Preparation of Rat Brain Aggregate Cultures for Neuron and Glia Development Studies

We introduce a system for culturing CNS neuro progenitors as they develop and differentiate into mature neurons and glia. The procedure begins with isolating four brains from E 16 rat fetuses and digesting them with propane. The tissues are ated into a single cell suspension, filtered, counted, and dispersed into a six.

Well plate cells are gently resuspended once per day for two to three days to allow aggregate formation. Aggregates are then collected and sied through a 200 micrometer mesh and small clumps and cell debris are removed by washing. Finally, aggregates are counted and plated onto matrigel coated cover slips.

Hi, I'm Coto from the laboratory of Dr.John L in the Department of Veterinary Integrated Biosciences at the Texas a and m University. Today we will show you a procedure for brain aggregate cultures. We use this procedure in our level three to study endocyte lineage, cell development and accelerate interactions.

So let's get started. Before you start, have ready on the bench sterilize tools, 70%ethanol a diaper pad to work on and 10 centimeter petri dishes of dissection. Medium on ice.

In the hood, prepare a dissecting microscope with an ice platform and several 10 centimeter dishes of dissecting medium also on ice. Lastly, make up the solutions you will use and warm the plating medium to 37 degrees in a water bath. You begin this protocol by harvesting embryonic day 16 E 16 embryos from a Sprague dolly rat.

First euthanize a pregnant female according to a procedure approved by your institutional animal care and use committee. Lay the euthanized rat on a diaper pad and spray its abdomen with alcohol to clean it. Next, use tweezers to pinch the abdominal skin and with the other hand, use scissors to cut only the skin, spread apart the skin, and use another set of sterilized tweezers and scissors to cut through the muscle layer.

Next, remove the uterus and transfer it into a 10 centimeter dish containing ice cold dissection medium. The next step takes place in the hood using micros dissecting scissors. Carefully remove each embryo from the uterus.

Now it is time to harvest and begin. Work with the embryonic brains. Harvest the brain from each embryo and place the freed brains into a clean 10 centimeter dish of dissection medium on an ice platform Under the dissecting microscope, finish dissecting the brains.

Remove the midbrain and hind brain sections. Use fine forceps to remove the menes transfer what remains the forebrain to a 50 milliliter falcon tube containing dissection medium. Now the tissue is ready for the next step.

Dissociation of cells by enzymatic digestion. Just before you are ready to begin the digest, make up fresh solutions of propane and trypsin inhibitor filter. Sterilize them and warm them to 37 degrees Celsius in a water bath.

Remove excess medium from the dissected brains. Then add four milliliters of the prepared propane digestion solution to your 50 milliliter tube. Incubate the tube in a 37 degree Celsius water bath for exactly five minutes.

When time is up, use a pipette to remove propane's solution. Next, add five milliliters of freshly prepared trypsin inhibitor solution and put the tube back in the 37 degree Celsius water bath for two to three more minutes. When time is up, remove the trypsin inhibitor solution.

Repeat the incubation with trypsin inhibitor solution three more times. After you remove the trypsin inhibitor solution for the last time, add 20 milliliters. Warm plating medium pm to your tube.

Now it is time to iterate and wash your cells. Iterate the cells with a 10 milliliter pipette until all clumps have disappeared approximately 20 to 30 times. Then centrifuge at a hundred GS for seven minutes.

At this point, your cells should be well separated from each other. After seven minutes, remove the super Dayton and resuspend the pellet. In 10 milliliter plating medium to wash the cells.

Then spin them down again. Remove the supernatant and repeat the washing and centrifugation. Step after the second wash.

Resuspend the pellet in 10 milliliters of aggregate culture. Medium, pass the cells through a 70 micrometer cell sieve. Finally count live cells using a hemo cytometer.

And now you are ready to allow the dissociated brain cells to form aggregates. After counting, suspend the dissociated cells in aggregate culture medium at a density of two times 10 to the sixth cells per milliliter. Transfer two milliliter of cell suspension into each well of an uncoated six well plate culture.

The cells for three nights each day gently resuspend the cells. Once using a P 1000 pipette men, the cells will begin to form aggregates on the day of aggregate plating. After the cells have grown for three nights, gently resuspend cells again and CVE the suspension through a 200 micrometer mesh into a 50 milliliter falcon tube.

Allow aggregates to settle to the bottom of the tube for approximately three to five minutes. Then carefully remove the supernatant, add medium, gently resuspend the aggregates and let them settle again. Repeat this procedure several times.

To remove dead cells, individual cells and debris, use the microscope to check whether the supernatants still contains un aggregated cells and debris. Once you have thoroughly washed the aggregates, gently resuspend them in the same medium and count aggregates to determine their concentration. Then adjust the density of aggregates to around 25 to 30 aggregates per 50 microliters.

Now it is time to seed cover slips with aggregates. Transfer 1000 microliter aliquots of the aggregate suspension to two milliliter eend Dwarf tubes since aggregates tend to settle down to the bottom of tubes, it is crucial to make multiple tubes of the aggregate suspension for plating. The cover slips used in this step are coated in a cell culture substrate matrigel by an overnight incubation at 37 degrees Celsius.

After you have made Alec watts of the aggregate suspension, take the culture plate containing the matrigel coated cover slips out of the incubator. Remove the solution from each. Well wash the cover slips with warm PBS and then DD H2O.

They are now ready for aggregate seeding to seed the cover slip with aggregates. Invert the tube of aggregate suspension to mix and evenly distribute the aggregates. Then load 50 microliters of aggregate suspension into the center of each cover slip.

Gently put the plate back to incubator without any further disturbance to allow aggregates to attach evenly to the cover slip. The density of aggregates is crucial if aggregates are seated too close to each other or if the seating density is high, they tend to merge together as they grow four to six hours after initial seating. Add 500 microliters of culture medium to each well.

To maintain the aggregate culture, half change the medium every three to four days. When you change the medium, add it along the sidewall of the well to avoid disrupting the aggregates. A few hours after aggregates attached to the cover slip, neite outgrowth from the aggregates can be observed.

Axons grow rapidly in the first two weeks and form abundant connections among aggregates. Ggl progenitors migrate radially outta the aggregates and differentiate over time into astrocytes and mature oligodendrocytes. We've just shown you how to culture CS aggregate.

When doing this procedure. It's important to remember to quote capacity properly and see the aggregate evenly. So that's it.

Thanks for watching and good luck with your experiment.