Fluorescence Quenching of Quantum Dots by Gold ... - ACS Publications

Aug 1, 2014 - Department of Chemistry, University of Central Florida, 4104 Libra Drive, Orlando, Florida 32816-2366, United States. •S Supporting ...
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Fluorescence Quenching of Quantum Dots by Gold Nanoparticles: A Potential Long Range Spectroscopic Ruler Anirban Samanta,† Yadong Zhou,‡ Shengli Zou,‡ Hao Yan,† and Yan Liu*,† †

Department of Chemistry and Biochemistry, and Center for Molecular Design and Biomimicry, Biodesign Institute at Arizona State University, 1001 South McAllister Avenue, Tempe, Arizona 85287-5601, United States ‡ Department of Chemistry, University of Central Florida, 4104 Libra Drive, Orlando, Florida 32816-2366, United States S Supporting Information *

ABSTRACT: The dependence of quantum dot (QD) fluorescence emission on the proximity of 30 nm gold nanoparticles (AuNPs) was studied with controlled interparticle distances ranging from 15 to 70 nm. This was achieved by coassembling DNA-conjugated QDs and AuNPs in a 1:1 ratio at precise positions on a triangular-shaped DNA origami platform. A profound, long-range quenching of the photoluminescence intensity of the QDs was observed. A combination of static and timeresolved fluorescence measurements suggests that the quenching is due to an increase in the nonradiative decay rate of QD emission. Unlike FRET, the energy transfer is inversely proportional to the 2.7th power of the distance between nanoparticles with half quenching at ∼28 nm. This long-range quenching phenomena may be useful for developing extended spectroscopic rulers in the future. KEYWORDS: Quantum dots, gold nanoparticles, FRET, NSET, DNA, self-assembly

F

useful for measuring intermolecular distances in the small window between 1 and 10 nm. This range is sufficient for observing dynamic interactions between proteins, nucleic acids, and cell membranes.1−6 However, many other biomolecular processes occur over longer distances and their dynamics and interactions are difficult to follow using FRET. This has motivated researchers to develop longer range “spectroscopic rulers”. In the past decade, nanometal surface energy transfer (NSET) has emerged as a spectroscopic ruler technique that can measure intermolecular distances up to 50 nm, depending on the size of the metallic nanoparticle involved.7−10 NSET is based on the phenomenon in which the lifetime of an oscillating dipole is damped when it is located at a certain distance from a metal surface. Experiments have shown that the proximity of a metal nanoparticle to an organic fluorophore quenches its fluorescence at longer ranges. Although some reports have suggested that the efficiency of NSET energy transfer is inversely proportional to the fourth power of the distance between particles, the transfer efficiency is also dependent on the size of the particles.9,11 For FRET, electrodynamic coupling exhibits short-range distance dependence because it involves the interaction between two point dipoles. For NSET, the large size of the metal nanoparticles and

örster resonance energy transfer (FRET) has become a well-utilized tool to measure the distance between molecules because FRET efficiency is sensitive to small changes in distance. FRET is an electrodynamic phenomenon where two nearby oscillating dipoles interact with one another and energy is transferred from the excited state of a donor molecule to the ground state of an acceptor molecule in a nonradiative fashion. The rate of FRET is inversely proportional to the sixth power of the distance between the dipoles. The efficiency of energy transfer is described by 1

E FRET = 1+

6

( ) r R0

(1)

where r1 is the distance between the interacting dipoles, and R0 is the Förster radius that is defined as the distance at which FRET efficiency equals 50%. R0 depends on various factors including the spectral overlap of the donor emission spectrum and the acceptor absorption spectrum, the quantum yield of the donor in the absence of the acceptor, and the relative orientation of the donor emission and acceptor absorption dipole moments. For commonly used organic fluorophore pairs, R0 is generally in the range of 2−6 nm. FRET energy transfer efficiencies are extremely sensitive to changes in distance when operating close to the Förster radius. For example, at r = 0.8R0 and r = 1.2R0, the energy transfer efficiencies are ∼80% and ∼25%, respectively. In fact, it is impractical to examine intermolecular distances that fall outside the range of r = 0.5R0 to 2R0 using FRET. Thus, FRET is only © 2014 American Chemical Society

Received: May 7, 2014 Revised: July 25, 2014 Published: August 1, 2014 5052

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higher polarizability relative to the smaller fluorescent molecule produces longer range effects.12 The underlying mechanism of NSET is still under debate.13−20 In the seminal work by CPS-Kuhn, the quenching effect was supposed to be due to absorption by the metal nanoparticle upon the formation of an image dipole on the metal nanoparticle surface.14 Although this theory explains experimental results corresponding to fluorescence quenching by a 2 nm AuNP, it fails to predict the quenching behavior when the size of the nanoparticle changes.8 Other theories that claim to predict and explain the experimental NSET results of different sized AuNPs have emerged, including the GersteinNitzan model,19 but an adequate depiction of the real picture has yet to be unveiled. Recently, Breshike et el. attempted to improve the model by introducing empirical corrections to adjust the absorptivity and dielectric constant terms as the size of the AuNP changed.8 Their theoretical predictions match fairly well with experimental results from different-sized AuNPs. To date, NSET experiments have focused on the interaction of AuNPs with organic dyes. Meanwhile, the interaction of metallic nanoparticles with photoluminescent semiconductor nanoparticles, another type of quantum emitter also known as quantum dots, has not been extensively explored.21−23 QDs offer some distinctive advantages over organic dyes. Their broad absorption and narrow emission spectra, high quantum yield and excellent chemical- and photostability have facilitated their emergence as reliable labeling and imaging agents. Although there are numerous reports of QDs serving as universal donors for FRET-like energy transfer to organic dyes,24−27 systematic studies of distance-dependent quenching of QD emission by plasmonic nanoparticles is very scarce. Pons et al. investigated the nature of distance dependent quenching of semiconductor QDs by 1.4 nm (diameter) AuNPs and observed NSET type quenching behavior.21 However, AuNPs smaller than 2 nm do not have well-defined surfaces and consequently do not exhibit a clear surface plasmon band. This indicates that the mechanism of quenching might be different for larger AuNPs. Here, we were motivated to investigate the nature of distance-dependent quenching by larger AuNPs that display prominent surface plasmon bands with the hope of developing an optical ruler that can be used for measuring longer range intermolecular distances. We used the bottom up approach of DNA nanotechnology to assemble AuNP-QD heterodimers and subsequently studied energy transfer between the particles. In recent years, DNA nanostructures have emerged as reliable scaffolds for spatially organizing nanoparticles and biomolecules with nanometer precision.28−32 Among all the reported DNA nanostructures, DNA origami is particularly well suited for fine-tuning the distance between nanoparticles. DNA functionalized AuNPs and AgNPs of various shapes and sizes have successfully been organized into unique patterns by DNA origami.33−40 In addition, DNA origami has served as a scaffold for the organization of DNA conjugated- and streptavidin coatedQDs.31,41 Moreover, the addressability of DNA origami makes it an ideal platform to coassemble multifunctional components.42−46 Recently, several groups have demonstrated the attachment of discrete numbers of AuNPs and QDs on a DNA origami platform.47,48 However, the streptavidin-coated QDs used in these reports are very large due to the cross-linked polymers and streptavidin protein coating on their surface. As a

result, they perform poorly in distance-dependent energy transfer studies compared to DNA functionalized QDs.49 We recently developed a method for synthesizing stable DNA-conjugated water-soluble core/shell quantum dots with varying elemental compositions and site-specifically displayed them from DNA origami.31 Here, we displayed one 30 nm AuNP and one red light emitting QD (CdTe/CdS core/shell) from a triangular DNA origami structure at five different controlled interparticle distances, ranging from 15 to 70 nm. Static fluorescence spectroscopy and dynamic lifetime measurements were employed to study the distance dependent quenching of QD fluorescence. The triangle origami structure (∼115 nm outer edges) was formed by mixing approximately 200 unique, short staple strands with a long single-stranded DNA scaffold (from M13mp18).50 Binding sites for DNA-conjugated AuNPs and QDs were created at specific positions by extending selected staple strands with a DNA sequence complementary to the binding domains of the DNA displayed from the AuNPs and QDs. To reduce the translational freedom of the bound NPs, each binding site was designed as a cluster with three capture strands located ∼6 nm apart, at the vertices of a roughly equilateral triangle. The resulting nanostructures were purified by washing with 1× TAE-Mg2+ buffer and filtration using 100 KD molecular weight cut off (MWCO) Amicon filters to remove excess staple and capture strands. Atomic force microscopy (AFM) confirmed the yield and integrity of the purified origami structure. DNA-conjugated CdTe/CdS core/shell QDs with emission maxima at ∼645 nm were synthesized and functioned as quantum emitters. Commercially available citrate stabilized spherical AuNPs (30 ± 1 nm diameter) served as the quencher. Thirty nanometer AuNPs were utilized because of their large extinction cross-section, yet relatively low scattering cross section. Besides suffering from significant scattering as particle size increases, the extinction spectra of AuNPs larger than 30 nm are dominated by quadrupole and octapole resonances, which make the system more complicated to explain theoretically. Five different constructs were assembled; the position of the AuNP was fixed while the QD was systematically moved farther from the AuNP. The assembly of the constructs is performed in several steps (Figure 1). First, the DNA functionalized AuNPs are incubated overnight at room temperature with DNA origami. The Mg2+ ion concentration is reduced from a standard level of 12.5 to 6.25 mM due to the intrinsic instability of the large AuNPs in high salt conditions. This change in the ionic strength does not significantly reduce the stability of the 2D DNA origami. In order to encourage maximum assembly yield, the preassembled origami are mixed with an excess of AuNPs. Thus, the AuNP decorated DNA origami require subsequent isolation from the excess, unbound AuNPs and any other higher order structures that form during assembly, by agarose gel electrophoresis (Supporting Information Figure S8). A high yield of the desired structure is confirmed by transmission electron microscopy (TEM) imaging before proceeding to the next step (Supporting Information Figure S1). Next, the DNA functionalized QDs are incubated with the purified AuNP-origami structures. Finally, the excess, unbound QDs are removed by gentle centrifugation (g = 4000 for 5 min). During centrifugation, the high molecular weight AuNPorigami-QD structures are deposited as a pellet in the bottom of the test tube. The supernatant solution containing the 5053

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luminescent intensity of each sample was obtained from static fluorescence measurements and normalized by comparison to the corresponding control sample. The most appropriate control sample for these experiments is one in which the QDs, AuNPs, and DNA are not bound together, but are present in the same concentration. We followed a rather simple but elegant strategy employed by Pal et al. to prepare the control sample.51 Origami structures with bound QDs and AuNPs were heated above their melting temperature (65 °C for 5 min) and quickly immersed into an ice bath. Thus, the origami structures are melted and all the component DNA strands are dissociated such that the particles are released into solution. Quick immersion into ice ensures that the melted origami does not refold and the nanoparticles are not coassembled. Considering the subnanomolar concentration of the particles, we assume that they are well dispersed following treatment and do not interact with one another with an average intermolecular distance on the micrometer scale. Having the same concentration of AuNPs in the control sample also helps to correct the extinction coefficient of the AuNPs for fluorescence intensity measurements at both excitation and emission wavelengths. The quenching efficiency, defined as

Figure 1. Schematic depicting the stepwise assembly of 30 nm gold nanoparticles and CdTe/CdS Core/shell QDs (red) on DNA origami (gray). The gold NP and QD capture strands displayed from the DNA origami triangle are displayed in red and orange, respectively. (1) The scaffold (M13 viral DNA) and staple strands (including the capture strands, three per particle) are mixed together in 1:5:10 molar ratio in 1× TAE-Mg buffer and annealed (heated to 90 °C and cooled to 4 °C over 12 h). The excess staple strands are removed with an Amicon centrifugal filter device (MWCO 100 kDa). (2) DNA functionalized gold nanoparticles are mixed with the DNA origami (1:5 ratio) and incubated overnight to ensure capture of the nanoparticles. Purification is performed by native agarose gel electrophoresis to remove the free gold NPs and higher order structures. (3) DNA functionalized QDs (emission peak at 645 nm) are mixed with the purified DNA origami-gold NP conjugates, incubated to ensure QD capture, and subsequently purified by gentle centrifugation to remove the free QDs.

ET = 1 −

PLsample PLcontrol

(2)

was plotted against the AuNP-surface to QD-center interparticle distance (Figure 2B). It is clear from Figure 2 that the 30 nm AuNP reduces the fluorescent intensity of the proximal QD with the level of quenching strongly dependent on the interparticle distance. Theoretical calculations at 520 and 645 nm (red and purple traces, respectively) emission wavelengths, based on an electrodynamic model (more details in the Supporting Information), are plotted together with the experimental data (black circles). The theoretical calculations involve extension of a dipole−dipole interaction model by including higher order excitation, since the size of the particles are larger than previously reported.39 We found that the quenching rate calculated at 645 nm is not well represented by the experimental data. However, the predicted quenching rate at 520 nm is in agreement with the experimental measurements. The green trace represents a data fit based on the equation below, where D0 and n are the fitting parameters,

unbound QDs is removed and discarded. The pellet is then resuspended in 0.5× TAE-Mg2+ buffer by gentle agitation. This process does not cause any notable damage to the final structure, although some loss of the product is inevitable. Assuming that all three capture strands in a cluster hybridize to three DNA strands displayed from the same nanoparticle, the center of the particle should be located in the center of the triangle formed by the three captures strands, with a vertical distance from the 2D origami plane equal to the radius of the particle plus the height of the dsDNA projected in the vertical direction. The length of the DNA displayed from the QDs and the AuNPs are equal, thus, the center-to-center distance between the two particles in the vertical direction (perpendicular to the origami plane) is the difference in their radii. The horizontal center-to-center distance between the two particles (along the origami plane) is calculated from the geometry of the designed DNA origami and the locations of the capture strands. On the basis of these assumptions, the center-tosurface interparticle distances were calculated using simple geometry (Supporting Information Figure S9). The main source of error in this calculation is the uncertainty in the size of the particles and the rigidity of the DNA origami structure. The calculated distances were compared to experimental distances measured from TEM images of the samples. Since TEM images are 2D projections of 3D objects, we introduced a correction that took the height difference of the two nanoparticles into consideration (Supporting Information Figure S9). The experimentally determined distances between the AuNP surface and the center of the QDs in the five different constructs were 14.7 ± 3.3, 29.6 ± 2.7, 39.0 ± 3.3, 55.0 ± 3.1, and 69.8 ± 3.6 nm. All five purified DNA origami structures with one QD and one AuNP at precisely controlled interparticle distances were subjected to static fluorescence and dynamic lifetime measurements (Supporting Information Figures S10−12). The photo-

1

Y=1− 1+

n

( ) d D0

(3)

Analysis of the fit yields n = 2.72 ± 0.08 and D0 = 27.8 ± 0.4 nm. For the infinite surface model pioneered by Person, the value of n is expected to be 4. We also fit the calculated data using the previously mentioned electrodynamic model to the same equation, which yields n = 3.05 ± 0.03 and D0 = 22.9 ± 0.1 nm for 520 nm emission, and n = 3.16 ± 0.06 and D0 = 12.7 ± 0.1 nm for 645 nm emission. We speculate that there are two reasons for the discrepancy between theory and experiment. The first is likely because of a false assumption that the supposed 2D DNA origami triangle structure is rigid and therefore flat. Thus, the expected interparticle distance that we calculated may not be accurate. In particular, the presence of a heavy 30 nm AuNP (also functionalized with a large number of DNA molecules, thus highly charged) on the origami may cause slight bending in the 5054

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Figure 2. (A) Absorption spectra of 30 nm AuNPs and PL emission spectra of QDs with emission maxima at 645 nm. (B) Quenching efficiency (defined as 1- (PL sample/PL control)) is plotted against the interparticle distance. The green trace corresponds to predicted values. (C) Life time decay of each construct shown in D. (D) Design of the five constructs and the corresponding TEM images. Scale bar 100 nm. The distance between AuNP surface to the center of the QDs in the five different constructs are (i) 69.8 ± 3.6, (ii) 55.0 ± 3.1, (iii) 39.0 ± 3.3, (iv) 29.6 ± 2.7, and (v) 14.7 ± 3.3 nm.

primarily caused by an increase in the rate of nonradiative decay, we expect to observe an interparticle distance dependent decrease in the lifetime. The data was analyzed by a custom software package, ASUFIT, and was fit by a sum of multiexponential decay model, (URL http://www.public.asu. edu/∼laserweb/asufit/asufit.html)

structure, reducing the actual distance between the particles. For example, for the construct shown in Figure 2Di, a 10° deviation from the 2D plane in any of the arms will cause the particles to be ∼5−6 nm closer together than expected. Another potential cause for the discrepancy between theory and experiment is related to the properties of the QDs themselves. Unlike small organic dyes, the fluorescent properties of QDs are strongly dependent on their surface properties, which may affect the corresponding fluorescence decay mechanisms in unexpected ways. Also, because the state density of a QD is much higher than that of a dye molecule, an excited QD can experience a different decay mechanism than a dye molecule with a very limited number of states. These effects will need further investigation in the future. We also examined the photoluminescence lifetimes of the assembled QDs using the time-correlated single photon counting (TCSPC) method. Assuming the quenched PL is

F (t ) =

∑ Aie−t/τ τ

(4)

where F(t) is the obtained kinetic decay curve, Ai is the amplitude of the ith decay channel and τi is the corresponding lifetime. Three exponentials were required to fit the decay data, thus, the average lifetime was calculated using 3

⟨τ ⟩ = 5055

∑i = 1 Ai τi 3

∑i = 1 Ai

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Table 1. Average Lifetime ⟨τ⟩ of QD Emission in the Five Constructs Shown in Figure 2C free QDs

construct (i)

construct (ii)

construct (iii)

construct (iv)

construct (v)

23.9

69.8 ± 3.6 23.3 23.6

55.0 ± 3.1 19.8 23.1

39.0 ± 3.3 17.1 21.3

29.6 ± 2.7 14.1 18.6

14.7 ± 3.3 8.1 4.8

interparticle distance (nm) experimental (ns) theoretical (ns)

The experimentally obtained average lifetime and the theoretically predicted values are listed in Table 1. The steady state fluorescence measurements and the average lifetime were used to calculate the average radiative and nonradiative decay rates. The quantum efficiency and the average lifetime of a fluorophore are expressed as

QE =

kr k r + k nr

(6)

and τav =

1 k r + k nr

(7)

respectively. In the presence of a metallic nanoparticle, additional pathways are introduced in the relaxation mechanism of the excited states. The additional radiative and nonradiative rate constants corresponding to these pathways are krm and knrm, respectively, such that the effective rate constants of the radiative and nonradiative pathways become (kr + krm) and (knr + knrm), respectively. Thus, the modified expressions for QE and average lifetime are QE m =

k r + k rm k r + k rm + k nr + k nrm

Figure 3. Average radiative and nonradiative decay rates of QD photoluminescence from each of the five constructs shown in Figure 2D.

relatively heavy nanoparticles or by unexpected electrostatic interactions and/or decay mechanisms of QDs relative to small dye molecules. Additional studies with more rigid scaffold structures will provide insight into the discrepancies. In the future, the long-range quenching effects present within this biocompatible framework can be engineered to serve as a kind of molecular ruler in vitro and in vivo.

(8)

and τav =

k r + k rm

1 + k nr + k nrm

■

(9)

respectively. Steady state fluorescence measurements are used to calculate the modified quantum yields of individual constructs, considering the quantum yield of the free QDs is 45%. The results show that the nonradiative decay rate constant increased by more than 10 folds as the distance between the particles decreased (Figure 3). Meanwhile, the radiative decay rate constant remained the same, except for when the QD and AuNP were ∼15 nm apart and a slight drop in the radiative decay rate constant was observed. In summary, we developed a reliable method for coassembling metallic and semiconducting nanoparticles using DNA directed self-assembly. This method provided unprecedented control over the relative stoichiometry and interparticle distance and allowed us to perform bulk measurements and enhance our understanding of the plasmonic interactions between a gold nanoparticle and a photoluminescent semiconductor nanoparticle. We observed long-range quenching of the photoluminescence of a QD driven by the proximity of a large AuNP displayed from the same DNA origami structure. The quenching effect that we observed exceeds traditional dye−dye or QD−dye FRET, or NSET quenching of an organic dye by an AuNP. We determined that quenching of QD fluorescence is primarily due to increased nonradiative decay pathways that are induced by the presence of proximal AuNPs. We also found that the experimentally measured quenching efficiencies were always higher than the predicted values. This is probably due to bending of the 2D DNA scaffold by the

ASSOCIATED CONTENT

S Supporting Information *

Materials and methods, details of theoretical calculations, supplementary figures, and DNA sequences. This material is available free of charge via the Internet at http://pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS We thank financial support from ONR, NSF, ARO and Arizona State University. We also thank Dr. Jeanette Nangreave for proofreading the manuscript.

■

REFERENCES

(1) Stryer, L. Annu. Rev. Biochem. 1978, 47, 819−846. (2) Sekar, R. B.; Periasamy, A. J. Cell Biol. 2003, 160, 629−633. (3) Dosremedios, C. G.; Moens, P. D. J. J. Struct. Biol. 1995, 115, 175−185. (4) Lilley, D. M. J.; Wilson, T. J. Curr. Opin. Chem. Biol. 2000, 4, 507−517. (5) Selvin, P. R. Nat. Struct. Mol. Biol. 2000, 7, 730−734. (6) Parkhurst, L. J.; Parkhurst, K. M.; Powell, R.; Wu, J.; Williams, S. Biopolymers 2001, 61, 180−200. 5056

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Letter

(42) Fu, J.; Liu, M.; Liu, Y.; Woodbury, N. W.; Yan, H. J. Am. Chem. Soc. 2012, 134, 5516−5519. (43) Torring, T.; Voigt, N. V.; Nangreave, J.; Yan, H.; Gothelf, K. V. Chem. Soc. Rev. 2011, 40, 5636−5646. (44) Pilo-Pais, M.; Watson, A.; Demers, S.; Labean, T. H.; Finkelstein, G. Nano Lett. 2014, 14, 2099−2104. (45) Acuna, G. P.; M?ller, F. M.; Holzmeister, P.; Beater, S.; Lalkens, B.; Tinnefeld, P. Science 2012, 338, 506−510. (46) Lin, C.; Jungmann, R.; Leifer, A. M.; Li, C.; Levner, D.; Church, G. M.; Shih, W. M.; Yin, P. Nat. Chem. 2012, 4, 832−839. (47) Ko, S. H.; Du, K.; Liddle, J. A. Angew. Chem., Int. Ed. 2013, 52, 1193−1197. (48) Wang, R.; Nuckolls, C.; Wind, S. J. Angew. Chem., Int. Ed. 2012, 51, 11325−11327. (49) Boeneman, K.; Deschamps, J. R.; Buckhout-White, S.; Prasuhn, D. E.; Blanco-Canosa, J. B.; Dawson, P. E.; Stewart, M. H.; Susumu, K.; Goldman, E. R.; Ancona, M.; Medintz, I. L. ACS Nano 2010, 4, 7253−7266. (50) Rothemund, P. W. K. Nature 2006, 297−302. (51) Pal, S.; Dutta, P.; Wang, H.; Deng, Z.; Zou, S.; Yan, H.; Liu, Y. J. Phys. Chem. C 2013, 111, 12735−12744.

(7) Yun, C. S.; Javier, A.; Jennings, T.; Fisher, M.; Hira, S.; Peterson, S.; Hopkins, B.; Reich, N. O.; Strouse, G. F. J. Am. Chem. Soc. 2005, 127, 3115−3119. (8) Breshike, C. J.; Riskowski, R. A.; Strouse, G. F. J. Phys. Chem. C 2013, 117, 23942−23949. (9) Jennings, T. L.; Singh, M. P.; Strouse, G. F. J. Am. Chem. Soc. 2006, 128, 5462−5467. (10) Chowdhury, S.; Wu, Z.; Jaquins-Gerstl, A.; Liu, S.; Dembska, A.; Armitage, B. A.; Jin, R.; Peteanu, L. A. J. Phys. Chem. C 2011, 115, 20105−20112. (11) Griffin, J.; Singh, A. K.; Senapati, D.; Rhodes, P.; Mitchell, K.; Robinson, B.; Yu, E.; Ray, P. C. Chem.Eur. J. 2009, 15, 342−351. (12) Chance, R. R.; Prock, A.; Silbey, R. J. Chem. Phys. 1975, 62, 2245−2253. (13) Persson, B. N. J.; Lang, N. D. Phys. Rev. B 1982, 26, 5409−5415. (14) Kuhn, H. J. Chem. Phys. 1970, 53, 101−108. (15) Singh, M. P.; Strouse, G. F. J. Am. Chem. Soc. 2010, 132, 9383− 9391. (16) Pustovit, V. N.; Shahbazyan, T. V. J. Chem. Phys. 2012, 136, 204701. (17) Carminati, R.; Greffet, J. J.; Henkel, C.; Vigoureux, J. M. Opt. Commun. 2006, 261, 368−375. (18) Ruppin, R. J. Chem. Phys. 1982, 76, 1681−1684. (19) Gersten, J.; Nitzan, A. J. Chem. Phys. 1981, 75, 1139−1152. (20) Saini, S.; Srinivas, G.; Bagchi, B. J. Phys. Chem. B 2009, 113, 1817−1832. (21) Pons, T.; Medintz, I. L.; Sapsford, K. E.; Higashiya, S.; Grimes, A. F.; English, D. S.; Mattoussi, H. Nano Lett. 2007, 7, 3157−3164. (22) Li, X.; Qian, J.; Jiang, L.; He, S. Appl. Phys. Lett. 2009, 94, 063111. (23) Gueroui, Z.; Libchaber, A. Phys. Rev. Lett. 2004, 93, 166108. (24) Medintz, I. L.; Uyeda, H. T.; Goldman, E. R.; Mattoussi, H. Nat. Mater. 2005, 4, 435−446. (25) Medintz, I. L.; Clapp, A. R.; Mattoussi, H.; Goldman, E. R.; Fisher, B.; Mauro, J. M. Nat. Mater. 2003, 2, 630−638. (26) Boeneman, K.; Prasuhn, D. E.; Blanco-Canosa, J. B.; Dawson, P. E.; Melinger, J. S.; Ancona, M.; Stewart, M. H.; Susumu, K.; Huston, A.; Medintz, I. L. J. Am. Chem. Soc. 2010, 132, 18177−18190. (27) Clapp, A. R.; Medintz, I. L.; Mattoussi, H. ChemPhysChem. 2006, 7, 47−57. (28) Sharma, J.; Chhabra, R.; Liu, Y.; Ke, Y. G.; Yan, H. Angew. Chem., Int. Ed. 2006, 45, 730−735. (29) Sharma, J.; Chhabra, R.; Andersen, C. S.; Gothelf, K. V.; Yan, H.; Liu, Y. J. Am. Chem. Soc. 2008, 130, 7820−7821. (30) Ding, B. Q.; Deng, Z. T.; Yan, H.; Cabrini, S.; Zuckermann, R. N.; Bokor, J. J. Am. Chem. Soc. 2010, 132, 3248−3249. (31) Deng, Z.; Samanta, A.; Nangreave, J.; Yan, H.; Liu, Y. J. Am. Chem. Soc. 2012, 134, 17424−17427. (32) Lin, C.; Liu, Y.; Yan, H. Biochemistry 2009, 48, 1663−1674. (33) Pal, S.; Deng, Z. T.; Ding, B. Q.; Yan, H.; Liu, Y. Angew. Chem., Int. Ed. 2010, 49, 2700−2704. (34) Ding, B.; Deng, Z.; Yan, H.; Cabrini, S.; Zuckermann, R. N.; Bokor, J. J. Am. Chem. Soc. 2010, 132, 3248−3249. (35) Pal, S.; Deng, Z.; Wang, H.; Zhou, S.; Liu, Y.; Yan, H. J. Am. Chem. Soc. 2011, 133, 17606−17609. (36) Shreiber, R.; Do, J.; Roller, E. M.; Zhang, T.; Schuller, V. J.; Nickels, P. C.; Feldman, J.; Liedl, T. Nat. Nanotechnol 2014, 9, 74−78. (37) Lan, X.; Chen, Z.; Dai, G.; Lu, X.; Ni, W.; Wang, Q. J. Am. Chem. Soc. 2013, 135, 11441−11444. (38) Shen, X.; Asenjo-Garcia; Liu, Q.; Jiang, Q.; Garcia de Abajo, F. J.; Liu, Na.; Ding, B. Nano Lett. 2013, 13, 2128−2133. (39) Shen, X.; Song, C.; Wang, J.; Shi, D.; Wang, Z.; Liu, N.; Ding, B. J. Am. Chem. Soc. 2012, 134, 146−149. (40) Kuzyk, A.; Schreiber, R.; Fan, Z.; Pardatscher, G.; Roller, E. M.; Hogele, A.; Simmel, F. C.; Govorov, A. O.; Liedl, T. Nature 2012, 483, 311−314. (41) Bui, H.; Onodera, C.; Kidwell, C.; Tan, Y.; Graugnard, E.; Kuang, W.; Lee, J.; Knowlton, W. B.; Yurke, B.; Hughes, W. L. Nano Lett. 2010, 10, 3367−3372. 5057

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