Controlled Loading of Building Blocks into Temporary Self-Assembled

Sep 10, 2008 - Ben Anderson,‡ Andrew Richter,‡ and Eugene Pinkhassik*,†. Institute for Nanomaterials DeVelopment and InnoVation and Department o...
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Langmuir 2008, 24, 11464-11473

Controlled Loading of Building Blocks into Temporary Self-Assembled Scaffolds for Directed Assembly of Organic Nanostructures L. Todd Banner,† Delia C. Danila,† Katie Sharpe,† Melissa Durkin,† Benjamin Clayton,† Ben Anderson,‡ Andrew Richter,‡ and Eugene Pinkhassik*,† Institute for Nanomaterials DeVelopment and InnoVation and Department of Chemistry, The UniVersity of Memphis, 213 Smith Chemistry Building, Memphis, Tennessee 38152-3550, and Department of Physics and Astronomy, Valparaiso UniVersity, Valparaiso, Indiana 46383 ReceiVed June 6, 2008. ReVised Manuscript ReceiVed August 11, 2008 Using temporary self-assembled scaffolds to preorganize building blocks is a potentially powerful method for the synthesis of organic nanostructures with programmed shapes. We examined the underlying phenomena governing the loading of hydrophobic monomers into lipid bilayer interior and demonstrated successful control of the amount and ratio of loaded monomers. When excess styrene derivatives or acrylates were added to the aqueous solution of unilamellar liposomes made from saturated phospholipids, most loading occurs within the first few hours. Dynamic light scattering and transmission electron microscopy revealed no evidence of aggregation caused by monomers. Bilayers appeared to have a certain capacity for accommodating monomers. The total volume of loaded monomers is independent of monomer structure. X-ray scattering showed the increase in bilayer thickness consistent with loading monomers into bilayer interior. Loading kinetics is inversely proportional to the hydrophobicity and size of monomers. Loading and extraction kinetic data suggest that crossing the polar heads region is the rate limiting step. Consideration of loading kinetics and multiple equilibria are important for achieving reproducible monomer loading. The total amount of monomers loaded into the bilayer can be controlled by the loading time or length of hydrophobic lipid tails. The ratio of loaded monomers can be varied by changing the ratio of monomers used for loading or by the time-controlled replacement of a preloaded monomer. Understanding and controlling the loading of monomers into bilayers contributes to the directed assembly of organic nanostructures.

Introduction Controlling the structure of materials in the (sub)nanometer scale is the key to achieving nonlinear performance improvement.1-3 A growing number of papers report nanoscale functional materials with superior properties. Practical methods for the synthesis of materials with well defined structure are increasingly important for further progress in the field. In the “traditional” self-assembly of nanomaterials,4 the inherent entropy loss accompanying aggregation of several molecules is offset by either large enthalpy gain due to multiple binding interactions, typically with hydrogen bonds,5 or by entropy gain associated with interactions with solvent molecules.6 Both methods impose stringent requirements on the structure of individual building blocks and limit options for precise structural control of the resulting materials. Furthermore, it may not always be beneficial for the final structure to retain moieties that enabled self-assembly. Directed assembly is an emerging paradigm for * To whom correspondence should be addressed. E-mail: epnkhssk@ memphis.edu. † University of Memphis. ‡ Valparaiso University. (1) Ozin, G. A.; Arsenault, A. C. Nanochemistry - A Chemical Approach to Nanomaterials; The Royal Society of Chemistry: Cambridge, UK, 2005; p 628. (2) Rao, C. N. R.; Muller, A.; Cheetham, A. K. The Chemistry of Nanomaterials: Synthesis, Properties and Applications; Wiley-VCH: Weinheim, 2004; Vol. 1, p 761. (3) Cao, G. Nanostructures and Nanomaterials: Synthesis, Properties & Applications; Imperial College Press: London, 2004; p 451. (4) Fendler, J. H. Chem. Mater. 1996, 8, 1616–1624. (5) Valdes, C.; Spitz, U. P.; Toledo, L. M.; Kubik, S. W.; Rebek, J. J. Am. Chem. Soc. 1996, 117, 12733–12745. (6) Messmore, B. W.; Hulvat, J. F.; Sone, E. D.; Stupp, S. I. J. Am. Chem. Soc. 2004, 126, 14452–14458. (7) Sanchez, C.; Boissiere, C.; Grosso, D.; Laberty, C.; Nicole, L. Chem. Mater. 2008, 20, 682–737. (8) Bae, C.; Yoo, H.; Kim, S.; Lee, K.; Kim, J.; Sung, M. M.; Shin, H. Chem. Mater. 2008, 20, 756–767.

efficient and scalable creation of nanostructures with well controlled shape and size. A few examples of templated materials include periodically organized mesoporous thin films,7 oxide nanotubes,8 and nanofibrous materials templated with organogelators.9 So far, a majority of reports have focused on inorganic materials. Fabrication of nanostructured organic materials is largely done by layer-by-layer (LBL) assembly and related methods.10,11 Other emerging approaches include bioinspired assembly such as functionalized viral particles.12 There is a clear need for exploring new methods for the synthesis of organic nanostructured materials to fully realize their potential. Our work focuses on materials fabrication within the welldefined environment of temporary self-assembling scaffolds. In this approach, a self-assembled scaffold is loaded with building blocks to arrange them into a desired shape before forming covalent bonds. The scaffolds can then be removed and reused for templating a new batch of nanostructures. Compared with “traditional” self-assembly, this method allows for a far greater flexibility in varying the structure of building blocks facilitating fine structural control and programming specific functional performance. This makes the method especially attractive since inexpensive materials can be used for nanostructures fabrication which alleviates the high costs of self(9) Llusar, M.; Sanchez, C. Chem. Mater. 2008, 20, 782–820. (10) Decher, G. Science 1997, 277, 1232–1237. (11) Quinn, J. F.; Johnston, A. P. R.; Such, G. K.; Zelikin, A. N.; Caruso, F. Chem. Soc. ReV. 2007, 36, 707–718. (12) Lugwig, C.; Wagner, R. Curr. Opin. Biotech. 2007, 18, 537–545. (13) Boerakker, M. J; Botterhuis, N. E.; Bomans, P. H. H.; Frederik, P. M.; Meijer, E. M.; Nolte, R. J. M.; Sonunerdijk, N. A. J. M. Chem. Eur. J. 2006, 12, 6071–6080. (14) Stendahl, J. C.; Rao, M. S.; Guler, M. O.; Stupp, S. I. AdV. Funct. Mater. 2006, 16, 499.

10.1021/la801755b CCC: $40.75  2008 American Chemical Society Published on Web 09/10/2008

Directed Assembly of Organic Nanostructures

assembling components. Recent advances in controlling the shape and size of noncovalent assemblies of amphiphiles promise excellent versatility for this approach.13,14 Recently, we used this approach to prepare subnanometer thin organic materials with programmed size nanopores from readily available components.15 We used templated polymerization in the interior of phospholipid bilayers to fabricate materials with 0.8 and 1.3 nm pores in high (>80%) yield from t-butyl styrene and divinylbenzene. This was achieved by localizing nonpolymerizable, molecular-sized pore forming templates (porogens) in the bilayer along with monomers before polymerization. Following polymerization, the bilayer scaffold and porogens were removed to yield a nanothin, nanoporous material. Precise permeability control was achieved by using different porogens. We expanded this method to control the chemical environment of nanopores.16 Previous studies have clearly demonstrated the feasibility of using self-assembled aggregates formed by amphiphilic molecules as templating media for the synthesis of nanostructures. Most of the studies used liposomes that have been thoroughly investigated in the past.17 Primarily, liposomes were used as a template for the synthesis of polymer capsules.18-24 Admicelles were also used as a two-dimensional solvent for the organization of monomers.25-29 Styrene was polymerized in the interior of wormlike micelles to form nanofibers.30 Although liposomes have received the most attention for use as a bilayer template the scope of the technique is potentially very broad because it may be adapted to other surfactant-based aggregate systems that have different three-dimensional architectures. These include surface bilayers,31,32 hybrid bilayers,33,34 supported bilayers,35,36 admicelles,25-29 spheres and rods,30 and bicelle disks.37 Biotechnology applications of bilayer-templated nanomaterials may include nanobioreactors38 as well as therapeutic and pharmaceutical

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delivery vehicles.39 Other applications include anticorrosion, solid lubrication, optical coatings, electrochemical sensors, and electronics.40 To date, no report systematically examined loading of monomers into the bilayer interior and methods to control the quantity and ratio of monomers. Previous papers also included some contradictory data. For example, Gomes et al. reported that adding monomers to the aqueous solution of liposomes did not lead to the formation of nanocapsules,24 while this method was used successfully by others.18,19 The ability to preprogram templated materials will only come from a thorough understanding of the parameters that control how monomers and other functional molecules are loaded into and organized within the templating scaffold. We believe it is timely to explore this method in detail due to its enormous potential in the synthesis of nanomaterials. In this work we report significant progress toward understanding the fundamental parameters that govern this system. We have developed and validated an analytical method for quantifying monomers loaded within a liposome bilayer. Using this method we have studied the loading kinetics of a variety of styrenes and methacrylates as a function of the template phospholipids’ acyl chain length and symmetry. We show that the saturated phospholipids have a certain intrinsic capacity for accommodating monomers independent of monomer structure. Furthermore, we present strategies for controlling loaded monomer composition so that functionality may be highly controlled. Although we chose unilamellar liposomes as the bilayer model to study, its results can likely be applied to other bilayer systems such as those mentioned above. The knowledge reported herein will provide a foundation by which organic nanostructures can be rationally designed and fabricated using amphiphilic bilayers as temporary self-assembled scaffolds.

Experimental Details (15) Danila, D. C.; Banner, L. T.; Karimova, E. J.; Tsurkan, L.; Wang, X.; Pinkhassik, E. Angew. Chem., Int. Ed. 2008, 47, 7036–7039. (16) Dergunov, S. A.; Pinkhassik, E. Angew. Chem., Int. Ed. 2008, DOI: 10.1002/anie.200803261. (17) Torchilin, V.; Weissig, V., Liposomes - A Practical Approach, 2nd ed.; Oxford University Press; Oxford, NY, 2003; Vol. 264, p 396. (18) Hotz, J.; Meier, W. Langmuir 1998, 14, 1031–1036. (19) Poulain, N.; Nakache, E.; Pina, A.; Levesque, G. J. Polym. Sci., Part A: Polym. Chem. 1996, 34, 729–737. (20) Nardin, C.; Hirt, T.; Leukel, J.; Meier, W. Langmuir 2000, 16, 1035– 1041. (21) Kurja, J.; Noelte, R. J. M.; Maxwell, I. A.; German, A. I. Polymer 1993, 34. (22) McKelvey, C. A.; Kaler, E. W.; Zasadzinski, J. A.; Coldren, B.; Jung, H. T. Langmuir 2000, 16, 8285–8290. (23) Jung, M.; Huber, D. H. W.; Bomans, P. H. H.; Frederic, P. M.; Meuldijk, J.; Herk, A. M. v.; Fischer, H.; German, A. I Langmuir 1997, 13, 6877–6880. (24) Gomes, J. F. P. d. S.; Sonnen, A. F.-P.; Kronenberger, A.; Fritz, J.; Coelho, M. A. N.; Fournier, D.; Noel, C. F.; Mauzac, M.; Winterhalter, M Langmuir 2006, 22, 7755–7759. (25) Wu, J.; Harwell, J. H.; O’Rear, E. A. Langmuir 1987, 3, 531–537. (26) Wu, J.; Harwell, J. H.; O’Rear, E. A. J. Phys. Chem. 1987, 91, 623–634. (27) Wu, J.; Harwell, J. H.; O’Rear, E. A. Am. Inst. Chem. Eng. J. 1988, 34, 1511–1518. (28) Sakhalkar, S. S.; Hirt, D. E. Langmuir 1995, 11, 3369–3373. (29) See, C. H.; O’Haver, J. J. Appl. Polym. Sci. 2003, 89, 36–46. (30) Becerra, F.; Soltero, J. F. A.; Puig, J. E.; Schulz, P. C.; Esquena, J.; Solans, C. Colloid Polym. Sci. 2003, 282, 103–109. (31) Tajima, K.; Gershfeld, N. L Biophys. J. 1985, 47, 203–209. (32) Gershfeld, N. L.; Tajima, K. Nature 1979, 279, 708–709. (33) Plant, A. L. Langmuir 1993, 9, 2764–2767. (34) Hubbard, J. B.; Silin, V.; Plant, A. L. Biophys. Chem. 1998, 75, 163–176. (35) Howland, M. C.; Szmodis, A. W.; Sanii, B.; Parikh, A. N. Biophys. Chem. 2007, 92, 1306–1317. (36) Sackmann, E. Science 1996, 271, 43–48. (37) Sanders, C. R.; Schwonek, J. P. Biochemistry 1992, 31, 8898–8905. (38) Graff, A.; Winterhalter, M.; Meier, W. Langmuir 2001, 17, 919–923. (39) Sukhorukov, G. B.; Rogach, A. L.; Zebli, B.; Liedl, T.; Skirtach, A. G.; Kohler, K.; Antipov, A. A.; Gaponik, N.; Susha, A. S.; Winterhalter, M.; Parak, W. J. Small 2005, 1, 194–200. (40) Broz, P.; Driamov, S.; Ziegler, J.; Ben-Haim, N.; Marsch, S.; Meier, W.; Hunziker, P. Nano Lett. 2006, 6, 2349–2353.

Chemicals. All solvents were HPLC grade and were not further purified before use. All phospholipids were purchased from Avanti Polar Lipids, Inc. as a chloroform solution and were not further purified (Chart 1): 1,2-didecanoyl-sn-glycero-3-phosphocholine (DDPC), 1,2-dilauroyl-sn-glycero-3-phosphocholine (DLPC), 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC), 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC), 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC), and 1-stearoyl-2-myristoyl-snglycero-3-phosphocholine (SMPC). The monomers used are shown in Chart 2, were purchased from Sigma-Aldrich, and were not further purified before use. Literature data for molecular square areas of phospholipids were used for calculations: DDPC, 0.670 nm2; DLPC, 0.640 nm2; DMPC, 0.617 nm2; DPPC, 0.486 nm2; DSPC, 0.516 nm2; SMPC, 0.486 nm2.41-45 Thin layer chromatography (TLC) was performed with silica G TLC plates with UV254 aluminum backed, 200 µm purchased from Sorbent Technologies. Flash chromatography was performed with an Analogix pump and Analogix Superflash silica columns purchased from Sorbent Technologies. Semipreparative HPLC (high-performance liquid chromatography) was performed with a Waters 600 pump and controller. A Waters 2487 detector was used equipped with a semiprep flow cell. The detection wavelength used was 255 nm. The column was a Nova-Pak Silica, 6 µm, 19 × 300 mm2. The flow rate was 10 mL/min. Elemental analyses were performed by Desert Analytics (Tucson, Arizona). (41) Lis, L. J.; McAlister, M.; Fuller, N.; Rand, R. P Biophys. J. 1982, 37, 657–666. (42) Joos, P.; Demel, R. A. Biochim. Biophys. Acta 1969, 183, 447–457. (43) Weschayanwiwat, P.; Scamehorn, J. F.; Reilly, P. J. J. Surfactants Deterg. 2005, 8, 65–72. (44) McIntosh, T. J.; Simon, S. A.; J.C; Ellington, J.; Porter, N. A. Biochemistry 1984, 23, 4038–4044. (45) Hui, S. W.; Mason, J. T.; Huang, C.-h. Biochemistry 1984, 23, 5570– 5577.

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Banner et al. Chart 1. Structures of Phospholipids Used

Synthesis of p-Propylstyrene and p-Hexylsytrene. 46 Aluminum chloride (15.0 g, 110 mmol) was added to methylene chloride (100 mL) under an argon atmosphere. The reaction flask was cooled in an ice bath. Acetic anhydride (5.6 g, 55 mmol) in methylene chloride (5 mL) was added dropwise over 30 min. The mixture was stirred for an additional 15 min. n-Alkylbenzene (25 mmol) in methylene chloride (5 mL) was as added to the reaction mixture over 30 min. The mixture was stirred for an additional 2 h while cooled in an ice bath. The mixture was slowly poured onto crushed ice (200 g). The organic phase was washed with 10% HCl (3 × 50 mL), saturated NaHCO3 (3 × 50 mL), and saturated NaCl (3 × 50 mL) in that order. The solvent was removed under vacuum at 30 °C to recover crude p-alkylacetophenone as an orange liquid. Crude p-alkylacetophenone (22 mmol) was diluted with methanol (60 mL) under argon atmosphere. The reaction flask was cooled in an ice bath. Sodium borohydride (0.83 g, 22 mmol) was added to the mixture over 5 min. The mixture was stirred for an additional 2 h. Saturated sodium chloride (70 mL) was added to the mixture and the product was extracted with methylene chloride (2 × 20 mL). The organic phase was washed with saturated sodium chloride (3 × 50 mL). The solvent was removed under vacuum at 40 °C to recover the crude (p-alkylphenyl)methylcarbinol as a colorless liquid. Crude (p-alkylphenyl)methylcarbinol (20 mmol) was diluted with toluene (200 mL) in a reaction flask under argon atmosphere. p-Toluenesulfonic acid (0.125 g, 0.7 mmol) was added. The reaction mixture was refluxed for 3 h in argon atmosphere. The mixture was washed with water (3 × 50 mL) and the solvent was removed under vaccum at 55 °C to give crude p-alkylstyrene. The crude product was purified by flash chromatography using 90% hexane/10% ethyl acetate (Rf ) 0.75 by TLC) and then semipreparative HPLC using (46) p-Propylstyrene and p-hexylstyrene were prepared using a synthetic method based upon the work of Quirk and Ok: Quirk, R. P.; Ok, M.-A. Macromolecules 2004, 37, 3976–3982.

99% hexane and 1% ethyl acetate. The solvent was removed under vacuum at 55 °C to give p-propylstyrene (33% overall yield from propylbenzene) or p-hexylstyrene (26% overall yield from hexylbenzene), both as colorless liquids. p-Propylstyrene. 1H NMR (270 MHz, CDCl3) δ 7.33 (d, 2 H, J ) 7.9 Hz), 7.13 (d, 2 H, J ) 7.9 Hz), 6.69 (dd, 1 H, J ) 16.2 Hz, J ) 10.8 Hz), 5.75-5.65 (d, 1 H, J ) 16.2 Hz), 5.22-5.14 (d, 1 H, J ) 10.8 Hz), 2.57 (t, 2 H, J ) 7.4 Hz), 1.71-1.56 (m, 2 H), 0.94 (t, 2 H, J ) 7.4 Hz); 13C NMR (270 MHz, CDCl3); 142.54, 136.85, 135.17, 128.70, 126.18, 112.85, 37.86, 24.57, and 13.87 ppm. Anal. Calcd for C11H14: C, 90.35; H, 9.65. Found: C, 90.50; H, 9.70. p-Hexylstyrene. 1H NMR (270 MHz, CDCl3) δ 7.32 (d, 2 H, J ) 7.9 Hz), 7.13 (d, 2 H, J ) 7.9 Hz), 6.69 (dd, 1 H, J ) 16.2 Hz, J ) 10.8 Hz), 5.74-5.65 (d, 1 H, J ) 16.2 Hz, J ) 10.8 Hz), 5.21-5.15 (m, 1 H), 2.58 (t, 2 H, J ) 7.7 Hz), 1.68-1.51 (m, 2 H), 1.40-1.21 (m, 6 H), 0.87 (t, 3 H, J ) 6.4 Hz); 13C NMR (270 MHz, CDCl3): 142.82, 136.85, 135.13, 128.63, 126.18, 112.83, 35.78, 31.80, 31.47, 29.0, 22.68, and 14.15 ppm. Anal. Calcd for C14H20: C, 89.29; H, 10.71. Found: C, 89.55; H, 10.51. Small Unilamellar Liposome Preparation. Unilamellar liposomes were prepared by the extrusion method.47 Ten milligrams (500 µL of 20 mg/mL in chloroform) of phospholipids was pipetted into a glass culture tube. The solvent was evaporated using a stream of argon gas to give a phospholipid film on the walls of the tube. Residual chloroform was removed from the film by placing the tube under high vacuum for at least 30 min. Deionized water (1000 µL) was then added to the tube and it was vortexed until no phospholipid film could be seen on the sides of the tube to produce a solution of multilamellar liposomes. This solution was then extruded through (47) MacDonanld, R. C.; MacDonald, R. I.; Menco, B. P.; Takeshita, K.; Subbarao, N. K.; Hu, L.R. Biochim. Biophys. Acta 1991, 1061, 297–303.

Directed Assembly of Organic Nanostructures Chart 2. Structures of Monomers Used

a 0.1 µm polycarbonate membrane using a Mini-Extruder (Avanti Polar Lipids). Liposome Loading. Using a glass syringe, 10 µL of each monomer was added into separate 4 mL glass vials containing 900 µL of unilamellar liposome solution and a 5 × 2 mm2 PTFE coated stir bar. All experiments were conducted at least in duplicates. Gas chromatography was later used to determine that 10 µL was enough monomer to exceed the loading capacity of the liposomes’ bilayers. Furthermore, residual oil monomer droplets were observed in the samples (with exception to styrene and butylmethacrylate which is discussed later). The vials were sealed with the cap (black phenolic closure with rubber liner) and were placed on a stir plate inside a refrigerator at 6 °C for 1, 4, 16, 24, and 96 h (other times are experiment specific and are detailed in the results section). The solution was gently stirred (60-90 rpm) to avoid emulsion formation. Dynamic Light Scattering (DLS). DLS measurements were performed on a Malvern Nano-ZS zetasizer (Malvern Instruments Ltd., Worcestershire, U.K.). Sample aliquots of 100 µL were diluted with 4 mL of water and placed in a disposable polystyrene cuvette. The measurements were done at room temperature. At least 11 scans were collected from each sample. Transmission Electron Microscopy (TEM). TEM images were acquired on a JEOL JEM1200EXII microscope. Samples were negatively stained with phosphotungstic acid (pH 5.9) on a carbon grid. Monomer Separation and Extraction. Using a plastic pipet, 400 µL of the loaded liposome solution was transferred to the top of a Pasteur pipet (9 in. clear borosilicate glass), stoppered on the bottom with folded parafilm to provide a microseparatory funnel. Care was taken to avoid the monomer oil droplet and only collect the aqueous liposome phase. The Pasteur pipet, containing the liposome solution was held in place for three minutes to allow any excess monomer to phase separate with the bottom/aqueous phase. The top of the Pasteur pipet was then stoppered with a finger and then raised from the Parafilm. About 200 µL of the bottom, aqueous liposome solution was released into a separate glass sample vial. A plastic pipet was then used to transfer 100 µL of this solution into 500 µL of 0.4 mg/mL toluene in hexane. A 5 × 2 mm2 PTFE coated stir bar was added and the vial was capped. The vial was placed on

Langmuir, Vol. 24, No. 20, 2008 11467 a stir plate inside a refrigerator at 6 °C and gently stirred for the specified time. Following extraction the organic phase was analyzed by gas chromatography. Gas Chromatography Analyses. Gas chromatography analyses were performed with a Hewlett-Packard 5890 Series II Gas Chromatograph with a flame ionization detector (FID). The column was a HP-1 Cross-linked Methyl Silicone Gum, 12 m × 0.2 mm × 0.33 µm film thickness. Helium was the carrier gas and the detector gases were air and hydrogen. Ten µL of sample (or standard) was injected into the split-mode injector. The inlet temperature was 150 °C and the detector temperature was 250 °C throughout the analysis. The initial temperature of the oven was 90 °C and after 1.5 min the temperature was ramped to 250 °C at a rate of 50 °C/min. Data acquisition was performed with a Peak Simple Chromatography Data System and Peak Simple software. Analytical standards for gas chromatography were prepared using serial dilution. A stock solution was prepared by adding 10 µL of monomer to 900 µL hexane. Working standards were prepared by adding 100 µL of the stock solution to 500 µL of 0.4 mg/mL toluene in hexane. Monomer and toluene peaks were integrated to obtain peak areas. Toluene was used as the internal standard. Combined Monomer Loading. Monomer loading was carried out in duplicate as described above for 24 h. The exception was that two monomers were added to unilamellar DLPC liposomes. Combined loading was performed using the following monomer volumes (mole ratio). Ten µL styrene/10 µL DVB (1.2/1.0), 10 µL hexylstyrene/10 µL DVB (0.7/1.0), 20 µL styrene/5 µL DVB (5.0/ 1.0), 20 µL hexylstyrene/5 µL DVB (2.6/1.0), 10 µL butylmethacrylate/10 µL EGDM (1.2/1.0). The separation, extraction, and gas chromatography analysis procedures were then performed as described above. Replacement Monomer Loading. Monomer loading of DVB was carried out as described above for 24 h with four unilamellar DLPC liposome samples. The separation procedure was then performed on the samples as described above. The four separated loaded liposome samples were combined into two separate samples. The extraction and gas chromatography analysis procedures were then performed using aliquots of the combined samples. Monomer loading of p-propylstyrene was then carried out for 24 h with 900 µL of the remaining aliquots. The extraction and gas chromatography analysis procedures were then performed as described above. This process was repeated except p-propylstyrene was loaded for 5 h. Small Angle X-Ray Scattering (SAXS). The sample cell consisted of a 10 mm thick Teflon ring with inlet and outlet valves, sandwiched between Kapton-lined beryllium windows, held in place with screws, such that it is airtight. The total volume of the sample cell was about 10 mL, with a path length along the direction of the beam of 10 mm. The positioning of the sample cell is precise, such that successive scans with different samples have the same background. After each sample was loaded into the cell, it was mounted and scanned. X-ray scans were taken of water, solutions of nonloaded DLPC liposomes, and solutions of DLPC liposomes loaded with DVB monomers. The X-ray data was taken at Sector 9, ID-C, at the Advanced Photon Source, Argonne National Laboratory. The X-ray energy was chosen to be fairly high, 22.8 keV in order to reduce absorption within the liquid, thereby also reducing background scatter. The beam measured 1 mm wide by 0.05 mm tall, with the detector slit set large enough to capture the entire beam (1 × 0.4 mm2). Data were taken using a NaI point detector, positioned 1 m from the sample, scanning from q ) 0.05-0.70 Å-1, where q ) (4π sin θ)/λ, θ is the angle between the detector direction and the incident beam direction, and λ is the X-ray wavelength, 0.544 Å. Scans were repeated twice to check for reproducibility. Since the repeated scans were identical, they were combined to improve the statistical uncertainty. The water background was subtracted from the liposome scans, yielding nearly zero intensity away from the observed features, indicating that the background was reproducible from sample to sample.

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Results and Discussion Measurement of Loading. We developed and validated an analytical method to quantify the amount of monomers localized within the bilayers. Typically, an excess of monomers was added to 1 mL of aqueous unilamellar liposome solution, and the mixture was gently stirred for 24 h. To determine the amount of monomers in the bilayer, aqueous solution was first separated from remaining free monomers. We found that the separation within a Pasteur pipet works best for 1 mL volume of aqueous solution. Next, monomers were extracted from the liposome bilayer with hexane containing 0.4 mg/mL toluene for 24 h to allow for complete extraction. The extract was analyzed by gas chromatography using toluene as an internal standard. Blank control experiments were performed as described above in the absence of liposomes. Amounts of extracted monomers correlated well with published values for aqueous solubility of styrene48 and ethylene glycol dibutyrate,49 which is structurally similar to EGDM. In all cases except EGDM, the total amount of monomers extracted from water was at least 30 times lower than the amount of monomers extracted from liposome solution. This shows that the monomers are predominantly associated with the liposomes. With possible exception of EGDM, which was determined to have relatively high water solubility (5.5 mM), we conclude that the method is practical for nanostructure synthesis since polymerization in bulk water is negligible compared with polymerization in the bilayer. Kinetics. Time-resolved loading of different monomers, derivatives of styrene and various methacrylates, into DLPC liposomes revealed that the quantity of monomers localized within the bilayer is a logarithmic function of loading time (Figure 1; individual loading curves for each monomer are shown in Supporting Information). Maximum loading values were observed in the curves, which indicate that the bilayer has a maximum capacity for accommodating monomers (1). In control experiments, styrene reached the maximum concentration in water within the first 15 min of exposure suggesting that dissolving monomers in water is not the rate limiting step. Since the aqueous concentration of monomers rapidly reaches the saturation limit and remains constant during the loading period, the logarithmic shape of the loading curves is likely due to conformational changes in the bilayer to accommodate monomers, with the most energetically favorable occurring first. It is likely that the kinetics of loading has at least two mutually dependent components: crossing the polar heads region and rearranging the bilayer to accommodate monomers within the hydrophobic region. As monomers are loaded into the bilayer, it becomes increasingly harder to accommodate more monomers until the saturation limit is reached. Larger molecules exhibited slower loading kinetics (Figure 2), likely due to steric factors. Also, more hydrophobic and less water-soluble molecules (as estimated by log P, where P is the partition coefficient between 1-octanol/water) exhibit slower loading (Figure 3) suggesting that crossing the polar heads region may be the rate limiting step. The data can be used to make preliminary estimates about loading kinetics of other molecules. Excluding molecules that may not load due to large size, those with a log P below 3.0 will load very rapidly. In contrast, molecules with log P values above 5.0 will take several days to reach maximum capacity, which may be impractical for nanomaterials synthesis. In fact, p-hexylstyrene loading required at least 10 days to achieve the maximum capacity (Figure 1). (48) Ley, G. J. M.; Hummel, D. O.; Schneider, C. AdV. Chem. Ser. 1967, 66, 184–202. (49) Funasaki, N.; Hada, S.; Kawamura, K. Nippon Kagaku Kaishi 1976, 12, 1944–1946.

Figure 1. Loading kinetics of styrenes and methacrylates in DLPC. Each data point is an average of three measurements. Error bars are omitted for clarity.

Figure 2. Plot of initial rate (0-1 h) of monomer loading vs molecular volume. Molecular volume was calculated from the bulk density and molecular weight. Larger molecules exhibit slower loading rates.

Extraction of monomers from the bilayer into hexane followed kinetics similar to loading. Extraction became slower as the monomer became more hydrophobic. For all monomers in all phospholipids, extraction appears nearly complete after 24 h. Figure 4 provides representative curves of the extraction kinetics for some styrenes. Experimental conditions are especially important for loading of lower-boiling monomers (bp < 170 °C) such as styrene and butylmethacrylate. Evaporation of such monomers from the water surface and their condensation on the vial walls in the headspace may compete with their loading into the bilayers. We found that in a typical setup, exposure of 10 µL of styrene to 1 mL of liposome solution (20 mg of DLPC) for 24 h resulted in 90% of the loading occurs within 4-24 h. Both loading into the bilayer and extraction from the bilayer occur on a comparable time scale. Consideration of loading kinetics and multiple equilibria are important for selecting reproducible experimental conditions.

Directed Assembly of Organic Nanostructures

In the case of lipids with saturated acyl chains, bilayers fill to a certain capacity measured to be (1.9 ( 0.2) × 10-20 L for a 100 nm unilamellar liposome, which is virtually independent of monomer structure. The bilayer capacity is greatly affected by the aggregation state of lipids. Above the phase transition temperature, when bilayers are at the fluid liquid crystalline phase, bilayers accommodate 5-10 times more monomers than in the gel phase below the phase transition. Loading bilayers with monomers greatly reduces the phase transition temperature (by at least 11 °C for DPPC), which is consistent with previously reported data on solubilization of small organic molecules in the bilayer interior.52 Exposing liposomes made from saturated lipids to various styrene derivatives and acrylates does not lead to aggregation as evidenced by DLS and TEM data. X-ray scattering showed the increase of bilayer thickness by 0.4 nm consistent with loading monomers into the bilayer interior. The total amount of monomers loaded into the bilayer can be varied by changing the length of acyl chains, using asymmetric

Langmuir, Vol. 24, No. 20, 2008 11473

lipids, or by varying the loading time. The ratio of different monomers can be controlled by either varying the ratio of monomer mixture used for loading or by partial replacement of a preloaded monomer. These findings offer a broad set of methods for controlling the placement of hydrophobic building blocks in the interior of temporary self-assembled scaffolds. Considering a wide range of self-assembled structures formed from amphiphilic molecules, we anticipate that this templated assembly method will be highly versatile. Acknowledgment. This work was supported by the National Science Foundation CAREER award (CHE-0349315) and the FedEx Institute of Technology Innovation Award. Supporting Information Available: Additional information regarding the synthesis and characterization of compounds, analytical method development and validation is provided along with individual loading and extraction curves. This material is available free of charge via the Internet at http://pubs.acs.org. LA801755B