LETTER pubs.acs.org/Langmuir
DNA Concentration Modulation on Supported Lipid Bilayers Switched by Surface Acoustic Waves Martin Hennig,†,^ Manuel Wolff,† J€urgen Neumann,‡ Achim Wixforth,§ Matthias F. Schneider,|| and Joachim O. R€adler*,† †
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Center for NanoScience, Ludwig-Maximilians-Universit€at, Fakult€at f€ur Physik, Geschwister Scholl Platz 1, D-80539 M€unchen, Germany ‡ Center for NanoScience, Biozentrum der LMU M€unchen, Großhaderner Straße 2, D-82152 Planegg-Martinsried, Germany § Center for NanoScience, Universit€at Augsburg, Institut f€ur Physik, Universit€atsstraße 1, D-86159 Augsburg, Germany Mechanical Engineering, Boston University, 110 Cummington Street, Boston, Massachusetts 02215, United States
bS Supporting Information ABSTRACT: Spatially addressable arrays of molecules embedded in or anchored to supported lipid bilayers are important for on-chip screening and binding assays; however, methods to sort or accumulate components in a fluid membrane on demand are still limited. Here we apply in-plane surface acoustic shear waves (SAWs) to laterally accumulate double-stranded DNA segments electrostatically bound to a cationic supported lipid bilayer. The fluorescently labeled DNA segments are found to segregate into stripe patterns with a spatial frequency corresponding to the periodicity of the standing SAW wave (∼10 μm). The DNA molecules are accumulated 10-fold in the regions of SAW antinodes. The superposition of two orthogonal sets of SAW sources creates checkerboard like arrays of DNA demonstrating the potential to generate arrayed fields dynamically. The pattern relaxation time of 0.58 s, which is independent of the segment length, indicates a sorting and relaxation mechanism dominated by lipid diffusion rather than DNA self-diffusion.
’ INTRODUCTION Supported lipid bilayers (SLBs) provide a versatile biomimetic platform for studying membrane-binding proteins and cell substrate interactions.13 The properties and functionalities of SLBs can be tailored using defined lipid compositions and engineered surfaces. Many fundamental properties of lipid membranes including diffusion and their interaction with solid surfaces have thus been studied in great detail.4,5 The incorporation or association of membrane proteins to SLBs has been developed and used to study processes of the plasma membrane in a well-controlled and easily accessible model system. Many biotechnological applications have emerged using SLBs, in particular, for biosensing, membrane-based immunoassays, and cell adhesion.6 An important improvement in this context is the use of polymer-brush-supported membranes for the incorporation of bulky transmembrane proteins.1,7 Furthermore, the structuring of SLBs with lateral geometries has been achieved by many approaches (e.g., by substrate micropatterning,8 substrate topography,911 and UV lipid cross-linking12). Recently, DNA molecules have also been grafted or adsorbed to SLBs in order to exploit their inherent structural functionality, such as their defined molecular spacing or sequence specific recognition.1318 DNA can be electrostatically bound to cationic lipid membranes13 or covalently grafted to lipids.14,15 In the r 2011 American Chemical Society
former case, the attached DNA is only loosely bound to cationic lipids via electrostatic forces, remains laterally mobile, and can be moved by external electrical fields.1618 Additionally, the possibility to align DNA by microtextured surfaces, microchannels, or fluid flow has been demonstrated.1921 In principle, electrical fields also provide a switch with which to control local membrane accumulations;7,8 however, there still remains a demand for the easy and dynamic manipulation of macromolecules such as DNA on solid-supported membranes. Surface acoustic waves have been employed to manipulate molecules at surfaces. They are used to drive liquids and accumulate or separate particles in microfluidics.22,23 However, Rayleighwave substrates, often made of quartz or lithium niobate (LiNbO3), strongly couple to the liquid (acoustic streaming)24 and cannot be used to move molecules at surfaces because of their strong attenuation.25 We have recently shown that, in contrast, in-plane shear SAWs are able to cause modulations within SLBs observed through the redistribution of fluorescent lipid probes.26 The SAW-induced patterns can be switched on and off and are in agreement with a model assuming local lipid membrane density Received: August 31, 2011 Revised: November 11, 2011 Published: November 11, 2011 14721
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Figure 2. (a) Chip design with four crossed transducers exciting two orthogonal standing SAWs. Because every transducer excites waves in both directions, wave breakers are placed behind each transducer to avoid unwanted reflections. The direction of the main SAW propagation is indicated by an arrow. (b) AFM images of the SAW substrate in deflection mode with no (upper left), one (upper right, lower left), or both (lower right) pathways activated. (c) Accumulation of fluorescent DNA attached to an SLB in different operating modes.
Figure 1. (a) Diagrammatic representation of the setup used. A LiTaO3 chip has two interdigital transducers (IDTs) on opposite sides to generate a standing SAW. The supported membrane and partially accumulated macromolecules on top, which are between the IDTs, can be observed by fluorescence microscopy through the chip. (b) DNA accumulation (red) is shown at the antinode position where cationic lipid DOTAP (orange) likely accumulates. (c) Distinct, large-scale DNA stripe pattern. The DNA concentration in the sharply separated depleted and accumulated regions differs by approximately a factor 10. (bright value 299, dark value 91, background 68).
modulations. The lipid density modulations relax on timescales in accordance with lipid diffusion. These SAW-induced density modulations within SLBs can be used to accumulate, transport, and separate membrane-bound proteins.27 In this letter, we report on the application of SAWs to generate, control, and move DNA patterns adsorbed to cationic SLBs. Using fluorescence microscopy, we study the reversible accumulation of DNA in regions of standing SAW shear waves. Crossed transducers enable accumulation in switchable checkerboard patterns of ∼10 μm size. To investigate the mechanism of the substrateSAWSLB interaction, we measured the response rates of accumulation as well as redistribution times of DNA after the SAW pulse. The DNA pattern relaxation times are compared to the self-diffusion constants of DNA of different lengths independently measured by FRAP (fluorescence recovery after photobleaching).
’ EXPERIMENTAL SECTION SAWs are generated on piezoelectric crystals using interdigital transducers (IDTs)28 as shown in Figure 1a. The specially prepared lithium tantalate (LiTaO3) substrates used are 200400 μm thick, transparent, and mirror polished on both sides, allowing for high optical
resolution by fluorescence microscopy (Roditi Ltd., London). A crystal cut of 36Y is employed that generates in-plane shear waves.25 Standing acoustic waves were excited by continuous and simultaneous stimulation on both opposed IDTs (Figure 1a). The standing shear SAW reaches over the whole distance between both transducers (3.5 mm) without significant observable damping even in the presence of water. Supported lipid bilayers were prepared by the vesicle fusion method.1 SLBs contained 20% cationic lipid DOTAP in a neutral DOPC bulk environment. This lipid composition was found to be well suited for binding an observable amount of DNA while still providing reliable vesicle fusion. Cy5-labeled double-stranded DNA segments ranging from 20 to 110 base pairs in length were added in excess and bound to the membrane electrostatically. After 15 min, when the surface coverage reached saturation, unbound DNA was removed by rinsing with buffer, whereby the remaining density of adsorbed molecules depends on the percentage of cationic DOTAP within the membrane. Here, we reached an estimated diluted DNA density of less than 1000 molecules/μm2. When SAWs are switched on, the DNA accumulates in the antinodes of the standing SAW (Figure 1b). Using this technique, we can create very homogeneous DNA stripe patterns over macroscopic distances (several millimeters) (Figure 1c). When one of two coupled wave generators creating a standing SAW is slightly detuned (e.g., 0.1 Hz at an operating frequency of 153 MHz), the generated beat starts moving across the chip, resulting in a continuous shift of the DNA pattern (movie in the Supporting Information). When four transducers and two split-wave generators are used (Figure 2a), standing SAWs orthogonal to each other can be excited and directly observed on the SAW substrate by AFM in deflection mode (Figure 2b). It was necessary to rotate the chip design by 45 against the main wave propagation direction (as defined by the crystal cut) so that equally efficient wave stimulation could be achieved in both pathways, although it should be noted that the stimulation of each of the two crossed waves was still less efficient than that along the principal direction as shown in Figure 1c. When both SAWs are switched on, checkerboard patterned “fields” of bound DNA can be observed (Figure 2c, right bottom). The checkerboard pattern arises from the superposition of the patterns that active sound paths form separately along orthogonal axes. In all cases, the induced pattern is reversible independent of the order of SAW channel activation. The SAWs hence reversibly accumulate DNA in fluid 2D microfields.
’ DISCUSSION OF EXPERIMENTAL FINDINGS To investigate the dynamics of the switchable DNA accumulation in more detail, we analyzed the main Fourier component of the periodic pattern for DNA segments of different lengths (204075110 bp) as a function of time. A movie of DNA 14722
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Figure 3. (a) Image series of DNA pattern decay after the SAW has been switched off. (b) SAW-induced membrane density pattern following exp(sin2 x). (For details, see the main text.) (c) Fast Fourier transformation of vertically summed intensity data from image a plotted logarithmically. Its main feature is the dominant peak at a transducer distance of 13.3 μm (half the SAW wavelength). Higher harmonics are also visible. (d) Height of the main Fourier component over time in a typical experiment. The pattern is very distinct the two times that the SAW is switched on and decays exponentially when it is switched off. (e) Plot of the decay of the main peak in c for samples of 75 bp DNA yields the diffusion time D 1/τq02 as a fit parameter, which is visible as the slope in a log I/t diagram. The red line corresponds to an average value of τ = 0.54 s. (f) Diffusion times of DNA segments with lengths of 20, 40, 75, and 110 base pairs (10, 7, 7, and 4 data points, respectively). Error bars depict the standard deviation of the data points. No strong length dependence is observed as theoretically expected. (see the explanation in the main text), and a typical average value of 0.58 s is shown (solid line).
pattern formation and relaxation was recorded, and images were cropped and turned such that the DNA accumulation stripes appear to be vertical (Figure 3a). In a next step, gray values of the images were summed up vertically (Figure 3b), zero padded, and fast Fourier transformed (Figure 3c). It should be noted that the Fourier spectrum reveals that the intensity contour in Figure 3b is not described by a pure sinusoidal intensity distribution. Instead, higher harmonics are visible in the intensity spectrum (Figure 3c). In fact, we have previously shown that the SAW-induced intensity is well described by the phenomenological expression26,27 ! α sin2 kx IF ðxÞ ¼ IB þ I0 exp ð1Þ kB T where IB is the background fluorescence intensity, I0 is the amplitude of the modulation, k is the spatial wave vector, and α is a coupling coefficient. The expression results from a simple model that assumes that the mechanical coupling of the SAW to
the SLB causes density modulations within the membrane, which leads to a redistribution of lipids. We now follow the time course of the main Fourier component at q1 = 0.47 μm1 (corresponding to the IDT distance of 13.3 μm) depicted in Figure 3d. The plot shows a steep rising and falling of the main Fourier component according to the switching on and off of the SAW transducer. The onset occurs on a timescale of approximately 1 s, after which half of the full intensity is reached. The relaxation process appears to be well described by a single exponential in all cases. Figure 3e shows logarithmic plots of experimental intensity courses for several measurements each using 75 bp DNA, displaying an average relaxation time of 0.54 s. There is, however, a considerable sample-to-sample variance, which gives a large relative standard deviation of roughly 50% that is likely due to unavoidable differences in cleaning, focus, temperature, and the fact that the excited membrane is far from its equilibrium state—membranes were observed to degrade at intense SAW exposure. The 14723
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Figure 4. (a) FRAP measurement of double-stranded DNA bound to an SLB. Homogeneously distributed DNA molecules are bleached by a short, intense Gaussian laser pulse, and then the bleached spot evolves by diffusion. The diffusion constant can be calculated from D = σ2(t2) σ2(t1)/2Δt whereas σ represents the width of the Gaussian bleaching profile at time t. (b) Diffusion constants of Cy5-labeled DNA show a segment length dependence. To guide the eye, x1.74 is fitted. The overall values are below the values measured by SAW pattern decay (Figure 3d). For comparison, the independently measured labeled lipid (TR-DHPE) diffusion constant (solid line) is one order of magnitude higher than the DNA diffusion constants.
experiments were repeated for DNA segments with lengths of 20, 40, 75, and 110 bp. We find relaxation times distributed within the experimental error around an average relaxation time of τ1 = 0.58 s, as indicated by the solid line in Figure 3f.
’ PROPOSITION OF A PATTERN-FORMATION MODEL In the following text, we discuss the possible mechanisms of the observed pattern relaxation. Assuming that relaxation occurs on a purely diffusive basis, we expect the main Fourier component intensity to decay according to t ð2Þ Iðq1 , tÞ ¼ A exp τ1 with τ1 = 1/Dq21. Hence the relaxation time τ1 = 0.58s measured for the main Fourier component yields an effective diffusion constant of D = 7.7 μm2/s. To interpret this diffusion constant, we independently measured the diffusion constant of the lipids within the supported bilayer as well as that of the adsorbed double-stranded DNA segments using fluorescence recovery after photobleaching (FRAP). Figure 4a shows an example of an experimental preparation of DNA adsorbed to the SLB before, shortly after, and a few minutes after bleaching. The image processing and data evaluation follows standard FRAP protocols.29 The measured diffusion constants are summarized in Figure 4b. The diffusion of DNA strongly decreases with increasing length. In comparison, the diffusion of a lipid probe (TR-DHPE) in a lipid SLB with the same cationic lipid composition is about 2.8 μm2/s (solid line in Figure 4b). Remarkably, the DNA diffusion constants are significantly lower than the effective diffusion constant retrieved from the SAW relaxation experiment. Furthermore, the SAW experiments did not show any dependence on the DNA segment length. However, the lipid diffusion constant is of the same order as the effective diffusion constant. For this reason, we assume that the relaxation of accumulated DNA after SAW-induced pattern formation is dominated by the lipid mobility rather than the self-diffusion of the adsorbed DNA. However, there remains a small discrepancy because the effective
diffusion appears to be slightly faster than the free lipid diffusion. There indeed remains the possibility that the relaxation of the DNA pattern is to some extent forced by electrostatic repulsion rather than being solely due to free lipid diffusion. Such a driven relaxation seems plausible considering that the pattern is associated with electrostatic repulsion from both the cationic lipids and negatively charged DNA that are proposed to be accumulated within the observed stripes. In this context, it is unfortunate that second-order relaxation times τ2 cannot be measured with the necessary accuracy, allowing us to distinguish a free diffusion process (eq 2) from driven relaxation.
’ CONCLUSIONS We have shown that standing surface acoustic shear waves applied to DNA-decorated supported lipid membranes are able to accumulate DNA into switchable linear stripes or 2D corrals. In this regard, short DNA segments behave like membranebound proteins, which we previously observed to accumulate on SAWSLB devices.27 DNA was found to concentrate in the SAW antinodes, which are assumed to exhibit higher membrane density than the SAW node regions.26 We propose that cationic lipid DOTAP is pushed into the dense phase because of its small headgroup being more favorable to close packing. Because the cationic DOTAP lipids are colocalized with DNA, as shown in ref 16, this leads to the observed DNA accumulation in the antinodes. In contrast, Texas red-labeled lipid (TR-DHPE), which contains a larger headgroup, migrates to node regions of lower membrane density.26,30 In addition, we also determined the kinetic rates of DNA segregation and dissipation after SAW switching on and off and compared this with the diffusion dynamics as determined via FRAP. Dynamic modulations of DNA concentrations on membrane surfaces could be widely used to direct DNA-based reactions or DNA-bound vesicle cargo: in principle, SAW devices could be used in this context for spatial manipulation as well as the detection of artificial reaction systems. In particular, DNA-based vesicle fusion or DNA-mediated structural reactions such as hybridization and strand displacement could be localized. The 14724
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Langmuir reversible patterning and accumulation technique, shown here for membrane-bound DNA, can also be generalized for the concentration modulation of other adsorbed macromolecules or membrane-anchored proteins.27 Striped patterns with repeat distances ranging from 100 nm up to 100 μm are technically feasible simply by changing the wavelength and frequency of the SAWs respectively. In particular, this means that local gradients of surface-bound molecules could be created with varying steepness and extension of the gradient. These could be used to monitor molecular binding or the cellsubstrate interaction as a function of the position-dependent concentration. Moreover, artificial surface gradients play an increasing role as tools in the study of cell migration and cytokine-mediated chemotaxis. In future work, SAW-driven concentration modulations could possibly allow the separation, sorting, and controlled switching of macromolecules and hence could contribute to the development of integrated lab-on-a-chip devices.
’ ASSOCIATED CONTENT
bS
Supporting Information. Detailed materials and methods; experiment on SAW with lipids and DNA labeled; a movie showing DNA stripes moving in a SAW beat pattern. This material is available free of charge via the Internet at http://pubs.acs.org.
’ AUTHOR INFORMATION Corresponding Author
*E-mail:
[email protected]. Present Addresses ^
Current address: Infineon Technologies AG, Am Campeon 1-12, D-85579 Neubiberg, Germany.
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’ ACKNOWLEDGMENT Financial support by the Excellence Cluster “Nanosystems Initiative Munich (NIM)” and the Center for NanoScience (CeNS) is gratefully acknowledged. M.S. acknowledges support from the Center of Nanoscience and Nanotechnology (CNN) of BU. We thank D. Smith for proofreading the letter and S. B€ossinger for AFM expertise. ’ REFERENCES (1) Sackmann, E. Science 1996, 271, 43–48. (2) Chan, Y. M.; Boxer, S. G. Curr. Opin. Chem. Biol. 2007, 11, 581–587. (3) Czolkos, I; Jesorka, A.; Orwar, O. Soft Matter 2011, 7, 4562. (4) Stelzle, M.; Miehlich, R.; Sackmann, E. Biophys. J. 1992, 63, 1346–1354. (5) Nagle, J. F.; Tristram-Nagle, S. Biochim. Biophys. Acta 2000, 1469, 159–195. (6) Smith, A.; Sengupta, K.; Goennenwein, S.; Seifert, U.; Sackmann, E. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 6906–6911. (7) Tanaka, M; Kaufmann, S; Nissen, J; Hochrein, M Phys. Chem. Chem. Phys. 2001, 3, 4091–4095. (8) Groves, J. T.; Boxer, S. G.; McConnell, H. M. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 935–938. (9) Sanii, B.; Smith, A. M.; Butti, R.; Brozell, A. M.; Parikh, A. N. Nano Lett. 2008, 8, 866–871. (10) Rossetti, F. F.; Bally, M.; Michel, R.; Textor, M.; Reviakine, I. Langmuir 2005, 21, 6443–6450. (11) St€ogbauer, T.; Hennig, M.; R€adler, J. O. Biophys. Rev. Lett. 2010, 5, 153–161. 14725
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