This protocol offers a standardized method for determining immune cell identity and purity, and addresses limitation of flow cytometry such as large sample requirement, high user to user variability, and subjective data analysis. This assay includes an automated data analysis template which allows for users to assess immune cell identity and purity objectively. This, along with the multiple assay controls offers a turnkey, easily standardized method for assessing immune cell identity and purity.
Demonstrating this technique will be Jerry Guzman, a scientist in our laboratory. After genomic DNA isolation from the hematopoietic cell sample of interest, use one microliter of the genomic DNA to measure the purity of the sample on a spectrophotometer by obtaining the optical density at 260, 280 and 230 nanometers. Next dilute 400 to 1200 nanograms of genomic DNA sample to a final volume of 142 microliters with elution buffer from a genomic DNA isolation kit.
Add 75 microliters of the calibrator to 67 microliters of elution buffer. And dilute 1200 nanograms of the reference genomic DNA to a total volume of 142 microliters with elution buffer. Then incubate the tubes for five minutes at 56 degrees Celsius and 900 revolutions per minute in a thermal mixer.
For bisulfite conversion, add 270 microliters of ammonium bisulfite and 90 microliters of THFA to all of the tubes. And vortex the tubes to ensure a thorough mixing. Briefly spin down the samples to collect the liquid and place the tubes in the thermal mixer for 45 minutes at 80 degrees Celsius and 900 revolutions per minute.
Then briefly spin down the samples again and allow them to cool to room temperature for three to five minutes. For DNA purification following bisulfite conversion, vortex the DNA isolation paramagnetic beads for 30 seconds. Add ethanol and isopropanol to the wash buffers according to the volumes listed on the bottles.
Vortex again until the beads are completely resuspended and at 870 microliters of lysis binding buffer and 105 microliters of the DNA binding paramagnetic beads to each tube. Vortex the tubes to mix before briefly spinning down the samples. Add 570 microliters of 2-propanol to each tube and mix by vortexing.
Incubate the tubes for seven minutes at room temperature and 50 RPM per minute before briefly spinning down the tubes again and placing them onto a magnetic rack for five minutes. At the end of the incubation remove the supernate without disturbing the beads. Wash the sample with buffer 1 and vortex the tubes.
After a brief centrifugation place the tubes back in the rack for three minutes and remove the supernate without disturbing the beads. Wash the sample twice with wash buffer 2 and dry the tubes at 65 degrees Celsius for 15 minutes. Next, add 60 microliters evolution buffer to each tube.
After a seven minute incubation at room temperature and 1400 rotations per minute, briefly spin down the tubes and return the tubes to the magnet for two minutes. At the end of the incubation transfer 55 microliters of the bisulfite converted DNA containing to a new tube without disturbing the beads. To run quantitative PCR on the samples, assign each standard the Task Standard in the quantitative PCR software and specify the final copy numbers according to the table.
Prepare serial dilutions of standard DNA and prepare one quantitative PCR master mix cocktail for the target cell type. And one quantitative PCR master mix cocktail for GAPDH, according to the table. Load three microliters of the template DNA into the appropriate wells of a 96 well rounded bottom plate followed by seven microliters of master mix.
When all of the wells have been loaded seal the plate with quantitative PCR film and briefly spin the plate before loading it onto the quantitative PCR instrument. Then run the reaction as indicated in the table. At the run completion, export the quantitative PCR data as a txt or xlsx file.
After purifying the bisulfite converted DNA with paramagnetic beats as demonstrated, elute 25 microliters of the DNA. Transfer the bisulfite converted DNA into a new tube and add two microliters of the sample into individual PCR strip tubes. Add two microliters of water to a no template control tube and prepare the preamplification master mix for all of the samples according to the table.
Add 23 microliters of master mix to each tube and cap, vortex, and briefly spin down the tubes. Then run the preamplification protocol on a thermo cycler using a heated lid. After the run is complete, briefly spin down the tubes and transfer to microliters of the amplified DNA to new tubes.
Then dilute the samples with 78 micro liters of water to a one to 40 concentration and run a quantitative PCR of the samples as demonstrated. Here, representative data from a CD8+T cell assay obtained from analyzing both peripheral blood, mononuclear cells, and T cells from three different donors using both flow cytometry and methylation assays are shown. The trends between the two methods across all donors in both cell types were similar.
These representative data from the Treg assay also show similar results between methods. However, in all cases, this methylation assay yielded higher values under both stimulated and unstimulated conditions. The sensitivity and specificity of each assay was demonstrated by the Limit of Blank, Limit of Detection, and Limit of Quantitation values.
Because this assay offers a automated data analysis template, it improves objective analysis and increases standardization capabilities. This is ideal for cell therapy manufacturing where standardized methods are crucial for creating a consistent cell therapy product.
