Whole mount immunostaining of organoids and embryonic lungs

Wholemount antibody staining of organoids.

Before you start

• Prepare sufficient quantities of ice-cold wash medium.
• Prepare fresh 4% PFA.
   

Things you will need

15ml falcon tubes

Plastic Pasteur pipettes

Pipettes and filter tips (p1000, p200, p20)

Cold Advanced DMEM/F12 (ThermoFisher Scientific 12634010) for washing.

Cold Advanced DMEM/F12 (ThermoFisher Scientific 12634010) containing 10% FBS. 

Corning Cell Recovery Solution (Corning 354243)

Freshly made 4% paraformaldehyde on ice.

PBS

Wash buffer: PBS containing 0.5% BSA and 0.2% Triton X-100

Blocking buffer: PBS containing 1% BSA, 5% normal donkey serum and 0.2% Triton X-100

Appropriate primary and secondary antibodies

Mounting medium e.g. Vectashield (Vector Laboratories H-1000)

Slides

Coverslips

Grace Bio-labs Secure-seal Imaging spacers 20 mm Diameter x 0.12 mm Depth 654006 (Sigma-Aldrich GBL654006)

Wholemount antibody staining of organoids

1. Cut the end of a plastic Pasteur pipette to increase diameter.
2. Coat the Pasteur pipette with medium containing 10% FBS.
3. Add excess cold FBS-containing medium to the well containing the organoids.
4. Remove matrigel containing the organoids with the Pasteur pipette and transfer to a fresh falcon tube.
5. Add 10 ml cold Advanced DMEM/F12 (FBS is not required here).
6. Incubate on ice 10 min. Invert every ~2 mins. (This removes most of the matrigel).
7. Incubate on ice for 5 mins to allow the organoids to sediment.
8. Remove the medium (leave ~500 μl behind).
9. Repeat the washing steps (numbers 5-9) two more times until the matrigel is dissolved.
10. Add 3 ml Corning Cell Recovery Solution (Corning 354243) and incubate on ice 30 mins. Invert every 10 mins.
11. Spin 200 r.c.f.  5 mins, 4°C.
12. Remove corning solution and wash in 10 ml PBS. Spin 200 r.c.f.  5 mins, 4°C.
13. Remove PBS and add 3 ml 4% PFA and incubate 30 mins on ice.
14. Discard PFA by local approved disposal route.
15. Add 10 ml wash buffer and spin 200 rcf, 5 mins, 4°C.
16. Wash again in 10 ml wash buffer
    If necessary store at 4°C in wash buffer up to 72 hours before beginning wholemount staining. 
17. Transfer organoids to a conical-bottomed 96 well plate to begin wholemount antibody staining protocol using a cut P1000 tip which is coated in wash buffer. 
    All subsequent wash steps are performed on a rotating platform and wash medium is removed from the organoids slowly under a dissecting microscope to avoid aspiration of organoids.
18. Perform 3x 5 minute washes at room temperature in wash buffer.
    Do not try to remove all the buffer after each wash to avoid aspirating the organoids.
19. Add blocking buffer and incubate at 1 hour at room temperature.
    Can incubate in block at 4°C overnight.
20. Replace the block with primary antibodies diluted to appropriate concentrations in blocking buffer. Incubate overnight at 4°C.
    Ensure the organoids are fully covered. 100-200 μl primary antibody mix is usually sufficient.
21. The next morning wash 3x briefly in wash buffer.
22. Wash 3x 10-20 minutes at room temperature in wash buffer.
23. Replace wash buffer with secondary antibodies diluted to appropriate concentration in PBS containing 0.2% triton and 5% normal donkey serum. Incubate room temperature for 2-3 hours.
    Can incubate in secondary antibodies at 4°C overnight if convenient.
24. Wash 3x briefly in wash buffer.
25. Wash 3x 10-20 minutes at room temperature in wash buffer.
26. Wash in wash buffer containing DAPI for 30 minutes at room temperature.
27. Wash 3x briefly in wash buffer.
28. Wash 3x 10-20 minutes at room temperature in wash buffer.
29. Attach a Grace BioLabs imaging spacer to the centre of a microscope slide.
30. Using a cut P200 tip slowly suck up the organoids and allow them to drop to the end of the tip. Mount organoids in the centre of the imaging spacer in a small volume of wash buffer. Remove excess wash buffer with a P20 tip.
31. Gently pipette 50 μl of fluorescent mounting medium on top of the organoids and mount a coverslip on top of the imaging spacer.
32. Seal with nail polish and store at 4°C until imaging.

Wholemount antibody staining of embryonic or fetal human lungs

1. Up to 10 pcw fix submerged in 4% PFA for at least 1 hour on an orbital shaker. Beyond 10 pcw separate the lobes, or cut the lobes into smaller pieces, and fix for 2-3 hours. Smaller pieces such as micro-dissected airways should be fixed for 30 minutes. Note: fixation is the most critical step to be optimised and very much depends on the source of the tissue and how it is collected.
2. Discard PFA by local approved disposal route.
3. Rinse 3x 15 minutes in wash buffer at 4 degrees.
   If necessary store at 4°C in wash buffer up to 72 hours before beginning wholemount staining. 
4. Optional permeabilization step for larger samples: 0.5% triton in PBS for 30 mins at room temperature
5. Perform 3x 5 minute washes at room temperature in wash buffer.
6. Add blocking buffer and incubate at 1 hour at room temperature.
   Can incubate in block at 4°C overnight.
7. Replace the block with primary antibodies diluted to appropriate concentrations in blocking buffer. Incubate overnight at 4°C.
   Ensure the lungs are fully covered. 
8. The next morning wash 3x briefly in wash buffer.
9. Wash 3x 10-20 minutes at room temperature in wash buffer.
10. Replace wash buffer with secondary antibodies diluted to appropriate concentration in PBS containing 0.2% triton and 5% normal donkey serum. Incubate room temperature for 2-3 hours.
    Can incubate in secondary antibodies at 4°C overnight if convenient.
11. Wash 3x briefly in wash buffer.
12. Wash 3x 10-20 minutes at room temperature in wash buffer.
13. Wash in wash buffer containing DAPI for 30 minutes at room temperature.
14. Wash 3x briefly in wash buffer.
15. Wash 3x 10-20 minutes at room temperature in wash buffer.
16. Depending on the size of your sample you may need to add a clearing protocol and mount under a raised coverslip e.g. using imaging spacers as above, or plasticene/Vaseline to raise the coverslip.