Confocal Microscopy of Drosophila Embryos
The genetically tractable organism Drosophila melanogaster
is proving to be an excellent model system
for cell biological analysis in the context of the whole
organism. The relative ease with which embryos can
be obtained in large numbers and processed for highresolution
light microscopy has facilitated many recent
advances at the interface between cell and developmental
biology. Fine subcellular structures previously
impossible to visualise by conventional fluorescence
microscopy, on account of high noise resulting from
out-of-focus signals, are revealed with clarity on a confocal
There are several reasons why scientists who have
not used Drosophila
before may wish to use Drosophila
embryos for the analysis of protein localisation and
expression. The embryo contains representatives of
each cell type and is small enough (500 x 100 µm) to fit
within the field of view of a 20X objective lens. The
embryos are nearly transparent, permitting visualisation
of all cells in whole mount preparations. These
features allow one to assay the tissue distribution of a
particular protein in a single specimen. The tissues
have a relatively simple structure, with the epithelia
being made up of a single layer of cells. In general,
there are fewer copies of each protein encoded by the
genome compared with vertebrates, e.g., one α-actinin
rather than four, further simplifying the analysis of the
distribution of a particular kind of protein. Injection of
double-stranded RNA can be an effective way to knock
down protein expression, provided the bulk of the
protein in the embryo comes from new synthesis.
Sophisticated manipulation of the proteins is possible using the powerful molecular genetic techniques available
in this organism.
has many advantages for cell biological
analysis, it also has some drawbacks. The cells
are small: an embryonic epidermal cell, for example,
has dimensions of only 2 x 5 µm compared to a vertebrate
epidermal cell of 10 x 20µm, which can make it
difficult to resolve different intracellular compartments.
The embryo is the only stage where the whole
animal can be stained in its entirety; at late stages of
development, antibody penetration is blocked by
the secreted exoskeleton [although the use of proteins
tagged with green fluorescent protein (GFP) circumvents
this problem]. Therefore, the most easily
generated samples for analysis are the embryo or
tissues, that are easily dissected from the larva or
adult, such as the imaginal discs and ovaries. Only a
small number of Drosophila
cell lines are available for in vitro
culture and experimentation. These represent
just a few cell types and are also small. Finally,
antibodies raised against vertebrate proteins rarely
bind to Drosophila
orthologues, even when they are
highly conserved. Therefore, new antibodies need to
be raised to see the distribution of a given protein in Drosophila
. An important exception to this are antibodies
raised to specific motifs of proteins, such as
those recognising particular phosphorylated residues.
A growing number of antibodies against Drosophila
proteins are available, either from the Developmental
Studies Hybridoma Bank or directly from the laboratory
that has generated them; few are available
This article describes the techniques used to look
at cell junctions and cytoskeletal structures in the
developing embryo, using both antibodies against Drosophila
proteins and GFP-tagged proteins
expressed from transgenes.
The embryo is surrounded by two proteinaceous
layers: the egg shell or chorion, which is removed
easily; and the vitelline membrane, which is impermeable
and more difficult to remove. The vitelline
membrane can be permeabilised by treatment with
heptane, permitting fixatives to reach the embryo.
The vitelline membrane is generally removed by
"popping" the embryos with a rapid osmotic shock
produced by adding methanol to create a
heptane-methanol interface and shaking vigorously.
The fixation/devitellinisation steps are the part of the
procedure most likely to go wrong, but with practice
become routine. Once the embryos are fixed and the
vitelline membrane removed, antibody staining is
straightforward. Simple mounting of embryos is sufficient
in most cases, which, due to the elongated shape
of the embryo, allows many optical section views
except a cross section of the embryo (i.e., cutting a slice
from the middle of loaf of bread). As this can be very
informative, we also describe a more labour-intensive
method that allows such a view. We also make some
recommendations for getting the best confocal images
embryos. Additional references describing
these methods are found in Davis (1999),
Hazelrigg (2000), and Rothwell (2000).
II. INSTRUMENTATION AND
Fly bottles or vials (250-ml bottles and 30/40ml vials,
Scientific Lab Supplies)
Egg collection cages (see Fig. 1, I)
Apple juice agar plates (see solutions)
Embryo collection baskets (see Fig. 1, I)
Scintillation vials (4ml, 908-054, Jencons; 20ml, 215/0079/00, BDH)
Rollers (Denley Spiramix 5 or equivalent)
Rotary shakers (Labinco BV or equivalent)
Microscope glass slides 76 x 26mm (Menzel Glaeser,
Coverslips 22 x 22mm, 22 x 40mm (Menzel Glaeser)
Forceps (Watkins and Doncaster)
A001 size dissecting tungsten needles (E6871, Watkins
Needle holder (Watkins and Doncaster)
, 0.45 x 10 syringe needle (Industrial and
Artist's paintbrushes 00, 000
Heptane (27051-2, Sigma)
Methanol (M/4000/PC17, Fischer Chemicals)
Ethanol (E/0555DF/17, Fischer Chemicals)
Formaldehyde (37-40%, 101134-A, AnalaR/BDH,
Paraformaldehyde (40% EM grade, 15715, Electronmicroscopy
Triton X-100 (T-9284, Sigma)
Bovine serum albumin (BSA) fraction IV (A-2153,
NaCl (102415K, AnalaR/BDH)
KCl (101984L, AnalaR/BDH)
(63064 Fluka Biochemika)
EGTA (E-4378, Sigma)
PIPES (P-8203, Sigma)
SDS (Ultrapure, Melford Labs Ltd.)
Glycerol (101184K, AnalaR/BDH)
Sucrose (S-0389, Sigma)
Apple juice (commercial)
Bactoagar (0140-01, Difco laboratories)
Nipagin (Nipa Laboratories Inc.)
Baker's yeast blocks (Commercial)
Vectashield (H-1000,Vector Labs, Burlingame)
Voltalef H10S Halocarbon oil (Elf Atochem, France)
Fly food (Instant Drosophila
medium, Phillip Harris
A. Producing Embryos for
- 10× phosphate-buffered saline (PBS) (1 litre) (adapted
from Sambrook and Russell, 2001): 80g (137mM final)
NaCl, 2g (2.7mM final) KCl, 14.4g (10mM final)
Na2HPO4, 2.4g (2mM final) KH2PO4, and distilled
water up to 1 litre. Dissolve all components in 800ml
of H2O and make volume up to 1 litre. Adjust pH to
7.4 with HCl/NaOH. Sterilise by autoclaving.
- PBT (500ml) PBS with 0.3% Triton X-100: 50ml
10x PBS, 750 µl Triton X-100 (0.3% v/v), 0.1 g (optional)
sodium azide, and distilled water to 500ml
- PBTB (500ml) PBS with 0.3% Triton X-100 and
0.5% BSA: 50ml 10x PBS, 2.5g BSA (0.5%w/v), 750µl
Triton X-100 (0.3% v/v), 0.1 g (optional) sodium azide,
and distilled water to 500ml.
- PB: 77.4ml (100mM final) Na2HPO4 (1M) and
22.6ml (100mM final) NaH2PO4 (1M). Dissolve in distilled
water to 1000ml. Adjust pH to 7.4.
- PEM (4 ml) (adapted from Ashburner, 1989): 400µl
(100mM final) 1M PIPES (pH 6.9), 4µl (1mM final) 1M MgCl2, and 20µl (1mM final) 200mM EGTA (pH
8.0). Adjust to pH 6.9 with 10M KOH to final volume
- 4% formaldehyde in PBS (100ml): 10ml 10x PBS,
10.8 ml 37% formaldehyde, and 79.2ml distilled water.
- Apple juice agar plates (1.6 litre): 36g Bactoagar
and 1200ml distilled water. Dissolve the agar in a
microwave oven or on a heating pad until the solution
is clear. Dissolve 40 g sucrose in 400 ml apple juice in a
microwave oven or on a heating pad until the solution
is clear. Mix the two solutions once clear on a stirrer.
Add 20ml of 20% Nipagin when the solution has
cooled below 60°C. Stir again and pour while still
liquid into 5-cm/9-cm petri dishes. When set, store at
4°C. Allow to reach room temperature before use.
B. Removal of Chorion (Dechorionation)
- Grow up flies. If you are not a fly laboratory, try
to obtain a few bottles from a fly colleague. Alternatively,
make simple bottles with instant fly food,
topped with cotton or a foam plug. Try to get the
density of flies such that they are neither under- nor
overcrowded. For 250-ml bottles, 20 females and 10
males will lay the ideal number of eggs per bottle in a
24-h period, after an initial lag of a couple of days
while they feed and mate. Such a bottle will produce
about 500 flies after 10 days at 25°C, depending on the
food and the fly strain. Ideally, collect adult flies within
1-2 days after eclosion from the pupal case. For the egg
collection cages described later, 25-200 females and
about half the number of males is a good number.
- Set up an egg collection cage containing an apple
juice agar plate at the bottom, fixed on with tape, and
a ventilation screen at the top (Fig. 1, IA). The plate
should have an approximately 50 µl dab of yeast paste
in the centre. Make yeast paste by stirring in a bit of
water into the baker's yeast to make it the consistency
of toothpaste. This is the food for the flies and is essential
for the females to lay large numbers of eggs.
Ensure that there is no excess moisture in the cage or
plate (usually resulting from condensation on the
plate). Put the flies into the cage and incubate at the
desired temperature (usually 25°C). It will take a
couple of days before the females lay substantial
numbers of eggs, but once they get going each female
can lay 100 eggs a day. Replace the yeasted apple juice
plates at least once a day. To do this, invert the cage and peel back the tape holding the plate. Gently tap
the flies down on the bench and quickly swap the
plate. Even for the most practiced of us, this usually
results in a few flies gaining freedom.
- Collect embryos for analysis. Place a new yeasted
apple juice plate on the cage. After the appropriate time
period (e.g., 4 hs), remove the plate from the cage, replacing
with a fresh one, and age the plate containing the
embryos at 25°C for the desired length of time. Such short
collections are an advantage when one is interested
in a particular developmental event (for staging, see
Wieschaus and Nuesslein-Volhard, 1998; Campos-
Ortega and Hartenstein, 1997), and for the novice who
has not yet learned how to stage embryos by visual
examination. Embryogenesis lasts 22h at 25°C, but the
exoskeleton becomes secreted at about 16h, making
embryos older than this inaccessible to antibody staining
using standard methods. Development takes twice as
long at 18°C, allowing some adjustment to more convenient
hours. The embryo plates can also be stored at 4°C for up to 24h, but this has been known to cause subtle
phenotypes. Overnight collections allow the examination
of all stages of embryogenesis, but the distribution of
embryos at the different stages is rarely even.
In this step, the embryos are collected from the
plate, washed well to remove yeast and other detritus
from the plate and the eggshell (chorion) removed.
You should prepare vials ready for the fixation step
(Cl) before starting this step.
- Add about 2ml water to each plate and, with the
aid of a paintbrush, gently release the embryos from
the agar (the eggs are usually partially pushed into the
agar when they are laid).
- Rinse/pour embryos into a meshed basket, made
as shown in Fig. 1, IB.
- Place the basket in a dish (the lids from the apple
juice plates are useful for this) containing 50% bleach
(1:1, water:commercial bleach). It should be made
fresh and can be used for a maximum of 2 days. If
older, the bleach still removes the chorion, but affects
devitellinisation adversely. Swirl the basket in the dish
and add more bleach into the basket. After 2-5 mins,
most embryos will have risen to the surface, indicative
of successful dechorionation. Do not extend the period
in bleach beyond 5min; prolonged exposure can
destroy tissue architecture.
- Wash the embryos thoroughly while in the
basket by swirling embryos under running tap water
or water from a squirt bottle.
- Briefly dry embryos by placing them on a paper
towel. Do not overdry.
|FIGURE 1 Schematic showing the parts that make up an egg-laying cage (IA). Apple juice agar is poured
and allowed to set in petri plates. A dab of yeast is placed on warmed plates that are placed on top of cages
containing flies. The plate is strapped onto the cage and the contraption is placed inverted in an incubator.
The parts that make up an egg collection basket are shown in IB. A nylon mesh/sieve is screwed onto a cut
50-ml Falcon tube by the cap in which a hole has been cut using a hot scalpel blade. A 15-ml cut Falcon tube
can also be used to make smaller baskets using metal wire mesh that is cut into rounds of the appropriate
diameter and fastened onto the tube by melting the end of the tube with a hot scalpel blade. (II) Schematic
showing how sections are cut.
See text for details.
C. Standard Fixation and Removal of Vitelline
- Label a scintillation vial for each different sample
of embryos (stage or genotype) and add 0.8ml of 4%
formaldehyde in PBS.
- Transfer the basket of embryos into a dish containing
heptane. Using a Pasteur pipette, transfer the
embryos with the heptane into the scintillation vial
containing 4% formaldehyde in PBS. The amount of
heptane should be equal to or greater than the volume
of fixative solution. Mix for 20-30mins at room temperature
or at 37°C on a roller or other device.
- Remove the aqueous phase from the bottom with
a Pasteur pipette and then remove the heptane. Add
1ml of new heptane and then rapidly add 1-3 ml of
methanol and shake by vigorous inverting/vortexing
for 1 min. (If you want to stain the embryos with phalloidin,
use 80-90% ethanol rather than methanol at this
and the next two steps; this has the disadvantage
that reduced number of embryos are successfully
- Let the embryos settle for about 10s and then
transfer the embryos from the bottom of the vial to an
Eppendorf tube with a Pasteur pipette. Any embryos
still at the interface between the heptane and the
methanol have not popped out of the vitelline membrane
and therefore will not stain well and should be
left behind. An alternative method for this step is
remove all of the liquid phase, including embryos that
have not settled, leaving only the devitellinised
embryos at the bottom of the vial. Add 1-2ml of
methanol to the embryos and vortex/shake for 30s.
- Wash in methanol three times. For each wash,
squirt in 1ml of methanol so that the embryos disperse.
Let the embryos settle by gravity and remove
the methanol (this is called a gravity wash). Either
proceed to the next step or add 1ml methanol and
store at -20°C (for embryos that will be stained with
phalloidin store in 100% ethanol).
Embryos can be stored in methanol or ethanol
at -20°C for several months and will continue to be
successfully stained by most antibodies. However,
detection of some antigens and GFP fluorescence are
affected adversely by exposure to ethanol or methanol.
In these cases, do steps 3-5 as rapidly as possible and
proceed quickly to step D1. Some antibodies will not
recognise their antigen in embryos fixed by this
method, but will when one of the alternative methods
described in Section III, G is used.
D. Antibody Staining of Embryos
E. Staining with Phalloidin to Visualise Actin
- Wash embryos in PBT (PBS with 0.3% Triton
X-100): First give three to five quick washes, where
1ml is squirted in so the embryos are dispersed, the
embryos are allowed to settle by gravity, and then the
PBT is removed (a gravity wash) and then give one
longer wash (15mins) with mixing. We use a device
that gently inverts the tubes for this and all subsequent
steps, but a variety of mixing devices should work. If
mixing is not sufficient, it will result in variable staining
of embryos in the tube.
- (Optional) Staining with some antibodies is
improved by a 15-min incubation in 0.1% SDS with
mixing, followed by three washes in PBT. This may
help those antibodies raised against bands from SDS
- Block nonspecific binding sites on the embryos
by incubating in PBTB for 30min. Transfer 15µl (5-
20µl) of embryos to a 0.5-ml Eppendorf tube for each
staining with a Pipetteman and a yellow tip with the
last couple of millimeter of the end cut off with a razor
blade. Rinse the tip with some PBT before taking up
the embryos. While the volume of embryos is not critical,
larger volumes can result in uneven staining. The
volume of 15µl of embryos corresponds to about 100
embryos, which is sufficient for most purposes. If more
embryos are desired, use multiple tubes or a larger
tube for staining.
- Incubate embryos with 250-500µl of PBTB containing
an appropriate dilution of each primary antibody
at 4°C overnight. Optimal timing depends on the
antibody and whether staining of internal tissues is
important, in which case longer incubations are recommended.
We have successfully used a range from
2h at room temperature to 2 days at 4°C. If the appropriate
dilution is not known, a good starting point is 1
:1000 for antisera and 1:10 for monoclonal antibody
- Wash off the primary antibody with PBT with
three to five gravity washes and one to three longer (at
least 15 min) washes.
- Incubate with fluorochrome-conjugated secondary
antibodies diluted in PBTB as suggested by the
manufacturer (usually 1:100-300) at room temperature
for 2h or overnight at 4°C. The choice of fluorochrome
coupled will depend on the laser lines of
the confocal you are using. Our preferred ones are
Alexa488, Alexa568 or Cy3, and Cy5. We tend to keep
the embryos in the dark during the incubation.
- Wash secondary antibodies off with three to five
gravity washes and three longer (5-30min each)
washes in PBT. Washing overnight is also fine. The
embryos can be left for a day or two in PBT at 4°C, but
it is better to store them after step F1 or once mounted
on microscope slides.
F. Standard Mounting on a Microscope Slide Steps
- Embryos that have been exposed to methanol
will not stain with phalloidin, so use 90% ethanol
rather than methanol during devitellinisation.
- Dissolve rhodamine-conjugated phalloidin in
methanol (methanol harms the actin, not the phalloidin)
and store at -20°C in aliquots of 6 units. To stain
embryos, vacuum dessicate the required number of
aliquots and resuspend in PBTB at 1 unit/100µl.
- Follow sections D1 and D2. (If phalloidin staining
is to be combined with antibody staining, proceed
through sections D3 to D5 and then go to E4. Phalloidin
can be combined with the secondary antibodies
at D6, at the concentration described later.
- Incubate embryos with 1 unit of phalloidin
(100 µl of the aforementioned solution, plus 400 µl PBT
or PBTB)/0.5 ml tube containing approximately 20µl
of embryos for at least 30min.
- Wash twice with PBT (if combined with secondary
antibodies, wash more thoroughly).
- After the final wash, replace PBT with 50-200µl
of Vectashield or similar glycerol-based mounting medium containing antibleaching agents. The
embryos can be stored at 4°C for a couple of days or,
for longer periods, allow 30min to equilibrate in the
Vectashield and store at -20°C.
- Using a Pipetteman and a yellow tip with the end
cut off, transfer some of the embryos onto a slide. First
suck up approximately 5µl of Vectashield alone and
then the embryos, as this reduces sticking of the
embryos to the inside of the yellow tip. For a 22 x
22-mm coverslip, use 20-30µl per slide. The lower
volume will lead to a mild flattening of the embryos,
which can be an advantage, but is trickier to mount
without air bubbles. As you add the embryos to the
slide, move the tip to spread them out over a square
region a bit smaller than the coverslip. Using a needle
or similar instrument (such as plastic Pipetteman tip
with a very small diameter), gently distribute the
embryos more evenly and separate them from each
other. (Do not fiddle too long with this, as they will
generally distribute into a monolayer when the coverslip
is placed on top, but this step helps reduce or eliminate
the number of embryos lying on top of each
other.) The goals of the experiment will dictate the
optimal number of embryos per slide. For staged
embryos, it is better to have fewer embryos, less than
30 per slide. You can then in a single session on the
confocal completely examine all embryos on the slide
and start a new session with a new slide. This is
helpful because it is easier to find the best embryos at
low power and then add oil for the image collection.
If you have embryos from an overnight collection, a
larger number of embryos improves the chance of
finding each stage on a given slide. With a forceps,
gently place a coverslip over the embryos, starting at
one end to avoid introducing air bubbles and gently
let go of the forceps as the Vectashield spreads. If a
small volume has been used to flatten the embryos, let
sit for 10min and then if there is still air under the
coverslip, add a little more Vectashield to an edge.
- (Optional) Seal the edges with nail varnish (but
see Section III,G,5).
Almost every aspect of this procedure will vary
from fly laboratory to fly laboratory and even within
our laboratory there are a number of variations. None
of the following are critical: which recipe is used for
the egg collection plates (some laboratories prefer
grape juice), the tubes used for fixation and staining,
the solutions used for antibody staining, the timing of
the incubations and the number of washes, and the
medium for mounting the embryos. As mentioned
at the beginning, the key steps are fixation and
devitellinisation. Again there are many variants of this, and the important thing is to get a method that works
well for the antibodies you are using. Once you have
a method that works, try to do this step as identically
as possible each time. This will give you reproducibly
good staining. Generally, the first time a new person
in the laboratory does this technique it does not work
well. However, after a few times, it is hard to understand
how you could have failed the first time. A
simple check is to look at a few embryos at step C4 or
D1 down a dissecting microscope. If the embryos are
still surrounded by the transparent shiny vitelline
membrane, throw them away and try again. Before
moving onto microscopy, we will mention some useful
variants of this technique.
G. Alternative Methods for Embryo Fixation
1. Alternative Fixation Solutions
In these the method is identical to that described
earlier, but instead of 1:1 4% formaldehyde in PBS:
heptane, and 20-30min at room temperature, use the
1:1 4% formaldehyde in PEM:heptane, 20-30min at
1:1 4% formaldehyde in PB :heptane, 20-30 minutes
(Uemura et al.
1:1 37% formaldehyde:heptane, 2-5min at 25-37°C with gentle mixing
This can improve the preservation of cytoskeletal
structures present in membrane compartments [e.g.,
scribble, β-heavy spectrin (Bilder et al.
, 2000)] and
gives the best preservation of microtubules (Foe et al.
2. Heat Fixation (Muller et al., 1996)
Heat fixation improves the penetration of antibodies
into late stage embryos and larvae (when the
exoskeleton normally blocks antibodies).
3. Fixation Conditions for Visualising Nonmuscle
Myosin (Foe et al., 2000)
- With a paintbrush, transfer dechorionated embryos
from the dechorionation basket into a scintillation
vial containing 1-2ml of 0.4% NaCl and 0.3% Triton
X-100 solution that has been heated in a boiled, but
no longer boiling, water bath with its cap half
- Pull the vial out of the hot water bath, screw cap
tightly, and shake once vigorously.
- Uncap the vial and fill up with ice-cold 0.4% NaCl
and 0.3% Triton X-100.
- Leave on ice until cooled.
- Pour off the solution, add heptane and methanol in
equal volumes, and vortex to devitellinize.
- Remove embryos from bottom, transfer to new
tube, and wash twice with methanol.
- Incubate in methanol for an additional hour and
proceed as usual for antibody staining.
4. Methanol Fixation
- Transfer dechorionated embryos to 4ml of fixation
solution: 1 volume 40% formaldehyde (EM grade)
and 3 volumes PBS.
- Vortex for 45 s and add 4ml heptane.
- Shake vigorously for 25 min.
- To devitellinise the embryos, use either 80-90%
ethanol (as for phalloidin described earlier) or hand
peel embryos after replacing the fixation solution
with PBS many times (see Section III,J,1).
- Transfer dechorionated embryos to 1:1 heptane:
(97% methanol, 3% 0.5 M NaEGTA, pH 8.0)
- Shake hard for 2min.
- Transfer embryos from the bottom of the vial to a
1.5-ml Eppendorf tube.
- Wash three times with methanol
- Incubate in methanol overnight at 4°C.
This is a good alternative when a new antibody fails
to stain embryos fixed under standard conditions, as a
number of antibodies have been found to only work
on embryos prepared by this method. Other antibodies
will work with both this method and standard
fixation, while some will not stain with this method.
5. Fixation Conditions for Visualising GFP
If you are planning to visualise GFP fluorescence,
keep exposure of the embryos to methanol or ethanol
to a minimum and do not seal the slides with clear nail
varnish. Organic solvents quench GFP fluorescence or
severely affect its ability to fluoresce.
H. Embryo Thick Sections (Adapted from
Grosshans and Wieschaus, 2000)
To get a good cross-sectional view of the embryo by
confocal microscopy it is necessary to cut sections.
They give far better resolution than optical X
(see later and Figs. 2E and 2F). We have used a relatively
crude method that does not require a microtome
and is compatible with our standard methods for antibody staining. These sections are particularly useful
for the visualisation of morphogenetic events that
occur in the midline and in the interior of the embryo,
such as invagination of the mesoderm, formation of
the midgut, and dorsal closure.
I. Confocal Microscopy
- Fix and stain the embryos according to the
desired protocol. It is hard to combine this method
on phalloidin-stained embryos, because the ethanol
(rather than methanol) treatment results in embryos
that are harder to cut and the morphology is not as
- Follow the antibody staining protocol described
in Section III, C. Keep embryos in PBT after the
secondary antibody has been washed off.
- Refix the stained embryos in 4% formaldehyde in
PBT for 30-60min at 37°C.
- Wash the embryos in PBT, do three gravity
washes, and one 15-min wash with mixing.
- Take the embryos through a glycerol series
comprising of 10, 20, and 30% glycerol in PBT for 1h
each followed by 40% overnight. (This is the optimal
concentration, do not go higher.)
- On clean dusted microscope slides, make bridges
with strips of coverslips cut using a diamond knife.
These are stuck to the slide using clear nail varnish.
- Place an embryo on the slide in a drop of
Vectashield between the bridges (see Fig. 1, IIA).
- To cut the embryo, use a 26G3/8 hypodermic
needle fitted to a 2.5-ml syringe whose plunger has
been removed. Cut at one end in one clean sweep with
the bevelled edge facing away from the slice you wish
to keep. Turn the embryo around with the dissecting
needle. Once again, with the bevelled edge of the
needle pointing away from the slice you want to keep,
cut the embryo to obtain a slice of the desired region
(Fig. 1, IIB). The thickness of the section must be
approximately the same thickness as the bridges; that
is the thickness of a coverslip, which is about one-fifth
the length of the embryo. If the sections are thinner,
they are hard to orient and most of the section is
beyond the depth of focus of the 60x objective; if they
are thicker, the morphology is distorted by the resulting
squashing. With practice you can cut more than
one section from the same embryo, but each should be
mounted on a separate slide due to slight differences
- Remove the undesired embryo pieces with the
dissecting needle (Fig. 1, IIC) and orient the desired
slice so that it lies at the bottom of the drop of
Vectashield with its anterior face up.
- Place a 22 x 22-mm coverslip over the section
and the bridges (Fig. 1, IID). Do not seal the sides to allow for reorientation. If the section is slightly thinner
than the thickness of the coverslip, it may move/turn
in the drop of Vectashield. Slight pressure on the
corner of the coverslip can then bring the section back
to its right orientation. If the section moves under the
coverslip during imaging, abandon that section and
proceed to the next.
- View under a 60x oil immersion objective lens.
embryos can be examined with a variety
of confocal microscope setups. Both inverted and
upright microscopes are appropriate. The key variable
is the available laser lines; this will dictate the optimal
choice of fluorochrome-conjugated secondary antibodies
that one should use. A detailed description of
confocal microscopy is clearly beyond the scope of this
article (a good reference is Centonze and Pawley,
1995), but we hope to provide a few helpful hints.
Once one has obtained an initial quick view of the
result of the staining, which is often possible by conventional
fluorescence microscopy if the signal is
strong, the usual goal is to produce a set of images
for publication. The biggest challenge is to produce
a set of comparable images of different embryos. For example, for a general description of the distribution
of a new protein one would like comparable views of
embryos at different stages; generally a lateral view.
There is a strong convention in the fly field regarding
images of embryos: they should always have anterior
to the left and dorsal up. Similarly, for a comparison
of staining in different mutants, it is essential to obtain
a suitable micrograph from wild type and each mutant
at the same stage and with the same view: this can
often require looking through many embryos. Clearly,
to be able to achieve this you must be able to tell the
stage of the embryo and which way is up, which is not
as easy as it sounds. The only way to achieve this is to
spend time looking at embryos down the microscope.
To learn your way around embryonic development, it
helps to examine embryos that are stained with antibodies
that highlight the development of a particular
tissue. The following antibodies are recommended for
this: Fasciclin III (available from the Developmental
Studies Hybridoma Bank) and phosphotyrosine (available
from Sigma). For an overall view of the embryo,
phalloidin (rhodamine conjugated, Molecular Probes)
is very useful (see Fig. 2).
In general, we start our confocal session by first
scanning through the slide by conventional fluorescence,
taking note of the approximate position on the
slide of embryos of the right stage and orientation that
have stained well. Next we find a well-stained embryo
that is not one of the best ones and use it to work out
the appropriate settings for each channel on the confocal.
When it is essential that there is minimal mixing
between the outputs of each channel, then it is advisable
to do sequential scanning rather than simultaneous.
Use the channel with the brightest signal to get
started. In most cases we use a collection area of 1024
x 768 pixels. Using rapid scanning, find the best focal
plane, adjusting the focus by hand (adjustments to the
eyepieces often result in the view by eye being at a different
focal plane than the resultant scan). Using intermediate
scan rates, set the laser power, iris, gain, and
background levels appropriately for the first channel
and then proceed to the next channel. We keep the iris
similar in the different channels so that they all capture
the same depth of focus. Some confocal setups have
this feature built into the software.
Once all channels are set up, adjust the focus to
obtain the optimal image. Collect images using
Kalman scanning to improve the signal-to-noise ratio.
We use either 166 lines per second with three to five
scans or 50 lines per second and two scans. Make a
note of the orientation of the embryo, especially if you
are just focused on a small region.
Because the embryo is curved, in many cases a
single section will not capture all of the relevant tissue.
In this case, collect a series of sections at different focal
planes and combine them into a single image using the
projection software. The first step is to find the top and
bottom of the series you wish to collect. Turn on the
focus motor, doing fairly rapid scans in a channel with
the most robust signal, and vary the focal plane to find
the top and bottom. Be aware that with an oil objective
lens, the new focal plane can take a minute to "settle."
Set up the software to collect the Z series. Generally
we use a step of 1 µm, with a range from 0.5 to 2.0 µm.
Collect the Z series and project the image. For
some tissues, e.g., the somatic muscles, it helps if
the embryos are mildly flattened by mounting in a
minimal amount of Vectashield, as described in Section
III,F,2; then fewer scans are required to visualise all the
muscles (Figs.2A and 2B).
One other variable to keep an eye on is the zoom.
We standardly keep this at 1 (no zoom), and it is
helpful to check that it has not been varied by a previous
worker, as your images from session to session
will not be comparable. Some judicious use of the
zoom function can improve resolution. In practice we
have found that a zoom factor of greater than two provides
no further improvement in resolution and may
even lead to loss of resolution.
J. Examination of GFP in Living Embryos
|FIGURE 2 A projection of four optical sections of a late Drosophila embryo stained with phalloidin to highlight
actin in somatic muscles (A) is compared to a single optical section (B). Note that some muscles (asterisk
in A) are not visualised in the single optical section. A projection of 42 sections of a Drosophila embryo
stained with phalloidin is used to visualise actin in the whole embryo (C). A single optical section through
the same embryo (D) reveals staining in internal structures that are obscured by the extensive surface staining
in C. A confocal generated
X-Z section along the Z axis depicted by the white line in D is shown in F.
Such a scan provides very poor resolution. A single optical section of a thick section cut (as described in
Section III,H) to include the same region of the embryo as shown in D is shown in E. Such a section resolves
the apicobasal extension of actin especially in internal organs that are obscured in whole embryos. In all
images, the dorsal surface of the embryo is on top.
In A-D, the anterior end of the embryo is on the left.
IV. COMMON PROBLEMS
A. Nonstaining Embryos
- For imaging live embryos, briefly dry embryos
over a paper towel after dechorionation. Alternatively,
the embryos can be dechorionated by hand by gently
rolling on double-stick 3MM Scotch tape with a forceps.
- Transfer embryos in their vitelline membranes
with a paintbrush or forceps to a drop of halocarbon
oil. Cover with a coverslip and leave the slides
unsealed. For time-lapse analysis it is essential to use
a method that allows better access of air to the embryo.
We use a microscope slide consisting of an airpermeable
Teflon membrane in a holder (Edwards et al., 1997; Kiehart et al., 1994).
The most common causes are (i) use of inappropriate
fixation method, (ii) inefficient devitellinisation,
(iii) inappropriate primary and/or secondary antibody,
and (iv) faulty laser, either nonfunctioning or
unsuitable for the fluorochrome coupled to the secondary
This is a common problem inherent to confocal
microscopy, especially during the acquisition of a Z
series at high laser power and high magnification. It
can be reduced by ensuring that the laser power is set
to the minimum value that is optimal for the image.
C. Low-Resolution Images
Low-Resolution Images can result from faulty
image processing of the original image. Obtain the
highest resolution image possible on the confocal.
Information that does not exist in the acquired data
cannot be added by image processing software. Too
high a gain can result in too much noise. Such noisy
images may be improved by increasing either the iris
or the laser power. Out of focus signals resulting from
inappropriate mounting and from inappropriate thickness
of sections are the cause of low-resolution images
We thank Sylvia Erhardt, Anja Hagting, Jasmin
Kirchner, Jordan Raft, Katja Roeper, Alex Sossick, and
Christos Zervas for comments and suggestions.
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