Letter pubs.acs.org/NanoLett
Simultaneous Single-Molecule Force and Fluorescence Sampling of DNA Nanostructure Conformations Using Magnetic Tweezers Felix E. Kemmerich,‡,§ Marko Swoboda,† Dominik J. Kauert,‡,§ M. Svea Grieb,† Steffen Hahn,† Friedrich W. Schwarz,†,∥ Ralf Seidel,*,‡,§ and Michael Schlierf*,† †
B CUBE - Center for Molecular Bioengineering, TU Dresden, 01307 Dresden, Germany Institute for Molecular Cell Biology, University of Münster, 48149 Münster, Germany § Institute of Experimental Physics I, Universität Leipzig, 04103 Leipzig, Germany ∥ cfaed - Center for Advancing Electronics Dresden, TU Dresden, 01307 Dresden, Germany ‡
S Supporting Information *
ABSTRACT: We present a hybrid single-molecule technique combining magnetic tweezers and Förster resonance energy transfer (FRET) measurements. Through applying external forces to a paramagnetic sphere, we induce conformational changes in DNA nanostructures, which are detected in two output channels simultaneously. First, by tracking a magnetic bead with high spatial and temporal resolution, we observe overall DNA length changes along the force axis. Second, the measured FRET efficiency between two fluorescent probes monitors local conformational changes. The synchronized orthogonal readout in different observation channels will facilitate deciphering the complex mechanisms of biomolecular machines. KEYWORDS: Magnetic tweezers, FRET, single-molecule, correlated measurements, force-fluorescence microscopy
S
provide deeper insight into the mechanisms of such molecular machines. The potential of combined measurements has recently been realized and has stimulated the development of a number of hybrid approaches. This includes the combination of mechanical manipulation with single-molecule fluorescence detection8−15 as well as the coupling of different fluorescence detection channels.16 The first simultaneous and fully synchronized execution of force-based methods and singlemolecule fluorescence experiments was accomplished by optical tweezers.17 However, these experiments were often limited by a reduced fluorophore lifetime due to the generation of oxygen radicals18−20 and by the lacking capability to parallelize the experiment. A force-based single-molecule technique that can overcome these limitations is magnetic tweezers, which do not use lasers that would lead to oxygen radicals. Furthermore, they expand the manipulation toolkit by the ability to apply and quantify torsion on single biomolecular tethers21,22 and are massively parallelizable providing an increased throughput of singlemolecule measurements.23 However, for magnetic tweezers a simultaneous execution of a force-based experiment combined with single-molecule fluorescence is more difficult to realize.
ingle-molecule methods have emerged as valuable tools to study nanoscale biomolecular systems. They allow the detection and quantification of conformational states of individual biomolecular complexes in real-time.1 At the same time, they permit the manipulation of single molecules by external mechanical stimuli such as tension and torque.2,3 In this way, unique insights into the dynamics of biomolecular systems can be obtained, which often presents a challenge for established ensemble methodologies. Most single-molecule techniques fall into one of two categories:4 force-based manipulation experiments2 and lightbased observation via fluorescence or plasmonics.5 While in the first category mechanical stimuli are applied to probe mechanical properties or to regulate kinetic processes, experiments of the second category track unconstrained molecular behavior and allow localizing and distinguishing molecular species. Typically single-molecule techniques are applied in isolation, that is, either force or fluorescence. However, in these kinds of experiments the accessible information space is often limited to a simplifying one-dimensional coordinate. Thus, the content available from single-molecule measurements can be significantly enhanced by a combination of orthogonal methods. For example, tracking the movement of single motor proteins under force (e.g., cytoskeletal or DNA interacting motors6,7) and correlating the measured trajectories with a particular conformational state of the enzyme, could © XXXX American Chemical Society
Received: September 29, 2015 Revised: December 2, 2015
A
DOI: 10.1021/acs.nanolett.5b03956 Nano Lett. XXXX, XXX, XXX−XXX
Letter
Nano Letters The position detection of the magnetic particles in these instruments typically relies on real-time analysis of particle images acquired by a camera. Integrating such a particle tracking and synchronous fluorescence detection is technically challenging due to the limited wavelength range for imaging and fluorescence excitation and emission, as well as due to limited computational power. Previous realizations of magnetic tweezers with single molecule fluorescence therefore focused on simplified schemes such as single color detection13 or serial execution, where first the magnetic forces were calibrated and then the fluorescence measurements were performed in isolation.24−26 Alternatively, a pseudosynchronization was applied that uses an independent acquisition and processing unit for each detection channel,27 which is only reliable for slow processes. Here, we combined high-resolution magnetic tweezers and dual-color single-molecule fluorescence detection into a single instrument. We demonstrate the simultaneous and fully synchronized operation of both orthogonal channels and provide a proof-of-principle for the detection of more than one molecule at a time. The constructed instrument conveniently supports magnetic tweezers experiments with subnanometer accuracy at acquisition rates of up to 2 kHz, while the dual-color fluorescence imaging of single organic dye molecules enables simultaneous FRET measurements over extended periods of time. An overview of the setup is depicted in Figure 1A. The key is the spectral separation of both the tracking microscope for magnetic tweezers and an objective-based total internal reflection fluorescence (TIRF) microscope for FRET. The high-speed tracking for the magnetic tweezers requires relatively high illumination intensities; therefore an infrared light source with a low photon energy was used to avoid excitation and damage of the fluorophores or the sample by the tracking light. In particular, a spectrally filtered (770−950 nm wavelength) output of a laser-ignited Xe-plasma lamp was focused on the sample chamber to illuminate 1 μm superparamagnetic beads attached to DNA. A pair of magnets above the flow chamber enabled the stretching and twisting of tethered molecules.28 During the measurements, images of the beads were recorded with a CCD camera or with a CMOS camera. The CCD camera was employed for bead imaging at a standard frame rate of 200 Hz, while the CMOS camera was used for rates >200 Hz in particular also for the high resolution measurements recorded at 2 kHz.29 The bead diffraction images (Figure 1B) were used to calculate the position of the bead with respect to the objective focal point in real time.29,30 Forces were calibrated using the lateral fluctuations of the DNA tethered bead.31 Fluorescence excitation was implemented in objective-type TIRF configuration by focusing 532 and 642 nm lasers onto the backfocal plane of the high-numerical-aperture objective. Total internal reflection conditions were achieved by lateral displacement of the excitation beams from the optical axis. This generated an evanescent field and limited the fluorescence excitation to a small region above the sample surface. The fluorescence channel was optically separated from the bead-tracking channel using a dichroic mirror. Further laser rejection filters blocked both, scattered excitation light and residual tracking light. The emission light was then divided into two channels (for donor and acceptor emission) using a dichroic mirror. Each channel was imaged on one-half of an electron-multiplying CCD camera and mapped onto each
Figure 1. Setup overview and sample design: (A) Sketch depicting the general layout of the instrument. An infrared light source illuminates the flow cell and contained sample, which is imaged by a tracking camera. Lasers for fluorescence excitation are coupled in through the objective in TIRF configuration and fluorescence is separated from the infrared illumination light for the bead tracking by dichroic mirrors (DM, see Supporting Information for specifications of dichroic mirrors and filters). Images from the different detection paths are recorded by an electron-multiplying CCD camera (up to 47.5 Hz) or a CMOS camera (up to 2 kHz), respectively. Magnets mounted above the flow cell allow exertion of force and torque to tethered molecules within the flow cell. (B) Two exemplary bead diffraction images as taken by the tracking camera (CCD) with 3 μm difference in height are depicted, whose radial profiles are evaluated during real-time position tracking.
other.32 A full description of the setup including the optical layout is given in the Supporting Information. At first, we studied a DNA hairpin structure consisting of a 40 basepair (bp) double-stranded DNA (dsDNA) stem and a 4 nucleotide (nt) apical loop (Figure 2A). The 5′ end of the hairpin is flanked by 10 nt single-stranded DNA (ssDNA) with a Cy3 fluorophore attached to the fourth base followed by a terminal biotin modification. The 3′ end is followed by an 11 nt ssDNA linker with a Cy5 dye attached to the sixth base. This linker is attached to a 5.9 kbp long dsDNA spacer and a 660 bp dsDNA fragment containing multiple digoxigenins. The digoxigenin-containing fragment allows binding of the DNA construct to an antidigoxigenin coated magnetic bead, while the 5.9 kbp spacer positions the bead out of the evanescent excitation field (see Supporting Information for a full description of sample preparation and experimental conditions). The DNA constructs with attached magnetic beads were flushed into the sample chamber and allowed to bind with the biotinylated DNA end to neutravidin, which was immobilized on the glass slide. When stretching the molecules by lowering the magnets, a mean contour length of L0 = 2.14 ± 0.05 μm was found in good agreement with the ∼6 kbp total length of the construct. At a force of 10.6 ± 0.2 pN, the hairpin underwent spontaneous and reversible opening and closing B
DOI: 10.1021/acs.nanolett.5b03956 Nano Lett. XXXX, XXX, XXX−XXX
Letter
Nano Letters
fluorescence signal was possible for several tens of seconds before photobleaching occurred (Figure 2B). To further challenge the spatiotemporal resolution achievable with our instrument, we repeated the experiment using highspeed positional tracking of the magnetic bead at 2 kHz with simultaneous fluorescence acquisition at 47.5 Hz. To cope with the high data throughput while maintaining real-time capability, the tracking of the magnetic bead was carried out in the GPU of the computer as recently described.29 Simultaneously recorded FRET efficiencies and DNA length values were again fully synchronous (Supporting Information Figures S2−S4). These frame-rates allow resolving very rapid transitions in particular for the DNA length measurements (Supporting Information Figures S3 and S4). Again, recording times of tens of seconds of the fluorescence signal were typical before photobleaching (Supporting Information Figures S3 and S4). To quantify the accuracy of the measured DNA length during the synchronous acquisition, we recorded several time trajectories of the differential coordinates between the DNAbound and surface-attached reference beads. At a force of 12.5 pN, the DNA length signal exhibited a root-mean-square (RMS) amplitude of 4 nm at the tracking rate of 2 kHz that decreased to 0.2 nm when averaged over 1 s, as revealed by calculating the Allan deviation (Supporting Information Figure S5B,D). This shows that simultaneous to FRET monitoring sub-nm resolution (corresponding to single base-pair resolution for the hairpin configuration) is achievable for DNA length measurements at sufficiently large forces. We note that the noise in the DNA length measurement is dominated by the thermal fluctuations of the DNA attached bead, because the difference coordinate between two surface attached particles was only 2 nm at 2 kHz and 0.07 nm when averaging over 1 s (Supporting Information Figure S5A,C). Consequently the accuracy of DNA length measurements will be reduced at lower forces, where thermal fluctuations are increased.28 In the previous experiments, both detection channels probed the same conformational change in order to verify their synchronous operation. In a second approach, we demonstrate that also independent, orthogonal information can be readily obtained. To this end, we studied the structural transitions within a DNA construct containing a four-strand Holliday Junction (HJ) (Figure 3A) at varying external forces. In absence of force, such a junction exhibits spontaneous conformational changes34 between two energetically equal conformers, socalled stacked-X structures. Notably, the relative occupancy of the two conformational states can be influenced by applying forces in the low pico-Newton range.8 To probe the conformation of the junction, two of the DNA strands were modified to incorporate FRET labels. A third arm comprised a terminal digoxigenin modification to allow attachment to the flow cell surface, while the fourth arm was attached to a 9.7 kbp dsDNA spacer with biotinylated nucleotides incorporated at the end, to facilitate an attachment to streptavidin-coated superparamagnetic beads (see Supporting Information). The conformational transition between the two conformers is relatively minute and in an orthogonal direction to the applied force (Figure 3A), rendering their observation with magnetic tweezers difficult if not impossible. We simultaneously monitored the DNA length as well as the FRET signal (at 32 Hz) of the Holliday junction construct similar to the previously described experiments. Applying alternating forces between 1.1 and 0.2 pN (Figure 3B) changes both the DNA extension and the FRET state distributions. The DNA was less elongated at
Figure 2. Spontaneous opening and closing of a DNA hairpin detected by simultaneous magnetic tweezers and FRET measurements. The hairpin structure is held under constant tension of 10.6 pN at which the hairpin fluctuates between a fully closed and a fully open state. (A) Illustration of the DNA sample, consisting of a 40 bp hairpin that is flanked by a 5.9 kb double-stranded DNA spacer. Donor and acceptor fluorophores for FRET detection are shown in green and red, respectively. At high force, the hairpin opens, while it is closed at low forces. (B) DNA length measured simultaneously with the fluorescence data shown at the acquisition rate of 300 Hz (in gray) and after adjacent average filtering to 10 Hz (in black). (C) Offsetcorrected fluorescence intensities over time of the donor and the acceptor fluorophores that are attached at the ssDNA ends of the hairpin recorded at a rate of 10 Hz. (D) Apparent FRET efficiency calculated from the trajectories in C. (E) The strict anticorrelation of the FRET and the length data verifies the fully synchronous acquisition of both signals.
transitions. This was evident in our positional tracking data of the magnetic bead (carried out in real-time at 300 Hz within the CPU of the computer), as abrupt changes of the DNA length of 37 ± 1.3 nm (Figure 2B), which is in agreement with earlier work.33 Simultaneously to the DNA length, FRET between the two dyes at the hairpin ends was measured with an acquisition rate of 10 Hz (Figure 2C,D). The observed FRET efficiency exhibited abrupt changes concurrently to the DNA length changes. When the hairpin was open (elongated DNA), the FRET efficiency was low (Eapp = 0.10 ± 0.01), while for the closed hairpin state the FRET efficiency was high (Eapp = 0.34 ± 0.01). This anticorrelation between DNA length and FRET efficiency was expected, since in the closed hairpin conformation the two fluorophores are in close proximity, while they are far apart in the open conformation. We verified the synchronicity of the two signals by calculating the crosscorrelation between both signals, which provided strong anticorrelation centered at 0 frames (Figure 2E). These results demonstrate that our CPU-based tracking is compatible with the simultaneous fluorescence acquisition, and that the two data streams are fully synchronous. Notably, the acquisition of the C
DOI: 10.1021/acs.nanolett.5b03956 Nano Lett. XXXX, XXX, XXX−XXX
Letter
Nano Letters
Figure 4. Force-dependent Holliday Junction dynamics: (A) Apparent FRET efficiency (gray) and force (red) applied of a HJ molecule. The red arrow at 76 s indicates the final bleaching event. (B) Forcedependent FRET efficiency histograms of the constant-force subsections from A, showing a decreasing population of the highFRET state with increasing externally applied force. (C) Stateprogression path for the 0.6 pN subsection determined by vbFRET to determine force-dependent rates (apparent FRET efficiency in gray, state transition path in red). (D) Force-dependence of the state transition rates between the two conformations of the Holliday junction. Red and gray circles represent rates for the low and the high FRET state are shown as red and gray circles, respectively. Solid lines show exponential fits to the data. Triangles at zero force represent force-free rates from a tether-free molecule under the same experimental conditions. All data (except zero-force data in D) in this figure was obtained on a single DNA molecule.
Figure 3. Simultaneous length and FRET measurements on a DNA construct containing a Holliday junction. (A) Sketch of the DNA Holliday Junction construct, which exhibits salt- and force-dependent conformational oscillations. (B) Relative extension of the DNA molecule over time shown at the acquisition rate of 300 Hz (gray) and after adjacent average filtering to 32 Hz (black). The amplitude of the extension fluctuations is force-dependent (forces indicated within each section are separated by dashed lines) with greater amplitudes corresponding to lower applied forces. (C) Apparent FRET efficiency measured simultaneously with the DNA extension. (D) FRET efficiency histograms of the four individual regions of high and low applied forces, showing reproducible manipulation of Holliday Junction oscillations.
transition rates between the high and low FRET state were obtained. The state transition rates obtained from a single molecule were strongly force dependent and in good agreement with previously determined energy landscape parameters obtained from multiple molecules8 (Figure 4D). As fluorophore lifetimes are statistically distributed and single-molecule experiments are by definition based on individual stochastic events, maximizing the number of investigated molecules is highly desirable. Magnetic tweezers and single-molecule FRET measurements are both suited for parallelization.23,36 While for parallel FRET detection only a sufficiently large field of view needs to be recorded, parallel magnetic tweezers detection requires a parallelized particle tracking.29,37 Here we demonstrate such a parallelized mechanical experiment with simultaneous FRET detection and monitor the force-dependent conformational changes of two Holliday junctions simultaneously (Figure 5). Histograms of the FRET efficiency for the trajectory subsections with equal force showed for both molecules the expected force dependency of the Holliday junction conformations (see above). Interestingly, for molecule 1 (Figure 5A) the high FRET state was more favored at low force, than for molecule 2 (Figure 5B). This difference was reproducibly observed when applying the low force a second time (Figure 5). At around t = 15 s, molecule 2 showed a spike in the FRET efficiency, beyond which the FRET histogram significantly changed (blue arrows/
low force and more elongated at high force, which is in agreement with the typical force−extension behavior of DNA. The length change of the DNA tether between the two applied forces was about 1 μm for this particular construct (Figure 3B). While the DNA length signal monitors the global conformation of the whole construct, the simultaneously recorded FRET signal revealed the local conformation of only the Holliday junction (Figure 3C). At 0.2 pN applied force, the Holliday junction showed a two-state behavior with a high-FRET efficiency peak around E = 0.57 ± 0.09 and a low FRET efficiency peak around E = 0.19 ± 0.06 (Figure 4B). Increasing the force to 1.1 pN shifted the equilibrium toward the low FRET efficiency peak (Figure 3D) as previously described.8 Next, we explored the photostability of FRET pairs in our instrument showing that photobleaching in magnetic tweezers is reduced compared to optical tweezers. To illustrate the benefit, we subsequently applied four different forces on a Holliday junction molecule before photobleaching occurred at 76 s to obtain force-dependent conformational kinetics on a single molecule (Figure 4A), a big improvement compared to previous studies, where only a single force was probed per molecule.8 We analyzed the FRET efficiency time trajectories at a given force using a variational Bayesian analysis (vbFRET)35 to extract the corresponding two-state transition paths (Figure 4C). This analysis yielded the transition matrices from which D
DOI: 10.1021/acs.nanolett.5b03956 Nano Lett. XXXX, XXX, XXX−XXX
Letter
Nano Letters
themselves limit the resolution of DNA length measurements8 or interleaved trapping and excitation.11 In contrast to previous approaches,27 we achieved synchronization down to 3 ms, a fraction of a single camera frame of the fluorescence detection camera (see Supporting Information and Figures S2 and S4). This allows to quantify even minute delays of subsequent steps in biomolecular reactions, such as the binding of a protein to DNA (FRET detection) and the subsequent DNA processing (tweezers detection). We also demonstrated that our hybrid measurements can be parallelized with higher degrees of parallelization achievable by higher surface densities of the tethers.23 Taken together, these achievements maximize the amount of information attainable from one single-molecule experiment. In addition, we note that in our hybrid instrument twisting of single tethers and torque measurements are readily supported and even minute structural changes can readily be observed by the combination with single-molecule FRET. Overall, we anticipate many new applications coming within reach by simultaneous magnetic tweezers and FRET detection.
■
ASSOCIATED CONTENT
S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.nanolett.5b03956. Detailed method description as well as high-resolution magnetic tweezers data. (PDF)
■
Figure 5. Simultaneous recording of force-dependent oscillations for two Holliday junction constructs molecule 1 (A) and molecule 2 (B). Shown are apparent FRET efficiency trajectories (gray lines) and histograms including double-Gaussian fits. The dashed lines in the trajectory plots separate sections of constant force (as indicated in the plots). Histograms for these sections are provided directly above. In B, the histogram on the right side (blue fit curve) indicates the changed behavior after the donor spike at 15 s (between the blue arrows, and before the fluorophore bleaching event marked by the red arrow).
AUTHOR INFORMATION
Corresponding Authors
*E-mail:
[email protected]. *E-mail:
[email protected]. Author Contributions
D.J.K., F.E.K., M.S., F.W.S., R.S., and M.S. designed and constructed the hybrid instruments; F.E.K., M.S., R.S., and M.S. designed experiments, F.E.K. and M.S. performed experiments, M.S.G. provided samples, S.H. developed software, F.E.K., M.S., R.S., and M.S. analyzed results and wrote the manuscript. All authors commented on the manuscript. F.E.K. and M.S contributed equally to this work.
lines in Figure 5) and subsequently resembled the histogram of molecule 1 at 0.2 pN. The initially altered FRET state of molecule 2 may have originated from an additional interaction that suddenly changed. Our technique thus allows tracking minute details in the behavior of multiple individual molecules and to extract information about heterogeneities. Here, the maximum number of parallel measurements is limited by the number of tethered DNA molecules bearing an intact FRET pair rather than by the software (note that a total of N = 7 fieldof-views with two force-dependent Holliday junctions were studied), which supports full frame fluorescence image acquisition at 32 Hz and parallel tracking of up to 10 magnetic beads per field of view. In conclusion, we have demonstrated the simultaneous manipulation and detection of single molecule conformation using a hybrid experimental setup, employing the capabilities of both high-resolution magnetic tweezers and single-molecule FRET. Using DNA nanostructures, we could either steer the conformational ticking of a Holliday Junction, with FRET providing information orthogonal to the DNA tether extension, or show DNA hairpin opening both in tracking and FRET experiments. Because magnetic tweezers avoid the generation of oxygen radicals and avoid high laser powers, long lifetimes of the fluorophores are obtained, without any workarounds as required for optical tweezers such as long DNA molecules that
Funding
This work was supported by an ERC starting Grant (No. 261224) to R.S. and the German Ministry for Science and Education (BMBF 03Z2EN11) to M.S.. Notes
The authors declare no competing financial interest.
■ ■
ACKNOWLEDGMENTS We thank all members of the Seidel and Schlierf groups for helpful comments and discussions. REFERENCES
(1) Ha, T. Nat. Methods 2014, 11 (10), 1015−8. (2) Bustamante, C.; Cheng, W.; Mejia, Y. X. Cell 2011, 144 (4), 480−97. (3) De Vlaminck, I.; Dekker, C. Annu. Rev. Biophys. 2012, 41, 453− 72. (4) Hohng, S.; Lee, S.; Lee, J.; Jo, M. H. Chem. Soc. Rev. 2014, 43 (4), 1007−13. (5) Kozankiewicz, B.; Orrit, M. Chem. Soc. Rev. 2014, 43 (4), 1029− 43. E
DOI: 10.1021/acs.nanolett.5b03956 Nano Lett. XXXX, XXX, XXX−XXX
Letter
Nano Letters (6) Klaue, D.; Kobbe, D.; Kemmerich, F.; Kozikowska, A.; Puchta, H.; Seidel, R. Nat. Commun. 2013, 4, 2024. (7) Yodh, J. G.; Schlierf, M.; Ha, T. Q. Rev. Biophys. 2010, 43 (2), 185−217. (8) Hohng, S.; Zhou, R.; Nahas, M. K.; Yu, J.; Schulten, K.; Lilley, D. M.; Ha, T. Science 2007, 318 (5848), 279−83. (9) van Mameren, J.; Modesti, M.; Kanaar, R.; Wyman, C.; Peterman, E. J.; Wuite, G. J. Nature 2009, 457 (7230), 745−8. (10) Gross, P.; Farge, G.; Peterman, E. J.; Wuite, G. J. Methods Enzymol. 2010, 475, 427−53. (11) Comstock, M. J.; Ha, T.; Chemla, Y. R. Nat. Methods 2011, 8 (4), 335−40. (12) Zhou, R.; Schlierf, M.; Ha, T. Methods Enzymol. 2010, 475, 405−26. (13) Brutzer, H.; Schwarz, F. W.; Seidel, R. Nano Lett. 2012, 12 (1), 473−8. (14) Schwarz, F. W.; Toth, J.; van Aelst, K.; Cui, G.; Clausing, S.; Szczelkun, M. D.; Seidel, R. Science 2013, 340 (6130), 353−6. (15) Graves, E. T.; Duboc, C.; Fan, J.; Stransky, F.; Leroux-Coyau, M.; Strick, T. R. Nat. Struct. Mol. Biol. 2015, 22 (6), 452−7. (16) Ghoneim, M.; Spies, M. Nano Lett. 2014, 14 (10), 5920−31. (17) Lang, M. J.; Fordyce, P. M.; Block, S. M. J. Biol. 2003, 2 (1), 6. (18) Landry, M. P.; McCall, P. M.; Qi, Z.; Chemla, Y. R. Biophys. J. 2009, 97 (8), 2128−36. (19) Vogelsang, J.; Cordes, T.; Tinnefeld, P. Photochem. Photobiol. Sci. 2009, 8 (4), 486−96. (20) Swoboda, M.; Henig, J.; Cheng, H. M.; Brugger, D.; Haltrich, D.; Plumere, N.; Schlierf, M. ACS Nano 2012, 6 (7), 6364−9. (21) Strick, T. R.; Allemand, J. F.; Bensimon, D.; Croquette, V. Biophys. J. 1998, 74 (4), 2016−28. (22) Lipfert, J.; Kerssemakers, J. W.; Jager, T.; Dekker, N. H. Nat. Methods 2010, 7 (12), 977−80. (23) De Vlaminck, I.; Henighan, T.; van Loenhout, M. T.; Pfeiffer, I.; Huijts, J.; Kerssemakers, J. W.; Katan, A. J.; van Langen-Suurling, A.; van der Drift, E.; Wyman, C.; Dekker, C. Nano Lett. 2011, 11 (12), 5489−93. (24) Bae, W.; Kim, K.; Min, D.; Ryu, J. K.; Hyeon, C.; Yoon, T. Y. Nat. Commun. 2014, 5, 5654. (25) Long, X.; Parks, J. W.; Bagshaw, C. R.; Stone, M. D. Nucleic Acids Res. 2013, 41 (4), 2746−55. (26) Shroff, H.; Reinhard, B. M.; Siu, M.; Agarwal, H.; Spakowitz, A.; Liphardt, J. Nano Lett. 2005, 5 (7), 1509−14. (27) Lee, M.; Kim, S. H.; Hong, S. C. Proc. Natl. Acad. Sci. U. S. A. 2010, 107 (11), 4985−90. (28) Klaue, D.; Seidel, R. Phys. Rev. Lett. 2009, 102 (2), 028302. (29) Huhle, A.; Klaue, D.; Brutzer, H.; Daldrop, P.; Joo, S.; Otto, O.; Keyser, U. F.; Seidel, R. Nat. Commun. 2015, 6, 5885. (30) Gosse, C.; Croquette, V. Biophys. J. 2002, 82 (6), 3314−29. (31) Daldrop, P.; Brutzer, H.; Huhle, A.; Kauert, D. J.; Seidel, R. Biophys. J. 2015, 108 (10), 2550−61. (32) Roy, R.; Hohng, S.; Ha, T. Nat. Methods 2008, 5 (6), 507−16. (33) Woodside, M. T.; Behnke-Parks, W. M.; Larizadeh, K.; Travers, K.; Herschlag, D.; Block, S. M. Proc. Natl. Acad. Sci. U. S. A. 2006, 103 (16), 6190−5. (34) Buranachai, C.; McKinney, S. A.; Ha, T. Nano Lett. 2006, 6 (3), 496−500. (35) Bronson, J. E.; Fei, J.; Hofman, J. M.; Gonzalez, R. L., Jr.; Wiggins, C. H. Biophys. J. 2009, 97 (12), 3196−205. (36) Ribeck, N.; Saleh, O. A. Rev. Sci. Instrum. 2008, 79 (9), 094301. (37) Ramanathan, S. P.; van Aelst, K.; Sears, A.; Peakman, L. J.; Diffin, F. M.; Szczelkun, M. D.; Seidel, R. Proc. Natl. Acad. Sci. U. S. A. 2009, 106 (6), 1748−53.
F
DOI: 10.1021/acs.nanolett.5b03956 Nano Lett. XXXX, XXX, XXX−XXX
