Whole-mount Immunofluorescence Imaging: A Fixing and Staining Technique to Evaluate Intact Organoids

To perform whole-mount immunofluorescence staining of prostate organoids, first, add 500 microliters of 4% paraformaldehyde in PBS. Incubate the organoids for 2 hours at room temperature with gentle shaking. After washing the pellet as described in the text protocol, add 1 microgram per milliliter of DAPI stain in blocking solution and incubate for 2 hours at room temperature. Protect the sample from light during incubation from this step forward. Following centrifugation of the organoids as before, add primary antibody and blocking solution and incubate overnight at 4 degrees Celsius with gentle shaking.

Pellet the organoids again and wash the pellet with 1 milliliter of PBS for 15 minutes with gentle shaking. Repeat this washing procedure two times. Then, add secondary antibody and blocking solution and incubate overnight at 4 degrees Celsius with gentle shaking. Following incubation, pellet the organoids and wash the pellet for an additional two times. Add 1 milliliter of 30% sucrose in PBS with 1% Triton X-100 to the pelleted organoids. Then, incubate for 2 hours at room temperature with gentle shaking.

After pelleting the organoids again, add 1 milliliter of 45% sucrose in PBS with 1% Triton X-100 and gently shake for 2 hours at room temperature. Then, repeat the procedure except add 1 milliliter of 60% sucrose in PBS with 1% Triton X-100. Pellet the organoids by centrifugation at 800 x g for 3 minutes at room temperature and remove 95% of the supernatant. Observe the pellet under UV light to confirm that it was not lost during removal of the supernatant. The pellet becomes looser as the concentration of sucrose becomes higher. Transfer 10 to 20 microliters of the remaining suspension to a chambered coverslip and proceed to confocal microscopy.