Direct Printing of Self-Assembled Lipid Tubules on Substrates

Nov 20, 2007 - curved substrates from the recessed channels of the PDMS stamp by bringing the tubule-inked PDMS stamp into contact with these substrat...
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Langmuir 2008, 24, 5113-5117

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Direct Printing of Self-Assembled Lipid Tubules on Substrates Yue Zhao and Jiyu Fang* AdVanced Materials Processing and Analysis Center and Department of Mechanical, Materials, and Aerospace Engineering, UniVersity of Central Florida, Orlando, Florida 32816 ReceiVed NoVember 20, 2007. In Final Form: January 9, 2008 Lipid tubules formed by rolled-up bilayer sheets have shown promise in drug delivery systems, nanofluidics, and microelectronics. Here we report a method for directly printing lipid tubules on substrates. Preformed lipid tubules of 1,2-bis(tricosa-10,12-diynoyl)-sn-glycero-3-phosphocholine are aligned in the recessed channels of a thin poly(dimethylsiloxane) (PDMS) stamp. The aligned lipid tubules then serve as an “ink” for microcontact printing. We demonstrate that two-dimensional (2-D) arrays of aligned lipid tubules can be transferred onto planar, patterned, and curved substrates from the recessed channels of the PDMS stamp by bringing the tubule-inked PDMS stamp into contact with these substrates. We show that the 2-D array of aligned lipid tubules can be transcribed into a 2-D array of aligned silica cylinders through templated sol-gel condensation of tetraethoxysilane.

Introduction Molecular self-assembly is becoming increasingly popular as a “bottom up” approach to synthesize supramolecular architectures.1 One of the most attractive aspects of this approach is the prospect of assembling structures with molecular precision under experimentally straightforward and inexpensive conditions. Recently, there has been great interest in using biomolecules including lipids,2-17 peptides,18-22 proteins,23-24 and DNA25-28 * To whom correspondence [email protected].

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(1) Whitesides, G. M.; Mathias, J. D.; Seto, C. Science 1991, 254, 1312-1319. (2) Spector, M. S.; Selinger, J. V.; Singh, A.; Rodriguez, J. M.; Prince, R. R.; Schnur, J. M. Langmuir 1998, 14, 3493-3500. (3) Matsui, H.; Golongan, B. J. Phys. Chem. B 2000, 104, 3383-3386. (4) John, G.; Masuda, M.; Okada, Y.; Yase, K.; Shimizu, T. AdV. Mater. 2001, 13, 715-718. (5) Spector, M. S.; Singh, A.; Messersmith, P. B.; Schnur, J. M. Nano Lett. 2001, 1, 375-378. (6) Thomas, B. N.; Lindemann, C. M.; Corcoran, R. C.; Cotant, C. L.; Kirsch, J. E.; Persichini, P. J. J. Am. Chem. Soc. 2002, 124, 1227-1233. (7) Singh, A.; Wong, E. M.; Schnur, J. M. Langmuir 2003, 19, 1888-1898. (8) Lee, S. B.; Koepsel, R.; Stolz, D. B.; Warriner, H. E.; Russell, A. J. J. Am. Chem. Soc. 2004, 126, 13400-13405. (9) Masuda, M.; Shimizu, T. Langmuir 2004, 20, 5969-5977. (10) Jean, B.; Oss-Ronen, L.; Terech, P.; Talmon, Y. AdV. Mater. 2005, 17, 728-731. (11) Douliez, J. P.; Gaillard, C.; Navailles, L.; Nallet, F. Langmuir 2006, 22, 2942-2945. (12) Mahajan, N.; Zhao. Y.; Du, T.; Fang, J. Y. Langmuir 2006, 22, 19731975. (13) Zhao, Y.; Mahajan, N.; Fang, J. Y. Small 2006, 2, 364-367. (14) Ambrosi, M.; Fratini, E.; Alfredsson, V.; Ninham, B. W.; Giorgi, R.; Lo Nostro, P.; Baglioni, P. J. Am. Chem. Soc. 2006, 128, 7209-7214. (15) Iwaura, R.; Shimizu, T. Angew. Chem., Int. Ed. 2006, 45, 4601-4604. (16) Motoyanagi, J.; Fukushima, T.; Ishii, N.; Aida, T. J. Am. Chem. Soc. 2006, 128, 4220-4221. (17) Brizard, A.; Aime, C.; Labrot, T.; Huc, I.; Berthier, D.; Artzner, F.; Desbat, B.; Oda, R. J. Am. Chem. Soc. 2007, 129, 3754-3762. (18) Gao, X.; Matsui, H. AdV. Mater. 2005, 17, 2037-2050. (19) Lu, K.; Jacob, J.; Thiyagarajan, R.; Conticello, V. P.; Lynn, D. G. J. Am. Chem. Soc. 2003, 125, 6391-6393. (20) Bong, D. T.; Clark, T. D.; Granja, J. R.; Ghadiri, M. R. Angew. Chem., Int. Ed. 2001, 40, 988-1011. (21) Reches, M.; Gazit, E. Science 2003, 300, 625-627. (22) Hartgerink, J. D.; Beniash, E.; Stupp, S. I. Science 2001, 294, 16841688. (23) Graveland-Bikker, J. F.; Schaap, I. A. T.; Schmidt, C. F.; de Kruif, C. G. Nano Lett. 2006, 6, 616-621. (24) Hou, S.; Wang, J.; Martin, C. R. Nano Lett. 2005, 5, 231-234. (25) Hou, S.; Wang, J.; Martin, C. R. J. Am. Chem. Soc. 2005, 127, 85868587. (26) Ke, Y.; Liu, Y.; Zhang, J.; Yan, H. J. Am. Chem. Soc. 2006, 128, 44144421.

as building blocks to assemble tubule structures because of their applications in drug delivery systems, nanofluidics, sensors, and electronics. Self-assembled lipid tubules have been extensively studied.29-32 However, due to the high aspect ratio, the construction of ordered arrays of lipid tubules on solid substrates still remains challenging. Recently, Orwar and co-workers33 reported the formation of an ordered tube array by wiring the fluidic lipid tubes, which are pulled from lipid vesicles, around microfabricated SU-8 pillars with a micropipet technique. They demonstrated that this lipid tube network was able to support electrophoretic transport of colloidal particles contained in the lipid tubes down to the limit of single particles. Shimizu and collaborators34 described an approach in which a crystalline lipid tube of synthetic cardanyl β-D-glucopyranoside was aligned by microextruding an aqueous dispersion onto a glass substrate with a needle. Brazhnik et al.35 and Dittrich et al.36 studied the direct self-assembly of lipid tubules within microfluidic channels. Bundles of aligned lipid tubules were found to grow along the microchannels. We developed a method that combines microfluidic networks and dewetting to produce two-dimensional (2-D) ordered arrays of aligned 1,2-bis(tricosa-10,12-diynoyl)-sn-glycero-3-phosphocholine (DC8,9PC) lipid tubules on glass substrates.13,37 In this process, preformed individual lipid tubules were pulled into microfluidic networks by the capillary force and then dried on the substrate. We also showed that the alignment and positioning of DC8,9PC tubules on a patterned Au substrate with alternating bare Au stripes and thiol monolayer stripes could be achieved (27) Rothemand, P. W. K.; Ekani-Nkodo, A.; Papadakis, N.; Kumar, A.; Fygenson, D. K.; Winfree, E. J. Am. Chem. Soc. 2004, 126, 16344-16452. (28) Mitchell, J. C.; Harris, J. R.; Malo, J.; Bath, J.; Turberfield, A. J. J. Am. Chem. Soc. 2004, 126, 16342-16343. (29) Spector, M. S.; Selinger, J. V.; Schnur, J. M. In Materials-Chirality; Green, M. M., Nolte, R. J. M., Meijer, E. W., Eds.; Topics in Stereochemistry: Chiral Molecular Self-Assembly, Vol. 24; Wiley: Hoboken, NJ, 2003. (30) Brizard, A.; Oda, R.; Huc, I. Top. Curr. Chem. 2005, 256, 167-218. (31) Shimizu, T.; Masuda, M.; Minamikawa, H. Chem. ReV. 2005, 105, 14011444. (32) Fang, J. Y. J. Mater. Chem. 2007, 17, 3479-3484. (33) Hurtig, J.; Gustafsson, B.; Tokarz, M.; Orwar, O. Anal. Chem. 2006, 78, 5281-5288. (34) Frusawa, H.; Fukagawa, A.; Ikeda, Y.; Araki, J.; Ito, K.; John, G.; Shimizu, T. Angew. Chem., Int. Ed. 2003, 42, 72-74. (35) Brazhnik, K. P.; Vreeland, W. N.; Hutchison, J. B.; Kishore, R.; Well, J.; Helmerson, K.; Locascio, L. E. Langmuir 2005, 21, 10814-10817. (36) Dittrich, P. S.; Heule, M.; Renaud, P.; Manz, A. Lab Chip 2006, 6, 488493. (37) Mahajan, N.; Fang, J. Y. Langmuir 2005, 21, 3153-3157.

10.1021/la703634t CCC: $40.75 © 2008 American Chemical Society Published on Web 03/27/2008

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Figure 1. (a-c) Schematic illustration of the microfluidic technique used to align lipid tubules in the recessed channels of a thin PDMS stamp. (a) Rectangular capillaries were formed by placing a PDMS stamp on a glass substrate. A drop of tubule solution was placed along one of the open ends. (b) Alignment of lipid tubules in the rectangular capillaries by capillary action. (c) The PDMS stamp was removed from the glass substrate before the tube solution-filled capillaries were dried. (d) Optical image of the aligned lipid tubules confined in the recessed channels of the PDMS stamp. The optical image was taken at transmission mode.

by withdrawing the patterned Au substrate from a tubule solution.38 The DC8,9PC lipid tubules were found to selectively adsorb on the bare Au stripes and then were aligned along the stripe direction by a moving contact line during the withdrawing process. Microcontact printing (µCP), which utilizes a molded elastomeric stamp, has been proven to be a useful tool to directly print molecular “inks” such as thiols and silanes on surfaces upon contact to form patterned self-assembled monolayers with submicrometer resolution.39 µCP has also attracted great interest in directly printing macromolecular inks including dendrimers,40 DNA oligonucleotides,41 proteins,42 lipid vesicles,43 polymers,44 and nanoparticles.45 Recently, direct printing of elongated nanostructures on substrates has also been reported. For example, Rogers and co-workers46 directly printed carbon nanotubes on planar and curved surfaces with µCP. Li et al.47 used the fluidicalignment method to align acid-treated carbon nanotubes on a smooth PDMS stamp followed by printing of the aligned carbon tubes onto gold electrodes. Guan and Lee48 reported an approach in which DNA molecules aligned on the protruded regions of a PDMS stamp with shear flow were used as an ink. Highly (38) Zhao, Y.; Fang, J. Y. Langmuir 2006, 22, 1892-1896. (39) Xia, Y.; Whitesides, G. M. Angew. Chem., Int. Ed. 1998, 37, 550-575. (40) Li, H.; Kang, D. J.; Blamire, M. G.; Huck, W. T. S. Nano Lett. 2002, 2, 347-349. (41) Lange, S. A.; Benes, V.; Kern, D. P.; Ho¨rber, J. K. H.; Bernard, A. Anal. Chem. 2004, 76, 1641-1647. (42) Renault, J. P.; Bernard, A.; Bietsch, A.; Michel, B.; Bosshard, H. R.; Delamarche, E.; Kreiter, M.; Hecht, B.; Wild, U. P. J. Phys. Chem. B 2003, 107, 703-711. (43) Mahajan, N.; Lu, R.; Wu, S. T.; Fang, J. Y. Langmuir 2005, 21, 31323135. (44) Wang, M.; Braun, H. G.; Kratzmuller, T.; Meyer, E. AdV. Mater. 2001, 13, 1312-1317. (45) Wu, X. C.; Bittner, A. M.; Kern, K. AdV. Mater. 2004, 16, 413-417. (46) Meitl, M. A.; Zhou, Y.; Guar, A.; Jeon, S.; Usrey, M. L.; Strano, M. S.; Rogers, J. A. Nano Lett. 2004, 4, 1643-1647. (47) Li, S.; Yan, Y.; Liu, N.; Chan-Park, M. B.; Zhang, Q. Small 2007, 3, 616-621. (48) Guan, J.; Lee, L. J. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 1832118325.

ordered arrays of stretched DNA molecules were generated on substrates with µCP. In most of the cases, ink molecules were transferred onto the surfaces from the protrusions of the PDMS stamp during the printing process. In this paper, we present a method in which preformed lipid tubules are aligned in the recessed channels of a PDMS stamp with capillary action. We demonstrate that the aligned lipid tubules in the recessed channels can be transferred onto different types of substrates with µCP to form 2-D arrays of aligned lipid tubules. Through the templated sol-gel condensation of tetraethoxysilane (TEOS), 2-D arrays of aligned lipid tubules can be transcribed into 2-D arrays of aligned silica cylinders. Experimental Section Lipid tubules were prepared by thermal cycling of a 5 mg/mL suspension of DC8,9PC (Avanti Polar Lipids, Alabaster, AL) in ethanol/water (70:30, v/v) from 60 °C to room temperature at a rate of ∼0.5 °C/min.49 The polymerization of the tubule suspension was performed with UV light (254 nm) for 20 min at room temperature. The Au-coated mica with the (111) surface was purchased from Molecular Imaging Inc. Patterned Au thin films were deposited onto Si wafers through a transmission electron microscopy (TEM) grid with square pores with a JEOL vacuum evaporator (model JEE-4X). Poly(dimethylsiloxane) (PDMS) stamps were made from Sylgard 184 (Dow Corning Corp.) by casting the Sylgard agents (the weight ratio of component A to component B was 1:10) against a master with stripe patterns. The PDMS stamps having parallel channels (0.8 µm high and 1.0 µm wide) were then treated with oxygen plasma for 30 s, rendering the polymer hydrophilic due to the formation of a thin silicon oxide layer. The separation of the parallel channels in the PDMS stamp is 4 and 7 µm, respectively. Contact angles of a water droplet (20 µL) on the smooth side of the plasmatreated PDMS stamp were measured with a goniometer (RameHart, Inc.) in air at room temperature. The precision of the contact angle measurement is (3°. TEOS from Aldrich was used as received. The TEOS solution was made by adding 20.6 µL of TEOS into 1.8 (49) Yager, P.; Schoen, P. E. Mol. Cryst. Liq. Cryst. 1984, 106, 371-381.

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Figure 2. (a) Tapping-mode AFM images of parallel aligned lipid tubules with a constant separation of 5 µm. (b) High profile along the line shown in (a). The image was taken in air at room temperature. The PDMS stamp with a separation of 4 µm between parallel channels was used to print the 2-D ordered array. mL of a solution, which was prepared from H2O (50 mL) and aqueous HCl (10 µL). The deposition and condensation of TEOS on the tubule arrays were carried out by exposing them to the TEOS solution at room temperature for a week. An Olympus BX 40 microscope with a digital camera (Olympus C2020 Zoom) and an atomic force microscope (Dimension 3100, Wecco) were used to image the lipid tubules printed on substrates. For atomic force microscopy (AFM) measurements, silicon nitride cantilevers (Nanosensors) with normal spring constants of about 30 N/m and resonant frequencies between 250 and 330 kHz were used. The cantilever was excited just below its resonant frequency. All AFM measurements were performed in tapping mode at a scan rate of 0.5 Hz. Scanning electron microscopy (SEM) measurements of silica-coated tubular arrays were performed on a JEOL 6400F microscope equipped with an energy-dispersive X-ray analyzer at 200 kV.

Results and Discussion Figure 1 illustrates our approach, which is used to align lipid tubules inside the recessed channels of a thin PDMS stamp. In our experiment, a freshly prepared PDMS stamp with parallel channels was placed on a planar glass slide to form hydrophilic rectangular capillaries (Figure 1a). The water contact angles on the glass slide and the plasma-treated PDMS are about 8° and 12°, respectively. A drop (100 µL) of preformed tubule suspension was placed along one of the opened ends of the hydrophilic rectangular capillaries, and lipid tubules were pulled into individual capillaries by capillary action and aligned along the channel direction (Figure 1b). The PDMS stamp was then quickly peeled from the glass slide (Figure 1c). As can be seen from the optical microscopy image shown in Figure 1d, the aligned lipid tubules are kept inside the recessed channels of the PDMS stamp by surface tension after the stamp is removed from the glass substrate. There are no bundles of aligned lipid tubules observed in individual channels. We also note the incomplete filling of some channels. The filling of aligned lipid tubules could be further improved by pulling the preformed tubule solution through the channels with a glass pipet, which connects to a moderate vacuum. It is known that DC8,9PC tubules formed in ethanol/

Figure 3. Tapping-mode AFM images of 3-D cross-bar junctions of aligned lipid tubules printed on a Au-coated mica surface. The images were taken in air at room temperature. The PDMS stamp with a separation of 7 µm between parallel channels was used to print these 3-D cross-bar junctions.

water mixtures have a characteristic length of 20-200 µm. The variation of tubule lengths is also visible for the aligned tubules inside the recessed channels. The PDMS stamp with lipid tubule inks provides a simple approach to directly print lipid tubules onto different types of substrates. By bringing the tubule-inked PDMS stamp into contact with Au-coated mica for 2 h in air at room temperature, we find that the aligned lipid tubules are transferred onto the substrate from the recessed channels of the PDMS stamp. After removal of the PDMS stamp from the substrate, a 2D ordered array of aligned lipid tubules is left behind (Figure 2a). The transfer mechanism of lipid tubules might be a result of the hydrophobicity change of the plasma-treated PDMS stamp surface during the printing process. It has been shown that the hydrophilic glassy silicate layer on the PDMS stamp generated by a brief oxygen plasma treatment is not stable in an ambient environment.50 During the 2 h contacting, the plasma-treated PDMS stamp becomes hydrophobic, and lipid tubules deposit from the low free energy PDMS surface onto the high-energy Au-coated mica. The high profile along the line marked in Figure 2a shows that the height of the aligned lipid tubules on the Au-coated mica is about 400 nm (Figure 2b), which agrees with the diameter of single DC8,9PC lipid tubules. The apparent width of the aligned lipid tubules in the AFM image is broadened by the geometry of the AFM tip. The separation of the parallel-aligned tubules is about 5 µm. (50) Delamarche, E.; Geissler, M.; Bernard, A.; Wolf, H.; Michel, B.; Hilborn. J.; Donzel, C. AdV. Mater. 2001, 13, 1164-1167.

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Figure 4. (a) Optical microscopy image of a 2-D ordered array of aligned lipid tubules printed on a square Au film. (b) Tapping mode AFM image of an aligned lipid tubule across the edge of the Au film. (c, d) High profiles along the lines shown in (b). The optical image was taken at reflection mode. The AFM image was taken at room temperature in air.

After the first layer of aligned lipid tubules is printed, the second layer of aligned lipid tubules can be printed by rotating the tubule-inked PDMS stamp 90° for constructing a 3D crossbar junction of aligned lipid tubules (Figure 3a). In this case, a PDMS stamp with a 7 µm separation between parallel channels was used. The separation of the parallel-aligned lipid tubules in the 3D cross-bar junction is about 8 µm. Some varieties in the tubule diameter are also visible in Figure 3a. As can be seen from Figure 3b the upper tubule bends when it flexes around the lower tubule, leading to a raised area at the junction. There are no breaks observed for the bent upper tubule. The lower tubule is stable on the Au-coated mica surface and not destroyed by the second printing. The printing of a 3D cross-bar junction of aligned lipid tubules with the PDMS stamp with a 4 µm separation between parallel channels was not successful. It is likely that the denser packing of the first layer of aligned tubules with a separation of 5 µm prevents the transfer of the second layers of the aligned tubules due to the repulsive force between two densely packed tubule arrays. Figure 4a shows a 2D ordered array of aligned lipid tubules that are printed on a patterned Au square film with a thickness of ∼250 nm on a SiO2 substrate. As can be seen, a number of parallel aligned lipid tubules are across a protruded Au square film in an ordered array. The separation of aligned lipid tubules is about ∼5 µm. The ends of the aligned lipid tubules are also visible in the 2D ordered array. Figure 4b is an AFM image of a printed tubule across the edge of the Au film. High profiles along the lines shown in Figure 4b reveal that the heights of the printed lipid tubule on the SiO2 substrate and the protruded Au film are fairly constant (Figure 4c,d), suggesting that the tubule is in contact with both the SiO2 substrate and the protruded Au film. The interaction with the pattern surface forces the lipid tubule to bend at the edge of the Au film to maximize their contact area with the surface and conform to the topography of the patterned substrate. The ability of directly printing ordered tubule arrays on patterned surfaces suggests that our approach is capable of being integrated with current fabrication technology.

To further test the feasibility of this approach, we print the aligned lipid tubules on curved substrates. A glass capillary tube with a 1 mm diameter was used in our experiments. A thin PDMS stamp inked with aligned lipid tubules was brought to contact with the glass tube by bending the stamp around the tube surface under the pressure generated by the Scotch tape (Figure 5a). After 2 h of contact in air at room temperature, the PDMS stamp was peeled from the glass tube (Figure 5b). We find that the aligned lipid tubules are transferred onto the curved glass tube from the recessed channels of the PDMS stamp. By changing focal planes, we observe that the alignment of lipid tubules can be achieved over a large area on the curved glass tube (Figure 5c,d). Again, due to the variation of tubule lengths, there are disconnections in the linearly aligned tubules observed. DC8,9PC lipid tubules have been used as a template for the synthesis of silica cylinders by the sol-gel condensation of deposited TEOS51,52 in solutions. Here we carried out the deposition and condensation of TEOS on 2-D tubule arrays formed on Au-coated mica in the TEOS solution at pH 2.0 at room temperature for a week. The sample was then treated at 110 °C in air for 2 h before imaging. SEM reveals that the 2-D array of aligned lipid tubules is transcribed into a 2-D array of aligned silica cylinders (Figure 6a). The surface of the lipid-templated silica cylinders is continuous and smooth. The control experiments showed that when pristine DC8,9PC tubules adsorbed on substrates were heated in air at 110 °C for 2 h, they broke into small pieces. The chemical composition of thermally treated silica cylinders in the 2-D ordered arrays shown in Figure 6a was analyzed by energy-dispersive X-ray (EDX) microanalysis. The presence of Si is confirmed (Figure 6b). We also find the presence of O, C, and P from lipid molecules in the EDX spectrum, suggesting that the lipid template is not completely removed from the silica cylinder by the thermal treatment. Our previous FT-IR studies showed that the crystalline ordering of lipid bilayer walls confined (51) Seddon, A. M.; Patel, H. M.; Burkett, S. L.; Mann, S. Angew. Chem., Int. Ed. 2002, 41, 2988-2990. (52) Zhao, Y.; Liu, J.; Sohn, Y.; Fang, J. Y. J. Phys. Chem. B 2007, 111, 6418-6421.

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Figure 5. (a, b) Schematic illustration of printing 2-D ordered tubule arrays on a glass tube with a thin PDMS stamp inked with aligned lipid tubules. (c, d) Optical microscopy images of 2-D tubule arrays on the glass tube at different focal planes. These optical images were taken at transmission mode.

Figure 6. (a) SEM image and (b) EDX spectrum of 2-D arrays of lipid-templated silica cylinders on Au-coated mica. The sample was treated at 110 °C for 2 h in air before imaging.

in the hybrid silica-lipid cylinders is stable up to 110 °C.52 Baral and Schoen53 found that the thermal decomposition of lipid molecules confined in silica cylinders took place at a temperature above 250 °C. It is known that the DC8,9PC lipid is zwitterionic rather than charged, so the deposition of TEOS on DC8,9PC tubules cannot be explained by charge interactions. The curvature of a cylindrical lipid tubule can lead to a polarization (53) Baral, S.; Schoen, P. Chem. Mater. 1993, 5, 145-147.

of ∼10-12 C/m.54 The flexoelectric effect of zwitterionic lipid tubules has been used to explain the adsorption of charged nanoparticles on tubule surfaces. It is known that under acidic conditions the alkoxide groups of TEOS are protonated.55 We speculate that the deposition of positively charged TEOS on the zwitterionic DC8,9PC tubules at pH 2.0 might be due to the flexoelectric effect. The condensation of the deposited TEOS leads to the formation of silica films on the DC8,9PC tubules. The diameters of aligned silica cylinders shown in Figure 6a are about 0.5 µm. The separations of the aligned silica cylinders are about 5 µm. The advantage of our approach is that the synthesis and alignment of the silica cylinders can be achieved in a singlestep process. We also noted that some aligned lipid tubules come out from the Au-coated mica during the deposition and condensation of TEOS. In conclusion, a method for directly printing lipid tubules into 2-D arrays on planar, curved, and patterned substrates has been demonstrated. In the process, the aligned lipid tubules which are aligned in the recessed channels of a thin PDMS stamp serve as an ink in the printing process. By printing the aligned lipid tubules onto substrates with µCP, we have constructed 2D ordered arrays and 3D cross-bar junctions. We also show that the 2-D arrays of aligned lipid tubules can be transcribed into the 2-D arrays of aligned silica cylinders through the templated sol-gel condensation. It is believed that our approach can be used to print ordered arrays of other elongated biomolecules on patterned and curved substrates as well, which will open up a simple way to integrate them into microdevices. Acknowledgment. This work was supported by the National Science Foundation (Grants ECS 0521497 and CMMI 0726478). LA703634T (54) Lvov, Y. M.; Price, R. R.; Selinger, J. V.; Singh, A.; Spector, M. S.; Schnur, J. M. Langmuir 2000, 16, 5932-5935. (55) Brinker, C. J.; Schere, C. W. Sol-Gel Science. The Physics and Chemistry of Sol-Gel Processing; Academic Press: New York, 1990.