Theory of Confocal Microscopy
Spectral Bleed-Through Artifacts in Confocal Microscopy
Bleed-through (often termed crossover or crosstalk) of fluorescence emission, due to the very
broad bandwidths and asymmetrical spectral profiles exhibited by many of the common
fluorophores, is a fundamental problem that must be addressed in both widefield and laser
scanning confocal fluorescence microscopy. The phenomenon is usually manifested by the
emission of one fluorophore being detected in the photomultiplier channel or through the filter
combination reserved for a second fluorophore. Bleed-through artifacts often complicate the
interpretation of experimental results, particularly if subcellular co-localization of fluorophores is
under investigation or quantitative measurements are necessary, such as in resonance energy
transfer (FRET) and photobleaching (FRAP) studies.
Imaging specimens having two or more fluorescent labels (or plant tissue sections exhibiting a
high degree of autofluorescence) is often complicated by the bleed-through or crossover of
fluorescence emission, unless the spectral profiles of the fluorophores are very well separated.
For example, when double labeling with the traditional green and red probes, fluorescein and
rhodamine, bleed-through can only be reduced by using optimized fluorescence filter sets (and/or
photomultiplier detector slit widths), but is never completely eliminated. This effect is due to the
fact that these dyes have very broad absorption and emission spectra that exhibit a significant
degree of overlap. Thus, excitation of fluorescein using the 488-nanometer spectral line of an
argon-ion laser will also produce excitation of rhodamine, although to a lesser degree.
Furthermore, fluorescein emission will be detected in the photomultiplier channel or widefield
filter set reserved for rhodamine.
Spectral bleed-through in a thin section of mouse intestine labeled with Alexa Fluor 488 and Cy3
(a cyanine dye), and imaged with a laser scanning confocal microscope having adjustable
photomultiplier detector slits is illustrated in Figure 1. These fluorophores exhibit absorption and
emission spectra similar to fluorescein and rhodamine, although with slightly different peak
wavelengths and somewhat narrower bandwidths. The pair of images illustrated in Figures 1(a)
and 1(b) were obtained by simultaneous lateral scanning of the specimen using an argon-ion
laser (488 nanometers; Figure 1(a)) and a green helium-neon laser (543 nanometers; Figure
1(b)). Note the Alexa Fluor 488 fluorescence bleed-through apparent in the Cy3 detection
channel (Figure 1(b)), which is manifested by yellow overlap regions in the final merged image
(Figure 1(c)). This artifact can be easily confused with co-localization of the fluorophores. By
sequentially scanning the specimen with the individual lasers and detecting fluorescence in each
channel to coincide with laser illumination (Figure 1(d) and 1(e); discussed in more detail
below), spectral bleed-through is minimized (compare Figure 1(c) to Figure 1(f)) to produce a
more accurate merged image of fluorophore distribution.
The Alexa Fluor 488 detection channel (see Figures 1(a) and 1(d)) photomultiplier slit has been
set to a 30-nanometer bandwidth (ranging from 500 to 530 nanometers) that encompasses the
primary probe emission peak, but does not capture a significant amount of fluorescence from
Cy3 emission. As a result, the Alexa Fluor 488 channel does not detect Cy3 fluorescence
emission at normal voltage and gain settings, regardless of whether the instrument is scanning
simultaneously or sequentially. In contrast, to capture sufficient fluorescence emission from the
cyanine dye, the Cy3 detection channel photomultiplier slit must be set to a wider range (555 to
625 nanometers), which also allows the longer wavelengths of Alexa Fluor 488 emission to
register on the detector. Thus, adjusting the detector slits (or interference filters) alone will not
adequately reduce bleed-through with fluorophores having this high degree of spectral overlap.
In many cases, bleed-through can be controlled by judicious choice of fluorophores having wellseparated absorption and emission spectra. Substituting Alexa Fluor 568 for Cy3 in this instance
(a 35-nanometer difference in emission peak wavelength) would lead to slightly less efficient
excitation with the helium-neon laser, but would significantly reduce bleed-through.
A comparison of spectral overlap for a series of Alexa Fluor dye combinations potentially useful
in dual color labeling experiments is presented in Figure 2. All of the emission spectra are
normalized for comparison, and the overlap regions are indicated by gray shading. In Figure
2(a), the emission spectra for the green fluorescent Alexa Fluor 488 and yellow-orange
fluorescent Alexa Fluor 555 indicate clear separation of the peak wavelengths, which are also
easily distinguished by the human eye. However, the moderate level of spectral overlap (gray
shaded area) illustrates that there is a considerable amount of emission from Alexa Fluor 488 at
the peak emission wavelength of Alexa Fluor 555 (denoted by a black line running from the
emission peak to the abscissa). This high level of signal bleed-through renders separation of the
probes difficult in situations where the fluorescence emission intensity of Alexa Fluor 488 is
significantly greater than that of Alexa Fluor 555, which can occur due to a number of factors
including large differences in label target population. These probes are efficiently excited by the
488-nanometer and 543-nanometer spectral lines of the argon-ion and green helium-neon lasers,
respectively.
The spectral overlap between Alexa Fluor probes decreases as the bandwidth between emission
maxima increases, as illustrated in Figure 2(b). In this case, Alexa Fluor 488 and orange
fluorescent Alexa Fluor 594 demonstrate a reduced level of overlap when compared to Figure
2(a). Both dyes are easily distinguishable to the human eye and the low degree of spectral
overlap should yield good results with minimal bleed-through in dual labeling experiments,
provided the concentrations of each probe are similar in the specimen. Alexa Fluor 594 is most
efficiently excited by the 568-nanometer line of a krypton-argon laser or the 594-nanometer line
of a yellow helium-neon laser. Perhaps the best spectral separation in visible-light emitting Alexa
Fluor dyes is the comparison between Alexa Fluor 488 and Alexa Fluor 633 depicted in Figure
2(c). There is virtually no spectral overlap between these dyes and bleed-through artifacts should
be absent, even in specimens containing excessive levels Alexa Fluor 488. The Alexa Fluor 633
probe is efficiently excited by the 633-nanometer spectral line of the red helium-neon laser or the
647-nanometer line of the krypton-argon laser.
In describing spectral overlap artifacts, the terms bleed-through, crossover, and crosstalk are
often used interchangeably. Although bleed-through and crossover refer to the spillover of
fluorescence emission (or excitation) from one filter set or photomultiplier detection channel into
another, crosstalk is widely employed in the filter industry to describe the minimum attenuation
level of two filters placed together in series. The crosstalk value of a fixed filter combination is
important for manufacturers when matching fluorescence excitation, emission, and dichromatic
beamsplitter filters in a set, but is somewhat different from spectral bleed-through in confocal
microscopy. Therefore, care should be used in selecting a term to describe the overlap of
fluorescence emission from one detector channel into another, and the investigator should be
aware of differences in terminology.
Spectral crossover can occur during both excitation and emission of the synthetic fluorophores
and fluorescent proteins commonly utilized in confocal microscopy. In general, crossover
between fluorophore absorption (or excitation) spectral profiles occurs toward the blue end
(shorter wavelengths) of the spectrum, whereas crossover between fluorescence emission spectra
occurs in the red (longer wavelengths) region. For example, emission from a green fluorophore
can often be detected through red emission filters, but a red dye is only seldom imaged through a
green emission filter. This is due to the fact that the absorption and emission spectra for most
dyes are not symmetrical, but usually display long, skewed tails that cover regions of tens to
hundreds of nanometers. Absorption spectra are generally skewed towards shorter wavelengths
whereas emission spectra are skewed towards longer wavelengths. For this reason, multicolor
fluorescence imaging should be conducted with the reddest (longest wavelength peak emission)
dye imaged first, using excitation wavelengths that are only minimally absorbed by the skewed
spectral tails of the bluer dyes.
Specimen Labeling Precautions to Reduce Bleed-Through
For determining the physical and spatial location, as well as association between biomolecules
and subcellular structures of interest, labeling specimens with two or more probes is a highly
effective technique in fluorescence microscopy. In this regard, confocal microscopy (when using
two or more lasers) is well-suited to multiple labeling techniques because of the ability to
differentiate between fluorescence emission spectra of individual fluorophores by directing the
signals to several detection channels. There are, however, numerous limitations that must be
considered when performing multiple labeling experiments, either with confocal or traditional
widefield fluorescence microscopy.
The fluorescence emission spectral profiles of common fluorophores differ significantly with
regards to bandwidth, peak emission wavelength, symmetry, and number of maxima. In multiply
labeled specimens, if the degree of labeling and the intensity of fluorescence emission from the
fluorophores is not equally balanced, brighter signals can overwhelm and penetrate the barrier
filters of channels reserved for weaker fluorophores or those with less abundant targets. The
result is too often a significant contribution from the overstained fluorophore to the image
recorded in the channel reserved for a lower intensity probe. The fluorescence intensity from
probes such as fluorescein and rhodamine (as well as their relatives) should be similar and are
adjusted according to the quantity of dye in the specimen and the illumination source. For
example, at equal concentrations, rhodamine is excited more effectively than fluorescein in
widefield fluorescence (by a factor of 10) due to the bright 546-nanometer emission line in the
mercury arc spectrum.
Careful balancing of fluorophore emission cannot be overemphasized during specimen
preparation. During imaging, the apparent balance can be adjusted by altering the gain,
photomultiplier voltage, or laser power for the individual detection channels in confocal
microscopy or through the use of neutral density filters with arc-discharge lamps in widefield
fluorescence. However, simply balancing the amount and color of the fluorophore with target
abundance during specimen preparation will alleviate many problems when imaging and, in
general, lead to superior images. This concept is illustrated in Figure 3 for a dual label situation
using fluorescein and rhodamine (only the fluorescence emission spectra are presented). Each
detection channel window is outlined with a colored box in Figure 3, with the fluorescein
channel collecting signal between 490 and 555 nanometers (red box) and the rhodamine channel
set to a range of 570 to 665 nanometers (blue box). Note that the rhodamine channel has a
significantly wider bandwidth in order to gather signal from the skewed long-wavelength tail
region of the emission spectrum.
If the fluorophore concentrations are balanced such that both fluorophores produce similar
emission intensities, then the amount of bleed-through from one channel into another is shown as
the gray overlap areas labeled 1 and 2 in Figure 3. Clearly, a significantly higher level of the
fluorescein emission crosses into the rhodamine channel than vice versa. The degree of bleedthrough could be reduced, in this case, by decreasing the concentration of fluorescein while
maintaining that of rhodamine constant. In situations where the level of fluorescein labeling
greatly exceeds that of rhodamine, the amount of bleed-through becomes more severe, as
illustrated by the increasing overlap in areas labeled 3 and 4 (note that the concentration of
fluorescein is doubled and quadrupled, respectively). At the highest fluorescein concentration
illustrated in Figure 3, the amount of bleed-through emission (area 4) collected in the rhodamine
channel almost equals that of the target fluorophore emission itself.
When selecting fluorescent probes for multiply labeled specimens, the brightest and most
photostable fluorophores should be reserved for the least abundant cellular targets. In general,
dyes with green and red fluorescence emission tend to be brighter than those emitting in the blue
and far-red portions of the spectrum. However, even when probe concentrations are carefully
controlled with regards to quantum yield, photostability, illumination source, and target
abundance, the level of signal crossover and bleed-through generally remains between 10 and 15
percent unless the emission spectral maxima are separated by 100 to 150 nanometers or more.
Furthermore, the localized environment significantly influences fluorophore absorption and
emission spectral profiles, so the spectra published by manufacturers of isolated fluorophores in
solution can differ markedly from those observed under actual experimental conditions. In
experiments involving two or more fluorophores, the investigator should always examine singlestained control specimens using the filter sets for the other fluorophores to assure that the level of
bleed-through is minimal. In many cases, the primary cause of bleed-through is unequal staining
by the fluorophores, which should be corrected by altering the staining protocol rather than
adjusting exposure times in the microscope or performing extensive image rehabilitation with
post-acquisition processing.
Instrumental Approaches to Fluorophore Emission Separation
The traditional fluorescein and rhodamine fluorophores, along with their Alexa Fluor and
cyanine dye relatives, are popular combinations for dual labeling in both widefield and confocal
fluorescence microscopy. In fact, the fluorescein and rhodamine probes have been so widely
used that most microscope and aftermarket filter manufacturers have specialized filter sets
named after their reactive isothiocyanate intermediates (FITC and TRITC, respectively).
Widefield microscopes equipped with arc-discharge lamps generally excite these fluorophores
using the 495 and 546-nanometer spectral peaks from mercury burners, while modern confocal
microscopes employ the Argon-ion laser 488-nanometer spectral line and the 543-nanometer line
of the green helium-neon laser.
The fluorescein and rhodamine fluorophore combination, however, often yields less than optimal
results in confocal microscopes that are equipped with only an argon-ion or krypton-argon laser,
neither of which emit the appropriate spectral lines at wavelengths (between 530 and 560
nanometers) for the most efficient excitation of rhodamine-class fluorophores. Instead,
fluorophores with absorption peaks residing at longer wavelengths, such as Alexa Fluor 568,
Alexa Fluor 594, or Texas Red (578, 590, and 596-nanometer absorption maxima, respectively)
should be employed with the krypton-argon laser. In some cases, microscopes equipped with a
multi-line argon-ion laser are utilized to excite fluorescein and rhodamine derivatives with the
488 and 514-nanometer lines. However, only fluorescein and its derivatives are efficiently
excited at 488 nanometers, and most have highly skewed emission tails that exhibit significant
overlap with the emission spectrum of rhodamine-class probes. In addition, at 514 nanometers,
both fluorescein and rhodamine (and their relatives) are almost equally excited, which leads to a
significant amount of spectral bleed-through.
Choosing two or more fluorophores for simultaneous excitation by a single laser wavelength
places severe restrictions on the experimental parameters. The fluorophore absorption spectra
must overlap significantly to permit simultaneous excitation, but for most combinations, one
fluorophore will be excited to a much higher degree than the others. In addition, the emission
spectra should have a minimum degree of overlap in order for the signal from each fluorophore
to be effectively separated with carefully chosen bandpass filters. Furthermore, the emission
spectra of the fluorophores having the shortest excitation wavelengths should not overlap with
the absorption spectra of other fluorophores to avoid energy transfer artifacts. Due to the these
strict requirements, and the general lack of suitable fluorophores, most investigators choose
confocal microscopes with two or more lasers for multiple labeling experiments.
Both widefield and confocal microscopes use filters constructed of coated glass or multiple thinfilm interference layers to separate fluorescence emission signals. In these instruments, a
dichromatic beamsplitter diverts light into two pathways, one reflected onto the specimen and
the other transmitted to the detector. A bandpass barrier or emission filter refines light
transmitted by the beamsplitter before it is gathered by the detector, either a charge-coupled
device (CCD) or photomultiplier. The dichromatic mirror and barrier filter perform the functions
of limiting excitation light from reaching the detector and collecting the maximum amount
emission signal from a single fluorophore without allowing emission from other fluorophores to
contaminate the signal. Thus, the separation of fluorescence emission is highly dependent upon
filter characteristics. Well-spaced and/or excessively broad emission spectral profiles can take
advantage of wide bandpass emission filters covering a hundred nanometers or more, and
produce bright images due to the high levels of fluorescence signal passing through the filter.
These high signal levels can be controlled through the use of neutral density filters in widefield
fluorescence or by reducing laser power in confocal microscopy. In contrast, closely overlapping
emission spectral profiles require very narrow bandpass filters for optimum separation, but at the
cost of lower signal levels.
The major disadvantage of using filter sets to detect fluorescence emission is that all photons
collected by a filter are treated in the same manner, regardless of source. Transmission of
unwanted wavelengths through a typical filter set is generally about 10 percent of the total
number of photons passing through. In order to combat this dilemma, confocal manufacturers are
developing multispectral instruments that are capable of distinguishing the source of
fluorescence emission based on the spectral profiles of the individual fluorophores (a technique
often referred to as emission fingerprinting). These advanced instrumental designs, which will
probably become the workhorses of modern confocal microscopy, utilize one or more of several
available options to discriminate between emission in experiments where it is not possible or
feasible to choose non-overlapping fluorescent probes. The simplest approach in multispectral
imaging is to dissect fluorescence emission into its component wavelengths with a finely lined
dispersion grating and limit collection to specific regions using a movable slit. A second method
uses a prism or acousto-optic deflector instead of the dispersion grating to perform the same
function, while a third design couples the dispersion grating to a multi-channel photomultiplier
that collects a series of narrow (10-nanometer) wavelength bands. Each multispectral microscope
configuration has its advantages and limitations, but all three are capable of wavelength
discrimination down to a resolution of only a few nanometers.
Multispectral confocal microscopy is the method of choice when imaging multiple fluorescent
protein variants in a single experiment or when naturally occurring and fixative-induced
fluorescence (usually collectively termed autofluorescence) masks the signals of target
fluorophores. Autofluorescence and other forms of unwanted emission signal often cross over
into several channels, a factor that can complicate quantitative analysis. Furthermore, when four
or more fluorophores are used in a single experiment, a significant degree of spectral overlap,
along with the resulting bleed-through artifacts, is inevitable. In this case, multispectral imaging
is one of the most promising techniques for eliminating bleed-through. Among the drawbacks of
multispectral imaging is the reduction in signal that accompanies the narrow detection windows
often required for separation of fluorescence emission. One of the emerging instrumental
approaches to eliminate bleed-through (termed fluorescence lifetime imaging microscopy, or
FLIM) is based on gating fluorescence output in lifetime studies where fluorophores with
similar spectral profiles can be distinguished due to their individual decay characteristics.
Although multispectral imaging is a promising new methodology for dealing with spectral bleedthrough, the instrumentation required is currently very expensive. Confocal microscopes
equipped with acousto-optic tunable filters (AOTFs) or similar rapid laser line switching devices
can be effectively utilized to sequentially scan specimens using a technique known as high-speed
channel switching or multitracking. This approach enables the sequential excitation and
collection of emission from individual fluorophores in order to reduce or eliminate bleedthrough, but does not solve the problem when both emission and absorption spectra overlap
extensively. In practice, the rapid switching of laser lines, either individually or for the entire
frame, is coupled to serial detection by each channel as its target fluorophore is stimulated (see
Figure 4). For the example illustrated in Figure 4, using Alexa Fluor 488 and Cy3, the 488nanometer argon-ion laser line employed to excite Alexa Fluor 488 is turned on as the laser scans
across the line in the x-direction while simultaneously collecting Alexa Fluor 488 emission using
the appropriate filter set for the channel detector. On the return path, the 488-nanometer line is
turned off and the Cy3 probe in the specimen is excited with the 543-nanometer helium neon
laser line, again collecting only fluorescence emission using a Cy3-compatible filter set for the
second channel detector. This scanning sequence avoids exciting both fluorophores
simultaneously and is one of the most effective methods available to control bleed-through,
particularly when the choice of emission filters is limited.
Sequential frame scanning with the concurrent collection of fluorescence signal for each
individual channel is an excellent method to generate very nice images with fixed specimens that
are immobilized. However, when collecting images from living cells that have been labeled with
two or more fluorophores, even a small degree of motion by subcellular structures will
compromise the images and decrease the accuracy of co-localization investigations. Therefore,
for live-cell investigations it is more prudent to use fast multitrack line scanning to rapidly
switch between individual laser lines, collecting image information for each channel one line at a
time as the frame is being recorded. Confocal microscopes equipped with AOTF laser controllers
are capable of alternate line scan image collection at speeds that rival or exceed those obtained
when scanning two or three channels simultaneously on older instruments.
Bleed-Through in Fluorescent Proteins
Fluorescent proteins, which are now widely available in emission wavelengths ranging from blue
to the far red, are very effective for live-cell imaging studies using confocal microscopy, albeit
with some trade-offs. For example, the rate of data acquisition is reduced when more than two
species are employed in an experiment, and each fluorescent protein usually requires
substantially different image collection conditions. Addressing spectral bleed-through is often far
more complicated with fluorescent proteins than traditional synthetic probes, especially because
the emission spectra of the former tend to be quite broad. In addition, because different
fluorescent proteins vary in relative brightness, each color may require different signal
integration times. The enhanced cyan and blue varieties (ECFP and EBFP) are very dim
compared to green and yellow fluorescent proteins, requiring longer collection times that saturate
the brighter fluorophores. Thus, when using fluorescent proteins, the experimental requirements
must be balanced with the choice of fluorophore to determine combinations that have similar
emission intensities with well-separated spectral characteristics.
An elegant example of bleed-through management in confocal imaging using fluorophores with
highly overlapping spectra is the simultaneous detection of enhanced green fluorescent protein
(EGFP) and enhanced yellow fluorescent protein (EYFP) in living cells. In this case, the yellow
fluorescent protein image should be collected first using the 514-nanometer spectral line of an
Argon-ion laser, which only marginally excites the green fluorescent protein (see Figure 5). For
optimal detection, the photomultiplier detector slits should be set to a narrow bandwidth region
extending only 20 nanometers (530 to 550 nanometers; Figure 5(b)) or a similar bandpass barrier
filter can be employed. However, the exact size of the emission filter bandwidth is not critical
because only the yellow fluorescent protein is excited.
In the second sequential scan, the 477-nanometer line of the argon-ion laser is used to image the
green fluorescent protein together with a very narrow 10-nanometer bandpass (490-500
nanometers; Figure 5) emission filter or slit bandwidth. Because the emission collection is
critical in this instance, it is far easier to perform this imaging sequence using slits to exclude
EYFP bleed-through at the expense of signal intensity. Note that although the 488-nanometer
argon-ion laser spectral line is closer to the excitation maximum of enhanced green fluorescent
protein, it is too close to the emission filter (or slit) bandpass region to exclude reflected laser
light that will interfere with image collection.
Currently, the brightest fluorescent proteins are green and yellow, but successful discrimination
between these species is often hampered by the fact that their spectral peaks are separated by
only 25 nanometers. As discussed above, dual or multicolor experiments using the green and
yellow fluorescent combination are possible, but bleed-through between filter sets and/or the
necessity to radically reduce detection slit bandwidths presents a significant problem. More
commonly, yellow fluorescent protein is coupled with cyan fluorescent protein for dual color
imaging experiments. Although cyan fluorescent protein is not much brighter than the blue
species, it has two advantages in that it can be excited with the 458-nanometer argon-ion laser
line commonly available on confocal microscopes, and is much more resistant to photobleaching
than blue fluorescent proteins.
Combinations of DsRed with green or yellow fluorescent proteins also yield adequate spectral
separation, but biological artifacts, such as slow maturation and aggregate formation, often
complicate studies with red fluorescent protein species. Second-generation red fluorescent
proteins should solve the biological problems to produce excellent candidates for multi-labeling
experiments. Newer fluorescent proteins with emission profiles extending into the far red and
near-infrared are on the horizon. These fluorophores should be able to take advantage of the red
helium-neon 633-nanometer spectral line currently available in many high-end confocal systems.
Quantum Dots
Quantum dots, which are semiconductor nanocrystals coated with a hydrophilic polymer shell
and conjugated to antibodies or other biologically active moieties, enjoy several advantages not
shared by most traditional fluorophores. Unlike typical organic fluorochromes or fluorescent
proteins, which display highly defined spectral profiles, quantum dots have an absorption
spectrum that increases steadily with decreasing wavelength. Also in contrast, the fluorescence
emission intensity is confined to a symmetrical peak with a maximum wavelength that is
dependent on the dot size, but independent of the excitation wavelength. As a result, the same
emission profile is observed regardless of whether the quantum dot is excited at 300, 400, 500, or
600 nanometers, but the fluorescence intensity increases dramatically at shorter excitation
wavelengths. For example, the extinction coefficient for a typical quantum dot conjugate that
emits in the orange region (605 nanometers) is approximately 5-fold higher when the
semiconductor is excited at 400 versus 600 nanometers. The full width at half maximum value
for a typical quantum dot conjugate is about 30 nanometers, and the spectral profile is not
skewed towards the longer wavelengths (having higher intensity "tails"), such is the case with
most organic fluorochromes. The narrow emission profile enables several quantum dot
conjugates to be simultaneously observed with a minimal level of bleed-through.
In confocal microscopy, quantum dots are excited with varying degrees of efficiency by most of
the spectral lines produced by the common laser systems, including the argon-ion, heliumcadmium, krypton-argon, and the green helium-neon. Particularly effective at exciting quantum
dots in the ultraviolet and violet regions are the new blue diode and diode-pumped solid-state
lasers that have prominent spectral lines at 442 nanometers and below. The 405-nanometer blue
diode laser is an economical excitation source that is very effective for use with quantum dots
due to their high extinction coefficient at this wavelength. Another advantage of using these
fluorophores in confocal microscopy is the ability to stimulate multiple quantum dot sizes (and
spectral colors) in the same specimen with a single excitation wavelength, making these probes
excellent candidates for multiple labeling experiments. Quantum dots are currently available
with emission maxima extending from 525 to 705 nanometers in 20 to 40-nanometer increments,
and a near-infrared probe having an emission peak centered at 800 nanometers. By carefully
choosing the appropriate quantum dot combination (for example, 525, 585, and 655; see Figure
6), confocal experiments can be conducted with a single laser and dramatically reduced bleedthrough artifacts when compared to traditional fluorophores.
Bleed-Through Correction
Because the number of fluorescent signals in multi-labeling experiments can easily exceed the
discrimination capabilities of the detection system, regardless of the instrument sophistication
level, post-acquisition image processing is often the only alternative to bleed-through correction.
A series of control specimens should be prepared when conducting multiple labeling with two or
more fluorophores in order to minimize the confusion in experimental results due to bleedthrough artifacts. The most important controls are preparing the specimen without secondary
antibodies or synthetic fluorophores (the background control) and labeling the specimen with
each fluorophore separately (the bleed-through controls). The background control should be
examined independently with each laser and detection channel to set the limits of signal gain and
offset that should be employed for the final imaging series. All channels that will be used to
image a multiply labeled specimen must be subjected to an independent background correction
because the level of autofluorescence in each channel can vary substantially. Generally,
autofluorescence is greater for shorter excitation wavelengths, such as those emitted by
ultraviolet, 405-nanometer diode, and 488-nanometer argon-ion lasers. The green, yellow,
orange, and red laser channels are far less prone to suffer autofluorescence artifacts.
The bleed-through controls are necessary to determine the amount of signal gain possible in each
channel without initiating bleed-through into adjacent channels. For example, when examining
dual-labeled specimens fluorescein and rhodamine, specimens containing both fluorescein and
rhodamine alone should be prepared. To establish the level of bleed-through, the fluorescein
control is imaged with an argon-ion 488-nanometer laser under optimum conditions and the
amount of signal present in the rhodamine channel recorded (note there is no rhodamine dye in
the specimen). This signal represents fluorescein bleed-through combined with background
autofluorescence. The procedure is repeated with the rhodamine control, this time exciting with
the helium-neon 543-nanometer line and examining bleed-through into the fluorescein channel.
This approach to bleed-through correction requires that the relationship between fluorescence
emission intensity and fluorophore concentration be constant for all pixels in the image. The
photomultipliers used for confocal microscopy exhibit good linearity when imaging at a single
wavelength, but large changes in the observed signal levels can be obtained with individual
channels due to the highly nonlinear relationship between photomultiplier quantum efficiency
and wavelength of the detected light (especially in the extremes of the response curve). In
addition, the influence of localized environmental variables and photobleaching in the region
being examined must be taken into consideration. Shifts in the emission spectra wavelength
profile of fluorophores, although unlikely with most of the popular probes, will affect the relative
distribution of fluorescence signal acquired between the detectors.
Once the appropriate controls have been prepared and analyzed, it is possible to estimate the
amount of signal bleed-through that is likely to occur when fluorophore pairs in the target
specimen are imaged. As discussed above, even with filter bandwidth optimization, a portion of
the fluorescein emission will bleed into the rhodamine channel and some of the rhodamine
emission will bleed into the fluorescein channel, although rhodamine bleed-through will be much
less than that observed with fluorescein. Several commercial software packages are available to
process bleed-through data using linear unmixing algorithms, and confocal microscope
manufacturers often bundle similar software with their instruments. This software can apply the
simultaneous linear equation correction algorithms for each pixel in the image to determine the
amount of bleed-through based on the ratio of intensities recorded with the specimen and
controls.
Spectral bleed-through artifacts are easily confused with resonance energy transfer, colocalization, and non-specific background staining. If the emission spectrum of one fluorophore
overlaps significantly with the absorption spectrum of a second probe, then regions where the
two fluorophores are co-localized may undergo resonance energy transfer. Bleed-through can be
distinguished from resonance energy transfer by performing control measurements, as described
above, for specimens labeled with the individual fluorophores alone. Resonance energy transfer
can be observed (after bleed-through correction) by exciting the fluorophore with the lowest
absorption maxima, and then detecting signal in the channel of the fluorophore whose emission
spectrum overlaps with the excited probe. As an example using fluorescein and rhodamine, after
exciting fluorescein with the 488-nanometer laser, fluorescence is monitored in the rhodamine
channel. Any signal recorded in the rhodamine channel could be due to resonance energy
transfer, but only in regions where the two probes are co-localized.
Conclusions
In summary, the best fluorophores for confocal or widefield fluorescence microscopy have
absorption maxima that closely match the laser or arc-discharge spectral lines utilized to excite
the probe. Choosing fluorophores with the highest quantum yields for the least abundant targets
will assist in balancing overall fluorescence emission. In addition, probes with narrow emission
spectra may dramatically reduce the problem of bleed-through, but will not eliminate it
altogether. The optical filter sets chosen to examine fluorophore emission should be closely
matched to the spectral profiles of the probe with regards to bandwidth size and location. Also,
interference filter blocking levels often vary by manufacturer and should be checked to ensure
that unwanted fluorescence emission is excluded by the filter set used for imaging.
Many of the newer synthetic fluorophores exhibit dramatically reduced susceptibility to
photobleaching, which allows longer integration periods with CCD cameras or photon collection
times with reduced laser power in confocal microscopy. This results in higher quality images and
far less cell damage during live-cell imaging experiments. Newer fluorescent proteins are more
photostable than previous versions, and far more likely to not disrupt cellular metabolic
processes than synthetic probes. However, it should be noted that all fluorophores have the
potential to affect cell behavior to some extent, especially if the chemical and physical
characteristics of the probe are significantly altered by environmental variables, such as ion
concentration, pH, and hydrophobicity.
Regardless of the fluorophore physical characteristics, a high degree of target specificity is
necessary to obtain suitable signal levels and reduce bleed-through artifacts, especially with
secondary antibodies in immunofluorescence preparations. Even a small amount of non-specific
labeling will produce a high degree of background that both degrades the specimen image and
confuses quantitative interpretation. Many of the probes designed to highlight subcellular
structures (such as the Golgi complex or mitochondria) have a relatively low level of specificity,
but are still useful because of the known geometry of these organelles.
As manufacturers develop more advanced fluorophores and cheaper diode-based laser systems
having spectral lines throughout the visible and near-infrared regions, the choice of probes for
multiple labeling experiments in confocal and widefield fluorescence microscopy without bleedthrough artifacts will become easier. For example, confocal microscopes equipped with a 405nanometer diode laser, a 543-nanometer helium-neon laser, and a 650-nanometer red diode laser
can excite fluorophores with emission spectra having wavelength maxima separated by over 100
nanometers, thus minimizing the potential for bleed-through. In addition, new fluorescent
proteins with emission spectral profiles in the far red and infrared will require laser spectral lines
that are less damaging to cellular metabolic processes than the current violet, blue, and green
systems.
Contributing Authors
Douglas B. Murphy - Department of Cell Biology and Anatomy and Microscope Facility, Johns Hopkins University School of Medicine, 725
N. Wolfe Street, 107 WBSB, Baltimore, Maryland 21205.
David W. Piston - Department of Molecular Physiology and Biophysics, Vanderbilt University, Nashville, Tennessee, 37232.
Stuart H. Shand and Simon C. Watkins - Center for Biologic Imaging, University of Pittsburgh, S233 Biomedical Science Towers, 3500
Terrace Street, Pittsburgh, Pennsylvania, 15261.
Michael W. Davidson - National High Magnetic Field Laboratory, 1800 East Paul Dirac Dr., The Florida State University, Tallahassee,
Florida, 32310.
Argon-Ion Lasers
As a distinguished member of the common and well-explored family of ion lasers, the argon-ion
laser operates in the visible and ultraviolet spectral regions by utilizing an ionized species of the
noble gas argon. Argon-ion lasers function in continuous wave mode when plasma electrons
within the gaseous discharge collide with the excited laser species to produce light.
The tutorial initializes with the gas laser tutorial speed set to Medium, a level that enables the
visitor to observe the slow build-up of light in the laser cavity as it is reflected back and forth
through the Brewster windows and mirrors. In order to operate the tutorial, translate the Laser
Wavelength slider between the various available laser spectral lines (351, 364, 457, 488, and
514 nanometers), and observe how the color of the output beam changes with wavelength. Use
the Tutorial Speed slider to adjust the speed of light oscillations within the laser cavity and the
level of light emitted through the output lens.
The argon-ion laser is capable of producing approximately 10 wavelengths in the ultraviolet
region and up to 25 in the visible region, ranging from 275 to 363.8 nanometers and 408.9 to
686.1 nanometers, respectively. In the visible light spectral region, the lasers can produce up to
100 watts of continuous-wave power with the output concentrated into several strong lines
(primarily the 488 and 514.5 nanometer transitions). The gain bandwidth on each transition is on
the order of 2.5 gigahertz. Argon-ion gas laser discharge tubes have a useful life span ranging
between 2000 and 5000 hours, and operate at gas pressures of approximately 0.1 torr.
An unfortunate side effect of the high discharge currents and low gas pressure employed by
argon-ion lasers is an extremely high plasma electron temperature, which generates a significant
amount of heat. In most cases, high power (2 to 100 watts) argon-ion laser systems are watercooled through an external chiller, but lower power (5-150 milliwatts) models can be cooled with
forced air through an efficient fan. Argon-ion lasers utilized in confocal and other fluorescence
microscopy techniques are generally of the lower power variety, which produce between 10 and
100 milliwatts of power in TEM(00) mode at 488.0 nanometers. The laser cavity for these
smaller systems is approximately 35 to 50 centimeters in length and about 15 centimeters in
diameter, and can be housed in a small cabinet with an integral fan to supply fresh, cool air.
Fluorescent Probe Excitation Efficiency
The absorption and fluorescence emission spectral profiles of a fluorophore are two of the most
important criteria that must be scrutinized when selecting probes for applications in laser
scanning confocal microscopy. In addition to the wavelength range of the absorption and
emission bands, the molar extinction coefficient for absorption and the quantum yield for
fluorescence emission should be considered. At laser excitation levels that do not saturate the
fluorophore, fluorescence intensity is directly proportional to the product of the extinction
coefficient and the quantum yield. This interactive tutorial examines how this relationship can be
utilized to match fluorophores with specific lasers for confocal microscopy.
The tutorial initializes with a spectral wavelength versus relative intensity plot appearing in the
window, and the absorption and fluorescence emission spectral profiles for a randomly selected
fluorophore displayed on the graph. Superimposed over the fluorophore excitation spectrum is
the closest common confocal laser spectral line to the center of the dye absorption maximum. At
this wavelength, fluorescence emission is theoretically nearest the maximum value possible with
the laser, and the fluorophore emission curve in the tutorial contains a fill indicative of the color
spectrum at the intensity level permitted by the product of the fluorophore extinction coefficient
and the emission intensity. The Emission Intensity box in the lower left-hand corner of the
window presents the color most closely associated with fluorescence emission at the maximum
wavelength.
In order to operate the tutorial, use the Wavelength slider to adjust the laser selection over a
region dictated by the absorption spectral profile of the fluorophore. The slider is limited in a
range by the ends of the spectra, and can only travel to the closest laser line past these
boundaries. As the virtual laser line is translated along the wavelength axis, the tutorial calculates
the product of the fluorophore molar extinction coefficient and fluorescence quantum yield and
continuously updates the resulting value as a percentage beneath the Emission Intensity box to
provide a qualitative measure of the relative fluorescence emission that can be expected at the
corresponding excitation wavelength. Likewise, the intensity of the color displayed in the
Emission Intensity box changes to reflect the relative intensity of fluorescence emission at the
chosen laser source excitation wavelength. The laser source associated with each wavelength is
displayed in a yellow box above the Wavelength slider, and the specific spectral line from that
laser is displayed above the slider bar. Fluorophore classes can be selected using the radio
buttons on the right-hand side of the window, and individual fluorophores can be loaded into the
display graph using the Choose A Dye pull-down menu.
The number of fluorescent probes currently available for confocal microscopy runs in the
hundreds, with many dyes having absorption maxima closely associated with common laser
spectral lines. An exact match between a particular laser line and the absorption maximum of a
specific probe is not always possible, but the excitation efficiency of lines near the maximum is
usually sufficient to produce a level of fluorescence emission that can be readily detected. For
example, in Figure 1 the absorption spectra of two common probes are illustrated, along with the
most efficient laser excitation lines. The green spectrum is the absorption profile of fluorescein
isothiocyanate (FITC), which has an absorption maximum of 495 nanometers. Excitation of the
FITC fluorophore at 488 nanometers using an argon-ion laser produces an emission efficiency of
approximately 87 percent. In contrast, when the 477-nanometer or the 514-nanometer argon-ion
laser lines are used to excite FITC, the emission efficiency drops to only 58 or 28 percent,
respectively. Clearly, the 488-nanometer argon-ion (or krypton-argon) laser line is the most
efficient source for excitation of this fluorophore.
The red spectrum in Figure 1 is the absorption profile of Alexa Fluor 546, a bi-sulfonated
alicyclic xanthene (rhodamine) derivative with a maximum extinction coefficient at 556
nanometers, which is designed specifically to display increased quantum efficiency at
significantly reduced levels of photobleaching in fluorescence experiments. The most efficient
laser excitation spectral line for Alexa Fluor 546 is the yellow 568-nanometer line from the
krypton-argon mixed gas ion laser, which produces an emission efficiency of approximately 84
percent. The next closest laser spectral lines, the 543-nanometer line from the green helium-neon
laser and the 594-nanometer lines from the yellow helium-neon laser, excite Alexa Fluor 546
with an efficiency of 43 and 4 percent, respectively. Note that the 488-nanometer argon-ion laser
spectral line excites Alexa Fluor 546 with approximately 7-percent efficiency, a factor that can
be of concern when conducting dual labeling experiments with FITC and Alexa Fluor 546
simultaneously.
Colocalization of Fluorophores in Confocal Microscopy
Two or more fluorescence emission signals can often overlap in digital images recorded by
confocal microscopy due to their close proximity within the specimen. This effect is known as
colocalization and usually occurs when fluorescently labeled molecules bind to targets that lie in
very close or identical spatial positions. This interactive tutorial explores the quantitative
analysis of colocalization in a wide spectrum of specimens that were specifically designed either
to demonstrate the phenomenon, or to alternatively provide examples of fluorophore targets that
lack any significant degree of colocalization.
The tutorial initializes with a randomly chosen confocal microscopy dual or triple labeled
fluorescence image appearing in the Specimen Image window and the accompanying red-green
(or red-blue) scatterplot graphed two-dimensionally in the adjacent Colocalization Scatterplot
coordinate system. Plots of the available channel permutations (Red-Green, Red-Blue, and
Green-Blue) can be displayed using the Colocalization Channels set of radio buttons. In
addition, each channel in the Specimen Image window can be toggled on or off using the check
boxes in the Channels menu. A three-dimensional rendering of the colocalization scatterplot
(number of pixels plotted on the z axis) can be obtained by activating the 3D radio button. This
view can be rotated within the window using the mouse cursor. Colocalization coefficients
automatically displayed beneath the scatterplot graph include Pearson's, Overlap, and Global
(k1 and k2), as described below.
In order to operate the tutorial, use the mouse cursor to draw a region of interest in the
Colocalization Scatterplot graph. The default area selection tool generates rectangular regions,
but elliptical and freehand areas can be chosen with the appropriate Region of Interest radio
buttons. Once a region has been selected, an overlay of the colocalized pixels is displayed in the
Specimen Image window and the Global colocalization coefficient display changes into the
Local (M1 and M2) value calculated within the region of interest. The image Colocalization
Overlay view can be toggled between Full Color and Binary views using check boxes. At any
point, a new specimen can be selected using the Choose A Specimen pull-down menu. Details
of the specimen fluorophore staining protocol and the potential for colocalization are described
in the yellow text box at the bottom of the tutorial window.
A quantitative assessment of fluorophore co-localization in confocal optical sections can be
obtained using the information obtained from scatterplots and selected regions of interest.
Several values are generated using information from the entire scatterplot, while others are
derived from pixel values contained within a selected region of interest. Among the variables
used to analyze the entire scatterplot is Pearson's correlation coefficient (R(r)), which is one of
the standard techniques applied in pattern recognition for matching one image to another in order
to describe the degree of overlap between the two patterns. Pearson's correlation coefficient is
calculated according to the equation:
(1)
where S1 is the signal intensity of pixels in the first channel and S2 is the signal intensity of
pixels in the second channel. The values S1(average) and S2(average) are the average values of
pixels in the first and second channel, respectively. In Pearson's correlation, the average pixel
intensity values are subtracted from the original intensity values. As a result, the value of this
coefficient ranges from -1 to 1, with a value of -1 representing a total lack of overlap between
pixels from the images, and a value of 1 indicating perfect image registration. Pearson's
correlation coefficient accounts only for the similarity of shapes between the two images, and
does not depend upon image pixel intensity values. When applying this coefficient to colocalization analysis, however, the potentially negative values are difficult to interpret, requiring
another approach to clarify analysis results.
A simpler technique often employed to calculate an alternative correlation coefficient involves
eliminating the subtraction of average pixel intensity values from the original intensities. Defined
formally as the Overlap coefficient (R), this value ranges between 0 and 1 and is not sensitive to
intensity variations in the image analysis. The Overlap coefficient is defined as:
(2)
The product of channel intensities in the numerator returns a significant value only when both
values belong to a pixel involved in co-localization (if both intensities are greater than zero). As
a result, the numerator in equation (2) is proportional to the number of co-localizing pixels. In a
similar manner, the denominator of the Overlap equation is proportional to the number of pixels
from both components in the image, regardless of whether co-localization is present (Note: the
components are defined as the red and green images or the pixel arrays from channel 1 and
channel 2, respectively). A major advantage of the Overlap coefficient is its relative insensitivity
to differences in signal intensities between various components of an image, which are often
produced by fluorochrome concentration fluctuations, photobleaching, quantum efficiency
variations, and non-equivalent electronic channel settings.
The most important disadvantage of using the Overlap coefficient is the strong influence of the
ratio between the number of image features in each channel. To alleviate this dependency, the
Overlap coefficient is divided into two different sub-coefficients, termed k(1) and k(2) in order
to express the degree of co-localization as two separate parameters:
(3)
The overlap coefficients, k(1) and k(2), describe the differences in intensities between the
channels, with k(1) being sensitive to the differences in the intensity of channel 2 (green signal),
while k(2) depends linearly on the intensity of the pixels from channel 1 (red signal). The
equations described thus far are able to generate information about the degree of overlap and can
account for intensity variations between the color channels. In order to estimate the contribution
of one color channel in the co-localized areas of the image to the overall amount of co-localized
fluorescence, an additional set of co-localization coefficients, m(1) and m(2), are defined:
(4)
The co-localization coefficient m(1) is employed to describe the contribution from channel 1 to
the co-localized area, while the coefficient m(2) is used to describe the same contribution from
channel 2. Note that the variable S1(i,coloc) is equal to S1(i) if S2(i) is greater than zero and vice
versa for the variable S2(i,coloc). These coefficients are proportional to the amount of
fluorescence of the co-localizing fluorophores in each channel of the composite image, relative
to the total fluorescence in that channel. Co-localization coefficients m(1) and m(2) can be
determined even when the signal intensities in the two image channels have significantly
different levels.
A second pair of co-localization coefficients can be calculated for pixel intensity ranges defined
by an area of interest delineated on the scatterplot. The coefficient M(1) is utilized to describe
the contribution of the channel 1 fluorophore to the co-localized area, while M(2) is used to
describe the contribution of the channel 2 fluorophore. These co-localization coefficients are
defined as:
(5)
where S1(i,coloc) equals S1(i) if S2(i) lies within the region of interest thresholds (left and right
sides of a rectangular ROI) and equals zero if S2(i) represents a pixel outside the threshold
levels. Similarly, S2(i,coloc) equals S2(i) if S1(i) lies within the region of interest thresholds (top
and bottom sides of a rectangular ROI) and equals zero if S1(i) is outside the region of interest.
In other words, for each channel, the numerator represents the sum of all pixel intensities in that
channel that also have a component from the other channel, whereas the denominator represents
the sum of all intensities from the channel. These coefficients are proportional to the amount of
fluorescence of co-localizing objects in each channel of the composite image, relative to the total
fluorescence in that channel.
A majority of the co-localization software analysis programs available commercially are able to
calculate the parameters described above, including Pearson's correlation coefficient, the total
overlap coefficient, as well as the individual k(x), m(x), and M(x) co-localization coefficients. In
addition, many programs contain algorithms to apply background subtraction corrections,
generate scatterplots of the entire image, and/or perform the calculations using selected regions
of interest on single dual channel composite images or optical stacks along the axial plane. The
most important data output from these software packages is the co-localization coefficient, which
indicates the relative degree of overlap between signals. For example, a co-localization
coefficient value of 0.75 for the fluorophore in channel 1 indicates that the ratio for all channel 1
intensities that have a channel 2 component, divided by the sum of all channel 1 intensities, is 75
percent. This is a relatively high degree of co-localization. Likewise, a value of 0.25 for the
channel 2 fluorophore indicates a significantly diminished level of co-localization (equal to onethird of the channel 1 fluorophore).
Contributing Authors
Will Casavan and Yuri Gaidoukevitch - Media Cybernetics, 8484 Georgia Avenue, Suite 200, Silver Spring, Maryland, 20910.
Matthew J. Parry-Hill, Nathan S. Claxton, and Michael W. Davidson - National High Magnetic Field Laboratory, 1800 East Paul Dirac Dr.,
The Florida State University, Tallahassee, Florida, 32310.
Theory of Confocal Microscopy
Laser
scanning confocal microscopy represents one of
the most significant advances in optical microscopy
ever developed, primarily because the technique
enables visualization deep within both living and fixed
cells and tissues and affords the ability to collect
sharply defined optical sections from which threedimensional renderings can be created. The principles
and techniques of confocal microscopy are becoming
increasingly available to individual researchers as new
single-laboratory microscopes are introduced.
Development of modern confocal microscopes has
been accelerated by new advances in computer and
storage technology, laser systems, detectors,
interference filters, and fluorophores for highly
specific targets.
Introduction to Confocal Microscopy - Confocal microscopy offers several advantages over
conventional widefield optical microscopy, including the ability to control depth of field,
elimination or reduction of background information away from the focal plane (that leads to
image degradation), and the capability to collect serial optical sections from thick specimens.
The basic key to the confocal approach is the use of spatial filtering techniques to eliminate outof-focus light or glare in specimens whose thickness exceeds the immediate plane of focus.
There has been a tremendous explosion in the popularity of confocal microscopy in recent years,
due in part to the relative ease with which extremely high-quality images can be obtained from
specimens prepared for conventional fluorescence microscopy, and the growing number of
applications in cell biology that rely on imaging both fixed and living cells and tissues. In fact,
confocal technology is proving to be one of the most important advances ever achieved in optical
microscopy.
Fluorescence Excitation and Emission Fundamentals - Fluorescence is a member of the
ubiquitous luminescence family of processes in which susceptible molecules emit light from
electronically excited states created by either a physical (for example, absorption of light),
mechanical (friction), or chemical mechanism. Generation of luminescence through excitation of
a molecule by ultraviolet or visible light photons is a phenomenon termed photoluminescence,
which is formally divided into two categories, fluorescence and phosphorescence, depending
upon the electronic configuration of the excited state and the emission pathway. Fluorescence is
the property of some atoms and molecules to absorb light at a particular wavelength and to
subsequently emit light of longer wavelength after a brief interval, termed the fluorescence
lifetime. The process of phosphorescence occurs in a manner similar to fluorescence, but with a
much longer excited state lifetime.
Fluorophores for Confocal Microscopy - Biological laser scanning confocal microscopy relies
heavily on fluorescence as an imaging mode, primarily due to the high degree of sensitivity
afforded by the technique coupled with the ability to specifically target structural components
and dynamic processes in chemically fixed as well as living cells and tissues. Many fluorescent
probes are constructed around synthetic aromatic organic chemicals designed to bind with a
biological macromolecule (for example, a protein or nucleic acid) or to localize within a specific
structural region, such as the cytoskeleton, mitochondria, Golgi apparatus, endoplasmic
reticulum, and nucleus. Other probes are employed to monitor dynamic processes and localized
environmental variables, including concentrations of inorganic metallic ions, pH, reactive
oxygen species, and membrane potential. Fluorescent dyes are also useful in monitoring cellular
integrity (live versus dead and apoptosis), endocytosis, exocytosis, membrane fluidity, protein
trafficking, signal transduction, and enzymatic activity. In addition, fluorescent probes have been
widely applied to genetic mapping and chromosome analysis in the field of molecular genetics.
Interference Filters for Fluorescence Microscopy - The performance of high-resolution
fluorescence microscopy imaging systems and related quantitative applications, especially as
applied in living cell and tissue studies, requires precise optimization of fluorescence excitation
and detection strategies. Fluorescence microscopy techniques could not have advanced so
dramatically in recent years without significant developments in every dimension of the current
state of the art, including the optical microscopes, the biology and chemistry of fluorophores, and
perhaps most important, filter technology. The utilization of highly specialized and advanced
thin film interference filters has enhanced the versatility and scope of fluorescence techniques,
far beyond the capabilities afforded by the earlier use of gelatin and glass filters relying on the
absorption properties of embedded dyes.
Spectral Bleed-Through Artifacts in Confocal Microscopy - The spectral bleed-through of
fluorescence emission (often termed crossover or crosstalk), which occurs due to the very broad
bandwidths and asymmetrical spectral profiles exhibited by many of the common fluorophores,
is a fundamental problem that must be addressed in both widefield and laser scanning confocal
fluorescence microscopy. The phenomenon is usually manifested by the emission of one
fluorophore being detected in the photomultiplier channel or through the filter combination
reserved for a second fluorophore. Bleed-through artifacts often complicate the interpretation of
experimental results, particularly if subcellular co-localization of fluorophores is under
investigation or quantitative measurements are necessary, such as in resonance energy transfer
(FRET) and photobleaching (FRAP) studies.
Resolution and Contrast in Confocal Microscopy - All optical microscopes, including
conventional widefield, confocal, and two-photon instruments are limited by fundamental
physical factors in the resolution that they can achieve. In a perfect optical system, resolution is
limited by numerical aperture of the optical components and by the wavelength of the light, both
incident and detected. The concept of resolution is inseparable from contrast, and is defined as
the minimum separation between two points that results in a certain contrast between them. In a
real fluorescence microscope, contrast is determined by the number of photons collected from
the specimen, the dynamic range of the signal, optical aberrations of the imaging system, and the
number of picture elements (pixels) per unit area.
Introduction to Lasers - Ordinary natural and artificial light is released by energy changes on
the atomic and molecular level that occur without any outside intervention. A second type of
light exists, however, and occurs when an atom or molecule retains its excess energy until
stimulated to emit the energy in the form of light. Lasers are designed to produce and amplify
this stimulated form of light into intense and focused beams. The word laser was coined as an
acronym for Light Amplification by the Stimulated Emission of Radiation. The special nature of
laser light has made laser technology a vital tool in nearly every aspect of everyday life including
communications, entertainment, manufacturing, and medicine.
Laser Systems for Confocal Microscopy - The lasers commonly employed in laser scanning
confocal microscopy are high-intensity monochromatic light sources, which are useful as tools
for a variety of techniques including optical trapping, lifetime imaging studies, photobleaching
recovery, and total internal reflection fluorescence. In addition, lasers are also the most common
light source for scanning confocal fluorescence microscopy, and have been utilized, although
less frequently, in conventional widefield fluorescence investigations.
Acousto-Optic Tunable Filters (AOTFs) - Several benefits of the AOTF combine to greatly
enhance the versatility of the latest generation of confocal instruments, and these devices are
becoming increasing popular for control of excitation wavelength ranges and intensity. The
primary characteristic that facilitates nearly every advantage of the AOTF is its capability to
allow the microscopist control of the intensity and/or illumination wavelength on a pixel-bypixel basis while maintaining a high scan rate. This single feature translates into a wide variety
of useful analytical microscopy tools, which are even further enhanced in flexibility when laser
illumination is employed.
Non-Coherent Light Sources for Confocal Microscopy - The traditional illumination system
in the modern widefield microscope utilizes a tungsten-halogen source for transmitted light and a
short-arc lamp for fluorescence excitation. Various lasers have been utilized as a light source for
widefield observation by a few investigators, but the advent of the confocal microscope vastly
increased laser use in microscopy. This discussion reviews the merits and limitations of noncoherent (or non-laser) light sources in confocal microscopy, both as light sources for confocal
illumination and as secondary sources for widefield microscopy in confocal microscopes. Two
initial issues frequently arise when illumination systems for confocal microscopes are
considered, and these have a direct bearing on the choice of light sources for a particular
instrument.
Confocal Microscope Objectives - For any conventional optical microscope configuration, the
objective is the most critical component of the system in determining the information content of
the image. The contrast and resolution of fine specimen detail, the depth within the specimen
from which information can be obtained, and the lateral extent of the image field are all
determined by the design of the objective and its performance under the specific conditions
employed for the observation. Additional demands are imposed on the objective in scanning
confocal techniques, in which this crucial imaging component also serves as the illumination
condenser and is often required to perform with high precision at a wide range of wavelengths
and at very low light levels without introducing unacceptable image-degrading noise.
Confocal Microscope Scanning Systems - Confocal imaging relies upon the sequential
collection of light from spatially filtered individual specimen points, followed by electronic
signal processing and ultimately, the visual display as corresponding image points. The point-by-
point signal collection process requires a mechanism for scanning the focused illuminating beam
through the specimen volume under observation. Three principal scanning variations are
commonly employed to produce confocal microscope images. Fundamentally equivalent
confocal operation can be achieved by employing a laterally translating specimen stage coupled
to a stationary illuminating light beam (stage scanning), a scanned light beam with a stationary
stage (beam scanning), or by maintaining both the stage and light source stationary while
scanning the specimen with an array of light points transmitted through apertures in a spinning
Nipkow disk. Each technique has performance features that make it advantageous for specific
confocal applications, but that limit the usefulness in others.
Signal-to-Noise Considerations - In any quantitative assessment of imaging capabilities
utilizing digital microscopy techniques, including confocal methods, the effect of signal
sampling on contrast and resolution must be considered. The measured signal level values do not
directly represent the number of photons emitted or scattered by the specimen, but are
proportional to that number. Furthermore, each individual sample of signal intensity is only an
approximation of the number of collected photons, and will vary with repeated measurement.
The variation, referred to as noise, imparts an uncertainty in the quantification of intensity, and
therefore in the contrast and resolution of the image data.
Electronic Light Detectors: Photomultipliers - In modern widefield fluorescence and laser
scanning confocal optical microscopy, the collection and measurement of secondary emission
gathered by the objective can be accomplished by several classes of photosensitive detectors,
including photomultipliers, photodiodes, and solid-state charge-coupled devices (CCDs). In
confocal microscopy, fluorescence emission is directed through a pinhole aperture positioned
near the image plane to exclude light from fluorescent structures located away from the objective
focal plane, thus reducing the amount of light available for image formation. As a result, the
exceedingly low light levels most often encountered in confocal microscopy necessitate the use
of highly sensitive photon detectors that do not require spatial discrimination, but instead
respond very quickly with a high level of sensitivity to a continuous flux of varying light
intensity.
Electronic Imaging Detectors - Over the past several years, the rapidly growing field of
fluorescence microscopy has evolved from a dependence on traditional photomicrography using
emulsion-based film to one in which electronic images are the output of choice. The imaging
device is one of the most critical components in fluorescence microscopy because it determines
at what level specimen fluorescence may be detected, the relevant structures resolved, and/or the
dynamics of a process visualized and recorded.
Fluorochrome Data Table - As a guide to fluorophores for confocal and widefield fluorescence
microscopy, the table presented in this section lists many commonly-used fluorochromes, with
their respective peak absorption and emission wavelengths and suggested laser illumination
sources. While the authors assume responsibility for the accurate reporting of the data as
published in various reliable sources, several caveats must be given. Fluorochrome dyes are
environmentally sensitive and different results will be obtained with different solvents and
applications. In reviewing the literature, one will frequently find somewhat different data
supplied for the identical fluorochrome. There are also, in many instances, several sub-varieties
of a fluorochrome.
Glossary of Terms in Confocal Microscopy - The complex nomenclature of fluorescence
microscopy is often confusing to both beginning students and seasoned research microscopists
alike. This resource is provided as a guide and reference tool for visitors who are exploring the
large spectrum of specialized topics in fluorescence and laser scanning confocal microscopy.
Recommended Books on Confocal Microscopy - A surprisingly limited number of books
dealing with various aspects of laser scanning and spinning disk confocal microscopy and related
techniques are currently available from the booksellers. This section lists the FluoView Resource
Center website development team's top 12 recommended books. Although the volumes listed in
this section deal pricipally with confocal microscopy and related methodology, there exist a
number of additional books that contain focused treatments of the materials described below, and
these should also be consulted for specific techniques and timely review articles.
Basic Concepts in Laser Scanning Confocal Microscopy (PDF; 2.8 Mb) - Laser scanning
confocal microscopy has become an invaluable tool for a wide range of investigations in the
biological and medical sciences for imaging thin optical sections in living and fixed specimens
ranging in thickness up to 100 micrometers. Modern instruments are equipped with 3-5 laser
systems controlled by high-speed acousto-optic tunable filters (AOTFs), which allow very
precise regulation of wavelength and excitation intensity. Coupled with photomultipliers that
have high quantum efficiency in the near-ultraviolet, visible and near-infrared spectral regions,
these microscopes are capable of examining fluorescence emission ranging from 400 to 750
nanometers. Download this review article to learn more.
Interactive Java Tutorials - Explanations for many of the exceedingly complex concepts in
laser scanning confocal microscopy can significantly benefit from the assistance of interactive
tutorials that enable the student to obtain instanteous (real-time) response to changes in variables.
The tutorials in section address the basic aspects of confocal microscopy instrumentation, laser
systems, detectors, image processing, resolution, contrast, and many other aspects of the
technique. All interactive Java tutorials require the Java Virtual Machine, which is available
without cost as a browser plug-in from Sun Microsystems.