Using Human Induced Pluripotent Stem Cell-derived Intestinal Organoids to Study and Modify Epithelial Cell Protection Against Salmonella and Other Pathogens

1. Culturing and Passaging of Induced Pluripotent Stem Cells

NOTE: All methods described here use commercially available human cell lines. All tissue culture work detailed below should be done in a Class II laminar flow hood. iPSCs are routinely maintained in stem cell culture medium (see the Table of Materials), as per the manufacturer’s instructions, which allows a weekend-free culture of iPSCs. iPSCs can be adapted from other iPSC culture systems with relative ease.

1. Passage the cells once the colonies cover approximately 80%–90% of the plate surface.
2. Prepare plates for the passage 1 h prior to use by adding vitronectin 10 μg/mL diluted in Dulbecco’s phosphate-buffered saline (DPBS; without calcium [Ca] or magnesium [Mg]) to tissue-culture-treated plates. The volumes for coating are dependent on the plate size and can be found in the manufacturer’s instructions. During this time, warm stem cell culture medium to room temperature (RT).
3. Remove the media from the iPSCs ready for passage and wash the cells 2x with DPBS (without Ca or Mg).
4. Add EDTA solution (see the Table of Materials) to the plates, making sure the entire surface is coated and incubate at RT for 5–8 min. When holes start to appear in the center of the iPSC colonies, aspirate and discard the EDTA solution.
5. Add stem cell culture medium to the wells; dislodge the iPSCs by gently washing media over the plate surface a few times. The iPSCs are passaged as clumps and not as single cells, so make sure that the EDTA solution is not left on for too long. Move any dislodged iPSCs to a 15 mL conical tube.
6. Aspirate the vitronectin from the precoated plates and replace it with stem cell culture medium. Invert the iPSC suspension a number of times to make sure the iPSCs have not settled at the bottom of the conical tube, and add the appropriate volume of suspension to give a 1:10 dilution of cells on a new plate. Split ratios may be adjusted depending on the iPSC growth rate, which may vary between iPSC lines.
7. Rock the plate to disperse the iPSCs over the surface and place it in an incubator at 37 °C/5% CO₂. Feed the iPSCs the day after passaging.

2. Differentiation from iPSCs to the Hindgut

1. On Day 0, split the iPSCs on a 10 cm tissue-culture-treated dish, precoated with vitronectin as described in step 1.2, into 10 mL of stem cell culture medium supplemented with activin A (10 ng/mL) + basic fibroblast growth factor (bFGF; 12 ng/mL). Add the growth factors to the media directly before use; do this in all subsequent steps.
2. On Day 1, change the media (10 mL of stem cell culture medium with activin A [10 ng/mL] + bFGF [12 ng/mL]).
3. On Day 2, begin the differentiation by changing the media to 10 mL of stem cell culture medium supplemented with the following growth factors: activin A (100 ng/mL), bFGF (100 ng/mL), bone morphogenetic protein 4 (BMP-4; 10 ng/mL), phosphoinositol 3-kinase inhibitor LY294002 (10 μM), and GSK3 inhibitor CHIR99021 (3 μM).
4. On Day 3, change the media to 10 mL of stem cell culture medium supplemented with activin A (100 ng/mL), bFGF (100 ng/mL), BMP-4 (10 ng/mL), and LY294002 (10 μM). Endoderm specification induced by this media should result in visible changes to the iPSC colony morphology over the next 24 h.
5. On Day 4, change the media to 10 mL of RPMI/B27 media supplemented with activin A (100 ng/mL) and bFGF (100 ng/mL).
   NOTE: RPMI/B27 media contains 500 mL of RPMI medium with L-glutamine supplement (see Table 1), 10 mL of B27 supplement (50x, serum free), and 5 mL of nonessential amino acids. Optional: add 5 mL of penicillin-streptomycin (10,000 U/mL). Filter-sterilize before use.
6. On Day 5, change the media to 10 mL of RPMI/B-27 media supplemented with activin A (50 ng/mL).
7. On Day 6, to begin patterning the posterior endoderm to the hindgut, change the media to 10 mL of RPMI/B27 media supplemented with CHIR99021 (6 μM) + retinoic acid (3 μM).
8. On Days 7, 8, and 9, repeat step 2.7. During these steps, visible 3-D structures of the hindgut should become apparent, covering the surface of the plate.
9. On Day 10, embed the resulting hindgut in basement membrane matrix (see the Table of Materials).

3. Embedding of the Hindgut in Basement Membrane Matrix

1. Make up iHO base growth media (500 mL of advanced Dulbecco’s modified Eagle’s medium [DMEM]/F12, 10 mL of B27 supplement [50x, serum free], 5 mL of N2 supplement [100x, serum free], 5 mL of 1 M 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), and 5 mL of nonessential amino acids [100x]. Optional: add 5 mL of penicillin-streptomycin [10,000 U/mL]; filter-sterilize before use. See Table 1).
2. Remove the media from the hindgut plate, and wash the plate 1x with DPBS (without Ca or Mg). Add 5 mL of collagenase solution to the plate and incubate at 37 °C for 5 min.
   1. Produce collagenase solution by adding 500 mg of collagenase IV powder to 400 mL of advanced DMEM/F12. Following this, add 100 mL of serum replacement (see the Table of Materials), 5 mL of L-glutamine (200 mM), and 3.5 μL of 2-mercaptoethanol, and swirl the solution to mix. Filter-sterilize the solution once the collagenase powder is fully dissolved.
      NOTE: This can be stored at -20 °C for up to 6 months in smaller aliquots.
3. Inactivate the collagenase by adding 5 mL of iHO base growth media to the plate and scrape off the hindgut cells using a cell scraper, collecting the hindgut suspension in a 15 mL conical tube.
4. Centrifuge at 240 x g for 1 min and pipette off the supernatant.
5. Add 10 mL of media, break up the hindgut into smaller pieces by gently pipetting, and centrifuge again at 95 x g for 1 min.
6. Wash the cells 2x in iHO base growth media by repeating step 3.5. Resuspend the cells in a small volume of base growth medium (~300–500 μL) and add around 100 μL of this solution to 1.5 mL of basement membrane matrix. The matrix must remain on ice during this time as it will begin to solidify rapidly at RT.
7. Set up a 24-well plate on a plate heater at 37 °C and spot out 60 μL into one well of the 24-well plate. Allow it to set briefly and check the density under a microscope.
   1. If required, add more hindgut solution to the basement membrane matrix in increments until the desired concentration is achieved, and spot out the solution into the remaining wells.
8. Incubate at 37 °C for 10 min; then, add 800 μL of iHO base growth media containing growth factors to each well of the 24-well plate at the following concentrations (see Table 1): 500 ng/mL R-spondin-1, 100 ng/mL Noggin, 100 ng/mL epidermal growth factor (EGF), 3 μM CHIR99021, 2.5 μM prostaglandin E2, and 10 μM Y-27632 dihydrochloride monohydrate (ROCK inhibitor).
9. Change the iHO base growth media every 2–3 days, or immediately if the media begins to discolor. After the initial seeding into basement membrane matrix, allow the iHOs to develop for 7 days before splitting them. By day 3–4, distinct spheres should be visible in the culture.
   1. For media change alone, omit Y-27632, as this is only required when splitting/seeding.

4. Maintenance and Passage of iHOs

1. To allow gradual thawing, put out the required volume of basement membrane matrix in a covered ice bucket at 4 °C overnight, 24 h prior to splitting.
2. Remove the media from the iHOs and replace it with 500 μL of cell-lifting solution (see the Table of Materials) per well. Incubate at 4 °C for 40–50 min, at which point the iHOs should be floating in the solution.
   1. Optional: use an in-hood imaging system to select only iHOs with the desired morphology (see the Table of Materials).
3. Gently pipette the iHO/cell lifting solution suspension into 15 mL conical tubes, trying not to break up the iHOs. Allow the IHOs to settle for 3–5 min and remove the supernatant and single cells.
4. Resuspend the iHOs in 5 mL of iHO base growth medium and pipette them gently to wash. Centrifuge at 95 x g for 2 min.
5. Set up a 24-well plate on a plate heater at 37 °C within the hood.
6. Remove the supernatant and resuspend the iHOs in ~300–500 μL of base growth media, Using a P1000 pipette to break up the iHOs into smaller chunks. Note that the force that needs to be applied will vary dependent on the iHO line and maturation state, so start gently, increasing the force if required.
7. Place ~100 μL of the iHOs (the volume is dependent on the density of the solution) into 1.5 mL of basement membrane matrix and pipette briefly to mix.
8. Spot out 1 x 60 μL of basement membrane matrix into one well of the 24-well plate, leave it to solidify for ~30 s, and then check the density under the microscope. If the density is too low, add more iHOs to the matrix.
9. Repeat step 4.8 until the correct density is acquired, and then spot out the rest of the matrix into a 24-well plate.
10. Place it in an incubator at 37 °C for 10 min and, then, overlay it with 800 μL of base growth medium with growth factors, as described in step 3.7.
11. To prepare the iHOs for an invasion assay experiment (outlined below), passage the iHOs 4–5 days prior to the experiment as described in steps 4.1–4.10, but place 120 μL of the matrix iHO solution generated in step 4.7 into 5 mm glass-bottomed microinjection dishes.
    1. Rather than leaving the iHO suspension in a droplet as with routine passaging, spread the droplet over the bottom of the dish to create a thin layer of matrix. Cover it with 2.5 mL of base growth medium plus growth factors.
       NOTE: If antibiotics have been used in culture medium, these must be removed and replaced with nonantibiotic supplemented media for microinjection experiments.

5. Prestimulation of iHOs with rhIL-22

1. Aspirate the media from the iHOs and replace it with fresh base growth media (which must not contain antibiotics) 18 h prior to the invasion assay.
2. Add rhIL-22 to culture media to a final concentration of 100 ng/mL.

6. Microinjection of iHOs and Intracellular Invasion Assays

1. On the day prior to the experiment, set up S. Typhimurium SL1344 culture in 10 mL of Luria-Bertani broth and incubate at 37 °C overnight with shaking.
2. On the day of the experiment, if a microscope with an enclosed heat chamber is available, turn it on and allow the temperature to reach 37 °C prior to starting the assay.
3. Dilute overnight bacterial cultures in DPBS (containing Ca and Mg) to an optical density of 2 at 600 nm (OD₆₀₀) and, then, mix it 1:1 with phenol red.
4. Load the microinjection dish containing iHOs onto the microscope stage, remove the lid, and bring the iHOs into focus, ready for the injection to begin.
5. Turn the injector and the arm control stations on. Ensure the injector is set to a pressure of 600 kPa and an injection time of 0.5 s. If it is not already backed away from the microscope stage, rotate the injection arm to make sure it is.
6. Set up a 6 μm microinjection drill tip by removing the wrapping and the plastic cylinder from the needle. Remove the grip head from the injecting arm.
7. Load the drill tip with 10 μL of the inoculum, gripping the drill tip gently at its blunt end. Place the drill tip into the grip head and reattach it to the microinjection arm.
8. Gently move the arm into position so that the needle is situated 1–2 cm above the microinjection dish. Use the arm control to position the needle tip in the center of the dish and lower it until it is just over the surface of the media.
   1. Program the arm control station to return the needle to this point after all injections.
9. Focus the microscope on the iHOs and select the target to inject. Position the needle just above and to the right of the iHO to be injected and move the needle downward and laterally into the iHO lumen.
10. Press the inject button on the microinjector; the phenol-stained bacterial mixture will emerge from the needle. Inject each iHO 3x. Inject at least 30 iHOs per condition.
    NOTE: Due to heterogeneity in iHO size and structure within a culture, it is necessary to inject a large number of iHOs to control for variation.
11. When all required iHOs are injected, remove the microinjection plate from the stage, replace the lid, and incubate the plate at 37 °C for 90 min.
12. After 90 min, aspirate the growth media and replace it with 3 mL of cell lifting solution; incubate at 4 °C for 45 min.
13. Gently move the iHOs/cell lifting solution to a 15 mL conical tube containing 5 mL of DPBS. Ensure that all injected iHOs have been removed from the plate (rinse the plate with 1 mL of the DPBS if required). Centrifuge at 370 x g for 3 min.
14. Remove the supernatant and resuspend the iHOs in base growth media containing gentamicin at 0.1 mg/mL (add 1 mL of media; then, use a P1000 pipette ~50x to break up the iHOs, and add 4 mL of further media).
15. Incubate at 37 °C for 1 h to kill extracellular bacteria.
16. Centrifuge the iHOs at 370 x g for 3 min and aspirate the supernatant, leaving as little as possible. Wash the iHOs 1x with DPBS and centrifuge again.
    NOTE: This step removes gentamicin, which is important as any remaining gentamicin may kill intracellular bacteria once the cells are lysed.
17. Resuspend the iHOs in 500 μL of lysis buffer (see the Table of Materials) and manually dissociate the organoids by pipetting ~50x. Leave this mixture for 5 min at RT.
18. Serially dilute the resulting solution 10-fold in DPBS to generate 10^-1, 10^-2, and 10^-3 concentrations. Pipette 3 x 20 μL droplets of the neat and diluted solutions onto prewarmed LB agar plates.
19. Incubate overnight at 37 °C and perform colony counting and calculation of the colony-forming units (CFU). Colony counts will reflect the numbers of intracellular bacteria which were released during the cell lysing process.

7. Cell Freezing and Recovery

NOTE: As noted earlier, it is possible to cryogenically preserve iHOs and reconstitute them when desired. The freezing and thawing processes are outlined below.

1. Select the wells of iHOs you wish to freeze. Add cell lifting solution to the wells and incubate for 40–50 min at 4 °C. The iHOs should be floating in the solution.
2. Gently pipette the iHOs into a 15 mL conical tube and allow them to settle. Remove the media and wash the iHOs 1x with base growth media (no growth factors).
3. Centrifuge at 95 x g for 2 min, remove the supernatant, and replace it with an appropriate volume of cell freezing medium (see the Table of Materials; use the medium as per the manufacturer’s instructions), decanting the iHOs into cryogenic vials.
   1. Store the vials in a -80 °C freezer overnight to allow more gradual freezing; then, transfer them to liquid nitrogen storage.
4. To reconstitute the iHOs, rapidly defrost a cryogenic vial at 37 °C using a water/bead bath; then, gently pipette its contents into 10 mL of base growth media (no growth factors). Allow the iHOs to settle and replace the media with ~300 μL of fresh base growth media.
   1. Do NOT manually dissociate iHOs. Add iHOs to basement membrane matrix and plate them out as described in section 3.