Dynamics and Stability of DNA Mechano-Nanostructures: Energy

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Dynamics and Stability of DNA Mechano-Nanostructures: Energy-Transfer Investigations B. Kolaric*,† and R. A. L. Valle´e‡ Laboratoire Interfaces & Fluides Complexes, Centre d’InnoVation et de Recherche en Mate´riaux Polyme`res, UniVersite´ de Mons Hainaut, 20 Place du Parc, 7000 Mons, Belgium, and Centre de Recherche Paul Pascal (CNRS), 115 aVenue du docteur Albert Schweitzer, 33600 Pessac, France ReceiVed: August 10, 2009; ReVised Manuscript ReceiVed: October 26, 2009

In this article, the assembly of double-stranded DNA (dsDNA) into a DNA parallelogram is described. The stability and dynamics of the corresponding superstructure were investigated by using fluorescence resonance energy transfer (FRET). For these purposes two dyes, Cy3 and Cy5, were covalently attached at the extremity of one junction. Fluorescence investigations revealed a clear difference of the dynamics and stability of the parallelogram compared to a single junction and show that FRET can be used to determine stability and dynamics of mechanically interlocked compounds, which is crucial for a fundamental understanding of their behavior. Introduction Nanoscience is the emerging science of objects that are intermediate in size between 10 and 100 nm. Today, nanoscience is a very diverse field, ranging from extensions of conventional device physics, to completely new approaches based on selfassembly, to the development of new materials with dimensions on the nanoscale. DNA, whatever its biological significance, is one of the favorite building blocks for designing complex nanostructures.1-15 Almost 50 years after the famous Feynman lectures,15 different nanostructures such as nanorods, dendrimers, quantum dot, and buckytubes have been successfully synthesized1-13,16 and have attracted the attention of the physical chemistry community because of the unusual electrical and optical behavior of their electrons and photons. It is important to notice that life science is also a nanoscience, considering the dimensions of cellular compartments, as well as at least one dimension of biomolecules such as DNA, proteins, and their conjugates.1-14 The ultimate goal of structural DNA nanotechnology is to build nanoscale objects and devices from DNA, as well as to achieve their self-assembly into periodic arrays. The formation and stability of any self-assembled structure are based on minimizing intermolecular and intramolecular interactions1-6,9 between the constitutive units. DNA nanotechnology takes advantage of the diversity of DNA intermolecular interactions, which are highly specific and very well programmed through Watson-Crick complementarity.1-6 The advantage of fusing DNA for constructing the parallelogram structure lies in the specificity of base pairing and DNA’s robust physicochemical structure Understanding the dynamics of interlocked DNA structures is essential because of their possible applications. Future applications of such interlocked structures range from macromolecular crystallography and new materials to molecular electronics and DNA-based computation.2,4,6,10 In the case of either the DNA parallelogram or the single DNA junction nanostructures (Figure 1), these interactions are noncovalent and highly directional in nature (electrostatic, hydrogen,

Figure 1. Schematic representation of the DNA parallelogram (R) and the DNA junction (J). The sticks represent the DNA double strands; blue and violet balls represent Cy3 and Cy5 molecules, respectively.

steric, and van der Waals), such that they depend strongly on the orientation between the constitutive molecules.1-6,17-19 Taking into account the number of bases per oligonucleotide strand and the amount of counterions used to stabilize the structures, the charges located on the backbone are partially screened, and the junction appears to be bendable from a structural point of view. Atomic force microscopy (AFM) images published elsewhere34 clearly confirm that these junctions are quite flexible. The DNA parallelogram structure described in this article is simply the juxtaposition of four Holliday junctions, with rigidity too low to be used for geometrically controlled assemblies. Nevertheless, the extra rigidity exhibited by the parallelogram structure is probably an effect of the mechanical coupling of all of the constituent junctions. An increase of the temperature generates additional energy17-19 within the system, which could cause a change of the arrangement of the single junctions within the structure of the overall parallelogram or even cause a rupture of the structure. Note also that the presented results clearly demonstrate that the stability and dynamics of self-assembled structure can be investigated by fluorescent methods. Results and Discussion

* Corresponding author. E-mail: [email protected]. † Universite´ de Mons Hainaut. ‡ Centre de Recherche Paul Pascal (CNRS).

The investigation of the mechanical rigidity of the parallelogram structure of DNA as a function of temperature by

10.1021/jp907704e  2010 American Chemical Society Published on Web 01/05/2010

Dynamics and Stability of DNA Mechano-Nanostructures

Figure 2. Fluorescence emission spectra of the FRET pair in the parallelogram structure as a function of temperature; the structure was excited at a wavelength of 530 nm.

fluorescence spectroscopy is one of the key points of this article. Because stability is an essential issue for self-assembled structures, we have investigated the influence of temperature on the dynamics and stability of the DNA parallelogram by fluorescence resonance energy transfer (FRET).2,13,20,,,23 The established link between FRET response and thermodynamic stability suggests the broader application of FRET and offers the possibility of using photophysical methods to study the dynamics of interlocked compounds at the bulk and singlemolecule levels. Long-range energy transfer from the Cy3 donor to the Cy5 acceptor through dipole-dipole interactions was initially described by Fo¨rster.20-22 The corresponding dyes are not the best fluorescent probes,32 but they were the most suitable for our investigations because of their small size (which will not disturb the dynamics of the system) and simple labeling procedure. The sensitivity of the energy-transfer process to the nanoscale environment21-24 of the donor-acceptor system allows the dynamics of the interconnected system to be probed efficiently. This efficiency is known to depend crucially on the orientation angle between the emission transition dipole moment of the donor and the absorption transition dipole moment of the acceptor, as well as on the donor-acceptor distance.20,22 In the case of a charged polymer such as DNA, a cloud of counterions within the polymer redistributes as a function of the donor-acceptor separation, making the FRET process in such a complex structure a superposition of many contributions that cannot be easily separated.22-25 Figure 2 shows the fluorescence emission spectra of the Cy3 donor-Cy5 acceptor system related to the parallelogram structure, observed by steady-state measurements as a function of temperature. Photoexcitation of the system at a wavelength of 530 nm primarily induces a transition from the ground state to the first excited state of the Cy3 molecule. Direct emission from this state leads to an emission spectrum that peaks at 560 nm, which constitutes the first part (shorter-wavelength side) of the spectra shown in Figure 2. Alternatively, the energy-transfer process, which is faster than the primary radiative deactivation process of Cy3 (the donor), can proceed to populate the first excited state of the Cy5 molecule (the acceptor). The radiative emission of Cy5 results in the observation of the second part of the emission spectra (longer-wavelength peaks shown in Figure 2). The dominance of this second part in the observed spectra clearly indicates an efficient energy-transfer process for the Cy3-Cy5 system in the DNA parallelogram structure. The same experiment was performed for the single junctions (Figure 3). In this case, the energy-transfer process is less efficient, as indicated by the lessened reduction of intensity of

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Figure 3. Fluorescence emission spectra of the FRET pair in the singlejunction structure as a function of temperature; the excitation wavelength was 530 nm.

Figure 4. Calculated efficiency of Cy3-Cy5 energy transfer for both parallelogram and single-junction double-stranded DNA (dsDNA) nanostructures as a function of temperature.

the band corresponding to the donor emission in the spectra (Figure 3). For calculation of the FRET efficiency (eq 1), the apparent energy-transfer efficiency was used. Spectra were normalized at the maximum. In both cases, the relative intensities of the two parts of the spectra allow for the calculation of the energy-transfer efficiency as

Eeff )

IA ID + IA

(1)

where IA is the normalized fluorescence emission intensity of the acceptor and ID is the normalized fluorescence emission intensity of the donor. The calculated energy-transfer efficiencies for the two types of structures are plotted as a function of temperature in Figure 4. Figure 4 clearly shows that the energy-transfer efficiency is systematically larger in the parallelogram structure than in the single-junction structure (at least up to about 50 °C). Furthermore, Figure 4 shows that, in the case of DNA single junctions, the Cy3-Cy5 energy transfer drops dramatically and abruptly at around 32 °C (taken in the middle of the curve). In the case of the DNA parallelogram, a smoother drop is observed, at higher temperature (around 42 °C). From steady-state spectra, the Fo¨rster distance has been determined20,22,23 to be around 6 nm. Taking into account the excellent overlap between the donor emission and acceptor absorption spectra, the systematically smaller energy-transfer efficiency exhibited by the FRET pairs in the single-junction structures has to be related to the higher mobility of these structures as compared to the parallelograms, with the consequent disordered motions of their linkers causing

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Kolaric and Valle´e

Figure 5. Time-resolved fluorescence decay profile and corresponding fit (red curve) of the donor of the FRET pair, taken at the emission wavelength of 562 nm in the parallelogram structure at 25 °C. The excitation wavelength was 488 nm.

the occurrence of distributions of distances and orientations between the FRET pairs.20-24 Furthermore, the clear and abrupt reduction of the energy-transfer efficiency for the junction seems to indicate a transition from an ordered structure, in which motions of the junctions are correlated to motions of the linkers, to a more disordered structure, in which these collective motions lose their coherence, at a critical temperature around 32 °C. Indeed, the DNA single junction is not a rigid-rod cylinder but a bendable structure with motions whose amplitudes vary as a function of the applied energy and the sequence of base pairs.26 A small increase of only 15 °C (from 20 to 35 °C) is enough, in the case of the junction, to overcome the favorable interaction that stabilizes the structure. In the case of the parallelogram, the decrease of the energy-transfer efficiency as a function of temperature is smooth, which indicates a higher rigidity of the parallelogram structure compared to the single junction, as well as synchronous motions of the constitutive units within the parallelogram (see Figure 4). Here, it should be emphasized that static quenching22 can be excluded as a dominant cause of the decrease of the FRET efficiency with increasing temperature because, in the case of static quenching, there would be no difference in the quenching of the FRET signal between the parallelogram and the junction due to the fact that the concentrations of the probes (parallelogram and junction) and quality (viscosity and ionic strength) of the solvent used to dissolve the two structures were the same. To further investigate the influence of temperature on the dynamics and stability of these two nanostructures, we performed time-resolved measurements of the fluorescence decay profiles of the FRET pairs in the structures. The decay profiles (Figure 5) were fit best with a multiexponential function (as described in the Experimental Details) consisting of three decay times. The fitting of the fluorescence decay profiles, best performed by multiexponential functions (Figure 5), has to be attributed to the complexity of the DNA-dye dynamics.12,21,22 Observed decay times can be related to three different processes: The fastest decay is related to the fast dynamics of the linkers and

side groups at the backbone, the intermediate process is related to the dynamics of the backbone itself, and longest decay is related to the decay time of the single dye (Cy3). Increasing the temperature modifies the relaxation modes because of the large fluctuations occurring in the system. These decay times and their populations for various temperatures are summarized in Table 1. The results presented in Table 1 show that values of all decay times did indeed decrease with temperature, but the populations showed the opposite trend. Note that contributions of τ2 and τ3 decrease as a function of temperature, in contrast to the contribution of τ1. The influence of temperature on the decay time of Cy3 attached to the parallelogram is also graphically presented in Figure 6a. Because τ2 and τ3 are related to the dynamics of the parallelogram and the dye itself, changes in the parallelogram structure as a function of temperature affect these two decay times, as was observed. Furthermore, the decay times (τ1, τ2, τ3) decrease faster as a function of temperature in the case of the single-junction nanostructure than in the case of the parallelogram nanostructure (Table 1). On the basis of the literature,12,21-23,35 we attribute the largest decay time τ3 to the fluorescence decay time of the Cy3 donor. Given the opposite trends shown by τ1 and τ2 as functions of temperature, we can tentatively assign the fastest decay components τ1 to the dynamics of the linkers and side groups and τ2 to the dynamics of the DNA backbone. Indeed, the increased contribution of molecules, together with the reduced value of the decay time τ1 observed as a function of temperature, suggests the occurrence of increased mobility of the linkers and/or small segments in the DNA structures.12,21,22 On the contrary, although the reduction of decay time τ2 as a function of temperature is also a sign of increased mobility of the structures, the reduced number of molecules participating in these motions as a function of temperature indicates a loss of concerted dynamics of larger parts of the DNA backbones.27 The observed difference in the decay-time contributions for the single-junction and parallelogram nanostructures is a consequence of their different rigidities (with increased mobility of the single-junction structure with respect to the parallelogram structure). The small variation of the τ1 decay time occurring at different temperatures in the parallelogram structure indicates the relatively good conservation of the synchronous motion of the side groups and small segments within this rigid structure. In contrast to the behavior of this fastest τ1 decay time in the parallelogram structure, the two other decay times show a linear decrease with increasing temperature. By increasing the temperature, one assists faster motions of the molecules, more collisions between them, and faster conformational changes, which causes a quenching of the donor emission and a reduction of the donor-acceptor energy transfer because of an unfavorable orientation between donor-acceptor pairs. A similar influence of the rigidity of the backbone on the macromolecular dynamics for various synthetic polyelectrolytes is very well described elsewhere.28,29 For the single-junction nanostructure, on the contrary, both τ1 and τ3 decrease faster as a function of temperature, which provides further evidence for the higher

TABLE 1: Decay Times and Respective Contributions of the Donor Measured at the Emission Wavelength of 562 nm for the Single-Junction (J) and Parallelogram Nanostructures (r) as a Function of Temperature τ1 (ns) R R R R

/J, /J, /J, /J,

25 35 45 55

°C °C °C °C

0.22 0.23 0.15 0.16

(30) (50) (50) (65)

τ2 (ns) 0.27 0.19 0.17 0.14

(28) (44) (55) (60)

0.78 0.65 0.51 0.50

(41) (35) (40) (29)

τ3 (ns) 0.82 0.61 0.54 0.44

(48) (41) (35) (33)

1.83 1.71 1.64 1.57

(29) (15) (10) (6)

1.82 1.64 1.5 1.40

(24) (15) (10) (7)

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Figure 6. (a) Decay times of Cy3 attached at the parallelogram as a function of temperature. (b) Decay times (τ3 of Cy3) as a function of temperature for the (2) parallelogram and (b) junction. (c) Average lifetime as a function of temperature for the (2) parallelogram and (b) junction.

conformational mobility of this structure compared to the parallelogram structure. The faster dynamics of the junction in comparison to the parallelogram is visible in Figure 6b,c, where the decay times of the donor (Cy3) and the overall average decay times, respectively, are plotted as a function of temperature for both the junction and parallelogram structures. Figure 6b clearly shows the effect of temperature on the lifetime of the dye (Cy3) attached to the backbones of the different DNA structures. The longest decay time corresponds to the lifetime of the dye, and because the solvent quality and concentration of the dye attached to the backbone were the same, the observed difference can be explained in terms of the different rigidities of the parallelogram and junction. Note that an increase of the temperature causes bending of the structure, simultaneously changing the nanoscale environment around the dye (affecting the photoresponse of the single dye) and the spectral overlap between the donor and acceptor molecules, which affects the FRET response. Because of the different rigidities, which affect the bending dynamics, the corresponding decay-time profiles of Cy3 are different for the parallelogram and for the junction (Figure 6b). For the two structures, the average decay times are also different and decrease with increasing temperature. To further illustrate the temperature sensitivity of the FRET efficiency in the parallelogram and junction structures, we determined the speed of rupturing, that is, the speed of loss of mechanical stability (LMS) as the slope from the experimentally measured average decay time as a function of heating (Figure 6c). The slope of LMS shows how fast (easily) the FRET efficiency changes with increasing temperature.30 The latter30,31 can be related to the loss of correlations between the motions of the linkers and the motions of the DNA structures. In the case of the junction, the slope is 11 ps °C-1, which is almost 50% higher than for the parallelogram (6 ps °C-1). Clearly, the parallelogram structure better preserves its rigidity as a function of temperature than the DNA single junction. Note that an investigation of the thermodynamic stability of a small DNA hexagon using fluorescence spectroscopy was recently published elsewhere.32 Our work can be understood as an extension of the published method because measuring FRET efficiency as a function of temperature allows for the simultaneous investigation

of the mechanical stability and dynamics of changes induced by external factors, such as temperature. Conclusions In conclusion, the data presented in this article clearly show differences between the mobilities and rigidities of singlejunction and parallelogram DNA nanostructures. Both FRET measurements and time-resolved decay profile analysis, performed at different temperatures, allowed the dynamics and stability of these different DNA nanostructures to be distinguished. The energy-transfer efficiency is almost 25% larger for the parallelogram than for the single-junction structure at ambient temperature (20 °C) because of the rigid interlocked parallelogram structure and synchronous motion of its subunits. This energy-transfer efficiency reduces smoothly as a function of temperature in the case of the parallelogram nanostructure, in contrast to the case of the single-junction nanostructure. The DNA parallelogram structure preserves its rigidity and keeps the synchronous motion of the constitutive segments even at high temperatures (higher than 42 °C). This interpretation is further confirmed by the evolution of the various decay times of the donor linked to each type of nanostructure as a function of temperature. The rigidity and stability of the parallelogram at higher temperature will allow for the use of this structure as a building block in complex arrays or supramolecular systems. Tuning the structures as function of temperature is not possible because corresponding self-assembly structures are formed by optimizing the electrostatic, steric, and hydrogen interactions between constituents. The annealing procedure is needed to reform the parallelogram from the loose relaxed parallelogram structure. In summary, the fluorescence lifetime of labeled DNA has been shown to provide a useful, generalizable method for measuring temperature changes and determining the thermodynamic and mechanical stability of the system. This method is completely noninvasive and can be applied to microliter-sized volumes and for investigations of the dynamics and stability of various interlocked compounds.33 Also, considering fluorescence detection, the corresponding method can be scaled up to the

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SCHEME 1: Parallelogram and Holliday Junction Sequences

single-molecule level, which offers the possibility of studying the dynamics and stability of interlocked compounds at the single-molecule level.

Kolaric and Valle´e was directed into a frequency-doubled optical parametric oscillator to deliver the 488-nm wavelength used for excitation. Emission light, passed through a sheet polarizer set to the magic angle (54.7°), was focused on a monochromator (9030DS, Sciencetech) to obtain spectral separation and detected with a microchannel plate (R3809U, Hamamatsu). Acquisition was done using a TCSPC PC card (SPC 430, Becker & Hickl GmbH, Berlin, Germany). The experimental instrument response function was on the order of 50 ps. Decays were collected in 4096 channels. Data from single-photon timing were analyzed using a homemade program based on the Marquardt algorithm, by convoluting the experimental instrument response function with a sum of exponentials. The fitting was judged by the value of the reduced χ2 parameter and also by the appearance of the residuals and the autocorrelation function. The fluorescence intensity decays were analyzed in term of a multiexponential model12,22,35

Experimental Details Materials. The oligonucleotide structures described in this article were a gift from the Samori group (University of Bologna, Bologna, Italy) To provide more information to the readers about the oligonucleotides, the following information was taken from Marco Brucale’s Ph.D. thesis:34 The sequences of the oligonucleotides used in this work were designed with the program SEQUIN. Oligonucleotides were purchased from MWG (Ebersberg, Germany), HPLC-purified and lyophilized by the supplier, and suspended in Milli-Q water. The final concentration of each strand was 0.1 µM. The concentration of each strand was estimated by UV spectroscopy by measuring OD260. The resulting sequences are shown in Scheme 1. Both DNA structures were assembled in solution by mixing stoichiometric quantities of each component strand in Trisacetate-EDTA (TAE, where EDTA is ethylenediaminetetraacetic acid)/Mg2+ buffer (20 mM Tris, 2 mM EDTA, 12.5 mM MgCl2, pH 8.4). The mixtures were then heated to 90 °C for 5 min and cooled to 10 °C in a polymerase chain reaction (PCR) thermocycler (PCR Sprint, Thermo Electron Corp., Waltham, MA) at a rate of 0.01 °C s-1. The correct formation of the structures was verified with native polyacrylamide gel electrophoresis (PAGE) experiments (data not shown) in which the complete structures ran as a single band. In our case, the two dyes, Cy3 and Cy5, meet all the required conditions for FRET. First, they have a significant spectral overlap. Although they are not perfectly free to rotate in space, the relative orientations of their dipole moments vary constantly because they are tethered to the DNA structure with quite flexible linkers (C6 linkers). The structures to which they are tethered keep them at a distance of a few nanometer (4.5 nm). The linkers used to anchor the dyes to the DNA are also identical. Methods. Fluorescence emission spectra were recorded using a FluoroMax-3 spectrofluorometer (SPEX Instruments, Edison, NJ) at an excitation wavelength of 530 nm and a band pass of (1 nm. Fluorescence emission was resolved in wavelength by use of a band-pass filter of (2 nm between 550 and 700 nm. The temperature was controlled by an LFI-3751 thermoelectric temperature controller from Wavelength Electronics. The fluorescence decay times and the time-resolved fluorescence spectra in all solvents were determined by using a timecorrelated single-photon-counter (TCSPC) setup, which is described in detail elsewhere.12,35 In brief, linearly polarized light of a mode-locked Ti:sapphire laser (Tsunami 3950D, Spectra Physics, 1.2 ps pulse, 4.09 MHz repetition rate, λpump ) 976 nm) pumped by a diode laser (Millennia Xs, Spectra Physics)

I(t) )

∑ Ri exp(-t/τi)

(2)

i

where Ri represents the pre-exponential factors and τi represents the lifetimes. This model is described elsewhere.22 The contribution of each lifetime component to the steady-state fluorescence is given by the formula

fi )

Riτi

∑ Rjτj

(3)

Acknowledgment. The authors acknowledge support from a Eurocores grant (ESF-Bionics). The Fonds voor Wetenschappelijk Onderzoek Vlaanderen is thanked for a postdoctoral fellowship for R.A.L.V. and for Grants G.0421.03 and G.0458.06. INPAC is thanked for a postdoctoral grant for B.K. The authors warmly acknowledge Bruno Samori, Giampaolo Zuccheri, and Marco Brucale (Department of Biochemistry “G. Moruzzi” and National Institute for the Physics of the Matter, University of Bologna, Bologna, Italy) for providing compounds. Marco Brucale is additionally thanked for participation in preliminary steady-state investigations. The authors also warmly acknowledge Frans de Schryver and Johan Hofkens for providing the fluorescence setup, in order to perform the experiments during their postdoctoral stay in the Laboratory of Photochemistry and Spectroscopy at KU Leuven (Belgium). References and Notes (1) Niemeyer, M. C. Angew. Chem., Int. Ed. 2001, 40, 4128–4158. (2) Seeman, C. N. Biochemistry 2003, 42, 7259–7269. (3) Simonsson, T.; Sjoback, R. J. Biol. Chem. 1999, 274, 17379–17383. (4) Seeman, C. N.; Lukeman, P. S. Rep. Prog. Phys. 2005, 68, 237– 270. (5) Samori, B.; Zuccheri, G. Angew. Chem., Int. Ed. 2005, 44, 1166– 1181. (6) Brucale, M.; Zuccheri, G.; Samori, B. Trends Biotechnol. 2006, 24, 235–243. (7) Rothemund, K. W. P. Nature 2006, 440, 297–302. (8) Becker, W. F. C.; Wacker, R.; Bouschen, R.; Seidel, R.; Kolaric, B.; Lang, P.; Schroeder, H.; Muller, O.; Niemeyer, M. C.; Spengler, B.; Goody, S. R.; Engelhard, M. Angew. Chem., Int. Ed. 2005, 44, 7635–7639. (9) Whitesides, M. G. Nanoscience, Nanotechnology, and Chemistry. Small 2005, 1, 172–179. (10) Liao, P. S.; Mao, D. C.; Birktoft, J. J.; Shuman, S.; Seeman, N. C. Biochemistry 2004, 43, 1520–1531. (11) Xu, Q.; Wang, S.; Korystov, D.; Mikhailovsky, A.; Bazan, C. G.; Moses, D.; Heeger, J. A. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 530– 535.

Dynamics and Stability of DNA Mechano-Nanostructures (12) Kolaric, B.; Sliwa, M.; Brucale, M.; Valle´e, L. A. R.; Zuccheri, G.; Samori, B.; Hofkens, J.; de Schryver, C. F. Photochem. Photobiol. Sci. 2007, 6, 614–618. (13) Shi, L.; Rosenzweig, N.; Rosenzweig, Z. Anal. Chem. 2007, 79, 208–214. (14) Mergny, L. J.; Maurizot, C. J. ChemBioChem 2001, 2, 124–132. (15) Feynman, R. There’s Plenty of Room at the Bottom: An Invitation to Enter a New Field of Physics. http://www.zyvex.com/nanotech/feynman.html. (16) Grimsdale, C. A.; Mullen, K. Angew. Chem., Int. Ed. 2005, 44, 5592–5629. (17) Isrealchvilli, J. Intermolecular and Surface Forces; Academic Press: London, 1998. (18) Evans, F. D.; Wennerstro¨m, H. Colloidal Domain; Wiley-VCH: New York, 1999. (19) Shakhnovich, E. Chem. ReV. 2006, 106, 1559–1588. (20) Stryer, L. Annu. ReV. Biochem. 1978, 47, 819–846. (21) Sabanayagam, R. C.; Eid, S. J.; Meller, A. J. Chem. Phys. 2005, 122, 061103. (22) Lackowicz, R. Principles of Fluorescence Spectroscopy, 2nd ed.; Kluwer Academic/Plenum Publishers: New York, 1999, and references therein. (23) Kolaric, B.; Baert, K.; Valle´e, L. A. R.; van der Auweraer, M.; Clays, K. Chem. Mater. 2007, 19, 5547–5552. (24) Ying, L.; Wallace, I. M.; Balasubramanian, S.; Klenerman, D. J. Phys. Chem. B 2000, 104, 11551–115555.

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