Polyelectrolyte-Coated Unilamellar Nanometer-Sized Magnetic

Mar 24, 2009 - Jacobs University Bremen, Campus Ring 1, 28725 Bremen, Germany ... Albert-Ludwigs-University of Freiburg Hermann-Herder-Strasse 9, ...
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Polyelectrolyte-Coated Unilamellar Nanometer-Sized Magnetic Liposomes :: Joana Filipa Pereira da Silva Gomes,† Anja Rank,‡ Astrid Kronenberger,† Jurgen Fritz,† † ,† Mathias Winterhalter, and Yannic Ramaye* †

Jacobs University Bremen, Campus Ring 1, 28725 Bremen, Germany, and ‡Institute for Pharmaceutical Sciences Department of Pharmaceutical Technology and Biopharmacy, Albert-Ludwigs-University of Freiburg Hermann-Herder-Strasse 9, 79104 Freiburg, Germany Received January 24, 2009. Revised Manuscript Received February 23, 2009 Superparamagnetic nanoparticles were encapsulated in liposomes prior to the stepwise adsorption of polyelectrolytes of opposite charges commonly known as the layer-by-layer (LbL) technique. Magnetic fields allow a fast separation of coated liposomes from unbound polyelectrolytes. The coated particles were characterized by dynamic light scattering (DLS), cryo-TEM, AFM, and zeta-potential techniques. The presence of magnetic nanoparticles and the polyelectrolyte shell opens the possibility of their magnetic manipulation and targeting by applying an external magnetic or electric field.

Introduction Nanocapsules provide a compartmentalized volume in which reactions can take place, thus protecting unstable molecules from hostile environments. For example, in cosmetics, these systems are used as protective shells for the encapsulation of labile substances such as antioxidants. In pharmacology, there is an increasing need for nanometer-sized carriers to deliver drugs at the site of interest. Moreover, the encapsulation of drugs in nanospheres, nanocapsules, or liposomes may increase their biodistribution, stability, and solubility.1,2 Liposomes or lipid vesicles are formed spontaneously by the selfassembly of phospholipid molecules because of their amphiphilic character.3 The interest in liposomes as a carrier is based on their broad potential to enclose and protect molecules of biological interest and to deliver them functionally intact and in significant quantities.5-8 Nevertheless, often liposomes have limitations: they can be degradated chemically by oxidative and hydrolytic pathways of the liposomal phospholipid molecules, physically and biologically.9-11 One approach to stabilizing vesicles is to replace lipids by *To whom correspondence should be addressed. Tel: +49 421 200-3556. Fax: + 49 421 200-3249. E-mail: [email protected]. (1) Mainardes, R. M.; Silva, L. P. Curr. Drug Targets 2004, 5, 449–455. (2) El Maghraby, G. M. M.; Williams, A. C.; Barry, B. W. Int. J. Pharm. 2005, 292, 179–185. (3) Robinson, B. H.; Bucak, S.; Fontana, A. Langmuir 2000, 16, 8231–8237. (4) Schneider, M. F.; Marsh, D.; Jahn, W.; Kloesgen, B.; Heimburg, T. Proc. Natl. Acad. Sci. U.S.A. 1999, 96, 14312–14317. (5) Walde, P.; Ichikawa, S. Biomol. Eng. 2001, 18, 143–177. (6) Lamprecht, A.; Saumet, J. L.; Roux, J.; Benoit, J. P. Int. J. Pharm. 2004, 278, 407–414. (7) Mahato, R. I. Adv. Drug Delivery Rev. 2005, 57, 699–712. (8) Kokkona, M.; Kallinteri, P.; Fatouros, D.; Antimisiaris, S. G. Eur. J. Pharm. Sci. 2000, 9, 245–252. (9) Sulkowski, W. W.; Pentak, D.; Nowak, K.; Sulkowska, A. J. Mol. Struct. 2005, 744, 737–747. (10) Grit, M.; Crommelin, D. J. A. Chem. Phys. Lipids 1993, 64, 3–18. (11) Heurtault, B.; Saulnier, P.; Pech, B.; Proust, J. E.; Benoit, J. P. Biomaterials 2003, 24, 4283–4300. (12) Mayer, C. Int. J. Artif. Organs 2005, 28, 1163–1171. (13) Funhoff, A. M.; Van Nostrum, C. F.; Lok, M. C.; Kruijtzer, J. A. W.; Crommelin, D. J. A.; Hennink, W. E. J. Controlled Release 2005, 101, 233–246. (14) Meier, W.; Graff, A.; Diederich, A.; Winterhalter, M. Phys. Chem. Chem. Phys. 2000, 2, 4559–4562. (15) Graff, A.; Winterhalter, M.; Meier, W. Langmuir 2001, 17, 919–923. (16) Ruysschaert, T.; Sonnen, A.F.-P.; Haefele, T.; Meier, W.; Winterhalter, M.; Fournier, D. J. Am. Chem. Soc. 2005, 127, 6242–6247. (17) Gomes, J.F.P.S.; Sonnen, A.F.-P.; Kronenberger, A.; Fritz, J.; Coelho, M. :: A. N.; Fournier, D.; Fournier-Noel, C.; Mauzac, M.; Winterhalter, M. Langmuir 2006, 22, 7755–7759.

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polymeric vesicles.12-16 A different concept is using liposomes as a template for a 2D polymer support inside a hydrophobic core.17 Another approach to stabilization is the use of charged polymers such as polyelectrolytes. Concerning liposomes/polyelectrolyte systems, previous work indicated an increase in the thermal and detergent stability of l-R-phosphatidic acid (DMPA) liposomes by the coverage of only one layer of polycation PAH and the increase in stability by the coverage of several layers of cross-linkable polyelectrolytes.18,19 Recent studies reported on cationic lipid vesicles coated with linear polyions and demonstrated their relevance to gene transfection.20 Investigations of the complexation of 1,2-dioleoyltrimethylammoniumpropane (DOTAP) vesicles and charged linear polyions with different charge density and flexibility revealed the formation of large aggregates close to charge neutralization in a lowdensity colloidal system.21 The ionic conductivity of lipid polyelectrolyte composed capsules can be controlled by the presence of cholesterol.22 Recently, Michel and co-workers described the possibility of building a submicrometer reactive system composed of large unilamellar lipid vesicles encapsulating active species embedded/ immobilized within polyelectrolyte multilayers in order to enhance their loading capacity and protection.23 The concept of stepwise adsorption of opposite charged polyelectrolytes onto a surface was introduced by Decher et al. and is known as the layer-by-layer (LbL) technique.24-27 Later, this approach was implemented on a colloidal spherical core surface that can be subsequently decomposed after the polymer coating.16,22,28,29 Such polymer hollow shells are permeable to :: (18) Ge, L. Q.; Mohwald, H.; Li, J. B. Colloids Surf., A 2003, 221, 49–53. (19) Germain, M.; Grube, S.; Carriere, V.; Richard-Foy, H.; Winterhalter, M.; Fournier, D. Adv. Mater. 2006, 18, 2868–2871. (20) Ciani, L.; Ristori, S.; Salvati, A.; Calamai, L.; Martini, G. Biochim. Biophys. Acta 2004, 1664, 70–79. (21) Sennato, S.; Bordi, F.; Cametti, C.; Di Biasio, A.; Diociaiuti, M. Colloids Surf., A 2005, 270-271, 138–147. :: (22) Georgieva, R.; Moya, S. E.; Baumler, H.; Mohwald, H.; Donath, E. J. Phys. Chem. B 2005, 109, 18025–18030. (23) Michel, M.; Arntz, Y.; Fleith, G.; Toquant, J.; Haikel, Y.; Voegel, J.-C.; Schaaf, P.; Ball, V. Langmuir 2006, 22, 2358–2364. (24) Decher, G. Science 1997, 277, 1232–1236. (25) Dobrynin, A. V.; Rubinstein, M. Prog. Polym. Sci. 2005, 30, 1049–1118. (26) Rojas, O. J.; Claesson, P. M.; Berglund, K. D.; Tilton, R. D. Langmuir 2004, 20, 3221–3230. (27) Ubbink, J.; Khokhlov, A. R. J. Chem. Phys. 2004, 120, 5353–5365. (28) Vinogrova, O. I.; Lebedeva, O. V.; Kim, B.-S. Anuu. Rev. Mater. Res. 2006, 36, 147–178. (29) Zhu, Y. H.; Yang, X. L.; Li, P. L.; Ying, H. Prog. Chem. 2003, 15, 512–517.

Published on Web 03/24/2009

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Figure 1. Scheme of magnetic liposomes coated with polyelectrolyte layers with opposite charges.

small (ca. 1 to 2 nm) polar molecules but are more stable against chemical and physical influences. Parameters such as size, surface charge, membrane fluidity and stability, and the presence of coupling groups on the surface can be used to design the carrier to be adapted to a wide range of experimental conditions. This approach provides several advantages such as the protection of the encapsulated substance as an enzyme or drug.29 The permeability of the polyelectrolyte shell membrane can be modified by pH change or by magnetic field modulation when light-adsorbing magnetic particles, such as ferromagnetic gold-coated cobalt (Co@Au) nanoparticles are incorporated inside the polymer shell.30 Permeability control is extremely important for the use of this polymer capsule as a delivery vehicle. Macromolecules can be encapsulated and can be delivered as a result of the pH dependence of the permeability of the polymer.31 Upon irradiation or pH change, the microcapsules will release the encapsulated macromolecules. The shell permeability can also be modified by the coverage of lipid molecules as an outside layer. The outer capsule wall can be functionalized with fused virions and recrystallized S layers.32 For targeting, ligands such as antibodies can be attached to the lipid molecules.31 The encapsulation/incorporation of magnetic nanoparticles in lipid vesicles, called magnetic liposomes, was currently employed in drug delivery,33,34 such as toxin delivery systems in tumor tissues, as mediators of intercellular hyperthermia,35 and as a separation technique for proteins by high gradient magnetic filtration.36 These vesicles can guide and transport biologically active substances, allowing an optimum therapeutic concentration of the drug in the desired tissue of the organism, while keeping the total injection dose low, providing site specificity and selectivity.37-42 Magnetic liposomes have a high in vivo potential use as a magnetic resonance imaging (MRI) contrast. These vesicles encapsulating the magnetic contrast agents avoid the local dilution of those agents and their interactions with the (30) Hodenius, M.; De Cuyper, M.; Desender, L.; Muller-Schulte, D.; Steigel, A.; Lueken, H. Chem. Phys. Lipids 2002, 120, 75–85. (31) Angelatos, A. S.; Radt, B.; Caruso, F. J. Phys. Chem. B 2005, 109, 3071– 3076. (32) Moya, S. E.; Toca-Herrera, J. L. J. Nanosci. Nanotechnol. 2006, 6, 2329– 2337. (33) Jain, S.; Mishra, V.; Singh, P.; Dubey, P. K.; Saraf, D. K.; Vyas, S. P. Int. J. Pharm. 2003, 261, 43–55. (34) Kuznetsov, A. A.; Filipov, V. I.; Alyautdin, R. N.; Torshina, N. L.; Kuznetsov, O. A. J. Magn. Magn. Mater. 2001, 225, 95–100. (35) Hotani, H.; Inaba, T.; Nomura, F.; Takeda, S.; Takiguchi, K.; Itoh, T. J.; Umeda, T.; Ishijima, A. Biosystems 2003, 71, 93–100. (36) Saiyed, Z.; Telang, S.; Ramchand, C. Biomagn. Res. Technol. 2003, 1, 2. (37) Elmi, M. M.; Sarbolouki, M. N. Int. J. Pharm. 2001, 215, 45–50. (38) Giri, J.; Thakurta, S. G.; Bellare, J.; Nigam, A. K.; Bahadur, D. J. Magn. Magn. Mater. 2005, 293, 62–68. (39) Grabielle-Madelmont, C.; Lesieur, S.; Ollivon, M. J. Biochem. Biophys. Methods 2003, 56, 189–217. (40) Freisleben, H. J.; Zwicker, K.; Jezek, P.; John, G.; Bettin-Bogutzki, A.; Ring, K.; Nawroth, T. Chem. Phys. Lipids 1995, 78, 137–147. (41) Menager, C.; Cabuil, V. J. Phys. Chem. B 2002, 106, 7913–7918. (42) Sangregorio, C.; Wiemann, J. K.; O’Connor, C. J.; Rosenzweig, Z. J. Appl. Phys. 1999, 85, 5699–5701. (43) Martina, M. S.; Fortin, J. P.; Menager, C.; Clement, O.; GrabielleMadelmont, C.; Gazeau, F.; Cabuil, V.; Lesieur, S. J. Am. Chem. Soc. 2005, 127, 10676–10685.

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biological environment and may combine both diagnosis and therapeutics.43 Liposomes are filled with a ferrofluid that is composed of superparamagnetic particles dispersed in a carrier fluid. Lesieur et al. described a method of producing large unilamellar liposomes encapsulating maghemite nanoparticles and have shown the surfactant-induced formation of transient pores in the liposomes bilayer.44 The surface modification of the magnetic nanoparticles by organic molecules allows their stabilization in a biological suspension at physiological pH and high salt concentrations and also provides functional groups at the surface for further use.36,38,45,46 Here we combined both nanometer-sized liposomes with polyelectrolyte shells to achieve highly stable and selective nanocapsules. The aim is to protect the lipid bilayer with a polymer shell by covering the liposomes with one or more layers of polyelectrolytes to maintain the liposomes intact and to obtain biologically and chemically stable nanometer-sized capsules. The method that is implemented consists of the adsorption of an outer shell of polyelectrolytes of opposite charges on the surface of the liposome by the LbL technique. To separate the polyelectrolyte-coated liposomes from the free polyelectrolyte, magnetic nanoparticles were encapsulated. This approach, which is schematically shown in Figure 1, is straightforward and very efficient. The results are discussed in terms of the size, shape, and morphology of the liposomes/polyelectrolyte nanocapsules and their permeability. As a possible application, these polyelectrolyte-coated unilamellar magnetic liposomes could be delivered to living cells by electroporation or microinjection, allowing us to study, for example, intracellular pH changes by confocal microscopy.47 In addition, because these liposomes are both charged and magnetic they can be easy manipulated by an external magnetic or electric field.

Experimental Section Materials. Egg yolk L-R-phosphatidylcholine (egg-PC, MW = 760.08 g/mol) was purchased from Lipoid (Ludwigshafen, Germany), and L-R-phosphatidylethanolamine-N-(lissamine rhodamine B sulfonyl) (ammonium salt) (PE-lissamine RhodB, MW = 1301.73 g/mol) was provided by Avanti Polar Lipids Inc. (Alabaster, AL). Poly(sodium 4-styrenesulfonate) (PSS, MW = 70 000 g/mol), poly(allylamine hydrochloride) (PAH, MW = 70 000 g/mol), poly(fluorescein isothiocyanate allylamine hydrochloride) (FITC-PAH, MW = 3355 g/mol), sepharose-4B, chloroform (CHCl3, MW = 119.38 g/mol), sodium citratre tribasic dihydrate (HOC(COONa)(CH2COONa)2 3 2H2O, MW = 294.13 g/mol), calcein (C30H26N2O13, MW = 622.53 g/mol,) and Triton X-100 (TX-100, 4-(C8H17) C6H4(OCH2CH2)nOH, n ≈ 10) were obtained from SigmaAldrich (Munich, Germany). Ferrous chloride (FeCl2, (44) Lesieur, S.; Grabielle-Madelmont, C.; Menager, C.; Cabuil, V.; Dadhi, D.; Pierrot, P.; Edwards, K. J. Am. Chem. Soc. 2003, 125, 5266–5267. (45) Neuberger, T.; Schopf, B.; Hofmann, H.; Hofmann, M.; von Rechenberg, B. J. Magn. Magn. Mater. 2005, 293, 483–496. (46) Gupta, A. K.; Gupta, M. Biomaterials 2005, 26, 3995–4021. (47) Ramaye, Y. To be submitted for publication.

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Figure 2. Picture of the magnetic separation of polyelectrolytecoated magnetic liposomes from free electrolyte. (A) Polyelectrolyte-coated magnetic liposomes and free polyelectrolyte with a 0.7 T rare earth magnet. (B) Separation of the magnetic pellet (magnetic liposomes) and supernatant (free polyelectrolyte), t < 1 min. MW = 126.75 g/mol), ferric chloride (FeCl3, MW = 162.20 g/ mol), ferric nitrate (Fe(NO3)3, MW = 241.88 g/mol), ammonium hydroxide (NH4OH, MW = 35.05 g/mol), sodium chloride (NaCl, MW = 58.44 g/mol), and the Tris buffer 2amino-2-hydroxymethyl-1,3-propanediol (NH2C(CH2OH)3, MW = 121.14 g/mol) were purchased from Fluka (Seelze, Germany). Magnetic Fluid Preparation. The magnetic fluid was constituted of citrate-coated maghemite magnetic particles (citrateγ-Fe2O3), which was 1 M in Fe concentration. The method of synthesis was the coprecipitation method, first described in 1981 by Massart.48,49 Aqueous solutions of ferric and ferrous salts (FeCl2 and FeCl3), Fe(III)/Fe(II) with a molar ratio of 2, are mixed with an alkali solution of ammonium hydroxide (NH4OH) to induce precipitation. Then the precipitate, magnetite (Fe3O4), is separated from the solution by magnetic separation. Afterward, magnetite is oxidized to maghemite by ferric nitrate (1.3 mol of Fe(NO3)3) in nitric acid solution (1 L, 2 N HNO3) under boiling. As these particles aggregate in a pH range of 5-9, they are coated with citrate ions, and a stable ionic ferrofluid solution at pH 7 is obtained. For this, the maghemite precipitate is dispersed in a sodium citrate solution (0.25 M), held for 30 min at 80 °C, and finally precipitated in acetone at room temperature. Finally, the particles are separated from acetone by magnetic separation and redispersed in water. The magnetic nanoparticles have superparamagnetic behavior and a mean size of 8 nm.39,41 After the external magnetic field is removed, the magnetic particles can be resuspended in an aqueous medium without aggregation and changes in their initial magnetic properties. This behavior is essential for magnetic separation techniques. Magnetic Liposome Preparation. Magnetic liposomes were prepared by the thin film hydration method. Lipids in chloroform stock solutions were mixed and dried. Five milligrams of a 50 mg/mL egg-PC stock solution and 10 μL of a 2.5 mg/mL PElissamine RhodB stock solution in chloroform were mixed in a Pyrex vial. The lipid film was made by removing the chloroform under a stream of purified nitrogen or argon flux. To remove traces of chloroform, the vial was placed under vacuum for at least 3 h. The dried lipid film was then redispersed in 200 μL of magnetic fluid in order to encapsulate the magnetic nanoparticles inside the lipid vesicles. The suspension was homogenized by vortex mixing. Then 800 μL of buffer (10 mM Tris, 2 mM NaCl, pH 7.3) or 800 μL of the fluorescent probe calcein at 50 mM in the same buffer was added to the suspension, followed again by vortex mixing. Highly polydisperse multilamellar vesicles were (48) Massart, R. IEEE Trans. Magn. 1981, 17, 1247–1248. (49) Massart, R.; Dubois, E.; Cabuil, V.; Hasmonay, E. J. Magn. Magn. Mater. 1995, 149, 1–5.

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Figure 3. TEM picture of magnetic citrated nanoparticles. obtained. The hydration method was followed by 10 freeze (liquid nitrogen)-thaw (37 °C) cycles to achieve a suspension of unilamellar but polydisperse vesicles. A highly monodisperse unilamellar vesicle solution was obtained by applying the extrusion method with a miniextruder (Avanti Polar Lipids). The suspension was passed 10 times through a 200 nm pore size polycarbonate nucleopore filter (Millipore). Nonentrapped magnetic nanoparticles were removed by size exclusion chromatography (SEC). The suspension was poured into an 8  500 mm2 Sepharose-4B column washed and eluted with the liposome preparation buffer (10 mM Tris, 2 mM NaCl, pH 7.3). Fractions of 500 μL were collected, and the size distribution of these fractions was characterized by dynamic light scattering. Polyelectrolyte Coating. Polyelectrolyte-coated magnetic liposomes were prepared by using the LbL method associated with a magnetic separation technique. The polyelectrolytes used were PAH as the polycation and PSS as the polyanion. First, 500 μL of a 10 mg/mL aqueous solution of PAH was added to 500 μL of a magnetic liposomes suspension in a 2 mL microtest Eppendorf tube. Then the suspension was vortex mixed vigorously. To separate the PAH-coated magnetic liposomes from the free PAH, magnetic separation was performed with the help of a 0.7 T rare earth magnet for less than 1 min (Figure 2). Then the supernatant (nonmagnetic - free PAH) was removed, and the pellet (magnetic - PAH-coated magnetic liposomes) was redispersed in 1 mL of buffer solution (10 mM Tris, 2 mM NaCl, pH 7.3). Afterwards, a second layer of PSS was absorbed using a similar procedure to that used for the first layer of PAH coating. A 10 mg/mL aqueous solution (500 μL) of PSS was added to 500 μL of a PAH-coated magnetic liposomes suspension in a 2 mL microtest Eppendorf tube prior to vortex mixing. Nonabsorbed PSS was removed using magnetic separation. Layer forming continued until four layers of PAH/PSS were absorbed on the surface. Transmission Electron Microscopy (TEM). A drop of a diluted solution of the citrated magnetic nanoparticles was deposited on the carbon membrane of a copper grid (Plano, Germany). After drying, the grid was transferred to a transmission electron microscope (Zeiss EM 900, Oberkochen), and the sample was examined at an accelerating voltage of 80 kV at room temperature. Magnetic Measurements. Magnetic measurements of the citrated magnetic nanoparticles were performed with a vibrating DOI: 10.1021/la9003142

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Figure 4. Average hydrodynamic radius (average ( standard deviation, n = 10) of magnetic egg-PC liposomes, extruded with a 200 nm pore filter, as determined by dynamic light scattering measurements. (A) Uncoated liposomes, (B) (PAH/PSS)-coated liposomes, (C) (PAH/PSS/ PAH)-coated liposomes, and (D) (PAH/PSS)2-coated liposomes.

sample magnetometer (Montage Foner) at LI2C (Laboratoire des Liquides Ioniques Interfaces Chargees, Paris VI, France). Dynamic Light Scattering (DLS) and Zeta-Potential Measurements. The size and the surface charge of our particles were characterized by dynamic light scattering (ZetaSizer Nano Series, Nano ZS, Malvern Instruments). To measure the size of the lipid/polyelelectrolyte vesicles, 50 μL of sample and 400 μL of buffer (10 mM Tris, 2 mM NaCl, pH 7.3, carefully degassed) were poured into a semimicro polystyrene cuvette (Sarstedt, Germany), and the suspension was homogenized prior to measurement. The zeta potential of the lipid/polyelelectrolyte vesicles was investigated using 10 μL of sample and 740 μL of buffer (10 mM Tris, 2 mM NaCl, pH 7.3) in the standard cuvette provided by Malvern. Cryogenic-Transmission Electron Microscopy (CryoTEM) Measurements. Cryogenic-transmission electron microscopy is ideal for visualizing colloidal nanostructures. It is suited to the visualization of structures in the size range of 5-500 nm, and detailed information about local amphiphilic nanostructures in solution can be obtained. A drop of the sample was placed on a copper grid (Quantifoil S7/2, Jena). The excess liquid was removed with filter paper, leaving a sample film stretched over the holes of the grid. The samples were immediately shock frozen by shooting them into liquid ethane at 90 K. After the ethane was blotted, the samples were transferred to the microscope (TEM LEO 912 Omega, Zeiss Oberkochen) with a liquid-nitrogen-cooled transfer device. The specimens were viewed in the transmission electron microscope device at 90 K with an accelerating voltage of 120 kV under zero-loss conditions. Atomic Force Microscopy (AFM) Measurements. The atomic force microscope can be used to image particles on surfaces and characterize them in terms of size, morphology, surface texture, and roughness. A Multimode atomic force microscope (Veeco, Mannheim, Germany) was used together with a Nanoscope IIIa controller to image the polyelectrolyte-coated magnetic liposomes in buffer solution. For all samples, a 40 μL portion of a polyelectrolyte-coated magnetic liposomes solution was incubated for 30 min on freshly cleaved mica and then rinsed five times with 100 μL of buffer solution. To improve the immobilization of the negatively charged polymer liposomes on a negatively charged surface, the mica was pretreated with positively charged divalent ions in a 200 mM MgCl2 solution. The best and most stable imaging conditions were obtained in contact mode in buffer on mica using a fluid cell sealed with an O-ring. Other imaging conditions, that is, tapping mode in liquid or imaging in air, did not produce successful results. The imaging cantilever was an unmodified silicon nitride tip immersed in buffer solution with a spring constant of 0.03 or 0.1 N/m; applied scanning forces varied between 150 and 500 pN. 6796 DOI: 10.1021/la9003142

Figure 5. Zeta potential (average ( standard deviation, n = 10) of PAH/PSS multilayers on magnetic liposomes as a function of the number of layers. Fluorescence Measurements. Leakage tests of the polyelectrolyte-coated magnetic liposomes were performed with a self-quenching calcein assay.17 The release of calcein is initiated by the addition of 0.1% Triton X-100 solubilizing the liposome membranes. The dilution of released calcein increased the fluorescence intensity. Fluorescence measurements were performed in a Varian Cary Eclipse fluorescence spectrophotometer at excitation at 495 nm in buffer (10 mM Tris, 2 mM NaCl, pH 7.3). Because of the sedimentation of magnetic liposomes caused by the magnetic stirring system of the fluorimeter, kinetic experiments were not performed.

Results and Discussion Coprecipitation of iron salts is a well-known and well-characterized method of obtaining magnetic nanoparticles. As expected, the TEM picture (Figure 3) showed nanoparticles with a large size distribution (average size 8 nm), and the nanoparticles showed superparamagnetic behavior. (For additional information, see Supporting Information.) Size Distribution. Figure 4 shows the average hydrodynamic radius (average ( standard deviation) measured by dynamic light scattering of 200 nm extruded magnetic liposomes before and after the deposition of one, two, three, and four polyelectrolyte layers. The average and standard deviation values were calculated from the experimental data of 10 experiments. EggPC magnetic liposomes show an average size of 164 nm prior to coating. Although the average value differed slightly from batch to batch, coating with PAH leads always to a more narrow distribution in the range of 140-300 nm. It is interesting that the Langmuir 2009, 25(12), 6793–6799

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Figure 7. (A) Representative contact-mode AFM topography of PAH-coated magnetic egg-PC liposomes. (B) Cross-section of the same sample.

Figure 6. Cryo-TEM micrographs of magnetic egg-PC liposomes. (A) Uncoated liposomes, (B) (PAH/PSS)-coated liposomes, (C) (PAH/PSS/PAH)-coated liposomes, and (D) (PAH/PSS)2-coated liposomes.

polyelectrolyte coating as well as the presence of magnetic nanoparticles changes the refractive index, which might modify the size revealed from the DLS measurements. The adsorption of another polyelectrolyte layer does not vary the average size distribution. After the adsorption of (PAH/PSS/PAH) and (PAH/PSS)2, the average size of the liposome/polyelectrolyte vesicles also does not change significantly. Assembly of Polyelectrolytes Multilayers. The assembly of (PAH/PSS)4 multilayers on magnetic liposomes was characterized by zeta potential measurements before and after each polyelectrolyte adsorption step, and the results are summarized in Figure 5. The initial value of approximately 0 mV corresponds to the uncovered egg-PC magnetic liposomes. The zeta-potential average value for the first layer (31 mV) corresponds to the Langmuir 2009, 25(12), 6793–6799

adsorption of the polycation PAH, the second (-40 mV) corresponds to the adsorption of the polyanion PSS, the third (42 mV) corresponds to another layer of PAH, and the fourth zeta-potential average value (-37 mV) corresponds to a new layer of PSS. Figure 6 shows alternating values of the zeta potential as expected for polyelectrolyte stepwise adsorption. The alternating zeta potential depending on whether vesicles are covered with PAH or PSS as outer layers demonstrates a successful stepwise adsorption. The adsorption of polyelectrolytes onto charged or neutral surfaces is mainly driven by electrostatic attraction and the entropically favored release of the counterions.50,51 Other forces such as van der Waals attraction or hydrogen bonding may contribute.51 The quality of the coating can be readily tested using the zeta potential. Structure of the Polyelectrolyte-Coated Magnetic Liposomes. Cryo-TEM revealed spherically shaped liposomes before and after coating. The identification of magnetic nanoparticles inside the vesicles indicates a successful encapsulation (Figure 6A) as seen inside the lipid vesicles. After the first two coated layers (second layer of PSS), the liposomes were of similar sizes and had a homogeneous cover absorbed on the surface (Figure 6B). Because of the sample preparation for cryo-TEM, some magnetic liposomes can be damaged, leading to the release of magnetic particles to the surrounding medium (Figure 6B). In Figure 6D, the liposomes were coated with four layers (fourth-layer PSS). The morphology was quite similar to that of the two-layer-coated magnetic liposomes, and the size did not change significantly. However, they seemed to superpose each other. A possible explanation can be the sample preparation for cryo-TEM studies. These structures were also studied by AFM, and the results show structures with similar dimensions. (50) Kotov, N. A. Nanostruct. Mater. 1999, 12, 789–796. (51) Laugel, N.; Betscha, C.; Winterhalter, M.; Voegel, J. C.; Schaaf, P.; Ball, V. J. Phys. Chem. B 2006, 110, 19443–19449.

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Figure 8. Representative fluorescence traces. (A) Egg-PC liposomes before and after membrane disruption with 0.1% TX-100. (B) (PAH/ PSS)2-coated egg-PC magnetic liposomes before and after addition of 0.1% TX-100.

The AFM images generally confirm the DLS measurements (Figure 7). The images show spherical polymer-coated vesicles. Their diameters range from 35 to 260 nm whereas their respective heights vary from 5 to 150 nm. When using AFM to determine the vesicle size, it is important to consider the vesicle-substrate interaction, the tip shape, and the applied scanning force that may influence the results (e.g., flatten the liposomes). Therefore, the detected height and diameter might differ between surface-based methods (AFM) and solutionbased methods (DLS). Furthermore, it was not possible to distinguish between the different states of polymer coating because the thickness of the polymer coating is negligible to the size variation of the template liposomes. Stability Tests Against Detergent and Phospholipases. Stability tests in the presence of detergent Triton TX-100 and phospholipases revealed no significant changes on the four-layer-polyelectrolyte-coated magnetic liposomes. Both detergent/surfactant molecules and the phospholipases enzymes could not penetrate through the polyelectrolyte shell that covers the magnetic liposomes, thus no hydrolysis of the phospholipid molecules occurred. To evaluate the integrity of the polyelectrolyte-coated magnetic liposomes in the presence of TX-100, fluorescence studies were performed. Figure 8 shows the fluorescence of liposomes with encapsulated calcein: egg-PC liposomes (Figure 8A) and (PAH/PSS)2-coated egg-PC magnetic liposomes (Figure 8B) before and after the addition of 0.1% 6798 DOI: 10.1021/la9003142

TX-100. Obviously, calcein leakage occurs only during lipid membrane disruption. After lipid shell disruption by the presence of TX-100, calcein is released. The calcein concentration is decreased, leading to an increase in fluorescence intensity. This is illustrated clearly in Figure 8A, with an increase of 63% in the fluorescence intensity after the addition of the detergent. The same behavior could not be seen for the (PAH/PSS)2-coated eggPC magnetic liposomes (Figure 8B). In fact, no significant changes were observed after the addition of the detergent. The two layers ((PAH/PSS)2) of polyelectrolytes are sufficient to protect the lipid shell against membrane disruption by detergents.

Conclusions The present investigation demonstrates a simple method for polyelectrolyte coating of nanometer-sized unilamellar liposomes by the stepwise adsorption of oppositely charged polyelectrolytes PAH and PSS. In this context, superparamagnetic nanoparticles were encapsulated in the liposomes prior to polyelectrolyte coating. This encapsulation permits the separation of polyelectrolyte-coated magnetic liposomes from the free polyelectrolyte. Cryo-TEM studies revealed the morphological change of the outside layer of the polyelectrolyte-coated magnetic liposomes compared with that of the noncoated magnetic liposomes. This change is attributable to the polymer layer(s). Cryo-TEM and AFM investigations did not reveal any significant changes in size and shape after layer deposition. These structures are spherical with average sizes ranging from approximately 200 to 400 nm. Langmuir 2009, 25(12), 6793–6799

Pereira da Silva Gomes et al.

The encapsulation of superparamagnetic nanoparticles does not disrupt or modify the lipid membrane structure. Stability tests with Triton TX-100 have shown that two polyelectrolyte layers are sufficient to protect lipid vesicles. This detergent cannot pass through the (PAH/PSS) layer and disrupt the lipid bilayer. Our results show that the presence of a polyelectrolyte shell together with the encapsulated magnetic nanoparticles is a promising approach to the simple manipulation of nanocapsules by electric and/or magnetic fields.

Langmuir 2009, 25(12), 6793–6799

Article

Acknowledgment. We thank Delphine Talbot from LI2C (Laboratoire des Liquides Ioniques Interfaces Chargees, Paris VI, France) for magnetic measurements. This research was supported by the Volkswagen Foundation (project I/80 051-054). Supporting Information Available: AFM data and magnetic curve. This material is available free of charge via the Internet at http://pubs.acs.org.

DOI: 10.1021/la9003142

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