FRET - Förster Resonance Energy Transfer || Single-Molecule Applications

8Single-Molecule ApplicationsThomas Pons

8.1Introduction

Recent years have witnessed major progress in fluorescence microscopy instru-mentation, raising its detection sensitivity to the single-molecule level. Observationof single molecules has since brought a wealth of information and allowed a betterunderstanding of many physical, chemical, and biological processes [1–6]. Single-molecule fluorescence has become in particular a powerful tool to study bio-molecular functions [7–12]. Indeed, these functions most often involve conforma-tional changes and/or multimolecular association/dissociation. This implies that, inthe absence of an external synchronization, biomolecules in solution fluctuatebetween different states independently of each other. Whereas ensemble experi-ments provide only average measurements over all these different states, single-molecule measurements can reveal both heterogeneity in the population (in theequilibrium distribution of states) and dynamics (i.e., sequence of transitions,frequencies, rates, etc.).Single-molecule F€orster (or Fluorescence) resonance energy transfer is certainly

one of the most fertile single-molecule fluorescence techniques [13–19]. The vastmajority of smFRETstudies involve labeling of the target biomolecules with a donorand an acceptor fluorophore at specific sites. The FRET distance dependencetranslates changes in donor–acceptor separation distance into measurable photo-physical parameters, such as donor/acceptor emission ratios, lifetimes, and anisot-ropy. This in turn provides a tool to follow conformational changes in a singlebiomolecule or association/dissociation dynamics in a single complex of interactingpartners.Observation of single-molecule fluorescence signals raises several difficulties,

including weak fluorescence signals and the need to isolate the signal from onemolecule from the large background of other molecules, without perturbing thefunctional integrity of the biomolecule. These challenges can be overcome usingmainly two categories of microscopy modalities. The first modality relies onimmobilization of molecules on a substrate. A time trace of fluorescence signals

FRET – Förster Resonance Energy Transfer: From Theory to Applications, First Edition.Edited by Igor Medintz and Niko Hildebrandt.� 2014 Wiley-VCH Verlag GmbH & Co. KGaA. Published 2014 by Wiley-VCH Verlag GmbH & Co. KGaA.

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can then be recorded, giving access to the full dynamics ofmolecular conformationalchanges. The second modality uses molecules dissolved in solution and freelydiffusing in and out of a confocal detection volume, resulting in the rapid acquisitionof series of short fluorescence bursts from many individual molecules. Analysis ofthese bursts reveals sample heterogeneity and allows identification of subpopula-tions and their repartition under equilibrium conditions. For each modality, we willbriefly present the corresponding experimental techniques and analysis methods,discuss their capabilities, and present a few examples to illustrate their potentialapplications, with a strong focus on biophysical studies. Finally, we will presentother single-molecule FRET schemes that involve multiple interacting FRETpartners.

8.2Single-Molecule FRET of Immobilized Molecules

This section presents an overview of experimental techniques used to immobilizebiomolecules on substrates, with the standard data analysis methods and someillustrative examples of applications. An advanced trace analysis technique is finallypresented.

8.2.1Experimental Setup

8.2.1.1 Molecule ImmobilizationProtocols used for molecule immobilizationmust be carefully designed to attach thebiomolecule without interfering with its functionality and to avoid its nonspecificinteractions with the substrate. One of the most common methods used forbiomolecule immobilization is the biotinylation of the biomolecule and its subse-quent attachment to surface-bound streptavidin [16]. This provides a highly specificinteraction with a high affinity and allows subsequent washing of the surroundingsolution without risking the detachment of the biomolecule of interest. However,extreme care must be taken to minimize potential interactions between thebiomolecule and the rest of the substrate surface. Single DNA and RNA studiesare usually performed using substrates coated with biotinylated BSA and thenstreptavidin [20,21]. Several studies have indeed verified that the conformation ofoligonucleotides is not perturbed when immobilized on these surfaces [20,22,23],probably thanks to the electrostatic repulsion between the negatively chargedoligonucleotides and the negatively charged glass, BSA, and streptavidin at neutralpH.In contrast, single proteins tend to present much more pronounced nonspecific

adsorption on these surfaces and require better passivated substrates. Severalmethods have been proposed to reduce these undesired interactions. Most ofthem require the glass slide to be activated with aminosilanes first, followed bythe covalent coupling of molecules to form a “furtive” coating, with a few biotin

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groups for further specific biomolecule attachment. Covalently surface-linked andcross-linked BSA layers present a more uniform and robust surface coveragecompared to their adsorbed counterparts, which result in a lower level of nonspecificprotein adsorption [24]. Dense layers of polyethylene glycol are another popularcoating and can indeed strongly reduce nonspecific adsorption when sufficientlylong PEG chains are used [25]. These flexible PEG brushes have, however, beenshown to intermingle with attached proteins and modify their conformation after acycle of denaturation and refolding [26]. In comparison, cross-linked star-shapedPEG coatings seem to reduce these interactions since individual immobilizedproteins refolded in their initial conformation after successive exposition to dena-turation and refolding buffers [26,27].Finally, an interesting alternative to direct attachment of biomolecules to the

substrate is their confinement in lipidic nanovesicles [28]. The biomolecules areencapsulated within large (typically 100 nm) unilamellar vesicles containing a fewbiotinylated lipids, which allow subsequent immobilization of the vesicles on avidin-functionalized supported lipid bilayers [28–31]. This method is attractive since thebiomolecule remains in solution but is confined in a volume smaller than thediffraction limit of the microscope, and is therefore available for prolongedobservation. The lipid vesicle is impermeable to the biomolecule of interest butmay be made permeable to other smaller molecules by incorporating small pores inits membrane. This allows the controlled modification of the chemical environmentof the biomolecule (ions, nucleotides, etc.) from the outside of the nanovesiclecontainer [32–34]. Detailed sample preparation protocols are available in Ref. [33]and references therein. Irrespective of the immobilization technique used, appro-priate control experiments must be performed to ensure that the biomolecules areindeed not perturbed. These may include testing different immobilization tech-niques, measuring their enzymatic activity, checking for dye anisotropy, and so on(see Section 8.2.3).

8.2.1.2 Fluorophore PhotostabilityThe choice of fluorescent labels must be optimized to provide strong and stablesignals. The fluorophores should possess high extinction coefficients, fluorescencequantum yields, and photostability to allow long observation times and high signal-to-noise ratios, and present limited photophysical effects (e.g., transient blinking).The most common single-molecule fluorophore pairs include Cy3–Cy5 and theirAtto and AlexaFluor equivalents. The excitation rate of dyes in single-moleculeexperiments is much higher compared to ensemble measurements, leading tomuch faster photobleaching. Several reactants should therefore be added to thebuffer solution to enhance the dye photostability. Molecular oxygen should beremoved since it accelerates photobleaching through the formation of radicalspecies. This is usually obtained using an enzymatic oxygen scavenger systemcomposed of glucose oxidase, catalase, and b-D-glucose [35]. Other antioxidants mayalso be used, such as propyl gallate, ascorbic acid (vitamin C) [36], and cysteamine[37]. Oxygen is, however, also an efficient quencher of the triplet state, and removingit increases the lifetime of this dark state. Other triplet state quenchers thus need to

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be added in the buffer solution, such as Trolox [37], an analogue of vitamin E, ormercaptoethylamine [36]. The nature and optimal concentration of oxygen andradical scavengers and triplet state quenchers may depend on the type of dye used[36]. Alternatively, oxygen removal may be coupled with the addition of an appro-priate combination of reducing and oxidizing agents, such as ascorbic acid andmethylviologen [38].

8.2.1.3 Optical SetupThe solution above the surface is easily washed from residual fluorophore-labeledmolecules; however, fluorescence from the immobilized smFRETmolecules muststill be isolated from out-of-focus background to improve the signal-to-noise ratio.This can be achieved by confocal microscopy or most frequently by total internalreflection fluorescence (TIRF) microscopy [9,39,40]. Confocal detection consists infocalizing a laser beam through an objective using a pair of excitation and detectionpinholes to eliminate fluorescence photons from outside a three-dimensional,diffraction-limited (<1 mm3) confocal volume. The fluorescence signals frommolecules inside this volume are detected using photomultipliers or avalanchephotodiodes, allowing high acquisition speed. However, these detectors only acquiredata from onemolecule at a time, andmeasuring a statistically significant number ofsingle molecules becomes very time consuming. In contrast, TIRFmicroscopy useswide-field illumination with an excitation beam that hits the substrate with an anglelarger than the total internal reflection angle [40]. No light thus propagates into themedium above but the excitation light is confined to a thin (<200 nm) evanescentlayer above the surface, and the intensity decays exponentially from the surface.Under these conditions, excitation is confined to fluorophores on or immediatelyabove the substrate, and eliminates background from the solution above. In practice,this may be realized by focusing a laser beam on the edge of a high-NA objectiveback pupil to create a parallel beam tilted with respect to the objective optical axis. Inthis case, fluorescence photons are collected through the same objective. Alterna-tively, the laser may be focused on a prism placed on top of a thick quartz slide tocreate the evanescent wave (see Figure 8.1), and a high NA objective is placed belowthe sample to collect fluorescence photons. Fluorescence light is then split intodonor and acceptor channels using dichroic and bandpass filters to form twoseparate images on a high-sensitivity, low-noise cooled EM-CCD camera. Thisallows parallel imaging of typically up to a few tens of single FRET pairs.

8.2.2Data Analysis

The first step in the analysis of single-molecule FRET data obtained by wide-fieldTIRF microscopy is the isolation of pixels and group of pixels containing singledonor–acceptor pair signals. This is usually performed by selecting pixels above apredefined threshold, averaging a small area (e.g., 5� 5 or 7� 7 pixels) around thecenter pixel to integrate the point spread function of the microscope, and removingan average background value corresponding to neighboring “empty” pixels.


Molecules with only one active fluorophore present donor-only or acceptor-onlyfluorescence signals and may then be discarded. The time traces obtained fromsignal intensities in the donor and acceptor channels, SD and SA, and their ratio,Eapp¼SA/(SAþSD), qualitatively reflect the evolution of the separation distancebetween the two fluorophores with time. Low Eapp values correspond to long

Figure 8.1 (a) SmFRET setup for theobservation of molecules immobilized on asubstrate, using prism-based (i) or objective-based (ii) TIRF microscopy. The donor andacceptor fluorescence images are separatedinto two halves. (b) Example of smFRET donor

and acceptor and FRET time traces showingacceptor blinking and photobleaching eventsand three distinct FRET states. (Reproducedwith permission from Ref. [16]. Copyright 2008,Macmillan Publishers Ltd.)


separation distances, while high Eapp values correspond to shorter distances. Thereare, however, several factors that should be taken into account for a more quantita-tive analysis and calculation of true FRET efficiencies E, including direct acceptorexcitation, spectral cross-talks of the donor emission into the acceptor channel andvice versa, and differences in fluorescence quantum yields and collection efficien-cies between the two fluorophores. The signals in the donor and acceptor channelsare related to the real molecular fluorescence intensities IA and ID and to thecollection efficiencies through

SD ¼ RDDID þ RADIA þ BA; ð8:1ÞSA ¼ RAAIA þ RDAID þ BD; ð8:2Þ

where Rij describes the instrument response and collection efficiency of signal i intochannel j (i, j¼D, A) and BA,D represents the background signal from the detector(dark signal) and from spurious photons. The Rij elements may be determined bycareful calibration using samples composed of donor-only and acceptor-only singlefluorophores or fluorescent beads to correctly evaluate the corresponding ID and IAintensities. Due to the spectral shape of organic fluorophores, usually only the donorsignal leaks into the acceptor channel, and the signals may be corrected by

ID ¼ aDSD � BD; ð8:3ÞIA ¼ aASA � BA � bSD; ð8:4Þ

where a and b constants take into account effects of channel cross-talks anddetection efficiencies. Fluorescence intensities of the donor and acceptor moleculesdepend on their respective absorption cross sections, fluorescence quantum yield,and FRET efficiency as follows:

ID ¼ sDIexcð1� EÞWD; ð8:5ÞIA ¼ ðsDE þ sAÞIexcWA; ð8:6Þ

where sD(A) and WD(A) are the donor (acceptor) excitation cross section at the laserexcitation wavelength and fluorescence quantum yield, respectively, Iexc is the laserexcitation intensity, and E is the FRET efficiency. Assuming that the acceptor directexcitation is negligible (sDE � sA) or correctly accounted for, the FRET efficiencycan then be evaluated as

E ¼ IA=ðIA þ cIDÞ; ð8:7Þ

c ¼ WA

WD: ð8:8Þ

However, this assumes that c is identical for all individual FRET pairs. This is notnecessarily true because of inhomogeneity in the fluorophore environment due tothe substrate or the macromolecule conformation. However, the values of thec correction factor and the FRET efficiency may often be determined for eachindividual FRET pair. Indeed, the limited donor and acceptor photostability finallyleads to permanent photobleaching of both emitters. In the case of the common


Cy3–Cy5 pair, the Cy5 acceptor often photobleaches before the Cy3 donor does. Thisleads to an instantaneous suppression of FRET processes and a correspondingrecovery of donor emission. The amplitude of the donor recovery is directly relatedto the FRET efficiency before photobleaching since E¼ (ID0� ID)/ID0, where ID0 isthe final donor intensity after photobleaching, in the absence of any acceptor. Inaddition, the c factor corresponds to the ratio of the intensity changes before andafter photobleaching DID and DIA: c¼DIA/DID. It can therefore be useful tocompare the c values obtained for each individual molecule to the value obtainedby average measurements.Single dyes often display complex photophysics that need to be taken into account

for a correct single-molecule FRET interpretation. For example, organic dyes tendto “blink” and present short dark periods attributed to a triplet state (Figure 8.1b)[41–43]. Moreover, additional donor dye–acceptor dye interactions may take place atshort separation distances [44]. These effects should not be confused with abruptchanges of FRET efficiencies. A good safeguard against incorrect interpretation ofsmFRET data is to look at the weighted sum of the fluorescence signals c IDþ IA,which should remain constant when FRET is the sole source of fluorescencefluctuations.Analysis of single-molecule FRET trajectories usually starts with identifying the

different FRET states presented by the observed biomolecules. This is most oftenperformed using simple thresholding, that is, defining a specific FRET efficiencyrange for each state (e.g., 0<E< 0.2 for state s1, 0.35<E< 0. 5 for state s2, and soon; see Figure 8.1). The first available information is the average distribution ofFRET values (often presented as a histogram) or of states (e.g., at any given timemolecules have a P1¼ 50% probability of being in state s1, P2¼ 20% probability ofbeing in state s2, and so on). The free energy Gi of each conformation may then besimply evaluated using Gi¼�kT ln(Pi) [45]. The second important informationavailable is the sequence of conformational changes, for example, determiningwhether transitions always occur from s1 to s2, and then to s3, or directly from s1 tos3, and the frequency of the different transitions. Finally, another importantparameter is the distribution of dwell time of each state, as this can be directlyrelated to the corresponding transition rates. In particular, for states involved insingle-rate kinetics, the distribution of dwell times t may be fitted with a mono-exponential decay, exp(�kt), where k is the transition rate out of the state. In thefollowing section, we will present a few typical examples to illustrate the potentialand limitations of smFRET on immobilized molecules.

8.2.3Applications

Since its demonstration using near-field scanning microscopy [46] in 1996, soonfollowed by its application to confocal microscopy [41,47], single-pair FRET mea-surements have been applied to a wide range of immobilized molecules, includingoligonucleotides and proteins, to study various problems such as folding kineticsand bimolecular interactions.


The hairpin ribozyme has been extensively studied by smFRET and provides agood illustration of what can be achieved using smFRETon immobilized molecules:identifying subpopulations, measuring transition rates between these subpopula-tions, and the sequence of events occurring during a particular reaction. Thisribozyme is an RNA enzyme capable of cleaving a specific RNA substrate into twoproducts. The proposed reaction pathway starts with binding of the substrate RNA tothe ribozyme. The hairpin ribozyme then consists in a four-arm DNA junctioncontaining two internal loops. The ribozyme fluctuates between an extendedunfolded conformation, in which the loops are far apart, and an active foldedconformation, in which the two internal loops are interacting and close to each other.In the folded state, the cleavage reaction occurs and the resulting products finallydissociate from the ribozyme [48]. These conformational fluctuations may thus beobserved by labeling one loop with a donor fluorophore and the other loop with anacceptor, and measuring changes in the FRET interactions on single molecules. In2002, smFRET experiments were performed on a minimal form of the hairpin.Several FRETstates were identified with high FRETvalues corresponding to “folded”conformations and lower FRET values to “unfolded” conformations [49]. Thecleavage reaction of the substrate finally occurs leading to a third distinct “cleaved”state. It was shown in particular that 90–95% of the single ribozyme moleculesjumped to the cleaved state from the folded state, not from the unfolded state. Theremaining 5–10% of FRET trajectories were attributed to contributions of short-livedfolded state that were not observable with the limited (2 s) time resolution. SmFRETmeasurements thus allowed verification of the proposed reaction pathway, withcleavage occurring only in the folded state. In addition, measuring the dwell times inboth folded and unfolded states revealed simple rate kinetics for the foldingtransition, but more complex multiexponential dynamics for unfolding transitions.This suggested the existence of different folded states. Moreover, a large heteroge-neity in the unfolding kinetics was observed, with some time traces showingpredominantly fast unfolding and some other slow unfolding.This conformational heterogeneity was confirmed in subsequent observations by

Tan in 2003 on the natural form of the hairpin [50]. These experiments also revealedthat the system did not show a single unfolded but rather two rapidly interconvertingunfolded states. Transitions to the folded state occurred from the unfolded statespossessing the higher FRET efficiency, therefore called a “proximal” state. Thedynamics of conformational fluctuations depended strongly on Mg2þ concentra-tions. When the transitions were slow enough, it was possible to construct ahistogram of dwell times of the molecule in the proximal state (Figure 8.2a).This histogram could be fitted with a monoexponential decay, with a decay timecharacteristic of the transition between the proximal and the distal states(Figure 8.2b). At low Mg2þ concentrations, however, the fluctuations became toofast to be clearly directly measured from single-molecule traces (Figure 8.2c). FastsmFRETdynamics may be more clearly resolved by cross-correlating the donor andacceptor fluorescence intensity traces. Indeed, changes in FRET efficiency result inanticorrelated intensity variations of the donor and acceptor fluorophores. The cross-correlation function could then be fitted in turn with a monoexponential decay. In


this study, this analysis yielded transition rates of up to 1000s�1 between the twounfolded states (Figure 8.2d). Finally, cleavage reaction kinetics were examined on asingle hairpin basis from smFRET measurements and also showed a highheterogeneity.SmFRET data analysis must provide strong evidence that the molecule

immobilization on the substrate does not modify its conformational dynamics.To test whether the previously observed heterogeneity could originate from non-specific interactions between the immobilized hairpin and the substrate, theseexperiments were reproduced on single hairpins encapsulated inside 100–200 nmlipid vesicles [30]. This conformation is very different since the RNA is confined nearthe surface but not attached to it. Again, 50-fold variations in the dwell times offolded and unfolded states were observed. This confirmed that folding heterogeneitywas indeed intrinsic to the hairpin and possibly attributable to heterogeneity in theconformation of the loop substructures, not to nonspecific interactions with thesurface.While oligonucleotides present in general low levels of nonspecific interactions

with the substrate, possibly due to repulsive electrostatic interactions, proteins aremuch more complex macromolecules that interact more strongly with the substrateand its functionalization layer. For example, Rhoades had studied the foldingfluctuations of vesicle-encapsulated adenylate kinase (AK), a 214-amino acid protein

Figure 8.2 (a) SmFRET time trace showingtransitions between folded (F), proximal (Up),and distal (Ud) states. When transitions areclearly resolved, the histogram of dwell times instate Up reveals the transition time constant(here, 96ms). (b–d) When transitions are faster,

cross-correlation of donor and acceptor tracesallows access to short transition timeconstants, down to 1ms. (Reproduced withpermission from Ref. [50]. Copyright 2003,National Academy of Sciences, USA.)


[29]. To test whether AK proteins were interacting nonspecifically with the vesiclewalls, he first observed the fluorescence polarization values of AK labeled with thedonor fluorophore only. All observedmolecules showed very low polarization values,due to the fast rotational motion of the fluorophore that randomizes the polarizationof consecutive fluorescence photons. On the contrary, proteins adsorbed on glassshowed a much broader polarization distribution due to immobilization of theprotein. In addition, he found that the distribution of FRET values was significantlydifferent between vesicle-encapsulated and surface-immobilized proteins, suggest-ing a partial denaturation of the protein when adsorbed on the glass substrate andemphasizing the importance of the immobilization strategy. Most of the observedtime traces showed single FRET values due to the folding transition being slowerthan the average fluorophore lifetime (10–20 s due to photobleaching). However, insmFRET traces showing at least one transition, histograms of FRET values showedmainly two subpopulations corresponding to folded and unfolded states. Analyzingthe amplitude of changes in FRET values showed a large spread of the transitions,indicating a large heterogeneity of the folding reaction, and the existence of severalintermediate folded states. In addition, the authors show that many moleculesexhibit slow transitions (>1 s). These slow transitions were interpreted as continu-ous directed conformational changes, possibly slowed down by local traps.Intermolecular, not only intramolecular, interactions between DNA, RNA, and

proteins may also be probed by smFRET to examine reaction pathways and kinetics.Ha et al. have, for example, examined DNAunwinding by the Rep helicase [21]. DNAprobes were composed of an acceptor strand attached to a polymer-coated surfaceand a complementary donor strand. The two strands form a junction between single-stranded and double-stranded DNA (dsDNA). Unwinding of the dsDNA portion byhelicases increased the separation between the dyes and thus reduced FRETinteractions. Some time traces showed complete unwinding with a completesuppression of FRET due to fast diffusion of the donor strand away from theimmobilized acceptor strand. However, some time traces show stalls in the DNAunwinding, with occasional rewinding. These transient events could not have beendetected in ensemble experiments. Further experiments examining the kinetics ofhelicase binding and DNA unwinding under different concentration of Rep heli-cases were able to show that DNA unwinding must involve the interaction of morethan one Rep protein.In the preceding example, only one of the interacting partners, the dsDNA, was

labeled with fluorophores. This allowed tracking conformational changes of a fewDNA molecules in the presence of a high concentration of Rep helicases (up to100 nM) to increase the probability of Rep–DNA intermolecular interactions.However, situations where the molecules do not undergo drastic conformationalchanges upon interaction require labeling of both interacting molecules to followtheir binding and unbinding kinetics. In that case, one is confronted with twoseemingly incompatible constraints: the need for a low concentration to ensuredetection of isolated FRET pairs (typically <0.1 nM) and the need for a highconcentration to ensure efficient interaction (>1 mM depending on association–dissociation constants). Vesicle co-encapsulation provides a way to reconcile these


two aspects [34]. The effective concentration inside the nanovesicle may be as highas tens of mM due to the confined space (1 molecule complex in a 100 nm sphere).Vesicles may, however, be immobilized on a substrate with a sufficient spatialseparation to allow visualization of single vesicles. This experimental setup is thusparticularly interesting to study weakly interacting molecules. Benitez et al., forexample, have studied interactions between two copper-binding proteins, Hah1 andWDP, encapsulated in lipid nanovesicles (Figure 8.3a) [31]. Each protein was labeledat a C-terminal cysteine residue with a donor or acceptor fluorophore. The presenceof only one protein pair was verified by examining photobleaching steps. FRETtraces showed three distinct FRET states attributed to one unbound state and twodifferent bound states. Transition rates (binding, dissociation, and conformationalchange) for each state were then derived from histograms of the different dwelltimes (Figure 8.3b). In another study, Cisse et al. developed methods to introducenanometer-sized pores into the vesicle walls [32]. These pores were created either byincorporation of bacterial toxins or by bringing the vesicle at the lipid phasetransition temperature, triggering lipid packing defects. These pores wereimpermeable to large molecules and maintained confinement of the studiedDNA and protein molecules, but allowed modulating the concentration of ATP.The authors could then follow the interaction dynamics of the same interactingmolecules under different ATP conditions. The change in the chemical environ-ment indeed induced an observable change in the interaction between proteins andDNA. Finally, it should be noted that the vesicle lipid membrane constitutes apromising platform to study membrane-anchored proteins [34].

Figure 8.3 (a) Schematics showing twointeracting proteins trapped inside a lipidnanovesicle. (b) Model for the proteininteractions with one unbound and two distinct

bound states. (c–e) Histograms of dwell timeswith the corresponding derived transition rates.(Adapted with permission from Ref. [31].Copyright 2009, American Chemical Society.)


8.2.4Analyzing Complex FRET Trajectories

In most of the above examples, FRET trajectories were decomposed into two or threedistinguishable FRET states and corresponding macromolecular conformations.This is easily achieved when the number of states is low and when FRET values canbe clearly separated by a manually set threshold to discriminate between FRETstates. This method is, however, ill adapted to situations with a higher number ofstates, or when it becomes difficult to discriminate the effects of noise or photo-physical changes from a true conformational transition. These more complex FRETtrajectories may be instead analyzed using hidden Markov modeling (HMM). Thisalgorithm has been used in various applications such as speech recognition,cryptanalysis, and biophysics, including single-molecule FRET [51– 53]. Its parame-ters consist of the emission probability functions and the transition probabilitymatrix. The emission probability functions epA(B . . . )(g ) describe the probability of aspeci fic FRET ratio value g being detected when the system is in state A (B . . . ).Gaussian functions are generally used to include effects of real (conformational,photophysical) and noise-induced fluctuations. The transition probability matrix tpdescribes the probability of the system changing from a FRET state to another in thesubsequent step and is directly related to the different transition monoexponentialrates. The cumulated probability that a given FRET ratio sequence {g1, g 2, g 3}corresponds to a specific conformation trajectory {A ! A ! B} is then determinedfrom the product of all corresponding emission and transition probabilities: epA(g 1)epA(g 2)epB(g3)tp(A ! A)tp(A ! B). Similar probabilities are calculated for allpossible trajectories to determine the most probable trajectory. The Viterbi algo-rithm may help here to reduce computation costs [51,54]. In most experimentalcases, neither the emission probability functions nor the transition matrix is knownbeforehand. These parameters are thus varied and optimized over a set of manydifferent single-molecule FRET trajectories to obtain a maximized total probability.This finally yields the most probable emission probability functions for each stateand the corresponding transition probability matrix. Several publicly availableprograms have been developed that allow application of HMM to single-moleculeFRET data, such as HaMMy [51] (http://bio.physics.illinois.edu/HaMMy.html),QuB [52] (http://www.qub.buffalo.edu/wiki/index.php/Main_Page), and vb-FRET[55] (http://vbfret.sourceforge.net/).Hidden Markov modeling has been used, for example, by McKinney et al. to

analyze binding of RecA proteins to single DNA molecules [51]. Since several RecAproteins can bind to a single DNA strand, different FRET states are observed,corresponding to different RecA:DNA ratios. While the high number of statesprecluded analysis using manually set thresholds between the different FRETstates,HMM revealed the existence of five different states (0, 1, 2, 3, and 4 associated RecAproteins). Transition probabilities were high only between neighboring states,suggesting that RecA proteins bind and dissociate one by one to the DNA strand.Further analysis showed that association rates increased with RecA concentration,while dissociation rates were independent.


http://bio.physics.illinois.edu/HaMMy.html

http://www.qub.buffalo.edu/wiki/index.php/Main_Page

http://vbfret.sourceforge.net/

In another report, Abelson et al. study the conformational changes of a smallmRNA strand during in vitro splicing [56]. This complex multistep reaction involvesan ensemble of small ribonucleoproteins and leads to the removal of specific intronsequences from the RNA strand. Figure 8.4a shows a typical fluorescence and FRETratio time trace and the corresponding most probable sequence of FRET states.Figure 8.4b shows the transition density plot, representing the number of transi-tions observed from an initial FRET state (horizontal axis) to a particular final FRETstate (vertical axis) in a collection of single-molecule FRET trajectories. This plot isused as a visual representation of the different observed transitions. Sometimes,however, a small number of molecules exhibit an unusually high number of fasttransitions, which become therefore strongly emphasized in this representation,while slow transitions may be underrepresented. To avoid this problem, anotherrepresentation may be used with population-weighted and kinetically indexedtransition density (POKIT) plots. POKIT plots represent as concentric circles thefraction of molecules undergoing a specific transition at least once. This avoidsgiving too much importance to a small number of molecules exhibiting a largenumber of transitions. The average dwell time in an initial state before undergoing aspecific transition is coded with different colors (e.g., red corresponding to fasttransitions, green to slower ones, and so on). In the particular example shown inFigure 8.4c, these graphs summarize several important characteristics of the studied

Figure 8.4 (a) Typical donor and acceptorfluorescence and FRET time traces in thesplicing buffer, with the corresponding HMM-derived traces; corresponding TDP (b) and

POKIT (c) plots; (d) POKIT plot in ATP-depletedbuffer. (Reproduced with permission from Ref.[56]. Copyright 2010, Macmillan PublishersLtd.)


molecular system. It allows a quick evaluation of the number of distinct FRETstates.HMM analysis of this system revealed the existence of up to 10 or 11 states buttransitions occur predominantly between 5 distinct states. One can also immediatelysee that transitions occur mostly between neighboring FRETstates, implying a step-by-step reaction. Finally, the graph is symmetric with respect to the diagonal, whichmeans that most transitions work in both directions and that they are reversible.These graphs also facilitate comparisons of different reaction conditions. Forexample, Figure 8.4d shows the POKIT plot under ATP-depleted conditions: boththe nature and the dynamics of the observed transitions are modified.

8.3Single-Molecule FRET of Freely Diffusing Molecules

This section presents the second family of single-molecule FRET techniques,consisting in the detection of freely diffusing molecules in solution. We presentthe experimental techniques, along with a few examples to illustrate what can beachieved with solution smFRET experiments. Finally, we present advanced tech-niques that take advantage of the large volume of data available with solutionsmFRET to perform more refined data analysis.

8.3.1Experimental Setup

Detecting single molecules freely diffusing in solution presents several advantagesand drawbacks. It is both simpler and more robust than imaging immobilizedmolecules. This is due in part to the absence of any substrate surface susceptible tointerfere with the conformation of the molecule of interest, which greatly simplifiessample preparation and reduces possible sample-to-sample variations. On the otherhand, the need to isolate the fluorescence signal of an individual molecule from itsneighbors puts strong constraints on the detection scheme and chromophoreconcentration. The detection volume must indeed be sufficiently small to containat most one molecule at a time. This is usually realized by confocal or two-photonexcited fluorescence microscopy. Typically, an excitation laser is focused by a highnumerical aperture objective into the sample solution a few microns above a glasscoverslip. Fluorescence photons are collected through the same objective andseparated from the excitation beam by a dichroic mirror and emission filters.Donor and acceptor photons are then separated by a second dichroic mirror, passedthrough additional emission filters, and finally detected on avalanche photodiodedetectors (APDs). Appropriate pinholes are placed in the excitation and detectionpath at the image conjugate planes to ensure confocality. All detected fluorescencephotons originate from molecules located inside the small (few femtoliters)diffraction-limited confocal volume. Alternatively, the sample solution may beintroduced in a small glass capillary and flowed through the confocal observation


volume. This capillary may be integrated into a microfluidic device to probe, forexample, chemical reactions at different times after mixing or the effects of heatingby changing the location of the detection spot along a microfluidic channel [57].Finally, zero-mode waveguides present an interesting alternative to isolate fluores-cence signals from single molecules [58,59]. This technique uses nanometer-scaleapertures fabricated in thin metal films on a transparent substrate. Light intensitydecays very rapidly at the entrance of the apertures, creating effective detectionvolumes that are three orders of magnitude smaller than diffraction-limited confocalvolumes. In practice, this enables to probe more concentrated solutions but reducesthe time spent by the molecule in the detection spot.In concentrated solutions, Brownian motion and flow induce fluctuations in the

detected fluorescence signals, whichmay be analyzed using fluorescence correlationspectroscopy (FCS) [60,61]. However, when the observed solution is dilute enough(typically a few tens of pM for confocal detection schemes), most of the time theconfined observation volume does not contain any fluorophore, and for brief periodsit contains a single biomolecule. The resulting traces thus present long periods ofbackground noise interrupted by fluorescence bursts. The duration of these burstscorresponds to the time necessary for the biomolecule to diffuse out of theobservation volume. Their intensity depends on the intrinsic fluorescence parame-ters described in Equations 8.3 and 8.4 for the observation of single immobilizedmolecules (excitation cross sections, fluorescence quantum yields, FRET efficiency,etc.). APD signals are collected by counting boards and time traces are thenregistered, which can be binned using different time resolutions. Fluorescencebursts above a predefined threshold value are then selected from the backgroundnoise and corrected from detection efficiencies and spectral cross-talks as describedin Equations 8.1 and 8.2. The most simple and straightforward analysis of single-molecule FRETdata consists in displaying histograms of the number or populationfraction of bursts as a function of the emission ratio g¼ IA/(IAþcID), where IA andID are the acceptor and donor burst intensities, respectively, and c¼WA/WD is thecorrection factor already described above (Figure 8.5). The emission ratio g is closelyrelated to the FRET efficiency provided all corrections are properly performed(Equation 8.5). It is, however, difficult to ensure that each fluorescence burst isappropriately corrected, due to the possible heterogeneity in the environment andphotophysical properties of each fluorophore pair, and the emission ratio is oftenreferred to in practice as the apparent FRET efficiency, Eapp.

8.3.2Applications

The first ratiometric measurements of single freely diffusing biomolecule FRETwere developed practically simultaneously to those on surface-bound molecules[63–65]. Immediate advantages of solution measurements were to eliminate possi-ble artifacts due to nonspecific interactions with the surface and to rapidlymeasure alarge number of events to enable statistical analysis. Solution smFRET was rapidlyapplied to identify subpopulations in a heterogeneous ensemble. In particular,

8.3 Single-Molecule FRET of Freely Diffusing Molecules j337

biomolecule labeling with donor and acceptor fluorophores is often incomplete. Inaddition, a portion of the acceptor dyes may be nonfluorescent and/or nonabsorbingdue to photobleaching and blinking. This often leads to a nonnegligible populationof donor-only biomolecules. These donors are not quenched by FRET, and this maylead to underestimate FRET efficiencies in ensemble measurements. SolutionsmFRET allows an easy identification of those donor-only molecules as a zero-centered peak in apparent FRETefficiency histograms. Deniz et al. demonstrated in1999 that they were able to isolate this population and take into account only signalsfrom dual-labeled DNA double strands for sufficiently high average Eapp (�>0.4)[64]. In addition, they varied the distance between the donor and acceptor dyes andobserved a progressive decrease in FRET efficiency for larger separation distances.They showed that they were able to separate two dsDNA populations correspondingto different interdye distances. The sequence with the longer separation distance,DNA17, contained an enzyme target sequence between the donor and acceptorfluorophores, while the other, DNA7, did not. Cleavage of the DNA strands by theenzyme led to the separation of the fluorophores in the DNA17 population. Theauthors showed that the DNA17 peak in the Eapp histogram indeed disappearedwhile the peak around zero, corresponding to isolated donor molecules, increasedaccordingly. The peak corresponding to the DNA7 sequence remained unchangedin the process.Protein folding studies have benefited from smFRET as a tool to separate folded

from unfolded species. The cold shock protein, for example, has served as an

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Figure 8.5 Typical solution smFRET opticalsetup and experimental fluorescence burst timetraces. Bursts are selected (arrows) when thesum of their donor and acceptor photons islarger than the predefined threshold (dashedline). The donor and acceptor burst intensities

are then analyzed to provide a FRET histogram,showing, for example, donor-only (D), unfolded(U), and folded (F) species. (Adapted withpermission from Ref. [62]. Copyright 2006,American Chemical Society.)


excellent model system for protein folding due to its simple structure and its two-state behavior [66,67]. Under normal conditions, FRET efficiency histograms ofdual-labeled protein solutions show the usual donor-only peak around zero and apeak at high FRET efficiency, corresponding to the compact folded state. Underdenaturating conditions, this peak disappears while a new peak arises at lower FRETefficiencies, corresponding to unfolded states (Figure 8.6a) [68]. In addition, thissecond peak shifts toward lower FRET efficiencies as the concentration of denatur-ant increases, indicating that the protein further unfolds. Revealing this type ofcomplex behavior is a unique force of single-molecule studies and would have beenvery difficult to extract from FRETensemble measurements. In this simple solutionexperiment, one observes the equilibrium of folded and unfolded populations, butdoes not have access to out-of-equilibrium conditions such as transient states or theunderlying folding and unfolding dynamics. This may be achieved using micro-fluidic devices in which different solvents, proteins, andmolecules may be flowed inchannels and mixed where the channels merge under controlled conditions of flow.Locating the detection volume at different points of the output channel allowsprobing different times after mixing of the reactants, depending on the flow speedand the distance between the detection volume and themixer. Lipman et al. used this

Figure 8.6 (a) Histograms of measured FRETefficiencies at various denaturantconcentrations for labeled cold shock protein.(Reproduced with permission from Ref. [68].Copyright 2002, Macmillan Publishers Ltd.) (b)Varying the distance between the mixing pointof the microfluidic channel and the laser

confocal volume allows probing different timesafter mixing. FRET histograms obtained fordifferent times after mixing. (Reproduced withpermission from Ref. [57]. Copyright 2003,American Association for the Advancement ofScience.)


principle to probe cold shock protein folding kinetics by smFRET [57]. They inducedan abrupt drop of denaturant concentration in the protein solution by dilution. Theyobserved that the unfolded proteins rapidly switch to a more compact (higher FRET)unfolded state before slowly reaching the new equilibrium between folded andunfolded states (Figure 8.6b). This experiment thus gives access to transient speciesthat are not strongly represented under equilibrium conditions.The ability to eliminate artifacts due to imperfect biomolecule labeling and

measure FRET efficiencies with a good precision was used, for example, to revisitpolyproline peptides as rigid spectroscopic rulers. These studies provide a goodillustration of what information can be extracted from smFRET measurements interms of subpopulations and dynamics, and of the possible caveats. Polyprolinesadopt a type II helix in aqueous solution [69] and have been used as rigid spacers toverify the distance dependence of FRET in the range of 1–12 prolines per peptide,

corresponding to a �20–45A�range [70]. Schuler examined polyprolines in a larger

size range, from 6 to 40 proline residues using a FRETpair with a larger R0 distance

(54A�instead of 35A

�in the earlier work) [71]. He found that solution of longer

peptides contained a nonnegligible fraction of donor-only molecules, which couldlead to errors in ensemble measurements. However, the authors observed asignificant discrepancy between the observed FRET efficiencies, Eapp, and thosetheoretically predicted assuming a rigid peptide conformation. Shorter peptidesshowed lower Eapp than predicted; this discrepancy has been attributed to failure ofthe point-dipole FRETmodel at short interdye distances and to the absence of fastorientational averaging, and consequently an error in the estimation of the k2 factor.This was supported by polarization measurements. Long peptides showed a lowresidual steady-state polarization value of 0.05, corresponding to a near totalreorientation of the dipole between excitation and emission. However, for shorterpeptides, the donor decay was significantly accelerated due to FRET and becamecomparable to the reorientation speed, as shown by the increase of steady-statepolarization value to 0.11. Replacing the usual k2¼ 2/3 value corresponding to therapid orientational averaging by an approximation of random, but static orienta-tions, the authors found theoretical FRET efficiencies compatible with the observedvalues.Longer peptides, on the contrary, showed higher FRETefficiencies than expected.

These longer peptides must thus adopt different conformations, bringing the twodyes closer to each other, and allow for higher FRET rates. Molecular dynamicsshowed that the timescale of these fluctuations (0.1–10 ns) was much shorter thanthe fluorescence burst duration (1ms). In this regime, the effect of conformationalfluctuations on the apparent FRET efficiency depends on the comparison betweenthe dynamics of the peptide conformation, the rotational correlation time of thefluorophores, and their fluorescence lifetime. In this study, fluorescence andanisotropy decay measurements showed that the dye rotation (�0.3 ns) was fasterthan both the fluorescence lifetime (1–10 ns) and the expected conformationalfluctuations (0.1–10 ns). The obtained theoretical estimations of FRET efficiencieswere in the same range as the experimental values, confirming that these peptides


indeed adopt flexible conformations and showing the importance of consideringdynamics for extracting structural information.Another regime exists when conformational fluctuations change slowly compared

to the duration of fluorescence burst, that is, the time spent by the biomolecule in thedetection spot before diffusing away. In that case, eachmolecule presents a different,static conformation when passing through the confocal volume. This results in abroadening of the FRET histograms. The width of FRET histogram peaks thuscontains information about dynamics of conformational changes. Its analysis is,however, complicated by additional contributions from shot noise and photophysicaleffects. The shot noise standard deviation ssn arises from the finite number ofphotons in each time bin and provides a fundamental statistical lower limit to thewidth of the histogram peaks. It is defined by

s2sn ¼ �Eð1� �EÞ=ðnA þ nDÞ; ð8:9Þ

where �E is the average FRETefficiency and nA þ nD is the total number of photons inthe donor and acceptor channel. A limit on this number is provided by consideringthe threshold value below which fluorescence bursts are rejected. Additional effectsof dye molecular fluctuations are difficult to isolate and characterize. A referencemolecule with the same fluorophores but no conformational fluctuations slowerthan the time bins considered may be used to estimate a standard width s0. Anyexcess width, defined by s2 � s2

0, may then be attributed to slow conformationalfluctuations. Theoretical FRET histogram widths have been derived by Gopich andSzabo as a function of fluctuation dynamics and acquisition speed [72,73]. Schuleret al. used this method to examine the folding kinetics of cold shock protein. Theyanalyzed FRET histograms obtained under different denaturing conditions, wherethe protein fluctuated between unfolded and folded configurations (Figure 8.6a).They then observed that the FRETpeak widths were not much larger than those for ashort rigid polyproline peptide [67,68]. This allowed them to put an upper limit of30 ms to the protein reconfiguration time. This conformational dynamics can beused to obtain information about the free energy barrier for folding, as discussed inRefs [67,68].Best et al. have further used the distributions in FRET histograms to obtain

information about conformational flexibility of long polyprolines [74]. They haveused a pulsed excitation source and recorded the arrival time of each photon after thecorresponding excitation pulse. Interestingly, the authors were able to rule outpossible contributions to the distribution widths from acceptor photobleaching, byobserving that histograms constructed from the first and second halves of the timebins were identical. If photobleaching had any influence, histograms from thesecond half of the bins would have been shifted to lower FRET values. They alsoruled out contributions from acceptor random blinking by examining the statisticsof strings of consecutive donor or acceptor photons. If there is no blinking, theprobability of observing a sequence of n consecutive donor photons varies as(1�Eapp)

n. In contrast, blinking of the acceptor would give rise to a higherprobability of observing long strings of donor photons. The distributions of


consecutive sequences of donor photons perfectly matched theoretical predictionsin the absence of blinking.The ability to record the arrival times of photons after each excitation pulse

allowed them to reconstruct fluorescence decay curves from selected molecularsubpopulations [74,75]. They were able to show that, for polyproline peptides inwater, photons from molecules showing up in the higher side of the FREThistogram peak yielded a faster decay curve compared to photons from moleculesshowing lower FRET values (Figure 8.7). This indicated that FRET value heteroge-neity indeed originated from differences in FRET rates, that is, to (slow) conforma-tional fluctuations, not from noise. In contrast, fluorescence decay curves wereidentical throughout the smFRET histogram measured in trifluoroethanol (TFE),indicating that conformational fluctuations were smaller in this solvent compared towater. The authors were further able to use fluorescence decay data as an additionaltool to evaluate FRETefficiencies. To construct the fluorescence decay of donor-onlyspecies, they used photons from bursts showing a near-zero emission ratio. Theythen calculated frommolecular simulations the predicted fluorescence decay curvestaking into account FRET processes corresponding to different peptide conforma-tions. This analysis, together with NMR measurements, allowed them to identifydifferent peptide conformations in TFE and water due to isomerization of prolineresidues.

Figure 8.7 SmFRET results for polyproline-20in TFE (a) and water (b). The yellow dashed lineindicates shot noise-limited width of thedistribution. Insets: The donor fluorescencedecays for donor photons from the

subpopulations with corresponding colors inthe efficiency histograms. (Reproduced withpermission from Ref. [74]. Copyright 2007,National Academy of Sciences, USA.)


8.3.3Advanced Solution smFRET Methods

Single-molecule FRETmethods described above are powerful tools to study hetero-geneous populations but suffer from several limitations, due to sample preparationand complex fluorophore photophysics. For example, smFRET cannot distinguishbetween donor-only species and dual-labeled species with low FRET efficiencies: inboth cases, only the donor fluorophore is detected. More generally, it cannot detectacceptor-only species, which limits studies of the interactions of a donor-labeledmolecule A with an acceptor-labeled molecule B. To overcome these limitations,several techniques were introduced, based on alternate laser excitation (ALEX) andmultiparameter fluorescence detection (MFD). These schemes may also be appliedto surface-immobilized molecules, but are more adapted to solution diffusingprobes, thanks to the large volume of data accessible in solution measurements.

8.3.3.1 Alternate Laser ExcitationThe ALEX method has been developed by Kapanidis et al. as an extension ofsmFRET, in which the excitation consists of interleaved pulses of two differentwavelengths: one for donor excitation and the other for acceptor excitation [76,77].The excitation wavelength switchesmore rapidly than the average fluorescence burstduration, so that many excitation pulses are used for each single diffusing molecule.Photon detection is synchronized with the excitation pattern providing four sets ofdata after cross-talk and background corrections. Two of these fluorescence time

traces correspond to standard smFRET measurements, denoted FDemDexc and FAem

Dexc,respectively, for the donor and acceptor emissions under excitation of the donor. Theother fluorescence time traces correspond to the emission of the donor and the

acceptor under direct acceptor excitation, FDemAexc and FAem

Aexc (in practice, FDemAexc � 0).

The first two signals provide the usual apparent FRET efficiency, Eapp, usingEquation 8.7. In addition, the sum of all donor and acceptor emissions under

donor excitation,FDexc ¼ cFDemDexc þ FAem

Dexc, and that under acceptor excitation,

FAexc ¼ cFDemAexc þ FAem

Aexc, are calculated. One can now also define a stoichiometryratio, S, as

S ¼ FDexc

FDexc þ FAexc: ð8:10Þ

This ratio is independent of FRET efficiency, since FDexc sums all photons emittedafter donor excitation and is corrected for the difference in quantum yields anddetection efficiency between the two fluorophores. The laser excitation intensitiesare usually adjusted to yield FDexc � FAexc. In that case, donor-only species displayS� 1, acceptor-only species display S� 0, and dual-labeled molecules assumeintermediate values. Kapanidis et al. demonstrated that this method enabledseparation of subpopulations based on apparent FRET efficiency and stoichiometry[77]. They were, for example, able to correctly take into account acceptor-only species,and also the presence of dimers (e.g., two donor–one acceptor macromolecules).


In general, Smay be directly used to determine the ratio of the number of donor andacceptor fluorophores in each fluorescence burst [78]:

nAnD

� 1SIAexcsA

Aexc

IDexcsDDexc

� 1

� �; ð8:11Þ

where IAexc and IDexc are the two laser excitation intensities and sDDexc and sA

Aexc arethe excitation cross sections of the donor and acceptor molecules at their respectivedirect excitation wavelengths.It was also used to discriminate donor quenching due to increased FRETor due to

a change in the donor local environment. In the first case, E changes, not S, while inthe second case both E and S change. In a standard smFRET setup, these twosituations would have been indistinguishable. In addition, ALEX measurementsallow a more accurate determination of the c factor (Equation 8.6) using severalsamples or a standard pair [78]. The authors were then able to validate ALEX-baseddistance measurements on dsDNA samples of different lengths (see Figure 8.8) andmonitor protein–DNA association–dissociation equilibrium and kinetics [77].

8.3.3.2 Multiparameter Fluorescence DetectionMost of the experiments and methods described above used fluorescence intensityin the donor and acceptor channels. In fact, fluorescence photons carry additionalinformation that can provide useful information, help identify possible artifacts, andlead to a better characterization of the observed system. MFD typically uses photonpolarization and delay between dye excitation and fluorescence emission as twoother particularly useful parameters [79–82]. Fluorescence polarization measure-ments require the use of linearly polarized excitation light and two detectors peremission channel placed behind polarizers oriented parallel and perpendicular,respectively, to the excitation polarization direction. In addition, using laser

Figure 8.8 Typical example of an ALEXsmFRET experiment, with two dsDNA strandswith different interdye distances. The S versus Eplot reveals donor-only species (upper left),acceptor-only species (bottom right), and

different FRET pairs (middle). The relationshipbetween S and E for FRET pairs enables anaccurate determination of the c correctionfactor. (Reproduced with permission fromRef. [78]. Copyright 2005, Elsevier.)


excitation pulses shorter than the fluorescence decay of fluorophores allows regis-tering the arrival time of each photon using standard time-correlated single photoncounting (TCSPC) techniques. Complete data sets then consist of four streams ofphotons (donor and acceptor channels with two polarizations each) with twoattached time stamps. The first time is a “micro” time and corresponds to thedelay between photon emission and excitation by the laser pulse (up to a fewnanoseconds). The second time stamp is a “macro” time and corresponds to thenumber of the excitation pulse or, equivalently, to the time elapsed, since thebeginning of the acquisition sequence (seconds to minutes to hours). This exper-imental scheme may be combined with ALEX by alternating excitation pulses atwavelengths corresponding to the donor and acceptor excitation, respectively (seeabove). Fluorescence bursts corresponding to diffusion of single molecules in theconfocal spot are identified as in a normal smFRET experiment and are character-ized by a large amount of data. Several analytical techniques may then be utilized tocombine these data in order to characterize smFRET populations more accuratelyand avoid possible misinterpretation related to complex photophysics or restricteddye mobility [82].In particular, apparent FRET efficiencies may be estimated from both the

emission ratios and donor fluorescence lifetime. If FRET is the only processresponsible for molecular fluorescence changes, then Eapp values estimated fromthese two methods should coincide:

Eapp ¼ IAIA þ cID

¼ 1� tDðAÞtDð0Þ

; ð8:12Þ

where tDðAÞ and tDð0Þ represent the donor fluorescence lifetime in the presence and

in the absence of the acceptor, respectively. A standard technique then consists inrepresenting fluorescence bursts in a 2D histogram of emission ratio versus changein donor fluorescence lifetime. All fluorescence bursts should be located along asingle curve determined by Equation 8.12, and molecules with fluorescence burstsoutside this curve must necessarily undergo additional photophysical processes. Forexample, quenching of the donor caused by its local environment would accelerateits fluorescence decay without increasing the emission ratio, resulting in a deviationfrom the theoretical curve. MFD is thus able to identify and correctly take intoaccount the effect of the local environment of the donor and acceptor dyes [83].MFD is also able to check for any variation in the rotational diffusion of the dyes.

Indeed, FRETanalysis often relies on the assumption that the k2 factor, related to theorientation correlation of the dyes, remains constant within the whole molecularpopulation. Measuring the donor polarization anisotropy for all fluorescence burstsis a good way to verify this assumption. In the absence of any uncontrolled variationof dye rotational diffusion, the anisotropy should follow the Perrin equation:

r ¼ r0=ð1þ tDðAÞ=rÞ; ð8:13Þwhere r0 is a fundamental dye anisotropy and r is the dyemean rotational correlationtime. Similarly to emission ratio versus fluorescence lifetime histograms described


above, 2D histograms comparing donor anisotropy and fluorescence lifetimemay beused to monitor any undesired modification in the orientational dynamics of thedonor fluorophore [82]. For example, Figure 8.9 shows typical results of smFRETexperiments with a mixture of same dsDNA labeled with three distinct interdyedistances (5, 11, and 19 bp separation) and donor-only species [84]. All data pointsremain close to the theoretical relation curves between the emission ratio, the donorfluorescence lifetime, and the donor anisotropy. This shows that no dye quenchingor change in dye rotational correlation time due to the local environment isoccurring, and that observed photophysical changes can safely be attributed to FRET.

8.4Single-Molecule FRET Studies Involving Multiple FRET Partners

Sections 8.2 and 8.3 were dedicated to smFRET methods applied to single FRETpairs, that is, most of the time, to individual (macro-)molecular complexes contain-ing a single donor and a single acceptor. These techniques have provided a greatwealth of information about intermolecular interactions and changes in molecularconformations that were not accessible by ensemble measurements. However, theresulting information is limited to the distance between two labels placed at strategiclocations on the molecules of interest. This single inter- or intramolecular dis-tance is insufficient to completely resolve three-dimensional molecular structures.

Figure 8.9 Typical results from an smFRET experiment analyzed using MFD, showing theexpected relation between donor lifetime, emission ratios, and donor anisotropy as red lines.(Reproduced with permission from Ref. [84]. Copyright 2008, Elsevier.)


In addition, complex systems composed of FRET interactions between multipledonors and/or multiple acceptors require specific single-molecule FRETmethods.In the following section, we will explore expansions of single-molecule methods tomulticolor and multiple FRET interaction systems.

8.4.1Multistep FRET

The first advantage of increasing the number of labels involved in FRET is that itprovides access to larger distances [85]. Since the FRETrate decreases as the inversesixth power of the donor–acceptor separation distance, inserting an intermediatedye greatly enhances the total FRET rate between the initial donor and the finalacceptor dye. This intermediate “jumper” dye serves as an acceptor for the initialdonor and as a donor for the final acceptor, so special care should be taken to selectdyes with appropriate spectral features. The range of accessible FRET distances

increases to more than 100A�instead of typically 70A

�for standard two-color FRET

[85]. This concept may be extended with the use of several jumper dyes to form“photonic wires” capable of transporting energy over large molecular distances [86].Garcia-Parajo et al. studied such a system composed of five distinct fluorophoresplaced along a double-stranded DNA molecule from the bluest at one end to thereddest at the other end [87]. Single-molecule fluorescence spectroscopy showed thatthese systems exhibited complex fluctuations in FRET rates due to dye blinking andspectral and/or orientational fluctuations. Interestingly, about 10% of the wires

showed FRET efficiencies of up to 90% over a total distance of more than 130A�.

An alternative configuration consists in labeling a macromolecule site specificallywith a single donor and two distinct acceptors. FRETmay then occur from the bluestdye to each of the two other, and also from the intermediate dye to the reddest one.Monitoring emission intensity ratios corresponding to each acceptor now providesinformation about two independent distances. These observations offer betterinsight into three-dimensional structures and into correlations between differentconformational changes [88–90]. For example, Hohng examined surface-immobi-lized DNA four-arm Holliday junctions as a model system [89]. He labeled thejunction with a donor on one arm and two different acceptors on the two adjacentarms. He was able to confirm previous data showing that the junction adopts twoconfigurations corresponding to the donor arm being stacked with either one of theacceptor arms. The two acceptor signals were anticorrelated: when one was strongthe other was weak, while the donor signal was almost quenched with a constantrate. This suggested that the two acceptor arms do not approach the donorsimultaneously, but do so in an alternate manner [89]. A three-color ALEX methodhas also been developed, using three alternate excitation sources to improve theaccuracy of distance estimations [91,92]. In this case, it is necessary to have threefluorophores that are spectrally distinct enough so that each of them may beselectively excited independently of the others. Lee et al. demonstrated thatthree-color ALEX enabled separation of different triply labeled species and the

8.4 Single-Molecule FRET Studies Involving Multiple FRET Partners j347

simultaneous determination of all three interdye distances in each subpopulation(Figure 8.10).Finally, the use of multicolor FRET to investigate biomolecular structures has

been pushed one step further with the development of four-color FRETALEX at thesingle-molecule level [93,94], with the use of three different excitation lasers or asupercontinuum laser for free excitation wavelength selection. Four is, however,probably the maximum number of dyes that can be used in such schemes due to thedifficulty to maintain spectral separation.

8.4.2Multi-Acceptor and Multi-Donor Systems

Other interesting configurations with multiple interacting FRETpartners consist ofa single donor linked to a large number of acceptors, or vice versa. This is, forexample, the case of nanoparticle FRET systems such as semiconductor quantumdots (QDs) [95,96]. QDs display optical properties that make them interesting FRETdonors: high quantum yields, spectrally narrow emission spectra tominimize donorleaking into the acceptor channel, and finally high excitation cross sections andbroad excitation spectra to excite the QD donor far from the acceptor excitationspectrum and minimize direct acceptor excitation (see Chapter 12 for a completediscussion on QD-based FRET). Many acceptor-labeled molecules may assemble atthe surface of the QDs, increasing the overall FRETquenching efficiency of the QDdonor. When n acceptor dyes surround the QD donor, separated from the QD centerby an equal distance r, FRET efficiency can be expressed as

EðnÞ ¼ n

nþ ðr=R0Þ6: ð8:13Þ

Near-unity FRETefficiencymay thus be attained by simply increasing the number ofacceptors per QD. These characteristics led Zhang et al. to propose a highly sensitivesensor for the detection of DNA targets based on single quantum dot detection

Figure 8.10 Three-dimensional FRET efficiency histograms for triply labeled species, B-G-R. One-dimensional FRET histograms are obtained after collapsing the three-dimensional histogram oneach axis. (Reproduced with permission from Ref. [91]. Copyright 2007, Elsevier.)


(Figure 8.11a) [97]. All DNA strands in a sample were labeled with an acceptor dye,but only specific target DNAmolecules were assembled at the surface of streptavidinQDs using biotinylated complementary DNA sequences. The QD played two roles inthis scheme. Thanks to the proximity-driven FRET process between the QD and its

Figure 8.11 (a) FRET histograms for QD aloneand QD with on average 12 acceptor-labeledDNA strands per QD, showing the QD-only andthe QD–DNA populations. (b) FRET histogramfor 0.5 acceptor-labeled proteins per QD onaverage, showing the QD-only, QD-1 protein,QD-2 protein, . . . subpopulations. (c)Fraction of QDs without any acceptor (E< 10%;red squares) and engaged in FRET (E> 15%;

black triangles) as a function of N, the averagenumber of acceptors per QD, obtained fromsmFRET measurements. The fit corresponds tothe Poisson distribution p(N, 0)¼ exp(�N).(Reproduced with permission from Ref. [97],copyright 2005, McMillan Publishers, and Ref.[62], copyright 2006, American ChemicalSociety.)

8.4 Single-Molecule FRET Studies Involving Multiple FRET Partners j349

conjugated acceptor-labeled DNA strands, it allowed selective detection of capturedDNA target molecules and rejection of the signal from other unconjugated DNAstrands. It also played the role of a nanoconcentrator by bringing up to several tens ofacceptor-labeled DNA strands at its surface, thus increasing the local target concen-tration and detection sensitivity. The resulting assay was able to detect specific DNAstrands with a higher sensitivity than a standard molecular beacon detectionscheme, thanks to a very low background and a high responsivity [97]. In anotherstudy, solution smFRET was used to probe the heterogeneity within a populationof QDs self-assembled with acceptor-labeled proteins [62]. Different sampleswere studied with different average ratios of acceptor-labeled proteins per QD.Single FRET QD histograms showed that each sample consisted of several distinctsubpopulations, corresponding to n¼ 0, 1, 2, . . . acceptors per QD (Figure 8.11b).It was shown that the fraction of QDs assembled with exactly n proteins, p(n),followed a Poisson distribution as a function of the average number N of proteinsper QD in the solution (Figure 8.11c):

pðnÞ ¼ Nnexp ð�NÞn!

: ð8:14Þ

The authors were then able to probe conformational changes of a maltose-bindingprotein (MBP) conjugated to the QD surface, as a function of the maltose concen-tration, and to discriminate between QD–MBP conjugates bound to maltose or not.Single QD FRET is therefore a powerful tool to characterize the composition andfunctionality of a QD bioconjugate population, which is critical to further applica-tions for biological imaging or sensing.The use of QD donors to measure intramolecular conformational dynamics at the

single-molecule level has been demonstrated by Hohng and Ha in 2005 [98] onsurface-immobilized QD–DNA junction conjugates. The authors were able toanalyze the dynamics of conformational changes of the DNA junction. Again,the optical properties of the QD donor allowed for limited direct acceptor excitationand efficient spectral separation. However, this type of experiment is confrontedwith several limitations. The large size of the functionalization coating of commer-cial quantum dots causes significant increase in the QD donor–dye acceptordistance, resulting in low FRET efficiencies in single QD–single acceptor pairs.In addition, a quantum dot population is composed of quantum dots of differentsizes and hence different emission wavelengths. This spectral inhomogeneityresults in a difference of spectral overlap and FRET efficiencies among the QDpopulation [99]. Furthermore, control over the exact number and orientation ofbiomolecules conjugated to the QD surface remains a challenge that needs to beaddressed before QDs can be reliably used for estimating intramolecular distanceswith the same level of accuracy as organic dye donors at the single-molecule level.Symmetrically opposite FRETsystems composed of multiple donors surrounding

a single acceptor also exist, such as QD acceptors coupled to several terbium chelatedonors [100] or dendritic organic chromophores [101,102]. Single-molecule spec-troscopy has been used to characterize photophysics and FRET interactions inorganic dendrimers composed of four or eight donor fluorophores surrounding a


single acceptor fluorophore. These studies confirmed the influence of the isomericconformations on the decay rates of dyes and revealed that these different confor-mations also strongly influenced the FRETefficiency from the peripheral donors tothe central acceptor. The authors also showed the existence of excited donor dimers,when two donors are excited simultaneously in the same molecule, and of “excitonblockade,” when donor and acceptor chromophores are excited in the same mole-cule, leading to simultaneous dual emission. Single-molecule spectroscopy there-fore represents a powerful tool to characterize the photophysics of complexfluorophore systems.Finally, FRET may also occur between multiple identical chromophores, and is

often called in this case homo-FRET or donor–donor energy migration (DDEM)[103,104]. This may be particularly interesting since labeling a macromolecule withseveral identical fluorophores at several specific locations is much easier thanattaching one donor and one acceptor molecule. However, the overlap betweenthe absorption and emission spectra is not as important as in standard donor–acceptor pairs, and thus FRET rates are smaller. Furthermore, FRET cannot beobserved using spectral separation of fluorescence signals. Instead, fluorescencedepolarization and lifetimes are used to characterize FRET interactions and estimatedye separation distances. Single-molecule homo-FRET measurements have beenperformed on multiple-fluorophore-labeled antibodies, not to derive a three-dimen-sional structure but to characterize homo-FRET and self-quenching effects andoptimize single-antibody fluorescence brightness [105]. The authors measured thelifetimes, fluorescence intensities, and polarizations from single antibodies labeledwith average ratios of up to 10 dyes per antibody. Fluorescence time traces showedseveral distinct steps corresponding to successive bleaching of individual dyes onthe same antibody. Measuring the fluorescence lifetime from each of these stepsrevealed that the average lifetime slows down as the number of dyes decreases onindividual antibodies. This has been attributed to the presence of homo-FRET: oneexcitation quantum can visit several dyes, including some with fast nonradiativedecays and with fast bleaching. As a consequence, the overall quantum yield is lowerand bleaching occurs faster in the initial steps when the antibody is labeled with ahigh density of fluorophores.

8.5Conclusions and Perspectives

Recent progress in both instrumentation (avalanche photodiodes, low-noise sensi-tive CCDs, time-correlated single photon counting, etc.) and biomolecule labelingtechniques has allowed a rapid development of powerful single-molecule FRETmethods. The distance dependence of FRETprocesses gives access to the separationdistance between two dye molecules in individual macromolecules. This allows inturn the identification of subpopulations that would be indistinguishable inensemble measurements and access to the dynamics of conformational fluctua-tions. A wide variety of smFRET experimental formats may be used depending on

8.5 Conclusions and Perspectives j351

specific constraints and the information seeked. Solution measurements are fastestand easiest to perform, but several techniques have also been developed toimmobilize the labeled molecules on a substrate with minimal perturbation oftheir function and conformation characteristics. Time trajectories of FRET signalsare then accessible, enabling access to the sequence and dynamics of conforma-tional changes without having to synchronize a large molecular population. Finally,the ability to detect photophysical properties of FRET complexes one molecule at atime provides a way to correctly take into account effects of imperfect dye conjuga-tion schemes, dye environment, orientation heterogeneity, and complex photo-physics through appropriate analysis strategies.One of the most obvious limitations specific to smFRETmeasurements lies in the

limited number of photons that can be emitted by a single fluorophore beforephotobleaching. This limits the precision of the measured parameters and theduration of single-molecule observations. Recently though, improved photoprotec-tion strategies have been developed to increase the photostability of commercial dyesand reduce their photoblinking. Campos et al. optimized the combination of oxygenradical scavengers and triplet quenchers to increase the number of available photonsper single dye in solution-phase smFRETmeasurements [37]. They were thus able touse high-power illumination to increase the fluorescence photon flux and ultimatelythe time resolution, down to 50 ms. As demonstrated by Campos et al., this givesaccess to fast conformational dynamics that were previously masked by the longeracquisition time binning.Single-molecule FRET techniques are still rapidly evolving. One of the most

promising challenges consists in translating smFRETmeasurements from in vitro tothe intracellular environment. FRET data may then be coupled to single-moleculetracking to fully characterize single-molecule conformation, dynamics, and inter-actions with the complex intracellular medium. Important challenges to smFRET inliving cells reside in higher backgrounds (thus lower signal-to-noise ratios) and inlabeling the biomolecules of interest. FRET pairs composed of two fluorescentproteins are difficult to observe; therefore, the experimenter has to introduce at leastone FRETpartner labeled with an organic fluorophore. Murakoshi, for example, hadused FRET to measure the intracellular binding of microinjected acceptor-labeledGTP on a YFP-fused G protein [106]. He correlated FRETsignals with the binding ofGTP on single proteins and with the measured diffusion coefficients. He found thatthis activation was correlated with the immobilization of the protein in the cellularmembrane, which could be induced by formation of a signaling complex. Furtherprogress toward the application of smFRET in vivo will require the development ofnew labeling techniques and adapted analysis methods, such as multiparameterfluorescence image spectroscopy (MFIS) [107,108].Finally, smFRETmay also be combined with other biophysical techniques, such as

in simultaneous force and smFRETmeasurements. Applying and measuring forceson a single molecule enables controlled manipulation of the molecular structure,while smFRET detection of the same single molecule gives access to smallconformational changes at specific locations. These methods have, for example,been applied using optical or magnetic tweezers on dual-labeled immobilized


oligonucleotides [109–111] and could give rise to the design of novel single-moleculeforce sensors to probe the relationship between mechanical properties, molecularstructure, and biochemical function.

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