Scanning Single-Molecule Fluorescence Correlation Spectroscopy

May 3, 2016 - To effectively bridge this millisecond gap, we developed a single-molecule fluorescence correlation spectroscopy (smFCS) method that wor...
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Scanning Single-Molecule Fluorescence Correlation Spectroscopy Enables Kinetics Study of DNA Hairpin Folding with a Time Window from Microseconds to Seconds Huimin Bi, Yandong Yin,† Bailong Pan, Geng Li, and Xin Sheng Zhao* Beijing National Laboratory for Molecular Sciences, State Key Laboratory for Structural Chemistry of Unstable and Stable Species, Department of Chemical Biology, College of Chemistry and Molecular Engineering, and Biodynamic Optical Imaging Center (BIOPIC), Peking University, Beijing 100871, China S Supporting Information *

ABSTRACT: Single-molecule fluorescence measurements have been widely used to explore kinetics and dynamics of biological systems. Among them, single-molecule imaging (SMI) is good at tracking processes slower than tens of milliseconds, whereas fluorescence correlation spectroscopy (FCS) is good at probing processes faster than submilliseconds. However, there is still shortage of simple yet effective single-molecule fluorescence method to cover the time-scale between submilliseconds and tens of milliseconds. To effectively bridge this millisecond gap, we developed a single-molecule fluorescence correlation spectroscopy (smFCS) method that works on surface-immobilized single molecules through surface scanning. We validated it by monitoring the classical DNA hairpin folding process. With a wide time window from microseconds to seconds, the experimental data are well fitted to the two-state folding model. All relevant molecular parameters, including the relative fluorescence brightness, equilibrium constant, and reaction rate constants, were uniquely determined.

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To deal with this situation, efforts have been made in both SMI and FCS. In SMI, millisecond time resolution has been recently realized9 using scientific complementary metal-oxide semiconductor (sCMOS) as detectors, but further improvements to submilliseconds and faster are still challenging. In FCS, efforts to extend the time window to milliseconds and slower have been made in two ways. One is to slow down the free diffusion. The trials include mixing molecules in agarose,10 attaching big objects to the target biomolecules,11,12 using dual beam fluorescence cross-correlation spectroscopy,13 and so on. However, they all have different deficiencies. For example, in our previous work,14 we extended the diffusion time to seconds by attaching DNA molecules to microspheres, so that the dynamics of spontaneous single base pair flipping with relaxation times on the order of ten milliseconds were resolved. Nevertheless, we found that time and efforts were needed in practice to keep experimental conditions stable to collect statistically sufficient data. The other way is to get rid of the free diffusion by molecular trapping or surface-immobilization. The anti-Brownian electrokinetic (ABEL) trap15 enables observation in solution up to seconds. It can be implemented in the FCS studies. The idea of immobilizing molecules on surface16 is also facile and effective. In order to realize high time and spatial resolution, APD detectors are used together with a singlemolecule positioning apparatus,17 which scans the surface to search for the molecules first and then jumps from one

vercoming ensemble averaging, single molecule detection (SMD) is powerful to display molecular diversity and to provide dynamic information without synchronization.1 In recent years, the SMD techniques, especially the singlemolecule fluorescence measurements, have been widely used to deal with kinetic and dynamic questions in biological systems.2−7 Biological processes, including conformational changes and molecular interactions, occur in a wide time range, from below nanoseconds to beyond minutes.8 With this concern, choosing a method with sufficient time resolution as well as an appropriate time window is vital for rational kinetic and dynamic measurements. In this respect, the two prevalent categories of single molecule fluorescence methods, the single molecule imaging (SMI)3−5 and the fluorescence correlation spectroscopy (FCS),6,7 both have their own merits and limitations. SMI, working with wide-field illumination and electron multiplying charge coupled device (EMCCD) sensitivity, usually needs at least tens of milliseconds time binning to achieve satisfactory S/N ratio, and is therefore applicable for tracking processes slower than tens of milliseconds. On the other hand, FCS is generally used to study processes in solution with nanosecond time resolution using avalanche photon diodes (APDs) as detectors, but its longest possible observation time is limited by the molecular diffusion. With a typical dwell time of submilliseconds for most biomolecules in the confocal detection volume, it is hard for a conventional FCS to track processes slower than that. Clearly, in the toolbox of SMD, there is still a shortage of techniques that fill the gap between submilliseconds and tens of milliseconds. © XXXX American Chemical Society

Received: March 31, 2016 Accepted: May 3, 2016

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Figure 1. Scanning confocal fluorescence microscope platform. (a) Diagram of the scanning smFCS platform. (b) A typical image (10 μm × 10 μm) of TMR-labeled DNA. Every single light spot corresponds to one molecule. (c) The time traces collected by the two APDs during a line scan. The density of the immobilized molecules was kept on a single-molecule level. (d) The time window of the scanning smFCS is 4 orders of magnitude longer than that of the conventional FCS. The sample used here is polyT-4G-24. The conventional FCS was conducted in solution, so that the time window is limited by the diffusion (τD = 740 μs in this example). While in the scanning smFCS, the time window is controlled by the scan speed (τS = 3.8 s in this example).

Biomolecules labeled with fluorophores were immobilized on the coverslip surface, which was functionalized with biotinPEG.25 It should be assured that the probability of more than one molecule in the laser focus was negligible. The piezo stage carrying the coverslip was scanned across the laser focus linearly so that biomolecules were illuminated one after another. The fluorescence was split to two beams and then detected by two APDs. The cross-correlation of the two beams was in situ calculated with a correlator (Flex 02−01D) as the conventional FCS did.12 In the conventional FCS, an FCS curve, G(τ), is described by26

molecule to another to record FCS individually. Although this search-and-positioning strategy has a wide time window from microseconds to seconds, it is time-consuming and may suffer from bleaching. In a more efficient way, FCS data can be collected while scanning through the surface-immobilized molecules as the scanning FCS (sFCS) does. The idea of sFCS was presented almost at the same time as the FCS technique was invented,18,19 and it has been applied with various purposes.20−22 But to our knowledge, the application to study dynamics of immobilized molecules was only introduced by Xiao et al.23 They exhibited the microsecond dynamics of surface-bound fluorescent species. However, in this time range, the conventional FCS actually works better than the sFCS. Here, we tried to apply the sFCS to the millisecond region and therefore developed the scanning single-molecule FCS (scanning smFCS). There are two key points in this method: (1) single-molecule and (2) controllable scan. Our previous work found that the amplitudes of the relaxations are inversely proportional to the number of fluorophores moving together.24 Therefore, keeping the experiment on single molecule condition is critical to ensure that the FCS data are not distorted and easy to analyze. With controllable scan over immobilized molecules, we can prolong the time window of FCS observation in an effective and efficient manner. The experimental platform is based on a home-built scanning confocal fluorescence microscope (Figure 1a). We equipped a piezo stage on the basis of a home-built conventional FCS apparatus to perform the scanning.17 Certainly, a commercial scanning confocal fluorescence microscope will also be usable.

G(τ ) = G D(τ ) ·G R (τ )

(1)

−1/2 −1 1 ⎛ τ ⎞ ⎛ τ ⎞ G D(τ ) = ·⎜1 + ⎟ · ⎜1 + 2 ⎟ N ⎝ τD ⎠ ⎝ w τD ⎠

(2)

where GD(τ) is the contribution from the free diffusion, GR(τ) is that from chemical reactions, conformational fluctuations, or other possible kinetic processes, N denotes the average number of molecules in the laser focal volume, τD is the characteristic diffusion time, and ω is the ellipsoid parameter of the laser focus. In the scanning smFCS,19 G(τ ) = GS(τ ) ·G R (τ ) 1866

(3) DOI: 10.1021/acs.jpclett.6b00720 J. Phys. Chem. Lett. 2016, 7, 1865−1871

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Figure 2. Hairpin design and scanning smFCS under different NaCl concentrations. (a) Diagram of hairpin folding equilibrium. (b) As the concentration of NaCl increases, the fluorescence intensity of hp-4G-polyT-24 decreases, but that of 4G-polyT-24 remains constant. Error bars denote the s.d. of three independent experiments. In some cases, error bars are smaller than the symbols. (c,d) Scanning smFCS curves of hp-4GpolyT-24 and 4G-polyT-24. The scan speed (v) was the same for all the curves, at 0.1 μm/s. Because the laser was focused on the surface by the naked eyes, the laser width (ωxy) on the surface altered slightly every time, so that the trailing edges do not overlap due to different τS (see eq 4).

Figure 3. (a) GR(τ) from conventional FCS. (b) GR(τ) from scanning smFCS with double-exponential-decay fitting curves. (c−f) Fitting results of the two reaction components in GR(τ). Parameters of the faster one on microsecond time-scale are shown in (c) and (d), and those of the slower one on hundreds of microsecond time-scale are shown in (e) and (f). Error bars denote the s.d. of three independent experiments.

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⎛ ⎛ ⎞2 ⎞ τ 1 ·exp⎜⎜ −⎜ ⎟ ⎟⎟ , τ N ⎝ S⎠ ⎠ ⎝

τS =

(Figure 2d, Figure S2a). This indicates that the obvious changes from microseconds to submilliseconds are related to the folding process rather than other photophysical processes. In the conventional FCS, the correlation curve of hp-4GpolyT-24 is divided by that of 4G-polyT-24 under the same salt concentration to get GR(τ)6 (Figure 3a). Because τD is submilliseconds (Figure S2b), the S/N ratio worsens quickly when τ is larger than 1 ms. Under this condition, it is not clear whether there are kinetic reactions in the millisecond region. We also tried to get GR(τ) directly by least-squares fitting of the FCS curves using eq 1 and eq 2. The fitting results are shown in Figure S2 and Figure S3, but they are unacceptably off from the true values (see discussion in the Supporting Information), and therefore will not be further discussed. In contrast to τD in the conventional FCS, τS in the scanning smFCS is well separated from the characteristic times of kinetic relaxation, so that the influence of GS(τ) on GR(τ) is negligible. A clear and reliable GR(τ) is therefore obtained using the dividing method. The S/N ratio is good from microseconds to seconds. Comparing Figure 3b with 3a, it is assured that the extra component in the millisecond range in Figure 3a is not a true portion of GR(τ) but the residue of incomplete removal of GD(τ). If there were no experimental evidence from wider time window, the residue might be incorrectly attributed to an additional chemical process. This is a good example to show where the conventional FCS may fail. Based on eq 2, it is easy to see that the conventional FCS is bound to fail to work somewhere τ > τD, which is on the order of milliseconds, because G(τ)s of both the sample and control go to 0. Similarly, the scanning smFCS is bound to fail, too, somewhere τ > τS (eq 4), but that happens when τ is on the order of seconds (see Supporting Information for more discussion). Figure 3 indicates that there are two relaxation processes. To quantitatively fit them, we used a double-exponential function for GR(τ):12

ωxy v

(4)

where GS(τ) is the contribution due to the scan, τS is the characteristic time of the scan, ωxy is the width of the laser focus on the surface, and v is the scan speed. For the scanning smFCS, N has to be controlled at a level less than one. As eq 2 shows, GD(τ) is related to τD of the target molecule, which is intrinsic to the system and the experimental arrangement. Typically, τD is hundreds of microseconds for biomolecules (Figure 1d). Unlike GD(τ), GS(τ) is determined by τS, which is controlled by the scan speed, as shown in eq 4. By changing the scan speed, we can easily regulate the time window (Figure 1d and Figure S1) for a specific process. We then tried to apply the method on folding kinetics of DNA hairpins. DNA hairpins play important roles in biological processes such as DNA replication, transcription, and translation.27 Lots of works6,10,12,13,28−33 have been carried out to explore the hairpin folding kinetics through different ways, but there are still controversial issues regarding the folding and unfolding rates. Furthermore, some researchers found that the hairpin folding process obeys a two-state model,6 whereas others found more intermediates.12,13,29,30 In our opinion, these discrepancies result from different molecular compositions and experimental arrangements, so that different aspects of the process are probed. Here we designed our experiments with two considerations: (1) Choosing a four-stem hairpin with long loops to study. In this way, the time window of the conventional FCS is still suitable but at the margin to fail, so we can check the validity of the scanning smFCS with the established technique and simultaneously show the limit of the conventional FCS. (2) Using a simple fluorophore-quencher design to report the hairpin conformation.34 Quenching TMR by natural guanine instead of an extra labeled dye,6,10,13 possible extraneous interactions between the fluorophores are avoided. In addition, because the photoinduced electron transfer (PET) quenching of TMR by guanine is a short-distance interaction,35 only the folded hairpin is registered as the dark state. The hairpin design is shown in Figure 2a and the DNA sequences are in Table S1. To realize a uniform quenching effect, we designed the stem to be four G-C pairs. In this way, when the hairpin is unfolded, TMR is fluorescent. Once it is folded, TMR is quenched by the adjacent four guanines. In practice, adding dGs together does not complicate the quenching process but provides stronger quenching effect,36 which is handy for a kinetic data analysis. A linker of 24 thymines between the hairpin stem and the surface is used to minimize the influence of immobilization. Figure 2b shows that the fluorescence intensity of hp-4G-polyT-24 decreases as the NaCl concentration increases, whereas in the control group with similar sequence but no pairing stem, 4G-polyT-24, the fluorescence intensity keeps constant. It indicates that at higher NaCl concentration, more hairpin molecules prefer the folded state. Then, we applied both scanning smFCS (Figure 2c,d) and conventional FCS (Figure S2a) to explore the system. The conventional FCS was performed in free-diffusing solution, and the scanning smFCS was performed on DNA molecules immobilized on the coverslip of a flow cell.37 In both methods, the correlation curves of hp-4G-polyT-24 change along with the NaCl concentration (Figure 2c, Figure S2a), whereas the control group (4G-polyT-24) without paring bases changes little under all NaCl concentrations from 0 mM to 500 mM

⎛ τ⎞ ⎛ τ⎞ G R (τ ) = 1 + α1·exp⎜ − ⎟ + α2·exp⎜ − ⎟ ⎝ τ2 ⎠ ⎝ τ1 ⎠

(5)

The fitting parameters, including amplitudes (α) and characteristic relaxation times (τ) of the two reaction components, are shown in Figure 3c−f. Considering the interference of diffusion, exponential fitting in the conventional FCS needs special care.38 The good agreement within the experimental errors between the two methods verifies each other. It also indicates that the immobilization in the scanning smFCS has not influenced the folding kinetics. More discussions about the effect of linkers and surface are in the Supporting Information. The above two processes can be further identified. For the fast process, α1 increases as the salt concentration increases, but τ1 remains constant on microsecond time scale. In the Supporting Information we show that the contribution from the triplet−singlet transition in this region is removed from the GR(τ) curve by the dividing treatment. We assign this remaining process to the TMR swing motion after the DNA paired stem is formed. The swing motion is the motion between TMR capped on the terminal of the dsDNA and TMR moved away from the capped position. It results in the distance change between TMR and guanines, so does the fluorescence.35 As the salt concentration increases, the kinetics of the TMR swing is not notably affected, whereas α1 increases as a result of increased population of the folded hairpin. 1868

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quantitative agreement with eq 6 with Qfit = 0.389 ± 0.006 (Figure S6a). The less efficient fluorescence quenching in hp4G-polyA than in hp-4G-polyT-24 implies that with the larger and stiffer polyA linker, TMR caps less tightly at the end of hairpin stem. With the known Q and K, we are now at the position to factor out the folding and unfolding reaction rate constants according to the relationship

The slow component corresponds to the folding reaction. As there is one exponential component for the hairpin folding, a two-state folding model should be sufficient to describe the process: kf

U⇌F ku

where U is the unfolded state with a relative fluorescence brightness of 1, F is the folded state with a relative fluorescence brightness of Q, kf and ku represent the folding and unfolding reaction rate constants, respectively. In the two-state model, α is related to both Q and equilibrium constant K by29,39 α=

K (1 − Q )2 (1 + KQ )2

⎧ kf ⎪ K= ku ⎪ ⎨ 1 ⎪ ⎪kf + ku = τ ⎩ 2

(6)

(7)

The ku and kf under different NaCl concentrations are shown in Figure 4b for hp-4G-polyT-24 and in Figure S6b for hp-4GpolyA. When the NaCl concentration is lower than 75 mM, the unfolding reaction of hp-4G-polyT-24 is faster than the folding reaction, so the hairpin prefers unfolded state. As the NaCl concentration increases, the folding rate increases rapidly and the unfolding rate decreases slowly, so the hairpin prefers folded state. Compared with hp-4G-polyT-24, the folding rate of hp-4G-polyA is obviously slower, whereas the unfolding rate of hp-4G-polyA is a little bit faster at low NaCl concentration but gradually approaches that of hp-4G-polyT-24 as NaCl concentration increases (Figure S6b). These results agree with previous work of Bonnet et al.,6 and the rate difference is because the polyA loop is more rigid than polyT loop. The loop rigidity affects the folding appreciably but its effect on unfolding is less prominent. In this work, we presented the idea of using scanning smFCS to explore biophysical or biochemical kinetics. The feasibility of the scanning smFCS was illustrated on a classical system of the hairpin folding. We showed that the scanning smFCS not only bridges the millisecond gap left over by conventional FCS and SMI but also works well in the domains they apply. It can study processes in a broad time range from microseconds to seconds simultaneously. Moreover, it is stable, efficient and easy to work with. We believe that scanning smFCS will present itself a promising technique to study the kinetics of many biological systems, including inter- and intramolecular conformational changes or motions, protein−protein interactions, protein− DNA recognitions, enzymatic reactions, and others.

To determine Q and K simultaneously using eq 6 and the measured fluorescence intensity in Figure 2b, we fit the experimental data of α2 versus the salt concentration iteratively to reach the optimal setting of Qfit = 0.224 ± 0.006. The fitting result is shown in Figure 4a. It is also easy to derive from eq 6

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Figure 4. Two-state model fit and calculated reaction rate constants. (a) The amplitude of scanning smFCS is fitted to the two-state model. (b) Folding and unfolding reaction rate constants change over NaCl concentrations. Error bars denote the s.d. of three independent experiments. In some cases, error bars are smaller than the symbols.

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jpclett.6b00720. Experimental methods, supporting discussions, DNA sequences (Table S1), and supporting figures (Figures S1−S8) (PDF)

that when K = 1/Q, α reaches maximum as αmax = (1 − Q)2/ (4Q). In this way, Q = 0.227 ± 0.015 is directly read out from Figure 3e at αmax without seeking any curve fitting. We note that Q is a crucial parameter in obtaining accurate rate constants from the FCS data, but it is very difficult to determine. To our knowledge, this is the first time that eq 6 is quantitatively verified by an experiment. The good agreement between the experimental results and theoretical prediction not only proves the validity of eq 6 but also confirms the two-state model on the hairpin folding. To further test the scanning smFCS method and verify eq 6, we studied another hairpin hp-4G-polyA, with thymines being replaced by adenines (Figure S5). We found that the folding process of hp-4G-polyA also follows the two-state model, in a

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Present Address †

Postdoctoral Fellow, Department of Biochemistry and Molecular Pharmacology, New York University School of Medicine, New York, NY 10016.

Notes

The authors declare no competing financial interest. 1869

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(20) St.-Pierre, P. R.; Petersen, N. O. Average Density and Size of Microclusters of Epidermal Growth Factor Receptors on A431 Cells. Biochemistry 1992, 31, 2459−2463. (21) Skinner, J. P.; Chen, Y.; Müller, J. D. Position-Sensitive Scanning Fluorescence Correlation Spectroscopy. Biophys. J. 2005, 89, 1288−1301. (22) Petrásě k, Z.; Ries, J.; Schwille, P. Scanning FCS for the Characterization of Protein Dynamics in Live Cells. Methods Enzymol. 2010, 472, 317−343. (23) Xiao, Y.; Buschmann, V.; Weston, K. D. Scanning Fluorescence Correlation Spectroscopy: A Tool for Probing Microsecond Dynamics of Surface-Bound Fluorescent Species. Anal. Chem. 2005, 77, 36−46. (24) Yin, Y.; Yuan, R.; Zhao, X. S. Amplitude of Relaxations in Fluorescence Correlation Spectroscopy for Fluorophores That Diffuse Together. J. Phys. Chem. Lett. 2013, 4, 304−309. (25) Ha, T.; Rasnik, I.; Cheng, W.; Babcock, H. P.; Gauss, G. H.; Lohman, T. M.; Chu, S. Initiation and Re-Initiation of DNA Unwinding by the Escherichia coli Rep Helicase. Nature 2002, 419, 638−641. (26) Krichevsky, O.; Bonnet, G. Fluorescence Correlation Spectroscopy: The Technique and Its Applications. Rep. Prog. Phys. 2002, 65, 251−297. (27) Bikard, D.; Loot, C.; Baharoglu, Z.; Mazel, D. Folded DNA in Action: Hairpin Formation and Biological Functions in Prokaryotes. Microbiol. Mol. Bio. Rev. 2010, 74, 570−588. (28) Grunwell, J. R.; Glass, J. L.; Lacoste, T. D.; Deniz, A. A.; Chemla, D. S.; Schultz, P. G. Monitoring the Conformational Fluctuations of DNA Hairpins Using Single-Pair Fluorescence Resonance Energy Transfer. J. Am. Chem. Soc. 2001, 123, 4295−4303. (29) Jung, J.; Van Orden, A. A Three-State Mechanism for DNA Hairpin Folding Characterized by Multiparameter Fluorescence Fluctuation Spectroscopy. J. Am. Chem. Soc. 2006, 128, 1240−1249. (30) Nayak, R. k.; Peersen, O. B.; Hall, K. B.; Van Orden, A. Millisecond Time-Scale Folding and Unfolding of DNA Hairpins Using Rapid-Mixing Stopped-Flow Kinetics. J. Am. Chem. Soc. 2012, 134, 2453−2456. (31) Narayanan, R.; Zhu, L.; Velmurugu, Y.; Roca, J.; Kuznetsov, S. V.; Prehna, G.; Lapidus, L. J.; Ansari, A. Exploring the Energy Landscape of Nucleic Acid Hairpins Using Laser Temperature-Jump and Microfluidic Mixing. J. Am. Chem. Soc. 2012, 134, 18952−18963. (32) Balkwill, G. D.; Williams, H. E. L.; Searle, M. S. Structure and Folding Dynamics of a DNA Hairpin with a Stabilising d(GNA) Trinucleotide Loop: Influence of Base Pair Mis-Matches and Point Mutations on Conformational Equilibria. Org. Biomol. Chem. 2007, 5, 832−839. (33) Nayak, R. K.; Van Orden, A. Counterion and Polythymidine Loop-Length-Dependent Folding and Thermodynamic Stability of DNA Hairpins Reveal the Unusual Counterion-Dependent Stability of Tetraloop Hairpins. J. Phys. Chem. B 2013, 117, 13956−13966. (34) Piestert, O.; Barsch, H.; Buschmann, V.; Heinlein, T.; Knemeyer, J.-P.; Weston, K. D.; Sauer, M. A Single-Molecule Sensitive DNA Hairpin System Based on Intramolecular Electron Transfer. Nano Lett. 2003, 3, 979−982. (35) Qu, P.; Chen, X. D.; Zhou, X. X.; Li, X.; Zhao, X. S. Fluorescence Quenching of TMR by Guanosine in Oligonucleotides. Sci. China, Ser. B: Chem. 2009, 52, 1653−1659. (36) Qu, P.; Yang, X.; Li, X.; Zhou, X.; Zhao, X. S. Direct Measurement of the Rates and Barriers on Forward and Reverse Diffusions of Intramolecular Collision in Overhang Oligonucleotides. J. Phys. Chem. B 2010, 114, 8235−8243. (37) Kim, S.; Blainey, P. C.; Schroeder, C. M.; Xie, X. S. Multiplexed Single-Molecule Assay for Enzymatic Activity on Flow-Stretched DNA. Nat. Methods 2007, 4, 397−399. (38) In conventional FCS, sometimes the millisecond residues interfered with the fitting process, we then added a third exponential component to accomplish the fitting. The third component did not have physical significance, so we will not discuss it further.

ACKNOWLEDGMENTS We thank NKBRSF (2012CB917304) and NSFC (21233002 and 21521003) for the financial support.

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REFERENCES

(1) Lakowicz, J. R. Principles of Fluorescence Spectroscopy, 3rd ed.; Springer Science+Business Media: New York, 2006. (2) Lu, H. P.; Xun, L.; Xie, X. S. Single-Molecule Enzymatic Dynamics. Science 1998, 282, 1877−1882. (3) Yildiz, A.; Forkey, J. N.; McKinney, S. A.; Ha, T.; Goldman, Y. E.; Selvin, P. R. Myosin V Walks Hand-Over-Hand: Single Fluorophore Imaging with 1.5-nm Localization. Science 2003, 300, 2061−2065. (4) Yu, J.; Xiao, J.; Ren, X.; Lao, K.; Xie, X. S. Probing Gene Expression in Live Cells, One Protein Molecule at a Time. Science 2006, 311, 1600−1603. (5) Roy, R.; Kozlov, A. G.; Lohman, T. M.; Ha, T. SSB Protein Diffusion on Single-Stranded DNA Stimulates RecA Filament Formation. Nature 2009, 461, 1092−1097. (6) Bonnet, G.; Krichevsky, O.; Libchaber, A. Kinetics of Conformational Fluctuations in DNA Hairpin-Loops. Proc. Natl. Acad. Sci. U. S. A. 1998, 95, 8602−8606. (7) Riedel, C.; Gabizon, R.; Wilson, C. A. M.; Hamadani, K.; Tsekouras, K.; Marqusee, S.; Pressé, S.; Bustamante, C. The Heat Released During Catalytic Turnover Enhances the Diffusion of an Enzyme. Nature 2015, 517, 227−230. (8) Henzler-Wildman, K.; Kern, D. Dynamic Personalities of Proteins. Nature 2007, 450, 964−972. (9) Juette, M. F.; Terry, D. S.; Wasserman, M. R.; Altman, R. B.; Zhou, Z.; Zhao, H.; Blanchard, S. C. Single-Molecule Imaging of NonEquilibrium Molecular Ensembles on the Millisecond Timescale. Nat. Methods 2016, 13, 341−344. (10) Santoso, Y.; Kapanidis, A. N. Probing Biomolecular Structures and Dynamics of Single Molecules Using In-gel Alternating-Laser Excitation. Anal. Chem. 2009, 81, 9561−9570. (11) Rogers, J. M.; Polishchuk, A. L.; Guo, L.; Wang, J.; DeGrado, W. F.; Gai, F. Photoinduced Electron Transfer and Fluorophore Motion as a Probe of the Conformational Dynamics of Membrane Proteins: Application to the Influenza A M2 Proton Channel. Langmuir 2011, 27, 3815−3821. (12) Yin, Y.; Wang, P.; Yang, X. X.; Li, X.; He, C.; Zhao, X. S. Panorama of DNA Hairpin Folding Observed Via DiffusionDecelerated Fluorescence Correlation Spectroscopy. Chem. Commun. 2012, 48, 7413−7415. (13) Jung, J.; Ihly, R.; Scott, E.; Yu, M.; Van Orden, A. Probing the Complete Folding Trajectory of a DNA Hairpin Using Dual Beam Fluorescence Fluctuation Spectroscopy. J. Phys. Chem. B 2008, 112, 127−133. (14) Yin, Y.; Yang, L.; Zheng, G.; Gu, C.; Yi, C.; He, C.; Gao, Y. Q.; Zhao, X. S. Dynamics of Spontaneous Flipping of a Mismatched Base in DNA Duplex. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 8043−8048. (15) Wang, Q.; Goldsmith, R. H.; Jiang, Y.; Bockenhauer, S. D.; Moerner, W. E. Probing Single Biomolecules in Solution Using the Anti-Brownian Electrokinetic (ABEL) Trap. Acc. Chem. Res. 2012, 45, 1955−1964. (16) Wennmalm, S.; Edman, L.; Rigler, R. Conformational Fluctuations in Single DNA Molecules. Proc. Natl. Acad. Sci. U. S. A. 1997, 94, 10641−10646. (17) Watkins, L. P.; Chang, H.; Yang, H. Quantitative SingleMolecule Conformational Distributions: A Case Study with Poly-(Lproline). J. Phys. Chem. A 2006, 110, 5191−5203. (18) Weissman, M.; Schindler, H.; Feher, G. Determination of Molecular Weights by Fluctuation Spectroscopy: Application to DNA. Proc. Natl. Acad. Sci. U. S. A. 1976, 73, 2776−2780. (19) Magde, D.; Webb, W. W.; Elson, E. L. Fluorescence Correlation Spectroscopy. III. Uniform Translation and Laminar Flow. Biopolymers 1978, 17, 361−376. 1870

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The Journal of Physical Chemistry Letters (39) Zhu, R.; Li, X.; Zhao, X. S.; Yu, A. Photophysical Properties of Atto655 Dye in the Presence of Guanosine and Tryptophan in Aqueous Solution. J. Phys. Chem. B 2011, 115, 5001−5007.

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