This protocol is significant in that it allows for high-throughput ChIP-Seq analysis of primary samples to better understand cis-regulatory mechanisms, and its implication on disease. The main advantages of this method are the reproducibility of the data, and the drastic reduction of hands-on lab time. For chromatin shearing, add 70 microliters of fresh, room-temperature, complete lysis buffer to a glycine-fixed, snap-frozen and thawed cell pellet of interest, for a one-minute incubation at room temperature, followed by one minute of resuspension without bubbles.
Incubate the sample for another minute at room temperature, before placing the cells on ice. Transfer the resuspended pellet into a 0.65 milliliter low-binding tube on ice, before loading the tube onto the tube holder of a sonicator. Fill any gaps with balance tubes containing 70 microliters of water, and let the cells stabilize in the water bath for about one minute before starting the sonication.
After three cycles, remove the samples from the sonicator for gentle vortexing, and pulse-spin the tubes before placing them back into the holder. For histone modification, add 100 microliters of complete tC1 buffer to each tube of two chip eight-tube strips, and transfer 20 microliters of each of 16 chromatin-shared samples into individual tubes of the chip eight-tube strips. Wash the chromatin tubes with 80 microliters complete tC1 buffer, and pool the washes in the appropriate tubes of the chip eight-tube strips for a final volume of 200 microliters of solution per tube.
Add the appropriate volume of antibody to 500 microliters of TBW1 buffer, and quickly vortex, and pulse spin. Then add 70 microliters of tBW1, and 30 microliters of antibody solution to each tube of a new chip eight-tube strip. Next, thoroughly vortex a container of protein A bead solution, and add five microliters of beads per 0.5 micrograms of antibody to a new tube strip.
Pulse spin the strip of beads, and fill the last row of the chip liquid handler with labeled, empty, chip eight-tube strips. Then follow the chip 16-I pure 200 D program specifications for the placement of the rest of the strips, and load the buffers into the correct positions. After sequencing, capture the beads in an eight-tube strip magnet, and transfer 25 microliters of tagmentation buffer to each tube.
With the tubes removed from the magnet, mix gently, until the bead solutions are homogenous, before placing the re-capped tubes into a preheated ThermoMixer for three minutes. At the end of the incubation, remove the chip eight-tube strips in the last row of the chip liquid handler, and add two microliters Rnase A to each sample. Then pulse spin the re-capped tubes, and gently mix, until the bead solutions are homogenous.
For a purification of the tagmented DNA fragments, transfer 100 microliters of the D cross-linked DNA samples into individual 1.5 milliliter tubes, and wash the tubes with 100 microliters of DNA binding buffer per tube. pulse the washes in the appropriate tubes, and load the samples onto columns. Place the columns into new 1.5 milliliter collection tubes, and add nine microliters of 55 degrees Celsius Tris-EDTA buffer directly to the column matrices.
After one minute, centrifuge the columns, and transfer the nine microliters of eluate from each column into a new eight-tube strip on ice. Elute the DNA fragments again as demonstrated, but with eight microliters of Tris-EDTA buffer, and pool the eluate with the previously-harvested DNA fragment solution. To amplify the purified DNA fragment samples, mix the samples with a multi-channel pipette, and perform an amplification program with the appropriate number of cycles.
After the amplification, use a plate magnet to capture the beads, and discard the supernatant. Wash the beads three times with 200 microliters of fresh 80%ethanol per wash, without disrupting the bead pellet. After the last wash, use a 20-microliter pipette tip to remove any excess ethanol, and allow the beads to dry for 10 minutes, or until cracks appear in the bead pellets.
Add 40 microliters of pre-warmed water to each dried sample, and seal the plate before thoroughly vortexing, and briefly pulse spinning. Then quantify the DNA, using a fluorescence-quantifying assay, according to standard protocols. Sheared chromatin fragment size measurements demonstrate great reproducibility with greater than 70%of the samples being observed between 100 and 500 base pairs for 14 cycles.
As illustrated, the cycle at which the intensity of the tagmented sample is half the average maximum is optimal for cycle determination. The best sequencing data are obtained when more than 85%of the DNA fragments range between 200 to 1000 base pairs. In these representative enrichment tracks for four gene loci, individual tracks for each sample show a high mapping quality, and signal-to-noise ratio.
The first two loci harbor well-expressed genes in these cell types, while the genes in the last two loci are not expressed, and serve as background controls. For the majority of the pairwise comparisons, Pearson correlation indexes show more than a 90%correlation, suggesting a high level of consistency between the biological replicates. While cell type-specific loci exhibit a high enrichment in the appropriate cells, a housekeeping gene shows very consistent histone modification.
Correlation analysis between ChIP-Seq dataset performed from samples with less than 100, 000 cells still demonstrate a high reproducibility and correlation down to 10, 000 cells. However, there is an increased background as the cell numbers are reduced, as well as a decrease in their correlation coefficients. Proper lysing of the cells is essential for generating high-quality data, as we have found that incomplete cell lysis affects the rest of the protocol.
Using this method, many different histone modifications and transcription factors can be tested on various cell types of interest to further our understanding of cis-regulatory elements.
