Spinal Cord Neurons Isolation and Culture from Neonatal Mice

The overall goal of this procedure is to be able to repeatedly and consistently isolate spinal cord neurons that can be used in a variety of experiments. This method can allow us to gain better insight into the biology of spinal cord neurons and hopefully understand their response to a variety of stressors. The main advantage of this technique is, that it allows for the isolation of a significant number of neurons, allowing those neurons to be used in a wide variety of experiments.

Visual demonstration of this method is critical as the trituration step where neurons are mechanically isolated from the tissue, requires some time to master, in order to ensure healthy neuron isolation, without killing them. To begin this procedure, separate the head from the body of the euthanized mouse pups, using scissors. Next, stabilize the tail and arms on the procedure table with the dorsal side facing up.

Then, cut the skin off using the curved iris scissors. After that, cut the spinal cord from the lumbar region, just above the hips and then cut both sides of the thorax to separate it from the body. Wash the samples sequentially in three 10 centimeter petri dishes, containing five milliliters of sterilized PBS for 10 seconds each, to remove excess tissue.

Subsequently, insert a 22 gauge needle and syringe filled with five milliliters of sterilized PBS into the caudal end of the spinal column and flush cranially, allowing the cord to exit into a fourth petri dish. Transfer the spinal cord in a 15 milliliter tube with five milliliters of HABG on ice. Avoid crushing the spinal cord and repeat the procedures for the rest of the pups.

Ideally, this process should take no more than 30 minutes for one group of pups, to ensure neuron isolation. To prepare the density gradient, prepare each of the four layers in four 15 milliliter tubes. Then, add one milliliter of each layer into a new 15 milliliter tube.

Start with layer one at the bottom, and add sequentially, until reaching layer four at the top. Avoid disturbing the layers while adding and any vigorous movements, that may cause mixing. Next, add the digestion media to the minced tissue.

Place the sample in a shaker in the water bath at 30 degrees celsius. Afterward, remove the tissue from the water bath and allow it to settle for a few minutes. To perform trituration, aspirate the excess digestion medium.

Suspend the tissue in two milliliters of HABG, using a narrow-bore pipette, triturate 10 times in 45 seconds and avoid introducing air, as it would significantly decrease the viable yield. Trituration is one of the most critical steps of the procedure, the tissue should be carefully drawn into and emptied from the fire polished pipette, avoiding excess vigor which can lead to neuronal death or being too gentle, which would result in a low yield. Following that, aspirate the top two milliliters of the supernatant and transfer it into a new 15 milliliter tube labeled, collection.

Repeat the collection procedures two more times, to yield six milliliters of supernatant. Next, slowly transfer the collection tube contents into the gradient tube prepared previously. Avoiding the disruption of the gradient.

To purify the neurons, centrifuge the gradient tube for 15 minutes at 800 times G, at 22 degrees celsius. Collect the desired layers and transfer them in a new 15 milliliter tube. For the highest purity neuron isolation, collect layer three.

For more yield with less purity, collect layers two to three. Next, dilute the density gradient by adding five milliliters of HABG to the newly collected layers. Centrifuge it, at 200 times G for two minutes at 22 degrees celsius.

After two minutes, discard the supernatant and re-suspend the cells in five milliliters of HABG, by flicking the palate. Subsequently, centrifuge the sample at 200 times G for two minutes at 22 degrees celsius again. Then, discard the supernatant and re-suspend the cells in three milliliters of neurobasal medium, by flicking the palate.

After that, determine the cell density using a cell counter and then, dilute the suspension to 300, 000 cells per one milliliter of neurobasal medium. Shake gently to distribute the cells and solution and add one milliliter of the sample to each well in the coded 24 well plates. Shown here, is the time course of the neurons in culture, after seeding onto the wells.

The cells will typically start adhering to the surface within the first couple of hours. Axons will begin to sprout within the first 24 to 48 hours, connections between various neurons and culture, typically reach maturity at seven days, at which point, experiments are typically carried out on the neurons. In this figure, nuclear staining is shown in blue, with cytoplasmic staining in green and here, the neuron cyctoskeletal protein microtubular associated protein two is stained, showing the outlines and axonal projections of the neurons after a week in culture.

The merge images are combined, showing the relative abundance of the neurons. Once mastered, this technique can be done in three to four hours, if it is perform properly. After its development, this technique paves the way for researchers, studying the spinal cord physiology and pathology, to explore the behavior of spinal cord neurons under normal conditions and in response to for pharmacological agents or stressors such as, oxygen and glucose deprivation, just mimicking our surgical procedures.

While taking on this procedure, it is important to avoid delaying or prolonging any of the steps to prevent isolation of unhealthy neurons. After watching this video, you should have a good understanding of how to isolate healthy and viable neurons from the spinal cords of neonatal mice that can be cultured in serum free media.