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Two-Dimensional Ordered Arrays of Aligned Lipid Tubules on Substrates with Microfluidic Networks Nidhi Mahajan 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 December 14, 2004. In Final Form: January 31, 2005 Microfluidic networks is a powerful tool for aligning one-dimensional materials over a large area on solid substrates. Here we show that lipid nano- and microtubules can be assembled into two-dimensional (2-D) parallel arrays with controlled separations by combining fluidic alignment with dewetting, which occurs within microchannels. We also demonstrate that lipid tubules can be bent into a well-defined shape at the entrance of the channels by the capillary force. Atomic force microscopy is used to study the structure and stability of the aligned lipid tubules on substrates. The deposition experiments with silica colloidal particles show that the 2-D parallel-aligned tubules can be used as a template to synthesize silica films with controlled morphologies and patterns on substrates in a single-step process.
Introduction The self-assembled hollow cylindrical tubules of amphiphilies have attracted considerable attention due to their interesting supramolecular structures and technological applications.1-2 It has been found that a variety of amphiphilies including alkylaldonamides,3 diacetylenic phospholipids,4 and diphenylglycoluril-, amino acid-, and gemini-based surfactants5-7 self-assemble into cylindrical tubules in solutions. The hollow cylindrical shape and the crystalline molecular order of the bilayer walls make the tubules attractive as a template for the synthesis of onedimensional inorganic materials,8-15 a substrate for the crystallization of proteins,16-17 a controlled release system for drug/gene delivery,18-20 and a colorimetric material for chemical sensors.21-23 * To whom correspondence should be addressed. E-mail: jfang@ mail.ucf.edu. (1) Schnur, J. M. Science 1993, 262, 1669-1676. (2) Fuhrhop, J. H.; Helfrich, W. Chem. Rev. 1993, 93, 1565-1582. (3) Fuhrhop, J. H.; Schnieder, P.; Boekema, E.; Helfrich, W. J. Am. Chem. Soc. 1988, 110, 2861-2867. (4) Yager, P.; Schoen, P. E. Mol. Cryst. Liq. Cryst. 1984, 106, 371381. (5) van Nunen, J. L. M.; Stevens, R. S. A.; Picken, S. J.; Nolte, R. J. M. J. Am. Chem. Soc. 1994, 116, 8825-8826. (6) Imae, T.; Takahashi, Y.; Muramatsu, H. J. Am. Chem. Soc. 1992, 114, 3414-3419. (7) Oda, R.; Huc, I.; Schmutz, M.; Candau, S. J.; MacKintosh, F. C. Nature 1999, 399, 566-569. (8) Archibald, D. D.; Mann. S. Nature 1993, 364, 430-433. (9) Baral, S.; Schoen, P. Chem. Mater. 1993, 5, 145-147. (10) Lvov, Y. M.; Price, R. R.; Selinger, J. V.; Singh, A.; Spector, M. S.; Schnur, J. M. Langmuir 2000, 16, 5932-5935. (11) Seddon, A. M.; Patel, H. M.; Burkett, S. L.; Mann, S. Angew. Chem., Int. Ed. 2002, 41, 2988-2991. (12) Price, R. R.; Dressick, W. J.; Singh, A. J. Am. Chem. Soc. 2003, 125, 11259-11263. (13) Patil, A. J.; Muthusamy, E.; Seddon, A. M.; Mann, S. Adv. Mater. 2003, 15, 1816-1819. (14) Jung, J. H.; Lee, S. H.; Yoo, J. S.; Yoshida, K.; Shimizu, T.; Shinkai, S. Chem.sEur. J. 2003, 9, 5307-5313. (15) Yang, B.; Kamiya, S.; Shimizu, Y.; Koshizaki, N.; Shimizu, T. Chem. Mater. 2004, 16, 2826-2831. (16) Wilson-Kubalek, E. M.; Brown, R. E.; Celia, H.; Milligan, R. A. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 8040-8045. (17) Melia, T. J.; Sowa, M. E.; Schutze, L.; Wensel, T. G. J. Struct. Biol. 1999, 128, 119-130. (18) Schnur, J. M.; Price, R. R.; Rudolph, A. S. J. Controlled Release 1994, 28, 3-13. (19) Meilander, N. J.; Yu, X.; Ziats, N. P.; Bellamkonda, R. V. J. Controlled Release 2001, 71, 141-152. (20) Zarif, L. J. Controlled Release 2002, 81, 7-23.
Recently, advances have been made toward the rational control of the tubule structures by adjusting the chemical compositions and the conditions under which self-assembly occurs.24-34 But, as a result of the high aspect ratio, the alignment and ordered arrays of these self-assembled tubules on solid substrates still remain challenging. Rosenblatt et al.35 reported the magnetic alignment of the lipid tubules in dilute solutions. They found that the lipid tubules were oriented with their axes parallel to the field direction when the magnetic field was larger than 4 T. But there is no control in the separations of the aligned lipid tubules. More recently, Shimizu and collaborators36 developed an approach in which a single lipid nanotube could be aligned by microextruding an aqueous dispersion on glass substrates with a needle. Microfluidic networks (µFN)37 have been demonstrated to be a powerful tool to align inorganic nanowires on solid (21) Song, J.; Cheng, Q.; Kopta, S.; Stevens, R. C. J. Am. Chem. Soc. 2001, 123, 3205-3213. (22) Lee, S. B.; Koepsel, R.; Stolz, D. B.; Warriner, H. E.; Russell, A. J. J. Am. Chem. Soc. 2004, 126, 13400-13405. (23) Song, J.; Cisar, J. S.; Bertozzi, C. R.J. Am. Chem. Soc. 2004, 126, 8459-8465. (24) Thomas, B. N.; Safinya, C. R.; Plano, R. J.; Clark, N. A. Science 1995, 267, 1635-1638. (25) Spector, M. S.; Selinger, J. V.; Singh, A.; Rodriguez, J. M.; Price, R. R.; Schnur, J. M. Langmuir 1998, 14, 3493-3500. (26) Thomas, B. N.; Lindemann, C. M.; Clark, N. A. Phys. Rev. E 1999, 59, 3040-3047. (27) Sevenson, S.; Messersmith, P. B. Langmuir 1999, 15, 44644471. (28) John, G.; Mssuda, M.; Okada, Y.; Yase, K.; Shimizu, T. Adv. Mater. 2001, 13, 715-718. (29) Spector, M. S.; Singh, A.; Messersmith, P. B.; Schnur, J. M. Nano Lett. 2001, 1, 375-378. (30) John, G.; Jung, J. H.; Minamikawa, H.; Yoshida, K.; Shimizu, T. Chem.sEur. J. 2002, 8, 5494-5500. (31) Yang, B.; Kamiya, S.; Shimizu, Y.; Koshizaki, N.; Shimizu, T. Chem. Mater. 2004, 16, 2826-2831. (32) Thomas, B. N.; Corcoran, R. C.; Cotant, C. L.; Lindemann, C. M.; Kirsch, J. E.; Persichini, P. J. J. Am. Chem. Soc. 2002, 124, 68666871. (33) Jung, J. H.; John, G.; Yoshida, K.; Shimizu, T. J. Am. Chem. Soc. 2002, 124, 10674-10675. (34) Singh, A.; Wong, E. M.; Schnur, J. M. Langmuir 2003, 19, 18881898. (35) Rosenblatt, C.; Yager, P.; Schoen, P. E. Biophys. J. 1987, 52, 295-301. (36) Frusawa, H.; Fukagawa, A.; Ikeda, Y.; Araki, J.; Ito, K.; John, G.; Shimizu, T. Angew. Chem., Int. Ed. 2003, 42, 72-74. (37) Xia, Y.; Whitesides, G. W. Angew. Chem., Int. Ed. 1998, 37, 550-575.
10.1021/la046928c CCC: $30.25 © 2005 American Chemical Society Published on Web 02/19/2005
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Figure 1. Schematic illustration of the microfluidic technique used to form 2-D ordered arrays of aligned lipid tubules on glass substrates. Parallel microchannels were formed by bringing a PDMS mold to contact with a glass substrate. The alignment of lipid tubules was carried out by pulling a tubule suspension into the channels with capillary action.
substrates. For example, Lieber and co-workers38 have used the fluid flows occurring within the µFN to form two-dimensional (2-D) parallel arrays of nanowires on solid substrates. They have found that once forming the first layer of the aligned nanowires on a substrate, the second layer of the aligned nanowires can be added on it by repeating the alignment process at different directions to form three-dimensional complex crossed arrays. Yang and collaborators39 have used the dewetting occurring within the µFN to align nanowires at the edges of the microchannels and form 2-D parallel arrays on solid substrates. This result suggests the possibility to downsize the patterned features with respect to the size of the channels with the dewetting. Here we show that the µFN can be used to align soft lipid tubules over a large area on glass substrates. Atomic force microscopy (AFM) is used to study the structure and stability of the aligned lipid tubules on substrates. The deposition experiments with silica colloidal particles show that the aligned lipid tubules can be used as a template for the synthesis of silica films with controlled morphologies and patterns on solid substrates in a single-step process. Experimental Section Lipid microtubules were prepared by controlling the cooling process of a 5 mg/mL suspension of 1,2-bis(tricosa-10,12-diynoyl)sn-glycero-3-phosphochloline (DC8,9PC; Avanti Polar Lipids, Alabaster, AL) in ethanol/water (70:30, v/v), following the standard procedure.4 The polymerization of the microtubule suspension was performed with the UV irradiation (254 nm) for 20 min at room temperature. Lipid nanotubules were made by mixing DC8,9PC with 1,2-dinonanoyl-sn-glycero-3-phosphocholine (DNPC) as described previously.27 The UV irradiation of the nanotubule suspension was conducted at 10 °C to avoid the lipid phase transition. The polymerized nanotubule suspension was immediately used after the UV irradiation. Glass substrates were cleaned in 1:1 methanol-hydrochloric acid for 30 min, followed by boiling in distilled water at 100 °C for 15 min and washing with water. The cleaned glass substrates are hydrophilic with a water contact angle of about 8°. (38) Huang, Y.; Duan, X. F.; Wei, Q. Q.; Lieber, C. M. Science 2001, 291, 630-633. (39) Messer, B.; Song, J. H.; Yang, P. D. J. Am. Chem. Soc. 2000, 122, 10232-10233
Figure 2. Optical microscopy image of parallel arrays of the aligned lipid tubules with separations of 2 and 5 µm on a glass substrate (a). AFM images of parallel-aligned lipid tubules with separations of 2 (b) and 5 µm (c). The samples were dried in air at room temperature for 6 h before imaging. In our experiments, µFN were used to align lipid tubules on glass substrates. In the µFN technique, an oxygen plasma treated poly(dimethylsiloxane) (PDMS) stamp having parallel channels was carefully placed on a cleaned glass substrate to form rectangular capillaries (Figure 1a). The capillaries have a height of 0.8 µm and widths of 1 and 2 µm. They are separated by 2 and 4 µm, respectively. The water contact angle on the treated PDMS stamp is about 12°. A droplet of tubule suspensions was pulled into the rectangular capillaries from one of the open ends by capillary action. The tubule solution-filled capillaries were dried in air at room temperature. After removal of the PDMS stamp, the aligned lipid tubules were left on the glass substrate (Figure 1b). The deposition experiments of silica colloidal particles was carried out by exposing the aligned lipid tubules to a Ludox solution (Du Pont Chemical Co., Wilmington, DE). The silica particles are about 10 nm in diameter. An atomic force microscope (Dimension 3100, Digital Instruments) was used to study the structures and stability of the aligned lipid tubules on glass substrates. Silicon nitride cantilevers (Nanosensors) with a normal spring constant of about 30 N/m and a resonant frequency of about 260 kHz were used. The cantilever was excited just below its resonant frequency. All AFM measurements were performed in the tapping mode at a scan rate of 0.5 Hz in air and a humidity chamber. Optical images of the aligned tubules on glass substrates were taken with an Olympus BX41 microscope in air at room temperature.
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Figure 3. (a and b) Low-resolution AFM images of parallel arrays of the aligned lipid tubules on glass substrates. The samples were dried in air at room temperature for a week before imaging. (c) A higher-resolution AFM image taken from the part of the tubule marked in part b. (d) Cross section along the line drawn in part c.
Results and Discussion Figure 2a is an optical microscopy image of 2-D parallel arrays of the aligned microtubules on a glass substrate. It can be seen that the microtubules with different lengths are aligned parallel along one direction, that is, the flow direction over a large area on the glass substrate. The separations of the parallel-aligned tubules are about 2 and 4 µm, respectively. Figure 2b is an AFM image of the parallel-aligned microtubules with a separation of about 2 µm. This AFM image was taken 6 h after the tubules were dried in air at room temperature. These parallelaligned tubules show a cylindrical external surface with a height of about 470 nm, which agrees with the diameter of single lipid microtubules. The apparent width of the microtubules in the AFM image is broad by the geometry of the AFM tip. The ends of the parallel-aligned tubules with a separation of about 4 µm are visible in the AFM image (Figure 2c). Here the tubules, which are aligned within single channels, are not connected. There are gaps between them. The edges of the helical bilayer are visible near the ends of the tubules. These aligned tubules deform on glass substrates through flattening after being dried in air at room temperature for a week. Some of them are only 210 nm high with a flattened top surface (Figure 3a). Despite the significant deformation, there are no cracks and breaks observed on the flattened tubules. Occasionally, the alignment of double tubules in a single channel (1 µm wide) is observed (Figure 3b). By imaging the end of the lipid tubule highlighted in Figure 3b at a high resolution, we observe helical markings (Figure 3c). They are the edges of the lipid bilayer wrapped around the inner tubule core. The cross-section measurement along the line drawn in Figure 3c shows that the edges are about 14.3 and 43.2 nm high (Figure 3d), corresponding to stacks of two and six lipid bilayers, respectively. It has been reported that the thickness of the lipid bilayer in the tubules is about 6.6 nm.24
Figure 4. AFM images of parallel arrays of the aligned lipid tubules on glass substrates. The sample was kept in air (a) and in a humidity chamber (b) for 1 month.
After being dried in air at room temperature for 1 month, we find that these aligned microtubules collapse in the middle. The collapsed tubules show “double peak” morphologies (Figure 4a). A few of the cracks, which are perpendicular to the tubule axis, are visible on the collapsed tubule surfaces. If the sample is sealed in a chamber locked in the humidity from the evaporating
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Figure 5. AFM images of lipid helical (a) and cylindrical nanotubules (b) immobilized on glass substrates. The freshly prepared samples were quickly dried on the glass substrates in air at room temperature.
water for 1 month, we find that the high humidity can prevent the tubules from the collapse (Figure 4b). If we place the flattened tubules in water, they resume their cylindrical shapes, suggesting the elastic deformation of the tubule walls. Figure 5 shows AFM images of the polymerized nanotubules made of DC8,9PC and DNPC on glass substrates. Here a drop of a diluted tubule solution was quickly dried on glass substrates in air at room temperature. Both helical (Figure 5a) and cylindrical nanotubules (Figure 5b) are observed on the glass substrates. The measured height of the lipid nanotubules is about 55 nm, which is close to the diameter of single lipid nanotubules. The average width of the nanotubules measured in the AFM images is about 130 nm. Much of the difference in the widths can be attributed to the geometrical effects of the AFM tip. It has been reported that the nanotubules are not stable and undergo the phase transition in solutions as the temperature rises.27 In our case, the quick immobilization of the lipid nanotubules on the glass substrate might freeze the phase transition. By pulling a dilute nanotubule solution into the microchannels with capillary action, we find that the edges of the channels can act as pinning sites for the immobilization of the nanotubules. Figure 6a is an AFM image of parallel-aligned nanotubules which are formed by dewetting within the hydrophilic microchannels. In our experiments, the solution-filled microchannel was dried in air at room temperature. When solvent evaporates rapidly, the residual solution is expected to recede into two corners of the hydrophilic channel. As a result, the flow-aligned nanotubules are transferred to the edges of the channels via the capillary force (Figure 6b). The parallel-aligned nanotubules shown in Figure 6a are separated by 2 µm, which reflect the feature of the microchannels (width, 2 µm; spacing, 2 µm). This also confirms that the nanotubules are indeed aligned along
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Figure 6. (a) AFM image of a parallel array of the aligned nanotubules on a glass substrate. (b) Schematic illustration of receding of the tubule solution within a channel during dewetting. The tubules are confined at the two edges of the channel.
Figure 7. (a) AFM image of a bent nanotubule on a glass substrate. (b) Schematic illustration of bending of a nanotubule at a channel entrance.
the edges of the channels. Other groups have also used the dewetting occurring within microchannels to align inorganic nanowires39-41 and position block-copolymer micelles.42 (40) Ko, H.; Peleshanko. S.; Tsukruk, V. V. J. Phys. Chem. B 2004, 108, 4385-4393. (41) Chen, J.; Weimer, W. A. J. Am. Chem. Soc. 2002, 124, 758-759. (42) Levi, S. A.; Mourran, A.; Spatz, J. P.; Van Veggel, F. C. J. M.; Reinhoudt, D. N.; Mo¨ller, M. Chem.sEur. J. 2002, 8, 3808-3814.
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Pc ) γ
Figure 8. AFM images of the deposition of silica particles on the aligned lipid tubules at pH 8.4 (a) and pH 2.0 (b). The samples were dried in air for 4 h before imaging.
Occasionally, we find that the nanotubules are bent at the capillary entrance. The bent tubule shows a welldefined shape (Figure 7a), which reflects the structure of the capillary entrance. In this situation, the microchannel flow induced by the capillary action applies a pressure on the portions of the nanotubule, which are exposed to the capillary entrance, in a direction orthogonal to the nanotubule (Figure 7b). The capillary pressure can be calculated by the Young-Laplace equation.43 In our case, the channel is opened at both ends. Given its small dimension, the capillary force Pc can be estimated according to the equation:44 (43) Adamson, A. W.; Gast, A. P. Physical Chemistry of Surface, 6th ed.; Wiley: New York, 1997; pp 4-16. (44) Dimalanta, E. T.; Lim, A.; Runnheim, R.; Lamers, C.; Churas, C.; Forrest, D. K.; de Pablo, J. J.; Graham, M., D.; Coppersmith, S. N.; Goldstein, S.; Schwartz, D. C. Anal. Chem. 2004, 76, 5293-5301.
(
)
cos θglass + cos θpdms 2 cos θpdms + a b
where γ is the surface tension of the liquid, θglass and θPDMS are the contact angles of the channel, and a and b are the height and width of the channel. The channel used in our experiments to bend the nanotubule is 1 µm wide and 0.8 µm high. For water at 20 °C, we have γ ) 72.8 dyn/cm, θglass ) ∼8°, and θPDMS ) ∼12°, which gives a capillary pressure of about 3.2 × 105 N/m2. This pressure bends the nanotubule into a half-circular shape with a radius of about 0.41 µm (Figure 7a). But there are no breaks observed along the bent nanotubule. We explore the application of these aligned lipid tubules as a template to synthesize the silica films. In our experiments, the tubule-coated substrates were immersed in the silica colloidal solution for 6 days without stirring and then gently rinsed with water. The rinsed samples were dried in air before imaging. At pH 8.4, the deposition of the silica particles on the aligned lipid tubules is observed. The gel of the deposited silica particles through the neutralization of their charges leads to the formation of the silica films on the aligned tubules (Figure 8a). The average thickness of the silica films measured from the partially covered tubules is about 420-470 nm. While at pH 2.0, there is no deposition of the silica particles observed on the aligned tubules (Figure 8b). The lipid tubuletemplated silica-full cylinders in solutions have been reported by Baral and Schoen.9 In our case, the deposition of the silica particles only occurs on the tubule surface, which is exposed to the solution. The templated silica films are expected to be hollow, half-cylinders. The advantage of our approach is that the morphology and pattern of the templated silica films are achieved in a single-step process. Conclusions We demonstrate that lipid micro- and nanotubules with high aspect ratios can be aligned and manipulated on glass substrates with µFN. 2-D parallel arrays of the aligned lipid tubules with controlled separations are achieved by combing fluidic alignment and dewetting occurring within µFN. We also show that the lipid nanotubules can be bent into a well-defined shape at the channel entrance by the capillary pressure. By using the aligned lipid tubules as a template, we synthesize silica thin films controlled morphologies and patterns on solid substrates in a single-step process. LA046928C
