Isolation and Culture of Dissociated Sensory Neurons From Chick Embryos

The overall goal of this procedure is to culture an enriched population of dissociated primary sensory neurons from chicken embryos. This is accomplished by collecting intact dorsal root ganglia from the embryonic day, seven to 10 chicken embryos into a conical tube, and subsequently breaking them up into its dissociated cell suspension. As a second step dissociated, DRG cells are plated on a culture dish for incubation.

Afterward, the culture dish is gently rinsed to preferentially dislodged the neurons. The final step includes plating the dissociated sensory neurons on the well-defined substrata, such as acid washed and baked cover slips pre-coded with high or low concentrations of laminin. One immuno cyto chemistry is used to show the presence of NCA and activated beta one integrins in the cell bodies, neurites and growth cones of the dissociated cultured embryonic chick sensory neurons.

This method is a powerful tool to elucidate numerous aspects of neurobiology, including mechanisms of axon guidance, growth cones, cytoskeletal dynamics, as well as cell surface composition and regulation. This technique allows researchers to rapidly obtain the cultures highly enriched with dissociated sensory neurons, which contain very few glial cells. This is distinct from the intact dorsal root gangling cultures that contain both neurons and non neuronal cells.

Furthermore, minor variations of this protocol should allow high yield of neuronal cultures from sources other than dorsal root ganglia, including embryonic forebrain. In a laminar flow hood crack a staged fertile white leghorn chick egg and place its contents into a 100 millimeter Petri dish. Subsequently transfer the embryo to another 100 millimeter Petri dish.

Add warmed one XPBS to the Petri dish to keep the tissue wet. The embryo is decapitated and the body is transferred to another clean Petri dish. Warmed F 12 HS 10 is added to the tissue.

Place the Petri dish with embryonic tissue under the dissecting microscope in a laminar flow hood. After that, make a vertical midline incision from the tail to the neck by pinching the developing dermis to expose the heart, lung, and other organs. With the fine forceps, gently grasp the neck and the aorta or other tissue immediately superior to the heart and pull towards the tail in order to eviscerate the animal.

Next, apply warm F 12 HS 10 to the tissue to further clean the preparation and keep the tissue moist and repeat the applications of F 12 HS 10 throughout dissection as necessary. Then remove any remaining cardiovascular, respiratory, or digestive organs to expose the developing ribs vertebral column and DRGs to collect the DRGs along both sides of the lumbar vertebral column, inferior to the ribs, pluck the DRGs from the embryo. This is done by pinching and severing the roots that reach from the Dr.G towards the spinal cord and grasping the developing nerve from the DRG towards the periphery or vice versa.

Afterward, place the intact DRGs in a 35 millimeter culture dish filled with warm F 12 HS 10. Remove any excess tissue such as the dorsal or ventral roots that may adhere to DRGs. To obtain DRGs superior to the lumbar region, remove the ventral half of the thoracic and cervical vertebral column.

This is done by making a cut in the developing vertebral column perpendicular to the spinal cord axis in the upper lumbar region, and another cut in the upper cervical region. Then make cuts along the right and left sides of the vertebral column. Collectively, these cuts will detach the ventral half of the vertebral column from the embryo, and the column is then removed from the embryo to allow easy access to the DRGs.

Remove the thoracic and cervical spinal cord by grasping the spinal cord with fine forceps. Next, collect the intact thoracic and cervical DRGs as done previously, and place them in warm F 12 HS 10 with other DRGs. Remove excess tissue that may adhere to the DRGs.

Then rinse the interior of a new glass paster pipette with F 12 HS 10 To help prevent DRGs from sticking to the wall of the pipette. Use the rinsed pipette to transfer all the DRGs and F 12 HS 10 from the small Petri dish to a 15 milliliter sterile conical tube, additional embryos can be dissected to increase the number of DRGs in this procedure. Place the conical tube with intact DRGs into the centrifuge and gently spin at 200 times G for two to three minutes.

Confirm that the DRGs have sunk to the bottom of the tube. Then gently remove F 12 HS 10, leaving the DRG pellet undisturbed. Next, add two milliliters of one X-C-M-F-P-B-S to dislodge the pellet of DRGs.

Spin again and remove the C-M-F-P-B-S while retaining DRGs. Repeat this wash step for a total of three washes. After the washes are complete and C-M-F-P-B-S is removed, add two milliliters of warmed trypsin to the 15 milliliter tube and gently dislodge the DRGs.

Then place the tube in a 37 degrees Celsius water bath for 10 to 15 minutes to digest the DRGs into dissociated cells. Gently tate the DRGs in trypsin solution and visually inspect for the absence of intact DRGs. Continue tation until intact DRGs are no longer seen by eye and or allow for longer tripsin incubation in the 37 degrees Celsius water bath until intact DRGs are dissociated.

Next centrifuge the dissociated DRGs and trypsin at 200 times G for three to five minutes in order to form a pellet. Spin longer if needed until the pellet is formed. Then carefully aspirate the trypsin without disrupting the pellet.

Add two milliliters of F 12 hs 10 to the pellet and gently tri the DRGs. Subsequently transfer all the contents from the conical tube to a 100 millimeter culture dish. Rinse the tube with eight milliliters of F 12 HS 10 and transfer it to the culture dish two milliliters at a time for a total of 10 milliliters.

After that, place the culture dish in a humidified 37 degrees Celsius incubator for three hours. Now add 400 microliters of prepared laminin, one to each acid washed and baked glass cover slip and spread it with a pipette tip to ensure full coverage. Surface tension will allow laminin one to remain on the cover slip and not spread onto the bottom of the culture dish.

Place a lid over the dish and allow Lamin one to incubate for two to three hours at room temperature. Afterward, rinse the laminin coated cover slips with sterile PBS in a laminar flow hood three times. Leave the PBS on the cover slips after the last rinse.

Do not lose surface tension during rinsing steps. After incubation, use a 10 milliliter sterile pipette to gently rinse the bottom of the dish containing dissociated DRG cells. Hold the dish at a 45 degree angle.

Withdraw approximately seven milliliters of F 12 HS 10. Then gently expel it onto one third of the dish to dislodge the neurons. Repeat this five more times.

Next, rotate the dish approximately 120 degrees and repeat the rinsing procedure for another six rinses. Then repeat the procedure again for the remaining one third of the dish. Now using the same pipette transfer F 12 HS 10 containing neurons from the dish to a conical tube.

Centrifuge it for five to 10 minutes at 200 G to pellet. The cells subsequently remove F 12 HS 10, leaving the pellet undisturbed. Then resuspend the pellet with two milliliters of warmed F 12 H plus supplements.

Afterward, determine the cell density with a hemo cytometer dilute the cells as necessary with F 12 H plus supplements to obtain the desired plating concentration. Then remove PBS from the laminin coated cover Slips immediately. Place 400 microliters of cells onto them.

Carefully transfer the dish to a 37 degree Celsius cell culture incubator and allow to incubate for two to three hours under sterile conditions. Gently flood the 35 millimeter dishes containing neurons with 1.6 milliliters of F 12 H plus supplements. Subsequently return the cells to the incubator and incubate them overnight.

This figure shows cultured dissociated primary sensory neurons immuno labeled for NCA and activated beta one integrins. The embryonic chick neurons grow robustly on high concentrations of laminin one neuronal cell bodies. Neurites and growth cones are immuno positive for NCA and activated beta one integrin.

The merged image of the two stains reveals that most of the cells are positive for both N cam and activated beta one integrin. As expected for embryonic chick sensory neurons, less than 5%of the cells harvested by this technique are non neuronal. In this image, there is a cell with a glial like morphology that is NCA M negative and beta one integrin positive.

Consistent with a non neuronal cell. A higher magnification of a cultured sensory neuron clearly shows the neurites extending from the cell body, and this inset shows higher magnification of the growth cone at the tip of a neurite. While attempting this procedure, it's important to remember to carefully and fully dissociate intact dorsal root ganglia and plate the neurons and appropriate density on the defined substrata, such as acid washed and baked covered slips coded with known concentrations of laminate one Following the culture procedure.

Live imaging methods such as time-lapse microscopy can be performed to evaluate growth coone, motility phylo, POAL dynamics, and other growth cone behaviors on defined substrata and in response to molecules applied to the culture media. After watching this video, you should have a solid understanding of how to produce a robustly growing culture that is highly enriched for primary dissociated neurons from chick embryonic dorsal root ganglia.