www.sciencemag.org/cgi/content/full/338/6106/506/DC1 Supplementary Materials for Fluorescence Enhancement at Docking Sites of DNA-Directed Self- Assembled Nanoantennas G. P. Acuna,* F. M. Möller, P. Holzmeister, S. Beater, B. Lalkens, P. Tinnefeld* *To whom correspondence should be addressed. E-mail: [email protected] (G.P.A.); [email protected](P.T.) Published 26 October 2012, Science 338, 506 (2012) DOI: 10.1126/science.1228638 This PDF file includes: Materials and Methods Figs. S1 to S9 Table S1 Full Reference List
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Supplementary Materials for...AHF) after the pinhole onto a second detector (τ-SPAD-100, Picoquant) with appropriate filters (Brightline HC582/75, AHF and RazorEdge LP 532, Semrock).
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Published 26 October 2012, Science 338, 506 (2012) DOI: 10.1126/science.1228638
This PDF file includes:
Materials and Methods Figs. S1 to S9 Table S1 Full Reference List
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Materials and Methods Numerical simulations:
Numerical simulations were performed using commercial FDFD software (www.cst.com). The relative change in the excitation rate was estimated by simulating the electric field intensity in the vicinity of the nanoparticle system at the fluorophore position and averaged over the tangential and radial incident polarization. The relative change of the radiative and non radiative decay as well as the relative change in the lifetime was estimated following ref. (30), where the dye is modeled by a dipole current source with a determined orientation. The total power radiated into the far field and dissipated by the metallic objects is computed and normalized to the total power radiated into the far field in the absence of metallic objects. For these calculations, the intrinsic quantum yield of the ATTO647N dye of 0.65 was considered according to (31). All simulations were performed assuming a planar illumination with water as medium, mimicking the buffer conditions and effects arising from the glass interface were neglected. The error bars take into account the size distribution of the nanoparticles, but the deviation of particles from the spherical shape, the angular distribution of the DNA origami pillar or the possibility that not all three capturing strands are binding to a nanoparticle are not considered.
Functionalization of gold nanoparticles:
Gold nanoparticles of five different diameters (20, 40, 60, 80 and 100 nm) were purchased from BBInternational (www.bbi-gold.com). TEM measurements revealed the following size distribution, (19.4 ± 1.7 nm, 43.5 ± 4.6 nm, 61.5 ± 7 nm, 80.1 ± 7.4 nm, 105 ± 10.4 nm). Additionally some particles showed a considerable deviation from a spherical shape, see Fig. S3.
The DNA functionalization was performed as described in ref (32) with the following DNA sequence from VBC Biotech (www.vbc-biotech.com) containing a thiol modification at the 5’ end: 5’-Thiol-TTTTTTTTTTTTTTT-3’. The binding of the gold nanoparticles to the DNA origami structure was included in ref (19). The immobilized DNA origami pillars were incubated with DNA modified gold nanoparticles until the desired binding yield was achieved. This incubation was usually carried out on the microscope for several hours. Determination of intensity and fluorescence lifetime:
Fluorescence intensity and lifetime measurements were carried out on a custom-built confocal setup based on an Olympus IX-71 inverted microscope. For excitation, the light of an 80 MHz pulsed diode laser (640nm, LDH-D-C-640, Picoquant) was coupled into an oil-immersion objective (UPlanSApo 60XO / 1.35 NA, Olympus). A linear polarizer, an electrooptical modulator (EOM, LM 0202, Qioptiq) and a quarter wave plate (AQWP05M-600, Thorlabs) in the correct orientation with respect to each other allowed control over the excitation polarization (33). Depending on the rotation speed, we either measured with rotating linear polarization or average over all directions which for our measurements is equivalent to measurements with circular polarized light. Excitation and emission light were separated by a dual-band dichroic beam splitter (z532/633, AHF) and subsequently focused onto a 50 µm pinhole (Linos) before detection with an avalanche
photo diode (τ-SPAD-100, Picoquant) with appropriate spectral filtering (ET 700/75m, AHF and RazorEdge LP 647, Semrock). The detector signal was registered with a single photon counting PC card (SPC-830, Becker&Hickl) and further analyzed using custom-made LabView code.
For FRET measurements, an additional 532 nm laser (TECGL-30, World Star Tech) was used to excite Cy3 and the emitted light was split spectrally at 640 nm (640DCXR, AHF) after the pinhole onto a second detector (τ-SPAD-100, Picoquant) with appropriate filters (Brightline HC582/75, AHF and RazorEdge LP 532, Semrock). The two lasers were alternated with 1 ms period by use of an acousto-optical tunable filter (AOTFnc-VIS, AA optoelectronic) to separate FRET sensitized from direct Cy5 excitation.
We employed the reconvolution algorithm of the FluoFit software (Picoquant) to obtain the fluorescence lifetime from the measured decay and the instrument response function (IRF) of the setup. We used an IRF acquired at an appropriate intensity to account for count rate dependence of the detector and included the periodicity of the excitation as well as scattering in the analysis. The width of the IRF (FWHM=650 ps) limits the temporal resolution to approximately 100-200 ps. All decays could be fitted with the convolution of a monoexponential decay and the IRF.
The Holliday junction:
The Holliday junction consisted of four strands (28) called R, H, X and B with the following sequences R: 5’- ACA AAT ATC CTT GCC CCA GCA GGC GAA TTT CCC ACC GCT CGG CTC AAC TGG G -3’, H: 5’-Cy3-CCG TAG CAG CGCG AGC GGT GGG-3’, X: 5’-CCC AGT TGA GCG CTT GCT AGG G-3’ and B: 5’-Cy5-CCC TAG CAA GCC GCT GCT AGG G-3’. The first 27 base pairs of the strand R bind to the origami pillar. All four strands were incorporated together with the staples prior to the folding process.
DNA origami pillar structures:
Unmodified and modified staple strands (see Table S1) were purchased from MWG (Munich, Germany) or IBA (Göttingen, Germany) at a concentration of 100 μM and were used without further purification. DNA origamis are formed with a molar ratio of 1:30 between the viral DNA and the unmodified staple strands and 1:100 between the viral DNA and the modified staple strands. For preparation of the scaffold strands Escherichia coli strain K91 was infected with the respective M13mp18 phage (p8634) at a Multiplicity of Infection of ~1. After amplification, the phage particles were separated, purified and their ssDNA was extracted and purified similar as described before (17). The concentration was adjusted to 100 nM using a molecular weight of 330 g/mol per base and an extinction coefficient = 33 mg/ml for A260 = 1 in a NanoDrop Spectrophotometer (Peqlab, Erlangen, Germany). The DNA origami design was performed with the open-source software caDNAno (www.cadnano.org)(17). The folding buffer contained 12.5 mM MgCl2 as well as 5 mM Tris + 1 mM EDTA (pH 7.9 at 20°C). A TEM image of the DNA origami pillar is included in Fig. S8. Folding time was three days.
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Fig. S1. Sketch of the top-view of the DNA origami pillar. The numbered circles represent DNA helices. Helices 0 to 11 form the central 12-helix bundle. The remaining helixes form the extra 6-helix bundles of the base. The helix center-to-center distance is 3 nm (16). The positions of the single dye (ATTO647N), the capturing paint strands, the Holliday Junction and the nanoparticles (NP) are also included.
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Fig. S2. Fluorescence lifetime imaging microscopy of DNA origami pillars with 80 nm nanoparticles. Orange-red spots represent ATTO647N dyes on DNA origami pillars without nanoparticles. The shortened fluorescence lifetime (blue spots) indicates binding of nanoparticles. Commonly, intermediate binding yields were intended to use the fluorescence from DNA origami pillars without nanoparticles as internal reference (A). On the other hand, yields exceeding 70% for the dimer could be reached even for 80 nm particles (B). (C) Histogram of yields of dimers, monomers and DNA origamis without a nanoparticle for the measurement included in Fig. 3A.
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Fig. S3. (A)-(E) TEM images of the gold nanoparticles of five different sizes. Analysis of the images revealed some deviation from perfect spherical shapes as well as a distribution of sizes (19.4 ± 1.7 nm, 43.5 ± 4.6 nm, 61.5 ± 7 nm, 80.1 ± 7.4 nm, 105 ± 10.4 nm).
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Fig. S4. (A)-(E) Fluorescence intensity versus lifetime plot of the DNA origami pillar with binding sites for one (monomer) and two (dimer) particles of different diameters.
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Fig. S5. Simulated quantum yield (A) and fluorescence lifetime (B) as a function of the nanoparticle diameter for the monomer and dimer system at different dye orientations, radial and tangential. The error bars take into account the size distribution of the nanoparticles. The quantum yield and fluorescence lifetime is normalized to the properties of the dye in the absence of nanoparticles. A quantum yield of 0.65 for ATTO647N was considered for both the quantum yield and fluorescence lifetime calculations.
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Fig. S6. Time gating for signal to noise improvement at elevated background fluorescence from freely diffusing dye molecules. (A) Fluorescence decay of the ATTO647N dye located within the DNA origami pillar with two 80 nm nanoparticles (dimer). Measurements were carried out at a concentration of 0.5 µM of ATTO647N dyes in solution. The decay shows a bi-exponential behavior due to the combination of the quenched dye in the hotspot and the unquenched dyes of the background. (B) Corresponding intensity transient showing that the single ATTO647N in the hotspot is easily detected despite the high background concentration. For (C) and (D) the intensity transients were reconstructed by time gating allowing only photons arriving 1.5-6 ns and 1.5-3 ns in the fluorescence decay, respectively. The time gating further increased the signal-to-noise by removing the largest fraction of the background fluorescence.
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Fig. S7. High count rates for fast dynamics. (A) Intensity transient at 1 µs binning. Off-states as short as 10 µs are clearly visualized. Therefore, the excitation intensity was increased to 7 µW and 100 µM ascorbic acid and 100 µM of methylviologen were added to the buffer to induce fast blinking of the dye molecule (34). At these high count rates, saturation effects of the setup and not by the organic dye limit photon counts. To estimate the actual rate of photons arriving at the detector in the on-state, we corrected for the dead time of the setup (B,C). Therefore, we measured the detected count rate of a micromolar concentration of ATTO647N in solution for different excitation intensities in an excitation regime where fluorescence from ATTO647N is linearly depending on excitation intensity (B). Photon count rate vs excitation intensity for a µM concentration of ATTO647N. (C) Correction factor dependence on the measured count rate. Hyperbolic fitting yields a saturation at Nmax = 6.8 ± 0.2 MHz which corresponds to a dead time of τD = 1/Nmax = 147 ± 4 ns. This yields a correction factor of f=1/(1-N τD) as displayed in (C). Accordingly, the count rate of the on-state for the transient displayed in (A) is 10.6 MHz. It should be emphasized that the blinking of the dye was purposely induced for this measurement and that all measurements discussed before were taken at lower excitation intensities to avoid saturation effects.
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Fig. S8. Transmission electron microscopy image of the DNA origami pillar.
Table S1. List of unmodified and modified oligonucleotides from the 5’ to the 3’ used for the different DNA pillar structures sketched in Fig. 1A, Fig. 4A and E. The nomenclature follows the numbering used in Fig. S9.
References and Notes 1. C. Joo, H. Balci, Y. Ishitsuka, C. Buranachai, T. Ha, Advances in single-molecule
2. P. Tinnefeld, M. Sauer, Branching out of single-molecule fluorescence spectroscopy: Challenges for chemistry and influence on biology. Angew. Chem. Int. Ed. 44, 2642 (2005). doi:10.1002/anie.200300647
3. J. Eid et al., Real-time DNA sequencing from single polymerase molecules. Science 323, 133 (2009). doi:10.1126/science.1162986 Medline
4. B. Huang, H. Babcock, X. Zhuang, Breaking the diffraction barrier: super-resolution imaging of cells. Cell 143, 1047 (2010). doi:10.1016/j.cell.2010.12.002 Medline
5. M. J. Levene et al., Zero-mode waveguides for single-molecule analysis at high concentrations. Science 299, 682 (2003). doi:10.1126/science.1079700 Medline
6. S. Uemura et al., Real-time tRNA transit on single translating ribosomes at codon resolution. Nature 464, 1012 (2010). doi:10.1038/nature08925 Medline
7. L. Novotny, N. van Hulst, Antennas for light. Nat. Photonics 5, 83 (2011). doi:10.1038/nphoton.2010.237
8. J. A. Schuller et al., Plasmonics for extreme light concentration and manipulation. Nat. Mater. 9, 193 (2010). doi:10.1038/nmat2630 Medline
9. T. H. Taminiau, F. D. Stefani, F. B. Segerink, N. F. van Hulst, Optical antennas direct single-molecule emission. Nat. Photonics 2, 234 (2008). doi:10.1038/nphoton.2008.32
10. A. G. Curto et al., Unidirectional emission of a quantum dot coupled to a nanoantenna. Science 329, 930 (2010). doi:10.1126/science.1191922 Medline
11. A. Kinkhabwala et al., Large single-molecule fluorescence enhancements produced by a bowtie nanoantenna. Nat. Photonics 3, 654 (2009). doi:10.1038/nphoton.2009.187
12. M. Ringler et al., Shaping emission spectra of fluorescent molecules with single plasmonic nanoresonators. Phys. Rev. Lett. 100, 203002 (2008) and references therein. doi:10.1103/PhysRevLett.100.203002 Medline
13. H. Lin et al., Mapping of surface-enhanced fluorescence on metal nanoparticles using super-resolution photoactivation localization microscopy. ChemPhysChem 13, 973 (2012) and references therein. doi:10.1002/cphc.201100743 Medline
14. H. Cang et al., Probing the electromagnetic field of a 15-nanometre hotspot by single molecule imaging. Nature 469, 385 (2011) and references therein. doi:10.1038/nature09698 Medline
15. M. P. Busson, B. Rolly, B. Stout, N. Bonod, S. Bidault, Accelerated single photon emission from dye molecule-driven nanoantennas assembled on DNA. Nat. Commun. 3, 962 (2012) and references therein. doi:10.1038/ncomms1964 Medline
16. P. W. Rothemund, Folding DNA to create nanoscale shapes and patterns. Nature 440, 297 (2006). doi:10.1038/nature04586 Medline
17. S. M. Douglas et al., Self-assembly of DNA into nanoscale three-dimensional shapes. Nature 459, 414 (2009). doi:10.1038/nature08016 Medline
18. See the supplementary materials on Science Online.
19. G. P. Acuna et al., Distance dependence of single-fluorophore quenching by gold nanoparticles studied on DNA origami. ACS Nano 6, 3189 (2012). doi:10.1021/nn2050483 Medline
20. E. A. Coronado, E. R. Encina, F. D. Stefani, Optical properties of metallic nanoparticles: Manipulating light, heat and forces at the nanoscale. Nanoscale 3, 4042 (2011). doi:10.1039/c1nr10788g Medline
21. P. Anger, P. Bharadwaj, L. Novotny, Enhancement and quenching of single-molecule fluorescence. Phys. Rev. Lett. 96, 113002 (2006). doi:10.1103/PhysRevLett.96.113002 Medline
22. A. Bek et al., Fluorescence enhancement in hot spots of AFM-designed gold nanoparticle sandwiches. Nano Lett. 8, 485 (2008). doi:10.1021/nl072602n Medline
23. S. Kühn, U. Håkanson, L. Rogobete, V. Sandoghdar, Enhancement of single-molecule fluorescence using a gold nanoparticle as an optical nanoantenna. Phys. Rev. Lett. 97, 017402 (2006). doi:10.1103/PhysRevLett.97.017402 Medline
24. J. Vogelsang et al., A reducing and oxidizing system minimizes photobleaching and blinking of fluorescent dyes. Angew. Chem. Int. Ed. 47, 5465 (2008). doi:10.1002/anie.200801518
25. N. Di Fiori, A. Meller, The effect of dye-dye interactions on the spatial resolution of single-molecule FRET measurements in nucleic acids. Biophys. J. 98, 2265 (2010). doi:10.1016/j.bpj.2010.02.008 Medline
26. R. Jungmann et al., Single-molecule kinetics and super-resolution microscopy by fluorescence imaging of transient binding on DNA origami. Nano Lett. 10, 4756 (2010). doi:10.1021/nl103427w Medline
27. S. A. McKinney, A.-C. Déclais, D. M. J. Lilley, T. Ha, Structural dynamics of individual Holliday junctions. Nat. Struct. Biol. 10, 93 (2003). doi:10.1038/nsb883 Medline
28. A. Gietl, P. Holzmeister, D. Grohmann, P. Tinnefeld, DNA origami as biocompatible surface to match single-molecule and ensemble experiments. Nucleic Acids Res. 40, e110 (2012). doi:10.1093/nar/gks326 Medline
29. H. S. Chung, K. McHale, J. M. Louis, W. A. Eaton, Single-molecule fluorescence experiments determine protein folding transition path times. Science 335, 981 (2012). doi:10.1126/science.1215768 Medline
30. T. H. Taminau, F. D. Stefani, N. F. Van Hulst, Single emitters coupled to plasmonic nano-antennas: Angular emission and collection efficiency. New J. Phys. 10, 105005 (2008).
31. P. Bharadwaj, L. Novotny, Spectral dependence of single molecule fluorescence enhancement. Opt. Express 15, 14266 (2007). doi:10.1364/OE.15.014266 Medline
32. C. A. Mirkin, R. L. Letsinger, R. C. Mucic, J. J. Storhoff, A DNA-based method for rationally assembling nanoparticles into macroscopic materials. Nature 382, 607 (1996). doi:10.1038/382607a0 Medline
33. K. D. Weston, L. S. Goldner, Orientation imaging and reorientation dynamics of single dye molecules. J. Phys. Chem. B 105, 3453 (2001). doi:10.1021/jp001373p
34. J. Vogelsang, T. Cordes, C. Forthmann, C. Steinhauer, P. Tinnefeld, Controlling the fluorescence of ordinary oxazine dyes for single-molecule switching and superresolution microscopy. Proc. Natl. Acad. Sci. U.S.A. 106, 8107 (2009). doi:10.1073/pnas.0811875106 Medline