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A thermal shift assay (TSA) measures how much heat a purified protein tolerates before it unfolds, and whether a buffer, additive or small molecule changes that. The protein is heated in the presence of a fluorescent dye, the fluorescence rises as the protein unfolds, and the midpoint of that rise is the melting temperature (Tm). A higher Tm in the presence of a ligand suggests stabilization.
Short answer
A thermal shift assay, also called differential scanning fluorimetry (DSF) or ThermoFluor, is a rapid and inexpensive assay to identify stabilizing solution conditions, additives and small-molecule ligands for purified recombinant proteins (Huynh and Partch, 2015). It runs in a 96-well plate on an ordinary real-time PCR instrument, so many labs already own the hardware.
Definition and scope
The names TSA, DSF and ThermoFluor describe the same experiment. The method reports a protein’s thermal stability, not its function. Its typical uses are finding buffers and additives that keep a protein folded and screening small molecules that bind and stabilize it. It works on purified protein, so it sits after expression and purification in a protein-characterization workflow.
Two terms matter when reading results. Tm is the temperature at the midpoint of the unfolding transition. dTm (the shift) is the Tm of a sample minus the Tm of a control that contains the same buffer and solvent but no ligand.
How it works in practice
The dye most often used is SYPRO Orange. It fluoresces more when it binds hydrophobic regions that become exposed as the protein unfolds, and it is largely quenched while the protein is folded (Huynh and Partch, 2015; Analytik Jena application note). Its excitation and emission properties fit the filter sets common on real-time PCR instruments.
The plate is heated gradually while fluorescence is read at each step. The result is a sigmoidal melting curve. Huynh and Partch describe fitting a Boltzmann sigmoid to the curve to obtain Tm, after removing the region past the fluorescence peak, where protein aggregation quenches the signal. The first-derivative plot is an alternative way to locate the transition. The Analytik Jena note describes the same readout: the midpoint or melt peak of the curve corresponds to the melting temperature.
The ligand-binding logic is simple: ligand binding tends to rigidify proteins and so to increase their thermal stability (Huynh and Partch, 2015). The Analytik Jena note shows buffer screening as a use: a model protein, alpha-chymotrypsinogen A, is run in Tris-buffered saline at three NaCl concentrations in a 96-well format to find better buffer conditions.
For orientation on the instrument side, a 96-well run from 25 to 95 °C took about 1 hour 45 minutes on a Peltier-controlled real-time PCR instrument in the Huynh and Partch method. For how melt curves behave on that class of instrument for DNA, see the CASRAI guide on qPCR melt curve analysis; the protein assay uses the same instrument function with a different dye and a different molecule.
Reading the curve: what each shape means
This table summarizes curve quality and fitting guidance from Huynh and Partch (2015). It is a reading aid, not a pass/fail rule.
| What you see | What the source says it reflects | What to do |
|---|---|---|
| Clear sigmoid with a single transition | A cooperative unfolding event that yields a usable Tm | Fit a Boltzmann sigmoid or use the first derivative |
| Signal falls after the peak | Aggregation quenching the fluorescence | Remove the post-peak region before fitting |
| High starting fluorescence and no clear transition | A non-ideal curve; typical causes include a protein with no hydrophobic core or an intrinsically disordered protein | The protein may not suit this assay; the authors estimate 15 to 25 percent of recombinant proteins give non-ideal curves |
| Tm lower than the control | Some ligands lower stability, particularly for proteins with a Tm above 75 °C | Treat it as a possible destabilizing effect, not automatically as noise |
Related concepts
DSF, TSA and ThermoFluor
Three names, one method. Search results and vendor pages use them interchangeably, so a protocol titled DSF and one titled thermal shift assay are normally the same experiment.
Thermal shift assay versus differential scanning calorimetry
Huynh and Partch contrast the assay with differential scanning calorimetry (DSC), which needs dedicated, expensive equipment and has low throughput. The fluorescence assay uses a standard real-time PCR instrument and a 96-well plate. That difference is the main reason the assay is used for screening conditions and ligands. Binding kinetics are a different question; for a method that measures them directly, see the CASRAI guide on bio-layer interferometry.
Where it matters for labs and equipment decisions
The platform requirement is modest: a real-time PCR instrument with Peltier temperature control and a filter set compatible with the dye (Huynh and Partch, 2015). Labs that already run qPCR can use the same instrument family; the CASRAI entry on thermal cycler specifications explains the block and temperature-control terms, and the qPCR guide covers the instrument family. The assay itself needs the dye, the purified protein and the buffers or ligands being tested.
Checklist for reading a result
- Include a control with the same buffer and solvent as the sample so that dTm has a baseline.
- Compute dTm against that solvent control, not against a different buffer.
- Huynh and Partch treat a dTm above 2.0 °C over the solvent control as a potential hit in a first-pass screen. That is that paper’s convention, not a universal cutoff.
- Check each curve for a clear sigmoid before trusting the fitted Tm.
Common misunderstandings
- Every protein gives a usable curve. Not so; the authors estimate that 15 to 25 percent of recombinant proteins give non-ideal curves.
- A positive shift always means a good binder. The assay reports stabilization of the folded protein. It is a screening readout, so hits are things to follow up, and some ligands lower stability rather than raise it.
- It needs special equipment. It runs on a real-time PCR instrument with Peltier control.
- The peak is the end of the curve. Signal after the peak can be quenched by aggregation, so that region is removed before fitting.
FAQ
What is a thermal shift assay used for?
Finding stabilizing buffer conditions, additives and small-molecule ligands for purified recombinant proteins.
Is a thermal shift assay the same as differential scanning fluorimetry?
Yes. TSA, DSF and ThermoFluor are names for the same dye-based melting experiment.
What dye does it use?
SYPRO Orange is the dye described in the sources cited below; it binds the hydrophobic regions exposed on unfolding.
What does Tm mean in this assay?
The temperature at the midpoint of the protein’s unfolding transition, read from the melt curve.
How long does a run take?
In the Huynh and Partch method, a 96-well run from 25 to 95 °C took about 1 hour 45 minutes.
Sources
- Huynh K, Partch CL. Analysis of protein stability and ligand interactions by thermal shift assay. Current Protocols in Protein Science. 2015;79:28.9.1-28.9.14. DOI 10.1002/0471140864.ps2809s79. https://pmc.ncbi.nlm.nih.gov/articles/PMC4332540 (checked 2026-10-29).
- Analytik Jena. Thermal Shift Assay using SYPRO Orange to Detect Protein Melting Temperatures (application note). https://www.analytik-jena.com/knowledge/applications/application-teasers/thermal-shift-assay-using-sypro-orange-to-detect-protein-melting-temperatures (checked 2026-10-29).
- Niesen FH, Berglund H, Vedadi M. The use of differential scanning fluorimetry to detect ligand interactions that promote protein stability. Nature Protocols. 2007;2(9):2212-2221. DOI 10.1038/nprot.2007.321. https://doi.org/10.1038/nprot.2007.321 (cited as background).
