Immunostaining of frozen sections of mouse embryos

From Susan Wheatley

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Fixation

Dissect into PBS and fix in 4% paraformaldehyde in PBS 4 h - o/n, depeindiingon the size of the embryo.

Wash out pfa with PBS. Can store at this stage by dehydrating through 25, 50 to 75% EtOH in PBS and keeping at -2OoC. Reverse the series when ready to use.

Embedding

Saturate embryos with 15% sucrose in PBS, by incubating embryos with this solution, up to o/n

Make mixtures of 25, 50 and 75% OCT compound (TissueTek) in sucrose solution. Incubate embryos 20 min to 1 h, depending on size, in each one. Place embryo in cryomould (TissueTek), remove excess OCT/sucrose. Fill up mould with neat OCT and freeze on dry ice. To store, wrap in parafilm and keep at -7OoC.

Sectioning

Cut 10 um sections on cryostat. Stick onto charged slides, or TESPA coated slides (have not tried this yet, but some sections have fallen off charged slides). Probably best to avoid gelatine coated or other types of protein coating as this may interfere with staining.

Air dry and store at -2OoC in sealed box with silica gel inside.

Immunohistochemistry

Slides should be dry before starting staining as they will drop off the slides if not. Should be fine after storage with silica gel. It may be worth placing in a warm, dry area for a while if staining immediately after cutting, eg. on top of an incubator or oven.

Wash off OCT with PBS.

Bleach embryos with 0.6% H202 in PBS, 20 - 30 rmn.

Block sections with serum (foetal calf serum or goat serum), 5-10% in PBS/0.1% Triton-X-l00. 20-30 min.

Add primary ab @ 1/100. Dilute in PBS/serum[I'riton. I make up 250 ~ for each slide, dry the wrong side of the ~ide with tissue and drain the right side, but don't let it dry out, then drop the ab carefully onto the slide with a Gilson tip. Then lower a coverslip gently over. Incubate om in a sealed, humidified sandwich box, at 4oC. I put the slides in large weighing trays in a box and pour water in the bottom.

Wash with PBS/Triton, 2 changes about 15 min each.

Add secondary ab @ 1/100 in exactly the same way as primary ab. I use Vector Laboratories goat anti-rabbit Ig, HRP conjugated.

Wash with PBS/Triton.

Rinse with PBS.

Make up a solution of 250 ug/ml diaminobenzidine (DAB) and 0.08% nickel chloride and incubate sections in this for 10 min. Add H202 to 0.03% and watch the colour reaction. You can monitor the colour development under the microscope with a cover slip on the slide to prevent it drying out. When the desired colour is reached, wash out with PBS. I have been leaving my sections to develop for between 10 and 20 min.

To mount sections permanently, dehydrate through 75, 85 then 100% EtOH in water. 10 sec, 15 sec, 2 min in each. Then dip in Histoclear until the EtOH and Histoclear have mixed. Drain excess and mount in DPX.

Immunofluorescence

I have also tried immunofluorescence and this has worked well with one fluorescent ab that I used. It was CY-3 conjugated (looks red). There was some autofluorescence in the liver and heart, so I will have to look into improving this. The protocol was the same, except that I did not bleach the sections. The secondary ab was also diluted to 1/100.