Standardization and Maintenance of 3D Canine Hepatic and Intestinal Organoid Cultures for Use in Biomedical Research

The research was approved and performed in compliance with the Institutional Animal Care and Use Committee of Iowa State University (IACUC-19-337; IACUC-18-065; IACUC-19-017).

NOTE: The following section (steps 1-3) describes the Canine Organoid Isolation Protocol.

1. Preparation for isolation

1. Transport tubes: Before organoid isolation (typically 24 h prior), fill a 50 mL conical tube with 10 mL of Dulbecco's Modified Eagle Medium/Nutrient Mixture F-12 (Advanced DMEM/F12) enriched with 0.2 mL of Pen Strep.
2. For laparoscopic, incisional, or excisional biopsies, prepare three additional 50 mL conical tubes. Fill these tubes with 10 mL of complete chelating solution (1x CCS; see Table 1).
   NOTE: For step 1.2, 2 mM N-Acetylcysteine (NAC) in Phosphate Buffered Saline (PBS) can also be used as a traditional solution used for stem cell harvesting. There were no differences observed when using 1x CCS or NAC in PBS. Both solutions are added to release cells in the solution.
3. Keep the tubes at 4 °C overnight and transport the tubes on ice for the remainder of the protocol.
4. Prepare five 15 mL centrifuge tubes with 5 mL of 1x CCS, one 15 mL centrifuge tube with 3 mL of 1x CCS, one empty 15 mL centrifuge tube (supernatant tube), and one 15 mL centrifuge tube with 5 mL of 1x CCS and 2 mL of fetal bovine serum (FBS).
   NOTE: The above can be prepared prior to the day of isolation if multiple samples will be processed. For illustration, see the isolation tube layout in Figure 2.
5. On the day of isolation, prepare a Petri dish, scalpel, ice bucket, and cold Advanced DMEM/F12 in the biosafety cabinet. Place the required number of 24-well cell culture plates in the incubator (37 °C; 5% CO₂ atmosphere) to pre-warm.
6. Place solubilized extracellular membrane matrix (ECM; see Table of Materials) on ice to begin thawing.
   NOTE: Submersion in ice protects against rapid thawing and helps avoid solidification. A box of pipette tips can be placed in the freezer to assist in plating the solubilized ECM.
7. Prechill a centrifuge to 4 °C.
8. Move Complete media with growth factors "CMGF+" or "organoid media" (see Table 1 for composition) from the freezer/refrigerator to a 37 °C water bath. Avoid direct light exposure when possible.

Figure 2: Isolation tube layout. Recommended setup for tissue harvesting includes 10 mL of Advanced DMEM/F12 and 0.2 mL of Pen Strep in a 50 mL centrifuge tube. Additionally, three 50 mL tubes filled with 10 mL of 1x CCS are required for surgical biopsies or necropsies. Recommended tube layout for isolation steps includes five tubes containing 5 mL of 1x CCS to be used during CCS wash steps. The first tube is used as a sample tube containing the minced tissue, and the remaining tubes serve as reservoirs of 1x CCS to be added to the first tube. The sixth tube contains 3 mL of 1x CCS for flushing the remaining tissue from the sample tube when transferring to a 6-well plate. These six tubes will be used before the EDTA incubation step. A tube with 5 mL of 1x CCS and 2 mL of FBS serves as a sample tube after EDTA incubation, and supernatant from this tube is transferred with stem cells into an empty tube for the rest of the isolation. Keep tubes at 4 °C before beginning isolation. Please click here to view a larger version of this figure.

2. Tissue harvesting

1. Intestinal endoscopic, and laparoscopic biopsies (diameter 2.8 mm) can be acquired using large biopsy forceps. Harvest at least eight endoscopic samples per intestinal site.
2. Collect samples directly into transport tubes and place them on ice.
3. For surgical biopsies and necropsies, harvest tissue pieces with a size of 0.5 cm x 0.5 cm and place them in the first 1x CCS tube.
   NOTE: For intestinal biopsies, remove any remaining intestinal content and scrape the mucosal layer with a scalpel to remove villi. If samples are plentiful, additional biopsies can be collected in cryovials containing RNA storage reagent (1 mL) or paraffin-embedded for future comparison between organoids and their tissue of origin.In the case of taking biopsies for TEM analysis, do not scrape the villi and store sample in preservative (3% paraformaldehyde and 3% glutaraldehyde in phosphate-buffered saline (PBS)) and store at 4 °C.
4. Shake the 1x CCS tube vigorously for ~30 s, and then transfer the sample to a new 1x CCS tube using forceps. Repeat this process twice.
5. Transfer the sample from the last 1x CCS tube to the transport tube (Advanced DMEM/F12 + Pen Strep) and bring the sample back to the laboratory.
   NOTE: Samples pre-treated this way can also be shipped overnight on ice (do not ship on dry ice).

3. Organoid isolation

NOTE: Perform isolation using aseptic techniques in a biosafety cabinet. See Figure 3 for canine organoid isolation workflow.

1. Shake tissue sample in the transport tube for ~30 s and remove excessive supernatant until there is 0.5 mL left in the tube by slow pipetting near the fluid surface. Be sure not to discard any tissue.
2. Transfer tissue and remaining supernatant to a sterile Petri dish. Using a disposable scalpel (or sterilized forceps and scissors), cut and mince the tissue into smaller pieces (size of 1 mm²) resembling a mash consistency for approximately 5 min.
3. Pipet the minced tissue with liquid from the Petri dish to the first CCS tube. Add 2 mL of Advanced DMEM/F12 to the Petri dish, flush the remaining tissue, and transfer to the first CCS tube.
4. Vortex the 1x CCS tube for 5 s approximately five times. Allow biopsies to settle to the bottom of the 15 mL tube (approximately 1 min) and remove the supernatant until there is 5 mL left in the tube. Transfer the 1x CCS from the new tube to the sample tube.
5. Repeat the previous step for the next two tubes. On the final two washes, remove the supernatant down to 3 mL remaining in the tube.
6. Transfer the biopsies and 1x CCS from the sample tube to one well of a 6-well plate. Then, add 3 mL of 1x CCS to the sample tube, gently swirl to collect any remaining tissue, and transfer to the same well of the plate.
7. Add 150 µL of 0.5 M EDTA (to achieve a total volume of 6.15 mL in a well). Place the 6-well plate on a 20°, 24-rpm nutating mixer/rocker at 4 °C. Incubate the liver samples for 10 min and the intestinal samples for 1 h on the moving rocker.
8. Transport the 6-well plate back to the biosafety cabinet. Transfer minced tissue and liquid to a 1x CCS/FBS tube and allow the tissues to settle. Transport the supernatant (this portion now includes free stem cells) and approximately 0.2 mL of the upper portion of the tissue to the empty tube.
9. Spin down the tube containing the sample (700 x g for 5 min at 4 °C). Stem cells are now pelleted along with the minced tissue. Remove and discard the supernatant carefully to not disturb the pellet.
10. Resuspend the pellet in Advanced DMEM/F12 and spin the tube again (700 x g for 5 min at 4 °C). Aspirate the supernatant and do not disturb the pellet.
11. Calculate the volume of solubilized ECM needed for seeding the dissociated cells and tissue. Use 30 µL of solubilized ECM per well of a 24-well plate to achieve proper seeding density.
    NOTE: Embed a sample post isolation in 4 to 6 wells of a 24-well plate (i.e., based on the amount of minced tissue in the sample).
12. Add the calculated volume of solubilized ECM to the sample tube and slowly pipette up and down to avoid the formation of bubbles. Seed the suspension in the middle of the wells so that the solubilized ECM can form a dome-like structure.
    NOTE: Using pipette tips from a -20 °C freezer assists in plating solubilized ECM. If tissue chunks are larger than the P200 pipette tip, use wide cold tips or cut cold P1000 tips to assist in plating. Keep the sample with the solubilized ECM on ice whenever possible.
13. Transport the plate to an incubator (37 °C; 5% CO₂ atmosphere) and allow the solubilized ECM to solidify for ~30 min.
14. Mix ROCK inhibitor and GSK3β in CMGF+ (concentrations in Table 1). Add 500 µL of this solution (CMGF+ R/G) to every well. Place the plate in the incubator (37 °C; 5% CO₂ atmosphere).

Figure 3: Canine organoid isolation workflow. The harvested tissue sample is transferred to a Petri dish and minced properly. The sample is then transferred to a 1x CCS tube, and wash steps are performed. For the EDTA incubation in a 6-well plate, the sample is then transferred to a tube containing 1x CCS and FBS. After the tissue settles, the supernatant with a small amount of tissue is transferred to an empty tube. The sample is consequently spun, the supernatant is removed, and the pellet is resuspended in Advanced DMEM/F12. The tube is spun again, and the supernatant is aspirated and discarded. Solubilized ECM is added to the tube, mixed, and the sample is plated in a 24-well plate. The plate is consequently incubated (37 °C; 5% CO₂ atmosphere) for 30 min, and media is then added. Please click here to view a larger version of this figure.

NOTE: The following section (steps 4 and 5) describes the Canine Organoid Maintenance Protocol. Change media according to Table 2 and check the organoids daily for signs of apoptosis, contamination, overcrowding, and detachment of solubilized ECM. Daily notes must be taken according to Figure 4 to accurately monitor conditions and experimental effects on the cultures. See Supplementary Table 1 for a template allowing for accurate and reproducible organoid culture-related notetaking. For hepatic organoids, use CMGF+ enhanced with ROCK inhibitor and GSK3β.

Figure 4: Organoid sizing and density guide. (A) Organoid size chart for accurate tracking of organoid growth. The Sizing Guide includes extra-small (XS), small (S), medium (M), large (L), and extra-large (XL) categories. (B) The Density Guide consists of very low density (VLD), low density (LD), medium density (MD), high density (HD), and very high density categories (VHD). (C) Representative images of bacterial and fungal contamination of the organoid sample. (D) Representative images of organoid overcrowding and apoptosis. The objective magnification is indicated in every panel. Please click here to view a larger version of this figure.

Table 2: Media change recommendation. A recommended timeline of weekly media change. Media (500 µL of CMGF+ for intestinal organoids, or 500 µL of CMGF+ R/G for hepatic organoids) is recommended to be changed every other day. To account for the extra hours on a weekend, 750 µL of media is added on Friday afternoons, with media being refreshed on Monday mornings.

4. Organoid cleaning

NOTE: Organoid cleaning or passaging must be performed regularly to upkeep the health of the organoid culture. Perform the cleaning procedure whenever apoptosis in the culture, presence of debris, overcrowding of the organoids, or detachment of solubilized ECM is noticed. Refer to Figure 4D.

1. Remove the media from the wells while tilting the plate to avoid destruction of the solubilized ECM.
   NOTE: If solubilized ECM detachment is considerable, it is advised to transfer the media to the 15 mL tube to avoid losing fragments of solubilized ECM containing the sample.
2. Add 0.5 mL of prechilled Advanced DMEM/F12 to each well to dissolve the matrix domes by repeated pipetting using a P1000 tip (avoid creating excessive bubbles). Transfer the organoid-containing dissolved matrix to a 15 mL centrifuge tube. Keep the tube on ice and if the volume is lower than 6 mL, slowly fill the tube with Advanced DMEM/F12 to reach a total volume of 6 mL.
3. Spin the tube (700 x g for 5 min at 4 °C) and remove all the supernatant while making sure not to disturb the pellet.
4. Add the required volume of solubilized ECM (30 µL per well to achieve proper seeding density) and slowly resuspend the pellet by pipette mixing. Plate the suspension in the middle of the 24-well plate to form a dome.
5. Place the plate in an incubator (37 °C; 5% CO₂ atmosphere) for ~30 min, and then add the appropriate volume of media (Table 2).

5. Organoid passaging

NOTE: Passaging is typically performed 5-7 days after initial culture to expand the organoid cell line. Organoid cultures can typically be expanded in a 1:3 ratio. Images of healthy cultures ready for passage can be seen in Figure 4. Organoids have to be at least medium in size.

1. Perform steps 4.1 to 4.3.
2. Remove the supernatant leaving 0.5 mL in the tube. Make sure not to disturb the pellet.
3. Add 0.5 mL of trypsin-like protease (see Table of Materials), mix properly by aspirating with a pipette and incubate in the 37 °C water bath (incubate intestinal organoids for 8 min or hepatic organoids for 10 min). Continue to mix the solution by flicking the tube several times at the half-time point of the incubation.
4. Move the tube containing the sample back to a biosafety cabinet and slowly add 6 mL of prechilled Advanced DMEM/F12 to inactivate trypsin-like protease and stop the dissociation of the cells.
5. Spin the tube (700 x g for 5 min at 4 °C) and remove the supernatant. Make sure not to disturb the pellet.
6. Perform steps 4.4. and 4.5.
   NOTE: The following section (steps 6-9) describes Organoid Harvesting and Biobanking Protocol. Organoid cultures have to be free of tissue that has been removed during previous passages. Cultures should also be healthy, at least large in size, and medium to high in density. Images of healthy cultures ready for downstream applications can be seen in Figure 4. Moving downstream with preservation and harvesting of sub-optimal organoid cultures can negatively impact characterization results and the biobank's viability. Review the recommended 24-well plate layout in Figure 5.

Figure 5: Recommended 24-well plate layout. The recommended layout of the 24-well plate for basic characterization after expansion of the organoid culture. Paraffin-embedding of six wells (medium to large organoids of medium to high density) will typically allow for a high concentration of organoids in a histology block. Four wells of organoids can be pooled to one cryovial with RNA storage reagent for downstream applications. Fourteen wells are used for biobanking the organoid samples and provide material for up to seven cryopreserved vials. Please click here to view a larger version of this figure.

6. Fixation of organoids

1. Remove media from the well and make sure not to disturb the solubilized ECM dome.
2. Add 500 µL of Formalin-Acetic Acid-Alcohol solution serving as a fixative (FAA; composition in Table 1).
3. Store organoids at room temperature. After 24 h, aspirate FAA and fill the well with 70% ethanol. Wrap plates with a laboratory flexible film tape (see Table of Materials) to avoid rapid evaporation. The organoids are now ready for paraffin-embedding. The paraffin-embedding of the organoid culture is performed in traditional metal base molds.

7. RNA preservation

1. Remove media from the wells and make sure not to disturb the solubilized ECM dome.
2. Use 0.5 mL of prechilled Advanced DMEM/F12 per well to dissolve solubilized ECM domes by repeatedly pipetting up and down (avoid creating excessive bubbles). Transfer the organoids to a 15 mL centrifuge tube. Keep the tube on ice and if the volume is lower than 6 mL, slowly fill the tube with Advanced DMEM/F12 to reach 6 mL of total volume.
3. Spin the tube (700 x g for 5 min at 4 °C) and remove all the supernatant. Make sure not to disturb the pellet.
4. Add 100 µL of PBS to the sample tube and resuspend the pellet by gentle pipetting. Transfer the contents of the sample tube to a cryovial.
5. Add 900 µL of RNA storage reagent (see Table of Materials) to the sample tube and mix to collect any remaining organoids. Transfer these residual organoids to the cryovial and store them at -80 °C (typically four wells pooled to one cryovial will be sufficient for downstream applications, including qPCR and RNA sequencing).
   NOTE: One cryovial typically produces a total of 4,000 ng of RNA (measured via spectrophotometer analysis).

8. Organoid biobanking

NOTE: Biobanking usually occurs 3-4 days after passage. The signs of apoptosis should not be present in the culture to perform this method. Refer to Figure 4 for referencing size and density appropriate for freezing. Biobank medium to extra-large organoids at medium to very high densities. An emergency freeze step can be followed if an organoid cell line is especially rare or further viability is not guaranteed. Follow the same steps for normal organoid biobanking (steps 8.1 to 8.4). Emergency freezing is done with smaller and less dense cultures. Pool as many wells of growing organoids into one cryovial following steps 8.1 to 8.4. Keep in mind that a sufficient number of organoids must be kept alive in an attempt to expand the culture (emergency freezing is simply a backup procedure to protect against possible loss of culture via contamination or other unexpected occurrences).

1. Follow steps 7.1 to 7.3.
2. Add 1 mL of freezing media per cryovial (composition in Table 1) to the sample tube and gently resuspend the pellet by pipetting up and down.
3. Transfer 1 mL of the solution to one cryovial (ratio of two wells/cryovial) and keep the cryovials on ice.
4. Transfer the cryovials from ice to a freezing container (regularly refill the reservoir with isopropanol) and immediately transfer to -80 °C. Move the samples to liquid nitrogen (-196 °C) for long-term storage after 24 h.
   NOTE: An alcohol-free cell freezing container can also be used instead of a traditional one. Make sure the samples do not thaw during transport from the -80 °C to the long-term liquid nitrogen storage. Repeated thawing decreases the viability of the cell culture.

9. Revival from liquid nitrogen storage

NOTE: When choosing to thaw an organoid line, a subset of the revived organoids must be refrozen and replaced in the biobank as quickly as possible with at least one (preferably more) cryovials.

1. Place solubilized ECM on ice to slowly thaw, put a 24-well plate in the incubator (37 °C; 5% CO₂ atmosphere), and prepare the required reagents such as a 15 mL tube and Advanced DMEM/F12.
2. Recover a cryovial containing an organoid sample from liquid nitrogen storage and immediately transfer it to a heat bath (37 °C) for 2 min.
3. Transfer contents of the cryovial to a 15 mL centrifuge tube in a biosafety cabinet. Slowly add prechilled Advanced DMEM/F12 to reach a total volume of 6 mL.
4. Spin the tube (700 x g for 5 min at 4 °C). Remove the supernatant and make sure not to disturb the pellet.
5. Add 180 µL (30 µL per well) of solubilized ECM to the pellet and plate this suspension in the pre-heated 24-well plate. One cryovial can be plated to six wells of a 24-well plate.
6. Place the 24-well plate in the incubator (37 °C; 5% CO₂ atmosphere) for ~30 min and add CMGF+ R/G (for intestinal organoids, switch to CMGF+ in 24 to 48 h).
   NOTE: When a sample is plated and growing in the 24-well plate, it must be allowed to recover for at least 2 days before passaging to increase viability.