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Immunofluorescence


Fixation:
  1. fix the samples either in ice-cold methanol, acetone(1-10min) or in 3-4% paraformaldehyde in PBS pH 7.4 for 15mins at room temperature
  2. wash the samples twice with ice cold PBS
=4% paraformaldehyde=
fixing in paraformaldehyde for more than 10-15min will cross link the proteins to the point where antigen retrieval may be required to ensure the antibody has free access to bind and detect the protein
=ethanol=
add 100-200uL per slide of cooled 95% ethanol, 5% galcial acetic acid for 5-10min
=methanol=
add 100-200uL per slide of ice cold methanol
place at -20℃ for 10min
methanol will also permeabilize. some epitopes are very sensitive to methanol as it can disrupt epitope structure. can try acetone instead for permeabilization if required.
=acetone=
add 100-200 uL perslice ice cold acetone
place at -20℃ for 5-10min
acetone will also permeabilize. Consequently, no further perpeabilitzation step is required.
=

Permeabilization
If the target protein is expressed intracellularly, it is required to permeabilize the cells
acetone fixed samples do not require permeabilization

  1. incubate the samples for 10mins with PBS containing 0.25% triton X-100 ( or 100uM digitonin or 0.5% saponin). Triton X-100 is the most popular detergent for improving the penetration of the antibody. However, it is not appropritate for the use of membrane-associated antigens since it destroys membranes.
  2. wash cells in PBS three times for 5min
alternative permeabilization concentration


Blocking and simultaneous incubation
  1. incubate cells with 1% BSA in PBST for 30mins to block unspecific binding of the antibodies. (alternative blocking solution: 1% gelatin or 10% serum from the species that the secondary antibody was raised in).
  2. Incubate cells in the mixture of two primary antibodies in 1% BSA in PBST in a humidified chamber for 1hr at room temperature or overnight at 4℃.
  3. decant the mixture solution and wash the cells three times in PBS, 5min each wash
  4. incubate the cells with the mixture of two secondary antibodies in 1% BSA for 1hr at room temperature in dark.
  5. decant the mixture of the secondary antibody solution and wash three times with PBS for 5min each in dark.
Blocking from Abcam
 1% FBS in PBST
Blocking from U.S. Biological
1-2% BSA, FBSA, or 2% non fat milk in PBS with or without 0.2% Tween 20, 0.02% sodium azide



Counter staining
  1. incubate cells on 0.1-1ug/mL hoechst or DAPI (DNA stain) for 1 min
  2. rinse with PBS
Mounting
  1. Mount coverslips with a drop of mounting medium
  2. seal coverslip 
  3. store in dark at -20℃ or 4℃


37℃ warm paraformaldehyde fixation for preserving microtubules in lamond lab.com






making 4% paraformaldehyde
  1. Fill a 2000ml Beaker with about 400ml H2O, put a thermometer into the beaker. Heat beaker at hot stir plate to reach temperature at 62-64°C.
  2. 980ml RNAse free PBS in 1000ml bottle. Put a stir bar into PBS bottle. Add 100ul 10N NaOH (7 drops 10N NaOH) and 40g paraformaldehyde into PBS.
  3. Put bottle into beaker and stir the paraformaldehyde.
  4. Watch the temperature while stiring the paraformaldehyde. When the temperature reach to 62-64°C, the fixative will be clear gradually.
  5. Remove fixative from hot plate, cool down to room temperature.
  6. Filte fixative.
    The pH of this fixative should be around 7.3; check pH is not necessary.
  7. Aliquot fixative and store at -20 °C.

1. For 100mL. Best to use glassware, stir bar set aside for making paraformaldehyde only.

2.  Put 50mL ddH2O in a glass beaker with a stir bar in the bottom. Put stirring on hot plate in the hood. Set hot plate to setting 3-4 (about 60 degrees, make sure the solution does NOT boil).

3. Measure out 4g of paraformaldehyde (Sigma, stored on the bottom left in the cold box). Measure IN THE HOOD (move the scale in there). Add the paraformaldehyde to the heater stirring water.

4. You need to raise the pH until the solution clears. I do this by adding a few drops (2-3) of 1N NaOH and watch the solution. Gabrielle does with 10uL of 10N NaOH and that is it. If the solution fails to clear, check the pH by pipetting a small amount of the solution onto a pH strip. If you are too high (>8) or too low (<7) keep pHing. You cannot pH on the pH meter as it will be ruined by the paraformaldehyde.

5. When dissolved, cool, then filter the solution by pouring through a piece of filter paper inside a funnel. Pour into a graduated cylinder to check the volume.

6. Add 10mL of 10X PBS. Add ddH2O to 100mL.

7. Recheck the pH with paper. Adjust carefully with small amounts of dilute HCl if needed.

8. Aliquot to use.

http://www.westlab.org/protocols/protocols/pF.htm


Subpages (2): Current protocols mountant
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Paul, Hsieh-Fu Tsai,
Jan 26, 2012, 6:19 AM
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