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A practical guide to single-molecule FRET

Rahul Roy^1, 2, Sungchul Hohng³ & Taekjip Ha^1, 2, 4

¹ Department of Physics, University of Illinois at Urbana-Champaign, 1110 West Green Street, Urbana, Illinois 61801, USA

² Center for Biophysics and Computational Biology, University of Illinois at Urbana-Champaign, 1110 West Green Street, Urbana, Illinois 61801, USA.

³ Department of Physics and Astronomy, Seoul National University, San 56-1 Sillim 9-dong, Gwanak-gu, Seoul 151-747, Korea.

⁴ Howard Hughes Medical Institute, 1110 West Green Street, Urbana, Illinois 61801, USA.

Figure 1. smFRET description. (a) FRET efficiency, E, as a function of inter-dye distance (R) for a R0 = 50 Å. Donor dye directly excited with incident laser either fluoresces or transfers energy to acceptor dye, depending upon its proximity. At R = R0, E = 0.5, but at smaller distances, it is >0.5 and vice versa, according to the function shown by the blue line. Notice the linearity of the E values adjacent to R0. (b) Example of a two-color smFRET data. Data are acquired in the form of intensities of donor and acceptor (top), from which apparent FRET efficiency (bottom) is calculated. A mutant hairpin ribozyme93 that carries the donor and acceptor on different arms of the same molecule undergoes transitions between three FRET states (E1, E2 and E3). The anti-correlated nature of the donor and acceptor signal indicates that these intensity changes are due to energy transfer. Dye molecules also show transitions to dark states—for example, acceptor intensity transiently drops to zero (9 s) or completely photobleaches (18.5 s). Full Figure and legend (67K)

Figure 2. Single-molecule FRET schemes. Full Figure and legend (29K)

Single-molecule fluorescence dyes. An ideal fluorophore for single-molecule studies must be bright (extinction coefficient, , > 50,000 M^{- 1} cm^{- 1}; quantum yield, QY, > 0.1), photostable with minimal photophysical or chemical and aggregation effects, small and water-soluble with sufficient forms of bio-conjugation chemistries. Additionally, an excellent smFRET pair should have (i) large spectral separation between donor and acceptor emissions and (ii) similar quantum yields and detection efficiencies. Although fluorescent proteins have been used for smFRET studies²³, low photostability and photoinduced blinking have hindered further applications. Semiconductor quantum dots (QDs) have also been used as a smFRET donor²⁴ with their blinking chemically suppressed²⁵, but the large size (>20 nm diameter for commercial QDs) and the lack of a monovalent conjugation scheme limit their use. Consequently, the most popular single-molecule fluorophores are small (<1 nm) organic dyes²⁶. We compared three FRET pairs with absorbance from 500–700 nm from different vendors (Cyanine, Alexa and Atto dyes; Table 1). Though cyanine dyes (Cy3 and Cy5; donor and acceptor, respectively) have long been the favorites, their counterparts seem to have comparable relevant properties. None of the bluer dyes—for example, those that can be excited at 488 nm—were as photostable as the original cyanine dyes. But a Cy3 replacement for custom RNA synthesis, Dy547, is even more photostable than Cy3 (S. Myong; personal communication). Also, tetramethylrhodamine is a viable alternative to Cy3 and has an almost identical spectrum but with a lower extinction coefficient. However, it has the tendency to change its intensity between three different levels spontaneously (T.H.; unpublished observations). Near-infrared dyes such as Cy5.5 and Cy7 also serve as efficient single-molecule dyes and can be used in multicolor schemes (discussed below).

Table 1. Comparison of single-molecule FRET dyes Full Table

Enhancing photostability. Molecular oxygen is an efficient quencher of a dye's unfavorable triplet state, but is also a source of a highly reactive oxygen species that ultimately causes photobleaching²⁷. Although oxygen removal reduces photobleaching, it prolongs the residence time in the triplet dark state²⁸ causing millisecond or longer fluorescence intermittency or an early onset of signal saturation. A vitamin E analog named Trolox (2 mM; 100 stock prepared in dimethyl sulfoxide; pH adjustment and filtration required) is an excellent triplet-state quencher, which suppresses blinking and stimulates long-lasting emission of the popular cyanine dyes in combination with an enzymatic oxygen-scavenging system²⁸. Trolox should also provide a major improvement in time resolution as it delays the onset of emission saturation with increasing laser intensity. The reductant -mercaptoethanol (ME; 142 mM) is a less efficient triplet-state quencher, and it induces long-lived dark states of Cy5 (ref. 28), a side effect that has been exploited for super-resolution photoswitching imaging applications²⁹. A plethora of other triplet-state quenchers or antifade agents exists that may prove beneficial for non-cyanine dyes³⁰.

The most popular enzymatic oxygen scavenging system³¹ is a mix of glucose oxidase (165 U/ml), catalase (2,170 U/ml) and -D-glucose (0.4% wt/wt) (or 0.8% wt/wt dextrose monohydrate). Glucose oxidase must be added just before imaging, and the sample must be kept isolated from air such that the solution pH does not drop substantially as a result of gluconic acid production, a byproduct of the reaction. Other options include the use of the protocatechuic acid and protocatechuate-3,4-dioxygenase combination³², which provides 40% improvement over the glucose oxidation system (N. Walter, personal communication). If highly purified forms of these enzymes are not commercially available, care must be taken to ensure the absence of contaminating activity, especially that of RNases.

Table 2. Conjugation strategies for dyes or biotin Full Table

Total internal reflection spectroscopy. In TIR microscopy^{40, 41}, one creates an evanescent field of excitation light that extends only 100–200 nm from the surface to which the sample is bound, which greatly reduces background fluorescence (Fig. 3a,b). A solid-state laser at 532 nm (50 mW) is suitable for the FRET pairs discussed here, and a HeNe laser at 633 nm (30 mW) or diode lasers of similar wavelengths can be added for checking the presence of the acceptor. The laser intensity is attenuated using a half-waveplate and a polarizing beam-splitting cube (or with neutral density filters). By exciting a large area (0.05 mm² in size) and using camera-based detection, hundreds of molecules are imaged in parallel. A laser table with air-floated legs is typically used, but a 25 mm thick breadboard with uniformly spaced threaded holes mounted on a regular laboratory bench or desk is stable enough for TIR smFRET. There are two types of TIR, prism-type (PTIR) and objective-type (OTIR) (Fig. 3a,b).

Figure 3. Schematic for smFRET spectroscopy. (a) TIR excitation and single FRET pair emission detection. Tethered single molecules are either excited by PTIR or OTIR. Fluorescence is collected using the objective, and the slit generates a final imaging area that is half of the CCD imaging area. The collimated image is split into the donor and acceptor emissions and imaged side by side on the CCD camera (camera image in inset). (b) TIR excitation schemes (boxed region in a). In PTIR, a laser beam focused at a large incident angle (c > 68°) on the prism placed on the top of the sample creates an evanescent field at the quartz-water interface on the slide. Alternately in OTIR excitation, the focused laser beam strikes at the periphery of the objective back focal plane causing TIR at the glass-water interface on the coverslip close to the objective. (c) Emission detection for three-color scheme. Image is framed using a slit such that final image size covers one-third of the CCD chip. A set of dichroics allow separation of the individual emissions from the three fluorophores and imaging them simultaneously. /2, half waveplate; PBS, polarizing beam splitter; L1–6, lenses; EF, evanescent field; DM1–5, dichroic mirrors; BE, beam expander; LP, long pass filter; C, incident angle. Full Figure and legend (78K)

In PTIR, an inverted microscope is adapted to hold a fused silica prism on top of the sample channel, and the fluorescence is collected from the objective below^{40, 41} (Fig. 3a). After the incident laser beam is focused using a long focal length lens, it enters the prism, passes through refractive index–matching oil and is internally reflected at the quartz-water interface (Fig. 3a,b and Supplementary Methods online). The fluorescence signal is collected using a long working distance water immersion objective (60, 1.2 numerical aperture (NA)). PTIR is necessary for fluid-injection experiments because the imaging surface, made of a 1-mm thick slide, does not bow upon pressure change during flow. Expensive (but recyclable) quartz slides are needed to minimize fluorescence background, and the prism needs to be reassembled every time a new sample is loaded. The need to readjust the illumination path is minimized if the prism is reproducibly placed in the same location relative to the microscope body.

OTIR relies on using a high-NA oil objective to create an evanescent field. Focusing the beam at the back focal plane generates a parallel beam exiting the objective, and then translating it to the periphery of the objective produces TIR at the glass-water interface^{40, 41} (Fig. 3a,b). Fluorescence from the molecules tethered to the coverslip surface is collected using the same objective. OTIR has higher photon-collection efficiency, frees up the space above the sample for additional sample control and has commercial options. Aided largely by the use of a low-fluorescence objective, UPlanSApo (100, 1.4 NA; Olympus), we can acquire smFRET data of Cy3-Cy5 pair with signal-to-noise ratio and imaging area comparable to that of prism-type TIR (T.H.; unpublished results).

Emission detection. TIR microscopy has gained popularity partly spurred by the new wave of highly sensitive and fast frame transfer electron-multiplying charge-coupled device (EM-CCD) cameras^{3, 42}. smFRET setups usually use EM-CCDs that have high quantum efficiency (85–95%) in the 450–700 nm range, low effective readout noise (<1 electron r.m.s.) even at the fastest readout speed (10 MHz), fast vertical shift speeds (1 s/row; to achieve faster frame rates) and a low multiplication noise. Using a 512 512 pixel EM-CCD (referred to hereafter as CCD), we can acquire data at 33 Hz (at full frame) to 125 Hz (with 2 2 binning).

The scattered light of the excitation laser is rejected from the fluorescence collected with the objective using a long pass filter. A vertical slit is introduced at the imaging plane just outside the microscope side port to limit the image area such that the final image incident on the CCD is half the size of the CCD chip (Fig. 3a). Donor and acceptor fluorescence emission is then split using a dichroic mirror. By adjusting an offset with the dichroic and an additional mirror, both donor and acceptor emission can be imaged side by side on the CCD camera (Fig. 3a). This optical layout maps the imaging area of 75 m 37 m on the CCD chip. A straightforward extension using a narrower slit corresponding to one-third of the CCD area enables three-color FRET detection (Fig. 3c). Without additional bandpass filters, there is a sizeable cross-talk between the detection channels (for example, 40% of Cy5 emission leaks into the Cy5.5 channel if Cy5.5 is used as a second acceptor), but careful calibration and correction can recover the original intensities⁴³. Our recent test indicates that Cy7 is a promising fluorophore that can replace Cy5.5 in the original single-molecule three-color FRET scheme; photostability and brightness of Cy7 were comparable to those of Cy5.5, whereas Cy7's emission can be better discriminated from Cy5 emission. A relatively low detection efficiency of CCD camera at the Cy7 emission wavelength is currently an issue, but this should not be a problem for confocal measurements because the silicon avalanche photodiode maintains high quantum yield of detection at these wavelengths. Reduced transmission in this spectral region can be alleviated to some degree with infrared-optimized objectives and optics.

Figure 4. Surface immobilization strategies for smFRET experiments. (a) Biotinylated BSA proteins bound nonspecifically to the surface tether biotinylated molecules with the aid of multivalent avidin proteins (for example, neutravidin). (b) Mixture of biotin-PEG and PEG is covalently attached to amino-silanized slide surface. Biotins on the PEG can bind DNA (or protein) engineered with a biotin moiety with the help of a sandwiched neutravidin protein. PEG coating prevents the nonspecific binding. (c) PEG-coated surfaces can be engineered to carry Ni2+ or Cu2+ chelated on iminodiacetic acid (or nitrilotriacetic acid) groups that bind efficiently to 6His-tagged proteins while retaining functional activity. (d) Using dimyristoyl phospatidylcholine (DMPC) at room temperature, vesicles selectively permeable to small molecules can be used to trap biomolecules. A fraction of biotinylated lipid allows specific tethering of the vesicles to the biotin-PEG surface. Full Figure and legend (85K)

The sample chamber. An air-tight sample chamber is created by sandwiching double-sided tape or parafilm between a precleaned slide and a coverslip (Supplementary Protocol) and by applying epoxy as necessary. Simple pipetting or pumping through two holes pre-drilled in the slide allows exchange of solution without drying (Fig. 5). The assembled chamber is checked for nonspecific binding before application of streptavidin by imaging the surface in the presence of 1 nM labeled DNA and/or protein. If the nonspecifically bound fluorescent spot density is <10% of specifically tethered molecules (typically, specific attachment with 'good' density provides 0.1–0.2 spots/m²), the slide preparation is deemed acceptable. After streptavidin, biotinlyated biomolecules are added in low concentrations (20–100 pM in buffer containing 0.1 mg/ml BSA to reduce loss of molecules to other surfaces) to achieve immobilization at the desired single-molecule density. Higher density increases the chance of overlapping neighboring molecules. Movies of such tethered molecules are acquired and saved on the hard disk directly using a custom program written in Visual C++.

Figure 5. Sample chamber. (a) A sample chamber is made by sandwiching a microscope slide and a coverslip with double-sided tape and by sealing with epoxy. The holes on the slide are used as the inlet and outlet for solution exchange. (b) A syringe is connected to the chamber through tubing, and a pipette tip that contains a solution is snugly plugged into an inlet hole. When the syringe is pulled, the solution is introduced into a chamber. Reproduced from ref. 7 with permission from Cold Spring Harbor Laboratory Press. Full Figure and legend (36K)

Data processing algorithms. To convert the movie files into single-molecule time trajectories, we first identify isolated single-molecule peaks from a composite image of donor and acceptor channels (average of first 10 frames) built using the overlap mapping generated (see above). Donor and acceptor spots that colocalize are selected for final analysis to eliminate partially labeled molecules. Next the local background is subtracted from the peaks and intensities from 7 7 pixels (depending on overall magnification) surrounding the peak (corresponding to 1 m² area) are integrated to recover donor and acceptor intensities for each single molecule for every frame. In experiments for which only transient presence of the donor-labeled molecule is anticipated⁵⁴, the first step can be carried out for each set of ten successive frames.

FRET efficiency. After correcting for the cross-talk and background (determined from intensity traces after dye photobleaching) in both channels, apparent FRET efficiency is calculated as E_app = I_A / (I_A + I_D), where I_A and I_D represent acceptor and donor intensities, respectively. E_app provides only an approximate indicator of the inter-dye distance because of uncertainty in the orientation factor ² between the two fluorophores and the required instrumental corrections. As a rule of thumb, if fluorescence anisotropy, r, of both fluorophores is less than 0.2, ² is close to 2/3 (refs. 34,55). We found that in general r > 0.2 for popular smFRET fluorophores conjugated to nucleic acids or proteins, so care must be taken in extracting the absolute distance information. Nevertheless, in every case we tested, apparent FRET was a monotonic function of distance. To determine actual FRET efficiency, one has to determine the correction factor, , which accounts for the differences in quantum yield and detection efficiency between the donor and the acceptor. is calculated as the ratio of change in the acceptor intensity, I_A to change in the donor intensity, I_D upon acceptor photobleaching ( = I_A / I_D)^{21, 56}. Corrected FRET efficiency is then calculated using the expression,

Proteins can change the photophysical properties of a fluorophore. For example, we and others have observed that the fluorescence of DNA-conjugated Cy3 increases when a protein binds nearby^{57, 58}. This effect changes the FRET efficiency through a change in . Similarly, photoinduced electron transfer between DNA bases and dye can quench dyes, and this phenomenon can be influenced by protein binding in the vicinity^{59, 60, 61}.

Although the analysis of single-molecule data is system-dependent, some commonly used tools are worth mentioning. Information on equilibrium properties is best gleaned through the smFRET histogram generated by averaging each molecule's FRET efficiency over the first 3–10 data points. We typically obtain 10–20 short (5 s) movies of different areas of the sample for an unbiased overview of population distribution. Time trajectories of individual molecules (from long movies) in equilibrium convey valuable kinetics information of the system via dwell times in each conformational state. For example, for a two-state system undergoing stochastic transitions, the transition rates are determined from exponential decay fits to the dwell-time distribution of each state⁶³ or by performing the autocorrelation analysis combined with equilibrium determination if the transitions are too fast for reliable dwell-time determination⁶⁴. For more complicated transitions between distinctly identifiable multiple states, a hidden Markov model analysis can be used for an unbiased estimate of the number of FRET states populated to estimate the rates of interconversion among them and to determine the most likely time evolution of each state⁶⁵ (an executable version of hidden Markov model–based FRET time trajectory analysis program, HaMMy is available for download at http://bio.physics.uiuc.edu/HaMMy.html). A transition density plot of initial to final FRET values for each transition obtained from hidden Markov model (or other) analysis provides a visually attractive and objective compilation of the data^{57, 65, 66, 67}. Other information theory based approaches are also available for confocal single-molecule data obtained photon by photon^{68, 69, 70}. The advantages of single-molecule experiments become more obvious in nonequilibrium conditions where one can follow, for example, the reaction pathway of a single enzyme undergoing a series of transitions to achieve its catalytic cycle. A powerful, visual method of presenting the non-equilibrium smFRET data of many molecules has been developed for ribosome studies⁷¹.

It is crucial to point out some of the limitations of the technique before one designs a first smFRET experiment. (i) smFRET requires attachment of at least two extrinsic dyes to the molecule(s) of interest because (semi-) intrinsic chromophores such as 2-aminopurine and tryptophan are not sufficiently bright or photostable for single-molecule measurement. In some cases site-specific labeling is not trivial, especially for large RNA molecules^{72, 73} and many proteins^{74, 75}. Note that smFRET is relatively insensitive to incomplete labeling. If the donor is missing, the molecule is simply not observed; if the acceptor is missing, this donor-only species shows up as a zero-FRET population. (ii) As single-molecule detection is achieved via spatial separation, weakly interacting fluorescent species are difficult to study (but not always; see above). (iii) FRET is insensitive to distance changes outside the 2–8 nm inter-dye distance range for R₀ = 5 nm. However, distance changes as small as 0.3 nm can be detected from single molecules between 0.6R₀ and 1.5R₀, where FRET is almost a linear function of R. (iv) To achieve adequate signal-to-noise ratio, 100 total photons need to be detected. Considering that more than 10⁵ photons can be collected from single dye molecules before photobleaching, more than 10³ data points can be obtained. (v) Time resolution is limited by the frame rate of the CCD camera (in best case = 1 ms). (vi) Absolute distance estimation is challenging because of the dependence of the fluorescence properties and energy transfer on the environment and orientation of the dyes. So, smFRET has been used mostly in situations where precise distance information is not vital. Nevertheless, reasonable estimates of orientation factors can be made⁷⁶, and control experiments with each dye can provide good approximations of the inter-dye distances^{77, 78}. Concerns about these issues could be further alleviated by using a redundant number of distance constraints in triangulation studies^{79, 80}.

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