I am struggling with getting immunofluorescence working on my mouse brain slices. It seems that most antibodies, despite testing various concentrations, are not working out, so I was hoping for a better protocol or some advice.
My samples are mouse brains of various developmental ages (E16-p40). I flash-froze them straight without OCT, so there is no going back there, and I hope I can make these samples work without repeating the sacrifices/sectioning.
10 um frozen sections, thawed for 20 minutes at RT and fixed in 4% PFA at RT for 20 minutes. Samples are permed in .2% Triton-PBS for 1 hour, and blocked in 10% horse serum .05% Triton-PBS for 1 hour, and placed in primary overnight. Wash steps are in .05% Triton-PBS. Secondary is for two hours at RT, washed and mounted in ProLong Gold.
So far I've tried Tau, beta-3-tubulin, calbindin, NeuN, and GFAP with no luck on any front. What I naively thought might have been axons in 568 (where I usually stick Tau or Tuj1) now appear to be autofluorescing vasculature. Dapi works great, so there is actually a brain slice there.
Any advice is appreciated. Do I need antigen unmasking? Are flash-frozen, OCT-less brains unusable?
Usually antibodies are working better on tissues that are just frozen and not chemically modified. So you should be able to work with your frozen brain specimens. The only disadvantage is that the morphology of frozen tissue is usually not as good as the one of paraffin embedded tissue or tissue that is fixated by perfusion.
There might, however, be a lot of different reasons why your immunofluorescence does not work.
Are you sure that your primary antibodies are generally working in immunofluorescence or immunohistochemistry? Most of the commercially available antibodies are made and screened against peptides, meaning that many of them will work quite well in Western Blotting but not in immunohistochemistry or immunofluorescence. Did you already test your antibodies on other tissues that might serve as a positive control?
Are you sure that your secondary antibody ist still working and are you using an appripriate dilution? Maybe you might want to try doing an immunohistochemistry instead of an immunofluorescence for controlling the system. This might also serve as a control for autofluorescence as PFA can cause severe autofluorescence in different tissues.
To test if the PFA-fixation causes to much crosslinking, so that your antibodies do not have free epitopes for binding anymore you could try to use aceton instead (5-10 min. at room temperature). The morphology will not be as good as with PFA as brain is a very fat tissue, but you will avoid crosslinking of the proteins. In case you want to try this, don't use triton for permeabilization (in aceton fixation you don't need permeabilization anyway) and don't use triton in your washing puffer.
Finally you can also try to use a different mount like moviol to test if the mounting medum is quenching your signals.
Thanks for your help. I'll give acetone a try. Some of the primaries I've used in neurons on coverslips, like Tau and Tuj1, and they work amazingly well! I'm using AlexaFluor secondaries at 1:1000, which works great on cells.
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