Antigen Retrieval (Optional)

Antigen retrieval is optional, but highly recommended. Cross-linkages formed by PFA fixation may interfere or block antibody binding to the epitope of interest so detection of certain samples will require an additional antigen retrieval step before incubation with primary antibodies.

1. Place slides in a heat-resistant plastic Coplin staining jar and fill with citrate buffer.
2. Place Coplin jar in food steamer and steam for 20 minutes.
3. Carefully remove hot citrate buffer and wash slides 3X in PBS-T for 10 minutes each.

Blocking

1. Wash slides with PBS-T at 37°C for 10 minutes to fully remove gelatin from slides.
2. Lay slides flat and pipette enough blocking solution to completely cover sections. The borders formed by the wax pen will hold blocking solution on the sections of tissue.
3. Incubate at room temperature in a humidified chamber for 1 hour.

Primary Staining

1. Prepare primary antibody solution by adding primary antibodies at the appropriate dilution factor to the primary dilution buffer.
2. Pour blocking solution off of slides and replace with primary antibody solution.
3. Incubate at room temperature in a humidified chamber overnight (16 hours).

Secondary Staining

1. Prepare secondary antibody mixes in PBS-T.
2. Pour primary antibody mix off of slides and place slides in a Coplin staining jar.
3. Wash slides 3X with PBS-T.
4. Lay slides flat and pipette secondary antibody mix onto the slides. Incubate at room temperature in a humidified chamber for 2 hours.

Coverslips

1. Pour off secondary antibody mix and place slides in a Coplin staining jar.
2. Wash slides 3X with PBS-T for 30 minutes each.
3. Air-dry slides for 5 minutes.
4. Add PermaFluor™ to slides and add coverslip. Store at 2 – 8°C and allow to dry before imaging.

Antigen Retrieval (Optional)

Antigen retrieval is only meant for samples fixed in PFA or other cross-linking agents. It is an essential step when staining for some antigens (e.g. keratin 8/18) but is optional for other antigens (e.g. acetylated tubulin). Samples fixed in methanol or other organic solvents should not undergo antigen retrieval. Samples fixed in alcohol or other dehydration-type fixatives should not be boiled.

1. Aspirate PBS and add 1 mL of citrate buffer to the organoids.
2. Set a 1.7 mL-tube heating block to 98°C. Once the heating block reaches 96 – 98°C, place the tube in the heating block and incubate for 20 minutes.
3. Turn off the heating block. Allow the organoids to sit in the heating block for an additional 20 minutes while it cools down.
   Caution the tubes can be hot at this point. If needed, use tweezers to remove the tubes.
4. Proceed to permeabilization and blocking steps.

Cell Permeabilization and Blocking

Permeabilization of cells is only needed for intracellular targets. If assessing the surface markers, there is no need for permeabilization. Methanol-fixed samples may not require permeabilization.

1. Create the permeabilization / blocking solution by adding 5% normal serum (v/v) to the permeabilization solution.
   The animal serum used for blocking should be the same as the host of the secondary antibody. For example, if you are planning on using donkey secondary antibodies, donkey serum should be used for this step. Add fresh serum each time; do not store permeabilization / blocking solution long term with serum.
2. Aspirate the citrate buffer and add 1 mL of permeabilization / blocking solution to the organoids. Place the tube on its side on a tilting platform and incubate at room temperature with agitation for 1 – 72 hours.
   The incubation period may need to be optimized for each tissue, antigen, and antibody. Often, however, the incubation period may range from 1 – 72 hours with little to no difference in background signal.
   If the cells have been fixed in PFA, consider washing the organoids with 0.3 M glycine for 30 minutes before blocking. Glycine will bind to unreacted aldehyde groups to quench PFA autofluorescence and to prevent free aldehydes from reacting with primary or secondary antibodies, thereby causing a high background signal.

Primary Staining

Typical primary antibody dilutions for organoid whole mount are 1:400 – 1:800; titrate as needed per antibody. User optimization for dilutions may be required depending on the antibody. Use as little antibody as possible to mitigate background signal. Antibody incubation periods may need to be optimized for each tissue, antigen, and/or antibody.

1. Aspirate the permeabilization / blocking solution and wash 3 times in immunofluorescence buffer. Allow the organoids to sit for 5 minutes between each wash.
2. Add 0.5 mL – 1 mL of the primary antibody solution. The solution should include primary antibodies appropriately diluted in the immunofluorescence buffer.
   Some organoids should be set aside at this point as an isotype control and/or a secondary only control, especially if the antigen is rare or the antibody is new.
3. Place the tube on its side on a tilting platform and incubate at room temperature for 16 – 72 hours with gentle agitation.

Secondary Staining

The secondary antibody must be raised against the same isotype as the primary. For example, if using mouse IgM as your primary you will need an anti-mouse IgM for your secondary. Typical antibody dilutions for secondary antibodies are between 1:500 and 1:1000. User optimization may be required depending on the antibody.

To minimize background fluorescence or false signals, it is important that the emission spectra of your conjugated-secondary antibodies do not extensively overlap with one another. Consult the manufacturer’s specifications to determine the excitation and emission spectra of your desired fluors.

1. Aspirate the primary antibody solution and wash 3x in immunofluorescence buffer. Allow the organoids to sit for 5 minutes between each wash.
2. Add 0.5 mL – 1 mL of the secondary antibody cocktail. The solution should include secondary antibodies appropriately diluted in the immunofluorescence buffer supplemented with 10% normal serum.
3. Place the tube on its side on a tilting platform and incubate at room temperature for 16 – 72 hours with gentle agitation. Avoid exposing the organoids to light during this incubation.
   At any point after the primary or secondary antibody incubations, organoids can be stored in immunofluorescence buffer without antibodies at 2 – 8°C for 2 days or over the weekend (in the dark, without rocking). For best results, continue with the next steps as soon as possible.

Counterstaining

1. Add DAPI directly to the secondary antibody solution / organoids to a final concentration of 2 – 4 µg/mL. Place the tube on its side and incubate at room temperature with gentle agitation for 15 – 20 minutes in the dark.
   Samples can be washed before the addition of DAPI but is not always necessary.
   If counterstaining with DAPI, use red or far-red secondaries to detect nuclear antigens to prevent DAPI signal spillover into downstream channels (i.e. to avoid a false positive).
2. Aspirate the DAPI and secondary antibody solution. Wash the organoids once in water and once in PBS. Allow the organoids to sit for 5 minutes between each wash.
   At this stage, organoids may be transferred to glass-bottom chamber slides (e.g. 8-well Ibidi® chamber slide) and imaged in PBS without clearing. However, clearing is recommended for best results. Organoids may also be stored for 2 – 3 days in PBS at 2 – 8°C, although image as soon as possible for best results.

Clearing and Mounting

Optical clearing is highly recommended. Not only does the process increase the overall signal-to-noise ratio by reducing light scatter, it also protects fluorophores and dyes from quenching (fading).

In order to reduce light scatter during acquisition, cleared organoids should be imaged in solutions that roughly match the refractive index (RI) of proteins (~1.35-1.6). Prolong™ Gold Antifade Mountant (RI=1.47) is one example of an index-matching solution. Other examples of RI-matching solutions include BABB (2 parts benzyl benzoate with 1 part benzyl alcohol; RI=1.56) and fructose-glycerol (60% glycerol supplemented with 2.5 M fructose; RI=1.469).

Organoids immersed in BABB must be imaged on the same day as clearing. On the other hand, immunolabeled and cleared organoids can be stored in ProLong™ Gold Antifade Mountant at room temperature for as long as 6 months with minimal loss in signal intensity.

1. Aspirate the supernatant and resuspend the organoid pellet in 1 mL of 50% methanol in PBS. Place the tube on its side on a tilting platform and incubate at room temperature with gentle agitation for at least 1 hour in the dark.
2. Remove the 50% methanol solution and add 1 mL of 100% methanol to the organoids. Place the tube on its side on a tilting platform and incubate at room temperature with gentle agitation for at least 1 hour in the dark.
3. Use a 1 mL pipette tip followed by a 200 µL pipette tip to aspirate as much of the methanol solution as possible.
4. Resuspend the organoid pellet in 50 µL ProLong™ Gold Antifade Mountant.
5. Dispense the organoid/mountant suspension onto the centre of a glass microscope slide.
   Mountant is quite viscous. Aspirate and dispense slowly to avoid bubbles. For larger organoids, you may need to cut the 200 µL pipette tip before aspirating.
6. Place a coverslip on top of the organoid/mountant suspension.
   The weight of the coverslip on the organoids will compress organoids in the Z plane. If planning on acquiring Z stacks, resuspend in a larger volume of mountant (e.g. 100 µL) and transfer the organoids into a chamber slide or use imaging spacers o confine organoids without compression.
7. Allow the mountant to cure for at least 24 hours before imaging.
   You may want to use clear nail polish to seal the coverslip onto the slide before placing the slide into long-term storage.

1. Aspirate medium from the well containing kidney organoids. Carefully add 200 µL D-PBS to the well. Remove D-PBS.
2. Fix: Add 85 µL 4% PFA in D-PBS to the well. Incubate at room temperature (15 – 25°C) for 15 minutes. Remove the solution.
3. Wash the well 3X with 200 µL D-PBS. Remove D-PBS.
   Note: For later immunostaining, keep 200 µL D-PBS in the well, wrap plate with Parafilm®, and store at 2 – 8°C. Remove remaining D-PBS before proceeding to Step 4.
4. Permeabilize: Add 200 µL PBS-T to the well. Incubate at room temperature (15 – 25°C) for 15 minutes. Remove PBS-T.
5. Block: Add 200 µL 10% donkey serum in PBS-T to the well. Incubate at room temperature (15 – 25°C) for at least 30 minutes.
6. Primary antibody staining: While incubating, prepare primary antibody solution by diluting each primary antibody in the blocking buffer. Remove blocking buffer from the well and add 80 µL primary antibody solution.
7. Incubate at 2 – 8°C overnight with low shaking.
8. Primary antibody wash: Add 200 µL D-PBS to the well and incubate at room temperature (15 – 25°C) for 5 minutes. Remove D-PBS. Repeat this wash step 5X for a total of 6 washes.
9. Prepare secondary antibody solution by diluting each secondary antibody in blocking buffer with 1 µg/mL DAPI.
10. Secondary antibody staining: Add 80 µL secondary antibody solution to the well.
11. Incubate in the dark at room temperature (15 – 25°C) overnight with low shaking.
12. Secondary antibody wash: Remove secondary antibody solution. Add 200 µL D-PBS to the well and incubate at room temperature (15 – 25°C) for 5 minutes. Remove D-PBS. Repeat this wash step 5X for a total of 6 washes.
13. Add 200 µL D-PBS to stained cells; they are now ready for immunofluorescent imaging.
    Note: If not used immediately for imaging, wrap plate with Parafilm® and store in the dark at 2 – 8°C.

Blocking

1. Gently wipe the bottom of each Transwell® insert with a PBS-soaked Kimwipe™.
2. Wash each insert 3X with PBS for 5 minutes each, adding 200 μL PBS to the apical compartment and 500 μL PBS to the basal compartment. After each wash, use an aspirator to carefully remove the PBS.
3. Incubate the inserts with the blocking buffer at room temperature for 1 hour. Rocking the plates is not necessary, but beneficial when possible.

Primary Antibody Addition

1. Wash each insert 3X with PBS for 5 minutes each.
2. Add primary antibodies of your choice to the primary antibody buffer at the desired dilution. Add 200 μL of this diluted mixture to the apical compartment and 500 μL to the basal compartment.
3. Incubate overnight at 2 – 8°C. Rocking the plates is not necessary, but beneficial when possible.

Secondary Antibody Addition

1. Remove the primary antibody buffer from each insert by aspiration, and then wash them again 3X with PBS, for 5 minutes each.
2. Add relevant secondary antibodies into the secondary antibody buffer at a dilution of 1:1000. Add 200 μL of this diluted mixture to the apical compartment and 500 μL to the basal compartment.
3. Incubate the inserts at room temperature for 2 hours in the dark. After adding the conjugated secondary antibody, avoid exposing the plate to light to prevent any fluorescence quenching.

Mounting and Visualization

1. Wash the inserts 3X with PBS, for 5 minutes each.
2. Add DAPI staining stock solution into PBS at a dilution of 1:1000. Add 200 μL of this diluted mixture to the apical compartment and 500 μL to the basal compartment.
3. Incubate at 2 – 8°C for 10 – 15 minutes.
4. Wash each insert once with PBS for 5 minutes.
5. Aspirate the PBS from the individual inserts, then peel or cut the polyester membrane from the Transwell® hanger (Figure 1). Once separated from the hanger, use surgical scissors to cut away the curved membrane edge. This will allow the stained culture membrane to sit flat under the coverslip during the mounting procedure.
6. Using a plastic Pasteur pipette, add one droplet of liquid mountant to a microscope slide for every stained culture membrane to be mounted.
   Do not exceed three culture membranes to be mounted on a single slide.
   To prevent air bubbles being caught under the coverslip from step 24, add a small volume of liquid mountant in a line to connect the existing droplets of the solution.
7. Using forceps, place one stained culture membrane, cell-side up, on a droplet of liquid mountant. Add extra liquid mountant on top of the membrane, so as to completely cover the membrane’s surface. Carefully place a coverslip on top of the membranes covered in liquid mountant.
8. Allow the slides to cure at room temperature overnight in the dark, sitting horizontally flat.
9. On the following day, visualize and take images of the stained membranes using a confocal microscope at 63X oil immersion.