Shining
Fluorescence Details
34 ˇ Basics of light Microscopy & iMaging shining fluorescence Details
A first introduction
Most of the newly developed microscopic
techniques make use of fluorescence.
Fluorescence microscopy is more than
'just making colourful images in one,
two, three or even more colours`, it is an
enormously powerful tool for investigations
in the biological field. Fluorescence
techniques place numerous benefits in
the hands of researchers wishing to exploit
the upper limits of sensitivity and
resolution in microscopy. Beyond the scientific
benefits, just studying the fluorescence
images can sometimes offer a new
insight into a reality which is usually hidden
from the view of the world.
Fundamentals ­ a development both
remarkable and ongoing
Within the last few decades numerous
new techniques such as confocal, deconvolution,
ratio-imaging, total internal reflection
and applications such as the use
of fluorescent proteins (e.g. GFP) have
initiated a real renaissance in the microscopy
field. All of these techniques make
use of fluorescence, a phenomenon first
observed by Sir George Gabriel Stokes in
1852 and physically described by Alexander
Jablonski in 1935 (see box 13).
Compared with today, the number of
widely used fluorochromes was restricted
to just a few in the 1990`s. For example,
nowadys the fluorochrome FITC filter set
for fluorescence microscopy can also be
used for a wide range of different fluorochromes
with green emission spectra.
Why use fluorescence?
Using fluorescence can be compared to
the situation where a teacher asks if the
students have done their homework. The
rapidly changing facial colours of the
"guilty" students provide conclusive "results".
However, fluorescence techniques
are not really for answering questions
such as the above. They help to address
specific questions regarding life science
or materials science specimens and to
visualise the result in a specific colour.
For example, to identify the distribution
of a specific protein within a tissue, a
fluorochrome can be used to mark the
protein via an antibody (immunohisto-
chemistry).
Histological staining procedures for
transmission light microscopy have a long
history in microscopy. One essential advantage
of fluorescence microscopy, however,
is the presence of fluorescent molecules
themselves. Even if a structure is
too small to be resolved by a light microscope,
the emission light remains visible.
Fluorescent molecules act like light
sources that are located within specific
areas of a specimen, indicating their location
with light of a specific colour.
These indicators require energy to emit
light and this is given to the fluorochrome
by the excitation light, provided by the
microscope light source. A specific range
of wavelengths is needed to excite a specific
fluorochrome. For example, a range
of blue wavelengths around 480 nm can
excite the FITC fluorochrome. This involves
using two different light beams
and having to separate them. On the one
hand, we need to direct the light of the
microscope light source onto the specimen
and on the other hand we have to
observe the light that is originating from
the fluorochromes. This separation is
possible due to the "Stokes shift", which
describes the fact that the wavelength of
fluorescent light (emission) is always
longer than that of the excitation. Using
a blue excitation light will thus result in a
green emission for the FITC fluorochrome.
Every fluorochrome has its own
excitation and emission spectra. The microscope
must be perfectly equipped to
visualise this fluorescence accordingly.
Fluorescent molecules
There are two options for using fluorescent
microscopy, depending on what is
Fig. 44 a: Spectral diagram
of a typical fluorochrome
that can be
excited using blue light
and then emit green
fluorescence. This example
shows that both
curves are close to each
other (small Stokes shift)
and also show an area of
spectral overlap.
Fig. 44 b: Cracks within
the resin of an electrical
circuit board.
shining fluorescence Details Basics of light Microscopy & iMaging ˇ 35
being investigated: either the specimen
itself already contains molecules that
show fluorescence; or specific fluorochromes
have to be added to the specimen.
Autofluorescence is widely found in
materials such as plant sections or electrical
circuits, for example. The resin on
circuits is fluorescent and can easily be
inspected under blue excitation (fig. 44b).
The green emission of the resin enables
the detection of the tiniest cracks which
may influence material quality.
Fluorochromes themselves can be divided
into at least three groups. The first
are fluorochromes that require other
molecules, such as antibodies or lectins,
to bind to specific targets. This rapidly
growing group of fluorochromes includes
longstanding ones such as FITC and
TRITC. Most of these fluorochromes are
sold together with the specific targetfinding
molecule (e.g. a goat anti-mouse
IgG antibody Cy5 labelled). Quantum dots
are also partial members of this group
but different in structure and theory.
They are nanometre-sized crystals of purified
semiconductors and exhibit longterm
photo stability, as well as bright
emission. The main difference featurewise
is their capacity to be excited by
wavelengths up to the blue range and
having different emission colours depending
on their size. Due to their flexible
capabilities they can also be used for
direct staining of cells (e.g. cell viability).
The second group contains fluorochromes
that have inherent binding capacities,
such as the DAPI nucleic acid
stain or the DiI anterograde neuron stain.
This group also contains fluorochromes
that change their fluorescent properties
when bound to different amounts of molecules
such as calcium (e.g. Fura-2). This
means that these fluorochromes are used
directly and do not necessarily require a
transportation system such as an anti-
body.
The third group contains fluorescent
proteins produced by organisms themselves
such as GFP. This makes it possible
to set up experiments in an entirely
different way. It is most often used for
live cell imaging or developmental studies
and molecular biology. All fluorochromes
show distinct spectral properties
(fig. 44a) and can often be combined
for a multicolour specimen analysis.
Requirements for a fluorescence
microscopy system
The Light Source
To excite the fluorescence of a fluorochrome,
an intense light source is needed
that provides the necessary excitation
wavelengths to excite the particular
fluorochrome. In the first chapter we described
the most frequently used light
sources for light microscopy and their
alignment. A correctly aligned burner
plays an essential role in creating good
fluorescent images. If the burner is not
correctly aligned, the signal from the
fluorochrome may be excited in a very
weak way and the field of view will not
be homogeneously illuminated, or can
show a bad signal to noise ratio.
Due to the very different specimens
and applications that can be analysed using
fluorescent techniques, there is no
one-size-fits-all strategy. All fluorochromes
are subject to the process of
photo-bleaching, which is the chemical
destruction which takes place during excitation.
Living cells, too, may be damaged
by the intense light. This makes it of
supreme importance to restrict the excitation
brightness and duration to the exact
amount needed. The amount of light
can be efficiently modified with neutral
density filters or a motorised attenuator.
When the light is not needed for excitation,
the shutter is closed. These features
Box 13: What is fluorescence exactly?
Fluorescence activity can be schematically illustrated using the familiar Jablonski diagram, where absorbed
light energy excites electrons to a higher energetic level in the fluorochrome.These lose a proportion of their
energy by vibration and rotation and the rest is then given off as fluorescent light as they return to their original
state after only about 10 ns.
Fig. 45: Light path on a microscope equipped for
fluorescence.
Fig. 46: Characteristics of an Olympus
HQ filter set optimised for GFP. By
using up to 100 layers of an ion
deposition technique with new
substrates and coating materials,
filters can be created with high transmission
and exceptionally sharp cutoff
(tolerances < 2 nm).Autofluorescence
is significantly reduced and the
filters are equipped with a stray light
noise destructor to enhance the
signal-to-noise ratio.
36 ˇ Basics of light Microscopy & iMaging shining fluorescence Details
Fig. 47: Upper row: Images of a double-labelled specimen (DAPI for DNA staining and Alexa 488labelled
antibody for F-Actin staining). The first image shows the detection of the fluorescence using a
filter set with the appropriate excitation filter for DAPI and with a long pass emission filter. Analysed
with a monochrome camera, even the Alexa-488-labelled structures are visible and prevent effective
analysis. The blue and the green image show the same specimen with optimised emission filters,
allowing the separation of each signal.
The lower row shows an example of an image area analysed with an achromat (left), a fluorite (centre)
and an apochromat objective (right) at the same magnification.
Note that the signal intensity and colour transmission increases from achromat to apochromat due to
the increasing NA and quality of the objectives, whereas the background intensity in the upper left
corner remains unchanged.
Box 14: How are filters described?
Filters for fluorescence microscopy are described
using letters and numbers: e.g. BA 460-480. In
this case, BA stands for bandpass filter and the
numbers indicate the 50% cut on and the 50%
cut off (fig. 46, box 14 ­ Figure 1). For a longpass
filter, just the number for the 50% cut on is indicated.
Some companies use a different description
for the same overall filter characteristics (e.g.
470/20), 470 indicating the central wavelength
and 20 indicating the range of the full width at
half maximum (FWHK). The correct transmission
characteristics of the filter can only be provided
using a transmission/wavelength diagram (box 14
­ Figure 1).
can be predefined in motorised microscopes
and help to optimise experi-
ments.
The Filter Sets
In addition to the special light path within
a fluorescence microscope (fig. 45), another
necessity is having a filter which
only permits the required range of excitation
wavelengths to pass through. This
is achieved using an exciter filter with
what are referred to as bandpass filter
characteristics (box 14). After restricting
the light to the particular colour that is
needed to excite the fluorochrome, the
light is directed to the specimen via a dichromatic
mirror (fig. 45).
As indicated by its name, the dichromatic
mirror treats different colours differently.
It reflects light below a given
wavelength and is able to let longer
wavelengths pass through. The excitation
light travels through the objective to
the specimen, acting like a condenser.
This is where the fluorescence phenomenon
takes place. Excited by the light,
the fluorochromes emit the fluorescence
light of longer wavelengths. This is captured
by the objective, moving on to the
dichromatic mirror, now letting the
longer wavelengths pass through. The
last step of filtering is undertaken by the
emission filter (also called a barrier filter,
see fig. 46). This filter restricts the
light colour to best fit the fluorochrome
emission and the question being investigated.
It ensures that no unwanted wavelengths
are observed and analysed. The
emission filter can be designed as a bandpass
filter (precisely restricted to one
spectrum) or as a longpass filter (brighter
in effect, but less optimal due to a reduction
in restricted wavelengths). To help
find the best filter combination for the
fluorochromes in use and the analysis in
mind, a variety of web pages is available
(e.g. www.olympusmicro.com/primer/
java/fluorescence/matchingfilters/index.
html). A straightforward example (below)
will demonstrate how the combination
of filters may differ depending on
the context.
Using the correct filter set ­ an example
If you wish to make a vital analysis of a
cell culture you may choose a Fluorescein-diacetate
(FDA) to stain vital cells.
This fluorochrome is excited by blue light
and will have a green emission. The vital
cells will be the ones appearing green.
The filter set could thus be chosen as follows:
an exciter with BP460-490 bandpath
characteristics, a dichromatic mirror
with DM505 characteristics and an
emission filter with LP510 long path
characteristics. This will result in a bright
green image of all green fluorescent molecules.
So far, so good. The vital cells are
stained. To verify that non-labelled cells
are in fact dead, propidumiodide (PI) dye
may be used. This dye cannot pass
through intact cell membranes. The DNA
of dead cells only will be labelled and appear
red. This means it can be used along
with the FDA. When doing so, however,
the excitation of the FDA with the filter
mentioned will cause some problems. PI
will already be excited by the blue light
and the red emission is also visible. This
is caused by the emission filter because
in this set up, all wavelengths above 510
nm are allowed to pass through. Both
dyes are thus excited and visible. A definite
separation of both signals is not possible
and as we will see later on, this can
cause problems during imaging.
To separately identify both signals
from the cell culture, the emission filter
required for FDA is a bandpass filter (e.
g. BP510-550). This filter will only allow
the green emission of the FDA to pass
through and the emission of the PI will
be blocked. A second filter set can then
be used to analyse the PI signal efficiently
shining fluorescence Details Basics of light Microscopy & iMaging ˇ 37
(e.g. BP530-550 excitation filter, LP575
emission filter, DM570 dichromatic mirror).
This principle also applies to other
multicolour staining procedures. Best results
are achieved when the emission filter
for the detection of the fluorochrome
with the shorter wavelength is a band
pass filter (fig. 47 upper row).
The objectives
The light gathering capacity of the objective
plays an essential role in fluorescence
microscopy. For optimal signal
strength, a high-numerical aperture
(high NA) and the lowest useable magnification
should be employed. For example,
using a 1.4 NA aperture lens instead
of a 1.3 NA lens of the same magnification
will result in a 35% increase in light
intensity, assuming that all other factors
are the same. Furthermore, the type of
glass that is used requires good transmission
for the wavelengths used. Fluorite
or apochromat objectives are used for
that reason. Further enhancement can
be achieved by selecting objectives with
extremely low autofluorescence of the
glass material used (Olympus UIS2 Objectives).
When all of these factors are
provided for ­ high NA, good transmission
and low autofluorescence ­ this ensures
a perfect signal-to-noise ratio (i.e.
a strong signal with low background intensity
(fig. 47 lower row)). Background
noise can also be introduced by the specimen
itself due to fixation, autofluorescence
of the specimen or non-optimised
staining procedures.
Type of camera
The imaging device is one of the most
critical components in fluorescence microscopy
analysis. This is because the
imaging device used determines at what
level specimen fluorescence may be detected,
the relevant structures resolved
and/or the dynamics of a process visualised
and recorded. Numerous properties
are required to use fluorescence microscopy
effectively. These include: high resolution,
extreme sensitivity, cooling, variable
exposure times and an external
trigger function. Generally no single detector
will meet all these requirements in
fluorescence microscopy. Consequently,
the fluorescence microscopist frequently
has to compromise. However, the cameras
used in fluorescence microscopy
should at the very least offer high signal
sensitivity, low noise and the ability to
quantify intensity of intensity distribu-
tion.
Colour cameras are less sensitive than
their monochrome counterparts because
of the additional beam-splitting and
wavelength selection components. Therefore,
monochrome cameras are preferable.
They image the fluorescence intensity
of each fluorochrome separately and
can handle the respective images later on
within a specific colour space using the
appropriate software. The resulting multicolour
images can be displayed, printed
out and analysed or further processed.
Every channel of colour represents the
result of one fluorochrome. This result
can be obtained if the fluorescence filters
are chosen correctly. Returning to our example
of the FDA and PI double labelling:
when using the longpass emission filter to
detect the FDA signal, the red emission of
the PI will also contribute to the resulting
digital image. A monochrome camera
would not differentiate between red and
green or between blue and green as
shown in fig. 47 (first image) ­ it will only
show intensity in grey values. Therefore,
the image will represent the resulting distribution
of both fluorochromes within
the same colour. The use of the bandpass
emission filter as described above will
help in this respect.
Software-tools: Spectral unmixing
A further problem can occur when using
different fluorochromes with overlapping
Box 15: How to measure FRET.
Ratio measurement
An epifluorescence microscope
equipped with highly specific fluorescence
filter sets, a digital camera
system and image analysis software
is required.The ratio between the intensity
of the sensitised acceptor
emission (= FRET signal) and the intensity
of the donor emission is calculated
(box 15, fig. 1). The results
have to be corrected for spectral
cross-talk by spectral unmixing soft-
ware.
Acceptor photo-bleaching.
This effect occurs when the acceptor
is irreversibly destroyed by extensive
illumination from excitation light.After
bleaching of the acceptor fluorophore, FRET is stopped. Excitation of the donor will result in donor fluorescence
emission.After the acceptor photo-bleaching, the donor shows more intensity than before.
Donor-bleaching.
The sample is irradiated with the specific excitation wavelength of the donor. Images of the donor are continuously
acquired and the intensity decay is quantified. FRET decreases the mean lifetime of the donor and
protects the donor from photo damage. As a result, when using FRET, the rate of photo-bleaching is lower;
without FRET, the rate of photo-bleaching is higher.
Fig. 48: Multi-colour image of a GFP/YFP double-labelled sample, before (left) and after spectral
unmixing (right). In this sample, GFP was fused with the H2B histone protein and YFP with tubulin. The
visible result is a pronounced chromatic resolution of both structures now displayed in green and red.
The tubulin on top of the bright green nuclei is even detectable.
38 ˇ Basics of light Microscopy & iMaging shining fluorescence Details
spectra within one sample. The considerable
overlap of excitation and emission
spectra of different fluorochromes is exemplified
by the spectra of enhanced
green fluorescence protein (eGFP) and
enhanced yellow fluorescent protein
(eYFP), two commonly used variants of
the green fluorescent protein (fig. 48 left
side). Even with the use of high quality
optical filters it is not possible to separate
the spectral information satisfactorily.
This means that YFP-labelled structures
are visible with a GFP filter set and
vice versa, affecting the resulting images
significantly and detrimentally. This phenomenon,
known as "bleed-through",
strongly reduces colour resolution and
makes it difficult to draw accurate conclusions.
The solution to overcome this
effect is called spectral imaging and linear
unmixing.
Spectral imaging and linear unmixing
is a technique adapted from satellite imaging
to wide-field fluorescence microscopy.
Using this highly effective method,
it becomes possible to ascertain the specific
emission of the different fluorochromes
to the total signal and to restore
a clear signal for each colour channel,
unaffected by emission from the other
fluorochrome. This is achieved by redistribution
of the intensity (fig. 38 right
side). It is important to note that original
data is not lost during linear unmixing
nor is any additional data added to the
image. The original image information is
all that is used in this procedure. The
overall intensity of pixels is maintained.
Thus, the technique does not result in artificially
embellished images. After unmixing,
quantification analysis not only
remains possible, but also becomes more
precise.
How to use light as a tool
After having laid the fundamental principles
and the microscopic requirements
we can now go deeper into different
state-of-the-art qualitative and quantitative
imaging methods for fluorescence
microscopy. These approaches require
advanced microscopic and imaging
equipment which varies from case to
case. So only general statements can be
made here about the hardware and software
used, along with a description of
the applications and methods.
Life under the microscope
Wherever there is life there are dynamic
processes ­ observation of living cells
and organisms is a tremendous challenge
in microscopy. Microscopy offers different
techniques to reveal the dynamic
processes and movements of living cells.
The use of fluorescent proteins and live
fluorescent stains enable the highly specific
labelling of different organelles or
molecules within a living cell. The intensity
of the emission light of these markers
can be used to image the cells. In addition
to the application protocol and the
fluorescent technique to be used there
are further general considerations to be
aware of.
First of all, there is the definition of
the needs for the specific environmental
conditions and controlling these with regard
to specimen handling. Assuming
you are using a cell line and would like to
analyse processes over time, it may be
necessary to provide appropriate environmental
conditions for these cells. Dynamic
processes in single cells can occur
within the millisecond range ­ such as
shifts in ion concentrations. Or they may
take minutes ­ such as the active or passive
transport of proteins or vesicles. Microscopes
can be equipped with heating
stages and/or minute chambers, or with
complete cultivation chambers to ensure
cultivation of living cells with all the appropriate
parameters on the microscope
while observation is conducted for hours
or days (fig. 49).
Where do all the ions go?
Fluorescent dyes such as FURA, INDO or
Fluo show a spectral response upon binding
Ca2+
ions and are a well established
tool to investigate changes in intracellular
Ca2+ concentration. The cells can be
loaded with a salt or dextran conjugate
form of the dye ­ e.g. by microinjection,
electroporation or ATP-induced permeabilisation.
Furthermore, the acetoxymethylester
of the dyes can be added
to the medium, loaded passively into the
cells and cleaved enzymatically to produce
cell-impermeable compounds.
For the analysis of a typical two channel
FURA experiment it is necessary to
switch between the excitation of 340nm
and 380nm. When observing FURA
loaded cells without any stimulus, Ca2+ is
bound in cell compartments. The FURA
molecules show strong fluorescence at
an emission of 510nm when excited with
380nm, and weak fluorescence when excited
with 340nm. As the cell releases
Ca2+ from storage compartments due to a
reaction to a stimulus, FURA molecules
form complexes with these released Ca2+
ions. The fluorescence signal in the emission
channel of 510nm increases when
excited with 340nm, and decreases when
excited with 380nm. The ratio between
the signals of the two excitation channels
is used to quantify the change of inten-
sity.
Why use a ratio? Lamp fluctuations or
other artificial intensity changes can
cause a false signal which can be misinterpreted
as a change in ion concentration
when intensity is measured in one
channel only. The second drawback of a
single channel analysis is that it displays
the amount of the fluorescence only.
Therefore, thicker parts may look
brighter than smaller parts of a cell;
however, they simply contain more fluorochromes
due to the larger volume. Physiologically
relevant changes in ion concentration
in small areas such as growth
cones of a neuron may then not be visible
over time because they are too dim in
fluorescence compared to the bright centre
of the cell. After background subtraction,
the calculation of a ratio between
two channels corrects the result for overall,
artificial fluctuations and specimen
thickness. Following a calibration procedure,
even quantitative results are obFig.
49: A climate chamber offers features like
control of temperature, humidity and CO2 content
of the atmosphere.
Fig. 50: Autofluorescence of an apple slice
(Lieder). The image was taken with an Olympus
FluoView FV1000 and the TCSPC by PicoQuant.
The colour bar in the lower right corner is a key,
showing the distribution of the various lifetimes
within the image.
shining fluorescence Details Basics of light Microscopy & iMaging ˇ 39
tainable. There is a range of fluorochromes,
with different spectral
properties for numerous different ions,
available on the market. Microscopical
systems with fast switching filter wheels
and real time control permit investigations
even in the millisecond range.
Light as a ruler
Many processes in a cell are controlled by
inter- and intra-actions of molecules: e.g.
receptor-ligand interactions, enzyme-substrate
reactions, folding/unfolding of molecules.
Receptor-ligand interactions, for
example, occur in the very close proximity
of two proteins in the Angstrm range.
Colocalisation studies do not reveal interactions
of molecules in the Angstrm
range because the spatial resolution of a
light microscope is limited to 200 nm.
When using a light microscope, how can
the proximity of two molecules in the Angstrm
range be proven beyond the physical
limits of light microscopy? Fluorescence
Resonance Energy Transfer (FRET)
helps to find an answer to this question.
FRET is a non-radiative energy transfer
between two different fluorophores. The
first fluorophore (the donor) is excited by
light. The donor transfers its energy to the
second fluorophore (the acceptor) without
radiation, meaning without any emission
of photons. As a result, the acceptor is excited
by the donor and shows fluorescence
("sensitised emission"). The donor is
quenched and does not show any fluorescence.
This radiation-free energy transfer
occurs within the very limited range of 1­
10 nm distances between the donor and
the acceptor. A positive FRET signal provides
information about the distance between
the FRET partners and can be
quantified as FRET efficiency. When no
FRET signal is achieved, there may be
many reasons for that: e.g. too much distance
between the FRET partners, insufficient
steric orientation, insufficient dipole
orientation, insufficient spectral
overlap between the emission spectrum of
the donor and the excitation spectrum of
the acceptor. See box 15.
How long does a fluorochrome live? count
the photons!
When a fluorochrome is excited, it is
shifted to a higher energy level. The lifetime
of a fluorophore is the average
amount of time (in the nanosecond/picosecond
range) that it remains at the higher
energy level before it returns to the
ground state. A fluorochrome`s lifetime is
a highly specific parameter for that particular
fluorochrome. It can be influenced
easily by changes of environmental parameters
(e.g. pH, ion concentration, etc.),
by the rate of energy transfer (FRET) or
by interaction of the fluorochrome with
quenching agents. Fluorochromes often
have similar or identical spectral properties,
therefore, analysing a fluorochrome`s
lifetime is critical to distinguishing the localisation
of those fluorochromes in a cell
(fig. 50). Here, the different experimental
setups are referred to as FLIM ­ Fluorescence
Lifetime Imaging Microscopy.
Fluorescence Lifetime Imaging Microscopy ­
FLIM.
There are different techniques for fluorescence
lifetime imaging microscopy
available on the market, for both widefield
and confocal microscopy. Here we
focus on Time-Correlated Single Photon
Counting (TCSPC). The fluorochrome is
excited by a laser pulse emitted by a
pulsed laser diode or femto-second
pulsed Ti:Sa laser. A photon-counting
photo- multiplier or single photon avalanche
diode detects the photons emitted
from the fluorophore. The time between
the laser pulse and the detection of a
photon is measured. A histogram accumulates
the photons corresponding to
the relative time between laser pulse and
detection signal. Every pixel of the FLIM
image contains the information of a complete
fluorescence decay curve. If an image
is composed of three fluorochromes
with different lifetimes, distribution of all
Fig. 51: Photoconversion: In Miyawaki`s lab, Ando et al [1] succeeded in cloning the gene that encodes
the Kaede fluorescence protein of a coral. By irradiation with UV light, the green fluorescence can be
converted to red fluorescence. By using excitation wavelengths in the blue and green spectral range,
the movement and distribution of the protein can be observed in a living cell. The figure shows the
protein diffusion in Kaede-expressing HeLa cells. Stimulation via 405 nm laser in the region of interest
converts the Kaede protein from green to red emission. Confocal images were recorded every three
seconds using 488/543 nm laser excitation, showing the activated Kaede protein spreading throughout
the HeLa cell. By using two synchronised scanners (one for photo-conversion, one for image acquisition),
fluorescence changes that occur during stimulation can be observed.
Data courtesy of: R. Ando, Dr A. Miyawaki, RIKEN Brain Science Institute Laboratory for Cell Function
Dynamics.
Fig. 52: Photoactivation of PAGFP
expressing HeLa cells. The
PA-GFP was activated by 405
diode laser (ROI) and images of
the PA-GFP distribution within
the cell were acquired at 1 second
intervals using the 488nm
laser.
Data courtesy: A. Miyawaki, T.
Nagai, T. Miyauchi,
RIKEN Brain Science Institute
Laboratory for Cell Function
Dynamics.
40 ˇ Basics of light Microscopy & iMaging shining fluorescence Details
bleached, the other remains undamaged
and acts as a reference label. The population
of bleached fluorochromes can be
identified by subtraction of the image of
the bleached dye from the image of the
unbleached dye.
FAUD = Fluorescence Application Under
Development!
We are looking for inventors of new fluorescence
techniques ­ please keep in
touch!
From black to green, from green
to red ...
If we sunbathe for too long, the colour of
our skin changes from white to red and
is referred to as sunburn. If we take a UV
laser and irradiate a cell with Kaede protein,
the colour of the Kaede protein
changes from green to red ­ and we call
this photo-conversion (fig. 51). A confocal
microscope can be used to stimulate
fluorochromes in selected parts of the
cell. The Olympus FluoView FV1000 even
offers stimulation of one area with one
laser whilst observing the result in a different
channel with a second laser simultaneously.
A recently developed photoactivatable
GFP mutant, PA-GFP, can be
dyes can be shown as three different colours
(fig. 50).
Time-resolved FRET microscopy
With FRET, the lifetime of the donor depends
on the presence or absence of radiation-free
energy transfer (see above).
Therefore, time-resolved FRET microscopy
is a technical approach for quantitative
measurement of the FRET efficiency.
FRAP, FLIP and FLAP
FRAP, FLIP and FLAP are all photobleaching
techniques. By using the laser
scanner of a confocal microscope, fluorochromes
(which are bound to a specific
protein, for example) in a selected region
of a stained cell can be bleached (destroyed).
As a result, the fluorochrome
does not show any appropriate fluorescence.
Other proteins that are also labelled,
but where the fluorescence was
not bleached, can now be observed during
movement into the previously
bleached area. Dynamic processes, such
as active transport or passive diffusion in
a living cell cause this movement. Therefore
the intensity of the fluorochrome recovers
in the bleached area of the cell.
FRAP = Fluorescence Recovery After
Photobleaching
After bleaching of the fluorochrome in a
selected region of a cell, a series of images
of this cell is acquired over time.
New unbleached fluorochromes diffuse
or are transported to the selected area.
As a result, a recovery of the fluorescence
signal can be observed and measured.
After correction for the overall
bleaching by image acquisition, a curve
of the fluorescence recovery is obtained.
FLIP = Fluorescence Loss In Photobleaching
The fluorophore in a small region of a
cell is continuously bleached. By movement
of unbleached fluorophores from
outside of the selected region into the
bleaching area, the concentration of the
fluorophore decreases in other parts of
the cell. Measuring the intensity loss,
outside the bleached area results in a decay
curve. A confocal laser scanning microscope
which incorporates one scanner
for observation and one scanner for
light stimulation is especially appropriate
for this technique. A simultaneous
scanner system allows image acquisition
during continuous bleaching.
FLAP = Fluorescence Localization After
Photobleaching
The protein of interest is labelled with
two different fluorochromes. One is
activated by irradiation using a 405 nm
diode laser (fig. 52). The intense irradiation
enhances the intensity of the fluorescence
signal of PA-GFP by 100 times. PAGFP
is a powerful tool for tracking
protein dynamics within a living cell
(fig. 52).
Imaging at the upper limits of
resolution
The bleaching and photoactivation techniques
described above can be undertaken
easily using a suitable confocal laser
scanning microscope. The laser is a
much more powerful light source than a
fluorescence burner based on mercury,
xenon or metal halide. The galvano mirror
scanners, in combination with acousto-optically
tuneable filters (AOTF), permit
concentration of the laser onto one
or more selected regions of the cell without
exciting any other areas in that cell.
This makes the laser scanning microscope
the preferred tool for techniques
which use light as a tool. Additionally,
the confocal principle removes out-of-focus
blur from the image, which results in
an optical section specifically for that focal
plane. This means that high resolution
along the x, y and z axes can be obtained.
A confocal microscope is an ideal
system solution for researchers who
want to use light not only for imaging,
but also as a tool for manipulating fluor-
ochromes.
To achieve an even thinner optical section
(for a single thin section only and not
a 3-D stack), Total Internal Reflection Microscopy
(TIRFM) may be the technique of
choice (for more information on TIRFM,
see box 4 or at www.olympusmicro.com/
primer/techniques/fluorescence/tirf/tirfhome.html).
The evanescent field used for
this technique excites only fluorochromes
which are located very closely to the coverslip
(approx. 100­200 nm). Fluorochromes
located more deeply within the
cell are not excited (fig. 53). This means
that images of labelled structures of membranes,
fusion of labelled vesicles with the
plasma-membrane or single molecule interactions
can be achieved with high z
resolution (200 nm or better, depending
on the evanescent field).
References
[1] R. Ando, H. Hama, M. Yamamoto-Hino, H. Mizuno,
A. Miyawaki , PNAS Vol. 99, No. 20,
12651­12656 (2002)
[2] G. H. Patterson, J. Lippincott-Schwartz, Science,
Vol 297, Issue 5588, 1873­1877 (2002)
Fig. 53: Imaging at the Outer Limits of Resolution.
(a) Dual emission widefield
(b) TIRF images acquired with a 488nm
laser
green Fitc-Actin
red DyeMer 605-EPS8.
Courtesy of M. Faretta, Eup. Inst. Oncology, Dept.
of Experimental Oncology, Flow Cytometry and
Imaging Core, Milan, Italy.
shining fluorescence Details Basics of light Microscopy & iMaging ˇ 41