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All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Removal of P0-P2 mouse brain
- Right before starting the procedure, fill several petri dishes with chilled, carboxygenated sucrose artificial cerebrospinal fluid (sACSF). Also fill the collection tube(s) with ~25 mL sACSF.
- Wrap a postnatal day 0 to 2 (P0-P2) pup in some parafilm or a glove and place it well covered under the ice. Set a timer for 5 minutes and start it. After that time, remove the pup and check that it does not respond to pinching of the hind paw.
- Decapitate the pup and collect the head in a petri dish filled with sACSF. Cut the skin along the midline to expose the skull.
- Locate the opening in the skull at the hindbrain/spinal cord and insert one end of the forceps along dorsal portion of the skull. Gently grasp the skull at the midline and 'peel' away pieces laterally to expose the brain, being careful not to damage the brain.
- Upon removal of the dorsal and lateral portions of the skull, use the forceps or the spatula to gently remove the brain from the skull by scooping underneath the ventral surface of the brain, trying not to damage any area. Place the brain into another petri dish filled with carboxygenated sACSF. NOTE: We present two different strategies for dissecting out the hippocampus, striatum and cortex. The first strategy (steps 2.1-2.10) is useful for any mouse line whereas the second strategy (steps 3.1-3.7) would need a fluorescent reporter mouse to cleanly separate the striatum from the globus pallidus and other brain structures. If striatum is not desired, then ignore the striatum-specific portions of the two techniques
2. Harvest Striatum, Hippocampus and Cortex, Technique #1
- Place whole brain on sectioning matrice mold, ventral side up. Place 1 razor blade through the most anterior portion of the brain (through anterior olfactory tract).
- Insert another razor blade just posterior to the first one to generate a 0.5 mm slice and place an additional razor blade posterior to this one.
- Transfer these slices to a petri dish with carboxygenated sACSF and place the rest of the brain to a separate dish with sACSF. Transfer the striatal slices to a dissecting scope to visualize striatum. These slices should contain large portions of the anterior striatum and lack hippocampus. Use a fluorescent dissecting scope with Nkx2.1-Cre+/-;Ai9+/- slices to differentiate the weak striatal tdTomato signal from the strong tdTomato signal of the globus pallidus.
- With or without fluorescence, pinch out the striatum from both hemispheres on all the sections and transfer striatal chunks to properly labeled 50 mL tube with sACSF, store on ice.
- Hemisect the brain along the midline using forceps or a razor blade and lay the hemisphere so that the medial surface is facing up.
- Remove the ventral brain tissue (thalamus, basal ganglia, etc) by pinching off this tissue with forceps. When complete, the hippocampus (a sausage shaped structure spanning the anteroposterior axis along the dorsal cortex) and ventricular side of the cortex should be clearly visible.
- To remove the hippocampus, insert the tips of the forceps in front of the anterior hippocampus and gently separate the hippocampus from the cortex by moving the forceps posteriorly and pinching along the hippocampal-cortical border. Transfer the isolated hippocampus to properly labeled 50 mL tube with sACSF, store on ice.
- To dissect the cortex, place the hemisected cortex medial side down. Then use the forceps to pinch the most dorsal, ventral, anterior and posterior portions of the cortex to leave a square chunk of medial somatosensory cortex, ~2 mm x 2 mm. Flip the cortex so medial side is up and clean of any excess non-cortical tissue (thalamus, basal ganglia, etc.) on the medial surface if necessary.Transfer the cortex chunk to properly labeled 50 mL tube with sACSF, store on ice.
- Repeat the procedure with the other hemisphere to collect both hippocampi and cortex regions.
- Repeat this procedure for all pups, making sure to replace the sACSF in the petri dishes between pups to ensure that the sACSF remains cold and fresh.
3. Harvest Striatum, Hippocampus and Cortex, Technique #2
- Place the brain ventral side down in a sylgard-coated petri dish containing sACSF and pin it down through the cerebellum and anterior cortex or olfactory bulbs.
- Starting with one hemisphere, use curved forceps to gently separate the posterior cortex from underlying hippocampus and other tissue. Gently lay the peeled cortex flat on petri dish. Repeat for the other hemisphere. Hippocampi and striatum should be clearly visible in exposed brain.
- Use forceps to pinch one hippocampus at the midline and peel hippocampus laterally to remove. Place in collection vial on ice and repeat with other hippocampus.
- For the striatum, gently scrape around the edges of the striatum to loosen it from the surrounding tissue. Then, remove the striatum by pinching it off from underneath and add it to the collection tube on ice. Repeat for the other striatum.
- Cortical sections can be harvested as described in step 2.8.
- If using a transgenic reporter mouse line (e.g., Nkx2.1-Cre;Ai9, Lhx6-GFP, etc.), transfer the striatum to a petri dish with sACSF and view under a fluorescent dissecting scope. Use forceps to remove any non-striatal tissue from the striatum (in Nkx2.1-Cre;Ai9 mice, remove all 'bright' tissue, as globus pallidus has much higher medial ganglionic eminence or MGE-derived, tdTomato+ cell density compared to striatum). Repeat for all striatum.
- Repeat this procedure for all pups, making sure to replace the sACSF in the petri dishes between pups to ensure that the sACSF remains cold and fresh.
4. Generating Single Cell Dissociations
- After the dissection is complete, prepare a 1 mg/mL Pronase solution by weighing 10 mg Pronase and dissolving in 10 mL carboxygenated sACSF.
- Transfer the tissue from the 50 mL collection vials to 5 mL round bottom tubes containing 2 mL of the Pronase-sACSF solution. Incubate the tissue for 20 min at room temperature, shaking/flicking the tube 3-4 times with your finger during incubation to mix the samples.
- During this incubation, prepare Reconstitution solution: 1% fetal bovine serum (FBS, 100 μL) + DNAse (1 μL of 1:10,000 stock DNAse) in 10 mL carboxygenated sACSF.
- After the 20 mins, carefully remove the pronase solution to not disturb the tissue at the bottom and replace it with 1-2 mL of the Reconstitution solution (total volume is dependent on starting amount of tissue).
- Mechanically dissociate the tissue by triturating the tissue with the previously prepared fire-polished Pasteur pipettes. Starting with the large bore (~600 µm) pipette, aspirate and expel the tissue solution at least ten times to break up the tissue. Do not introduce bubbles into the solution. Repeat this process for the medium bore pipette and the small-bore pipette. The solution should be cloudy, and no clear piece of tissue should be visible in the collection tubes.
- Pipet the cell lysate solution through a 50 µm filter into 5 mL conical tubes to remove any cell clumps. Proceed with these single cell solutions to flow cytometry (or potentially directly to cell counting if no sorting is needed).
5. Preparing Fluorescence-Activated Cell Sorting (FACS)-purified Cell Solutions for Transplantation
- Upon receiving the cell solutions from the flow cytometer, transfer the solution to one (or multiple) 1.5 mL conical tubes and spin the cells at 500 g for 5 min at 4oC. After centrifugation, remove media so that ~20-40 µL remain in the tube. Reconstitute cells in this remaining media (and combine solution if multiple tubes were needed).
- On a small piece of parafilm, mix together 2 µL of cell solution + 8 µL sACSF + 10 µL of trypan blue stain. Pipet 10 µL into a hemocytometer with slide cover.
- Count the number of live cells in each of the 4 x 4 square grids in the corners of the hemocytometer using a standard laboratory hand tally counter (dead cells will be blue from the dye) and average these 4 counts. Multiply this number by 100 to determine the number of cells per µL.
- If needed, adjust volume so the cells are at a concentration of 10,000-30,000 cells/µL in Reconstitution solution. Final concentration is dependent on total amount of cells and desired number of transplantations. Keep cells on ice for transplantation
6. Transplantation Into P0-2 WT Pups
- Wrap a wild-type (WT) pup in some parafilm or a glove and place it well covered under the ice. Set a timer for 5 min and start it. After that time, remove the pup and check that it does not respond to pinching of the hind paw.
- While the pup is on ice, mix the cell solution by pipetting it several times to ensure the cell suspension is well mixed prior to loading (mix the cells prior to loading the micropipette for every injection). Then empty the micropipette of the mineral oil and fill it completely with the cell suspension.
- Place the anesthetized pup on the petri dish with his head lying on the adhesive putty so that the top of the head is relatively flat. Place the tape with the diamond hole across the head of the pup, pulling it taut to ensure the skin is stretched and the head is firm but that the mouse is comfortable and breathing well. Lambda should be visible through the diamond hole.
- Move the secured pup under the Nanoject and lower the micropipette so that the tip of the micropipette is on directly above lambda. Record the x-y coordinates on the manipulator.
- Adjust the manipulator knobs to move the micropipette to the desired coordinates along the mediolateral and the anteroposterior axis of the head: primary somatosensory cortex or S1 Cortex → 1.0 mm anterior and 1.0 mm lateral, Hippocampus → 0.75-1.0 mm anterior and 1.0-1.2 mm lateral, Striatum → 2.0-2.2 mm anterior and 1.5 mm lateral. Coordinates can be adjusted slightly to compensate for age of the pup (e.g., P0 vs. P2) or the strain of mice (e.g., CD1 mice are larger than and C57/B6 at P2).
- Lower the micropipette until it forms a small concavity on the skin. Then turn the z-axis manipulator knob firmly but gently to drive the micropipette through the skin and skull to enter the brain. When the micropipette enters the brain, the pressure on the skull will be released and the concavity will disappear.
- Retract the micropipette slightly until the tip is surrounded by a cone of skin, shaped like a tent. Read the coordinates along the z axes: This will be the z-axis zero point.
- Lower the micropipette to the desired depth: Cortex → 0.75-1.0 mm, Hippocampus → 1.2-1.5 mm, Striatum → 2.0-2.5 mm. Depth can be adjusted slightly to compensate for age or strain of the pup.
- Inject the cell suspension using the Nanoject III program described above. After injection program is complete, wait 15-20 s before slowly retracting the micropipette to minimize solution leaking out of the injection site.
- If performing bilateral injections, re-zero the micropipette above lambda and move to the proper coordinates in the other hemisphere. Alternatively, one can make two injections into the cortex of the same hemisphere by moving the micropipette 1 mm anterior of the first injection.
- Once the transplantation is complete, remove the tape and transfer the pup onto the heating pad. If necessary, tag the pup by tail or toe clipping. Once the pup has reacquired a red color and is moving, place it back in the cage with the mother.