Hello everyone!

I am new to IF imaging and I was wondering if I could get clarification on this protocol for whole organoid imaging.

Below is the protocol outlined from the paper: Human bone marrow organoids for disease modelling, discovery and validation of therapeutic targets in hematological malignancies

Sections were blocked using 2% Goat Serum (Thermo Fisher Scientific, Cat#31872) 1% Bovine Serum Albumin (BSA) (Sigma, Cat#A9418) prior to primary antibody labelling with antibody diluted in 1% BSA, sequential PBS washes, and finally secondary labelling with AlexaFluor conjugates. Whole organoid blocking solution was further supplemented with Triton X100, Tween, and Sodium deoxycholate as described by Wimmer et al (3).

Sprouting organoids were imaged within hydrogels in 8-well microslides (Ibidi, Cat#80806), whole organoids were labelled in 15mL Falcons before embedding in 0.5% Agarose within 8-well microslides. Whole organoids were subject to serial dehydration (50%, 70%, 90%, 100%) within microslides before clearance with Ethyl Cinnamate and subsequent imaging. Sections were prepared by embedding fixed organoids in Optimal Cutting Temperature compound (OCT, VWR Cat#361603E) before sectioning onto Poly-L-Lysine covered slides. Slides were washed in Acetone before immunofluorescence labelling.

I was wondering how I would go about creating a step-wise protocol for myself for the whole organoid (d18). Would it go something like this?

1. Fixation

2. Blocking and staining

3. Embedding in 0.5% agarose in microslides

4. Dehydration

5. Sectioning