The overall goal of this procedure is to produce healthy pure populations of cerebellar granule neurons and to genetically manipulate them and model different mechanisms of neuronal injury in the primary culture in vitro. This method can help answer key questions in the field of neuronal injury. We use this protocol to investigate the underlying molecular mechanisms of neural injuries following acute brain damage and neuro-generative diseases.
The main advantage of this protocol is that we can model different mechanisms of cell death such as excitotoxicty, oxidative stress, DNA damage, and developmental event using the culture system. Although this method can provide insights into neuronal injury it can also be used with rat's cerebellar neurons to study neuronal activity and morphogenesis in response to growth factors and side events. To extract the brain of a six to seven day old mouse, use a pair of forceps to grasp a head, and with micro-dissection scissors, cut the skin anteriorly towards the head.
Push back the skin to reveal the skull. Next, penetrate the skull with the tip of the scissors and cut anteriorly and laterally. Taking care not to damage the cerebellum, and facilitate identification and removal of meninges, then use forceps to peel back the skull exposing the brain.
With a pair of forceps or spatula, gently tease out the brain into cool dissection solution. To isolate the cerebellum, place the brain in the dissection solution supplemented with magnesium sulfate and keep the solution and the brain on ice. Under the dissection microscope, remove the meninges by using fine forceps, then dissect out the cerebellum from the brain in magnesium sulfate supplemented dissection solution.
This helps in peeling off the remaining meninges and allows one to go in between layers to clean the cerebellar folds. The presence of meninges is neuronal culture results in unhealthy cells and eventual cell death. As such, it's important to ensure complete removal of meninges before proceeding with culture.
After that, turn the cerebellum to its ventral side and ensure the removal of the choroid plexus. Next, pull the cerebella into a 35 millimeter dish containing one milliliter of magnesium sulfate supplemented dissection solution. Chop the tissues into small pieces and transfer them into a 50 milliliter tube containing 30 milliliters of magnesium sulfate buffered dissection solution.
In this step, centrifuge the 15 milliliter tube containing the chopped cerebral tissue for five minutes at 644 times g, and four degrees Celsius. After that remove the supernatant, and add 10 milliliters of trypsin dissection solution. Then shake the table at high speed for 15 minutes at 37 degrees Celsius.
Add two milliliters of trypsin inhibitor solution one to the tube and rock gently for two minutes. Subsequently, centrifuge the tube for five minutes at 644 times g and four degrees Celsius. After five minutes, remove the supernatant, and add two milliliters of trypsin inhibitors solution two before transferring it to a 15 milliliter tube.
Then triturate the tissue in the 15 milliliter tube until the solution becomes murky. Let it settle for five minutes. Then remove the clear supernatant and transfer it to a new tube containing one milliliter of calcium chloride supplemented dissection solution.
Add another two milliliters of trypsin inhibitor solution two to the bottom of the tube containing the pellet. Triturate it again and let it settle for five minutes. Remove the supernatant and add it to the tube containing supernatant from the previous step.
Repeat this process until most of the tissue is mechanically dissociated. Add 0.3 milliliters of calcium chloride supplemented dissection solution to the supernatant collection for every milliliter of supernatant. Mix the contents of the tube and then centrifuged for five minutes at 644 times G at room temperature.
Following that, remove the supernatant. Add 10 milliliters of fresh media to the pellet and mix. Then count the live cells and dilute them to a concentration of 1.5 times 10 to the six cells per milliliter.
Plate the cells on the previously prepared poly D-lysine plates. For four well plates, plate 0.5 millimiters of the sample giving 7.5 times 10 to the fifth cells per well. For 35 millimeter dishes, plate four milliliters of the sample, giving six times 10 to the sixth cells per plate.
For cover slips, plate 0.5 milliliters giving 7.5 times 10 to the fifth cells per well. After 24 hours, add AraC to the plates to reduce glial contamination. If the cells are to be maintained for seven to eight days, repeat this treatment on day three and maintain the cultures in a 5%CO2 incubator at 37 degrees celsius.
For the cultures maintained longer than five days feed the culture with glucose every two days starting on day five. To induce neuronal excitotoxicity with NMDA, following seven days in vitro, treat the cerebellar granule neurons with 100 micromolar NMDA and 10 micromolar glycine for one hour. Then, replace the medium with conditioned medium from the parallel cultures with no treatment.
This concentration results in 50%cell death at 24 hours post treatment. For ROS induced cell death, treat the neurons with hydrogen peroxide at 75 to 100 micromolar for five minutes. After five minutes, switch it to the conditioned media from parallel cultures.
Due to the instability of hydrogen peroxide, the concentration must be optimized to a level that induces between 50 and 70%cell death after 24 hours. This concentration in usually between 75 and 100 micromolar. To induce cell death by DNA damage, treat the cerebellar granule neurons with 10 micromolar camptothecin to induce more than 50%cell death within 24 hours.
For low potassium induced neuronal apoptosis in cerebellar granule neurons, change the media containing 25 millimolar potassium to low potassium medium with five millimolar potassium following seven days in vitro. Neurons were transduced with lentivirus for RFP at MOI of three at the time of plating. Fixed and stained at seven days in vitro.
The co-localization of RFP signal MAP2 and Hoechst is shown to demonstrate healthy neurons that are fully transduced by lentiviruses. The images shown here represent live-dead assay analyses of neurons infected with different MOI of adenovirus to measure toxicity. Cerebellar granule neurons infected with adenovirus expressing LacZ at an MOI between 25 and 50 allows for maximal efficiency while maintaining minimal toxicity.
Less than a 1%difference in cell survival compared to control is seen when infecting at this MOI. These figures show the Hoechst staining of control cerebellar granule neurons and cerebellar granule neurons treated with NMDA to induce cell death. Note the formation of pyknotic nuclei with NMDA treatment.
This is an indication of cell death and can be seen in approximately 50%of the culture 24 hours after treatment with 100 micromolar NMDA and 10 micromolar glycine. Once mastered, this technique can be performed in two and a half hours with six mouse pops, if performed properly. When performing this technique, it's important to use aseptic technique and chained surgical instruments as required in order to minimize risk of culture contamination.
After its development, this technique paved the way for researchers in the field of neuroscience to study neuronal development and neuronal injury in primary mouse neurons. After watching this video, you should be able to culture cerebellar granular neurons, genetically manipulate them using viruses and induce varying mechanisms of neural injury. Following this procedure, other methods like immunofluorescence or cell viability assays can be performed to answer questions like whether a gene of interest enhances cell survival in response to excitotoxicity, oxidative stress, or DNA damage.
