Freezing Tissues for Sectioning

1. Cut fresh moist tissue into 2mm cubes. (Stomach, esophagus and intestine from big animals can be opened, cut into strips and rolled. In the case of small animals roll intestine or esophagus inside strips of stomach).

2. Wet forceps with PBS and throw tissue cubes individually into a beaker of liquid nitrogen — don’t put too many in at once or they will stick together.

3. Place cubes into cryovials precooled in liquid nitrogen. Take care not to have liquid nitrogen in the tubes when you seal the lids or they will become dangerous bombs.


Sectioning Frozen Tissues

4. Mount tissue cubes in a small blob of cryosupport medium on the chucks 30min before you want to cut sections.

5. Use 0 degree angle with knife. Advance block towards the knife. Trim block face using 30mm sections. When the desired area is produced change to 5mm and cut and discard next three sections checking that the roll plate is positioned correctly to give sections that are the same shape as the block and free of wrinkles.

6. Cut 5mm sections and press onto a multiwell glass slide. The chuck must be below the knife while doing this. Brush away section imprint on the knife firmly.

7. Sections in wells 4 and 8 will not photograph well because this region of the slide is not supported on the microscope.

8. Sections of stratified epithelia benefit from air drying overnight otherwise they tend to come off the slide. Other sections should be air dried for 2hrs then used or stored at -20° C (with silica gel crystals to keep them dry). Some tissues such as liver and kidney always look best when cut, stained and photographed on the same day. Tissues vary in their degree of autofluorescence; liver is particularly bad and gives an orange glow.




Tissue sections can be directly stained or first fixed for 5mins in -20C acetone. If the antigen is masked try using saponin in the wash buffers.

Cultured cells should be rinsed in PBS to remove FCS then dipped very quickly in H2O to remove salt and fixed for 10mins in -20C methanol

The sections should never be allowed to dry out during the immunofluorescence procedure; for this reason, set up a humid chamber in which to keep slides during staining (rest the slides on a layer of wet paper towels in a pan and keep them covered during incubation periods).

1. Apply 10-15ul 1X PBS (alone) to each well. (This lowers non-specific binding.) Flick off, then dry between wells with a Kimwipe or remove liquid with vacuum aspirator.

2. Apply 10-15ul of antibody diluted in 1XPBS/1:10,000 azide for 30mins at RT (antibody dilution 1:50 for sera).

3. Shake off antibody and dip slide in 400mls 1XPBS. Wash 3X 10mins in 1XPBS.

4. Dilute secondary antibody in 1XPBS/1:10,000 azide according to manufacturer's instructions and spin for 5mins in a microfuge to pellet aggregates of free fluorescein. Dry around wells with a Kimwipe or use vacuum aspirator and apply 10-15ul secondary antibody for 30 min at RT.

5. Shake off antibody and dip slide in 400mls 1XPBS. Wash 3X 10mins in 1XPBS.

6. Mount slides with coverslips in Gelvatol while still wet (Gelvatol is PBS based) or use Gelmount and in this case seal with nail polish. Store in dark at RT.

7. Controls include:

no antibodies –for autofluorescence

PBS + secondary antibody–background due to secondary antibody

Known positive primary ab + secondary ab–test of technique



To enhance nuclear localization

Make all solutions and dilute all antibodies in PBS containing 2mM MgCl and 0.5mM CaCl (PBS-MC)

Rinse cells in PBS-MC

Fix as usual in -20C methanol/acetone 50:50 for 5 mins

Air dry

Apply antibodies diluted in PBS-MC for only 15 mins

Wash for only 3mins in PBS-MC

Apply secondary antibody diluted in PBS-MC for only 15 mins

Wash for only 3mins in PBS-MC

View while wet under the microscope to determine background if acceptable mount slides in gelvatol.