Success in obtaining high-quality ChIP data is critically dependent on good primer design. In general, primers should be 20 to 30 bases long with a Tm between 55° and 60°C. Most primers require no purification or special treatment prior to PCR. Amplification products should be 75 to 350 bp; longer products should be avoided, as the amplification efficiency is substantially lower. A final primer concentration of 1 μM works well for most primers, but in some instances, improved product specificity may be obtained by lowering the final primer concentration 5- to 10-fold.
High-quality primer pairs should result in ~1.9-fold amplification/cycle (this can be determined from quantitative analysis of raw fluorescence data for each cycle, which is generally available on commercial instruments). Amplified material at the completion of the PCR should contain only one band (as assayed on high-percentage agarose or polyacrylamide gels). Specificity information may also be obtained by running dissociation curves on reactions following the conclusion of the qPCR run.
In general, individual samples should be run in triplicate. For each primer pair examined, the input DNA samples should be run alongside the immunoprecipitated samples. Amplification efficiencies among different primer pairs vary slightly on a per-cycle basis, but those slight variations in efficiency translate into substantially different amounts of amplified material in the cycle range used for analysis. Precise quantitation of relative binding cannot be accurately performed without a primer pair–specific input signal.
We have validated ChIP primer pairs for various promoter regions for validation of ChIP-qPCR. Primers were validated by testing 5-point dilutions of input DNA from MCF7 and PC3 cells, with primer efficiency and slope assessed for each pair.