Single-Molecule Positioning in Zeromode Waveguides by DNA

Apr 29, 2014 - Nanotechnology is challenged by the need to connect top-down produced nanostructures with the bottom-up world of chemistry. A nanobiote...
1 downloads 12 Views 3MB Size
Download PDF
Letter pubs.acs.org/NanoLett

Single-Molecule Positioning in Zeromode Waveguides by DNA Origami Nanoadapters Enrico Pibiri, Phil Holzmeister, Birka Lalkens, Guillermo P. Acuna,* and Philip Tinnefeld* NanoBioSciences Group, Institute for Physical and Theoretical Chemistry Braunschweig, Hans-Sommer-Strasse 10, 38106 Braunschweig, Lower Saxony, Germany S Supporting Information *

ABSTRACT: Nanotechnology is challenged by the need to connect top-down produced nanostructures with the bottomup world of chemistry. A nanobiotechnological prime example is the positioning of single polymerase molecules in small holes in metal films, so-called zeromode waveguides (ZMWs), which is required for single-molecule real-time DNA sequencing. In this work, we present nanoadapters made of DNA (DNA origami) that match the size of the holes so that exactly one nanoadapter fits in each hole. By site-selective functionalization of the DNA origami nanoadapters, we placed single dye molecules in the ZMWs, thus optimizing the hole usage and improving the photophysical properties of dyes compared to stochastically immobilized molecules. KEYWORDS: Fluorescence, DNA origami, single molecule, zeromode waveguides, nanophotonics

S

metal cladding can, for example, lead to altered (plasmonic) excitation, and the fluorescence process can be modified by quenching through energy or electron transfer to the metal. Furthermore, the radiative rate of the fluorophores can be reduced or enhanced. A recent study has indicated that these processes are strongly spatially dependent, and the brightest fluorescence in nanoapertures of 375 nm diameter was obtained in their center.12 Obviously, a method is needed for the paralleled positioning of molecules in the center of the ZMWs. Here we introduce DNA origami nanoadapters to overcome Poissonian statistics for filling ZMWs and to position molecules close to the centers of ZMWs with homogenized spectroscopic properties. We designed a disk-shape DNA origami of ∼62 nm diameter that fits into the nanoapertures and that blocks further binding of other nanoadapters in the ZMW (see Figure 1a and Figure S1b). We used commercially available ZMWs with holes in a 100 nm aluminum coating ranging from 85 to 200 nm in diameter. Because the DNA origami disk is smaller in diameter than the smallest ZMWs (see Figure 1b), we also used a larger rectangular DNA origami to demonstrate the size exclusion (Figure 1a and Figure S1a). To control the orientation of the DNA origami nanoadapters when binding in the ZMWs, we developed a steric hindrance strategy that only allows the DNA origami to bind irreversibly when they are oriented flat on the bottom of the ZMW (Figure 1c). We attached the fluorescent dye ATTO647N to the DNA

ingle-molecule techniques have a substantial impact on nanobiotechnological developments such as superresolution microscopy or single-molecule DNA sequencing.1−3 It has been argued that the key to success for such breakthrough technologies is less the sophistication to detect a single molecule, but rather the challenge not to detect the molecules around it.4−6 One successful approach to exclude background molecules has been the use of tiny holes in a ∼100 nm thick metal layer deposited on a glass coverslip.7,8 These holes of subwavelength diameter do not support light propagation and have therefore been denoted zeromode waveguides (ZMWs or nanoapertures).7 Within ZMWs, evanescent light is confined to a minimal volume close to the glass coverslip so that single fluorescent molecules can be observed at biologically relevant concentrations in the nano- to micromolar range. These nanophotonic structures have enabled breakthroughs in singlemolecule real-time DNA sequencing and in the study of translation.3,9−11 To exploit the full potential of ZMWs, however, two challenges have to be met. First, single-molecule techniques commonly require exactly one molecule such as one polymerase molecule for sequencing per ZMW. More than one polymerase will quickly lead to dephasing so that the sequence information gets lost. With the exception of serial loading by an atomic force microscope,12 the loading of ZMWs is controlled by Poissonian statistics which implies a theoretical maximum of 37% single occupancy. This problem is connected to the general nanotechnological challenge of connecting top-down production with the bottom-up molecular world of chemistry and biology. Second, the fluorescence of single molecules inside ZMWs is not well-understood with most of the studies focusing on an FCS analysis averaging over the ZMW volume.13,14 The © 2014 American Chemical Society

Received: March 21, 2014 Revised: April 17, 2014 Published: April 29, 2014 3499

dx.doi.org/10.1021/nl501064b | Nano Lett. 2014, 14, 3499−3503

Nano Letters

Letter

Figure 1. (a) Sketches of the top view and bottom view of the DNA origami disk and the rectangular-shaped DNA origami employed. Selected staples were modified to incorporate a single ATTO647N dye or biotin strands as indicated in red and yellow, respectively. (b) Sketch of the ZMWs with diameters between 85 and 200 nm. Due to the size of the structures both the disk-shaped DNA origami and the ds-DNA sample can enter all of the ZMWs; however, the rectangular structure has a diagonal of 122 nm, bigger than the smaller nanoapertures. (c) Immobilization strategy. The biotin modifications close to the center of the rigid DNA origami guarantee irreversible binding to a neutravidin-functionalized glass surface at the desired position and orientation.

Figure 2. (a) Intensity transients of dyes on rectangular DNA origamis within the ZMWs with one and two bleaching steps. The inset shows the confocal fluorescence image with the corresponding spots highlighted. (b) Single and double occupation as a function of ZMW diameter. The cutoff diameter for the rectangular DNA origami is indicated. (c) Comparison between optimal Poissonian and measured occupation distribution for the 200 nm ZMWs.

selected staple strands that terminate on the opposing site of the DNA origami were modified with biotins that bind to the neutravidin modified surfaces. ZMWs were used as obtained and according to the suggestions of the manufacturer.9 The walls of the ZMWs are coated with polyvinylphosphonic acid (PVPA) to protect against unspecific adsorption.21 The ZMW bottom glass surface is derivatized with biotin−PEG (poly(ethylene glycol)) to mediate immobilization of the target molecules. The approach of placing exactly single objects in ZMWs is based on using DNA origamis as adapters so that exactly one DNA origami fits into one ZMW by size exclusion (see scheme in Figure 1b). This idea allows using a high concentration of DNA origami carrying the biomolecular assay of interest so that the ZMW occupation can be saturated. On the other hand, the sizing unit and the immobilization method have to ensure that the biomolecular assay is placed close to the glass surface and optimally at the center of the ZMW. This implies that also the orientation of the DNA origami in the ZMW has to be controlled. To achieve this, we used a steric hindrance approach by employing rigid sizing units such as a rectangular DNA origami or a DNA origami disk that can be site-specifically modified.18,22 We attached the biotin anchors for surface

origami that reports on the presence of the DNA origami in the ZMWs. We compared the properties of stochastically immobilized dye molecules (see Figure 1b) in the ZMW with those on the DNA origami and found a clear distinction and an interesting size dependence of the photophysical properties of individual dyes. Electromagnetic simulations and geometrical considerations were used to explain the results and underline the success of the immobilization strategy. Results and Discussion. For the DNA origami nanoadapters, a new disk-shaped DNA origami was constructed. DNA origami are DNA nanostructures which are folded from a ∼8000 nucleotides (nt) long M13mp18 single strand with the aid of ∼200 so-called staple strands15,16 and were designed with the software caDNAno17 (see the Supporting Information for sequences). The DNA origami disk consists of an array of sixhelix bundles (6hb) which makes it rigid and slightly elliptical with a 65 nm long axis and a 57 nm short axis (see Figure 1a). Alternatively, we used a larger, rectangular DNA origami with dimensions of 100 × 70 nm2.15,18 As a functional unit, we attached a fluorescent dye on the DNA origami by using a dyelabeled staple strand (ATTO647N). Alternatively, staple strands could be modified to host biomolecular assays such as a polymerase for DNA sequencing.3,19,20 For immobilization, 3500

dx.doi.org/10.1021/nl501064b | Nano Lett. 2014, 14, 3499−3503

Nano Letters

Letter

molecules by a factor of ∼10 comparable to previously reported results12 and a generally reduced fluorescence lifetime. These were lowest for the stochastically immobilized ds-DNA (1.5 ± 0.4 ns, Figure 3a) and longer for dyes immobilized via the DNA

immobilization only close to the center of the DNA origami nanoadapters so that irreversible neutravidin−biotin bond formation can only occur when the orientation of the DNA origami structure is almost parallel with respect to the surface (see Figure 1c). This also ensures that functional units are placed at specific positions and that the bottom of the ZMWs is blocked against other DNA origami binding events. Because the DNA origami disk has a smaller diameter than the smallest available ZMWs and can enter all of the ZMWs (Figure 1b), we used the rectangular DNA origami to demonstrate the size exclusion effect of DNA origami placement. ZMWs were incubated for 30 min with a 45 nM solution of rectangular DNA origami carrying one fluorescent dye and two biotins as indicated in Figure 1a. We determined the occupation by counting the number of occupied ZMWs with respect to all ZMWs in the area studied (see scan image in the inset of Figure 2a). After each scan, each occupied ZMW was placed in the laser focus to determine the number of dye molecules by counting bleaching steps (see exemplary transients in Figure 2a). When no molecule was detected in the transient after it was detected on the scan image, this ZMW was assigned to the one-molecule population as the simultaneous photobleaching of two molecules during the scan or while approaching the ZMW is unlikely. Below 120 nm diameter, ZMWs remained completely empty as expected for a 122 nm diagonal of the rectangular DNA origami (Figure 2b). Above 120 nm, the occupation quickly increased and saturated above 60% for 200 nm ZMWs. At above 140 nm diameter, we also note a fraction of doubly occupied ZMWs that increases up to 13% for the largest ZMW diameter. Still, this fraction is substantially smaller than what is expected from Poissonian statistics, and single-molecule occupancy can clearly be increased not only compared to previously reported stochastic immobilization in the range between 20 and 35%7,12 but also beyond the Poissonian limit (Figure 2c). As control, we carried out the same experiment but without biotin modifications in the DNA origami. In this case, no binding was detected (no molecule in 225 ZMWs) assuring that the DNA origamis are specifically bound via the biotins. We do not know why we could not load all ZMWs even with longer incubation time but speculate that this might be related to the density of biotins on the glass surface of the ZMWs. The DNA origami with two biotins can only sample a small fraction of the ZMWs’ glass surface, and on this area there might not be any neutravidin anchoring point. This idea is supported by the fact that occupation could be saturated with double-stranded DNA as well as with the smaller DNA origami disk for which subPoissonian binding distributions could also be obtained (see Figure S2). Next, we studied how the immobilization via the DNA origami disk nanoadapters influences the spectroscopic properties of the dyes in the ZMWs. As the dyes were placed ∼15 nm away from the edge of the DNA origami the accessible area for the dyes was reduced, and we therefore expected more homogeneous spectroscopic properties. For the characterization of the dye’s photophysics, we recorded fluorescence transients of the dyes immobilized in the ZMWs (Figure 2a) and concurrently recorded fluorescence lifetime decays.23 By deconvolution with the instrument response function, we determined the fluorescence lifetime. On glass coverslips covered with BSA−biotin−neutravidin, ATTO647N on DNA exhibited a fluorescence lifetime of 4.0 ± 0.3 ns (see Figure S3). For dyes in ZMWs, we found lower brightnesses of the

Figure 3. Scatter plot of fluorescence intensity versus fluorescence lifetime with the corresponding histograms for the DNA origami disk (black) and the ds-DNA (red) immobilized in ZMWs of 100 nm (a) and 200 nm (b) diameter.

origami disk. A lifetime of 2.4 ± 0.3 ns was found in the ZMWs of 200 nm diameter (Figure 3b) and 3.2 ± 0.5 ns for the 100 nm ZMWs (Figure 3a). Usually, fluorescence lifetime distributions are broader for longer decay times,23 but in this case the fluorescence lifetime exhibit a similar homogeneity. The main surprising findings are that the fluorescence lifetime populations of ds-DNA immobilization and DNA origami disk immobilization exhibit a fairly small overlap and that the fluorescence lifetime is smaller in the 200 nm ZMWs than in the 100 nm ZMWs. To verify that the change in the fluorescence lifetime is not related to the immobilization on the DNA origami, we compared the properties of the dye attached to glass coverslips via ds-DNA or via the DNA origami and found no differences (see Figure S3). To understand the different properties of dyes immobilized via the DNA origami and those stochastically distributed in ZMWs, we carried out electromagnetic FDTD simulations24 in frequency mode. These simulations yielded the fluorescence lifetime normalized to the fluorescence lifetime of the dye 3501

dx.doi.org/10.1021/nl501064b | Nano Lett. 2014, 14, 3499−3503

Nano Letters

Letter

Figure 3. For both ZMWs diameters considered, the shorter fluorescence lifetimes for the stochastic ds-DNA immobilization were reproduced, and the separation of the fluorescence lifetime distribution could partially be explained. The cutoff at short fluorescence lifetimes for the experimental ds-DNA distribution can be explained by a minimum spacing of the dsDNA to the metallic walls due to the PVPA metal coating and because very dim molecules might not be detected. Strikingly, the simulations also reproduce the nonintuitive, longer fluorescence lifetimes in the 100 nm ZMWs compared to the 200 nm ZMWs which can qualitatively be explained by the reduced photonic mode density in the smaller ZMWs. Conclusion. Zeromode waveguides are a key technology to take single-molecule studies to the main stream of biological interaction and dynamics studies. In this work, we present DNA origami nanoadapters that overcome the Poisson-limited positioning of single molecules in these nanophotonic structures. The practical usage of ZMWs could thus be improved by a factor of about two. In addition, the DNA origami nanoadapters restrict the positioning of functional units to a more defined area. This ability was exploited to learn about the nanophotonic environment in ZMWs, and we found unexpected changes of the fluorescence properties of the DNA origami nanoadapters compared to stochastically immobilized dye molecules. The different fluorescence lifetime distribution and increased fluorescence lifetimes in smaller ZMWs were rationalized using electromagnetic simulations. The DNA origami nanoadapters bridge the gap between top-down nanolithography and bottom-up chemical synthesis,27 and ideas developed here including the steric hindrance approach for controlling orientation might be applicable to a wider range of nanotechnological problems including molecular electronics and nanoscale light control.28,29 Materials and Methods. Sample Preparation. For details of the design and folding of the DNA-origami see the SI. We used ZMWs from Pacific Biosciences (noncommercial zeromode waveguide arrays), where the array chip is mounted on a conventional microscope coverslip. Each chip provides six different ZMW diameters ranging from 85 to 200 nm. The glass bottom of the ZMW chip is already functionalized with PEG− biotin to allow immobilization of neutravidin.30 To prevent nonspecific binding the aluminum surface is passivated with a phosphonate.9 The chip is washed twice with MOPS before incubation with 1 mg/mL neutravidin in MOPS buffer for half an hour (50 mM MOPS, 75 mM KOAc, and 12.5 mM MgOAc, pH 7). After five more washing steps to remove unbound neutravidin, the DNA origami sample is added, and the binding process is controlled by confocal imaging. Once a reasonable occupancy is reached, the sample is immediately washed to remove unbound DNA structures. The arrays of ZMWs were used no longer than 10 h after incubation with neutravidin. After that time, we have noticed that the apertures start to degrade, despite our choice of a chloride free buffer. Confocal Measurement and Analysis. All samples were measured with a custom built confocal setup based on an inverted microscope (IX71, Olympus) with a high NA oil immersion objective (100×/NA 1.40, UPLSAPO100XO, Olympus). DNA origami with the dye ATTO647N were excited at 640 nm with an 80 MHz pulsed laser diode (LDH-DC-640, Picoquant, 20 μW). The polarization of the excitation light was aligned to circular polarization by a quarter wave plate (AQWP05M-600, Thorlabs) in order to avoid polarization effects. The chip with immobilized DNA origamis was imaged

immobilized on glass for a dye placed at different positions within the ZMW and averaged for all three orientations with equal weight25,26 (see the SI for details on the simulations). To compare the spatially dependent properties of the dye in the ZMW with the experimental data, we additionally have to account for the probability distribution of finding a dye at a certain distance to the ZMW center. For ds-DNA immobilization, the dye can be found at each position in the ZMW at equal probability, yielding a linear probability function (inset in Figure 4a,b, red data points). The size exclusion by the DNA

Figure 4. Histogram of the simulated fluorescence lifetime distribution for the DNA origami disk and the ds-DNA immobilized in ZMWs of 100 nm (a) and 200 nm (b) diameter, normalized to the reference fluorescence lifetime. Both insets show the normalized probability distribution of finding the dye at a certain distance to the ZMW center when the dye is immobilized via the DNA origami disk (black) and the ds-DNA (red). The dyes belonging to the ds-DNA and the DNA origami disk are assumed to be placed at a depth of 5 and 15 nm, respectively.

origami disk prohibits the dye to be closer than ∼15 nm to the metal walls and also yields a more complex probability function with a maximum probability of finding the dye at a distance of ∼20 nm (for the 100 nm ZMWs, Figure 4a) or ∼50 nm from the center of the ZMW (for the 200 nm ZMWs, Figure 4b). By combining the spatially dependent properties of the dye with the probability distribution of finding the dye at a certain position within the ZMW, we obtained theoretical fluorescence lifetime distributions (Figure 4a,b) that can be compared to the experimental fluorescence lifetime distributions presented in 3502

dx.doi.org/10.1021/nl501064b | Nano Lett. 2014, 14, 3499−3503

Nano Letters

Letter

(7) Levene, M. J.; Korlach, J.; Turner, S. W.; Foquet, M.; Craighead, H. G.; Webb, W. W. Science 2003, 299 (5607), 682−686. (8) Zhu, P.; Craighead, H. G. Annu. Rev. Biophys. 2012, 41, 269−93. (9) Flusberg, B. A.; Webster, D. R.; Lee, J. H.; Travers, K. J.; Olivares, E. C.; Clark, T. A.; Korlach, J.; Turner, S. W. Nat. Methods 2010, 7 (6), 461−5. (10) Chen, J.; Dalal, R. V.; Petrov, A. N.; Tsai, A.; O’Leary, S. E.; Chapin, K.; Cheng, J.; Ewan, M.; Hsiung, P. L.; Lundquist, P.; Turner, S. W.; Hsu, D. R.; Puglisi, J. D. Proc. Natl. Acad. Sci. U.S.A. 2014, 111 (2), 664−9. (11) Tsai, A.; Petrov, A.; Marshall, R. A.; Korlach, J.; Uemura, S.; Puglisi, J. D. Nature 2012, 487 (7407), 390−3. (12) Heucke, S.; Baumann, F.; Acuna, G. P.; Severin, P. M.; Stahl, S. W.; Strackharn, M.; Stein, I.; Altpeter, P.; Tinnefeld, P.; Gaub, H. E. Nano Lett. 2014, 14 (2), 391−395. (13) Wenger, J.; Gerard, D.; Dintinger, J.; Mahboub, O.; Bonod, N.; Popov, E.; Ebbesen, T. W.; Rigneault, H. Opt. Express 2008, 16 (5), 3008−20. (14) Rigneault, H.; Capoulade, J.; Dintinger, J.; Wenger, J.; Bonod, N.; Popov, E.; Ebbesen, T. W.; Lenne, P. F. Phys. Rev. Lett. 2005, 95 (11), 117401. (15) Rothemund, P. W. Nature 2006, 440 (7082), 297−302. (16) Douglas, S. M.; Dietz, H.; Liedl, T.; Hogberg, B.; Graf, F.; Shih, W. M. Nature 2009, 459 (7245), 414−8. (17) Douglas, S. M.; Marblestone, A. H.; Teerapittayanon, S.; Vazquez, A.; Church, G. M.; Shih, W. M. Nucleic Acids Res. 2009, 37 (15), 5001−6. (18) Schmied, J. J.; Raab, M.; Forthmann, C.; Pibiri, E.; Wünsch, B.; Dammeyer, T.; Tinnefeld, P. Nat. Protocols 2014, accepted. (19) Rajendran, A.; Endo, M.; Sugiyama, H. Angew. Chem. 2012, 51 (4), 874−90. (20) Gietl, A.; Holzmeister, P.; Grohmann, D.; Tinnefeld, P. Nucleic Acids Res. 2012, 40 (14), e110. (21) Korlach, J.; Marks, P. J.; Cicero, R. L.; Gray, J. J.; Murphy, D. L.; Roitman, D. B.; Pham, T. T.; Otto, G. A.; Foquet, M.; Turner, S. W. Proc. Natl. Acad. Sci. U.S.A. 2008, 105 (4), 1176−1181. (22) Steinhauer, C.; Jungmann, R.; Sobey, T. L.; Simmel, F. C.; Tinnefeld, P. Angew. Chem. 2009, 48 (47), 8870−8873. (23) Tinnefeld, P.; Herten, D. P.; Sauer, M. J. Phys. Chem. A 2001, 105 (34), 7989−8003. (24) Acuna, G. P.; Holzmeister, P.; Möller, F. M.; Beater, S.; Lalkens, B.; Tinnefeld, P. J. Biomed. Opt. 2013, 18 (6), 65001. (25) Acuna, G. P.; Möller, F. M.; Holzmeister, P.; Beater, S.; Lalkens, B.; Tinnefeld, P. Science 2012, 338 (6106), 506−10. (26) Taminiau, T. H.; Stefani, F. D.; van Hulst, N. F. New J. Phys. 2008, 10, 105005. (27) Kershner, R. J.; Bozano, L. D.; Micheel, C. M.; Hung, A. M.; Fornof, A. R.; Cha, J. N.; Rettner, C. T.; Bersani, M.; Frommer, J.; Rothemund, P. W.; Wallraff, G. M. Nat. Nanotechnol. 2009, 4 (9), 557−61. (28) Hung, A. M.; Micheel, C. M.; Bozano, L. D.; Osterbur, L. W.; Wallraff, G. M.; Cha, J. N. Nat. Nanotechnol. 2010, 5 (2), 121−6. (29) Eskelinen, A. P.; Kuzyk, A.; Kaltiaisenaho, T. K.; Timmermans, M. Y.; Nasibulin, A. G.; Kauppinen, E. I.; Torma, P. Small 2011, 7 (6), 746−50. (30) Korlach, J.; Bjornson, K. P.; Chaudhuri, B. P.; Cicero, R. L.; Flusberg, B. A.; Gray, J. J.; Holden, D.; Saxena, R.; Wegener, J.; Turner, S. W. Methods Enzymol. 2010, 472, 431−55.

by scanning the sample with a piezo stage (P-517.3CL, Physik Instrumente). From the scan image, molecules are selected and then automatically placed in the laser focus for time-resolved analysis. The resulting fluorescence was collected by the same objective and separated from the excitation light by appropriate spectral filtering.25 The signal is detected by a single photon counting module (τ-SPAD 100, Picoquant) and a PC-card for time-correlated single-photon counting (SPC-830, Becker & Hickl). The data are analyzed with custom-made software (LabView2009, National Instruments) and the commercial FluoFit software (Picoquant).

■

ASSOCIATED CONTENT

S Supporting Information *

Details on the preparation of the different samples, AFM images of the rectangle and the disk structure (Figure S1), the dependence with the ZMW diameter of the single and double occupation of the origami disk together with a comparison with the Poisson distribution (Figure S2), fluorescence reference measurements on glass slides (Figure S3), details on the electromagnetic simulations, and a list of the staples strands employed for the origami structures. This material is available free of charge via the Internet at http://pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected] (G.P.A.). *E-mail: [email protected] (P.T.). Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The authors are grateful to Andreas Gietl for stimulating discussion and to Frank Demming from CST for assistance with the numerical simulations. This work was supported by a starting grant (SiMBA, EU 261162) of the European Research Council (ERC) and the Volkswagen Foundation (8641586416). P.H. was supported by Studienstiftung des Deutschen Volkes. TU Braunschweig, E.P., G.P.A., and P.T. have filed a provisional patent application on the described method of positioning molecules using DNA origami nanoadapters.

■

REFERENCES

(1) Hell, S. W. Science 2007, 316 (5828), 1153−1158. (2) Harris, T. D.; Buzby, P. R.; Babcock, H.; Beer, E.; Bowers, J.; Braslavsky, I.; Causey, M.; Colonell, J.; Dimeo, J.; Efcavitch, J. W.; Giladi, E.; Gill, J.; Healy, J.; Jarosz, M.; Lapen, D.; Moulton, K.; Quake, S. R.; Steinmann, K.; Thayer, E.; Tyurina, A.; Ward, R.; Weiss, H.; Xie, Z. Science 2008, 320 (5872), 106−9. (3) Eid, J.; Fehr, A.; Gray, J.; Luong, K.; Lyle, J.; Otto, G.; Peluso, P.; Rank, D.; Baybayan, P.; Bettman, B.; Bibillo, A.; Bjornson, K.; Chaudhuri, B.; Christians, F.; Cicero, R.; Clark, S.; Dalal, R.; Dewinter, A.; Dixon, J.; Foquet, M.; Gaertner, A.; Hardenbol, P.; Heiner, C.; Hester, K.; Holden, D.; Kearns, G.; Kong, X.; Kuse, R.; Lacroix, Y.; Lin, S.; Lundquist, P.; Ma, C.; Marks, P.; Maxham, M.; Murphy, D.; Park, I.; Pham, T.; Phillips, M.; Roy, J.; Sebra, R.; Shen, G.; Sorenson, J.; Tomaney, A.; Travers, K.; Trulson, M.; Vieceli, J.; Wegener, J.; Wu, D.; Yang, A.; Zaccarin, D.; Zhao, P.; Zhong, F.; Korlach, J.; Turner, S. Science 2009, 323 (5910), 133−8. (4) Holzmeister, P.; Acuna, G. P.; Grohmann, D.; Tinnefeld, P. Chem. Soc. Rev. 2014, 43 (4), 1014−28. (5) van Oijen, A. M. Nat. Chem. Biol. 2008, 4 (8), 440−3. (6) Wenger, J.; Rigneault, H. Int. J. Mol. Sci. 2010, 11 (1), 206−21. 3503

dx.doi.org/10.1021/nl501064b | Nano Lett. 2014, 14, 3499−3503