Spinal Cord Neuron Isolation: A Density Gradient Procedure to Isolate Neonatal Spinal Cord Neurons for In Vitro Studies

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Harvesting Spinal Cords

NOTE: All instruments should be autoclaved (135 °C and 30 psi for 4 x 7 min cycles) for sterility.

1. Euthanize 1–3 day-old C57BL/6 mouse pups in a chamber with isoflurane. Wait 30 s after the cessation of movement and pinch the leg to confirm a lack of response.
2. Separate the head from the body using scissors, with pup in the prone position.
3. Stabilize the hind legs or tail and arms on the procedure table, with dorsal side facing the user.
4. Cut the skin off using curved iris scissors.
5. Cut the spinal cord from the lumbar region just above the hips and proceed to cut both sides of the thorax to separate it from the body.
   NOTE: This requires the careful dissection of the spinal cord from visceral organs to avoid inadvertent damage to other organs (Figure 1a).
6. Wash sequentially for 10 s in 3 x 10 cm Petri dishes containing 5 mL of 0.2 μm filter-sterilized Phosphate-Buffered Saline (PBS) to remove excess tissue.
7. Insert a 22G needle and syringe filled with 5 mL of filter-sterilized PBS into the caudal end of the spinal column and flush cranially, allowing the cord to exit into a fourth Petri dish (Figure 1b).
8. Collect the spinal cord in a 15 mL tube with 5 mL of HABG (Table 1) on ice. Use care to avoid crushing the spinal cord.
9. Repeat steps 1.1–1.8 for each pup in the litter.
   NOTE: Ideally, this process should take less than 30 min per spinal cord to ensure healthy neuron isolation.

2. Isolating Neurons

NOTE: The following step should be performed in a laminar flow hood. Familiarity with basic sterile technique is expected.

1. Tissue Mincing
   1. Take the tube containing the spinal cords and shake lightly to suspend the tissue.
   2. Pour tissue from the tube into a 60 mm glass Petri dish and dice with a razor blade to create fine pieces ~0.5 mm in size.
   3. Transfer the tissue with a wide-bore pipette into a 15 mL tube containing 5 mL of HABG.
   4. Place it in a 30 °C water bath for 30 min to allow the cells to equilibrate at this temperature. Keep the cells on a shaker just enough to allow them to be suspended in the fluid.
      NOTE: This step is done to avoid shocking the cells upon transfer from ice to the digestion medium. Keeping the cells at 30 °C helps to decrease cell death associated with an otherwise increased metabolism at 37 °C.
2. Prepare the digestion medium (Table 1).
3. Prepare the density gradient (Table 1).
   1. Prepare each of the 4 layers in 4 separate 15 mL tubes, as outlined in Table 1.
   2. Add 1 mL from each layer into a new 15 mL tube. Start with layer 1 at the bottom and sequentially add until reaching layer 4 at the top. Avoid disturbing the layers while adding.
4. Wash the PDL-coated plates. Wash with sterile water for a few minutes before removing any remaining water and allowing them to dry for 1 h.
5. Transfer the tissue to digestion medium.
   1. Remove the tissue-containing tube from the shaker water bath at 30 °C and allow it to settle for a few minutes.
   2. Remove the digestion medium tube from the 37 °C water bath and aspirate it into a leur-lock syringe.
   3. Aspirate off the excess HABG from the tissue-containing tube.
   4. Use a leur-lock 0.2 μm filter on the syringe to add digestion medium to the tissue-containing tube.
   5. Place the tube in a 30 °C water bath for 30 min. Keep the cells shaking just enough to allow them to be suspended in the fluid.
      NOTE: It is important not to keep the cells in the digestion medium for too long or to let the temperature get too high, which could lead to excessive digestion and result in the tissue becoming suspended in a gelatinous mixture.
6. During this period, coat the laminin.
7. Perform trituration (i.e. separating the cells from the tissue).
   1. Remove the tube from the shaking 30 °C water bath and allow it to settle for a few min.
   2. Aspirate excess digestion medium.
   3. Suspend the tissue in 2 mL of HABG.
   4. Using a narrow-bore pipette, triturate 10x for 45 s.
      NOTE: This is probably the single most crucial step and can significantly decrease the yield if not done properly.
      1. Aspirate the tissue into the pipette and immediately empty the contents back.
      2. Avoid introducing air, as it will significantly decrease the viable yield.
         NOTE: The ideal pipette is a 9" glass pipette. The tip of the pipette should be fire polished to smooth out rough surfaces. It should then be siliconized by placement in a 1:20 solution of dichlorodimethylsilane (DMDCS) in chloroform and left O/N. The pipette should then be removed and allowed to air dry. Subsequently, it should be autoclaved for sterilization.
         CAUTION: DMDCS and chloroform are highly flammable, and siliconizing should be carried out in a fume hood.
   5. Aspirate the top 2 mL of supernatant and place it into a new 15 mL tube labeled "collection".
   6. Repeat steps 2.7.3–2.7.5 two additional times (the cell collection tube should have 6 mL by the end).
   7. Slowly transfer the collection tube contents into the gradient tube prepared in step 2.2, avoiding the disruption of the gradient.
8. At this point, remove the previously prepared neurobasal medium (Table 1) from the refrigerator and allow it to warm in a 37 °C bath.
9. Purify the neurons.
   1. Centrifuge the gradient tube for 15 min at 800 x g and 22 °C.
   2. Collect the desired layer(s) with a pipette (Figure 2) and place in a new 15 mL tube. For the highest-purity neuron isolation (i.e. >90%), collect layer 3. For more yield with less purity (i.e. >70–80%), collect layers 2 & 3.
   3. Dilute out the density gradient by adding 5 mL of HABG to the newly collected layers.
   4. Centrifuge at 200 x g for 2 min at 22 °C.
   5. Discard the supernatant, resuspend in 5 mL of HABG, and flick the pellet to suspend the cells.
   6. Centrifuge at 200 x g for 2 min at 22 °C.
   7. Discard the supernatant, resuspend in 3 mL of neurobasal medium, and flick the pellet to resuspend the cells.

Table 1: Preparation of the media and solutions used in this protocol.

Figure 1: Spinal column dissected and spinal cord released from a 3 day-old neonatal mouse. The arrow in (a) points towards the caudal end of the spine, where a needle is inserted to release the spinal cord. A released spinal cord (b) is shown submerged in PBS.

Figure 2: Density gradient, with cells added-on after centrifugation. The layers are outlined and better visualized on the cartoon depiction. The most superficial layer (0) is generally debris from the spinal cord tissue. Layer 1 is rich in oligodendrocytes and astrocytes. Layers 2 and 3 contain the majority of neurons. While layer 3 contains neurons with the highest purity, some supporting cells (i.e., astrocytes and oligodendrocytes) are found in layer 2. The pellet at the bottom mainly contains microglial cells.