Frontiers in fluorescence microscopy
JOSÉ RINO*, JOSÉ BRAGA, RICARDO HENRIQUES and MARIA CARMO-FONSECA
Instituto de Medicina Molecular, Faculdade de Medicina, Universidade de Lisboa, Lisboa, Portugal
ABSTRACT  How we see organisms and cells depends on the tools at our disposal. For over 150
years, biologists were forced to rely on fixed, dehydrated and stained specimens in order to guess
how the living cells could function. It all changed abruptly during the last two decades when the
rapid development of novel imaging techniques revolutionized the way scientists look at the
structures of life alive.
KEY WORDS: fluorescence, photobleaching, photoactivation, FRET, FLIM
A brief history of light microscopy
More than 300 years ago, Antony van Leeuwenhoek observed
for the first time bacteria and other unicellular organisms using
simple microscopes that he himself designed and constructed.
Leeuwenhoek’s skills enabled him to build single-lens micro-
scopes that magnified over 200 times, with clearer and brighter
images than any of the existing compound microscopes could
achieve. Although combining multiple lenses could in theory yield
higher magnification, the early compound microscopes invented
near the end of the 16th century were limited by defects in the
manufacturing of lenses, which greatly distorted the images.
During the 19th century the compound microscope finally evolved
to become a reliable instrument allowing biologists to make the
seminal discoveries that led to the formulation of the cell theory by
Schleiden and Schwann in 1838.
The ultimate limit of all microscopes is resolution, or the
capacity to clearly distinguish two separate point objects. Be-
cause of the wave nature of light, a single point object seen
through a microscope appears as a blurred disc (the so-called
Airy pattern, Fig. 1), and the discs produced by two point objects
close together may overlap into a single merged image (Fig. 1C).
Ernst Abbe was the first to demonstrate, in 1873, how the
diffraction of light by both the specimen and the lenses affect
image resolution. Theoretically, said Abbe, objects less than 200
nm apart cannot be resolved using visible light. This limit of
resolution was nearly achieved by microscope makers at the end
of the 19th century.
The concept of resolution relates to the level of structural detail
that can be seen when a sample is imaged. In a wide-field
microscope, resolution is affected by light scattering: because the
whole sample is illuminated, more than 90% of the light collected
is out-of-focus. This greatly reduces image detail. “An ideal
Int. J. Dev. Biol. 53: 1569-1579 (2009)
doi: 10.1387/ijdb.072351jr
THE INTERNATIONAL JOURNAL OF
DEVELOPMENTAL
BIOLOGY
www.intjdevbiol.com
*Address correspondence to:  José Rino. Instituto de Medicina Molecular, Faculdade de Medicina, Av. Prof. Egas Moniz, 1649-028 Lisboa, Portugal.
Fax: +351-21-799-9412. e-mail: joserino@fm.ul.pt
Final author-corrected PDF published online:  28 November 2008.
ISSN: Online 1696-3547, Print 0214-6282
© 2009 UBC Press
Printed in Spain
microscope would examine each point in the specimen and
measure the amount of light scattered or absorbed by that point”
noted Marvin Minsky, the inventor of the confocal microscope
(Minsky, 1988). Minsky’s invention in 1953 was far sighted and
remained largely unnoticed until the development of lasers and
the widespread use of fluorescent biological markers turned the
confocal microscope into one of the most exciting instruments
available to biologists toward the end of the 1980s.
The key feature of confocal microscopy is its ability to produce
images of specimens at different depths with excellent contrast.
Confocal laser scanning microscopes (CLSMs) are capable of
slicing optical sections from thin biological samples, allowing
them to be rendered in 3D reconstruction images. Moreover,
because most light coming from regions above or bellow the focal
plane is blocked, out-of-focus light is not added to the image. This
greatly improves depth discrimination and contrast (Stelzer, 1995).
Additionally, the design of CLSMs provides a theoretical improve-
ment of approximately 30% in resolution. However, this does not
hold in practice for the observation of biological samples due to
noise, pixelation and image aberrations of the system (Hell et al.,
1993, Hell and Stelzer, 1995). Indeed, for the current generation
of confocal microscopes the limit of resolution has been reached
at around 200 nm in the lateral (x,y) direction and 600 nm in the
axial (z) direction. Axial resolution can be increased with 4Pi-
confocal microscopes, which use two lenses instead of one,
placed on both sides of the sample. This two-lens system was
called 4Pi in reference to 4πr2 (the surface of a sphere) and the
attempt to visualize a sample from all angles (Hell and Stelzer,
Abbreviations used in this paper: CLSM, confocal laser scanning microscope;
FRAP, fluorescence recovery after photobleaching; FRET, Förster resonance
energy transfer; GFP, green fluorescent protein.

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1570    J. Rino et al.
1992b). Much like astronomers, microscopists then used the
interference between two wave fronts originating from both objec-
tives to yield more image information (Gustafsson et al., 1999,
Hell and Stelzer, 1992a).
A drawback to the traditional CLSM is speed of image acqui-
sition. Because the excitation laser source is scanned across the
specimen in a point-by-point raster pattern, just one to three high
resolution images can be captured per second. This will not be
enough to image processes that occur in fractions of a second.
This disadvantage is overcome with the spinning-disk (also called
Nipkow-disk) confocal microscope that uses disks containing
multiple pinholes to simultaneously illuminate and collect light
from several points of the specimen. Imaging rates of up to 30
frames per second are obtained by rotating the disks at high
speed. The main caveat of this approach however, is the highly
inefficient excitation of the sample, as the vast majority of excita-
tion light available is rejected by the pinholes (Nakano, 2002).
Additional disadvantages include worse axial resolution, lesser
homogeneity of field of illumination and less flexibility to changes
in objectives and experimental settings (Wang et al., 2005).
Another drawback of CLSMs is their inability to image large,
thick living specimens. This is because large specimens absorb
and scatter a considerable amount of light. The advent of Multi-
Photon Microscopy (MPM), introduced in 1990 by the work of W.
Denk, J. Strickler and W. Webb (Denk et al., 1990) was the
breakthrough in optical microscopy that allowed to fulfill the
challenging task of imaging living organs such as brain and liver.
MPM relies on the quasi-simultaneous absorption of two or more
photons by a molecule (i.e., the fluorescent marker). The multi-
photon absorption process, first predicted in 1930 by Maria
Göppert-Mayer, needs a very high density of photons from a
pulsed light source. Two-photon fluorescence microscopy (2PFM)
is currently the most common multi-photon fluorescence applica-
tion in biology. Commercially available instruments use a laser
operating in the range of 720-920 nm. While photodamage is
lower and penetration depth is higher, the resolution obtained with
these long illumination wavelengths is necessarily less good than
in the traditional confocal microscope.
An alternative to visualize large living specimens with higher
resolution was more recently developed. The so-called selective
plane illumination microscopy (SPIM) is a system that combines
two-dimensional illumination with orthogonal camera-based de-
tection. SPIM generates multidimensional images of samples up
to a few millimeters in size with high-resolution and minimal
photodamage (Huisken et al., 2004, Verveer et al., 2007).
Of all the numerous developments introduced in light micros-
copy over the past century, the most spectacular breakthrough
was the demonstration that the resolution of images obtained with
a fluorescence microscope is no longer limited by the wavelength
of the light, as postulated by Abbe. This ingenious trick makes use
of the nonlinear excitation properties of fluorophores. The prin-
ciple of stimulated emission depletion (STED) microscopy is to
make the fluorescent spot smaller by quenching the excited
molecules at the rim. Two synchronized trains of laser pulses are
employed for this purpose. A visible pulse excites the fluorophores
in the focus and a subsequent red-shifted pulse performs the
quenching. By using enough photons, the quenching process is
saturated and this “helps to squeeze the spot down to a very small
scale, in principle infinitely” says Stephen Hell, the inventor of
STED (Dyba and Hell, 2002, Hell and Wichman, 1994, Klar et al.,
2000). This novel microscopy, which has recently become com-
mercially available, has already proved to be applicable to live
specimens, resolving the fate of individual synaptic vesicles
(approximately 40 nm in diameter) after neurotransmitter release
(Willig et al., 2006).
Fluorescence and fluorophores
Fluorescence is the process by which molecules become
excited and emit light with defined wavelengths as a consequence
of having absorbed photons with shorter wavelengths (Stokes,
1852). The difference between the exciting and emitted wave-
lengths, known as the Stokes shift, depends on the outmost
electron orbitals of the fluorescent molecule, or fluorophore.
Fluorescent compounds at room temperature are normally in the
lowest vibrational levels of the ground state (S0 where S stands for
singlet state). When a photon is absorbed, all its energy is
transferred to the fluorescent molecule. However, for fluores-
cence to occur the minimum energy of an incoming photon must
correspond to the energy difference between the lowest energy
level of the first excited state (S1) and S0  (Fig. 2A). Photons with
higher energies induce transitions into higher vibrational and
rotational levels or to higher energy states (S2, S3,…). The energy
(E) of a photon is E = h x c / λ (where h is Plank’s constant, c is the
speed of light and λ is the photon’s wavelength in vacuum), and
each fluorophore can be excited by a range of wavelengths that
define its absorption spectrum.
Excited states are usually very unstable and the molecule
θ 
spherical 
wavefronts 
Objective 
Point  
Source 
Image 
Plane 
B
C
A
Fig. 1. The Airy pattern.
(A) A point source emits
light waves in all direc-
tions. A fraction of these
is captured by the objec-
tive lens, at a cone with a
half-angle θ. The light
waves are then diffracted
by the lens but instead of
being focused into an infi-
nitely small point, they
converge and interfere
with each other at the
image plane, producing a diffraction Airy pattern. (B) Airy patterns for two point objects which are sufficiently separated in order to be clearly resolved.
(C) Two point objects are not resolved if their separation is not sufficient to produce a contrast between them.

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Frontiers in fluorescence microscopy   1571
will tend to return to its low-energy ground state. Rotation,
vibrational relaxation and “internal conversion” will bring the
excited molecule to the lowest energy level of S1 in picosec-
onds, without emission of radiation. Only when the molecule
returns to any of the vibrational levels of S0 does fluorescence
occur, with emission of a photon with energy equivalent to this
transition. The wavelength range of an emission spectrum thus
reflects the different transitions from the lowest level of S1 to the
different rotation and vibrational levels of the ground state.
The process just described involves the absorption of one
single photon at a time. However, with very high photon densi-
ties it is possible that a molecule absorbs two (or more) photons
almost instantaneously. In this case, a transition to an excited
state occurs if the sum of energies of the photons is at least
equal to the energy gap between S1 and S0. This implies that
multi-photon fluorescence can be obtained using less ener-
getic, longer wavelengths for excitation.
The time it takes before an excited fluorophore undergoes
the transition from the lowest level of S1 to the ground state is
called the excited state lifetime. The lifetime of a fluorophore,
which is usually in the nanosecond range, can be reduced if
another nearby molecule absorbs the energy in a non-radiative
manner, preventing the fluorophore from emitting a photon
(quenching). These lifetime reductions due to intermolecular
interactions can be measured by a technique called Fluores-
cence Lifetime Imaging Microscopy (FLIM). If the nearby inter-
acting molecule is itself a fluorophore, the absorbed energy can
in turn lead to its excitation and subsequent fluorescence. This
phenomenon, known as Fluorescence Resonance Energy
Transfer (FRET) can be probed by a variety of techniques,
providing information on intramolecular distances far below the
resolution of an optical microscope.
During the process of relaxation, the molecule may make a
transition to unlikely states called triplet states. In these states,
the excited electron inverts its spin state (Fig. 2B) and thus the
overall magnetic moment of a molecule will change. These
excited states usually last for a long time because further
relaxation can only occur if the molecule undergoes another
unlikely spin-inverting transition. Triplet states can sometimes
reach the ground state by emitting a photon, in a process called
phosphorescence. Alternatively, they may undergo triplet-trip-
let transitions into higher triplet states if another photon is
absorbed, further delaying any light emission by the molecule.
The intrinsic brightness of a fluorophore is determined by its
photostability, quantum yield and molar extinction coefficient
(Shaner et al., 2005). The quantum yield (Φ) is a measure of the
fluorophore’s total light emission over the entire spectral range.
It is defined as the ratio between the numbers of emitted and
total absorbed photons and has a value between 0 and 1. The
molar extinction coefficient (μ), or molar absorption coefficient,
corresponds to the absorbance of light per unit path length and
per unit of concentration of a given fluorophore. Its value,
calculated as μ = A / C (where A is the absorbance and C the
fluorophore’s concentration) reflects the probability of a photon
to be absorbed by the fluorophore. The higher the value of
quantum yield and molar extinction coefficient, the brighter is
the fluorophore. Stability relates to the fact that fluorophores
can only undergo a limited number of cycles of absorption and
emission. All fluorophores will eventually photobleach by mecha-
nisms that are not completely elucidated but most probably
involve triplet states. As triplet states are long-lived, the excited
molecules have more time to interact with other molecules such
as oxygen. Molecular oxygen is a ground state triplet which, by
interaction with a molecule in a triplet state, may make a
transition to a highly reactive excited singlet state. This excited
molecule in turn can react with either another fluorophore (chang-
Fig. 2. Excitation and emission of light by a fluorophore. (A) The energy states of a fluorophore molecule are represented in this Jablonski diagram
together with the times that the various steps in fluorescence excitation, emission and phosphorescence take. The electronic levels are indicated by
thick black horizontal lines, whereas the vibration and rotational levels are represented by thinner lines. When a photon with the appropriate wavelength
is absorbed by the molecule, it causes a transition to a high vibrational level of the excited electronic state S2 (dark blue arrows). The molecule then
undergoes rotational and vibrational relaxations (dotted dark red arrows) and internal conversion (red arrow) before it reaches the lowest energy level
of S1. Transition to the ground state usually occurs via emission of a photon, by fluorescence (green arrows). Alternatively the molecule might undergo
intersystem crossing (brown arrow) and arrive at a triplet state, where it may cause phosphorescence emission (orange arrows) or triplet-triplet
transitions (light blue arrows) that further delay the emission of light or prevent it altogether. (B) Electrons are usually paired with anti-parallel spins
in a singlet state. If an electron is excited, the system maintains their anti-parallel configuration; however, there is a low probability that an excited
electron reverses its spin. In this situation the system is in a triplet state T1.
S0 
S1 
S2 
T1 
ground state 
Singlet 
T2 
Triplet 
vibrational levels 
rotational levels 
Excitation 
(fs) 
Fluorescence (ns) 
Internal conversion 
Relaxation 
Intersystem 
crossing 
Phosphorescence (>μs) 
triplet-triplet 
transition 
S 
S 1 
0 
ground  
state 
excited 
state 
T 1 
excited  
state 
S 0 
S 
S 1 
0 
(ps)
B
A

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1572    J. Rino et al.
ing irreversibly its structure) or other organic molecules causing
phototoxicity (Lichtman and Conchello, 2005).
The GFP revolution
The discovery and development of the Green Fluorescent
Protein (GFP) and related fluorescent proteins (FPs) originated
an extraordinary revolution in biological research, which was
recently acknowledged with the attribution of the Nobel Prize in
Chemistry 2008 to Osamu Shimomura, Martin Chalfie and Roger
Y. Tsien. With these genetically encoded markers it became
possible to fluorescently tag almost any protein with minimal
invasiveness (Heim et al., 1995). However, not all fluorescent
chimeras are successful. FP tagging can sometimes alter the
endogenous protein folding, rendering it non-functional. There-
fore, it is crucial to perform careful and detailed biological controls
to ensure the functionality of the fusion protein.
The GFP from the jellyfish Aequorea victoria (avGFP) was the
first fluorescent protein to be cloned (Tsien, 1998). The protein is
a 27 kDa monomer with a cylindrical shape composed of a 11-
stranded β-can, 42 Å long and 24 Å in diameter, with both ends
closed by short helical segments (Fig. 3A). The chromophore (p-
hydroxybenzylideneimidazolidinone) is almost perfectly buried in
a α-helix at the center of the cylinder. This configuration is thought
to be responsible for the high stability of GFP. Indeed, the protein
is highly resistant to denaturation, proteolysis and quenching by
molecular oxygen or pH changes (Ormo et al., 1996). The mature
chromophore is formed autocatalytically by a series of intramo-
lecular reactions involving amino acids 65-67 (Ser-Tyr-Gly) and
the only exogenous reagent needed is oxygen (Prendergast,
1999, Reid and Flynn, 1997).
Despite its great potential, the wild type avGFP was not an
ideal fluorophore. The excitation spectrum had two peaks (at 396
and 475 nm) and the molar extinction coefficient was low (22,000
M-1cm-1, compared to fluorescein’s 80,000 M-1cm-1). Moreover,
the protein took a relatively long time to mature in cells and had
a tendency to dimerize at high concentrations (Cubitt et al., 1995).
Serious mutagenesis efforts followed by rational fluorescent
protein design led to the progressive development of improved
versions of the protein, such as the enhanced GFP (EGFP),
enhanced blue fluorescent protein (EBFP), enhanced cyan fluo-
rescent protein (ECFP) and enhanced yellow fluorescent protein
(EYFP) (Lippincott-Schwartz and Patterson, 2003, Shaner et al.,
2005, Tsien, 1998).
A variety of additional mutations aiming to reduce dimerization
and improving chromophore maturation have culminated in the
development of vastly improved fluorophores. These include
monomeric versions of the blue, cyan, green and yellow versions
of GFP (mBFP, mCFP, mGFP, mCitrine and mVenus (Zhang et
al., 2002)) and also mCerulean (excitation: 433 nm; emission: 475
nm) (Rizzo et al., 2004) and T-Sapphire (excitation: 399 nm;
emission: 511 nm) (Zapata-Hommer and Griesbeck, 2003), which
are specially suited for FRET applications.
On the other side of the spectrum, a red fluorescent protein
from the coral Discosoma, called DsRed (Matz et al., 1999), was
also mutated to generate a monomeric red fluorescent protein
(mRFP1) (Campbell et al., 2002). More recent mRFP1 mutagen-
esis improved brightness and photostability leading to a plethora
of new monomeric spectral mutants (Shaner et al., 2004), which
range from the greenish mHoneydew to the far red mPlum and
include the highly photostable and fast maturing mCherry (Fig.
3C).
Further innovation was introduced by the development of
photoactivatable and photoswitchable fluorescent proteins
(Lukyanov et al., 2005). These fluorophores are capable of
pronounced changes in their spectral properties once irradiated
with light of a specific wavelength and intensity. Photoactivatable
GFP (PA-GFP) and photoswitchable CFP (PS-CFP) were both
developed from avGFP, taking advantage of the wild type protein
double excitation peaks. PA-GFP is the result of a single point
mutation (Thr203_His) in avGFP (Patterson and Lippincott-
Schwartz, 2002). The non-photoactivated form of the protein is
characterized by 400 nm excitation and 515 nm emission peaks,
with almost no fluorescence emission when excited at 480 – 510
nm. Irreversible photoactivation by intense UV light (~ 400 nm)
results in a 100-fold increase of green fluorescence (excitation:
504 nm; emission: 517 nm). Non-photoactivated PS-CFP shows
cyan fluorescence instead, with an emission peak at 468 nm
(Chudakov et al., 2004). Irreversible photoconversion with in-
tense UV light then provokes a 300-fold increase in green fluores-
cence (excitation peak: 490nm; emission peak: 511 nm) accom-
panied with a 5-fold reduction in cyan fluorescence.
Photoactivatable red fluorescent proteins have also been devel-
Fig. 3. The Green Fluorescent Protein and variants. (A) GFP folds into
a β-can structure with the chromophore buried inside the cylinder-shaped
molecule, which has a diameter of about 25 Å and a length of 40 Å. (B)
Enhanced GFP (EGFP) expressed in a transfected human cell, visualized
by confocal microscopy. Bar: 5 μm. (C) Engineered fluorescent proteins,
derived from GFP and the monomeric Red Fluorescent Protein (mRFP1),
covering the full visible emission spectrum. Protein samples are shown
in purified forms inside Eppendorf tubes. Adapted from (Tsien, 2005).
mHoneydew 
EBFP 
ECFP 
EGFP 
YFP 
mBanana 
mOrange 
tdTomato 
mStrawberry 
mCherry 
mGrape1 
mRaspberry 
mGrape2 
mPlum 
mTangerine 
GFP-derived 
mRFP1-derived 
B
C
A

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Frontiers in fluorescence microscopy   1573
oped, such as the monomeric PA-mRFP (Verkhusha and Sorkin,
2005), which requires intense 380 nm activation to exhibit a 70-
fold increase in red fluorescence (excitation: 578 nm; emission:
605 nm). Other photoconvertible proteins able to convert irrevers-
ibly from green to red upon UV illumination include Kaede (Ando
et al., 2002), mEosFP and KikGR (Lukyanov et al., 2005). Revers-
ible conversion, on the other hand, can be achieved with the
tetrameric kindling fluorescent protein (KFP1), converted by 488
nm illumination (Chudakov et al., 2003) or Dronpa, (Ando et al,
2004) and its fast-switchable variant rsFastLime (Stiel, Trowitzsch
et al., 2007) which can both be activated by 405 nm UV light and
deactivated with 488 nm illumination. The recent development of
Dendra (Gurskaya et al., 2006) resulted in the first monomeric
protein capable of green (488 nm) photoactivation. Both Dendra
and PS-CFP have recently been optimized to generate Dendra2
and PS-CFP2 respectively, both of them monomeric derivatives
which display enhanced brightness and accelerated maturation
rates at 37ºC (Chudakov et al., 2007).
Quantum dots
To overcome the limitations imposed by the photostability of
standard dyes, alternatives have been sought. Quantum dots
(QDs) are small nanocrystals, 2-8 nm in diameter, which have
very interesting optical properties (Gao et al., 2005). Excitation
and absorption spectra are dependent on the size and composi-
tion of QDs. Usually, excitation spectrum is broad and the emis-
sion spectrum narrow, thus QDs with different emission spectra
can be excited with only one wavelength, which makes them
specially suited for multiple color applications. Due to a high
extinction coefficient QDs are much brighter than most organic
fluorophores, though their main advantage is that they are much
more resistant to bleaching. Another advantage is that they are
non-toxic to cells. To date, QDs in cell biology are most often used
as labels for immunofluorescence (Michalet et al., 2005). How-
ever, QDs have already been applied in vivo to track single-
molecule movement inside cells (Dahan et al., 2003) and to image
lymph nodes and tumours at the whole-organism level (Gao et al.,
2005). Blinking of QDs, which can occur in a wide range of times
(~100 μs -100 s), is one the major disadvantages of these probes
because it restricts quantitative analysis (Arya et al., 2005).
Photobleaching methods
The advent of live cell imaging by time-lapse microscopy using
fluorescent proteins as molecular tags triggered a large number
of studies on the localization and dynamics of cells and sub-
cellular structures. However, in order to obtain information on the
dynamics of the components of sub-cellular structures at the
molecular level, new methods had to be developed.
Although photobleaching of fluorophores is usually undesir-
able in most fluorescence microscopy applications, more than
thirty years ago it was devised purposefully as a tool to study
molecular dynamics (Cone, 1972, Edidin et al., 1976, Peters et al.,
1974, Poo and Cone, 1973). This technique was to become
known as fluorescence recovery after photobleaching (FRAP).
FRAP is based on the local perturbation of the fluorescence
steady-state by inducing irreversible photobleaching with an
intense light source at a selected region of interest (Fig. 4). Then,
due to the motion of unbleached fluorophores from regions not
affected by bleaching, fluorescence relaxes to a new steady-
state. The rate by which this relaxation occurs is related with the
overall mobility of the fluorescently-tagged molecule: a higher
mobility implies a faster recovery of fluorescence inside the
bleached region (Fig. 4D, E). Monitoring of recovery should be
performed with minimal light intensity to avoid significant and
global unspecific bleaching of the sample. In the case that all
fluorescent molecules are mobile (Fig. 4A, D), the fluorescence
intensity will reach a plateau that will correspond to the initial pre-
bleach fluorescence value, provided that the bleach region is
small (typically with a diameter of ~1 μm; Braga et al., 2004). If
fluorescence recovery does not return to the initial pre-bleach
value (Fig. 4E) then either there is an immobilized sub-population
of fluorescent molecules in the bleach region (Fig. 4B) or a
significant amount of the total number of fluorescent molecules
was lost during bleaching (Fig. 4C).
FRAP is easily implemented in most standard CLSMs, which
largely contributed to the increased availability and popularity of
Fig. 4. Fluorescence Recovery After Photobleaching (FRAP). (A) In
FRAP, a cell is imaged before and after bleaching a small spot. As all the
molecules are mobile, fluorescence in the bleach region eventually
recovers to its initial state. (B) If some molecules are stalled in the bleach
region, final fluorescence will be lower than initial fluorescence. (C) Due
to the finite size of the cell (or the nucleus, in the case depicted) bleaching
a large region will deplete a significant proportion of the fluorescent pool,
thus leading to incomplete recovery. (D) Simulated FRAP recovery
curves show that species with different rates of movement will yield
different recovery curves. Fluorescence values are normalized for the
initial intensity. (E) Same as in (D), but with 25% of the molecules
immobilized in the bleached region.
with immobile fraction 
with immobile fraction 
no immobile fraction 
Normalized Fluorescence 
no immobile fraction 
large bleach region 
Bleaching 
Bleaching 
Bleaching 
Time 
2 
4 
6 
8 
0.2 
0.4 
0.6 
0.8 
1 
2 
4 
6 
8 
0.2 
0.4 
0.6 
0.8 
1 
Time (s) 
Time (s) 
D = 5 μm  s 
2 -1 
D = 0.5 μm  s 
2 -1 
D = 5 μm  s 
2 -1 
D = 0.5 μm  s 
2 -1 
B
C
D
E
A

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1574    J. Rino et al.
the technique (Fig. 5). FRAP has been extensively used in a wide
variety of biological applications and the information extracted
from these experiments range from purely qualitative to sophisti-
cated quantification of diffusion and binding parameters. The
technique can be used to evaluate if a protein is a stable compo-
nent of a cellular structure or otherwise is in constant exchange
between different pools (Boisvert et al., 2001, Desterro et al.,
2003, McNally et al., 2000, Tavanez et al., 2005, Trinkle-Mulcahy
et al., 2001). A direct comparison of recovery curves from the
same molecule under different experimental conditions may
already show biologically relevant differences in mobility.
Extracting quantitative parameters from FRAP recovery curves
is not a trivial task. Simple approaches consist in calculating the
time it takes to reach a defined percentage of recovery (Cheutin
et al., 2003, Harrer et al., 2004, Sunn et al., 2005), measure the
percentage of recovery observed after a fixed time period
(McDonald et al., 2006), and fit recoveries to either one exponen-
tial (Belgareh et al., 2001, Lam et al., 2002, Lang et al., 1986) or
a sum of exponentials (Handwerger et al., 2003, Kimura and
Cook, 2001). However, care must be taken as diffusion-like
recoveries are apparently properly fitted with 2 or more
exponentials, but this type of fitting gives incorrect information
about the underlying process (Sprague et al., 2006, Sprague et
al., 2004).
More accurate, yet much more laborious methods involve
solving the diffusion equation. After the pioneering work devel-
oped thirty years ago by Axelrod and co-workers (Axelrod et al.,
1976), several mathematical approaches have been devised to
extract mobility parameters (Blonk et al., 1993, Braeckmans et al.,
2003, Braeckmans et al., 2006, Braga et al., 2004, Kubitscheck et
al., 1998) and binding parameters from biological FRAP data
(Beaudouin et al., 2006, Braga et al., 2007, Carrero et al., 2003,
Sprague et al., 2006, Sprague et al., 2004). The choice of a
method for analysis of FRAP data should be guided by the
level of precision desired and the practicality of its imple-
mentation, However, and most importantly, the underlying
assumptions of the models (such as negligible beach time,
two-dimensional diffusion, very fast diffusion compared to
binding) should be analyzed carefully in order to check the
applicability of the method to the situation being studied.
Fluorescence loss in photobleaching (FLIP) consists on
the repetitive photobleaching of a defined cellular region
(Fig. 6). Fluorescence intensity is monitored over regions of
interest distant from the bleached region (Cole et al., 1996).
FLIP is mainly used to assess whether the tagged molecule
shuttles between the bleached and imaged regions (Goodwin
and Kenworthy, 2005, Lippincott-Schwartz et al., 2003,
Lippincott-Schwartz et al., 2001).
Inverse FRAP (iFRAP, Fig. 6) consists in photobleaching
an entire cell or sub-cellular structure with the exception of
a region of interest. Fluorescence loss in the region of
interest is subsequently monitored over time. This method
yields information on the mobility rate and residence time of
the tagged molecule in the region of interest (Dundr et al.,
2002, Lippincott-Schwartz et al., 2003). However, because
a considerable amount of time is required to bleach the
whole cell, iFRAP can only be applicable to molecules with
slow kinetics, such as nucleoporins at the nuclear pore
complex (Rabut et al., 2004). Photoactivation (PA) or
LASER
beam
Bleaching
No Bleaching
Line start
Line end
Bleaching Phase
Imaging Phase
1 st
2 nd
i th
n th
1 st
2 nd
N th
Fig. 5. Schematics of bleaching with a Confocal Laser Scanning Micro-
scope (CLSM). The software of a CLSM allows for arbitrary 2D shapes to be
drawn in the sample. A laser beam scans the region (comprising a total of n lines,
numbered in italic), for a certain number of iterations. The laser beam has a
certain width (depicted as the blue transparent circle). Laser intensity is high
inside the user defined region (thick lines) and nearly zero outside (dashed blue
lines). After bleaching, the entire region is imaged. For imaging, the monitoring
beam scans the whole image with constant illumination intensity. The resulting
image contains the bleached region of interest (ROI) and consists of N lines
(numbered in normal lettering), with N>n. (adapted from Braga et al., 2004)
photoconversion methods circumvent this limitation (see bellow).
FRAP for immobilization measurement (FRAP-FIM, Fig. 6)
specifically aims at quantifying the percentage of immobile mol-
ecules in a given region of the cell (Houtsmuller et al., 1999,
Rademakers et al., 2003). In this method, spot bleaching is
performed by a relatively long bleach pulse (~ 5s) at low laser
intensity, allowing for diffusion of molecules to occur during the
bleach phase. Confocal images acquired before and after bleach-
ing are used to calculate the fluorescence ratio before and after
bleaching as a function of the distance to the bleached spot.
Transiently immobilized molecules at the bleached region can be
followed by recording post-bleach images at increasing time
intervals. Plotting the average immobile fraction as a function of
time allows to estimate the average binding time of the molecules
at the region of interest (Houtsmuller and Vermeulen, 2001).
Photoactivation and photoconversion
Photoactivation or photoconversion of fluorophores at a spe-
cific region of interest allows for a selected population of tagged
molecules to be rendered visible (Fig. 6). The dynamics of the
selected pool of molecules can then be directly monitored over
time (Chapman et al., 2005, Deryusheva and Gall, 2004). Thus,
compared to photobleaching, the main advantage of these meth-
ods is that they allow the identification and tracking of a specific
population of molecules. By photoactivation, an initially faint
fluorophore is illuminated with specific wavelengths that cause an
increase in its fluorescence. For example, after UV irradiation, the
fluorescence of photoactivatable GFP (PA-GFP) increases by a
factor of 100 (Patterson and Lippincott-Schwartz, 2002). By
photoconversion, the absorption and emission spectra of a
fluorophore change upon illumination with specific wavelengths.

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Frontiers in fluorescence microscopy   1575
Photoswitchable fluorophores that convert from green to red
under UV illumination have already been used to study the
dynamics of proteins inside cells (Ando et al., 2002, Deryusheva
and Gall, 2004). However, care must be taken with choosing a
photoactivatable/photoswitchable fluorescent protein, as only the
most recently developed monomeric ones can be used for indi-
vidual protein tagging. Non-monomeric proteins, such as tet-
rameric KFP1 should only be used to label whole cells or or-
ganelles (Lukyanov et al., 2005).
Fluorescence correlation spectroscopy
Contrasting with the previously described techniques that
involve photobleaching or photoactivation of tagged molecules, in
fluorescence correlation spec-
troscopy (FCS) the primary pa-
rameter of interest is not the in-
tensity of fluorescence emission
(Bacia and Schwille, 2003, Elson
and Magde, 1974, Elson, 2004,
Gosch and Rigler, 2005, Kim et
al., 2005, Schwille, 2001). FCS
consists in measuring, with high
temporal resolution, fluorescence
fluctuations in the diffraction-lim-
ited volume defined by a focused
laser beam (Fig. 6). Unlike
photobleaching or photoactivation
microscopy, which uses micro-
molar to milimolar concentrations
of fluorescent molecules, FCS
typically uses femtoliter detection
volumes and nanomolar concen-
trations of tagged molecules
(Schwille et al., 1999). Within the
observation volume only a few
molecules are present. Because
the molecules are moving in and
out of that volume, the number of
molecules detected is stochasti-
cally fluctuating around a mean
value. Presumably the fluores-
cence measured at one particu-
lar instant is strongly correlated
with the fluorescence measured
shortly after but will possibly be
weakly correlated with fluores-
cence at distant times. These
correlations will vanish more rap-
idly in the case of a fast moving
species. By computing the
autocorrelations of the fluctua-
tions it is possible to deduce the
diffusion coefficient of the mol-
ecules (Bacia and Schwille,
2003). FCS further allows bind-
ing studies to be performed, by
tagging two interacting molecules
Fig. 6. FRAP variants and other methods for measuring molecular mobility. (A) Schematics of the
Fluorescence Loss In Photobleaching (FLIP), inverse FRAP (iFRAP), photoactivation (PA), FRAP for
immobilization measurement (FRAP-FIM) and fluorescence correlation spectroscopy (FCS) techniques.
The regions delimited by the red line indicate bleaching regions, yellow circles photoactivation regions.
Monitored regions are outlined by blue circles (FLIP, iFRAP and PA) or by a blue line (FRAP-FIM). In FCS,
the monitored region is limited to a focal volume of less than one femtoliter (hourglass-shaped volumes).
(B) Normalized fluorescence curves obtained from the different techniques are plotted over time (iFRAP and
PA) or over the bleach radius (FRAP-FIM). In FCS, the fluorescence fluctuation curves (red graph) are used
to calculate the autocorrelation function (blue graph). See text for details.
iFRAP 
Time 
Bleaching
photoactivation 
Bleaching
0.2 
0.4 
0.6 
0.8 
1 
0.2 
0.4 
0.6 
0.8 
1 
PA 
Time 
0.2 
0.4 
0.6 
0.8 
1 
Radius 
FRAP-FIM 
Time 
FCS 
iFRAP 
PA 
FRAP-FIM 
FCS 
Bleaching 
Bleaching 
Time 
FLIP 
FLIP 
Time 
0.2 
0.4 
0.6 
0.8 
1 
B
A
with different fluorophores (Bacia and Schwille, 2003).
Förster resonance energy transfer (FRET)
Protein-protein interactions are not directly discernible by light
microscopy. The relative proximity of two proteins tagged with
different fluorophores can only be determined by conventional
fluorescence microscopy to the scale of roughly ~ 200 nm, the
limit of optical resolution imposed by the diffraction barrier.
However, protein-protein interactions require proximity distances
within the range of 1 – 10 nm (Heim et al., 1995). This degree of
resolution can only be achieved in light microscopy through the
use of Förster resonance energy transfer (FRET) methods
(Wallrabe and Periasamy, 2005, Wouters et al., 2001).
Förster resonance energy transfer, also called fluorescence

----!@#$NewPage!@#$----
1576    J. Rino et al.
resonance energy transfer, is a phenomenon that occurs when
two different fluorophores (called donor and acceptor) with over-
lapping emission/absorption spectra are in close proximity to
each other and in a suitable orientation (Selvin, 2000, Truong and
Ikura, 2001, Voss et al., 2005). FRET involves the non-radiative
transfer of energy (no photons are emitted) from an excited state
in the donor fluorophore to the nearby acceptor.
The energy transfer efficiency E is related to the distance r
between the donor and acceptor fluorophores by
0)6
/
(
1 / 1
R
r
E =
+
, where R0 is the Förster radius, the dis-
tance at which the efficiency of energy transfer is 50% of maxi-
mum. The Förster radius depends on the extent of overlap
between the donor emission and the acceptor excitation spectra,
the absorption coefficient of the acceptor, the quantum yield of the
donor and the relative orientation of the donor and acceptor (the
donor and acceptor transition dipoles must be aligned relative to
each other).
The value of R0 effectively defines the resolution of FRET,
which is typically < 10 – 100 Å. Because FRET falls off as the sixth
power of the distance between the donor and acceptor, when
these molecules are separated by distances greater than 2R0 no
FRET occurs. The phenomenon of FRET can thus be applied in
fluorescence microscopy to distinguish proteins that are merely
co-localized in the same compartment from those that are under-
going protein-protein interactions (Kenworthy, 2001).
The earliest use of GFP in a FRET pair involved a BFP donor
and an EGFP acceptor (Heim and Tsien, 1996), a combination
that was later replaced by the ECFP/YFP pair due to the improved
brightness and photostability of ECFP when compared to BFP
(Pollok and Heim, 1999). ECFP/YFP quickly became the most
commonly used FRET pair, with ECFP being currently replaced
by its improved version Cerulean, which has a higher quantum
yield, extinction coefficient and, most importantly, a single fluo-
rescence lifetime (Rizzo et al., 2004). Other popular FRET pairs
include the recently developed mOrange and T-Sapphire (Shaner
et al., 2004), Cerulean and Dronpa (Lukyanov et al., 2005) and
EYFP and mCherry (Shaner et al., 2004). Of particular interest are
the recently developed monomeric optimized versions of cyan
and yellow super fluorescent proteins, termed SCFP3A and
SYFP2 respectively, which fold faster and more efficiently at 37ºC
and are brighter than their predecessors CFP and YFP (Kremers,
Goedhart et al., 2006). Both of them constitute an obvious FRET
pair, with SCFP3A as a donor and SYFP2 as an acceptor, but
SYFP2 can also act as a donor for mCherry.
FRET microscopy approaches can be divided into intensity
based methods and fluorescence decay kinetics based methods
(Wouters et al., 2001). Intensity based FRET techniques take
advantage of the fact that excitation of a donor fluorophore in
FRET results in quenching of donor emission and in an increased,
sensitized acceptor emission. Detection of FRET through sensi-
tized emission of the acceptor is a complex task that requires
correction of both leak-through of the donor emission and direct
excitation of the acceptor (Day, 1998, Gordon et al., 1998, Nagy
et al., 1998, Xia and Liu, 2001). These corrections often involve
the acquisition of images from samples with the donor alone and
the acceptor alone, a pitfall that can lead to errors in the estimation
of the correction parameters because the quantum yields of the
donor and the acceptor might vary in the different samples
(Wouters et al., 2001).
Another approach consists in measuring the donor fluores-
cence specifically, by using appropriate emission filters that
eliminate the leak-through of acceptor emission (Bastiaens and
Pepperkok, 2000). FRET can then be detected by comparing the
quenched with the unquenched donor emission after specific
photobleaching of the acceptor fluorophore (Fig. 7) (Bastiaens et
al., 1996, Wouters et al., 1998). The principle behind acceptor
photobleaching FRET is that energy transfer is reduced or elimi-
nated when the acceptor is irreversibly bleached, thereby causing
an increase in donor fluorescence.
Kinetic-based approaches for FRET determination measure
the excited state decay kinetics of the donor or the acceptor,
instead of their fluorescence intensity. The most widely used of
these approaches is fluorescence lifetime imaging microscopy
(FLIM) (Gadella and Jovin, 1995, Wallrabe and Periasamy, 2005,
Wouters and Bastiaens, 1999), which detects the decrease in
fluorescence lifetime of the donor due to the depopulation of its
excited state by FRET. FLIM determines a single lifetime value for
each position in an image. This measured lifetime is a nonlinear
function of the true lifetimes and the populations of bound and
Fig. 7. Acceptor Photobleaching FRET. (A) FRET between a donor and
an acceptor molecule only occurs if the distance between them is in the
order of a few nanometers. FRET causes the donor fluorescence to be
quenched while the acceptor undergoes sensitized emission. (B) Accep-
tor photobleaching FRET assesses protein-protein interactions by bleach-
ing the acceptor molecules with a high intensity laser (yellow arrow). (C)
If the donor fluorescence was being quenched by FRET, it will increase
after the acceptor has been bleached.
no FRET 
FRET 
donor 
acceptor 
donor 
acceptor 
acceptor is 
photobleached 
donor fluorescence 
is not altered 
donor fluorescence 
increases 
B
C
A

----!@#$NewPage!@#$----
Frontiers in fluorescence microscopy   1577
unbound donor molecules in that position (Wouters et al., 2001).
Two different FLIM implementations can be distinguished. In
time-domain FLIM, a short laser pulse excites the fluorophore and
the subsequent emission is measured time-resolved, thus origi-
nating a decay curve which can be fitted to a lifetime value. In
frequency-domain FLIM, a modulated excitation light is used to
excite the fluorophore and the lifetime is determined by measuring
either the phase shift or the decrease in modulation depth in the
emission. Time-domain and frequency domain FLIM have been
applied to both widefield (Oida et al., 1993, Squire and Bastiaens,
1999, van Munster and Gadella, 2005) and scanning microscopy
(Carlsson and Liljeborg, 1997, Sanders et al., 1995).
Towards super-microscopy
More than 400 years after the advent of the compound micro-
scope, light microscopy is still a field of intensive research and
further developments will certainly provide biologists with novel
tools to explore the cell. In parallel with the groundbreaking
demonstration that the resolution of images obtained with a
fluorescence microscope is no longer limited by the wavelength of
the light, a revolution in optical components is currently emerging.
Super-lenses with negative refraction indexes, a concept initially
proposed by J. Pendry in 2000 (Pendry, 2000) have proven to
image bellow the diffraction limit (Fang et al., 2005). Conventional
lenses made of positive refraction index media limit resolution
essentially because evanescent waves containing high spatial
frequency information are lost. Meta-materials (reviewed in Smith
et al., 2004) are artificially designed materials with unusual
electromagnetic properties in which a negative refraction index is
attainable. In those materials, the evanescent wave decay is
compensated, thus allowing for sub-wavelength image recon-
struction. A super-lens made from a thin silver slab, working at
optical frequencies, was able to produce a non-magnified image
with 60 nm resolution (Fang et al., 2005). More recently, super-
lenses were shown to produce magnified images with 70 nm-130
nm resolution (Liu et al., 2007, Smolyaninov et al., 2007). Super-
microcopy is visibly on the horizon.
Acknowledgments
We thank Dorus Gadella for stimulating discussions and critical
comments. We are also grateful to Fundação para a Ciência e Tecnologia,
Portugal (POCI/SAU-MMO/57700/2004), and the European Commission
(LSHG-CT-2003-503259).
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