Generation and Manipulation of Rat Intestinal Organoids

NOTE: All cell culture should be handled using proper aseptic technique in a tissue culture hood. All animal work in this study was approved by Yale's Institutional Animal Care and Use Committee (IACUC).

1. Preparation of cell culture reagents

1. Prepare R-spondin 1 conditioned media following the manufacturer's instructions. Prepare Wnt3a conditioned media following the manufacturer's instructions. Prepare AdDMEM+ as described in Table 1.
2. Resuspend gastrin in sterile dH₂O to prepare a 100 µM stock. Aliquot and store at -80 °C. Resuspend N-acetylcysteine in sterile water to prepare a 100 mM stock. Aliquot and store at -20 °C for up to 1 month.
3. Resuspend recombinant human Noggin in phosphate buffered saline (PBS) + 0.1% bovine serum albumin (BSA) to prepare a 250 µg/mL stock. Aliquot and store at -80 °C. Resuspend recombinant mouse epidermal growth factor (EGF) in PBS + 0.1% BSA to prepare a 100 µg/mL stock. Aliquot and store at -80 °C.
4. Dilute recombinant human IGF-1 in PBS + 0.1% BSA to prepare a 100 µg/mL stock. Aliquot and store at -80 °C. Resuspend recombinant human FGF-2 in 5 mM Tris, pH 7.6, to prepare a 100 µg/mL stock. Aliquot and store at -80 °C.
   ​NOTE: For all the growth factors, use an intermediate dilution in PBS + 0.1% BSA to 100x the final concentration in the culture media. Store at -20 °C.

2. Establishment of rat small intestine organoids

NOTE: This protocol was modified from two previously published protocols for rat intestinal organoids^23,24.

1. Prepare rat intestinal organoid media (rIOM) as per Table 2 and set a water bath at 37 °C. This complete medium is stable for 5 days at 4 °C.
2. Prepare 10 mL of 3 mM ethylenediaminetetraacetic acid (EDTA) in PBS in a 15 mL conical tube and keep on ice. Thaw 250 µL of extracellular matrix extract (EME) on ice.
3. Fast a rat overnight with access to water ad libitum. Euthanize the rat according to the IACUC-approved protocol. In this protocol, adult male Sprague Dawley rats (weighing ~200 g) were euthanized via CO₂ inhalation (Table of Materials). Cervical dislocation was used as a secondary method of euthanasia.
4. Use sterile autoclaved forceps and dissection scissors for dissection. Place the euthanized rat onto the dissection surface ventral side up. Pinch the skin layer with forceps; the following cuts should be done at the surface level, so they are only cutting through this skin layer and not deep enough to damage internal organs.
5. To open the abdominal cavity, using large, sharp dissection scissors, cut through the skin layer in a large, longitudinal, surface-level cut in the center of the abdomen. Next, stemming from that cut, make two shorter horizontal cuts, one on each side. Use forceps to peel the skin away to expose the abdominal cavity. Cut through the peritoneal membrane to fully expose the internal organs in the abdominal cavity with ready access to the intestine.
6. Using scissors and forceps, locate the stomach and identify the duodenum approximately 2-3 cm distal to it, which appears as a yellowish segment. The proximal jejunum is located approximately 4-5 cm distal to the ligament of Treitz, which serves as a landmark between the duodenum and jejunum.
7. Place the isolated intestinal fragment into a 10 cm Petri dish. Clean the desired intestinal segment of mesentery as much as possible. Flush with 10 mL of ice-cold PBS until cleared of luminal contents. On a paper towel, cut the intestinal segment into ~2 cm long pieces. Open each intestinal piece longitudinally to expose the epithelium.
8. Using a glass microscope slide, scrape the exposed luminal surface to remove villi. Place intestinal pieces into the prepared EDTA solution on ice. Rotate at 4 °C for 30 min on a tube revolver set to 10 rpm.
9. At the dissecting microscope, pour the contents of the conical tube into a 10 cm Petri dish. Add in an additional ~5 mL of ice-cold PBS.
10. Using fine forceps, hold an intestinal segment and shake vigorously. It will be possible to see the epithelium release into the PBS. At first, the PBS will primarily contain villi.
11. Continue to shake. Periodically discard the PBS containing villi and add 10 mL of fresh ice-cold PBS to the intestinal fragments. Continue to shake the fragments and repeat this washing step until villi are no longer released into the PBS, but instead the PBS primarily contains crypts. Do not exceed 15 min for crypt isolation, so that cell viability is not compromised.
12. Discard the remaining intestinal fragments. Enrich the remaining PBS in the Petri dish for intestinal crypts. In a tissue culture hood, collect the PBS-containing crypts and filter through a 70 µm cell strainer (Table of Materials).
13. Centrifuge at 250 x g for 5 min. Remove the supernatant and resuspend the pellet in 5 mL of AdDMEM+. Centrifuge again at 250 x g for 5 min.
    NOTE: It is recommended to use a centrifuge with a swing bucket rotor.
14. Remove the supernatant, leaving ~50 µL of media with the pellet. Resuspend the pellet in the remaining media and add to the aliquot of the EME on ice. Gently pipette up and down to suspend the crypts evenly throughout the EME. Avoid introducing bubbles.
15. Plate 50 µL of EME domes into a 35 mm tissue culture dish and incubate for 20 min in a 37 °C, 5% CO₂ tissue culture incubator.
16. Add 2 mL of rIOM (Table 2) containing 10 µM Y27632 and 10 µM CHIR99021 (Table 3). After passaging, Y27632 and CHIR99021 can be discontinued.
17. Change the rIOM every 2-3 days. Passage as needed, usually between 3-7 days, depending on the initial number of organoids, size, and growth rate.

3. Passaging rat intestinal organoids

1. Thaw a 250 µL aliquot of EME on ice and prewarm the rIOM to 37 ˚C.
2. Aspirate the medium from a 35 mm plate containing organoids and add 1 mL of dissociation reagent to immediately release the organoids from the EME domes.
3. Immediately transfer dissociation reagent with fragmented organoids to a 15 mL conical tube. Wash the culture plate with 2 mL of AdDMEM+ and add to the 15 mL conical tube containing fragmented organoids.
4. With a glass Pasteur pipette, gently pipette up and down 15-20 times to fragment the organoids. Centrifuge at 350 x g for 2 min.
5. Add ~50 µL of solution from the bottom of the conical tube into the EME. Gently pipette up and down to mix. Avoid making bubbles.
6. Plate 50 µL of EME domes into a 35 mm dish and incubate for 20 min in a 37 °C, 5% CO₂ tissue culture incubator.
7. Add 2 mL of rIOM with 10 µM Y27632 and 10 µM CHIR99021. Change the growth media every 2-3 days. When changing media, Y27632 and CHIR99021 can be omitted.

4. Cryopreservation and thawing of rat intestinal organoids

1. Cryopreserving rat intestinal organoids
   NOTE: The cryopreservation protocol was modified from a previous protocol for human and mouse organoids²⁵. Prior to freezing, the organoids must have at least two passages in primary culture. It is advisable to grow organoids to the point of large spheroids or slightly budded organoids prior to cryopreservation, as this will result in a higher yield of viable organoids post-thawing. This can be accomplished by increasing the R-spondin conditioned media to 15% and/or by adding 10 mM nicotinamide (Table 3) to the organoid culture.
   1. Using a microscope, count the number of organoids on the 35 mm dish. Label the cryovials such that 200 organoids will be aliquoted into each cryovial.
   2. Remove the rIOM from the 35 mm dish. Replace with 2 mL of cold tissue culture grade PBS.
   3. Using a P1000 tip, release the EME domes from the bottom of the culture dish into the PBS by pipetting up and down. Continue to pipette up and down ~20 times to break up the EME and release the organoids. Collect the organoids and PBS into a 15 mL conical tube.
   4. Add 2 mL of cold PBS to the culture dish and pipette up and down to release any remaining organoids into the PBS. Transfer the PBS to the 15 mL conical tube.
   5. Pellet the organoids by centrifuging at 290 x g for 5 min. Remove and discard the supernatant without disrupting the organoid pellet.
   6. Wash the pellet by gently resuspending in 5 mL of cold AdDMEM+. Centrifuge at 200 x g for 4 min. Carefully remove and discard the supernatant.
   7. Resuspend the organoid pellet in 1 mL of cold freezing medium per 200 organoids. Aliquot 1 mL of organoids in freezing medium per labeled cryovial. Place the cryovials in a freezing container.
   8. Store the organoids in a freezing container at -80 °C for 24 h, then transfer the cryovials to liquid nitrogen for long-term storage.
2. Thawing rat intestinal organoids
   NOTE: This protocol was modified from a previous protocol for thawing human and mouse intestinal organoids²⁶.
   1. Thaw a 250 µL aliquot of EME on ice. Prepare rIOM supplemented with 15% R-spondin conditioned media, 10 µM Y27632, and 10 µM CHIR99021. Warm at 37 °C.
   2. Add 2 mL of defrosting medium (Table 3) to a 15 mL conical tube at room temperature.
   3. Retrieve and thaw a vial of organoids from liquid nitrogen by placing the vial in a 37 °C water bath until the vial is almost completely thawed.
   4. Add 1 mL of defrosting medium to the vial and transfer all of the contents to the conical tube containing defrosting medium. Wash the vial twice with 1 mL of defrosting medium and transfer to the conical tube.
   5. Centrifuge at 200 x g for 5 min. Aspirate the media, leaving ~50 µL of medium with the organoids. Transfer the media containing organoids to 250 µL of EME.
   6. Evenly distribute the organoids through the EME by pipetting up and down, avoiding bubbles. Pipette six 50 µL domes into a 35 mm tissue culture dish.
   7. Incubate in the tissue culture incubator for 15-20 min to allow the EME to polymerize. Add 2 mL of the prepared rIOM to the dish.
   8. After 2 days, replace the media with rIOM. The use of Y27632 and CHIR99021 can be suspended. Organoid growth may be slow in the first passage after thawing. It is advised to passage the organoids twice after thawing before beginning experiments.

5. Generation of rat intestinal 2D monolayers from 3D organoids

NOTE: The following protocol describes the volumes required to generate 24 wells of a 48-well plate coated with EME, starting with six domes of 50 µL (35 mm dish) containing ~300 intestinal organoids/dome (scale: one dome generates four wells), but can be scaled up or down as needed. As written, this protocol achieves ~80% confluence in 4-5 days. At higher confluency, the cells begin to acquire 3D organoid structures again. At low confluence (≤40%) , the cells remain as monolayers and are viable for ~14 days. If the purpose of the study is to use 2D monolayers, scale down so that one dome generates eight wells of a 24-well plate. The wells can also be coated with collagen I to form monolayers.

1. Preparation of coated surfaces
   1. To coat the plate with EME, dilute EME 1:20 in cold AdDMEM+ (Table 1). For coating with collagen, prepare collagen I according to the manufacturer's instructions. Dilute 5 mg/mL collagen I in AdDMEM+ to 100 µg/mL (1:50 in this case).
   2. Coat the plate with 200 µL of diluted EME or collagen to cover the well surface entirely. Incubate for 1-2 h at 37 °C in a tissue culture incubator. Prepare rat intestinal organoid medium for 2D monolayer culture (rIOM2D) as per Table 4.
2. Generation of monolayers
   1. Aspirate the media from a 35 mm plate containing organoids. Add 1 mL of PBS.
   2. Disrupt the EME in the wells by scratching with a P1000 tip. Pipette up and down approximately 20 times to loosen all the EME. Transfer everything to a 15 mL conical tube.
   3. Add 1 mL of PBS to the 35 mm plate to recover any additional organoids and transfer to the same 15 mL conical tube.
   4. Centrifuge at 350 x g for 2 min and aspirate the supernatant, including EME residue. Add 1 mL of trypsin solution to the organoid pellet and incubate at 37 °C for 2 min.
   5. Pipette up and down 10 times using a P1000 tip and add 2 mL of AdDMEM+ to neutralize trypsin.
   6. Centrifuge at 350 x g for 5 min. Aspirate the supernatant and add 4.8 mL of rIOM2D (Table 4). Resuspend the cell pellet.
   7. Before placing the organoids, take out any excess EME or collagen in AdDMEM+ from the wells. Then, add 200 µL of organoids in rIOM2D and 10 µM Y27632 to each precoated well.
   8. After 4-16 h, collect the media and centrifuge at 1,000 x g for 1 min. Transfer the supernatant to a new 15 mL conical tube and discard the pellet.
   9. Wash each well with 300 µL of PBS and add 200 µL of the centrifuged rIOM2D to each well. Change the rIOM2D every 2-3 days, suspending the use of Y27632.
3. Reforming 3D organoids from 2D monolayers
   NOTE: Monolayers grown on EME can be induced to reform 3D organoids, while monolayers grown on collagen I do not efficiently return to 3D organoids. Usually, 2-3 wells of organoids generated from monolayers can be used to make one dome of 50 µL of EME at day 5.
   1. Dilute the EME 1:4 in rIOM2D. When the monolayers reach ~80% confluency, carefully aspirate the media and add 100 µL of diluted EME to the wells containing monolayers.
   2. Incubate at 37 °C, 5% CO₂ in a tissue culture incubator for 20 min. Afterward, add 100 µL of rIOM2D and return to the tissue culture incubator. 3D organoids will be generated within 5 days of adding diluted EME.
   3. After small organoids have reformed (~day 5), prepare rIOM. Collect all the well contents, pipetting up and down to disrupt the EME, and transfer to a 15 mL conical tube.
   4. Centrifuge at 350 x g for 2 min. Aspirate the supernatant and EME residue.
   5. Add 1 mL of cold AdDMEM+ and centrifuge at 350 x g for 2 min. Remove the supernatant, leaving ~50 µL of media.
   6. Transfer the 50 µL of media and organoids to a 250 µL EME aliquot. Pipette up and down to distribute the organoids throughout the EME.
   7. Pipette 50 µL of EME domes into a 35 mm dish and incubate for 20 min in a tissue culture incubator.
   8. Add 2 mL of rIOM plus 10 µM Y27632. Change the growth media every 2-3 days. When changing the media, Y27632 can be omitted.

6. Genetic manipulation

1. Transient transfection of 2D monolayers
   CAUTION: Prepare rat intestinal epithelial monolayers following section 5.1 in a 48-well plate. Transfection must be performed at 70%-80% confluency. Always replace the media in the wells with 200 µL of fresh rIOM2D before transfecting. Calculate 260/280 nm and 260/230 nm absorbance ratios of the plasmid DNA; these must be above 1.8 to ensure good transfection results.
   NOTE: Use a plasmid control to calculate the efficiency of transfection. A plasmid that encodes a fluorescent protein is recommended for ease. In this protocol, pLJM1-EGFP²⁷ was used.
   1. Prepare 1 mg/mL of 20 kDa polyethylenimine (PEI; Table 3). For each well, prepare tube A (0.6 µg of plasmid + 50 µL of reduced serum medium) and tube B (1.8 µL of PEI + 50 µL of reduced serum medium). Maintain a DNA:PEI ratio of 1:3.
   2. Vortex both tubes for 30 s. Combine tube A and tube B, and vortex again for 30 s. If necessary, use a centrifuge to spin down. Incubate for 20 min at room temperature.
   3. Gently add the DNA/PEI complex drop-wise to 2D monolayer. Gently swirl the plate to mix. Incubate at 37 °C, 5% CO₂. The expression can usually be detected after 24 h.
      NOTE: If it is necessary to return to 3D organoid structures, wait for 48 h after transfection for adding diluted EME.
2. Lentiviral transduction of rat intestinal organoids
   NOTE: Before working with lentiviruses, obtain proper authorization and specialized training from the institution. Always wear appropriate personal protective equipment (PPE) when handling lentiviruses. While not outlined here, high-quality, concentrated lentivirus is essential for the successful infection of organoids. This protocol utilized empty pLJM1-EGFP vector²⁷ to express soluble GFP. This protocol is modified from a previously published protocol²⁸. The transduction efficiency depends on the quality and concentration of viral particles, efficient dissociation of organoids into small cell clusters, and the gene being expressed. When calculated 5 days after infection, the mean transduction efficiency prior to selection was 19.4% (± 6.5% standard deviation).
   1. At 2 days prior to lentiviral infection, plan to passage (section 3) two dense wells of a 24-well plate for each lentivirus that will be transduced. Each well of a 24-well plate can accommodate one 50 µL dome of EME.
   2. Once the EME solidifies, add 0.5 mL of rIOM supplemented with 10 mM nicotinamide, 10 µM Y27632, and 2.5 µM CHIR99021. This induces large spheroid morphologies, which is favorable for the efficiency of lentiviral transduction. Within 2 days of plating, rat organoids should be large spheroids. If significant differentiation (i.e., budding) is observed, repassage the organoids.
   3. Thaw the concentrated virus on ice. Prepare fresh transduction media, as per Table 3.
   4. Using a P1000 tip, release the EME domes from the bottom of the cell culture dish into the media by pipetting up and down. Continue to pipette up and down 20 times to break up the EME and release the organoids.
   5. Pipette the organoids and media into a 15 mL conical tube. All the wells can be pooled together, provided the organoids have the same genotype and are from the same line.
   6. Wash each well twice with 1 mL of cold AdDMEM+. Collect the AdDMEM+ and transfer it to the same 15 mL conical tube.
   7. Using a glass Pasteur pipette, mechanically disrupt the rat organoids. This is a critical step, as the aim is small multicellular clusters of cells. Pipette up and down ~30 times. Check for efficiency of the disruption under the microscope using a 4x objective. Continue this process until the cell suspension is primarily comprised of cell clusters with few organoids remaining.
      NOTE: Alternatively, organoids can be centrifuged at 200 x g for 5 min at room temperature, the supernatant carefully removed, and resuspended in 1 mL of recombinant enzyme replacing conventional trypsin/EDTA, prewarmed to 37 °C in the tissue culture incubator. Incubate the organoids in trypsin replacement for 2 min in a 37 °C water bath, with regular vortexing to promote dissociation. Avoid prolonged incubation with trypsin replacement, as this can promote cell death. Regularly check for efficiency of disruption under the microscope using a 4x objective. When the suspension is composed of primarily single cells with few cell cultures, dilute the trypsin replacement by adding 4 mL of AdDMEM+ and proceed with the following step.
   8. Spin the 15 mL conical tube containing the cell clusters at 200 x g for 5 min at 4 °C. Carefully remove and discard the supernatant, being careful not to disturb the cell pellet.
   9. Resuspend the cell clusters in 230 µL of transduction medium per well to be infected. For the infection, use one well of a 48-well plate for each lentivirus.
   10. Plate 230 µL of cell suspension into each well of a labeled 48-well plate. Add 20 µL of concentrated virus to each well. Use a P1000 tip to mix the virus/cell solution in each well and seal the plate with a transparent film.
   11. Perform spinoculation: centrifuge the plate at 600 x g for 1 h at 32 °C. Unseal the plate and incubate at 37 °C, 5% CO₂ for 6 h.
   12. Preheat a 24-well plate in the 37 °C incubator.
   13. Following incubation, pipette each well up and down and transfer the contents to a labeled 1.5 mL tube. Wash each well with 750 µL of AdDMEM+ and transfer to the tube. Spin the tubes at 600 x g for 5 min at 4 °C.
   14. Remove the tubes from the centrifuge and store on ice. Use a P1000 tip to carefully remove and properly dispose of the supernatant.
   15. Resuspend the cell pellet in EME and plate 50 µL domes into the preheated 24-well plate. Incubate for 15-20 min at 37 °C until the EME is polymerized.
   16. To each well, add 500 µL of rIOM supplemented with 10 mM nicotinamide, 10 µM Y27632, and 2.5 µM CHIR99021.
   17. Replace the media with rIOM plus 10 µM Y27632 1 day after infection. Change the media every 2-3 days. If selection will be performed, add selection 48-72 h after infection. For puromycin selection, use 2 µg/mL puromycin.

7. Immunofluorescence whole mount staining of organoids

1. Aspirate the media and add 4% paraformaldehyde (PFA) in PBS-Tween 20 (PBS-T) (Table 3) to the organoids in a cell culture dish. Incubate at room temperature for 10 min.
2. Release the organoids from the EME by pipetting up and down. Collect the organoids into a 0.75 mL tube. The organoids will settle to the bottom of the tube by gravity within a few minutes. If needed, centrifuge at 100 x g for 1 min.
3. Using a transfer pipette, remove the PFA and resuspend the organoids in PBS-T. Let the organoids settle to bottom of the tube. Remove the PBS-T and resuspend in 200 µL of block solution (Table 3).
4. Incubate at room temperature on a rocker or nutator for 45 min. Organoids can clump and settle to the bottom of the tube. Periodically finger flick the tube to resuspend and disperse throughout the solution.
5. Let the organoids settle to the bottom of the tube. If needed, centrifuge at 100 x g for 1 min. Remove block solution.
6. Add primary antibody diluted in 100 µL of block solution. Incubate at room temperature on a nutating mixer for 45 min at 24 rpm. Antibody concentrations vary depending on the antibody used.
   NOTE: This step can be extended based on the primary antibody, up to overnight at 4 °C.
7. Let the organoids settle to the bottom of the tube. If needed, centrifuge at 100 x g for 1 min.
8. Wash in PBS-T five times using a transfer pipette. Incubate at room temperature for 5 min on nutating mixer at 24 rpm. Repeat this step twice.
9. Add secondary antibody diluted 1:200 and 50 µg/mL 4′,6-diamidino-2-phenylindole (DAPI) diluted in 100 µL of block solution. Incubate at room temperature on a nutating mixer for 30 min at 24 rpm.
10. Let the organoids settle to the bottom of the tube. If needed, centrifuge at 100 x g for 1 min. Remove secondary antibody. Wash five times in PBS-T using a transfer pipette.
11. Incubate at room temperature for 5 min on a nutating mixer at 24 rpm. Repeat the washing step twice.
12. While the organoids are washing, heat an aliquot of VALAP sealant (Table 3) at 40-50 °C to liquify. Using a paintbrush, paint a thin square of VALAP on a microscope slide approximately the size of a coverslip. Use a 22 mm x 22 mm No. 1.5 coverslip .
13. Using scissors, cut the end of a P200 pipette tip. Transfer organoids to the VALAP well on the slide. Using a tissue wipe, carefully wick away the PBS-T. Do not let the organoids dry out.
14. Flood the VALAP well with an antifade mounting medium (Table 3). For a ~22 mm x 22 mm square, this will require 100-150 µL of antifade.
15. The organoids can cluster; swirl the antifade with a pipette tip to redistribute the organoids, if necessary. Mount with 22 mm x 22 mm No. 1.5 coverslip, avoiding air bubbles. Seal the coverslip by painting a thin layer of VALAP onto the edges.
    ​NOTE: For inverted microscopes, the organoids can also be mounted in a 35 mm glass-bottom dish.

8. Forskolin-induced swelling of rat intestinal organoids

1. Grow the organoids for 3-5 days after passaging in rIOM. It is advisable to grow organoids in a 24-well plate to ensure that the same region can be easily reimaged. Capture images before the addition of forskolin (T0).
2. Add forskolin (Table 3) directly to the organoid media to a final concentration of 10 µM. Add the same volume of dimethyl sulfoxide (DMSO) to the control wells.
3. Image the control and forskolin-treated wells at regular intervals, every 15-30 min. When not imaging, keep organoids in the incubator or use a controlled imaging system with incubation. Maximum swelling should be observed by 120 min.
4. Follow standard protocols to calculate relative swelling from the acquired images²⁹.