Intestinal Organoid Culture

Notes

• This media is extremely expensive, 500 ml will cost $3,750 or $7.50 per ml
• Can also make your own 1M HEPEs by adding 238.3 g HEPES to 700 ml water, add approximately 5.5 g NaOH pellets to bring pH to 7.5, then add H2O to 1 L
• To make stock solution of mouse recombinant EGF, resuspend lyophilized powder in sterile PBS to a stock concentration of 50 µg/ml (2 ml PBS in a 100 µg bottle). Filter sterilize through 0.22 µm filter. Aliquot in 100 µl volumes and store at -20ºC.
• To make stock solution of mouse recombinant noggin, resuspend lyophilized powder in sterile PBS to a stock concentration of 100 µg/ml (1 ml PBS in a 100 µg bottle). Filter sterilize through 0.22 µm filter. Aliquot in 100 µl volumes and store at -20ºC.
• To make stock solution of human recombinant RSPO1, resuspend lyophilized powder in sterile PBS to a stock concentration of 100 µg/ml (1 ml PBS in a 100 µg bottle). Filter sterilize through 0.22 µm filter. Aliquot in 100 µl volumes and store at -20ºC. Get this from cell line that produces it. Optionally, use conditioned media from ? cell line, strain the media and use at 1:10. When switching to new batch of media, compare head-to-head with old media using 1:5, 1:10, and 1:20
• Optional for RSPO1 use filtered media from a cell line that expresses RSPO1. To normalize between batches, grow organoids with the old batch and with various concentrations using the new batch.

Crypt culture protocol

• Coat all pipettes with FCS before using them up until you get the pellet of crypts
• Prepare enough 2.5mM EDTA/PBS for the number of samples you are preparing (25 mls/sample = 50 mls/mouse). Add 250 µl of 0.5 M EDTA to 50 ml PBS (final concentration 2.5 mM) and chill on ice in 50 ml conical.
• Place 500 ml bottle of PBS on ice
• Place 1.5 ml tubes with enough matrigel on ice. You will need 200 µl for each prep.
• Place 24-well plate in 37ºC incubator
• Euthanize mouse, open up chest cavity
• Cut stomach in half. grasp the lower half of the stomach connected to the intestine and slowly lift, using scissors to remove fat/mesentery as you pull the intestines up. Cut small intestine when you reach cecum and place the entire small intestine in a 10 cm petri dish containing 10 ml PBS (optional, can be on ice, but does not have to be).
• Cut the large intestine where it attaches to the cecum, lift and remove fat as before. Cut at anus and place large intestine in 10 cm petri dish containing 10 ml PBS. (optional, can be on ice, but does not have to be).
• Remove all fat from intestines. If fat remains, some of the pieces will float.
• Cut intestines longitudinally with special intestine scissors (Made in Germany FST 14080-11). Make sure forceps are flat forceps. Insert blunt blade of intestine scissors through the pyloris into the upper small intestine and start cutting. Do not cut all the way, but pull the intestines up the scissors blade using the forceps so the blunt end always remains inside the intestines. Keep scissors pointed down to avoid tearing. Cut through the entire length of the intestines. Go slowly.
• Wash the open intestine in the petri dish by gently, but rapidly making swishing movements (u-turns) and slowly lifting the intestine out of the petri dish. The goal is to get all the intestinal debris to stay in the petri dish. Transfer the clean intestine to a fresh petri dish with 10 ml PBS. Repeat the lifting/swishing procedure a few times in this clean petri dish.
• Lay the intestine on a hard surface, graspend with forceps and use scissors to flatten out the intestines so that the villi are pointing upwards.
• Scrape off the villi using a glass microscope slide by firmly drawing the edge of the slide along the entire length of the intestines. Do this repeatedly. Each time you do it, villi and other gunk will come off onto the edge of the slide, which you can wipe off on a kim wipe. Repeat until very little material comes off. Presence of differentiated cells of villi in crypt culture will inhibit organoid formation.
• After cleaning the intestines, use a razor blad to cut the intestines into 2 mm slices.
• Use the razor blade push all the pieces into a pile and then use the forceps to transfer all the pieces to a 50 ml conical containing 10 ml of ice cold PBS
• Pipette entire 10 mls up and down 3x using a 10 ml pipette (don't forget to coat with FBS). The purpose here is to dislodge all the single cells and other debris, while keeping the crypts attached to the small pieces of intestine. Hold the 50 ml conical at an angle while you are performing the procedure.
• Let pieces settle for 10 to 30 seconds, then carefully remove as much supernatant as possible with the pipette and discard the supernatant.
• Refill tube with ice cold 10 ml PBS and repeat the process at least 10 times until PBS is clear.
• After removing supernatant from the last wash, resuspend in 25 ml of 2.5mM EDTA/PBS (which you prepared at the start and left on ice).
• Rotate 50 ml conical at 4ºC for 30 minutes. Do not leave in EDTA/PBS for too long because the purpose of this step is to "loosen" the crypts from the pieces of intestine, but not to dislodge them, so longer incubations will cause you to lose crypts.
• Remove conical from rotator, let intestine pieces settle, remove supernatant using a pipette and discard EDTA/PBS
• Resuspend with 10 ml PBS (optional: switch to ADF instead of PBS for greater survival). This is considered the first fraction, or wash step. Pipette entire volume up and down 3x with a strong pipette, try and get pieces to rub against side of tube for mechanical shearing. Let pieces settle for 10 to 30 seconds then remove and discard supernatant.
• Resuspend in 10 ml PBS (or ADF) and pipette 3x again. Let pieces settle, then remove and save supernatant in a clean 50 ml conical. You can usually remove about 6 to 8 mls without getting any of the intestinal pieces. At this point, the crypts are being dislodged from the pieces and are floating in the supernatant, making it cloudy. The crypts are too small to see with the eye, so you shouldn't see chunks floating in the supernatant. This is considered fraction 2.
• Repeat two more times, each time adding the supernatant to the 50 ml conical containing fraction 2. These are considered fractions 3 and 4. Save the conical containing the pieces of intestine, in case you need to return for fraction 5, which usually is not necessary.
• For one mouse at this point, you will have two 50 ml conicals containing roughly 18 to 24 mls of PBS (or ADF) containing crypts. Add enough PBS (or ADF) to the tubes so that they contain equal volumes.
• Centrifuge for 5 min at 1200 rpm (290 rcf) at 4ºC. You should see a small pellet which contains crypts and single cells. Pour off the supernatant carefully, leaving pellet in tube.
• At this point you can stop coating pipettes in FBS before using.
• Resuspend crypt/cell pellet in 10 ml ADF (or PBS). Pipette entire volume up and down at least 10x to reduce clumping. Make sure cells are resuspended before straining in next step, as crypts are around 40 - 70 µm in size.
• Pass through 70 µm cell strainer into a clean 50 ml falcon tube. Transfer entire volume to a 15 ml conical.
• Centrifuge for 2 min at 600 rpm at 4ºC so that single cells will not be included in the pellet. If there is no pellet at 600 rpm, increase to 800 rpm. Gently pour off supernatant.
• Perform the next step 2 or 3 times until supernatant is fairly clear. Remember to pipette vigorously to disrupt cell clumps.
• Resuspend pellet in 10 ml ADF and centrifuge again for 2 min at 600 rpm at 4ºC. Pour off supernatant.
• After washing 2 to 3 times, resuspend pellet in 5 to 10 ml ADF (small pellets, use less, big pellets use more) and pipette up and down another 10x.
• Calculate number of crypts. Mix 10 µl of crypts with 10 µl of trypan blue. Place all 20 µl on a microscope slide (no coverslip) and count all the crypts in the entire drop using a quality inverted scope. This is a difficult step because cell clumps can look like crypts, even though they are not. Basically trying to find small clumps that are test-tube in shape, with an open end and a round bottom. The other clue is the presence of a gray/black line surrounding clump, which represents the epithelium. If the preparation was good you should expect roughly 50 crypts in the drop.Calculate number of crypts in full volume. For example, if you counted crypts in 10 µl from a 5 ml sample, multiply the number of crypts you counted by 500 to get total number of crypts.
• Aliquot enough media/crypts into one tube so that there are 1,500 crypts in the tube. Aliquot enough media/crypts into a second tube so that there are 3,000 crypts.
• Centrifuge both tubes for 2 min at 600 rpm at 4ºC.
• Remove supernatant with a pasteur pipette attached to a vacuum, being careful not to touch the pellet. Leave ~50 µl of media in tube.
• Resuspend crypt pellet in the 50 µl that remained in the tube
• Add an additional 100 µl Matrigel and mix well with the pipette, but avoid making any bubbles. Remember to place the tube with the matrigel back on ice immediately after using it.
• Draw up the entire 150 µl volume with crypts into a pipette, being careful to avoid any bubbles. Carefully pipette the matrigel into three wells of a pre-warmed 24-well plate. Place matrigel drops in middle of wells and try to evenly aliquot into each well so that each well receives about 50 µl. Do not let matrigel touch side of plate and avoid leaving any bubbles. If there is a bubble, remove it by sucking it up with a pipette. Incubate 24-well plate at 37ºC for 5-10 min until the matrigel is solidified. For each prep, there should be six wells, three with 500 crypts each and three with 1,000 crypts each. Plating at a higher density actually inhibits crypts growth.
• Add 500 µl Crypt culture medium to each well and incubate at 37ºC 5%CO2.
• Typically, crypts start to bud after 2-4 days in culture.
• To measure crypt efficiency, count crypts on day 2 and day 5. Published results usually report day 5 efficiency only.
• Add EGF/Noggin/RSPO1/Jagged1/Y27632 every other day.
• Exchange whole culture medium every four days.

Passaging organoids

• Organoids should grow enough to passaged in 7-10 days
• Remove media from wells.
• Add 200 µl ice cold ADF to each well. This well melt the matrigel. Remove contents from the three replicate wells and combine in a single 1.5 ml tube.
• Put a pipette tip on a pipetter and press against a hard surface to bend the tip of the pipette. Then use this pipetter to pipette the organoids at least 100x. This should disrupt the organoids into small pieces and single cells.
• Transfer to a 15 ml conical and spin for 2 min at 600 - 800 rpm at 4ºC. Remove supernatant, leaving crypt pellet.
• Proceed with plating as above. Note that you will just plate this back into three wells. You will not get more organoids than when you started, and may actually get fewer. It is not like cell culture, you are not splitting the cells.

Single sorted cell organoid culture protocol

• Single sorted cell culture is much more difficult than whole crypt culture. Therefore I recommend to do the culture after succesfully culturing whole crypts or dissociated cell culture. Dissociated cells die easily, so be sure to do the procedure as quickly as possible and keep them as cold as possible.
• Need to change media, using Jagged1 and Rho kinase inhibitor Y27632
• Perform all steps at room temperature, when doing single cell for flow sorting. Washing steps always with media and not PBS, and can even keep the media at 37ºC. Single cell disruption using 3 ml media + 1 ml TripleLE at 37ºC for 30 minutes, pipetting up and down every few minutes.

Freezing organoids

Human intestinal organoids

This protocol was developed by Pat Scott and our lab and is based mostly on Clevers lab protocols (see Sato, 2011, Medema, 2013, Mahe, 2015, and Clevers, 2015)

Human Protocol

Generating human colon organoids

• Follow BSL2 safety precautions
• Place biopsy in 50 ml steril conical containing 10-40 ml of PBS or saline solution at 4ºC
• Transport to lab on ice
• Pinch biopsies using large forceps generally capture the mucosal layer only and can be directly processed following the next steps. Larger tumor specimens may contain multiple layers of the colon. One layer is generally smooth, tan and does not contain blood vessels. The other layer can be red and white with lots of blood vessels. These specimens need to be processed. See Mahe, 2015 et al for detailed pictures of layers and an excellent Journal of Visual Experimentation video describing this process.
• Rinse biopsy with 4ºC PBS until PBS runs clear.
• Remove muscle fat layer with scissors or other instrument.
• If there is enough sample, save sections for RNA, DNA, protein, and formalin fixation
• If the sample is from a tumor, cut the tumor into pieces and digest in colon tumor digestion buffer for 30 min at 37ºC
• Cut into 1-2 mm strips and place in 15 ml conical with chelation solution (no EDTA) on ice
• Wash 3x with chelation solution (no EDTA) by spinning for 5 minutes at low speed, and replacing chelation solution
• Incubate in 10 ml chelation buffer (+ EDTA, + DTT) for 30 minutes on ice. Periodically disrupt by pipetting with 1 ml pipette tip
• Shake vigorously to release crypts from tissue
• Allow large fragments to settle for 1-2 min (crypts will, hopefully, remain in suspension)
• Transfer ~10 µl of supernatant to a microscope slide and inspect for presence of crypts. Proceed if crypts are present, otherwise continue incubating and disrupting.
• Place supernatant in a clean 15 ml
• Optional: Repeat process with pellet to release more crypts and then combine supernatants
• Pellet the crypts by centrifuging 650 rpm (100-200 x g) for 3 min at 4ºC
• Wash 2x in 40 ml basal ADF media (+HEPES, +Glut). Make final resuspension in ~5 ml. Count crypts using hemocytometer
• Pellet again and resuspend at 200-1,000 crypts/ml. Aliquot 6 mls into a fresh tube. This will be enough crypts to plate into 6 wells of a 24 well plate.
• Pellet again, remove supernatant and resuspend in ~100 µl matrigel
• Place 2-3 drops of cell/matrigel mixture into the middle of 6 wells of a 24 well plate
• Incubate ~20 min at 37ºC to allow materiel to solidify
• Place 500 µl of complete ADF media in each well
• Change media every 2 days

Passaging and collecting organoids

• Remove media and place 24 well plate on ice
• Add 500 µl cell dissociation media (Corning Cat # 35423: Cell Recovery Solution for Dissolving Matrigel)
• Incubate 15 min at 4ºC. Gently disrupt matrigel with 1 ml pipette
• Incubate another 15 minutes
• Scrape matrigel using pipette and remove with 1 ml pipette and place in 15 ml conical
• Combine 6 wells in one 15 ml conical and add basal ADF media (+HEPES, +Glut)
• Pellet by spinning 100 x g for 3 min at 4ºC and remove media
• Note: If there is a fuzzy layer, repeat process to get rid of the fuzzy layer
• Wash 1x with basal ADF media (+HEPES, +Glut)
• At this point pellet can processed for RNA, DNA or protein
• To passage, resuspend in matrigel and follow protocol above
• To freeze, resuspend in freezing media (6 ml media, 3 ml FBS, 1 ml DMSO)

Reagents

Chelation Buffer (500 ml)

To create chelation buffer + EDTA +DTT, combine 25 ml chelation buffer with 100 µl 0.5 M EDTA (pH 8.0) and 10 µl 1.0 M DTT. The final concentration of EDTA is 2 mM.

Tumor Digestion Buffer (10 ml)