Isolation of single nucleated cells from human intestinal mucosa
Author: J.E. Colligan et al, Translated by Xuitao Cao et al. This experiment is from the "Compendium of Immunology Laboratory Guide".
Operation method
Isolation of human intestinal mucosal single nucleated cells Move Basic program to isolate human intestinal mucosal single nucleated cells Material Fresh samples of large or small intestine, obtained by post-surgical isolation H B S S , without calcium and magnesium, p H 7. 2 D T T - H B S S solution: 75m g D T T dissolved in 50 ml H B S S , freshly prepared E D T A - H B S S solution. Digestive enzyme solution, freshly prepared R P M I -IO complete medium VFicoll isolate, density L 076 to 1.078 g/ml 30 % Percoll solution Disposable 250 m l flat bottom plastic container with screw cap, elbow forceps and elbow ophthalmic shears IOO m m and 50 m m Plastic Petri dishes 5c m magnetic stirrer, sterile or aseptic Magnetic stirrer, room temperature and 37°C Optical microscope 250 ml - secondary Erlenmeyer flask with screw cap, sterile Surgical sterile gloves I O c m Sterile glass funnel 50 m l sterile conical-bottomed centrifuge tube Polystyrene ring (5 cm diameter, I cm height) Nitex screen (100 mesh; Tetko), cut to IO cm square size 1 . Thoroughly rinse surgically isolated samples of large or small bowel with cold tap water to remove blood and luminal contents, cut vertically along the lumen, and visualize the mucosa. According to the clinical diagnosis, anatomical localization, size and morphological features, cut an appropriate area [samples from sigmoidectomy or ileocolic resection, with an average size of (2~3) c m X (2~10) c m and a weight of about 10 g]. If the block of tissue obtained is too large, it is cut into uniformly sized samples and processed separately. 2 . The excised tissue was immersed in 100 ml of H B S S solution, placed in a disposable 250 m l flat-bottomed plastic container and quickly transferred to the laboratory. 3 . Rinse the sample with IOOml of fresh H B S S solution to remove necrotic tissue and feces. Gently blot the mucosal surface to remove excess mucus. Wash the sample again with HBCS. 4 . Lay the tissue flat on the paper with the mucosa side facing up. Gently lift the mucosa from the edge of the sample with curved-tip forceps. Starting at the edge of the lifted sample, cut the mucosa and muscle layers with curved-tip scissors. If possible, cut the muscle vertically at the location of the annular fold to obtain a sample 5 m m long and wide. Examine the sample visually to see if the muscle has been completely removed. Place the mucosal slice in a petri dish containing 100m m H B S S. 5 . Thoroughly wash the mucosal pieces in the petri dish containing fresh HBSS and transfer them to a 250-ml flat-bottomed plastic container containing 50 ml of DTT/HBSS with a tight-fitting lid. Place the container on a stirrer and stir at 60r/m i n room temperature for 30m i n to loosen residual mucous membranes and debris (the solution will gradually become mildly cloudy and contain some small floating debris). 6 . Remove the mucous membrane debris and stirrer, wash in a petri dish containing fresh HBSS, and transfer to another container containing 100 ml EDTA/HBSS. Stir at room temperature for 90 min, allowing the tissue in solution to vibrate vigorously to dislodge epithelial cells from the basement membrane. 7 . Repeat stirring once or twice, depending on the situation, until the epithelial cells are basically detached. If the solution becomes very cloudy during agitation, it should be replaced promptly. 8 . Transfer the mucous membrane sheet and stirrer to a new container containing 100 ml of HBS. Wash the mucosal slice progressively for 30 millimeters with a change of HBS until the wash solution becomes clear (typically 2 to 6 washes). Some samples of heavily inflamed tissue may show some uncertainty, and it is necessary to distinguish between expected shed epithelial cells, cells that continue to be shed, and debris to avoid loss of epithelial cells due to excessive agitation. If suspicious material appears, use a microscope to carefully examine the floating material. 9 . In a sterile 50 m m petri dish containing 5 m l of digestive enzyme solution cut mucosal pieces to a size of 5 m m with ophthalmic scissors . Pour the entire contents of the petri dish into a sterile 250 ml Erlenmeyer flask containing IOOml digestive enzyme solution and a magnetic stirrer. If the mucosal piece is large, use two or more Erlenmeyer flasks. Unscrew the cap and place on a magnetic stirrer at 37°C , 60r/m i n stirring for > 8 h . 1 0 . Wearing surgical gloves, place an IO cm diameter glass funnel on a 50 ml conical centrifuge tube. Place an IO cm2 Nitex screen under the polystyrene ring and make it secure. If two flasks are used in Step 9, two separate filter sets are required. 1 1 . Check for bacterial contamination in the Erlenmeyer flasks depending on the color and odor of the supernatant and reprepare the samples if microscopy confirms contamination. 1 2. Tilt the Erlenmeyer flask at an angle of 450 for 30 s to allow the tissue fragments to settle. Slowly pour the supernatant onto a filter. Avoid pouring tissue into the filter to avoid clogging. Replace the centrifuge tube if necessary. Repeat the filtration if the fluid is thick or contains too much debris (as seen when large samples are removed from repeatedly inflamed intestinal mucosa). 1 3 . Centrifuge rapidly at 400 g at room temperature and discard the supernatant. Resuspend all sediments (including the samples obtained in Steps 1 and 9) in RPMI-10 complete medium. The sediment tends to be large and has a reddish color due to contamination with red blood cells. 14. Count the cells (Appendix 3A) and determine the viability of the cells using the trypan blue reject assay (Appendix 3C). Note the density of cells, the proportion of dead cells, the presence of epithelial cells that may be contaminated, and the presence or absence of cell clumps. If the percentage of viable cells is <50% or there are many clumps (which may be present in inflammatory samples), filter the sample through a nylon column (see Supporting Solutions). 15 . Separate the cell suspension into several aliquots in 50 ml centrifuge tubes (each aliquot should be separated with Ficoll Density Solution, samples with 60% to 70% cell viability should be separated with 4 to 8 gradients), each aliquot should be diluted with 25 ml of HBSS. 16 . Slowly add 12 m l of Flcoll Density Solution (to avoid damaging the liquid level), cover tightly, and centrifuge at 2100 g for 5 min at room temperature. 17 . Discard the uppermost layer. There is a layer of cells at the junction of the liquid surfaces, white and loose, wider and less clearly demarcated than after P B M C separation. Carefully aspirate the cells of the junction layer with a 5 m l pipette and add them to a new 50 m l centrifuge tube, using one tube for each of the two gradients. Avoid aspirating the lower layer of Ficoll density fluid and clumps of cells on the tube wall. 1 8 . Fill the centrifuge tube with H B S S, cover tightly, turn to mix, and centrifuge at 400 g for 5 min at room temperature, discarding the supernatant. Transfer all the sediment into a centrifuge tube and repeat the centrifugation. Resuspend the sediment with 5~IOml of RPMI-IO complete medium. Cells were counted and the viability of the cells was determined by trypan blue rejection. 19. Optional: If the final cell suspension contains >20% dead cells, the amount of Ficoll's Isolation Solution is insufficient, or the cells need to be prefiltered on a nylon column prior to isolation. To obtain more viable cells, the cells can be divided into aliquots and added to 2 to 4 times the volume of Percoll (Unit 2.5) and centrifuged. For each portion, in a 50 m l centrifuge tube, the cells should be centrifuged. 3 Tubing 20m l nylon capillary 1 . Prior to filtration (see basic protocol, step 14) transfer the cells into 50 ml centrifuge tubes (< 2 X 108 cells/column). Fill with HBSS and mix with repeated inversions. Remove large cell clumps to prevent excessive cell adhesion that could clog the nylon column. 2 . Attach a 20mL nylon capillary column to a 3-way tube and secure to an upright ring stand. A new 50 ml centrifuge tube is attached to the bottom of the column to collect the cells. 3 . Open the 3-way tube and allow the cell suspension to enter the nylon column in an orderly fashion. Wash the column with HBSS and replace the tube with a full one if it is full until a total volume of 100mL has been collected. The nylon wool matrix should be kept completely moist. If the flow rate is slow, agitate the top of the column with a sterile Pasteur pipette (to avoid resuspending the entire column matrix). 4 . Centrifuge at 400 ppm for 5 min at room temperature, discard supernatant, and resuspend precipitate with HBSS. Adjust the volume for Ficoll separation (see basic protocol, step 15). 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2 0 . Use the cells immediately or resuspend the cells in R PMI-IO complete medium and store the cells temporarily at 4°C (not overnight).