To begin, prepare a 1.8%agarose plate with HEPES Buffered E3 medium or E3.While working under a dissection microscope, place six to 10 anesthetized larvae in a row on the agarose plate. Put one insect pin on the head of each larvae. Then remove the remaining E3 from the plate using a tissue.
Using another insect pin, isolate the intestine from a larvae without disturbing any other organs. Ensure proper removal of yolk. Once isolated, inspect the intestine and remove all non-intestinal material, such as skin, fat, or liver.
Use tweezers to collect the intestine and transfer it to PBS containing 10%FCS in a microcentrifuge tube placed on ice. Immediately after the dissection of all intestines, centrifuge the microcentrifuge tube at 13, 800 g for 30 seconds. Remove the supernatant, leaving about 100 microliters in the tube to prevent the intestines from drying out.
Add 500 microliters of papain solution to the intestines in the tube. Activate the papain by adding 2.5 microliters of one molar cysteine. Then incubate the tube containing the intestines in a water bath at 37 degrees Celsius for 10 minutes.
Pipette the contents up and down halfway after five minutes to stimulate enzymatic tissue digestion. Transfer the dissociated cells into a fluorescence-activated cell sorting, or FACS, tube through a pre-wedded 35-micron cell strainer. Wash the strainer several times by adding 0.5 milliliters of PBS containing 10%FCS to a total wash volume of two milliliters.
Centrifuge the collected filtrate at 700 g for five minutes at 4 degrees Celsius, and remove the supernatant. Add one microgram per milliliter DAPI to 300 microliters of PBS containing 10%FCS. Add this mixture to the pellet and resuspend to label the dead cells.
Then incubate for five minutes on ice to allow their exclusion during subsequent FACS analysis.
