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,‡
Jürgen Fritz,‡ Manuel A
Ä lvaro Neto Coelho,§ Didier Fournier,† Clara Fournier-Nöel,#
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, Laboratório 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 ReactiVité 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 Reactivité 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 © xxxx American Chemical Society
Published on Web 07/28/2006
PAGE EST: 4.7
B Langmuir
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.
Gomes et al.
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.
Stable Hollow Polymethacrylate Nanocapsules
Langmuir C
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
D Langmuir
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
Gomes et al.
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
Stable Hollow Polymethacrylate Nanocapsules
PAGE EST: 4.7
Langmuir E
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.