A Quantitative Cell Migration Assay for Murine Enteric Neural Progenitors

Ethics statement

Experiments involving mice were performed following Canadian Council of Animal Care guidelines for the care and manipulation of animals used in medical research. Protocols involving the manipulation of animals were approved by the institutional ethics committee of the University of Quebec in Montreal (Comité Institutionnel de Protection des Animaux; reference number 0512-R3-650-0513).

1. Preparation of Collagen Gels

Work in a sterile fashion, under a tissue culture hood.

1. Prepare complete 5x DMEM (Eagle’s modified essential medium) including standard antibiotics. Dissolve 3.37 g of DMEM powder and 0.925 g of NaHCO₃ in 20 ml water. Sterilize by passing through a 0.22 μm filter. Add 2.5 ml of sterile 100x penicillin/streptomycin and 25 ml of sterile heat-inactivated fetal bovine serum. Store at 4 °C.
2. On ice, mix 800 µl of Collagen I solution (3.77 mg/ml in 0.02 N acetic acid, filter-sterilized), 600 µl of complete 5x DMEM and 17 µl of 1 N NaOH. Dilute with sterile water to a final volume of 3 ml. Include relevant growth factors in the mix. We typically use GDNF at 10 ng/ml to stimulate enteric NCC migration.
3. Deposit about 480 µl in each well of a single row in a 24-well plate. Avoid bubbles. The remaining rows can be used to test the effect of other growth factors on cell migration.
4. Let the collagen polymerize at least 1 hr in a sterile incubator at 37 °C.

2. Dissection of Animals¹⁶

1. Set up matings and check for vaginal plugs the next morning. Day e0.5 being noon on the day the vaginal plug is found, isolate the female and wait 12 days until e12.5.
2. Anesthetize pregnant mice with isoflurane and euthanize by CO₂ inhalation.
3. Spray the mouse with 70% ethanol. Lift the abdomen skin and open the abdomen cavity with dissecting scissors.
4. Remove the uterus into a glass Petri dish filled with ice-cold PBS (phosphate-buffered saline). Cut the uterus transversely between individual deciduum swellings to isolate each embryo implantation site.
5. Work on each implantation site separately in another glass Petri dish filled with ice-cold PBS. Under a dissecting microscope, use fine forceps to remove the muscle layer of the uterus.
6. Open the visceral yolk sac and amnion to reveal the embryo. Take care when severing the blood vessels joining the embryo to the placenta/visceral yolk sac, as they are intertwined with the developing intestines.
7. Sever the embryo's head at the neck.
8. Insert a closed forceps in the abdominal cavity of an embryo, just above the dark red-colored liver and let the forceps open of itself (stop applying pressure) to make a transversal opening in the abdomen cavity. Pull the opened forceps down towards the posterior end of the embryo to open the abdomen completely.
9. Grab the connective tissue behind the liver and pull the guts out of the abdomen, being careful not to break the intestines (the colon is attached to the anus).
10. Cut the colon to free the guts from the rest of the embryo. The cut can be made at any location along the colon. Reserve the tail portion for later.
11. Tease out the connective tissue from the cecum, then the rest of the intestines. Be careful not to wound the intestines while doing so.
12. Cut away the liver and stomach (on the rostral end of the small intestine), as well as the mesonephros and genital ridges if some are present.
13. Isolate the small intestine. Again, be careful not to nip the intestine. From now on, the rosrrocaudal orientation of the gut tissue can be tracked using the sharp curvature present on the rostral end. Leave the small intestine in PBS at room temperature for as short a time as possible before embedding (see step 3.3).
14. Finally, record the number of tail somites for each embryo to accurately determine the stage of embryonic development.

3. Sectioning of Embryonic Intestines

1. Before proceeding with embryo dissections, melt 1.5 g agarose in 100 ml PBS (phosphate-buffered saline) and keep at 50 °C.
2. Pour melted agarose into an embedding mold (e.g. a closed 2 ml microcentrifuge tube that has been cut lengthwise to excise about 1/4 of the tube wall). Let the agarose cool down to approximately 42-45 °C, it should be only slightly warm to the touch.
3. Embed the embryonic intestine just before the agarose solidifies (this can be evaluated with forceps tips and will occur around 36-38 °C). Holding the intestine with forceps by the folded rostral end, pull it very slowly through the agarose along the length of the mold. This helps to keep the gut straight while the agarose sets. Release the tissue as soon as it begins to resist being moved. Keep track of the rostral-caudal orientation of the gut.
4. Put the mold in a refrigerator 2-3 minutes, to ensure the agarose has completely set.
5. Take out the agarose from the mold (by sliding it out of the opened Eppendorf tube). With a blade, take out the excess agarose at both ends, making cuts perpendicular to the intestine.
6. Glue the rostral end of the intestine/agarose block down on the metal stage of a microtome with vibrating blade. Trim the excess agarose on the sides of the intestine/agarose block.
7. Mount the metal stage on the vibrating microtome chamber. If necessary, adjust the angle of the stage so the intestine is as vertical as possible (hence perpendicular to the blade). Cover the specimen with ice-cold PBS. Bring the microtome blade down to a few millimeters under the buffer surface.
8. Make 200 μm vibratome transverse cuts of the caudal-most small intestine, ensuring that each agarose slice contains a full intestinal section.

4. Culture of Intestinal Explants

1. Gently deposit the freshly cut intestine/agarose slices flat onto the collagen gels with forceps, placing one slice towards the middle of each well.
2. Incubate 3 days at 37 °C, in a humid 5% CO₂ atmosphere, to allow migration of NCC out of the explant.
3. Take the intestine/agarose slices off the collagen gel very gently with forceps. Take care not to damage the gel below.
4. Avoid touching the collagen gel directly during the subsequent incubation and wash steps in order to ensure that the cell migration pattern is not disturbed. Fix with 500 µl of 4% PFA (paraformaldehyde) (in PBS) per well 1 hr at room temperature.
5. Replace the fixative with 500 µl of DAPI (4′,6-diamidino-2-phenylindole) solution (5 µg/ml in PBS) per well and incubate 10 min at room temperature.
6. Wash each well 3x with 500 µl of PBS for 5 min.
7. Photograph the fluorescent cells (GFP and DAPI channels) embedded in the collagen gel within each well.

5. Image Analysis

We made extensive use of ImageJ¹⁷ to process and quantify the images generated after explant culture.

1. Start by imaging a micrometer slide at the same magnification as the fluorescent cell photographs. Measure the length of one micron in pixels (using the Straight Line tool).
2. Set the scale (Analyze/Set Scale; input number of pixels/micron).
3. For each GFP fluorescence photograph, change the image format to 8-bit grayscale (Image/Type/8-bit).
4. Adjust intensity of the signal (Image/Adjust/Brightness/Contrast).
5. Subtract the background noise if necessary (Process/Subtract Background; adjust the rolling ball radius).
6. Set a threshold to highlight cells and cell clumps (Image/Adjust/Threshold), then apply a watershed to divide the clumps (Process/Binary/Watershed).
7. Analyze particles to specify regions of interest (ROI) and generate cell number statistics (Analyze/Analyze Particles; set size minimum to exclude remaining pixels).
8. Set measurement options to include Feret's diameter (Analyze/Set Measurements).
9. Group ROIs and measure as a whole to determine the Feret's diameter, an indication of cell spread (Analyze/Tools/ROI Manager/More/OR, then ROI Manager/Add, and ROI Manager/Measure).