Hi all,
I’m looking for some advice to optimize my experiment. I have a muscle orgnanoid tissue fixed (4% PFA) in a organ-on-chip device that I want to perform whole-mount immunostaining on in combination with confocal imaging. The optical path goes through a 200 micron layer of PDMS (RI unknown but I assume similar to publshed values => around 1.44). The sample can be close to the substrate or could be suspended up to 150 micron above. The tissue itself is likely around 200 micron thick.
Imaging the whole tissue with a 10x objective is not too much of an issue. However I also need to zoom in a bit on some structural features.
Up until now I have used a 40x 0.75 NA ELWD air objective due to the working distance. In this case the light goes form air → PDMS → medium → Sample. I have mounted my sample with ibidi Mounting medium (RI 1.42-1.44) such that the light only changes RI once. However, here I already notice that I cannot image too deep into the tissue.
Would it be better to instead use an aqueous mounting medium and use a WI objective?
My last resort would be to remove the tissue from the device and mount it on a (depressed) glass slide, which would open up the option for the usual oil objective, although given the thickness of the tissue I would not be able to image very deep either.
Thanks for the advice.