Grant sponsor: National Institutes of Health; Grant number: R37 NS041435.

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 studies12,13. Alternative media formulations may also be compatible with this procedure.
<ol>
	<li>Resuspend poly-L-lysine (PLL) to a concentration of 10 &#181;g/ml in sterile deionized water. Pipette 400 &#956;l of the diluted PLL into each well of a black-walled, clear bottom 24 well tissue culture grade dish.</li>
	<li>Coat tissue culture dishes for 2 hr in a 37 &#176;C tissue culture incubator or overnight in a 4 &#176;C refrigerator.</li>
	<li>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.</li>
	<li>Prepare N-Acetyl-L-cysteine (NAC) stock solution (1,000x): Dissolve 100 mg N-Acetyl-L-cysteine (NAC) in 20 ml of Dulbecco&#39;s modified Eagle&#39;s medium (DMEM). Aliquot and store at -20 &#176;C.</li>
	<li>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 &#176;C. The addition of hydrocortisone to the base media has been shown to enhance the survival of glial cells14.</li>
	<li>Prepare d-Biotin stock solution (5,000x): Dissolve 2.5 mg of d-biotin in 50 ml of PBS. Add 1-2 x 5 &#181;l drops of 0.1 N NaOH to aid in dissolution. Aliquot and store at -20 &#176;C.</li>
	<li>Prepare Insulin stock solution (100x): Dissolve 25 mg Insulin in 50 ml of tissue culture grade water. Add 250 &#181;l of 1 N HCl and mix until solution is clear. Sterilize with a 0.22 &#181;m filter and store at 4 &#176;C for up to 6 weeks.</li>
	<li>Prepare Sato stock solution (100x):
	<ol>
		<li>Prepare progesterone stock solution (25 &#181;g/&#181;l): Dissolve 2.5 mg progesterone in 100 &#181;l of ethanol.</li>
		<li>Prepare sodium selenite stock solution (400 ng/&#181;l): Dissolve 4 mg sodium selenite in 100 &#181;l of 0.1 N NaOH. Add 10 ml of DMEM and mix.</li>
		<li>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 &#181;l of progesterone stock solution, 1,000 &#181;l of sodium selenite stock solution, and mix. Aliquot and store at -20 &#176;C.</li>
	</ol>
	</li>
	<li>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 &#181;l NAC, 100 &#181;l hydrocortisone, 20 &#181;l d-Biotin, 1 ml insulin, 1 ml Sato stock, 100 &#181;l trace elements B, 2 mL B-27 supplement (50x), and 1 ml of penicillin/streptomycin (100x). Sterilize with a 0.22 &#181;m filter and store at 4 &#176;C.</li>
	<li>Prepare PDGF-AA stock solution (1,000x): Dissolve 250 &#956;g in 12.5 ml of PBS with 0.1% bovine serum albumin (BSA) to make a 20 &#956;g/ml solution. Aliquot and store at -80 &#176;C.</li>
	<li>Prepare OPC proliferation media: OPC base media supplemented with 20 ng/ml PDGF-AA.</li>
	<li>Prepare OPC differentiation media: OPC base media supplemented with 45 nM triiodothyronine.</li>
	<li>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 &#181;m filter and store at 4 &#176;C.</li>
</ol>
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.
<ol>
	<li>Sterilize all dissection equipment using a 250 &#176;C heated bead sterilizer.</li>
	<li>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.</li>
	<li>Add cold PBS to a 10 cm Petri dish. This dish will be used for dissection under the light microscope.</li>
	<li>Sacrifice rat pups by decapitation with large surgical scissors. Spray head with 70% ethanol.</li>
	<li>Extract the Whole Brain
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>Carefully peel each half of the cranium back. Remove the whole brain and place in the 10 cm Petri dish filled with cold PBS.</li>
	</ol>
	</li>
	<li>Under the dissection light microscope, remove the olfactory bulbs and cerebellum with fine surgical forceps.</li>
	<li>Flip the brain so that the ventral surface is visible under the light microscope.</li>
	<li>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.</li>
	<li>Tease the cortex from the hypothalamus and midbrain regions.</li>
	<li>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.</li>
	<li>Remove the hippocampus by pealing it outward and then severing its connection to the cortex using fine bent dissection forceps.</li>
	<li>Remove remaining striatum by scrapping it from the underlying cortex in an outward diagonal motion using fine bent dissection forceps.</li>
	<li>Clear any blood vessels and meninges from the ventral surface of the cortex.</li>
	<li>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.</li>
	<li>Complete steps 2.4-2.14 for a total of 3-4 rat pups.</li>
	<li>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.</li>
</ol>
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.
<ol>
	<li>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 &#181;l of enzyme mix. Place in the enzyme mix in a 37 &#176;C bath for 10 min. 
	&#160;&#160;&#160;&#160; 1 Sample:&#160;&#160; 1,900 &#181;l of Buffer + 50 &#181;l of papain enzyme</li>
	<li>Spin all 50 ml conical tubes containing 3-4 diced rat pup cortices at 300 x g for 3 min.</li>
	<li>Aspirate the supernatant and add 1,950 &#181;l of enzyme solution to each sample tube and break apart the pellet by inverting the tube and shaking gently.</li>
	<li>Incubate in a 37 &#176;C tissue culture incubator for 15 min with continuous tube rotation.</li>
	<li>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 &#181;l will be added to each 50 ml conical sample tube. 
	&#160;&#160;&#160;&#160; 1 Sample:&#160;&#160; 20 &#181;l of Buffer + 10 &#181;l of enzyme</li>
	<li>Once incubation is completed add 30 &#181;l of the second enzyme mix to each tube and invert to mix.</li>
	<li>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 &#176;C tissue culture incubator for 15 min with continuous rotation.</li>
	<li>Pipette each tissue homogenate sample 10-15 times using a 1 ml pipette.</li>
	<li>Pipette each tissue homogenate sample 15-20 times using a 200 &#181;l pipette</li>
	<li>Return all samples to the 37 &#176;C tissue culture incubator for 10 min with continuous rotation.</li>
	<li>Add 10 ml of PBS (with Ca2+ and Mg2+) to each sample and filter the homogenate through a 40 &#181;m pore filter placed over a 50 ml tube. Rinse the filter with an additional 1-2 ml of PBS.</li>
	<li>Pool all of the homogenate samples into one 50 ml conical tube and acquire an overall cell count.</li>
	<li>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.</li>
</ol>
4. Anti-A2B5 Bead Application, Column Purification, and Plating
<ol>
	<li>Perform calculations for column buffer and anti-A2B5 microbeads. Add 7 &#956;l Column Buffer for every 1 x 106 cells. Add 2 &#956;l Anti-A2B5 microbeads for every 1 x 106 cells.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Incubate the cell suspension for 10 min at 4 &#176;C.</li>
	<li>Add anti-A2B5 microbeads (calculated in step 4.1), invert several times and incubate at 4 &#176;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.</li>
	<li>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.</li>
	<li>Centrifuge for 10 min at 300 x g.</li>
	<li>Determine how many LS columns will be needed for the purification. One LS column is sufficient for 15-20 x 106 total cells.</li>
	<li>Resuspend the cell pellet in 1,000 &#181;l of column buffer per LS column being used.</li>
	<li>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 &#181;l 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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Since the volume of cell suspension will be much greater, evenly apply 1,000 &#181;l 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 &#181;l.</li>
	<li>Once the cell suspension has been completely applied to the second round of columns, wash 2-3 times with 3 ml of column buffer.</li>
	<li>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.</li>
	<li>Centrifuge the cell suspension at 300 x g for 12-15 min. The pellet should appear compact.</li>
	<li>Resuspend the cell pellet in 5-10 ml of pre-warmed OPC proliferation media (OPC base media supplemented with PDGF-AA 20 ng/ml).</li>
	<li>Count the cells using a hemocytometer.</li>
	<li>Plate the cells. Dilute the cells to 60,000 cells/ml with OPC proliferation media. Into each black, clear bottom 24 well, pipet 500 &#181;l 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.</li>
	<li>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.</li>
	<li>Culture the cells in OPC proliferation media at 37 &#176;C for 3 days with no media change.</li>
</ol>
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 &#176;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).
<ol>
	<li>Pre-warm OPC base media to 37 &#176;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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Aspirate the media from the culture wells one treatment group at a time so as to avoid drying the cells. Use a 200 &#181;l 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.</li>
	<li>Slowly add 500 &#181;l of treatment mastermix along the side of the well using a 1 ml pipet.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Aspirate the media and slowly add 400 &#181;l of PFA to the side of the well to fix the cells. Incubate the plate for 20 min at room temperature.</li>
	<li>Aspirate the PFA and slowly add 500 &#181;l 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.</li>
	<li>Wrap the plate in parafilm and store at 4 &#176;C for later processing or proceed directly to the next step. Plates can be stored for several weeks.</li>
</ol>
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.
<ol>
	<li>Bring the plate to room temperature, aspirate the wells, and add 250 &#956;l 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.</li>
	<li>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 &#181;l per well.</li>
	<li>Incubate primary antibodies at 4 &#176;C overnight for 16-18 hr with gentle orbital shaking.</li>
	<li>Aspirate the primary incubation solution and wash the cells 3 x 10 min with 500 &#181;l of D-PBS (with Ca2+ and Mg2+) at room temperature with gentle orbital shaking.</li>
	<li>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.</li>
	<li>Aspirate the final wash and add 250 &#181;l of secondary incubation solution. Protect the plate from light and incubate it at room temperature for 1 hr with gentle orbital shaking.</li>
	<li>Aspirate the secondary incubation solution and wash the wells 3 x 10 min with 500 &#181;l of D-PBS (with Ca2+ and Mg2+) at room temperature with gentle orbital shaking.</li>
	<li>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.</li>
	<li>Quantify the extent of oligodendrogenesis using semi-automated software-based methods.
	<ol>
		<li>Using the software, set the plate analysis mode to &#34;24 Well&#34; and align circles around the outer perimeter of the scanned wells. Set the normalization channel to &#34;800&#34; and export the total and relative fluorescence intensities for the 700 nm (MBP) and 800 nm (Olig2/Actin) channels.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>

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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Defining+glial+cells+during+CNS+development.">Defining glial cells during CNS development.</a> Nat Rev Neurosci. 2 (11), 840-843 (2001).</li><li>O'Meara, R. W., Ryan, S. D., Colognato, H., Kothary, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Derivation+of+enriched+oligodendrocyte+cultures+and+oligodendrocyte/neuron+myelinating+co-cultures+from+post-natal+murine+tissues.">Derivation of enriched oligodendrocyte cultures and oligodendrocyte/neuron myelinating co-cultures from post-natal murine tissues.</a> J Vis Exp. (54), (2011).</li><li>Dugas, J. C., Emery, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Purification+of+oligodendrocyte+precursor+cells+from+rat+cortices+by+immunopanning.">Purification of oligodendrocyte precursor cells from rat cortices by immunopanning.</a> Cold Spring Harb Protoc. 2013 (8), 745-758 (2013).</li><li>Fletcher, J. M., Lalor, S. J., Sweeney, C. M., Tubridy, N., Mills, K. H. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=T+cells+in+multiple+sclerosis+and+experimental+autoimmune+encephalomyelitis.">T cells in multiple sclerosis and experimental autoimmune encephalomyelitis.</a> Clin Exp Immunol. 162 (1), 1-11 (2010).</li></ol>

The authors have nothing to disclose.

The protocol presented here describes a fast and reliable method for isolating rat OPCs and quantifying in vitro oligodendrogenesis in response to experimental factors. Using positive selection, high yields of viable and pure OPCs are used directly for experimental purposes. This method streamlines the isolation, culturing, and differentiation steps into a 1 week time frame, and fixed cells can be conveniently stored at 4 &#176;C for several weeks without any loss in assay sensitivity.
A major advantage with scanner-based fluorescence assays is that data acquisition and analysis is greatly expedited versus traditional camera-based microscopy techniques. Although microscopy has been used successfully in high-throughput screens to identify inducers of OPC differentiation, data acquisition is inherently slower because it requires programming of image capture sequences, several hours of data collection, analysis of hundreds to thousands of image files, and user-defined algorithms to distinguish individual cell bodies for normalization or analysis18,19. Furthermore, with camera-based microscopy systems it is not feasible to acquire data from the entire surface of the culture vessel. Instead, representative images must be captured from different areas within a single well, which may lead to positional bias and false-positive hits. By comparison, this assay detects MBP expression from all cells within a given well, and multiple plates can be scanned simultaneously. While this method can be adapted for use in 96-well or 384-well formats, the 24-well plate may be preferred when fewer treatments are being tested. We have observed that cell loss in the 24-well plate due to liquid handling is minimized, which is a complication that should be considered when designing higher-throughput experiments.&#160;
A critical parameter of the assay is the density of the cells when differentiation is initiated at Day 0. Cultures with too few cells appear to differentiate more slowly and die off, while high-density cultures exhibit spontaneous differentiation. Both situations may reduce the dynamic range and reliability of the assay. To achieve an optimal balance between cell health and cell density, we culture the OPCs for 3 days without a media exchange or supplementation of fresh PDGF-AA. We hypothesized that since the concentration of PDGF-AA in the media declines by Day 3 in vitro (Day 0) as a result of cellular catabolism, the proliferative rate of the cells will slow such that proliferation is reduced through the first day of PDGF-AA withdrawal. This technique works best when the cells are evenly distributed during plating. Pipetting the cells along the side of the well and mixing them using a figure eight motion should promote an even distribution. Pipetting into the center of the well tends to produce clusters of cells around the edges of the well with fewer cells adhering to the center. This phenomenon may be due to disruption of the freshly dried PLL layer by liquid pipetting forces. Accumulation of cells around the perimeter may reduce assay sensitivity because of density-dependent differentiation. It is important to note that significant differences in the intensity of the Actin/Olig2 signal can occur between drug-treated and control, PDGF withdrawal cultures. Significantly decreased Actin/Olig2 signal may indicate drug toxicity, whereas significantly increased Actin/Olig2 signal may indicate drug-induced proliferation. We normally observe a slightly higher Actin/Olig2 signal from cells treated with pro-differentiation drugs compared to the PDGF withdrawal controls. We attribute this difference to a higher rate of spontaneous apoptosis in the PDGF withdrawal group over the 4 day differentiation procedure. When assessing oligodendrogenesis, normalization of MBP to Actin/Olig2 should correct for cell number differences. To assess toxicity and proliferation using this assay, the anti-MBP antibody can be exchanged for primary antibodies directed against the appropriate markers.
When experiments require both quantitative and qualitative assessment, it is important to limit the exposure of the fixed rat oligodendrocyte cultures to Triton X-100 when using 4% PFA to fix the cells. Other immunocytochemistry protocols for oligodendrocyte cultures, which typically include Triton X-100 during both of the antibody incubation steps to improve antibody epitope accessibility, may be used successfully depending on experimental requirements. On the other hand, although MBP is an intracellular protein it may be possible to exclude Triton X-100 entirely if alternative fixation methods are employed that adequately permeabilize the cells. Using the conditions described here, we have observed that Triton X-100 affects the observed morphology of the oligodendrocytes as well as the localization of MBP staining when used at high concentrations. We do not see a significant decline in antibody staining when Triton X-100 is omitted from the antibody incubation steps. As shown in Figure 6, including 0.4% Triton X-100 in the permeabilization and blocking step, and 0.1% Triton X-100 during antibody incubation, reduces the detection of myelin processes and results in discontinuous MBP staining. Therefore, for quantitative and qualitative assessment it may be sufficient to use a lower concentration of Triton X-100 as described here.
While the use of A2B5-selected OPCs from rats can be advantageous, we have observed comparable results in the dual-infrared fluorescence-scanning (DIFS) assay using A2B5-selected OPCs from C57BL/6 mice, although cell yield per mouse is typically much lower (data not shown). Other protocols for OPC purification may also be compatible with this assay. For example, oligodendrocyte marker O4 may be used as an antigen to purify intermediate stage oligodendrocyte precursors using magnetic beads20. As an alternative to magnetic-based methods, OPCs can be enriched from 9 day old mixed glial cultures containing astrocytes and microglia by hi-speed orbital shaking and differential adhesion methods21. Furthermore, immuno-panning procedures which incorporate both negative and positive cell selection may be used if culture purity is preferred over high cell yields22. In addition to examining how therapeutics may directly enhance the differentiation of OPCs in pure cultures, it is sometimes necessary to consider the indirect effects of drugs through other cell types in a co-culture system. In MS, the infiltration of autoreactive T effector cells into the CNS is thought to play a role in disease progression23. Effector T cells may secrete pro-inflammatory factors that may inhibit the differentiation of OPCs at sites of demyelination. It is therefore of interest to identify therapeutics that may protect the OPCs and block these pro-inflammatory factors from exacerbating injury. The DIFS assay may be ideal for modeling this activity as T cells grow in suspension and may be washed away from the oligodendrocyte culture prior to MBP quantification. For co-culture studies involving adherent cells such as neurons, astrocytes, and microglia, Olig2 normalization may be employed to collect data from the oligodendrocytes specifically. Thus, the flexibility of the DIFS assay may facilitate a variety of experimental designs.
In conclusion, this protocol provides a fast and reliable method to quantify the oligodendrogenesis of A2B5-positive rat OPCs in vitro. This isolation method consistently delivers high yields of viable OPCs that are responsive to established differentiation cues. Sensitive analysis can be performed in multi-well format vessels using a variety of experimental paradigms, which extends the utility of this system to many different applications. Furthermore, the DIFS assay can be adapted for high-throughput drug screening, or may be used to verify results of low-throughput assays such as quantitative PCR and Western blot. Importantly, this system may offer a simple yet effective means for the identification of OPC targeted therapies in demyelinating diseases such as multiple sclerosis.

Efficient nerve conduction in the mammalian central nervous system (CNS) requires myelination of axons by oligodendrocytes. During development, oligodendrocytes arise from a pool of oligodendrocyte progenitor cells (OPCs) that are thought to migrate from the ventricular zones of the developing forebrain and neural tube1. After migration OPCs differentiate into mature, myelinating oligodendrocytes that not only facilitate efficient impulse propagation, but also provide axons with trophic support2. The adult CNS maintains an abundant population of OPCs that are dispersed throughout the gray and white matter comprising approximately 5-8% of all cells3. Following a demyelinating insult, OPCs migrate to the site of injury, proliferate, and differentiate to replace lost or damaged myelin sheaths on exposed axons. However, in some disease/injury settings, this process is found to be inefficient or can fail completely4. While chronic demyelination is thought to add to the burden of disease, efficient oligodendrogenesis and remyelination may alleviate symptoms5. It has therefore been of interest to study OPCs in vitro to determine the effect of experimental factors on oligodendrogenesis.
Insight into the different phases of oligodendrocyte differentiation has been made possible by the identification of stage specific cell markers. Self-renewing early progenitor cells are defined by their expression of platelet derived growth factor receptor alpha (PDGFR&#945;), neural/glia antigen 2 (NG2) and A2B56-8. As OPCs initiate their differentiation program and withdraw from the cell cycle, they downregulate expression of these markers and begin to express proteins indicative of premyelinating oligodendrocytes including Cyclic-nucleotide 3&#39;-phosphodiesterase (CNPase) and O4. Finally, their differentiation into more mature oligodendrocytes is characterized by the expression of myelin-associated proteins, including myelin-associated glycoprotein (MAG), proteolipid protein (PLP), and myelin basic protein (MBP). MBP is an intracellular peripheral membrane protein and a major component of the myelin sheath. Mice lacking MBP develop a severe phenotype in which CNS myelination is significantly decreased leading to tremors, convulsions and early death9,10. This important role of MBP in myelination has led to its use as a marker of oligodendrocyte differentiation both in vitro and in vivo11.
Quantification of MBP can be achieved using several different methodologies. RT-PCR and Western blot analysis allow for quantification of MBP levels under different treatment regimens. Immunocytochemistry is a common qualitative approach that can also be quantitative when camera-based microscopy approaches are used. Although these systems are reliable and fundamental to the study of oligodendrocyte differentiation, they each have their own disadvantages that limit their use in drug screening. First, the amount of primary OPCs that are needed for RT-PCR and Western blot analysis reduces the number of variables that can be examined simultaneously. While the cell requirement for immunocytochemistry is much lower, substantial time must be devoted to each experiment if quantification is the goal. Numerous images must be captured and then quantified to facilitate experimental analysis. These caveats become important obstacles to consider for studies that require high throughput assessment. Here we describe a method that utilizes fundamental aspects from both the immunocytochemistry and Western blot methodologies for myelin quantification, while significantly reducing both the number of cells required and time to complete quantitative analysis.

<table border="0" cellpadding="0" cellspacing="0" width="691">
	<tbody>
		<tr height="20">
			<td colspan="4" height="20" style="height:20px;width:691px;">Dissection Materials</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">PBS without Ca2+ and Mg2+</td>
			<td>Mediatech</td>
			<td>21-040-CV</td>
			<td>1 x</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">10 cm Polystyrene Petri Dishes</td>
			<td>Fisherbrand</td>
			<td>0857512</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Light Dissection Microscope</td>
			<td>Motic</td>
			<td>SM2-168</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;"></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td colspan="4" height="21" style="height:21px;">Dissociation Materials</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Tube Rotator</td>
			<td>Miltenyi Biotec</td>
			<td>130-090-753</td>
			<td>Dissociation Tube Rotator</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Neural Tissue Dissociation Kit (P)</td>
			<td>Miltenyi Biotec</td>
			<td>130-092-628</td>
			<td>Enzymatic Dissociation Kit</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PBS with Ca2+ and Mg2+</td>
			<td>Corning</td>
			<td>20-030-CV</td>
			<td>1 x</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">40 &#956;m Cell Strainer</td>
			<td>Corning</td>
			<td>352340</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;"></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td colspan="4" height="21" style="height:21px;">A2B5 Column Purification&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Anti-A2B5 Microbeads</td>
			<td>Miltenyi Biotec</td>
			<td>130-097-864</td>
			<td>Bead Bound A2B5 Antibody</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">LS Columns</td>
			<td>Miltenyi Biotec</td>
			<td>130-042-401</td>
			<td>Magnetic Selection Columns</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">EDTA</td>
			<td>Corning</td>
			<td>46-034-Cl</td>
			<td>2mM</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PBS with Ca2+ and Mg2+</td>
			<td>Corning</td>
			<td>20-030-CV</td>
			<td>1 x</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Bovine Serum Albumin&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>A9647</td>
			<td>0.50%</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;"></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Culture Materials</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PDGF-AA</td>
			<td>PeproTech Inc</td>
			<td>100-13A</td>
			<td>20 ng/mL</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Dimethyl sulfoxide</td>
			<td>Sigma-Aldrich</td>
			<td>D2650</td>
			<td>Hybri-Max 100 mL</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Triiodothyronine&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>T6397</td>
			<td>45 nM</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Clemastine fumarate salt</td>
			<td>Sigma-Aldrich</td>
			<td>SML0445-100MG</td>
			<td>1 &#956;M</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Quetiapine hemifumarate salt</td>
			<td>Sigma-Aldrich</td>
			<td>Q3638-10MG</td>
			<td>1 &#956;M</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">N-acetyl cysteine</td>
			<td>Sigma-Aldrich</td>
			<td>A9165</td>
			<td>5 &#956;g/mL</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Progesterone</td>
			<td>Sigma-Aldrich</td>
			<td>P8783</td>
			<td>60 ng/mL</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Poly-L-lysine</td>
			<td>Sigma-Aldrich</td>
			<td>P1524</td>
			<td>10 &#956;g/mL</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Putrecine</td>
			<td>Sigma-Aldrich</td>
			<td>P7505</td>
			<td>16 &#956;g/mL</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sodium Selenite</td>
			<td>Sigma-Aldrich</td>
			<td>S5261</td>
			<td>40 ng/mL</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Hydrocortisone</td>
			<td>Sigma-Aldrich</td>
			<td>H0135</td>
			<td>50 ng/mL</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">d-Biotin</td>
			<td>Sigma-Aldrich</td>
			<td>B4639</td>
			<td>10 ng/mL</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Insulin</td>
			<td>Sigma-Aldrich</td>
			<td>I6634</td>
			<td>5 &#956;g/mL</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Apo-Transferrin</td>
			<td>Sigma-Aldrich</td>
			<td>T1147</td>
			<td>100 &#956;g/mL</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Bovine Serum Albumin</td>
			<td>Sigma-Aldrich</td>
			<td>A4161</td>
			<td>100 &#956;g/mL</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Trace Elements B</td>
			<td>Corning</td>
			<td>25-022-Cl</td>
			<td>1 x&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">B-27 Supplement</td>
			<td>Life Technologies</td>
			<td>17504044</td>
			<td>1 x&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Penicillin/Streptomycin</td>
			<td>Life Technologies</td>
			<td>15140122</td>
			<td>1 x&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">DMEM</td>
			<td>Life Technologies</td>
			<td>11995-065</td>
			<td>1 x&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">24-well Black Visiplate with lid</td>
			<td>Perkin Elmer</td>
			<td>1450-605</td>
			<td>Tissue culture grade</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;"></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td colspan="4" height="21" style="height:21px;">Immunocytochemistry and Quantification</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PFA</td>
			<td>Sigma-Aldrich</td>
			<td>100-13A</td>
			<td>4%</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Triton X-100</td>
			<td>Sigma-Aldrich</td>
			<td>78787</td>
			<td>0.10%</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Normal Goat Serum</td>
			<td>Jackson Immuno Research</td>
			<td>005-000-121</td>
			<td>5%</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">mouse monoclonal anti-MBP</td>
			<td>Biolegend</td>
			<td>SMI-99P</td>
			<td>1:1000 Dilution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">rabbit polyclonal anti-Actin</td>
			<td>Santa Cruz Biotecnology</td>
			<td>SC-7210</td>
			<td>1:200 Dilution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">rabbit polyclonal anti-Olig2</td>
			<td>Millipore</td>
			<td>AB9610</td>
			<td>1:1000 Dilution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">goat anti-mouse 680RD</td>
			<td>Licor</td>
			<td>926-68070</td>
			<td>1:500 Dilution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">goat anti-rabbit 800CW</td>
			<td>Licor</td>
			<td>926-32211</td>
			<td>1:500 Dilution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Alexa Fluor 594 goat anti-mouse</td>
			<td>Life Technologies</td>
			<td>A11032</td>
			<td>1:000 dilution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Alexa Fluor 488 goat anti-rabbit</td>
			<td>Life Technologies</td>
			<td>A11008</td>
			<td>1:000 dilution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Prolong Gold antifade with DAPI</td>
			<td>Life Technologies</td>
			<td>P36931</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">DPBS with Ca2+ and Mg2+</td>
			<td>Corning</td>
			<td>21-030-CV</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Licor Odyssey Clx Infared Imaging System</td>
			<td>Licor</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

preparation of rat oligodendrocyte progenitor cultures and quantification of oligodendrogenesis using dual-infrared fluorescence scanning

Efficient oligodendrogenesis is the therapeutic goal of a number of areas of research including spinal cord injury, neonatal hypoxia, and demyelinating diseases such as multiple sclerosis and transverse myelitis. Myelination is required to not only facilitate rapid impulse propagation within the central nervous system, but also to provide trophic support to underlying axons. Oligodendrocyte progenitor cells (OPCs) can be studied in vitro to help identify factors that may promote or inhibit oligodendrocyte differentiation. To date, many of the methods available to evaluate this process have either required large numbers of cells, thus limiting the number of conditions that can be investigated at any one time, or labor-intensive methods of quantification. Herein, we describe a protocol for the isolation of large numbers of highly pure OPCs together with a fast and reliable method to determine oligodendrogenesis from multiple conditions simultaneously. OPCs are isolated from P5-P7 neonatal rat cortices and grown in vitro for three days prior to differentiation. Four days after differentiation, oligodendrogenesis is evaluated using a dual-infrared fluorescence-scanning assay to determine expression of the myelin protein.

A2B5-positive OPCs are isolated through positive cell selection using a magnetic column separation system. Prior to the isolation procedure, the whole brain is removed from rat pups that are between P5 and P7. Once the whole brain is successfully detached from the skull, the olfactory bulbs and cerebellum are removed using fine surgical forceps (Figure 1A). In order to isolate cerebral cortical tissue the hypothalamus, thalamus and midbrain are excised by careful dissection (Figure 1B). Next, the hippocampus and striatum are removed using bent surgical forceps (Figure 1C-D). The isolation process efficiently eliminates meningeal and fibroblast cells, but enzymatic digestion is improved when meninges and vasculature are removed (Figure 1E). Finally, tissue blocks of 1 mm x 1 mm are prepared for the subsequent tissue dissociation steps (Figure 1F).
A good OPC isolation should result in a compact cell pellet after the final spin step (Figure 2A). Once plated these cultures will be relatively free of cellular debris when viewed under a microscope (Figure 2B). A suboptimal isolation may result in a large and fluffy pellet due to the presence of large amounts of cellular debris. This debris will adhere strongly to the PLL-coated vessel and may interfere with the culture (Figure 2C). If a large and fluffy pellet materializes, it may help to resuspend the cells in 10 ml of PBS and repeat the final spin for 10 min at &#60;300 x g. Once the isolation is complete, the purity of the cultures can be assessed by flow cytometry to identify the percentage of A2B5-positive cells. A successful isolation should result in a pool of cells that are &#62;95% positive for A2B5 (Figure 2D-F). Compared to cells stained with a fluorescent dye-conjugated anti-A2B5 antibody, unstained cells should appear as a negative population and can be used as a baseline reference for gating. Moreover, we have previously shown that selection of OPCs using the A2B5 antigen yields cultures that stain positive for PDGFR&#945; by immunocytochemistry11.
As demonstrated by immunocytochemistry, OPCs at the start of treatment (Day 0) are Olig2+/MBP- whereas cells are Olig2+/MBP+ by Day 4 (Figure 3). The optimal confluency of OPCs and oligodendrocytes at Day 0 and Day 4 are shown in representative bright field images (Figure 4A). The absence of MBP expression at Day 0 serves as a baseline for quantifying oligodendrogenesis. Spontaneous differentiation as a result of PDGF-AA withdrawal serves as a negative control, whereas 45 nM thyroid hormone (triiodothyronine; T3) can be used as a positive control11,15. Quetiapine, also known as Seroquel, and clemastine, also known as Tavist, are FDA-approved drugs that have been shown to accelerate in vitro oligodendrogenesis and can be used as additional positive controls16,17. Figure 4B shows a representative two-channel infrared fluorescent scan demonstrating the expected results for each of these conditions in triplicate. Actin and MBP signals were pseudocolored green and red respectively so that highly differentiated cultures appear yellow in merged images. Results show that MBP expression at Day 4 in the PDGF withdrawal condition was 4.5 &#177; 1.4-fold higher compared to Day 0, whereas the fold changes in MBP due to treatment with thyroid hormone, quetiapine, and clemastine were 33.9 &#177; 4.1, 33.4 &#177; 1.0, and 32.8 &#177; 0.5 respectively (Figure 4C). The robustness and accuracy of the assay are demonstrated by the small standard deviations among the triplicate samples and the large treatment activation window that is about 20 standard deviations above the PDGF withdrawal control.
While Actin serves as a reliable reference gene for normalization in oligodendrocyte cultures, alternative reference genes can also be used. Olig2 is a nuclear transcription factor that is constitutively expressed in cells of the oligodendrocyte lineage. Its specific and constitutive expression pattern thus may provide a more preferable normalization strategy in heterogeneous culture situations. Figure 5 shows that in OPC cultures differentiated with 10 nM thyroid hormone, the calculated fold change in MBP expression relative to PDGF withdrawal is the same when using either Olig2 or Actin for normalization. Presumably, since the OPC cultures in this experiment were very pure, both proteins can be used reliably for normalization. However, we often observe a small population of Olig2-negative cells by Day 4, but the ratio of this population to the Olig2-positive population does not seem to significantly differ between the PDGF withdrawal and T3 treated group. Thus, the fold-changes are not affected. Given a scenario where a treatment might cause proliferation of Olig2-negative cell types, the choice of normalization antibody should be carefully considered.
<img alt="Figure 1" src="/files/ftp_upload/53764/53764fig1.jpg" /> 
Figure 1: Rat brain dissection procedure. Stepwise description of the dissection procedure. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53764/53764fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53764/53764fig2.jpg" /> 
Figure 2: Flow cytometry analysis of isolated OPCs. (A) After magnetic column purification, the bound cell population is plunged out of the column and centrifuged to form a compact pellet. The amount of debris is indicated by the size of the pellet. (B) If the pellet is compact, then debris will be minimal in the culture. (C) A large and fluffy pellet typically results in a culture with heavy debris. The purity of the culture is determined by flow cytometry using a rat anti-mouse A2B5 antibody that is conjugated to APC. (D) Dead cells and debris are excluded from the analysis as shown in forward scatter and side scatter plots. (E) The A2B5 positive population can be determined by using an unstained sampled as a baseline reference for gating. (F) Successfully isolated OPCs should be &#62;95% positive for A2B5. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53764/53764fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53764/53764fig3.jpg" /> 
Figure 3: OPC culture and differentiation procedure. The dual-infrared fluorescence-scanning (DIFS) assay begins with freshly isolatedA2B5-positive rat OPCs plated on PLL-coated culture vessels (Day 3). These cultures are proliferated for 3 days in the presence of 20 ng/ml PDGF-AA, and then treated with experimental factors in fresh PDGF-free media on Day 0. Treatment media is fully replenished on Day 2, and the cells are fixed with 4% paraformaldehyde on Day 4. Immunocytochemistry for Olig2 (green) and MBP (red) was performed on representative cultures at Day 0 and Day 4 using Dapi (blue) as a nuclear counter stain. These cultures exhibit constitutive staining for the pan oligodendrocyte lineage cell marker Olig2 at Days 0 and 4, whereas staining for the mature oligodendrocyte marker MBP is only evident by Day 4. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53764/53764fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/53764/53764fig4.jpg" /> 
Figure 4: MBP quantification using the dual-infrared fluorescence-scanning (DIFS) assay. (A) The density of OPC cultures at Day 0 and Day 4 required for optimal assay performance. (B) Representative dual-infrared fluorescence scans. OPCs were differentiated in 24 well plates for 4 days in the presence of 45 nM T3, 1 &#181;M quetiapine, 1 &#181;M clemastine, or 0.1% DMSO in triplicate. A separate plate containing OPCs that were fixed on day 0 (+PDGF) was processed in parallel. Actin (pseudocolored green) and MBP (pseudocolored red) were simultaneously quantified using a Licor Odyssey scanner set to detect 700 nm and 800 nm emissions. (C) MBP signals were normalized to Actin signals, and results were scaled to the Day 0 +PDGF control. The mean relative fluorescence units &#177; SD were plotted using GraphPad Prism 6. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53764/53764fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/53764/53764fig5.jpg" /> 
Figure 5: Olig2 is a reliable reference protein for normalizing MBP expression in oligodendrocytes. OPCs were differentiated in triplicate in the presence of 10 nM T3 or 0.1% DMSO vehicle control in 24 well plates for 4 days. The dual-infrared fluorescence-scanning assay was performed using anti-Olig2 or anti-Actin primary antibodies for normalization. Olig2/Actin and MBP were simultaneously quantified using a Licor Odyssey scanner set to detect 700 nm and 800 nm emissions. MBP expression was normalized to either Olig2 or Actin expression, and results were scaled to the 0.1% DMSO control. The mean relative fluorescence units &#177; SD were plotted using GraphPad Prism 6. Results show that the calculated fold-change is the same when normalizing to either of the two reference proteins.
<img alt="Figure 6" src="/files/ftp_upload/53764/53764fig6.jpg" /> 
Figure 6: Effect of Triton X-100 on MBP staining. OPCs were differentiated in 24 well plates for 4 days in the presence of 45 nM T3 or 0.1% DMSO. Cultures were then fixed with 4% PFA and stained for Olig2 and MBP using two different immunocytochemistry protocols. For the high Triton X-100 sample, the cells were permeabilized for 1 hr with 0.4% Triton X -100 and all antibody incubation steps were performed in the presence of 0.1% Triton X-100. For the low Triton X-100 sample, the cells were permeabilized for 1 hr with 0.1% Triton X-100, while antibody incubations were performed without Triton X-100. Olig2 and MBP were visualized with fluorescent dye-conjugated secondaries and images were acquired using a camera-based fluorescence microscope. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53764/53764fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Preparation of Rat Oligodendrocyte Progenitor Cultures and Quantification of Oligodendrogenesis Using Dual-infrared Fluorescence Scanning at JoVE.com

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

Oligodendrocytes are the myelinating cells of the central nervous system. This protocol describes a method for the isolation and culture of their precursors, oligodendrocyte progenitor cells, from rat cortices, as well as a fast and reliable quantitative method to evaluate oligodendrogenesis in vitro in response to experimental factors.

Efficient oligodendrogenesis is the therapeutic goal of a number of areas of research including spinal cord injury, neonatal hypoxia, and demyelinating ...

preparation-rat-oligodendrocyte-progenitor-cultures-quantification

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

11.8K Views.  Johns Hopkins University, School of Medicine. Oligodendrocytes are the myelinating cells of the central nervous system. This protocol describes a method for the isolation and culture of their precursors, oligodendrocyte progenitor cells, from rat cortices, as well as a fast and reliable quantitative method to evaluate oligodendrogenesis in vitro in response to experimental factors.

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

Organotypic Slice Cultures for Studies of Postnatal Neurogenesis

Here we describe a technique for studying hippocampal postnatal neurogenesis in the rodent brain using the organotypic slice culture technique. This method maintains the characteristic topographical morphology of the hippocampus while allowing direct application of pharmacological agents to the developing hippocampal dentate gyrus. Additionally, slice cultures can be maintained for up to 4 weeks and thus, allow one to study the maturation process of newborn granule neurons. Slice cultures allow for efficient pharmacological manipulation of hippocampal slices while excluding complex variables such as uncertainties related to the deep anatomic location of the hippocampus as well as the blood brain barrier. For these reasons, we sought to optimize organotypic slice cultures specifically for postnatal neurogenesis research.

Here we describe a technique for studying hippocampal postnatal neurogenesis using the organotypic slice culture technique. This method allows for in vitro manipulation of adult neurogenesis and allows for the direct application of pharmacological agents to the cultured hippocampus.

Here we describe a technique for studying hippocampal postnatal neurogenesis in the rodent brain using the organotypic slice culture technique. This ...

Delivery of In Vivo Acute Intermittent Hypoxia in Neonatal Rodents to Prime Subventricular Zone-derived Neural Progenitor Cell Cultures

Extended culture of neural stem/progenitor cells facilitates in vitro analyses to understand their biology while enabling expansion of cell populations to adequate numbers prior to transplantation. Identifying approaches to refine this process, to augment the production of all CNS cell types (i.e., neurons), and to possibly contribute to therapeutic cell therapy protocols is a high research priority. This report describes an easily applied in vivo &#8220;pre-conditioning&#8221; stimulus which can be delivered to awake, non-anesthetized animals. Thus, it is a non-invasive and non-stressful procedure. Specifically described are the procedures for exposing mouse or rat pups (aged postnatal day 1-8) to a brief (40-80 min) period of intermittent hypoxia (AIH). The procedures included in this video protocol include calibration of the whole-body plethysmography chamber in which pups are placed during AIH and the technical details of AIH exposure. The efficacy of this approach to elicit tissue-level changes in the awake animal is demonstrated through the enhancement of subsequent in vitro expansion and neuronal differentiation in cells harvested from the subventricular zone (SVZ). These results support the notion that tissue level changes across multiple systems could be observed following AIH, and support the continued optimization and establishment of AIH as a priming or conditioning modality for therapeutic cell populations.

Delivery of In Vivo Acute Intermittent Hypoxia in Neonatal Rodents to Prime Subventricular Zone-derived Neural Progenitor Cell Cultures

This article describes the methodology for administering short periods of intermittent hypoxia to postnatal day 1-8 mouse or rat pups. This approach effectively elicits a robust tissue level &#8220;priming effect&#8221; on cultured neural progenitor cells that are harvested within 30 min of hypoxia exposure.

Extended culture of neural stem/progenitor cells facilitates in vitro analyses to understand their biology while enabling expansion of cell populations to ...

Isolation of Neural Stem/Progenitor Cells from the Periventricular Region of the Adult Rat and Human Spinal Cord

Adult rat and human spinal cord neural stem/progenitor cells (NSPCs) cultured in growth factor-enriched medium allows for the proliferation of multipotent, self-renewing, and expandable neural stem cells. In serum conditions, these multipotent NSPCs will differentiate, generating neurons, astrocytes, and oligodendrocytes. The harvested tissue is enzymatically dissociated in a papain-EDTA solution and then mechanically dissociated and separated through a discontinuous density gradient to yield a single cell suspension which is plated in neurobasal medium supplemented with epidermal growth factor (EGF), basic fibroblast growth factor (bFGF), and heparin. Adult rat spinal cord NSPCs are cultured as free-floating neurospheres and adult human spinal cord NSPCs are grown as adherent cultures. Under these conditions, adult spinal cord NSPCs proliferate, express markers of precursor cells, and can be continuously expanded upon passage. These cells can be studied in vitro in response to various stimuli, and exogenous factors may be used to promote lineage restriction to examine neural stem cell differentiation. Multipotent NSPCs or their progeny can also be transplanted into various animal models to assess regenerative repair.

The adult mammalian spinal cord contains neural stem/progenitor cells (NSPCs) that can be isolated and expanded in culture. This protocol describes the harvesting, isolation, culture, and passaging of NSPCs generated from the periventricular region of the adult spinal cord from the rat and from human organ transplant donors.

Adult rat and human spinal cord neural stem/progenitor cells (NSPCs) cultured in growth factor-enriched medium allows for the proliferation of ...

Generation of Genetically Modified Organotypic Skin Cultures Using Devitalized Human Dermis

Organotypic cultures allow the reconstitution of a 3D environment critical for cell-cell contact and cell-matrix interactions which mimics the function and physiology of their in vivo tissue counterparts. This is exemplified by organotypic skin cultures which faithfully recapitulates the epidermal differentiation and stratification program. Primary human epidermal keratinocytes are genetically manipulable through retroviruses where genes can be easily overexpressed or knocked down. These genetically modified keratinocytes can then be used to regenerate human epidermis in organotypic skin cultures providing a powerful model to study genetic pathways impacting epidermal growth, differentiation, and disease progression. The protocols presented here describe methods to prepare devitalized human dermis as well as to genetically manipulate primary human keratinocytes in order to generate organotypic skin cultures. Regenerated human skin can be used in downstream applications such as gene expression profiling, immunostaining, and chromatin immunoprecipitations followed by high throughput sequencing. Thus, generation of these genetically modified organotypic skin cultures will allow the determination of genes that are critical for maintaining skin homeostasis.

The goal of this paper is to provide a comprehensive and detailed protocol on how to generate genetically modified human organotypic skin from epidermal keratinocytes and devitalized human dermis.

Organotypic cultures allow the reconstitution of a 3D environment critical for cell-cell contact and cell-matrix interactions which mimics the function ...

Large-scale Scanning Transmission Electron Microscopy (Nanotomy) of Healthy and Injured Zebrafish Brain

Large-scale 2D electron microscopy (EM), or nanotomy, is the tissue-wide application of nanoscale resolution electron microscopy. Others and we previously applied large scale EM to human skin pancreatic islets, tissue culture and whole zebrafish larvae1-7. Here we describe a universally applicable method for tissue-scale scanning EM for unbiased detection of sub-cellular and molecular features. Nanotomy was applied to investigate the healthy and a neurodegenerative zebrafish brain. Our method is based on standardized EM sample preparation protocols: Fixation with glutaraldehyde and osmium, followed by epoxy-resin embedding, ultrathin sectioning and mounting of ultrathin-sections on one-hole grids, followed by post staining with uranyl and lead. Large-scale 2D EM mosaic images are acquired using a scanning EM connected to an external large area scan generator using scanning transmission EM (STEM). Large scale EM images are typically ~ 5 - 50 G pixels in size, and best viewed using zoomable HTML files, which can be opened in any web browser, similar to online geographical HTML maps. This method can be applied to (human) tissue, cross sections of whole animals as well as tissue culture1-5. Here, zebrafish brains were analyzed in a non-invasive neuronal ablation model. We visualize within a single dataset tissue, cellular and subcellular changes which can be quantified in various cell types including neurons and microglia, the brain&#39;s macrophages. In addition, nanotomy facilitates the correlation of EM with light microscopy (CLEM)8 on the same tissue, as large surface areas previously imaged using fluorescent microscopy, can subsequently be subjected to large area EM, resulting in the nano-anatomy (nanotomy) of tissues. In all, nanotomy allows unbiased detection of features at EM level in a tissue-wide quantifiable manner.

Large-scale Scanning Transmission Electron Microscopy Nanotomy of Healthy and Injured Zebrafish Brain

Large-scale 2D electron microscopy (EM), or nanotomy, is the tissue-wide application of nanoscale resolution EM. Here we describe a universal method for nanotomy applied to investigate the zebrafish larval brain in health and upon non-invasive brain injury.

Large-scale 2D electron microscopy (EM), or nanotomy, is the tissue-wide application of nanoscale resolution electron microscopy. Others and we previously ...

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM compared to confocal systems are its low phototoxicity, gentle mounting strategies, fast acquisition with high signal to noise ratio and the possibility of imaging samples from various angles (views) for long periods of time. Imaging from multiple views unleashes the full potential of LSFM, but at the same time it can create terabyte-sized datasets. Processing such datasets is the biggest challenge of using LSFM. In this protocol we outline some solutions to this problem. Until recently, LSFM was mostly performed in laboratories that had the expertise to build and operate their own light sheet microscopes. However, in the last three years several commercial implementations of LSFM became available, which are multipurpose and easy to use for any developmental biologist. This article is primarily directed to those researchers, who are not LSFM technology developers, but want to employ LSFM as a tool to answer specific developmental biology questions. 
Here, we use imaging of zebrafish eye development as an example to introduce the reader to LSFM technology and we demonstrate applications of LSFM across multiple spatial and temporal scales. This article describes a complete experimental protocol starting with the mounting of zebrafish embryos for LSFM. We then outline the options for imaging using the commercially available light sheet microscope. Importantly, we also explain a pipeline for subsequent registration and fusion of multiview datasets using an open source solution implemented as a Fiji plugin. While this protocol focuses on imaging the developing zebrafish eye and processing data from a particular imaging setup, most of the insights and troubleshooting suggestions presented here are of general use and the protocol can be adapted to a variety of light sheet microscopy experiments.

Light sheet fluorescence microscopy is an excellent tool for imaging embryonic development. It allows recording of long time-lapse movies of live embryos in near physiological conditions. We demonstrate its application for imaging zebrafish eye development across wide spatio-temporal scales and present a pipeline for fusion and deconvolution of multiview datasets.

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM ...

Using Primary Neurosphere Cultures to Study Primary Cilia

The primary cilium is fundamentally important for the proliferation of neural stem/progenitor cells and for neuronal differentiation during embryonic, postnatal, and adult life. In addition, most differentiated neurons possess primary cilia that house signaling receptors, such as G-protein-coupled receptors, and signaling molecules, such as adenylyl cyclases. The primary cilium determines the activity of multiple developmental pathways, including the sonic hedgehog pathway during embryonic neuronal development, and also functions in promoting compartmentalized subcellular signaling during adult neuronal function. Unsurprisingly, defects in primary cilium biogenesis and function have been linked to developmental anomalies of the brain, central obesity, and learning and memory deficits. Thus, it is imperative to study primary cilium biogenesis and ciliary trafficking in the context of neural stem/progenitor cells and differentiated neurons. However, culturing methods for primary neurons require considerable expertise and are not amenable to freeze-thaw cycles. In this protocol, we discuss culturing methods for mixed populations of neural stem/progenitor cells using primary neurospheres. The neurosphere-based culturing methods provide the combined benefits of studying primary neural stem/progenitor cells: amenability to multiple passages and freeze-thaw cycles, differentiation potential into neurons/glia, and transfectability. Importantly, we determined that neurosphere-derived neural stem/progenitor cells and differentiated neurons are ciliated in culture and localize signaling molecules relevant to ciliary function in these compartments. Utilizing these cultures, we further describe methods to study ciliogenesis and ciliary trafficking in neural stem/progenitor cells and differentiated neurons. These neurosphere-based methods allow us to study cilia-regulated cellular pathways, including G-protein-coupled receptor and sonic hedgehog signaling, in the context of neural stem/progenitor cells and differentiated neurons.

The primary cilium is fundamentally important in neural progenitor cell proliferation, neuronal differentiation, and adult neuronal function. Here, we describe a method to study ciliogenesis and the trafficking of signaling proteins to cilia in neural stem/progenitor cells and differentiated neurons using primary neurosphere cultures.

The primary cilium is fundamentally important for the proliferation of neural stem/progenitor cells and for neuronal differentiation during embryonic, ...

Single-Cell Quantification of Protein Degradation Rates by Time-Lapse Fluorescence Microscopy in Adherent Cell Culture

Proteins are in a dynamic state of synthesis and degradation and their half-lives can be adjusted under various circumstances. However, most commonly used approaches to determine protein half-life are either limited to population averages from lysed cells or require the use of protein synthesis inhibitors. This protocol describes a method to measure protein half-lives in single living adherent cells, using SNAP-tag fusion proteins in combination with fluorescence time-lapse microscopy. Any protein of interest fused to a SNAP-tag can be covalently bound by a fluorescent, cell permeable dye that is coupled to a benzylguanine derivative, and the decay of the labeled protein population can be monitored after washout of the residual dye. Subsequent cell tracking and quantification of the integrated fluorescence intensity over time results in an exponential decay curve for each tracked cell, allowing for determining protein degradation rates in single cells by curve fitting. This method provides an estimate for the heterogeneity of half-lives in a population of cultured cells, which cannot easily be assessed by other methods. The approach presented here is applicable to any type of cultured adherent cells expressing a protein of interest fused to a SNAP-tag. Here we use mouse embryonic stem (ES) cells grown on E-cadherin-coated cell culture plates to illustrate how single cell degradation rates of proteins with a broad range of half-lives can be determined.

This protocol describes a method to determine protein half-lives in single living adherent cells, using pulse labeling and fluorescence time-lapse imaging of SNAP-tag fusion proteins.

Proteins are in a dynamic state of synthesis and degradation and their half-lives can be adjusted under various circumstances. However, most commonly used ...

Three-dimensional Organotypic Cultures of Vestibular and Auditory Sensory Organs

The sensory organs of the inner ear are challenging to study in mammals due to their inaccessibility to experimental manipulation and optical observation. Moreover, although existing culture techniques allow biochemical perturbations, these methods do not provide a means to study the effects of mechanical force and tissue stiffness during development of the inner ear sensory organs. Here we describe a method for three-dimensional organotypic culture of the intact murine utricle and cochlea that overcomes these limitations. The technique for adjustment of a three-dimensional matrix stiffness described here permits manipulation of the elastic force opposing tissue growth. This method can therefore be used to study the role of mechanical forces during inner ear development. Additionally, the cultures permit virus-mediated gene delivery, which can be used for gain- and loss-of-function experiments. This culture method preserves innate hair cells and supporting cells and serves as a potentially superior alternative to the traditional two-dimensional culture of vestibular and auditory sensory organs.

Three-dimensional organotypic cultures of the murine utricle and cochlea in optically clear collagen I gels preserve innate tissue morphology, allow for mechanical stimulation through adjustment of matrix stiffness, and permit virus-mediated gene delivery.

The sensory organs of the inner ear are challenging to study in mammals due to their inaccessibility to experimental manipulation and optical observation. ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...