Hi all
I am having a trouble with my immunohistochemistry. I am trying to detect a secreted protein in FFPE mouse tissue sections, and I use concentration-matched goat IgG as my negative control. The secondary antibody is biotinylated rabbit Anti-Goat IgG Antibody. I block with 3% H2O2 for 20 minutes. After secondary antibody incubation, I treat the tissue sections with TSA, and the colorimetric detection is performed by AEC.
My expectation with the negative control would be such that there isn't any signals at all, but I always end up getting unknown staining.
Does it have anything to do with treatment with TSA after secondary antibody incubation? Any advice would be much appreciated.
Have you tried adding serum of host species of secondary Ab to your blocking solution to prevent any potential non-specific binding? Also, I would highly suggest doing a trial run using one slide with just primary (no secondary), one slide with just secondary (no primary), and one slide performed normally with both, just to confirm there isn't an issue with any of the antibodies.
My guess would be one of the following may the culprit:
1) cross-reactivity of labeled secondary with endogenous immunoglobulins of tissue
2) inadequate washing or blocking time
3) secondary antibody not diluted enough, or possibly contaminated
Primary antibody tag with inert protein like KLH, so sometimes cross reactivity. You do one thing, preincubation with TSA and then primary anitibody and follow the same protocol.
Have you tried adding serum of host species of secondary Ab to your blocking solution to prevent any potential non-specific binding? Also, I would highly suggest doing a trial run using one slide with just primary (no secondary), one slide with just secondary (no primary), and one slide performed normally with both, just to confirm there isn't an issue with any of the antibodies.
My guess would be one of the following may the culprit:
1) cross-reactivity of labeled secondary with endogenous immunoglobulins of tissue
2) inadequate washing or blocking time
3) secondary antibody not diluted enough, or possibly contaminated
i have a similar conclusion like Blair Austin: it can be that the concentration is not the optimal one so you need to make a series of decreasing concentrations and choose the one with best signal:background ratio.
also you may need to choose another blocking buffer and use it also to dilute the secondary antibody or increase the blocking time and do not wash after blocking
another thing is that you should, as mentioned by Blair, make control staining using the usual protocol you use but skip the secondary antibody or the primary antibody
i hope that may help
Your 3% hydrogen peroxide block step serves to block the activity of endogenous peroxidases, which is critical given your detection method. However from the description you provided from your protocol, it looks like you do not include a block step for non specific binding of primary antibody. You can do this after you rinse off your slides from your peroxidase blocking step. As Blair Austin said, this is usually conducted using normal serum from the species your secondary antibody was raised in (in your case rabbit) to which you can add .1 to 5% of something like BSA.
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