This method can help answer key questions in our understanding of genome regulation, such as how disease associated sequence variants in non-coding regions affect gene expression and regulation by altering protein binding, and it can help identify those proteins. The main advantage of this technique is that it does not require any sort of genetic manipulation of cells or tissues, nor antibodies, and no prior knowledge of DNA binding proteins that might interact with DNA at that locus. Demonstrating this procedure will be Danu Perumalla, a technician from my laboratory.
After designing oligonucleotides according to the text protocol, plate 2 to 5 times 10 to the 5th lymphoblastoid cells, or other cell lines, in a T25 flask with 6 milliliters of RPMI 1640 medium, supplemented with 1%Pen Strep, 2 mil or more L-glutamine, and 5%FBS. Incubate the cells at 37 degrees Celsius, and 5%carbon dioxide. Use a hemocytometer, or an automated cell counter, to measure the cell density.
Before the cells reach 1 times 10 to the 6 cells per milliliter, spin them down at 100 times G for five minutes, and transfer them to a T75 flask with 25 milliliters of the RPMI 1640 medium prepared earlier in this video. Grow the cells until the reach a cell density of 1 times 10 to the 6 cells per milliliter, then transfer the cells to a T150 flask in 38 milliliters of the same medium, and grow them for an additional two days. Next, spin down the cells at 100 times G for five minutes, and resuspend them in 10 milliliters of RPMI 1640 medium.
Then pour the suspension into an 1850 square centimeter roller bottle containing 500 milliliters of medium, and incubate the cells under the same conditions as before, except with constant rotation. Once the desired cell count is reached, spin down the culture at 100 times G for five minutes, and resuspend the cells in 36 milliliters of RPMI 1640 medium. Then transfer the cell suspension to a 50 milliliter conical tube, and remove small aliquots for cell counting and DNA extraction.
To cross-link the cells, add 1 milliliter of 37%formaldehyde to reach a final concentration of 1%Then incubate the cells at 10 to 30 RPM in room temperature for 10 minutes. Quench the cross-linking reaction by adding 2 milliliters of a 2.5 molar glycine solution for a final concentration of 125 millimolar. Then incubate the cells with rotation for five minutes.
After pelleting the cells by centrifugation, use ice-cold 1X PBS to wash the cells twice. Continue with lysis and sheering of the cells, or resuspend the pellet in 5 milliliters of cell lysis buffer, and snap-freeze the suspension drop-by-drop in liquid nitrogen. Then store the samples at minus 80 degrees Celsius.
To carry out lysis and sheering, resuspend the pellet in 20 milliliters of cell lysis buffer. The centrifuge the cells at 400 times G, in 4 degrees Celsius for five minutes. Discard the supernatant, and resuspend the cells in 6 milliliters of nuclei lysis buffer with protease inhibitors.
In preparation for sonication, divide the sample evenly into six to eight Microfuge tubes. Place the samples on ice or a cold rack, and use a sonicator with a microtip to sonicate them at 65%amplitude with five 20-second constant bursts, allowing the suspension to cool down for at least 40 seconds between pulses. Centrifuge the cells at 12, 000 times G, and 4 degrees Celsius for 10 minutes, then transfer the supernatant to a new tube, and discard the pellets.
After estimating the total recovery using fluorometry, snap freeze the samples. Following the preparation of hybridization and wash buffers according to the text protocol, use 1, 200 microliters of 1X hybridization buffer to wash 600 microliters of beads three times. Then use 1, 200 microliters of 2X hybridization buffer to resuspend the beads.
In 1.5 milliliter microcentrifuge tubes, add 600 microliters of sample, 120 microliters of washed beads suspended in 2X hybridization buffer, and 480 microliters of 2X hybridization buffer. Incubate the tubes at 31 degrees Celsius, and 30 to 60 RPM for 10 minutes. Then place the tubes on a magnet, until the beads are immobilized, and transfer the samples to new tubes before discarding the beads.
Resuspend the capture oligonucleotides to a working solution of 10 picomoles per microliter, and add 4 microliters of this solution to each tube. Incubate the tubes at 42 degrees Celsius in 30 to 60 RPM for 40 minutes. During the incubation, set aside the beads needed, wash the beads three times as before, but this time use 1X hybridization buffer to resuspend them.
Add the washed beads to the tubes, and incubate the samples at 30 to 60 RPM in room temperature for 30 minutes. Place the tubes on the magnet, and remove the supernatant once the beads have been immobilized. After washing the beads, elute the sample by adding 40 microliters of molecular grade water, and incubate the tubes at 94 degrees Celsius for five minutes.
Place the tubes on the magnet, and wait a couple of minutes until the solution clears, then transfer the supernatant into a new tube, and discard the beads. Store the sample at minus 80 degrees Celsius, or proceed to the proteomic analysis according to the text protocol. To evaluate the capture yield, use molecular grade water to dilute an aliquot of the eluate, one to ten.
After extracting the DNA from an aliquot taken earlier in the video, use the DNA sample and molecular grade water to prepare a standard curve, with five serial one to ten dilutions. Prepare a qPCR master mix adding primers and probes, and dispense the appropriate amount in each well of the reaction. To each well, add 5 microliters of either the diluted sample, the standard curve, or molecular grade water to serve as no template control.
Seal the plate and spin it at 100 times G for one minute. Then run the samples in a qPCR system using the parameters found in the text protocol. Finally, use the standard curve to assess yields, and scrambled capture to assess capture specificity.
Shown here are hybridization captures using a titration of oligonucleotides. The number of oligonucleotides is gradually increased, while the total concentration is kept constant. This experiment helps to determine how many different oligonucleotides are needed to reach optimal hybridization efficiencies.
It also reveals any obvious detrimental hybridization interactions between a particular set of oligonucleotides, and if there are any oligos with poor specificity, resulting in high background. A large proportion of fragments in the cross linked chromatin will not be amenable for hybridization capture. If the hybridization capture is successfully run, a subsequent capture experiment of the same chromatin material, using a different set of capture oligonucleotides, will result in comparable yields as when using fresh chromatin.
However, in a second capture experiment using the original oligos and leftover chromatin from the first experiment, the yield will decrease approximately 90%if most of the hybridization amenable material had been captured in the first experiment. This figure shows the hybridization capture of different chromosomal target regions, with each region serving as a negative control for the other. A scrambled sequence also serves as a true negative control.
Once mastered, this technique can be done in two days, if it is performed properly. While attempting this procedure, it's important to remember to test the capture oligonucleotides first, to assure that the desired region is amenable to hybridization capture. Following this procedure, other methods, like chromatin immunoprecipitation, can be performed to answer additional questions, like where else in the genome the identified DNA interacting proteins bind.
After watching this video, you should have a good understanding of how to perform HyCCAPP in order to enrich genomic regions of interest with the purpose of de novo identification of local protein interactions.
