Capsosomes with “Free-Floating” Liposomal Subcompartments

4082

www.advmat.dewww.MaterialsViews.com

CO

MM

UN

ICATI

ON

Leticia Hosta-Rigau , Shiow Fong Chung , Almar Postma , Rona Chandrawati , Brigitte Städler , and Frank Caruso *

Capsosomes with “Free-Floating” Liposomal Subcompartments

Biological cells are able to perform multiple complex reactions within confi ned environments, owing to their structures com-prising internal subcompartments (e.g., cell organelles). [ 1 ] Arti-fi cial cells, [ 2–4 ] although far less complex than their biological counterparts, can exhibit a hierarchical structure with a large number of subcompartments confi ned within a structurally stable scaffold. Such systems are engineered for therapeutic cell mimicry via the encapsulation and/or conversion of biologically active materials. [ 5 ]

Reported examples of subcompartmentalized micrometer-sized assemblies include multiple subcompartments in the form of micelles, [ 6 ] liposomes, [ 1 , 7–9 ] or polymersomes [ 10 ] entrapped within a carrier vehicle; dual compartmentalized polymeric cap-sules; [ 11 , 12 ] three-compartment microparticles; [ 13 ] polymer cap-sules containing smaller polymer capsules, [ 14 ] cubosomes, [ 15 ] or polymersomes; [ 16 ] capsules embedded in a polyelectrolyte multilayer-stabilized gel bead; [ 17 ] and cellosomes, prepared by the assembly of polyelectrolyte-coated yeast cells onto a tem-plate that is subsequently removed. [ 18 ] Recently, we reported the assembly of capsosomes–polymer (hydrogel) carrier capsules containing liposomal subcompartments. [ 19 ] While the liposomes provide protection for encapsulated fragile biological or small hydrophobic and hydrophilic material, the polymeric capsule provides the structural scaffold, and the semipermeable nature of the polymer membrane enables interaction with the external milieu. We have demonstrated that capsosomes are an effi cient functional encapsulation platform with potential use in drug delivery or as artifi cial mimics of cells. In particular, we have described capsosomes containing multilayers of different types of liposomes (zwitterionic or negatively charged, saturated or unsaturated) embedded within either nondegradable [ 20 ] or (bio)degradable [ 21 ] polymer carrier capsules. Hydrophilic [ 21–23 ] and hydrophobic [ 24 , 25 ] cargo have been encapsulated within the lipo-somal subcompartments embedded within disulfi de-stabilized

© 2011 WILEY-VCH Verlag Gwileyonlinelibrary.com

Dr. L. Hosta-Rigau , S. F. Chung , Dr. A. Postma, R. Chandrawati , Dr. B. Städler, [+] Prof. F. Caruso Department of Chemical and Biomolecular EngineeringThe University of MelbourneParkville, Victoria 3010, Australia E-mail: [email protected] Dr. A. Postma CSIRO Materials Science and EngineeringClayton, Victoria 3168, Australia [ + ] Present address: Interdisciplinary Nanoscience Center (iNANO), Aarhus University, Aarhus 8000, Denmark

DOI: 10.1002/adma.201100908

poly(methacrylic acid) (PMA) carrier capsules, and preservation of cargo functionality has been confi rmed through enzymatic assays [ 21 , 23 ] and cell viability studies. [ 24 , 25 ] In our previous work, all of the capsosomes contain liposomal subcompartments attached to the polymeric capsule wall. While the loading effi ciency of the subunits is not expected to be affected, the functional activity of capsosomes could benefi t from spatial control over the positioning of the subcompartments, that is, membrane-associated or “free-fl oating” liposomes in the cap-sule interior (similar to the organelles in biological cells). This is benefi cial especially when fast mixing is essential or equal access of reagents to the subcompartments is required, in par-ticular when channel-gated liposomes are considered. Further, controlling multistep reactions may be facilitated when the subunits containing the reagent for the initial step are located close to the scaffold membrane, while those loaded with rea-gents for subsequent conversion and/or synthesis are situated in the interior of the assembly.

With the goal to further advance our cell mimicry approach, herein, we report the assembly of capsosomes with controlled positioning of the liposomal subunits within the polymer hydrogel carrier capsules via the pH-dependent activity of hydrogen bonds between poly( N -vinyl pyrrolidone) (PVP) and PMA ( Scheme 1 ). In particular, we: i) synthesized poly( N -vinyl pyrrolidone)- block -(cholesteryl acrylate) (PVP c ), a block copolymer consisting of a short block of cholesteryl acrylate and a longer PVP segment, via reversible addition-fragmentation chain transfer [ 26 ] (RAFT) and confi rm the selective adsorption of liposomes to PVP c ; ii) demonstrate the effect of sacrifi cial polymer layers on the loading effi -ciency of liposomes into capsosomes; iii) confi rm the pres-ence of “free-fl oating” liposomes within capsosomes at pH 7.4; iv) assembled capsosomes containing both “free-fl oating” and membrane-associated liposomes; and v) demon-strate preservation of cargo functionality by performing a triggered enzymatic reaction encapsulating a protease in the liposomal subcompartments.

Electrostatic interactions alone can be limiting in the choice of liposomes and polymers that can be assembled into a cap-sosome due to restrictions in affi nity between the liposomes and the precursor and/or capping polymer layers. Hence, we pioneered a non-covalent linkage concept based on the use of cholesterol-modifi ed polymers, which promotes successful anchoring of the liposomes to a polymer fi lm and does not induce rupturing or displacement of the liposomes from the surface upon deposition of the polymer capping layer. [ 21 ] We previously introduced two cholesterol-containing polymers, poly(methacrylic acid)- co -(cholesteryl methacrylate) (PMA c )

mbH & Co. KGaA, Weinheim Adv. Mater. 2011, 23, 4082–4087


CO

MM

UN

ICATIO

N

Scheme 1 . Schematic illustration of a capsosome containing “free-fl oating” liposomal subcompartments. The fi rst inset shows the block copolymer PVP c coating a liposome (i), while the second inset is an illus-tration of the intercalation of the cholesteryl moieties attached to the PVP c into the lipid bilayer of a liposome (ii).

and cholesterol-modifi ed poly(L-lysine) (PLL c ). [ 21 ] Both of these polymers lead to the formation of capsosomes but neither

Figure 1 . a) Fluorescence intensity of silica particles pre-coated with a variable number of PVP/PMA bilayers and PVP c or PVP upon the adsorption of fl uorescently labeled liposomes (L u, − NBD ). b) The fl uorescence intensity of particles or capsosomes due to the presence of L u, − NBD at different stages of the assembly: after the initial liposome deposition (black), after the assembly of the core-shell particles (dark grey), capsosomes in NaOAc buffer (light grey) and in HEPES buffer (striated light grey). Red arrows indicate the decrease in fl uorescence intensity of the capsosomes upon buffer change. c) CLSM images of capsosomes (PVP c /L u, − NBD /PVP c /PMA/PVP/(PMA SH /PVP) 4 ) showing the liposomes associated with the polymer membrane in NaOAc buffer (i) and “free-fl oating” within the capsules in HEPES buffer (ii). In HEPES buffer, the PMA SH membrane has been post-labeled with Alexa Fluor-633 maleimide.

of them promotes the formation of “free-fl oating” liposomes in the capsule interior. This is not surprising for PLL c because of its electrostatic attraction with the PMA of the carrier capsule. PMA c , on the other hand, is negatively charged and is not cross-linked to the polymer membrane of the carrier, but the liposomes remain associated with the capsule wall, most likely due to physical entanglement of the copolymer with the building blocks of the polymer membrane. We therefore utilize a tailor-made cholesterol-containing block-copolymer that is (a) uncharged and (b) contains cholesterol moieties in a short block to minimize the entanglement of the liposome-anchoring polymer with the PMA capsule; namely PVP c . This polymer can anchor liposomes via its cholesteryl-block while the uncharged PVP-block forms hydrogen bonds with PMA only during the assembly of the carrier membrane (pH 4) but not at physiological conditions (pH 7.4).

The RAFT synthesized block copolymer, PVP c , was found to have a total molecular weight of 10.9 kDa, with 3 mol% of choles-terol and a narrow polydispersity of 1.22 (for more details see the Supporting Infor-mation, Scheme S1, Figure S1 and S2). We examined the ability of PVP c to adsorb lipo-somes onto 3- μ m-diameter silica particles via noncovalent anchoring of the cholesterol moieties to the lipid membrane. Addition-ally, we compared the amount of adsorbed liposomes onto particles with different numbers of preadsorbed bilayers of PVP and PMA. Negatively charged fl uorescently labeled unsaturated liposomes (L u,− NBD ),

© 2011 WILEY-VCH Verlag GmAdv. Mater. 2011, 23, 4082–4087

1,2-dioleoyl- sn -glycero-3-phosphocholine (DOPC) and 1,2-dioleoyl- sn -glycero-3-(phospho-L-serine) (DOPS) (DOPC:DOPS = 4:1 wt/wt, L u,− ), and fl uorescent lipids (1 wt% 1-myristoyl-2-[12-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]-hexanoyl]- sn -glycero-3-phosphocholine (NBD-PC)), were employed to maxi-mize the electrostatic repulsion between the negatively charged disulfi de-stabilized PMA carrier capsule and the liposomes at physiological conditions. The carboxylic groups of the PMA chains are deprotonated at pH 7.4, and this should facilitate detachment of the liposomes from the polymer membrane. The fl uorescence intensity of silica particles due to adsorp-tion of fl uorescently labeled liposomes was monitored by fl ow cytometry ( Figure 1 a). The largest amount of liposomes depos-ited was observed when the liposomes were adsorbed onto silica particles only precoated with PVP c . The adsorption of PVP/PMA bilayers prior to PVP c reduced the amount of lipo-somes adsorbed by at least 70%. This observation is attributed to a reduced number of cholesteryl units protruding from the surface due to the presence of underlying polymer bilayers and, hence, less units being accessible for anchoring the liposomes. Importantly, there was negligible adsorption of liposomes to the particles coated with unmodifi ed PVP as a precursor layer,

4083bH & Co. KGaA, Weinheim wileyonlinelibrary.com

4084


CO

MM

UN

ICATI

ON
demonstrating the need for the cholesteryl moieties of PVP c to
anchor the liposomes. After having identifi ed PVP c as a specifi c and suitable pre-

cursor polymer layer for L u, − NBD adsorption (without the need for PVP/PMA bilayers), capsosomes were assembled by cap-ping the liposomes with PVP c and subsequently adsorbing a variable number of PMA/PVP separation/sacrifi cial bilayers. These polymer bilayers are expected to serve as sacrifi cial layers because they disassemble under specifi c conditions, con-sequently assisting in detachment of the liposomes from the polymer membrane. The polymer carrier capsules were subse-quently created by the sequential deposition of four bilayers of thiol-modifi ed PMA (PMA SH ) and PVP in sodium acetate buffer (20 mM NaOAc, pH 4). [ 27 ] The thiol groups in the polymer multilayer fi lm of these core–shell particles were cross-linked into disulfi de linkages using chloramine T prior to dissolution of the template particles in buffered hydrofl uoric acid (HF) to yield capsosomes. The retention effi ciency of the fl uorescently labeled liposomes within the polymer multilayer fi lm as a func-tion of the number of sacrifi cial separation bilayers was assessed by fl ow cytometry. The fl uorescence intensity of the particles due to the initial adsorption of fl uorescent liposomes was set to 100% and compared to the fl uorescence intensity of the core–shell particles after cross-linking as well as the capsosomes (after core dissolution) in NaOAc buffer and in HEPES buffer [10 m M HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid), 150 m M NaCl, pH 7.4] (Figure 1 b). The core–shell particles and the capsosomes in NaOAc buffer showed similar fl uorescence intensities for all of the different assemblies, suggesting that the different number of separation PMA/PVP bilayers did not affect liposome retention. However, upon changing the environment of the capsosomes from NaOAc to HEPES buffer, the number of stably trapped liposomes varied, depending on the number of separation PMA/PVP bilayers. Increasing the number of sacri-fi cial separation bilayers caused a decrease of the fl uorescence intensity of the capsosomes upon buffer change (Figure 1 b). While in the absence of PMA/PVP bilayers a reduction of only ≈ 10% was observed, the presence of two PMA/PVP bilayers gave rise to ≈ 45% lower intensity. The increase in pH results in the disruption of the hydrogen bonds, and thus the (sacrifi cial) PVP and nonthiolated PMA were destabilized and released, possibly either dragging the liposomes along or temporarily increasing the permeability of the polymer membrane, which might facili-tate loss of the liposomes. Release of PVP from the multilayer fi lm has previously been shown, [ 28 ] and the release of PMA at pH 7.4 was confi rmed (Supporting Information, Figure S3). The yellow signal arising from the superposition of green fl uo-rescently labeled liposomes (L u, − NBD ) and red PMA AF633 was observed (Supporting Information, Figure S3a). Upon buffer change, the sacrifi cial layers were disrupted and PMA AF633 was released (Supporting Information, Figure S3b). Capsosomes were assembled via this process (i.e., using PVP c as precursor and capping layers without additional PMA/PVP sacrifi cial bilayers) with the goal to achieve the highest liposome reten-tion and were characterized both in NaOAc buffer and under physiological conditions using optical and fl uorescence micro-scopy. Differential interference contrast images (Supporting Information, Figure S4) demonstrated that the capsosomes obtained were structurally intact and non-aggregated under

© 2011 WILEY-VCH Verlag wileyonlinelibrary.com

both conditions. Confocal laser scanning microscopy (CLSM) images of PVP c /L u, − NBD /PVP c /PMA/PVP/(PMA SH /PVP) 4 in NaOAc buffer showed regular green fl uorescence associated with the capsosome membrane, which arose from the green labeled liposomes (Figure 1 c(i)). Upon buffer change, the hydrogen bonds between the PVP c and PMA are disrupted and thus the liposomes trapped between the PVP c layers were detached from the membrane of the carrier capsule. This is shown by the green fl uorescence homogeneously distributed within the capsosomes (Figure 1 c(ii)). The “free-fl oating” green liposomes were stably entrapped within the red-labeled PMA hydrogel carrier capsules (the polymer membrane was post-labeled with (Alexa Fluor-633)-maleimide in HEPES buffer). Further, detachment of the liposomes was found to be revers-ible. Upon reverting back to NaOAc buffer, the liposomes reat-tached themselves to the polymer membrane, suggesting that the PVP c was not released from the capsosomes, but rather that the cholesterol-modifi ed polymer remained largely associated with the liposomes (Supporting Information, Figure S5). From fl uorescence analysis, we estimate that each 3- μ m-diamater capsosome contains ≈ 5000 “free-fl oating” liposomal subcom-partments (see the Supporting Information).

We next sought to obtain capsosomes comprising both “free-fl oating” and membrane-associated liposomes. We assembled liposomes in two deposition steps, expecting the inner lipo-some layer to become “free-fl oating” while the outer layer to remain associated with the polymer membrane. The fi rst layer of liposomes was adsorbed onto PVP c pre-coated silica par-ticles while the second layer was anchored to (PVP c /L u, − NBD /PVP c /PMA/PVP/PMA c )-modifi ed particles. The measured fl uorescence intensity of the silica particles after the fi rst and the second liposome deposition steps showed that a similar number of liposomes adsorb in each step (Supporting Infor-mation, Figure S6), demonstrating that the initial layers did not hinder subsequent liposome assembly. Capsosomes were then formed by capping with PMA c and assembling the PVP/PMA SH polymer membrane, followed by cross-linking of the thiols in the fi lm and removing the core. CLSM images of these capsosomes containing NBD-labeled (L u, − NBD ) and rhodamine-labeled (L u, − Rhd ) liposomes consisting of DOPC:DOPS = 4:1 wt/wt and fl uorescent lipids 1 wt% 1,2-dipalmitoyl- sn -glycero-3-phosphoethanolamine-N-(lissamine Rhodamine B sulfonyl) (Rhd-PE) showed both types of liposomes associated with the polymer membrane in NaOAc buffer (assembly conditions) ( Figure 2 a(i) and 2 a(i ′ ). Upon changing the buffer to HEPES, the green fl uorescently labeled liposomes were observed to be “free-fl oating” inside the capsosomes, as shown by the homo-geneously distributed green emission (Figure 2 a(ii)). The red-fl uorescently labeled liposomes remained attached to the shell (Figure 2 a(ii ′ )), as shown by the red emission emerging from the walls of the capsules. This demonstrates spatial control over the positioning of the liposomal subcompartments, a crucial step forward in assembling confi ned hierarchical structures. How-ever, when assembling the same system but containing unla-belled liposomes in the inner layer (L u, − ) and green labeled lipo-somes in the outer layer (L u, − NBD ), the green labeled liposomes became “free-fl oating” in the capsule interior in HEPES buffer (Supporting Information, Figure S7). These results suggest that control over the position of the liposomes is a combination of

GmbH & Co. KGaA, Weinheim Adv. Mater. 2011, 23, 4082–4087


© 2011 WILEY-VCH Verlag Gm

CO

MM

UN

ICATIO

N

Adv. Mater. 2011, 23, 4082–4087

Figure 2 . a) CLSM images of capsosomes (PVP c /L u, − NBD /PVP c /PMA/PVP/PMA c /L u, − Rhd /PMA c /PVP/(PMA SH /PVP) 4 ) containing a layer of green (i), (ii) and red (i ′ ), (ii′) labeled liposomes in NaOAc buffer and in HEPES buffer (left and right, respectively). b) Fluorescence microscopy images of (i) PVP/PMA c /L u, − NBD /PMA c /PVP/PMA/PVP/(PMA SH /PVP) 4 -capsosomes and (ii) PVP c /L u, − NBD /PVP c /PMA/PVP/(PMA SH /PVP) 4 -capsosomes at physiological conditions. c) Fluorescence microscopy images of PVP c /L s, − NBD /PVP c /PMA/PVP/(PMA SH /PVP) 4 -capsosomes in (i) NaOAc buffer and (ii) in HEPES buffer.

(i) the different properties of PVP c and PMA c and (ii) the dif-ferent fl uorophore-conjugated lipids (i.e., NBD-PC and Rhd-PE). For the former case, the assembly with a PMA c capping layer resulted in membrane-associated liposomes (Figure 2 b(i)), while when assembling the same capsosomes but using PVP c as a capping layer, the liposomes became “free-fl oating” at phys-iological conditions (Figure 2 b(ii)). For the latter case, the lipid NBD-PC contains the fl uorophore attached to the hydrophobic tail of the phospholipids (Supporting Information, Figure S8a), while the Rhd-PE lipid contains the fl uorophore anchored to the hydrophilic head of the phospholipid (Supporting Infor-mation, Figure S8b). Thus, when assembling liposomes con-taining NBD-PC lipids, the fl uorophore will be enclosed within the lipid membrane of the liposomes and therefore is unlikely to interact with other compounds, while the fl uorophore of the Rhd-PE lipids will remain exposed for interaction with other species.

To examine the functionality of capsosomes containing “free-fl oating” liposomes, we encapsulated the protease subtilisin in the subunits of the capsosomes and performed a triggered enzy-matic reaction by converting the coumarine peptidase substrate N -CBZ-L-phenylalanyl- L -arginine 7-amido-4-methylcoumarin (AMC s ) into its hydrolyzed product 7-amino-4-methylcoumarin (AMC p ). Depending on the purpose of the capsosomes, different types of liposomes might be employed. For the encapsulation of enzymatic cargo, saturated liposomes are preferably used due to their higher phase-transition temperature ( T m ). Nega-tively charged saturated liposomes (L s, − NBD ), 1,2-dimyristoyl- sn -glycero-3-phosphocholine (DMPC), 1,2-dipalmitoyl- sn -glycero-3-phosphocholine (DPPC) and 1,2-dipalmitoyl- sn -glycero-3-(phospho-L-serine) (DPPS) (DMPC:DPPC:DPPS = 8:1:1 wt/wt, T m ≈ 28 ° C) and fl uorescent lipids (1 wt% of NBD-PC) were employed to assemble PVP c /L s, − NBD /PVP c /PMA/PVP/(PMA SH /PVP) 4 , and after cross-linking and core removal capso-somes were observed by fl uorescence microscopy. As depicted in Figure 2 c(i), in NaOAc buffer a regular green fl uorescence signal arising from the shell of the capsosomes was observed, while upon changing to HEPES buffer, the green fl uorescence signal was homogeneously distributed throughout the capso-some cavity (Figure 2 c(ii)). This demonstrates “free-fl oating” saturated liposomes in the capsule interior. The choice of the enzyme subtilisin and the AMC s made it possible to follow and quantify the conversion of the substrate into the product by fl uorescence spectrophotometry (AMC p excitation/emission maxima at ≈ 351/430 nm, Supporting Information, Figure S9). Saturated negatively charged liposomes loaded with subtilisin (L s, − Sub ) were assembled in two different ways, one consisting of PVP c /L s, − Sub /PVP c /PMA/PVP/(PMA SH /PVP) 4 -capsosomes, which promotes “free-fl oating” of the liposomal subcompart-ments, and PLL/L s, − Sub /PMA c /PVP/(PMA SH /PVP) 4 -capsosomes, which generates capsosomes containing liposomes attached to the inside of the polymer shell ( Figure 3 a). We used the enhanced permeability of the liposomal membrane at elevated temperature to trigger the enzymatic reaction. Equal numbers of subtilisin-containing capsosomes were incubated with AMC s and heated at T = 23 ° C (RT, room temperature) or at T = 37 ° C (above the T m of the liposomes). AMC s was also incubated with free subtilisin at T = 23 ° C (Figure 3 b, column 1) or at T = 37 ° C (Figure 3 b, column 2) at a similar concentration as the subtilisin


4086


CO

MM

UN

ICATI

ON

Figure 3 . a) Schematic illustration of the enzymatic conversion of AMC s into AMC p using capsosomes loaded with the protease subtilisin upon increasing the temperature above T m . b) Fluorescence intensity of the AMC p after 4, 17, and 24 h due to the enzymatic activity by: free subtilisin at a similar concentration as the subtilisin entrapped within the liposomal compartments of the capsosomes incubated at 23 ° C ( 1 ) or 37 ° C ( 2 ); capsosomes containing the subtilisin-loaded membrane-associated lipo-somal compartments at 23 ° C ( 3 ) or 37 ° C ( 4 ); and capsosomes con-taining the subtilisin-loaded “free-fl oating” liposomal compartments incubated at 23 ° C ( 5 ) or 37 ° C ( 6 ).

encapsulated in the capsosomes. As depicted in Figure 3 b, the fl uorescence intensity of AMC p was measured at 436 nm after 4 h (white columns), 17 h (light grey columns) and 24 h (dark grey columns) of incubation. After 24 h of incubation, both cap-sosomes containing liposomes attached to the polymer shell and capsosomes containing “free-fl oating” subcompartments at 37 ° C showed a normalized fl uorescence intensity of ≈ 100%, a value close to the fl uorescence intensity of the AMC p converted

© 2011 WILEY-VCH Verlag Gwileyonlinelibrary.com

by the free subtilisin in solution ( ≈ 90%) (Figure 3 b, dark grey columns 4, 6, and 2). Upon increasing the temperature the liposomal membrane permeability is enhanced, and the small substrate AMC s is able to cross through the liposomes, thus interacting with the enzymes and being converted into AMC p (Figure 3 a). This result demonstrates the effi ciency in the encapsulation of the enzyme within the liposomal subcompart-ments and preservation of cargo functionality (we achieved a similar conversion of AMC s when the enzyme is trapped in the liposomal subcompartments and when the enzyme is free in solution). Although no difference was observed between capso-somes containing “free-fl oating” or membrane-associated sub-compartments when incubated at T = 37 ° C, the control sam-ples left at RT showed a difference. After 24 h it was observed that capsosomes containing membrane-associated liposomes showed ≈ 10% conversion of the substrate (Figure 3 b, column 3, dark grey) while the “free-fl oating” liposomes converted ≈ 50% of the substrate into its fl uorescent product (Figure 3 b, column 5, dark grey). This confi rms that temperature can be used to trigger the enzymatic reaction. In addition, this result gives rise to the possibility of using capsosomes with “free-fl oating” subunits in a continuous manner, that is, without the need of a trigger to initiate the reaction. The differences between the two types of capsosomes can be explained due to the different poly-mers used for their assembly. When assembling capsosomes containing the liposomal subcompartments attached to the shell walls we used PMA c , a graft copolymer. In contrast, when assembling capsosomes containing “free-fl oating” subcompart-ments, a block copolymer PVP c was used. The different distri-bution of the cholesteryl pendant units along the polymer chain as well as the type of polymer is likely to affect the liposomal membrane and is currently part of a more detailed investiga-tion. Further, we incubated the same capsosomes with AMC s solution one week after the assembly and performed the same enzymatic assay with the goal to demonstrate preservation of enzyme activity since the proteases are prone to self-degradation. As depicted in Figure S10 (Supporting Information), the enzy-matic activity of subtilisin is preserved when it is stored in solution or encapsulated in capsosomes containing either “free-fl oating” or membrane-associated liposomes.

In conclusion, the synthesis and use of a novel block copolymer PVP c to anchor liposomes to a polymer fi lm allows control over the spatial positioning of liposomes entrapped within a PMA hydrogel carrier capsule. We demonstrated that with the appropriate use of cholesterol-modifi ed polymers and PMA/PVP sacrifi cial layers, capsosomes with “free-fl oating” and polymer membrane-associated subcompartments can be prepared. The controlled enzymatic conversion of AMC s into its hydrolyzed product dem-onstrates the preservation of cargo functionality when the enzyme subtilisin is encapsulated into the “free-fl oating” or membrane-associated liposomal compartments of capsosomes. These are pivotal features on the way to further developing capsosomes into a functional biomedical platform and as mimics for cells.

Experimental Section Capsosomes with “free-fl oating” liposomal subcompartments were assembled via the layer-by-layer technique. A suspension of 3.25- μ m-diameter silica particles (5 wt%) in NaOAc buffer was incubated with

mbH & Co. KGaA, Weinheim Adv. Mater. 2011, 23, 4082–4087


CO

MM

UN

ICATIO

N

PVP c (1 mg mL − 1 , 15 min), washed three times, suspended in the liposome solution (1.25 mg mL − 1 , 40 min), and washed again three times, followed by incubation with PVP c (1 mg mL − 1 , 15 min). A sacrifi cial layer PMA (1 mg mL − 1 , 10 min) was adsorbed and the polymer membrane of the carrier capsules was assembled by sequential deposition of four bilayers of PVP and PMA SH (1 mg mL − 1 , 10 min). The thiols within the polymer layers were cross-linked with 1,8-bismaleimidodiethyleneglycol (BM(PEG) 2 ) (2 m M , 15 h) in MES buffer (50 m M MES (2-(N-morpholino)-ethanesulfonic acid), pH 6.0), and the cores were removed using a 2 M HF/8 M NH 4 F solution for 2 min, followed by several washing cycles in NaOAc buffer (4500 g, 3 min). A CyFlow Space (Partec Flowmax, Münster, Germany) fl ow cytometer using an excitation wavelength of 488 nm was used for all of the fl ow cytometry experiments. The fl uorescently labeled capsosomes were imaged using a Leica TCS SP2 SE confocal microscope. The conversion of the AMC s into its product was monitored over time by fl uorescence spectrophotometry (Fluorolog-3 Model FL3-22 spectrofl uorometer) using an excitation wavelength of 351 nm and recording the fl uorescence intensity at 436 nm.

Supporting Information Supporting Information is available from the Wiley Online Library or from the author.

Acknowledgements This work was supported by the Australian Research Council under the Federation Fellowship (F.C.) and Discovery Project (F.C.) schemes. We thank Dr. Alexander N. Zelikin (Chemistry Department, Aarhus University, Denmark) and Siow-Feng Chong (The University of Melbourne, Australia) for their input in assembling the PMA hydrogel carrier capsules.

Received: March 10, 2011Published online: July 27, 2011

[ 1 ] E. T. Kisak , B. Coldren , C. A. Evans , C. Boyer , J. A. Zasadzinski , Curr. Med. Chem. 2004 , 11 , 199 .

[ 2 ] D. Deamer , Trends Biotechnol. 2005 , 23 , 336 . [ 3 ] D. A. Drubin , J. C. Way , P. A. Silver , Genes Dev. 2007 , 21 , 242 . [ 4 ] S. Mann , Angew. Chem. Int. Ed. 2008 , 47 , 5306 . [ 5 ] T. M. S. Chang , Eur. J. Pharm. Biopharm. 1998 , 45 , 3 .

© 2011 WILEY-VCH Verlag GmAdv. Mater. 2011, 23, 4082–4087

[ 6 ] Z. B. Li , E. Kesselman , Y. Talmon , M. A. Hillmyer , T. P. Lodge , Science 2004 , 306 , 98 .

[ 7 ] W. T. Al-Jamal , K. Kostarelos , Int. J. Pharm. 2007 , 331 , 182 . [ 8 ] P. Y. Bolinger , D. Stamou , H. Vogel , Angew. Chem. Int. Ed. 2008 , 47 ,

5544 . [ 9 ] A. M. Yashchenok , M. Delcea , K. Videnova , E. A. Jares-Erijman ,

T. M. Jovin , M. Konrad , H. Möhwald , A. G. Skirtach , Angew. Chem. Int. Ed. 2010 , 49 , 8116 .

[ 10 ] H. C. Chiu , Y. W. Lin , Y. F. Huang , C. K. Chuang , C. S. Chern , Angew. Chem. Int. Ed. 2008 , 47 , 1875 .

[ 11 ] O. Kreft , M. Prevot , H. Möhwald , G. B. Sukhorukov , Angew. Chem. Int. Ed. 2007 , 46 , 5605 .

[ 12 ] O. Kreft , A. G. Skirtach , G. B. Sukhorukov , H. Möhwald , Adv. Mater. 2007 , 19 , 3142 .

[ 13 ] H. Baumler , R. Georgieva , Biomacromolecules 2010 , 11 , 1480 . [ 14 ] O. Kulygin , A. D. Price , S.-F. Chong , B. Städler , A. N. Zelikin ,

F. Caruso , Small 2010 , 6 , 1558 . [ 15 ] C. D. Driever , X. Mulet , A. P. R. Johnston , L. J. Waddington ,

H. Thissen , F. Caruso , C. J. Drummond , Soft Matter 2011 , 7 , 4257 .

[ 16 ] H. Lomas , A. P. R. Johnston , G. K. Such , Z. Zhu , K. Liang , M. P. van Koeverden , S. Alongkornchotikul , F. Caruso , Small 2011 , DOI: 10.1002/smll.201100744.

[ 17 ] B. G. De Geest , S. De Koker , K. Immesoete , J. Demeester , S. C. De Smedt , W. E. Hennink , Adv. Mater. 2008 , 20 , 3687 .

[ 18 ] R. F. Fakhrullin , V. N. Paunov , Chem. Commun. 2009 , 2511 . [ 19 ] B. Städler , A. D. Price , R. Chandrawati , L. Hosta-Rigau , A. N. Zelikin ,

F. Caruso , Nanoscale 2009 , 1 , 68 . [ 20 ] B. Städler , R. Chandrawati , K. Goldie , F. Caruso , Langmuir 2009 , 25 ,

6725 . [ 21 ] B. Städler , R. Chandrawati , A. D. Price , S.-F. Chong , K. Breheney ,

A. Postma , L. A. Connal , A. N. Zelikin , F. Caruso , Angew. Chem. Int. Ed. 2009 , 48 , 4359 .

[ 22 ] R. Chandrawati , L. Hosta-Rigau , D. Vanderstraaten , S. A. Lokuliyana , B. Städler , F. Albericio , F. Caruso , ACS Nano 2010 , 4 , 1351 .

[ 23 ] R. Chandrawati , B. Städler , A. Postma , L. A. Connal , S.-F. Chong , A. N. Zelikin , F. Caruso , Biomaterials 2009 , 30 , 5988 .

[ 24 ] L. Hosta-Rigau , R. Chandrawati , E. Saveriades , P. D. Odermatt , A. Postma , F. Ercole , K. Breheney , K. L. Wark , B. Städler , F. Caruso , Biomacromolecules 2010 , 11 , 3548 .

[ 25 ] L. Hosta-Rigau , B. Städler , Y. Yan , E. C. Nice , J. K. Heath , F. Albericio , F. Caruso , Adv. Funct. Mater. 2010 , 20 , 59 .

[ 26 ] G. Moad , E. Rizzardo , S. H. Thang , Aust. J. Chem. 2009 , 62 , 1402 . [ 27 ] A. N. Zelikin , J. F. Quinn , F. Caruso , Biomacromolecules 2006 , 7 , 27 . [ 28 ] A. N. Zelikin , Q. Li , F. Caruso , Chem. Mater. 2008 , 20 , 2655 .


Capsosomes with “Free-Floating” Liposomal Subcompartments

Documents

Transcript of Capsosomes with “Free-Floating” Liposomal Subcompartments