what is fret
TRANSCRIPT
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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
specimen using donor
excitation and acceptor
emission filter set.
cAcceptor
OnlyDonor Acceptor
Signal from an acceptor only
specimen using donor
excitation and acceptor
emission filter set.
dAcceptor
OnlyAcceptor Acceptor
Signal from an acceptor only
specimen using acceptor
excitation and acceptor
emission filter set.
eDonorand
Acceptor
Donor Donor
Signal from donor-and-
acceptor specimen using donor
excitation and donor emissionfilter set.
fDonor and
AcceptorDonor Acceptor
Signal from donor-and-
acceptor specimen using donorexcitation and acceptor
emission filter set.
gDonorand
AcceptorAcceptor Acceptor
Signal from donor-and-
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.