Jump to content

User:Structurebiologist/sandbox

From Wikipedia, the free encyclopedia
This is an old revision of this page, as edited by Structurebiologist (talk | contribs) at 16:56, 6 June 2015. The present address (URL) is a permanent link to this revision, which may differ significantly from the current revision.

A thermal shift assay quantifies the change in thermal denaturation temperature of a protein under varying conditions. The differing conditions that can be examined are very diverse, e.g. pH, salts, additives, drugs, drug leads, oxidation/reduction, or mutations. The binding of low molecular weight ligands can increase the thermal stability of a protein, as described by Koshland (1958)[1] and Linderstrom-Lang and Schellman (1959).[2] Almost half of enzymes require a metal ion co-factor.[3] Thermostable proteins are often more useful than their non-thermostable counterparts, e.g. DNA polymerase in the polymerase chain reaction,[4] so protein engineering often includes adding mutations to increase thermal stability. 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.[5] 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.

Thermofluor

Diagram

The technique was first described by Semisotnov et al. (1991)[6] using 1,8-ANS and quartz cuvettes. Millenium Pharmaceuticals were the first to describe a high-throughput version using a plate reader[7] and Wyeth Research published a variation of the method with SYPRO Orange instead of 1,8-ANS.[8] SYPRO Orange has an ex/em (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[9] 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. 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.

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.[10]

Distinguishing features relative to other thermal shift assays

  • Low barrier of entry

Concerns relative to other thermal shift assays

  • Protein of interest must be purified and free of fusion proteins such as GST, MBP, or GFP
  • High background signal with detergent concentrations higher than cmc (critical micelle concentration)
  • High background signal with very high concentrations of some organic molecules
  • Some proteins cause very high background signals from the fluorophore in their folded state
  • Some proteins cause a shift in ex/em wavelengths
  • Some proteins fibrillate instead of going to a molten globule state
  • DMSO and fluorophore dye added to experiment

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.[11]
  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.[11]
  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.[11]
  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).[11]
  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.[11]

CPM, thiol-specific dyes

Alexandrov et al. (2008)[12] 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 ex/em wavelengths for CPM are 387 nm/ 463 nm so a fluorescence plate reader or a qPCR machine with specialised 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.

Distinguishing features relative to other thermal shift assays

  • Works with detergents
  • Works with (some) fibrillating proteins

Concerns relative to other thermal shift assays

  • Protein of interest must be purified and free of fusion proteins such as GST, MBP, or GFP
  • Few qPCR machines can be fitted with appropriate filters and many fluorescence plate readers are not capable of heating to a high temperature
  • Some proteins do not have cysteines
  • Some proteins have cysteines on their surface
  • Some proteins some proteins have all their cysteines engaged in disulphide bonds
  • Thiols and other strong nucleophiles can not be added to the experiment
  • CPM will over time react with other nucleophiles so measurements must be performed straight after mixing protein and dye
  • DMSO and fluorophore dye added to experiment

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.[13]

Distinguishing features relative to other thermal shift assays

  • Works with detergents
  • Low barrier of entry

Concerns relative to other thermal shift assays

  • Protein of interest must be purified and free of fusion proteins such as GST, MBP, or GFP
  • Uses extremely high concentrations of protein
  • Few examples of use in the literature
  • DMSO and fluorophore dye added to experiment

Intrinsic fluorescence lifetime

The lifetime of tryptophan fluorescence differs between folded and unfolded protein and by measuring the lifetime of UV-excited fluorescence at temperature intervals you can get a measurement of the melting temperature of your protein. The NovaFluorPR Fluorescence Lifetime Plate Reader is an example of a machine that can perform this measurement.

A prominent advantage is that no reporter dyes are required to 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 em/ex wavelength of 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 NanoTemper Prometheus and the Unchained Labs UNit.

Technique-wise, the advantages and disadvantages are basically the same as for fluorescence lifetime except that there are more examples in the scientific literature of use.

Static light scattering

Static light scattering allows monitoring of the sizes of the species in solution. Since proteins typically aggregate upon denaturation (or form fibrils) the detected species size will go up. Both Epiphyte3's Stargazer-2 and Unchained Labs' UNit measure SLS.

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 (thermolysin) and takes out samples regularly while heating.[14] A Western blot is then run to determine at what temperature a protein becomes degraded. FastPP exploits that proteins become extra vulnerable to proteolysis when unfolded and that thermolysin cleaves between hydrophobic residues which are typically found in the core of proteins.

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 80 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.

CETSA

CETSA stands for cellular thermal shift assay.[15] Very similar to fastPP but instead of adding a protease samples are centrifuged after heating and the supernatant is run on Western blots or two target-directed antibodies are added and their relative proximity is detected through FRET(?). When a protein denatures it aggregates and becomes part of the pellet and can therefore not be found in the supernatant.

It is a powerful technique that works on both complex mixtures and whole cells but requires antibodies. The protein of interest should not be fused to other proteins such as MBP or GFP as they could keep a denatured protein in solution.

FSEC-TS

In fluorescence-detection size exclusion chromatography the protein of interest is fluorescently tagged, e.g. with GFP, and run through a gel filtration column on an FPLC system equipped with a fluorescence detector. The resulting chromatogram allows allows the researcher to estimate the monodispersity and expression level of the tagged protein in the current buffer.[16] Since only fluorescence is measured only the tagged protein is seen in the chromatogram. FSEC is typically used to compare membrane protein orthologs and screen detergents to solubilise specific membrane proteins in.

For fluorescence-detection size-exclusion chromatography-based thermostability assay (FSEC-TS) the samples are heated in the same manner as in FastPP and CETSA and after centrifugation to clear away precipitate the supernatant is treated in the same manner as FSEC.[17] 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. Of all the techniques mentioned it may have the lowest throughput but it is free of antibody requirements and works on complex mixtures of proteins.

Comparisons of the various approaches

The table's caption
blank? Thermofluor CPM DCVJ Tryptophan fluorescence lifetime Tryptophan fluorescence wavelength Static light scattering FastPP CETSA FSEC-TS
Purification Pure protein Pure protein Pure protein at very high concentration Pure protein Pure protein Pure protein Complex mixture Complex mixture Complex mixture
Detergents Below cmc Yes Yes Yes Yes Yes Yes Yes Yes
Buffer Avoid fluorophores, very high conc. organic solvents Avoid thiols, possibly other nucleophiles - - - - - - -
Thoughtput Highest Highest Highest Intermediate Intermediate Intermediate Slowest Slowest Slowest
Row heading A Below cmc Yes Yes Yes Yes Yes Yes Yes Yes
Row heading A Cell B Cell C
Row heading A Cell B Cell C
Row heading A Cell B Cell C

References

  1. ^ Koshland, DE (February 1958). "Application of a Theory of Enzyme Specificity to Protein Synthesis". Proceedings of the National Academy of Sciences of the United States of America. 44 (2): 98–104. PMID 16590179.
  2. ^ Linderstrøm-Lang, K., and Schellman, J. A. (1959). "Protein structure and enzyme activity". The Enzymes. 1(2) 443-510.
  3. ^ Waldron, KJ; Rutherford, JC; Ford, D; Robinson, NJ (13 August 2009). "Metalloproteins and metal sensing". Nature. 460 (7257): 823–30. PMID 19675642.
  4. ^ Saiki, RK; Gelfand, DH; Stoffel, S; Scharf, SJ; Higuchi, R; Horn, GT; Mullis, KB; Erlich, HA (29 January 1988). "Primer-directed enzymatic amplification of DNA with a thermostable DNA polymerase". Science (New York, N.Y.). 239 (4839): 487–91. PMID 2448875.
  5. ^ Grøftehauge, MK; Hajizadeh, NR; Swann, MJ; Pohl, E (1 January 2015). "Protein-ligand interactions investigated by thermal shift assays (TSA) and dual polarization interferometry (DPI)". Acta crystallographica. Section D, Biological crystallography. 71 (Pt 1): 36–44. PMID 25615858.
  6. ^ Semisotnov, GV; Rodionova, NA; Razgulyaev, OI; Uversky, VN; Gripas', AF; Gilmanshin, RI (January 1991). "Study of the "molten globule" intermediate state in protein folding by a hydrophobic fluorescent probe". Biopolymers. 31 (1): 119–28. PMID 2025683.
  7. ^ Pantoliano, MW; Petrella, EC; Kwasnoski, JD; Lobanov, VS; Myslik, J; Graf, E; Carver, T; Asel, E; Springer, BA; Lane, P; Salemme, FR (December 2001). "High-density miniaturized thermal shift assays as a general strategy for drug discovery". Journal of biomolecular screening. 6 (6): 429–40. PMID 11788061.
  8. ^ Lo, MC; Aulabaugh, A; Jin, G; Cowling, R; Bard, J; Malamas, M; Ellestad, G (1 September 2004). "Evaluation of fluorescence-based thermal shift assays for hit identification in drug discovery". Analytical biochemistry. 332 (1): 153–9. PMID 15301960.
  9. ^ Niesen, FH; Berglund, H; Vedadi, M (2007). "The use of differential scanning fluorimetry to detect ligand interactions that promote protein stability". Nature protocols. 2 (9): 2212–21. PMID 17853878.
  10. ^ Ciulli, A; Abell, C (December 2007). "Fragment-based approaches to enzyme inhibition". Current opinion in biotechnology. 18 (6): 489–96. PMID 17959370.
  11. ^ a b c d e Cite error: The named reference first was invoked but never defined (see the help page).
  12. ^ Alexandrov, AI; Mileni, M; Chien, EY; Hanson, MA; Stevens, RC (March 2008). "Microscale fluorescent thermal stability assay for membrane proteins". Structure (London, England : 1993). 16 (3): 351–9. PMID 18334210.
  13. ^ Menzen, T; Friess, W (February 2013). "High-throughput melting-temperature analysis of a monoclonal antibody by differential scanning fluorimetry in the presence of surfactants". Journal of pharmaceutical sciences. 102 (2): 415–28. PMID 23212746.
  14. ^ Minde, DP; Maurice, MM; Rüdiger, SG (2012). "Determining biophysical protein stability in lysates by a fast proteolysis assay, FASTpp". PloS one. 7 (10): e46147. PMID 23056252.
  15. ^ Jafari, R; Almqvist, H; Axelsson, H; Ignatushchenko, M; Lundbäck, T; Nordlund, P; Martinez Molina, D (September 2014). "The cellular thermal shift assay for evaluating drug target interactions in cells". Nature protocols. 9 (9): 2100–22. PMID 25101824.
  16. ^ Kawate, T; Gouaux, E (April 2006). "Fluorescence-detection size-exclusion chromatography for precrystallization screening of integral membrane proteins". Structure (London, England : 1993). 14 (4): 673–81. PMID 16615909.
  17. ^ Hattori, M; Hibbs, RE; Gouaux, E (8 August 2012). "A fluorescence-detection size-exclusion chromatography-based thermostability assay for membrane protein precrystallization screening". Structure (London, England : 1993). 20 (8): 1293–9. PMID 22884106.

Category:Microbiology techniques

Retrieved from "https://en.wikipedia.org/w/index.php?title=User:Structurebiologist/sandbox&oldid=665778201"