Many readers will be familiar, at least in principle, with the "thermal shift assay". It goes by other names as well, but the principle is the same. The idea is that when a ligand binds to a protein, it stabilizes its structure to some degree. This gets measured by watching its behavior as samples of bound and unbound proteins are heated up, and the most common way to detect those changes in protein structure (and stability) is by using a fluorescent dye. Thus another common name for the assay, DSF, for Differential Scanning Fluorimetry. The dye has a better chance to bind to the newly denatured protein once the heat gets to that point, and that binding even can be detected by increasing fluorescence. The assay is popular, since it doesn't require much in specialized equipment and is pretty straightforward to set up, compared to something like SPR. Here's a nice slide presentation that's up on the web from UC Santa Cruz, and here's one of many articles on using the technique for screening.
I bring this up because of this paper last suumer in Science, detailing what the authors (a mixed team from Sweden and Singapore) called CETSA, the cellular thermal shift assay. They trying to do something that is very worthwhile indeed: measuring ligand binding inside living cells. Someone who's never done drug discovery might imagine that that's the sort of thing that we do all the time, but in reality, it's very tricky. You can measure ligand binding to an isolated protein in vitro any number of ways (although they may or may not give you the same answer!), and you can measure downstream effects that you can be more (or less) confident are the result of your compound binding to a cellular target. But direct binding measurements in a living cell are pretty uncommon.
I wish they weren't. Your protein of interest is going to be a different beast when it's on the job in its native environment, compared to sitting around in a well in some buffer solution. There are other proteins for it to interact with, a whole local environment that we don't know enough to replicate. There are modifications to its structure (phosphorylation and others) that you may or may not be aware of, which can change things around. And all of these have a temporal dimension, changing under different cellular states and stresses in ways that are usually flat-out impossible to replicate ex vivo.
Here's what this new paper proposes:
We have developed a process in which multiple aliquots of cell lysate were heated to different temperatures. After cooling, the samples were centrifuged to separate soluble fractions from precipitated proteins. We then quantified the presence of the target protein in the soluble fraction by Western blotting . . .
Surprisingly, when we evaluated the thermal melt curve of four different clinical drug targets in lysates from cultured mammalian cells, all target proteins showed distinct melting curves. When drugs known to bind to these proteins were added to the cell lysates, obvious shifts in the melting curves were detected. . .
That makes it sound like the experiments were all done after the cells were lysed, which wouldn't be that much of a difference from the existing thermal shift assays. But reading on, they then did this experiment with methotrexate and its enzyme target, dihydrofolate reductase (DHFR), along with ralitrexed and its target, thymidylate synthase:
DHFR and TS were used to determine whether CETSA could be used in intact cells as well as in lysates. Cells were exposed to either methotrexate or raltitrexed, washed, heated to different temperatures, cooled, and lysed. The cell lysates were cleared by centrifugation, and the levels of soluble target protein were measured, revealing large thermal shifts for DHFR and TS in treated cells as compared to controls. . .
So the thermal shift part of the experiment is being done inside the cells themselves, and the readout is the amount of non-denatured protein left after lysis and gel purification. That's ingenious, but it's also the sort of idea that (if it did occur to you) you might dismiss as "probably not going to work" and/or "has surely already been tried and didn't work". It's to this team's credit that they ran with it. This proves once again the soundness of Francis Crick's advice (in his memoir What Mad Pursuitand other places) to not pay too much attention to your own reasoning about how your ideas must be flawed. Run the experiment and see.
A number of interesting controls were run. Cell membranes seem to be intact during the heating process, to take care of one big worry. The effect of ralitrexed added to lysate was much greater than when it was added to intact cells, suggesting transport and cell penetration effects. A time course experiment showed that it took two to three hours to saturate the system with the drug. Running the same experiment on starved cells gave a lower effect, and all of these point towards the technique doing what it's supposed to be doing - measuring the effect of drug action in living cells under real-world conditions.
There's even an extension to whole animals, albeit with a covalent compound, the MetAP2 inhibitor TNP-470. It's a fumagillin derivative, so it's a diepoxide to start off, with an extra chloroacetamide for good measure. (You don't need that last reactive group, by the way, as Zafgen's MetAP2 compound demonstrates). The covalency gives you every chance to see the effect if it's going to be seen. Dosing mice with the compound, followed by organ harvesting, cell lysis, and heating after the lysis step showed that it was indeed detectable by thermal shift after isolation of the enzyme, in a dose-responsive manner, and that there was more of it in the kidneys than the liver.
Back in the regular assay, they show several examples of this working on other enzymes, but a particularly good one is PARP. Readers may recall the example of iniparib, which was taken into the clinic as a PARP-1 inhibitor, failed miserably, and was later shown not to really be hitting the target at all in actual cells and animals, as opposed to in vitro assays. CETSA experiments on it versus olaparib, which really does work via PARP-1, confirm this dramatically, and suggest that this assay could have told everyone a long time ago that there was something funny about iniparib in cells. (I should note that PARP has also been a testbed for other interesting cell assay techniques).
This leads to a few thoughts on larger questions. Sanofi went ahead with iniparib because it worked in their assays - turns out it just wasn't working through PARP inhibition, but probably by messing around with various cysteines. They were doing a phenotypic program without knowing it. This CETSA technique is, of course, completely target-directed, unless you feel like doing thermal shift measurements on a few hundred (or few thousand) proteins. But that makes me wonder if that's something that could be done. Is there some way to, say, impregnate the gel with the fluorescent shift dye and measure changes band by band? Probably not (the gel would melt, for one thing), but I (or someone) should listen to Francis Crick and try some variation on this.
I do have one worry. In my experience, thermal shift assays have not been all that useful. But I'm probably looking at a sampling bias, because (1) this technique is often used for screening fragments, where the potencies are not very impressive, and (2) it's often broken out to be used on tricky targets that no one can figure out how to assay any other way. Neither of those are conducive to seeing strong effects; if I'd been doing it on CDK4 or something, I might have a better opinion.
With that in mind, though, I find the whole CETSA idea very interesting, and well worth following up on. Time to look for a chance to try it out!