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The effect of P-glycoprotein inhibition and ultrasound exposure on the cytotoxicity of taxane loaded… Wan, Chung Ping Leon 2014

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The effect of P-glycoprotein inhibition and ultrasound exposure on the cytotoxicity of taxane loaded diblock copolymer nanoparticles in multidrug resistant cells  by Chung Ping Leon Wan  B.Sc. (Molecular Biology and Biochemistry), Simon Fraser University, 2004 M.Sc. (Chemistry and Biochemistry), California State University, Long Beach, 2008  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Pharmaceutical Sciences)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  September 2014  © Chung Ping Leon Wan, 2014   ii Abstract One of the major mechanisms of multidrug resistance involves an efflux protein, P-glycoprotein (Pgp), which pumps commonly used anticancer drugs such as taxanes out of cells, leading to a decrease in cellular drug accumulation. The overall goal of this project was to develop strategies to enhance intracellular drug accumulation and cytotoxicity of nanoparticulate taxanes in multi-drug resistant (MDR) cell lines. Paclitaxel (PTX) loaded nanoparticles fabricated from micelle forming MePEG114-b-PCL19 and nanosphere forming MePEG114-b-PCL104 were compared for drug and block copolymer uptake, and cytotoxicity in drug sensitive MDCKII and drug resistant MDCKII-MDR1 cell lines. PTX loaded micelles were more cytotoxic than PTX loaded nanospheres. Co-administration of the known Pgp inhibitor, MePEG17-b-PCL5, with PTX loaded micelles or nanospheres significantly increased drug cytotoxicity in MDCKII-MDR1 cells.    Mixed molecular weight (MW) PCL200/PCL5 nanoparticles composed of long hydrophobic block, MePEG114-b-PCL200, and MePEG17-b-PCL5, were developed and characterized for the co-delivery of taxanes and Pgp inhibitor. Both PTX and docetaxel (DTX) loaded mixed MW PCL200/PCL5 nanoparticles were demonstrated to release MePEG17-b-PCL5 in a controlled release manner and increase drug cytotoxicity in MDR cells as compared to the drug loaded MePEG114-b-PCL200 nanoparticles in the absence of MePEG17-b-PCL5. The mixed MW nanoparticles remained in the plasma for longer than the drugs with approximately 3% of the injected dose remaining 24 hrs post injection.  Ultrasound irradiation was investigated as a potential strategy to enhance the cytotoxicity of PTX loaded MePEG-b-PDLLA micelles in MDR cells. Using an ultrasound regime of a single 10-second burst of high frequency (4 MHz) and high intensity (32 W/cm2) ultrasound, it was shown that ultrasound irradiation resulted in a two-fold increase in intracellular uptake of PTX in  iii drug sensitive MDCKII and MCF-7 cell lines and their respective Pgp-overexpressing MDCKII-MDR1 and NCI-ADR counterparts as compared to untreated cells (no ultrasound). The enhanced accumulation and retention of PTX resulting from ultrasound treatment translated into greater cytotoxicity in both drug sensitive and resistant cell lines. In conclusion, we have demonstrated two promising strategies for enhancing MDR cellular drug accumulation and effectiveness: the use of mixed molecular weight taxane loaded nanoparticles and ultrasound irradiation.   iv Preface  This thesis is comprised of the following 3 manuscripts, for which I am the principal author:  1. Wan, C.P., Letchford, K., Jackson, J.K., and Burt H.M. (2013). The combined use of paclitaxel-loaded nanoparticles with a low molecular weight copolymer inhibitor of P-glycoprotein to overcome drug resistance. Int. J. Nanomedicine. 9: 379-91. 2. Wan, C.P., Letchford, K., and Burt H.M. (2014). Mixed molecular weight copolymer nanoparticles for the treatment of drug resistant tumors: formulation development, cytotoxicity and pharmacokinetics. (The manuscript is in preparation for submission). 3. Wan, C.P., Jackson, J.K., Pirmoradi, F.N., Chiao, M., and Burt H.M. (2012). Increased accumulation and retention of micellar paclitaxel in drug-sensitive and P-glycoprotein- expressing cell lines following ultrasound exposure. Ultrasound Med Biol. 38(5): 736-44.  Chapter 2 is based on manuscript 1. I was the primary individual responsible for the design and conduct of the research experiments and preparation of the manuscript. Dr. Kevin Letchford helped with experimental design, conduct of experiments, and preparation of the manuscript.  Mr. Jackson helped with preparation of the manuscript. The contribution of all co-authors was through the provision of intellectual discussion and editorial assistance.  Chapter 3 is based on manuscript 2. I was the primary individual responsible for the design and conduct of the research experiments and preparation of the manuscript. Dr. Kevin Letchford performed characterization of nanoparticles and helped with manuscript preparation. The  v contribution of all co-authors was through the provision of intellectual discussion and editorial assistance.  Chapter 4 is based on manuscript 3. I was the primary individual responsible for the design and conduct of the research experiments and preparation of the manuscript. Mr. Jackson helped with experimental design and manuscript preparation. Dr. Pirmoradi and Dr. Chiao helped manufacture of the ultrasound device. All co-authors contributed through intellectual discussion and editorial assistance.  All animal work was carried out in accordance with the Canadian Council on Animal Care (CCAC) guidelines, and the animal care protocol was approved by the Animal Care Committee from the University of British Columbia (Animal Care Certificate A09-0519).  vi Table of Contents  Abstract .......................................................................................................................................... ii Preface ........................................................................................................................................... iv Table of Contents ......................................................................................................................... vi List of Tables ................................................................................................................................ xi List of Figures .............................................................................................................................. xii List of Abbreviations ...................................................................................................................xx Acknowledgements .................................................................................................................. xxiv Dedication ...................................................................................................................................xxv Chapter  1: Introduction ...............................................................................................................1 1.1 Thesis overview .............................................................................................................. 1 1.2 Multidrug resistance........................................................................................................ 6 1.2.1 Mechanisms of drug resistance ................................................................................... 7 1.2.1.1 ABC drug efflux transporters.............................................................................. 9 1.2.1.2 The structure and function of P-glycoprotein ................................................... 12 1.2.1.3 Taxanes: chemistry and pharmacology ............................................................. 15 1.3 Modulation of P-glycoprotein ....................................................................................... 17 1.3.1 Pharmacological agents as P-glycoprotein inhibitors ............................................... 18 1.3.2 Pharmaceutical excipients as P-glycoprotein inhibitors ........................................... 20 1.3.2.1 Pluronic® triblock copolymer surfactants ......................................................... 21 1.3.2.2 MePEG17-b-PCL5 diblock copolymer ............................................................... 23 1.4 Nanoparticulate taxane formulations ............................................................................ 25 1.4.1 Polyether-polyester based block copolymers............................................................ 25  vii 1.4.1.1 Taxane loaded MePEG-b-PCL nanoparticles ................................................... 27 1.4.1.2 Pharmacokinetics and biodistribution of taxane loaded nanoparticles ............. 30 1.4.1.3 Tumor uptake of taxane loaded nanoparticles .................................................. 32 1.4.1.4 Intracellular uptake and subcellular distribution of nanoparticles .................... 34 1.5 Ultrasound in cancer treatment ..................................................................................... 37 1.5.1 Ultrasound irradiation ............................................................................................... 37 1.5.2 Ultrasound mediated drug delivery ........................................................................... 39 1.5.3 Mechanisms of ultrasound enhanced cytotoxicity from drug loaded nanoparticles . 42 1.5.4 Effect of ultrasound on cytotoxicity of drug loaded nanoparticles in MDR cells .... 45 1.5.5 Ultrasound device design and characteristics ........................................................... 46 1.6 Thesis goal and research objectives .............................................................................. 47 Chapter  2: The combined use of paclitaxel-loaded nanoparticles with a low molecular weight copolymer inhibitor of p-glycoprotein to overcome drug resistance ..........................50 2.1 Introduction ................................................................................................................... 50 2.2 Materials and methods .................................................................................................. 52 2.2.1 Materials ................................................................................................................... 52 2.2.2 Synthesis and characterization of MePEG-b-PCL diblock copolymers ................... 53 2.2.3 Fluorescent labeling of MePEG-b-PCL copolymers ................................................ 56 2.2.4 Preparation of MePEG-b-PCL nanoparticles............................................................ 56 2.2.5 Immunodetection of P-glycoprotein ......................................................................... 57 2.2.6 Cytotoxicity of blank MePEG-b-PCL block copolymers ......................................... 58 2.2.7 Cytotoxicity of PTX loaded MePEG-b-PCL nanoparticles ...................................... 58 2.2.8 Drug accumulation .................................................................................................... 59 2.2.9 Polymer uptake and fluorescence confocal microscopy ........................................... 60  viii 2.2.10 Statistical analysis ................................................................................................. 61 2.3 Results ........................................................................................................................... 62 2.3.1 Immunodetection of P-glycoprotein ......................................................................... 62 2.3.2 Cytotoxicity of diblock copolymers.......................................................................... 62 2.3.3 Influence of MePEG17-b-PCL5 on cell viability ....................................................... 65 2.3.4 Cytotoxicity of PTX loaded micelles and nanospheres ............................................ 67 2.3.5 Drug accumulation studies ........................................................................................ 69 2.3.6 Polymer uptake and fluorescence confocal microscopy ........................................... 69 2.3.7 The effect of MePEG17-b-PCL5 on the cytotoxicity of PTX loaded nanoparticles .. 73 2.4 Discussion ..................................................................................................................... 78 2.4.1 Cellular biocompatibility of MePEG-b-PCL ............................................................ 78 2.4.2 Cytotoxicity of PTX loaded nanoparticles ................................................................... 79 2.4.3 The effect of MePEG17-b-PCL5 on the cytotoxicity of PTX loaded nanoparticles ..... 81 2.5 Conclusions ................................................................................................................... 83 2.6 Acknowledgments......................................................................................................... 84 Chapter  3: Mixed molecular weight copolymer nanoparticles for the treatment of drug resistant tumors: formulation development, cytotoxicity and pharmacokinetics .................85 3.1 Introduction ................................................................................................................... 85 3.2 Materials and methods .................................................................................................. 88 3.2.1 Materials ................................................................................................................... 88 3.2.2 Synthesis and characterization of copolymers .......................................................... 88 3.2.3 Preparation and characterization of nanoparticles .................................................... 89 3.2.4 Drug loading ............................................................................................................. 90 3.2.5 In vitro drug and MePEG17-b-PCL5 release.............................................................. 90  ix 3.2.6 Cytotoxicity of MePEG-b-PCL nanoparticulate formulations ................................. 91 3.2.7 Pharmacokinetics and biodistribution of nanoparticulate formulations ................... 93 3.2.8 Statistical analysis ..................................................................................................... 94 3.3 Results ........................................................................................................................... 95 3.3.1 Synthesis and characterization of copolymers .......................................................... 95 3.3.2 Formation and characterization of nanoparticles ...................................................... 95 3.3.3 Drug loading ............................................................................................................. 97 3.3.4 In vitro drug and MePEG17-b-PCL5 release.............................................................. 99 3.3.5 Cytotoxicity of MePEG-b-PCL nanoparticulate formulations ............................... 101 3.3.6 Pharmacokinetics and biodistribution of nanoparticulate formulations ................. 108 3.4 Discussion ................................................................................................................... 112 3.5 Conclusion .................................................................................................................. 119 3.6 Acknowledgments....................................................................................................... 120 Chapter  4: Increased accumulation and retention of micellar paclitaxel in drug sensitive and P-glycoprotein expressing cell lines following ultrasound exposure ..............................121 4.1 Introduction ................................................................................................................. 121 4.2 Materials and methods ................................................................................................ 124 4.2.1 Materials ................................................................................................................. 124 4.2.2 MePEG-b-PDLLA micelle preparation .................................................................. 124 4.2.3 Ultrasound treatment ............................................................................................... 125 4.2.4 Physical stability of MePEG-b-PDLLA micelles ................................................... 125 4.2.5 Fluorescence resonance energy transfer (FRET) and drug release study ............... 126 4.2.6 PTX accumulation and PTX efflux study ............................................................... 128 4.2.7 Cell proliferation study ........................................................................................... 128  x 4.2.8 Statistical analysis ................................................................................................... 129 4.3 Results ......................................................................................................................... 129 4.3.1 Physical stability, FRET analysis and drug release ................................................ 129 4.3.2 Cellular PTX accumulation..................................................................................... 133 4.3.3 Cellular PTX efflux................................................................................................. 134 4.3.4 Cell proliferation ..................................................................................................... 135 4.4 Discussion ................................................................................................................... 137 4.4.1 Physical stability, FRET analysis and drug release ................................................ 137 4.4.2 PTX accumulation .................................................................................................. 137 4.4.3 PTX efflux .............................................................................................................. 140 4.4.4 Cell proliferation ..................................................................................................... 140 4.5 Conclusion .................................................................................................................. 141 4.6 Acknowledgments....................................................................................................... 142 Chapter  5: Summarizing discussion, conclusions, and suggestions for future work .........143 5.1 Summarizing discussion ............................................................................................. 143 5.2 Summary and conclusions .......................................................................................... 154 5.3 Suggestions for future work ........................................................................................ 155 References ...................................................................................................................................157 Appendix: Supporting data .......................................................................................................184   xi List of Tables  Table 1.1 Anticancer drugs known to be substrates of ATP-binding cassette transporters (Patel et al., 2011; Szakacs et al., 2006; Gillet et al., 2007; Coley, 2010; Glavinas et al., 2004; Sharom, 2006; Ozben, 2006). Shaded cells represent drugs that are the substrates of the transporters. .... 11 Table 2.1 Characterization of synthesized MePEG-b-PCL diblock copolymers. ........................ 55 Table 3.1 Characterization of synthesized MePEG-b-PCL diblock copolymers. ........................ 96 Table 3.2 Pharmacokinetic parameters for PTX, DTX and MePEG114-b-PCL200 ...................... 110   xii List of Figures  Figure 1.1 A) The membrane topology of P-glycoprotein showing two homologous halves, each with a transmembrane domain and a nucleotide-binding domain (NBD). B) A simplified representation of the proposed mechanisms: I) Hydrophobic Vacuum Cleaner model and II) Flippase model. Pgp substrate (green triangles) partition into lipid bilayer, binds to transmembrane domain and ATP hydrolysis results in substrate efflux (I) or flipping substrate from the inner to the outer leaflet and rapid efflux (II). These figures are adapted from Sharom, 2014............................................................................................................................................... 14 Figure 1.2 Chemical structures of paclitaxel (A) and docetaxel (B) ............................................ 18 Figure 1.3 Micelles and Nanospheres formed by MePEG-b-PCL diblock copolymers and their general characteristics. .................................................................................................................. 29 Figure 1.4 Schematic representation of enhanced permeability and retention of drug delivery carriers in solid tumors.  Tumor blood vessels are characterized by leaky vasculature and malfunctioning lymphatic system, allowing drug loaded nanoparticulates to extravasate from tumor blood vessels and be retained in the tumor tissues. The figures are adapted from (Maeda, 2010). ............................................................................................................................................ 33 Figure 1.5. Cavitation effect induced by ultrasound. A) stable cavitation can form, in which bubbles oscillate without collapsing, resulting in a mild perturbation of cell membranes. B) inertial cavitation occurrs when cavitation bubbles collapse producing enormous amounts of pressure (shown by black arrows) and shock waves that can disrupt cell membranes. ................ 39 Figure 1.6 Proposed mechanisms of ultrasound irradiation mediated enhanced drug uptake. A) enhancement of drug release, B) increased rates of endocytosis, C) enhancement of membrane permeability by the formation of transient membrane pores. ....................................................... 44  xiii Figure 1.7. A) A schematic representation of a 48-well cell culture plate, adapted for exposure with ultrasound from a custom-designed ultrasonic-transducer. The height of cell culture medium in wells is around 8mm. B) A schematic representation of 4 MHz, burst-mode ultrasound waveform with 50 ms burst period. The diagrams were published (Jackson et al., 2011). .......... 47 Figure 2.1 Detection of Pgp expression in MDCKII and MDCKII-MDR1 cells by Western blot analysis. ......................................................................................................................................... 62 Figure 2.2 Cell viability of a) MDCKII and b) MDCKII-MDR1 cells in the presence of various concentrations of () MePEG114-b-PCL19 or (☐) MePEG114-b-PCL104 diblock copolymer nanoparticles. Cells were seeded in 96-well plates and treated with various concentrations of the nanoparticles for 3 days at 37°C. Cell viability was determined using an MTS assay. Indicated values are mean (±SEM) of five independent experiments (n=5). ............................................... 63 Figure 2.3 Cell lysis of (a) MDCKII and (b) MDCKII-MDR1 cells treated with () MePEG114-b-PCL19 and () MePEG114-b-PCL104 diblock copolymer nanoparticles at varying concentrations for 90 minutes. Cell lysis was measured by LDH release assay. The indicated values are the mean of three independent experiments (±SEM). ........................................................................ 64 Figure 2.4 a) Cell viability of (●) MDCKII and ()  MDCKII-MDR1 in the presence of MePEG17-b-PCL5. The cells were incubated with various concentrations of MePEG17-b-PCL5 for three days in culture medium followed by the determination of cell viability by MTS assay. b) LDH release of (●) MDCKII and () MDCKII-MDR1 in the presence of MePEG17-b-PCL5. The cells were incubated with various concentrations of MePEG17-b-PCL5 for 90 minutes in HBSS followed by the LDH release assay. The indicated values are the mean of three independent experiments (±SEM). ............................................................................................... 66 Figure 2.5 Cytotoxicity of PTX loaded () MePEG114-b-PCL19 micelles and (☐) MePEG114-b-PCL104 nanospheres in (a) MDCKII and (b) MDCKII-MDR1 cell lines after 3 days incubation.  xiv The cell viability was determined by an MTS assay. The indicated values are the mean of three independent experiments (±SEM). ............................................................................................... 68 Figure 2.6 The intracellular accumulation of PTX delivered by MePEG114-b-PCL19 micelles (black) and MePEG114-b-PCL104 nanospheres (white) in MDCKII and MDCKII-MDR1 cells.  Cells were incubated with nanoparticulate PTX (32 μg/ml) for 90 min at 37°C followed by washing and lysis using 2% TritonX-100 in HBSS.  Intracellular PTX levels were measured using liquid scintillation counting of cell lysates.  The intracellular PTX content was normalized to the total protein content as determined by BCS assay. The indicated values are the mean of three independent experiments (±SEM). ...................................................................................... 69 Figure 2.7 Intracellular concentration of TMRCA-labeled copolymers.  Cells were exposed to TMRCA-labeled micelles (white) or nanospheres (black) for 90 min at 37 °C.  After the incubation period, cells were washed and lysed with 2% TritonX-100 in HBSS.  The amount of polymer-bound or uptake into cells was determined by fluorescence spectroscopy. The indicated values are the mean of three independent experiments (±SEM). ................................................. 71 Figure 2.8 Confocal fluorescence imaging of MDCKII (a,b,e,f) and MDCKII-MDR1 (c,d,g,h) cells illustrating uptake of TMRCA-labeled MePEG114-b-PCL19 (a-d) and MePEG114-b-PCL104 (e-h) diblock copolymers after 90 min incubation.  The images on the left column are an overlay of a direct contrast signal (which shows the contour of the cells) and the fluorescence of the nucleus (blue).  The images on the right column demonstrate the overlay of fluorescence signal of the nucleus and the TMRCA-labeled diblock copolymers (red). ............................................. 72 Figure 2.9 Cell viability of MDCKII (white) or MDCKII-MDR1 (black) cells treated for 3 days with MePEG-b114-PCL19 micelles or MePEG-b114-PCL104 nanospheres loaded with 16 µg/ml PTX with or without concurrent treatment with (0.05% w/v) MePEG-b17-PCL5 or (5 μM) CsA.  xv The cell viability was determined by an MTS assay. The indicated values are the mean of three independent experiments (±SEM). * and *** indicate statistical significance (p<0.05).............. 75 Figure 2.10 The effect of MePEG17-b-PCL5 diblock copolymer on the cytotoxicity of PTX loaded- (a) MePEG114-b-PCL19 micelles and (b) MePEG114-b-PCL104 nanospheres. (●) MDCKII and () MDCKII-MDR1 were incubated with PTX loaded nanoparticles at a drug concentration of 32 μg/ml and copolymer concentration of 0.2% w/v for 90 min followed by removal of the drug loaded nanoparticle and a subsequent 3 day incubation in various concentrations of MePEG17-b-PCL5.  The cell viability was determined by an MTS assay. The indicated values are the mean of three independent experiments (±SEM) (c) A schematic illustration of experimental sequences. ..................................................................................................................................... 76 Figure 2.11 The cytotoxicity of PTX- loaded (a) MePEG114-b-PCL19 micelles and (b) MePEG114-b-PCL104 nanospheres in MDCKII and MDCKII-MDR1 cells with and without subsequent incubation with MePEG17-b-PCL5. Cells were treated with PTX-loaded micelles and nanospheres with varying concentrations of PTX for 90 min followed by removal of the drug loaded nanoparticles and a subsequent 3 day incubation with media with or without 0.05% w/v MePEG17-b-PCL5. Cell viability was determined by an MTS assay. The indicated values are the mean of three independent experiments (±SEM). (c) A schematic illustration of experimental sequences. ..................................................................................................................................... 77 Figure 3.1 (A) Mass of polymer in pellet after centrifugation of PCL200/PCL5 nanoparticles at 9500 xg for 5 minutes as a function of MePEG17-b-PCL5 added to nanoparticle dispersions. (B) Hydrodynamic diameter (▲) and zeta potential (□) of PCL200/PCL5 nanoparticles as a function of MePEG17-b-PCL5 added during formation of nanoparticles. Each point represents the average of 3 replicates  SD. ...................................................................................................................... 97  xvi Figure 3.2 PTX or DTX solubilization () and loading efficiency () for PCL200/PCL5 nanoparticles (A and B) composed of 1% w/v MePEG114-b-PCL200 prepared by emulsification with 1% w/v MePEG17-b-PCL5 or PCL200 nanoparticles prepared by nanoprecipitation and dialysis (C and D) composed of 1% w/v MePEG114-b-PCL200. Each point represents the average of 3 samples  SD. ........................................................................................................................ 98 Figure 3.3 (A) In vitro release of PTX from mixed MW PCL200/PCL5 nanoparticles () or PCL200 nanoparticles  (□) or DTX release from mixed MW PCL200/PCL5 nanoparticles (◆) or PCL200 nanoparticles (○). Release experiments were conducted in 0.01M PBS at pH 7.4 and 37C. Each point represents the average of 4 samples  SD. (B) In vitro release of MePEG17-b-PCL5 from PCL200/PCL5 nanoparticles (○ ) or MePEG17-b-PCL5 micelles (■ ). Release experiments were conducted in distilled water at 37C. Each point represents the average of 3 samples  SD. ............................................................................................................................. 100 Figure 3.4 The cytotoxicity of PCL200 nanoparticles (A) or mixed MW PCL200/PCL5 nanoparticles (B) on MDCKII cells (○) and MDCKII-MDR1 cells (■). Cells were incubated with various concentrations of nanoparticles in the presence of culture medium for 3 days. The cell viability were determined by the MTS assay. Each point represents the mean of three independent experiments (±SEM). ............................................................................................. 104 Figure 3.5 The cytotoxicity of free drug or nanoparticle encapsulated PTX or DTX on MDCKII (A and B) or MDCKII-MDR1 (C and D). Treatment groups were drug loaded mixed MW PCL200/PCL5 nanoparticles (■) or PCL200 nanoparticles (▲) or free drug (○). Cells were incubated with various concentrations of free drug or drug encapsulated in different nanoparticle formulations for 90 min followed by washing. After washing, the cells were incubated for 3 days in culture medium. The cell viabilities were determined by the MTS assay. Each point represents  xvii the mean of three independent experiments (±SEM). (E) A schematic illustration of experimental sequences. ................................................................................................................................... 105 Figure 3.6 (A) Cell viability of MDCK-MDR1 cells after 3 day incubation with free PTX (○) or free DTX (■). The effect of concentration of PCL200 or mixed MW PCL200/PCL5 nanoparticles on the cytotoxicity of (B) PTX or (C) DTX in MDCKII-MDR cells. Cells were treated with varying concentrations of PCL200/PCL5 (■ ) or PCL200 (○ ) nanoparticles (expressed as concentration of MePEG114-b-PCL200) loaded with a fixed concentration of 500 ng/ml of PTX (B) or 50 ng/ml of DTX (C) for 3 days. The cell viability was measured using MTS assay. Each point represents the average of 3 samples  S.E.M. (D) A schematic illustration of experimental sequences for panels B and C. .................................................................................................... 106 Figure 3.7 The effect of the concentration of blank (no drug) PCL200 or mixed MW PCL200/PCL5 nanoparticles on the cytotoxicity of MDCKII-MDR1 after pretreatment with PTX or DTX loaded nanoparticles. MDCKII-MDR1 cells were pre-treated with mixed MW PCL200/PCL5 nanoparticles with (A) PTX or (B) DTX loadings of 25 µg/ml or 6 µg/ml, respectively, for 90 minutes followed by washing and subsequent incubation in culture media for 3 days with different concentrations of either blank (no drug) mixed MW PCL200/PCL5 (■) or blank (no drug) PCL200 nanoparticles (○) (expressed as concentration of MePEG114-b-PCL200). The cell viability was measured using MTS assay. Each point represents the mean of three independent experiments (±SEM). (C) A schematic illustration of experimental sequences. ........................ 107 Figure 3.8 Plasma drug or copolymer concentration vs. time profiles for (A) PTX formulated as Paclitaxel for Injection (▽ ), or mixed MW PCL200/PCL5 nanoparticles () or (B) DTX formulated as Taxotere® (□) or mixed MW PCL200/PCL5 nanoparticles (■) or (C) MePEG114-b- xviii PCL200 in PTX loaded mixed MW PCL200/PCL5 nanoparticles () or DTX loaded mixed MW PCL200/PCL5 nanoparticles (■). Each point represents average of 4 samples  SD. ................ 109 Figure 3.9 Biodistribution in organs 24 hours post injection of (A) PTX formulated in mixed MW PCL200/PCL5 nanoparticles (diagonal stripes) and Paclitaxel for Injection (white) or DTX formulated in mixed MW PCL200/PCL5 nanoparticles (grey) and Taxotere® (black) or (B) MePEG114-b-PCL200 in PTX loaded mixed MW PCL200/PCL5 nanoparticles (white) or DTX loaded mixed MW PCL200/PCL5 nanoparticles (black). Each bar represents the average of 4 samples ±SD. .............................................................................................................................. 111 Figure 4.1 A schematic representation of a 48-well cell culture plate, adapted for exposure with ultrasound from an ultrasonic-transducer.  The height of cell culture medium in wells is around 8mm.  The ultrasound applied to the well resulted in the formation of stable cavitation. .......... 126 Figure 4.2 The effect of ultrasound (4MHz, 32 W/cm2, 10 second exposure) on the hydrodynamic diameter (determined using DLS) of PTX loaded MePEG-b-PDLLA micelles.  The PTX loaded micelles were treated with (solid triangle) or without (solid square) ultrasound (US).  All data points represent mean ± SD, (n=3). .................................................................... 131 Figure 4.3 a) FRET pair ratio of DiIC18 and DiOC18 within micellar core as a function of time with or without ultrasound (US) treatment (4MHz, 32 W/cm2, 10 second exposure) in release medium. The MePEG-PDLLA micelles were loaded with DiOC18 and DiIC18 and were treated with (solid triangle) or without (solid square) ultrasound. Albumin at 40 mg/ml was used in the release medium for binding FRET probes. The micellar dispersions were incubated at 37°C and the FRET ratio, IDiI/(IDiI + IDiO), was measured over time. b) The effect of ultrasound irradiation on PTX release from micelles. Micellar PTX (100 μg/ml) were exposed to ultrasound irradiation (4MHz, 32 W/cm2, 10 second exposure) and immediately placed in a dialysis insert. At different  xix time points, the PTX contents were measured using beta-scintillation counter. All data points represent mean ± SD, (n=3). ....................................................................................................... 132 Figure 4.4 Intracellular PTX accumulation with (solid bar) or without (open bar) ultrasound (US) treatment (4MHz, 32 W/cm2, 10 second exposure) in (a) MDCKII and MDCKII-MDR1 cell lines & (b) MCF-7 and NCI-ADR cell lines. Cells were incubated with micellar PTX (100 μg/ml) for 90 min at 37°C prior to ultrasound treatment. Intracellular PTX levels were measured using liquid scintillation counting. All data points represent mean ± SEM, (n=8). *Statistically significant compared to no ultrasound treatment (P<0.05). ........................................................ 133    xx List of Abbreviations  °C Degrees Celsius 14C Carbon-14 3H Tritium ABC ATP-binding cassette ACE Acetone ACN Acetonitrile ANOVA Analysis of variance Ara-c Cytosine arabinoside AUC Area under the curve ATP Adenosine triphosphate CL Clearance CMC Critical micelle concentration CrEL Cremophor-EL® CsA Cyclosporin A CME Clathrin-mediated endocytosis  Da Daltons DAPI 4’, 6-diamino-2-phenylindole dihydrochloride DiIC18 Dioctadecyl-tetramethylindocarbocyanine perchlorate DiOC18 Dioctadecyloxacarbocyanine perchlorate DMEM Dulbecco’s modified eagle’s media DMF N,N,-dimethyl formamide DMSO Dimethyl sulfoxide  xxi DPM Disintegrations per minute DTX Docetaxel EPR Enhanced permeation and retention RES Reticuloendothelial system FBS Fetal bovine serum FDA Food and Drug Administration FRET Fluorescence resonance energy transfer g Gram GPC Gel permeation chromatography HBSS Hank’s balanced salt solution HIFU High-intensity focused ultrasound H2O Water HPLC High performance liquid chromatography HUVEC Human umbilical vein endothelial cell IC50 Half maximal inhibitory concentration ISATA Spatial average temporal average intensity IV Intravenous KHz Kilohertz LDH Lactate dehydrogenase MDCKII Madin-Darby Canine Kidney MDR Multi-drug resistance MDR1 Multi-drug resistance gene MePEG Methoxy poly(ethylene glycol) MePEG-b-PCL Methoxy poly(ethylene glycol)-block-poly(caprolactone)  xxii mg Milligram MHz Megahertz ml Milliliter mol Mole ms Millisecond MW Molecular weight MWCO Molecular weight cut off NMR Nuclear magnetic resonance spectroscopy nm Nanometer PBS Phosphate buffered saline PCL poly(ε-caprolactone) PCL5 MePEG17-b-PCL5 PCL200 MePEG114-b-PCL200 PDI Polydispersity index PDLLA Poly(D,L-lactic acid) PEO Poly(ethylene oxide) Pgp P-glycoprotein PLGA Poly(lactic-co-glycolic acid) PPO Poly(propylene oxide) PTX Paclitaxel PVP Poly(N-vinyl- pyrrolidone) PZT Lead zirconate titanate R-123 Rhodamine 123 R-6G Rhodamine 6G  xxiii S.D. Standard deviation S.E.M. Standard error of the mean t1/2 Half-life TMRCA Tetramethylrhodamine-5-carbonyl azide TPGS d-alpha-tocopheryl polyethylene glycol-1000 succinate Vd Volume of distribution  % w/v Percent weight in volume % w/w Percent by weight W/cm2 Watts per square centimeter μCi Microcurie μg Microgram μl Microliter μm Micrometer μM Micromolar          xxiv Acknowledgements  This thesis would not have been possible without the support of many people. First and foremost, I would like to thank my supervisor, Dr. Helen Burt who has provided me with a stimulating lab environment in which to grow and learn, and an unwavering commitment to the success of this project. Her knowledge, dedication, enthusiasm and support have been instrumental in both my personal and professional development. I could not have asked for a better mentor. Thank you to John and Dr. Kevin Letchford, who have always been there to be my supportive friends and provide technical expertise and insightful scientific discussion. My sincere thanks to my committee members, Dr. Marcel Bally, Dr. Urs Hafeli, Dr. Ujendra Kumar, and Dr. Kishor Wasan for their effort, thoughtful discussions and time.  I am grateful for the technical expertise of Irina Manisali for her help with the pharmacokinetic studies, Dr. Padmesh Rajput for help with the cell uptake studies, Dr. Steve Yin for his help with the synthesis of the polymers, Dr. Markus Heller for his help with the NMR, and the Center for Drug Research and Development (CDRD) for allowing me access to their equipment. Thank you to Karanvir Sall, Nathan Wong, Winnie Ye, and Donna Leung for their hard work through the summers. To all my past and present lab mates: Sam Gilchrist, Clement Mugabe, David Plackett, Chiming Yang, Rakhi Pandey, and Lucy Ye. I thank you all for your friendship, support, and making the Burt Lab a great place to be. Thank you to Dr. Barbara Conway and Rachel Wu for their friendship and the great effort in keeping this faculty afloat.  To my wife, thank you for unselfishly sacrificing so much to allow me to pursue my goals and my passion.  Finally, special thanks are owed to my parents, for their unconditional support and love over the years.   xxv Dedication      To my parents Pen Mao Wan and Ching Yun Wang    1 Chapter  1: Introduction 1.1 Thesis overview There has been a great deal of interest in the development of amphiphilic block copolymer micelles as drug delivery vehicles for use with the anticancer taxane drugs, paclitaxel (PTX) and docetaxel (DTX) (Ernsting et al., 2012a; Letchford et al., 2009; Mugabe et al., 2011b; Sarisozen et al., 2012; Zhang et al., 1996). Due to their amphiphilic nature, diblock copolymers are capable of self-assembling at the critical micelle concentration to form micelles characterized by a hydrophobic core surrounded by a hydrophilic corona. The core is usually composed of biocompatible, biodegradable polyesters including, poly(lactic acid) or poly(ε-caprolactone) (PCL) that solubilize water insoluble drugs such as the taxanes and the hydrophilic corona is typically methoxy poly(ethylene glycol) (MePEG) (Gaucher et al., 2005). The unimers of the micellar structure in solution are in equilibrium with those participating in the nanoparticle structure, resulting in a “fluid-like” core. These micellar systems offer several formulation and pharmacokinetic advantages for the delivery of taxanes, and this subject has been extensively reviewed (see (Gaucher et al., 2010)). Elimination of the use of the solubilizing agent, Cremophor-EL® (CrEL) (currently used in the PTX commercial formulation, Taxol®) would reduce the known hypersensitivity reactions associated with the use of this agent (Gelderblom et al., 2001).  One of the early micellar compositions of PTX developed by Burt and coworkers was based on methoxy poly(ethylene glycol)-block-poly (D, L-lactic acid) (MePEG-b-PDLLA) (Burt et al., 1999). In vitro and in vivo studies showed enhanced cytotoxicity and efficacy of PTX loaded MePEG-b-PDLLA nanoparticles compared to the commercial Taxol® formulation (Lee et al., 2011a).  Currently, a MePEG-b-PDLLA formulation of PTX is marketed as Genexol-PM® by Samyang (Lee et al., 2011a).   2 Aside from the poor aqueous solubility of taxanes, another significant barrier to the effective killing of cancer cells with these agents is drug efflux from cells by membrane bound transporter proteins, particularly P-glycoprotein (Pgp). Drug efflux proteins are known to reduce the intracellular concentration of drugs and therefore reduce cytotoxicity (Wong et al., 2014). Pgp is a transmembrane drug efflux protein, a member of the ATP-binding cassette superfamily, and is one of the most characterized drug transporters responsible for cellular multi-drug resistance (MDR) to a wide variety of anticancer drugs, including PTX and DTX (Szakacs et al., 2006; Ambudkar et al., 1999). Numerous strategies have been investigated to overcome the efflux of drugs that are Pgp substrates. These strategies include the administration of agents like verapamil or cyclosporin A which inhibit Pgp function in conjunction with taxanes (Tolcher et al., 1996; van Asperen et al., 1998). Other approaches include the encapsulation or conjugation of Pgp substrate drugs in nanoparticles to enable the entry into cells by endocytosis (therefore bypassing Pgp), or even co-formulation of the drugs and Pgp inhibitors in the same nanoparticles (Jabr-Milane et al., 2008; Chen et al., 2010; Vauthier et al., 2003; Yan et al., 2010; Lee et al., 2011b; Patel et al., 2011). Substantial work in this area has been conducted by Kabanov and co-workers who have demonstrated that certain commercially available Pluronic® amphiphilic agents are capable of inhibiting Pgp (Kabanov et al., 2002b). This led to several studies that evaluated the use of mixed polymeric nanoparticles incorporating one or more Pluronic® agents which served to both solubilize a Pgp substrate drug, doxorubicin, and inhibit Pgp function (Mei et al., 2009; Mu et al., 2010; Zhang et al., 2011; Zhang et al., 2010c). However, a potential drawback of using Pluronic® agents is the non-degradability and lack of renal excretion for high molecular weight members of this copolymer class (Besheer et al., 2009; Ruel-Gariepy and Leroux, 2004).    3 Previous studies by Burt and coworkers have demonstrated that a low molecular weight MePEG-b-PCL copolymer (MePEG17-b-PCL5) is an effective modulator of Pgp function. We reported on the ability of MePEG17-b-PCL5, used at concentrations both above and below the critical micelle concentration, to enhance the accumulation of Pgp substrates, including PTX and doxorubicin, in Pgp overexpressing Caco-2 and MDCKII-MDR1 cells (Elamanchili et al., 2009; Zastre et al., 2008). Our laboratory has investigated the use of nanoparticles composed of MePEG-b-PCL for the delivery of taxanes for a number of years. These studies have shown that as the molecular weight of the hydrophobic block increases, the nanoparticles shift from a fluid-like to a frozen core structure with corresponding changes in the performance of these nanoparticles as drug delivery systems (Letchford et al., 2009). In the studies described in this thesis, three types of nanoparticles were fabricated, keeping the MePEG block length at a constant of 114 ethylene glycol repeat units and conjugated to either a very short PCL block length of 19 caprolactone repeat units (MePEG114-b-PCL19) to form micelles, or a longer PCL block lengths of 104 (MePEG114-b-PCL104) and 200 (MePEG114-b-PCL200) repeat units to form kinetically “frozen core” structures which we have termed nanospheres. MePEG114-b-PCL104 nanospheres have been shown to solubilize more PTX, release the drug at a more sustained rate, and are more stable in the presence of human plasma as compared to their micelle counterparts, likely due to the “solid-like” core of these nanospheres (Letchford et al., 2009). However, whether the differences in the physicochemical properties and the stability of these nanoparticles might result in differences in cellular uptake and cytotoxicity of PTX loaded nanoparticles was not known. Our studies have shown that the MePEG-b-PCL copolymers are very versatile as both a delivery system for taxanes and an inhibitor of Pgp mediated drug efflux in resistant cells, depending on the length of the hydrophilic and hydrophobic blocks. Accordingly, they can   4 function as a micellar-type drug carrier (MePEG114-b-PCL19), a nanosphere-type drug carrier (MePEG114-b-PCL104 and MePEG114-b-PCL200) or a Pgp inhibitor (MePEG17-b-PCL5). Hence, we hypothesized that cellular uptake and cytotoxicity of taxane loaded MePEG-b-PCL nanoparticles would be dependent on the nanoparticle composition, that is, the length of hydrophobic PCL block (micelles versus nanospheres) and also on the co-administration (or co-encapsulation) with the Pgp inhibitor, MePEG17-b-PCL5. In Chapter 2 of this thesis, we describe the evaluation of the cellular uptake and cytotoxicity of PTX formulated in MePEG114-b-PCL19 micelles or MePEG114-b-PCL104 nanospheres incubated with drug sensitive (MDCKII) and drug resistant (MDCKII-MDR1) cell lines, with or without exposure to the Pgp inhibitor, MePEG17-b-PCL5. Our results show that PTX loaded micelles are more effective than nanospheres in inhibiting the proliferation of MDCKII-MDR1 cells, but have similar activity against MDCKII cells. Furthermore, the inclusion of MePEG17-b-PCL5 greatly enhanced the cytotoxicity of the nanospheres in MDCKII-MDR1 cells, probably via the inhibition of Pgp and the reversal of chemoresistance. Although differences between MePEG114-b-PCL19 micelles and MePEG114-b-PCL104 nanospheres were observed in in vitro cell studies, the pharmacokinetics of PTX loaded into these two formulations and administered intravenously at the same dose into healthy mice were similar (Letchford and Burt, 2012). In the studies described in Chapter 3, we manufactured a copolymer with a higher PCL block length, MePEG114-b-PCL200 and prepared PTX and DTX loaded nanospheres using this copolymer, with or without the coencapsulation of the low molecular weight, Pgp inhibitor copolymer, MePEG17-b-PCL5. The resulting nanospheres were termed “Mixed MW PCL200/PCL5” nanospheres and MePEG17-b-PCL5 was found to release in a controlled manner   5 from the nanospheres at sufficient concentrations to result in a dramatic increase in cytotoxicity in a drug resistant cell line. The development of ultrasound-enhanced chemotherapeutic strategies has resulted in a number of reported effects, including increased diffusion of drugs through tumors, increased tumor uptake through the leaky vasculature of tumors, increased cellular uptake of drugs (sonoporation) and localized release of drugs from controlled release nanoparticles at target sites after systemic administration (Nelson et al., 2002; Nomikou et al., 2010; Thakkar et al., 2013). In collaboration with Dr. Mu Chiao from the Department of Mechanical Engineering at UBC, we have developed a device that delivers short burst (10 s), high intensity (32 W/cm2) and high-frequency (4 MHz) ultrasound irradiation. Studies have shown that such ultrasound treatment increases the cellular uptake of many agents including antisense oligo-nucleotides, micellar paclitaxel (PTX) and doxorubicin in both PC3 prostate cancer (PC3) and human umbilical vein endothelial (HUVEC) cell lines (Jackson et al., 2011; Siu et al., 2007a; Siu et al., 2007b). Chapter 4 of this thesis describes the effect of this ultrasound regime on the cellular accumulation of PTX loaded in MePEG-b-PDLLA micelles in both drug-sensitive and Pgp overexpressing cell lines. There was an approximately two-fold increase in intracellular PTX accumulation for all ultrasound-treated drug-sensitive cell lines and their respective drug-resistant counterparts as compared to cells not treated with ultrasound. Significant decreases in drug efflux rates were also observed for both drug-sensitive and resistant cell lines receiving ultrasound. The enhanced accumulation and retention of PTX following ultrasound treatment resulted in greater cytotoxicity in both MDCKII and MDCII-MDR1 cell lines. These data suggest that ultrasound may facilitate the uptake of intact PTX-loaded micelles into cells, allowing greater retention of drug in both drug sensitive and resistant cells.     6 1.2 Multidrug resistance Drug resistant tumors are formed when tumor cells develop resistance to various drugs usually following exposure to chemotherapy (Gillet and Gottesman, 2010). Some tumors may also grow with a pre-existing multidrug resistance (MDR) without any prior exposure to drugs (Baguley, 2010a). Drug resistance is one of the major contributing factors to the failure of many chemotherapy regimes and is common in blood cancer and solid tumors including breast, ovarian, lung, and lower gastrointestinal tract cancers (Persidis, 1999). Initial studies on tumor chemoresistance were strongly influenced by well established studies in bacterial resistance which demonstrated many different mechanisms of resistance (Bolhuis et al., 1997). However, since the discovery of drug efflux transporters in chemoresistant cancer cells some 30 years ago, the increased expression of transmembrane drug efflux proteins is now established as the dominant cause of drug resistance (Baguley, 2010a; Borst et al., 2007). More recently, it has become clear that tumors are not composed of homogeneous cell populations but may mirror the heterogeneity of the originating tissues, including the presence of stem cells, rapidly dividing transit cells, differentiated cells and stromal cells (Borst et al., 2007; Dean et al., 2005; Heddleston et al., 2010). In general, cancer cells remain unaffected by extremely adverse conditions including inflammation, infection and toxins so it is not surprising that tumor cells are thought to survive in protected microenvironments. This includes conditions of reduced perfusion, high pre-existing expression levels of membrane efflux transporters (to reduce exposure to toxins or cytotoxic drugs), as well as functional prosurvival signal transduction pathways (Baguley, 2010a; Dean et al., 2005; Heddleston et al., 2010). It may be that the fate of tumors lies in the survival of just a few cells and may explain the recurrence of tumors from seemingly eliminated tumor masses.      7 1.2.1 Mechanisms of drug resistance  The mechanisms of drug resistance in tumors may be classified by the physiological or cellular factors causing resistance. The physiological factors causing drug resistance are the tumor physiology and the microenvironment of the solid tumors including high interstitial fluid pressure, hypoxia and low extracellular pH (Minchinton and Tannock, 2006). These factors are highly interconnected and limit the distribution of drugs in tumors. An indirect form of drug resistance may result from poor transport of drugs to target cells in tumors, due to poor perfusion resulting from a disorganized vasculature, the absence of lymphatic drainage (Leu et al., 2000), increased rigidity of extracellular matrices and high interstitial fluid pressure (Heldin et al., 2004), which in turn reduces convection and therefore inhibits the distribution of anticancer drugs (Reed et al., 2001). Nanoparticles move through tumors largely by convection arising from hydrostatic pressure (Campbell, 2006; Holback and Yeo, 2011). This means that the majority of nanoparticles that may enter the tumor via the disorganized vasculature, largely remain clustered in the tumor tissue around the blood vessels at the lower pressure periphery of the tumor due to an outward convection gradient (Holback and Yeo, 2011; Jang et al., 2003; Campbell, 2006; Jabr-Milane et al., 2008). Poor transport may result in drug localization close to blood vessels in the more vascularized peripheral regions only, with poor distribution in more central regions (Minchinton and Tannock, 2006; Holback and Yeo, 2011; Kuh et al., 1999). Poorly organized vasculature also limits the transport of oxygen and nutrients to cells that are distant from functional blood vessels, resulting in hypoxia, a well known feature of cancer (Minchinton and Tannock, 2006). It has been reported that hypoxic cells may be the source of tumor recurrence and metastasis (Holback and Yeo, 2011), and are resistant to chemotherapy as most anticancer drugs target proliferative aspects of cell growth (Subarsky and Hill, 2003). Hypoxic cells are also resistant to radiotherapy as this treatment is associated with the formation of reactive oxygen   8 species, which are reduced in hypoxic conditions due to the lack of an oxygen supply (Harris, 2002). The absence of a lymphatic system also leads to the build-up of acidic metabolic products, which lowers the extracellular pH (Tannock and Rotin, 1989; Gatenby and Gillies, 2004). Studies have shown that low pH reduce the permeability of cell membranes to basic anticancer drugs like anthracyclines and vinca alkaloids due to ionization effects (Wojtkowiak et al., 2011).  The cellular mechanisms of multidrug resistance include a reduction in drug influx, increased drug efflux, increased drug metabolism, increased DNA repair, lack of apoptotic machinery or an increased anti-apoptotic machinery (Patel et al., 2013; Baguley, 2010b; Fallica et al., 2011). It has been suggested that the binding affinity of microtuble binding agents such as taxanes to microtubules is dependent on the presence of specific β-tubulin isotypes (Hammond et al., 2008). The high expression of the β-tubulin isotype, βIII-tubulin, has been reported to associate with the drug resistance to taxanes likely due to weak binding affinity to microtubules (Kavallaris et al., 1997; Seve and Dumontet, 2008). However, the mechanisms on the up-regulation of βIII-tubulin expression in drug resistant cells are still unclear. Enhanced drug metabolism in tumors is another cellular method of drug resistance which inactivates anti-cancer drugs (Krishna and Mayer, 2000). Glutathione S-transferase is an enzyme involved in metabolic biotransformation of drugs or xenobiotics by glutathione conjugation, resulting in detoxification of the molecules. High levels of glutathione and increased glutathione S-transferase activity have been shown in many resistant cell lines and thus are promising therapeutic targets (Tew and Townsend, 2012). Apoptosis is programmed cell death that can be initiated by two pathways, 1) the mitochondrial intrinsic pathway involving the release of cytochrome c from mitochondria and 2) an extrinsic pathway involving the activation of death receptors on the cell membrane in response to ligand binding (Indran et al., 2011). The intrinsic pathway is regulated by Bcl-2   9 family members, that can be classified as pro-apoptotic (Bax and Bak) and anti-apoptotic (Bcl-2 and Bcl-xL) proteins (Youle and Strasser, 2008). In multidrug resistant cancer cells, the anti-apoptotic proteins are up-regulated, thus preventing the apoptotic process (Mashima and Tsuruo, 2005; Rodriguez-Nieto and Zhivotovsky, 2006; Viktorsson et al., 2005). However, one of the major cellular mechanisms of multidrug resistance involves the expression of drug efflux transporters in cancer cells which limit intracellular accumulation of chemotherapeutic agents, necessitating the administration of high doses (Patel et al., 2011). These mechanisms will be discussed in the following section.  1.2.1.1 ABC drug efflux transporters Membrane transporters are integral transmembrane proteins involved in the selective absorption of endogenous substances and in the elimination of toxic substances (Khurana et al., 2013). The adenosine triphosphate (ATP)-binding cassette (ABC) transporters are members of a protein superfamily that is found in all organisms and is the most well studied of transporter systems related to cancer due to their role in multidrug resistance (Wong et al., 2014). The ABC transporters share a high degree of sequence homology and organization, and are characterized by a highly conserved ATP binding domain located at the cytoplasmic side of the membrane. Most of the ABC transporters contain a minimum of two transmembrane domains and two nucleotide-binding domains, and function to transport a wide range of drugs and xenobiotics out of cells (Massey et al., 2014).   Around 50 ABC transporters have been identified and grouped into seven subfamilies, designated ABCA through ABCG, based on their nucleotide binding domain homology (Massey et al., 2014). P-glycoprotein (Pgp), multidrug resistant protein 1 (MRP1), and breast cancer resistant protein (BCRP) from ABC subfamilies B, C and G, respectively, are all known to be   10 involved in drug transport out of cells (Cole et al., 1992; Doyle et al., 1998; Ueda et al., 1987). Pgp is one of the best characterized human efflux transporters and is known to confer the strongest resistance to the widest variety of anticancer agents (Sharom, 2014). The ABCC subfamily comprises 12 members and includes 9 MRPs, of which MRP1 has been shown to transport a wide spectrum of substrates. MRP1 is the founding member of the C-subclass of ABC transporters and was first identified by Cole et al. in H69AR human drug-resistant cancer cell lines (Cole et al., 1992). The structural analysis information of MRP1 showed that MRP1 contains three transmembrane domains and two nucleotide-binding domains (Cole, 2014; Rosenberg et al., 2010). BCRP was first identified by Doyle et al. as a major factor mediating drug resistance in a human breast cancer cell line (MCF-7/AdrVp) selected for anthracycline resistance (Doyle et al., 1998). This group reported an ATP-dependent reduction in the intracellular accumulation of anthracycline anticancer drugs in MCF-7/Adr Vp cells which did not express Pgp. BCRP is also referred to as the mitoxantrone resistance factor as Miyake et al. described this transporter to be involved in mitoxantrone resistance in S1-M1-80 human colon carcinoma cells (Miyake et al., 1999). BCRP is referred to as a half–transporter, which contains only one transmembrane and one ATP-binding domain, and which undergoes homodimerization to generate functional activity (Meyer zu Schwabedissen and Kroemer, 2011). A list of common ABC transporters and their anti-cancer drug substrates are provided in Table 1.1.  The most prevalent ABC transporter in MDR is Pgp, which has broad poly-specificity, recognizing hundreds of compounds as small as 300 daltons up to 4000 daltons; thus, it has become an important drug efflux transporter to study in cancer research (Wong et al., 2014).      11 Table 1.1 Anticancer drugs known to be substrates of ATP-binding cassette transporters (Patel et al., 2011; Szakacs et al., 2006; Gillet et al., 2007; Coley, 2010; Glavinas et al., 2004; Sharom, 2006; Ozben, 2006). Shaded cells represent drugs that are the substrates of the transporters.  Drug Class Drug  ABC Family ABCA ABCB ABCC ABCG A2 A3 B1 Pgp B4 B5 B11 C1 MRP1 C2 C3 C4 C5 C6 C10 C11 G2 BCRP Vinca alkaloids Vinblastine                Vincristine                Anthracyclines Daunorubicin                Doxorubicin                Epirubicin                Epipodophyllotoxins Etoposide                Teniposide                Taxanes Paclitaxel                Docetaxel                Kinase inhibitors Imatinib                Flavopiridol                Camptothecins Irinotecan                SN-38                Topotecan                Thiopurines 6-Mercaptopurine                6-Thioquanine                5-Fu                Antracenes Bisantrene                Mitoxantrone                Anti-tumor antibiotics ActinomycinD                Mitomycin C                Other Cisplatin                Arsenite                Colchicine                Estramustine                Methotrexate                Saquinivir                PMEA                AZT                Cytarabine  (Ara-C)                  12 1.2.1.2 The structure and function of P-glycoprotein There are a number of potential mechanisms of MDR, but overexpression of P-glycoprotein (or Pgp) is perhaps the best characterized. Pgp is a 170-kDa transmembrane efflux transporter protein encoded by the multidrug resistance gene (MDR1)(Sharom, 2014). The crystallographic structure of mouse Pgp has been resolved and suggests that this protein is composed of two homologous halves, each containing a transmembrane domain and a cytoplasmic nucleotide-binding domain (Fig 1.1 A) (Aller et al., 2009). The structural models depict the 12 transmembrane domains forming a ring, with the nucleotide (ATP) binding domains positioned on the cytoplasmic side (Jin et al., 2012). Three glycosylation branches are attached to the first extracellular loop between the first transmembrane helix bundle and the second transmembrane helix (Schinkel et al., 1993). Although glycosylation of Pgp was found not to affect drug transport activity of Pgp, it was shown to be important for proper quality control of the processing of Pgp in the endoplasmic reticulum and proper membrane routing and stability (Loo and Clarke, 1998; Schinkel et al., 1993). Site-directed mutagenesis studies demonstrated that the drug binding site was located at the transmembrane domains of Pgp, which was later confirmed by photoaffinity labeling in conjunction with high-resolution mass spectroscopy (Parveen et al., 2011; Loo and Clarke, 2008). Studies have indicated that these binding sites are also the transport and modulating sites to accommodate Pgp substrates and inhibitors (Hennessy and Spiers, 2007). In general, Pgp substrates are structurally diverse but are hydrophobic in nature. These substrates are proposed to bind to the Pgp drug binding domain via hydrophobic and van der Waal’s interactions (Massey et al., 2014). Fluorescence resonance energy transfer studies showed that the binding sites of the Pgp substrates, Hoechst 33342 and LDS0751, were located within the transmembrane domain of Pgp, in the cytoplasmic leaflet of the membrane (Sharom, 2014). Loo and Clarke constructed deletion mutants of Pgp with both   13 nucleotide binding domains removed, and showed that the transmembrane domains of the transporter were sufficient to bind drug substrate, suggesting that cytoplasmically located nucleotide binding domains were not required for binding (Loo and Clarke, 2008). However, the Pgp efflux activity was abolished for the deletion mutants, suggesting that the nucleotide binding domains were required for displacement of bound substrates. These findings have led to the most widely accepted model for Pgp mediated efflux called the Hydrophobic Vacuum Cleaner model, which proposed that substrates bind to the lipid bilayer due to their partition coefficients, interact directly with the transmembrane domain of Pgp and are then transported back to the extracellular spaces (Fig. 1.1 BI) (Gottesman and Pastan, 1993). Interaction of substrates with the binding sites within the transmembrane domain of Pgp along with ATP hydrolysis, results in a conformational change in the transmembrane domains of Pgp. Based on the assumption that the substrates have to partition into the lipid bilayer prior to Pgp binding, the release of substrate into the extracellular space by Pgp may occur via a “flippase” activity that moves the substrates from the inner to the outer leaflet of the membrane followed by rapid partitioning into the extracellular medium (Figure 1.1 BII) (Sharom, 2014). This mechanism requires substrates to be localized in the inner leaflet or outer leaflet of the phospholipid bilayer rather than being randomly distributed in the hydrophobic core. Siarheyeva et al. supported this hypothesis using NMR analysis and showed that Pgp substrates concentrated between the phosphate of the lipid headgroup and the upper segments of the lipid acyl chains (Siarheyeva et al., 2006).    14    Figure 1.1 A) The membrane topology of P-glycoprotein showing two homologous halves, each with a transmembrane domain and a nucleotide-binding domain (NBD). B) A simplified representation of the proposed mechanisms: I) Hydrophobic Vacuum Cleaner model and II) Flippase model. Pgp substrate (green triangles) partition into lipid bilayer, binds to transmembrane domain and ATP hydrolysis results in substrate efflux (I) or flipping substrate from the inner to the outer leaflet and rapid efflux (II). These figures are adapted from Sharom, 2014.    ATP ATPIIIExtracellularCytoplasmATP ATPA) B)   15 1.2.1.3 Taxanes: chemistry and pharmacology PTX and DTX belong to the taxane family and are effective anticancer agents for treating a wide range of malignancies (Crown and O'Leary, 2000; Pazdur et al., 1993). PTX was first extracted from the bark of the Western Yew tree, Taxus brevifolia (Wani et al., 1971). The extraction process recovered only low yields of PTX so that a semisynthetic approach using a precursor, 10-deacetyl baccatin III, that is readily available from the needles of the European Yew tree, Taxus baccata L, was developed to increase the production of PTX (Denis et al., 1988). DTX is a semisynthetic analogue of PTX from the precursor 10-deacetyl baccatin III (Subrahmanyam Duvvuri et al., 1998). PTX has been used as an anticancer agent for treating a range of cancers including breast, ovarian, lung, prostate, melanoma, leukemia, and bladder cancers (Galsky, 2005; Rowinsky and Donehower, 1995). On the other hand, DTX was approved by the Food and Drug Administration (FDA) of the United States for the treatment of advanced ovarian, breast, head and neck, prostate, gastric, and non-small cell lung cancer (Ramaswamy and Puhalla, 2006). The primary mechanism of action of the taxanes is to inhibit cell proliferation by binding to β-tubulin, forming stable microtubules, and arresting the cell cycle at the G2-M phase, leading to apoptosis (Rowinsky and Donehower, 1995; Ringel and Horwitz, 1991).  PTX is a large molecule (853.9 g/mol) with the chemical formula C47H51NO14 (Fig 1.2 A). Due to its hydrophobic taxane ring and relatively few polar groups, PTX is extremely water insoluble. Liggins et al. reported the aqueous solubility of PTX to be around 1 μg/ml (Liggins et al., 1997). The commercially available PTX formulation is Taxol® in which 6mg/ml PTX is solubilized in Cremophor® EL (CrEL) (polyethoxylated castor oil) and anhydrous ethanol. The administration of CrEL is associated with serious adverse effects such as acute hypersensitivity reactions, including symptoms of dyspnoea, flushing, rash, chest pain, tachycardia, hypotension,   16 and generalized urticaria (Gelderblom et al., 2001). CrEL is also linked to abnormal lipoprotein patterns, hyperlipidaemia, and neurotoxicity (Gelderblom et al., 2001). Given these problems associated with CrEL, nanoparticulate drug delivery systems for PTX have been the subject of much research interest.  DTX differs from PTX at the C-10 and C-3 positions of the side chain and has a chemical formula of C43H53NO14 with a molecular weight of 808 g/mol (Fig 1.2 B). Similar to PTX, DTX has a very low aqueous solubility of approximately 7 μg/ml (Du et al., 2007). DTX is commercially available as Taxotere® in which Tween 80 (polysorbate 80) and ethanol are used to solubilize DTX. The formulation is diluted 5-20 fold with saline prior to intravenous administration.  Both PTX and DTX are substrates for Pgp mediated efflux, and their cytotoxicity is thus compromised in MDR cells that overexpress Pgp (Geney et al., 2002; Shirakawa et al., 1999; Stordal et al., 2012). While overexpression of ABCB11, ABCC2 and ABCC10 is also related to taxane resistance, Pgp is the most studied member of ABC superfamily for the taxane resistance (Hopper-Borge et al., 2011; Huisman et al., 2005; Massey et al., 2014). Ehrlichova et al. reported that Pgp-overexpressing MDR breast cancer cells (NCI/ADR-RES) were 1,000-fold less sensitive to PTX than the PTX sensitive cell counterpart (MDA-MB-435) with up to 20-fold lower uptake of the drug in the resistant cells which overexpressed Pgp (Ehrlichova et al., 2005). The National Cancer Instiute has conducted studies in which 60 cell lines from a broad range of malignancies were measured for their mdr-1/Pgp expression using polymerase chain reaction. It was found that high Pgp expression correlated with a greater resistance of the cell lines to PTX (Alvarez et al., 1995). A recent study reported that in resistant MDA-MB-231 human breast cancer cells, the IC50 of DTX was increased 25-fold as compared to the parent line, and Pgp   17 expression was upregulated by more than 20,000-fold in the DTX resistant cells (Roy et al., 2014).   Interestingly, taxanes have low oral bioavailability since Pgp is expressed in the intestine and these drugs are rapidly metabolized by cytochrome P450 enzymes. This phenomenon has led to a great interest in developing oral formulations of taxanes using the coadministration of  agents that can inhibit Pgp (such as cyclosporin A) or cytochrome P4503A (such as ritonavir) (Hendrikx et al., 2013; Jibodh et al., 2013).   1.3 Modulation of P-glycoprotein Because only minor levels of Pgp upregulation in cancer cells can confer full drug resistance (Borst et al., 2007), Pgp has been established as the dominant drug efflux transporter implicated in MDR. Accordingly, inhibition of Pgp function has become the major chemosensitizing strategy for most drug companies over the last 20 years (Baguley, 2010a; Borst et al., 2007). Inhibitory strategies fall into three categories: blocking drug binding to Pgp, inhibiting Pgp-ATPase activity or modulating the protein’s lipid microenvironment in the membrane (Palmeira et al., 2012).        18  Figure 1.2 Chemical structures of paclitaxel (A) and docetaxel (B)  1.3.1 Pharmacological agents as P-glycoprotein inhibitors The development of Pgp modulators or inhibitors has received much attention. In general, Pgp inhibitors can be classified into first, second and third generation agents. First generation compounds such as cyclosporin A, verapamil and several calmodulin antagonists were identified in the 1980s (Hait et al., 1989; Tsuruo et al., 1981). Verapamil was discovered by Tsuruo and co-workers to enhance intracellular accumulation of vincristine in P388 leukemia cells and its vincristine-resistant subline by reversing MDR (Tsuruo et al., 1981). It was later demonstrated that other calcium channel blockers such as felodipine, isradipine, nicardipine, nifedipine, bepridil and diltiazem demonstrated a similar multidrug resistance reversing property (Krishna and Mayer, 2000). However, these agents reverse MDR at very high concentrations ranging from 5 to 50 μM, which are generally cytotoxic to normal cells when administered intravenously (Krishna and Mayer, 2000). The clinical trials using verapamil for reversing MDR were OOOOOHOOOHONHOHOOOOOOOOHNO OOOOOHOOOOHOHHHA) B) Chemical Formula: C47H51NO14 Molecular Weight: 853.9 g/mol  Chemical Formula: C43H53NO14 Molecular Weight: 807.9 g/mol    19 discontinued due to significant cardiac toxicity (Benson et al., 1985). Cyclosporin A, a commonly used immunosuppressant for organ transplantation showed great promise as a first generation inhibitor. Cyclosporin A was shown to increase the cytotoxicity and intracellular levels of anthracyclines and vinca alkaloids by binding to the drug recognition sites on Pgp (Coley et al., 1993; Tamai and Safa, 1990). However, the phase III clinical studies showed that cyclosporin A had no positive effect in cancer patients (Sonneveld et al., 2001; Tallman et al., 1999). In general, the first generation inhibitors were effective only at relatively high concentrations due to low binding affinities to Pgp, resulting in unacceptable toxicity (Coley, 2010). In addition, many of the modulators were themselves substrates and/or inhibitors of drug transporters and/or CYP450 that are intrinsically expressed in the organs responsible for drug metabolism and elimination; thus, considerable pharmacokinetic interactions of the modulating agents with cytotoxic agents were observed.  The second generation Pgp inhibitors were designed to reduce the toxicity associated with the first generation inhibitors. Among the second generation compounds, valspodar (a derivative of cyclosporin D) was studied extensively. Twentyman and Bleehen reported that valspodar (PSC-833) had 20 fold improved binding compared to the first generation Pgp inhibitors (Twentyman and Bleehen, 1991). Lo et al. investigated the effect of valspodar on the intracellular accumulation of liposomal formulated epirubicin in Caco-2 cells.  The study showed that both free and liposomal valspodar reduced basolateral to apical efflux of epirubicin across Caco-2 monolayers by inhibiting Pgp (Lo et al., 2001). However, administration of valspodar failed to show any advantages in acute myeloid leukemia patients in clinical trials using daunorubicin, cytarabine and etoposide which are all Pgp substrates (Balayssac et al., 2005; Machaczka et al., 1998). In addition, valspodar was shown to interact with CYP450 and thus   20 altered the pharmacokinetics of co-administered drugs, resulting in increased cytotoxicity in the patients.  The third generation Pgp inhibitors include small molecule inhibitors such as elacridar and tariquadar. Elacridar, an acridone-carboxamide derivative, is a Pgp inhibitor, active in the nanomolar concentration range, and reverses drug resistance by binding to the allosteric site of Pgp (de Bruin et al., 1999; Hyafil et al., 1993). The potency of tariquadar was shown to be about 30 times greater than the second generation inhibitors, and several logarithmic orders greater than the first generation inhibitors (Pajeva and Wiese, 2009). Tariquadar inhibits Pgp by binding to its substrate binding site and inhibiting ATPase activity (Fox and Bates, 2007). The clinical trial of tariquadar was suspended in Phase III due to unfavorable toxicity in non-small cell lung cancer patients (Fox and Bates, 2007). In general, third generation inhibitors demonstrate enhanced potency with minimal pharmacokinetic interactions but lack comprehensive clinical trial data (Baguley, 2010a; Borst et al., 2007; Coley, 2010).  1.3.2 Pharmaceutical excipients as P-glycoprotein inhibitors Significant attention is being paid to the use of pharmaceutical excipients as Pgp inhibitors (Seelig and Gerebtzoff, 2006). Pharmaceutical excipients such as several polysorbates (Tween® 20 and 80), Triton® X-100, Cremophor® EL, PEG-400, several poloxamers (Pluronic® P85, P123, and F127), Solutol® HS-15, Peceol®, Gelucire® 44/14, Labrasol® and vitamin E tocopheryl-PEG-1000-succinate (TPGS) have been demonstrated to enhance the cellular accumulation of Pgp substrates (Woodcock et al., 1992; Bogman et al., 2003; Bittner et al., 2002; Johnson et al., 2002; Hugger et al., 2002; Sachs-Barrable et al., 2007; Lin et al., 2007; Rege et al., 2002; Wei et al., 2012). Most of the pharmaceutical excipients mentioned above are surfactants. With respect to their application as Pgp inhibitors, non-ionic surfactants have   21 structures consisting of a long hydrophobic chain or tail for membrane anchoring and a high proportion of ester and/or ether groups, which act as hydrogen bonding sites with membrane components. These hydrogen acceptors, often present as ester groups, are abundant in Pgp modulating surfactants such as vitamin E TPGS and MePEG-b-PCL (Seelig and Gerebtzoff, 2006). Hydrogen bonding is believed to result in a close association of the modulator hydrogen acceptors sites with donor sites in the Pgp protein molecule with possible subsequent modulation of transport function (Seelig and Gerebtzoff, 2006). Overall, it has been proposed that surfactants inhibit/modulate Pgp by various mechanisms including, altering membrane fluidity and permeability, interaction with Pgp substrate binding sites, inhibition of Pgp-ATPase activity, and ATP depletion (Dudeja et al., 1995; Regev et al., 1999; Kabanov et al., 2003). Rege et al. reported that Tween® 80 and CrEL increased apical-to-basolateral and decreased the basolateral-to-apical permeability of the Pgp substrate rhodamine 123 in Caco-2 cells and these surfactants also increased membrane fluidity (Rege et al., 2002). On the other hand, vitamin E TPGS and PEG surfactants that reversed MDR, decreased lipid fluidity in plasma membrane from MDR cells, whereas surfactants that failed to inhibit Pgp (octyl-β-D-glucoside) had no effect on membrane fluidity, evaluated using several fluorescent probes that were Pgp substrates (Rege et al., 2002; Dudeja et al., 1995). It is evident that changes in the lipid membrane microenvironment of Pgp influences Pgp function with variable effects of modulators on membrane fluidity.  1.3.2.1 Pluronic® triblock copolymer surfactants Among all the surface active agents showing Pgp inhibitory function, the copolymer series known as Pluronic® agents have been the most extensively studied. These Pluronic® agents are block copolymers consisting of hydrophilic poly(ethylene oxide) (PEO) and hydrophobic   22 poly(propylene oxide) (PPO) blocks arranged in an A-B-A triblock (PEO-PPO-PEO) structure (Batrakova and Kabanov, 2008). The use of Pluronic® copolymers to overcome MDR was first reported in the early 90’s by Paradis et al., where Pluronic® P85 micelles loaded with daunorubicin were shown to increase cellular accumulation, resulting in a greater cytotoxic effect (lower IC50) of the drug in an MDR ovarian carcinoma cell line (SKVLB) as compared to free daunorubicin (Paradis et al., 1994). Similar results were obtained with other Pgp substrates such as doxorubicin, epirubicin, vinblastine and mitomycin C in SKVLB cells (Alakhov et al., 1996). The Pluronic® P85 copolymer was shown to be a potent Pgp inhibitor capable of sensitizing MDR cancer cells and enhancing drug transport across cellular membranes of Caco-2 cells and brain endothelium (Batrakova et al., 2001c; Kabanov et al., 2002a). It was reported that Pgp inhibition caused by Pluronic® P85 was most effective at concentrations below the CMC suggesting that the unimers were responsible for the Pgp inhibition (Batrakova et al., 1999).  Studies using Pluronic® agents have shown that additional factors likely contribute to the mechanisms of Pgp inhibition by amphiphilic block copolymers. Batrakova et al. examined the mechanisms by which Pluronic® P85 inhibited Pgp using bovine brain microvessel endothelial cells (BBMEC) (Batrakova et al., 2001b). Following exposure of BBMEC to Pluronic® P85, the intracellular ATP levels were depleted, the membrane fluidity was increased, and a decreased Pgp-ATPase activity was observed (Batrakova et al., 2001b). The depletion in intracellular ATP level was not due to leakage of intracellular ATP out of the cells but resulted from lowered metabolic mitochondrial activity and inactivation of mitochondrial oxidative phosphorylation as confirmed by mitochondrial membrane potential determination (Hong et al., 2013; Li et al., 2014; Batrakova et al., 2001b). It was suggested that since Pgp depends on ATP hydrolysis for pump activity, it is likely that ATP depletion in combination with a reduced ATPase activity, may contribute to the ability of Pluronic® agents to inhibit Pgp efflux activity (Batrakova et al.,   23 2001b; Batrakova et al., 2001a; Batrakova et al., 2004). The authors also suggested that Pluronic® agents might reduce ATP levels in the cells via an interaction with mitochondrial membranes and subsequent disruption of the electron transport chain process (Batrakova and Kabanov, 2008). In order for the Pluronic® agents to interact with mitochondrial membranes, the Pluronic ® agents must enter cells. Batrakova et al. labeled Pluronic® P85 with fluorescein and showed a strong intracellular fluorescein level in the cytoplasm of BBMEC cells, suggesting that Pluronic® P85 was able to pass through the cell membrane and enter the cytoplasm (Batrakova et al., 2001b).   In other studies Minko et al. investigated the effect of Pluronic® agents on drug induced changes in signal transduction related to apoptosis (Minko et al., 2005). Various cells lines including the MDR subline of human epithelial cells, KBV and MDR human breast carcinoma cells and MCF-7/ADR were treated with Pluronic® P85 and doxorubicin and the gene expression of various apoptosis related proteins was studied (Minko et al., 2005). Using real time polymerase chain reaction analysis, the MDR cells treated with Pluronic® P85 and doxorubicin showed enhanced proapoptotic responses, and decreased antiapoptotic cellular defenses (Minko et al., 2005). In contrast, the MDR cells that were treated with doxorubicin alone showed the activation of both proapoptotic and antiapoptotic signals (Minko et al., 2005). An enhanced proapoptotic signal by Pluronic® P85 is likely to result in an enhanced cytotoxicity of doxorubicin in MDR cells but the molecular mechanism by which Pluronic® P85 alters apoptotic signal transduction of doxorubicin in MDR cells is still unknown.  1.3.2.2 MePEG17-b-PCL5 diblock copolymer Our research group has explored the use of MePEG17-b-PCL5 to enhance the cellular accumulation of the Pgp substrates, rhodamine-123 (R-123), rhodamine-6G (R-6G), PTX and   24 doxorubicin (Zastre et al., 2002; Zastre et al., 2004; Zastre et al., 2007; Wan et al., 2013; Elamanchili et al., 2009). The mechanisms of increased R-123 accumulation in Caco-2 cells in the presence of MePEG17-b-PCL5 were suggested to involve inhibition of Pgp and increased membrane permeabilization resulting in increased transmembrane diffusion of R-123 (Zastre et al., 2004). Directional flux studies were conducted with Caco-2 cell monolayers on transwell plates and showed that the apical to basolateral flux of R-123 in the presence of MePEG17-b-PCL5 and verapamil was similar to R-123 alone. However, the basolateral to apical flux of R-123 was significantly decreased with MePEG17-b-PCL5 and verapamil when compared to R-123 alone, suggesting that MePEG17-b-PCL5 inhibited the drug efflux pump on the apical side of the cells (Zastre et al., 2004). Fluorescence anisotropy measurements of membrane fluidity demonstrated that MePEG17-b-PCL5 decreased Caco-2 cell membrane fluidity (Zastre et al., 2007). In the same study, MePEG17-b-PCL5 stimulated Pgp-ATPase activity approximately threefold at concentrations at which maximum intracellular accumulation of R-123 into Caco-2 cells was observed, suggesting that MePEG17-b-PCL5 may interact directly with Pgp leading to its inhibition (Zastre et al., 2007). Clear differences in accumulation enhancement in Caco-2 cells were observed when comparing hydrophobic Pgp substrates, R-6G and PTX with more hydrophilic Pgp substrates, R-123 and doxorubicin. Maximum enhancement of accumulation for the hydrophobic substrates was found to be below the CMC of MePEG17-b-PCL5, as compared to above the CMC for the more hydrophilic substrates (Zastre et al., 2008). Elamanchili et al. conducted experiments to evaluate the effect of MePEG17-b-PCL5 in enhancing intracellular accumulation of Pgp substrates in Pgp expressing human ovarian cancer NCI/ADR-RES and canine kidney MDCKII-MDR1 cells using tritium-labeled PTX and doxorubicin (Elamanchili et al., 2009). MePEG17-b-PCL5 enhanced accumulation of PTX and doxorubicin in Pgp overexpressing MDR cells but did not influence substrate accumulation in their non-Pgp   25 expressing counterpart cells (Elamanchili et al., 2009). Drug retention studies demonstrated that doxorubicin was retained inside MDR cells whereas PTX was rapidly effluxed from the cells upon removal of MePEG17-b-PCL5, suggesting that a sustained inhibition of Pgp in the presence of MePEG17-b-PCL5 was required to retain PTX in MDR cells and enhnace cytotoxicity (Elamanchili et al., 2009). However, mechanisms by which MePEG17-b-PCL5 inhibits Pgp in MDR cells and enhances intracellular accumulation of Pgp substrates remains poorly understood.  1.4  Nanoparticulate taxane formulations 1.4.1 Polyether-polyester based block copolymers  Amphiphilic block copolymer nanoparticles have garnered a great deal of attention as drug delivery systems due to their ability to solubilize hydrophobic drugs and, in some cases, alter the pharmacokinetic and biodistribution of their drug payload (for reviews, see (Gaucher et al., 2010) and (Mikhail and Allen, 2009)). Due to their well established biocompatibility and biodegradability, and an extensive history of approved use in biomedical applications, block copolymers composed of hydrophilic blocks such as PEO and MePEG and a variety of polyester hydrophobic blocks based on PDLLA (Zhang et al., 1997; Zhang et al., 1996), poly(lactic-co-glycolic acid) (PLGA) (Hu and Zhang, 2010; Yoo and Park, 2001), and PCL (Bruce et al., 1992; Letchford et al., 2009) have been extensively investigated as nanoparticulate delivery vehicles. In other studies, poly(N-vinyl-pyrrolidone) (PVP) (Le Garrec et al., 2004) has served as the hydrophilic block and conjugated to PDLLA hydrophobic blocks, for the delivery of taxanes. The cellular toxicity of unloaded PVP-b-PDLLA micelles is considerably less than CrEL or polysorbate 80 alone. This has also been demonstrated by other groups using polyether-polyester-based nanoparticles (Gaucher et al., 2010).   26 In general, PTX or DTX-loaded polyether-polyester diblock copolymer nanoparticles show slightly less cytotoxicity in cell studies when compared to Taxol® or Taxotere®, respectively, at low drug concentrations (≤1 μg/mL) (Gaucher et al., 2010). It is thought that taxanes are rapidly dissociated from CrEL or polysorbate 80 and enter the cells via diffusion, resulting in greater cytotoxicity. Diblock copolymers nanoparticles with PDLLA or PLGA hydrophobic blocks have been demonstrated to release drug payload more slowly and are thus less cytotoxic than the commercial formulations at low concentrations (Xu et al., 2009). However, at high drug concentrations, PTX and DTX loaded polyether-polyester nanoparticles show greater cytotoxicity than Taxol® and Taxotere®. Dong reported that using a high PTX concentration (10 μg/mL), PTX loaded MePEG-b-PLGA nanoparticles were more cytotoxic than Taxol® against giloma C6 cells (Dong and Feng, 2007). Similarly, greater cytotoxicity of PTX-loaded MePEG-b-PLGA nanoparticles at high PTX concentration (25 μg/mL) was observed against human small lung cancer cell line (NCI-H69 SCLC) compared to Taxol® (Fonseca et al., 2002).   A number of non-polyether-polyester based nanoparticles have been demonstrated as the promising delivery systems for taxanes. Several years ago, a new intravenous formulation of PTX, marketed as Abraxane® was approved, in which albumin is used to coat nanoparticulate solid drug. Upon IV injection of Abraxane®, PTX quickly partitions out of nanoparticles and binds to albumin and alpha-1-acid glycoprotein in the blood, followed by subsequent tumor uptake via albumin-specific receptor, glycoprotein 60, mediated endocytosis (Yardley, 2013). Currently, Abraxane® is recommended for the treatment of metastatic breast cancer after the failure of first-line treatment with anthracycline (Petrelli et al., 2010). Cellax™, a DTX conjugated carboxymethylcelluose nanoparticle developed by Li and coworkers, has been shown to outperform Abraxane® and Taxotere® in mouse tumor models with signifcant control of   27 metastases and long blood circulation (Ernsting et al., 2012a; Ernsting et al., 2012b). The clinical trials of Cellax™ are anticipated to begin in 2015.  1.4.1.1 Taxane loaded MePEG-b-PCL nanoparticles The use of MePEG-b-PCL diblock copolymer in delivering hydrophobic drugs has received increasing attention (for review, see (Jin et al., 2014)). MePEG-b-PCL nanoparticles with MePEG molecular weights ranging from 2999-5000 g/mol and 14-114 PCL repeat units have been evaluated as nanocarriers of hydrophobic drugs, including indomethacin (Shin et al., 1998), PTX (Kim and Lee, 2001), dihydrotestosterone (Allen et al., 2000), ellipticine (Liu et al., 2004), cyclosporin A (Binkhathlan et al., 2012), and DTX (Jin et al., 2014). Forrest et al. synthesized hydrophobic prodrugs of PTX to increase PTX loading in PEG114-b-PCL92 nanoparticles, demonstrating 17-22% (w/w) prodrug loading with a mean diameter of less than 50 nm (Forrest et al., 2008). The functional groups at the end of the PCL hydrophobic block are important for taxane loading capacity in MePEG-b-PCL nanoparticles (Carstens et al., 2008). Carstens et al. synthesized MePEG17-b-PCL5 diblock copolymer with hydroxyl, benzoyl, or naphthoyl end groups, and showed that MePEG17-b-PCL5 micelles with aromatic end groups such as benzoyl and naphthoyl had a better compatibility between the micellar core and the encapsulated taxanes as compared to the MePEG17-b-PCL5 copolymers with hydroxyl end groups (Carstens et al., 2008). Shahin and Lavasanifar reported good drug loading and sustained drug release from MePEG114-b-PCL18 nanoparticles in which PTX was chemically conjugated to the PCL block (Shahin and Lavasanifar, 2010). However, the cytotoxicity of the PTX conjugated MePEG114-b-PCL18 nanoparticles against human breast cancer MDA-MB-435 cells was reported to be significantly lower than the cytotoxicity of free PTX (Shahin and Lavasanifar, 2010). Using similar approaches, Mikhail and Allen synthesized a PEG-b-PCL-DTX conjugate, which   28 demonstrated more than a 1000-fold increase in the aqueous solubility of DTX and sustained drug release over the course of one week (Mikhail and Allen, 2010).  Letchford et al. synthesized a series of MePEG-b-PCL diblock copolymers with the same hydrophilic block length but with different hydrophobic block lengths. The MePEG-b-PCL diblock copolymer with short hydrophobic block length, MePEG114-b-PCL19, was shown to form micelles using a direct dissolution technique. The long hydrophobic block length, MeEPG114-b-PCL104, was not water-soluble and therefore required nanoprecipitation and dialysis techniques in order to form stable nanoparticulates, termed nanospheres. The hydrophobic core of micelles was more fluid-like, suggesting that the aggregated copolymer chains were self-assembled at the CMC and were in dynamic equilibrium with unassociated unimers (Riley et al., 2001). On the other hand, there was no distinct CMC for nanospheres and the larger, kinetically frozen, solid-like hydrophobic core solubilized greater amounts of PTX (3% (w/w)) (Letchford et al., 2009). Similar results were reported by Gellert’s group who investigated the effect of increasing the hydrophobic block length of MePEG-b-PDLLA copolymers on the physicochemical properties of resulting nanoparticles (Heald et al., 2002; Riley et al., 2001). Gellert and coworkers showed that as the PDLLA block length increased, the hydrophobic core of the nanoparticles became more solid, resembling nanospheres, whereas the shorter PDLLA block produced nanoparticles resembling micelles (Heald et al., 2002; Riley et al., 2001). In this thesis work, three compositions of MePEG-b-PCL with constant MePEG block length and varying PCL block lengths were used to fabricate taxane loaded nanoparticles; MePEG114-b-PCL19, MePEG114-b-PCL104 and MePEG114-b-PCL200. The MePEG114-b-PCL19 copolymer formed micelles, with diameters of 30-40 nm and displayed low core microviscosity, whereas MePEG114-b-PCL104 formed nanoparticles with diameters of 50-70 nm, and possessed a higher core microviscosity than MePEG114-b-PCL19 micelles (Letchford et al., 2009). The   29 resulting MePEG-b-PCL micelles and nanospheres demonstrated distinct physicochemical characteristics (Figure 1.3) (Letchford et al., 2009). Both types of micelle- and nanosphere-like nanoparticles demonstrated controlled and sustained PTX release. For MePEG114-b-PCL19, about 90% PTX release was observed after 7 days in phosphate buffered saline (PBS), and 60% release for MePEG114-b-PCL104 over the same period of time (Letchford et al., 2009). Due to the larger hydrophobic block, MePEG114-b-PCL104 nanospheres solubilized a maximum of 3% w/w of PTX compared to 1.6% w/w PTX in MePEG114-b-PCL19 micelles (Letchford et al., 2009).  Figure 1.3 Micelles and Nanospheres formed by MePEG-b-PCL diblock copolymers and their general characteristics.  MePEG114-b-PCL19 micelle characteristics Size: 30-40nm Aggregated copolymers in dynamic equilibrium with unimers. Mobile fluid-like core MePEG114-b-PCL104 nanosphere characteristics Size: 50-70nm Copolymer unimers in a “frozen” state Phase-separated solid matrix core   30 1.4.1.2 Pharmacokinetics and biodistribution of taxane loaded nanoparticles The nanoparticle core of polyether-polyester based copolymers may be exploited as a cargo space and has been demonstrated to provide enhanced solubilization and controlled release of a variety of hydrophobic drugs. Detailed NMR studies have demonstrated that the PEGylated surface of amphiphilic block copolymer nanoparticles consists of PEG chains that extend into the aqueous milieu and exhibit significant chain mobility (Hrkach et al., 1997). This highly water-bound and mobile corona acts to sterically hinder the adsorption of plasma proteins such as opsonins and lipoproteins, which otherwise would lead to the rapid uptake and clearance of nanoparticles by the reticuloendothelial system, disruption of the nanoparticle and/or extraction of the encapsulated drug (Jeschke et al., 1978; Savic et al., 2006). Numerous studies have demonstrated that PEGylated nanoparticles with sufficient PEG shielding significantly increase the circulation time of the carrier (Bazile et al., 1995; Gref et al., 1994; Dunn et al., 1994). The size of the nanoparticles also has an impact on the pharmacokinetics and biodistribution of the carrier and its drug payload. The kidneys and spleen filter and remove nanoparticles from the blood that are less than about 5.5 nm and greater than 200 nm, respectively (Mikhail and Allen, 2009) whereas, the endothelial fenestrae in the liver range from 100 to 150 nm (Igarashi, 2008). This size dependent clearance was demonstrated by Liu et al. who showed that liposomes between 100-200 nm in diameter demonstrated prolonged circulation with minimal splenic and hepatic uptake as compared to particles outside this size range (Liu et al., 1992).  Many groups have shown that upon IV administration of taxane loaded low molecular weight polyether-polyester nanoparticles, there is a rapid dissociation and elimination of the drug payload, resulting in no advantage over the commercial formulation with respect to prolonged circulation and potential passive targeting of the drug (Burt et al., 1999; Kim et al., 2001; Le Garrec et al., 2004; Lee et al., 2011b; Letchford and Burt, 2012). Burt et al. reported that   31 MePEG-b-PDLLA diblock copolymer with an average molecular weight of 3333 g/mol demonstrated good PTX loading and physical stability (Burt et al., 1999). Biodistribution studies using dual-labeled radiolabeled PTX loaded MePEG-b-PDLLA micelles in rats showed that PTX rapidly dissociated from the micellar components in the blood and that greater than 95% of the administered MePEG-b-PDLLA diblock copolymer was eliminated within 15 hours. Letchford and Burt also reported similar rapid clearance of PTX from PTX loaded MePEG114-b-PCL19 and MePEG114-b-PCL104 nanoparticles (Letchford and Burt, 2012).  There is some evidence that larger taxane loaded polymeric nanoparticles composed of high molecular weight hydrophobic cores, with nanoparticle size greater than 100nm may circulate longer than Taxol® or Taxotere® (Gaucher et al., 2010). Using PLGA nanoparticles (200-400nm), surface stabilized with Vitamin E TPGS, Win and Feng showed a five-fold increase in the plasma AUC of PTX as compared to Taxol® even though there was an immediate rapid drop in plasma PTX concentration following injection (Win and Feng, 2006). Gaucher et al. demonstrated the prolonged circulation of PTX loaded PDLLA nanoparticles (180 nm in diameter) coated with low molecular weight PEG-b-PDLLA after IV administration to rats, as compared to Taxol® (Gaucher et al., 2010).  In general, significantly higher PTX concentrations were observed in the liver and spleen for drug loaded polyether-polyester nanoparticles when compared to Taxol® (Vonarbourg et al., 2006; Gaucher et al., 2010). The mononuclear phagocyte system is comprised primarily of the Kupffer cells and hepatocytes of the liver and the macrophages of the spleen. The binding of opsonins such as complement proteins C3b, immunoglobulin G and M, fibronectin and apolipoproteins to the surface of nanoparticles mediate recognition and removal of nanoparticles from the bloodstream by phagocytotic cells. Size and surface properties of nanoparticles are the two factors influencing the extent of opsonization. Some studies have shown a size dependence   32 in the clearance of nanoparticles, with the smaller nanoparticles less than 200 nm circulating for a longer time in the blood, suggesting that small size of nanoparticles is advantageous (Vonarbourg et al., 2006). This is because small nanoparticles have a high degree of curvature, making it more difficult for complement proteins to bind (Vonarbourg et al., 2006). The corona-forming block of nanoparticles is also an important factor for the clearance of nanoparticles by the reticuloendothelial system. Coating the surface of nanoparticles with hydrophilic polymer chains such as PEG has been shown to prolong blood circulation and prevent the binding of opsonins (Owens and Peppas, 2006).  1.4.1.3 Tumor uptake of taxane loaded nanoparticles The small and variable size of polymeric nanoparticles provides a mechanism by which disease sites such as solid tumors and areas of inflammation can be passively targeted (for review, see (Igarashi, 2008)). Many solid tumors are characterized by rapid angiogenesis resulting in a poorly formed hypervasculature with abnormal vessel wall structure (Folkman, 1971). This “leaky” vasculature in combination with poor lymphatic drainage has been demonstrated to lead to the accumulation of macromolecules and nanoparticles that are able to circulate in the blood for extended times (Matsumura and Maeda, 1986). This passive targeting mechanism is referred to as the enhanced permeation and retention effect (EPR) (Figure 1.4) and has been shown to be a key benefit for the delivery of anticancer agents as well as imaging agents to tumors using nanoparticulate delivery systems (Fang et al., 2011). Two prerequisites have been proposed for the occurrence of the EPR effect. First, the molecules must have a sufficient mass (> 40 kDa) to prevent diffusion back to the general circulation (Vonarbourg et al., 2006). Nanoparticles or macromolecules that are greater than 40 kDa have been demonstrated to evade renal clearance and leak into tumors (Ernsting et al., 2013). The other   33 prerequisite is that the drug-loaded nanoparticles must be able to achieve long circulation for at least 6 hours, allowing sufficient time for passive targeting to occur (Vonarbourg et al., 2006). As anticipated from EPR theory, 60 nm radiolabeled PEG114-b-PCL48 micelles may accumulate in tumors and was demonstrated by Lee et al who showed up to 5% of the injected dose of polymer accumulated in MCF-7 tumors up to 48 hours after injection (Lee et al., 2011a). After IV administration of PTX loaded PVP-b-PDLLA nanoparticles into female Balb/C mice, Le Garrec et al. showed that the tumour PTX concentration at the maximum tolerated dose was significantly higher than for Taxol® (Le Garrec et al., 2004).  However, the plasma levels of PTX administered via PVP-b-PDLLA nanoparticles were lower than for Taxol®, suggesting that the nanoparticles were passively targeting the tumour (Le Garrec et al., 2004). The enhanced tumor uptake and efficacy of taxane nanoparticulate formulations in mice models has also been observed for nanoparticles composed of PEG-b-PDLLA (Kim et al., 2001), PEG-b-PLGA (Cheng et al., 2007), and PEO-b-modified poly(β-amino ester) (Shenoy et al., 2005). Figure 1.4 Schematic representation of enhanced permeability and retention of drug delivery carriers in solid tumors.  Tumor blood vessels are characterized by leaky vasculature and malfunctioning lymphatic system, allowing drug loaded nanoparticulates to extravasate from tumor blood vessels and be retained in the tumor tissues. The figures are adapted from (Maeda, 2010). Healthy Blood Vessel Tumor Blood Vessel   34 1.4.1.4 Intracellular uptake and subcellular distribution of nanoparticles Internalization of drug-encapsulated nanoparticles into cells might be critical to evade membrane drug transporters related to MDR and form a drug depot inside the cells. Endocytosis is a process by which cells take up molecules or nanoparticles by engulfment following interaction with the cell membrane. The two types of endocytosis are phagocytosis for particles >1 micron and pinocytosis for smaller particles <1 micron. Pinocytosis can occur via four mechanisms; macropinocytosis, clathrin-mediated endocytosis, caveolae-mediated endocytosis, and clathrin- and caveolae- independent endocytosis. Macropinocytosis involves the formation of actin-filament driven membrane extensions which package cargo molecules or particles and form large endocytic vesicles named macropinosomes. It has been suggested that nanoparticles >200nm are internalized by cells through macropinocytosis (Walsh et al., 2006). Caveolae-mediated endocytosis is the dominant transendothelial pathway for the engulfment of cargo molecules (Oh et al., 2007). Membrane bound caveolin is a hairpin-like dimeric protein which inserts its loop into the inner leaflet of the cell membrane and forms an invaginated pouch engulfing cargo molecules (Monier et al., 1995). Clathrin-mediated endocytosis (CME) (also termed receptor mediated endocytosis) is an inherently active mechanism in all mammalian cells and is responsible for the uptake of essential nutrients, such as cholesterol and transferrin (Sahay et al., 2010a). CME involves the binding of cargo molecules to receptors in coated pits (a protein complex composed of clathrin and assembly proteins) which pinch off from the plasma membrane and form endocytic vesicles (Rejman et al., 2004). Clathrin- and caveolae- independent endocytosis occurs in cells devoid of clathrin and caveolin and are involved in the internalization of glycosylphosphatidylinositol linked proteins, extracellular fluid and growth hormones (Sahay et al., 2010a). The mechanism of clathrin- and caveolin- independent endocytosis remains poorly understood.    35 Numerous studies have reported the cellular uptake of nanoparticles by endocytosis. The internalization process may be highly dependent on factors such as size, shape, surface charge, and ligands (Harush-Frenkel et al., 2007; Iversen et al., 2011; Jin et al., 2009; Rejman et al., 2004). In general, nanoparticles composed of PEG-b-PLA, PLGA, silica nanotubes, and chitosan are subjected to clathrin-mediated endocytosis (Panyam and Labhasetwar, 2003; Qaddoumi et al., 2004; Nan et al., 2008; Ma and Lim, 2003). Allen et al. studied the cellular internalization of PEO44-b-PCL20 diblock copolymer micelles in rat pheochromocytoma PC12 cells and found that the endocytotic uptake of micelles was time, temperature, pH and energy dependent (Allen et al., 1999). Lavasanifar and coworkers studied the effect of PEO and PCL molecular weights on the internalization of PEO-b-PCL nanoparticles into MCF-7 cells, reporting that PEO and PCL molecular weights of 5000 and 13,000 g/mol, respectively, showed maximum cellular uptake of nanoparticles (Mahmud and Lavasanifar, 2005). Endocytosis inhibition studies showed that chlorpromazine reduced the cellular uptake of PEO-b-PCL micelles in a manner that was independent of the block copolymer composition, suggesting that CME was involved in the uptake of PEO-b-PCL micelles by MCF-7 cells (Mahmud and Lavasanifar, 2005). In contrast, the cellular internalization of PEO-b-poly(methacrylate) nanoparticles, liposomal doxorubicin (DOXIL®), and quantum dots nanoparticles occurs via caveolae-mediated endocytosis (Ma and Lim, 2003; Vogel et al., 1998; Nishikawa et al., 2009; Zhang and Monteiro-Riviere, 2009). However, most of the nanoparticles can gain cellular entry using more than one pathway. For example, Pluronic® P85 unimers or micelles can enter cells through CME, caveolae-mediated, and clathrin- and caveolin- independent endocytosis depending on the aggregation state (Sahay et al., 2008).  Following endocytosis, nanoparticles are internalized into early endosomes which mature as the lumen of the endosomal organelles is acidified via a vacuolar ATPase driven proton pump   36 (Lafourcade et al., 2008). The late endosomes then fuse with lysosomes which contain a variety of enzymes that may degrade the nanoparticles or drugs. The acidic environment of the endo-lysosomal compartment can be used as a strategy to trigger release of drugs by conjugating drugs to nanoparticle components through pH sensitive linkages (Shen and Ryser, 1981; Bae et al., 2005; Sahay et al., 2010b). Many drug loaded nanoparticles avoid endo-lysosomal degradation by escaping from early endosomes during the cellular internalization process. They subsequently localize with other cytoplasmic organelles inside the cell, such as mitochondria, the endoplasmic reticulum, and the Golgi apparatus (Shen and Ryser, 1981; Bae et al., 2005; Sahay et al., 2010b). Using rhodamine labeled PEG45-b-PCL23 micelles, Savic et al. provided evidence that these PEG-b-PCL micelles were internalized and localized mainly in the cytoplasm (within cytoplasmic organelles) of PC12 cells (Savic et al., 2003). In vitro subcellular distribution studies of radiolabeled PEG114-b-PCL48 nanoparticles in MDA-MB-478 and MCF-7 cells indicated that most of these nanoparticles were found to be associated with the cytoplasm, with very few nanoparticles associated with the nucleus (Lee et al., 2011a). Interestingly, it has been reported that Pluronic® P85 micelles localize within mitochondria and lead to depletion of ATP, a mechanism by which Pgp may be inhibited in MDR cancer cells (Alakhova et al., 2010). Cationic nanoparticles have been reported to bind well to plasma membranes and following endocytosis, may disrupt the endosomal membrane (due to the positive charge) and release drug into the cytoplasm (Munoz Javier et al., 2006; Hafez et al., 2001). However, the cationic charge might also lead to general membrane potential perturbation effects and therefore have background toxicity issues (Dobrovolskaia et al., 2008; Arvizo et al., 2010). Functionalization of nanoparticles may also affect the intracellular fate of nanoparticles. De La Fuente and Berry reported that following endocytosis, gold nanoparticles conjugated with tat peptide (nuclear signaling peptide) allowed translocation into the cell nucleus (de la Fuente and Berry, 2005).   37 Rapoport et al. compared the cellular uptake of the anthracyclin drug, ruboxyl, in the free form to the encapsulated micellar form in MDR cells (Rapoport et al., 2002). The free drug was sequestered in cytoplasmic vesicles and was excluded from nuclei, whereas the micellar drug was endocytosed and accumulated in cell nuclei, suggesting that the intracellular fate of the drug can be largely affected by its carrier.   1.5 Ultrasound in cancer treatment  Over the last decade, there has been significant research interest in combining ultrasound with nanoparticulate drug delivery system to achieve tumor specific release and intracellular uptake (for review, see (Rapoport, 2012)). The advancement in clinical ultrasound imaging and focusing techniques, and in combination with nanoparticulate drug delivery system may be a promising strategy for targeting and eliminating MDR cancer cells.   1.5.1 Ultrasound irradiation Ultrasonic technology has been widely used in diagnostic medicine for many years. The main advantages of ultrasound are that it is non-invasive, can penetrate tissues and focus on the interior of the body. Ultrasound is defined as acoustic waves with a frequency above the human hearing range of 20 kHz. The acoustic waves can be generated by ultrasound transducers through the piezoelectric effect, which arises from a mechanical deformation of a piezoelectric substance following application of an electric field (Meyer, 1969). Lead zirconate titanate (PZT) is the most commonly used piezoelectric material for ultrasound transducers. The polarized crystals of PZT are compressed or expanded when an electric field is applied, causing mechanical vibration. When the frequency of the vibration is above 20 kHz, an ultrasound wave is formed. Similar to optical or audio waves, ultrasonic waves can be focused, reflected and refracted in a medium   38 (Zagzebski, 1996). Clinically, ultrasound frequency has been classified into three categories: low (0-20 kHz), medium (20 kHz-1 MHz) and high (greater than 1 MHz) frequency, with intensities (W/cm2) described as low (0-0.5 W/cm2), medium (0.5-3 W/cm2) and high (greater than 3 W/cm2). A wide range of ultrasound regimes are used for either diagnostic or therapeutic applications. These include, high frequency with low intensity for imaging, high frequency, medium intensity for physiotherapy or high frequency (3 MHz) with very high intensity (greater than 1000 W/cm2) for tissue destruction (Matsunaga, 2005; Schroeder et al., 2009). Ultrasound is known to generate rapid cyclical changes in pressure resulting in biomechanical effects such as shear stress in tissues (Grainger et al., 2010). These effects are the result of acoustic cavitation, a physical phenomenon in which gas-filled cavitation bubbles are created in the liquid by a large negative pressure produced by ultrasound (Figure 1.5). The cavitation bubbles can be expanded or compressed depending on the surrounding pressures (Mason and Lorimer, 2002). In some cases, stable cavitation (Figure 1.5 A) can form, in which bubbles oscillate without collapsing, resulting in a mild perturbation of cell membranes (Williams and Miller, 1980; Rooney, 1970; Husseini and Pitt, 2008b). Deng et al. reported that stable cavitation could temporarily disrupt the cell membranes, leading to the formation of pores (Deng et al., 2004). When the cavitation bubbles collapse in the case of inertial cavitation (Figure 1.5 B), enormous amounts of pressure exceeding 104 atmospheres are released over a short time, producing shock waves that can impact the surrounding environment (Leighton, 1994; Pecha and Gompf, 2000). The inertial cavitation effect is thought to occur at low frequencies and Schlicher et al. demonstrated transient pores (<28 nm diameter) with lifetimes of less than 1 min in the plasma membrane of cells, following exposure to low frequency ultrasound (24 kHz) (Schlicher et al., 2006). The likelihood of inertial cavitation is thought to depend on the mechanical index (MI), which has been described as:   39 MI = 𝑃𝑁𝑃√𝐹𝑐 where PNP is the peak negative pressure (MPa) of the ultrasound wave and Fc is the center frequency (MHz) of the ultrasound wave (Apfel and Holland, 1991). It has been reported that the inertial cavitation occurs when the mechanical index is greater than 0.5 (Apfel and Holland, 1991).  Figure 1.5. Cavitation effect induced by ultrasound. A) stable cavitation can form, in which bubbles oscillate without collapsing, resulting in a mild perturbation of cell membranes. B) inertial cavitation occurrs when cavitation bubbles collapse producing enormous amounts of pressure (shown by black arrows) and shock waves that can disrupt cell membranes.  1.5.2 Ultrasound mediated drug delivery Therapeutic applications of ultrasound in drug delivery have been studied for several decades. An early application of ultrasound was in conjunction with transdermal delivery to increase the permeability of skin to high molecular weight proteins such as insulin, interferon-gamma, and erythropoietin (Ng and Liu, 2002; Tachibana, 1992; Mitragotri et al., 1995). It was also reported that ultrasound could be utilized to enhance the cytotoxic effect of various drugs.   40 The cytotoxicity of cytosine arabinoside (Ara-c) incubated with Chinese hamster lung fibroblasts and human leukemia cells was increased by exposing the cells to ultrasound with 48 kHz-2.6 MHz frequencies and power intensity of between 0.3-2.3 W/cm2 (Loverock et al., 1990) (Tachibana et al., 2000). Ultrasound irradiation with a frequency of 70 kHz has been shown to enhance antibiotic activity in bacteria and reduce bacterial resistance to antibiotics (Rediske et al., 1998; Rediske et al., 1999). Rapoport and co-workers showed that 70 kHz frequency ultrasound increased the intracellular uptake of doxorubicin loaded Pluronic® P105 micelles in HL-60 cells and substantially lowered IC50 values for doxorubicin (Rapoport et al., 1999; Munshi et al., 1997). These studies were the first to show the use of ultrasound to enhance the cytotoxicity of drugs loaded in polymeric micelles. In another study, the intracellular uptake of PTX loaded in methoxy poly(ethylene oxide)-co-poly(L-lactide)-tocopherol micelles was increased substantially when incubated with drug resistant human MCF-7 breast cancer cells treated with ultrasound (1 MHz, 1.7 W/cm2) (Howards et al., 2006). The degree of enhancement in drug accumulation by ultrasound is thought to be carrier-dependent. Howards et al demonstrated only minor increases in PTX accumulation in MCF-7 cancer cells with ultrasound treatment (1MHz, 1.7W/cm2, 30 seconds) using Taxol® but demonstrated 20 fold increases when using a poly(ethylene oxide)-co-poly(L-lactide)-tocopherol polymeric micelle formulation of the drug (Howards et al., 2006). High intensity focused ultrasound (HIFU) is used to thermally ablate tumors (Schroeder et al., 2009). HIFU provides millimeter level accuracy in beam focus (typically at 1Mhz) and may be combined with simultaneous image guidance to allow for precise targeting of diseased tissues in most regions of the body. Adjunct chemotherapy is usually delivered, before, during, or after HIFU to kill cancer cells in regions adjacent to the ablated regions (Mo et al., 2012). Real time imaging of blood flow may also be achieved using microbubble contrast agents with   41 ultrasound (Schroeder et al., 2009). These advances in both diagnostic and therapeutic applications of ultrasound, together with the increased clinical use of focused ultrasound equipment have led to the search for pharmaceutical applications for ultrasound. These include, the encapsulation of drugs in microbubbles, liposomes or micelles for ultrasound- induced localized drug release in target tissues (Schroeder et al., 2009; Eisenbrey et al., 2010; Husseini and Pitt, 2008b; Xing et al., 2008). For the past decades, microbubbles have been used extensively in ultrasound application as the ultrasound contrast agent (Dijkmans et al., 2004). Recently, there has been significant research interest in the development of drug encapsulated microbubbles with the goal of targeting tumor specific cells with real time imaging (Kooiman et al., 2014). Ji et al. encapsulated curcumin in ultrasound-activated perflurocarbon microbubbles stabilized by PEG44-b-PCL22 block copolymers and demonstrated that upon ultrasound irradiation (20 kHz for 60 seconds), 90% of encapsulated curcumin was immediately released from the microbubbles (Ji et al., 2014). After administration of the microbubbles into tumor bearing mice, a real time ultrasound contrast imaging of tumour was generated allowing for ultrasound guided drug delivery (Ji et al., 2014). The tumor inhibition observed for mice treated with curcumin-loaded microbubbles in combination with ultrasound irradiation was significantly higher than the non-ultrasound group (Ji et al., 2014). Eisenbrey et al. encapsulated doxorubicin in microbubbles stabilized by PLA (Eisenbrey et al., 2009). When irradiated by ultrasound (5 MHz), it was shown that the formulation was more effective in inhibiting MDA-MB-231 breast cancer cell growth when compared to cells without ultrasound treatment. The abundance of advanced ultrasonic equipment and devices in hospitals and clinical environments suggests that applications of this technology should be readily transferable to the field of localized nanoparticulate drug delivery to tumors. Using non-focused ultrasound from a commercial sonoporator (Sonidel®) at 1MHz frequency with contact gel to externally sonicate   42 subcutaneous tumors in mice, Nomikou et al. demonstrated improved dispersion of the dye, Evan’s Blue, and camptothecin throughout the subcutaneous tumor masses (Nomikou et al., 2010). Grainger et al. used non-focused ultrasound (1MHz), to increase the penetration of fluorescently labeled polystyrene nanoparticles with sizes ranging from 20-100 nm into MCF-7 breast cancer spheroids (Grainger et al., 2010). Other workers have shown that ultrasound (690 kHz-1.7 MHz) may enhance the permeability of tumor vasculatures to macromolecules and that the blood brain barrier may be disrupted using magnetic resonance guided, focused ultrasound to effectively deliver liposomes or immunotherapeutics (Park, 2010; Treat et al., 2007; Raymond et al., 2008).     1.5.3 Mechanisms of ultrasound enhanced cytotoxicity from drug loaded nanoparticles Three theories have been proposed in an attempt to explain enhanced cellular accumulation and cytotoxicity caused by ultrasound irradiation of cells incubated with drug loaded nanoparticles, as shown schematically in Figure 1.6. Firstly, ultrasound may trigger drug release from nanoparticles at the cell membrane (Figure 1.6 A). By using fluorescence detection techniques, it was found that doxorubicin release was enhanced from Pluronic® P105 micelles when triggered by ultrasound (70 KHz), resulting in enhanced doxorubicin uptake in HL-60 cells (Marin et al., 2001a). Husseini et al. used ultrasound of a fixed frequency (70 kHz) but at different power densities ranging from 0.01 to 0.75 W/cm2 for studying the release of doxorubicin from Pluronic® P105 micelles (Husseini et al., 2005). The results of the studies showed a strong correlation between percent drug released and occurrence of collapse cavitation. The authors attributed the drug release to collapse cavitation that perturbed the structure of the micelle and released drug (Husseini et al., 2005). A second mechanism has been suggested by Rapoport et al. and Sheikov et al (Figure 1.6 B) (Rapoport, 2004; Sheikov et al., 2006) which   43 proposes that ultrasound (70 kHz-0.69 MHz) may enhance the rate of endocytosis of doxorubicin loaded polymeric micelles and horseradish peroxidase into cells. However, the actual mechanism of the ultrasound effect on endocytosis is unknown. The third proposed mechanism of ultrasound enhanced drug delivery is via enhancement of membrane permeability (Figure 1.5 C) (Husseini and Pitt, 2009). It has been suggested that the formation of membrane pores is a result of acoustic cavitation which produces shockwaves that temporarily disrupt the cell membranes, leading to the formation of transient pores (Deng et al., 2004). Nanoparticles can then pass through these pores into the cytoplasm of the cell. The oscillatory motion of the medium created by ultrasound may also enhance the transport of free drugs, increasing the rate of drug transport into the cell (Starritt et al., 1989). It is also possible that all three mechanisms may be responsible for ultrasound enhanced drug delivery into cells.     44 Figure 1.6 Proposed mechanisms of ultrasound irradiation mediated enhanced drug uptake. A) enhancement of drug release, B) increased rates of endocytosis, C) enhancement of membrane permeability by the formation of transient membrane pores.   A) B) C)   45 1.5.4 Effect of ultrasound on cytotoxicity of drug loaded nanoparticles in MDR cells Ultrasound has been shown to increase cellular accumulation of Pgp substrates in various MDR cell lines (Ng and Liu, 2002). Liu et al. reported that ultrasound (1 MHz, 0.4 W/cm2) increased cellular uptake and cytotoxicity of doxorubicin in human metastatic lung carcinoma MV522/Q6 MDR cancer cell lines and that drug accumulation was higher as compared to cells treated with verapamil and cyclosporin A analog, suggesting that ultrasound treatment could be a potential strategy to enhance drug accumulation in MDR cells (Liu et al., 2001a; Liu et al., 2001b). A similar result was published by Shao et al, showing enhanced cellular uptake and retention of R-123 and adriamycin in ultrasound (0.8 MHz, 0.43 W/cm2) treated HepG2/MDR (human hepatocarcinoma) cells (Shao et al., 2008). Rapoport found that the cytotoxicity of Pluronic®P105 encapsulated doxorubicin was enhanced by ultrasound in ovarian carcinoma MDR cells (Rapoport, 2004). It was also demonstrated that ultrasound (1MHz, 1.7W/cm2, 30 seconds) enhanced the drug accumulation of poly(ethylene oxide)-co-poly(L-lactide)-tocopherol polymeric micellar PTX in drug resistant breast cancer cells (Howards et al., 2006). Mohan and Rapoport prepared doxorubicin encapsulated PEG45-b-PCL23 micelles and incubated with drug resistant A2780 ovarian cancer cells. Upon ultrasound treatment (3 MHz, 2 W/cm2), more doxorubicin was observed in the nucleus as compared to non-ultrasound treated groups, resulting in enhanced cytotoxicity of the drug (Mohan and Rapoport, 2010). It has been proposed that the hyperthermia generated by ultrasound may enhance membrane permeation of Pgp substrates, leading to greater cellular drug accumulation (Ng and Liu, 2002; Liu et al., 2001a; Liu et al., 2001b). The ultrasound enhanced uptake of nanoparticles in MDR cells has been proposed to by-pass drug efflux transporters on the MDR cell membranes, since the internalization of nanoparticles proceeds via membrane pores induced by ultrasound irradiation, avoiding contact with the transmembrane drug efflux transporters (Rapoport, 2012). Sun et al. showed that the   46 expression of Pgp in SK-N-SH/MDR1 cells (human neuroblastoma cell line) was reduced following exposure to low-intensity pulsed ultrasound (0.3 MHz, 1W/cm2). It was proposed that the reduction in protein expression level was related to the disruption of the cell membrane (Sun et al., 2013). Ultrasound irradiation (0.8 MHz, 0.43 W/cm2) also reversed MDR in HepG2/ADM hepatocarcinoma cells by decreasing mRNA expression of Pgp and MRP, and increasing the expression of the anti-apoptotic proteins, Bax (Wu et al., 2011). However, possible mechanisms for ultrasound induced alteration in protein expression are unknown.   1.5.5 Ultrasound device design and characteristics In this work, a micro-ultrasonic transducer device was designed and developed by our group and collaborators, that provides ultrasonic exposure to cells within the wells of a 48 well plate in a highly controlled fashion (Figure 1.7 A). This ultrasound device provides us with an opportunity to investigate the influence of ultrasound on the cellular accumulation of nanoparticulate PTX in MDR cell lines.  The PZT transducer was made from a lead zirconate titanate ceramic. The electrical signal from a signal generator can be amplified by a power amplifier before being applied to the transducers. The transducer is actuated at 4 MHz frequency resulting in a sine wave form with a period of 0.25 μs. To avoid overheating the ultrasound transducer, burst mode is used to allow for intervals between waves. In burst mode, the system generates 4 MHz frequency waveform for 12.5 ms in every 50 ms repetition period, resulting in 50,000 cycles in each burst. A schematic of the ultrasound output is illustrated in Figure 1.7 B. Overall, this system delivers: 4 MHz ultrasound, 50 ms repetition period, 25% duty cycle with a power intensity at specimen of 32 W/cm2. There is no significant increase in temperature (less than 1°C) caused by the transducer at the transducer-water interface as measured by a thermocouple system.   47  Figure 1.7. A) A schematic representation of a 48-well cell culture plate, adapted for exposure with ultrasound from a custom-designed ultrasonic-transducer. The height of cell culture medium in wells is around 8mm. B) A schematic representation of 4 MHz, burst-mode ultrasound waveform with 50 ms burst period. The diagrams were published (Jackson et al., 2011).  1.6 Thesis goal and research objectives The overall goal of the work described in this thesis was to evaluate two different strategies to increase cellular accumulation of nanoparticulate taxanes in both drug sensitive and drug resistant cell lines. The first strategy was to determine whether a difference in   48 nanoparticulate structure (“fluid-like” core micelles versus “frozen core” nanospheres) along with exposure of resistant cells to a Pgp inhibitor, influenced drug accumulation and cytotoxicity. The second strategy employed the use of a custom-fabricated device able to expose cells to a very short burst of ultrasound irradiation to determine whether ultrasound irradiation influenced the cellular uptake and cytotoxicity of drug loaded nanoparticles. The MePEG-b-PCL diblock copolymers used in this work have been shown to possess Pgp inhibitory activity and to form nanoparticles well-suited to drug encapsulation. We also reported ultrasound mediated intracellular nanoparticulate drug accumulation. Taken together with the evidence that nanoparticles and ultrasound irradiation may provide a means of by-passing Pgp, and their ability to passively target tumors via the enhanced permeability and retention effect, we believe that the proposed studies will lead to the rational design of a diblock copolymer-based nanoparticulate drug delivery system with enhanced efficacy against drug resistant tumors. The working hypotheses for this research project were as follows: 1. Taxane loaded in fluid-core MePEG114-b-PCL19 micelles will be more effective in sensitizing MDR cells than the solid-core MePEG114-b-PCL104 nanospheres.  2. The release of Pgp inhibitor, MePEG17-b-PCL5 from the taxane loaded PCL200/PCL5 mixed molecular weight nanoparticles following cellular uptake will inhibit Pgp mediated drug efflux and enhance cytotoxicity in MDR cells.  3. The application of ultrasound irradiation to MDR or drug sensitive cells will result in enhanced cellular accumulation and cytotoxicity of taxane loaded nanoparticles. The objectives of this research project were as follows: 1. To evaluate the cellular accumulation and cytotoxicity of PTX formulated in MePEG114-b-PCL19 micelles and MePEG114-b-PCL104 nanospheres incubated with drug sensitive   49 (MDCKII) and drug resistant (MDCKII-MDR1) cell lines with or without exposure to the Pgp inhibitor, MePEG17-b-PCL5 (Chapter 2).  2. To determine the cytotoxicity of PTX and DTX loaded nanospheres composed of MePEG114-b-PCL200 with or without coencapsulation of the Pgp inhibitor, MePEG17-b-PCL5, to form “mixed MW PCL200/PCL5 nanoparticles” (Chapter 3).   3. To determine the effect of an optimized ultrasound irradiation regime on the cellular accumulation and cytotoxicity of PTX loaded in MePEG-b-PDLLA micelles in drug sensitive and Pgp overexpressing cell lines (Chapter 4).     50 Chapter  2: The combined use of paclitaxel-loaded nanoparticles with a low molecular weight copolymer inhibitor of p-glycoprotein to overcome drug resistance 3 2.1 Introduction  There has been a great deal of interest in the development of amphiphilic block copolymer micelles for the delivery of the hydrophobic anticancer agent, PTX (Letchford et al., 2009; Sarisozen et al., 2012; Zhang et al., 1996). Due to their amphiphilic nature, diblock copolymers are capable of self-assembling at the critical micelle concentration to form micelles characterized by a hydrophobic core surrounded by a highly water bound corona. The core is usually composed of biocompatible, biodegradable polyesters such as PLA or PCL that solubilize PTX and the hydrophilic polymer is typically MePEG (Gaucher et al., 2005). The micellar structure is considered dynamic, as there is an equilibrium between unimers in solution and those participating in the nanoparticle structure resulting in a “fluid-like” core. These systems offer several formulation advantages for the delivery of PTX. By eliminating the use of the solubilizing agent, CrEL, currently used in the commercial formulation, Taxol®, any toxicity associated with the use of this agent is prevented. Furthermore, copolymer micelles are capable of enhancing PTX solubility from 1µg/ml in water to over 5mg/ml in a micellar formulation (Zhang et al., 1996). Although these nano-sized structures might offer the potential for prolonged circulation and passive targeting of the drug (Iyer et al., 2006), dynamic micelles are usually disrupted in the presence of biological milieu resulting in the rapid release and elimination of their hydrophobic drug payload (Chen et al., 2008a; Savic et al., 2006; Chen et al., 2008b; Letchford and Burt, 2012).                                                  3A version of this chapter has been published. Wan et al., (2013) Int. J. Nanomedicine. 8, 379-391.   51 Our group has investigated the use of the nanoparticles formed from MePEG-b-PCL for the delivery of PTX. We have demonstrated that there is a shift in the physicochemical properties and hence, the performance of these nanoparticles as drug delivery systems, as the molecular weight of the hydrophobic block increases (Letchford et al., 2009). Two types of nanoparticles were fabricated using the same MePEG block length of 114 ethylene glycol repeat units, with either a short PCL block length of 19 caprolactone repeat units (MePEG114-b-PCL19) to form micelles, or a longer PCL block length of 104 repeat units (MePEG114-b-PCL104) to form kinetically “frozen core” structures termed nanospheres. We have shown that MePEG114-b-PCL104 nanospheres solubilize more PTX, release the drug at a more sustained rate and are more stable in the presence of human plasma as compared to their micelle counterparts, likely due to the “solid-like” core of these nanoparticles (Letchford et al., 2009). However, whether the differences in the physicochemical properties and the stability of these nanoparticles affect the cellular uptake of loaded PTX or the copolymer themselves is not known. Aside from the poor aqueous solubility, another significant barrier in the treatment of cancer with PTX is its efflux by Pgp. Pgp is a transmembrane drug efflux protein and a member of the ATP-binding cassette superfamily. Pgp is responsible for cellular MDR to a wide variety of anticancer drugs, including PTX (Szakacs et al., 2006). Numerous strategies have been investigated to overcome the efflux of Pgp substrates, including the encapsulation of these drugs in nanoparticles to enable the entry into cells by endocytosis and therefore bypassing Pgp (Yan et al., 2010; Lee et al., 2011b). Additionally, the extensive work by Kabanov and co-workers has demonstrated that certain commercially available Pluronic™ amphiphilic agents are capable of inhibiting Pgp (Kabanov et al., 2002b). This led to several reports evaluating multi-component or mixed polymeric nanoparticles incorporating one or more Pluronics for both solubilizing taxanes and inhibiting Pgp function (Mei et al., 2009; Mu et al., 2010; Zhang et al., 2011; Zhang et al.,   52 2010c). However, the potential drawback of using Pluronics is the non-degradability and lack of renal excretion for high molecular weight members of this copolymer class (Besheer et al., 2009; Ruel-Gariepy and Leroux, 2004). Previous work by our group has demonstrated that a very low molecular weight MePEG-b-PCL copolymer (MePEG17-b-PCL5) is an effective modulator of P-gp. We reported on the ability of MePEG17-b-PCL5 used at concentrations above and below the critical micelle concentration to enhance the accumulation of Pgp substrates, including PTX, in Pgp over-expressing cells (Elamanchili et al., 2009; Zastre et al., 2008). Given the dual functionality of MePEG-b-PCL as micellar or nanosphere PTX carriers and Pgp modulator, depending only on block lengths, we postulated that the uptake and cytotoxicity of PTX-loaded MePEG-b-PCL nanoparticles in drug sensitive or multi-drug resistant (MDR) cell lines would be dependent on the composition of the diblock copolymer carrier and co-administration of MePEG17-b-PCL5.  The objectives of this work were to evaluate the cellular uptake and cytotoxicity of PTX formulated in MePEG114-b-PCL19 micelles or MePEG114-b-PCL104 nanospheres and administered to drug sensitive (MDCKII) and drug resistant (MDCKII-MDR1) cell lines, with or without exposure to the Pgp inhibitor, MePEG17-b-PCL5. Our results show that PTX loaded micelles are more effective than nanospheres in inhibiting proliferation in MDCKII-MDR1 cells, but are not different in MDCKII cells, and that there are dramatic effects on reversing chemoresistance in MDCKII-MDR1 cells in the presence of MePEG17-b-PCL5.    2.2 Materials and methods 2.2.1 Materials Paclitaxel (PolymedT Inc, Houston, TX), radioactive paclitaxel (Moravek Radiochemicals, Brea, CA), tetramethylrhodamine-5-carbonyl azide, 4’, 6-diamino-2-  53 phenylindole dihydrochloride (DAPI) (Invitrogen, Grand Island, NY), Dulbecco’s modified eagle’s media (DMEM) supplemented with 5% fetal bovine serum (FBS) and 1% penicillin/streptomycin, Hank’s buffered salt solution (HBSS) (Invitrogen), MTS cell proliferation assay kit (Promega, Madison, WI), CytoTox 96® NonRadioactive Cytotoxicity Assay (Promega, Madison, WI), methoxy poly(ethylene glycol) (Fluka, Bucks SG, Switzerland), BCA protein assay kit (Thermo Scientific, Rockford, IL), ε-caprolactone (Fluka), micro-BCA protein assay kit (Pierce, Rockford, IL), stannous octoate (Sigma-Aldrich Canada Ltd, Oakville, ON, Canada), N, N-dimethylformamide (DMF) (Fisher Scientific Co., Ottawa, On, Canada) were used as supplied without further purification. MDCKII and MDCKII-MDR1 cells were kind gifts from Dr. Piet Borst (National Cancer Institute, the Netherlands). Amphiphilic diblock copolymers composed of methoxy poly(ethylene glycol) and poly(caprolactone) were synthesized and characterized as previously described (Letchford and Burt, 2012; Letchford et al., 2004). These copolymers are abbreviated MePEGm-b-PCLn where m and n are the number of repeat units of ethylene glycol and caprolactone, respectively. In these studies three copolymers were investigated, MePEG114-b-PCL104, MePEG114-b-PCL19 and MePEG17-b-PCL5.  2.2.2 Synthesis and characterization of MePEG-b-PCL diblock copolymers  Methoxy terminated poly(ethylene glycol) (MePEG) with a molecular weight of either 750 g/mol (MePEG17-b-PCL5) or 5000 g/ mol (MePEG114-b-PCL19 and MePEG114-b-PCL104) was combined with ε-caprolactone in weight ratios of either 60:40 (MePEG17-b-PCL5), 30:70 (MePEG114-b-PCL19), or 70:30 (MePEG114-b-PCL104) with a total mass of 50 g. The reactants were reacted at 140 °C for 4 h (MePEG17-b-PCL5) or 24 h (MePEG114-b-PCL19 and MePEG114-b-PCL104) in a sealed round-bottomed flask containing 35 ml of anhydrous toluene (Sigma-  54 Aldrich) using 0.15 ml of stannous octoate as a catalyst. The products were purified by dissolving the copolymers in chloroform followed by precipitation with a 70/30 mix of hexane and diethyl ether. Using gel permeation chromatography (GPC) with polyethylene glycol standards in the range of 400 – 22800 g/mol (Polymer Laboratories Inc., Amherst, MA, USA) for the low molecular weight copolymer or polystyrene standards in the range of 4000 – 100000 g/mol (Polysciences Inc., Warrington, PA, USA) for the high molecular weight copolymer, the molecular weights and molecular weight polydispersity indexes were determined. Waters Styragel columns (HR3 and HR1) connected in series were used and the detection was through a Waters model 2410 refractive index detector. The compositions of the copolymers were determined by proton NMR spectra of 10% w/v solutions of the copolymers in deuterated chloroform, obtaining from a 400 MHz Bruker Advance II+ spectrometer (Bruker Corporation, Milton, ON). The spectra were analyzed using MestRe-C 2.3a software. The degree of polymerization (DPn) of copolymers were calculated using peaks situated around 1.3 and 1.55 ppm from the caprolactone methylene protons and the peaks at 3.55 ppm from the MePEG methylene protons. The characterization of synthesized MePEG-b-PCL diblock copolymers is included in Table 2.1.     55 Table 2.1 Characterization of synthesized MePEG-b-PCL diblock copolymers. Feed ratioa DPnb MW.Theoc (g/mol) MW.NMRd (g/mol) MW.GPCe (g/mol) PDIf Diameterg (nm) 70:30 MePEG114-b-PCL19 7143 6955 7698 1.17 32.45±0.03h 30:70 MePEG114-b-PCL104 16651 17460 14532 1.3 55.48±0.62i 60:40 MePEG17-b-PCL5 1250 1320 1539 1.07  a. Feed weight ratio of MePEG:caprolactone.  b. Degree of polymerization (DPn) of MePEG:MWMePEG/44, DPn of PCL: rounded off value determined by NMR.  c. Theoretical MW based on feed weight ratio.  d. MW determined by NMR.  e. MW determined by GPC. f. Polydispersity index determined by GPC.  g. Nanoparticle hydrodynamic diameter determined by PCS based on scattering intensity at 37 °C at [copolymer] of 0.1 %(w/v).  Diameters did not change upon loading of PTX. h. A representative size distribution by intensity diagram of MePEG114-b-PCL19 nanoparticles.  i. A representative size distribution by intensity diagram of MePEG114-b-PCL104 nanoparticles.      56 2.2.3 Fluorescent labeling of MePEG-b-PCL copolymers  The copolymers MePEG114-b-PCL19 and MePEG114-b-PCL104 were fluorescently labeled with tetramethylrhodamine-5-carbonyl azide (TMRCA) as described previously (Luo et al., 2002). Briefly, 20 mg of MePEG114-b-PCL19 or MePEG114-b-PCL104 copolymers were reacted with 6 mg of TMRCA in 20 ml of anhydrous toluene under a nitrogen atmosphere for 24 hours at 80°C in a sealed round bottom flask. After termination of the reaction, the reaction mixture was placed in 3500 MWCO SnakeSkin® dialysis membranes (Thermo Scientific) and dialyzed against DMF for one week, with frequent exchanges of fresh DMF, to remove unreacted TMRCA. The dialysis tubing is made of high performance regenerated-cellulose, which has fair resistance to DMF. The dialysis tubing was changed several times to avoid the leakage of polymers. The mixture was then dialyzed against distilled water for 3 days to remove any organic solvent. The dialysis tube and distilled water were changed frequently and the dialysis was terminated when no trace of rhodamine in the external solvent was detected. The aqueous mixture was then lyophilized. A small amount of the labeled copolymers was dissolved in a DMF and the TMRCA labeling efficiency of the diblock copolymers was determined by spectrofluorometry against a calibration curve of free TMRCA in DMF using a Synergy MX (Biotek, Winooski, VT) fluorescence plate reader and a Nunc 96-well plate (Thermo Scientific, Rochester, NY) using an excitation wavelength of 548 nm and an emission wavelength of 572 nm. The fluorescent labeling efficiency for MePEG114-b-PCL19 and MePEG114-b-PCL104 copolymers was approximately 50.3 and 63.3%, respectively.  2.2.4 Preparation of MePEG-b-PCL nanoparticles   A previously described nanoprecipitation technique was used to prepare nanoparticles (Letchford and Burt, 2012; Letchford et al., 2009). Briefly, 30 mg of either MePEG114-b-PCL104   57 or MePEG114-b-PCL19 were dissolved in 0.5 ml of DMF. In formulations in which PTX was to be included, 0.5 mg of PTX was also added to the DMF solution. This solution was added drop wise to 2 ml of rapidly stirring phosphate buffered saline (PBS) (0.01 M, pH 7.4). The DMF was removed from the nanoparticle dispersion by dialysis overnight against 4L of PBS using 3500 MWCO Spectra/Por® dialysis membranes (Spectrum Laboratories, Inc.). Following dialysis, the nanoparticle dispersions were made up to 3 ml with PBS. The drug loading was determined using a previously described HPLC assay (Liggins et al., 2000). As discussed in our previous publications, nanoparticles formed from MePEG114-b-PCL19 are referred to as micelles, whereas those formed from MePEG114-b-PCL104 are referred to as nanospheres.  2.2.5 Immunodetection of P-glycoprotein Cells were centrifuged and resulting cell pellets were lysed for 30 min using ice-cold RIPA buffer. Samples were centrifuged at 500 g for 10 min at 4°C and the supernatants were recovered. Protein concentration of the supernatant was determined using the micro-BCA protein assay kit (Pierce, Rockford, IL). An aliquot of supernatant (containing 40 μg of protein) was resolved on a 10% SDS-polyacrylamide gel and electroblotted onto a 0.45 μm polyvinylidene difluoride membrane. The membrane was incubated in block solution for 45 min (1% nonfat dried milk and 0.1% Tween-20 in PBS) then incubated with 1:200 dilution of C210 primary antibody and a 1:4000 dilution of anti-actin antibody AC-40, followed by an incubation with 1:2000 dilution of rabbit IgG anti-mouse HRP-conjugated antibody. The Pgp was visualized using chemiluminescence (Perkin Elmer, Woodridge, On, Canada) as per manufacturer’s protocol.     58 2.2.6 Cytotoxicity of blank MePEG-b-PCL block copolymers Madin-Darby Canine Kidney (MDCKII) and MDCKII-MDR1 cells were seeded at 1500 cells/well in 96-well plates and allowed to grow overnight (5% CO2/95% humidity at 37°C) in DMEM. To investigate the long-term cytotoxicity of the MePEG114-b-PCL19 or MePEG114-b-PCL104 nanoparticles or MePEG17-b-PCL5, with no drug present, the cells were incubated for 3 days in the presence of 200 μl of MePEG114-b-PCL19 or MePEG114-b-PCL104, ranging in concentration from 0-0.4% w/v (0-4 mg/ml) or MePEG17-b-PCL5 from 0-5% w/v (0-50 mg/ml), prepared in culture medium. As a control, cells were grown in culture medium in the absence of copolymer. Cell proliferation was measured using the MTS cell proliferation assay kit according to the manufacturer’s instruction. Cell viability was expressed as a percentage of the ratio of absorbance after treatment as compared to the absorbance of control (no treatment). To assess any immediate cytotoxicity of MePEG114-b-PCL19, MePEG114-b-PCL104 or MePEG17-b-PCL5 diblock copolymers due to cell lysis, cells were incubated for 90 minutes with copolymers ranging in concentration from 0 – 5% w/v. Cytotoxicity was measured using the lactate dehydrogenase (LDH) release assay kit and 2% TritonX-100 was used as the positive control.   2.2.7 Cytotoxicity of PTX loaded MePEG-b-PCL nanoparticles Nanoprecipitation was used to prepare MePEG114-b-PCL19 micelles and MePEG114-b-PCL104 nanospheres, at a copolymer concentration of 1% w/v with a drug loading of 166 μg/ml PTX in PBS. These nanoparticles were subsequently diluted in culture media to prepare nanoparticle dispersions with PTX concentrations ranging from 2 - 32 μg/ml with a maximum copolymer concentration of 0.2% w/v. MDCKII and MDCKII-MDR1 cells were incubated in the presence of the PTX loaded nanoparticles with these varying PTX loadings in culture media for 90 minutes or 3 days. Nanoparticles without drug were used as the control. To investigate   59 whether co-administration of PTX loaded micelles or nanospheres with MePEG17-b-PCL5 inhibits the proliferation of MDCKII and MDCKII-MDR1 cells, these cell lines were incubated with micelles or nanospheres with a PTX loading of 16 µg/ml and MePEG17-b-PCL5 at a concentration of 0.05% w/v in culture media for 3 days followed by determination of cell viability by MTS assay. The effect of a sustained 3 day incubation of MePEG17-b-PCL5 at varying concentration after an initial 90 minute incubation with drug loaded MePEG114-b-PCL19 micelles and MePEG114-b-PCL104 nanospheres was also investigated. Micelles and nanospheres prepared with a copolymer concentration of 0.2% w/v and a PTX loading of 32 μg/ml were first incubated with MDCKII and MDCKII-MDR1 cells for 90 minutes and then removed and the cells were washed with warm HBSS. Subsequently, the cells were incubated with MePEG17-b-PCL5 at concentrations between 0 - 0.25% w/v in culture media for 3 days to determine the concentration of MePEG17-b-PCL5 that was most effective at inhibiting cell proliferation after this short exposure to PTX loaded nanoparticles. Cell viability was determined using the MTS cell proliferation assay kit as described above. The IC50 of PTX loaded nanoparticles was then determined in the presence of MePEG17-b-PCL5 at this concentration. In order to do so, cells were first incubated for 90 minutes in the presence of nanoparticles loaded with PTX ranging in concentration from 0 – 60 µg/ml. The drug-loaded formulations were then removed and the cells washed with HBSS at 37°C followed by incubation with 0.05% w/v MePEG17-b-PCL5 in culture media for 3 days. Cell viability was determined by MTS assay.  2.2.8 Drug accumulation Micelles and nanospheres at a copolymer concentration of 0.2% w/v and a PTX concentration of 32 μg/ml, along with a trace amount of 3H-PTX, were prepared using the methods described above. MDCKII and MDCKII-MDR1 cells were plated in 48-well plates at a   60 density of approximately 10,000 cells/well and grown for 2 days. Drug loaded micelle and nanosphere dispersions were equilibrated for 30 min in a 37°C water bath followed by addition to the cells and a 90 min incubation. Following the incubation period, the supernatant was removed and the cells washed with ice-cold PBS to remove extracellular PTX. The cells were lysed with PBS containing 1.33% Triton X-100 and 33% DMSO and the cell lysate was analysed by β scintillation counting using a LS6000TA scintillation counter (Beckman Instruments Inc., Fullerton, CA). A set of 3 wells per experimental plate was collected individually for total protein content as determined by BCA assay. The intracellular accumulation of PTX quantified using calibration curves prepared from known concentration of 3H-PTX and the concentration of PTX was normalized to total protein content.  2.2.9 Polymer uptake and fluorescence confocal microscopy  Drug loaded nanoparticles composed of TMRCA labeled and unlabeled MePEG114-b-PCL19 and MePEG114-b-PCL104 diblock copolymers with the same concentrations of TMRCA (20 μM) were prepared using a nanoprecipitation method as described above. MDCKII and MDCKII-MDR1 cells were seeded on poly-L-lysine-coated coverslips in 12 well plates at approximately 5000 cells/well and incubated overnight. TMRCA-labelled nanoparticle dispersions (300 μl of a 1 mg/ml dispersion) were incubated with cells for 90 mins and washed once with an acidified wash (0.5 M NaCl, 0.2 M CH3COOH, pH 2.5) and once with HBSS (pH 7.4) to remove any surface associated copolymer. The coverslips with the adhered cells were removed from the plates and the cells were fixed with paraformaldehyde (4%) on ice for 20 min and the nuclei were stained with DAPI according to manufacturer’s instructions. The coverslips were then mounted cell-side-down on glass slides and microscopy studies were performed on a Fluoview® FV-1000 (Olympus, Japan) inverted confocal microscope. The laser wavelengths   61 used were 568 nm and 405 nm for imaging of TMRCA and DAPI, respectively. Differential interference contrast was also performed to visualize cell membranes and was activated with the 405 nm laser. The fixed cells were observed using objective lenses at 40X or 100X magnification. All images were generated under the same microscope settings to allow for consistent comparison. The figures were created using FV10-ASW Ver3.0 software (Olympus, Japan). Approximately 50 confocal sections from the basal to the apical cell side with an increment of 0.2 µm were captured for z-stacking analysis. To quantitate the amount of polymer taken up by cells, cells were seeded in 12 well culture plates as described above but without coverslips. After incubation with TMRCA labeled nanoparticles, the cells were lysed with 1% TritonX-100 in HBSS. TMRCA copolymer was quantified by analyzing the cell lysate in a Synergy MX fluorescence plate reader (Biotek, Winooski, VT) with excitation at 548 nm and emission at 572 nm, using a calibration curve of TMRCA labeled copolymer in cell lysate solution over a concentration range of 25 to 150 μg/ml. The intracellular content of the fluorescent-labeled polymer was expressed as the amount of polymer (mg) over the cellular protein content (mg) determined by BCA assay as described above.    2.2.10 Statistical analysis The experiments were performed in three individual experiments. The data are presented as mean ± standard error of the mean. The data were analyzed by Student’s t-test using Prism 5.0 software (GraphPad Software Inc.) with p < 0.05 considered to be statistically significant.       62  2.3 Results 2.3.1 Immunodetection of P-glycoprotein The expression of Pgp in MDCKII and MCKII-MDR1 was determined by Western blot analysis. As shown in Figure 2.1, high expression of Pgp was measured for MDCKII-MDR1 cells whereas no Pgp expression was measured for MDCKII.        Figure 2.1 Detection of Pgp expression in MDCKII and MDCKII-MDR1 cells by Western blot analysis.  2.3.2 Cytotoxicity of diblock copolymers The cytotoxicity of MePEG114-b-PCL19 micelles and MePEG114-b-PCL104 nanospheres over the concentration range of 0-0.4 % (w/v) in the absence of drug was investigated for MDCKII and MDCKII-MDR1 cells. As shown in Figure 2.2, the cell viability of MDCKII and MDCKII-MDR1 cells after 3 days incubation with the nanoparticulate formulations decreased slightly, to approximately 90% cell viability, with an increase in copolymer concentration above 0.1% w/v. However, this decrease in cell viability was not statistically different to the cell viability values determined at concentrations below 0.1% w/v copolymer. The short term cytotoxicity of both MePEG114-b-PCL19 micelles and MePEG114-b-PCL104 nanospheres on β-Actin Pgp MDCKII MDCKII-MDR1    63 MDCKII and MDCKII-MDR1 cell lines was determined using an LDH-release assay which indicates the amount of cell lysis over 90 minutes.  Figure 2.2 Cell viability of a) MDCKII and b) MDCKII-MDR1 cells in the presence of various concentrations of () MePEG114-b-PCL19 or (☐) MePEG114-b-PCL104 diblock copolymer nanoparticles. Cells were seeded in 96-well plates and treated with various concentrations of the nanoparticles for 3 days at 37°C. Cell viability was determined using an MTS assay. Indicated values are mean (±SEM) of five independent experiments (n=5). As shown in Figure 2.3, less than 3% cell lysis was measured for all polymers and cell types up to a copolymer concentration of 1% w/v. Residual DMF in the formulations may be of concern when formulating nanoparticles by the nanoprecipitation and dialysis method which may impact cell viability. In our formulations the final concentration of DMF would be very low (calculated to be less than 0.005%) due to rapid exchange of DMF with buffer since the volume of buffer 0.0 0.1 0.2 0.3 0.4020406080100MePEG114-b-PCL19MePEG114-b-PCL104Diblock Copolymer Concentration (% w/v)% Cell Viability0.0 0.1 0.2 0.3 0.4020406080100MePEG114-b-PCL19MePEG114-b-PCL104Diblock Copolymer Concentration (% w/v)% Cell Viabilitya)b)  64 used was very large and the high molecular weight cut off of the dialysis membrane relative to the molecular weight of DMF. This small residual amount of DMF in the formulations did not appear to have a negative effect on the cells as cell viability remained high across all concentrations tested (Figure 2.2).   Figure 2.3 Cell lysis of (a) MDCKII and (b) MDCKII-MDR1 cells treated with () MePEG114-b-PCL19 and () MePEG114-b-PCL104 diblock copolymer nanoparticles at varying concentrations for 90 minutes. Cell lysis was measured by LDH release assay. The indicated values are the mean of three independent experiments (±SEM).    0.0 0.5 1.00246810MePEG114-b-PCL19MePEG114-b-PCL104Diblock Copolymer Concentration (%w/v)% LDH Release0.0 0.5 1.00246810MePEG114-b-PCL19MePEG114-b-PCL104Diblock Copolymer Concentration (%w/v)% LDH Releasea)b)  65 2.3.3 Influence of MePEG17-b-PCL5 on cell viability The influence of MePEG17-b-PCL5 on MDCKII and MDCKII-MDR1 cell viability was investigated by incubating the diblock copolymer at various concentrations with cells in the presence of culture medium for 3 days. The cell viability was measured using an MTS assay. As shown in Figure 2.4A, the viability of cells was unaffected by incubation with MePEG17-b-PCL5 at concentrations up to 1% (w/v) beyond which, cell viability rapidly declined. The immediate cytotoxicity of MePEG17-b-PCL5 on MDCKII and MDCKII-MDR1 cells was measured using the LDH release assay. As shown in Figure 2.4B, for both cell lines the amount of cell lysis was below 5% at diblock copolymer concentrations at or below 1% w/v beyond which the cell lysis gradually increased. No difference in cytotoxicity of MePEG17-b-PCL5 diblock copolymer between MDCKII and MDCKII-MDR1 cell lines was observed at concentrations up to 1%. However, it appeared that MDCKII-MDR1 cell line was more resistant to MePEG17-b-PCL5 at higher concentrations when compared to MDCKII cell line.     66   Figure 2.4 a) Cell viability of (●) MDCKII and ()  MDCKII-MDR1 in the presence of MePEG17-b-PCL5. The cells were incubated with various concentrations of MePEG17-b-PCL5 for three days in culture medium followed by the determination of cell viability by MTS assay. b) LDH release of (●) MDCKII and () MDCKII-MDR1 in the presence of MePEG17-b-PCL5. The cells were incubated with various concentrations of MePEG17-b-PCL5 for 90 minutes in HBSS followed by the LDH release assay. The indicated values are the mean of three independent experiments (±SEM).   0 1 2 3 4 5050100MDCKIIMDCKII-MDR1Concentration of MePEG17-PCL5 (%w/v)% Cell Viability0 1 2 3 4 501020304050MDCKIIMDCKII-MDR1Concentration of MePEG17-b-PCL5 (%w/v)% LDH Releasea)b)  67 2.3.4 Cytotoxicity of PTX loaded micelles and nanospheres PTX was loaded into MePEG114-b-PCL19 micelles and MePEG114-b-PCL104 nanospheres using a previously reported nanoprecipitation and dialysis method (Letchford et al., 2009). It was previously determined that the maximum solubilization of PTX by MePEG114-b-PCL19 micelles was 1.7% (w/w) at an approximate 84% loading efficiency, whereas MePEG114-b-PCL104 nanospheres solubilized PTX at 2.5% (w/w) with an approximate 85% loading efficiency (Letchford and Burt, 2012). The cytotoxicity of PTX loaded nanoparticles was investigated over a PTX concentration range from 2 μg/ml to 32 μg/ml with a maximum copolymer concentration of 0.2 w/v over a 3 day incubation period. The IC50 values of PTX loaded micelles and nanospheres for MDCKII cells was found to be 1.4 and 1.9 μg/ml, respectively (Figure 2.5a). The PTX loaded MePEG114-b-PCL19 micelles showed greater inhibition of proliferation of MDCKII-MDR1 as compared to PTX loaded MePEG114-b-PCL104 nanospheres (approximately 60% and 90% cell viability, respectively at a 32 µg/ml PTX loading) (Figure 2.5b). Due to the inability of these formulations to effectively inhibit the proliferation of MDCKII-MDR1 over the PTX concentration range tested, the IC50 for both nanoparticulate formulations could not be determined. The presence of the P-gp inhibitor, cyclosporin A (CsA), resulted in greater than 90% inhibition of proliferation of MDCKII-MDR1 cell lines.     68      Figure 2.5 Cytotoxicity of PTX loaded () MePEG114-b-PCL19 micelles and (☐) MePEG114-b-PCL104 nanospheres in (a) MDCKII and (b) MDCKII-MDR1 cell lines after 3 days incubation. The cell viability was determined by an MTS assay. The indicated values are the mean of three independent experiments (±SEM).   0 10 20 30050100MePEG114-b-PCL19MePEG114-b-PCL104Concentration of PTX (mg/ml)% Cell Viability0 10 20 30050100 MePEG114-b-PCL19MePEG114-b-PCL104MePEG114-b-PCL19+CsAMePEG114-b-PCL104+CsAConcentration of PTX (mg/ml)% Cell Viabilitya)b)  69 2.3.5 Drug accumulation studies The intracellular PTX accumulation in both MDCKII and MDCKII-MDR1 were investigated for incubations with both micellar and nanosphere PTX (Figure 2.6). Although it appeared that cells treated with PTX loaded nanospheres showed slightly higher PTX accumulation when compared to micelles, there was no statistical difference between micelles and nanospheres in both MDCKII and MDCKII-MDR1 cell lines (P values = 0.34 and 0.20, respectively).  Figure 2.6 The intracellular accumulation of PTX delivered by MePEG114-b-PCL19 micelles (black) and MePEG114-b-PCL104 nanospheres (white) in MDCKII and MDCKII-MDR1 cells.  Cells were incubated with nanoparticulate PTX (32 μg/ml) for 90 min at 37°C followed by washing and lysis using 2% TritonX-100 in HBSS.  Intracellular PTX levels were measured using liquid scintillation counting of cell lysates.  The intracellular PTX content was normalized to the total protein content as determined by BCS assay. The indicated values are the mean of three independent experiments (±SEM).   2.3.6 Polymer uptake and fluorescence confocal microscopy  To quantify the amount of binding and/or cellular uptake of the two different copolymers, TMRCA labeled copolymers were incubated with the two cell lines followed by quantification of the fluorescence using fluorescence spectroscopy. The association of MePEG114-b-PCL19 MDCKII MDCKII-MDR1050010001500MePEG114-b-PCL19MePEG114-b-PCL104PTX (ng/mg protein)  70 micelles with MDCKII and MDCKII-MDR1 cells (0.11 and 0.13 mg polymer/mg protein, respectively) appeared to be higher than values for MePEG114-b-PCL104 nanospheres (0.08 and 0.11 mg polymer/mg protein, respectively), but there was no statistical significant difference (Figure 2.7). To evaluate whether the nanoparticles were internalized by cells or simply bound to the cell membrane of MDCKII and MDCKII-MDR1, confocal fluorescent microscopy was used. The presence of TMRCA-labeled MePEG114-b-PCL19 micelles (Figure 2.8. b and d) and MePEG114-b-PCL104 nanospheres (Figure 2.8. f and h) in the cytoplasm was shown by the red fluorescence of the polymer. The images on the left column of Figure 7 represented the cell contour using direct contrast. The internalization of nanoparticles was confirmed by the z-stack analysis, demonstrating that the red fluorescent polymer is present throughout the cytoplasm rather than being only on the cell membrane. No autofluorescence of the cells was observed and a negative control was performed in which the cells were treated with unlabeled nanoparticles, showing no red fluorescence signal (data not shown).      71    Figure 2.7 Intracellular concentration of TMRCA-labeled copolymers.  Cells were exposed to TMRCA-labeled micelles (white) or nanospheres (black) for 90 min at 37 °C.  After the incubation period, cells were washed and lysed with 2% TritonX-100 in HBSS.  The amount of polymer-bound or uptake into cells was determined by fluorescence spectroscopy. The indicated values are the mean of three independent experiments (±SEM).   MDCKII MDCKII-MDR10.000.050.100.150.20MePEG114-b-PCL19MePEG114-b-PCL104mg diblock polymer/ mg protein  72   Figure 2.8 Confocal fluorescence imaging of MDCKII (a,b,e,f) and MDCKII-MDR1 (c,d,g,h) cells illustrating uptake of TMRCA-labeled MePEG114-b-PCL19 (a-d) and MePEG114-b-PCL104 (e-h) diblock copolymers after 90 min incubation.  The images on the left column are an overlay of a direct contrast signal (which shows the contour of the cells) and the fluorescence of the nucleus (blue).  The images on the right column demonstrate the overlay of fluorescence signal of the nucleus and the TMRCA-labeled diblock copolymers (red).     73 2.3.7 The effect of MePEG17-b-PCL5 on the cytotoxicity of PTX loaded nanoparticles To investigate the ability of the Pgp modulator, MePEG17-b-PCL5 to enhance the cytotoxicity of PTX loaded nanoparticles in MDCKII cell lines, cells were treated with PTX loaded micelles or nanospheres concurrently with MePEG17-b-PCL5 at a concentration of 0.05% w/v (Figure 2.9). As a positive control MePEG17-b-PCL5 was substituted with 5µM CsA. Regardless of the treatment, the proliferation of MDCKII cells was effectively inhibited by PTX loaded nanoparticles with cell viability values in the range of 10-30%. In the absence of MePEG17-b-PCL5 or CsA, MDCKII-MDR1 cell viability was found to be approximately 80% and 60% for treatment with PTX loaded nanospheres and micelles, respectively. As a positive control MDCKII-MDR1 cells were treated with PTX loaded nanospheres or micelles with 5 μM CsA resulting in cell viability values of approximately 19% and 14%, respectively. Concurrent treatment with PTX loaded micelles or nanospheres with 0.05% w/v MePEG17-b-PCL5 resulted in MDCKII-MDR1 viability of approximately 40%.  Cells were then incubated with PTX (32 μg/ml) loaded MePEG114-b-PCL19 micelles or MePEG114-b-PCL104 nanospheres at a copolymer concentration of 0.2 % w/v, for 90 minutes followed by extracellular washing/removal of the nanoparticulate dispersions and subsequent incubation with MePEG17-b-PCL5 diblock copolymer at concentrations up to 0.25% w/v in culture medium for 3 days. At this high concentration of PTX, both micellar and nanosphere formulations strongly inhibited MDCKII cell proliferation in the absence of MePEG17-b-PCL5, with a slight increase in cell viability as the MePEG17-b-PCL5 concentration increased (Figure 2.10a,b). The drug resistant MDCKII-MDR1 cells displayed a higher degree of cell viability in the absence of MePEG17-b-PCL5 as compared to MDCKII. However, with an increase in MePEG17-b-PCL5 concentration, there was a significant decrease in the viability of these cells (Figure 2.10) with minimum cell viability occurring at MePEG17-b-PCL5 diblock copolymer   74 concentrations above 0.05% w/v for both the micelles and nanospheres. This low cell viability of MDCKII-MDR1 cells was similar to that of MDCKII at the same MePEG17-b-PCL5 concentration. Low concentrations of MePEG17-b-PCL5 resulted in a higher degree of cell viability for MePEG114-b-PCL104 nanospheres as compared to cells incubated in the presence MePEG114-b-PCL19 micelles until MePEG17-b-PCL5 concentrations above 0.05% were reached at which point cell viability remained at approximately 20%.  To determine the IC50 of PTX loaded nanoparticles in the presence of MePEG17-b-PCL5 at the concentration of maximal inhibition of cell proliferation found in the previous experiment (0.05% w/v), both MDCKII and MDCKII-MDR1 were pre-treated with PTX-loaded micelles or nanospheres at various drug concentrations for 90 minutes followed by removal of the drug-loaded formulations and subsequent incubation with MePEG17-b-PCL5 at 0.05% w/v in the presence of culture medium for 3 days. The presence or absence of MePEG17-b-PCL5 did not significantly affect the IC50 for MDCKII treated with PTX loaded MePEG114-b-PCL19 micelles (Figure 2.11a). Interestingly, incubation of MDCKII-MDR1 cells with PTX loaded MePEG114-b-PCL19 micelles followed by subsequent incubation with MePEG17-b-PCL5 resulted in the significant decrease in IC50 of this formulation to a level that was comparable to that obtained with treatment of MDCKII with the same formulation (Figure 2.11a). Treatment of MDCKII with PTX loaded MePEG114-b-PCL104 nanospheres with subsequent incubation with MePEG17-b-PCL5 resulted in a slightly higher IC50 than treatment without MePEG17-b-PCL5 (Figure 2.11b). Incubation of MDCKII-MDR1 cells with PTX loaded MePEG114-b-PCL104 nanospheres did not result in any decrease in cell viability; however, subsequent incubation with MePEG17-b-PCL5 resulted in significant inhibition of cell proliferation (Figure 2.11b). The IC50 of this group was not as low as the IC50 for the group first incubated with MePEG114-b-PCL19 micelles followed by incubation with MePEG17-b-PCL5.    75    Figure 2.9 Cell viability of MDCKII (white) or MDCKII-MDR1 (black) cells treated for 3 days with MePEG-b114-PCL19 micelles or MePEG-b114-PCL104 nanospheres loaded with 16 µg/ml PTX with or without concurrent treatment with (0.05% w/v) MePEG-b17-PCL5 or (5 μM) CsA. The cell viability was determined by an MTS assay. The indicated values are the mean of three independent experiments (±SEM). * and *** indicate statistical significance (p<0.05).   MePEG114-b-PCL104MePEG114-b-PCL19MePEG114-b-PCL104+MePEG17-b-PCL5MePEG114-b-PCL19+MePEG17-b-PCL5MePEG114-b-PCL104+CsAMePEG114-b-PCL19+CsA020406080100MDCKIIMDCKII-MDR1********% Cell Viability  76   Figure 2.10 The effect of MePEG17-b-PCL5 diblock copolymer on the cytotoxicity of PTX loaded- (a) MePEG114-b-PCL19 micelles and (b) MePEG114-b-PCL104 nanospheres. (●) MDCKII and () MDCKII-MDR1 were incubated with PTX loaded nanoparticles at a drug concentration of 32 μg/ml and copolymer concentration of 0.2% w/v for 90 min followed by removal of the drug loaded nanoparticle and a subsequent 3 day incubation in various concentrations of MePEG17-b-PCL5.  The cell viability was determined by an MTS assay. The indicated values are the mean of three independent experiments (±SEM) (c) A schematic illustration of experimental sequences.   0.00 0.05 0.10 0.15 0.20 0.25020406080100MDCKIIMDCKII-MDR1Concentration of MePEG17-b-PCL5 (%w/v)% Cell Viability0.00 0.05 0.10 0.15 0.20 0.25020406080100MDCKIIMDCKII-MDR1Concentration of MePEG17-b-PCL5 (%w/v)% Cell Viabilityb)a)  77   Figure 2.11 The cytotoxicity of PTX- loaded (a) MePEG114-b-PCL19 micelles and (b) MePEG114-b-PCL104 nanospheres in MDCKII and MDCKII-MDR1 cells with and without subsequent incubation with MePEG17-b-PCL5. Cells were treated with PTX-loaded micelles and nanospheres with varying concentrations of PTX for 90 min followed by removal of the drug loaded nanoparticles and a subsequent 3 day incubation with media with or without 0.05% w/v MePEG17-b-PCL5. Cell viability was determined by an MTS assay. The indicated values are the mean of three independent experiments (±SEM). (c) A schematic illustration of experimental sequences.    0204060050100MDCKIIMDCKII-MDR1MDCKII+PCL5MDCKII-MDR1+PCL5Concentration of PTX (mg/ml)% Cell Viability0 20 40 60050100MDCKIIMDCKII-MDR1MDCKII+PCL5MDCKII-MDR1+PCL5Concentration of PTX (mg/ml)% Cell Viabilitya)b)  78 2.4 Discussion Our previous reports have focused on the physicochemical characterization of MePEG114-b-PCL19 and MePEG114-b-PCL104 diblock copolymers, and their performance as drug delivery vehicles for PTX (Letchford and Burt, 2012; Letchford et al., 2009). We demonstrated that MePEG114-b-PCL104 nanospheres solubilized a greater amount of PTX, released the drug at a slower rate, and exhibited enhanced stability and retention of PTX in human plasma (Letchford et al., 2009). The current work expands upon the previous reports and investigates the differences in these PTX loaded nanoparticulate carriers with respect to their ability to inhibit cell proliferation of drug sensitive and resistant Pgp overexpressing cell lines MDCKII and MDCKII-MDR1, respectively. Additionally, we investigated whether concurrent and subsequent administration of MePEG17-b-PCL5 with these drug-loaded nanoparticles resulted in enhanced inhibition of cellular proliferation. For these studies we evaluated our formulations using the MDCKII and MDCKII-MDR1 cell lines as this pair is a good model for the study of Pgp substrates/inhibitors (Tang et al., 2002; Taub et al., 2005).  2.4.1 Cellular biocompatibility of MePEG-b-PCL The determination of the biocompatibility of the three MePEG-b-PCL diblock copolymers (MePEG114-b-PCL104, MePEG114-b-PCL19, and MePEG17-b-PCL5) in MDCKII and MDCKII-MDR1 cells was required to define the concentration of polymer tolerated by the cells in order to evaluate the cytotoxicity of drug loaded formulations in subsequent experiments. Our results demonstrated that MePEG-b-PCL diblock copolymers are well tolerated by the MDCKII and MDCKII-MDR1 cell lines with little or no cytotoxicity of either cell line for concentrations of MePEG114-b-PCL104 or MePEG114-b-PCL19 up to 0.4% w/v (Figure 2.2). This high degree of biocompatibility of MePEG-b-PCL diblock copolymers was also reported by other groups in   79 several different cell lines including erythrocytes, breast cancer, pheochromocytoma, and cervical cancer cells (Allen et al., 1998; Letchford et al., 2009; Zhang et al., 2010a). The short-term cytotoxicity study also suggested that the MePEG114-b-PCL19 and MePEG114-b-PCL104 copolymers did not cause a high degree of cell lysis as indicated by low LDH release (Figure 2.3). Similar to MePEG114-b-PCL19 and MePEG114-b-PCL104 copolymers, both short and long term cell viability assays demonstrated that the MePEG17-b-PCL5 copolymer did not cause cell death at concentrations up to 1% (w/v), suggesting that this copolymer is also highly biocompatible with MDCKII and MDCKII-MDR1 cell lines (Figure 2.4a and b). In previous studies using NCI/ADR-RES cells we demonstrated that cell death occurred beyond a MePEG17-b-PCL5 concentration of 0.1% w/v (Elamanchili et al., 2009) whereas in Caco-2 cells, lysis was not observed until concentrations exceeded 0.5% (Zastre et al., 2002). These previous results, in combination with our current results, indicate that the cellular biocompatibility of these copolymers may vary dependent on the particular cell line used.  2.4.2 Cytotoxicity of PTX loaded nanoparticles PTX-loaded nanoparticles effectively inhibited the proliferation of MDCKII cells (Figure 2.5a); but MDCKII-MDR1 cells demonstrated significant resistance to the drug as exhibited by the high degree of cell viability, regardless of the concentration of PTX used (Figure 2.5b). However, treatment of MDCKII-MDR1 cells with PTX loaded nanoparticles in the presence of the recognized Pgp inhibitor CsA (Szakacs et al., 2006) resulted in a rapid decline in cell viability with increased PTX loadings, establishing that the high degree of resistance to PTX treatment was due to the up-regulation of Pgp in this cell line. Figure 2.5b shows that PTX loaded MePEG114-b-PCL19 micelles were more effective than PTX loaded MePEG114-b-PCL104 nanosphere at inhibiting the proliferation of MDCKII-MDR1 cells. This was unlikely due to any   80 differences in the total amount of PTX delivered to the cells as there was no statistical difference between drug accumulation levels in cells incubated with either nanosphere or micelle formulations (Figure 2.6).  In the present study, the fact that the nanospheres were shown to have a higher IC50, yet delivered approximately the same amount of total drug as the micelles, leads us to speculate that a fraction of the drug is still associated with intracellular MePEG114-b-PCL104 nanospheres and therefore not available to bind to β-tubulin, the intracellular target of PTX. In order for this concept to be feasible, cellular uptake of the copolymer is a prerequisite. Cellular uptake of the copolymers was quantified using a spectrofluorometric assay (Figure 2.7) and observed visually by confocal microscopy (Figure 2.8). Both MePEG114-b-PCL19 and MePEG114-b-PCL104 copolymers were associated with both MDCKII and MDCKII-MDR1 cells (Figures 2.7 and 2.8) providing some evidence for intact nanoparticulate uptake. It is not known why the MDR cells took up more copolymer than the MDCKII cells. However, while MDCKII and MDCKII-MDR1 are very similar in nature there are likely large differences in protein content between the cells and the copolymer uptake in Figure 2.7 is expressed as a function of protein content. For both the nanospheres and micelles, MDCKII and MDCKII-MDR1 cells clearly displayed red cytoplasmic staining due to the TMRCA labeled copolymer, indicating association of the copolymers with the cells (Figure 2.8). Z-stacking verified that that this staining was located in the cytosol and not located in the plasma membrane or in the nuclear compartment of the cells.  Consistent with our findings, Savic et al. also demonstrated that PEO45-b-PCL23 labeled with TMRCA was localized in the cytoplasm (Savic et al., 2003). Additional studies using a fluorogenic dye conjugated to PEO45-b-PCL21, demonstrated that micelles formed from these copolymers, for the most part, retained their integrity upon uptake by T24 bladder carcinoma cells (Savic et al., 2006). Several groups have also demonstrated that block copolymers are   81 internalized by cells, primarily via endocytotic mechanisms (Luo et al., 2002; Sahay et al., 2008). On the other hand, in a series of FRET studies using KB cells and MePEG-b-PLA and MePEG-b-PCL micelles, Chen et al. provided evidence that hydrophobic fluorescent probes solubilized in these micelles rapidly partitioned into the plasma membrane of KB cells followed by uptake of FRET by endocytosis (Chen et al., 2008a). Similar findings were reported for the mechanism of uptake of the hydrophobic dye Nile Red by Xiao et al. but with some evidence of copolymer association with the membrane (Xiao et al., 2011). It must be kept in mind that different polymers, cell lines and surface charges of nanoparticles may result in varying degrees and mechanisms of uptake (Sahay et al., 2010a).  2.4.3 The effect of MePEG17-b-PCL5 on the cytotoxicity of PTX loaded nanoparticles We investigated the ability of PTX loaded MePEG114-b-PCL19 micelles or MePEG114-b-PCL104 nanospheres to inhibit cell proliferation of MDCKII cell lines when concurrently administered with the Pgp modulators MePEG17-b-PCL5 or CsA, for 3 days (Figure 2.9). Treatment of MDCKII cells resulted in low cell viability regardless of the nanoparticulate carrier used or whether a Pgp modulator was present, as expected. MDCKII-MDR1 cells treated with, either PTX-loaded micelles or nanospheres in the absence of MePEG17-b-PCL5 or CsA resulted in considerably higher cell viability values as compared to the MDCKII group. The presence of CsA or MePEG17-b-PCL5 decreased the cell viability of MDCKII-MDR1 to about the same extent, confirming the usefulness of MePEG17-b-PCL5 as a Pgp inhibitor when used in combination with micelles and nanospheres. The short-term (90 min) incubation of PTX loaded micelles and nanospheres, with a 32µg/ml drug loading, followed by washing, produced a rapid and high intracellular loading of PTX. Subsequent incubation in drug free culture media for 3 days without MePEG17-b-PCL5,   82 effectively inhibited the proliferation of MDCKII cells (Figure 2.10). The slight increase in the MDCKII cell viability when the copolymer concentration was above 0.05% w/v was not found to be significant and the cytotoxicity of PTX was determined to be more than 80% over the full MePEG17-b-PCL5 concentration range. In the absence of MePEG17-b-PCL5, MDCKII-MDR1 cell viability remained relatively high, and treatment with micelles was slightly more effective at inhibiting cell proliferation than nanospheres (Figure 2.10). Subsequent incubation of MDCKII-MDR1 cells with increasing concentrations of MePEG17-b-PCL5 resulted in a copolymer dose dependent decrease in cell viability with the most effective MePEG17-b-PCL5 concentrations above 0.05% w/v (Figure 2.10). The key finding presented in the current study is that PTX loaded MePEG-b-PCL nanoparticles are capable of delivering enough PTX during a short 90 min incubation period to effectively inhibit cell proliferation of MDCKII-MDR1 cells provided that MePEG17-b-PCL5 is present continuously throughout the subsequent 3 day incubation period at a concentration above 0.05% w/v. Although the mechanism by which PCL inhibits Pgp is not entirely clear, previous studies indicate that MePEG17-b-PCL5 can penetrate cell membranes and increase Pgp ATPase activity (Zastre et al., 2007). It was speculated that the stimulation of the ATPase might reflect an ability of the copolymer to interact directly with Pgp leading to inhibition. It has been proposed that non-ionic surfactants inhibit/modulate Pgp by various mechanisms including, changing membrane fluidity, inhibiting Pgp ATPase activity, and depleting ATP stores (Dudeja et al., 1995; Regev et al., 1999; Kabanov et al., 2003; Collnot et al., 2006; Collnot et al., 2010). Interestingly, as we have previously demonstrated, if MePEG17-b-PCL5 is not maintained above an inhibitory concentration, PTX rapidly effluxes from the MDCKII-MDR1 cells, resulting in reduced inhibition of cell proliferation (Elamanchili et al., 2009). An additional experiment was conducted to determine the IC50 of PTX loaded   83 nanospheres or micelles in the presence of MePEG17-b-PCL5 at its most effective concentration (0.05%). The results from Figure 2.11 show that the subsequent incubation with MePEG17-b-PCL5 copolymer did not affect the IC50 in MDCKII cells treated with either PTX-loaded micelles or nanospheres. Treatment of MDCKII-MDR1 cells with PTX-loaded micelles or nanospheres along with subsequent incubation with 0.05% MePEG17-b-PCL5 resulted in a significant reduction in the cell viability of these cells. PTX loaded micelles resulted in a lower IC50 than drug loaded nanospheres which we speculate is due to a higher fraction of PTX bound to an intracellular pool of MePEG114-b-PCL104 nanospheres and less free intracellular PTX available to bind to β-tubulin in the nanospheres formulation. The use of mixed nanoparticle systems that carry both a cytotoxic drug and a polymeric Pgp inhibitor to overcome MDR has been previously reported using a variety of Pluronic block copolymers (Mei et al., 2009; Zhang et al., 2010c; Zhang et al., 2011; Mu et al., 2010). However, in recent studies by Zhang et al. (Zhang et al., 2011), blank Pluronic P123/F127 mixed polymeric micelles were demonstrated to be toxic at relatively low concentrations (0.1% w/v) compared to the copolymers investigated in this study which, do not induce cytotoxicity below 1% w/v. We suggest that the greater cellular biocompatibility along with the biodegradability of MePEG-b-PCL may be a potential advantage allowing for higher dosing of drug and copolymer.  2.5 Conclusions In this study, we demonstrated that the cytotoxicity of PTX loaded MePEG-b-PCL nanoparticles is dependent on the hydrophobic block length of the carrier, with shorter hydrophobic block length copolymer (MePEG114-b-PCL19) micelles demonstrating greater cytotoxicity in MDR cells. Importantly, MePEG17-b-PCL5 co-administered with PTX loaded nanoparticles dramatically enhanced the cytotoxicity of PTX in drug resistant cells. Further   84 studies are in progress to formulate mixed molecular weight copolymer nanoparticles, in which MePEG-b-PCL serves as the carrier and both MePEG17-b-PCL5 and PTX are co-loaded in the nanoparticles to achieve intracellular sustained release of both components with the goal of overcoming drug resistance in cancer cells.   2.6 Acknowledgments These studies were financially supported by an operating grant from the Canadian Institutes of Health Research (CIHR) awarded to Dr Helen Burt. The authors would like to thank Dr. Padmesh Rajput, Lindsay Heller and The Center for Drug Research and Development (CDRD) for the use of their laser confocal microscope. We would also like to thank Karanvir Sall, Nathan Wong and Winnie Ye for their assistance in sample preparation.      85 Chapter  3: Mixed molecular weight copolymer nanoparticles for the treatment of drug resistant tumors: formulation development, cytotoxicity and pharmacokinetics4 3.1 Introduction The taxanes, PTX and DTX are widely used antineoplastics approved for use in the treatment of a variety of cancers including breast, ovarian and non-small cell lung cancer. Due to their low aqueous solubility, the commercial formulations of PTX (Taxol®) and DTX (Taxotere®) contain large amounts of the surfactants CrEL and Tween 80, respectively. Life threatening hypersensitivity reactions, leaching of plasticizers from infusion sets, vesicular degeneration and the alteration of the pharmacokinetics and disposition of PTX have been associated with the use of these surfactants (van Zuylen et al., 2001). Furthermore, these drugs are substrates for Pgp, a transmembrane efflux protein that is overexpressed in multi-drug resistant cancer cells and hence provides a significant barrier to successful cancer treatment (Gottesman et al., 2002). Amphiphilic block copolymers composed of a hydrophilic block of methoxy poly (ethylene glycol) (MePEG) covalently bound to a hydrophobic, biodegradable polyester block such as poly(D,L-lactic acid) (Zhang et al., 1997), poly(caprolactone) (Aliabadi et al., 2005; Letchford et al., 2008) or poly(lactic-co-glycolic acid) (Hu and Zhang, 2010; Yoo and Park, 2001) (abbreviated MePEG-b-PDLLA, MePEG-b-PCL and MePEG-b-PLGA, respectively) have been extensively explored for the delivery of taxanes and other hydrophobic drugs, providing many advantages in the formulation of these compounds. These materials form core-shell                                                  4 The data presented in this chapter are in preparation for submission for publication. Wan C.P. and Letchford K.J. et al. (2014). Mixed molecular weight copolymer nanoparticles for the treatment of drug resistant tumors: formulation development, cytotoxicity and pharmacokinetics.    86 nanoparticles that are capable of dramatically enhancing the aqueous solubility of taxanes and may increase the maximum tolerated dose through the use of less toxic pharmaceutical excipients, thereby improving the safety profile and efficacy compared to the commercial formulations (Kim et al., 2004). The highly water bound PEG corona is thought to minimize opsonization, delay capture by the RES and increase the circulation time of the nanoparticles, promoting accumulation in solid tumors by the enhanced permeation and retention effect. Furthermore, our group has demonstrated that low molecular weight MePEG-b-PCL referred to as MePEG17-b-PCL5 is an effective modulator of Pgp resulting in the enhanced uptake of Pgp substrates including PTX (Elamanchili et al., 2009; Zastre et al., 2002; Zastre et al., 2004; Zastre et al., 2007; Zastre et al., 2008). Most studies investigating the delivery of taxanes by block copolymer nanoparticles have focused on the strategy of drug solubilization for systemic administration with prolonged circulation times for improved tumour uptake after intravenous administration (Gaucher et al., 2010). However, we have recently demonstrated that taxane loaded nanoparticles offer another potentially important delivery strategy, based on our finding that intracellular accumulation of drug and cytotoxicity were enhanced in both drug sensitive and drug resistant (Pgp over-expressing) cells following the application of ultrasound to the cells (Wan et al., 2012). We suggest that if stable, taxane loaded nanoparticles were taken up into cells to form a controlled release intracellular depot of drug, then co-encapsulation of the Pgp inhibitor, MePEG17-b-PCL5, within the nanoparticle may also inhibit drug efflux from drug resistant cells and enhance cytotoxicity. Using a similar strategy, doxorubicin has been co-formulated in a variety of nanoparticles with a number of Pgp inhibitors including verapamil, cyclosporine, siRNA and G918 and thus allowing for greater doxorubicin accumulation in drug resistant cells (Jabr-Milane et al., 2008; Chen et al., 2010; Vauthier et al., 2003). Similarly, PTX encapsulated in lipid-based   87 nanoparticles or formulated as nanocrystals, both including surfactant-type Pgp inhibitors, have been shown to reduce the mass of drug resistant tumors in vivo by a mechanism proposed to include endocytosis of the nanoparticles and likely, the intracellular release of the surfactant after nanoparticulate uptake (Dong et al., 2009; Liu et al., 2010). In recent years, a number of taxane loaded Pluronic® containing mixed nanoparticle systems have been investigated for the treatment of drug resistant cancers (Mei et al., 2009; Mu et al., 2010; Zhang et al., 2011). Due to a well-documented ability to inhibit Pgp, Pluronic® agents have been demonstrated to significantly decrease the IC50 of the encapsulated drug, as compared to free drug or the commercial formulation. However, in some cases, the carrier itself was shown to be cytotoxic at relatively low copolymer concentrations and in most reports, the viability of the drug resistant cells remained relatively high despite treatment with high concentrations of drug (Li et al., 2010; Shah et al., 2009; Zhang et al., 2011; Zhang et al., 2010b). In this work, we developed a mixed molecular weight nanoparticulate system referred to as “mixed MW PCL200/PCL5” nanoparticles that are composed entirely of biodegradable and biocompatible MePEG-b-PCL. A high molecular weight diblock copolymer, MePEG114-b-PCL200, was used to maximize taxane loading and stability, and a low molecular weight MePEG17-b-PCL5, was used as a Pgp inhibitor. The in vitro physicochemical characteristics, cytotoxicity and in vivo stability of these nanoparticles loaded with PTX and DTX were determined. We demonstrated cytotoxic effects of these nanoparticles on a MDR cell line that were markedly superior to previously reported Pluronic® mixed micelles using similar or lower drug concentrations.      88 3.2 Materials and methods 3.2.1 Materials -caprolactone was purchased from Alfa Aesar (Ward Hill, MA, USA). Stannous octoate and methoxy poly(ethylene glycol) (MePEG) were obtained from Sigma Aldrich Canada Ltd. (Oakville, ON, Canada), paclitaxel from Polymed Therapeutics Inc. (Houston, TX, USA), docetaxel from Natural Pharma (Vancouver, BC, Canada), Paclitaxel for Injection (Equivalent to Taxol®) from Biolyse Pharma Corp. (St. Catherines, ON, Canada), Taxotere® from Sanofi Aventis (Laval, PQ, Canada), 3H and 14C paclitaxel and docetaxel from Moravek Biochemicals Inc. (Brea, CA, USA) and 3H methyl iodide from American Radiolabeled Chemicals, Inc. (St. Louis, MO, USA), Dulbecco’s modified eagle’s media (DMEM) supplemented with 5% FBS and 1% penicillin/streptomycin, and Hank’s buffered salt solution (HBSS) from Invitrogen (Grand Island, NY, USA). The solvents chloroform, ethyl acetate, anhydrous toluene, hexane, diethyl ether, acetonitrile, methanol, N, N-dimethylformamide were all purchased from Fisher Scientific Co. (Ottawa, ON, Canada). Deuterated chloroform was obtained from Cambridge Isotope Laboratories (Andover, MA, USA). The MTS cell proliferation assay kit and BCA protein assay kit were purchased from Promega (Madison, WI, USA) and Thermo Scientific (Rockford, IL, USA), respectively. MDCKII and MDCKII-MDR1 cells were kind gifts from Dr. Piet Borst (National Cancer Institute, the Netherlands).  3.2.2 Synthesis and characterization of copolymers A high molecular weight MePEG-b-PCL copolymer, termed MePEG114-b-PCL200, was synthesized by reacting methoxy poly(ethylene glycol) (MePEG) (MW 5000 g/mol) with ε-caprolactone in a weight ratio of 18:82. Synthesis and characterization of MePEG-b-PCL diblock copolymers have been described in Section 2.2.2 of this thesis. The high molecular weight   89 MePEG-b-PCL was radiolabeled by reacting the terminal hydroxyl group with 3H methyl iodide (American Radiolabeled Chemicals, Inc. St Louis, MO, USA). The copolymer (1g) was dissolved in 30 ml of anhydrous dioxane followed by addition of 150 mg of potassium hydride and allowed to react for 30 minutes with stirring. Subsequently, 2.5 mCi of 3H methyl iodide in DMF was added and the mixture stirred overnight. The reaction mixture was transferred to a 3500 MWCO dialysis tube (Thermo Scientific, Rockford, IL, USA) and dialysed against distilled water for 4 days with continuous water changes until the dialysate was no longer radioactive. The water was then exchanged for methanol through dialysis with four solvent changes and the solvent evaporated in a fume hood and then dried under vacuum.  3.2.3 Preparation and characterization of nanoparticles Nanoparticles were prepared by an oil-in-water emulsion technique or a nanoprecipitation and dialysis method. In the emulsification method, MePEG114-b-PCL200 was dissolved in 500 l of ethyl acetate at a concentration of 100 mg/ml with a specified amount of MePEG17-b-PCL5 (ranging from 0 to 100 mg/ml) to stabilize the emulsion. The oil phase was emulsified in 5 mL of aqueous buffer (10mM phosphate buffered saline (PBS) for drug release and pharmacokinetic studies or HBSS for cell studies) using a 100W tip sonicator (Sonic Dismembrator, Fisher Scientific, Ottawa, ON, Canada) for 1 minute in an ice bath. The nanoparticulate dispersions were stirred overnight to allow the ethyl acetate to evaporate. The dispersions were diluted to a final volume of 5 ml with the appropriate buffer and then centrifuged at 9500 xg for 5 minutes to remove any precipitate prior to further studies. The final concentration of MePEG114-b-PCL200 in the dispersion was 1% w/v with the MePEG17-b-PCL5 concentration ranging from 0 to 1% w/v. The hydrodynamic diameter and zeta potential of nanoparticles was determined using a Malvern Zetasizer Nano ZS (Malvern, UK). Nanoparticles were also prepared in the absence of   90 MePEG17-b-PCL5 using a nanoprecipitation and dialysis technique and are referred to as PCL200 nanoparticles. In this method, 30 mg of MePEG114-b-PCL200 was dissolved in 1 ml of DMF and this was added drop-wise to 1 ml of rapidly stirring PBS or HBSS (for cell studies). This dispersion was then transferred to a 3500 MWCO dialysis tube and dialysed against 4L of buffer overnight. The final nanoparticle dispersion was made up to 3 ml with the appropriate buffer.  3.2.4 Drug loading To determine the maximum loading and loading efficiency of PTX or DTX in the nanoparticles, several nanoparticle dispersions were prepared, as outlined above, with increasing amounts of drug in the ethyl acetate or DMF copolymer solution. After centrifugation, an aliquot of the nanoparticle dispersions was collected and dried under a stream of nitrogen gas and reconstituted in acetonitrile. This solution was analyzed for drug content by HPLC. The loading efficiency of the nanoparticles was calculated as:         (3.1)  Where [Drug]solubilized is the concentration of PTX or DTX found in solution after encapsulation in nanoparticles as determined by HPLC and [Drug]added is the concentration of PTX or DTX added to the solution during the formation of nanoparticles.  3.2.5 In vitro drug and MePEG17-b-PCL5 release Nanoparticles were prepared as described above with a PTX or DTX concentration of 100 g/ml and a PCL200 or mixed MW PCL200/PCL5 concentration of 1% w/v. A trace amount    Loading Efficiency = [Drug]so lub ilized[Drug]added ´100  91 of 3H PTX or 3H DTX was added to the drug/copolymer solution prior to emulsification or nanoprecipitation to enable quantification of released drug. A 40 l aliquot of drug loaded nanoparticles was pipetted into 7000 MWCO Slide-A-Lyzer® mini dialysis units (Thermo Scientific, Rockford, IL, USA), and the samples were dialysed against 6 L of 10 mM PBS at pH 7.4 and 37C with constant stirring. Release into this large volume ensured that sink conditions were maintained throughout the experiment. At predetermined time points, the contents of 4 dialysis units were removed with 3 washings with distilled water and the drug content analyzed by -scintillation counting. The cumulative percent drug released was calculated by subtracting the amount of drug remaining from the initial amount of drug in the dialysis unit at the beginning of the experiment. The data were expressed as cumulative percentage of drug released as a function of time. The release of MePEG17-b-PCL5 from mixed MW PCL200/PCL5 nanoparticles was determined by monitoring the loss of MePEG17-b-PCL5 from a 100k MWCO dialysis membrane (Spectrum Labs, Rancho Dominguez, CA, USA). A 1% w/v dispersion of mixed MW PCL200/PCL5 nanoparticles was added to dialysis bags followed by incubation in 4 L of distilled water at 37C with constant stirring. At predetermined intervals the contents of three of the bags were removed with washings with distilled water and freeze dried. The lyophilized powder was dissolved in chloroform followed by analysis by GPC as outlined above. The ratio of the peak area attributed to MePEG17-b-PCL5 to that from the MePEG114-b-PCL200 was monitored. As a control, the release of MePEG17-b-PCL5 from a micellar dispersion of 1% w/v MePEG17-b-PCL5 was monitored over time as described above. In this case, the decrease in peak area of MePEG17-b-PCL5 over time was determined by GPC. 3.2.6 Cytotoxicity of MePEG-b-PCL nanoparticulate formulations For the cytotoxicity studies, MDCKII and MDCKII-MDR1 cells were seeded at a density of 1500 cells/well in 96-well plates and allowed to grow overnight (5% CO2/95% humidity at   92 37°C). The cytotoxicity of the nanoparticles in the absence of drug was first investigated. Nanoparticles were prepared in culture medium at varying concentrations and incubated with the cells for 3 days. Cytotoxicity studies for both types of nanoparticles were performed over a MePEG114-b-PCL200 concentration range of 0-0.5% w/v. It should be noted that copolymer concentrations investigated for mixed MW PCL200/PCL5 nanoparticles also ranged from 0-0.5% w/v MePEG114-b-PCL200; however, these nanoparticles were prepared with a 1:1 weight ratio of MePEG114-b-PCL200 to MePEG17-b-PCL5. As a control, cells were grown in culture medium in the absence of copolymer. Cell proliferation was measured using the MTS cell proliferation assay kit according to the manufacturer’s instructions. Cell viability was expressed as a percentage of the ratio of absorbance after treatment as compared to the absorbance of the control.  Cell viability in the presence of free drug or drug encapsulated in nanoparticles was investigated. PTX or DTX loaded PCL200 nanoparticles were prepared with a MePEG114-b-PCL200 concentration of 1% w/v or mixed MW PCL200/PCL5 nanoparticles with 1% w/v of MePEG114-b-PCL200 and a 1:1 weight ratio of MePEG114-b-PCL200 to MePEG17-b-PCL5 with a drug loading of 400 μg/ml. The nanoparticles were subsequently diluted in HBSS to produce nanoparticle dispersions with drug concentrations ranging from 0.4 to 100 μg/ml with the same drug to copolymer ratio. Due to the high degree of resistance of the MDCKII-MDR1 cells to PTX, these cells were treated with PTX at higher concentrations (1.56 to 400 μg/ml). Cells were also incubated in the presence of free drug ranging in concentration from 0.03-10 μg/ml. Cells were incubated for 90 minutes with various concentrations of free drug or PTX or DTX loaded nanoparticles followed by removal of the drug solutions or nanoparticle dispersions. The cells were immediately washed with HBSS and incubated in culture media for 3 days in a biological incubator. The cell viability was measured using the MTS cell proliferation assay kit.   93 Cell viability of MDCKII-MDR1 in the presence of a continuous 3-day incubation with free drug or drug-loaded nanoparticles with varying concentrations of copolymer was investigated. Cells were incubated for 3 days in culture media containing free PTX or DTX ranging from 0.03 to 2.5 μg/ml followed by assessment of cell viability by the MTS cell proliferation assay. In a subsequent experiment to investigate the effect of copolymer concentration on the viability of MDCKII-MDR1 cells over a 3-day incubation period, cells were incubated with a range of concentrations of either PCL200 or mixed MW PCL200/PCL5 nanoparticles with a fixed drug concentration of 500 ng/ml of PTX or 50 ng/ml of DTX. In this experiment the copolymer concentration ranged from 0.25% w/v to 0.00195% w/v of MePEG114-b-PCL200 with a 1:1 weight ratio of MePEG17-b-PCL5 for the mixed MW PCL200/PCL5 nanoparticles. Assessment of cell viability was by the MTS cell proliferation assay. An investigation of the ability of blank nanoparticles to inhibit cell proliferation after an initial incubation of drug-loaded mixed MW PCL200/PCL5 nanoparticles was conducted. Cells were first incubated for 90 minutes with mixed MW PCL200/PCL5 nanoparticles with a relatively low concentration of drug loaded mixed MW PCL200/PCL5 nanoparticles (either 6.25 μg/ml of DTX or 25 μg/ml of PTX). Immediately after this incubation, the nanoparticle dispersions were removed, the cells were washed with HBSS and subsequently incubated for 3 days with either PCL200 nanoparticles or mixed MW PCL200/PCL5 nanoparticles, both with a MePEG114-b-PCL200 concentration range of 0-0.25% w/v prepared in culture media. The cell viability was determined using the MTS assay.  3.2.7 Pharmacokinetics and biodistribution of nanoparticulate formulations Animal studies were carried out in accordance with the Canadian Council on Animal Care and using protocols approved by the Animal Care Committee at the University of British   94 Columbia. Nanoparticles were prepared using 1% w/v MePEG114-b-PCL200 with a 1:1 weight ratio of MePEG114-b-PCL200 to MePEG17-b-PCL5 with 1% of 3H labeled MePEG114-b-PCL200 and loaded with 100 g/ml of PTX or DTX and 1% of either 14C PTX or 14C DTX. Paclitaxel for Injection or Taxotere® were prepared at drug concentrations of 100 g/ml of PTX and DTX, respectively. An aliquot of 3H PTX or 3H DTX was added to the commercial formulations during their preparation. Doses of 100 mg/kg copolymer and 1 mg/kg of drug (approximately 200 µl) were administered slowly to mature CD-1 mice at a rate of 30 µl/second (average weight 30g) via tail vein injection using a 25 G needle gauge. Mice were terminated by CO2 inhalation at 5, 15, 30, 90 min, 3, 6 10 and 24 h and blood was collected by cardiac puncture with immediate separation of the plasma by centrifugation at 2400 xg for 15 minutes at 5C. 4 mice were sacrificed at each time point. The major organs (liver, spleen, kidneys, lungs and heart) were harvested from one group of animals 24 h post injection, rinsed with ice-cold saline, blotted dry and weighed. All organs and plasma samples were digested overnight at 60 °C in 1 ml of Solvable (Perkin Elmer, Woodbridge, ON, Canada) followed by addition of 50 l of 200 mM EDTA and 200 l of 30% hydrogen peroxide to de-color the samples. Drug and copolymer levels in plasma and organs were analyzed by β scintillation counting and the data were fit to a non-compartmental model using WinNonlin® 6.2 (Phoenix™) (Pharsight, Sunnyvale, CA, USA).  3.2.8 Statistical analysis The in vitro and in vivo experiments were performed in three individual experiments and four mice, respectively. The data are presented as mean ± standard deviation or standard error of the mean. The data were analyzed by Student’s t-test using Prism 5.0 software (GraphPad Software Inc.) with p < 0.05 considered to be statistically significant.    95 3.3 Results 3.3.1 Synthesis and characterization of copolymers The NMR chemical shifts (spectra not shown) and peak assignments for the synthesized copolymers were as follows: 4.07 ppm (2H CO-CH2-CH2-CH2-CH2-CH2), 3.65 ppm (4H O-CH2-CH2-O), 3.40 ppm (3H CH3-O), 2.3 ppm (2H CO-CH2-CH2-CH2-CH2-CH2), 1.66 ppm (4H CO-CH2-CH2-CH2-CH2-CH2), 1.4 ppm (2H CO-CH2-CH2-CH2-CH2-CH2). Using the sum of the integrated peak areas of the peaks at 1.66 ppm and 1.4 ppm from the caprolactone block and the peak area of the peak at 3.65 ppm from the MePEG, the degree of polymerization (DPn) for the two copolymers were calculated as 5 and 200 caprolactone repeat units and the copolymers were subsequently termed MePEG17-b-PCL5 and MePEG114-b-PCL200, respectively. Using these DPn values, the calculated molecular weights were 1320 g/mol and 27800 g/mol, respectively. The Mn and Mw were determined by GPC and were found to be 1434 g/mol and 1539 g/mol, respectively for MePEG17-b-PCL5 and 33867 g/mol and 40477 g/mol, respectively for MePEG114-b-PCL200. The molecular weight polydispersity indices were calculated as 1.07 and 1.20 for MePEG17-b-PCL5 and MePEG114-b-PCL200, respectively (Table 3.1).  3.3.2 Formation and characterization of nanoparticles Mixed molecular weight nanoparticles were manufactured using an oil-in-water emulsification technique. The addition of MePEG17-b-PCL5 to the oil phase served to stabilize the emulsion as well as act as a Pgp inhibitor. With an increased MePEG17-b-PCL5 concentration, the amount of precipitated polymer collected by centrifugation after solvent evaporation dramatically decreased from approximately 60% down to less than 1% (Figure 3.1A). Additionally, the hydrodynamic diameter of the nanoparticles also significantly decreased from 163 nm to 80 nm as the concentration of MePEG17-b-PCL5 increased from 0 to 1% w/v   96 (Figure 3.1B). This increase in MePEG17-b-PCL5 concentration also resulted in an decrease in the zeta potential from -21 mV to -5 mV. Nanoparticles prepared by nanoprecipitation and dialysis without MePEG17-b-PCL5 had a hydrodynamic diameter of 100 nm and a zeta potential of -19.1 mV.  Table 3.1 Characterization of synthesized MePEG-b-PCL diblock copolymers. Feed Ratio DPn MW (g/mol) Mn (g/mol) Mw (g/mol) PDI 18:82 MePEG114-b-PCL200 27800 33867 40477 1.20 60:40 MePEG17-b-PCL5 1320 1434 1539 1.07  Notes: Reaction feed ratio of MePEG:caprolactone Abbreviations: DPn, Degree of polymerization determined by 1H NMR; MW, Molecular weight calculated through DPn; Mn, number average molecular weight; Mw, weight average molecular weight; PDI, polydispersity index.        97  Figure 3.1 (A) Mass of polymer in pellet after centrifugation of PCL200/PCL5 nanoparticles at 9500 xg for 5 minutes as a function of MePEG17-b-PCL5 added to nanoparticle dispersions. (B) Hydrodynamic diameter (▲) and zeta potential (□) of PCL200/PCL5 nanoparticles as a function of MePEG17-b-PCL5 added during formation of nanoparticles. Each point represents the average of 3 replicates  SD.   3.3.3 Drug loading From these characterization experiments, the mixed MW PCL200/PCL5 nanoparticle formulation manufactured by emulsification of 1% MePEG114-b-PCL200 w/v with 1% w/v MePEG17-b-PCL5 (i.e a 1:1 weight ratio of MePEG114-b-PCL200 to MePEG17-b-PCL5) was selected for all further experimentation. For PTX, the maximum drug loading was 500 g/ml with a 100% loading efficiency before there was any noticeable drug precipitation or decrease in the loading efficiency (Figure 3.2A). The same mixed MW PCL200/PCL5 nanoparticles were capable of solubilizing considerably more DTX than PTX, solubilizing up to 1300 g/ml with approximately 87% loading efficiency of DTX before any noticeable drug precipitation or decrease in loading efficiency was observed (Figure 3.2B). Similarly, PCL200 nanoparticles solubilized more DTX than PTX; however, the loading efficiency was less than that achieved by the mixed MW PCL200/PCL5 nanoparticles. PCL200 nanoparticles solubilized a maximum of 500 g/ml of PTX with an efficiency of 65%, whereas 980 g/ml of DTX was solubilized with approximately a 65% loading efficiency (Figures 3.2C and 3.2D).  0.00 0.25 0.50 0.75 1.00 1.250102030405060708090MePEG17-b-PCL5 added (% w/v)% of Total Polymerin PelletA0.0 .2 0.50 0.75 1.005075100125150175-25-20-15-10-50MePEG17-b-PCL5 added (% w/v)Diameter  (nm)Zeta Potential (mV)B  98   Figure 3.2 PTX or DTX solubilization () and loading efficiency () for PCL200/PCL5 nanoparticles (A and B) composed of 1% w/v MePEG114-b-PCL200 prepared by emulsification with 1% w/v MePEG17-b-PCL5 or PCL200 nanoparticles prepared by nanoprecipitation and dialysis (C and D) composed of 1% w/v MePEG114-b-PCL200. Each point represents the average of 3 samples  SD.   0 250 500 750 1000503005508000255075100[PTX] added (mg/ml)[PTX] solubilized (mg/ml) Loading efficiency (%)0 250 500 750 100001002003004005006000255075100[PTX] added (mg/ml)[PTX] solubilized (mg/ml) Loading efficiency (%)500 750 1000 1250 1500 1750 2000 22500500100015000255075100[DTX] added (mg/ml)[DTX] solubilized (mg/ml)Loading efficiency (%)500 1000 1500 200050010000255075100[DTX] added (mg/ml)[DTX] solubilized (mg/ml)Loading efficiency (%)A BC D  99 3.3.4 In vitro drug and MePEG17-b-PCL5 release There was little difference in the release profile of DTX solubilized in either mixed MW PCL200/PCL5 or PCL200 nanoparticles (Figure 3.3A). The release of DTX from both these types of nanoparticles was characterized by an initial rapid release of approximately 20% of the encapsulated drug over the first 8 hours. A slower release rate lasting up to 3 days resulted in approximately 80% of the DTX released, followed by a decrease in release rate lasting up to 14 days until all of the drug was released. The release profile of PTX from both types of nanoparticles was similar for both formulations but the rate of release was considerably slower than that observed for DTX. Release of PTX was characterized by an initial rapid release of approximately 10% over the first 8 hours followed by a slower, sustained release of drug. By 14 days, approximately 70% of PTX solubilized in PCL200 nanoparticles was released whereas approximately 60% was released from the mixed MW PCL200/PCL5 nanoparticles over the same time period. The mixed MW PCL200/PCL5 nanoparticles displayed a burst release of MePEG17-b-PCL5 over the first 6 hours followed by a slow release lasting for the remaining 6 days with a total of 85% of the associated MePEG17-b-PCL5 released by six days (Figure 3.3B). The MePEG17-b-PCL5 micelles displayed rapid copolymer release from the dialysis membrane with complete release within one day.     100   Figure 3.3 (A) In vitro release of PTX from mixed MW PCL200/PCL5 nanoparticles () or PCL200 nanoparticles  (□) or DTX release from mixed MW PCL200/PCL5 nanoparticles (◆) or PCL200 nanoparticles (○). Release experiments were conducted in 0.01M PBS at pH 7.4 and 37C. Each point represents the average of 4 samples  SD. (B) In vitro release of MePEG17-b-PCL5 from PCL200/PCL5 nanoparticles (○) or MePEG17-b-PCL5 micelles (■). Release experiments were conducted in distilled water at 37C. Each point represents the average of 3 samples  SD.   0 2 4 6 8 10 12 14020406080100ATime (days)Cumulative Release (%)0 1 2 3 4 5 60255075100Time (days)Cumulative Release (%)B  101 3.3.5 Cytotoxicity of MePEG-b-PCL nanoparticulate formulations The influence of 3-day incubations of PCL200 or mixed MW PCL200/PCL5 nanoparticles on the proliferation of MDCKII and MDCKII-MDR1 cells, in the absence of drug, was investigated. The PCL200 nanoparticles were well tolerated by MDCKII and MDCKII-MDR1 cells at all concentrations tested (Figure 3.4A). The cell viability of both MDCKII and MDCKII-MDR1 cells decreased significantly in the presence of mixed MW PCL200/PCL5 nanoparticles at MePEG114-b-PCL200 concentrations above 0.25% w/v (Figure 3.4B). In the presence of drug-loaded formulations or free drug during a 90-minute incubation, MDCKII cell proliferation was dramatically inhibited (Figure 3.5A and 3.5B). There was no difference in the proliferation of these cells when incubated for 90 minutes in the presence of PTX loaded mixed MW PCL200/PCL5 or PCL200 nanoparticles with both formulations having an IC50 value of approximately 7 µg/ml of PTX (Figure 3.5A). Incubations with DTX loaded formulations were more effective at inhibiting cell proliferation of MDCKII cells than PTX formulations with similar cell proliferation profiles for both nanoparticulate formulations and an IC50 value below the minimum tested DTX concentration of 0.4 µg/ml (Figure 3.5B). Treatment with free PTX or DTX was found to be slightly more effective at inhibiting cell proliferation of MDCKII than the nanoparticle encapsulated drug. Substantially higher drug concentrations were required to inhibit proliferation of MDCKII-MDR1 cells after a 90-minute incubation for all formulations tested (Figures 3.5C and 3.5D). The IC50 values for PTX loaded mixed MW PCL200/PCL5 or PCL200 nanoparticles were both approximately 350 µg/ml (Figure 3.5C). Treatment with DTX loaded formulations resulted in lower IC50 values than the PTX formulations; however, there was little difference in the inhibition of proliferation with IC50 values of approximately 30 and 40 µg/ml, for DTX loaded mixed MW PCL200/PCL5 or PCL200 nanoparticles, respectively (Figure 3.5D). Free PTX was unable to reduce the cell viability of MDCKII-MDR up to the maximum   102 concentration tested (10 µg/ml) whereas free DTX was more effective at inhibiting proliferation than the nanoparticulate DTX with an IC50 of approximately 6 µg/ml. In the subsequent experiment, MDCKII-MDR1 cells were incubated for 3-days with free PTX or DTX demonstrating improved efficacy with prolonged exposure to the drug with IC50 values of approximately 1 µg/ml for PTX and 0.1 µg/ml for DTX (Figure 3.6A). In the next experiment, viability of MDCK-MDR1 cells was assayed after a three day incubation with PCL200 or PCL200/PCL5 nanoparticles with varying copolymer concentrations loaded with drug at a concentration half that of the IC50 of free drug (i.e. 500 ng/ml PTX or 50 ng/ml DTX) (Figure 3.6B and 3.6C). At a PTX loading of 500 ng/ml, cell viability of MDCKII-MDR1 cells was inhibited 50% by mixed MW PCL200/PCL5 nanoparticles at a copolymer concentration of approximately 0.03% w/v (Figure 3.6B). Likewise, for an even lower drug concentration of 50 ng/ml, DTX loaded nanoparticles inhibited cell viability by 50% at a copolymer concentration of 0.008% w/v (Figure 3.6C). Treatment of MDCKII-MDR1 cells with drug loaded PCL200 nanoparticles resulted in significantly higher cell viability with values remaining at approximately 90% and 80% for PTX or DTX loaded nanoparticles, respectively (Figures 3.6B and 3.6D). Profound differences in cytotoxicity of mixed MW PCL200/PCL5 nanoparticles compared to PCL200 nanoparticles were found when MDCKII-MDR1 cells were initially incubated for 90 minutes with relatively low concentrations of PTX or DTX (25µg/ml and 6 µg/ml, respectively) solubilized in mixed MW PCL200/PCL5 nanoparticles, followed by removal of the formulations and a subsequent 3-day incubation with blank mixed MW PCL200/PCL5 or PCL200 nanoparticles, (i.e. in the absence of drug loading) (Figure 3.7). Pretreatment with DTX loaded nanoparticles was more cytotoxic than pretreatment with PTX loaded nanoparticles with nearly complete inhibition of cell proliferation after the subsequent incubation with 0.125 % w/v blank mixed   103 MW PCL200/PCL5 nanoparticles (Figure 3.7B) whereas pretreatment with a higher loading of PTX and subsequent incubation with the same concentration of mixed MW PCL200/PCL5 nanoparticles led to 60% cell viability (Figure 3.7A). Treatment of cells with blank PCL200 nanoparticles after pretreatment with PTX loaded nanoparticles did not result in inhibition of the proliferation of MDCKII-MDR1 cells whereas for DTX, cell viability initially decreased but then remained constant at approximately 70% cell viability, regardless of the concentration of PCL200 nanoparticles.     104         Figure 3.4 The cytotoxicity of PCL200 nanoparticles (A) or mixed MW PCL200/PCL5 nanoparticles (B) on MDCKII cells (○) and MDCKII-MDR1 cells (■). Cells were incubated with various concentrations of nanoparticles in the presence of culture medium for 3 days. The cell viability were determined by the MTS assay. Each point represents the mean of three independent experiments (±SEM).   0.0 0.2 0.4 0.6050100A[PCL200] (%w/v)% Cell Viability 0 2 4 0.6050100B[PCL200/PCL5] (%w/v)% Cell Viability   105                  Figure 3.5 The cytotoxicity of free drug or nanoparticle encapsulated PTX or DTX on MDCKII (A and B) or MDCKII-MDR1 (C and D). Treatment groups were drug loaded mixed MW PCL200/PCL5 nanoparticles (■) or PCL200 nanoparticles (▲) or free drug (○). Cells were incubated with various concentrations of free drug or drug encapsulated in different nanoparticle formulations for 90 min followed by washing. After washing, the cells were incubated for 3 days in culture medium. The cell viabilities were determined by the MTS assay. Each point represents the mean of three independent experiments (±SEM). (E) A schematic illustration of experimental sequences.   0.01 0.1 1 10 100 1000020406080100[PTX]  mg/mlMDCKII% Cell Viability0.01 0.1 1 10 100 1000020406080100120[PTX]  (mg/ml)MDCKII-MDR1% Cell Viability0.01 0.1 1 10 100 1000020406080100120[DTX]  (mg/ml)MDCKII-MDR1 % Cell ViabilityABC D0.01 0.1 1 10 100 1000020406080[DTX]  (mg/ml)MDCKII% Cell Viability  106           Figure 3.6 (A) Cell viability of MDCK-MDR1 cells after 3 day incubation with free PTX (○) or free DTX (■). The effect of concentration of PCL200 or mixed MW PCL200/PCL5 nanoparticles on the cytotoxicity of (B) PTX or (C) DTX in MDCKII-MDR cells. Cells were treated with varying concentrations of PCL200/PCL5 (■) or PCL200 (○) nanoparticles (expressed as concentration of MePEG114-b-PCL200) loaded with a fixed concentration of 500 ng/ml of PTX (B) or 50 ng/ml of DTX (C) for 3 days. The cell viability was measured using MTS assay. Each point represents the average of 3 samples  S.E.M. (D) A schematic illustration of experimental sequences for panels B and C.   0.01 0.1 1 10050100A[Drug] mg/mlMDCKII-MDR% Cell Viability0.0 0.1 0.2 0.3050100B[MePEG114-b-PCL200] (%w/v)% MDCKII-MDR1 Cell Viability0.0 0.1 0.2 0.3050100C[MePEG114-b-PCL200] (%w/v)% MDCKII-MDR1Cell Viability  107       Figure 3.7 The effect of the concentration of blank (no drug) PCL200 or mixed MW PCL200/PCL5 nanoparticles on the cytotoxicity of MDCKII-MDR1 after pretreatment with PTX or DTX loaded nanoparticles. MDCKII-MDR1 cells were pre-treated with mixed MW PCL200/PCL5 nanoparticles with (A) PTX or (B) DTX loadings of 25 µg/ml or 6 µg/ml, respectively, for 90 minutes followed by washing and subsequent incubation in culture media for 3 days with different concentrations of either blank (no drug) mixed MW PCL200/PCL5 (■) or blank (no drug) PCL200 nanoparticles (●) (expressed as concentration of MePEG114-b-PCL200). The cell viability was measured using MTS assay. Each point represents the mean of three independent experiments (±SEM). (C) A schematic illustration of experimental sequences.   A0.0 0.1 0.2 0.30255075100125[MePEG114-b-PCL200] (%w/v)MDCKII-MDR1% Cell ViabilityB0.0 0.1 0.2020406080[MePEG114-b-PCL200] (%w/v)MDCKII-MDR1% Cell Viability  108 3.3.6 Pharmacokinetics and biodistribution of nanoparticulate formulations Using radiolabeled drug and copolymer, the pharmacokinetics and biodistribution of PTX and DTX loaded in mixed MW PCL200/PCL5 nanoparticles as well as solubilized in the commercial formulations was determined. The plasma concentration versus time profiles for PTX and DTX indicate that, regardless of the formulation, the drug was cleared very rapidly from the circulation with rapid distribution phases and less than 1% of the administered dose remaining in the plasma after only 30 minutes (Figure 3.8A and 3.8B). PTX formulated as Paclitaxel for Injection had a higher area under the curve (AUC) and longer half-life (t1/2), lower clearance (CL) and volume of distribution (Vd) than the nanoparticles (Table 3.2). Similar to the PTX formulations, DTX was rapidly eliminated when administered as mixed MW PCL200/PCL5 nanoparticles, resulting in a lower AUC and higher CL, and Vd values as compared to the commercial formulation, Taxotere®. The mixed MW PCL200/PCL5 nanoparticles displayed similar plasma concentration versus time profiles regardless of whether PTX or DTX was formulated in the nanoparticles (Figure 3.8C). The copolymer remained in the plasma for longer than the drugs with approximately 3% of the injected dose remaining 24 hrs post injection with significantly higher AUC values and lower CL and Vd values compared to PTX or DTX (Table 3.2). Very low levels of PTX and DTX were detected in the organs 24 hrs after administration for the nanoparticles with the most drug found in the liver. Considerably more PTX and DTX formulated as the commercial formulations was found in the liver 24 hours after injection (Figure 3.9A). The majority of the copolymer was found in the liver for both nanoparticulate formulations with more of the polymer from the DTX formulation found in all organs as compared to that of the PTX nanoparticles (Figure 3.9B).     109     Figure 3.8 Plasma drug or copolymer concentration vs. time profiles for (A) PTX formulated as Paclitaxel for Injection (▽), or mixed MW PCL200/PCL5 nanoparticles () or (B) DTX formulated as Taxotere® (□) or mixed MW PCL200/PCL5 nanoparticles (■) or (C) MePEG114-b-PCL200 in PTX loaded mixed MW PCL200/PCL5 nanoparticles () or DTX loaded mixed MW PCL200/PCL5 nanoparticles (■). Each point represents average of 4 samples  SD.     110    Table 3.2 Pharmacokinetic parameters for PTX, DTX and MePEG114-b-PCL200 Drug or Copolymer t1/2 (h) AUC0-∞ (h∙µg/ml) CL (ml/h/kg) Vd (ml/kg) MRT (h) PTX in mixed MW PCL200/PCL5 10.04±0.20 0.47±0.01 1885.61±20.15 15875.76±143.75 4.70±0.058 PTX in Paclitaxel for Injection 18.67±0.74 1.66±0.07 394.31±15.76 8993.55±359.72 7.91±0.32 DTX in mixed MW PCL200/PCL5 14.99±0.69 0.901±0.04 858.76±39.50 13109.63±603.01 6.38±0.29 DTX in Taxotere® 8.63±1.90 1.57±0.34 542.94±119.46 5643.04±1241.46 5.61±1.23 MePEG114-b-PCL200 (PTX nanoparticles) 14.78±0.63 3110.44±143.06 22.92±1.05 429.18±19.74 8.01±0.37 MePEG114-b-PCL200 (DTX nanoparticles) 13.90±0.20 4313.65±15.15 17.47±0.25 290.35±3.15 7.65±0.09  Abbreviations: t1/2, elimination half life; AUC, area under the curve; CL, clearance; Vd, volume of distribution at steady state; MRT, mean residence time.     111   Figure 3.9 Biodistribution in organs 24 hours post injection of (A) PTX formulated in mixed MW PCL200/PCL5 nanoparticles (diagonal stripes) and Paclitaxel for Injection (white) or DTX formulated in mixed MW PCL200/PCL5 nanoparticles (grey) and Taxotere® (black) or (B) MePEG114-b-PCL200 in PTX loaded mixed MW PCL200/PCL5 nanoparticles (white) or DTX loaded mixed MW PCL200/PCL5 nanoparticles (black). Each bar represents the average of 4 samples ±SD.   spleen liver kidney heart lung02681012A% Injected Dose of Drugspleen liver kidney heart lung01020304050B% Injected Dose of Copolymer  112 3.4 Discussion Through a ring opening polymerization of -caprolactone using the hydroxyl group of MePEG as the macroinitiator and stannous octoate as a catalyst, we synthesized both MePEG17-b-PCL5 and MePEG114-b-PCL200 (Table 3.1). In agreement with previous reports by our group, the synthesis of MePEG17-b-PCL5 resulted in a copolymer with a predictable molecular weight and composition controlled by the feed weight ratio of the initiator and monomer as well as a relatively narrow molecular weight distribution (Letchford and Burt, 2012; Letchford et al., 2008; Zastre et al., 2002). The molecular weight distribution of MePEG114-b-PCL200 was considerably broader than that of MePEG17-b-PCL5, which is typical for a polymer of this molecular weight synthesized by this method. The molecular weight, calculated using the NMR spectra, indicated that the composition of the copolymer was that which was predicted based on the feed weight ratio of MePEG to caprolactone. Due to the poor water solubility of the MePEG114-b-PCL200, a nanoprecipitation and dialysis technique was used to form nanoparticles, referred to as PCL200 nanoparticles. To form mixed MW PCL200/PCL5 nanoparticles, the addition of MePEG17-b-PCL5 allowed for the oil-in-water emulsification method to be employed. As shown in Figure 3.1, an increase in the amount of MePEG17-b-PCL5 used during the emulsification process led to a decrease in the amount of precipitated polymer (and therefore an increase in the yield of nanoparticles) and a decrease in the diameter of nanoparticles. As the MePEG17-b-PCL5 was initially dissolved in the oil phase, these results suggest that this low molecular weight copolymer migrates to the oil/water interface where it acts to stabilize the emulsion, allowing for the formation of smaller, more stable nanoparticles. Nanoparticles formed in the absence of MePEG17-b-PCL5, either by emulsification or nanoprecipitation, had highly negative zeta potentials. This negative charge is likely due to the presence of carboxylate groups present from hydrolyzed poly(caprolactone) and an insufficient   113 PEG coating to shield this charge (Gref et al., 2000; Radovic-Moreno et al., 2012). Increasing the MePEG17-b-PCL5 concentration used during emulsification decreased the zeta potential likely due to the increased encapsulation/association of the lower molecular weight copolymer with the nanoparticles and thus shielding the negative charge of the nanoparticle (Figure 3.1B). These changes in the physicochemical properties of the mixed MW PCL200/PCL5 nanoparticles provide evidence of the co-encapsulation of the low molecular weight MePEG17-b-PCL5 in the nanoparticles.  Both PCL200 and mixed MW PCL200/PCL5 nanoparticles were capable of solubilizing relatively high concentrations of PTX and DTX (Figure 3.2). In comparison to our previous work, in which we investigated the solubilization of PTX using MePEG114-b-PCL104 nanospheres, the nanoparticles reported in this work were capable of solubilizing considerably more PTX (Letchford and Burt, 2012). This difference may be due to the increased PCL block length, which in turn produce nanoparticles with a larger hydrophobic core, thus, providing a greater cargo space for the loading of the drug. Additionally, the emulsification procedure for the formation and loading of mixed MW PCL200/PCL5 nanoparticles allowed for a significant increase in the drug encapsulation efficiency which was likely due to avoidance of drug loss caused by the dialysis procedure (only required for nanoprecipitation method). Interestingly, significantly more DTX was solubilized by these nanoparticles as compared to PTX. Carstens et al found similar results in their investigation of the solubilization of PTX and DTX by MePEG-b-PCL micelles. This group attributed the solubilization differences between the two drugs to the slightly lower hydrophobicity of DTX and more favorable interactions between the micellar core and the drug (Carstens et al., 2008). PTX and DTX were released from the nanoparticles in a sustained fashion with a small burst release of both drugs, followed by a slower release rate (Figure 3.3). The burst release may   114 be attributed to weakly bound drug that may be located near the particle surface. DTX loaded nanoparticles had a larger burst release and a faster overall release throughout the experiment. Furthermore, there was very little difference in the release profile between PCL200 and mixed MW PCL200/PCL5 nanoparticle formulations for each drug. It is speculated that the larger burst release of DTX, as compared to PTX, was due to a larger amount of DTX at the MePEG/PCL interface as well as the higher aqueous solubility of DTX. Mugabe et al, who compared the release of PTX and DTX from hyperbranched polyglycerol nanoparticles, also noted a faster DTX release rate, which they attributed to the 7-fold difference in aqueous solubility between the two drugs (Mugabe et al., 2011a). Release of MePEG17-b-PCL5 from mixed MW PCL200/PCL5 nanoparticles was characterized by an initial rapid release likely due to release of loosely associated or free MePEG17-b-PCL5. After this initial rapid release, there was a prolonged controlled release of MePEG17-b-PCL5 due to the copolymer encapsulated or tightly associated with the mixed MW PCL200/PCL5 nanoparticles. As a control, release of MePEG17-b-PCL5 from a micelle dispersion was monitored demonstrating that release of MePEG17-b-PCL5 on its own was rapid and complete by one day. To assess the cytotoxicity of the taxane formulations, we chose MDCKII and MDCKII-MDR1 cells, as this pair has been demonstrated to be a good model for studying the modulation of Pgp over-expressing cells (Tang et al., 2002; Taub et al., 2005). Initial studies demonstrated that both nanoparticles had a high degree of biocompatibility with these cells at concentrations up to 0.25% w/v of MePEG114-b-PCL200 (Figure 3.4). Additional experiments investigating the cytotoxicity of drug-loaded formulations were all kept below this copolymer concentration. Both free and nanoparticulate encapsulated PTX and DTX effectively inhibited the proliferation of MDCKII cells after a 90-minute incubation with DTX formulations having lower IC50 values (Figure 3.5A and 3.5B). This trend was also seen in the MDCKII-MDR1 cells although the IC50   115 for both drugs was dramatically increased due to the drug efflux effect of Pgp in these cells (Figure 3.5C and 3.5D). Several reasons for this enhanced cytotoxicity of DTX over that of PTX are possible including the enhanced binding of DTX to β- tubulin as compared to PTX, which has been shown for several cell lines (Kelland and Abel, 1992; Garcia et al., 1994). Additionally, as demonstrated in our release study, the higher aqueous solubility of DTX may lead to a greater amount of free drug available for binding to β- tubulin either released in the external growth media or released within cells after uptake of intact nanoparticles. This is further supported by the lower IC50 values for free PTX and DTX as compared to the nanoparticulate formulations for the MDCKII cells indicating that likely a large portion of the drug is sequestered in the nanoparticles and not freely available for diffusion through the cell membrane or binding to the cellular target during the short 90-minute incubation. However, it was shown that the PTX loaded nanoparticulate formulations were able to solubilize PTX at concentrations high enough to inhibit proliferation of MDCKII-MDR1 cells whereas, the maximum concentration of free PTX used to treat the cells was limited by the solubility of the drug in the media (maximum concentration of 10 µg/ml) which was not high enough to inhibit cell growth (Figure 3.5C). Our group has previously demonstrated the ability of MePEG17-b-PCL5 to modulate the efflux of a variety of Pgp substrates and therefore enhance the uptake of these compounds in Pgp overexpressing cell lines (Elamanchili et al., 2009; Zastre et al., 2008). After a 90-minute incubation, there was no effect of the incorporation of MePEG17-b-PCL5 in the mixed MW PCL200/PCL5 nanoparticles, as shown by very little difference in the IC50 values of MDCKII-MDR cells for the PCL200 and mixed MW PCL200/PCL5 nanoparticle formulations (Figure 3.5C and 3.5D). As demonstrated in previously published studies, after an initial incubation with PTX, it was essential to maintain the presence of MePEG17-b-PCL5 in the media to enable inhibition of Pgp and hence retention of PTX within MDCKII-MDR1 cells (Wan et al., 2013). Therefore, it is   116 possible that after incubation with the cells and subsequent removal of the formulations, the concentration of MePEG17-b-PCL5 associated with the cells did not remain high enough to prevent drug efflux by Pgp. In light of the results presented in Figure 3.5 it was apparent that long-term treatment with the Pgp inhibitor was required to inhibit the growth of MDCKII-MDR1 cells. Therefore, we investigated the effect of continuous 3-day incubation of MDCKII-MDR1 cells with free drug or drug loaded nanoparticles with varying concentrations of copolymer (Figure 3.6). As expected, prolonged incubation with free drug resulted in improved inhibition of growth of MDCKII-MDR1 cells compared to the short 90-minute incubation with either PTX or DTX (Figure 3.6A). MDCKII-MDR1 cells were then incubated for 3 days in the presence of mixed MW PCL200/PCL5 or PCL200 nanoparticles with a drug loading half that of the free drug (50 and 500 ng/ml for DTX and PTX, respectively) and varying concentrations of copolymer. Treatment with drug loaded PCL200 nanoparticles did not lead to significant inhibition of cellular proliferation due to the inability of these nanoparticles to inhibit drug efflux by Pgp. The inclusion of MePEG17-b-PCL5 in mixed MW PCL200/PCL5 nanoparticles resulted in significant cytotoxicity of MDCKII-MDR1 demonstrating that, even with a low drug loading, mixed MW PCL200/PCL5 nanoparticles were capable of delivering MePEG17-b-PCL5 at a concentration high enough to modulate Pgp and inhibit the efflux of PTX or DTX. However, it was required that the mixed MW PCL200/PCL5 nanoparticles were maintained at a minimum concentration and were present for the entire duration of the 3-day incubation. In additional studies we demonstrated that even a short term exposure (90 minutes) of drug loaded mixed MW PCL200/PCL5 nanoparticles, followed by 3-day incubation of blank (no drug) mixed MW PCL200/PCL5 nanoparticles were highly effective at inhibiting proliferation of MDCKII-MDR1 cells (Figure 3.7). In these experiments drug loadings of PTX and DTX were well below cytotoxic levels for MDCKII-MDR1 cells for this short incubation period (25 and   117 6.25 µg/ml, respectively); however when combined with incubation with blank (no drug) mixed MW PCL200/PCL5 nanoparticles for 3 days, proliferation of these highly drug resistant cells was effectively inhibited (Figure 3.7). Clearly, this was the effect of the inhibition of Pgp by MePEG17-b-PCL5 as incubation with blank (no drug) PCL200 nanoparticles did not lead to cytotoxicity. Again, it was found that DTX was more effective at inhibiting cellular proliferation than PTX, as cells were initially incubated with less DTX and cell proliferation was inhibited at a lower concentration of blank (no drug) mixed MW PCL200/PCL5 nanoparticles. As mentioned above, this could be the result of several factors including the enhanced binding of DTX to β-tubulin, as well as the faster release of DTX from the nanoparticles. Since studies by Kabanov and co-workers demonstrated the effectiveness of Pluronic® agents on the inhibition of Pgp (Alakhova et al., 2010; Batrakova et al., 2004) there have been numerous reports of the formulation of taxanes in mixed nanoparticle systems for the treatment of drug resistant cancers utilizing a variety of Pluronic® agents (Ma et al., 2010; Mei et al., 2009; Mu et al., 2010; Zhang et al., 2011; Zhang et al., 2010b). These studies showed that these formulations are effective at increasing the cytotoxicity of taxanes in MDR cells; however, in some cases it was demonstrated that the blank formulations have significant cytotoxicity at relatively low polymer concentrations. Clearly, an advantage of the mixed molecular weight nanoparticles reported in the current study is the low cytotoxicity of the blank formulations. Cell proliferation was not affected by our blank mixed MW PCL200/PCL5 nanoparticles until concentrations above 2.5 mg/ml were reached (Figure 3.4), whereas, for example, the Pluronic P123/F127 mixed micellar systems studied by Zhang et al. began to show significant signs of cytotoxicity at concentrations above 0.1 mg/ml (Zhang et al., 2011; Zhang et al., 2010b). The high degree of cellular biocompatibility in our work allowed us to maintain relatively high, non-cytotoxic concentrations of the copolymers enabling the inhibition of Pgp resulting in the   118 complete killing of the MDR cell line at low drug concentrations (500 ng/ml and 50 ng/ml for PTX and DTX, respectively). In many studies using mixed micellar formulations, complete cytotoxicity of the MDR cell line was not achieved resulting in cell viability values of 20% or more in some cases at drug concentrations of 10µg/ml or higher (Li et al., 2010; Shah et al., 2009; Xiao et al., 2011; Zhang et al., 2011). Upon IV administration to mice, drug solubilized by mixed MW PCL200/PCL5 nanoparticles was eliminated more rapidly than from commercial formulations (Figure 3.8) as evidenced by the lower AUC values and higher CL of the nanoparticulate formulations as compared to the commercial formulations, Paclitaxel for Injection and Taxotere® (Table 3.2). The copolymer remained in the circulation for an extended period of time, eventually accumulating, for the most part, in the liver (Figure 3.9B). However, the organ biodistribution after 24 hours shows that very little drug, originally encapsulated in the nanoparticles, was found in the organs (Figure 3.9A). Therefore, it is likely that upon injection, the drug rapidly partitioned out of the mixed MW PCL200/PCL5 nanoparticles and was quickly metabolized and eliminated from the body. The commercial formulations displayed considerably more drug accumulation in the liver 24 hrs post injection, due to the longer circulation time of the drug than the nanoparticle formulations. The longer circulation of PTX formulated in Paclitaxel for Injection is likely due to the ability of Cremophor EL® to sequester PTX in micelles and thus retaining the drug in the central compartment resulting in prolonged circulation (Sparreboom et al., 1999). Additionally, Tween 80 present in Taxotere®, has been shown to influence the pharmacokinetics of the drug, with low levels of Tween 80, such as those used in this study, resulting in a decreased unbound fraction of drug and hence, a decreased clearance (Loos et al., 2003). It is interesting to note that our previous studies demonstrated enhanced stability of the nanoparticulate systems in vitro in the presence of plasma and whole blood as the molecular   119 weight of the hydrophobic core-forming block increased. However, this enhanced blood or plasma in vitro stability did not translate to prolonged drug circulation upon IV administration (Letchford and Burt, 2012). The current system also did not demonstrate prolonged circulation over that of the commercial formulations, even despite the fact that the core-forming block had nearly twice the molecular weight of the previously tested MePEG-b-PCL nanoparticles. Several groups have reported prolonged circulation of taxanes solubilized by Pluronic or high molecular weight polyester homopolymer nanoparticles (Han et al., 2006; Mu et al., 2010; Win and Feng, 2006; Zhang et al., 2011; Zhang et al., 2008). However, in several instances, encapsulation of taxanes in polyester based diblock copolymer nanoparticles has not produced a pharmacokinetic benefit over that of the commercial formulations (Lee et al., 2011a; Li et al., 2010; Wang et al., 2011). In light of the current results along with those found in our previous study, it is clear that an increase in PCL block length of MePEG-b-PCL does not improve the in vivo retention of taxanes with the nanoparticle.  3.5 Conclusion In this work we demonstrated that mixed MW nanoparticles composed of MePEG114-b-PCL200 and MePEG17-b-PCL5 formed through an emulsification process are capable of solubilizing high levels of PTX or DTX and are characterized by relatively small diameters, amenable to IV administration of these drugs for the treatment of solid tumors. In vitro, both drugs were released at a controlled rate with faster release for DTX than PTX regardless of whether MePEG17-b-PCL5 was present in the formulation. Furthermore, we demonstrated that these systems are capable of overcoming Pgp mediated efflux of PTX and DTX resulting in cytotoxicity of Pgp overexpressing MDCKII-MDR1 cells at low drug concentrations; however it was essential for the mixed MW copolymer nanoparticles to be incubated with the cells for   120 prolonged periods of time. Further studies are ongoing to optimize the encapsulation and release of both taxanes and MePEG17-b-PCL5 in nanoparticles that might allow for the intracellular controlled release of both agents for improved cytotoxicity against drug resistant cancer cells. Furthermore, future studies are ongoing to modify the MePEG-b-PCL copolymers to better retain the taxane payload to provide increased blood circulation times.  3.6 Acknowledgments This work was supported by an operating grant awarded to Helen M. Burt by the Canadian Institutes of Health Research.      121 Chapter  4: Increased accumulation and retention of micellar paclitaxel in drug sensitive and P-glycoprotein expressing cell lines following ultrasound exposure5 4.1 Introduction The clinical use of ultrasound includes diagnostic sonography, real time blood flow imaging and therapeutic applications such as heating for physiotherapy and the thermal ablation of tumors in oncology (Schroeder et al., 2009). Ultrasound equipment found in hospital settings such as HIFU for thermal ablation of tumors provides millimeter level accuracy in focused beams and may be combined with image guidance to allow for precise targeting of diseased tissues.   Furthermore, the development of ultrasound-enhanced chemotherapy approaches has resulted in increased diffusion of drugs through tumors, increased cellular uptake of drugs (sonoporation) and localized release of drugs from controlled release nanoparticles at target sites following systemic administration (Nelson et al., 2002; Nomikou et al., 2010). Many studies of ultrasound-mediated drug effects have used the anticancer drug doxorubicin, due to its various properties, including the ability to encapsulate within micellar nanoparticles, accumulation in tumor tissues via the enhanced permeation and retention effect and evidence of release from micelles by ultrasound exposure with subsequent increased tumor cell uptake (Husseini et al., 2000; Marin et al., 2001b; Nelson et al., 2002; Rapoport, 2004).     We have developed a sonoporation method using short burst (10 seconds), high intensity (32 W/cm2), and high frequency (4MHz) ultrasound to increase the cellular uptake of many agents including antisense oligonucleotides, micellar PTX and doxorubicin in both PC3 prostate cancer and human umbilical vein endothelial (Huvec) cell lines (Siu et al., 2007a; Siu et al.,                                                  5 A version of this chapter has been published. Wan C.P. et al. (2012) Ultrasound Med Biol. 38(5): 736-44.   122 2007b; Jackson et al., 2011). PTX is a highly effective drug for treating cancer with an IC50 (inhibitory concentration that kills 50% of cells) in the low nanomolar range (approximately 10 fold lower than doxorubicin) for most proliferating cells including Huvec cells (Jackson et al., 2005). However, PTX has an extremely low water solubility (approximately 1 μg/ml) so that micellar delivery systems are typically used for administration of this drug. Micellar delivery systems are composed of amphiphilic agents which self-assemble into micelles with a hydrophobic core, containing the drug, and a hydrophilic corona, when placed in an aqueous environment above the CMC (Letchford and Burt, 2007). We recently demonstrated that ultrasound exposure (4MHz, 32 W/cm2, 10 second exposure) had little effect on the cellular uptake of free PTX but greatly enhanced the uptake of micellar PTX in PC3 and Huvec cell lines (Jackson et al., 2011). Using xenograft tumors grown in mice, other studies have demonstrated that externally applied ultrasound, using frequency 1MHz, intensity 1.7W/cm2, and sonication duration up to 90 seconds, may enhance the antitumor action of PTX following the intravenous administration of micellar paclitaxel (Howards et al., 2006).   Some evidence suggests that doxorubicin is released from Pluronic® based micelles following ultrasound exposure, to create high local concentrations of the drug (Rapoport et al., 1999; Pruitt and Pitt, 2002). We believe that the increased cellular PTX levels caused by sonoporation effects for micellar PTX may arise from either increased uptake of intact micelles or micelle disruption at the cell membrane with increased drug penetration into cells (Jackson et al., 2011). Indeed, many studies suggest that ultrasound may increase the uptake of intact doxorubicin micelles into cancer cells as well as increasing drug release rates (Husseini et al., 2000; Marin et al., 2001b; Rapoport, 2004). The pharmacological site of action of some chemotherapeutic drugs may further affect retention in cells. For example, anthracyclines, like doxorubicin, cross the nuclear membrane and bind to DNA so there is little efflux of doxorubicin   123 from the cytoplasm. Therefore, doxorubicin is effectively retained within the cell.  On the other hand, when PTX is inside the cells, it binds reversibly to β-tubulin in the cytoplasm and is more readily transported actively or passively out of the cell (Elamanchili et al., 2009; Rowinsky et al., 1990; Zastre et al., 2008). This is particularly important in chemoresistant cells expressing drug efflux proteins (especially P-glycoprotein (Pgp)) in the plasma membrane, which greatly increase the rate of efflux of drugs such as doxorubicin and PTX from cells via an active process (Szakacs et al., 2006). It has been reported that ultrasound may enhance the uptake of drugs that are Pgp substrates in chemoresistant cancer cells, presumably by overwhelming Pgp-mediated efflux in the face of a sudden increased influx of drug (Cho et al., 2002; Ng and Matsunaga, 2005). Interestingly, the addition of a small molecule Pgp inhibitor in those studies allowed for the increased retention of doxorubicin in the cells with resulting decreases in cell viability (Ng and Matsunaga, 2005).   The objective of this study was to investigate the effect of ultrasound on the uptake and retention of PTX loaded amphiphilic diblock copolymer micelles in Pgp expressing cells and their non-Pgp expressing counterparts. PTX was solubilized using a previously established micellar drug delivery system (Zhang et al., 1996) composed of  methoxy poly(ethylene glycol)-block-poly(D,L-lactide) (MePEG-b-PDLLA) diblock copolymer.  Using either MDCKII and MDCKII-MDR1 (Pgp expressing version of MDCKII) or MCF-7 and NCI/ADR (Pgp expressing match to MCF-7) cell lines, with a 4MHz, 32 W/cm2 intensity, 10 second ultrasound exposure, we found enhanced uptake of drug in all cells. The efflux rates of PTX from ultrasound treated cells were slower than those for non-treated cells but no difference was observed in efflux rates between Pgp expressing cells and their non-Pgp counterparts. These data suggest that ultrasound may facilitate the uptake of intact paclitaxel loaded micelles into cells allowing greater retention of drug in both Pgp and non-Pgp expressing cells.    124 4.2 Materials and methods 4.2.1 Materials Paclitaxel was obtained from PolymedT Inc. (Miami, FL, USA). Radioactive paclitaxel (3H) (1μCi /μl was obtained from Moravek Radiochemicals (Brea, CA, USA). MDCKII and MDCKII-MDR1 cells were kind gifts of Dr. Piet Borst (National Cancer Institute, Netherlands). MCF-7 and NCI-ADR were generous gifts from Dr. Marcel Bally (British Columbia Cancer Research Centre, Canada). The MTS assay kit was purchased from Promega (Madison, WI). 1, 1’-dioctadecyl-3,3,3’,3’-tetramethylindocarbocyanine perchlorate (DiIC18), 3,3’-dioctaecyloxacarbocyanine perchlorate (DiOC18), Dulbecco’s modified eagle’s medium (DMEM) media supplemented with 5% FBS and 1% penicillin/streptomycin and Hank’s buffered salt solution (HBSS) were purchased from Invitrogen (Grand Island, NY, USA).  Bovine serum albumin was obtained from Sigma Aldrich (St. Louis, MO, USA). The diblock copolymer (MePEG-b-PDLLA) was composed of MePEG (molecular weight of 2000 g/mol) and 60:40 MePEG/D,L-lactide with a calculated molecular weight of 3333 and was a kind gift from Angiotech Pharmaceuticals (Vancouver, Canada). Slide-A-Lyzer dialysis cassettes (3.5K MWCO) were purchased from Thermo Scientific (Rockford, IL, USA).  4.2.2 MePEG-b-PDLLA micelle preparation Micellar PTX was prepared by the film rehydration method as reported previously (Zhang et al., 1996). Briefly, 80 mg MePEG-b-PDLLA and 24 mg PTX were dissolved in 3ml of acetonitrile and dried under nitrogen at 40°C to form a film. The micellar formulation was made up by rehydrating the film with 80 ml warm HBSS (pH 7.4) and vortexing vigorously to give a final PTX concentration of 300 μg/ml. This micellar dispersion was further diluted as necessary. For drug accumulation studies, micellar PTX was prepared as described above except that a trace   125 amount of radioactive 3H-PTX was added to the dissolved diblock copolymer and cold PTX to give final radioactivity of 1 μCi/ml. It was not necessary to remove free PTX from the medium as the micelles would only release a small percentage of their payload because the solubility limit of PTX (1 μg/ml) would prevent any further release from micelles containing encapsulated PTX at 75-300 μg/ml.  4.2.3 Ultrasound treatment  A custom manufactured ultrasound apparatus consisting of a sealed transducer attached to the base of a shallow acrylic bath was used for ultrasound treatment (Figure 4.1). The details of this apparatus and parameters used were previously reported (Jackson et al., 2011). The bath contained water at 37 °C and allowed for placement of a well of choice directly over the ultrasonic transducer. The distance between the transducer and cell well was 15 mm and the height of the medium in the cell well was 8 mm. The piezoelectric transducer (PZT) was made from a lead zirconate titanate ceramic (PZT-8, material:841, flat, thickness=0.5 mm, rectangular 5 mm x 5 mm). Each well received a 10 second ultrasound exposure at the frequency of 4MHz and an intensity of 32 W/cm2. To avoid continuous ultrasound exposure at this frequency, burst mode (25% duty cycle and 50,000 cycles in each burst) was used to allow for intervals between waves. The spatial-average-temporal-average (ISATA) intensity was measured using an ultrasound power meter (Ohmic Instruments C., Easton, MD, USA). The electrical signal from a function generator (Agilent Inc., Palo Alto, CA, USA) was amplified by a 50 dB broadband power amplifier (Electronic Navigation Industries, Inc., Rochester, NY, USA) before being applied to the transducer. There was no significant increase in temperature (less than 1 °C) caused by the transducer at the transducer-water interface as measured by a thermocouple system. 4.2.4 Physical stability of MePEG-b-PDLLA micelles   126 PTX loaded micelles in PBS (pH 7.4) with or without ultrasound treatment were transferred into a cuvette and incubated in a water bath at 37°C. At each time point, the hydrodynamic diameter of micelles was determined by light scattering measurements carried out on a Malvern NANO-ZS Zetasizer (Malvern Instruments Ltd, Malvern, UK) with a He-Ne laser (632.8 nm) and 12.8°-175° measuring angles.  All the measurements were made at 37°C.   Figure 4.1 A schematic representation of a 48-well cell culture plate, adapted for exposure with ultrasound from an ultrasonic-transducer.  The height of cell culture medium in wells is around 8mm.  The ultrasound applied to the well resulted in the formation of stable cavitation.  4.2.5 Fluorescence resonance energy transfer (FRET) and drug release study The FRET pair DiIC18 and DiOC18 are highly lipophilic due to their 18-carbon alkyl chains.  FRET pair micelles were prepared by a nanoprecipitation and membrane dialysis method as reported previously (Chen et al., 2008a). Briefly, 24 mg of MePEG-b-PDLLA, 0.19 mg of DiIC18 (λEm:501nm) and 0.19 mg of DiOC18 (λEm:565nm), were dissolved in 0.5 ml of acetone.  The solution was added drop-wise into 2 ml of rapidly stirred Phosphate Buffered Saline (PBS)(pH7.4). After 2 h stirring, the solution was transferred into a 1000 MWCO Spectra/Por® Acrylic PlateAmplifierSignal GeneratorUltrasoundTransducerDegassed WaterCell Monolayer48 well plate Micellar drug dispersionAluminum CasingPZTWires to Amplifier  127 dialysis membrane (Spectrum Laboratories, Inc., Rancho Dominguez, CA, USA) and dialyzed against 4 L of PBS overnight to remove unloaded DiIC18 and DiOC18.  250 μl of FRET pair micelles were added to 1.25 ml of release medium, containing PBS and bovine serum albumin (40 mg/ml) at 37°C. At various time points (15, 30, 45, 60, 75, 120min), the fluorescent intensity of the samples was measured using a Varian Cary Eclipse fluorescence spectrophotometer (Varian Inc., Pal Alto, CA). The release of FRET pairs was determined by calculating as follows: FRET Ratio =  IDiIIDiI + IDiO                                                                  (4.1) Where IDiI and IDiO represent the fluorescent intensity of DiI and DiO, respectively.  Nanoparticles with a copolymer concentration of 0.3% (w/v) were loaded with cold PTX and 3H-PTX as described above. The cold PTX loading of micelles was 100 μg/ml. Drug loaded micelles were exposed to ultrasound irradiation (4MHz, 32 W/cm2, 10-second exposure) then added to 3500 MWCO Slide-A-Lyzer dialysis cassettes and dialysed against 4 L of PBS at 37°C with shaking. As a control, PBS in dialysis cassettes was spiked with an aliquot of cold PTX in DMSO containing a trace amount of 3H-PTX so that the resulting drug concentration was 1 μg/ml, equivalent to the aqueous solubility of PTX. At time points of 0, 2, 6, 8, 24, 48 hours, the drug content in the dialysis cassettes was determined by beta scintillation counting. The cumulative percent PTX released from micelles was calculated as follows: (1 −𝑎𝑚𝑜𝑢𝑛𝑡 𝑜𝑓 𝑑𝑟𝑢𝑔 𝑟𝑒𝑚𝑎𝑖𝑛𝑖𝑛𝑔𝑖𝑛𝑖𝑡𝑖𝑎𝑙 𝑎𝑚𝑜𝑢𝑛𝑡 𝑜𝑓 𝑑𝑟𝑢𝑔) ×  100%        (4.2) where amount of drug remaining is the amount of PTX remaining in the dialysis cassette at different time points, and initial amount of drug is the amount of PTX placed in the dialysis cassette at the beginning of the experiment. The data were expressed as cumulative percentage of drug released as a function of time.     128 4.2.6 PTX accumulation and PTX efflux study Cells were plated onto 48-well plates at a density of approximately 10,000 cells/well and grown for two days. Micellar PTX (500 μl, 100 μg/ml) with 1% of 3H-PTX was incubated with cells for 90 min prior to ultrasound treatment. After washing with cold HBSS, cells were lysed using the lysis buffer composed of (2% Triton X-100 in water (60% v/v) and DMSO (40% v/v)) and the intracellular PTX content was measured using a scintillation counter (LS6000TA Beckman Instruments Inc., Fullerton, CA). For drug efflux studies, cells and micellar PTX were prepared as described above. After 90 min incubation with or without ultrasound treatment, cells were washed and incubated with HBSS (37°C) for up to 2 h. At various time points (20, 40, 60 min), cells were washed and lysed as described above. The cellular drug content was measured using liquid-scintillation counting. The percentage of PTX efflux was calculated using the disintegrations per minute (DPM) values from cells lysed at t=0 as the 100% PTX content in the cells.    4.2.7 Cell proliferation study Cells were cultured onto 48-well plates at a density of approximately 3,000 cells/well and grown overnight. Cells were incubated with various concentrations of micellar PTX (75, 150, 300 μg/ml) for 90 min prior to ultrasound treatment. After treatment, cells were washed and allowed to grow in DMEM media for 2 days and the cell viability was measured using the MTS cell proliferation assay kit according to the manufacturer’s instructions. Cell proliferation was expressed as a percentage of absorbance in the control well to the absorbance after treatment.      129 4.2.8 Statistical analysis Statistical significance was analyzed by two-way ANOVA using the Bonferroni post-tests with p<0.05 considered to be statistically significant.  4.3 Results 4.3.1 Physical stability, FRET analysis and drug release Our ultrasound conditions were 4MHz, 32W/cm2 and 10 second exposure. Frequencies in the Megahertz range were used because they offer better resolution and focusing in tissues or cells. Unfortunately, high frequencies and high intensities which are necessary for effective tissue penetration come with an associated danger of heating and damage to normal tissues.  Therefore, it is important to reduce the time of exposure to minimize such effects. Our previous work described the optimization process that led to the use of the described ultrasound regime that allowed for normal cell proliferation without any cytotoxic effect (Siu et al., 2007a; Siu et al., 2007b). PTX is soluble in aqueous media at a concentration of about 1 μg/ml whereas the drug can be solubilized to a much greater concentration (up to 5 mg/ml) using a MePEG-b-PDLLA micellar formulation (Zhang et al., 1996). In this study, PTX was loaded into MePEG-b-PDLLA micelles using a rehydration method. To investigate the effect of ultrasound on the physical stability of the micelles, a dynamic light scattering technique was used to compare the difference between the hydrodynamic diameters of micelles with or without ultrasound exposure. No difference in the particle size of micelles was observed between micelles receiving ultrasound exposure and untreated micelles over 48 hours (Figure 4.2). The drug dispersions were clear with absence of any precipitate during the experiment. Hydrophobic FRET pairs (DiOC18 and DiIC18) were incorporated into the core of micelles. DiIC18 is a red-orange fluorescent lipophilic probe as an acceptor, whereas DiOC18 is a green fluorescent lipophilic   130 probe as a donor. When the two probes are in close proximity (as in the cores of micelles) fluorescent energy is transferred, resulting in high FRET efficiency. In conditions when the two probes are released from the core of micelles, the FRET efficiency decreases as a result of increasing distance between probes. To investigate whether ultrasound treatment would result in any difference in FRET pair release from micelles, a FRET pair release study was carried out in albumin containing release medium. This release medium mirrors the tonicity of physiological fluid. Bovine serum albumin was added to the release medium to increase the aqueous solubility of FRET pairs, so that their release was not slowed by the solubility of the FRET pairs in aqueous media (Magenheim et al., 1993). At various time points, the release of FRET pairs from micelles was detected using fluorescent spectrometry. Both untreated and ultrasound treated groups showed similar changes in FRET pair ratios as a function of time (Figure 4.3a). The effect of ultrasound irradiation on PTX release from micelles was investigated (Figure 4.3b) and showed that PTX release from micelles was not affected by ultrasound irradiation. For both ultrasound treated and untreated groups, PTX was released in a controlled and sustained fashion.  Free PTX was released rapidly with complete release of the drug by 10 hours (data not shown).     131       Figure 4.2 The effect of ultrasound (4MHz, 32 W/cm2, 10 second exposure) on the hydrodynamic diameter (determined using DLS) of PTX loaded MePEG-b-PDLLA micelles.  The PTX loaded micelles were treated with (solid triangle) or without (solid square) ultrasound (US).  All data points represent mean ± SD, (n=3).   0 10 20 30 40 50010203040-US+USTime (hours)Hydrodynamic Diameter (nm)  132     Figure 4.3 a) FRET pair ratio of DiIC18 and DiOC18 within micellar core as a function of time with or without ultrasound (US) treatment (4MHz, 32 W/cm2, 10 second exposure) in release medium. The MePEG-PDLLA micelles were loaded with DiOC18 and DiIC18 and were treated with (solid triangle) or without (solid square) ultrasound. Albumin at 40 mg/ml was used in the release medium for binding FRET probes. The micellar dispersions were incubated at 37°C and the FRET ratio, IDiI/(IDiI + IDiO), was measured over time. b) The effect of ultrasound irradiation on PTX release from micelles. Micellar PTX (100 μg/ml) were exposed to ultrasound irradiation (4MHz, 32 W/cm2, 10 second exposure) and immediately placed in a dialysis insert. At different time points, the PTX contents were measured using beta-scintillation counter. All data points represent mean ± SD, (n=3).   0.0 0.5 1.0 1.5 2.0020406080100+US-USTime (hours)IDiI/(IDiO+IDiI)0 10 20 30 40 50020406080100-US+USTime (hours)Cumulative PTX Release (%)a) b)   133 4.3.2 Cellular PTX accumulation The intracellular accumulation of micellar PTX in untreated cells and cells receiving ultrasound exposure was investigated.  Ultrasound irradiation on its own did not affect the cell proliferation (see Figure S1 and S2 as supporting data in Appendix). MePEG-b-PDLLA micelles were loaded with PTX (final aqueous concentration of 100 μg/ml) and a trace amount of 3H-PTX in HBSS. The drug sensitive MDCKII and MCF-7 cells, and their respective resistant cou-nterparts MDCKII-MDR1 and NCI-ADR were incubated with micellar PTX and then irradiated with ultrasound.  Both untreated and ultrasound treated cells were washed and lysed. The intracellular PTX content was measured using liquid scintillation counting. The drug resistant cell lines accumulated less PTX than drug sensitive cell lines (Figure 4.4). However, both drug sensitive and resistant cell lines showed significant increases (~2 fold) in intracellular PTX levels when treated with ultrasound.   Figure 4.4 Intracellular PTX accumulation with (solid bar) or without (open bar) ultrasound (US) treatment (4MHz, 32 W/cm2, 10 second exposure) in (a) MDCKII and MDCKII-MDR1 cell lines & (b) MCF-7 and NCI-ADR cell lines. Cells were incubated with micellar PTX (100 μg/ml) for 90 min at 37°C prior to ultrasound treatment. Intracellular PTX levels were measured using liquid scintillation counting. All data points represent mean ± SEM, (n=8). *Statistically significant compared to no ultrasound treatment (P<0.05).   MDCKII MDCKII- MDR10500010000150002000025000+US-USn=8**PTX (ng/mg protein)MCF7 NCI-ADR05000100001500020000+US-USn=8**PTX (ng/mg protein)) b)  134 4.3.3 Cellular PTX efflux The effect of ultrasound on cellular PTX efflux was investigated by incubating PTX loaded MePEG-b-PDLLA micelles with both drug sensitive (MCF-7 and MDCKII) and their drug resistant (NCI-ADR and MDCKII-MDR1) counterparts for 90 minutes. After incubation, both untreated and ultrasound treated cells were washed and incubated in HBSS for up to 1 hour. At various time points (0, 20, 40, 60 min), cells were lysed and intracellular PTX content was measured. For both untreated and ultrasound treated groups in both drug sensitive and resistant cell lines, a burst efflux was observed at 20 min, followed by a steady efflux over the next 40 min (Figure 4.5a-d). The percent efflux at 60 min for MDCKII, MDCKII-MDR1, and MCF-7 was approximately 60% (Figure 4.5a-c) and 45% for NCI-ADR cells (Figure 4.5d). For all ultrasound treated groups, the initial burst efflux at 20 min for ultrasound treated groups was less pronounced than that of no ultrasound groups. All ultrasound treated cell lines showed significantly slower drug efflux when compared to untreated groups (Figure 4.5a-d).     135  Figure 4.5 The effect of ultrasound (4MHz, 32 W/cm2, 10 second exposure) on the efflux of PTX from (a) MDCKII, (b) MDCKII-MDR1, (c) MCF-7, and (d) NCI-ADR cell lines. Cells were incubated with micellar PTX (100 μg/ml) for 90 min at 37°C followed by ultrasound (solid triangle) or no ultrasound treatment (solid square).  After washing with HBSS, cells were incubated in HBSS for 20, 40, or 60 min. At each time point, cells were lysed and intracellular PTX levels were measured using liquid scintillation counting. All data points represent mean ± SEM, (n=5 for (a) and (b) and n=7 for (c) and (d)). *Statistically significant compared to no ultrasound treatment (P<0.05).   4.3.4 Cell proliferation Since ultrasound treatments resulted in enhanced intracellular PTX accumulation and prolonged drug retention in all cell lines as shown above, cytotoxicity studies were then performed to examine if this induced a greater anti-proliferative effect of PTX in cells.  Since there was no difference in the uptake or retention effect between cell lines, only MDCKII and MDCKII-MDR1 cell lines were used in this cytotoxicity study. With no ultrasound treatment, 0 20 40 60020406080MDCKII -USMDCKII +US***Efflux Time (min)Efflux PTX (%)0 20 40 60020406080MCF7 -USMCF7 +US**Efflux Time (min)Efflux PTX (%)0 20 40 60020406080MDCKII-MDR1 -USMDCKII-MDR1 +US**Efflux Time (min)Efflux PTX (%)0 20 40 60020406080NCI-ADR -USNCI-ADR +US***Efflux Time (min)Efflux PTX (%)a) b)c) d)  136 PTX inhibited the proliferation of MDCKII cells in a concentration dependent manner up to 300 μg/ml (Figure 4.6a, solid square). However in the MDCKII cells treated with ultrasound, PTX demonstrated an enhanced cytotoxic effect at all concentrations of drug (Figure 4.6a, solid triangle). Regardless of ultrasound treatment, PTX had a reduced cytotoxic effect in drug resistant MDCKII-MDR1 cells as compared to MDCKII cells which are highly sensitive to the drug (Figure 4.6a-b). However, the cytotoxicity of PTX on both MDCKII and MDCKII-MDR1 was significantly enhanced by ultrasound treatment. In the absence of PTX, there was no difference in the cell viability of ultrasound treated and untreated groups indicating that the ultrasound regime did not induce any cytotoxicity.   Figure 4.6 The effect of ultrasound (4MHz, 32 W/cm2, 10 second exposure) on the cytotoxicity of micellar PTX in (a) MDCKII and (b) MDCKII-MDR1 cell lines. Cells were incubated with differing concentrations of micellar PTX for 90 min at 37°C prior to ultrasound treatment (solid triangle) or no ultrasound treatment (solid square).  Immediately, cells were washed with HBSS and incubated in culture medium for 2 days. The cell viability was measured using the MTS assay. All data points represent mean ± SEM, (n=3). *Statistically significant compared to no ultrasound treatment at same concentration (P<0.05).   0 100 200 300050100 MDCKII -USMDCKII +US***PTX Concentration (mg/ml)% Cell Viability0 100 200 300050100 MDCKII-MDR1 -USMDCKII-MDR1 +US* **PTX Concentration (mg/ml)% Cell Viability) b)  137 4.4 Discussion 4.4.1 Physical stability, FRET analysis and drug release The effectiveness of micellar drug delivery systems as controlled release nanoparticles depends significantly on the kinetic stability and integrity of the micelles and the retention of drug with the micelle core (Savic et al., 2006). In this study, dynamic light scattering measurements demonstrated that ultrasound exposure did not alter the integrity of PTX loaded MePEG-b-PDLLA micelles. In the FRET pair release study, both untreated and ultrasound treated groups showed similar changes in FRET pair ratios as a function of time, suggesting that ultrasound treatment did not alter either the integrity of the micelles or the release of the hydrophobic FRET probe payload. The use of ultrasound exposure for enhancing drug release from nano-carriers has been reported. It has been shown that ultrasound at 80 kHz can trigger the release of doxorubicin, a more hydrophilic anticancer drug, from Pluronic® P-105 micelles (Munshi et al., 1997). Other studies have shown that ultrasound (at 20 kHz and 3.3 W/cm2) may trigger the release of drugs encapsulated in liposomes (Schroeder et al., 2007).  Similarly, Xing et al. reported that ultrasound (at 1 MHz and 0.5 W/cm2, with the field on for 10 seconds followed by 5 seconds off, for a total of 2 minutes) triggered PTX release from lipid microbubbles (Xing et al., 2008). However, in our work, ultrasound treatment did not accelerate the release of the hydrophobic FRET pairs and PTX indicating a strong binding association between hydrophobic molecules such as FRET pairs or PTX with the core of the MePEG-b-PDLLA micelles.   4.4.2 PTX accumulation The diameter of MePEG-b-PDLLA micelles used in this study was approximately 20 nm.  It has been shown that nanoparticles with this size may be internalized into cells through   138 clathrin-mediated endocytosis mechanisms and are localized in defined cytoplasmic organelles (Luo et al., 2002; Mahmud and Lavasanifar, 2005; Jiang et al., 2008; Savic et al., 2003; Liaw et al., 1999). In all cell lines, ultrasound induced an approximately 2 fold increase in intracellular PTX levels. These levels of enhanced uptake were similar to those previously reported using the same ultrasound regime and the same micellar PTX formulation in PC3 and Huvec cell lines (Jackson et al., 2011). In addition, ultrasound irradiation had little effect on the intracellular uptake of free PTX on MDCKII and MDCII-MDR1 as published previously (see Figure S3 in supporting data in Appendix). In another study, reported by Howard et al. 2006, the intracellular uptake of PTX loaded in methyl poly(ethylene oxide)-co-poly(L-lactide)-tocopherol micelles (mPEG-PLA-tocopherol) was increased substantially when ultrasound (1MHz, 1.7W/cm2, 30 second exposure) was applied (Howards et al., 2006). Figure 4.4 shows that the cellular PTX levels in sensitive cell lines were higher than their resistant counterparts.  This might be due to the expression of efflux transporter proteins on resistant cell lines, which actively pumps drugs out of cells.  It is well known that PTX is a Pgp substrate and is efficiently removed from the cytoplasm through Pgp mediated efflux (Zhan et al., 1997). The mechanisms of ultrasound-mediated cellular uptake have been suggested to occur via either increased drug release from the carrier, disruption of cell membranes or increased uptake of intact drug/micelle complexes (Husseini and Pitt, 2008a). For doxorubicin loaded Pluronic® micelles, the increased uptake of doxorubicin was due to ultrasound triggered release from micelles (Husseini et al., 2000). However, the ultrasound enhanced PTX accumulation in our study was unlikely to be due to any increase in drug release from the carrier since our experiments with FRET pair release rates do not support this (Figure 4.3a-b). Therefore, the observed enhanced PTX accumulation in this study was likely due to transient disruption of cell membranes and either increased diffusion of free drug from micelles into the cells or increased uptake (endocytosis or pinocytosis) of intact   139 micelles into the cytoplasm. Acoustic cavitation is a physical phenomenon in which gas-filled cavitation bubbles are created in the liquid by a large negative pressure produced by ultrasound (Mason and Lorimer, 2002). This type of cavitation generally occurs with the use of low frequency ultrasound in combination with contrast agent microbubbles. When the cavitation bubbles collapse in the case of inertial cavitation, enormous amounts of pressure exceeding 104 atmospheres are released over a short time, producing shock waves that can impact the surrounding environment (Leighton, 1994; Pecha and Gompf, 2000). Husseini et al. reported that shock waves resulting from the collapse of microbubbles of inertial cavitation at low frequency (70 kHz) were required for release of doxorubicin from Pluronic® micelles (Husseini and Pitt, 2008b). Inertial cavitation was unlikely to occur in our system due to the use of high frequency (4MHz) ultrasound, short pulse length, and long burst cycle (Daniels et al., 1987; Brennen, 1995). The likelihood of inertial cavitation is thought to depend on a mechanical index which has been described as the ratio of acoustic pressure to the square root of the frequency (Apfel and Holland, 1991). It has been reported that the inertial cavitation occurs when the mechanical index is greater than 0.5. In our study, the mechanical index is around 0.3, which further suggests that the collapse cavitation was unlikely to occur. In some cases, stable cavitation can form, in which bubbles oscillate without collapsing. This cavitation may result in a mild perturbation of cell membranes and increased drug transfer through a more porous membrane (Williams and Miller, 1980; Rooney, 1970; Husseini and Pitt, 2008b). Deng et al. reported that stable cavitation could temporarily disrupt the cell membranes, leading to the formation of pores (Deng et al., 2004). In this study, stable cavitation may have allowed for the increased accumulation of micellar and/or free PTX.      140 4.4.3 PTX efflux We have previously reported the rapid efflux of free PTX from drug resistant cell lines (NCI-ADR and MDCKII-MDR1) (Elamanchili et al., 2009) so that only 30% of intracellular PTX was retained in the cells after 30 minute incubation in HBSS. On exposure to ultrasound, this study shows a slightly slower efflux of PTX so that approximately 50% of accumulated PTX was retained in the cells after a 30 minute of incubation (Figure 4.5). Furthermore, despite the fact that all cell lines receiving ultrasound treatment had higher intracellular PTX levels than untreated cells, slower PTX efflux rates were shown for ultrasound treated cells. If ultrasound treatment resulted in enhanced intracellular uptake of PTX loaded micelles and the PTX sequestered in the micelles did not diffuse out of cells as rapidly as free drug, it is conceivable that the decreased efflux of PTX from cells treated with ultrasound was due to a lower intracellular concentration of free drug. Since the intracellular concentration of free PTX was low in micellar-containing cells, drug efflux by diffusion of free drug may decreased. In this study, there was no difference in efflux rates in ultrasound treated drug sensitive and Pgp expressing cell lines, indicating that diffusion of drug out of the cells was the major transport mechanism for the drug. This also suggests that the use of micellar PTX in combination of ultrasound might be a good strategy for treating both drug sensitive and Pgp expressing cell lines.  4.4.4 Cell proliferation There was no cell death associated with the application of our ultrasound regime alone. For both untreated and ultrasound treated cells, micellar PTX had a greater cytotoxic effect on MDCKII cells than on MDCKII-MDR1 cells. This is consistent with previous studies that the IC50 of free PTX in MDCKII cells was more than 10 fold lower than in MDCKII-MDR1 cells   141 due to the expression of Pgp in MDR cells which reduces the intracellular free drug concentration (Elamanchili et al., 2009). Therefore, in MDCKII-MDR1 cell lines, free drug is continuously effluxed from the cells so that a much higher initial concentration is needed to cause any cytotoxic effect. On the other hand, MDCKII cell lines, lacking the expression of Pgp, allow greater retention of PTX in the cytoplasm, and greater cell cytotoxicity. The cytotoxicity of micellar PTX was enhanced by ultrasound treatment for both MDCKII and MDCKII-MDR1 cell lines (Figure 4.6). This effect was likely due to the increased accumulation and the increased retention of micellar PTX in ultrasound treated cells as discussed above. Therefore, as drug releases from the micelles intracellularly, it may then undergo transport out of the cell by diffusion. Sequestration of micellar drug therefore delays the efflux of the drug from cells and maintains a higher intracellular concentration of drug for longer periods ensuring a greater cytotoxic profile of the drug. It has been previously reported that hydrophobic molecules such as PTX loaded in the core of micelles are released from nanoparticles prior to cellular internalization (Chen et al., 2008a). In this study, we hypothesize that the ultrasound treatment may lead to a transient and increased permeability of cell membranes, resulting in internalization of intact micelles loaded with PTX from which PTX could be released in a controlled manner by diffusion as reported by various groups (Gaucher et al., 2010; Kim et al., 2011; Letchford et al., 2009).  4.5 Conclusion Overall, these data support the use of ultrasound to increase the accumulation and retention of micellar PTX in both drug sensitive and resistant cell lines. The resulting increases in the cytotoxic effect of PTX delivered in this manner may improve drug efficacy. Further   142 studies are needed to investigate the effect of ultrasound parameters, ultrasound focusing techniques and the route of administration of micellar PTX on drug efficacy in vivo.   4.6 Acknowledgments This work was supported by a Canadian Institutes of Health Research (CIHR) grant (to HMB), Natural Sciences and Engineering Research Council of Canada (NSERC) Strategic Grant (to MC) and NSERC Discovery Grant (to HMB). Mu Chiao is supported by Canada Research Chairs program, Canada Foundation for Innovations (CFI) and UBC’s Institute for Computing, Information and Cognitive Systems (ICICS).       143 Chapter  5: Summarizing discussion, conclusions, and suggestions for future work 5.1 Summarizing discussion Over the past two decades, there has been significant research interest in the development of novel nanoparticulate formulations for PTX and DTX, with the goal of achieving improved delivery and safer and more effective treatment of various cancers including breast, ovarian, lung, prostate, melanoma, leukemia, and bladder cancers. Abraxane®, Genexol-PM®, and Cellax™ are examples of new taxane nanoparticulate formulations showing promising performance when compared to Taxol® and Taxotere® (Galsky, 2005; Rowinsky and Donehower, 1995; Ernsting et al., 2012a; Yardley, 2013; Lee et al., 2012). Due to the hydrophobicity and very limited water solubilities of PTX and DTX, the commercial parenteral formulations, Taxol® and Taxotere®, contain high concentrations of the nonionic surfactants, CrEL and polysorbate 80, respectively, which form micelles in solution, solubilizing PTX and DTX in the hydrophobic core of the micelles. However, the use of CrEL in Taxol is associated with life threatening hypersensitivity reactions characterized by dyspnea, hypotension, angioedema, and generalized urticaria (Szebeni et al., 1998). This excipient is also incompatible with the poly(vinyl chloride) infusion sets and causes the leaching of the plasticizers such as di(2-ethylhexyl) phthalate (Kim et al., 2005). Thus, the development of a PTX formulation that could eliminate the use of CrEL has been one of the factors driving the search for new delivery approaches for this drug.  In addition to an improved toxicity profile, nanoparticulate taxane formulations have been investigated with the goal of altering the pharmacokinetics, biodistribution and tumor uptake of the nanoparticulate drug payload (Gaucher et al., 2010). There are several reports demonstrating increased circulation lifetime and enhanced tumor uptake, likely due to a passive   144 targeting mechanism and the EPR effect in mouse tumor models for PTX loaded nanoparticles (Cheng et al., 2007; Kim et al., 2011; Shenoy et al., 2005). On the other hand, some groups, including ours have shown that upon IV administration of taxane loaded nanoparticles, there is rapid dissociation and elimination of the drug payload, resulting in no advantage over the commercial formulation with respect to prolonged circulation and passive tumor targeting (Burt et al., 1999; Kim et al., 2001; Le Garrec et al., 2004; Lee et al., 2011b; Letchford and Burt, 2012).  Another very significant problem associated with the effective delivery of taxanes to cancer cells is that PTX and DTX are subject to efflux transport by Pgp which is a transmembrane protein present in resistant cells that reduces intracellular drug accumulation and therefore prevents these drugs from reaching pharmacologically active concentrations within the cells (Geney et al., 2002; Shirakawa et al., 1999; Stordal et al., 2012). Pgp is overexpressed in MDR cancer cells and hence is a significant barrier to successful cancer treatment (Gottesman et al., 2002).   One of the early micellar nanoparticulate formulations for PTX developed by Burt and coworkers was based on the amphiphilic diblock copolymer, MePEG-b-PDLLA and in vitro and in vivo studies showed controlled drug release and enhanced efficacy of PTX loaded MePEG-b-PDLLA nanoparticles compared to Taxol® in P388 leukemia tumour bearing mice (Zhang et al., 1997; Burt et al., 1999). We also began investigating polymeric micelles based on amphiphilic diblock copolymers with varying core-forming hydrophobic blocks, PCL being selected since it is biocompatible, biodegradable and approved by the FDA for biomedical applications (Nair and Laurencin, 2007). Solubilization of several hydrophobic drugs, including PTX, by a series of MePEG-b-PCL diblock copolymers with varying MePEG and PCL block lengths showed that the amount of drug solubilized increased with increasing PCL block length and was also   145 dependent on the compatibility between the drug and hydrophobic core-forming block (Letchford et al., 2008). We also demonstrated that there is a shift in the physicochemical properties and hence, the performance of MePEG-b-PCL nanoparticles as drug delivery systems as the molecular weight of the hydrophobic block increases (Letchford et al., 2009). Finally, an important finding by our group was that a low molecular weight diblock copolymer of MePEG-b-PCL, termed MePEG17-b-PCL5, was an effective modulator of Pgp function and increased the intracellular accumulation of Pgp substrates in MDR cell lines (Zastre et al., 2002; Zastre et al., 2004; Zastre et al., 2007; Wan et al., 2013; Elamanchili et al., 2009). Hence, the selection of MePEG-b-PCL for the work described in chapters 2 and 3 of this thesis was based on the fact that we could combine the functions of both a nanoparticle delivery system for taxanes, with variable physicochemical properties, and a Pgp inhibitor using the same amphiphilic excipient, MePEG-b-PCL. Three compositions of MePEG-b-PCL with constant MePEG block length and increasing PCL block lengths were used to fabricate taxane loaded nanoparticles: MePEG114–b-PCL19, MePEG114–b-PCL104 and MePEG114–b-PCL200. The cell line, MDCKII-MDR1 was used as a Pgp overexpressing model to evaluate the cellular accumulation and cytotoxicity of PTX and DTX loaded nanoparticles, with or without exposure or co-encapsulation of the Pgp inhibitor, MePEG17-PCL5. This cell line is not a cancer cell line but is derived from a canine kidney source and was selected since it is an established model for studying Pgp. For the purposes of this work, it was critical to use a cell line that specifically overexpressed Pgp to avoid the complexities of multiple drug transporters.  In chapter 2, two MePEG-b-PCL diblock copolymers with very different hydrophobic block lengths were synthesized and studied: the fluid-core, micelle-forming short hydrophobic block, MePEG114-b-PCL19 and the higher core microviscosity, nanosphere-forming long hydrophobic block, MePEG114-b-PCL104. PTX-loaded micelles and nanospheres were evaluated   146 for their cytotoxicity, cellular polymer uptake, and drug accumulation in drug-sensitive MDCKII and Pgp overexpressing MDCKII-MDR1 cell lines. Comparing PTX loaded micelles and PTX loaded nanospheres following incubations in MDCKII and MDCKII-MDR1 cells, there was no difference in PTX or copolymer uptake into cells, but PTX loaded micelles were more cytotoxic than PTX loaded nanospheres in both cell lines, likely due to a faster release of PTX from the micelles. The cytotoxicity of PTX loaded MePEG114-b-PCL19 micelles in MDCKII and MDCKII-MDR1 cells was similar to the Taxol® formulation (Figure S4 a) which was also reported to release drug rapidly. After 3 day incubation, free PTX (Figure S5 a) in the absence of drug carrier was more cytotoxic than PTX loaded nanospheres and micelles, and Taxol® formulation at the concentrations tested in MDCKII and MDCKII-MDR1 cells. This finding is not surprising as free PTX can diffuse freely into cells. However due to the aqueous solubility of PTX, the highest concentration of PTX can only be achieved at 10 μg/ml in the culture medium. We evaluated cell binding and uptake of rhodamine labeled MePEG114-b-PCL19 micelles and MePEG114-b-PCL104 nanospheres in both MDCKII and MDCKII-MDR1 by fluorescence spectroscopy. Rhodamine was covalently linked to MePEG-b-PCL diblock copolymers using tetramethylrhodamine-5-carbonyl azide (TMRCA, MW 455 Da). This conjugation had no significant effect on the molecular weight of MePEG-b-PCL copolymers as it represented only a small fraction of the overall molecular weight of MePEG114-b-PCL19 (MW 7143 Da) and MePEG114-b-PCL104 (MW 16651 Da). Using the fluorescent-labeled diblock copolymers and radiolabeled PTX, we found that the intracellular accumulation of both PTX and copolymers were similar for both nanoparticles (Figure 2.6 and 2.7), further suggesting that the difference in cytotoxicity might be due to the different drug release profiles.  The mechanism by which these MePEG-b-PCL nanoparticles are internalized by MDCKII cells has not been established. However, it has been shown that other nanoparticles,   147 such as liposomes, micelles, dendrimers, and quantum dots are generally internalized by cells via endocytosis mechanisms, typically, caveolae- and clathrin- mediated endocytosis (Sahay et al., 2010a). Since MDCKII cells are devoid of caveolae on the apical surface, it is believed that the nanoparticles used in this work are likely internalized via clathrin-mediated endocytosis (Lahtinen et al., 2003; Vogel et al., 1998).  The cytotoxicity of PTX loaded micelles and PTX loaded nanospheres was significantly enhanced in MDR cells when co-administered with MePEG17-b-PCL5 diblock copolymer (Figures 2.9 and 2.10). The MePEG17-b-PCL5 concentration at 0.05% (w/w) was most effective in inhibiting Pgp, resulting in the maximum cytotoxicity of PTX loaded micelles and PTX loaded nanospheres in MDCKII-MDR1 cells, but also providing that MePEG17-b-PCL5 was present continuously throughout the 3 day incubation. The MDCKII-MDR1 cells treated with PTX loaded MePEG114-b-PCL104 nanospheres along with subsequent incubation with 0.05% MePEG17-b-PCL5 were shown to have a higher IC50 when compared to PTX loaded MePEG114-b-PCL19 micelles (Figure 2.11), yet delivered approximately the same amount of PTX as the PTX loaded MePEG114-b-PCL19 micelles, suggesting that a greater fraction of intracellular PTX was still associated with intracellular MePEG114-b-PCL104 nanospheres and less free intracellular PTX was available to bind to β-tubulin, the intracellular target of PTX. Hence, these data indicate that drug release rate from intracellular nanoparticles appears to be an important factor in determining cytotoxicity in a drug resistant cell line. Using PTX loaded MePEG114-b-PCL19 micelles and MePEG114-b-PCL104 nanospheres, our group investigated the stability characteristics in the presence of blood components and the pharmacokinetic properties following IV administration into healthy mice. MePEG114-b-PCL104 nanospheres showed better retention of their drug payload when incubated in vitro in human plasma as compared to MePEG114-b-PCL19 micelles (Letchford and Burt, 2012). Using H3-  148 labeled copolymers and C14-labeled PTX, it was shown that upon injection into the tail vein of mice, MePEG114-b-PCL104 nanospheres circulated for longer than MePEG114-b-PCL19 micelles (Letchford and Burt, 2012). However, PTX was rapidly eliminated from the blood regardless of the formulation and no difference in the pharmacokinetics of the drug encapsulated in micelles and nanospheres was observed (Letchford and Burt, 2012).  Literature reports suggest that factors leading to enhanced circulation time for nanoparticulates in the blood include, the steric hindrance properties of the hydrophilic corona and larger taxane loaded nanoparticles composed of high molecular weight hydrophobic cores (Owens and Peppas, 2006) (Gaucher et al., 2010). Accordingly, we synthesized another block copolymer, keeping an identical hydrophilic corona size of 114 MePEG repeat units but almost doubling the hydrophobic PCL block length to 200 repeat units. We hypothesized that the larger volume core of MePEG114-b-PCL200 would create more stable taxane loaded nanoparticles, retaining the drug within the core over a more extended time period following IV administration. In chapter 3, we describe the synthesis of MePEG114-b-PCL200 through a ring opening polymerization of -caprolactone using the hydroxyl group of MePEG as the macroinitiator and stannous octoate as a catalyst. Due to the poor water solubility of MePEG114-b-PCL200, a nanoprecipitation and dialysis technique was used to form nanoparticles, referred to as PCL200 nanospheres. Based on the data obtained in chapter 2 showing the importance of maintaining a continuous level of MePEG17-b-PCL5 in contact with PTX loaded MePEG114-b-PCL104 nanospheres in order to achieve maximum cytotoxicity effects in MDR cells, our next approach to nanoparticulate formulation development was to investigate the potential of developing nanoparticles of co-encapsulated MePEG114-b-PCL200 and the Pgp inhibitor, MePEG17-b-PCL5. We hypothesized that release of MePEG17-b-PCL5 from the taxane loaded, co-encapsulated MePEG114-b-PCL200 and MePEG17-b-PCL5 nanoparticles following cellular uptake would inhibit   149 Pgp mediated drug efflux and enhance cytotoxicity in MDR cells. Using emulsification methods, we were able to produce novel “mixed MW PCL200/PCL5 nanoparticles” in which MePEG17-b-PCL5 diblock copolymer was added in a 1 to 1 (PCL200:PCL5) weight ratio. In Chapter 3, we evaluated the physicochemical properties and performance characteristics of mixed MW PCL200/PCL5 nanoparticles as PTX or DTX carriers. Size, surface charge, yield and physical stability of mixed MW PCL200/PCL5 nanoparticles were dependent on the amount of MePEG17-b-PCL5 diblock copolymer used to produce the nanoparticles (Figure 3.1). An increase in the amount of MePEG17-b-PCL5 diblock copolymer used in the manufacture of mixed MW PCL200/PCL5 nanoparticles led to a decrease in the amount of precipitated polymer and a decrease in the diameter of nanoparticles, allowing for the formation of more stable nanoparticles. The addition of MePEG17-b-PCL5 diblock copolymer during emulsification, dramatically decreased the zeta potential of the nanoparticles likely due to the increased association of MePEG17-b-PCL5 diblock copolymers with the nanoparticles, thus shielding the negative charges of carboxylate groups on PCL (Figure 3.1B). Using a dialysis method, it was shown that MePEG17-b-PCL5 was associated with the nanoparticles and was released in a controlled manner. These MePEG17-b-PCL5 release data, along with the change in the physicochemical properties of the mixed MW PCL200/PCL5 nanoparticles, provide some evidence that MePEG17-b-PCL5 was co-encapsulated in the nanoparticles and not just surface-associated. Both PCL200 and mixed MW PCL200/PCL5 nanoparticles were capable of solubilizing relatively high concentrations of PTX and DTX (up to 500 µg/mL and 1300 µg/mL, respectively), likely resulting from a larger hydrophobic core, providing a larger cargo space for loading the drug (Figure 3.2). The solubilization differences between the two drugs might be attributed to the slightly lower hydrophobicity of DTX and more favorable interactions between the hydrophobic core and the drug (Carstens et al., 2008). PTX and DTX were released from the   150 nanoparticles in a controlled manner with a small burst phase of release, followed by a slower sustained release over 14 days (Figure 3.3). DTX loaded nanoparticles had a faster release rate in comparison to PTX loaded nanoparticles, likely due to the 7-fold difference in aqueous solubility between the two drugs and potentially more DTX at the MePEG/PCL interface as compared to PTX. A similar difference in drug release was also reported by Mugabe et al, who compared the release of PTX and DTX from hyperbranched polyglycerol nanoparticles (Mugabe et al., 2011a). The cytotoxicities of the PTX or DTX loaded PCL200 and mixed MW PCL200/PCL5 nanoparticles on MDCKII and MDCKII-MDR1 were evaluated using the MTS assay. Due to the co-encapsulation of MePEG17-b-PCL5 diblock copolymer, these drug loaded mixed MW PCL200/PCL5 nanoparticles dramatically reduced the viability of Pgp overexpressing MDCKII-MDR1 cells since MePEG17-b-PCL5 was present at sufficient concentration over a 3 day incubation to inhibit Pgp mediated drug efflux. As discussed in chapter 3, we believe our novel nanoparticulate formulation may possess significant potential advantages over other drug loaded co-encapsulated Pgp inhibitor nanoparticulate formulations such as mixed Pluronic® micellar systems.  Pharmacokinetic studies in mice using dual radiolabeled drug and copolymer demonstrated prolonged circulation of the MePEG114-b-PCL200 polymer present in the mixed MW PCL200/PCL5 nanospheres; however, the encapsulated drug was rapidly eliminated from the blood. The organ biodistribution after 24 hours showed that very little drug was found in the organs (Figure 3.9A). It is likely that upon injection the drug partitioned out of the nanoparticles and was quickly metabolized and eliminated from the body. Despite the fact that the hydrophobic block length of the MePEG114-b-PCL200 diblock copolymer was increased significantly from previous pharmacokinetic studies with PTX loaded MePEG114-b-PCL19 and MePEG114-b-PCL104 nanoparticles, nevertheless, this increased core volume did not translate to   151 improved in vivo retention of taxanes in the nanoparticles or prolonged drug circulation upon IV administration. However, in light of the prolonged circulation of mixed MW PCL200/PCL5 nanoparticles in the blood, the nanoparticles hold out some promise for being able to deliver the co-encapsulated Pgp inhibitor, MePEG17-b-PCL5 to tumor sites through passive targeting and the EPR effect, despite taxanes not being retained within the nanoparticles. We speculate that once at the tumor site, and provided effective concentrations of PTX or DTX are present in the tumor tissue, the MePEG17-b-PCL5 might be released at sufficient concentrations and over a prolonged period to inhibit Pgp mediated efflux of taxanes from cancer cells to enhance cytotoxicity and potentially be used as a strategy to overcome MDR.  Non-invasive ultrasonic technology has been widely used in diagnostic medicine for many years. Recent work using ultrasound triggered anti-cancer drug delivery from polymeric micelles composed of Pluronic® P-105 has demonstrated the potential of applying ultrasound for controlled and targeted tumor chemotherapy (Husseini and Pitt, 2009; Rapoport, 2004). Ultrasound may be focused on a tumor site very precisely, to trigger the controlled release of drug from micelles and/or to perturb cell membranes to enhance the intracellular uptake of nanoparticulate drugs (Husseini and Pitt, 2009; Rapoport, 2004). The proposed mechanisms for ultrasound effects on delivery of nanoparticulate drugs have been discussed and include enhancement of drug release from nanoparticles at the cell membranes, increased rate of endocytosis of nanoparticles, and enhancement of membrane permeability by the formation of transient membrane pores (Chapter 1.5.3).  Previous work by our group carried out ultrasound optimization studies to develop the ultrasonic irradiation regime used in this thesis. A single 10 second burst of high frequency (4 MHz), high intensity (32 W/cm2) ultrasound was produced by a custom-made device and can be directed to a single well at a time, of a 48 well plate (Figure 1.7 A) (Jackson et al., 2011).   152 Although the ultrasound power intensity used in this study was high compared to values used in other reports, it is likely that some of the ultrasound energy was absorbed by the polystyrene cell culture plate, the water in the water bath (beneath the cell culture plate) and the aqueous medium in the cell culture well. In addition, the power cycle used in the ultrasound experiment was 12.5 ms followed by a 37.5 ms rest interval, which reduce the likelihood of heat generation by the ultrasound transducer. This ultrasound irradiation regime produced no significant temperature effects and there was no background cell lysis when used with a burst mode protocol which allowed for intervals between ultrasound waves. To avoid the problem of the ultrasound irradiation to one well inadvertently irradiating cells in an adjacent well, the adjacent well was kept empty. Thus, alternating wells only were seeded with cells.  In the previous work using the same ultrasound regime (4 MHz, 32 W/cm2, 10 second burst), we compared the effects of ultrasound on the cellular uptake of free PTX and PTX loaded MePEG-b-PDLLA micelles (Jackson et al., 2011). We found little to no enhancement of uptake of free PTX whereas the intracellular accumulation of micellar PTX was significantly enhanced in HUVEC and PC3 cells (Jackson et al., 2011). In the studies described in chapter 4, we used PTX loaded in MePEG-b-PDLLA micelles to remain consistent with our previous work. In addition, this micellar system has been well-characterized by our group. In Chapter 4, we evaluated the effect of this ultrasound regime on the cellular accumulation of PTX loaded in copolymer micelles of MePEG-b-PDLLA in both drug sensitive (MDCKII and MCF-7) and Pgp overexpressing (MDCKII-MDR1 and NCI-ADR) cell lines. There was no effect of ultrasound on the hydrodynamic diameters of micelles or the release of FRET pairs indicating that the integrity of micelles was unaffected by ultrasound (Figures 4.2 and 4.3). Compared to no ultrasound treatment, the drug release study showed that PTX was released from the micelles in a controlled manner and no difference in drug release was observed for micelles irradiated by ultrasound. The   153 drug accumulation study showed that there was a 2-fold increase in intracellular PTX for all ultrasound treated drug sensitive cell lines and their respective drug resistant counterparts compared to no ultrasound (Figure 4.4). Significant decreases in drug efflux rates were observed for both drug sensitive and resistant cell lines receiving ultrasound (Figure 4.5). The enhanced accumulation and retention of PTX by ultrasound resulted in greater cytotoxicity in both MDCKII and MDCKII-MDR1 cell lines as indicated by the MTS assay (Figure 4.6). These data suggest that ultrasound may facilitate the uptake of micellar PTX into cells allowing greater retention of the drug in both Pgp and non-Pgp expressing cells. Although the cytotoxicity of micellar PTX in MDCKII-MDR1 cells was enhanced significantly, there was no evidence to suggest that Pgp function was inhibited by ultrasound irradiation. In order to further enhance the cytotoxicity of micellar PTX by ultrasound treatment, it may be critical to co-administer Pgp inhibitors to allow for the inhibition of Pgp function. It is unclear whether the use of ultrasound in vivo for enhancing nanoparticulate drug uptake into tumors would require a formulation that demonstrates prolonged blood circulation of drug. However, it is possible that ultrasound might enhance intracellular accumulation of plasma protein bound drugs as lots of drugs are transport by plasma proteins in the blood (Karmali and Simberg, 2011). The mechanical index of the ultrasound parameters described in this work was determined to be less than 0.5, below the threshold of inertial cavitation (Apfel and Holland, 1991). Therefore, the ultrasound enhanced intracellular micellar drug accumulation was unlikely due to the inertial cavitation effect. It was speculated that stable cavitation may be induced by the ultrasound irradiation to create oscillatory bubbles in the medium and increase the permeability of cell membranes to the PTX loaded micelles by forming transient pores on the membranes. Schlicher et al. reported that the stable cavitation effect could induce pore formation in cell membranes with diameters of 28 nm and lifetimes of less than 1 minute (Schlicher et al., 2006).   154 Guy and coworkers reported similar findings showing that the transient pores in cell membranes induced by ultrasound allowed the passage of nanoparticles with diameters up to 37 nm (Mehier-Humbert et al., 2005). The ultrasound data presented in chapter 4, demonstrate that ultrasound irradiation enhanced the intracellular uptake and retention of nanoparticulate PTX and resulted in a greater drug cytotoxicity in MDR cells. These results, taken together with the data from chapter 3, suggest that mixed MW PCL200/PCL5 nanoparticles together with an ultrasound irradiation regime such as that presented in this work, may be a potentially exciting drug delivery strategy for future treatment of MDR cancer cells.     5.2 Summary and conclusions Major findings in this work were:  1. Nanoparticles composed of MePEG-b-PCL with shorter  hydrophobic PCL block lengths, produced “fluid-like core” PTX loaded MePEG114-b-PCL19 micelles whereas nanoparticles made from intermediate length PCL chains produced “solid-like core” PTX loaded MePEG114-b-PCL104 nanospheres. The MePEG114-b-PCL19 fluid core micelles released PTX more rapidly than the solid core MePEG114-b-PCL104 nanospheres. However, the MePEG114-b-PCL104 nanospheres were superior in terms of PTX loading as compared to the MePEG114-b-PCL19 micelles. There was no difference in PTX or copolymer uptake into MDCKII and MDCKII-MDR1 cells following incubation with either PTX loaded micelles or nanospheres. However, PTX loaded micelles were more cytotoxic than PTX loaded nanospheres in both cell lines, likely due to a faster release profile.   155 2. Co-administration of the known Pgp inhibitor, MePEG17-b-PCL5 with PTX loaded MePEG114-b-PCL19 micelles or MePEG114-b-PCL104 nanospheres significantly increased drug cytotoxicity in MDR cells. 3. Both PTX and DTX loaded mixed MW PCL200/PCL5 nanoparticles showed increased cytotoxicity in MDR cells as compared to PTX and DTX loaded in nanoparticles of PCL200 (in the absence of MePEG17-b-PCL5) due to the Pgp inhibition by the controlled release of MePEG17-b-PCL5 from the mixed nanoparticles. 4. Ultrasound irradiation was investigated as a potential strategy to enhance the cytotoxicity of MePEG-b-PDLLA nanoparticulate PTX formulation in MDR cells. This effect probably arose from increased micellar uptake into cells so that drug might avoid membrane interactions with Pgp and might further be sequestered intracellularly in micelles, again avoiding Pgp efflux. 5.3 Suggestions for future work The drug accumulation, efflux, and cytotoxicity studies on the combination of mixed MW PCL200/PCL5 nanoparticles with ultrasound irradiation for treating MDR cells should be investigated as mentioned in Chapter 4. This might open an opportunity for an alternative approach to overcome MDR based on a combined strategy of the inhibition of Pgp by MePEG17-b-PCL5 and the enhancement of drug uptake by ultrasound irradiation.  In our nanoparticle uptake study, fluorescent-labeled nanoparticles were observed in the cells. However, the mechanisms of internalization of these nanoparticles are still unknown. It may be possible to study the mechanisms using different types of endocytosis inhibitors on cells that have defined endocytosis pathways. Although the preliminary pharmacokinetic data showed that the drugs were rapidly partitioned out of the mixed MW PCL200/PCL5 nanoparticles upon injection, a prolonged   156 circulation of copolymers in the blood was observed, suggesting a potential application of the nanoparticles for delivery of MePEG17-b-PCL5 to the cancer cells at least. Given that the MePEG-b-PCL diblock copolymers are very well tolerated in mice, it would be possible to deliver high concentrations of blank mixed MW PCL200/PCL5 nanoparticles to MDR tumor bearing mice in combination with taxanes to evaluate the effectiveness of the mixed MW nanoparticles as a Pgp inhibitor carrier. MDCKII and MDCKII-MDR1 cells were used in this study because this pair is a well-established model for the study of Pgp substrates/inhibitors. A further cell line that provides a Pgp overexpressing clone and a wild type line would be ideal to differentiate the Pgp inhibition effects by MePEG17-b-PCL5. 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The MDCKII (white) and MDCKII-MDR1 (black) cells were subjected to ultrasound irradiation for up to three daily exposures (4MHz, 32 W/cm2, 10 second exposure). The cell proliferation was measured by MTS assay. All data points represent mean ± SEM, (n=3).   0 exposure1 exposure2 exposures3 exposures050100MDCKIIMDCKII-MDR1% Cell Viability  185  Figure S2 The effect of ultrasound irradiation on cell proliferation. The MDCKII (white) and MDCKII-MDR1 (black) cells were exposed to ultrasound irradiation for 10, 20 and 30 seconds in burst mode (4MHz, 32 W/cm2). The cell proliferation was measured by MTS assay. All data points represent mean ± SEM, (n=3).   Figure S3 The effect of ultrasound irradiation on the intracellular uptake of free PTX (1 μg/mL) in MDCKII (black) and MDCKII-MDR1 (white) cells. Cells were incubated with free PTX for 90 mins prior to ultrasound irradiation (4MHz, 32 W/cm2, 10-second exposure). Intracellular PTX levels were measured using liquid scintillation counting. All data points represent mean ± SEM, (n=3).  0 10 20 300255075100MDCKIIMDCKII-MDR1Ultrasound Exposure (s)% Cell Viability+US -US0200004000060 00MDCKIIMDCKII-MDRPTX (DPM)  186  Figure S4 The cytotoxicity of Taxol® (a) and Taxotere® (b) in MDCKII and MDCKII-MDR1 cells. Cells were incubated at different concentrations of drug for 3 days followed by MTS assay. All data points represent mean ± SEM, (n=3). 0.1 1 10 100050100MDCKIIMDCKII-MDR1[PTX] (mg/ml)% Cell Viability 0.1 1 10 100050100MDCKIIMDCKII-MDR1[DTX]  (mg/ml)% Cell Viability a)b)  187  Figure S5 The cytotoxicity of free PTX (a) and free DTX (b) in MDCKII and MDCKII-MDR1 cells. Cells were incubated at different concentrations of drug for 3 days followed by MTS assay. All data points represent mean ± SEM, (n=3).  0.01 0.1 1 10 100020406080100MDCKIIMDCKII-MDR1[PTX] (mg/ml)% Cell Viability0.01 0.1 1 10 100020406080100MDCKIIMDCKII-MDR1[DTX] (mg/ml)% Cell Viabilitya)b)

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