This article describes a protocol for differentiating human induced pluripotent stem cells into neuronal and glial cells using embryoid bodies (EBs). The process involves culturing EBs in a matrix-coated dish and utilizing a neuroepithelial induction medium to facilitate cell differentiation.
Take embryoid bodies, or EBs, derived from human induced pluripotent stem cells. These EBs are composed of three germ layers.
Transfer the EBs to a matrix-coated culture dish containing a suitable medium and incubate.
The matrix enables the EBs to attach to the culture dish.
The growth factors and nutrients in the medium facilitate the ectodermal cells of the EBs to form neuroepithelial aggregates arranged in a circular pattern termed a rosette.
Cut the rosette fragments and detach them from the dish.
Collect the fragments and spin them down.
Discard the supernatant and resuspend the fragments.
Using a pipette, move the fragments up and down to dissociate them.
Add the desired number of cells to a matrix-coated well plate containing a differentiation medium.
The components of the differentiation medium facilitate the neuroepithelial cells to transform into fully functional neuronal and glial cells, the primary cells of the nervous system.
On day two, aspirate standard matrix coating solution from a previously prepared 60-milliliter dish, and add 5 milliliters of complete neuroepithelial induction medium or NRI to the dish. Working under the stereoscopic microscope at 4x magnification, and using a 200-microliter pipette, collect around 50 floating embryoid bodies and transfer to one coated dish. Then, incubate the dish at 37 degrees Celsius and 5% carbon dioxide.
The next day, day three of the differentiation procedure, check the dish under the microscope at 10x magnification to ensure that the embryoid bodies are all attached. Then, gently perform a total medium change with complete NRI medium. Change the NRI medium every other day up to day seven when rosette-like neuroepithelial aggregates should be visible.
On day seven, coat a 96-well plate by pipetting 100 microliters of standard matrix diluted in culture medium into each well. On day eight, cut the rosette-like aggregates into fragments under a stereoscopic microscope at 10x magnification in sterile conditions.
Note that the rosettes tend to easily detach from the dish when touched with the needle. In this case, partially disaggregate the detached rosette with the needle tip. If the rosettes will remain partially attached, use a 200-microliter pipette to complete the detachment of the rosette fragments.
Rosette cuttings require good manual skills and precision in order to avoid cutting non-neuroectodermal derivatives.
Next, collect the rosette fragments and their medium into a 15-milliliter conical tube. Rinse the dish with 2 milliliters of NRI medium to recover all fragments. Then, spin down the rosette fragments at 112 times g for one or two minutes.
After aspirating the supernatant, gently resuspend the pellet in 1 milliliter of pre-warmed 1x DPBS without calcium and magnesium. Gently pipette the rosette fragments up and down to partially dissociate them. Then, add 4 milliliters of complete NRI medium and count the cells using trypan blue and an automated cell counter.
Next, aspirate the standard matrix coating solution from the 96-well plate, and deposit the cells into the wells at a density of around about 15,000 cells per square centimeter in NRI medium. Incubate the plate overnight at 37 degrees Celsius and 5% carbon dioxide. On day 10, perform a total medium change using complete neuronal differentiation medium.
繁體中文
