Chromatin Fractionation for Protein Analysis

Chromatin fractionation was performed as described^{63 (link)}. Briefly, cells were washed in PBS and resuspended in 1 ml buffer A (10 mM HEPES pH 7.9, 10 mM KCl, 1.5 mM MgCl₂, 0.34 M sucrose, 10% glycerol, 1 mM DTT and 1X protease inhibitor cocktail). Triton X-100 was added to 0.1% and the cells are incubated on ice for 10 min. Nuclei were collected by centrifugation at 4000 rpm at 4 °C. The supernatant was taken as the cytosolic fraction. Nuclei were washed once with buffer A and then lysed for 30 min in ‘No Salt’ buffer (3 mM EDTA, 0.2 mM EGTA, 1 mM DTT, and 1X protease inhibitor cocktail) on ice. Chromatin was pelleted by centrifugation at 4000 rpm at 4 °C and supernatant was enriched in soluble nuclear proteins. For western blotting, equal amounts of isolated chromatin, estimated by amido black (Sigma) staining, were run on an 8%, 15% or 4-15% SDS-PAGE gel, then transferred to PVDF membranes (Millipore). After blocking with Intercept® (PBS) Blocking Buffer (LI-COR) for 1 h at room temperature, the membrane was incubated with primary antibodies at 4 °C overnight. The membrane was then washed three times with PBST for 10 min and then incubated for 1 h at room temperature with appropriate secondary antibodies conjugated with Dylight (Invitrogen). After extensive washing, fluorescent detection was performed using the Odyssey® Fc imaging system (Li-Cor Biosciences). Alternatively, immunoblotting was performed as described^{63 (link)}. For the quantification of the bands obtained by Western blot experiments, the relative density of the band obtained from the Odyssey® Fc imaging system after blotting with antibodies of interest was normalized to the relative density of the bands obtained by blotting with antibodies against housekeeping proteins (H3 or H4). This ratio was used to compare expressions between conditions.