Our method is significant because it allows researchers to isolate and manipulate different cell types for investigation of their individual lineage contributions to tissue form and function. This technique is a gentle and efficient method for separating cell types and as a result, the integrity of the cells is preserved for 3D culture. This technique can also be applied to other systems to isolate cells that don't have well-characterized biomarkers.
To harvest the number two, three, four, and five mammary glands from a 10 to 14-week-old female mouse, first make a one centimeter incision at the midline between the two hindlimbs. Extend the cut up to the neck and make small lateral cuts from the midline incision toward the limbs. Then stretch the skin taught before securing it with a pin on each side of the animal.
Using forceps, collect the lymph nodes from the number four glands. To remove the mammary glands, use sharp scissors to cut under each region of tissue, placing the tissues in 50 milliliters of four degrees Celsius DMEM-F12 supplemented with 5%FBS and antibiotic-antimycotic as they are harvested. When all of the tissues have been collected, chop the glands until the pieces can fit easily through a one milliliter pipette tip rotating the plate as necessary so that all of the tissues are minced evenly.
Then transfer the fragments in three milliliters of digestion medium into a single well of a six-well low adhesion dish for 14 hours at 37 degrees Celsius and 5%carbon dioxide. The next morning, gently pipette the tissues 10 times with a one milliliter micropipette and transfer the tissue fragments into a 15 milliliter tube. Collect the tissue by centrifugation and resuspend the pellet in five milliliters of fresh DPBS.
Filter the suspension through a pre-wet 70 micrometer nylon cell strainer, rinsing the tube in the strainer four times with 10 milliliters of 37 degrees Celsius DMEM-F12 per wash. To release the tissue fragments, hold the strainer tab with gloved fingers and invert the strainer over a 60 millimeter tissue culture dish. Pass four one milliliter aliquots of maintenance medium through the bottom of the strainer and quickly examine the dish for single cells, fat droplets, and contaminating tissue under an inverted microscope.
Then place the plate in the cell culture incubator for 24 hours to allow the tissue fragments to adhere to the dish and to generate bilayered fragments. To separate the myoepithelial cells from the luminal epithelial cells, rinse the dish with one milliliter of DPBS before adding one milliliter of fresh 0.5%trypsin-EDTA to the cells. Carefully monitor the digestion under an inverted microscope.
The outer layer of myoepithelial cells will begin to detach within three to six minutes. When the myoepithelial cells have detached, transfer the supernatant into a 15 milliliter tube containing two milliliters of 10%FBS in DPBS. Without disturbing the luminal epithelial cells, gently rinse the dish with two milliliters of DPBS before adding a fresh milliliter of trypsin-EDTA to the remaining cells.
After seven to 15 minutes, quench the enzymatic reaction with two milliliters of 10%FBS in PBS and transfer the cells to a new 15 milliliter tube. It's critical to closely monitor the cells during the differential trypsinization and to not over digest the cells. When both fractions have been collected, centrifuge the cells to remove any residual dissociation reagent and resuspend the pellets in 250 microliters of maintenance medium per tube.
After counting, dilute each cell population to a 1.2 times 10 to the four cells per well concentration in fresh maintenance medium. Add 90 microliters of 50%extracellular matrix in DMEM-F12 without phenol red to each well of an eight-well chamber slide. When all of the wells have been coated, solidify the base layer in the cell culture incubator for 30 minutes.
During this polymerization, collect the cell fractions by centrifugation and resuspend each population in 100 microliters of 10%extracellular matrix in 90%growth medium per well. Next, add 100 microliters of each cell suspension to each well and allow the co-cultures to settle for 20 minutes in the cell culture incubator. At the end of the incubation, carefully add 100 microliters of growth medium down the side of each chamber wall and return the slide to the cell culture incubator.
Image the cells every 24 hours to track their growth and gently renew the growth medium every two to three days. To fix the organoids for immunostaining, at the appropriate experimental time point, carefully aspirate the medium from each slide to be imaged and rinse each well with 200 microliters of DPBS per well. Fix the organoids with 200 microliters of four degrees Celsius paraformaldehyde for 10 minutes at room temperature before treating the organoids with 200 microliters of 0.2%glycine in DPBS per well for 30 minutes at room temperature.
At the end of the incubation, permeabilize the organoids with 0.25%triton X-100 in DPBS for 10 minutes at room temperature followed by the blocking of nonspecific binding with 5%donkey serum or the appropriate serum that matches the species of the secondary antibody in DPBS for one hour with rocking. Then label the cells with 125 to 200 microliters of the appropriate primary antibodies of interest in 1%donkey serum in DPBS overnight at four degrees Celsius with rocking. The next morning, wash each well two times with 200 microliters of DPBS plus triton X-100 before labeling the organoids with the appropriate fluorescence conjugated secondary antibodies for 45 minutes at room temperature with rocking protected from light.
Then wash each well two times with fresh DPBS plus triton X-100 as demonstrated and image the organoids by fluorescence microscopy according to standard protocols. After 24 hours of incubation, purified epithelial fragments adhered to the bottom of the culture dish forming flat pancake-like structures with an outer layer of myoepithelial cells encircling an inner layer of luminal epithelial cells. Trypsin treatment differentially detaches the cells with the myoepithelial cells detaching first and appearing as bright rounded cells that encircle the core of remaining cuboidal luminal epithelial cells.
The overall purity of the two cell compartments is about 90%as assessed by the keratin-14 and E-cadherin expression within the two populations. After 10 days of culture in 10%extracellular matrix on a 50%extracellular matrix base as demonstrated, the myoepithelial luminal epithelial cell organoids formed large branched structures with well-developed lumens. Differentiation at day five with the addition to alveologenesis medium induces the formation of larger, more branched milk-containing organoids.
It is important to remember to wash the strainer carefully when collecting the epithelium to ensure that there are no stromal contaminants. To query the changes in gene expression, researchers can collect RNA from the organoids by releasing them from the extracellular matrix using a recovery solution. Paraformaldehyde is considered hazardous and researchers should use personal protective equipment and work inside a fume hood when using this reagent.
