Formation of Stable Nanocapsules from Polymerizable Phospholipids

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Formation of Stable Nanocapsules from Polymerizable Phospholipids† Glenn E. Lawson, Yongwoo Lee, and Alok Singh* Center for Bio/Molecular Science and Engineering, Code 6930, Naval Research Laboratory, Washington, D.C. 20375 Received March 13, 2003. In Final Form: April 30, 2003 The formation of lipid-based hollow, spherical nanocapsules is reported. Polymerizable phospholipids equipped with divinylbenzoyl functionality in their headgroup region are synthesized to explore their polymerization behavior in vesicles. Polymerizable phospholipids alone or as mixtures with phosphatidylcholine produced vesicles as an aqueous dispersion and produced stable nanocapsules upon polymerization at room temperature. These structures survived lyophilization and a redispersion cycle and maintained their structural integrity in nonaqueous solvent. Vesicles were used for encapsulating enzymes in their hydrophilic cores.

Introduction Vesicles represent a unique class of technologically relevant structures by virtue of their ever-increasing utility across disciplines. Extensive research activities involving vesicles began after the appearance of a report by Bangham1 showing electron microscopic evidence to reveal the presence of concentric, bimolecular layers producing vesicular morphologies. Published reports on the formation and properties of vesicles revealed their application potential involving their encapsulation and trans-bilayer transportation capabilities.2,3 Despite their technological marvels, vesicles suffer a major drawback of limited stability in working environments. Subsequent research efforts to overcome the stability problem led to development of the following three strategies: (a) crosslinking the amphiphiles after the formation of vesicles by polymerization;4-7 (b) formulating lipid compositions including involvement of lipids containing poly(ethylene glycol) (PEG) in their headgroups;8-10 and (c) developing new, nonlipid materials to form stable vesicles.11,12 The approach involving PEG lipid is mainly directed to drug delivery applications involving stable vesicles with prolonged circulation life.8-10,13-14 The approach involving * To whom correspondence should be addressed. E-mail: asingh@ cbmse.nrl.navy.mil. † Part of the Langmuir special issue dedicated to David O’Brien. (1) Bangham, A. D. Adv. Lipid Res. 1963, 1, 65-104. (2) Juliano, R. L. In Liposomes: from physical properties to therapeutic applications; Knight, C. G., Ed.; Elsevier/North-Holland: New York, 1981; Chapter 14, pp 391-407. (3) Mayhew, E.; Papahadjopoulos, D. Liposomes; Ostro, M. J., Ed.; Marcel Dekker: New York, 1983; Chapter 7, pp 289-341. (4) Regen, S. L.; Czech, B.; Singh, A. J. Am. Chem. Soc. 1980, 102, 6638-6640. (5) Hub, H.; Hupfer, B.; Koch, H.; Ringsdorf, H. Angew. Chem., Int. Ed. Engl. 1980, 19, 938-940. (6) Johnston, D. S.; Sanghera, S.; Pons, M.; Chapman, D. Biochim. Biophys. Acta 1980, 602, 57-69. (7) O’Brien, D. F.; Klingbiel, R. T.; Whitesides, T. H. J. Polym. Sci., Polym. Lett. Ed. 1981, 19, 85-101. (8) Lasic, D. D. Angew. Chem., Int. Ed. Engl. 1994, 33, 1685-1698. (9) Lasic, D. D.; Needham, D. Chem. Rev. 1995, 95, 2601-2628. (10) Lin, H. Y.; Thomas, J. L. Langmuir 2003, 19, 1098-1105. (11) Discher, D. E.; Eisenberg, A. Science 2002, 297, 967-973. (12) Nardin, C.; Thoeni, S.; Widmer, J.; Winterhalter, M.; Meier, W. Chem. Commun. 2000, 15, 1433-1434. (13) Allen, T. M.; Hansen, C.; Martin, F.; Redemann, C.; Yau-Young, A. Biochim. Biophys. Acta 1991, 1066, 29-36. (14) Needham, D.; Dewhirst, M. W. Adv. Drug Delivery Rev. 2001, 53, 285-305.

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cross-linking of monomers after the formation of vesicles received much attention because of the applicability of such systems in both biological and nonbiological systems.15-17 The versatility of the polymerization approach lies in the fact that it provides selection of monomer functionalities18 and choice of their placement18-21 in the lipid molecules allowing vesicles to be for desired applications including extending vesicle utility beyond biological applications.22-24 Despite their superior technical advantage and versatility, the higher cost of polymerized vesicles has restricted their overall usage in technology development. The focus of the current study is on the development of polymerized vesicles that can be polymerized efficiently and effectively while sustaining their membrane properties. In particular, our goal is to incorporate polymerization capability to the headgroup region leaving the acyl chain region intact. Previously, formation of vesicles from oligomerizable phospholipids containing β-nitrostyrene25 and styryl26 functionality has been reported. Lipid monomer with β-nitrostyryl functionality polymerizes effectively to form oligomers in self-organized assemblies but reacts with water in a basic medium to produce aldehyde functionality. Monomer reactivity in a basic medium limits their utility. Lipids with a styryl group polymerize well, but the polymerization process takes about 90 min of UV exposure at 25 °C to achieve complete participation of monomers. From an application point of (15) Ringsdorf, H.; Schlarb, B.; Venzmer, J. Angew. Chem., Int. Ed. Engl. 1988, 27, 113-158. (16) Singh, A.; Schnur, J. M. Polym. Adv. Technol. 1994, 5, 358-373. (17) Mueller, A.; O’Brien, D. F. Chem. Rev. 2002, 102, 727-757. (18) Singh, A.; Schnur, J. M. Phospholipids Handbook; Cevc, G., Ed.; Marcel Dekker: New York, 1993; pp 233-291. (19) Ringsdorf, H.; Schlarb, B. Makromol. Chem. 1988, 189, 299315. (20) Regen, S. L.; Shin, J.; Yamaguchi, K. J. Am. Chem. Soc. 1984, 106, 2446-2447. (21) Ringsdorf, H.; Schlarb, B.; Tyminski, P. N.; O’Brien, D. F. Macromolecules 1988, 21, 671-677. (22) Stanish, I.; Lowy, D. A.; Tender, L. M.; Singh, A. J. Phys. Chem. B 2002, 106, 3503-3509. (23) Markowitz, M. A.; Chow, G. M.; Singh, A. Langmuir 1994, 10, 4095-4102. (24) Singh, A.; Chow, G. M.; Chang, E. L.; Markowitz, M. A. CHEMTECH 1995, 25, 38-43. (25) Ravoo, B. J.; Engberts, B. F. N. J. Chem. Soc., Perkin Trans. 2001, 2, 1869-1886. (26) Lawson, G. L.; Breen, J. J.; Marquez, M.; Singh, A.; Smith, D. Langmuir 2003, 19, 3557-3560.

This article not subject to U.S. Copyright. Published 2003 by the American Chemical Society Published on Web 05/28/2003

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Figure 1. Structures of headgroup-polymerizable phospholipids.

view, there is need for a polymerized vesicle system, which is convenient to synthesize and whose precursor monomers are chemically inert toward the dispersion medium and the molecules intended for encapsulation, and the polymerization process should be complete within a few minutes. In this paper, we report the synthesis and selfassembling behavior of two new phospholipids, 1 and 2, by incorporating 3,5-divinylbenzoyl functionality in their headgroup region (Figure 1). These phosphatidylethanolamine-based lipids are designed considering their inertness against other chemical species present in the dispersion medium including those selected for encapsulation. Moreover, two active vinyl species should allow complete participation of active vinyl groups available to form a cross-linked polymer network to produce stable vesicle structures without affecting the morphologies. Experimental Section General. 1,2-Dipalmitoyl-sn-glycero-3-phosphoethanolamine (DPPE), 1,2-dilauryl-sn-glycero-3-phosphoethanolamine (DLPE), and dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) were purchased from Avanti Polar Lipids. Unless stated otherwise, all solvents and reagents were purchased from Aldrich Chemical Co. and used as received. CHES buffer was prepared by making a 10 mM solution of 2-cyclohexylamino-1-ethanesulfonic acid in deionized water, and its pH was adjusted to 8.6 with 1 M NaOH. Wild-type phosphotriesterase (PTE) was received from Professor Frank Raushel at Texas A&M University and was used after diluting with CHES buffer to attain a protein concentration of 100 µg/mL. For thin-layer chromatography (TLC), glass plates (coated with silica gel-60 with fluorescent indicator) were used. For spectral characterization and analysis, an Agilent 8453 UVvis spectrophotometer, a Bruker DRX-400 nuclear magnetic resonance spectrometer (400 MHz for proton and 100 MHz for carbon), and a Tensor 27, Bruker infrared spectrophotometer were used. Mass spectra of synthetic intermediates and lipids were recorded on an Applied Biosystems, MDS Sciex, API Qstar Pulsar instrument. Synthesis of Phospholipids and Their Intermediates. Synthesis of 3,5-Divinylbenzoic Acid (DVBA). Solid N-bromosuccinimide (2.37 g, 13.3 mmol) was added to a stirred solution of 3,5-dimethylbenzoic acid (1.0 g, 6.6 mmol) in carbon tetrachloride (10 mL). The reaction mixture was heated to dissolve all the components and refluxed after the addition of catalytic amounts (73 mg) of benzoyl peroxide until the starting material 3,5-dimethyl benzoic acid was consumed in the reaction and a new spot due to reaction product was observed on thin-layer chromatography (5% ethyl acetate/hexane, Rf ) 0.4). A 1-3 h refluxing is sufficient. The reaction mixture was cooled to room temperature, and the solid byproduct succinimide was filtered off. Solid 3,5-bis(bromomethyl) benzoic acid was collected after removing the solvent from the filtrate. Final traces of the solvent were removed under high vacuum affording 2.05 g of product in 99% yield. Before proceeding to the next step, disappearance of methyl protons was confirmed by NMR spectrum. This product

Lawson et al. was used without further purification to perform the next synthetic step. Preparation of 3,5-carboxybenzyltriphenylphosphoniumdibromide salt was carried out by the slow addition of solid triphenylphosphine (3.49 g, 13.3 mmol) to a solution of 3,5-bis(bromomethyl) benzoic acid (2.05 g, 6.6 mmol) in 25 mL of acetone and refluxing the mixture overnight. Formation of a precipitate from solution indicated the salt formation. The reaction mixture was then cooled to room temperature, and the salt was collected by filtration. The solid was washed repeatedly with hexane and dried under vacuum to afford 2.42 g (45%) of ylide. This salt is good enough to serve as the precursor for the next step. To a 30 mL formaldehyde solution (240 mmol) prepared by diluting 20 mL of 37% aqueous solution with 10 mL of water, 2.42 g (3.1 mmol) of the salt was added. The solution was efficiently stirred to make a suspension followed by a dropwise addition of 5 M solution of NaOH (7.5 mL, 37.5 mmol). This resulted in a clear solution. A white precipitate was observed after stirring the reaction mixture for 3 h at room temperature. This precipitate was collected by filtration and treated with 10% aqueous HCl until a pH of 2 was achieved. This resulted in the liberation of free divinylbenzoic acid as a white solid. The free acid is distinctly different than its sodium salt in texture and dispersing behavior. This solid was collected by suction filtration and dried under high vacuum to yield 310 mg (62%) of the divinylbenzoic acid. The acid was analyzed as a chloroform solution on silica gel coated TLC plates (CHCl3/CH3OH, 95:5 v/v). A single spot on TLC (Rf ) 0.24) and a sharp melting point at 142-143 °C revealed the homogeneity of the product. 1H NMR (400 MHz CDCl3) δppm: 7.9 (s, 2H), 7.4 (s, 1H), 6.7 (dd, 2H, J ) 11 Hz), 5.8 (d, 2H, J ) 11 Hz), 5.3 (d, 2H, J ) 18 Hz). 13C NMR (100 MHz CDCl3): 115.0, 126.0, 128.0, 130.0, 135.0, 137.0, 166.0. TOF HRMS: calculated for C11H10O2, 174.07; found, 173.1. FTIR (KBr pellet): 3435, 2912, 2599, 1704, 1600, 1589, 1457, 1313, 1241 cm-1. Synthesis of 1,2-Dipalmitoyl-sn-glycero-3-phospho-N-(2hydroxyethyl)-3,5-divinylbenzamide (DPPE-DVBA, 1). In a roundbottom flask equipped with a magnetic stirring bar and an inlet and outlet for dry nitrogen were placed equimolar amounts of DVBA (25 mg, 0.143 mmol), 1-hydroxybenzotriazole (19 mg, 0.144 mmol), and N-ethyl-3-(dimethylaminopropyl) carbodiimide hydrochloride (EDC) (28 mg, 0.145 mmol), and the compounds were dissolved in 40 mL of reagent grade chloroform. The reaction mixture was protected from light by wrapping the flask with aluminum foil and stirring for 30 min at room temperature under an inert atmosphere to activate the acid. To this solution, DPPE (100 mg, 0.172 mmol) dissolved in chloroform (1 mL) was added in small portions followed by slow addition of 20 µL of triethylamine over a period of 1 min. The reaction mixture was stirred overnight maintaining the inert atmosphere. TLC analysis using a CHCl3/CH3OH (80:20, v/v) solvent mixture indicated complete disappearance of the starting materials (DVBA and DPPE) and formation of a new spot. The reaction was stopped by taking it off the stirrer, washed sequentially using 15 mL portions each of 4% sodium carbonate solution and water, and dried over anhydrous sodium sulfate. Solid material collected after the removal of solvent was chromatographed on a silica gel column. Elution with a CHCl3/CH3OH (80:20, v/v) solvent mixture afforded 88 mg of pure material in 73% yield. A single spot on TLC (Rf ) 0.35) was detected using a CHCl3/CH3OH (80:20, v/v) solvent system. Ninhydrin spray did not produce any stain on the plate, indicating the absence of any free DPPE in the product collected after column chromatography. 1H NMR (400 MHz CDCl3) δppm: 7.90 (s (b), 2H), 7.40 (s (b), 1H), 6.61 (d, 2 H, J ) 11 Hz), 5.82 (d, 2 H, J ) 11 Hz), 5.32 (d, 2 H, J ) 18 Hz), 4.10-3.51 (8H), 2.18 (s (b), 4H), 1.52-1.28 (s (b), 52 H), 0.92 (t, 6H). 13C NMR (100 MHz CDCl3): 17.0, 21.0, 22.0, 34.0-36.0, 37.0, 41.0, 61.0, 62.0, 118.0, 128.0, 130.0, 132.0, 138.0, 141.0, 168.0, 172.0. TOF HRMS: calculated for C48H8NO9P, 846.56; found, 847.8. FT-IR (KBr pellet): 3380, 2942, 2858, 1746, 1656, 1595, 1547, 1247, 1079 cm-1. Synthesis of 1,2-Dilauroyl-sn-glycero-3-phospho-N-(2-hydroxyethyl)-3,5-divinylbenzamide (DLPE-DVBA, 2). Following the general procedure described in the previous section, coupling of DVBA with DLPE (25 mg, 0.043 mmol) provided 20 mg (63%) of pure DLPE-DVBA (Rf ) 0.64, silica gel, CHCl3/CH3OH (80:20,

Nanocapsules from Polymerizable Phospholipids v/v)). 1H NMR (400 MHz CDCl3) δppm: 7.91 (d, 2H), 7.54 (d, 2H), 6.63 (dd, 2H, J ) 11.0, 11.0 Hz), 5.81 (d, 2H, J ) 18.0 Hz), 5.3 (d, 2H, J ) 11.0 Hz), 5.22 (s (b), 1H), 4.33, 4.12, 3.97, 3.79 (m, 8H), 2.18 (s (b), 4H), 1.52-1.28 (s (b), 36H), 0.92 (t, 6H). 13C NMR (100 MHz CDCl3): 17.0, 21.0, 22.0, 34.0-36.0, 37.0, 41.0, 61.0, 62.0, 118.0, 128.0, 130.0, 132.0, 138.0, 168.0, 172.0. TOF HRMS: calculated for C40H65NO9P, 734.44; found, 735.7. FT-IR (KBr pellet): 3374, 2930, 2858, 1740, 1638, 1583, 1541, 1475, 1240, 1081 cm-1. Differential Scanning Calorimetry (DSC) Studies. Calorimetric studies on hydrated phospholipids and their mixtures were performed on a model 2920 Modulated DSC from TA Instruments. Phospholipid samples were weighed directly in the DSC pans and dissolved in chloroform to spread as a uniform thin film, and the solvent was removed under a gentle stream of nitrogen followed by applying high vacuum to remove traces of solvent. Thin lipid film was covered with 10 µL of deionized water, and the pans were sealed and left in an oven set at 70 °C for 5 h to allow complete hydration. For polymerized samples, freeze-dried polymerized vesicles were weighed into DSC pans and hydrated with water before running DSC scans. Preparation and Characterization of Vesicles. Preparation and Visualization of Vesicles. Calculated aliquots of chloroform solution of lipid were removed to dispense 5 mg of the desired lipid into a test tube. A thin lipid film was coated on the walls of the tube by removing solvent under a gentle stream of nitrogen followed by thorough drying of the film under high vacuum. Thin lipid films were hydrated in deionized water and buffers at pH values ranging from 4.0 to 9.3 in order to optimize vesicle formation. Lipid dispersions were hydrated by heating at 50 °C for 1 h and dispersed in the medium by intermittent vortex mixing followed by sonication at 50 °C using a Branson Sonifier model 450. A cup horn device equipped with water intake and outlet connections was used for sonicating the sample at the desired temperature. In most of the cases at 50% power and 80% duty cycle 1 h of sonication produced suspensions of constant turbidity leading to the formation of uniform-sized vesicles. A similar protocol has been used for making vesicles from 1 and DPPC mixtures in different ratios. Vesicles images were acquired using a Zeiss EM-10 transmission electron microscope at 60 kV. Typically, a drop of vesicle suspension was placed on a 200 mesh copper Formvar/carbon grid. Vesicles on the grid were stained by placing a drop of 1% uranyl acetate in water followed by removal of excess solution by wicking it with a piece of filter paper. Polymerization of Vesicles. All polymerization experiments were carried out at room temperature. Vesicles derived from either single components or mixed lipid systems were polymerized using a combination of radical initiation and photopolymerization, and redox polymerization. Irrespective of the method of polymerization, the polymerization extent was monitored using TLC and NMR techniques. For all NMR studies, the vesicles were prepared in 0.7 mL of buffer prepared by mixing a D2O solution of 0.05 M NaHCO3/0.1 M NaOH and adjusting the pH to 9.3. For radical-initiated polymerization, a water-soluble initiator, 2,2′-azobis(2-methylpropionamidine) dihydrochloride (AAHP), was used alone or in combination with UV irradiation at 254 nm. AAHP dissolved in water was added to the vesicle dispersions, and the [phospholipids]/[initiator] ratio was adjusted to 7:1 (mol/ mol). Photochemical reactions were carried out in a quartz cell placed at 12 cm distance from the light source using a Rayonet photochemical reactor. Vesicle dispersions were thoroughly purged with nitrogen to exclude oxygen. Redox polymerization was carried out by adding 10 µL of freshly prepared aqueous solution of ammonium persulfate and 4 µL of N,N,N′,N′tetramethylenediamine (TEMED) to oxygen-free vesicle dispersions. The polymerization progress was monitored by recording spectra on 1H NMR at different intervals. Stability of Polymerized Vesicles. Dispersions of polymerized vesicles were quickly frozen by immersing them in a dry ice/ isopropanol bath and subjected to high vacuum using a Labconco freeze-dry system (model freezone 4.5). Polymerized, freeze-dried vesicles from 1 were resuspended in water with intermittent vortex mixing and examined by electron microscopy. A portion of freeze-dried sample was treated with 95% ethanol for 2 h. The ethanol was removed under a gentle stream of dry nitrogen, and

Langmuir, Vol. 19, No. 16, 2003 6403 the sample was dried for 2 h under high vacuum and then redispersed in water before examining under the transmission electron microscope. Encapsulation and Activity of Phosphotriesterase in Vesicles. Encapsulation. For the encapsulation of the enzyme, a slightly modified protocol was used. Freeze-dried, unpolymerized vesicles were used for the enzyme encapsulation experiment. PTE (200 µg) dissolved in 2 mL of CHES buffer (pH 8.6) was added to 4 mg of powdered, freeze-dried vesicles from a 1/DPPC (80:20 w/w) mixture. The vesicles were incubated at 40 °C and brought to suspension with occasional vortex agitation while maintaining the dispersion temperature to produce a stable vesicle dispersion. In the meantime, 4.0 g of Sephadex gel (G 50-150) was soaked in deionized water, degassed, and poured into a glass column to form a column of gel (20 cm × 1 cm). The Sephadex column was primed with CHES buffer before loading the vesicle-enzyme dispersion. Exogenous enzyme was separated from the vesicle-encapsulated enzyme by gel filtration. A 0.5 mL volume was collected in each fraction, and the presence of vesicles and the enzyme was confirmed by monitoring PTE at 279 nm and vesicles at 400 nm. The gel-filtered enzyme-encapsulated vesicles were polymerized by exposing the vesicles to UV irradiation for 5 min after the addition of radical initiator AAHP. Polymerization was confirmed by TLC analysis. Polymerized vesicles were used without gel filtration. Enzymatic Activity of PTE Encapsulated in Vesicles. From the combined enzyme-positive vesicle fractions collected from gel filtration, a 100 µL aliquot was withdrawn and placed in a quartz cuvette. A 500 µL, 100 µM methyl parathion (MPT) solution in 25% aqueous methanol was added to the vesicle dispersion. The contents in the cuvette were thoroughly mixed, and the generation of para-nitrophenol (pNP) was monitored for 120 s by continuously recording absorbance at 405 nm. Based on the production of pNP over time, enzyme velocity was measured. A similar protocol was used for determination of enzyme activity in polymerized vesicles. As a control, the activity of enzyme dissolved in buffer was determined before and after exposing to UV irradiation for 5 min.

Results and Discussion The goal of this study is to make stable vesicles by crosslinking their phospholipid building blocks. Stable morphologies including vesicles are the key for developing novel applications. Our ongoing efforts on preservation of enzyme activity under harsher environments led us to explore vesicles as potential carriers for active enzymes.27 One of the criteria for vesicles to be useful is their ability to polymerize quickly and effectively at room temperature so that fragile molecules including enzymes could be captured inside of polymerized capsules. For the current study, we selected two phosphatidylethanolamines consisting of 1,2-dilauroyl and -dipalmitoyl acyl chains to incorporate polymerizable functionality to the amine. Polymerization in the headgroup region was considered to facilitate efficient polymerization caused by activation of monomers by unrestricted contact with water-soluble radical initiator. Selection of palmitoyl or lauroyl acyl chains was based on the common occurrence of their phosphocholine analogues in biological membranes. Both lipids, 1 and 2, were dispersible in water. Their dispersibility in an aqueous medium is enhanced with a rise in the pH of the dispersion medium to 8.0 and above. While a higher pH (9.4 or higher) of the medium further facilitated their dispersion, some hydrolysis was observed upon prolonged contact with the medium at higher temperatures. For enzyme encapsulation purposes, a mixed lipid system involving 1 with DPPC was used for improving the dispersibility of lipid in water. Lipid Synthesis. As illustrated in Scheme 1, the phospholipid synthesis was performed by a straightfor(27) Markovac, A.; LaMontagne, M. P. J. Med. Chem. 1980, 23, 11981201.

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Scheme 1. Synthesis of 1,2-Diacyl-sn-glycero-3phospho-N-(2-hydroxyethyl)-3,5-divinylbenzamidea

a Reagents: (a) N-bromosuccinimide, (b) triphenylphosphine/ acetone, (c) formaldehyde (37 wt %)/NaOH/reflux, (d) 1-hydroxybenzotriazole/1-(3-dimethylaminopropyl)-3-ethylcarbodiimide/CHCl3.

ward, high-yield reaction between phosphatidylethanolamine and DVBA monomer. Formation of activated carboxylic acid involving EDC and its coupling to primary amines in the presence of 1-hydroxybenzotriazole is a wellestablished technique. The overall yields of 1 or 2 including the steps involved in the synthesis of DVBA fell in the range of 27%. For an unknown reason, the yields of ylide formation were low (45%) in our hand and need optimization. The bromination step involving reaction of Nbromosuccinimide (NBS) with 3,5-dimethylbenzoic acid leads to monosubstituted bromomethyl product and proceeds further substituting an additional proton producing gem-dibromomethyl product. Careful monitoring of the course of the reaction helped alleviate this problem. We used the NMR technique to monitor the conversion of methyl groups (2.36 ppm) to bromomethyls (4.49 ppm) to optimize reflux time. A previous report relied on protecting the carboxylic acid to perform the NBS reaction.28 We have eliminated this step without compromising the yields. Vesicle Formation and Characterization. The dispersion behavior of polymerizable phospholipids 1 and 2 was studied by hydrating them in a series dispersion medium. The presence of a hydrophobic aromatic ring and the hydrogen bond promoting amide group in the lipid headgroup present a unique situation that may influence the lipid self-assembly. The effect of pH on the formation of vesicles was examined in water and in buffers of varying pH values at 9.3, 8.6, 7.0, and 4.0. Vesicle formation was visually observed after hydrating the lipids at 70 °C followed by occasional vortex mixing. The suspension was visually observed for the ability of vesicles to stay as suspension. The vesicle suspension was most stable at basic pH. Stability was observed in the following order: 9.3 > 8.6 >7.0 > 4.0. At pH 9.3, lipid dispersed easily into the aqueous phase producing vesicles within 30 min. Vesicles prepared at this pH produced decent spectral features in the NMR spectrum allowing the noninvasive role of this technique to study polymerization extent in the vesicles. Deionized water has a pH that is suitable to aid in the formation of polydispersed vesicles by vortex mixing at 50 °C and was used in most of the vesicle preparations. Additional proof about strong lipid-lipid interaction at pH 9.3 came from their UV spectrum. Lipid 1 dissolved in organic solvent or dispersed in water showed a strong absorption maximum at 220 nm. In buffer at pH 9.3, it showed an absorption peak at 200 nm, a hypsochromic shift presumably caused by strong lipid-lipid interaction in dispersions.29 (28) Liu, S.; O’Brien, D. F. J. Am. Chem. Soc. 2002, 124, 6037-6042. (29) Singh, A.; Tsao, L. I.; Markowitz, M.; Gaber, B. P. Langmuir 1992, 8, 1570-1577.

Figure 2. 1H NMR spectra of vesicles from 1 in D2O at 25 °C as a function of UV irradiation time. The proton signals denoted as (a), (b), and (c) are the vinyl protons of DPPE-DVBA.

Figure 3. 1H NMR spectra of vesicles from 2 in D2O at 25 °C as a function of UV irradiation time. The proton signals denoted (a), (b), and (c) are the vinyl protons of DLPE-DVBA.

Polymerization. Figures 2 and 3 show the NMR spectral traces collected after UV irradiation at different time intervals. Polymerization of vesicles was performed in D2O. Vesicles produced from lipids 1 and 2 were polymerized at room temperature by the addition of watersoluble radical initiator, AAHP, followed by ultraviolet irradiation at 254 nm. A reduction in the intensity of proton resonance peaks due to vinyl protons at 6.6, 5.8, and 5.3 ppm was monitored as a function of time to evaluate the extent of polymerization. Figure 2 shows that the vesicle from lipid 1 containing 16-carbon long, dipalmitoyl chains completely polymerized within 5 min of UV irradiation. On the other hand, Figure 3 showed that only 48% lipid 2 in the vesicles polymerized. An additional 15 min of UV exposure brought the polymerization extent to a technologically acceptable number, 95%. Thereafter it took another 15 min for complete disappearance of proton signals at 6.6, 5.8, and 5.3 ppm to be observed indicating completion of polymerization. Previously, we have reported a styryl system containing one vinyl moiety, which took 90 min of UV irradiation at 25 °C to achieve complete polymerization.26 An incomplete polymerization was observed when polymerization was initiated either with AAHP (60 °C) or UV exposure (up to 5 h) alone irrespective of polymerization time. Redox polymerization of 1 in vesicles carried out by the addition of redox initiators ammonium persulfate and tetramethylene ethylenediamine at room temperature led to the complete consumption of monomers in just 1 h as indicated by TLC analysis at different time intervals and confirmed by 1H NMR. A previous report on polymerization of lipids containing β-nitrostyrene25 in their headgroup region showed UV irradiation alone as an effective means of polymerization. The focus of the study is on the formation of stable capsules by polymerization of vesicles under mild conditions including least exposure to UV. Since 1 polymerized

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Figure 4. Transmission electron micrograph of vesicles of 1 dispersed in water by sonicating the lipid at 50 °C for 1 h: (A) nonpolymerized; (B) polymerized. The bar represents 500 nm.

Figure 5. Transmission electron micrograph of polymerized vesicles from 1 after freeze-drying and redispersion: (A) freeze-dried and redispersed in water; (B) freeze-dried, washed with 95% ethanol, and redispersed in water. The bar represents 250 nm. Table 1. Thermotropic Behavior of Hydrated Phospholipids heating transition (°C) phospholipid(s)

nonpolymerized

polymerized

cooling transition (°C) nonpolymerized

1

37.5

29.4

32.8

1 + DPPC DPPC

35.2 43.2

31.6

27.9 41.3

polymerized 25.9 (main), 28.8 not observed

within 5 min, further studies were conducted focusing on DPPE-DVBA only. Thermotropic Behavior of Hydrated Phospholipids. Table 1 shows the results obtained by scanning the hydrated lipids using a differential scanning calorimeter. Lipid was directly weighed into the DSC pans for nonpolymerized samples, and for polymerized samples freeze-dried samples of polymerized vesicles were used. Scans on DPPC are reported in the table being the route lipid for DPPE-DVBA and used for comparison. Lipid 1 showed a sharp chain-melting transition temperature at 37.5 °C and an even sharper transition peak at 32.8 °C upon cooling. These transition temperatures are very close to those of its route lipid, DPPC, which has 43.2 and 41.3 °C heating and cooling transition temperatures, respectively. The polymerized vesicles showed cooling and heating transition temperatures at 29.4 and 25.9 °C, respectively, which were about 7 °C lower than those of their nonpolymerized counterpart. Closeness in transition temperatures between polymerized and nonpolymerized lipid bilayers indicates that the membrane properties are not affected by polymerization. An 80:20 (w/w) mixture of 1 and DPPC showed broad, single transition peaks in the heating and cooling cycle centering at 35.2 and 27.9 °C. No transition due to individual lipids was observed. These results are indicative of complete mixing of two lipids upon hydration. Visualization. Figure 4 provides visual proof of the presence of vesicle structures in sonicated lipid suspensions hydrated in water. Transmission electron microscopy (TEM) images clearly show the presence of almost uniform vesicles in the dispersion. Both nonpolymerized (Figure 4A) and polymerized (Figure 4B) vesicles from lipid 1 showed an average diameter of 70 nm. Because vesicle

Figure 6. Simultaneous monitoring of phosphotriesterase and vesicles separated by gel filtration. Closed circles (b) represent the absorption at 400 nm due to vesicles, and open circles (O) represent the absorption at 279 nm due to enzyme encapsulated in vesicles.

sizes differ from preparation to preparation, vesicles from a single batch were used for comparing their size before and after polymerization. The vesicles shown in Figure 4A,B are for making a comparison between polymerized and nonpolymerized vesicles. The similarity in vesicle size before and after polymerization demonstrates the utility of the polymerization approach for making stable structures. Stability. The mechanical and chemical stability of polymerized vesicles was determined by TEM after subjecting them to conditions under which regular vesicles lose their structural integrity. Figure 5 shows TEM images of polymerized vesicles redispersed after a freeze-drying cycle. Polymerized vesicles from 1 were prepared in water and freeze-dried to obtain a solid mass. Part of these vesicles were redispersed in water and brought into dispersion by vortex mixing. Figure 5A shows vesicles averaging 36 nm in diameter with a size range between 31 and 55 nm. Vesicles from nonpolymerizable analogues usually produce aggregated large structures. The other part of the vesicles was treated with 95% ethanol and

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Figure 7. Transmission electron micrograph of PTE encapsulated vesicles made from 1/DPPC (80:20 w/w) in CHES buffer: (A) before polymerization; (B) polymerized vesicles. The bar represents 250 nm.

then redispersed in water after removing ethanol by evaporation under a gentle stream of nitrogen. As shown in Figure 5B, intact vesicles were observed ranging from 25 to 55 nm. Some vesicles show aggregation, which was due to weak electrostatic interaction between aromatic moieties available on vesicle surfaces containing polydivinyl benzene moieties. Interactions between aromatic rings are well-known. Agitation assisted through a common laboratory bath sonicator separated such aggregated vesicles. Sustainment of vesicle turbidity after the addition of ethanol has been previously demonstrated as a proof for vesicle stability.30 Encapsulation and Activity of Phosphotriesterase in Vesicles. For the enzyme encapsulation studies, we decided to use DPPC mixed with 1 to aid efficient vesicle formation. DSC studies suggested that DPPC is miscible with 1. Out of several mixed lipid compositions examined for the ease of vesicle formation, two compositions comprised of 1 and DPPC in 80:20 and 70:30 w/w ratios produced vesicle dispersions easily by gentle vortex mixing of the hydrated sample. An 80:20 (1/DPPC) lipid mixture was used for this study. Vesicles for enzyme encapsulation were prepared by sonicating the vesicles until a constant turbidity at 400 nm was observed. The small unilamellar vesicles thus produced were freeze-dried and redispersed after addition of enzyme dissolved in CHES buffer at pH 8.6. Hydration at 50 °C and vortex mixing produced small vesicles. Vesicles produced from a freeze-thaw cycle are reported to form uniform dispersions with high encapsulation efficiency.31 Formation of giant unilamellar vesicles is reported by partitioning lipid in a chloroformmethanol-water medium and removing the organic solvent using a rotary evaporator.32 Giant vesicles are ideal to encapsulate higher concentrates of large molecules. We did not use that technique because of deactivation of enzyme in organic solvent. The enzyme PTE was selected for the current study because of its potential application in pesticide detoxification.33 Figure 6 shows a profile of vesicle separation from the exogenous protein by monitoring each fraction collected during gel filtration by the UV-vis spectrophotometer. Enzyme shows an absorption maximum at 279 nm, and the vesicles were monitored at 400 nm. A graph plotting the absorptions at 279 and 400 nm against fractions shows the presence of PTE in the fractions containing vesicles (Figure 6). Figure 7 illustrates the morphological features of enzymeencapsulated vesicles collected after removing the excess enzyme by gel filtration as well as after subjecting to the polymerization process. Vesicles of ∼46 nm diameter were observed in both cases. (30) Lee, Y.; Stanish, I.; Rastogi, V.; Cheng, T. C.; Singh, A. Langmuir 2003, 19, 1330-1336. (31) Coldren, B.; Zasadzinski, J. A. Langmuir 2002, 18, 284-288. (32) Moscho, A.; Orwar, O.; Chiu, D. T.; Modi, B. P.; Zare, R. N. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 11443-11447. (33) Tuovinen, K.; Kaliste-Korhonen, E.; Raushel, F. M.; Hanninen, O. Fundam. Appl. Toxicol. 1994, 23, 578-584.

Figure 8. Bar graph showing enzyme activity of phosphotriesterase encapsulated in vesicles before and after polymerization. The activity of free enzyme before and after UV irradiation is shown for comparison. Nonirradiated (light gray); UV irradiated (dark gray).

Enzyme Activity. Both polymerized and nonpolymerized PTE-encapsulated vesicles were brought in contact with pesticide, MPT, and the progress of MPT hydrolysis was observed. Chemical agents have been reported to permeate through membranes of enzyme-encapsulated vesicles.34 In this system, the presence of aromatic moieties on the vesicle surface provides an added advantage by attracting MPT molecules to the vesicles. The generation of hydrolysis product, pNP, was monitored at 405 nm over a time period of 120 s, and the velocity of enzyme catalysis was calculated. The results are compiled in Figure 8. The gel-filtered, enzyme-encapsulated vesicles before polymerization showed an initial catalytic velocity of 1.8 µM/s. However, the catalytic velocity calculated after polymerization by UV irradiation was 0.95 µM/s (52% retention of activity). To determine the extent of enzyme deactivation by UV irradiation, free PTE in solution was subjected to 5-min UV exposure similar to that of vesicles. To make a fair comparison, the enzyme was further diluted to match its activity with that of vesicle-encapsulated enzymes. As shown in Figure 8, the catalytic activity of free enzyme was reduced from 1.5 to 0.32 µM/s, a significant 80% decrease in the velocity. A higher retention of enzyme activity may be due to the presence of lipid bilayers and monomers, which blocked the UV light reaching encapsulated PTE. Experiments are in progress to minimize the deactivation of PTE during polymerization by changing the vesicle size and lamellarity. Conclusions Headgroup-polymerizable phospholipids derived from 1,2-dipalmitoyl- and 1,2-dilauroyl-sn-glycero-3-phosphoethanolamine consisting of divinylbenzoyl units produced stable vesicles in water and at basic pH. Efficient polymerization of the vesicles at room temperature involved a combination of water-soluble free radical (34) Petrikovics, I., et al. Toxicol. Sci. 2000, 57, 16-21.

Nanocapsules from Polymerizable Phospholipids

initiator and UV irradiation producing stable vesicles. Efficient polymerization by a 5-min UV exposure made the system technologically attractive for encapsulation of fragile molecules such as enzymes with retention of the bulk of their catalytic activity. Bilayers of polymerized vesicles protected the enzyme phosphotriesterase entrapped within the vesicle core from deactivation due to UV exposure. Differences in the polymerization behavior of lipids 1 and 2 are interesting, and further investigations are in progress. These findings have implications for the

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applicability of encapsulated polymerized vesicles in the development of multicomponent bioreactors. Acknowledgment. Financial support for this work from the Office of Naval Research through an NRL base program is acknowledged. We thank Professor Frank Raushel (Texas A&M) for the generous gift of the enzyme phosphotriesterase. G.L. is an NRC research associate, and Y.L. is an ASEE fellow. LA034434U