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

The overall goal of this procedure is to isolate and culture rat oligodendrocyte progenitor cells and quantify the oligodendrogenesis in response to experimental factors. Efficient oligodendrogenesis is the therapeutic goal in numerous areas of research, including spinal cord injury, neonatal hypoxia, and demyelinating diseases such as transverse myelitis and multiple sclerosis. The main advantage of this technique is that it combines a protocol for the isolation of large numbers of OPCs with a fast and reliable method to determine oligodendrogenesis across multiple experimental conditions simultaneously.

To begin this procedure, sterilize all the dissection equipment using a heated bead sterilizer. Then, add 15 to 20 milliliters of PBS to a 50 milliliter conical tube. Place the conical tube on ice, and use it to hold the diced cortex tissue during dissection.

Next, add cold PBS to a 10 centimeter Petri dish which will be used for dissection under the light microscope. After sacrificing the rat pups, cut the skin behind the ears along the midline using small scissors. And peel the skin flaps back.

Next, cut the cranium along the midline from the brain stem to the eyes. And be careful not to cut too deep in order to preserve the structure of the underlying cortex. After that, 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. Carefully peel each half of the cranium back. Remove the whole brain and place it in a 10 centimeter Petri dish filled with cold PBS.

Under a dissection light microscope, remove the olfactory bulbs and cerebellum with fine surgical forceps. Flip the brain so that the ventral surface is visible. Then, using fine straight forceps, perform a blunt dissection by placing closed forceps tips between the cortex and hypothalamus to a depth of about two-thirds of the brain.

Open the forceps once they are in place, then repeat the procedure for the other hemisphere. After that, tease the cortex from the hypothalamus and midbrain regions. Remove the hypothalamus, thalamus, and midbrain by holding the midbrain at its posterior surface and peeling it toward the anterior end of the brain.

Subsequently, sever the anterior connections. Using fine bent dissection forceps, remove the hippocampus and sever it's connection to the cortex. Then, remove the remaining striatum by scraping it from the underlying cortex in an outward diagonal motion.

Clear the blood vessels and meninges from the ventral surface of the cortex. Next, flip the brain so that the dorsal surface is visible. Peel the meninges from the underlying cortex where the olfactory bulb is located.

Repeat the procedures for a total of three to four rat pups. And place all the dissected cortices into a Petri dish. Chop them with a sterilized razor blade until one millimeter by one millimeter chunks are achieved.

Then collect the tissues by rinsing the dish with PBS before placing it back on ice. Now, using a glass Pasteur pipette attached to a pipette controller, mechanically dissociate the tissue by pipetting ten to fifteen times or until the tissue pieces are reduced in size and the solution has become cloudy. Next, return the sample to the 37 degree Celsius tissue culture incubator for 15 minutes with continuous rotation.

After that, add 10 milliliters of PBS to each sample and filter the homogenate through a 40 micrometer pore filter placed over a 50 milliliter tube. Then, rinse the filter with an additional one to two milliliters of PBS. Pool all the homogenate samples into one 50 milliliter conical tube and acquire an overall cell count.

After performing the cell count, split the homogenate evenly into the 15 millileter conical tubes, and centrifuge them for 10 to 12 minutes at 300 gs. Prime each LS column by adding five milliliters of column buffer. And collect the flow through a 15 milliliter conical tube.

Once the column buffer has completely passed through the upper chamber, apply 1000 microliters of the cell suspension to each column and allow the cells to run through by gravity. Next, remove each column and place it into a 15 milliliter conical tube. Add five milliliters of column buffer and plunge the buffer though the column at a fast rate to dislodge the bound cells and release them from the column.

To plate the cells, dilute them with OPC proliferation media to 60, 000 cells per milliliter. Pipette 500 microliters of cells along the side of each black, clear bottom well of a 24 well plate. Mix the cells by shaking the plate horizontally, using a figure eight motion.

In this procedure, place a 200 microliter pipette tip over a glass Pasteur pipette to ensure that the well is not damaged during aspiration. Then, aspirate the media from the culture wells one treatment group at a time to avoid drying the cells. After that, slowly add 500 microliters of treatment master mix along the side of the well using a one milliliter pipette.

Incubate the cultures and perform a full media exchange with fresh treatment media every two to three days. At the end of the desired treatment period, aspirate the media. Slowly add 400 microliters of PFA to the side of the well to fix the cells.

And incubate the plate for 20 minutes at room temperature. After 20 minutes, aspirate PFA and slowly add 500 microliters of sterile D-PBS to wash the cells. Repeat the wash two more times and do not aspirate the final wash.

Now, bring the plate to room temperature, aspirate the wells, and add 250 microliters of D-PBS containing five percent NGS and 0.1 percent Triton x-100. Incubate the plate at room temperature for one hour with gentle orbital shaking. Prepare the primary incubation solution by diluting mouse monoclonal anti MBP antibody and rabbit polyclonal anti olig2 antibody in D-PBS containing five percent NGS.

Then, incubate the primary antibodies at four degrees Celsius overnight for 16 to 18 hours with gentle orbital shaking. While washing the plate, prepare the secondary incubation solution by diluting the anti mouse 680, and anti rabbit 800 antibodies in D-PBS containing five percent NGS. Next, aspirate the final wash and add 250 microliters of secondary incubation solution.

Protect the plate from light and incubate it at room temperature for one hour with gentle orbital shaking. After that, scan the plate using an imaging system capable of detecting 700 nanometer and 800 nanometer fluorescence emissions. As a starting point, set the focal offset to three, and sensitivities to one point five for MBP, five point zero for actin, and three point five for olig2.

Adjust the sensitivity values as needed to avoid signal overexposure. In this schematic, the DIFS assay begins with freshly isolated A2B5 positive rat OPCs plated on PLL coded culture vessels. These cultures are proliferated for three days in the presence of 20 nanograms per milliliter PDGFAA, and then treated with experimental factors in fresh PDGF-free media on day zero.

Treatment media is fully replenished on day two. And the cells are fixed with 4 percent paraformaldehyde on day four. Here, immunocytochemistry for olig2 and MBP was performed on the representative cultures on day zero and day four using Dapi as a nuclear counter stain.

These cultures exhibit constitutive staining for the pan oligodendrocyte lineage cell marker, Olig2, on days zero and four. Whereas staining for mature oligodendrocyte marker MBP was only evident by day four. After watching this video you should have a good understanding of how to purify, culture and differentiate A2B5 positive rat OPCs in order to quantify the effect of experimental factors on oligodendrogenesis.

The identification of factors that promote oligodenrogenesis is of particular relevance to the study of demyelinating diseases such as multiple sclerosis. Once mastered, the techniques described here may help to identify such factors and ultimately develop better therapeutics.