Thermal shift assay
A thermal shift assay measures changes in the thermal denaturation temperature and hence stability of a protein under varying conditions such as variations in drug concentration, buffer pH or ionic strength, redox potential, or sequence mutation. The most common method for measuring protein thermal shifts is differential scanning fluorimetry or thermofluor, which utilizes specialized fluorogenic dyes.
The binding of low molecular weight ligands can increase the thermal stability of a protein, as described by Daniel Koshland and Kaj Ulrik Linderstrøm-Lang and Schellman. 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
DSF-GTP
The DSF-GTP 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. 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.Thermofluor
The technique was first described by Semisotnov et al. 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 :de:SYPRO Orange|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 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 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 ; 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 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, followed by an additional centrifugation step.
- Instrumental set up: A typical temperature :wikt:ramp|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.
CPM, thiol-specific dyes
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.
DCVJ, rigidity sensitive dyes
4-julolidine 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 fluorescer in the buffer could drown out the signal and few articles are published using the technique for thermal shift assays.Intrinsic tryptophan fluorescence wavelength
The excitation and emission wavelengths of tryptophan are dependent on the immediate environment and therefore differs between folded and unfolded protein, just as the fluorescence lifetime. Currently there are at least two machines on the market that can read this shift in wavelength in a high-throughput manner while heating the samples. The advantages and disadvantages are the same as for fluorescence lifetime except that there are more examples in the scientific literature of use.Static light scattering
allows monitoring of the sizes of the species in solution. Since proteins typically aggregate upon denaturation the detected species size will go up.This is label-free and independent of specific residues in the protein or buffer composition. The only requirement is that the protein actually aggregates/fibrillates after denaturation and that the protein of interest has been purified.
FastPP
In fast parallel proteolysis the researcher adds a thermostable protease and takes out samples in parallel upon heating in a thermal gradient cycler. Optionally, for instance for proteins expressed at low levels, a western blot is then run to determine at what temperature a protein becomes degraded. For pure or highly enriched proteins, direct SDS-PAGE detection is possible facilitating Commassie-fluorescence based direct quantification. FastPP exploits that proteins become increasingly susceptible to proteolysis when unfolded and that thermolysin cleaves at hydrophobic residues which are typically found in the core of proteins.To reduce the workload, western blots could be replaced by SDS-PAGE gel polyhistidine-tag staining, provided that the protein has such a tag and is expressed in adequate amounts.
FastPP can be used on unpurified, complex mixtures of proteins and proteins fused with other proteins, such as GST or GFP, as long as the sequence that is the target of the western blot, e.g., His-tag, is directly linked to the protein of interest. However, commercially available thermolysin is dependent on calcium ions for activity and denatures itself just above 85 degrees Celsius. So calcium must be present and calcium chelators absent in the buffer - other compounds that interfere with the function of the protease could also be problematic.
FASTpp has also been used to monitor binding-coupled folding of intrinsically disordered proteins.
CETSA
Cellular thermal shift assay is a biophysical technique applicable on living cells as well as tissue biopsies. CETSA is based on the discovery that protein melting curves can also be generated in intact cells and that drug binding leads to very significant thermal stabilization of proteins. Upon denaturation, proteins are aggregated and can thus be removed by centrifugation after lysis of the cells. The stable proteins are found in the supernatant can be detected; e.g., by western blot, alpha-LISA, or mass spectrometry. The CETSA-technique is highly stringent, reproducible, and not prone to false positives. However, it is possible for a sample, or small molecule compound, to bind a protein in a given target's pathway. If that protein induces further stabilization of the original target protein through a cascade event, it could manifest as direct target engagement. An advantage of this method is that it is label-free and thus applicable for studies of drug binding in a wide range of cells and tissues. CETSA can also be conducted on cell lysates versus intact cells, helping to determine sample penetration of the cell membrane.ThermoFAD
Thermofluor variant specific for flavin-binding proteins. Analogous to thermofluor binding assays, a small volume of protein solution is heated up and the fluorescence increase is followed as function of temperature.In contrast to thermofluor, no external fluorescent dye is needed because the flavin cofactor is already present in the flavin-binding protein and its fluorescence properties change upon unfolding.
SEC-TS
can be used directly to access protein stability in the presence or absence of ligands. Samples of purified protein are heated in a water bath or thermocycler, cooled, centrifuged to remove aggregated proteins, and run on an analytical HPLC. As the melting temperature is reached and protein precipitates or aggregates, peak height decreases and void peak height increases. This can be used to identify ligands and inhibitors, and optimize purification conditions.While of lower throughput than FSEC-TS, requiring large amounts of purified protein, SEC-TS avoids any influence of the fluorescent tag on apparent protein stability.
FSEC-TS
In fluorescence-detection size exclusion chromatography the protein of interest is fluorescently tagged and run through a gel filtration column on an FPLC system equipped with a fluorescence detector. The resulting chromatogram allows the researcher to estimate the dispersity and expression level of the tagged protein in the current buffer. Since only fluorescence is measured, only the tagged protein is seen in the chromatogram. FSEC is typically used to compare membrane protein orthologs or screen detergents to solubilize specific membrane proteins in.For fluorescence-detection size-exclusion chromatography-based thermostability assay the samples are heated in the same manner as in FastPP and CETSA and following centrifugation to clear away precipitate the supernatant is treated in the same manner as FSEC. Larger aggregates are seen in the void volume while the peak height for the protein of interest decreases when the unfolding temperature is reached.
GFP has a Tm of ~76 °C so the technique is limited to temperature below ~70 °C.
Radioligand binding thermostability assay
s are pharmacologically important transmembrane proteins. Their X-ray crystal structures were revealed long after other transmembrane proteins of lesser interest. The difficulty in obtaining protein crystals of GPCRs was likely due to their high flexibility. Less flexible versions were obtained by truncating, mutating, and inserting T4 lysozyme in the recombinant sequence. One of the methods researchers used to guide these alterations was radioligand binding thermostability assay.The assay is performed by incubating the protein with a radiolabelled ligand of the protein for 30 minutes at a given temperature, then quench on ice, run through a gel filtration mini column, and quantify the radiation levels of the protein that comes off the column. The radioligand concentration is high enough to saturate the protein. Denatured protein is unable to bind the radioligand and the protein and radioligand will be separated in the gel filtration mini column. When screening mutants selection will be for thermal stability in the specific conformation, i.e., if the radioligand is an agonist, selection will be for the agonist binding conformation and if it is an antagonist, then the screening is for stability in the antagonist binding conformation.
Radioassays have the advantage of working with minute amounts of protein. But it is work with radioactive substances and large amount of manual labour is involved. A high-affinity ligand has to be known for the protein of interest and the buffer must not interfere with the binding of the radioligand. Other thermal shift assays can also select for specific conformations if a ligand of the appropriate type is added to the experiment.
Comparisons of the various approaches
Applications
Label-free drug screening
Thermofluor has been extensively used in drug screening campaigns.Because thermofluor detects high affinity binding sites for small molecules on proteins, it can find hits that bind to active site subsites, cofactor sites, or allosteric binding sites with equal efficacy. The method typically requires the use of screening compound concentrations at >10x the desired binding threshold. Setting 5 μM as a reasonable hit threshold consequently requires a test ligand concentration of 50 to 100 μM in the sample well. For most drug compound libraries, where many compounds are not soluble beyond ~100 μM, screening multiple compounds is consequently not feasible owing to solubility issues. Thermofluor screens do not require the development of custom screening reagents, do not require any radioactive reagents, and are generally less sensitive to the effects of compounds that are chemically reactive with protein active site residues, and that consequently show up as undesirable hits in enzyme activity screens.