Immunocytochemistry

Immunocytochemical studies are easily performed on adherent cultured cells. Cells are generally grown in culture dishes, on glass coverslips, or in chambers on slides. Immobilisation of the antigen varies depending on its subcellular localisation. Different fixation techniques must be employed if the antigen is considered to be associated with the cell surface membrane, or whether it may be contained within the cell cytoplasm. Low concentrations of paraformaldehyde are used to fix surface antigens. This is compatible with antigenicity and will usually preserve the cellular morphology. If it is determined that fixation destroys the antigenicity, one may eliminate the fixation step, provided that all incubations are performed at 4°C.

If instead, cytoplasmic antigens are to be detected, the membrane must be permeabilised to allow antibodies into the cell. Two methods have been developed and are in common use to disrupt the lipid bilayer of the plasma membrane and fix the cytoplasmic antigen. The dehydration method uses methanol to extract the lipid bilayer and concommitantly fix the protein of interest, while the cross-linking method uses a two step protocol to first fix cellular proteins with paraformaldehyde, followed by extraction of the plasma membrane with detergent.

NB -There are many permutations of the immunolabelling protocols described below. Ask three people how to do immunofluorescence and you will likely get 3 widely varying answers. The details in the protocols below are meant as starting points only and may be required to be modified depending on your particular cell type and/or experiment. For example, labelling cells with a fluorochrome conjugated phalloidin to visualise the actin cytoskeleton requires that the cells be fixed with PFA and not methanol, as methanol fixation destroys the binding between actin and phalloidin. The protocols below were adapted by Steve through many years of trial and error in the Lamond lab from the “Immunohistochemistry” section in Current Protocols in Molecular Biology, originally contributed by Simon Watkins.

IMMUNOFLUORESCENT LABELLING OF ADHERENT CELL MONOLAYERS

  1. Fix cells as desired (downloadable paraformaldehyde and methanol fixation protocols available on our website).
  2. Permeabilize with 1% Triton X-100 in PBS for 10 minutes at room temperature (omit if cells are fixed using methanol).
  3. Block by incubating cells for 10 min in blocking buffer (see below). We perform all blocking and antibody incubation steps in a humidified chamber (fancy name for a box with a lid and a water- saturated tissue in it…line the bottom with parafilm to put the coverslips on).
    1. [important]Some 2° antibodies are exquisitely sensitive to the reagent used for blocking. This will normally be indicated on the product information sheet of the secondary. Pay particular attention to this![/important]
  4. Incubate cells with primary antibodies diluted in blocking buffer for 35mins to 1 hour. If you have never used the antibody before, try several dilutions (low of 1:100 to high of 1:1000, for example). Check the antibody data sheet if possible for starting antibody concentrations.
  5. Wash coverslips 3 X 10 minute with PBS (can do washes in 6-well plates, then transfer back to humidified chamber for secondary antibody incubation).
  6. Incubate coverslips for 35mins to 1 hour with the appropriate secondary antibodies diluted in blocking buffer, then wash 3 X 10 minutes with PBS. I use fluorophore-conjugated secondary antibodies from Jackson Immunochemicals, see Antibodies.
  7. If desired, stain DNA with DAPI (1:15,000 dilution in water of a 5 mg/ml stock). A good single stranded nucleic acid stain, which fluoresces in the red channel, is Pyronin Y (1:10,000 dilution in water of a 10 mM stock). If used subsequently to DAPI, it will specifically label the ribosomal RNA in nucleoli.
  8. Wash well with PBS and mount cells as desired (mounting media recipes described here). Slides can be stored in the fridge or freezer until analyzed by microscopy.

Blocking Buffer (also used to prepare all antibody dilutions)
1X PBS
1% serum
0.1% Tween-20

Tips:
[important]Note: Appropriate serum must be chosen for blocking. Try to use serum from the animal used to raise the secondary antibody[/important]

[important]Microcentrifuge the diluted antibody solutions (both primary and secondary) for 2 min at 13,500 X g prior to use[/important]

[important]Do not store or mount cells in water as the osmotically competent intracellular membrane systems will be compromised [/important]

IMMUNOFLUORESCENT LABELLING OF SUSPENSION CELLS

Cells may be labelled in suspension, and then layered onto poly-L-lysine coated coverslips, or first layered onto the coverslips and then labelled as described above for monolayer cells. The procedure in solution is essentially the same as for monolayer cells, except that the washing steps are replaced by gentle centrifugation, replacement of the supernatant, and resuspension of the cells. Fixation with PFA/Triton X-100 is preferred, as methanol fixation renders the cells too fragile to be centrifuged reliably.

  1. Start with 0.5-1.0 X 107 cells in media in a 15ml Falcon tube.
  2. Cool cells on ice.
  3. Spin 5 min 800 X g, 4°C in a tabletop centrifuge (for lymphocytes).
    1. Speed is adjusted according to cell type, but must be slow enough to prevent cell damage
  4. Pipet off culture media and resuspend cells in ice cold PBS.
  5. Spin again, pipet off PBS, and fix by resuspending cells in 1-2 ml of fixative (4% PFA in PBS) for 10 min at 4°C.
  6. Spin, pipet off fixative and resuspend cells in 15ml of PBS at 4°C. Incubate 5 min, repeat with a second PBS wash. Use of a large volume of PBS allows a minimal number of washes.
  7. Permeabilise cells by resuspension in 1-2 ml of 1% Triton X-100 in PBS for 10 min at 4°C.
  8. Spin, pipet off fixative and resuspend cells in 15ml of PBS at 4°C. Incubate 5 min, repeat with a second PBS wash. Use of a large volume of PBS allows a minimal number of washes.
  9. Spin cells to a pellet, pipet off PBS wash, resuspend cells in 250µL blocking buffer per 0.5E6 cells (see above for recipe). Incubate 30 min to 1 hr at 4°C.
  10. Spin cells to a pellet, pipet off blocking solution and resuspend cells in 250µL per 0.5E6 cells of primary antibody dilution. Spin the primary antibody at 13,500 X g for 2 min prior to use. Incubate cells in primary antibody for 1 hr at 4°C.
  11. Dilute cell/antibody suspension to 15 ml with PBS. This serves as an initial wash, minimising centrifugation steps.
  12. Spin, remove supernatant, resuspend cells in 15 ml 4°C PBS. Repeat
  13. Pipet off PBS wash, resuspend cells in 250µL per 0.5E6 cells diluted secondary antibody. Spin the secondary antibody at 13,500 X g for 2 min prior to use. Incubate cells in secondary antibody for 1 hr at 4°C.
  14. Repeat steps 11 and 12.
  15. Spin cells to a pellet, resuspend in 100µL of PBS, pipet onto a prepared poly-L-lysine coated coverslip. Let cells adhere for 30 min. Gently aspirate cell suspension.
  16. Stain with DAPI as in the adherent cell protocol if desired.
  17. Wash well with PBS and mount cells as desired (mounting media recipes described here). Slides can be stored in the fridge or freezer until analyzed by microscopy.