Enzymatic Assay for dNTPase Activity

dNTPase assays were carried out in a reaction buffer containing 10 mm Tris-HCl, pH 7.5, 50 mm NaCl, 5 mm MgCl₂, appropriate dNTPs each at 100 μm, and 56 nm (28 pmol) recombinant SAMHD1 at 25 °C. Aliquots collected at various time points were diluted into 9 volumes of ice-cold PBS to stop the reaction and spun through an Amicon Ultra 0.5-ml 10 kDa filter (Millipore) at 14,000 × g for 20 min. Deproteinized samples were analyzed by HPLC using an Eclipse XDB-C18 4.6- × 150-mm column (Agilent). The column was equilibrated in 0.1 m KH₂PO₄, 10 mm tetrabutylammonium bromide, pH 6.5 (Solvent A). Injected samples were eluted with a linear gradient of 0–7.5% acetonitrile in solvent A, over 30 min followed by 7.5% acetonitrile in solvent A for 20 min at flow rate of 1 ml/min. Alternatively, deoxyribonucleosides and dNTPs were separated over a CapCell Pak C18 4.6- × 250-mm column (Phenomenex) pre-equilibrated with 10 mm ammonium phosphate, pH 7.8, and 4.8% methanol. Then they were eluted with a linear gradient of 4.8–19.2% methanol over 22.5 min at a flow rate of 1.5 ml/min. The amounts of substrates and products were quantified by peak integration of the absorbance data recorded at 260 nm. For chemical cross-linking experiments, the reactions were assembled in the absence or presence of the indicated dGTP concentrations and preincubated on ice for 30 min, diluted with equal volume of 2% formaldehyde solution in PBS to a final concentration of 1%, followed by incubation for 15 min at room temperature. The cross-linking reactions were quenched with 0.25 m glycine for 15 min at room temperature, resolved by SDS-PAGE, and SAMHD1 was revealed by silver staining.