Basic immunofluorescence

Cells should be grown on acid-washed coverslips (see below). Six-well or twelve-well culture dishes make the washes easier. We mainly use two fixation protocols. Fixation 1 tends to preserve structure better. Fixation 2 works really well for microtubules and can sometimes uncover epitopes left hidden by Fixation 1 (e.g. for the E10 anti-phospho erk antibody). We use Fixation 3 for some epitopes buried inside organelles, especially cholesterol-rich membranes like late endosomes (e.g. LAMP-1 staining).

Coverslips

Coverslips should be acid-washed to remove spots of dirt or detergent. Place the coverslips in a glass beaker and cover with concentrated nitric acid. Swirl for 5 min. Discard the acid and rinse the coverslips with copious distilled water and then once with methanol. Sterilise the coverslips by baking for 4h at about 80oC. Most cells are fine plated directly onto glass. During plating these coverslips will become coated with vitronectin from the serum in the media. Other cells (e.g. HEK 293) will survive processing for immunofluorescence much better if the coverslips are coated before the cells are plated.

Fixation 1

Wash x1 PBS
Fix for 15min in fresh 4% paraformaldehyde
Wash x1 PBS
Permeabilise for 5min in 0.2% Triton X-100
Wash x1 PBS
Treat 5-10min with fresh 0.1% sodium borohydride
Wash x3 PBS

All solutions are made in PBS. Dissolve paraformaldehyde in PBS by heating to 65oC and leaving to cool while stirring. Do this in a fume cupboard – hot PFA is nasty. Sodium borohydride quenches any remaining PFA and also removes most of the cellular autofluorescence. The fresh solution should bubble, or the borohydride is too old.

Fixation 2

Wash x1 PBS
Dip for 2min in -20oC methanol
Wash x1 PBS

Fisher sell alumina coverslip racks that make the methanol dip a lot easier to handle.

Fixation 3

Wash x1 PBS
Fix for 15min in fresh 4% paraformaldehyde
Wash x1 PBS
Permeabilise with 0.5% saponin for 5 min
Wash x1 PBS
Treat 5-10min with fresh 0.1% sodium borohydride/ 0.1% saponin
Wash x3 PBS

Make up PFA as for Fixation 1. All solutions are in PBS. Saponin permeabilisation is reversible and so all subsequent antibody steps must contain 0.1% saponin.

Staining

Incubate 1h with primary antibody in 1% BSA/PBS
Wash x3 PBS
Incubate 45min with secondary antibody in PBS
Wash x3 PBS

Centrifuging the diluted primary and secondary antibodies in a microfuge for 15min, 4oC removes any precipitated material and gives cleaner coverslips. An alternative blocking agent to BSA is 1% fish skin gelatin, which may give better results with cow or sheep antibodies. A good way to save on antibody is to incubate with a small (80ul) drop placed on the coverslip. Try a starting dilution of 500ng/ml for primary antibodies. For a long time we used Cy2 (green), Cy3 (red) and Cy5 (far red) conjugated secondary antibodies from Jackson. These are very good, but the Alexa range of dyes are better and so now we mainly buy Alexa conjugated secondaries from Invitrogen.  For multiple labeling, use antibodies raised in donkey that have been precleared against other species. We reconstitute as directed and then add an equal volume of glycerol and store the antibodies at -20oC. Use the reconstituted antibodies at 1:200.

Mounting

Dip the coverslip briefly in water, blot the edge on a piece of tissue and quickly mount onto about 10ul of Mowiol mount on a microscope slide. Leave the slides at room temperature in the dark for about 2h until the Mowiol has set. Store at 4oC.

Mowiol mount is a PVA-based mount which sets as a hard resin. It has the advantage that cells under the coverslip do not dry out and the slips do not require sealing with nail varnish.

In a 50ml Falcon tube, mix 6ml glycerol and 2.4g of Mowiol 4-88 (Merck #475904) and 6ml water. Vortex and then shake for 2h. Add 12ml of 200mM Tris-HCl, pH 8.5 and incubate at 50°C with occasional vortexing until the Mowiol dissolves (approximately 3h). Filter through a 0.45um syringe filter and store in aliquots at 4°C. Just before use it is a good idea to add an antifade reagent to prevent photobleaching of the green signal: add 2.5% (w/v) 1,4-diazabicyclo-[2.2.2]octane (DABCO, Sigma #D2522) and vortex for about 30s to dissolve.

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