DNA nanotechnology as a tool to manipulate lipid bilayer membranes
Meng, Zhuojun
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DNA nanotechnology as a tool to
manipulate lipid bilayer membranes
DNA nanotechnology as a tool to manipulate lipid bilayer membranes
Zhuojun Meng PhD thesis
University of Groningen October 2017
Zernike Institute PhD thesis series 2017-22 ISSN: 1570-1530
ISBN: 978-90-367-9976-8(printed version) ISBN: 978-90-367-9975-1(electronic version)
The research described in thesis was carried out in Polymer Chemistry and Bioengineering group at Zernike Institute for Advanced Materials, University of Groningen, The Netherlands. This work was financially supported by the Chinese Scholarship Council (CSC), the University of Groningen and the Netherlands Organization for Science Research (NWO).
Cover design by: Zhuojun Meng Printed by: Ridderprint BV
DNA nanotechnology as a tool to
manipulate lipid bilayer membranes
PhD thesis
to obtain the degree of PhD at the University of Groningen
on the authority of the Rector Magnificus Prof. E. Sterken
and in accordance with the decision by the College of Deans. This thesis will be defended in public on
Friday 13 October 2017 at 16.15 hours
by
Zhuojun Meng
born on 5 May 1987 in Henan, China
Supervisor
Prof. A. Herrmann
Assessment committee
Prof. S. Vogel
Prof. A. M. van Oijen Prof. D. J. Slotboom
Dedicated this book to myself
and my best friend
Contents
Chapter 1
Functionalization of LipidBilayer Membranes ... 9
1. 1 Lipid bilayer membranes ... 10
1.2 Classification and Preparation of Liposomes ... 12
1.3 Modification and Applications of liposomes ... 14
1.4 Motivation and Thesis Overview ... 24
References ... 26
Chapter 2 Stability Study of Lipid-DNA on the Liposomal Membrane ... 31
2.1 Introduction ... 32
2.2 Results and Discussion ... 35
2.3 Conclusion ... 42
2.4 Experimental Section ... 42
References ... 48
Chapter 3 Efficient Fusion of Liposomes by Nucleobase Quadruple-Anchored DNA .. 51
3.1 Introduction ... 52
3.2 Results and Discussion ... 54
3.3 Conclusion ... 63
3.4 Experimental Section ... 65
Chapter 4
DNA Replacement and Hybridization Chain Reaction on the Surface of
Liposome Membrane ... 73
4.1 Introduction ... 74
4.2 Results and Discussion ... 76
4.4 Experimental Section ... 84
References ... 88
Chapter 5 Performing DNA Nanotechnology Operations on a Zebrafish Surface ... 91
5.1 Introduction ... 92
5.2 Results and Discussion ... 94
5.3 Conclusion ... 101 5.4 Experiment Section ... 103 References ... 105 Summary ... 108 Samenvatting ... 114 Acknowledgements ... 119
Chapter 1
Functionalization of
Lipid Bilayer Membranes
10
1. 1 Lipid bilayer membranes
Lipids play an important role in the physiology and pathophysiology of living systems why they are produced, transported, and recognized by the concerted actions of numerous enzymes, binding proteins, and receptors.1
Micelles are formed by the aggregation of single-chain lipids in a polar solvent (such as water) beyond a particular concentration, known as Critical Micelle Concentration (CMC) (Fig. 1.1A). Therefore, the micelle formation and stability are highly dependent on the lipid concentration and solvent composition (Fig. 1.1B).
Two-chain lipids can hardly be packed into micelles due to the bulky hydrophobic part. They usually form a lipid bilayer membrane, which is a thin polar sheet made of two layers of lipid molecules and is characterized by hydrophobic tails facing inwards towards each other and hydrophilic head groups facing outwards to associate with aqueous solution.2 At this
moment, the hydrophobic parts of the molecules are still in contact with water, which leads to an energetically unfavorable state of the bilayer. This is overcome through folding of the bilayer membrane into a liposome with closed edges (Fig. 1.1C).3,4
Functionalization of Lipid Bilayer Membranes
11 Fig. 1.1 (A) Surface tension as a function of the surfactant concentration. Schematic structure of a micelle (B) and a liposome (C). (Fig. 1.1 C was adapted from reference 4)
12
1.2 Classification and Preparation of Liposomes
Depending on the number of bilayers, liposomes can be classified into two categories: unilamellar vesicles (ULV) and multilamellar vesicles (MLV). Unilamellar vesicles can also be classified into three categories on the basis of their sizes, which can vary from nanometer to micrometer range: small unilamellar vesicles (SUV), large unilamellar vesicles (LUV) and giant unilamellar vesicles (GUV). GUVs also include other morphologies such as multilamellar vesicles (MLV), which consist of SUVs or multiple concentric bilayers (Fig. 1.2).
Fig. 1.2 Schematic structure of unilamellar and multilamellar liposomes.
There are four classical methods to prepare liposomes, differing in the way how the lipids are dried from organic solvent and then redispersed in aqueous buffer.5 These steps can be performed individually or jointly.6
These four methods are:
1. Hydration of a Thin Lipid Film.7
2. Reverse-Phase Evaporation Technique.8
3. Solvent (Ether or Ethanol) Injection Technique.9,10
Functionalization of Lipid Bilayer Membranes
13 Since the “Hydration of a Thin Lipid Film” method, is widespread used and easy to handle, it is explained here in more details. Firstly, the lipids are dissolved and mixed in an organic solvent to assure a homogeneous mixture. Once the lipids are thoroughly dispersed in the organic solvent, the solvent is removed using a dry nitrogen stream in a fume hood to yield a lipid film. The lipid film is dried to remove residual organic solvent by using a vacuum desiccator overnight. Afterwards, hydration of the dry lipid film is accomplished by stirring in an aqueous buffer. The temperature of the hydrating buffer should be higher than the gel-liquid crystal transition temperature (Tc) of the lipid. Subsequently, several stirring (above the Tc) and freeze-thawing cycles of the swelling multilayer sample results in MLVs. Finally, the sample is extruded multiple times using an extruder and polycarbonate membranes to obtain unilamellar vesicles (LUVs or SUVs).
Fig. 1.3 shows the classical hydration method of liposome preparation.
Fig. 1.3 Schematic diagram of liposome preparation method. (Schematic obtained from www. avantilipids.com)
14
1.3 Modification and Applications of liposomes
The first discovery of liposomes in 1964 by A. D. Bangham12 was the
starting point for these self-assembled containers to become a multifunctional tool in biology, biochemistry and medicine today. Because of the structure, charge, chemical composition and colloidal size can be well controlled by preparation methods, liposomes can be useful in various applications. Vesicles can also be prepared from natural substances and are therefore in many cases nontoxic, biodegradable, biocompatible, targetable and non-immunogenic.13 Due to these properties, liposomes can be used as
drug14-16 protein, plasmid17 and gene18-21 delivery vehicles in medicine and
diagnosis.
1.3.1 Loading and surface modification
Molecular interactions between the cargo and the lipid bilayer membrane play an important role on liposome formation and cargo encapsulation.22
Liposomes consist of an aqueous core surrounded by a lipid bilayer, sectioning off two separate inner areas. They can carry hydrophobic molecules in their hydrocarbon tail region (between the phospholipid bilayer), or hydrophilic molecules in the core and direct the cargo to the required diseased site in the body with some targeting moieties on the surface.23 The thickness of the lipid bilayer is around 4 to 10 nm, which is a
natural barrier for many substances such as sugars and proteins.24 But
small hydrophilic substances such as water, gases, ammonia and glycerol can penetrate freely through the bilayer.25-27 Some large hydrophilic
substances can be encapsulated in the water core of the liposome during liposome preparation using the common thin layer hydration method. Cationic liposomes, which are made of positively charged lipids, appear to be better suited for DNA delivery due to the natural charge-charge interaction between the positively charged lipid head groups and the negatively charged phosphate groups of the DNA-backbone.28,29 Due to
their favorable interactions with negatively charged DNA and cell membranes,30-33 cationic liposome–DNA complexes are increasingly being
researched for their use in gene therapy and nucleic acid release.34,35 In
Functionalization of Lipid Bilayer Membranes
15 tissues, the use of targeted liposomes with surface modification has been suggested.
Surface modification of liposomes with controlled propertied requires the chemical conjugation of peptides, DNA, antibodies or other targeting molecules. Moreover, some “smart” vesicle designs allow the release of the encapsulated cargo by incorporation of transport channels.36-39 Both
chemical attachment and physical interactions can be used to achieve surface modification (Fig. 1.4A).
Fig. 1.4 (A) Schematic representation of liposomes surface modifications. (B) Interaction of the particle with cell surface antigens and receptors.40 (C) Scheme of tetrac tagged liposome and
enhanced delivery by the ligand-mediated targeting strategy.41(Fig. 1.4 B was adapted from
reference 40. Fig. 1.4 C was adapted from reference 41)
To realize active targeting, the liposome surface can be coated with ligands or antibodies that will confer cell type-specificity to ensure that the liposomes are internalized and that their content is released, improving the efficacy and reducing side effects over non-targeted cells (Fig. 1.4B).40 For
16
to integrin αvβ3, was used for the surface modification of liposomes and successfully enhanced the tumor-targeting ability of PEGylated liposome (Fig. 1.4C).41 Although the physical properties of liposomes were not
significantly changed, tetrac-tagged liposomes showed significantly higher cancer cell localization than the unmodified PEGylated liposome, and tumor growth was effectively retarded. The ligand-mediated targeting strategy could provide better therapeutic effects with more accurate delivery of nanoparticles.
1.3.1.1 Membrane fusion
Surface modification of lipid bilayers can also be used for membrane fusion which is an essential process of life resulting in the highly regulated transport of bio-molecules both between and within cells.42-44 Membrane
fusion is an essential but not a spontaneous process as free energy is required to overcome the electrostatic and steric repulsions between two merging membrane surfaces and to break the hydration shell.45,46 A highly
conserved protein machinery, known as SNARE proteins (soluble N-ethylmaleimide sensitive factor attachment protein receptors), facilitates the communication within a cell.47-49 The SNAREs from synaptic vesicles
interact with the SNAREs from the target membrane to form a coiled-coil bundle of four helices, pulling the membranes tightly together and initiating fusion.
Design and construction of simplified artificial model systems mimicking natural systems are one of the most promising approaches for studying complex biological mechanisms.50 Several of these systems have been
reported for realizing membrane fusion, such as DNA51-53, peptides54, 55,
enzymes56 and polymers57. Yang et al. designed an artificial biorthogonal
targeting system that was able to target liposomes and other nanoparticles efficiently to the tissue of interest by using coiled coil forming peptides, E4[(EIAALEK)4] (E4) and K4[(KIAALKE)4] (K4) (Fig. 1.5C), which are
known to trigger liposomal membrane fusion when tethered to lipid vesicles in the form of lipopeptides.58 The same group proved that E4
peptide-modified liposomes could deliver far-red fluorescent dye TOPRO-3 iodide (E4-Lipo-TP3) and doxorubicin (E4-Lipo-DOX) into HeLa cells
Functionalization of Lipid Bilayer Membranes
17 expressing K4 peptide (HeLa-K) on the surface. Then, Lipo-TP3 and E4-Lipo-DOX were injected into zebrafish xenografts of HeLa-K (Fig. 1.5A, B). The results showed that E4-liposomes delivered TP3 to the implanted HeLa-K cells (Fig. 1.5D), and E4-Lipo-DOX could suppress cancer proliferation in the xenograft when compared to nontargeted conditions. These data demonstrated that coiled-coil formation enables drug selectivity and efficacy in vivo.
Fig. 1.5 Drug Delivery by E4/K4 Coiled-Coil Formation in Cells (A) and Zebrafish (B). (C) Schematic representation of coil structure between peptides E and K. (D) E4/K4 coiled-coil formation allows delivering the content in the liposome to cancer cells in the xenograft zebrafish.58 (This figure was reproduced with permission from reference58)
1.3.1.2 Controlled release
Conventional liposomes (Fig. 1.6A) are easily recognized by the mononuclear phagocyte system and are rapidly cleared from the blood stream.59 Many methods have been suggested to achieve long circulation of
18
polymers to delay the elimination process, such as coating the surface of liposomes with biocompatible polymers like poly(ethylene glycol) (PEG) linked phospholipids. These can be incorporated into the liposomal bilayer to form a hydrophilic polymer shield over the liposome surface, protecting the liposome from penetration or disintegration by plasma proteins60-65
(Fig. 1.6B). While many varieties have been synthesized by using chemically modified forms of PEG, in some cases it’s necessary to make the liposomes shed their cloak of modified PEG molecules when they reach their target (Fig. 1.6C). In this way they can interact with the target and release their payload. Using imaging technologies, visual evidence of the effect of PEGylation on the circulation kinetics of the liposomes was provided (Fig. 1.6D).66 The images clearly demonstrate that PEGylation
significantly enhances the persistence of liposomes in the blood stream. At the same time, the uptake of PEGylated liposomes in organs (liver and spleen) responsible for particle clearance decreased.
Fig. 1.6 Schematic representation of (A) conventional liposome, (B) PEG-liposome and (C) chemically modified PEG-liposome. (D) The effect of PEGylation on the circulation persistence of liposomes. The liposomes were labeled with Tc-99m, administered in rats, and the rats were imaged with a gamma camera over 24 h. As is evident from the heart (H) image signal, the PEG-liposomes remained in circulation even 24 h post-injection. The accumulation in the liver (L) and the spleen (S) was also lower in the case of PEG-liposomes, as compared to the plain liposomes.66 (Fig. 1.6 D was adapted from reference66)
Functionalization of Lipid Bilayer Membranes
19 Ligands conjugated with hydrophobic molecules form amphiphiles. The hydrophobic part can insert into the liposome bilayer, exposing the ligand outside of the liposomes for being recognized or for other interactions. For instance, DNA-b-polypropyleneoxide (DNA-ppo) has proven to be stably anchored into the lipid membrane for over at least 24 h. In this way, the containers are encoded with sequence information. The DNA-ppo present on the surface was used for anchoring a photosensitizer by hybridization. Upon light irradiation the PPO was oxidized leading to cargo release (Fig.
1.7).67
Fig. 1.7 Illustration of selective cargo release from DNA block copolymer (DBC) -decorated phospholipid vesicles. (1) DNA-ppo is stably anchored in unilamellar lipid vesicles; (2) DBC-decorated vesicles are functionalized with conjugated DNA-photosensitizers by hybridization; (3) singlet oxygen is generated by light irradiation; and (4) selective cargo release is induced by the oxidative effect of singlet oxygen.67 (This figure was reproduced with permission from
reference 67)
1.3.2 Stimuli-responsive liposomes
Liposomes can suspend cargos with their peculiar solubility properties and act as a sustained-release system for microencapsulated molecules. After modification, liposomes can be used as stimuli-responsive nanoparticles, which are visionary concepts to deliver and release a drug exactly where it is needed.68,69 There are several ways to trigger cargo release, such as
20
to be combined to appropriately improve the cargo release kinetics and distribution to reduce side effects.
1.3.2.1 Light responsive vesicle systems
Methods of sensitizing liposomes to light have progressed from the use of organic molecule moieties to the use of metallic plasmon resonant structures which can be broadly categorized as photochemical or photophysical release. Photochemical release can be achieved via photoisomerization, photocleavage and photopolymerization, which all lead to destabilization of the liposome bilayer and release of encapsulated contents (Fig. 1.8A-C).
Fig. 1.8 Release from liposomes mediated by photochemical responses: photoisomerization (A), photocleavage (B), or photopolymerization (C); and photophysical responses: molecular absorbers (D) and gold nanoparticles (E).
Functionalization of Lipid Bilayer Membranes
21 On the other hand, photophysical release from liposomes does not rely on any chemical changes of structures within or associated with the bilayer membrane. Examples of photophysical release discussed here take advantage of photothermal conversion of absorbed light with ensuing thermal and/or mechanical processes in the lipid membrane and the surrounding medium. The methods for achieving photophysical release are developed around molecular absorbers (Fig. 1.8D) or gold nanoparticles (Fig. 1.8E).70
1.3.2.2 Temperature responsive vesicle systems
Temperature-responsive liposomes are classified into two types: traditional temperature-responsive liposomes and liposomes modified with responsive polymers. Traditional temperature-responsive liposomes which are composed of temperature-temperature-responsive lipids show the greatest permeation of the lipid membrane at its gel-to-liquid crystalline phase transition temperature.
Moreover, liposomes modified with temperature-responsive polymers exhibit a lower critical solution temperature (LCST) behavior. These polymers are soluble in an aqueous solution below this temperature but dehydrate and aggregate if heated above the LCST. This behavior induces the release of a drug within a polymer-modified liposome. For instance, a temperature-responsive polymer, poly (N-isopropylacrylamide)-co-N,N'-dimethylaminopropylacrylamide (P(NIPAAm-co-DMAPAAm)) was synthesized and used for liposome modification. This research showed that the polymer underwent dehydration and aggregation above 40 °C and that temperature-responsive polymer-modified liposomes had faster cellular uptake and release compared to non-modified liposomes (Fig. 1.9).74
22
Fig. 1.9 Liposomes modified with temperature-responsive polymers are used for cellular uptake. The copolymer displayed a thermosensitive transition at a lower critical solution temperature (LCST) that is higher than body temperature. Above the LCST, the temperature-responsive liposomes started to aggregate and release their content. The liposomes showed a fixed aqueous layer thickness (FALT) at the surface below the LCST, and the FALT decreased with increasing temperature. Above 37°C, cytosolic release from the temperature-responsive liposomes was higher than that from the PEGylated liposomes, indicating intracellular uptake.74 (This figure was adapted from reference 74)
1.3.2.3 Magnetic responsive vesicle systems
Magnetoliposomes are composed of a lipid bilayer surrounding superparamagnetic iron oxide nanoparticles. Due to the biocompatibility, size, material-dependent physicochemical properties and potential applications as alternative contrast enhancing agents for magnetic resonance imaging, magnetoliposomes are ideal candidates to achieve a spatial and temporal control over drug release.75,76 Superparamagnetic iron
oxide nanoparticles (SPION) can be guided to their site of action using an externally applied magnetic field. The subsequent accumulation of SPION in the target site can be exploited for simultaneous drug delivery, MR imaging or hyperthermia therapy of cancer (Fig. 1.10).
Functionalization of Lipid Bilayer Membranes
23 Fig. 1.10 Superparamagnetic iron oxide nanoparticles can be guided to the site of action using an externally applied magnetic field.77 (This figure was adapted from reference 77)
In the beginning, liposomes were studied only for their physicochemical properties as models of membrane morphology. Today, they are used as delivery devices to encapsulate cosmetics, drugs, fluorescent detection reagents, and as vehicles to transport nucleic acids, peptides, and proteins to specific cellular sites in vivo. Advances in therapeutic applications of liposomes have been achieved through surface modifications. With these surface modifications, their biological stability could be increased, which includes reduced constituent exchange and leakage as well as reduced unwanted uptake by cells of the mononuclear phagocytic system.78
Targeting components such as antibodies can be attached to liposomal surfaces and were used to create large antigen-specific complexes. In this sense, liposomal derivatives are being used to target cancer cells in vivo, to enhance detectability in immunoassay systems.
24
1.4 Motivation and Thesis Overview
The overall goal of the work described in this thesis was to use DNA nanotechnology as a tool to manipulate lipid bilayer surfaces. Our group synthesized and characterized a new family of DNA amphiphiles containing modified nucleobases. The modification is introduced in uracil and consists of hydrophobic moieties. Through solid phase synthesis, the modified nucleotides can be incorporated in any desired position and several modifications per DNA strands can be introduced.79 The resulting DNA
sequences still undergo specific Watson-Crick base pairing. This property combined with the amphiphilic nature of this lipid-DNA qualifies the material as appealing candidate to interact with and manipulate biological membrane structures.
In chapter 2, a powerful new approach was introduced by modifying DNA with lipid chains at four nucleobases to tightly anchor the nucleotide to the lipid membrane. This strategy allows highly stable incorporation of DNA into the liposomal bilayer, thereby limiting dissociation. Several assays were employed proving the incorporation and stable anchoring in the phospholipid bilayer. These measurements involve small vesicles and fluorescence energy transfer. These experiments allow to measure how long the DNA amphiphiles remain in the bilayer.
In chapter 3, efficient fusion of liposomes was studied using lipid-DNA introduced in the chapter before. While the orientation of DNA hybridization played a significant role in the efficacy of full fusion of DNA-grafted vesicles, the number of anchoring units was found to be a crucial factor as well. As compared to vesicles functionalized with single-anchored or double-anchored DNA, liposomes containing quadruple-anchored oligonucleotides were found to be highly fusogenic, achieving considerable full fusion of up to 29% without notable leakage. This study demonstrates the importance of the DNA-anchoring strategy in hybridization-induced vesicle fusion, as not only the structural properties of the unit itself, but also the number of anchoring units determines its favorable fusion-inducing properties. Several fluorescence assays, dynamic light scattering and cryogenic transmission electron microscopy were utilized to prove these results.
Functionalization of Lipid Bilayer Membranes
25 In chapter 4, we expand the functionality of DNA encoded vesicles significantly. It was demonstrated that strand replacement can be carried out. In this chapter it will be outlined what sequences and what DNA amphiphiles are needed to reach this goal, i.e. changing the surface functionalities of liposomes by the simple addition of oligonucleotides. Moreover, it will be detailed how such a surface modification can be amplified by a simple DNA-triggered supramolecular polymerization. In chapter 5, we investigated whether it is possible to insert the lipid-modified DNA sequences into the membrane of live zebrafish to function as artificial receptor. We demonstrate that oligonucleotides functionalized with a membrane anchor can be immobilized on a zebrafish. Protruding single-stranded DNA atop the fish was functionalized by Watson-Crick base pairing employing complementary DNA sequences. In this way, small molecules and liposomes were guided and attached to the fish surface. The anchoring process can be designed to be reversible allowing exchange of surface functionalities by simple addition of DNA sequences. To achieve this on a fish surface, the strand exchange experiments established in
chapter 4 on simple vesicles as model were crucial. Finally, a DNA based
amplification process was performed atop of the zebrafish enabling the multiplication of surface functionalities from a single DNA anchoring unit.
26
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42. Ma, M.; Bong, D.; Controlled Fusion of Synthetic Lipid Membrane Vesicles. Acc Chem Res.
2013, 46, 2988-2997.
43. Kumar, P.; Guha, S.; Diederichsen, U.; SNARE protein analog-mediated membrane fusion. J. Pept. Sci. 2015, 21, 621-629.
44. Kong, L.; Askes, S. H. C.; Bonnet, S.; Kros, A.; Campbell, F.; Temporal Control of Membrane Fusion through Photolabile PEGylation of Liposome Membranes. Angew. Chem. Int. Ed. 2016, 55, 1396-1400.
45. Chernomordik, L. V.; Kozlov, M. M.; Protein-lipid interplay in fusion and fission of biological membranes. Annu. Rev. Biochem. 2003, 72, 175-207.
46. Cohen, F.S.; Melikyan, G. B.; The energetics of membrane fusion from binding, through hemifusion, pore formation and pore enlargement. J. Membr. Biol. 2004, 199, 1-14.
47. Weber, T.; Zemelman, B. V.; Mcnew, J. A.; Westermann, B.; Gmachl, M.; Parlati, F.; Söllner, T. H.; Rothman, J. E.; SNAREpins: minimal machinery for membrane fusion. Cell 1998, 92, 759-772.
48. Jahn, R.; Scheller, R. H.; SNAREs–engines for membrane fusion. Nat. Rev. Mol. Cell Biol.
2006, 7, 631-643.
49. Hong, W. J.; Lev, S.; Tethering the assembly of SNARE complexes. Trends Cell Biol. 2014, 24, 35-43.
50. Kumar, P.; Guha, S.; Diederichsen, U.; SNARE protein analog-mediated membrane fusion. J. Pept. Sci. 2015, 21, 621-629.
Functionalization of Lipid Bilayer Membranes
29 51. Stengel, G.; Zahn, R.; Höök, F.; DNA-induced programmablefusionof phospholipidvesicles. J. Am. Chem. Soc. 2007, 129, 9584-9585.
52. Chan, Y. H. M.; van Lengerich, B.; Boxer, S. G.; Lipid-anchored DNAmediates vesicle fusion as observed by lipid content mixing. Biointerphases 2008, 3,17-21.
53. Chan, Y. H. M.; van Lengerich, B.; Boxer, S. G.; Effects of linker sequences on vesicle fusion mediated by lipid-anchored DNA oligonucleotide. Proc. Natl. Acad. Sci. U.S.A. 2009, 106, 979-984.
54. Zheng, T.; Voskuhl, J.; Versluis, F.; Zope, H. R.; Tomatsu, I.; Marsden, H. R.; Kros, A. Controlling The Rate of Coiled Coil Driven Membrane Fusion. Chem. Commun. 2013, 49, 3649-3651.
55. Kong, L.; Askes, S. H.; Bonnet, S.; Kros, A.; Campbell, F. Temporal Control of Membrane Fusion through Photolabile PEGylation of Liposome Membranes. Angew. Chem. Int. Ed. 2016, 55, 1396-1400.
56. Mukai, M.; Sasaki, Y.; Kikuchi, J.; Fusion-Triggered Switching of Enzymatic Activity on an Artificial Cell Membrane. Sensors 2012, 12, 5966-5977.
57. Su, W.; Luo, Y.; Yan, Q.; Wu, S.; Han, K.; Zhang, Q.; Gu, Y.; Li, Y.; Photoinduced Fusion of Micro-Vesicles Self-Assembled from Azobenzene-Containing Amphiphilic Diblock Copolymers. Macromol. Rapid Commun. 2007, 28, 1251-1256.
58. Jian Yang, Yasuhito Shimada, René C. L. Olsthoorn, B. Ewa Snaar-Jagalska, Herman P. Spaink, and Alexander Kros, Application of Coiled Coil Peptides in Liposomal Anticancer Drug Delivery Using a Zebrafish Xenograft Model. ACS Nano. 2016, 10, 7428-7435.
59. Nag, O. K.; Awasthi, V.; Surface Engineering of Liposomes for Stealth Behavior. Pharmaceutics 2013, 5, 542-569.
60. Papahadjopoulos, D.; Allen, T. M.; Gabizon, A.; Sterically stabilized liposomes-improvements in pharmacokinetics and antitumor therapeutic efficacy. Proc. Natl. Acad. Sci. U.S.A. 1991, 88, 11460-11464.
61. Pashin, Y. V.; Bakhitova, L. M.; Bentkhen, T. I.; Antimutagenic activity of simple phenols and its dependence on the number of hydroxyl groups. Bull Exp Biol Med. 1986, 102, 1121-1123.
62. Woodle, M. C.; Lasic, D. D.; Sterically stabilized liposomes. Biochim. Biophys. Acta 1992, 1113, 171-199.
63. Woodle, M. C.; Newman, M. S.; Cohen, J. A.; Sterically stabilized liposomes: physical and biological properties. J Drug Target 1994, 2, 397-403.
64. Woodle, M. C.; Newman, M. S.; Collins, L. R.; Efficient evaluation of long circulating or stealth liposomes by studies of in vivo blood-circulation kinetics and final organ distribution in rats. Biophys J. 1990, 57, A261.
65. Çağdaş, M.; Sezer, A.D.; Bucak, S.; Liposomes as Potential Drug Carrier Systems for Drug Delivery. Application of Nanotechnology in Drug Delivery. 2014 Chapter 1.
66. Nag, O. K.; Yadav, V. R.; Hedrick, A.; Awasthi, V.; Post-modification of preformed liposomes with novel non-phospholipid poly(ethylene glycol)-conjugated hexadecylcarbam -oylmethyl hexadecanoic acid for enhanced circulation persistence in vivo. Int. J. Pharm.
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67. Rodríguez-Pulido, A.; Kondrachuk, A. I.; Prusty, D. K.; Gao, J.; Loi, M. A.; Herrmann, A.; Light-Triggered Sequence-Specific Cargo Release from DNA Block Copolymer–Lipid Vesicles. Angew. Chem. Int. Ed. 2013, 52, 1008-1012.
68. Chiu, H. C.; Lin, Y. W.; Huang, Y. F.; Chuang, C. K.; Chern, C. S.; Polymer Vesicles Containing Small Vesicles within Interior Aqueous Compartments and pH-Responsive Transmembrane Channels. Angew. Chem. Int. Ed. 2008, 47, 1875-1878.
69. Volodkin, D. V.; Skirtach, A. G.; Möhwald, H.; Near-IR Remote Release from Assemblies of Liposomes and Nanoparticles. Angew. Chem. Int. Ed. 2009, 48, 1807-1809.
70. Leung, S. J.; Romanowski, M.; Light-Activated Content Release from Liposomes. Theranostics 2012, 2, 1020-1036.
71. Li, L.; Hagen, ten T. L.M.; Hossann, M.; Süss, R.; Rhoon, van G. C.; Eggermont, A. M.M.; Haemmerich, D.; Koning, G. A.; Mild hyperthermia triggered doxorubicin release from optimized stealth thermosensitive liposomes improves intratumoral drug delivery and efficacy. J Control Release 2013, 168, 142-150.
72. Tai, L.; Tsai, P.; Wang, Y.; Wang, Y.; Lo, L.; Yang, C.; Thermosensitive liposomes entrapping iron oxide nanoparticles for controllable drugrelease. Nanotechnology 2009, 20, 1-9.
73. Nappini, S.; Bombelli, F. B.; Bonini, M.; Nord, B.; Baglioni. P.; Magnetoliposomes for controlled drug release in the presence of low-frequency magnetic field. Soft Matter 2010, 6, 154-162.
74. Wang, J.; Ayano, E.; Maitani, Y.; Kanazawa, H.; Tunable Surface Properties of Temperature-Responsive Polymer-Modified Liposomes Induce Faster Cellular Uptake, ACS Omega 2017, 2, 316-325.
75. Monnier, C. A.; Burnand, D.; Rutishauser, B. R.; Lattuada, M.; Petri-Fink, A.; Magnetoliposomes: opportunities and challenges. Eur J Nanomed. 2014, 6, 201-215. 76. Amstad, E.; Kohlbrecher, J.; Müller, E.; Schweizer, T.; Textor, M.; Reimhult, E.; Triggered Release from Liposomes through Magnetic Actuation of Iron Oxide Nanoparticle Containing Membranes. Nano Lett. 2011, 11, 1664-167.
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78. Woodle, M. C.; Surface-modified liposomes: assessment and characterization for increased stability and prolonged blood circulation. Chem. Phys. Lipids 1993, 64, 249-262. 79. Anaya, M.; Kwak, M.; Musser, A. J.; Müllen, K.; Herrmann, A.; Tunable Hydrophobicity in DNA Micelles: Design, Synthesis, and Characterization of a New Family of DNA Amphiphiles. Chem. Eur. J. 2010, 16, 12852-12859.
Chapter 2
Stability Study of Lipid-DNA on
the Liposomal Membrane
32
2.1 Introduction
Deoxyribonucleic acid (DNA) is a macro molecule that carries hereditary information of all known living organisms and many viruses. Its double-stranded helix structure was discovered by Watson & Crick in 1953,1 which
has greatly fueled many technologies dealing with DNA and hence revolutionized modern science. In recent years DNA has become a valuable functional building block and tool in nanotechnology and material science due to the unique nature and properties of DNA and DNA hybrid materials. A wide variety of products and applications have been realized using DNA technologies among which is incorporating DNA with a functional group and utilizing its information-carrying capability to develop DNA detection systems. For instance, fluorescent dye-labeled DNA was used as probe monitor in PCR2 or for sequence analysis.3,4 Additionally, coupling DNA
strands with moieties like polymers or nanoparticles changes the morphological structure and introduces new functionalities, which are
Stability Study of Lipid-DNA on the Liposomal Membrane
33 different from conventional polymers. For instance, DNA conjugated gold nanoparticles were used in DNA microarray technology.5-7 Another
functional moiety chemically conjugated with DNA consists of hydrophobic molecules, such as long alkyl chains, cholesterol, or fatty acids, resulting in amphiphiles, which spontaneously form nanoparticles in solution and enhance the pharmacokinetic behavior and trans-membrane delivery. Their amphiphilic nature arises from the hydrophilic DNA backbone containing charged phosphodiester bonds and the hydrophobicity of attached alkyl chains.8 These nanoparticles can be further functionalized
through hybridization of a modified complimentary DNA or internalization of payloads in the hydrophobic core.9
Our group reported the synthesis and characterization of a family of DNA amphiphiles containing hydrophobically modified nucleobases.10,11
Specifically, 1-dodecyne (C12H22) was attached to a uracil base which was
further attached to the 5’ or 3’ position of a DNA sequence (Fig. 2.1A). In aqueous environment, due to their amphiphilic nature, lipid-DNA self-assembles into micelles whereby the hydrophilic DNA strands shield the hydrophobic lipid core. These DNA micelles can be loaded with cargo by hydrophobic interactions or hybridization with functionalized complementary DNA (Fig. 2.1B). The aggregation properties of lipid-DNA can be relatively easy manipulated by changing the length of the lipid part or the number and position of the modified uracil bases within the DNA sequence. Fig. 2.1C shows three different lipid-DNAs. U2M and U2T are lipid DNA with two modified uracil bases either in the middle or at the terminus and U4T represents lipid DNA with four modified uracil bases at the 5’ end.10
Because of the amphiphilic and sequence specific properties, lipid-DNA can be used for liposome surface modification by insertion of the hydrophobic part into the membrane while the hydrophilic DNA is exposed to the aqueous medium. Compared with existing terminal modifications, our design allows the precise and easy introduction of hydrophobic units at arbitrary positions and numbers in a DNA sequence through conventional solid-phase synthesis. In this chapter, DNA was modified with four lipid
34
chain modified nucleobases at both terminals and it was used to anchor it to phospholipid membranes.
Fig. 2.1 Structure of lipid modified nucleotide and representation of lipid–DNAs. (A) Chemical structure of the lipid-modified uracil nucleobase. (B) Lipid-DNAs self-assemble to form DNA micelles due to their amphiphilic nature. These self-assembled structures can carry cargo by hydrophobic interaction (1) or by hybridization with a functionalized complementary DNA (2). (C) Schematic representation of the ss and ds lipid–DNA amphiphiles (U2M, U2T, and U4T) and their propensity to undergo Watson-Crick base pairing.10
Stability Study of Lipid-DNA on the Liposomal Membrane
35
2.2 Results and Discussion
2.2.1 Lipid-DNA design and characteristics
To obtain stable incorporation of DNA into the liposomal bilayer, we use lipid-DNA (U4T-18), which has been designed to contain four modified uracil nucleobases at the 5’ position of a 18-mer oligonucleotide (including the 4 lipid modified uracil bases). CrU4T-18 is complementary to U4T-18 with the lipid anchor at the opposite terminus (i.e. the 3’ position). Cr-ATTO488 is a 14-mer DNA complementary to U4T-18 and was covalently attached an ATTO488 dye to the 3’ end (Table 2.1).
Table 2.1 Sequences of modified DNA.
Name Sequence (5’→ 3’)*
U4T-18 UUUUGCGGATTCGTCTGC
CrU4T-18 GCAGACGAATCCGCUUUU
14mer GCGGATTCGTCTGC
Cr-ATTO488 GCAGACGAATCCGC-ATTO488
*: U represents the lipid-modified uracil base.
U4T-18 can be attached to the liposome surface by insertion of four lipid-modified nucleobases into the lipid membrane while the remaining 14mer DNA part is protruding into the aqueous medium. This DNA unit can hybridize with the DNA part from CrU4T-18 or Cr-ATTO488 (Fig. 2.2A). According to the results from polyacrylamide gel electrophoresis (PAGE), a lower electrophoretic mobility of hybridized lipid-DNA (lane 2) is observed compared to ssDNA controls (lane 1 and lane 3), indicating successful Watson-Crick base pairing (Fig. 2.2B).
After confirming hybridization, the melting temperature (Tm) of the
ds-lipid-DNA (U4T-18+Cr-ATTO488) was determined. The ds-ds-lipid-DNA and ds14mer (14mer+Cr-ATTO488) were heated at 0.5 °C/min while measuring the absorption at 260 nm. Afterwards the first derivative of the curve was calculated and Tm of the ds DNA was taken at maximum slope.
36
The Tm value of lipid-DNA (62.5 °C) is very close to that of 14mer (63.6 °C)
(Fig 2.2C, D). The result indicates that lipid chains have little influence on the melting temperature.
Fig. 2.2 (A) Schematic representation of U4T-18 hybridization with Cr-ATTO488 on the surface of liposomes. (B) Native PAGE characterization of lipid-DNA (20% TBE gel, 100V, 80min). Lane 1: U4T-18, lane 2: U4T-18 + Cr-ATTO488, lane 3: Cr-ATTO488. (C) Melting curve of dsDNA, U4T-18 + Cr-ATTO488. (D) Melting curve of dsDNA, 14mer + Cr-ATTO488. Melting curve (black squares, left Y-axis) and calculated
derivative
for corresponding sample (red circle, right Y-axis).2.2.2 Characterization of the incorporation of lipid-DNA in liposomal
bilayer.
After synthesis of the nucleobase-modified DNA hybrids and testing their ability for Waston-Crick base pairing, the lipid DNAs were stably anchored into the membrane of DOPC:DOPE:cholesterol lipid vesicles, while the oligonucleotides remained available for hybridization, as demonstrated by a Fluorescence Resonance Energy Transfer (FRET) assay.12
Stability Study of Lipid-DNA on the Liposomal Membrane
37 Since ATTO488 and rhodamine dyes show energy transfer when there is a sufficiently short distance between them,13 ATTO488 was covalently
attached to the 3’ end of a 14-mer DNA complementary to U4T-18 (Cr-ATTO488) to act as a donor. In parallel, rhodamine-functionalized phospholipid (Rh-DHPE) was incorporated in the liposomal bilayer to function as an acceptor (Fig. 2.3A). To observe the dynamic emission changes of donor and acceptor after adding Cr-ATTO488, fluorescence emission spectra with excitation at 470 nm of Cr-ATTO488 (donor, emission maximum 520 nm) and Rh-DHPE (acceptor, emission maximum 592 nm) were recorded over 30 min (Fig. 2.3B). The fluorescence of donor significantly decreased by adding Cr-ATTO488 and the fluorescence of acceptor slightly increased at the same time, illustrating that FRET is induced by DNA hybridization.
Fig. 2.3 (A) Schematic of FRET assay demonstrating that oligonucleotides anchored into liposomal bilayers via lipid-DNA remain available for hybridization. (B) Fluorescence emission of Cr-ATTO488 (donor, emission maximum 520 nm) and Rh-DHPE (acceptor, emission maximum 592 nm) followed over 30 min after adding Cr-ATTO488.
Meanwhile, as demonstrated by the increase in the maximum intensity ratio I592/I520 (acceptor/donor peak) (Fig. 2.4D), hybridization only
occurred upon mixing of Cr-ATTO488 with U4T-18-grafted Rh-DHPE-containing vesicles, positioning both dyes sufficiently close to each other to achieve FRET (Fig. 2.4A), whereas for vesicles containing non-complementary lipid-DNA, CrU4-18, (Fig. 2.4B) or no lipid-DNA at all (Fig.
38
Fig. 2.4 Anchoring of lipid-DNA in the membrane and hybridization on the vesicle surface leads to Fluorescence Resonance Energy Transfer (FRET) upon hybridization of donor-modified complementary DNA with DNA-functionalized, acceptor-containing vesicles. (A) FRET is achieved when complementary Cr-ATTO488 DNA hybridizes with U4T-18 and brings the donor close to the acceptor, rhodamine, positioned in the membrane. If hybridization is not possible, either due to mismatch of the two DNA strands (B) or the absence of membrane-grafted DNA (C) FRET does not occur. (D) Fluorescence spectra of systems capable of FRET (red) and non-FRET controls, either due to DNA mismatch (blue) or absence of membrane-grafted DNA (green).
Disruption of vesicles by addition of Triton X-100 to a final concentration of 0.3% (v/v) resulted in a drop in FRET in the U4T-18 vesicles hybridized with Cr-ATTO488 (Fig. 2.5A vs Fig 2.4D), confirming that FRET was indeed caused by bringing the donor in close vicinity to the acceptor dye located in the liposomal membrane. As expected, in two control non-FRET systems in which DNA hybridization could not occur, either due to absence of DNA in the membrane (Fig. 2.5B) or the presence of non-complementary DNA (Fig. 2.5C) energy transfer from donor to acceptor
Stability Study of Lipid-DNA on the Liposomal Membrane
39 was prevented. Therefore, similar spectra were observed before and after liposomal disruption.
Fig. 2.5 To further investigate the engraftment of the lipid-DNA hybrids into the membrane, FRET liposomes were disrupted with Triton X-100 at a final concentration of 0.3 % (v/v). Fluorescence spectra of FRET liposomes before and after adding Triton X-100 (A). Similar spectra were observed in control experiments before and after liposomal disruption, either due to the absence of DNA on the membrane (B) or the presence of non-complementary DNA (C).
2.2.3 Temporal stability of lipid-DNA in the liposomal membrane.
To study whether the incorporation of U4T-18 in the membrane is stable overtime, FRET (U4T-18/Cr-ATTO488/Rh-DHPE) liposomes were incubated with non-FRET (NF) liposomes (Fig. 2.6A) at different ratios (1:1, 1:5 and 1:10).
40
Fig. 2.6 Measurement of stability of lipid-DNA in liposomes over time. FRET (U4T-18/Cr-ATTO488/Rh-DHPE) liposomes were incubated with non-FRET (NF) liposomes (A) at different ratios (1:1, 1:5 and 1:10), and the relative Rh-DHPE/ATTO488 (IA/ID) emission intensity ratio
was monitored over 24 h after mixing (B). Fluorescence spectra of Cr-ATTO488/Rh-DHPE pair in FRET liposomes mixed with NF liposomes at different ratio (v/v): 1:1(red line), 1:5(blue line), 1:10(green line) (C). Solid and dashed lines represent the spectra of the mixed systems before and after adding Triton X-100, respectively.
The relative Rh-DHPE/ATTO488 (IA/ID) emission intensity ratio of the
three systems was monitored over 24 h after mixing (Fig. 2.6B). If lipid-DNA redistributes from FRET liposomes to NF liposomes, a decrease in relative fluorescence of acceptor peak would be observed. After 24 h, some of the acceptor intensity had dropped, but the relative fluorescence IA/ID of
the mixture remained at a similar value as that during the initial measurement before non-FRET liposomes were added. The results demonstrate that the lipid–DNA is stably anchored in the liposomes over at last 24 hours. Fig. 2.6C shows the fluorescence spectra of Cr-ATTO488/Rh-DHPE pair in FRET liposomes mixed with NF liposomes at different ratio before and after liposomal disruption.
Stability Study of Lipid-DNA on the Liposomal Membrane
41 Moreover, lipids were mixed with U4T-18 at different molar ratios (5000, 1000, 100, 62.5). The final concentration of Cr-ATTO488 and lipid-mixture (DOPC+DOPE) were kept at 7.32 µM and 0.45 mM, respectively, in all FRET experiments. The results show the I592/I520 ratio increased markedly with
higher U4T-18 densities in the membrane (Fig. 2.7, Table 2.2). These results demonstrate that when more lipid DNA is incorporated into the membrane more DNA strands can be attached to this vesicle surface by hybridization (Table 2.2).
Fig. 2.7 U4T-18/Rh-DHPE fluorescence spectra of FRET liposomes mixed with Cr-ATTO488 at different lipid/U4T-18 ratios. The inset shows a zoom-in of the acceptor Rh-DHPE peak. Solid lines and dashed lines represent the spectra of the FRET system before and after adding Triton X-100, respectively. Lipids were mixed with U4T-18 at different molar ratios (5000, 1000, 100, 62.5).
Table 2.2 The acceptor/donor fluorescence intensity ratios (I592/I520) at different lipid/U4T-18
ratios.
Lipid : U4T-18 ratio
U4T-18:liposome
ratio I592/I520 FRET system
5000 8 0.22
1000 38 0.24
100 380 0.31
42
2.3 Conclusion
In conclusion, we proposed a powerful new approach employing lipid-DNA which contains four lipid chains modified nucleobases to tightly anchor the nucleotide to the lipid membrane. The incorporation and stability of lipid-DNA on the liposomal membrane were proved by FRET. FRET was achieved when the hybridization occurred between Cr-ATTO488 and U4T-18, which brought the donor (Cr-ATTO488) close to the acceptor (rhodamine) that was positioned in a U4T-18 functionalized membrane. Meanwhile, the I592/I520 (acceptor/donor peak) ratio increased markedly
with higher U4T-18 densities in the membrane, and disruption of vesicles by addition of Triton X-100 resulted in a drop of FRET vesicles system, confirming that FRET was indeed caused by bringing the donor in close vicinity to the acceptor dye located in the liposomal membrane. Finally, the lipid–DNA remained stably anchored in the liposomes for at least 24 hours.
2.4 Experimental Section
2.4.1 Materials
Cholesterol (Chol), 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE) and 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) were purchased from Avanti Polar Lipids (Alabaster, USA) (Fig 2.8A-C, purity >99%) and used without further purification. Headgroup-labeled phospholipid, Lissamine rhodamine B 1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine (triethylammonium salt) (Rh-DHPE) was purchased from Invitrogen (Amsterdam, Netherlands), and used as received (Fig.
2.8D). The DNA-dye conjugate Cr-ATTO488 was purchased from
Biomers.net GmbH (Ulm, Germany). Trition X-100 (10% in water), and Tris/HCl buffer were purchased from Sigma-Aldrich (St. Louis, United States). Anhydrous CHCl3 was purchased from Acros Organics (Geel,
Belgium) and stored over molecular sieves. For all experiments, ultrapure water (specific resistance > 18.4 MΩ cm) was obtained by a Milli-Q water purification system (Sartorius).
Stability Study of Lipid-DNA on the Liposomal Membrane
43 Fig. 2.8 Structures of lipids: (A) DOPC, (B) DOPE, (C) Cholesterol; fluorescent lipids: (D) Rh-DHPE.
2.4.2 Synthesis and characterization of amphiphilic oligonucleotides
The synthesis of 5-(dode-1-cynyl) deoxyuracil and 5-(dode-1-cynyl) deoxyuracil phosphoramidite were reported previously (Fig. 2.9).10,11 In
short, the modified uracil phosphoramidite was dissolved in CH3CN to a
final concentration of 0.15 M in the presence of 3 Å molecular sieves and the prepared solution was directly connected to a DNA synthesizer (ÄKTA oligopilot plus, GE Healthcare (Uppsala, Sweden)). Oligonucleotides were synthesized on a 10 μmol scale using standard β-cyanoethylphosphoramidi -te coupling chemistry. Deprotection and cleavage from the PS support was carried out by incubation in concentrated aqueous ammonium hydroxide solution for 5 h at 55 °C. Following deprotection, the oligonucleotides were purified by using reverse-phase chromatography, using a C15 RESOURCE RPCTM 3 mL reverse phase column (GE Healthcare) through a custom gradient elution (A: 100 mM triethylammonium acetate (TEAAc) and 2.5% acetonitrile, B: 100 mM TEAAc and 65% acetonitrile). Fractions were
44
desalted using centrifugal dialysis membranes (MWCO 3000, Sartorius Stedim). Oligonucleotide concentrations were determined by UV absorbance using extinction coefficients. Finally, the identity and purity of the oligonucleotides was confirmed by RPC-HPLC (Fig. 2.10) and MALDI-TOF mass spectrometry (Fig. 2.11).
Fig. 2.9 Synthesis of 5-(dode-1-cynyl) deoxyuracil 2 and 5-(dode-1-cynyl) deoxyuracil phosphoramidite 3.
Stability Study of Lipid-DNA on the Liposomal Membrane
45 Fig. 2.11 RPC HPLC analysis of purified lipid-DNAs: (A) U4T-18, (B) CU4T-18 and (C) CrU4T-18. Numbers beside the elution peaks represent the buffer B contents when lipid-DNAs were eluted.
2.4.3 Preparation and characterization lipid-DNA liposomes
An appropriate amount of freeze-dried lipid-DNA was mixed with DOPC:DOPE:Cholesterol (50:25:25 mol% in chloroform), to obtain the required lipid:lipid-DNA ratio. Afterwards, chloroform was removed by evaporation under an air stream and then under vacuum overnight. An aqueous buffer (100 mM NaCl, 20 mM Tris, pH 7.5) was added to the flask and the solution was vortexed and freeze-thawed 5 times. Subsequently, the dispersion was extruded 21 times, using an extruder and 100 nm polycarbonate membranes (Whatman), to obtain unilamellar vesicles. After extrusion, external buffers of each sample were removed by size exclusion chromatography. The column was filled with Sephadex G-75 (GE Healthcare Life Sciences) and equilibrated with buffer (100 mM NaCl, 20 mM Tris, pH 7.5). Lipid-DNA liposomes were used within one day. All liposomal formulations had an average diameter of around 130 nm as determined by DLS (ALV/CGS-3 ALV-Laser Vertriebsgesellschaft mbH, Langen, Germany). The ratio between lipid and U4T-18 was 500:1, unless stated otherwise.
46
2.4.4 Calculation of lipid-DNA/liposome ratio.
The amount of lipid-DNAs per liposome was calculated using the equation:
where Φ is the number of lipids per liposome which can be calculated from geometrical considerations:
where Souter and Sinner are the outer and inner surface area of the spherical
liposomes. Assuming the thickness of the lipid bilayer is 5 nm.14,15 α is the
average cross-sectional area of the lipid headgroups, which is assumed to be (2*80+65)/3=75 Å for DOPC:DOPE(2:1 molar ratio).16 Router is the
averaged radius of spherical liposomes, which was determined by DLS.
2.4.5 Characterization of lipid-DNA incorporation in liposomes
measured by Fluorescence Resonance Energy Transfer (FRET) assay
Fluorescence emission spectra of Cr-ATTO488 (donor) and Rh-DHPE (acceptor) in the 500–700 nm region were recorded with excitation at 470 nm using a SPECTRAMAX M2 (Molecular Devices) fluorescence spectrophotometer. Measurements were carried out at constant temperature of 25.0 °C, using a 100 mM NaCl, 20 mM Tris, pH 7.5 buffer.
2.4.6 FRET assay via DNA hybridization
U4T-18 was incorporated in Rh-DHPE/(DOPC+DOPE) (3:97 molar ratio) liposomes to obtain U4T-18 liposomes with a lipid to U4T-18 ratio of 500:1. Subsequently, an aliquot of these liposomes was mixed with a small amount of Cr-ATTO488 such that [U4T-18] = [Cr-ATTO488] = 0.906 μM and with a final lipid (DOPC+DOPE) concentration of 0.45 mM. Then, U4T-18 and Cr-ATTO488 were hybridized using an Eppendorf Mastercycler (Germany). The protocol consisted of heating the mixture 15 min to 40 °C
Stability Study of Lipid-DNA on the Liposomal Membrane
47 and slowly cooling to 4 °C over a period of 140 min. Afterwards, the emission spectra of Cr-ATTO488/Rh-DHPE pair were measured.
Author contributions
Meng Z designed and conducted the experiments, performed data analysis and wrote the manuscript. Liu Q and de Vries JW synthesized lipid-DNA. Herrmann A supervised the project.
48
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