Preparation and Morphological Analysis of Chick Cranial Neural Crest Cell Cultures

Neural crest cells emigrate from cranial neural folds in embryos or when placed in culture. This provides a convenient method to isolate primary migratory neural crest cells without cell dissociation. While neural crest migration typically occurs within the three-dimensional embryonic environment, two-dimensional culture enables visualization and investigation of migration-related processes.

Although cranial neural fold culture protocols exist, this method requires training and organism-specific steps. This video demonstrates techniques to facilitate the reproducible preparation of chick cranial neural fold cultures. Beginners often struggle with dissecting and plating neural folds and assessing culture outcomes.

Following the dissection guidelines and applying the morphometric analysis described here allows consistent, quantitative results to be obtained. To begin, place the fertilized eggs in an upright position inside a humidified incubator at 38 degrees Celsius. After removing the eggs from the incubator, disinfect the shells by thoroughly spraying 70%ethanol and letting them dry.

For fixing and staining the cultures, use glass cover slips placed into a multi-well tissue culture dish, and pipette enough fibronectin solution to cover the bottom of the well. Next, replace the lid, and incubate the plate with fibronectin solution in a humidified incubator at 38 degrees Celsius for at least one hour while dissecting neural folds. Use blunt forceps to poke a small hole in the shell at 1/4 to 1/3 of the egg's length.

Carefully insert the blunt forceps into the small hole, ensuring not to disrupt the yolk. Then, cut through the eggshell around the egg, remove the top of the eggshell, and position the embryo for isolation. Prepare the embryo by gently using the flat edge of closed blunt forceps to wipe away any excess albumin on the yolk's surface covering the embryo.

Use forceps to place a filter paper support frame over the embryo, with the embryo in the frame's window. Gently press down on the filter paper to adhere to the yolk. Cut around the outside of the filter paper frame with dissection scissors.

Use forceps or scissor tips to grasp the edge of the frame, and gently lift the embryo away from the yolk. Place the embryo with the paper frame side down into a 60 or 100-milliliter Petri dish filled with Ringer's with pen-strep. Keep the dish of embryos on ice if collecting them for an RNA or protein-sensitive downstream application.

Transfer an embryo to a clean dish containing Ringer's pen-strep solution. Gently swish the embryo back and forth to clear away any yolk that obscures the view, and exchange the Ringer's pen-strep solution or transfer it to a fresh dish if it becomes cloudy. Position the embryo dorsal and frame side up under a dissecting microscope.

Leave the embryo on the paper frame to keep it stretched and held in place. Then, remove the vitelline membrane, using forceps to expose the neural folds. Include tissue caudal to the expanding optic vesicles and rostral to the hindbrain, where rhombomere constrictions are just beginning to appear.

Using spring scissors, carefully excise the midbrain neural folds. Take care to excise the dorsal-most aspect of the neural fold with minimal contamination of the neural tube and non-neural ectoderm, and transfer the neural folds to a clean dish containing Ringer's pen-strep solution using a P20 pipettor or a sterile glass Pasteur pipette rinsed with yolky Ringer's pen-strep solution. Store collected folds on ice while dissecting additional folds.

After removing the culture dish from the incubator, use a pipettor or Pasteur pipette to remove the fibronectin solution from cover slips, dish, or well. After rinsing the fibronectin-coated substrate with Ringer's pen-strep solution, add an appropriate volume of complete culture media to the dish or well. To block the plastic and prevent the tissue from sticking, use a P20 or P200 pipettor, and rinse the pipette tip with yolky Ringer's pen-strep solution.

Then, transfer the isolated neural folds towards the center of the fibronectin-coated cover slip, taking care to transfer as little Ringer's pen-strep solution as possible. After allowing the explants to settle for 10 to 15 minutes, place them into a humidified chamber at 38 degrees Celsius by slowly and carefully carrying the culture dish with plated neural folds. Remove the culture media with a Pasteur pipette, and rinse the wells with filter-sterilized PBS.

Add 4%paraformaldehyde, and keep it on a platform shaker for 15 minutes at room temperature. At the end of the incubation, remove paraformaldehyde, rinse the wells three times with PBS, and add PBST containing 5%serum. Now, pipette 30 microliters of 200-nanomolar Oregon Green conjugated phalloidin onto a smooth surface for each cover slip.

Using a pair of forceps, remove the cover slip from the PBST containing 5%serum, ensuring the orientation of the cells facing upward. Wick off excess liquid by briefly touching the cover slip's edge to a delicate task wiper. Gently place each cover slip cell side down onto the drop of diluted phalloidin, and incubate for 30 minutes at room temperature.

After the incubation period, lift the cover slip from the staining solution and place it back into the culture dish, flipping it over so the cell side is up. Ensure to cover the cover slip with PBST, and place it on a platform shaker for 10 minutes, keeping the cover slips covered and in the dark. Remove PBST, and repeat washing the cover slips twice for a total of three 10-minute washes.

Place one drop of mounting media onto a microscope slide. Mount the cover slip cell side down by slowly lowering the cover slip at an angle onto the mounting media to avoid creating bubbles, and allow the media to set before imaging. Finally, image the stained cells, and export them as TIFF files.

The neural crest cells began to emerge from adherent neural fold explants within three to four hours of incubation, and migration was completed after approximately 20 hours. The migratory neural crest cells were labeled using HNK-1, and most cells appeared to be HNK-1 positive. The migrated neural crest cells were visualized by staining filamentous actin with phalloidin.

Images of the phalloidin-stained cells were processed using ImageJ to produce a threshold image in which cells appeared black with a white background. The threshold image was analyzed and various measurements displayed in contrasting bar graphs or violin plots. For example, the average area of the 69 cells analyzed was approximately 802.11 square micrometers, ranging from 60.27 to 2, 664.53 square micrometers.

Circularity reflects the cell's protrusiveness and ranges from 0.101 to 0.875. The lower values indicate an elongated shape, while a value of one indicates a perfect circle. Most cells exhibited an elongated shape, which is most easily seen in the violin plot.

Migratory neural crest cells in this field had an average aspect ratio of approximately 2.13, and an aspect ratio of one indicates a symmetrical shape. Careful excision and explant plating are crucial for successful cultures. Allow neural folds to settle cut side down for 10 to 15 minutes, and then slowly transfer to the incubator to encourage adhesion.

Variations on these techniques include transient genetic manipulation prior to culturing and time-lapse imaging during incubation. Additionally, this protocol allows for collection of premigratory and migratory neural crest cell populations for omics-level analysis.