Specifically how do i make 3D structure adhered onto the glass surface without losing its structural integrity?
Hi Mohit Jadli
A little more information on your sample to be imaged will help with understanding how to stain it. Size, cell type?
One can stain 3D structures in solution as well and mount onto glass or another embedding media for microscopy.
We usually put thick samples (stained in free floating way) with mounting media and border it with double sided tape whose thickness we know, to a level where the sample will not be compressed when the coverslip is put. This is mainly when we do not want the lens to touch the tissue directly.
If you are alright with the lens touching the sample, embed it it agarose and mount the sample on a glass slide (fixing block with superglue). If you are using lightsheet microscopes there are either suitable sample carriers or the agarose embedding can be done in capillaries of appropriate thickness to be inserted in the window of the microscope.
Hope this helps
Best
Aparajita
Thanks, Dr. Aparajita Lahree . The sample is a breast organoid of around 100um size. How do I stain 'in solution'? Do I need to spin after each and every step of ICC?
Additionally, if I try to fix it beforehand (like in the chamber slide) will it be superior over the free-floating way (as in a free-floating way I might loose a few of organoid in every successive stage)
I appreciate your valuable time.
Best,
Mohit Jadli
We use 100-150 um liver sections and stain then in 48-well plates in. Free-floating sections. Now we do fixation through transcardial perfusion, which in your case is not possible.
Hence, I would recommend that the fixation be done in a well of 24-well plate (one sample per well, submerged with fixative), with mild rocking RT for 2h followed by 4C overnight.
Rinse in PBS in the same plate and then transfer organoids to 48-well plate (one sample per well) to reduce reagent usage.
You can permeabilize, block and do all the staining and rinsing in this plate (48-well). We used 0.3% Triton X-100 at RT for 60 min for permeabilization and blocked in 3%BSA-PBS for 60 min at RT. All incubations are with rocking (mild)
Primary incubation was for 24h in blocking buffer, RT, followed by 5 washes in 0.3% triton-PBS. Secondary staining was for 48h in blocking buffer, folloowed by 5 washes in 0.3% triton-PBS and 3 washes in PBS. All incubations are with mild rocking.
Samples were then mounted for imaging.
Now if your organoid is fragile, so to prevent cell detachment you can perform fixation followed by mild-dip coating in 2-4% agarose PBS solution (45-50 C max to avoid completely cooking the cells). The antibodies can move through these, the cells will remain in the structure.
Hope this helps
Best
Aparajita
Thanks, Aparajita Lahree for your help.
I will try it. Parallelly, I plan to perform 'in-solution' staining, and then I will fix the stained 3D structure onto the slide for ICC.
Best,
MJ
Hi Mohit Jadli ,
Growing organoids embedded in solid-ECM can be tricky. And immunofluorescence protocols are usually lengthy, and require organoid release with the risk of losing the microtissues, and/or modifying their original morphology. We have developed a platform to overcome those challenges, Gri3D https://sunbioscience.ch/products/
With these microwell plates, you can grow organoids of any type in little (or no) presence of ECM, in suspension-like conditions. A pipetting port on the side allows media/buffer exchange without disrupting the organoids. You can do your immunofluorescence protocol directly on the plate (from fixation with PFA to antibody incubation), and image without having to do any transfer steps!
Check out how Gri3D works in the following publication:
Article High-throughput automated organoid culture via stem-cell agg...
If you have any questions, free to contact me, cheers!
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