Preparation of Rat Oligodendrocyte Progenitor Cultures and Quantification of Oligodendrogenesis Using Dual-infrared Fluorescence Scanning

Procedures involving animal subjects have been approved by Institutional Animal Care and Use Committee (IACUC) and Johns Hopkins School of Medicine.

1. Preparation of Assay Plates, Stock Solutions, and OPC Base Media

Note: The rat OPC base (Sato) media described here has been derived from previously published studies^12,13. Alternative media formulations may also be compatible with this procedure.

1. Resuspend poly-L-lysine (PLL) to a concentration of 10 µg/ml in sterile deionized water. Pipette 400 μl of the diluted PLL into each well of a black-walled, clear bottom 24 well tissue culture grade dish.
2. Coat tissue culture dishes for 2 hr in a 37 °C tissue culture incubator or overnight in a 4 °C refrigerator.
3. Aspirate PLL from coated dishes at least 20 min prior to plating. Allow the remaining PLL to dry from the well by placing 24 well plates with the lid partially ajar in a tissue culture hood. Verify that the wells are completely dry before plating isolated OPCs. Cells will not adhere to plastic that has liquid PLL present.
4. Prepare N-Acetyl-L-cysteine (NAC) stock solution (1,000x): Dissolve 100 mg N-Acetyl-L-cysteine (NAC) in 20 ml of Dulbecco's modified Eagle's medium (DMEM). Aliquot and store at -20 °C.
5. Prepare Hydrocortisone stock solution (1,000x): Add 1 ml of ethanol to 1 mg Hydrocortisone and swirl to dissolve. Add 19 ml of DMEM, mix the solution, aliquot and store at -20 °C. The addition of hydrocortisone to the base media has been shown to enhance the survival of glial cells¹⁴.
6. Prepare d-Biotin stock solution (5,000x): Dissolve 2.5 mg of d-biotin in 50 ml of PBS. Add 1-2 x 5 µl drops of 0.1 N NaOH to aid in dissolution. Aliquot and store at -20 °C.
7. Prepare Insulin stock solution (100x): Dissolve 25 mg Insulin in 50 ml of tissue culture grade water. Add 250 µl of 1 N HCl and mix until solution is clear. Sterilize with a 0.22 µm filter and store at 4 °C for up to 6 weeks.
8. Prepare Sato stock solution (100x):
   1. Prepare progesterone stock solution (25 µg/µl): Dissolve 2.5 mg progesterone in 100 µl of ethanol.
   2. Prepare sodium selenite stock solution (400 ng/µl): Dissolve 4 mg sodium selenite in 100 µl of 0.1 N NaOH. Add 10 ml of DMEM and mix.
   3. To 100 ml of DMEM, add 1 g of human apo-Transferrin, 1 g of bovine serum albumin, and 160 mg of putrescine and mix to fully dissolve. Add 25 µl of progesterone stock solution, 1,000 µl of sodium selenite stock solution, and mix. Aliquot and store at -20 °C.
9. Prepare OPC Base Media: per 100 ml of DMEM (with 4.5 g/L D-glucose, L-glutamine, and 110 mg/L sodium pyruvate), add 100 µl NAC, 100 µl hydrocortisone, 20 µl d-Biotin, 1 ml insulin, 1 ml Sato stock, 100 µl trace elements B, 2 mL B-27 supplement (50x), and 1 ml of penicillin/streptomycin (100x). Sterilize with a 0.22 µm filter and store at 4 °C.
10. Prepare PDGF-AA stock solution (1,000x): Dissolve 250 μg in 12.5 ml of PBS with 0.1% bovine serum albumin (BSA) to make a 20 μg/ml solution. Aliquot and store at -80 °C.
11. Prepare OPC proliferation media: OPC base media supplemented with 20 ng/ml PDGF-AA.
12. Prepare OPC differentiation media: OPC base media supplemented with 45 nM triiodothyronine.
13. Prepare Column Buffer: To 500 ml of PBS, add 2.5 g of BSA and 2 ml of 0.5 M ethylenediaminetetraacetic acid (EDTA) and adjust the pH to 7.2. Sterilize with a 0.22 µm filter and store at 4 °C.

2. Rat Brain Dissection

Note: P5-P7 rat pups are used in this protocol. Each rat pup cortex yields approximately 1.5-2.0 x 10⁶ A2B5 positive cells.

1. Sterilize all dissection equipment using a 250 °C heated bead sterilizer.
2. Add 15-20 ml of PBS (without Ca²⁺ and Mg²⁺) to 50 ml conical tubes. One 50 ml conical tube is sufficient for 3-4 rat pup cortices. Place 50 ml conical tubes on ice. Use 50 ml conical tubes to hold diced cortex tissue during dissection.
3. Add cold PBS to a 10 cm Petri dish. This dish will be used for dissection under the light microscope.
4. Sacrifice rat pups by decapitation with large surgical scissors. Spray head with 70% ethanol.
5. Extract the Whole Brain
   1. Using small scissors cut the skin down the midline and behind the ears and peel skin flaps back. Cut the cranium down the midline starting from the brainstem and ending at the eyes. Be careful not to cut too deeply in order to preserve the structure of the underlying cortex.
   2. Make two lateral cuts inferior to the cerebellum by inserting small surgical scissors into the foramen magnum. Make an additional cut between the eyes, anterior to the olfactory bulbs.
   3. Carefully peel each half of the cranium back. Remove the whole brain and place in the 10 cm Petri dish filled with cold PBS.
6. Under the dissection light microscope, remove the olfactory bulbs and cerebellum with fine surgical forceps.
7. Flip the brain so that the ventral surface is visible under the light microscope.
8. Using fine straight forceps perform a blunt dissection by placing closed forceps tips between the cortex and hypothalamus to a depth about 2/3 of the brain. Once forceps are in place allow the ends to open. Repeat for the other hemisphere.
9. Tease the cortex from the hypothalamus and midbrain regions.
10. Remove the hypothalamus, thalamus, and midbrain by holding the midbrain at its posterior surface and peeling it towards the anterior end of the brain. Sever the anterior connections.
11. Remove the hippocampus by pealing it outward and then severing its connection to the cortex using fine bent dissection forceps.
12. Remove remaining striatum by scrapping it from the underlying cortex in an outward diagonal motion using fine bent dissection forceps.
13. Clear any blood vessels and meninges from the ventral surface of the cortex.
14. Flip the brain so that the dorsal surface is visible. Meninges and blood vessels should be apparent. Peel meninges from the underlying cortex using fine dissection forceps. Start peeling from the olfactory bulb attachment location.
15. Complete steps 2.4-2.14 for a total of 3-4 rat pups.
16. Place 3-4 dissected rat pup cortices into a dry petri dish and chop with a sterilized razor blade until 1 mm x 1 mm chunks are achieved. Collect tissue by rinsing dish with PBS from one 50 ml conical tube (step 2) then place back on ice.

3. Enzymatic and Mechanical Tissue Dissociation

Note: For this step, a papain-based neural dissociation kit is recommended. Using Neural Dissociation Kit (P), special steps that are optimal for OPC isolation are described.

1. Prepare the first dissociation enzyme by diluting enzyme P with the appropriate buffer. The contents of each 50 ml conical (1 sample) will be resuspended with a total of 1,950 µl of enzyme mix. Place in the enzyme mix in a 37 °C bath for 10 min.
        1 Sample:   1,900 µl of Buffer + 50 µl of papain enzyme
2. Spin all 50 ml conical tubes containing 3-4 diced rat pup cortices at 300 x g for 3 min.
3. Aspirate the supernatant and add 1,950 µl of enzyme solution to each sample tube and break apart the pellet by inverting the tube and shaking gently.
4. Incubate in a 37 °C tissue culture incubator for 15 min with continuous tube rotation.
5. During the last 5 min of the incubation, prepare the second enzyme mix A. Dilute the enzyme in its appropriate buffer. A total of 30 µl will be added to each 50 ml conical sample tube.
        1 Sample:   20 µl of Buffer + 10 µl of enzyme
6. Once incubation is completed add 30 µl of the second enzyme mix to each tube and invert to mix.
7. Using a glass Pasteur pipette attached to a pipette controller, mechanically dissociate the tissue by pipetting 10-15 times or until the tissue pieces are reduced in size and the solution has become cloudy. Return the sample to the 37 °C tissue culture incubator for 15 min with continuous rotation.
8. Pipette each tissue homogenate sample 10-15 times using a 1 ml pipette.
9. Pipette each tissue homogenate sample 15-20 times using a 200 µl pipette
10. Return all samples to the 37 °C tissue culture incubator for 10 min with continuous rotation.
11. Add 10 ml of PBS (with Ca²⁺ and Mg²⁺) to each sample and filter the homogenate through a 40 µm pore filter placed over a 50 ml tube. Rinse the filter with an additional 1-2 ml of PBS.
12. Pool all of the homogenate samples into one 50 ml conical tube and acquire an overall cell count.
13. After performing the cell count, split the homogenate evenly into 15 ml conical tubes (1 tube for each sample). Centrifuge for 10-12 min at 300 x g.

4. Anti-A2B5 Bead Application, Column Purification, and Plating

1. Perform calculations for column buffer and anti-A2B5 microbeads. Add 7 μl Column Buffer for every 1 x 10⁶ cells. Add 2 μl Anti-A2B5 microbeads for every 1 x 10⁶ cells.
2. Combine all of the samples by resuspending the cells in a total volume of 10 ml of column buffer and centrifuge at 300 x g for 10-12 min.
3. Resuspend the cell pellet in the appropriate amount of column buffer as calculated in step 4.1. If the pellet is large and loose, only add approximately half of the volume of column buffer to keep the antibody at the appropriate concentration in the cell resuspension.
4. Incubate the cell suspension for 10 min at 4 °C.
5. Add anti-A2B5 microbeads (calculated in step 4.1), invert several times and incubate at 4 °C for 30-60 min. Gently invert the tube several times every 10 min. Longer incubation times may increase cell yields but may increase non-specific binding.
6. Add 5 volumes of column buffer. If the cells are resuspended in a volume of 1 ml, add 5 ml of column buffer, whereas if the cells are resuspended in 2 ml, add 10 ml of column buffer.
7. Centrifuge for 10 min at 300 x g.
8. Determine how many LS columns will be needed for the purification. One LS column is sufficient for 15-20 x 10⁶ total cells.
9. Resuspend the cell pellet in 1,000 µl of column buffer per LS column being used.
10. Prime each LS column by adding 5 ml of column buffer. Collect the flow through in a 50 ml conical tube. Once the column buffer has completely passed through the upper chamber, apply 1,000 µl of the cell suspension to each column and allow the cells to run through by gravity.
    Note: If the density of cells is too high, the column may run slow.
11. Immediately after the cell suspension has passed through the column, slowly add 3 ml of column buffer to each column to wash the unbound cells. If the wash is readily running through the column (1 drop for every 30-45 sec), wash two additional times with 3 ml of column buffer.
12. Remove each column and place it on a 15 ml conical tube. Add 5 ml of column buffer and plunge the buffer through the column at a fast rate using the plastic plunger that is packaged with the column. Plunging will dislodge the bound cells releasing them from the column.
13. Increase purity by running the sample through a second round of LS columns. Use one third of the number of columns used during the first pass. Prime the columns with 5 ml of column buffer. Allow the column buffer to pass through the upper chamber.
14. Since the volume of cell suspension will be much greater, evenly apply 1,000 µl of cell suspension to each column and allow the suspension to pass through the upper chamber of the column before replenishing with an additional 1,000 µl.
15. Once the cell suspension has been completely applied to the second round of columns, wash 2-3 times with 3 ml of column buffer.
16. Remove each column and place it over a sterile 15 ml conical tube. Add 5 ml of column buffer and plunge the buffer using the plastic plunger that is packaged with the column. Plunging will dislodge the bound cells releasing them from the column.
17. Centrifuge the cell suspension at 300 x g for 12-15 min. The pellet should appear compact.
18. Resuspend the cell pellet in 5-10 ml of pre-warmed OPC proliferation media (OPC base media supplemented with PDGF-AA 20 ng/ml).
19. Count the cells using a hemocytometer.
20. Plate the cells. Dilute the cells to 60,000 cells/ml with OPC proliferation media. Into each black, clear bottom 24 well, pipet 500 µl of cells along the side of the well to obtain 30,000 cells per well. Mix the cells by shaking the plate horizontally using a figure eight motion. If performing the assay in 48, 96, or 384-well plates, add 15,000, 7,500, or 1,500 cells per well, respectively. These numbers may be adjusted depending on individual plate specifications.
21. Prepare a separate plate for the Day 0 baseline control cultures. This control plate should be fixed with 4% paraformaldehyde as described in section 5.7, when differentiation is initiated on Day 0.
22. Culture the cells in OPC proliferation media at 37 °C for 3 days with no media change.

5. Induction of OPC Differentiation and Fixation with 4% Paraformaldehyde

Note: When testing the effect of small molecules on oligodendrogenesis, we routinely dissolve drugs in dimethyl sulfoxide (DMSO) to prepare 1,000x stocks that can then be stored at -80 °C and diluted directly into SATO media to obtain a final concentration of 0.1% DMSO. We have observed no changes in either OPC proliferation or differentiation using this concentration of DMSO (data not shown).

1. Pre-warm OPC base media to 37 °C and thaw drug treatment stocks to room temperature. When performing treatments in duplicate, prepare approximately 1.25 ml of media per treatment group in a 1.5 ml tube. For triplicate samples, use 2 ml capacity centrifuge tubes to account for slack.
2. Prepare the treatment master mixes for replicate wells by diluting treatment stocks directly into OPC base media. Briefly vortex or gently mix each mastermix to ensure homogeneous distribution of the treatment.
3. Prepare 45 nM triiodothyronine (T₃) as the positive control and the appropriate vehicle as the negative control. Dilute 10 mM T₃ stock 1:1,000 in 1 ml of OPC base media and then further dilute in the treatment master mix to reduce the final concentration of DMSO if necessary.
4. Aspirate the media from the culture wells one treatment group at a time so as to avoid drying the cells. Use a 200 µl pipet tip on the end of the glass Pasteur pipet to avoid scraping off black material from the sides of the well and into the culture.
5. Slowly add 500 µl of treatment mastermix along the side of the well using a 1 ml pipet.
6. Incubate the cultures for the desired timeframe, performing a full media exchange with fresh treatment media every 2-3 days. We routinely observe a 30-40% MBP⁺ culture after 4 days in differentiation media.
7. At the end of the desired treatment period, prepare fresh 4% paraformaldehyde (PFA) or thaw a frozen stock to room temperature. CAUTION: PFA is extremely toxic and must be prepared in a fume hood.
8. Aspirate the media and slowly add 400 µl of PFA to the side of the well to fix the cells. Incubate the plate for 20 min at room temperature.
9. Aspirate the PFA and slowly add 500 µl of sterile D-PBS (with Ca²⁺ and Mg²⁺) to the side of the well to wash the cells. Repeat the wash a total of two more times. Do not aspirate the final wash.
10. Wrap the plate in parafilm and store at 4 °C for later processing or proceed directly to the next step. Plates can be stored for several weeks.

6. Immunocytochemistry and Quantification of Myelin Basic Protein

Note: When performing liquid handling procedures in the 24 well plates, it is recommended to work fast to avoid drying the cells. Here, the use of a multi-channel vacuum aspirator and multi-channel pipet are recommended.

1. Bring the plate to room temperature, aspirate the wells, and add 250 μl of D-PBS (with Ca²⁺ and Mg²⁺) containing 0.1% Triton X-100 and 5% normal goat serum (NGS). Incubate the plate at room temperature for 1 hr with gentle orbital shaking.
2. Prepare the primary incubation solution by diluting mouse monoclonal anti-MBP antibody 1:1,000 and rabbit polyclonal anti-Actin antibody 1:200 in D-PBS (with Ca²⁺ and Mg²⁺) containing 5% NGS. Alternatively, rabbit polyclonal anti-Olig2 at 1:1,000 can be used for oligodendrocyte specific normalization in mixed glial cultures. Prepare enough primary solution to accommodate 250 µl per well.
3. Incubate primary antibodies at 4 °C overnight for 16-18 hr with gentle orbital shaking.
4. Aspirate the primary incubation solution and wash the cells 3 x 10 min with 500 µl of D-PBS (with Ca²⁺ and Mg²⁺) at room temperature with gentle orbital shaking.
5. While washing the plate, prepare the secondary incubation solution by diluting anti-mouse 680 and anti-rabbit 800 antibodies 1:500 in D-PBS (with Ca²⁺ and Mg²⁺) containing 5% NGS. Minimize ambient light exposure when preparing this solution.
6. Aspirate the final wash and add 250 µl of secondary incubation solution. Protect the plate from light and incubate it at room temperature for 1 hr with gentle orbital shaking.
7. Aspirate the secondary incubation solution and wash the wells 3 x 10 min with 500 µl of D-PBS (with Ca²⁺ and Mg²⁺) at room temperature with gentle orbital shaking.
8. Scan the plate using an imaging system capable of detecting 700 nm and 800 nm fluorescence emissions. As a starting point, set the focal offset to 3 and sensitivities to 1.5 for MBP, 5.0 for Actin, and 3.5 for Olig2. Adjust the sensitivity values as needed to avoid signal overexposure.
9. Quantify the extent of oligodendrogenesis using semi-automated software-based methods.
   1. Using the software, set the plate analysis mode to "24 Well" and align circles around the outer perimeter of the scanned wells. Set the normalization channel to "800" and export the total and relative fluorescence intensities for the 700 nm (MBP) and 800 nm (Olig2/Actin) channels.
   2. Divide the intensity at 700 nm by the intensity at 800 nm to give the normalized MBP expression in relative fluorescence units. Calculate the average of replicate measurements and scale the expression to the Day 0 +PDGF controls as needed for the analysis.