what is fret

8/13/2019 What is FRET

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What is FRET?

Fluorescence resonance energy transfer (FRET) is a process involving the radiationless

transfer of energy from a donor fluorophore to an appropriately positioned acceptor

fluorophore. FRET can occur when the emission spectrum of a donor fluorophore

significantly overlaps (>30%) the absorption spectrum of an acceptor (seeFigure 1),

provided that the donor and acceptor fluorophores dipoles are in favorable mutual

orientation. Because the efficiency of energy transfer varies inversely with the sixth

power of the distance separating the donor and acceptor fluorophores, the distance over

which FRET can occur is limited to between 1-10 nm. When the spectral, dipole

orientation, and distance criteria are satisfied, illumination of the donor fluorophore

results in sensitized fluorescence emission from the acceptor, indicating that the tagged

proteins are separated by


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approach of fluorescence resonance energy transfer (FRET) microscopy, this

information can be obtained from single living cells with nanometer resolution.

FRET microscopy relies on the ability to capture weak and transient fluorescent signals

efficiently and rapidly from the interactions of labeled molecules in single living or

fixed cells. The occurrence of FRET signal (sensitized signal) can be verified by

acquiring the two-emission signal bands of the double labeled cells excited with donor

wavelength. If FRET occurs the donor channel signal will be quenched and the acceptorchannel signal will be sensitized or increased. In principle, the measurement of FRET in

a microscope can provide the same information that is available from the more common

macroscopic solution measurements of FRET; however, FRET microscopy has the

additional advantage that the spatial distribution of FRET efficiency can be visualized

throughout the image, rather than registering only an average over the entire cell or

population. Because energy transfer occurs over distances of 1-10 nm, a FRET signal

corresponding to a particular location within a microscope image provides an additional

magnification surpassing the optical resolution (~0.25 mm) of the light microscope.

Thus, within a voxel of microscopic resolution, FRET resolves average donor-acceptor

distances beyond the microscopic limit down to the molecular scale. This is one of the

principal and unique benefits of FRET for microscopic imaging: not only colocalization

of the donor- and acceptor-labeled probes within ~0.09 mm2 can be seen, but intimate

interactions of molecules labeled with donor and acceptor can be demonstrated. Several

FRET techniques exist based on wide-field, confocaland 2p microscopy as well as

FRET/FLIM, each with its own advantage and disadvantage. All FRET microscopy

systems require neutral density filters to control the excitation light intensity, a stable

excitation light source (Hg or Xe or combination arc lamp; UV, Visible or Infrared

lasers), a heated stage or a chamber to maintain the cell viability and appropriate filter

sets (excitation, emission, and dichroic) for the selected fluorophore pair. It is important

to carefully select filter combinations that reduce the spectral bleed through (SBT) to

improve the signal-to-noise (S/N) ratio for the FRET signals.

FRET PAIR

The widely used donor and acceptor fluorophores for FRET studies come from a class

of autofluorescent proteins, called Green Fluorescent Proteins (GFPs). The

spectroscopic properties that are carefully considered in selecting GFPs as workable

FRET pairs include: sufficient separation in excitation spectra for selective stimulation

of the donor GFP, an overlap (>30%) between the emission spectrum of the donor and

the excitation spectrum of the acceptor to obtain efficient energy transfer and reasonable

separation in emission spectra between donor and acceptor GFPs to allow independent

measurement of the fluorescence of each fluorophore. GFP-based FRET imaging

methods have been instrumental in determining the compartmentalization and

functional organization of living cells and for tracing the movement of proteins inside

cells.

There are number of combination of FRET pair can be used depending on the biological

applications. Selected popular FRET pair fluorophore are - CFP-YFP, CFP-dsRED,

BFP-GFP, GFP or YFP-dsRED, Cy3-Cy5, Alexa488-Alexa555, Alexa488-Cy3, FITC-

Rhodamine (TRITC), YFP-TRITC or Cy3, etc. You can find the FRET Pair Spctra from

here.


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Problems with FRET microscopy Imaging

One of the important conditions for FRET to occur is the overlap of the emissionspectrum of the donor with the absorption spectrum of the acceptor. As a result of

spectral overlap, the FRET signal is always contaminated by donor emission into the

acceptor channel and by the excitation of acceptor molecules by the donor excitation

wavelength (see Figure 1). Both of these signals are termed spectral bleed-through

(SBT) signal into the acceptor channel. In principle, the SBT signal is same for 1p- or

2p-FRET microscopy. In addition to SBT, the FRET signals in the acceptor channel

also require correction for spectral sensitivity variations in donor and acceptor c

hannels, autofluorescence, and detector and optical noise, which contaminate the FRET

signal. How to correct the contaminated signals is explained in data process part.

FIGURE1


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WIDE-FIELD FRET (W-FRET) MICROSCOPY

Any fluorescence microscope (inverted or upright) can be converted to W-FRET

microscopy. There are number of papers listed in the literature for various protein

studies using W-FRET system (Day, 1998; Day et al, 2003; Gordon et al, 1998; Jovinand Arndt-Jovin, 1989; Kam et al, 1995; Kraynov et al., 2000; Periasamy and Day,

1999; Varma and Mayor, 1998). For W-FRET it is advisable to use a single dichroic to

acquire the donor (D) and acceptor (A) images for the donor excitation wavelength in

the double-labeled specimen. This can be achieved by using excitation and emission

filter wheels in the microscope system. This option helps to reduce any spatial shift of

donor and acceptor channel images, since the processed FRET image is obtained

through pixel-by-pixel calculation as described in the FRET data analysis section.

Even though W-FRET microscopy is the simplest and most widely used technique,

there is a major limitation to W-FRET in that the emission signals originating from

above and below the focal plane contribute to out-of-focus signals that reduce the

contrast and seriously degrade the image. Digital deconvolutionmicroscopy in the W-FRET system helps to localize the proteins at different optical sections, but this requires

intensive computational process to remove the out-of-focus information from the optical

sectioned FRET images (Periasamy and Day, 1998 and 1999). For protein interactions

taking place homogeneously over a wider area of a cell (e.g. nucleus), W-FRET is an

entirely suitable technique.

Confocal Theory

Confocal Microscopy is rapidly gaining acceptance as an important technology owing

to its capability to produce images free of out-of-focus information. In a conventional

epi-fluorescence microscope, the entire object is exposed to excitation light and the

emission collected by high NA objectives comes from throughout the specimen,whether above or below the focal plane. This seriously degrades the image by reducing

the contrast and sharpness. In confocal microscopy out-of-focus information (blur) is

removed. In addition, confocal microscopy provides a significant improvement in

lateral resolution and the capacity for direct, non-invasive serial optical sectioning of

intact, thick living specimens.

Confocal Microscopy was introduced in 1957. Most confocal microscopes are of two

types: (1) stage-scanning (SSCM), and (2) laser scanning (LSCM). The SSCM is

assembled on an epi-illuminated microscope employing a stationary laser as an

excitation source, a photomultiplier as the detector and a specimen holder (stage) which

moves and thus allows the specimen to be "rapidly" scanned in the X-Y plane. A pin-

hole in the emission path coupled with a high NA (1.4) objective lens removes out-of-

focus information and sharply improves the contrast. However, SSCM requires a

relatively long period of time (~10 sec) to acquire a single image. Thus the SSCM can

be used satisfactorily for fixed specimens or microelectronic circuits, but not for the live

specimens where dynamic events are occuring.


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Laser Scanning Confocal Microscopy

Many investigators designed confocal microscopes for use with live specimens to image

dynamic events in which a fixed microscope stage is scanned by a laser beam using a

rotating disk or mirror galvanometers. LSCM generates a clear, thin image (512 X 512)free from out-of-focus information within 2 or 3 seconds. A single diffraction-limited

spot of light is projected on the specimen using a high numerical aperture objective lens

and the light reflected or fluoresced by the specimen is collected by the objective and

focused upon a pinhole aperture and the signal detected by a photomultiplier. Light

originating from above or below the image plane strikes the walls of the pinhole and is

not transmitted to the detector. To generate a two-dimensional image, the laser beam is

scanned across the specimen pixel-by-pixel. To produce an image using LSCM, the

laser beam must be moved in a regular two-dimensional raster scan across the specimen

and the instantaneous response of the photomultiplier must be displayed with equivalent

spatial resolution and relative brightness at all points on the synchronously scanned

phosphor screen of a CRT monitor.

For a three-dimensional projection of a specimen one needs to collect a series of images

at different Z-axis planes. The vertical spatial resolution is approximately 0.5um for a

40X 1.3 NA objective. Three-dimensional image reconstruction can be accomplished

with many commercially available software systems.

The photomultiplier tube (PMT) used in LSCM has highly desirable characteristics

compared to video cameras: (1) stability; (2) low noise; (3) very large dynamic range (>

1 million fold); (4) sensitivity; (5) wide range of spectral resonse; (6) rapid response;

and (7) small physical size. The PMT has a low quantum efficiency (QE) of about 30%

and in the red wavelengths about 3% and produces very low background noise signal.

The alternative, a cooled-PIN photodiode, has a QE of 60-80% but an equivalent noise

level of about 100 photons/pixel so that it is not useful for weak signals. The optimumselection of pinhole size is important in the compromise between intensity (brightness)

and thickness of the slice observed. For instruments with variable pinholes, an optimum

pinhole diameter should be determined empirically to provide the best combination of

brightness and slice thickness.

Laser scanning confocal FRET (C-FRET) microscopy overcomes the limitation of out-

of focus information owing to its capability of rejecting signals from outside the focal

plane and acquire the signal in real-time (Kenworthy et al, 2000; Pozo et al, 2002;

Wallrabe et al, 2003). This capability provides a significant improvement in lateral

resolution and allows the use of serial optical sectioning of the living specimen (Pawley,

1995; Lemasters et al, 2001). By selecting appropriate filter combinations one can

configure any commercially available confocal microscopy system for FRET imaging.


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Disadvantage of LSCM

A disadvantage of this technique is that the wavelengths available for excitation ofdifferent fluorophore pairs is limited to standard lasers lines. Standard laser lines do

allow C-FRET to be used for a number of fluorophore combinations including CFP-

YFP or ds-RED, GFP-Rhodamine or Cy3, FITC or Alexa488-Cy3, Alexa488-Alexa555

and Cy3-Cy5 (Day et al., 2003; Elangovan et al., 2003; Kenworthy et al, 2000; Mills et

al, 2003; Periasamy, 2001; Wallrabe et al, 2003).

Also, in one-photon wide-field or confocal microscopy, illumination occurs throughout

the excitation beam path, in an hourglass-shaped pattern. This results in absorption

along the excitation beam path, giving rise to substantial fluorescence emission both

below and above the focal plane. Excitation of other focal planes contributes to

photobleaching and photodamage in the specimen planes that are not being involved in

imaging. This can be ameliorated by Multi-photon/2-photon microscopy.

How to collect FRET Images

In Confocal FRET imaging, we select the appropriate filters and high sensitivity

photomultiplier tubes (PMTs) to acquire donor and acceptor images. It is important to

note that appropriate average power should be used to reduce photobleaching.

The background subtraction of the image is important to remove the autofluorescence,

detector and optical noise. The SBT correction should be implemented as discussed in

the data process part. Seven imagesare required. In brief, (1) single labeled donor cells

should be excited with donor molecule excitation wavelength and D- and A- channel

images are acquired. (2) Single labeled acceptor molecule should be excited with donorand acceptor wavelength and the A- channel images are acquired. (3) Double labeled

(D+A) cell should be excited with donor excitation wavelength and the D- and A-

channel images are acquired. Acceptor excitation wavelength will be used to excite the

D+A labeled cells and collect the A-channel image. These seven images are used

toprocess to obtain the processed or precision FRET (PFRET) image.

The laser power for excitation for donor and the acceptor may be different. But once

you adjust the donor laser power (say 10%) and that should be used whenever you use

the donor excitation wavelength. The same way the acceptor excitation wavelength, if

you use, say 5% or 10% for the acceptor excitation wavelength then, the same acceptor

power (5% or 10%) should be used whenever you use the acceptor excitation

wavelength.

The same theory for PMT gain adjust for donor and acceptor emission. It is important

not to saturate the pixel intensity.


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Seven Images Required for FRET Data Process

SymbolFluorophone

or Sample

ExcitationFilter

Excitation

Wavelength

Emission Filter

Emission

Wavelength

Meaning

a Donor Only Donor Donor

Signal from a donor only

specimen using donor

excitation and donor emission

filter set.

b Donor Only Donor Acceptor

Signal from a donor only


excitation and acceptor

emission filter set.

cAcceptor

OnlyDonor Acceptor

Signal from an acceptor only




dAcceptor

OnlyAcceptor Acceptor

Signal from an acceptor only

specimen using acceptor



eDonorand

Acceptor

Donor Donor

Signal from donor-and-

acceptor specimen using donor

excitation and donor emissionfilter set.

fDonor and

AcceptorDonor Acceptor


acceptor specimen using donorexcitation and acceptor


gDonorand

AcceptorAcceptor Acceptor


acceptor specimen using

acceptor excitation and

acceptor emission filter set.


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FRET Data Analysis

What is SBT?

One of the important conditions for FRET to occur is the overlap of the emission

spectrum of the donor with the absorption spectrum of the acceptor . As a result of

spectral overlap, the FRET signal is always contaminated by donor emission into the

acceptor channel(DSBT) and by the excitation of acceptor molecules by the donor

excitation wavelength(ASBT) (see Figure left). Both of these signals are termed spectral

bleed-through (SBT) signal into the acceptor channel. In principle, the SBT signal is

same for 1p- or 2p-FRET microscopy. In addition to SBT, the FRET signals in the

acceptor channel also require correction for spectral sensitivity variations in donor and

acceptor channels, autofluorescence, and detector and optical noise, which contaminate

the FRET signal.

Algorithm

The details of the algorithm to remove SBT and the relevant biological applicationshave been listed in the literature (Elangovan et al. 2003; Mills et al., 2003; Wallrabe et

al., 2003; www.circusoft.com).

In brief, to remove the spectral bleed-through or cross-talk for 1p- or 2p-FRET, seven

imagesare acquired. Our approach works on the assumption that the double-labeled

cells and single-labeled donor and acceptor cells, imaged under the same conditions,

exhibit the same SBT dynamics. The hurdle we had to overcome was the fact that we

had three different cells (D, A, and D+A), where individual pixel locations cannot be

compared. What could be compared, however, were pixels with matching fluorescence

levels. Our algorithm follows fluorescence levels pixel-by-pixel to establish the level ofSBT in the single-labeled cells, and then applies these values as a correction factor to

the appropriate matching pixels of the double-labeled cell.

We use a,b,c,d,e,f,g to represent the seven images. The following equations are used to

remove the spectral bleed-through signal from the FRET channel image.

Where j is the jth range of intensity, m is the number of pixel in a and d, n is the number

of pixel in e and g, DSBTi is the donor bleed-through of the pixel (i) in f, ai is the


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intensity of pixel i, so is bi, and ei, k is the number of range, DSBT is the total donor

bleed-through, ASBTi is the acceptor bleed-through of the pixel (i) in f, ci is the

intensity of pixel i, so is di, and gi.k is the number of range, ASBT is the total acceptor

bleed-through.

The precision FRET (PFRET) is calculated using following equation where uFRET

represents uncorrected FRET which is image f:

PFRET=uFRET-DSBT-ASBT

Energy transfer efficiency(E)

Conventionally, energy transfer efficiency (E) is calculated by ratioing the donor image

in the presence (IDA) and absence (ID) of acceptor. When using the algorithm asdescribed, we indirectly obtained the IDimage by using the PFRET image (Elangovan et

al., 2003). ID=IDA+PFRET where IDAis image e. The efficiency calculation is shown in

following equation:

E=1-[IDA/(IDA+PFRET)]

It is important to note that there are a number of other processes involved in the excited

state during energy transfer. The new efficiency (En) is calculated by generating a new

ID image by including the detector spectral sensitivity of donor and acceptor channel

and the donor quantum yield with PFRET signal as shown in following equation

Software

Based on the algorithm described above, our center developed a user friendly PFRET

software to process FRET data. Details about the software, please click hereor check

circusoft web site. You can look at the seven images we collected from Wide-Field

Microscope. The pseudo color, PFRET image , efficiency image and histogram were

produced by the PFRET software.


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Filter configurations for confocal image acquisition forselected fluorophore pairs.

Fluorophore Excitation wavelengh (nm) Emission filter (nm)

Alexa 488 or GFP Argon 488 515/30 or 535/50

Cy3 or Phod-2 Green HeNe 543 590/70

CFP Argon 457 485/30

dsRED1 HeNe 543 590/70

CFP Argon 457 485/30

YFP Argon 514 528/50

Cy3 Green HeNe 543 590/70

Cy5 HeNe 633 or HeNe 594 660LP


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Confocal FRET Images

MDCK cells were internalized with pIgA-R ligand-Alexa488 and pIgA-R ligand-Cy3,

for 4h at 17degrees C, from the apical and basolateral PM, respectively. Fluorescence

confocal images were taken at ~3.5microns below the apical PM to evaluate FRET.

Schematic representation of a polarized MDCK epithelial cell, showing apical and

basolateral PM. The enclosed star is the basolaterally internalized Cy3-pIgA-R-ligand

complex. The open star is the apically internalized Alexa488-pIgA-R-ligand complex.

Co-localization occurs in the apical endosome in the sub-apical region.


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Confocal FRET Images

This is a two-color fluorescent RNA in situ on an early neural groove stage Xenopus

laevis embryo. The green channel shows the expression of epidermal keratin (xk81) in

the prospective epidermis. The red channel shows the expression of Xslug a marker for

prospective neural crest cells. Notice the overlap (yellow) between the neural crest and

epidermal markers

The C-FRET systems were used for CFP-RFP pair to visualize the C/EBP proteins in a

single cell using Nikon PCM2000 laser scanning confocal microscopy. The excitation

wavelength used to excite the donor molecule (CFP-C/EBP) was 457 nm from an argon

laser. A HeNe green laser line (543 nm) was used to acquire the acceptor (RFP-C/EBP)

image. Using the excitation wavelength 457 nm both the donor and acceptor images


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were acquired from the cells expressed both the proteins (CFP-RFP-C/EBP). The

energy transfer signals were processed from these images using the software by

removing the entire noise and bleed-through signal as shown in Figure 2F (CFPem-

480/30; RFPem-610/60). The respective histograms below the figures clearly

demonstrate the noise signal in the acceptor channel (Figure 2AH) and the processed

true FRET signals (Figure 2FH). Adapted from A. Periasamy, Journal of Biomedical

Multiphoton Introduction

General Information

Conventional one-photon confocal laser scanning microscopy (CLSM) often provides

high resolution but is limited in sensitivity and spatial resolution by background flare

noise resulting from "out-of-focus" fluorescence. In CLSM, the repeated scanning of

UV light greatly reduces cell or tissue viability. The two- or three-photon excitation

laser scanning microscope (nonlinear microscope), however, circumvents this limitation

by using two or three red-wavelength photons to obtain both sensitivity and depth

resolution without a confocal aperture. The MEFIM technique considerably reduces

autofluorescence and photodamage above and below the focal plane, and the volume ofthe focal plane depends on a diffraction spot created by the objective lens.

Two-photon absorption was theoretically predicted by Goppert-Mayer in 1931, and it

was experimentally observed for the first time in 1961 by using a ruby laser as the light

source (Kaiser and Garrett, 1961). The original idea of two-photon fluorescence

scanning microscopy was proposed by Sheppard et al. (1977) and was experimentally

demonstrated for biological imaging by Winfried Denk and Watt Web (1990).

Physics of Two-Photon Excitation

The probablility of two-photon absorption depends on the co-localization of two

photons within the absorption cross section of the fluorophore, and the rate of excitationis proportional to the square of the instantaneous intensity. Two-photon excitation is

made possible by the very high local instantaneous intensity that is provided by a

combination of diffraction-limited focusing of a single laser beam in the specimen plane

and the temporal concentration of a femtosecond (fsec) mode-locked laser. The two-

photon advantage is roughly proportional to the inverse excitation duty cycle, for

example, a 100,000-fold improvement over CW illumination is achieved by using 100-

fsec pulses at 76 MHz repetition rate. The use of such short pulses and small duty cycles

is, in fact, essential for image acquisition in a reasonable time while using "biologically

tolerable" power levels.

Advantages of MEFIM

(i) In one-photon excitation CLSM, photobleaching occurs well above and below the

focal (volume) plane; in MEFIM, the photobleaching is considerably reduced, and

illumination of laser light occurs only at the focal plane.

(ii) Repeated scanning on the specimen in CLSM, particularly with UV light, induces

rapid photoisomerization and high background autofluorescence. MEFIM reduces these

complications, providing better penentration at infrared wavelengths and thus

prolonging cell viability during image acquisition.


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(iii) The CLSM technique requires special UV optics for the UV excitation probe.

MEFIM uses conventional microscope optics.

(iv) In CLSM, the emission wavelength is close to the excitation wavelength (about 50-

200nm). In MEFIM, the fluorescence emission occurs at a wavelength substantially

shorter than the excitation wavelength.

Multiphoton FRET Images

Localization of BFP- and RFP-C/EBP protein expressed in mouse 3T3 cells using 2p-

FRET microscopy. The doubly expressed cells (BFP-RFP-C/EBP) were excited by 740


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nm and the donor (A) and acceptor (B) images of proteins localized in the nucleus of a

single living cell were acquired by single scan (slow scan). As explained in the text, the

bleed-through (or cross talk) correction was implemented and then ratioed to obtain a

FRET image is shown in C. The respective histograms are shown below the images (D,

E, F). The higher gray level intensity distribution in Figure C compared to E indicates

the importance of bleed-through correction and ratioing of the corrected D and A (A/D)

images to localize the proteins. (Donor-blue color, Uncorrected FRET-pink color, and

Corrected FRET-red color dots) Adapted from A. Periasamy, Microscopy andMicroanalysis, In Press, 2001

FLIM Theory

Time-resolved fluorescence emission spectroscopy of a photoexcited sample is a

powerful tool for the study of intricate living cells in both space and time of their

internal biochemistry. The experimental challenge of actually visualizing the complex

reaction kinetics is feasible by using the state-of-the-art imaging system and the design

and synthesis of new fluorescent probes. Fluorescence measurements in the time-

domain possess much greater information content about the rates and kinetics of intra-and intermolecular processes than is afforded by wavelength spectroscopy alone.

WHAT IS FLUORESCENCE LIFETIME?

The fluorescence lifetime is defined as the average time that a molecule remains in an

excited state prior to returning to the ground state. For a single exponential decay, the

fluorescence intensity as a function of time after a brief pulse of excitation light is

described as

I (t) = I0 exp (-t/)

where I0 is the initial intensity immediately after the excitation pulse.

In practice, the fluorescence lifetime (tau) is defined as the time in which the

fluorescence intensity decays to 1/e of the intensity immediately following excitation.

Fluorescence decay is often multiexponential, leading to complex decay curves.

Instrumental methods for measuring fluorescence lifetimes are divided into two major

categories, frequency-domain and time-domain. Frequency-domain fluorometers excite

the fluorescence with light, which is sinusoidal and modulated at radio frequencies (for

nanosecond decays), and then measure the phase shift and amplitude attenuation of the

fluorescence emission relative to the phase and amplitude of the exciting light. Thus,

each lifetime value will cause a specific phase shift and attenuation at a given

frequency.

In time-domain methods, pulsed light is used as the excitation source, and fluorescence

lifetimes are measured from the fluorescence signal directly or by photon counting.

WHAT IS FLIM?

Many currently available fluorescence microscopic techniques, such as confocal or

multi-photon excitation, cannot provide detailed information about the organization and


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dynamics of complex cellular structures. In contrast, time-resolved fluorescence

microscopy allows the measurement of dynamic events at very high temporal resolution

and can monitor interactions between cellular components with very high spatial

resolution as well. To date, most measurements of fluorescence lifetimes have been

performed in solution or cell suspensions. Fluorescence lifetime imaging was developed

to overcome this drawback and still provide the ability to use the power of fluorescence

lifetime measurements in a single living cell.

FLIM-FRET

The combination of lifetime and FRET (FLIM-FRET) provides high spatial

(nanometer) and temporal (nanoseconds) resolution (Bacskai et al., 2003; Elnagovan et

al., 2002; Krishnan et al., 2003). The presence of acceptor molecules within the local

environment of the donor that permit energy transfer will influence the fluorescence

lifetime of the donor. By measuring the donor lifetime in the presence and the absence

of acceptor one can accurately calculate the distance between the donor- and acceptor-

labeled proteins. While 1p-FRET produces 'apparent' E%, i.e. efficiency calculated on

the basis of all donors (FRET and non-FRET), the double-label lifetime data in 2p-

FLIM-FRET usually exhibits 2 peaks of donor lifetimes (FRET and non-FRET),

allowing a more precise estimate of distance based on FRET donors only. The former

may be sufficiently accurate for many situations; the latter may be vital for establishing

comparative distances of several proteins from a protein of interest.

what is fret

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