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Isolation of mouse intestinal crypts and organoid culture, adapted from (Sato and Clevers 2013) is schematically represented in Fig. 22.2. A detailed protocol of the culture protocol is described below.
Small Intestinal Crypt Isolation
Thaw aliquots of Matrigel™ Basement Membrane Matrix (BD Biosciences) on ice before isolation and pre-incubate a 24-well plate in a CO2-incubator (5 % CO2, 37 °C). Matrigel™ is a solubilized gelatinous protein mixture and is derived from the Engelbreth–Holm–Swarm (EHS) mouse sarcoma cells. The extract resembles the complex extracellular environment found in many tissues and is commonly used as a substrate for culturing cells. At room temperature, Matrigel™ polymerizes into a three dimensional structure that is useful for both cell culture and studying cellular processes in three dimensions, including cell migration.
Fig. 22.2 Schematic summary of intestinal epithelial crypt isolation and subsequent organoid cultures
Isolate a piece of mouse small intestine (at least 5 cm) and open longitudinally. Wash the intestine with ice cold PBS until most of the luminal contents are cleared and scrape off the villi using a coverslip. After another washing with ice-cold PBS, cut the intestine into 2–4 mm pieces with scissors and transfer them to a 50-mL tube. Add 30 mL ice-cold PBS and wash the fragments by gently pipetting the fragments up and down with a 10 mL pipette and discard supernatant after settling down. Repeat this step 5–10 times until the supernatant is almost clear. Next, add 30 mL of ice-cold crypt isolation buffer (5 mM EDTA in PBS; one can also use higher EDTA concentration, up to 10 mM if yields are low) and gently rock the tube at 4 °C for 30 min. Keep pipetting up and down the tissue in solution every 10 min in order to release as much epithelium as possible. After settling down the fragments, the supernatant can be removed and 20 mL of ice-cold PBS can be added to wash the fragments using a pipet. Repeat this procedure when the fragments are settled down, in order to release most of the crypts, that will also settle down after some time. Villous fractions present in the supernatant are discarded and crypts fractions are passed through a 70-mm cell strainer and collected into 50 mL tube(s), after adding 5 % FBS to a crypt solution. Spin down the crypt fractions at 300×g 5 min and resuspend the pellets in 10 mL of ice-cold basal culture medium (advanced Dulbecco's modified Eagle medium/F12 supplemented with penicillin/streptomycin, 10 mmol/L HEPES, Glutamax, 1 × N2, 1 × B27, and 1 mmol/L N-acetylcysteine). The suspension is transferred to a 15 mL Falcon tube and centrifuged at 150–200×g for 2 min, to remove single cells (mostly lymphocytes), which end up in the supernatant.
This washing step should be repeated 2–3 times until most single cells are cleared. After the washing steps, the number of crypts can be calculated (take 10 μL of the crypts suspension and count numbers of crypts using a heamocytometer by inverted microscopy; the total number of crypts = the number of crypts counted × 1,000). It should be noted that at this point it is difficult to distinguish true crypts from other epithelial debris, hence the count is only a rough estimate.
Small Intestinal Organoid Culture
Centrifuge small intestinal crypts, and remove supernatant as much as possible to avoid dilution of Matrigel™ in the next step. Keep the tube at 4 °C and resuspend the crypts pellet in Matrigel™ (200–500 crypts/50 μL matrigel). Pipet 50 μL of the crypt-Matrigel™ suspension into the pre-warmed 24-well plate. The suspension should be applied on the center of the well so a hemispherical droplet can be formed. Transfer the plate back into the CO2 incubator (5 % CO2, 37 °C) as soon as possible after the seeding. Allow the Matrigel™ to solidify for 5–10 min and add 500 μL complete organoid culture medium per well. For preparation of the organoid culture medium (e.g. amount of growth factors) check Table 22.1. Place the 24-wells plate in a CO2-incubator and refresh the culture medium every 4–6 days.
Small Intestinal Organoid Passage
Crypt organoids can be passaged 7–14 days after seeding. Place the plate with organoids on ice to allow the matrigel to dissolve. Remove the culture medium and gently break up the Matrigel™ by pipetting with a 100–1,000 μL-pipette. Resuspend the organoids in 1–2 mL basal culture medium and transfer them to a 15 mL-tube. Gently disrupt the organoids using a fire-polished Pasteur pipette to remove released dead cells and single cells. Pellet everything by centrifuging at 300×g for 5 min, and wash the organoids afterwards with 10 mL basal culture medium. Pellet again at the same speed and add the correct amount of Matrigel™, namely 50 μL per well that you would like to plate. Count the “crypts/immature organoids” as described before (by hemocytometer) and pipet the Matrigel™ suspension with crypts (50 μL per well) into a new 24 culture plate and place the plate back in CO2 incubator (5 % CO2, 37 °C) to culture the organoids.
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