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Stable Polymethacrylate Nanocapsules from Ultraviolet Light-Induced Template Radical Polymerization of Unilamellar Liposomes Joana Filipa Pereira da Silva Gomes,*,†,‡, Andreas F.-P. Sonnen,‡,⊥ Astrid Kronenberger,‡ Ju¨rgen Fritz,‡ Manuel A Ä lvaro Neto Coelho,§ Didier Fournier,† Clara Fournier-No¨el,# Monique Mauzac,# and Mathias Winterhalter†,‡ Institut de Pharmacologie et Biologie Structurale CNRS UMR5089, Toulouse F-31077, France, International UniVersity Bremen, Campus Ring 1, D-28759 Bremen, Germany, Laborato´ rio de Engenharia de Processos, Ambiente e Energia, Faculdade de Engenharia da UniVersidade do Porto, Porto, Portugal, The DiVision of Structural Biology, UniVersity of Oxford, RooseVelt DriVe, OX3 7BN, United Kingdom, and Laboratoire Interactions Moleculaires et ReactiVite´ Chimique et Photochimique UPS/CNRS UMR5623, Toulouse, France ReceiVed May 12, 2006. In Final Form: June 14, 2006 We employed UV-induced template polymerization to create hollow nanometer-sized polymer capsules. Homogeneous, unilamellar liposomes served as a two-dimensional template for the cross-linking of either butyl methacrylate or hydroxyethyl methacrylate with the bifunctional ethyleneglycol dimethacrylate. Different molar ratios of lipid/hydrophobic monomer/bifunctional monomer/photoinitiator were tested and dynamic light scattering revealed negligible changes of size at a defined molar ratio of 2/1/10/20, respectively. Cryo-transmission electron microscopy provided clear evidence that incorporation of the methacrylate monomers into and polymerization in the hydrophobic bilayer phase does not disrupt vesicle integrity. Moreover, after solubilization of the lipids, the polymethacrylate nanocapsules were stable at conditions needed for negative staining and could be visualized by atomic force microscopy. In contrast to previous findings, the nanocapsule size and shape did not change considerably after removal of the template phase, and the size distribution remained strictly monomodal. The employed method is not only an advance to fortify liposomes, but the nanocapsules themselves can be functionalized.
1. Introduction Hollow nanometer-sized containers are of increasing interest in nanotechnology, since they can protect proteins, enzymes, or drugs from hostile surroundings and provide an optimal microenvironment different from the bulk medium. Such nanocontainers may be used in drug delivery, in medical diagnostics, or as intracellular reporters. Drugs or enzymes may be hidden from the outside, protected against chemical and biological degradation, targeted to specific cells, and released in a controlled manner.1 Liposomes made from natural or synthetic lipids are a typical and widespread example of such nanocontainers. They are closed, vesicle-like aggregates usually composed of phospholipids or surfactants. When perforated, they close rapidly, since the energy loss associated with opening of the lipid bilayer is far greater than the thermal energy gain. As a functional material, liposomes can carry hydrophilic as well as lipophilic cargo, their size can be easily adjusted, and they are biocompatible.1-3 However, liposomes are not suited for many material applications because of their sensitivity toward environmental factors such as pH changes, osmotic stress, lipases, and the presence of detergents. Many different approaches have been devised to fortify liposomes. Notable examples are the use of polymerizable sur* To whom correspondence should be addressed. Telephone: +494212003151; fax: +494212003249; e-mail:
[email protected]. † Institut de Pharmacologie et Biologie Structurale. ‡ International University of Bremen. § Universidade do Porto. ⊥ University of Oxford. # Laboratoire Interactions Moleculaires et Reactivite ´ Chimique et Photochimique. (1) Heurtault, B.; Saulnier, P.; Pech, B.; Proust, J.-E.; Benoit, J.-P. Biomaterials 2003, 24, 4283-4300. (2) Graff, A.; Winterhalter, M.; Meier, W. Langmuir 2001, 17, 919-923. (3) Antonietti, M.; Forster, S. AdV. Mater. 2003, 15, 1323-1333.
factants, the incorporation of polymers during the formation of vesicles, and surface grafting with water-soluble polymers.4-13 Template polymerization in surfactant phases is a versatile method in polymer chemistry to obtain ordered nanostructured materials. The general idea is to turn a dynamic, self-organized molecular assembly into a mechanically and chemically stable supramolecular material. In the case of so-called direct templating, the morphology of the polymeric product resembles the structure of the template.1,10,14 The appealing character of these reactions is explained by the rather simple concept to construct ordered phases: a lyotropic liquid crystalline phase as a template is loaded with monomer molecules, and, subsequently, a polymerization reaction is initiated with the intention to produce a polymeric material that, in an ideal case, would preserve and stabilize the structure of the employed template.1 With liposomes as templates, hydrophobic monomers can be dissolved in the hydrophobic part of the bilayer, and their radical (4) Ruysschaert, T.; Sonnen, A. F. P.; Haefele, T.; Meier, W.; Winterhalter, M.; Fournier, D. J. Am. Chem. Soc. 2005, 127, 6242-6247. (5) Poulain, N.; Nacjache, E.; Pina, A.; Levesque, G. J. Polym. Sci., Part A: Polym. Chem. 1996, 34, 729-737. (6) Meier, W.; Graff, A.; Diederich, A.; Winterhalter, M. Phys. Chem. Chem. Phys. 2000, 2, 4559-4562. (7) McKelvey, C. A.; Kaler, E. W.; Zasadzinski, J. A.; Coldren, B.; Jung, H.-T. Langmuir 2000, 16, 8285-8290. (8) Kurja, J.; Nolte, R. J. M.; Maxwell, I. A.; German, A. L. Polymer 1993, 34, 2045-2049. (9) Krafft, M. P.; Schieldknecht, L.; Marie, P.; Giulieri, F.; Schmutz, M.; Poulain, N.; Nakache, E. Langmuir 2001, 17, 2872-2877. (10) Hentze, H.-P.; Kaler, E. W. Curr. Opin. Colloid Interface Sci. 2003, 8, 164-178. (11) Decker, C. Prog. Polym. Sci. 1996, 21, 593-650. (12) Co, C. C.; Cotts, P.; Burauer, S.; de Vries, R.; Kaler, E. W. Macromolecules 2001, 34, 3245-3254. (13) Bronich, T. K.; Ouyang, M.; Kabanov, V. A.; Eisenberg, A.; Szoka, F. C., Jr.; Kabanov, A. V. J. Am. Chem. Soc. 2002, 124, 11872-11873. (14) Jung, M.; Hubert, D. H. W.; Bomans, P. H. H.; Frederik, P.; van Herk, A. M.; German, A. L. AdV. Mater. 2000, 12, 210-213.
10.1021/la0613575 CCC: $33.50 © 2006 American Chemical Society Published on Web 07/28/2006
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polymerization can be induced by UV light, leading to the formation of a polymer network. The main advantage of using UV radiation to initiate the chain reaction lies in the very high polymerization rates that can be reached under intense irradiation, so that the liquid-to-solid phase transition, that is, the monomerto-polymer transition, takes place within a fraction of a second. Longer time intervals would promote the phase separation of lipids and polymerizable monomers. Only a homogeneous distribution of mono- and bifunctional monomers within the lipid bilayer can lead to a polymer hollow sphere. An incoherent distribution of monomers or an excess of the monofunctional composite will not feature a closed polymer shell. The netlike polymer scaffold inside the vesicle bilayer does not restrict the lateral mobility of the lipids and the transbilayer diffusion of low molecular weight substances, which is important for investigating the interaction of pharmaceutically active substances with biological membranes.2,6,11 Earlier studies by Meier et al. introduced new nanometer-sized bioreactors based on liposomes that could be functionalized by incorporating β-lactamase in the hydrophilic interior and the channel protein OmpF into the bilayer.2 The functionalized liposomes could be stabilized by the cross-linking polymerization of methacrylate monomers in the interior of the membranes. Notably, the enzyme and the membrane channel preserved their activity, even in the presence of these monomers and after their polymerization. In strong contrast to these findings, Jung et al. reported polymer shell structures with a morphology that did not resemble the vesicle template; instead, vesicle-polymer hybrid morphologies were obtained.14-19 The polymerization of styrene or alkyl methacrylates was not able to overcome the phase separation between polymer and surfactant, thus leading to the formation of several individual polymer loci on a vesicle. Interestingly, the polymerization of styrene within the bilayers of dioctadecyldimethylammonium bromide vesicles leads to peculiar polymer colloid morphologies: small polymer latex beads attach to the membrane to form vesicle-polymer hybrid particles, producing parachute architectures. The addition of a cross-linker or copolymerization of styrene with butyl methacrylate (BMA) resulted in the formation of several polymer beads attached to one vesicle bilayer giving a necklace-like morphology.18 Here we present a novel experimental approach to stabilize liposomes and to achieve polymerization of hydrophobic methacrylate monomers inside a liposome bilayer by a radical mechanism. The use of mono and bifunctional monomers leads to the formation of a two-dimensional (2D) polymer network, which maintains its size and shape after the removal of lipids. We elucidate the crucial factors needed to obtain stable, nanometersized polymethacrylate capsules: the optimal point of monomer addition, the molar ratio of lipid to monomer, the overall concentrations, and the intensity of UV radiation. The size-distribution was determined before and after UV polymerization as well as after lipid removal. Cryo-transmission electron microscopy (CryoTEM), negative-stain TEM and atomic force microscopy (AFM) were used to visualize the structure of the capsules at different preparative stages. (15) Jung, M.; Hubert, D. H. W.; van Veldhoven, E.; Frederik, P. M.; Blandamer, M. J.; Briggs, B.; Visser, A. J. W. G.; van Herk, A. M.; German, A. L. Langmuir 2000, 16, 968-979. (16) Jung, M.; Hubert, D. H. W.; van Veldhoven, E.; Frederik, P.; van Herk, A. M.; German, A. L. Langmuir 2000, 16, 3165-3174. (17) Jung, M.; Hubert, D. H. W.; van Herk, A. M.; German, A. L. Macromol. Symp. 2000, 151, 393-398. (18) Jung, M.; German, A. L.; Fischer, H. R. Colloid Polym. Sci. 2001, 279, 105-113. (19) Hubert, D. H. W.; Jung, M.; German, A. L. AdV. Mater. 2000, 12, 12911294.
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2. Experimental Section 2.1. Materials. Lipid egg phosphatidylcholine (egg-PC) was purchased from Avanti Polar Lipids Inc. (Alabaster, AL). BMA, hydroxyethyl methacrylate (HEMA), and the cross-linker ethyleneglycol dimethacrylate (EGDMA) were obtained from Sigma-Aldrich (Munich, Germany). The photoinitiator Irgacure 90720-22 (λoptimal ) 365 nm) was used as received from Ciba (Lampertheim, Germany). The monomers BMA, HEMA, and EGDMA were used as received, without further purification. They were not purified by liquidliquid extraction in the presence of NaOH to remove the inhibitors. The detergent Triton X-100 (TX-100) was obtained from Merck (Darmstadt, Germany). 2.2. Preparation of Lipid/Polymer Capsules. A chloroform solution containing 12.6 mol of BMA, 1.26 mol of EGDMA, and 0.63 mol of Irgacure 907 was added to 6.25 mol of egg-PC in chloroform solution. Chloroform was evaporated to form a lipid/ monomer film on the wall of a glass tube using a flux of purified nitrogen or argon. Further traces of solvent were removed by drying the film under vacuum for at least 2 h. The dried film was dispersed in a buffer containing 10 mM Tris-HCl and 100 mM NaCl at pH 7.4, giving a dispersion of multilamellar, polydisperse vesicles. Unilamellar but still polydisperse vesicles were obtained after 10 freeze (liquid nitrogen)-thaw (27 °C water bath) cycles, and the extrusion method was used to calibrate and control the size of the vesicles. The suspension was passed 10 times through a polycarbonate Nucleopore filter (Millipore) of 200 nm pore size, followed by a 2 h long agitation. Liposomes only composed of egg-PC were prepared in the same manner. Prior to UV polymerization, oxygen was removed by bubbling purified nitrogen or argon through the solution. Radical polymerization took place during a 3 h long incubation with UV light (λ ) 350 nm) in a photochemical reactor equipped with a constant rotation device (16 lamps of 6 W power each; the distance of the lamps to the sample was 6 cm). 2.3. Dynamic Light Scattering. Dynamic light scattering (DLS) measurements were performed in a DynaPro molecular sizing instrument (Protein Solutions Inc., USA). 2.4. Cryo-TEM and Negative-Stain TEM. Control liposomes composed only of the template lipids, liposomes containing the polymerized monomers, and the hollow nanocapsules were imaged on a Technai F30 microscope operated at 300 kV under liquid nitrogen temperatures. The control liposomes and liposomes with the polymerized 2D network were vitrified in liquid ethane on lacey carbon-coated copper grids and imaged at defoci between 1.5 and 3.0 µm to enhance the contrast. The polymeric nanocapsules were stained with 2% uranyl acetate. For this, 4 µL of the sample was incubated for 30 s on a standard carbon-coated copper grid, which was then placed upside-down onto 150 µL filtered stain drops (sample side facing the stain). Two short 3 s long incubations on clean drops preceded the actual 30 s long staining incubation to remove most of the Triton-solubilized lipids. 2.5. Atomic Force Microscopy. A Multimode atomic force microscope (AFM) (Veeco, Mannheim, Germany) was used together with a Nanoscope III controller to image the nanocapsules under ambient conditions. A 10 µL portion of a solution of nanocapsules was incubated for 10 min on freshly cleaved mica, then rinsed three times with buffer and three times with deionized water, and finally dried in a weak flow of nitrogen. The best and most stable imaging conditions were obtained in contact mode in air on mica. Other imaging conditions, that is, tapping mode in liquid, or substrates, that is, highly oriented graphite, could not produce successful results. The imaging cantilever was an unmodified silicon nitride tip with a spring constant of 0.01 N/m. (20) Masson, F.; Decker, C.; Andre, S.; Andrieu, X. Prog. Org. Coat. 2004, 49, 1-12. (21) Masson, F.; Decker, C.; Jaworek, T.; Schwalm, R. Prog. Org. Coat. 2000, 39, 115-126. (22) Moon, J. H.; Shul, Y. G.; Han, H. S.; Hong, S. Y.; Choi, Y. S.; Kim, H. T. Int. J. Adhes. Adhes. 2005, 25, 301-312.
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Figure 1. Scheme of nanocapsule formation. The template liposome with homogeneously distributed monomers is irradiated with UV light, resulting in the formation of a fortified liposome. After lipid removal, the 2D polymer network constitutes an intact hollow nanocapsule.
Figure 2. Scheme of the UV-induced radical polymerization of BMA, HEMA, and EGDMA. Also shown is the chemical structure of the photoinitiator Irgacure 907.
3. Results and Discussion The formation of hollow polymethacrylate nanocapsules is essentially a two-step process with three distinct stages (Figure 1). BMA, HEMA, and EGDMA (Figure 2) incorporate as monomers into the hydrophobic part of the bilayer and polymerize to a 2D closed network upon intensive UV irradiation. The nature of the individual stages can be characterized by standard biophysical techniques and is an indication of the success of the preparative method. 3.1. Formation of Hollow Polymer Nanocapsules. Representative DLS traces at the different preparative stages are depicted in Figure 3. As expected, liposomes extruded through a 200 nm pore filter showed a typical monomodal size distribution centered around a hydrodynamic radius, RH, of 100. Not only do the liposomes retain the expected monomodality when monomers are incorporated, but UV irradiation and polymerization do not alter the size of the aggregates as well (cf.
Figure 3. Representative DLS traces. The hydrodynamic radius is plotted against the percentage of total scattered intensity. (A) Liposomes after extrusion through a 200 nm nucleopore filter. (B) Mixed liposome/polymer capsules after polymerization. (C) Polymeric nanocapsules after removal of lipid with TX-100. The second peak at approximately 4 nm corresponds to mixed TX-100/lipid micelles.
Figure 3A,B). The size distribution remains monomodal with an average vesicle size of about 100 nm in radius. To investigate the quality of the stabilizing polymer network, we dissolved the liposomes after polymerization by intensive vortexing with 1% TX-100. Notably, DLS could detect two distinct populations of aggregates (Figure 3C). The peak at a radius of approximately 4 nm corresponds to the TX-100/lipid micelles that are formed after removal of lipids from the polymer network. The peak at 100 nm indicates that the polymerization led to a stable and intact 2D network and that closed capsules formed. An open
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Figure 4. Hydrodynamic radius determined by DLS as a function of molar ratio. Upper panel: BMA/lipids; lower panel: HEMA/ lipids. (A) Before UV polymerization, (B) after polymerization, and (C) after removal of lipids with TX-100.
network with defects would not be stable without the lipid environment and would either collapse or deflate. It is important to state that the size distribution shown in Figure 3C can only be accomplished if we add the polymer monomers and the photoinitiator directly to the chloroformic solution of lipids, prior to the drying of the film. It was absolutely essential to use UVinduced polymerization with the apparatus described above. If this experimental procedure was not followed, most of the particles were destroyed after the addition of TX-100, and a clear bimodal size distribution originating from lipid/TX-100 micelles and the polymeric nanocapsules could not be found. Preliminary experiments following exactly the procedures described by Meier et al. and Jung et al. were also not successful.14-20 One of the most crucial points in the overall preparative procedure is the moment of the addition of the monomers and the initiator to the lipids and the molar ratio of lipids/monofunctional monomer/bifunctional monomer/photoinitiator. The monomers and photoinitiator should not be added after the liposome preparation, that is, to a dispersion of liposomes. DLS measurements revealed (data not shown) that they could not insert into the lipid bilayer. The addition of TX-100 disrupted all structures, not leading to liposome stabilization or polymer capsule formation. 3.2. Effect of Monomer Concentration. In a series of measurements, we varied the molar ratio of monomer to lipid and used the monofunctional monomers BMA and HEMA. The molar ratio of monofunctional monomer/cross-linker/photoinitiator was 1/10/20 for all the experiments. Figure 4 depicts the average RH for monomer-to-lipid ratios between 1 and 4, showing liposomes before polymerization (A), after polymerization (B), and after liposomes lysis (C). The molar ratio of zero corresponds to the native, template liposomes. The structures do not significantly change size during the three-step procedure. Importantly, the cross-linking does not modify the liposome size, and most of the particles kept the initial template size after the addition of detergent. For smaller and bigger molar ratios (data not shown), the results were not satisfactory. Higher monomer concentrations led to a heterogeneous sample after UV irradiation and ratios smaller than 1 could not cover the bilayer surface entirely, leading
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Figure 5. Cryo-TEM micrographs of egg-PC/BMA/EGDMA/ Irgacure vesicles at a molar ratio of 2/1/10/20. (A) Control liposomes only composed of egg-PC. (B) Mixed liposome/polymer capsules after UV polymerization.
Figure 6. Negative-stain TEM micrographs of polymeric nanocapsules after removal of lipids.
to incomplete polymerization. Specifically, increasing the amounts of the monofunctional monomer resulted in the formation of heterogeneous samples with aggregates of a mean diameter between 100 and 500 nm, sometimes even larger. This phenomenon can easily be explained by considering the overall available space for the hydrophobic monomers inside the bilayer. At a certain point, no more space is available, and the monomer will aggregate inside the aqueous solution and polymerize into larger structures. 3.3. Shape of the Nanocapsules. In addition to DLS, we performed cryo-TEM and negative-stain TEM to investigate the shape and integrity of the different structures. We first imaged control liposomes only composed of egg-PC and the stabilized liposomes containing the polymer network (Figure 5A,B). Clearly, the polymerization process did not influence the shape and integrity of the liposomes. Negatively stained hollow polymer nanocapsules without the surrounding lipid are depicted in Figure 6. We could not image these particles under cryo-conditions since the solubilized lipid and the presence of TX-100 prevented the formation of clean vitreous ice. However, the polymer nanocapsules could withstand the conditions needed for negative staining, and spherical particles of about 100 nm radius could
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Figure 7. (A) Representative contact-mode AFM topography of BMA/EGDMA/Irgacure vesicles at a molar ratio of 1/10/20. (B) Crosssection of same sample.
be visualized. No phase separation was observed, and we conclude that the present experimental procedure allows all monomers to homogeneously distribute within the bilayer. In accordance with the TEM and DLS results, AFM images revealed circular, flattened spheres of polymeric nanocapsules. A representative topography image and cross-section are shown in Figure 7. The image clearly shows a circular shape (Figure 7A), with a 97 nm diameter and 13 nm height (Figure 7B), which was verified with different AFM tips. The size distributions of the nanocapsules are in agreement with the TEM images. The AFM images exclude an elongated or angular structure of the nanocapsules, and the images are not influenced by a staining procedure. Nevertheless, the nanocapsules might collapse or shrink when dried on a surface, resulting in a smaller height than in solution. The height of nanocapsules in AFM images might indicate an upper limit of twice the thickness of the polymer membrane.23-25
bilayer was maintained. It is especially important to tune the molar ratio of polymerizable monomer to lipid, as otherwise undesired structures or aggregates might be obtained. Interestingly, this procedure might be a new way of encapsulating active agents into non-liposome-based polymer nanocontainers, since the polymer network remained stable during negative staining and AFM imaging. Moreover, the incorporation of hydrophobic functional constituents that are apt to undergo radical polymerization inside the bilayer is a new step toward the development of stable nanocapsules with functionalized walls. In this way, capsules with magnetic nanoparticles anchored inside the 2D polymer network can be obtained.26 The fortified lipid/polymer hybrid capsules combine the advantages of liposomes, that is, biocompatibility, and polymer nanocapsules, that is, stability. However, to envisage new applications and encapsulation processes of active molecules, such as enzymes, more information will be needed about the wall thickness, permeability, and porosity of the capsules in different media and under biological conditions.
4. Conclusion Our investigations feature the possibility of template polymerization as a method to prepare hollow lipid/polymer capsules of a defined size, which was previously not possible. Notably, no significant size and shape changes were observed at the different preparation steps, and the integrity of the liposomal template (23) Hotz, J.; Meier, W. Langmuir 1998, 14, 1031-1036. (24) Li, S. L.; Palmer, A. F. Langmuir 2004, 20, 7917-7925. (25) Wang, T.; Deng, Y.; Geng, Y.; Gao, Z.; Zou, J.; Wang, Z. Biochim. Biophys. Acta 2006, 1758, 222.
Acknowledgment. This research was supported by the European project Nanocapsules with Functionalized Surfaces and Walls (HPNR CT200000159) and by AC NanosciencesNanotechnologies (NN082). A.F.P.S. acknowledges funding from the Wellcome Trust. LA0613575 (26) Gomes, J. F. P. d. S. International University of Bremen, Bremen, Germany. To be submitted for publication.
