DOI: 10.3791/51397-v
Embryonic neurons are born in the ventricular zone of the neural tube, but migrate to reach appropriate targets. Facial branchiomotor (FBM) neurons are a useful model to study neuronal migration. This protocol describes the wholemount ex vivo culture of mouse embryo hindbrains to investigate mechanisms that regulate FBM migration.
The overall goal of this procedure is to culture embryonic mouse hind brain's ex vivo, to study candidate molecules that may regulate the migration of facial brachi neuromotor neurons. This is accomplished by first micro dissecting embryonic day 11.5 mouse hind brains for culture in an open book preparation. The second step is to implant heparin beads coated with molecules of interest into the hind brains or to prepare media with pharmacological compounds.
Next, the dissected hind brain is carefully placed onto a culture insert. The final step is to incubate the explants for 24 hours at 37 degrees Celsius to allow the neurons to migrate ultimately immunofluorescence microscopy on whole mount hind brainin. Explants is used to reveal the effect of the candidate molecules on facial brachial motor neuron migration.
This method can help answer key questions in the neuronal migration field, such as understanding the attractive and repulsive signals that guide the migration of developing bronchi motor neurons, and identifying new guidance molecules On the day of ex planting. First, prepare the culture inserts in a flow hood by washing the inserts in sterile PBS for five minutes and then trying them for five to 10 minutes. Then place one culture insert into each well of a 12 Well plate pushing down as needed so they fit tightly into the well.
Cover the culture inserts with 10 to 20 micrograms per milliliter of mouse laminin in neuro basal medium, and place them into a tissue culture incubator. For one hour. After euthanizing a pregnant female mouse, place the uterus containing embryonic day 11.5 embryos into a 100 millimeter plastic dish with ice cold L 15 medium under a dissecting microscope.
Use forceps to tear the uterine muscle wall and expose the embryos. After releasing each embryo, sever the umbilical cord and carefully remove the yolk sack using a plastic pipette with a wide bore opening, transfer each embryo into a clean plastic dish with ice cold L 15. If the experiment requires genotyping of the embryos, collect a small piece of tail tissue for genomic DNA isolation.
Next, use forceps to decapitate the embryo just above the four limbs. Then use forceps to turn the head dorsal side up and identify the fourth ventricle, which is covered by a thin tissue layer. Carefully pierce the roof plate with the forceps and begin to peel it away.
Coddly along the midline over the posterior hind brain and spinal cord, and roly over the midbrain. The hind brainin should now be exposed. Carefully tease away the remaining head mesenchyme and any meninges that are attached to the peel side of the hind brain.
Remove the midbrain and spinal cord tissue so that the hind brainin unfurls and lies flat. In an open book preparation using a wide bore plastic paster pipette, transfer the dissected hind brain to a 12 well plate containing ice cold L 15, and store it on ice until all hind brains have been dissected. After all the hind brains have been dissected, use a wide bore plastic pasture pipette to transfer a single hind brain to an empty dish.
Keeping the hind brainin in an open book preparation ventricular side up. Add an approximately 100 microliter droplet of L 15. Retrieve a suspension containing previously prepared heparin beads, and transfer a few microliters of the bead suspension to the droplet containing the hind brain.
The suspension may contain approximately 10 to 100 beads. Now make a small tear in the hind brainin tissue at the level of the R five six boundary about halfway between the midline and the lateral edge of the hind brain. Now carefully insert one to three beads with an estimated diameter of approximately 200 microns by lowering them into the tissue.
Each bead should be positioned just beneath the hind brain surface so that it will not float away during subsequent culture. Continue to insert beads into all but two of the remaining hind brains, which will be cultured under normal growth conditions to serve as controls. Work quickly during the dissection so as to begin culturing the hind brains no later than three hours postmortem.
To prepare the hind brain X explan cultures. First, remove the plate containing the culture inserts from the incubator and aspirate the laminin coating solution. Place one culture insert into a separate culture dish filled with ice cold L 15, and using a wide boar plastic pasture pipette transfer each hind brain ventral side up onto the culture insert.
The hind brain should lie completely flat on the insert membrane. Carefully lift the culture insert from the dish and dab it several times on clean tissue paper to remove excess liquid. This process ensures that the hind brainin adheres to the culture.
Insert in a flat open book preparation check that the hind brain remains flat. If it curls up, flatten it by manipulating it gently with the pipette. Add 500 microliters of pre-war explant culture media to each well of the original 12 well plate, and then place the insert containing the hind brain back into its.
Well carefully adjust the volume with another 400 to 600 microliters of xplan culture media to just cover the hind brain, ensuring that the hind brain does not float off the membrane. At this stage, add any biological inhibitors of interest to study their effect on neuronal migration. Now incubate the explants for 24 to 30 hours in a tissue culture incubator.
Following incubation, aspirate the media from each well Then rinse the hind brains in PBS and fix in freshly prepared ice cold. 4%formaldehyde for two hours at four degrees Celsius with gentle agitation after fixation. Rinse each hind brain three times with PBS.
Now under the dissecting microscope, use forceps to carefully peel the hind brains from the culture inserts. If an explan is difficult to peel off, apply gentle pressure by repeatedly expelling PBS from a pipette. Transfer the hind brains to 2.0 milliliter round bottom tubes for immunofluorescence labeling.
After labeling and fixation, cover a glass slide with three layers of black electrical tape and use a scalpel to remove a small square of the layered tape to create a pocket for the x explants. Next, use a wide bore past pipette to transfer each hind brain into one pocket and remove excess PBS with a pipette. Next, cover each hind brain with slow fade reagent and then a glass cover slip being careful.
Avoid trapping air bubbles image the immuno labeled hind brain using a laser scanning confocal microscope. The facial brachi neuromotor neurons in explanted hind brains from day 11.5, mouse embryos undergo a tangential migration into streams on each side of the hind brain. As shown in this ventricular view, they then begin to assemble into the facial motor nuclei as shown in this peel view.
Similar to their behavior in utero, the midline is indicated with an asterisk in each panel. Embryonic day 11.5, litter mate hind brains were cultured in the presence of implanted heparin beads that had been soaked in either PBS or coated with V EEG F1 65. Note that the facial brachi neuromotor neurons on the hindbrain side containing the V EEG F1 65 bead migrated towards and onto the bead and the migrating stream, therefore extended further coddly compared to the untreated side of the same hind brain or the hind brain containing the control bead.Shown.
Here are examples of unsatisfactory E 11.5 hind brainin explants, in which the facial brachial motor neurons have not emigrated from R four or in which the hindbrain tissue folded during culture. While attempting this procedure, it's important to remember to dissect the hindbrain tissue carefully to avoid damaging it and to set up the explant cultures quickly to ensure that the neurons in the tissue remain viable.
