Isolating Fluorescently Labeled Neurons from a Murine Brain

Take a transgenic mouse cortex containing fluorescent protein-expressing neurons.

Secure it on a sample holder.

Mount the holder on the vibratome tray containing oxygenated aCSF to maintain the tissue's physiological conditions.

Using set parameters, obtain coronal slices.

Place the slices in a meshed holder containing oxygenated aCSF and activity blockers to stabilize them.

Treat the slices with proteases to digest the extracellular matrix and loosen cells.

Wash with aCSF to remove the proteases.

Transfer the slices to a culture plate with aCSF and serum.

Under a fluorescence microscope, dissect slice regions containing fluorescent neurons.

Transfer the dissected fragments to a tube containing aCSF and serum.

Mechanically dissociate the tissue to form a single-cell suspension. Transfer the cells to a culture plate containing aCSF.

Under a fluorescence microscope, use a capillary pipette to collect fluorescent neurons.

Dispense these neurons into a culture plate containing aCSF for further analysis.

Dissect the brain from a euthanized mouse by opening the cranium with a small scissor, and extracting the fresh brain using fine forceps without damaging the cortex. Place the brain in chilled and oxygenated aCSF, leaving an air stone attached to 5% carbon dioxide-balanced oxygen during the entire duration of the sectioning.

Position the brain on the vibratom chuck, and cut coronal or sagittal sections at 300-micron thickness. Collect as many sections as needed to obtain a minimum of 10 to 50 labeled cells. Put slices on a cotton-meshed slice holder placed inside a beaker so they are bathed in oxygenated aCSF.

Move the slices into the beaker containing aCSF with activity blockers, and block for 15 to 20 minutes at room temperature while bubbling oxygen using an air stone. Move the slices to the beaker containing aCSF with the protease solution to perform mild digestion at room temperature. Following the digestion, wash out the protease by moving slices back to the beaker containing aCSF with activity blocker solution for 5 to 10 minutes at room temperature. Keep bubbling oxygen.

Next, prepare a 100-millimeter Petri dish containing aCSF with 1% FBS at room temperature and move individual sections into the dish for microdissection. Under a fluorescent dissection scope, use a pair of fine forceps to micro-dissect areas and layers of interest having a minimum of 10 to 50 cells.

Using a Pasteur pipette, move the micro-dissected pieces to a 2-milliliter microcentrifuge tube containing approximately 0.8 milliliters of 1% FBS in aCSF solution. Triturate the dissected tissue in the microfuge tube at room temperature.

Perform approximately 10 strokes with each of the flamed Pasteur pipettes, starting with the biggest and ending with the smallest exit diameter. Dispense the dissociated cells into a 100-milliliter Petri dish containing oxygenated aCSF, and wait 5 to 7 minutes for the cells to gradually settle.

Observe the GFP and RFP signal under a dissection microscope. Identify debris by examining the morphology under brightfield illumination. Once an area of cells with little debris has been identified, use the capillary and attached tubing to pick cells by blocking the end of the tubing valve with the tongue. Then, position the capillary close to the cells. Release the block on the capillary and quickly block again to capture the cell using capillary action.

Move the capillary to a 100-millimeter Petri dish with fresh oxygenated aCSF and gently blow into the tube while observing the capillary tip under fluorescence optics to dispense the cells into the dish. Repeat the collection procedure until 100 to 150 cells are collected, making sure that contaminants such as debris are minimal in brightfield differential interference contrast optics.