Isolating and Culturing Chick Embryonic Neurons on a Multi-Electrode Array

Take a fertilized chicken egg containing an early-stage embryo.

Sterilize the shell and carefully open it.

Remove the embryo. Dissect and transfer the head to a culture plate containing a buffer. Remove the eyeball.

Make an incision and remove the skin layers, exposing the forebrain and midbrain.

Remove the inner membranous layer and blood vessels. Dissect and transfer the forebrain to a culture plate with a buffer.

Cut the tissue into smaller fragments and transfer them to a conical tube. Once the fragments settle, remove the buffer.

Add a proteolytic enzyme solution. Incubate to digest the tissue's extracellular matrix and loosen neurons.

Remove enzyme-rich media, wash with neurobasal media, removing any remaining enzymes.

Add neurobasal media. Mechanically dissociate the tissue to obtain a neuronal suspension.

Seed these neurons on a multi-electrode array system coated with an extracellular matrix. Add neurobasal media and incubate.

Neurons attach and extend projections, forming a neuronal network.

On the day of neuron plating, remove a vial of extracellular matrix from the -20 degrees Celsius freezer. Spray with 70% ethanol and place on ice. Be sure to thaw out the extracellular matrix on ice. Do not thaw out at room temperature, as the ECM polymerizes above 0 degrees Celsius.

Work in the BSL-2 hood to dilute the extracellular matrix to 25% by adding 300 microliters of cold neurobasal medium to the 100-microliter aliquot. Then, use a P200 pipette to add 100 microliters of 25% extracellular matrix solution to the center of the multielectrode array, taking care not to touch the electrodes. Immediately remove the ECM, leaving a thin film on the surface.

Cover the multielectrode array, and place in the tissue culture incubator until ready to plate the neurons. Begin chick neuron isolation by sterilizing the outer shell of an E7 egg with 70% ethanol. It's important to maintain sterile conditions from this point onward. Ensure all instruments are sterilized with 70% ethanol and solutions are sterile. It is helpful to use a dissection hood but is not absolutely necessary for dissections, if it is performed quickly.

Then, after retrieving and decapitating the embryo, cut around the eyes and remove the eyeballs. Next, use Dumont number 5 fine forceps and spring scissors to make an incision on the ventral side and remove the outer layers of skin to expose the forebrain and optic tectum. Peel and remove the peel membrane carefully.

Transfer the forebrain into another Petri dish containing HBS and cut it into small pieces of about 2 millimeters with spring scissors. Use a sterile transfer pipette to transfer the pieces of forebrain into a 50-milliliter centrifuge tube. After the pieces of forebrain sink to the bottom of the centrifuge tube, remove as much HBS as possible.

Next, add 1 milliliter of pre-warmed 0.5% trypsin-EDTA and incubate at 37 degrees Celsius for 15 minutes. Use a Pasteur pipette to carefully remove the trypsin without disturbing the pieces of tissue. Add 1 milliliter of neurobasal medium and let the pieces of tissue sink to the bottom. After removing the medium and repeating the wash, add 2 milliliters of neurobasal medium and triturate gently until no more pieces of tissue are seen.

Dilute the resuspended cells 1 to 10 with neurobasal medium, and count viable cells using trypan blue dye and a hemocytometer. After counting, plate the associated cells on ECM-coated multielectrode arrays at a density of 2,200 cells per square millimeter.