Obtaining Various Neural Cell Populations from a Rat Hippocampus

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

1. Tissue Collection

1. Prepare Buffer X from the Neural Dissociation Kit by adding beta-mercaptoethanol to Buffer X to a final concentration of 0.067 mM (for 4 samples, add 13.5 µl of 50 mM beta-mercaptoethanol to 10 ml of Buffer X).
2. Prepare Enzyme Mix 1 by mixing 50 µl of Enzyme P with 1,900 µl of Buffer X for each sample as described in the protocol for the Neural Dissociation Kit For Example, for 4 samples, mix 200 µl of Enzyme P with 7,600 µl of Buffer X.
3. Place Buffer X in the water bath at 37 ºC.
4. Place 2 ml of cold Hank's Buffered Salt Solution (HBSS) (without Ca⁺⁺ and Mg⁺⁺) into one well of a sterile 6-well culture plate for each sample that will be collected and kept on ice for tissue dissection.

2. Neural Dissociation (1 hr)

1. Dice each tissue sample into small pieces using sterile razor blades on the lid of the 6-well culture plate. This protocol has been used for tissue samples of approximately 30 – 100 mg, but is approved for tissue samples of up to 400 mg. For tissue samples larger than 400 mg, the reagent volumes must be scaled up appropriately.
2. Use 1 ml of fresh HBSS without Ca²⁺ and Mg²⁺ to transfer the diced pieces of tissue with a pipette into a sterile 2 ml centrifuge tube. Repeat steps 2.1 and 2.2 for all sample until all samples have been transferred into a 2 ml centrifuge tube.
3. Centrifuge the samples at 300 x g for 2 min at RT and aspirate supernatant.
4. Add 1,900 µl of warm Enzyme Mix 1, as prepared in Steps 1.2 and 1.3, to each sample and close the tubes.
5. Incubate the samples for 15 min in the 37 ºC water bath, inverting the tubes several times every 5 min to re-suspend the settled pieces of tissue.
6. Meanwhile, prepare Enzyme Mix 2 by mixing 20 µl of Buffer Y with 10 µl of thawed Enzyme A (for 4 samples, mix 80 µl of Buffer Y with 40 µl of thawed Enzyme A).
7. Add 30 µl of Enzyme Mix 2 to each sample and invert gently. Do not vortex.
8. Dissociate each tissue sample with Pasteur pipette "1", triturating up and down 30 times.
9. Incubate the samples for 15 min in the 37 ºC water bath; invert the tubes several times every 5 min to re-suspend the settled pieces of tissue.
10. Dissociate each tissue sample with Pasteur pipette "2," triturating up and down 30 times, avoiding generating bubbles.
11. Dissociate each tissue sample with Pasteur pipette "3," triturating up and down 30 times, avoiding generating bubbles.
12. Incubate the samples for 10 min in the 37 ºC water bath; invert the tubes several times every 5 min to re-suspend the settled cells.
13. Apply single-cell suspension to an 80-micron cell strain placed on top of a 15 ml falcon tube to remove any large pieces of debris and wash the filter and cells with 10 ml of HBSS (with Ca²⁺ and Mg²⁺) to stop the enzyme reactions. Repeat until each sample is transferred into a clean 15 ml falcon tube.
14. Centrifuge the samples at 300 x g for 10 min at RT and aspirate off the supernatant.

3. Myelin Depletion (45 min)

1. Thoroughly re-suspend the pellet of cells with 400 µl of myelin removal buffer.
2. Add a specified amount of the myelin removal beads and pipette up and down to thoroughly mix.
   NOTE: The volume of myelin removal beads used will depend upon the size of the tissue, the age of the animal, and the amount of myelin expected in that particular brain region. For example, for an adult rat hippocampus sample (approximately 300 mg), incubate the sample with 100 µl of myelin removal beads.
3. Incubate for 15 min in the refrigerator at 4 ºC.
4. Wash each sample with 5 ml of myelin removal buffer.
5. Centrifuge the samples at 300 x g for 10 min at RT.
6. While centrifuging the samples, place 1 column for each sample in the magnetic field of the magnetic sorter and place a clean 80-micron filter on top of each column.
7. Prepare the columns and filter by rinsing each column with 1 ml of myelin removal buffer three times (3 ml total). Collect all flow-through in a waste container, such as the bottom of an empty tip box.
8. Position a 5 ml polystyrene round bottom tube directly underneath each column in preparation of sample collection.
9. When the cells have finished centrifugation, aspirate the supernatant and thoroughly re-suspend it in 500 µl of myelin removal buffer. Immediately apply the cell suspension to the column and collect the cells in the tube below.
10. Wash the filter and column with 1 ml myelin removal buffer, 4 times (4 ml total).
11. Centrifuge the cell collection at 300 x g for 10 min at RT.
12. Aspirate the supernatant and the cells are now ready for staining.

4. Staining Live Cells for FACS (1 hr for staining)

1. Briefly vortex (2 sec) the cells to separate the pellet and immediately add 5 µl of Fc block (anti-clusters of differentiation 32, CD32), vortex again, and incubate in the refrigerator at 4 ºC for 5 min.
2. Meanwhile, prepare the primary antibody mixture by adding the three primary antibodies in wash buffer as follows: 0.125 µl of allophycocyanin (APC) -conjugated CD11b antibody (1:800), 1 µl of anti-GLT1 antibody (1:100), and 0.4 µl of Fluorescein isothiocyanate (FITC) – conjugated Thy1 antibody (1:250) for every 100 µl of wash buffer. Each sample will receive 100 µl of primary antibody mixture for incubation; however, if your sample is much larger than 400 mg of tissue, you may want to consider increasing the volume of your primary antibody mixture accordingly
3. Prepare tubes for single staining controls by removing just 2 µl of cells from each sample tube and adding it to each single stain control tube. The "single stain" controls include: APC-CD11b only, GLT1 only, FITC-Thy1 only, PE secondary antibody only, and the "blank" or unstained control.
   NOTE: All samples should be represented in all staining controls, thus 2 µl of cells from each sample should be added to each staining control tube.
4. Briefly vortex the tubes (2 sec) and add 100 µl of primary antibody mixture to each sample tube and 100 µl of appropriate primary antibody to each staining control tube, vortex the tubes again, cover the tubes with aluminum foil to protect the fluorescent antibodies from any light, and incubate the tubes in the refrigerator at 4 ºC for 20 min. The samples should be kept cool (at 4 ºC) for the remainder of the procedure to maintain the integrity of the samples and the antibodies.
5. Briefly vortex the tubes and add 2 ml of wash buffer to each tube.
6. Centrifuge tubes at 350 x g for 5 min at 4 ºC.
7. Meanwhile, prepare the secondary antibody mixture by adding 0.2 µl of PE-conjugated anti-rabbit secondary antibody for every 100 µl of wash buffer. Each sample will get 100 µl of secondary antibody mixture; however, if your sample is much larger than 400 mg of tissue, you may want to consider increasing the volume of your primary antibody mixture accordingly.
8. When the centrifugation is complete, dump the supernatant off of each sample into the sink and blot the tubes upside-down on a paper towel.
9. Briefly vortex the tubes and add 100 µl of secondary antibody mixture to each sample tube and the appropriate secondary antibody to each staining control. Then, vortex the tubes again, cover them with aluminum foil, and incubate them in the refrigerator at 4ºC for 15 min.
   NOTE: The GLT1-only staining control tube should get the secondary antibody mixture.
10. After the incubation, briefly vortex the tubes and add 2 ml of wash buffer to each tube.
11. Centrifuge the tubes at 350 x g for 5 min at 4ºC.
12. When the centrifugation is complete, dump the supernatant off of each sample into the sink and blot the tubes upside-down on a paper towel.
13. Vortex the tubes and add 0.25 ml of sterile PBS to each tube. The samples are ready for sorting.
    NOTE: More cells may require 0.5 ml of sterile PBS; however it is best to begin with a lower volume as the sample can always be diluted if it is too concentrated with cells.
    NOTE: Optionally add DNase to the tube to prevent the cells from clumping and thus clogging the sorter. Samples can also be filtered using a sterile 80 micron filter just prior to sorting to avoid clogging the sorter.