A thermal shift assay (TSA) measures the stability of a protein by the change in its thermal denaturation temperature. This is done under varying conditions, such as variations in drug concentration, buffer formulation (pH or ionic strength), redox potential, or sequence mutation. The most common method for measuring protein thermal shifts is differential scanning fluorimetry (DSF). DSF methodology includes techniques such as nanoDSF, which relies on the intrinsic fluorescence from native tryptophan or tyrosine residues, and Thermofluor, which utilizes extrinsic fluorogenic dyes.
The binding of low molecular weight ligands can increase the thermal stability of a protein, as described by Daniel Koshland (1958) and Kaj Ulrik Linderstrøm-Lang and Schellman (1959). Almost half of enzymes require a metal ion co-factor. Thermostable proteins are often more useful than their non-thermostable counterparts, e.g., DNA polymerase in the polymerase chain reaction, so protein engineering often includes adding mutations to increase thermal stability. Protein crystallization is more successful for proteins with a higher melting point and adding buffer components that stabilize proteins improve the likelihood of protein crystals forming. If examining pH then the possible effects of the buffer molecule on thermal stability should be taken into account along with the fact that pKa of each buffer molecule changes uniquely with temperature. Additionally, any time a charged species is examined the effects of the counterion should be accounted for.
Thermal stability of proteins has traditionally been investigated using biochemical assays, circular dichroism, or differential scanning calorimetry. Biochemical assays require a catalytic activity of the protein in question as well as a specific assay. Circular dichroism and differential scanning calorimetry both consume large amounts of protein and are low-throughput methods. The Thermofluor assay was the first high-throughput thermal shift assay and its utility and limitations has spurred the invention of a plethora of alternate methods. Each method has its strengths and weaknesses but they all struggle with intrinsically disordered proteins without any clearly defined tertiary structure as the essence of a thermal shift assay is measuring the temperature at which a protein goes from well-defined structure to disorder.
Methods
nanoDSF nano-Differential scanning fluorimetry, or nanoDSF, is a biophysical characterization technique used for assessing the conformational stability of a biological sample, typically a protein. Samples are subjected to either temperature ramps or gradients of chemical denaturant, and the intrinsic fluorescence is measured and fit to determine the melting point (Tm). Applications include formulation ranking, protein engineering (comparing mutants to wild type), and ligand binding (quantification of affinity constants). A prerequisite of the technique is that the protein must contain an intrinsically fluorescent residue, typically tryptophan or tyrosine residues. Benefits include tag-free analysis, avoidance of extrinsic fluorophores, low sample consumption, easy of use, amenity to automation, and high screening throughput. Drawbacks include a propensity for false positives and negatives, usually necessitating follow-up screening with a potentially lower-throughout orthogonal technique to confirm. Current commercial instruments employ either proprietary capillaries or generic high-throughput 384-well plates for sample analysis.
Thermofluor
The technique was first described by Semisotnov et al. (1991) using 1,8-ANS and quartz cuvettes. 3 Dimensional Pharmaceuticals were the first to describe a high-throughput version using a plate reader and Wyeth Research published a variation of the method with SYPRO Orange instead of 1,8-ANS. SYPRO Orange has an excitation/emission wavelength profile compatible with qPCR machines which are almost ubiquitous in institutions that perform molecular biology research. The name differential scanning fluorimetry (DSF) was introduced later but Thermofluor is preferable as Thermofluor is no longer trademarked and differential scanning fluorimetry is easily confused with differential scanning calorimetry.
SYPRO Orange binds nonspecifically to hydrophobic surfaces, and water strongly quenches its fluorescence. When the protein unfolds, the exposed hydrophobic surfaces bind the dye, resulting in an increase in fluorescence by excluding water. Detergent micelles will also bind the dye and increase background noise dramatically. This effect is lessened by switching to the dye ANS; however, this reagent requires UV excitation. The stability curve and its midpoint value (melting temperature, Tm also known as the temperature of hydrophobic exposure, Th) 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 ΔTm is calculated. The method may not work very well for protein-protein interactions if one of the interaction partners contains large hydrophobic patches as it is difficult to dissect prevention of aggregation, stabilization of a native folds, and steric hindrance of dye access to hydrophobic sites. In addition, partly aggregated protein can also limit the relative fluorescence increase upon heating; in extreme cases there will be no fluorescence increase at all because all protein is already in aggregates before heating. Knowing this effect can be very useful as a high relative fluorescence increase suggests a significant fraction of folded protein in the starting material.
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.
A typical assay Materials: A fluorometer equipped with temperature control or similar instrumentation (qPCR machines); suitable fluorescent dye; a suitable assay plate, such as a 96-well qPCR plate. 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 wells, comprising 11 different concentrations of a test compound with a single negative control well. 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. Centrifugation and oil dispense: Brief centrifugation (~1000 × g, 1 min) of the assay plate to mix compounds into the protein solution, 1–2 μL of silicone oil to prevent the evaporation during heating is overlaid onto the solution (some systems use plastic seals instead), followed by an additional centrifugation step (~1000 × g, 1 min). Instrumental set up: A typical temperature ramp rates range from 0.1 to 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.
CPM, thiol-specific dyes Alexandrov et al. (2008) published a variation on the Thermofluor assay where SYPRO Orange was replaced by N-[4-(7-diethylamino-4-methyl-3-coumarinyl)phenyl]maleimide (CPM), a compound that only fluoresces after reacting with a nucleophile. CPM has a high preference for thiols over other typical biological nucleophiles and therefore will react with cysteine side chains before others. Cysteines are typically buried in the interior of a folded protein as they are hydrophobic. When a protein denatures cysteine thiols become available and a fluorescent signal can be read from reacted CPM. The excitation and emission wavelengths for reacted CPM are 387 nm/ 463 nm so a fluorescence plate reader or a qPCR machine with specialized filters is required. Alexandrov et al. used the technique successfully on the membrane proteins Apelin GPCR and FAAH as well as β-lactoglobin which fibrillates on heating rather than going to a molten globule.
DSF-GTP The DSF-GTP (GFP-Tagged Protein-of-Interest) technique was developed by a team led by Patrick Schaeffer at James Cook University and published in Moreau et al. 2012. The development of differential scanning fluorimetry and the high-throughput capability of Thermofluor have vastly facilitated the screening of crystallization conditions of proteins and large mutant libraries in structural genomics programs, as well as ligands in drug discovery and functional genomics programs. These techniques are limited by their requirement for both highly purified proteins and solvatochromic dyes, prompting the need for more robust high-throughput technologies that can be used with crude protein samples. This need was met with the development of a new high-throughput technology for the quantitative determination of protein stability and ligand binding by differential scanning fluorimetry of proteins tagged with green fluorescent protein (GFP). This technology is based on the principle that a change in the proximal environment of GFP, such as unfolding and aggregation of the protein of interest, is measurable through its effect on the fluorescence of the fluorophore. The technology is simple, fast and insensitive to variations in sample volumes, and the useful temperature and pH range is 30–80 °C and 5–11 respectively. The system does not require solvatochromic dyes, reducing the risk of interferences. The protein samples are simply mixed with the test conditions in a 96-well plate and subjected to a melt-curve protocol using a real-time thermal cycler. The data are obtained within 1–2 h and include unique quality control measures through the GFP signal. DSF-GTP has been applied for the characterization of proteins and the screening of small compounds.
DCVJ, rigidity sensitive dyes 4-(dicyanovinyl)julolidine (DCVJ) is a molecular rotor probe with fluorescence that is strongly dependent on the rigidity of its environment. When protein denatures, DCVJ increases in fluorescence. It has been reported to work with 40 mg/ml of antibody.
Intrinsic tryptophan fluorescence lifetime The lifetime of tryptophan fluorescence differs between folded and unfolded protein. Quantification of UV-excited fluorescence lifetimes at various temperature intervals yields a measurement of Tm. A prominent advantage of this technique is that no reporter dyes need be added as tryptophan is an intrinsic part of the protein. This can also be a disadvantage as not all proteins contain tryptophan. Intrinsic fluorescence lifetime works with membrane proteins and detergent micelles but a powerful UV fluorophore (e.g. auto-fluorescent small molecule) in the buffer could drown out the signal.
Intrinsic tryptophan fluorescence wavelength