The overall goal of this procedure is to study global methylation profiling and identify differentially methylated CpG-rich regions in chronic lymphocytic leukemia patients using next-generation sequencing. This method can help answer key questions in the epigenetics field such as identification of the potential CLL-specific signature genes disease pathogenesis, and prognosis. The main advantage of this technique is that it provides complete genome-wide coverage of CpG methylation in an unbiased and PCR-independent manner.
To begin, use a commercially available DNA extraction kit to isolate genomic DNA from the patient and normal peripheral blood mononuclear cell samples according to the manufacturer's protocol. After quantifying the genomic DNA as described in the text protocol, dilute 5 micrograms of genomic DNA from each sample, to a total of 200 microliters using TE buffer, giving a final concentration of 25 nanograms per microliter. Next, perform sonication in specially-designed tubes, using a sonicator, for a total of 30 cycles of 30 seconds on and 30 seconds off.
After every five cycles, briefly spin the tubes to collect the samples to the bottom. Check the sonication range for all samples using DNA electrophoresis as described in the text protocol. To prepare the magnetic streptavidin beads, first re-suspend them from the stock tube provided by the kit.
Gently pipette the beads up and down to obtain a homogenous suspension. For each five micrograms of fragmented DNA sample, place 50 microliters of the beads into separate, clean and labeled 1.5-milliliter tubes. Add 50 microliters of 1X bead wash buffer to reach a final volume of 100 microliters.
Place the tubes on a magnetic stand for one minute to allow all the magnetic beads to concentrate on the inner wall of the tube facing the magnet. Remove the liquid without touching the beads, using a 200 microliter pipette. Then, remove the tubes from the magnetic stand, add 250 microliters of 1X bead wash buffer, and mix the beads gently with a pipette.
After repeating these steps at least four times for all samples, finally resuspend the samples in 250 microliters of 1X bead wash buffer. Keep the samples on ice. To bind the MBD-biotin protein to the washed beads, first add 35 microliters of protein to separate tubes and bring the total volume up to 250 microliters using 1X bead wash buffer.
Add 250 microliters of diluted MBD protein to the 250 microliters of washed beads and leave them on end-to-end rotation at room temperature. After mixing the beads in protein for one hour, wash the protein-bound magnetic streptavidin beads by placing the tubes on the magnetic stand for one minute. Remove the liquid without touching the beads using a pipette.
Then, add 250 microliters of 1X bead wash buffer and place the tubes on a rotation mixer for five minutes at room temperature. Repeat the washing step two more times before re-suspending the washed MBD-biotin beads in 200 microliters of 1X bead wash buffer, making the beads ready for methylated DNA capture. In a clean 1.5-milliliter DNase-free tube, add 100 microliters of milliliter bead wash buffer and 180 microliters of the fragmented genomic DNA.
Bring the final volume to 500 microliters using DNase-free water. Add 380 microliters of DNase-free water to the remaining 20 liters of the fragmented genomic DNA. Freeze the samples to later process them further and use them as input DNA controls.
Place the tubes containing washed MBD-biotin beads on a magnetic stand for one minute, and remove the liquid without disturbing the beads. Then, add 500 microliters of fragmented genomic DNA diluted in bead wash buffer. Seal all the tubes tightly with paraffin film and leave them overnight at 4 degrees Celsius on an end-to-end rotation stand at 8 to 10 rotations per minute.
After the DNA and MBD bead binding reaction, place the tubes on the magnetic rack for one minute to concentrate all the beads on the inner wall of the tube. Remove the supernatant liquid with a pipette without touching the beads, and save this unbound DNA sample fraction on ice. Then add 200 microliters of 1X bead wash buffer to the beads and place the tubes on the rotating stand for three minutes at room temperature.
After removing the liquid using the magnetic stand, repeat the wash another two times to remove the residual unbound DNA. After the final wash, add 200 microliters of high-salt elution buffer provided in the kit to elute the DNA. Then, place the tubes on the rotating stand for 15 minutes at room temperature.
Following incubation, place them on the magnetic stand for one minute and use a pipette to carefully transfer the supernatant to a new, clean 1.5-milliliter tube. Next, add 200 microliters of high-salt elution buffer to the beads, and repeat the elution by rotating the tubes at room temperature for an additional 15 minutes. Add the second elute to the same tube containing the 200 microliters of the first elute.
To precipitate the DNA, add one microliter of glycogen, 40 microliters of three molar sodium acetate, pH 5.2, and 800 microliters of ice-cold absolute ethanol to 400 microliters of eluted DNA. Also add the reagents to the previously frozen 400 microliters of input DNA control samples. Mix the tubes well by vortexing before incubating them at minus 80 degrees Celsius overnight.
The next day, centrifuge the tubes at 12, 000 times g and 4 degrees Celsius for 30 minutes. Discard the supernatant carefully without disturbing the pellet. Then, add 500 microliters of 70%ethanol and vortex the tubes.
After centrifuging the tubes again using the same conditions but for 15 minutes, remove the supernatant by carefully using a pipette. Then, centrifuge the tubes at maximum speed for one minute at room temperature, and remove the residual ethanol completely using a pipette tip. Air-dry the pellet for five minutes at room temperature.
Finally, add 10 microliters of DNase-free water to the DNA pellet before proceeding to the methylated DNA quantification, MBD sequencing, and analysis as described in the text protocol. The analysis identified several differentially hyper-and hypomethylated regions enriched in CLL samples compared to normal controls. These differentially methylated regions were mapped to different classes of protein coding and noncoding genes.
Finally, the data analysis showed significant overlap between the CLL-associated differentially methylated regions obtained from two different normal control comparisons. Here, all the CpG sites located in target regions of two differentially methylated genes were validated in a large number of CLL samples using pyrosequencing. The optimum range of sonication required for this method is illustrated here.
This range of fragmented DNA is ideal and more suitable for next-generation sequencing purposes. Once mastered, this technique can be done in two days if it is done properly. While attempting this procedure, the crucial factors that affect the final DNA recovery are the quality and the amount of the input DNA used and the range of the fragmented DNA after sonication.
We decided to use this method because it is the first CLL global methylation study that is based on the immunoprecipitation for enriching methylated DNA covering all the methylated regions across the genome. The implications of this technique extend towards prognosis or therapy because the identified differentially methylated signature genes could serve as global normal biomarkers and epigenetic targets for therapy. Following this procedure, the enriched methylated DNA can be used for other downstream applications like hybridizing or doing microarrays to easily compare the metalomes between two samples or disease entities using a fixed set of CpG sites present on the arrays.
Finally, this is a cost-effective technique to analyze large number of patient samples for methylated sequences across the genome, including annotated sequences spanning protein-coding genes as well as non-annotated sequences spanning repetitive elements and long non-coding RNAs.
