Hydrophobic Gold Nanoparticle Self-Assembly with

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Hydrophobic Gold Nanoparticle Self-Assembly with Phosphatidylcholine Lipid: MembraneLoaded and Janus Vesicles Michael R. Rasch,† Emma Rossinyol,‡ Jose L. Hueso,† Brian W. Goodfellow,† Jordi Arbiol,§ and Brian A. Korgel*,† †

Department of Chemical Engineering, Texas Materials Institute, and Center for Nano and Molecular Science and Technology, The University of Texas at Austin, Austin, Texas 78712-1062, ‡ Servei de Microscopia de la UAB, Universitat Autonoma de Barcelona, 08193, Bellaterra, Catalonia, Spain, and § Institucio Catalana de Recerca i Estudis Avanc¸ats (ICREA) and Institut de Ciencia de Materials de Barcelona, CSIC, Campus de la UAB, 08193, Bellaterra, Catalonia, Spain ABSTRACT Hybrids of hydrophobic sub-2-nm-diameter dodecanethiol-coated Au nanoparticles and phosphatidylcholine (PC) lipid vesicles made by extrusion were examined by cryogenic transmission electron microscopy (cryoTEM). The nanoparticles loaded the vesicles as a dense monolayer in the hydrophobic core of the lipid bilayer, without disrupting their structure. Nanoparticle-vesicle hybrids could also be made by a dialysis process, mixing preformed vesicles with detergent-stabilized nanoparticles, but this approach led to vesicles only partially loaded with nanoparticles that segregated into hemispherical domains, forming a Janus vesicle-nanoparticle hybrid structure. KEYWORDS Gold nanocrystal, nanoparticles, vesicles, liposomes, cryoTEM, self-assembly, Janus particle, detergent dialysis, extrusion, lipid assembly

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materials with the highest quality properties are obtained with hydrophobic capping ligands from synthetic reactions carried out in organic high boiling point solvents.40,41 Therefore, the ability to load hydrophobic nanoparticles into the lipid bilayer of vesicles provides a quite general route to vesicle-nanoparticle hybrids, making available a wide range of nanoparticle chemistry, including the prospect of loading different combinations of nanoparticles within each vesicle. There are many examples in the literature of hydrophobic nanoparticles, including Au,35,37,42 Ag,43,44 CdSe,3,36,38,39,45 Si,46 and Fe2O3,47,48 being dispersed in water with vesicleforming surfactants. The structures of the resulting lipidnanoparticle assemblies, however, are typically not well characterized, leaving fundamental questions about how hydrophobic nanoparticles really associate with liposomes and affect their stability. It is known that nanoparticles can disrupt vesicle formation under certain conditions; for example, hydrophobic nanoparticles larger than about 6 nm in diameter tend to induce lipid adsorption around the nanoparticles into micelle-like structures instead of vesicles.56,57 Lipid adsorption around smaller nanoparticles on the other hand, leads to too much curvature of the lipid membrane,49,56,58 and vesicles form instead of micelles, reportedly with hydrophobic nanoparticles embedded within the lipid bilayer.56,59,60 Molecular dynamics simulations have predicted that very small (∼1 nm diameter) hydrophobic fullerenes can disperse in a lipid bilayer without significant distortion of the lipid bilayer,53 but spherical nanoparticles of slightly larger size (i.e., 2-4 nm diameter) should distort the packing of the hydrophobic

ybrids of vesicles, or liposomes, and inorganic nanoparticles have interesting implications for medical imaging, drug delivery, and nanotoxicology.1 They should exhibit the well-established properties of vesicles, like the ability to encapsulate drugs for targeted delivery in the body,2 while imparting new functionality, such as imaging contrast for real-time tracking in vivo,3,4 or a therapeutic response to external electromagnetic stimulation by heating or cooling.5-8 Studies of vesicle-nanoparticle hybrids could also provide fundamental insight about how nanomaterials distribute in live cells and organisms, with vesicles serving as models of naturally occurring biological membranes.9,10 Vesicle-nanoparticle hybrids can be created using several strategies.1 For example, nanoparticles can be precipitated in-place within vesicles11-26 or they can be synthesized and then associated with vesicles, either by electrostatic adsorption27-33 or encapsulation34,35 of hydrophilic nanoparticles or by insertion of hydrophobic nanoparticles into the nonpolar interior of the lipid bilayer.3,36-39 Precipitation within the vesicles limits the nanoparticle chemistry to a relatively narrow range of materials because the synthesis must be carried out near room temperature in aqueous media.11-21,24,26 The reliance on hydrophilic nanoparticles is also limiting, as a much wider range of nanoparticle

* Corresponding author: [email protected]; telephone, (512) 471-5633; fax, (512) 471-7060. Received for review: 07/8/2010 Published on Web: 08/23/2010 © 2010 American Chemical Society

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FIGURE 1. Two methods used to form nanoparticle-vesicle hybrids: (top) lipids and nanoparticles were combined and coextruded through a 50 nm pore diameter filter; (bottom) vesicles were formed first by extrusion and then dialyzed in the presence of Au nanoparticles dispersed with detergent. The lipid bilayer is represented in green for the hydrophobic lipid tails and blue for the hydrophilic lipid headgroups; dodecanethiol-passivated Au nanoparticles are tan spheres, and octylglucoside is represented by the red hydrophilic headgroup connected to the purple hydrophobic octyl tail.

chains. This is different than alkanes or membrane-spanning proteins that can simply insert in a bilayer parallel to the lipid molecules;61-64 nanoparticles should force a molecular “unzipping”46,56 of the lipid bilayer that could also disrupt the integrity of the vesicles. Therefore, we sought to study a model system, with the help of cryogenic transmission electron microscopy (cryoTEM) imaging, of phosphatidylcholine (PC) vesicles with dodecanethiol-coated gold nanoparticles that are smaller than the lipid bilayer thickness, to understand in detail how hydrophobic nanoparticles tend to interact with liposomes. We found that vesicles are indeed stable with dodecanethiol-coated 2 nm diameter Au nanoparticles embedded in the lipid bilayers of vesicles. CryoTEM provides clear visualization of the vesicle-nanoparticle hybrids, with sufficient resolution to view individual nanoparticles and their location within the vesicles in their native environment.18,63,65-67 The images showed unexpectedly that the vesicle membranes are completely loaded with a dense monolayer of nanoparticles. Others have performed cryoTEM imaging of vesicles hybridized with hydrophobic CdSe and Fe2O3 nanoparticles;38,39,47 however, these reports have primarily shown vesicles associated with only a few welldispersed nanoparticles in the lipid bilayer. In our case of 2 nm diameter dodecanethiol-coated Au nanoparticles in PC vesicles, we observed only vesicles that were fully loaded with nanoparticles or completely free of nanoparticles. When liposomes were observed with only partial loading of nanoparticles, the nanoparticles tended to cluster and form islands in the bilayer. Dodecanethiol-coated Au nanoparticles with diameters between 1.6 and 1.8 nm were synthesized for the experiments. (See Supporting Information for Experimental Details and Nanoparticle Sizing.) The nanoparticles were associated with PC vesicles using either the extrusion or a detergent dialysis approach as illustrated in Figure 1. In one case, © 2010 American Chemical Society

vesicles were formed by coextrusion with the nanoparticles. In the other case, nanoparticles were dispersed with detergent and then combined with vesicles formed by extrusion and dialyzed to remove the detergentsas the detergent is removed, the nanoparticles associate with the vesicles. Both methods enabled loading of vesicles with the nanoparticles. (See Supporting Information for Experimental Details.) Figure 2 shows images of dispersions of PC vesicles prepared with and without nanoparticles. Samples A and B were prepared by extruding either pure lipid (A) or lipid with nanoparticles (B) through a membrane with 50 nm diameter pores. Both samples exhibit the characteristic opalescence of a vesicle dispersion, but sample B has a brown color from the Au nanoparticles. The fact that the hydrophobic nanoparticles remain dispersed in the aqueous media implies that the lipid is associated with the nanoparticles. When samples were prepared with lipid-nanoparticle ratios lower than 1500:1, there was observable sedimentation within a couple of days, though most of the nanoparticles remained dispersed by the lipid for weeks. CryoTEM was always performed no more than 1 day after preparing the samples. Vesicle-nanoparticle hybrids made with lipid-nanoparticle ratios higher than about 1500 were stable, without any sedimentation, for many weeks. CryoTEM of nanoparticle-vesicle hybrids made by lipid-nanoparticle coextrusion (i.e., the vial in Figure 2B) showed that the vesicles were predominantly unilamellar, intact, and densely loaded with nanoparticles in the lipid bilayer. The nanoparticles in the bilayer appear to form close-packed monolayers with an interparticle spacing close to the dodecanethiol layer thickness of about 1-2 nm. For comparison, Figure 3A shows cryoTEM images of vesicles made by extrusion in the absence of nanoparticles. The average diameter of the vesicles was similar for those prepared with and without nanoparticles (60 ( 10 nm) which is slightly larger than the 50 nm pores in the polycar3734

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FIGURE 2. Aqueous dispersions of (A, C) vesicles, (D) nanoparticles, and (B, E) nanoparticle-vesicle hybrids. The vesicles in (A) and (C) were prepared by extrusion without nanoparticles. In (B), the nanoparticle-vesicle hybrids were prepared by coextrusion using 1500 lipid molecules per nanoparticle. In (D), dodecanethiol-coated Au nanoparticles were dispersed in deionized (DI) H2O with OCG detergent. In (E), nanoparticle-vesicle hybrids were prepared by mixing samples (C) and (D) with 5000 lipid molecules per Au nanoparticle and then dialyzing. (F and G) Samples obtained after dialyzing OCG-stabilized nanoparticles with and without vesicles, respectively: (F1, G1) nanoparticle-vesicle solutions in dialysis tubing before dialysis; (F2, G2) solutions at the end of dialysis; (F3, G3) dialysis tubing after removing the solutions in (F2, G2).

bonate filter used for extrusion. The nanoparticles did not appear to influence the vesicle diameter. The vesicles prepared by coextrusion with nanoparticles either were loaded with nanoparticles or were free of nanoparticles. Panels C and G of Figure 3 show examples of this. Vesicles having only a few nanoparticles were not observed. A higher fraction of vesicles could be loaded with Au nanoparticles (Figures S4-S6) when the lipid-nanoparticle ratio was decreased, but empty vesicles were still observed, even when vesicles were extruded with only 100 lipid molecules per Au nanoparticle. It is possible that the apparent clustering of nanoparticles is due to incomplete mixing of the starting lipid-nanoparticle film; however, the lipid-nanoparticle dispersion was well-sonicated prior to extrusion (Figure S2 in Supporting Information). When high concentrations of nanoparticles were addedswith lower than 1500 lipid molecules per nanoparticlesworm-shaped agglomerates of Au nanoparticles were also observed (average length 100 ( 50 nm, average width 30 ( 10 nm), as shown in Figure 3F. In Figure 3G, the nanoparticle-vesicle hybrid population made with a lipid-nanoparticle ratio of 100sthe lowest molar ratio usedscontains a mixture of nanoparticle agglomerates, vesicles fully loaded with Au nanoparticles, and vesicles without any Au nanoparticles in the bilayer. Other vesicle structures were observed in some instances as well, like the bilamellar vesicles shown in Figure 3D and the two separate nanoparticle-loaded vesicles in Figure 3E that are enveloped in a single lipid bilayer. In the bilamellar vesicle in Figure 3D, the separation between the concentric rings of nanoparticles is about 3.9 nm, which is near the expected thickness of a PC bilayer. The nanoparticles do not bridge the concentric bilayers of the bilamellar vesicle, which © 2010 American Chemical Society

contrasts recent observations of much larger 8-14 nm diameter hydrophobic iron oxide nanoparticles spanning membranes when embedded in multilamellar vesicles.47 The enveloped nanoparticle shells in Figure 3E nearly touch but are separated by about 3.9 nm, which again corresponds to the thickness of a lipid bilayer or, equivalently, two lipid monolayers comprising the outer layer of each vesicle bilayer. Clearly, the hydrophobic nanoparticles are associating with the hydrophobic tails of the lipid and embedding within the bilayer. Vesicles could also be loaded with nanoparticles by a dialysis process. Initially, we attempted to form vesicles by dialyzing mixed micelle solutions of lipid, sodium cholate detergent, and nanoparticles, but the ionic detergent did not disperse the nanoparticles in aqueous solution. Nonionic octylglucoside (OCG) detergent disperses nanoparticles in aqueous solution, as demonstrated with other micelle-forming surfactants,68-71 but the vesicles formed by dialyzing OCG-PC mixed micelle solutions were very large (100-200 nm diameter) with broad size distribution.72-74 The most effective approach that we found was to make 50 nm diameter PC vesicles by extrusion75 (Figure 2C) and then combine them with gold nanoparticles dispersed in water with OCG detergent.76 Figure 2D shows a typical optically clear dispersion of dodecanethiol-coated gold nanoparticles using OCG detergent.77 Care must be taken to avoid excessive detergent-lipid ratios, which destroy the vesicles.81 The vesicle dispersions had PC concentrations of 8.6 mM, and OCG concentrations needed to be less than 25-30 mM.67,78,79 After membrane dialysis,80 a vesicle dispersion is obtained like the one shown in Figure 2E. If the detergent-stabilized nanoparticles are dialyzed in the absence of vesicles, the 3735

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FIGURE 3. CryoTEM images of vesicles prepared by extrusion in the (A) absence and (B-G) presence of Au nanoparticles. Imaging was performed 1 day after preparing the vesicles. The samples imaged in (B) and (C) were made with a lipid-nanoparticle ratio of 1500, and the samples imaged in (D-G) were prepared with a lipid-nanoparticle ratio of 100. The arrows in (C) highlight vesicles that are free of nanoparticles surrounding one nanoparticle-loaded vesicle. The vesicle in (D) is bilamellar with both lipid bilayers loaded with nanoparticles. In (E), two nanoparticle-loaded vesicles are encapsulated within a lipid bilayer, labeled by blue arrows. In (F), two Au-loaded vesicles are accompanied by a nanoparticle agglomerate. The field of vesicles in (G) shows a mixture of nanoparticle agglomerates, nanoparticle-loaded vesicles, and nanoparticle-free vesicles.

high OCG concentration,72-74 and increasing the number of nanoparticles with respect to OCG does not work because a minimum amount of detergent (per nanoparticle) is needed to disperse the nanoparticlessa ratio of OCG-nanoparticles of less than 10000 gave cloudy dispersions. Therefore, the total loading of vesicles with nanoparticles was more limited using the dialysis process than the coextrusion process. The observation of vesicles with lipid membranes fully loaded with hydrophobic nanoparticles is unprecedented, as far as we know. Hydrophobic molecules like dodecane generally have a low solubility in lipid bilayers due to the hydrated membrane interface;61 however, the relatively large dodecanethiol-coated Au nanoparticles appear to insert rather freely into the hydrocarbon core of the lipid bilayer. There is indeed a strong thermodynamic driving force for nanoparticle insertion into the bilayer, but this also creates a mechanical deformation in the bilayer. The free energy change ∆Gsolv, to place the hydrophobic sphere into the

nanoparticles aggregate and do not remain dispersed in water. The series of images in panels F and G of Figure 2 show the difference when the nanoparticles are dialyzed with and without the vesicles. Figure 4 shows cryoTEM images of the nanoparticlevesicle hybrids obtained after dialysis. When the nanoparticles are in focus in the image, as in Figure 4A, the lipid membrane of the vesicle is difficult to see, but when the vesicle is imaged slightly out of focus, as in Figure 4B, the lipid bilayer is clearly visible and intact. The vesicles are heavily loaded with nanoparticles, but only about half of the bilayer is filled with nanoparticles. The nanoparticles cluster into hemispherical nanoparticle-rich domains, forming Janus-type nanoparticle-vesicle hybrids. Attempts were made to increase the nanoparticle loading of the vesicles by the dialysis process. However, increasing the number of OCG-nanoparticle micelles added per vesicle eventually causes the vesicles to disassemble because of the © 2010 American Chemical Society

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FIGURE 4. Janus-type nanoparticle-vesicle hybrids formed by dialyzing vesicles and detergent-dispersed nanoparticles at a lipid-nanoparticle ratio of 5000. Imaging was performed 1 day after preparing the vesicles. (A, B) Images taken in focus and slightly out of focus, respectively, on the same region of the sample. (C) Schematic illustration of how the insertion of a hydrophobic nanoparticle into the lipid bilayer leads to a membrane deformation and an “unzipping” of the bilayer. In the situation where two nanoparticles are present in the bilayer, as in II, the total membrane curvature is reduced by lateral aggregation of the nanoparticles as shown in III. The strained regions of the lipid bilayer are circled in red in II.

hydrophobic membrane from pure water is πD2γ, where γ is the liquid-vapor surface tension of water (72.0 mN/m)82 and D is the nanoparticle diameter including the hydrophobic ligands.83 Fully extended dodecanethiol ligands should add about 1.8 nm to the particle radius. Therefore, with D ) 5.5 nm diameter, ∆Gsolv ) 1650kT, where k is Boltzmann’s constant. Wi et al. has calculated the energy penalty to deform a dioleoylphosphatidylcholine lipid bilayer ∆Gdef, when a hydrophobic sphere that is 5.5 nm in diameter is situated in the center of the bilayer: ∆Gdef ) 240kT.56 ∆Gsolv is nearly an order of magnitude greater than ∆Gdef. However, the membrane must “unzip” when the spherical nanoparticle is located at the center of the bilayer, as illustrated in Figure 4C. This unzipping creates void space around the nanoparticle within the bilayer (red circles, Figure 4C) that draws other nanoparticles together in the membrane. Clustering of the nanoparticles in the lipid bilayer reduces the total void space around the nanoparticles. The minimization of ∆Gdef may also explain why nanoparticles tend to insert into some vesicles-forming densely packed layers-and not into others. It also explains the formation of the Janus-type nanoparticle-vesicle hybrids, as opposed to vesicles with uniformly dispersed nanoparticles in the membrane.84 The cryoTEM images of the nanoparticle-vesicle hybrids prepared here provide new insight into how hydrophobic nanoparticles with size similar to the lipid bilayer thickness © 2010 American Chemical Society

interact with vesicles. The observation of dense monolayers of tightly packed nanoparticles in the lipid bilayers of vesicles is unprecedented; however, it makes sense when one considers the significant energy penalty that occurs when nanoparticles insert into the bilayer and deform the lipid membrane.56,85 The clustering of the nanoparticles, as in the rather extreme case of the Janus-like nanoparticle-vesicle hybrids, contrasts previous observations of CdSe and Fe2O3 nanoparticle-loaded vesicles,3,39,47 in which the nanoparticles were apparently uniformly dispersed in the bilayer. The observed coexistence of “bare” vesicles and vesicles fully loaded with nanoparticles was also unexpected. These results provide evidence that membrane-bound species, like small proteins and other molecules, might interact much more strongly than anticipated before.86 Further work is now needed to determine how the nanoparticle size, hydrophobic ligand layer thickness and chemistry, and lipid composition influence the structure of nanoparticle-vesicle hybrids. Acknowledgment. We thank Angela Holmberg and Josep M. Rebled for insightful discussions. We also thank the Robert A. Welch Foundation (F-1464), the National Science Foundation (DMR-0807065), and the National Institutes of Health (Grant No. R01 CA132032) for financial support. J.L.H. acknowledges financial support by a Fundacion Alfonso Martin Escudero Postdoctoral Fellowship, and J.A. 3737

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acknowledges financial support by the Spanish Government MICINN project Consolider Ingenio 2010 CSD2009 00013 IMAGINE and the CSIC project NEAMAN.

(33) Dietrich, C.; Angelova, M.; Pouligny, B. J. Phys. II 1997, 7, 1651– 1682. (34) Beaune, G.; Menager, C.; Cabuil, V. J. Phys. Chem. B 2008, 112, 7424–7429. (35) Paasonen, L.; Laaksonen, T.; Johans, C.; Yliperttula, M.; Kontturi, K.; Urth, A. J. Controlled Release 2007, 122, 86–93. (36) Feng, L. Y.; Kong, X. G.; Chao, K. F.; Sun, Y. J.; Zeng, Q. H.; Zhang, Y. L. Mater. Chem. Phys. 2005, 93, 310–313. (37) Binder, W. H.; Sachsenhofer, R.; Farnik, D.; Blaas, D. Phys. Chem. Chem. Phys. 2007, 9, 6435–6441. (38) Gopalakrishnan, G.; Danelon, C.; Izewska, P.; Prummer, M.; Bolinger, P. Y.; Geissbuhler, I.; Demurtas, D.; Dubochet, J.; Vogel, H. Angew. Chem., Int. Ed. 2006, 45, 5478–5483. (39) Mueller, W.; Koynov, K.; Fischer, K.; Hartmann, S.; Pierrat, S.; Basche, T.; Maskos, M. Macromolecules 2009, 42, 357–361. (40) Van Zanten, J. H.; Monbouquette, H. G. J. Colloid Interface Sci. 1994, 165, 512–518. (41) Huang, C.; Mason, J. T. Proc. Natl. Acad. Sci. U.S.A. 1978, 75, 308– 310. (42) Park, S. H.; Oh, S. G.; Mun, J. Y.; Han, S. S. Colloids Surf., B 2006, 48, 112–118. (43) Park, S. H.; Oh, S. G.; Suh, K. D.; Han, S. H.; Chung, D. J.; Mun, J. Y.; Han, S. S.; Kim, J. W. Colloids Surf., B 2009, 70, 108–113. (44) Bothun, G. D. J. Nanobiotechnol. 2008, 6, 13. (45) Bothun, G. D.; Rabideau, A. E.; Stoner, M. A. J. Phys. Chem. B 2009, 113, 7725–7728. (46) Jang, H.; Pell, L. E.; Korgel, B. A.; English, D. S. J. Photochem. Photobiol., A 2003, 158, 111–117. (47) Krack, M.; Hohenberg, H.; Kornowski, A.; Lindner, P.; Weller, H.; Forster, S. J. Am. Chem. Soc. 2008, 130, 7315–7320. (48) Zhang, X. W.; Yang, Y. F.; Tian, J.; Zhao, H. Y. Chem. Commun. 2009, 3807–3809. (49) Deserno, M.; Gelbart, W. M. J. Phys. Chem. B 2002, 106, 5543– 5552. (50) Li, Y.; Gu, N. J. Phys. Chem. B 2010, 114, 2749–2754. (51) Lipowsky, R.; Dobereiner, H. G. Europhys. Lett. 1998, 43, 219– 225. (52) Li, Y.; Chen, X.; Gu, N. J. Phys. Chem. B 2008, 112, 16647–16653. (53) Wong-Ekkabut, J.; Baoukina, S.; Triampo, W.; Tang, I. M.; Tieleman, D. P.; Monticelli, L. Nat. Nanotechnol. 2008, 3, 363–368. (54) Le Bihan, O.; Bonnafous, P.; Marak, L.; Bickel, T.; Trepout, S.; Mornet, S.; De Haas, F.; Talbot, H.; Taveau, J. C.; Lambert, O. J. Struct. Biol. 2009, 168, 419–425. (55) Roiter, Y.; Ornatska, M.; Rammohan, A. R.; Balakrishnan, J.; Heine, D. R.; Minko, S. Langmuir 2009, 25, 6287–6299. (56) Wi, H. S.; Lee, K.; Pak, H. K. J. Phys.: Condens. Matter 2008, 20, 494211. (57) Levin, C. S.; Kundu, J.; Janesko, B. G.; Scuseria, G. E.; Raphael, R. M.; Halas, N. J. J. Phys. Chem. B 2008, 112, 14168–14175. (58) Roiter, Y.; Ornatska, M.; Rammohan, A. R.; Balakrishnan, J.; Heine, D. R.; Minko, S. Nano Lett. 2008, 8, 941–944. (59) Ginzburg, V. V.; Balijepailli, S. Nano Lett. 2007, 7, 3716–3722. (60) Alexeev, A.; Uspal, W. E.; Balazs, A. C. ACS Nano 2008, 2, 1117– 1122. (61) Aagaard, T. H.; Kristensen, M. N.; Westh, P. Biophys. Chem. 2006, 119, 61–68. (62) McIntosh, T. J.; Simon, S. A.; MacDonald, R. C. Biochim. Biophys. Acta 1980, 597, 445–463. (63) Saxena, K.; Shipley, G. G. Biochemistry 1997, 36, 15940–15948. (64) Nelson, P. Biological Physics: Energy, Information, Life. W. H. Freeman: New York, 2004. (65) Jung, H. T.; Coldren, B.; Zasadzinski, J. A.; Iampietro, D. J.; Kaler, E. W. Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 1353–1357. (66) Johnsson, M.; Edwards, K. Biophys. J. 2003, 85, 3839–3847. (67) Vinson, P. K.; Talmon, Y.; Walter, A. Biophys. J. 1989, 56, 669– 681. (68) Dubertret, B.; Skourides, P.; Norris, D. J.; Noireaux, V.; Brivanlou, A. H.; Libchaber, A. Science 2002, 298, 1759–1762. (69) Fan, H. Y.; Leve, E. W.; Scullin, C.; Gabaldon, J.; Tallant, D.; Bunge, S.; Boyle, T.; Wilson, M. C.; Brinker, C. J. Nano Lett. 2005, 5, 645– 648. (70) Erogbogbo, F.; Yong, K. T.; Roy, I.; Xu, G. X.; Prasad, P. N.; Swihart, M. T. ACS Nano 2008, 2, 873–878.

Supporting Information Available. Detailed experimental procedures, Au nanoparticle characterization, SAXS model equations, low-resolution TEM imaging of the vesicles, and photographs of the dialysis results. This material is available free of charge via the Internet at http://pubs.acs.org. REFERENCES AND NOTES (1) (2) (3) (4) (5) (6) (7) (8) (9) (10) (11) (12) (13) (14) (15) (16) (17) (18) (19) (20) (21) (22) (23) (24) (25) (26) (27) (28) (29) (30) (31) (32)

Al-Jamal, W. T.; Kostarelos, K. Nanomedicine (London, U.K.) 2007, 2, 85–98. Torchilin, V. P. Nat. Rev. Drug Discovery 2005, 4, 145–160. Al-Jamal, W. T.; Al-Jamal, K. T.; Tian, B.; Lacerda, L.; Bomans, P. H.; Frederik, P. M.; Kostarelos, K. ACS Nano 2008, 2, 408–418. McCarthy, J. R.; Kelly, K. A.; Sun, E. Y.; Weissleder, R. Nanomedicine (London, U.K.) 2007, 2, 153–167. Duguet, E.; Vasseur, S.; Mornet, S.; Devoisselle, J. Nanomedicine (London, U.K.) 2006, 1, 157–168. Park, J.; Estrada, A.; Sharp, K.; Sang, K.; Schwartz, J. A.; Smith, D. K.; Coleman, C.; Payne, J. D.; Korgel, B. A.; Dunn, A. K.; Tunnell, J. W. Opt. Express 2008, 16, 1590–1599. Huang, X. H.; Jain, P. K.; El-Sayed, I. H.; El-Sayed, M. A. Lasers Med. Sci. 2008, 23, 217–228. Pankhurst, Q. A.; Thanh, N. K. T.; Jones, S. K.; Dobson, J. J. Phys. D: Appl. Phys. 2009, 42, 220301. Olbrich, K.; Rawicz, W.; Needham, D.; Evans, E. Biophys. J. 2000, 79, 321–327. Evans, E.; Heinrich, V.; Ludwig, F.; Rawicz, W. Biophys. J. 2003, 85, 2342–2350. Liu, H.; Graff, G. L.; Hyde, M.; Sarikaya, M.; Aksay, I. A. Mater. Res. Soc. Symp. Proc. 1991, 218, 115–121. Yaacob, I. I.; Bhandarkar, S.; Bose, A. J. Mater. Res. 1993, 8, 573– 577. Tricot, Y.; Fendler, J. H. J. Phys. Chem. 1986, 90, 3369–3374. Heywood, B. R.; Fendler, J. H.; Mann, S. J. Colloid Interface Sci. 1990, 138, 295–298. Hutchinson, J. L.; Mann, S.; Skarnulis, A. J.; Williams, R. J. P. J. Chem. Soc., Chem. Commun. 1980, 634–635. Mann, S. Nature 1993, 365, 499–505. Mann, S.; Hannington, J. P. J. Colloid Interface Sci. 1988, 122, 326– 335. Kennedy, M. T.; Korgel, B. A.; Monbouquette, H. G.; Zasadzinski, J. A. Chem. Mater. 1998, 10, 2116–2119. Korgel, B. A.; Monbouquette, H. G. Langmuir 2000, 16, 3588– 3594. Watzke, H.; Fendler, J. H. J. Phys. Chem. 1987, 91, 854–861. Korgel, B. A.; Monbouquette, H. G. J. Phys. Chem. 1996, 100, 346– 351. Song, Y. J.; Garcia, R. M.; Dorin, R. M.; Wang, H. R.; Qiu, Y.; Shelnutt, J. A. Angew. Chem., Int. Ed. 2006, 45, 8126–8130. Bota, A.; Varga, Z.; Goerigk, G. J. Appl. Crystallogr. 2007, 40, S259– S263. Mann, S.; Hannington, J. P.; Williams, R. J. P. Nature 1986, 324, 565–567. Yang, P.; Lipowsky, R.; Dimova, R. Small 2009, 5, 2033–2037. Chang, A.; Pfeiffer, W.; Guillaume, B.; Baral, S.; Fendler, J. H. J. Phys. Chem. 1990, 94, 4284–4289. Al-Jamal, W. T.; Al-Jamal, K. T.; Bomans, P. H.; Frederik, P. M.; Kostarelos, K. Small 2008, 4, 1406–1415. Wang, M. F.; Zhang, M.; Siegers, C.; Scholes, G. D.; Winnik, M. A. Langmuir 2009, 25, 13703–13711. Pornpattananangkul, D.; Olson, S.; Aryal, S.; Sartor, M.; Huang, C. M.; Vecchio, K.; Zhang, L. ACS Nano 2010, 4, 1935–1942. Zhang, L. F.; Granick, S. Nano Lett. 2006, 6, 694–698. Volodkin, D. V.; Skirtach, A. G.; Mohwald, H. Angew. Chem., Int. Ed. 2009, 48, 1807–1809. Mornet, S.; Lambert, O.; Duguet, E.; Brisson, A. Nano Lett. 2005, 5, 281–285. © 2010 American Chemical Society

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(71) Park, J. H.; von Maltzahn, G.; Ruoslahti, E.; Bhatia, S. N.; Sailor, M. J. Angew. Chem., Int. Ed. 2008, 47, 7284–7288. (72) Schwendener, R. A.; Asanger, M.; Weder, H. G. Biochem. Biophys. Res. Commun. 1981, 100, 1055–1062. (73) Mimms, L. T.; Zampighi, G.; Nozaki, Y.; Tanford, C.; Reynolds, J. A. Biochemistry 1981, 20, 833–840. (74) Paternostre, M. T.; Roux, M.; Rigaud, J. L. Biochemistry 1988, 27, 2668–2677. (75) Korgel, B. A.; van Zanten, J. H.; Monbouquette, H. G. Biophys. J. 1998, 74, 3264–3272. (76) On mixing the vesicles and Au-loaded micelles, the OCG and PC concentrations drop to 20 and 8.6 mM, respectively, while yielding 5000 lipid molecules per nanoparticle in solution . (77) The final OCG concentration in the aqueous phase was 66 mM, with 10000 OCG molecules per Au nanoparticle. (78) Jackson, M. L.; Schmidt, C. F.; Lichtenberg, D.; Litman, B. J.; Albert, A. D. Biochemistry 1982, 21, 4576–4582. (79) Ollivon, M.; Eidelman, O.; Blumenthal, R.; Walter, A. Biochemistry 1988, 27, 1695–1703. (80) The dialysis membrane consisted of regenerated cellulose dialysis tubing having a molecular weight cutoff of 6-8 kDa. On the basis of a technical note from Bang Laboratories (http://www.bangslabs. com/files/bangs/docs/pdf/203.pdf), this cutoff should correspond to a pore size of ∼2 nm. (81) When introducing detergent, the vesicle stability depends on the ratio of detergent to lipid molecules in solution at a particular lipid concentration. For a vesicle solution with 8.5 mM PC, the vesicles will rupture and disassemble if the total OCG concentration exceeds 25-30 mM as the OCG completely solubilizes the PC. If the OCG concentration is kept below ∼25 mM though, the vesicles do not rupture and the OCG simply partitions from the aqueous phase into the PC lipid bilayers.67,78,79 This detergent

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dilution requirement places a limit on the maximum number of Au nanoparticles that can be added per vesicle, since the initial detergent/nanoparticle mole ratio (∼104 OCG molecules/nanoparticle) is kept constant in these experiments. Ideally, both Au nanoparticles and detergent will partition into the lipid bilayers when mixed with the vesicles, similar to the reported PC vesicle embedding of OCG-solubilized lipoprotein receptor.63 In 90th ed.; CRC Handbook of Chemistry and Physics, 90th ed.; Lide, D. R., Ed.; CRC Press/Taylor and Francis: Boca Raton, FL, 2010. Chandler, D. Nature 2005, 437, 640–647. Israelachvili points out that the aggregation of membrane-spanning proteins in a lipid bilayer can be driven by minimization of the number of boundary lipidssdeformed lipids at the interface between the lipid bilayer and membrane proteins.87 However, the lipid deformation in this case has been attributed to formation of a bilayer meniscus at the lipid-protein interface, causing protein aggregation through lateral capillary forces.86 Because the Au nanoparticles are purely hydrophobic, and not amphiphilic like bilayer-spanning proteins, the nanoparticles are expected to stay completely immersed in the fluid bilayer without forming a meniscus with the bilayer surface. Lateral capillary forces are not expected to act on the nanoparticles in the lipid bilayer, rather the nanoparticles are expected to cluster together in the bilayer in order to minimize elastic deformation of the lipid membrane. Monticelli, L.; Salonen, E.; Ke, P. C.; Vattulainen, I. Soft Matter 2009, 5, 4433–4445. Kralchevsky, P. A.; Nagayama, K. Adv. Colloid Interface Sci. 2000, 85, 145–192. Israelachvili, J. N. Intermolecular and Surface Forces. Academic Press: San Diego, CA, 1992.

DOI: 10.1021/nl102387n | Nano Lett. 2010, 10, 3733-–3739