DNA Origami Nanoneedles on Freestanding Lipid Membranes as a

Letter pubs.acs.org/NanoLett

DNA Origami Nanoneedles on Freestanding Lipid Membranes as a Tool To Observe Isotropic−Nematic Transition in Two Dimensions Aleksander Czogalla,†,‡,§ Dominik J. Kauert,∥ Ralf Seidel,∥ Petra Schwille,*,⊥ and Eugene P. Petrov*,⊥ †

Laboratory of Membrane Biochemistry, Paul Langerhans Institute, Technische Universität Dresden, Fetscherstraße 74, 01307 Dresden, Germany ‡ German Center for Diabetes Research (DZD), 85764 Neuherberg, Germany § Department of Cytobiochemistry, Faculty of Biotechnology, University of Wrocław, ul. F. Joliot-Curie 14a, 50383 Wrocław, Poland ∥ Institute for Molecular Cell Biology, University of Münster, Schlossplatz 5, 48149 Münster, Germany ⊥ Department of Cellular and Molecular Biophysics, Max Planck Institute of Biochemistry, Am Klopferspitz 18, 82152 Martinsried, Germany ABSTRACT: We introduce a simple experimental system to study dynamics of needle-like nanoobjects in two dimensions (2D) as a function of their surface density close to the isotropic−nematic transition. Using fluorescence correlation spectroscopy, we find that translational and rotational diffusion of rigid DNA origami nanoneedles bound to freestanding lipid membranes is strongly suppressed upon an increase in the surface particle density. Our experimental observations show a good agreement with results of Monte Carlo simulations of Brownian hard needles in 2D. KEYWORDS: DNA origami, lipid membrane, fluorescence correlation spectroscopy, translational diffusion, rotational diffusion, isotropic−nematic transition

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membrane-mediated self-organization of membrane-bound protein molecules into a nematic domain. Brownian motion in dense ensembles of elongated particles can provide important complementary information on the behavior of the system. Translational and rotational diffusion of elongated particles of various aspect ratios in (quasi-) 2D geometry has been extensively studied experimentally during the last years in the dilute regime. These experiments were carried out using ellipsoids,20,21 actin filaments,22 carbon nanotubes23 and nanofibers,24 and germanium nanowires25 in a narrow slit between two walls mimicking 2D geometry. More recently, DNA origami nanoneedles26 and fd-virus particles27 adsorbed on the surface of a freestanding lipid membrane have been studied. At the same time, the very few theoretical and experimental studies that addressed the effect of the surface density of elongated particles in 2D on their diffusion properties, employed particles with the aspect ratios not exceeding 9 (see refs 6, 28, 29, 30, and 31), which is not enough to capture the collective behavior of thin needle-like objects. Thus, both theoretical and experimental results on diffusion properties of 2D ensembles of elongated particles in the hard needle limit are currently missing. In the present Letter, we experimentally study the isotropic− nematic transition in an ensemble of DNA origami nanoneedles bound to a 2D freestanding lipid membrane. Specifically, we

s a result of steric interactions, a two-dimensional (2D) system of hard needle-like objects can undergo the isotropic−nematic (IN) transition.1 Theory predicts1−4 that the IN transition takes place at the reduced surface density of needles ρ = σL2 ≈ 7. Here, L is the needle length, and σ = ⟨n⟩/ A is the surface density, that is, the average number of needles per unit surface area. As it is typical for 2D systems, the resulting nematic phase shows only a quasi-long-range order.1 Similar behavior was found for 2D ensembles of semiflexible filaments5 and strongly elongated hard ellipses with aspect ratios larger than 9 (see ref 6). The importance of the IN transition of strongly elongated particles in 2D has been demonstrated for a wide range of systems, including surface assembly of inorganic nanowires,7−10 spontaneous assembly of rodlike virus particles on polymer surfaces,11 and self-organization of amyloid fibrils at liquid interfaces.12,13 However, the IN transition of strongly elongated particles in 2D may also be of more fundamental biological relevance. In particular, it has been argued that emergence of the nematic phase at high surface densities of strongly elongated particles in 2D can affect the character of binding of long peptide molecules to lipid membranes.14 Furthermore, coupling between the local membrane curvature and nematic ordering of membrane-bound fibers or rodlike biopolymers has been predicted.15 Recent theoretical, simulation-based, and experimental studies16−19 suggest that protein-controlled membrane shape transformation by, for example, BAR domain superfamily proteins, reticulons, and caveolin, which all are characterized by a strongly elongated shape, starts with © XXXX American Chemical Society

Received: October 29, 2014 Revised: November 27, 2014

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monitor the dynamics of their translational and rotational Brownian motion by fluorescence correlation spectroscopy (FCS) and compare the experimental findings with results of Monte Carlo simulations of Brownian hard needles in 2D. The nanoscale folding of DNA into custom-shaped objects (DNA origami) has revolutionized the field of nanotechnology.32 In the DNA origami approach, a large number of short “staple” oligonucleotides bind to defined segments of singlestranded “scaffold” DNA molecule in a sequence-specific manner,33,34 which allows one to construct two- and threedimensional close-packed DNA bundles with predesigned twist and curvature.35 Another crucial feature of DNA origami nanostructures is the possibility to introduce chemically or structurally functional groups with nanometer precision.36,37 Functionalization with lipid moieties can result in DNA origami nanostructures capable of binding to lipid membranes, which has been recently demonstrated experimentally by several groups.26,38−42 In particular, using this approach in our previous work,26 we created DNA origami nanoneedles with the aspect ratio of ∼70 that readily bind to lipid membranes and exercise Brownian motion on the membrane surface. In the present study, in order to understand the primary events in proteinassisted membrane shape transformation we experimentally study diffusion of membrane-bound DNA origami nanoneedles as a function of their surface density, thereby following the IN transition in this system. Amphipathic stiff DNA origami nanoneedles in the form of six-helix bundles (6HB) with cholesteryl-triethylene glycol membrane anchors and fluorescent labels at defined positions have been recently described.26 In the present study, similar structures based on 7560 nucleotide long scaffold strand were designed, produced, and purified according to previously established protocols.26,43 Their correct assembly was confirmed by transmission electron microscopy imaging.26 These structures have a contour length of 0.42 μm, a diameter of 6 nm, and a persistence length of 1.8 μm.43 The amphipathic character of DNA origami nanoneedles was achieved by attaching either two or eight cholesteryl-triethylene glycol membrane anchors on one of the facets along the DNA bundle26 (see Figure 1). These structures readily bind to freestanding lipid membranes and exercise Brownian motion on the membrane surface (for other remarkable properties of these DNA origami nanoneedles, see ref 26). We note that oligonucleotides functionalized with a cholesteryl anchor connected via an ethylene glycol chain efficiently bind to lipid membranes and at the same time virtually do not perturb the lipid bilayer.44,45 In our experiments, even at the highest surface densities of DNA origami nanoneedles on the membrane, the molar fraction of cholesteryl moieties in the membrane leaflet was below 0.01 mol %, which guaranteed that the membrane properties are not affected by the membranebound nanoneedles. To ensure fluorescence detection of membrane-bound nanoneedles, the opposite facet of the 6HB was labeled with a fluorescent marker by choosing appropriate staple oligonucleotides for functionalization. Two labels were used in our experiments: green emitting Alexa 488 and red emitting Alexa 647. The green fluorescent label Alexa 488 was used to label the nanoneedles at either the center or one of the ends, whereas the red fluorescent label Alexa 647 was used to produce only center-labeled DNA origami nanoneedles. Our model lipid membrane system is based on giant unilamellar vesicles (GUVs) composed of 1,2-dioleoyl-sn-

Figure 1. Cholesteryl-triethylene glycol-anchored DNA origami nanoneedles. Schematics of six-helix bundles (a) with eight and two membrane anchors and a fluorescent label at one of the ends or at the center of mass (b). (c) Detailed structure of a cholesteryl-triethylene glycol-modified nucleotide used to produce the membrane anchors. (d) Representative transmission electron microscopy image of a DNA origami nanorod (scalebar: 100 nm). B

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nanoneedles shows a faster decay compared to that of centerlabeled ones (Figure 2a).

glycero-3-phosphocholine (DOPC) (Avanti Polar Lipids, Alabaster, U.S.A.). GUVs were produced by electroformation46 in a polytetrafluoroethylene chamber with Pt electrodes as described previously.26 The electroformation chamber was filled with 320 mOsm/kg aqueous solution of sucrose. GUVs were placed into a bovine serum albumin-coated MatriCal 384 MicroWell Plate observation chambers (MatriCal, Spokane, U.S.A.) filled with iso-osmolar 10 mM HEPES, 150 mM NaCl, 10 mM MgCl2, pH 7.5 buffer. Fluorescently labeled DNA origami nanoneedles in buffer (see above) were added to the observation chambers at a final concentration of 10−500 pM. After that, the samples were incubated for at least 6 h at 4 °C in sealed chambers. Before the measurement, the samples were equilibrated at room temperature for 30 min. No DNA origami particles diffusing freely in bulk solution could be detected when the above protocol was followed. FCS measurements were carried out using an LSM 780 ConfoCor 3 system (Carl Zeiss, Jena, Germany) equipped with a C-Apochromat 40×/1.2W water immersion objective. Laser lines with wavelengths of 488 nm (excitation of Alexa 488) and 633 nm (excitation of Alexa 647) were used at low power (below 2 μW) to avoid photobleaching. The pinhole position and the correction collar of the objective were adjusted to maximize the detected fluorescence signal and photon count rate per molecule, respectively. The lateral size of the FCS detection volume r0 (190 ± 4 and 241 ± 4 nm in the green and red detection channels, respectively) was determined using fluorescent dyes with known diffusion coefficients in water, as described previously.26 To ensure proper determination of the concentration of membrane-bound DNA origami particles, amplitudes of FCS autocorrelation functions were corrected for the presence of a noncorrelated background signal. All experiments were carried out at room temperature (21.5 ± 1 °C). To measure fluorescence intensity fluctuations due to Brownian motion of membrane-bound nanoneedles, the center of the confocal detection volume was placed at the upper pole of a GUV with the diameter of at least 20 μm. The very large size of a GUV compared to the size of the detection spot ensures that the system can be considered as essentially twodimensional. The translational and rotational diffusion coefficients of fluorescently labeled membrane-bound nanoneedles, DT and DR, were determined from the FCS autocorrelation curves using the approach described previously.26 Briefly, in the absence of photophysical and photochemical processes the autocorrelation function of fluorescence fluctuations reflects the Brownian motion of a fluorescent label. Under certain assumptions, the FCS autocorrelation function for a particle exercising Brownian motion in 2D within a focal plane of the confocal detection volume can be written as follows:47 G(τ) = [⟨N⟩(1 +⟨r2(τ)⟩/r20)]−1, where ⟨r2(τ)⟩ is the mean square displacement (MSD) of the fluorescent label attached to the particle, and ⟨N⟩ is the mean number of particles in the confocal detection spot. If a fluorescent label is attached to the center of mass of a rodlike particle, FCS gives information only on the translational diffusion: ⟨r2(τ)⟩ = 2 (τ)⟩ = 4D T τ. In case of end-labeling, the FCS ⟨r CM autocorrelation function reflects both the translational and rotational diffusion of the rodlike particle:26 ⟨r2(τ)⟩ = ⟨r2end(τ)⟩ = 4DTτ + 1/2L2(1 − exp(−DRτ)). As a result, the FCS autocorrelation function of the end-labeled DNA origami

Figure 2. Diffusion dynamics of DNA origami nanoneedles bound to a freestanding lipid membrane as detected by FCS. Representative FCS autocorrelation curves for membrane-bound nanoneedles with eight membrane anchors fluorescently labeled at the center of mass or at one of the ends (symbols) along with their fits using the model described in the text (solid curves). (a) Effect of the label position at a low surface density of nanoneedles: (□) center-labeled, σ = 0.82 μm−2 (ρ = 0.15) and (○) end-labeled, σ = 0.96 μm−2 (ρ = 0.17). The graphical insets illustrate the effect of the fluorescent label position on the character of fluorescence fluctuations for a particle moving in a Gaussian detection spot. (b) Effect of the surface density for endlabeled nanoneedles: (○) σ = 0.96 μm−2 (ρ = 0.17), (Δ) σ = 2.7 μm−2 (ρ = 0.48), (▽) σ = 5.3 μm−2 (ρ = 0.94), and (◊) σ = 8.3 μm−2 (ρ = 1.46). The number of fluorescent particles in the detection volume of the green (red) channel in these measurements was 0.11 (0), 0.31 (0), 0.09 (0.84), and 0.07 (1.41), respectively.

In the present study, we are interested in the self-diffusion properties of membrane-bound origami nanostructures. In the dilute regime (surface density below 2 μm−2), FCS measurements were carried out using DNA origami nanoneedles that were labeled at either the center or one of the ends with a green dye and bound to freestanding lipid membranes. The surface density of particles was determined from the amplitude of the FCS autocorrelation function. In order to detect only selfdiffusion of particles and avoid the unwanted contribution of collective (mutual) diffusion to the FCS signal, at higher surface densities of membrane-bound fluorescent particles, the greenlabeled DNA origami nanoneedles constituted only a small fraction of membrane-bound particles, whereas the majority of the membrane-bound particles were labeled with the red dye. For particle densities of 4−7, 8−10, and >11 μm−2 (ρ = 0.7− 1.2, 1.4−1.8, and >1.9) the fraction of the origami particles labeled with the green label was in the range of 0.1−0.4, 0.04− C

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0.2, and 0.01−0.08, respectively. We found that for particle densities σ < 5 μm−2 (ρ < 0.88) samples containing only greenlabeled particles and mixtures of green- and red-labeled particles gave identical results within the experimental uncertainty. On the one hand, this approach allowed us to estimate the total surface density of membrane-bound particles from the amplitudes of the fluorescence autocorrelation functions in the red and green detection channels and, on the other hand, to correctly measure fluorescence fluctuations due to translational (and rotational, in case of end-labeling) self-diffusion of green-labeled DNA origami nanoneedles in a dense 2D ensemble. First, we briefly discuss the Brownian motion of DNA origami nanoneedles bound to lipid membranes in the dilute regime. We estimated the diffusion coefficients at infinite dilution, DT0 and DR0, by averaging the values for surface densities σ < 2 μm−2 (ρ < 0.35). For DNA origami nanoneedles functionalized with eight cholesteryl anchors we found DT0 = 1.48 ± 0.06 μm2/s and DR0 = 39.0 ± 2.6 rad2/s, which is about factor of 1.5 higher than predictions of the hydrodynamicsbased theory of diffusion of a needle-like membrane inclusion.48,49 At the same time, nanoneedles bound to the lipid membrane with two cholesteryl anchors show faster translational and rotational diffusion: DT0 = 1.68 ± 0.07 μm2/s and DR0 = 68.8 ± 7.4 rad2/s. Taken together, these results allow us to conclude that in both cases the nanoneedles are not embedded into the lipid membrane bilayer but rather glide over its surface, being held by the membrane anchors buried in the lipid membrane. When the surface density of membrane-bound nanoneedles exceeds 2 μm−2 (ρ > 0.35), we observe that both the translational and rotational diffusion coefficients show a pronounced drop with increasing the particle surface density (see Figures 2b and 3). Note that the rotational diffusion coefficient shows a stronger dependence on the surface density of membrane-bound particles (Figure 3). These observations are in agreement with the progressively stronger steric interactions of the nanoneedles, which is illustrated by Figure 4. Remarkably, the translational and rotational diffusion coefficients normalized by their extrapolated values at infinite dilution, DT0 and DR0, show essentially identical dependences on the surface density of particles for the nanoneedles with eight and two cholesteryl anchors (Figure 5). This provides evidence that the dependence is due to steric interactions of membrane-bound particles, and, irrespective of the number of membrane anchors, the same phenomenon of the onset of the isotropic−nematic transition in 2D is observed (Figure 5). To verify this conclusion, we carried out Monte Carlo (MC) simulations of a system resembling the one studied experimentally, albeit under a number of simplifying assumptions. First, we represent membrane-bound origami DNA particles as a collection of hard needles in 2D. This assumption is justified because of the very high aspect ratio of the DNA origami particles (their contour length is 70 times larger than their diameter) and large persistence length (which is ca. 4.5 times larger than their contour length).41 Second, we assume that the Brownian motion of the membrane-bound origami particles is completely overdamped, which is fully justified for the range of time scales under consideration (from microseconds to seconds). Third, to further simplify the description of the translational and rotational Brownian motion of membrane-bound origami particles, we assume that their translational diffusion coefficients along and normal to their axis

Figure 3. Translational (a) and rotational (b) diffusion coefficients obtained from experimental FCS curves, as a function of membrane surface concentration of DNA origami nanoneedles attached to the membrane with two (○, ●) and eight (□, ■) membrane anchors. Open and filled symbols denote data obtained from center- and endlabeled nanoneedles, respectively.

Figure 4. Representative configurations of hard needles obtained in Monte Carlo simulations for a range of reduced density values: ρ = 0.0625 (a), 0.25 (b), 1 (c), 2 (d), 4 (e), and 6.25 (f). The particle density is controlled by appropriately choosing the simulation box size.

are equal (as discussed previously,26 for a membrane-bound particle of a size of the 6HB and the typical viscosity of the DOPC membrane, the ratio is 1.6 or lower). Under this assumption, the translational and rotational Brownian motions of a needle are decoupled, which considerably simplifies the computations. We carry out simulations with n needles in a square box with periodic boundary conditions using a D

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At short times, when needles move freely without interaction with their neighbors, the MSD curves show faster diffusion corresponding to the infinite dilution regime. At later times the MSD curves cross over to long-time slower diffusion controlled by steric interactions in the ensemble of needles (Figure 6).

Figure 5. Reduced translational (a) and rotational (b) diffusion coefficients as a function of the surface density of DNA origami nanoneedles attached to the membrane with two (○, ●) and eight (□, ■) membrane anchors (as determined from FCS data) and for Brownian hard needles in 2D (as determined from Monte Carlo simulations) (⧫). Open and filled symbols representing experimental data denote results obtained for center- and end-labeled nanoneedles, respectively.

Figure 6. Representative MSD plots for Monte Carlo simulations of translational (a) and rotational (b) self-diffusion of Brownian hard needles in 2D for several reduced densities of needles: ρ = 0.82 (red), 1.98 (blue), 3.27 (orange), and 4.44 (green). The solid black lines represent the behavior at infinite dilution, corresponding to the dependences MSDT(τ) = 4DT0τ and MSDR(τ) = 2DR0τ. DT0 and DR0 denote translational and rotational diffusion coefficients at ρ = 0, respectively. Dashed lines illustrate the linear time dependence of the MSD at long times.

modification of the event-driven Brownian dynamics algorithm.50 The particle density is controlled by appropriately choosing the simulation box size. Unlike in the original algorithm,50 where all n particles are simultaneously assigned translational and rotational displacements, we split an MC step into n substeps. During each substep, a single randomly chosen needle attempts to make a random displacement. In case this substep results in an intersection with a neighbor, the motion of a needle during a substep is treated as in the original algorithm.50 To adequately describe the time scales of the translational and rotational motion in the simulation, we choose the rotational and translational diffusion coefficients such that the quantity L2DR0/DT0 is the same as in the experiment. The MC simulation time step was chosen depending on the surface density of the needles to provide, on the one hand, a reasonable computation speed and, on the other hand, to make sure that the results are not influenced by the particular choice of the time step. After equilibration of the system, position and orientation trajectories of a subset of the needles were recorded, and translational and rotational MSDs were calculated. Consistent results were obtained in simulations with n = 40 and n = 100 at the same particle densities.

These long-time tails of the translational and rotational MSDs were used to estimate the translational and rotational diffusion coefficients of the hard needles to compare with the experimental data. This is justified, because the crossover times estimated from the simulations range from 3−5 ms at low surface densities to 0.5−1 ms at high surface densities. At the same time, the important part of the FCS autocorrelation curves corresponds to times longer than 1 ms. Therefore, under our experimental conditions they are mainly sensitive to the long-time diffusion dynamics. In agreement with the theoretical work1−4 predicting that the isotropic−nematic transition in a system of hard needles should take place at a reduced surface density values of ρ = σL2 ≈ 7, we observe that at the reduced surface densities approaching this value the rotational diffusion coefficient drops dramatically by a factor of ∼100, whereas the translational diffusion coefficient is reduced by a factor of ∼2.5 compared to its infinite dilution value (Figure 5). These results indicate the formation of a strongly locally ordered phase with a pronounced coalignment of the needles in which their rotational diffusion as well as their E

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(10) Hore, M. J. A.; Composto, R. J. ACS Nano 2010, 4, 6941−6949. (11) Yoo, P. J.; Nam, K. T.; Qi, J.; Lee, S.-K.; Park, J.; Belcher, A. M.; Hammond, P. T. Nat. Mater. 2006, 5, 234−240. (12) Isa, L.; Jung, J. M.; Mezzenga, R. Soft Matter 2011, 7, 8127− 8134. (13) Jordens, S.; Isa, L.; Usov, I.; Mezzenga, R. Nat. Commun. 2013, 4, 1917. (14) Almeida, P. F. F.; Wiegel, F. W. J. Theor. Biol. 2006, 238, 269− 278. (15) Murugesan, Y. K.; Pasini, D.; Rey, A. D. Soft Matter 2011, 7, 7078−7093. (16) Lipowski, R. Faraday Discuss. 2013, 161, 305−331. (17) Ramakrishnan, N.; Sunil Kumar, P. B.; Ipsen, J. H. Biophys. J. 2013, 104, 1018−1028. (18) Cui, H.; Mim, C.; Vázquez, F. X.; Lyman, E.; Unger, V. M.; Voth, G. A. Biophys. J. 2013, 104, 404−411. (19) Simunovic, M.; Mim, C.; Marlovits, T. C.; Resch, G.; Unger, V. M.; Voth, G. A. Biophys. J. 2013, 105, 711−719. (20) Han, Y.; Alsayed, A. M.; Nobili, M.; Zhang, J.; Lubensky, T. C.; Yodh, A. G. Science 2006, 314, 626−630. (21) Han, Y.; Alsayed, A. M.; Nobili, M.; Yodh, A. G. Phys. Rev. E 2009, 80, 011403. (22) Li, G.; Tang, J. X. Phys. Rev. E 2004, 69, 061921. (23) Duggal, R.; Pasquali, M. Phys. Rev. Lett. 2006, 96, 246104. (24) Bhaduri, B.; Neild, A.; Ng, T. W. Appl. Phys. Lett. 2008, 92, 084105. (25) Marshall, B. D.; Davis, V. A.; Lee, D. C.; Korgel, B. A. Rheol. Acta 2009, 48, 589−596. (26) Czogalla, A.; Petrov, E. P.; Kauert, D. J.; Uzunova, V.; Zhang, Y.; Seidel, R.; Schwille, P. Faraday Discuss. 2013, 161, 31−43. (27) Herold, C. PhD Thesis, Technische Universität Dresden, Dresden, 2012. (28) Lahtinen, J. M.; Hjelt, T.; Ala-Nissila, T.; Chvoj, Z. Phys. Rev. E 2001, 64, 021204. (29) Maeda, H.; Maeda, Y. Nano Lett. 2007, 7, 3329−3335. (30) Zheng, Z.; Han, Y. J. Chem. Phys. 2010, 133, 124509. (31) Zheng, Z.; Wang, F.; Han, Y. Phys. Rev. Lett. 2011, 107, 065702. (32) Saccà, B.; Niemeyer, C. M. Angew. Chem., Int. Ed. 2012, 51, 58− 66. (33) Rothemund, P. W. Nature 2006, 440, 297−302. (34) Castro, C. E.; Kilchherr, F.; Kim, D. N.; Shiao, E. L.; Wauer, T.; Wortmann, P.; Bathe, M.; Dietz, H. Nat. Methods 2011, 8, 221−229. (35) Dietz, H.; Douglas, S. M.; Shih, W. M. Science 2009, 325, 725− 730. (36) Jahn, K.; Torring, T.; Voigt, N. V.; Sorensen, R. S.; Bank Kodal, A. L.; Andersen, E. S.; Gothelf, K. V.; Kjems, J. Bioconjugate Chem. 2011, 22, 819−823. (37) Simmel, F. C. Curr. Opin. Biotechnol. 2012, 23, 516−521. (38) Börjesson, K.; Lundberg, E.; Woller, J.; Nordén, B.; Albinsson, B. Angew. Chem., Int. Ed. 2011, 50, 8312−8315. (39) Langecker, J.; Arnaut, M. V.; Martin, T. G.; List, J.; Renner, S.; Mayer, M.; Dietz, H.; Simmel, F. C. Science 2012, 338, 932−936. (40) List, J.; Wever, M.; Simmel, F. Angew. Chem., Int. Ed. 2014, 53, 4236−4239. (41) Johnson-Buck, A.; Jiang, S.; Yan, H.; Walter, N. G. ACS Nano 2014, 8, 5641−5649. (42) Conway, J. W.; Madwar, C.; Edwardson, T. G.; McLaughlin, C. K.; Fahkoury, J.; Lennox, R. B.; Sleiman, H. F. J. Am. Chem. Soc. 2014, 136, 12987−12997. (43) Kauert, D. J.; Kurth, T.; Liedl, T.; Seidel, R. Nano Lett. 2011, 11, 5558−5563. (44) Pfeiffer, I.; Hook, F. J. Am. Chem. Soc. 2004, 126, 10224−10225. (45) Bunge, A.; Loew, M.; Pescador, P.; Arbuzova, A.; Brodersen, N.; Kang, J.; Dahne, L.; Liebscher, J.; Herrmann, A.; Stengel, G.; Huster, D. J. Phys. Chem. B 2009, 113, 16425−16434. (46) Angelova, M.; Dimitrov, D. S. Prog. Colloid Polym. Sci. 1988, 76, 59−67. (47) Petrov, E. P.; Schwille, P. In Standardization and Quality Assurance in Fluorescence Measurements II: Bioanalytical and Biomedical

translational diffusion in the direction normal to the needle direction is strongly inhibited. In other words, we observe here the onset of the isotropic−nematic transition of Brownian hard needles in 2D. At the same time, based on the results of ref 1 and our simulations (see Figure 4), the characteristic sizes of the locally ordered nanoneedle domains are not expected to exceed 1 μm under conditions of our experiments. Remarkably, the dependences of the reduced translational and rotational diffusion coefficients as a function of the surface density of particles obtained from the simulations agree well with the results of the FCS measurements (Figure 5). Taking into account all the simplifying assumptions used in the simulations, we conclude that the 2D IN transition under the conditions of our experiment is mainly caused by the direct steric interactions of DNA origami nanoneedles on the membrane.

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AUTHOR INFORMATION

Corresponding Authors

*(P.S.) E-mail: [email protected]. *(E.P.P.) E-mail: [email protected]. Author Contributions

The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by the Deutsche Forschungsgemeinschaft “Transregio 83” Grant TRR83 to A.C. and P.S. and a starting grant of the European Research Council (No. 261224) to R.S. E.P.P. and R.S. gratefully acknowledge the support by the Deutsche Forschungsgemeinschaft via the Research Group FOR 877 “From local constraints to organized transport”. We thank Veselina Uzunova and Yixin Zhang from B CUBE, Dresden, Germany for providing cholesteryl-triethylene glycol modified staple oligonucleotides and Gokcan Aydogan for his assistance in FCS measurements. E.P.P. appreciates helpful and illuminating discussions with Ralf Stannarius from the Otto von Guericke University, Magdeburg, Germany.

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ABBREVIATIONS 2D, two-dimensional, two dimensions; 6HB, six-helix bundle; DOPC, 1,2-dioleoyl-sn-glycero-3-phosphocholine; FCS, fluorescence correlation spectroscopy; GUV, giant unilamellar vesicle; IN, isotropic−nematic; MSD, mean square displacement; MC, Monte Carlo

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REFERENCES

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