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The overall goal of this protocol is to use x laevis blastocoel roof or animal cap explants for testing the molecular mechanisms and cellular processes underlying neural development in vitro and in vivo. These methods can help answer key questions in the developmental and stem cell biology fields about fate acquisition, cell migration, and the cell autonomous and non-autonomous properties of neural cells. The presented techniques provide simple, cheap, versatile, and efficient tools for exploring the fundamental mechanism of key neural cells behaviors.

These assays allow the testing of neural fate determinals for reprogramming induced pluripotent stem cells and the mature generation of neural derived cell ties furtherized screening. Generally, individuals new to this method will struggle as the techniques require manual dexterity. I first had the idea for these methods when I was thinking of analyzing cell segregation behavior.

To flat-mount a xenopus laevis anterior neural tube use plastic transfer pipette to transfer a fixed, dehydrated and rehydrated embryo into a 60 millimeter petri dish filled to the top with PBT. Next, insert a pair of fine forceps between the eyes and the neural tube at the level of the optic stalk and detach the eyes. Then, carefulLy introduce the forceps below the ectoderm overlying the neural tube behind the head and carefully peel off the ectoderm.

After performing the appropriate NC2 hybridization labeling transfer the embryo into a new 60 milliliter petri dish filled with PBT and use fine forceps to grossly isolate the anterior neural tube from the embryo. Then, carefully remove the loosely attached notochord from below the neural tube. If necessary, the otic vesicles and the remaining parts of the mesoderm.

When all of the neural tube appendices have been severed, separate the anterior neural tube from the embryo. Then, transfer the isolated neural tube into a 1.5 milliliter conical tube filled with 50%glycerol and PBS for overnight treatment at four degrees Celsius. After a second overnight treatment in 90%glycerol, transfer the neural tube onto a microscope slide in a small amount of glycerol and use Tungsten needles to dissect the neural tubes along the dorsal and ventral midlines.

Then use reinforcement rings covered with a glass cover slip to mount the two sides of the neural tube in 90%glycerol and PBS. To prepare bead for induction of animal cap explant use a P200 micropipette to split half an aliquot of BSA treated resin beads into a new 1.5 milliliter tube. Then, replace the sterile distilled water in the new tube with 200 microliters of medium containing the experimental molecule of interest for two hours at four degrees Celsius.

At the end of the incubation, transfer blastula or very early gastrula embryos into PBS with calcium and magnesium supplemented with 0.2%BSA. Next, under a stereo microscope, pinch the vitelline membrane with the side of one pair of forceps and use a second pair of forceps to slowly peel the membrane from the vegetal side of the embryo taking care not to damage the animal side. Then, using fine forceps, cut out a small square from the animal pole of each embryo.

Using a BSA-coated pipette tip, transfer the caps into individual wells of a Terasaki multiwell plate in 0.5X modified Barth's saline, placing the pigmented animal side in contact with the round bottom of the well. Using a P20 micropipette, select the darkest pink beads from the treatment tube and place one bead onto each cap, using forceps to center the beads on the caps. When the last bead has been placed, set the plate on top of water-soaked papers and cover the plate with a plastic container before incubation.

To prepare dissociated and reaggregated animal cap explants, isolate 15 to 30 animal caps as just demonstrated. Using fine forceps to cut out a small square of tissue from the animal poles. Transfer the animal caps into an agarose coated well filled with calcium-free Holtfreter saline with their pigmented sides facing up.

After a few minutes, disaggregation should be observed. Use fine forceps to separate the pigmented layers from the rest of the tissue. And then, use a P20 micropipette to discard the pigment.

When most of the pigmented layers have been removed and the cells are completely dissociated, swirl the plate to center the cells and use a P1000 micropipette to remove as much of the medium as possible, taking care not to touch the cells. Then, add one milliliter of Holtfreter's saline with calcium to the well. Transfer the dissociated animal caps into a 1.5 milliliter tube.

After spinning down the cells, use a P1000 micropipette to carefully remove the supernatant and re-suspend the pellet in 20 microliters of Holtfreter's saline with calcium. Allow the cells to reaggregate for three to six hours at room temperature. Then slowly add one milliliter of 3/4 NAM to carefully detach the explants from the bottom of the tube and use a plastic transfer pipette to distribute the explants into individual uncoated plates.

To explant animal cap explants onto xenopus laevis neural plates, first coat 60 millimeter petri dishes with 3%agarose and 3/4 NAM into which small holes have been made. Next, transfer early neurula embryos into PBS with calcium and magnesium supplemented with a 0.2%BSA for removal of the vitelline membrane. Then fill the agarose-coated dissection dish with fresh 3/4 NAM and transfer the embryos into the dissection dish dorsal side up.

Now, transfer an animal cap reaggregated explant to the dissection plate allowing the explant to slowly sink down into the dish by gravity once the pipette tip has breached the NAM surface. Then under a stereo microscope, use rounded forceps to hold the embryo in place while using a Tungsten needle to make an incision on the neural plate for the explant. Cut out a small piece of the reaggregated animal cap explant using rounded forceps and an eyebrow knife.

Use the tissue fragment to size a piece of explant for the graft. This figure shows X laevis and X tropicalis anterior neural tubes flat-mounted as just demonstrated at stages 30, 32, and 35 after whole mount double NC2 hybridization. As observed in these images, the expression of xenopus sonic hedgehog is detected in cells in contact with beads soaked in conditioned medium supplemented with the end terminus of the sonic hedgehog gene, but not in cells in contact with beads treated with control medium.

After dissociation and reaggregation as just demonstrated, these explants with anteriorized animal cap cells were mixed with anteriorized animal cap cells expressing Otx2. As observed, these Otx2 expressing cells did not segregate from the non-Otx2 expressing cells. Both types of cells intermingled freely throughout the explant.

In contrast, in explants where Otx2 expressing cells were mixed with BAR HL-2, Otx2 and IRX3 expressing cells the multigene expressing cells formed cohesive patches that did not spread uniformly within the reaggregated explant. When grafted into the anterior neural plate of x laevis embryos as just demonstrated, mixed explants composed of cells coexpressing BAR HL-2, Otx2, and IRX3 mixed with N-terminus sonic hedgehog expressing cells exhibited indogenous sonic hedgehog expression and segregated from the anterior neural epithelial cells. Whereas, when mixed explants composed of Otx2 and N-terminus sonic hedgehog expressing cells were grafted into the x laevis neural plate, the grated cells neither expressed sonic hedgehog nor segregated from their neighbors.

Once mastered, grafting the animal cap explant onto embryonic neural plates can be performed in two to three hours depending on the number of embryos grafted. It is a very robust and very rewarding technique. When attempting this procedure, it's important to be extremely cautious about bacterial and fungal contamination.

Following the grafting procedures, other methods like whole mount in situ hybridization can be performed to follow the fates and the behaviors of the grafted cells. The developmental programs underlying neural tube organogenosis are largely conserved especially in vertebrates. Thus, information acquired in amphibians has furthered our understanding of the cellular and molecular processes underlying vertebrate development.