EP2575896A2 - Magnetically responsive membrane structures - Google Patents

Abstract

The present invention is directed towards magnetically responsive vesicular compositions comprising (a) a vesicular structure having a membrane enclosing a cavity and (b) at least one stabilized magnetic nanoparticle embedded in said membrane, and in particular towards lipid and/or polymeric vesicular compositions comprising stabilized superparamagnetic iron oxide nanoparticles. The vesicular compositions may further be functionalized with reactive groups for attachment of active agents or tethers for attachment to surfaces. Methods of preparation and their use in (targeted) delivery of an active agent, as a nanoreactor, for imaging purposes and combinations thereof are also disclosed.

Description

Magnetically Responsive Membrane Structures

Field of the invention

The present invention is directed towards a magnetically responsive composition comprising a membrane and at least one stabilized magnetic nanoparticle embedded in said membrane, wherein the membrane is in form of a magnetically responsive vesicular structure or in form of a magnetically responsive supported lipid bilayer, as well as derivatives of said composition, methods of preparation and use in (targeted) delivery of an active agent, as a nanoreactor, for imaging purposes and combinations thereof.

Background of the Invention

Nanoscale vesicles to store, react and release compounds is a large and hot field of research in itself and the by far largest sub-field with possibly the highest societal and economic impact is drug delivery. The obvious advantages of handling molecules in nanoscale containers are the small and concentrated amounts of materials that can be used and dimensions that are of the size typically encountered for transporting and handling materials in biological systems on close to the molecular size scale. Thus, e.g., drug delivery vehicles are generally regarded as having to be maximum 100 ran in diameter.

Injecting drug loaded vesicles into an in vivo environment exposes them to tremendous challenges. They should among other things i) be small enough to pass through the smallest blood vessels; ii) not be recognized by the immune system or too quickly be enzymatically degraded; iii) release the compounds they carry quickly and efficiently at the desired site of action without leaking during systemic circulation. To date, this area remains a hot and challenging topic of research both in academia and industry since the trade-off between the desired properties above is essentially unresolved. In particular, it has in practice been

l extremely difficult to design vesicles which circulate for a long time with low leakage of compounds but still can release drug cargo efficiently at the required destination.

One of the most researched drug delivery systems and currently the research benchmark in terms of performance and assays for testing are poly(ethylene glycol)-shielded liposomes (PEG-liposomes). These vesicles can be formed at a size of 50-400 nm with standard methods like extrusion and techniques have been developed for efficient loading with drugs. They can in principle be loaded with both hydrophilic drugs in the lumen and with hydrophobic drugs in the membrane interior, or as water-soluble hybrid composite materials in the vesicle lumen. The PEG coating provides them with stealth properties that significantly prolongs the circulation time in vivo and it is easy to functionalize them with ligands for targeting specific sites in the body. Thanks to the properties of the lipid membrane they are very stable also in dilute and physiological conditions, while retaining flexible permeability and being completely biodegradable. However, the high stability beneficial for circulation and drug retention, compromises efficiency of cargo release at the desired location and it has been difficult to tune the release to occur at sufficient rate at the desired site of action without causing high leakage during circulation.

Further developments of vesicular delivery and release systems include polymersomes and polymer micelles. These combine the useful hydrophobically driven self-assembly property of liposomes with the greater synthetic flexibility of polymer systems, and in particular vesicular structures that are susceptible to triggers for release. The latter could be used to enhance desired properties such as suppressing leakage, make the vehicle more susceptible to triggers for release or modify the release mechanism as described below. Polyelectrolyte multilayer vesicles and crosslinked polymer and block copolymer vesicles formed around micro and nano particles, droplets and bubbles also belong to this category. Such vesicles do not necessarily contain a hydrophobic polymer core, but can also derive stability from physical or chemical crosslinking of polymers in the shell. Furthermore, such complex vesicular systems can allow encapsulation of multiple solvents, e.g. including a hydrophobic solvent encapsulated by an amphiphilic membrane.

Different approaches have been used to give PEG-liposomes as well as polymersomes better release properties. The most promising approaches focus on using changes in vesicle stability through environmental triggers affecting permeability to which the vesicle is only subjected at the desired site of release. Approaches that have been explored are e.g. enhanced release through higher temperature in sick, e.g. cancerous, tissue and degrading and release through a lowering of pH in, e.g., the endosome after internalization. Yet, although these are approaches of continued interest, their success so far has been limited.

An attractive alternative approach to relying on weak stimuli from the biological environment is to use triggers that can be externally actuated from outside the body. Vesicular systems, which can release their cargo by external triggers, such as ultrasound activation, laser induced release, temperature change (Bedard, M.F., et al., Acs Nano, 2008. 2(9): 1807), etc., may find various uses, ranging from drug delivery vehicles to reactants in nanoscale chemical and biochemical reactions However, current approaches are likely to also significantly disturb the membranes and internal processes of tissue cells through heating or other energy transfer to cell components.

As was described above it has been realized that nanoparticles (NPs) possess the ability to couple strongly to external electromagnetic fields. There are some attempts to exploit this for drug delivery in the literature but most do not fulfill the basic requirements for drug delivery vehicles or nanoreactors as outlined above, and none demonstrate a high-performance system suitable for drug delivery applications. One attempt of incorporating NPs into organic membranes to make responsive capsules make use of polyelectrolyte multilayers (PEMs). The PEMs are formed onto sacrificial microparticle scaffolds into which functional NPs have been physisorbed, typically by charge interaction (Andreeva, D.V. et al., Macromolecular Rapid Communications, 2006, 27: 931-936; Angelatos, A.S. et al., Journal of Physical Chemistry B, 2005, 109 : 3071-3076; Gorin, D.A. et al, Physical Chemistry Chemical Physics, 2008, 10: 6899-6905). This approach has allowed large micron-sized capsules to be made which incorporate Au or magnetic iron oxide NPs. It has furthermore been shown that such capsules can be moved by magnetic fields when magnetic NPs (Andreeva, D.V., et al., Macromolecular Rapid Communications, 2006, 27: 931-936; Gorin, D.A., et al., Physical Chemistry Chemical Physics, 2008, 10: 6899-6905) are incorporated and that encapsulated dye can be released by laser irradiation when plasmonically active Au NPs (Angelatos, A.S., et al., Journal of Physical Chemistry B, 2005, 109: 3071-3076; Gorin, D.A., et al., Physical Chemistry Chemical Physics, 2008, 10: 6899-6905) have been incorporated. It has also been shown that incorporation of NPs can significantly enhance the mechanical stability of the membranes (Bedard, M.F. et al., Soft Matter, 2009, 5: 148-155) and that close packing of the plasmonic NPs greatly enhances the efficiency of energy transfer from the NPs to the polymer matrix. (Bedard, M.F. et al., Acs Nano, 2008, 2: 1807-1816) Yet, multiple process steps of both deposition and dissolution may result in membranes of significant thickness and large capsule size and the use of plasmonic heating as well as the highly charged nature of the obtained membranes may limit their in vivo compatibility and thus in vivo applicability. Triggered release has also recently been demonstrated actuated by magnetic fields for silica nanospheres encapsulating Fe₃0₄ NPs (Hu, S.H. et al., Langmuir, 2008, 24: 239-244; Hu, S.H. et al., Advanced Materials, 2008, 20: 2690-+). The particles were heated by applying high-frequency magnetic fields which caused pore formation and release of drugs. In an attempt to combine thermoresponsive polymer membranes with magnetic heating through NPs, Hoare et al. recently described the encapsulation of aggregated superparamagnetic particles in 0.1-3 μιη clusters into PNIPAM hydronanogels (Hoare, T. et al., Nano Letters, 2009, 9: 3651-3657). By AC magnetic heating the nanogel could be collapsed and let model drugs penetrate from one side to the other of the thick gel. This was however accomplished for macroscopically thick membranes meant for large implanted capsules and not applicable for circulating drug delivery vehicles and nanoreactors.

There are only few reports on incorporation of NPs, primarily quantum dots (QDs) and metal NPs, directly in the membrane of liposomes.

Theoretical studies indicate that lipid membranes can accommodate quite large NPs without an unfavorable increase in energy. Wi et al. have predicted that NPs at least up to 6.5 nm in diameter should favorably embed into the hydrophobic core of a lipid membrane of typical thickness by employing a simple model for symmetric elastic deformation of the membrane around an embedded NP (Wi, H.S. et al., Journal of Physics-Condensed Matter, 2008, 20). English and coworkers were the first to show incorporation of NPs into a lipid membrane of liposomes in the size range 60-300 nm (both embedded in the hydrophobic core of the membrane as well as exposed at the surface) without causing changes in the liposome permeability (Jang, H. et al., Journal of Photochemistry and Photobiology a-Chemistry, 2003, 158: 11 1-1 17). Incorporation of NPs in the liposome membrane was shown and there was no significant change detected in the liposome permeability due to the incorporation. Fluorescence quenching experiments suggested that NPs could both completely embed in the hydrophobic core of the membrane and be exposed at the surface. Yet, neither size nor colloidal properties of these weakly stabilized NPs were well characterized.

Vogel and co-workers showed stained liposome membranes using TOPO stabilized CdSe (3 nm core, 5 nm total diameter) QDs. The QDs were first dried with lipids to form a thin film, which upon hydration on an electrode at applied bias formed giant unilamellar liposomes with the QDs dispersed in the membrane (Gopalakrishnan, G. et al., Angewandte Chemie- International Edition, 2006, 45. 5478-5483). A low QD to lipid ratio was used, but the formation of liposomes with QDs incorporated was claimed to work over a large range of lipid compositions and vesicle sizes without noticeable change to the melting temperature, T_m, of the lipid membrane. Single particle tracking of QDs was performed which revealed a diffusion coefficient lower than that of the lipids, but with a large uncertainty in value due to the experimental method. The liposomes were used to label cells, but it remained unclear whether the liposomes stained the membrane of the cell through fusion or association.

Al Jamal et al. also incorporated ultra-small, 2 nm in diameter, TOPO coated, commercial CdSe/ZnSe QDs into liposome membranes, but used sonication to produce 100 nm in diameter liposomes (Al- Jamal, W.T. et al., Acs Nano, 2008, 2: 408-418). The QD-liposomes were shown to be stable over time, but cryo-transmission electron microscopy (cryo-TEM) revealed clustering of QDs in the membrane and a high frequency of joined liposomes, possibly due to the weak stabilization of TOPO coated QDs. No further characterization was made but the labeled liposomes were used for in vitro and in vivo imaging. This system was also used by Bothun et al. who did not perform any additional characterization (Bothun, G.D. et al, Journal of Physical Chemistry B, 2009, 113: 7725-7728).

Binder et al. recently performed a study of the incorporation of hydrophobic Au and CdSe NPs about 2 nm in size into lipid and polymer vesicle membranes (Binder, W.H. et al., Physical Chemistry Chemical Physics, 2007, 9: 6435-6441). A high NP to lipid ratio of the weakly stabilized CdSe particles was shown to disrupt the formation of liposomes, while more thoroughly stabilized hydrophobic Au NPs were shown to exist in a large fraction outside the liposome membrane. A non-conclusive observation of small hydrophobic NPs having a larger detrimental effect on liposome formation was proposed by comparing to integration of virus particles. In contrast, Park et al. incorporated a high concentration of hydrophobic 4 nm Ag NPs in the liposome membrane and claimed increased membrane fluidity through fluorescence anisotropy measurements (Park, S.H. et al., Colloids and Surfaces B-Biointerfaces, 2006, 48: 112-118).

Finally, Paasonen et al. showed both incorporation of NPs into the membrane of liposomes and were the first to show triggered release from liposomes using plasmonic particles (Paasonen, L. et al., Journal of Controlled Release, 2007, 122: 86-93). However, release could only be achieved by irradiation of a UV laser due to the small size of the NPs. While liposome stability, stable encapsulation and efficient release were demonstrated. Other more recent attempts of incorporating magnetic NPs covered by oleic acid into lipid membranes (Chen, Y.J. et al, Acs Nano, 2010, 4: 3215-3221) showed high passive leakage of incorporated dyes from the liposome interior and the necessity to apply high alternating magnetic field densities to effect actuated release of encapsulated dyes.

Other reports disclose multi-lamellar liposomes incorporating iron oxide NPs based on similar older work by synthesizing the NPs in situ using co-precipitation of iron salts (Faure, C. et al., Journal of Physical Chemistry B, 2009, 113: 8552-8559). The large onion-like liposomes showed aggregates of iron oxide NPs intercalated in the aqueous space between the membranes, where the individual particles in the aggregates were 3-6 nm. These magnetosomes could be aggregated using application of a permanent magnet. Beaune et al. loaded hydrophilic superparamagnetic iron oxide nanoparticles (SPIONs) into giant unilamellar liposomes similar to others, which could be deformed from spherical shape by applied magnetic fields if the ionic strength of the ferrofluid was high enough (Beaune, G. et al, Journal of Physical Chemistry B, 2008, 1 12: 7424-7429). They have also shown that QDs inserted in the membrane and SPIONs in the lumen can be combined in a single vesicle at high particle to lipid ratio with the vesicles stable for months (Beaune, G. et al., Angewandte Chemie- International Edition, 2007, 46: 5421-5424). Magnetic release using extruded liposomes containing SPIONs in the lumen has been shown by a couple of groups using high- frequency magnetic fields, e.g. (Tai, L.A. et al., Nanotechnology, 2009, 20; Babincova, M. et al., Bioelectrochemistry, 2002, 55: 17-19). The stability and detailed characterization for these systems are lacking and high power has typically been applied. A very recent example using charge stabilized CoFe₂0₄ NPs inside the liposome lumen but applying low-frequency magnetic fields for the release was demonstrated by Nappini et al. (Nappini, S. et al., Soft Matter, 2010, 6: 154-162).

Clearly there are still disadvantages relating to stability, actuation (release mechanism), etc. associated with the currently known systems uding NPs. Applicants have now found that magnetic NPs, which are stabilized using high-affinity anchors to establish a thin but very dense hydrophobic coating, can be incorporated into a membrane of choice without affecting important parameters such as permeability and phase transition temperatures to form membrane systems of high stability. Applicants have found that these systems can be used for spatially and temporally localized release by minimally invasive means using application of externally applied magnetic fields and thus are particularly useful for various applications such as therapeutic and diagnostic drug delivery as well as nanoreactor applications. In addition, magnetic NP containing membranes can be selected such that pulsed application of alternating magnetic fields can be used to release compounds encapsulated in vesicles without environmental heating degrading the vesicle, thermosensitive compounds or surrounding cells or tissue.

Summary of the Invention

The present invention relates to a magnetically responsive composition comprising a membrane and at least one stabilized magnetic nanoparticle embedded in said membrane, wherein the membrane is in form of a magnetically responsive vesicular structure or in form of a magnetically responsive supported lipid bilayer.

Thus, in one aspect the magnetically responsive composition is in form of a magnetically responsive vesicular composition which comprises (a) a vesicular structure having a membrane enclosing a (optionally partially or completely tilled) cavity and (b) at least one stabilized magnetic nanoparticle embedded in said membrane.

Depending on the nature of the membrane, one (or more) permeable, semi-permeable, and or non-permeable cavity (cavities) may be formed (and/or can change from one state to another). The cavity may be optionally partially or completely filled, with a solid, liquid or gaseous cargo, in form of emulsions, droplets, bubbles and the like.

In one embodiment the vesicular structure is a liposome comprising at least one lipid type. In another embodiment the vesicular structure is a polymersome based on at least one synthetic and/or natural polymer, preferably amphiphilic synthetic polymers such as synthetic (co-)polymers, proteins (protein layers) or mixtures thereof. In a specific embodiment the polymersome comprises at least one block copolymer type.

In another aspect the magnetically responsive composition is in form of a magnetically responsive supported lipid bilayer which comprises (a) a planar bilayer of lipids or lipid- related materials including polymeric materials and (b) at least one stabilized magnetic nanoparticle embedded in said bilayer.

Preferably, the stabilized magnetic nanoparticle is selected from the group consisting of iron, cobalt or nickel, alloys thereof, preferably oxides or mixed oxides/hydroxides, nitrides, carbides or sulfides thereof. In a preferred embodiment the stabilized magnetic nanoparticles are superparamagnetic iron oxide nanoparticles (SPIONs).

In some embodiments, stabilization is achieved through association with a dispersant. Preferably, the dispersant comprises a catechol derivative anchor group covalently bound to a hydrophobic spacer. In specific embodiments, the hydrophobic spacer is a polymeric spacer and may be selected from the group consisting of linear, branched or dendritic hydrocarbon chains.

In further embodiments the dispersant further comprises at least one functional group, including but not limited to imaging groups, such as fluorophores, chelates binding MR (magnetic resonance)-active ions such as Gd³⁺ or radiotracers or a chemically reactive group for coupling of hydrophobic or hydrophilic moieties extending into or outside the host membrane.

In yet further embodiments the magnetically responsive composition further comprises an active agent, which is covalently linked to the membrane of a vesicular structure or embedded in the membrane of a vesicular structure or enclosed in the cavity of a vesicular structure in form of a vesicle.

The active agent may be a therapeutic or diagnostic agent, a nutritional agent or a targeting group or combinations thereof.

In other embodiments the magnetically responsive composition further comprises a polymeric coat of a stealth polymer tethered to the membrane.

In yet other embodiments the membrane of the vesicular structure further comprises a functinal group, such as NHS ester, maleimide, azide, acrylates, methacrylates, amines, thiols, carboxy groups, photoinitiators or physically strongly interacting groups such as chelates, charged groups or specifically binding protein ligands. The functional group may be linked to the lipid or block-co-polymers of the membrane.

In further embodiments the magnetically responsive composition may be encapsulated in a matrix such as a physically or chemically crosslinked hydrogel or embedded in a matrix tethered to a substrate. In some embodiments the encapsulating matrix may be a fibrous or membranous tissue engineering scaffold promoting tissue and cell growth and adhesion, or a polymer layer bound to an implant surface.

In yet another aspect the invention is directed towards magnetically responsive compositions of the invention for use in (targeted) delivery of an active agent, as a nanoreactor, or for imaging purposes.

In another aspect the invention is directed towards a method of locally accumulating a magnetically responsive composition of the invention by application of a magnetic field. In specific embodiments a magnetically responsive composition may be tethered to a liquid crystalline surface such as a lipid bilayer through mobile linkers and optionally can be magnetically locally accumulated in the surface plane. In specific embodiments the actuation is of magnetic NPs moving within the plane of a membrane in response to applied magnetic fields.

In another aspect the invention is directed towards a method of changing the permeability of a magnetically responsive composition of the invention, said method comprising the step of exposing said magnetically responsive vesicular composition to an alternating magnetic field, whereby permeability of the magnetically responsive vesicular composition is increased through local heat generation.

In another aspect the invention is directed towards a method of changing the permeability of a magnetically responsive vesicular composition of the invention, said method comprising the step of exposing said magnetically responsive vesicular composition to a direct or slowly varying magnetic field, whereby permeability of the magnetically responsive vesicular composition is increased through mechanical deformation.

In another aspect the invention is directed towards a method of locally changing the fluidity of magnetically responsive membranes (including associated or non-associated, i.e. freely moveable compounds) by application of magnetic fields.

In another aspect the invention is directed towards a method for delivery of an active agent in a subject, comprising the steps of: (a) associating at least one active agent with a magnetically responsive vesicular composition of the invention, (b) administering said composition to a subject; and (c) exposing said composition to a direct or alternating magnetic field, whereby either the permeability of said composition is increased such that said active agent is released in said subject (and the vesicular structure is retained during release), or the vesicular structure is destroyed by the response of the magnetic NPs which are embedded in the membrane, releasing said active agent.

In another aspect the invention is directed towards a method for delivery of an active agent in cell or tissue culture, comprising the steps of: (a) associating at least one active agent with a magnetically responsive vesicular composition of the invention, (b) incorporating said composition in the cell or tissue culture directly or as part of the scaffold; and (c) exposing said composition to a direct or alternating magnetic field, whereby either the permeability of said composition is increased such that said active agent is released in said subject or the vesicular structure is destroyed by the response of the magnetic NPs which are embedded in the membrane, releasing said active agent.

In specific embodiments the magnetically responsive vesicular composition further comprises at least one targeting moiety to achieve delivery of an active agent to a target site of choice. Brief Description of the Figures

Figure 1. Schematic of the self-assembled a) magnetoliposome and b) supported lipid bilayer (SLB) structure incorporating magnetic nanoparticles. Alternating magnetic fields produce heat or distortions in the liposome or supported lipid bilayer membranes to release compounds, while constant magnetic fields are used to probe the mechanics of particle movement in membrane and membrane buckling inSLBs or liposomes.

Figure 2. Stabilization of iron oxide NPs. a) Schematic of nitroDOPA-PEG assembled onto a Fe₃0₄ core; b) and c) TEM and size distribution of stabilized NPs of 3.5 and 6.6 nm in radius respectively; d) nitroDOPA-PEG used for stabilizing NPs including PEG used so far for stabilization; e) Volume weighted diameters of iron oxide NPs stabilized with nitroDOPA- PEG(5) (-a-), nitrodopamine-PEG(5) (-□-), DOPA-PEG(5) (-A-), dopamine-PEG(5) (-Δ-), mimosine-PEG(5) (-·-), hydroxypyrrolidone-PEG(5) (-0-) and hydroxydopamine-PEG(5) (-V -); f) TEM of nitroDOPA-palmityl stabilized, Fe₃0₄ cores synthesized in the oil bath showing higher yield, and more uniform morphology and size than NPs synthesized in the microwave.

Figure 3: Thermogravimetry analysis (TGA) of palmityl-nitroDOPA stabilized 2.5 nm (red circles) and 5 nm (black squares) core radius iron oxide NPs.

Figure 4: DLS measurements of DSPC liposomes containing 5 mol% PEG(2)-PE. DLS measurements of DSPC liposomes containing 5 mol% PEG(2)-PE in their membranes and hosting small, 2.5 nm core radius (red squares) and large, 5 nm core radius (blue triangles) palmityl-nitroDOPA stabilized iron oxide NPs in their membranes. Liposomes that were formed through sonication (empty symbols) primarily assembled into micelles while liposomes that were extruded (filled symbols) assembled into vesicular structures.

Figure 5: Effect of the lipid composition of liposomes on their size, a) DLS measurements of l-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) (black circles), l-stearoyl-2- oleoyl-sn-glycero-3-phosphocholine (SOPC) (blue triangles) and l,2-Distearoyl-sn-glycero-3- phosphocholine (DSPC) (red squares) liposomes containing 2.5 nm core radius palmityl- nitroDOPA stabilized iron oxide NPs in their membranes (filled symbols) and control liposomes that do not contain any NPs in their membranes (empty symbols). All liposomes contained 5 mol% PEG(2)-PE in their membranes. DLS measurements were performed at 25 °C. The chemical structure of b) DSPC, c) SOPC and d) POPC is shown. Figure 6 TEM micrographs of trehalose fixed liposomes. DSPC liposomes containing 2.5 nm core radius palmityl-nitroDOPA stabilized iron oxide NPs in their membranes. A) While liposomes fixed with high amounts of trehalose remained intact upon drying, b) and c) liposomes that were fixed with smaller amount of trehalose partially ruptured resulting in liposome membranes that contain individually dispersed iron oxide NPs. No agglomerated NPs could be seen on these TEM micrographs indicating that individually palmityl- nitroDOPA stabilized NPs do not agglomerate if embedded in the hydrophobic part of liposome membranes.

Figure 7. a) cryo-TEM of DSPC liposomes containing 5 mol% PEG(2)-PE. Cryo-TEM images of DSPC liposomes containing 5 mol% PEG(2)-PE a) without any iron oxide NPs, with b) oleic acid and c) stabilized 2.5 nm core radius NPs and d) palmityl-nitroDOPA stabilized 5 nm core radius NPs.

Figure 8: Chemical analysis of liposomes. A) Photographs of DSPC liposome dispersions without NPs, and liposomes containint small, 2.5 nm core radius iron oxide NPs stabilized with palmityl-nitroDOPA and oleic acid respectively. All liposomes contained 5 mol% PEG(2)-PE in their membranes. STEM micrographs of DSPC liposomes containing 5 mol% PEG(2)-PE in their membranes and functionalized with small, 2.5 nm core radius palmityl- nitroDOPA stabilized iron oxide NPs in their membranes detected with a b) high angle annular dark field (HAADF) and c) secondary electron (SE) detector, d) Electron diffraction X-ray (EDX) measurements performed on the location shown in b and c.

Figure 9: SANS measurements on DSPC liposomes. SANS performed at a) 25 °C and b) 60 °C was measured on DSPC liposomes without NPs (black) and on liposomes hosting 2.5 nm core radius (red) and 5 nm core radius (blue) palmityl-nitroDOPA stabilized iron oxide NPs in their membranes.

Figure 10: SANS measurements of DSPC liposomes containing 5 mol% PEG(2)-PE. SANS was measured at a) 25 °C and b) 60 °C on DSPC liposomes containing 5 mol% PEG(2)-PE in their membranes. Small, 2.5 nm core radius (red) and large, 5 nm core radius (blue) palmityl- nitroDOPA stabilized iron oxide NPs were embedded in the liposome membrane. As a control, SANS was measured on DSPC/PEG(2)-PE liposomes without NPs (black).

Figure 11 : SANS measurements of POPC liposomes. SANS was measured at 25 °C on POPC liposomes without NPs (black), POPC liposomes containing palmityl-nitroDOPA stabilized small, 2.5 nm core radius (red) and large, 5 nm core radius (blue) NPs in their membranes. Figure 12: DSC measurements of DSPC liposomes. DSC measurements of a) DSPC liposomes and b) DSPC liposomes containing 5 mol% PEG(2)-PE in their membranes. DSC was measured on liposomes that did not contain any iron oxide NPs in their membranes (black), liposomes that hosted palmityl-nitroDOPA stabilized small, 2.5 nm (red) and large, 5 nm (blue) core radius iron oxide NPs in their membranes.

Figure 13: The relaxivity r₂ ^* of PEG(5 kDa)-nitroDOPA stabilized, 5 nm core diameter SPIONs as was measured with MRI. The same cores stabilized with palmityl-nitroDOPA instead of PEG(5 kDa)-nitroDOPA were incorporated into vesicle membranes.

Figure 14: QCM-D kinetics demonstrating formation of SLBs from POPC liposomes containing SPIONs (Af- solid symbols; AD - open symbols).

Figure 15. Quartz crystal microbalance with dissipation monitoring (QCM-D) measurements of a) SPION functionalized POPC liposomes that b) have been actuated with an external small magnet, c) Control measurements of pure POPC SLBs d) where no magnetic response was seen when a small external magnet was approached. Frequency changes (solid line) can be translated into changes in the adsorbed mass and dissipation changes (dotted line) indicate changes in the viscoelastic behavior of adsorbed films.

Figure 16: a) Reflection image of a nanoporous silicon nitride substrate with 100 nm in diameter pores onto which a supported lipid bilayer has been formed by liposome fusion of 132 nm diameter liposomes, b) The corresponding image of water soluble carboxyfluorescein dye captured and contained in the fraction of pores that could be shown to be spanned by a lipid bilayer (non-fluorescent pores have the membrane following the walls of the pore).

Figure 17: DSPC liposomes which have SPIONs in their membrane were loaded with a) carboxy- fluorescein and b) calcein. These liposomes were externally heated. They started to leak at temperatures -54 °C, close to their phase transition temperature which can be seen in an increase in fluorescence for carboxy-fluorescein loaded liposomes and in a shift of the absorption maxima for calcein loaded liposomes.

Figure 18: Alternating magnetic field triggered release. Release from DSPC liposomes containing 5 mol% PEG(2)-PE loaded with self-quenched calcein was measured by monitoring the fluorescence, (a) AMF treatment for 6 x 5 min followed by 1 min equilibration between every AMF exposure did not release calcein from unmodified DSPC liposomes containing 5 mol% PEG(2)-PE (red filled circles). However, liposomes hosting small 2.5 nm core radius (red filled squares) and large 5 nm core radius (blue filled triangles) palmityl- nitroDOPA stabilized iron oxide NPs in their membranes efficiently released their cargo. The release from liposomes functionalized with palmityl-nitroDOPA stabilized iron oxide NPs was 180% more efficient compared to that of liposomes prepared with oleic acid coated NPs (magenta filled stars) or liposomes that were loaded with hydrophilic PEG(1.5)-nitroDOPA stabilized small (red empty squares) and large (blue empty triangles) iron oxide NPs. (b) Photograph of dispersions containing DSPC/PEG(2)-PE liposomes that hosted palmityl- nitroDOPA stabilized small iron oxide NPs in their membranes before and after they were treated 6 x 5 min with an AMF. The system was equilibrated for 1 min in between the AMF pulses, (c) DLS measurements of DSPC/PEG(2)-PE liposomes (black filled circles) and DSPC/PEG(2)-PE liposomes functionalized with palmityl-nitroDOPA stabilized small NPs (red square) before (filled) and after (empty) AMF exposure are shown. They reveal that the liposome structure is retained upon AMF treatment indicating that cargo is released by the enhanced permeability of liposomes close to T_m. (d) DSPC/PEG(2)-PE liposomes functionalized with small palmityl-nitroDOPA stabilized NPs only started to release significant amounts of calcein at T > 50 °C if externally heated (red filled squares). As a comparison, the normalized fluorescence of liposomes that were exposed to an AMF as a function of T is shown for unmodified liposomes (black filled circles) and liposomes hosting small 2.5 nm core radius palmityl-nitroDOPA stabilized NPs in their membranes red empty squares).

Figure 19: Influence of the AMF sequence on the release efficiency, (a) Calcein release of DSPC/PEG(2)-PE liposomes functionalized with small 2.5 nm core radius iron oxide NPs stabilized with palmityl-nitroDOPA was tested for different sequences of the AMF. Release was less efficient if the system was equilibrated for 5 min (red crossed squares) in between each 5 min long AMF cycle (AMF sequence 2) compared to the release of liposomes equilibrated only for 1 min (AMF sequence 1) (red filled squares) but still significantly above the zero release of unmodified liposomes (black filled circles) treated with the AMF pulse sequence 1. Furthermore, release of NP modified liposomes was insignificant if these liposomes were subjected to 10 cycles of 1 min AMF pulses followed by 1 min equilibration time (AMF sequence 3) (red empty squares), (b) Bulk temperatures of liposome dispersions subjected to the respective AMF sequence used in (a).

Figure 20: a) cryo-TEM micrograph of PMCL-PDMAEMA polymersomes containing iron oxide NPs in their membranes. Volume weighted diameters of PMCL-PDMAEMA polymersomes b) containing iron oxide NPs with core diameters <4 nm in their membranes and c) PMCL-PDMAEMA block-co-polymers which were mixed with iron oxide NPs with a core diameter >10 nm prior to extrusion. Both shown before and after exposure to an AMF (100 kA/m) for lO min.

.Figure 21: SANS measurements of polymersomes. SANS was performed on a) PMOXA-b- PDMS-b-PMOXA and b) PMCL-PDMAEMA polymersomes. Measurements were performed on polymersomes that did not contain any NPs (black) and polymersomes containing palmityl-nitroDOPA stabilized 5 nm core radius NPs in their membrane (red), c) Additionally, SANS was measured on PMCL-PDMAEMA polymersomes containing 5 nm core radius palmityl-nitroDOPA stabilized iron oxide NPs at 25 °C (blue) and at 60 °C (red) respectively.

Figure 22: UV/VIS spectra of POPC vesicles where palmityl-nitroDOPA stabilized SPIONs (black) and SPIONs that were surface modified with a mixture of FITC-nitroDOPA and palmityl-nitroDOPA (red) and incorporated into the liposome bilayer. The absorption around 490 nm of liposomes loaded with FITC-labeled SPIONs shows feasibility to label liposomes by incorporating labeled SPIONs in their membrane.

Figure 23: FITC labeled dextran was loaded into SPION functionalized MPPC liposomes (black) and PCML-PEO polymersomes (red) resulting in fluorescently labeled vesicles.

Figure 24: QCM-D measurements of DNA tethered, SPION functionalized POPC liposomes that bound to POPC SLBs surfaces presenting the complementary DNA. The frequency (solid line) and dissipation shift (dashed line) upon addition of DNA tethered liposomes shows strong binding of tagged, SPION functionalized liposomes.

Figure 25: a) Build-up of a polyelectrolyte multilayer embedded vesicle layer on an inorganic substrate recorded by quartz crystal microbalance with dissipation monitoring. The process can be repeated to create multilayers of liposomes, b) Fluorescence microscopy image of 5,6- Carboxyfluorescein encapsulated in the liposome lumen. The square bleach spot in the image shows the contrast afforded by the dye. The dye intensity showed no decrease over lh while pH-induced destruction of the liposomes gave complete removal of the fluorescence signal.

Detailed Description of the Invention

The present invention is directed towards a magnetically responsive composition comprising a membrane and at least one stabilized magnetic nanoparticle embedded in said membrane, wherein the membrane is in form of a magnetically responsive vesicular structure or in form of a magnetically responsive supported lipid bilayer.

Figure 1 illustrates schematically a magnetically responsive composition in form of a magnetically responsive vesicular structure (a) or in form of a magnetically responsive supported lipid bilayer (b). Figure la shows schematically how alternating magnetic fields produce heat or distortions in the liposome membrane to release compounds. Figure lb shows schematically how constant magnetic fields may be used to probe the mechanics of particle movement in a membrane and membrane buckling in magnetically responsive supported lipid bilayers.(Rossetti, F.F. et al, Langmuir, 2005, 21: 6443-6450).

The term "magnetic nanoparticle(s)" as used herein (also termed nanoparticles of the invention or NPs of the invention) refers to any particle having a size in the nanometer scale that is magnetically responsive and exhibits superparamagnetic, paramagnetic, ferromagnetic or ferrimagnetic properties (i.e. that orients in a magnetic field along the magnetic field). For the purposes of the present invention, which is to incorporate the nanoparticles of the invention into a membrane, it is understood, that "a size in the nanometer scale" depends on both magnetic properties of the nanoparticle as well as the nature of the vesicular system of choice. An optimal size range allows to maximize the magnetic response of the nanoparticles and thus vesicles which are functionalized with those nanoparticles under the applied external magnetic field as well as colloidal nanoparticle incorporation efficiency within the membrane. It further minimizes mechanical instability of the vesicle membrane caused by nanoparticles from choice of nanoparticles larger in size than the vesicle membrane, and/or aggregation caused by heavy distortion of the membrane also leading to leakage, etc.).

The magnetic nanoparticle may either be paramagnetic, ferromagnetic, ferrimagnetic or superparamagnetic (or may show an intermediate characteristic), preferably superparamagnetic.

In specific embodiments, the magnetic nanoparticle is a member selected from the elements in the fourth row of the periodic table (i.e. chrome, manganese, iron, cobalt, and nickel), preferably of the group consisting of iron, cobalt or nickel, alloys thereof, more preferably oxides or mixed oxides/hydroxides, nitrides, carbides or sulfides thereof. In a specific embodiment, the magnetic nanoparticle is an iron oxide, nitride or an iron sulfide, preferably an oxide, mixed oxide/hydroxide, nitride or sulfide of Fe (II) and/or Fe (III), e.g. in the form of a nanocrystal. Preferably, the magnetic nanoparticle is Fe₃0₄ (magnetite). The magnetic nanoparticle can be prepared by methods known in the art in various shapes and sizes (see e.g. method described in this patent, Hyeon T.; The Royal Society of Chemistry 2003, Chem. Commun., 2003, 927-934 or US 4,810,401, incorporated herein by reference). For example, iron oxide nanoparticles may typically be prepared by a non-aqueous sol-gel method, Alternatively, the addition of base, such as ammonium hydroxide, to an aqueous mixture of ferrous and ferric sulfate or chloride, e.g. a mixture of FeCl₃ and FeCl₂. The molar ratio of the divalent to the trivalent salts may be varied form 0.0-3.0, preferable from 0.3 to 0.7, to obtain the desired size and magnetic characteristics of the magnetic nanoparticles. Divalent transitional metal salts such as cadmium, cobalt, copper, magnesium, manganese, nickel, zinc salts and their mixtures, may be substituted for some or all of the ferrous salt or ferrous precursors if nanoparticles are synthesized through a sol-gel method.

Preferred methods to obtain magnetic nanoparticles with core sizes sufficiently small to match the dimension of the hydrophobic domain of a membrane of choice, such as a lipid or amphiphilic polymer membrane typically include any synthesis route which results in a monomodal, preferentially monodisperse core size distribution, e.g. synthesis using an oil bath using a precursor such as Fe(ac)₂, Fe(ac)₃, Fe(acac)₂ or Fe(acac)₃ which is dissolved in an oxygen containing solvent such as benzyl-alcohol.

Typically the core size radii of nanoparticles range from 1 to 8 nm, preferably 1.5 to 5 nm, more preferably 1.5 to 2.5 nm. NPs having a core size radius of about 2.5 nm may be referred to as "small NPs", while NPs having an average radius of about 5 nm or larger may also be referred to as "large NPs".

The term "stabilized" as used herein in general but in particular with reference to a magnetic nanoparticle of the invention refers to a nanoparticle that is associated with a dispersant conferring steric stability to the nanoparticles.

The term "dispersant" as used herein refers to an anchor or an anchor group, covalently bound to a hydrophobic spacer. In this particular case, dispersants comprise an anchor group that allows irreversible binding of the dispersant to the nanoparticle surface which is covalently bound to a hydrophobic spacer. Irreversible binding of anchors to the nanoparticle surface as used herein refers to an adsorption constant k₀„ » than the desorption constant k₀ff of the dispersant to the nanoparticle surface. Typically, the dispersants comprise a terminal anchor group selected from unsubstituted or substituted catechol groups (i.e. 1 ,2-dihydroxybenzene groups) and derivatives thereof, preferably nitrocatechols, preferably 4-nitro-substituted catechol groups and derivatives thereof, such as 6-nitro-DOPA and 6-nitrodopamine [28]. The anchor groups of the present invention are effective to irreversibly immobilize dispersants on magnetic nanoparticles, thereby achieving good NP stability in dilute and high salt aqueous environments up to temperatures above 90°C (see Figure 2). Typically the dispersant provides a dense, thin layer, which is sufficiently thick to prevent NP agglomeration. More specifically, the term "dense" as used in this context refers to a packing density of dispersant typically above 0.5 dispersant/nm , preferably above 1 molecule/nm most preferably above 1.5 dispersant/nm². In specific embodiments, the dispersant should be larger than a single catechol such as nitrocatechols but can be as short as palmityl-nitroDOPA or shorter. In a preferred embodiment, the dispersant layer thickness is palmityl-nitroDOPA which provides a thin dispersant layer with a comparably low mass fraction of dispersants (typically around 10-20 wt%).

Preferred dispersants are dispersants with molecular weights < 10 kDa consisting of a well- suited anchor covalently linked to a hydrophobic spacer, preferably the anchor consists of a moiety with a high affinity towards iron oxide such as electronegetively substituted catechols that are covalently linked to a hydrophobic spacer where the spacer molecular weight is < 2 kDa, most preferably the dispersants consist of nitrocatechols covalently linked to hydrophobic spacers with molecular weights < 1 kDa.

A skilled person will understand that parameters such as length, chemical composition and end-functionality of the dispersants or anchor-spacer groups may be chosen very flexibly and adjusted to the specific applications of choice.

Preferred polymeric ligands are hydrocarbon groups, which encompass any polymers soluble in organic solvents in which lipids and block-co-polymers are also soluble. Typically, "hydrocarbon chains" or "hydrophobic spacers" include linear, branched or dendritic structures. Different forms of hydrocarbon chains may differ in molecular weights, structures or geometries (e.g. branched, linear, forked hydrocarbon chains, multifunctional, and the like). Hydrocarbon chains for use in the present invention may preferably comprise one of the two following structures: -(CH₂)_m- or -[(CH)_n-(CH₂)_m]o-[(CH)p-(CH₂)_q]_r, dendrimers of generations 1 to 10 where m is 3 to 3000 and n-r is 0 to 3000 and the terminal groups and architecture of the overall hydrocarbon chains may vary. This description includes any linear or branched hydrocarbon chains with ratios of unsaturated : saturated bonds varying from 0 : 100 to 100 : 0.

In some embodiments the hydrophobic spacer comprises e.g. > 50% of subunits that are - CH₂-. Non-limiting examples of hydrophobic spacers include e.g. fatty acids, such as acids having from 14 to 20 carbon atoms, e.g. stearic, palmitic, lauric, octadecaonic, mystric acids or spacers with saturated and unsaturated bonds such as oleyl and linoleic acids and the like.

In further embodiments the preferably hydrophobic spacers may be end-functionalized or end- capped.

When hydrocarbon chain is defined as -(CH₂)_m- the end capping group may generally be a carbon-containing group typically comprised of 1-50 carbons, nitrogen and/or sulphur atoms, preferably alkyl (e.g., methyl, ethyl or benzyl) although saturated and unsaturated forms thereof, as well as aryl, heteroaryl, cyclyl, heterocyclyl, and substituted forms of any of the foregoing are also envisioned.

In some embodiments, end-capping groups of the dispersants may be second labels such as fluorophores, which refers to a molecule or moiety, generally a polyaromatic hydrocarbon or heterocycle, that has the ability to fluoresce, and/or chelating agents (binding MR (magnetic resonance)-active ions such as Gd³⁺ or radiotracers). Any metal capable of accepting electron pairs from a chelating agent can bind the chelating agents of the invention. However, certain metals coordinate bond more strongly with sulfur containing substituents, and these metals are preferred. Preferably the metal is a radiometal, i.e., a radioactive isotope of a coordinate metal. Such metals are useful as imaging agents in diagnosis, and as therapeutic agents for targeted radiotherapy.

In other embodiments end-capping groups of the dispersants further comprises at least one (chemically) reactive group for coupling of hydrophobic or hydrophilic moieties extending into or outside the host membrane. As used throughout the text, a "reactive group" refers to any group that is capable of reacting with another chemical group to form a covalent bond, i.e. is covalently reactive under suitable reaction conditions, and generally represents a point of attachment for another substance, e.g. such as NHS ester, maleimide, azide, acrylates, methacrylates, amines, thiols, carboxy groups.

Thus, the invention also provides a magnetically responsive composition according to the invention, wherein one or more of the at least one dispersant adsorbed on the nanoparticle surface further comprises a reactive group.

When the hydrocarbon chains is defined as -[(CH)_n-(CH₂)_m]o-[(CH)p-(CH2)q]r, the end capping group is generally a carbon-containing group typically comprised of 1-50 carbons, nitrogen and/or sulphur atoms which optionally chelate metal ions such as Gd or radiotracers covalently bonding to one terminus of the hydrocarbon chain. In this case, the group is typically alkoxy (e.g., methoxy, ethoxy or benzyloxy) and with respect to the carbon- containing group which optionally can have nitrogen, sulphur, metal ion or radiotracers. The end-group can optionally be saturated and unsaturated, as well as aryl, heteroaryl, cyclyl, heterocyclyl, and substituted forms of any of the foregoing. In addition, the end-capping group can also be a silane. The other ("non-end-capped") terminus consists of an anchor group which has a optimized binding affinity to the oxides such as nitrocatechols to iron oxide surfaces. A review for the preparation of various end-group functionalized or activated PEG is known in the art (see for example Zalipsky S., Bioconjug. Chem., 6, 150-165 (1995)). For specific and preferred methods of stabilization and dispersants as well as specific and preferred methods of preparation of stabilized nanoparticles we refer to the detailed descriptions in copending European application No. 09 01 1190.7, which is incorporated herein by reference.

In a first aspect, the present invention provides a magnetically responsive composition comprising a membrane and at least one stabilized magnetic nanoparticle embedded in said membrane, wherein the membrane is in form of a magnetically responsive vesicular composition.

The term "vesicle" or "vesicular structure" as used herein refers to a roughly spherical membrane of lipids or lipid-related materials, preferably a roughly spherical, free-standing bilayer consisting of lipids or lipid-related materials, or polymeric materials. The term "vesicular composition" refers to vesicular structures comprising within their membrane at least one nanoparticle for the invention. A vesicle is unilamellar if it contains a single bilayer or multilamellar if it contains several bilayers. A vesicle is typically a closed surface so that the vesicle contents and molecules outside the vesicle do not exchange if the temperature is far away from the lipid phase transition. Depending on the nature of the membrane, one (or more) permeable, semi-permeable, and or non-permeable cavity (cavities) may be formed (and/or can change from one state to another). The cavity may be optionally partially or completely filled, with a solid, liquid or gaseous cargo, in form of emulsions, droplets, bubbles and the like.

Vesicles can be prepared by sonication of dispersions of lipid or co-polymer components in water or buffer or by extrusion of such solutions through membranes with defined pore sizes. In addition to lipid content, the vesicle bilayer can contain proteins, glycolipids and other biological molecules that are typically associated with biological membranes. As the vesicle is mimicking a normal cell membrane, proteins are typically fully functional. Proteins can also form additional supporting membranes around a vesicular structure, such as S-layer protein and annexin V layers. The inside of the vesicle can be used to trap molecules providing a probe for the integrity of the vesicle enclosure, as sensors for changes in properties of the interior (e.g. pH, ion concentrations and the like) or for studies of content mixing upon vesicle fusion or rupture.

Suitable vesicular structures for use in the present invention include, but are not limited to, liposomes (comprising at least one lipid type), polymersomes (based on at least one synthetic and/or natural polymer, such as synthetic (co-)polymers, proteins or mixtures thereof), porous nano- and microparticles or nano- and microsized droplets and bubbles surrounded by phospholipid and/or block-co-polymer bilayers.

Vesicles can be prepared with diameters from tens of nm to tens of mm. Typically, the vesicles of the present invention are less than 250 nm, preferably less than 200 nm, more preferably in the range from about 50 to 150 nm. Thus, preferably, the vesicles of the present invention are in the range of 20 to 250 nm, 30 to 250 nm, 10 to 200 nm, preferably about 20 to 200 nm, 30 to 200 nm, or 50 to 150 nm, with a mean and/or average size of the vesicles preferably being 100 nm.

The term "liposome" as used herein is a vesicular structure with uni- or multilamellar lipid membanes. In the case multilamellar liposomes, bilayers are generally concentric. Thus, the present compounds (and optionally colipids) may be used to form a unilamellar liposome (comprised of one bilayer), an oligolamellar liposome (comprised of two or three bilayers) or a multilamellar liposome (comprised of more than three bilayers). Lipids, which may be used in the formation of liposomal vesicular compositions of the invention may include acyclic and cyclic, saturated or unsaturated lipids of natural or synthetic origin. As used herein a lipid may be a neutral lipid, a cationic lipid or an anionic lipid. A cationic lipid has a positive net charge and may include lipids such as N-[l-(2,3-dioleoyloxy)propyl]-N,N,N-trimethyl ammonium salts, e.g. the methylsulfate (DOTAP), DDAB, dimethyldioctadecyl ammonium bromide; l,2-diacyloxy-3-trimethylammonium propanes, (including but not limited to: dioleoyl, dimyristoyl, dilauroyl, dipalmitoyl and distearoyl; also two different acyl chain can be linked to the glycerol backbone); N-[l-(2,3-dioloyloxy)propyl]-N,N-dimethyl amine (DODAP); 1 ,2-diacyloxy-3 -dimethyl ammonium propanes, (including but not limited to: dioleoyl, dimyristoyl, dilauroyl, dipalmitoyl and distearoyl; also two different acyl chain can be linked to the glycerol backbone); N-[l-(2,3-dioleyloxy)propyl]-N,N,N- trimethylammonium chloride (DOTMA); l,2-dialkyloxy-3-dimethylammonium propanes, (including but not limited to: dioleyl, dimyristyl, dilauryl, dipalmityl and distearyl; also two different alkyl chain can be linked to the glycerol backbone); dioctadecylamidoglycylspermine (DOGS); 3 -[N-(N',N'-dimethylamino- ethane)carbamoyl]cholesterol (DC-Choi); 2,3-dioleoyloxy-N-(2-(sperminecarboxamido)- ethyl)-N,N-dimethyl-l-propanam-inium trifluoro-acetate (DOSPA); β-alanyl cholesterol; cetyl trimethyl ammonium bromide (CTAB); diC14-amidine; N-tert-butyl-N'-tetradecyl-3- tetradecylamino-propionamidine; 14Dea2; N-(alpha-trimethylammonioacetyl)didodecyl-D- glutamate chloride (TMAG); 0,0'-ditetradecanoyl-N-(trimethylammonio- acetyl)diethanol amine chloride; 1 ,3-dioleoyloxy-2-(6-carboxy-spermyl)-propylamide (DOSPER); N,N,N',N'-tetramethyl-N,N'-bis(2-hydroxylethyl)-2,3-dioleoyloxy-l,4-butan- ediammonium iodide; 1 -[2-(acyloxy)ethyl]2-alkyl(alkenyl)-3-(2-hydroxyethyl)- imidazolinium chloride derivatives (as described by Solodin et al. (1995) Biochem. 43:13537- 13544), such as l-[2-(9(Z)-octadecenoyloxy)ethyl]-2-(8(Z)-heptadecenyl-3-(2-hydroxyethyl) imidazolinium chloride (DOTIM), l-[2-(hexadecanoyloxy)ethyl]-2-pentadecyl-3-(2- hydroxyethyl)imidazolinium chloride (DPTIM), 2,3-dialkyloxypropyl quaternary ammonium compound derivatives, containing a hydroxyalkyl moiety on the quaternary amine (see e.g. by Feigner et al. J. Biol. Chem. 1994, 269, 2550-2561), such as: l,2-dioleoyl-3-dimethyl- hydroxyethyl ammonium bromide (DORI), l,2-dioleyloxypropyl-3-dimethyl-hydroxyethyl ammonium bromide (DORIE), 1 ,2-dioleyloxypropyl-3 -dimethyl -hydroxypropyl ammonium bromide (DORIE-HP), l,2-dioleyloxypropyl-3-dimethyl-hydroxybutyl ammonium bromide (DORIE-HB), l,2-dioleyloxypropyl-3-dimethyl-hydroxypentyl ammonium bromide (DORIE- Hpe), 1 ,2-dimyristyloxypropyl-3 -dimethyl -hydroxylethyl ammonium bromide (DMRIE), 1,2- dipalmityloxypropyl-3-dimethyl-hydroxyethyl ammonium bromide (DPRIE), 1,2- disteryloxypropyl-3-dimethyl-hydroxyethyl ammonium bromide (DSRIE); cationic esters of acyl carnitines (as reported by Santaniello et al. U.S. Pat. No. 5,498,633); cationic triesters of phospahtidylcholine, i.e. l,2-diacyl-sn-glycerol-3-ethylphosphocholines, where the hydrocarbon chains can be saturated or unsaturated and branched or non-branched with a chain length from C₆ to C₂₄, the two acyl chains being not necessarily identical. Neutral or anionic lipids have a neutral or anionic net charge, respectively. These can be selected from sterols or lipids such as cholesterol, phospholipids, lysolipids, lysophospholipids, sphingolipids or pegylated lipids with a neutral or negative net change. Useful neutral and anionic lipids thereby include: phosphatidylserine, phosphatidylglycerol, phosphatidylinositol (not limited to a specific sugar), fatty acids, sterols, containing a carboxylic acid group for example, cholesterol, cholesterol sulfate and cholesterol hemisuccinate, 1 ,2-diacyl-sn-glycero- 3-phosphoethanolamine, including, but not limited to, DOPE, l,2-diacyl-glycero-3- phosphocholines and sphingomyelin. The fatty acids linked to the glycerol backbone are not limited to a specific length or number of double bonds. Lipids with a number of acyl chains different from two can also be used, including, but not limited two lysolipids and lipopolysaccharides. Lipids can be synthetically derived, but they can also be purified from natural membrane extract and, if required, modified. Naturally occurring and post-modified lipopolysaccharides that are used can have up to eight acyl chains.

Other materials for use in the preparation of liposomal vesicular compositions of the invention, in addition to those exemplified above, e.g. stabilizing material for liposomal structures, would be apparent to one skilled in the art based on the present disclosure. The selection of suitable lipids and e.g. stabilizing material in the preparation of liposomal vesicular compositions of the invention would be apparent to a person skilled in the art and can be achieved without undue experimentation, based on the present disclosure. Suitable nano- and microparticle substrates for formation of a lipid membrane for encapsulation are for example porous Si0₂ and Ti0₂ particles.

A wide variety of methods are available in connection with the preparation of liposomal vesicular compositions of the invention. Accordingly, the liposomes may be prepared using any one of a variety of conventional liposome preparatory techniques which will be apparent to those skilled in the art. These techniques include ethanol injection, thin film technique, electroswelling, homogenizing (extrusion), solvent dialysis, forced hydration, reverse phase evaporation, simple freeze-thawing, microemulsification and microfluidics using e.g. conventional microemulsification equipment. Additional methods for the preparation of liposomal vesicular compositions of the invention from the compounds of the present invention include, for example, sonication, chelate dialysis, homogenization (extrusion), solvent infusion, spontaneous formation, solvent vaporization, controlled detergent dialysis, and others, each involving the preparation of liposomes in various ways. Typically, methods which involve ethanol injection, thin film technique, homogenizing and extrusion are preferred in connection with the preparation of liposomal compositions of the invention from the compounds of the present invention.

The size of the liposomes can be adjusted, if desired, by a variety of techniques, including extrusion, filtration, sonication and homogenization. Other methods for adjusting the size of the liposomes and for modulating the resultant liposomal biodistribution and clearance of the liposomes would be apparent to one skilled in the art based on the present disclosure. Preferably, the size of the liposomes is adjusted by extrusion under pressure through pores of a defined size. The liposomal compositions of the invention may be of any size, preferably less than about 200 nanometer (nm) in outside diameter.

The term "polymersome' as used herein refers to vesicles defined by a membrane formed from natural or synthetic polymers, preferably amphiphilic synthetic polymers, more preferably block-co-polymers. Structurally, polymersomes resemble liposomes, the only difference being that liposomes are based on lipids. Polymersomes possess most of the properties of liposomes, but in addition they typically have greater stability and depending on the membrane thickness, adjustable permeability. Typically, polymersomes have a unilamellar membrane. This unilamellar membrane is termed "symmetrical" when the two superposed layers that form it are constituted by identical copolymers. In contrast, an "asymmetrical" unilamellar membrane has two superposed layers that are distinguished from each other by the specific natures of the copolymers that constitute them or are comprised of an asymmetric triblock or higher order copolymers assembled into an asymmetrical membrane. This difference between said two types of copolymer may reside in the nature of the hydrophobic block and/or in the nature of the hydrophilic block forming the copolymers.

In some embodiments, polymersomes of the invention comprise amphiphilic block-co- polymers or amphiphilic block-co-polymer mixtures, preferably amphiphilic di-and/or tri- block-co-polymers which have a low critical micelle concentration (CMC). These block-co- polymers may consist of a wide variety of polymers and can but do not have to be responsive. Prominent examples for thermoresponsive blocks of block-co-polymers are polymers, where the hydrophilic block consists of poly(2 -dimethyl amino ethyl) methacrylate (PDMAEMA), Poly(N-isopropylacrylamide) (PNIPAAM) or other thermoresponsive polymers. Hydrophilic blocks can also be pH-sensitive such as poly(acrylic acid) (PAA), poly(L-lysine) (PLL) and poly(L-glutamic acid) (PGA) resulting in pH responsive polymersomes. Next to poly(methyl carpolactone) (PMCL) and poly(carpolactone) (PCL), poly(ethylethylene) (PEE), poly(dimethyl siloxan) (PDMS), polystyrole (PS), poly(N-vinyl 2-pyrrolidone) (PVP), poly(propylene oxide) (PPO) and polybutadiene (PBD) are prominent examples for hydrophobic blocks of responsive and non-responsive polymersomes. The wall thickness, which determines the maximum SPION core size that can be incorporated into the polymersome membrane and that determines the inherent polymersome leakiness, can be adjusted by tuning the number of repeat units of the hydrophobic block. Furthermore, the thickness of the hydrophilic shell inside and outside the vesicle can be tuned independently from the thickness of the hydrophobic wall by controlling the number of repeat units of the hydrophilic block(s). Preferably, the number of hydrophobic repeat units should be small enough so that the block-co-polymer still can be solubilized in aqueous media. Thus, in specific embodiments, the number of repeat units of the hydrophobic block may range between 5 and 100, more preferably between 7 and 60, more preferably between 7 and 40, most preferably between 10 and 20. Prominent examples for hydrophilic blocks of non- responsive block-co-polymers well suited for polymersome formation especially but not exclusively in the biomedical field are poly(ethylene glycol) (PEG) (also called poly(ethylene oxide) (PEO)), poly(2-methyl-2-oxozaline) (PMOXA) and poly(lactic-co-glycolic acid) (PLGA). Amphiphilic block-co-polymers which are the building blocks of polymersomes can consist of combinations of the above mentioned polymers or any other polymer which is suited for a specific application. Prominent examples of block-co-polymers are poly(butadiene)-PEO (PB-PEO), poly(D, L- lactide)-PEG (PDLLA-PEG), PEG-PLA, PEG- polypropylene sulfide)-PEG (PEG-PPS-PEG), PEG-disulfide polypropylene sulfide) (PEG- SS-PPS), PEO-PCL, PEO-PCL-PLA, PEO-PDEAMA, PEO-PNIPAm, PEO-PCL-PAA, PLA- PEG-PLA, PMOXA-PCL, PMOXA-PDMS-PMOXA or poly(2-methacryloyloxy)ethyl- phosphorylcholine)-poly(2-(diisopropylamino)-ethyl methacrylate) (PMPC-PDPA)(Onaca, O. et al., Macromolecular Bioscience, 2009, 9: 129-139). Other suitable block-co-polymers may include oxidation sensitive polymersomes such as PEO-poly(propylene sulphide)-PEO triblocks, (Napoli, A. et al, Langmuir, 2004, 20: 3487-3491).

Furthermore, block-co-polymers may be functionalized e.g. with peptide functionalized block-co-polymers where one of the hydrophilic blocks of a tri- or multi block copolymer or the hydrophilic block of a di-block-co-polymer is replaced with a peptide. (Christian, D.A. et al., European Journal of Pharmaceutics and Biopharmaceutics, 2009, 71: 463-474) Furthermore, the hydrophilic and hydrophobic blocks can be replaced by oppositely charged peptides or polymers resulting in pH responsive peptide functionalized or unfunctionalized polymersomes(Christian, D.A., et al., European Journal of Pharmaceutics and Biopharmaceutics, 2009, 71: 463-474). The polymer wall can also be comprised of a protein shell with charged and hydrophobic pockets. Such vesicles or polymersomes comprising predominantly peptides or proteins in the membrane may also be termed proteosomes. In some embodiments proteosomes may comprise S-layer protein membranes, amphiphilic alpha-helix or beta-sheet forming peptide bilayers, or annexin coated vesicles. Furthermore, to target polymersomes, block-co-polymers may be end-functionalized with functional groups like avidins, antibodies or other biologically relevant ligands(Discher, D.E. et al., Annual Review of Biomedical Engineering, 2006, 8: 323-341). End-functionalization of the block-co-polymers may be achieved through their terminal group, such as a terminal -OH group (in case of PEG blocks), or through NHS-ester, amine, thiol, maleimide, acrylate, methacrylate, carboxy and other chemically reactive groups present on the block-co-polymer of choice.

Thus, in some embodiments the invention is directed towards a magnetically responsive composition comprising a membrane and at least one stabilized magnetic nanoparticle embedded in said membrane, wherein the membrane is a liposome and wherein the membrane comprises at least one lipid type which forms the lipid bilayer. For incorporation into a lipid bilayer, the stabilized, optionally further functionalized nanoparticles have core sizes of less than 10 nm, preferably between 2 and 8 nm, most preferably between 2 and 5 nm. Incorporation into the lipid bilayer is performed using standard liposome formation protocols through extrusion, sonication or freeze- thaw cycles.

Typically, liposomes comprising stabilized magnetic NPs of the invention, hereinafter also called magnetic liposomes of the invention, are smaller than 1 μηι, preferably in the size range between 50 nm and 400 nm, preferably between 50 and 200 nm, most preferably around 100 nm. The size of liposomes containing magnetic NPs in their membrane according to the invention may be controlled using standard procedures, such as extrusion, sonication, where extrusion is the preferred procedure. They are typically assembled by mixing of the at least one lipid of choice in a volatile organic solvent with stabilized, magnetic NPs, preferably stabilized SPIONs (optionally functionalized with additional labels such as fluorophores, radiotracers and the like), which are dispersed in an organic solvent such as chloroform or dichlormethane. Typically the weight ratio of stabilized SPIONs : lipids ranges between 0 and 1, preferably between 0.1 and 0.5, most preferably around 0.3. The resulting mixture is dried into a thin film at the bottom of a flask and the organic solvent is completely removed under a flow of inert atmosphere. The lipid film is rehydrated in the desired buffer to form multilamellar magnetic liposomes, which may optionally be reshaped by extrusion to the desired size by pushing the sample through pores of the same nominal size track-etched polycarbonate membranes. Larger vesicles can be formed if the liposome/stabilized NP dispersion has gone through multiple freeze-thaw cycles without having been extruded before, while sonication leads to a large size distribution but rather unilamellar vesicles. Typically, for release of a hydrophilic cargo unilamellar vesicles are preferred, while for release of a hydrophobic cargo multilamellar vesicles may be preferred.

Typically, incorporation into the liposome membrane was achieved using standard liposome formation protocols, such as through extrusion, sonication or freeze-thaw cycles.(Barenholz, Y. et al, Biochemistry, 1977, 16: 2806-2810; MacDonald, R. et al., Biochim Biophys Acta, 1991, 1061: 297-303; Reimhult, E. et al., Langmuir, 2003, 19: 1681-1691; Reimhult, E. et al., International Journal of Molecular Sciences, 2009, 10: 1683-1696) Larger vesicles may be formed by subjecting a liposome/ P dispersion to multiple freeze-thaw cycles without prior extrusion while sonication leads to a large size distribution of unilamellar vesicles and a large fraction of micelles. Typically, unimellar vesicles may be suitable for release of hydrophilic cargo, while multilamellar vesicles may be suitable for release of hydrophobic cargos.

In specific embodiments, the invention provides magnetic liposomes of the invention wherein the stabilized magnetic NPs are nitroDOPA-palmityl stabilized SPIONs with core diameters between 2 and 15 ran, preferably between 3 and 10 ran, most preferably between 3 and 5 ran. As indicated hereinabove, the magnetic liposomes of the invention may further include end- functionalized, stabilized magnetic NPs, i.e. wherein the NPs have been functionalized, e.g. with reporter groups such as fluorescent dyes, radiotracers or chelated magnetic ions such as Gd3+, FITC, rhodamine, CY-dyes, NBD, Alexa fluorophores, radioactive elements and paramagnetic ions.

In preferred embodiments, magnetic liposomes of the invention comprise Fe₃0₄-NPs stabilized with palmityl-nitroDOPA (Figure 2), optionally in combination with Fe₃0₄-NPs stabilized with palmityl-nitroDOPA and labeled with an additional tag such as fluorophores (FITC, rhodamine, Alexa dyes, CY dyes etc), radiotracers, chelators etc..

Another preferred embodiment comprises Fe₃0₄-NPs stabilized with palmityl-nitrodopamine, optionally in combination with Fe₃0₄-NPs stabilized with palmityl-nitroDOPA and labeled with an additional tag such as fluorophores (FITC, rhodamine, Alexa dyes, CY dyes etc), radiotracers, chelators etc.

In other preferred embodiments, magnetic liposomes of the invention comprise Fe₃0₄-NPs stabilized with oleyl-nitroDOPA optionally in combination with Fe₃0₄-NPs stabilized with oleyl-nitroDOPA and labeled with an additional tag such as fluorophores (FITC, rhodamine, Alexa dyes, CY dyes etc), radiotracers, chelators etc.. The magnetic liposomes of the invention including PEG-lipid inclusion may be obtained by simple mixing of different lipid species in an organic solvent (typically chloroform) and removal of that solvent before rehydration to form liposomes . A high enough fraction of lipids with PEG attached to the headgroups to yield liposome surface densities of the PEG in the range of polymer brushes provides the liposome with stealth-like properties for application in biofiuids and in vivo. This yields low protein adsorption, protection from enzymatic degradation and thus longer circulation times in vivo. Any of the typically used lipids as mentioned hereinabove may be incorporated in this way into liposomes to tune the vesicle mechanical, physical and chemical properties, including phospholipids, sphingolipids, lysolipids, glycolipids, saccharolipids, glycophospholipids, cholesterol, PEG-lipids and others using standard procedures see for example formation of phospholipid unilamellar vesicles of various charge (Rossetti, F.F., et al., Langmuir, 2005, 21: 6443-6450; Khan, T.R. et al., Biointerphases, 2008, 3: FA90-FA95; Kumar, K. et al, Lab on a Chip, 2009, 9: 718-725; Mashaghi, A. et al, Analytical Chemistry, 2008, 80: 3666-3676; Reimhult, E. et al., Langmuir, 2006, 22: 3313-3319), bacterial lipid compositions (Merz, C. et al., Biointerphases,

2008, 3: FA41-FA50) and PEG-liposomes and glycolipid/lipopolysaccharide liposomes (Kaufmann, S. et al., J. Am. Chem. Soc, in preparation; Kaufmann, S. et al., Soft Matter,

2009, 5: 2804-2814)). Since the main interaction properties of the liposome is controlled by its surface presentation of functional groups, while the described incorporation of magnetic NPs is dependent mainly on the fatty acid chain internal region of the membrane, any of these compositional variations in the membrane headgroup region, is applicable to the vesicular compositions of the invention. Likewise it has been shown that chain length and saturation of the hydrophobic core of the lipid membrane may be changed without noticeable effect on the incorporation of magnetic NPs into the membrane (Figure 5), since both (at room temperature typically) non-saturated liquid membranes and both (at room temperature typically) saturated gel membranes have been used for NP incorporation and actuation (e.g. POPC which has unsaturated and MPPC and DSPC which both have saturated hydrocarbon chains of different lengths).

In other embodiments the invention is directed towards a magnetically responsive composition comprising a membrane and at least one stabilized magnetic nanoparticle embedded in said membrane, wherein the membrane is a polymersome formed from natural or synthetic polymers, preferably amphiphilic synthetic polymers, more preferably block-co- polymers as defined hereinabove. In specific embodiments, a polymersome comprising at least one stabilized magnetic nanoparticle of the invention (also called magnetic polymersome of the invention) comprises at least one natural or synthetic polymers, preferably amphiphilic synthetic polymer, more preferably block-co-polymer as defined hereinabove and palmityl-nitroDOPA or oleyl- nitroDOPA stabilized iron oxide NPs approximating the size of the hydrophobic core of the polymersome membrane. Typically, SPIONs were dispersed in an organic solvent, e.g. CHC1₃ or another solvent in which SPIONs and the at least one polymer, preferably block copolymer of choice are soluble. Preferably, SPIONs were dissolved with poly(2-dimethyl amino ethyl) methacrylate (PMCL- PDMAEMA) block-co-polymers at a weight ratio of block-co-polymers: SPIONs up to 3:1 (see e.g. Figure 20, Example 6). The solvent was dried under steady N₂ flow and the block-co-polymer/iron oxide NP mixture swollen between 2 and 10 days in an aqueous buffer or water to form a magnetically responsive composition comprising a block co-polymer membrane incorporating palmityl-nitroDOPA stabilized iron oxide NPs.

The size of polymersomes in the hydrated dispersion may be controlled after rehydration by extrusion.

Actuation and disruption of the obtained magnetic polymersomes may be achieved by applying an alternating magnetic field (AMF) (Figure 20b).

In a further aspect, the present invention provides a magnetically responsive composition comprising a membrane and at least one stabilized magnetic nanoparticle embedded in said membrane, wherein the membrane is in form of a magnetically responsive supported lipid bilayer.

The term "supported lipid bilayers (SLBs)" as used herein refers to a planar bilayer of lipids or lipid-related materials including polymeric materials on a solid support such as glass, mica, or oxidized polydimethylsiloxane (PDMS), and the like. Typical lipids or lipid-related materials or polymeric materials are the same as those forming liposomes or polymersomes as defined hereinabove. In a specific embodiment the supported lipid bilayer is comprised of lipids with different melting temperatures.

The procedure for obtaining magnetic supported lipid bilayers is analogous to the procedure for obtaining magnetic liposomes or magnetic polymersomes as defined hereinabove.

In some embodiments, supported lipid bilayers of the invention may comprise additional molecules such as polymers and proteins of similar size to small NPs. Thus in other embodiments, the supported lipid bilayers (SLBs) of the invention may comprise proteins selected from, but not limited to, glycoprotein and protein membranes such as S-layers which form crystalline, self-assembled membranes. More specifically, S-layers may be crystallized e.g. on a lipid membrane or emulsion templates. Upon removal of the template stable, and free-standing planar or vesicular membranes incorporating magnetic NPs according to the invention may be obtained.

It is understood that any membranes mimicking biological membranes and containing mixtures of lipids, proteins and polymers (optionally including functional molecules spanning the thickness of the membrane, such as ion channel and transporter membrane proteins) may be amenable to functionalization with stable SPIONs.

Supported lipid bilayers comprising magnetic NPs according to the invention (hereinafter also called supported lipid bilayers of the invention) can be used in various applications. Thus in some embodiments, supported lipid bilayers of the invention may be used for actuation of a surface coating both in terms of deformation (Figure 15) and in terms of changes to permeation.

Thus in one embodiment the magnetically responsive supported lipid bilayer of the invention may be used to span an aperture or cavity comprising one or more encapsulated agents. Subsequently release of said agents can be actuated through changing the permeability of the aperture or cavity spanning bilayer of the invention (Figure 16)

In some embodiments the cavity is a nanocavity or nanopore, in other embodiments the aperture is a larger aperture, where the bilayer may be formed by e.g. the Montal-Mueller method (Montal, M. et al, Proc. Natl Acad Sci USA, 1972, 6: 3561-3566) from mixtures of lipids and stable hydrophobic NPs as those used to assemble the magnetoliposomes, or by microfluidic flow driven membranes across nanostructures and nanopores as recently demonstrated by H5ok and coworkers (Jonsson, P. et al., Nano Letters, 2010, 10: 1900-1906). Thus, in a specific embodiment the supported lipid bilayer of the invention is covering a porous substrate, where the bilayer is spanning the pores

In other embodiments the supported lipid bilayer of the invention may be used to induce convection by magnetic fields in said lipid bilayer.

In yet other embodiments the supported lipid bilayer of the invention may be used to induce movement of a species associated with the membrane (i.e. embedded within the membrane or attached to the NPs within the membrane). As indicated above, all of the magnetically responsive compositions of the invention (in all their embodiments) are susceptible to magnetic fields (alternating or direct), which results in change of permeability, change of local fluidity, etc. Thus in one embodiment, the present invention provides a method of changing the permeability of a magnetically responsive composition comprising the step of exposing said magnetically responsive composition to an alternating magnetic field, whereby permeability of the magnetically responsive structure is increased through local heat generation. In another one embodiment, the present invention provides a method of changing the permeability of a magnetically responsive composition comprising the step of exposing said magnetically responsive composition to a direct magnetic field, whereby permeability of the magnetically responsive composition is increased through mechanical deformation. In yet another embodiment, the present invention provides a method of locally changing the fluidity of magnetically responsive membranes (including associated or non-associated, i.e. freely moveable compounds) by application of magnetic fields.

In a further aspect the vesicular compositions may further comprise at least one active agent. In one embodiment, the at least one active agent may be attached or adsorbed onto the surface of the vesicular compositions, which may be achieved e.g. by covalently linking the at least one active agent to at least one constituent of the vesicular composition (i.e. a lipid or a polymer). In another embodiment, the at least one active agent may be embedded into the membrane (as the magnetic nanoparticles). In yet another embodiment the at least one active agent may be incorporated into the cavity or lumen of the vesicular composition.

A suitable active agent may be any molecule or macromolecule having a therapeutic or diagnostic utility, or producing a desired biological response. More specifically the at least one active agent is a therapeutic or diagnostic agent, a nutritional agent, an enzyme, a growth factor or a targeting group or combinations thereof. A suitable therapeutic agent may be selected from the group consisting of antineoplastic agents, anti- inflammatory agents, antitumor agents, immunosuppressive agents, antibiotic agents and anti-infective agents. A suitable diagnostic agent may be selected from the group consisting of fluorophores, QDs and radioactive tracers. A suitable targeting group may be selected from the group consisting of an antibody, Fab fragments, nucleic acid (including DNA, NA) aptamers, sugars (including saccharides), proteins, fusion proteins, cross-linking agent, or combination thereof.

Selected examples of such active agents include but are not limited to a nucleic acid, a protein, a peptide, an oligonucleotide, an antibody, an antigen, a viral vector, a bioactive polypeptide, a polynucleotide coding for the bioactive polypeptide, a cell regulatory small molecule, a gene therapy agent, a gene transfection vector, a receptor, a cell, a drug, a drug delivering agent, an antimicrobial agent, an antibiotic, an antimitotic, an antisecretory agent, an anti-cancer chemotherapeutic agent, steroidal and non- steroidal anti-inflammatories, a hormone, a proteoglycan, a glycosaminoglycan, a free radical scavenger, an iron chelator, an antioxidant, or a tracer agent, such as an imaging agent, a fluorophore and a radiotherapeutic agent.

Furthermore, the magnetic liposomes of the invention may be subjected to further liposome functionalization, such as changing the lipid composition, adding polymer shielding to the surface of the liposome, incorporating membrane proteins or attaching peptides, proteins, sugars and DNA to the membrane, which keep the integrity of the membrane structure is compatible with magnetic NP functionalization of the vesicle through membrane insertion according to the invention.

In some embodiments, addition of DNA strands to the vesicle membrane for targeting or tethering of magnetic liposomes of the invention may be performed by post-modification after liposome assembly through addition of DNA with an inserting hydrophobic anchor, e.g. double cholesterol (Pfeiffer, I. et al., Journal of the American Chemical Society, 2004, 126: 10224-10225). Figure 24 shows the binding kinetics of DNA tethered SPION functionalized POPC (Palmitoyl oleoyl phosphatidyl choline) vesicles, which were bound to a POPC SLB which had been functionalized with the complementary DNA prior to the addition of DNA tethered POPC vesicles containing SPIONs in their membrane. Such DNA tethers may be made reversibly or irreversibly anchored in the membrane depending on the size and solubility of the hydrophobic anchor, typically a double cholesterol derivative.

In other embodiments, membrane protein functionality may be conferred to a magnetic liposome of the invention by reconstitution of protein into said liposomes to form proteoliposomes. This is typically achieved by detergent dilution using a detergent of choice, e.g. Triton X. Stabilized membrane proteins may then insert into preformed liposomes as the detergent concentration is brought down successfully by introduction and separation of biobeads. This can be done into small and large unilamellar liposomes at controlled membrane protein concentrations (Graneli, A. et al., Langmuir, 2003, 19: 842-850) and combined with NP containing liposomes.

In yet other embodiments, functional ligands, such as proteins, Fab fragments, peptides, aptamers, and the like may be attached to the surface of magnetic liposomes of the invention through binding groups present on the constituent head groups of lipids used for the liposome formation. Several functional groups have been used for such attachment of functional groups to liposomes and supported lipid membranes, e.g. biologically inspired such as biotin- avidin,(Salaita, K. et al, Science, 2010, 327: 1380-1385) chelates such as NTA-Ni-His(Nye, J.A. et al., Langmuir, 2008, 24: 4145-4149) and covalent such as maleimid-thiol(Svedhem, S. et al., Langmuir, 2003, 19: 6730-6736).

It is understood that, in addition to functionalizing a preformed liposome, such ligands may also be attached to an individual lipid headgroup before assembly of the liposomes (using standard coupling chemistries, such as coupling to NHS-esther, acrylates, azide, methacrylates, amine, thiol and carboxy groups).

Furthermore, a lipid membrane (in form of a vesicle or a supported lipid bilayer) may also be used as a template for assembly of protein layers, including protein crystals, formed to cover the surface of the lipid membrane, thereby providing higher stability, robustness, specific binding interactions stealth properties and additional functions to the lipid membrane. Examples of such surface modifications include modification with e.g. streptavidin (Reviakine, I. et al., Langmuir, 2001, 17: 8293-8299), annexin V (Reviakine, I. et al., Journal of Structural Biology, 1998, 121: 356-362) and S-layer proteins (Sara, M. et al., Journal of Nanoscience and Nanotechnology, 2005, 5: 1939-1953).

It is understood, that all the above modifications (i.e. changes in surface functionality and/or changing the chemical and physical interaction properties of a magnetically responsive composition according to the invention) are variations to the underlying concept of the invention to enable or optimize their use in various applications (where specific characteristics are desired such as e.g. free circulation of liposomes; or targeting of cells and tissues in vivo; or binding and assembling of vesicles into matrices, patterns and other structures in a specific way on scaffolds and pre-patterned substrates).

In further embodiments the magnetically responsive composition may be encapsulated in a matrix such as a physically or chemically crosslinked hydrogel or embedded in a matrix tethered to a substrate. The composition may be encapsulated or embedded through non covalent forces or physically or chemically crosslinked through covalent linkages.

In some embodiments the encapsulating matrix may be a fibrous or membranous tissue engineering scaffold promoting tissue and cell growth and adhesion, or a polymer layer bound to an implant surface. More specifically, the magnetically responsive compositions may be linked or adsorbed to or incorporated (or encapsulated) into surface architectures, such as hydrogels, membranes and fibers for tissue engineering, cell culture or implants using standard procedures, including tethering into matrices (GRANELI, A. et al., 2004), encapsulation into electrolyte and other polymer layers (Example 6, Figure 25), binding to protein surfaces, direct adsorption onto a substrate without membrane rupture (Rossetti, F.F., et al, Langmuir, 2005, 21 : 6443-6450; Khan, T.R., et al, Biointerphases, 2008, 3: FA90- FA95; Kumar, K., et al, Lab on a Chip, 2009, 9: 718-725; Reimhult, E., et al, Langmuir, 2006, 22: 3313-3319; Merz, C, et al, Biointerphases, 2008, 3: FA41-FA50; Reimhult, E. et al., Journal of Chemical Physics, 2002, 117: 7401-7404) optionally in combination with other linking strategies to form various scaffolds of choice (e.g. layers on a substrate, fibers, membranes) for delivery and release applications, for tissue engineering, as a tool to control cell and tissue growth and differentiation. In some embodiments, polymer and inorganic substrates may be first surface modified with, e.g., a thin self-assembled monolayer, magnetically responsive composition of the invention providing binding groups for further binding of liposomes with any of the methods outlined above and below. In other embodiments, magnetically responsive vesicles of the invention tethered with mobile linkers, such as the described cholesterol-DNA, to a liquid crystalline surface, such as a lipid bilayer or a block-co-polymer membrane, will retain lateral mobility and can thus be used for magnetically actuated movement, concentration and separation applications. In yet other embodiments, magnetically responsive vesicles of the invention encapsulating dyes may be embedded into polyelectrolyte multilayers suitable for cell culture (Figure 25, Example 11).

Thus in another aspect, the invention provides a matrix comprising at least one magnetically responsive composition of the invention.

As indicated above, in a further aspect, the present vesicular compositions of the present invention are particularly suitable for use as carriers for a targeted delivery of (diagnostic and therapeutic) active agents. The vesicular compositions of the present invention are not limited to the specific embodiments described herein. In contrast, any kind of substrate providing a physical and chemical binding site may be used to build up layers of magnetically functionalized liposomes for release applications through at least one of the above disclosed methods of immobilization. Such magnetic particle functionalized vesicles adsorbed on a substrate or in a matrix or scaffold may then be used to trigger release of substances in time and space for a variety of applications. Such applications are particularly useful in cell cultures platform for release of triggers and cues for growth and differentiation, similar release for vesicles embedded in surface layers of implants and to release reactants in miniaturized devices.

Thus, the magnetically responsive compositions, in particular the magnetically responsive vesicular compositions of the present invention are particularly applicable for use in vitro and/or in vivo in methods for the treatment of diseases, for which a targeted delivery of one or more specific active agents is desirable or required, as well as for use in methods in vitro/in vivo diagnostic applications.

In one embodiment, the magnetically responsive compositions, in particular the magnetically responsive vesicular compositions of the present invention comprise at least one targeting moiety to direct the vesicular composition to a target site and at least one active agent to be released upon detection of local accumulation of the vesicular compositions.

Thus, the invention provides a method for targeted delivery of a magnetically responsive vesicular composition to a biological target of choice, said method comprising: providing a magnetically responsive vesicular composition of the invention with a ligand group to obtain a functionalized magnetically responsive vesicular composition which is able to couple through the ligand group with the biological target; entrapping an active agent within said functionalized magnetically responsive vesicular composition; exposing the functionalized magnetically responsive vesicular composition to the biological target and allowing binding of ligand group to biological target to occur; and releasing the active agent by external application of an alternating or strong direct magnetic field.

The vesicular structure may or may not be retained during release

Thus, in one embodiment, the invention provides a method for delivery of an active agent in a subject, comprising the steps of: (a) associating at least one active agent with a magnetically responsive vesicular composition of the invention (to obtain an functionalized composition), (b) administering said (functionalized) composition to a subject; and (c) exposing said (functionalized) composition to a direct or alternating magnetic field, whereby the permeability of said (functionalized) composition is increased such that said active agent is released in said subject (and the vesicular structure is retained during release). In another embodiment, the invention is directed towards a method for delivery of an active agent in a subject, comprising the steps of: (a) associating at least one active agent with a magnetically responsive vesicular composition of the invention (to obtain an functionalized composition), (b) administering said (functionalized) composition to a subject; and (c) exposing said (functionalized) composition to a direct or alternating magnetic field, whereby the vesicular structure is destroyed by the response of the magnetic NPs which are embedded in the membrane, releasing said active agent.

In another aspect the invention provides methods of delivery of an active agent in vitro, i.e. in ell or tissue culture. Thus, in one embodiment, the invention is directed towards a method for delivery of an active agent in cell or tissue culture, comprising the steps of: (a) associating at least one active agent with a magnetically responsive vesicular composition of the invention (to obtain an functionalized composition), (b) incorporating said (functionalized) composition in the cell or tissue culture directly or as part of the scaffold; and (c) exposing said (functionalized) composition to a direct or alternating magnetic field, whereby the permeability of said (functionalized) composition is increased such that said active agent is released in said subject. In another embodiment, the invention is directed towards a method for delivery of an active agent in cell or tissue culture, comprising the steps of: (a) associating at least one active agent with a magnetically responsive vesicular composition of the invention (to obtain an functionalized composition), (b) incorporating said (functionalized) composition in the cell or tissue culture directly or as part of the scaffold; and (c) exposing said (functionalized) composition to a direct or alternating magnetic field, whereby the vesicular structure is destroyed by the response of the magnetic NPs which are embedded in the membrane, releasing said active agent.

It is understood that the concept underlying the invention may easily be combined with any magnetic targeting technology, whereby the externally focused magnetic fields are used to direct the accumulation of the therapeutic agent instead of affinity ligands. Furthermore, surface-functionalization of either the SPIONs or the magnetically responsive vesicular compositions with additional labeling groups such as fluorophores, or other tracers, their localization can be determined also by other means than magnetic contrast imaging prior to release.

In a further aspect the invention is directed towards the use of magnetically responsive compositions, in particular the magnetically responsive vesicular compositions as nanoreactors. Combining ultra-small volumes of reactants in confined environments, e.g. mimicking the chemistry in biological systems, may be useful in a wide variety of application including e.g. in high-yield and low environmental impact chemical production.

However, control of reactants and mixing volumes on this size scale is very difficult since they cannot be manipulated with conventional technologies, and even new technologies like micro and nanofluidic chips do not approach the dimensions of the small volumes and confined environments ideal for these applications.

However, there are a lot of emulsion techniques that can be used to assemble capsules and vesicles close to or layer by layer inside each other. This precludes external manipulation by other means than penetrating fields, and our described invention can be used to externally control the mixing of two or more reactants in a localized volume by controlling the permeability of vesicles in which they are encapsulated.

With self-assembled lipid membranes being attractive, non-fouling, biocompatible coatings and selective permeation barriers, actuation of liposomes and membranes covering channels in nanofluidic devices could be an attractive way of locally releasing (bio)chemical compounds and open nanoscopic valves in lab-on-chip devices.

In one embodiment fusion of magnetoliposomes with cell membranes may be used to label membranes of a cell. Since the labels will be magnetic nanoparticles the same actuation may be applied to the cell membrane as to the vesicular delivery systems described above. Thus, cells may move by magnetic fields but also may deform in shape and the local permeation of the cell membranes can be altered, providing a new manipulation tool for cells.

Additionally, polymersomes are more robust compared to liposomes which renders them easier to handle and more versatile to apply. Therefore, polymersomes which have magnetic NPs in or associated with their membrane are attractive alternatives to magnetic NP functionalized liposomes especially for nanoreactor applications or release of higher molecular weight compounds. The thickness of the hydrophobic core of the polymersome membrane can be tailored by adjusting the number of hydrophobic repeat units of the block- co-polymer used to assemble such polymersomes. Thus, larger magnetic NPs can likely also be incorporated into polymersomes in contrast to liposomes. This could massively increase the responsiveness of magnetic NP functionalized polymersomes to external magnetic fields and thus render magnetic actuation easier compared to of magnetic NP functionalized liposomes. Additionally, there is a lot of flexibility in designing the blocks of the block-co- polymers. Thus, if certain reactions need to be carried out under solvent or other conditions for which liposomes are not stable, polymerosomes can be designed such that they remain stably assembled under said conditions. Additionally, the hydrophilic blocks of polymersomes can be optimized for the cargo they are loaded with. Because iron oxide NPs only locally heat and the bulk temperature does not rise significantly if subjected to external alternating magnetic fields, thermoresponsive reactions can be performed even if reagents are involved which are temperature sensitive. Furthermore, the chemical flexibility to design block-co- polymers allows to further surface-functionalize polymersomes e.g. with ligands and labels with more chemical flexibility and at lower cost than can be afforded by lipid vesicles. Thus all the advantages described above for liposome vesicles in terms of multi-modality apply at least as well to NP-polymersome systems.

The present invention is further described in the following non-limiting examples.

Examples

Materials:

Substrates: The build-up of polyelectrolyte multilayers(e.g. Figure 25) was performed on indium tin oxide (ITO) coated glass (for the confocal laser scanning microscopy (CLSM) experiments) and ITO coated QCM quartz crystals (for quartz crystal microbalance experiments). All the ITO samples were produced by MicroVacuum (Hungary). The CLSM substrates were cleaned by ultrasonication (Elna Transsonic Digital S, Iswork, Singapore) in Cleaner (Cobas Integra Cleaner, Roche), Isopropanol (puriss., Sigma Aldrich) and ultrapure water (resistivity = 18.2 ΜΩ/cm, Milli-Q gradient A 10 system, Millipore Corporation) for 10 minutes in each subsequent solvent. The samples were rinsed with ultrapure water and dried with nitrogen after every cleaning step. Finally, they underwent UV/Ozone cleaning (Uvo Cleaner 42-220, Jelight Company, USA) for 30 minutes prior to spraying. The QCM sensors were immersed for 30 minutes in sodium dodecyl sulfonate (SDS; Sigma Aldrich), rinsed with ultrapure water and then cleaned by UV/Ozone for 30 minutes before spraying.

Nanostructured substrates (pore sizes: 40, 80, 100, 200 and 500nm) were produced by colloidal lithography and etching through 350 nm of PECVD grown silicon nitride (n«2.16) deposited on #1 thickness borosilicate microscope glass cover slips (Isa, L. et al., ACS Nano, 2010, 4: 5665-5670; Reimhult, E. et al., Nanotechnology, 2007, 18: 275303) . The average nanopore size was determined by the diameter of the colloids (e.g. Figure 16). The pore-to- pore separation was controlled during the colloidal lithography step by particle self-assembly at liquid-liquid interfaces (SALI) (Isa, L., et al, ACS Nano, 2010, 4: 5665-5670) or by random adsorption of charged colloids (Reimhult, E., et al., Nanotechnology, 2007, 18: 275303; Hanarp, P. et al., Colloid Surface A, 2003, 214: 23-36). Chemicals: l,2-distearoyl-sn-glycero-3-phosphocholine (DSPC), l-stearoyl-2-oleoyl-sn- glycero-3 -phosphochline (SOPC), 1 -palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC), l,2-Dioleoyl-sn-Glycero-3-Phosphoethanolamine-N-[Methoxy(Polyethylene glycol)2000 Da] (Ammonium Salt) (PEG(2)-PE) and l,2-Dioleoyl-sn-Glycero-3-[Phospho-L- serine] (DOPS, Sodium Salt), l-01eoyl-2-[12-[(7-nitro-2-l,3-benzoxadiazol-4- yl)amino]lauroyl]-sn-Glycero-3-Phosphocholine (NBD-POPC), l-palmitoyl-2-oleoyl-5^,n- glycero-3-phosp o-L-serine (sodium salt) (POPS) and l^-dioleoyl-sn-glycero-S- phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) (ammonium salt) (DOPE-RhoB) were purchased from Avanti Polar Lipids (Alabama, USA).

Calcein was obtained from ABCR (Kahlsruhe, Germany), D₂0, palmitic acid N- hydroxysuccinimide ester (palmityl-NHS), Fe(ac)₂ (batch 517933, Lot 03901JJ), N-N dimethylfonnamide (DMF), benzylalcohol, fuming HC1, Sephadex G75 (superfine) and phosphate buffered saline (PBS) (pH=7.4) tablets from Sigma (Buchs, Switzerland), ethanol (absolute) from Scharlau (Barcelona, Spain), chloroform from ecsa (Pennsylvania, USA) and nitroDOPA and PEG(1.5)-nitroDOPA were synthesized as described earlier.(Amstad, E. et al., Nano Letters, 2009, 9: 4042-4048)

Buffers: All buffers were made using ultrapure water (Millipore, Switzerland; i?=18.2 Ω, TOC < 6 ppb). Phosphate buffered saline (PBS, Fluka) was used for preparation of all liposome preparations for liposomes with magnetic particles incorporated. Tris buffered saline (TBS) was used for liposome stock solutions for the experiments on nanopore spanning lipid bilayers: 10 mM Tris(hydroxymethyl)-aminomethan (Tris), 150 mM NaCl and pH 7.4 set by HC1 (all chemicals from Fluka, Switzerland). Liposome adsorption experiments on nanoporous substrates were performed in TBS with additionally 3 mM CaCl₂ added (TBSCa). Liposomes for adsorption into polyelectolyte multilayers were formed in 150 mM NaCl in ultrapure water without buffer. Dyes such as carboxyfluorescein were added to the buffer in some experiments as detailed below.

Poly electrolytes: Polyelectrolyte multilayers were made from: PGA: Poly-L-Glutamic-Acid (PGA, SIGMA), PLL: Poly-L-lysine hydrobromide, Sigma Aldrich P7890, MW = 24000 and PSS: poly(sodium 4-styrene sulfonate), Sigma Aldrich 24,305, MW = 70000 dissolved in 150 mM NaCl solution. (PLL/PSS)n means n layer pairs of the polyelectrolyte couple PLL and PSS. The PEM directly on the substrate is denoted as underlying PEM or short uPEM. The vesicles were adsorbed onto the uPEM. The PEM layer covering the liposomes is accordingly denoted cPEM. Synthesis and assembly:

Palmityl-nitroDOPA synthesis: 1.8 g nitroDOPA and 2 g palmityl-NHS (molar ratio of nitroDOPA : palmityl-NHS = 1.1 : 1) were dissolved in 60 ml DMF before 1.8 ml morpholine was added. The solution was magnetically stirred for 19 h in air at room temperature before 1 ml fuming HC1 was added to acidify the solution. After removing DMF, the products were re- dispersed in Millipore water and extracted with CH₂CI₂ before CH₂CI₂ was evaporated. To ensure complete removal of unreacted nitroDOPA, the product was five times thoroughly washed with Millipore water before it was freeze-dried (freeze dryer ALPHA 1-2 / LDplus, Kuhner LabEquip, Switzerland) and analyzed by 1H- and 13C-NMR, MALDI-tof and microelement analysis.

Iron oxide NP synthesis: Iron oxide NPs were synthesized by a non-aqueous sol-gel route.(Bilecka, I. et al., Chemical Communications, 2008, 886-888) 1 mmol Fe(ac)2 was dispersed in 5 ml benzylalcohol. To ensure complete dispersion of the precursor, the dispersion was magnetically stirred for 1 h at 70 °C in an oil bath before the temperature was raised to 150 °C for NPs with an average core diameter of 5 nm and 180 °C for NPs with an average core diameter of 10 nm respectively. NPs were grown at the respective temperature for 24 h before they were washed twice with 10 ml ethanol and re-dispersed in 10 ml fresh ethanol. As-synthesized NPs were stabilized with palmityl-nitroDOPA within 2 h after synthesis.

Iron oxide NP stabilization: 6 mg palmityl-nitroDOPA dissolved in DMF at a concentration of 100 mg/ml was added to 0.5 ml ethanol before 0.5 ml of the as-synthesized iron oxide NPs were added. Palmityl-nitroDOPA was adsorbed for 24 h at 50 °C under constant mechanical stirring (Thermomixer comfort, Vaudaux-Eppendorf, Switzerland). To remove excessive dispersants, NPs were washed three times by centrifuging them for 30 min at 14000 rpm (MiniSpin, Vaudaux Eppendorf, Switzerland) before the supernatant was exchanged with 1 ml fresh ethanol. NPs were centrifuged a fourth time where ethanol was exchanged with 1 ml Millipore water before these NPs were freeze-dried.

Liposome preparation: 886 μg PEG(2)-PE dissolved in CHC1₃ at a concentration of 25 mg/ml was added to 5 mg DSPC, SOPC or POPC, respectively, dispersed in 0.5 ml CHCI₃ before optionally 1.5 mg palmityl-nitroDOPA stabilized iron oxide NPs, dispersed in 150 μg CHC1₃ was added. Chloroform was removed during 1 h under a constant N2 stream before the lipid film optionally functionalized with iron oxide NPs was swollen in Millipore water (i?=18.2 Ω, TAC<6 ppb), PBS or PBS containing calcein. Liposomes loaded with PEG(1.5)-nitroDOPA stabilized iron oxide NPs were swollen in PBS containing calcein where the iron oxide concentration was kept constant at 0.5 mg/ml for both core diameters. To prevent filter clogging by oversaturated calcein solutions, calcein dispersed in PBS at 3 mg/ml was filtered using syringe filters (Sartorius, Germany) before it was added to the dried lipid film. The lipid film was swollen for 1 h at 65 °C before it was sequentially extruded 10 times through 200 nm and 31 times through 100 nm polycarbonate filters using a hand extruder (Avestin, Mannheim, Germany). Extrusion was performed at 65 °C. Liposomes where calcein, FITC or PEG(1.5 kDa)-nitroDOPA stabilized iron oxide NPs were embedded in the lumen were run through a Sephadex column (Sephadex G75) to remove not encapsulated fluorophores and NPs respectively. Calcein loaded liposomes were used within 24 h after column separation. No change in size and morphology of extruded liposomes containing 5 mol% PEG(2)-PE could be determined with DLS and cryo-TEM if liposomes were stored at 4 °C for more than 4 weeks.

Liposomes prepared for encapsulation in polyelectrolyte multilayers were formed from lipids dissolved in chloroform and stored at -20°C. Membrane-labeled vesicles were consisting of 98wt% DOPS and 2wt% fluorescently labeled NBD-POPC. 5 mg of the lipids were dried under a constant flow of nitrogen for 30 minutes and then rehydrated in 150 mM NaCl, no buffer used (pH 5-6). Cargo-labeled vesicles consisting of 100wt% DOPS were used and 5 mg of the lipids were dried and rehydrated in 50 mM 5,6-Carboxyfluorescein. Then, the suspensions were each extruded through a double-stacked polystyrene membrane with a pore size of 200 nm. The extruded vesicles were diluted to a concentration of 0.5 mg/ml and stored in the fridge for maximum 14 days before use. Non-labeled vesicles consisting of 100% DOPS lipids, produced in exactly the same manner as explained above, were characterized at 20° with respect to their hydrodynamic radius by dynamic light scattering (170 ± 33 nm) and ζ-potential (-45 mV).

Fluorescent anionic lipid vesicles were prepared for the experiments demonstrating nanopore- spanning lipid membranes according to the Barenholz method (Barenholz, Y., et al., Biochemistry, 1977, 16: 2806-2810) with the following lipid constituents: 69.8 wt% POPC, 30 wt% POPS and 0.2 wt% DOPE-RhoB. Monodisperse liposomes were prepared for these experiments by extruding through double-stacked 200 nm polycarbonate filters (Avestin, Germany) according to the protocols of Macdonald et al. (MacDonald, R., et al, Biochim Biophys Acta, 1991, 1061: 297-303), leading to a diameter of 132.3 ± 1.1 nm determined by dynamic light scattering. Liposomes were suspended in 5 mg/ml stock solutions in TBS and used within 2 weeks from preparation. Solutions were diluted in TBSCa immediately prior to use.

DNA tethering: Liposomes dispersed in PBS at a concentration of 5 mg/ml were tethered with DNA according to the protocol of Hook et al.(Tabaei, S.R. et al, Journal of Structural Biology, 2009, 168: 200-206). Briefly, liposomes were incubated with 100 nM of DNA (Lazerlab, Sweden) for 1 h at 25 °C.

Polymersome preparation: Similar protocols to those for the preparation of liposomes were used for assembling polymersomes. Palmityl-nitroDOPA stabilized iron oxide NPs were dispersed in CHC1₃ and poly(2-dimethyl amino ethyl) methacrylate (PMCL- PDMAEMA) block-co-polymers were dissolved in CHC1₃. After adding iron oxide NPs dispersion to the PMCL-PDMEMA CHC1₃ solution in a weight ratio of block-co-polymers :SPIONs up to 3:1, CHC1₃ is dried under steady N₂ flow for 1 h. The block-co-polymer/iron oxide NP mixture is swollen between 2 and 10 days in an aqueous buffer or water before the hydrated dispersion was sequentially extruded through two stacked polycarbonate filters using a hand extruder. Polymersomes were extruded 20 times through 400 nm pore size filters followed by 31 time extrusion through 200 nm pore size filters.

Spraying of polyelectrolyte multilayers: The spraying process as described in (Izquierdo, A. et al., Langmuir, 2005, 21 7558-7567) has been automated in our laboratory by a home-built spraying robot (SI 2). The custom made program first wets the substrate with 150 mM NaCl solution for 5 seconds. After a pause of 5 seconds the PLL solution (0.5 mg/ml) was sprayed for 5 seconds. After a pause of 15 seconds the substrate was rinsed with buffer or 150 mM NaCl solution for 5 seconds. After another 5 seconds break, the PSS solution (0.5 mg/ml) was sprayed for 5 seconds, followed by 15 seconds pause and rinsing of the substrate with buffer or 150 mM NaCl solution for 5 seconds. This procedure was repeated 9.5 times in order to obtain a (PLL/PSS)9-PLL multilayer (Guillaume-Gentil, O. et al, Soft Matter, 2010, 6: 4246- 4254). The PEM coated samples were stored at 4°C in 150 mM NaCl solution for maximal 4 weeks until use.

Methods

TGA of palmityl-nitroDOPA stabilized NPs: 3-5 mg stabilized NPs were analyzed on a NETZSCH STA 449 C Jupiter TGA. Samples were heated from 35 °C to 600 °C at 10 °C/min using a flow rate of 47.4 seem Ar and 12.6 seem 0₂. TGA of liposomes: TGA analysis was performed on a TA Q500 instrument (TA instruments). Millipore based liposome solutions were analyzed. Liposomes were extruded at a lipid concentration of 5 mg ml where the liposome concentration was increased by partially evaporating water under a constant N₂ flow were analyzed. 50 μΐ liposome dispersions were analyzed at a time. Water was evaporated in situ at 50 °C before the sample was heated at 20 °C/min to 600 °C in a N₂ atmosphere. The measurement was repeated for statistics on 2-4 independent identical samples.

DSC of liposomes: DSC was measured on a Perkin Elmer instrument (Perkin Elmer DSC7). The same Millipore based liposome solutions as analyzed with TGA were used to measure DSC. Samples were analyzed from 25 °C to 80 °C before they were cooled to 25 °C using a heating and cooling rate of 10 °C/min. This temperature cycle was repeated twice. 40 μΐ of liposome solution was analyzed at a time in pressure tight steel crucibles. The measurement was repeated for statistics on 2-4 independent identical samples.

TEM: cryo-TEM was performed on Millipore based liposome dispersion where the liposome concentration was 1 mg/ml. These solutions were adsorbed on Quantifoil holey carbon films R3.5/1 (Electron Microscopy Sciences, PA, USA) and plunged into liquid ethane using a Vitrorobot (FEI) and analyzed on a Philips CM 12 microscope operated at 100 kV at liquid N₂ temperatures. Conventional TEM was done on liposomes that were fixed with 1 wt% trehalose and air dried on a Quantifoil holey carbon films R3.5/1 (Electron Microscopy Sciences, PA, USA) using a Philips CM12 microscope operated at 100 kV.

STEM: Millipore water based liposome dispersions were freeze-dried on a TEM grid that was supported by an 8 nm thick, glow discharged carbon film. The HAADF STEM images as well as the EDX-measurements were performed on a HD-2700 Cs-corrected dedicated STEM (Hitachi), operated at 80 kV and equipped with an additional secondary electron (SE) detector.

DLS and ζ-potential: DLS experiments were performed on a Zetasizer Nano ZS (Malvern, UK) in the 173° backscattering mode for liposomes and polymersomes for experiments related to loading of magnetic nanoparticles and adsorption onto nanoporous substrates, and at 20° for liposomes used in the polyelectrolyte multilayer experiments. Liposomes or polymersomes were diluted in PBS or Millipore water respectively, and analyzed at a concentration of 50 μg/ml. Because intensity weighted diameters scale with r⁶ and thus agglomerates predominate, volume weighted diameters that scale with r³ are shown.^(Mie' ^G"

Annalen der Physik, 1908, 25: 377-445) ^ _{analyzed u§ing mg multiple nam)w mod}es evaluation incorporated into the Malvern software, ζ-potential measurements were performed on the same machine using standard gold plated electrode cuvettes in the buffer used for liposome preparation.

Small angle neutron scattering (SANS): SANS measurements were performed at SANS-I at the Paul-Scherrer-Institute (PSI, Villigen, Switzerland). For SANS experiments, liposomes were extruded in D₂0. Samples were analyzed at a liposome concentration of 5 mg/ml, corresponding to 0.6 vol% liposomes assuming liposomes have a diameter of 76 nm at 25 °C and 60 °C. For these measurements, liposome dispersions were filled between two quartz plates that were separated and sealed by an O-ring. The distance between these glass plates was 3 mm. Data were acquired on a two dimensional ³He detector at distances of 2 m, 6 m and 15 m and a neutron wavelength of 0.5 nm. Additionally, the neutron wavelength was increased to 1.3 nm at 15 m detector distance covering a -range of 0.02 nm^"1 < q < 3 nm^"1. After correcting the data for background scattering, empty cell scattering and detector efficiency, the data were radially averaged.

SANS Data analysis: Data were analyzed by with the SasFit software ( ohlbrecher, Paul- Scherrer Institute, Switzerland). A form factor for vesicles was combined with one that describes scattering of a core-shell NP. These form factors were adapted from Pedersen et /.(Pedersen, J.S., Journal of Applied Crystallography, 2000, 33: 637-640) as described in (Amstad, E. et al., Nano Letters, 2011, 11: 1664-1670). To account for agglomeration of DSPC liposomes at 25 °C, a constantly increasing background with a slope of -3.5 was assumed. For these fits, a phosphocholine head group size of 0.5 nm(Nagle, J.F. et al., Biochimica Et Biophysica Acta-Reviews on Biomembranes, 2000, 1469: 159-195) was assumed.

Alternating magnetic field (AMF) measurements: Samples comprising 1.5 ml self-quenched calcein loaded liposomes dispersed at a liposome concentration of ~ 0.5 mg/ml in PBS were subjected to an AMF. The alternating magnetic field was induced by running 450 A at a frequency of 230 kHz through a 3.5 cm diameter coil with 6 loops where the sample was localized within only 2 loops (Ambrell, Netherlands). Unless stated otherwise, samples were exposed to the AMF for 6 times 5 min and equilibrated between every cycle for 1 min. Fluorescence was quantified using a Fluorispectrometer at an excitation and emission wavelength of 488 nm and 520 nm respectively. The fluorescence was normalized to the fluorescence of samples that were not exposed to an AMF and to the volume of the vesicle lumen calculated from DLS results. The measurements were performed on 3-6 independent identical batches for statistics. Control samples were externally heated at a heating rate of 1 °C/min.

Magnetic resonance imaging (MRI): The relaxivities r₂ ^*, and thus the performance of Fe₃0₄ NPs as MR contrast agents, were tested in vitro with MRI. PEG-nitroDOPA stabilized Fe₃0₄ NPs, dispersed in Millipore water were embedded in a hydrogel to mimic their binding to tissue in vitro. Therefore, NPs were dispersed in Millipore water at a final Fe₃0₄ concentration of 20-100 μg/ml. A hydrogel was formed by radical initiated polymerization of methacrylates.(Murthy, P.S.K. et al, Reactive & Functional Polymers, 2006, 66: 1482-1493) From a Millipore water based 100 mg/ml N, N'-methylene-bis-acrylamide (MBA) and ammonium persulfate (APS) stock solutions, 10 mg MBA and 5 mg APS were added to eppendorf containing Millipore water based Fe₃0₄ NP solutions. The total volume of Millipore water was 0.5 ml. The radical reaction was initiated by adding 7 μΐ N, N, N¹, N¹- tetramethylethylenediamine (TMEDA). Eppendorfs containing NP loaded hydrogels were inserted into a 50 ml Falkon tube which was filled with Millipore water to avoid discontinuities of the permeability of the hydrogels at the interface of the eppendorf tubes which could cause artifacts in MRI and thus influence the measured relaxivities. As a comparison, PEG-nitroDOPA stabilized in the oil bath synthesized NPs dispersed in PBS were analyzed.

MRI on in hydrogels embedded NPs was measured on a 4.7 T Bruker instrument using a volume resonator coil with a diameter of 31 mm. A field of view of 3.5 cm x 4.0 cm was imaged. The repetition time was 5000 ms where spins were flipped 90 ° with an echo spacing of 6 ms. 16 echos were measured per repetition, r measurements as a function of the PEG molecular weight were performed on 3 independent identical samples at room temperature.

Quartz-crystal microbalance with dissipation (QCM-D): QCM-D studies were carried out on a Q-Sense E4 equipment (Q-Sense AB). 50 μg ml liposomes dispersed in PBS were added to a Si0₂-coated quartz crystals (Q-Sense AB). The formation of supported lipid bilayers (SLBs) was followed in real time by monitoring changes in frequency (Af) and dissipation {AD).

UV/VIS experiments: UV/VIS experiments on liposomes loaded with 3 mg/ml calcein or FITC respectively were performed on a Cary El spectrometer. The absorbance of these liposomes was measured between 400 and 600 m at a step size of 2 ran. Samples were heated with a thermostate within the spectrometer at a heating rate of 1 °C/min. Confocal laser scanning microscopy: Confocal microscopy experiments onnanoporous substrates were conducted in a custom made open microscopy cell. Before being placed in the microscopy cell, nanopore substrates were thoroughly cleaned by ultrasonication in ethanol and water followed by 30 minutes of UV-ozone cleaning; substrates were then used within 45 minutes of removal from the UV chamber. Vesicle solutions were diluted to 0.5 mg ml in TBSCa and allowed to adsorb directly on the nanopore arrays for at least 15 min, before thoroughly rinsing with TBSCa. The dye encapsulation experiment shown in Figure 16 was conducted using 5(6)-carboxyfluorescein (CF). CF was used as it has a low permeability through planar lipid bilayers. 1 mg/ml CF was added to TBSCa and filtered to produce a saturated solution of CF-TBSCa. Vesicle solutions were diluted directly into CF-TBSCa as above prior to use. CF experiments were typically conducted within 20 minutes of initial adsorption. All confocal microscopy experiments were conducted on a Leica SP5 (Leica Microsystems, Germany) inverted setup with an oil immersion 100X N/A 1.4 objective using the 488 nm and 514 nm lines of the Ar laser and the 561 nm He-Ne laser. Two images were recorded simultaneously: the reflection of the incident laser and the corresponding fluorescence signal of the labelled lipid bilayers. The large difference in refractive index between the silicon nitride (n « 2.16) and underlying glass substrate (n « 1.52) and the refractive index of the buffer (n « 1.33) caused the incident laser light to scatter at the positions of the nanopores. Although all the recorded images are subject to the diffraction limit of the incident laser line, each nanopore could be accurately located from their respective scattered dark spot since their spacing was larger than the diffraction limit.

Electrochemical Confocal Laser Scanning Microscope (EC-CLSM): EC-CLSM observations of liposomes embedded in polyeletrolyte multilayers were carried out on a Zeiss LSM 510 microscope. NBD, fluorescence was detected after excitation at 488 nm with 4% laser power, cutoff dichroic mirror 488 nm, and emission band pass filter 505-530 nm (green). Bleaching of fluorescence was accomplished by exposure to maximum laser intensity in the designated area before renewed imaging at 4% excitation intensity.

The uPEM covered samples were mounted after quick rinsing with ultrapure water (Resistivity = 18.2 ΜΩ/cm) and drying with nitrogen. The used flowcell was the same that was also used for the AFM experiments. The teflon cell was provided with a silver wire which was anodized in chloride to be an Ag/AgCl reference electrode and with a platinum wire as counter electrode. A copper spring established the electrical contact with the ITO surface of the substrate used as working electrode. The vesicles were then adsorbed in situ on the pre-sprayed (PLL/PSS)9-PLL PEM for approximately 30 minutes. The substrate was extensively rinsed (7 ^χ 400 μΐ) after adsorption of the vesicles. The integrity of the membrane-labeled vesicles adsorbed on the PEM layer was assured by means of fluorescence recovery after photobleaching (FRAP).

Electrochemical Quartz Crystal Microbalance with Dissipation Monitoring (EC-QCMD): The QCM measurements on embedding of liposomes in polyelectrolyte multilayers were carried out with a QCM-D system from Q-Sense (Sweden), described in detail (Rodahl, M. et al., Review of Scientific Instruments, 1996, 67: 3238-3241). Briefly, QCM measures the changes in the resonance frequency (Δί) of a quartz crystal when material is adsorbed onto it. The quartz crystal was excited at its fundamental frequency (about 5 MHz) and at the third, fifth, and seventh overtones (n). A home-built, three-electrode, electrochemistry flow cell was used for the EC-QCM-D measurements (Grieshaber, D. et al., Langmuir, 2008, 24: 13668- 13676). The ITO coated surface of the quartz crystal sensor was the working electrode whereas the Ag/AgCl reference and the platinum counter electrodes were situated on the upper side of the flow cell.

Shortly before mounting into the EC-QCM flow cell, the uPEM-covered quartz crystal was rinsed with ultrapure water and dried with nitrogen followed by rehydration in 150 mM NaCl solution within a few minutes. This procedure assured that no salt crystals formed upon drying the samples. The vesicle solution (0.5 mg/ml) was then injected and incubated for 30 minutes. After that, the flow cell was rinsed with 150 mM NaCl solution. Two additional layer pairs of (PLL/PGA) were then injected as described in (Guillaume-Gentil, O. et al., Advanced Materials, 2008, 20: 560-+).

Example 1 : Synthesis of Fe₃Q NPs (SPIONs) in an oil bath

A solution of 1 mmol Fe(ac)₂ (173 mg) precursors dissolved in 5 ml benzylalcohol was heated in an oil bath to temperatures in the range of 140-180 °C for 1-3 days. The obtained particles with core diameters between 2 and 15 nm were then washed twice with two times 10 ml EtOH (to remove impurities and excessive dispersants) through centrifugation. Washed nanoparticles were re-dispersed in 10 ml fresh ethanol.

Example 2: Stabilization of Fe₃Q₄ NPs with PEG-nitroDOPA dispersants (see also Figure 2, (Amstad, E., et al., Nano Letters, 2009, 9: 4042-4048)) PEG-nitroDOPA was dispersed in dimethylformamide (DMF) at a concentration of 100 mg/ml. 60 μΐ of this stock-solution was added to 0.5 ml ethanol before 0.5 ml of the ethanol based iron oxide nanoparticle dispersion obtained in Example 1 was added. Nanoparticles were stabilized at 50 °C for 24 h under constant mechanical stirring at 500 rpm (Thermomixer comfort, Vaudaux-Eppendorf, Switzerland). The so obtained stabilized NPs were dialyzed against, Millipore water using dialysis tubes with a cut-off of 14-16 kDa (Spectra/Por dialysis membrane, spectrum labs, Netherlands) for 1 d. Alternatively, the obtained stabilized NPs were centrifuged for 30 min at 13500 rpm (MiniSpin, Vaudaux Eppendorf, Switzerland), to remove agglomerates. The supernatant was freeze-dried (freeze dryer ALPHA 1-2 / LDplus, Kuhner LabEquip, Switzerland) and run through a Sephadex column (Sephadex G75 superfine) using Millipore water as an eluent to remove excessive dispersants. The purified stabilized nanoparticles (or PEG-nitroDOPA SPIONs) were freeze-dried again and stored as a powder or re-dispersed in aqueous media at concentrations ranging from 10 μg/ml to 10 mg/ml. The purified stabilized nanoparticles had an core size diameter ranging from 2 nm to 13 nm, were of high crystallinity and consisted of a single domain, leading to the expected superparamagnetic behavior and high magnetization values for pure magnetite NPs at room temperature Stabilized iron oxide NPs are schematically illustrated in Figure 2(a). Figure 2 (b) and c) show TEM and size distribution of stabilized NPs of 3.5 and 6.6 nm in radius respectively. Figure 2(d) illustrates nitroDOPA-PEG used for stabilizing NPs. Figure 2(e) shows volume weighted diameters of iron oxide NPs stabilized with nitroDOPA-PEG(5) (-■- ), nitrodopamine-PEG(5) (-□-), DOPA-PEG(5) (-A-), dopamine-PEG(5) (-Δ-), mimosine- PEG(5) (-·-), hydroxypyrrolidone-PEG(5) (-0-) and hydroxydopamine-PEG(5) (-V-). Figure 2(f) shows TEM of nitroDOPA-palmityl stabilized, Fe₃0₄ cores synthesized in the oil bath showing higher yield, and more uniform morphology and size than NPs synthesized in the microwave.

As expected, PEG(5 kDa)-nitroDOPA stabilized, 5 nm core diameter SPIONs increased the relaxivity r₂ ^' as was shown with MRI measurements (Figure 13).

Example 3: Stabilization of Fe^Q NPs by nitroDOPA-alkyl dispersants

NitroDOPA-palmityl was dissolved in DMF at a concentration of 100 mg/ml. 60 μΐ from this stock solution (corresponding to 6 mg palmityl-nitroDOPA) is added to 0.5 ml ethanol before 0.5 ml of the iron oxide nanoparticle dispersion obtained in example 1 is added.. The obtained mixture was incubated at 50 °C for 24 h under mechanical stirring. To remove excessive dispersants, the nitroDOPA-palmityl SPIONs were centrifuged three times at 13500 rpm for 30 min (MiniSpin, Vaudaux Eppendorf, Switzerland). After each time, the supernatant (about

1 ml) was exchanged with 1 ml pure EtOH. The washed nitroDOPA-palmityl SPIONs were again centrifuged at 13500 rpm for 30 min, the supernatant was decanted and 1 ml of Millipore water was added before the nitroDOPA-palmityl SPIONs were freeze dried for storage. Prior to any intended use the freeze-dried particles were re-dispersed in a suitable solvent, e.g., chloroform. TGA analysis results showed that this procedure resulted in a palmityl-nitroDOPA packing density of 1.5 molecule/nm², irrespective of the iron oxide core size as shown in Figure 2: for 2.5 nm (red circles) and 5 nm (black squares) core radius SPIONs.Example 4: Liposome assembly:

(a): Assembly of DSPC liposomes containing nitroDOPA-palmityl SPIONs

5 mg of l,2-distearoyl-sn-glycero-3-phosphocholine (DSPC) (Avanti polar lipids) dissolved in chloroform at a concentration of 50 mg/ml and 886 μg l,2-Dioleoyl-sn-Glycero-3- Phosphoethanolamine-N-[Methoxy(Polyethylene glycol)2000 Da] (ammonium salt) (0.25 mmol) (PEG(2)-PE) dissolved in chloroform at a concentration of 25 mg/ml were mixed before 3 mg of palmityl-nitroDOPA stabilized iron oxide nanoparticles described in Example

2 dissolved in 100 μΐ of chloroform was added. Chloroform was removed by constantly flowing N2 through the vial for 2-3 h at 25 °C resulting in a thin lipid film at the bottom of the glass vial. The lipid film was rehydrated with water, PBS or HEPES respectively at 60 °C to form the desired multi-lamellar liposomes. These liposomes were sequentially extruded at 60 °C first through two stacked 200 nm and then two 100 nm pore size polycarbonate filters. In extruded form, the desired liposomes could be stored at 4 °C for at least 4

(b) : Assembly of SOPC liposomes containing nitroDOPA-palmityl SPIONs:

l-stearoyl-2-oleoyl-sn-glycero-3-phosphochline (SOPC) (Avanti polar lipids) liposomes were assembled identical to the DSPC liposomes described in example 4a except that the lipid DSPC was exchanged by SOPC and they were rehydrated and extruded at 25 °C.

(c) : Assembly of POPC liposomes containing nitroDOPA-palmityl SPIONs:

l-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) (Avanti polar lipids) liposomes were assembled identical to SOPC liposomes described in example 4b. Only the lipid SOPC was replaced by POPC

(d): Assembly of MPPC liposomes containing nitroDOPA-palmityl SPIONs: 1 -myristoyl-2-palmitoyl-CT-glycero-3 -phosphocholine (MPPC) (Avanti polar lipids) liposomes were assembled identical to SOPC liposomes described in example 4b. Only the lipid SOPC was replaced by MPPC.

(e) : Assembly of DSPC liposomes containing FITC-nitroDOPA functionalized SPIONs.

Iron oxide nanoparticles synthesized as described in Example 1 were stabilized similar to the procedure described in Example 3. However, iron oxide nanoparticles synthesized as described in example 1 were stabilized according to the procedure described in example 3. However, instead of stabilizing them with 5 mg/ml palmityl-nitroDOPA, they were stabilized with 3.5 mg/ml FITC-nitroDOPA for 1 h before they were back-filled by adding 2.5mg/ml palmityl-nitroDOPA. Nanoparticles were purified as described in example 3 before they were freeze-dried. 3 mg of the resulting nanoparticles were re-dispersed in chloroform and added to the DSPC/PEG(2)-PE lipid mixture identical to what was described in Example 4a. Liposomes were prepared identical to the procedure described in Example 4a.

UV/VIS spectra of POPC vesicles where palmityl-nitroDOPA stabilized SPIONs (black) and SPIONs that were surface modified with a mixture of FITC-nitroDOPA and palmityl- nitroDOPA (red) and incorporated into the liposome bilayer are shown in Figure 22. The absorption around 490 nm of liposomes loaded with FITC-labeled SPIONs shows feasibility to label liposomes by incorporating labeled SPIONs in their membrane.

(f) : Assembly of DSPC liposomes containing palmityl-nitroDOPA stabilized iron oxide nanoparticles in their membranes as described in Example 4a, loaded with fiuorophores in their cavities.

Liposomes loaded with fiuorophores were prepared according to the procedure described in Example 4a to obtain a dried lipid film. The dried lipid film was re-hydrated in PBS that was saturated with calcein or FITC respectively (corresponding to about 3 mg fiuorophore per ml buffer). After extrusion of the liposomes using the procedure described in Example 4a, not- encapsulated fiuorophores were removed by running the liposome dispersion through a Sephadex coloumn (G75, Superfine) using PBS as an eluent.

(g) : Assembly of liposomes using sonication. DSPC, PEG(2)-PE and palmityl-nitroDOPA stabilized iron oxide nanoparticles were mixed and dried and re-hydrated as described in Example 4a. Liposomes were then sonicated using a Branson sonicator for 30 min.

(h) : Assembly of liposomes containing no iron oxide nanoparticles: Liposomes that did not contain any nanoparticles were assembled according to the procedure described in Example 4a. However, no iron oxide nanoparticles were added to the DSPC/PEG(2)-PE solution prior to drying.

(i): Assembly of liposomes loaded with FITC-dextran. Liposomes loaded with FITC-dextran were prepared according to the protocol described in Example 4e. Instead of dissolving calcein or FITC in the buffer as described in Example 4e, FITC-dextran was dissolved in HEPES at a concentration of 1 mg/ml. After extrusion of liposomes, unencapsulated FITC- dextran was removed by runnding the liposome dispersion through a Sephadex column (G75, Superfine) according to the procedure described in example 4f.

(j): Assembly of liposomes containing PEG-nitroDOPA stabilized iron oxide nanoparticles in their lumen: Liposomes loaded with PEG-nitroDOPA stabilized iron oxide nanoparticles were assembled as described in Example 4f. Instead of fluorophores, PEG(1.5 kDa)-nitroDOPA stabilized iron oxide nanoparticles (preparaed as described in Example 2) were dispersed in PBS at a concentration of 10 mg ml. Unencapsulated iron oxide nanoparticles were removed by running the liposome solution through a sephadex column as described in Example 4f. (k): Assembly of liposomes containing oleic acid stabilized iron oxide nanoparticles. Iron oxide nanoparticles synthesized as described in Example 1 were coated with oleic acid by adding 100 μΐ oleic acid to 1 ml of the iron oxide nanoparticle solution described in example 1. Oleic acid was adsorbed on the iron oxide nanoparticles for 24 h at 50 °C under constant mechanical stirring alike the stabilization of iron oxide nanoparticles with PEG-nitroDOPA described in example 2. Non-adsorbed oleic acid was removed by centrifuging iron oxide nanoparticles for 30 min at 13500 rpm (MiniSpin, Vaudaux Eppendorf, Switzerland). After decanting the supernatant, 1 ml of Millipore water was added before the nanoparticles were freeze-dried as described in Example 2. 1.5 mg of these nanoparticles were re-dispersed in 100 μΐ chloroform and added to the DSPC/PEG(2)-PE mixture as was described for palmityl- nitroDOPA stabilized iron oxide nanoparticles in example 4a. Liposomes were then extruded as described in example 4a.

Example 5: Characterization of liposomes prepared as described in example 4

The size of liposomes obtained in Example 4 was measured with DLS. Liposomes were dispersed in PBS at a concentration of 50 μg/ml.

Liposomes that were formed through sonication primarily assembled into micelles while liposomes that were extruded assembled into vesicular structures as was exemplified on DSPC liposomes shown in Figure 4. DSPC liposomes containing 5 mol% PEG(2)-PE in their membranes and hosting small, 2.5 nm core radius (red squares) and large, 5 nm core radius (blue triangles) palmityl-nitroDOPA stabilized iron oxide NPs in their membranes that were formed through sonication (empty symbols) primarily assembled into micelles while liposomes that were extruded (filled symbols) assembled into vesicular structures.

However, the liposome composition did significantly affect the liposome size (Figure 5). However, liposomes containing iron oxide nanoparticles in their membranes were considerably smaller compared to control liposomes that did not contain any nanoparticles in their membranes. This is shown in Figure 5 for l-palmitoyl-2-oleoyl-sn-glycero-3- phosphocholine (POPC) (black circles), l-stearoyl-2-oleoyl-sn-glycero-3-phosphocholine (SOPC) (blue triangles) and l,2-Distearoyl-sn-glycero-3-phosphocholine (DSPC) (red squares) liposomes; Liposomes containing 2.5 nm core radius palmityl-nitroDOPA stabilized iron oxide NPs in their membranes (filled symbols) were considerably larger compared to control liposomes that did not contain any NPs in their membranes (empty symbols). All liposomes contained 5 mol% PEG(2)-PE in their membranes. DLS measurements were performed at 25 °C. Furthermore, the chemical structure of DSPC (Figure 5b), SOPC (Figure 5c) and POPC (Figure 5d) is shown.

Furthermore, individually dispersed palmityl-nitroDOPA stabilized iron oxide nanoparticles could be seen on TEM images of liposomes that were fixed with 1 wt% trehalose and dried in air (Figure 6). However, if the concentration of Trehalose was lowered to 0.5 wt%, some of the liposomes ruptured during drying resulting in the lipid bilayer films seen in Figures 5b and 5 c. No agglomerated NPs could be seen on these TEM micrographs indicating that individually palmityl-nitroDOPA stabilized NPs do not agglomerate if embedded in the hydrophobic part of liposome membranes. Cryo-TEM images revealed no significant change in the overall shape of DSPC liposomes upon loading them with palmityl-nitroDOPA stabilized iron oxide nanoparticles as can be seen by comparing Figure 7, a cryo-TEM of DSPC liposomes containing 5 mol% PEG(2)-PE without SPIONs in their bilayers to images of DSPC liposomes containing 5 mol% PEG(2)-PE containing palmityl-nitroDOPA stabilized 2.5 nm core radius NPs (Figure 7c) and palmityl-nitroDOPA stabilized 5 nm core radius NPs (Figure 7d). Furthermore, cryo-TEM revealed that oleic acid coated SPIONs agglomerated and did not efficiently embed in the DSPC membrane containing 5 mol% PEG(2)-PE (Figure 7b).

Figure 8a shows photographs of DSPC liposome dispersions without NPs, and liposomes containing small, 2.5 nm core radius iron oxide NPs stabilized with palmityl-nitroDOPA and oleic acid respectively. All liposomes contained 5 mol% PEG(2)-PE in their membranes. Furthermore STEM micrographs of DSPC liposomes containing 5 mol% PEG(2)-PE in their membranes and functionalized with small, 2.5 nm core radius palmityl-nitroDOPA stabilized iron oxide NPs in their membranes detected with a high angle annular dark field (HAADF) (Figure 8b) and secondary electron (SE) (Figure 8c) detector revealed that electron dense particles that were assigned to iron oxide nanoparticles were covered by an organic layer. The latter was assigned to the phospholipid bilayer. This interpretation was supported by electron diffraction X-ray (EDX) measurements performed on the location shown in Figure 8b and c and shown in Figure 8d.

To characterize membrane distortions caused by the incorporation of palmityl-nitroDOPA stabilized NPs into the liposome membrane, SANS measurements were performed on DSPC liposomes (Figure 9) at a) 25 °C and b) 60 °C. While DSPC liposomes without NPs (black) showed the for liposome expected constant increase in the scattering curve towards lower q- values in the range of 0.5 nm^"1 > q > 0.1 nm^"1, SANS curves measured on liposomes hosting 2.5 nm core radius (red) and 5 nm core radius (blue) palmityl-nitroDOPA stabilized iron oxide NPs in their membranes were undulated in this region. These undulations can be assigned to the distortions of the liposome membrane caused by iron oxide nanoparticles.

However, DSPC liposomes that had no PEG(2)-PE in their membranes agglomerated at temperatures below their transition temperature (55 °C) as was seen in the constant increase of the SANS scattering curve at low q- values. The addition of 5 mol% PEG(2)-PE prevented liposome agglomeration as can be seen in Figure 10. Figure 10 presents SANS measurements performed on DSPC liposomes containing 5 mol% PEG(2)-PEat a) 25 °C and b) 60 °C. All liposomes containing PEG(2)-PE in their membranes, regardless if they had no (black), small, 2.5 nm core radius (red) and large, 5 nm core radius (blue) palmityl-nitroDOPA stabilized iron oxide NPs were embedded in the liposome membrane were stable at room temperature.

Furthermore, SANS measurements performed at 25 °C on POPC liposomes without NPs (black), POPC liposomes containing palmityl-nitroDOPA stabilized small, 2.5 nm core radius (red) and large, 5 nm core radius (blue) NPs in their membranes revealed that these liposomes are stable at room temperature in the absence of PEG(2)-PE (Figure 11). This showed that liposomes start to agglomerate if stored at temperatures below their transition temperature, irrespective if they have SPIONs in their membrane. Despite the massive membrane distortions caused by SPIONS that are incorporated in the liposome bilayer, DSC measurements of Figure 12 a) DSPC liposomes and Figure 12 b) DSPC liposomes containing 5 mol% PEG(2)-PE in their membranes revealed no significant change in the liposome phase transition temperature upon embedding iron oxide nanoparticles in the membrane as a comparison of DSC graphs was measured on liposomes that did not contain any iron oxide NPs in their membranes (black), liposomes that hosted palmityl- nitroDOPA stabilized small, 2.5 nm (red) and large, 5 nm (blue) core radius iron oxide NPs in their membranes reveals (Figure 12).

Example 6: Synthesis of polymersomes

(a): Synthesis of PMCL-PDMEMA polymersomes

3 mg of palmityl-nitroDOPA stabilized iron oxide NPs obtained in Example 3 were dispersed in 100 μΐ in CHC1₃ andadded to a solution of 5 mg poly(2-dimethyl amino ethyl) methacrylate (PMCL- PDMAEMA) block-co-polymers that were dissolved in 1 ml CHC1₃ . After the addition was completed, CHC1₃ was dried off under steady N₂ flow for 1 h. The so obtained block-co-polymer/iron oxide NP mixture was swollen for 10 days in water or PBS respectively to obtain a hydrated dispersion. Subsequently, the hydrated dispersion was extruded 20 times through two stacked 400 nm pore size polycarbonate filters followed by 31 times extrusion through two stacked 200 nm pore size polycarbonate filters to obtain the desired polymersomes (Figure 20).

Cryo-TEM micrograph of the resulting PMCL-PDMAEMA polymersomes containing iron oxide NPs in their membranes are shown in Figure 20a. Furthermore, these polymersomes were characterized with DLS. The volume weighted diameters of PMCL-PDMAEMA polymersomes containing iron oxide NPs with core diameters <4 nm in their membranes (Figure 20b) and PMCL-PDMAEMA block-co-polymers which were mixed with iron oxide NPs with a core diameter >10 nm prior to extrusion (Figure 20c) are shown. Significant changes in the polymersome shape of PMCL-PDMAEMA polymersomes containing palmityl-nitroDOPA stabilized SPIONS with core diameters < 4 nm indicate that polymersomes can be actuated if exposed to alternating magnetic fields (100 kA/m) for 10 min.

Furthermore, these polymersomes were characterized with SANS. SANS was performed on PMOXA-b-PDMS-b-PMOXA (Figure 21a) and PMCL-PDMAEMA (Figure 21b) polymersomes. Measurements were performed on polymersomes that did not contain any NPs (black) and polymersomes containing palmityl-nitroDOPA stabilized 5 nm core radius NPs in their membrane (red). Additionally, SANS was measured on PMCL-PDMAEMA polymersomes containing 5 nm core radius palmityl-nitroDOPA stabilized iron oxide NPs at 25 °C (blue) and at 60 °C (red) respectively (Figure 21c).

(b): Assembly of PMCL-PDMAEMA liposomes loaded with FITC-dextran.

PMCL-PDMAEMA liposomes were assembled as described in Example 6a. Instead of hydrating them in water, they were hydrated in water containing 1 mg/ml FITC-dextran. After polymersomes were extruded as described in Example 6a, unencapsulated FITC-dextran was removed by running the samples through a Sephadex column (G75, Spuerfme) using Millipore water as an eluent.

These polymersomes and MPPC liposomes assembled as described in Example 4i using MPPC as a lipid were characterized with UV/VIS spectroscopy on MPPC liposomes (black) and PCML-PEO polymersomes (red) loaded with FITC-dextran (Figure 23).

Example 7: Membrane actuation through of liposome bilayers containing iron oxide nanoparticles in their membranes.

Calcein is a self-quenching dye. Therefore, fluorescence increases upon release of calcein initially located in the lumen of vesicles and/or in the vesicle bilayer at a concentration where self-quenching occurs, when vesicles become leaky.

Figure 17 illustrates control experiments of DSPC liposomes which have SPIONs in their membrane and were loaded with a) carboxy-fluorescein and b) calcein. These liposomes were externally heated using a thermostate and their fluorescence was monitored. They started to leak at temperatures ~54 °C, close to their phase transition temperature which can be seen in an increase in fluorescence for carboxy-fluorescein loaded liposomes and in a shift of the absorption maxima for calcein loaded liposomes. This supports DSC measurements (Figure 12) where no significant change in the transition temperature of liposomes could be measured upon adding palmityl-nitroDOPA stabilized SPIONs in DSPC liposome membranes.

However, release of calcein embedded in the cavity of DSPC liposomes containing 5 mol% PEG(2)-PE that were loaded with palmityl-nitroDOPA stabilized SPIONs in their membranes could be triggered with an alternating magnetic field (AMF). This is shown in Figures 18 and 19. 1.5 ml solution of DSPC liposomes that were loaded with the self-quenched calcein prepared as described in Example 4f dispersed at a liposome concentration of ~ 0.5 mg/ml in PBS were AMF treated. An alternating magnetic field was induced by running 450 A at a frequency of 230 kHz through a 3.5 cm diameter coil with 6 loops where the sample was localized within only 2 loops. Unless stated otherwise, samples were exposed to the AMF for 6 times 5 min and equilibrated between every cycle for 1 min. Fluorescence was quantified using a Fluorispectrometer at an excitation and emission wavelength of 488 nm and 520 nm, respectively. The fluorescence was normalized to the fluorescence of samples which were not exposed to an AMF and to the volume of the vesicle lumen calculated from DLS results. For statistics the measurements were performed on 3-6 independent identical batches. Control samples were externally heated at a heating rate of 1 °C/min.

Release from DSPC liposomes containing 5 mol% PEG(2)-PE loaded with self-quenched calcein was measured by monitoring the fluorescence (Figure 18). 1.5 ml of a PBS based 0.5 mg/ml liposome solution was AMF treatment for 6 x 5 min followed by 1 min equilibration between every AMF exposure (Figure 18a). No calcein release could be seen for unmodified DSPC liposomes containing 5 mol% PEG(2)-PE (red filled circles) that were prepared as described in Example 4h. Liposomes without NPs in their membranes also did not leak when subjected to AMF sequence 1 (Figure 18a). The low leakiness of the liposomes shown here can be explained by the high stability against agglomeration of liposomes sterically shielded with PEG(2)-PE compared to DSPC liposomes that do not contain PEGylated lipids in their membrane if stored at T < T_m. However, liposomes hosting small 2.5 nm core radius (red filled squares) and large 5 nm core radius (blue filled triangles) palmityl-nitroDOPA stabilized iron oxide NPs in their membranes, prepared as described in example 4a, efficiently released their cargo. The release from liposomes functionalized with palmityl- nitroDOPA stabilized iron oxide NPs was 180% more efficient compared to that of liposomes prepared with oleic acid coated NPs (magenta filled stars) prepared as described in Example 4k or liposomes that were loaded with hydrophilic PEG(1.5)-nitroDOPA stabilized small (red empty squares) and large (blue empty triangles) iron oxide NPs Example 4j. Figure 18b shows photograph of dispersions containing DSPC/PEG(2)-PE liposomes that hosted palmityl-nitroDOPA stabilized small iron oxide NPs in their membranes before and after they were treated 6 x 5 min with an AMF. The system was equilibrated for 1 min in between the AMF pulses. In Figure 18c, DLS measurements of DSPC/PEG(2)-PE liposomes (black filled circles) and DSPC/PEG(2)-PE liposomes functionalized with palmityl-nitroDOPA stabilized small NPs (red square) before (filled) and after (empty) AMF exposure are shown. They reveal that the liposome structure is retained upon AMF treatment indicating that cargo is released by the enhanced permeability of liposomes close to T_m. Importantly, the possibility to release cargo with pulsed AMF sequences without destroying the vesicles paves the way to not only trigger release but also to control the dose of released cargo and to slowly release cargo over prolonged times. It additionally allows preventing bursts that could lead to a temporary and local overdose of cargo when this is not desired.

Figure 18d reveals that DSPC/PEG(2)-PE liposomes functionalized with small palmityl- nitroDOPA stabilized NPs only started to release significant amounts of calcein at T > 50 °C if externally heated (red filled squares). As a comparison, the normalized fluorescence of liposomes that were exposed to an AMF as a function of T is shown for unmodified liposomes (black filled circles) and liposomes hosting small 2.5 nm core radius palmityl- nitroDOPA stabilized NPs in their membranes red empty squares).

The influence of the AMF sequence on the release efficiency is shown in Figure 19. The calcein release of DSPC/PEG(2)-PE liposomes functionalized with small 2.5 nm core radius iron oxide NPs stabilized with palmityl-nitroDOPA was tested for different sequences of the AMF (Figure 19a). Release was less efficient if the system was equilibrated for 5 min (red crossed squares) in between each 5 min long AMF cycle (AMF sequence 2) compared to the release of liposomes equilibrated only for 1 min (AMF sequence 1) (red filled squares) but still significantly above the zero release of unmodified liposomes (black filled circles) treated with the AMF pulse sequence 1. Furthermore, release of NP modified liposomes was insignificant if these liposomes were subjected to 10 cycles of 1 min AMF pulses followed by 1 min equilibration time (AMF sequence 3) (red empty squares). Figure 19b indicates the bulk temperatures of liposome dispersions subjected to the respective AMF sequence used in Figure 19a.

The greatly improved release efficiency of liposomes containing individually stabilized hydrophobic iron oxide NPs in their membranes can be assigned to a direct transfer of heat, generated by the iron oxide NPs upon subjection to an AMF, into the liposome membrane. In contrast, hydrophilic and with oleic acid coated agglomerated iron oxide NPs have to heat bulk water to temperatures approaching T,„ to release cargo. The need to strongly heat bulk water prevents that thermally sensitive chemicals, drugs and proteins can be incorporated into such magnetoliposomes without risking thermal degradation and loss of functionality of the cargo during release. It would also preclude the use in cell cultures and tissue where heating of the bulk liquid would damage or kill surrounding cells.

A prerequisite for the assembly of NPs in liposome vesicle membrane is that the core size is below 6.5 nm. This is only possible if NP cores are individually stabilized. Thus, controlled surface modification of SPIONs is key. A dense, thin dispersant layer where dense refers to a dispersant packing density above 0.2 dispersant/nm , preferably above 0.5 molecule/nm most preferably above 1 dispersant/nm², which is thick enough to prevent NP agglomeration is therefore a prerequisite to successfully functionalize liposomes with NPs. Therefore, the dispersant should be larger than a single catechol such as nitrocatechols but can be as short as palmityl-nitroDOPA or oleyl-nitroDOPA or shorter. The preferred dispersant layer thickness is palmityl-nitroDOPA or oleyl-nitroDOPA which results in sufficient NP stability while it still leads to a thin dispersant layer and a low mass fraction of dispersants (around 10-20 wt%) (Figure 12). Dispersants have to be hydrophobic, preferably linear and can comprise unsaturated bonds such as oleyl-nitroDOPA to incorporate such stabilized NPs into the membrane interior of liposomes and polymersomes.. However, dispersants can also consist of dendritic structures.

The magnetic response (heating or force) of the NP scales with the SPION volume and actuation is much facilitated if the SPION diameter is maximized. Therefore, it is desirable to precisely control the SPION size such that it is as big as possible while still giving a high yield of incorporation into a non-leaky membrane. Thus, if the dispersant layer thickness can be minimized by closely controlling the SPION surface chemistry, the SPION core diameter and with it the magnetic response of an individual nanoparticle can be maximized.

Example 8: Acuation of supported lipid bilayers containing iron oxide nanoparticles in their membrane.

Liposomes prepared according to Example 4e were adsorbed in Si02 coated QCM-D crystals (qsense, Sweden) at a concentration of 50 μg/ml. Liposomes spontaneously formed supported lipid bilaysers (SLBs) (Figure 15). These SLBs could be actuated by approaching a small table-top magnet if they contained palmityl-nitroDOPA stabilized iron oxide nanoparticles in their membranes (Figure 15).

Example 8: Co-attachment of FITC-nitroDOPA and palmityl-nitroDOPA

The fluorescence of liposomes loaded with FITC -labeled iron oxide nanoparticles prepared as described in Example 4e was measured with a UV/VIS spectrometer (Cary El, Elmer Perkins). Figure 22 illustrates the co-attachment of FITC-nitroDOPA and palmityl- nitroDOPA on the magnetic particles leading to fluorescently and magnetically labeled liposome membranes. Preferably the molar ratio of palmityl-nitroDOPA is such to provide sufficient steric stability to the SPIONs. Thus, the molar ratio of palmityl-nitroDOPA:FITC- nitroDOPA may preferably be smaller than 1:1. Alternatively, fluorophores such as FITC can be attached to the end of the hydrophobic dispersant resulting e.g. in FITC-palmityl- nitroDOPA or similar dispersants where the length of the hydrophobic chain is changed.

Example 9: Loading of SPION functionalized MPPC liposomes

The fluorescence of FITC labeled dextran loaded liposomes assembled as described in Example 4i and FITC labeled dextran loaded PMCL-PEO polymersomes assembled as described in Example 6b was measured with a UV/VIS spectrometer (Cary El, Elmer Perkins). (Figure 23).

Example 10: SLB formation on porous SiO^nanoparticles

POPC liposomes containing palmityl-nitroDOPA stabilized SPIONs in their bilayer spontaneously formed SLBs if adsorbed on Si02 surfaces as quartz crystal microbalance with dissipation monitoring (QCM-D) measurements revealed (Figure 14). Furthermore, Figure 15 reveals that a, b) SPION functionalized POPC liposomes can be actuated with an external small magnet. This is in contrast to POPC SLBs that have no SPIONs in their bilayer as c, d) control measurements of pure POPC SLBs reveal. Frequency changes (solid line) can be translated into changes in the adsorbed mass and dissipation changes (dotted line) indicate changes in the viscoelastic behavior of adsorbed films. Furthrmore, POPC vesicles containing 6 mol% PEG(2)-PE formed SLBs on 200 nm core diameter Si0₂ nanoparticles if incubated at RT for 12 h as was shown with cryo-TEM showing feasibility of coating porous Si0₂ nanoparticles with SLBs.

Additionally, 123 nm diameter liposomes form SLBs on nanoporous silicon nitride substrate with 100 nm in diameter pores (Figure 16a). In fact, the SLB spans these pores as indicated by reflection images water soluble carboxyfluorescein dye was captured and contained in the fraction of pores that could be shown to be spanned by a lipid bilayer (non-fluorescent pores have the membrane following the walls of the pore) (Figure 16b).

Example 11 : Assembly of layer-bv-laver structure of liposomes.

Feasibility to build layer-by-layer structures was shown in QCM-D measurements (Figure 24) QCM-D measurements were performed on DNA tethered, SPION functionalized POPC liposomes. These liposomes were bound to POPC SLBs surfaces presenting the complementary DNA sequence. The frequency (solid line) and dissipation shift (dashed line) upon addition of DNA tethered liposomes shows strong binding of tagged, SPION functionalized liposomes. Furthermore, polyelectrolyte multilayers incorporating intact liposomes which incorporate dyes over long periods of time without passive leakage can be assembled by layer-by-layer method, as described in (Graf, N. et al, Advanced Functional Materials, 2011, 21: 1666-1672). The content of the liposomes could later be release by increasing the local pH by application of current to an underlying electrode. Figure 25a shows the build-up of a polyelectrolyte multilayer embedded vesicle layer on an inorganic substrate recorded by quartz crystal microbalance with dissipation monitoring. The process can be repeated to create multilayers of liposomes. Figure 25b showns fluorescence microscopy image of 5,6-Carboxyfluorescein encapsulated in the liposome lumen. The square bleach spot in the image shows the contrast afforded by the dye. The dye intensity showed no decrease over lh while pH-induced destruction of the liposomes gave complete removal of the fluorescence signal. Substrates for these experiments were prepared as follows: Polyelectrolyte multilayers were made from Poly-L-Glutamic-Acid (PGA) (SIGMA), Poly-L- lysine hydrobromide (PLL) (Sigma Aldrich P7890, MW = 24000) and poly(sodium 4-styrene sulfonate) (PSS) (Sigma Aldrich 24,305, MW = 70000). Polymers were dissolved in 150 mM NaCl solution. (PLL/PSS)n means n layer pairs of the polyelectrolyte couple PLL and PSS.

The substrates were cleaned by ultrasonication (Elna Transsonic Digital S, Iswork, Singapore) in Cleaner (Cobas Integra Cleaner, Roche), Isopropanol (puriss., Sigma Aldrich) and ultrapure water (resistivity = 18.2 ΜΩ/cm, Milli-Q gradient A 10 system, Millipore

Corporation) for 10 minutes in each subsequent solvent. The samples were rinsed with ultrapure water and dried with nitrogen after every cleaning step. Finally, they underwent UV/Ozone cleaning (Uvo Cleaner 42-220, Jelight Company, USA) for 30 minutes prior to spraying. The QCM sensors were immersed for 30 minutes in sodium dodecyl sulfonate (SDS; Sigma Aldrich), rinsed with ultrapure water and then cleaned by UV/Ozone for 30 minutes before spraying.

The spraying process as described in (Izquierdo, A., et al., Langmuir, 2005, 21 7558-7567) has been automated by a home-built spraying robot. The custom made program first wets the substrate with 150 mM NaCl solution for 5 seconds. After a pause of 5 seconds the PLL solution (0.5 mg/ml) was sprayed for 5 seconds. After a pause of 15 seconds the substrate was rinsed with buffer or 150 mM NaCl solution for 5 seconds. After another 5 seconds break, the PSS solution (0.5 mg/ml) was sprayed for 5 seconds, followed by 15 seconds pause and rinsing of the substrate with buffer or 150 mM NaCl solution for 5 seconds. This procedure was repeated 9.5 times in order to obtain a (PLL/PSS)g-PLL multilayer (Guillaume-Gentil, O., et al., Soft Matter, 2010, 6: 4246-4254) on both EC-QCM crystals and microscopy slides. The PEM coated samples were stored at 4°C in 150 mM NaCl solution for maximal 4 weeks until use.

EC-CLSM observations were carried out on a Zeiss LSM 510 microscope. NBD, fluorescence was detected after excitation at 488 nm with 4% laser power, cutoff dichroic mirror 488 nm, and emission band pass filter 505-530 nm (green). Bleaching of fluorescence was accomplished by exposure to maximum laser intensity in the designated area before renewed imaging at 4% excitation intensity.

QCM measurements were carried out with a QCM-D system from Q-Sense (Sweden), described in detail (Rodahl, M., et al., Review of Scientific Instruments, 1996, 67: 3238- 3241). Briefly, QCM measures the changes in the resonance frequency (Δί) of a quartz crystal when material is adsorbed onto it. The quartz crystal was excited at its fundamental frequency (about 5 MHz) and at the third, fifth, and seventh overtones (n). A home-built, three- electrode, electrochemistry flow cell was used for the EC-QCM-D measurements (Grieshaber, D., et al., Langmuir, 2008, 24: 13668-13676). The ITO coated surface of the quartz crystal sensor was the working electrode whereas the Ag AgCl reference and the platinum counter electrodes were situated on the upper side of the flow cell.

Shortly before mounting into the EC-QCM flow cell, the PEM-covered quartz crystal was rinsed with ultrapure water and dried with nitrogen followed by rehydration in 150 mM NaCl solution within a few minutes. This procedure assured that no salt crystals formed upon drying the samples. The vesicle solution (0.5 mg/ml) was then injected and incubated for 30 minutes. After that, the flow cell was rinsed with 150 mM NaCl solution. Two additional layer pairs of (PLL/PGA) were then injected as described in (Guillaume-Gentil, O., et al., Advanced Materials, 2008, 20: 560-+). The adsorption of the each layer was monitored and shown in Figure 25 a. The PEM covered samples were mounted after quick rinsing with ultrapure water (Resistivity = 18.2 ΜΩ/cm) and drying with nitrogen. The teflon flow cell was provided with a silver wire which was anodized in chloride to be an Ag/AgCl reference electrode and with a platinum wire as counter electrode. A copper spring established the electrical contact with the ITO surface of the substrate used as working electrode. The liposomes were then adsorbed in situ on the pre-sprayed (PLL/PSS)9-PLL PEM for approximately 30 minutes. The substrate was extensively rinsed (7 x 400 μΐ) after adsorption of the liposomes. The integrity of the membrane-labeled liposomes adsorbed on the PEM layer was assured by means of fluorescence recovery after photobleaching (FRAP).

FRAP measurements revealed that the dye 5,6-carboxyfluorescein that was encapsulated in the liposome lumen (Figure 25b). The square bleach spot in the image shows the contrast afforded by the dye. The dye intensity showed no decrease over lh while pH-induced destruction of the liposomes gave complete removal of fluorescence signal.

Claims

Claims

1. Magnetically responsive composition comprising a membrane and at least one stabilized magnetic nanoparticle embedded in said membrane, wherein the membrane is in form of a magnetically responsive vesicular structure or in form of a magnetically responsive supported lipid bilayer.

2. Magnetically responsive composition according to claim 1, wherein the magnetically responsive vesicular structure comprises (a) a vesicular structure having a membrane enclosing a cavity and (b) at least one stabilized magnetic nanoparticle embedded in said membrane.

3. Magnetically responsive composition according to claim 2, wherein the vesicular structure is a liposome comprising at least one lipid type or a polymersome comprising at least one synthetic and/or natural polymer.

4. Magnetically responsive composition according to any preceding claim wherein the stabilized magnetic nanoparticle is selected from the group consisting of iron, cobalt or nickel, alloys thereof, preferably oxides or mixed oxides/hydroxides, nitrides, carbides or sulfides thereof.

5. Magnetically responsive composition according to any preceding claim wherein the stabilized magnetic nanoparticles are superparamagnetic iron oxide nanoparticles

6. Magnetically responsive composition according to any preceding claim wherein the core diameter of the stabilized magnetic nanoparticles is between 2 and 10 nm.

7. Magnetically responsive composition according to any preceding claim wherein the stabilized magnetic nanoparticle is a magnetic nanoparticle associated with at least one dispersant, preferably a catechol derivative anchor group covalently bound to a hydrophobic spacer.

8. Magnetically responsive composition according to claim 7, wherein the hydrophobic spacer is a polymeric spacer and is selected from the group consisting of linear, branched and dendritic hydrocarbon chains.

9. Magnetically responsive composition according to claim 7, wherein the hydrophobic spacer comprises a functional group such as a fluorophore or chelate binding MR-active ions or radiotracers.

10. Magnetically responsive composition according to claim 7, wherein one or more of the at least one dispersant adsorbed on the nanoparticle surface further comprises a chemically reactive group.

11. Magnetically responsive composition according to any preceding claim further comprising at least one active agent.

12. Magnetically responsive composition according to claim 11, wherein the at least one active agent is a therapeutic or diagnostic agent, a nutritional agent, an enzyme, a growth factor or a targeting group or combinations thereof.

13. Magnetically responsive composition according to any preceding claim further comprising a polymeric coat of a stealth polymer tethered to the membrane

14. Magnetically responsive composition according to any preceding claims further comprising at least one functional group such as NHS ester, maleimide, photoinitiator, acrylates, methacrylates, amines, carboxy groups or physically strongly interacting groups such as chelates, charged groups or specifically binding protein ligands.

15. Magnetically responsive composition according to claim 14, wherein the at least one reactive group is linked to the membrane, preferably to the at least one lipid or the at least one synthetic and/or natural polymer of the membrane.

16. Magnetically responsive composition according to any preceding claim which is encapsulated in a matrix such as a physically or chemically crosslinked hydrogel or embedded in a matrix tethered to a substrate.

17. Magnetically responsive composition according to any preceding claim to for use in (targeted) delivery of an active agent, as a nanoreactor, or for imaging purposes.

18. A method of locally accumulating a magnetically responsive composition according to to any one of claims 1-17 by application of a magnetic field.

19. A method according to claim 18, wherein the magnetically responsive composition is tethered to a liquid crystalline surface such as a lipid bilayer through mobile linkers and optionally can be magnetically locally accumulated in the surface plane.

20. A method of changing the permeability of a magnetically responsive composition according to any one of claims 1-17, said method comprising the step of exposing said magnetically responsive composition to an alternating or direct magnetic field.

21. A method for delivery of an active agent in a subject, comprising the steps of: (a) associating at least one active agent with a magnetically responsive vesicular composition according to any one of claims 1-17, (b) administering said composition to a subject; and (c) exposing said composition to a direct or alternating magnetic field to effect release of said active agent in said subject. A method for delivery of an active agent in cell or tissue culture, comprising the steps of: (a) associating at least one active agent with a magnetically responsive vesicular composition according to any one of claims 1-17, (b) incorporating said composition in the cell or tissue culture directly or as part of the scaffold; and (c) exposing said composition to a direct or alternating magnetic field to effect release of said active agent.