A modified 3C assay protocol was conducted as described previously (67, 68). Briefly, 1 × 107 cells were washed in cold PBS buffer. Cells were cross-linked with a final concentration of 2% formaldehyde for 10 min at room temperature, and cross-linking was stopped with glycine (final concentration, 125 mM). Nuclei were collected from the cross-linked cells and then digested with Eco RI or Hind III at 37°C overnight. The restriction enzymes were heat inactivated, and the reaction mixture was diluted in the ligation buffer to favor intramolecular ligation of cross-linked chromatin segments, and the DNA was subjected to ligation with T4 DNA ligase at 16°C for 3 days. The ligation reaction mixtures were incubated overnight at 65°C with the reverse buffer containing proteinase K (final concentrations at 200 μg/ml) to reverse the cross-links and digest the proteins. After the cross-links were reversed, DNA was purified by phenol chloroform extraction and ethanol precipitated. 3C yields a genome-wide ligation product library in which each ligation product corresponds to a specific interaction between the two corresponding loci. The frequency with which a specific 3C ligation product occurs in the library is a measure of the frequency with which the loci are sufficiently close in space to be cross-linked. Real-time PCR amplification with primers across the restriction sites in the specific 3C ligation products was carried out to quantify the frequency with which the loci interact. Loading controls represent total DNA concentrations between 3C library samples (using PCR primers that do not amplify across the restriction sites used during the 3C assays). The PCR products were also analyzed by agarose gel electrophoresis, purified, and verified by DNA sequencing.

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