First of all, which antibody you are using? We don't have a problem with mouse antibody in mouse brain. The protocol we use is 1% goat serum with goat anti-mouse secondary. It would be nice if you can provide more details then we can really have a look what is going wrong.

How is your tissue prepared? Is it fixed? If so, in what? For how long? Is the tissue paraffin embedded? What antibodies are you trying to use? What staining protocol? If you are using a donkey secondary I would suggest using Donkey serum in your blocking instead of horse serum, FBS or BSA. Are you getting any staining at all? If so, is it non-specific? More information is needed to help with your problem.

I too faced so many problems when using mouse on mouse antibodies. mIgGk-BP-FITC is a very good alternative to traditional secondary antibodies. You can get this from santa cruz biotechnology inc. sc-516140

All the best

Simon Chang Thank you for your suggestion! I should probably purchase new goat serum. Since none of my coworkers are using goat serum, all I could find is an aliquot prepared back in 2013 at this moment.

I am the only one who does neuroscience research in diabetes lab.

Renee R Donahue Thanks for your comment! Please refer to my description again. (I added more information.) It is difficult for me to get suggestions from my coworkers since I am the only one who does neuroscience research in diabetes lab.

I frequently use epigenetic markers (ie. rabbit host H3K4 methylation).

I'd like to co-stain with rabbit host antibodies, and hence I need to use mouse host antibodies (popular species), like NeuN from Millipore.

I see non-specific binding. There is no detection under confocal microscope.

Siva Ramakrishna Uppalapati Thank you for your suggestion! Which blocking (and concentration) are you using?

Yuka Obayashi I would suggest for long term storage of your samples, keep them in anti-freezer in -20. And for the NeuN antibody from Millipore you can try 1: 400 in 0.5% triton PBS with 1% goat serum.

Simon Chang Thank you! Do you use agar/agarose embedded samples (prior to sectioning)? I figured out that my samples got a lot of winkles after transferred to -20oC. I thought that it was due to agarose.

Oh I just used 1:1000. I should probably increase concentration.

Yuka Obayashi Yes, we use 6% agarose. I think it has nothing to do with the agar but with the buffer you store your samples.

Yuka,

I have a couple of suggestions. First, we typically use a 4 hour post-fix for CNS samples. However, most antibodies will work with an overnight post-fix. After the post-fix, transfer your samples to a 30% buffered sucrose solution. The samples should remain in this solution until they sink to ensure proper freezing (this may be why your samples wrinkled after transfer). Once the samples sink, they are ready for freezing and cutting. Once tissues are cut, follow this protocol:

Day 1

Day 2

CNS usually does not require antigen retrieval. However, the type and length of fixation can affect this and mask your antigen. I think overnight with 4% PFA may be causing a problem masking your antigen. I can suggest some antibodies that are tried and true for us as well if you would like. Alternatively, you could try using a method of antigen retrieval to try unmasking your antigen from the paraformaldehyde. I would also try to run a positive control and a negative control with your IHC each time as it can help you answer some questions about what is going on. Hope this helps.

Do you have any chance to test it on fresh sample? the antigen might be masked.

see below:

https://bitesizebio.com/13454/antigen-retrieval-techniques-for-immunohistochemistry-unmask-that-antigen/

Renee R Donahue Thank you very much for detailed instruction! I'll try your methodology on my samples next time!

Junjie Shao I never tried 'completely' fresh sample. I'll let my fresh samples to be exposed to pfa overnight.