A Modified Roller Tube Method for the Long-Term Culture of Rodent Brain Slices

All procedures involving sample collection have been performed in accordance with the institute's IRB guidelines.

NOTE: The protocol below describes the preparation and culture method for the long-term incubation and intermittent imaging of hippocampal slices. A single hippocampal slice is attached to a specially prepared photoetched coverslip using a plasma clot, and then the coverslips are sealed onto the flat side of a drilled-out roller tube, which is maintained in a roller incubator.

1. Preparation of Roller Tube Rack

1. Use the template shown in Figure 1A printed to the size shown on the scale bar. With a nail, punch small holes (large enough for a fine point marker) in the template centered on the holes.
2. Set the template on the bottom of a 15 cm tissue culture dish (nominal diameter of 14 cm) and mark the position of the holes. Repeat this on a second dish.
3. With drill bits designed for use on plastic, drill six 1.5 cm diameter holes on each dish in a hexagonal array (4.8 cm center-to-center) with hole centers 2.5 cm from the edge of the dish. Drill three holes (3 mm in diameter) 12 mm from the edge that are placed equidistantly between two of the larger holes as shown in Figure 1A.
4. With the bottoms of each dish facing each other, place a 2.5-inch-long machine screw (3/16-inch diameter) with a flat washer through one of the small holes followed by a second flat washer, a piece of polyethylene tubing (spacer, 4.7 cm), another flat washer, the second tissue culture dish, another flat washer, a locking washer, and a nut.
5. Repeat step 1.4 on the other two machine screws and tighten only loosely until all machine screws are in place. Then tighten the nuts securely.
6. Work the grommets (5/16 inch thick, 5/8-inch hole diameter) into the holes of the bottom dish to obtain the final roller tube rack (Figure 1B; shown with two tubes in place). Place a sticker on each rack with a unique number.

2. Preparation of Roller Tubes and Coverslips

1. Making the jig for drilling the hole in roller tubes
   1. Drill a 1.5 cm hole 8 cm deep in the center side of a 2 x 4 x 5.5-inch wooden block at an angle such that the flat side of the roller tube will be nearly parallel with the block when inserted (Figure 2A).
   2. Enlarge the hole using a round wood file to both widen and taper the hole to allow tube insertion (roller tubes are slightly larger in diameter near the cap end) (Figure 2B).
   3. Drill a 1.5 cm diameter vertical hole, 5.5 cm from the side of the block, and centered over the side hole (Figure 2C).
   4. When the side hole is tapered enough, insert a roller tube that is marked at the desired spot to center the hole for the slice and position the tube so the marked spot is centered in the 1.5 cm vertical hole.
   5. Remove the tube and measure the distance from the spot to the end of the tube. Mark this distance from the center of the hole in the jig and insert a nail to provide a stop to correctly position the tube for drilling (arrow in Figure 2C).
   6. Use a hacksaw to cut the nail off flush with the surface of the wood block to prevent injury.
   7. Add spring clips on the bottom of the jig if there is a drill press with slots that allow it to be anchored (Figure 2D black arrow). Otherwise, use C-clamps to hold the jig securely onto the drill press.
2. Using the jig described above to hold and position a flat sided 11 cm plastic culture tube with the flat side up (Figure 2A), drill a 6 mm diameter hole with the center 1.0 cm from the bottom and centered between the sides of the tube.
   NOTE: A drill bit designed for plastic should be used.
3.  With a swiveling deburring tool, smooth the edges of the hole (Figure 2E) and make 4 grooves on the inside edge of the hole (Figure 2E, inset) to facilitate draining of the hole during rotation.
4. With a 12 mm hole punch, cut 12 mm diameter disks from non-toxic double sided adhesive silicon rubber sheets. Using a standard one-hole paper punch (6 mm diameter), make a hole in the center of each disk.
5. Rinse the drilled tubes with 70% ethanol, air dry them in a biological safety cabinet, and sterilize the tubes and the punched adhesive discs for 40 min under the ultraviolet (UV) lamp (30 W at 70 cm average distance) in the biological safety cabinet.
6. Reposition the tubes and discs after 20 min so that all exposed surfaces are sterilized. Under sterile conditions, peel off the white backing from an adhesive disc and affix the silicone rubber to the outside of a tube, aligning the holes (Figure 2F).
   Caution: To avoid UV exposure, wear eye protection and close the cabinet before turning on the UV lamp.
7. Clean 12 mm diameter photoetched (100 center numbered 1 mm squares) German glass coverslips. Hold the coverslips gently with forceps and dip in absolute ethanol, followed by water, followed by absolute ethanol again, and finally dip the coverslips in a flame to burn off the ethanol. Allow coverslips to cool.
8. Holding the coverslips with forceps, dip into 2% 3-aminopropyltriethoxysilane in acetone for 10 s. Rinse the coverslips with ultrapure water and allow to air dry.
9. Set the coverslips on sterile filter paper inside of a biological safety cabinet and turn on the UV light. Expose each side of the coverslips for 20 min.
   Caution: To avoid UV exposure, wear eye protection and close the cabinet before turning on the UV lamp.

3. Hippocampal Slice Preparation

1. Before starting the dissection, prepare halves of double-edged razor blades for the tissue chopper. Fold the blades lengthwise carefully with fingers and snap in half.
2. Rinse the blade halves with acetone using a cotton swab to clean them, followed by rinsing in absolute ethanol and air drying. Before mounting a half blade on the tissue chopper, sterilize it by rinsing with 70% ethanol.
3. Follow protocols approved by the institutional Animal Care and Use Committee; after isoflurane anesthesia, euthanize a 4-7 day old mouse or rat pup by decapitation and remove the head with scissors or guillotine.
   NOTE: The brain slice culture protocol is independent of mouse or rat strain or genotype. Many transgenic mouse lines with different genetic backgrounds have been used.
4. Rinse the head with 70% ethanol, and place it in a 60 mm Petri dish. Until the final mounting of the brain slice onto the roller tube, all of the following steps are performed in a laminar flow hood to maintain sterility.
5. Using a #21 surgical blade, make a sagittal cut through the skin and skull. With a #5 Dumont forceps, peel back the skin and skull to expose the brain.
6. With closed forceps, gently tease out the whole brain, releasing it by pinching with the forceps through the brain stem behind the cerebellum. Place the brain in a sterile 60 mm dish containing 4 °C Gey's Balanced Salt Solution/0.5% glucose (GBSS/glucose).
7. Using a dissection microscope to visualize the brain (Figure 3A), place the brain dorsal side up and cut off the front third of the brain and the cerebellum with a surgical blade (Figure 3B).
   1. With forceps, hold the trimmed brain posterior side up and ventral side against the side of the Petri dish for stability. Gently tease away meninges around the sagittal midline and remove the midbrain tissue using a fine tipped Dumont #5 forceps (Figure 3C, dashed circle).
   2. Make two cuts along the side of the brain to spread it open (Figure 3C, dashed lines). Once the brain is placed dorsal side down and spread open, the hippocampal fissure should be visible (Figure 3D, arrow).
   3. Transfer the spread open brain to a piece of polychlorotrifluoroethylene plastic film and position it for slicing on the stage of a tissue chopper. Wet the blade with GBSS/glucose and chop the hippocampus into ~300 µm thick slices.
   4. With a transfer pipette, flush the sliced brain off the plastic film into a fresh 60 mm dish containing GBSS/glucose (Figure 3E). Gently pinch off and tease away, with fine tipped forceps, the remaining meninges and other non-hippocampal tissue (Figure 3F) from the slices (Figure 3G, H).

4. Plating Slices

1. Once slices have been obtained, place 2 µL of chicken plasma on the center of the photoetched side of a prepared coverslip. Spread the plasma slightly to achieve a 3-4 mm diameter spot.
   NOTE: The photoetched side is the top side of the coverslip when viewed through a dissection microscope such that the numbers are oriented correctly.
2. Transfer 1 brain slice with a sterile narrow-tip spatula (Figure 4A) to the plasma spot (Figure 4B). Use closed forceps to keep the slice on the spatula tip while lifting the slice from the GBSS/glucose.
3. Touch the spatula to the plasma spot on the coverslip, and with closed forceps, push the slice onto the coverslip.
4. Mix 2.5 µL of plasma with 2.5 µL of thrombin in a separate tube. Quickly place 2.5 µL of this mixture over and around the slice and pipet up and down gently to mix it (Figure 4B).
   NOTE: The plasma will clot within 10-15 s, so this must be done quickly. If slice adhesion is a problem, mix 5 µL plasma with 5 µL thrombin and use 4-5 µL on the slice, removing some after mixing so that the slice lies flat on the coverslip.
5. Remove the clear plastic covering from the exposed side of the silicone rubber adhesive previously affixed to a roller tube and place the coverslip with the brain slice onto the adhesive aligning the slice within the hole (Figure 4C).
6. To ensure adhesion, apply soft, even pressure to the coverslip with the thumb by pressing the coverslip down evenly and holding it for about 1 min while transferring it to the biological safety cabinet.
7. In a biological safety cabinet, add 0.8 mL of complete Neurobasal A culture medium (Table of Materials) to each tube (Figure 4D).
8. Flow a 5% CO₂/95% air mixture through a sterile cotton-plugged Pasteur pipette held securely by a clamp. Flush the roller tube with the gas mixture and rapidly cap the tube as it is withdrawn from around the pipette.
9. Label the tubes with the slice number and rack number. Insert tubes into a roller rack, ensuring they are geometrically balanced. If there is an odd number of tubes, add tubes to balance.
10. Place the racks in a 35 °C roller incubator with rollers turning the roller rack at about 10-13 Revolutions Per Hour (RPH) (Figure 4E). To keep the medium in the bottom of the tubes, tilt the incubator back approximately 5° by raising its front on a board.