Amphiphilic Block Copolymer Nanotubes and Vesicles Stabilized by

Apr 26, 2007 - A. Jofre*, J. B. Hutchison, R. Kishore, L. E. Locascio, and K. Helmerson ... Ruud J. R. W. Peters , Madhavan Nallani , Roeland J. M. No...
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J. Phys. Chem. B 2007, 111, 5162-5166

Amphiphilic Block Copolymer Nanotubes and Vesicles Stabilized by Photopolymerization A. Jofre,*,† J. B. Hutchison,‡ R. Kishore,† L. E. Locascio,‡ and K. Helmerson† Physics Laboratory, National Institute of Standards and Technology, 100 Bureau DriVe, Gaithersburg, Maryland 20899, and Chemical Science and Technology Laboratory, National Institute of Standards and Technology, 100 Bureau DriVe, Gaithersburg, Maryland 20899 ReceiVed: February 22, 2007; In Final Form: March 18, 2007

We report on a new method to stabilize nanotube and vesicle structures created from amphiphilic diblock copolymers by means of photopolymerization. Cross-linking with UV light exposure minimizes fluid disruption during stabilization. Additionally, the spatial control afforded by focusing or masking the initiating light source enables stabilization of distinct segments of individual nanostructures. This contribution demonstrates (1) that vesicles and nanotubes formed from poly(ethylene oxide)-block-polybutadiene are stabilized by exposure to UV light in the presence of a water-soluble photoinitiator and (2) that new nanotube geometries can be constructed by means of spot-curing, and (3) it reveals an application for photopolymerized nanotubes by showing electrophoresis of DNA through a UV-stabilized nanotube.

The assembly of amphiphilic molecules, such as phospholipids or block copolymers, into vesicles provides an avenue to construct tubes with sub-micron diameters and exquisitely welldefined chemistry at the membrane surface. Nanotubes and nanotube networks have been constructed from liposomes1-4 and polymer vesicles (i.e., polymersomes),5-9 by directly pulling on the membrane with a micropipette tip or optical tweezers. A number of review articles describe the evolution of knowledge associated with the synthesis and assembly of amphiphilic (macro)molecules, as well as the characterization, manipulation, and application of the resulting micelles, vesicles, and tubes.2,6,10-15 The utility of nanotubes and vesicles is often limited by the fragility of the fluid membranes. Therefore, a crucial step in developing applications for nanostructures is stabilizing the organized amphiphilic (macro)molecules. Although it is possible to cross-link phospholipids nanostructures by functionalizing them with polymerizable groups, adding cross-linking components after assembly, or templating nonamphiphilic monomers,16 block copolymers are advantageous for two specific reasons. First, the concentration of polymerizable groups can be much higher. Second, the associated tubes and vesicles are often more mechanically and chemically robust than structures assembled from small molecule amphiphiles.17,18 Commercially available block copolymers with poly(ethylene oxide) (PEO) and polybutadiene (PB) blocks yield mechanically robust nanostructures with membranes containing unsaturated moieties that can be converted into covalent bonds between adjacent molecules. Previously, the cross-linking reaction in polymersomes formed from PB-containing block copolymers have been stabilized by inducing cross-linking via a redox initiation system.18 However, the addition of the redox initiator components moves and deforms nanotubes and disrupts networks of vesicles and tubes, so a practical alternative method for initiating cross-linking is highly desirable. * Corresonding autor. † Physics Laboratory, National Institute of Standards and Technology. ‡ Chemical Science and Technology Laboratory, National Institute of Standards and Technology.

Cross-linking with UV light has significant advantages over stabilizing structures by addition of a redox initiating solution. Upon photopolymerization, nanotubes and vesicles can be minimally disturbed. Additionally, the rate of initiation and degree of polymerization can be controlled by shuttering and/ or tuning the intensity of the initiating light source. Finally, the spatial control afforded by focusing or masking the initiating light source offers a dramatic advantage for stabilizing distinct segments of individual nanostructures and/or well-defined regions of a network of tubes and vesicles. Only a few previous demonstrations of photopolymerizing block copolymers to stabilize nanostructures have been reported, and the methods require synthesis of amphiphiles containing specific crosslinkable repeat units or end groups.19,20 We report a facile technique to photopolymerize commercially available block copolymers that readily assemble into vesicles and nanotubes. We also provide some insight into the cross-linking chemistry, and reveal an application for photopolymerized nanotubes by demonstrating electrophoresis of DNA through a stabilized nanotube integrated with a spherical vesicle reservoir. As a precursor to forming nanotubes and for polymerization experiments, we made giant polymer vesicles (i.e., diameter greater than 1 µm) using an electroformation technique.7,17,21 Two platinum electrodes, spaced by 3 mm, were coated with the amphiphilic block copolymer, polybutadiene(1800 Da)block-poly(ethylene oxide)(900 Da) with 94% 1,2- (i.e., vinyl) groups (PB-b-PEO or B35-b-EO22; Polymer Source Inc., Dorval, PQ, Canada), shown below. The electrodes were immersed in DI water and an oscillating voltage was applied (10 V at 10 Hz) for 12 or more hours

After collecting the vesicle suspension from the electroformation chamber, we added an aqueous photoinitiator solution containing a mass fraction of 0.5% 1-[4-(2-hydroxyethoxy)phenyl]-2-hydroxy-2-methyl-1-propane-1-one (I-2959; Ciba Spe-

10.1021/jp071503v CCC: $37.00 © 2007 American Chemical Society Published on Web 04/26/2007

Amphiphilic Block Copolymer Nanotubes and Vesicles

Figure 1. Time sequences showing effects of a focused UV pulse (optical scalpel) on polymersomes. Time t ) 0 denotes the time the pulse is applied. Top row: vesicle is not cross-linked. The membrane vibrates and reorders within 6 s. Bottom row: vesicle is cross-linked. The initiator mass fraction used to cross-link the vesicle is 0.05% and the exposure time is 60 s at an intensity of 18 mW/cm2 in a 1 mm deep chamber, which corresponds to ∼4 µmol/L activated photoinitiator. There are no fluid-like undulations in the membrane, and the membrane does not reorder itself. A hole created by the optical scalpel remains in the vesicle indefinitely.

cialty Chemicals, Basel, Switzerland), such that the photoinitiator was diluted to a mass fraction of 0.01-0.05%. The samples were then placed in a 1-mm deep chamber, and exposed to about 18 mW/cm2 of 365-nm UV light for 10 to 180 s. The light source used in these experiments was Omnicure Series 1000 UV/vis spot curing system (EXFO, Mississauga Ontario). We confirmed the stabilizing effect of photopolymerizing the polymersome membrane by monitoring the membrane fluidity. When a polymer vesicle is not stabilized, a disrupted membrane will rapidly reorder itself such that the hydrophobic region is not exposed to the aqueous buffer. Figure 1 reveals a distinct difference between the fluidity of uncross-linked and crosslinked polymersomes. The aqueous buffer used to cross-link the polymersomes consists of DI water with 0.05% mass fraction of photoinitiator. We used a 10 ns, 355 nm focused UV laser pulse (i.e., an optical scalpel7) to create localized damage in the vesicle membrane. In the case of the uncross-linked vesicle displayed in the top sequence, the membrane reorders rapidly, and a fully closed vesicle reforms within seconds or less. In contrast, when the optical scalpel breaks the membrane of a cross-linked vesicle, the membrane does not reorder, and the defect remains in the vesicle indefinitely. The mass fraction of photoinitiator used to cross-link the vesicle shown in Figure 1 was 0.05% and the exposure time was 60 s at an intensity of 18 mW/cm2 in a 1 mm deep chamber. We have also observed that vesicles photopolymerized under these conditions maintain their integrity under a single dehydration-rehydration cycle. Figure 2 displays qualitative and quantitative evidence of cross-linking during photopolymerization of vesicles. The images of polymersomes in Figure 2 reveal typical changes in morphology that occur upon exposure to UV light. The first figure in the sequence, Figure 2a, is unexposed. After short exposure times, as in the 10 s shown in Figure 2b, the membrane flexibility is reduced, which means that nanotubes cannot be pulled out from the vesicle at this stage in the photopolymerization. We also observe at this stage that any vesicles that were asymmetric prior to exposure (which is often the case) become spherical. The image in Figure 2c was captured at a moderate exposure time (20 s) when the membrane exhibits the level of rigidity demonstrated in Figure 1, but significant morphological changes (e.g., wrinkles and creases) are not apparent. After an exposure of 60 s, we begin to observe morphological changes in many of the vesicles as shown in Figure 2d. It should be

J. Phys. Chem. B, Vol. 111, No. 19, 2007 5163 noted that at this stage of photopolymerization, there is always some portion of vesicles in the sample that show no significant morphological changes. Finally, Figure 2e (180 s) is a typical image of a polymersome after a long period of exposure. The creases and wrinkles that form in the membrane have been attributed to changes in osmolarity that occur upon polymerization of the double bonds in the PB block.18 We employed 1H NMR spectroscopy as a first step in analyzing the changes in molecular structure of the PB-b-PEO that occur during exposure of vesicles to UV light. In short, 30-50 mL of solution containing vesicles suspended in water after UV exposure was lyophilized; the residue was resuspended in deuterated chloroform (though not necessarily dissolved since the exposed vesicles are most likely cross-linked), and the relative concentration of unreacted double bonds was measured using a 270 MHz NMR spectrometer. The conversion of double bonds was determined from the area beneath the peaks (Figure 2, bottom graph) centered at 4.9 and 5.4 ppm normalized by the signal at 3.6 ppm from the protons in each PEO block. The signal at 4.9 ppm is from the two protons at the terminus of each unreacted vinyl group (-CHdCH2) and the signal at 5.4 ppm is from the individual protons present at the other end of each vinyl group (-CHdCH2) and both ends of each 1,4unsaturated group (-CHdCH-). Other peaks apparent in the spectra correspond to photoinitiator (4.0 and 4.2 ppm), methine and methylene protons adjacent to alkene carbons in vinyl and 1,4-groups, respectively (∼2.1 ppm, broad), methylene protons from vinyl PB units (1.2 ppm, broad), and methyl protons from the hydrophobic terminus of PB-b-PEO (0.8 ppm). Finally, the narrow signal at 1.6 ppm (in the lower spectrum of Figure 2) is likely residual water, and the broad signal at 1.6 ppm indicates the possible presence of oxidation products of polybutadiene (including carboxylic acids, aldehydes, and ethers). Specifically, a signal at 1.6 ppm corresponds to methylene protons in the β-position to typical acids, aldehydes, and ethers. The plot in Figure 2 separately describes the conversion of vinyl and 1,4-double bonds in vesicle samples exposed to different doses of UV light and initial concentrations of photoinitiator. By plotting the conversion as a function of the cumulative amount of activated photoinitiator, the data from samples with different levels of photoinitiator collapse to two distinct trends (for vinyl and 1,4-double bonds). Equation 2 describes the relationship between exposure time and cumulative activated initiator concentration, [PI]activated:

[PI]activated ) [PI]o{1 - exp(-2fIot)}

(2)

where t is the exposure time, [PI]o the initial concentration of initiator, Io the intensity of the light,  the extinction coefficient of the initiator, and f the efficiency.22 [PI]o is an experimental variable; Io was measured to be ∼18 mW/cm2 (which converts to ∼5.5 × 10-5 mol photons cm L-1-s-1); and the product of  and f for the initiator used in this study is ∼1 L mol-1 cm-1.23 The primary observations made from the data presented in Figure 2 are that the rate of double bond consumption decreases after activation of ∼4 µmol of initiator per liter of vesicle suspension, and the maximum conversion is ∼15%. Also, the apparent reactivity of 1,4-double bonds is significantly higher than the reactivity of vinyl bonds. Finally, the conversion of vinyl bonds in the unexposed vesicle samples is nonzero. In fact, the average conversion is approximately 5%, which is onethird of the maximum conversion of vinyl bonds. We also note the large spread in the 1,4-data of unexposed vesicles, which can be used to estimate the standard deviation of each of the points on the graph.

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Figure 2. Top: time sequence of a polymer vesicle exposed to UV light in a buffer containing 1.2 mmol/L photoinitiator. (a) No photoinitiator, no UV exposure. (b) 10 s UV exposure. (c) 20 s. (d) 60 s. (e) 180 s. The dotted lines match the exposure times to the concentration of activated photoinitiator. Middle: conversion of pendant vinyl double bonds (filled symbols) and 1,4-polybutadiene double bonds (open symbols) as a function of the concentration of activated initiator. Bottom: NMR spectra of the unreacted polymer vesicles (dark trace) and photopolymerized vesicle sample (gray trace; 2 µmol/L activated photoinitiator).

In the case of radical chain polymerization, the pendant vinyl bond is more reactive than the 1,4-double bond, which is part of the main chain backbone.24 This fact suggests that the observed disappearance of double bonds is likely not primarily due to chain polymerization. In fact, the more likely mechanism for cross-linking is photo-oxidation.25-28 1,4-double bonds are approximately 5 times more susceptible to oxidation than pendant vinyl groups. Photoinitiation in the presence of oxygen generates a large number of oxygen radicals, which participate in hydrogen abstraction reactions followed by chain transfer and/ or radical-radical combination, leading to cross-linking. Further evidence for oxidation is apparent in the comparison of NMR spectra included in Figure 2. The top spectrum (dark line) is the block copolymer. The gray line was measured from a UVexposed vesicle sample. The broad peak at 1.6 ppm matches the resonance that could be expected for oxidized polybutadiene. Although the preliminary data regarding the molecular structure of the exposed vesicles suggests photo-oxidation as the mechanism for cross-linking, further exploration of the kinetics and mechanism of stabilization are warranted. Using conditions that maximize the PB double bond conversion, yet minimize morphological deformations of the membrane (1.2 mmol/L photoinitiator and 20 s of UV exposure at 18mW/ cm2), we stabilized nanotubes using UV-induced cross-linking. Interest in studying biological macromolecules in highly confined geometries has driven many of the advances in the

fabrication of new nanofluidic devices.29-31 In particular, strong interest in examining the elongation and transport of DNA molecules within nanochannels32-34 and nanotubes35,36 has been motivated by fundamental studies of transport phenomena in biological systems,37 by the elucidation of the effects of entropic forces in highly confined geometries,38 and by the possibility of capturing the macromolecule’s sequence and/or structural information in its elongated configuration.39 Figure 3 demonstrates the transport of λ-DNA through a nanotube stabilized by photopolymerization. The λ-DNA (New England Biolabs, Ipswich, MA) was diluted 1:10 (from an initial concentration of 500µg/mL) with 10 mmol/L Tris-HCl (pH 8.0) and 1 mmol/L EDTA. YOYO-1 dye (Invitrogen, Carlsbad, CA) was intercalated at a ratio of 1 dye molecule to 5 base pairs. The dye was excited with the 488 nm line of an argon-Ion laser for fluorescence imaging. Note that the membranes of the polymer vesicle and nanotube do not contain any dye. The configuration of the experiment is shown in the top panel of Figure 3. Two micropipette tips were used to manipulate the vesicles; one tip was for pulling out the nanotube and the other tip was used to electro-inject the DNA into the vesicle.40,41 Each micropipette tip contained an electrode to generate a field across the nanotube. After inserting the tip containing DNA into a vesicle and pulling a nanotube from the same vesicle with the other micropipette tip, the system was stabilized by 20 s of UV exposure at 18 mW/cm2. Shortly following photopolymerization, a 5 V potential

Amphiphilic Block Copolymer Nanotubes and Vesicles

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Figure 3. Top: experimental setup for the DNA transport experiment. Two micropipette tips, each containing an electrode, were used to manipulate the vesicle and nanotube. One micropipette tip, inserted into the vesicle, was used to inject the λ-DNA into the vesicle, and the other was used to pull out a nanotube. The vesicle, which was suspended in 1.2 mmol/L photoinitiator, was subsequently exposed to 20 s of UV light to stabilize the structure. Once the nanotube and vesicle were cross-linked, an electric field was applied between the tips to electrophorese DNA through the nanotube. Middle sequence: cluster of DNA fluorescently labeled with YOYO-1 dye entering a nanotube. We applied 5 V between the micropipette tips to electrophorese the DNA through the tube. Bottom picture: results of the experiment when the nanotube and vesicle are not cross-linked. As the λ-DNA material fills the nanotube, the vesicle deforms and the tube expands. The dashed lines indicate the original size and shape of the vesicle and nanotube.

was applied to the micropipette tips and the negatively charged DNA was electrophoresed through the nanotube. As shown in Figure 3, the DNA is observed to move into a photopolymerized nanotube without any visible effect on the nanotube diameter. In contrast, a nanotube that was not stabilized by cross-linking expanded to an increasingly larger diameter as DNA solution entered the tube, as shown in the bottom panel of Figure 3. We used a relatively high concentration of DNA (50µg/mL), and therefore, the pressure inside the nanotube increased rapidly as more material entered the tube. The tube that is not cross-linked stretched and deformed under such pressure, while the crosslinked tube maintained its integrity. Photopolymerization facilitates microscopic spot-curing to cross-link well-defined regions of the sample. To demonstrate microscopic spot-curing, we used a continuous-wave 365 nm 10mW diode laser, coupled through the objective lens of the microscope, to create bent nanotubes. A free-standing bend in a bilayer polymer nanotube cannot be created unless the membrane is stabilized by cross-linking, because the selfassembled fluid membrane always rearranges itself to minimize its energy. On the other hand, if one segment of a nanotube is fixed by photopolymerization, then one can reposition the other nanotube segment at ninety degrees from the first, creating a ninety degree bend in the nanotube. This bent region can then be fixed by photopolymerization to create a stable free-standing bent nanotube. Figure 4a schematically illustrates this procedure. Figure 4b reveals bends in nanotubes created by sequentially spot-curing and manipulating the orientation of the nanotube with a micropipette tip. Specifically, the bend was made by first cross-linking one segment, which makes it rigid. Subsequently, we used a micropipette tip to reposition the uncross-linked end of the tube, then cross-linked the next segment. This process was repeated to make the second bend. In this manner, complex geometries and configurations of stable nanotubes can be constructed.

Figure 4. Bends in nanotubes were created by cross-linking part of the tube, then bending the remaining flexible region of the tube with a micropipette tip. The bent shape is then spot-cured, and the process is repeated to create a second kink in the tube. A CW UV laser, coupled into the microscope objective, was used for spot-curing small regions of the nanotube. Figure 4a is a diagrammatic representation of how the bends in Figure 4b were created. After the nanotube is pulled out of the vesicle with a micropipette tip, a portion of the nanotube near the vesicle is stabilized by UV exposure, as shown in the top panel of Figure 4a. This creates a new pivot point in the nanotube, so that when the micropipette tip is repositioned, the nanotube bends as illustrated in the middle panel of Figure 4a. At this point, a second portion of the nanotube is exposed to UV light to stabilize the bent structure. In the bottom panel of Figure 4a, the micropipette tip is once again repositioned and the bends remain stably in the nanotube. The experimental equivalent of this picture is presented in the top panel of Figure 4b. The procedure is repeated to produce the second bend shown in the lower panel of Figure 4b.

In summary, polymersome membranes containing unsaturated groups were cross-linked by UV exposure in the presence of a water-soluble photoinitiator. We also used photopolymerization

5166 J. Phys. Chem. B, Vol. 111, No. 19, 2007 to stabilize nanotubes pulled from polymersome membranes. The distinct advantage of spot curing (or masking a flood exposure) to cross-link nanotubes in well-defined regions was demonstrated by selectively stabilizing individual segments of a single nanotube to create permanent kinks and bends. As an application of our technique for photopolymerization of supermolecular structures from the self-assembly of amphiphilic diblock copolymers, we demonstrate the electrophoresis of DNA in a UV cross-linked nanotube. Acknowledgment. This work was supported by the ONR (Office of Naval Research). Certain commercial equipment, instruments, or materials are identified herein to foster understanding. Such identification does not imply recommendation or endorsement by the National Institute of Standards and Technology, nor does it imply that the materials or equipment identified are necessarily the best available for the purpose. References and Notes (1) Evans, E.; Bowman, H.; Leung, A.; Needham, D.; Tirrell, D. Biomembrane templates for nanoscale conduits and networks. Science 1996, 273 (5277), 933-935. (2) Karlsson, M.; Davidson, M.; Karlsson, R.; Karlsson, A.; Bergenholtz, J.; Konkoli, Z.; Jesorka, A.; Lobovkina, T.; Hurtig, J.; Voinova, M.; Orwar, O. Biomimetic nanoscale reactors and networks. Annu. ReV. Phys. Chem. 2004, 55, 613-649. (3) Karlsson, M.; Sott, K.; Davidson, M.; Cans, A. S.; Linderholm, P.; Chiu, D.; Orwar, O. Formation of geometrically complex lipid nanotubevesicle networks of higher-order topologies. Proc. Natl. Acad. Sci. U.S.A. 2002, 99 (18), 11573-11578. (4) Brazhnik, K. P.; Vreeland, W. N.; Hutchison, J. B.; Kishore, R.; Wells, J.; Helmerson, K.; Locascio, L. E. Directed growth of pure phosphatidylcholine nanotubes in microfluidic channels. Langmuir 2005, 21 (23), 10814-10817. (5) Grumelard, J.; Taubert, A.; Meier, W. Soft nanotubes from amphiphilic ABA triblock macromonomers. Chem. Commun. 2004, (13), 1462-1463. (6) Hamley, I. W. Nanoshells and nanotubes from block copolymers. Soft Matter 2005, 1 (1), 36-43. (7) Reiner, J. E.; Wells, J. M.; Kishore, R. B.; Pfefferkorn, C.; Helmerson, K. Stable and robust polymer nanotubes stretched from polymersomes. Proc. Natl. Acad. Sci. U.S.A. 2006, 103 (5), 1173-1177. (8) Stewart, S.; Liu, G. Block copolymer nanotubes. Angew. Chem., Int. Ed. 2000, 39 (2), 340. (9) Won, Y. Y.; Davis, H. T.; Bates, F. S. Giant wormlike rubber micelles. Science 1999, 283 (5404), 960-963. (10) Antonietti, M.; Forster, S. Vesicles and liposomes: A self-assembly principle beyond lipids. AdV. Mater. 2003, 15 (16), 1323-1333. (11) Discher, D. E.; Eisenberg, A. Polymer vesicles. Science 2002, 297 (5583), 967-973. (12) Forster, S.; Konrad, M. From self-organizing polymers to nanoand biomaterials. J. Mater. Chem. 2003, 13 (11), 2671-2688. (13) Hamley, I. W. Nanostructure fabrication using block copolymers. Nanotechnology 2003, 14 (10), R39-R54. (14) Kita-Tokarczyk, K.; Grumelard, J.; Haefele, T.; Meier, W. Block copolymer vesicles - using concepts from polymer chemistry to mimic biomembranes. Polymer 2005, 46 (11), 3540-3563. (15) Lazzari, M.; Lopez-Quintela, M. A. Block copolymers as a tool for nanomaterial fabrication. AdV. Mater. 2003, 15 (19), 1583-1594. (16) Hotz, J.; Meier, W. Vesicle-templated polymer hollow spheres. Langmuir 1998, 14 (5), 1031-1036. (17) Discher, B. M.; Won, Y. Y.; Ege, D. S.; Lee, J. C. M.; Bates, F. S.; Discher, D. E.; Hammer, D. A. Polymersomes: Tough vesicles made from diblock copolymers. Science 1999, 284 (5417), 1143-1146. (18) Discher, B. M.; Bermudez, H.; Hammer, D. A.; Discher, D. E.; Won, Y. Y.; Bates, F. S. Cross-linked polymersome membranes: Vesicles

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