An Explant Assay for Assessing Cellular Behavior of the Cranial Mesenchyme

1. Preparation of ECM Coated Culture Dishes or Coverslips

1. Prepare ECM stock solution by dissolving ECM substrate in PBS (Dulbecco’s Phosphate-Buffered Saline, Invitrogen, #14040-133). Filter using a 0.2 mm syringe filter (VWR, #28145-495) and freeze in aliquots at -80 °C.
2. Dilute ECM stock solution in PBS to desired concentration. For MaxGel, dilute in DMEM (Dulbecco’s Modified Eagle Medium, Invitrogen, #11995-065).
3. If immunofluorescence will be performed, sterilize 12 mm circular glass coverslips (VWR, #89015-724) by dipping in ethanol and allow to air dry. This should be done in the sterile laminar hood. If immunofluorescence will not be performed skip to step 1.5.
4. Place sterilized coverslips in each well of a 24 well plate (Corning, #3526).
5. Add 0.5 ml of diluted substrate solution to each well of 24 well plate and incubate at room temperature for 3 hr.
6. Aspirate off the ECM solution and wash three times in PBS.
7. Aspirate off PBS and use immediately or wrapped in parafilm and stored at 4 °C for up to 2 weeks.

2. Preparation of Dissection Tools

Sylgard plates with added charcoal are prepared per the manufacture’s protocol. The resistant black background of these plates provides support for pinning down the embryo and the color offers good contrast which facilitates visualizing the translucent embryo.

1. To prepare plates weigh directly into a tri-pour beaker on a balance 150 g Part A liquid 15 g Part B and 1 g charcoal powder (World Precision Instruments, #SYLG184). Mix together using a wooden popsicle stick or tongue depressor. Pour into glass or plastic 100 cm dishes. Use a small butane torch held about 10 to 12 inches above the dishes to burn off any bubbles. Place lids on plates and let sit for at least 24 hr to set.
2. To prepare dissection tools insert a Minutien pin (0.1 mm diameter) into a pin holder (Fine Scientific Tools, #26002-10 and #26018-17, respectively) alternatively 26 gauge needles (BD, # 309597) bent at right angles with the aid of forceps may be used as dissecting instruments. Sharpened Student Dumont #5 Forceps (Fine Science Tools, #91150-20) using a sharpening stone (Fine Scientific Tools, #29008-22).
3. Sterilize all dissection tools with 70% ethanol before use.

3. Collection of Embryos

1. Euthanize the timed pregnant mouse at 8.5 days post coitum according to animal use committee regulations and remove uterus.
2. Place uterus in a plastic Petri dish with sterile PBS.
3. Wash uterus once with sterile PBS.
4. Under a dissection microscope carefully remove embryo from deciduas using a pair of forceps.
5. Remove extraembryonic membranes and save for genotyping if needed.
6. Stage embryo by counting somites. The ideal stages to make explants to analyze cellular behavior during neural fold elevation would be between the 6 and 10 somite stages when the neural folds are just beginning to elevate and before the major morphogenesis of the cranial mesenchyme takes place.
7. Wash embryo in PBS. Place in a black sylgard plate with DMEM that contains no phenol red (Invitrogen, #21063-029). Remove the extraembryonic membranes and save for genotyping if needed.

4. Preparation of Explants

1. Refocus the microscope at the highest possible magnification over the head region of the embryo.
2. Make cuts with a dissection needle in each hand. Use one needle to hold the tissue in place and the other to make cuts.
3. Cut off the head of the embryo anterior to the rhombomeres.
4. Cut along the midline to bisect the head into two equal halves.
5. Cut around the outer fringes of the head and midline. These cuts will remove the premigratory crest and surface ectoderm at the outer border and the prechordal plate at the midline. The remaining tissue will be the explant and will contain the migrating cranial neural crest, paraxial mesoderm and surface ectoderm.
6. As the tissue is cut, remove dissected debris from the dish using a sterile pasteur pipette. This prevents confusing the explant with the dissected debris.
7. Gently wash the explant by pipeting up and down with a 20 μl pipette tip and pipetman. This will remove loosely attached cells and prevent jagged edges on the explants.
8. Carefully pick up each explant in approximately 10 μl of liquid using a 20 μl pipette tip and pipetman and place at the center of the coated culture well.
9. Add 20 μl of culture media supplemented with 15% FBS to cover the explant.
10. Place plate in a humidified tissue culture incubator (5% CO₂, 37 °C) for 1-2 hr to allow the explant to attach to the bottom of the dish. View periodically to ensure that the explant does not dry out.
11. When the explant has attached, carefully add to each well 250 μl DMEM supplemented with 10% FBS that has been warmed to 37 °C in a waterbath. Once explants have attached they will not move when plate is gently shaken under microscope.
12. Return dish to tissue culture incubator.
13. After 24 hr, the explants will be firmly attached and cells will begin to emigrate. At this time pharmacological agents can be added to the media.
14. 48 hr after plating, cells will have migrated from the explants. The distance migrated can be visualized and imaged. We typically stop the culture at this point since explants begin to show signs of decreased viability with longer incubations. A vital stain such as acridine orange or trypan blue can also be used to check viability of the explant.
15. At this point images can be acquired to document the distance and number of cells that migrate from the explant. Alternatively, if explants were plated on coverslips, they can be fixed and immunohistochemical analyses performed to visualize proteins of interest.

Critical steps in the protocol

1. Cut explants of approximately the same size.
2. Clear all dissection debris off the dissecting plate as you are dissecting to avoid confusing debris with the desired explanted tissue.
3. When plating explants in wells or on coverslips be sure to place droplet of media with explant in center of well.
4. Allow explant sufficient time to attach before flooding the well with media.
5. Treatment conditions should be standardized for every test substrate and pharmacological agent used.
6. Two explants can be made from each embryo (one from the left and one from the right of the neural tube).
7. When testing migration of cells from mutant mouse lines under different experimental conditions, put one mutant explant in the control group and the other in the treated group.

Troubleshooting

1. If explants fail to attach, increase the allotted time for attachment. In our experience, approximately half of the explants do not attach and larger explants adhere more efficiently.
2. If explants attach to the periphery of the well they will be hard to image. Be sure to place explants in center of the well.
3. Ideally explants will be approximately the same size. However, discrepancies in explant size can be corrected for using the diameter of the explants as a correction factor when quanitating migrations of cells from explants.
4. If contamination occurs be sure sterile working conditions are being used. All tools and work surfaces should be sterilized with 70% ethanol. Pipets and Petri dishes should be new and sterile.
5. If explants are getting lost during the dissection, periodically remove debris to prevent losing explant.
6. Mastering the dissection is time consuming and takes practice. Optimal magnification, sharp dissection tools and needles help.