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
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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
30
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.
32
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
40
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
41
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.
43
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.
48
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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
53
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
54
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.
55
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
56
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
57
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).
58
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
59
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-
60
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.
61
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.
62
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
63
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
64
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).
65
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
66
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.
67
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.
68
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.
69
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),
70
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
73
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
74
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
76
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
77
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,
78
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
79
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
80
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.
81
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.
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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
84
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.
85
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)
87
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
88
~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
89
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
91
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
92
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
93
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
94
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.
95
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.
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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.
98
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
99
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.
100
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
101
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.
102
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.
103
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.
104
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).
105
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
106
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.
107
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
108
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.
113
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
115
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
117
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,
118
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-
119
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
123
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
126
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).
127
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).
128
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.
131
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
134
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.
136
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.
138
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
139
(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
140
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
141
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.
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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
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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
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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
0
10
20
30
Time (hours)
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
174
(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.
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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
ent I
nten
sity
Time (mins)
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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