Blocking Staining with Immunogen Protocol
This protocol can be used to prepare “blocked” antibody for use in either western blotting or immunohistochemistry.
- 1. Determine the amount of antibody that is needed for your blocking experiment. For example, if your normal experiment is done at a 1:1000 dilution and you need 300ul to stain one slide you would need 0.3 ul of antibody in 299.7 ul of buffer, 300 ul for each slide.
- Dilute the appropriate amount of antibody in 100 μl PBS/TBS. Make 2 tubes. To one tube add the blocking peptide solution at a 5-10:1 ratio to your antibody, i.e. if you added 1 ug of antibody, add 5-10 ug of blocking peptide. Bring the volume to 200ul with PBS/TBS. Add the same volume of saline/PBS (no peptide/antigen) to the antibody in the other tube and label as “no peptide”. Mix gently.
- Incubate both tubes at room temperature for 1hr or overnight at 4°C.
- After incubating bring the volume of supernatant to 300 ul (or what is necessary depending upon the amount of antibody used initially) with buffer (PBS-Tween or your normal antibody staining buffer). Use both antibodies (with and without peptide) for staining using your normal staining protocol.
- Compare staining patterns between the blocked and un-blocked antibodies.
*The above information is only intended as a guide. The researcher should determine what protocol best meets their needs. Please follow safe laboratory procedures.