The overall goal of this procedure is to isolate and reprogram murine amniotic fluid cells using a transposon system. The main advantage of this technique is that it is easily performed. In a therapeutic setting, cells from human disease fetuses can be differentiated to cells of interest for drug testing or tissue engineering, in order to prepare another way the patient specific therapy before childbirth.
This method can help answer key questions of programming. The transposon system is efficient in different cell types. It's more suitable for clinical approach with respect to viral vectors and allow the production of oxino-free IPS cells.
To begin the isolation, clean the abdominal wall of the dam using 70 percent ethanol. Using scissors, perform a midline laparotomy, incising the length of the abdominal wall to gain access to the abdominal cavity. Use forceps to expose the uterus and then excise it using scissors.
Transfer the uterus into a 100 millimeter dish filled with sterile 1xPBS and keep on ice. Place the dish under a stereomicroscope at 10X magnification and hold the uterus with forceps whilst removing the uterine wall with scissors, taking care to avoid damaging the fetal membranes. Collect the fetuses into a 100 millimeter dish filled with sterile 1XPBS and place on ice.
Grasp the placenta of one fetus with the fine point forceps and move it to a clean 100 millimeter dish. Here, carefully remove the yolk sack. Disrupt the amnion using the forceps and using an insulin syringe, collect 30 microliters of amniotic fluid.
Transfer the fluid to a 15 milliliter conical vial. Repeat this process for each fetus, collecting the fluid into the same vial each time. Next, wash each fetus with sterile 1XPBS supplemented with one percent fetal bovine serum and collect this into the 15 milliliter conical tube along with the amniotic fluid.
Centrifuge the tube at 145 times G for five minutes and then under a laminar flow hood, remove the supernatant. Re-suspend the pelleted cells in two milliliters of amniotic fluid culture medium. Take a zero point one percent gelatin-coated six well plate containing myomycin-C treated mouse embryonic fibroblasts or MEF and then seed two milliliters of cells onto the plate.
Cultivate the cells at 37 degrees Celsius and five percent carbon dioxide, changing the medium every other day. Continue culture for seven days until the cells are ready for transfection. To begin transfection, in a one point five millimiter microcentrifuge tube, mix one microgram of transposon plasmid with zero point five micrograms of transposase expression plasmid and zero point five micrograms of reverse tetracycline transactivator plasmid.
Dilute to 100 microliters using d-man. Add eight microliters of transfection reagent into the microcentrifuge tube containing the DNA. Incubate this mix for 15 minutes at room temperature.
Taking the six well plate with mouse AF cells, dropwise add 100 microliters of the transfection agent DNA mixture per well, distributing by gentle swirling with a final volume of two milliliters per well. Leave cells for 24 hours at 37 degrees Celsius in five percent carbon dioxide. The following day, remove the culture medium and then feed each well of cells with two milliliters of fresh iPS AF medium supplemented with one point five micrograms per milliliter of doxycycline to induce the expression of Yamanaka's factors.
Feed the cells daily with fresh dox-containing medium without passage from this point forward. Observe the cells under the florescence microscope within 24 hours of the first doxycylcine treatment to look for any mCherry expression and daily from this point on for colony formation. Once colony formation is observed, pick single colonies into 50 microliters of cell culture medium.
Next, transfer them sequentially into separate wells of a 96 well tissue culture plate with mytomycin c treated MEF. The following day, add 50 microliters of trypsin to each well and incubate for five minutes at 37 degrees Celsius to dissociate the colonies into single cells. Following incubation, pipette the liquid up and down to further disaggregate the colonies.
Transfer 50 microliters of the trypsin cell mix into a new well containing inactivated MEF and then return the cells to the incubator. When cells reach 60 to 70 percent confluence, split them into a new well of a 24 well plate and then continue incubation at room temperature. Once these cells reach 60 to 70 percent confluence, flip the cells one to two into two wells, one containing doxycycline and one without to verify that colonies will also grow in the absence of doxycycline.
Continue to maintain the IPS AF colonies and doxycycline independent medium on inactivated MEF at 37 degrees Celsius in five percent carbon dioxide. Change the medium daily until cells are ready for staining, imaging, and RNA extraction. Here, the stable doxycycline independent induced pleuripotent mouse amniotic GFP positive cells express the pleuripotency markers required to confirm successful reprogramming.
RTPCR analysis using R1 ES cells as a control showed expression of the induced pleuripotency markers in the presence or absence of doxycycline. The expression of the housekeeping gene, beta two microglobulin, was also observed in all three cell types. Teratomas isolated from mice injected with the induced pleuropotent cells were immunostained.
Detection of alpha fetal protein, alpha smooth muscle actin, and beta three tubulin in the masses confirm cell differentiation into all three germ layers, ectoderm, mesoderm, and endoderm. Further, RTPCR analysis of in vitro embryoid bodies and in vivo teratoma cells confirm the expression of germ layer specific marker genes for the mesoderm, endoderm, and ectoderm. After watching this video, you should have a good understanding of how to isolate and reprogram murine amniotic fluid cells.
This is a simple, safe procedure that can be applied also to human amniotic fluid cells from a sample obtained from routine diagnoses or cesarean sections.
