Biology
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Isolation of Myoepithelial Cells from Adult Murine Lacrimal and Submandibular Glands
Chapters
Summary June 11th, 2019
The lacrimal gland (LG) has two cell types expressing α-smooth muscle actin (αSMA): myoepithelial cells (MECs) and pericytes. MECs are of ectodermal origin, found in many glandular tissues, while pericytes are vascular smooth muscle cells of endodermal origin. This protocol isolates MECs and pericytes from murine LGs.
Transcript
This protocol is significant because it allows the separation and isolation of two smooth muscle cell lines, the myoepithelial cells and pericytes. This method combines genetic labeling of smooth muscle cells via cell surface markers to purify populations of myoepithelial cells and pericytes. This protocol allows comparison the function and regenerative abilities of healthy and diseased myoepithelial cells and processing these cells for gene expression studies.
Myoepithelial cells exist in other exocrine glands such as mammary, salivary, and pancreas, which allows this protocol to be adapted by isolation of myoepithelial cells and pericytes from any other tissue. To label the alpha smooth muscle actin or SMA expressing cells, inject three to four-week-old tamoxifen-inducible alpha SMA driven reporter mice with 100 microliters of tamoxifen per 20 grams of body weight intraperitoneally once a day for two days. For lacrymal gland collection, two to three days after the last injection, use tweezers to gently pull one gland while at the same time scratching the connective tissue around the gland with the sharp tip of a pair of small scissors to free the gland.
When the lacrymal and parotid glands have been separated, use scissors to cut the lacrymal gland out and place the glands into a 35 millimeter dish with two milliliters of cold PBS on ice. Place the dish under a dissecting microscope and trim any surrounding fat and connective tissue. Then remove the lacrymal gland capsule with two forceps.
When all of the glands have been harvested, check a small piece of tissue under a fluorescent microscope to confirm cell labeling. To prepare a single-cell suspension, transfer all of the lacrymal glands to a 35 millimeter dish containing 0.5 milliliters of room-temperature digestion medium and use small scissors to mince the glands into 0.2 to one micrometer square pieces. Using a wide bore pipette filter tip transfer the minced tissue into a two milliliter round-bottom tube and bring the volume in the tube to two milliliters with digestion medium.
Invert the tube to mix, and place the tube into a 37 degree celsius shaking incubator for 90 minutes at 100 to 120 rotations per minute. Every 30 minutes slowly triturate the gland pieces 20 to 30 times using 1000 microliter filter tips with a decreasing bore size. At the end of each trituration, inspect a 10 microliter aliquot under a microscope for clusters.
After 90 minutes pass the sample two to three times through an insulin syringe needle to further release the cells into suspension and transfer the cell solution to a 15 milliliter tube. Add Blocking Medium Type 1 to a total of five milliliters and mix the tube two to three times by inversion. Pass the cells through a 70 micrometer cell strainer into a 50 milliliter tube, and wash the strainer with one milliliter of Blocking Medium Type 1, before straining the cells through the filter one more time.
Next collect the cells by centrifugation and resuspend the pellet in two milliliters of blocking Blocking Medium Type 2. Transfer the cells to a two milliliter microcentrifuge tube and collect the cells by centrifugation. Resuspend the cells in one milliliter of Accutase solution for a two to three minute incubation at 37 degrees celsius and 100 to 120 rotations per minute.
At the end of the incubation, transfer the cells to a 50 milliliter tube and add up to 10 milliliters of Blocking Medium Type 1. After centrifugation resuspend the cells in six milliliters of recovery medium for a 30 minute incubation at room temperature. At the end of the incubation, check a 10 microliter sample under the microscope to ensure complete cell dissociation.
After counting, collect the cells by centrifugation. To stain the cells for Fluorescence-activated Cell Sorting, or FACS, add up to five times 10 to the five cells per two milliliter tube containing 400 microliters of FACS buffer and add five microliters of Brilliant Violet 421 anti mouse CD326 and 0.5 microliters of Ghost Red 780 Viability Dye. After thorough mixing, wrap the tubes with foil for a 45 minute incubation at 4 degrees celsius with rotation.
At the end of the incubation, collect the cells by centrifugation and resuspend the pellets in one milliliter of FACS buffer per sample. Transfer the cell samples to individual five milliliter FACS tubes on ice and adjust the compensation using single color controls. Then sort the cells at 20 pounds per square inch through a 100 micrometer nozzle.
Microvascular pericytes develop around the walls of capillaries and demonstrate a square shape. Myoepithelial cells surround the lacrymal secretory acini, have long processes, and occupy a relatively large area. Here, cells dissociated from a lacrymal gland in preparation for Fluorescence-activated Cell Sorting, as just demonstrated, are shown.
For flow cytometric sorting of myoepithelial cells and pericytes the cells should first be gated by their forward and side-scatter areas before excluding any doublets. After gating to exclude the dead cells, control samples are used to determine the background noise and to establish a proper compensation for an optimum separation between signals. Remember to keep the number of the glands and digestion medium volume as described in protocol, and to check the efficiency of tissue digestion at all steps.
The isolated cells can be used for multiple downstream application including in vivo/in vitro experiments, for example cell culture, transplantation, metabolomics, proteomics, and single-cell analysis.
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