Alexandra S.
Angelatos
,
Kiyofumi
Katagiri†
and
Frank
Caruso
*
Centre for Nanoscience and Nanotechnology, Department of Chemical and Biomolecular Engineering, The University of Melbourne, Victoria 3010, Australia. E-mail: fcaruso@unimelb.edu.au
First published on 25th November 2005
This paper provides an overview of our recent work in the area of bioinspired colloidal particles. We highlight how modifying the basic polyelectrolyte multilayer shell with materials such as light-absorbing gold nanoparticles, lipid bilayer membranes, and targeting ligands can functionalize colloids prepared via the layer-by-layer assembly technique. These nanoengineered colloids are expected to show promise in areas ranging from drug and gene delivery to cell membrane modeling.
![]() Alexandra Angelatos | Alexandra Angelatos (born in Melbourne, Australia in 1979) is a PhD student at the Centre for Nanoscience and Nanotechnology, Department of Chemical and Biomolecular Engineering, The University of Melbourne. She completed her BEng/BSc degree at The University of Melbourne in 2003, and commenced her postgraduate studies under the supervision of Professor Frank Caruso in 2004. |
![]() Kiyofumi Katagiri | Kiyofumi Katagiri (born in Gifu, Japan in 1975) is a postdoctoral research fellow in the Department of Materials Science, Toyohashi University of Technology (supervisor: Professor A. Matsuda). He received his BEng degree from Osaka Prefecture University (supervisors: Professors M. Tatsumisago and T. Minami), and his MEng and PhD degrees from Nara Institute of Science and Technology (supervisors: Professors K. Ariga and J. Kikuchi). He spent two years as a postdoctoral research fellow under the supervision of Professor Frank Caruso at the Centre for Nanoscience and Nanotechnology, Department of Chemical and Biomolecular Engineering, The University of Melbourne. He was awarded a Research Fellowship for Young Scientists from the Japan Society for the Promotion of Science. His current research interests include biomimetic materials and organic–inorganic nanohybrid materials based on supramolecular chemistry, colloid and surface science, and sol–gel science and technology. |
![]() Frank Caruso | Frank Caruso (born in Platania, Italy in 1968) is a professor, Australian Research Council Federation Fellow, and Director of the Centre for Nanoscience and Nanotechnology in the Department of Chemical and Biomolecular Engineering at The University of Melbourne. He received his PhD degree from The University of Melbourne in 1994. He then moved to the CSIRO Division of Chemicals and Polymers in Melbourne studying the interfacial alignment of receptor molecules for biosensor applications. In 1997 he became an Alexander von Humboldt Research Fellow at the Max Planck Institute (MPI) of Colloids and Interfaces (Germany), and from 1998–2002 was group leader at the MPI. His main research interests are polymers at interfaces, biomaterials, nanostructured colloidal systems, and nanocomposite thin films. He has published over 150 papers in peer-reviewed scientific journals and is co-inventor of more than 15 patents. |
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Fig. 1 Illustration of bioinspired colloidal systems prepared via LbL assembly: (a) polyelectrolyte multilayer capsules; (b) light-responsive polyelectrolyte–gold nanoparticle capsules for controlled delivery; (c) polyelectrolyte-supported lipid vesicles for cell membrane modeling and drug/gene delivery; (d) biofunctionalized colloids for targeted delivery. |
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Fig. 2 (a) Illustration of laser-induced drug release from biofunctionalized polyelectrolyte–gold nanoparticle capsules (diagram courtesy of New Scientist, Issue 8, January 5, 2005). (b) Extinction at 450 nm of the bacterium micrococcus lysodeikticus as a function of time after mixing with various poly(sodium 4-styrenesulfonate) (PSS)–poly(allylamine hydrochloride) (PAH)–gold nanoparticle-encapsulated lysozyme crystals: intact encapsulated crystals (closed circles); irradiated encapsulated crystals (open circles); crushed encapsulated crystals (triangles). The initial data point (0 min, 0.82), denoted by a cross, applies to all three systems. The bacterium is a substrate for the enzymatic action of lysozyme, with its extinction being a measure of the amount of bacteria in the sample. Hence, the amount of bacteria digested by the lysozyme correlates to the activity of the released lysozyme. When no release is induced, the extinction is essentially constant over the time range investigated; this implies that the lysozyme-loaded PSS–PAH–gold nanoparticle capsules are effectively leak-proof. When release is induced via laser irradiation, the extinction decreases abruptly and then levels out; this indicates that a significant proportion of the bacteria are digested and that the lysozyme activity remains stable, following the laser-induced release. When release is induced via crushing, the decay in the extinction is very similar to that for the laser-induced release case; this indicates that any loss in lysozyme activity induced by the laser pulse is comparable to that caused by mechanical rupturing of the capsules. |
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Fig. 3 (a) (i) Scanning electron microscope (SEM) and (ii) transmission electron microscope (TEM) images of PSS–PAH–gold nanoparticle capsules prior to laser exposure. The dried capsules resemble deflated balloons, and there is a homogeneous dense packing of gold nanoparticles (which appear bright in TEM) within the capsule shells. (b) (i) SEM and (ii) TEM images of PSS–PAH–gold nanoparticle capsules following laser exposure. The outlines of individual capsules can no longer be readily discerned, and there is evidence of gold nanoparticle fusion among the remnants of the capsules. (c) (i) SEM and (ii) TEM images of PSS–PAH capsules prior to laser exposure. These images are indistinguishable from those of PSS–PAH capsules following laser exposure, indicating that the laser irradiation has no apparent effect on the morphology of PSS–PAH capsules without the light-absorbing gold nanoparticles in their shell. |
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Fig. 4 (a) Relative N-(7-nitro-2,1,3-benzoxadiazol-4-yl)-1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine (NBD-DPPE) fluorescence at 533 nm as a function of the TX-100/lipid concentration ratio for PS core–PDDA–PSS shell particles coated with Si-lipid (containing 3 mol% NBD-DPPE) (triangles) or DMPA (containing 3 mol% NBD-DPPE) (circles). In the case of the core–shell particles coated with DMPA, the lipid probe fluorescence decreases, indicating disassembly of the DMPA membrane when the particles are exposed to five or more equivalents of TX-100. In the case of the core–shell particles coated with Si-lipid, the lipid probe fluorescence remains constant, and hence the Si-lipid membrane remains stable in the presence of up to 30 equivalents of TX-100. (b) Relative NBD-DPPE fluorescence at 533 nm as a function of EtOH concentration for PS core–PDDA–PSS shell particles coated with Si-lipid (containing 3 mol% NBD-DPPE) (squares) or DMPA (containing 3 mol% NBD-DPPE) (circles). In the presence of pure EtOH, the decrease in the lipid probe fluorescence for the core–shell particles coated with DMPA is more than three times that for the core–shell particles coated with Si-lipid, demonstrating that the Si-lipid membrane is considerably more resistant to EtOH than the DMPA membrane. |
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Fig. 5 (a) Illustration of a templating core particle LbL-coated with several precursor polyelectrolyte layers, followed by an asymmetric lipid bilayer membrane (i.e., a lipid membrane where the inner and outer layers of the bilayer comprise different lipid molecules), followed by several additional polyelectrolyte layers. (b) Relative NBD-DPPE fluorescence at 533 nm as a function of time for MF particles coated with a polyelectrolyte-supported asymmetric lipid bilayer where DHP (containing 3 mol% NBD-DPPE) forms the inner (open circles) or outer (closed circles) layer of the bilayer membrane. In both cases, NBD-DPPE is embedded in the DHP layer and DDAB forms the alternate lipid layer. Sodium hydrosulfite (Na2S2O4), a compound capable of quenching the fluorescence of lipid probe molecules located in the outer layer of lipid membranes only as it cannot diffuse across lipid bilayers, was added to the particle dispersions after 1 min. For the particles where DHP forms the outer layer of the asymmetric lipid bilayer, the lipid probe fluorescence undergoes a rapid decrease upon Na2S2O4 addition (ca. 80% within 4 min), which confirms that the majority of the NBD-DPPE, and in turn the DHP, resides in the outer layer of the lipid membrane. For the particles where DHP forms the inner layer of the asymmetric lipid bilayer, the lipid probe fluorescence undergoes a relatively minor decrease upon Na2S2O4 addition (ca. 20% within 4 min), confirming that the majority of the NBD-DPPE, and hence the DHP, resides in the inner layer of the lipid membrane. No apparent change in the emission properties of the two systems was observed after 72 h, indicating that the asymmetry of the lipid bilayers is maintained for days after preparation. |
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Fig. 6 Fluorescence microscope images of MF core–PSS–PAH shell particles coated with DLPE and functionalized with antibodies, mouse IgG (primary antibody) and FITC-labelled rabbit anti-mouse IgG (secondary antibody). The fluorescence observed originates from the FITC label on the secondary antibody. |
Footnote |
† Current address: Department of Materials Science, Toyohashi University of Technology, Toyohashi, Aichi 441-8580, Japan |
This journal is © The Royal Society of Chemistry 2006 |