Functional Characterization and Visualization of Esophageal Fibroblasts Using Organoid Co-Cultures

Animal experiments for this study were approved by Stockholms Norra djurförsöksetiska nämnd (ethical permit no 14051-2019). Animals were housed in pathogen-free conditions according to the recommendations of the Federation of European Laboratory Animal Science Association.

1. Preparation

1. Thaw the enzymatic stock solutions used for dissociation (see Table of Materials) on ice. Thaw an aliquot of growth factor reduced (GFR) basement membrane matrix (matrix) at 4 °C.
2. Prepare the culture medium. Use the medium described in Table 1 for organoid cultures and co-cultures and prepare it before starting the protocol.

2. Dissection and separation of the esophageal epithelium and stroma

NOTE: Ensure that all instruments used for dissection and tissue processing are sterile. Prepare 2 mL of dissociation solution (see Table of Materials) in Hanks' balanced salt solution (HBSS) per three esophagi.

1. Use a mouse strain of choice, such as C57BL/6J mice. Esophageal development in mice finishes after post-natal day (p) 70, so it is recommended to use mice that are older than p70¹³. Use four or five mice as they provide sufficient material to establish 8-10 organoid co-cultures.
   NOTE: Organoid-forming efficiency decreases with the age of the mice. If specific fibroblast subpopulations are of interest, it is possible that the low fibroblast yield is limiting the number of organoid co-cultures that can be established.
2. Use genetically modified (e.g., Pdgfrα^H2BeGFP) or wild-type (WT) mice for the isolation of fibroblast populations. When using WT mice, perform antibody staining to isolate specific fibroblast subpopulations from the stroma via FACS.
3. Euthanize the mice by CO₂ asphyxiation. Dissect the esophagus using forceps and dissection scissors. To remove the whole esophagus, cut the distal end of the esophagus directly above the stomach and the proximal end at the beginning of the trachea. Submerge the esophagus in PBS and put it on ice.
4. Mechanically remove the muscularis externa using a dissection microscope (total magnification range of 8x-40x) and forceps. Hold the distal end of the dissected esophagus using one pair of forceps and use the other forceps to grab and pull the muscle from the distal to the proximal end of the dissected esophagus. Remove and discard the muscle layer.
5. Open the esophagus longitudinally. This works best using microdissection spring scissors with a ball tip to prevent tissue damage. Hold one end of the esophagus in order to insert the ball of the spring scissors into the lumen of the esophagus and cut open the esophagus while holding on to the end.
6. Place the esophagus in a 1.5 mL micro-centrifuge tube or a 24-well plate. Incubate the opened esophagus in 0.5 mg/mL thermolysin in HBSS at 37 °C on a rocker-shaker for 15 min. Submerge the esophagus completely in thermolysin solution.
   NOTE: The volume of thermolysin solution used depends on the well or tube-size. Several esophagi can be placed and submerged in the same well or tube.
7. Take out the esophagus from the thermolysin solution. Use a dissection microscope to carefully separate the esophageal epithelium from the stroma. Using fine forceps, grab both the epithelial side and the stroma side of the tissue and slowly pull them apart to separate the two layers.
   NOTE: The stroma is identified by its white and opaque appearance, in contrast to the transparent epithelium. The stroma contains the lamina propria and submucosal layer.
8. Transfer the epithelial and stromal layer to two separate 1.5 mL micro-centrifuge tubes with 200 µL of dissociation solution in HBSS. Put on ice.

3. Isolation of esophageal progenitor cells

NOTE: The isolation of esophageal progenitor cells (step 2) and fibroblasts (step 3) can be performed simultaneously. Prepare a 50 mL tube of 1% FBS in HBSS (1% FBS).

1. Transfer the separated esophageal epithelium from the 1.5 mL micro-centrifuge tube (step 2.8) to a clean Petri dish and use a sharp scalpel to mince. Collect the minced tissue from the Petri dish with 200 µL of dissociation solution and transfer it back to the 1.5 mL micro-centrifuge tube.
   NOTE: The epithelium is minced properly when the pieces can be placed back into the 1.5 mL micro-centrifuge tube using a 200 µL pipette tip.
2. Add 800 µL of fresh dissociation solution to the 1.5 mL micro-centrifuge tube to a total volume of 1 mL.
   NOTE: It is important to add a sufficient volume of dissociation solution. Per one to three esophagi, 1 mL of solution is recommended. Scale up when more esophagi are processed at once.
3. Place the tube with the minced epithelial layer on a rocker-shaker at 37 °C for 60 min. Pipette the solution up and down approximately 20 times using a 200 µL pipette tip every 15 min to enhance digestion.
4. After 60 min, pipette up and down another 20 times using a 200 µL pipette tip. Pass the solution through a 40 µm cell strainer into a new 1.5 mL micro-centrifuge tube.Centrifuge at 300 x g for 10 min at 4 °C.
   NOTE: Wet the cell strainer with 1% FBS before straining the cells to minimize adherence of the cells to the filter. The epithelium will not be completely digested, and pieces of tissue will still be visible. Longer incubation will, however, decrease cell survival and not result in a higher yield of viable cells.
5. Discard the supernatant by removing excess liquid with a 1 mL pipette and resuspend the pellet in 1 mL of 1% FBS. Centrifuge at 300 x g for 10 min at 4 °C.
   1. During centrifugation, prepare the conjugated antibody mix for FACS of esophageal progenitor cells.
   2. Mix 1 μL of CD324-PE-Cy7 (ECADHERIN) and CD104-A647 (INTEGRIN-β4) in 200 μL of 1% FBS per one million cells.
      NOTE: 1 µL of antibody (200 µL final volume) is sufficient for one or two esophagi. When processing more esophagi at once, increase the volume of the antibody staining solution.
6. Resuspend the pellet in 200 µL of antibody mix and transfer to a flow cytometry tube. After adding fluorescent antibodies, keep the cell suspensions in the dark to avoid bleaching of signal. Incubate the cells for 30 min at 4 °C. Add 3 mL of 1% FBS and centrifuge at 300 x g for 10 min at 4 °C, and then resuspend the cells in a minimum of 200 µL of 1% of FBS.
   NOTE: 500 µL of 1% FBS is used for up to four or five esophagi. Increase the volume when more esophagi are processed at once, so a FACS flow rate of 100-300 events/s can be achieved. More events/s will decrease sorting efficiency, and an increase in the flow rate will decrease cell survival.
7. Add dead cell stain marker to a final concentration of 1:10,000, 5 minutes prior to FACS sorting to isolate live cells. Sort the progenitor cells using a FACS machine (see Figure 1 for gating strategy). Collect the cells in 1.5 mL micro-centrifuge tubes filled with 200 µL of basic organoid medium (Table 1).

4. Isolation of fibroblasts from the stromal layer

1. Cut the stomal layer in fine pieces in a 1.5 mL tube containing 200 µL of dissociation solution (step 2.8) using dissection scissors. The tissue is properly minced once the solution can be pipetted up and down using a 200 µL pipette tip.
2. Add 800 µL of fresh dissociation solution to the 1.5 mL micro-centrifuge tube to a total volume of 1 mL.
   NOTE: It is important to add a sufficient volume of dissociation solution. Per one to three esophagi, 1 mL of solution is recommended. Scale up when more esophagi are processed at once.
3. Place the tube on a rocker-shaker at 37 °C for 30 min. After 15 min, pipette the solution up and down approximately 20 times using a 200 µL pipette tip to enhance digestion.
4. After 30 min of enzymatic digestion, pipette up and down another 20 times using a 200 µL pipette tip. Strain the solution through a 70 µm cell strainer into a new 1.5 mL micro-centrifuge tube. Centrifuge at 300 x g for 10 min at 4 °C.
   NOTE: Wet the cell strainer with 1% FBS before straining the cells to minimize adherence of cells to the filter.
5. Discard the supernatant by removing excess liquid with a 1 mL pipette and resuspend the pellet in 1 mL of 1% FBS. Centrifuge at 300 x g for 10 min at 4 °C.
   NOTE: When using a genetically modified mouse strain containing flourescent-labeled fibroblasts antibody stainings are optional. If antibody stainings are not required, continue with step 3.7 and transfer the sample to a flow cytometry tube.
   1. During centrifugation, prepare the conjugated antibody mix for the FACS isolation of fibroblasts. Mix 1 µL of CD26-APC (DPP4) in 200 µL of 1% FBS per one million cells.
      NOTE: 1 µL of antibody is sufficient for one or two esophagi. When processing more esophagi at once, increase the volume of the antibody staining solution.
6. Resuspend the pellet in 200 µL of conjugated antibody mix in 1% FBS and transfer to a flow cytometry tube. Incubate the cells for 30 min at 4 °C.
7. Add 3 mL of 1% FBS and centrifuge at 300 x g for 10 min at 4 °C. Resuspend the cells in a minimum of 200 µL of 1% of FBS.
   NOTE: 500 µL of 1% FBS is used for up to four or five esophagi. Increase the volume when more esophagi are processed at once, so a flow rate of 100-300 events/s can be achieved. More events/s will decrease sorting efficiency, and increasing the flow rate will decrease cell survival. After adding fluorescent antibodies, cell suspensions should be kept dark to avoid bleaching of signal.
8. Add dead cell stain marker to a final concentration of 1:10,000, 5 minutes prior to FACS sorting to isolate live cells. Sort the cells using a FACS machine (see Figure 1 for gating strategy). Collect the cells in 1.5 mL micro-centrifuge tubes filled with 200 µL of basic organoid medium (Table 1).

5. Establishment and culture of esophageal organoids

NOTE: Prewarm ER_low (organoid co-culture), ENR (organoid) medium (see Table 1 for description), and a 48-well plate at 37 °C. Place the thawed matrix (prepared in step 1) aliquot on ice. It is recommended to use the matrix provided here (see Table of Materials) for mouse esophageal organoid culture, since other brands of matrix negatively affect organoid forming efficiency.

1. For the organoid co-culture, mix the sorted epithelial cells and fibroblasts at a ratio of 1:2 in a tube. For each matrix dome, use 5,000 epithelial cells and 10,000 fibroblasts. When preparing for more domes, add a multiple of 5,000 epithelial cells and 10,000 fibroblasts to one tube. For organoid cultures without fibroblasts, use 5,000 epithelial cells per matrix dome.
2. Centrifuge the mixed cell population at 300 x g for 5 min. Discard the supernatant by carefully removing it with a 200 µL pipette.
3. Wash the cells once by resuspending the pellet in cold basic organoid medium and centrifuge at 300 x g for 5 min. Place the cells on ice.
4. Prepare a mix consisting of 80% matrix and 20% cold basic organoid medium. Place everything on ice as matrix solidifies at room temperature (RT).
5. Discard the supernatant after centrifugation by carefully removing it with a 200 µL pipette. Resuspend the cells in 10 µL of matrix mix/dome and put back on ice.
6. Take the prewarmed 48-well plate from the 37 °C incubator and make one matrix dome per well using a 20 µL pipette. Each dome contains 10 µL of 80% matrix, 5,000 epithelial cell,s and 10,000 fibroblasts. Transfer the plate upside down to an incubator and let the domes solidify for another 20-30 min at 37 °C.
   NOTE: A decrease in matrix dome volume and/or an increase in cell number will affect the organoid forming efficiency.
7. Add 200 µL of prewarmed ER_low medium (Table 1) to the matrix domes containing organoid co-cultures and ENR medium (Table 1) to the respective matrix domes containing only epithelial organoids and place the plate in the incubator.
8. Grow the organoids at 37 °C and 5% CO₂. For the first 2 days, supplement the medium with 10 µM Rock-inhibitor (Y-27623). Rock inhibitor prevents stress induced cell death and increases the chance of a successful establishment of organoid cultures.
9. Refresh the medium every 2-3 days. Make sure that the medium is warm to prevent the dissociation of the temperature sensitive matrix.
10. Perform analyses of the experiment 6 to 8 days after plating. The organoids can be kept in culture for up to 14 days. Around day 14, loss of dome integrity is observed.

6. Passaging of organoids

NOTE: Passaging of organoids grown in co-culture results in the loss of fibroblasts. Therefore, it is recommended to use ENR medium for all organoids when passaged. Prewarm ENR, PBS, and a 48-well plate at 37 °C.

1. Remove medium and wash the well containing the matrix dome with pre-warmed PBS. Add 200 µL of cold 0.25% trypsin solution and pipette up and down to break the matrix dome.
   NOTE: The use of cold 0.25% trypsin is recommended, as the matrix is temperature sensitive and this aids in breaking down the matrix domes.
2. Incubate the organoids with trypsin at 37 °C for ~20 min. Pipette up and down after 10 min to increase the dissociation of the organoids. Monitor the dissociation process with a microscope every 5-10 min. Since trypsin decreases cell viability, this can help to identify the ideal dissociation time.
3. After 20 min, pipette the organoids up and down with a 200 µL pipette to dissociate the organoids. Collect the cells in a 1.5 mL micro-centrifuge tube, add 1 mL of basic organoid medium, and centrifuge at 300 x g for 5 min at RT.
4. Optional: To ensure best organoid growth conditions and that new organoids are derived from single cells, strain the cells through a pre-wetted 40 µm cell strainer. Filtering the cell suspension results in the removal of organoid cores and cell clumps that are hard to dissociate.
5. Prepare a matrix mix consisting of 80% matrix and 20% cold basic organoid medium. Place everything on ice, as the matrix solidifies at RT.
6. Discard the supernatant by carefully removing it with a 200 µL pipette, resuspend the cells in 10 µL of matrix mix/dome, and put the mixture back on ice.
   NOTE: Organoids can be split at a 1:5 to 1:10 ratio, depending on dome density. Epithelial cells can also be counted and re-plated at 5,000 cells/dome.
7. Take the prewarmed 48-well plate from the 37 °C incubator and make one dome per well. Transfer the plate upside down to an incubator and let the domes solidify for another 20-30 min at 37 °C.
8. Add 200 µL of prewarmed ENR medium to the respective organoid domes. Supplement the medium with 10 µM Rock-inhibitor (Y-27623) for the first 2 days.
9. Refresh the medium every 2-3 days. Make sure that the medium is warm to prevent dissociation of the temperature sensitive matrix.

7. Organoid processing for whole mount staining

NOTE: Coat the tips and tubes with 10% FBS in PBS before use to avoid organoids adhering to plastics. For pipette tips, it is sufficient to pipette once or twice up and down in 10% FBS/PBS solution before using the tip. For tubes, fill the tube with 10% FBS/PBS and then remove the solution.

1. Remove the organoid medium and add 200 µL of ice-cold PBS to the matrix domes. Place the plate on ice for 5-10 min.
2. Pipette up and down and transfer the solution to 0.6 mL micro-centrifuge non-adherent tubes. Centrifuge shortly for 30-60 s at 100 x g to let the organoids settle in the bottom.
   NOTE: Excess pipetting and long centrifugation breaks the organoids and disrupts the fibroblast-organoid interaction.
3. Remove excess liquid and add ice cold PBS. Centrifuge shortly for 30-60 s at 100 x g to let the organoids settle in the bottom.
4. Remove excess liquid and fix the organoid with 200 µL of cold 4% formaldehyde in PBS solution for 30 min on ice.
   CAUTION: Formaldehyde is toxic and must always be used in a chemical fume hood. Nitril gloves, safety glasses, and lab coats must always be worn.
5. Fixation of the organoid causes them to sink, and centrifugation is not needed anymore. Let the organoids sink by placing the tube upright and wait 2-3 min, remove the formaldehyde, and add 500 µL of cold PBS to wash away the formaldehyde.
6. Let the organoids sink, remove the excess PBS, and add 500 µL of fresh cold PBS. Let the organoids sink, remove the excess PBS, and add 500 µL of blocking buffer (5% normal donkey serum, 1% BSA, and 0.5% Triton X-100 in PBS). Put the tube on a rocker-shaker for 60 min at RT.
   NOTE: Blocking can also be done overnight (O/N) at 4 °C.
7. Let the organoids sink, remove the blocking buffer, and resuspend the organoids in 200 µL of blocking buffer with primary antibodies (see Table of Materials). Keep the organoids on a rocker-shaker O/N at 4 °C.
   NOTE: To improve the staining of nuclear proteins, low expressed proteins, or antibodies that show unspecific staining, the incubation time can be increased for 1 or 2 days at 4 °C. Placing the organoids on a rocker-shaker is essential as it prevents organoids from clumping together and increases staining efficiency.
8. Let the organoids sink, remove the primary antibody mix, and wash the organoids using 500 µL of 0.02% Triton X-100 in PBS (0.02% Tx) for 60 min at RT. Repeat three times.
   NOTE: When longer primary antibody incubation is needed, add a washing step with 0.02% Tx in PBS O/N at 4 °C.
9. Let the organoids sink, remove the washing buffer, and add 200 µL of fluorescence-conjugated secondary antibody in blocking buffer O/N at 4 °C. After adding fluorescent secondary antibodies, keep the organoids in the dark to avoid bleaching of signal.
10. Let the organoids sink, remove the secondary antibody mix, and wash the organoids using 500 µL of 0.02% Tx for 60 min at RT. Repeat three times.
11. Counterstain the organoids with 200 µL of DAPI (0.25 µg/mL) solution for nuclear staining if needed. Incubate the samples for 60 min at RT on a rocker-shaker.
12. Let the organoids sink, remove DAPI solution, and wash the organoids in 500 µL of PBS for 15 min at RT. Repeat three times.
13. Let the organoids sink and remove all excess liquid. Add 10 µL of clearing solution to the organoids and incubate for 15 min at RT.
    NOTE: The clearing solution is viscous, so cut off the outer part of a 20 µL pipette tip before pipetting the clearing solution. More clearing solution can be added if a bigger spacer is used for imaging. Mounting solution can be used instead of the clearing solution when not clearing the organoids. The clearing solution reduces the background when imaging and increases the imaging depth.
14. Place a 0.05 mm double-sided sticky 4-well spacer on a microscope slide. Add the 10 µL of clearing solution with the organoids in one well and place a cover slip on top of the spacer.
    NOTE: Organoids can also be mounted without the use of a spacer. The spacer keeps the organoid shape intact and prevents the organoids from being flattened.
15. Keep the slide at RT O/N to clear the organoids. For long term storage, keep the slides at 4 °C. Acquire images using a confocal microscopy system.