We focus on the seafaring the mechanisms, controlling early cell fate decisions during embryonic stem cell differentiation, both in vitro and in mouse models. Complex regulatory networks, sky differentiation during development, and we investigate how the interplay of signaling pathways, transcription factors, and epigenetic regulators promote different sulfates. Studying the dynamic and rapid process of gastrulation in mice is technically challenging due to the small number of cells per embryo, and the limited number of embryos with the desired genotype per litter.
While literature extensively shows the study of single cells using wild-type or fluorescent-activated cell sorted populations of embryos, a comprehensive analysis of embryos with lineage mutations is still limited. Our protocol offers the possibility of generating single-cell omic datasets from gastrulating embryos harboring lineage mutations. It will help scientists with none or limited mouse handling experience or who wanna learn early embryo manipulation or single-cell approaches.
Our protocol will help answer new scientific questions in gastrulation and early organogenesis, including heart development. We provide a pipeline to analyze lineage specific neuron embryos at single-cell solution, which will facilitate analysis or the role of a specific genes in lineage commitment. We hope to use this methodology to understand the regulatory mechanisms of sulfate decisions during the rapid and dynamic process of gastrulation.
Prior to starting the Murine Embryo isolation, clean the area thoroughly with 70%ethanol. Ensure all dissection tools are washed and sterilized. Obtain five milliliters of sterile DMEM, containing 10%FBS, and 20 milliliters of DPBS, and place them on ice.
Next, to perform the embryo isolation using a stereo microscope with a transmitted light stage and camera, place the properly euthanized pregnant dam on its back, and sterilize the area near the vaginal opening with ethanol. Using dissection scissors and tweezers, lift the skinfold near the vaginal opening and make a small V-shaped cut, slowly revealing the uterus of the pregnant dam. Then dissect out the uterine horn of the dam by holding one end of it with tweezers and cutting along it, making sure to remove the cervix.
Place the uterine horn into a 10-centimeter Petri dish containing DPBS on ice. With dissection scissors and tweezers, remove the uterine muscle along the implantation sites and cut each implantation site or pearls containing the deciduous swellings inside. Place it into fresh DPBS in a six-centimeter Petri dish on ice.
Take one implantation site, place it in a new six-centimeter Petri dish on top of the stereo microscope stage, and add 500 microliters of DPBS to it. Adjust the focus of the microscope and the light source. Hold down the implantation site with one set of tweezers in one hand.
And with the other hand, slowly insert another pair of forceps into the end of the implantation site, cut from the uterine horn, gradually revealing the deciduous swelling. To reveal the embryo, hold the antimesometrial end of the isolated deciduous swelling with one pair of forceps. And with the other pair, slowly make a horizontal cut, about one quarter of the size of the deciduous swelling from the mesometrial end.
Now with both forceps, slowly push from the anti-mesometrial and of the deciduous swelling until the embryo pops out from the freshly cut mesometrial end. Once the embryo is revealed, proceed to remove any extra embryonic tissues attached. If the parietal endodermal sac and ecto placental cone do not spontaneously come off the embryo, use a pair of forceps to remove them along with any associated maternal blood.
Then while holding down the embryo with one pair of forceps, slowly peel the visceral yolk sac from the embryo using another pair of forceps. Locate the parietal endodermal sac and ectoplacental cone. Using a P20 pipette, transfer the yolk sac with no more than 10 microliters of DPBS from the dish into an eight-strip PCR tube on ice.
Take bright field pictures of freshly isolated embryos to ensure the staging of the littermates is similar. With a P200 pipette, slowly transfer the embryo with 50 microliters of DMEM containing 10%FBS into a 1.5 milliliter tube kept on ice. Repeat the same for all deciduous swelling using clean tools and new plastics for each isolation.
After isolating the visceral yolk sac from the mouse embryo, proceed to digest each yolk sac in an eight-strip polymerase chain reaction or PCR tube. Using a P20 pipette, add 19.3 microliters of PCR template DNA lysis buffer, and 0.7 microliters of 0.2 micrograms per milliliter proteinase K to each yolk sac sample. Vortex the sample for 10 seconds and using a mini centrifuge with a strip adapter, spin down the samples at 1, 000 G for 10 seconds.
Place the eight-strip PCR tube in an 85-degree Celsius heat block for 45 minutes while vortexing for five seconds every five minutes. After 45 minutes, spin the tube strip down again at 1, 000 G for 10 seconds before proceeding with PCR for desired genetic identification. To perform the PCR reaction for Cre genotyping, prepare a PCR master mix containing the displayed components for eight-strip PCR tube for each yolk sac.
Proceed to run the PCR thermal cycle amplification program using the displayed cycle, then run the PCR products on a 1%agarose gel to draw genotyping conclusions. Store the remaining digested yolk sac samples in a minus 20-degree Celsius freezer for long-term storage. When the PCR product was separated on an agarose gel, expected fragment sizes for the locks p and wild-type alleles, as well as the Cre allele resin.
Proceed to perform the Cell Dissociation of Isolated Murine Embryos once the genotypes have been confirmed. Using a P200 pipette, add 50 microliters of DMEM with 10%FBS to a new 1.5 milliliter tube. After pooling embryos with the same genotype into the new tube, place it on ice.
Allow the pooled embryos to settle to the bottom of the tube, then wash the embryos using 50 microliters of DPBS and wait for them to settle before removing as much DPBS as possible without removing the embryos. Next, add 100 microliters of trypsin to the pooled embryos and incubate at 37 degrees Celsius in a heat block for five minutes. Gently flick the 1.5 milliliter tube every 30 seconds to help the cells dissociate.
After five minutes, neutralize the trypsin with 300 microliters of DMEM containing 10%FBS. Centrifuge the embryos at 100 G for four minutes at room temperature. Resuspend the obtain small palate in 40 microliters of DMEM with 10%FBS and place the tube on ice.
Mix five microliters of the cell suspension with five microliters of trip and blue in a new 1.5 milliliter tube, and pipette the mixture up and down thoroughly. Transfer the mixture onto a slide to determine the cell number and viability using an automated cell counter. Finally, proceed to single-cell partitioning using a microfluidic chip following the manufacturer's protocol.
