I am trying to stain paraffin-embedded tissue slides using primary + secondary antibodies, and also a phalloidin actin probe. My plan is to deparaffinize the tissue slides (in this case, mouse lung), do a heat-induced epitope recovery, a blocking step, incubation with the primary antibodies overnight, then secondaries... etc. The protocol I follow works well for me in terms of visualizing the structures I stain with the antibodies. I tried staining my slides with the actin probe separately after the secondaries, but I can't see any actin. My question is, should I incorporate the actin probe into my secondary antibody incubation step? Is there anything else I should do to make sure the probe shows up? I've never worked with a phalloidin probe before, only antibodies.
Hi Kathleen,
How well phalloidin works depends in how good your fixation protocol preserved actin structures. Phalloidin specifically decorates actin filaments. Very often, PFA-perfussion of animals isn't fast enough to preserve actin structures in tissues and therefore the use of phalloidin results in weird-looking backgrounds. Probably you'll get better results using antibodies to detect actin in your samples.
I had assumed PFA perfussion. Would you mind to comment the protocol you follow including temperatures?
Cheers,
The tissues I am using are from another institution and were prepared by an undergrad. But, I believe the mice were perfused with formalin, the lungs were harvested, mounted in paraffin blocks, and then sliced and placed on slides.
Hi again,
You can also check in this RG thread on the issue. https://www.researchgate.net/post/Phalloidin_and_DAPI_staining_for_paraffin-embedded_tissue
Maybe dissecting small sections of your tissue and immediately fixing them PFA-Glutaraldehyde buffers may help you preserve actin structures in your samples enough for successful phalloidin staining.
Cheers,
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