NPs in solid tumors is evidenced in Fig. 5.6a. We injected
near-infrared dye-labelled NPs into orthotopic EMT6 breast tumors.In vivo fluorescence imaging
data (Fig. 5.6a) showed substantial diffusion of NPs within the tumor tissue almost immediately
after the injection and prolonged retention of NPs within the tumors for at least 24 h, followed by
gradual clearance from the tumors over 24-48 h. The A-MnO2 NPs are expected to be cleared as
MnO2 NPs which can be completely consumed by H2O2 (see Supporting Information Fig. S3)
and thereafter the remaining BSA/PAH complex is expected to undergo enzymatic degradation
144
by specific proteases, via an already established mechanism for the fate of polyelectrolyte
complexes taken by cells [450, 451].
Figure 5.6.Tumor retention of A-MnO2 NPs and effect on tumor hypoxia, HIF-1α and
VEGF. (a) Representative optical images of EMT6 tumor-bearing mouse with i.t. injected
near infrared-labelled A-MnO2 NPs at various times. (b) Representative
immunohistochemistry in continuous sections from EMT6 tumors treated i.t. with saline
(control) or A-MnO2 NPs for 30 min, 60 min and 24 h. Tumor hypoxia was determined by
pimonidazole binding HIF-1α and VEGF antibody. Scale bars correspond to 85 µm. (c-e)
Quantification of tumor hypoxia, HIF-1 and VEGF after treatments, determined from
classified images (not shown). (n=3). Error bars represent standard errors of the mean.
*statistically significant difference (*p = 6.9E-5, **p = 0.003, ***p = 4.5E-4) as compared
to saline (control) treated group.
145
We then investigated the effect of A-MnO2 NPs on tumor hypoxia in vivo within the 24 h period
post NP-injection by immunohistochemistry to directly measure tissue hypoxia using a
pimonidazole marker [452] and the expression of HIF-1α and VEGF (Fig 5.6b) using antibodies.
Unlike the PA experiments that measure hemoglobin-related vascular sO2,
immunohistochemistry of ex vivo tissues from animals injected with pimonidazole prior to
sacrifice directly detects the presence of hypoxic tumor cells. We found that tumors treated with
A-MnO2NPs for 30 min, 60 min or 24 h showed 24, 45 and 37% less tissue hypoxia,
respectively, as compared with the saline control (Fig. 5.6c), suggesting a time-dependent and
sustained effect of NPs on tumor hypoxia. The same tumors also showed a 19, 21 and 10%
decrease in the expression of HIF-1 (Fig. 5.6d), and 7, 65 and 65% decrease in the expression
of VEGF (Fig. 5.6e), after 30 min, 60 min and 24h treatment with A-MnO2NPs, respectively.
HIF-1 is a master regulator of the transcriptional response to acute and chronic hypoxia [452,
453], while VEGF is involved in cancer cell metabolism, angiogenesis, invasion, metastasis and
apoptosis [60]. Over-expression of VEGF is a hallmark of tumor angiogenesis [60]. Based on the
impact of angiogenesis on cancer progression and treatment, several anti-angiogenic agents
including anti-VEGF molecules are now in clinical trials as a sole treatment or in combination
with conventional cancer chemotherapy [453]. Thus downregulation of HIF-1 and VEGF
expression would improve tumor prognosis. The results presented above show that the
effectiveness of A-MnO2 NPs on the regulation of the TME is not limited to the transient
increase of tumor oxygenation and pH; they also have an effect on the down-regulation of
hypoxia-responsive protein expression that plays an important role in biological behavior and
therapeutic response of many types of cancers [230, 419]. Because the expression of HIF-1 is
146
transient depending on the relative rate of synthesis (via an O2-independent pathway) and
degradation (via an O2-dependent pathway) [454], the sustained in situ production of O2 by A-
MnO2 NPs is beneficial to prolonged regulation of TME especially impacting on the expression
of downstream proteins, such as VEGF-Error! Reference source not found..
4.8 A-MnO2 NPs enhanced anti-tumor effect of radiation
Various studies have shown that hypoxic cells in solid tumors are two-to-three times more
resistant to a single dose of ionizing radiation than those with normal levels of oxygen [140,
418]. To explore whether in situ oxygen production by A-MnO2 NPs can enhance RT, we
conducted preliminary studies in an in vivo orthotopic murine breast tumor model. EMT6 tumors
were treated with A-MnO2 NPs or saline 30 min prior to irradiation. A significant tumor growth
delay was observed in mice treated with the combination of A-MnO2 NPs and RT (Fig. 5.7a)
compared to the control groups. The average tumor volume in the A-MnO2 NPs + RT group at
day 5 remained at ≈78 mm3 while the RT alone group (treated with saline + RT) reached an
average tumor volume of ≈ 231 mm3 at the end of day 5 after treatment. Tumor weight was also
significantly lower in the A-MnO2 NPs + RT group (Fig. 5.7b). Interestingly, a decreased tumor
growth was observed in the A-MnO2 NP alone treated group compared to the saline group (non-
irradiated controls) (Fig. 5.7a). This moderate antitumor effect may be attributable to the
manipulation of the TME by the A-MnO2 NP formulation which reduces VEGF levels by 65%.
A more in-depth study will be conducted in the future to further investigate this observation.
147
Fig. 5.7. Effect on tumor growth after treatment with radiation and A-MnO2 NPs: Tumors
(n=3/ group) were treated with intratumoral injection of 1) Saline, 2) Saline+RT, 3) A-
MnO2 NP, 4) A-MnO2 NP + RT. A radiation dose of 10Gy was given 30 minutes after
saline or A-MnO2 NP treatment. (a) Tumor volume measured over time after treatment. (b)
Ex vivo measurement of tumor weight at the end of Day 5. (c) Quantification of % necrotic
+ apoptotic area in tumors after treatment. (d) Quantification of DNA DSBs as measured by
γ-H2AX staining in tumors after treatment. (e) Quantification of DNA DSBs determined by
measuring % of positive γ-H2AX cells in tumor tissue implanted in dorsal window chamber
and treated with Saline and A-MnO2 NPs. (f) Representative image of tumor implanted in
dorsal window chamber and treated with Saline and A-MnO2 NPs, and
immunohistochemical image of tumor tissue stained with γ-H2AX.*statistically significant
difference (*p < 0.05) as compared to all other groups.
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To confirm that the effect of A-MnO2 NP on the enhancement of radiation response was
due to tumor cell cytotoxicity, the percentage of tumor apoptotic and necrotic areas was
determined. Tumors treated with A-MnO2 NP + RT showed a significantly higher tumor cell
death (71%) compared to the saline + RT treated group (40%) (Fig. 5.7c). We further evaluated
radiation-induced DNA double strand breaks (DNA DSBs) by using gammaH2AX staining to
stain for DNA DSBs. The DNA DSB is considered the most lethal type of damage induced by
ionizing radiation and is a major indicator of the efficacy of treatment [455]. Combined
treatment with A-MnO2 NPs and irradiation resulted in increased DNA damage (71%) compared
to the saline control with irradiation (28%) in the EMT6 solid tumor (Fig. 5.7d). A window
chamber mouse model bearing tumor [437] was employed to determine the induction of DNA
DSB after treatment with radiation combined with A-MnO2 NPs or saline (Fig. 5.7e-f). Spatially-
localized focal x-ray irradiation was performed on half of the tissue in the transparent window
chamber allowing us to determine the relative effect of treatment in the same mouse. Increased
DNA DSBs were observed when the tumor was treated with both A-MnO2 NP and RT versus
radiation alone. Oxygen close to DNA is known to react with radiation produced radicals in
DNA “fixing” them in a state that is difficult for the intrinsic cellular DNA repair mechanisms to
deal with [229]. Therefore, it is likely that oxygen generated by reaction of A-MnO2NP with
H2O2 in tumor tissue facilitated the oxygen effect causing more tumor cell death upon radiation
thereby leading to an enhanced delay in tumor growth.
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5 Conclusions
We have demonstrated a completely new and innovative application of A-MnO2 NPs for the
modulation of the TME. The intratumoral treatment of murine breast tumors with A-MnO2 NPs
resulted in simultaneous attenuation of hypoxia and acidosis in solid tumors in vivo. Moreover,
for the first time a bioinorganic nanoparticle system has been demonstrated to promote down-
regulation of crucial tumor progression-related factors, i.e., HIF-1α and VEGF. In addition we
have demonstrated the application of A-MnO2 NPs for enhancement of radiation induced tumor
growth delay and cancer cell death. This work suggests a great potential of the A-MnO2 NP
system to improve cancer therapy (i.e.,RT) by regulating multiple attributes of the TME
simultaneously.The in vivo results obtainedin the present work encourage continuing efforts for
the optimization and application of MnO2 NPs in combination with other cancer treatments such
as chemotherapy and photodynamic therapy (PDT). Such applications are the subject of future
research by our group.
6 Acknowledgements This work was financially supported in part by a Discovery Grant to X.Y.W. from the Natural
Sciences and Engineering Research Council (NSERC) of Canada (2008-2013 - #RGPIN 170460-
08). The radiation experiments were supported by an Innovation Grant from the Canadian
Cancer Society (2013_2015, Grant No. 702133) to X.Y.W. (PI), R.S.D.
(co-PI), and A.M.R. (co-applicant). The Ontario Graduate Scholarship (2012), Ben Cohen Fund,
University of Toronto Open Scholarship (2011-2012) and Pfizer Graduate Scholarship (2012) to
P.P., NSERC summer research to A.I. We also acknowledge Wendy Xiong and Dr. Ping Cai for
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the technical support. R.S.D. is supported, in part, by a Cancer Care Ontario Research Chair. The
authors declare no competing financial interests.
7 Supporting information Further experimental details regarding NP characterization and the pKa calibration for SNARF
in tissue like phantoms is described in the Supporting Information. This material is available free
of charge via the Internet at http://pubs.acs.org.
7.1 pKa Calibration forSNARF in tissue like phantoms.
SNARF-4F 5-(and-6)-carboxylic acid (Life Technologies, USA, #S23920) is dual imaging pH
sensitive fluorophorethat allows the measurement of pH values in solution as well as in
biological tissue. The typical fluorescence emission spectra of SNARF shift from green-yellow
to red when the pH changes from acidic to alkaline. The ratio between the two fluorescence
intensities, typically at 580 nm and 640 nm, provides quantitative information about the pH
values. For the quantitative measurement of pH using SNARF, calibration curves must be
obtained under the similar conditions at which the pH values are to be determined as the pKa
values of the dye are sensitive to the local environment. In the present studies, the calibration of
SNARF was performed using tissue-like phantoms prepared in pH range 4-10, following an
established protocol [434, 435].
In details, buffers were prepared using the following buffering systems: potassium hydrogen
phthalate/hydrochloric acid from pH 3-4, potassium hydrogen phthalate/sodium hydroxide from
pH 4.5-6, potassium dihydrogen phosphate/sodium hydro oxide from pH 6-8 and disodium
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hydrogen phosphate/hydrochloric acid from pH 8-10. pH values for all buffers were measured
using Acumet AB15 pH meter (Fisher Scientific, US).
Phantoms were prepared by heating 10%(wt/v) gelatine in deionised water at 500C. Once gelatin
melted, the temperature was reduced to 40°C and hemoglobin (Sigma #H2625) and intra lipid
(Sigma #I141) were added to give final concentration of 42.5µM and 1% (wt/v), respectively.
After few minutes of stirring, one part of gelatin solution was mixed with three parts of 1 mM
SNARF solution prepared in different pH buffers (pH 4-10) with a pH increment of 0.5 unit1. In
a last step, 200µL of the final mixture was transferred to a 96 wells microplate and placed in
fridge for solidification. Biological phantom were prepared in triplicate for each pH value.
Fluorescence images of the microplates were recorded using a Xenogen microscope (Xenogen
IVIS Spectrum, Caliper Life Sciences Inc., USA) with two different filter channels; the
excitation wavelength for each channel was 535 nm whereas the emission intensities were
recorded at 580 nm (green channel) and 640nm (red channel).
For the calculation of the pKa values, the fluorescence intensities Ig(580nm) and Ir(640nm) were
measured using Image-J programby drawing the region of interest (ROI) on the obtained image
for each pH value(Fig. S2a).The ratio (R) of the intensities Ig/Ir was then calculated and plotted
against the respective pH values (Fig. S2b), and the R curve was then fitted using the Boltzmann
function (Fig. S2.c).
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In the above equation (Eq.1), R is the measured ratio of fluorescence intensities at each pH
value, Ra is the value of the R curve at acidic pH which is considered as the acidic titration
endpoint, and Rb is the value of the R curve at alkaline pH which is considered as the basic
titration endpoint. The parameter pHinfl is the point of inflection of the R curve, i.e. the pH value
at which the slope of the curve is maximum, ΔpH is an indicator for the slope at the point of
inflection. From the fitting of the R curve the fit parameters Ra, Rb, pHinfl and ΔpH were
determined.
In the next step pKa value of SNARF was calculated using the following equation (Eq.2),
Eq. 2
The –log term in equation2 was calculated for all pH values from pH4-10, the value Rin –
log term correspond to each point in Boltzmann fitting curve, whereas parameter RaandRb
were obtained from fitting of equation1. The two other parametersIa(λ2)and Ib(λ2)in –log
term correspond to the fluorescence intensities of SNARF obtained from images using image-J
program at 640nm (red channel) at acidic and alkaline region respectively for the fitted Rcurve
(in the present study we took intensities at pH4.5 and pH 8.5). At the end, graph was plotted for–
b
inf
ba R
ΔpH
pHpHexp1
RRR(pH)
(2)
2
a
)b(
a
ba
I
Ix
RR
RRlogpKpH
Eq. 1
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log versus pH that would give a straight line (see Eq.1) and intercept of linear fit is pKa.
The calculated pKa value was 6.39.
Figure S2. Calculation of pKavalue for SNARF in biological phantoms.(a) Fluorescence images
of SNARF containing phantoms at different pH values, recorded using Xenogen microscope
with a excitation wavelength of 535nm (row(i) green channel, row(ii) red channel and row(iii) is
overlay of two channel).(b) The ratios (R=Ig/Ir) versus pH graph, the ratio R was calculated from
the values Ig and Ir which were obtained from fluorescent imagein (a) using image-J program. (c)
The Boltzmann fit of data points R using Eq.1, the values for fit parameter were Ra= 1.47,
Rb=0.60, pHinfl = 6.25, and ΔpH = 0.3.(d) Shows graph of - log(...) term versus pH, the ratio R
was obtained from the Boltzmann fit in (c) andIa(λ2) and Ib(λ2)are fluorescence intensities at
640nm (red channel) obtained from image in (a) at pH4.5 and pH8.5using image-J program,
Ia(λ2)= 41.92 and Ib(λ2)= 63.11. Finally the intercept of linear fit of data points in (d)ispKa
according to Eq. 2. The obtained pKa of SNARF was 6.39.
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7.2 Consumption of A-MnO2 Nanoparticles by H2O2:
Figure S3. Upon reaction with hydrogen peroxide for the production of molecular oxygen MnO2
nanoparticles are consumed. In the graph we show the consumption of the MnO2 NPs (90 µM)
by various endogenous concentrations of H2O2 (up to 1 mM). For the experiment, H2O2 was
added to A-MnO2 in saline, incubated for 5 min at room temperature and the absorbance of the
MnO2 was measured at 360 nm.
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7.3 Nanoparticle structure and MnO2 quantification:
Figure S4. Two different NP systems are described in the manuscript: NP #1 (MnO2) - small
(~15 nm) MnO2 NPs stabilized with a positively charged polyelectrolyte (PAH) and NP #2 (A-
MnO2) - complex formed between NP #1 and BSA (~50 nm). For all in vitro and in vivo
experiments, NP #2, named A-MnO2 NPs, were used. In NP #1 MnO2 is stabilized by PAH
polymer, while in NP #2 several NP #1 particles are entrapped in a PHA/BSA complex due to
strong electrostatic interaction between the protein and the polymer. The complex formation was
confirmed by zeta potential analysis and TEM. As shown in the TEM picture above, several
small MnO2 NPs can be identified within the protein/polymer complex.
We have estimated 100% loading of the MnO2 NPs in NP #2. UV-Vis spectrophotometry
analysis of the supernatant indicated the absence of free MnO2 NPs in the supernatant after
centrifugation of NP #2 emulsion. The absolute concentration of MnO2molecules in the emulsion
was quantified by ICP analysis to determine the concentration of Mn2+
ions, and thereby the
concentration of MnO2 in the emulsion. We then correlated the molar or weight ratios
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MnO2/PAH or MnO2/BSA in our formulation, as expressed in molar units or w/w ratios in the
manuscript.
CONTRIBUTIONS
P.P., C.R.G., R.S.D. and X.Y.W. conceived and designed the experiments. C.R.G. and A.Z.A.
prepared and characterized the nanoparticles. A.I. executed in vitro experiments. A.Z.A., P.P.
and C.R.G. performed the in vivo pH experiments. P.P. designed and performed in vivo
experiments. A.M. assisted with the photoacoustic measurements and RT experiments. A.M.R.
provided advice on the hypoxic chamber design, experiments with hypoxic cancer cells and RT.
R.S.D. provided advice on the photoacousticand RT experiments. C.R.G., A.Z.A. and P.P.
performed the data analysis. C.R.G. prepared the illustrations. C.R.G., P.P., A.Z.A., R.S.D. and
X.Y.W. wrote the manuscript. All authors have read the final manuscript.
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Chapter 6 Overall conclusions, major contributions and Future
Perspectives
1 Overall Conclusions
MDR of cancer cells is a potentially surmountable obstacle to effective chemo and radiation
therapy of cancer. The mechanisms of drug resistance are multiple and often combinatorial,
allowing the cancer cells many escape routes to survive cancer therapy. Resistance to therapies
may be caused by alterations in the intracellular machinery of cancers cells themselves or
associated with the anatomical and physiological properties of the tumor, resulting in decreased
sensitivity to anti-cancer agents or radiation [194, 195]. The goal of this work was to overcome
cellular and non-cellular MDR mechanisms to enhance chemo- and radiation therapy in breast
cancer.
The most frequent occurring causes of MDR include the up-regulation of membrane bound ATP-
binding cassette transporters including P-gp, MRP-1 and BCRP [195, 196, 285]. These transport
proteins serve as energy dependent drug efflux pumps exporting anticancer drugs out of cells
against a concentration gradient [195, 196, 285]. Previously, PLN, as a novel drug delivery
system was developed to overcome multiple membrane efflux pumps and enhance Dox-MMC
toxicity against multidrug MDR breast cancer cells [193, 278]. In this thesis, we further assessed
the ability of PLN loaded with Dox-MMC (DMsPLN) to overcome multiple types of MDR
efflux pumps resulting in the control and/or reduction of both solid tumor growth in human
breast cancer models and, for the first time, the control of tumor metastasis using RGD
conjugated PLN.
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In Chapter 2, DMsPLN were assessed in vitro against human MCF-7 breast cancer cells that
were either chemosensitive (MCF7/WT) or chemoresistant (MCF7/MX, BCRP+) and
(MCF7/VP, MRP1+) human breast cancer cells. BCRP and MRP1 drug efflux pumps which are
one of the most frequently occurring causes of drug resistance in cancer therapy are
overexpressed in the MCF7/MX and MCF7/VP cell lines, respectively. It was demonstrated
DMsPLN were 20-30 fold more effective in killing the resistant cells (MCF7/MX and
MCF7/VP) than free drugs, achieving 99% cell kill. Significant enhancement of cell kill in the
resistant cell lines using DMsPLN was attributable to the ability of the PLN to be efficiently
taken up by the cells and localized near the perinuclear region of the cells, allowing simultaneous
delivery of Dox and MMC near its intended site of action (i.e., DNA). Simultaneous delivery of
Dox and MMC is important as it is postulated that they produce synergism by induction of DNA
double strand breaks via a tri-valent topoisomerase IIα-Dox-DNA complex [191, 193].
Therefore, the use of the PLN delivery system protected the drugs from being pumped out of the
cell by membrane transporters and delivered cytotoxic levels of Dox and MMC to achieve high
cell kill. P-gp inhibitors have been developed for the circumvention of MDR in cancer patients;
however these inhibitors have not reached routine clinical use due to unacceptable toxicity [296,
298]. In addition, these inhibitors failed to demonstrate statistically significant positive outcomes
in overall survival in patients [296, 298]. Inhibition of one transporter may be insufficient to
reverse chemoresistance because a single anticancer agent can be a substrate of multiple efflux
transporters. Therefore the ability of DMsPLN to overcome multiple membrane transporters (P-
pg, BCRP and MRP) to deliver the synergistic anti-cancer agents Dox and MMC may be a more
159
beneficial therapeutic approach than inhibiting a single type of efflux pump to improve
chemotherapeutic efficacy.
In chapter 3, we utilized the anatomical and physiological characteristic of solid tumor which are
known to possess “leaky” tumor neovasculature and malfunctioning tumor lymphatics [14, 24],
which enables accumulation of nanoparticulate therapeutics in tumor tissue by passive targeting via
the enhanced permeability and retention (EPR) effect. Therapeutic efficacy of DMsPLN was
assessed relative to the clinically applied PEGylated liposomal Dox (PLD) formulation in sensitive
(MDA-MB 435/LCC6/WT) and P-gp overexpressing MDR (MDA-MB435/LCC6/MDR1) human
breast tumor xenografts. DMsPLN treatment resulted in significantly higher efficacy than clinically
used PLD in both tumor models. The mice bearing human breast tumors showed a 108 - 151%
increase in tumor growth delay, 210 – 316% increase in life span, and a 10-20% de facto cure after
treatment with single or 4 DMsPLN. The superior anti-tumor efficacy of the DMsPLN group is
likely attributable to both the passive targeting of the PLN to the tumor tissue [280, 345] as well as
the efficient cellular uptake and perinuclear trafficking of PLN and the synergistic action of Dox
and MMC in cancer cells [193, 328]. Encapsulation of Dox-MMC in the same nanoparticle carrier
allows the delivery of this synergistic drug combination to the tumor site at a predetermined drug
ratio, which cannot be done with free drug cocktails in vivo. The nanocarriers are able to bypass P-
gp efflux pumps and deliver Dox and MMC simultaneously to the site of drug action, i.e., DNA in
the nuclei, resulting in increased DNA double strand breaks thus overcoming several cellular
mechanisms of MDR [191, 193, 328].
In chapter 4, we have further functionalized the surface of PLNs with cyclic Arg-Gly-Asp
(cRGD) to interact with αvβ3 integrin receptors overexpressed on tumor neovasculature and
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breast cancer cells [377, 389], to enhance the targetability and tumor retention of the
nanoparticles. Biodstribution studies showed increased accumulation and retention of RGD
targeted nanoparticles compared to the untargeted particles in mouse lungs bearing metastatic
triple negative MDA-MB 231-luc-D3H2LN breast tumors. At a low dose (3 mg/kg), both the
nanoparticle formulations (DMsPLN and RGD-DMsPLN) showed modest effects in inhibiting
tumor growth. However, the synergistic cytotoxic effects of free Dox-MMC as observed in vitro
[191, 193, 328] did not show any advantage over the saline control treated mice, attributable to
the huge difference in their pharmacokinetics profile. Encapsulating Dox and MMC within the
same nanoparticle not only allowed simultaneous release of both drugs at the site of action but
also allowed mitigation of the systematic toxicity usually observed with Dox administration
[324]. Therefore, using nanoparticles we were able to deliver a Dox dose of 15mg/kg, however
owing to the extreme acute toxicity observed at higher doses of free Dox-MMC, we were only
able to evaluate the efficacy of free Dox-MMC at 3 mg/kg. Both nanoparticle systems (DMsPLN
and RGD-DMsPLN) demonstrated a significant higher efficacy in controlling the metastatic
growth of tumor at 15 mg/kg dose. RGD conjugated DMsPLN (RGD-DMsPLN), at a 15 mg/kg
Dox dose, had an even more profound effect and exhibited the highest therapeutic effect,
resulting in 106-fold and 2.6 fold improvement in controlling lung metastasis burden compared
to free Dox-MMC and DMsPLN treatment, respectively.
In addition to cellular mechanisms of resistance, there are factors related to the tumor
microenvironment that can lead to decreased efficacy of both chemotherapy and RT [225, 226].
Hypoxia (insufficient oxygenation), acidosis (low extracellular pH, pHe) and increased rates of
ROS are common characteristics of the solid TME and together are responsible for several
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factors related to cancer biology and cancer progression and resistance to therapies [226, 231,
232]. To date, there is no treatment available for comprehensive modulation of the TME.
Approaches so far have been limited to regulating hypoxia, acidosis or ROS individually,
without accounting for their interdependent effects on tumor progression and response to
treatments. Hence in chapter 5, we have engineered new nanoparticles composed of a
polyelectrolyte-albumin complex and MnO2 nanoparticles (A-MnO2 NPs) and utilized the
reactivity of MnO2 towards peroxides for modification of the TME with simultaneous oxygen
generation and pH increase. Intratumoral treatment with A-MnO2 NPs simultaneously increased
tumor oxygenation by 45% while increasing tumor pH from pH 6.7 to pH 7.2 in vivo.
Intratumoral treatment with NPs also led to the down-regulation of two major effectors of tumor
progression and aggressiveness in the tumor, i.e., HIF-1α and VEGF. The ability of A-MnO2 NPs
to generate oxygen was further proposed to enhance RT as the effect of RT on hypoxic cells can
be enhanced by molecular oxygen which potentiates radiation damage to DNA resulting in cell
death [138, 139]. Combination treatment of the tumors with NPs and ionizing radiation
significantly inhibited breast tumor growth, increased DNA double strand breaks and cancer cell
death as compared to RT alone in a murine breast tumor animal model.
2 Major contributions of this thesis
This thesis has proposed nanotechnology-based strategies to tackle the complex issue of tumor
resistance to therapies, thereby enhancing the effect of chemo and radiation therapy in breast
cancer. The present work has made several original contributions: (1) demonstration of
synergism between Dox and MMC resulting in enhanced cytotoxicity in MRP1 and BCRP-
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overexpressing MDR breast cancer cells, and the ability of DMsPLN to overcome multiple
membrane efflux pumps (MRP1 and BCRP), (2) demonstration of enhanced in vivo efficacy and
reduced system toxicity of Dox-MMC loaded PLN (DMsPLN) superior to clinically used
PEGylated liposomal doxorubicin (PLD) in both sensitive and resistant orthotopic breast tumor
model, (4) discovery of anti-angiogenic effect of DMsPLN in solid tumors, (5) demonstration of
the ability of integrin-targeted RGD-DMsPLN to significantly reduce lung metastases without
producing drug-associated systemic toxicity, (6) development and characterization of
multifunctional MnO2 nanoparticles (A-MnO2 NPs), (7) demonstration of ability of A-MnO2 NPs
to generate oxygen and decrease pH both in vitro and in vivo, to downregulate HIF-1 and VEGF
and to enhance radiation treatment in solid breast tumor model.
3 Future Directions
3.1 Delivery of Dox-MMC using nanoparticles in other metastatic
models of breast cancer
The validation of our findings in other models of metastatic breast cancer is warranted. Studies
presented in chapter 3, show the advantages of RGD-DMsPLN in a metastatic model of breast
cancer established in the lungs using MDA-MB 231-luc triple negative breast cancer cells. The
model employed here used the direct introduction of cancer cells into the blood circulation which
is not a true metastatic process. While this experimental model of lung metastasis is highly useful
for preliminary proof of concept studies , it does not replicate the early events in metastasis from
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the primary tumor site and, therefore, is might not encompass the molecular mechanisms by
which breast cancer cells spread to other organs.
Therefore, it will be important to establish metastasis using other methods and evaluate the
efficacy of RGD-DMsPLN. The bioluminescent MDA-MB 231-luc cells can be implanted
othoptopically in the mammary fat pad of the mice and allowed to develop spontaneous
metastasis to other organs as described previously [456]. Once metastasis has occurred, as
confirmed by bioluminescent imaging, ICG loaded RGD PLN can be injected intravenously to
determine co-localization of the nanoparticle with the metastatic site. After confirmation of
nanoparticle localization, drug loaded RGD-DMsPLN can be administered intravenously and
reduction in metastasis can be evaluated using bioluminescent and fluorescent imaging. Similar
studies can be done in a immunocompetent Balb/c mice using 4T1 cells which have been shown
to metastasize to lungs, liver, bone, and brain [457, 458]. This will be a more clinically relevant
model as it will allow us to evaluate the effect of nanoparticles on metastatic tumors developed
in an immune competent microenvironment, thus recapitulating the crosstalk between an
emerging tumor and its surroundings [457, 458]
3.2 Determine the distribution of Dox and MMC in vivo at both
macroscopic and microscopic level
Effective treatment of solid tumors with anti-cancer drug requires adequate concentrations of
drugs at the tumor site to be cytotoxic [151]. However, toxicity of anti-cancer drugs towards
164
normal tissue, limits the dose of the drug that can be administered. In order to accomplish high
tumor concentration with low systemic concentration of the drug, DMsPLN delivery system was
developed. In this thesis (Chapter 3 and 4), we have shown that encapsulation of Dox-MMC
within a nanoparticle system (DMsPLN), increased the efficacy of the drug towards the tumor
with low systemic toxicity. Previous studies have shown the distribution profile of PLN in vivo
[345, 346]; however, we don’t have any knowledge regarding the distribution and fate of Dox
and MMC once it’s administered via DMsPLN. It is important to assess the pharmacokinetic
profile and tissue distribution of Dox and MMC after the intravenous administration of DMsPLN
in vivo as the profile of the anti-cancer agents can be altered dramatically.
The pharmacokinetic profiles of Dox and MMC are completely different with elimination half-
lives (t1/2) being 7~20 hours for Dox and 7~90 minutes for MMC [403, 404]. Therefore, it is
necessary to confirm the distribution of Dox and MMC as simultaneous delivery of both drugs to
the same cancer cells are required for their synergistic action. The pharmacokinetic profile will
also allow understanding the low systemic toxicity observed with DMsPLN. The
pharmacokinetic profile of Dox and MMC can be determined by collecting blood and organ
samples (tumor, liver, spleen, heart, lung, kidney, and brain) at various time points after
administration of DMsPLN in mice bearing orthotopic breast tumor. The concentration of Dox
and its metabolite, doxorubicinol and MMC can be determined using high-performance liquid
chromatography technique.
The pharmacokinetic analysis estimates the mean concentration of the drug within the tissue.
However, it does not provide any information regarding the spatial distribution of the drug. Once
the anti-cancer drug reaches the solid tumor, it must penetrate the extravascular space to reach all
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cancer cells at a sufficient concentration to cause cytotoxicity [194, 218]. The autofluorescence
property of Dox can be used to determine the penetration and distribution of Dox in solid tumor
using immunohistochemistry as previously established [459, 460]. The distribution of Dox can
further be evaluated relative to the blood vessels and hypoxic areas in the tumor using CD31 and
EF5 markers for blood vessels and hypoxia, respectively. Studies of spatial distribution of Dox
and MMC are required to complement pharmacokinetic data in order to better understand and
predict drug effects and toxicities.
Therefore, the results from these studies will allow to further modify the nanoparticle system to
achieve high drug concentration within the tumor tissue resulting in increased cytotoxicity.
3.3 Optimizing the time of MnO2 NP administration prior to
irradiation
Hypoxia, a characteristic of the tumor microenvironment (TME), has been shown to contribute
to the resistance to RT [139, 418]. In chapter 4, we utilized the reactivity of MnO2 NPs towards
H2O2 for the simultaneous and sustained production of oxygen and pH increase. We further
demonstrated that Intratumoral treatment of MnO2 NPs led to the downregulation of two major
regulators in tumor progression and aggressiveness, i.e., hypoxia-inducible factor-1 alpha (HIF-
1α) and vascular endothelial growth factor (VEGF) in the tumor. In Chapter 4, we conducted
preliminary studies in an in vivo orthotopic murine breast tumor model to explore whether in situ
oxygen production by A-MnO2 NPs can enhance RT. We demonstrated that combination
treatment of the tumors with NPs and ionizing radiation significantly inhibited breast tumor
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growth, increased DNA double strand breaks and cancer cell death as compared to RT alone
when A-MnO2 NPs are administered 30 min prior to irradiation.
As a next step to further improve the efficacy of RT, it would be beneficial to determine the
optimal time of A-MnO2 NPs required prior to irradiation to get enhanced cell kill. The
optimization studies can be first conducted in murine breast cancer EMT6 cells grown in hypoxic
conditions to mimic tumor microenvironment. Cells will be treated with A-MnO2 NPs for 15
min, 30 min, 45 min, 60 min, or 120 min prior to irradiation at 2 Gy. Clonogenic assay can be
employed to evaluate the effect of RT + A-MnO2 NPs on cell kill.
3.4 Application of MnO2 nanoparticles for enhancement of
Chemotherapy Therapy
As previously discussed, the microenvironment of solid tumor is slightly acidic (extracellular
pH, pHe < 6.9) unlike normal tissues, influencing several factors related to cancer biology and
progression and resistance to therapies [226, 232]. The newly formed blood vessels of a growing
tumor is dilated, disorganized, irregular and tortuous with discontinued or absent basement
membrane, resulting in insufficient supply of various nutrients, including oxygen to tumor cells
[60, 220]. The low levels of oxygen, or , within the tumor not only causes the tumor to undergo
glucose metabolism through the glycolytic pathway, thereby producing lactic acid but also
activates carbonic anhydrase, which converts CO2 and H2O molecules to carbonic acid [56, 59,
140]. Both of these mechanisms culminate in the accumulation of acidic metabolic products in
the extracellular space (i.e., H+ and lactate), rendering a mildly acidic interstitial pH (pHe < 6.9)
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[55, 227]. The reduced pH within the tumor can cause resistance to chemotherapy
(chemoresistance) as many cancer drugs, such as Dox , are mildly basic (pKa > 7.5. Their
protonation in the extracellular space of tumors may decrease the ability of the drug to permeate
through the cell membrane (ion trapping phenomenon) [225]. In addition, the hypoxic
environment upregulates HIF1 which induces ABC transporter gene expression, allowing cancer
cells to efflux the anti-cancer agents from inside the cell, making the drug ineffective [56, 230].
In order to increase pH within the tumor, MnO2 NP will be utilized. They will increase local pH
by the consumption of H+ as demonstrated in Chapter 4. In addition, production of oxygen by
MnO2 NP will decrease hypoxia, and prevent the switch to anaerobic respiration, thereby
reducing production of lactic acid which also contributes to the acidic environment. Reduction in
the hypoxic environment will also decrease activation of HIF1 and further reduce expression of
the ABC transporter. MnO2 NP in combination with Dox will be utilized to determine the effect
of modulating tumor microenvironment on chemosensitivity. The proof of concept study should
be conducted in vivo by first administrating MnO2 NP intratumoraly, and if positive results are
obtained, followed by Dox injection i.v. The increase in pH with a reduction in ABC transporters
should result in increased cytotoxicity of Dox towards cancer cells.
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Appendix
Appendix 1: To elucidate the mechanism of cellular uptake and
intracellular transport of fatty acid-based nanoparticles in cells
The primary mechanisms of intracellular transport of most nanocarriers are the pathways of
endocytosis, including clathrin-mediated endocytosis, caveolae-mediated endocytosis,
macropinocytosis and other clathrin and caveolae-independent vesicular pathways, as well as
phagocytosis in specialized cells (e.g. macrophages) [461, 462]. Intracellular lipid chaperones
known as fatty acid binding proteins (FABPs) serve as transporters for fatty acids and other
hydrophobic molecules within specific compartment in the cells such as lipid droplets[463, 464].
FABPs are abundantly expressed in tumor. Elevated FABP levels have been shown to promote
cancer cell proliferation, migration and poor prognosis in patients [465, 466]. These proteins
reversibly bind hydrophobic ligands, such as saturated and unsaturated long-chain fatty acids,
eicosanoids and other lipids, with high affinity.
Previous research in our lab has suggested that SLN may overcome cellular drug resistance by
bypassing the membrane associated transporters via endocytosis [402]. The objective of this
study is to further elucidate the mechanisms of cellular uptake and determine the dynamics of
intracellular trafficking and delivery of SLN as drug efficacy relies on effective delivery of SLN
to cell interior. Since our SLN, consisting of large quantity of fatty acid, are taken up
significantly by various breast cancer cells even without specific targeting moieties, we
hypothesize that interaction between FABPs and the SLN may contribute to their cellular uptake
and intracellular trafficking
169
It was previously shown that fluorescent SLN are internalized in MDA-MB 435/LCC6/MDR1
cells after a1hour-incubation. To expand upon this initial observation, the intracellular
distribution of SLN was investigated using transmission electron microscope (TEM) on fixed
cells. Two prominent mechanisms were observed in the internalization of SLN (Fig. 6.1A). The
cell membranes formed protrusions, extended over and wrapped the SLN for uptake. Cell
membrane also formed flask shaped membrane invaginations for the internalization of SLN. This
suggests that macropinocytosis and/or phagocytosis may play a role in uptake of the SLN. We
also observed within the cell, the SLN appeared to be in membrane bound vesicles and the
vesicles appeared to move towards the perinucelar region. SLN within the vesicles were a bit
deformed once internalized. However, these vesicles were not present in cells cultured without
the SLN. In some cases, SLN were also internalized within the nucleus and they co-localized
with the nucleolus (Fig. 6.2).
Fig. 6.1 TEM images of MDA-MB 435/LCC6/MDR cells with 1h incubation with PLN at 37ºC.
Different stages of the cellular uptake process of SLN. A Arrival of nanoparticle at cell
membrane and membrane wrapping of PLN. B Internalization of SLN into the cell via cell
170
membrane penetration. C Internalization into the cell. D Movement of vesicle containing SLN
towards the nucleus. E Internalization of SLN into the nucleus. N depicts nucleus
Fig. 6.2 Transmission electron micrograph MDA-MB 435/LCC6/MDR1 cells with 1h incubation
with PLN at 37ºC. (A) SLN uptake by the nucleus (B) zoomed –in image of A to depict uptake
of SLN by the nucleus. N is the nucleus
Flouresceinamine was conjugated to myristic acid to formulate fluorescent nanoparticles. To
confirm whether the uptake of the nanoparticles was mediated by energy-dependent endocytosis,
the cells were incubated with fluorescent SLN at 37℃ and 4℃, respectively. As shown in Fig.
6.3 and 6.4, the uptake of SLN occurred successfully at 37℃ (Fig. 6.3A, 6.3B, 6.3C); however;
incubation of cells with SLN at 4℃ significantly impeded uptake (Fig. 6.4A, 6.4B, 6.4C). SLN
kept at 4℃ were not internalized and remained attached on the surface membrane while at 37℃
SLN were internalized into the cytoplasm. All three cell lines showed similar pattern of uptake.
Cellular uptake was also determined quantitatively using fluorescent spectrometer (Fig. 6.5A,
6.5B, 6.5C) which also depicted a much higher uptake at 37℃. Together, the results demonstrate
that uptake of SLN is an energy dependent process. Fig. 6.5 also depicts the difference in uptake
between different cell lines of breast cancer. MDA-MB 231 cells showed the highest uptake of
171
SLN after 1 hour incubation at 37℃.
Fig. 6.3 Cellular uptake and intracellular distribution of SLN (green) in (A) MDA-MB
435/LCC6/WT, (B) MDA-MB 435/LCC6/MDR1 and (C) MDA-MB 231 breast cancer cells.
Cells were incubated with SLN for 1 hour at 37°C and the uptake pattern was observed by
Confocal laser scanning microscope (CLSM). The left panel shows the optical microscopic
image, the middle panel shows the fluorescence image (FITC filter), and the right panel shows
172
the images merged together.
Fig. 6.4 Confocal images taken after 1 hour incubation of SLN at 4°C in (A) MDA-MB
435/LCC6/WT, (B) MDA-MB 435/LCC6/MDR1 and (C) MDA-MB 231 breast cancer cells.
The left panel shows the optical microscopic image, the middle panel shows the fluorescence
image (FITC filter), and the right panel shows the images merged together.
173
Fig. 6.5 Cellular association of SLN analyzed using fluorescent spectrometer. Graph represents
intensity of SLN fluorescence at 37°C or 4°C in (A) MDA-MB 435/LCC6/WT, (B) MDA-MB
435/LCC6/MDR1 and (C) MDA-MB 231 breast cancer cells. Higher SLN uptake was observed
in MDA-MB 231 cells at 37°C. Each data points represent mean ± SEM with n = 5.
To further understand the internalization mechanisms of SLN, we studied the effect of inhibition
of specific endocytic process on the uptake of particles and measured the internalized SLN using
fluorescent spectrometer (Fig. 6.6). When MDA-MB 231 cells were pre-treated with sucrose,
creating a hypertonic condition, known to disrupt the clathrin lattice, a 60% reduction in the
uptake of SLN was observed (Fig. 6.6). Cells were pretreated In the presence of
micropinocytosis inhibitor, amiloride (5 mM) which inhibits the Na+/ H+, SLN uptake was
reduced by almost 30% compared to the uptake by untreated cells (Fig. 6.6).
174
Intracellular localization of SLN was further investigated using confocal microscopy. The role of
lipid droplets in SLN transport was studied by using nile red which preferentially stains lipid
droplets. Nile red stained round structures within MDA-MB 231 cells (Fig. 6.7). SLN was
localized in the structures of similar size, shape, and location as those stained by nile red,
suggesting that SLN was primarily localized into lipid droplets within the cells. Co-localization
images at different optical slice (Fig. 6.7A and 6.7B) show localization of SLN with lipid
droplets.
Fig. 6.6 SLN uptake analyzed using
fluorescent spectrometer in the presence of
different specific inhibitors. MDA-MB 231
cells were pre-treated with 450mM of sucrose
for 60 min (clathrin inhibition), and 5mM of
amiloride for 10 min (inhibition of
macropinocytosis). After pre-incubation, cells
were incubated with fluorescent SLN at the
respective conditions for 1 hour at 37°C. Each
data points represent mean ± SEM with n = 5.
175
Fig. 6.7 Intracellular distribution of SLN and lipid droplets in MDA-MB 231 cells. Confocal
imaging was performed to visualize lipid droplets (red) and SLN (green) using the FITC filter.
SLN co-localizes with lipid droplets (yellow) as seen in the right panel. Images were acquired at
8µm (A) and 16 µm (B) optical slice.
B
A
176
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