The number of stories and journal articles about how CRISPR DNA-editing technology works, has worked, and is planned to work are beyond counting. How about an article about how to stop it in its tracks? That’s this one, just published in Cell from a multicenter team in Cambridge and New York. It describes a screening program for small-molecule inhibitors of S. pyogenes Cas9 (spCas9), because one would want some ways (not all of which currently exist) to turn its effects off in given places and at given times. There are already protein-based inhibitors, but to the best of my knowledge, this is the first report of small-molecule ones. And in a line I like very much, the paper states “Unsurprisingly, the pharmacological inhibition of intracellular proteins is usually accomplished using small molecules.”

Yes, it is! So how did they find these? Because setting up a Cas9 inhibitor screen is not so straightforward. For one thing, the enzyme binds its substrate with picomolar affinity. It’s also (obviously) a DNA-binding protein, and finding small-molecule inhibitors of that interaction has been. . .well, let’s say “challenging”, with all the baggage that adjective implies. What’s more, a direct inhibitor would need to deal with two nuclease domains, and the overall structure of the Cas9 protein has some unusual protein folds in general, so your chances of getting hits from a standard screening collection could be diminished for that reason, too. Finally, as the authors note, no one has yet reported a Cas9 screen that’s even suitable for high-throughput screening (in terms of miniaturization, signal/noise, readout, etc.) Thus all the co-authors, no doubt.

The protein-based inhibitors, though, are known to target the binding of the Cas9 protein to the protospacer adjacent motif (PAM) on DNA, making it an obvious choice for a screen (the affinity there is much less daunting, while still vital for CRISPR function). The team set up a fluorescence polarization assay, a DSF (differential scanning fluorimetry) assay, and a BLI (biolayer interference) assay as a suite of ways to look for inhibitors, and for people who don’t do this stuff for a living, I’ll briefly go into what the heck those are. All such assays are built around some readout that changes when a molecule binds to a protein, but man, are there ever a lot of ways to skin that particular cat.

FP assays depend on fluorescently labeled species, which emit light at specific wavelength when another specific wavelength excites them. Think of a fluorescent sticker, which might give off bright green light when ultraviolet light hits it. If you used polarized light to do this excitation, you get polarized fluorescent light back out – but as it happens, the kind you get varies. If you shoot vertically polarized light in and measure both vertical and horizontally polarized light on the way out (at the new fluorescent-emitter wavelength) you’ll see a mixture of the two: and that mixture depends on the physical rotation, in solution, of the fluorescent species. So if something has bound to it in your assay, its rotation rate will slow down, and the larger the binding species, the more it’ll slow. Changes in fluorescent polarization are thus a measure of binding events.

DSF is a variety of thermal shift assay. You add a particular type of dye to a sample of your protein target of interest, a dye that is fluorescent but whose fluorescence is quenched when the dye molecules are floating around in water solution. You warm up the sample while watching for the particular fluorescence wavelength of your dye to appear. At the melting point of the protein, you’ll see a pretty sharp increase in the signal, because the protein has now unfolded and thus exposed some new hydrophobic surfaces for the dye molecules to bind to – and now that they’re out of aqueous solution, they will fluoresce. The key thing is, when a ligand has bound to the protein, the resulting complex is generally more stable, and has a higher melting point. You test small molecules against the protein/dye system, looking for the ones that make the complex melt (and hence fluoresce) at higher temperatures. In other words, you’re looking for a thermal shift, and generally the larger the better. You can, of course, also look for compounds that are disrupting some complex that’s already bound (of two proteins, say, or of protein/DNA), in which case you’ll be watching for lower melting points.

Now, BLI is the only one of these three that I haven’t been involved with. The acronym is “bio-layer interferometry“, and it depends on being able to immobilize your protein target onto a small tip. Quite a few of these biophysical techniques depend on that sort of attachment; it has its good points and its bad ones. Chief among the latter is that you may find that you can’t seem to attach your protein in a useful manner, and it’s often impossible to say just why that’s happening, which leads you to try another attachment chemistry method, and then another, and there are a lot of choices. For BLI, you’re shining light on the spot of attached protein and reading out the resulting interference pattern. If ligands bind to that protein sample, though, the optical thickness of layer increases, and shining light on it now gives you a shifted interference pattern, and you’re measuring the difference. Optical interference patterns are brutally sensitive – in my weekend hobby of astronomy, that’s how you test telescope mirrors during their production, and it’s a very unforgiving test indeed. One nice thing about this instrument is that you can monitor the binding event (and its dissociation, if you want) in real time, a feature BLI shares with another common immobilized-protein biophysical assay, SPR.

Back to CRISPR inhibitors. Running three orthogonal technologies for a screen like this is highly recommended, because each of them (like any assay) is subject to its own false positive and false negatives. What you want, ideally, are compounds that work across several very different techniques, greatly increasing your confidence that real protein binding events are at the center of it all. And indeed, when this team ran a diverse set of about 10,000 molecules across the FP assay, they found particularly enriched hits in three different chemical classes. The first class, though, tended to give an FP signal even when you left out the Cas9 protein in the assay, which is why you had better run those control experiments! The second class of compounds tended to have fluorescence of their own, which always complicates the assay interpretation (and is one of the particular banes of FP and other fluorescence-driven assay techniques), and in addition were also cytotoxic when taken on into cells. So that left the third class.

That one made it through all the assay techniques, and was active in cells as well. Control experiments suggested that it was indeed affect the Cas9/PAM interaction, and in cells, the compounds were able to disrupt a model system where dCas9 caused upregulation of particular targeted genes (shown at right). So these really do look like Cas9 inhibitors, and it appears that they’re interacting first with the complex of Cas9 and its guide RNA. I’ve left out a lot of validation experiments, I should add:

Our screening strategy involved disrupting DNA binding by SpCas9, followed by demonstrating activity in multiple mammalian cell lines using gene or protein delivery. Furthermore, we demonstrated inhibition of SpCas9 nuclease and transcription activation in assays with gain of signal (e.g., eGFP-disruption assay), loss of signal (e.g., HiBiT assay), various DNA repair pathways, and a myriad of readouts (e.g., fluorescence, luminescence, next-generation sequencing [NGS], qPCR).

What exactly is going on – how compound binding disrupts the enzyme activity in detail – remains to be seen. It could be direct competition, could be an allosteric site, who knows. The SAR of the compounds, from what I can see, looks pretty tight. Small changes in the compound structure or in the protein target can have big effects, which suggests some sort of tight, specific interaction. The hope is that structural biology techniques will be able to shed some light on the actual structures of the bound forms. Along the way, we’ll probably learn quite a bit about CRISPR inhibition (and what it can do), as well as about the CRISPR enzymes themselves.