ARTICLE pubs.acs.org/ac
DNA Separation in Nanowall Array Chips Takao Yasui,*,†,‡ Noritada Kaji,†,‡,^ Ryo Ogawa,§ Shingi Hashioka,§ Manabu Tokeshi,†,‡ Yasuhiro Horiike,§ and Yoshinobu Baba*,†,‡,|| †
Department of Applied Chemistry, Graduate School of Engineering, Nagoya University, Nagoya 464-8603, Japan FIRST Research Center for Innovative Nanobiodevices, Nagoya University, Nagoya 464-8603, Japan § National Institute for Materials Science, Tsukuba 305-0044, Japan Health Research Institute, National Institute of Advanced Industrial Science and Technology, Takamatsu 761-0395, Japan
)
‡
bS Supporting Information ABSTRACT: A nanowall array structure was fabricated on a quartz chip as a separation matrix of DNA fragments, and a 30 s separation was realized for a mixture of DNA fragments (48.5 and 1 kbp fragments) by applying the electric voltage. A longer DNA fragment migrates faster than a shorter one in a nanowall array chip, and it is completely different from the separation of DNA based on gel electrophoresis, nanopillar chips, and nanoparticle array chips. Although the result is similar to DNA separation by entropic trapping, it could not be fully explained by entropic trapping phenomena. Direct observation of single-DNA molecular dynamics inside a nanowall array structure indicates that both confined elongation and relaxation recoiling of a DNA molecule occur, and an elongated DNA molecule migrates faster than a recoiled DNA molecule. Numerical fitting of DNA molecular dynamics reveals that the balance between times for the transverse of a DNA molecule in the nanowall array chip and the relaxation-recoiling of a DNA molecule governs the separation of DNA.
’ INTRODUCTION Recent developments of nanofabrication techniques have produced various nanostructures with highly controlled design as an alternative to conventional separation matrices, such as gels or polymers. These highly controlled nanostructures have also been applied to experimentally elucidate polymer dynamics in confined spaces that de Gennes1 predicted theoretically.29 From a practical standpoint, it is important to integrate microand nanostructures into micro total analysis systems (μTAS) as separation matrices and to apply size fractionation of biomolecules based on electrophoretic techniques. Separation capability has been demonstrated for nanopillar array structures inside microchannels,10,11 and even large DNA molecules that were difficult to separate under direct current (DC) electric fields could be separated in only 20 s.11 Fu et al.12 demonstrated size separation of DNA and protein molecules using patterned anisotropic nanofluidic sieving structures. They also studied separation mechanisms, i.e., Ogston sieving, entropic trapping, and electrostatic sieving. Recently, we fabricated two types of nanopillar array patterns on quartz substrates, including tilted and square array patterns along microchannels. We found that even the square array pattern, which resulted in fewer physical interactions between DNA molecules, could generate a molecular sieving effect, and we achieved DNA separations.13 On the basis of the ratio of the radius of gyration Rg of DNA to the pore size a of a matrix gel, which is equivalent to the nanopillar spacing, the separation r 2011 American Chemical Society
of DNA molecules could be classified into Ogston sieving (Rg/a < 1), 14entropic trapping (Rg/a ≈ 1), or biased reptation (Rg/a > 1). Considering these regimes as they have been established for gel electrophoresis, it can be said that each migrating DNA molecule in the square array pattern underwent a different molecular sieving effect leading to separation. However, the separation mechanism is still not clear, because physical collisions between DNA molecules and nanopillars are far fewer in number than in the tilted array pattern. Thus, it is unlikely that interactions in the nanopillar region are essential for separation. Unlike the demonstration of the entropic trapping technique,1517 the small number of interfaces between nanopillar-free regions and the nanopillar region might not play any role in DNA separation. Therefore, in this study, we fabricated multiple nanowalls into an array structure in order to focus on the contribution of nanospace to DNA separation and to reveal a separation mecphanism.
’ EXPERIMENTAL SECTION The nanowall array structure was fabricated on a quartz substrate basically using the reported procedures.11 Each fabricated nanowall was 500 nm thick, 5000 nm high, and 215 μm long, and a spacing between the nanowalls in the array structure Received: May 9, 2011 Accepted: July 19, 2011 Published: July 19, 2011 6635
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Figure 1. Schematic representation of the main microchannel with the nanowall array structure and SEM images (30 tilted angle) of the nanowall array structure etched into a quartz substrate. A nanowall is 500 nm thick, 5000 nm high, and 215 μm long, with a 200 nm spacing between the nanowalls in the array structure.
was 200 nm, as shown in Figure 1. Briefly, the fabrication process consisted of 11 steps. A 20 nm thick Pt/Cr layer was deposited on a 0.5 mm thick quartz substrate by sputtering, and then electron beam (EB) resist (ZEP-520A, ZEON, Tokyo, Japan) was coated onto the substrate to a thickness of about 1 μm by a spinner. Next, a pattern (500 nm wide and 1 μm deep) of nanowall arrays was delineated by electron beam lithography (EBL; ELS-7500, Elionix, Tokyo, Japan). Ni electroplating into the nanowall array pattern in the EB resist was employed to get a strong mask resistant to reactive ion etching. An array of Ni walls was formed after removal of the EB resist, and then a positive photoresist (OFPR8600, Tokyo Ohka Kogyo, Tokyo, Japan) was coated onto the quartz substrate. Standard photolithography was used to pattern 25 μm wide microchannels. Neutral loop discharge (NLD) plasma etching was applied to the substrate, in which case the Ni walls and photoresist were a mask for the CF4 etching. Inside one of the microchannels, the nanowalls were formed in arrays. Reservoirs were formed by punching with an ultrasonic drill, followed by a cleaning process to remove the mask and the metal layer. The quartz substrate was bonded to a 130 μm thick quartz cover plate by dipping both of them into H2SiF6 and then removing and bonding them at 5 MPa and 65 C for 12 h (Figure 2). DNA fragments of 100 bp (NoLimits, Fermentas Inc., Glen Burnie, MD), 1 kbp (NoLimits, Fermentas), and λDNA (48.5 kbp) (Nippon Gene Co., Ltd., Tokyo, Japan) were stained with bis-intercalating fluorescence dye, YOYO-1 (Invitrogen, Tokyo, Japan), at a dye-to-base pair ratio of 1:15 for DNA separation experiments. T4 DNA (166 kbp) (Nippon Gene Co., Ltd.) was stained with the dye at a dye-to-base pair ratio of 1:5 for single-DNA observations. To reduce photobleaching of DNA stained by YOYO-1, a concentrated buffer solution (3 TBE; 267 mM Tris-borate and 6 mM EDTA, pH 8.3, Sigma-Aldrich, Inc., Tokyo, Japan) containing 10 mM dithiothreitol (DTT, Sigma-Aldrich, Inc.) was used. All experiments were performed on a TE300 inverted microscope (Nikon, Tokyo, Japan) equipped with a high-voltage sequencer (HVS448-1500, Lab Smith, Livermore, CA) to apply the electric fields. DNA electrophoretic behavior was monitored with a C7190-43 EB-CCD camera (Hamamatsu Photonics K.K., Hamamatsu, Japan) through a 10/0.45 NA objective lens and a 100/1.40 NA objective lens (Nikon). The frame rate was 30 Hz. The images
were recorded on a DV tape (Sony, Tokyo, Japan) and then analyzed by image-processing software (Cosmos 32, Library, Tokyo, Japan).
’ RESULTS AND DISCUSSION The mixtures of DNA fragments were injected into the separation channel with the nanowall array structure using a cross-injection method under various electric field conditions. As shown in Figure 3, although the mixture of 48.5 and 1 kbp DNA fragments was successfully separated within 30 s, the mixture of 1 kbp and 100 bp DNA fragments was not. Furthermore, we found that longer DNA fragments migrated faster than shorter DNA ones, and DNA separation was achieved under electric fields of the more than 3.5 V/mm, as shown in Figure 3c. The features of these separation results were contrary to those of previously reported separation results such as when using nanopillars,11 nanoballs,18 nanoparticles,19 and conventional gels or polymers.14 Rather, our results closely resembled the entropic trapping system developed by Han and Craighead17 in which longer DNA fragments migrate faster than shorter ones in crossing nanofilter barriers because of their greater hernia nucleation possibility, which accompanies configurational entropy loss. However, our results could not be explained only by entropic trapping due to the small number of interfaces from the nanowall-free region to the nanowall region. On the basis of these related separation results, the present results can be understood by considering the relationship between the gyration radius (Rg) of a DNA fragment and the nanowall spacing. Considering the persistence length of a DNA molecule (ca. 50 nm), the 100 bp DNA fragment (contour length of 34 nm) acts as a rigid and rod-like polymer. Thus, its gyration radius (three-dimensional size) may be comparable to its contour length. The Rg values of other DNA fragments are estimated from the KratkyPorod model:14 78 nm for 1 kbp and 520 nm for 48.5 kbp. The separation regime will be determined from the relationship between the Rg and the nanowall spacing (G), 200 nm. In the separation of 100 bp and 1 kbp DNA fragments, as both their Rg values are smaller than the nanowall spacing, the separation regime is classified as the Ogston regime. In the Ogston regime, DNA molecules are regarded as spheres and 6636
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Figure 2. Illustration of the fabrication steps for a nanowall array chip. There are 11 steps: pretreating, sputtering, EB resist coating, EBL, Ni electroplating, forming Ni walls after resist removal, photoresist coating, photolithography, plasma etching, cleaning, and bonding.
physical interactions between DNA molecules and randomly placed gel fibers are essential for size-dependent separation.14 Unlike gel systems or entropic trapping arrays, the physical interaction sites in our nanowall array structure are limited to only 20 nanowall interfaces and inside the nanowall spacing. As Han and Craighead15 discussed in another paper, the limited number of interaction sites might make it difficult to separate DNA molecules that are classified in the same separation regime, and therefore, we could not achieve the separation of 100 bp and 1 kbp DNA. In the separation of 48.5 and 1 kbp DNA fragments, each fragment is classified into a different separation regime, i.e., biased reptation for 48.5 kbp DNA (Rg/G > 1) and Ogston sieving for 1 kbp DNA (Rg/G < 1). In this case, it can be thought that DNA molecules have enough interactions to generate different migration velocities, even if the total number of nanowall interfaces is limited. To elucidate the separation regimes of 48.5 and 1 kbp DNA fragments, understanding the confinement effect in nanowall spacing is necessary to estimate the contribution of nanospace during electrophoresis. When a DNA molecule is confined in a cylindrical tube of diameter D (D , Rg), the DNA must be elongated to a certain end-to-end length and this confined elongation keeps the DNA stretched in its equilibrium configuration without any external forces. In addition, physical properties of DNA, considered as a random walk polymer, have been extensively studied using the nanospace formed by nanofabricated structures, which resulted in verification of the scaling law of self-avoiding confined polymers developed by de Gennes.1 Although the nanowall spacing is of sufficient size to confine DNA molecules larger than ca. 15 kbp
(Rg ≈ 200 nm) one dimensionally, DNA molecules can be relaxed to their equilibrium length along perpendicular axes (x and z) to the electric field (y axis) even under electrophoretic migration because the nanospace formed by the nanowall array structure is 200 nm spacing, 5000 nm high, and 215 μm long. The time to achieve the equilibrium length is expressed by the relaxation time τ, and an understanding of this relaxation time τ directly leads to understanding the behavior of DNA molecules in the nanowall spacing for electrophoresis, and as a result the effect of the nanowall array on DNA fragment separation can be estimated. The relaxation time τ in a cylindrical tube of diameter D can be numerically estimated from the following equation4 involving de Gennes’ arguments1 τ¼
8π ηL2 ðpwÞ2=3 5 kB T D1=3
ð1Þ
where η is the viscosity of the solvent, L is DNA contour length, kB is the Boltzmann constant, p is the DNA persistence length, and w is the DNA molecule width. Using the relaxation time, the DNA length L(t) during the relaxation process at arbitrary time t can be roughly estimated by an exponential fitting function described by Mannion et al.20 t LðtÞ ¼ Le þ ðL0 Le Þexp τ ! 5tkB TD1=3 ¼ Le þ ðL0 Le Þexp ð2Þ 8πηL2 ðpwÞ2=3 6637
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Figure 3. Separations of (a) 48.5 (20 ng/μL) and 1 kbp (50.5 ng/μL) DNA fragments and (b) 1 kbp (40 ng/μL) and 100 bp (20 ng/μL) DNA fragments. Electropherograms were taken at 4115 μm from the entrance of the microchannel with the nanowall array structure. In both separations, the applied voltage in the separation channel was 14 V/mm. (c) Resolution vs the separation electric field. Circles and squares show the results from 48.5 vs 1 kbp and 1 kbp vs 100 bp DNA fragments, respectively. Each point is the average for 3 runs.
where Le is the equilibrium length in the nanowall spacing and L0 is the measured initial length. To examine which scale in a nanowall array corresponds to the cylindrical tube of diameter D in eq 1, T4 DNA, whose Rg is much larger than the nanowall spacing, was employed for measuring relaxation time in the nanowall spacing. The DNA molecule was pulled into the nanowall spacing by electrophoresis, and then the observed relaxation process was induced immediately after completion of DNA entry by switching off the voltage. Figure 4a shows the relaxation process of a T4 DNA molecule in the nanowall spacing (see also the Supporting Information movie), and Figure 4b shows the plot of DNA length as a function of time. Since the contour length of T4 DNA is 56.4 μm and our dyeto-base pair ratio (1:5) would increase the contour length by around 23%, resulting in a new contour length of 69.4 μm,20 the T4 DNA was almost fully stretched by the confined elongation as shown in Figure 4. This confined elongation in the nanowall spacing should be governed by the x axis dimension (200 nm) because we could not observe the confined elongation in the channel of 5000 nm width and 5000 nm height (data not shown). To clarify the dominant factor in the nanowall array which relaxes the fully stretched DNA, as a first approximation, we supposed
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Figure 4. (a) Relaxation process of a T4 DNA molecule in the 200 nm spacing between the nanowalls in the array structure was imaged under a weak bright field in order to illuminate the microchannel with the nanowall array structure. (b) Plot of DNA length L(t) as a function of time for the relaxation process of the T4 DNA molecule in the 200 nm spacing. Equation 2 was fitted to the data on the basis of the DNA length in a cylindrical tube with a diameter of 200, 1130, or 5000 nm to quantify this relaxation process.
three diameters of cylindrical tubes (200, 1130, and 5000 nm); 200 nm was the nanowall spacing, 1130 nm was the equivalent area of the nanowall spacing (106 nm2), and 5000 nm was the nanowall height. The relaxation time was calculated using each of the cylindrical diameters, and the obtained values are plotted in Figure 4b. The DNA length in the 5000 nm diameter tube gave the best fit to the experimental value. These results showed that the confined elongation in the nanowall spacing was governed by the x axis dimension (200 nm spacing), and the z axis dimension (5000 nm height) dominated the relaxation process of DNA for this time course (to ∼20 s). In a 5000 nm diameter tube, a DNA molecule can assume a random-coiled configuration if it is in the bulk phase, because the tube radius is much larger than the predicted Rg. Once a DNA molecule whose radius is larger than the nanowall spacing enters the nanowall spacing and confined elongation is achieved, it is expected that relaxation to the thermal equilibrium configuration is rapidly completed and then the DNA recoils to nearly the random-coiled configuration it would have if it were in the bulk phase. In the case of a 48.5 kbp DNA fragment, eq 1 gives only 6638
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Table 1. Transverse Time Per One Nanowall Region (215 μm) Ttrans at 48.5 and 1 kbp DNA Fragment, Separation Resolution Rs, and Migration Distance μ for Various Separation Voltages Ea E/V mm1
Ttrans of 48.5 kbp/s
Ttrans of 1 kbp/s
Rs
μ
28
0.509
0.597
0.712
2.46
14
1.17
1.32
0.648
2.10
7.0
2.49
2.66
0.545
1.19
3.5
7.43
7.52
0.340
0.315
0
0
2.0
10.7
10.7
a
Each value is the average of 3 runs. The transverse time was calculated as in Figure S1 (Supporting information).
to the hydrodynamic friction assumes that the DNA molecule does not accelerate in the nanowall spacing but has a constant velocity ν¼
Figure 5. (a) Schematic illustrations of the DNA migration for 48.5 and 1 kbp DNA fragments. (b) Migration velocity image for 48.5 (black dotted line) and 1 (black solid line) kbp DNA fragments, and separation results of 48.5 and 1 kbp DNA fragment (red curves). The upper graph shows the situation for electric fields over 3.5 V/mm, and the lower one shows it for the electric field of 2 V/mm.
0.5 s as a relaxation time after the confined elongation and some seconds are needed to recoil to nearly the initial random-coiled configuration. When the transverse time (Ttrans), which measures as shown in Figure S-1 in the Supporting Information, across a 215 μm long nanowall region is longer than the sum of the relaxation and recoil times, it is reasonable to assume that both the 48.5 and the 1 kbp DNA fragments would migrate with a random-coiled configuration, rather than an elongated one, during electrophoresis in the nanowall array. Under electrophoresis conditions with a high concentration of DNA molecules, however, this relaxation time is expected to be much longer, because thermal relaxation of a DNA molecule along the z axis should be suppressed by a number of effects which are never trivial and include the electric force, interference of other neighboring DNA molecules, and friction against the nanowall. In total, it would take a few seconds for the 48.5 kbp DNA fragment to recoil to nearly the initial random-coiled configuration. One possible reason why we could separate 48.5 and 1 kbp DNA fragments in the nanowall array structure is that the transverse time of the 48.5 kbp DNA fragment is shorter than the sum of the relaxation and recoil times. In that case, only 48.5 kbp DNA has a different configuration inside the nanowall spacing under electrophoresis, as shown in Figure 5a. Generally, a DNA molecule with charge q, charge per unit contour length ε, and contour length L experiences the electric force fe = qE = εLE under an external electric field E. When a DNA molecule migrates during electrophoresis, the migration of the randomcoiled DNA might be hindered by hydrodynamic friction, Fh ≈ 6πηRgv, where ν is the migration velocity and η is the buffer viscosity. The condition in which the electric force is equivalent
qE 6πηRg
ð3Þ
Because 48.5 kbp DNA should change its configuration inside the nanowall spacing, its configuration is an oval sphere and the 0 apparent Rg (Rg ) to the hydrodynamic friction is smaller than that in the nanowall-free region, unless the DNA has completely relaxed and recoiled (Figure 5a). This process results in a different velocity between that in the nanowall spacing and that in the nanowall-free region, and the DNA velocity eventually increases to some constant value inside the nanowall spacing due to the relatively small Rg generated by the confined elongation. On the other hand, 1 kbp DNA (Rg = 78 nm) does not change its configuration inside the nanowall spacing, and therefore, it has a constant velocity during electrophoresis even if it is in the nanowall spacing (Figure 5a). In Figure 5b we give a schematic diagram of the behavior of 48.5 and 1 kbp DNA fragments in response to electric fields. The migration velocity of 48.5 kbp DNA will change inside and outside the nanowall spacing in contrast to that of 1 kbp DNA which has a constant velocity at all points. For electric fields of 3.5 V/mm or larger in which the transverse time of the 48.5 kbp DNA fragment will be shorter than the sum of the relaxation and recoil times, the 48.5 kbp DNA fragment will have a faster migration velocity than that of the 1 kbp DNA fragment inside the nanowall spacing. We can separate 48.5 and 1 kbp DNA fragments. However, in the electric field of 2 V/mm in which the transverse time of the 48.5 kbp DNA fragment will be longer and the 48.5 kbp DNA fragment will have almost the same migration velocity as the 1 kbp DNA fragment inside the nanowall spacing, we cannot get separation. Due to the experimental difficulty in observing and quantifying relaxation and recoil processes in the nanowall spacing during electrophoresis, we cannot estimate the exact relaxation and recoil times presently. However, the field dependence of separation resolution as shown in Table 1 supports our speculation qualitatively, and the relaxation and recoil times of 48.5 kbp DNA in electric fields will be around 7.510 s. Another possible explanation for the slower migration velocity of the 1 kbp fragment is Brownian diffusion along the z axis in the nanowall structures during electrophoresis. In free solution electrophoresis, Nkodo et al.21 investigated the diffusion coefficient of DNA molecules and found that the scaling law, D ≈ 1/M0.5, could fit their results. If this scaling law is applicable 6639
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’ CONCLUSIONS In summary, we realized the separation of DNA fragments within 30 s using a nanowall array structure and carried out a single-molecule observation to measure the contribution of the nanospace. From the findings we identified the separation mechanism in the nanowall array structure chip and discussed it mainly from the viewpoint of molecular confinement in the nanowall spacing, utilizing observation of a single-DNA molecule. Considering all the various factors, not only the entropic trapping but also the balance between transverse and relaxationrecoil times and diffusion toward the z axis might be dominant factors for DNA separation in the nanowall array structure, which allowed larger DNA fragments to migrate faster than smaller ones. Since this highly controlled nanostructure can determine the separation mechanism by the structure design, we expect it to provide a core technology for further development of μTAS.
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’ REFERENCES (1) de Gennes, P.-G. Scaling Concepts in Polymer Physics; Cornell University Press: Ithaca, NY, 1979. (2) Reccius, C. H.; Mannion, J. T.; Cross, J. D.; Craighead, H. G. Phys. Rev. Lett. 2005, 95, 268101. (3) Reisner, W.; Morton, K. J.; Riehn, R.; Wang, Y. M.; Yu, Z.; Rosen, M.; Sturm, J. C.; Chou, S. Y.; Frey, E.; Austin, R. H. Phys. Rev. Lett. 2005, 94, 196101. (4) Tegenfeldt, J. O.; Prinz, C.; Cao, H.; Chou, S.; Reisner, W. W.; Riehn, R.; Wang, Y. M.; Cox, E. C.; Sturm, J. C.; Silberzan, P.; Austin, R. H. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 10979–83. (5) Turner, S. W.; Cabodi, M.; Craighead, H. G. Phys. Rev. Lett. 2002, 88, 128103. (6) Ueda, M.; Hayama, T.; Takamura, Y.; Horiike, Y.; Dotera, T.; Baba, Y. J. Appl. Phys. 2004, 96, 2937–2944. (7) Odijk, T. Phys. Rev. E 2008, 77, 060901. (8) Daoud, M.; Degennes, P. G. J. Phys. (Paris) 1977, 38, 85–93. (9) Salieb-Beugelaar, G. B.; Dorfman, K. D.; van den Berg, A.; Eijkel, J. C. Lab Chip 2009, 9, 2508–23. (10) Baba, M.; Sano, T.; Iguchi, N.; Iida, K.; Sakamoto, T.; Kawaura, H. Appl. Phys. Lett. 2003, 83, 1468–1470. (11) Kaji, N.; Tezuka, Y.; Takamura, Y.; Ueda, M.; Nishimoto, T.; Nakanishi, H.; Horiike, Y.; Baba, Y. Anal. Chem. 2004, 76, 15–22. (12) Fu, J.; Schoch, R. B.; Stevens, A. L.; Tannenbaum, S. R.; Han, J. Nat. Nanotech. 2007, 2, 121–8. (13) Ogawa, R.; Kaji, N.; Wakao, S.; Hashioka, S.; Baba, Y.; Horiike, Y. Proc. microTAS 2006 2006, 1, 404–406. (14) Viovy, J. L. Rev. Mod. Phys. 2000, 72, 813–872. (15) Han, J.; Craighead, H. G. Anal. Chem. 2002, 74, 394–401. (16) Fu, J.; Yoo, J.; Han, J. Phys. Rev. Lett. 2006, 97, 018103. (17) Han, J.; Craighead, H. G. Science 2000, 288, 1026–9. (18) Tabuchi, M.; Ueda, M.; Kaji, N.; Yamasaki, Y.; Nagasaki, Y.; Yoshikawa, K.; Kataoka, K.; Baba, Y. Nat. Biotechnol. 2004, 22, 337–40. (19) Zeng, Y.; Harrison, D. J. Anal. Chem. 2007, 79, 2289–2295. (20) Mannion, J. T.; Reccius, C. H.; Cross, J. D.; Craighead, H. G. Biophys. J. 2006, 90, 4538–45. (21) Nkodo, A. E.; Garnier, J. M.; Tinland, B.; Ren, H.; Desruisseaux, C.; McCormick, L. C.; Drouin, G.; Slater, G. W. Electrophoresis 2001, 22, 2424–32.
’ ASSOCIATED CONTENT
bS
Supporting Information. Figure S1 and movie. This material is available free of charge via the Internet at http:// pubs.acs.org.
’ AUTHOR INFORMATION Corresponding Author
*Phone: +81 52 789 3560. Fax: +81 52 789 4666. E-mail:
[email protected] (T.Y.), babaymtt@apchem. nagoya-u.ac.jp (Y.B.). Present Address ^
Graduate School of Science, ERATO Higashiyama Live-Holonics Project, Nagoya University, Japan.
’ ACKNOWLEDGMENT We gratefully acknowledge the Japan Society for the Promotion of Science (JSPS) for financial support through its “Funding Program for World-Leading Innovative R&D on Science and Technology (FIRST Program)” and the Grant-in-Aid for JSPS Fellows. 6640
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