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HOME > Protocols > Histology > Staining protocols > Protocol for Immunofluorescence staining of paraffin tissue sections

Protocol for Immunofluorescence staining of paraffin tissue sections

  1. Clean slides with 95% ethanol
  2. Treat with a solution containing 0.3% gelatin and 0.05% chromium poatassium sulfate
  3. Cut tissue sections using microtome and apply to slides
  4. Deparafinize in xylene for 10 minutes. Repeat twice for a total of three treatments
  5. Hydrate sections through graded alcohols
  6. Rinse three times with distilled water
  7. Incubate slides for 20 minutes in 10% goat serum in PBS (suppresses non-specific binding of IgG)
  8. Wash with PBS
  9. Incubate with primary antibody for 1 hour at room temperature or overnight at 4oC. Optimal antibody concentration is usually from 1-10ug/ml in PBS-BSA
  10. Wash three times with PBS
  11. Incubate with biotin-conjugated secondary antibody for 45 minutes. Again, optimal antibody concentration is usually from 1-10ug/ml in PBS-BSA
  12. Wash three times with PBS
  13. Incubate with streptavadin-fluorescein or streptavadin-texas red for 15 minutes in a dark chamber. This step should be titrated as excess streptavadin-label can lead to high background
  14. Wash a minimum of three times with PBS
  15. Mount aqueous coverslip

Note: Incubations are to be at room temperature or a 4oC humidified chamber. Antibody concentrations here may not hold for all sources. These concentrations should be titrated for each source.


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