This method can help answer key questions in the field of gene regulation, as it enables core promoter and enhancer discovery using low amounts of starting material. The main advantage of this technique is that it allows unbiased, genome-wide single nucleotide resolution mapping of RNA polymerase II transcriptional start sites using only 10 nanograms of total RNA. CAGE is proving to be invaluable for uncovering disease-associated transcription start sites that can be used as diagnostic markers.
SLIC-CAGE extends these discoveries to scale samples such as tissue biopsies. SLIC-CAGE is ideally suited for in-depth and high-resolution promoter analysis of frail cell types, including early embryonic developmental stages or embryonic tissue from a wide range of model organisms. Since there are numerous steps in this protocol, it is crucial to minimize sample loss in each step by taking care to retrieve as much material as possible when transferring between tubes.
Start by preparing the PCR mix for each template. Combine buffer, dNTPs, unique forward primer, template plasmid with the synthetic carrier gene, and fusion polymerase according to manuscript directions. Mix reagents by pipetting.
Add 90 microliters of the PCR mix to 10 microliters of each reverse primer and mix. Refer to the manuscript for thermocycling conditions. Perform reverse transcription or RT on the RNA of interest.
Combine one microliter of reverse transcription primer, 10 nanograms of RNA, and 4, 990 nanograms of carrier mix in a total volume of 10 microliters. Mix by pipetting up and down. Heat the mixture to 65 degrees Celsius for five minutes and then immediately place it on ice.
Prepare the RT mix according to the manuscript directions and add 28 microliters of the mix to 10 microliters of RNA. Mix the contents of the tube by pipetting until homogenous. Run RT in a thermocycler.
Once reverse transcription is complete, purify the product with DNase and RNase-free SPRI magnetic beads. Add the beads to the RT mix at a volume ratio of 1.8 to one and mix well by pipetting. Let the mixture sit at room temperature for five minutes.
Then, place the beads on a magnetic stand and let them separate for five minutes. Discard the supernatant and add 200 microliters of freshly-prepared 70%ethanol to the beads. Do not mix the beads or take them off the magnetic stand and remove the ethanol immediately after adding.
Keep the tube on the magnetic stand and remove all traces of ethanol by pushing any remaining droplets out with a P10 pipette. Immediate add 42 microliters of water, and pipette up and down 60 times to elute sample, taking care to not cause foaming. Incubate the tube at 37 degrees Celsius for five minutes without the lid and then separate the beads on the magnetic stand.
Then, transfer the supernatant to a new set of tubes, taking care to retrieve all supernatant but avoid bead carryover. After RNase I treatment, mix 45 microliters of sample with 105 microliters of prepared streptavidin beads and incubate at 37 degrees Celsius for 30 minutes. During the incubation, pipette the sample every 10 minutes to mix.
Then, separate the beads on the magnetic stand and remove the supernatant. Wash the beads with buffers A, B, and C.Starting with buffer A, resuspend the beads in 150 microliters of buffer. Separate on the magnetic stand for two to three minutes, and then remove the supernatant.
Preheat buffers B and C to 37 degrees Celsius and repeat the wash steps. To remove all non-capped RNA, it is crucial to wash the streptavidin beads and tube walls thoroughly and to completely remove the wash buffer before adding the next one. To release the cDNA, resuspend the beads in 35 microliters of 1X RNase I buffer and incubate at 95 degrees Celsius for five minutes.
After incubation, immediately place the beads on ice for two minutes. Separate the beads on a magnetic stand and transfer the supernatant to a new set of tubes. Resuspend the beads in 30 microliters of 1X RNase I buffer and then separate on a magnetic stand.
Combine the supernatant with the previously-collected supernatant for a total volume of about 65 microliters. Perform qPCR to determine the number of PCR cycles for target library amplification. Prepare the qPCR master mixes for amplifying whole libraries of DNA from the carrier.
Set thermocycling conditions according to manuscript directions and amplify the prepared sample. The SLIC-CAGE protocol makes it possible to obtain sequencing-ready libraries from nanograms of starting RNA material. The fragment length in the final library ranges between 200 and 2, 000 base pairs.
Shorter fragments are PCR artifacts and can cause sequencing problems down the line. Additional rounds of size selection can be performed to remove them. Compared to NanoCAGE, SLIC-CAGE exhibits significantly better performance at transcription start site identification, which is evident from the receiver operating characteristic curves of the two techniques performed on various quantities of S.cerevisiae total RNA.
While performing this protocol, it is important to keep in mind that sample loss may lead to low-complexity libraries. To prevent it, low-binding pipette tips and tubes must be used. Without SLIC-CAGE, promoter analysis of various cell types has so far been inaccessible.
This includes analysis of embryonic developmental stages or embryonic tissue from a wide range of model organisms.
