Whole mount IF staining

[Amritha Varshini]

Hi TooT team!

We are working on the whole mount IFs currently and we had a couple of questions.

1. At the end of the dehydration steps, do the samples stay in ethyl cinnamate until imaging? How long are they stable for, and how would we store them for imaging if we can't do it on the same day?

2. The first 3 dehydration steps specify pH9 for the ethanol. Do you adjust the pH using NaOH for this? How important is the pH for these steps?

Thanks!

Amritha

[Abdullah Khan]

Hi Amritha, 

Sorry replies a bit longer than usual particularly busy time. 

1. Yes the samples need to be completely dehydrated and then kept in ethyl cinnamate. Any hydration will mess up the clearance and you'll get precipitation and opaque nonsense. 

2. yes! Adjust with NaOH. The pH has been shown to be optimal for mEGFP stability at 9 - though to be honest how much it affects the function and stability of organic fluorophores... I don't know. I wouldn't risk it in the first instance in case you need to troubleshoot. 

I think this is the original paper: https://journals.biologists.com/dev/article/146/3/dev166884/48939/Broad-applicability-of-a-streamlined-ethyl 

Hope that helps! Thanks for TOOTing ;) 

Abs

[Amritha Varshini]

Thanks! We did end up adjusting the pH, so I'm glad to hear that! 

We did remove all the PBS before embedding in agarose and then added ethyl cinnamate. We found that the clearing process was not very efficient and/or the antibodies just don't permeate deep enough into the organoid since it is so dense (please see attached for an example of what we saw). Do you have any tips/tricks for this? I did overnight incubation for both primary and secondary as suggested, but maybe we will need to try longer times to get deeper into the organoid?

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Amritha Varshini

[Abdullah Khan]

Hi Amritha, 

Thanks for sharing the image. To be honest, there seems to be a labelling problem as the fluorescenec looks very grainy and punctate. Even if the clearing was inefficient, you should get a few tens of microns' worth of clear imaging at the very least from the bottom of the sample upwards. 

My thoughts/questions: 

1. How did you fix these? 

2.. Did you use the blocking buffer with detergents in ? (Triton etc) 

3. What concentration were your antibodies at ? 

4. When you cleared, did you visibly see the organoids turn translucent to some degree? 

5. Also temperature of primary/secondary incubation + was it on a rocker? 

Thanks! Sorry for the million and one quesitons. 

Abs

PS Also have you had these sectioned and stained yet? Curious as to how they look. 

[Karolin Stumpf]

Hi all,

we experienced similar issues using ECI clearing, testing both with and without agarose embedding. While seeing them turn translucent in ECI at RT quite fast, we do not seem to get antibody labeling deep into the organoids (primary antibodies 1:100, secondaries at 1:1000, each ON at 4 degrees), see attached pictures. We tried increasing dehydration times from 15 min per step to an hour, however with no change except damage to organoid structure.

Do you have any other idea why this might be the case?

thanks in advance

Karo:)

[Abdullah Khan]

Hey Karo 

What microscope are you using? 

If you're getting rapid ECI clearing at RT and they're visibly clear the problem is not the clearing. 

Your problem here is more likely the depth you're getting in your imaging set up (e.g. working distance of your objective). It might be that your antibody labelling steps need to be longer, but my feeling looking at your image is that's more likely a microscope thing! Are you using a confocal? 

Abs