Un sistema de Microbiomechanical para estudiar la formación de várices y recuperación en axones de la neurona Central

The overall goal of this pressurized fluid setup is to produce repeatable and consistent swellings along the axons of cultured neurons. This method can help answer key questions in the brain injury field such as the mechanisms behind subcellular morphological changes in neurons following injury. The main advantage of this technique is that the method produces consistent and repeatable morphological changes that recover over time.

This is very important technique we developed in the lab. This is because we can not only use the system to identify molecular mechanism governing mechanical-stress-induced axon morphology change as a new form of neuronal plasticity, but also use the system as a screening tool to identify new drugs and the treatment strategy for mild or traumatic brain injury. To begin this procedure, place one or more boxes of 12-millimeter or 25-millimeter coverslips into a glass beaker containing 70%nitric acid, and incubate the coverslips at room temperature overnight.

The next day, transfer all coverslips into a four-liter beaker filled with 2.5 liters of double-distilled water. Shake the beaker containing the coverslips for one hour at 100 RPM, and don’t forget to use rubber bands to hold the beaker during shaking. Subsequently, transfer the cleaned coverslips to a dry flask with the metal cover.

Bake the coverslips in an oven at 225 degrees Celsius for six hours. Then, allow them to cool to room temperature and store them in a sterilized plastic container until usage. Now, place one coverslip into each well of a 24-well plate.

Add 30 microliters of coverslip coating solution onto a 12-millimeter coverslip and swirl. Afterward, add one milliliter of sterile PBS to each well. The plate can be either used immediately or stored at four degrees Celsius in sterile PBS for up to one week.

On the day of dissection, remove PBS from the plate. Air dry and place the plate under UV light for two to four hours. To dissociate the hippocampal neurons for culture, thaw the protease enzyme solution and pre-warm the plating medium at 37 degrees Celsius.

Next, rinse the dissected hippocampus with SLDS. Then, remove it and add two milliliters of protease enzyme solution and incubate the sample at 37 degrees Celsius for 15 minutes. Afterward, wash the hippocampus explants with five to 10 milliliters of plating medium twice and add five milliliters of plating medium.

Dissociate the hippocampal explants into single cells by generating a small vortex using a pipette with a one-milliliter plastic tip. Pipette up and down about 40 times avoiding the formation of oxygen bubbles until the solution becomes cloudy and no large pieces of explant remain. Next, centrifuge the cells at 1, 125 times gravity for three minutes.

Remove the supernatant with the cells submerged in medium and re-suspend the cells in 145 milliliters of plating medium. Then, add one milliliter of the cell suspension per well to a 24-well plate. Incubate the cells in the plating medium at 37 degrees Celsius for two to four hours.

Afterward, remove the plating medium and replace it with fresh maintenance medium. After two days, replace half of the medium with two micromolar RSC dissolved in maintenance medium to inhibit growth of fibroblast, endothelial and glial cells. Then, after exposing the cells to RSC for two days, replace the medium with fresh maintenance medium.

To transfect for the visualization of cell morphology, dilute the construct from stock at one microgram per microliter in Opti-MEM medium and vortex. Then, dilute liposome-mediated transfection reagent in Opti-MEM media and vortex. Subsequently, prepare one construction dilution and one transfection dilution for each well.

Next, prepare a one-to-one mixture of the construct and Opti-MEM medium and transfection reagent and Opti-MEM medium and vortex. Incubate the mixture at room temperature for 20 minutes. After 20 minutes, remove half of the medium from each well and transfer this medium into a clean conical tube.

Keep the tube in the incubator, then add 100 microliters of the mixture prepared in the previous step to the remaining medium in the well. Allow the cells to incubate at 37 degrees Celsius for 20 to 30 minutes. During incubation, add one volume of fresh maintenance medium to the medium in the conical tube and place the mixture in the same incubator.

Following incubation, remove the mediums containing the transfection solution from the wells and replace it with the one-to-one mixture of old and fresh maintenance medium in the conical tube. In this procedure, insert the pipette into the screw top of the micromanipulator to hold it into place. Screw the metal flat-top screw attached to the micromanipulator arm so that it holds the unpulled end of the pipette just above where the rubber tubing is attached.

Angle the pipette at a 45 degree angle with the surface of the neuron containing coverslips. Turn on the florescence filter. Open the aperture, and ensure a fluorescent spot can be seen coming through the objective.

Using the micromanipulator, move the pipette into a position that lines up the tip with the fluorescent spot and lower the pipette into a position just above the height of the cell culture dish. Once it is in position, turn on the transmitted light and turn off the florescence. Using tweezers, transfer one coverslip with the cells facing toward the pipette from the cell culture plate to the cell culture dish containing 2 milliliters of Hank’s buffer at room temperature.

Using the fine focus knob and 20 fold objective, focus on a plane about four full turns above the cells. Subsequently, use the micromanipulator to position the pipette tip so that it is focused and in the center left-hand side of the plane as seen through the eyepieces. Once the pipette is in position, focus the microscope on a plane that contains the region of interest.

In the imaging of seven div, GFP transfected mouse hippocampal neurons showed an axon pre and post puffing, as well as a following 10 minute recovery period. Here is the Kymograph of time-lapse imaging of the same neuron. Red arrows indicate the varicosities that have formed to following 150 seconds of 190 milliliter H2O puffing, beginning at 30 seconds.

Once mastered, this technique can be done in three hours if it is performed properly. Following this procedure, other methods like whole cell recording, time-lapse FRET and TIRF imaging, and calcium imaging can be performed in order to answer additional questions like the impacts of mechanical stress on neural and electrical and calcium signaling, protein trafficking and interactions. After its development, this technique paved the way for researchers in the photo traumatic brain injury to explain molecular mechanisms underlying axonal varicosity formation in living neurons.

After watching this video, you should have a good understanding of how to perform the experiments to study mechanosensations of central neurons.