Dimethyl sulphate (DMS) footprinting

DMS footprinting was conducted using previously described methods⁵⁹, with some modifications. Briefly, 5′-FAM labelled ssDNAs (Table ^S1) were diluted to concentration of 0.2 μM in 50 mM Tris·HCl (pH 7.4) buffer containing 100 mM LiCl or 100 mM KCl plus 40% (w/v) PEG 200. After heating at 95 °C for 5 min and slowly cooling to 25 °C at 0.01 °C/s, 150 μL of water were added to the annealed DNA sample to obtain a total volume of 200 μL. Then, the samples were incubated with 4 μL of 10% (v/v) dimethyl sulphate (DMS) in ethanol at room temperature for 6 min. The reaction was stopped by the addition of 200 μL of stop buffer (0.1 M β-mercaptoethanol and 20 μg of sperm DNA). After phenol/chloroform extraction and ethanol precipitation, the oligonucleotides were dissolved in 50 μL of water and mixed with 50 μL of 20% (v/v) piperidine in water. The samples were heated at 90 °C for 30 min and dried in a Concentrator Plus (Eppendorf, Germany); the pellets were resuspended in 100 μL water and the sample was dried again. The precipitated DNAs were dissolved in 95% (v/v) deionized formamide in water containing 5 mM EDTA, heat-denatured at 95 °C for 5 min and resolved on a denaturing 16% polyacrylamide gels containing 7 M urea. Gels were scanned using an Amersham Imager 600 (GE Healthcare).