1) Dissect eye imaginal discs; from larvae in 0.1M NaPO4 (pH 7.2). It is convenient for subsequent handling to leave the discs attached to the mouthparts. (See detailed instructions.)
2) Fix the discs. Usually we use PLP for 45 min on ice. PLP gives fixation results comparable to glutaraldehyde but preserves antigenicity. Of course the interaction of each antibody with its antigen is unique and may require individual treatment. Less commonly we use 4% paraformaldehyde in PEM for 30 min at room temperature, particularily with nuclear antigens (PLP is a better membrane fix and we guess may reduce accessibility of the nucleus to antibodies). (See detailed instructions.)
3) Wash discs in 0.1M NaPO4 (pH7.2), 0.1% saponin for 15 min on ice.
4) Dissect away the peripodial membrane. Removing the peripodial
membrane increases accessibility of the epithelium to antibody and visibility
of the stained tissue under the microscope. For some antibodies the improvement
is neglegible (eg: Mab BP104). For others this step is essential (eg: Mab22C10).
(See detailed instructions.)
5) Incubate in primary antibody, 1-2 hr on ice. Make up antibody in
0.1M NaPO4 (pH7.2), 5% normal goat serum, 0.1% saponin (store frozen or filter
sterilized at 4°C). Use monoclonal supernatants at 1:1 dilution (but
Mab22C10 at 90%), ascites or sera at 1/250 - 1/5000 as appropriate. Final
saponin concentrations of 0.01% - 0.1% are satisfactory for light microscopy.
(See detailed instructions.)
6) Wash 3X in NaPO4 + serum + saponin.
7) Incubate in secondary antibody, 1-2 hr on ice. Generally we use HRP-coupled goat anti mouse/rabbit from BioRad at 1/200 - 1/500.
8) Wash 3X in NaPO4 + serum + saponin.
9) Incubate in diaminobenzidine (for HRP coupled secondary). The reaction is generally complete after 15-30 min, and for many antibodies staining can be much briefer. Staining solution is 0.5 mg ml-1 DAB, 0.1% saponin, 0.1 M NaPO4 pH7.2, 0.003% H2O2. (See detailed instructions.)
10) Wash in 0.1M NaPO4 .
11) Transfer to a screw-cap vial in 0.1M NaPO4, such as the 1 dram/3.7 ml bottles from Fisher.
12) Postfix and intensify in 2% OsO4 in 0.1M NaPO4 , from 2min on ice to 10 min at room temperature, as desired. (See detailed instructions.)
13) Wash in several changes of 0.1M NaPO4. Discs can be stored at 4°C at this stage.
14) Using a pasteur pipet, transfer to 50% ethanol in a petri-dish (60x15mm). After 5 min transfer to 100% ethanol, and then again to fresh 100% ethanol.
15) Mount in DPX. (See detailed instructions.)
Detailed Instructions for steps numbered:
1) To remove eye-antennal discs, hold the larva firmly about halfway
down the body, using forceps. With a second pair of forceps, grasp the
mouthparts and pull them out of the head. Usually the eye-antennal discs
come out with the mouthparts. Other tissues may also remain attached; still
holding on to the mouthparts, remove the brain and salivary glands, if present.
Then grab the internal part of the mouthparts (forceps B in Fig.
1) and remove this and the eye-antennal discs from the external
part of the mouthparts (still held by forceps A in Fig. 1). All other tissues
should get left behind at this step.
To make 1 liter 0.1 M NaPO4 pH7.2 use:
10.22 g Na2HPO4.H2O OR
12.82 g Na2HPO4.2H2O OR
19.30 g Na2HPO4.7H2O OR
25.79 g Na2HPO4.10H2O plus 3.86 g NaH2PO4.H2O OR
4.37 g NaH2PO4.2H2O
5) Antibody incubations, washes, and DAB reactions are carried out
in the 60-well (conical bottomed) microtest plates manufactured by Nunc.
Use a wire hook to transfer the discs from one solution to the next. Each
well takes 13 µL. Incubations can be left overnight at 4°C if the
lids are sealed with parafilm to reduce evaporation.
9) Unsubstituted benzidene is one of the most potent chemical carcinogens
known. Diaminobenzidene is believed to be safer but should be handled
carefully. Sigma sells DAB in sealed "isopacks" which are convenient
for making stock solution. Aliquots should be stored at -70°C. All
materials that contact DAB should be soaked with bleach prior to disposal
(I like to handle DAB inside a heatseal bag, which the waste never leaves).
12) Osmium intensification produces a background staining of the tissue
that we find useful as a counterstain. Cleaner intensification can be obtained
by adding Co and Ni to the DAB reaction (add 30 µl 50 mM CoCl2 50 mM
NiCl2 to 1 ml of buffer immediatly before the DAB. Ignore the clouding due
to precipitation of the phosphates)(Adams, J.C., J. Histochem. Cytochem.
29, 775, 1981). In this case a different postfixation will be required
before mounting. 2% glutaraldehyde can be used (and results in slight destaining).
Alternatively a graded series of ethanols is possible (30%, 50%, 70%, 90%,
100%), but this will not preserve delicate structures such as the apical
photoreceptor-cell membranes. Of course EM-intensification protocols can
also be used, such as the silver-gold method (Liposits et al., Neuroscience
, 13, 663, 1984).
SAFETY. Both osmium tetroxide and glutaraldehyde are volatile neurotoxins
with cumulative effects, and should be handled in a fume-hood. Osmium
tetroxide fixes the brain via the nasal mucosa, repeated glutaraldehyde exposure
can cause blindness. Osmium tetroxide has a pungent odour, somewhat
like garlic. Glutaraldehyde has only a mild smell. Waste solutions should
be inactivated by pouring onto excess "protein" (eg glycine or powdered milk).
OsO4 is sufficiently volatile that solid waste, eg gloves, is clean if
left overnight.
15. Mount as follows: Take up discs in 100% ethanol using pasteur and
drop onto a clean microscope slide. Wipe away excess ethanol with a tissue.
Add one drop of DPX mountant (FLUKA) before the discs dry, but when little
ethanol remains. Remove the mouthparts and orient the discs apical side upwards
using fine forceps. If desired, remaning peripodial membrane or other debris
can be knocked off at this point. Carefully lower a 22x40 mm coverslip down
from one side, watching all the while to see that the discs do not turn over.
To some extent disc rotation can be controlled by moving the coverslip.
If this is unsuccessful, remove the coverslip and reorient the discs. Another
drop of DPX will be necessary before replacing the coverslip. As soon as
you are satisfied, flatten the discs with smooth firm pressure on the coverslip
(eg: with the butt end of a pair of forceps). Releasen the pressure equally
smoothly. Do not attempt to remove the coverslip once the discs are flattened.
B. Cobalt Sulphide staining:
CoS staining offers a rapid method for visualizing the apical borders of disc cells. The protocol given here is that of Tomlinson & Ready (Dev. Biol., 120, 366, 1987).
1) Dissect discs in 0.1 M NaPO4 pH7.2.
2) Fix in 2% glutaraldehyde in 0.1 M NaPO4 pH7.2 (handle in the fume
hood), 30 min, room temperature.
3) Wash in 0.1 M NaPO4 pH7.2
4) Wash in H2O.
5) Dissect away the peripodial membrane as for antibody staining, except
that the discs are still in H2O. Also it is necessary to remove
the membrane, not just peel it back, or else it will snap back over the disc
in subsequent steps.
6) Incubate in 2% CoNO3 in H2O. 96-well microtest plates are useful
here too.
7) Transfer to 1% (NH4)2S in H2O, until the disc is coal-black (typically
about 30s).
8) Transfer to H2O. Watch the disc destain. It is about done as the
black becomes dark grey and some details can be seen (typically about 30
s). Slightly before this point, transfer to a drop of glycerol on
a microscope slide and mount under a coverslip. Do not squash the disc,
allow it to be gently flattened by the cover-slip. The staining is not stable
and data should be recorded as soon as possible. If the coverslip is sealed
with nail polish and the slide stored at 4°C the staining seems to be
OK for a few weeks, otherwise 24 hr is about the safe limit.
The stain is essentially a precipitate coating the apical disc
surface and is very delicate. It is easily destroyed if the discs are handled
roughly. Once the peripodial membrane has been removed it is important not
to handle the apical surface during transfers. Make sure to get all the
glycerol off the wire hook before starting each new prep.
C. Basic Fuchsin staining
Basic fuchsin is a nuclear stain. Mitotic chromosomes and, at other
stages of the cell cycle, spindle pole body associated material are especially
well stained. Basic fuchsin is useful for following cell division and cell
death (in dying cells the whole cell is labelled). The version used here
was adapted from Weischaus and Nüsslein-Volhard, p 223 in Drosophila,
a Practical Approach , (ed D.B. Roberts), IRL, Oxford/Washington, 1986.
Stainless steel mesh baskets, of the kind used for embryonic cuticle preps.
are useful for handling the discs.
1) Dissect discs in 0.1 M NaPO4 pH7.2.
2) Transfer to Fix A (4 ml 95% EtOH, 0.5 ml CH3COOH, 1.6 ml 20% paraformaldehyde)
at room temperature for 10 min.
3) Transfer to Fix B (4 ml 95% EtOH, 0.5 ml CH3COOH, 0.4 ml 20% paraformaldehyde)
at room temperature for 1 h.
4) Wash thoroughly in 70% ethanol.
5) Incubate in 2 M HCl for 10 min at 60 °C.
6) Wash with H2O.
7) Wash with 5% acetic acid.
8) Stain in 1% basic fuchsin in 2.5% acetic acid for 20 min at room
temperature. The fuchsin solution should be made up fresh every few weeks.
9) Destain in 5% acetic acid. Do not overdo this step - usually
a few minutes is enough. A few salivary glands can be included as a control,
as their nuclear staining is easily visible with the dissecting microscope.
10) Dehydrate in 70 %, 95% and 100% EtOH, and mount in DPX.