NOTE: Ethics Statement: The protocol for animal handling was approved by the Institutional Animal Care and Use Committee (IACUC) of the University of California, Irvine, protocol #2010-2943.

1. Removal of Spinal Cord

1. Place paper towels wetted with ~100% liquid isoflurane, USP in euthanizing chamber and place dry paper towels on top. Place mouse in chamber on top of dry paper towels so mouse is not touching the isoflurane and make sure the chamber is covered. Wait at least one minute after cessation of breathing to ensure that the mouse is euthanized.
2. Perform a spinal transection of the neck to ensure the mouse is euthanized. Do not perform a cervical dislocation as this could damage the spinal cord.
3. Spray the mouse with 70% ethanol to wet the hair.
4. Use sharp Fine scissors to remove the hair and skin from the back of the mouse to expose the spinal column from approximately cervical lamina 1 (C1) to sacral lamina 4 (S4).
5. Using a scalpel with a #10 blade, make incisions to the left and right of the spinal column to separate it from muscle and fat. This will make removal of the spinal cord much easier.
6. optional: a curved Luer rongeurs can be used to scoop away additional flesh.
7. While holding the top of the spinal vertebrae with serrated Graefe forceps, insert titanium curved Vannas scissors with the curved side up into the exposed spinal column at C1, being careful not to touch the spinal cord.
8. Slide the titanium curved Vannas scissors all the way to the right and make one small cut. A small crunch sound should be heard and felt as the scissors cut the vertebrae. Repeat this cut on the far left side of the cord. Be careful not to go too quickly or make too large of a cut as this will risk damaging the cord with the scissors.
9. The single lamina should be able to be lifted with the forceps once the sides of the vertebrae are cut. Repeat this step for each lamina until S4, continuing to hold and pull back the spinal cord vertebrae.
   NOTE: the vertebrae above the transplant site will likely come off once it is cut down to the transplant site. Continue the procedure starting just below the transplant site.
10. optional: lay the vertebrae down to see where the transplant site is and measure and cut ~20 mm rostral and caudal to the transplant site.
    NOTE: When transplanting NPCs more will not be needed as the transplanted NPCs generally do not migrate farther than 15 mm. The amount of spinal cord needed will be dependent on experiment and migration characteristics of cell type transplanted. Making rostral and caudal cuts that are equidistant to the transplant site will enable the transplant site to be located more easily during imaging.
11. Using a scalpel with a #11 blade carefully cut the ganglia on the right and left of the ventral side of the spinal cord starting at the rostral end. This will enable the spinal cord to be removed from the ventral vertebrae more easily.
    NOTE: Done quickly (steps 1.7-1.11), the spinal cord will not dry out. However, 1x PBS can be administered to the cord to keep it moist.
12. Invert the mouse and hold the mouse up so the spinal cord is facing down. Carefully peel out the spinal cord using closed serrated Graefe forceps. Without nicking the spinal cord, carefully cut any remaining ganglia to allow the spinal cord to be lifted out in a single intact piece.
13. Make sure the spinal cord is in RPMI-1640 and place it on ice for transport to the 2P microscope.

2. Preparation of Spinal Cord for Imaging

1. Embedding spinal cord in agarose
   1. Keep the isolated spinal cord in chilled RPMI-1640 on ice prior to imaging.
   2. Weigh out agarose (low gelling temperature) and prepare a 5% solution in 5 ml of 1x PBS. Microwave the 5% agarose solution for 15 sec to dissolve the agarose and let the solution cool to 37 °C.
   3. Prepare the spinal cord by placing it on a sheet of Parafilm with the ventral side facing up.
   4. Pipet approximately 5 ml of 5% agarose solution over the ventral side of the spinal cord. Let the agarose solidify by cooling to room temperature. Full solidification takes approximately 5 min.
   5. Apply a light coat of tissue adhesive to a 22 mm square cover slip.
   6. Invert the embedded spinal cord, placing the ventral side on Parafilm. The dorsal side will now be facing up. Adhere the cover slip to the dorsal side, and submerge the agarose embedded spinal cord/cover slip preparation in RMPI to solidify the adhesive. Remove excess solidified agarose using a razor blade.
2. Mounting the spinal cord on the microscope stage
   1. Apply petroleum jelly to the bottom of the cover slip. This will stabilize the preparation during perfusion.
   2. Place the spinal cord preparation in an imaging well on the microscope stage with the ventral side facing up toward the 25X dipping objective. The custom-built imaging well is approximately 20 mm deep, 50 mm long, and 50 mm across.
   3. Image while superfusing the preparation with warmed (37 °C), oxygenated (95:5 oxygen:carbon dioxide-carbogen) RPMI-1640 medium without serum.
      1. Pre-warm media to 37 °C in the water bath and connect it to the tubing pump by inserting the tubing into the media bottle.
      2. Retain the media temperature at 37 °C a heater device at the connection of the tubing to the chamber. Superfuse media through well at 3 ml/min using tubing connected to the tubing pump (Figure 1B).

3. 2P Imaging of Ventral Spinal Cord

1. Microscope Setup
   1. Acquire images from Thy1-eYFP spinal cords using a Chameleon Ultra Ti: Sapphire laser tuned to 900 nm. Attenuate laser power at the specimen to <5% to ensure minimal phototoxicity¹⁶. Maintain temperature by perfusing oxygen-perfused RPMI-1640 at a constant 37 °C using a single inline solution heater to ensure stable imaging, and to prevent tissue drift and cellular damage.
   2. To separate eGFP and eYFP fluorescent signal, place a 520 nm single-edge dichroic and a 560 nm single-edge dichroic beam splitter in series to separate 2P emission into three channels, purposely splitting the green emission to enhance visibility.
      NOTE: These channel are referred to as blue-green (emission <520 nm), green-yellow (520–560 nm), and red (emission >560 nm). Photomultiplier tubes detect emitted light in each channel.
2. Locating imaging areas of interest
   1. Using the eyepiece and a bright field light source, focus the dipping objective at the ventral edge of the spinal cord to set a reference point.
   2. Make sure ambient light source is off and switch to 2P excitation by opening the laser shutter.
   3. If available, when searching tissue for areas of interest, use a lower resolution setting, higher volume without digital zoom, and higher scan rate than when acquiring final images. Typical settings with this system when searching tissue for a region of interest are: resolution: 256 pixels; volume: x = 600 μm, y = 600 μm, z = 0-300 μm.
   4. Observe the eYFP axons near the ventral edge of the spinal cord. Second harmonic signal from collagen (blue) will be brightest at the spinal cord tissue edge. Locate the eYFP axons just dorsal to the second harmonic signal from the collagen and eYFP signal is visualized in the green-yellow channel.
   5. Locate the transplant site in the longitudinal center of the spinal cord preparation¹⁴. For mice 1 day following eGFP-NPC transplantation, locate the transplant site by focusing the beam path in the z plane deep into the tissue. The transplant site will vary slightly between animals but is generally located ~200 μm from the ventral edge. Observe the clusters of eGFP-NPCs in the white matter tracts, closer to the ventral and lateral edges in mice after day 1 post-transplantation.
3. Acquiring final images
   1. Acquire image resolution of 512 pixels; volume: x = 270 μm, y =212 μm, and z = 100 μm using Slidebook 6 software. Compile z-stacks by acquiring sequential focal planes in 2.5 μm increments.
   2. Perform a bidirectional scan with proper interlace offset to ensure rapid quantification of any migrating objects in the spinal cord. Ensure that the ideal acquisition frame rate to determine cellular velocity within the spinal cord is approximately 1 frame/sec.
      NOTE: Imaging settings can be changed to individual user preferences, such as acquisition frame rate, imaging resolution and imaging volumes. Larger imaging volumes will generally take longer to acquire at a given resolution.
   3. Analyze, crop, smooth, and pseudocolor Slidebook image files with image analysis software such as Bitplane Imaris 7.7. Produce final time-lapse videos by combing consecutive imaging volumes using an automated process in Slidebook and Imaris.