Co-Culture of Murine Small Intestine Epithelial Organoids with Innate Lymphoid Cells

All experiments must be completed in accordance and compliance with all relevant regulatory and institutional guidelines for animal use. Ethical approval for the study described in the following article and video was acquired in accordance and compliance with all relevant regulatory and institutional guidelines for animal use.

All mice were culled by cervical dislocation according to the standard ethical procedure, conducted by trained individuals. Before organ and tissue harvesting, slicing of the femoral artery or decapitation was conducted (as appropriate to the protocol in hand) as confirmatory assessments of death. Animals were housed under specific-pathogen-free conditions (unless stated otherwise) at an accredited commercial unit and King's College London animal unit in accordance with the UK Animals (Scientific Procedures) Act 1986 (UK Home Office Project License (PPL:70/7869 to September 2018; P9720273E from September 2018).

1. Establishing murine small intestinal organoids

NOTE: This section of the protocol describes the generation of intestinal organoids from the murine small intestine. Crypts are first isolated from tissue, resuspended in TBEM, and then incubated with media containing EGF, Noggin, and R-Spondin (ENR). Establishing murine small intestinal organoids has also been well-described elsewhere^24,25,27.

1. Place TBEM on ice to thaw (40 µL/well) and put a standard tissue-culture treated (refers to plates with a hydrophilic and negatively charged surface) 24-well plate in a 37 °C incubator to pre-warm.
   NOTE: 500 µL of TBEM will thaw in approximately 2-4 h; do not leave it at room temperature.
2. Prepare ~4 mL or 550 mL per well of ENR medium by adding EGF, Noggin, and R-Spondin to basal medium (Table 1) and place in a 37 °C incubator.
3. Cull a 6-12-week-old animal and dissect out the small intestine using forceps and microdissection scissors. Place it in 15 mL of Phosphate-Buffered Saline (PBS) on ice.
4. Using the stomach and caecum as points of orientation, isolate the desired region of the small intestine (duodenum, jejunum, or ileum). In this protocol, the ileum is isolated.
5. Submerge the intestine in PBS on a 10 cm² Petri dish on ice. Using forceps, remove fat gently, but completely, from the intestine.
6. Cut the tissue longitudinally using microdissection scissors (preferably with rounded tips to avoid perforation). Keeping the intestine submerged in PBS, shake to remove fecal matter, and maintain the mucous side up to track tissue orientation.
7. Transfer tissue to the dry lid of a Petri dish on ice, placing the intestine mucous/villus side up. Holding one end of the intestine with forceps, gently scrape away mucus using the angled edge of a clean glass slide.
8. Fill the plate with ice-cold PBS and rinse the tissue.
9. Pre-coat a 50 mL tube with PBS2 (PBS + 2% FCS; see Table 1) to prevent adhesion of the tissue to the plastic and add 10 mL of ice-cold PBS to this tube. Cut the intestine into small (~2 mm) segments and transfer them into the 50 mL tube pre-coated with PBS2.
10. Using a 25 mL pipette pre-coated with PBS2, pipette the segments up and down 5-10 times to clean the tissue fragments.
11. Allow the segments to settle for 15-30 s, remove and discard the PBS supernatant, and repeat steps 1.10 and 1.11 until the solution is clear (approximately 3-4 times).
12. Allow the segments to settle and discard the PBS supernatant. Add 30 mL of ice-cold crypt isolation buffer (Table 1).
13. Place the tubes on a horizontal roller or rocker at 30-60 rpm (or gentle) for 30 min at 4 °C. Do not incubate at faster speeds, higher temperatures, or longer duration as the crypts will prematurely dislodge and become single cells with lower viability and yield.
    NOTE: From this stage onward, all procedures should be conducted in an aseptic environment using sterile materials and reagents.
14. Allow intestinal segments to settle at the base of the 50 mL tube. Remove crypt isolation buffer without disrupting the settled segments and transfer it into a 50 mL tube on ice.
    NOTE: If the crypt isolation process was too rigorous, crypts may be seen in this fraction. It can therefore be kept as a reserve but will optimally be discarded at the end of the protocol.
15. Add 20 mL of ice-cold PBS to intestinal segments. Using a 25 mL pipette pre-coated with PBS2, pipette intestinal segments up and down 5-8 times.
16. Let the segments settle for 30 s. Pre-coat a 50 mL tube and 25 mL pipette with PBS2. Using this pipette, collect the supernatant (fraction 1) into the pre-coated 50 mL tube and place the tube on ice.
    NOTE: This fraction serves to rinse away the remaining crypt isolation buffer. However, it also serves as an additional quality control step; if pipetting was too vigorous, crypts will be abundant in this rinse fraction, and if it was too gentle, there will be very little debris in this fraction. Optimally, fraction 1 is discarded at the end of the protocol, but it can be used to make organoids if issues arise with fractions 2-4 obtained below.
17. Repeat steps 1.15 and 1.16 but add 10 mL of ice-cold PBS rather than 20 mL and place the supernatant in a fresh 50 mL tube pre-coated with PBS2 (fraction 2).
18. Repeat step 1.17 twice more but pool the resulting supernatant with fraction 2 (into the same 50 mL tube from step 1.17) to obtain fractions 2-4. This single 50 mL tube should contain the dislodged crypts in 30 mL of ice-cold PBS.
19. Remove a 50 µL aliquot from the pooled 2-4 fractions and place it onto a coverslip. Assess for the presence of epithelial crypts using an inverted light microscope.
    NOTE: If crypts are not present, repeat steps 1.17 and 1.18 using more force during pipetting until crypts are released. Ensure that crypts were not prematurely dislodged in the crypt isolation buffer from step 1.14 or in fraction 1 from step 1.16.
20. Pre-coat a 100 µm strainer and 50 mL tube with PBS2. Pass the pooled crypt fractions through this strainer and into the pre-coated 50 mL tube.
21. Spin down strained crypt fractions at 300 x g for 5 min at 4 °C.
22. Discard the supernatant and resuspend the crypts in 10 mL of ice-cold Advanced DMEM/F12. Move the crypts to a 15 mL tube.
23. Spin down crypts at 210 x g for 5 min at 4 °C to remove single cells and lymphocytes.
24. Resuspend crypts in 1 mL of ice-cold Advanced DMEM/F12.
25. Remove a 10 µL aliquot and place it onto a coverslip. Using an inverted light microscope, count the number of crypts to determine crypt concentration.
26. Calculate the volume required for ~400 and ~1,500 crypts. Pre-coat a p1000 tip with PBS2 and use this tip to pipette the required volumes (containing ~400 and ~1,500 crypts) into separate 1.5 mL tubes.
27. Spin down the crypts at 300 x g for 5 min at 4 °C.
28. Remove as much supernatant as possible, and then resuspend the crypts in 80 µL of TBEM placed on ice (pre-cool pipette tips at 4 °C to prevent matrix gelation, which occurs at 12 °C).
29. Remove the 24-well plate, placed in step 1.1, from the incubator. Gently pipette 40 µL of crypts into the center of a well in a slow, circular motion to form a flat but 3D dome structure, or into three separate small domes.
    NOTE: The viability of the organoids is the lowest in the center of the dome where nutrients and gases permeate less effectively³¹; thus, maximizing the surface area exposed to media while maintaining clear 3D structures is critical.
30. Repeat step 1.29 to create a total of two wells with a density of ~200 crypts per well and two more wells with a density of ~750 crypts per well. Directly place the plate in an incubator (37 °C and 5% CO₂) for 15-20 min, taking care to not disrupt the viscous but still liquid matrix domes.
31. Add 550 µL of pre-warmed ENR media per well (from step 1.2) and incubate for 24 h at 37 °C and 5% CO_2. At this stage, it is suggested to supplement ENR medium with 5 µM of Wnt agonist (CHIR 99021) and 5 µM Rho Kinase inhibitor (Y-27632) for the first 24 h of culture post isolation to improve the yield and efficiency of organoid generation of crypts freshly isolated from tissue.
    NOTE: Crypts should close to form rounded structures within 24 h, and crypt buds should appear within 2-4 days (Figure 1A).
32. Replace with fresh ENR medium at the end of the incubation in step 1.31, and then every ~2 days or when the phenol pH indicator in the culture medium turns pale orange but before it turns yellow.

2. Maintenance of murine small intestinal organoids

NOTE: This section of the protocol describes the maintenance and passaging of murine small intestinal organoids. Organoids are first harvested, and then mechanically disrupted using a bent p1000 tip. This process breaks large organoids consisting of numerous crypts into multiple smaller fragments for expansion and releases dead cells that have accumulated in the pseudolumen. Murine small intestinal organoid maintenance has also been well-described elsewhere^24,25,27. All procedures should be conducted in an aseptic environment using sterile materials and reagents. Passage or expand organoids once every 4-5 days, before bursting of the organoids occurs from substantial accumulation of debris in the organoid lumen. Organoids can be passaged at a ratio of 1:2-1:3 depending on organoid density, which will optimally be between 100-200 organoids per well.

1. As in steps 1.1 and 1.2, thaw TBEM on ice (40 µL/well) and prepare ENR medium (550 µL per well). Place the medium and standard tissue culture treated 24-well plate in a 37 °C incubator.
2. Remove the plate containing organoids from the incubator. Discard the media from the well to be passaged.
3. Add 500 µL of ice-cold Advanced DMEM/F12 to the well. Using a p1000 tip (pre-coated with media to avoid organoid sticking to the tip interior), harvest the organoids into a 15 mL tube.
4. Rinse the bottom of the well with 250-300 µL of ice-cold Advanced DMEM/F12 ensuring no organoids remain in the well, and pool into the 15 mL tube containing harvested organoids from step 2.3. Repeat steps 2.3 and 2.4 when passaging multiple wells and pool the organoids from multiple wells into the same 15 mL tube.
5. Spin down organoids at 300 x g for 3 min at 4 °C.
6. Upon centrifugation, ensure that there are four visible fractions: a base fraction with healthy organoids, a central and clear matrix fraction, a matrix fraction containing single and dead cells, and an upper supernatant layer containing single and dead cells. Remove the supernatant, the debris fraction, and as much of the clear matrix fragment as possible without disrupting the pellet of organoids.
7. Gently bend the tip of a p1000 pipette tip (approximately 2-5 mm bend). Pre-coat this tip in PBS2 or Advanced DMEM/F12. Using this tip, pipette the organoids up and down 10-20 times to mechanically disrupt the organoids and the remaining matrix.
8. Spin down at 210 x g for 3 min at 4 °C.
9. As in step 2.6, remove the supernatant, the debris fraction, and as much of the clear matrix fragment as possible without disrupting the pellet of dissociated crypts. If healthy crypts or small organoids have not formed a clear pellet, centrifuge again at 300 x g for 3 min at 4 °C.
10. Calculate the required volume of TBEM (80-120 µL per well of harvested organoids depending on whether a 1:2 or 1:3 passage ratio is used).
11. Resuspend the pellet in the calculated volume of TBEM and apply 40 µL per well to the pre-warmed 24-well plate to form a dome. Directly place the plate in the incubator (37 °C; 5% CO₂) for 15-20 min.
12. Add 550 µL of ENR medium per well and incubate at 37 °C and 5% CO₂. Check cultures for organoid formation after 24 h_. Replace with fresh ENR medium every 2-3 days post passage.

3. Isolation of small intestinal lamina propria innate lymphoid cells

NOTE: This section of the protocol describes the isolation of ILC1 from the murine small intestine lamina propria of the RORγt^GFP reporter mice. This involves epithelial cell removal, tissue digestion, density gradient separation of lymphocytes, and ILC1 isolation via fluorescence-activated cell sorting (FACS). FACS isolation following the gating strategy in Figure 2 requires the extracellular staining mastermix (Table 4), with the additional staining controls described in Table 2 and Table 3 for the machine (Table 2) and gating (Table 3) set up. The RORγt^GFP reporter mice are used to isolate live, pure ILC1 and gate out RORγt^GFP+ ILC3. Tissue processing for isolation of lamina propria lymphocytes has also been well-described elsewhere³².

1. Tissue processing
   NOTE: Tissue from individual biological animal replicates is kept separate in this protocol. These samples should be appropriately labeled and kept on ice whenever possible. All reagents except for digestion enzymes should be allowed to reach room temperature to ensure rapid tissue digestion.
   1. Place TBEM on ice to thaw (40 µL/well). Keep a standard tissue-culture treated 24-well plate in a 37 °C incubator to pre-warm.
   2. Prepare fresh epithelial removal buffer and digestion buffer (see Table 1). Prepare isotonic low viscosity density gradient medium (LVDGM) (90% LVDGM and 10% 10x PBS), 40% isotonic LVDGM, and 80% isotonic LVDGM in neutralizing buffer (Table 1).
   3. Cull a 6-12-week-old animal, dissect out the small intestine using forceps and microdissection scissors, and place the intestine in 15 mL of PBS on ice.
   4. Submerge the intestine in PBS on a 10 cm² Petri dish on ice and use forceps to remove fat gently but completely from the intestine.
   5. Remove Peyer's patches (~5-10; running in a line opposite to the line of fat tissue) using microdissection scissors to deplete lymphoid tissue inducer cells and B-cells.
      NOTE: Peyer's patches are absent in Rag^-/- and other lineage-depleted animals. Unlike air bubbles, Peyer's patches will remain in place if nudged.
   6. Cut the tissue longitudinally using microdissection scissors (preferably with rounded tips to avoid perforation). Keeping the intestine submerged in PBS, shake to remove fecal matter.
   7. Cut the tissue into 2-4 cm length pieces and transfer them into a 50 mL tube.
   8. Add approximately 20-40 mL of ice-cold PBS and shake vigorously for 5-15 s to remove mucous and debris.
   9. Discard the contents of the tube onto a fresh Petri dish and replace intestinal fragments into the same 50 mL tube using forceps.
   10. Repeat steps 3.1.8 and 3.1.9 3-4 times, or until the PBS is clear.
2. Epithelium removal
   1. Place intestinal fragments into a fresh Petri dish on ice. Using forceps, pick up intestinal segments and remove excess liquid by tapping on a dry surface.
   2. Cut the tissue into ~0.75-1 cm pieces and transfer them into a fresh 50 mL tube. Add 5-7 mL of epithelial removal buffer (Table 1). Vortex the tube and ensure that all the intestinal segments are submerged in the buffer.
   3. Incubate the samples at 37 °C with rocking (100-150 rpm) for 12-15 min. Vortex the tube for 20-30 s.
   4. Repeat steps 3.2.1-3.2.3 once more, discarding the cloudy supernatant (fraction contains epithelial and intra-epithelial cells) and replacing it with fresh epithelial removal buffer.
3. Digestion of the tissue
   1. Tip contents onto a fresh Petri dish. Using forceps, pick up the intestinal segments and tap on a dry surface to discard excess liquid. Place the segments into a fresh Petri dish on ice.
   2. Cluster the segments into the center of the Petri dish into a dense mass. Using either two scalpels or sharp scissors, finely shred the tissue until it reaches a viscous consistency that could pass through a p1000 pipette tip.
   3. Using tweezers, place the minced tissue into a clean 50 mL tube. Rinse the Petri dish with 1-2 mL of digestion buffer (Table 1) to collect any remaining tissue and pool it into the 50 mL tube with the shredded tissue.
   4. Add 5-7 mL of digestion buffer to the shredded tissue and ensure that all tissue is collected at the bottom of the tube.
   5. Incubate the samples at 37 °C, with medium rocking (~100-150 rpm) for 15 min. Vortex the tube for 20-30 s every 5 min to aid digestion.
   6. While the samples are incubating, place a 40 µm cell strainer on top of a fresh 50 mL tube for each sample. Coat the filters with 1-2 mL of neutralizing buffer (Table 1).
   7. Vortex the samples for 20-30 s after the incubation has finished.
   8. Filter the partially digested tissue through the coated 40 µm filter into the 50 mL tube containing neutralizing buffer.
   9. Use tweezers to collect undigested tissue from the 40 µm filter and place it back into the 50 mL tube that the digestion was performed in, for the second round of digestion. Remove filters and rinse them with digestion buffer to dislodge any undigested tissue adhering to the filter.
   10. Once as much undigested tissue as possible has been removed from the filter and placed back into the 50 mL tube that digestion was performed in, rinse the filter with 1-2 mL neutralizing buffer over the 50 mL tube containing the filtered supernatant.
   11. Add 20-25 mL of neutralizing buffer to the filtered supernatant and place it on ice to maintain the viability of isolated cells during the second round of tissue digestion.
   12. Add another 5-10 mL of digestion buffer to the undigested tissue and repeat steps 3.3.5-3.3.10, ensuring to filter the digested tissue into the same tube as used in step 3.3.8 (the same filter can also be reused). Rinse the filter with neutralizing buffer until the 50 mL line is reached, and then discard any remaining undigested tissue.
4. Lymphocyte isolation by density gradient
   1. Spin the collected supernatants for each biological replicate at 500 x g for 10 min.
   2. During step 3.4.1, add 5 mL of 80% isotonic LVDGM to a 15 mL tube for each biological replicate.
   3. Following centrifugation, discard the supernatant from the filtered, digested tissue. Resuspend the pellet in 10 mL of 40% isotonic LVDGM, ensuring that the pellet is well homogenized, and no large chunks remain.
   4. Setting the pipette aid to its slowest setting, tilt the 15 mL tube containing 80% isotonic LVDGM, and very gently overlay the 10 mL of cell suspension in 40% isotonic LVDGM ensuring that the cell suspension does not mix with the 80% fraction.
      NOTE: If the fractions should ever accidentally mix, rapidly distribute lymphocytes that reached the 80% fraction between two 50 mL tubes filled with PBS, and centrifuge for 10 min at 500 x g. Then, discard the supernatant, and repeat steps 3.4.3-3.4.4.
   5. Spin the tubes at 900 x g and 20 °C, with the acceleration and deacceleration set to the lowest setting to ensure that the fractions are not disrupted. Centrifuge for 20 min (not including break time).
      NOTE: All the procedures from this step onward should be conducted in an aseptic tissue culture hood using sterile materials and reagents.
   6. Pre-coat a 50 mL tube for each sample with PBS2 and add 45 mL of ice-cold PBS.
   7. After centrifugation, gently remove the upper debris layer from the 15 mL tube. Coat a p1000 tip with PBS2 and use it to collect the lymphocyte-laden interphase between the 40%-80% gradients into the 50 mL tube containing 45 mL of ice-cold PBS.
   8. Spin the tubes at 300 x g for 5 min at 4 °C.
   9. Discard the supernatant and resuspend the pellet in 500 µL of FACS buffer (PBS, 2% FCS, 0.5 mM EDTA, and 10 mM HEPES; see Table 1). Pre-coat a 40 µm filter with FACS buffer and use it to filter the cell suspension into a flow tube. Rinse the 50 mL tube with an additional 500 µL of FACS buffer and filter into the same flow tube.
   10. Remove a 10 µL aliquot from the resulting 1 mL of cell suspension. Count the lymphocyte yield using a cell counter or hemocytometer and calculate the cell concentration for each sample.
5. Sample preparation for sorting – extracellular staining
   NOTE: The antibodies used in the extracellular staining mastermix, as well as the reagents to prepare the fixable viability dye and Fc block solution, are used in concentrations enough for up to 5 x 10⁶ cells per 100 µL. Volumes should be adjusted accordingly. For FACS machine compensation to sort ILC1, single-color controls for each of the fluorophores in the extracellular staining mastermix are required. For the antibodies, compensation beads that contain a positive antibody binding bead population and a negative antibody non-binding bead population can be used. For the ultraviolet (UV) fixable viability dye single color control, compensation beads that contain amine-reactive (for positive signal) and non-reactive (for negative signal) populations can be used. It is not recommended to use cells from the RORγt^GFP reporter mice for the UV fixable viability dye single color control, as these cells will contain a GFP⁺ signal.
   1. Remove a small aliquot of ~10 µL from each sample and pool it into a separate flow tube containing 250 µL of FACS buffer. Place the tube on ice. This will be used for the unstained control.
   2. Add 1 mL of PBS to the remaining volume in the sample tubes. Centrifuge at 300-400 x g for 3-5 min.
   3. Prepare 200 µL of UV fixable viability dye solution per sample. Dilute UV fixable viability dye (resuspended in DMSO following the manufacturer's instructions) 1:500 in PBS (or following the manufacturer's instructions).
   4. Discard the supernatant and resuspend the samples in 200 µL of UV fixable viability dye solution. Vortex the tubes and incubate in the dark at 4 °C for 10-15 min.
   5. Add 2 mL of FACS buffer to the tubes to quench the UV fixable viability dye, vortex for 10 s, and centrifuge the tubes at 300-400 x g for 3-5 min at 4 °C. Discard the supernatant from the centrifuged tubes.
   6. Prepare 200 µL of Fc block solution per sample (0.25 mg/mL of Fc block in the FACS buffer).
   7. Add 200 µL of Fc block solution to each of the samples from step 3.5.5, vortex for 10 s, and incubate in the dark at 4 °C for 10 min.
   8. Add 500 µL of FACS buffer to a fresh flow tube. Remove a 2-5 µL aliquot from each sample and pool it into the tube for the fluorescence minus one (FMO) controls.
   9. Vortex the FMO tube for 10 s and distribute evenly (100 µL per tube) into fresh flow tubes labeled for each of the FMO controls (Lineage cocktail FMO, CD127 FMO, KLRG1 FMO, NKp46 FMO, and NK1.1 FMO; see Table 3). Add all the antibodies except the antibody of interest to each tube (as described in Table 3) and vortex for 10 s. Place the FMO controls aside in the dark at 4 °C.
   10. Centrifuge the samples (not FMO controls) at 300-400 x g for 3-5 min at 4 °C.
   11. Prepare extracellular staining mastermix (200 µL per sample) with the final antibody dilutions as described in Table 4. Adjust the staining volume for the appropriate cell concentration (up to 5 x 10⁶ cells per 100 µL) depending on the cell counts from step 3.4.10.
   12. Add extracellular staining mastermix to the samples, vortex for 10 s, and place them aside in the dark at 4 °C.
   13. Prepare a fresh single color control for each antibody intended to be used in the gating strategy (Figure 2). Vortex antibody compensation beads for 20 s, add 1 drop of beads to a flow tube, stain with 0.5 µL of antibody (as shown in Table 2, or following the manufacturer's instructions), and vortex for 10 s.
   14. Prepare a UV fixable viability dye single color control using an amine-reactive compensation bead kit. Vortex amine-reactive compensation beads for 20 s, add 1 µL of Live/Dead UV dye (as shown in Table 2, or following the manufacturer's instructions) and vortex for 10 s.
   15. Incubate the samples, FMOs, and single color controls in the dark for 30 min at 4 °C.
   16. During this time, prepare tubes to collect the sorted cells. Add 400-500 µL of 10% FBS in Advanced DMEM/F12 to 1.5 mL tubes for each sample.
   17. After the incubation, add 2 mL of PBS2 to each of the samples, FMO controls, and single color controls.
   18. Centrifuge the samples, FMO controls, and single color controls at 300-400 x g for 3-5 min at 4 °C.
   19. Discard the supernatant from all the centrifuged tubes. Resuspend the samples and FMO controls in 250 µL of FACS buffer and vortex for 10 s.
   20. Resuspend the single color controls in 250 µL of PBS2. Add 1 drop of amine non-reactive beads to UV fixable viability dye single color control only. Vortex all controls for 10 s.
   21. The cells are now ready to be sorted. Keep samples, FMO controls, and single color controls on ice in the dark whenever possible to improve cell viability and prevent loss of signal from photobleaching.

4. Co-culture of small intestinal organoids with innate lymphoid cells

NOTE: This section describes the co-culture of sorted murine small intestinal ILC1 (isolated following the protocol in section 3) with murine small intestinal organoids (described in sections 1 and 2). Organoids should optimally be used 1-2 days following passage. Co-culture involves harvesting the organoids, adding the appropriate number of ILC1, centrifuging to pellet organoids and ILC1 together, and resuspending in TBEM. Complete this section as soon as possible once ILC1 have been isolated. All the procedures should be conducted in an aseptic environment using sterile materials and reagents.

1. Ensure that TBEM from step 3.1.1 has been thawed.
2. Harvest 1-2-day old organoids as described in steps 2.3 and 2.4.
3. Resuspend the organoid pellet in 1 mL of cold ice-cold Advanced DMEM/F12. Do not bend the p1000 tip as done when passaging the organoids, as the intention here is to maintain the organoid structure for co-culture. If required, place the organoids in separate PBS2 coated 1.5 mL tubes on ice for a short period to be distributed between different co-culture conditions.
   NOTE: Only begin preparation of organoids once ILC samples are ready for co-culture; work on ice and rapidly as soon as organoids have been harvested to minimize epithelial cell death.
4. Pre-coat one 1.5 mL tube with PBS2 per ILC replicate sample. Distribute ~100-200 organoids (approximately 1 well of a 24-well plate) into each tube.
5. Pre-coat a p1000 tip in PBS2 and use it to assess the volume of ILC1s after FACS purification (250-300 µL + sorted volume). Determine the concentration of ILC1 per mL using the number of cells recorded from the sort and determine the required volume.
6. Using the same PBS2 coated p1000 tip, add no fewer than 500 ILC1s to the PBS2 coated 1.5 mL tube containing the organoids. If the number of ILC1s yielded from the sort is less than 500, add the organoid directly to the tube containing the original sort to minimize the loss of cells from the transfer.
7. Spin down ILC1 and organoids at 300 x g for 5 min at 4 °C. If possible, avoid small diameter tabletop centrifuges, as these will pool the cells along the edge of the tube interior as opposed to creating a pellet at the tip of the tube. Since cell numbers are very low, it is imperative to minimize the loss of cells during these steps.
8. Slowly and gently remove as much of the supernatant as possible without disturbing the pellet (which may not be visible by eye, especially in no-organoid ILC-only controls). Place the sample on ice.
9. Resuspend the cold pellet in 30 µL per well of TBEM. While holding the tube on a cold surface (e.g., a small box of ice placed in the tissue culture hood), mix the organoids and ILC at least 10-15 times to ensure even distribution. Triturate frequently but gently, to avoid damaging the organoids and the formation of bubbles.
10. Apply 30 µL per well of ILC-organoids in TBEM to a pre-warmed 24- or 48-well plate to form a single dome. Directly place the plate in the incubator (37 °C and 5% CO₂) for 10-20 min.
    NOTE: It is strongly recommended that ILC-only and organoid-only controls are set up for downstream analysis. ILC1 are rare, but GFP^– ILC2 can be substituted for immunofluorescence or FSC/SSC flow cytometry controls where scientifically appropriate.
11. Add 550 µL (per well) of complete ILC1 medium (ENR + IL-2 + IL-7 + IL-15 + 2-Mercaptoethanol; see Table 1) with any desired experimental cytokines or blocking antibodies and incubate at 37 °C and 5% CO₂ for 24 h.
12. After 24 h, gently remove the plate from the incubator and allow the plate to sit in a tissue culture hood for 1 min to ensure the lymphocytes are settled.
13. Remove 200-250 µL of media and place it into an empty well of a 24-well plate. Check this supernatant with an inverted microscope to ensure that no lymphocytes were removed.
    1. If the supernatant is clear, add 300 µL of fresh ILC1 medium to the co-culture in the original well. If lymphocytes are present in the supernatant, centrifuge at 300-400 x g for 3-5 min at 4 °C and resuspend in 300 µL of fresh ILC1 medium and add this to the remaining 200-250 µL of media in the original well. Ensure to replenish media every 1-2 days or when media becomes pale-orange/yellow.
       NOTE: Do not allow media to evaporate sufficiently that the tip of the matrix dome breaks the surface of the media. Always ensure that matrix is completely submerged. Excess evaporation can be avoided by using central wells in the tissue culture plates and adding ~600 µL of PBS to the surrounding wells. Co-cultures with adult ILC are stable for 1-4 days, after which point the organoids that were seeded without major disruption will rupture and reseed as new crypts. If analyzing the epithelium, it is recommended that cultures are analyzed within 1-4 days of establishing co-cultures.
    2. Perform downstream analysis using immunofluorescence, flow cytometry, or FACS purification of target populations into lysis buffer for gene expression analysis by single cell or bulk RNA-seq or RT-qPCR, as described in reference⁸.