Detecting DNA/RNA Hybrids via Antibody Staining

To stain DNA/RNA hybrids, we modified the protocol described by Lisa Prendergast et al. (27 (link)) Cells were fixed in ice-cold methanol overnight at –20°C. The following day, cells were washed with PBS and permeabilized using 0.1% Triton X-100 in PBS for 5 min at RT. After washing with PBS, permeabilization was quenched with 50 mM NH₄Cl for 10 min at RT. Cells were then blocked in a solution of 3% BSA and 0.1% Triton X-100 in PBS for 30 min at RT. The S9.6 antibody (Millipore, MABE1095) was applied at a 1:200 dilution in a solution of 1% BSA and 0.1% Triton X-100 in PBS overnight at 4°C. After incubation, cells were washed three times with 0.05% PBST and then treated with the secondary antibody, Alexa Fluor 488 goat anti-mouse IgG conjugate, at a 1:250 dilution for 1 h at RT. Subsequently, nuclei were stained using 1 μg/ml of DAPI in PBS for 10 min at RT. After a final set of three washes with PBS, the cells were mounted onto glass slides using Vectashield mounting medium (Vector Labs). Images of the stained cells were captured under a Nikon AlR confocal microscope.