Fluorescence thermal shift (FTS) aka Differential Scanning Fluorimetry (DFS) aka Thermal Melt basics
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- Опубликовано: 9 июл 2024
- Fluorescence Thermal Shift (FTS), aka Differential Scanning Fluorimetry (DSF), is a way to measure the thermal stability of a protein by seeing how much heat the protein can withstand before unfolding.
Blog form: bit.ly/dsfscience
Basic idea:
* Hydrophobic (water-avoiding) parts of protein are hidden in the center of folded (water soluble) proteins
* Hydrophobic parts of protein are exposed when the protein unfolds
* Heat lets proteins wiggle, eventually causing them to unfold
* The more stable the protein is, the more heat it can withstand before unfolding
* The dye binds to hydrophobic parts of the protein
* The dye fluoresces (gives off light when you shine a specific wavelength of light on it) when bound to the protein
* The dye’s fluorescence is quenched by water when it isn’t bound to protein (I know, how rude, right?!)
* You can measure dye fluorescence as a measure of protein unfolding, since you will only see fluorescence when the dye is bound to the protein
* The maximum fluorescence is when the protein is all unfolded
* After all the protein unfolds (max fluorescence) it starts clumping together (aggregating) kicking out the dye so you stop seeing fluorescence
* The Tm is the melting temperature, which is the temperature at which half the protein is unfolded
* The peak of the slope of the curve gives you the Tm
* Complicated unfolding might give you weird peak patterns
More detailed:
Proteins are held together by various intramolecular forces, including hydrophobic interactions, which basically means that there are parts of proteins that water avoids because they’re not very interesting for the water - they just are hydrocarbony, without charged or partially charged (polar) parts that water likes. Since water avoids them, they get pushed into the center of the protein, leaving only hydrophilic (water-loved) groups on the outs.
But, when you heat up protein what’s gonna happen? As you heat the protein, you give it more wiggling energy. The more tightly held together the protein’s structure is, the more it can wiggle before it comes it unfolds, and we can kind of measure how much heat it takes to get the protein to unfold.
Specifically we measure the Tm or the melting temperature which is the temperature at which like half the as unfolded. We know when the protein unfolds the protein they’re going to be exposed to the water. If there’s nothing else around, the exposed hydrophobic parts of one copy of the protein will bind to those parts in another copy. That is, you’ll get aggregation (clumping up).
But, before that happens, you have an opportunity to sneak something else in there that likes to bind hydrophobic things. Something like our SYPRO Orange dye.
The dye will give off light when it’s bound to the protein. And it will “only” bind to the protein when the protein is unfolded. And it will have its light stolen (quenched) by water before we can see it when it’s not bound to protein.
So, we don’t see light if the protein is folded, then we do see light if the protein is unfolded, but not aggregated, then we don’t see light as the protein is aggregated (because the protein’s binding other copies of itself rather than the dye).
So what we can do is we can take the protein and we can heat it up gradually. Initially you won’t see light (because the protein is folded and the hydrophobic parts thus hidden). Then, you’ll start seeing light (because the hydrophobic parts of the protein are exposed to the dye), and then you stop seeing light as the protein aggregates, kicking out the dye.
How will we heat it? Here the thermal cycler (PCR machine) comes to the rescue. In PCR, what you do is you heat a sample of DNA over and over and over again and you change the temperature so you heat it up to separate the strands then you cool it down so the primers can bind, then then you heat it up a little more so that the copies can get made and then you heat it up more so that the strands come apart and you keep doing that over and over and over again. To do all that, you have to be able to change the temperature really quickly and go through a wide range of temperatures, so these machines are great for being able to heat up your protein for this assay (experiment).
Instead of using a normal PCR machine, you need to use a real-time PCR machine, which is able to measure fluorescence. And instead of using normal PCR tubes, you use like tubes with white walls and you need to have optically clear lids so that basically what can happen is that take a picture from the top in order to see how much fluorescence is being given off. Then you can figure out the Tm. The easiest way is to find the peak of the derivative (slope) of the fluorescence vs. temperature curve.
The top of the peak is correspond to the slope going from positive to negative, but kind of halfway to the point the slope starts slowing so you actually gonna get a peak in your derivative plot at your Tm.
Finished in comments 
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Thank you ❤
If your protein was sturdier, that is was more tightly folded, your Tm will be higher. But if your protein is not that stable is unfold or easily well then you’re gonna have a lower Tm. This sturdiness is condition-dependent, so you can measure the TM of the protein under various conditions like different pH, in the presence of various salt concentrations, etc. to see how that affects its stability. You can even make mutations to the protein and see if that changes the Tm.
One thing that often changes the Tm is binding to something (a ligand). We can introduce the ligand to the protein and what’s gonna happen is often when the protein binds, it’s going to become sturdier because it has additional kind of molecular interactions holding it together going to be pulled up tightly and therefore it’ll raise the Tm. So you can actually use this essay as a way to kind of see the binding.
It often takes some troubleshooting to get things in the right concentration range getting the ratio of protein so you might need to play around a little bit.
Here’s the protocol we’ve been going off of. Thanks’ MCC! www.mdhcures.org/s/MCC-MDH-Fast-Thermal-Melt-V3.pdf
And here’s an article that might be of interest: Wu, T., Hornsby, M., Zhu, L., Yu, J. C., Shokat, K. M., & Gestwicki, J. E. (2023). Protocol for performing and optimizing differential scanning fluorimetry experiments. STAR protocols, 4(4), 102688. doi.org/10.1016/j.xpro.2023.102688
Happy experimenting!
More about protein structure: bit.ly/proteinstructure & ruclips.net/video/V2pStHenOJo/видео.html
More about fluorescence & FRET: bit.ly/FRET_2 & ruclips.net/video/crA5f1ZGCLo/видео.html
More about PCR: bit.ly/pcrtrain
More about qPCR: bit.ly/rtrtqpcrprimer
more about all sorts of things: #365DaysOfScience All (with topics listed) 👉 bit.ly/2OllAB0 or search blog: thebumblingbiochemist.com
#scicomm #biochemistry #molecularbiology #biology #sciencelife #science #realtimechem
nice explanation 🙂
Thank you 🙂
Hi Dr. Bibel,
Thank you for the excellent content. I would like to know if I can use DSF to check the integrity of membrane proteins, such as GPCRs. Does the protein need to be purified, or can I perform the assay using a crude extract obtained after ultracentrifugation (membrane preparation), where the lipid bilayer remains intact?
I think having the lipids will cause high background and prevent it from working well
@@thebumblingbiochemist Thank you!
I was thinking how would you know the fluorescent dye will be reliable; any controlled experiment to confirm?
You should include a negative control without your protein. In terms of positive controls, that's hard because it's protein-dependent. But if you have another higher-tech technique you can do to compare and ensure, then you can use this?
What if a ligand binds to both the folded and unfolded forms with the same or similar energy? or better to the unfolded form?
Things would get complicated, I suppose