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Home > Fluorescence Microscopy > FRET Microscopy

Fundamental Principles of Förster Resonance Energy Transfer (FRET)Microscopy with Fluorescent Proteins

In living cells, dynamic interactions between proteins are thoughtto play a key role in regulating many signal transductionpathways, as well as contributing to a wide spectrum of othercritical processes. In the past, classical biochemistry approachesto elucidating the mechanism of such interactions were commonplace, but weakor transient interactions that might occur within the natural cellular milieu areusually transparent to these techniques. For example, co-localization of suspectedprotein partners using immunofluorescence microscopy in fixed cells has been apopular method for examining interactions in situ, and numerous literature reportshave been presented based on this technique. However, because the resolutionof a fluorescence microscope is several hundred times less than the size of atypical protein, co-localization often leads to questionable results. An excellentanalogy is that fluorescence microscopy yields information equivalent to theknowledge that two students are present in a large lecture hall. It doesn't offer theresolution necessary to determine whether the students are in the sameclassroom or, better yet, if they are sitting in adjacent desks.

Typical fluorescence microscopy techniques rely upon the absorption by a

fluorophore of light at one wavelength (excitation), followed by the subsequentemission of secondary fluorescence at a longer wavelength. The excitation andemission wavelengths are often separated from each other by tens to hundreds ofnanometers. Labeling of cellular components, such as the nuclei, mitochondria,cytoskeleton, the Golgi apparatus, and membranes, with specific fluorophoresenables their localization within fixed and living preparations. By simultaneouslylabeling several sub-cellular structures with individual fluorophores havingseparated excitation and emission spectra, specialized fluorescence filtercombinations can be employed to examine the proximity of labeled moleculeswithin a single cell or tissue section. Using this technique, molecules that arecloser together than the optical resolution limit appear to be coincident (and aresaid to co-localize). This apparent spatial proximity implies that a molecularassociation is possible. In most cases, however, the normal diffraction-limitedfluorescence microscope resolution is insufficient to determine whether aninteraction between biomolecules actually takes place.

Co-localization measurements are suggestive at best and misleading at worst,especially considering that many signaling pathways use the same cellularstructure, as for example, clathrin-coated pits that are utilized for internalization ofmany receptor complexes. The knowledge that two molecules or proteins are infact adjacent, and not just residing in the same neighborhood, provides a

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fact adjacent, and not just residing in the same neighborhood, provides asignificantly more reliable determination of their potential for interactions. Thetime-honored technique of electron microscopy has ample resolution to meet theneeds of high-precision localization, but simply lacks the precise labelingmethodology necessary to produce reliable results. Furthermore, many co-localization techniques are generally applied for use within fixed cells, whichprecludes the highly desirable dynamic measurements attainable by assays inliving cells. Fluorescence imaging with multi-color fluorescent proteins readilypermits experimentation with live cells, which are necessary for assays oftransient interaction, but the approach suffers from having a relatively poor spatialresolution limited to approximately 200 nanometers.

Limitations in determination of the spatial proximity of protein molecules can beovercome by applying Förster (or Fluorescence) Resonance Energy Transfer(FRET) microscopy techniques. FRET occurs between two appropriatelypositioned fluorophores only when the distance separating them is 8 to 10nanometers or less. Thus, FRET is well-suited to the investigation of proteininteractions that occur between two molecules positioned within severalnanometers of each other. Over the past ten years, FRET approaches havegained popularity due to the rise in applications requiring genetically targeting ofspecific proteins and peptides using fusions to green fluorescent protein (GFP)and its mutated derivatives. FRET between two spectrally distinct fluorescentproteins (known as FP-FRET) has been widely applied for two distinctly separateexperimental techniques, as discussed below. Presented in Figure 1 is aJablonski energy diagram illustrating the coupled excited state transitionsinvolved between the donor emission and acceptor absorbance in FRET.Absorption and emission transitions are represented by straight vertical arrows(blue, green and red), while vibrational relaxation is indicated by wavy yellowarrows. The coupled transitions are drawn with dashed lines that suggest theircorrect placement in the Jablonski diagram should they have arisen from photon-mediated electronic transitions. In the presence of a suitable acceptor, the donorfluorophore can transfer excited state energy directly to the acceptor withoutemitting a photon (illustrated by a violet arrow in Figure 1). The resultingfluorescence sensitized emission has characteristics similar to the emissionspectrum of the acceptor.

One of the major obstacles to the widespread implementation of FRETinvestigations in living cells has been the lack of suitable methods for labelingspecific intracellular proteins with the appropriate fluorophores. The recentdevelopment of fluorescent proteins possessing a wide array of spectral profilesand the increasing sophistication of protein chimeras (fusions as well asbiosensors) has resulted in a number of potential fluorescent protein pairs that areuseful in FRET experiments. Application of fluorescent proteins to FRET involveseither integrating a selected pair into a biosensor (a single genetically-encodedconstruct) or conducting intermolecular measurements between two separateproteins, each fused to a different fluorescent protein. The latter approach hasbeen employed to image a variety of protein interactions, includingoligomerization of receptors and elucidating the functions of transcription factors.However, conducting FRET assays on independently expressed protein chimerasis far more difficult due to the variable stoichiometry that inevitably occurs whenseparate fluorescent entities are expressed in living cells. Regardless of thedifficulty, experiments of this nature can yield informative results when appropriatecontrols are installed and the investigation is conducted with exacting precision.

Fluorescent Protein Biosensors

Fluorescent protein biosensors have found widespread utility in reporting on adiverse array of intracellular processes. By creatively fusing pairs of fluorescentproteins to biopolymers that perform critical functions involved in various aspectsof physiological signaling, research scientists have developed a host of newmolecular probes that are useful for optical live-cell imaging of importantprocesses such as calcium wave induction, cyclic nucleotide messenger effects,pH changes, membrane potential fluctuations, phosphorylation, and intracellularprotease action. An alternative, but quite useful, strategy to biosensor constructioninvolves modifications to the fluorescent protein backbone structure itself, either tosplit the peptide into individual units that are combined in vivo to producefluorescence (a technique termed Bi-Molecular Fluorescence Complementation;;BiFC) or to join the natural amino and carboxy termini together and create aninsertion site within the molecule for a sensor peptide.

The first fluorescent protein biosensor was a calcium indicator named cameleon,constructed by sandwiching the protein calmodulin and the calcium calmodulin-binding domain of myosin light chain kinase (M13 domain) between enhanced



binding domain of myosin light chain kinase (M13 domain) between enhancedblue and green fluorescent proteins (EBFP and EGFP). In the presence ofincreasing levels of intracellular calcium, the M13 domain binds the calmodulinpeptide to produce an increase in FRET between the fluorescent proteins.Unfortunately, this sensor was hampered by a very low dynamic range (a 1.6-foldincrease in fluorescence) and was difficult to visualize due to lack of brightnessand poor photostability of EBFP. Improved versions using the same templateincorporated the cyan and yellow variants ECFP and EYFP to yield higher signallevels, and even better results were obtained when YFP derivatives (termedcamgaroos) were generated by inserting the calcium-sensitive peptides at the

beginning of the seventh beta-strand in the fluorescent protein backbone. Sensorpeptides situated at this unusual position are quite well tolerated with regards tomaintaining high levels of fluorescence. Yet another strategy takes advantage ofthe unique barrel structure common in fluorescent proteins to reconfigure the endsof the protein by linking the natural N and C termini and creating a new start codonin one of several locations within the central region of the structure (usually in theloops). Termed circularly permuted fluorescent proteins, these structurallymodified derivatives can be fused to calmodulin and M13 to produce excellentcalcium biosensors.

Calcium biosensors were quickly followed by genetic indicators for pH,phosphorylation, and protease activity. Two general approaches can be used toadapt fluorescent proteins as sensors of pH. The first relies on the fluorescencesensitivity of EGFP (pKa = 5.9) and EYFP (pKa = 6.5) to acidic environmentscoupled to the relative insensitivity of other proteins, such as ECFP (pKa = 4.7) orDsRed (pKa = 4.5). Fusions of EGFP or EYFP with a less sensitive fluorescentprotein create a ratiometric probe that can be used to measure the acidity ofintracellular compartments. The second approach relies on protonation changesof native (wild-type) GFP that result in a shift in the bimodal spectral profiles of thenative protein. A class of probes named pHluorins, derived from wtGFP, exhibitsa shift in the excitation peak from 470 to 410 nanometers as the pH decreases.Dual-emission pH sensors have also been developed, which have peaks in thegreen and blue spectral regions. Although unable to report kinase activity in realtime, phosphorylation biosensors consist of a peptide containing aphosphorylation motif from a specific kinase and a binding domain for aphosphopeptide sandwiched between two FRET-capable fluorescent proteins.When the biosensor is phosphorylated by the kinase, the phosphopeptide bindingdomain binds to the phosphorylated sequence, thus invoking or destroying FRET.This simple strategy has proven to generate robust and highly specific biosensors.As with many other biosensors, the major drawback is reduced dynamic range.

Perhaps the most widely used biosensor design to screen new or improved FRET

pairs involves a protease cleavage assay (see Figure 2). The simple motifconsists of two fluorescent proteins linked together by a short peptide thatcontains a consensus protease cleavage site. In general, the sensor exhibits verystrong energy transfer that is completely abolished upon cleavage of the linkersequence. Because the technique usually features high dynamic range levels, itcan be used to screen new cyan and green FRET donors with yellow, orange,and red acceptors. The largest family of protease biosensors incorporates acleavage site sensitive to one of the caspase family of proteases, which enablesthe sensor to be examined during induction of apoptosis. Over the past severalyears, a large number of novel biosensors using both sensitized fluorescent



years, a large number of novel biosensors using both sensitized fluorescentproteins and FRET pairs have been reported. Despite the continued limitations indynamic range of FRET sensors using ECFP and EYFP derivatives, this strategyhas been widely adopted, probably due to the simplicity of ratiometricmeasurements and ease of probe construction. New strategies will no doubtemerge using more advanced fluorescent protein combinations that serve toincrease the dynamic range and other properties of this highly useful class ofprobes.

Basic Principles of FRET

The fundamental mechanism of FRET involves a donor fluorophore in an excitedelectronic state, which may transfer its excitation energy to a nearby acceptorfluorophore (or chromophore) in a non-radiative fashion through long-rangedipole-dipole interactions. The theory supporting energy transfer is based on theconcept of treating an excited fluorophore as an oscillating dipole that canundergo an energy exchange with a second dipole having a similar resonancefrequency. In this regard, resonance energy transfer is analogous to the behaviorof coupled oscillators, such as a pair of tuning forks vibrating at the samefrequency or a radio antenna. In contrast, radiative energy transfer requiresemission and re-absorption of a photon and depends on the physical dimensionsand optical properties of the specimen, as well as the geometry of the containerand the wavefront pathways. Unlike radiative mechanisms, resonance energytransfer can yield a significant amount of structural information concerning thedonor-acceptor pair.

Resonance energy transfer is not sensitive to the surrounding solvent shell of afluorophore, and thus, produces molecular information unique to that revealed bysolvent-dependent events, such as fluorescence quenching, excited-statereactions, solvent relaxation, or anisotropic measurements. The major solventimpact on fluorophores involved in resonance energy transfer is the effect onspectral properties of the donor and acceptor. Non-radiative energy transferoccurs over much longer distances than short-range solvent effects and thedielectric nature of constituents (solvent and host macromolecule) positionedbetween the involved fluorophores has very little influence on the efficacy ofresonance energy transfer, which depends primarily on the distance between thedonor and acceptor fluorophore.

The phenomenon of FRET is not mediated by photon emission, and furthermore,does not even require the acceptor chromophore to be fluorescent. In mostapplications, however, both donor and acceptor are fluorescent, and theoccurrence of energy transfer manifests itself through quenching of donorfluorescence and a reduction of the fluorescence lifetime, accompanied also by anincrease in acceptor fluorescence emission. The theory of resonance energytransfer was originally developed by Theodor Förster and, in honor of hiscontribution, has recently been named after him. The Förster theory shows thatFRET efficiency (E) varies as the inverse sixth power of the distance between thetwo molecules (denoted by r):

E = 1/[1 + (r/R ) ]

where R(0) is the characteristic distance where the FRET efficiency is 50 percent,

FRET 0 6



where R(0) is the characteristic distance where the FRET efficiency is 50 percent,which can be calculated for any pair of fluorescent molecules (this variable is alsotermed the Förster radius and is discussed below in greater detail). The FRETefficiency of a theoretical fluorophore pair (enhanced cyan and yellow fluorescentproteins) is graphically demonstrated in Figure 3(a). Because of the inverse sixthpower dependence on the distance between the two molecules (r), the curve hasa very sharp decline. For distances less than R(0), the FRET efficiency is close tomaximal, whereas for distances greater than R(0), the efficiency rapidlyapproaches zero. The useful range for measuring FRET is indicated by the redshaded region in Figure 3(a) with limits of 0.5 and 1.5 x R(0). FRET can beeffectively used as a molecular ruler for those distances close to R(0), andindeed FRET has been adapted for such purposes in structural biology by usingprecision spectroscopic approaches. For most applications in cell biology,however, the signal-to-noise ratios available limit FRET experiments to a morebinary readout. In effect, a measurement will often be only able to distinguishbetween high-FRET and low-FRET, or simply between the presence andabsence of FRET.

As previously discussed, R(0) can be readily calculated for any pair of fluorescentmolecules. The value of R(0) in an aqueous (or buffered) solution is determinedby a fairly simple equation with the well-established input parameters:

R = [2.8 x 10 × Κ × Q × Ε × J(λ)] nanometers

where Κ(2) or kappa squared represents the orientation factor between the twofluorophore dipoles (see Figure 3(b) for a summary of angles used to calculate theorientation factor), Q(D) is the donor quantum yield, Ε(A) is the maximal acceptorextinction coefficient in reciprocal moles per centimeter, and J(λ) is the spectraloverlap integral (see Figure 4) between the normalized donor fluorescence, F(D)(λ), and the acceptor excitation spectra, E(A)(λ), according to the equation:

J(λ) = F (λ) × Ε (λ) × λ dλ

Although the mathematics may appear complicated, most of the parameters areconstants that are easily found in the literature. The two most important terms thatgenerally require further explanation are Κ(2) and J(λ), the overlap integral. Theorientation angle variable (Κ(2)) simply indicates that the FRET coupling dependson the angle between the two fluorophores in much the same manner as theposition of a radio antenna can affect its reception. If the donor and acceptor arealigned parallel to each other, the FRET efficiency will be higher than if they areoriented perpendicular. This degree of alignment defines Κ(2). Although Κ(2) canvary between zero and 4, it is usually assumed to be 2/3, which is the averagevalue integrated over all possible angles. For almost any realistic situation Κ(2) isclose to 2/3, and there is usually nothing that an investigator can do to adjust thisvalue (although some have attached fluorescent proteins rigidly to their targetproteins of interest, which could lead to dramatic effects). The overlap integral,J(λ), is the region of overlap between the two spectra, as illustrated in Figure 4.The other parameters that can affect FRET are the quantum yield of the donor andthe extinction coefficient of the acceptor. Thus, in order to maximize the FRETsignal, the researcher must choose the highest quantum yield donor, the highestabsorbing acceptor, and fluorophores having significant overlap in their spectralprofiles. This theory has been repeatedly verified by experiment, and there are noother mechanisms to maximize FRET for non-aligned fluorescent probes.

0 17 2 D A 1/6

D A 4



It should be noted that each of the parameters discussed above affects the Försterradius calculation only by the sixth power. Thus, doubling of the donor quantumyield results in only a 12.5 percent change in R(0). Because almost allfluorophores used in FRET imaging experiments have high quantum yields(greater than 0.5) and extinction coefficients (greater than 50,000), the range ofpossible Förster radius values is limited to between 4 and 6 nanometers, andmost FRET pairs have an average value of R(0) ~ 5 nanometers. Given that FRETefficiency is strongly dependent on the distance separating the FRET pair as wellas the relative orientation of the fluorophores, FRET can be used to detectchanges in protein-protein interactions that arise from changes in the affinitybetween the two proteins or changes in the conformation of their binding. It isworth repeating that, for most FRET imaging applications in cell biology,experiments generally differentiate only between two states (FRET and no FRET)and additional information is necessary to aide in the molecular interpretation ofthe observed FRET changes.

Factors Affecting FRET Measurements

In practice, a wide spectrum of issues can complicate and/or compromise FRETmeasurements, ultimately leading to ambiguous or meaningless results. One ofthe principal issues is that the donor and acceptor fluorophores might exhibitsignificantly different brightness levels when imaged together. Although in theorythis discrepancy should not be a problem, however in practice because mostinstruments can measure only a limited dynamic range, dual fluorophore imagingmay result in one channel that is saturated (for the brighter fluorophore) while theother channel is dominated by systematic noise (for the dimmer fluorophore).Thus, whenever possible it is best to use a donor and acceptor that are ofcomparable brightness.

Another factor that can limit the detection of FRET is a donor-to-acceptorstoichiometry that lies outside the range between 10:1 and 1:10. This factor canbe a serious limitation in FRET measurements of protein-protein interactions inwhich one partner might be in excess concentration. The primary problem is themeasurement of a small level of FRET against a background of fluorescent labelsthat are not undergoing FRET. Due to the fact that there is really nothing that canbe done to improve this situation, a host of possible protein-protein interactionexperiments falling into this category are simply unsuitable for examination byFRET techniques. For the fluorescent protein biosensors described above, whichare constructed with only a single donor and acceptor, the stoichiometry is fixedand guaranteed to be 1:1;; thus, this issue never arises and the signal levelremains constant, regardless of the biosensor concentration.

The presence of bleed-through (also termed crosstalk and crossover) and crossexcitation between spectrally overlapping fluorophores are also important issuesthat can hamper FRET investigations (see Figure 5). In some cases, the acceptorcan be directly excited with light in the wavelength region chosen to excite thedonor (Figure 5(a)). Additionally, fluorescence from the donor can leak into thedetection channel for the acceptor fluorescence, especially when the emissionspectral profiles of the donor and acceptor exhibit significant overlap (Figure 5(b)).Because these two sources of crosstalk arise from the photophysics of organicfluorophores and will most certainly be present for any FRET pair, they must beaddressed when FRET is measured. Choosing fluorophores that are well-separated spectrally is an excellent mechanism to reduce crosstalk. However, inmost cases the increased spectral separation also reduces the overlap integral,(J(λ)), which in practice usually translates to a reduced ability to detect the FRETsignal.

Finally, the level of a FRET signal can be reduced if the two fluorophores are notproperly aligned (for instance, having a Κ(2) value of approximately zero) or if theyare simply not positioned within the Förster radius (greater than 6 nanometers). Asan example, if two labeled proteins interact, but the fluorescent labels are locatedon opposite sides of the complex, then there might not be a detectable FRETsignal, even though the proteins of interest are bound. In general practice, thistype of false negative is quite common, especially with fluorescent protein FRETpartners. Often, several labeling strategies are required before a sufficient andreliable FRET signal is detected. However, each of the issues described abovecan be mitigated (or partially so) by an informed choice of the fluorophore pair tobe used prior to making vector constructs or conducting synthetic labelingexperiments.



Presented in Figure 5 is the overlap in the excitation and emission spectral

profiles of ECFP and mVenus, currently one of the most preferred fluorescent

proteins pairs for FRET investigations. These two proteins exhibit considerable

overlap in both the excitation (Figure 5(a)) and emission (Figure 5(b)) spectra.

Direct excitation of the FRET acceptor (mVenus;; red curve) can be significant

depending on the wavelength used for excitation of the donor (ECFP;; cyan curve

or mCerulean;; blue curve) due to the higher extinction coefficient of the yellow

protein as compared to the cyan proteins. This overlap is especially problematic

when ECFP is used as the donor and can be partially offset by using CFP

variants with high extinction coefficients, such as mCerulean. Note that the

excitation curves in Figure 5(a) are drawn to scale in order to reflect the

differences in extinction coefficient between the yellow and cyan proteins.

Excitation at 458 nanometers produces a much higher level of mVenus excitation

crosstalk than does excitation at 405 or 440 nanometers. The broad fluorescence

emission spectrum of ECFP (Figure 5(b)) exhibits considerable intensity overlap

throughout the region of mVenus emission.

FRET Techniques in Cell Biology Applications

Investigators employing fluorescent protein biosensors, or attempting to match the

stoichiometry of fluorescent probes fused to separate interacting targets, should

use as many different FRET analysis techniques as feasible to establish the

methodology for a given experiment. Such an effort is warranted because each of

the fluorescent protein FRET pairs exhibits a distinct pathology that complicates

its use, necessitating a clear understanding of the optical microscopy parameters

applied to measuring the relatively small signal differences produced in most

FRET assays. Once the system and the possible results are well established,

then the simplest approaches can be used for ongoing procedures. The list of

techniques that have been developed to image FRET is quite extensive. In

general, all of the existing strategies for measuring FRET can be applied to

fluorescent protein experiments but, on the basis of practical considerations, five

general approaches have proven particularly useful:

Sensitized Emission - Two-channel imaging using an algorithm thatcorrects for excitation and emission crosstalk

Acceptor Photobleaching - Also known as donor dequenching, thistechnique measures increased donor emission when the acceptor is

photobleached

Fluorescence Lifetime Imaging Microscopy (FLIM) - Fluorescentprotein (or other fluorophore) donor lifetime measurement changes

Spectral Imaging - Exciting at one or two wavelengths and measuringthe complete spectral profiles of donor and acceptor

Fluorescence Polarization Imaging - Measure polarization parallel andperpendicular to excitation with high signal-to-noise

Each of the FRET approaches listed above has strengths and weaknesses. For

example, on one hand, two-channel imaging is the simplest method, but requires

the most complicated set of controls. On the other hand, FLIM can yield an

unambiguous measurement of FRET efficiency, but instrumentation to measure

the nanosecond lifetimes is expensive and not yet widely available to most cell

biology laboratories.

Sensitized Emission



Also commonly referred to as two-color ratio imaging with controls, sensitizedemission is perhaps the simplest method of imaging FRET. The donor fluorophore

is excited by a specific wavelength (in a widefield or confocal microscope), and

the signal is collected by using emission filters chosen for the donor fluorescence

and the acceptor fluorescence. In the (unrealistic) absence of crosstalk between

the excitation and fluorescence of the two fluorophores, then sensitized emission

would be a perfect method. However, crosstalk between fluorescent proteins is a

significant problem and extensive control experiments are usually required to

establish the presence or absence of FRET. Thus, it is difficult to obtain

quantitatively accurate FRET data with this approach. Sensitized emission is

relatively simple to configure on a widefield fluorescence microscope, available in

many laboratories, but the necessary control experiments require considerable

image processing to subtract crosstalk components, which significantly increases

the noise level and uncertainty in the measurements.

A variety of corrective approaches have been developed for sensitized emission

FRET imaging. The basic concept involves the use of different filter combinations

with multiple samples that contain: only the donor, only the acceptor, and the

putative FRET sample with both the donor and acceptor. The emission values

from these samples permit the investigator to determine the amount of expected

crosstalk in both excitation and emission channels and to subtract it from the

FRET measurement. In theory this approach works nicely, but the requirement for

image processing unfortunately increases the noise level in all of the images.

Thus, if the FRET signal is minute, then it may be difficult to measure FRET using

this approach.

Despite the difficulties mentioned above, sensitized emission measurements can

be useful for rapid dynamic experiments in which FRET signals are large due to

the ability to acquire both images simultaneously. Sensitized emission is a

particularly attractive technique when examining fluorescent protein biosensors

where the FRET dynamic range is large and the stoichiometry of the donor and

acceptor is fixed in a 1:1 ratio. A good example is the protease biosensor

illustrated in Figure 2. This chimera has been engineered to have a high FRET

efficiency that drops essentially to zero when the peptide linker is enzymatically

cleaved. The result is a large and readily measurable FRET change that

demonstrates a specific protease activity at a given time and region within the

living cell.

Acceptor Photobleaching

Although limited to only a single measurement, acceptor photobleaching (or donor

dequenching) is also a simple technique that often yields excellent results. The

underlying concept takes advantage of the fact that donor fluorescence is

quenched during FRET because some of the donor fluorescence energy is

channeled to the acceptor. Photobleaching the acceptor fluorophore irreversibly

eliminates the quenching effect and increases the level of donor fluorescence. If

FRET is occurring between the fluorophores, the donor fluorescence must

increase when the acceptor is removed. In general, it is important to ensure that

acceptor photobleaching does not degrade the donor fluorescence, and that the

acceptor is photobleached to approximately 10 percent of its initial value. Both of

these constraints are easily met with a laser scanning confocal microscope, but



these constraints are easily met with a laser scanning confocal microscope, butcan also be accomplished with widefield or spinning disk microscopes equippedwith a specialized illumination system.

The acceptor photobleaching technique has the advantage of being verystraightforward, quantitative, and performed using only a single sample. TheFRET efficiency can be calculated by subtracting the donor intensity in thepresence of the acceptor from its intensity after photobleaching the acceptor, andthen normalizing this value to the donor intensity after bleaching. The primarydisadvantage is that acceptor photobleaching is destructive and can be used onlyonce per cell, limiting its application to those experiments not involved withdynamic measurements. Furthermore, photobleaching is a relatively slow processthat often requires several minutes or longer. Nevertheless, it is almost alwaysworthwhile to perform an acceptor photobleaching measurement at the end of anexperiment, regardless of whichever methods are being used to assay FRET.

Presented in Figure 6 are examples of sensitized emission and acceptorphotobleaching FRET assays using live cell imaging. Figure 6(a) illustrates ahuman cervical carcinoma epithelial cell (HeLa line) expressing a cameleonbiosensor comprised of mCerulean and mVenus fused together with anintervening calcium-sensitive peptide containing calmodulin and the M13 domain(described above). Prior to the addition of a calcium-inducing agent (ionomycin),excitation of the cell with 440-nanometer illumination produces cyan fluorescenceindicating a lack of FRET between the cyan and yellow fluorescent proteins(Figure 6(a)). Upon addition of ionomycin, time lapse two-color ratio imaging(sensitized emission) records a calcium wave traversing the cytoplasm as thebiosensor responds with an increase in the level of FRET between the fluorescentproteins (Figures 6(b) and 6(c);; FRET is pseudocolored yellow-red). The Africangreen monkey kidney cell (COS-7 line) in Figure 6(d)-(f) was labeled with thesynthetic cyanine dyes, Cy3 (Figure 6(d);; green) and Cy5 (Figure 6(e);; red),conjugated to cholera toxin B-subunit and targeting the plasma membrane. Withinthe membrane, the close proximity of the two dyes produces a high level of FRET.Photobleaching Cy5 in a selected region of the cell (white box in Figure 6(e))increases the donor dequenching (increase in green fluorescence in Figure 6(f))in a corresponding area when viewing fluorescence in the donor channel only.

Fluorescence Lifetime Imaging Microscopy (FLIM)

Lifetime measurements are by far the most rigorous method for determining FRET;;furthermore, they are also less prone to crosstalk artifacts due to the fact that onlythe donor fluorescence is monitored. All fluorescent molecules exhibit anexponential decay in their fluorescence on a nanosecond timescale, and the rate

of this decay is sensitive to environmental variables that quench the fluorescence.Thus, the basic concept of FLIM is somewhat related to that of acceptorphotobleaching. The donor fluorescence is quenched by the FRET interaction,and the amount of quenching can be determined by measuring the decrease influorescence decay time of the donor in the presence of FRET. In this manner,FLIM provides an unambiguous value of the FRET efficiency. Among theadvantages of combined FLIM-FRET measurements is their insensitivity to directacceptor excitation artifacts as well as the fact that fluorescent donors can becoupled to acceptors that are not themselves fluorescent. Both of these aspectsserve to expand the number of useful fluorescent protein FRET pairs available toinvestigators.

FLIM has several limitations that prevent it from being the dominant approach inFRET imaging. Primarily, measurements in the nanosecond lifetime region arecomplex and the instrumentation is expensive to obtain and maintain. Also, this



complex and the instrumentation is expensive to obtain and maintain. Also, this

type of sophisticated equipment is not widely available. Additionally, FLIM is

usually among the slower imaging methodologies, potentially requiring several

minutes to acquire each image, which limits its utility in many FRET experiments.

These constraints might be lifted in the future as more user-friendly and faster

turnkey commercial systems are developed by the manufacturers. Another

significant downside is that the lifetimes of fluorescent proteins in live cells often

display multi-exponential decays that require more comprehensive data analysis

for quantitative FRET assays. Furthermore, localized environmental factors, such

as autofluorescence or a change in pH, can also shorten the measured

fluorescence lifetime, leading to artifacts. Thus, a great deal of care must be taken

in the interpretation of FLIM-FRET data in living cells.

Spectral Imaging

The technique of spectral imaging is a variation on the sensitized emission FRET

detection method, but instead of acquiring data through two individual channels,

the entire emission spectrum containing both donor and acceptor fluorescence is

collected upon excitation of the donor. Recording of the entire spectrum is a

typical approach used for spectroscopy experiments, but is a relatively recent

addition to the tool palette in widefield and confocal microscopy. The concept

centers on the premise that collection of the entire fluorescence spectrum enables

overlapping spectra to be separated by using not just the emission peaks but also

the distinct shapes of the spectral tails. In gathering the spectrum from both the

donor and acceptor fluorophore, it is possible to determine the relative levels of

donor and acceptor fluorescence.

The spectral imaging technique requires specialized equipment, but excellent

systems are readily available on many commercial confocal microscopes and can

be added onto a conventional fluorescence microscope at modest cost.

Conducting a quantitative analysis of the level of crosstalk due to direct excitation

of the acceptor, or the use of two excitation wavelengths in confocal microscopy,

permits an accurate determination of the amount of FRET. The principal drawback

of this approach is the reduced signal-to-noise ratio associated with acquiring the

complete spectrum rather than collecting it through two channels with a filter-

based system. As more commercial systems are being developed and installed,

however, the application of spectral imaging in FRET assays is increasing. In the

near future, it is quite possible that spectral imaging will become one of the

primary methods for performing FRET imaging experiments.

Illustrated in Figure 7(a) are changes in the donor lifetime decay (mCerulean

fluorescent protein) of a pseudo-FRET biosensor consisting of mCerulean and

mVenus fluorescent proteins fused together with a 10-amino acid linker. The blue

decay curve shows the lifetime observed in cells expressing mCerulean alone,

whereas the red decay curve presents the mCerulean lifetime obtained when cells

express the concatenated proteins. Note the decrease in mCerulean lifetime when

the protein is involved in resonance energy transfer. The area between the curves

represents the energy that is transferred through FRET from mCerulean (donor) to

mVenus (acceptor) in the FRET pairing. The emission profile from 450 to 650

nanometers of mCerulean-mVenus in the same pseudo-biosensor when excited

at 405 nanometers in live cells is depicted by the red curve in Figure 7(b). Transfer

of energy from mCerulean to mVenus results in a substantial emission peak at

529 nanometers (the mVenus emission maximum), with a much lower value

(approximately 25 percent) at 475 nanometers, the peak emission wavelength of

mCerulean. After photobleaching mVenus with a 514-nanometer laser and

repeating the spectral scan, the emission profile shifts to lower wavelengths and

closely resembles the spectrum of mCerulean in the absence of a FRET partner.

The difference in intensities at 475 and 529 nanometers of these spectral profiles

is related to the FRET efficiency between the coupled proteins.

Polarization Anisotropy Imaging

Measurements of fluorescence polarization offer particular advantages for high-

contrast discrimination of fluorescent protein FRET. The concept is based on the

fact that excitation with polarized light selects a population of fluorescent

molecules whose absorption vectors are aligned parallel to the polarization vector

of the exciting light. Immediately after excitation, most of the fluorescence

emission will remain polarized parallel to the excitation so that the fluorescence

can be considered anisotropic in terms of polarization. The anisotropy will

disappear if the molecules rotate during the nanosecond fluorescent lifetime.

However, because fluorescent proteins are large and rotate slowly, their

fluorescence does not depolarize to any great degree during the measurement

time course. If FRET occurs between two fluorescent proteins that are slightly



time course. If FRET occurs between two fluorescent proteins that are slightly

misaligned, then the polarized fluorescence emission will emerge at a different

angle (from the excitation vector), which simulates a rotation of the fluorescent

protein.

The primary strength of this approach is the ease of measuring fluorescence

polarization parallel and perpendicular to the excitation vector with high signal-to-

noise. Because polarization anisotropy data can be acquired rapidly and with

minimal image processing requirement, the technique is well-suited for

applications in high-content screening. Direct excitation of the acceptor must be

avoided, however, because it can decrease the donor signal and reduce the

signal-to-noise ratio of the measurement. In addition, although this technique is

superb in discriminating between the presence and absence of FRET, it is not a

good approach for differentiating between strong and weak FRET. Finally,

polarization can be degraded in high numerical aperture objectives, so polarized

FRET experiments should be limited to imaging with objectives having a

numerical aperture of 1.0 or less.

Presented in Figure 8 is a graphic illustration of polarization anisotropy using

fluorescent proteins as a model system. When a randomly oriented population of

fluorescent proteins (Figure 8(a)) is excited with linearly polarized light (cyan

wave), only those molecules whose absorption dipole vector is oriented parallel to

the polarization azimuth are preferentially excited. Emission from properly

oriented fluorescent proteins can be observed as a signal using an analyzer that

is also parallel to the excitation light polarization vector (green wave). The

resulting anisotropy, which is an indicator of the degree of orientation, can be

determined by measuring and comparing the emission intensity through the

vertically and horizontally oriented analyzers. The anisotropy signal level will

decrease if the fluorescent protein rotates in the timescale of the experiment

(Figure 8(b)) or if it transfers excitation energy due to FRET to a neighboring

protein (Figure 8(c)) having a different orientation. As described above, due to the

fact that resonance energy transfer can occur far more rapidly than molecular

rotation for large fluorescent protein molecules, depolarization due to FRET can

be readily distinguished from the loss of anisotropy that occurs during rotation.

Considerations for Using Fluorescent Proteins in FRET

The choice of suitable probes for examining FRET in living cells is limited.

Synthetic fluorophores, ideal for resonance energy transfer investigations in fixed

cells, are difficult to administer and target in live cells. Likewise, quantum dots can

be utilized to label membrane components for examination of phenomena on the

exterior of a cell, but they too are unable to penetrate the membrane and,

consequently, of little use in intracellular compartments such as the nucleus,

mitochondria, or endoplasmic reticulum. Genetically encoded fluorescent proteins

currently represent the best candidates for high-resolution imaging of FRET in live

cells, as evidenced by the volume of literature that is published in this arena on a

yearly basis. However, many of the typical artifacts that are encountered in

measuring FRET with synthetic fluorophores and quantum dots are particularly

acute when applied to fluorescent proteins. For example, contrary to the 30-40

nanometer bandwidth of emission spectral profiles in synthetics, those in

fluorescent proteins range from approximately 60 nanometers to 100 nanometers,

often leading to significant overlap when attempting to segregate donor and

acceptor fluorescence. The broad spectra of fluorescent proteins also limit the

number of probes that can be used together in FRET and other types of imaging



number of probes that can be used together in FRET and other types of imagingexperiments. Furthermore, fluorescent proteins exhibit a wide variation inbrightness levels. For example, one of the most popular donor proteins, ECFP,has fivefold less brightness than its common yellow acceptor partner, EYFP.

Surrounding the fluorescent protein chromophore is a 220+ amino acidpolypeptide wound into a three-dimensional cylindrical structure approximately2.4 by 4.2 nanometers in dimension (termed a beta-barrel or beta-can), andcomposed of extensively hydrogen-bonded polypeptide beta-sheets that surroundand protect a central alpha-helix containing the chromophore (see Figure 9). Theends of the barrel are capped with semi-helical peptide regions that serve to blockentry of ions and small molecules. The interior of the protein is so tightly packedwith amino acid side chains and water molecules that there is little room fordiffusion of oxygen, ions, or other intruding small molecules that manage to passthrough the ends of the barrel. These favorable structural parameters, which arepartially responsible for the resilient photostability and excellent performance offluorescent proteins, also contribute to a reduction in FRET efficiency. The largesize of the barrel effectively shields adjacent fluorescent protein chromophoreswith peptide residues (to a limiting close approach distance of 2 to 3 nanometers;;indicated by the red line in Figure 9), resulting in a reduction of the maximumFRET efficiency to approximately 40 percent of the theoretical value. Regardless,the numerous benefits of using fluorescent proteins for live cell FRET imaging faroutweigh the costs.

Compounding the high degree of spectral bandwidth overlap and size problemsthat occur with fluorescent proteins is their tendency to oligomerize. Almost all ofthe fluorescent proteins discovered to date display at least a limited degree ofquaternary structure, as exemplified by the weak tendency of native Aequoreavictoria green fluorescent protein and its derivatives to dimerize whenimmobilized at high concentrations. This tendency is also verified by the stricttetramerization motif of the native yellow, orange, and red fluorescent proteinsisolated in reef corals and anemones. Oligomerization can be a significantproblem for many applications in cell biology, particularly in cases where thefluorescent protein is fused to a host protein that is targeted at a specificsubcellular location. Once expressed, the formation of dimers and higher orderoligomers induced by the fluorescent protein portion of the chimera can produceatypical localization, disrupt normal function, interfere with signaling cascades, orrestrict the fusion product to aggregation within a specific organelle or thecytoplasm. This effect is particularly marked when the fluorescent protein is fusedto partners which themselves participate in natural oligomer formation. Fusionproducts with proteins that form only weak dimers (in effect, most Aequoreavictoria variants) may not exhibit aggregation or improper targeting, provided thelocalized concentration remains low. However, when weakly dimeric fluorescent



localized concentration remains low. However, when weakly dimeric fluorescentproteins are targeted to specific cellular compartments, such as the plasmamembrane, the localized protein concentration can, in some circumstances,become high enough to permit dimerization. This can be a particular concernwhen conducting intermolecular FRET experiments, which can yield complexdata sets that are sometimes compromised by dimerization artifacts. On the otherhand, the naturally occurring weak dimerization in Aequorea proteins can be, insome cases, utilized to increase the FRET signal in biosensors that otherwisewould exhibit limited dynamic range.

Toxicity is an issue that occurs due to excessive concentrations of synthetic

fluorophores and the over-expression or aggregation of poorly localizedfluorescent proteins. Furthermore, the health and longevity of optimally labeledmammalian cells in microscope imaging chambers can also suffer from a numberof other deleterious factors. Foremost among these is the light-induced damage(phototoxicity) that occurs upon repeated exposure of fluorescently labeled cells toillumination from lasers and high-intensity arc-discharge lamps. In their excitedstate, fluorescent molecules tend to react with molecular oxygen to produce freeradicals that can damage subcellular components and compromise the entire cell.Fluorescent proteins, due to the fact that their fluorophores are buried deep withina protective polypeptide envelope, are generally not phototoxic to cells. Indesigning FRET experiments, fluorescent protein combinations that exhibit thelongest possible excitation wavelengths should be chosen in order to minimizedamage to cells by short wavelength illumination, especially in long-term imagingexperiments. Thus, rather than creating fusion products and biosensors with blueor cyan fluorescent proteins (excited by ultraviolet and blue illumination,respectively), variants that emit in the yellow, orange, and red regions of thespectrum would be far more ideal.

Investigators should take care to perform the necessary control experiments whenusing new fluorescent protein biosensors and cell lines to ensure that cytotoxicityand phototoxicity artifacts do not obscure FRET results or other importantbiological phenomena. In some cases, lipophilic reagents induce deleteriouseffects that may be confused with fluorescent protein toxicity during imaging in celllines following transient transfections. Oligomeric fluorescent proteins (discussedabove) from reef corals have a far greater tendency to form aggregates (combinedwith poor subcellular localization) than do the monomeric jellyfish proteins, butimproperly folded fusion products can occur with any variant. Recently, afluorescent protein capable of generating reactive oxygen species (ROS) uponillumination with green light has been reported as an effective agent forinactivation of specific proteins by chromophore-assisted light inactivation (CALI).Appropriately named KillerRed, this genetically encoded photosensitizer iscapable of killing both bacteria and eukaryotic cells upon illumination in themicroscope. Previous studies on EGFP phototoxicity indicate that even throughthe chromophore is capable of generating singlet oxygen, the fluorescent proteinis relatively inefficient as a photosensitizer. However, prolonged illumination ofcells expressing EGFP and its variants can result in physiological alterations andeventual cell death, a definite signal of the potential for phototoxicity in long-termimaging experiments.

In live-cell experiments, fluorescent proteins are highly advantageous forextended imaging due to their reduced rate of photobleaching when compared tosynthetic fluorophores. Although there is a high degree of uncorrelated variabilitybetween fluorescent proteins in terms of photostability, most variants are useful forshort-term imaging (from 1 to 25 captures), while several of the more photostableproteins can be employed in time-lapse sequences that span periods of 24 hoursor longer (in which hundreds to thousands of images are gathered). The long termstability of any particular protein, however, must be investigated for everyillumination scenario (widefield, confocal, multiphoton, swept-field, etc.) becausedifferences in photostability are often observed with the same protein whenillumination is produced by an arc-discharge lamp versus a laser system. Thus, interms of photostability, the selection of fluorescent proteins is dictated bynumerous parameters, including the illumination conditions, the expressionsystem, and the effectiveness of the imaging setup.

Potential Fluorescent Protein FRET Partners

Over the past few years, a wide variety of new fluorescent protein variants havebeen developed and refined to feature emission profiles spanning a 200-nanometer range (from approximately 450 nanometers to 650 nanometers), thusfilling many gaps to provide potentially useful FRET partners in every color class.



Recent advances in developing proteins in the blue (440 nanometers to 470

nanometers) and cyan (470 nanometers to 500 nanometers) spectral regions have

yielded several new probes that may be of use for imaging and FRET

investigations. Three protein engineering groups have reported improved blue

Aequorea fluorescent protein variants that feature significantly higher brightness

and photostability compared to EBFP. Named Azurite, SBFP2 (strongly

enhanced blue FP), and EBFP2 (see Table 1), these proteins offer the first real

hope for successful long-term imaging of live cells in the blue spectral region, and

all have significant applicability to combination with EGFP and derivatives in

FRET biosensors. The brightest and most photostable of the new blue reporters,

EBFP2, exhibits typical GFP-like behavior in fusions and has been demonstrated

to be an excellent FRET donor for proteins in the green spectral class. All of the

blue fluorescent proteins can be readily imaged in a fluorescence microscope

using standard DAPI filter sets or proprietary BFP sets available from aftermarket

manufacturers.

Fluorescent proteins in the cyan spectral region have been widely applied as

FRET donors when paired with yellow-emitting proteins, and were dominated by

variants of the original Aequorea ECFP until the introduction of a monomeric teal-colored reporter, known as mTFP1. Teal fluorescent protein exhibits higher

brightness and acid stability compared to Aequorea CFPs, and is far morephotostable. The high emission quantum yield of mTFP1 (see Table 1) provides

an excellent alternative to the cyan derivatives, mECFP and mCerulean, as a

FRET donor when combined with either yellow or orange fluorescent proteins.

Additional investigations have produced useful proteins in the cyan spectral class.

Among the improved cyan fluorescent proteins that have recently been

introduced, CyPet and the enhanced cyan variant termed Cerulean show the

most promise as candidates for fusion tags, FRET biosensors, and multicolor

imaging. Cerulean is at least 2-fold brighter than ECFP and has been

demonstrated to significantly increase contrast as well as the signal-to-noise ratio

when coupled with yellow-emitting fluorescent proteins, such as Venus (see

below), in FRET investigations. The CFP variant named CyPet (from the acronym:

Cyan fluorescent Protein for energy transfer) was derived through a unique

strategy utilizing fluorescence-activated cell sorting (FACS) to optimize the cyan

and yellow pairing for FRET. CyPet is about half as bright as EGFP and two-thirds

as bright as Cerulean, but expresses relatively poorly at 37 degrees Celsius.

However, CyPet has a more blue-shifted and narrower fluorescence emission

peak than CFP, which greatly increases its potential for multicolor imaging.

The introduction of beneficial folding mutations into monomeric variants of ECFP

has resulted in the production of new variants featuring enhanced brightness,

folding efficient, solubility, and FRET performance. Termed super CFPs

(SCFPs), the new reporters are significantly brighter than the parent protein when

expressed in bacteria and almost two-fold brighter in mammalian cells. These

high-performance probes should be useful both for routine fusion tags and in

creating new CFP-YFP FRET biosensors exhibiting high dynamic range. Another

new monomeric cyan reporter, TagCFP, was derived from a GFP-like protein from

the jellyfish Aequorea macrodactyla. Specific details about the protein areunavailable in the literature, but it is commercially available as mammalian

cloning vectors and fusions from Evrogen. TagCFP is reported to be brighter than

ECFP and Cerulean, but of similar acid resistance. Another cyan-emitting protein,

Midoriishi-Cyan (abbreviated MiCy) was originally designed as the donor in a

new FRET combination with the monomeric Kusabira Orange (mKO;; see Table

1) to generate a biosensor with high spectral overlap (Förster distance of 5.3).

This protein features the longest absorption and emission wavelength profiles

(472 and 495 nanometers, respectively) reported for any probe in the cyan

spectral region. The high molar extinction coefficient and quantum yield exhibited

by MiCy render the protein of equal brightness to Cerulean.

Properties of Selected Fluorescent Protein FRET Pairs

Protein Pair

Donor

Excitation

Maximum

(nm)

Acceptor

Emission

Maximum

(nm)

Donor

Quantum

Yield

Acceptor

Molar

Extinction

Coefficient

Förster

Distance

(nm)

Brightness

Ratio

EBFP2-

mEGFP383 507 0.56 57,500 4.8 1:2

ECFP-EYFP 440 527 0.40 83,400 4.9 1:4

Cerulean-

Venus440 528 0.62 92,200 5.4 1:2

MiCy-mKO 472 559 0.90 51,600 5.3 1:2

TFP1-

mVenus492 528 0.85 92,200 5.1 1:1

CyPet-YPet 477 530 0.51 104,000 5.1 1:4.5



CyPet-YPet 477 530 0.51 104,000 5.1 1:4.5

EGFP-mCherry 507 610 0.60 72,000 5.1 2.5:1

Venus-mCherry 528 610 0.57 72,000 5.7 3:1

Venus-tdTomato 528 581 0.57 138,000 5.9 1:2

Venus-mPlum 528 649 0.57 41,000 5.2 13:1

Table 1

The best current choice for live-cell imaging of FRET reporters in the green colorclass (500 nanometers to 525 nanometers) is the GFP derivative Emerald, whichhas properties similar to its EGFP parent. Emerald contains the F64L and S65Tmutations featured in EGFP, but the variant also has four additional pointmutations that improve folding, expression at 37 degrees Celsius, and brightness.Recently, a new addition to the green spectral region has been coinedsuperfolder GFP, which is brighter and more acid resistant than either EGFP orEmerald and has similar photostability. Therefore, the superfolder variant shouldbe an excellent candidate for fusions with mammalian proteins and theconstruction of FRET biosensors, especially those that demonstrate foldingproblems with standard GFP derivatives. Another brightly fluorescent reporter,which may be a good FRET candidate, is termed Azami Green and has beenisolated from the stony coral Galaxeidae and demonstrated to mature rapidlyduring expression in mammalian cell lines. In addition, two bright, monomericGFP reporters obtained through site-directed and random mutagenesis incombination with library screening in cyan proteins have been reported. Derivedfrom the Clavularia coral genus, mWasabi is a potential alternative green-emitting FRET partner for blue fluorescent proteins due to negligible absorbanceat 400 nanometers and lower where blue variants are often excited. The newgreen reporter is commercially available (Allele Biotechnology) and should beparticularly useful for two-color imaging in conjunction with long Stokes shiftproteins (such as T-Sapphire) as well as a localization tag in fusions withtargeting proteins. A derivative of TagCFP, named TagGFP, is a bright andmonomeric green variant having an absorption maximum at 482 nanometers andemission at 505 nanometers. TagGFP, which is only slightly brighter than EGFP,is available as cloning vectors and fusion tags from Evrogen, but has not beenthoroughly characterized in literature reports.

Yellow fluorescent proteins (525 nanometers to 555 nanometers) are among themost versatile genetically-encoded probes yet developed and should providecandidates acting as both donors and acceptors in FRET pairings. The variantsknown as Citrine and Venus are currently the most useful proteins in this spectralclass (see Table 1), but neither is commercially available. Another variant, namedafter the birthstone Topaz, is available from Invitrogen and has been of service infusion tag localization, intracellular signaling, and FRET investigations. A newmember of the Evrogen “Tag” commercial series of localization reporter proteins,TagYFP, is a monomeric jellyfish (Aequorea macrodactyla) derivative that isslightly less bright than EYFP, but an order of magnitude more photostable.Similar to its partners, TagYFP (emission peak at 524 nanometers) has not beencharacterized in the literature, but can be purchased as mammalian cloningvectors or fusion tags.

During the same fluorescence-activated cell sorting investigation that led to thegeneration of CyPet (discussed above), the evolutionary optimizedcomplementary FRET acceptor, termed YPet, was also obtained. Named after itsproficiency in FRET (YFP for energy transfer), YPet is the brightest yellow variantyet developed and demonstrates reasonable photostability. The resistance toacidic environments afforded by YPet is superior to Venus and other YFPderivatives, which will enhance the utility of this probe in biosensor combinationstargeted at acidic organelles. However, although the optimized CyPet-YPetcombination should be the preferred starting point in the development of newFRET biosensors, there remains a serious doubt as to the origin of YPet'sincreased performance, which is likely due simply to enhanced dimerization withits co-evolved partner, CyPet. Likewise, the suitability of CyPet and YPet in fusiontags for localization experiments, bimolecular complementation analysis, andother applications has yet to be established. Both proteins exist in solution asweak dimers, but presumably can be converted to true monomers using theA206K mutation that has worked so well with other Aequorea variants (althoughthis apparently destroys their advantages in FRET).

Orange fluorescent proteins, all of which have all been isolated from coral reefspecies, have the potential to be useful in a variety of FRET imaging scenarios.



species, have the potential to be useful in a variety of FRET imaging scenarios.Perhaps the most versatile of these is monomeric Kusabira Orange, a proteinoriginally derived as a tetramer from the mushroom coral Fungia concinna(known in Japanese as Kusabira-Ishi). A monomeric version of Kusabira Orange(abbreviated mKO) was created by introducing over 20 mutations through site-directed and random mutagenesis. The monomer (commercially available fromMBL International) exhibits similar spectral properties to the tetramer and has abrightness value similar to EGFP, but is slightly more sensitive to acidicenvironments than its parent. The photostability of this reporter, however, isamong the best of any protein in all of the spectral classes, making mKO anexcellent choice for long-term imaging experiments. Furthermore, the emissionspectral profile is sufficiently well separated from cyan fluorescent proteins toincrease the FRET efficiency in biosensors incorporating mKO, and the probe isuseful in multicolor investigations with a combination of cyan, green, yellow, andred probes.

Illustrated in Figure 10 are spectral profiles of ECFP (Figure 10(a)), EGFP (Figure10(b)), EYFP (Figure 10(c)), and mOrange (Figure 10(d)), each acting as a FRETdonor to mPlum, a far-red emitting fluorescent protein acceptor. As the emissionspectral profiles of the donors shift to longer wavelengths (from cyan to orange),the spectral overlap (filled gray region) and calculated Förster distance (R(0))increases correspondingly. Similarly, the excitation and emission crosstalk (redand blue hatched regions, respectively) also increases as the wavelengthseparation between the donor and acceptor emission peaks decreases. Note thatECFP and mPlum exhibit only a limited degree of overlap in the excitation spectraand virtually none in the emission spectra. In contrast, there is a high level of bothexcitation and emission crosstalk when mOrange is paired with mPlum. As thefluorescent protein color palette continues to expand, a wide spectrum of newFRET pairs should become readily available to investigators.

The mRFP1 derivative, mOrange, is slightly brighter than mKO, but has less than10 percent the photostability, thus severely compromising its application forexperiments that require repeated imaging. However, mOrange remains one ofthe brightest proteins in the orange spectral class and is still an excellent choicewhere intensity is more critical than long-term photostability. In addition, combinedwith the green-emitting T-Sapphire, mOrange is a suitable alternative to CFP-YFPproteins as a FRET pair to generate longer wavelength biosensors, and can becoupled with proteins in other spectral regions for multicolor investigations. Animproved version of mOrange (named mOrange2) featuring dramaticallyincreased photostability is now available. A bright new monomeric orange protein,named TagRFP has recently been introduced as a candidate for localization andFRET studies and may prove to be effective in a wide number of biosensorconstructs. The brightest fluorescent protein in any spectral class is the tandemversion of dimeric Tomato (dTomato), an orange derivative that was one of theoriginal Fruit proteins. The Tomato protein contains the first and last seven aminoacids from GFP on the N- and C- termini in an effort to increase the tolerance tofusion proteins and reduce potential artifacts in localization as well as enhancethe possibility of its use in FRET biosensors. A tandem-dimer version (effectively



a monomer) was created by fusing two copies, head-to-tail, of dTomato with a 23-amino acid linker. Due to the presence of twin chromophores, the resultingtdTomato is extremely bright and has exceptional photostability. The majordrawback in the use of this protein is the larger size (twice that of a monomericprotein), which may interfere with fusion protein packing in some biopolymers.

The search for an ideal red-emitting fluorescent protein has long been the goal forlive-cell and whole animal imaging using FRET biosensors and fusions, primarilydue to the requirement for probes in this spectral region in multicolor imagingexperiments as well as the fact that longer excitation wavelengths generate lessphototoxicity and can probe deeper into biological tissues. To date, a widespectrum of potentially useful red probes has been reported (emission at 590nanometers to 650 nanometers), many of which still suffer from some degree ofthe obligatory quaternary structure bestowed by their species of origin. Unlike thejellyfish proteins, most of the native and genetically engineered variants of coralreef proteins mature very efficiently at 37 degrees Celsius. Extensive mutagenesisresearch efforts, including newly introduced methodology, have successfully beenapplied in the search for yellow, orange, red, and far-red florescent proteinvariants that further reduce the tendency of these potentially efficacious biologicalprobes to self-associate while simultaneously pushing emission maxima towardlonger wavelengths. The result has been improved monomeric proteins thatfeature increased extinction coefficients, quantum yields, and photostability,although no single variant has yet been optimized by all criteria.

The red mFruit proteins, mApple, mCherry and mStrawberry (emission peaks at592, 610, and 596 nanometers, respectively), have brightness levels ranging from50 percent to 110 percent of EGFP, but mApple and mCherry are far morephotostable than mStrawberry and are the best probe choices to replace mRFP1for long-term imaging experiments. Further extension of the mFruit protein spectralclass through iterative somatic hypermutation has yielded two new fluorescentproteins with emission wavelength maxima of 625 and 649 nanometers,representing the first true far-red genetically engineered probes. The most

potentially useful probe in this pair was named mPlum, which has a rather limitedbrightness value (10 percent of EGFP), but excellent photostability. Thismonomeric probe should be useful in combination with variants emitting in thecyan, green, yellow, and orange regions for multicolor imaging experiments andas a biosensor FRET partner with green and yellow proteins, such as Emeraldand Citrine (see Figure 10). Several additional red fluorescent proteins showingvarying degrees of promise have been isolated from the reef coral organisms. Theapplication of site-specific and random mutagenesis to TurboRFP variants,followed by screening for mutations exhibiting far-red fluorescence, resulted in adimeric protein named Katushka (emission maxima of 635 nanometers).Although only two-thirds as bright as EGFP, Katushka exhibits the highestbrightness levels of any fluorescent protein in spectral window encompassing 650to 800 nanometers, a region that is important for deep tissue imaging. Introductionof the four principal Katushka mutations into TagRFP generated a monomeric, far-red protein named mKate that has similar spectral characteristics. Thephotostability of mKate is reported to be exceptional and the protein displaysbrightness similar to that of mCherry, which makes it an excellent candidate forlocalization and FRET experiments in the far-red portion of the spectrum.

Despite significant advances in expanding the fluorescent color palette into theorange, red, and far-red regions of the spectrum, cyan and yellow Aequoreaderivatives remain the most useful pairing scenario for developing usefulbiosensors. This unforeseen discrepancy occurs because most of the orange andred coral-derived proteins exhibit a relatively broad absorption spectral profilehaving a long excitation tail that extends into the violet and cyan regions, thusproducing direct acceptor excitation. Another factor that might come into play isthe relative maturation rates of the fused fluorescent protein partners. In mostcases, variants derived from Aequorea proteins mature far more rapidly thanthose obtained from reef corals so it is possible that immature acceptors contributeto the poor sensitized emission exhibited by many of the coral-derived proteins. Inaddition, the broad adsorption spectra of the orange and red proteins, combinedwith the reduced quantum yields of the monomeric versions, likely render themdifficult for use in FRET. Future success in fluorescent protein FRET experimentaldesign will focus on, among other factors, matching the maturation rates of thepaired proteins.

Conclusions

Although FRET experiments based on the ubiquitous fluorescent protein familyoffer tremendous potential to reveal molecular dynamics in living cellular systems,



offer tremendous potential to reveal molecular dynamics in living cellular systems,

as yet there is not a perfect FRET pair. The optimized versions of CFP and YFP

still provide the most effective pair for general use, although better combinations

may loom over the horizon. Likewise, there is no perfect technique with which to

measure FRET, although the approaches described above all have strengths that

can be leveraged depending on the particular experimental situation under

investigation. As more optimized fluorescent proteins become available, including

bright red variants that might be appropriate as acceptors for GFP or YFP donors,

FRET using fluorescent proteins should become even more useful for protein-

protein interaction investigations in live cells. As discussed, the broad absorption

spectra of the current palette of red fluorescent proteins, in addition to the lower

quantum yields of the monomeric versions, make these candidates difficult to

employ in FRET. However, the rapid pace of fluorescent protein improvements

lends optimism that such proteins will be available in the near future and will help

to further revolutionize this new approach to elucidating intracellular biochemical

mechanisms.

BACK TO FLUORESCENCE MICROSCOPY

Contributing Authors

Gert-Jan Kremers and David W. Piston - Department of Molecular

Physiology and Biophysics, Vanderbilt University, 702 Light Hall, Nashville,

Tennessee, 37232.

Michael W. Davidson - National High Magnetic Field Laboratory, 1800

East Paul Dirac Dr., The Florida State University, Tallahassee, Florida,

32310.

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