Facile Directed Assembly of Hollow Polymer Nanocapsules within

Dec 16, 2013 - Department of Physics and Astronomy, Valparaiso University, Valparaiso, ... neously formed surfactant vesicles in the synthesis of poly...
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Facile Directed Assembly of Hollow Polymer Nanocapsules within Spontaneously Formed Catanionic Surfactant Vesicles Mariya D. Kim,† Sergey A. Dergunov,† Andrew G. Richter,‡ Jeffrey Durbin,† Sergey N. Shmakov,† Ying Jia,† Saltanat Kenbeilova,§ Yerbolat Orazbekuly,§ Aigerim Kengpeiil,§ Ernö Lindner,∥ Sai Venkatesh Pingali,⊥ Volker S. Urban,⊥ Steven Weigand,# and Eugene Pinkhassik*,† †

Department of Chemistry, Saint Louis University, 3501 Laclede Avenue, St. Louis, Missouri 63103, United States Department of Physics and Astronomy, Valparaiso University, Valparaiso, Indiana 46383, United States § Department of Chemical Technology of Processing of Petroleum, Gas, and Polymers, Kazakh National Technical University, 22 Satpaev St., Almaty 050013, Kazakhstan ∥ Department of Biomedical Engineering, University of Memphis, Memphis, Tennessee 38152, United States ⊥ Center for Structural Molecular Biology, Oak Ridge National Laboratory, P.O. Box 2008 MS-6430, Oak Ridge, Tennessee 37831-6430, United States # DND-CAT, Advanced Photon Source, ANL Bldg. 432, 9700 S. Cass Ave., Argonne, Illinois 60439, United States ‡

ABSTRACT: Surfactant vesicles containing monomers in the interior of the bilayer were used to template hollow polymer nanocapsules. This study investigated the formation of surfactant/monomer assemblies by two loading methods, concurrent loading and diffusion loading. The assembly process and the resulting aggregates were investigated with dynamic light scattering, small angle neutron scattering, and small-angle X-ray scattering. Acrylic monomers formed vesicles with a mixture of cationic and anionic surfactants in a broad range of surfactant ratios. Regions with predominant formation of vesicles were broader for compositions containing acrylic monomers compared with blank surfactants. This observation supports the stabilization of the vesicular structure by acrylic monomers. Diffusion loading produced monomer-loaded vesicles unless vesicles were composed from surfactants at the ratios close to the boundary of a vesicular phase region on a phase diagram. Both concurrent-loaded and diffusion-loaded surfactant/monomer vesicles produced hollow polymer nanocapsules upon the polymerization of monomers in the bilayer followed by removal of surfactant scaffolds.

1. INTRODUCTION Using self-assembled scaffolds to template the synthesis of organic nanostructures is an attractive way to create materials with new shapes and properties. In this directed assembly approach, building blocks are loaded into a supramolecular scaffold and are then linked by covalent bonds. The scaffold can be removed and reused. The shape and size of the selfassembled scaffold determines the shape and size of nanomaterials. In contrast with self-assembled materials, use of scaffolds permits creating nanostructures from common building blocks, such as simple monomers, that would not assemble into a welldefined structure in the absence of the templating scaffold. Templating is common in the creation of inorganic materials, for example, biomineralization.1 Until recently, few papers reported the synthesis of organic nanomaterials using this method.2−7 In the recent years, this general approach led to the creation of hollow nanocapsules,8−12 nanorattles,13 nanodisks,14,15 nanotubes, and nanorods.16−18 Better understanding of interactions between scaffolds and building blocks is important for programming properties of organic nanomaterials.19−21 Nanocapsules are particularly interesting nanostructures due to diverse potential applications. They can serve as vehicles for © 2013 American Chemical Society

delivery of drugs and imaging contrasts or as semipermeable containers for creating nanoreactors or nanosensors. Recently, encapsulation of indicator dyes addressed the long-standing challenge of combining the fast response times and long-term stability of optical chemosensors.22 Vesicle-templated nanocapsules showed good control of permeability, ultrafast transport rates, and excellent long-term stability.9,10,23,24 Because of the high potential utility, a simple and scalable method for the synthesis of nanocapsules from inexpensive materials is likely to have a broad impact. Spontaneously formed vesicles are attractive as temporary scaffolds. Previous reports showed that different cationic and anionic surfactants spontaneously formed vesicles when mixed in certain ratios.25−29 Surfactant vesicles were used to template polymer nanocapsules.7,30 In these experiments, monomers were added to the aqueous solution of vesicles and allowed to diffuse into the interior of bilayers. The most pressing question in using surfactant templates is whether loaded vesicles can be Received: October 21, 2013 Revised: December 6, 2013 Published: December 16, 2013 7061

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polyester Nucleopore membrane (Sterlytech) with 0.2 μm pore size using a Lipex stainless steel extruder (Northern Lipids). Diffusion Loading of Monomers. Stock solutions of cationic and anionic surfactant were prepared by hydrating respective surfactant in deionized water. Stock solution of each surfactant was incubated at 40 °C for 1 h. Samples were prepared by mixing the stock solutions at the desired ratio at room temperature, and compositions reported are on a weight percent basis. After brief vortex mixing, the solution was extruded five times at 25 °C through a track-etched polyester Nucleopore membrane (Sterlytech) with 0.2 μm pore size using a Lipex stainless steel extruder (Northern Lipids). Five milliliters of vesicles was transferred into a 20 mL glass vial containing a 5 × 2 mm PTFE coated stir bar. Vials were placed in a water bath at 30 °C. The solution of vesicles was allowed approximately 30 min to equilibrate with its temperature-controlled environment. After temperature equilibration, 500 μL of DVB was added in each vial. The vial was sealed with the cap (polypropylene with pulp-backed metal foil liner). All experiments were shielded from light during monomer loading, except for taking aliquots for analysis. Excess DVB was clearly visible floating atop the aqueous vesicle solution throughout every experiment. Each sample was stirred gently (40−60 rpm) to prevent the formation of a monomer emulsion in the surfactant solution. Temperature was checked every hour throughout the procedure. Synthesis of Nanocapsules. The sample was irradiated for 1.5 h with UV light (λ = 254 nm) in a photochemical reactor (10 lamps, 32 W each; the distance between the lamps and the sample was 10 cm) using a quartz tube with path length of light of approximately 3 mm. Short path length is important for efficient polymerization in the presence of dyes. Following the polymerization, a solution of NaCl (0.1 mL of 3N) in methanol (10 mL) was added to the reaction mixture to precipitate the nanocapsules. The nanocapsules were separated from the reaction mixture and purified in repeated centrifugation and resuspension steps using methanol (3−5 drops of NaCl (0.1 mL of 3 N) were added for better precipitation), water− methanol mixture, and water as washing solutions. Dynamic Light Scattering (DLS). Hydrodynamic diameter and polydispersity index (PDI) measurements were performed on a Malvern Zetasizer Nano ZS (Malvern Instruments Ltd., Worcestershire, U.K.). The helium−neon laser, 4 mW, operated at 633 nm, with the scatter angle fixed at 173° and the temperature at 25 °C. Samples of 80 μL were placed into disposable cuvettes without dilution (8.5 mm center height BRANDmicro UV cuvette). Each data point was an average of 16 scans. High Performance Liquid Chromatography (HPLC). An aliquote of 50 μL of sample was carefully taken from the bottom of the vial to avoid contamination with excess monomer and mixed with 950 μL of methanol to lyze liposomes. Then 20 μL of the resulting mixture sample was injected into a split-mode injector. Analytical HPLC was performed with a Waters 600 pump and Waters 2487 dual wavelength UV−vis detector. The detection wavelengths used were 250 and 270 nm. The column was a Nova-Pak C18, 3.9 mm diameter × 150 mm length. HPLC grade methanol was used as the mobile phase. The flow rate was 2 mL/min. Samples and concentration standards were run at least five times for each measurement, and the data were averaged. When necessary, the experimental samples were diluted further so that the signals would fall within a range of concentrations where the detector response was highly reproducible and the relationship between area and concentration could be modeled with a simple polynomial. Standard samples of known concentrations of DVB were prepared by serial dilution of a stock DVB solution in methanol and were analyzed by HPLC as described above. A calibration curve was produced by fitting HPLC data from DVB solutions in methanol at different concentrations via the least-squares fitting method, and was used to interpolate between the points. TEM and SEM Studies. Electron microscopy images were obtained with a FEI Inspect F50 STEM scanning electron microscope (Hillsboro, OR) at a working voltage of 30 kV. To prepare the sample for transmission electron microscopy (TEM) analysis, a drop of sample was carefully placed on a 200-mesh carbon grid and excess sample was wiped away with filter paper. Then a drop of 2% uranyl

formed spontaneously upon mixing of surfactants and monomers. Recently, we reported synergistic self-assembly of surfactants and monomers resulting in vesicles containing monomers in the bilayer.31 Monomer-loaded vesicles produced hollow polymer nanocapsules upon the polymerization of monomers and removal of surfactant scaffold. This study prompted us to investigate the surfactant/monomer assemblies in detail. In this work, we investigated the application of spontaneously formed surfactant vesicles in the synthesis of polymer nanocapsules (Figure 1). We examined two methods for

Figure 1. Schematic representation of the synthesis of polymer nanocapsules. (a) Spontaneous formation of loaded bilayer from a mixture of surfactants and monomers (concurrent loading) followed by the polymerization and scaffold removal. (b) Diffusion loading of the preformed bilayer followed by the polymerization and scaffold removal.

placing monomers into the bilayer, concurrent loading (loading monomers simultaneously with the formation of the bilayer) and diffusion loading (loading of monomers into preformed bilayer).

2. EXPERIMENTAL SECTION 2.1. Chemicals. Hexadecyltrimethylammonium p-toluenesulfonate (CTAT), hexadecyltrimethylammonium bromide (CTAB), sodium dodecylbenzenesulfonate (SDBS), sodium dodecylsulfate (SDS), tertbutyl methacrylate (t-BMA), butyl methacrylate (BMA), and tert-butyl styrene (TBS), used as monomers, and ethylene glycol dimethacrylate (EGDMA) and divinylbenzene (DVB), used as a cross-linking agents, were purchased from Sigma-Aldrich and were passed through an aluminum oxide column to remove the inhibitor shortly before sample preparation. The photoinitiator 2,2-dimethoxy-2-phenyl-acetophenone (DPA), purchased from Sigma-Aldrich, was used without any additional purification. The solvents and other chemicals used in this study were HPLC and ACS reagent grade, respectively and were used as received. 2.2. Methods. Concurrent Loading of Monomers into Surfactant Vesicles. Stock solutions of cationic surfactant with monomers were prepared by mixing 100 mg of CTAT with t-BMA (32 μL, 0.193 mmol), BMA (32 μL, 0.199 mmol), EGDMA (32 μL, 0.166 mmol), and initiator 2,2-dimethoxy-2-phenyl-acetophenone (1 mg, 0.003 mmol) in a test tube and hydrating the mixture in 4 mL of deionized water. To prepare a stock solution of anionic surfactant, 100 mg of SDBS was mixed with t-BMA (32 μL, 0.193 mmol), BMA (32 μL, 0.199 mmol), EGDMA (32 μL, 0.166 mmol), and initiator 2,2dimethoxy-2-phenyl-acetophenone (2 mg, 0.01 mmol) in a test tube and hydrating the mixture in 10 mL of deionized water. Each stock solution was equilibrated at 40 °C during 30 min. Samples were prepared by mixing the stock solutions at proper ratio followed by additional incubation at 25 °C for 1 h. After brief vortex mixing, the solution was extruded five times at 25 °C through a track-etched 7062

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acetate was added to the grid to negatively stain the sample. After 2 min, the excess liquid was wiped off. To prepare the sample for scanning electron microscopy (SEM) analysis, a drop of sample was placed on SEM pin stub specimen mount covered with double coated carbon conductive tabs and dried under vacuum. The studied samples were coated with a 5 nm gold layer using EMS 590 X sputter (Hatfield, PA). Small Angle X-ray Scattering (SAXS). SAXS was performed at 5-ID at the Advanced Photon Source, Argonne National Laboratory. The energy was set to 17 keV, and the detector distance was 1.5 m. Samples were flowed through 1 mm diameter quartz capillaries at 4 mL/s for total exposure times of 50 s, collected as five 10 s scans with an area detector. Using the data reduction software at Sector 5, these scans were corrected for gain and dark current, azimuthally averaged, and combined into a single absolute intensity vs Q data set ranging from 0.0175 to 0.73 Å−1. X-ray damage was observed to occur when not flowing the samples, but was not evident when flowing. Small-Angle Neutron Scattering (SANS). SANS measurements were performed with the CG-3 Bio-SANS instrument32 at the High Flux Isotope Reactor (HFIR) facility of Oak Ridge National Laboratory and at the NC-7 SANS instrument at the National Institute of Standards and Technology’s Center for Neutron Research (NCNR).33 Quartz cells of 1 mm thickness were used to hold the liquid samples. At both facilities, we collected data with three different detector-distance/neutron-wavelength configurations. At NCNR, we used 1 m/6 Å, 4 m/6 Å, and 15.3 m/8 Å configurations, giving a Qrange, where Q = (4π/λ)sin θ and 2θ is the scattering angle, of 0.0014−0.53 Å−1. At HFIR, we used 1.7 m/6 Å, 6 m/6 Å, and 14.5 m/ 12 Å configurations, giving a Q-range of 0.0015−0.38 Å−1. The wavelength spread, Δλ/λ, at NCNR is 12% and at HFIR 15%. Both beamlines use Ordela area detectors. In order to maximize contrast, fully hydrogenated surfactants were dispersed in 100% D2O. The samples were held at 30 °C. The scattering intensity profiles, I(Q) versus Q, were obtained by azimuthally averaging the processed 2D images, which were normalized to incident beam monitor counts, and corrected for detector dark current, pixel sensitivity, and empty beam scattering background.34

3. RESULTS AND DISCUSSION 3.1. Concurrent Loading of Monomers. Our investigation probed the question: if hydrated together, will surfactants with monomers form vesicles with loaded bilayers (Figure 1a)? We found that monomer-loaded vesicles can indeed be formed in a broad range of surfactant ratios (Figure 2). The implication of this finding is that the vesicle-templated nanocapsules can be formed simply by hydrating surfactants with monomers followed by a polymerization. To investigate the monomer/surfactant assemblies, we performed a series of experiments where mixtures of surfactants and monomers at different ratios were agitated in water and allowed to equilibrate. We found that mixing monomers with each surfactant separately worked best. These surfactant/ monomer stock solutions did not show the presence of vesicles. The results were analyzed with dynamic light scattering and plotted on phase diagrams (Figure 2). Each line (surfactants to water ratio) in the phase diagram was assembled from 21 data points. The increments were 5%. Areas labeled as V and M showed predominantly vesicles and micelles, respectively. DLS data for these samples showed objects with 220 ± 40 nm in size. Samples that formed a visible precipitate were labeled as P on the phase diagram. The appearance of the precipitate ranged from visible white clumps to highly turbid suspensions. All of them showed large objects in DLS. The areas labeled as L were attributed to a lamellar phase, in accordance with previous reports.26,27 In these regions, samples show no visible precipitate of turbid suspensions; however, the DLS analysis

Figure 2. CTAT−SDBS−water phase diagrams without monomers (a) and with acrylic (b) and styrene (c) monomers at 25 °C. The concentration of monomers was constant. One-phase regions represent vesicles (V) and micelles (M). Two-phase regions represent precipitates (P) and lamellar phases (L).

shows large aggregates, and the samples are more viscous than all the other samples. Often, samples show phase separation after standing for several hours. Predominant structures in these samples appear to be wormlike micelles and bilayer sheets.27 Figure 2a shows the phase diagram for surfactants in the absence of monomers. The results are consistent with previously published literature data.27 Figure 2b and c shows the behavior of surfactant/monomer mixtures in water at constant concentration of monomers (Figure 2b) and constant surfactant/monomer ratio (Figure 2c). 7063

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aging of loaded samples up to 24 h or after polymerization of monomers under appropriate conditions. In nonpolymerized vesicles, after 24 h both PDI and size of vesicles increased substantially, indicating formation of aggregates. Our first study of the use of surfactants as templates involved mixtures of CTAB and SDS in water (1% w/v), evaluated with SAXS. The most prominent feature of the unloaded CTAB/ SDS samples was a broad, low intensity feature centered around 0.15 Å−1 and a sharp, intense Bragg peak located at 0.169 Å−1 (Figure 4). The broad feature is located in the region associated

The most important outcome of these experiments was the spontaneous formation of vesicles loaded with monomers in a broad range of concentrations of monomers and surfactants (Figure 2b and c, red area). To put the concentrations of surfactant vesicles in perspective, at 4% by weight, 200 nm vesicles would occupy more than 30% of the total volume. This concentration is near the practical limit of vesicle formation. Monomers generally did not negatively affect the ability of surfactants to form vesicles. In fact, over a significant range of concentrations, the presence of acrylate monomers favored the formation of vesicles over lamellar or micellar phases. The polydispersity index (PDI) remained within the 0.05− 0.2 range in the middle of the vesicle regions (Figure 3). Figure

Figure 4. SAXS measurement of CTAB-SDS surfactant aggregates interacting with monomers. Without monomers, a Bragg peak was seen, indicating a lamellar phase. When concurrently loaded with monomers, the Bragg peak disappears and lower Q features (broad feature with small narrow peaks superposed), presumably due to vesicles, appear. The inset shows similar low Q features for 1,2dimyristoyl-sn-glycero-3-phosphocholine (DMPC) vesicles for comparison.

with the thickness of the vesicle shell,21 while the Bragg peak is indicative of intermolecular structure. The lack of additional Bragg peaks in the region studied does not allow for definitive determination of the lattice structure, but the repeat distance given by the single peak, 2π/0.169 Å−1 = 37 Å, fits well with the expected size of the lamellar structure of CTAB.39 Additionally, the location in the phase diagram of the mixture studied is near the boundary of the V and Lα phases for CTAB−SDS mixtures. Therefore, we suggest that we are observing both the form factor of vesicle shells and the structure factor of lamellar sheets for the unloaded surfactants. When monomers were added, the Bragg peak quickly, though not completely, disappeared, indicating that the lamellar structure was greatly disrupted. Concurrently, the broad formfactor feature rose significantly in intensity, suggesting that more vesicles are formed. Therefore, the addition of monomers apparently converts the lamellar phases into the vesicle phase. Additional peaks are superposed on the broad peak, similar to those seen for 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC) vesicles,21 shown in the inset of Figure 4. These are presumably due the counterions associated with the surfactants. Figure 5 shows SANS scans of CTAB−SDBS samples as they vary in SDBS concentration. This figure provides confirmation that coassembly of loaded vesicles is occurring only at low SDBS concentration, outside of the normal region for vesicle formation; and that at higher concentrations, within the vesicle phase, a more standard empty vesicle to loaded vesicle process

Figure 3. CTAT−SDBS−water phase diagrams with (a) acrylic and (b) styrene monomers at 25 °C and representation of polydispersity (PDI), concentration of monomers was constant. Two-phase regions represent precipitates (P). Dashed lines show vesicle regions (region V in Figure 2).

3 shows the portions from the phase diagram in Figure 2 that include the vesicle regions (up to 3% surfactant content for acrylic monomers and up to 1.5% surfactant content for styrene monomers). We chose square representation instead of more traditional triangular shape to highlight the features in the phase diagram at low concentration of surfactants. PDI values below 0.2 have been traditionally associated with narrow monodisperse distributions.35−38 In both monomeric systems, PDI increased near the boundary on the phase diagram with regions where vesicles did not predominate. This observation is expected since the transition between regions on the phase diagram is not likely to be very sharp, and a mixture of aggregates with different shapes and sizes exist at the boundary. The polydispersity index did not change noticeably during the 7064

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micelles aggregate, giving rise to correlation peaks around ∼0.02−0.03 Å−1 (for the low SDBS concentration) and a nonzero slope at the lowest −Q values. At an intermediate concentration without monomers, 20% (red data points), micelles are present along with some nonzero volume fraction of vesicles. These vesicles are similar to those seen in the 30% sample. Addition of monomers leads to weak correlation peaks and a fair amount of aggregation, like the 15% sample, but the vesicle feature behaves similarly to the 30% sample. Thus, this sample appears to be in a coexistence region between a pure micelle phase and a pure vesicle phase; adding monomers pushes it more into the vesicle phase. In our previous studies of lipid vesicle loading, we found that the monomers swelled the bilayer by partially phase separating into the interleaf region.20 The same type of analysis was performed for the surfactant vesicles by analyzing data in the bilayer-thickness Guinier region. At these Q values (∼0.04− 0.08 Å−1), the bilayer appears as a flat sheet with thickness d. For QRg < 1, where Rg is related to the thickness as d = Rg(12)1/2, the intensity of scattering from a sheet goes as I ∝ exp(−Q2Rg2)/Q2. Therefore, a plot of ln(IQ2) versus Q2 will give line with slope −Rg2, from which the thickness can be determined.43−46 Fits to unloaded and concurrently loaded CTAT-SDBS samples are shown in Figure 6, showing a swelling of about 0.88

Figure 5. SANS study of the effect of concurrent loading on vesicle formation near the vesicle phase boundary of CTAB-SDBS samples. All samples were at 1% total surfactant concentration. At 15% SDBS, without monomers the surfactants produce only micelles; addition of monomers converts some of the micelles into large vesicles. At 30% SDBS, the system is firmly within the vesicle phase, even without monomers; addition of monomers does not affect the vesicle volume fraction.

occurs. In the SANS scans, large vesicles are indicated by a low Q shoulder (a change in the slope of the curve, the position of which depends on the diameter of the vesicle) and then a Q−2 dependence until reaching the Guinier region due to the bilayer thickness, after which the intensity goes as Q−4.40 Micelles, the predominant structures at low SDBS concentration, being smaller and compact, are indicated only by a mid- to high-Q feature, depending on their diameters. For the samples without monomers (Figure 5, left side), as the SDBS concentration is increased from 15% to 30% the low Q intensity due to vesicles clearly increases, while the higher Q features due to micelles diminish. At 30% (green data points), only vesicles are present in large quantity, as can be seen by the Q−2 dependence of the intensity from ∼0.01 Å−1 up to ∼0.06 Å−1 where the Guinier region for the thickness of the vesicle shell begins. When the surfactants are mixed in the presence of monomers, the SANS curves are altered (Figure 5, right side). The low SDBS concentration (black data points) shows that where there had been no large length-scale scattering,41,42 the addition of monomers creates large vesicles. DLS gives the diameter of the vesicles as ∼220 nm. Modeling the low Q region as a vesicle with this length-scale gives a shoulder feature at around 0.002 Å−1, which compares well with what we observe in the scan. In contrast, within the normal vesicle phase (at 30% SDBS), the addition of monomers does not appreciably change the low Q intensity (green data points), nor does it change the position of the shoulder, meaning that the vesicles remain roughly the same size, though the bilayer does swell as will be shown below. Interestingly, the vesicles are smaller in diameter in the normal vesicle phase region, as can be seen by the higher Q value of the shoulder feature, at ∼0.005 Å−1. Since this Q value is ∼2.5 times higher than that for the 220 nm vesicles, this suggests the vesicles have a diameter of around 85 nm, as was confirmed by DLS (not shown). Therefore, while coassembly of surfactants with monomers leads to vesicle formation even outside of the regular vesicle phase region, the vesicles have a larger diameter than when formed within the vesicle phase. In all the loaded samples, regardless of SDBS concentration, it appears that the remaining

Figure 6. Modified Guinier analysis for sheetlike forms of SANS data of a concurrently loaded CTAT-SDBS sample. Linear fits give bilayer thickness values that increase by 8.77 ± 0.25 Å when loaded with acrylate monomers.

nm. This is significantly larger than we saw for the lipid vesicles (0.28 nm). More studies will need to be performed to understand the reason for this difference. It is possible that the surfactant vesicles accommodate more monomers than the lipid vesicles, or that more monomers phase separate into the interleaf region, which would give important information about monomer/surfactant interactions. Simultaneous HPLC/SANS measurements, in development, are needed to be able make any definitive statements. These observations validate and expand our recent report on synergistic self-assembly of surfactants and monomers.31 Data presented on phase diagrams suggest that monomers may play an active role in assembling a bilayer. Acrylic monomers expanded the range of surfactant ratios that resulted in predominant formation of vesicles (Figure 2). These results support the hypothesis that certain monomers stabilize the bilayer structure by increasing van der Waals interactions. At 7065

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the same time, styrene monomers did not result in stable vesicles above 1.5% surfactant concentration in water (Figure 2). The active role of monomers in coassembly of loaded vesicles is an interesting phenomenon that merits further investigation in future studies. In summary, surfactant vesicles containing monomers inside the bilayer can be formed by the hydration of a surfactant/ monomer mixture. Monomers appear to have a stabilizing effect on the vesicles as judged by the broader area on the phase diagram corresponding to spontaneous vesicle formation. 3.2. Diffusion Loading of Monomers. In diffusion loading, monomers are added to an aqueous solution of preformed vesicles (Figure 1b). Monomer molecules are allowed to diffuse through water into the hydrophobic interior of the bilayer. This method has been used in previous studies using vesicles as templates for nanocapsules. Vesicles formed by phospholipids and nonlipid surfactants were used in combination with diffusion loading. In previous studies of diffusion loading of vesicles prepared from saturated phospholipids, we found that lipid bilayers have a certain capacity for accommodating monomers. This loading capacity was independent of the structure of monomers or the curvature of vesicles but increased with increased temperature.19−21 We also found that due to the kinetics of diffusion, the time allowed for loading was important and likely explained the discrepancies reported earlier.21 A potential benefit of diffusion loading is that the loading procedure is the same, regardless of the type of monomers. We showed previously that, with lipids as scaffolds, it is possible to control the amount and ratio of monomers loaded into the bilayer by adjusting the loading time, initial ratio of monomers, or replacing one monomer with another.21 In diffusion loading experiments, we added neat monomers to the aqueous suspension of vesicles. We monitored the size of the vesicles with dynamic light scattering and measured the amount of monomers with HPLC following a method described previously.19 Figure 7 shows time-resolved data on the size of vesicles and the amount of monomers in the bilayer. Diffusion loading experiments were performed with formulations of surfactants in the middle of the vesicle region in the phase diagrams (Figure 2) and close to the boundary between the vesicle and micelle or lamellar phases. Representative data are shown for vesicle formulations with different surfactant concentrations. Other surfactant formulations revealed the same behavior as described in Figure 1b (data not shown). A possible reason for greater loading capacity of vesicles with higher SDBS content is lower stiffness of bilayers as could be inferred from previously published data.29 In addition, CTAT has a large counterion that may hinder the diffusion of monomers into the vesicles. Vesicles prepared with 2% surfactant concentration showed precipitation upon absorption of a moderate amount of monomers. The position of the surfactant mixture on the phase diagram (Figure 2a) had no effect on the stability of vesicles. These findings are in contrast with the observation of stable monomer-loaded vesicles prepared with 2% and greater surfactant concentration (Figure 2b,c). It appears that the diffusion loading causes the collapse of surfactant vesicles at higher surfactant concentrations, while concurrent loading, described above, can lead to stable monomer-loaded vesicles. In previous studies, the synthesis of nanocapsules was done using vesicles prepared with 1% concentration of surfactants31 and

Figure 7. Time-resolved loading of DVB into CTAT-SDBS-water bilayers at 30 °C. Measured by HPLC (a) and DLS (b). Each series is labeled with the composition of the bilayers. Most of the error bars have been omitted for clarity.

higher concentrations of surfactant vesicles were not used for templating the nanocapsules. Vesicles prepared at 1% total concentration of surfactants in water were stable upon addition of monomers. Here, monomers were absorbed until a certain amount has been reached. This behavior of vesicles is similar to what we observed previously with saturated phospholipids. We used divinylbenzene as a standard monomer for these loading experiments, similarly to our previous studies of loading phospholipid vesicles. The kinetics of loading observed here was faster than what we found with phospholipids. In the present study, it took approximately 3−4 h to reach the saturation capacity of the surfactant bilayer, while time-resolved loading of DVB into DMPC liposomes showed that the capacity has been reached within 10−15 h.19 SANS scans of fully loaded acrylate diffusion-loaded CTAT− SDBS samples were performed to measure the amount of bilayer swelling in comparison to concurrently loaded samples and previously studied lipid vesicle samples. Figure 8 shows a modified Guinier analysis, indicating a swelling of 0.660 ± 0.044 nm. This is less swelling than for the concurrently loaded samples, but more than for lipid vesicles. Further planned experiments will determine if this indicates that diffusion loaded samples accommodate fewer monomers than concurrently loaded samples or if this indicates a different proportion of the monomers are phase separated into the interleaf region. 3.3. Synthesis of Vesicle-Templated Nanocapsules. We used monomer-loaded vesicles to create polymer capsules. The polymerization was initiated with UV irradiation using a photoinitiator. The polymerization was done in a similar way to 7066

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Figure 8. Modified Guinier analysis of SANS data of a diffusion-loaded CTAT−SDBS sample. Linear fits give bilayer thickness values that increase by 6.60 ± 0.44 Å when loaded with acrylate monomers.

the previously reported synthesis of nanocapsules in liposomes9,47 and surfactants.31 In a typical setup, photochemically induced polymerization was accomplished in a quartz vessel with the depth of approximately 3 mm.9 To evaluate the permeability properties of nanocapsule shells, we prepared nanocapsules with entrapped dyes.10,23,24 High concentration of dyes in the original solution can affect the polymerization by blocking UV light in the areas of the sample away from the interface with the quartz vessel. To minimize the effect of the dyes, one could either ensure adequate agitation of the sample or use a vessel with a short path length for light. In our setup, a quartz test tube was placed inside a larger diameter quartz vessel so that the gap between the walls was approximately 3 mm. The aqueous solution of monomer-loaded vesicles was poured into this gap, and the quartz assembly was placed into a photochemical cabinet with cylindrical placement of UV lamps. Surfactant scaffold was removed by washing nanocapsules in methanol. Upon mixing aqueous suspension with methanol, nanocapsule form a gel-like precipitate. The precipitate can be centrifuged and washed repeatedly with methanol to remove surfactant molecules. Nanocapsules can be freeze-dried48 from water, benzene or tert-butanol. Dried nanocapsules show predominantly spherical structures in TEM and SEM images (Figure 9). The dye retention test allowed us to estimate the fraction of nanocapsules without pinhole defects exceeding approximately 1 nm as shown previously.31 Most formulations of monomer-loaded vesicles produced polymer nanocapsules that showed retention of dye in the inner compartment.

Figure 9. Electron microscopy images of nanocapsules. SEM images of polymer nanocapsules (a, b, d, e, f), after surfactant removal, precipitation, multiple washings, resuspension, and freeze-drying. (c) TEM image of polymer nanocapsules, after surfactant removal, precipitation, multiple washings, and resuspension. All samples 1% (w/v) solution in water; (a−c) SDBS/ CTAT = 80:20; (d) SDBS/ CTAT = 90:10; (e) SDBS/CTAT = 70:30; (f) SDBS/CTAT = 20:80. Acrylic monomers/surfactant ratio = 2:1.

synergistic self-assembly of surfactant and monomers into vesicles.31 Preassembled vesicles can be loaded with monomers, depending on the position of the surfactant composition on a phase diagram. For both concurrent loading and diffusion loading, neutron scattering data confirmed an increase in thickness of the bilayer in agreement with the amount of accommodated monomers. As shown previously for the phospholipid vesicles, surfactant vesicles have a limited capacity for accommodating monomers via diffusion loading. Polymerization of monomers with cross-linkers in the interior of surfactant vesicles resulted in the formation of hollow polymer nanocapsules. Spontaneous formation of predominantly unilamellar vesicles loaded with monomers without the need for further processing, such as extrusion, coupled with the use of inexpensive materials, greatly simplifies the synthesis of hollow nanocapsules, versatile platforms with diverse documented applications.

4. CONCLUSIONS This study investigated the formation of vesicles loaded with different monomers produced by two methods, concurrent loading and diffusion loading. In concurrent-loading method, monomer-loaded vesicles were formed in a broad range of ratios of anionic and cationic surfactants. For acrylic monomers, the vesicle regions were broader for monomer-loaded vesicles compared with vesicles with no monomers. These observations suggest that accommodation of monomers may stabilize the bilayer. X-ray scattering data supported the transition of a lamellar phase into a vesicular phase. Neutron scattering data also supported the stabilization of vesicle structures with monomers. These data validate and expand recent report on

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Corresponding Author

*E-mail: [email protected]. Author Contributions

The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes

The authors declare no competing financial interest. 7067

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ACKNOWLEDGMENTS This work was supported by FedEx Institute of Technology, National Science Foundation (CHE-1316680, CHE-1012951, and CHE-0933363), and Saint Louis University Presidential Research Fund award. Work was partially supported by grant from the Committee of Science of the Ministry of Education and Science of Republic of Kazakhstan (No. 0547/GF2). Portions of this work were performed at the DuPontNorthwestern-Dow Collaborative Access Team (DND-CAT) located at Sector 5 of the Advanced Photon Source (APS). DND-CAT is supported by E.I. DuPont de Nemours & Co., the Dow Chemical Company and Northwestern University. Use of the APS, an Office of Science User Facility operated by the U.S. Department of Energy (DOE) Office of Science by Argonne National Laboratory, was supported by the U.S. DOE under Contract No. DE-AC02-06CH11357. Portions of this work were performed at Oak Ridge National Laboratory’s Center for Structural Molecular Biology and High Flux Isotope Reactor, sponsored by the Office of Biological and Environmental Research and the Scientific User Facilities Division, Office of Basic Energy Sciences, U.S. Department of Energy. V.U. acknowledges support by the U.S. Department of Energy, Basic Energy Sciences, Materials Sciences and Engineering Division. We acknowledge the support of the National Institute of Standards and Technology, U.S. Department of Commerce, in providing the neutron research facilities used in portions of this work.

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