Functional Anchoring Lipids for Drug Delivery Carrier Fabrication and Cell

FUNCTIONAL ANCHORING LIPIDS FOR

DRUG DELIVERY CARRIER FABRICATION AND

CELL SURFACE RE-ENGINEERING APPLICATIONS

PRATIMA VABBILISETTY

Bachelor of Pharmacy Acharya Nagarjuna University, India March 2006

Master of Science in Chemistry Youngstown State University, Ohio, USA December 2008

Submitted in partial fulfillment of requirements for the degree of

DOCTOR OF PHILOSOPHY IN CLINICAL-BIOANALYTICAL CHEMISTRY

at the

CLEVELAND STATE UNIVERSITY

December 2014

We hereby approve this thesis/dissertation for

Pratima Vabbilisetty Candidate for the Doctoral degree for the Department of Chemistry and Cleveland State University’s College of Graduate Studies by

Xue-Long Sun, Ph.D. Dissertation Committee Chairperson Professor, Department of Chemistry, 12/10/2014

Aimin Zhou, Ph.D. Professor, Department of Chemistry, 12/11/2014

Bin Su, Ph.D. Associate Professor, Department of Chemistry, 12/10/2014

John F. Turner II, Ph.D. Associate Professor, Department of Chemistry, 12/10/2014

Petru S. Fodor, Ph.D. Associate Professor, Department of Physics, 12/10/2014

Date of Defense December 10th , 2014

Dedicated to my parents, my sister and my best pal Anand for their unconditional love

and constant support

ACKNOWLEDGEMENTS

This dissertation would not have been possible without the help and support of many people. I would like to express my immense gratitude to all of them for their support and encouragement throughout my academic career.

Firstly, I would like to thank my advisor Dr. Xue-Long Sun for giving me the opportunity to join his lab and work on this research. His expertise, understanding, patience, constant support and encouragement have been undoubtedly valuable both academically and personally. This thesis would not have been completed successfully without his guidance and effort and motivation.

I would like to thank my committee members Dr. Aimin Zhou, Dr. Bin Su, Dr.

John F Turner and Dr. Petru S Fodor for their valuable advice, support and assistance towards the progress of my work. Special thanks to Dr. Jerry Mundell for his support and understanding throughout the years at Cleveland State University.

I would like to thank all my lab members Dr. Yong Ma, Dr. Rui Jiang, Dr. Libo

Wang, Dr. Jinshan Tang, Dr. Srinivas Chalagalla, Dr. Satya Nandana Narla, Valentinas

Gruzdys, Lin Wang, Joshua Whited, Dan Wang, Jayasree Kakarla, Poornima

Pinnamaneni, Vinay Kodithyala, Kishan Yalamarthy, Swetha Patel and Rachel Troyan for their understanding and helpful nature throughout my graduate study. I thank them for their friendship and the memorable moments spent together in our lab. Particularly, I am very thankful to Dr. Hailong Zhang, Dr. Jacob Weingart and Dr. Huan Nie for their help

in taking my research to the next step. I would also like to thank Dr. Evgeny Ozhegov for helping with the confocal microscopy. Additionally, I would like to give special regards to my friends Leela Immadisetty and Sangeetha Chari for their moral support in all aspects of my life. I would like to specially thank Vinay Kodithyala, Snigdha

Chennamaneni, Venkat Bobba, Ramakrishna Reddy Voggu, Ravali Alagandula and

Prasad Gobburi for the wonderful memories spent together at Cleveland State University.

I would also like to thank the Department of Chemistry, its faculty and staff and the Graduate School, Cleveland State University for providing funding throughout my

Doctoral program.

Last but not least, I would not have completed and achieved this work without the unwavering love and encouragement of my family. I am very grateful to my parents Mrs.

Ramani, Mr. Krishna Rao and my sister Deepthi for all the support and advice they have given me throughout my life. I am especially grateful to my best friend Anand, who has always been there with me in every walk of my life.

FUNCTIONAL ANCHORING LIPIDS FOR

DRUG DELIVERY CARRIER FABRICATION AND CELL SURFACE

RE-ENGINEERINGAPPLICATIONS

PRATIMA VABBILISETTY

ABSTRACT

For decades, lipid vesicular bodies such as liposomes have been widely used and explored as biomimetic models of cell membranes and as drug/gene delivery carrier systems. Similarly, micellar iron oxide nanoparticles have also been investigated as potential MRI agents as well as drug delivery carrier systems. Cell surface carbohydrate- protein interactions allow them to serve as markers for recognition of many molecular and cellular activities thereby, are exploited as attractive molecules for surface modification of nanocarrier systems with purpose for tissues specific targeting and biocompatibility. In addition, the cell lipid membrane serves as an important platform for occurrence of many biological processes that are governed and guided by cell surface receptors. Introduction of chemoselective functional groups, via bio-orthogonal conjugation strategies, at the cell surface facilitates many cellular modifications and paves path for novel and potential biomedical applications. Anchoring lipids are needed for liposome surface functionalization with ligands of interest and play important roles in ligand grafting density, liposomes stability and biological activity. On the other hand,

anchoring lipids are also needed for cell surface re-engineering by lipid fusion approach and have high impact for ligand insertion efficiency and biological activity. Overall, in this dissertation study, functional anchoring lipids for glyco-functionalized carrier systems and for efficient cell surface re-engineering applications were systematically investigated, respectively.

Firstly, investigation of the synthesis of glyco-functionalized liposome systems based on phosphatidylethonalamine (PE) and cholesterol (Chol) anchoring lipids, prepared by post chemically selective functionalization via Staudinger ligation were carried out. The effect of anchor lipids on the stability, encapsulation and releasing capacity of the glycosylated liposomes were investigated by dynamic light scattering (DLS) technique and by entrapping 5, 6-carboxyfluorescein (CF) dye and monitoring the fluorescence leakage, respectively. Overall, the Chol-anchored liposomes showed faster releasing rate than

DSPE-anchored liposomes. This could be due to the increase in rigidity of the lipid membrane upon inclusion of Chol, thereby, leading to fast leakage of liposomes.

Second, the potential effects of phospholipid (PE) and cholesterol (Chol)-based anchor lipids on cell surface re-engineering via copper free click chemistry were assessed with

RAW 264.7 cells as model. The confocal microscopy and flow cytometry results indicated the successful incorporation of biotinylated Chol-based anchor lipids after specific streptavidin-FITC binding onto the cell surface. Higher fluorescence intensities from the cell membrane were observed for Chol-based anchor lipids when compared to

DSPE as anchoring lipid. Furthermore, cytotoxicity of the synthesized biotinylated anchor lipids on the RAW 264.7 cells was assessed by MTT assay. The MTT assay

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results further confirmed that cell surface re-engineering via lipid anchoring approach strategy has very little or negligible amount of cytotoxicity on the cell viability. Thus, this study suggests the possible use of these lipids for potential cell surface re-engineering applications.

In addition, synthesis of lipid coated iron oxide nanoparticles via dual solvent exchange approach and their glyco-functionalization via Staudinger ligation were investigated and characterized by FT-IR and TEM techniques. The stability of iron oxide nanoparticles with varying compositions of lipid anchors was evaluated by dynamic light scattering technique.

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TABLE OF CONTENTS

ABSTRACT ...... vi

LIST OF TABLES ...... xvii

LIST OF FIGURES ...... xviii

I. INTRODUCTION ...... 1

1.1 Drug delivery...... 1

1.1.1 Classical drug delivery ...... 2

1.1.2 Targeted drug delivery ...... 2

1.2 Nanocarriers as drug delivery systems ...... 5

1.2.1 Liposomes as drug delivery systems...... 8

1.2.1.1 Preparation of liposomes...... 11

1.2.1.1.1 Thin-film hydration and extrusion ...... 12

1.2.1.1.2 Ethanol injection ...... 13

1.2.1.1.3 Reverse-phase evaporation ...... 13

1.2.2 Iron oxide nanoparticles as drug delivery systems ...... 14

1.2.2.1 Synthesis of Iron oxide nanoparticles ...... 18

1.2.2.1.1 Co-precipitation method ...... 19

1.2.2.1.2 Microemulsion method ...... 20

1.2.2.1.3 Thermal decomposition method ...... 21

1.2.2.2 Lipid coating techniques ...... 22

1.2.2.2.1 Thin-film hydration and sonication ...... 23

1.2.2.2.2 Dual solvent exchange method ...... 26

1.3 Carbohydrate mediated targeted delivery ...... 27

1.3.1 Cell surface carbohydrates ...... 28

1.3.2 Glyco-functionalized liposomes ...... 30

1.3.2.1 Preparation of glyco-functionalized liposomes ...... 30

1.3.2.1.1 Direct liposome formulation approach ...... 31

1.3.2.1.2 Post liposome functionalization approach...... 32

1.3.3 Glyco-functionalized iron oxide nanoparticles ...... 33

1.4 Targeting bioconjugation strategies for drug delivery systems ...... 34

1.4.1 Conventional bioconjugation strategies ...... 35

1.4.2. Bioorthogonal conjugation strategies ...... 39

1.4.2.1 Staudinger ligation ...... 41

1.4.2.2 Click chemistry ...... 41

1.5 Cell surface re-engineering applications ...... 42

1.6 Research design and rationale ...... 44

1.7 References ...... 47

II. LIPOSOME SURFACE FUNCTIONALIZATION BASED ON DIFFERENT

ANCHORING LIPIDS VIA STAUDINGER LIGATION ...... 74

2.1 Introduction ...... 74

2.2 Experimental ...... 78

2.2.1 Materials and methods ...... 78

2.2.2 Syntheses of DSPE–PEG2000–TP and Cholesterol-PEG2000–TP anchor lipids for

chemoselective surface functionalization via Staudinger ligation ...... 79

2.2.2.1 Synthesis of cholesterol–PEG2000-triphenylphosphine (Chol-PEG2000-TP)80

2.2.2.2. Synthesis of DSPE-PEG2000-triphenylphosphine (DSPE-PEG2000–TP) ... 81

2.2.3 Determination of critical micelle concentration (CMC) of the anchor lipids .... 82

2.2.4 Liposome preparation ...... 82

2.2.5 Conjugation of lactose onto liposomes surface ...... 83

2.2.6 Determination of concentration of lactose on the liposome surface ...... 83

2.2.7 Characterization of specific binding between lactose on the liposome surface

and lectin ...... 85

2.2.8 5, 6-Carboxyfluorescein (5, 6-CF) encapsulation efficiency ...... 85

2.2.9 5, 6-CF dye leakage assay ...... 87

2.3 Results and discussion ...... 87

2.3.1 Syntheses and characterization of DSPE–PEG2000–TP and Cholesterol-PEG2000–

TP anchor lipids ...... 88

2.3.2 Characterization and evaluation of the stability of glyco-functionalized

liposomes via dynamic light scattering technique (DLS) and fluorescence

spectroscopy ...... 90

2.3.3 Measurement of critical micelle concentration (CMC) of anchor lipids before

and after glyco-functionalization ...... 94

2.3.4 Quantification of lactose on the surface of liposome ...... 99

2.3.5 Accessibility of surface coated lactose molecules on liposome formulations via

lectin binding assay ...... 99

2.4 Conclusion ...... 102

2.5 References ...... 103

III. CELL SURFACE RE-ENGINEERING VIA EFFICIENT LIPID ANCHORING

APPROACH ...... 107

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3.1 Introduction ...... 107

3.2 Experimental ...... 111

3.2.1 Materials and methods ...... 111

3.2.2 Syntheses of DSPE–PEG2000–DBCO and Cholesterol-PEG2000–DBCO anchor

lipids for chemoselective surface functionalization via copper free click chemistry 112

3.2.2.1 Synthesis of Chol-PEG2000–DBCO ...... 113

3.2.2.2 Synthesis of DSPE-PEG2000–DBCO ...... 113

3.2.3 Determination of critical micellar concentration (CMC) of the anchor lipids . 114

3.2.4 Cell culture ...... 115

3.2.5 Preparation of aqueous solutions of DSPE–PEG2000–DBCO and Cholesterol-

PEG2000–DBCO anchor lipids ...... 115

3.2.6 Cell surface re-engineering of RAW 264.7 cells with DSPE and CHOL based

anchor lipids via copper-free click chemistry ...... 116

3.2.6.1 Two-step method ...... 116

3.2.6.2 Direct/one-pot method ...... 116

3.2.6.2.1 Syntheses of lipid-biotin conjugate via copper-free click chemistry 116

3.2.6.2.2 Anchoring of biotinylated lipid conjugates into the cell membranes 117

3.2.7 Liposome preparation ...... 117

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3.2.8 Cell surface labeling of biotinylated lipid conjugates on RAW 264.7 cells with

streptavidin-FITC and imaging by confocal laser scanning microscopy ...... 118

3.2.9 Flow cytometry analysis of streptavidin-FITC labeled biotinylated–lipid

anchored cells...... 119

3.2.10 In vitro cytotoxic effect of the biotinylated lipid conjugates: MTT assay ..... 119

3.3 Results and discussion ...... 120

3.3.1 Syntheses of biotinylated anchor lipids via copper-free click chemistry for cell

surface re-engineering ...... 121

3.3.2 Determination of critical micelle concentration of Cholesterol-PEG2000-DBCO

and DSPE-PEG2000-DBCO anchor lipids...... 122

3.3.3 Incorporation efficiency of biotinylated lipid conjugates into the membrane of

cells ...... 124

3.3.4 Comparative biotinylation efficiencies of two-step and one-pot step on RAW

264.7 cells via copper-free click chemistry ...... 130

3.3.5 Rate of anchoring of biotinylated lipid conjugate (Chol-PEG2000-biotin) on the

cell membrane ...... 135

3.3.6 Rate of anchoring and incorporation efficiency of liposomal biotinylated lipid

conjugates into the cell membrane ...... 139

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3.3.7 Effect of aqueous solutions of biotinylated lipid conjugates and liposomes on

cell viability: MTT assay ...... 148

3.4 Conclusion ...... 154

3.5 References ...... 156

IV. BIOMIMETIC FUNCTIONALIZATION OF IRON OXIDE NANOPARTICLES 162

4.1 Introduction ...... 162

4.2 Experimental ...... 164

4.2.1 Materials and methods ...... 164

4.2.2 Synthesis of iron oxide nanoparticles via thermal decomposition method ...... 165

4.2.3 Lipid coating of iron oxide nanoparticles ...... 166

4.2.3.1 Thin-film hydration and sonication ...... 167

4.2.3.2 Dual solvent exchange method ...... 167

4.2.4. Glyco-functionalization of iron oxide nanoparticles via staudinger ligation .. 168

4.3 Results and discussion ...... 168

4.3.1 Synthesis of iron oxide nanoparticles via thermal decomposition method ...... 168

4.3.1.1 Characterization of iron oxide nanoparticles by FT-IR ...... 170

4.3.1.2 Characterization of iron oxide nanoparticles by TEM ...... 172

4.3.2 Lipid coating of iron oxide nanoparticles ...... 173

4.3.3 Glyco-functionalization of iron oxide nanoparticles via staudinger ligation ... 173

4.3.4 Stability evaluation of lipid coated and glyco-functionalized iron oxide

nanoparticles via DLS ...... 174

4.4 Conclusion ...... 177

4.5 References ...... 178

V. SUMMARY ...... 181

VI. FUTURE PERSPECTIVE ...... 184

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LIST OF TABLES

Table 1. Conventional bioconjugation strategies for nanoparticle surface functionalization...... 36

Table 2. Linker chemistry strategies for nanoparticle surface functionalization...... 38

Table 3. Liposomes with different anchoring lipids and their sizes determined by DLS. 92

Table 4. Critical micelle concentration (CMC) determination of anchor lipids

Cholesterol–PEG2000–TP (A) and DSPE-PEG2000-TP (B)...... 95

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LIST OF FIGURES

Figure 1. Schematic diagram for targeted drug delivery systems...... 4

Figure 2. Types of nanocarrier systems...... 7

Figure 3. Schematic representation of the structural composition of different types of liposomes...... 10

Figure 4. Graphic illustration of the structure of multifunctional iron oxide nanoparticles.

...... 17

Figure 5. Lipid coating techniques for iron oxide nanoparticles [1, 2] Traditional thin-film and hydration method. [3, 4] Dual exchange solvent method...... 25

Figure 6. Cell surface carbohydrates mediating biological processes and glyco- functionalized drug delivery systems...... 29

Figure 7. Bioorthogonal conjugation strategies for surface functionalization...... 40

Figure 8. Liposome surface glyco-functionalization based on two kinds of anchoring lipids via Staudinger ligation...... 77

Figure 9. Synthesis of anchoring lipids Chol-PEG2000-TP and DSPE-PEG2000-TP...... 79

Figure 10. Standard curve for lactose assay...... 84

Figure 11. Encapsulation efficiency of 5, 6-carboxyfluorescein dye...... 86

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Figure 12. NMR spectra of Chol-PEG2000-TP (A) and DSPE-PEG2000-TP (B) in CDCl3. 89

Figure 13. DLS monitoring of liposomes before and after glyco-functionalization: Chol-

PEG2000-TP anchored liposome (A) and DSPE-PEG2000-TP anchored liposome (B) and their stabilities monitored with DLS...... 93

Figure 14. Critical micelle concentration (CMC) determination of anchor lipids

Cholesterol–PEG2000–TP (A) and DSPE-PEG2000-TP (B)...... 95

Figure 15. 5, 6-CF releasing kinetics of lactosylated liposomes with anchoring lipid in different ratios: Cholesterol–PEG2000-TP (A) and DSPE–PEG2000-TP (B)...... 98

Figure 16. DLS monitoring of agglutination due to multivalent lectin binding of lactosylated liposomes with anchor lipid of Cholesterol–PEG2000-TP (A) and DSPE–

PEG2000-TP (B)...... 101

Figure 17. Schematic illustration of cell surface re-engineering via Chol-PEG2000–DBCO and DSPE-PEG2000-DBCO lipid anchoring approach...... 110

Figure 18. Schemes for the syntheses of Cholesterol–PEG2000–DBCO and DSPE-

PEG2000–DBCO anchor lipids for cell surface re-engineering applications...... 112

Figure 19. Measurement of critical micelle concentration (CMC) of anchor lipids:

Cholesterol–PEG2000–DBCO (A) and DSPE-PEG2000-DBCO (B)...... 123

Figure 20. Confocal microscopy images of Raw 264.7 macrophage cells treated for 20 mins at 37 ºC, in PBS buffer, pH 7.4. Control 1 & Control 2 (panels A, B) showing no fluorescence signal...... 126

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Figure 21. Confocal microscopy images of biotinylated Chol-DBCO (panels A, C) and

DSPE-DBCO ( panels B, D) treated with Raw 264.7 macrophage cells with varying concentrations (5 µM, 10 µM) for 20 mins at 37 ºC, in PBS, pH 7.4...... 127

Figure 22. Flow cytometry images of biotinylated Chol (panels C, E) & DSPE anchor lipids (panels D, E) treated with Raw 264.7 macrophage cells with varying concentrations

(5 µM, 10 µM) for 20 mins at 37 ºC, in PBS, pH 7.4. Control cells (A, B)...... 129

Figure 23. Confocal microscopy images of Raw 264.7 macrophage cells treated for 20 mins at 37 ºC, in PBS buffer, pH 7.4. Control 1 & Control 2 (panels A, B) showing no fluorescence signal...... 132

Figure 24. One-pot biotinylated Chol (panels A, C) & DSPE ( panels B, D) lipid conjugates treated with Raw 264.7 cells with varying concentration (5, 10 µM ) for 20 mins at 37 ºC, in PBS, pH 7.4...... 133

Figure 25. One-pot biotinylated lipid conjugate treated RAW 264.7 cells with different concentrations (5 µM, 10 µM Chol-PEG2000–DBCO–C, E), (5 µM, 10 µM DSPE-

PEG2000–DBCO–D, F) for 20 mins at 37 ºC, PBS, pH 7.4. (Control cells A, B)...... 134

Figure 26. One-pot biotinylated cholesterol lipid conjugate (5 µM) treated RAW 264.7 cells) at different time points at 37 °C, PBS pH 7.4. (5 mins–A), (10 mins–B), (20 mins–

C)...... 136

Figure 27. One-pot biotinylated cholesterol lipid conjugate (10 µM) treated RAW 264.7 cells) at different time points at 37 °C, PBS pH 7.4. (5 mins–A), (10 mins–B), (20 mins–

C)...... 137 xx

Figure 28. One-pot biotinylated CHOL conjugates treated with RAW 264.7 cells at different concentrations (5, 10 µM) and at varying incubation periods at 37 °C, PBS pH

7.4 (panels A, B-5mins), (panels C, D-10 mins), (panels E, F–20 mins)...... 138

Figure 29. Confocal microscopy images of Raw 264.7 macrophage cells treated for 20 mins at 37 ºC, in PBS buffer, pH 7.4. Control 1 & Control 2 (panels A, B) showing no fluorescence signal...... 140

Figure 30. One-pot biotinylated liposomal cholesterol (A, B, C) lipid conjugate treated

RAW 264.7 cells with varying concentrations (5, 10, 25 µM) at 20 mins time point. ... 141

Figure 31. One-pot biotinylated liposomal cholesterol (A, B, C) lipid conjugate treated

RAW 264.7 cells with varying concentrations (5, 10, 25 µM) at 2 hr time point...... 142

Figure 32. One-pot biotinylated liposomal cholesterol (A, B, C) lipid conjugate treated

RAW 264.7 cells with varying concentrations (5, 10, 25 µM) at 4 hr time point...... 143

Figure 33. One-pot biotinylated liposomal DSPE (A, B, C) lipid conjugate treated RAW

264.7 cells with varying concentrations (5, 10, 25 µM) at 20 mins time point...... 144

Figure 34. One-pot biotinylated liposomal DSPE (A, B, C) lipid conjugate treated RAW

264.7 cells with varying concentrations (5, 10, 25 µM) at 2 hr time point...... 145

Figure 35. One-pot biotinylated liposomal cholesterol (A, B, C) lipid conjugate treated

RAW 264.7 cells with varying concentrations (5, 10, 25 µM) at 4 hr time point...... 146

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Figure 36. One-pot biotinylated liposomal cholesterol (C, E, G) & DSPE (D, F, H) lipid conjugate treated RAW 264.7 cells at varying concentrations (5, 10, 25 µM) for 4 hr time point. Control cells (A, B)...... 147

Figure 37. Viability of RAW 264.7 cells (A) during DBCO-lipid conjugate treatment at varying time points (20 min, 16 hr and 24 hr) at 37 °C in DMEM medium as indicated.

(B) after biotinylation of DBCO-lipid conjugate treatment for 20 min at 37 °C in DMEM medium...... 150

Figure 38. Viability of RAW 264.7 cells (A) during one-pot biotinylated lipid conjugate treatment at varying time points (20 mins, 16 hr and 24 hr) at 37 ºC, in DMEM medium as indicated. (B) after biotinylation of DBCO-lipid conjugate treatment for 20 min. .... 151

Figure 39. Viability of RAW 264.7 cells (A) during one-pot biotinylated liposomal conjugate treatment at varying time points (2 hr, 4 hr, 16 hr and 24 hr) at 37 °C, in

DMEM medium as indicated. (B) after treatment with biotinylated liposomes for 20 min at 37 °C...... 153

Figure 40. Schematic illustration of lipid coating of oleic acid capped iron oxide nanoparticles...... 166

Figure 41. Schematic illustration of synthesis of oleic acid capped iron oxide nanoparticles by thermal decomposition of iron oleate complex with high boiling points.

...... 169

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Figure 42. FT-IR spectra of oleic acid capped iron oxide nanoparticles (A) oleic acid (B) synthesized iron oxide nanoparticles (C) commercially synthesized oleic acid capped iron oxide nanoparticles...... 171

Figure 43. TEM images of (A) laboratory synthesized iron oxide nanoparticles (B) commercially synthesized iron oxide nanoparticles. (bar scale: 20 nm)...... 172

Figure 44. DLS spectra of (2:1 wt. ratio) (2% Chol-PEG2000–TP+DPPC): iron oxide nanoparticles. (A) thin-film hydration approach (B) dual solvent exchange approach (C) azido-lactose conjugated iron oxide nanoparticles...... 175

Figure 45. DLS spectra of (2:1 wt. ratio) (2% DSPE-PEG2000–TP+DPPC): iron oxide nanoparticles. (A) thin-film hydration approach (B) dual solvent exchange approach (C) azido-lactose conjugated iron oxide nanoparticles...... 176

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ABBREVIATIONS

5, 6-CF Carboxyfluorescein

CDCl3 Deuterated chloroform

CMC Critical Micelle concentration

TP Triphenylphosphine

DBCO Dibenzocyclooctyne

DLS Dynamic Light Scattering

DPPC 1, 2-Dipalmitoyl-sn-glycero-3-phosphocholine

DSPE 1, 2-distearoyl-sn-glycero-3-phosphoethanolamine

Et3N Triethylamine

EPR Enhanced permeability and retention effect

FITC Fluorescein isothiocyanate

NHS N-hydroxysuccinimide

PEG Poly (ethylene glycol)

SPION Super paramagnetic iron oxide nanoparticles

RES Reticuloendothelial system

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CHAPTER I

INTRODUCTION

1.1 Drug delivery

Drug delivery is the process of transporting and delivering of a pharmaceutical drug of interest to obtain a therapeutic effect in human and animals with diseased conditions. Drug delivery is an important factor for everyone since it has the capacity to deliver newer treatments for many diseases which are currently under investigation and also to deliver the existing drugs more effectively with minimal side effects. The extensive use of different types of drug delivery systems has changed significantly in the past few decades. The rate of drug release and the site of action are controlled by the drug delivery systems. Therefore, the major goal of drug delivery systems is to deliver the drug to the site of action with minimal side effects and in a controlled manner. Many

physiological factors such as movement of drug through cells and tissues, their transport in the blood need to be considered for development of efficient drug delivery systems.

1.1.1 Classical drug delivery

Though there is advancement in the discovery of new drugs and development of newer strategies, systemic side effects [1] and low bioavailability are of major concern for classical drug delivery systems. This is due to the fact that most therapeutic drugs are water insoluble which results in poor water solubility and membrane permeability.

Furthermore, this type of classical or traditional drug delivery systems also leads to cytotoxicity to certain extent (they attack both primary target cells as well as the healthy cells too) and thus causing several side effects. Due to this, the quality of the patient’s life as well as the time of treatment is affected. Especially, due to the poor biological stability and bioavailability of certain biomacromolecules like peptides, proteins and nucleotides, it is much more challenging for the development of efficient and safe drug products [2].

Due to these reasons there arises a need for specific drug targeting and its delivery with reduced toxicity while maintaining the therapeutic effects [3]. In addition increased bioavailability and prevention from degradation of drug before reaching its target site is very necessary.

1.1.2 Targeted drug delivery

In general accumulation of drugs or nanocarriers at specific active sites occurs based on their own or depending upon the carrier system’s physicochemical properties. 2

Passive targeting of drugs is a result of enhanced permeability and retention (EPR effect)

[1] caused by the leaky vasculature of the tumor sites, further leading to enhanced accumulation of drug or drug carrier systems. Though the EPR effect leads to more than

50-fold increase in accumulation of nanoparticles in the tumor sites, passive targeting is typically less efficient [4]. On the other hand, active targeting of drugs or the nanocarrier systems can achieved by targeting to specific disease cells or tissues by employing suitable affinity ligands that have specificity [1, 5, 6].

In order to overcome these drawbacks extensive investigation of targeted drug delivery systems has been carried out in the past decades. Targeted drug delivery mainly involves in capturing or encapsulating the drug of interest in a suitable carrier system and by finally targeting the carrier system to a specific site of action with the help of specific targeting ligands on their surface [2]. The principle of targeted drug delivery systems depends mainly on the coordinating behavior of three components (Figure 1.)[2], wherein the targeting moiety recognizes and binds to target; the carrier entraps the drug, and most importantly the therapeutic drug which has therapeutic action at the specific site. Based on these properties, targeted drug delivery systems can play a significant role in increasing the drug or gene efficacy with minimal side effects to a large extent especially in cancer therapy [2]. In addition, simplification of the drug or gene administration methods and also can result in therapy cost reduction by reducing the amount of gene or drug required. Targeted drug or gene delivery can be achieved by using specific targeting ligands such as antibodies, carbohydrates, polypeptides and endogenous hormones.

Figure 1. Schematic diagram for targeted drug delivery systems. Reproduced from [2].

1.2 Nanocarriers as drug delivery systems

Owing to their size-dependent physicochemical and biological properties, most of the nanoparticles existing in the rapidly growing field of nanotechnology, have demonstrated remarkable potential for a variety of biomedical applications. In particular, the biological compatibilities and functions of nanoparticles as nanocarriers have been widely investigated for further increasing their potential in areas of biomedical application such as bioimaging, biosensing and drug/gene delivery [7, 8, 9, 10, 11]. Of the many nanoparticles explored so far, most of them possess large functional surfaces which can be used as cargo units for targeted delivery of many bioactive molecules of interest using targeting ligands including polypeptides, antibodies, carbohydrates, proteins, and molecules of endogenous origin [12]. The different types of nanocarriers of major biomedical importance are liposomes, magnetic nanoparticles, quantum dots, metal nanoparticles, silica nanoparticles, and polymeric nanoparticles (Figure 2) [13]. However, the nanoparticles alone cannot be directly used, since some of them pose certain harmful effects to the surrounding biological environment. In order to address these problems, effective surface functionalization of nanoparticles has been widely explored in the past decades and has demonstrated a key role in nanoparticle modification and development for many practical applications. For example, different types of ligands or coatings containing bulky hydrophobic molecules may be conjugated to nanomaterial surfaces to prevent agglomeration of the nanoparticle core, while surfaces for use in aqueous environments can be coated with water-soluble polymers such as poly (ethylene glycol)

(PEG) to enhance solubility and biocompatibility [14, 15] of the carrier systems. Certain types of ligands can also be conjugated to the nanoparticle surface to serve as “tags” for monitoring the molecular interaction properties that can further be used in drug targeting and bio-imaging applications [16].

Figure 2. Types of nanocarrier systems. Modified and reproduced from [13].

Additionally, in order to determine the properties of nanocarrier itself, some ligands can be attached to its surface. Particularly lipid-coated nanocarriers have been proven successful in the delivery of many chemotherapeutic agents and are currently used in many cancer therapies [17]. The main factors that need to be taken into consideration for appropriate carriers for drug delivery systems include drug incorporation and release properties, biocompatibility with surrounding environments, stability and shelf life of carrier systems, their targeting and biodistribution properties and functionality of the carrier system itself. Most importantly, the effects and clearance of residual material after drug delivery should be taken into consideration as well [3].

1.2.1 Liposomes as drug delivery systems

For decades, lipid vesicular bodies such as liposomes have been widely used and explored as biomimetic models of cell membranes and as drug/gene delivery carrier systems. Liposomes are self-assembling lipid bilayers that comprises of an outer lipid bilayer enclosing an inner aqueous pool [18]. They have the capability of encapsulating both hydrophobic as well as hydrophilic drugs. Liposome encapsulation technique has been demonstrated to enhance their compatibility with the biological milieu in vitro and in vivo. Liposomes have been have been extensively studied for the past decades and they serve as attractive carrier systems that can improve the therapeutic efficiency, stability along with increasing the pharmacokinetic properties of drugs while minimizing their side effects. Liposomes are also known for their biodegradability and non-toxic property.

Early traditional liposomes or also known as the first generation liposomes were mainly made up of natural phospholipids such as dipalmitoyl phosphatidylcholine

(DPPC, PC, egg PC), or monosialogangliosides. Though these liposomes have gained a lot of importance in basic science research applications [19], due to poor stability and shorter circulation times their application in pharmaceutical industry was limited.

Therefore, in order to overcome such drawbacks, many attempts such as inclusion of cholesterol in lipid bilayers for stable bilayer membranes [20]; preparation of different size ranges of liposomes to increase circulation time [21]; further improvement of drug encapsulation efficiency by insertion of negative or positive charged lipids [22] was investigated. However, still certain major issues pertaining to faster elimination and non- targeting properties of liposomes needed to be investigated further.

For this purpose, a stealth liposome approach was achieved by coating the surface of liposomes with a hydrophilic and biocompatible polymer such as poly ethylene glycol

(PEG) [23] (Figure 3.). The PEG polymer acts a steric barrier which further prevents opsonization of proteins and liposome aggregation [24]. Similarly, various other hydrophilic polymers such as chitosan [25] and polyvinyl alcohol (PVA) [26], have been employed to increase the half-time and non-toxicity of liposomes in vivo. In spite of their low immunogenicity, non-toxicity and biodegradability properties, the use of PEG has been considered as the most efficient coating for liposome protection and increased circulation time.

Surface functionalization of such carrier systems further facilitates for enormous potential applications, such as enhanced stability, bioactive liposomal conjugates, and targeted drug, gene and imaging agent delivery [27].

Figure 3. Schematic representation of the structural composition of different types of liposomes. Reproduced from [23]

Additionally, surface modifications and change in lipid compositions can allow liposomes for use in various biological applications. Anchoring lipids are necessary for grafting ligands of interest and also they play vital roles in ligands grafting density, liposome stability, and liposome chemical and physical characteristics as well. Targeted liposomes not only significantly increase the local drug administration in to the cells of interest, but also have shown less or negligible systemic side effects [28, 29]. Many types of targeting ligands such as antibodies [30], peptides [31], carbohydrates [32], growth factors and certain receptors have been extensively investigated for conjugation to liposomes making them potential targeted delivery systems [33]. Among them, carbohydrates have been widely used for liposome surface modification (glycoliposomes) for variety of biomedical research and applications.

1.2.1.1 Preparation of liposomes

Liposomes with varying sizes and bilayers can be prepared by employing different types of methods. Generally the liposomes are classified as unilamellar vesicles

(ULVs, 25 nm-1µM), multilamellar vesicles (MLVs, 0.1-15 µM), large unilamellar vesicles (LUVs, 100 nm–1 µM) and small unilamellar vesicles (SUVs, 25-50 nm). The most common preparation methods include thin-film hydration, reverse-phase evaporation, polyol dilution, ethanol injection, freeze-thaw, double emulsion, detergent removal and high-pressure homogenization. [34, 35, 36, 37]. These types of methods typically result in the formation of LUVs and MLVs. Further, mechanical dispersion

methods such as freeze-thawing of liposomes, sonication, and extrusion can be employed to obtain unilamellar or small unilamellar vesicles. [38].

1.2.1.1.1 Thin-film hydration and extrusion

This method involves the dispersion or solubilization of lipids in volatile organic solvents, particularly in chloroform, methanol or ether. Briefly, the organic solvents consisting of dissolved lipids are evaporated under reduced pressure by means of a rotary evaporator and the lipids are formed as a thin film on the wall of the round bottom flask.

Further, hydration of the deposited lipids can be achieved with an aqueous buffer at a temperature above the transition phase of the lipids. Therefore, formation of stable and favorable rigid structure of liposomes is mainly dependent on the phase transition temperature of the lipids. The usual limitations associated with this type of preparation of liposomes are low encapsulation efficiency and difficulty in nano scale production of liposomes. Additional processes like sonication or extrusion [38, 39] through different sizes of polycarbonate membranes can be used in order to obtain SUVs or ULVs. In comparison to reverse-phase and ethanol injection methods, results obtained by many researchers suggest that liposomes prepared by the thin-film hydration method have shown to have better quality and stability. [40]

1.2.1.1.2 Ethanol injection

Briefly, in this method, the lipids dissolved in ethanol are rapidly injected into an aqueous buffer solution, wherein spontaneous formation of SUVs occurs. By this method, the sizes of the liposomes formed can be increased by increasing the lipid concentrations.

[41]. Certain chemical or physical treatment of lipids can be avoided by this liposome preparation method. However, this method usually results in the production of low concentration of vesicles and moreover it needs an additional step for removal of ethanol.

[36].

1.2.1.1.3 Reverse-phase evaporation

This method briefly involves the formation of liposomes from water-in-oil emulsions of aqueous buffers and phospholipids surrounded in an excess organic phase environment. [34]. Similar to the thin-film hydration technique, initially the phospholipids are dissolved in organic solvents, followed by thin film formation by evaporation of solvents. Further, followed by re-suspension of the thin-film is in diethyl ether and addition of water. The above formed suspension is subjected to sonication for a brief period of time resulting in a homogeneous emulsion. The remaining organic solvents are removed by rotary evaporation under reduced pressure, forming a viscous gel like dispersion representing the formation of LUVs. Though, this method can allow for the encapsulation of larger macromolecules with higher encapsulation efficiency, the major disadvantage is the exposure of the encapsulated material to organic solvents and

to sonication processes, wherein denaturation of certain sensitive molecules such as proteins or DNA may occur. [41].

1.2.2 Iron oxide nanoparticles as drug delivery systems

Magnetic nanoparticles of iron oxides, that exhibit magnetic moments in the vicinity of an external magnetic field, have attracted increasing interest and have been widely explored in the life sciences [42]. In view of the fact that the magnetic nanoparticles obey the Coulomb’s law, they can be guided to a specific target site by means of an external magnetic field. This unique property of magnetic nanoparticles makes them applicable in the transportation and delivery of molecular markers, various drugs and can also facilitate in biological purifications [42, 43]. Fe3O4 nanoparticles are the most promising type of magnetic nanoparticles and have already been approved by

FDA (i.e. Feridex I.V.®) for usage in liver imaging [44]. The chemical and physical properties of the iron oxide nanoparticles play an important role in their usage. By controlling and adjusting the particle core size, particle shape, bio-distribution and magnetic properties, one can meet specific parameters necessary for a variety of applications [45]. Therefore, suitable techniques must be chosen for the synthesis of magnetic iron oxide nanoparticles. There are many synthetic techniques available for the production of magnetic iron oxide nanoparticles. The most commonly utilized techniques are (i) co-precipitation of iron salts [46, 47], (ii) thermal-decomposition of iron precursors

[48, 49, 50, 51, 52] and (iii) micro-emulsion formation [53, 54].

Although there have been many significant developments in the synthesis of magnetic nanoparticles, maintaining the stability of these particles for a long time without agglomeration or precipitation is an important issue. Surfactants or polymers are often employed to passivate the surface of the nanoparticles during or after the synthesis to avoid agglomeration. In addition, for applications in biomedicine, it is necessary to functionalize nanoparticles with appropriate biocompatible coatings because iron oxide nanoparticles on their own, can pose certain harmful effects to the surrounding biological environment. This is due to the large surface to volume ratios and hydrophobic interactions between the unmodified bare iron oxide nanoparticles that leads to aggregation of magnetic nanoparticles forming larger agglomerates, opsonization, and, if not rapidly cleared by the reticulo-endothelial system (RES), inflammation and potentially cellular damage [55]. Therefore, it is necessary to surface coat the iron oxide nanoparticles with organic or inorganic monolayers in order to reduce their toxicity and to further stabilize the nanoparticles by preventing aggregation. The monolayer coating acts as a barrier between the inner iron oxide core and the surrounding environments.

They also govern factors like solubility, reactivity, interactions with targeting biomolecules, and also determine the biological function of the nanoparticles. The introduction of different functional groups/linkers onto the surface of the nanoparticles

(Figure 4.) [56] can enable the conjugation of different biomolecules such as antibodies, carbohydrates, peptides, enzymes, etc., making them applicable for many biomedical applications [57] such as magnetic resonance imaging (MRI) [58, 59], drug delivery [10,

60], hyperthermia [61], in vivo monitoring of tumor cell growth, and cell labeling [62].

For example, Hultman et al. demonstrated an application of immunotargeted

superparamagnetic iron oxide nanoparticles (ITSIONs), with in vivo magnetic resonance diagnostic and potential drug delivery capability for kidney disease [63].

Figure 4. Graphic illustration of the structure of multifunctional iron oxide nanoparticles.

Reproduced from [56]

1.2.2.1 Synthesis of Iron oxide nanoparticles

Over the past years, the synthesis of iron oxide nanoparticles has been studied intensively in order to make them suitable for many biomedical applications such as targeted drug delivery, contrast agents in magnetic resonance imaging (MRI), cell separation and hyperthermia. The tailoring of the monodisperse size of iron oxide nanoparticles is very important since, the properties of these nanoparticles mainly rely upon their dimensions. In general, most of these applications necessitate that the nanoparticles are within a size of 100 nm, having a homogeneous and narrow particle size distribution along with high magnetization values. In addition, suitable surface coating of magnetic iron oxide nanoparticles further enhances their biocompatibility and non-toxicity thereby, allowing for targeted delivery of biomolecules with particle localization in a specific area. [46, 47]. Various methods involving physical methods such as mechanical grinding and biomineralization processes along with many chemical methods have been developed and established to synthesize magnetic iron oxide nanoparticles. Chemical methods such as co-precipitation method, microemulsion, hydrothermal synthesis, sol-gel synthesis and thermal decomposition methods have been used to synthesize iron oxide nanoparticles. Of these co-precipitation methods, microemulsion and thermal decomposition have been widely employed for synthesis of iron oxide nanoparticles.

1.2.2.1.1 Co-precipitation method

The co-precipitation method is one of the simplest and the most widely employed method for synthesis of iron-oxide based MRI contrast agents. Iron oxide nanoparticles are usually formed by reaction between stoichiometric mixture of ferrous and ferric salts

(1:2 ratio) in an aqueous environment, at a pH 8-14 and in a non-oxidizing oxygen atmosphere. However, the formation of magnetite (Fe3O4) is not stable for long and is susceptible to oxidation and gets converted to maghemite (γFe3O4). For this purpose, different types of polymers or small molecules such as citric acid, tartaric acid and phosphoryl choline are sometimes used as precipitating or stabilizing agents for achieving colloidal stability. For example, dextran-coated ferromagnetic colloids were synthesized by the precipitation method wherein, the polymer coating was achieved after precipitation step [64]. On the other hand, Cox et al. have reported an iron-dextran complex formation in 1965, in which the iron oxide nanoparticles were synthesized directly in the presence of polymers [65].

Generally, the co-precipitation method is based on a complex approach, which involves complicated hydrolysis equilibria of ferric and ferrous ions. Moreover many parameters such as pH, ratio and concentration of iron salts, reaction conditions and temperature play an important role in governing the shape and size distributions of the synthesized iron oxide nanoparticles. In spite of these limitations, co-precipitation technique is well employed when large amounts of nanoparticles are required.

1.2.2.1.2 Microemulsion method

Microemulsion method is mainly useful in synthesizing iron oxide nanoparticles with controlled particle size and distributions. This technique mainly depends on the occurrences of chemical reactions in constrained environments namely, aqueous microdroplets in oil and amphoteric surfactants to form reverse micellar structures in non-polar solvents. The size of the nanoparticles can be tailored by altering the size of aqueous droplet core, reactions conditions, concentration of metal and base employed.

Microemulsions of iron oxide nanoparticles can be produced by dispersion of nanodroplets in aqueous phase by the surfactant molecules assembled in a continuous hydrocarbon phase [66]. Numerous types of surfactants such as cationic, anionic, or non- ionic can be employed in preparation of microemulsions of iron oxide nanoparticles.

Some of the commonly used surfactants include sodium dodecylsulfate (SDS), cetyltrimethylammonium bromide (CTAB), sodium bis(2-ethylhexylsulfosuccinate)

(AOT), TritonX-100 and Brij-97.

Though, this method can be used to synthesize size controlled iron oxide nanoparticles, the major disadvantage lies in dealing with complicated procedures for removal of surfactants employed for emulsification of the immiscible systems. In addition, when compared to the co-precipitation method the yield of nanoparticles synthesized is much lower.

1.2.2.1.3 Thermal decomposition method

High quality iron oxide nanoparticles can be synthesized via the thermal decomposition method. This method is achieved by pyrolysis of metal salts, metal- surfactant complexes and some organometallic compounds. High-temperature decomposition of certain iron organic precursors such as Fe (Cup)3, Fe (CO)5, or Fe

(acac)3 in the presence of organic solvents and surfactants leads to the formation of monodisperse iron oxide nanoparticles with a very narrow size variation thereby eliminating further size selective processes [67]. For the first time in 2002, Sun et al. have reported the synthesis of iron oxide nanoparticles via thermal decomposition method which resulted in monodisperse sizes below 20 nm. Briefly, thermal decomposition of Fe

(acac)3 dissolved in phenyl ether was carried out in the presence of 1, 2 hexadecanediol, oleic acid, and oleylamine. They have further studied that higher temperatures would result in larger particle formation providing scope for synthesis of different sizes of nanoparticles [22, 68].

In order to synthesize aqueous soluble iron oxide nanoparticles, use of non-polar solvents such as 2-pyrrolidone was employed [69]. Use of 2-pyrrolidone facilitates thermal decomposition reactions with organometallic precursors such as Fe (acac)3. On the other hand use of non-toxic iron chloride salts as precursors has been investigated by many researchers [70]. Though these methods produce water dispersible magnetite nanoparticles, their solubility at neutral pH is of concern. In order to resolve this issue, pyrolysis of Fe (acac)3 dissolved in 2-pyrrolidone was carried out and in the presence of monocarboxyl –terminated poly(ethylene glycol) (MPEG-COOH). PEG modified Fe3O4

nanoparticles have exhibited excellent solubility in waster as well as in physiological solutions. Therefore, the above mentioned methods can serve as excellent methods for producing water soluble and biocompatible iron oxide nanoparticles that can be applicable for detection of many diseases.

1.2.2.2 Lipid coating techniques

Lipids such as phospholipids, glycolipids, and cholesterol are naturally occurring amphiphillic molecules that constitute the major structural elements of biological membranes. Many phospholipid-based membrane mimetic systems, such as liposomes, have been widely developed for biomedical applications. A liposome encapsulation technique, in which iron oxide nanoparticles, quantum dots, silica and polystyrene nanoparticles are encapsulated into liposomes, have been investigated for a variety of different applications [71]. On the other hand, lipid membrane as a micellular shell has been employed to nanoparticle surface functionalization for various biomedical applications. In general, hydrophobic-ligand stabilized metal cored nanoparticles such as iron oxide, quantum dots, and gold, can be entrapped in a micellular shell of lipids- poly(ethylene glycol) (PEG) derivatives. The bulky hydrophillic PEG molecule acts as a steric barrier and prevents the adsorption of the plasma proteins and uptake by the macrophages, thus facilitating longer circulation times of nanoparticles with enhanced overall biocompatibility and biodistribution times of nanoparticles [55, 72, 73, 74, 75]. In addition, incorporation of lipid-PEG provides the nanoparticle system drug encapsulation capacity, making it applicable for therapeutic/imaging purposes. Depending on the type

of application, lipid-PEG derivatives are frequently mixed with different ratios of other lipids such as DOPC, DOPS, DOTAP, DOPE, DPPC or DPPE to form a membrane mimetic system. In some cases, the PEG end chain can be modified with certain targeting moieties such as antibodies, drug molecules, and other biological entities, dependent on the desired application [48, 61, 76, 77]. So far, two major methods have been developed for membrane-mimetic surface functionalization of magnetic nanoparticles.

1.2.2.2.1 Thin-film hydration and sonication

First, a thin-film hydration and sonication process was reported by Bao and co- workers [58], in which amphiphillic lipids self-assemble on the hydrophobic ligand- coated iron oxide nanoparticles surface in aqueous conditions. As shown in (Figure 5)

[78], In general, a mixture of lipids is added to the hydrophobic iron oxide nanoparticles in chloroform or other organic solvent to allow even mixing. Later, the organic solvent is completely removed by either rotary evaporation or evaporation under an inert gas to afford a thin film of lipids and nanoparticles. This thin film mixture is then hydrated with distilled water and subjected to sonication at 60 °C for 20 min to ensure the removal of any residual organic solvent, if present. After sonication the lipid-coated iron oxide nanoparticles are collected by either centrifugation or by means of magnetic field attraction. Finally, the obtained pellet via centrifugation or the attracted lipid coated nanoparticles are re-suspended in distilled water and are further filtered through a 0.22

µm filter and stored at 4 °C in argon gas [58, 76, 79]. The main advantages of this technique is the easiness of lab production under mild conditions, use of biocompatible

amphiphillic lipids, and the probability to obtain high encapsulation and loading capacity for lipophillic drugs.

Figure 5. Lipid coating techniques for iron oxide nanoparticles [1, 2] Traditional thin- film and hydration method. [3, 4] Dual exchange solvent method. Reproduced from [78].

1.2.2.2.2 Dual solvent exchange method

Recently, Bao and co-workers developed a solvent exchange method (Figure 5.)

[78], to obtain a very stable DSPE-PEG coated magnetic nanoparticle [78, 80]. Briefly, iron oxide nanoparticles (in toluene) were mixed with DSPE-PEG in chloroform (1:1 ratio). To the solution, DMSO was gradually added. Later, the toluene and chloroform were completely vaporized under vacuum and therein DMSO was substituted by water using an ultra-filtration centrifugal device with a polyethersulfone membrane. This technique is advantageous when compared with the traditional thin film hydration and sonication technique as it does not require extensive sonication and heating steps and also helps in preventing aggregation of the nanoparticles.

Recently, lipid-functionalized nanoparticles have been investigated for contrast- enhanced MRI and molecular imaging [81]. For example, enhanced colloidal stability and high r2 relaxivity was observed for water-dispersible ferromagnetic iron oxide nanocubes (WFIONs) when coated with lipid membrane [58]. Thereby, allowing much scope for the magnetic nanoparticles to play crucial roles in the early diagnosis and detection of tumor metastasis [58].

Another promising application of magnetic iron oxide nanoparticles is the site– specific delivery of both hydrophobic and hydrophilic drugs and genes [82]. Many anticancer drugs have been shown to exhibit nonspecific cytotoxicities that limit their therapeutic potential. Therefore, surface modified iron oxide nanoparticles have been investigated as drug carriers for efficient drug delivering to the pathological site and limiting the negative systemic toxicity effects. Most likely, a pharmaceutical drug of interest can be encapsulated by or entrapped in the iron oxide nanoparticles that could be

driven to the target organ by means of an external magnetic field where they can be released. Prasad and co-workers reported the magnetophoretically guided drug delivery of light-activated photodynamic therapy using a lipid micelle-magnetic hybrid [10]. The nanocarrier showed excellent stability and activity and efficient cellular uptake confirmed via confocal laser scanning microscopy. Furthermore, the magnetic response of the nanocarriers was demonstrated by their magnetically directed delivery to tumor cells in vitro. These multifunctional nanocarriers opened many new doors in the development of much more effective drug delivery systems.

1.3 Carbohydrate mediated targeted delivery

“Glycotargeting” was first demonstrated in 1971 where carbohydrates were used as targeting ligands to target protein receptors expressed at the active sites

[83].Carbohydrate modified liposomes, polymers, iron-oxide nanoparticles and other nanocarriers have been widely explored in the past decades in order to deliver drugs or genes to specific cells. For example, very high density asialoglycoprotein receptors are expressed on the surface of hepatocytes making them attractive targets for targeting by galactose modified nanocarriers which have high specificity towards those receptors [84].

Similarly, one of the important characteristic sign of liver fibrosis is increased deposition of extracellular matrix which is a result of activated hepatic stellalte cells (HSC) [85, 86].

Targeted drug delivery of mannose-6-phosphate (M6P)-modified human serum albumin

(HSA) to activated HSC in rats with liver fibrosis was first demonstrated by Poelstra et

al. This approach further led to the discovery and development of improved methods for selective delivery of antifibrotic drugs [87, 88, 89, 90].

1.3.1 Cell surface carbohydrates

The cell surface comprises of bilayers of amphiphillic lipids and cholesterol along with glycolipids, glycoproteins and proteoglycans that are richly coated with carbohydrate molecules [91]. Cell surface carbohydrates serve as remarkable models for surface modification with carbohydrates owing to their biodegradability, low toxicity, and mainly due to their specific binding interactions with the receptors present at the targeted cell’s surface. Carbohydrates can lead to the recognition of many medicinally important physiological processes in which they are a part of. They have specific interactions with proteins, which play important roles in various processes such as fertilization, , inflammations and host-pathogen adhesion [92], cell-cell interaction [93,

94], tumor metastasis [95], molecular and cellular targeting as shown in (Figure 6), thereby serving as attractive molecules for surface functionalization of nanocarriers with purpose for tissues specific and biocompatibility. The strong interactions/attractions between the carbohydrates and their receptors can be significantly increased through the multivalent exhibition of carbohydrates on nanomaterials.

Figure 6. Cell surface carbohydrates mediating biological processes and glyco- functionalized drug delivery systems.

1.3.2 Glyco-functionalized liposomes

Cell surface carbohydrates have specific interactions with proteins, which play an important role in various biological recognition processes such as fertilization, metastasis, inflammations and host–pathogen adhesion [93], thereby they can serve as attractive molecules for surface modification of liposomes with purpose for specific biomedical applications [96]. For example, monosialoganglioside (GM1) can enhance circulation lifetimes of liposomes comparable to that observed for PEG [97]. Mannosylated liposomes revealed superior uptake by macrophages both in vitro and in vivo after intratracheal administration and show great potential for targeted antigen delivery for immnune therapy [98]. On the other hand, carbohydrate-decorated liposomes have been used as a multivalent platform of carbohydrates to inhibit carbohydrate-mediated cell adhesion. For example, sialic acid-decorated liposomes have showed strong inhibitory activity against influenza hemagglunitin and neuraminidase [99].

1.3.2.1 Preparation of glyco-functionalized liposomes

Many strategies for surface glyco-functionalization of liposomes have been reported in the past decades, of which two strategies/approaches have been commonly used. The first approach, named as direct liposome formulation approach, is to synthesize the glyco-lipid ligand first, followed by preparation of liposomes in combination with other principal lipids [32, 100, 101, 102, 103]. The second approach named as post- functionalization approach, mainly involves grafting the carbohydrate ligand onto the preformed liposomes via various bioconjugation chemistry [33, 104, 105, 106].

1.3.2.1.1 Direct liposome formulation approach

The direct liposome formulation approach is mostly suitable for conjugation with small ligands such as saccharides [107] and vitamins [108] that are cheap and available in larger quantities. Various neoglycolipids have been synthesized by either chemical or enzymatic strategies. Traditional NHS coupling reaction is often used for the synthesis of neoglycolipids, such as maltose-DSPE (M2-DSPE) and oligomaltose-DSPE (M15-DSPE) glycolipids, for preparation of oligosaccharide-coated liposomes in combination with

DSPC for use in potential drug delivery applications. [109]. BPC NeuAc-pegylated lipids for incorporation into doxorubicin loaded liposomes were synthesized by NHS based coupling reaction between NHS activated pegylated lipids with BPC NeuAc (Sialoside)

[110]. Reductive amination is another practical method to synthesize neoglycolipids for glyco-liposome preparation. Ikehara et al. synthesized the neoglycolipids (Man3-DPPE) by conjugation of DPPE with oligosaccharides (Mannotriose) via reductive amination, followed by preparation of the liposomes with these neoglycolipids by thin-film extrusion method [111, 112]. In addition, chemo-enzymatic strategies were used for the synthesis of neoglycolipids such as lactosylceramide mimetics. Harada et al. prepared glycoliposomes incorporated with (Lac-DPPA, Lac-DPPC, and Lac-DPPE) neoglycolipid conjugates, which were synthesized by reacting phospholipids and the respective disaccharide glycosides [113].

1.3.2.1.2 Post liposome functionalization approach

Though direct liposome formation approach has been widely used for glyco- liposome formation, one drawback is that some of the carbohydrate targeting ligands can inevitably face towards the interior aqueous compartment of the formed liposomes and thus become unavailable for the interaction with the respective target molecules. Many post liposome functionalization methods have been developed for glyco-liposome preparations. This approach allows for maximum surface functionalization of liposomes with carbohydrates, most importantly on the exterior part allowing for more efficient delivery of drugs/gene/antigens. The most well-known methods are mainly based on the coupling reactions between maleimide and thiol groups [114]. A study done by the Rice group focuses on the chemo-enzymatic strategy for synthesis of tyrosine-linked oligosaccharides, bearing terminal sialyl or GalNac-Lex along with vinyl sulfonate ending poly(ethyleneglycol) spacer arm. This enables for further coupling reactions between thiol bearing liposomes incorporated with PC, Chol, and phosphatidylthioethanol lipids [115]. Neo-mannosylated liposomal vesicles were constructed by covalent coupling reaction between preformed vesicles containing 4-(p- maleimidophenyl) butyryl phosphatidylethanolamine (MPB-PE) lipids and α-D- thiomannopyranoside residues bearing sulfhydryl group [116]. Chemoselective conjugation between carbohydrate ligands and preformed liposomes require mild aqueous conditions in order to prevent or minimize any liposomal disruptions in the process. Recent advances led to the development of Click chemistry and Staudinger ligation methods that have high specificity, chemoselectivity, and efficiency. Hassane et al. employed click chemistry to conjugate an unprotected α-D-mannosyl derivative

carrying an azide-armed spacer with the surface of liposomes comprising of a synthetic lipid bearing a terminal alkyne functional group [117]. We demonstrated a chemoselective glyco-functionalization of preformed liposomes via Staudinger ligation, in which azide-functionalized lactose molecules were conjugated onto liposomes incorporated with synthetic anchor lipids bearing triphenylphosphine moiety [118].

Though the above mentioned strategies have been widely used and are highly efficient, many factors need to be considered for glyco-functionalization of liposomes. The affinity of the glyco lipids towards their target is mainly based on the type of PEG linker employed which tailors the accessibility and availability of the surface bound carbohydrate ligand for many interactions [119].

1.3.3 Glyco-functionalized iron oxide nanoparticles

Glyco-functionalized nanoparticles have been shown to increase the specific binding interactions between certain glycans and lectins, especially for biosensing application purposes. In particular, owing to their highly rich and complex structural variations, the carbohydrate molecules are very well suited for encoding biological information. [120]. The structural properties such as high surface area and spherical shape of the iron oxide nanoparticles allows for attachment of multiple carbohydrate molecules, thereby leading to increased avidity with their respective carbohydrate receptors via multivalent interactions.[121]. For example, monosaccharides such as mannose have been used as targeting ligands for sensing proteins and bacteria. Liu et al. have developed a strategy for immobilization of unmodified monosaccharide onto the

surface of iron oxide nanoparticles. Glyco-functionalized iron oxide nanoparticles were obtained by covalent coupling of D-mannose onto iron oxide nanoparticles by CH insertion reaction of photochemically activated phosphate-functionalized perfluorophenylazide moieties. [122]. Based on these properties of iron oxide nanoparticles and carbohydrates El-Boubbou et al. have reported magnetic glyco- nanoparticle based nanosensor systems to detect and differentiate cancer cells as well as to quantitatively profile their individual carbohydrate binding capacities via magnetic resonance imaging.[123]. Similarly, lactobionic acid coated superparamagnetic iron oxide nanoparticles were synthesized in order to enhance the efficacy of probe binding to hepatocytes. [124]. Mannose coated iron oxide nanoparticles have been shown to exhibit potential properties for use as a macrophage-targeting MRI contrast agent. [125]. Various types of conjugation strategies such as NHS/DCC chemistry [124], click chemistry, amide coupling reactions[123] and chemo-enzymatic approaches[126] have been developed for preparation of glyco-functionalized iron oxide nanoparticles.

1.4 Targeting bioconjugation strategies for drug delivery systems

In the past several years the investigation and development of many targeting moieties/ligands has gained lot of importance. Surface modification of nanocarrier systems can be achieved by means of conjugation with targeting moieties such as carbohydrates, antibodies, peptides, small molecules, etc in order to provide for multifunctional nanoparticles. A variety of chemical strategies have been used to conjugate targeting ligands, contrast agents, fluorophores, therapeutic agents, diagnostic

agents to nanoparticles surfaces. The major role of the targeted bioconjugation techniques is to bind the targeting moiety to the surface of nanoparticles without any loss or change in its functionality upon its attachment [127].

1.4.1 Conventional bioconjugation strategies

This type of bioconjugation strategy usually involves in either direct conjugation of the nanoparticle surfaces with different functional moieties of interest or by the use of linker molecules to provide for conjugation of targeted moieties to the nanoparticle surfaces. The direct conjugation method usually is preferred for conjugation with fluorescence dyes, chelators, imaging agents or drugs.

Table 1. Conventional bioconjugation strategies for nanoparticle surface functionalization. Reproduced from [127].

In general, this strategy allows for surface functionalization of nanoparticles with maleimide, aldehyde, amine, or active hydrogen groups (Table 1) [128, 129, 130, 131,

132]. The coupling reaction between aldehyde and hydrazide functional group on the surface of liposomes was demonstrated by Chua et al. wherein the reaction between aldehyde group in the immunoglobin and hydrazide on the surface of liposome resulting in a stable amide bond formation allows for attachment of immunoglobulin onto liposome surface [133]. On the other hand, the conjugation of targeting moieties on the nanoparticle surface via the linker chemistry is more beneficial since it can control the binding orientation of targeting ligands. This targeting strategy can be used for conjugation of many antibodies, small molecules and peptides. The reaction between amine functionalized nanoparticles and sulfhydryl bearing biomolecules is one of the most commonly employed linker chemistry strategy used for bioconjugation. Linker molecules such as (N-succinimidyl iodoacetate) SIA, (succinimidyl-4-(N- maleimidomethyl) cyclo-hexane-1-carboxylate) SMCC, (N-succinimidyl-3-(2- pyridyldithio)-propionate) SPDP, or certain heterobifunctional PEG molecules (NHS-

PEG-MAL) have been extensively used for bioconjugation to surface of nanoparticles

(Table 2). In general, the heterobifunctional molecules include thiol-reactive iodoacetate, maleimide, pyridyldithio groups at one end and some amine reactive succinimydyl esters on the other end of the molecule. Successful attachment of various protein molecules such as wheat germ agglutinin, avidin, antimyosin antibody 2G4 and concavalin onto the surface of liposomes via a stable amide bond was demonstrated by Torchillin et al [134].

Table 2. Linker chemistry strategies for nanoparticle surface functionalization.

Reproduced from [127]

But, one of the major drawbacks of using heterofunctional ligands involves the possibility of cross-linking reactions. These reaction methods usually form covalent bonds between ligands and nanoparticles, due to which nanoparticle modifications before ligand attachment or purification between each step is required [135]. Similarly,

EDC/NHS linkers can be used to covalently bond carboxyl group-bearing nanoparticles to amine functional moieties via amide bond formation. For example, EDC/NHS linker strategy was used to synthesize hepatocyte-targeted LBA-immobilized SPIONs via an amide bond linkage between dopamine modified SPION bearing a primary amine and lactobionic acid [124]. However this strategy is more efficient for conjugation of molecules bearing a single amine group only, thereby leading to loss of functionality of targeting ligands having multiple amine groups.

1.4.2. Bioorthogonal conjugation strategies

So far, the most available conjugation techniques are less selective and nonspecific leading to poorer efficiency. Due to which the use of these methods is less reliable for bioconjugation to different nanocarrier systems. In the past few years, many biocompatible and bioorthogonal methods have been developed and investigated (Figure

7). When compared to the conventional conjugations methods, these bioorthogonal reactions [136] offer excellent biocompatibility and high specificity and selectivity in living systems.

Figure 7. Bioorthogonal conjugation strategies for surface functionalization.

1.4.2.1 Staudinger ligation

Recent studies suggest the successful use of Staudinger ligation in living systems without any physiological harm, leading to potential applications in cell surface functionalization. Staudinger ligation involves in the reaction between an azide functional group and a triphenylphosphine moiety that selectively react to form a stable amide bond

[137]. Most importantly, the reaction is known to occur at room temperature in aqueous environment and without the use of any catalyst. The reaction is said to occur at high yield and is compatible with unprotected functional groups involving a broad range of biomolecules of interest. In our lab we have demonstrated a highly efficient and chemically selective liposome surface functionalization with carbohydrate molecule via the Staudinger ligation method [118, 138]. Because of the high specificity, high selectivity and high yield along with biocompatible native reaction conditions, Staudinger ligation strategy can serve as an effective surface functionalization method for various nanocarrier systems.

1.4.2.2 Click chemistry

The 1, 3-dipolar cycloaddition also termed as the “click reaction” was developed by Sharpless et al. in 2001 [139]. This reaction involves the conjugation between azide and alkyne groups in the presence of Cu (I)-catalyst, resulting in the formation of a stable triazole ring. The reaction can be performed under aqueous conditions with high efficiency and is very biocompatible. Both the azide and alkyne groups are very inert and orthogonal to other chemical groups due to which this reaction results in high specificity

and selectivity. Moreover this feature of click chemistry provides for highly oriented linkages, thus making the reactions more suitable for conjugation of targeting ligands to the nanoparticles. Despite the attractive features, the use of Cu (I) catalyst can cause toxicity leading to neurological disorders, hepatitis or kidney diseases in vivo [140].

Alternatively, recent studies have led to the development and used of highly strained alkynes which prevent the need of Cu (I) catalysts in click chemistry reactions, wherein cyclooctynes react with azide nearing molecules forming a stable triazole moiety. This strategy is known as copper free click chemistry, which occurs at a faster rate due to the strain activation inherent to the cyclooctyne, making it a suitable method for in vivo labeling. Recently, in our lab Zhang et al have developed liposome surface functionalization via copper-free click chemistry, thereby investigating the site-specific liposomal conjugation of recombinant TM at C-terminus [141]. These features of copper free click chemistry reactions make it a potential method for cell surface labeling, imaging, drug or gene delivery via many modified nanoparticles like liposomes.

1.5 Cell surface re-engineering applications

The surface of cell membrane serves as a platform for the occurrence of many biologically significant events. The cell surface is decorated with many biomolecules that participate in various extracellular communications and recognition processes. Therefore, modifications made at cell surface can further enhance the functioning of the cell allowing for many potential therapeutic applications [142, 143, 144, 145]. For example, macrophages are known to play a key role in the defense mechanism of an organism

[146]. Phagocytosis of pathogens such as viruses and microbes is mainly achieved by phagocytic cells such as dendritic cells (DC’s) and macrophages. Monocytes exist as circulating cells in blood with typical half-life of 48 hrs. Upon entering the tissues they exist as tissue /resident macrophages [147, 148] which distribute mostly to, all organs of mammals [149]. They are the major cells of the innate immune system, which mainly involve in capturing and destruction of microbes/pathogens, antigen processing and its presentation at the cell surface, apoptotic cell clearance and tissue repair. Due to this characteristic feature of travelling to and accumulating at different pathological sites and its central role in inflammation, infection, cancer and other diseased conditions [150], macrophages can serve as important drug, gene and antigen delivery targets or as drug delivery vehicles to treat many diseases. Thus, further investigation of different strategies focusing at targeting the macrophages is highly desirable.

In recent years, significant progress has been achieved in introduction of various chemical species or biomolecules of interest to the cell membrane, thereby allowing a wide range of applications in biology, drug delivery, tissue engineering and grafting.

Different types of approaches such as metabolic engineering [151, 152], direct chemical or enzymatic modification [153, 154, 155] and membrane fusion [156] have been studied and investigated for cell surface re-engineering. For example, metabolic glyco engineering method was employed for the generation of unnatural sialic acids with azide groups on the surface of cells. This strategy provides for conjugation with dibenzylcyclooctyne-conjugated Cy5 (DBCO-Cy5) fluorophore via copper free click chemistry, which further helps in cell labeling and tracking [151].

Different types of targeting ligands can be introduced on to the surface of target cells, of which, lipid anchors have shown to have promising cell surface re-engineering applications. For the past years, liposomes are the most widely explored delivery systems employed for targeting to macrophages [110, 157]. Owing to their biocompatibility, low immunogenicity and cell specificity characteristics they can serve as good targeting systems. In general, macrophages express certain carbohydrate receptors which have affinity towards mannose. Ikhera et al. have demonstrated the delivery of anticancer drugs to metastatic sites through uptake of oligomannose-coated liposomes (OML’s) by macrophages [111] Similarly, liposomal antigen delivery was also reported by Chen et al. wherein liposomal nanoparticles decorated with specific high affinity glycan ligands of

Sialoadhesin/ CD 169 (Sn/CD 169) was employed to target macrophages [158] In addition, targeting efficiency of liposomes to macrophages can be further enhanced by altering the lipid composition and by introducing various targeting ligands such as proteins, peptides, glycolipids, polysaccharides and certain lectins.

1.6 Research design and rationale

Delivery of drugs to the target area with minimal side effects and with high therapeutic index is of major concern. For this purpose, efficient and targeted drug delivery systems are necessary. Various types of carriers systems such as liposomes and iron oxide nanoparticles have been extensively investigated for over a decade and have shown promising results.

Owing to their biocompatibility, liposomes have been widely employed as biomimetic models of cell membrane as well as potential carrier systems for delivery of many molecules such as drugs, antigens, vaccines or genes to the targeted area. Similarly, nanoparticles such as magnetic nanoparticles have been showing great potential to revolutionize biological imaging and drug delivery applications, but their clinical usages have been limited by difficulties in obtaining nanoparticles that are biocompatible and tissue specific. To address these problems, surface modification of nanoparticles has been widely explored. Among them, lipid coating, micelle encapsulation and liposome encapsulation have been demonstrated to enhance their compatibility with the biological milieu in vitro and in vivo. Anchoring lipids serve as biomimicking models, as they naturally constitute the major structural elements of the biological membrane.

Incorporation of lipid anchors for preparation of nanocarrier systems allows for biocompatibility and non-toxicity. Furthermore, anchoring lipids facilitate conjugation with targeting ligands on to surface of nanocarrier systems, thereby increasing the efficiency of targeted delivery. On the other hand, cell surface carbohydrates mediate many biologically important processes, hence, carbohydrate molecules can serve as attractive molecules for surface modification of different nanoparticles with purpose for tissues specificity and biocompatibility.

Therefore, based on the physical properties of liposomes and iron oxide nanoparticles, significant roles of anchoring lipids, carbohydrates and the efficiency of bioorthogonal conjugation techniques, in this study, we hypothesized that, preparation of glyco functionalized liposomes as well as iron oxide nanoparticles with different anchor

lipid compositions through bio orthogonal conjugation techniques provides an efficient strategy for development of targeted carrier systems.

In recent years, considerable advancement has been made in introducing many biologically important molecules to the cell surface, permitting a wide range of biomedical applications. Surface modifications made at the cell surface further enhances the cellular functions. In particular, since macrophages play significant roles in the body’s defense mechanism, surface modification of macrophages makes them ideal candidates as cellular vehicles carrying therapeutic molecules. Therefore, we hypothesized that surface re-engineering of macrophages with different types of lipid anchors and other biologically significant molecules via bioorthogonal conjugation approach, significantly offers newer engineering strategies for many applications in biology, tissue engineering and targeted delivery of various molecules.

1.7 References

[1] Z. Cheng, A. Al Zaki, J. Z. Hui, V. R. Muzykantov and A. Tsourkas,

"Multifunctional nanoparticles: cost versus benefit of adding targeting and imaging

capabilities," Science, vol. 338, no. 6109, pp. 903-910, 2012.

[2] H. Zhang, Y. Ma and X.-L. Sun, "Recent developments in carbohydrate-decorated

targeted drug/gene delivery," Medicinal Research Reviews, vol. 30, no. 2, pp. 270-

289, 2010.

[3] W. H. De Jong and P. J. Borm, "Drug delivery and nanoparticles: applications and

hazards," International Journal of Nanomedicine, vol. 3, no. 2, p. 133, 2008.

[4] D. Peer, J. M. Karp, S. Hong, O. C. Farokhzad, R. Margalit and R. Langer,

"Nanocarriers as an emerging platform for cancer therapy," Nature

Nanotechnology, vol. 2, no. 12, pp. 751-760, 2007.

[5] N. Kamaly, Z. Xiao, P. M. Valencia, A. F. Radovic-Moreno and O. C. Farokhzad,

"Targeted polymeric therapeutic nanoparticles: design, development and clinical

translation," Chemical Society Reviews, vol. 41, no. 7, pp. 2971-3010, 2012.

[6] E. A. Sykes, J. Chen, G. Zheng and W. C. W. Chan, "Investigating the impact of

nanoparticle size on active and passive tumor targeting efficiency.," ACS Nano,

vol. 8, no. 6, pp. 5696-5706, 2014.

[7] A. Senpan, S. D. Caruthers, I. Rhee, N. A. Mauro, D. Pan, G. Hu, M. J. Scott, R.

W. Fuhrhop, P. J. Gaffney, S. A. Wickline and G. M. Lanza, "Conquering the dark

side: colloidal iron oxide nanoparticles.," ACS Nano, vol. 3, no. 12, pp. 3917-3926,

2009.

[8] B. Dubertret, P. Skourides, D. J. Norris, V. Noireaux, A. H. Brivanlou and A.

Libchaber, "In vivo imaging of quantum dots encapsulated in phospholipid

micelles.," Science, vol. 298, no. 5599, pp. 1759-1762, 2002.

[9] Y. Liu, K. Li, J. Pan, B. Liu and S.-S. Feng, "Folic acid conjugated nanoparticles of

mixed lipid monolayer shell and biodegradable polymer core for targeted delivery

of Docetaxel.," Biomaterials, vol. 31, no. 2, pp. 330-338, 2010.

[10] L. O. Cinteza, T. Y. Ohulchanskyy, Y. Sahoo, E. J. Bergey, R. K. Pandey and P. N.

Prasad, "Diacyllipid micelle-based nanocarrier for magnetically guided delivery of

drugs in photodynamic therapy," Molecular Pharmaceutics, vol. 3, no. 4, pp. 415-

423, 2006.

[11] X. Wang, L.-H. Liu, O. Ramstrom and M. Yan, "Engineering nanomaterial

surfaces for biomedical applications," Experimental Biology and Medicine, vol.

234, no. 10, pp. 1128-1139, 2009.

[12] S. P. Vyas, A. Singh and V. Sihorkar, "Ligand-receptor-mediated drug delivery: an

emerging paradigm in cellular drug targeting," Critical Reviews™ in Therapeutic

Drug Carrier Systems, vol. 18, no. 1, pp. 1-76, 2001.

[13] J. Weingart, P. Vabbilisetty and X.-L. Sun, "Membrane mimetic surface

functionalization of nanoparticles: methods and applications.," Advances in Colloid

and Interface Science, Vol. 197-198, pp. 68-84, 2013.

[14] M. L. Immordino, F. Dosio and L. Cattel, "Stealth liposomes: review of the basic

science, rationale, and clinical applications, existing and potential," International

Journal of Nanomedicine, vol. 1, no. 3, p. 297, 2006.

[15] S. M. Moghimi, A. C. Hunter and J. C. Murray, "Long-circulating and target-

specific nanoparticles: theory to practice," Pharmacological Reviews, vol. 53, no.

2, pp. 283-318, 2001.

[16] W. J. Mulder, G. J. Strijkers, G. A. Van Tilborg, D. P. Cormode, Z. A. Fayad and

K. Nicolay, "Nanoparticulate assemblies of amphiphiles and diagnostically active

materials for multimodality imaging," Accounts of Chemical Research, vol. 42, no.

7, pp. 904-914, 2009.

[17] G. A. Koning and G. C. Krijger, "Targeted multifunctional lipid-based nanocarriers

for image-guided drug delivery.," Anticancer Agents in Medicinal Chemistry, vol.

7, no. 4, pp. 425-440, Jul 2007.

[18] G. Sessa and G. Weissmann, "Phospholipid spherules (liposomes) as a model for

biological membranes.," Journal of Lipid Research, vol. 9, no. 3, pp. 310-318,

1968.

[19] R. Langer and M. Chasin, Biodegradable Polymers as Drug Delivery Systems,

Marcel Dekker, New York, 1990.

[20] M. A. Tran, R. J. Watts and G. P. Robertson, "Use of liposomes as drug delivery

vehicles for treatment of melanoma," Pigment Cell & Melanoma Research, vol. 22,

no. 4, pp. 388-399, 2009.

[21] H. Harashima, K. Sakata, K. Funato and H. Kiwada, "Enhanced hepatic uptake of

liposomes through complement activation depending on the size of liposomes,"

Pharmaceutical Research, vol. 11, no. 3, pp. 402-406, 1994.

[22] D. Rajendran, S. S. Kumar, S. Dhanaraj, R. Dube and B. Suresh, "Comparative

evaluation of targeting efficiency of charged and neutral liposomes of 5-

fluorouracil," Drug Development and Industrial Pharmacy, vol. 23, no. 11, pp.

1099-1104, 1997.

[23] S. Ait-Oudhia, D. E. Mager and R. M. Straubinger, "Application of

Pharmacokinetic and Pharmacodynamic Analysis to the Development of

Liposomal Formulations for Oncology," Pharmaceutics, vol. 6, no. 1, pp. 137-174,

2014.

[24] D. C. Drummond, O. Meyer, K. Hong, D. B. Kirpotin and D. Papahadjopoulos,

"Optimizing liposomes for delivery of chemotherapeutic agents to solid tumors,"

Pharmacological Reviews, vol. 51, no. 4, pp. 691-744, 1999.

[25] R. Liu, L. Gan, X. Yang and H. Xu, "Chitosan as a condensing agent induces high

gene transfection efficiency and low cytotoxicity of liposome," Journal of

Bioscience and Bioengineering, vol. 111, no. 1, pp. 98-103, 2011.

[26] K. Nakano, Y. Tozuka and H. Takeuchi, "Effect of surface properties of liposomes

coated with a modified polyvinyl alcohol (PVA-R) on the interaction with

macrophage cells," International Journal of Pharmaceutics, vol. 354, no. 1, pp.

174-179, 2008.

[27] V. P. Torchilin, "Recent advances with liposomes as pharmaceutical carriers,"

Nature reviews Drug discovery, vol. 4, no. 2, pp. 145-160, 2005.

[28] C. Falciani, A. Accardo, J. Brunetti, D. Tesauro, B. Lelli, A. Pini, L. Bracci and G.

Morelli, "Target-Selective Drug Delivery through Liposomes Labeled with

Oligobranched Neurotensin Peptides," ChemMedChem, vol. 6, no. 4, pp. 678-685,

2011.

[29] N. Zhang and A. F. Palmer, "Liposomes surface conjugated with human

hemoglobin target delivery to macrophages," Biotechnology and Bioengineering,

vol. 109, no. 3, pp. 823-829, 2012.

[30] A. S. Manjappa, K. R. Chaudhari, M. P. Venkataraju, P. Dantuluri, B. Nanda, C.

Sidda, K. K. Sawant and R. S. Ramachandra Murthy, "Antibody derivatization and

conjugation strategies: application in preparation of stealth immunoliposome to

target chemotherapeutics to tumor," Journal of Controlled Release, vol. 150, no. 1,

pp. 2-22, 2011.

[31] K. Takara, H. Hatakeyama, N. Ohga, K. Hida and H. Harashima, "Design of a

dual-ligand system using a specific ligand and cell penetrating peptide, resulting in

a synergistic effect on selectivity and cellular uptake," International Journal of

Pharmaceutics, vol. 396, no. 1, pp. 143-148, 2010.

[32] C. K. Song, S. H. Jung, D.-D. Kim, K.-S. Jeong, B. C. Shin and H. Seong,

"Disaccharide-modified liposomes and their in vitro intracellular uptake,"

International Journal of Pharmaceutics, vol. 380, no. 1, pp. 161-169, 2009.

[33] R. I. Jolck, L. N. Feldborg, S. Andersen, S. M. Moghimi and T. L. Andresen,

"Engineering liposomes and nanoparticles for biological targeting," in

Biofunctionalization of Polymers and their Applications, Springer, 2011, pp. 251-

280.

[34] F. Szoka and D. Papahadjopoulos, "Procedure for preparation of liposomes with

large internal aqueous space and high capture by reverse-phase evaporation,"

Proceedings of the National Academy of Sciences, vol. 75, no. 9, pp. 4194-4198,

1978.

[35] H. Kikuchi, H. Yamauchi and S. Hirota, "A polyol dilution method for mass

production of liposomes," Journal of Liposome Research, vol. 4, no. 1, pp. 71-91,

1994.

[36] S. Batzri and E. D. Korn, "Single bilayer liposomes prepared without sonication,"

Biochimica et Biophysica Acta (BBA)-Biomembranes, vol. 298, no. 4, pp. 1015-

1019, 1973.

[37] U. Pick, "Liposomes with a large trapping capacity prepared by freezing and

thawing of sonicated phospholipid mixtures," Archives of Biochemistry and

Biophysics, vol. 212, no. 1, pp. 186-194, 1981.

[38] R. C. MacDonald, R. I. MacDonald, B. P. M. Menco, K. Takeshita, N. K. Subbarao

and L.-R. Hu, "Small-volume extrusion apparatus for preparation of large,

unilamellar vesicles," Biochimica et Biophysica Acta (BBA)-Biomembranes, vol.

1061, no. 2, pp. 297-303, 1991.

[39] A. Gomez-Hens and J. Manuel Fernandez-Romero, "The role of liposomes in

analytical processes," TrAC Trends in Analytical Chemistry, vol. 24, no. 1, pp. 9-

19, 2005.

[40] S. Yang, J. Chen, D. Zhao, D. Han and X. Chen, "Comparative study on

preparative methods of DC-Chol/DOPE liposomes and formulation optimization by

determining encapsulation efficiency," International Journal of Pharmaceutics,

vol. 434, no. 1, pp. 155-160, 2012.

[41] F. Szoka Jr and D. Papahadjopoulos, "Comparative properties and methods of

preparation of lipid vesicles (liposomes)," Annual Review of Biophysics and

Bioengineering, vol. 9, no. 1, pp. 467-508, 1980.

[42] M. Arruebo, R. Fernandez-Pacheco, M. R. Ibarra and J. Santamaria, "Magnetic

nanoparticles for drug delivery," Nano Today, vol. 2, no. 3, pp. 22-32, 2007.

[43] Q. A. Pankhurst, J. Connolly, S. Jones and J. Dobson, "Applications of magnetic

nanoparticles in biomedicine," Journal of physics D: Applied Physics, vol. 36, no.

13, p. R167, 2003.

[44] Y.-X. J. Wang, S. M. Hussain and G. P. Krestin, "Superparamagnetic iron oxide

contrast agents: physicochemical characteristics and applications in MR imaging,"

European Radiology, vol. 11, no. 11, pp. 2319-2331, 2001.

[45] A. Roca, M. Morales, K. O’Grady and C. Serna, "Structural and magnetic

properties of uniform magnetite nanoparticles prepared by high temperature

decomposition of organic precursors," Nanotechnology, vol. 17, no. 11, p. 2783,

2006.

[46] R. Qiao, C. Yang and M. Gao, "Superparamagnetic iron oxide nanoparticles: from

preparations to in vivo MRI applications," Journal of Materials Chemistry, vol. 19,

no. 35, pp. 6274-6293, 2009.

[47] S. Laurent, D. Forge, M. Port, A. Roch, C. Robic, L. Vander Elst and R. N. Muller,

"Magnetic iron oxide nanoparticles: synthesis, stabilization, vectorization,

physicochemical characterizations, and biological applications," Chemical Reviews,

vol. 108, no. 6, pp. 2064-2110, 2008.

[48] E. V. Shtykova, X. Huang, N. Remmes, D. Baxter, B. Stein, B. Dragnea, D. I.

Svergun and L. M. Bronstein, "Structure and properties of iron oxide nanoparticles

encapsulated by phospholipids with poly (ethylene glycol) tails," The Journal of

Physical Chemistry C, vol. 111, no. 49, pp. 18078-18086, 2007.

[49] J. Park, K. An, Y. Hwang, J.-G. Park, H.-J. Noh, J.-Y. Kim, J.-H. Park, N.-M.

Hwang and T. Hyeon, "Ultra-large-scale syntheses of monodisperse nanocrystals,"

Nature Materials, vol. 3, no. 12, pp. 891-895, 2004.

[50] Y. Sahoo, A. Goodarzi, M. T. Swihart, T. Y. Ohulchanskyy, N. Kaur, E. P. Furlani

and P. N. Prasad, "Aqueous ferrofluid of magnetite nanoparticles: fluorescence

labeling and magnetophoretic control," The Journal of Physical Chemistry B, vol.

109, no. 9, pp. 3879-3885, 2005.

[51] L. M. Bronstein, X. Huang, J. Retrum, A. Schmucker, M. Pink, B. D. Stein and B.

Dragnea, "Influence of iron oleate complex structure on iron oxide nanoparticle

formation," Chemistry of Materials, vol. 19, no. 15, pp. 3624-3632, 2007.

[52] C. Yue-Jian, T. Juan, X. Fei, Z. Jia-Bi, G. Ning, Z. Yi-Hua, D. Ye and G. Liang,

"Synthesis, self-assembly, and characterization of PEG-coated iron oxide

nanoparticles as potential MRI contrast agent," Drug Development and Industrial

Pharmacy, vol. 36, no. 10, pp. 1235-1244, 2010.

[53] M. M. Husein and N. N. Nassar, "Nanoparticle preparation using the single

microemulsions scheme," Current Nanoscience, vol. 4, no. 4, pp. 370-380, 2008.

[54] J. Lopez Perez, M. Lopez Quintela, J. Mira, J. Rivas and S. Charles, "Advances in

the preparation of magnetic nanoparticles by the microemulsion method," The

Journal of Physical Chemistry B, vol. 101, no. 41, pp. 8045-8047, 1997.

[55] F. M. Kievit and M. Zhang, "Surface engineering of iron oxide nanoparticles for

targeted cancer therapy," Accounts of Chemical Research, vol. 44, no. 10, pp. 853-

862, 2011.

[56] C. Fang and M. Zhang, "Multifunctional magnetic nanoparticles for medical

imaging applications," Journal of Materials Chemistry, vol. 19, no. 35, pp. 6258-

6266, 2009.

[57] Y.-w. Jun, J.-H. Lee and J. Cheon, "Chemical Design of Nanoparticle Probes for

High-Performance Magnetic Resonance Imaging," Angewandte Chemie

International Edition, vol. 47, no. 28, pp. 5122-5135, 2008.

[58] N. Nitin, L. LaConte, O. Zurkiya, X. Hu and G. Bao, "Functionalization and

peptide-based delivery of magnetic nanoparticles as an intracellular MRI contrast

agent," JBIC Journal of Biological Inorganic Chemistry, vol. 9, no. 6, pp. 706-712,

2004.

[59] N. Lee, Y. Choi, Y. Lee, M. Park, W. K. Moon, S. H. Choi and T. Hyeon, "Water-

dispersible ferrimagnetic iron oxide nanocubes with extremely high r 2 relaxivity

for highly sensitive in vivo MRI of tumors," Nano letters, vol. 12, no. 6, pp. 3127-

3131, 2012.

[60] P. Mohan and N. Rapoport, "Doxorubicin as a molecular nanotheranostic agent:

effect of doxorubicin encapsulation in micelles or nanoemulsions on the

ultrasound-mediated intracellular delivery and nuclear trafficking," Molecular

Pharmaceutics, vol. 7, no. 6, pp. 1959-1973, 2010.

[61] G. Zhang, Y. Liao and I. Baker, "Surface engineering of core/shell iron/iron oxide

nanoparticles from microemulsions for hyperthermia," Materials Science and

Engineering: C, vol. 30, no. 1, pp. 92-97, 2010.

[62] G. A. van Tilborg, D. P. Cormode, P. A. Jarzyna, A. van der Toorn, S. M. van der

Pol, L. van Bloois, Z. A. Fayad, G. Storm, W. J. Mulder, H. E. de Vries and others,

"Nanoclusters of iron oxide: effect of core composition on structure,

biocompatibility, and cell labeling efficacy," Bioconjugate Chemistry, vol. 23, no.

5, pp. 941-950, 2012.

[63] K. L. Hultman, A. J. Raffo, A. L. Grzenda, P. E. Harris, T. R. Brown and S.

O’Brien, "Magnetic resonance imaging of major histocompatibility class II

expression in the renal medulla using immunotargeted superparamagnetic iron

oxide nanoparticles," Acs Nano, vol. 2, no. 3, pp. 477-484, 2008.

[64] W. Elmore, "Ferromagnetic colloid for studying magnetic structures," Physical

Review, vol. 54, no. 4, p. 309, 1938.

[65] C. Ricketts, J. Cox, C. Fitzmaurice and G. Moss, "The iron-dextran complex.,"

Nature, vol. 208, no. 5007, p. 237, 1965.

[66] M. A. Lopez-Quintela, C. Tojo, M. Blanco, L. Garc{\i}a Rio and J. Leis,

"Microemulsion dynamics and reactions in microemulsions," Current Opinion In

Colloid & Interface Science, vol. 9, no. 3, pp. 264-278, 2004.

[67] T. Hyeon, S. S. Lee, J. Park, Y. Chung and H. B. Na, "Synthesis of highly

crystalline and monodisperse maghemite nanocrystallites without a size-selection

process," Journal of the American Chemical Society, vol. 123, no. 51, pp. 12798-

12801, 2001.

[68] S. Sun and H. Zeng, "Size-controlled synthesis of magnetite nanoparticles,"

Journal of the American Chemical Society, vol. 124, no. 28, pp. 8204-8205, 2002.

[69] Z. Li, H. Chen, H. Bao and M. Gao, "One-pot reaction to synthesize water-soluble

magnetite nanocrystals," Chemistry of Materials, vol. 16, no. 8, pp. 1391-1393,

2004.

[70] M. Iijima, K. Sato, K. Kurashima, T. Ishigaki and H. Kamiya, "Low-temperature

synthesis of redispersible iron oxide nanoparticles under atmospheric pressure and

ultradense reagent concentration," Powder Technology, vol. 181, no. 1, pp. 45-50,

2008.

[71] W. Al-Jamal and K. Kostarelos, "Liposome-nanoparticle hybrids for multimodal

diagnostic and therapeutic applications," Nanomedicine, vol. 2, no. 1, pp. 85-98,

2007.

[72] Y. Zhang, N. Kohler and M. Zhang, "Surface modification of superparamagnetic

magnetite nanoparticles and their intracellular uptake," Biomaterials, vol. 23, no. 7,

pp. 1553-1561, 2002.

[73] A. K. Gupta and S. Wells, "Surface-modified superparamagnetic nanoparticles for

drug delivery: preparation, characterization, and cytotoxicity studies,"

NanoBioscience, IEEE Transactions on, vol. 3, no. 1, pp. 66-73, 2004.

[74] J. V. Jokerst, T. Lobovkina, R. N. Zare and S. S. Gambhir, "Nanoparticle

PEGylation for imaging and therapy," Nanomedicine, vol. 6, no. 4, pp. 715-728,

2011.

[75] K. Kaaki, K. Herve-Aubert, M. Chiper, A. Shkilnyy, M. Souce, R. Benoit, A.

Paillard, P. Dubois, M.-L. Saboungi and I. Chourpa, "Magnetic nanocarriers of

doxorubicin coated with poly(ethylene glycol) and folic acid: relation between

coating structure, surface properties, colloidal stability, and cancer cell targeting.,"

Langmuir, vol. 28, no. 2, pp. 1496-1505, Jan 2012.

[76] H.-C. Huang, P.-Y. Chang, K. Chang, C.-Y. Chen, C.-W. Lin, J.-H. Chen, C.-Y.

Mou, Z.-F. Chang and F.-H. Chang, "Formulation of novel lipid-coated magnetic

nanoparticles as the probe for in vivo imaging," Journal of Biomedical Science,

vol. 16, no. 1, p. 86, 2009.

[77] P. A. Jarzyna, L. H. Deddens, B. H. Kann, S. Ramachandran, C. Calcagno, W.

Chen, A. Gianella, R. M. Dijkhuizen, A. W. Griffioen, Z. A. Fayad and others,

"Tumor angiogenesis phenotyping by nanoparticle-facilitated magnetic resonance

and near-infrared fluorescence molecular imaging," Neoplasia, vol. 14, no. 10, pp.

964-973, 2012.

[78] S. Tong, S. Hou, B. Ren, Z. Zheng and G. Bao, "Self-assembly of phospholipid--

PEG coating on nanoparticles through dual solvent exchange," Nano letters, vol.

11, no. 9, pp. 3720-3726, 2011.

[79] L. E. LaConte, N. Nitin, O. Zurkiya, D. Caruntu, C. J. O'Connor, X. Hu and G.

Bao, "Coating thickness of magnetic iron oxide nanoparticles affects R2 relaxivity,"

Journal of Magnetic Resonance Imaging, vol. 26, no. 6, pp. 1634-1641, 2007.

[80] S. Tong, S. Hou, Z. Zheng, J. Zhou and G. Bao, "Coating optimization of

superparamagnetic iron oxide nanoparticles for high T2 relaxivity," Nano letters,

vol. 10, no. 11, pp. 4607-4613, 2010.

[81] W. J. Mulder, G. J. Strijkers, G. A. van Tilborg, A. W. Griffioen and K. Nicolay,

"Lipid-based nanoparticles for contrast-enhanced MRI and molecular imaging,"

NMR in Biomedicine, vol. 19, no. 1, pp. 142-164, 2006.

[82] S. A. Wahajuddin, "Superparamagnetic iron oxide nanoparticles: magnetic

nanoplatforms as drug carriers," International Journal of Nanomedicine, vol. 7, p.

3445, 2012.

[83] J. C. Rogers and S. Kornfeld, "Hepatic uptake of proteins coupled to fetuin

glycopeptide.," Biochemical and Biophysical Research Communications, vol. 45,

no. 3, pp. 622-629, 1971.

[84] J. Wu, M. H. Nantz and M. A. Zern, "Targeting hepatocytes for drug and gene

delivery: emerging novel approaches and applications.," Frontiers in Bioscience,

vol. 7, pp. d717-d725, 2002.

[85] L. Beljaars, G. Molema, B. Weert, H. Bonnema, P. Olinga, G. M. Groothuis, D. K.

Meijer and K. Poelstra, "Albumin modified with mannose 6-phosphate: A potential

carrier for selective delivery of antifibrotic drugs to rat and human hepatic stellate

cells.," Hepatology, vol. 29, no. 5, pp. 1486-1493, 1999.

[86] L. Beljaars, P. Olinga, G. Molema, P. de Bleser, A. Geerts, G. M. Groothuis, D. K.

Meijer and K. Poelstra, "Characteristics of the hepatic stellate cell-selective carrier

mannose 6-phosphate modified albumin (M6P(28)-HSA).," Liver, vol. 21, no. 5,

pp. 320-328, 2001.

[87] W. I. Hagens, L. Beljaars, D. A. Mann, M. C. Wright, B. Julien, S. Lotersztajn, C.

Reker-Smit and K. Poelstra, "Cellular targeting of the apoptosis-inducing

compound gliotoxin to fibrotic rat livers," Journal of Pharmacology and

Experimental Therapeutics, vol. 324, no. 3, pp. 902-910, 2008.

[88] J. M. Luk, Q.-S. Zhang, N. P. Lee, J. Y. Wo, P. P. Leung, L.-X. Liu, M.-Y. Hu, K.-

F. Cheung, C.-K. Hui, G. K. Lau and S.-T. Fan, "Hepatic stellate cell-targeted

delivery of M6P-HSA-glycyrrhetinic acid attenuates hepatic fibrogenesis in a bile

duct ligation rat model.," Liver Int, vol. 27, no. 4, pp. 548-557, 2007.

[89] J. E. Adrian, K. Poelstra and J. A. A. M. Kamps, "Addressing liver fibrosis with

liposomes targeted to hepatic stellate cells.," Journal of Liposome Research, vol.

17, no. 3-4, pp. 205-218, 2007.

[90] J. E. Adrian, K. Poelstra, G. L. Scherphof, D. K. F. Meijer, A.-M. van Loenen-

Weemaes, C. Reker-Smit, H. W. M. Morselt, P. Zwiers and J. A. A. M. Kamps,

"Effects of a new bioactive lipid-based drug carrier on cultured hepatic stellate cells

and liver fibrosis in bile duct-ligated rats.," Journal of Pharmacology and

Experimental Therapeutics, vol. 321, no. 2, pp. 536-543, 2007.

[91] M. N. Jones, "Carbohydrate-mediated liposomal targeting and drug delivery,"

Advanced Drug Delivery Reviews, vol. 13, no. 3, pp. 215-249, 1994.

[92] K.-A. Karlsson, "Microbial recognition of target-cell glycoconjugates," Current

Opinion in Structural Biology, vol. 5, no. 5, pp. 622-635, 1995.

[93] H. Lis and N. Sharon, "Lectins: carbohydrate-specific proteins that mediate cellular

recognition," Chemical Reviews, vol. 98, no. 2, pp. 637-674, 1998.

[94] S. L. Flitsch and R. V. Ulijn, "Sugars tied to the spot," Nature, vol. 421, no. 6920,

pp. 219-220, 2003.

[95] K. J. Lee, S. Mao, C. Sun, C. Gao, O. Blixt, S. Arrues, L. G. Hom, G. F.

Kaufmann, T. Z. Hoffman, A. R. Coyle, J. Paulson, B. Felding-Habermann and K.

D. Janda, "Phage-display selection of a human single-chain fv antibody highly

specific for melanoma and breast cancer cells using a chemoenzymatically

synthesized G(M3)-carbohydrate antigen.," Journal of American Chemical Society,

vol. 124, no. 42, pp. 12439-12446, 2002.

[96] N. Jayaraman, K. Maiti and K. Naresh, "Multivalent glycoliposomes and micelles

to study carbohydrate--protein and carbohydrate--carbohydrate interactions,"

Chemical Society Reviews, vol. 42, no. 11, pp. 4640-4656, 2013.

[97] R. Mehvar, "Recent trends in the use of polysaccharides for improved delivery of

therapeutic agents: pharmacokinetic and pharmacodynamic perspectives," Current

Pharmaceutical Biotechnology, vol. 4, no. 5, pp. 283-302, 2003.

[98] J. M. Irache, H. H. Salman, C. Gamazo and S. Espuelas, "Mannose-targeted

systems for the delivery of therapeutics," Expert Opinion Drug Delivery, vol.5, no.

6, pp. 703-724, 2008.

[99] X.-L. Sun, Y. Kanie, C.-T. Guo, O. Kanie, Y. Suzuki and C.-H. Wong, "Syntheses

of C-3-Modified Sialylglycosides as Selective Inhibitors of Influenza

Hemagglutinin and Neuraminidase," European Journal of Organic Chemistry, vol.

2000, no. 14, pp. 2643-2653, 2000.

[100] A. M. Haywood and B. P. Boyer, "Ficoll and dextran enhance adhesion of Sendai

virus to liposomes containing receptor (ganglioside GD1a)," Biochemistry, vol. 25,

no. 13, pp. 3925-3929, 1986.

[101] J. S. Slama and R. R. Rando, "Lectin-mediated aggregation of liposomes

containing glycolipids with variable hydrophilic spacer arms," Biochemistry, vol.

19, no. 20, pp. 4595-4600, 1980.

[102] S. Zalipsky, N. Mullah, A. Dibble and T. Flaherty, "New chemoenzymatic

approach to glyco-lipopolymers: practical preparation of functionally active

galactose--poly (ethylene glycol)--distearoylphosphatidic acid (Gal-PEG-DSPA)

conjugate," Chemical Communications, no. 7, pp. 653-654, 1999.

[103] J. Zhu, J. Xue, Z. Guo, L. Zhang and R. E. Marchant, "Biomimetic glycoliposomes

as nanocarriers for targeting P-selectin on activated platelets," Bioconjugate

Chemistry, vol. 18, no. 5, pp. 1366-1369, 2007.

[104] P. Sapra and T. Allen, "Ligand-targeted liposomal anticancer drugs," Progress in

Lipid Research, vol. 42, no. 5, pp. 439-462, 2003.

[105] J. Kunimasa, K. Inui, R. Hori, Y. Kawamura and K. Endo, "Mannan-coated

liposome delivery of gadolinium-diethylenetriaminepentaacetic acid, a contrast

agent for use in magnetic resonance imaging.," Chemical & Pharmaceutical

Bulletin, vol. 40, no. 9, pp. 2565-2567, 1992.

[106] S. Vyas, Y. Katare, V. Mishra and V. Sihorkar, "Ligand directed macrophage

targeting of amphotericin B loaded liposomes," International Journal of

Pharmaceutics, vol. 210, no. 1, pp. 1-14, 2000.

[107] S. Chono, T. Tanino, T. Seki and K. Morimoto, "Uptake characteristics of

liposomes by rat alveolar macrophages: influence of particle size and surface

mannose modification," Journal of Pharmacy and Pharmacology, vol. 59, no. 1,

pp. 75-80, 2007.

[108] A. Gabizon, A. T. Horowitz, D. Goren, D. Tzemach, F. Mandelbaum-Shavit, M. M.

Qazen and S. Zalipsky, "Targeting folate receptor with folate linked to extremities

of poly (ethylene glycol)-grafted liposomes: in vitro studies," Bioconjugate

Chemistry, vol. 10, no. 2, pp. 289-298, 1999.

[109] Z. Xu, J. Jayaseharan and R. E. Marchant, "Synthesis and characterization of

oligomaltose-grafted lipids with application to liposomes," Journal of Colloid and

Interface Science, vol. 252, no. 1, pp. 57-65, 2002.

[110] G.-J. Boons, "Liposomes modified by carbohydrate ligands can target B cells for

the treatment of B-cell lymphomas," Expert Review of Vaccines, vol. 9, no. 11, pp.

1251-1256, 2010.

[111] Y. Ikehara, T. Niwa, L. Biao, S. K. Ikehara, N. Ohashi, T. Kobayashi, Y. Shimizu,

N. Kojima and H. Nakanishi, "A carbohydrate recognition--based drug delivery

and controlled release system using intraperitoneal macrophages as a cellular

vehicle," Cancer Research, vol. 66, no. 17, pp. 8740-8748, 2006.

[112] Y. Ikehara, N. Shiuchi, S. Kabata-Ikehara, H. Nakanishi, N. Yokoyama, H. Takagi,

T. Nagata, Y. Koide, K. Kuzushima, T. Takahashi and others, "Effective induction

of anti-tumor immune responses with oligomannose-coated liposome targeting to

intraperitoneal phagocytic cells," Cancer Letters, vol. 260, no. 1, pp. 137-145,

2008.

[113] Y. Harada, T. Murata, K. Totani, T. Kajimoto, S. M. Masum, Y. Tamba, M.

Yamazaki and T. Usui, "Design and facile synthesis of neoglycolipids as

lactosylceramide mimetics and their transformation into glycoliposomes,"

Bioscience, Biotechnology, and Biochemistry, vol. 69, no. 1, pp. 166-178, 2005.

[114] F. Said Hassane, A. Phalipon, M. Tanguy, C. Guerreiro, F. Belot, B. Frisch, L. A.

Mulard and F. Schuber, "Rational design and immunogenicity of liposome-based

diepitope constructs: Application to synthetic oligosaccharides mimicking the

Shigella flexneri 2a O-antigen," Vaccine, vol. 27, no. 39, pp. 5419-5426, 2009.

[115] V. H. Thomas, J. Elhalabi and K. G. Rice, "Enzymatic synthesis of N-linked

oligosaccharides terminating in multiple sialyl-Lewis(x) and GalNAc-Lewis(x)

determinants: clustered glycosides for studying selectin interactions.,"

Carbohydrate Research, vol. 306, no. 3, pp. 387-400, 1998.

[116] C. Muller and F. Schuber, "Neo-mannosylated liposomes: synthesis and interaction

with mouse Kupffer cells and resident peritoneal macrophages," Biochimica et

Biophysica Acta (BBA)-Biomembranes, vol. 986, no. 1, pp. 97-105, 1989.

[117] F. Said Hassane, B. Frisch and F. Schuber, "Targeted liposomes: convenient

coupling of ligands to preformed vesicles using "click chemistry".," Bioconjugate

Chemistry, vol. 17, no. 3, pp. 849-854, 2006.

[118] H. Zhang, Y. Ma and X.-L. Sun, "Chemically-selective surface glyco-

functionalization of liposomes through Staudinger ligation," Chemical

Communications, no. 21, pp. 3032-3034, 2009.

[119] A. Engel, S. K. Chatterjee, A. Al-arifi, D. Riemann, J. Langner and P. Nuhn,

"Influence of spacer length on interaction of mannosylated liposomes with human

phagocytic cells," Pharmaceutical Research, vol. 20, no. 1, pp. 51-57, 2003.

[120] H.-J. Gabius, "Glycans: bioactive signals decoded by lectins.," Biochemical Society

Transactions, vol. 36, no. Pt 6, pp. 1491-1496, 2008.

[121] Y. C. Lee and R. T. Lee, "Carbohydrate-protein interactions: basis of

glycobiology," Accounts of Chemical Research, vol. 28, no. 8, pp. 321-327, 1995.

[122] L.-H. Liu, H. Dietsch, P. Schurtenberger and M. Yan, "Photoinitiated coupling of

unmodified monosaccharides to iron oxide nanoparticles for sensing proteins and

bacteria," Bioconjugate Chemistry, vol. 20, no. 7, pp. 1349-1355, 2009.

[123] K. El-Boubbou, D. C. Zhu, C. Vasileiou, B. Borhan, D. Prosperi, W. Li and X.

Huang, "Magnetic glyco-nanoparticles: a tool to detect, differentiate, and unlock

the glyco-codes of cancer via magnetic resonance imaging," Journal of the

American Chemical Society, vol. 132, no. 12, pp. 4490-4499, 2010.

[124] C.-M. Lee, H.-J. Jeong, E.-M. Kim, D. W. Kim, S. T. Lim, H. T. Kim, I.-K. Park,

Y. Y. Jeong, J. W. Kim and M.-H. Sohn, "Superparamagnetic iron oxide

nanoparticles as a dual imaging probe for targeting hepatocytes in vivo," Magnetic

Resonance in Medicine, vol. 62, no. 6, pp. 1440-1446, 2009.

[125] M. K. Yoo, I. Y. Park, I. Y. Kim, I. K. Park, J.-S. Kwon, H.-J. Jeong, Y. Y. Jeong

and C. S. Cho, "Superparamagnetic iron oxide nanoparticles coated with mannan

for macrophage targeting," Journal of Nanoscience and Nanotechnology, vol. 8,

no. 10, pp. 5196-5202, 2008.

[126] S. I. van Kasteren, S. J. Campbell, S. Serres, D. C. Anthony, N. R. Sibson and B.

G. Davis, "Glyconanoparticles allow pre-symptomatic in vivo imaging of brain

disease," Proceedings of the National Academy of Sciences, vol. 106, no. 1, pp. 18-

23, 2009.

[127] M. K. Yu, J. Park and S. Jon, "Targeting strategies for multifunctional

nanoparticles in cancer imaging and therapy.," Theranostics, vol. 2, no. 1, pp. 3-44,

2012.

[128] R.-M. Lu, Y.-L. Chang, M.-S. Chen and H.-C. Wu, "Single chain anti-c-Met

antibody conjugated nanoparticles for in vivo tumor-targeted imaging and drug

delivery.," Biomaterials, vol. 32, no. 12, pp. 3265-3274, 2011.

[129] Y. Ge, Y. Zhang, S. He, F. Nie, G. Teng and N. Gu, "Fluorescence Modified

Chitosan-Coated Magnetic Nanoparticles for High-Efficient Cellular Imaging.,"

Nanoscale Research Letters, vol. 4, no. 4, pp. 287-295, 2009.

[130] H. Lee, M. K. Yu, S. Park, S. Moon, J. J. Min, Y. Y. Jeong, H.-W. Kang and S.

Jon, "Thermally cross-linked superparamagnetic iron oxide nanoparticles: synthesis

and application as a dual imaging probe for cancer in vivo.," Journal of American

Chemical Society, vol. 129, no. 42, pp. 12739-12745, 2007.

[131] F. M. Brunel, J. D. Lewis, G. Destito, N. F. Steinmetz, M. Manchester, H.

Stuhlmann and P. E. Dawson, "Hydrazone ligation strategy to assemble

multifunctional viral nanoparticles for cell imaging and tumor targeting.," Nano

Letters, vol. 10, no. 3, pp. 1093-1097, 2010.

[132] J. Xie, K. Chen, H.-Y. Lee, C. Xu, A. R. Hsu, S. Peng, X. Chen and S. Sun,

"Ultrasmall c (RGDyK)-coated Fe3O4 nanoparticles and their specific targeting to

integrin αvβ3-rich tumor cells," Journal of the American Chemical Society, vol.

130, no. 24, pp. 7542-7543, 2008.

[133] M.-M. Chua, S.-T. Fan and others, "Attachment of immunoglobulin to liposomal

membrane via protein carbohydrate," Biochimica et Biophysica Acta (BBA)-

General Subjects, vol. 800, no. 3, pp. 291-300, 1984.

[134] V. Torchilin, T. Levchenko, A. Lukyanov, B. Khaw, A. Klibanov, R. Rammohan,

G. Samokhin and K. Whiteman, " p-Nitrophenylcarbonyl-PEG-PE-liposomes: fast

and simple attachment of specific ligands, including monoclonal antibodies, to

distal ends of PEG chains via p-nitrophenylcarbonyl groups," Biochimica et

Biophysica Acta (BBA)-Biomembranes, vol. 1511, no. 2, pp. 397-411, 2001.

[135] O. Veiseh, J. W. Gunn and M. Zhang, "Design and fabrication of magnetic

nanoparticles for targeted drug delivery and imaging," Advanced Drug Delivery

Reviews, vol. 62, no. 3, pp. 284-304, 2010.

[136] X. Zhang and Y. Zhang, "Applications of Azide-Based Bioorthogonal Click

Chemistry in Glycobiology," Molecules, vol. 18, no. 6, pp. 7145-7159, 2013.

[137] E. Saxon and C. R. Bertozzi, "Cell surface engineering by a modified Staudinger

reaction," Science, vol. 287, no. 5460, pp. 2007-2010, 2000.

[138] P. Vabbilisetty and X.-L. Sun, "Liposome surface functionalization based on

different anchoring lipids via Staudinger ligation.," Organic and Biomolecular

Chemistry, vol. 12, no. 8, pp. 1237-1244, 2014.

[139] H. C. Kolb, M. G. Finn and K. B. Sharpless, "Click Chemistry: Diverse Chemical

Function from a Few Good Reactions.," Angew Chem Int Ed Engl, vol. 40, no. 11,

pp. 2004-2021, 2001.

[140] C. D. Hein, X.-M. Liu and D. Wang, "Click chemistry, a powerful tool for

pharmaceutical sciences.," Pharmaceutical Research, vol. 25, no. 10, pp. 2216-

2230, 2008.

[141] H. Zhang, J. Weingart, R. Jiang, J. Peng, Q. Wu and X.-L. Sun, "Bio-inspired

liposomal thrombomodulin conjugate through bio-orthogonal chemistry.,"

Bioconjugate Chemistry, vol. 24, no. 4, pp. 550-559, 2013.

[142] M. T. Stephan and D. J. Irvine, "Enhancing cell therapies from the outside in: Cell

surface engineering using synthetic nanomaterials," Nano today, vol. 6, no. 3, pp.

309-325, 2011.

[143] J. M. Baskin, J. A. Prescher, S. T. Laughlin, N. J. Agard, P. V. Chang, I. A. Miller,

A. Lo, J. A. Codelli and C. R. Bertozzi, "Copper-free click chemistry for dynamic

in vivo imaging," Proceedings of the National Academy of Sciences, vol. 104, no.

43, pp. 16793-16797, 2007.

[144] D. Dutta, A. Pulsipher, W. Luo, H. Mak and M. N. Yousaf, "Engineering cell

surfaces via liposome fusion," Bioconjugate Chemistry, vol. 22, no. 12, pp. 2423-

2433, 2011.

[145] C. A. Holden, Q. Yuan, W. A. Yeudall, D. A. Lebman and H. Yang, "Surface

engineering of macrophages with nanoparticles to generate a cell--nanoparticle

hybrid vehicle for hypoxia-targeted drug delivery," International Journal of

Nanomedicine, vol. 5, p. 25, 2010.

[146] D. Ringler, "Inflammation and repair," Veterinary pathology. 6th Edition, TC

Jones, pp. 113-158, 1997.

[147] D. W. Kennedy and J. L. Abkowitz, "Mature monocytic cells enter tissues and

engraft," Proceedings of the National Academy of Sciences, vol. 95, no. 25, pp.

14944-14949, 1998.

[148] M. Naito, S. Umeda, T. Yamamoto, H. Moriyama, H. Umezu, G. Hasegawa, H.

Usuda, L. D. Shultz and K. Takahashi, "Development, differentiation, and

phenotypic heterogeneity of murine tissue macrophages.," Journal of Leukocyte

Biology, vol. 59, no. 2, pp. 133-138, 1996.

[149] P. J. Murray and T. A. Wynn, "Protective and pathogenic functions of macrophage

subsets.," Nature Reviews Immunology, vol. 11, no. 11, pp. 723-737, 2011.

[150] C. Kelly, C. Jefferies and S.-A. Cryan, "Targeted liposomal drug delivery to

monocytes and macrophages," Journal of Drug Delivery, vol. 2011, 2010.

[151] S.-W. Kang, S. Lee, J. H. Na, H. I. Yoon, D.-E. Lee, H. Koo, Y. W. Cho, S. H.

Kim, S. Y. Jeong, I. C. Kwon and others, "Cell Labeling and Tracking Method

without Distorted Signals by Phagocytosis of Macrophages," Theranostics, vol. 4,

no. 4, p. 420, 2014.

[152] J. Du, M. A. Meledeo, Z. Wang, H. S. Khanna, V. D. Paruchuri and K. J. Yarema,

"Metabolic glycoengineering: sialic acid and beyond," Glycobiology, vol. 19, no.

12, pp. 1382-1401, 2009.

[153] J. L. Panza, W. R Wagner, H. L. Rilo, R. Harsha Rao, E. J. Beckman and A. J.

Russell, "Treatment of rat pancreatic islets with reactive PEG," Biomaterials, vol.

21, no. 11, pp. 1155-1164, 2000.

[154] P. De Bank, B. Kellam, D. Kendall and K. Shakesheff, "Surface engineering of

living myoblasts via selective periodate oxidation," Biotechnology and

Bioengineering, vol. 81, no. 7, pp. 800-808, 2003.

[155] C. L. Stabler, X.-L. Sun, W. Cui, J. T. Wilson, C. A. Haller and E. L. Chaikof,

"Surface re-engineering of pancreatic islets with recombinant azido-

thrombomodulin," Bioconjugate Chemistry, vol. 18, no. 6, pp. 1713-1715, 2007.

[156] H. R. Zope, F. Versluis, A. Ordas, J. Voskuhl, H. P. Spaink and A. Kros, "In Vitro

and In Vivo Supramolecular Modification of Biomembranes Using a Lipidated

Coiled-Coil Motif," Angewandte Chemie International Edition, vol. 52, no. 52, pp.

14247-14251, 2013.

[157] S. Kawakami, A. Sato, M. Nishikawa, F. Yamashita and M. Hashida, "Mannose

receptor-mediated gene transfer into macrophages using novel mannosylated

cationic liposomes.," Gene Therapy, vol. 7, no. 4, pp. 292-299, 2000.

[158] W. C. Chen, N. Kawasaki, C. M. Nycholat, S. Han, J. Pilotte, P. R. Crocker and J.

C. Paulson, "Antigen delivery to macrophages using liposomal nanoparticles

targeting sialoadhesin/CD169," PloS one, vol. 7, no. 6, p. e39039, 2012.

CHAPTER II

LIPOSOME SURFACE FUNCTIONALIZATION BASED ON DIFFERENT

ANCHORING LIPIDS VIA STAUDINGER LIGATION

2.1 Introduction

Liposomes as self-assembled lipid bi-layers have been widely investigated as biomimetic models of cell membranes, biomimetic conjugates, and drug, gene and image agent delivery vehicles [1, 2]. Liposome surface functionalization facilitates enormous potential applications of liposomes, such as enhanced stability, bioactive liposome conjugates, and targeted delivery [3, 4]. Various biomolecules have been conjugated onto liposome surface for variety of biomedical applications. Among them, carbohydrate molecules, particularly which play an important role in various biological recognition processes such as fertilization, metastasis, inflammations and host–pathogen adhesion

[5], serve as attractive molecules for surface modification of liposomes with purpose for

targeted drug delivery [6] and as bioactive membrane mimetic conjugates [7]. Many methods have been developed to prepare surface functionalized liposomes, of which the most commonly used method involves the initial synthesis of an anchor lipid-ligand conjugate, followed by formulation of the liposome with all lipid components or post- insertion into the preformed liposome. Alternatively, chemical modifications of the surface of preformed vesicles that carry functionalized anchoring lipids with biomolecules have been well explored [8, 9]. So far, variable conjugation methods such as using amide [10] or thiol-maleimide coupling [11] as well as by imine [12] or hydrazone linkage, [13] have been developed for liposome surface functionalization.

However, in many cases there is a lack of specificity resulting in the uncontrolled formation of the number of covalent bonds between liposome and ligands of interest. In addition, the reactive anchoring lipid might react with other membrane components or the encapsulated drugs as well in this conventional conjugation chemistry. Recently, bioorthogonal chemistry such as click chemistry [14] and Staudinger ligation [15] have been investigated as novel generic chemical tools for the facile in situ surface modification of liposomes. Particularly, the Staudinger ligation reaction is known to occur in high yield at room temperature in aqueous solvents and without any catalyst, and is compatible with the unprotected functional groups of wide range of biomolecules.

The anchoring lipids are essential for liposome surface functionalization and play important roles in ligand grafting density, liposome stability, and liposome chemical and physical characteristics. In the present study, two anchor lipids, phosphatidylethonalamine (PE) and cholesterol (Chol) are investigated for liposome surface glyco-functionalization via Staudinger ligation, with the consideration of stability

and grafting carbohydrate density as well as activity of glyco-liposome conjugates, as shown in (Figure 8.). Briefly, glyco-functionalized liposome systems were prepared by chemically selective glyco-functionalization of liposomes carrying anchoring lipids namely Chol-PEG2000-TP and DSPE-PEG2000-TP via Staudinger ligation, respectively.

The size and stability of the liposomes were confirmed by dynamic light scattering

(DLS). Also, the impact of anchoring lipids on encapsulation and releasing capacity of the glycosylated liposomes were investigated by entrapping 5, 6-carboxyfluorescein dye and monitoring the fluorescence dye leakage, respectively. Furthermore, the density and accessibility of grafted carbohydrate residues on the liposome surface were evaluated with lectin binding studies for DSPE-PEG and Chol-PEG-derived liposome, respectively.

Figure 8. Liposome surface glyco-functionalization based on two kinds of anchoring lipids via Staudinger ligation.

2.2 Experimental

2.2.1 Materials and methods

All solvents and reagents were purchased from commercial sources and were used as received, unless otherwise noted. Deionized water was used as a solvent in all procedures. Monocholesteryl-PEG2000-amine was synthesized as literature method [17].

3-Diphenylphosphino-4-methoxy-carbonylbenzoic acid NHS active ester (TP-NHS) was synthesized as literature method [18].

1H NMR spectra were recorded with Bruker 400 MHz spectrometer. In all cases, the sample concentration was 10 mg/mL, and the appropriate deuterated solvent was used with TMS as an internal standard. Dynamic Light Scattering (DLS) was recorded with 90

Plus particle size analyzer (BIC). Fluorescent spectrum was measured with FluoroMax-2

(ISA). UV-Vis Spectroscopy was recorded with Cary 50 UV-Vis spectrophotometer

(VARIAN).

2.2.2 Syntheses of DSPE–PEG2000–TP and Cholesterol-PEG2000–TP anchor lipids for chemoselective surface functionalization via Staudinger ligation

Figure 9. Synthesis of anchoring lipids Chol-PEG2000-TP and DSPE-PEG2000-TP.

2.2.2.1 Synthesis of cholesterol–PEG2000-triphenylphosphine (Chol-PEG2000-TP)

Et3N (14 µL, 0.1 mmol) was added to Monocholesteryl-PEG2000-amine (50 mg,

20.1 µmol) in anhydrous CH2Cl2 (4 mL) and was stirred for 30 min. at room temperature under Argon gas atmosphere, then TP-NHS (14 mg, 30.1 µmol, 1.5 equiv.) dissolved in anhydrous CH2Cl2 (1 mL) was added and the reaction solution was stirred overnight under darkness. TLC analysis was used to check the formation of Chol-PEG2000-TP

(visualized under UV lamp and Ninhydrin staining). The reaction mixture was concentrated by rotary evaporator and then the reaction mixture residual was separated by passing through Sephadex LH-20 column with MeOH as eluent. The fractions containing

Cholesterol-PEG2000-TP were collected and evaporated to dryness under vacuum (32 mg,

1 50.1% yield). H NMR (CDCl3, 400 MHz): 8.09 (dd, J = 8.0, 3.6 Hz, 1H), 7.81 (dd, J =

8.0, 1.4 Hz, 1H), 7.37-7.29 (m, 8H), 6.60 (m, 1H), 5.38 (m, 3H), 5.19 (m, 2H), 4.50 (m,

3H), 3.85 (m, 2H), 3.75 (s, 3H), 3.80-3.50 (br. m, 100H, O-CH2-CH2-O), 3.57-3.55 (m,

8H), 3.49-3.34 (m, 2H), 3.37-3.36 (m, 5H), 2.38 (dd, J = 7.8, 1.9 Hz, 2H), 2.23 (m, 2H),

2.04-1.80 (m, 7H), 1.89 (s, 3H), 1.62-0.95 (m, 25H), 1.02 (s, 6H), 0.93 (d, J = 6.6 Hz,

13 3H), 0.87 (d, J = 6.7 Hz, 6H) ppm, C NMR (CDCl3, 100 MHz) (δ): 166.70, 166.68,

166.38, 156.22, 139.88, 137.39, 137.28, 137.17, 134.00, 133.79, 132.79, 130.79, 128.96,

128.66, 128.58, 126.72, 122.42, 70.57, 56.69, 56.14, 52.17, 50.03, 42.31, 40.70, 39.86,

39.74, 39.5038.59, 37.00, 36.56, 36.18, 35.77, 31.88, 28.21, 28.19, 27.99, 24.27, 23.81,

31 21.03, 19.32, 18.71, 11.85 ppm, and P NMR (CDCl3, 170 MHz) (δ): -3.67 ppm.

2.2.2.2. Synthesis of DSPE-PEG2000-triphenylphosphine (DSPE-PEG2000–TP)

0.2 mL of Et3N was added to DSPE-PEG2000-NH2 (25 mg, 8.9 µmol) dissolved in

10 mL anhydrous CH2Cl2 and was stirred for 30 min at room temperature under Argon gas atmosphere. Then a solution of succinimidyl-3-diphenylphosphino-4- methoxycarbonylbenzoate (TP-NHS) (8 mg, 17.9 µmol) dissolved in 5 mL of anhydrous

CH2Cl2 was added to the above solution under Argon gas. The reaction mixture was further allowed to stir for 24 hr at room temperature under Argon gas and then the solution was concentrated under vacuum to give a residue. TLC analysis (chloroform: methanol 1:10, v/v, visualized under UV lamp and Ninhydrin staining) was used to check the reaction. The reaction mixture was concentrated by rotary evaporator and then the concentrated reaction mixture was separated by passing through Sephadex LH-20 column with MeOH as eluent. The fractions containing DSPE-PEG2000-TP were collected and

1 evaporated to dryness under vacuum (10 mg, 30.1 % yield). H NMR (CDCl3, 400 MHz)

(δ): 8.09 (dd, J = 8.0, 3.6 Hz, 1H), 7.81 (dd, J = 8.0, 1.4 Hz, 1H), 7.36-7.28 (m, 8H), 6.65

(m, 1H), 5.22 (m, 1H), 4.40 (m, 1H), 4.20-4.15 (m, 3H), 4.00-3.95 (m, 3H), 3.75 (s, 3H),

3.80-3.50 (br. m, 100H, O-CH2-CH2-O), 3.58-3.54 (m, 1H), 3.39-3.38 (m, 1H), 3.09-3.06

(m, 1H), 2.29 (m, 4H), 2.28 (br.s, 1H), 1.59 (br.s, 2H), 1.34 (t, J = 6.9 Hz, 2H), 1.66 (br.

13 s, 32H), 0.89 (t, J = 6.9 Hz, 6H) ppm, C NMR (CDCl3, 100 MHz) (δ): 173.38, 172.99,

166.70, 166.39, 156.54, 141.28, 137.39, 137.28, 137.17, 136.54, 134.79, 133.80, 132.81,

130.79, 128.96, 128.66, 128.59, 126.73, 71.57, 70.41, 70.25, 70.08, 69.61, 63.38, 62.63,

52.19,45.62, 39.86, 34.28, 34.10, 31.91, 29.70, 29.65, 29.52, 29.34, 29.17, 29.15, 24.92,

31 24.88, 22.67, 14.10 8.55 ppm, and P NMR (CDCl3, 170 MHz) (δ): -3.68 ppm.

2.2.3 Determination of critical micelle concentration (CMC) of the anchor lipids

A series of anchor lipid (DSPE-PEG2000-DBCO and Chol-PEG2000-DBCO) solutions, ranging from 1 µM to 100 µM, containing 1 µM pyrene were prepared [21].

Briefly, 100 µL of pyrene solution was added to diﬀerent vials and the solvent was evaporated, to which 500 µL of diﬀerent concentrations of the anchor lipids were added.

The solutions were stirred overnight at room temperature. Finally, the emission spectra at two wavelengths (I385 and I374) were recorded using a Hitachi FluoroMax-2 (ISA) spectrophotometer. The graphs were plotted by taking the intensity ratios of I385 and I374, which correspond to the third and first peaks of the pyrene molecule. The CMC values of the two anchor lipids were determined by taking the intersection of the straight lines of the graphs.

2.2.4 Liposome preparation

DPPC (30 mg, 40.9 µmol) and different ratios of DSPE-PEG2000-TP or

Cholesterol-PEG2000-TP (0.5%-2.0%) were dissolved in 3.0 mL anhydrous chloroform.

The solvent was slowly removed on a roto-evaporator under reduced pressure in order to form a thin lipid film on the flask wall that was further dried in vacuum chamber overnight. Then, the lipid film was swelled in the dark with 3 mL PBS buffer (pH 7.4) to form multilamellar vesicle suspension. It was further subjected to 10 freeze-thaw cycles involving quenching in liquid N2 and then immersed in a 65 °C water-bath. The crude lipid suspension thus formed was passed through an extruder with different

polycarbonate membranes (pore sizes 800 nm, 600 nm, 400 nm, 200 nm and finally 100 nm) to obtain the liposomes.

2.2.5 Conjugation of lactose onto liposomes surface

Azido-Lactose (0.5 mg) in 0.5 mL PBS (pH 7.4) buffer (Argon bubbled before use) was added into 1 mL of liposomes obtained as above. The Staudinger ligation reaction was conducted at room temperature for 6 h under Argon atmosphere, after which the unreacted azido-lactose was removed by gel filtration (1.5 x 20 cm column of

Sephadex G-75). The size of liposomes during the Staudinger ligation was monitored over time by using 90 Plus particle analyzer. Similarly, control experiments were conducted in the absence of azido-lactose.

2.2.6 Determination of concentration of lactose on the liposome surface

A standard curve was generated as described by Lactose Assay kit (Bioassay systems) with free lactose solution. To 20 µL of lactose-grafted liposome obtained above,

80 µL of reaction mix (85 µL Assay buffer+1 µL Lactase enzyme+1 µL dye reagent) was added and mixed. The mixture was kept on a shaker, and was allowed to stand for 30 min at room temperature. The optical readings were taken at 570 nm. The amount of lactose conjugated onto the liposome surface was thus calculated from the standard calibration curve. (Figure 10.)

Figure 10. Standard curve for lactose assay.

2.2.7 Characterization of specific binding between lactose on the liposome surface and lectin

To study the accessibility of lactose grafted on to the liposome surface, 100 µL lectin (β-galactose binding lectin, Arachis hypogae, 120 kDa, Sigma) in PBS buffer solution (1 mg/mL, pH 7.4) was added into 100 µL solution of lactose grafted liposomes.

The stability and size of the liposomes were monitored with DLS over time. Control was conducted with liposome without lactose in the same manner above.

2.2.8 5, 6-Carboxyfluorescein (5, 6-CF) encapsulation efficiency

5, 6-CF-encapsulated liposomes were prepared in the same conditions as above along with 85 mM 5, 6-CF. Separation of the 5, 6-CF encapsulated liposomes from the non-encapsulated 5, 6-CF dye was achieved by size-exclusion chromatography that involved passage through a 1.5 x 20 cm column of Sephadex G-75. The 5, 6-CF encapsulation efficiency (EE %) was determined by measuring the amount of encapsulated dye relative to the total initial amount, using FluoroMax-2 (ISA). The free dye was removed from the dye encapsulated liposomes by passing through a 1.5 x 20 cm column of Sephadex G-75. The amount of encapsulated dye was determined after disrupting the liposome using 5 µL Triton-X 1%. The readings were obtained by adding

20 µL of reaction solution + 1980 µL PBS buffer (pH 7.4). The EE% is calculated by using the formula (Figure 11.)

Where V and C are volume and concentration and i represents initial values taken from the original preparation and f represents values taken from the encapsulated dye liposomes.

Figure 11. Encapsulation efficiency of 5, 6-carboxyfluorescein dye.

2.2.9 5, 6-CF dye leakage assay

5, 6-CF-encapsulated liposomes were prepared in the same conditions as above along with 85 mM 5, 6-CF. The fluorescent leakage was measured using FluoroMax-2

(ISA) to monitor the stability of liposomes. Briefly, 20 µL reaction solution and 1980 µL

PBS buffer (pH 7.4) were mixed together and then the fluorescent intensity was measured. 5 µL solution of Triton-X 1% was added to obtain the total amount of encapsulated dye present in the liposome, from which the % leakage of 5, 6-CF dye was determined. Similarly, the control experiments were conducted with the plain liposome without lactose modification.

2.3 Results and discussion

The aim of this study was to investigate the anchoring lipid effects on liposome stability, ligand grafting density and liposome chemical and physical characteristics upon liposome surface glyco-functionalization and their lectin binding activity. In this study, two anchoring lipids namely Chol-PEG2000-TP and DSPE-PEG2000-TP were proposed for liposome surface glyco-functionalization with azide derivative of lactose as a model carbohydrate via Staudinger ligation. The major difference between these two anchoring lipids is their respective hydrophobic molecules inserted in the lipid bilayer of liposomes, a sterol in the case of Chol-PEG, while a phospholipid with long saturated fatty acid chains in the case of DSPE-PEG. In addition, sterol is a neutral molecule which stabilizes liposomes and prevents liposome aggregation, while phospholipid imparts negative charge to the liposome surface, which may lead to additional binding interactions with

plasma proteins or the drugs encapsulated and released [16]. Therefore, it is expected that these anchoring lipids will have impact on both the chemistry upon liposome surface modification and their chemical and physical characteristics, in vitro and in vivo behavior as well.

2.3.1 Syntheses and characterization of DSPE–PEG2000–TP and Cholesterol-

PEG2000–TP anchor lipids

First, the terminal triphenylphosphine carrying anchoring lipids were synthesized by amidation of synthetic Chol-PEG2000-NH2 [17] and commercially available DSPE-

PEG-NH2 (Avanti Polar Lipids) with 3-diphenylphosphino-4-methoxycarbonylbenzoic acid NHS active ester [18] in good yield, respectively (Figure 9). The resultant anchoring lipids were characterized by 1H, 13C and 31P NMR spectra (Figure 12). As previously reported, the triphenylphosphine is air sensitive, which is the drawback of Staudinger ligation [15]. However, there is no oxidized product formed for both Chol-PEG2000-TP

31 and DSPE-PEG2000-TP after purification in the present study, as shown in P NMR spectra, in which the phosphine in both compounds gave a chemical shift at -3.74 ppm

(Figure 12 A and 12 B).

Figure 12. NMR spectra of Chol-PEG2000-TP (A) and DSPE-PEG2000-TP (B) in CDCl3.

Next, the azido-reactive liposomes composed of saturated phospholipid DPPC and the anchoring lipid in different lipid ratios (see Table 3) were prepared by thin-film hydration and extrusion through polycarbonate membranes with pore size of 800 nm, 600 nm, 400 nm, 200 nm, and 100 nm sequentially at 65 °C. This produced predominantly small unilamellar vesicles, which displayed different average mean diameters of the liposomes of different lipids used, which were confirmed by DLS. A liposome with Chol-

PEG2000-TP anchoring lipid is relatively larger than liposome with DSPE-PEG2000-TP anchoring lipid in the same percentage in the liposomes (Table 3). Glyco-surface modification of the preformed liposomes with lactosyl azide [19] as a model was performed in PBS buffer (pH 7.4) at room temperature under an argon atmosphere for 6 hours (Figure 8).

2.3.2 Characterization and evaluation of the stability of glyco-functionalized liposomes via dynamic light scattering technique (DLS) and fluorescence spectroscopy

DLS was used to verify the integrity of the vesicles during and after the coupling reaction. As a result, there was 10 to 20 nm size increase in the average mean diameter for all liposomes of different lipids and anchoring lipids (Table 3). There was no liposome aggregate formation during and after the conjugation reaction as monitored by

DLS (Figure 13, liposomes with 1% anchoring lipid). Therefore, the reaction conditions described above do not alter the integrity of the liposomes. The continued stability of the glycosylated liposomes were investigated by DLS as well. As shown in Figure 13

(liposomes with 1% anchoring lipid), the glycosylated liposomes with either Chol-PEG or DSPE-PEG anchoring lipids showed good stability during the 14 days period.

However, the liposome with and without anchoring lipids began to collapse and aggregate since 7th and 5th day, respectively (Figure 13). There is no apparent difference between the two kinds of anchoring lipids at different concentrations. These results demonstrated that the presence of lactose on the liposome surface provides a steric barrier that prevents liposome aggregation.

Table 3. Liposomes with different anchoring lipids and their sizes determined by DLS.

Mean Diameter (nm)

Anchoring lipid/DPPC Before Conjugation After Conjugation

0.5% 96 102

1.0% 122 129

DSPE-PEG2000-TP 1.5% 145 156

2.0% 168 186

0.5% 100 110

CHL-PEG2000-TP 1.0% 120 123

1.5% 149 161

2.0% 174 195

Figure 13. DLS monitoring of liposomes before and after glyco-functionalization: Chol- PEG2000-TP anchored liposome (A) and DSPE-PEG2000-TP anchored liposome (B) and their stabilities monitored with DLS.

2.3.3 Measurement of critical micelle concentration (CMC) of anchor lipids before and after glyco-functionalization

The PEG linkers also stabilize the liposomes as well since PEG polymers are often used to stabilize liposomes [20]. In addition, critical micelle concentration (CMC) was measured for these two anchor lipids by using the pyrene assay [21]. As a result,

Chol-PEG2000-TP had a 3.4 times higher critical micelle concentration (CMC) value than that of DSPE-PEG2000-TP and conjugation with lactose resulted in only 1.1 and 1.4 times increments in CMC for Chol-PEG2000-TP and DSPE-PEG2000-TP, respectively, as CMC is strongly dependent on the alkyl chain length (hydrophobicity) (Table 4). (Figure 14.)

Table 4. Critical micelle concentration (CMC) determination of anchor lipids Cholesterol–PEG2000–TP (A) and DSPE-PEG2000-TP (B).

CMC (µM)

Anchor lipid Before Conjugation After Conjugation

DSPE-PEG2000 –TP ~8 µM ~11 µM

CHL-PEG2000-TP ~27 µM ~31 µM

Figure 14. Critical micelle concentration (CMC) determination of anchor lipids Cholesterol–PEG2000–TP (A) and DSPE-PEG2000-TP (B).

The drug encapsulation capacity and releasing property of liposomes over time are important factors for its biomedical applications, especially in drug/gene delivery application. Therefore, in our study, both DSPE and Chol-based anchoring lipids were investigated consideration the drug encapsulation capacity and releasing properties of glycosylated liposomes. The same liposomes with 1% anchoring lipid and having encapsulated self-quenching concentrations of 5, 6-carboxyfluorescein (5, 6-CF, 85 mM) were used as a model compound to determine encapsulation capacity and to monitor the releasing kinetics over a two week period. Briefly, 5,6-CF-encapsulated liposomes containing TP were prepared under the same conditions as above along with 85 mM 5,6-

CF, followed by glycosylation with lactosyl azide and separation from unreacted lactosyl azide and the non-encapsulated 5,6-CF dye. As a result, encapsulation efficiency of conventional liposome (DPPC:Chol (2:1) was found to be 2.25% ± 0.05 while that of

Chol-PEG and DSPE-PEG containing liposomes were found to be 1.77% ± 0.05 and

1.98% ± 0.02, respectively. These decreases in encapsulation efficiency by incorporation of Chol-PEG and DSPE-PEG are probably due to the reduction in internal vesicular volume by bulky PEG chains covering both the inner and outer surfaces of liposomes.

The results are in agreement with previous reports by Schneider et al. who observed concentration dependent reduction in encapsulation efficiencies of water soluble contrast agents in liposomes containing different concentrations of cholesteryl hemisuccinate and distearoyl phosphatidylethanolamine [22].

Next, the releasing behaviors of glycosylated liposomes with Chol-PEG and

DSPE-PEG anchoring lipid and plain liposome of DPPC and Cholesterol (2:1) were investigated. As shown in (Figure 15 A), the cumulative release of 5, 6-CF in plain

liposomes was 5% daily and sped up after 5 days due to its instability, while, 5, 6-CF releasing was consistent in a reduced rate (1% daily) for two weeks in the liposomes with the presence of Chol–PEG in 0.5, 1.0, 1.5, and 2%. Also, liposomes with the DSPE-PEG anchoring lipid in 0.5, 1.0 and 1.5% released 5, 6-CF at a consistently reduced rate (less

1% daily) for two weeks. However, liposome with the DSPE-PEG anchoring lipid in

2.0% has the same 5, 6-CF releasing pattern of plain liposome (5% daily and sped up after 5 days). Overall, Chol–PEG anchored liposome showed relatively faster releasing pattern (Figure 15 A) than DSPE-PEG anchored liposomes (Figure 15 B). The difference could be explained by the fact that Chol increases rigidity of the lipid membrane and thus leads to fast leakage of liposomes [23].

Figure 15. 5, 6-CF releasing kinetics of lactosylated liposomes with anchoring lipid in different ratios: Cholesterol–PEG2000-TP (A) and DSPE–PEG2000-TP (B).

2.3.4 Quantification of lactose on the surface of liposome

Furthermore, the grafted lactose on the surface of liposome was quantified by using the Lactose Assay kit [24]. Briefly, a reaction mix (85 µL Assay buffer+1 µL

Lactase enzyme + 1 µL dye reagent) was added to samples containing lactose and were incubated for 30 min at room temperature. The optical density readings were obtained at

570 nm using UV-Vis Spectroscopy. As a result, 90% glyco-functionalization yield was determined for Chol-PEG2000-TP anchored liposome, while 35% yield was obtained for

DSPE-PEG2000-TP anchored liposome with 1% anchoring lipids in both liposome preparations. The reason for this lower coupling efficiency is unknown; however, it might be due to the incorporation of less phospholipid anchoring lipid in the outer leaf surface of the reactive liposome.

2.3.5 Accessibility of surface coated lactose molecules on liposome formulations via lectin binding assay

To determine whether the grafted lactose residues are easily accessible at the surface of liposomes, lectin binding assay was conducted by incubating lactosylated liposome in the presence of β-galactose binding lectin (Arachis hypogae, 120 kDa,

SIGMA) in PBS (pH 7.4). As a result, apparent visible aggregation was observed and was monitored in DLS experiment for glycosylated liposome with Chol-PEG anchoring lipid after 25 minutes (Figure 16 A), while, it took 75 minutes (Figure 16 B) to observe the similar apparent visible aggregation that was also monitored in DLS experiment for glycosylated liposome with DSPE-PEG anchoring lipid. In contrast, neither aggregation

nor size change was observed with control liposomes without lactose. Furthermore, the presence of free lactose (5.0 mM) prevented aggregates formation for both glycosylated liposomes, confirming that the agglutination was due to a specific recognition of the lactose residues on the surface of the liposomes by lectin. The lectin binding difference might be due to the lactose density on the liposome surface, in which multivalent interactions contribute to the lectin induced glycosylated liposome aggregation.

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Figure 16. DLS monitoring of agglutination due to multivalent lectin binding of lactosylated liposomes with anchor lipid of Cholesterol–PEG2000-TP (A) and DSPE– PEG2000-TP (B).

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2.4 Conclusion

In this study, we have developed an efficient and chemoselective conjugation method for liposome surface glyco-functionalization based on two kinds of anchoring lipid via Staudinger ligation. The reaction could be performed under mild conditions in aqueous buffers without catalyst and in high yields under reasonable reaction times. The reaction conditions developed in the present work did not alter the integrity of the bilayers, in terms of liposome sizes and leakiness, and provided perfectly functional vesicles. In this study, the anchoring lipid effect on liposome size, stability, encapsulation capacity, and glycosylation efficiency and their lectin binding property were investigated.

This versatile approach, which is particularly suitable for the ligation of water soluble molecules and which can accommodate many chemical functions, is anticipated to be useful in the coupling of many other ligands onto liposomes for a wide range of applications.

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2.5 References

[1] J. Philippot, and F. Schuber (Eds.), Liposomes as Tools in Basic Research and

Industry, CRC Press, Boca Raton, 1995.

[2] T. M. Allen, and P. R. Cullis, "Drug Delivery Systems: Entering the Mainstream,"

Science, vol. 303, no. 5665, pp. 1818-1822, 2004.

[3] V. P. Torchilin, "Recent advances with liposomes as pharmaceutical carriers,"

Nature Reviews Drug Discovery, vol. 4, pp. 145-160, 2005.

[4] T. M. Allen, C. Hansen, F. Martin, C. Redemann, A. Yau-Young, "Liposomes

containing synthetic lipid derivatives of poly(ethylene glycol) show prolonged

circulation half-lives in vivo," Biochimica et Biophysica Acta, 1066, pp. 29-36,

1991.

[5] H. Lis, N. Sharon, Lectins: "Carbohydrate-specific proteins that mediate cellular

recognition," Chemical Reviews, vol. 98, no. 2, pp. 637-674, 1998.

[6] H. Zhang, Y. Ma, X.-L. Sun, "Recent developments in carbohydrate-decorated

targeted drug/gene delivery," Medicinal Research Reviews, vol. 30, no. 2, pp.

270-289, 2010.

[7] X.-L. Sun, Y. Kanie, C.-T. Guo, O. Kanie, Y. Suzuki, and C.-H. Wong,

"Syntheses of C-3 modified sialylglycosides as selective inhibitors of influenza

hemagglutinin and neuraminidase," European Journal of Organic Chemistry, vol.

2000, no. 14, pp. 2643-2653, 2000.

[8] P. Sapra, and T. M. Allen, "Ligand-targeted liposomal anticancer drugs,"

Progress in Lipid Research, vol. 42, no. 5, pp. 439-462, 2003.

103

[9] L. Nobs, F. Buchegger, R. Gurny, and E. Allemann, "Current methods for

attaching targeting ligands to lipoosmes and nanoparticles," Journal of

Pharmaceutical Sciences, vol. 93, no. 8, pp. 1980-1992, 2004.

[10] V. T. Kung, and C. T. Redemann, "Synthesis of carboxyacyl derivatives of

phosphatidylethanolamine and use as an efficient method for conjugation of

protein to liposomes," Biochimica et Biophysica Acta, vol. 862, no. 2, pp. 435-

439, 1986.

[11] P. Schelte, C. Boeckler, B. Frisch, F. Schuber, "Differential reactivity of

maleimide and bromoacetyl functions with thiols: application to the preparation of

liposomeal diepitope constructs," Bioconjugate Chemistry, vol. 11, no. 1, pp.118-

123, 2000.

[12] Y. Nakano, M. Mori, S. Nishinohara, Y. Takita, S. Naito, H. Kato, M. Taneichi,

K. Komuro, T. Uchida, "Surface-linked liposomal antigen induces IgE-selective

unresponsiveness regardless of the lipid components of liposomes," Bioconjugate

Chemistry, vol. 12, no. 3, pp. 391-395, 2001.

[13] L. Bourel-Bonnet, E. I. Pecheur, C. Grandjean, A. Blanpain, T. Baust, O. Melnyk,

B. Hoflack, and H. Gras-Masse, "Anchorage of synthetic peptides onto liposomes

via hydrazone and α-oxo hydrazone bonds. Preliminary functional investigators,"

Bioconjugate Chemistry, vol. 16, no. 2, 450-457, 2005.

[14] F. Said Hassane, B. Frisch, F. Schuber, "Targeted liposomes: convenient coupling

of ligands to preformed vesicles using “Click Chemistry”," Bioconjugate

Chemistry, vol. 17, no. 3, pp. 849-854, 2006.

104

[15] H. Zhang, Y. Ma, X.-L. Sun, "Chemically-selective surface glyco-

functionalization of liposomes through Staudinger ligation," Chemical

Communications, no. 21 , pp. 3032-3034, 2009.

[16] F. T. Presti, R. J. Pace, S. I. Chen, "Cholesterol-phospholipid interaction in

membranes. 2. Stoichiometry and molecular packing of cholesterol-rich

domains," Biochemistry, vol. 21, no. 16, pp. 3831-3835, 1982.

[17] X. Pan, G. Xu, W. Yang, R. F. Barth, W. Tjarks, R. J. Lee, "Synthesis of

cetuximab-immunoliposomes via a cholesterol-based membrane anchor for

targeting of EGFR," Bioconjugate Chemistry, vol. 18, no. 1, pp. 101-108, 2007.

[18] C. C.-Y. Wang, T. S. Seo, Z. Li, H. Ruparel, J. Ju, "Site-specific fluorescent

labeling of DNA using Staudinger ligation," Bioconjugate Chemistry, vol. 14, no.

3, pp. 697-701, 2003.

[19] X.-L. Sun, D. Grande, S. Baskaran, E. L. Chaikof, "Glycosaminoglycan mimetic

biomaterials.4. Synthesis of sulfated lactose-based glycopolymers that exhibit

anticoagulant activity," Biomacromolecules, vol. 3, no.5, pp. 1065-1070, 2002.

[20] M. L. Immordino, F. Dosio and L. Cattel, "Stealth liposomes: review of the basic

science, rationale, and clinical applications, existing and potential," International

Journal of Nanomedicine, vol. 1, no. 3, 297–315, 2006.

[21] S. Song, L.-Z. Cheong, M. Falkeborg, L. Liu, M. Dong, H. M. Jensen, K.

Bertelsen, M. Thorsen, T. Tan, X. Xu and Z. Guo, "Facile Synthesis of

Phosphatidyl Saccharides for Preparation of Anionic Nanoliposomes with

Enhanced Stability," PLoS One, vol. 8, no. 9, e73891, 2013.

105

[22] T. Schneider, A. Sachse, J. Leike, G. Roβling , M. Schmidtgen, M. Drechsler, M.

Brandl, "Surface modification of continuously extruded contrast carrying

liposomes: effect on their physical properties," International Journal of

Pharmaceutics, vol. 132, no. 1-2, pp. 9–21, 1996.

[23] S. C. Semple, A. Chonn, P. R. Cullis, "Influence of Cholesterol on the

Association of Plasma Proteins with Liposomes," Biochemistry, vol. 35, no. 8,

2521–2525, 1996.

[24] B. Priem, M. Gilbert, W. W. Wakarchuk, A. Heyraud, E. Samain, "A new

fermentation process allows large-scale production of human milk

oligosaccharides by metabolically engineered bactteria," Glycobiology, vol. 12,

no.4, pp. 235-240, 2002.

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CHAPTER III

CELL SURFACE RE-ENGINEERING VIA EFFICIENT LIPID ANCHORING

APPROACH

3.1 Introduction

The cell membrane mainly comprises of a bilayer of amphiphillic phospholipids and cholesterol molecules. The cell surface is decorated with many proteins and carbohydrates that aid in many biological recognition processes, thus making the cell membrane an important interface for extracellular and intracellular communications.

Surface re-engineering of cell membranes with biologically important molecules has scope for potential applications such as cell labeling [1], glycan trafficking [2], efficient carrier systems for drug/gene/antigen delivery [3, 4], and in vivo imaging [2]. Due to the growing importance for cell surface re-engineering and its promising applications, development of different types of strategies or approaches is very necessary. In recent years, many approaches such as direct chemical modification [5], membrane fusion [6],

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and metabolic engineering methods [7] have been widely used for the introduction of bioactive molecules into the surface of the cell membrane. Antigen presenting cells

(APC’s) such as dendritic cells and macrophages play a pivotal role in innate immunity and apoptotic cell clearance and most importantly in antigen processing and its presentation on cell surface. Due to their key role during inflammation, tissue injury, repair processes and in immune system, macrophages can serve as important drug delivery targets [8] or as drug delivery vehicles [9, 4] for treatment of many disease conditions [10]. Recently, macrophages have been explored widely as drug/antigen delivery targets [11, 12, 13, 14], drug delivery carrier systems [15, 16] and also in transplantation/grafting applications [17, 4]. However, macrophages or other APC’s are considered as difficult targets especially, when intracellular delivery of active substances is required. In this study, we proposed a surface re-engineering strategy of macrophages aimed to bringing biomolecules directly onto the cell surface and paving path for potential biomedical applications.

Cell surface modifications have been successfully achieved by using derivatives of phospholipids [18, 19], cholesterol [20], oleyl [21], stearyl [22] as lipid anchors.

Furthermore, the use of PEGylated lipid derivatives for surface re-engineering of cells has been extensively investigated for achieving increased lifetime and biocompatibility of the biomolecules in vivo [19, 23, 24, 25]. Similarly, liposome systems have also been effectively employed for cell surface modifications [26]. In addition, they have been extensively explored for targeted delivery applications, especially for drug [27], antigen

[13], and gene [28] delivery purposes. Targeting of surface decorated liposomes to their

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respective receptors on cells can be achieved by conjugation of liposomes with antigens, antibodies, carbohydrates and recombinant proteins [12, 14].

In the present study, we have developed a simple and efficient method for cell surface re-engineering of RAW 264.7 macrophage cells as a model macrophage cell

(Figure 19). Two types of lipid anchors namely; phospholipid (DSPE-PEG2000-DBCO) and cholesterol (CHOL-PEG2000-DBCO) derivatives were synthesized and investigated for their effectiveness in surface modification of macrophages. The incorporation efficiency of both anchor lipids and their respective liposomal forms onto the cell surface were explored at different concentrations, incubation times and at 37 °C, in PBS buffer, pH 7.4 and were determined via confocal microscopy and flow cytometry analysis.

Further, biotinylation of the anchored phospholipid and cholesterol anchor lipids was achieved by incubation of with azide-biotin biomolecule for 1 hr via copper-free click chemistry. This facilitates for further labeling of the cells with streptavidin-FITC via the strong binding interactions between biotin and streptavidin molecules. Overall, when compared to DSPE lipid, cholesterol anchor lipid could rapidly anchor into the cell membranes without any negative effects and cytotoxicity. MTT assays were performed to evaluate the cytotoxic effects of the two types of anchor lipids.

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Figure 17. Schematic illustration of cell surface re-engineering via Chol-PEG2000–DBCO and DSPE-PEG2000-DBCO lipid anchoring approach.

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3.2 Experimental

3.2.1 Materials and methods

All solvents and reagents were purchased from commercial sources and were used as received, unless otherwise noted. Deionized water was used as a solvent in all procedures. The synthesis of monocholesteryl-PEG2000-amine was performed using a previously described method [29]. N-hydroxysuccinimide (NHS-ester) and Cholesteryl chloroformate were purchased from Sigma-Aldrich (St. Louis, MO). 1, 2-dipalmitoyl-sn- glycero-3-phosphocholine (DPPC), 1, 2-distearoyl-sn-glycero-3-phosphoethanolamine-

N-[amino (polyethylene glycol)-2000] (ammonium salt) (DSPE-PEG2000-NH2) were purchased from Avanti Polar Lipids, Inc (Alabaster, AL). DBCO-PEG4-NHS ester was purchased from Click Chemistry tools (Scottsdale, AZ). N3-biotin, Streptavidin- fluorescein isothiocyanate (Streptavidin-FITC) was purchased from Biolegend (San

Diego, CA), Sodium dodecyl sulfate (SDS) was purchased from Sigma-Aldrich (St.

Louis, MO). Dulbecco’s modified Eagle medium (DMEM) medium, (3-(4, 5- dimethylthiazol-2-yl)-2, 5-diphenyltetrazolium bromide) MTT reagent were purchased from Life Technologies (Grand Island, NY).

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3.2.2 Syntheses of DSPE–PEG2000–DBCO and Cholesterol-PEG2000–DBCO anchor lipids for chemoselective surface functionalization via copper free click chemistry

Figure 18. Schemes for the syntheses of Cholesterol–PEG2000–DBCO and DSPE-

PEG2000–DBCO anchor lipids for cell surface re-engineering applications.

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3.2.2.1 Synthesis of Chol-PEG2000–DBCO

Triethylamine, Et3N (50 µL, 0.35 mmol) was added to Monocholesteryl-PEG2000- amine (100 mg, 40.4 µmol) in anhydrous CH2Cl2 (3 mL) and was stirred for 30 min at room temperature under argon gas atmosphere, then to the above mixture DBCO-PEG4-

NHS ester (31 mg, 48.48 µmol, 1.2 equiv.) dissolved in anhydrous CH2Cl2 (1 mL) was added and the reaction solution was stirred for 48 hrs at RT. TLC analysis was performed to monitor the reaction. The pure Chol-PEG2000-PEG4-DBCO fractions were collected by passing through silica gel column and eluted with a gradient solvent system (CHCl3:

MeOH- 95:5~80:20 v/v). The fractions containing Cholesterol-PEG2000-PEG4-DBCO were collected and evaporated to dryness under vacuum (92 mg, 70.22 % yield). (1H

NMR, CDCl3, 600 MHz) (:δ): 7.65 (d, 1H), 7.40-7.28 (d, 8H), 6.53 (br, 1H), 5.35 (br,

1H), 5.22 (br, 1H), 5.14 (d, 1H), 4.46 (br, 1H), 3.81 (t, 3H), 3.75-3.69 (m, 3H), 3.76-3.40

(m, 180 H), 3.36-3.27 (m, 4H), 3.26-3.21(m, 2H), 2.87 (t, 3H), 2.80 (br, 6H), 2.60-2.20

(m, 10 H), 2.00-1.75 (m, 10 H), 1.50-0.95 (m, 30 H), 0.89 (d, 3H), 0.84 (dd, 6H), 0.65 (d,

13 3H) ppm, ( C NMR, CDCl3, 100 MHz): (:δ): 169.62, 168.69, 166.58, 164.36, 138.5,

129,75, 126.73, 126.22, 125.96, 125.89, 125.46, 124.84, 123.23, 120.16, 119.33, 105.41,

69.42, 68.34, 68.20, 68.11, 67.87, 63.36, 54.33, 53.12, 47.65, 39.95, 37.15, 34.52, 32.39,

29.79, 29.54, 29.29, 23.22, 20.46, 20.20, 16.35, 9.50, 9.38 ppm.

3.2.2.2 Synthesis of DSPE-PEG2000–DBCO

DSPE-PEG2000–PEG4-DBCO was synthesized by following the method from previous literature [30]. Briefly, Triethylamine, Et3N (50 µL, 0.35 mmol) was added to

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DSPE-PEG2000-amine (95 mg, 33.6 µmol) in anhydrous CH2Cl2 (3 mL) and was stirred for 30 min at room temperature under argon gas atmosphere, then to the above mixture

DBCO-PEG4-NHS ester (23.30 mg, 33.6 µmol, 1.0 equiv.) dissolved in anhydrous

CH2Cl2 (1 mL) was added via syringe drop wise slowly. The reaction mixture was stirred for 48 hrs at RT, under Argon gas atmosphere. TLC analysis was performed to check for the product formation (CHCl3 : MeOH, 10:1). The pure DSPE-PEG2000-PEG4-DBCO fractions were collected by passing through silica gel column column chromatography and eluted with (CHCl3 : MeOH, 10:1 v/v) solvent system. The fractions containing pure

DSPE-PEG2000-PEG4-DBCO were collected and evaporated to dryness under vacuum (78

1 mg, 71.55% yield). ). ( H NMR, CDCl3, 400 MHz) (:δ): 7.75-7.3 (d, 8H), 6.71 (br, 1H),

6.57 (br, 1H), 5.22 (s, 1H), 5.15 (d, 1H), 4.40 (d, 2H), 4.24 (br, 2H), 4.18 (t, 2H), 3.98

(br, 4H), 3.85-3,28 (m, 173 H), 2.48 (t, 2H), 2.49-2.27 (m, 4H), 1.27 (s, 47 H), 0.90 (t,

13 6H) ppm, ( C NMR, CDCl3, 100 MHz) (:δ):).173.33, 172.95, 148. 08, 132.13, 129.08,

128.57, 128.31, 128.24, 127.19, 125.56, 123.04, 70.57, 70.26, 70. 21, 69.80, 67.38, 39.16,

34.24, 34.06, 31.89, 29.69, 29.63, 29.55, 29.30, 29.35, 29.32, 29.19, 29.17 ppm.

3.2.3 Determination of critical micellar concentration (CMC) of the anchor lipids

A series of anchor lipid (DSPE-PEG2000-DBCO and Chol-PEG2000-DBCO) solutions, ranging from 1 µM to 100 µM, containing 1 µM pyrene were prepared [31].

Briefly, 100 µL of pyrene solution was added to diﬀerent vials and the solvent was evaporated, to which 500 µL of diﬀerent concentrations of the anchor lipids were added.

The solutions were stirred overnight at room temperature. Finally, the emission spectra at

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two wavelengths (I385 and I374) were recorded using a Hitachi FluoroMax-2 (ISA) spectrophotometer. The graphs were plotted by taking the intensity ratios of I385 and I374, which correspond to the third and first peaks of the pyrene molecule. The CMC values of the two anchor lipids were determined by taking the intersection of the straight lines of the graphs.

3.2.4 Cell culture

The mouse macrophage cell line RAW264.7 cells were used in our study. The cells were cultured in DMEM medium supplemented with 10% fetal bovine serum, penicillin (100 units / ml) and streptomycin (100 µg / ml) at 37 °C in 5% CO2 conditions.

The cells were further sub cultured by trypsinization with (0.25% trypsin, 0.02% EDTA) after cells became confluent.

3.2.5 Preparation of aqueous solutions of DSPE–PEG2000–DBCO and Cholesterol-

PEG2000–DBCO anchor lipids

Stock solutions of 1 mg/1 mL anchor lipids in CHCl3 were prepared to obtain a thin film, by evaporation under a stream of N2 gas. The thin film was further put under vacuum for overnight in order to ensure complete removal of any residual solvent. The thin film was hydrated with PBS buffer, pH 7.4 with vigorous sonication and vortexing for about 2-3 hrs to form aqueous solutions. Serial dilutions from the above solution resulted in 5 µM, 10 µM aqueous solutions of anchor lipids for further studies.

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3.2.6 Cell surface re-engineering of RAW 264.7 cells with DSPE and CHOL based anchor lipids via copper-free click chemistry

3.2.6.1 Two-step method

The RAW 264.7 cells were washed twice with PBS buffer and incubated with different concentrations (5µM, 10µM) of Chol-PEG2000- DBCO & DSPE-PEG2000-

DBCO anchor lipids for different incubation times (5 min, 10 mins, 20 mins) in PBS buffer, pH 7.4 at 37 °C. After further washing twice with PBS buffer, the cells were treated with 750 µL N3-Biotin (thrice the concentration of lipids) (50mM stock solution,

PBS, pH 7.4) for 1 hr at 37 °C. After 1 hr, the cells were washed again with PBS buffer and incubated with 750 µL FITC-streptavidin (1:800 dilution, 0.5 mg/ mL stock solution) for 5 mins, in darkness. The treated cells were washed, fixed by addition of 4% PFE solution and were set aside for 10 mins, gently aspirated, washed 2-3 times with PBS buffer and finally mounted onto glass slides for viewing under a confocal laser scanning microscope.

3.2.6.2 Direct/one-pot method

3.2.6.2.1 Syntheses of lipid-biotin conjugate via copper-free click chemistry

To the above mentioned aqueous lipid solutions, twice the concentration of N3-

Biotin (50 mM stock solution, PBS, pH 7.4) was added and the reaction was carried out in eppendorf tubes at room temperature for 1 hr. The synthesized lipid-biotin conjugates

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were directly incubated with the RAW 264.7 cells for different incubation times as mentioned below.

3.2.6.2.2 Anchoring of biotinylated lipid conjugates into the cell membranes

The RAW 264.7 cells were incubated with different concentrations of biotinylated lipid conjugates (5 µM, 10 µM) at different incubation times (5 mins, 10 mins, 20 mins) in PBS buffer, pH 7.4 at 37 °C. After each time point the medium was gently aspirated, followed by washing 2-3 times with PBS buffer, pH 7.4. The treated cells were further incubated with 750 µL FITC-Streptavidin (1:800 dilution, 0.5 mg/ mL stock solution) for

5 mins, in darkness and finally mounted onto glass slides for viewing under a confocal laser scanning microscope in a similar manner as above.

3.2.7 Liposome preparation

DPPC (30 mg, 40.87 µmol), and different concentrations (5 µM, 10 µM, 25 µM) of DSPE-PEG2000-DBCO & Cholesterol-PEG2000-DBCO were dissolved in 3.0ml chloroform. The solvent was slowly removed on a roto-evaporator under reduced pressure in order to form a thin lipid film on the flask wall and further dried in vacuum chamber overnight. Then, the lipid film was swelled in the dark with 3 ml PBS buffer

(pH 7.4) to form multilamellar vesicle suspension. It was further subjected to 10 freeze- thaw cycles involving quenching in liquid N2 and then immersed in a 65 °C water-bath.

The crude lipid suspension thus formed was sent through an extruder with different

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polycarbonate membranes (pore sizes 800 nm, 600 nm, 400 nm, 200 nm and finally 100 nm) to obtain the liposomes. The resultant liposomal solutions were also reacted in a similar manner with twice the concentration of N3-Biotin (50 mM stock solution, PBS, pH 7.4) and the reaction was carried out in eppendorf tubes at room temperature for 1 hr.

Similar procedures were used to incubate the biotinylated liposomal solutions with RAW

264.7 cells, but at different incubation times (20 mins, 2 hr, and 4 hr). After, each time point the treated cells were washed with PBS buffer, pH 7.4, further incubated with streptavidin-FITC and finally analyzed under confocal laser scanning microscope.

3.2.8 Cell surface labeling of biotinylated lipid conjugates on RAW 264.7 cells with streptavidin-FITC and imaging by confocal laser scanning microscopy

RAW 264.7 cells with the incorporated biotinylated lipid conjugates were incubated with 750 µL streptavidin-FITC (1:800 dilution from 0.5 mg/ mL stock solution, PBS buffer, pH 7.4) for 5 mins in darkness. The cell nuclei were stained by adding 3 µL of 4’, 6-diamidino -2- phenylindole (DAPI) solution. The buffer medium was gently aspirated, followed by washing 2-3 times with PBS buffer, pH 7.4 between each step. The cover slips were mounted onto glass slides using Anti-fade gold mounting medium (Life Technologies). The sections were imaged using a Nikon A1RSI Confocal

Microscope with a 60 x oil objective.

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3.2.9 Flow cytometry analysis of streptavidin-FITC labeled biotinylated–lipid anchored cells

Raw 264.7 cells were (1 x 106 cells/test tube) cultured overnight were washed 2-3 times with PBS buffer via centrifugation (1200 rpm, 5 mins) and treated with different concentrations (500 µL /test tube, 5, 10 and 25 µM) of biotinylated anchor lipids in PBS buffer medium and further incubated for different time intervals (5 mins, 10 mins, 20 mins) at 37 °C. The cells were once again washed twice with PBS buffer via centrifugation and then incubated with Streptavidin-FITC for 5 mins at room temperature, in darkness. Similarly, cells were treated with (5 µM, 10 µM and 25 µM) biotinylated liposomal solutions for different incubation times (20 min, 2 hr, and 4 hr) at

37 °C. After further washing steps, the fluorescence signal of 20 x 103 cells were measured using a flow cytometer (BD-Canto).

3.2.10 In vitro cytotoxic effect of the biotinylated lipid conjugates: MTT assay

Cell viability was carried out using MTT assay (Vybrant® MTT Cell

Proliferation Assay Kit). In this experiment, RAW264.7 were seeded at a density of 2.5×

104 cells per well in 96-well plates overnight, then the completed medium was removed, after which the cells were washed with PBS buffer, pH 7.4. The effect on cell viability during the biotinylation process was assessed by addition of FBS-free culture medium with compounds to different wells and incubated for different time points (20 mins, 16 hr,

24 hr) at 37 °C. In addition, to investigate the cell viability after biotinylation, the cells were incubated for a period of 20 mins, after which the cells were washed with PBS

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buffer, pH 7.4, and placed back in medium at 37 °C under 5% CO 2, further followed by assessment of the cell viability at 16 hr and 24 hr. The medium was removed by gentle aspiration, to which 100 µL of fresh culture medium and 10 µL of the 12 mM MTT were added and incubated for an additional 4 hr. Subsequently, 100 µL of the SDS-HCl solution was added (10% SDS in 0.01 M HCl) and incubated for another additional 1-3 hr. Similar protocols were followed for cell viability assessments of biotinylated liposomes at different concentrations (5 µM, 10 µM and 25 µM) and incubation times.

The cytotoxicity of the liposomal solutions at both longer incubation times (2 hr, 4 hr, 16 hr and 24 hr) and after biotinylation process for 20 mins (further washed and incubated at

16 hr and 24 hrs) was also investigated. Finally, the measurements were read at 570 nm using a UV-Vis multi plate reader.

3.3 Results and discussion

The present work demonstrated a simple and efficient strategy to modify the surface of the RAW 264.7 macrophage cells using two types of anchor lipids. We investigated the effects of the incorporation efficiencies of biotinylated CHOL- and

DSPE-based anchor lipids for surface modification of RAW 264.7 cells. In addition, the viability of the RAW 264.7 cells was evaluated by MTT assay.

The basic structure of the cell membrane is mostly made up of lipids in the form of lipid bilayers, due to which the anchoring lipids developed in our work allows them to serve as biomimicking models. In this study, we have developed (Figure 18) an efficient and bioorthogonal conjugation method for cell surface re-engineering applications based 120

on two kinds of anchoring lipids namely Chol-PEG2000-DBCO & DSPE-PEG2000-DBCO via copper-free click chemistry. The terminal DBCO moiety serves for reaction with

Azide-biotin (N3–biotin) biomolecule via the copper-free click chemistry. A PEG (polyethylene glycol) molecule acts as the spacer between the lipid head group and the terminal DBCO functional group which makes the anchor lipids water soluble and facilitates for use in many biomedical applications.

Copper-free click chemistry is a widely used biocompatible and bioorthogonal technique employed for metabolic labeling of cells which allows for cellular imaging, surface modifications with different biomolecules of interest such as proteins, peptides, carbohydrates, antigens. The reaction could be performed under mild conditions in aqueous buffers without catalyst and in high yields under reasonable reaction times.

Therefore, this versatile approach, which is particularly suitable for the conjugation of water soluble molecules and which can accommodate many chemical functional groups, is anticipated to be useful in the coupling of many other ligands onto cell surface, facilitating for many biomedical applications.

3.3.1 Syntheses of biotinylated anchor lipids via copper-free click chemistry for cell surface re-engineering

The synthesis of DBCO-terminal end bearing anchor lipids play an important role in facilitating for bioconjugation with many significant N3- bearing biomolecules via copper-free click chemistry. For this purpose, in this study CHOL-PEG2000-DBCO and

DSPE-PEG2000-DBCO anchor lipids were synthesized. The syntheses of these anchor 121

lipids was carried out by amidation reaction between DBCO-PEG4-NHS ester group bearing molecule and NH2-ending CHOL and DSPE anchor lipids. The resultant anchor lipids were purified via silica gel column chromatography, using a gradient mobile phase comprising of CHCl3 : MeOH respectively. Further, the collected eluents were pooled and dried under vacuum for overnight resulting in approximately 70.22% (CHOL-PEG

2000–DBCO) and 71.55% (DSPE-PEG2000-DBCO) yields. The obtained products were characterized by both 1H NMR and 13C NMR. The synthesized DBCO-terminal end anchor lipids facilitated for biotinylation of the anchor lipids with N3-biotin via copper- free click chemistry.

3.3.2 Determination of critical micelle concentration of Cholesterol-PEG2000-DBCO and DSPE-PEG2000-DBCO anchor lipids

Critical micelle concentration is the property that determines the concentration above which the individual molecules tend to start forming into micelles. The critical micelle concentration of the DBCO anchor lipids were measured by the standard pyrene assay. Briefly, aqueous solutions of CHOL and DSPE anchor lipids in PBS buffer, pH

7.4, were reacted together for overnight with pyrene (dissolved in methanol and evaporated). The CMC value determined for Chol-PEG2000-DBCO from the pyrene assay was 20 µM, whereas, the CMC value for DSPE-PEG2000-DBCO was approximately 8

µM. (Figure 19.)

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Figure 19. Measurement of critical micelle concentration (CMC) of anchor lipids:

Cholesterol–PEG2000–DBCO (A) and DSPE-PEG2000-DBCO (B).

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3.3.3 Incorporation efficiency of biotinylated lipid conjugates into the membrane of cells In order to investigate the anchoring ability of both CHOL & DSPE anchor lipids in the cell membranes, RAW 264.7 macrophage cells were either directly incubated with different concentrations (5 µM, 10 µM) of aqueous solutions of biotinylated lipid conjugates (Chol-PEG2000-biotin & DSPE-PEG2000-biotin) for 20 mins at 37 ºC or by treatment of N3-biotin to cells pre-incubated with anchor lipids according to the procedure mentioned above. In a similar manner, anchoring of liposomal solutions consisting of biotinylated anchor lipids was also studied. The incorporation of the biotinylated lipid conjugates into the cell membrane was detected by cell surface labeling with fluorescein labeled streptavidin (streptavidin-FITC). The successful incorporation of biotinylated anchor lipids was confirmed by confocal microscopy and flow cytometry after specific streptavidin-FITC binding into the cells.

Confocal microscopy images indicated higher fluorescence signals for RAW

264.7 cells (pre-incubated with anchor lipids and further reacted with N3-biotin) treated with CHOL anchoring lipids for 20 mins (Figure 20 A, C) when compared to DSPE as anchoring lipid (Figure 20 B, D). From the negative control experiments it is clearly evident that the cell membrane did not exhibit any fluorescence signal indicating that biotin-(FITC–streptavidin) was not non-specifically bound or incorporated into the cell membrane and also that the cells treated without lipid or N3-biotin had no affinity for

FITC-streptavidin (Figure 20 A, B).

Mostly the cell membrane is made up of lipid bilayers which mainly act as the backbone of the cell membrane. The cellular composition would vary from cell to cell

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and also depends on the diseased/pathological condition of the cells. Due to this possible reason, in this work, biotinylated CHOL anchor lipids have shown higher incorporation efficiencies into the cell membrane of RAW 264.7 cells in comparison to DSPE as anchoring lipid. [18].

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Figure 20. Confocal microscopy images of Raw 264.7 macrophage cells treated for 20 mins at 37 ºC, in PBS buffer, pH 7.4. Control 1 & Control 2 (panels A, B) showing no fluorescence signal.

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Figure 21. Confocal microscopy images of biotinylated Chol-DBCO (panels A, C) and

DSPE-DBCO ( panels B, D) treated with Raw 264.7 macrophage cells with varying concentrations (5 µM, 10 µM) for 20 mins at 37 ºC, in PBS, pH 7.4.

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Similarly, flow cytometry analysis demonstrated higher fluorescence intensities for cells treated with (5 µM & 10 µM) CHOL anchor lipids followed by biotinylation.

The data suggests that nearly 78-81% of the cells were incorporated with biotinylated

CHOL lipids (Figure 22 C, E). Almost negligible amounts of fluorescence signal was observed for the biotinylated DSPE anchor lipids indicating very low amounts of incorporated lipids (Figure 22 D, F). Control experiments showed no fluorescence signals indicating the specific binding interactions between the biotinylated lipids and the streptavidin-FITC (Figure 22 A, B). These results were well in agreement with the confocal data and also further indicated higher fluorescence intensities resulting from higher incorporation efficiency of the CHOL-based anchor lipids.

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Figure 22. Flow cytometry images of biotinylated Chol (panels C, E) & DSPE anchor lipids (panels D, E) treated with Raw 264.7 macrophage cells with varying concentrations

(5 µM, 10 µM) for 20 mins at 37 ºC, in PBS, pH 7.4. Control cells (A, B).

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3.3.4 Comparative biotinylation efficiencies of two-step and one-pot step on RAW

264.7 cells via copper-free click chemistry

Furthermore, we next compared the biotinylation efficiencies based on two-step and one-pot step procedures via copper-free click chemistry on RAW 264.7 cells. For this purpose, initially the cells were treated based on a two-step procedure according to the protocol mentioned earlier, wherein the cells were first incubated with just the anchor lipids (CHOL & DSPE) at different concentrations and incubation times, then treated with N3-biotin for 1 hr, and then labeled with streptavidin-FITC. However, the flow cytometry data for the two-step procedure further suggested that though most cells were labeled with streptavidin-FITC, the fluorescence signals were relatively lower.

Alternatively, cells were also treated directly with the biotinylated lipids by following the one-step procedure for biotinyaltion. In this experiment, varying concentrations of anchor lipids were reacted with N3-biotin for 1 hr at RT in eppendorf tubes, followed by direct incubation of the biotinylated lipids with the RAW 264.7 cells at different incubation times. Both the confocal microscopy and flow cytometry implied that most of the cells incorporated with biotinylated CHOL lipids exhibited much stronger fluorescence signals at 20 mins (5 µM & 10 µM, Fig 24 A, C) compared to biotinylated phospholipid (DSPE) as anchoring lipid, which on the other hand has almost no detectable signals (Fig 24 B,

D). These results indicated that there is uniform incorporation of the biotinylated cholesterol on the cell membrane with almost negligible signals from within the cytoplasm. Overall, from the comparison studies between the two-step and one-pot biotinylation process, the confocal and the flow cytometry data suggest that direct

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biotinylation process via the one-pot step method has resulted in higher incorporation efficiency of CHOL-based anchor lipids.

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Figure 23. Confocal microscopy images of Raw 264.7 macrophage cells treated for 20 mins at 37 ºC, in PBS buffer, pH 7.4. Control 1 & Control 2 (panels A, B) showing no fluorescence signal. 132

Figure 24. One-pot biotinylated Chol (panels A, C) & DSPE ( panels B, D) lipid conjugates treated with Raw 264.7 cells with varying concentration (5, 10 µM ) for 20 mins at 37 ºC, in PBS, pH 7.4.

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Figure 25. One-pot biotinylated lipid conjugate treated RAW 264.7 cells with different concentrations (5 µM, 10 µM Chol-PEG2000–DBCO–C, E), (5 µM, 10 µM DSPE-

PEG2000–DBCO–D, F) for 20 mins at 37 ºC, PBS, pH 7.4. (Control cells A, B).

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In addition, it was observed from flow cytometry data that the fluorescent intensity was exhibited from almost 99% of RAW 264.7 cells directly treated with biotinylated CHOL anchor lipids indicating complete incorporation into the cell membrane (Fig 25 C, E). Due to its highly efficient surface anchoring ability, further rate of anchoring studies were conducted with CHOL lipids. Based on this fact, further studies using liposomal solutions were conducted using the one-pot step procedure.

Moreover, this procedure also avoids extra washing steps and the biotinyaltion process can be performed at room temperature.

3.3.5 Rate of anchoring of biotinylated lipid conjugate (Chol-PEG2000-biotin) on the cell membrane

The rate of anchoring of the biotinylated CHOL lipid conjugates on the cell membranes was also investigated at different periods of time. For this purpose, the cells were incubated with different concentrations of anchor lipid (5 & 10 µM) and for different incubation times (5, 10, 20 mins) at 37 °C. As shown in (Figure 26 & 27) confocal microscopy and flow cytometry data suggests that incorporation of CHOL based anchor lipid was observed within 5 mins and their intact incorporation on to the cell membrane. However, a higher fluorescent intensity signal was observed at 20 mins

(Figure 26 & 27 C) indicating more lipid incorporation. Additionally, the rate of anchoring was also determined by flow cytometry analysis (Figure. 28), which also resulted in high fluorescence intensities, thereby confirming that almost 99.9 % of the

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cells were incorporated with the biotinylated CHOL anchor lipids synthesized by the one- pot step procedure.

Figure 26. One-pot biotinylated cholesterol lipid conjugate (5 µM) treated RAW 264.7 cells) at different time points at 37 °C, PBS pH 7.4. (5 mins–A), (10 mins–B), (20 mins–

C).

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Figure 27. One-pot biotinylated cholesterol lipid conjugate (10 µM) treated RAW 264.7 cells) at different time points at 37 °C, PBS pH 7.4. (5 mins–A), (10 mins–B), (20 mins–

C).

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Figure 28. One-pot biotinylated CHOL conjugates treated with RAW 264.7 cells at different concentrations (5, 10 µM) and at varying incubation periods at 37 °C, PBS pH

7.4 (panels A, B-5mins), (panels C, D-10 mins), (panels E, F–20 mins).

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3.3.6 Rate of anchoring and incorporation efficiency of liposomal biotinylated lipid conjugates into the cell membrane

The potential anchoring ability and effects of biotinylated liposomal anchor lipids was further investigated in our study. Similar protocols were followed for preparation of liposomal solutions where in the cells were incubated with biotinylated liposomal anchor lipids (5 µM, 10 µM, 25 µM) for different incubation times at 37 °C. After washing twice with PBS buffer, pH 7.4, the treated cells were analyzed by confocal microscopy and flow cytometry. Biotinylation of the liposomal solutions was carried out at RT for 1 hr, after liposome preparation. Initial studies performed at 20 mins, showed low fluorescence signals, due to which further experiments were carried out at higher incubation times (2 hr, 4 hr). In comparison to biotinylated DSPE lipids (Figure 33, 34 & 35), confocal images for RAW 264.7 cells treated with liposomal solutions consisting of biotinylated

CHOL lipids (Figure 30, 31 & 32) exhibited higher fluorescence signals at 20 mins, 2 hr and 4 hrs incubation times and in addition there is almost negligible fluorescence at lower concentrations (5 µM & 10 µM).

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Figure 29. Confocal microscopy images of Raw 264.7 macrophage cells treated for 20 mins at 37 ºC, in PBS buffer, pH 7.4. Control 1 & Control 2 (panels A, B) showing no fluorescencesignal.

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Figure 30. One-pot biotinylated liposomal cholesterol (A, B, C) lipid conjugate treated

RAW 264.7 cells with varying concentrations (5, 10, 25 µM) at 20 mins time point.

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Figure 31. One-pot biotinylated liposomal cholesterol (A, B, C) lipid conjugate treated

RAW 264.7 cells with varying concentrations (5, 10, 25 µM) at 2 hr time point.

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Figure 32. One-pot biotinylated liposomal cholesterol (A, B, C) lipid conjugate treated

RAW 264.7 cells with varying concentrations (5, 10, 25 µM) at 4 hr time point.

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Figure 33. One-pot biotinylated liposomal DSPE (A, B, C) lipid conjugate treated RAW

264.7 cells with varying concentrations (5, 10, 25 µM) at 20 mins time point.

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Figure 34. One-pot biotinylated liposomal DSPE (A, B, C) lipid conjugate treated RAW

264.7 cells with varying concentrations (5, 10, 25 µM) at 2 hr time point.

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Figure 35. One-pot biotinylated liposomal cholesterol (A, B, C) lipid conjugate treated

RAW 264.7 cells with varying concentrations (5, 10, 25 µM) at 4 hr time point.

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The fluorescence intensities of the treated RAW 264.7 cells were additionally analyzed by flow cytometry, which further demonstrated that nearly most of the RAW

264.7 cells were incorporated with higher concentrations of biotinylated CHOL lipids (25

µM, 99% cells gave fluorescence signals), suggesting the successful anchoring of the biotinylated CHOL lipids on the cell membrane (Figure 36 G). Partial fluorescence intensities indicated that about 35 %-66% incorporation efficiency was observed at lower concentrations (Figure 36 C, E) at 4 hr time point. In contrast, lower fluorescence intensities were observed for different concentrations of liposomal solutions consisting biotinylated CHOL lipids when treated at lower incubation times (20 mins, 2 hr) and with liposomes made of DSPE lipids (Fig 36 D, F, H).

3.3.7 Effect of aqueous solutions of biotinylated lipid conjugates and liposomes on cell viability: MTT assay

To evaluate the effect of the individual anchoring lipid aqueous solutions and liposomal solutions of biotinylated Chol-PEG2000-DBCO and DSPE-PEG2000-DBCO on cell viability, MTT assay was performed on the RAW 264.7 cell lines at different incubation times. The cell viability was assessed for 20 mins, and for longer incubation times such as 16 hrs, 24 hrs at varying concentrations of the biotinylated lipid conjugates.

In addition, effect of varying concentrations of the biotinylated lipid conjugates after biotinyaltion on the cell viability was also explored for 20 mins incubation time, after which the cells were washed twice with PBS buffer (pH 7.4) and placed back in medium for 16 hr and 24 hrs at 37 °C and 5% CO2. As shown in Figure 37. A, B and Figure 38. A,

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B), the biotinylated lipid conjugates during and after biotinyaltion for both two step and one-pot biotinylation methods did not show any significant cell toxicity except that low cytotoxicity was seen at 24 hrs.

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Figure 37. Viability of RAW 264.7 cells (A) during DBCO-lipid conjugate treatment at varying time points (20 min, 16 hr and 24 hr) at 37 °C in DMEM medium as indicated.

(B) after biotinylation of DBCO-lipid conjugate treatment for 20 min at 37 °C in DMEM medium.

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Similarly, cell viability for liposomal solutions was performed at longer incubation times such as 2 hr, 4 hr, 16 hr and 24 hr. In addition the cell viability was also assessed after the biotinylation for 20 mins. The MTT assay for liposomal solutions of biotinylated CHOL and DSPE anchor lipids demonstrated high cell viability (5 µM and

10 µM), with slight cytotoxicity at higher concentration and incubation time. (25 µM, 24 hrs) (Figure 39. A, B).

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Figure 39. Viability of RAW 264.7 cells (A) during one-pot biotinylated liposomal conjugate treatment at varying time points (2 hr, 4 hr, 16 hr and 24 hr) at 37 °C, in

DMEM medium as indicated. (B) after treatment with biotinylated liposomes for 20 min at 37 °C.

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3.4 Conclusion

In summary, the present study demonstrates an efficient and chemoselective conjugation method for surface modification of cells with Cholesterol- and DSPE-based anchor lipids. Biotinylation of the anchor lipids was achieved via Cu-free click chemistry.

The successful incorporation of the biotinylated anchor lipids was confirmed by specific interactions with Streptavidin-FITC. The findings from the confocal microscopy and flow cytometry studies indicated that the lipid fusion and biotinylation process for the purpose of cell surface modification did not have any considerable negative effects on the modified RAW 264.7 cells. The results indicated the successful incorporation of CHOL based anchor lipids with high fluorescence intensities from the cell membrane and with little or negligible internalization when compared to DSPE as anchoring lipid. This difference in the incorporation efficiencies of the anchoring lipids can be attributed to the difference in their structural characteristics. Furthermore, the cell viability assay (MTT) study also reveals that the incorporation of the CHOL and DSPE based anchor lipids into the surface of cell membrane did not cause any significant toxicity to the modified RAW

264.7 cells. These results suggest the potential use of this cell surface modification approach for many cell surface re-engineering applications.

This process not only allows for biotinylation alone but instead the Cu-free click chemistry can be used for conjugation of many important biomolecules such as peptides, proteins, carbohydrates which allows for glycan trafficking, cell labeling and for various cell surface re-engineering applications. This methodology also suggests the potential use

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of liposomes in many bio-medical applications especially in drug delivery, antigen delivery and in other diagnostic applications.

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3.5 References

[1] V. Hong, N. F. Steinmetz, M. Manchester and M. G. Finn, "Labeling live cells by

copper-catalyzed alkyne-azide click chemistry.," Bioconjugate Chemistry, vol. 21,

no. 10, pp. 1912-1916, 2010.

[2] J. M. Baskin, J. A. Prescher, S. T. Laughlin, N. J. Agard, P. V. Chang, I. A. Miller,

A. Lo, J. A. Codelli and C. R. Bertozzi, "Copper-free click chemistry for dynamic in

vivo imaging.," Proceedings of the National Academy of Sciences, vol. 104, no. 43,

pp. 16793-16797, 2007.

[3] M. T. Stephan and D. J. Irvine, "Enhancing Cell therapies from the Outside In: Cell

Surface Engineering Using Synthetic Nanomaterials.," Nano Today, vol. 6, no. 3, pp.

309-325, 2011.

[4] C. Sarkis, G. Gras, F. Sanchez, J. Mallet and C. Serguera, "Long term survival and

limited migration of genetically modified monocytes/macrophages grafted into the

mouse brain," Journal of Biomedical Science and Engineering, vol. 6, p. 561, 2013.

[5] C. L. Stabler, X.-L. Sun, W. Cui, J. T. Wilson, C. A. Haller and E. L. Chaikof,

"Surface re-engineering of pancreatic islets with recombinant azido-

thrombomodulin," Bioconjugate Chemistry, vol. 18, no. 6, pp. 1713-1715, 2007.

[6] H. R. Zope, F. Versluis, A. Ordas, J. Voskuhl, H. P. Spaink and A. Kros, "In Vitro

and In Vivo Supramolecular Modification of Biomembranes Using a Lipidated

Coiled-Coil Motif," Angewandte Chemie International Edition, vol. 52, no. 52, pp.

156

14247-14251, 2013.

[7] S.-W. Kang, S. Lee, J. H. Na, H. I. Yoon, D.-E. Lee, H. Koo, Y. W. Cho, S. H. Kim,

S. Y. Jeong, I. C. Kwon and others, "Cell Labeling and Tracking Method without

Distorted Signals by Phagocytosis of Macrophages," Theranostics, vol. 4, no. 4, p.

420, 2014.

[8] S. Gordon and S. Rabinowitz, "Macrophages as targets for drug delivery," Advanced

Drug Delivery Reviews, vol. 4, no. 1, pp. 27-47, 1989.

[9] Y. Ikehara, T. Niwa, L. Biao, S. K. Ikehara, N. Ohashi, T. Kobayashi, Y. Shimizu,

N. Kojima and H. Nakanishi, "A carbohydrate recognition--based drug delivery and

controlled release system using intraperitoneal macrophages as a cellular vehicle,"

Cancer Research, vol. 66, no. 17, pp. 8740-8748, 2006.

[10] C. Kelly, C. Jefferies and S.-A. Cryan, "Targeted liposomal drug delivery to

monocytes and macrophages," Journal of Drug Delivery, vol. 2011, 2010.

[11] N. K. Jain, V. Mishra and N. K. Mehra, "Targeted drug delivery to macrophages,"

Expert Opinion on Drug Delivery, vol. 10, no. 3, pp. 353-367, 2013.

[12] N. Kojima, M. Ishii, Y. Kawauchi and H. Takagi, "Oligomannose-Coated Liposome

as a Novel Adjuvant for the Induction of Cellular Immune Responses to Control

Disease Status," BioMed Research International, vol. 2013, 2013.

[13] N. Kawasaki, J. L. Vela, C. M. Nycholat, C. Rademacher, A. Khurana, N. van

157

Rooijen, P. R. Crocker, M. Kronenberg and J. C. Paulson, "Targeted delivery of

lipid antigen to macrophages via the CD169/sialoadhesin endocytic pathway induces

robust invariant natural killer T cell activation," Proceedings of the National

Academy of Sciences, vol. 110, no. 19, pp. 7826-7831, 2013.

[14] W. C. Chen, N. Kawasaki, C. M. Nycholat, S. Han, J. Pilotte, P. R. Crocker and J. C.

Paulson, "Antigen delivery to macrophages using liposomal nanoparticles targeting

sialoadhesin/CD169," PloS one, vol. 7, no. 6, p. e39039, 2012.

[15] M. J. Haney, Y. Zhao, E. B. Harrison, V. Mahajan, S. Ahmed, Z. He, P. Suresh, S.

D. Hingtgen, N. L. Klyachko, R. L. Mosley and others, "Specific transfection of

inflamed brain by macrophages: a new therapeutic strategy for neurodegenerative

diseases," PloS One, vol. 8, no. 4, p. e61852, 2013.

[16] N. L. Klyachko, M. J. Haney, Y. Zhao, D. S. Manickam, V. Mahajan, P. Suresh, S.

D. Hingtgen, R. L. Mosley, H. E. Gendelman, A. V. Kabanov and others,

"Macrophages offer a paradigm switch for CNS delivery of therapeutic proteins,"

Nanomedicine, no. 9, pp. 1403-1422, 2014.

[17] H. Hattori, S. Suzuki, Y. Okazaki, N. Suzuki and M. Kuwana, "Intracranial

transplantation of monocyte-derived multipotential cells enhances recovery after

ischemic stroke in rats.," Journal of Neuroscience Research, vol. 90, no. 2, pp. 479-

488, 2012.

[18] K. Tatsumi, K. Ohashi, Y. Teramura, R. Utoh, K. Kanegae, N. Watanabe, S.

158

Mukobata, M. Nakayama, H. Iwata and T. Okano, "The non-invasive cell surface

modification of hepatocytes with PEG-lipid derivatives.," Biomaterials, vol. 33, no.

3, pp. 821-828, 2012.

[19] S. Miura, Y. Teramura and H. Iwata, "Encapsulation of islets with ultra-thin polyion

complex membrane through poly(ethylene glycol)-phospholipids anchored to cell

membrane.," Biomaterials, vol. 27, no. 34, pp. 5828-5835, 2006.

[20] B. Schafer, E. Orban, A. Borics, K. Huszar, A. Nyeste, E. Welker and C. Tomboly,

"Preparation of semisynthetic lipoproteins with fluorescent cholesterol anchor and

their introduction to the cell membrane with minimal disruption of the membrane.,"

Bioconjugate Chemistry, vol. 24, no. 10, pp. 1684-1697, 2013.

[21] K. Kato, C. Itoh, T. Yasukouchi and T. Nagamune, "Rapid protein anchoring into

the membranes of Mammalian cells using oleyl chain and poly(ethylene glycol)

derivatives.," Biotechnology Progress, vol. 20, no. 3, pp. 897-904, 2004.

[22] N. Takemoto, Y. Teramura and H. Iwata, "Islet surface modification with urokinase

through DNA hybridization," Bioconjugate Chemistry, vol. 22, no. 4, pp. 673-678,

2011.

[23] H. A. Chung, K. Tajima, K. Kato, N. Matsumoto, K. Yamamoto and T. Nagamune,

"Modulating the Actions of NK Cell-Mediated Cytotoxicity Using Lipid-PEG (n)

and Inhibitory Receptor-Specific Antagonistic Peptide Conjugates," Biotechnology

Progress, vol. 21, no. 4, pp. 1226-1230, 2005.

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[24] Y. Teramura and H. Iwata, "Islet encapsulation with living cells for improvement of

biocompatibility," Biomaterials, vol. 30, no. 12, pp. 2270-2275, 2009.

[25] Y. Teramura, Y. Kaneda and H. Iwata, "Islet-encapsulation in ultra-thin layer-by-

layer membranes of poly (vinyl alcohol) anchored to poly (ethylene glycol)--lipids in

the cell membrane," Biomaterials, vol. 28, no. 32, pp. 4818-4825, 2007.

[26] D. Dutta, A. Pulsipher, W. Luo, H. Mak and M. N. Yousaf, "Engineering cell

surfaces via liposome fusion," Bioconjugate Chemistry, vol. 22, no. 12, pp. 2423-

2433, 2011.

[27] G.-J. Boons, "Liposomes modified by carbohydrate ligands can target B cells for the

treatment of B-cell lymphomas," Expert Review of Vaccines, vol. 9, no. 11, pp.

1251-1256, 2010.

[28] M. Kulkarni, U. Greiser, T. O’Brien and A. Pandit, "Liposomal gene delivery

mediated by tissue-engineered scaffolds," Trends in Biotechnology, vol. 28, no. 1,

pp. 28-36, 2010.

[29] X. Pan, G. Wu, W. Yang, R. F. Barth, W. Tjarks and R. J. Lee, "Synthesis of

cetuximab-immunoliposomes via a cholesterol-based membrane anchor for targeting

of EGFR," Bioconjugate Chemistry, vol. 18, no. 1, pp. 101-108, 2007.

[30] H. Zhang, J. Weingart, R. Jiang, J. Peng, Q. Wu and X.-L. Sun, "Bio-inspired

liposomal thrombomodulin conjugate through bio-orthogonal chemistry,"

160

Bioconjugate Chemistry, vol. 24, no. 4, pp. 550-559, 2013.

[31] S. Song, L.-Z. Cheong, M. Falkeborg, L. Liu, M. Dong, H. M. Jensen, K. Bertelsen,

M. Thorsen, T. Tan, X. Xu and others, "Facile synthesis of phosphatidyl saccharides

for preparation of anionic nanoliposomes with enhanced stability," PloS One, vol. 8,

no. 9, p. e73891, 2013.

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CHAPTER IV

BIOMIMETIC FUNCTIONALIZATION OF IRON OXIDE NANOPARTICLES

4.1 Introduction

The study of the interactions occurring at cellular and molecular levels has been of importance over the past few decades, leading to the development of more specific and targeted nano-therapies. In recent years, nanoparticles such as quantum dots (QDs), gold, silica, and superparamagnetic iron oxide nanoparticles have exhibited a great potential for use in a variety of biomedical applications including cell separation [1], magnetic resonance imaging (MRI) [2], drug/gene delivery [3] and cell tracking [4].The nanoparticles mainly consist of an inner core which is the basis for its physical functionality and properties. But, the nanoparticles alone cannot be directly used for biomedical applications. Therefore, the use of a coating layer is very necessary. It plays a vital role in the biomedical applications wherein it provides for increased stability and shelf-life, enhanced biocompatibility, non-toxicity and further prevents aggregation of

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nanoparticles. In addition, the coating layer/material offers a surface for conjugation of specific targeting ligands and other biomolecules of interest. Moreover, the type of targeting ligands [5, 6] conjugated onto the nanoparticle surface further controls and guides the interactions between the nanoparticles and their respective targets [7, 8]. Many types of amphiphillic polymers such as lipid-PEG copolymers have been employed to form stable hydrophilic layer around the hydrophobic core of nanoparticles [9, 10].

Till date, many nanoparticles have been explored widely and in particular, iron oxide nanoparticles have shown great potential as drug/gene delivery or imaging agents.

Magnetic nanoparticles have a number of physical characteristics that make them attractive tools for use in therapeutic and biomedical applications. The various sizes of these particles allow them to gain access and interact with the tissue cells in the human body. Their high surface-to-volume ratio makes them capable of accommodating large payloads of drugs and molecular markers necessary for the recognition of many cellular processes. Lipid coating of (lipid–PEG polymer conjugates) nanoparticles allows for aqueous dispersion and for prolonged blood circulation times [10, 11, 12, 13].

Furthermore, cell surface carbohydrates have specific interactions with proteins such as lectins, which play an important role in various biological recognition processes.

Therefore, cell surface carbohydrates serve as remarkable models for surface modification of iron oxide with carbohydrates owing to their biodegradability, low toxicity, and their specific binding interactions with the receptors present at the targeted cell’s surface [14]. Bioorthogonal approaches serve as a chemically selective and biocompatible method for nanoparticle surface functionalization [15].

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Therefore, in our present study we have reported the synthesis and preparation of carbohydrate/lipid–coated magnetic nanoparticle hybrid systems. Particularly, magnetic nanoparticles were synthesized by the thermal-decomposition of iron oleates in high– boiling solvents containing surfactants in order to obtain tunable, high quality and monodisperse iron oxide nanoparticles. Lipid–coated magnetic nanoparticles were prepared by standard thin-film hydration method and sonication process as well as dual solvent exchange method. Chemically selective glyco-functionalization of lipid–coated magnetic nanoparticles was conducted via Staudinger ligation. The structural characteristics of the nanoparticles were confirmed by IR, TEM and DLS techniques.

4.2 Experimental

4.2.1 Materials and methods

All solvents and reagents were purchased from commercial sources and were used as received, unless otherwise noted. Deionized water was used as a solvent in all procedures. N-hydroxysuccinimide (NHS-ester) and Cholesteryl chloroformate were purchased from Sigma-Aldrich (St. Louis, MO). 1, 2-dipalmitoyl-sn-glycero-3- phosphocholine (DPPC), 1, 2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[amino

(polyethylene glycol)-2000] (ammonium salt) (DSPE-PEG2000-NH2) were purchased from Avanti Polar Lipids, Inc (Alabaster, AL). Iron oxide nanocrystals (10 nm size) in chloroform was purchased from Ocean NanoTech LLC (Springdale, AR).

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4.2.2 Synthesis of iron oxide nanoparticles via thermal decomposition method

The synthesis of iron oxide nanoparticles was synthesized by modified from previously published method [10]. Briefly, 3.24g of FeCl3.6H2O (12mmol) was dissolved in 12 ml of distilled water, filtered and mixed with 10.95g of sodium oleate (40mmol),

24ml of ethanol, 6 ml of distilled water, and 42ml of hexane. The solution was then heated to 70 °C and stirred for 4hr under an argon flow. An upper red-brownish organic layer which contains the iron oleate complex was removed and then washed with 9 ml of distilled water for 3 times using a separatory funnel. Then the hexane was removed by evaporating the solution by means of a rotary evaporator to obtain a viscous oily substance. This viscous oil was extracted with ethanol and acetone for 2 times and then it was dried. 2.78g (3mmol) of iron oleate complex (thermally treated at 30 °C in a vacuum oven for 24h), 0.96ml of oleic acid (3mmol), and 10ml of octadecane were mixed in a three neck round- bottom reaction flask. The mixture was first heated to 600C to melt the solvent, and then the reactants were dissolved with vigorous stirring. Then the temperature was increased to 318 °C with a heating rate of 3.3 °C / min under stirring and refluxing for 3 mins. There appears a color change from initial reddish–brown to brownish–black color during this time. The solution was cooled down to 50 C and then a mixture of 10ml of hexane and 40ml of acetone was added to precipitate the nanoparticles. Then these nanoparticles were separated by centrifugation and washed 3 times with the hexane and acetone mixture. Again the nanoparticles were centrifuged and dissolved in chloroform for long–term storage.

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4.2.3 Lipid coating of iron oxide nanoparticles

Lipid coating of the oleic acid capped hydrophobic iron oxide nanoparticles was performed by the thin film hydration and sonication process as well as dual solvent exchange method. A schematic illustration of lipid coated iron oxide nanoparticles is shown in (Figure.40).

Figure 40. Schematic illustration of lipid coating of oleic acid capped iron oxide nanoparticles.

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4.2.3.1 Thin-film hydration and sonication

Iron oxide nanoparticles coated with oleic acid (hydrophobic nanoparticles) were prepared via the thermal decomposition method. Further, lipid coated iron oxide nanoparticles were prepared by standard thin-film hydration method and sonication process. DPPC and Chol-PEG2000-TP were dissolved at 2:1 (weight ratio) in 10ml chloroform, mixed with the hydrophobic nanoparticles (3750µg/10ml) and finally the chloroform was evaporated under reduced pressure. The resultant nanoparticle – lipid film was hydrated in 10 ml distilled water to give the final concentrations of lipids at

200µM and iron oxide at 375µg/ml. The mixture was sonicated at 60 ºC for 20mins. The lipid coated iron oxide nanoparticles were separated by a magnet, filter sterilized through a Millex 0.22µm–diameter filter and finally stored at 4 °C in argon gas. The size distribution of the prepared lipid coated iron oxide nanoparticles was monitored by DLS.

4.2.3.2 Dual solvent exchange method

Recently, Tong et al. [11] have demonstrated a better and efficient lipid coating technique for iron oxide nanoparticles. Briefly, iron oxide nanoparticles and lipids (2% anchor lipid + DPPC) were mixed (1:2 wt. ratio) and dissolved in 1 mL CHCl3. After which 4 mL of DMSO was slowly added and incubated for 30 mins on a shaker. Further, followed by evaporation of CHCl3 for about 30-40 mins. Finally, 20 mL of deionized water was added and followed by removal of DMSO by centrifugation (10 kDa cutoff filter, 6000 g, 3 times).

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4.2.4. Glyco-functionalization of iron oxide nanoparticles via staudinger ligation

Azido-sugars (4 mg, 0.01 mmol) in 0.2 mL PBS buffer (Argon bubble before use) were added to 2 mL of obtained lipid coated iron oxide nanoparticles. The Staudinger ligation was performed at room temperature for 6 hr under an argon atmosphere; later lactose conjugated iron oxide nanoparticles were separated from unreacted azido-lactose by means of an external magnet. The size of nanoparticles before and after Staudinger ligation was monitored over time by using 90 Plus particle analyzer. Control experiments were conducted in the absence of azide-lactose.

4.3 Results and discussion

4.3.1 Synthesis of iron oxide nanoparticles via thermal decomposition method

The desire to synthesize monodisperse and highly stable iron oxide nanoparticles with hydrophobic surfaces poses number of obstacles. Therefore, the use of high temperature thermal decomposition methods was chosen for the synthesis process.

(Figure 41). The main advantage of this synthetic approach is that highly crystalline and monodisperse iron oxide nanoparticles can be obtained in a wide range of sizes from 6 nm–30 nm, just by varying the heating conditions and the temperature of the solvent and also that the iron oleate complex obtained as the intermediate is non-toxic and inexpensive compounds are used for the synthesis.

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Figure 41. Schematic illustration of synthesis of oleic acid capped iron oxide nanoparticles by thermal decomposition of iron oleate complex with high boiling points.

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Briefly, iron chloride hexahydrate is used as the metal precursor and was added to oleic acid which acts as the stabilizer surfactant to form the iron oleate complex. The iron oleate complex was confirmed by FT-IR spectrum. The iron oleate was further reacted with oleic acid and octadecane to finally obtain the iron oxide nanoparticles. The nanoparticles were confirmed by FT-IR spectrum and their size was determined by (high resolution transmission electron microscopy) HRTEM images. The TEM images showed that the core particle sizes were ranging from 3 nm-30 nm.

4.3.1.1 Characterization of iron oxide nanoparticles by FT-IR

The FT-IR spectrum of pure oleic acid was obtained as shown in (Figure 42), where a characteristic peak was observed at 1710 cm-1 exhibiting the presence of C=O

-1 stretch. Symmetric and asymmetric CH2 stretching was observed at 2922cm and 2853 cm-1 respectively and the band at 1285 cm-1 exhibits the presence of C-O stretch. The FT-

IR spectrum of the Oleic acid capped iron oxide nanoparticles showed the shift in bands to a lower frequency region which indicated the adsorbed state of the surfactant molecules. This indicated that the hydrocarbon chains in the monolayer surrounding the nanoparticles were in a closed-packed state. The appearance of two new peaks at around

1541 cm-1 and at 1645 cm-1 exhibited the presence of asymmetric and symmetric COO- stretching.

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Figure 42. FT-IR spectra of oleic acid capped iron oxide nanoparticles (A) oleic acid (B) synthesized iron oxide nanoparticles (C) commercially synthesized oleic acid capped iron oxide nanoparticles.

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4.3.1.2 Characterization of iron oxide nanoparticles by TEM

HRTEM images of the oleic acid capped nanoparticles were obtained by placing a drop of dilute sample solution on the carbon-coated Cu TEM grid, and allowed to dry.

From the image as shown below, it is seen that nanoparticles of size ranges from 3 nm-30 nm were obtained. As shown in (Figure 43.) when compared to the commercially synthesized iron oxide nanoparticles, homogeneous size distribution was not achieved for the laboratory scale synthesized iron oxide nanoparticles. For this purpose, commercially synthesized iron oxide nanoparticles were employed for further experiments.

Figure 43. TEM images of (A) laboratory synthesized iron oxide nanoparticles (B) commercially synthesized iron oxide nanoparticles. (bar scale: 20 nm).

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4.3.2 Lipid coating of iron oxide nanoparticles

The commercial iron oxide nanoparticles synthesized via the thermal decomposition method results in the production of oleic acid capped iron oxide nanoparticles which are hydrophobic and are not dispersible in aqueous solutions. Due to this purpose efficient lipid coating of such nanoparticles is highly essential for use in biomedical applications. Initial lipid coating of iron oxide nanoparticles was achieved by the traditional thin film and sonication process. However, efficient coating of iron oxide nanoparticles was not achieved by the conventional thin film hydration approach due to which it resulted in early formation of aggregates. For this purpose further lipid coating experiments were carried out by employing the dual solvent exchange method.

4.3.3 Glyco-functionalization of iron oxide nanoparticles via staudinger ligation

Cell surface carbohydrates serve as remarkable models for the iron oxide surface modification with carbohydrates owing to their biodegradability, low toxicity, and mainly due to their specific binding interactions with the receptors present at the targeted cell’s surface. Glyco-functionalization of iron oxide nanoparticles can be achieved by

Staudinger ligation. Staudinger ligation is one of the latest mechanisms of surface conjugation techniques, which involves the interactions between an azide group and a triphosphine group to selectively react to form an amide bond. This mechanism offers an efficient and chemoselective conjugation method for surface functionalization of iron oxide nanoparticles.

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4.3.4 Stability evaluation of lipid coated and glyco-functionalized iron oxide nanoparticles via DLS

Lipid coating of iron oxide nanoparticles was achieved by both thin-film hydration and sonication process as well as dual exchange solvent method. Two anchor lipids, namely Chol-PEG2000–TP and DSPE-PEG2000–TP were mixed with DPPC and further coated onto iron oxide nanoparticles. The lipid coating occurs through the hydrophobic interactions between the oleic acid capped iron oxide nanoparticles and the hydrophobic regions of the lipid molecules. As observed from (Figure. 37 and Figure.

38), it is seen that the traditional thin-film hydration and sonication process lead to early formation of aggregates. The stability of the lipid coated iron oxide nanoparticles was lost from 7th day itself. For this reason, more efficient approach such as the dual solvent exchange method was employed for lipid coating onto the hydrophobic iron oxide nanoparticles. As observed from (Figure. 44 and 45) lipid coating via dual exchange solvent approach allowed for stability of iron oxide nanoparticles for almost 14 days.

Further glyco-functionalization of these lipid coated iron oxide nanoparticles via

Staudinger ligation strategy was achieved. The glyco-functionalized iron oxide nanoparticles were stable and water dispersible without any aggregate formation for almost more than a month with a size distribution of about 95 nm-130 nm for both Chol-

PEG2000–TP and DSPE-PEG2000–TP anchor lipids coated iron oxide nanoparticles.

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4.4 Conclusion

Overall, in this study, we investigated the anchor lipid effects on the iron oxide nanoparticle surface functionalization via Staudinger ligation. Two lipid anchors namely,

Chol-PEG2000-TP and DSPE-PEG2000-TP were synthesized and further evaluated for their efficiency for nanoparticle surface functionalization by means of a lactose moiety via

Staudinger ligation. Lipid coating of iron oxide nanoparticles was achieved by thin film hydration and sonication approach as well as dual solvent exchange method. The stability and size of the glyco-functionalized iron oxide nanoparticles were monitored by the dynamic light scattering technique. Lipid coating and further glyco-functionalization (via

Staudinger ligation) of iron oxide nanoparticles via the dual solvent exchange method resulted in longer shelf-life and more stability when compared to the traditional thin film hydration and sonication approach. Therefore, the reported anchor lipids and the chemoselective bioorthogonal strategy employed for preparation of lipid coated and glyco-functionalized iron oxide nanoparticles can serve as efficient coupling strategies for attachment of many biomolecules of interest, thereby further enhancing the biocompatibility of nanoparticles in many biomedical applications. However, further studies such as drug encapsulation, cytotoxicity, cellular uptake and imaging studies needs to be further investigated, for demonstrating the novelty and efficiency of iron oxide nanocarrier systems developed in our study.

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4.5 References

[1] O. Olsvik, T. Popovic, E. Skjerve, K. S. Cudjoe, E. Hornes, J. Ugelstad and M.

Uhlen, "Magnetic separation techniques in diagnostic microbiology.," Clinical

Microbiology Reviews, vol. 7, no. 1, pp. 43-54, 1994.

[2] N. Nitin, L. LaConte, O. Zurkiya, X. Hu and G. Bao, "Functionalization and peptide-

based delivery of magnetic nanoparticles as an intracellular MRI contrast agent,"

JBIC Journal of Biological Inorganic Chemistry, vol. 9, no. 6, pp. 706-712, 2004.

[3] M. K. Yu, Y. Y. Jeong, J. Park, S. Park, J. W. Kim, J. J. Min, K. Kim and S. Jon,

"Drug-loaded superparamagnetic iron oxide nanoparticles for combined cancer

imaging and therapy in vivo," Angewandte Chemie International Edition, vol. 47,

no. 29, pp. 5362-5365, 2008.

[4] E. Bull, S. Y. Madani, R. Sheth, A. Seifalian, M. Green and A. M. Seifalian, "Stem

cell tracking using iron oxide nanoparticles," International Journal of

Nanomedicine, vol. 9, p. 1641, 2014.

[5] L. Yang, H. Mao, Z. Cao, Y. A. Wang, X. Peng, X. Wang, H. K. Sajja, L. Wang, H.

Duan, C. Ni and others, "Molecular imaging of pancreatic cancer in an animal model

using targeted multifunctional nanoparticles," Gastroenterology, vol. 136, no. 5, pp.

1514-1525, 2009.

[6] L. Yang, H. Mao, Y. A. Wang, Z. Cao, X. Peng, X. Wang, H. Duan, C. Ni, Q. Yuan,

G. Adams and others, "Single chain epidermal growth factor receptor antibody

178

conjugated nanoparticles for in vivo tumor targeting and imaging," Small, vol. 5, no.

2, pp. 235-243, 2009.

[7] D. Artemov, N. Mori, B. Okollie and Z. M. Bhujwalla, "MR molecular imaging of

the Her-2/neu receptor in breast cancer cells using targeted iron oxide

nanoparticles," Magnetic Resonance in Medicine, vol. 49, no. 3, pp. 403-408, 2003.

[8] L. Yang, Z. Cao, H. K. Sajja, H. Mao, L. Wang, H. Geng, H. Xu, T. Jiang, W. C.

Wood, S. Nie and others, "Development of receptor targeted magnetic iron oxide

nanoparticles for efficient drug delivery and tumor imaging," Journal of Biomedical

Nanotechnology, vol. 4, no. 4, pp. 439-449, 2008.

[9] S. M. Moghimi, A. C. Hunter and J. C. Murray, "Long-circulating and target-

specific nanoparticles: theory to practice," Pharmacological Reviews, vol. 53, no. 2,

pp. 283-318, 2001.

[10] E. V. Shtykova, X. Huang, N. Remmes, D. Baxter, B. Stein, B. Dragnea, D. I.

Svergun and L. M. Bronstein, "Structure and properties of iron oxide nanoparticles

encapsulated by phospholipids with poly (ethylene glycol) tails," The Journal of

Physical Chemistry C, vol. 111, no. 49, pp. 18078-18086, 2007.

[11] S. Tong, S. Hou, B. Ren, Z. Zheng and G. Bao, "Self-assembly of phospholipid--

PEG coating on nanoparticles through dual solvent exchange," Nano Letters, vol. 11,

no. 9, pp. 3720-3726, 2011.

179

[12] H.-C. Huang, P.-Y. Chang, K. Chang, C.-Y. Chen, C.-W. Lin, J.-H. Chen, C.-Y.

Mou, Z.-F. Chang and F.-H. Chang, "Formulation of novel lipid-coated magnetic

nanoparticles as the probe for in vivo imaging," Journal of Biomedical Science, vol.

16, no. 1, p. 86, 2009.

[13] A. A. Allam, M. E. Sadat, S. J. Potter, D. B. Mast, D. F. Mohamed, F. S. Habib and

G. M. Pauletti, "Stability and magnetically induced heating behavior of lipid-coated

Fe3O4 nanoparticles," Nanoscale Research Letters, vol. 8, no. 1, pp. 1-7, 2013.

[14] C. Tassa, S. Y. Shaw and R. Weissleder, "Dextran-coated iron oxide nanoparticles: a

versatile platform for targeted molecular imaging, molecular diagnostics, and

therapy," Accounts of Chemical Research, vol. 44, no. 10, pp. 842-852, 2011.

[15] J. I. Cutler, D. Zheng, X. Xu, D. A. Giljohann and C. A. Mirkin, "Polyvalent

oligonucleotide iron oxide nanoparticle “click” conjugates," Nano Letters, vol. 10,

no. 4, pp. 1477-1480, 2010.

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CHAPTER V

SUMMARY

Functional lipids play important roles in lipid-based nanoparticles for targeted drug delivery and cell surface re-engineering applications. In this dissertation study, two functional lipids, phospholid- and cholesterol-based anchoring lipids were designed, synthesized and evaluated for efficient and bioorthogonal liposomes and lipid-coated iron oxide nanoparticles surface functionalization via Staudinger ligation and for potential cell surface re-engineering via copper-free click chemistry.

First, a chemoselective and bioorthogonal glyco-liposome surface functionalization through Staudinger ligation reaction was developed. Briefly, the surface of liposomes decorated with a lipid anchor bearing a terminal triphenylphosphine moiety is conjugate to a carbohydrate molecule with azide functionality. The stability of the

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glyco-functionalized liposomes was assessed by comparing the effects of two types of anchors lipids, namely Chol-PEG2000-TP and DSPE-PEG2000-TP. The integrity of the liposome vesicles before and after glyco-functionalization was measured by dynamic light scattering and fluorescent dye releasing kinetic studies of entrapped 5, 6

Carboxyfluorescein dye. Furthermore, the density and accessibility of the grafted carbohydrate molecules on the liposome surface was evaluated with specific lectin binding interactions. Therefore, this simple and efficient approach, which is mainly effective for conjugation of many aqueous soluble biomolecules and in the absence of catalyst, was developed and is anticipated to allow coupling of many targeting ligands onto liposomes, which paves path for many potential applications.

In addition, stable lipid coated and glyco-functionalized iron oxide nanoparticles were developed via dual solvent exchange and Staudinger ligation methods. The iron oxide nanoparticles were synthesized via thermal-decomposition method and characterized by FT-IR spectroscopy. Glyco-functionalization of iron oxide nanoparticles was achieved by conjugation of an azide bearing lactose molecule to triphenylphosphine carrying lipid coated iron oxide nanoparticles via Staudinger ligation. The stability of the glyco-functionalized iron oxide nanoparticles with time was evaluated by dynamic light scattering technique. Further studies involving the assessment of the biocompatibility and non-toxicity of iron oxide nanoparticles need to be investigated by MTT assay.

Second, a simple, rapid and efficient approach for cell surface re-engineering of

RAW 264.7 cells via lipid anchoring approach was investigated. The effective lipid anchoring of phospholipid (DSPE-PEG2000-DBCO) and cholesterol (CHOL-PEG2000-

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DBCO) based anchor lipids was designed for conjugation to azide-biotin molecules via bioorthogonal copper-free click chemistry method. Briefly, the RAW 264.7 cells were treated with varying concentrations of biotinylated anchor lipids or their respective liposomal formulations, and at different incubation times at pH 7.4. The successful incorporation of biotinylated anchor lipids was confirmed by confocal microscopy and flow cytometry after specific streptavidin-FITC binding onto the cells. Cholesterol-based anchor lipids afforded relatively higher cell membrane incorporation efficiency with less internalization compared to phospholipid as an anchoring lipid. Furthermore, the cytotoxic effect of the synthesized biotinylated anchor lipids on the RAW 264.7 cells was assessed with less negative effect. Thus, this study suggests the possible usage of these lipids for potential cell surface re-engineering applications.

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CHAPTER VI

FUTURE PERSPECTIVE

Overall, in this study, stable and biocompatible lipid coated and glyco- functionalized nanocarrier systems have been successfully developed through chemoselective and bioorthogonal approaches. The effect of different types of anchor lipids on the stability of liposomes as well as their cell membrane incorporation efficiencies was investigated and demonstrated. These systems pose significant potential to be employed as targeted delivery systems as well as for many cell surface re- engineering applications. However, further studies with cell lines still needs to be investigated in order for these systems to be recognized as novel carrier systems.

Additional investigations will seek to identify the possible use of liposomes and iron oxide nanoparticles in many animal studies.

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Further, evaluation of glyco-functionalized liposomes as drug carriers for selective and active targeting can be determined by exploiting the carbohydrate-protein recognition events. Moreover, use of immunoliposomes incorporating suitable therapeutic agents will further enhance the selectivity and therapeutic index of encapsulated drugs.

In addition, role of macrophages (RAW 264.7 cells) in antigen processing and its presentation, using our proposed cell surface re-engineering via lipid anchoring approach needs to be explored in future. Also, the integrity of the structure and activity of macrophages upon modifications with different anchoring lipids needs to be explored and can be determined by ELISA assay.

Similarly, drug loading capacity, delivery and cytotoxicity of glyco-functionalized iron oxide nanoparticles with varying compositions of lipid anchors needs to be explored further.

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