Thermal shift assay

Diagram

A thermal shift assay, also called differential scanning fluorimetry (DSF) is a thermal-denaturation assay that measures the thermal stability of a target protein and a subsequent increase in protein melting temperature upon binding of a ligand to the protein.^[1]

The binding of low molecular weight ligands can increase the thermal stability of a protein, as described by Koshland (1958)^[2] and Linderstrom-Lang and Schellman (1959).^[3] The thermal stability change is measured by performing a thermal denaturation curve in the presence of a fluorescent dye, such as Sypro Orange.^[4] Sypro orange binds nonspecifically to hydrophobic surfaces, and water strongly quenches the fluorescence of Sypro Orange.^[5] When the protein unfolds, the exposed hydrophobic surfaces bind the dye, resulting in an increase in fluorescence. The stability curve and its midpoint value (melting temperature, T_m) are obtained by gradually increasing the temperature to unfold the protein and measuring the fluorescence at each point. Curves are measured for protein only and protein + ligand, and ΔT_m is calculated.

A fluorescence-based thermal shift assay can be performed on instruments that combine sample temperature control and dye fluorescence detection, such as readily available real-time polymerase chain reaction (RT-PCR) machines.^[5] This assay allows high-throughput screening of ligands to the target protein and it is widely used in the early stages of drug discovery in the pharmaceutical industry, structural genomics efforts, and high-throughput protein engineering.^[6]

Thermal shift assays can also be used to identify optimal protein buffer conditions. ^[7]

A related method to screen for protein buffer conditions is nanoDSF. It measures protein stability in thermal and chemical denaturation experiments by detecting changes in intrinsic tryptophan fluorescence upon unfolding.^{[8] [9]}

A typical assay

1. Materials: A fluorometer equipped with temperature control or similar instrumentation (RT-PCR machines); suitable fluorescent dye; a suitable assay plate, such as 96 well RT-PCR plate.^[5]
2. Compound solutions: Test ligands are prepared at a 50- to 100-fold concentrated solution, generally in the 10-100 mM range. For titration, a typical experimental protocol employs a set of 12 well, comprising 11 different concentrations of a test compound with a single negative control well.^[5]
3. Protein solution: Typically, target protein is diluted from a concentrated stock to a working concentration of ~0.5-5 μM protein with dye into a suitable assay buffer. The exact concentrations of protein and dye are defined by experimental assay development studies.^[5]
4. Centrifugation and oil dispense: A brief centrifugation (~1000 × g-force, 1 min) of the assay plate to mix compounds into the protein solution, 1-2 μl silicone oil to prevent the evaporation during heating is overlaid onto the solution, followed by an additional centrifugation step (~1000 × g-force, 1 min).^[5]
5. Instrumental set up: A typical temperature ramp rates range from 0.1-10 °C/min but generally in the range of 1 °C/min. The fluorescence in each well is measured at regular intervals, 0.2-1 °C/image, over a temperature range spanning the typical protein unfolding temperatures of 25-95 °C.^[5]

References