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Hybrid lipid-based nanostructures
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Hybrid lipid-based nanostructures
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Content
HYBRID LIPID-BASED NANOSTRUCTURES
by
Yasaman Dayani
________________________________________________
A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(CHEMICAL ENGINEERING)
August 2014
Copyright 2014 Yasaman Dayani
ii
Dedication
To my parents, Azita and Shahrokh, my husband, Alireza,
and my beloved son, Kian
iii
Acknowledgements
First and foremost, I would like to sincerely thank my advisor, Dr. Noah Malmstadt, for
his continuous support, guidance and encouragement throughout my PhD program. His
thoughtful advice and suggestions have been precious toward completion of this work. I
am grateful for the opportunities he has provided me to conduct my research in his
motivating research group. Thank you so much for believing in my abilities, supporting
me, and making this journey such a great experience for me. Without your help, this
would not have been possible.
I would also like to thank my doctoral committee members, Dr. Pin Wang and Dr. Moh
El-Naggar, for their efforts in evaluating my research and giving me valuable advice. I
would like to express my special thanks to Dr. Katherine Shing who kindly supported me
in the past few years at USC. I will never forget your kindness. Many thanks to Dr.
Malancha Gupta and Dr. Andrea Armani for their time and support during my PhD
program.
I am greatly thankful of Dr. Jae Jung and members of his research group, Dr. Hye-Ra Lee
and Dr. Samad Amini-Bavil-Olyaee, for collaboration in doing intracellular delivery
studies. I would also like to thank Dr. Terry Takahashi for his scientific advice and
guidance over these years at USC. I am thankful of Dr. Ralph Langen's group and also
Douglas Hauser in USC Norris Comprehensive Cancer Center for assistance in TEM
imaging. Many thanks to Dr. Nickolas Chelyapov and Dr. Shuxing Li for their guidance
and technical assistance as USC Nanobiophysics Core Facility managers.
iv
I am grateful of the current and former members of Dr. Malmstadt's group: Carson Riche,
Peichi Hu, Shalene Sankhagowit, Kristina Runas, Astro Yang, Gertrude Gutierrez, Krisna
Bhargava, Dr. Su Li, Dr. James Thompson, Dr. Jesper Hansen and Dr. Celine Billerit for
their support and friendship. You all made a memorable time for me at USC.
Finally I am extremely grateful of my dearest parents for their unconditional love and
support. They have always encouraged me to pursue my goals and provided me the best
opportunities in my life. Words cannot express what they have done for me all through
my life. I would like to express my appreciation to my husband, Alireza, for his continual
support, patience and love through this journey. I could not have made it through without
you by my side. You are a great support in my life.
v
Table of Contents
Dedication.............................................................................................................................. ii
Acknowledgements................................................................................................................ iii
List of Figures......................................................................................................................... ix
Abstract................................................................................................................................. xii
Chapter 1: Introduction..................................................................................................... 1
1.1 Overview and Motivation................................................................................................. 1
1.2 Biomimetic Lipid Bilayer.................................................................................................. 4
1.2.1 Vesicles....................................................................................................................5
1.2.1.1 Liposomes Importance and a Drawback in Using Them.................................... 7
1.2.2 Planar Supported Lipid Membranes........................................................................... 8
1.2.3 Tethered Planar Lipid Bilayers.................................................................................. 10
1.2.4 Tethered Liposomes.................................................................................................. 11
1.2.4.1 Progress toward Tethering of Liposomes Using DNA Anchors.......................... 12
1.3 Biohybrid Nanostructures................................................................................................. 13
1.3.1 Fabrication of Hybrid Nanostructures....................................................................... 14
1.3.2 Classification of Hybrid Structures........................................................................... 16
1.4 Lipid bilayer-based Hybrid Nanostructures....................................................................... 17
1.5 Poly(ethylene) glycol....................................................................................................... 20
1.5.1 Poly(ethylene) glycol Free Radical Polymerization Mechanism............................... 22
1.6 Carbon Nanotubes............................................................................................................. 23
vi
Chapter 2: Lipid Bilayers Covalently Anchored to Carbon Nanotubes...................... 25
2.1 Introduction...................................................................................................................... 25
2.2 Materials and Methods..................................................................................................... 28
2.2.1 Materials.................................................................................................................. 28
2.2.2 Synthesis of Lipid-modified Multi-walled Carbon Nanotubes................................ 29
2.2.3 FTIR Analysis.......................................................................................................... 30
2.2.4 Liquid-Liquid Extraction......................................................................................... 30
2.2.5 Lipid Bilayer Fabrication......................................................................................... 30
2.2.6 Protein Incorporation in Lipid Bilayer-coated Nanotubes....................................... 31
2.2.7 Sample Observation with Fluorescence Confocal Microscopy............................... 31
2.2.8 Fluorescence Quenching.......................................................................................... 32
2.2.9 Transmission Electron Microscopy (TEM)............................................................. 33
2.2.10 Fluorescence Anisotropy Measurements............................................................... 33
2.3 Results and Discussion.................................................................................................... 34
2.3.1 Solubility of Lipid-modified MWCNTs.................................................................. 35
2.3.2 FTIR Spectroscopy.................................................................................................. 37
2.3.3 Lipid Bilayer Formation on MWCNTs.................................................................... 39
2.3.4 Alpha-hemolysin Insertion in Lipid Bilayers on Nanotube Surfaces...................... 42
2.3.5 Lipid Bilayer Fluorescence Quenching................................................................... 44
2.3.6 Lipid Bilayer Fluidity.............................................................................................. 45
2.4 Conclusions...................................................................................................................... 47
vii
Chapter 3: Liposomes with Double-stranded DNA Anchoring the Bilayer to
Hydrogel Core.................................................................................................................... 48
3.1 Introduction....................................................................................................................48
3.2 Materials and Methods....................................................................................................... 50
3.2.1 Materials................................................................................................................ 50
3.2.2 PEG Hydrogel Formation in Liposomes.................................................................. 51
3.2.3 Conjugation of Lipid Bilayer to PEG Hydrogel Using DNA Linkages.................. 52
3.2.4 Size Exclusion Chromatography (SEC)................................................................... 53
3.2.5 Confirming DNA Location...................................................................................... 54
3.2.6 Transmission Electron Microscopy (TEM)............................................................. 54
3.2.7 Dynamic Light Scattering (DLS)............................................................................. 55
3.2.8 Size Changes of Nanogels Exposed to Organic Solvent............................................. 55
3.2.9 DNase Treatment of Cholesterol-dsDNA-anchored Liposomes................................. 56
3.2.10 Release Study......................................................................................................... 56
3.3 Results and Discussion.................................................................................................... 57
3.3.1 Fabrication of Hydrogel-anchored Liposomes with dsDNA Anchors........................ 57
3.3.2 Characterization of Fabricated Nanoparticles........................................................... 59
3.3.2.1 SEC and TEM.................................................................................................. 59
3.3.2.2 DLS.................................................................................................................. 62
3.3.3 Surface Accessibility of Cholesterol-dsDNAs........................................................... 62
3.3.4 dsDNAs Localization on Nanogel Surface................................................................. 63
3.3.5 Stability of Hydrogel-anchored Liposomes............................................................... 65
3.3.6 Encapsulation Efficiency in Hydrogel-containing Liposomes................................... 67
viii
3.3.7 Conclusions.............................................................................................................. 68
Chapter 4: pH sensitive, Hydrogel-anchord Liposomes for Peptide eptide
Delivery ................................................................................................................................ 70
4.1 Introduction...................................................................................................................... 70
4.2 Materials and Methods.................................................................................................... 73
4.2.1 Materials.................................................................................................................. 73
4.2.2 Fabrication of Hydrogel-anchored Liposomes........................................................... 74
4.2.3 vif2 Peptide Encapsulation in Hydrogel-anchored Liposomes................................... 75
4.2.4 Size Exclusion Chromatography of vif2-containing Liposomes ............................... 76
4.2.5 Determination of pH Sensitivity of Liposomes......................................................... 76
4.2.6 Lipid Bilayer Lysis by Cholesterol-dsDNA-nanogels................................................ 77
4.2.7 Cellular Uptake of Hydrogel-anchored Liposomes.................................................... 78
4.2.8 Confocal Microscopy of Cells................................................................................... 79
4.3 Results and Discussion..................................................................................................... 80
4.3.1 vif2 Peptide and Its Mechanism of Action................................................................ 81
4.3.2 vif2 Encapsulation in Liposomes............................................................................... 82
4.3.3 Destabilization of pH-sensitive Liposomes at Acidic pH.......................................... 83
4.3.4 Lipid Bilayer Lysis by Cholesterol-dsDNA-nanogels............................................... 85
4.3.5 Cellular Internalization of Peptide-containing Liposomes........................................ 87
4.3.6. Conclusions............................................................................................................. 90
Chapter 5: Conclusions and Recommendations.............................................................. 92
References............................................................................................................................. 97
ix
List of Figures
Figure 1.1 Models of artificial lipid bilayers...................................................................... 5
Figure 1.2 Schematic representation of basic structures and various types of
liposomes……………………………………………………………….......... 6
Figure 1.3 Strategies for preserving transmembrane proteins intact in planar lipid
bilayers.............................................................................................................. 9
Figure 1.4 Schematic diagram of the tethering of DNA-tagged liposomes to
supported lipid bilayer presenting complementary DNA………………......... 13
Figure 1.5 Schematic of different techniques used for fabrication of hybrid
structures........................................................................................................... 17
Figure 1.6 Lipid-based hybrid nanostructures composed of synthetic metarials, such as
hydrogel and carbonnanotubes and lipid bilayers……………………........ 19
Figure 1.7 Structure of poly(ethylene glycol)………………………………………......... 20
Figure 1.8 Free radical polimerization mechanism of PEG-DA using APS and
TEMED…………………………………………………………………........ 22
Figure 1.9 Structure of single-walled carbon nanotubes……………………………........ 23
Figure 2.1 Synthesis of lipid-MWCNT conjugates in a carbodiimide-mediated
reaction…………..……………………………………………………......... 35
Figure 2.2 Liquid-liquid extraction of MWCNTs in a CHCl
3
/H
2
O system…………........ 36
Figure 2.3 Schematic drawing of the insertion of protein into the bilayer of a lipid-
modified MWCNT (Front view)…………………………………………....... 37
Figure 2.4 FTIR spectra of (A) COOH-MWCNTs; (B) DPPE; (C) MWCNT-DPPE
conjugate…………………………………………………………………....... 38
Figure 2.5 Photographs of sonicated lipid bilayer-coated MWCNTs in buffer after
(A) 1 day, and (B) 60 days………………………………………………........ 39
Figure 2.6 Confocal images of MWCNTs coated with NBD-DPPC-containing
lipid bilayers in buffer solution…………………………………………......... 40
Figure 2.7 TEM images of (A) COOH-MWCNTs and (B and C) bilayer-coated
MWCNTs after removal of excess lipid. (D) Closer view of COOH
MWCNTs; (E) an enlarged image of part of the nanotube wall…………....... 41
x
Figure 2.8 Confocal fluorescence images of (A) NBD-labeled lipids on MWCNTs
coated with lipid bilayers by sonication………………………………......….. 43
Figure 2.9 Normalized fluorescence intensity of bilayer-coated MWCNTs before
and after the addition of sodium dithionite quencher…………………......…. 45
Figure 2.10 Fluorescence anisotropy of lipid bilayers conjugated to the surface of
MWCNTs as a function of temperature……………………………......…... 46
Figure 3.1 Schematic representation of a lipid bilayer anchored to a PEG hydrogel by a
DNA linker……….……………………………………………………......….. 57
Figure 3.2 (A) Normalized SEC chromatograms tracking rh-MA in different
liposome species……………………………………………………......……. 60
Figure 3.3 Normalized SEC chromatograms tracking rh-DPPE fluorescence…......……. 61
Figure 3.4 Normalized chromatograms tracking H33342 in dsDNA-anchored
liposomes…………………………………………………………......…….... 63
Figure 3.5 Changes in fluorescence intensity (ΔFI, red symbols) and maximum
emission wavelength (Δλ, blue symbols) of 0.02 nM H33342 dye in the
presence of ssDNA-nanogels (triangle) and dsDNA-nanogels (circle) as a
complimentary DNA species (DNA-1) is added to the system…...……......... 64
Figure 3.6 Dye dequenching experiment that shows stability of hydrogel-anchored
liposomes…………………………………………………………………...... 65
Figure 3.7 (A) Dye release profiles from hydrogel-anchored liposomes, hydrogel
liposomes without anchors, and liposomes without hydrogel (plain
liposomes) incubated in 20% serum…………………………………......…... 66
Figure 3.8 Liposomes size changes in 20% FBS………………………………….....….. 67
Figure 4.1 Cell internalization of pH-sensitive liposomes carrying vif2 peptide…......…. 81
Figure 4.2 Normalized SEC chromatograms tracking FITC-vif2 in various
liposomes with dsDNA anchors………………………………………......….. 82
Figure 4.3 pH-induced leakage of nano-scale liposomes composed of 70% (wt%)
DOPE and 30% (wt%) DOGS...………………………………………......…. 85
Figure 4.4 Release of dye from plain DOPE/DOPC liposomes incubated with chol-
dsDNA-nanogels made of different concentrations of dsDNAs, at 37°C and
pH4.……………………………………………………………….................... 86
xi
Figure 4.5 Fluorescence confocal microscopy images of A549-IFITM3 cells
incubated with liposomes with Rhod-PE lipid at various times……......……. 88
Figure 4.6 Fluorescence confocal microscopy images of A549-IFITM3 cells
incubated with hydrogel-anchored liposomes encapsulating FITC-
peptide at various times………………………………………………......….. 88
Figure 4.7 Fluorescence confocal microscopy of A549-IFITM3 cells incubated for 3h
with NBD-labeled liposomes encapsulating rh-MA crosslinked to PEG
hydrogel............................................................................................................ 89
Figure 4.8 Fluorescence confocal microscopy of A549-IFITM3 cells incubated with
NBD-labeled liposomes encapsulating rhodamine-hydrogels……………….. 90
xii
Abstract
Biological membranes serve several important roles, such as structural support of
cells and organelles, regulation of ionic and molecular transport, barriers to non-mediated
transport, contact between cells within tissues, and accommodation of membrane proteins.
Membrane proteins and other vital biomolecules incorporated into the membrane need a
lipid membrane to function. Due to importance of lipid bilayers and their vital function in
governing many processes in the cell, the development of various models as artificial
lipid membranes that can mimic cell membranes has become a subject of great interest.
Using different models of artificial lipid membranes, such as liposomes, planar
lipid bilayers and supported or tethered lipid bilayers, we are able to study many
biophysical processes in biological membranes. The ability of different molecules to
interact with and change the structure of lipid membranes can be also investigated in
artificial lipid membranes. An important application of lipid bilayer-containing interfaces
is characterization of novel membrane proteins for high throughput drug screening
studies to investigate receptor-drug interactions and develop biosensor systems.
Membrane proteins need a lipid bilayer environment to preserve their stability and
functionality. Fabrication of materials that can interact with biomolecules like proteins
necessitates the use of lipid bilayers as a mimic of cell membranes.
The objective of this research is to develop novel hybrid lipid-based nanostructures
mimicking biological membranes. Toward this aim, two hybrid biocompatible structures
are introduced: lipid bilayer-coated multi-walled carbon nanotubes (MWCNTs) and
hydrogel-anchored liposomes with double-stranded DNA anchors. These structures have
xiii
potential applications in biosensing, drug targeting, drug delivery, and biophysical studies
of cell membranes.
In the first developed nanostructure, lipid molecules are covalently attached to the
surfaces of MWCNTs, and then, using a sonication process, a uniform lipid bilayer that
supports the incorporation of membrane proteins is formed. These bilayer-coated carbon
nanotubes are highly dispersible and stable in aqueous solution, and they can be used in
development of various biosensors and energy producing devices.
In the other hybrid nanostructure, the lipid bilayer of a liposome is covalently
anchored to a biocompatible poly(ethylene) glycol (PEG) hydrogel core using double-
stranded DNA (dsDNA) linkers. Release studies shows that nano-size hydrogel-anchored
liposomes are exceptionally stable, and they can be used as biomimetic model
membranes that mimic the connectivity between the cytoskeleton and the plasma
membrane. After lipid bilayer removal, dsDNA linkers can provide programmable
nanogels decorated with oligonucleotides with potential sites for further molecular
assembly. These stable nanostructures can be useful for oligonucleotide and drug delivery
applications.
The developed hydrogel-anchored liposomes are exploited for encapsulation and
intracellular delivery of therapeutic peptide. Peptides with anti-cancer properties are
successfully encapsulated in hydrogel core of pH-sensitive liposomes during rehydration
process. Liposomes release their cargo at acidic pH. Confocal microscopy confirms the
intracellular delivery of liposomes through an endocytotic pathway.
1
Chapter 1: Introduction
1.1 Overview and Motivation
Constructing advanced nanoscale structures that can mimic biological systems has
been a subject of great interest in recent years. Since many biomolecules are functional in
the presence of lipid membranes, we require biomimetic structures that support these
molecules in their functional form. Phospholipid bilayers mimicking the cellular
membrane are the only structures that can incorporate biomolecules and provide
integrating platforms with nanomaterials. These nanoscale, biocompatible structures
usually show specific physical and chemical properties that could lead us to develop new
devices for potential use in biosensing and drug delivery.
The focus of this dissertation is developing and characterizating hybrid
nanostructures derived from biomimetic lipid membranes and synthetic materials, such as
carbon nanotubes and hydrogels.
Carbon is one of the most versatile materials showing a variety of stable forms with
unique properties allowing for various applications. The unique physical and electrical
properties of carbon nanotubes make them exciting materials for applications in various
fields such as bioelectronics and biosensing. Due to the poor water solubility of carbon
nanotubes, functionalization for such applications has been a challenge. Of particular
need are functionalization methods for integrating carbon nanotubes with biomolecules
and constructing novel hybrid nanostructures for bionanoelectronic applications. In this
dissertation, we introduce a novel method for the fabrication of dispersible,
biocompatible carbon nanotube-based materials using biomimetic lipid bilayers (Chapter
2). Multi-walled carbon nanotubes (MWCNTs) are covalently modified with primary
2
amine-bearing phospholipids in a carbodiimide-activated reaction. These modified
carbon nanotubes have good dispersibility in nonpolar solvents. Fourier transform
infrared (FTIR) spectroscopy shows peaks attributable to the formation of amide bonds
between lipids and the nanotube surface. Simple sonication of lipid-modified nanotubes
with other lipid molecules leads to the formation of a uniform lipid bilayer coating the
nanotubes. These bilayer-coated nanotubes are highly dispersible and stable in aqueous
solution. Confocal fluorescence microscopy shows labeled lipids on the surface of
bilayer-modified nanotubes. Transmission electron microscopy (TEM) shows the
morphology of dispersed bilayer-coated MWCNTs. Fluorescence quenching of lipid-
coated MWCNT confirms the bilayer configuration of the lipids on the nanotube surface.
The membrane protein α-hemolysin spontaneously inserts into the MWCNT-supported
bilayer, confirming the biomimetic membrane structure. Fluorescence anisotropy
measurements demonstrate the fluidity of the CNT-bound lipid bilayer. These membrane
biomimetic nanostructures are a promising platform for the integration of CNT-based
materials with biomolecules.
Liposomes—spherical lipid bilayers—are widely used as models for biological
membranes. They are important biomolecular nanostructures for handling membrane-
associated molecules in the lab and delivering drugs in the clinic. In addition to their
biomedical applications, they have been widely used as model cell membranes in
biophysical studies. They are potential structures for drug delivery and sustained release
of therapeutic materials. Surface modification of liposomes allows targeted and efficient
delivery of drugs. However, fragility and short lifetime limit the applicability of
liposomes. To date, there have been efforts to develop hybrid nanostructures that
3
combine advantages of hybrid materials and lipid bilayers to produce more robust
liposomes. For example, lipid-polymer hybrid nanostructures merge the strength of each
individual material in one structure. Those polymer-lipid structures mostly suffer multi-
step procedures for fabrication and lack of biocompatibility due to toxic polymers. In
chapter 3, we develop a stable liposome-based hybrid nanostructure that mimics the
attachment of membrane-resident molecules to the cytoskeleton. In a self-assembly
procedure, the liposome bilayer membrane is covalently anchored to a biocompatible
poly(ethylene) glycol (PEG) hydrogel core using short double-stranded DNA (dsDNA)
linkers. Using DNA as a linker between the bilayer and a hydrogel core allows for
temperature-dependent release of the anchoring interaction, produces polymer nanogels
with addressable hybridization sites on their surface, and provides a prototype structure
for potential future oligonucleotide drug delivery applications. Size exclusion
chromatography (SEC) of intact and surfactant-treated nanoparticles confirms the
formation of anchored hydrogel structures. The location of dsDNA groups at the
hydrogel-bilayer interface is confirmed with a fluorescence assay. Fluorescence
dequenching experiments demonstrate the high stability of hydrogel-anchored liposomes
even in serum.
In chapter 4, we describe the intracellular delivery of hydrogel-anchored
liposomes encapsulating a therapeutic peptide. The pH sensitive liposomes that enhance
intracytoplasmic drug delivery through an endocytotic pathway are developed and
characterized. These liposomes protect a peptide during cell internalization process and
allow the release of their cargo in a pH-triggered pathway at low pH sites. Fluorescence
4
confocal microscopy shows successful delivery of peptide-containing liposomes through
an endocytotic pathway.
1.2 Biomimetic Lipid Bilayers
Cell membranes composed of different types of lipids in the form of two leaflets
plus functional proteins have important roles in many cellular processes (Lee et al. 2013).
In addition to separating organelles in a cell, they allow regulated transport of vital
compounds, signal transduction, trafficking, intracellular organiztion and response to the
extracellular matrix (Ariga et al. 2006; Lee et al. 2013). The functions of membrane
proteins can be reproduced in the laboratory using biomimetic interfaces that consist of
artificial bilayer membranes with embedded membrane proteins (Chan and Boxer 2007).
The architectural components of biological membranes are lipid molecules consisting of
hydrophilic head groups and hydrophobic tails. These amphiphilic molecules form closed
spherical structures in an aqueous environment by self-assembly. Due to the complexity
of natural membranes, there have been many efforts to develop simpler model systems
whose size, geometry, and composition can be controlled accurately (Chan and Boxer
2007). These model membranes can retain the structure of biological membranes, but by
simplifying the system, they allow studies of individual components and visualization of
their organization and dynamics in lipid bilayer membranes (Chan and Boxer 2007).
Model lipid bilayers allow investigation of biological processes and provide information
for processes such as ligand-receptor interactions and cellular signaling (Castellana and
Cremer 2006). As shown in Figure 1.1, to date several model membranes such as vesicles,
planar supported lipid membranes and tethered lipid membranes have been introduced
5
(Figure 1.1). Vesicle-like bilayers can be free-standing or tethered to supports; planar
supported bilayers can either directly interact with a solid substrates or be tethered to the
substrate (Chan and Boxer 2007) (Figure 1.1). This wide range of assemblies allows the
study of bilayers using different techniques and methods.
Figure 1.1. Models of artificial lipid bilayers. (A) Planar supported lipid bilayer;
(B) Tethered planar lipid bilayer; (C) Vesicle; and (D) Tethered vesicles.
1.2.1 Vesicles
Vesicles are commonly used as models of biological lipid membranes. They form
when phospholipid molecules are exposed to an aqueous solution. The polar, hydrophilic
head groups of phospholipids are in contact with aqueous solution, while hydrocarbon
tails organize to prevent contact with aqueous phase. As shown in Figure 1.2, there are
various size ranges of vesicles as model systems for natural membranes: small
unilamellar vesicles (SUVs) with diameters of tens of nanometers, large unilamellar
vesicles (LUVs) with diameters bigger than 100 nm, and giant unilamellar vesicles
(GUVs) with diameters of tens of microns. These vesicles can be unilamellar or
A B
C
D
A B
C
D
6
multilamellar. Unilamellar vesicles contain a single lamella with the hydrophobic tails
from each leaflet in contact, while multilamellar vesicles (MLVs) contain multiple
concentric lamellar layers with a thin water layer between lipid layers (Figure 1.2).
Figure 1.2. Schematic representation of basic structures and various types of liposomes. The size of small
unilamellar vesicles (SUVs), large unilamellar vesicles (LUVs) and giant unilamellar vesicles (GUVs) are
tens of nanometer, >100 nm, and tens of microns, respectively.
SUVs are commonly prepared by sonication of MLVs and extrusion of them
through polycarbonate filters; GUVs are usually formed by the electroformation method;
and LUVs are formed by combination of freeze/thaw cycles followed by extrusion. LUVs
along with GUVs are appropriate model membranes for reconstitution of membrane
proteins due to the lower membrane curvature (Olson et al. 1979). GUVs are also used
for study of diffusion of various components across membrane (Kahya et al. 2003) and
lipid phase behavior in membrane (Korlach et al. 1999).
7
1.2.1.1 Liposomes Importance and a Drawback in Using Them
The unique structure of liposomes—small vesicles,with an amphiphilic lipid
bilayer and hydrophilic aqueous core—makes them interesting tools for encapsulating
hydrophilic compounds in the aqueous area, hydrophobic compounds within the lipid
bilayer and attaching amphiphilic compounds at the aqueous-lipid bilayer interface.
Liposomes possess several other important advantages. They are flexible structures
because different types of lipids such as cationic, anionic and neutral can be used in their
formulation (MacKinnon et al. 2009). Liposome surface modification with biomolecules
such as antibodies affords targeted drug delivery while capturing large volumes of
components in their cores (MacKinnon et al. 2009). Generally, liposomes provide a free-
standing membrane for the study of lipid properties by various techniques. They can be a
good environment for studying reconstituted protein function because they avoid direct
contact with a solid substrate, and they allow solution phase access to all surfaces of the
membrane (Castellana and Cremer 2006). Recently, liposomes have been used to handle
integral membrane proteins such as G protein coupled receptors (GPCRs), which are
important targets for pharmaceutical treatments. They also have many applications in
drug delivery, gene therapy and drug targeting (Hellwich and Schubert 1995; Lasic 1998;
Leserman 2004). An important drawback of liposomes is their limited stability and
susceptibility to mechanical and chemical disruption. Different methods have been
recommended to increase the lifetime of classical liposomes. These methods include the
use of photopolymerizable phospholipids in the lipid mixture (Fendler 1984; Guo et al.
2009; Hub et al. 1980), coating the liposome surfaces with polymer compounds (Hayashi
et al. 1999; Ringsdorf et al. 1993; Yamazaki et al. 1999) and encapsulating polymers in
8
the interior of liposomes (Schillemans et al. 2006). Stauch et al. have shown that creating
a cross-linked polymer network inside the liposomes can significantly increase the
stability of liposome in sodium cholate solution if the polymer is linked to the inner
monolayer of the bilayer by anchoring group (Stauch et al. 2002). Other research groups
reported longer circulation time for polymerized liposomes in blood and higher stability
for them against Triton X-100 (Guo et al. 2009; Papahadjopoulos et al. 1991). In chapter
3, we discuss other methods that have been developed for improving the stability of
liposomes, and we introduce our new approach for fabricating liposomes with higher
stability.
1.2.2 Planar Supported Lipid Membranes
Planar supported lipid bilayers which are usually formed by small vesicle fusion on
hydrophilic solid surfaces such as glass, quartz, or gold, have some advantages, such as
ease of preparation, stability, and availability to different kinds of surface sensitive
techniques for characterization (Chan and Boxer 2007). McConnell and coworkers first
deposited lipid membranes directly on solid supports in early 80s (Tamm and McConnell
1985). Although supported lipid bilayers (SLBs) have many advantageous, and they are
excellent sensor platforms, their inability to incorporate membrane proteins in their
functional form limits their application (Chan and Boxer 2007). Direct contact between
transmembrane proteins, especially those with large peripheral domains, and the solid
support causes protein denaturation and dysfunction (Castellana and Cremer 2006).
Several approaches have been developed to overcome this problem including assembling
bilayers on softer supports like polymer cushions (Castellana and Cremer 2006; Spinke et
9
al. 1992; Tanaka 2006), tethering of the membrane to a lipid-presenting polymer
(Naumann et al. 2002; Purrucker et al. 2004; Seitz et al. 2001) and using long-chain
tethers (Figure 1.3) (Atanasov et al. 2006; Yoshina-Ishii and Boxer 2003). In 1992, Knoll
et al. developed a method to appropriately incorporate transmembrane proteins into the
supported bilayer and avoid direct contact of lipid membrane to solid support (Spinke et
al. 1992). They used thin polymer film which could couple bilayers with solid substrates
composed of various materials. In 1997, Boxer et al. lead the new way to partition
supported phospholipid bilayer in micro-meter scale corrals of lithographically patterned
grids of aluminum oxide or gold on oxidized silicon substrates (Groves et al. 1997). Then,
Cremer et al. developed a method for generation of rapid screening arrays and sensor
devices for SLBs containing biological molecules such as peptides, receptors and
membrane proteins to mimic many of the properties of the cell surface (Cremer and Yang
1999).
Figure 1.3. Strategies for preserving transmembrane proteins intact in planar lipid bilayers. (A) Denatured
protein, which is in direct contact with solid support; (B) Polymer cushion for preserving nature of
membrane protein; and (C) Bilayers tethered to solid support (Castellana and Cremer 2006; Chan and
Boxer 2007).
10
In the most recent studies, Boxer et al. have developed methods for tethering lipid
membranes using DNA linkers. In one approach, DNA tethers are directly coupled to the
solid substrate by click chemistry, and GUVs containing complementary DNA are
hybridized with DNA on the surface (Chung et al. 2009). In another approach, DNA-
conjugated lipids are incorporated into an SLB on the solid substrate, and GUVs
containing complementary DNA hybridizes with these mobile DNAs (Chung et al. 2009).
Using metal supports in planar supported bilayers allows performing electrical
measurements to monitor activity of embedded proteins or ion channels into the bilayar
(Chan and Boxer 2007).
1.2.3 Tethered Planar Lipid Bilayers
In physisorbed systems such as assembly of lipid bilayers on polymer cushions
used for decoupling the lipid membrane from a solid substrate, weak interactions between
the phospholipid bilayer and the polymer support result in an unstable system (Castellana
and Cremer 2006). This could be overcome by either covalently attaching the polymer
layer to the bilayer or using anchor lipids that effectively tether the membrane to the
polymer layer (Castellana and Cremer 2006). As mentioned in section 1.2.2, a tethered
bilayer membrane provides a gap between the substrate and membrane proteins
associated with the lipid membrane and allows proteins to have similar lateral diffusion
coefficients to membrane proteins in cell membranes (Wagner and Tamm 2000). Several
strategies have been used for assembly of tethered bilayer membranes. The most common
one is adsorbing or binding of polymer cushion to the prefunctionalized solid support
with a reactive layer that form a covalent bond with corresponding reactants on the
11
polymer (Naumann et al. 2002). Tethered bilayer membranes provide a potential platform
for very detailed studies of the structure-function relationships of model membrane and
reconstituted proteins (Bally et al. 2010). This model facilitates investigation of the
electrical and transport properties of the membrane and components within the membrane
(Meier et al. 2010).
1.2.4 Tethered Liposomes
Liposomes with self-assembled lipid bilayer structure, allowing protein flexibility
and movement, can perform as supports for membrane proteins. In this type of lipid
bilayer, membrane proteins are preserved from denaturation resulting from direct contact
with solid supports. To date, several techniques, such as using self-assembled lipid
monolayers, poly(ethylene glycol) (PEG)-based polymeric layers or supported lipid
bilayers have been introduced to tether liposomes to solid substrates (Bally et al. 2010).
Tethered liposomes give us the opportunity to study individual vesicles in a parallel
manner via to binding to specific locations on a substrate (Bolinger et al. 2004). They
could also provide appropriate environment for investigation of controlled interaction
between vesicles and membrane proteins.
Surface coupling using the anchoring molecules, such as biotin-BSA and biotin-
DNA is one of the usual techniques used for formation of tethered lipid bilayers
(Svedhem et al. 2003; Vockenroth et al. 2007; Yoon et al. 2006). Biotin-avidin chemistry
has been used for tethering of small vesicles to a supported lipid bilayer to study single
protein molecules (Boukobza et al. 2001; Rhoades et al. 2003; Stamou et al. 2003). In an
alternative approach, a biotin-PEG-lipid anchor has been used to tether LUVs to a
12
neutravidin-coated solid surface (Bolinger et al. 2004). In the most recent approaches,
lipid vesicles have been tethered to solid supported lipid bilayers using DNA anchor
molecules (Castellana and Cremer 2006; Yoon et al. 2006). Tethered liposome
architectures enable us to take advantages of a supported lipid bilayer and free standing
vesicles in a single, individual structure.
1.2.4.1 Progress toward Tethering of Liposomes Using DNA Anchors
The goal to immobilize liposomes using tag molecules, connecting them to the
solid substrates, has motivated researchers to create appropriate linkers. In one recent
strategy, DNA linkers have been used for tethering of liposomes to the solid substrates. In
this approach, liposomes with specific DNA sequences can recognize linkers with
specific sequence on the substrate. Due to self-assembly and molecular recognition,
oligonucleotides have been the subject of much interest in recent years. They can act as
controllable and programmable building blocks for the accurate spatial arrangement of
biomolecules such as proteins and ligands, cells and nanoparticles on artificial organs
(Bunge et al. 2009).
Because of high applicability of liposomes as drug delivery systems, different
methods for tagging of liposomes with DNA have been introduced. The Boxer group has
pioneered constructing of DNA-containing liposomes tethered to supported bilayers by
DNA hybridization (Yoshina-Ishii et al. 2005). The sequence specificity of DNA allows
fabrication of arrays of vesicles using patterned bilayers (Chan and Boxer 2007). Since
DNA-tethered liposomes can diffuse in two dimensions in a plane parallel to the surface,
the interaction between the vesicle and reactive membrane components such as DNA or
13
proteins can be studied as vesicles diffuse or collide with each other (Chan and Boxer
2007). Using DNA linkers results in sequence specific tethering of vesicles to the SLB
and enables visualization of vesicles individually by fluorescence microscopy (Figure
1.4) (Yoshina-Ishii and Boxer 2003).
Figure 1.4. Schematic diagram of the tethering of DNA-tagged liposomes to supported lipid bilayer
presenting complementary DNA (Yoshina-Ishii and Boxer 2003).
The Boxer group also developed a method for covalent conjugation of liposomes
with DNA linkers. In this approach, liposomes displaying sense and antisense DNAs
were covalently conjugated to supporting lipid bilayer using a click chemistry reaction
(van Lengerich et al. 2010).
1.3 Biohybrid Nanostructures
Biohybrid nanomaterials are derived by the assembly of molecular species of
biological origin and synthetic substrates through interactions on the nanometer scale
(Ruiz-Hitzky et al. 2008). The development of these materials is result of combination of
fields such as the life sciences, material science, and nanotechnology. There are number
of differences between properties of synthetic substances and biological materials.
14
Synthetic materials can be mechanically robust and provide an overall structure to the
natural objects, while biological materials are more soft and flexible, and they are capable
of forming everchanging assemblies (Ariga et al. 2006). Biohybrid nanomaterials can
provide new structures with special functionalities. Therefore, assembly of structures that
combine both synthetic and biological materials to take advantage of both substances has
taken special attention. These nanostructure assemblies with unique physical, chemical,
optical, magnetic and mechanical properties have broad range of applications in tissue
engineering, drug delivery systems, biosensing devices, biocatalysis, and green
nanocomposites (Pandey et al. 2005; Patil and Mann 2008; Ruiz-Hitzky et al. 2010).
Recently, there have been efforts to create advanced hybrid nanostructures with improved
functional and structural properties mimicking biological systems in nature.
1.3.1 Fabrication of Hybrid Nanostructures
Several interesting approaches have been introduced for the synthesis of biohybrid
structures. These structures can be made by the combination of natural polymers, such as
polysaccharides, biologically produced polyesters, RNA and DNA, polypeptides,
globular proteins, and enzymes, with synthetic substrates, such as silica and
phyllosilicates, layered double hydroxides (LDHs), phosphates, metals and carbon
nanotubes (Ruiz-Hitzky et al. 2008; Ruiz-Hitzky and Darder 2006). Recently, there have
been attempts to construct biohybrid nanostructures by immobilization of biomolecules
on nanoscale synthetic substrates. These hybrid assemblies have practical applications
including entrapment of materials, bone tissue regeneration, bone implant fabrication,
controlled release and injection of bioactive molecules such as drugs and genes (Bonderer
15
et al. 2008; Firkowska et al. 2008; Ruiz-Hitzky et al. 2008). Important factors that should
be considered in designing of biohybrid nanomaterials are appropriate synthetic route and
chemical composition in respect to the desired use. There is a huge variety of
nanostructured materials that can be used as appropriate supports for biomaterial
immobilization (Ruiz-Hitzky et al. 2010; Sanchez et al. 2011). The three major classes of
support materials are porous solids, micro or nanoparticulated solids and soft networks
(Ruiz-Hitzky et al. 2010). The first two classes of substrates are rigid. These types of
substrates can provide nanostructures with well-defined dimensions and geometries
incorporating biorelated components in nanoscale spaces in which biomolecules have
restricted molecular motion (Ruiz-Hitzky et al. 2010). Construction of composite
materials using peptides or amino acids and mesoporous silicates is a good example for
inclusion of biological species into porous solids (Zhang et al. 2004). Single and multi-
walled carbon nanotubes assembled with different biological molecules are also
functional constructions in category of nanoparticulated solids (Ruiz-Hitzky et al. 2010).
Compared to rigid structures, soft networks have more flexibility in adapting various
morphologies and structures increasing the biocompatibility of these nanostructures.
These soft structures can preserve a partially hydrated environment, similar to hydrogels,
and provide more biocompatible structures (Fennouh et al. 2000). The sol-gel process
based on polymerization of silica precursors is an example of processes used for
formation of soft networks (Boettcher et al. 2007). Cell-like structures (cerasome) is a
type of bio-hybrid structures have been developed using lipid and surfactant-bearing
alkoxysilanes to produce biomimetic silica nanohybrids (Kim et al. 1998). The most
common synthetic components of bionanohybrids are carbon particles, metal oxides and
16
hydroxides, silica, silicates, carbonates and phosphates (Desimone et al. 2005; Gu et al.
2005; Ruiz-Hitzky et al. 2008). Important factors in design of nanostructured materials as
functional platforms include surface functionalization, assembly, orientation and
alignment (Kickelbick 2007). Diverse mechanisms have been used to assemble hybrid
nanostructures with specific applications. Natural hybrid nanostructures are usually
formed by biomineralization or self-assembly processes (Ariga et al. 2007) in which
useful techniques such as Langmuir-Blodgett (Acharaya et al. 2009), layer-by-layer
adsorption (Ariga et al. 2007) and sol-gel fabrication (Boettcher et al. 2007) are used.
Related synthetic, so-called biomimetic, approaches include crystallization on self-
assembled monolayers (Aksay et al. 1996; Bunker et al. 1994), supramolecular self-
assembly (Aksay et al. 1996) and sequential deposition (Keller et al. 1994).
1.3.2 Classification of Hybrid Structures
Hybrid materials are classified based on their composition, the type of
physiochemical interactions between their components, or the nature of chemical bonding
in their structure (Kickelbick 2007). Due to the importance of type of interaction between
the synthetic and the biological component in changing their characteristics, biohybrid
structures are usually classified based on this criterion. In the first class, one of the
biologic or synthetic components is entrapped within a network of the other component
(Figure 1.5) (Kickelbick 2007). In this class of hybrid structures, biologic and synthetic
components are bound together through mechanisms such as hydrogen bonding,
electrostatic interactions, ion-dipole coordination, proton and electron transfer processes,
and van der Waals forces (Ruiz-Hitzky et al. 2010 ; Sanchez et al. 2011). Soft chemistry-
17
based methods such as the sol-gel process are adaptable techniques in the first class
providing several possibilities for constructing biohybrid nanostructures with different
physico-chemical characteristics (Avnir et al. 2006; Barbadillo et al. 2011; Kim et al.
1998; Pandey and Mishra 2011; Sanchez et al. 2005). In the second class of hybrid
structures, biomolecules and a synthetic component are chemically bonded by a covalent
bond, providing platforms with enhanced and unique functional properties (Figure 1.5)
(Kickelbick 2007; Ruiz-Hitzky et al. 2010; Ruiz-Hitzky et al. 2011). Covalent
immobilization can enhance the stability of biohybrid nanostructures against different
solvents, high temperature and extreme pH conditions. Sometimes combination of two of
these mechanisms is used to provide the hybrid structure with the desired structural
properties (Judeinstein and Sanchez 1996; Kickelbick 2007; Ruiz-Hitzky et al. 2010).
Figure 1.5. Schematic of different techniques used for fabrication of hybrid structures.
1.4 Lipid bilayer-based Hybrid Nanostructures
Developing analytical devices for detecting and monitoring biological and
chemical analytes have taken a great interest from few years ago. For example,
18
fabricating sensitive and selective biosensors for high-throughput screening of bio-
compounds has an important role in modern medical care (Bally et al. 2010). As
mentioned before, biological molecules need a biomimetic environment to keep their
natural structure and functionality. Among bioinspired hybrid nanostructures,
phospholipid bilayers are one of the best options that can be used along with synthetic
materials to produce hybrid systems for studying of many biological processes in cell
(Figure 1.6). Assembly of lipid bilayers on nano-scale solid substrates results in
formation of versatile hybrid structures that allow investigation of biological phenomena
at a subcellular level (Lee et al. 2013). Simplified models of the cell membrane are
prepared through the self-assembly of phospholipid molecules in either form of vesicles
or lipid films. These biomimetic lipid membranes control all the communications of the
cell with external environment and provide an appropriate environment for understanding
biological function. The synthesis of lipid bilayer-based hybrid nanostructures allows the
fabrication of biomimetic lipid membrane on the surface of nanomaterials that can keep
biological activity, molecular structure and orientation of biomolecules (Lee et al. 2013).
19
Figure 1.6. Lipid-based hybrid nanostructures composed of synthetic metarials, such as hydrogel and
carbonnanotubes and lipid bilayers. These biomimetic structures accomodate bilogical molecules.
There have been many studies reporting the function and application of lipid
bilayers in biosensors (Bally et al. 2010; Jimenez et al. 2006; Tien 1990; Ye et al. 2003).
Two different areas for sensing applications based on modified or unmodified lipid
bilayers have been reported. In modified lipid bilayers, enzymes, antibodies, receptor
proteins, or DNA probes are immobilized in the bilayer, while in the unmodified form the
sensing ability just originates from the interaction between an analyte and the lipid
bilayer (Bally et al. 2010; Chalkias and Giannelis 2007). Biomimetic lipid bilayers
produce other biohybrid platforms by placing on metal surfaces (solid-supported)
(Rossetti et al. 2006; Tamm and McConnell 1985), polymer matrices (gel-supported)
(Spinke et al. 1992) and porous membranes (filter supported) (Schmitt et al. 2009). The
self-assembly of amphiphilic lipid molecules on surface of carbon nanotubes (CNTs) for
biosensing applications has been also previously reported (Artyukhin et al. 2005).
20
Due to important role of artificial lipid bilayers in producing simple and
biomimetic structures with different size, shape and compositions, we have used them in
combination with synthetic structures to fabricate new biomimetic hybrid nanostructures
that have various applications in biosensing, drug and gene delivery, and biophysical
studies of cell membrane. The developed lipid-based hybrid nanostructures exhibit new
structures and properties that cannot be obtained with each individual component. We
have developed two hybrid nanostructures using multi-walled carbon nanotubes
(MWCNTs) and poly(ethylene) glycol (PEG) hydrogel. These structures produce
artificial cell membrane environments on the surface of hybrid materials that allow us to
exploit them as novel biomimetic structures with unique properties. In the following
sections, we will describe characteristics and properties of hybrid materials we used for
fabrication of lipid-hybrid nanostructures. Then, in chpater 2 and 3 we will discuss about
new hybrid nanostructures we have constructed using MWCNTs, PEG hydrogel and lipid
bilayers.
1.5 Poly(ethylene) glycol
Poly(ethylene) glycol (PEG) is a linear or branched polymer composed of ethylene
oxide molecules. Each molecule of PEG contains hydroxyl group which makes it reactive
to form other polymer structures and enables it for chemical modification and covalent
attachment to other surfaces and molecules (Figure 1.7) (Harris 1992).
Figure 1.7. Structure of poly(ethylene glycol).
21
This neutral polymer with various molecular weights is soluble in both aqueous
solutions and most of organic solvents. This solubility property of PEG makes it as a
good choice for synthesis of PEG derivatives that need different environment for
processing the reaction (Harris 1992). PEG has wide range of biomedical and
biotechnical applications. Due to nontoxicity and biocompatibility of PEG, approved by
FDA for internal use (Herold et al. 1989), it has many applications in drug delivery,
making cosmetic products, personal care products, and PEG-proeins (Harris 1992). In
addition, poor immunogenicity of PEG allows development of PEG-proteins as drugs
(Fuertges and Abuchowski 1990). Studies have shown that incorporation of lipid-PEG in
liposome's bilayer can increase the serum stability of liposomes and protect them from
clearance via immune system (Senior et al. 1991). The stealth properties of PEG lead to
not only a decrease in release rate of components, but also a reduction in uptake of
harmful immunoglobins. On the other hand, PEG hydrogels are usually used as systems
for controlled drug delivery (sustained drug release) which result in achievement of
constant plasma level of the therapeutic agent from delivery system. PEG hydrogels can
be fabricated by different techniques. In a common approach called free radical
polymerization, hydroxyl group of PEG can be acrylated to form PEG-acrylate molecules.
Acrylate monomers are esters containing vinyl group, double bonded two carbon atoms,
attached to the carbonyl group. These acrylate-functionalized monomers are crosslinked
using free radical generating initiators. Free radicals are created from initiator molecules
upon exposure of them to UV radiation and heat, or through chemical reactions, such as
reduction. The initiating free radicals transfer to monomer molecules and grow them to
the longer chains.
22
1.5.1 Poly(ethylene) glycol Free Radical Polymerization Mechanism
As mentioned in the previous section, free radical polymerization of acrylate-
modified PEG molecules can be achieved by UV radiation or chemical reaction. Here we
will discuss the detail of chemical polymerization mechanism which is focus of our
research for making PEG hydrogel in liposome core. Chemical free radical
polymerization of PEG-diacrylate (PEG-DA) molecules is initiated by ammonium
persulfate (APS) as an initiator and TEMED (tetramethylethylenediamine). TEMED
behaves as catalyst and accelerate the rate of formation of free radicals from ammonium
persulfate. The persulfate free radicals attack double bonded carbon atoms of acrylate
groups and convert PEG-DA monomers to free radicals which later react with other
monomers to start polymerization chain reaction (Figure 1.8).
Figure 1.8. Free radical polimerization mechanism of PEG-DA using APS and TEMED. In a initiation step,
free radicals of persulfate are formed by APS as an initiator and TEMED as an accelerator. Persulfate free
radicals convert PEG monomer to free radical which initiates the polymerization chain reaction.
23
1.6 Carbon Nanotubes
Carbon nanotubes, discovered in 1991 by a group at the Naval Research
Laboratory, consist of one or more rolled up sheets of carbon with hexagonal network
(Iijima 2002). Those are usually longer than micrometer and their diameters are in range
of 1-20 nm (Iijima 2002). The nanotubes have been categorized to two groups: single-
walled carbon naotubes (SWCNTs) and multi-walled carbon nanotubes (MWCNTs). The
first group consists of single layer graphene cylinder with diameter of 1-2 nm, while the
second group are made of cocenteric graphene cylinders placed around one central
hollow (Ajayan 1999). The diameter range of MWCNTs is from 2 to 25 nm and spacing
between layers is about 0.34-0.36 nm (Ajayan 1999).
Figure 1.9 Structure of single-walled carbon nanotubes (A) and multi-walled carbon nanotubes (B) (Iijima
2002; Reilly 2007).
The carbon nanotubes unique structure, dimension and topology make them
special material for various applications. The basic structure of carbon nanotubes is C-C
covalent bond which is the strongest structure in the nature (Ajayan 1999). Extremely
small size, high mechanical stability, conductivity and incredible electrical properties of
24
carbon nanotubes allow using them as potential nanostructures in different technologies.
Due to the above- mentioned properties of carbon nanotubes, they have been used as
quantum wires (Tans et al. 1998), nanoprobes (Dai et al. 1998), and energy producer
devices (Che et al. 1998), such as fuel cells. High surface specificity of carbon nanotubes
have motivated scientists to exploit them in electron transfer reactions (Ajayan 1999).
The high electron density of nanotubes originates from their aromatic structure and
makes them observable by transmission electron microscopy (Klumpp et al. 2006).
Moreover, nanotubes have been widely used for fabricating biocompatible
nanostructures. The combination of nanotubes with biological molecules, such as proteins,
nucleic acids and polysaccharides convert them to the biocompatible structures with
potential applications in drug delivery and biosensing (Klumpp et al. 2006). Carbon
nanotubes can also form biomimetic structures with lipid bilayers. However, insolubility
of carbon nanotubes in water and most of organic solvents is a noticeable problem for
their biological applications. Studies showed that functionalization of carbon nanotubes
with desired functional groups improve their solubility in both aqueous and organic
solutions. To date several noncovalent and covalent strategies, such as using surfactants,
polymers, biopolymers and lipids or modification of sidewall of nanotubes by hydrophilic
species have been introduced for nanotubes functionalization and solubilization.
Interfacing nanotubes with lipids not only enhance the solubility of them, but also give us
the opportunity to study many important biological processes via developing functional
hybrid devices.
25
Chapter 2: Lipid Bilayers Covalently Anchored to Carbon Nanotubes
(This work has been published in Langmuir, 2012, 28 (21), pp 8174–8182,
DOI: 10.1021/la301094h)
2.1 Introduction
Carbon nanotubes (CNTs) have received significant attention because of their
unique structural, physical and electrical properties (Artyukhin et al. 2004; Baughman et
al. 2002; Dresselhaus et al. 2001; Richard et al. 2003). They have a wide range of
applications in biosensing; gene and drug delivery in cells; intracellular transport of
oligonucleotides, proteins and peptides; biophysical studies; and nanodevices (Gruner
2006; Heller et al. 2008; Kam and Dai 2005; Kam et al. 2004; Kong et al. 2000; Lei and
Ju 2010; Polizu et al. 2006; Shim et al. 2002; Wang 2005). However, the hydrophobic
surface of CNTs and their resulting insolubility in water limits their potential
biomolecular applications. Carbon nanotubes have a tendency to aggregate in most
solvents, leading to difficult-to-disperse bundles. Hydrophobic CNTs do not provide an
appropriate environment for integration of biomolecules such as proteins, and they are
highly resistant to any coupling with biomolecules (Artyukhin et al. 2005; Huang et al.
2010). Here, we address these issues by fabricating a novel lipid-nanotube bioconjugate
system in which lipid bilayers are covalently anchored to the surfaces of multi-walled
carbon nanotubes (MWCNTs) to support the insertion of membrane proteins. This
approach will not only facilitate the integration of CNTs with other biomolecules, but
also establish a biocompatible platform for molecular devices and nanoscale architectures.
Several approaches have been developed for functionalization and solubilization of
carbon nanotubes, including covalent and non-covalent surface modifications. Non-
26
covalent functionalization has been based on surface adsorption of surfactants, polymers
and biomolecules to CNTs and formation of CNT adduct nanostructures, such as CNT-
crown ethers, using attractive electrostatic forces (Chen et al. 2002; Islam et al. 2003;
Kahn et al. 2002; Moore et al. 2003; Tu et al. 2009; Yurekli et al. 2004; Zhang et al.
2010). Covalent modifications involve direct reaction of CNTs to decorate their side
walls with a desired functional group (Baker et al. 2002; Bianco et al. 2005; Hazani et al.
2003; Huang et al. 2002b; Pham et al. 2010; Singh et al. 2006; Tasis et al. 2003; Zhu et al.
2003). Many of the functionalization reactions are based on amiditaion and esterification
of carboxylic acid-activated CNTs (Huang et al. 2002). Although many covalent
modifications allow for attachment of biomolecules to 1D nanomaterials, they offer a
limited capacity to orient biomolecules and provide a biomimetic environment that
facilitates biomolecular function (Noy et al. 2009). Lipid bilayers attached to CNT
surfaces can serve as a biomimetic environment for incorporating functional membrane
proteins and other biomolecules.
To date, noncovalent modification of CNTs with single-tailed, double-tailed,
dextran-containing phospholipids and polymerized lipids has been reported (Contal et
al. ; Douroumis et al. 2007; Goodwin et al. 2009; Lin et al. 2006; Richard et al. 2003; Wu
et al. 2006). Due to the amphiphilic nature of lipid molecules, they can self-assemble on
the hydrophobic surface of CNTs in aqueous solution and provide a soluble
supermolecular structure with wide range of applications (Contal et al. 2010; Douroumis
et al. 2007; Goodwin et al. 2009; Lin et al. 2006). Although adsorption of lipids to CNTs
leads to soluble, stable aqueous suspensions, the structures thereby formed lack the
27
biomimetic structure of a true lipid bilayer. Direct self-assembly of lipid molecules on
hydrophobic nanotubes instead forms a lipid monolayer (Richard et al. 2003).
Several routes to the surface modification of CNTs with lipid bilayers have been
proposed. Artyukhin et al. modified single-walled carbon nanotubes (SWCNT) with
polyelectrolyte multilayers to make a hydrophilic polymer surface for lipid self-assembly
(Artyukhin et al. 2004; Artyukhin et al. 2005). This modified SWCNT was incubated
with lipid vesicles containing anionic and zwitterionic lipids leading to spontaneous
bilayer formation by vesicle fusion (Artyukhin et al. 2005; Zhang et al. 2000). In other
approaches lipid bilayers were formed on the hydrophilized surface of SWCNTs through
fusion of small or giant vesicles (Gagner et al. 2006; Huang et al. 2010a; Huang et al.
2010b; Zhou et al. 2007). Self-assembly of bilayers on the hydrophilic surface of
MWCNTs was also reported by Ye et al. (Ye et al. 2005). There have also been studies
attempting covalent lipid modification of both SWCNTs and MWCNTs resulting in
improved aqueous dispersion (He and Urban 2005; He and Zhu 2008); however, these
reports do not describe the formation of well characterized bilayers.
The covalent modification of CNTs provides an advantage over other methods in
that it allows for durable attachment with higher stability (Baker et al. 2002; Gao and
Kyratzis 2008). Such structures should be robust to changes in environmental conditions
such as pH and temperature.
In this study, we introduce a novel approach for lipid bilayer formation on
MWCNTs. MWCNTs have outer diameters in the range of 20-30 nm, providing a
sufficiently large radius of curvature to allow for assembly of true molecular bilayers
(Artyukhin et al. 2005b; Huang and Mason 1978). Bilayer-coated MWCNTs were
28
fabricated in a two-step process. First, lipid head groups were attached to the nanotube
surfaces in a carbodiimide-mediated reaction. This creates a covalently modified
nanotube surface that allows for the self-assembly of second lipid layer in a simple
sonication process. The bilayer-coated MWCNT suspension is well dispersed and is
stable for at least 60 days. The biomimetic cell membrane-like structure of the bilayer on
the nanotube surface is demonstrated by the stable insertion of the bacterial membrane
protein α-hemolysin (Bakas et al. 1996; Schindel et al. 2001; Valeva et al. 1996),
fluorescence quenching of dye-labeled lipid, and fluorescence anisotropy measurements
of bilayer fluidity. These biocompatible carbon nanotubes will allow for the creation of
novel nanoscale structures for biosensing and bioelectronics.
2.2 Materials and Methods
2.2.1 Materials
COOH-functionalized multi-walled carbon nanotubes were obtained from
Nanostructured and Amorphous Materials (Houston, TX, USA). 1,2-dipalmitoyl-sn-
glycero-3-phosphoethanolamine (DPPE), 1-palmitoyl-2- 6-[(7-nitro-2-1,3-
benzoxadiazol-4-yl)amino]hexanoyl -sn-glycero-3-phosphocholine (NBD-PC), 1,2-
dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-(7-nitro-2-1,3-benzoxadiazol-4-yl)
(NBD-PE), 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphocholine (lyso-PC) and 1,2-
dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) were purchased from Avanti Polar
Lipids (Alabaster, AL, USA). N, N′-diisopropylcarbodiimide (DIC), N-
Hydroxysuccinimide (NHS), 5(6)-carboxytetramethylrhodamine N-succinimidyl ester
(NHS-rhodamine), Triton X-100, sodium dodecyl sulfate (SDS), sodium hydrosulfite
29
(sodium dithionite) and all solvents were from Sigma-Aldrich (St. Louis, MO, USA).
Sodium bicarbonate and sodium chloride were obtained from J.T. Baker (Phillipsburg, NJ,
USA). Sodium phosphate was from EMD chemicals (Gibbstown, NJ, USA).
2.2.2 Synthesis of Lipid-modified Multi-walled Carbon Nanotubes
For covalent conjugation of carbon nanotubes to DPPE, 1 mg of COOH-
functionalized MWCNTs was suspended in 2 mL of a 2:1:1 (volume ratio) mixture of
DMF, chloroform, and methanol and sonicated for 1 h. 10 mg DPPE along with a 10×
mixture, sonicated for 1 h and stirred for 48 h at 50°C. The reaction mixture was
centrifuged for 20 min at 4000 rpm to separate MWCNTs and remove unbound lipid
molecules. The resulting supernatant was discarded and modified MWCNTs were
washed with 1:1 (volume ratio) mixture of chloroform and methanol 3 times to remove
free lipids. For complete removal of physisorbed lipids from the MWCNT surface, a 2%
solution of triton X-100 in water was added to the MWCNTs, mixed thoroughly, and
centrifuged for 15 min at 4000 rpm to pellet the MWCNTs. To remove any trace of
detergent, MWCNTs obtained from this process were washed with pure water and
centrifuged to pellet (4000 rpm, 15 min) 3 times. Then, a mixture of 1:1 chloroform and
methanol was added to MWCNTs, mixed well and centrifuged for 2 more cycles. Lipid
modified-MWCNTs were dried in vacuum overnight at 30 °C. As a control for
physisorption, 1 mg COOH-functionalized MWCNTs were mixed with 10 mg DPPE in
the absence of DIC and NHS. Otherwise all reaction and washing conditions were as
applied in the synthesis of lipid-modified MWCNTs.
30
2.2.3 FTIR Analysis
The covalent conjugation of DPPE lipid molecules with MWCNTs was
confirmed using a Vertex 80 FTIR spectrometer (Bruker, Germany). Infrared spectra
were recorded at 2 cm
-1
resolution at room temperature over a wavenumber range of
1000-4000 cm
-1
. A ZnSe window was used for recording of IR spectra of all samples. To
obtain the IR absorption of samples, COOH-MWCNTs were dispersed in dimethyl
sulfoxide (DMSO), and DPPE and MWCNT-lipid conjugate solutions were prepared in
chloroform. Samples were observed directly in the ZnSe liquid cell. Spectra of neat
solvents were background-subtracted from sample spectra.
2.2.4 Liquid-Liquid Extraction
To confirm the successful conjugation of DPPE with MWCNTs in the
carbodiimide-mediated reaction, lipid-modified MWCNTs were dispersed in a vessel
containing 2 mL of chloroform and 2 mL of pure water, shaken very well for 10 min and
left for an hour to partition into either the chloroform or the water phase.
2.2.5 Lipid Bilayer Fabrication
To form a lipid bilayer on lipid-modified MWCNTs, a solution of 2 mg DPPC in
chloroform was mixed with 50 μL DPPE-MWCNT dispersion as prepared in chloroform
at total volume of 1 mL. 1 wt% of either tail-labeled NBD-PC or headgroup-labeled
NBD-PE was mixed with the DPPC as a fluorescence probe. Chloroform was evaporated
under an argon flow and the resulting MWCNT-lipid film was dried in vacuum at room
temperature for 3 h to remove any trace of chloroform. 1 mL of 10 mM phosphate buffer,
31
pH 7.5, containing 150 mM NaCl was added to the MWCNT-lipid film and sonicated for
30 min in a bath sonicator. To remove excess lipid from the sonicated suspension of
lipid-coated MWCNTs, it was centrifuged for 10 min at 3500 g, the supernatant was
discarded, and the nanotubes were resuspended in fresh buffer. Nanotubes were observed
using fluorescence confocal microscopy.
2.2.6 Protein Incorporation in Lipid Bilayer-coated Nanotubes
To observe insertion of α-hemolysin into the lipid bilayer with fluorescence
confocal microscopy, α-hemolysin from Staphylococcus aureus (Sigma) was labeled with
NHS-rhodamine. 250 µL of 0.5 mg/mL protein solution in 50 mM sodium bicarbonate
buffer, pH 8.5, was incubated with a 5× molar excess of NHS-rhodamine for 2 h at room
temperature. To remove excess dye, the reaction mixture was centrifuged using Zeba spin
desalting column with 7K MWCO (Thermo Scientific, Pierce). 1 μM labeled α-
hemolysin solution was then added to the lipid bilayer-coated MWCNT suspension and
incubated at 37°C for 1 h. Unbound α-hemolysin molecules were removed by
centrifugation of the suspension for 10 min at 3500 g and resuspension of the resulting
nanotube-containing pellet. As control experiments, 1 μM labeled α-hemolysin was
incubated with a MWCNT-COOH suspension in PBS buffer or a MWCNT-COOH
suspension in PBS buffer with 2.5 mg/mL lyso-PC and mixed well for 1h at 37°C. Free
α-hemolysin molecules were removed by centrifugation as described above.
2.2.7 Sample Observation with Fluorescence Confocal Microscopy
To observe MWCNTs, we used a Nikon TI-E inverted microscope (Tokyo, Japan)
equipped with a Yokagawa CSUX confocal head (Tokyo, Japan). All images were taken
32
using a 60× oil-immersion objective (Apo TIRF). NBD and rhodamine were excited with
50 mW solid-state lasers at 491 nm and 561 nm, respectively, and the emission signals
were recorded at 535 and 595 nm. The suspension of nanotubes was transferred into a
Sykes-Moore microscopy chamber (Bellco, Vineland, NJ) containing a #1 glass coverslip.
Prior to use, coverslips were first sonicated in 80°C Millipore water for 30 min and then
immersed in pure sulfuric acid for 2 h. After rinsing with water thoroughly, coverslips
were sonicated in water for 30 min and then sonicated in methanol for 15 min. Cleaned
coverslips were dried in a 60°C oven.
2.2.8 Fluorescence Quenching
To confirm the formation of true lipid bilayers on MWCNTs, the total fluorescence
of lipid bilayers labeled with 1 wt% of either tail-labeled NBD-PC or head-labeled NBD-
PE was measured using a spectrofluorophotometer (RF-5301 PC, Shimadzu, Japan). The
samples were excited at 460 nm, and emission signals were recorded at 530 nm. Then, 20
µL of 1 M sodium dithionite in 10 mM PBS buffer with pH 9, prepared fresh and purged
with argon, was added to each suspension. One minute after addition of sodium dithionite,
the fluorescence of each sample was measured and compared with the initial fluorescence
value. Additional aliquots of quencher were then added to the suspension to confirm that
no further decrease in fluorescence intensity could be observed, and then, SDS surfactant
was added to the suspension to lyse the membrane and expose the inner leaflet of the
bilayer to the quencher.
33
2.2.9 Transmission Electron Microscopy (TEM)
Thirty microliters of MWCNT suspension in water and 30 µL of lipid bilayer-
coated MWCNT suspension in 10 mM phosphate buffer, pH 7.5 were pipetted on
paraffin wax surfaces separately. Formvar coated-copper grids (TED Pella, Redding, CA)
were placed on the sample droplets for 3-4 minutes and the excess fluid was removed by
filter paper. TEM images were taken with a JEOL JEM-2100 LaB6 microscope operating
at 200 KV.
2.2.10 Fluorescence Anisotropy Measurements
To verify the fluidity of lipid bilayers bound to the surface of MWCNTs, the
fluorescence anisotropy of NBD-labeled DPPC included in the lipid bilayer was
measured as a function of temperature using a QuantaMaster QM-4SE
spectrofluorometer from PTI (Photon Technology International, Birmingham, NJ, USA).
A temperature range of 4-55°C was examined, allowing for the gel-to-liquid phase
transition of DPPC to be observed. To equilibrate the cuvette temperature after each
temperature change, it was incubated at the target temperature for 30 min, and a pair of
measurements was taken to confirm no changes in fluorescence anisotropy. Excitation
was at 460 nm and emission was measured at 530 nm.
The sample containing the fluorophore probe was excited with linearly polarized
light and the intensities of the vertical and horizontal components of the emitted light
were measured to calculate anisotropy (r) using the following equation:
r = (I
VV
-GI
VH
)/ (I
VV
+2GI
VH
)
34
where I
VV
and I
VH
are the emitted intensities polarized parallel and perpendicular to the
excitation light when the excitation light is polarized vertically. G is a correction factor
defined as G = I
HV
/I
HH
where
I
HV
and I
HH
are vertical and horizontal fluorescence
emission intensities when the excitation light is polarized horizontally.
2.3 Results and Discussion
Due to the hydrophobic nature of carbon nanotubes, lipid molecules can self-
assemble on the nanotube surface via hydrophobic-hydrophobic interactions. This self-
assembly, however, does not lead to formation of a lipid bilayer. To fabricate a
membrane-like bilayer on a nanotube surface, we covalently conjugated 1, 2-dipalmitoyl-
sn-glycero-3-phosphoethanolamine (DPPE), a lipid bearing a primary amine on its head
group, with COOH-functionalized MWCNTs in a carbodiimide-mediated reaction using
N, N′-diisopropylcarbodiimide (DIC) (Figure 2.1). N-Hydroxysuccinimide (NHS) was
added to the reaction mixture to increase the stability of the active reaction intermediate
(Hermanson 1996). Since single-walled carbon nanotubes (SWCNTs), with outer
diameters of approximately 1-2 nm, are too highly curved to support the formation of
closed shell lipid bilayers (the smallest reported inner radius for lipid bilayers is about 5
nm) (Artyukhin et al. 2005; Huang and Mason 1978), MWCNTs with diameters in the
range of 20-30 nm were used. MWCNTs have some advantages for covalent
modification. They have higher mechanical stability than SWCNTs, and since
conjugation occurs only on the external walls of the multi-walled nanotubes, the
mechanical and electronic properties of the inner walls are unaffected by conjugation
(Lamprecht et al. 2009).
35
Figure 2.1. Synthesis of lipid-MWCNT conjugates in a carbodiimide-mediated reaction (not to scale).
2.3.1 Solubility of Lipid-modified MWCNTs
To confirm that covalent modification of water-dispersible COOH-functionalized
MWCNTs with DPPE converts them into a species that can be dispersed in nonpolar
solvents, liquid-liquid extraction of lipid-modified MWCNTs was performed between
chloroform and water. The excellent dispersibility of lipid-MWCNT conjugates in
chloroform is convincing evidence for the successful conjugation of lipid to MWCNT
surfaces (Figure 2.2). A control experiment in the absence of DIC and NHS resulted in
partitioning of the nanotubes to the water phase, confirming lack of any covalent bond
between lipids and MWCNTs. A comparison between extraction results for COOH-
MWCNTs as supplied by the vendor (Figure 2.2A), MWCNTs treated with lipids without
DIC (Figure 2.2B) and lipid-conjugated MWCNTs (Figure 2.2C) clearly establishes the
covalent conjugation of lipids to the nanotube surface. The nanotubes in Figures 2.2A
and 2.2B partition to the water phase due to hydrophilic nature of carboxylic acid group
36
on the oxidized MWCNTs. Covalently modified nanotubes were stable in chloroform
and no aggregation was observed even after 3 months.
Figure 2.2. Liquid-liquid extraction of MWCNTs in a CHCl
3
/H2O system. The upper phase is water and
the lower phase is chloroform. Images show partitioning of MWCNTs to either the chloroform or water
phase after shaking the mixtures for 10 min and separating for 1 h. (A) COOH-MWCNTs; (B) A negative
control consisting of physically mixed MWCNTs and lipid reagents with no DIC or NHS; (C)
Conjugated lipid-MWCNTs from the carbodiimide-mediated reaction. All unbound and physisorbed lipid
molecules have been removed by treatment with surfactant. It is clear that MWCNTs in the negative
control behave like COOH-functionalized MWCNTs and partition to the water phase, while covalently
modified lipid-MWCNTs partition to the chloroform phase.
Our results contrast those of He et al., who reported that lipid-modified CNTs are
dispersible in both chloroform and water (He and Urban 2005; He and Zhu 2008). We
observed similar behavior in CNTs immediately after covalent modification, but when
these CNTs were treated with surfactant to remove physisorbed lipid species, they
partitioned exclusively to the chloroform phase, as described above. These hydrophobic-
surface CNTs, modified with a single layer of lipids, serve as the basis of our bilayer
fabrication technique (see Figure 2.3).
37
Figure 2.3. Schematic drawing of the insertion of protein into the bilayer of a lipid-modified MWCNT
(Front view). (A) Covalent modification of a COOH-MWCNT after reaction with DPPE; (B) Lipid
bilayer formation during sonication of lipid-modified MWCNT with DPPC molecules in phosphate buffer
and (C) insertion of α-hemolysin in the lipid bilayer-coated MWCNT. Drawing not to scale.
2.3.2 FTIR Spectroscopy
In order to further confirm the reaction between MWCNTs and lipid molecules,
Fourier Transform Infrared (FTIR) spectra of a COOH-functionalized MWCNT
suspension, DPPE solution, and DPPE-MWCNT conjugate suspension were analyzed. As
can be seen in Figure 2.4, there is a sharp peak at 1662 cm
-1
in the COOH-functionalized
MWCNT IR spectrum (Figure 2.4A) corresponding to the carboxylate group (COO
-
) of
deprotonated COOH, as reported previously by Singh et al (Singh et al. 2006). The peaks
at 1989 and 2149 cm
-1
are indicative of aromatic combination bands of carbon nanotubes
(Meyers 2000). The strong peak at 3475 cm
-1
in this spectrum corresponds to the
hydroxyl (OH) stretch of COOH-MWCNTs. The DPPE spectrum (Figure 2.4B) shows
peaks at 2855 and 2927 cm
-1
for C-H stretch vibrations in the alkyl chain of saturated
lipid tails (Meyers 2000). Other peaks at 1737 and 1056 cm
-1
in this spectrum can be
attributed to the N-H bend vibration of the primary amine group of DPPE and C-N
stretch vibration of the same group in the DPPE, respectively. In the lipid-MWCNT
conjugate spectrum (Figure 2.4C), the two new peaks at 3008 and 1653 cm
-1
correspond
38
to N-H and C=O vibration of an amide bond. This clearly confirms the formation of
covalent amide bonds between DPPE and COOH-functionalized MWCNTs. These two
peaks are only seen in the conjugate spectrum.
Figure 2.4. FTIR spectra of (A) COOH-
MWCNTs; (B) DPPE; (C) MWCNT-
DPPE conjugate. Appearance of NH
stretch (3008 cm
-1
) and C=O stretch
(1653 cm
-1
) peaks in figure C correspond
to amide bond formation, and the
disappearance of NH
3
+
and OH peaks in
this figure confirms the conversion of
primary amine and carboxylic acid to the
amide linking the lipid to the MWCNT
surface.
This result for amide peak frequency is in good agreement with result of Singh et
al. for formation of an amide bond between a peptide nucleic acid and SWCNTs (Singh
et al. 2006). Disappearance of the primary amine peak and hydroxyl peak in the
conjugate spectrum is verifying evidence for conversion of the lipid primary amine and
carboxylic acid to an amide. Appearance of a peak at 1262 cm
-1
in the lipid-MWCNT
39
conjugate spectrum also corresponds to the C-N stretch vibration of secondary amine of
amide bond, and the peaks in the range of 2800-2960 cm
-1
are indication of the C-H
stretch vibration of the conjugated lipid alkyl chain. The peaks at 3604 and 3693 cm
-1
in
the spectra of both the lipid and lipid-MWCNT conjugate are due to the aqueous nature
of sample and existence of non-bonded hydroxyl groups.
2.3.3 Lipid Bilayer Formation on MWCNTs
Formation of lipid bilayers on MWCNTs was accomplished by sonication of the
zwitterionic DPPC lipid with DPPE-modified MWCNTs in 10 mM phosphate buffer
containing 150 mM NaCl (pH 7.5). Sonication of lipid-MWCNT conjugates with free
lipid molecules is a novel method for the formation of bilayer-coated nanotubes. To
visualize the structure of lipid-coated MWCNTs, fluorescently labeled lipid (NBD-PC or
NBD-PE) was included in the lipid mixture. Figure 2.5 shows that the sonicated
suspension of MWCNT-lipid conjugates has good dispersibility in buffer solution after
the removal of excess lipid, and demonstrates that no nanotube sediment can be observed
even after two months.
Figure 2.5. Photographs of sonicated lipid bilayer-coated MWCNTs in buffer after (A) 1 day, and (B) 60
days. Lipid bilayer-coated MWCNTs are well dispersed in buffer.
40
Dispersibility of nanotubes in buffer solution is a strong evidence for fabrication of
lipid bilayers on MWCNT surfaces, because liposome-like lipid bilayers are stably
suspended in aqueous solutions (Basu and Basu 2002; Weissig and Torchilin 2003).
Similar behavior has been observed for self-assembled photopolymerized lipid micelles
in buffer solution (Contal et al. 2010).
Fluorescence confocal images show linear fluorescent features in buffer solution
corresponding to the lipid bilayer coated-MWCNTs (Figure 2.6). Since the diameter of
MWCNTs used in this study was in the range of 20-30 nm, this result is in good
agreement with report of Roiter et al. in which lipid bilayers can easily envelop
nanoparticles diameters greater than 22 nm (Roiter et al. 2008). Moreover, the cylindrical
geometry of double-chained phospholipids like DPPC means they easily form lamellar
bilayers (Hamai et al. 2006; Wu et al. 2006) and adapt to the curvature of 1D nanotubes
(Noy et al. 2009).
Figure 2.6. Confocal images of MWCNTs coated with NBD-DPPC-containing lipid bilayers in buffer
solution. Arrows indicate linear nanotubes. Scale bars: 10 µm.
41
The structure of COOH-functionalized and lipid bilayer-coated MWCNTs was also
investigated with transmission electron microscopy (TEM). As shown in Figure 2.7,
COOH-functionalized MWCNTs have a tubular shape, and they are entangled (Figure
2.7A), while the lipid-coated MWCNTs are more loosely associated and their length is
shorter compared to COOH-functionalized nanotubes (Figures 2.7B, C).
Figure 2.7. TEM images of (A) COOH-MWCNTs and (B and C) bilayer-coated MWCNTs after removal
of excess lipid. (D) Closer view of COOH-MWCNTs; (E) an enlarged image of part of the nanotube wall.
Lipid-coated MWCNTs are shorter than COOH-MWCNTs, and are less tangled. COOH-MWCNTs were
deposited from suspension in pure water and lipid-coated MWCNTs were deposited from suspension in
phosphate buffer solution. Dark areas in (B) and (C) are salt crystals formed after drying the buffer-
containing sample on formvar-coated grids. Scale bars: (A and C) 20 nm; (B) 50 nm; (D) 5 nm; and (E) is
an expanded view of the portion of D in the white box.
The walls of MWCNTs are clearly observed in Figures 2.7D and 2.7E. Although
there is no major difference in morphology between COOH-functionalized and lipid
bilayer-coated MWCNTs, the shorter length and improved dispersion confirms the
42
modification and coating of nanotube surface with a lipid bilayer. Shortening of
nanotubes in a carbodiimide-mediated reaction has been reported earlier (Huang et al.
2002).
To confirm the formation of true lipid bilayers rather than multilayers on
MWCNTs after sonication, two experiments performed. In one experiment, α-hemolysin
was inserted into lipid-coated MWCNTs; in another, fluorescence from one leaflet of the
lipid bilayer was selectively quenched.
2.3.4 Alpha-hemolysin Insertion in Lipid Bilayers on Nanotube Surfaces
The α-hemolysin protein inserts spontaneously into lipid bilayers, forming
heptameric pores through which ions and small molecules can pass (Aksimentiev and
Schulten 2005; Dinges et al. 2000; Song et al. 1996). Spontaneous insertion of α-
hemolysin into the lipid structures on the surface of our modified MWCNTs is evidence
that these lipid structures are true molecular bilayers. Alpha-hemolysin from
Staphylococcus aureus labeled with rhodamine was added to the lipid bilayer-coated
MWCNT solution and incubated for 1 h (Figure 2.3). The images obtained from
fluorescence confocal microscopy with excitation at 561 nm and emission at 595 nm
confirmed the insertion of rhodamine-labeled α-hemolysin in the lipid membrane (Figure
2.8). Overlaying images of rhodamine and NBD fluorescence emission clearly shows that
α-hemolysin is exactly co-localized with lipid-coated MWCNTs (Figure 2.8C).
In a control experiment, COOH-functionalized MWCNTs with no lipid on them
were incubated with labeled α-hemolysin. No α-hemolysin adsorbed to nanotube surfaces
(Figure 2.8D) in the absence of a lipid bilayer. In another control experiment, COOH-
43
functionalized MWCNTs were mixed with lysophosphatidylcholine (lyso-PC), a single-
chain lipid with a polar headgroup, in PBS buffer (pH 7.5), and then incubated with
labeled α-hemolysin. In this control, a small number of MWCNTs in suspension
colocalized with a low level of α-hemolysin fluorescence, demonstrating weak non-
specific adsorption (Figure 2.8E).
Figure 2.8. Confocal fluorescence images of (A) NBD-labeled lipids on MWCNTs coated with lipid
bilayers by sonication (ex: 491 nm, em: 535 nm); (B) spontaneous insertion of rhodamine-labeled α-
hemolysin after incubation of bilayer-coated MWCNTs with protein at 37ºC for 1 h (ex: 561 nm, em: 595
nm); (C) Overlay of figures (A) and (B), which shows colocalization of α-hemolysin with bilayer-coated
MWCNTs; (D) COOH-MWCNTs incubated with rhodamin-labeled α-hemolysin (negative control); and
(E) COOH-MWCNTs coated with single-chain lyso-PC lipid and incubated with rhodamine-labeled α-
hemolysin (negative control). No rhodamine fluorescence emission was observed on MWCNTs in (D). A
very low rhodamine fluorescence emission from a few MWCNTs in figure (E) is the result of nonspecific
adsorption of α-hemolysin to lipid monolayer-coated MWCNTs. Scale bars: 10 µm.
Lyso-PC molecules can self-assemble on the surface of carbon nanotubes as a
monolayer forming a micelle-like surface, but they cannot form membrane-like bilayers
44
because of their conical molecular shape and curvophilicity.(Richard et al. 2003; Wu et
al. 2006) These results show that α-hemolysin interacts only weakly with this lipid
monolayer. Together, these observations indicate that the lipids assembled on the surface
of MWCNT by covalent conjugaction and sonication form a molecular bilayer rather than
a monolayer or disorganized structure.
2.3.5 Lipid Bilayer Fluorescence Quenching
In a complementary experiment, we confirmed the formation of lipid bilayers on
MWCNTs by fluorescence quenching. In this experiment, sodium dithionite, which can
quench the NBD fluorescence by chemically reducing the dye, was added to a suspension
of lipid bilayer-coated MWCNTs in which 1 wt% of either the lipid tails or lipid
headgroups were labeled with NBD. As McIntyre and Sleight have shown previously, the
reaction of NBD with dithionite results in reduction of the nitro group to an amine
without any side reactions or ring opening (McIntyre and Sleight 1991). This reduction
reaction is irreversible and makes NBD nonfluorescent (Kamal et al. 2009). One minute
following quencher addition to the suspensions, a 55.42% (head-labeled) or 47.53% (tail-
labeled) decrease in total fluorescence intensity was observed (Figure 2.9). Dithionite is a
charged molecule (dithionite anion) and has very low permeability across lipid bilayers
(Moreno et al. 2006); therefore, it can only quench the fluorescence of lipid molecules
placed in the outer leaflet of the bilayer. No further decrease in fluorescence intensities
was observed upon addition of more quencher to the suspensions. Upon addition of 10
mM SDS to the suspension, the fluorescence signal immediately dropped to near zero due
to exposure of the inner leaflet of the bilayer to the quencher. This observation
45
demonstrates that the initial drop (about 50%) in fluorescence is due to the quenching of
the outer leaflet and that the lipid molecules on the MWCNTs take on a molecular bilayer
structure. Other reports have demonstrated this technique as a confirmation of the bilayer
nature of lipid membranes (Gruber and Schindler 1994; Hu et al. 2011; Kamal et al.
2009; Pautot et al. 2003).
Figure 2.9. Normalized fluorescence intensity of bilayer-coated MWCNTs before and after the addition of
sodium dithionite quencher. The total fluorescence intensity dropped 55.42% and 47.53% for head-labeled
NBD-PE (dark gray) and tail-labeled NBD-PC (light gray), respectively, after addition of quencher.
Sodium dithionite only quenches lipid molecules in the outer leaflet of bilayer. Addition of 10 mM SDS
surfactant reduced the fluorescence intensities to about zero. These data demonstrate that only half of the
lipid population is in the outer leaflet of the bilayer, as expected for a molecular bilayer of lipids.
2.3.6 Lipid Bilayer Fluidity
Since lipid bilayer fluidity has an important influence on membrane functionality,
we confirmed fluidity of our MWCNT-anchored membranes by measuring fluorescence
anisotropy (r) of NBD-PC in these membranes as a function of temperature. Fluorescence
anisotropy has long been used as a measure of membrane fluidity; as molecular motions
46
are constrained in low-fluidity conditions (e.g. gel phase), fluorescence anisotropy
increases (Lentz et al. 1976; Marczak 2009; Mesquita et al. 2000). A true lipid bilayer
would be expected to demonstrate a significant change in fluidity at the lipid gel-to-liquid
phase transition. Therefore, we measured fluorescence anisotropy from temperatures
across the phase transition temperature of DPPC (~41 °C) (Lentz et al. 1976). Figure 2.10
shows the effect of temperature on the fluorescence anisotropy of bilayer-coated
MWCNTs. As expected, anisotropy values decrease from 4 °C to 55 °C as lipid rotational
motion increases. A drastic change in fluorescence anisotropy can be observed near the
expected phase transition temperature of DPPC. This change verifies the fluidity of the
bilayers near physiological temperatures and the mobility of lipids within the bilayers.
Figure 2.10. Fluorescence anisotropy of lipid bilayers conjugated to the surface of MWCNTs as a function
of temperature. Decreasing the temperature from 55 to 4 °C results in an increase in fluorescence
anisotropy. The gel-to-liquid phase transition is clear at ~40 ºC.
47
2.4 Conclusions
The need for biocompatible nanotube-based materials, which have potential
applications in biosensing and in energy-producing devices, motivated us to make lipid
bilayer-coated MWCNTs. Bilayers are formed in a two-step process. First, DPPE is
covalently anchored to the surface of COOH-functionalized MWCNTs in a carbodiimide-
mediated reaction. These lipid-modified MWCNTs then serve as a substrate for lipid
bilayer assembly via co-sonication with DPPC. Modification of MWCNTs with a
covalently anchored bilayer of lipids was confirmed by liquid-liquid extraction, FTIR,
fluorescence microscopy, and differential fluorescence quenching. Lipid bilayer fluidity
was verified with fluorescence anisotropy measurements. We also demonstrated that the
membrane protein α-hemolysin can spontaneously insert into the bilayers on the
MWCNT surfaces. Lipid bilayers can provide a biomimetic environment for integrating
proteins with carbon nanotubes, leading to a broad range of analytical and bioelectronic
applications.
48
Chapter 3: Liposomes with Double-stranded DNA Anchoring the
Bilayer to a Hydrogel Core
(This work has been publishd in Biomarcomolecules, 2013, 14 (10), pp 3380–3385,
DOI: 10.1021/bm401155a)
3.1 Introduction
Liposomes are well-studied nanostructures with a variety of biomedical
applications, useful for drug encapsulation and molecular targeting (Cassidy and
Schatzlein 2004; Hamad and Moghimi 2008). In addition to their biomedical
applications, liposomes have long been used as model cell membranes to study such
biophysical properties and phenomena as membrane permeability (Nichols and Deamer
1980), mechanical stress relaxation (Bruckner et al. 2009), and fusion (Torres and Bong
2011). In real cell plasma membranes, covalent anchoring between the bilayer and the
underlying polymeric cytoskeleton plays an essential role in biophysical processes,
radically affecting lipid diffusion (Murase et al. 2004), phase structure (Head et al. 2014;
Lillemeier et al. 2006), and mechanics (Chernomordik and Kozlov 2008). Constructing
model liposome membranes that can recapitulate the effects of cytoskeletal anchoring
requires forming both a biomimetic “cytoskeleton” in the core of the liposome and a
mode of anchoring between the membrane and this cytoskeleton.
There have been several efforts to construct liposomes with hydrogel cores that
mimic the cytoskeleton. In early work, Torchilin et al. reported a free radical
polymerization method for making acrylamide gel in ~600 nm vesicles (Torchilin et al.
1987). In 1995, Monshipouri and Rudolph cross-linked alginate in large liposomes
(Monshipouri and Rudolph 1995). Anchoring of the bilayer to an underlying hydrogel in
liposomes was reported by Stauch and coworkers in 2002: they used a membrane-
49
inserted anchor monomer with a polymerizable head group (Stauch et al. 2002). Several
other approaches have been proposed for anchoring a liposome's lipid bilayer to an
interior hydrogel (Gutmayer et al. 2006; Kazakov et al. 2003; Kazakov et al. 2002).
Phospholipid bilayers have also been self-assembled on preformed lipid anchor-
containing hydrogel beads (Ng et al. 2004) and fabricated on hydrophobically modified
core-shell hydrogel spheres (Saleem et al. 2011). Immobilization of liposomes onto
hydrogel microbeads via avidin-biotin binding has also been previously reported
(MacKinnon et al. 2009).
Anchoring the membrane to a hydrogel can greatly improve membrane stability.
This is seen, for instance, in the radical increase of stability of hydrogel-anchored planar
bilayers (Malmstadt et al. 2008).
Other efforts to improve liposome stability have focused
on introducing synthetic components that allow for bilayer polymerization or provide a
steric buffer between the bilayer and its environment (Guo et al. 2009; Gutmayer et al.
2006; Papahadjopoulos et al. 1991). Generally methods for increasing liposome stability
include the use of photopolymerizable phospholipids (Fendler 1984; Guo et al. 2009;
Hub et al. 1980), making a polymer scaffold in the lipid bilayer (Roberts et al. 2009),
coating the liposome surface with various polymers or nanoparticles (Hayashi et al. 1999;
Pornpattananangkul et al. ; Ringsdorf et al. 1993; Ruysschaert et al. 2006; Yamazaki et
al. 1999), encapsulating polymer in the interior of liposomes (Hong et al. ; Schillemans et
al. 2006), and fabrication of hybrid phospholipid-block copolymer structures (Cheng et al
2011. ; Nam et al. 2011 ; Ruysschaert et al. 2005).
Here we present a liposome/nanogel structure in which anchoring of the bilayer
to the hydrogel core is accomplished by a double-stranded DNA linkage. These structures
50
present remarkably stable intact bilayers, and they can be used as biomimetic model
membranes that controllably recapitulate the connectivity between the cytoskeleton and
the plasma membrane. The connectivity is thermally reversible, as the bilayer-anchoring
cholesterol molecule is released from the cytoskeleton above the DNA melting
temperature. By using the bilayer as an organizing architectural principle, we are able to
construct nanogels in which the location of the oligonucleotide is well defined. After
stripping the lipid from the nanogels, we are left with polymer nanostructures decorated
specifically on their surfaces with oligonucleotides. These are potential sites for further
molecular assembly and can be used to facilitate directed assembly of nanogels, allowing
for mesoscale networks of networked nanoparticles (Park et al. 2008). These structures
also represent potential prototypes for oligonucleotide drug delivery vehicles, building on
significant previous work that has gone into designing polymeric and liposomal
particulate drug delivery systems (Kazakov et al. 2002; Park et al. 2008; Zhang et al.
2008).
3.2 Materials and Methods
3.2.1 Materials
1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), cholesterol, 1,2-dipalmitoyl-
sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) (Rhodamine-
DPPE, rh-DPPE), mini extruder and accessories were obtained from Avanti Polar Lipids.
Poly(ethylene glycol) diacrylate (PEG-DA) (M
n
575), Poly(ethylene glycol) methacrylate
(PEG-MA) (M
n
360), 3-[(3-Cholamidopropyl)dimethylammonio]-1-propanesulfonate
hydrate (CHAPS) and Triton X-100 (TX-100) surfactants, Sephacryl 200, ammonium
51
persulfate (APS), N,N,N′,N′-Tetramethylethylenediamine (TEMED) and all solvents were
obtained from Sigma-Aldrich. Sodium phosphate was obtained from EMD chemicals.
Sodium chloride was purchased from J. T. Baker. Cholesteryl-TEG-DNA (chol-DNA-1)
was from Trilink Biotechnologies. Acrydite-modified DNA (acry-DNA-2) and the
complement of acry-DNA-2 without cholesterol (DNA-1) were purchased from IDT.
DNA sequences are given below. Hoechst 33342 fluorescent dye (H33342), 8-
aminonaphthalene-1,3,6-trisulfonic acid, disodium salt (ANTS), and p-xylene-bis-
pyridinium bromide (DPX) were purchased from Invitrogen. Fetal bovine serum was
obtained from Gibco. Methacryloxyethyl thiocarbonyl rhodamine B (rhodamine-
methacrylate, rh-MA) was from Polysciences. RNase-free DNase Ι and
ethylenediaminetetraacetic acid (EDTA) were from Pierce (Thermo Scientific).
DNA Sequences
chol-DNA-1:
5’-Cholesteryl-tetra(ethylene glycol)- CTCCCTTCTCTCTCT-3’
acry-DNA-2: 5’-Acrydite- AGAGAGAGAAGGGAG-3’
DNA-1: 5’- CTCCCTTCTCTCTCT-3’
3.2.2 PEG Hydrogel Formation in Liposomes
To prepare hydrogel-containing liposomes, a solution of DOPC in chloroform (2
mg/mL) was prepared and 200 µL of this solution was evaporated under argon flow in a
glass test tube to form a dry lipid film. The film was kept under vacuum for at least 3 h to
remove any trace of chloroform. A mixture of 10% (v) PEG-DA, 5% (v) PEG-MA, 0.2%
(w/v) ammonium persulfate and 0.2% (v) TEMED in PBS containing 10 mM phosphate
52
and 150 mM NaCl at pH 7.5 was added to the lipid film and sonicated for 5-10 seconds.
The solution was placed in an ice bath to prevent polymerization prior to liposome
formation. Small unilamellar liposomes were formed by 50 × extrusion of the PEG-
containing hydrated lipid solution through a 0.08 µm polycarbonate filter in a cold room.
To prevent polymerization exterior to the liposomes, the suspension was diluted 5 ×
with a solution of 2% (v) TEMED in PBS. Since TEMED molecules are small and
sufficiently nonpolar to cross lipid bilayers, the presence of TEMED in the dilution buffer
is necessary to maintain the TEMED concentration in the liposome interior. The diluted
suspension was then incubated in a ~40°C water bath for 15 minutes to polymerize the
hydrogel. The hydrogel-liposome suspension was concentrated using an Amicon
ultracentrifuge tube with 10 kDa MWCO (EMD Millipore), and dialyzed against PBS
buffer using a dialysis cassette with a 20 kDa MWCO (Pierce) for 20 h to remove extra
monomer and initiator. To track hydrogel inside the liposomes, 24 µM rhodamine-
methacrylate (rh-MA), which can cross-link with PEG monomers during free radical
polymerization, was added to the monomer mixture prior to liposome formation.
3.2.3 Conjugation of Lipid Bilayer to PEG Hydrogel Using DNA Linkages
Lipid bilayers were anchored to the hydrogel surfaces by including cholesterol-
conjugated dsDNA in the lipid mixture during lipid film formation. Briefly, chol-DNA-1
was mixed with acry-DNA-2 at equimolar concentrations (20 µM) in a solution of 1:2
(v:v) acetonitrile : PBS. Oligonucleotides were annealed by incubating the solution at
90°C for 3-5 min and cooling it down to room temperature gradually. Annealed DNA
molecules were added to the solution of DOPC in 1:1 (v:v) chloroform/methanol. In
53
some experiments, a 0.01 μM Hoechst 33342 dye (H33342) was added to the hybridized
DNAs to detect liposomes with dsDNA anchors. The rest of procedure was as for
formation of hydrogel-containing liposomes. Based on the stoichiometry in the liposome
fabrication protocol, there were on average ~1100 cholesterol-conjugated dsDNA
molecules per liposome. This number was obtained by calculating the total number of
DOPC lipid molecules in a liposome considering average liposome hydrodynamic radius
(obtained by DLS) with 4 nm bilayer thickness (Banchelli et al. 2009). In these
calculations, we assumed an area per DOPC molecule of 0.7 nm
2
(Pan et al. 2008).
3.2.4 Size Exclusion Chromatography (SEC)
To confirm the formation of hydrogels in liposomes and show the presence of
dsDNA anchors, CHAPS solution in PBS was added to the liposome suspension to a final
concentration of 15 mM to strip off the lipid bilayer. The liposome suspension was then
pumped over a C-type SEC column (GE Healthcare) (packed with Sephacryl 200 (Sigma)
and prewashed with CHAPS solution) at a 0.2 mL/min flow rate to separate nanogels
from lipid or lipid/DNA micelles. A Shimadzu fluorescence detector (RF-535) integrated
with the SEC column was used to record fluorescence intensity (FI) chromatograms of
labeled nanogels and dsDNAs. All FI values were normalized to the maximum FI
recorded in each chromatography run. The Hoechst dye, complexed with dsDNA, was
excited at 350 nm and the emission intensity was recorded at 460 nm. The excitation and
emission for rh-MA were 548 and 570 nm, respectively. To remove surfactant from
hydrogel suspensions following SEC, hydrogels collected from the column were dialyzed
against PBS buffer with 150 mM NaCl for 3 days with threefold buffer exchange. For
54
control experiments, liposome species without PEG monomers and initiator were
prepared and examined in the presence of surfactant at the above-mentioned conditions.
3.2.5 Confirming DNA Location
A Shimadzu spectrofluorophotometer (RF-5301 PC) was used in H33342/DNA
hybridization experiments. Surfactant-treated liposomes with dsDNA anchors were
pumped over a surfactant-prewashed SEC column incubated either at room temperature
(RT) or at a temperature above dsDNA melting temperature (T
m
). Collected
dsDNA/ssDNA-conjugated nanogels were dialyzed for 3 days with threefold buffer
exchange to remove surfactant. 0.02 nM H33342 dye was added to the DNA-conjugated
nanogels and change in FI of dye was measured upon addition of increasing
concentrations of single-stranded, unmodified DNA-1. After addition of dye and DNA-1,
the DNA-liposome suspension was left for 10-15 minutes for dye equilibration. The dye
was excited at 350 nm and the emission was recorded at 400-600 nm. FI values are
reported at the emission maximum. The total number of DNA molecules anchored to a
nanogel was estimated by measuring the total concentration of rh-MA encapsulated in
liposomes by absorbance to estimate total volume of hydrogel. Together with the
liposome diameter (from DLS) this allowed for an estimate of the number of liposomes.
DNA concentration was then given by the saturation concentration in the experiment
described above.
3.2.6 Transmission Electron Microscopy (TEM)
The structure and morphology of hydrogel-containing liposomes and nanogels were
observed using a JEOL JEM-1400 microscope operating at 100 KV. An aqueous
55
suspension of nanoparticles was negatively stained with 1% (w/v) solution of uranyl
acetate (UA) before observation. Briefly, 12 µL of suspensions were absorbed onto the
formvar-coated copper grids, and then, extra solution was removed with filter paper.
To stain the nanoparticles, sample-loaded grids were placed on few microliters of
UA solution for a few seconds. Extra stain was removed, and the grids were air-dried.
3.2.7 Dynamic Light Scattering (DLS)
The hydrodynamic radius (R
H
) of hydrogel-liposomes and hydrogels was
determined using a DynaPro dynamic light scattering instrument (Wyatt technology)
using a 10 sec acquisition time for 10 acquisitions at 25°C. Nanoparticle suspensions
were filtered into a glass cuvette using a 0.45 μm filter. The results have been processed
with the DYNAMICS software program. The radii and size distribution has been
calculated with the using second order cumulant analysis. R
H
and polydispersity of all
species was reported based on weighted averages of several measurements for 3 distinct
samples. To measure the R
H
of dsDNA-conjugated nanogels, lipid bilayers were removed
by surfactant and hydrogels collected from SEC column were dialyzed in PBS buffer to
prevent interference of surfactant micelles in measurements.
3.2.8 Size Changes of Nanogels Exposed to Organic Solvent
To confirm the stability of fabricated nanogels in organic solvent, we studied size
changes after exposing them to chloroform. Briefly, nanogels suspended in PBS were
mixed with chloroform, shaken well and left for an hour. After separating the two phases,
56
rhodamine acrylate-labeled nanogels were collected in the PBS phase. Nanogels were
dried and redissolved in PBS again, and the size of nanoparticles was measured by DLS.
3.2.9 DNase Treatment of Cholesterol-dsDNA-anchored Liposomes
To determine whether any of the dsDNA anchor molecules were retained in the
outer leaflet of the liposomes, we treated the liposomes with DNase I. Liposomes were
incubated with DNase for 1h at 37 ºC in 10 mM Tris-HCl (pH 7.5), 0.1 mM CaCl
2,
and
10 mM MnCl
2
. The DNase was then deactivated by addition of EDTA to 5 mM and
incubation for 10 min at 65 ºC. The DNase-treated liposomes were then incubated with
0.1 µM H33342. Since this is a membrane-permeable dye, it would be expected to label
DNA molecules on both the interior and the exterior of the liposomes. Finally, the
liposomes were subjected to SEC analysis.
3.2.10 Release Study
To examine the stability of liposomes, 12.5 mM ANTS dye and 40 mM DPX
quencher were loaded in liposomes during their preparation. The maximum quenching
was observed at these concentrations of dye and quencher. Unencapsulated dye and
quencher were removed by passing the liposome suspension over a Zeba spin column
with 7kDa MWCO (Pierce). Then, liposomes were incubated in 20% fetal bovine serum
(FBS) at 37°C, and release of dye was measured by fluorescence spectrophotometry at
various times. The dye was excited at 360 nm and emission was recorded at 510 nm. The
release percentage was calculated based on the fluorescent intensity following lysis of
liposomes with 1% TX-100.
57
3.3 Results and Discussion
3.3.1 Fabrication of Hydrogel-anchored Liposomes with dsDNA Anchors
DNA-directed organization and manipulation of liposomes has recently been
implemented as a tool for controlling interactions between nanoscale lipid bilayer
structures (Banchelli et al. 2008; Chandra et al. 2006; Yoshina-Ishii et al. 2005). DNA
molecules can act also as controllable and programmable building blocks for the accurate
spatial arrangement of biomolecules such as proteins (Gartner and Bertozzi 2009). DNA-
lipid conjugates, such as cholesterol-tagged DNA, have been recently used for liposome
tethering and studies of interactions between individual lipid vesicles (Beales and
Vanderlick 2007; Chan et al. 2008; van Lengerich et al. 2010). In this study, cholesterol-
conjugated double-stranded DNA (dsDNA) is incorporated into the bilayer. The
complement of the cholesterol-conjugated strand is terminated with a vinyl group that is
incorporated in the hydrogel during free radical polymerization of PEG-diacrylate (PEG-
DA) macromonomers (Figure 3.1).
Figure 3.1. Schematic representation of a lipid bilayer anchored to a PEG hydrogel by a DNA linker.
Cholesterol-conjugated dsDNA is covalently anchored to the hydrogel during free radical polymerization.
58
The dsDNA anchor molecule was composed of two complementary single-stranded
DNA (ssDNA) molecules: Cholesterol-tetra(ethylene glycol)-modified ssDNA (chol-
DNA-1) and acrydite-modified ssDNA with a sequence complimentary to that of DNA-1
(acry-DNA-2) (each 15 bases long, sequences in materials and methods). To hybridize,
ssDNAs were dissolved in a 1:2 (v:v) mixture of acetonitrile and phosphate buffered
saline (PBS, 150 mM NaCl, 10 mM phosphate, pH 7.5). Oligonucleotides were annealed
by incubating an equimolar mixture at 90°C for 3-5 min and gradually cooling to room
temperature prior to addition to the lipid solution. Hydrogel-anchored liposomes were
prepared by dissolving cholesterol-conjugated dsDNA and dioleoylphosphatidylcholine
(DOPC) in a chloroform/methanol solution at molar ratio of 1:125. The mixture of
DOPC/dsDNA anchor was dried and the resulting lipid film was rehydrated in a hydrogel
precursor solution containing poly(ethylene) glycol diacrylate (PEG-DA, 10% (v)) and
poly(ethylene) glycol methacrylate (PEG-MA, 5% (v)) along with ammonium persulfate
(APS) initiator (0.2% (w/v)) and TEMED (0.2% (v)) in PBS. Rhodamine-methacrylate
(rh-MA) was included in the monomer mixture at 24 μM to provide a covalently
incorporated fluorescent tracker. The dried lipid film was rehydrated by sonicating (10 s)
at reduced temperature (4 ºC) to prevent polymerization.
Following lipid rehydration, the liposomes were extruded at 4 ºC through a
polycarbonate filter to produce a monodisperse sample. The liposome suspension was
diluted to prevent polymerization of PEG-DA and PEG-MA macromonomers outside of
the liposomes. Since these monomers are relatively large and hydrophilic, they are
trapped in the liposome interior. Their concentration inside the liposomes remains
constant while they are diluted to a non-polymerizing concentration outside of the
59
liposomes. The liposome suspension was brought to 40 ºC to trigger polymerization of
the acrylate and methacrylate groups inside the liposomes. This thermally controlled
method for forming hydrogels in liposomes has not been previously reported; in fact, we
know of only two previous reports of PEG-based hydrogels in liposome cores (An et al.
2009; Murphy et al. 2011). The vinyl group of the cholesterol-dsDNA anchor was
incorporated into the hydrogel during polymerization.
3.3.2 Characterization of Fabricated Nanoparticles
The nanoparticles were characterized using size exclusion chromatography
(SEC), transmission electron microscopy (TEM), and dynamic light scattering (DLS).
3.3.2.1 SEC and TEM
Figure 3.2A shows chromatograms of liposomes formed by the temperature-
controlled technique described above, but without the cholesterol-dsDNA anchors. These
chromatograms track the fluorescence of the rh-MA included in the liposome core. SEC
was performed with a Sephacryl 200 gel that allowed liposome-sized structures to elute
in the void volume while retaining smaller species. As can be seen in Figure 3.2A, intact
hydrogel-liposomes, liposomes without hydrogel, and hydrogel-liposomes treated with a
high concentration of CHAPS surfactant have almost the same retention time in the
column (and therefore a hydrodynamic radius (R
H
) larger than the fractionation range of
the column). On the other hand, the fluorescence from a surfactant-treated liposome
sample prepared without monomer or initiator has a much longer retention time due to
release of rh-MA from surfactant-lysed liposomes. These results indicate that hydrogels
60
are formed by the process described above and that even after stripping the lipid bilayer
with surfactant these hydrogels migrate as large particles, like the intact liposomes. TEM
images also show the presence of nanoscale hydrogels (~80 nm) associated both with
intact hydrogel liposomes and surfactant-treated nanogels. These images of partially
dried hydrogels do not show the associated lipid bilayer; rather, they establish the
successful fabrication of hydrogel cores (Figure 3.2B). The size of these nanoparticles is
in good agreement with pore size of polycarbonate filter used for extrusion of liposomes.
Figure 3.2. (A) Normalized SEC chromatograms tracking rh-MA in different liposome species. Intact and
surfactant-treated hydrogel liposomes elute in the void volume, showing shorter retention time than
surfactant-treated liposomes with no hydrogel, indicating that a surfactant-resistant hydrogel is formed. (B)
TEM images of negatively-stained hydrogel-liposomes (left) and bare hydrogels formed by stripping away
the bilayer with surfactant (right). Scale bars are 50 nm. (C) Normalized chromatograms tracking H33342
in various liposome species with dsDNA linkers. The retention time is the same for dsDNA anchor
molecules in intact liposomes and in surfactant-stripped hydrogels.
61
The insertion of the dsDNA anchor molecule in the lipid bilayer and conjugation
of the anchor to the hydrogel were confirmed by SEC of liposomes in which the dsDNA
anchor molecule had been dyed with a dsDNA-sensitive dye, Hoechst 33342 (H33342).
H33342 complexes with the minor groove of dsDNA, resulting in a significant increase
of the dye’s quantum yield (Wiederholt et al. 1996). As shown in Figure 3.2C, intact
anchored hydrogel-liposomes and surfactant-treated anchored hydrogel-liposomes in
which unanchored lipids were stripped off had the same retention time in SEC column,
indicating that the dsDNA is associated with the hydrogel. Control experiments with
surfactant-treated dsDNA-liposomes show the fluorescent species with a much higher
retention time, corresponding to the surfactant-solubilized DNA anchor molecule with a
size much smaller than that of the intact liposomes.
Association of the lipid bilayer with hydrogel in samples without surfactant
treatment was confirmed by tracking rhodamine-labeled lipids (rh-DPPE) included in the
lipid bilayer (Figure 3.3). Chromatography was performed both in the presence and
absence of surfactant (15 mM CHAPS). As shown in Figure 3.3, in the surfactant-free
case, labeled lipids elute in the void volume with the nanogels. When surfactant is added,
the lipids are retained on the column, likely in the form of mixed lipid-surfactant micelles.
Figure 3.3. Normalized SEC
chromatograms tracking rh-DPPE
fluorescence. Lipid molecules
colocalize with nanogels and elute in
the void volume unless they are
stripped off with surfactant.
62
3.3.2.2 DLS
The size of the dsDNA-conjugated nanogels that had been treated with surfactant
to strip the bilayer was compared to that of the intact dsDNA-conjugated hydrogel
liposomes by DLS. The measured R
H
of surfactant-stripped dsDNA-conjugated nanogels
(53±31 nm) was slightly smaller than that of the intact dsDNA-anchored hydrogel-
liposomes (64±10 nm), but the size distributions overlapped substantially. No significant
change in size of anchored hydrogel-liposomes was observed after 5 weeks. We also
exposed surfactant-stripped hydrogels to organic solvent to assure that the hydrogel
remained intact (Section 3.2.8). Following treatment with chloroform, the size of the
particles remained essentially unchanged. We measured particles with R
H
=57±35 nm.
This is within the sizes of liposome-nanogel particles we regularly observed.
3.3.3 Surface Accessibility of Cholesterol-dsDNAs
We also exposed the intact liposomes to DNase to evaluate whether there were any
surface-accessible dsDNA molecules present (Section 3.2.9). DNAse-treated liposomes
were analyzed by SEC. The resulting chromatogram, based on H33342 fluorescence, is
shown in Figure 3.4, along with a chromatogram of H33342 liposomes that have not been
treated with DNase. Our results indicate that no H33342-associated low-molecular-
weight species (i.e. free DNA) is generated upon DNase treatment, suggesting that all
dsDNA molecules are located on the interior of the liposomes. This is likely due to the
strongly polar nature of the DNA moiety leading to partitioning of surface-accessible
dsDNA anchor molecules into bulk solution.
63
Figure 3.4. Normalized chromatograms
tracking H33342 in dsDNA-anchored
liposomes with and without DNase
treatment.
3.3.4 dsDNAs Localization on Nanogel Surface
We used H33342 fluorescence to probe the position and accessibility of dsDNA
species on nanogels. The quantum yield of H33342 increases about 10 times when it
complexes with dsDNA (Cosa et al. 2001). We prepared surfactant-stripped nanogels
from dsDNA-anchored hydrogel liposomes. SEC was used to separate these nanogels
from the lipid/surfactant mixture that results from the surfactant treatment. SEC was
performed either at room temperature or above the melting temperature (T
m
) of the DNA-
1/DNA-2 duplex (65 ºC, compared to a theoretical melting temperature of 55 ºC). SEC
above the T
m
leads to DNA melting, separation of the chol-DNA-1 species from the
nanogels, and a nanogel product containing only single-stranded acry-DNA-2. We refer
to this product as an ssDNA-nanogel while the product containing the hybridized duplex
(prepared by SEC at room temperature following surfactant stripping) is referred to as a
dsDNA-nanogel.
To confirm that the DNA anchor molecules are at the surface of the nanogels and
solvent-accessible, we pre-incubated both nanogel species with 0.02 nM H33342 and
64
then added increasing concentrations of DNA-1. As shown in Figure 3.5, there was a
significant increase in H33342 fluorescence intensity (FI) upon addition of DNA-1 to the
ssDNA-nanogel species. This was accompanied by a blueshift of the H33342 emission
maximum wavelength (λ
max
), as is known to occur upon the dye’s binding to
dsDNA.(Cosa et al. 2001)
Above a concentration of ~ 9 µM DNA-1, no further changes
in FI or λ
max
were observed, indicating that all available single-stranded acry-DNA-2 was
hybridized. Similar shifts were not observed with the addition of DNA-1 to the ds-DNA
nanogel because the H33342 in these experiments was bound to the dsDNA on the
nanogel species; further hybridization could not be observed. These results indicate that
the DNA anchor molecules are localized to the nanogel-bilayer interface and that when
the bilayer is stripped off of the liposomes, DNA anchors are accessible and available for
further assembly processes via melting and hybridization. Based on the saturation
concentration of the complementary oligonucleotide strand, the measured absorbance of
nanogel-associated rh-MA, and the DLS-determined average nanogel diameter, we
estimate that ~20 DNA molecules are solvent-accessible on the surface of each nanogel
particle (Section 3.2.5).
Figure 3.5. Changes in
fluorescence intensity (ΔFI,
red symbols) and maximum
emission wavelength (Δλ, blue
symbols) of 0.02 nM H33342
dye in the presence of ssDNA-
nanogels (triangle) and
dsDNA-nanogels (circle) as a
complimentary DNA species
(DNA-1) is added to the
system.
65
3.3.5 Stability of Hydrogel-anchored Liposomes
The stability of hydrogel-anchored liposomes was examined with a fluorescence
dequenching experiment. A membrane impermeable dye (ANTS) and fluorescent
quencher (DPX) were co-encapsulated in liposomes prepared as described above. This
dye-quencher combination has been utilized widely for liposome leakage studies
(Pornpattananangkul et al. 2011); ANTS quantum yield increases as it leaks from
liposomes and the average dye-quencher distance increases. Liposomes were incubated
in 20% fetal bovine serum at 37°C (Figure 3.6).
Figure 3.6. Dye dequenching experiment that shows stability of hydrogel-anchored liposomes.
Figure 3.7A shows that hydrogel-anchored liposomes were stable up to 136 h in
serum, with ~22% release observed over 184 h. In contrast, unanchored liposomes
showed rapid release of dye upon incubation in serum, with 100% release achieved after
184 h. Liposomes containing hydrogels but without anchors showed significant release
after ~90 h and complete release after 184 h. These observations demonstrate the role of
anchoring in the successful formation of stable liposomes as a model for biological
membranes studies. To confirm that the stability we observed in our chol-DNA-1-
containing liposomes was a result of anchoring between the hydrogel and bilayer rather
66
than a result of increased stability due to the presence of cholesterol, we repeated the
release experiments with plain (unanchored, no hydrogel) liposomes containing the same
mole fraction of cholesterol as the anchored liposomes contained. Results shown in
Figure 3.7A and 3.7B confirms that cholesterol does not significantly stabilize the
liposomes compared to hydrogel anchoring.
Figure 3.7. (A) Dye release profiles from hydrogel-anchored liposomes, hydrogel liposomes without
anchors, and liposomes without hydrogel (plain liposomes) incubated in 20% serum. Release profiles
shows that hydrogel-anchored liposomes are significantly more stable than other types. The 100% point for
each experiment is based on fluorescence intensity after total liposome lysis with surfactant. Each release
experiment was performed three times; each data point shows the average over these three trials with error
bars showing one standard deviation. (B) Dye release profile of plain liposomes composed of same mole
fraction of cholesterol as that of dsDNA-anchored liposomes and incubated in 20% serum at 37°C.
In a complementary experiment, DLS showed a significant increase in the size
of
plain liposomes and hydrogel-liposomes incubated in 20% serum for 130 h (Figure 3.8).
No significant size change was observed for hydrogel-anchored liposomes. Covalent
anchoring likely increases the energy necessary to delaminate the bilayer from the
hydrogel surface, making it more difficult to form aggregated structures. Dye release data
67
(Figure 3.7A) and the time course of vesicle size as observed by DLS (Figure 3.8)
indicate that membrane instability leading to dye release is accompanied by vesicle
aggregation.
Figure 3.8. Liposomes size changes in 20% FBS. Plain liposomes, hydrogel-anchored liposomes, and
hydrogel-liposomes with no anchor were incubated in serum for 130 h. Changes in R
H
of different
liposomes were measured over 130 h.
3.3.6 Encapsulation Efficiency in Hydrogel-containing Liposomes
To demonstrate that the presence of the hydrogel did not impact the ability to
package a payload in the liposomes, we encapsulated a fluorescently labeled peptide
(sequence GTRQVTQASSFTWRVPG) (0.1 mg/mL) in core of liposomes both with and
without hydrogel precursors during the liposome hydration process. Free peptides were
removed by overnight dialysis in PBS and passing the liposomes over a Zeba spin
column (40kDa MWC). The concentration of encapsulated peptides was calculated by
measuring the absorbance of the dye in the same volume of liposomes. No significant
difference in encapsulation efficiency could be observed (10% and 8% encapsulation for
68
hydrogel-liposomes and plain liposomes, respectively). The hydrogel precursor solution
is only 15 wt% monomer, and all monomers have molecular weights less than 600 Da.
This solution would not be expected to exclude a relatively low-molecular-weight peptide
cargo from the vesicle interior during liposome formation.
3.3.7 Conclusions
In conclusion, we have fabricated stable nanogel liposomes in which dsDNA
anchor the bilayer to the underlying hydrogel. SEC and TEM demonstrated the formation
of hydrogels, and a dsDNA-binding dye confirmed the location of the DNA anchors at
the surface of the hydrogel. In this work, we have demonstrated temperature control of
the polymerization process to construct complex supermolecular polymer-based
structures without the use of photointiators. These structures incorporate a unique
acrylate-DNA-cholesterol macroamphiphile that both participates in lipid bilayer self
assembly and serves to stabilize lipid bilayers once formed. This allows for precise
localization of the DNA molecules within the nanostructure. This DNA-programmable
~100 nm, highly stable liposome structure, mimicking cytoskeletal connectivity to the
plasma membrane, has potential applications in biophysical studies of biomembranes.
DNA melting can be used to remove the anchoring, allowing for the effects of anchoring
to be specifically studied. These structures are potential prototypes for drug delivery
vehicles. In a drug delivery application, the eventual fate of the hydrogel core would be
important to the performance of the particles. This fate could be controlled by modifying
the core to be a degradable PEG hydrogel, subject to hydrolysis or degradation of cross-
linking disulfides in the cytosolic environment (Zustiak and Leach). With the bilayers
69
stripped off, the nanoparticles produced here have well-defined surface-localized DNA
hybridization sites that can be used for the directed assembly of additional DNA-
conjugated molecules or the assembly of the nanogels themselves into mesoscalar
structures.
70
Chapter 4: pH sensitive, hydrogel-anchord liposomes for peptide
delivery
4.1 Introduction
Liposomes are important vehicles for pharmaceutical delivery. To date, several
liposomal drugs, such as doxorubicin-loaded PEGylated liposomes (DOXIL) have been
clinically approved (Torchilin 2005). Clinical research has shown the effectiveness of
doxorubicin encapsulated in PEG-liposomes on hepatocellular carcinoma (Schmidinger
et al. 2001), cutaneous T-cell lymphoma (Wollina et al. 2003) and sarcoma (Skubitz
2003). Liposomes can be also used for delivery of DNA (Fraley et al. 1980) and nucleic
acid-based therapeutics, such as antisense oligonucleotides and siRNA (Straubinger and
Papahadjopoulos 1983). In recent years, biologically active compounds originating from
proteins and peptides have been broadly used as drugs for treatment of various diseases
(Torchilin 2005). There are reports for incorporation of insulin (Iwanaga et al. 1999),
interlukin-2 (Kanaoka et al. 2001) and recombinant human EGF (Li et al. 2003) in
liposomes coated with PEG. Due to unique structure of liposomes composed of
hydrophobic bilayer and hydrophilic core, they are potential vehicles for delivery of
different types of drugs. Liposomal drug delivery systems enable us to deliver higher
concentration of drugs while preventing distribution of them in non-targeted tissues
(Woodle and Storm 1998).
An important drawback of liposomes as drug carriers is their fast clearance from
the circulatory system and their capture by the reticulo-endothelial system cells
(Torchilin 2005). Therefore, for in vivo application, there is difficulty in retaining of
entrapped compound in core of liposome. As mentioned previously (chapter 3), several
71
attempts have been exploited to improve the therapeutic lifetime of liposomes. These
methods include the use of photopolymerizable phospholipids (Fendler 1984; Guo et al.
2009; Hub et al. 1980), making a polymer scaffold in the lipid bilayer (Roberts et al.
2009), coating the liposome surface with various polymers and nanoparticles (Hayashi et
al. 1999; Pornpattananangkul et al. 2011; Ringsdorf et al. 1993; Ruysschaert et al. 2006;
Yamazaki et al. 1999), and fabrication of hybrid phospholipid-block copolymer structure
(Cheng et al. 2011 ; Nam et al. 2011 ; Ruysschaert et al. 2005). Anchoring of lipid bilayer
to the polymer is another approach that increases the liposome stability significantly. In
chapter 3, we introduced a novel method for improving the sability of liposomes by
anchoring the lipid bilayer to the hydrogel core using dsDNA anchors.
Another concern in using liposomes as drug delivery vehicles is the delivery
mechanism of therapeutic compound across cell membrane. Receptor-mediated
endocytosis of ligand-targeted liposomes is one of the common ways of introducing
liposomes and their cargo into the cell (Allen and Cullis 2013). Antibodies are one of the
most widely used targeting moieties for liposomes (Torchilin 2005). They can be
attached to liposomes without changing the liposome or antibody properties (Torchilin
2005). However, few targeted liposomes have stepped forward into the clinical use. In
addition to the targeted delivery, triggered release is another approach for release of
therapeutic compound from liposomes (Allen and Cullis 2013). Remote triggers, such as
heat, ultrasound and light, and local triggers, such as enzymes and pH are two major
types of triggers have been known (Allen and Cullis 2013). pH-triggered release of
liposome content are achieved by fabrication of liposomes composed of pH-sensitive
components. These liposomes are stable at physiological pH (pH 7.5) but release their
72
cargo in the late-stage endosome with acidic pH after being endocytosed in the intact
form (Allen and Cullis 2013). pH-sensitive liposomes can be composed of either lipids
that confer pH-sensitivity to liposomes or pH-sensitive polymers (Torchilin 2005).
Among different classes of pH-sensitive liposomos, categorized based on mechanism of
triggering pH sensitivity (Drummond et al. 2000; Venugopalan et al. 2002), liposomes
composed of phosphatidylethanolamine (PE) or its derivatives and compounds containing
an acidic group, such as carboxylic group that act as a stabilizer at neutral pH are the
most common ones for drug delivery (Ellens et al. 1984). Due to the minimally hydrated
and small headgroup of PE compared to its hydrocarbon chains, it presents a cone shape
that hinders the formation of the lamellar phase like other types of lipids with cylindrical
shape (Cullis and Dekruijff 1979). Including amphiphilic molecules with a protonatable
acidic group into a PE lipid mixture results in electrostatic repulsion between molecules
and allows the formation of bilayer structure at physiological pH (Duzgunes et al. 1985).
Acidification triggers protonation of the carboxylic group and inverts PE molecules into
their inverted hexagonal phase leading to destabilization of liposome and therefore
release of its content (Lasic 1998). In addition, PE molecules are crucial components for
cell internalization of liposomes. Since PE molecules have low hydration on their
headgroups, they have high tendency to adhere to the cell membranes (Simoes et al.
2004). Following binding to cells, liposomes are internalized via the endocytotic pathway,
destabilized in late endosomes that have acidic pH, and finally release their content into
the cytoplasm (Simoes et al. 2004).
Here we have developed a stable, pH-sensitive liposome in which the lipid bilayer
is anchored to the hydrogel core by DNA linkages (chapter 3). The goal of this study is
73
intracellular delivery of therapeutic peptide using liposomes that have high stability in
serum. Plain peptides are suceptible to degradation in serum under physiological
conditions; therefore, they cannot have their desired effect on action sites. As shown in
chapter 3, our fabricated hydrogel-anchored liposomes have high stability in serum at
physiological pH. We show upon decreasing the pH to an acidic pH, liposomes start
releasing their encapsulated content gradually. We used these liposomes for
encapsulation and intracellular delivery of peptides that have anticancer properties. This
peptide has been discovered by Lee et al. in Dr. Jung's group, and the details of this study
were published elsewhere (Lee et al. 2011). We demonstrate the pH-sensitivity of
liposomes by dye dequenching. Size exclusion chromatography (SEC) confirms the
successful encapsulation of peptides in the hydrogel cores, and confocal microscopy
shows successful intracellular delivery of peptide-containing liposomes. These pH-
sensitive, stable nanostructures have potential applications in protected delivery of
therapeutic compounds, such as peptides and proteins, into the cells.
4.2 Materials and Methods
4.2.1 Materials
1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE), 1,2-dioleoyl-sn-glycero-
3-succinate (DOGS), 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1-palmitoyl-2-
[12-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]dodecanoyl]-sn-glycero-3-
phosphoethanolamine (NBD-PE), 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-
(lissamine rhodamine B sulfonyl) (ammonium salt) (Rhod-PE), mini extruder and
accessories were obtained from Avanti Polar Lipids. Poly(ethylene glycol) diacrylate
74
(PEG-DA) (M
n
575), Poly(ethylene glycol) methacrylate (PEG-MA) (M
n
360), 3-[(3-
Cholamidopropyl)dimethylammonio]-1-propanesulfonate hydrate (CHAPS) and Triton
X-100 (TX-100) surfactants, Sephacryl 200, ammonium persulfate (APS), N,N,N′,N′-
Tetramethylethylenediamine (TEMED) and all solvents were obtained from Sigma-
Aldrich. Sodium phosphate was obtained from EMD chemicals. Sodium chloride was
purchased from J. T. Baker. Cholesteryl-TEG-DNA (chol-DNA-1) was from Trilink
Biotechnologies. Acrydite-modified DNA (acry-DNA-2) was purchased from IDT.
FITC-labled peptide (vif2-FITC) was from Biomatic. DNA and peptide sequences are
given below. 8-aminonaphthalene-1,3,6-trisulfonic acid, disodium salt (ANTS), and p-
xylene-bis-pyridinium bromide (DPX) were purchased from Life Technologies
(Invitrogen). Methacryloxyethyl thiocarbonyl rhodamine B (rhodamine-methacrylate, rh-
MA) was from Polysciences.
DNA Sequences
chol-DNA-1:
5’-Cholesteryl-tetra(ethylene glycol)- CTCCCTTCTCTCTCT-3’
acry-DNA-2: 5’-Acrydite- AGAGAGAGAAGGGAG-3’
Peptide sequence
vif2-FITC: GTRQVTQASSFTWRVPG-Lys(FITC)-OH
4.2.2 Fabrication of Hydrogel-anchored Liposomes
Hydrogel-anchored liposomes were prepared as described in sections 3.2.2 and
3.2.3. Briefly, a solution of DOPE (70 wt%) and DOGS (30 wt%) at total concentration
of 2 mg/mL was prepared in 1:2 (v:v) solution of chloroform/methanol. Cholesterol-tetra
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(ethylene glycol)-modified ssDNA (Chol-DNA-1) was mixed with acrydite-modified
ssDNA (acry-DNA-2) that have a complementary sequence to that of DNA-1 at
equimolar concentrations (20 µM) in a solution of 1:2 (v:v) acetonitrile : PBS.
Oligonucleotides were annealed by incubating the solution at 90°C for 3-5 min and
cooling it down to room temperature gradually, and then, added to the solution of
DOPE/DOGS lipids. A dry lipid film was formed by evaporation of chloroform with
argon flow and keeping the lipid film under vacuum for at least 3 h. Then, a mixture of
10% (v) PEG-DA, 5% (v) PEG-MA, 0.2% (w/v) ammonium persulfate and 0.2% (v)
TEMED in PBS containing 10 mM phosphate and 150 mM NaCl at pH 7.5 was added to
the lipid film and sonicated for few seconds. To track nanogels, rh-MA was included in
the monomer mixture at 24 µM in some experiments. The solution was placed in an ice
bath to prevent polymerization prior to liposome formation. Liposomes were formed by
50× extrusion of the PEG-containing lipid solution through a 0.08 µm polycarbonate
filter in a cold room. To prevent polymerization exterior to the liposomes, the suspension
was diluted 5× with a solution of 2% (v) TEMED in PBS and incubated in a ~40°C
water bath for 15 minutes to polymerize the hydrogel. The hydrogel-liposomes were
dialyzed against PBS using dialysis tubing (300 KDa MWCO) for about 20 h and
concentrated with an Amicon ultracentrifuge tube with 100 kDa MWCO.
4.2.3 vif2 Peptide Encapsulation in Hydrogel-anchored Liposomes
To encapsulate the FITC-labeled peptide in hydrogel-anchored liposomes, FITC-
vif2 dissolved in 8% DMF in water was added to the PEG solution at final concentration
of 167 µM. The rest of procedure is exactly similar to that described in section 4.2.2. Free
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vif2-FITC was removed during dialysis of liposomes in dialysis tubing (300 KDa
MWCO).
4.2.4 Size Exclusion Chromatography of vif2-containing Liposomes
To confirm successful encapsulation of vif2 peptide in hydrogel core of liposomes,
CHAPS in PBS was added to the liposome suspension to a final concentration of 15 mM
CHAPS to strip off the lipid bilayer. As described in section 3.2.4, the liposome
suspension was then pumped over a C-type SEC column (GE Healthcare) (packed with
Sephacryl 200 (Sigma) and prewashed with CHAPS solution) at a 0.2 mL/min flow rate
to separate nanogels from lipid or lipid/DNA micelles. The SEC column packed with
Sephacryl 200 separates nanogels that have same hydrodynamic radius (R
H
) as liposomes
in the void fraction, while micelles with much smaller R
H
remain for a longer period of
time in column. A Shimadzu fluorescence detector (RF-535) integrated with the SEC
column was used to record fluorescence intensity (FI) chromatograms of the vif2 peptide
label. All FI values were normalized to the maximum FI recorded in each
chromatography run. The FITC dye conjugated to the peptide as a fluorescence probe
was excited at 490 nm and emission intensity was recorded at 520 nm. In control
experiments, surfactant-treated dsDNA-liposomes and plain liposomes both without
hydrogel were passed over the SEC column.
4.2.5 Determination of pH Sensitivity of Liposomes
The pH sensitivity of liposomes was confirmed by coencapsulation of 12.5 mM
ANTS, a pH-insensitive fluorophore, and 40 mM DPX quencher in liposomes and
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measuring the release of the dye into the medium at acidic pH. After encapsulation of dye
and quencher in the core of liposomes during rehydration process, free dye and quencher
were removed by passing the liposome suspension over a Zeba spin column with 7kDa
MWCO (Pierce). Then, liposomes were incubated in either 20% fetal bovine serum in
PBS or PBS at 37°C, and release of dye was measured by fluorescence
spectrophotometry at pH 7.5 and acidic pH at various times. The pH was lowered by
highly concentrated acetic acid solution. The dye was excited at 360 nm and emission
was recorded at 510 nm. Dye release was calibrated with 0% set to the residual
fluorescence of the liposomes at pH 7.5, and with 100% set to the fluorescence intensity
following lysis of liposomes with 1% TX-100.
4.2.6 Lipid Bilayer Lysis by Cholesterol-dsDNA-nanogels
To evaluate the lysis of intact liposomes with cholesterol-dsDNAs anchored to the
nanogel surfaces, hydrogel-anchored liposomes with dsDNA anchors prepared in section
4.2.2 were treated with 15 mM CHAPS surfactant and dialyzed against PBS (10 mM
sodium phosphate, 150 mM NaCl) for 4 days with two buffer exchanges. Various
concentrations of dsDNAs from 20 µM to 80 µM were used during liposome preparation.
Plain liposomes composed of 1:1 (molar ratio) DOPC/DOPE and encapsulated 12.5 mM
ANTS dye and 40 mM DPX quencher were prepared by extrusion of liposomes through
0.08 µM polycarbonate filter. No PEG and dsDNA anchors were used in fabrication of
plain liposomes. Unencapsulated dye/quencher were removed by passing the liposome
suspension over a Zeba spin column with 7kDa MWCO. Finally, cholesterol-dsDNA-
nanogels (chol-dsDNA-nanogels) and plain liposomes in PBS were mixed at a 1:1 (v:v)
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ratio. The pH of the suspension was lowered to acidic pH (4-5), and then, release of the
dye was measured by fluorescence spectrometer at 37°C at different time points. The
release percentage was calculated based on the fluorescent intensity following lysis of
liposomes with 1% TX-100.
As a control, nanogels with no dsDNA anchors were prepared by surfactant
treatment of hydrogel-liposomes, incubated with DOPC/DOPE plain liposomes with
ANTS/DPX at 1:1 (v:v) ratio, and dye release was measured over the time at 37°C.
4.2.7 Cellular Uptake of Hydrogel-anchored Liposomes
A549 lung carcinoma cells were induced either by interferon-inducible
transmembrane protein (IFITM3) cDNA or empty Vector to generate a stable cell line to
produce IFITM3 protein (A549-IFITM3) or A549-Vector (control cell line with no
induced protein). Cells were cultured in DMEM media supplemented with puromycin,
10% FBS, penicillin-streptomycin antiobiotic and non-essential amino acid (NEAA).
IFITM3 protein mainly localizes at late endosomal compartment and may reduce fluidity
of the endosomal membrane. Stable induction of IFITM3 impairs endosomal function
through generation of massive accumulation of cholesterol and rigid membrane in the late
endosomal compartment. Therefore, cellular internalization through receptor mediated
endocytosis (like clathrin-coated pit endocytosis) from early endosome to late endosomal
compartment is impaired in this cell line. This cell line has been generated by Prof. Jae U.
Jung's laboratory and described previously (Amini-Bavil-Olyaee et al. 2013). The A549-
IFITM3 expressing cell line can be an appropriate system for studying liposomal entry,
since it produces large endosomal compartemts. Since in this cell line endocytosed
79
matterials is stopped and accumulated in the enlarged endosomal compartment, they can
be tracked and montiroed by confocal microscopy.
For determining intracellular delivery of peptide-containing liposomes, liposomes
labeled with Rhod-PE and liposomes carrying FITC- peptide were freshly prepared and
then 40 µL of those added to the media of the A549-IFITM3 stable cell line. At different
time points, the treated cells were subjected to confocal microscopy as described below.
In a separate experiment, simultaneous delivery of encapsulated hydrogel with liposome
was confirmed by including crosslinkable rh-MA in PEG core of liposome labeled with
NBD-PE. To verify that liposomes travel to late endosomal compartment, CD63 protein,
a late endosome marker, was simultaneously probed by mouse anti-CD63 antibody
during confocal microscopy.
4.2.8 Confocal Microscopy of Cells
To observe internalization of liposomes, A549-IFITM3 cells were cultured on a
round thin glass coverslip a day before experiment. The cells were treated with either
Rhod-labeled liposomes or vif2-FITC-containing liposomes at different time points.
Cells were washed with PBS three times to remove unbound liposomes and fixed with
4% paraformaldehyde in PBS. Then cells were stained by DAPI to visualize the nucleus.
Coverslips were mounted on the slides and fluorescence images were captured by
confocal microscopy (Nikon, Eclipse Ti). The intracellular delivery of Rhod-labeled
liposomes was confirmed by excitation of rhodamine at 560 nm and emission of that at
590 nm. To observe vif2-FITC liposomes, FITC was excited at 490 nm and the emission
signal was recorded at 530 nm.
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4.3 Results and Discussion
Liposomes are promising nanostructures for delivery of therapeutics, especially
anticancer drugs. From few years ago, liposomes serve as carriers of different types of
proteins and peptides. Incorporation of protein and peptide drugs in liposomes improves
their therapeutic activity (Torchilin 2005), and prevents their rapid elimination from the
blood circulation, their enzymatic degradation, their uptake by the reticulo-endothelial
system (RES) and their non-selective accumulation (Banga and Chien 1988; Ducat et al.
2011; Katanasaka et al. 2008; Torchilin 2006; Zhou and Li Wan Po 1991). Due to
biocompatibility and biodegradability, liposomes cause no or mild antigenic, pyrogenic,
allergic and toxic reactions (Ducat et al. 2011). In this study, we encapsulated an
anticancer peptide with 17 amino acids called vif2 in stable liposomes developed in
chapter 3 and used them as delivery vehicles for cancer therapy. The vif2 peptide that
regulates p53 activity leading to apoptosis in cancer cells has been discovered by Lee and
coworkers in Prof. Jae U. Jung's laboratory (Lee et al. 2011). Liposomes provide
protected cellular delivery of peptides by preventing their degradation. In order to use
pH-triggered mechanism for cellular delivery of encapsulated peptide, we fabricated
liposomes composed of DOGS, a pH-sensitive lipid, and DOPE which enhances the cell
internalization process. During endocytosis, the pH decreases within the endosome. Upon
pH decrease to acidic pH, the carboxylic groups of DOGS lipids are protonated, DOPE
molecules convert into their inverted hexagonal phase, and liposomes destabilize and
cause gradual release of peptides from nanogels (Figure 4.1). These pH-sensitive
liposomes have previously been shown to be effective, nontoxic carriers for intracellular
delivery of various types of drugs (Liu and Huang 1990).
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Figure 4.1. Cellular internalization of pH-sensitive liposomes carrying vif2 peptide.
4.3.1 vif2 Peptide and Its Mechanism of Action
vif2 peptide which has been discovered as an anticancer peptide by Lee and
coworkers,(Lee et al.) is composed of 17 amino acids (see materials for sequence). This
peptide has been derived from Kaposi's sarcoma-associated herpesvirus viral interferon
(vIRF4). vif2 restores the p53 pathway which has an important role in regulating the cell
cycle, and therefore, tumor suppression and apoptosis (Lee et al. 2011). The p53 pathway
has been a prime target for new cancer-drug developments. It has been known that the
vif2 peptide binds to the catalytic domains of Herpesvirus-associated ubiquitin-specific
protease (HAUSP) enzyme and robustly suppress its deubiquitination activity (Lee et al.
2011). Ubiquitin is a small protein that binds to proteins and labels them for destruction.
Blocking HAUSP by vif2 results in p53-dependent cell-cycle arrest and apoptosis.
HAUSP is a well-characterized deubiquitinase enzyme and it is capable to remove
ubiquitin moieties from ubiquitinated substrates.
82
4.3.2 vif2 Encapsulation in Liposomes
Encapsulation of vif2 peptide in the polymeric core of liposomes was demonstrated
by size exclusion chromatography (SEC) of vif2-FITC-containing liposomes. Since vif2
solution was miscible with monomer solution, we expected to encapsulate peptide in the
cores of liposomes during rehydration of the lipid film. As shown in Figure 4.2, intact
and surfactant-treated vif2-containing hydrogel-anchored liposomes have the same
retention time in an SEC column and they both eluted in the void volume. Sephacryl 200
allows liposome-sized structures to elute in the void volume, while retaining smaller size
particles, such as lipid/surfactant micelles in the column. Since these chromatograms
track conjugated FITC to vif2 peptide, these results confirm the successful encapsulation
of vif2 in nanogels that have similar hydrodynamic radius (R
H
) as liposomes.
Figure 4.2. Normalized SEC chromatograms tracking FITC-vif2 in various liposomes with dsDNA
anchors. The retention time is the same for vif2-FITC peptide in intact liposomes and surfactant-stripped
nanogels. Surfactant-treated dsDNA-liposomes and plain liposomes with no hydrogel have longer retention
time, indicating that a smaller size peptide micelles are formed.
83
Control experiments with surfactant-treated dsDNA-liposomes and plain
liposomes that do not contain hydrogel show fluorescence species with longer retention
time and therefore smaller R
H
, corresponding to the free peptide with smaller size than
intact liposomes (Figure 4.2). Chromatogram of Rhod-PE included in liposomes confirms
successful removal of lipid bilayer after treatment of liposomes with CHAPS surfactant
(data not shown).
4.3.3 Destabilization of pH-sensitive Liposomes at Acidic pH
Since pH sensitivity of liposomes is important for intracellular delivery of vif2
peptide, we fabricated pH-sensitive liposomes composed of DOGS and DOPE. Several
studies have shown that small liposomes composed of DOPE and a double chain
amphiphile such as DOGS are pH-sensitive and useful for the cytoplasmic delivery to
target cells (Collins et al. 1990; Leventis et al. 1987; Liu and Huang 1990). Although
other types of amphiphiles with a single chain, such as fatty acids have been used for
fabrication of pH-sensitive liposomes, they usually do not have serum stability due to
rapid transfer of fatty acids to serum proteins (Collins et al. 1990). In contrast, liposomes
with double-chain amphiphiles are relatively serum stable at physiological pH and show
less leakiness comparing with those composed of single chain lipids (Collins et al. 1990;
Leventis et al. 1987). These liposomes can still maintain their pH sensitivity (Collins et al.
1990).
To study pH sensitivity of fabricated liposomes, we measured leakage of ANTS
dye from various types of liposomes at low pH. ANTS fluorescence is quenched with
coencapsulated DPX quencher by collisional energy transfer (Ellens et al. 1984). Since
84
ANTS is a charged molecule, it cannot pass the intact liposomal membrane. Therefore,
release of the dye is evidence for destabilization of liposomes at low pH. As shown in
Figures 4.3A-C, no significant dye release was observed from plain liposomes, hydrogel-
liposomes and hydrogel-anchored liposomes incubated at pH 7.5 and 37°C in PBS buffer.
Upon decrease of pH at 37°C, all types of liposomes show destabilization and 80-90%
dye release was observed over 120 min. These observations demonstrate successful
fabrication of pH sensitive liposomes that are stable at physiological pH while releasing
their content at low pH environment.
According to Figure 4.3D, all three types of liposomes showed pH sensitivity in
20% serum. Double-stranded hydrogel-anchored liposomes and hydrogel-liposomes
ultimately showed ~30% dye release over 120 min. Plain liposomes showed ~60%
release within 120 min incubation at pH 4.6 and 37°C. As shown by other research
groups, the lower pH sensitivity of small liposomes in serum is probably due to the
insertion of plasma proteins in the bilayer of liposomes (Collins et al. 1990; Leventis et al.
1987). No significant release was observed in different liposome species incubated in
serum at pH 7.5. These results indicate that DOPE/DOGS liposomes are appropriate
nanostructures for intracellular delivery of vif2 peptide.
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Figure 4.3. pH-induced leakage of nano-scale liposomes composed of 70% (wt%) DOPE and 30% (wt%)
DOGS. Dye release was measured at different time points at physiological pH (pH 7.5) and acidic pH (pH
4-5) in liposomes incubated at 37°C. (A), (B) and (C) Dye leakage in PBS buffer; and (D) Dye leakage in
20% FBS serum.
4.3.4 Lipid bilayer Lysis by Cholesterol-dsDNA-nanogels
In order to show the potential lysis of the endosomal membrane with chol-dsDNA-
nanogels, we studied lysis of plain liposomes encapsulating ANTS/DPX with dsDNA-
anchored nanogels. As mentioned before, dsDNA-nanogels are prepared by surfactant
86
treatment of hydrogel-anchored liposomes and removal of lipid bilayer. Due to the
presence of cholesterol moieties, acrylate-DNA-cholesterol macroamphiphile that
participates in lipid bilayer self assembly and serves to stabilize lipid bilayers can
facilitate cellular internalization of nanogels. To show this, liposomes composed of pH-
insensitive DOPE and DOPC lipids and encapsulated ANTS/DPX were prepared in PBS
buffer, incubated with chol-dsDNA-nanogels at acidic pH and 37°C, and release of the
dye from liposomes was measured over the time (Experimental section). Since liposomes
were intact, release of the dye was evidence for lysis of them by chol-dsDNA-nanogels.
Figure 4.4 shows that increasing concentration of chol-dsDNA anchors from 20 µM to 80
µM increases the amount of dye released from intact liposomes. Increasing chol-dsDNAs
concentration resulted in faster dye release within first few hours of leakage observation.
Figure 4.4. Release of dye from plain DOPE/DOPC liposomes incubated with chol-dsDNA-nanogels made
with different concentrations of dsDNAs, at 37°C and pH 4.6. Higher concentration of chol-dsDNA
anchors on nanogels increases the leakage of dye.
87
In a control experiment, nanogels with no dsDNA anchors, obtained from
surfactant treatment of hydrogel-liposomes, were incubated with dye/quencher-
containing liposomes at 37°C and pH 4.6. No significant dye release was observed within
140 h (Figure 4.4). These observations indicate the important role of cholesterol-
conjugated dsDNAs in lipid membrane lysis. Liposome-templated nanogels surface-
modified with amphiphilic, programmable dsDNAs have potential application for
intracellular delivery of therapeutic compounds.
4.3.5 Cellular Internalization of Peptide-containing Liposomes
To observe cell internalization of hydrogel-anchored liposomes and their
encapsulated vif2 peptide, we used fluorescence confocal microscopy. The images were
obtained from A549-IFITM3 cells incubated with liposomes containing rhodamine-
labeled lipid (Rhod-PE) and liposomes encapsulating FITC-labeled peptide
(Experimental section). Since A549-IFITM3 cells have impaired late edosomal function
that can stack liposomes entered by endocytotic pathway, using them can prove
successful internalization of liposomes. Confocal images shown in Figures 4.5 and 4.6
confirm efficient and successful internalization of vif2-containing liposomes into cells.
In those images, increasing fluorescence signals of rhodamine and FITC can be
observed over longer incubation time of liposomes with cells. As can be seen in Figure
4.5A, liposomes have been accumulated on plasma membrane after immediate incubation
with cells, while by increasing incubation time higher concentration of liposomes has
been endocytosed and accumulated in the endosomal compartment (Figures 4.5 B-F).
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Figure 4.5. Fluorescence confocal microscopy images of A549-IFITM3 cells incubated with liposomes
with Rhod-PE lipid at various times. (A) Enlarged image of cell after immediate incubation with
liposomes; (B) 1h; (C) 2h; (D) 3h; (E) 4h; and (D) Overnight incubation with cells. These images show
successful internalization of liposomes into cells. Scale bars: 10 µm.
Figure 4.6. Fluorescence confocal microscopy images of A549-IFITM3 cells incubated with hydrogel-
anchored liposomes encapsulating FITC-peptide at various times. (A) Enlarged image of cell shows
successful delivery of encapsulated peptide after 30 min; (B) 3 h; (C) and (D) Overnight incubation of cells
with liposomes. Scale bars: 10 µm.
Overlaying images of cells incubated with hydrogel-anchored liposomes, labeled
with rh-MA crosslinked to hydrogel and NBD included in the lipid bilayer, showed
89
delivery of encapsulated nanogels along with liposomes through endocytotic pathway
(Figure 4.7).
Figure 4.7. Fluorescence confocal microscopy of A549-IFITM3 cells incubated for 3 h with NBD-labeled
liposomes encapsulating rh-MA crosslinked to PEG hydrogel. (A) Nuclei blue staining with DAPI; (B)
Rhodamine signal from nanogels labeled with rh-MA; (C) NBD signal from liposomes labeled with NBD-
PE; (D) overlaying images of rhodamine and NBD. Scale bars are 5 µm.
As mentioned in section 4.2.7, the correct internalization of liposomes in late
endosomal compartment was determined by CD63 protein marker. The A549-IFITM3
cells were fixed and subjected for confocal microscopy after internalization of NBD-
labeled liposomes encapsulating rh-MA hydrogels.
Figure 4.8 shows delivery of liposomes in late endosomal compartment stained
with anti-CD63 antibody. Overlaying images of NBD, rhodamine and CD63 flurescence
emissions demonstrate delivery of hydrogel-anchored liposomes in late endosome
through endocytosis. Results obtained from confocal images shows that hydrogel-
anchored liposomes are well-designed nanocarriers for protected intracellular delivery of
vif2-petide.
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Figure 4.8. Fluorescence confocal microscopy of A549-IFITM3 cells incubated with NBD-labeled
liposomes encapsulating rhodamine-hydrogels. (A) Nuclei blue staining with DAPI; (B) CD63 stained
cells; (C) NBD signal from NBD-liposomes; (D) Rhodamine signal from rh-MA hydrogels; and (E)
overlaying images of rhodamine,NBD and CD63. Scale bars are 20 µm.
4.3.6. Conclusions
We used stable hydrogel-anchored liposomes with dsDNA anchors to carry and
deliver anticancer peptide. For pH-triggered release of peptide, liposomes composed of
pH-sensitive lipid were fabricated. FITC-labeled peptide was encapsulated in hydrogel
core of liposome during rehydration of lipid film with PEG monomer solution. SEC of
intact and surfactant-treated liposomes confirmed successful encapsulation of peptide in
liposomes. Release studies of liposomes, coencapsulated dye and quencher, showed pH
sensitivity of them at low pHs while preserving their stability at physiological pH.
Intracellular delivery of peptide-containing liposomes was shown by fluorescence
confocal microscopy. Images were taken from liposomes with Rhod-labeled lipid and
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FITC-labeled peptide. Overlapping signals from rhodamine and NBD was confirming
evidence for intracellular delivery of nanogels with liposomes. Liposome lysis with chol-
dsDNA-nanogels was also shown by dye dequenching experiment. We observed
increasing dye release from intact liposomes incubated with chol-dsDNA-nanogels
contained various concentrations of chol-dsDNAs. This result confirmed potential
application of chol-dsDNA-nanogels for cellular delivery. The hydrogel-anchored
liposomes with higher stability are promising nanostructures for safe and protected
delivery of biomolecules, such as proteins and peptides.
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Chapter 5: Conclusions and Recommendations
We have presented studies for the development and fabrication of biomimetic,
lipid-based hybrid nanostructures that have potential applications in biosensing,
bioelectronics, drug delivery and biophysical studies of cell membranes. We integrated
lipid membranes with synthetic materials such as carbon nanotubes and hydrogels to
construct new lipid-hybrid nanostructures that have hybrid properties of both lipid
membranes and nanomaterials.
We presented a novel method for the fabrication of dispersible, biocompatible
carbon nanotubes using lipid bilayers. First MWCNTs were covalently conjugated to
amine-bearing lipid molecules in a carbodiimide-mediated reaction. Lipid-conjugated
nanotubes were highly dispersible in organic solvent. FTIR demonstrated the covalent
conjugation of lipid molecules to the nanotube surface. In a sonication process, a second
monolayer of lipid was self-assembled on the first monolayer, forming a biomimetic lipid
bilayer on the surface of MWCNTs. These bilayer-coated nanotubes were observed by
fluorescence confocal microscopy. Insertion of the membrane protein α-hemolysin into
the lipid bilayer and a fluorescence quenching experiment using dithionite showed the
successful formation of biomimetic lipid bilayers on surface of nanotubes. The fludity of
the conjugated lipid bilayer was confirmed by fluorescence anisotropy measurements.
These stable, biomimetic nanostructures are fascinating platforms for integration of
carbon nanotubes with various types of biomolecules.
Liposomes have various biomedical applications. They can be used as models of
biological lipid membranes for biophysical studies of cells. Since liposomes generally
unstable in long-term storage and in biological fluids, we developed a stable, biomimetic
93
nanoscale liposome that mimics the attachment of membrane-resident molecules to the
cytoskeleton. In a self-assembly procedure, the lipid bilayer of a liposome was covalently
anchored to a biocompatible poly(ethylene) glycol (PEG) hydrogel core using
programmable double-stranded DNA (dsDNA) linkers. These dsDNA linkers are
composed of two 15-mer complementary single-stranded DNAs (ssDNAs) modified with
cholesterol and vinyl groups. Cholesterol moieties of dsDNAs reside in the lipid bilayer
while vinyl groups incorporate in the cross-linked hydrogel backbone. A novel thermally
controlled method was used to form nanogels in the liposomes. Size exclusion
chromatography (SEC) of intact and surfactant-treated liposomes confirmed the
formation of nanogels anchored to the lipid membrane. Transmission electron
microscopy (TEM) showed ~100 nm nanoparticles before and after removal of
unanchored lipids, confirming the formation of PEG nanogels in liposomes. Dynamic
light scattering (DLS) also showed ~120 nm size nanoparticles corresponding to the
hydrogel-anchored liposomes. Upon lipid bilayer removal, we obtained dsDNA-
decorated nanogels which have potential sites for further molecular assembly.
Using the lipid bilayer to direct assembly, we could organize oligonucleotides on
the surface of nanogels. A dsDNA sensitive dye, Hoechst 33342, verified the location of
dsDNA anchors on the surface of nanogels. A dye dequenching assay was used to
demonstrate the vastly improved stability of hydrogel-anchored nanostructures versus
both standard liposomes and liposomes containing a hydrogel core without anchoring. A
dye/quencher pair, ANTS/DPX, was coencapsualted in liposomes and release of dye was
measured upon incubation of liposomes in 20% serum. The fabricated lipid-polymer
94
nanostructures can provide robust drug delivery vehicles with advantages of sustained
drug release and the potential for targeted delivery.
We have used stable hydrogel-anchored liposomes (chapter 3) for the intracellular
delivery of a peptide with anticancer properties. These stable nanostructures represent a
unique platform for protected delivery of oligonocleotides and therapeutic compounds
and allow formation of templated nanogels with programmable molecular recognition
sites. We included a pH-sensitive lipid, DOGS, into the lipid membrane to trigger release
in and disruption of late-stage endosomes. Release kinetics studies of these pH-sensitive
liposomes showed that liposomes are stable at neutral pH while rapidly releasing cargo at
acidic pH. We traced intracellular delivery of the nanoparticles and their cargo, the vif2
peptide, by fluorescent confocal microscopy. Confocal images showed successful
intracellular delivery of vif2-containing hydrogel-anchored liposomes. In this pathway,
cholesterol-dsDNA moleculess arranged on surface of nanogels can facilitate the
intracellular delivery of peptide-encapsulated nanogels by disrupting late stage
endosomes.
As mentioned in chapter 4, Lee and coworkers have shown that the vif2 peptide
has a cytotoxic effect on lymphoma tumor cell lines, and it can robustly suppress cell
proliferation and induce cell death to a significant extent (Lee et al. 2011). The vif2
peptide prohibits cell proliferation by regulating P53 protein activity, resulting in
apoptotic cell death. In vivo studies have also shown the efficient and powerful tumor
regression by vif2 in a mouse model. Due to vulnerability of peptide to digestion and
degradation, encapsulation and delivery of the peptide in developed stable liposomes was
suggested. Up to now, we have shown the successful encapsulation and intracellular
95
delivery of vif2 using hydrogel-anchored liposomes (chapter 4), but overall efficiency of
this lipid-based nanostructure in cellular delivery of vif2 should be examined by using the
functional assay developed by Lee and coworkers (Lee et al. 2011) in Jung's research
group. It is important to find out how peptide-containing liposomes differ from plain
peptide in cancer cell growth regression and apoptosis. The success of liposomes in
protected and safe delivery of peptide should be studied.
In addition, release kinetics of the encapsulated peptide should be examined over
time both in vitro and in vivo. Since a limited concentration of peptide is encapsulated in
hydrogel core of liposomes, it is also necessary to find the optimum concentration of
peptide required for desired and efficient biological effect. Following in vitro studies of
hydrogel-anchored liposomes in peptide delivery, in vivo studies will be also suggested to
show the efficient application of liposomes in delivery of peptide.
PEG has been widely used as hydrogel for drug delivery purposes because of their
excellent biocompatibity (Bell and Peppas 1996; Leach and Schmidt 2005; Nie et al.
2007). Different types of chemistries have been used to create reactive PEGs with
functionalized end-groups, such as acrylate, thiol, amine, and maleimide that can
crosslink and form hydrogels (Zustiak and Leach 2010). However, the crosslinked
network of PEG is nondegradable in physiological conditions (Zustiak and Leach 2010).
Degradable PEG hydrogels are desirable for delivery applications and controlled release
of the delivered component. A number of strategies has been used to make biodegradable
PEG-based hydrogels, especially in a hydrolytic manner (Zustiak and Leach 2010).
Hydrolysis of PEG hydrogels is more desirable than other strategies for delivery, because
it does not need biological molecules, such as enzymes for degradation, and it can
96
produce low molecular weight compounds after degradation that can be cleared from the
body easily (Zustiak and Leach 2010). To make PEG nanogels in our hydrogel-anchored
liposome, we used PEG-diacrylate and PEG-methacrylate monomers that cannot form a
degradable crosslinked structure. Therefore, delivery of encapsulated compounds is
limited to diffusion through hydrogel pores that is not really efficient way for delivery.
For the most favorable drug delivery purposes, it is recommended to synthesize
degradable PEG hydrogel and allow for release studies of model drugs or biomolecules,
such as peptides. Moreover, by synthesizing dsDNA anchors with a degradable anchoring
group, we will be able to use dsDNA-nanogels for oligonucleotide delivery and gene
therapy.
97
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Abstract (if available)
Abstract
Biological membranes serve several important roles, such as structural support of cells and organelles, regulation of ionic and molecular transport, barriers to non‐mediated transport, contact between cells within tissues, and accommodation of membrane proteins. Membrane proteins and other vital biomolecules incorporated into the membrane need a lipid membrane to function. Due to importance of lipid bilayers and their vital function in governing many processes in the cell, the development of various models as artificial lipid membranes that can mimic cell membranes has become a subject of great interest. ❧ Using different models of artificial lipid membranes, such as liposomes, planar lipid bilayers and supported or tethered lipid bilayers, we are able to study many biophysical processes in biological membranes. The ability of different molecules to interact with and change the structure of lipid membranes can be also investigated in artificial lipid membranes. An important application of lipid bilayer‐containing interfaces is characterization of novel membrane proteins for high throughput drug screening studies to investigate receptor‐drug interactions and develop biosensor systems. Membrane proteins need a lipid bilayer environment to preserve their stability and functionality. Fabrication of materials that can interact with biomolecules like proteins necessitates the use of lipid bilayers as a mimic of cell membranes. ❧ The objective of this research is to develop novel hybrid lipid‐based nanostructures mimicking biological membranes. Toward this aim, two hybrid biocompatible structures are introduced: lipid bilayer‐coated multi‐walled carbon nanotubes (MWCNTs) and hydrogel‐anchored liposomes with double‐stranded DNA anchors. These structures have potential applications in biosensing, drug targeting, drug delivery, and biophysical studies of cell membranes. ❧ In the first developed nanostructure, lipid molecules are covalently attached to the surfaces of MWCNTs, and then, using a sonication process, a uniform lipid bilayer that supports the incorporation of membrane proteins is formed. These bilayer‐coated carbon nanotubes are highly dispersible and stable in aqueous solution, and they can be used in development of various biosensors and energy producing devices. ❧ In the other hybrid nanostructure, the lipid bilayer of a liposome is covalently anchored to a biocompatible poly(ethylene) glycol (PEG) hydrogel core using double‐stranded DNA (dsDNA) linkers. Release studies shows that nano‐size hydrogel‐anchored liposomes are exceptionally stable, and they can be used as biomimetic model membranes that mimic the connectivity between the cytoskeleton and the plasma membrane. After lipid bilayer removal, dsDNA linkers can provide programmable nanogels decorated with oligonucleotides with potential sites for further molecular assembly. These stable nanostructures can be useful for oligonucleotide and drug delivery applications. ❧ The developed hydrogel‐anchored liposomes are exploited for encapsulation and intracellular delivery of therapeutic peptide. Peptides with anti‐cancer properties are successfully encapsulated in hydrogel core of pH‐sensitive liposomes during rehydration process. Liposomes release their cargo at acidic pH. Confocal microscopy confirms the intracellular delivery of liposomes through an endocytotic pathway.
Linked assets
University of Southern California Dissertations and Theses
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Asset Metadata
Creator
Dayani, Yasaman
(author)
Core Title
Hybrid lipid-based nanostructures
School
Viterbi School of Engineering
Degree
Doctor of Philosophy
Degree Program
Chemical Engineering
Publication Date
07/11/2014
Defense Date
05/22/2014
Publisher
University of Southern California
(original),
University of Southern California. Libraries
(digital)
Tag
biomimetic lipid bilayers,carbon nanotube,DNA anchors,Hybrid nanostructures,Hydrogel‐anchored liposomes,Intracellular delivery,OAI-PMH Harvest,PEG nanogels
Format
application/pdf
(imt)
Language
English
Contributor
Electronically uploaded by the author
(provenance)
Advisor
Malmstadt, Noah (
committee chair
), El-Naggar, Mohamed Y. (
committee member
), Wang, Pin (
committee member
)
Creator Email
ydayani@gmail.com,ydayani@usc.edu
Permanent Link (DOI)
https://doi.org/10.25549/usctheses-c3-436011
Unique identifier
UC11286857
Identifier
etd-DayaniYasa-2656.pdf (filename),usctheses-c3-436011 (legacy record id)
Legacy Identifier
etd-DayaniYasa-2656.pdf
Dmrecord
436011
Document Type
Dissertation
Format
application/pdf (imt)
Rights
Dayani, Yasaman
Type
texts
Source
University of Southern California
(contributing entity),
University of Southern California Dissertations and Theses
(collection)
Access Conditions
The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the a...
Repository Name
University of Southern California Digital Library
Repository Location
USC Digital Library, University of Southern California, University Park Campus MC 2810, 3434 South Grand Avenue, 2nd Floor, Los Angeles, California 90089-2810, USA
Tags
biomimetic lipid bilayers
carbon nanotube
DNA anchors
Hybrid nanostructures
Hydrogel‐anchored liposomes
Intracellular delivery
PEG nanogels