Mouse- and Human-derived Primary Gastric Epithelial Monolayer Culture for the Study of Regeneration

To avoid contamination, perform protocol in its entirety within a sterile tissue culture hood. Human fundic tissue was collected from patients undergoing sleeve gastrectomy according to the approved University of Cincinnati IRB protocol (IRB protocol number: 2015 – 4869).
All mouse studies were approved by the University of Cincinnati Institutional Animal Care and Use Committee (IACUC) that maintains an American Association of Assessment and Accreditation of Laboratory Animal Care (AAALAC) facility.

1. Establishing Human-derived 3D Gastric Fundic Organoids

NOTE: This protocol is based upon research previously published in this lab⁸.

1. Prepare 3D organoid media which consists of 50% Wnt-conditioned and 20% R-spondin conditioned media.
   1. Prepare Wnt conditioned media, culture the L cells in Wnt growth medium (Dulbecco's modified Eagle Medium (DMEM), 10% fetal bovine serum (FBS), 1% Penicillin/streptomycin) for 7 days, harvest the medium, and filter using a 0.22 µm filter.
   2. Prepare R-spondin conditioned media. Grow a modified HEK-293T R-spondin secreting cell line in R-spondin growth medium (Dulbecco's modified Eagle Medium (DMEM), 10% fetal bovine serum (FBS), 1% Penicillin/streptomycin). Upon attachment after 2 h, change the medium from these cells to OPTIMEM growth medium, 1% Penicillin/Streptomycin. Culture the cells for 7 days. Then harvest the R-spondin conditioned media and filter using a 0.22 µm filter.
      NOTE: This protocol has been published previously and for a more detailed protocol refer to ^6,7,8.
2. Thaw the growth factor reduced, phenol red-free basement membrane matrix for 16 h on the ice at 4 °C. Collect the tissue in the ice cold Ca²⁺/Mg²⁺-free DBPS in a large plastic container. Store on the ice until beginning step 1.4.
   NOTE: Tissue is collected from patients undergoing sleeve gastrectomy.
3. Using forceps, wash the tissue vigorously in a sterile beaker containing 100 mL Ca²⁺/Mg²⁺-free Dulbecco's Phosphate Buffered Saline (DPBS). Wash for approximately 30 s or until the debris and the blood has been removed. Next, using sterile gauze, wipe away the mucous layer from the epithelium.
4. Using large forceps, firmly grasp the muscle layer of the tissue and with force, scrape away the epithelium using the large curved hemostatic forceps. Collect the scraped epithelial fragments into a sterile, polystyrene petri dish.
5. Mince the epithelial tissue into approximately 2.0 x 2.0 mm² sized fragments using razors to optimize tissue digestion. Wash the tissue fragments with Ca²⁺/Mg²⁺ -free DBPS supplemented with antibiotics (Ca²⁺/Mg²⁺ -free DPBS, 1% Penicillin/Streptomycin, 0.25 mg/mL Amphotericin B /10 mg/mL Gentamicin, 50 mg/mL Kanamycin), until the wash is free of blood. Perform multiple washes if needed. Discard off the wash by carefully filtering the wash through a sterile gauze into a waste beaker.
6. Collect washed fragments into a 125 mL glass round bottom flask with a 25 mm stir bar and pre-warmed incubation media (basal media supplemented with 2 mM L-glutamine, 1% Penicillin/Streptomycin, 10 mM HEPES Buffer, 1 mg/mL Collagenase Type 1, 2 mg/mL cell culture grade Bovine Serum Albumin). Seal the round bottom flask with rubber septa.
7. Insert a 20 G spinal needle into the septa and connect it to an oxygen tank with rubber hosing. To filter out any contaminants, insert tissue paper into the tubing to create a filter. Turn on the oxygen outflow on low. Insert 10 – 15 outflow needles into the septa to avoid the rupture of septa.
8. Secure the set up to a ring stand using clamps and place it in a water bath calibrated to 37 °C with a stir plate for 30 – 45 min. After the tissue has been incubating for 15 min, remove 50 µL of incubation media and check visually for dissociated glands. If glands are not dense or have not separated from the tissue, leave for an additional 5 – 10 min.
9. Immediately following incubation, add 50 mL pre-heated DMEM/F12 to incubation media using a sterile serological pipette or by pouring directly into the incubation mixture.
10. Filter the gland mixture through a sterile gauze into four 50 mL conical tubes. Keep the filtrate on ice for 15 min to allow extracted glands to settle to the bottom of the conical tube. Be careful not to disturb the settled glands, remove and discard off the top 40 mL of supernatant using a serological pipette. Resuspend the remaining 10 mL in 10 mL of DPBS supplemented with antibiotics.
11. Distribute the mixture evenly into 5 mL cell culture test tubes. Centrifuge for 5 min at 65 x g at 4 °C and remove the supernatant carefully using a pipette. Resuspend the gland pellet in thawed basement membrane matrix using either a pipette or a 200 µL wide pipette tip. Mix until homogeneous.
    NOTE: It is crucial to perform this step on the ice and to avoid polymerization of basement membrane matrix while mixing. Additionally, it is important to visually inspect the gland density by sampling for 50 µL. The optimal density is ~70% confluency.
12. Next, plate the glands in 50 µL of basement membrane matrix using a wide-tipped pipette. Avoid producing any bubbles while plating.
13. To allow basement membrane matrix to polymerize, incubate for 10 – 15 min at 37 °C. If plating in a 12-well culture plate, add 1 mL of hFGO media (Advanced Dulbecco's modified Eagle medium/F12 medium supplemented with 2mM L-glutamine, 1% Penicillin/Streptomycin, 0.25 mg/mL Amphotericin B /10 mg/mL Gentamicin, 50 mg/mL Kanamycin, 10 mM HEPES Buffer, 1 mM n-Acteylcystine, 1 x N2, 1 x B27, 50% Wnt-conditioned medium, 20% R-spondin-conditioned medium supplemented with 100 ng/mL bone morphogenetic protein inhibitor, 1 nM gastrin, 50 ng/mL Epidermal Growth Factor, 200 ng/mL Fibroblast growth factor 10, 10 mM Nicotinamide, and 10 µM Y-27632 ROCK inhibitor) per well. If not using a 12-well culture plate add enough media to submerge the basement membrane matrix bubble.
14. Culture the cells at 37 °C in a 5% CO₂ humidified cell culture incubator. Change media every 4-5 days.

2. Establishing Mouse-derived 3D Gastric Fundic Organoids

NOTE: This protocol has been previously published ⁶. Thaw basement membrane matrix on the ice and prepare all reagents before beginning.

1. Sacrifice mice using a method approved by the research institution where research is being performed. Remove the stomach from the mouse and dissect according to the previously published protocols ⁶. Wash in 20 mL ice cold Ca²⁺/Mg²⁺-free DBPS.
   NOTE: C57BL/6 mice, aged 8 – 10 weeks, are used for fundic gastric organoid cultures. Mice were euthanized by inhalation of carbon dioxide followed by manual cervical dislocation before stomach removal.
2. Strip the muscle layer from the stomach and any visible blood vessels as previously published ⁶.
3. Separate the fundus from the antrum and forestomach using a sterile razor blade. Cut fundus using small surgical scissors into fragments approximately 2.0 x 2.0 mm. Using forceps, collect the fragments into a 15 mL conical tube containing EDTA incubation buffer (Ca²⁺/Mg²⁺-free DPBS, 5mM EDTA) and incubate at 4 °C for 2 h with gentle agitation.
4. In a sterile hood, leave conical tube and fragments on ice for approximately 5 min. Use a pipette to remove as much incubation buffer as possible leaving fragments untouched. Add 5 mL shaking buffer (1 g D-Sorbitol, 1.47 g sucrose in 100 mL Ca²⁺/Mg²⁺-free DPBS) and shake by hand with a force for 2 min. Allow the large fragments to settle and with haste, using a 1000p pipette, remove the small fragments that have yet to settle in the top layer.
5. Spin the removed fragments for 5 min at 4 °C at 65 x g.
6. While avoiding air bubbles, remove the supernatant, resuspend in the basement membrane matrix and mix until it is homogenous.
7. Use a wide tip pipette to plate the basement membrane matrix at 50 µL per well and then incubate at 37 °C for 20 min.
8. Add enough mFGO growth media (Advanced Dulbecco's modified Eagle medium/F12 medium supplemented with 2mM L-glutamine, 1% Penicillin/Streptomycin, 10 mM HEPES Buffer, 1mM n-Acteylcystine, 1 x N2, 1 x B27, 50% Wnt-conditioned medium, 20% R-spondin-conditioned medium supplemented with 100 ng/mL bone morphogenetic protein inhibitor, 10 nM gastrin, 50ng/mL Epidermal Growth Factor, 10 ng/mL Fibroblast growth factor 10, and 10 µM Y-27632 ROCK inhibitor) to submerge the bubble.
9. Culture the cells at 37 °C, 5% CO₂ humidified cell culture incubator. Change the media every 4 – 5 days.

3. Collagen Coating for 2D Transfer

NOTE: Perform all procedures in a sterile tissue culture hood unless otherwise specified. Begin collagen coating 24 h before transferring 3D organoids to the monolayer. Keep the rat tail collagen on the ice and shielded from light. Collagen coated plates are good for up to 2 weeks. If not using the coated plates immediately, wrap the plate in parafilm and store at 4 °C until needed.

1. Dilute the rat tail collagen to 50 µg/mL using 20 mM acetic acid.
2. Sufficiently cover the surface of the well with diluted collagen. For one well in a standard 12-well plate, coat with approximately 1 mL of collagen. Leave overnight at room temperature in a sterile hood.
3. Wash twice with 1 mL Ca²⁺/Mg²⁺-free DPBS per well and leave uncovered for 2 h at room temperature in a sterile hood.

4. Basement Membrane Matrix Coating for 2D Transfer

NOTE: Perform all procedures in a sterile tissue culture hood unless otherwise specified. Begin basement membrane matrix coating 2 h before transferring 3D organoids to the monolayer. Keep the basement membrane matrix on ice. Coated plates are to be used immediately and cannot be stored for an extended period of time.

1. Dilute the basement membrane matrix in a 15 mL conical tube with a cell culture grade water at a 1:10 ratio. Dilute on ice.
2. Using a pipette add 100 µL of diluted basement membrane matrix to each well of a standard 12-well plate. Using a cell scraper, evenly distribute the diluted basement membrane matrix throughout the well for an even coat.
3. Incubate at 37 °C for 1 h.
4. Remove the residual water using a pipette and dry at room temperature for 1 h before transferring 3D organoids.

5. Transfer of 3D m/hFGOs to 2D Gastric Epithelial Cell Monolayers

1. After 3D mouse- or human-derived FGOs have been cultured for 6 – 7 days, begin the transfer of 3D organoids to the 2D monolayer.
   NOTE: For each well of a monolayer required it is recommended to use 300 intact organoids or single cells derived from 300 organoids. In a robust culture it is expected to obtain approximately 150 organoid/well within a 12 well culture plate.
2. Be sure plates are coated before transferring.
   NOTE: The basement membrane matrix coated plates have a translucent gel layer, however, collagen coated plates will not have any obvious indication of the coating.
3. Dislodge the 3D basement membrane matrix bubble from the plate by directly pipetting 1 mL sterile Ca²⁺/Mg²⁺-free DBPS onto the center of the bubble.
4. Using a pipette, collect the basement membrane matrix and organoid mixture into cell culture test tubes. Collect at a ratio of 2 basement membrane matrix bubbles per tube. Centrifuge for 5 min at 40 x g and 4 °C.
5. Using a vacuum system, remove the supernatant and as much basement membrane matrix as possible. Resuspend in 4 mL of Ca²⁺/Mg²⁺-free DBPS, remove the supernatant, and repeat 2 – 3 times. For the whole organoid transfer protocol, proceed to step 5.10. Continue to step 5.7 for the single cell suspension protocol.
6. Pre-warm cell detachment solution  to 37 °C and add 1 mL to each test tube. Gently mix by either inverting tubes or pipetting up and down with a pipette. Incubate for 10 min at 37 °C.
7. Add 2 mL of Ca²⁺/Mg²⁺-free DBPS directly to each test tube.
8. Use a 26 G needle and syringe the mixture 2 – 3 times. Visually check for single cells. Syringe 1 – 2 more times if there are still organoids or clusters of cells.
9. Centrifuge at 40 x g for 5 min at 4 °C. Resuspend the pellet in 2D FGO media (2D mouse FGO growth medium: Advanced Dulbecco's modified Eagle medium/F12 medium supplemented with 2mM L-glutamine, 10 mM Penicillin/Streptomycin, 10% Fetal Calf Serum, 10 mM HEPES Buffer, 1 x N2, 1 x B27, supplemented with 10 nM gastrin, 50 ng/mL Epidermal Growth Factor, 1 µM TGF-β inhibitor, and 10 µM Y-27632 ROCK inhibitor. 2D human FGO growth medium: Advanced Dulbecco's modified Eagle medium/F12 medium supplemented with 2mM L-glutamine, 1% Penicillin/Streptomycin, 10 mM HEPES Buffer, 10% Fetal Calf Serum, 10 mM Nicotinamide, 1 x N2, 1 x B27, supplemented with 50 mg/mL Kanamycin, 50 ng/mL Epidermal Growth Factor, and 10 µM Y-27632 ROCK inhibitor, 1 µM TGF-β inhibitor).
10. Plate the resuspension on coated plates. Incubate plates at 37 °C. Replace media every 4 days or earlier if media turns yellow. Monolayer requires approximately 4 days to form.

6. Scratch Wound Assay Using 2D Gastric Epithelial Cell Monolayers

1. Using a razor blade, scratch the monolayer when the culture has reached 100% confluency.
2. Fix monolayers using 3.7% formaldehyde for 15 min at room temperature. Permeabilize with 0.5% non-ionic detergent/PBS for 20 min at room temperature.
3. Next block monolayers with 2% donkey serum for 1 h at room temperature and incubate with 1:1000 dilution of H⁺,K⁺-ATPase primary antibody overnight at 4 °C, wash with 0.1% non-ionic detergent/PBS and incubate with 1:100 dilution of donkey anti-mouse 488 secondary antibody and 10μg/mL of Hoechst cell nuclei stain for 1 h at room temperature.
4. To identify the surface mucous pit cells in 2D human-derived gastric monolayer cultures, stain cells with 20 μg /mL of Ulex europaeus (UEAI) FITC conjugate. Image monolayers on a confocal microscope.
5. Take images of wound every few h. Scratch will re-epithelialize between 24 – 48 h depending on the quality of the monolayer and variation between patients.