Isolation and Expansion of the Adult Mouse Neural Stem Cells Using the Neurosphere Assay

Overall goal of the following experiments is to isolate and expand adult mouse neuro stem cells using the neuro sphere assay. This is achieved by first obtaining adult mouse brains, followed by dissection of the brains under the microscope, removal of the septum and isolation of periventricular tissue. Then the tissue is minced and tryps inized to achieve single cell suspension.

The cells are then plated in neuro stem cell media for culture. Hi, my name is Hassan Ali and I'm working with Dr.Brent Reynolds at the University of Florida McKnight Brain Institute. Today I'm gonna demonstrate how to isolate and expand neural stem cells from the adult mouse brain.

Let's go to the lab and get started. Anesthetize two to four adult mice, perform cervical dislocation and rinse the head with 70%ethanol. Decapitate the animal and use small pointed scissors to make a median codal rostral cut and expose the skull.

After removing excess cervical tissue, a coronal cut is made between the orbits by placing each blade of the pointed scissors in the orbital cavity. Using the foramen magnum as an entry point, make a longitudinal cut through the skull along the sagittal suture. Be cautious not to damage the underlying brain by ensuring the blades stay as shallow as possible.

Again, using the frame and magnum, make a cut along the lateral aspects of the skull as well. Using curved forceps, grasp and peel the skull overlying each hemisphere outward to expose the brain. Then using a small spatula, scoop the brain into a 50 ml tube containing hem.

Repeat this procedure until all of the brains have been harvested. After transferring the brains to the PC two hood, remove the hem from the 50 ml tube and replace with fresh hem to remove contaminants like blood and hair. Perform three washes like this.

Then transfer the brains to a 10 centimeter Petri dish containing hem. Place the brains under the dissecting microscope and position each brain flat on its ventral surface by holding it from the coddle side with fine curved forceps. Using another set of forceps, remove olfactory bulbs.

Then rotate the brain to expose the ventral aspect and make a coronal cut perpendicular to the brain at the level of the optic chiasm. Place the roel part of the brain in a separate Petri dish with hem. Place the rostral part of the brain so you can see the cut surface.

Using pointed forceps, hold the brain and with another set of forceps, remove the septum. Identify the striatum corpus callosum and periventricular area using fine forceps. Dissect out the thin layer of tissue surrounding the lateral ventricles from each side.

Be sure to exclude the sal parenchyma and the corpus callosum. Pull the dissected tissue in a newly labeled 10 centimeter sterile glass petri dish. Repeat this procedure until all brains are micro Dissected.

Mince the Pooled tissue for one to two minutes using a scalpel blade until only very small pieces remain. Using one ml of prewarm trypsin, wash the scalpel blade to collect any attached tissue. With another ml of trypsin.

Wash the base of the Petri dish to collect the tissue. Transfer the tissue to a 15 ml conical two in order to leave no tissue behind. With the third ml of trypsin, wash the base of the Petri dish for any additional tissue to transfer to the 15 ml Tube.

Place the tube in a 37 degree water Bath for seven minutes. At the end of the enzymatic incubation, add an equal volume of trypsin inhibitor to the digestive tissue, pipette the suspension up and down to ensure trypsin, inactivation, and then pellet the tissue suspension by centrifugation at 700 RPM or 110 G for five minutes. Vacuum off the supernatant and discard it.

Then resuspend the cells in 150 microliters of sterile neuro stem cell basal media. Reset the pipette to 200 microliters, placing the tip against the bottom of the tube pipette up and down three to seven times to break up the clumps until you achieve a milky suspension. Over pipetting can result in cell Death.

Add another 9.8 mls of medium to reach a Total volume of 10 mls. Pass the suspension through a 40 micrometer cell strainer into a 50 ml tube so as to remove debris or unassociated pieces. Then pellet the cells by centrifugation at 700 RPM are 110 G for five minutes.

Vacuum off the S natin and resus. Suspend the cells in one ml of complete neuro stem cell medium. As we have cells from two brains, take nine mls of complete neuro stem cell medium and transfer to a 15 ml tube.

Mix the one ml of cells that were obtained from the two mouse brains. Then transfer to the 15 ml tube with a nine mls of complete neuro stem cell medium. Then add BFGF for a final concentration of 10 nanograms per ml EGF for a final concentration of 20 nanograms per ml and one microliter per ml of 0.2%heparin.

Then mix the cells that are in complete neuro stem cell medium. With growth factors, well transfer five mls of the suspension into each T 25 flask. Normally, the cells obtained from one brain are plated into one T 25 flask with five mls of complete neuro stem cell medium.

With growth factors. Label each flask And transfer to an incubator at 37 degrees Celsius with 5%CO for seven to 10 days. After seven To 10 days, evaluate the flask under the microscope.

By this time, the sphere should be 150 to 200 micrometers in diameter and ready to be passaged. Collect and transfer the contents of each flask into a 15 ml conical tube. Centrifuge the tube, remove the supernatant, Mix the pellet with one ml of prewarm trypsin and place in a 37 degree water bath for three minutes.

After trypsin, Add an equal volume of trypsin inhibitor and mix well centrifuge. Again, remove the supernatant and resuspend the cells in one ml of complete north stem cell medium. Take 10 microliters of the cell suspension and add to 90 microliters of triam blue.

Mix the suspension well and transfer 10 microliters of this to the hemo cytometer. Perform a cell Count. Transfer a total of 10 mls of complete nurse stem cell medium to a 15 ml tube to be able to plate two T 25 Flasks.

Then Mix the cells and add an appropriate amount of cells for a total density of 50, 000 cells per ml or 250, 000 cells per T 25 flask. Then add the growth factors as described before, which include E-G-F-B-F-G-F, and heparin. Mix the suspension and transfer to the T 25 flasks for a total volume of five MLS per flask.

Finally, Place the flasks in an incubator at 37 degrees Celsius with 5%CO2 to generate spheres. This is an example of primary culture three days after plating. This is an example of primary neuros sphere culture After seven days.

As you can see, there are spheres of various sizes. The neurospheres are phased bright or luminous, and become more spherical. As size increases.

There's also debris. The amount of debris depends on multiple factors such as microdissection and tissue preparation. Debris is commonly found in primary adult neuros sphere culture.

These are examples of passage. One adult neurospheres after seven days. Again, there are spheres of varying sizes, but there is much less debris.

The periphery of the spheres demonstrate micro spikes, which are evident in healthy spheres. If the spheres are allowed to grow too large, they may become dark in the center due to cell death and difficult To dissociate. We hope you'll find this video Useful.

Remember, do not let the sphere grow too large as they might attach to the substrate and differentiate, and also it would be difficult to subculture them. You need to pass your spears every seven to 10 days depending on their size and appearance. Good luck with your experiment.