Modification of Superparamagnetic Nanoparticles for

Biomedical Applications

By

Chenjie Xu

B.S., Nanjing University, 2002

M.Phil., Hong Kong University of Science & Technology, 2004

A Dissertation Submitted in Partial Fulfillment of the

Requirements for the Degree of Doctor of Philosophy

in the Department of Chemistry at Brown University

Providence, Rhode Island

May 2009

© Copyright 2009 by CHENJIE XU

iii

This dissertation by Chenjie Xu is accepted in its present form

by the Department of Chemistry as satisfying the

dissertation requirement for the degree of Doctor of Philosophy

Date__________________ ____________________

Shouheng Sun, Advisor

Recommend to the Graduate Council

Date__________________ ____________________

Matthew Zimmt, Reader

Date__________________ ____________________

Jeffrey Morgan, Reader

Approved by the Graduate Council

Date__________________ ____________________

Sheila Bonde, Dean of the Graduate School

iv

VITA

Chenjie Xu was born on October 8, 1979, in Yangzhou city of Jiangsu province in China.

He went to the Department for Intensive Instruction in Nanjing University (Nanjing,

China) for undergraduate study starting in 1998, graduating with his B.Sc. in Chemistry

in 2002. In 2002, he was admitted to the Department of Chemistry, Hong Kong

University of Science & Technology (Kowloon, Hong Kong) and obtained his M. Phil.

under the supervision of Prof. Bing Xu (2004). After a year short stay as a visiting

scholar at Molecular Imaging Program at Stanford University (Palo Alto, California,

2005), he joined Prof. Shouheng Sun’s group as a graduate student at Brown University

(Providence, RI). His research interests are in magnetic probes for cancer imaging and

drug delivery. He is the recipient of Vince Wernig Fellowship at 2008.

Publications

25) Xu, Chenjie; Weifeng, Shen; Gang, Xiao; Sun, Shouheng. “Size Effect of Magnetic

Nanoparticles for Magnetic Detection”, in preparation

24) Xu, Chenjie; Sun, Shouheng. “Gold Cluster Doped Magnetic Nanoparticles

Synthesis and Their Catalysis for Oxygen Reduction”, in preparation

23) Xu, Chenjie; Yuan, Zhenglong; Kim, Jaemin; Chung, Maureen A.; Sun, Shouheng.

“Controlled Release of Fe from FePt Nanoparticles for Tumor Inhibition”, Submitted.

22) Young, Kaylie L.; Xu, Chenjie; Xie, Jin; Sun, Shouheng. “Conjugating Methotrexate

to Magnetite (Fe3O4) Nanoparticles via Trichloro-s-Triazine”, Submitted.

v

21) Xu, Chenjie; Wang, Baodui; Sun, Shouheng. “Dumbbell-Like Au-Fe3O4

Nanoparticles for Target-Specific Platin Delivery”, Journal of the American Chemical

Society, 2009, 131(12), 4216-4217.

20) Wang, Chao; Xu, Chenjie; Zeng, Hao; Sun, Shouheng. “Recent Progress in

Syntheses and Applications of Dumbbell-like Nanoparticles”, Advanced Materials, 2009,

in press

19) Xu, Chenjie; Sun, Shouheng. “Superparamagnetic Nanoparticles as Targeted Probes

for Diagnostic and Therapeutic Applications”, Dalton Transactions, 2009, DOI:

10.1039/b900272n.

18) Wang, Baodui; Xu, Chenjie; Xie, Jin; Yang, Zhengyin; Sun, Shouheng. “pH

Controlled Release of Chromone from Chromone-Fe3O4 Nanoparticles”, Journal of the

American Chemical Society, 2008, 130(44), 14436-14437.

17) Lee, Ha-Young; Li, Zibo; Chen, Kai; Hsu, Andrew R.; Xu, Chenjie; Xie, Jin; Sun,

Shouheng; Chen, Xiaoyuan. “PET/MRI Dual-modality Tumor Imaging Using Arginine-

glycine-aspartic (RGD)-conjugated Radiolabeled Iron Oxide Nanoparticles”, Journal of

Nuclear Medicine, 2008, 49(8), 1371-1379

16) Xu, Chenjie; Tung, Glenn A.; Sun, Shouheng. “Size and Concentration Effect of

Gold Nanoparticles on X-ray Attenuation as Measured on Computed Tomography”,

Chemistry of Materials, 2008, 20(13), 4167-4169

15) Shen, Weifeng; Schrag, Benaiah D.; Carter, Matthew J.; Xie, Jin; Xu, Chenjie; Sun,

Shouheng; Xiao, Gang. “Detection of DNA Labeled with Magnetic Nanoparticles Using

vi

MgO-based Magnetic Tunnel Junction Sensors”, Journal of Applied Physics, 2008, 103(7,

Pt. 2), 07A306/1-07A306/3

14) Xie, Jin; Chen, Kai; Lee, Ha-Young; Xu, Chenjie; Hsu, Andrew R.; Peng, Sheng;

Chen, Xiaoyuan; Sun, Shouheng. “Ultrasmall c(RGDyK)-Coated Fe3O4 Nanoparticles

and Their Specific Targeting to Integrin αvβ3-Rich Tumor Cells”, Journal of the

American Chemical Society, 2008, 130(24), 7542-7543.

13) Lee, Ha-Young; Lee, Sang-Hoon; Xu, Chenjie; Xie, Jin; Lee, Jin-Hyung; Wu, Bing;

Koh, Ai Leen; Wang, Xiaoying; Sinclair, Robert; Wang, Shan X; Nishimura, Dwight G;

Biswal, Sandip; Sun, Shouheng; Cho, Sun Hang; Chen, Xiaoyuan. “Synthesis and

Characterization of PVP-coated Large Core Iron Oxide Nanoparticles as an MRI Contrast

Agent”, Nanotechnology, 2008, 19, 165101

12) Xu, Chenjie; Xie, Jin; Kohler, Nathan; Walsh, Edward; Chin, Y. Eugene; Sun,

Shouheng. “Monodisperse Magnetite Nanoparticles Coupled with Nuclear Localization

Signal Peptide for Cell-Nucleus Targeting”, Chemistry-an Asian Journal, 2008, 3, 548-

552

11) Xie, Jin; Xu, Chenjie; Young, Kaylie; Sun, Shouheng. “Controlled pegylation of

monodisperse magnetic nanoparticles for biomedical applications”, PMSE Preprints,

2008, 98 291

10) Xu, Chenjie; Xie, Jin; Don, Ho; Wang, Chao; Kohler, Nathan; Walsh, Edward;

Morgan, Jeffrey; Chin, Y. Eugene; Sun, Shouheng. “Au-Fe3O4 Dumbbell Nanoparticles

as Dual-Functional Probes”, Angewandte Chemie International Edition, 2008, 47(1),

173-176.

vii

9) Xu, Chenjie; Sun, Shouheng. “Monodisperse Magnetic Nanoparticles for Biomedical

Applications”, Polymer International, 2007, 56(7), 821-826.

8) Xie, Jin; Xu, Chenjie; Kohler, Nathan; Hou, Yanglong; Sun, Shouheng. “Controlled

PEGlation of Monodisperse Fe3O4 Nanoparticles for Reduced Non-specific Uptake by

Macrophage Cells”, Advanced Materials, 2007, 19(20), 3163-3166.

7) Xie, Jin; Xu, Chenjie; Xu, Zhichuan; Hou, Yanglong; Young, Kaylie L., Wang, S. X.;

Pourmond, Nader; Sun, Shouheng. “Linking hydrophilic macromolecules to

monodisperse magnetite (Fe3O4) nanoparticles via trichloro-s-triazine”, Chemistry of

Materials, 2006, 18(23), 5401-5403

6) Xu, Chenjie; Xing, Bengang; Rao, Jianghong. “A Self-assembled Quantum Dot Probe

for Detecting Beta-lactamase Activity”, Biochemical and Biophysical Research

Communications, 2006, 344(3), 931-935

5) So, Min-Kyung*, Xu, Chenjie*; Loening, Andreas M.; Gambhir, Sanjiv S.;

Rao,Jianghong. “Self-illuminating Quantum Dot Conjugates for In Vivo Imaging”,

Nature Biotechnology, 2006, 24(3), 339-343 (* Same Contribution)

4) Gu, Hongwei; Xu, Keming; Xu, Chenjie, Xu, Bing. “Biofunctional Magnetic

Nanoparticles for Protein Separation and Pathogen Detection”, Chemical Communication,

2006, (9), 941-949.

3) Xu, Chenjie; Xu, Keming; Gu, Hongwei; Zheng, Rongkun; Liu, Hui; Zhang, Xixiang;

Guo, Zhihong; Xu, Bing. “Dopamine as a Robust Anchor to Immobilize Functional

Molecules on the Iron Oxide Shell of Magnetic Nanoparticles”, Journal of the American

Chemical Society, 2004, 126(32), 9938-9939

viii

2) Xu, Chenjie; Xu, Keming; Gu, Hongwei; Zhong, Xiaofen; Guo, Zhihong; Zheng,

Rongkun; Zhang, Xixiang; Xu, Bing. “Nitrilotriacetic Acid-modified Magnetic

Nanoparticles as a General Agent to Bind Histidine-tagged Proteins”, Journal of the

American Chemical Society, 2004, 126(11), 3392-3393

1) Gu, Hongwei; Xu, Chenjie; Weng, Lu-Tao; Xu, Bing, “Solventless Polymerization:

Spatial Migration of a Catalyst to Form Polymeric Thin Films in Microchannels”,

Journal of the American Chemical Society, 2003, 125 (31), 9256-9257.

ix

ACKNOWLEDGEMENTS

There are many people I would like to thank for their help and encouragement during

the past four years. First of all, I would like to thank my advisor, Prof. Shouheng Sun. I

appreciate that he gave me the valuable opportunity to come to Brown and join his group

for my Ph.D. studies. I also appreciate his patience and everlasting support during my

research. The freedom to pursue my own ideas during my graduate studies is the best

thing I have enjoyed here at Brown.

Secondly, I would like to thank my committee members, Prof. Matthew Zimmt and

Prof. Jeffrey Morgan (Department of Molecular Pharmacology). They provided valuable

suggestions during my RP and ORP defenses, which are the essential steps to become a

Ph. D.

Thirdly, I want to thank Prof. Eugene Y. Chin (Rhode Island Hospital) for letting us

use his biology facilities and Prof. Gang Xiao (Department of Physics) for using the

magnetic microscope in his lab. Dr. Glenn A. Tung (Rhode Island Hospital) and Dr.

Edward Walsh (Department of Neuroscience), thank you for helping me acquiring and

analyzing data with computed tomography (CT) and magnetic resonance imaging (MRI).

In addition, I want to express my appreciation to Prof. Peter Weber, the chair of

Chemistry. You gave me a lot of encouragement and suggestions to begin a small

business. Although I haven’t done it, I do have the plan and courage to realize this dream

in the future.

I also feel fortunate to have many professional members here at Brown I can count on.

Dr. Zhenglong Yuan (Rhode Island Hospital), thank you for helping me do those cell

x

biological experiments. Dr. Tun-Li Shen, thank you for your efforts in helping me

acquire and analyze mass spectrum. Mr. Eric Friedfeld and Mr. Robert Wilson, thank for

helping us order instruments for our bio-lab. Dr. David Murray and Mr. Joe Orchardo

(Department of Geological Sciences), thank you for the help with element analysis. Dr.

Anthony McCormick (Engineering), thank you for helping me with high resolution TEM.

It has been a pleasure to interact with all the members in Sun’s group. I would

especially thank Dr. Jin Xie, a post-doctor at Stanford University now. All the

achievements in our bio-lab now are based on our cooperation and discussions when

there was no space for us to do experiment. And thanks Dr. Nathan Kohler, for the great

advices you gave. Thanks Dr. Sheng Peng, Dr. Chao Wang and Dr. Jaemin Kim, it has

been great four years working with you. Thanks Kaylie Young, and wish you have a

great time in Northwestern. And thanks for all the people who have worked in this group.

Finally I would like to thank my dear wife, Ms. Hong Qian who always loves and

supports me at any time. I also want to thank my mother, Meifang Xue for bringing me

up and supporting my study.

Dad, I achieved my dream.

xi

To My Love, HONG QIAN

xii

Abstract of “Modification of Superparamagnetic Nanoparticles for Biomedical

Applications” by CHENJIE XU, Ph. D., Brown University, May 2009

Superparamagnetic nanoparticles (NPs) have been attractive for medical diagnostics

and therapeutics due to their unique magnetic properties and their ability to interact with

various biomolecules of interest. The solution phase based chemical synthesis provides a

near precise control on NP size, and monodisperse magnetic NPs with standard deviation

in diameter of less than 10%, which are now routinely available. Upon controlled surface

functionalization and coupling with fragments of DNA strands, proteins, peptides or

antibodies, these NPs can be well-dispersed in biological solutions and used for drug

delivery, magnetic separation, magnetic resonance imaging contrast enhancement and

magnetic fluid hyperthermia.

This dissertation begins with an overview of the background, common syntheses and

controlled surface functionalization of monodisperse superparamagnetic nanoparticles.

Then the detailed examples are offered in each chapter to explain the efforts I spent in the

past four years exploring the functionalization and biomedical applications of magnetic

nanoparticles.

xiii

Table of Contents

Chapter 1: Background and Fundamental Theory of Superparamagnetic Nanoparticle

Synthesis, Characterization and Biomedical Applications ……………1

1. Introduction ………………………………………………………2

2. Fundamental Properties of Nanomaterials ………………………… 4

2.1. Surface-to-volume Ratio …………………………4

2.2. Quantum Effects …………………………5

3. Superparamagnetic Nanoparticles ………………………… 6

3.1. History and Mechanism of Magnetism …………………………6

3.2. Types of Magnetism …………………………8

3.3. Superparamagnetic Nanoparticles (SPM NPs) …………………11

4. Requirements of SPM NPs for Biomedical Applications ……………14

5. Synthesis of SPM NPs ………………………… 16

5.1. MFe2O4 NPs Synthesis ……………………………………18

5.2. Fe3O4–Based Bifunctional NPs ………………………………22

5.3. FePt and FeAu NPs ………………………………………24

6. Surface Functionalization of SPM NPs ………………………………27

7. Biomedical Applications of SPM NPs ………………………………… 29

7.1. SPM NPs as Contrast Agent in MRI …………………………32

7.2. SPM NPs as Drug Delivery Platform for Cancer Therapy .……38

7.3. SPM NPs as Mediators for Magnetic Hyperthermia ……………46

8. Summary and Conclusion ………………………………………… 48

9. Reference ……………………………………………………………… 49

xiv

Chapter 2: Synthesis and Surface Modification of Magnetite (Fe3O4) Nanoparticle……53

1. Background ………………………………………………………53

2. Synthesis and Modification …………….………………………… 54

2.1. Ligand addition with phospholipid or oleylamine modified

poly(acrylic acid) ..…………………….………………………57

2.2. Ligand exchange with dopamine modified poly(ethylene glycol)

with TsT as a linker ....…………………….…………………62

2.3. Ligand exchange with dopamine modified bifunctional

poly(ethylene glycol) ....…………………….…………………68

3. Conclusion ………………………………………………………… 77

4. Experimental…………………………………………………………….. 78

5. References …………………………………………………………….. 82

Chapter 3: Magnetite (Fe3O4) Nanoparticle for Cell Nucleus Labeling …….……84

1. Background ………………………………………...……………84

2. Fe3O4 NPs modification and functionalization .………………………… 86

3. NLS-Fe3O4 NPs for nucleus targeting ………………………………… 89

4. Summary …………………………………………………………….. 93

5. Experimental…………………………………………………………….. 94

6. References …………………………………………………………….. 96

Chapter 4: pH Controlled Release of Chromone from Chromone-Fe3O4 Nanoparticles for

Cancer Cell Growth Inhibition ……………….…….…….…….…….……98

1. Background ………………………………………...……………98

2. Fe3O4 NPs modification and functionalization .…………………………101

xv

3. Controlled chromone release from Chromone-Fe3O4 NPs …………..103

4. Summary ……………………………………………………………..106

5. Experimental……………………………………………………………..107

6. References ……………………………………………………………..113

Chapter 5: Conjugating Methotrexate to Magnetite (Fe3O4) Nanoparticles via

Trichloro-s-Triazine for Cancer Inhibition …………………….…….…115

1. Background ………………………………………...…………..115

2. Results and Discussion …………………….…………………………...118

3. Summary ……………………………………………………………..126

4. Experimental……………………………………………………………..127

5. References ……………………………………………………………..132

Chapter 6: Au-Fe3O4 Dumbbell NPs as Dual-functional Probes …………….……134

1. Background ..………………………………………...…………134

2. Results and Discussion ………………….…………………………...136

3. Summary ……………………………………………………………..144

4. Experimental……………………………………………………………..145

5. References ……………………………………………………………..149

Chapter 7: Au-Fe3O4 Dumbbell NPs for Target Specific Platin Delivery ……….……151

1. Background ..………………………………………...…………151

2. Results and Discussion ………………….…………………………...153

3. Summary ……………………………………………………………..161

4. Experimental……………………………………………………………..161

5. References ……………………………………………………………..166

xvi

Chapter 8: Controlled Release of Fe from FePt Nanoparticles for Tumor Inhibition …167

1. Background ..………………………………………...…………167

2. Results and Discussion ………………….…………………………...170

3. Summary ……………………………………………………………..180

4. Experimental……………………………………………………………..180

5. References ……………………………………………………………..186

xvii

List of Tables

2-1. Zeta potentials PEGylated Fe3O4 NPs and dextran coated Fe3O4 NPs 71

4-1. The ε values of Chromone at different pHs 111

6-1. Relaxivities r1 and r2 of Fe3O4 and Au-Fe3O4 nanoparticles with various Au core

sizes for the same Fe3O4 size at 3T (T=25˚) 142

7-1. ICP-AES analytical results in Au-Fe3O4 NPs for platin loading with or without platin

binding ligand 156

xviii

List of Figures

1- 1. Relative size of nanomaterials compared with familiar items 3

1- 2. Evolution of the dispersion F as a function of n for cubic clusters up to n=100 5

1- 3. A compass in the traditional Chinese design 7

1- 4. Diagram to show the magnetic moment produced by an electron orbiting the

nucleus and that produced by the spin of the electron 8

1- 5. Schematic illustration of different magnetism 9

1- 6. Hysteresis curve and a typical ZFC-FC magnetization measurement 10

1- 7. Nanoscale transition of magnetic nanoparticles from ferromagnetism to

superparamagnetism 12

1- 8. Illustration of the concept of superparamagnetism 13

1- 9. The preparation of monodisperse NPs in the framework of the La Mer model 18

1- 10. TEM images of Fe3O4 with different size 20

1- 11. Schematic illustration of the liquid–solid–solution (LSS) phase transfer synthesis

of various NPs 21

1- 12. Schematic illustration of the growth of Au-Fe3O4 NPs 23

1- 13. Schematic illustration of the formation of Ag-Fe3O4 NPs 23

1- 14. Schematic illustration of surface coating of Fe3O4 NPs with Au 24

1- 15. Schematic illustration of the unit cell of fcc and fct FePt 25

1- 16. TEM images of typical FePt NPs 26

1- 17. TEM images of typical FeAu NPs 27

1- 18. Schematic illustration of NP surface functionalization 28

1- 19. Macrophage uptake assay of the Fe3O4 NPs 31

xix

1- 20. MRI theory 33

1- 21. Relationship between, T2 contrast and SPM NPs. 35

1- 22. Size-dependent MR contrast effect of MnFe2O4 and Fe3O4 NPs 37

1- 23. Color maps of T2-weighted MR images of a mouse implanted with the cancer cell

line NIH3T6.7 at different time points after injection of MnFe2O4-Herceptin 38

1- 24. Schematic representation of different mechanisms by which nanocarriers can

deliver drugs to tumor 39

1- 25. A hypothetical magnetic drug delivery system shown in cross-section 42

1- 26. Surface modification of Fe3O4 NPs with MTX 44

1- 27. Schematic Release of doxorubicin in vitro from drug-loaded OA-Pluronic-

stabilized iron oxide nanoparticles. 45

1- 28. Device for magnetically induced hyperthermia 47

2- 1. Crystal structure and magnetization hysteresis curves of magnetite 54

2- 2. Mechanism of magnetite NPs formation from aqueous methods 55

2- 3. TEM images of Fe3O4 NPs synthesized with Fe(acac)3 as precursor 56

2- 4. Illustration of surface modification 57

2- 5. Structure of DSPE-PEG(2000)carboxylic acid 58

2- 6. Hydrodynamic diameter of Fe3O4 NPs 59

2- 7. Illustration of Fe3O4 NPs modified with OPA 61

2- 8. Hydrodynamic diameter change of Fe3O4 NPs in the dispersion 62

2- 9. Structure of dopamine and its proposed binding configurations with Fe 63

2- 10. 1H NMR of Dopamine, mPEG 64

2- 11. Hydrodynamic diameter of 9 nm Fe3O4 NPs 65

xx

2- 12. TEM images of 9 nm Fe3O4 NPs before and after modification 66

2- 13. Hydrodynamic diameter of Fe3O4 NPs in borate buffer at different pH values after

incubation at 70 ˚C 67

2- 14. Surface modification of Fe3O4 NPs via DPA-PEG-COOH. X=CH2NHCOCH2CH2

for PEG3000, PEG6000, PEG20000 69

2- 15. TEM images of NPs before and after ligand exchange with DPA-PEG-COOH 70

2- 16. Hydrodynamic sizes of the Fe3O4 NPs coated with different surfactants 71

2- 17. TGA analysis of Fe3O4 NPs after modification with DPA-PEG ligands 72

2- 18. IR study of Fe3O4 NPs before and after modification with DPA-PEG-COOH

ligand 73

2- 19. Size change monitoring of PEGylated Fe3O4 NPs by DLS 74

2- 20. Macrophage cell uptake of Fe3O4 NPs 76

2- 21. Ex cellular phantom study of Fe3O4 NPs’ T2 reducing effect 77

3- 1. Schematic illustration and image of NAv-Fe3O4 NPs 86

3- 2. Hydrodynamic diameters of NAv-Fe3O4 NPs 87

3- 3. Gel electrophoresis of NAv-Fe3O4 NPs 88

3- 4. Hydrodynamic diameters change of the Fe3O4 NPs in buffers 89

3- 5. Fluorescent images of NPs in HeLa cells 91

3- 6. TEM images of NPs in one HeLa cell 92

3- 7. Fluorescent microscopic images of the HeLa cells incubated with NaN3 93

4- 1. Schematic illustration and image of Chromone-Fe3O4 NPs 100

4- 2. Illustration of synthesis of DPA-PEG-NH2 101

4- 3. Fluorescent spectra of Chromone-Fe3O4 NPs 102

xxi

4- 4. IR spectra of Chromone-Fe3O4 NPs 103

4- 5. Chromone release 104

4- 6. Stability of Fe3O4-DAP-PEG-N-chromone and Fe3O4-DAP-PEG-NH2 in 1x PBS

buffer plus 10% FBS under 37 oC with pH=5 104

4- 7. HeLa cell uptake comparison of Fe3O4-DAP-PEG-N-chromone and Fe3O4-DAP-

PEG-NH2 through Fluorescent imaging 105

4- 8. Viability of HeLa cells 106

5- 1. Modification of Fe3O4 NPs MTX via TsT 117

5- 2. TEM images of Fe3O4 NPs and MTX-conjugated Fe3O4 NPs 119

5- 3. UV-Visible absorption spectra of free MTX, NH2-terminated NPs, and MTX-

conjugated NPs in water 120

5- 4. Hydrodynamic diameter of NH2–terminated NPs and MTX-conjugated NPs 121

5- 5. Cell viability for MTX-conjugated NPs, NH2-terminated NPs 122

5- 6. Intracellular uptake of MTX-conjugated NPs and NH2-terminated NPs 123

5- 7. Fluorescence images of 9L cells transfected with Rab5 to dye the early/sorting

endosomes 125

6- 1. Schematic illustration of surface functionalization of the Au-Fe3O4 NPs 136

6- 2. TEM images of Au, Fe3O4 and Au-Fe3O4 NPs 137

6- 3. MALDI mass spectra of PEG2000-Au-Fe3O4-PEG3000-EGFRA 138

6- 4. Hydrodynamic sizes of the nanoparticles shown in Figure 6-1 139

6- 5. Magnetic hysteresis loops and reflection spectra of Au-Fe3O4 NPs 139

6- 6. UV-vis spectra of Au and Au-Fe3O4 nanoparticles in water 140

6- 7. T2-weighted MRI images and reflection images of dumbbell labeled cells 142

xxii

6- 8. Reimage Figure 6-7 after three days 143

6- 9. Viability of A431 Cells with PEG-Au-Fe3O4-EGFRA 144

7- 1. Schematic illustration of Platin-Au-Fe3O4-Herceptin 153

7- 2. Images of different Au-Fe3O4 Dumbbell NPs 154

7- 3. Images of Au-Fe3O4 nanoparticles as synthesized and after modification 154

7- 4. MALDI Mass Spectra of the Au-Fe3O4 NPs before and after coupling with

Herceptin 155

7- 5. EDS characterization of S to Pt ratio for platin-Au-Fe3O4-Herceptin NPs 156

7- 6. Hydrodynamic diameter of Au-Fe3O4 nanoparticles at various functionalization

stages 157

7- 7. Reflection images of Sk-Br3 cells and MCF-7 cells after incubation with the same

concentration of platin-Au-Fe3O4-Heceptin NPs; Cisplatin release curves 158

7- 8. TEM image of the platin-Au-Fe3O4-Heceptin nanoparticles in Sk-Br3 cells 158

7- 9. Viability of Sk-Br3 cells 159

7- 10. Viability of Sk-Br3 cells and p53 expression in Sk-Br3 cells after incubation with

cisplatin or different NPs 160

8- 1. Fe release from fcc-Fe53Pt47 NPs in 0.1M HClO4 solution 168

8- 2. FePt NPs uptake by a cell through endocytosis followed by Fe release from FePt

NPs in lysosome 169

8- 3. TEM images of the as synthesized Fe40Pt60 NPs and Fe3O4 NPs 170

8- 4. Hydrodynamic diameter change of Fe40Pt60 NPs 171

8- 5. Schematic illustration of DCFH-DA conversion to DCF 172

8- 6. Fe release from FePt with different composition in PBS under different pHs 173

xxiii

8- 7. Fluorescent images of A2780 cells incubated with H2O2 and Fe3O4 NPs 174

8- 8. Schematic illustration of ROS initiated oxidation of C11-BODIPY into BODIPY

and fluorescent images of the A2780 cells 175

8- 9. TEM image of the internal part of an A2780 cell labeled with Fe3O4 NPs,

showing the intact endosome/lysosome 176

8- 10. Viability of several different cell lines incubated with Fe40Pt60 NPs 177

8- 11. Viability of A2780 cells incubated with 2,2’-bipyridine 177

8- 12. Specific targeting of LHRH labeled FePt NPs 179

8- 13. MALDI Mass Spectra of FePt-LHRH 185

  

Chapter I

Overview for the Background and Fundamental Theory of

Superparamagnetic Nanoparticle Synthesis, Characterization,

and Biomedical Applications

Abstract.

Over the last decade, there has been increased interest in “nanomaterial”, which

describes materials with structure on one dimension between 1nm and 100nm. A variety

of supermolecular ensembles, multifunctional supermolecule, carbon nanotubes, and

metal or semiconductor nanostructure have been synthesized and proposed as potential

building blocks for information storage media, cell imaging and bioprocessing devices,

and magnetic carriers. This has arisen for a variety of reasons, not the least of which is

technological advance, and the promise of control over material and device structure at

length scales far below conventional lithographic patterning technology.

Superparamagnetic nanoparticles (SPM NPs) and nanostructure are particularly

interesting and promising because those related studies provide not only information

about the structural and magnetic properties of the materials but also the opportunity to

find the potential applications in biomedical field.

This chapter will begin with the definition of nanotechnology and a short explanation

of fundamental mechanism of nanomaterials. Then we will focus on the theoretical

definition of SPM NPs, followed by a detailed explanation about the synthesis of SPM

NPs. More importantly, this chapter reveals the underground theories for several

biomedical applications together with some examples.

1  

  

1. Introduction to Nanoscience and Nanotechnology

On 29 December 1959, at the annual meeting of the American Physical Society, Richard

Feynman addressed the audience with his visionary and by now historical and legendary

lecture under the title – There is plenty of room at the bottom: Invitation to Enter a New

Field of Physics.1 With this talk on the problem of manipulating things on a small scale,

Feynman opened the field of nanotechnology.

Today, nanotechnology is already a commonly used buzzword in numerous fields of

science and everyday life. Numerous definitions have been coined to describe

nanotechnology and nanoscience and these are often used interchangeably. Based on the

explanation of U.S. Department of Energy, nanoscale science, engineering, and

technology are fields of research in which scientists are engineers are manipulating

matter at the atomic and molecular level in order to obtain materials and systems with

significantly improved properties to change the world we live in.

Ten nanometers is equal to one-thousandth the diameter of human hair. Materials

with at least one dimension between 1 and 100 nanometer scale are normally regarded as

nanomaterials. The scale of some nanomaterials systems is compared to some other

easily recognizable objects in Figure 1-1.

In general, nanomaterials may have globular, plate-like, rod-like or more complex

geometries. Once the materials’ size falls into the nanoscale, big changes corresponding

to size happen to the properties such as melting point, color (e.g. band gap and

wavelength of optical transitions), ionization potential, hardness, catalytic activity, or

magnetic properties such as coercivity, permeability and saturation magnetization.

2  

  

Figure 1-1. Relative size of nanomaterials compared with familiar items (courtesy of the Office of Basic Energy Sciences, Office of Science, U.S. Department of Energy, http://www.sc.doe.gov/bes/scale_of_things.html)

3  

  

2. Fundamental Properties of Nanomaterials

Two aspects of nanomaterials render them fundamentally different in their behavior

compared to bulk systems: surface-related properties and quantum properties.2

2. 1. Surface-to-volume ratio (A/V)3

The surface-to-volume ratio of nanoscale materials is significantly larger than that of

their bulk counterparts. If a cube is taken, it would be seen that its surface is scaled with

its radius, r2; however its volume scales with r3. The fraction of atoms at the surface is

known as the dispersion, F, and the dispersion scales with the ratio of surface area to

volume and therefore the inverse radius. Thus, we easily have

6 6

6 12 8

The r-1-dependence holds for simple geometries such as sphere and cubes, but for

complicated structures the relation is less straightforward. We need to go deeply and find

the relationship between atoms number and surface-to-volume ratio.

Let us focus on cubic clusters this time. For cubic clusters with n atoms of radius r0

along the edge the total number of atoms is N = n3, the number of atoms at the surface is

given by 6n2 for the six faces, corrected for double counts of the 12 edges (12n) and

reinstalling the 8 corners, so that the dispersion F becomes3

61

2 8

6

6

The conclusion from two equations is that all properties that are related to the

dispersion of surface groups will result in a dependence on the inverse radius of the

4  

  

particle and also on the number of the atoms by N-1/3 as depicted in Figure 1-2.3 On the

basis, we obtain F=0.4 for N=103, and F=0.04 for N=106.3

Figure 1-2. Evolution of the dispersion F as a function of n for cubic clusters up to n=100 (N=106). The structure of the first four clusters is displayed.3

The dispersion of surface atoms is also known as the coordination number, <NN>,

and describes the number of nearest neighboring atoms. Atoms or molecules at and near

the surface, and even more at edges and corners, have fewer neighbors and are therefore

less strongly bound than those in bulk. This is why the surface has a higher energy, why

often melts first, and why it affects many other properties of the particles.

2. 2. Quantum Effects

It has been found that the electronic structure of small particles is generally discrete

and not overlapping as is the case with bulk material phases. This is due to confinement

of the electron wave functions of certain physical dimensions of the nanoparticles.

As with most orbital systems, electrons can be found at different energy levels, and

the average spacing of this energy level is known as the Kubo gap, δ. By considering the

5  

  

lowest unoccupied energy state of the electronic system of a bulk material, the Femi

energy, Ef, could be incorporated in describe the Kubo gap:

4 /3

where n is representing the number of valence electrons in the nanosystems.4 In the case

where the thermal energy of systems exceeds the Kubo gap, they will behave metallically,

and if the thermal energy does not exceed this value, they will behave non-metallically.

This change is especially prevalent in small systems at the nanoscale and explains why

certain materials become magnetic or electrically conductive at the nanoscale.

Differences in optical properties are also noted for nanosystems that are observed as

luminescence and size-dependent color changes of certain metallic nanoparticles.

3. Superparamagnetic Nanoparticles

3. 1. History and Mechanism of Magnetism

In early as 4th century BC, the ancient Chinese had described the magnet in Book of the

Devil Valley Master: “The lodestone makes iron come or it attracts it”. Later, the Chinese

people began to use the magnetic needle compass to improve the accuracy of navigation

by employing the astronomical concept of true north (Figure 1-3).5 However, an

understanding of the relationship between electricity and magnetism just began in 1819

with work by Han Christian Oersted at University of Copenhagen, who discovered more

or less by accident that an electric current could influence a compass needle (Oersted’s

Experiment). Osersted’s discovery later led to Ampere observing that the magnetic field

of a solenoid being identical to that of a magnet.

6  

  

Figure 1-3. a) A compass in the traditional Chinese design (courtesy from Dr. Siry’s website, http://web.rollins.edu/~jsiry/)

Ampere then hypothesized that all magnetic effects were due to current loops and that

the magnetic effects in materials must be due to “molecular currents”, attributed to the

movement of electrons. But his model was not enough to explain why the predicted

current is larger than the actual one. Dirac in 1928 postulated electron spin also

contributed to the magnetism.

The spin of an electron is hard to visualize, but has the properties of a small magnetic

moment pointing either “up” or “down”. Within an atom, electrons are arranged in

orbitals, with a maximum of two electrons with opposite spin occupying each orbital

(due to the Pauli Exclusion Principle). The orbitals are further grouped into shells. In all

atoms except for hydrogen there is more than one electron and these electrons can

interact with each other as well as with the nucleus, leading to “coupling”.

In summary, the total magnetic moment of a free atom has two contributions from

each electron (Figure 1-4):

1. The angular momentum as the electron orbits the nucleus (strictly, the momentum

of the nucleus relative to the orbiting electron). This is effectively Ampère’s

molecular current and is known as the orbital contribution.

2. The ‘spin’ of the electron itself

7  

  

Figure 1-4. Diagram to show the magnetic moment produced by an electron orbiting the nucleus and that produced by the spin of the electron.

3. 2. Types of Magnetism

The overall magnetic behavior of a material can vary widely, depending on the structure

of the material, and particularly on its electron configuration. Several forms of magnetic

behavior have been observed in different materials, including: paramagnets, ferromagnets,

superparamagnets, antiferromagnets and ferrimagnets.

Paramagnetism. In a paramagnet, the magnetic moments tend to be randomly orientated

due to thermal fluctuations when there is no magnetic field. In an applied magnetic field

these moments start to align parallel to the field such that the magnetization of the

material is proportional to the applied field.

Ferromagnetism. The magnetic moments in a ferromagnet have the tendency to become

aligned parallel to each other under the influence of a magnetic field. However, unlike the

moments in a paramagnet, these moments will then remain parallel when a magnetic field

is not applied.

Antiferromagnetism. Adjacent magnetic moments from the magnetic ions tend to align

anti-parallel to each other without an applied field. In the simplest case, adjacent

magnetic moments are equal in magnitude and opposite therefore there is no overall

magnetisation.

8  

  

Ferrimagnetism. The aligned magnetic moments are not of the same size; that is to say

there is more than one type of magnetic ion. An overall magnetisation is produced but not

all the magnetic moments may give a positive contribution to the overall magnetisation.

Superparamagnetism. A superparamagnetic material is composed of small

ferromagnetic clusters, but where the clusters are so small that they can randomly flip

direction under thermal fluctuations. As a result, the material as a whole is not

magnetized except in an externally applied magnetic field (i.e. it is like paramagnetism).

  

Figure 1-5. a) Schematic showing the magnetic dipole moments randomly aligned in a paramagnetic sample; b) Schematic showing the magnetic dipole moments aligned parallel in a ferromagnetic material; c) Schematic showing adjacent magnetic dipole moments with equal magnitude aligned anti-parallel in an antiferromagnetic material. This is only one of many possible antiferromagnetic arrangements of magnetic moments; d) Schematic showing adjacent magnetic moments of different magnitudes aligned anti-parallel.

a b

c d

The Basic Parameters and Measurements

The relevant properties are the strength of the applied magnetic field, H, the magnetic

flux density inside a medium, B and the magnetization, M. They are related via

1

where μ0 = 12.566 x 10-7 VsA-1m-1 is the vacuum permeability and μr is the dimensionless

relative permeability which gives the enhancement factor of B over μ0H due to

magnetization of the medium. In the regime where the magnetization scales linearly with

H, it is useful to define the magnetic susceptibility, X

9  

  

Magnetisation and susceptibility are given per volume, per mass unit or per mol. μeff is

the effective atomic or molecular magnetic moment.

Two principal measurements are normally carried out: M(H), the magnetization as a

function of applied field at a given temperature (hysteresis loop, Figure 1-6a), and M(T),

the magnetization as a function of temperature at a fixed field (zero-field-cooled and field

cooled magnetization curves, Figure 1-6b).

Figure 1-6. (a) Hysteresis curve of the magnetization M(H) of a ferromagnetic material. The intrinsic properties are the saturation magnetization Ms, the remanent magnetization Mr and the coercive field Hc. The arrows give the cycling direction. (b) A typical ZFC-FC magnetization measurement

Figure 1-6a shows the typical symmetric hysteresis behavior of M(H) that is obtained

on cycling the external field to values beyond the magnetic fields where the

magnetization reaches its saturation value, ±Ms(T). The curves cross zero external field at

the remanent magnetizations, ±Mr(T), and the magnetizations become zero at the

coercive fields, ±Hc(T). The first application of an external field starts at zero and results

in the virgin curve. The ratio of the remanent magnetization to the saturation

magnetization, Mr/Ms, is called the remanence ratio and varies from 0 to 1.

The temperature-dependent magnetization data measured in zero-field-cooled (ZFC)

and field cooled (FC) procedures are usually used to obtain the information of the energy

10  

  

barriers (Figure 1-6b). The ZFC-FC magnetization measurement is carried out as follows.

For the ZFC curve, the sample is first cooled in a zero field from a high temperature well

above blocking temperature (TB), where nanoparticles are in a superparamagnetic state,

down to a low temperature well below TB, where nanoparticles are in a ferromagnetic

state. Then a magnetic field is applied and the magnetization as a function of temperature

is measured in the warming process to a temperature well above the blocking temperature.

The FC curve is obtained by measuring the magnetization when cooling the sample to the

low temperature in the same field. In the ZFC and FC measurements the field must be

weak enough in comparison with the anisotropy field to guarantee that the ZFC-FC curve

reflects the intrinsic energy barrier distribution.6

3. 3. Superparamagnetic (SPM) Nanoparticles (NPs)

The underlying physics of superparamagnetism is founded on an activation law for the

relaxation time τ of the new magnetization of the particle:

exp ∆

∆ KV

where ∆E is the energy barrier to moment reversal, and kBT is the thermal energy. For

non-interacting particles the pre-exponential factor τ0 is often of the order 10-10 – 10-12 s

and weakly dependent on temperature. The energy barrier has several origins, including

both intrinsic and extrinsic effects such as magnetocrystalline and shape anisotropies.

However for uniaxial anisotropies, ∆E is equal to the product of the anisotropy constant

and the volume.

11  

  

This direct proportionality between ∆E and V is the reason that superparamagnetism –

the thermally activated flipping of the net moment direction – is important for small

particles, since for them ∆E is comparable to kT at room temperature.

Under certain temperature, bulk materials have magnetic anisotropic energies that are

much larger than the thermal energy (kT) (Figure 1-7a, blue line). Thus the thermal

energy of the nanoparticle is insufficient to readily invert the magnetic spin direction, so

the material is ferromagnetic. However, the reduced size of nanoparticles results a much

smaller anisotropic energy compared with thermal energy, which is sufficient to invert

the spin direction (Figure 1-7a, red line). Such magnetic fluctuation leads to a net

magnetization of zero, which means superparamagnetism. For example, γ-Fe2O3

nanoparticles of 55nm exhibit ferromagnetic behavior with a coercivity of 52 Oe at 300k,

but smaller 12nm sized γ-Fe2O3 nanoparticles show superparamagnetism with no

hysteresis behavior (Figure 1-7b,c).

Figure 1-7. Nanoscale transition of magnetic nanoparticles from ferromagnetism to superparamagnetism: (a) energy diagram of magnetic nanoparticles with different magnetic spin alignment, showing ferromagnetism in a large particle (top) and superparamagentism in a small nanoparticle (bottom); (b, c) size dependent transition of iron oxide nanoparticles from superparamagnetism to ferromagnetism showing TEM images and hysteresis loops of (b) 55 nm and (c) 12 nm sized iron oxide nanoparticles. (Adopted from Reference7 )

12  

  

For a certain size of nanoparticles, the change of temperature can also induce the

transition between ferromagnetic and superparamagnetic. The transition temperature

from ferromagnetism to superparamagnetism is referred to as the blocking temperature

(Tb) and measured through zero-field cooled/field cooled set of measurement as

mentioned above. In ZFC curve, the peak temperature is normally the blocking

temperature TB.

However, we must realize that observations of superparamagnetism are dependent on

both temperature and measurement time τm of the experimental technique being used

(Figure 1-8). If τ << τm the flipping is fast relative to the experimental time window and

the particles appear to be paramagnetic; while if τ >> τm the flipping is slow and quasi-

static properties are observed – the so-called “blocked” state of the system. A “blocking

temperature” TB is defined as the mid-point between these two states, where τ = τm.

Figure 1-8. Illustration of the concept of superparamagnetism, where the circles depict three magnetic nanoparticles and the arrows represent the net magnetization direction in those particles. In case (a), at temperatures well below the measurement-technique-dependent blocking temperature TB of the particles, or for relaxation times τ (the time between moment reversals) much longer than the characteristic measurement time τm, the net moments are quasi-static. In case (b), at temperature well above TB, or for τ much shorter than τm, the moment reversals are so rapid that in zero external field the time-averaged net moment on the particles is zero.

13  

  

4. Importance and Requirements of SPM NP for Biomedical Applications

Superparamagnetic (SPM) nanoparticles (NPs) have been considered as attractive

magnetic probes for biological imaging and therapeutic applications due to two main

reasons. One is the high surface-to-volume ratio, which enables the maximum loading

molecules or largest interaction interface. Another one is the superparamagnetic property.

In normal biological conditions, these SPM NPs are not subject to strong magnetic

interactions in the dispersion due to the randomization of their magnetization and are

readily stabilized in physiological conditions. Under an external magnetic field, however,

they exhibit a magnetic signal far exceeding that from any of the known biomolecules

and cells. This makes SPM NPs readily identified by a magnetic sensing device from the

ocean of biomolecules. At a core diameter at less than 20 nm and overall hydrodynamic

diameter at less than 50 nm, these NPs have the size that is comparable to the nuclear

pore size (~50 nm) and is much smaller than a cell (normally 10 – 30 μm). Once coupled

with a target agent, they can serve as a nano-vector and interact specifically with

biomolecules of interest through well established biological interactions, providing

controllable means of magnetically tagging bio-identity. Under the normal range of

magnetic field strengths used in magnetic resonance imaging (MRI) scanners (usually

higher than 1 Tesla), these SPM NPs in the targeted area can be magnetically saturated,

establishing a substantial locally perturbing dipolar field that leads to a marked

shortening of proton relaxation (T2 relaxation) in MRI process and giving a “darker”

image of the targeted area over the biological background. The active investigation about

SPM NPs as MR imaging contrast agents has led several commercial products, i.e. bowel

14  

  

contrast agents (Lumiren® and Gastromark®) and liver/spleen imaging (Endorem® and

Feridex IV®).

Furthermore, under an alternating magnetic field with controlled field amplitude and

field reversal frequency, magnetization of the SPM NPs attached to the bio-identity can

be switched back and forth. This magnetization re-orientation may result from either a

physical rotation of the particle (Brownian relaxation), which creates frictions between

the NP and its surrounding liquid medium, or internal magnetization switching from one

direction to another (Néel relaxation).In both cases, these SPM NPs function as a heater

to heat the area they target to. This magnetic field induced NP heating has been known as

magnetic fluid hyperthermia and has been studied extensively for future cancer therapy.

As the last, but not the least application, SPM NPs have been evaluated extensively

for targeted delivery of pharmaceuticals through magnetic drug targeting and by active

targeting through the attachment of high affinity ligands. In the spirit of Ehrlich’s “Magic

Bullet”, SPM NPs have the potential to overcome limitations associated with systemic

distribution of conventional chemotherapies. With the ability to utilize magnetic

attraction and/or specific targeting of disease biomarkers, SPM NPs offer an attractive

means of remotely directing therapeutic agents specifically to a disease site, while

simultaneously reducing dosage and the deleterious side effects associated with non-

specific uptake of cytotoxic drugs by healthy tissue. Furthermore, the use of MNP as

carriers in multifunctional nanoparticles as a means of real-time monitoring of drug

delivery is of intense interest.

These potential biomedical applications of magnetic NPs require that SPM NPs are

monodisperse so that each individual nanoparticle has nearly identical physical and

15  

  

chemical properties for controlled biodistribution, bioelimination and contrast effects.8

The SPM NPs should also have high magnetic moment, and can be modified via surface

chemistry reactions so that they are capable of binding specifically to the bimolecular of

interest and able to with stand various physiological conditions.

A significant challenge associated with the application of these MNP systems is their

behavior in vivo.9 The efficacy of many of these systems is often compromised due to

recognition and clearance by the reticuloendothelial system (RES) prior to reaching target

tissue, as well as by an inability of to overcome biological barriers, such as the vascular

endothelium or the blood brain barrier. The fate of these MNP upon intravenous

administration is highly dependent on their size, morphology, charge, and surface

chemistry. These physicochemical properties of nanoparticles directly affect their

subsequent pharmacokinetics and biodistribution. To increase the effectiveness of MNPs,

several techniques, including reducing size and grafting nonfouling polymers, should be

employed to improve their “stealthiness” and increase their blood circulation time to

maximize the likelihood of reaching targeted tissues.

5. Synthesis of SPM NPs

SPM NPs can be synthesized by a variety of methods ranging from traditional co-

precipitation of metal salts in basic solution, high temperature organic phase

decomposition, and chemical vapor deposition. SPM NPs, iron oxide nanoparticles used

in biomedical applications are often synthesized by the co-precipitation of ferrous and

ferric ions at 1-to-2 ratio in an alkaline medium.10 In order to control the NPs’ growth and

stabilize the NPs from agglomeration, different kinds of polymers were added during the

16  

  

synthesis, such as dextran, dendrimer, and poly(aniline), which were coated onto NPs’

surface to create steric or statistic repulsion hence balancing the attraction forces among

NPs. In these hydrolytic processes, the control of the solution of pH value and the

presence of the coating material serving as a surfactant are critical to particles formation

and properties. However, the co-precipitation method can control NPs’ shape, size,

crystallinity and magnetic properties, which can vary vastly among synthesis methods

even within particles of similar size due to incorporation of impurities disrupting the

crystal structure, as well as the surface effects described previously.9

In recent years, high-temperature organic phase reductive decomposition of metal salt

or organometallic precursors has been applied to produce monodisperse SPM NPs.11

Classic studies by La Mer & Dinegar show that the production of monodisperse colloids

requires a temporally discrete nucleation event followed by slower controlled growth on

the existing nuclei (Figure 1-9).12 Rapid addition of reagents to the reaction vessel raises

the precursor concentration above the nucleation threshold. A short nucleation burst

partially relieves the supersaturation. As long as the consumption of feedstock by the

growing colloidal NPs is not exceeded by the rate of precursor addition to solution, no

new nuclei form. Since the growth of any one NC is similar to all others, the initial size

distribution is largely determined by the time over which the nuclei are formed and begin

to grow. If the percentage of NPs growth during the nucleation period is small compared

with subsequent growth, the NPs can become more uniform over time.

Many systems exhibit a second, distinct, growth phase called Ostwald ripening.13 In

this process, the high surface energy of the small NCs promotes their dissolution,

whereas material is redeposited on the larger NCs. The average NC size increases over

17  

  

time with a compensating decrease in NC number. Exploiting Ostwald ripening can

greatly simplify the preparation of a size series of NCs.14 Portions of the reaction mixture

can be removed at increments in time, as depicted in Figure 1-9.

Figure 1-9. (a) Cartoon depicting the stages of nucleation and growth for the preparation of monodisperse NPs in the framework of the La Mer model; (b) Representation of the simple synthetic apparatus employed in the preparation of monodisperse NPs samples.15

5.1 MFe2O4 NPs Synthesis

Magnetic ferrite MFe2O4 NPs, especially magnetite Fe3O4 NPs, are widely studied

due to their chemical and magnetic stability. This oxide represents a well-known and

important class of iron oxide materials where oxygen forms an fcc packing, and M2+

and Fe3+ occupy either tetrahedral or octahedral interstitial sites. By adjusting the

chemical identity of M2+, the magnetic configurations of MFe2O4 can be molecularly

engineered to provide a wide range of magnetic properties.

MFe2O4 NPs are commonly made by hydrolysis/condensation of M2+ and Fe3+

ions by a base, usually NaOH, or NH3•H2O, in an aqueous solution,16-18 or in reverse

micelles.19 Although this coprecipitation method is suitable for mass production of

18  

  

magnetic MFe2O4 ferrofluids, it requires careful adjustment of the pH value of the

solution for particle formation and stabilization, and it is difficult to control sizes and

size distributions, particularly for particles smaller than 20 nm. An alternative

approach to make monodisperse iron oxide NPs is via high temperature organic phase

decomposition of Fe(CO)5 in the presence of (CH3)3NO, air or 3-chloro-

peroxybenzoic acid.20-22

More conveniently, MFe2O4 NPs are synthesized by reductive decomposition of

metal acetylacetonates or carboxylates in an organic phase.23-25 For example,

monodisperse Fe3O4 NPs are prepared by high temperature (up to 305°C) reductive

decomposition of Fe(acac)3 in the presence of a long chain 1,2-hydrocarbon diol,

oleic acid and oleylamine.26 MFe2O4 NPs (with M = Co, Ni, Mn, etc) are made by

simply adding a different metal acetylacetonate precursor to the mixture of reactants

used for Fe3O4 synthesis.23 The size of the NPs is controlled by varying the reaction

temperatures or changing the concentrations of metal precursors. Alternatively, with

the smaller NPs as seeds, larger NPs up to 20 nm in diameter can be synthesized by

seed mediated growth.

Fe3O4 NPs can also be made by chemical conversion of FeO NPs.27 FeO NPs are

synthesized by heating the mixture of Fe(acac)3, oleic acid and oleylamine. When

treated under atmospheric pressure and air at 120°C for 90 min, the as-synthesized

FeO NPs are converted to Fe3O4 NPs. Using this method, large Fe3O4 NPs up to 100 nm

in diameter can be made.

By combining decomposition/oxidation of Fe(CO)5 and reductive decomposition

of iron oleate complex, 1 nanometer size control of iron oxide NPs can be

19  

  

achieved.20,28-30 Monodisperse NPs with particle sizes of 6, 7, 9, 10, 12, 13 and 15 nm

have been produced. The monodispersity of the NPs can be readily seen in the

representative TEM images of NPs in Figure 1-10.

Figure 1-10. TEM images of a) 6-, b) 7-, c) 8-, d) 9-, e) 10-, f) 11-, g) 12-, and h) 13-nm-sized air-oxidized iron oxide nanoparticles showing the one nanometer level increments in diameter Reproduced with permission from reference30.

Large-scale synthesis of iron oxide NPs is achieved through high temperature

decomposition of iron oleate.29 In the synthesis, iron chloride reacts with sodium

oleate to form a waxy iron-oleate complex that is subject to further thermal

decomposition at 320°C in 1-octadecene, leading to the formation of monodisperse

iron oxide NPs. Another method that has the potential for large scale synthesis of

magnetic NPs is via a liquid-solid-solution (LSS) phase transfer.31 The chemistry for

the synthesis is illustrated in Figure 1-11. It involves the reaction of metal precursor

at the interfaces of metal linoleate (solid), ethanol–linoleic acid liquid phase (liquid)

and water–ethanol solutions (solution) at different designated temperatures. A phase

transfer process occurrs spontaneously across the interface of the solid and the

20  

  

solution. The NPs generated at the interface are coated with a layer of linoleic acid,

resulting in a spontaneous phase-separation and the formation of hydrophobic NPs

that are easily collected at the bottom of the container.

Figure 1-11. Schematic illustration of the liquid–solid–solution (LSS) phase transfer synthesis of various NPs. Reproduced with permission from reference31.

Recently, the ultra-small Fe3O4 NPs ranging from 2.5 nm to 5 nm were made by

thermal decomposition of Fe(CO)5 in benzyl ether at 300°C followed by room

temperature air oxidation. Different from the previous synthesis methods, this preparation

used a small molecule 4-methylcatechol (4-MC) as the surfactant and the sizes of the NPs

were tuned by the MC/Fe ratio.32 More importantly, the 4-MC coated Fe3O4 NPs can be

directly conjugated with a peptide, c(RGDyK), via the Mannich reaction, rendering the

biocompatible SPM NPs with a hydrodynamic diameter of around 8 nm, suitable for

target-specific delivery and imaging applications.

21  

  

5.2 Fe3O4–Based Bifunctional NPs

Bifunctional NPs are those containing two different nanoscale functionalities within

one integrated identity. Dumbbell-like and core/shell NPs are two representative

bifunctional systems that have shown great potential for biomedical applications.

The dumbbell-like Au-Fe3O4 NPs are prepared via the decomposition of iron

pentacarbonyl, Fe(CO)5, at 300°C over the surface of the pre-formed Au NPs

followed by oxidation in air, as illustrated in Figure 1-12a.33 The Au NPs can be

either synthesized in situ by injecting HAuCl4 solution into the reaction mixture or

pre-made in the presence of oleylamine. The size of the Au NPs is tuned by

controlling the temperature at which the HAuCl4 solution is added, or by controlling

the HAuCl4/oleylamine ratio. The size of the Fe3O4 NPs is controlled by amount of

Fe(CO)5 added in the reaction mixture. Figure 1-12b shows the TEM image of the

Au-Fe3O4 NPs with Fe3O4 at around 14 nm and Au at 8 nm. Figure 1-12c is a typical

high-resolution TEM (HRTEM) image of a dumbbell-like NP with Fe3O4 at 12 nm

and Au at 8 nm. In the structure, a Fe3O4 (111) plane grows onto an Au (111) plane,

giving the dumbbell-like structure. These dumbbell-like NPs show a plasmonic

absorption at around 530 nm and have a suturation magnetization of 80 emu/g – a

value close to the pure Fe3O4 NPs.33

22  

  

Figure 1-12. (a) Schematic illustration of the growth of Au-Fe3O4 NPs. (b) TEM and (c) HRTEM images of the Au-Fe3O4 NPs. Reproduced with permission from reference33.

Different from Au-Fe3O4 NPs, the Ag-Fe3O4 NPs are made by controlled nucleation

of Ag on the pre-formed Fe3O4 NPs.34 In the synthesis, the as-prepared Fe3O4 NPs

dispersed in organic solution and AgNO3 dissolved in water are mixed and agitated by

ultrasonication. The sonication provides the energy required for the formation of a

microemulsion with the Fe3O4 NPs assembling at the liquid/liquid interface. Fe(II) on the

NPs acts as catalytic center for the reduction of Ag+ and nucleation/growth of Ag NPs, as

illustrated in Figure 1-13. The partial exposure of the NPs to the aqueous phase causes

the formation of Ag-Fe3O4 NPs. The NPs show a plasmonic absorption from Ag NPs and

the same magnetic hysteresis behavior as Fe3O4 NPs.

Figure 1-13. Schematic illustration of the formation of Ag-Fe3O4 NPs in a micellar structure by ultrasonication of a heterogeneous solution with as-prepared Fe3O4 NPs in the organic phase and AgNO3 in water. Reproduced with permission from reference34.

23  

  

Core/shell NPs are another group of NPs that can incorporate multifunctionality into

one structure. One recent example is Fe3O4/Au or Fe3O4/Au/Ag NPs.35 In the synthesis,

the pre-made Fe3O4 NPs are mixed with a solution of HAuCl4 and oleylamine. HAuCl4 is

reduced under this condition, forming a thin layer of Au shell over the Fe3O4 surface

(Figure 1-14a). The surface of the particles is then treated with sodium citrate and

cetyltrimethylammonium bromide (CTAB). NPs treated this way are water soluble and

can serve as seeds to grow more Au or Ag on their surface. The thicker coating is

achieved by mixing the seeding Fe3O4/Au NPs with HAuCl4 or AgNO3 and ascorbic acid

in the presence of CTAB, and incubating the mixture at 30°C (Figure 1-14b). The control

on shell thickness allows the tuning of plasmonic properties of the core/shell NPs to be

either red-shifted (to 560 nm with more Au coating) or blue-shifted (to 501 nm with more

Ag coating).

Figure 1-14. (a) Schematic illustration of surface coating of Fe3O4 NPs (i) with Au to form hydrophobic Fe3O4/Au NPs (ii) and hydrophilic Fe3O4/Au NPs (iii). (b) Schematic illustration of the formation of Fe3O4/Au and Fe3O4/Au/Ag and the control on the plasmonic properties. Reproduced with permission from reference35.

5.3 FePt and FeAu NPs

FePt NPs containing a near-equal atomic percentage of Fe and Pt are an important class

of magnetic nanomaterials. They are known to have a chemically disordered face-

24  

  

centered cubic (fcc) structure or a chemical ordered face-centered tetragonal (fct)

structure, as shown in Figure 1-15. The fcc-structured FePt has a small coercivity and is

magnetically soft. The fully ordered fct-structured FePt can be viewed as alternating

atomic layers of Fe and Pt stacked along the [011] direction. (c-axis in Figure 1-15b). The

anisotropy constant K, which measures the ease of magnetization reversal along the easy

axis, can reach as high as 107 Jm–3,36 a value that is one of the largest among all known

hard magnetic materials. This large K is caused by Fe and Pt interactions originating from

spin-orbit coupling and the hybridization between Fe 3d and Pt 5d states.37,38 These Fe–Pt

interactions further render the fct-FePt nanoparticles chemically much more stable than

the common high-moment nanoparticles of Co and Fe, as well as the large coercive

materials CoSm5 and Nd2Fe14B, making them especially useful for practical applications

in solid-state devices and biomedicine.

b a

Figure 1-15. Schematic illustration of the unit cell of (a) chemically disordered fcc and (b) chemically ordered fct FePt. Reproduced with permission from reference 39.

FePt NPs are normally synthesized through the thermal decomposition of iron

pentacarbonyl, Fe(CO)5, and reduction of platinum acetylacetonate, Pt(acac)2, in the

presence of 1,2-alkanediol.40 The synthetic chemistry is illustrated in Figure 1-16a.

Fe(CO)5 is thermally unstable and subject to decomposition at high temperature to carbon

monoxide and Fe. Pt(acac)2 is readily reduced by 1,2-alkanediol to Pt. A small group of

25  

  

Fe and Pt atoms combine to form Fe-Pt clusters that act as nuclei. The growth proceeds

as more Fe-Pt species deposit around the nuclei, forming FePt NPs (Figure 1-16b). Oleic

acid and oleyamine are used for surfactant. In the reaction, the composition or Fe-to-Pt

ratio in NPs is controlled by Fe(CO)5/Pt(acac)2 ratio. And the size of FePt NPs could be

achieved through seed-mediated growth or controlling the surfactant to metal ratio.41,42

A better and easier way to control the size of FePt NPs is obtained via a one-step

simultaneous thermal decomposition of Fe(CO)5 and reduction Pt(acac)2 in the absence

of 1,2-alkanedio.43 In this case, the reduction of Pt is much slower, which allows more

metal mixture to deposit onto the nuclei. Through controlling the heating rate, the size of

FePt could be tuned between 4nm to 12nm (Figure 1-16c,d).

Figure 1-16. (a) Schematic illustration of FePt NPs formation from the decomposition of Fe(CO)5 and reduction of Pt(acac)2; (b) 4nm FePt NPs; (c) 6nm FePt NPs; (d) 9nm FePt NPs. (Scale bar: 20nm) Reproduced with permission from references 40,43

Similar to FePt, monodisperse FeAu NPs with different Au/Fe ratio (Figure 1-17) can

be synthesized via the reduction of gold acetate by 1,2-hexdecanediol and the thermal

decomposition of iron pentacarbonyl in the presence of the stabilizers oleic acid and

oleyamine.44 The incorporation of Au into Fe NPs leads to a structural change from body-

centered cubic (bcc) to face-centered cubic (fcc). The resultant FeAu NPs possess of the

optical properties of Au NPs and the magnetic properties of Fe NPs. Alternatively, FeAu3

26  

  

alloy NPs could be made through HAuCl4 and Fe(acac)3 with n-butyllithium as reducing

agent.45

Figure 1-17. (a) TEM images of FeAu nanoparticles with a Au/Fe molar ratio of 0.5; Reproduced with permission from reference44 (b, c) 20nm Au3Fe NPs with their Powder XRD data and the fcc structure. Reproduced with permission from reference45

6. Surface Functionalization of Superparamagnetic Nanoparticles

The SPM NPs prepared above are normally stabilized by the surfactants like oleic

acid or oleylamine or their combination. The formation of metal carboxylate and/or

metal–amine bonds at the interface leaves NPs surrounded with a layer of

hydrocarbon, making them hydrophobic and only soluble in non-polar or weakly

polar organic solvents. For NPs to be useful in a biological system, they must be

water soluble and stable at various pH values ranging from 5 to 9, at salt

concentrations at hundreds of mM, and at various cell culture temperatures. They

must also achieve target-specific binding in biological systems.

27  

  

Figure 1-18. Schematic illustration of NP surface functionalization via (a) surfactant addition and (b) surfactant exchange. Reproduced with permission from reference 8

Surfactant addition and surfactant exchange are two general approaches for NP

surface functionalization, as illustrated in Figure 1-18.8 Surfactant addition is

achieved through the adsorption of amphiphilic molecules that contain both a

hydrophobic segment and a hydrophilic component. The hydrophobic segment forms

a double layer structure with the original hydrocarbon chain, while hydrophilic groups

expose to the outside of the NPs, rendering them water-soluble. The NP

biocompatibility can be further improved by using biodegradable amphiphilic

polymers originally developed for drug delivery applications.46,47 One of the most

widely utilized and successful polymers has been the functionalized

phospholipids.48,49 As various phospholipids or amphiphilic polymers are

28  

  

commercially available, this addition method offers a convenient approach to

functionalize NPs with biotin, –COOH, –SH and/or –NH2, facilitating their

conjugation with fragments of DNA, proteins, peptides, or antibodies.

Surfactant exchange is the direct replacement of the original surfactant with a new

bifunctional surfactant. This bifunctional surfactant has one functional group capable

of binding to the NP surface tightly via a strong chemical bond and the second

functional group at the other end with a polar character so that the NPs can be

dispersed in water or be further functionalized. Various monomeric species, such as

dimercaptosuccinic acid, dopamine and peptides have been applied for this NP

functionalization purpose.50,51

The functionalization described above offer the SPM NPs with robust colloidal

and bio-stability. The targeting agents coupled with these NPs allow facile

biorecognition event via strong biological interactions. These, plus their

multifunctional magnetic and optical properties, make SPM NPs ideal for diagnostic

and therapeutic applications.

7. Biomedical Applications of SPM NPs

For SPM NPs to be useful for biomedical applications, they should be first stabilized

against the absorption of plasma proteins and non-specific uptake by reticular-

endothelial system (RES), like macrophage cells.52 Due to their large surface area,

when exposed to a physiological environment, these NPs tend to interact with plasma

proteins, causing size increase that often results in serious agglomeration. They may

be also considered as an intruder by the innate immune system and be readily

29  

  

recognized and engulfed by the macrophage cells. In both cases, the particles will be

removed from the blood circulation and lose their function, leading to dramatic

reduction in efficiency in NP-based diagnostics and therapeutics. To inhibit the

plasma coating and to escape from the RES for longer circulation times, the NPs are

usually coated with a layer of hydrophilic and biocompatible polymer such as dextran,

dendrimers, polyethylene glycol (PEG), or polyethylene oxide (PEO).53,54

As an amphiphilic polymer and a non-specific interaction reducing reagent,54

polyethylene glycol (PEG) was used recently to functionalize monodisperse 9 nm

Fe3O4 NPs.55 In this study PEG is anchored on the Fe3O4 NPs through a covalent bond

as illustrated in Figure 1-19a. These PEG coated Fe3O4 NPs are incubated with the

RAW 264.7 cells - one kind of mouse macrophage cell line, at three different

concentrations: 0.1 mg Fe/ml, 0.01 mg Fe/ml and 0.001 mg Fe/ml. The results from

the 0.01mg Fe/mL of PEG-coated samples are shown in Figure 1-19b. It can be seen

that the dextran coated NPs give the highest uptake, followed by PEG600 coated NPs,

of which the uptake is about 30%-50% of that from the dextran coated ones. For

PEG3000, PEG6000 and PEG20000 coated NPs, their uptake are comparative with the

background, indicating negligible uptake of these NPs by the macrophage cells.55

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Figure 1-19. (a) Schematic of ligand exchange reaction on the surface of Fe3O4 NPs (b) Macrophage uptake assay of the Fe3O4 NPs in (a) with initial Fe concentration at 0.01 mg Fe/ml. Reproduced with permission from reference55.

To act as a sensitve probe for cell imaging, a NP must be taken up by cells to

“stain” the cells. As in normal biological trasport process, this uptake can be either

passive or active, or both. Passive uptake utilizes diffusion concept and is often

concentration driven and has no targeting capability. Active uptake, on the other

hand, involves receptor-mediated endocytosis. As fast grown cells, especially tumor

31  

  

cells, often over-express certain receptors of folic acid, sugars, peptides, proteins, or

antibodies. NPs coupled with these molecules tend to be recognized by these cells and

endocytosed for internalization, achieving target-specific binding.

7.1 SPM NPs as Contrast Agent in MRI

MRI is one of the most powerful non-invasive imaging modalities utilized in clinical

medicine. It’s based on the principle that protons align and precess along an applied

magnetic field (Figure 1-20a,b). Upon applying a transverse radiofrequency pulse,

these precessed protons are perturbed from the magnetic field direction (Figure 1-

20c,d). The subsequent process, through which the pulsing field is turned off to allow

protons to return to their original state, is referred to as relaxation. Two independent

relaxation processes, longitudinal relaxation (T1-recovery, Figure 1-20e) and

transverse relaxation (T2-decay, Figure 1-20f), are utilized to generate a bright and a

dark MR image respectively.

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Figure 1-20. (a) With no external magnetic field present, spins rotate about their axes in random direction. (b) In the presence of a magnetic field, slightly more spins align parallel to the main magnetic field, B0, and thus produce longitudinal magnetization, Mz. (c) A radiofrequency pulse tips the magnetization vector by exactly 90oC, causing the entire longitudinal magnetization to flip over and rotate into transverse magnetization, Mxy (d). (e) T1 relaxation. Decay of transverse magnetization and regrowth of magnetization along the z-axis require an exchange of energy. (f) T2 and T2* relaxation. Spins get out of phase (lose phase coherence), resulting in the loss of transverse magnetization without energy dissipation. Reproduced with permission from Reference 56.

33  

  

T1: Longitudinal Relaxation

Transverse magnetization decays and the magnetic moments gradually realign with

the z-axis of the main magnetic field B0, as discussed previously. The transverse

magnetization remaining within the xy-plane-strictly speaking the projection of the

magnetization vector onto the xy-plane (Figure 1-20e)-decreases slowly and the MR

signal fades in proportion. As transverse magnetization decays, the longitudinal

magnetization, Mz – the projection of the magnetization vector onto the z-axis – is slowly

restored. This process is known longitudinal relaxation or T1 recovery.

The nuclei can return to the ground state only by dissipating their excess energy to

their surroundings (the “lattice”, which is why this kind of relaxation is also called spin-

lattice relaxation). The time constant for this recovery is T1 and is dependent on the

strength of the external magnetic field, B0, and the internal motion of the molecules

(Brownian motion).

T2/T2*: Transverse Relaxation

To understand transverse relaxation, it is first necessary to know what is meant by

“phase”. As used here, phase refers to the position of a magnetic moment on its circular

precessional path and is expressed as an angle. Consider two spins, A and B, precessing

at the same speed in the xy-plane. If B is ahead of A in its angular motion by 10o, then we

can say that B has a phase of +10 relative to A. conversely, a spin C that is behind A by

30o has a phase of -30o.

Immediately after excitation, part of the spins precess synchronously. These spins

have a phase of 0o and are said to be in phase. This state is called phase coherence. Later

spins lose coherence due to 1) energy transfer between spins as a result of local changes

34  

  

in the magnetic field; 2) time-independent inhomogeneities of the external magnetic field

B0. Thus, the individual magnetization vectors begin to cancer each other out. The

resulting vector sum, the transverse magnetization, becomes smaller and smaller and

finally disappear, and with it the MR signal (Figure 1-20f).

SPM NPs as MR Contrast Agent

Upon accumulation in tissues, SPM NPs are magneticlly saturated in the normal range of

magnetic field strengths in MRI scanner and establish a substantial locally perturbing

dipolar field, which leads to a marked shortening of T2* along with a less marked

reduction of T1. Thus SPM NPs are a good candidate for T2 contrast agent to provide a

dark image and the contrast enhancement is proportional to the magnetization magnitude

(Figure 1-21).56,57

Figure 1-21. Relationship between, T2 contrast and SPM NPs. When there is no SPM NPs, the dephasing time, T2 is long. While T2 becomes shorten in the presence of SPM NPs, the image of tissue becomes dark due to the quick loss of signal. Reproduced with permission from Reference 56

As the magnetization value of a SPM NP at a certain magnetic field is dependent

on the size and magnetocrystalline anisotropy of the NP, SPM NPs with different

sizes and structures have been prepared and compared for their contrast effects in

MRI.50 In a recent comprehensive study on ferrite NPs for MRI application,25 the NPs

were fabricated by a high-temperature, nonhydrolytic reaction between divalent metal

chloride (MCl2) and iron tris-2,4-pentadionate in the presence of oleic acid and

35  

  

oleylamine as surfactants. The hydrophobic ligand was exchanged with 2,3-

dimercaptosuccinic acid (DMSA). The DMSA-coated NPs show high colloidal

stability at a salt (NaCl) concentration of 250 mM, across a wide pH range (pH 6–10)

and in serum. 12-nm MnFe2O4 NPs have the highest mass magnetization value of 110

(emu/mass of magnetic atoms). This value is reduced to 101, 99 and 85 (emu/mass of

magnetic atoms) for Fe3O4, CoFe2O4 and NiFe2O4, respectively. The spin-spin

relaxation time (T2)-weighted MR images for each sample at 1.5 tesla (T) are

consistent with the magnetization results with MnFe2O4 NPs show the strongest MR

contrast effect with a relaxivity value reaching 358 mM-1•s-1, much larger than 218,

172, 152 and 62 mM-1•s-1 for Fe3O4, CoFe2O4, NiFe2O4 NPs respectively. This size

and structure dependent relaxivity of MFe2O4 NPs can be seen in Figure 1-22, from

which one can conclude that larger NPs have large contrast effect, but at the same

size, MnFe2O4 NPs have the largest contrast enhancement due to the small

magnetocrystalline anisotropy and easy magnetization reversal in MnFe2O4 structure.

36  

  

 

Figure 1-22. Size-dependent MR contrast effect of MnFe2O4 and Fe3O4 NPs. (a) TEM images of MnFe2O4 NPs (scale bar, 50 nm), (b) T2-weighted MR images, (c) color maps of 6-, 9- and 12-nm MnFe2O4 NPs, and (d) a plot of NP size versus R2 relaxivity. Reproduced with the permission from reference 25

The cancer detection sensitivity of these ferrite NPs are further evaluated. In a recent

test, the MnFe2O4 NPs were coupled with the cancer-targeting Herceptin, an antibody

specifically binding to the HER2/neu marker over-expressed on the surface of breast and

ovarian cancers.58 Various cell lines with different levels of HER2/neu over-expression:

Bx-PC-3, MDA-MB-231, MCF-7 and NIH3T6.7 (relative HER2/neu expression levels

are 1, 3, 28 and 2,300, respectively) were used for the test. With the MnFe2O4-Herceptin

conjugates, the detection of the Bx-PC-3 cell line occurred with a noticeable MR contrast.

As the relative HER2/neu expression level increased to 3, 28 and 2,300, the MR contrast

increased consistently for the MDA-MB-231, MCF-7 and NIH3T6.7 cell lines,

respectively. In contrast, when Fe3O4-Herceptin conjugates were used, the only MR-

detectable cell line was NIH3T6.7. Figure 1-23 shows the color coded MRI of a mouse

37  

  

implanted with the cancer cell line NIH3T6.7 and treated with 50 mg of NP-Herceptin. It

can be seen that the tumor treated with the MnFe2O4-Herceptin NPs show color changes

from red to blue in the color-coded MR images (Figure 1-23a–c). In contrast, those

treated with the Fe3O4-Herceptin NPs at the same dosage have no apparent change in the

color-coded MR images (Figure 1-23d–f). These indicate that the high MR sensitivity of

MnFe2O4-Herceptin conjugates enables the MR detection of tumors.25

 

Figure 1-23. Color maps of T2-weighted MR images of a mouse implanted with the cancer cell line NIH3T6.7 at different time points after injection of MnFe2O4-Herceptin (a-c) and Fe3O4-Herceptin (d-e) conjugates (preinjection (a,d); and 1 h (b,e) or 2 h (c,f) after injection). Reproduced with permission from reference 25.

 

7.2 SPM NPs as Drug Delivery Platform for Cancer Therapy 

Cancer is a class of disease in which a group of cells display uncontrolled growth

(division beyond the normal limits), invasion (intrusion on and destruction of adjacent

tissues), and sometimes metastasis (spread to other locations in the body via lymph or

blood).59 These three malignant properties of cancers differentiate them from benign

tumors, which are self-limited, do not invade or metastasize.60 Most cancers form a tumor

but some, like leukemia, do not. Cancer is caused by both external factors (tobacco,

chemicals, radiation, and infectious organisms) and internal factors (inherited mutations,

hormones, immune conditions, mutations that occur from metabolism).61 These causal

38  

  

factors may act together or in sequence to initiate or promote carcinogenesis. The process

could take ten or more years between exposure to external factors and detectable cancer.

Current cancer treatments include surgical intervention, radiation and

chemotherapeutic drugs, which often kill healthy cells and cause toxicity to the patient. It

would therefore be desirable to develop chemotherapeutics that can either passively or

actively target cancerous cells.62 Passive targeting exploits the characteristic features of

tumor biology (leaky blood vessels and poor lymphatic drainage) that allow nanocarriers

to accumulate in the tumor by the enhanced permeability and retention (EPR) effect

(Figure 1-24).63

Figure 1-24. Schematic representation of different mechanisms by which nanocarriers can deliver drugs to tumours. Polymeric nanoparticles are shown as representative nanocarriers (circles). Passive tissue targeting is achieved by extravasation of nanoparticles through increased permeability of the tumor vasculature and ineffective lymphatic drainage (EPR effect). Active cellular targeting (inset) can be achieved by functionalizing the surface of nanoparticles with ligands that promote cell-specific recognition and binding. The nanoparticles can (i) release their contents in close proximity to the target cells; (ii) attach to the membrane of the cell and act as an extracellular sustained-release drug depot; or (iii) internalize into the cell. Reproduced with permission from reference62

39  

  

Although passive targeting approaches form the basis of current clinical therapy, they

suffer from several limitations. Ubiquitously targeting cells within a tumor is not always

feasible because some drugs cannot diffuse efficiently and the random nature of the

approach makes it difficult to control the process. The lack of control may induce

multiple-drug resistance – a situation where chemotherapy treatment fails owing to

resistance of cancer cells towards one or more drugs. The reason is that transporter

proteins that expel drugs from cells are overexpressed on the surface of cancer cells.64,65

The passive strategy is further limited because certain tumors do not exhibit the EPR

effect, and the permeability of vessels may not be the same throughout a single tumor.66

One way to overcome these limitations is to program the nanocarriers so they actively

bind to specific cells. The binding may be achieved by attaching targeting agents such as

ligands – molecules that bind to specific receptors on the cell surface – to the surface of

nanocarriers.67 Nanocarriers will recognize and bind to target cells through ligand-

receptor interactions, and bound carriers are internalized before the drug is released

inside the cell (Figure 1-24).62 The targeting agents can be broadly classified as proteins

(antibodies), nucleic acids (aptamers), or other receptor ligands (peptides, vitamins, and

carbohydrates).68

Back to nanocarriers for drug delivery, the current clinical delivery agents are based

on natural and synthetic polymers and lipids (e.g. dextran, poly(ethylene glycol)).69 But

there have lots of explorations about the possible application of dendrimers, carbon

nanotubes, gold nanovehicle (nanoparticle, nanoshells and nanocages) and magnetic

nanoparticles.64 The rationale for magnetic nanoparticle-based targeting and drug

delivery lies in the potential to reduce or eliminate the side effects of chemotherapy drugs

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by reducing their systemic distribution as well as the possibility of administering lower

but more accurately targeted doses of the cytotoxic compounds.

The idea of using magnetic particles to act as therapeutic drug carriers in order to

target specific sites in the body dates back to the late 1970s.70 The drug/carrier complex is

then injected into the subject either via intravenous or intraarterial injection. High-

gradient, external magnetic fields generated by rare earth permanent magnets are used to

guide and concentrate the drugs at tumor location (Figure 1-25). Once the magnetic

carrier is concentrated at the tumor or other target in vivo, the therapeutic agent is then

released from the magnetic carrier, either via enzymatic activity or through changes in

physiological conditions such as pH, osmolality, or temperature, leading to increased

uptake of the drug by the tumor cells at the target sites. In theory, magnetic targeting

offers some major advantages for drug delivery, in particular, the ability to target a

specific site, such as a tumor, in vivo thereby reducing the systemic distribution of

cytotoxic compounds, and enhancing uptake at the target site resulting in effective

treatment at lower doses.71

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Figure 1-25. A hypothetical magnetic drug delivery system shown in cross-section: a magnet is placed outside the body in order that its magnetic field gradient might capture magnetic carriers flowing in the circulatory system. Reproduced with permission from reference37.

In general, superparamagnetic nanoparticles have the following distinct advantages

over the other delivery systems: (1) the pathway of the drug can be readily tracked in the

biological systems through SPM NPs by MRI (more penetrative than optical based

detection methods); (2) the drug-NPs can be guided or held in place by an external

magnetic field; and (3) under an alternate magnetic field, the SPM NPs act as a heater

and can trigger controlled drug release. In the delivery study, therapeutic drugs are

normally coupled to SPM NPs via a covalent bond. Hydrophobic drugs can also be

adsorbed onto NP surface to be stored in the NP coating layer to preserve their activity.72

Ideally, the drug-NPs are introduced in the biological systems and concentrated in the

targeted area by an active targeting as described in Figure 1-24. Drug release can proceed

42  

  

by simple diffusion or through enzymatic activity or the changes in physiological

conditions such as pH or temperature.73

Here I present two examples on drug release from Fe3O4 NPs. Methotrexate (MTX), a

chemotherapeutic drug that can target many cancer cells whose surfaces are

overexpressed by folate receptors, can be conjugated with Fe3O4 NPs through an amide

bond, as shown in Figure 1-26.74,75 In the conjugation process, the NPs are first modified

with (3-aminopropyl)-trimethoxysilane and subsequently conjugated with MTX through

amidation between the carboxylic acid end groups on MTX and the amine groups on the

particle surface. Drug release experiments show that MTX can be cleaved from the NPs

under low pH conditions mimicking the intracellular conditions in the lysosome. Cellular

viability studies in human breast cancer cells (MCF-7) and human cervical cancer cells

(HeLa) further demonstrate that such chemical cleavage of MTX is very effective inside

the target cells through the action of intracellular enzymes. The intracellular trafficking of

NP-MTX can be monitored through MRI. The results show that the MTX-Fe3O4 NPs

target cells with folate receptors and inhibit their growth.

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Figure 1-26. Surface modification of Fe3O4 NPs with MTX. Reproduced with permission from reference75.

Doxorubicin is a representative anthracycline antibiotic and one of the most widely

used anticancer drugs.76 In the treatment of gliomas, very high doses of doxorubicin must

be administered systemically to exert any therapeutic benefit and these doses are highly

neurotoxic and therefore ineffective in treating central nervous system maliganancies.77

The limited efficacy when administered systemically can be explained by the poor

solubility in aqueous solution and poor penetration of the drug through the blood-brain

barrier. Thus an efficient drug delivery system is required for administration of

doxorubicin against malignant gliomas or even metastatic brain tumors.78 The efforts to

44  

  

minimize side effect and increase the administration have resulted in the developments of

various drug delivery systems, including micro encapsulation of drug79, conjugation of

drug with polymer80, and physically loading drug in hydrogel81. Labhasetwar et al72

developed a water-dispersible oleic acid (OA)-Pluronic-coated iron oxide magnetic

nanoparticle formulation for doxorubicin delivery.

In this case, drug partitions into the OA shell surrounding iron oxide nanoparticles,

and the Pluronic that anchors at the OA-water interface confers aqueous dispersity to the

formulation (Figure 1-27a). Neither the formulation components nor the drug loading

affected the magnetic properties of the core iron oxide nanoparticles. Doxorubicin

loading in formulation was around 10 wt% with an encapsulation efficiency of 82%. The

release of doxorubicin from nanoparticles was sustained, with about 28% cumulative

drug release occurring in 2 days and about 62% over 1 week (Figure 1-27b). The drug-

loaded nanoparticles had a dose-dependent cytotoxic effect, which was slightly lower

than that observed with equivalent doses of free doxorubicin. This is due to the sustained

drug-release property of nanoparticles.

Figure 1-27. (a) Schematic representing formulation of iron oxide nanoparticles and the process for drug loading. (b) Release of doxorubicin in vitro from drug-loaded OA-Pluronic-stabilized iron oxide nanoparticles.

45  

  

7.3 SPM NPs as Mediators for Magnetic Hyperthermia 

Instead of being a carrier and killing tumor cells by the loaded drugs, the SPM NPs can

also serve as colloidal mediators and help induce heat to the local tumors to make

damage, which is called hyperthermia. It is based on the theory that single-domain iron

oxide nanoparticles possess a global magnetic moment which undergoes orientational

thermal fluctuations due to either Brownian Fluctuations of the grain itself within the

carrier fluid or internal fluctuations of the magnetic moment with respect to the crystal

lattice (Néel Fluctuations). These fluctuations are responsible for the magnetization

relaxation that occurs in a suspension of SPM NPs when the magnetic field is removed.

An external AC magnetic field supplies energy that excites the magnetic moment

fluctuations, and this magnetic energy is converted into thermal energy. Hence,

nanomagnetis may serve as nanosources of heat within hybrid nanostructures such as

cells.

Hyperthermia also takes advantage of the fact that, tumor cells are more susceptible

to elevated temperatures in the range of 42-45oC than the normal cells, making it possible

to deliver magnetic materials specifically to tumor cells, and generate heat locally to

damage them, without influencing the normal tissues.

One of the most crucial parameters of such mediator is its specific absorption rate

(SAR), which indicates the heat evolution rate in hyperthermia. SAR values depend on a

large number of parameters (e.g. size, size distribution, shape, surface chemical

compositions, frequency and amplitude of the magnetic field viscosity of the surrounding

medium) and vary from a few tenths to a few hundreds of Watt per gram of magnetic

46  

  

materials. Nevertheless, SPM NPs appear to be the best compromise choice between

biocompatibility and adequate SAR values, and have thus been intensively studied.

A typical experiment setup is shown in Figure 1-28, in which colloidal γ-Fe2O3

maghemite NPs solution was inserted in a copper coil (diameter 16mm).82 And the coil

produced an alternating magnetic field in the frequency range 300-1.1 MHz and with an

amplitude of up to 27 kA/m. the nonane was used to obtain an equilibrium temperature of

37±0.5 oC. And temperature was probed with a fluorooptic fiber thermometer. It is

obviously that the presence of SPM NPs results the increasement of temperature.

Figure 1-28. Device for magnetically induced hyperthermia. The ferrofluid sample (Vs = 300 μL) is introduced into a copper coil, which is part of a resonant circuit producing an AC magnetic field in the frequency range 300-1.1 MHz and with amplitudes up to 27 kA/m. The coil was cooled by circulating nonane. Temperature was probed with a fluorooptic fiber thermometer placed in the center of the sample. Example of temperature growth in the ferrofluid (maghemite sample). The specific loss power (SLP) was deduced from the initial linear rise in temperature (plain line) versus time, dT/dt, normalized to the mass of magnetic material and the heat capacity of the sample.

47  

  

8. Summary and Conclusion

Recent research progress have indicated that monodisperse SPM NPs can be readily

synthesized and functionalized for biomedical applications. First, the monodipserse

NPs have been made with controlled magnetic properties and chemical stability.

Second, the initial research shows that the SPM NPs can be made stable in biological

environment to escape from RES. Third, various targeting agents, especially tumor

specific antibodies, have been coupled to SPM NPs for testing their specific targeting

in biological environments. Fourth, SPM NPs functionalized with targeting agents are

excellent contrast agent for MRI and the contrast effects can be optimized by

controlling NP size, stucture and coating thinkenss. Fifth, therapeutic drugs can be

coupled to the NP surface with enhanced stablity and solubility in biological systems

and can be released in a controlled manner. All these experimental results reveal that

it is now possible to explore fully the great potential of monodisperse SPM NPs for

early medical diagnostic and therapeutic applications. With effective targeting agent

and highly sensitive NP probes, one is able to study in detail site-specific targeting,

cell uptake and NP-drug pathway within biological systems. With the uniform control

in NP physical and chemical properties, one can also study in a more precise way the

biodistribution and bioelimination of the drug-NP conjugates in biological

circulations. These understandings will finally allow the therapeutic and toxic effects

of various drugs to be carefully evalulated. In a word, SPM NPs are ideal platforms

for future success in diagnostic medicine and therapy.

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Chapter II

Synthesis and Surface Modification of Magnetite (Fe3O4)

Nanoparticle

1. Background

Magnetite is a ferromagnetic mineral form of iron(II, III) oxide with the chemical

formula Fe3O4, one of several iron oxides and a member of the spinel group (MgAl2O4).

Magnetite is a common iron oxide mineral, named for an ancient region of Greece where

metal production was prominent. It is the only mineral that exhibits strong magnetism. A

chunk of crystallized magnetite is called a lodestone, which was the earliest form of the

sailor’s compass. A lodestone was mounted to a rod on cork and floated in a bowl of

water. When the rod aligns with the Earth’s magnetic field, it points roughly north-south,

which provides a useful but rather limited way of positioning.

Magnetite has the spinel structure, with a cubic close packed oxygen array, and iron

in both fourfold (tetrahedral sites) and sixfold (octrahedral sites) coordination (Figure 2-

1a). The tetrahedral and octahedral sites form the two magnetic sublattices, A and B,

respectively. The spins on the A sublattice are antiparallel to those on the B sublattice,

which is defined as ferrimagnetism. The two crystal sites are very different and result in

complex forms of exchange interactions of the iron ions between and within the two

types of sites.1

Bulk magnetite (Fe3O4) is famous for a high Curie temperature of 850 K and nearly

full spin polarization at room temperature. However, this is not the reason why magnetite

NPs become the focus in biomedical applications. In recent studies the magnetic

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properties of magnetite NPs of size between 5 and 150 nm has been investigated closely

(Figure 2-1b). A gradual evolution from bulk-like magnetite to single-domain behavior

has been observed with decreasing grain size.2(Figure 2-1b) More importantly, the

favorable biocompatibility and biodegradability of these NPs have contributed greatly to

their widespread use in biomedical applications. Upton metabolism, iron ions are added

to the body’s iron stores and eventually incorporated by erythrocytes as hemoglobin

allowing for their safe use in vivo.3

Figure 2-1. (a) Crystal structure of magnetite. Blue atoms are tetrahedrally coordinated Fe2+; red atoms are octahedrally coordinated, 50/50 Fe2+/Fe3+; white atoms are oxygen; (b) Magnetization hysteresis curves measured at 296 K for the Fe3O4 samples. Reproduced with permission from reference 2

2. Synthesis and modification

The most widely used synthesis routes for magnetite NPs are based on precipitation from

solution. In these processes, a nucleation phase is followed by a growth phase, affording

a fairly good control over the NPs size and polydispersity. Typically, magnetite is

precipitated from basic aqueous solution of ferric and ferrous salts (Figure 2-2a). While

some control over size and composition of the NPs can be achieved through changing the

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nature and ratio of ferric/ferrous salts as well as by controlling the reaction conditions

(e.g. pH, temperature), coprecipitation processes usually results in polydisperse

nanoparticle suspensions due to significant aggregation (Figure 2-2b).

Figure 2-2. (a) Scheme showing the reaction mechanism of magnetite NPs formation from an aqueous mixture of ferrous and ferric chloride by addition of a base. (b) A typical images of as-synthesized magnetite NPs.

Thermal decomposition processes have been recently developed to produce high

quality monodisperse and monocrystalline magnetite NPs. In these procedures, iron

precursors are decomposed in hot organic solvents in presence of stabilizing surfactants

such as oleic acid and oleylamine. Iron precursors include iron acetylacetonate, iron

cupferronates, and iron carbonyls. Considering the safety (Fe(CO)5 is toxic) and price

factors in the synthesis, I usually made magnetite (Fe3O4) NPs with iron acetylacetonate

under the protection of oleyamine and oleic acid4,5. The size of Fe3O4 NPs could be

controlled through the ratio between iron precursor and surfactant. Figure 2-3 is the

typical images of synthesized magnetite (Fe3O4) NPs.

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Figure 2-3. Typical images of Fe3O4 NPs synthesized with Fe(acac)3 as precursor (a) 4-6nm; (b) 6-8nm; (c) 12-14nm; (d) 20-25nm. (Scale bar: 20nm) (Unpublished results)

Surface coating is of great importance in determining the NPs’ stability under

physiology condition. Due to the strong magnetic dipole-dipole interaction among them,

the iron oxide nanoparticles (IONPs) tend to agglomerate if without a hydrophilic coating

layer. For those IONPs made by co-precipitation in water using hydrophilic polymer (like

dextran, dendrimer, PASP) as the capping agents, this might be a minor issue. But for

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those made from high temperature decomposition in organic solvent, a surface

modification step is necessary to render the particles water soluble, biocompatible and

functionalizable6. Various methods have been developed, which can be roughly divided

into three categories, i.e. 1) ligand exchange; 2) ligand addition, and 3) inorganic coating.

In this chapter I will discuss and present some of the effort I’ve made on developing

novel surface modification methods through ligand exchange and ligand addition (Figure

2-4).

Figure 2-4. Illustration of surface modification through A) ligand exchange, and B) ligand addition.

2. 1. Ligand addition with phospholipid or oleylamine modified poly(acrylic acid)

Ligand addition refers to the process of adding a new ligand which can be anchored onto

the NP surface and make the particle water soluble while keeping the original coating

intact. In this case, the newly added ligand needs to be amphiphilic, with one end being

hydrophobic and interacting with the inner hydrophobic NP cores, while sticking the

hydrophilic tail into aqueous solution to offer the NPs hydrophilicity and stability.7 One

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widely used ligand is the group of phospholipids, in most cases PEGlyated, which have

on one side two hydrocarbon chains, and on the other size, a long hydrophilic PEG chain,

terminated with various functional groups.8

Figure 2-5. Chemical structure of carboxylic phospholipid, DSPE-PEG(2000)carboxylic acid from Avanti Polar Lipids, Inc.

For example, we’ve applied carboxylic phospholipids to make our Fe3O4 NPs water

soluble (Figure 2-5). The ligand is composed of a hydrophobic hydrocarbon “tail” and a

hydrophilic PEG “head”, and has a carboxylic acid group on the end of the PEG “head”.

Once mixed with the hydrophobic NPs, the phospholipids assemble onto the NP surface

to form a double layer structure with the original surfactant molecules through

hydrophobic-hydrophobic interactions.9-11 With the PEG “head” pitching outward, the

NPs can be rendered water soluble and the functional “head” is suitable for bio-

conjugation. DLS measurements (Figure 2-6) of the dispersed 8 nm Fe3O4 NPs show that

before surface modification, the nanoparticles have an average hydrodynamic diameter of

11 nm that is close to a simple addition of the particle core diameter (8 nm) and the shell

coating (~3 nm). After phospholipid modification in water, the overall organic shell

coating is increased to ~30 nm. This, plus an 8 nm core, gives an overall diameter of 38

nm, while the hydrodynamic diameter of the structure from DLS measurement is at 39.6

nm (Figure 2-6).

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Figure 2-6. The hydrodynamic diameter distribution of the Fe3O4 NPs in the dispersion measured by DLS: the green line is the as-synthesized 8 nm Fe3O4 NPs in hexane (11nm). The red one is the particles in water after surface modification with phospholipid carboxylic acid illustrated in Figure 2-5 (39.6nm).

This method is easy to carry out, but these funtionalized and PEGylated

phospholipids are not commercially available ($490 for 100 mg for carboxylic acid

phospholipids, http://www.avantilipids.com/). Therefore, it is inevitable to develop a

similar but cheaper ligand for surface modification.

Several polymers have previously been reported for the surface modification of

semiconductor nanoparticles, including octylamine-modified poly(acrylic acid), block

copolymers, and amphiphilic polyanhydrides.9,12-14 Considering the surfactants on Fe3O4

NPs surface, oleylamine modified poly(acrylic acid) (MW=5,000) was tested for Fe3O4

NPs modification.

The modification of poly(acrylic acid) with oleylamine is following Zhou et al.15

Basically, 2.16 g of poly(acrylic acid) solid (0.03 mol of carboxylic acid, MW=5,000)

and 2.85 g of 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDAC,

0.015 mol) were transferred into a 100 mL round-bottom flask. 20 mL of DMF was

added. About 1.6 mL of oleylamine (0.0096mol) was added dropwise into the reaction

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flask. The clear solution was stirred overnight under N2. TLC (CH2Cl2/CH3OH=7:1,

Rf(octylamine)=0.55, Rf(product)=0.24) was used to monitor the reaction. When the reaction was

complete, DMF was removed by vacuum, and the residue was mixed with 10 mL of

acetone and transferred into a centrifuge tube. 25 mL of water was added, and the gummy

precipitated product was separated by centrifugation (3000 rpm for 5 min) and washed

with water (25 mL x 3). The solid product was dissolved in 40 mL of ethyl acetate (gentle

heating 40-50 °C was applied), and a tetramethylammonium hydroxide (6.4 g in 25 mL

water) solution was added to the polymer solution and stirred for 10 min, before being

transferred into a separation funnel. The yellowish aqueous layer was isolated in a

centrifugation tube and acidified by 1 N HCl to pH=2. The precipitate was separated by

centrifugation and washed with H2O (15 mL x 2). The sticky solid was redissolved in

ethanol, at which point the ethanol was removed under vacuum and 1.63 g of the yellow

solid product (OPA) was collected (around 45% of the carboxylic acids were converted

to octyl amide). 1H NMR (d-CH3OH): δ 0.85-0.92 (m, 3H), 1.20-1.35 (m, 10H), 1.40-

1.55 (m, 2H), 1.55-1.95 (m, 3.2H), 2.05-2.50 (m, 2.4H), 2.95-3.28 (m, 2H).

The synthesized polymer can have their hydrocarbon chains interact with the original

hydrophobic coating, while sticking the charged amine or carboxylate groups outside to

help stabilize the particles (Figure 2-7a). In detail, ~ 5 mg NPs are dispersed in

chloroform after precipitated out from Hexane solution with excessive ethanol. 50 mg

polymer (oleylamine modified poly(acrylic acid), or OPA) was dispersed in 4 mL of

chloroform. Teteramethylammonium hydroxide (25 wt% in methanol) was used to adjust

pH to 10, to which NPs solution in chloroform was added. The resulted solution was

mixed well by vortex and the solvent was evaporated by rotavapor, yielding black, wax-

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like solid. Water was subsequently added to redisperse the particles. The solution could

be dialysis to remove the base or other salt. The polymer could render different size of

Fe3O4 NPs water soluble (Figure 2-7b).

Figure 2-7. (a) Illustration of Fe3O4 NPs modified with oleylamine modified poly(acrylic acid). Adapt from reference16. (b) Picture of OPA modified 5-6nm, 12nm and 25nm Fe3O4 NPs in water.

The DLS analysis was conducted to test the 8nm Fe3O4 NPs’ overall size before and

after the modifications (Figure 2-8). One can see that, before adding surfactant in hexanes

solution, the particles are with an overall diameter around 11 nm, close to an estimation

of a 8 nm core, plus a 2~3 nm coating (i.e. one layer of oleic acid/oleylamine) (Figure 2-

6,8). After modification in aqueous solution, the overall size just goes up to around 13.6

nm, which is pretty similar to the as-synthesized one (Figure 2-8). Compared with

phospholipid modification, OPA definitely gives smaller hydrodynamic diameter. A

detailed examination about the stability and biocompatibility of the polymer modified

NPs is undergoing.

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Figure 2-8. The hydrodynamic diameter distribution of the Fe3O4 NPs in the dispersion measured by DLS: the green line is the as-synthesized 8 nm Fe3O4 NPs in hexane (11nm). The red one is the particles in water after surface modification with oleylamine modified poly(acrylic acid), or OPA (13.6nm).

2. 2. Ligand exchange with dopamine modified poly(ethylene glycol) with TsT as a linker

Ligand exchange refers to the approach of replacing the original hydrophobic surfactants

(usually alkylamine or alkylacid), with new, hydrophilic ones. This happens when the

new ligands have higher affinity to the particle surface than the original capping ligands.

One such example is the dopamine based ligands. Studies have shown that, the bidentate

enediol ligands such as dopamine could convert the under-coordinated Fe surface sites

back to a bulk-like lattice structure with an octahedral geometry for oxygen-coordinated

iron, which result in strong binding between dopamine moiety and the surface of the iron

oxide nanoparticles (Figure 2-9a).17-19 Therefore, the dopamine derivatives can help

anchor the ligand onto Fe3O4 NP surface.

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Figure 2-9. (a) Structure of dopamine (DA) and its proposed binding configurations with surface Fe sites on Fe3O4 NPs; (Adapted and reprinted with permission from reference17) (b) Synthesis of Dopamine-TsT-PEG and their replacing of oleic acid/oleylamine coating on Fe3O4 NPs (Adapted and reprinted with permission from reference20).

From the previous study we know that, only one charged group is not sufficient to

protect the particles from agglomeration in aqueous solution. So once again, we chose to

use PEG as the spacer to bring in steric repulsion forces so as to stabilize the particles.

Common PEG is terminated with hydroxyl group. In order to couple it with dopamine to

obtain anchorable ligand, we chose cyanuric chloride, also known as trichloro-s-triazine

(TsT) as the coupling agent, which is effective in activating OH group and is much less

inexpensive than other common coupling agents, like BS3 or sulfo-SMCC. TsT is a

symmetrical heterocyclic compound containing three acyl-like chlorines, as shown in

Figure 2-9b. In aqueous solution, these three chlorines show different reactivities toward

nucleophiles. For example, at pH=9, the first chlorine is reactive toward hydroxyls as

well as alkylamine groups at 4 ˚C. After this first chlorine is coupled to the nucleophile,

the second one requires at least room-temperature to do so. The third one is even more

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difficult to react, requiring at least 80 ˚C.21 Such reactivity feature of TsT allows it to be

sequentially labeled or coupled with different dyes or proteins.

Figure 2-10. 1H NMR of (a) Dopamine in D2O; (b) mPEG in CDCl3; (c) compound b in CDCl3 and (d) coumpound c in CDCl3.

The monodisperse Fe3O4 NPs (9 nm in diameter) used in this study were prepared by

one-pot high temperature (300 oC) reductive decomposition of Fe(acac)3 (2 mmol) in

oleylamine (10 mL) and benzyl ether (10 mL). And in order to couple dopamine with

mPEG by TsT, mPEG2000 (a) was first activated with TsT at room temperature in

anhydrous benzene (20 mL) to give b, as shown in Figure 2-9b. b further reacts with

dopamine, or 4-(2-aminoethyl)benzene-1,2-diol (adducted with hydrogen chloride) in

1,4-dioxane, forming compound c. The molecular structures of both b and c were

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confirmed by 1H NMR spectroscopy in CDCl3 (Figure 2-10). The catecol unit in

dopamine based molecule c is used to replace oleate/oleylamine around the Fe3O4 NPs,

which take place by mixing c with Fe3O4 NPs in CHCl3 overnight. Such replacement

gives mPEG coated Fe3O4 NPs (d). It can be seen from the structure of d that TsT acts as

a bridge to link mPEG and dopamine-Fe3O4 NPs. After evaporating out the solvent, the

PEG-dopamine-coated nanoparticles (d) are readily dispersed in water, PBS buffer

(pH=7.4, 137 mM NaCl, 2.7 mM KCl, and 10 mM Na2HPO4/KH2PO4) or borate buffer

(pH=8.5, 10 mM H3BO3).

DLS measurements on the dispersed 9 nm Fe3O4 NPs show that before surface

modification, the nanoparticles have an average hydrodynamic diameter of 11.9 nm

(Figure 2-11a). This is close to a simple addition of the particle core diameter (9 nm) and

the shell coating (~2-3 nm). After ligand exchange, the organic shell coating in d is

increased to about 30 nm (2×15 nm). This, plus a 9 nm core, gives an overall diameter of

39 nm, while the average hydrodynamic diameter measured from DLS is 40.3 nm (Figure

2-11b), a size that is close to the addition of mPEG-dopamine coating + radius of

nanoparticle.

Figure 2-11. Hydrodynamic diameter distribution of (a) 9 nm Fe3O4 NPs dispersion in hexane and (b) nanoparticles d (Figure 2-9b) in PBS measured by DLS. (Adapted and reprinted with permission from reference20).

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The TEM image of the NPs from hexane solution is shown in Figure 2-12a, and those

from aqueous solution (PBS) is given in Figure 2-12b. Comparing the two TEM images,

one can see that the Fe3O4 NPs are well dispersed in both hexane and PBS. However, the

morphology of the nanoparticles after dopamine replacement does change from

sphere/polyhedron-like to cube-like. This indicates slight Fe3O4 surface corrosion during

the exchange, which is different from the previous observation of particles decorated with

an additional layer, where particles remain unaltered after modification.

Figure 2-12. TEM images of (a) 9 nm Fe3O4 NPs coated with oleate/oleylamine from their hexane dispersion and (b) nanoparticles d in Figure 2-10b from PBS dispersion. (Adapted and reprinted with permission from reference20).

Stability of the mPEG-dopamine coated Fe3O4 NPs was tested in borate buffer (10

mM) at various pH values with an incubation temperature of 70 ˚C. DLS was used to

track the size change of the nanoparticles in these pH conditions during the incubation.

Figure 2-13 shows the variation of hydrodynamic diameters of the nanoparticles in borate

buffer incubated at 70 ˚C. It can be seen that the initial average size of the nanoparticles

is at ~40 nm. In the solutions with pH=7 or above, there is no size increase during the test

(24 h), indicating no particle aggregation/sintering. The TEM image of the nanoparticles

after incubation is similar to what is shown in Figure 2-12b (data not shown). On the

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other hand, at lower pH, e.g. pH=6, the particles are found stable for only 2 h before

serious aggregation occurs. After 15 h, the size of the clustered nanoparticles reaches 100

nm. Such sintering of the dopamine coated particles may result from the chemical bond

cleavage between iron oxide and the catecol unit under low pH, thus destabilizing the

nanoparticle dispersion. In neutral or basic conditions (tested from 7.0 to 8.5), the

nanoparticles are well stabilized. Their stability test in PBS showed similar result as that

in borate buffer.

Figure 2-13. DLS measured average hydrodynamic diameters of the nanoparticles d (Figure 2-9b) in borate buffer at different pH values after incubation at 70 ˚C. (Reprinted with permission from reference20).

This successful trial told us some important information in making Fe3O4 NPs water

soluble through liagnd exchange approach. First, dopamine derivatives can efficiently

take replace of the original coating, even after it was conjugated with macromolecules.

Second, similar as the case of ligand addition, incorporation of PEG did help to stabilize

the particles in aqueous solutions, at least in neutral or basic environment. However, in

this first trial, we used mPEG instead of PEG to eliminate the cross-linking possibility.

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Therefore after surface modification, since the PEG is methyl terminated, we cannot do

further conjugation with other species through mild chemistry.

In summary, there are three elements that are essential if we want to convert the

particles water soluble and functional through ligand exchange. First of all, we need an

anchoring moiety such as dopamine, which has high affinity to the particle surface thus

being able to take replace of the original organic coating. Second, it is necessary to have a

hydrophilic spacer which can bring steric repulsion forces so as to enhance the particles’

stability against agglomeration, e.g. PEG. Last but not least, a functional group, like

carboxyl, amine, thiol, etc, is necessary to be at the terminal of the ligand, which can be

used for further conjugation with other bimolecules, such as protein, DNA, peptide, etc,

at mild condition (4˚C to r.t., pH from 5 to 9).

2. 3. Ligand exchange with dopamine modified bifunctional poly(ethylene glycol)

Considering further functionalization with bio-molecules, we chose to use PEG diacid

which would be coupled with dopamine (DPA) in a 1:1 ratio by EDC/NHS chemistry to

form the new ligand (Figure 2-14). As discussed above, the dopamine moiety would help

the ligand to anchor onto the particles surface; the long PEG in the middle would help

stabilize the particles and gift the particles hydrophilicity; while the remaining carboxyl

at the end of the new ligand could be used for further conjugation.

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Figure 2-14. Surface modification of Fe3O4 NPs via DPA-PEG-COOH. X=CH2NHCOCH2CH2 for PEG3000, PEG6000, PEG20000. X is absent in PEG600, for in such case, the atoms on both sides of the X are directly linked. (Reprinted with permission from reference22)

The synthesis of modification was easy and straightforward. Taking DPA-PEG3000-

COOH synthesis for example, PEG3000 diacid was reacted in CHCl3/DMF 1:1 solution

with one equivalent dopamine by using EDC and NHS as the catalysts at room

temperature. Afterwards, Fe3O4 NPs in CHCl3 were added in, and the resulted mixture

was magnetically stirred overnight to proceed ligand exchange. Subsequently, the

products were precipitated out by adding hexane, collected by magnetic bar, and dried

out under vacuum. Afterwards, they were ready to be dispersed in water or PBS. We also

tried other PEG with different lengths, which are PEG600 diacid, PEG6000 diacid and

PEG20000 diacid. Coressponding synthesis and subsequent modification can be achieved

in a similar way.

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TEM images of the PEGylated nanoparticles were obtained by evaporating water

from the dispersion on amorphous carbon coated copper grid. Figure 2-15 shows TEM

image of IONPs before and after modifying by four kinds of PEGylated ligands. One can

see that there is no obvious change in core size after modification with DPA-PEG-COOH,

and in all cases the particle are dispersed well after modification.

Figure 2-15. TEM images of the IONPs before (a) and after ligand exchange with (b) DPA-PEG600-COOH, (c)DPA-PEG3000-COOH, (d) DPA-PEG6000-COOH and E) DPA-PEG-COOH 20000. (Adapted and Reprinted with permission from reference22)

The PEG coating thickness around the nanoparticles was characterized with DLS and

the results are shown Figure 2-16. It can be seen that before modification, the

nanoparticles in hexane have an overall size around 11 nm. This is close to the simple

addition of the dimensions from the core (9 nm) and the shell (~3 nm - the length of the

oleate and oleylamine molecules). After modification, the sizes of the particles increase

to around 40, 50, 70 and 90 nm for PEG600, PEG3000, PEG6000 and PEG20000 coated

nanoparticles, respectively, indicating successful PEGylation. Zeta potentials of these

PEGylated particles in water show that all of them are negatively charged (Table 2.1),

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probably due to the multiple carboxylate groups on the particle surface. As a comparison,

dextran coated NPs are also negatively charged but slightly more neutral.

Figure 2-16. Hydrodynamic sizes of the Fe3O4 NPs coated with different surfactants. The sizes were measured from the aqueous solution of the nanoparticles by DLS. (Reprinted with permission from reference22)

Table 2.1. Zeta potentials PEGylated Fe3O4 NPs and dextran coated Fe3O4 NPs (in water)

The surface coating of the PEG Fe3O4 NPs was further characterized by thermal

gravitational analysis (TGA) and infrared (IR) spectroscopy. In the TGA analysis, the as-

synthesized nanoparticles show two peaks at around 230 and 410 °C (Figure 2-17a),

accounting for the mass loss due to the evaporation of oleic acid or oleylamine on the

nanoparticle surface. But in the analysis for the PEGylated NPs, these two peaks

disappear and are replaced by a strong desorption peak at about 360 °C. Comparing with

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the TGA results of the free DPA-PEG-COOH ligand (Figure 2-17b), one can see that this

mass loss is caused by the evaporation of the DPA-PEG-COOH ligand anchored on the

surface of the particles.

Figure 2-17. TGA analysis of (a) the Fe3O4 NPs before and after modification with DPA-PEG ligands; (b) the DPA-PEG-COOH ligands alone. (Reprinted with permission from reference22)

The successful ligand exchange was also proven by IR analysis (Figure 2-18). The as-

synthesized nanoparticles show two main absorption peaks in IR spectrum: one is at

~3500 cm–1, which is contributed by the COOH and NH2 groups from oleic acid and

oleylamine; and the other one is around 2800 cm-1, arising from the stretching vibration

of C-H. After modification, however, all four sets of nanoparticles exhibit a new peak at

~1100 cm–1, which is due to the characteristic stretching vibration of C-O-C from PEG.

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Figure 2-18. IR study of the Fe3O4 NPs before and after modification with DPA-PEG-COOH ligands. (Reprinted with permission from reference22)

Stability of the PEG coated Fe3O4 NPs was analyzed in water as well as PBS plus 10%

FBS at 37 °C by monitoring the size change with DLS. These four kinds of PEGylated

NPs were sampled at 0, 2, 4, 6, 8, 16 and 24 h during the incubation and the results are

shown in Figure 2-19. Comparing two sets of data, it can be seen that, the Fe3O4 NPs

incubated in the mimic physiology environment (Figure 2-19b) are ~10-20 nm larger

those incubated in water (Figure 2-19a). Such size increase is attributed to the interaction

of nanoparticles with the FBS in the incubation medium. However, with the existence of

the dense PEG coating, this interaction did not cause further agglomeration and four

kinds of the nanoparticles all show excellent stability without obvious size increase

during an incubation period of 24 hours. No precipitation was found after the incubation

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at the vessel bottom, and the TEM study results are similar to those before the

modification (data not shown), indicating that DPA-PEG indeed offers a robust coating

around the Fe3O4 NPs, making them sustain from the cell culture condition.

Figure 2-19. Size change monitoring of PEGylated Fe3O4 NPs by DLS in (a) water and (b) PBS+10%FBS at 37 ˚C for 24 h. (Reprinted with permission from reference22)

As potential probes for MRI, those PEGylated Fe3O4 NPs’ uptake by

reticuloendothelial system (RES) is of great concern. To study that, we incubated these

particles with the RAW 264.7 cells, which are one kind of mouse macrophage cell line, at

three starting concentrations: 0.1 mg Fe mL–1, 0.01 mg Fe mL–1 and 0.001 mg Fe mL–1,

and measured Fe concentrations within the cells after 4h incubation by inductively

coupled plasma-atomic emission spectrometry (ICP-AES) analysis. For comparison,

uptake of dextran-coated Fe3O4 NPs with the same starting Fe concentration was also

tested. The RAW 264.7 cells grown without Fe3O4 NPs were used as control. The results

are shown in Figure 2-20 a-c. It is obvious that, the uptake is concentration dependent for

all the particles, i.e. higher concentration lead to higher particle uptake, which is

consistent with the previous observations.23 For example, at concentration of 0.1 mg Fe

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mL–1, the uptakes of all kinds of particles are about 20 times larger than those at 0.01 mg

Fe mL-1 (Figure 2-20a). At the same starting concentration, the coating nature determines

the uptake efficiency, which relies on the particle size, coating materials, charge, etc. One

can see that, at any concentration, the dextran coated nanoparticles shows much higher

uptake than the peers, followed by PEG600 Fe3O4 NPs, while the uptakes of PEG3000,

PEG6000, PEG20000 Fe3O4 NPs are comparable. For example, with a starting

concentration of 0.01 mg Fe mL–1 (Figure 2-20b), it can be seen that the dextran-coated

nanoparticles give the highest uptake, more than 3 times that of PEG600 Fe3O4 NPs. For

PEG3000, PEG6000, and PEG20000 coated Fe3O4 NPs, their uptakes are comparative

with the control, indicating negligible uptake of these nanoparticles by the macrophage

cells. Such uptake difference is attributed to the coating efficacy. PEG with molecular

weight higher than 3000 give dense coating over the surface of the nanoparticles, thus the

length of PEG chain becomes insignificant factor in terms of non-specific uptake.

However, less dense PEG600 coating nanoparticles is not sufficient enough to protect the

particles from protein adsorption therefore causing higher uptake.

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Figure 2-20. (a)-(c) Macrophage cell uptake of Fe3O4 NPs at different starting concentrations: (a) starting concentration of 0.1 mg Fe/ml (b) starting concentration of 0.01 mg Fe/ml (c) starting concentration of 0.001 mg Fe/ml. After incubation, those Fe3O4 NPs-bearing cells were collected and dispersed in 1% agarose gel and subjected to MR imaging, which is shown in (d). Each dot in D represents one sample. (Reprinted with permission from reference22)

After uptake, those cells, bearing with the internalized particles, were dispersed in

1ml 1% agarose gel and subjected to MR imaging, and the result is shown in Figure 2-

20d. Each dot represents one sample. One can see that, for each kind of particles, when

the starting concentration goes higher, the sample become “darker” under MRI,

indicating more particle internalization, which correlates with the ICP result. However,

when comparing the results horizontally, we found that, cells incubated with PEG 600

NPs are the “darkest” at all concentrations instead of dextran coated ones. This is

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explained by the fact that, our Fe3O4 NPs, synthesized from high temperature

decomposing, are higher in magnetization therefore having better T2 signal decrease

capability, which is confirmed by a later extra cellular study on all five kinds of particles.

In such study, all the particles were dispersed in agarose gel at different concentrations

and were subjected to MR imaging (Figure 2-21). It clearly shows that, with the same

Fe3O4 core, all PEGylated NPs have similar T2 contrast capability. However, dextran NPs

show much less impressive signal reduction at all tested concentrations. Such inefficacy

as T2 contrast agents, explains why in Figure 2-20d, the signal reduction caused by

dextran Fe3O4 NPs uptake is less prominent than those from PEGylated Fe3O4 NPs, even

if more dextran Fe3O4 NPs were uptaken.

 

Figure 2-21. Ex cellular phantom study of Fe3O4 NPs’ T2 reducing effect.

3. Conclusion

In summary, we’ve successfully demonstrated that, the Fe3O4 NPs synthesized from high

temperature decomposing, can be modified and converted water soluble. Starting from

inexpensive materials, we’ve successfully synthesized homemade ligands, such as

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oleylamine modified poly(acrylic acid) and PEGylated dopamine, that can efficiently

anchor onto as-synthesized NPs through either ligand addition or ligand exchange. The

resulted NPs were unalterted in core size and maintained their superior magnetism.

Specifically, for DOP-PEG modified Fe3O4 NPs, DLS studies showed that, they have an

over diameter from 40 to 90 nm which is tunable by changing the PEG length. By

monitoring the size change in a 24 hour interval, we’ve proved that they are very stable in

either pure water or in a mimic physiology environment. And from an in vitro uptake

experiment in which they were incubated with the macrophage cells, we found that, these

PEGylated Fe3O4 NPs have much lower macrophage uptake compared with the dextran

coated ones. And later MRI study shows that, our PEGylated Fe3O4 NPs have better T2

signal reduction ability than dextran coated ones, which is attributed from their higher

magnetization. All of the features are very important for the potential in vivo application.

Conjugated with appropriate biomolecules through the terminal carboxyl groups, these

particles can become superior specific targeting bullets in either MRI or drug delivery.

4. Experimental

Materials and Instruments: Fe(acac)3, αω-bis{2-[(3-carboxy-1-oxopropyl)amino]ethyl}

polyethylene glycol (M = 3000, 6000 and 20000), polyethylene glycol diacid (M = 600),

dopamine hydrochloride, sodium carbonate and organic solvents used in the syntheses

were purchased from Sigma Aldrich. N-hydroxysuccinimide (NHS) and N,N’-

dicyclohexylcarbodiimide (DCC) were from Pierce Biotechnology. All the buffers and

media were from Invitrogen Corp. Water was purified by Millipore Milli-DI Water

Purification System. All the dialysis bags were purchased from Spectrum Laboratories,

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Inc. All the other chemicals were from Sigma Aldrich. All the PEGylated phospholipids

were from Avanti Lipids.

 

Synthesis of Fe3O4 nanoparticles: 2 mmol of Fe(acac)3 was dissolved in a mixture of 10

ml benzyl ether and 10 ml oleylamine. The solution was dehydrated at 110°C for 1 h, and

was quickly heated to 300°C and kept at this temperature for 2 hours. 50 ml of ethanol

was added into the solution after it was cooled down to room temperature. The

precipitation was collected by centrifuge at 8000 rpm and was washed by ethanol for 3

times. Finally, the product (150 mg) was redispersed in hexane.

Surface modification of Fe3O4 nanoparticles by phospholipids: The solvent hexane

was evaporated from the hexane dispersion of the particles under a flow of nitrogen gas,

giving black solid residue of iron oxide nanoparticles. The residue was dissolved in

chloroform to form the chloroform dispersion at a concentration of 0.5 mg particles/mL

solution. 1 mL of chloroform solution of DSPE-PEG(2000)Carboxylic acid (10 mg/mL)

was added into a 2 mL of the nanoparticle dispersion. The mixture was shaken for 1 h,

the chloroform solvent was evaporated under nitrogen gas. The solid residue was

dispersed in phosphate buffered saline (PBS) solution for further test. A small portion of

undispersed residue was filtered off by a 0.2-μm syringe filter. The free lipid was

removed by a Nanosep 100 k Omega.

Synthesis of mPEG-TsT (b): 22 mg of thricholoro-s-triazine (TsT) was dissolved in 20

ml anhydrous benzene which contained 1g of anhydrous sodium carbonate. Then, 200 mg

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of monomethoxypolyethylene glycol 2000 (mPEG 2000) was added into the solution,

mixed well and reacted overnight at room temperature. The TsT-modified mPEG (TsT-

mPEG) was precipitated out by adding 30 ml petroleum ether and collected through

centrifuge. The product was washed with benzene for three times and then dried under

vacuum overnight.

Synthesis of mPEG-TsT-Dopamine: 1.7 mg of dopamine hydrochloride was dissolved

in 2 ml 1,4-dioxane which contained ~10 mg of sodium carbonate. 20mg of the as-

synthesized TsT-mPEG dissolved in 2ml 1,4-dioxane was added into the above solution

dropwisely in 5 minutes with stirring at room temperature. After 3 hours of reaction, the

product, c, was precipitated with petroleum ether and removed from the solution by

centrifuge.

Modification of Fe3O4 nanoparticles with mPEG-TsT-Dopamine: The compound c

was redissolved in CHCl3 and mixed with 5 mg of Fe3O4 nanoparticles. The mixture was

stirred under N2 at room temperature overnight. The final mPEG-TsT-Dopamine coated

Fe3O4 nanoparticles (d) was precipitated and washed with hexane. After being dried

under vacuum, the particles were able to be dissolved in DI water or buffer (PBS or

borate buffer).

Surface Modification of Fe3O4 Nanoparticles with DPA-PEG-COOH: PEG diacid 20

mg (this amount is for PEG diacid 3000; for other PEG diacids, same moles were used),

NHS (2 mg), EDC (3 mg) and dopamine hydrochloride (1.27 mg) were dissolved in a

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mixture solvent containing CHCl3 (2 mL), DMF (1 mL), and anhydrous Na2CO3 (10 mg).

The solution was stirred at room temperature for 2 h before Fe3O4 nanoparticles (5 mg)

were added, and the resulting solution was stirred overnight at room temperature under

N2 protection. The modified Fe3O4 nanoparticles were precipitated by adding hexane,

collected by a permanent magnet and dried under N2. The particles were then dispersed in

water or PBS. The extra surfactants and other salts were removed by dialysis using a

dialysis bag (MWCO = 10000) for 24 h in 1× PBS or water. Any precipitation was

removed by a 200 nm syringe filter. The final concentration of the particles was

determined by ICP-AES analysis.

Dextran coated Fe3O4 nanoparticles: were synthesized by co-precipitation of FeCl2 and

FeCl3 in aqueous solution with the presence of ammonium hydroxide according to an

earlier publication24.

Cell Uptake Experiment: Raw 264.7 cell lines were cultured in RPMI 1640 media (with

Glutamine and Phenol Red) with 10% FBS and 1% antibiotics in T25 culture flasks.

Before the test, the growth medium was removed. The cells were washed twice with PBS

before Fe3O4 nanoparticles coated with different PEG’s in growth media, each with

different concentrations (0.1 mg FemL–1, 0.01 mg FemL–1, 0.001 FemL–1) were added.

Cells grown without any particles were used as control. The cells were then incubated for

4 h at 37 °C, 5% CO2, washed with PBS twice and redispersed in RPMI. The cell

concentrations were determined by hemacytometry and the Fe concentrations were

determined by ICP-AES.

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Characterization: 1H NMR was operated at 400 MHz on a Bruker Avance NMR

Spectrometer. Nanoparticle samples for transmission electron microscopy (TEM)

analysis were prepared by drying the dispersion of the particles on amorphous carbon

coated copper grids. Images were taken on Philips EM 420 (120 kV). The size of the

nanoparticles in dispersion was evaluated using a Malvern Zeta Sizer Nano S-90

Dynamic light scattering (DLS) instrument. The measurements were done at 25°C. The

following parameters were used for size estimation: Refractive Index 2.420 (Fe3O4),

1.373 (hexane), 1.33 (water); Viscosity 0.3000 (hexane), 0.8872 (water); Absorption

0.010 (Fe3O4). Quantitative elemental analyses of the nanoparticles were carried out with

electron diffraction spectrum (EDS). X-ray powder diffraction patterns of the particle

assemblies were collected on a Bruker AXS D8 Advance diffractometer under Cu Ka

radiation (λ = 1.5405). Magnetic properties of the particles were studied using a

Lakeshore 7404 high-sensitivity vibrating sample magnetometer (VSM) with fields up to

1 T at room temperature. UV–vis analysis was performed on a PerkinElmer Lambda 35

UV–Vis spectrometer.

 

References:

1. Majewski, P.; Thierry, B. Critical Reviews in Solid State and Materials Sciences 2007, 32, 203-215.

2. Goya, G. F.; Berquo, T. S.; Fonseca, F. C.; Morales, M. P. Journal of Applied Physics 2003, 94, 3520-

3528.

3. Weissleder, R.; Stark, D. D.; Engelstad, B. L.; Bacon, B. R.; Compton, C. C.; White, D. L.; Jacobs, P.;

Lewis, J. American Journal of Roentgenology 1989, 152, 167-173.

4. Sun, S. H.; Zeng, H. Journal of the American Chemical Society 2002, 124, 8204-8205.

5. Xu, Z.; Hou, Y.; Sun, S. Journal of the American Chemical Society 2007, 129, 8698-8699.

6. Sun, C.; Lee, J. S. H.; Zhang, M. Q. Advanced Drug Delivery Reviews 2008, 60, 1252-1265.

7. Xu, C. J.; Sun, S. H. Polymer International 2007, 56, 821-826.

8. Medintz, I. L.; Uyeda, H. T.; Goldman, E. R.; Mattoussi, H. Nature Material 2005, 4, 435-446.

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9. Dubertret, B.; Skourides, P.; Norris, D. J.; Noireaux, V.; Brivanlou, A. H.; Libchaber, A. Science 2002,

298, 1759-1762.

10. Michalet, X.; Pinaud, F. F.; Bentolila, L. A.; Tsay, J. M.; Doose, S.; Li, J. J.; Sundaresan, G.; Wu, A.

M.; Gambhir, S. S.; Weiss, S. Science 2005, 307, 538-544.

11. Hultman, K. L.; Raffo, A. J.; Grzenda, A. L.; Harris, P. E.; Brown, T. R.; O'Brien, S. Acs Nano 2008, 2,

477-484.

12. Wu, X. Y.; Liu, H. J.; Liu, J. Q.; Haley, K. N.; Treadway, J. A.; Larson, J. P.; Ge, N. F.; Peale, F.;

Bruchez, M. P. Nature Biotechnology 2003, 21, 41-46.

13. Gao, X. H.; Yang, L. L.; Petros, J. A.; Marshal, F. F.; Simons, J. W.; Nie, S. M. Current Opinion in

Biotechnology 2005, 16, 63-72.

14. Pellegrino, T.; Manna, L.; Kudera, S.; Liedl, T.; Koktysh, D.; Rogach, A. L.; Keller, S.; Radler, J.;

Natile, G.; Parak, W. J. Nano Letters 2004, 4, 703-707.

15. Zhou, M.; Nakatani, E.; Gronenberg, L. S.; Tokimoto, T.; Wirth, M. J.; Hruby, V. J.; Roberts, A.;

Lynch, R. M.; Ghosh, I. Bioconjugate Chemistry 2007, 18, 323-332.

16. Yang, J.; Gunn, J.; Dave, S. R.; Zhang, M. Q.; Wang, Y. A.; Gao, X. H. Analyst 2008, 133, 154-160.

17. Chen, L. X.; Liu, T.; Thurnauer, M. C.; Csencsits, R.; Rajh, T. The Journal of Physical Chemistry B

2002, 106, 8539-8546.

18. Xu, C. J.; Xu, K. M.; Gu, H. W.; Zheng, R. K.; Liu, H.; Zhang, X. X.; Guo, Z. H.; Xu, B. Journal of

the American Chemical Society 2004, 126, 9938-9939.

19. Rajh, T.; Chen, L. X.; Lukas, K.; Liu, T.; Thurnauer, M. C.; Tiede, D. M. The Journal of Physical

Chemistry B 2002, 106, 10543-10552.

20. Xie, J.; Xu, C.; Xu, Z.; Hou, Y.; Young, K. L.; Wang, S. X.; Pourmand, N.; Sun, S. Chemistry of

Materials 2006, 18, 5401-5403.

21. Hermanson, G. T. Bioconjugate Techniques; Academic Press: New York, 1996.

22. Xie, J.; Xu, C.; Kohler, N.; Hou, Y.; Sun, S. Advanced Materials 2007, 19, 3163-3166.

23. Mosqueira, V. C. F.; Legrand, P.; Gref, R.; Heurtault, B.; Appel, M.; Barratt, G. Journal of Drug

Targeting 1999, 7, 65-78.

24. Kim, D. K.; Zhang, Y.; Kehr, J.; Klason, T.; Bjelke, B.; Muhammed, M. Journal of Magnetism and

Magnetic Materials 2001, 225, 256-261.

 

  

Chapter III

Magnetite (Fe3O4) Nanoparticle for Cell Nucleus Labeling

In the last several chapters, I’ve demonstrated how to synthesize Fe3O4 nanoparticles

with fine control over their core sizes, size distribution, crystallinity, magnetization, as

well as how to change their coating nature and hydrodynamic size. Especially, for DPA-

PEG coated Fe3O4 NPs, I’ve shown that they have much lower uptake by macrophage

cells due to the PEG coating, potentially could be good platform for specific targeting in

MRI and drug delivery. In this chapter, I will discuss one of the efforts I’ve made to

conjugate the functional peptide onto these PEGylated Fe3O4 NPs to target the cell

nucleus.

1. Background

Functionalization of monodisperse superparamagnetic magnetite (Fe3O4) nanoparticles

for cell specific targeting is crucial for cancer diagnostics and therapeutics1-4. Targeted

magnetic nanoparticles can be used to enhance the tissue contrast in magnetic resonance

imaging (MRI)5,6, to improve the efficiency in anticancer drug delivery7,8, and to

eliminate tumour cells via magnetic fluid hyperthermia9-11. Recent synthetic progress

makes it possible to produce monodisperse iron oxide nanoparticles with controlled sizes

and magnetic properties12-15, but interactions between these nanoparticles and

biomolecular entities, especially various tumour cells, are rarely studied due to the

challenge in nanoparticle functionalization and stabilization6,16. In this chapter, I will

show a robust surface functionalization approach to link monodisperse Fe3O4

nanoparticles with Nuclear Localization Signal (NLS) peptide and test their capability in

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targeting tumour cell nuclei. In vitro experiments showed that the uptake of the NLS

labelled nanoparticles by HeLa cells was increased up to 233% compared to the non-NLS

labelled nanoparticles. More importantly, the morphology of the nanoparticles during the

uptake process was unchanged. These nanoparticles and their presence in nuclei were

characterized by fluorescent microscopy, magnetic resonance imaging (MRI) and

transmission electron microscopy (TEM). The work demonstrates that, through proper

surface functionalization, it is possible to stabilize and deliver monodisperse Fe3O4

nanoparticles into tumour cell nuclei for sensitive diagnostic and efficient therapeutic

applications.

NLS represents a group of oligopeptides that contain a few short amino acid

sequences. It is known to act like a 'vector' to direct the protein into the cell nucleus

through the nuclear pore complex17,18, and has recently been applied to Au and dextran-

coated iron oxide nanoparticles for their targeting to cell nuclei19-22. Different from these

previous functionalization steps, my approach is to conjugate biotinylated NLS to

monodisperse Fe3O4 nanoparticles via NeutrAvidin (NAv) and a surfactant combination

in polyethylene glycol (PEG) and dopamine (DPA), or 4-(2-aminoehtyl)benzene-1,2-diol.

Neutravidin (NAv) is a derivative of avidin, but with a closer to neutral isoelectric point

(pI) (6.3 versus 10.5). Similar as avidin and streptavidin (pI=5.5), it is known for the

capability of interacting with biotin. And their paring is widely used as “glue” in

bioconjugation.23  DPA can form a strong chelate chemical bond with iron oxide

surface24,25, and PEG has been widely used to protect nanoparticles for their stabilization

in physiological conditions26,27.

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2. Fe3O4 NPs modification and functionalization

The monodisperse 9 nm Fe3O4 nanoparticles were prepared according to the method in

the previous chapter16 and coated with a hydrophobic layer of oleate and oleylamine

(Figure 3-1b). To render these nanoparticles hydrophilic, I first linked DPA with one

COOH group in α,ω-bis{2-[(3-carboxy-1-oxopropyl)amino]ethyl} polyethylene glycol

(Mr 3000) via conventional dimethylaminopropyl-ethylcarbodiimide/N-

hydroxysuccinimide ester (EDC/NHS) chemistry to synthesize NaOOC-PEG-CONH-

DPA. This NaOOC-PEG-CONH-DPA was then used to replace oleate/oleylamine

around the as-synthesized nanoparticles in CHCl3/DMF solution via the formation of a

chelate bond between Fe3O4 and DPA.25

Figure 3-1. Fe3O4 NPs used in the study: (a) Schematic illustration (not to scale) of the functionalized nanoparticles of NAv-Fe3O4 NPs; (b) TEM image of the 9-nm Fe3O4 nanoparticles coated with oleate/oleylamine; (c) TEM image of the 9-nm Fe3O4 nanoparticles coated with the surfactant shown in (a). (Reprinted with permission from reference28)

Thermogravimetric analysis revealed that each Fe3O4 nanoparticle contained about 32

PEG units. NAv was then conjugated to the -COONa group in NaOOC-PEG-DPA-Fe3O4

via EDC/NHS chemistry to give NAv-NHOC-PEG-DPA-Fe3O4, as illustrated in Figure

86  

  

3-1a,c. The NAv-PEG-DPA-Fe3O4 nanoparticles were further functionalized with a

biotinylated NLS peptide (KKKRKV) by conjugating the peptide to NAv via biotin-

avidin interaction. HeLa cells were chosen for functionalized nanoparticle penetration

and targeting.

Figure 3-1b and c show the TEM images of the monodisperse 9 nm Fe3O4

nanoparticles prior and subsequent to surface modification with DPA-PEG-NAv. Both of

the nanoparticles are well dispersed without any aggregation under both conditions. The

hydrodynamic sizes of the nanoparticles in the dispersions measured by dynamic light

scattering (DLS) (Figure 3-2) revealed that after ligand exchange the overall diameter of

the nanoparticles was increased from ~13 nm in the as-synthesized Fe3O4 to ~50 nm in

the functionalized NAv-PEG-DPA-Fe3O4 nanoparticles.

 

Figure 3-2. Hydrodynamic diameters of (a) the synthesized Fe3O4 NPs in hexane, (b) PEG-DPA-IONPs in water, (c) NAv-Fe3O4 NPs in PBS and (d) NLS-Fe3O4 NPs in PBS. The diameters were measured by DLS. The following parameters were used for size estimation: refractive index 2.420 (Fe3O4), 1.373 (hexane), 1.33 (water); viscosity 0.3000 (hexane), 0.8872 (water); absorption 0.010 (Fe3O4). (Reprinted with permission from reference28)

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Gel electrophoresis analyses on NaOOC-PEG-DPA-Fe3O4 and NAv-PEG-DPA-

Fe3O4 nanoparticle dispersions showed that NAv was closely associated with the

nanoparticles (Figure 3-3).

Figure 3-3. Gel electrophoresis of (a) PEG-Fe3O4 NPs and (b) NAv-Fe3O4 NPs. Both particle dispersions were run on agarose gel (0.5% w/v, 120 min, 100 V) in TAE buffer (40 mm Tris-acetate and 1 mm EDTA, pH 8.3). (Reprinted with permission from reference28)

The dispersion stability of the NAv-PEG-DPA-Fe3O4 and NLS-NAv-PEG-DPA-

Fe3O4 nanoparticles was tested by measuring their hydrodynamic size change during the

incubation in buffer solution. The nanoparticles were dispersed in phosphate buffered

saline (PBS), or PBS plus 10% fetal bovine serum (FBS) and were incubated under

ambient conditions at 37°C. The incubated dispersion was sampled at different time

periods and the average hydrodynamic size of the nanoparticles in each sample was

measured using DLS. Figure 3-4 gives the measurement results from NAv-PEG-DPA-

Fe3O4 and NLS-NAv-PEG-DPA-Fe3O4 nanoparticle dispersions. After incubation for 72

hrs, the average size of these NAv and NLS modified nanoparticles maintained a

hydrodynamic diameter of ~50 nm and ~60 nm for the dispersion in PBS and ~60 nm and

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~80 nm for the dispersion in PBS+10% FBS respectively (Figure 3-4). The increased size

of the functionalized nanoparticles in PBS+10% FBS was presumably due to the

interaction between the negatively charged FBS and the functionalized nanoparticle

surface that bears the positively charged NLS peptide.

 

Figure 3-4. Average hydrodynamic diameters of the Fe3O4 NPs in buffers: a) NAv-Fe3O4 NPs in PBS (pH 7.4), b) NLS-Fe3O4 NPs NPs in PBS, c) NAv-Fe3O4 NPs NPs in PBS+10% FBS, d) NLS-Fe3O4 NPs NPs in PBS + 10% FBS. (Reprinted with permission from reference28)

3. NLS-Fe3O4 NPs for nucleus targeting

To examine whether stable NLS peptide-nanoparticles are suitable for HeLa cell nuclear

targeting, we labelled the NAv first with Rhodamine B isothiocyanate (RA) prior to PEG-

DPA-Fe3O4 nanoparticle conjugation. RA-labelled VKRKKK-biotin-NAv-PEG-DPA-

Fe3O4 and RA-labelled NAv-PEG-DPA-Fe3O4 were then incubated with HeLa cells under

the same condition - 120 mins in Dulbecco's Modification of Eagle's Medium (DMEM)

buffer containing 3.7 mM NaHCO3 and 0.1% bovine serum albumin plus 10% FBS. The

cells were washed with PBS to remove extra nanoparticles. HeLa cells incubated with

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RA-labelled VKRKKK-biotin-NAv-PEG-DPA-Fe3O4 nanoparticles showed a much

brighter fluorescent image (Figure 3-5a) than those with the RA-labelled NAv-PEG-

DPA-Fe3O4 ones (Figure 3-5d), indicating the uptake enhancement for the NLS

nanoparticles due to the NLS-mediated internalization. To characterize the location of the

nanoparticles within the cells, we incubated the HeLa cells with DAPI (4', 6-diamidino-2-

phenylindole) - a blue-fluorescent molecule that can bind preferentially to dsDNA in cell

nucleus to produce a fluorescent enhancement for nuclear imaging29 (Figure 3-5b).

Overlaying of the images from the DAPI staining and the RA staining gives a pink image

in the nucleus (Figure 3-5c). In contrast, the cells incubated with RA-labelled NAv-PEG-

DPA-Fe3O4 nanoparticles do not show the pink area after the overlaying (Figure 3-5f).

These indicate that it is the NLS-coated nanoparticles, not the non-NLS nanoparticles,

which are enriched in the nuclei of the HeLa cells. The average iron concentration in each

cell that was incubated with NLS or non-NLS nanoparticles was measured by inductively

coupled plasma-atomic emission spectrometry (ICP-AES) (Figure 3-5g). For 0.01 mg

Fe/mL NLS-nanoparticle sample, the uptake was increased by 233%.

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Figure 3-5. Characterization of the nanoparticles in HeLa cells: (a) Fluorescent microscopic images of the HeLa cells incubated with RA-labeled NLS-IONPs (0.01 mgFemL-1) and (b) the cells counterstained with DAPI; (c) overlap image of (a) and (b); (d) fluorescence microscope images of the HeLa cells incubated with RA-labeled NAv-IONPs (0.01 mgFemL-1) and (e) the cells counterstained with DAPI; (f) overlap image of (d) and (e); (g) plot of the iron concentration within each HeLa cell that was incubated with NLS-IONPs (black column) and NAv-IONPs (white column) with different concentrations of iron: h) MRI of the HeLa cells containing NLS-IONPs (the first row), NAv-IONPs (the second row); and no nanoparticles (control, the third row). (Reprinted with permission from reference28)

The effect of these nanoparticles on the T2 relaxation of the protons within the

HeLa cells was analyzed with MRI. Figure 3-5h shows the MRI image obtained from the

HeLa cells that had been treated with the NLS-biotin-NAv-PEG-DPA-Fe3O4

nanoparticles (the first row) and the NAv-PEG-DPA-Fe3O4 nanoparticles (the second

row) at different concentrations as indicated. There is no apparent difference in image

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signal intensity between the cells containing NAv-PEG-DPA-Fe3O4 nanoparticles and the

cells containing no particles (control). In contrast, images from the cells containing NLS-

biotin-NAv-PEG-DPA-Fe3O4 nanoparticles are much darker, indicating that the

nanoparticles within the cells do offer a strong contrast enhancement in MRI.

Figure 3-6. TEM images of the nanoparticles in one HeLa cell: (a) The NLS-IONPs around cell membrane and cytoplasm area; (b) the NLS-IONPs in the cell nucleus; (c) a close-up view of the white box area in (b); (d) the NAv-IONPs enriched outside the nuclear membrane area. (Reprinted with permission from reference28)

 

The Fe3O4 nanoparticle uptake by HeLa cells was visualized by TEM to confirm

cellular localization and dispersion state of these nanoparticles. Figure 3-6 a-c shows the

NLS-peptide nanoparticle uptake by a single HeLa cell after incubation of the cells with

the NLS peptide-nanoparticle for a period of 2 hrs. It can be seen that the nanoparticles

are extensively dispersed in cytoplasm without apparent aggregation. For the NLS

peptide coated Fe3O4 nanoparticles, their nuclear accumulation was clearly observed in

Figure 3-6b and c. The nanoparticles are well dispersed and spread in the nuclear area. As

a comparison, most of the non-NLS coated nanoparticles, which entered the cells in much

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smaller amount compared with the NLS-nanoparticles, stay outside the nuclear envelope

(Figure 3-6d), indicating that the non-NLS nanoparticles are difficult to translocate into

the nucleus.

The particles can enter the cells via either endocytosis or diffusion process or both30.

To have a preliminary understanding in the uptake mechanism of the nanoparticles

reported in this work, the cells were incubated with sodium azide NaN3 - a well-known

metabolic inhibitor31, and DAPI as well as RA labelled NLS-NAv-PEG-DPA-Fe3O4

nanoparticles (0.01 mg Fe/mL). Fluorescent microscopic images of the so incubated cells

show that there were no particles in the cells (Figure 3-7a and b), indicating that the

nanoparticles enter the cell membrane through an endocytosis process. The presence of

the well-dispersed nanoparticles in the cytoplasm shown in Figure 3-6 suggests that our

nanoparticles survive and escape from the lysosome (or a similar organelle) environment

easily during their intracellular passage, which is an essential step for their targeting to

the nucleus.

Figure 3-7. Fluorescent microscopic images of the HeLa cells incubated with NaN3 (0.1% by wt) with (a) DAPI (30 nM) and (b) RA labeled NLS-NAv-PEG-DPA-Fe3O4 nanoparticles (0.01 mg Fe/mL). (Reprinted with permission from reference28)

4. Summary

I have shown that monodisperse Fe3O4 nanoparticles prepared from an organic phase

synthesis can be readily functionalized with hydrophilic DPA-PEG-based surfactant and

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stabilized in physiological conditions. The NLS-peptide coated nanoparticles show

preferred uptake by HeLa cell nuclei over the non-NLS labelled nanoparticles. Using the

similar synthetic strategy, one can coat the monodisperse iron oxide nanoparticles with

various signal peptides, genes, or drugs and target them into different cellular

compartments. These will allow the detailed studies in uptake mechanism (endocytosis)

and targeting of these particles in cells, especially tumour cells. The understanding will

help to create novel functional magnetic nanoprobes that are suitable for highly sensitive

medical diagnostics and highly efficient drug/gene delivery.

5. Experimental

Materials and Instruments: Fe(acac)3, α,ω-bis(2-carboxyethyl)polyethylene glycol

(MW = 3,000), dopamine hydrochloride, sodium azide, sodium carbonate and organic

solvents used in the syntheses were purchased from Sigma Aldrich. NeutrAvidin (NAv),

N-hydroxysuccinimide (NHS), N-(3-dimethylaminopropyl)-N’- ethylcabodiimide (EDC)

hydrochloride and 4',6-diamidino-2-phenylindole (DAPI) were from Pierce

Biotechnology. All the buffers and media were from Invitrogen Corp. Water was purified

by Millipore Milli-DI Water Purification System. Nano-sep 100k OMEGA was from

Fisher. All the dialysis bags were purchased from Spectrum Laboratories, Inc.

Synthesis of NaOOC-PEG-DPA-Fe3O4 NPs: α,ω-Bis{2-[(3-carboxy -1-

oxopropyl)amino]ethyl}polyethylene glycol (20 mg) , NHS (2 mg), EDC (3 mg) and

dopamine hydrochloride (1.27 mg) were dissolved in a mixture solvent containing CHCl3

(2 mL), DMF (1 mL), and anhydride Na2CO3 (10 mg). The solution was stirred at room

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temperature for 2 hrs before Fe3O4 nanoparticles (5 mg) was added, and the resulted

solution was stirred overnight at room temperature under N2 protection. The modified

Fe3O4 nanoparticles were precipitated by adding hexane, collected by a permanent

magnet and dried under N2. The particles were then dispersed in water or PBS. The extra

surfactants and other salts were removed by dialysis using a dialysis bag (MWCO =

10,000) for 24 hour in 1× PBS or water. Any precipitation (almost none in the synthesis)

was removed by a 200 nm syringe filter (MILLPORE Corp.). The final concentration of

the particles was determined by ICP-AES analysis.

Labeling NeutrAvidin (NAv) with Rhodamine B isothiocyanate (RA): NAv was

incubated with RA in Na2CO3/NaHCO3 (pH = 9) buffer at room temperature for 1 h. The

ratio between RA and NAv was 10:1. The final conjugate was purified to remove the

extra free RA by PD-10 column (GE Healthcare Corp.).

Conjugating RA-NAv to NaOOC-PEG-DPA-Fe3O4 NPs: NaOOC-PEG-DPA-Fe3O4 (2

mg) in PBS solution was incubated with EDC (0.1 mg) at room temperature for 15 mins.

Then RA-NAv (100 µg) in 1 x PBS was added. The mixture was incubated for 1 h. The

product was purified by dialysis over dialysis bag (MWCO = 100,000) in 1 x PBS for 24

hrs.

Conjugating biotin-KKKRKV to NAv-PEG-DPA-Fe3O4 NPs: Biotin-KKKRKV was

incubated with the NAv-PEG-DPA-Fe3O4 nanoparticles in 1× PBS (pH = 7.4) for 1 h

followed by filtering [Nanosep filter (MWCO = 100,000)] or dialyzing [Dialysis

Membranes (MWCO = 1000,000)] for 24 hrs in dark to remove the excessive peptides.

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Incubation of the RA-labeled NAv-PEG-DPA-Fe3O4 NPs and the RA-labeled

VKRKKK-biotin-NAv-PEG-DPA-Fe3O4 NPs with HeLa cells: HeLa cells were

cultured in glass bottom Petri dish (MatTek Corp.) with Dulbecco's Modified Eagle's

Medium (DMEM) with 10% FBS and 1% antibiotics. Before incubation with particles,

the cells were washed with 1 x PBS for 3 times. And then the particle solution in DMEM

media was incubated with cells for 2 hrs. For inhibition of the particles uptake by Sodium

Azide, HeLa cells were incubated with 0.1% NaN3 as well as 0.01mg/ml particles in

DMEM buffer for 120min.Then those cells were washed with PBS for 3 times and fixed

by 4% paraformadihyde solution. After 30 min fixation, the cells were washed by PBS 3

times and subjected to fluorescent microscope (Nikon Eclipse TE2000-U) or MRI. To

counterstain the cells with DAPI, a DAPI solution (30 nM) in PBS was mixed with the

cells for 5 min after paraformadihyde fixation and washed with PBS.

References:

1. Ferrari, M. Nature Reviews Cancer 2005, 5, 161-171.

2. Pankhurst, Q. A.; Connolly, J.; Jones, S. K.; Dobson, J. Journal of Physics D-Applied Physics 2003, 36,

R167-R181.

3. Mornet, S.; Vasseur, S.; Grasset, F.; Duguet, E. Journal of Materials Chemistry 2004, 14, 2161-2175.

4. Sunderland, C. J.; Steiert, M.; Talmadge, J. E.; Derfus, A. M.; Barry, S. E. Drug Development

Research 2006, 67, 70-93.

5. Bulte, J. W. M.; Kraitchman, D. L. Nmr in Biomedicine 2004, 17, 484-499.

6. Huh, Y. M.; Jun, Y. W.; Song, H. T.; Kim, S.; Choi, J. S.; Lee, J. H.; Yoon, S.; Kim, K. S.; Shin, J. S.;

Suh, J. S.; Cheon, J. Journal of the American Chemical Society 2005, 127, 12387-12391.

7. Brannon-Peppas, L.; Blanchette, J. O. Advanced Drug Delivery Reviews 2004, 56, 1649-1659.

8. Dobson, J. Drug Development Research 2006, 67, 55-60.

9. Jordan, A.; Scholz, R.; Wust, P.; Fahling, H.; Felix, R. Journal of Magnetism and Magnetic Materials

1999, 201, 413-419.

10. Ivkov, R.; DeNardo, S. J.; Daum, W.; Foreman, A. R.; Goldstein, R. C.; Nemkov, V. S.; DeNardo, G.

L. Clinical Cancer Research 2005, 11, 7093S-7103S.

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11. Hilger, I.; Hergt, R.; Kaiser, W. A. Journal of Magnetism and Magnetic Materials 2005, 293, 314-319.

12. Rockenberger, J.; Scher, E. C.; Alivisatos, A. P. Journal of the American Chemical Society 1999, 121,

11595-11596.

13. Hyeon, T.; Lee, S. S.; Park, J.; Chung, Y.; Bin Na, H. Journal of the American Chemical Society 2001,

123, 12798-12801.

14. Sun, S. H.; Zeng, H. Journal of the American Chemical Society 2002, 124, 8204-8205.

15. Sun, S. H.; Zeng, H.; Robinson, D. B.; Raoux, S.; Rice, P. M.; Wang, S. X.; Li, G. X. Journal of the

American Chemical Society 2004, 126, 273-279.

16. Xie, J.; Xu, C. J.; Xu, Z. C.; Hou, Y. L.; Young, K. L.; Wang, S. X.; Pourmond, N.; Sun, S. H.

Chemistry of Materials 2006, 18, 5401-5403.

17. Frankel, A. D.; Pabo, C. O. Cell 1988, 55, 1189-1193.

18. Vives, E.; Brodin, P.; Lebleu, B. Journal of Biological Chemistry 1997, 272, 16010-16017.

19. Josephson, L.; Tung, C. H.; Moore, A.; Weissleder, R. Bioconjugate Chemistry 1999, 10, 186-191.

20. Lewin, M.; Carlesso, N.; Tung, C. H.; Tang, X. W.; Cory, D.; Scadden, D. T.; Weissleder, R. Nature

Biotechnology 2000, 18, 410-414.

21. Tkachenko, A. G.; Xie, H.; Coleman, D.; Glomm, W.; Ryan, J.; Anderson, M. F.; Franzen, S.;

Feldheim, D. L. Journal of the American Chemical Society 2003, 125, 4700-4701.

22. de la Fuente, J. M.; Berry, C. C. Bioconjugate Chemistry 2005, 16, 1176-1180.

23. Wilchek, M.; Bayer, E. A.; Livnah, O. Immunology Letters 2006, 103, 27-32.

24. Rajh, T.; Chen, L. X.; Lukas, K.; Liu, T.; Thurnauer, M. C.; Tiede, D. M. Journal of Physical

Chemistry B 2002, 106, 10543-10552.

25. Xu, C. J.; Xu, K. M.; Gu, H. W.; Zheng, R. K.; Liu, H.; Zhang, X. X.; Guo, Z. H.; Xu, B. Journal of

the American Chemical Society 2004, 126, 9938-9939.

26. Storm, G.; Belliot, S. O.; Daemen, T.; Lasic, D. D. Advanced Drug Delivery Reviews 1995, 17, 31-48.

27. Gref, R.; Domb, A.; Quellec, P.; Blunk, T.; Muller, R. H.; Verbavatz, J. M.; Langer, R. Advanced

Drug Delivery Reviews 1995, 16, 215-233.

28. Xu, C. J.; Xie, J.; Kohler, N.; Walsh, E. G.; Chin, Y. E.; Sun, S. H. Chemistry-an Asian Journal 2008,

3, 548-552.

29. Kapuscinski, J. Biotechnic & Histochemistry 1995, 70, 220-233.

30. Gao, H. J.; Shi, W. D.; Freund, L. B. Proceedings of the National Academy of Sciences of the United

States of America 2005, 102, 9469-9474.

31. Torchilin, V. P.; Rammohan, R.; Weissig, V.; Levchenko, T. S. Proceedings of the National Academy

of Sciences of the United States of America 2001, 98, 8786-8791.

  

Chapter IV

pH Controlled Release of Chromone from Chromone-Fe3O4

Nanoparticles for Cancer Cell Growth Inhibition

In the chapter two, I’ve successfully demonstrated how to synthesize and modify Fe3O4

nanoparticles (NPs) with dopamine and bifunctional poly(ethylene glycol). In the

previous chapter, I also showed the functionalization of modified Fe3O4 NPs with nucleus

localization signal. The modified Fe3O4 NPs are stable and against macrophage cell

uptake before and after functionalization. Thus, the NPs are ready for the specific

application in MRI and drug delivery. In this chapter, I will discuss one of the

applications, Chromone delivery for the dopamine-PEG modified Fe3O4 NPs for cancer

cell growth inhibition.

1. Background

Chromones are an important class of compounds belonging to the flavonoid group that

occur naturally in plants. They are minor constituents of the human diet and have been

reported to exhibit a wide range of biological effects. These biological properties include

anti-inflammatory, antibacterial, antitumor, antioxidant, anti-HIV, vasodilator, antiviral

and antiallergenic. To date only a few flavonoids, such as Flavopiridol, have entered

clinical trials.1-4 Due to their low water-soluble ability and a short blood circulation time,

the usage of most flavonoids is still limited. Recent studies suggest that using water-

soluble drug delivery system can overcome the some drawbacks of anticancer drug, so as

to improve the therapy with these anti-cancer agents.

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As a good drug delivery vehicle, they must meet the following requests: (1) drug-

loading capacity; (2) desired release profile; (3) aqueous dispersion stability; (4)

biocompatibility with cells and tissue, and nontoxicity.5 Generally the drug can be

released through the following ways: (1) Ion concentration; (2) pH-responsive; (3)

Enzyme-Mediated.6 It is well-known that certain tissues of the body have a pH slightly

more acidic than the blood and normal tissue. Therefore, the carrier system based on

mildly acidic pH provides a safe and efficient way for drug release targeting specific sites

in the body, such as tumor and inflammatory tissues (pH 6.8), endosomes (pH 5.5-6),

and lysosomes (pH 4.5-5.0).

Magnetic Fe3O4 nanoparticles (NPs) are promising as drug delivery vehicles for both

diagnostic and therapeutic applications.7,8 The key to achieving these dual applications is

that the drug-Fe3O4 NPs are stable in biological circulation system, readily interact with

cells or other biological units of interest, and are capable of releasing the drug once the

selected targeting is realized.9,10 Currently, drug-Fe3O4 NP conjugates are made either by

embedding the drug in the hydrophobic media in the double-layer coating of Fe3O4

NPs,11 or by incorporating both drug and Fe3O4 NPs in the SiO2 matrix.12,13 Although the

conjugates prepared from these methods show enhanced dispersion stability, they have a

hydrodynamic diameter of 150 nm or larger. Such large NP delivery systems may have

very limited extravasation ability and may be subject to easy uptake by the

reticuloendothelial (RES) system,14,15 unsuitable for target-specific delivery applications.

In chapter two, I have described that Fe3O4 NPs coated with dopamine (DPA) and

COOH-terminated polyethylene glycol (PEG) are stable in cell culture media against

macrophage cell uptake.16 The hydrodynamic sizes of the NPs are tuned by the length of

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the PEG molecules. These PEG-DPA-Fe3O4 NPs offer an ideal platform for drug

coupling and delivery.

The plan for chromone delivery is that chromone, 6-hydroxy-chromone-carbaldehyde

(1a), can be readily coupled to these PEG-DPA-Fe3O4 NPs via a Schiff base bond, as

shown in Figure 4-1a, and released via a pH controlled manner. We demonstrate that 6-

hydroxy-chromone-carbaldehyde (1a) coupled to PEG-DPA-Fe3O4 (1c) show a dramatic

increase in solubility in cell culture medium, from less than 2.5 μg/mL for free chromone

to 633 μg/mL for chromone-PEG-DPA-Fe3O4 (1d). Such chromone-Fe3O4 NPs also

inhibit HeLa cell proliferation more efficiently than the free chromone. 1d is stable in

neutral pH condition but unstable in pH lower than 6 due to the fast hydrolysis of the

Schiff base bond, releasing free chromone. Due to the characteristic fluorescent

properties of chromone, 1d also acts as an optical probe for on-time tracking of the

chromone-NPs in cells. This, plus the intrinsic superparamagnetic properties of Fe3O4

NPs, renders 1d a powerful multifunctional delivery system for diagnostics and

therapeutic applications.

Figure 4-1. (a) Structure of chromone (1a) and the schematic illustration of the coupling between chromone and a Fe3O4 NP; TEM images of (b) the as-synthesized 12 nm Fe3O4 NPs from the hexane dispersion and (c) the chromone modified Fe3O4 NPs (1d) from water. Reprinted with permission from reference17.

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2. Fe3O4 NPs modification and functionalization

Fe3O4 NPs were synthesized through the decomposition of Fe(acac)3 with the core size

around 12 nm (Figure 4-1b).16 The as-synthesized NPs were coated with a layer of

oleate/oleylamine and are hydrophobic. The NPs were made biocompatible by replacing

oleate/oleylamin with DPA-PEG (1b).

PEG2000

BrCH2COCl

Et3N

K2CO3 KI

Dopamine

K2CO3 KI

BOCNHCH2CH2NH2

HCl

HOO

OHn OO

OnBr

OOBr

OO

OnBr

OO

NH

HO

HO

OO

On

HN

OO

NH

HO

HO NH

Boc

DPA-PEG-NH2HCl (1b)

OO

On

HN

OO

NH

HO

HO NH2 HCl

Figure 4-2. Synthesis of DPA-PEG-NH2 or 1b. Reprinted with permission from reference17.

To synthesize 1b, we treated poly(ethylene glycol) (MW=2000) with bromoacetyl

chloride to convert the OH’s to bromide, which could easily react with amine group

through a nucleophilic substitution reaction (Figure 4-2). Through two steps reaction, we

successively linked DPA to one end of HO-PEG-OH and n-tert-butoxycarbonyl-1,2-

ethanediamine to another. The protection group tert-butoxycarbonyl on the amide group

was removed by incubation with 4 M HCl/dichloromethane for 40 minutes. 1b was then

used to replace oleate/oleylamine from the as-synthesized Fe3O4 NPs, giving 1c. 1a was

loaded onto 1c via the formation of a Schiff-base bond between the primary amine group

present on 1c and the aldehyde group on 1a. The NPs of the 1d conjugate were readily

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dispersed in water and did not show observable change in core morphology after these

surface modification steps (Figure 4-1c). Compared with the original solubility of

chromone (less than 2.5 µg/ml), the new conjugates are highly soluble in aqueous

solution with chromone solubility reaching 633 µg/ml, equal to ~140 chromone

molecules per Fe3O4 NP.

Figure 4-3. Fluorescent spectra of 1d, 1c and 1a with the same Fe or chromone concentration. Reprinted with permission from reference17.

The presence of chromone on NPs was characterized by both fluorescence and

infrared (IR) spectra. 1d in Figure 4-1 shows fluorescent emission at 455 nm, blue-shifted

from the 1a emission at 468 nm (Figure 4-3). However, NPs without chromone exhibit no

fluorescent emission. IR spectrum of 1d (Figure 4-4) has a ketone (C=O) vibration at

1644 cm−1 that does not appear in 1c. This vibration is red-shifted from 1693 cm-1 of 1a,

demonstrating the covalent bonding between chromone and PEG. The vibration at 1589

cm−1 for 1d can be assigned to the vibration peak of C=N, suggesting that chromone in

1d is connected to Fe3O4 NPs through a Schiff-base bond.

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Figure 4-4. IR spectra of 1d, 1c and 1a. Reprinted with permission from reference17.

3. Controlled chromone release from Chromone-Fe3O4 NPs

The Schiff-base bond is biodegradable via hydrolysis and the process can be accelerated

at low pH conditions.18,19 To examine the pH controlled release of chromone in 1d, we

put the conjugate in the dialysis bag and incubated at 37oC in different buffer systems

with pH ranging from 3 to 9. The released chromone was quantified through its

fluorescent signal. Figure 4-5a shows the percentage of chromone released from 1d at

different pHs. It can be seen that few chromone is detached from 1d in pH > 7 conditions.

However, low pH (<6) leads to drastic increase in free chromone concentration,

indicating the increased release of free chromone from 1d. At pH 5 and 7, the incubation

temperatures (37oC and 20oC) have little influence on the chromone release, as shown in

Figure 4-5b.

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Figure 4-4. (a) Chromone release from 1d under different pH conditions at 37°C; (b) Chromone release from 1d under different incubation pH’s and temperatures. Reprinted with permission from reference17.

The hydrodynamic size of 1d is decreased in pH = 5, but those in pH = 7.4 are stable, as

shown in Figure 4-6. This proves that chromone is released from 1d at low pH but is stable in the

conjugate at pH > 7. From Figure 4-6, one can also see that 1c are stable in the incubation

conditions and show no statistical hydrodynamic size change over the incubation time. The

measured size increase from ~60 to ~110 nm in the presence of FBS is attributed to the

adsorption of FBS onto the NP surface as reported previously.20

Figure 4-6. (a) Stability of Fe3O4-DAP-PEG-N-chromone in 1x PBS buffer plus 10% FBS under 37 oC with pH=7.4 and 5; (b) Stability of Fe3O4-DAP-PEG-NH2 in 1x PBS buffer plus 10% FBS under 37 oC with pH=5. Reprinted with permission from reference17.

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The increased solubility of chromone present in 1d led to the enhanced uptake of 1d

by HeLa cells (Figure 4-7a). As we can see, HeLa cells show preferred uptake for

chromone-Fe3O4 NPs under three different concentrations. Figure 4-7b&c further shows

that at the same iron concentrations (7 μg/ml), more 1d than 1c are taken up by HeLa

cells, which corresponds to the higher fluorescent signal. Similar uptake enhancement is

also observed for 1d over 1a.

Figure 4-7. (a) HeLa cell uptake comparison of 1c and 1d NPs. (b) Fluorescent image of HeLa cells after incubated with Fe3O4-Chromone for 1 hour and (c) HeLa cells after incubated with PEG-DPA-Fe3O4 NPs for 1 h. Reprinted with permission from reference17.

Figure 4-8a&b are fluorescent images of the HeLa cells after their incubation with 1d

and 1a in the same chromone concentration at 15 μg/ml. Due to the high chromone

solubility in 1d, there exist more chromone molecules in solution interacting with HeLa

cells, leading to the enhanced uptake and brighter image of the cells in Figure 4-8a. In

contrast, the free chromone has very low solubility and with the same total amount of

chromone added, the majority of the free chromone stays in the solid form and can be

separated by centrifugation (8000 rpm). As a result, there is only small amount of free

chromone in solution interacting with the cells, resulting in fewer uptakes and much

darker fluorescent image (Figure 4-8b).

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Figure 4-8. Fluorescent images of HeLa cells after incubated with (a) 1d and (b) 1a for 1 h; and Viability of HeLa cells in the presence of (c) total iron concentration and (d) total chromone concentration.

The enhanced uptake of 1d leads to high toxicity to the HeLa cells. Figure 4-8c&d are

the HeLa cell viability data under the same iron (Figure 4-8c) and chromone

concentration (Figure 4-8d). It can be seen that both 1c and 1a have very limited toxicity

to HeLa cells while 1d are highly toxic with majority of the cells destroyed at ~100 ppm

iron concentration or at ~40 μg chromone/ml. Clearly this high toxicity of 1d to the HeLa

cells comes from their enhanced uptake by the HeLa cells and the controlled release of

free chromone from 1d in the low pH cellular environment.

 

4. Summary

The current work demonstrates that free chromone coupled to PEG-DPA-Fe3O4 NPs

show a dramatic increase in chromone solubility in cell culture medium from less than

2.5 μg/mL to 633 μg/mL, and the free chromone can be released in a controlled manner

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at low pH conditions. The high chromone solubility in the chromone-Fe3O4 conjugate

leads to the enhanced chromone uptake by HeLa cells and as a result, much more

efficient inhibition to the HeLa cell proliferation. With intrinsic fluorescent,

superparamagnetic and toxic properties, the chromone-Fe3O4 NPs should serve as a

powerful multifunctional delivery system for both chromone-based diagnostic and

therapeutic applications.

5. Experimental

Materials and Instruments: Iron (III) acetylacetonate was from Strem Chemicals, Inc..

All other chemicals including Triethylamine, potassium iodide, bromoacetyl chloride,

potassium carbonate, benzyl ether, oleic acid, oleylamine, poly(ethylene glycol)(Mol

MW=2000) etc were purchased from Sigma-Aldrich and used without further

purification. N-tert-Butoxycarbonyl-1,2- ethylenediamine and Chromone was synthesized

according to the published method.21 Deionized (DI) water was purified by a Millipore

Milli-DI Water Purification system. 1H NMR spectra were acquired with Varian 300

MHz NMR. TEM measurements were taken on a Philips EM 420 (120 kV). UV/Vis

absorption spectra of the samples were measured with a PerkinElmer Lambda 35 UV/Vis

spectrometer. The fluorescence spectra were acquired on Fluoromax 4 (HORIBA JOBIN

YVON Inc.) spectrofluorometer. Fluorescent pictures were taken on Zeiss Leica inverted

epifluorescence /reflectance laser scanning confocal microscope. Hydrodynamic sizes of

NPs were measured by Malvern Zeta Sizer S90 dynamic light scattering instrument.

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Synthesis of Fe3O4 Nanoparticles: Fe(acac)3 (0.706 g, 2 mmol) was dissolved in a

mixture of benzyl ether (10 mL) and oleylamine (10 mL). The above mixture solution

was dehydrated at 110℃ for 1 h under a flow of nitrogen, and quickly heated to 300℃

and kept at this temperature for 2 h under a blanket of nitrogen. The black-brown mixture

was cooled to room temperature later. Ethanol (40 mL) was added to the mixture and

precipitate was collected by centrifugation at 8000 rpm. Finally, the product was re-

dispersed in hexane.

Synthesis of O,O’-Bis(2-Bromoacetyl)polyethylene glycol (BBrAc-PEG):

Poly(ethylene glycol) (Mol MW=2000) (10 g, 5 mmol) was dissolved in anhydrous

dichloromethane (20 mL). Triethylamine (2.09mL, 15mmol) was added dropwise,

followed by addition of 1.249mL (15mmol) bromoacethyl chloride dropwise under

nitrogen. The reaction was stirred overnight in the dark. The product was purified by

precipitation in diethyl ether. After the product was dissolved in water, pH of the solution

was adjusted to 6. The compound was then extracted three times with 20 mL of

dichloromethane and precipitated out by addition of diethyl ether and stored at -20℃. 1H

NMR (300 MHz, chloroform-d6): δ 3.3-3.7 (232H, -O-CH2-CH2-O-), 4.15 (s, 4H, -CH2-

Br), 4.2 (t, 4H, -CH2-COO-).

Synthesis of O-(2-Bromoacetyl)-O’-(2-Dopamineacetyl)polyethylene glycol (DPA-

PEG-BrAc): BBrAc-PEG (448 mg, 0.2mmol) was dissolved in anhydrous

dichloromethane (20 mL). And then dopamine hydrochloride (41.58 mg, 0.22mmol), KI

(16.6 mg) and K2CO3 (70 mg) were added to the above solution. The mixture was stirred

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for 10 hrs at 25 ℃ under nitrogen. The insoluble compounds were filtered, and the filtrate

was added to diethyl ether (100 mL). The precipitation was collected by centrifugation

and dissolved in water. BrAc-PEG-DPA was extracted with dichloromethane (10mL x 3),

and precipitated out with diethyl ether (150 mL) on dry ice. The product was then stored

at -20℃. 1H NMR (300 MHz, chloroform-d6): δ 2.634 (t, 2H, -CH2-CH2N-), 2.86 (t, 2H,

-Ph-CH-CH2-), 3.4-3.6 (234H, -O-CH2-CH2-O), 4.07 (s, 2H, -CH2-Br), 4.24 (t, 4H, -CH2-

COO-), 6.54 (d, 1H, Ph), 6.75 (m, 2H, Ph).

Synthesis of O-[2’-(Boc-imino-ethylene-imino)acetyl-]-O’-(2-

Dopamineacetyl)polyethylene glycol (DPA-PEG-NHBoc): N-tert-Butoxycarbonyl-1,2-

ethylenediamine (16.0 mg, 0.1mmol) was dissolved in 20 mL dichloromethane. DPA-

PEG-BrAc (239.2 mg, 0.1mmol), KI (16.6mg) and K2CO3 (70 mg) were added later and

stirred for 10 hrs at 25℃ under nitrogen. Following the workup procedures described in

the synthesis of BrAc-PEG-DPA, the product was stored at -20 ℃. 1H NMR (300 MHz,

chloroform-d6): δ 1.41 (s, 9H, t-Bu), 2.68 (t, 2H, -CH2-CH2N-), 2.78 (t, 2H, -Ph-CH-

CH2-), 2.88 (t, 2H, CH2NH-BOC), 3.32 (t, 2H, CH2CH2NHCH2-)3.4-3.6 (234H, -O-CH2-

CH2-O), 4.07 (s, 2H, -CH2-NHCH2CH2-Ph), 4.24 (t, 4H, -CH2-COO-), 6.71 (d, 1H, Ph),

6.91 (m, 2H, Ph).

Synthesis of O-(2’-(Amino-ethylene-imino)acetyl-)-O’-(2-

Dopamineacetyl)polyethylene glycol (DPA-PEG-NH2⋅HCl): DPA-PEG-NHBoc (255.2

mg) was added to 4M HCl/Dichloromethane. After 40 minutes, the solvent was removed

under reduced pressure to obtain a light solid.

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Preparation of Fe3O4-DAP-PEG-NH2: DPA-PEG-NH2⋅HCl (50 mg) was dissolved in

dichloromethane (5 mL), and then Fe3O4 (10 mg) in 1 mL dichloromethane was added.

The mixture was stirred overnight at room temperature. The modified Fe3O4

nanoparticles were precipitated by adding hexane, and collected by centrifugation at 6000

rpm. After washed with dichloromethane and hexane (1/5, v/v) three times, the product

was re-dispersed in ethanol.

Preparation of Fe3O4-DAP-PEG-N-Chromone: Fe3O4-DAP-PEG-NH2 (10 mg) was

mixed with chromone (10 mg) in ethanol. The mixture was stirred for 5 hrs at room

temperature. The product was precipitated by adding hexane, and collected by

centrifugation at 6000 rpm. After washed with ethanol and hexane (1/5, v/v) 3 times, the

product was re-dispersed in DI water. And this final conjugates were filtered through

0.22 μm Millex@GP filter (Millipore Corp.) to remove aggregates.

Determination of Chromone Concentration: Chromone concentration was determined

based on Lambert-Beer law. The ε values at different pHs were obtained through

measuring the emission of chromone with different concentrations (2×10-5, 3 ×10-5, 4

×10-5, 5 ×10-5, 6 ×10-5 M) at different pH buffer with 20% ethanol. The ε values here are

listed in Table 4-1.

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Table 4-1. The ε values of chromone at different pHs.

pH ε (M-1·cm-1)

3 4.00×108

4 6.70 × 108

5 9.10 × 108

6 5.62× 109

7 7.40× 1010

8 3.80× 1010

9 3.20 × 1010

Chromone Release under Different pHs: In vitro chromone release properties from the

nanoparticles were determined as follows: 2 mL of Fe3O4-DAP-PEG-N-Chromone

(1.98×10-2 M, chromone) solution was put into dialysis bag (MWCO=100k, Spectrum

Laboratories, Inc) and each of them was immersed into 20 mL different buffers

containing 20% C2H5OH. At a definite time interval, 2 mL of the solution outside the

dialysis bag was sampled and the chromone concentration was determined through

Lambert-Beer law. Buffers with different pH values were prepared from borate buffer

(pH=9), phosphate Buffers (pH=8, 7 and 6) and acetate Buffers (pH=5, 4 and 3).

Cytotoxicity assay (MTT method): The experiments were performed using the

following human cancer cell lines: HeLa (cervical) was bought from ATCC. Cells were

cultured in 75 cm2 flasks (Corning) containing 10 mL DMEM with 10% fetal bovine

serum and 1% antibiotics. Cytotoxicity assay was performed in 96-wells microtiter plates

(Fisher Inc.) with seeding density, 4000 cells per well. Microtiter plates containing cells

were pre-incubated for 24 hours at 37oC in order to allow stabilization before the addition

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of the test substance. The plates were incubated with the test substance for 48 hours at 37

oC and 5% CO2. Then 5 μL MTT solution (5 mg/mL in PBS) was added to each well to

evaluate cell viability. After 2 h at 37oC, the solution was removed. 100 uL DMSO was

added to dissolve cells. After 30min incubation under 37 oC, the viability was measured

through microreader.

Stability Measurement: NPs were incubated PBS with 10% FBS in water bath. After

certain time, the samples were examined with dynamic light scattering. Each experiment

was repeated four times.

Uptake experiment for nanoparticles: HeLa cells were cultured in DMEM (containing

10% FBS and 1% antibiotics) in T25 flasks. For experiments, 200,000 HeLa cells were

seeded into each T25 flask. After 24 hours, Fe3O4-DAP-PEG-N-Chromone or Fe3O4-

DAP-PEG-NH2 NPs were incubated with cells for 4 hours. Then cells were washed with

PBS and detached with Trypsin-EDTA (0.25%Trypsin; 1mM EDTA4Na)(1×). After

collected with centrifugation, the cells were counted and dissolved with Aqua Regia. Fe

concentration was determined with ICP.

Uptake experiment for chromone. 200,000 HeLa cells were seeded into each T25 flask.

After 24 hours, Fe3O4-DAP-PEG-N-chromone or chromone were incubated with cells for

2 hours under the same chromone concentrations (110, 55 and 20 µg/ml). Then cells were

washed with PBS and detached with Trypsin-EDTA (0.25%Trypsin; 1mM

EDTA·4Na)(1×). After collected with centrifugation, the cells were counted, dispersed in

PBS and subject to fluorometer.

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Cell fluorescent images: HeLa cells were purchased from ATCC and cultured in glass

bottom Petri dish (MatTek Corp.) with Dulbecco's Modified Eagle's Medium (DMEM)

with 10% FBS and 1% antibiotics. And the particle solution in DMEM media was

incubated with cells for 1 hr. Then those cells were washed with PBS for 3 times and

fixed by 4% paraformadehyde solution. After 30 min fixation, the cells were washed by

PBS and subjected to fluorescent imaging.

References:

1. Edwards, A. M.; Howell, J. B. L. Clinical and Experimental Allergy 2000, 30, 756-774.

2. Mukherjee, A. K.; Basu, S.; Sarkar, N.; Ghosh, A. C. Current Medicinal Chemistry 2001, 8, 1467-

1486.

3. Barve, V.; Ahmed, F.; Adsule, S.; Banerjee, S.; Kulkarni, S.; Katiyar, P.; Anson, C. E.; Powell, A. K.;

Padhye, S.; Sarkar, F. H. Journal of Medicinal Chemistry 2006, 49, 3800-3808.

4. Pisco, L.; Kordian, M.; Peseke, K.; Feist, H.; Michalik, D.; Estrada, E.; Carvalho, J.; Hamilton, G.;

Rando, D.; Quincoces, J. European Journal of Medicinal Chemistry 2006, 41, 401-407.

5. Allen, T. M.; Cullis, P. R. Science 2004, 303, 1818-1822.

6. Peer, D.; Karp, J. M.; Hong, S.; Farokhzad, O. C.; Margalit, R.; Langer, R. Nat Nano 2007, 2, 751-760.

7. Xu, C. J.; Sun, S. H. Polymer International 2007, 56, 821-826.

8. Jun, Y. W.; Lee, J. H.; Cheon, J. Angewandte Chemie-International Edition 2008, 47, 5122-5135.

9. Arruebo, M.; Fernandez-Pacheco, R.; Ibarra, M. R.; Santamaria, J. Nano Today 2007, 2, 22-32.

10. Torchilin, V. P. Nature Reviews Drug Discovery 2005, 4, 145-160.

11. Jain, T. K.; Morales, M. A.; Sahoo, S. K.; Leslie-Pelecky, D. L.; Labhasetwar, V. Molecular

Pharmaceutics 2005, 2, 194-205.

12. Kohler, N.; Sun, C.; Fichtenholtz, A.; Gunn, J.; Fang, C.; Zhang, M. Small 2006, 2, 785-792.

13. Son, S. J.; Reichel, J.; He, B.; Schuchman, M.; Lee, S. B. Journal of the American Chemical Society

2005, 127, 7316-7317.

14. Hu, Y.; Xie, J. W.; Tong, Y. W.; Wang, C. H. Journal of Controlled Release 2007, 118, 7-17.

15. Fang, C.; Shi, B.; Pei, Y. Y.; Hong, M. H.; Wu, J.; Chen, H. Z. European Journal of Pharmaceutical

Sciences 2006, 27, 27-36.

16. Xie, J.; Xu, C.; Kohler, N.; Hou, Y.; Sun, S. Advanced Materials 2007, 19, 3163-3166.

17. Wang, B. D.; Xu, C. J.; Xie, J.; Yang, Z. Y.; Sun, S. L. Journal of the American Chemical Society

2008, 130, 14436.

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114  

18. Kratz, F.; Beyer, U.; Schutte, M. T. Critical Reviews in Therapeutic Drug Carrier Systems 1999, 16,

245-288.

19. Saito, H.; Hoffman, A. S.; Ogawa, H. I. Journal of Bioactive and Compatible Polymers 2007, 22, 589-

601.

20. Xu, C. J.; Xie, J.; Kohler, N.; Walsh, E. G.; Chin, Y. E.; Sun, S. H. Chemistry-an Asian Journal 2008,

3, 548-552.

21. Krapcho, A. P.; Kuell, C. S. Synthetic Communications 1990, 20, 2559 - 2564.

  

Chapter V

Conjugating Methotrexate to Magnetite (Fe3O4) Nanoparticles

via Trichloro-s-Triazine for Cancer Inhibition

In the previous two chapters, I’ve successfully demonstrated how to modify Fe3O4

nanoparticles (NPs) with dopamine and bifunctional poly(ethylene glycol). The

functionalization with nucleus localization signal or anti-cancer drug, chromone has

clearly revealed the possibility of using this chemistry for the biomedical application of

Fe3O4 NPs. However, bifunctional PEG is very expensive as I mentioned in chapter 2.

The synthesis (chapter 4) is laborious. In this chapter, I will discuss one of the efforts

we’ve made to use Trichloro-s-Triazine (TsT) for conjugating a chemotherapeutic drug,

methotrexate onto the Fe3O4 NPs through normal poly(ethylene glycol). The successful

application of TsT as a linker for functional molecules will

1. Background

Magnetic nanoparticles (NPs) with diameters below 20 nm exhibit interesting size-

dependent magnetic properties, including the phenomenon of superparamagnetism.1,2

Superparamagnetic magnetite NPs are ideal candidates for biomedical applications

because they are not subject to strong magnetic interactions between one another in the

dispersion state, facilitating their long-term stability in biological systems, and they

generate a large magnetic signal under an external magnetic field.3 Due their low

toxicity4 and stable magnetic properties, magnetite (Fe3O4) NPs have been investigated

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for potential applications in bio-separation, bio-sensing, drug delivery, magnetic fluid

hyperthermia, and magnetic resonance imaging (MRI) contrast enhancement.3,5-7

Linking hydrophilic macromolecules, especially biomolecules, to Fe3O4 NPs is a vital

step for producing water-based NPs to use in the abovementioned applications.5 The

synthesis procedures for monodisperse Fe3O4 NPs often result in a hydrocarbon-based

capping. As a result, the as-synthesized NPs are only soluble in non-polar or weakly polar

organic solvents.8 Existing linker chemistries to conjugate macromolecules to Fe3O4 NPs

are hard to work with and often not economically available.5 Thus two of the major

challenges in this field include (i) the ability to make high quality monodisperse water-

soluble Fe3O4 NPs and (ii) the lack of readily available linker chemistry that provides an

easy and inexpensive way of attaching functional molecules to Fe3O4 NPs. Previously

we reported an approach to conjugate Fe3O4 NPs with poly(ethylene glycol) (PEG)-based

hydrophilic macromolecules via the organic linker trichloro-s-triazine (TsT).5 Building

upon our previous work, we created a new inexpensive linking chemistry based on PEG

and TsT, allowing for the facile conjugation of biomolecules to Fe3O4 NPs.

The proposed ligand combination layer consists of dopamine (DA), PEG, and two

trichloro-s-triazine (TsT) molecules, as shown in Figure 5-1. DA has the ability to replace

the original capping ligand, olelyamine, on the Fe3O4 NPs surface and has been shown to

serve as a robust anchor on the surface of Fe3O4 NPs. Spectroscopic studies suggest that

catechol acts as a chelating agent, forming tight bonds with iron oxides by converting the

under-coordinated cationic iron surface sites to a bulk-like lattice structure.9 PEG is a

hydrophilic and biocompatible polymer often used to increase the stability of

nanomaterials in biological systems. It is commonly regarded as a non-specific

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interaction reducing agent and has been widely used to extend the circulation time of

proteins and nanomaterials in vivo as it can prevent aggregation and absorption by the

reticulo-endothelial system (RES) of the body.10 PEG chains are commercially available

in many molecular weights and with various functional groups. TsT, a symmetrical

heterocyclic compound, is a readily available organic linker molecule containing three

acyl-like chlorines which show different reactivities toward nucleophiles in aqueous

solution.5,11 The functionality provided by the second TsT molecule in the ligand allows

for the conjugation of various functional molecules to the NPs.

O

O NH

N

N

N

Cl

O O

N

N

N

Cl

ClOn

O

O NH

N

N

N

Cl

O O

N

N

N

Cl

On N

HNH2

OO

HN

NN

N

O ON

NN

Cl

On

HN N

H

N

N NN

NH2

NH2

NHN

O

OOH

O

Cl

Methotrexate

H2NNH2

O

O NH2

N

N

N

Cl

Cl O O

N

N

N

Cl

ClO

n

(a) (b)

(c)

(d)

 

Figure 5-1. Modification of Fe3O4 NPs MTX via TsT.

The chemotherapy drug, Methotrexate (MTX) was chosen to conjugate to Fe3O4 NPs

with the DA, PEG, and TsT-based ligand (Figure 5-1). MTX is an analogue of folic acid,

which blocks folate receptors from folic acid and inhibits dihydrofolate reductase

(DHFR), a critical enzyme in the folic acid cycle and key to regulating homeostasis,

leading to reduced cell viability and cell death. As folate receptors are overexpressed on

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the cell membranes of many types of cancer cells, MTX has proven to be an effective

targeting agent.12-14 MTX is one of the most widely used drugs for the treatment of many

forms of cancer, including tumors of the brain, breast, ovary, and several leukemias.

However, the clinical application of this drug is limited by its low solubility, short half-

life in the bloodstream and rapid diffusion throughout the body.13

The primary objectives of the present studies are (i) to present a newly designed

simple linker chemistry that makes the Fe3O4 NPs soluble in an aqueous environment and

allows for the conjugation of functional biomolecules, (ii) to demonstrate that the ligand

is stable under physiological conditions, and (iii) to demonstrate that MTX maintains its

anti-tumor activity after conjugation to the NPs. The conjugation of MTX to Fe3O4 NPs

provides the potential for a multifunctional entity which can take advantage of the

superparamagnetic nature of the NP core for imaging purposes while relying on the MTX

to be the targeting and therapeutic agent.

2. Results and discussion

Fe3O4 NPs were synthesized according to a previously reported method.5 The TEM

image in Figure 5-2a shows that as synthesized Fe3O4 NPs are nearly monodisperse with

an average diameter around 8 nm. The NPs were coated with a hydrophobic layer of

oleylamine. To functionalize these NPs with DA-PEG-TsT as illustrated in Figure 5-1,

the oleylamine ligand is replaced by DA through a ligand exchange reaction, resulting in

DA–capped NPs (a). The TsT-PEG-TsT precursor reacts with the amine group of

dopamine to form TsT-PEG-TsT-DA capped NPs (b). By controlling the ratio of TsT-

PEG-TsT and dopamine added, one end of TsT-PEG-TsT reacts with one DA-capped NP,

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leaving the other TsT end free for the further functionalization. Then ethylene diamine is

added to provide an amine group to the free TsT for further functionalization (c). Once

two of the chlorines of TsT have been coupled to nucleophilic groups, it is very difficult

for the third one to react.1 Thus only one ethylene diamine will react with each TsT.

Finally MTX was conjugated with NH2-terminated NPs through EDC/NHS chemistry. A

TEM image of the final MTX-conjugated NPs, shown in Figure 5-2b, indicates that the

NPs do not change in size or morphology during the modification steps.

Figure 5-2. TEM images of (a) oleylamine coated 8 nm Fe3O4 NPs in hexane and (b) MTX-conjugated Fe3O4 NPs in water.

The MTX-Fe3O4 NPs were analyzed by UV-Visible Spectroscopy and MALDI Mass

Spectrometry to confirm that MTX was successfully conjugated to the surface of the

Fe3O4 NPs. Figure 5-3 is the UV-Visible spectra of the MTX-conjugated NPs, the NH2-

terminated NPs, and an aqueous solution of free MTX. MTX has a characteristic UV

absorbance at 304 nm.13 The NH2-terminated NPs do not have such an absorbance peak.

The MTX-conjugated NPs show the characteristic absorbance peak at 304 nm,

confirming the presence of MTX on the NP surface. The MALDI spectra gave an average

molecular weight of 6,224 for the NH2-terminated NPs and 6,590 for the MTX-

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conjugated NPs, with a difference of 366 between the two. The majority of the molecular

weight is attributed to the PEG chain, which has an average molecular weight of 6,000.

Due to the variability in the weight of PEG chains, the difference of 366 provides further

evidence that MTX is attached to the NPs.

Figure 5-3. UV-Visible absorption spectra of free MTX, NH2-terminated NPs, and MTX-conjugated NPs in water. (MTX has a characteristic absorbance at 304 nm)

The stability of the MTX-NPs was tested in a PBS buffer solution with 10% FBS

over the course of 72 hours at an incubation temperature of 37°C to mimic physiological

conditions. Dynamic light scattering (DLS) was used to track the size change of the NPs

during the incubation. The DLS stability data for both the NH2-terminated and MTX-

conjugated NPs is shown in Figure 5-4. The hydrodynamic diameter, including the Fe3O4

core and the ligand coating, of both NPs started at 37 nm and leveled out at about 50 nm

after 24 hours, indicating very little particle aggregation/sintering. The slight increase in

size was most likely due to adsorption of FBS molecules onto the NPs. In the DLS

experiments, the NH2-terminated NPs were measured to have slightly smaller

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hydrodynamic diameters compared with the MTX-conjugated NPs, further supporting the

conjugation of MTX.

Figure 5-4. Hydrodynamic diameter of NH2–terminated NPs and MTX-conjugated NPs under physiological conditions over the course of 72 hours measured by DLS.

To demonstrate that MTX retained its anti-tumor activity after conjugation to the

Fe3O4 NPs, the conjugates were incubated with 9L rat glioma cells, a type of robust

tumor cell with a high metabolic activity leading to the overexpression of the folate

receptor on the cell surface.12,13 The toxicity of the MTX-conjugated NPs was compared

to the toxicity of the NH2-terminated NPs with equivalent iron concentrations as

determined by ICP, and the toxicity of free MTX at concentrations equal to those of the

MTX-conjugated NPs as determined by UV-Vis. The Fe3O4 NPs were incubated with the

9L cells at two different iron concentrations (0.01 mg/ml and 0.005 mg/ml) for 48 hours

and the live cells were counted to determine the cell viability. The cells were counted

using a hemocytometer because our data indicated that commercial cytoxicity assays

were inaccurate in assessing the viability of 9L cells in the presence of MTX. The 9L cell

viability data, shown in Figure 5-5a, indicates dose-dependent toxicity with the highest

concentration of iron and the highest concentration of MTX being the most toxic. The

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MTX-conjugated NPs killed 60% and 70% of the 9L cells at iron concentrations of 0.005

mg/mL and 0.01 mg/mL respectively over the course of 48 hours. This is in stark contrast

to the NH2-terminated NPs, which were observed to be non-toxic to the 9L cells, killing

less than 10% at an iron concentration of 0.01 mg/mL over 48 hours. This is in agreement

with the literature.4 Interestingly, the MTX conjugated Fe3O4 NPs was observed to be

slightly less toxic than the equivalent concentration of free MTX drug. This is likely

caused by the uptake difference as the bulkier MTX-Fe3O4 NPs may not be taken up as

easily as the free MTX at the same concentration, or by the fact that the MTX may need

to be cleaved from the NPs before it shows the toxicity.

Figure 5-5. 9L (a) and CPAE (b) cell viability data for MTX-conjugated NPs, NH2-terminated NPs of equivalent iron concentrations, and aqueous solutions of MTX corresponding to the concentration of MTX in the MTX-conjugated NPs samples. Iron concentrations of 0.005 mg/mL and 0.01 mg/mL were tested.

To demonstrate the specificity of the MTX-conjugated Fe3O4 NPs for cancer cells,

the cell viability experiments were repeated with CPAE (pulmonary artery endothelial

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cells) cells. The CPAE cell viability data, shown in Figure 5-5b, indicates that the MTX-

conjugated NPs were much less effective in reducing cell viability in CPAE cells

compared with 9L cells. At each iron concentration tested, less than 30% of the CPAE

cells died when incubated with MTX-conjugated NPs for a 48 hour period. The solutions

of free MTX showed similar results. Similar to the 9L cells, the NH2-terminated NPs

were nontoxic. The viability experiments with 9L cells and CPAE cells reinforce the

validity of the intracellular trafficking model proposed by Kohler et al. in which MTX

internalizes via the folate receptor and reduced folate carrier.13 Since 9L cells overexpress

the folate receptor and CPAE cells do not, free MTX and MTX conjugated to Fe3O4 NPs

can more efficiently internalize and cause reduced cellular viability in 9L cells. Thus the

data in Figure 5-5 demonstrates the potential for the MTX-conjugated NPs to be used as a

targeting agent as well as a cytotoxic entity.

Figure 5-6. Intracellular uptake of MTX-conjugated NPs and NH2-terminated NPs in 9L and CPAE cells for iron concentrations of 0.005 mg/mL and 0.01 mg/mL after 4 hours of incubation. The black bars represent 9L cells and the grey bars represent CPAE cells.

The targeting specificity of the MTX-conjugated NPs was further investigated

through cellular uptake studies of the Fe3O4 NPs conjugated with MTX compared to

those without MTX. The intracellular iron uptake for 9L and CPAE cells incubated with

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solutions of 0.01 and 0.005 mg Fe/ml MTX-conjugated NPs and NH2-terminated NPs is

shown in Figure 5-6. The uptake data demonstrates the specificity of the MTX-NPs for a

cancer cell line over a healthy cell line. At an Fe concentration of 0.01 mg/ ml, the

uptake of the MTX-conjugated NPs into 9L cells is almost twice that of the uptake of the

NH2-terminated NPs. At an Fe concentration of 0.005 mg/ml, the uptake of the MTX-

conjugated particles into 9L cells is four times that of the NH2-terminated particles. The

uptake data for the CPAE cells does not show a preference for uptake of either type of

NP, but instead the NPs seem to be nonspecifically internalized. The uptake data in

Figure 5-6 supports the trends seen in the cell viability data in Figure 5-5. The MTX-

conjugated NPs target the 9L cells resulting in more particles being internalized, thus

leading to a reduced cell viability. However, it must not be ruled out that the difference in

uptake between the 9L and CPAE cells could partially be due to differences in metabolic

activity of the two cell types.13 The cell viability experiments combined with the uptake

studies provide evidence that our newly developed linker chemistry does not alter the

biological activity of the conjugated drug.

To visualize the location of the NPs inside the cells after internalization, the green

fluorescent probe Fluorescein Isothiocynate (FITC) was used to label the MTX-Fe3O4

NPs. Because MTX possesses primary amine groups, FITC was conjugated to the NH2-

terminated NPs before MTX was conjugated to the particles to prevent FITC from

binding to MTX and affecting its ability to reduce cell viability. A small amount of FITC

was added to block approximately 10% of the NH2 group on NPs surface so that the

remaining ligands were available to attach to MTX. NPs conjugated with MTX and

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FITC were incubated with 9L cells and imaged over the course of 120 minutes using a

fluorescence microscope.

Figure 5-7. Fluorescence images of 9L cells transfected with Rab5 to dye the early/sorting endosomes red. The transfected cells were then incubated with MTX&FITC-conjugated NPs for 15 minutes (column A), 30 minutes (column B), and 60 minutes (column C). The first row (1) shows the green channel of the fluorescence image. This green fluorescence is due to the FITC conjugated to the NPs. The second row (2) shows the red channel of the fluorescence image. This red fluorescence comes from the early/sorting endosomes that fluorescence red due to the Rab5. The third row (3) shows the overlap of the red and green channels. The color yellow indicates red and green overlap.

In our preliminary data the MTX-conjugated NPs were observed in cellular

components believed to be early endosomes following their uptake into 9L cells. To

confirm that the cellular compartments containing the NPs were early endosomes, the

early/sorting endosomal marker Rab5 was used to label the early endosomes.15 The 9L

cells were transfected with a Rab5 plasmid causing their early endosomes to fluoresce red.

These transfected 9L cells were incubated with NPs conjugated with MTX and FITC and

images were collected over the course of 120 minutes. In order to capture both the green

fluorescence of the FITC and the red fluorescence of the Rab5, one image of the cells

125  

  

was taken with the green channel of the fluorescence microscope and a corresponding

image of the same cells was taken with the red channel of the fluorescence microscope.

These images were then overlayed using software.

The color green indicates the location of the FITC-conjugated NPs and the color red

indicates the location of the early endosomes. The color yellow indicates overlap of the

NPs and the early endosomes. Figure 5-7 shows images of the transfected 9L cells

collected at 15, 30, and 60 minutes of incubation with the MTX-conjugated NPs. While it

would be ideal to monitor the same cells over the course of the experiment, this was not

possible due to the fact that the cells has to be fixed with paraformaldehyde in order to be

imaged. Thus cells that were representative of the entire sample were chosen to be

imaged. Focusing on the overlay images in the third row of Figure 5-7, after 15 minutes

of incubation the NPs, indicated by green, are in the cytoplasm and some are in the early

endosomes, indicated by yellow. After 30 minutes, the majority of the particles are in the

early endosomes as yellow is the main color observable. After 60 minutes, the large

amount of red fluorescence from empty endosomes suggests that the NPs are no longer in

the early endosomes. This experiment provides further evidence to support the first step,

where NPs are transported to early endosomes following their uptake, of the intercellular

trafficking model proposed by Kohler et al.13

3. Summary

Monodisperse magnetite NPs have been conjugated with the anticancer drug

Methotrexate using a new linker trichloro-s-triazine (TsT). UV-Vis spectroscopy and

MALDI mass spectrometry were used to confirm that MTX was immobilized on the NP

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surface. Both the NH2-terminated and MTX-conjugated Fe3O4 NPs were found to be

stable under physiological conditions. The MTX-Fe3O4 NPs showed specificity and high

toxicity to 9L rat glioma cells rather than to CPAE cells. Finally, the intracellular

trafficking for MTX-conjugated NPs was investigated by attaching the fluorescent probe

FITC to the Fe3O4 NPs and visualizing the particles in 9L cells transfected with a Rab5

plasmid under a fluorescence microscope. These fluorescence experiments confirmed

that the particles do indeed enter the early/sorting endosomes following their uptake into

target cells. We hope to further extend this system to conjugate other types of

biomolecules, such as peptides, proteins, and DNAs to Fe3O4 NPs for biomedical

applications.

4. Experimental

Materials and Instruments: Reagents were purchased from Sigma-Aldrich and solvents

were purchased from Mallinckrodt Chemicals (Phillipsburg, NJ) unless otherwise noted.

NPs were imaged with transmission electron microscopy (TEM, Philips-EM 20) at 120 V.

UV-Visible spectra were measured with a Hewlett-Packard 8452 Diode Array

Spectrometer. Molecular weights were measured with matrix-assisted laser

desorption/ionization (MALDI) mass spectrometry (Voyager DE Pro, Applied

Biosystems). Iron concentrations were identified with inductively coupled plasma atomic

emission spectroscopy (ICP-AES, JY 2000).

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Modification of Fe3O4 NPs: Monodisperse Fe3O4 NPs (8 nm in diameter) were

synthesized by a previously reported one-pot high temperature method and stored in

hexane.16 The Fe3O4 NPs were conjugated with MTX following the reaction sequence

outlined in Figure 5-1. To prepare the TsT-PEG-TsT precursor, polyethylene glycol

(PEG, MW = 6000) was activated using trichloro-s-triazine by following a protocol based

on those of Abuchowski et al. and Gotoh et al.11 To modify the capping ligand on the

Fe3O4 NPs, 100 mg of dopamine hydrochloride (DA) dispersed in 2 mL N,N-

dimethylformamide (DMF) was added to 800 μl of the oleylamine-capped 8 nm Fe3O4

NPs dispersed in 2 ml chloroform (CHCl3) and sonicated for 30 minutes. Then 5 mL of

hexane was added the mixture and a permanent magnet was used to separate the magnetic

NPs out of solution. After repeating the wash step two more times, the DA-capped NPs

were redispersed in a 1:1 mixture of DMF and CHCl3. Next, 130 mg of the TsT-PEG-

TsT precursor and 10 mg of sodium carbonate were added and stirred for 24 hours. The

TsT-PEG-TsT-DA-capped NPs were separated from the excess TsT-PEG-TsT precursor

as above and redispersed in a 2:1 mixture of DMF and CHCl3. 150 μl of ethylene diamine

(NH2-C2H2-NH2) was added and the mixture was stirred for 24 hours. The NH2-TsT-

PEG-TsT-DA-capped nanoparticles were purified as above and dried under a gentle

stream of nitrogen. The modified Fe3O4 NPs were redispersed in deionized water. To

remove any remaining excess ethylene diamine, the NH2-terminated NPs were dialyzed

for 24 hours using molecular porous membrane tubing (MWCO = 12-14,000, Spectrum

Laboratories, CA).

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Conjugation of Methotrexate to NH2-terminated NPs: Methotrexate (MTX) was

conjugated to the amine-terminated NPs through NHS/EDC coupling chemistry. The

MTX conjugation reaction may occur through either the α or β carboxylic acid groups on

the glutamic acid residue.13 To remove the excess MTX, the NPs were separated with a

permanent magnet and washed with deionized water. The samples were further purified

using Centriprep 15 ml centrifugal filter devices (Amicon Bioseparations- Millipore).

The final MTX-conjugated NPs were stored in a 20 mL glass vial covered in aluminum

foil to avoid light exposure.

Characterization of MTX-conjugated NPs: To determine the concentration of MTX in

the MTX-conjugated NPs sample, a standard linear-fit curve of free MTX in water was

created by plotting the UV-Vis absorbance at 304 nm of several MTX solutions of known

concentrations. The concentration of MTX in the MTX-conjugated particles was then

determined by subtracting out the background absorbance of a sample of NH2-terminated

NPs with a normalized amount of iron at 304 nm. The accuracy of this method was

confirmed using the method developed by Kohler et al.12

Stability under physiological conditions: To test the stability of both the NH2-

terminated NPs and the MTX-conjugated NPs, the particles were placed in a PBS

(Dulbecco’s Phosphate Buffered Saline, Atlanta Biologicals) solution with 10% fetal

bovine serum (FBS, Atlanta Biologicals) and kept in an incubator at 37°C and 5% CO2.

The hydrodynamic diameters of NPs were measured using a Dynamic Light Scattering

(Malvern Instruments, Zetasizer Nano Series, Nano-S90) instrument over the course of

129  

  

72 hours. Data was collected for the hydrodynamic diameter of the particles at t = 0, 1, 2,

4, 8, 16, 24, 48, and 72 hours.

Cell culture: 9L rat glioma cells with the overexpression of the folate receptor and

mammalian cultured pulmonary artery endothelial (CPAE) cells with little folate receptor

were chosen to examine the targeting ability of MTX conjugated NPs. 9L cells and

CPAE cells were grown in 75 cm2 canted neck polystyrene culture flasks (Corning) in an

incubator kept at 37°C and 5% CO2. The medium used for culture was DMEM

(Dulbecco’s Modified Eagle Medium with D-glucose, L-glutamine, and sodium pyruvate,

Atlanta Biologicals) with 10 % FBS and 5% antibiotics. The cells were allowed to grow

to 80-90% confluency before they were split or harvested for experimental purposes.

Cell viability studies: 1X105 9L or CPAE cells were seeded into 25 cm2 polystyrene

culture flasks (Corning) containing 3 mL of DMEM and placed in an incubator for 24

hours. Solutions of MTX-conjugated NPs and NH2-terminated NPs with iron

concentrations of 0.01 and 0.005 mg Fe/mL were incubated with the cells for 48 hours.

The flasks were washed twice with PBS to remove any dead cells and 1 mL of 0.25%

Trypsin 1X (Gibco) was added. After collecting the cells from the flask, the cells were

counted using a hemacytometer (Hausser Scientific). As a control, cells were incubated

with DMEM. Cells were also incubated with MTX drug solutions with concentrations

corresponding to those in the MTX-conjugated NP samples. Each iron concentration was

repeated three times and the cells were counted three times. The average percentage of

living cells was calculated by comparison with the control.

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Intracellular uptake: The intracellular uptake of the MTX-conjugated NPs and the NH2-

terminated NPs was quantified by measuring the concentration of iron within the cells for

each sample using ICP-AES. 1.5x104 cells were seeded into 25 cm2 polystyrene culture

flasks (Corning) containing 3 mL of DMEM and placed in an incubator for 24 hours.

Solutions of MTX-conjugated NPs and NH2-terminated NPs with iron concentrations of

0.01 and 0.005 mg/mL were incubated with the cells for 4 hours. The NP solutions were

then removed from the flasks and 1 mL of 0.25% Trypsin 1X was added to detach the

cells. The cells were counted using the hemacytometer. After counting, the cell

suspensions were placed in 1.5 mL Eppendorf tubes and centrifuged at 10,000 RPM for 5

minutes. The supernatant was removed and 4-5 drops of concentrated HNO3 was added

to break down the cells. The iron uptake (pg per cell) was calculated by dividing the total

amount of iron by the total number of cells in each sample.

Tagging NPs with fluorescein isothiocynate (FITC): NH2-NPs were dispersed in a 0.1

M Na2CO3/NaHCO3 buffer with pH = 9. The total number of ligands on the surface of

each particle was estimated and the amount of FITC necessary to block 10% of these

ligands (2.177×10-7g/mL) was dissolved in DMSO. The FITC solution was then added to

the NP solution dropwise and stirred overnight under aluminum foil to prevent bleaching

of the FITC.11 Excess FITC was removed by using molecular porous membrane tubing

(MWCO = 12- 14,000). MTX was then conjugated to the FITC-conjugated NPs using the

procedure described previously.

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Particle internalization study: 9L cells were seeded in five small glass bottom culture

dishes (MatTek Corporation, MA) for 24 hours. Then 200 μL of DMEM solutions of

MTX&FITC-conjugated NPs with a concentration of 0.005 mg Fe/mL were added to

each dish and placed in the incubator. Over the course of 60 minutes the dishes were

removed from the incubator, washed 3 times with PBS, and the cells were fixed with 4%

paraformaldehyde in PBS. The samples were then imaged using a Nikon Eclipse

TE2000-U Fluorescence Microscope. The acquisition time was 5 ms for the bright field

and 500 ms for the fluorescence field.

The early/sorting endosomal marker Rab5 was used to further investigate the path of

MTX-conjugated NPs in 9L cells.15 A Rab5 plasmid was transfected into the 9L cells and

as a result the early/sorting endosomes within the 9L cells fluoresced red under a

fluorescence microscope. 9L cells were cultured in a Petri dish in DMEM medium. The

DMEM was removed and the cells were incubated with 1 ml of OPTI-MEM for 40

minutes. To transfect the cells, a solution containing 6 μl of the plasmid, 24μL of the

commercial transfection reagent Lipofectamine 2000 (Invitrogen), and 1.170 mL of

OPTI-MEM was incubated with the cells for 4 hours. The OPTI-MEM was then removed

and 2 mL of DMEM was added. After overnight culturing, the cells were incubated with

NPs and the imaging study described above was repeated.

References:

1. Jun, Y. W.; Seo, J. W.; Cheon, A. Accounts of Chemical Research 2008, 41, 179-189.

2. Xie, J.; Sun, S. H. Nanomaterials: Inorganic and Bioinorganic Perspectives- Encyclopedia in

Inorganic Chemistry; John Wiley & Sons, Ltd, 2008.

3. Xu, C. J.; Sun, S. H. Polymer International 2007, 56, 821-826.

4. Lewinski, N.; Colvin, V.; Drezek, R. Small 2008, 4, 26-49.

132  

  

133  

5. Xie, J.; Xu, C. J.; Xu, Z. C.; Hou, Y. L.; Young, K. L.; Wang, S. X.; Pourmond, N.; Sun, S. H.

Chemistry of Materials 2006, 18, 5401-5403.

6. Pankhurst, Q. A.; Connolly, J.; Jones, S. K.; Dobson, J. Journal of Physics D: Applied Physics 2003,

R167.

7. McNeil, S. E. J Leukoc Biol 2005, 78, 585-594.

8. Sun, S. H.; Zeng, H. Journal of the American Chemical Society 2002, 124, 8204-8205.

9. Xu, C. J.; Xu, K. M.; Gu, H. W.; Zheng, R. K.; Liu, H.; Zhang, X. X.; Guo, Z. H.; Xu, B. Journal of

the American Chemical Society 2004, 126, 9938-9939.

10. Xie, J.; Xu, C.; Kohler, N.; Hou, Y.; Sun, S. Advanced Materials 2007, 19, 3163-3166.

11. Hermanson, G. T. Bioconjugate Techniques; Academic Press: New York, 1996.

12. Kohler, N.; Sun, C.; Fichtenholtz, A.; Gunn, J.; Fang, C.; Zhang, M. Small 2006, 2, 785-792.

13. Kohler, N.; Sun, C.; Wang, J.; Zhang, M. Langmuir 2005, 21, 8858-8864.

14. Dhar, S.; Liu, Z.; Thomale, J.; Dai, H.; Lippard, S. J. Journal of the American Chemical Society 2008,

130, 11467-11476.

15. Elena S. Suvorova, J. M. G. H. M. M. Traffic 2005, 6, 100-115.

16. Zeng, H.; Li, J.; Liu, J. P.; Wang, Z. L.; Sun, S. H. Nature 2002, 420, 395-398.

 

Chapter VI

Au-Fe3O4 Dumbbell NPs as Dual-functional Probes

This chapter reports that through proper surface modification, the dumbbell-like

Au-Fe3O4 nanoparticles can be made biocompatible and suitable for A431 (human

epithelial carcinoma cell line) cell attachment. The particles are magnetically and

optically active and are useful for simultaneous magnetic and optical detection. The

T2 relaxivity (r2) of the 8 nm - 20 nm Au-Fe3O4 nanoparticles around the A431 cells is

80.4 s-1·mM-1 (r2/r1 = 37.1) and their optical detection limit reaches 90 pM Au. The

fact that the dumbbell nanoparticles are capable of imaging the exact same tissue area

through both magnetic resonance imaging (MRI) and an optical microscope implies

that they can be used to achieve high sensitivity in diagnostic imaging and therapeutic

applications.

1. Background

Synthesis of dumbbell-shaped nanoparticles containing different functionalities has

attracted much attention recently.1-6 In such a dumbbell structure, one nanoparticle is

linked to another, and electronic communication across the junction can drastically

change the local electronic structure, leading to an additional dimension of control in

catalytic, magnetic, and optical properties.7-9 Moreover, the dumbbell structure offers

two functional surfaces for the attachment of different kinds of molecules, making

such species especially attractive as multifunctional probes for diagnostic and

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therapeutic applications.10,11

The success of conjugating Fe3O4 NPs with peptide to achieve specific targeting

has encouraged us to further expand this DPA-PEG-COOH based modification

method to composites nanoparticles such as Au-Fe3O4 dumbbell nanoparticles. These

particles contain Au and Fe3O4 composites, both of which are known as

biocompatible materials and have been used extensively in biomedicine for their

optical and magnetic properties.12-16 Compared with the individual Au or Fe3O4

nanoparticles, the dumbbell-like Au-Fe3O4 system has distinct advantages in: 1) The

structure contains both a magnetic (Fe3O4) and an optically active plasmonic (Au) unit

therefore being suitable for simultaneous optical and magnetic detection. 2) The

presence of Fe3O4 and Au surfaces facilitates the attachment of different chemical

functionalities for target-specific imaging and delivery purposes. 3) The size of either

of the two nanoparticles can be controlled to optimize magnetic and optical properties,

and the small particle is only capable of accommodating a few DNA strands, proteins,

antibodies, or therapeutic molecules, thus facilitating kinetic studies in cell targeting

and drug release. In this section, we will demonstrate the dumbbell Au-Fe3O4

nanoparticles can be made biocompatible and used as magnetic and optical dual

functional probes for cell imaging.

135

2. Results and discussion

Figure 6-1. (a) Schematic illustration of surface functionalization of the Au-Fe3O4 nanoparticles. (b, c) TEM images of the 8-20 nm Au-Fe3O4 particles before (b) and after (c) surface modification.

Figure 6-1a illustrates the structure of the functionalized dumbbell-like 8-20 nm (core

particle diameter) Au-Fe3O4 nanoparticles used in this study. The dumbbell

nanoparticles were synthesized by decomposing iron pentacarbonyl on the surface of

pre-made Au nanoparticles in the presence of oleic acid and oleylamine, as described

previously.1 The hydrophobic coating of the as-synthesized nanoparticles was later

taken placed by DPA-PEG-COOH and SH-PEG-NH2, on Fe3O4 and Au surface,

respectively (Figure 6-1a). The morphology of NPs before and after surface change

was shown in Figure 6-1b&c. For control purposes, 8-nm Au, 20-nm Fe3O4, and 3-20

nm Au-Fe3O4 nanoparticles (Figure 6-2) were also prepared and modified by the same

manner.

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Figure 6-2. TEM images of (a) 8nm Au nanoparticles; (b) 20nm Fe3O4 nanoparticles; (c) 3 nm-20 nm Au-Fe3O4 nanoparticles.

Epidermal growth factor receptor antibody (EGFRA) was linked to the Fe3O4

surface through EDC/NHS coupling. Such antibody is capable of recognizing and

associating with EGFR that was found extensively expressed on many cancer cell

lines. The Au surface was passivated with HS-PEG-NH2 (Mr=2204) by thiol attaching.

Such treatment is to avoid particles’ nonspecific hydrophobic interactions with

proteins. Meanwhile, it allows the possibility of attaching other species onto the Au

surface (although we did not do that in this study). The functionalized nanoparticles

(Figure 6-1a) were characterized by matrix-assisted laser desorption/ionization

(MALDI) mass spectrometry (Figure 6-3), which proves the successful conjugation of

two kinds PEG and EGFR antibody (inset).

137

Figure 6-3. MALDI mass spectra of PEG2000-Au-Fe3O4-PEG3000-EGFRA (Mr = 2245, 3738, 150K).

These modified dumbbell nanoparticles are stable against aggregation in

phosphate buffered saline (PBS) or PBS containing 10% fetal bovine serum (FBS) at

37˚C during our test interval (12 h), as evidenced by their unchanged hydrodynamic

sizes (Figure 6-4). It is worth noting that there is slight size increase for the

EGFRA-DBNPs in PBS with 10% FBS, which is presumably due to the interaction

between the proteins and the nanoparticle surface. Transmission electron microscope

(TEM) images of the dumbbell nanoparticles showed a slight size reduction of Fe3O4

moiety after surface modification (Figure 6-1c). This effect is likely caused by the

corrosion by the catechol segment during the surfactant exchange process.

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Figure 6-4. Hydrodynamic sizes of the nanoparticles shown in Figure 6-1a measured by dynamic light scattering (DLS).

Magnetic measurements show that the nanoparticles are superparamagnetic at

room temperature before and after surface modification (Figure 6-5a).

Figure 6-5. (a) Magnetic hysteresis loops of the dumbbell nanoparticles before and after surface modification. The reduction of saturation magnetization is due largely to the weight contribution from the nonmagnetic Au particles. (b) Reflection spectra of 20-nm Fe3O4, 8-nm Au, 3-20 nm Au-Fe3O4, and 8-20 nm Au-Fe3O4 nanoparticles.

The nanoparticles also exhibit a plasmonic absorption in PBS at 525 nm for 8-nm

Au nanoparticles and at 530 nm for 8-20 nm Au-Fe3O4 dumbbell nanoparticles

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(Figure 6-6). The slight red shift is due to the junction effect in the dumbbell

structure.1

Figure 6-6. UV-vis spectra of Au and Au-Fe3O4 nanoparticles in water.

More interestingly, self-assembled nanoparticle on an aluminum substrate coated

with Teflon S (Boyd Coatings Research Co., Inc; the coating makes the reflection of

the substrate less than 5%) exhibit characteristic reflectance in the 590-650 nm range.

Figure 6-5b shows the reflectance spectra of 8-nm Au, 20-nm Fe3O4 nanoparticles as

well as 3-20 nm and 8-20 nm Au-Fe3O4 dumbbell nanoparticles. The relatively weak

reflectance from the dumbbell particles is likely caused by the dilution effect due to

the Fe3O4 presence. For comparison, Fe3O4 nanoparticles alone have no reflectance in

the same wavelength region. These magnetic and optical studies suggest that the

dumbbell nanoparticles are both magnetically and optically active and could be served

as dual functional probes for bimolecular imaging.

As an initial in vitro test, we demonstrate that the dumbbell nanoparticles can

specifically target to A431 (human epithelial carcinoma cell line, which is known to

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overexpression EGFR17,18) cell membrane and be imaged optically and magnetically.

Such study is meaningful in that, it can potentially be used for cancer diagnosis and

therapies, as EGFR overexpress is usually associated with tumor growth, like breast

and lung tumors.19,20 Briefly, we incubated the EGFRA-dumbbell nanoparticles with

A431 wells in Dulbecco’s Modified Eagle’s Medium (DMEM) containing 10% FBS

for 1 h and subsequently washed the cells three times with PBS. The binding between

EGFR and EGFRA enabled the dumbbell nanoparticles to be populated on the surface

or within the cytoplasm of A431 cells. Magnetic resonance imaging (MRI) analyses

revealed that 20-nm Fe3O4 particles, Au-Fe3O4 dumbbell nanoparticles, and A431

cells labeled with 8-20 nm Au-Fe3O4 nanoparticles could all shorten the T2 relaxation

time of the water molecules, as shown in the MRI phantom images in Figure 6-7a.

The iron content in all samples was determined by inductively coupled plasma atomic

emission spectrometry (ICP-AES) and used for calculating relaxivities. Table 6-1

gives the relaxivity data of r1, r2, and r2/r1. The slight increase in r1 and reduction in r2

with the increase in size of the Au core seems to indicate a larger junction effect

(reduced magnetization) on the dumbbell structure. Furthermore, 8-20 nm Au-Fe3O4

nanoparticles attached to A431 cells show smaller T1 and T2 relaxivities than the

Fe3O4 nanoparticles alone. This behavior is similar to what has been observed in the

iron oxide nanoparticle monocyte system, i.e. cellular compartmentalization of the

nanoparticles reduces proton relaxivity.21

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Figure 6-7. (a) T2-weighted MRI images of i) 20-nm Fe3O4, ii) 3-20 nm Au-Fe3O4, iii) 8-20 nm Au-Fe3O4 nanoparticles, and iv) A431 cells labeled with 8-20 nm Au-Fe3O4 nanoparticles. (b) Reflection images of the A431 cells labeled with 8-20 nm Au-Fe3O4 nanoparticles. c, d) Images of A431 cells labeled with 8-20-nm dumbbell particles, floating in the medium before (c) and after (d) an external magnetic field was applied (field gradient in the sample area was in 500-100 G). The dashed circles denote individual cells; the numbers label the same cells in (c) and (d); the arrow and H indicate the direction of the applied magnetic field.

Table 6-1. Relaxivities r1 and r2 of Fe3O4 and Au-Fe3O4 nanoparticles with various Au core sizes for the same Fe3O4 size at 3T (T=25˚)

A431 cells labeled with 8-20 nm Au-Fe3O4 nanoparticles were visualized with a

scanning confocal microscope. The wavelength used for the image was 594 nm,

which is close to the strong reflectance of the nanoparticles (Figure 6-5b). The

142

detected signals from the dumbbell nanoparticles reflect helps depict the typical

morphology of epithelial cells under the attachment conditions (1 mm Au and 8.8 mm

Fe, Figure 6-7b) and is much stronger in the cell-cell interacting region, suggesting

that EGFRA is involved in cell gap junction.22 Interestingly, by applying an external

magnetic field, the migration of the dumbbell particle labeled A431 cells can be

manipulated which is successfully tracked by the optical microscope (Figure 6-7c&d).

Figure 6-8. (a) Reflection image of the labeled cells used to obtain Figure 3b after three days. (b) Detection-limit examination of the 8-20 nm Au-Fe3O4-EGFRA labeled A431 cells. (c) Reflection image of Fe3O4-labeled A431 cells. (d) Reflection image of Au-Fe3O4 labeling without EGFR antibody.

The sample used for obtaining Figure 6-7b was re-imaged after three days and

showed no signal loss (Figure 6-8a). This result is extremely important for long-term

tracking of the nanoparticles in cellular structures. The detection limit for the 8-20 nm

dumbbell is about 90 pm Au (Figure 6-8b), which is 104 times lower than the normal

detection concentration (Figure 6-7b or Figure 6-8a). In contrast, Fe3O4 nanoparticles

yield much weaker reflectance signals (Figure 6-8c). As a control, we incubated A431

cells and the 8-20 nm Au-Fe3O4 nanoparticles without EGFRA (Figure 6-8d) under

143

the same concentration as shown in Figure 6-8a. The much higher signal-to-noise

ratio than that in 4.15 A proves that the targeting was specific and was EGFRA

directed. It is worth noting that the modified particles show negligible toxicity to

A431 cells at 0.01 mgFe mL-1 and 0.004 mg Au mL-1 (Figure 6-9).

Figure 6-9. Viability of A431 Cells with PEG-Au-Fe3O4-EGFRA at different concentrations

3. Summary

This work presented herein demonstrates that through proper surface functionalization,

the novel dumbbell Au-Fe3O4 nanoparticles can be made biocompatible and suitable

for linking different functional molecules to either end of the structure. The

EGFRA-conjugated dumbbell nanoparticles show higher internalization by A431 cells

than those without EGFRA. The nanoparticles are magnetically and optically active

and are therefore useful for simultaneous magnetic and optical detection. The fact that

the dumbbell nanoparticles are capable of imaging the exact same tissue area through

both MRI and an optical source without the fast signal loss observed in the common

fluorescent labeling implies that they can be used to achieve high sensitivity in

144

diagnostic imaging applications. Besides targeting agents, we can as well attach

therapeutic molecules to these dumbbell nanoparticles for site specific drug delivery

purpose. Related work is under way.

4. Experimental

Materials and Instruments: α,ω-Bis(2-carboxyethyl)polyethylene glycol

(MW=3,000), O,O’-bis(2-aminoethyl) poly(ethylene glycol) 2000, dopamine

hydrochloride, and sodium carbonate were purchased from Sigma-Aldrich.

NeutriAvidin (NAv), N-hydroxysuccinimide (NHS), and

N-(3-dimethylaminopropyl)-N’-ethylcarbodiimide (EDC) hydrochloride and

4’,6-diamidino-2-phenylindole (DAPI) were obtained from Pierce Biotechnology. All

organic solvents were purchased from Sigma-Aldrich Corp. All the buffers and media

used were acquired from Invitrogen Corp. The water was purified by a Millipore

Milli-DI Water Purification System. Nano-sep 100k OMEGA was purchased from

Fisher. All the dialysis bags were purchased from Spectrum Laboratories, Inc.

Synthesis of Au NPs for Au-Fe3O4 preparation: 1.0 g HAuCl4 · (H2O)3 (2.5 mmol)

was added to 100 ml tetralin, followed by 10 ml oleylamine (30 mmol) to form a red

solution. The solution was then heated at 65°C for 5 hrs and then cooled to room

temperature. Ethanol was added to the solution, and gold particles were separated by

centrifugation, washed by ethanol, and then redispersed in hexane.

145

Synthesis of Au-Fe3O4 NPs using pre-made Au NPs: A solution of 1ml oleic acid (3

mmol) in 20 ml Octadecene was heated at 120°C for 20 min under a flow of N2. Then

under a blanket of N2, 0.15 ml Fe(CO)5 was injected to the solution. After 5 min of

stirring, 0.5 ml oleylamine was injected to the reaction mixture, followed by 2 ml 8

nm Au colloidal dispersion (ca. 20 mg Au). The solution was heated to reflux (ca.

310°C) for 45 min. After cooled down to room temperature, the particles were

separated by adding iso-propanol, centrifuged and redispersed into hexane.

Modification of Au-Fe3O4 NPs : For modification of both Fe3O4 and Au-Fe3O4

particles, PEG diacid (20 mg), NHS (2 mg), DCC (3 mg), and dopamine

hydrochloride (1.27 mg) were dissolved in a mixture of CHCl3 (2 mL), DMF (1 mL),

and anhydrous Na2CO3 (10 mg). The solution was stirred at room temperature for 2 h

before nanoparticles (5 mg) were added, and the resulting solution was stirred

overnight at room temperature under N2. The modified nanoparticles were

precipitated by adding hexane (5 mL), collected by a permanent magnet and dried

under N2. The particles were then dispersed in water or PBS. The extra surfactants

and other salts were removed by dialysis using a dialysis bag (MWCO 10000) for 24

h in PBS or water. Any precipitate was removed by a 200-nm syringe filter

(MillexGP). The final concentration of the particles was determined by ICP-AES

analysis. To link EGFR antibody and nanoparticles, a solution of nanoparticles (1

nmol) in PBS was mixed with 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC;

1 mmol) for 15 min. After addition of EGFRA (4–5 nmol), the solution was rocked

146

for 1 h and separated from the unattached antibody with Nanosep (PALL Life Science

Corp.). For Au-Fe3O4 nanoparticles, HS-PEG-NH2 was added after EGFRA was

connected. After stirring for 3 h, the conjugates were subjected to dialysis to remove

free HS-PEG-NH2. The nanoparticles were analyzed by MS to confirm the

modification.

Synthesis of HS-PEG-NH2:12-(Acetylthio)dodecanoic acid was prepared according

to Xu et al.23 The thiol-protected compound was then mixed with one equivalent

O,O’-bis(2-aminoethyl) polyethylene glycol 2000 under EDC catalysis. Later, the

protecting group was removed by reduction with hydrazine acetate. (MALDI MS: m/z

2204).

Cell experiments: A431 cells were purchased from ATCC and cultured in a

glass-bottom Petri dish (MatTek Corp.) with Dulbecco’s modified Eagle’s medium

(DMEM) with 10% FBS and 1% antibiotics. Before incubation with particles, the

cells were washed with PBS three times. The particle solution in DMEM was

incubated with cells for 1 h. Then, those cells were washed with PBS three times and

fixed by 4% paraformaldehyde solution. After 30 min fixation, the cells were again

washed three times with PBS and subjected to reflection imaging using a Leica TCS

SP2 AOBS spectral confocal microscope.

Cell viability test: Viability of cells with particles was examined through WST1

147

assay. This cell viability test is based on the cleavage of the tetrazolium salt WST-1

(4-[3-(4-iodophenyl)-2-(4-nitrophenyl)-2H-5-tetrazolio]-1,6-benzene disulfonate) by

mitochondrial dehydrogenases in metabolically active cells. The cells were seeded

onto 96-well culture plates at a density of 105 cells per well in DMEM (100 mL)

containing 10% FBS. After 24 h incubation at 37˚C, nanoparticles in DMEM buffer at

different concentrations were added. The particles were washed away after 48 h

incubation. Then WST-1 solution (10 mL, Invitrogen) was added to each well to

evaluate cell viability. After 4 h at 37˚C, cell viability was measured using a

microplate reader.

MRI experiments for Au-Fe3O4 nanoparticles: Transverse T2-weighted spin echo

images were acquired using a 3 T Siemens Tim Trio MR Scanner. Echo times were

11-132 ms in 11-ms steps with a repetition time of 2000 ms. T1-weighted imaging was

performed using inversion recovery with 10 inversion times ranging from 100 ms to

2000 ms with a repetition time of 3000 ms. Gel preparations in 2-mL vials were

placed in a holder for insertion into the eight-channel volume head resonator. The

long axis of the vials was parallel to the static magnetic field, and a transverse

tomographic plane orientation was used. A gradient echo acquisition was used with a

repetition time of 2000 ms, an echo time of 1.8 ms, a slice thickness of 12 mm, and a

flip angle of 20˚. In-plane resolution was 0.41 mm. The normal first-order shim

process was applied, and the phantoms were imaged at room temperature (20 ˚C). For

A431cell experiments, 18000 A431 cells with attached dumbbell nanoparticles were

148

mixed into 4% agarose gel at 40 ˚C before imaging.

Characterizations: Reflection spectra were acquired on a UV/Vis/NIR bidirectional

spectrometer in the reflectance experiment laboratory (RELAB) of Brown University.

The hysteresis loop was obtained at 300 K with a LakeShore 7400 VSM system.

UV/Vis absorption spectra were obtained with a PerkinElmer Lambda 35 UV/Vis

spectrometer. Mass spectrometry of the modified nanoparticles was performed on a

matrix-assisted laser desorption ionization (MALDI) system. Optical images of A431

cells were obtained by a Zeiss Leica inverted epifluorescence/reflectance laser

scanning confocal microscope. The TEM image was taken on a Philips EM 420

instrument (120 kV). The hydrodynamic diameters of the nanoparticles were

measured using a Malvern Zeta Sizer Nano S-90 dynamic light scattering (DLS)

instrument.

References:

1. Yu, H.; Chen, M.; Rice, P. M.; Wang, S. X.; White, R. L.; Sun, S. H. Nano Letters 2005, 5,

379-382.

2. Gu, H. W.; Zheng, R. K.; Zhang, X. X.; Xu, B. Journal of the American Chemical Society 2004,

126, 5664-5665.

3. Teranishi, T.; Inoue, Y.; Nakaya, M.; Oumi, Y.; Sano, T. Journal of the American Chemical

Society 2004, 126, 9914-9915.

4. Shi, W. L.; Zeng, H.; Sahoo, Y.; Ohulchanskyy, T. Y.; Ding, Y.; Wang, Z. L.; Swihart, M.; Prasad,

P. N. Nano Letters 2006, 6, 875-881.

5. Yang, J.; Elim, H. I.; Zhang, Q. B.; Lee, J. Y.; Ji, W. Journal of the American Chemical Society

2006, 128, 11921-11926.

149

150

6. Glaser, N.; Adams, D. J.; Boker, A.; Krausch, G. Langmuir 2006, 22, 5227-5229.

7. Wood, A.; Giersig, M.; Mulvaney, P. Journal of Physical Chemistry B 2001, 105, 8810-8815.

8. Haruta, M. Gold Bulletin 2004, 37, 27-36.

9. Li, Y. Q.; Zhang, G.; Nurmikko, A. V.; Sun, S. H. Nano Letters 2005, 5, 1689-1692.

10. Gu, H. W.; Yang, Z. M.; Gao, J. H.; Chang, C. K.; Xu, B. Journal of the American Chemical

Society 2005, 127, 34-35.

11. Choi, J. S.; Jun, Y. W.; Yeon, S. I.; Kim, H. C.; Shin, J. S.; Cheon, J. Journal of the American

Chemical Society 2006, 128, 15982-15983.

12. Sokolov, K.; Follen, M.; Aaron, J.; Pavlova, I.; Malpica, A.; Lotan, R.; Richards-Kortum, R.

Cancer Research 2003, 63, 1999-2004.

13. Schultz, D. A. Current Opinion in Biotechnology 2003, 14, 13-22.

14. Pankhurst, Q. A.; Connolly, J.; Jones, S. K.; Dobson, J. Journal of Physics D-Applied Physics

2003, 36, R167-R181.

15. El-Sayed, I. H.; Huang, X. H.; El-Sayed, M. A. Nano Letters 2005, 5, 829-834.

16. Gupta, A. K.; Naregalkar, R. R.; Vaidya, V. D.; Gupta, M. Nanomedicine 2007, 2, 23-39.

17. Haigler, H.; Ash, J. F.; Singer, S. J.; Cohen, S. Proceedings of the National Academy of Sciences

of the United States of America 1978, 75, 3317-3321.

18. Kawamoto, T.; Sato, J. D.; Le, A.; Polikoff, J.; Sato, G. H.; Mendelsohn, J. Proceedings of the

National Academy of Sciences of the United States of America-Biological Sciences 1983, 80,

1337-1341.

19. Adams, G. P.; Weiner, L. M. Nature Biotechnology 2005, 23, 1147-1157.

20. Herbst, R. S. International Journal of Radiation Oncology Biology Physics 2004, 59, 21-26.

21. Simon, G. H.; Bauer, J.; Saborovski, O.; Fu, Y. J.; Corot, C.; Wendland, M. F.; Daldrup-Link, H. E.

European Radiology 2006, 16, 738-745.

22. Krutovskikh, V. A.; Troyanovsky, S. M.; Piccoli, C.; Tsuda, H.; Asamoto, M.; Yamasaki, H.

Oncogene 2000, 19, 505-513.

23. Xu, C. J.; Xu, K. M.; Gu, H. W.; Zheng, R. K.; Liu, H.; Zhang, X. X.; Guo, Z. H.; Xu, B. Journal

of the American Chemical Society 2004, 126, 9938-9939.

Chapter VII

Au-Fe3O4 Dumbbell NPs for Target-Specific Platin Delivery

This chapter describes the coupling of Herceptin antibody and platin complex to

Au-Fe3O4 nanoparticles. The platin-Au-Fe3O4-Herceptin NPs act as a target-specific

nanocarrier to deliver platin into Her2-positive breast cancer cells (Sk-Br3) with high

therapeutic effects. The conjugate has half maximal inhibitory concentration (IC50) to

Sk-Br3 cells at 1.76 μg Pt /ml, lower than that needed for cisplatin at 3.5 μg/ml. The

work demonstrates that the dumbbell-like Au-Fe3O4 nanoparticles are promising

nanocarriers for highly sensitive diagnostic and therapeutic applications.

1. Background

Pt-based platin complexes, such as cisplatin, carboplatin and oxaliplatin, as shown in

Figure 7-1a, are well-known generations of anticancer therapeutic agents.1 One

common feature of these square planar Pt complexes is that they all contain

coordination bonds of Pt-N/Pt-Cl, or Pt-N/Pt-O with two Pt-N bonds in cis-position.

Pt-Cl or Pt-O bonds in the complex are chemically much weaker than Pt-N bonds and

subject to facile hydrolysis in low Cl- and low pH conditions, giving charged

[cis-Pt(NH3)2(H2O)2]2+ that are highly reactive for DNA binding through the N7

atom of either an adenine or guanine base. This binding de-stacks the double helix

structure and interrupts with cell’s genetics/transcription machinery and repair

mechanism, leading to cell death.2,3 However, these powerful platin therapeutic agents

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have no capability of identifying the tumor cells from the healthy ones. As a result,

they tend to be taken up by any fast grown cells, tumorous and healthy ones alike,

causing the well-known toxic side effects.4,5

Here I want to show that dumbbell-like Au-Fe3O4 nanoparticles (NPs) can act as a

target-specific nanocarrier to deliver platin into Her2-positive breast cancer cells with

high therapeutic effects. Recent research progress has revealed that antigens are often

over-expressed on the surfaces of the fast growing tumor cells. These over-expressed

antigens provide obvious targets for specific binding as each type of antigens can be

selectively captured by a typical monoclonal antibody.6 Therefore, linked with a

monoclonal antibody, these carriers may achieve target-specific delivery through

strong antibody-antigen interactions and receptor-mediated endocytosis. The

dumbbell-like Au-Fe3O4 NPs offer an ideal platform for this delivery purpose. As

shown in Figure 7-1b, their core structure contains magnetic Fe3O4 NPs and optically

active Au NPs. Compared with the conventional single component iron oxide NPs

used for biomedical applications,7,8 the dumbbell-like Au-Fe3O4 NPs have the

following distinct advantages: (1) the presence of Fe3O4 and Au surfaces facilitates

the stepwise attachment of an antibody and a platin complex; (2) the structure can

serve as both magnetic and optical probes for tracking platin complex in cells and in

biological systems.

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Figure 7-1. (a) Structural illustration of the common therapeutic platin complexes; (b) Schematic illustration of the dumbbell-like Au-Fe3O4 NPs coupled with Herceptin and platin complex for target-specific platin delivery.

2. Results and discussion

To produce Au-Fe3O4 NPs for target-specific platin delivery, we first synthesized the

dumbbell-like Au-Fe3O4 NPs based on the published method.9 Briefly, Au NPs were

synthesized with size ranging from 4nm to 12nm based on the published methods

with oleylamine as surfactant.9,10 Then Au-Fe3O4 NPs were prepared via the

decomposition of iron pentacarbonyl, Fe(CO)5, over the surface of the Au NPs

followed by oxidation under air. The size of Fe3O4 NPs was tuned by controlling the

ratio between Au NPs and Fe(CO)5. A series of dumbbell-like NPs are shown in

Figure 7-2.

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Figure 7-2. Au-Fe3O4 Dumbbell NPs with different size of Au and Fe3O4 cores (a) 3nm-18nm; (b) 6nm-18nm; (c) 8nm-18nm; (d) 8nm-25nm. (Scale bar: 20nm)

In order to render the NPs water soluble, the oleate/oleylamine coated 8 nm – 18

nm Au-Fe3O4 NPs (Figure 7-3a) were modified by replacing oleate/oleylamine with

dopamine based surfactants (Figure 7-1b) following our published recipe.11 Then

Heceptin was linked with PEG through EDC/sulfo-NHS chemistry. In order to anchor

platin to Au side, Cisplatin-binding Ligand (L2H) was synthesized and replaced the

original surfactant on Au surface (Figure 7-1b). Later, platin was anchored on Au side

by reacting Au-S-CH2CH2N(CH2CH2COOH)2 with cisplatin. The final conjugate was

monodisperse in water (Figure 7-3b).

Figure 7-3. (a) Au-Fe3O4 nanoparticles as synthesized; (b) the final conjugates in water.

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The linkage of Au-Fe3O4-Heceptin was confirmed through matrix-assisted laser

desorption/ionization (MALDI) mass spectrometry (Figure 7-4). After conjugation,

the molecule peak at 150kDa corresponding to monoclonal antibody was clearly seen.

And the peak for subunit (75kDa) was also obvious.

Figure 7-4. Matrix-assisted laser desorption/ionization (MALDI) Mass Spectra of the Au-Fe3O4 NPs before (a) and after (b) coupling with Herceptin (Mr: 150 kDa).

Besides the evidence for Herceptin conjugation, the conjugation of platin-Au

was characterized by inductively coupled plasma atomic emission spectroscopy

(ICP-AES) and energy dispersive spectroscopy (EDS). The elemental analyses reveal

that the conjugate contains S/Pt at an atomic ratio of ~1/1 (Figure 7-5). This indicates

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that two carboxylic group’s replace two Cl’s in cisplatin, forming the platin complex

as shown in Figure 7-1b.

Figure 7-5. EDS characterization of S to Pt ratio for platin-Au-Fe3O4-Herceptin NPs.

The weight percentage of Pt/Au (~17.8%) was achieved through ICP analysis.

The difference between ICP and EDS came from the penetration problem in EDS

analysis. EDS could not reach the gold atom inside NPs. Based on ICP result, the

platinum number on each nanoparticle could be calculated out (Experimental part):

~2812 platin units are bound to each Au NP.

Table 7-1. ICP-AES analytical results in Au-Fe3O4 NPs for platin loading with or without platin binding ligand.

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We also characterized the size dependent platin loading on Au-Fe3O4 NPs. Among

the 3 nm-18 nm, 6 nm-18 nm, 8 nm-18 nm and 8 nm-25 nm Au-Fe3O4 NPs tested,

larger Au NPs were capable of incorporating more platin complexes, while the size of

the Fe3O4 had little effect on platin concentration (Table 7-1). This further proves that

platin binds to the Au side, not to the Fe3O4 side, as shown in Figure 7-1b. The final

conjugate can be dispersed in PBS. The 8 nm–18 nm Au-Fe3O4 NPs have a 32 nm

hydrodynamic diameter as measured by dynamic light scattering (DLS) (Figure 7-6).

10 1000

5

10

15

20

25

30

35

40

4030

Perc

enta

ge (%

)

Au-Fe3O4 in Hexane Au-Fe3O4 in Water Au-Fe3O4-Ab platin-Au-Fe3O4-Ab

20Size (nm)

Figure 7-6. Hydrodynamic diameter of Au-Fe3O4 nanoparticles at various functionalization stages

The specificity of the platin-Au-Fe3O4-Heceptin NPs was examined through their

preferred targeting to Sk-Br3 cells that are Her2-positive breast cancer cells

(Her2-negative breast cancer cells (MCF-7) were used as a control).12 Before

incubation with the platin-Au-Fe3O4-Heceptin NPs, Sk-Br3 and MCF-7 cells were

pre-blocked with 1% BSA. The cells were then incubated with the NPs in PBS for 1 h

and fixed with 4% paraformadehyde. The cells were later imaged using Leica TCS

SP2 AOBS spectral confocal microscope at 594 nm – the region where the Au NPs

show the strong reflection.11 Figure 7-7a&b show the reflection images of Sk-Br3

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cells (Figure 7-7a) and MCF-7 cells (Figure 7-7b). The brighter image (~1.5 times

brighter as measured through Image J) shown in Figure 7-7a indicates that more

platin-Au-Fe3O4-Heceptin NPs target to Sk-Br3 cells. We can conclude that under the

same incubation concentration, Herceptin helps the preferred targeting onto Sk-Br3

cells, not MCF-7 cells.

Figure 7-7. Reflection images of (a) Sk-Br3 cells and (b) MCF-7 cells after incubation with the same concentration of platin-Au-Fe3O4-Heceptin NPs. (c) Cisplatin and platin release curves at 37oC (pH = 7); (d) pH dependent Pt-release from platin-Au-Fe3O4-Herceptin at 37 oC.

TEM image analysis on the Sk-Br3 cells reveals the presence of NPs in

endosome/lysosome, which indicates that the NPs were up-taken through endocytosis

process (Figure 7-8).

Figure 7-8. TEM image of the platin-Au-Fe3O4-Heceptin nanoparticles in Sk-Br3 cells after two hour incubation.

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The platin release from the NP conjugate (100 μg Pt in 2 ml PBS) was analyzed in

a dialysis bag (MWCO = 1,000) that was put into a 30 ml PBS reservoir at 37 oC.

Cisplatin in the same Pt concentration was used as a control. The membrane of the

dialysis bag keeps the bound platin and the NPs inside the bag while the released

platin or free cisplatin can diffuse into the buffer reservoir from which the Pt

concentration was measured by ICP-AES. The platin release data are given in Figure

7-7c. It can be seen that 80% of free cisplatin diffuses through the dialysis bag in 1 h

while for the NP conjugate this release is reduced to only about 25% in the same

incubation time. Furthermore, the Pt-releases is pH dependent (Figure 7-7d). At pH =

6, 70% of platin is released from the platin-Au-Fe3O4-Heceptin NPs after 10 h while

at pH = 8, the amount of platin release is reduced to 40%. Clearly, lower pH

conditions accelerate the platin release from the conjugate shown in Figure 7-1b. As

endosome/lysosome has pH around 5, we can conclude that platin release will be

accelerated once the conjugate is inside the cells through endocytosis process.

Figure 7-9. Viability of Sk-Br3 cells after incubation with platin-Au-Fe3O4 NPs, platin-Au-Fe3O4-Herceptin NPs and free cisplatin.

The therapeutic effect of the platin-Au-Fe3O4-Heceptin NPs was studied by

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measuring the cell viability and p53 expression in Sk-Br3 cells. The control

experiments show that Au-Fe3O4 NPs without platin did not inhibit cell growth under

all Fe concentrations we tested (Figure 7-10a). Once coupled with platin, however, the

platin-Au-Fe3O4-Heceptin NPs have half maximal inhibitory concentration (IC50) to

Sk-Br3 cells at 1.76 μg Pt /ml (Figure 7-9), lower than that needed for cisplatin at

3.5μg/ml. Note that the platin-Au-Fe3O4 NPs without Herceptin is also toxic, but its

toxicity is less than cisplatin due to their non-specificity and the slow platin

hydrolysis in the conjugate. The highest toxicity to Sk-Br3 cells observed from the

platin-Au-Fe3O4-Heceptin NPs is clearly attributed to the specific targeting and

enhanced uptake of NPs by Sk-Br3 cells. In contrast, platin-Au-Fe3O4-Heceptin NPs

did not show obvious improvement in their toxicity to MCF-7 cells (Figure 7-10b).

Figure 7-10. (a) Viability of Sk-Br3 cells after incubation with Au-Fe3O4, Fe3O4-Au-platin and Herceptin-Fe3O4-Au-platin NPs under the same iron concentration; (b) Viability of MCF7 cells after incubation with Fe3O4-Au-platin, Herceptin-Fe3O4-Au-platin and cisplatin under the same platinum concentration; (c) p53 expression in Sk-Br3 cells after incubation with different concentrations of cisplatin; (d) p53 expression in Sk-Br3 cells after incubation with Au-Fe3O4, Fe3O4-Au-platin, Herceptin-Fe3O4-Au-platin or cisplatin under the same Pt concentration (1 μg/ml) or Fe concentration (45 μg/ml).

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The increase of Pt concentration within Sk-Br3 cells can also be monitored by the

accumulation of p53 - a tumor suppressor protein.13 This is easily seen in a control

experiment that more p53 are present with higher concentration of cisplatin added in

the cell culture medium with beta-actin as the loading control (Figure 7-10c). We

tested the p53 protein expression in Sk-Br3 cells after treatment with different NPs

and cisplatin. The cells treated with platin-Au-Fe3O4-Heceptin NPs have the highest

p53 expression (Figure 7-10d). This is consistent with what we observed in cell

toxicity data in Figure 7-9, indicating that Herceptin indeed induces more uptake of

platin into the Sk-Br cells, causing highly toxic effect to these cells.

3. Summary

al

In summary, I have demonstrated that the dumbbell-like Au-Fe3O4 NPs can serve as a

multifunctional platform for target-specific platin delivery. The release of the

therapeutic platin at a low pH condition render the NP conjugate more toxic to the

targeted tumor cells than the free cisplatin. The methodology developed here can be

generalized and the dumbbell-like Au-Fe3O4 NPs should have great potentials as

nanocarrriers for highly sensitive diagnostic and highly efficient therapeutic

applications.

4. Experiment

Materials and Instruments: All chemicals including α,ω-Bis11polyethylene glycol

(Mr = 3000) and (3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide)

161

(MTT) were purchased from Sigma Aldrich Corp. RIPA buffer (25mM Tris·HCl

pH=7.6, 150mM NaCl, 1% NP-40, 1% sodium deoxycholate, 0.1% SDS) was mixed

with HaltTM Protease inhibitor Cocktail before use (Pierce Corp.). Deionized (DI)

water was purified by a Millipore Milli-DI Water Purification system. UV/Vis

absorption spectra of the samples were measured with a PerkinElmer Lambda 35

UV/Vis spectrometer. Transmission electron microscopy (TEM) images were

acquired with Philips EM 420 (120kV) on amorphous carbon coated copper grids.

Cisplatin-binding Ligand (L2H) Synthesis: To a solution of cystamine

dihydrochloride (1.125 g, 5 mmol) in 100 mL acetone and 10 mL Et3N were added

ethyl bromoacetate (2.2 mL, 20 mL), KI (520 mg). After stirred for 6 h at room

temperature, the insoluble solid was removed by filtration. The filtrate was dried in a

rotavapor. The intermediate product was purified through flash chromatography

(petrol/EtOAc, 20:1). Yield: 80%. FAB-MS: m/z = 519[M+Na]+. 1H NMR (CDCl3,

300MHz): d 4.06-4.10 (8H, m, 4-H), 3.51 (8H, s, 5-H), 2.97-3.02 (4H, q, 2-H),

2.72-2.76 (4H, q, 3-H), 1.19-2.21 (12H, t, 1-H). The deprotection of carboxylic group

was carried in the methanol solution (0.496 g, 1 mmol). With 5 mL of 1 M NaOH

aqueous solution, the mixture was stirred for 30 min. If there was any precipitate,

small amount of water was needed. After 24 h of stirring, 20 mL distilled water was

added and the solution was acidified to pH = 3.0 with 1 M HCl (aq). The resulting

precipitate was collected by centrifugation and washed with EtOH/H2O (1:1).

FAB-MS: m/z = 385[M+H]+. 1H NMR (D2O, 300MHz): d 3.51 (8H, s, 3-H),

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2.97-3.02 (4H, t, 2-H), 2.72-2.76 (4H, t, 1-H).

Synthesis and Modification of Dumbbell Nanoparticles: Au-Fe3O4 nanoparticles

were synthesized and made water-soluble according to our previous work.9,11 To link

Her2 antibody (Herceptin) to nanoparticles, nanoparticles (5 mg) in water was mixed

with 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC, 1.1 mmol) and

sulfo-NHS (1 mmol) for 15 min. Then the conjugates run through PD-10 column

pre-washed with PBS to remove excessive EDC and sulfo-NHS. Then Herceptin

(100μg) was added into the PBS solution and shaked for 2 h. The final conjugates

were separated from unbound Herceptin with high speed centrifugation.

Linking Cisplatin onto Dumbbell Nanoparticles: The antibody-coupled Au-Fe3O4

nanoparticles (1 mg) were mixed with cisplatin binding ligand solution (10 mmol, 1

ml H2O) for 6 h. Later, the uncoupled ligand was removed through PD-10 column or

stirred cell (large amount synthesis). Cisplatin suspension (water, 20 mg/ml) was

added to the nanoparticles solution. After stirred for overnight in dark, free cisplatin

was separated from nanoparticles through low speed centrifugation (3000 rpm). The

NPs then run through PD-10 column to remove free cisplatin in solution. The amount

of platin was determined by ICP-AES.

Cisplatin Release: Herceptin-Fe3O4-Au-platin NPs with 100 Pt μg/2 ml were put into

dialysis bag (MWCO=1000, Spectrum Lab Corp), which was in 30 ml PBS bath at

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37oC. At certain time point, 1 ml PBS was sampled. The platinum concentration was

determined by ICP-AES.

Cell Experiments: Sk-Br3 cells were purchased from ATCC and cultured in a

glass-bottom Petri dish (MatTek Corp.) with Dulbecco’s Modified Eagle’s Medium

(DMEM) with 10% FBS and 1% antibiotics. Before incubation with nanoparticle

conjugates, the cells were washed with PBS two times and blocked with 1% bovine

serum albumin (BSA) in PBS. The nanoparticles solution in DMEM was incubated

with cells for 1 h. Then, those cells were washed with PBS three times and fixed by 4%

paraformaldehyde solution. After 30 min, the cells were washed with PBS for

reflection imaging using a Leica TCS SP2 AOBS spectral confocal microscope.

Cell Viability Test. Viability of Sk-Br3 cells incubated with Fe3O4-Au-platin,

Herceptin-Fe3O4-Au-platin or cisplatin were examined through MTT assay. This cell

viability test was based on the reduction of the tetrazolium salt MTT

(3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) by mitochondrial

reductase in metabolically active cells. The cells were seeded onto 96-well culture

plates at a density of 4000 cells per well in DMEM (200 μL) containing 10% FBS.

After 24 h incubation at 37oC, nanoparticles in DMEM buffer at different

concentrations were added. The particles were removed after 24 h incubation. Then

MTT solution (5 mg/ml in PBS) was added to each well to evaluate cell viability.

After 1 h at 37oC, the solution was removed. 100 μL DMSO was added to dissolve

164

cells. After 30 min incubation at 37 oC, the viability was measured by microreader.

Preparation of Sk-Br3 cell sample for TEM: Nanoparticles were dispersed in the

cell culture medium (DMEM with 10% FBS, 1% penicillin) at a concentration of 0.01

mg Fe/mL dispersion. The mixture was incubated for 2 h and washed twice with PBS

to remove the excess particles. The cells were detached with 0.05% trypsin EDTA and

fixed with modified Karnovsky’s Fixative (2% paraformaldehyde and 2%

gluteraldehyde in PBS) before they were post-fixed in 1% OsO4 for 1.5 h, stained

with 2% uranyl acetate for 2 h, and dehydrated in alcohol and propylene oxide. The

treated cells were then embedded in Eponate resin, sectioned with an ultramicrotome,

and mounted on the 150 mesh TEM grids. The sections were then stained again with

uranyl acetate (25 min) and lead citrate (10 min) for TEM image analysis. The images

were acquired from a Philips EM 420 at 80 kV.

p53 Protein Detection with Western Blot:

100,000 Sk-Br3 cells were plated in each well of a 6 well plate for 24 h. Au-Fe3O4,

platin-Au-Fe3O4, platin-Au-Fe3O4-Herceptin or cisplatin in 1mL DMEM were added.

After 16 h incubation, cells were collected and washed with cold PBS twice. Then

200 μL cold RIPA buffer was added to the cells and kept in ice for 40 min. The cell

lysate was gathered through centrifugation at 14,000 rpm for 15 min. The

concentration of protein for each sample was measured through Bradford protein

assay.14 20 μg protein was loaded in each well for SDS-PAGE. The proteins are

165

166

transferred to nitrocellulose membrane and blotted with Santa Cruz monoclonal p53

antibody (DO-1).

References:

1. Jamieson, E. R.; Lippard, S. J. Chemical Reviews 1999, 99, 2467-2498.

2. Kelland, L. Nature Reviews Cancer 2007, 7, 573-584.

3. Wang, D.; Lippard, S. J. Nature Reviews Drug Discovery 2005, 4, 307-320.

4. Siddik, Z. H. Oncogene 2003, 22, 7265-7279.

5. Gately, D. P.; Howell, S. B. British Journal of Cancer 1993, 67, 1171-1176.

6. Adams, G. P.; Weiner, L. M. Nature Biotechnology 2005, 23, 1147-1157.

7. Sun, C.; Lee, J. S. H.; Zhang, M. Q. Advanced Drug Delivery Reviews 2008, 60, 1252-1265.

8. Jun, Y. W.; Lee, J. H.; Cheon, J. Angewandte Chemie-International Edition 2008, 47, 5122-5135.

9. Yu, H.; Chen, M.; Rice, P. M.; Wang, S. X.; White, R. L.; Sun, S. H. Nano Letters 2005, 5,

379-382.

10. Peng, S.; Lee, Y.; Wang, C.; Yin, H.; Dai, S.; Sun, S. Nano Research 2008, 1, 229-234.

11. Xu, C.; Xie, J.; Ho, D.; Wang, C.; Kohler, N.; Walsh, E. G.; Morgan, J. R.; Chin, Y. E.; Sun, S.

Angewandte Chemie-International Edition 2008, 47, 173-176.

12. Daly, J. M.; Jannot, C. B.; Beerli, R. R.; GrausPorta, D.; Maurer, F. G.; Hynes, N. E. Cancer

Research 1997, 57, 3804-3811.

13. Yazlovitskaya, E. M.; DeHaan, R. D.; Persons, D. L. Biochemical and Biophysical Research

Communications 2001, 283, 732-737.

14. Sapan, C. V.; Lundblad, R. L.; Price, N. C. Biotechnology and Applied Biochemistry 1999, 29,

99-108.

Chapter VIII Controlled Release of Fe from FePt Nanoparticles for Tumor

Inhibition

This chapter describes that chemically disordered face centered cubic (fcc) FePt

nanoparticles (NPs) show pH-dependent release of Fe in low biological pH conditions.

The released Fe catalyzes H2O2 decomposition into reactive oxygen species, causing

fast oxidation and deterioration of lipid membrane. Functionalized with luteinizing

hormone-releasing hormone (LHRH) peptide via phospholipid, the fcc-FePt NPs can

bind preferentially to the human ovarian cancer cell line (A2780) that over-expresses

LHRH receptors, and exhibit high toxicity to these tumor cells. The work

demonstrates that once coupled with a targeting agent, the fcc-FePt NPs can be

delivered site-specifically to the tumor cells and function as a powerful therapeutic

agent.

1. Background

In the chapter one, I have discussed the solution phase synthesis and self-assembly of

FePt NPs. Due to their specific magnetic property and stable structure, those

monodisperse FePt nanoparticles (NPs) have been studied extensively for potential

applications in data storage1-3, exchange-spring nanocomposite magnet4, biodetection5,

and fuel cell catalyst6-8. As-synthesized, the FePt NPs adopt a chemically disordered

face centered cubic (fcc) structure that can be converted to face centered tetragonal

167

(fct) structure via high temperature annealing1. In the fcc-FePt, both Fe and Pt are

randomly positioned in the structure while in the fct-FePt, Fe and Pt form alternating

atomic layers stacked along the [001] direction9. Such structural difference in fcc-FePt

and fct-FePt NPs leads to distinctive property change not only in magnetism1 but also

in chemical stability. In the recent acid resistance test, we found that Fe in the

fcc-FePt NPs could be etched away in a dilute HCl solution while fct-FePt NPs were

stable against this etching in the same solution (Figure 8-1). We further noticed that

this Fe-release phenomenon was common for the fcc-FePt NPs in low pH solutions

and the released Fe could catalyze H2O2 decomposition into reactive oxygen species

(ROS) that are highly reactive for lipid membrane oxidation. Such Fe-catalyzed ROS

formation and its toxicity to cellular systems have long been known and an

uncontrolled accumulation of Fe in cellular environments can lead to serious cellular

damage and cell death10-12.

Figure 8-1. Fe release from fcc-Fe53Pt47 NPs in 0.1M HClO4 solution

Here I want to show that the Fe build-up and its therapeutic effect in cellular

system can be controlled through the fcc-FePt NPs. I will demonstrate that the

168

fcc-FePt NPs are readily transferred into water and are chemically stable in neutral pH

conditions. Once inside cells, these NPs release Fe in the low pH cellular environment.

The released Fe catalyzes the decomposition of H2O2 generated from mitochondria,

producing ROS and causing lipid membrane oxidation and cell death. These processes

are outlined in Figure 8-2a. More importantly, the fcc-FePt NPs can be further

functionalized with a targeting peptide, luteinizing hormone-releasing hormone

(LHRH) for their preferred uptake by the fast growing A2780 cells from human

ovarian carcinoma that over-express LHRH receptors (LHRHR), and not by cells with

low LHRHR expression13. After Fe-release, the remaining Pt-rich FePt NPs have

much less toxicity. This controlled delivery of catalytic Fe makes fcc-FePt NPs a

unique NP-based agent with powerful therapeutic capability.

Figure 8-2. (a) FePt NPs uptake by a cell through endocytosis followed by Fe release from FePt NPs in lysosome. The released Fe catalyzes the decomposition of H2O2 to form hydroxyl radicals. (b) the phospholipid molecule used for FePt NP functionalization. (c) FePt NPs modified by phospholipid addition.

169

2. Results and discussion

The fcc-FePt NPs14 and the Fe3O4 NPs15 were synthesized according to the published

methods. The sizes of the NPs were controlled to be around 9 nm as measured by

transmission electron microscopy (TEM). The composition of Fe and Pt was

controlled by Fe(CO)5 and Pt(acac)2 ratio during the synthesis and was measured

through inductively coupled plasma atomic emission spectroscopy (ICP-AES). TEM

images of the 9 nm fcc-Fe40Pt60 NPs and the 9 nm Fe3O4 NPs are given in Figure 8-3.

Figure 8-3. TEM images of the as synthesized (a) 9 nm Fe40Pt60 NPs and (b) 9 nm Fe3O4 NPs. (Scale bar: 20nm)

The as-synthesized NPs were coated with a layer of oleate and oleylamine and

were made water-soluble through surfactant addition of the commercially available

phospholipid (Figure 8-2b).16 In this functionalization process, DSPE-PEG(2000)

carboxylic acid (1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-

[carboxy(polyethylene glycol)2000] (ammonium salt)) was added to the surface of the

NPs. The hydrocarbon chains of oleate/oleylamine and the phospholipid molecule

lock together through hydrophobic interaction, forming a robust double layer that

efficiently stabilizes the NPs in aqueous solutions (Figure 8-2c). The free lipids were

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simply removed by dialysis and filtration. The phospholipid modified NPs have the

hydrodynamic diameters around 60 nm (Figure 8-4) in phosphate buffered saline

(PBS) as measured by dynamic light scattering (DLS). The phospholipid coated NPs

were also stable in PBS + 10% fetal bovine serum (FBS) (pH = 7.4) and their

hydrodynamic diameters were at 80 nm during the 72 hr incubation (37ºC) period

(Figure 8-4).

0 10 20 30 40 50 60 70 800

20

40

60

80

100

120

140

Hyd

ynam

ic D

iam

eter

(nm

)

Incubation Time (hours)

FePt in PBS FePt in PBS with 10% FBS

Figure 8-4. Hydrodynamic diameter change of Fe40Pt60 NPs in PBS and PBS with 10% FBS. The size of NPs was measured by Dynamic Light Scattering (Zeta Sizer NanoS90, Malvern Instruments). The following parameters were used for size estimation: refractive index 2.37 (FePt), 1.423 (PBS); viscosity 0.625 (PBS); absorption 0.4 (FePt).

To examine pH-dependent Fe release, we put the fcc-FePt NPs in a dialysis bag

and incubated them at 37 ºC in PBS with pH at 4.8 and 7.4 – two conditions that are

met in lysosomes and cytosol respectively within cells.17 The etching results are

shown in Figure 8-5a. At pH = 7.4, both 9 nm fcc-Fe40Pt60 NPs and the 9 nm Fe3O4

NPs NPs have no measurable Fe amount in the solution outside the dialysis bag

within 24 hrs. But at pH = 4.8, the fcc-FePt NPs have a drastic increase in Fe

concentration in the solution after ~8 hrs, while the Fe3O4 NPs have only very small

171

amount of Fe released within 24 hrs of incubation. Further analysis on the fcc-FePt

NPs with different Fe, Pt compositions shows that all Fe62Pt38, FePt54Pt46, and Fe27Pt73

NPs have negligible Fe-release at pH = 7.4 while at pH = 4.8, Fe-rich FePt NPs tend

to release more Fe (Figure 8-6a). It is worth to note that free Pt ion was not detected

in the same incubation conditions (Figure 8-6b). These indicate that (1) unlike easily

oxidized metallic Fe NPs18, the FePt alloy NPs act as a reservoir for Fe and (2) Pt is

tightly associated with the FePt NPs.

0 4 8 12 16 20 24

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10

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30

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Rel

ease

d Fe

(%)

Fe40Pt60 NPs in pH=7.4 Fe40Pt60 NPs in pH=4.8 Fe3O4 NPs in pH=7.4 Fe3O4 NPs in pH=4.8

0 100 200 300 400

4

8

12

16

Inte

nsity

( x

1000

)

Time (mins)

Fe3O4 NPs + A2780 cells Fe40Pt60 NPs + A2780 cells A2780 cells

DCFH‐DA DCFH DCF

a

b

c d

e

Figure 8-5. (a) Fe release from Fe40Pt60 and Fe3O4 NPs in PBS with different pHs at 37oC. (b) Schematic illustration of DCFH-DA conversion to DCF. (c) Fluorescent intensity (Excitation at 488nm and emission at 530 nm at 37ºC) from DCFH-DA labeled A2780 cells with different NPs (Fe concentration: 1.5 μg/ml) in HBSS. Green fluorescent images of DCFH-DA labeled A2780 cells incubated with (d) Fe40Pt60 NPs (Fe concentration: 1.5 μg/ml) and (e) with HBSS buffer only (Excitation at 488 nm and emission from 510-550 nm).

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Figure 8-6. (a) Fe release from FePt with different composition in PBS under different pHs, and (b) Pt release from Fe40Pt60 and Pt NPs in PBS under different pHs. (37C)

The ROS concentration increase in A2780 cells initiated by Fe releasing from the

fcc-FePt NPs was characterized by 2’,7’-dichlorodihydrofluorescein diacetate

(DCFH-DA) reaction19 as shown in Figure 8-5b. Due to the successive formation of

ROS and the reaction (Figure 8-5b), the fcc-FePt NP treated A2780 cells exhibited 1.5

times stronger fluorescent signal than Fe3O4 NPs after 6 hr incubation (Figure 8-5c).

Further fluorescent microscopic studies confirmed that the cells incubated with

DCFH-DA and the fcc-FePt NPs in Hank’s Buffered Salt Solution (HBBS) had

stronger green fluorescence from DCF (Figure 8-5d), similar to that from the cells

incubated with pure H2O2 (Figure 8-7a). In contrast, fluorescent intensity from the

cells incubated with Fe3O4 NPs was much weaker (Figure 8-5c and Figure 8-7b) and

close to that from the pure A2780 cells Figure 8-5c&e). These experiments confirm

that the fcc-FePt NPs induce faster H2O2 decomposition and formation of ROS in

A2780 cells.

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Figure 8-7. Green fluorescent images (Excitation at 488 nm and emission from 510-550 nm, Leica TCS SP2 AOBS spectral confocal microscope) of A2780 cells incubated with (a) H2O2 (25 μM); and (b) Fe3O4 NPs (Fe concentration: 1.5 μg/ml) in HBSS. Cells were pre-incubated with DCFH-DA (10 μM in HBSS, 0.5% DMSO (vol/vol)) for 30 mins.

The excessive production of ROS within cells causes cell death by inducing lipid

oxidation, DNA and protein damage11. Here we assessed the membrane lipid damage

via the oxidation of a fluorescent dye C11-BODIPY (Molecular Probes) to BODIPY20,

as illustrated in Figure 8-8a. C11-BODIPY molecule inserts into cell membrane and

allows for quantitative assessment of membrane lipid oxidation by ROS through

emission wavelength change during the oxidation21. In principle, oxidation of

C11-BODIPY in cell membrane decreases the emission at 595 nm and increases the

emission at 520 nm. In the experiment, the C11-BODIPY labeled cells treated with

fcc-FePt NPs show stronger emission at 520 nm (Figure 8-9a) and weaker emission at

595 nm (Figure 8-8b) compared with the untreated cells or those incubated with

Fe3O4 NPs. The emission changes at 520 nm and at 595 nm reveal the oxidation of

C11-BODIPY, which can be related to the damage to the lipid membrane of the

A2780 cells. The lipid membrane damage caused by fcc-FePt NPs was also evidenced

through the fluorescent images of the A2780 cells incubated only with C11-BODIPY

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(Figure 8-8c), which are brighter, and those incubated with both C11-BODIPY and

fcc-FePt NPs (Figure 8-8d), which are darker due to the C11-BODIPY oxidation.

Conversely, the image intensity from the BODIPY emission is increased (Figure

8-9b&c). The damage to the lipid membrane of endosome/lysosome within the A2780

cells can be visualized by TEM image (Figure 8-8e), in which two vesicles are shown

with one completely broken and the FePt NPs trapped inside spreading to larger area.

In contrast, endosomes/lysosomes of the A2780 cells treated with Fe3O4 NPs in the

same condition are clearly visible in the TEM image and no lipid membrane damage

can be observed (Figure 8-9d).

Figure 8-8. (a) Schematic illustration of ROS initiated oxidation of C11-BODIPY into BODIPY. Upon its oxidation, the maximum emission (595 nm) is shifted to 520 nm. (b) Fluorescent emission intensity detected at 595 nm from C11-BODIPY labeled A2780 cells treated with FePt or Fe3O4 NPs with a Fe concentration of 1.5 ppm in HBSS. Fluorescent images of the A2780 cells treated with (c) C11-BODIPY and those treated with (d) C11-BODIPY and Fe40Pt60 NPs. The images were collected from 580-610 nm with excitation at 534 nm. (e) TEM image of the two vesicles within an A2780 cell treated with Fe40Pt60 NPs, one vesicle is completely broken and the second one is partially damaged.

175

Figure 8-9. (a) Fluorescent emission signal at 520 nm from C11-BODIPY labeled A2780 cells treated with NPs (Fe: 1.5 ppm) in HBSS. Cells were pre-stained for 30 mins with a 10 μM solution of C11-BODIPY in HBSS (prepared from the stock solution 10mM C11-BODIPY in methanol) prior to NPs treatment. Fluorescent images of (b) the untreated A2780 cells and (c) the A2780 cells treated with Fe40Pt60 NPs. The images were collected from 510-540 nm with excitation at 488 nm. (d) TEM image of the internal part of an A2780 cell labeled with Fe3O4 NPs, showing the intact endosome/lysosome.

The increased ROS level caused by Fe-catalyzed H2O2 decomposition within

cells results in serious toxicity to A2780 cells and other cancer cells, including HeLa,

A431 (human epithelial carcinoma cell line), Sk-Br3 (human breast carcinoma cell

line), CPAE (Cultured Plumonary Artery Endothelial cell line) and HEK-293 cells

(human embryonic kidney cell line) (Figure 8-10a).

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Figure 8-10. (a) Viability of several different cell lines incubated with Fe40Pt60 NPs at different iron concentrations for 24 hrs. (b) A2780 cells were incubated with Fe40Pt60 NPs or with Fe40Pt60 NPs plus 200 μM of 2,2’-bipyridine at 37oC for 24 hrs before the plate was measured using 550 nm as test wavelength and 630 nm as the reference wavelength. Viability was calculated based on the recorded data.

For A2780 cells, the fcc-FePt NPs shown in Figure 8-2c have an IC50 of 1.25 μg

Fe/ml (Figure 8-10b). Characteristically, the generation of ROS catalyzed by the

fcc-FePt NPs, and therefore the toxicity of the NPs, can be blocked with

2,2’-bipyridine – an iron chelator that has been used to capture Fe within bacteria to

reduce the formation of ROS22. 2,2’-Bipyridine was non-toxic to A2780 cells under

200 μM concentration (Figure 8-11a). In the presence of 200 μM 2,2’-bipyridine, the

fcc-FePt NPs show much less toxicity to A2780 cells (Figure 8-10b). This indicates

that Fe released from the fcc-FePt NPs is chelated by 2,2’-bipyridine and is much less

active for H2O2 decomposition.

Figure 8-11. (a) Viability of A2780 cells incubated with 2,2’-bipyridine for 24 hrs; (b) viability of A2780 cells incubated with the pre-etched Fe40Pt60 NPs.

177

The high toxicity induced by the fcc-FePt NPs can be directed to specific tumor

cells once a targeting agent is coupled to the NPs. Here we choose luteinizing

hormone-releasing hormone (LHRH) peptide as such a targeting agent. It is known

that LHRH receptors (LHRHRs) are over-expressed on breast, ovarian, and prostate

cancer cells, and are not detectable on most visceral organs23. In this part of the

experiments, we first coupled LHRH peptide to the fcc-FePt NPs (Figure 8-2c) via the

common EDC/Sulfo-NHS chemistry. Using similar chemistry, we also deactivated

the –COOH group with CH3-PEG4-NH2 so that the fcc-FePt-CH3 NPs could be used

as a control. A2780 cells that over-express LHRHRs and HEK-293 cells (human

embryonic kidney) that have low LHRHR expression were chosen for cell targeting

demonstration13. ROS concentration increase in cells was characterized by the

fluorescent signal change from the oxidation of DCFH-DA to DCF as illustrated in

Figure 8-5b. Figure 8-12a&b are the results from A2780 cells and HEK-293 cells

respectively. It can be seen that fluorescent signal from the cells treated with

fcc-FePt-LHRH is much stronger than that from those treated with fcc-FePt-CH3

(Figure 8-12a). But the fluorescent signals from HEK-293 cells incubated with these

two kinds of particles are similar (Figure 8-12b). The conclusion is that

fcc-FePt-LHRH NPs can target specifically to A2780 cells, but not to HEK-293 cells,

and greatly increase the Fe-catalyzed ROS formation in cells. This is further

confirmed by the cell viability tests as over 50% of A2780 cells are dead at 0.4 μg

Fe/ml concentration (Figure 8-12c), much less than that needed for the

fcc-FePt-COOH NPs at 1.25 μg Fe/ml (Figure 8-10b). In contrast, most of the

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HEK-293 cells survive such treatment (Figure 8-12d). For the fcc-FePt-CH3 NPs, both

cells show only small amount of death due to the lack of preferred uptake24 (Figure

8-12c&d). It is also important to note that after Fe release in acidic buffer (pH=4.8)

for 1 hr, the Pt-rich fcc-FePt NPs show drastic decrease in toxicity to the A2780 cells

compared with those NPs in neutral buffer (Figure 8-11b). This controlled delivery of

catalytic Fe indicates that fcc-FePt NPs coupled with a target agent may serve as a

powerful agent for cancer therapy.

0 1 2 3

20

40

60

80

100

Viab

ility

(%)

Fe Concentration (μg/ml)

A2780 cells + FePt-LHRH A2780 cells + FePt-CH3

100 200 300 4000

400

800

1200

Fluo

resc

ent I

nten

sity

Time (mins)

A2780 cells A2780 cells + FePt-CH3

A2780 cells + FePt-LHRH

0 100 200 300 4000

400

800

1200

Fluo

resc

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HEK-293 cells HEK-293 cells + FePt-CH3 HEK-293 cells + FePt-LHRH

a b

c d

0 1 2 3

20

40

60

80

100

Viab

ility

(%)

Fe Concentration (μg/ml)

HEK-293 cells + FePt-LHRH HEK-293 cells +FePt-CH3

Figure 8-12. Fluorescent intensity of DCF from (a) A2780 and (b) HEK-293 cells: the cells were pre-incubated with DCFH-DA (10 μM in HBSS, 0.5% DMSO (vol/vol)) for 30 mins before their incubation with FePt-LHRH and FePt-CH3 at the Fe concentration of 1.5 ppm. Viability of (c) A2780 and (d) HEK-293 cells incubated with FePt-LHRH and FePt-CH3 NPs measured through MTT assay.

179

3. Summary

al

In summary, I have presented an exciting new property of fcc-FePt NPs - their

pH-dependent release of Fe in low pH conditions for catalytic H2O2 decomposition.

This property is successfully demonstrated in low pH cellular environments and the

resultant lipid membrane oxidation leads to fast cell death. Initial experiments indicate

the therapeutic effects of fcc-FePt NPs to cells can be regulated and specifically

targeted to human ovarian cancer cell line (A2780) when the fcc-FePt NPs are

coupled with LHRH peptide. The work suggests that high therapeutic efficiency can

be achieved by using fcc-FePt NPs as a drug. Furthermore, these fcc-FePt NPs are

superparamagnetic at room temperature and have been tested as a contrast

enhancement probes for magnetic resonance imaging (MRI)25,26. This, plus their

therapeutic effects, should warrant fcc-FePt NPs a promising agent for imaging

guided cancer therapy.

4. Experiment

Materials and Instruments:

LHRH peptides (Gln-His-Trp-Ser-Tyr-DLys(DCys)-Leu-Arg-Pro-NHEt, MW=1344.5)

were synthesized according to Minko’s design13 by American Peptide (Sunnyvale,

CA). Methyl-PEG4-Amine, 1-Ethyl-3-[3-dimethylaminopropyl]carbodiimide

hydrochloride (EDC) and N-Hydroxysulfosuccinimide (Sulfo-NHS) were purchased

from Thermo Scientific. 2’,7’-dichlorodihydrofluorescein diacetate (DCFH-DA)

and C11-BODIPY were purchased from Molecular Probes. All other chemicals were

180

purchased from Sigma-Aldrich and used without further purification. All the

biological buffers were purchased from Fisher Scientific. Deionized (DI) water was

purified by a Millipore Milli-DI Water Purification system. TEM images were taken

on a Philips EM 420 (120 kV). Fluorescent images were acquired on a Leica TCS SP2

AOBS spectral confocal microscope. Hydrodynamic sizes of NPs were measured by

Malvern Zeta Sizer S90 dynamic light scattering instrument. Iron concentration was

determined with inductively coupled plasma atomic emission spectroscopy

(Jobin-Yvon JY2000).

Cell culture: All human cancer cell lines were obtained from ATCC and cultured

(37oC, 5% CO2) in 75 cm2 flasks (Corning) containing Dulbecco's Modified Eagle

Medium (DMEM), 1% antibiotics and 10% FBS. Cells were used at 5-15 passages.

Coating NPs with phospholipid: FePt NPs were synthesized according to Chen et

al14. Fe3O4 NPs were prepared according to Sun et al15. After synthesis, NPs were

precipitated out with ethanol to remove the excessive surfactant and later dissolved in

1ml chloroform. 2-5 mg NPs in 1 ml chloroform were mixed with 10 mg

DSPE-PEG(2000)carboxylic acid lipid (1 ml chloroform solution) (MW=2847.779,

Avanti Polar Lipids Inc). The solvent was removed through rotavapor and DI water or

PBS buffer was added to disperse NPs. The NPs were later dialysis against 100k

dialysis bag (Spectrum Laboratories, Inc) and the final conjugates were filtered

through 0.22 μm Millex@GP filter (Millipore Corp.) to remove aggregates.

181

Concentration was determined with ICP-AES.

Measurement of ROS with DCFH-DA: A2780 cells were plated out at a density of

20,000 cells per well into black 96-well plates and were incubated at 37 oC, 5% CO2,

high humidity for 24 hrs. Cells were loaded with DCFH-DA [10 μM in HBSS, 0.5%

DMSO (vol/vol)] and were incubated at 37 oC, 5% CO2, high humidity for 30 mins. The

probe was removed and the cells were washed twice with PBS (200 μl). The cells were

then kept in Hanks’ Balanced Salt Solution (HBSS) and NPs were diluted with HBSS

and added to the wells. Hydrogen peroxide (25 μM) was added as a positive control and

the fluorescence was recorded every 15 min over a period of 6 hrs at 37 oC by excitation

at 480 nm and emission at 538 nm on a BMG FLUO star plate reader.

Lipid peroxidation: A2780 cells were stained for 30 mins with a 10 μM solution of

C11-BODIPY in HBSS (prepared from the stock solution 10 mM C11-BODIPY in

methanol) prior to NP treatment in HBSS buffer. The probe was removed and the cells

were washed twice with PBS (200 μl). The cells were then kept in HBSS and NPs

were diluted with HBSS (Fe: 1.5 ppm; Pt: 7.6 ppm) and added to the wells. The

fluorescence was recorded every 30 mins over a period of 6 hrs at 37 oC

(non-oxidized: λex 581 nm, λem 595 nm; oxidized: λex 485 nm, λem 520 nm). For the

fluorescent microscope examination, the similar steps were used as ROS

measurement except that DCFH-DA was replaced with C11-BODIPY.

182

Cytotoxicity assay (MTT assay):

Colorimetric MTT (3-(4,5-dimethylthiazol-2-yl)-2,5- diphenyl tetrazolium bromide,

Sigma) assays were performed to assess the mitochondrial activity of cells treated as

described above. Cytotoxicity assay was performed in 96-wells microtiter plates

(Fisher Inc.) with seeding density, 4000 cells per well. Microtiter plates containing cells

were pre-incubated for 24 hrs at 37 oC in order to allow stabilization before the addition

of the test substance. The plates were incubated with the test substance for 24 hrs at 37

oC and 5% CO2. Then 100 μg/ml MTT solution (DMEM) was added to each well to

evaluate cell viability after the NPs solution was removed. After 2 hrs at 37 oC, MTT

solution was removed. 100 μL DMSO was added to dissolve cells. After 30 mins

incubation under 37 oC, the plate was measured using 550 nm as test wavelength and

630 nm as the reference wavelength on microreader (SpectraMax 340PC384,

Molecular Devices). Viability was calculated based on the recorded data.

Fluorescent microscope examination of ROS with DCFH-DA as indicator: for the

fluorescent microscope examination, A2780 cells were plated onto cover slips

(Corning, Corning, NY) in 12-well plates. Cells monolayer grown on glass cover slips

were allowed to near confluence, and then placed on ice, the medium was aspirated,

and the cells were washed twice with PBS. Then cells were pre-stained with 20 μM

DCFH-DA in PBS for 30 mins and washed with PBS to remove free DCFH-DA.

Later, the cells were incubated with NPs or other reagents for 2 hrs before they were

washed with PBS for three times and fixed with 4% formaldehyde in PBS for 10 mins.

183

The cover slips with cells on the surface were removed from the wells and mounted

onto slides using 90% glycerol in H2O. The images were acquired by a Leica TCS

SP2 AOBS spectral confocal microscope with excitation at 488 nm and emission from

510-550 nm.

Fluorescent microscope examination of Lipid peroxidation with C11-BODIPY:

for the fluorescent microscope examination, the similar steps were used as ROS

measurement except that DCFH-DA was replaced with C11-BODIPY.

Coupling LHRH or methyl-PEG4-amine with FePt NPs: 0.4 mg EDC (2mM),

1.1mg sulfo-NHS (5 mM) were incubated with NPs (10 mg) in water for 15 mins.

Then 1.4 μL 2-mercaptoethanol was added to inactivate excessive EDC. The

Sulfo-NHS-NPs intermediate was separated from others with PBS pre-equilibrated

PD-10 column (GE Healthcare). Then 0.5 mg LHRH peptide or 0.08 mg

methyl-PEG4-amine was added to Sulfo-NHS-NPs intermediate and reacted for 2 hrs

at room temperature. The final product was purified with PD-10 column. The

successful coupling of LHRH was confirmed through Matrix-assisted laser

desorption/ionization (MALDI) Mass Spectra (Applied Biosystem, Voyager-DE PRO,

BioSpectrometry Workstation) (Figure 8-13).

TEM sample preparation: after incubated with NPs for 4 hours and washed with

PBS, the cells were detached with 0.05% trypsin EDTA and fixed with modified

184

Karnovsky’s Fixative (2% paraformaldehyde and 2% gluteraldehyde in PBS) before

they were post-fixed in 1% OsO4 for 1.5 hrs, stained with 2% uranyl acetate for 2 hrs,

and dehydrated in alcohol and propylene oxide. The treated cells were then embedded

in Eponate resin, sectioned with an ultramicrotome, and mounted on the 150 mesh

TEM grids. The sections were stained again with uranyl acetate (25 mins) and lead

citrate (10 mins) for TEM image analysis. The images were acquired from a Philips

EM 420 at 80 kV.

Figure 8-13. Matrix-assisted laser desorption/ionization (MALDI) Mass Spectra of FePt-LHRH with a-Cyano-4-hydroxycinnamic acid as calibration matrix, which clearly shows the LHRH peak

W=1345+1) (M

185

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