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Spectroscopic studies on interactions between cholesterol-end capped polyethylene glycol and liposome
Colloids and Surfaces B: Biointerfaces 97 (2012) 248–253
Contents lists available at SciVerse ScienceDirect
Colloids and Surfaces B: Biointerfaces journal homepage: www.elsevier.com/locate/colsurfb
Spectroscopic studies on interactions between cholesterol-end capped polyethylene glycol and liposome Zhi Rao a,b , Tetsushi Taguchi a,b,∗ a b
Graduate School of Pure and Applied Sciences, University of Tsukuba, 1-1-1 Tennodai, Tsukuba, Ibaraki 305-8577, Japan Biomaterials Unit, Nano-bio field, Materials Nanoarchitectonics (MANA), National Institute for Materials Science, 1-1 Namiki, Tsukuba, Ibaraki 305-0044, Japan
a r t i c l e
i n f o
Article history: Received 12 January 2012 Received in revised form 26 March 2012 Accepted 27 March 2012 Available online 3 April 2012 Keywords: Liposome Cholesterol end group Nile Red Pinacyanol chloride Bilayer microenvironment
a b s t r a c t In order to confirm that the cholesterol end groups of cholesterol-end capped polyethylene glycol really insert into the liposome bilayer and investigate how the incorporation affects the microenvironment of liposome bilayer, two kinds of molecular probes, namely Nile Red and pinacyanol chloride, were used. Their UV–visible and fluorescence spectrum were recorded before and after the addition of the polymer. Shifts of the maximum absorbance (max ) of Nile Red show that the bilayer microenvironment around Nile Red is becoming more polar with increasing polymer concentration while shifts of max of pinacyanol chloride indicate that the surrounding environment of pinacyanol chloride is becoming more apolar with addition of polymer. Effect of composition of liposome was also studied. With high ratio of dimethyldioctadecylammonium bromide (DODAB) fraction in liposome, max of Nile Red is more easily affected by the addition of Chol-PEG-Chol while liposome with cholesterol shows relatively high stability to the addition of Chol-PEG-Chol. © 2012 Elsevier B.V. All rights reserved.
1. Introduction Spherical bilayer lipid vesicles (liposome) have been used extensively for drug delivery because of their ability to solubilize both hydrophilic and hydrophobic substances in their inner aqueous phase and liposome bilayer. Drug encapsulation by liposome allows in vivo protection from enzymes and thus enhances circulation time and bioavailability [1–3]. However, a major problem with liposome is still their insufficient stability, which may limit the practical application. Nevertheless, the stability can be enhanced by the addition of surfactants or amphiphilic polymer [4–8]. Studies on liposome–amphiphilic polymer systems have become an important research area of colloid science, from both experimental and theoretical view points [9–11]. The liposome–polymer systems constitute good model system for living cells [12–14], and they also lead to important phase behavior phenomena, such as the formation of gels and networks [15–20], with potential industrial applications. Among the studies on interactions between amphiphilic polymer and liposome, some seek to identify polymers that, upon binding to liposomes, markedly affect the shape, curvature, stiffness, or stability of the bilayer [21–26], while others
∗ Corresponding author at: Biomaterials Unit, Nano-bio field, Materials Nanoarchitectonics (MANA), National Institute for Materials Science, 1-1 Namiki, Tsukuba, Ibaraki 305-0044, Japan. Tel.: +81 29 860 4498; fax: +81 29 860 4714. E-mail address: [email protected] (T. Taguchi). 0927-7765/$ – see front matter © 2012 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.colsurfb.2012.03.017
often aim at improving the stability and at controlling the permeability of liposomes for drug delivery or targeting, gene therapy, and so forth [27,28]. However, even for the simple lipid membrane system, there are still disputations over whether the hydrophobic groups really anchor into the lipid bilayer or not. Some previous studies indicated that hydrophobic groups directly penetrated into the liposome bilayer and the insertion profoundly altered the morphology or fluidity or of the liposome bilayer [10,29], while some others showed that the polymers were only present at the surface of the liposomes and do not interfere with the lipid bilayer microenvironment [30]. Various techniques have been used for the study of the liposome bilayer microenvironment. The use of molecular probes has proven particularly advantageous in providing information about the bilayer structure [30–34]. These molecules are either solubilized in the bilayer at optimum concentration or are covalently bonded to the phospholipids forming the bilayer. Investigations of the fluorescence and UV–visible spectrum of such molecular probes have provided valuable information about the electric, dynamic, conformational, and structural properties of the liposome bilayer. It is well known that the microenvironment surrounding the molecular probe influences its electronic structure and thus its photophysics. Changes in the local microenvironment can produce measurable spectrum shifts which can, in turn, be monitored spectroscopically. This property, known as solvatochromism, allows elucidation of the influence of the immediate environment of the molecules within the probed
Z. Rao, T. Taguchi / Colloids and Surfaces B: Biointerfaces 97 (2012) 248–253
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systems, and moreover, gives evidence of specific interactions [33]. We have previously reported on the self-assembled liposome gel from liposome and cholesterol-end capped polyethylene glycol (Chol-PEG-Chol) and carried out systematic studies on the gel mechanical properties by rheological method [20]. However, whether cholesterol end groups of Chol-PEG-Chol really insert into the liposome bilayer and how the incorporation affects the microenvironment of liposome bilayer is of great research interest and by far, no studies have yet been carried out on this aspect. In this work, we used two kinds of molecular probes, Nile Red and pinacyanol chloride (PIN) to investigate the interactions between Chol-PEG-Chol and liposome. By monitoring the changes in the UV–visible absorption and fluorescence emission spectrum of the molecular probes, information was obtained about their local microenvironment. The results may provide some insights on the interactions between the amphiphilic polymer Chol-PEG-Chol and liposome bilayer. 2. Materials and methods 2.1. Materials Dimyristoylphosphatidylcholine (DMPC) was purchased from NOF Corporation (Tokyo, Japan). Dimethyldioctadecylammonium bromide (DODAB, >98%) was purchased from Sigma–Aldrich. Cholesterol was purchased from Wako Pure Chemical Industrials Ltd. (Osaka, Japan). Diamino-polyethylene glycol (molecular weight: 30,000, >98%) was purchased from NOF Corporation (Tokyo, Japan). Cholesteryl chloroformate (>99%) was purchased from Fluka (Buchs, Switzerland). Dichloromethane was purchased from Nacalai Tesque, Inc. (Kyoto, Japan). Triethylamine was purchased from Wako Pure Chemical Industrials Ltd. (Osaka, Japan). Nile Red and pinacynol chloride (PIN) were purchased from Tokyo Chemical Industry Co., Ltd. (Tokyo Japan). All chemicals were used without further purification.
Fig. 1. Relation between solvent polarity and max of Nile Red in UV–visible and fluorescence emission spectrum in different organic solvent.
added to liposome solution in 5 mL tube, the mixture was shaken gently at room temperature (25 ◦ C) on a micromixer overnight until Chol-PEG-Chol fully dissolved. 2.5. UV–visible measurements UV–visible measurements were carried out using JASCO V660 UV–visible spectrophotometer (Tokyo, Japan), recording
2.2. Preparation of liposome solution In this work, four kinds of liposome of different composition were prepared. Liposome 1 (DMPC:DODAB molar ratio = 9:1); liposome 2 (DMPC:DODAB molar ratio = 1:1); liposome 3 (DMPC:DODAB:cholesterol molar ratio = 8:1:1); liposome 4 (DMPC:DODAB:cholesterol molar ratio = 6:1:3). The liposome was prepared by extrusion method [20]. The final liposome concentration was 1 mM. According to the dynamic light scattering measurement, the average liposome particle size was 109 nm, 193 nm, 111 nm, 129 nm respectively. Nile Red was introduced to the final liposome solution by injection of 0.3% (v/v) stock solution (0.25 mM) of the dye in ethanol. PIN was introduced to the final liposome solution by injection of 1% (v/v) of stock solution (1 mM) of the dye in DMSO. 2.3. Preparation of Chol-PEG-Chol Chol-PEG-Chol was prepared from diamino-polyethylene glycol and cholesteryl chloroformate following the procedure of Ref. [20]. 2.4. Preparation of mixture solution of Chol-PEG-Chol and liposome In this paper, Chol-PEG-Chol concentration varied from 0 to 5 mg/mL and liposome concentration was fixed at 1 mM. Based on our preliminary experiments, in this concentration range, the viscosity of the mixture of liposome and Chol-PEG-Chol was not too high to interfere with the measurement. After Chol-PEG-Chol was
Fig. 2. Relation between solvent polarity and max of bands I and II of PIN in UV–visible spectrum in different solvent.
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Fig. 3. (a) Effect of Chol-PEG-Chol concentration on max of Nile Red-loaded liposome in UV–visible spectrum at 25 ◦ C. (b) Schematic illustration of Nile Red local microenvironment change before and after the addition of Chol-PEG-Chol.
the absorbance spectrum between 400 and 700 nm wavelength directly. 2.6. Fluorescence measurements Fluorescence measurements were taken using JASCO FP-6500 fluorescent spectrometer (Tokyo, Japan). Excitation and emission slits were both fixed at 5 nm, and exc was 550 nm. 3. Results and discussion 3.1. UV–visible and fluorescence spectrum of molecular probes with relation to the solvent polarity Nile Red is an uncharged benzeophenoxazone dye, well soluble in organic solvents and lipids, but almost unsoluble and strongly quenches in water. It is photochemically stable and can be monitored by both UV–visible adsorption spectrum and fluorescence
spectrum [30]. Nile Red has been used as a fluorescent probe for intracellular lipids and as a polarity sensor deducing protein structure and conformation [35–37]. Being highly hydrophobic, both the UV–visible and fluorescence spectrum of Nile Red in water shows only flat line with no absorbance or intensity (supplementary materials, Fig. S1). However, when Nile Red was added to organic solvents, which have polarity lower than water, both the UV–visible and fluorescence spectrum of Nile Red in organic solvents show single broad peak (supplementary materials, Fig. S1). Fig. 1 shows the relation of the maximum wavelength (max ) of Nile Red UV–visible adsorption spectrum and fluorescence spectrum with the polarity of the organic solvent. It is obvious that max of both UV–visible adsorption spectrum and fluorescence spectrum increase with the increase of polarity of the organic solvent. This finding correlates well with previous work in which Nile Red max shifted to higher wavelength as the polarity of methanol/chloroform mixtures was increased [30].
Z. Rao, T. Taguchi / Colloids and Surfaces B: Biointerfaces 97 (2012) 248–253
When Nile Red is introduced into liposome solution, both UV–visible and fluorescence spectrum of liposome loaded with Nile Red show single broad peak, similar to that of Nile Red in organic solvent, as shown in Fig. S2 of supplementary materials. This shows that the hydrophobic dye molecules are easily solubilized inside the liposome bilayer due to the apolar microenvironment inside bilayer. With the increase of volume percentage of addition of Nile Red to liposome 1, intensity of both UV–visible adsorption spectrum and fluorescence spectrum increased accordingly and no obvious shift of max was observed, indicating that the polarity of Nile Red vicinity microenvironment remain unchanged in this concentration range of Nile Red. In this study, we choose 0.3% (v/v) as the volume percentage of addition of Nile Red. Pinacyananol chloride (PIN) is a cationic dye belonging to the class of conjugated cyanine dyes. Because of its amphiphilic nature, PIN is soluble in a wide range of solvents including water and chloroform [38]. The UV–visible spectrum of PIN in water is shown in supplementary materials, Fig. S3. The adsorption spectrum of PIN consists of three overlapping spectrum components, with the most intense band red-shifted relative to the others. The sharp peak at 600 nm is assigned to the absorbance of monomer dye molecules. The peak at 560 nm is due to absorbance of the dimeric form of PIN [39]. When PIN is added to solvents of different polarity, both max of bands I and II decrease with the increase of polarity of the solvent (Fig. 2). On the other hand, when PIN is added to liposome solution, band I of the UV–visible spectrum is also found at around 600 nm (Fig. S4 of supplementary materials). PIN is amphiphilic, so when being added to liposome solution, it may exist both inside and outside the liposome bilayer. However, PIN is also cationic, which is electrically expelled by positively charged DODAB in the liposome composition of our study, so the dye molecules are thought to exist in the outer aqueous phase, instead of being incorporated inside the hydrophobic liposome bilayer. From the above results, it is obvious that Nile Red molecules are successfully incorporated inside liposome bilayer while PIN molecules exist in the outer aqueous phase. Due to their sensitivity to the polarity, the molecular probes are suitable to be used as the indicators of the change of their local microenvironment respectively. 3.2. Effect of Chol-PEG-Chol concentration on max of Nile Red UV–visible spectrum Fig. 3a shows effect of Chol-PEG-Chol concentration on max of Nile Red UV–visible spectrum at 25 ◦ C. With the increase of CholPEG-Chol concentration, max of four kinds of liposome all showed blue shift (moving to higher wavelength) which suggests that the liposome bilayer microenvironment becomes more polar with the addition of Chol-PEG-Chol, especially in the case of liposome 2, which is composed of DMPC:DODAB (molar ratio 1:1). In previous studies [30], when tri-block copolymers (PEG-PPOPEG) were added to pre-prepared liposome solution, no obvious shift of max of Nile Red UV–visible spectrum was observed, indicating that the block copolymer molecules do not interfere with the liposome bilayer microenvironment and are only present at the surface of the liposomes. However, in our study, the considerable blue-shift of max of Nile Red confirmed that the amphiphilic polymers really interfere with the liposome bilayer microenvironment by the incorporation of cholesterol end groups into the bilayer and the insertion increased the polarity of the bilayer microenvironment. According to previous researches [40,41], the amphiphilic polymers grafted on the lipid membrane can exert lateral interactions of considerable strength that are likely to modify the properties of the host liposomal membrane. The lateral expansion of lipid membranes resulting from the lateral pressure exerted within the
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Fig. 4. Effect of Chol-PEG-Chol concentration on max of Nile Red-loaded liposome in fluorescence emission spectrum at 25 ◦ C.
surface polymer brush has been inferred directly by the increase in motional freedom of spin-labeled lipid chains in lipid membranes, on incorporating polymer-grafted lipids [42,43]. The steric repulsive interaction between the hydrophilic PEG chains of amphiphilic polymers contributes to membrane stretching and therefore lowers the packing density of the lipid molecules in liposome bilayer. After the incorporation of cholesterol end groups in the liposome bilayer, it tends to be easier for water molecules to enter inside the bilayer and the local microenvironment of Nile Red becomes more polar. With the increase of Chol-PEG-Chol concentration, more and more polymer molecules insert into the bilayer and the packing density of lipid molecules is further lowered, the local microenvironment of Nile Red becomes more polar with more water molecules entering inside the bilayer, which leads to the increase of max wavelength of Nile Red. Besides, cholesterol groups are usually embedded more deeply inside the bilayer than Nile Red, which are usually located in the vicinity of the hydrophilic headgroups of the phospholipids in liposome bilayer [44]. So when Chol-PEG-Chol is added to liposome solution, cholesterol end groups of the polymer will penetrate more deeply into the inner side of liposome bilayer than Nile Red, thus the local microenvironment of Nile Red is supposed to become more polar due to the existence of part of PEG chain which is connected with the cholesterol end-group (Fig. 3b). DODAB is a cationic lipid, with higher DODAB molar ratio in liposome bilayer, the electrostatic repulsion between the positively charged ammonium headgroups of DODAB molecules will make the spacers between headgroups bigger than those of neutral phospholipids molecules. Previous studies showed that DODAB molecules in liposome bilayer arrange spontaneously into a rippled structure. The hydrocarbon chain order is higher in the center of the DODAB membrane than in the region close to the ammonia head groups [45]. So it is obvious that liposome with higher DODAB molar ratio is easier for Chol-PEG-Chol to embed the cholesterol groups into the liposome bilayer due to the structure features. Therefore the local microenvironment of Nile Red in liposome 2 becomes more polar and the blue-shift of max of liposome 2 is much more obvious than the other three kinds of liposome when Chol-PEG-Chol is added. As a major component of the cell plasma membrane, cholesterol has unique cellular functions, such as stabilizing membrane fluidity and filling spaces between neighboring phospholipids [46]. There is a large body of evidence showing that the interactions
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Fig. 5. (a) Effect of Chol-PEG-Chol concentration on max of bands I and II of PIN-loaded liposome in UV–visible spectrum at 25 ◦ C. (b) Schematic illustration of PIN local environment change before and after the addition of Chol-PEG-Chol.
between guest molecules and cholesterol-contained lipid membranes are much more complex than those found from the neat lipid membranes [47]. It is well known cholesterol decreases the permeability of bilayers in the liquid crystalline state, because the cholesterol molecules incorporated into the bilayers make the upper portions of the lipid hydrocarbon chains adopt a more trans configuration, which decrease the number of kinks in the bilayer and increases the order of the lipid molecules. Therefore the inclusion of cholesterol leads to an increased lateral packing density of phosphatidaylcholine layers and decreases the free volume within the hydrophobic part of the layer [48,49]. In our present study, the temperature of the samples is fixed at 25 ◦ C, at which the main lipid composition DMPC (transition temperature Tm = 23 ◦ C) are in the liquid crystalline state. So the cholesterol content in the lipid membrane would impede the penetration of the cholesterol endgroups of Chol-PEG-Chol into the liposome bilayer. In previous studies on transport of chemical species across the lipid bilayer, similar impeding effect was also observed and it was found the
higher cholesterol content, the slower the transport rate [49]. So for Liposomes 3 and 4, effect of Chol-PEG-Chol concentration on max of Nile Red is not so obvious as compared with that of Liposomes 1 and 2.
3.3. Effect of Chol-PEG-Chol concentration on the Nile Red max of fluorescence spectrum Fig. 4 shows effect of Chol-PEG-Chol concentration on max of Nile Red fluorescence spectrum at 25 ◦ C. With the increase of CholPEG-Chol concentration, max of Nile Red fluorescence spectrum of liposome all showed moderate blue shift except liposome 2, in which max considerably increased from about 590 nm to 615 nm. These results of fluorescence spectrum correlate well with those of UV–visible spectrum, indicating that the bilayer microenvironment around Nile Red is becoming more polar with increasing polymer concentration, especially for liposome with high ratio of DODAB.
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3.4. Effect of Chol-PEG-Chol concentration on max of PIN UV–visible spectrum Fig. 5a shows effect of Chol-PEG-Chol concentration on max of bands I and II of PIN UV–visible spectrum at 25 ◦ C. For all four kinds of liposome, max of both bands I and II increased with the addition of Chol-PEG-Chol. The higher the polarity of the solvent, the lower the max of both bands I and II, we can see that the polarity of the local microenvironment of PIN becomes more apolar with the addition of Chol-PEG-Chol. As we described before, PIN is a positively charged, amphiphilic molecular dye and the four kinds of liposome in this study are all positively charged because of DODAB, so PIN molecules are thought to aggregate at the aqueous solvent outside of the liposome bilayer, instead of being entrapped in the inner side of the liposome bilayer. Before addition of CholPEG-Chol, PIN molecules are surrounded by aqueous environment outside of liposome, which has polarity same as that of water. However, when Chol-PEG-Chol is added to liposome and cholesterol end groups of Chol-PEG-Chol penetrate into the bilayer of liposome, PIN molecules are surrounded by PEG polymer solution (Fig. 5b). According to previous studies [50], the polarity of aqueous medium is found to decrease with increasing PEG size and PEG polymer concentration and the polarity decrease is due to the structuring action of the solutes on water. So in our experiment, with the addition of Chol-PEG-Chol, the PIN local environment becomes more apolar and max of both bands I and II of PIN increases accordingly. 4. Conclusion In this paper, we used two kinds of molecular probes to investigate the change of polarity of liposome bilayer microenvironment when Chol-PEG-Chol was added. By monitoring the shift of max of UV–visible spectrum and fluorescence spectrum of Nile Red, a highly hydrophobic molecular probe, it was confirmed that the cholesterol end groups of Chol-PEG-Chol really insert into the liposome bilayer and the incorporation increased the polarity of liposome bilayer microenvironment. On the other hand, the shift of max of UV–visible spectrum of PIN, a positively charged, amphiphilic molecular dye which is thought to exist in the aqueous solvent outside of the liposome bilayer, suggested that local microenvironment of PIN becomes more apolar with the addition of Chol-PEG-Chol. For liposome with different composition, it was found that with high ratio of DODAB, the polarity of liposome bilayer is more easily affected by the addition of Chol-PEG-Chol while liposome with cholesterol shows relatively high stability to the addition of Chol-PEGChol. Acknowledgements The authors are indebted to Dr. C. Kataoka, National Institute for Materials Science, for stimulating advices on liposome preparation. This work was financially supported in part by the Japan Society for the Promotion of Science (JSPS) through its Funding Program for World-Leading Innovation R&D on Science and Technology (FIRST Program), a Grant-in-Aid from the National Institute of Biomedical Innovation of Japan and the World Premier International Research Center (WPI) Initiative on Materials Nanoarchitectonics, MEXT, Japan.
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Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/ j.colsurfb.2012.03.017. References [1] T. Lian, R.J.Y. Ho, J. Pharm. Sci. 90 (2001) 667. [2] R.R. Sawant, V.P. Torchilin, Soft Matter 6 (2010) 4026. [3] V.P. Torchilin, V. Weissig, Liposomes: A Practical Approach, Oxford University Press, New York, 2003. [4] A. Caria, A. Khan, Langmuir 12 (1996) 6282. [5] A. Raudino, F. Castelli, Marcromolecules 30 (1997) 2495. [6] O. Regev, E.F. Marques, A. Khan, Langmuir 15 (1999) 642. [7] M.L. Hwang, R.K. Prud’homme, J. Kohn, J.L. Thomas, Langmuir 17 (2001) 7713. [8] S. Beugin-Deroo, M. Ollivon, S. Lesieur, J. Colloid Interface Sci. 202 (1998) 324. [9] J. Kevelam, J.F.L. van Breemen, W. Blokzijil, J.B.F.N. Engberts, Langmuir 12 (1996) 4709. [10] A. Polozova, F.M. Winnik, Biochim. Biophys. Acta 1326 (1997) 213. [11] P.G. de Gennes, J. Phys. Chem. 94 (1990) 8407. [12] H. Ringsdorf, B. Schlarb, J. Venzmer, Angew. Chem. Int. Ed. Engl. 27 (1988) 113. [13] T. Taguchi, Z. Rao, M. Ito, M. Matsuda, J. Mater. Sci. Mater. Med. 22 (2011) 2357. [14] M. Ito, T. Taguchi, Acta Biomater. 5 (2009) 2945. [15] W. Meier, J. Hotz, S. Gunther-Ausborn, Langmuir 12 (1996) 5028. [16] J.H. Lee, J.P. Gustin, T. Chen, G.F. Payne, S.R. Raghavan, Langmuir 21 (2005) 26. [17] B. Medronho, F.E. Antunes, B. Lindman, M.G. Miguel, J. Dispers. Sci. Technol. 27 (2006) 83. [18] C.C. Lin, C.H. Chang, Y.M. Yang, Colloids Surf. A: Physicochem. Eng. Aspects 346 (2009) 66. [19] H.S. Ashbaugh, K. Boon, R.K. Prud’homme, Colloid Polym. Sci. 280 (2002) 783. [20] Z. Rao, M. Inoue, M. Matsuda, T. Taguchi, Colloids Surf. B: Biointerfaces 82 (2011) 196. [21] G. Bai, M. Nichifor, M. Bastos, J. Phys. Chem. Lett. 1 (2010) 932. [22] M. Johnson, M. Silvander, G. Karlsson, K. Edwards, Langmuir 15 (1999) 6314. [23] C. Ladavière, M. Toustou, T. Gulik-Krzywicki, C. Tribet, J. Colloid Interface Sci. 241 (2001) 178. [24] J. Simon, M. Kühner, H. Ringsdorf, E. Sackmanna, Chem. Phys. Lipids 76 (1995) 241. [25] V. Frette, I. Tsafrir, M.-A. Guedeau-Boudeville, L. Jullien, D. Kandell, J. Stavans, Phys. Rev. Lett. 83 (1999) 2465. [26] J.C. Chung, D.J. Gross, J.L. Thomas, D.A. Tirrell, L.R. Opsahl-Ong, Macromolecules 29 (1996) 4636. [27] V.P. Torchilin, V.S. Trubetskoy, Adv. Drug Deliv. Rev. 16 (1995) 141. [28] K. Knop, R. Hoogenboom, D. Fischer, U.S. Schubert, Angew. Chem. Int. Ed. Engl. 49 (2010) 6288. [29] G. Liu, L. Fu, G. Zhang, J. Phys. Chem. B 113 (2009) 3365. [30] K. Kostarelos, P.F. Luckham, T.F. Tadros, J. Colloid Interface Sci. 191 (1997) 341. [31] P. Kaschny, F.M. Goni, J. Colloid Interface Sci. 160 (1993) 24. [32] M.E.C.D.R. Oliveira, G. Hungerford, M.G. Miguel, H.D. Burrows, J. Mol. Struct. 563–564 (2001) 443. [33] R. Sabate, J. Estelrich, J. Phys. Chem. B 107 (2003) 4137. [34] B.R. Lentz, Chem. Phys. Lipids 64 (1993) 99. [35] P. Greenspan, S.D. Fowler, J. Lipid Res. 26 (1985) 781. [36] M.K. McMillian, E.R. Grant, Z. Zhong, J.B. Parker, L. Li, R.A. Zivin, M.E. Burczynski, M.D. Johnson, In Vitro Mol. Toxicol. 14 (2001) 177. [37] D.L. Sackett, J. Wolff, Anal. Biochem. 167 (1987) 228. [38] K. Manna, A.K. Panda, Spectrochim. Acta A: Mol. Biomol. Spectrosc. 74 (2009) 1268. [39] R. Sabate, M. Gallardo, A. de la Maza, J. Estelrich, Langmuir 17 (2001) 6433. [40] C. Tribet, F. Vial, Soft Matter 4 (2008) 68. [41] O. Tirosh, Y. Barenholz, J. Katzhendler, A. Priev, J. Biophys. 74 (1998) 1371. [42] G. Montesano, R. Bartucci, S. Belsito, D. Marsh, L. Sportelli, J. Biophys. 80 (2001) 1372. [43] D. Marsh, J. Biophys. 81 (2001) 2154. ´ K. Nawalany, M. Kumorek, A. Kobierska, B. Jachimska, M. [44] M. Kepczynski, Nowakowska, Chem. Phys. Lipids 155 (2008) 7. [45] D. Jamroz, M. Kepczynski, M. Nowakowska, Langmuir 26 (2010) 15076. [46] P.L. Yeagle, Biochim. Biophys. Acta 822 (1985) 267. [47] T. Soderlund, J.Y.A. Lehtonen, P.K.J. Kinnunen, Mol. Pharmacol. 55 (1999) 32. [48] N.L. Gershfeld, R.E. Pagano, J. Phys. Chem. 76 (1972) 1244. [49] E.C.Y. Yan, K.B. Eisenthal, Biophys. J. 79 (2000) 898. [50] B.Y. Zaslavsky, L.M. Miheeva, E.A. Masimov, S.F. Djafarov, C. Reichardt, J. Chem. Soc. Faraday Trans. 86 (1990) 519.
Contents lists available at SciVerse ScienceDirect
Colloids and Surfaces B: Biointerfaces journal homepage: www.elsevier.com/locate/colsurfb
Spectroscopic studies on interactions between cholesterol-end capped polyethylene glycol and liposome Zhi Rao a,b , Tetsushi Taguchi a,b,∗ a b
Graduate School of Pure and Applied Sciences, University of Tsukuba, 1-1-1 Tennodai, Tsukuba, Ibaraki 305-8577, Japan Biomaterials Unit, Nano-bio field, Materials Nanoarchitectonics (MANA), National Institute for Materials Science, 1-1 Namiki, Tsukuba, Ibaraki 305-0044, Japan
a r t i c l e
i n f o
Article history: Received 12 January 2012 Received in revised form 26 March 2012 Accepted 27 March 2012 Available online 3 April 2012 Keywords: Liposome Cholesterol end group Nile Red Pinacyanol chloride Bilayer microenvironment
a b s t r a c t In order to confirm that the cholesterol end groups of cholesterol-end capped polyethylene glycol really insert into the liposome bilayer and investigate how the incorporation affects the microenvironment of liposome bilayer, two kinds of molecular probes, namely Nile Red and pinacyanol chloride, were used. Their UV–visible and fluorescence spectrum were recorded before and after the addition of the polymer. Shifts of the maximum absorbance (max ) of Nile Red show that the bilayer microenvironment around Nile Red is becoming more polar with increasing polymer concentration while shifts of max of pinacyanol chloride indicate that the surrounding environment of pinacyanol chloride is becoming more apolar with addition of polymer. Effect of composition of liposome was also studied. With high ratio of dimethyldioctadecylammonium bromide (DODAB) fraction in liposome, max of Nile Red is more easily affected by the addition of Chol-PEG-Chol while liposome with cholesterol shows relatively high stability to the addition of Chol-PEG-Chol. © 2012 Elsevier B.V. All rights reserved.
1. Introduction Spherical bilayer lipid vesicles (liposome) have been used extensively for drug delivery because of their ability to solubilize both hydrophilic and hydrophobic substances in their inner aqueous phase and liposome bilayer. Drug encapsulation by liposome allows in vivo protection from enzymes and thus enhances circulation time and bioavailability [1–3]. However, a major problem with liposome is still their insufficient stability, which may limit the practical application. Nevertheless, the stability can be enhanced by the addition of surfactants or amphiphilic polymer [4–8]. Studies on liposome–amphiphilic polymer systems have become an important research area of colloid science, from both experimental and theoretical view points [9–11]. The liposome–polymer systems constitute good model system for living cells [12–14], and they also lead to important phase behavior phenomena, such as the formation of gels and networks [15–20], with potential industrial applications. Among the studies on interactions between amphiphilic polymer and liposome, some seek to identify polymers that, upon binding to liposomes, markedly affect the shape, curvature, stiffness, or stability of the bilayer [21–26], while others
∗ Corresponding author at: Biomaterials Unit, Nano-bio field, Materials Nanoarchitectonics (MANA), National Institute for Materials Science, 1-1 Namiki, Tsukuba, Ibaraki 305-0044, Japan. Tel.: +81 29 860 4498; fax: +81 29 860 4714. E-mail address: [email protected] (T. Taguchi). 0927-7765/$ – see front matter © 2012 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.colsurfb.2012.03.017
often aim at improving the stability and at controlling the permeability of liposomes for drug delivery or targeting, gene therapy, and so forth [27,28]. However, even for the simple lipid membrane system, there are still disputations over whether the hydrophobic groups really anchor into the lipid bilayer or not. Some previous studies indicated that hydrophobic groups directly penetrated into the liposome bilayer and the insertion profoundly altered the morphology or fluidity or of the liposome bilayer [10,29], while some others showed that the polymers were only present at the surface of the liposomes and do not interfere with the lipid bilayer microenvironment [30]. Various techniques have been used for the study of the liposome bilayer microenvironment. The use of molecular probes has proven particularly advantageous in providing information about the bilayer structure [30–34]. These molecules are either solubilized in the bilayer at optimum concentration or are covalently bonded to the phospholipids forming the bilayer. Investigations of the fluorescence and UV–visible spectrum of such molecular probes have provided valuable information about the electric, dynamic, conformational, and structural properties of the liposome bilayer. It is well known that the microenvironment surrounding the molecular probe influences its electronic structure and thus its photophysics. Changes in the local microenvironment can produce measurable spectrum shifts which can, in turn, be monitored spectroscopically. This property, known as solvatochromism, allows elucidation of the influence of the immediate environment of the molecules within the probed
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systems, and moreover, gives evidence of specific interactions [33]. We have previously reported on the self-assembled liposome gel from liposome and cholesterol-end capped polyethylene glycol (Chol-PEG-Chol) and carried out systematic studies on the gel mechanical properties by rheological method [20]. However, whether cholesterol end groups of Chol-PEG-Chol really insert into the liposome bilayer and how the incorporation affects the microenvironment of liposome bilayer is of great research interest and by far, no studies have yet been carried out on this aspect. In this work, we used two kinds of molecular probes, Nile Red and pinacyanol chloride (PIN) to investigate the interactions between Chol-PEG-Chol and liposome. By monitoring the changes in the UV–visible absorption and fluorescence emission spectrum of the molecular probes, information was obtained about their local microenvironment. The results may provide some insights on the interactions between the amphiphilic polymer Chol-PEG-Chol and liposome bilayer. 2. Materials and methods 2.1. Materials Dimyristoylphosphatidylcholine (DMPC) was purchased from NOF Corporation (Tokyo, Japan). Dimethyldioctadecylammonium bromide (DODAB, >98%) was purchased from Sigma–Aldrich. Cholesterol was purchased from Wako Pure Chemical Industrials Ltd. (Osaka, Japan). Diamino-polyethylene glycol (molecular weight: 30,000, >98%) was purchased from NOF Corporation (Tokyo, Japan). Cholesteryl chloroformate (>99%) was purchased from Fluka (Buchs, Switzerland). Dichloromethane was purchased from Nacalai Tesque, Inc. (Kyoto, Japan). Triethylamine was purchased from Wako Pure Chemical Industrials Ltd. (Osaka, Japan). Nile Red and pinacynol chloride (PIN) were purchased from Tokyo Chemical Industry Co., Ltd. (Tokyo Japan). All chemicals were used without further purification.
Fig. 1. Relation between solvent polarity and max of Nile Red in UV–visible and fluorescence emission spectrum in different organic solvent.
added to liposome solution in 5 mL tube, the mixture was shaken gently at room temperature (25 ◦ C) on a micromixer overnight until Chol-PEG-Chol fully dissolved. 2.5. UV–visible measurements UV–visible measurements were carried out using JASCO V660 UV–visible spectrophotometer (Tokyo, Japan), recording
2.2. Preparation of liposome solution In this work, four kinds of liposome of different composition were prepared. Liposome 1 (DMPC:DODAB molar ratio = 9:1); liposome 2 (DMPC:DODAB molar ratio = 1:1); liposome 3 (DMPC:DODAB:cholesterol molar ratio = 8:1:1); liposome 4 (DMPC:DODAB:cholesterol molar ratio = 6:1:3). The liposome was prepared by extrusion method [20]. The final liposome concentration was 1 mM. According to the dynamic light scattering measurement, the average liposome particle size was 109 nm, 193 nm, 111 nm, 129 nm respectively. Nile Red was introduced to the final liposome solution by injection of 0.3% (v/v) stock solution (0.25 mM) of the dye in ethanol. PIN was introduced to the final liposome solution by injection of 1% (v/v) of stock solution (1 mM) of the dye in DMSO. 2.3. Preparation of Chol-PEG-Chol Chol-PEG-Chol was prepared from diamino-polyethylene glycol and cholesteryl chloroformate following the procedure of Ref. [20]. 2.4. Preparation of mixture solution of Chol-PEG-Chol and liposome In this paper, Chol-PEG-Chol concentration varied from 0 to 5 mg/mL and liposome concentration was fixed at 1 mM. Based on our preliminary experiments, in this concentration range, the viscosity of the mixture of liposome and Chol-PEG-Chol was not too high to interfere with the measurement. After Chol-PEG-Chol was
Fig. 2. Relation between solvent polarity and max of bands I and II of PIN in UV–visible spectrum in different solvent.
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Fig. 3. (a) Effect of Chol-PEG-Chol concentration on max of Nile Red-loaded liposome in UV–visible spectrum at 25 ◦ C. (b) Schematic illustration of Nile Red local microenvironment change before and after the addition of Chol-PEG-Chol.
the absorbance spectrum between 400 and 700 nm wavelength directly. 2.6. Fluorescence measurements Fluorescence measurements were taken using JASCO FP-6500 fluorescent spectrometer (Tokyo, Japan). Excitation and emission slits were both fixed at 5 nm, and exc was 550 nm. 3. Results and discussion 3.1. UV–visible and fluorescence spectrum of molecular probes with relation to the solvent polarity Nile Red is an uncharged benzeophenoxazone dye, well soluble in organic solvents and lipids, but almost unsoluble and strongly quenches in water. It is photochemically stable and can be monitored by both UV–visible adsorption spectrum and fluorescence
spectrum [30]. Nile Red has been used as a fluorescent probe for intracellular lipids and as a polarity sensor deducing protein structure and conformation [35–37]. Being highly hydrophobic, both the UV–visible and fluorescence spectrum of Nile Red in water shows only flat line with no absorbance or intensity (supplementary materials, Fig. S1). However, when Nile Red was added to organic solvents, which have polarity lower than water, both the UV–visible and fluorescence spectrum of Nile Red in organic solvents show single broad peak (supplementary materials, Fig. S1). Fig. 1 shows the relation of the maximum wavelength (max ) of Nile Red UV–visible adsorption spectrum and fluorescence spectrum with the polarity of the organic solvent. It is obvious that max of both UV–visible adsorption spectrum and fluorescence spectrum increase with the increase of polarity of the organic solvent. This finding correlates well with previous work in which Nile Red max shifted to higher wavelength as the polarity of methanol/chloroform mixtures was increased [30].
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When Nile Red is introduced into liposome solution, both UV–visible and fluorescence spectrum of liposome loaded with Nile Red show single broad peak, similar to that of Nile Red in organic solvent, as shown in Fig. S2 of supplementary materials. This shows that the hydrophobic dye molecules are easily solubilized inside the liposome bilayer due to the apolar microenvironment inside bilayer. With the increase of volume percentage of addition of Nile Red to liposome 1, intensity of both UV–visible adsorption spectrum and fluorescence spectrum increased accordingly and no obvious shift of max was observed, indicating that the polarity of Nile Red vicinity microenvironment remain unchanged in this concentration range of Nile Red. In this study, we choose 0.3% (v/v) as the volume percentage of addition of Nile Red. Pinacyananol chloride (PIN) is a cationic dye belonging to the class of conjugated cyanine dyes. Because of its amphiphilic nature, PIN is soluble in a wide range of solvents including water and chloroform [38]. The UV–visible spectrum of PIN in water is shown in supplementary materials, Fig. S3. The adsorption spectrum of PIN consists of three overlapping spectrum components, with the most intense band red-shifted relative to the others. The sharp peak at 600 nm is assigned to the absorbance of monomer dye molecules. The peak at 560 nm is due to absorbance of the dimeric form of PIN [39]. When PIN is added to solvents of different polarity, both max of bands I and II decrease with the increase of polarity of the solvent (Fig. 2). On the other hand, when PIN is added to liposome solution, band I of the UV–visible spectrum is also found at around 600 nm (Fig. S4 of supplementary materials). PIN is amphiphilic, so when being added to liposome solution, it may exist both inside and outside the liposome bilayer. However, PIN is also cationic, which is electrically expelled by positively charged DODAB in the liposome composition of our study, so the dye molecules are thought to exist in the outer aqueous phase, instead of being incorporated inside the hydrophobic liposome bilayer. From the above results, it is obvious that Nile Red molecules are successfully incorporated inside liposome bilayer while PIN molecules exist in the outer aqueous phase. Due to their sensitivity to the polarity, the molecular probes are suitable to be used as the indicators of the change of their local microenvironment respectively. 3.2. Effect of Chol-PEG-Chol concentration on max of Nile Red UV–visible spectrum Fig. 3a shows effect of Chol-PEG-Chol concentration on max of Nile Red UV–visible spectrum at 25 ◦ C. With the increase of CholPEG-Chol concentration, max of four kinds of liposome all showed blue shift (moving to higher wavelength) which suggests that the liposome bilayer microenvironment becomes more polar with the addition of Chol-PEG-Chol, especially in the case of liposome 2, which is composed of DMPC:DODAB (molar ratio 1:1). In previous studies [30], when tri-block copolymers (PEG-PPOPEG) were added to pre-prepared liposome solution, no obvious shift of max of Nile Red UV–visible spectrum was observed, indicating that the block copolymer molecules do not interfere with the liposome bilayer microenvironment and are only present at the surface of the liposomes. However, in our study, the considerable blue-shift of max of Nile Red confirmed that the amphiphilic polymers really interfere with the liposome bilayer microenvironment by the incorporation of cholesterol end groups into the bilayer and the insertion increased the polarity of the bilayer microenvironment. According to previous researches [40,41], the amphiphilic polymers grafted on the lipid membrane can exert lateral interactions of considerable strength that are likely to modify the properties of the host liposomal membrane. The lateral expansion of lipid membranes resulting from the lateral pressure exerted within the
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Fig. 4. Effect of Chol-PEG-Chol concentration on max of Nile Red-loaded liposome in fluorescence emission spectrum at 25 ◦ C.
surface polymer brush has been inferred directly by the increase in motional freedom of spin-labeled lipid chains in lipid membranes, on incorporating polymer-grafted lipids [42,43]. The steric repulsive interaction between the hydrophilic PEG chains of amphiphilic polymers contributes to membrane stretching and therefore lowers the packing density of the lipid molecules in liposome bilayer. After the incorporation of cholesterol end groups in the liposome bilayer, it tends to be easier for water molecules to enter inside the bilayer and the local microenvironment of Nile Red becomes more polar. With the increase of Chol-PEG-Chol concentration, more and more polymer molecules insert into the bilayer and the packing density of lipid molecules is further lowered, the local microenvironment of Nile Red becomes more polar with more water molecules entering inside the bilayer, which leads to the increase of max wavelength of Nile Red. Besides, cholesterol groups are usually embedded more deeply inside the bilayer than Nile Red, which are usually located in the vicinity of the hydrophilic headgroups of the phospholipids in liposome bilayer [44]. So when Chol-PEG-Chol is added to liposome solution, cholesterol end groups of the polymer will penetrate more deeply into the inner side of liposome bilayer than Nile Red, thus the local microenvironment of Nile Red is supposed to become more polar due to the existence of part of PEG chain which is connected with the cholesterol end-group (Fig. 3b). DODAB is a cationic lipid, with higher DODAB molar ratio in liposome bilayer, the electrostatic repulsion between the positively charged ammonium headgroups of DODAB molecules will make the spacers between headgroups bigger than those of neutral phospholipids molecules. Previous studies showed that DODAB molecules in liposome bilayer arrange spontaneously into a rippled structure. The hydrocarbon chain order is higher in the center of the DODAB membrane than in the region close to the ammonia head groups [45]. So it is obvious that liposome with higher DODAB molar ratio is easier for Chol-PEG-Chol to embed the cholesterol groups into the liposome bilayer due to the structure features. Therefore the local microenvironment of Nile Red in liposome 2 becomes more polar and the blue-shift of max of liposome 2 is much more obvious than the other three kinds of liposome when Chol-PEG-Chol is added. As a major component of the cell plasma membrane, cholesterol has unique cellular functions, such as stabilizing membrane fluidity and filling spaces between neighboring phospholipids [46]. There is a large body of evidence showing that the interactions
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Fig. 5. (a) Effect of Chol-PEG-Chol concentration on max of bands I and II of PIN-loaded liposome in UV–visible spectrum at 25 ◦ C. (b) Schematic illustration of PIN local environment change before and after the addition of Chol-PEG-Chol.
between guest molecules and cholesterol-contained lipid membranes are much more complex than those found from the neat lipid membranes [47]. It is well known cholesterol decreases the permeability of bilayers in the liquid crystalline state, because the cholesterol molecules incorporated into the bilayers make the upper portions of the lipid hydrocarbon chains adopt a more trans configuration, which decrease the number of kinks in the bilayer and increases the order of the lipid molecules. Therefore the inclusion of cholesterol leads to an increased lateral packing density of phosphatidaylcholine layers and decreases the free volume within the hydrophobic part of the layer [48,49]. In our present study, the temperature of the samples is fixed at 25 ◦ C, at which the main lipid composition DMPC (transition temperature Tm = 23 ◦ C) are in the liquid crystalline state. So the cholesterol content in the lipid membrane would impede the penetration of the cholesterol endgroups of Chol-PEG-Chol into the liposome bilayer. In previous studies on transport of chemical species across the lipid bilayer, similar impeding effect was also observed and it was found the
higher cholesterol content, the slower the transport rate [49]. So for Liposomes 3 and 4, effect of Chol-PEG-Chol concentration on max of Nile Red is not so obvious as compared with that of Liposomes 1 and 2.
3.3. Effect of Chol-PEG-Chol concentration on the Nile Red max of fluorescence spectrum Fig. 4 shows effect of Chol-PEG-Chol concentration on max of Nile Red fluorescence spectrum at 25 ◦ C. With the increase of CholPEG-Chol concentration, max of Nile Red fluorescence spectrum of liposome all showed moderate blue shift except liposome 2, in which max considerably increased from about 590 nm to 615 nm. These results of fluorescence spectrum correlate well with those of UV–visible spectrum, indicating that the bilayer microenvironment around Nile Red is becoming more polar with increasing polymer concentration, especially for liposome with high ratio of DODAB.
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3.4. Effect of Chol-PEG-Chol concentration on max of PIN UV–visible spectrum Fig. 5a shows effect of Chol-PEG-Chol concentration on max of bands I and II of PIN UV–visible spectrum at 25 ◦ C. For all four kinds of liposome, max of both bands I and II increased with the addition of Chol-PEG-Chol. The higher the polarity of the solvent, the lower the max of both bands I and II, we can see that the polarity of the local microenvironment of PIN becomes more apolar with the addition of Chol-PEG-Chol. As we described before, PIN is a positively charged, amphiphilic molecular dye and the four kinds of liposome in this study are all positively charged because of DODAB, so PIN molecules are thought to aggregate at the aqueous solvent outside of the liposome bilayer, instead of being entrapped in the inner side of the liposome bilayer. Before addition of CholPEG-Chol, PIN molecules are surrounded by aqueous environment outside of liposome, which has polarity same as that of water. However, when Chol-PEG-Chol is added to liposome and cholesterol end groups of Chol-PEG-Chol penetrate into the bilayer of liposome, PIN molecules are surrounded by PEG polymer solution (Fig. 5b). According to previous studies [50], the polarity of aqueous medium is found to decrease with increasing PEG size and PEG polymer concentration and the polarity decrease is due to the structuring action of the solutes on water. So in our experiment, with the addition of Chol-PEG-Chol, the PIN local environment becomes more apolar and max of both bands I and II of PIN increases accordingly. 4. Conclusion In this paper, we used two kinds of molecular probes to investigate the change of polarity of liposome bilayer microenvironment when Chol-PEG-Chol was added. By monitoring the shift of max of UV–visible spectrum and fluorescence spectrum of Nile Red, a highly hydrophobic molecular probe, it was confirmed that the cholesterol end groups of Chol-PEG-Chol really insert into the liposome bilayer and the incorporation increased the polarity of liposome bilayer microenvironment. On the other hand, the shift of max of UV–visible spectrum of PIN, a positively charged, amphiphilic molecular dye which is thought to exist in the aqueous solvent outside of the liposome bilayer, suggested that local microenvironment of PIN becomes more apolar with the addition of Chol-PEG-Chol. For liposome with different composition, it was found that with high ratio of DODAB, the polarity of liposome bilayer is more easily affected by the addition of Chol-PEG-Chol while liposome with cholesterol shows relatively high stability to the addition of Chol-PEGChol. Acknowledgements The authors are indebted to Dr. C. Kataoka, National Institute for Materials Science, for stimulating advices on liposome preparation. This work was financially supported in part by the Japan Society for the Promotion of Science (JSPS) through its Funding Program for World-Leading Innovation R&D on Science and Technology (FIRST Program), a Grant-in-Aid from the National Institute of Biomedical Innovation of Japan and the World Premier International Research Center (WPI) Initiative on Materials Nanoarchitectonics, MEXT, Japan.
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Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/ j.colsurfb.2012.03.017. References [1] T. Lian, R.J.Y. Ho, J. Pharm. Sci. 90 (2001) 667. [2] R.R. Sawant, V.P. Torchilin, Soft Matter 6 (2010) 4026. [3] V.P. Torchilin, V. Weissig, Liposomes: A Practical Approach, Oxford University Press, New York, 2003. [4] A. Caria, A. Khan, Langmuir 12 (1996) 6282. [5] A. Raudino, F. Castelli, Marcromolecules 30 (1997) 2495. [6] O. Regev, E.F. Marques, A. Khan, Langmuir 15 (1999) 642. [7] M.L. Hwang, R.K. Prud’homme, J. Kohn, J.L. Thomas, Langmuir 17 (2001) 7713. [8] S. Beugin-Deroo, M. Ollivon, S. Lesieur, J. Colloid Interface Sci. 202 (1998) 324. [9] J. Kevelam, J.F.L. van Breemen, W. Blokzijil, J.B.F.N. Engberts, Langmuir 12 (1996) 4709. [10] A. Polozova, F.M. Winnik, Biochim. Biophys. Acta 1326 (1997) 213. [11] P.G. de Gennes, J. Phys. Chem. 94 (1990) 8407. [12] H. Ringsdorf, B. Schlarb, J. Venzmer, Angew. Chem. Int. Ed. Engl. 27 (1988) 113. [13] T. Taguchi, Z. Rao, M. Ito, M. Matsuda, J. Mater. Sci. Mater. Med. 22 (2011) 2357. [14] M. Ito, T. Taguchi, Acta Biomater. 5 (2009) 2945. [15] W. Meier, J. Hotz, S. Gunther-Ausborn, Langmuir 12 (1996) 5028. [16] J.H. Lee, J.P. Gustin, T. Chen, G.F. Payne, S.R. Raghavan, Langmuir 21 (2005) 26. [17] B. Medronho, F.E. Antunes, B. Lindman, M.G. Miguel, J. Dispers. Sci. Technol. 27 (2006) 83. [18] C.C. Lin, C.H. Chang, Y.M. Yang, Colloids Surf. A: Physicochem. Eng. Aspects 346 (2009) 66. [19] H.S. Ashbaugh, K. Boon, R.K. Prud’homme, Colloid Polym. Sci. 280 (2002) 783. [20] Z. Rao, M. Inoue, M. Matsuda, T. Taguchi, Colloids Surf. B: Biointerfaces 82 (2011) 196. [21] G. Bai, M. Nichifor, M. Bastos, J. Phys. Chem. Lett. 1 (2010) 932. [22] M. Johnson, M. Silvander, G. Karlsson, K. Edwards, Langmuir 15 (1999) 6314. [23] C. Ladavière, M. Toustou, T. Gulik-Krzywicki, C. Tribet, J. Colloid Interface Sci. 241 (2001) 178. [24] J. Simon, M. Kühner, H. Ringsdorf, E. Sackmanna, Chem. Phys. Lipids 76 (1995) 241. [25] V. Frette, I. Tsafrir, M.-A. Guedeau-Boudeville, L. Jullien, D. Kandell, J. Stavans, Phys. Rev. Lett. 83 (1999) 2465. [26] J.C. Chung, D.J. Gross, J.L. Thomas, D.A. Tirrell, L.R. Opsahl-Ong, Macromolecules 29 (1996) 4636. [27] V.P. Torchilin, V.S. Trubetskoy, Adv. Drug Deliv. Rev. 16 (1995) 141. [28] K. Knop, R. Hoogenboom, D. Fischer, U.S. Schubert, Angew. Chem. Int. Ed. Engl. 49 (2010) 6288. [29] G. Liu, L. Fu, G. Zhang, J. Phys. Chem. B 113 (2009) 3365. [30] K. Kostarelos, P.F. Luckham, T.F. Tadros, J. Colloid Interface Sci. 191 (1997) 341. [31] P. Kaschny, F.M. Goni, J. Colloid Interface Sci. 160 (1993) 24. [32] M.E.C.D.R. Oliveira, G. Hungerford, M.G. Miguel, H.D. Burrows, J. Mol. Struct. 563–564 (2001) 443. [33] R. Sabate, J. Estelrich, J. Phys. Chem. B 107 (2003) 4137. [34] B.R. Lentz, Chem. Phys. Lipids 64 (1993) 99. [35] P. Greenspan, S.D. Fowler, J. Lipid Res. 26 (1985) 781. [36] M.K. McMillian, E.R. Grant, Z. Zhong, J.B. Parker, L. Li, R.A. Zivin, M.E. Burczynski, M.D. Johnson, In Vitro Mol. Toxicol. 14 (2001) 177. [37] D.L. Sackett, J. Wolff, Anal. Biochem. 167 (1987) 228. [38] K. Manna, A.K. Panda, Spectrochim. Acta A: Mol. Biomol. Spectrosc. 74 (2009) 1268. [39] R. Sabate, M. Gallardo, A. de la Maza, J. Estelrich, Langmuir 17 (2001) 6433. [40] C. Tribet, F. Vial, Soft Matter 4 (2008) 68. [41] O. Tirosh, Y. Barenholz, J. Katzhendler, A. Priev, J. Biophys. 74 (1998) 1371. [42] G. Montesano, R. Bartucci, S. Belsito, D. Marsh, L. Sportelli, J. Biophys. 80 (2001) 1372. [43] D. Marsh, J. Biophys. 81 (2001) 2154. ´ K. Nawalany, M. Kumorek, A. Kobierska, B. Jachimska, M. [44] M. Kepczynski, Nowakowska, Chem. Phys. Lipids 155 (2008) 7. [45] D. Jamroz, M. Kepczynski, M. Nowakowska, Langmuir 26 (2010) 15076. [46] P.L. Yeagle, Biochim. Biophys. Acta 822 (1985) 267. [47] T. Soderlund, J.Y.A. Lehtonen, P.K.J. Kinnunen, Mol. Pharmacol. 55 (1999) 32. [48] N.L. Gershfeld, R.E. Pagano, J. Phys. Chem. 76 (1972) 1244. [49] E.C.Y. Yan, K.B. Eisenthal, Biophys. J. 79 (2000) 898. [50] B.Y. Zaslavsky, L.M. Miheeva, E.A. Masimov, S.F. Djafarov, C. Reichardt, J. Chem. Soc. Faraday Trans. 86 (1990) 519.