colloidal microcapsules: surface engineering of
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5-13-2011
Colloidal Microcapsules: Surface Engineering of Nanoparticles for Colloidal Microcapsules: Surface Engineering of Nanoparticles for
Interfacial Assembly Interfacial Assembly
Debabrata Patra University of Massachusetts Amherst
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COLLOIDAL MICROCAPSULES: SURFACE ENGINEERING OF
NANOPARTICLES FOR INTERFACIAL ASSEMBLY
A Dissertation Presented
By
DEBABRATA PATRA
Submitted to the Graduate School of the University of Massachusetts Amherst in partial fulfillment
of the requirements for the degree of
DOCTOR OF PHILOSOPHY
May 2011
Chemistry
© Copyright by Debabrata Patra 2011
All Rights Reserved
COLLOIDAL MICROCAPSULES: SURFACE ENGINEERING OF NANOPARTICLES FOR INTERFACIAL ASSEMBLY
A Dissertation Presented
By
DEBABRATA PATRA
Approved as to style and content by: _______________________________________ Vincent M. Rotello, Chair _______________________________________ Dhandapani Venkataraman, Member _______________________________________ Michael Barnes, Member _______________________________________ T. J. Lakis Mountziaris, Member
____________________________________ Craig T. Martin, Department Head Chemistry
DEDICATION
To my parents and teachers
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ACKNOWLEDGEMENTS
I owe endless gratefulness to many people who pleasantly involved themselves in helping
me undertake this dissertation since I joined graduate school. It is a pleasure to convey
my gratitude to them all in my humble acknowledgement.
In first place, I want to express my deepest gratitude to my research advisor,
Professor Vincent Rotello, for his continuous support, patience and motivation to make
my Ph.D experience productive and stimulating. I am fortunate to have an advisor who
gave me the freedom to explore on my own. His wide knowledge and logical way of
thinking have been a great value for me. His mentorship provides me a well rounded
experience both in academics and for my long-term career goal.
I would also like to thank my dissertation committee members, Professor
Dhandapani Venkataraman, Professor Michael Barnes and Professor T. J. Lakis
Mountziaris, for their constructive comments which helped me a lot to think hard on
research.
I am in debt to many members of the past and present Rotello groups. My
research for this dissertation was made more efficient because of the friendly
environment of the group and helpful discussion with the senior members (Yuval, Hao,
Pops, Bappa, Mrinmoy and Partha). My thanks must go to my batch mates in our group
(Oscar, Sarit and Apiwat) for their help, support and friendship during graduate study. I
also want to mention few names (Chandra, Subinoy, Krish, Myoung Yu Xi, Youngdo,
Yi-Cheun, Xiaoning) for having all the fun together in the lab.
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I would like to thank Department of Chemistry and all the staff members for their
enormous help and support during my graduate study. My special thanks go to JMS and
Carol; they made my life easy and smooth from the very first day in my graduate school.
Most importantly, I would like to express my heart-felt gratitude to my parents
who always motivated me to pursue higher studies. None of this would have been
possible without their love, support and patience. I also want to thank my sister for her
love and concern.
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ABSTRACT
COLLOIDAL MICROCAPSULES: SURFACE ENGINEERING OF NANOPARTICLES FOR INTERFACIAL ASSEMBLY
MAY 2011
DEBABRATA PATRA
B.Sc., MIDNAPORE COLLEGE, WB, INDIA
M.Sc., INDIAN INSTITUTE OF TECHNOLOGY, BOMBAY
Ph.D., UNIVERSITY OF MASSACHUSETTS AMHERST
Directed by: Professor Vincent M. Rotello
Colloidal Microcapsules (MCs), i.e. capsules stabilized by nano-/microparticle shells are
highly modular inherently multi-scale constructs with applications in many areas of
material and biological sciences e.g. drug delivery, encapsulation and microreactors.
These MCs are fabricated by stabilizing emulsions via self-assembly of colloidal
micro/nanoparticles at liquid-liquid interface. In these systems, colloidal particles serve
as modular building blocks, allowing incorporation of the particle properties into the
functional capabilities of the MCs. As an example, nanoparticles (NPs) can serve as
appropriate antennae to induce response by external triggers (e.g. magnetic fields or
laser) for controlled release of encapsulated materials. Additionally, the dynamic nature
of the colloidal assembly at liquid-liquid interfaces result defects free organized
nanostructures with unique electronic, magnetic and optical properties which can be
tuned by their dimension and cooperative interactions. The physical properties of
colloidal microcapsules such as permeability, mechanical strength, and biocompatibility
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can be precisely controlled through the proper choice of colloids and preparation
conditions for their
This thesis illustrates the fabrication of stable and robust MCs through via
chemical crosslinking of the surface engineered NPs at oil-water interface. The chemical
crosslinking assists NPs to form a stable 2-D network structure at the emulsion interface,
imparting robustness to the emulsions. In brief, we developed the strategies for altering
the nature of chemical interaction between NPs at the emulsion interface and investigated
their role during the self-assembly process. Recently, we have fabricated stable colloidal
microcapsule (MCs) using covalent, dative as well as non-covalent interactions and
demonstrated their potential applications including encapsulation, size selective release,
functional devices and biocatalysts.
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TABLE OF CONTENTS
Page ACKNOWLEDGEMENTS .................................................................................................v
ABSTRACT ...................................................................................................................... vii
LIST OF FIGURES ......................................................................................................... xiv
CHAPTER
1. NANOPARTICLES AT LIQUID INTERFACES: AN INTRODUCTION…………...1
1.1 Self-assembly of nanoparticles….………………………………………….…1
1.2 Assembly in bulk…..….....................................................................................2
1.3 Assembly at surfaces ………….……………………………………………...4
1.4 Assembly at interface …………..…………………………………………….7
1.5 Colloidal microcapsules …..……………………………………………….....7
1.5.1 Stability of the emulsion based template……………………………..8
1.5.2 Effective radius of the particle (r)…………………………………….8
1.5.3 Wettablity of NPs……………………………………………………10
1.6 Dissertation overview………………………………………………………..11
1.7 References……………………………………………………………………14
2. STABILIZATION OF MICROCAPSULES VIA DATIVE NANOPARTICLE CROSSLINKING………………………………………………………………………..19
2.1 Introduction ......................................................................................................19
2.1.1 Metal directed assembly of nanoparticles……..…………….………20
2.2 Results and discussion………………………………………………………...22
2.2.1 Fabrication of colloidal microcapsules via coordination chemistry at liquid-liquid interface..……………………………….…22
2.2.2 Optical characterization………………….………………………….24
2.2.3 Electron microscopy characterization .……………….......................24
2.2.4 Magnetic membrane under external magnetic field…………………25
2.2.5 Colloidal shells stability study………………….……………………25
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2.2.6 Size variation in FePt NP colloidal shells with varying Fe (II) salt concentration……………………………..……….......................................27 2.2.7 Versatility of the assembly………….……………...............................27
2.3 Conclusion………………………………………………………………………..28
2.4 Experimental section……………………………………………………………...29
2.4.1 Synthesis of ligands…………………………………………………..29
2.4.2 Synthesis of FePt NPs………………………………………………...30
2.4.3 Terpyridine functionalized FePt NPs…………………………………31
2.4.4 Uv-Vis characterization of Terpy functionalized NPs………..………31
2.4.5 Fabrication of FePt colloidal microcapsules………….………………31
2.5 References…………………………………………………………………….…32
3. NANOWIRE MICROCAPSULES: SELF-ASSEMBLY OF ANISOTROPIC NANOPARTICLE AT LIQUID-LIQUID INTERFCAE……………………………….35
3.1 Introduction………………………………………………………….................35
3.1.1 Hierarchical assembly strategies……………………………………..35
3.1.2 Assembly at liquid-liquid interface…………………………………..37
3.2 Results and discussion..…………………………………………………………39
3.2.1Metal mediated assembly of NWs……………………………………39
3.2.2 Characterization of MCs……………………………………………..40
3.2.3 Size variation of NWs MCs………………………………………….41
3.2.4 Electron transport properties of NWs MCs………………………….42
3.3 Conclusion ……………………………………………………………………. .45
3.4 Experimental Section …………………………………………………………...45
3.4.1 Fabrication of Au NWs ……………………………………………..45
3.4.2 Ligand exchange reaction of Au NWs………………………………46
3.4.3 Synthesis of MCs.................................................................................46
3.4.4 Characterization ..................................................................................46
3.4.5 Electronic transport measurement……………………………………47
3.4.8 Size Distribution of the MCs with increasing NWs concentration……47
3.4.9 Set up used for electronic transport measurement……………………..50
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3.5 References……………………………………………………………………….51
4. COLLOIDAL MICROCAPSULE: A VERSTAILE SCAFFOLD FOR MOLECULAR RECOGNITION AT LIQUID-LIQUID INTERFACE…………………………………54
4.1 Introduction ……………………………………………………………………54
4.1.1 Towards effective hydrogen bonding in aqueous media………………54
4.2 Results and discussion………………………………………………………….56
4.2.1 Fabrication of the template………….…………………………………56
4.2.2 Characterization of oil-in-water emulsions….…………………………58
4.2.3 “Host” encapsulation and “Host-Guest” recognition at interface……...58
4.2.4 Determination of “Host-Guest” binding constant.....................................60
4.3 Conclusion………………………………………………………………………62
4.4 Experimental Section...............................................................................................62
4.4.1 Synthesis of Dopamine-C8-TEI Ligand……………………………..…62
4.4.2 Synthesis of DAP amphiphile…………………………………………..65
4.4.3 Functionalization of FePt NPs………………………………………….66
4.4.4 Fabrication of colloidal microcapsule…………………………………..67
4.4.5 Size distribution of the microcapsules………………………………….67
4.5 References………………………………………………………………………68
5. HOST-GUEST CHEMISTRY INSIDE LIVING CELLS: RECOGNITION-MEDIATED ACTIVATION OF THERAPEUTIC GOLD NANOPARTICLES……71
5.1 Introduction……………………………………………………………………..71
5.1.1 “Host-guest” chemistry-A noncovalent approach of nanoparticle
self-assembly………………………………..………………………72
5.2 Results and discussion……………………………………………….…………...74
5.2.1“Host-guest” recognition mediated microcapsules fabrication liquid-liquid Interface………………………………………………………...…74 5.2.2 Characterization of microcapsules………………………………….75
5.2.3 Recognition induced disassembly…………………………………76
5.2.4 Size tuning of microcapsules………………………………………78
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5.3 Conclusions……………………………………………………………………..79
5.4 Experimental section…………………………………………………………....80
5.4.1 Chemicals…………………………………………………………...80
5.4.2 Synthesis of ADA-disulfide ligand………...……………………….80
5.4.3 Ligand exchange reaction…………………………...………………81
5.4.3 Microcapsule fabrication……………………………..……………..81
5.4.4 Microcapsule characterization…………………………..…………..81
5.4.5 Size distribution of the microcapsules with increasing concentration of
ADA-TEG-OH……………………………………….........................82
5.5 References……………………………………………………………………..85
6. STABLE MAGNETIC COLLOIDOSOMES VIA “CLICK” MEDIATED CROSSLINKING OF NANOPARTICLES AT OIL-WATER INTERFACE…………89
6.1 Introduction…………………………………………………………………….89
6.1.1 “Click” chemistry at oil-water interface..............................................90
6.2Results and discussion…………………………………………………………..91
6.2.1 Magnetic Colloidosomes……………………………………………..91
6.2.2 Characterization of colloidosomes.......................................................93
6.2.3 Permeability of colloidosomes.............................................................95
6.3 Conclusion………………………………………………………………………97
6.4 Experimental section……………………………………………………………98
6.4.1 Synthesis of NPs...................................................................................98
6.4.2 Synthesis of 11-azidoundecanoic acid…………………….………….98
6.4.3 Place Exchange Reactions.....................................................................99
6.4.4 Synthesis of colloidosomes..................................................................100
6.4.5 Characterization...................................................................................100
6.4.6 Permeability test..................................................................................101
6.5 References…………………………………………………………………….101
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7. SELF-ASSEMBLY OF ENZYME-NANOPARTICLE CONJUGATES AT LIQUID-LIQUID INTERFACE……………………………………………………………..105
7.1 Introduction…………………………………………………………………..105
7.1.1 NPs-Protein Conjugation…………………………………………..…106
7.2 Results and discussion………………………………………………………...108
7.2.1 NP-enzyme assembly at oil-water interface………………..…………108
7.2.2 Characterization of enzyme immobilized microcapsules……..……….110
7.2.3 Enzymatic activity of microcapsules……………………………..……112
7.3 Conclusion………………………………………………………………….…113
7.4 Experimental Section………………………………………………………….114
7.4.1 Microcapsule preparation……………………………………………...114
7.4.2 Transmission electron microscopy (TEM)…………………………….115
7.4.3 Quantification of enzyme in water after microcapsule fabrication…....115
7.4.4 Activity Assays.......................................................................................115
7.4.5 Synthesis of trimethylammonium-tetraethylene glycol functionalize ~7 nm Au nanoparticles………………………………………………115
7.4.6 Labeling of β-gal with FITC………..….………………………………116
7.4.7 ζ-Potential measurements…………..……….…………………………116
7.4.8 Measurement of surface charge density of nanoparticle, β-gal and enzyme -NP conjugate…….……………………………………………………..116
7.5 References……………………………………………………………………..117
BIBLIOGRAPHY………………………………………………………………….121
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LIST OF FIGURES
Figure Page
1.1: Strategies for self-assembly of nanoparticles…………………………………….2
1.2: (a) “Bricks and mortar” assembly of Au NPs and polymer. (b) TEM images of the aggregates. (c) Magnetic Nanoparticles Assembled with Ferritin via Electrostatic Interaction (d) TEM images of NP-Ferritin assembly . .......................................................................................................... …3
1.3: (a) Schematic of NPs used for electrostatic assembly (b) Au-Ag NPs macroscopic crystals formed after assembly. (c) Au NPs-DNA conjugate programmed for crystallization of NPs. ........................................... 4
1.4: (a) Drying mediated self-assembly of colloidal particles (b) TEM micrographs of AB2 superlattices made of 11-nm g-Fe2O3 and 6-nm PbSe NPs . .......................................................................................................... ………5
1.5: (a) DTC mediated binding of Au NPs on amine terminated SAM (b) Electrostatic LBL of polymer and NPs ................................................................ 6
1.6: (a) Schematic of size selective displacement from interface. Confocal Microscopy images of (b) 2.8 nm CdSe stabilized droplets (c) Droplets after addition of 4.6 nm CdSe NPs ...................................................................... 9
1.7: (a) NPs of different wettability at the interface (b) reversible attachment/detachment of Au NPs (c) 2-bromo-2-methylpropionate functionalized NPs. (d) Emulsion stabilized by Au NPs of intermediate wettability . ......................................................................................................... 11
1.8: Dissertation overview: Fabrication of NPs stabilized MCs and their potential applications.. ........................................................................................ 13
2.1: Metal Mediated coordination of Au NPs. TEM micrograph showing metal induced aggregation with (a) Ag (CH3CN)4BF4 and (b) Fe (H2O)6(BF4)2 ...... 21
2.2 TEM micrographs of (a) hexagonal patterns of of Terpy-functionalized AuNPs on PS domain in a PS-b-PMMA film. (b) Fe-treated crosslinked samples (c) ethanol-treated samples after swelling in the chloroform vapor. ..................................................................................................... ………22
2.3 Illustration of (a) FePt NP colloidal shells formation at water-in-toluene interface. Cross-sectional view of the colloidal shells represents NP network formed by complexation of the Terpy ligand with Fe (II)
xv
tetrafluoroborate hexahydrate salt. (b) FePt NP membrane formation at water-toluene interface. .......................................................................... ………23
2.4: Optical micrographs (OMs) of (a) FePt NP colloidal shells fabricated at water-in-oil interface with complexation agent, Fe(II) metal salt Inset: Pink-colored shell around water droplets indicates NP network formation [Conc. of Fe(II) salt ~ 100 µM, scale bar 50 μm] (b) Crumbled colloidal shells after partial drying. (c) FePt NP colloidal shells fabricated at water-in-oil interface without Fe (II) salt . .................................................................... 24
2.5: TEM micrograph of (a) FePt colloidal shell (b) magnetic membrane. Inset: High magnification image of the membrane shows close-packed FePt NPs.............................................................................................................. 25
2.6: Photograph of the (a) FePt membrane at water-oil interface (b) Membrane in toluene subjected to magnetic field ............................................................... 26
2.7: Optical micrographs of FePt NP colloidal shells at (a) room temperature (b) 50° C, 1 minute (c) 50° C, 2 minutes. ........................................................... 26
2.8: Optical micrographs of FePt NP colloidal shells with varying Fe(II) salt concentration (a) 100 mM (b) 10 mM (c) 1 mM ............................................... 27
2.9: (a) Optical micrograph of Au NP colloidal shells (b) Au NP membrane at water-toluene interface ....................................................................................... 28
2.10: Scheme of terpyridne thiol for functionalization of FePt NPs . ...................... 29
2.11: TEM micrograph of FePt (7±1 nm) NPs……………………………………...30
2.12: Absorption spectrum of (a) the terpyridine thiol ligand (b) Terpy-SH functionalized FePt nanoparticles (c) the FePt complex with Fe (II) metal ion ...................................................................................................................... 31
3.1: SEM images of (a) Au-Ppy rods. The bright segments are gold, and the dark domains are Polypyrrole (b) A SEM image of aggregates formed from Au-Ppy rods with a 3:2 block length ratio. (Inset) An optical microscopy image of a 3D tubular superstructure formed from these rods [Reproduced from reference 9] (c) LB technique for NWs assembly (d) microfluidics assembly of NWs. ......................................................................... 37
3.2: (a) Stability diagram of different shape particles at fluid interface (b) Stability order ................................................................................................... ..38
3.3: (a) OM images of Hairy” colloidosomes produced by transferring the microrod-coated agarose beads in water (b) Scanning Force Microscopy
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(SFM) phase images of the surface of dried emulsions prepared by tobacco mosaic virus (TMV) ............................................................................. 39
3.4: (a) Fabrication of NWs MCs (b) Terpy-SH functionalized Au NWs (c) Cross- sectional view of single MCs . ................................................................ 40
3.5: (a) Scanning Electron Microscope images of NWs (b) OM micrograph of NWs MCs; Inset showing crumpled MCs (c) Dark field OM micrograph of NWs MCs (d) TEM micrograph of dried MCs ............................................. 41
3.6: Change in average MCs diameter with varying NWs concentration. The inset represents the OM micrograph of largest and smallest size MCs (Scale ~ 200 µm). Curve is leading the eye. ..................................................... 42
3.7: (a) I-V characteristic curve of MCs at low voltage (-0.3V to +0.3V) (b) I-
V characteristic curve of MCs at high voltage (-1.5V to +1.5 volt) ................... 43
3.8: Hysteresis behavior of I-V characteristic curve of MCs at high voltage (-1.5V to +1.5 volt) sweep . ................................................................................... 43
3.9: (a) Fowler-Nordheim tunneling at higher bias (1V-1.5V). (b) Direct tunneling at lower bias (0.5V-1V). ..................................................................... 44
3.10: Size distribution of the NW MCs with at 0.333 mg/mL NWs concentration (mean diameter: 660 μm, SD: 197). ………………………………………………...47 3.11: Size distribution of the NW MCs with at 0.666 mg/mL of NWs
concentration (mean diameter: 228 μm, SD: 31). ............................................... 48
3.12: Size distribution of the NW MCs with at 1.332 mg/mL of NWs concentration (mean diameter: 117 μm, SD: 13) ................................................ 48
3.13: Size distribution of the NW MCs with at 2.664 mg/mL of NWs concentration (mean diameter: 65 μm, SD: 7) .................................................... 49
3.14: Size distribution of the NW MCs with at 5.328 mg/mL of NWs concentration (mean diameter: 37 μm, SD: 6). ................................................... 49
3.15: Set up chamber for conductivity measurement of the NWs MCs…………….50
4.1: (a) Binding constants between phosphate and guanidinium for various aqueous media interfaces: (a) in aqueous solution (b) on the surface of micelles and bilayers (c) at air-water interface ................................................... 56
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4.2: Orthogonal functionalization of FePt NPs and DTC chemistry for further crosslinking at oil-water interface . ..................................................................... 57
4.3: Self-assembly and crosslinking of FePt NPs at oil-water interface…………….57
4.4: OM micrographs of (a) FePt microcapsules at oil-in-water interface using DTC crosslinking strategy. Inset shows crumbled microcapsules. (b) Low magnification TEM images of dried microcapsules showing film like texture. Inset represents higher magnification TEM. .......................................... 58
4.5: (a) Flavin encapsulated microcapsule and “host-guest” recognition at oil-water interface (b) Fluorescence microscopy images of Flavin encapsulated micro capsule and after addition of excess DAP (c) Three point hydrogen bonding interaction between flavin and DAP ............................ 59
4.6: Fluorescence images of (a) N-Methyl flavin encapsulated micro capsule (b) After addition of excess DAP. All scales bar represents 40 μm of length................................................................................................................... 60
4.7: (a) Binding plot shows dramatic quenching of flavin polymer fluorescence upon addition of DAP (Ka ≈ 250 M-1) whereas control N-Methylated flavin does not show any fluorescence quenching. (b) Fluorescence titration of flavin polymer with variable DAP equivalents at the flat interface. ............................................................................................... ..61
4.8: Schematic representation of ligand exchange reaction………………………...66 5.1: (a) Layer-by-Layer Assembly scheme for the alternating adsorption of
Adamantyl-Terminated PPI Dendrimer and CD Au NPs onto CD SAMs (b) Phase transfer of hydrophobic NPs to water via surface modification using CD (c) Self-assembly of ADA stabilized NPs at the water/oil interface by the inclusion of ADA-capped NCs from an oil phase into CDs dissolved in water. ............................................................... 73
5.2: (a) “Host” functionalized Au NPs (b) “Guest” functionalized Au NPs (c) NP- NP assembly via “Host-Guest” recognition ................................................ 74
5.3: Schematic of fabrication of colloidal microcapsules ......................................... 75
5.4: Optical microscopy image of (a) colloidal microcapsule (b) Fluorescence microscopy image of dye encapsulated microcapsule (c) Low resolution TEM image of single microcapsule (d) High resolution TEM image of the microcapsule wall. .............................................................................................. 76
xviii
5.5: (a) Schematic of recognition induced disassembly. Photographs of glass vial representing (b) recognition induced self-assembly of NP1 and NP2 at toluene-water interface (c) recognition induced disassembly in presence of external “guest” (NP1). Plot of (c) plasmon absorption band of NP2 in presence of increasing ADA-TEG-OH concentration ....................................... 77
5.6: (a) Size tunability of the colloidal microcapsule at different ADA-TEG-OH concentrations in presence of ~ 0.4 uM of NP2 and ~ 0.4 uM of NP1 (b-e) Representative fluorescence microscopy images of different size microcapsules ..................................................................................................... 79
5.7: Size distribution of the microcapsule after adding 0 μmoles of ADA-TEG-OH (mean size: 5.815 μm, SD: 2.1)........................................................... 82
5.8: Size distribution of the microcapsule after adding 2.5 μmoles of ADA-TEG-OH (mean size: 18.045 μm, SD: 6.889)..................................................... 83
5.9: Size distribution of the microcapsule after adding 5 μmoles of ADA-TEG-OH (mean size: 30.88μm, SD: 14.2).......................................................... 83
5.10: Size distribution of the microcapsule after adding 7.5 μmoles of ADA-TEG-OH (mean size: 98.05 μm, SD: 34.92)....................................................... 84
5.11: Size distribution of the microcapsule after adding 10 μmoles of ADA-
TEG-OH (mean size: 178.71μm, SD: 82.265).................................................... 84
6.1: Basic schematic of “click” reaction ................................................................... 90
6.2: “Click” reaction in different environments (a) In solution (b) Solution–Surface (c) Reagent-Stamping (d) Stamp Catalyst ............................................. 91
6.3: (a) Formation of magnetic colloidosomes synthesis through self-assembly at water-in-oil interfaces followed by interfacial crosslinking of NPs via click chemistry (b) Structures of the NPs with ligands structure used for crosslinking and cross-sectional view of a colloidosome ................................... 92
6.4: Optical micrographs of (a) crosslinked magnetic colloidosomes fabricated at water-in-oil interface using click reaction (b) crumbled colloidosomes after partial drying of same colloidosomes (c) 0.1 mM catalyst concentration (d) no catalyst. .............................................................................. 93
6.5: (a) FTIR spectra of crosslinked NPs (green) and uncrosslinked control sample (red): relevant peaks are marked (b) Photograph of the magnetic colloidosomes in toluene subjected to magnetic field. TEM images of a colloidosome (c) Low magnification (d) high magnification. ............................ 94
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6.6: Schematic illustration of close-packing arrays of colloidosomes for calculating their interstitial dimension . .............................................................. 95
6.7: Fluorescence microscopy images of magnetic colloidosomes loaded with riboflavin in a) toluene, and b) after transfer into water. Similarly, Fluorescence images of colloidosomes loaded with FITC-dextran (Mw 500k g mol-1) c) in toluene and d) after 30 min in water .................................... 96
6.8: Size dependent release of fluoroscent molecules from colloidosome in 10 mM aqueous NaCl solution . .............................................................................. 97
6.9: TEM images of a) alkyne functionalized iron oxide NPs b) azide functiona- lized iron oxide NPs…………………................................................................100
6.10: Dynamic light scattering analysis of FITC-dextran polymers with Mw 40k g mol-1 (solid line) and 500k g mol-1 (dashed line). . ............................ ….101
7.1: (a) NTA-Ni2+ functionalized magnetic nanoparticles selectively bind to histidine-tagged proteins (b) Ligands used for CdSe nanoparticles, and schematic depiction of protein−nanoparticle interactions ............................... 107
7.2: (a) Interfacial assembly of polystyrene-conjugated ChT and β-galactosidase at toluene-water interface (b) Schematic representation of ChT- NPs complex at air–water interface ........................................................ 108
7.3: (a) Structure of β-gal. (b) Chemical structure of cationic gold nanoparticle. (c) Formation of enzymatic microcapsules through electrostatic assembly of enzymes and nanoparticles in water followed by assembly of resulting enzyme-nanoparticle conjugates at oil-in-water interfaces. Bottom left: A cross-sectional view of an enzymatic microcapsule .................................................................................................... 110
7.4: (a) Optical micrograph of enzyme-nanoparticle conjugate stabilized microcapsules fabricated at oil-in-water interfaces. (b) TEM images of a microcapsule at low magnification and high magnification (inset). (c) Fluorescence microscopy image of microcapsules synthesized using Au nanoparticles and FITC-labelled β-gal in water. d) Free β-gal present in water after microcapsule construction using various molar ratios of nanoparticles to β-gal. ....................................................................................... 111
7.5: (a) Free β-gal present in water after microcapsule construction using various molar ratios of nanoparticles to β-gal. (b) ζ-Potential measurements of enzyme-nanoparticle conjugates synthesized by varying molar ratios of nanoparticles to β-gal . ............................................................. 112
xx
7.6: Activity assay of 0.5 nM β-gal in 5 mM phosphate buffer (a) free enzyme; (b) enzyme-nanoparticle conjugate; (c) enzyme-nanoparticle conjugate stabilized microcapsules. Inset: relative activity (free enzyme solution = 100%). (d) Enzymatic cleavage of yellow colored CPRG at microcapsules shell by β-gal to produce red colored chlorophenol-red; Stepwise color changes (from yellow to red) in chemical reaction by β-gal present in the microcapsule shell: microcapsules are settled down at bottom of glass vial . ......................................................................................... 113
CHAPTER 1
NANOPARTICLES AT LIQUID INTERFACES: AN INTRODUCTION
1.1 Self-assembly of nanoparticles
In past decades, nanoparticles (NPs) are the focus of much attention due to their
astonishing properties and numerous possibilities for applications in nanotechnology.i
Assembling NPs generates unique nanostructures, which have surprising collective, intrinsic
physical properties.ii Currently new synthetic methods have been developed to synthesize NPs
with different materials, different sizes and shapes.iii However, the most important aspect is their
organization at nanoscale. For pragmatic applications and versatile functions, assembly of NPs in
regular patterns on surfaces and at interfaces is crucial. The multi-dimensional assembly of NPs
with controlled morphology in highly ordered arrays is important for realizing novel
applications.
There is an intense search for a simple and general strategy to organize NPs
through self-assembly processes. NPs self-assembly is very interesting as one has the possibility
of exploiting spatial or temporal attributes of NPs to tailor assembly formation.iv Such processes
require adequate stabilization of the NPs with a high degree of organizational selectivity.
Sufficient dynamic freedom is necessary in the course of the ordering process and once
completed, the assembly must be stabilized. Three different approaches are currently explored to
effect ordering of NPs (Figure 1.1) by the self-assembly processes such as (i) assembly in bulk
(ii) assembly at surfaces (iii) assembly at the interfaces.
1
Figure 1.1: Strategies for self-assembly of nanoparticles
1.2 Assembly in bulk
Self-assembly of NPs in bulk can lead to a 3D ordering of colloidal aggregates; which are
of great importance for the development of new materials with potential application in sensing,v
catalysis,vi and data storage.vii In this process, noncovalent interactions play an important role in
controlling the free energy and kinetics of the assembly and led to create hierarchical, complex
architectures. In this context, exploitation of the spontaneous and directed assembly of NPs with
synthetic macromoclecules [polymers,viii dendrimersix] and biomoleculesx is well investigated.
Our group pioneered the use of “bricks and mortar” assembly for the ordering of NPs into
structured assemblies. In earlier work, Rotello et al have demonstrated the assembly of thymine
functionalized Au NPs with diaminotriazine functionalized polystyrene via three point hydrogen
bonding interaction (Figure 1.2a).xi This method has shown to generate discrete assembly with
network structure (Figure 1.2b). In another work, our group has shown the fabrication of
bionanocomposites based on ferritin and magnetic FePt NPs (Figure 1.2 c).xii This assembly
2
process provided discrete essentially monodisperse aggregates that feature controlled
interparticle spacing (Figure 1.2d). Most significantly, this assembly strategy provides
nanocomposites where the structural components are also functional, yielding a highly modular
approach to biomagnetic materials.
Figure 1.2: (a) “Bricks and mortar” assembly of Au NPs and polymer. (b) TEM images of the
aggregates. (c) Magnetic Nanoparticles Assembled with Ferritin via Electrostatic Interaction (d)
TEM images of NP-Ferritin assembly. [Reproduced from reference 11 and 12]
One major disadvantage of the above strategy is the lack of long range order in the
assembled aggregates. Despite the promising attempts, periodic organization or crystallization at
the nanoscale was a big issue in those cases. Later, these challenges have been addressed by
controlled aggregation of NPs. Mirkin and coworkers have used DNA linked NPs to synthesize
NPs lattices.xiii They demonstrated that DNA guided self-assembly of Au NPs can produce
different crystalline states such as face-centred-cubic or body-centred-cubic crystal structures of
NPs (Figure 1.3a). Similarly, Grzybowski et al fabricate diamond like lattice via electrostatic
interaction of NPs.xiv In their study they used oppositely charged metallic NPs of similar size for
self-assembly (Figure 1.3b). Crystallization of NPs into a diamond-like structure (Figure 1.3c) is
mediated by screened electrostatic interactions. Screening occurs because (i) the NP cores are
3
metallic and (ii) each charged NP is surrounded by a layer of counter ions. As a result, the
particles interact by short-range electrostatic potentials.
Figure 1.3: (a) Schematic of NPs used for electrostatic assembly (b) Au-Ag NPs macroscopic
crystals formed after assembly. (c) Au NPs-DNA conjugate programmed for crystallization of
NPs. [Reproduced from reference 13 and 14]
1.3 Assembly at surfaces
. Drying-mediated or evaporation-induced assembly of NPs is the simplest method of
assembling particles on surfaces (Figure 1.4a).xv The use of self-assembly strategies based on the
dispersive attractions of NPs was realized to be an attractive “bottom-up” chemical approach to
their superstructures. An advantage of this approach is its simplicity: both 2D and 3D assemblies
can be prepared simply by placing a drop of a colloidal solution of monodisperse NPs on a
4
suitable support and allowing the solvent to evaporate slowly. Self-assembled colloidal crystals
can also be formed from semiconductor or magnetic metal nanoparticles by gentle destabilization
of the colloidal dispersions. Using this approach, Murray and co-workers reported the
development of NPs based binary superlattices.xvi In this case, precisely ordered, large single
domains of AB2 superlattices were obtained by simply allowing the NPs mixture (11 nm Fe2O3
NPs and 6 nm PbSe NPs) to evaporate overnight (Figure 1.4b).
Figure 1.4: (a) Drying mediated self-assembly of colloidal particles (b) TEM micrographs of
AB2 superlattices made of 11-nm g-Fe2O3 and 6-nm PbSe NPs. [[Reproduced from reference 15
and 16]
Another approach of assembling NPs on surface is chemically directed assembly.
xviii
xvii It is
a powerful self-assembly approach to create highly ordered, specific nanoparticle assemblies or
patterns. In this approach, one can exploit covalent as well as non-covalent interactions of NPs
surface protecting groups. Different techniques such as electrostatic layer-by-layer assembly,
chemical templating, and self-assembled monolayers (SAMs) are preferred modes in this
approach. For example, our group described a new approach for binding monolayers of NPs on
surfaces via dithiocarbamate (DTC) chemistry.xix An amine terminated SAM was converted to
5
DTC in the presence of CS
xxiii
xxvii
2 solution affording the in-situ capturing of Au NPs from the solution.
The primary amine was converted to DTC, which can then place exchange with the existing
ligands on Au-NPs, effectively binding the NPs to the surface (Figure 1.5a). Another interesting
approach to create nanostructure on surface is layer-by-layer assembly (LbL). Using the LbL
approach, numerous materials with optical and electronic properties have been fabricated
including light emitting diodes, single electron charging devices, and sensors.xx This
methodology has been successfully adopted to incorporate a wide range of materials such as
polyelectrolytes,xxi DNA,xxii proteins, dendrimers.xxiv Decher first reported the LbL approach
for assembly using polyelectrolytes,xxv while Iler described it for macroscale colloids.xxvi
Subsequently, Kotov et al. developed this technique for nanoparticle assembly (Figure 1.5b).
Figure 1.5: (a) DTC mediated binding of Au NPs on amine terminated SAM (b) Electrostatic
LBL of polymer and NPs. [Reproduced from reference 19 and 27]
6
1.4 Assembly at interface
An elegant approach to assemble NPs is based on interfacial assembly. Interface of oil-
water offer an important alternative scaffold for the organization of NPs. This is an easy and fast
pathway for large-scale applications. In past years, approaches have been developed to assemble
colloidal particles at emulsion interface for designing novel materials with wide varieties of
applications ranging from food, cosmetic and drug industries. However, NPs stabilized
emulsions or capsules are still in its early stage. In the sections below, we will emphasize on the
factors which governs the interfacial stability of NPs.
1.5 Colloidal microcapsules
Colloidal microcapsules (MCs) are an emerging class of microcapsules where colloidal
particles are used as building blocks for the MC shell, providing a useful alternative to polymer
and lipid-based systems.xxviii These MCs are fabricated by stabilizing emulsions via self-
assembly of colloidal micro/nanoparticles at liquid-liquid interfaces. In these systems, colloidal
particles serve as modular building blocks, allowing incorporation of the particle properties into
the functional capabilities of the MCs. As an example, NPs can serve as appropriate antennae to
induce response by external triggers (e.g. magnetic fields or laser) for controlled release of
encapsulated materials.xxix Additionally, the dynamic nature of the colloidal assembly at liquid-
liquid interfaces can be used to provide defect-free organized nanostructuresxxx with unique
electronic, magnetic and optical properties that can be tuned by their dimension and cooperative
interactions. The physical properties of colloidal microcapsules such as permeability, mechanical
strength, and biocompatibility can be likewise precisely controlled through the proper choice of
colloids and preparation conditions for their assembly. Over the last decade, research on
7
nano/micro particle stabilized emulsions has evolved significantly due to advancements in the
understanding of the interfacial assembly at both the micro- and nanoscopic level.xxxi A variety
of interesting strategies have been developed to stabilize NPs at the oil-water interface, thus
providing long-term stability to these colloidal assemblies.
1.5.1 Stability of the emulsion based template
Colloidal particles tend to localize at liquid-liquid interfaces to minimize the interfacial
energy between immiscible liquids, e.g. oil-water (o/w) emulsions.xxxii
xxxiii
This phenomenon of
stabilizing emulsions via colloidal particles was observed almost a century ago and termed as
“Pickering emulsions”, named after its discoverer S. U. Pickering. Later pioneering work by
Pieranski showed that the decrease in interfacial energy is related to the effective radius (r) of
particle and surface tension of the particle with respect to oil (O) and water (W). The
simplified equation is
( )[ ]2////
2
opwpwor
woE γγγγ
π −−−=∆
(1)
Where γp/o = interfacial energy of particle-oil, γp/w= interfacial energy of particle-water and γo/w
= interfacial energy of oil-water systems. The subsequent sections illustrate the role played by
the size and wettability of the colloidal particles in governing the dynamic behavior of the
colloidosomes as predicted by Pirenski equation.
1.5.2 Effective radius of the particle (r)
Particle stabilization of the liquid-liquid interface is strongly size-dependent: ΔE is
proportional to r2, hence the energy gain is smaller and the assembly is less stable for smaller
NPs than for larger ones. Larger micron sized particles can irreversibly adsorb at the interface
and make stable emulsions but smaller NPs (<20 nm) adsorb much more weakly at the interface.
8
As a result, thermal fluctuation can easily displace the NPs from the interface, leading to particle
exchange equilibrium at the interface and opens up the avenues to size selective particle
assembly (Figure 1.6a). Russell et al demonstrated the size selective adsorption of NPs at the oil-
water interface by using fluorescent CdSe quantum dots.xxxiv Initially smaller size CdSe NPs
(~2.8 nm, green fluorescence) were used to stabilize water-in-oil type emulsion. Later, they
introduced larger size CdSe NPs (~4.6 nm, red fluorescence) to the toluene phase. As a result,
larger CdSe NPs assembled on the surface of existing stabilized droplets, displacing the smaller
2.8 nm NPs (Figure 1.6b-c). This displacement is unique to NPs when thermal fluctuation is
dominant and provides a strong validation of the Pieranski equation.
Figure 1.6: (a) Schematic of size selective displacement from interface. Confocal Microscopy
images of (b) 2.8 nm CdSe stabilized droplets (c) Droplets after addition of 4.6 nm CdSe NPs.
Scale Bar, 20 µm. [[Reproduced from reference 34]
1.5.3 Wettablity of NPs
9
Another key parameter that governs the interfacial assembly of NPs is wettability.xxxv
The relation between contact angle and interfacial tensions can be derived by Young’s equation;
cos𝜃 = 𝛾𝑝𝑜 − 𝛾𝑝𝑤
𝛾𝑜𝑤 (2)
Where θ = contact angle. The affinity of the particles for the respective solvents is a significant
determinant, as indicated by γ
xxxvi
xxxvii
p/o and γp/w of above equation. Generally, hydrophilic NPs (γp/o>
γp/w, θ < 90°) stabilize oil-in-water emulsion and hydrophobic NPs (γp/o> γp/w, θ > 90°) are used
to stabilize water-in-oil emulsions (Figure 1.7a). Only particles with intermediate wettabilty (γp/o
= γp/w, θ = 90°), however, produce thermodynamically stable emulsions. Extending this concept,
Reincke et. al. showed that 4-mercaptobenzoic acid capped Au NPs (~ 8nm) can be attached to
and detached from the liquid/liquid interface and subsequently redispersed in the aqueous bulk
phase by changing the pH. When NPs were introduced to the water-heptane interface, no
interfacial attachment of the NPs was observed due to their extreme hydrophilic nature. After
adjusting the pH (~2) by addition of dilute HCl, Au NPs became protonated and slowly migrated
to the interface and formed a purple film (Figure 1.7b). The reversible nature of the interfacial
self-assembly was demonstrated by adjusting the aqueous solution pH to 9. As a result the color
of the aqueous solution turned red, similar to the original solution, confirming the detachment of
the Au NPs from the interface. The wettability of NPs can also be tuned by proper selection of
capping ligands. Surface modification of NPs by a 2-bromo-2-methylpropionate ligand
(Figure 1.7c) produces particles with a contact angle very close to 90° and as a result the NPs
stay exactly at the middle of the interface and effectively stabilize the emulsions droplets (Figure
1.7d). Recently, Russell and Emrick demonstrated that mixed-monolayer protected NPs can be
used as a tool to tune the interfacial interactions of the NPs to provide stable water droplets in
oil. Gold particles coated with near equimolar amounts of n-dodecanethiol and 11-mercapto-1-
10
undecanol was found to stabilize the emulsions, whereas excess of either ligands on the surface
did not result in stable emulsions. This also provides a direct evidence of the importance of the
ligand shell composition in tuning particle wettability to achieve stable assembly at the
interface.xxxviii
Figure 1.7: (a) NPs of different wettability at the interface (b) reversible attachment/detachment
of Au NPs (c) 2-bromo-2-methylpropionate functionalized NPs. (d) Emulsion stabilized by Au
NPs of intermediate wettability. [Reproduced from reference 36 and 37]
1.6 Dissertation overview
Beside the above factors, interactions between adsorbed particles at the liquid-liquid
interface can play a decisive role during the self-assembly process. Various effective interfacial
forces such as capillary forces, solvation forces, electrostatic force, van der Waals forces, and
fluctuation forces are responsible for manipulating interparticle interactions and represent a
current challenge in this area.xxxix In addition to these physical forces, chemical interactions
between the adsorbed particles are also crucial to stabilize MCs at the curved interface required
to fabricate MCs. To prevent thermally activated escape, particles must be trapped at the
interface during the process of self-assembly.
11
This thesis work focuses on directed self-assembly of NPs at liquid-liquid interface. My
research illustrated the fabrication of stable and robust MCs through via chemical crosslinking of
the surface engineered NPs at oil-water interface. The chemical crosslinking assists NPs to form
a stable 2-D network structure at the emulsion interface, imparting robustness to the emulsions.
In brief, I was involved to develop strategies for altering the nature of chemical interaction
between NPs at the emulsion interface and investigated their role during the self-assembly
process. Recently, we have fabricated stable colloidal microcapsule (MCs) using covalent, dative
as well as non-covalent interactions and demonstrated their potential applications including
encapsulation, size selective release, functional devices and biocatalysts.
Our group’s initial research effort in this area was focused on the use of dative
interactions to stabilize NPs at oil-water interfaces. In chapter 2, we created stable water-in-oil
emulsions and membranes by cross-linking NPs at the liquid-liquid interface using coordination
chemistry. In this strategy, terpyridine thiol (Terpy-SH) functionalized FePt NPs were self-
assembled at water-toluene interfaces and cross-linked through complexation of terpyridine with
Fe (II) metal ions. Later this strategy was extended to assemble one dimensional nano objects at
the toluene-water interface. According to the theoretical prediction, anisotropic NPs with an
aspect ratio α larger than critical a value are not stable at the interface. Using metal mediated
crosslinking, we were able to self-assemble Au NWs at the interface. In this case NWs formed an
extended 2D network at emulsion interface and their electric conductivity is discussed in chapter
3. Molecular recognition is one of the most important chemical events in biological systems
occurs mostly at microscopic interfaces such as membrane surfaces, enzyme reaction sites, or at
12
Figure 1.8: Dissertation overview: Fabrication of NPs stabilized MCs and their potential
applications.
the interior of the DNA double helix. Oil-water emulsions can offer a unique microenvironment
where the larger interfacial area of emulsions can act as suitable platform to localize the “host-
guest” pair. In Chapter 4 we designed permeable colloidal microcapsules where hydrophobic
media is confined inside colloidal capsules and can act as a suitable template to encapsulate the
“host” molecule for further “host-guest” molecular recognition at oil-water interface. Molecular
recognition also provides a noncovalent coupling between two complementary functionalized
surfaces. The ‘tailored interactions’ between building blocks via molecular recognition along
with the ‘dynamic nature’ of self-assembly process provides a promising tool to create novel
functional materials. We employ this concept to assemble NPs at the interface. Chapter 5
13
illustrated the fabrication of MCs using noncovalent interaction between NPs and their stimuli
responsive behavior towards external stimuli. MCs designed by previous approaches are not
stable enough to sustain in homogenous environment. It tends to collapse after removing the
interfacial barrier. Chapter 6 demonstrated the synthesis of mechanically stable water-in-oil
emulsions which can be transfer into water. These colloidosomes shows excellent size selective
permeability towards releasing ingredients.
Finally, biomolecular assembly has been addressed in Chapter 7. We have shown that
protein and NPs can be co-assembled at oil-in-water emulsion interface. A hybrid thin layer of
enzyme-NPs conjugates on a template with high surface to volume ratio provides an ideal
geometry for the generation of biocatalysts for industrial applications. The assembly of the
enzymes and nanoparticles both stabilizes the emulsion and retains the surface availability of the
enzymes for catalytic reaction. These microcapsules were formed quickly and showed high
enzymatic activity, making them promising materials for biotechnology applications.
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18
CHAPTER 2
STABILIZATION OF MICROCAPSULES VIA DATIVE NANOPARTICLE CROSSLINKING
2.1 Introduction
Interactions between adsorbed particles at the liquid-liquid interface can play a
decisive role during self-assembly process. Various effective interfacial forces such as
capillary forces, solvation forces, electrostatic force, van der Waals forces, and
fluctuation forces are responsible for manipulating interparticle interactions and represent
a current challenge in this area.1 In addition to these physical forces, chemical
interactions between the adsorbed particles are also crucial to stabilize particles at the
curved interface for further fabrication of MCs. To prevent thermally activated escape,
particles must be trapped at the interface during the process of self-assembly.
Recently, significant advances have been made in the area of colloidal
microcapsule engineering through chemical crosslinking of the particles assembled at the
oil-water interface.2 Crosslinking helps NPs to form a stable 2-D network structure at the
emulsion interface, imparting robustness to the MCs. The most straightforward way of
crosslinking the NPs at the oil-water interface is to link their surface capping ligands via
chemical bond formation. Despite the versatility of interfacial assembly strategies, the
creation of stable membranes and capsules remains a challenge.3 The competition
between the interfacial energy and spatial fluctuations resulting from NP thermal energy
causes instability in as-formed membranes and capsules. In recent years, many
approaches have been developed to fabricate stable membranes and colloidosomes using
NPs of various sizes.4 In their work, Lin et al., has fabricated a stable ultrathin CdSe
membrane at the interface by cross-linking the ligands of CdSe NPs5 while Duan et al.
19
prepared stable magnetic colloidosomes at the interface of water-in-oil droplets using
agrose gelated water phase.6 In both cases the initially formed membrane and
colloidosomes were subjected to thermal treatment (≥ 60° C) to afford stability and also
the cross-linking of the NPs at the interfaces was slow (~ 8 hrs). Both, time and
temperature of thermal treatment can limit the tunability and applicability of these
methods. To overcome this challenge, a metal mediated crosslinking of NPs has been
introduced for instantaneous formation of robust structures under ambient condition.
2.1.1 Metal directed assembly of nanoparticles
The assembly of NPs into defined 2-D or 3-D structures via nanoscale
engineering provide the access to nanocomposites featuring useful chemical,7 electronic,8
and physical properties.9 General approaches to noncovalent assembly strategies employ
van der Waals/packing interactions,10 hydrogen bonding,11 ion pairing,12 and host-guest
inclusion chemistry.13 These self-assembly methods offer direct access to extended
structures from appropriately designed NP building blocks. Metal-ligand systems provide
a means of expanding the structural diversity of self-assembly processes,14 as well as
imparting functional attributes such as redox15 and photochemical16 properties to the
resulting constructs. To explore the application of this methodology to nanocomposite
fabrication, our group previously studied metal directed self-assembly of gold NPs in
solution using a variety of transition metals.17 The assembly process occurred due to the
formation of coordination complexes between two Terpyridine ligands (Figure 2.1)
attached to separate NPs. The NPs aggregates were studied by Transmission Electron
Microscopy (TEM, Figure 2.1a-b). Tunability of the interparticle spacing had also been
demonstrated using different alkyl spacer. In another work, complex 2D NP arrays have
20
Figure 2.1: Metal Mediated coordination of Au NPs. TEM micrograph showing metal
induced aggregation with (a) Ag (CH3CN)4BF4 and (b) Fe (H2O)6(BF4)2. [Reproduced
from reference 17]
been created onto a particular domain of a block copolymer thin film.18 Shenhar et al
used polystyrene-b-poly (methyl methacrylate) (PS-b-PMMA) block copolymer as a
template for NPs patterning. Hydrophobic nature of Terpyridine (Terpy) functionalized
AuNPs directed them to localize at relatively non-polar PS domain. Later, crosslinking of
the assembled AuNP film was performed by immersing the sample into an ethanol
solution of [Fe (H2O)6 (BF4)2], resulting in the formation of coordianation complex
between Au NPs (Figure 2.2 a-c). These simple approaches of constructing robust NPs
superstructures would enable to manipulate discrete aggregates for the creation of large
scale assemblies at different length scale.
21
Figure 2.2: TEM micrographs of (a) hexagonal patterns of of Terpy-functionalized
AuNPs on PS domain in a PS-b-PMMA film. (b) Fe-treated crosslinked samples (c)
ethanol-treated samples after swelling in the chloroform vapor.
2.2 Results and discussion
2.2.1 Fabrication of colloidal microcapsules via coordination chemistry at liquid-
liquid interface
Here, we reported a novel route to fabricate stable colloidal shells by cross-
linking NPs at the liquid-liquid interface using coordination chemistry. In this strategy,
terpyridine thiol (Terpy-SH) functionalized FePt NPs are self-assembled at water-toluene
interfaces and cross-linked through complexation of terpyridine with Fe (II) metal ions.
The major advantage of this assembly process is the simultaneous and self-assembly of
the NPs at the interface. Oleic acid and oleyl amine stabilized FePt magnetic NPs (7.0 ± 1
nm) were synthesized following a literature procedure and then functionalized with
Terpy-SH ligand via place exchange reaction. Aqueous Fe(II) tetrafluoroborate
22
hexahydrate (1M) droplets were then added to a toluene dispersion of the functionalized
FePt magnetic NPs. Vigorous mixing for several seconds resulted in the formation of
stable emulsions (Figure 2.3a). A similar strategy was used to prepare FePt membranes at
the water-toluene interface. Toluene dispersions of FePt NPs were allowed to stand over
aqueous solution containing 1M Fe (II) tetrafluoroborate hexahydrate. The dark toluene
solution became colorless with a concomitant formation of a pink membrane at toluene-
water interface (Figure 2.3 b).
Figure 2.3: Illustration of (a) FePt NP colloidal shells formation at water-in-toluene
interface. Cross-sectional view of the colloidal shells represents NP network formed by
complexation of the Terpy ligand with Fe (II) tetrafluoroborate hexahydrate salt. (b) FePt
NP membrane formation at water-toluene interface.
23
2.2.2 Optical characterization
The colloidal shells formed by this technique were analyzed by Optical
Microscope (OM). The diameter of those capsules was 35–60 µm (Figure 2.5a). The
appearance of pink colored (λmax = 540 nm) NP shells (Figure 2.4a, inset) confirms metal
to ligand charge transfer (MLCT) complex formation. The NP-stabilized colloidal shells
are very stable; evaporation of water over a 5 day period resulted in crumpled but still
intact colloidal shells (Figure 2.4 b). On the other hand, stabilization of emulsion without
Fe (II) salt resulted in formation of unstable structures (Figure 2.4 c).
Figure 2.4: Optical micrographs (OMs) of (a) FePt NP colloidal shells fabricated at
water-in-oil interface with complexation agent, Fe(II) metal salt Inset: Pink-colored shell
around water droplets indicates NP network formation [Conc. of Fe(II) salt ~ 100 µM,
scale bar 50 μm] (b) Crumbled colloidal shells after partial drying. (c) FePt NP colloidal
shells fabricated at water-in-oil interface without Fe (II) salt.
2.2.3 Electron microscopy characterization
To visualize the nanoscopic structure, dried MCs were drop casted on TEM grid
for characterization. TEM image of the dried shells revealed that the wall of the colloidal
shells is composed of FePt NPs in a network structure (Figure 2.5 a). Similarly, TEM
analysis of the dried membrane, (Figure 2.5 b) shows thin-film morphology, while the
24
high magnification image (inset) clearly reveals that the membrane is composed of
densely-packed FePt nanoparticles.
Figure 2.5: TEM micrograph of (a) FePt colloidal shell (b) magnetic membrane. Inset:
High magnification image of the membrane shows close-packed FePt NPs.
2.2.4 Magnetic membrane under external magnetic field
Preliminary experiment indicates that FePt membrane can be manipulated using
external magnetic fields. In this study, the biphasic system was homogenized by addition
of methanol. The free-floating membrane could then be moved through application of a
magnetic field using a permanent magnet (Figure 2.6a). The fabrication of NPs
membrane by this technique is general and can be extended to other nanoparticulate
system; Terpy-SH functionalized Au NPs (2±1 nm) provided essentially identical
membranes. In addition, FePt NP colloidal shells can also be manipulated using magnetic
field. When subjected to permanent magnet we noticed aggregation of colloidal shells
towards the magnet (Figure 2.7b)
25
Figure 2.6: Photograph of the (a) FePt membrane at water-oil interface (b) Membrane in
toluene subjected to magnetic field.
2.2.5 Colloidal shells stability study
The thermal stability of FePt colloidal shells were tested by heat treatment at 50°
C. The colloidal shells were transferred to UV cuvette and immersed in a water-bath
maintained at 50° C for 2 minutes. We found no change in size or stability of the shells
up to 1 minute (Figure 2.7b) but crumpling of colloidal shells were noticed if the
temperature was maintained for two or more minutes. Though the colloidal shell
crumpled due to solvent evaporation but they remain discrete and no coalescence was
observed suggesting their robustness at elevated temperature (Figure 2.7c).
Figure 2.7: Optical micrographs of FePt NP colloidal shells at (a) room temperature (b)
50° C, 1 minute (c) 50° C, 2 minutes.
26
2.2.6 Size variation in FePt NP colloidal shells with varying Fe (II) salt
concentration
The size variation of colloidal shells was done by varying salt concentration. FePt
NP colloidal shells were prepared by addition of 15 μL of aqueous Fe(II)
tetrafluoroborate hexahydrate droplets to a toluene dispersion of Terpy-SH functionalized
FePt NPs. Varying the Fe(II) salt concentration enabled tunning the size of the colloidal
shells as evident from figure S2. The use of 100 mM, 10 mM and 1 mM Fe(II) salt yield
colloidal shells with size 70-85 μm, 135-150 μm and 190-200 μm, respectively (Figure
2.8 a-c).
Figure 2.8: Optical micrographs of FePt NP colloidal shells with varying Fe(II) salt
concentration (a) 100 mM (b) 10 mM (c) 1 mM.
2.2.7 Versatility of the assembly
The versatility of this strategy has been demonstrated using Au NPs. Terpyridine
functionalized Au NPs was synthesized using similar protocol like FePt NPs. In the next
step, water droplets [that contain Fe (II) tetrafluoroborate hexahydrate (1M)] were added
to a toluene dispersion of 2±1 nm Au NPs. Vigorous mixing for several seconds resulted
in the formation of stable emulsion and the water droplets in such emulsion were
stabilized by thin Au NPs resulting desired colloidal shells (Figure 2.9 a). The size of as
formed colloidal shells was found to be ~ 55-90 μm. Similarly, mauve-colored Au NP
27
membranes were formed at the water-toluene interface by allowing toluene dispersion of
Au NPs to stand over aqueous solution containing 1M Fe (II) salt. The complexation of
Fe (II) salt with terpyridine stabilized NPs result in nanoparticle network at the interfaces
(Figure 2.9b).
Figure 2.9: (a) Optical micrograph of Au NP colloidal shells (b) Au NP membrane at
water-toluene interface.
2.3 Conclusion
In summary, we have developed a simple strategy for the creation of highly stable
NP self-assemblies at liquid-liquid interfaces. Stable water-in-oil colloidal shells and
membranes were prepared using Terpy-SH functionalized FePt NPs, with cross-linking
effected at room-temperature through Fe (II) metal ion complexation. These systems
could be manipulated using external magnetic fields, providing a convenient and gentle
means of positioning these materials. Tuning of the assembly process and application of
this methodology to the creation of functional devices might be an important finding for
future applications. Moreover, these MCs can be used as a catalyst micro container for
chemical reactions.
28
2.4 Experimental section:
2.4.1 Synthesis of ligands
Ligands have been synthesized following a literature reported procedure. The
schematic of synthesis has shown below.19
Figure 2.10: Synthesis scheme of terpyridne thiol for functionalization of FePt NPs.
29
2.4.2 Synthesis of FePt NPs
The synthesis of FePt NPs was done following a literature reported procedure.20 In a
typical process, Pt(acac)2 (0.5 mmol) was dissolved in benzyl ether ( 10 mL) and heated
to 100 °C in nitrogen atmosphere. Nitrogen flow was maintained throughout the reaction.
At this temperature oleic acid (4.0 mmol), oleyl amine (4.0 mmol) and Fe (CO)5 were
added. The temperature of the resulting solution was increased to 240 °C and the reaction
mixture was incubated at this temperature for 1 h. Then the reaction mixture was heated
further to reflux (300 °C) condition for 2 h. It was later allowed to cool down to room
temperature and the nitrogen flow was disconnected at this point. The black colored
product was precipitated by adding 20 mL ethanol and separated via centrifugation. The
precipitate was then dispersed in hexane (10 mL) in the presence of oleic acid (0.05 mL)
and oleyl amine (0.05 mL). These particles were re-precipitated by using ethanol (15 mL)
and collected using centrifugation. Finally the purified product was re-dispersed in
hexane (10 mL) and analyzed by TEM (Figure 2.11)
Figure 2.11: TEM micrograph of FePt (7±1 nm) NPs
30
2.4.3 Terpyridine functionalized FePt NPs
20 mg of 7 nm FePt nanocrystals were dispersed in 10 mL of dichloromethane. To this 60
mg of terpyridine thiol ligands were added. The reaction mixture was stirred for
overnight under inert atmosphere. In the next step, the reaction mixture was concentrated
and ethanol was added to precipitate the NPs. This cycle was repeated for three times to
remove excess ligands from the reaction mixture. Finally NPs were dissolved in toluene
for further experiment.
2.4.4 Uv-Vis characterization of Terpy functionalized NPs
Figure 2.12: Absorption spectrum of (a) the terpyridine thiol ligand (b) Terpy-SH
functionalized FePt nanoparticles (c) the FePt complex with Fe (II) metal ion.
2.4.5 Fabrication of FePt colloidal microcapsules
500 µL of Terpyridine functionalized FePt NPs (5 mg/mL) were taken in an
eppendorf tube. To this 10-15 µL of Fe (II) tetrafluoroborate hexahydrate (1M) was
added. Vigorous shaking for 30 sec resulted in formation of stable emulsion. The
31
emulsion appeared to be pink in color and settled down at the bottom. Later, excess NPs
were washed for several times by toluene and MCs were stored at room temperature for
further analysis.
2.5 References: [1] Bresme, F.; Oettel, M.; J. Phys. Condens. Matter. 2007, 19, 413101.
[2] Boker, A.; He, J.; Emrick, T.; Russell, T. P., Soft Matter 2007, 3, 1231.
[3] (a) Kumar, A.; Mandal, S.; Mathew, S. P.; Selvakannan, P. R.; Mandale, A. B.;
Chaudhari, R. V.; Sastry, M., Langmuir 2002, 18, 6478. (b) Duan, H.; Wang, D.; Kurth,
D. G.; Möhwald, H., Angew. Chem., Int. Ed. 2004, 43, 5639.
[4] (a) Lin, Y.; Skaff, H.; Boeker, A.; Dinsmore, A. D.; Emrick, T.; Russell, T. P., J. Am.
Chem
.Soc. 2003, 125, 12690. (b) Duan, H.; Wang, D.; Sobal, N. S.; Giersig, M.; Kurth, D. G.;
Mohwald, H., Nano Lett. 2005, 5, 949-952.
[5] (a) Lin, Y.; Skaff, H.; Boeker, A.; Dinsmore, A. D.; Emrick, T.; Russell, T. P., J. Am.
Chem. Soc. 2003, 125, 12690.
[6] Duan, H.; Wang, D.; Sobal, N. S.; Giersig, M.; Kurth, D. G.; Mohwald, H., Nano Lett.
2005, 5, 949.
[7] (a) Templeton, A. C.; Wuelfing, W. P.; Murray, R. W. Acc. Chem. Res. 2000, 33, 27.
(b) (b) Schmid, G.; Baumle, M.; Geerkens, M.; Heim, I.; Osemann, C.; Sawitowski,
T. Chem. Soc. Rev. 1999, 28, 179.
[8] Ung, T.; Liz-Marzán, L. M.; Mulvaney, P. J. Phys. Chem. B 2001, 105, 3441.
[9] (a) Puntes, V. F.; Krishnan, K. M.; Alivisatos, A. P. Science 2001, 291, 2115. (b) Sun,
S.; Anders, S.; Hamann, H. F.; Thiele, J.-U.; Baglin, J. E. E.; Thomson, T.; Fullerton, E.
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E.; Murray, C. B.; Terris, B. D. J. Am. Chem. Soc. 2002, 124, 2884. (c) Pileni, M. P. J.
Phys. Chem. B 2001, 105, 3328.
[10] (a) Martin, J. E.; Wilcoxon, J. P.; Odinek, J.; Provencio, P. J. Phys.
Chem. B 2000, 104, 9475. (b) Fink, J.; Kiely, J.; Bethell, D.; Schiffrin, D. J.
Chem. Mater. 1998,10, 922. (c) Wang, Z. L.; Harfenist, S. A.; Whetten, R. L.; Bentley,
J.; Evans, N. D. J. Phys. Chem. B 1998, 102, 3068. (d) Beomseok, K.; Tripp, S. L.; Wei,
A. J. Am. Chem. Soc. 2001, 123, 7955.
[11] (a) Boal, A. K.; Gray, M.; IIhan, F.; Clavier, G. M.; Kapitzky, L.; Rotello, V. M.
Tetrahedron 2002, 58, 765. (b) Boal, A. K.; Rotello, V. M. Langmuir 2000, 16, 9527. (c)
Simard, J.; Briggs, C.; Boal, A. K.; Rotello, V. M. Chem. Commun. 2000, 1943.(d)
Frankamp, B. L.; Uzan, O.; IIhan, F.; Boal, A. K.; Rotello, V. M. J. Am. Chem.
Soc. 2002, 124, 892.
[12] (a) Kolny, J.; Kornowski, A.; Weller, H. Nano Lett. 2002, 2, 361. (b) Boal, A. K.;
Galow, T. H.; IIhan, F.; Rotello, V. M. Adv. Funct. Mater. 2001, 11, 1. (c) Hao, E.; Yang,
B.; Zhang, J.; Zhang, X.; Sun, J.; Shen, J. Chem. Mater. 1998, 8, 1327.
[13] (a) Liu, J.; Mendoza, S.; Román, E.; Lynn, M. J.; Xu, R.; Kaifer, A. E. J. Am. Chem.
Soc. 1999, 121, 4304. (b) Liu, J.; Alvarez, J.; Ong, W.; Kaifer, A. E. Nano. Lett. 2001, 1,
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[14] (a) Leininger, S.; Olenyuk, B.; Stang, P. J. Chem. Rev. 2000, 100, 853. (b) Fujita, M.;
Ogura, K. Acc. Chem. Res. 1999, 32, 53. (c) Piguet, C.; Bernardinelli, G.; Hopfgartner,
G. Chem. Rev. 1997, 97, 2005. (d) Lehn, J.-M. Supramolecular Chemistry: Concepts and
Perspectives; VCH: Weinheim, 1995; p 144−160.
33
[15] (a) Sauvage, J.-P.; Collin, J.-P.; Chambron, J.-C.; Guillerez, S.; Coudret, C.; Balzani,
V.; Barigelletti, F.; De Cola, L.; Flamigni, L. Chem. Rev. 1994, 94, 993.(b) Constable, E.
C. Cargill Thompson, A. M. W. J. Chem Soc.Dalton. Trans. 1994, 1409.
[16] Norsten, T. B., Chichak, K.; Branda, N. R. Tetrahedron 2002, 58, 639. (b) Cárdenas,
D. J.; Collin, J.-P.; Gaviña, P.; Sauvage, J.-P.; De Cian, A.; Fischer, J.; Armaroli, N.;
Flamigni, L.; Vicinelli, V.; Balzani, V. J. Am. Chem. Soc. 1999, 121, 5481.
[17] Norsten, T. B.; Frankamp, B. L.; Rotello, V. M., Nano Lett. 2002, 2, 1345-1348.
[18] Shenhar, R.; Jeoung, E.; Srivastava, S.; Norsten, T. B.; Rotello, V. M., Adv. Mater.
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Rotello, V. M. J. Am. Chem. Soc. 2007, 129, 11776-11780. (b) Ofir, Y.; Samanta, B.;
Arumugam, P.; Rotello, V. M., Adv. Mater. 2007, 19, 4075-4079.
34
CHAPTER 3
NANOWIRE MICROCAPSULES: SELF-ASSEMBLY OF ANISOTROPIC NANOPARTICLE AT LIQUID-LIQUID INTERFCAE
3.1 Introduction
The significant development in nanotechnology over the past years has led to the
building of powerful device or functional system, ranging from nanoscale transitors,1
light-emitting diodes (LEDs),2 nanolasers3 and chemical/biological sensors.4 The
“bottom-up” self-assembly of nanomaterials has the great promise over traditional “top-
down” approach, leading to designing novel functional systems. The first and important
area is the synthesis of nanoscale building blocks with precisely controlled and tunable
chemical composition, structure, size, morphology, and correspondingly defined
electronic, optical/magnetic properties.5 Secondly, hierarchical assembly strategies must
be developed to organize those building blocks into highly integrated nanosystems with
predictable and versatile functions. In recent years, one-dimensional (1-D) nanostructures
(such as wires, rods, tubes, plates) have attracted considerable attention due to their size
and shape dependent physical properties (e.g. electronic, optical and mechanical
properties) relative to zero-dimensional nanostructures (nanoparticles or quantum dots).6
Considerable progress has been made in fabrication of metallic/semiconductor
nanowires7 and nanorods8 of different aspect ratios; however the hierarchical
organization of these nanoscale building blocks into 2D or 3D assemblies still remains a
challenge.
3.1.1 Hierarchical assembly strategies
The development and implementation of efficient and scalable strategies for
assembly and controlled organization of 1D nanostructure are critical to both
35
fundamental studies and for real world applications. The assembly approaches should be
able to hierarchically organize nanoparticles on multiple length scales and to make well-
defined interconnections between nano/micro and macroscopic worlds. Efforts have been
made to assemble nanowires (NWs) and nanotubes (NTs) in solution and on surfaces. In
their recent effort, Mirkin et al investigated the assembly of metal-polymer
multicomponent (Au-polypyrrole) building blocks.9 These structures behave like
amphiphiles and form superstructures consisting of bundles, tubes and sheets depending
upon the compositional periodicity for example rods with a 1:4 gold/Ppy ratio self-
assembled to form tubular structures (Figure 3.1a-b). Another approach of assembling
NWs on large scale with nanometer level separation is Langmuir-Blodgett (LB)
technique.10 In this case, an ordered monolayer of NWs is formed at air-water interface
and compressed on an aqueous sub phase to yield uniaxially aligned NWs on solid
support (Figure 3.1c). Microfluidic device has also been used to align NWs by passing
them through microfluidic channels.11 The shearing force along the flow direction
aligned the NWs and the density of NWs was controlled by concentration of NWs.
Complex architectures has been can also be formed through a layer-by-layer process.
(Figure 3.1d)
36
Figure 3.1: SEM images of (a) Au-Ppy rods. The bright segments are gold, and the dark
domains are Polypyrrole (b) A SEM image of aggregates formed from Au-Ppy rods with
a 3:2 block length ratio. (Inset) An optical microscopy image of a 3D tubular
superstructure formed from these rods [Reproduced from reference 9] (c) LB technique
for NWs assembly (d) microfluidics assembly of NWs. [Reproduced from reference 10
and 11]
3.1.2 Assembly at liquid-liquid interface
In this context, liquid-liquid interface provide a versatile platform for assembly of
anisotropic nanoparticles. Integrating nanocrystals into 2D assemblies gives rise to novel
multiscale materials with unique electronic, magnetic and optical properties that can be
tuned by their dimension and cooperative interactions.12 Most efforts in the area of liquid
interface assembly of nanomaterials have focused on assembly of spherical/low-aspect-
ratio colloidal particles.13 The assembly of non-spherical/high-aspect-ratio particles
represents a major challenge in the area of interfacial assembly.14 Thermodynamic
calculation by Faraudo et al15 shows the impact of particle geometry on free energy of
nanoparticles with same surface area but different shapes at liquid/liquid interfaces.
According to this model, the stability of NPs at fluid interface is determined by particle
geometry, particles with an aspect ratio α larger than a critical value are not stable at the
interface. Figure 3.2a shows the impact of particle geometry on the free energy of three
37
Figure 3.2: (a) Stability diagram of different shape particles at fluid interface (b)
Stability order [Reproduced from reference 15]
particles with same surface area but different shapes. According to this model, disk-
shaped particles would be the most stable, spherical particles will be metastable while
rod-shaped particles are quite unstable at the interface. Two approaches of interfacial
nanorod assembly have recently been reported. Paunov et al16 synthesized “hairy”
colloidosomes with a shell of polymeric rods, using gelation of the aqueous core to
stabilize the assembly (Figure 3.3a). In another example, He et al studied the self-
assembly and orientation of cylindrical virus particle17 and CdSe nanorods18 at the oil-
water interface, however the emulsions were structurally unstable (Figure 3.3b).
38
Figure 3.3: (a) OM images of Hairy” colloidosomes produced by transferring the
microrod-coated agarose beads in water (b) Scanning Force Microscopy (SFM) phase
images of the surface of dried emulsions prepared by tobacco mosaic virus (TMV)
[Reproduced from reference 16 and 17 respectively].
3.2 Results and Discussion
3.2.1 Metal mediated assembly of NWs
We developed here a facile approach for constructing stable colloidal
microcapsules by simultaneous self-assembly and crosslinking of Au NWs at oil-water
interfaces using coordination chemistry. In this strategy, terpyridine thiol (Terpy-SH)
functionalized Au-NWs are self-assembled at water-toluene interfaces and crosslinked
through complexation of terpyridine with Fe (II) metal ions (Figure 3.4a-c). The Au-NWs
(length ~ 3-5 µm and width ~ 250-300 nm; aspect ratio ~ 10-20; Figure 3.5a) used in this
study were synthesized following a literature reported procedure,19 and functionalization
with Terpy-SH was done by ligand exchange reaction.20 In the next step NWs were
transferred into 300 μL of toluene (continuous phase), and 10 µL of 1 (M) aqueous
solution of Fe (II) tetrafluoroborate hexahydrate (dispersed phase) was added. Vigorous
mixing by an amalgamator for 10 sec (~ 4000 rpm) resulted in formation of stable
emulsions (Scheme 1). The concomitant appearance of the pink color emulsions (Figure
3.5b) confirms the formation of metal-ligand charge transfer complex between Terpy and
Fe (II) salts as well as the crosslinking of NWs at the toluene-water interface.
39
Figure 3.4: (a) Fabrication of NWs MCs (b) Terpy-SH functionalized Au NWs (c)
Cross- sectional view of single MCs.
3.2.2 Characterization of MCs
Emulsions were then analyzed by optical microscopy (OM); the diameter of the
microcapsules ranged from 100-150 μm (Figure 3.5b). These emulsions are stable for 7-
10 days at room temperature, with slow evaporation of water resulting in crumbled
microcapsules (Figure 1b inset) as opposed to MC disruption. Dark field optical
microscopy images revealed that MCs were cover by Au NWs (Figure 3.5c). To visualize
the microstructure, emulsions were drop casted on carbon coated copper grids and dried
completely. Transmission Electron Microscopy (TEM) images confirmed that MCs were
composed of densely packed Au-NWs in a network structure (Figure 3.5d). On the other
hand, no droplets formed without Fe (II) in the toluene dispersion of NWs immediately
agglomerated, indicating that a stable and rapid crosslinking is necessary to interconnect
and stabilize NWs at the liquid-liquid interface.
40
Figure 3.5: (a) Scanning Electron Microscope images of NWs (b) OM micrograph of
NWs MCs; Inset showing crumpled MCs (c) Dark field OM micrograph of NWs MCs (d)
TEM micrograph of dried MCs.
3.2.3 Size variation of NWs MCs
The capsule diameter distribution of Pickering emulsions can be effectively varied
by adjusting the concentration of the particles while keeping the relative proportion of oil
and water constant. To assess the role of particle concentration on the size of the MCs,
we made five sets of samples with increasing NW concentration. In the next step, 300 µL
toluene dispersion of NWs were mixed with 15 µL of 1M Fe (II) solution and the
heterogeneous mixture emulsified using an amalgamator (~4000 rpm) for 10 sec. The
resultant emulsions were examined by OM and the size of the MCs was determined from
OM micrographs (see SI). In all cases, stable emulsions were formed, with the average
droplet diameter decreasing with higher NW concentration (Figure 3.6).
41
Figure 3.6: Change in average MCs diameter with varying NWs concentration. The inset
represents the OM micrograph of largest and smallest size MCs (Scale ~ 200 µm). Curve
is leading the eye.
3.2.4 Electron transport properties of NWs MCs
The electron transport properties of NWs MCs were measured using a two-probe
electrode configuration in which a bias voltage (Vb) is applied across two electrodes
attached to sample and the resulting current (I) measured. Electronic transport
measurements revealed that MCs are conductive over 50 µm gap electrodes. At low
voltages (-0.3V to +0.3V), linear I-V response was observed with conductance 8.89 ±
0.95 µS (1.7x102-fold higher than the background toluene) (Figure 3.7a). A hysteresis
behavior was observed in the reverse scan (Figure 3.8) presumably due to to Joule
heating and concomitant loss of ligand.21 I-V curves were obtained in the voltage range
of -1.5 V to +1.5V to investigate the mechanism of conduction. The curve followed linear
+ cubic barrier-bending (Figure 3.7 b) behavior expected for electron tunneling across
alkanethiol ligands.22 Based on applied bias, the tunneling through alkanethiol ligands
can be categorized into (a) direct (V<φB /e) and (b) Fowler-Nordheim (V> φB /e) through-
bond tunneling [where V is applied bias and φB is barrier height]. For direct tunneling,
42
Figure 3.7: (a) I-V characteristic curve of MCs at low voltage (-0.3V to +0.3V) (b) I-V
characteristic curve of MCs at high voltage (-1.5V to +1.5 volt).
Figure 3.8: Hysteresis behavior of I-V characteristic curve of MCs at high voltage (-
1.5V to +1.5 volt) sweep.
43
current scales linearly with voltage at low bias (as observed in our case, Figure 3.9 a)
whereas for Fowler-Nordheim tunneling, ln(I/V2) varies linearly with 1/V. Analysis of
ln(I/V2) versus 1/V in (Figure 3.9b) shows no significant voltage dependence, indicating
no obvious Fowler-Nordheim transport behavior in this bias range (0 to1.0 V) and thus
determining that the barrier height is larger than the applied bias: i.e., φB > 1.0 eV. The
transition from direct to Fowler-Nordheim through-bond tunneling was observed at
higher bias (1-1.5V) where linear relation of ln(I/V2) versus 1/V was observed. Therefore,
we conclude that the conduction mechanism through the nanowire microcapsule is
metallic conduction through the wires and tunneling of electrons from metal to ligands to
metal junctions. Direct tunneling was the conduction mechanism at low bias (< 1V) while
the transition from direct to Fowler-Nordheim through-bond tunneling was observed at
higher bias.
Figure 3.9: (a) Fowler-Nordheim tunneling at higher bias (1V-1.5V). (b) Direct
tunneling at lower bias (0.5V-1V).
44
3.3 Conclusion
In summary, we have developed a straightforward approach to self-assemble Au
NWs at liquid-liquid interface. Metal mediated crosslinking of NWs at the interface
provides stability to the MCs. The size variation of MCs has been demonstrated with
varying NWs concentration. We have also shown that our approach is highly attractive
for creating integrated and functional assemblies for nanoelectronics and device
applications. The electron transport measurement revealed that NWs MCs are
electronically conductive due to their interconnected network structure. Moreover, for the
future work, conductivity of the MCs can be tuned by fine tuning the assembly process
such as increasing the density of NWs on each MCs, mixing and matching with different
types of nanoparticles (metallic, semiconductor and insulator).
3.4 Experimental Section
3.4.1 Fabrication of Au NWs
A typical nanowire fabrication process is described as below. First, 200 nm thick
silver (Ag) was evaporated onto one face of an anodized aluminum oxide (AAO)
membrane (purchased from Whatman). The membrane was then placed in contact with a
copper plate and restrained by a glass joint with an O-ring. Before filling the electrolytic
solution into the glass joint, the AAO membrane was soaked by a small amount of water
for 5 min to pre-wet the membrane pores and then the water was got rid of. Before gold
nanowire plating, a sacrificial silver layer was plated first to improve the quality of the
gold nanowires. Silver and gold plating solutions were purchased from Technic, Inc. and
were utilized without further treatment. The length of the nanowires was controlled by
the duration of the applied current. Typical plating conditions were 0.8 to1.2 mA/cm2.
After electroplating, the Ag on the side of the AAO membrane was dissolved by 6 M
45
HNO3 solution. After washing HNO3 residue with water, the membrane was dissolved in
2M NaOH solutions to release the nanowires into suspensions. The nanowires were
rinsed repeatedly with water and ethanol (using sonication and centrifugation cycles) and
then stored in ethanol for further use or characterization.
3.4.2 Ligand exchange reaction of Au NWs
10 mg of the NWs was dispersed in 2 mL of DCM with 30 mg of Terpy-SH. The
reaction mixture was sonicated for 30 mins and stirred for overnight at room temperature
under argon atmosphere. Purification of the NWs was achieved by centrifugation and
redispersing in DCM. This step was done 3-4 times to remove excess ligands. Later
Terpy-SH functionalized Au NWs were dispersed in toluene.
3.4.3 Synthesis of MCs
300 µL of NWs dispersion in toluene (stock concentration 1.332 mg/mL) was
taken in an eppendorf tube. In the next step, 10 µL of 1 (M) aqueous solution of Fe (II)
tetrafluoroborate hexahydrate was added. Vigorous shaking was done by commercially
available amalgamator for 10 sec (~ 4000 rpm). After shaking, NWs segregated to the
water-oil interface to create microcapsule and appeared as pink colour dispersion, which
eventually settles down to the bottom of the organic solvents. Excess NWs were removed
by washing with toluene prior to any characterization.
3.4.4 Characterization
All the MCs characterization was done by Optical Microscope (Olympus IX51
microscope). NWs MCs were transferred to a cuvette filled with 200 µL of toluene for
imaging. For dried samples, MCs were drop casted on glass surface and dried completely
prior to imaging.
46
3.4.5 Electronic transport measurement
50 µl of NWs MCs were dropped on the electrode gap using a micropipette and
waited for 10-15 minutes to settle down before applying any voltage. Then, voltage was
applied across the gap using Keithley 2400 source meter. The electronic current
therefore passed through the gold electrodes, alkanethiol ligands and the gold nanowires
in the microcapsule. Voltage was scanned from -1.5V to +1.5V in steps of 0.1 V, then
from +1.5V to -1.5V. For each measurement, current was measured 10 s after setting the
voltage to allow the exponential decay of the transient ionic current in the gap and to
measure steady-state electronic current. Igor Pro (Wavemetrics Inc. OR USA) was used
for data fitting and analysis.
3.4.8 Size Distribution of the MCs with increasing NWs concentration
400 500 600 700 800 900 10000
1
2
3
4
5
6
7
Diameter of the MCs (µm)
Coun
t
Count
Figure 3.10: Size distribution of the NW MCs with at 0.333 mg/mL NWs concentration
(mean
diameter: 660 μm, SD: 197).
47
180 200 220 240 260 280 3000
1
2
3
4
5
6
7
Diameter of MCs (µm)
Coun
t
Count
Figure 3.11: Size distribution of the NW MCs with at 0.666 mg/mL of NWs
concentration (mean diameter: 228 μm, SD: 31).
80 90 100 110 120 130 140 150 1600
2
4
6
8
10
12
14
16
Diameter of MCs (µm)
Coun
t
Count
Figure 3.12: Size distribution of the NW MCs with at 1.332 mg/mL of NWs
concentration (mean diameter: 117 μm, SD: 13).
48
40 45 50 55 60 65 70 75 800
2
4
6
8
10
12
14
Diameter of MCs (µm)
Coun
t
Count
Figure 3.13: Size distribution of the NW MCs with at 2.664 mg/mL of NWs
concentration (mean diameter: 65 μm, SD: 7).
20 25 30 35 40 45 50 55 600
2
4
6
8
10
12
14
16
Diameter of MCs (µm)
Coun
t
Count
Figure 3.14: Size distribution of the NW MCs with at 5.328 mg/mL of NWs
concentration (mean diameter: 37 μm, SD: 6).
49
3.4.9 Set up used for electronic transport measurement
Electronic transport measurements of nanowire microcapsules were performed
using a setup shown in Fig. S6. To construct the electrodes, glass slides (12.7 mm X 10
mm) were cleaned ultrasonically using successive rinses of trichloroethylene, acetone,
and methanol and then blown dry with nitrogen. A 40 nm Au film atop a 10 nm Cr
adhesion layer was thermally evaporated on these substrates, at 10-6 mbar, at a deposition
rate 0.1 nm/s producing gold electrodes with a 50 µm non-conductive spacing using
standard photolithography processing. Optical microscopy revealed that the gap was
uniform, and resistance measurements assured that the electrodes were well insulated
from each other. Platinum wire was used for electrical connections. Methanol fuel cell
stacks (www.fuelcellstore.com) were used to construct the 7ml chamber (2.9 cm on each
side and 0.85 cm deep). Gaskets were butyl rubber. The chamber was filled with toluene
for all the time during the measurements. All measurements were performed under
ambient condition.
Figure 3.15: Set up chamber for conductivity measurement of the NWs MCs
50
3.5 References
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Jana, N. R.; Angew. Chem. Int. Ed. 2004, 43, 1536. (d) Wang, Z. L.; J. Mater. Chem.,
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(f) Kottmann, J. P.; Martin, O. J. F.; Smith, D. R.; Schultz, S., Chem. Phys. Lett. 2001,
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[7] (a) Foss, C. A..; Hornyak, G. L.; Stockert, J. A.; Martin, C. R., J. Phys. Chem. 1994,
98, 2963. (c) Yu, Y. Y.; Chang, S. S.; Lee, C. L.; Wang, C. R. C., J. Phys. Chem. B 1997,
101, 6661. (d) Pileni, M. P.; Ninham, B. W.; Gulik-Kryzwicki, T.; Tanori, J.; Lisiecki, L.;
Filankembo, A., Adv. Mater. 1999, 11, 1358. (e) Peng, X. G.; Manna, L.; Yang, W. D.;
Wickham, J.; Scher, E.; Kadavanich, A.; Alivisatos, A. P., Nature 2000, 404, 59. (f)
Filankembo, A.; Pileni, M. P., J. Phys. Chem. B 2000, 104, 5865. (g) Li, M.;
Schnablegger, H.; Mann, S., Nature 1999, 402, 393. (h) Duan, X.; Lieber, C. M., Adv.
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[8] a) Xia, Y.; Yang, P.; Sun, Y.; Wu, Y.; Mayers, B.; Gates, B.; Yin, Y.; Kim, F.; Yan,
H., Adv. Mater. 2003, 15, 353.
[9] Park, S.; Lim, J. H-.; Chung, S. W-.; Mirkin, C. A., Science, 2004, 303, 348.
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Goettmann, F.; Moores, A.; Floch, P.; Sanchez, C., J. Mater. Chem. 2007, 17, 3509. (c)
Binks, B. P.; Horozov, T. S., “Colloidal Particles at Liquid Interfaces” Cambridge
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Faraudo, J., J. Phys. Condens. Matter. 2007, 19, 375110.
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53
CHAPTER 4
COLLOIDAL MICROCAPSULE: A VERSTAILE SCAFFOLD FOR MOLECULAR RECOGNITION AT LIQUID-LIQUID INTERFACE
4.1 Introduction
Molecular recognition is one of the most important chemical events in biological
systems and is achieved through combined noncovalent interactions such as hydrogen
bonding, electrostatic interaction or hydrophobic interaction.1 The directionality in
intermolecular interaction is critical for specific molecular recognition between “host”
and “guest” pair. Hydrogen bonding, in particular, is highly directional compare to other
noncovalent interactions and plays decisive roles in biological system such as replication
of nucleic acids, maintenance of the tertiary structure of proteins and substrate
recognition of enzymes.2 Molecular recognition in living systems occurs mostly at
microscopic interfaces such as membrane surfaces, enzyme reaction sites, or at the
interior of the DNA double helix. In recent years, efforts have been made to understand
the basic concepts of biological systems and designing novel functional materials.3
Unlike biological systems, most of the artificial molecular recognition mediated
by hydrogen-bonding interactions takes place effectively in nonaqueous media due to the
high dielectric nature of water.4 To enhance the effectiveness of hydrogen bonding in
water, recent efforts have been made to create hydrophobic environment of different
dimensions in the vicinity of the “host-guest” functional pair.5
4.1.1 Towards effective hydrogen bonding in aqueous media
Several strategies have been explored for inducing effective hydrogen bonding
between artificial receptors in contact with aqueous media. One elegant approach is
creating unique microenvironment in the vicinity of “host-guest” pair. One may create
54
hydrophobic environments at three levels of dimensions (i) microscopic [molecular level]
(ii) mesoscopic (iii) macroscopic. In one approach, Rebek et al6 designed receptor
molecules that are capable of hydrogen bond mediated molecular recognition in aqueous
media. The hydrophobic contact between receptor and guest enhances the binding
efficiency and hydrogen bonding is made effective in such microscopic hydrophobic
environments. Hydrogen bonding interaction becomes more effective when the “host-
guest” pair is placed in a mesoscopic phase where water is not readily accessible. Nowick
et al7 showed binding of adenine and thymine moieties by burying them in the
hydrophobic core of aqueous micelles. In macroscopic regime, hydrogen bonding
interaction is made effective by placing host-guest functional pairs close to that of bulk
organic media. The interfacial molecular recognition can offer a unique scenario compare
to bulk media. The interface may be mesoscopic like surfaces of molecular aggregates
(micelles and bilayers) or it may be macroscopic like solid substrates or the air-water
interface. In his pioneer work, Kunitake et al compared the binding efficiency between
guanidium and phosphate in three different environments. The binding constant of
adenosine monophosphate (AMP) to guanidium functionality in micelle/bilayer vesicle
were determined at 102-104 M-1 (Figure 4.1a) where as moleculary dispersed guanidium
and phosphate in water was 1.4 M-1 (Figure 4.1b). Interestingly, a large enhancement in
binding constant was reported for binding of AMP to guanidinium groups when
embedded at a water surface. These results clearly indicate that molecular recognition can
be achieved much more efficiently at an appropriate interface. In this context, oil-water
emulsions can offer a unique microenvironment8 where the larger interfacial area of
emulsions can act as suitable platform to localize the “host-guest” pair. To encapsulate
the “host” molecule for further recognition at the interface, a stable emulsion is highly
55
desired. Microcapsules such as polymeric microcapsules,9 vesicles,10 colloidal
microcapsules11 can act as a suitable candiate due to their structural stability. Amongst,
colloidal microcapsules feature great advantages due to their easy fabrication, enhanced
mechanical stabilty and controlled permeability.12
Figure 4.1: (a) Binding constants between phosphate and guanidinium for various
aqueous media interfaces: (a) in aqueous solution (b) on the surface of micelles and
bilayers (c) at air-water interface. [Reproduced from reference 7]
4.2 Results and discussion 4.2.1 Fabrication of the template
Here, we employ a new approach to fabricate permeable colloidal microcapsules
where hydrophobic media is confined inside colloidal capsules and can act as a suitable
template to encapsulate the host molecule for further molecular recognition. Moreover,
these oil-in-water based emulsions are compatible with most “host-guest” system due to
mild crosslinking approach. In this strategy, orthogonally functionalized FePt
nanoparticles (NPs) are self-assembled at the oil-water interface and crosslinked via
56
Figure 4.2: Orthogonal functionalization of FePt NPs and DTC chemistry for further
crosslinking at oil-water interface.
dithiocarbamate (DTC) chemistry (Figure 4.2).13 The major advantage of this assembly
process is the strong binding affinity of DTC conjugated organic ligands to the metallic
and semiconductor NPs; this interaction helps to stabilize the NPs at oil-water interface.
For our studies we used triethylene imine (TEI) functionalized FePt NPs to fabricate
colloidal microcapsules. Initially oleic acid and oleyl amine coated FePt NPs (~7 nm)
were synthesized following a reported procedure.14 Functionalization of the NPs was
Figure 4.3: Self-assembly and crosslinking of FePt NPs at oil-water interface
accomplished by place exchange reaction with HS-C11-TEG-OH and dopamine-C8-TEI
lignads respectively. The modified NPs were dissolved in milliQ water and pH of the
solution was adjusted to ~ 9 by adding 0.1 M NaOH solution. In the next step, 250 μL of
the NPs solution was transferred to an Eppendorf tube and 10 μL of 0.5 M CS2 in 1,2,4-
Trichlorobenzene (TCB) was added to it. Vigorous shaking for 10 s gave stable
57
emulsions (Figure 4.3) and emulsions were kept for 30 min prior to analysis. Excess NPs
were washed two times by milliQ water and kept it for further characterization.
4.2.2 Characterization of oil-in-water emulsions
Oil-in-water based colloidal microcapsules visualized were by optical microscope
(OM) (Figure 4.4). This assembly strategy was further investigated by using different
nonpolar solvents such as dichloromethane and hexane. Low boiling organic solvents
resulted in formation of crumbled microcapsules due to rapid evaporation of the solvent.
(e.g. dichloromethane, Inset of Fig. 1a). Microcapsules were drop casted on carbon
coated copper grid for transmission (TEM) analysis. Low-magnification image of the
dried capsules shows (Fig. 1b) thin film like texture, while the high magnification image
(Inset of Fig. 1b) reveals that the capsule is composed of densely packed crosslinked FePt
NPs.
Figure 4.4: OM micrographs of (a) FePt microcapsules at oil-in-water interface using DTC
crosslinking strategy. Inset shows crumbled microcapsules. (b) Low magnification TEM
images of dried microcapsules showing film like texture. Inset represents higher
magnification TEM image.
58
4.2.3 “Host” encapsulation and “Host-Guest” recognition at interface
“Host-Guest” chemistry was demonstrated by encapsulation of the flavin polymer15
(“host”) inside the microcapsules to obtain three-point hydrogen bonding interaction at the
interface with a complementary diaminopyridine (DAP, “guest”) amphiphile (Figure 4.4 a).
“Host-Guest” interactions at the interface were then monitored by fluorescence quenching
of the flavin fluorophore.16 Successful encapsulation of the “Host” flavin polymer inside
the microcapsules during their formation was confirmed by fluorescence of the
microcapsules in aqueous medium To study the recognition process inside the
microcapsule, an excess equivalent of the guest DAP amphiphile was added to the aqueous
solution. Concomitant quenching of fluorescence (Fig. 2b) from the microcapsule was
observed due to three point hydrogen bonding interaction between
Figure 4.5: (a) Flavin encapsulated microcapsule and “host-guest” recognition at oil-water
interface (b) Fluorescence microscopy images of Flavin encapsulated micro capsule and
after addition of excess DAP (c) Three point hydrogen bonding interaction between flavin
and DAP. Scale bar represents 50 µm.
59
“host-guest” at the oil-water interface (Figure 4.4c). Titration with N-methylated flavin
polymer as a control shows no significant quenching upon addition of excess DAP (Figure
4.6 a-b) demonstrating that three point hydrogen bond recognion was responsible for
fluorescence quenching.
Figure 4.6: Fluorescence images of (a) N-Methyl flavin encapsulated micro capsule (b)
After addition of excess DAP. All scale bars represent 40 μm of length.
4.2.4 Determination of “Host-Guest” binding constant
Semiquantitative assessment of the binding constant between flavin and
complementary DAP was obtained by monitoring the fluorescence intensity of a single
microcapsule. Fluorescence titration of flavin in presence of increasing DAP equivalents
yielded a Ka ≈ 250 M-1 (Figure 4.7a) which is comparable to their solution study. Titration
with N-methylated flavin polymer as a control shows no significant quenching upon
addition of excess DAP (Figure 4.7 a) demonstrating that three point hydrogen bond
recognion was responsible for fluorescence quenching. The fact that DAP-flavin
60
recognition quenced fluorescence prevented direct determination of the localization of the
“host-guest” complexes within the microsphere. Localization can be inferred, however,
from experiments performed using a planar interface. In this experiment the fluorescence of
flavin polymer was recorded in TCB. In the next step, an aqueous solution of excess DAP
was added to the organic phase. No fluorescence quenching was observed after agitation,
indicating that DAP and flavin polymer are soluble only in water and oil respectively
(Figure 4.7 b) and recognition can occur only at the interface.
Figure 4.7: (a) Binding plot shows dramatic quenching of flavin polymer fluorescence
upon addition of DAP (Ka ≈ 250 M-1) whereas control N-Methylated flavin does not show
any fluorescence quenching. (b) Fluorescence titration of flavin polymer with variable
DAP equivalents at the flat interface.
61
4.3 Conclusion
In summary, we have developed a simple and mild way to fabricate stable colloidal
microcapsules at oil-water interface based on orthogonal functionalization of FePt NPs
followed by DTC mediated crosslinking. These capsules provide a suitable macroscopic
platform to confine the “host-guest” functional pair at the interface. We have exploited the
fluorescence properties of the flavin moieties to probe “host-guest” complexation with a
complementary DAP derivative. These studies pave the way for the formation of capsules
with interesting biomimetic and materials applications.
4.4 Experimental Section
4.4.1 Synthesis of Dopamine-C8-TEI Ligand
Step 1:
HOOH
O
OHO
OO
ON
O
O
NO
OH
DCC, DMAP, THF
O
A
Sebacic acid (2.00 g, 9.89 mmol) was dissolved in dry THF (25 mL). To this
solution N-hydroxysuccinimide (1.36 g, 11.87 mmol) was added followed by a solution
of DCC (2.02 g, 10.09 mmol) in 5 mL of dry THF at 0°C. The mixture was stirred for 15
min, then DMAP (0.01 g, 0.08 mmol) was added. The reaction mixture was allowed to go
to r.t. and stirred for 7 h. The white precipitate was filtered and the solvent was removed
from the filtrate to give a residue that was then re-dissolved in CH2Cl2 and washed with
water. The organic layer was dried over Na2SO4, filtered and evaporated and the crude
product obtained was purified by flash chromatography with 1/2=EtOAc/CH2Cl2 as
eluent to give activated acid in 72% yield.
62
1H NMR δ (400 MHz , CDCl3): 1.20-1.42 (m, 12H, CH2); 1.55 (quintet, J=7.2 Hz, 2H,
CH2); 1.70 (quintet, J=7.2 Hz, 2H, CH2); 2.30 (t, J=7.2Hz, 2H, CH2); 2.30 (t, J=7.2Hz,
2H, CH2); 2.56 (t, J=7.2Hz, 2H, CH2); 2.80 (s, 4H, CH2).
Step 2:
HOO
O
ON
O
O
H2NN
NNH
Boc
Boc
Boc
TEA, DMFHO
NHO
O
NN
NHBoc
BocBoc
A C
B
Compound B was synthesized according to the literature reported method.17 In the
next step, TEA was added to a solution of B (0.20 g, 0.45 mmol) in DMF/H2O (5 mL/5
mL), until pH 9 is reached. This basic solution was then added to another solution of A
(0.15 g, 0.45 mmol) in 11 mL of DMF. The reaction mixture was stirred for 18 h under
argon, then poured onto water and extracted with EtOAc (6x100 mL). The recollected
organic layers were washed with water, dried over Na2SO4, filtered and concentrated.
The residue was purified by flash chromatography with 50/1=CH2Cl2/MeOH as eluent to
yield amide C in 99% yield.
1H NMR δ (400 MHz , CDCl3): 1.20-1.30 (m, 12H, CH2); 1.43 (s, 9H, CH3); 1.47 (s, 18H, CH3); 1.52-1.61 (m, 1H, NH); 2.13 (at, J=7.2 Hz, 2H, CH2); 2.30 (t, J=7.2 Hz, 2H, CH2); 3.18-3.45 (m, 12H, CH2).
63
Step3:
HBTU, DIPEA, DMF/DMSO
HN
NHO
O
NN
NHBoc
BocBoc
HONH
O
O
NN
NHBoc
BocBoc
OH
OHH2N
HOOH
C D
Compound C (0.28 g, 0.44 mmol) and dopamine hydrochloride (0.08 g, 0.49
mmol) were dissolved in 8 mL of DMSO. The solution was degassed with argon and
cooled to 0 °C before adding HBTU (0.21 g, 0.56 mmol) and DIPEA (0.24 g, 1.79
mmol). The reaction mixture was then stirred at r.t. for 64 h, concentrated and, after the
addition of EtOAc, washed with HCl 1M and water. The resulting solution was dried
over Na2SO4, filtered and concentrated to give the desired dopamine derivative D in 99%
yield without further purification.
1H NMR δ (400 MHz , CDCl3): 1.21-1.29 (m, 12H, CH2); 1.35-1.59 (m, 31H, CH3 + CH2); 2.11-2.17 (m, 4H, CH2); 2.62 (at, J=7.2 Hz, 2H, CH2); 3.19 (bs, 2H, CH2); 3.27-3.41 (m, 10H, CH2); 6.51 (d, J=8.0 Hz, 1H, Harom); 6.66-6.75 (m, 2H, Harom). Step 4:
HN
NHO
O
NN
NHBoc
BocBoc
HOOH
HN
NHO
O
E
NHNH
NH2
HOOH
TFA
CH2Cl2D
Compound D (0.34 g, 0.45 mmol) was dissolved in 15 mL of CH2Cl2 and TFA (5.00
mL, 43.17 mmol) was added. The mixture was stirred under argon for 4 h. The solvent
was removed under vacuum and the residue was purified by washing with Hexane (3
times) and Et2O (twice) to obtain amine E in 99% yield. 1H NMR δ (400 MHz, CD3OD): 1.21-1.29 (m, 8H, CH2); 1.51-1.62 (m, 4H, CH2); 2.13 (t, J=7.2 Hz, 2H, CH2); 2.22 (t, J=7.6 Hz, 2H, CH2); 2.61-2.69 (m, 2H, CH2); 3.23 (bs, 2H, CH2); 3.27-3.41 (m, 12H, CH2); 6.51 (d, J=8.0 Hz, 1H, Harom); 6.66-6.75 (m, 2H, Harom).
64
4.4.2 Synthesis of DAP amphiphile
NH2N N
H
O
NN N
H
O
H
O
Br
The synthesis of ammonium was synthesized from a literature reported procedure. 18To a stirred solution of 2-ethylamido-6-aminopyridine (0.4g, 2.4 mmol) in CH2Cl2
(200 ml) was added 6-bromohexanoyl chloride (0.57 g. 2.7 mmol) and triethylamine
(0.35 ml). The reaction mixture was stirred for 24 h. at room temperature and then water
was cautiously added (100 ml). The organic layer was separated and the aqueous layer
was further extracted with CH2Cl2 (2 x 100 ml). The organic extracts were combined,
dried over MgSO4, filtered and concentrated under reduced pressure. The crude extract
was purified using silica gel column chromatography (eluent: CH2Cl2/ethyl acetate,
80/20 (v/v)) to afford the product as a white solid (0.5 g, 61 %).
Mpt. 93-95 oC.
1H NMR δ (400 MHz, CDCl3): δ 7.90 (2H, t); 7.70 (1H, t); 3.30 (2H, t); 2.30 (4H, m);
1.80 (2H, m); 1.65 (2H, m); 1.40 (2H, m); 1.10 (3H, m). 13C NMR (100 MHz, CDCl3): δ
173.03; 171.98; 149.78; 149.76; 140.41; 109.44; 50.08; 44.78; 36.97; 33.63; 32.28;
30.40; 27.57; 24.44; 9.38. MS (m/z, CI) 342/344 (100%) (M+H). Anal. Calc. for
C14H20N3O2Br C, 49.13, H, 5.89; N, 12.28; found C, 49.46, H, 5.78, N, 12.11.
NN NH
O
H
OBr
NN NH
O
H
ON
Br
To a stirred solution of 2-ethylamido-6-(6-bromohexylamido) pyridine (0.5 g, 1.5
mmol) in methanol (10 ml) was added trimethylamine (2 ml, 30 % solution in methanol).
The reaction was heater under reflux for 24 h. The solution was concentrated under
reduced pressure and precipitated into a vigorously stirred solution of diethyl ether (100
ml). The precipitate was filtered off and washed with ice-cold diethyl ether. The product
65
was dried under high-vacuum for 24 hrs to afford the product as a white solid (0.5 g.
83%).
Mp. 87-92 oC.
1H NMR δ (400 MHz, D2O): δ 7.70 (1H, t); 7.45 (2H, d); 7.37 (2H, m); 3.20 (2H, m);
3.00 (9H, s); 2.40 (4H, m); 1.75 (2H, m); 1.65 (2H, m); 1.35 (2H, m); 1.10 (3H, t). 13C
NMR (100 MHz, D2O): δ 141.22; 111.17; 111.03; 66.37; 52.80; 52.76; 52.72; 36.30;
30.02; 24.97; 24.39; 22.07; 8.96. MS (m/z, FAB) 322 (40 %) (M+ (-Br)). Anal. Calc. for
C17H29N4O2Br C, 50.87, H, 7.28; N, 13.96; found C, 50.66, H, 7.20, N, 13.69.
4.4.3 Functionalization of FePt NPs
Figure 4.8: Schematic representation of ligand exchange reaction
In a typical experiment 10 mg of oleic acid and oleylamine stabilized FePt NPs
were mixed with 30 mg of SH-C11-TEG-OH ligand in 5 mL of dichloromethane (DCM).
The mixture was stirred at RT for over night under inert atmosphere. Black precipitation
was observed after 12 hours. MeOH (5mL) was added to dissolve the resultant NPs. In
66
the next step, 7.5 mg of DOP-PEI (in 5 ml of MeOH) was added and the resulting
mixture stirred for 48 hours at 35 °C. After removing solvent, functionalized NPs were
dissolved back in MeOH and purified by precipitation using DCM. Finally, the purified
FePt NPs were dissolved in 2 mL of water. (Final concentration of the stock solution: 5
mg / mL).
4.4.4 Fabrication of colloidal microcapsule
In a typical procedure, 100 μL stock solutions of functionalized FePt NPs were
transferred in an eppendorf tube and diluted by adding 200 μL of milliQ water. The pH of
the NPs solution was adjusted to ~ 9 by adding 0.1 M NaOH solution. In the next step, 10
μL of oil (mixture of 0.5 M of CS2 and flavin polymer in TCB) were added to the
aqueous solution of the FePt NPs. The heterogeneous mixture was vigorously shaken by
hand for 10-15 seconds. As a result, the solution appeared to be cloudy due to the
formation of microemulsions. After 30 minutes, emulsions settled at the bottom of the
tube and the supernatant liquid was washed several times by milli Q water prior to
characterization.
4.4.5 Size distribution of the microcapsules (determined from optical microscopy
image)
Mean 12.529 μm
SD 5.061 μm
67
4.5 References [1] Alberts, B.; Bray, D.; Lewis, J.; Raff, M.; Roberts, K.; Watson, J. D. Molecular
Biology of the Cell, 2nd ed.; Garland Publishing: New York, 1989.
[2] (a) Ariga, K.; Anslyn, E.V.; J. Org. Chem. 1992, 57, 417. (b) Kneeland, D. M.;
Ariga, K.; Lynch, V.M.; Huang, C.Y.; Anslyn, E.V.; J. Am. Chem. Soc. 1993, 115, 10042
(c) Smith, J.; Ariga, K.; Anslyn, E.V.; J. Am. Chem. Soc. 1993, 115, 362.
[3] (a) Cram, D. J.; Science 1988, 240, 760; (b) Cram, D. J.; Angew. Chem., Int. Ed. Engl.
1988, 27, 1009. (c) Conn, M. M.; Rebek, J. Jr., Chem. Rev. 1997, 5, 1647.
[4] Fan, E.; Van Arman, S. A.; Kincaid, S.; Hamilton, A. D.; J. Am. Chem. Soc. 1993,
115, 369.
[5] (a) Bonar-Law, R. P.; J. Am. Chem. Soc., 1995, 117, 12397. (c) Ariga, K.; Kunitake,
T.; Acc. Chem. Res. 1998, 31, 378. (d) Sasaki, D. Y.; Kurihara, K.; Kunitake, T.; J. Am.
Chem. Soc. 1991, 113, 9685. (e) Ikeura, Y.; Kurihara, K.; Kunitake, T.; J. Am. Chem.
Soc. 1991, 113, 7342. (f) Cha, X.; Ariga, K.; Onda, M.; Kunitake, T.; J. Am. Chem. Soc.
1995, 117, 11833. (g) Oishi, Y.; Kato, T.; Kuramori, M.; Suehiro, K.; Ariga, K.; Kamino,
A.; Koyano, A.; Kunitake, T.; J. Chem. Soc., Chem. Commun. 1997, 1357. (h) Aoyama,
Y.; Tanaka, Y.; Sugahara, S.; J. Am. Chem. Soc. 1989, 111, 5397.
[6] Rotello, V. M.; Viani, E. A.; Deslongchamps, G.; Murray, B. A.; Rebek, J. Jr., J. Am.
Chem. Soc. 1993, 115, 797
[7] Nowick, J. S.; Cao, T.; Noronha, G.; J. Am. Chem. Soc. 1994, 116, 3285.
[8] (a) S. Ishizaka, S. Kinoshita, Y. Nishijima, N. Kitamura, Anal. Chem. 2003, 75, 6035.
(b) Onda, M.; Yoshihara, K.; Koyano, H.; Ariga, K.; Kunitake, T., J. Am. Chem. Soc.
1996, 118, 8524.
68
[9] (a) Opsteen, J. A.; Brinkhuis, R. P.; Teeuwen, R. L. M.; Lo¨wik, D. W. P. M.; van
Hest, J. C. M.; Chem. Commun. 2007, 3136. (b) Vriezema, D. M.; Garcia, P. M. L.; Oltra,
N. S.; Hatzakis, N. S.; Kuiper, S. M.; Nolte, R. L. M.; Rowan, A. E.; van Hest, J. C. M.;
Angew. Chem. Int. Ed. 2007, 46, 7378. (c) Broz, P.; Driamov, S.; Ziegler, J.; Ben-Haim,
N.; Marsch, S.; Meier, W.; Hunziker, P.; Nano Lett., 2006, 6, 2349.
[10] (a) Hotz, J.; Meier, W.; Langmuir, 1998, 14, 1031. (b) Stoenescua,R.; Meier, W.;
Chem. Commun., 2002, 3016.
[11] (a) Pieranski, P.; Phys. Rev. Lett. 1980, 45, 569; (b) Rautaray, D.; Banpurkar, A.;
Sainkar, S. R.; Limaye, A. V.; Ogale, S.; Sastry, M.; Cryst. Growth. Des. 2003, 3, 449.
(c) Wang, D.; Duan, H.; Mohwald, H.; Soft Matter, 2007, 3, 1231. (d) Boker, A.; He, J.;
Emrick, T.; Russell, T. P.; Soft Matter 2007, 3, 1231. (e) Skaff, H.; Lin, Y.; Tangirala, R.;
Breitenkamp, K.; Böker, A.; Russell, T. P.; Emrick, T.; Adv. Mater, 2005, 17, 2082. (f)
Arumugam, P.; Patra, D.; Samanta, B.; Agasti, S. S.; Subramani, C.; Rotello, V. M.; J.
Am. Chem. Soc. 2008, 130, 10046. (g) Glogowski, E.; Tangirala, R.; He, J.; Russell, T.
P.; Emrick, T.; Nano Lett. 2007, 7, 389. (h) Duan, H.; Wang, D.; Sobal, N. S.; Giersig,
M; Kurth, D. G.; Mohwald, H.; Nano Lett. 2005, 5, 949. (i) Samanta, B.; Patra, D.;
Subramani, C.; Ofir, Y.; Yesilbag, G.; Sanyal, A.; Rotello, V. M.; Small 2009, 5, 685.
[12] Wang, B.; Wang, M.; Zhang, H.; Sobal, N. S.; Tong, W.; Gao, C.; Wang, Y.;
Giersig, M.; Wang, D.; Mo¨hwald, H.; Phys. Chem. Chem. Phys. 2007, 9, 6313.
[13] (a) Sharma, J.; Chhabra, R.; Yan, H.; Liu, Y.; Chem. Commun. 2008, 2140. (b)
Dubois, F.; Mahler, B.; Dubertret, B.; Doris, E.; Mioskowski, C.; J. Am. Chem. Soc.
2007, 129, 482. (c) Park, M. –H.; Ofir, Y.; Samanta, B.; Arumugam, P.; Miranda, O. R.;
Rotello, V. M.; Adv. Mater. 2008, 20, 4185.
69
[14] (a) Srivastava, S.; Samanta, B.; Jordan, B. J.; Hong, R.; Xiao, Q.; Tuominen, M. T.;
Rotello, V. M.; J. Am. Chem. Soc. 2007, 129, 11776. (b) Ofir, Y.; Samanta, B.;
Arumugam, P.; Rotello, V. M.; Adv. Mater. 2007, 19, 4075.
[15] Carroll, J. B.; Jordan, B. J.; Xu, H.; Erdogan, B.; Lee, L.; Cheng, L.; Tiernan, C.;
Cooke, G.; Rotello, V. M.; Org. Lett. 2005, 7, 2551.
[16] (a) Caldwell, S. T.; Cooke, G.; Hewage, S. G.; Mabruk, S.; Rabani, G.; Rotello, V.
M.; Smith, B. O.; Subramani, C.; Woisel, P.; Chem. Commun. 2008, 4126.
[17] Srinivasachari, S.; Fichter, K. M.; Reineke, T. M.; J. Am. Chem. Soc. 2008, 130,
4618.
[18] Hirst, S. C.; Hamilton, A. D.; Tetrahedron Lett. 1990, 31, 2401.
70
CHAPTER 5
HOST-GUEST MOLECULAR RECOGNITION OF NANOPARTICLES AT LIQUID-LIQUID INTERFACE
5.1 Introduction
Self-assembled microcapsules have tremendous potential as multifunctional scaffolds for
microreactors and carriers for catalysts,1 enzymes2 and drugs.3 The essential prerequisite
for these applications is the fabrication of robust and stable microcapsules. Thus
designing the building blocks and their properties are of critical importance for further
advances. Various types of microcapsules (such as polymeric microcapsules,4 vesicles,5
colloidal microcapsules6) have been synthesized so far for this purpose. In this context,
colloidal microcapsules exhibit unique features highlighted with mechanical stability,
tunable permeability7 as well as optical, magnetic and fluorescent properties of their
precursor particle building blocks.
The generic approach to design colloidal microcapsules is using the emulsion
droplets as template to self-assemble and crosslink colloidal particles at the liquid-liquid
interface. Recently, significant advances have been done to engineer colloidal
microcapsules by using different assembly strategies.8 Although covalently built
microcapsules provide impressive robustness, lack of versatility such as structural
manipulation and stimuli responsive release of active ingredients restricts their potential
use. In comparison to the existing particle assembly approaches, molecular recognition
mediated self-assembly is an attractive alternative.9 Molecular recognition provides a
noncovalent coupling between two complementary molecules or functionalized surfaces.
The ‘tailored interactions’ between building blocks via molecular recognition along with
71
the ‘dynamic nature’ of self-assembly process provides a promising tool to create novel
molecular assemblies and functional materials.10
5.1.1 “Host-guest” chemistry-A noncovalent approach of nanoparticle self-assembly
One important focus in self-assembly of nanoparticles (NPs) is the tailoring of
supramolecular and interfacial forces for interactions. Monolayer protected nanoparticles
serves as a well-organized scaffold for the attachment of diverse functional molecular
units and large surface areas for molecular recognition.11 In addition, mutivalency of NPs
provides a higher degree of interaction compare to a single recognition element.12
Various types of noncovalent interactions have been employed (e.g., hydrogen bond,
electrostatic interaction, dipolar interactions, van der Waals forces and hydrophobic
interactions) to govern the self-assembly of nanoparticles.13 The key characteristics of
these interactions are high selectivity, directionality, reversibility and controlled affinity.
These interactions are considered to be an attractive pathway for engineering of new
functional materials with advanced and improved properties.14 Moreover, these
interactions help NPs to create controlled structures and lead to modulation of the
physical response.
In this context, “host-guest” mediated noncovalent interaction is an important
class of example such as cyclodextrin (CDs)-adamante (ADA) interaction because of
their site selective molecular recognition.15 CDs are well-known molecular hosts capable
of including small hydrophobic molecules inside their cavities in aqueous media. In
particular, β-CDs have cavities of around 7A° in size and can selectively incorporate
ADA molecules (~7 A°) inside the hydrophobic cavity via “host-guest” recognition.16
This system forms inclusion complexes with relatively high stability constants (104–105
M-1). In recent years, different approaches have been made to exploit CD-ADA
72
interaction for self-assembly of NPs. Huskens et al demonstrated the stepwise
construction of self-assembled organic/inorganic multilayers on the basis of multivalent
“host-guest” interactions between guest-functionalized dendrimers and host-modified Au
NPs (Figure 5.1a).17 In another work, Yang et al described a surface modification method
based on “host-guest” complex formation between organic monolayer of NPs and the
hydrophilic CD macromolecules (Figure 5.2b).18 Finally, it lead to the phase transfer of
hydrophobic NPs into aqueous phase. Recently, the concept of “host-guest” chemistry
was also employed at the oil-water interface to assemble NPs. The specific inclusion of
ADA by CD can direct hydrophobic adamantane carboxylic acid capped NCs to self-
assemble into macroscopic randomly close-pack monolayers at water/oil interfaces
(Figure 5.3c).19
Figure 5.1: (a) Layer-by-Layer Assembly scheme for the alternating adsorption of
Adamantyl-Terminated PPI Dendrimer and CD Au NPs onto CD SAMs (b) Phase
transfer of hydrophobic NPs to water via surface modification using CD (c) Self-
73
assembly of ADA stabilized NPs at the water/oil interface by the inclusion of ADA-
capped NCs from an oil phase into CDs dissolved in water. [Reproduced from references
17, 18 and 19 respectively]
5.2 Results and discussion
5.2.1 “Host-guest” recognition mediated microcapsules fabrication liquid-liquid
interface
In this study, we fabricate colloidal microcapsules via “host-guest” molecular
recognition at the liquid-liquid interface. Specifically, we crosslink β-cyclodextrin (β-CD,
“host”) and adamantane (ADA, “guest”) functionalized gold NPs at the “oil-water”
interface to create microcapsules. In the next step, recognition induced coalescence of the
microcapsule was achieved by sequential addition of ADA amphiphile as an external and
competing “guest” to the microcapsules. This approach allows us to modulate the “host-
guest” recognition at the interface.
Figure 5.2: (a) “Host” functionalized Au NPs (b) “Guest” functionalized Au NPs (c) NP-
NP assembly via “Host-Guest” recognition.
74
In the present study, we have synthesized hydrophilic β-CD functionalized Au NPs (NP1,
size ~ 3nm, Figure 5.2a) as a functional “host” according to the literature reported
procedure.20 The “guest” functionalized hydrophobic NPs (NP2, size ~ 6 nm, Figure
5.2b) were prepared by ligand exchange reaction of ADA-dithiol with 6 nm Au NPs.21 In
the next step, 300 μL of NP2 were dissolved in toluene to which 10 μL aqueous solution
of NP1 and fluorescence dye (fluorescein isothiocyanate functionalized dextran polymer;
Mw= 500,000 g mol-1, Rh = 9.4 nm,) were added. Vigorous shaking for 30 seconds
resulted in formation of stable emulsions (Figure 5.3).
Figure 5.3: Schematic of fabrication of colloidal microcapsules
5.2.2 Characterization of microcapsules
The supernatant liquid was washed several times by toluene and microcapsules
were analyzed by both optical (Figure 5.41a) and fluorescence (Figure 5.4b) microscope.
The size of the microcapsules fabricated by this procedure was varied from 15-30 μm.
The formation of stable emulsions (Figure 1a) confirmed sufficient and extended
crosslinking between NP1 and NP2 at the “oil-water” interface via “host-guest”
recognition. For further investigation, the microcapsules were drop casted on carbon
coated grid for transmission electron microscope (TEM) analysis. Low resolution TEM
75
image (Figure 5.4c) depicts the dried capsule structure where as high resolution TEM
image (Figure 5.4d) revealed that capsule is composed of multilayer of closely packed
crosslinked NPs.
Figure 5.4: Optical microscopy image of (a) colloidal microcapsule (b) Fluorescence
microscopy image of dye encapsulated microcapsule (c) Low resolution TEM image of
single microcapsule (d) High resolution TEM image of the microcapsule wall.
5.2.3 Recognition induced disassembly
The major advantage of noncovalent interaction is their reversible assembly and
disassembly. The assembly can be disrupted by adding external stimuli such as
temperature, pH, solvents or competitive recognition element. In this case we used
competitive “guest” molecules to disassemble the “host-guest” interaction between NPs.
Recognition induced deterioration of molecular recognition between NP1 and NP2 at the
“oil-water” interface was done by a simple set of experiment (Figure 5.5a). In this study,
1.5 mL of NP2 in toluene was placed on the top of 1.5 mL of NP1 in water. After
vigorous shaking, both the organic and aqueous phase became colorless and brown
76
colored emulsions, consisting of millimeter sized droplets formed at the interface (Figure
5.5c) due to recognition induced self-assembly between NP1 and NP2. When ADA-
Figure 5.5: (a) Schematic of recognition induced disassembly. Photographs of glass vial
representing (b) recognition induced self-assembly of NP1 and NP2 at toluene-water
interface (c) recognition induced disassembly in presence of external “guest” (NP1). Plot
of (c) plasmon absorption band of NP2 in presence of increasing ADA-TEG-OH
concentration.
-TEG-OH (external “guest” Figure 5.5 b) was successively added to the microcapsules
solution, both organic and aqueous phases turned back to brown color (Figure 5.5d). This
suggests the deterioration of molecular recognition between NP1 and NP2 in presence of
77
the external “guest”. The plasmon absorption of NP2 provides an efficient in situ
measurement to monitor the recognition induced assembly and disassembly process of
the NPs at the interface. The saturation of absorbance curve indicates that the maximum
number of NP2 redisperse to the organic phase after disassembly (Figure 5.5e).
5.2.4 Size tuning of microcapsules
An important aspect of “host-guest” chemistry is that interactions can be
disrupted by competing ligands, a possible means for weakening the assembly and
potentially controlling microcapsule size. Size tunability of the colloidal microcapsule
(Figure 2a) was demonstrated by sequential addition of an external amphiphilic “guest”
ADA-TEG-OH (Scheme 1d) to the microcapsule in toluene. Addition of ADA-TEG-
OH to the NP1-NP2 microcapsules interferes with the “host-guest” recognition between
NP1 and NP2. As a result, interfacial crosslinking is disrupted and microcapsules
coalesce with each other to form larger droplets (Figure 2b-2e).
\
78
Figure 5.6: (a) Size tunability of the colloidal microcapsule at different ADA-TEG-OH
concentrations in presence of ~ 0.4 uM of NP2 and ~ 0.4 uM of NP1 (b-e) Representative
fluorescence microscopy images of different size microcapsules.
5.3 Conclusions
In summary, we have developed a non-covalent approach to fabricate colloidal
microcapsule at the liquid-liquid interface based on molecular recognition. The reversible
and dynamic nature of specific recognition process provides a tool for structural
manipulation, specially tuning the size of the microcapsules. This approach might lead to
synthesize the stimuli responsive system for drug delivery applications. Further studies
79
aimed at controlling the size and release profile by external stimuli (i.e. temperature,
redox potential) are under investigation and will be reported in due course.
5.4 Experimental Section
5.4.1 Chemicals
Chemicals were obtained from commercial sources and were used as received
unless otherwise stated. Milli-Q water with a resistivity greater than 18 MΩ.cm was used
in all our experiments. Tetra (ethylene glycol) mono-adamantyl ether and β-CD
functionalized Au nanoparticles were synthesized as described before. NMR spectra were
recorded on Varian 400 MHz spectrometers. Elemental analysis results were obtained on
a FlashEA 1112 CHNS analyzer.
5.4.2 Synthesis of ADA-disulfide Ligand
To a solution of tetraethylene glycol mono-adamantyl ether22 (205 mg, 0.63
mmol) and thioctic acid (103 mg, 0.50 mmol) in dry CH2Cl2 (5 mL), were added EDC
(105 mg, 0.55 mmol) and DMAP (24 mg, 0.20 mmol). The reaction was allowed to stir
under argon overnight. The solvent was concentrated under reduced pressure and the
residue was purified by column chromatography (40% EtOAc in hexane) to afford the
ADA ligand as a yellow oil (196 mg, yield 76 %). 1H NMR (400 MHz, CDCl3) (ppm)
= 4.20 (2H, t), 3.67 (2H, t), 3.63 (8H, s), 3.56 (5H, m), 3.12 (2H, m), 2.43 (1H, m), 2.33
(2H, t), 2.11 (3H, s), 1.88 (1H, m), 1.72 (6H, s), 1.63 (10H, m), 1.44 (2H, m). 13C
NMR(400 MHz, CDCl3) δ(ppm): 173.43, 72.23, 71.27, 70.63, 70.58, 70.57, 70.55, 69.15,
63.46, 59.23, 56.30, 41.45, 40.19, 38.46, 36.44, 34.57, 33.92, 30.48, 28.71, 24.59. Anal.
Calcd for C26H44O6S2: C, 60.43; H, 8.58; S, 12.41. Found: C, 60.61; H, 8.62; S, 12.37.
80
5.4.3 Ligand Exchange Reaction
Reduction of ADA-disulfide to ADA-dithiol was performed before place
exchange reaction.23 In the next step, dodecanethiol-coated Au NPs (10 mg) were
dissolved in 5mL of DCM and 50 mg of ADA-dithiol was added to the NP solution.
Reaction mixture was purged with argon and stirred for 2 days at room temperature in a
closed vial. The mixture was precipitated with ethanol and centrifuged. Purification of the
NPs was achieved through redispersion in toluene.
5.4.3 Microcapsule Fabrication
In a typical procedure, 300 µL solution of NP2 (0.4 μM) in toluene was taken in
an eppendorf tube. In the next step, 10 μL aqueous solution (11 µM) of NP1 (final
concentration 0.4 uM) and FITC dextran (fluorescein isothiocyanate functionalized
dextran polymer; Mw= 500,000 g mol-1, Rh = 9.4 nm,) was added. The heterogeneous
mixture was vigorously shaken for 15 seconds using Amalgamator (blending speed: 4000
rpm). As a result, the solution appeared to be cloudy due to the formation of
microemulsions. After 10-15 minutes, emulsions settled at the bottom of the tube and the
supernatant liquid was washed several times by toluene prior to characterization.
5.4.4 Microcapsule characterization
TEM images were acquired on a JEOL 100CX operating at 100 keV. Samples
were drop casted onto a carbon-coated copper grid, dried, and imaged. Both optical and
fluorescence microscope images were taken on an Olympus IX51 microscope. For
fluorescence images, an excitation wavelength of 470 nm and emission wavelength above
515 nm (green fluorescence) were used.
81
5.4.5 Size distribution of the microcapsules with increasing concentration of ADA-
TEG-OH
Figure 5.7: Size distribution of the microcapsule after adding 0 μmoles of ADA-TEG-
OH (mean size: 5.815 μm, SD: 2.1)
82
Figure 5.8 : Size distribution of the microcapsule after adding 2.5 μmoles of ADA-TEG-
OH (mean size: 18.045 μm, SD: 6.889)
Figure 5.9: Size distribution of the microcapsule after adding 5 μmoles of ADA-TEG-
OH (mean size: 30.88μm, SD: 14.2)
83
Figure 5.10: Size distribution of the microcapsule after adding 7.5 μmoles of ADA-TEG-
OH (mean size: 98.05 μm, SD: 34.92)
Figure 5.11: Size distribution of the microcapsule after adding 10 μmoles of ADA-TEG-
OH (mean size: 178.71μm, SD: 82.265)
84
5.5 References
[1] (a) Shchukin, D. G.; Möhwald, H. Small 2007, 3, 926. (b) Buonomenna, M. G.;
Figoli, A.; Spezzano, I.; Morelli, R.; Drioli, D. J. Phys. Chem. B 2008, 112, 11264. (c)
Ryo Akiyama, R.; Kobayashi, S. Chem. Rev. 2009, 109, 594.
[2] (a) Zhu, H.; Srivastava, R.; Brown, J. Q.; McShane, M. J. Bioconjugate Chem. 2005,
16, 1451. b) Phadtare, S.; Kumar, A.; Vinod, V. P.; Dash, C; Palaskar, D. V.; Rao, M.;
Shukla, P. G.; Sivaram, S.; Sastry, M. Chem. Mater. 2003, 15, 1944. (c) Alves, C. S.;
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CHAPTER 6
STABLE MAGNETIC COLLOIDOSOMES VIA “CLICK” MEDIATED CROSSLINKING OF NANOPARTICLES AT OIL-WATER INTERFACE
6.1 Introduction Colloidosomes are generally hollow microcapsules with a shell composed of densely
packed colloidal particlesi and feature identical solvent inside and out (typically water).
The physical and chemical properties of colloidosomes (e.g. mechanical strength and
biocompatibility) can be tuned by proper selection of selecting colloidal particles with
desired surface functionalities. The hollow nature of colloidosomes provides to
encapsulate active ingredients which can be released in a controlled fashion. These
features make colloidosomes a potential candidate for macromolecular delivery,
cosmetics and food industries.ii
However, the formation of stable colloidosomes using nanoparticles (NPs) < 20
nm in diameter remains a challenge.iii The competition between the interfacial energy and
the spatial fluctuation of the NPs results from thermal energy causes instability of the
emulsions. Recent approaches to fabricate colloidosomes have included different types of
NPs, as well as the use of assembly strategies.iv For example, Duan et al. have used
agarose to gelate water at the water-in-oil interface and transferred them into water to
create stable colloidosomes.v In recent studies, we and others developed various
crosslinking reactions between NPs at water-in-oil droplet interfaces; however, to the best
of our knowledge there are no reports of successful transfer of these crosslinked droplets
into water to synthesize colloidosomes. Thus it is important to design a perfect and
simple chemistry at the interface to arrest the particle during the time of self-assembly
and to create dense packing of NPs for mechanical stability. vi
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6.1.1 “Click” chemistry at oil-water interface
The increasing need for materials with a high degree of structural order and well defined
properties will continue to stimulate new organic concepts in material science. In this
context, the discovery and selection of simpler and universal synthetic methods are
essential. One powerful example is the “click” chemistry. Sharpless and his coworkersvii
rediscover Huisgen 1,3-dipolar cycloaddition of azides and terminal alkynes in presence
of copper catalyst,
Figure 6.1: Basic schematic of “click” reaction
which makes the reaction highly efficient, fast and regioselective. Moreover, the copper-
catalyzed azide-alkyne cycloaddition can be performed in various solvents (including
water) and in the presence of numerous other functional groups under mild reaction
condition. The simplicity of the reaction makes it a versatile tool in material science.
Recently, few routes have been reported for the “click” functionalization of inorganic
NPsviii and carbon nanotubesix to make them more functional towards application.
Moreover, “click” chemistry has been shown to be an unprecedented tool for
functionalizing flat surfaces.x A very recent report described the spatial control of “click”
cycloadditions on flat surfaces by using microcontact printing creating various functional
patterns on azide terminated SAMs (Figure 6.2).xi The potential of “click” chemistry for
materials synthesis has been increasingly recognized and has already resulted in wide
90
range of beautiful manifestations. Ambient conditions and high yield of the reaction
allows us to perform “click” chemistry at oil-water interface where organic component
will be soluble in oil and catalyst will be taken in water.
Figure 6.2: “Click” reaction in different environments (a) In solution (b) Solution–
Surface (c) Reagent-Stamping (d) Stamp Catalyst [Reproduced from reference 11]
6.2Results and discussion
6.2.1 Magnetic Colloidosomes
Herein, we report the fabrication of stable magnetic colloidosomes by crosslinking
NPs at water-oil interface using “click” chemistry under ambient conditions. In this
strategy, alkyne and azide functionalized Fe3O4 NPs were co-assembled at the interface
and covalently linked using Cu (I) catalyzed Huisgen “click reaction”.xii There are two
major advantages for this interfacial crosslinking method. First, click chemistry involving
alkyne and azide functional groups is highly selective and essentially inert to the many
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functional groups and environmental conditions (e.g. pH and solvent).xiii Second, this
methodology provides dense packing of NPs on the colloidosome shell, resulting in high
stability of the colloidosomes. The alkyne (IO-1) and azide (IO-2) NPs used in this study
were formed by place-exchange of oleic acid from Fe3O4 NPs that were 11.3±2 nm in
diameter. These NPs were dissolved in equimolar ratio in oil (mixture of toluene and
methylene chloride with 7:1 ratio), and an aqueous solution of the Cu (I) catalyst
(mixture of CuSO4 and sodium L-ascorbate) was added with vigorous shaking for ~ 30
seconds (Figure 6.3).
Figure 6.3: (a) Formation of magnetic colloidosomes synthesis through self-assembly at
water-in-oil interfaces followed by interfacial crosslinking of NPs via click chemistry (b)
Structures of the NPs with ligands structure used for crosslinking and cross-sectional
view of a colloidosome.
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6.2.2 Characterization of colloidosomes
The colloidosomes formed by this technique were 49± 15 µm (Figure 6.4a-b) in
diameter, and required a crosslinking time of 30 mins with a catalyst concentration of 0.8
mM to form stable assemblies. The catalyst concentration had little effect on the shape
and size of the colloidosomes, however only unstable emulsions were observed in the
absence of catalyst or below 0.1 mM (Figure 6.4 c-d). Successful reaction of the terminal
azide and alkyne groups was confirmed by Fourier transform infrared (FTIR)
spectroscopy of the colloidosomes before and after the click reaction. After crosslinking,
the characteristic azide peak of IO-2 at 2094 cm-1 is significantly reduced with respect to
Figure 6.4 Optical micrographs of (a) crosslinked magnetic colloidosomes fabricated at
water-in-oil interface using click reaction (b) crumbled colloidosomes after partial drying
of same colloidosomes (c) 0.1 mM catalyst concentration (d) no catalyst.
93
alkyl hydrogen peal at 2920 cm-1 the successful reaction of the azide group. No decrease
in relative peak intensity is seen in a control sample, which contains only IO-1 and IO-2
NPs and CuSO4 (Figure 6.5 a). Low-magnification transmission electron microscope
(TEM) analysis of the dried shells (Figure 6.5c) indicates a thin-film morphology, while
the high magnification image (Figure 6.5 d) clearly reveals that the membrane is
composed of densely packed multiple layers of NPs. The enhanced stability of the
crosslinked colloidosomes is demonstrated by the uncollapsed structures seen even after
drying for 7 days (Figure 6.4b). Significantly, these crosslinked colloidosomes can be
transferred efficiently into water. Apart from their stability in water, the colloidosomes
are responsive towards external magnetic stimuli due to the inherent magnetic properties
of NP building blocks (Figure 6.5b).
Figure 6.5 (a) FTIR spectra of crosslinked NPs (green) and uncrosslinked control sample
(red): relevant peaks are marked (b) Photograph of the magnetic colloidosomes in toluene
subjected to magnetic field. TEM images of a colloidosome (c) Low magnification (d)
high magnification.
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6.2.3 Permeability of colloidosomes
The primary mode by which materials are released from colloidosomes is
diffusion through the interstitial pores between the colloidal particles. Sizes of these
interstitial pores are determined by the size and polydispersity of the colloidal particles.xiv
Thus, by controlling these parameters, the rate of diffusion through colloidosomes can be
finely tuned. As observed by TEM, NPs are randomly close packed in the colloidosomes
shell, so hexagonal, cubic and pentagonal packing arrays coexist. Based on simple
geometry analysis of the interstitial sizes of these three packing arrays, as shown in
Figure 6.6, the pentagonal arrangement gives the largest interstice, 0.7 of the NP
diameter, which is expected as the cutoff for permeation of ingredients through the
colloidosomes wall.xv In order to investigate the permeability of magnetic colloidosomes,
we incorporated fluorescent molecules of different sizes into the colloidosomes and
checked their
Figure 6.6: Schematic illustration of close-packing arrays of colloidosomes for
calculating their interstitial dimension. [Reproduced from reference 15]
95
diffusion surrounding solution using fluorescence microscopy to test the permeability of
the colloidosomes. Successful incorporation of fluorophores (e.g. riboflavin) into the
colloidosomes during their formation was verified by the fluorescence of the
colloidosomes in oil (Figure 6.7a). Riboflavin is a small dye (molecular weight: Mw 376
g mol-1, rh < 2 nm), and hence diffuses out to the outer aqueous dispersion medium
within a minute of solvent exchange (Figure 6.7b). In contrast, a high molecular weight
FITC-functionalized dextran polymers (Mw 500,000 g mol-1, rh 9.4 nm) did not diffuse
out from the colloidosomes, as no decrease in the fluorescence from colloidosome was
seen after transferring the colloidosomes from oil to water (Figure 6.7 c-d). An
intermediate molecular weight dextran polymer (Mw 40,000 g mol-1, rh 3.7 nm),
however, diffused slowly from colloidosome (Figure 6.74). These differences in the
permeability towards fluorophores are due to their size variation and demonstrate the
selective permeability of these colloidosomes. This observation is supported by a
previous report, where the pore size of colloidosomes shell was assumed to be ~70% of
the NP radius used for its assembly.
Figure 6.7 Fluorescence microscopy images of magnetic colloidosomes loaded with
riboflavin in a) toluene, and b) after transfer into water. Similarly, Fluorescence images
96
of colloidosomes loaded with FITC-dextran (Mw 500k g mol-1) c) in toluene and d) after
30 min in water.
Figure 6.8: Size dependent release of fluoroscent molecules from colloidosome in 10
mM aqueous NaCl solution.
6.3 Conclusion
In summary, we have presented a facile route to synthesize robust magnetic
colloidosomes based on interfacial self-assembly of Fe3O4 NPs followed by crosslinking
using click chemistry. Interfacial cross-linking at droplet surfaces enables the
encapsulation of water-soluble materials inside the resulting colloidosomes. Different
sized fluorescent molecules have been shown to have distinctive permeability out of the
colloidosomes in water based on size exclusion processes. By controlling the NPs size
and functionality, it will be possible to control the selectivity as well as the permeability
of colloidosomes, making them an attractive method for encapsulating and protecting
sensitive molecules, such as drugs and biologically relevant molecules. Additionally,
unreacted azide and alkyne sites on the particles will provide convenient attachment
points for functionality. Finally, the properties of these assemblies can be augmented by
97
chose of core materials for the constituent NPs, providing access to magnetic,
fluorescent, and photonic materials.
6.4 Experimental section
6.4.1 Synthesis of NPs
The Fe3O4 NPs precursors had a core size of ~10 nm and were synthesized using
a reported procedure.xvi In a typical procedure, 0.86 g (4.32 mmol) of FeCl2.4H2O and
2.34 g (8.66 mmol) of FeCl3.6H2O were dissolved under argon atmosphere in 80 mL of
milliQ water with vigorous stirring. The solution was purged with argon for 5 min to
remove any oxygen present in the milliQ water. A black precipitation was formed after
the addition of NH4OH (10 mL) to the reaction solution. The pH of the reaction mixture
was retained at 10-12. This precipitate was further incubated at 90 ºC for 20 min. The
reaction mixture was then cooled down to room temperature and washed with milli Q
water several times to remove any un-reacted chemicals and decrease the pH of the
solution to 7-8. 20 mL aqueous solution of oleic acid (1g) was added to the NP precursor
in argon atmosphere. Then the reaction mixture was sonicated for 20 min. A clear
reddish-brown color solution was formed. The reaction mixture was incubated at 100 °C
for 3h. Aqueous dispersion of the NPs was precipitated using acetone. The black colored
precipitation was redispersed in hexane. Excess ligands were removed by re-precipitating
using ethanol and dispersion in hexane. Purification was performed twice and NPs were
dispersed in hexane or dichloromethane.
6.4.2 Synthesis of 11-azidoundecanoic acid
11-bromoundecanoic acid
O
OHBrNaN3, KI
DMSO
O
OHN3
98
11-bromoundecanoic acid (3.77 mmol), sodium azide (37.7 mmol) and potassium iodide
(1.88 mmol) were mixed in dimethyl sulfoxide (60 mL). The resulting mixture was
heated to 80 °C for 2 days. After cooling down, 50 mL of water was added to the reaction
mixture and extracted with ethyl acetate (3×50 mL). The combined organic phase was
washed with brine (3×50 mL) and dried over anhydrous sodium sulfate.
NMR (1HNMR, 400 MHz, CDCl3): 3.22-3.18 (t, 2H), 2.25-2.21(t, 2H), 1.59-1.51(m,
4H), 1.32-1.21(m, 12H).
6.4.3 Place Exchange Reactions
IO-1: 40 mg of the oleic acid coated Fe3O4 NPs were dissolved in 5 mL DCM
and hexane mixture (9:1) with 200 mg of 10-undecynoic acid. The reaction mixture was
purged with argon and stirred for 2 days at room temperature in a closed vial. The
mixture was precipitated with ethanol and centrifuged. Purification of the NPs was
achieved through re-dispersion in DCM and precipitation in ethanol.
IO-2: 40 mg of the original Fe3O4 NPs were dissolved in 5 mL DCM and hexane
mixture (9:1) with 200 mg of 11-azidoundecanoic acid. The reaction mixture was purged
with argon and stirred for 2 days at room temperature in a closed vial. Purification of the
NPs was achieved through re-dispersion in DCM and precipitation in ethanol. Finally,
both IO-1 and IO-2 NPs were dispersed in DCM having ~10 mg mL-1 concentration.
99
Figure 6.9: TEM images of a) alkyne functionalized iron oxide NPs b) azide
functionalized iron oxide NPs
6.4.4 Synthesis of colloidosomes
35 µL of each NP solution were mixed in an Eppendorf tube and the resulting
mixture was diluted to 500 µL with toluene. Active click catalyst (Cu+1) was synthesized
by reducing CuSO4 (1 equivalent) with (+)- sodium L-ascorbate (10 equivalents) in 10
mM aqueous NaCl solution at room temperature. 25 µL of active catalyst solution was
added to the NPs solution and the resulting heterogeneous mixture was vigorously
shaken. After shaking, NPs segregated to the water-oil interface to create colloidosomes
and appeared as cloudy dispersion, which eventually settles down to the bottom of the
organic solvents mixture. Different dyes were added into catalyst solution to achieve dye
incorporation inside colloidosomes. Excess NPs were removed and fresh toluene was
added prior to imaging in the optical microscope.
6.4.5 Characterization
TEM images were acquired on a JEOL 100CX operating at 100 keV. Samples were
drop casted onto a carbon coated copper grid, dried, and imaged. Neat FTIR spectra of
the NPs were taken using a MIDAC M1200-SP3 spectrophotometer. Both optical and
100
fluorescence microscope images were taken on Olympus IX51 microscope. For
fluorescence images, excitation wavelength of 470±20 nm and emission wavelength
above 515 nm (green fluorescence) were used.
6.4.6 Permeability test
Permeability tests of the colloidosomes loaded with different sized fluorophores
were performed by monitoring the maximum fluorescence intensity of one colloidosome
over time. Dye loaded colloidosomes were transferred from organic solvent to 10 mM
aqueous NaCl solution. Since, colloidosomes-transferring process from oil to water takes
one minute, release rate was monitored after a minute.
Figure 6.10: Dynamic light scattering analysis of FITC-dextran polymers with Mw 40k g
mol-1 (solid line) and 500k g mol-1 (dashed line).
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104
CHAPTER 7
SELF-ASSEMBLY OF ENZYME-NANOPARTICLE CONJUGATES AT
LIQUID-LIQUID INTERFACE
7.1 Introduction
Enzymes catalyze chemical reactions with impressive levels of stereospecificity,
regioselectivity and chemoselectivity,1 making enzyme immobilization an important tool
for the fabrication of a diverse range of functional materials and devices.2 To date, a
variety of synthetic scaffolds and supports3 have been used for enzyme immobilization
including gels,4 macromolecules,5 vesicles,6 nanoreactors,7 carbon nanotubes,8
microspheres,9 and surface-anchored molecules.10 Recently, nanoparticles (NPs) have
been used to immobilize enzymes, harnessing the optical, fluorescence and magnetic
properties of the nanomaterials.11 The unique properties and utility of NPs arise from a
variety of attributes, including the similar size of NPs and biomolecules such as proteins
and polynucleic acids.12 The size of NPs cores can also be tuned from 1.5nm to more
than 20 nm depending on the core material, providing a larger surface area for the
interaction of NPs with proteins and other biomolecules. Additionally, NPs can be
created with a wide range of metals and semiconductors core materials that impart useful
properties such as fluorescence and magnetic behavior.13 Nonetheless, the NPs surface
can be precisely engineered by intelligent ligand design, offering versatility in the
creation of surface-specific receptors. Biomolecular surface recognition by NPs as
artificial receptors provides a potential tool for controlling cellular and extracellular
processes for numerous biological applications such as transcription regulation,
enzymatic inhibition, delivery and sensing.14 Hence, the modulation of structure and
105
function of biomolecules through engineered interactions with NPs is an important issue
in nano-biotechnology.
7.1.1 NPs-Protein Conjugation
NPs can be conjugated with proteins via two approaches (i) covalent interaction
(ii) noncovalent interaction. In first approach, NPs and proteins can be directly linked via
covalent bond formation.15 Proteins are often covalently linked to the ligand on NPs
surface via traditional coupling strategies. Moreover, proteins can be selectively bound to
the NPs through use of transition metal complex formation. Xu et al. fabricated FePt
magnetic NP with nickel terminated nitrilotriacetic acid (NTA).16 These NPs show high
affinity and specificity towards histidine-tagged proteins (Figure 7.1a). This concept can
be employed to manipulate the histidine-tagged recombinant proteins and bind other
biological substrates at low concentrations. In second approach, noncovalent
modification provides a highly modular approach for the biofunctionalization of NPs.
The complexation between protein and NPs can be achieved by noncovalent interaction
(e.g. electrostatic, hydrophobic, pi-pi stacking etc.) or by specific interaction (e.g.
antigen-antibody interaction).17 The noncovalent complexation can be achieved by
electrostatic interaction by using oppositely charged NPs with respect to proteins. One
important feature of NP-protein conjugation is to retain their native structure after
complex formation. The use of alkyl based monolayer results in protein denaturation. Our
group and others introduced oligoethyleneglycol unit in the monolayer of NPs to enhance
the stability of protein and to minimize nonspecific adsorption of other materials.18 In one
work, our group has shown three level of interaction of chymotrypsin (ChT) with cationic
CdSe NPs (Figure 7.1b): no interaction (OEG terminated with hydroxyl group),
inhibition with denaturation (carboxylate-terminated thioalkyl ligands), and inhibition
with retention of structure (carboxylate-terminated OEG). The binding can be tuned by
106
either varying the percentage of the recognition elements on the particles or increasing
the ionic strength of the solution.19
Figure 7.1: (a) NTA-Ni2+ functionalized magnetic nanoparticles selectively bind to
histidine-tagged proteins (b) Ligands used for CdSe nanoparticles, and schematic
depiction of protein−nanoparticle interactions. [Reproduced from references 16 and 19]
More importantly, the protein-NPs complex is still active towards neutral and
cationic substrates, facilitating the application of these protein-NPs conjugates as
biocatalysts. In another work, our group demonstrated the stabilization of ChT at air-
water interface via noncovalent interaction to negatively charged Au NPs.20 In this case,
Au NPs exhibit stronger segregation to air-water interface compare to ChT; as a result
protecting ChT from interfacial denaturation. In addition to air-water interface, oil-water
107
biphasic reactions are also preferred for the bioprocessing of water insoluble chemicals
and simplify the purification process. Wang et al has studied that enzymes can be self-
assembled at the oil-water interface for further biotransformation.21 They conjugated ChT
with hydrophobic polymer polystyrene which has shown improved stability and activity
than native enzyme at the interface. This study opens up the avenue for the real world
biocatalytic application of enzyme.
Figure 7.2: (a) Interfacial assembly of polystyrene-conjugated ChT and β-galactosidase
at toluene-water interface (b) Schematic representation of ChT- NPs complex at air–
water interface. [Reference reproduced from 20 and 21]
7.2 Results and discussion
7.2.1 NP-enzyme assembly at oil-water interface
Herein, we report a direct and versatile technique for the creation of catalytic
microcapsules based on assembly of enzyme-NPs conjugates at the interface of oil-in-
water emulsions. A hybrid thin layer of enzyme-NPs conjugates on a template with high
surface to volume ratio would provide an ideal geometry for the generation of
biocatalysts for industrial applications. The assembly of the enzymes and nanoparticles
both stabilizes the emulsion and retains the surface availability of the enzymes for
108
catalytic reaction. These microcapsules were formed quickly and showed high enzymatic
activity, making them promising materials for biotechnology applications.
In the current study, we used β-galactosidase (β-gal),[22] a large, tetrameric
enzyme (17.5nm × 13.5nm × 9nm), with an overall negative surface charge (-0.25 × 10-2
C m-2) at neutral pH (pI= 5.3), as the enzyme component (Figure 7.3a). The
trimethylammonium-tetraethylene glycol functionalized ~7 nm Au nanoparticles with
positive surface charge (+0.35 × 10-2 C m-2) were synthesized to bind with enzymes
through electrostatic charge complementarity. These particles feature a tetraethylene
glycol unit in the ligand shell to minimize denaturation of the bound protein (Figure
7.3b).[23] As depicted in Figure 7.3c, positively charged Au nanoparticles associate with
negatively charged β-gal enzymes to produce reduced-charge conjugates. The subsequent
addition of “oil” (a 23:77 mixture of toluene and 1,2,4-trichlorobenzene, chosen to
provide buoyancy to the microcapsules in water) and vigorous mechanical agitation
produced stable microcapsules as a result of entrapment of enzyme-nanoparticle
conjugates at the oil-water interfaces.
109
Figure 7.3: (a) Structure of β-gal. (b) Chemical structure of cationic gold nanoparticle.
(c) Formation of enzymatic microcapsules through electrostatic assembly of enzymes and
nanoparticles in water followed by assembly of resulting enzyme-nanoparticle conjugates
at oil-in-water interfaces. Bottom left: A cross-sectional view of an enzymatic
microcapsule.
7.2.2 Characterization of enzyme immobilized microcapsules
The resulting enzyme-NPs conjugate stabilized microcapsules were analyzed by
optical microscope (OM) and diameter was found to be 40±15 µm (Figure 7.4a).
Transmission electron microscopy (TEM) image confirmed the presence of densely
110
packed nanoparticles at the microcapsule surface (Figure 7.4b). When using a FITC-
labeled β-gal distinct green fluorescence signal was seen form the microcapsules,
confirming the presence of β-gal at the microcapsule surface (Figure 7.4c).
Figure 7.4: (a) Optical micrograph of enzyme-nanoparticle conjugate stabilized
microcapsules fabricated at oil-in-water interfaces. (b) TEM images of a microcapsule at
low magnification and high magnification (inset). (c) Fluorescence microscopy image of
microcapsules synthesized using Au nanoparticles and FITC-labelled β-gal in water. d)
Free β-gal present in water after microcapsule construction using various molar ratios of
nanoparticles to β-gal.
Microcapsule formation was optimized via varying the NPs:enzyme ratio.
Incorporation of β-gal into the microcapsules was quantified by measuring the amount of
residual enzyme left in the water after the construction of the microcapsules, using a
Coomassie (Bradford) protein assay. As seen in Figure 7.5a, increasing the NPs: enzyme
ratio results in decreasing free β-gal, with essentially no β-gal in the water above a 1:1
NPs to enzyme ratio. The interfacial entrapment of enzyme-NPs conjugates was further
verified by analyzing the ζ-potential of enzyme-nanoparticle conjugates in the water
(Figure 7.b). As expected, enzyme-NPs conjugates showed an increase in ζ-potential
(from negative to positive) with increase in NPs: enzyme ratio, eventually reversing the
111
overall charge of the enzyme-NPs conjugate. It was seen that the enzyme-NPs conjugates
with overall low surface charge (~0.30 ×10-4 C m-2, see supporting information for charge
calculation) successfully go to the interface. This observation is supported by the
previous reports of Reincke et al, where carboxylic acid functionalized Au NPs were used
to create nanoparticle sheets at the interface by controlling the charge on the Au NPs
surface.[24] Control experiments done with only one component (β-gal or NPs) provided
unstable microcapsules, suggesting that the surface charge of the enzyme-nanoparticle
conjugates plays a crucial role in the formation of stable microcapsules. The high surface
energy associated with emulsion required much lower charge dense materials (i.e.
enzyme-NPs conjugates) compare to flat oil-water interface for its stabilization.
Figure 7.5: (a) Free β-gal present in water after microcapsule construction using various
molar ratios of nanoparticles to β-gal. (b) ζ-Potential measurements of enzyme-
nanoparticle conjugates synthesized by varying molar ratios of nanoparticles to β-gal.
7.2.3 Enzymatic activity of microcapsules
The utility of the enzyme-NPs microcapsules as catalysts was demonstrated via
enzyme activity assay. The β-gal present on the microcapsule surface retained catalytic
activity for hydrolysis of chlorophenol red-β-D-galactopyranoside (CPRG) substrate
112
(Figure 7.6 a-c), with β-gal in the enzyme-NPs microcapsules retaining 76% enzymatic
activity as compared to free β-gal (Figure 3). This efficiency is similar to that observed
for the monophasic activity of enzyme-NPs conjugates (84%), demonstrating that the β-
gal in the enzyme-NPs conjugates retains its catalytic activity both in water and at the oil-
water interface.
Figure 7.6: Activity assay of 0.5 nM β-gal in 5 mM phosphate buffer (a) free enzyme;
(b) enzyme-nanoparticle conjugate; (c) enzyme-nanoparticle conjugate stabilized
microcapsules. Inset: relative activity (free enzyme solution = 100%). (d) Enzymatic
cleavage of yellow colored CPRG at microcapsules shell by β-gal to produce red colored
chlorophenol-red; Stepwise color changes (from yellow to red) in chemical reaction by β-
gal present in the microcapsule shell: microcapsules are settled down at bottom of glass
vial.
7.3 Conclusion
In summary, we have demonstrated the successful integration of hybrid enzyme-
NPs conjugates with microcapsule structures. The fabrication process rapidly and
quantitatively immobilizes the enzyme on the microcapsule surface, with retention of
high catalytic activity. The catalytic reaction can only be performed in aqueous medium.
113
However, to create versatile microcapsules that will operate in organic medium also, we
need to crosslink either microcapsule surface or oil phase. Our method can be applicable
to other enzymes or proteins. For example, immobilization of Candida antartica lipase B
(CALB) onto microcapsules surfaces followed by crosslinking the NPs or oil part of the
microcapsules. CALB possesses extraordinary potential as a catalyst for a wide-range of
chemical transformations on both small organic molecules24 and more recently, for
polymerization reactions. We will also explore other valuable enzymes such as amidase,
carbamoylase, hydantoinase etc due their industrial applications. Next, we will also
immobilize multiple enzymes for sequential reactions.
7.4 Experimental Section
7.4.1Microcapsule preparation
Microcapsule preparation is based on supramolecular assembly of β-gal and
nanoparticles in water followed by addition of oil with vigorous mechanical agitation. In
detailed, β-gal (60 nM) was incubated with varying concentrations of cationic Au
nanoparticles in 5 mM phosphate buffer for 5 min to create enzyme-nanoparticle
conjugates. Next, 5 µL oil (mixture of toluene and 1,2,4 trichlorobenzene in 23:77 ratio)
was added to the aqueous enzyme-nanoparticle conjugate solution (200 µL) and
vigorously shaken by hand for ~ 60 s. After shaking, enzyme-nanoparticle conjugates
were entrapped at oil-water interface and appeared as pink colored microcapsules, which
very slowly settle down to the bottom of the aqueous solution. These microcapsules were
washed 2 times to remove free enzyme-nanoparticle conjugates in water and fresh
phosphate buffer was added prior to imaging in the Olympus IX51 microscope.
114
7.4.2Transmission Electron Microscopy (TEM)
TEM images were acquired on a JEOL 100CX operating at 100 keV. Samples
were drop cast onto a 300-mesh carbon-coated Cu grid, dried, and imaged.
7.4.3 Quantification of free enzyme in water after microcapsule fabrication
As synthesized microcapsule solutions were centrifuge in 1000 rpm for 1 min
resulting in clear supernatant that was analyzed for residual enzyme quantification in
solution using commercially available Coomassie (Bradford) protein assay kit (Thermo
scientific). Enzyme-nanoparticle conjugates did not settle down after the centrifugation at
1000 rpm for 1 min. -gal was used to create standard curve for this assay.
7.4.4 Activity Assays
The activity assays of free β-gal, enzyme-nanoparticle conjugates, and enzyme-
nanoparticle conjugate stabilized micocapsules were carried out in sodium phosphate
buffer (5 mM, pH 7.4). The final β-gal concentration in all activity assays was 1.0 nM.
The enzymatic hydrolysis was initiated by adding a CPRG substrate stock solution (100
µL of 15 mM) in the same buffer to the 100 µL enzyme-nanoparticle conjugates solution.
Enzymatic activity was followed by monitoring the absorption of the product formation
in every 30 s for 10 min at 595 nm using a micro-plate reader (spectraMax M5 from
Molecular Devices). The assays were performed in duplicate or triplicate, and averages
are reported. All activity assay experiments were done at 25 °C.
7.4.5 Synthesis of trimethylammonium-tetraethylene glycol functionalized ~7 nm Au
nanoparticles
Synthesis of cationic Au nanoparticles was performed in two steps process. First,
hydrophobic Au-nanoparticles were synthesized using reported procedure. Second, place
exchange on these hydrophobic nanoparticles was performed following prior report. In
brief, 100 mg of thiol ligand i.e. HS-C11-tetra(ethylene glycol)lyated
115
trimethylammonium bromide in 3 mL of methylene chloride was added to the 20 mg of
gold nanoparticles dispersed in 15 mL of methylene chloride. The resulting mixture was
stirred for 2 days forming black precipitation. The black precipitate was then washed 7
times with methylene chloride and finally dissolved in MilliQ water.
7.4.6 Labeling of β-gal with FITC
250 mL of fluorescein isothiocyanate isomoer I (FITC) in dimethyl sulfoxide with
concentration of 4 mg mL-1 was mixed with 900 mL of β-gal solution (2.8 mg mL-1) in
100 mM sodium bicarbonate buffer (pH 9.0), and shaken at room temperature in dark for
2h. The resulting FITC labeled enzyme was purified by size exclusion chromatography
with Sephadex G-25 as stationary phase and phosphate buffer (5 mM, pH 7.4) as mobile
phase. UV absorption spectra revealed that 14 fluorescein molecules were attached to one
β-gal.
7.4.7 ζ-Potential measurements
ζ-Potential of enzyme-nanoparticle conjugates was measured on a Malvern
Zetasizer Nano ZS in 5 mM phosphate buffer at pH 7.4. Samples were prepared by
varying enzyme and nanoparticles (20 nM) ratios from 1.0:0 to 1.0:2.0.
7.4.8 Measurement of surface charge density of nanoparticle, β-gal and enzyme-NP
conjugate
(a) Surface charge density of β-gal
We have used actual charge (-24.3 at pH ~7.4)21 and surface area (1530 nm2) of
enzyme from previous reports to calculate surface charge density of enzyme. The surface
charge density of enzyme at pH 7.4 is -0.25 × 10-2 C m-2.
116
(b) Surface charge density of nanoparticles and enzyme-nanoparticle conjugate
The surface charge density (σ) was calculated using reported procedure by
Reincke et al based on Huckel equation.
with
With Where, ζ is the zeta potential, A the surface area, R the radius, η the
viscocity and constant for the medium. The core radius and zeta potential of cationic Au
NPs are 3.5 nm and +17.8 mV respectively. Therefore, σ of cationic nanoparticles is
+0.35 × 10-2 C m-2. The hydrodynamic radius and ζ-potential of the enzyme-NPs
conjugates with 1:1 nanoparticle to enzyme ratio are 229 nm and -10 mV respectively.
Therefore, the calculated σ of enzyme-nanoparticle conjugates is -0.30 × 10-4 C m-2.
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