Dear research community,
me and my colleagues are currently working on immunofluorescence staining of enteroids (ileum, colon). So far, our protocol works fine for targets at the cell borders and tight junctions, but we were not able to get a specific fluorescence for intracellular targets so far.
We already tried different methods for fixation (paraformaldehyde at room temperature or ice-cold with different durations up to over night; a methanol-formaldehyde combination; formalin-only will be tested this week), permeabilization (different concentrations of Triton X-100 in PBS, combinations with BSA and/or serum; no Triton) and blocking (different concentrations of BSA with/without Triton X-100/Serum). None of the named experiments had any affect regarding specific intracellular signals.
For the experiments, the enteroids are directly grown on sterile glass slides in a 12-Well-Plate and processed in the well plate. They are taken out in the final step and mounted on the slide then, so we don't use any cutting methods with freezing or paraffin.
Does someone have experience in intracellular staining of whole mount organoids/enteroids, who could help us with this problem?
Maybe try an antigen retrieval step? We usually do this only with paraffin sections but it may help.
Hello Petra Katebi Kashi
and Skylar Kingthank you for your support!
We partly solved our problem by using Formalin instead of Paraformaldehyde now, but it still doesn't work for every antibody we have (especially phosphorylated ones are difficult). During this process, some of the primary antibodies have been changed (different companies and/or lot numbers), but the problems still appear and we continue to feel like its a technical thing.
Petra Katebi Kashi
how exactly did you work with the parafilm? Could you provide a short description? Where do you see the benefit compared to glas slides?Skylar King As the enteroids are not in paraffin for staining, we can't use the common steps in antigen retrieval. We are working with Triton X-100 for permeabilisation, but if you have an idea on how to perform it with something else, please let me know.
Best wishes
Johanna Hagens
Hi Johanna,
we use (Co et al., 2019 Cell Reports) for organoids embedded in matrigel and for organoids in suspension the same procedure to stain them:
- Enteroids are in 2% paraformaldehyde in 100 mM phosphate buffer (pH 7.4) for 30 minutes at room temperature, then washed 1x with PBS. Enteroids are permeabilized and stained in phosphate-buffered saline with 3% bovine serum albumin, 1% Triton X-100, and 1% saponin.
https://pubmed.ncbi.nlm.nih.gov/30811997/
Doing this we are able to vizualize specific differentiation markers inside the organoid cells and others like ki67. Have you tried your antibodies in tissue as a positive control? It could be that your organoids are not expressing some of those markers you are looking for. Happy to help with the specifics if you need! Good luck!
Try fixing with methanol only. The methanol should help with phosphorylated proteins (or it does for me always). Fix at 4C, or on ice with ice cold Methanol for 20 minutes. Triton X-100 is not needed because the methanol will permeate the cells. Also, I usually dilute my serum in PBS-tween and use a 5% BSA solution for blocking. I also directly add the Triton-X 100 in the fixing solution. Generally, I use 4% PFA and add 0.2% of Triton X-100 directly into the PFA solution. This should also work to add in the formalin as well. How long are you incubating your primary antibody? I have found with large tissues (zebra fish embryos and C. elegans germlines, mouse aortic root) it is better to incubate overnight at 4C, then incubate in your secondary for 30 minutes at RT the following day. Make sure to use fresh blocking solution because if my block is even a few weeks old my results do not look good. I usually keep for a max 2 weeks (you can make a smaller amount to prevent waste, or freeze in aliquots). I also always dilute both my primary and secondary antibodies in block for incubation (some people use PBS-Tween for this step). I also rinse with PBS-T after the initial fixation method then everything after that step is done with block (rinses between primary etc. is all done with block). Hope this helps.
Hello again, I am sorry for my late reply to this, but currently we had some trouble with the organoid cultures.
I will be getting back to my staining protocol again during the next weeks and will try the suggested methods from this feed. Maybe they can fix my intracellular staining problem. I will keep you updated!
Hi All, I am new to colon organoid culturing. I am culturing the organoid successfully. However, When I am trying to fix it with a 4% PFA, It is not melting the matrigel. Did any of you face similar issues? Is there any way out of this?
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