Houston Methodist. Leading Medicine.
Houston Methodist. Leading Medicine




Immunofluorescence Staining Formaldehyde fixation for cultured/cytospun cells

All steps are performed at room temperature unless otherwise indicated. Square and rectangular coverslips may be fixed and washed in small Coplin jars, 35 mm petri dishes, or 6 well TC trays. Small round coverslips will fit into 24 well trays for processing. Since larger volumes of buffers, etc. are required for Coplin jars, using the "drop" method of immunostaining may conserve antibody.

Place the coverslips in a petri dish on a piece of parafilm, cell side up. A few drops of the diluted antibody can be added on the top. Alternatively, the coverslip may be inverted on a drop of antibody solution - however, loosely adherent cells may come off when the coverslip is pulled off the parafilm. Sealing the edge of the dish with tape or parafilm during incubation, plus placing a small wad of wetted kim-wipe in the dish, will keep the coverslips from drying out. Note: This protocol has been optimized for HeLa cells, and works well for many other cell lines. However, some cells may require adjustments in length of incubations, and concentration of formaldehyde or Triton X-100.

Part 1: Initial Fixation, with or without extraction

Aspirate or pipette off media. Wash X 2 with ice-cold PBS (containing Ca++ and Mg++). For "Whole fixation" fix the cells with 4% Formaldehyde (EM grade only) in PEM buffer for 30 min. on ice (1:4 dilution of 16% stock).

The fixation buffer should well cover the surface of the cells so that they do not dry out (about 250 m l buffer for 24 well trays, 1 ml for 6 well trays, etc.) Remove formaldehyde fix, discard in toxic waste bottle, and wash X 3 with PEM buffer, about 2-5 min. per wash.

Incubate 5-30 min. with PEM + 0.5% Triton X-100. This step permeablizes the cells which is essential for antibody penetration during immunostaining. Some cells are permeabilized fine in only 5 min, some take longer, e.g., 30 min; it should be determined empirically. Wash X 3 with PEM buffer, about 2-5 min. per wash.

Notes: If working with cells transfected with GFP plasmids that are not to be immunostained, you may skip step 5 and proceed to the final step in part 3, the DAPI counterstaining. Aspiration using the vacuum line may remove loosely adherent or mitotic cells. If the goal is to preserve the loose cells, pipetting solutions off manually will help.

Part 2: Immunostaining

Block with 5% powdered milk in TBS-T buffer plus 0.02% sodium azide (commonly called Blotto). Blocking time can be 30-60 min, or overnight at 4C. 3 percent serum from the 2nd antibody species, e.g., normal goat serum if using Goat anti-mouse IgG), can also be used. Remove blocking buffer and add Primary Antibody diluted in blocking buffer. Incubate either 30 min at 37 C, 1-2 hours at RT, or overnight at 4C. Remove Primary Antibody (it can be saved, frozen and reused a number of times). Wash 4 - 5 times with blocking buffer, 1 min. or more per wash. Can hold at this step if needed. Add Secondary Antibody diluted in blocking buffer. Incubate 30 min at RT protected from light (cover with foil). Remove and save Secondary Antibody. Wash X 5 with TBS-T, then wash with PEM. NOTE: Sodium azide is toxic and can cause corrosion of plumbing-flush sinks with large volumes of water. Do not use Sodium azide if the secondary antibody will be HRP labeled, since it inhibits enzyme activity.

Part 3: Post-Fixation and Quenching

The coverslips will need to be protected from light from now on (cover with foil during incubations; low ambient light is best). Fix 10 - 30 min. in 4% Formaldehyde in PEM buffer. (This is especially important for cells to be imaged at 100x with the deconvolution microscope; small structures may wiggle and result in blurry images, no kidding!). Remove fix (discard in toxic waste) and wash X 3 in PEM, 2-5 min per wash. To quench auto-fluorescence, add PEM buffer to pre-weighed NaBH4 in a PP conical tube. Mix up and down and add to coverslip(s). Incubate 5 min. Remove PEM/NaBH4, prepare a fresh tube and repeat. Check coverslips during this step to make sure they are not floating on top of the buffer due to the bubbling action. Note: The NaBH4 should be stored in a dessicator. The final concentration of the NaBH4 in PEM should be 1 mg/ml. It can be difficult to weigh out a specific amount of NaBH4 - it works best to just weight some out, note the amount of mgs. on the tubes, and then add the appropriate volume of PEM buffer. Plan on preparing double the volume used for fixative, etc. The tube should be a large one, since the NaBH4 will begin foaming, (if it does not foam, it should be discarded and new stock purchased). Wash X 2 @ 5 min. with PEM. Alternately, 0.1 M ammonium chloride diluted in PEM buffer can be used to quench. Incubate for 10 min. Wash X 2 with PEM following this treatment, then 1 X with TBS-T. Counterstain DNA with 1X DAPI diluted in TBS-T for 30 sec. to 1 min. Remove DAPI and add TBS-T.

Part 4: Mounting the Coverslips

Label slides using a marker designed for slides - pencil can be difficult to read in the microscope room, and immersion oil can smear some ink markers. If using SlowFade Kit (Molecular Probes), coverslips can (optionally) be incubated with 1-2 drops of Equilibration Buffer for 5-10 min. before mounting. Place a drop of mounting media on the slide for each coverslip to be mounted (two drops for square coverslips). One square or rectangular coverslip, or 3-4 round coverslips will fit on each slide (keep coverslips to the middle of the slide as much as possible; their edges can touch). Pick coverslip up with forceps, keeping the cell side toward you. Dip the coverslip in a beaker of water several times to rinse off the buffer salts. Carefully blot the edge of the coverslip with blotting paper or a kim-wipe to wick away the excess water. Alternatively, the coverslips may be propped against a box resting on a paper towel or kim-wipe and allowed to "drain" briefly, keeping track of the cell side orientation. Don't allow the coverslips to dry out. Slowly lay the coverslip on the mounting media, starting at one edge, to avoid creating bubbles. As the coverslips are mounted, place the slides in a holder, drawer, or other means of protecting from light. When finished mounting, excess mounting media will spread out from under the coverslips. Suction this off. The suctioning may need to be repeated several times - if you don't do this the nail polish won't stick. Seal the edges of the coverslips with nail polish. If coverslips need to be removed for some reason, you can peel off the nail polish and "float" them off with excess TBS-T buffer. Coverslips are now stained and ready to examine. Store at 4C to prolong fluorescence.

Reagents and Supplies:
PBS & TBS-T buffers:
PBS: Standard PBS for mammalian cells
TBS-T: Tris HCl 20 mM, pH 7.4 NaCl 137mM Tween-20 0.1%

PEM Buffer:
Stock Solution for 1 Liter:
Final Concentration: 400 mM Potassium PIPES, pH 6.8 200 mL. 80 mM 0.5 M EGTA, pH 7.0 10 mL. 5 mM 1.0 M MgCl2 2 mL 2 mM
Filter sterilize and store at 4 C.
Formaldehyde: Use EM grade formaldehyde - it can be obtained from the following suppliers:
Polysciences, Cat. #18814, 10 mL. ampules, 16%
EM Sciences, Cat. #15710, 10 mL ampules, 16%
Tousimis, Cat. #1008A, 10 mL ampules, 20%
Cover opened ampules with parafilm and store at 4C (best used fresh, but a few days to a week in at 4C, if sealed well, should ok. New testing of antibodies/etc should always use a new aliquot of formaldehye.

Triton X-100:
Sigma, Cat. #X-100-PC, Triton X-100 Peroxidase & Carbonyl Free, 5 ml ampule. Use during extractions on unfixed cells.
Sigma, Cat. #X-100, Triton X-100, 100 mls. Can use for permeablizing cells post fixation.
For ease of use, make a 10% stock solution of the Triton X-100 in PEM buffer and aliquot to microcentrifuge tubes covered with foil. Store the stocks at -20 C.

Round coverslips will fit in 24 well trays, and can be obtained from:
Fisher Scientific, Cat. #12-545-80, #1 thickness 12 mm circles
Thomas Scientific, Cat. #6672-A75, #1 thickness 13 mm circles
Rectangular coverslips will fit in small Coplin jars (available from Thomas Scientific) or two per well in a 6 well TC tray or 35 mm petri dish:
Thomas Scientific, Cat. #6663-F10, #1 thickness 11 X 22 mm rectangular coverslips.
22 X 22 mm coverslips may also be used, but will require larger volumes of reagents:
Fisher Scientific, Cat. #12-520-B, #1 1/2 thickness 22 X 22 mm square coverslips.

Note: # 1 1/2 thickness coverslips are preferred for microscopy, but are not widely available. The coverslips will need to be pre-cleaned with 100% ethanol and sterilized before use if cells are to be grown on them (see acid washing protocol). They can be sterilized by microwaving them at several short 30 sec bursts, or by speading them out under UV light. It may be necessary to acid wash them or coat them with poly-L-lysine to keep the cells from coming off.

Use pre-cleaned slides, such as Fisher Colorfrost slides, or clean them before use with ethanol.

Mounting Media:
For most microscopy: Vector Labs - Vectashield Mounting Media, Cat.# H-1000, 10 mls.
For deconvolution microscopy: Molecular Probes - SlowFade Anti-fade kit, Cat. #S2828. (Vectashield contains some non-soluble particles which can have a "lensing" effect at high resolution).



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