We developed a procedure for isolation of tracheal brush cells, rare specialized chemosensory epithelial cells from choline acetyltransferase fluorescent reporter mice. This method is based on an initial separation of tracheal epithelium from the submucosa allowing for a subsequent shorter incubation of the epithelial sheet with papain. This allows for robust recovery of tracheal brush cells and has been successfully used for isolation and transcriptional analysis of brush cells by RNA sequencing.
Long incubation with digestive enzymes can decrease cell viability and alter the transcriptional profile of cells isolated from tissues. Here we separate the tracheal epithelium with dispase and subsequently incubate it with papain for a short amount of time producing high yields of viable brush cells. Our method can contribute to studies of upper airway epithelium.
The same protocol can be applied to isolating brush cells from other mucosal sites such as nasal tissue. Demonstration of our highly reproducible technique will allow peer scientists to isolate and further investigate tracheal brush cells. To begin this procedure, add dispase and DNase I to PBS at the final concentrations shown here to prepare the dispase digestion solution.
Make sure that the dispase powder is fully dissolved before warming up the solution in a water bath at 37 degrees Celsius. Next, add 5%heat-inactivated FBS to DMEM to prepare a stopping solution. To prepare the Tyrode I buffer, add papain and L-cysteine to HEPES Tyrode's buffer without calcium.
To prepare the Tyrode II buffer, add leupeptin to HEPES Tyrode's buffer without calcium at a concentration of two microliters per milliliter. To prepare the FACS buffer, use Hank's Balanced Salt Solution without calcium, magnesium and phenol red supplemented with two millimolar EDTA and 2%FBS. After euthanizing the mouse, use 21 gauge needles to fix it on a surgical board in the supine position with extended upper and lower extremities.
Spray the fur with 70%ethanol to sanitize the area. Using straight forceps, lift the skin and fur of the abdomen and use dissecting scissors to make an incision in the center. Using the scissors, separate the skin from the subcutaneous tissue from the abdomen to the mandibula.
While holding the subcutaneous tissue up with the forceps, make a small incision with the scissors in the center of the abdominal wall. Then open the peritoneum with a V-shaped incision. Using the forceps, gently move the small intestine to the side.
Locate the abdominal aorta and vena cava and make an incision with the dissecting scissors to allow for rapid exsanguination. Locate the diaphragm. Using an 18 gauge needle, make an opening in the diaphragm just below the sternum to deflate the lungs.
Using sharp pointed straight dissecting scissors, cut along the base of the ribs to carefully separate the diaphragm from the ribcage. Use the forceps to lift the exposed end of the sternum and cut the sternum longitudinally from the base of the ribcage to the neck. Use short straight scissors to make a central cervical incision and separate the two lobes of the submandibular gland.
After this, use a pair of fine point high precision forceps to carefully remove the surrounding connective tissue and the thymus overlying the carina. Dissect the trachea free by first separating the proximal end at the level of the epiglottis and then dissecting the distal end at the level of the bifurcation of the trachea. Locate the epiglottis and cut the trachea longitudinally from the epiglottis to the carina.
To begin, place the trachea into a 1.5 milliliter tube containing 750 microliters of dispase digestion solution pre-warmed to 37 degrees Celsius. Cover the tube with aluminum foil to reduce the exposure to direct light and incubate on a shaker at 200 RPM and at room temperature for 40 minutes. Then add 750 microliters of DMEM with 5%FBS to stop the reaction and place the tube on ice.
Transfer the trachea to a Petri dish. Orient the trachea with the epithelial side facing up. The longitudinally dissected trachea has a semi-cylindrical shape maintained by the cartilaginous rings.
The epithelium is on the concave surface. Using straight forceps, tether the epiglottis area of the trachea to the Petri dish and use a size 22 disposal scalpel to scrape the epithelium off the trachea. The epithelial layer will separate as a translucent sheet.
Use the scalpel to mince the epithelium and transfer the epithelial layer to a two milliliter tube. Rinse the Petri dish with 750 microliters of Tyrode I buffer and transfer this to the tube containing the epithelial layer. Cover the tube with aluminum foil to reduce the exposure to direct light and incubate at 37 degrees Celsius on a shaker at 200 RPM for 30 minutes.
Then add 750 microliters of cold Tyrode II buffer. Vortex the digested tissue vigorously for 20 to 30 seconds. Using a syringe attached to an 18 gauge needle, triturate the homogenate 10 times.
After this, switch to a 21 gauge needle and triturate 10 to 20 more times. Filter the cells through a 100 micrometer strainer into a 50 milliliter conical tube. Add cold FACS buffer at a volumetric ratio of 30 to one.
Centrifuge at 350 times g and at four degrees Celsius for 10 minutes. Discard the supernatant and resuspend the pellet in cold FACS buffer. Transfer this suspension to a 12 by 75 millimeter polystyrene tube.
Centrifuge again at 350 times g and at four degrees Celsius for 10 minutes. Discard the supernatant and resuspend the pellet in 100 microliters of FACS buffer. Next, add one microliter of anti-mouse CD1632 blocking antibody to block non-specific binding and incubate on ice for 15 minutes.
Then add antibodies in the respective isotype controls as outlined in the text protocol. Incubate on ice for 45 minutes while protected from direct light. After this, add 4.5 milliliters of cold FACS buffer and mix.
Centrifuge at 350 times g and at four degrees Celsius. Discard the supernatant and resuspend the pellet in 300 microliters of cold FACS buffer. Add propidium iodide immediately before flow cytometric sorting.
In this study, tracheal brush cells from ChAT eGFP acetylcholine fluorescent reporter mice are successfully isolated for RNA sequencing. Cells are identified from debris by forward and side scatter angle. Doublets are excluded using forward scatter height and width and side scatter height and width.
The doublets are the cells that have high width values. Within the single cells, the live cells are identified as the population that is propidium iodide negative. Within the live single cells, the CD45 low to negative cells are identified based on the isotype control.
Within the CD45 low negative cells, the EpCAM positive cells that are also eGFP positive in the FITC channel are the population of brush cells. Brush cell numbers are altered by exposure to microbial metabolites and protozoa and therefore reflect the variability of institutional microbiota. Therefore, we would suggest estimating the number of brush cells by fluorescence microscopy to gauge the expected number of isolated brush cells by flow cytometric sorting.
Make sure to properly separate the epithelial layer of the trachea as it determines the yields of isolated brush cells. The isolated tracheal brush cells can be used in a broad array of assays such as RNA sequencing, cell culture and functional analysis.
