Reverse micelle encapsulation for measuring low affinities

NMR is among the more sensitive fragment-finding techniques: the starting point for clinical compound ASTX660 had low millimolar affinity at best. Now, three papers by A. Joshua Wand and colleagues at University of Pennsylvania have taken sensitivity to a new level, enabling the detection of fragments that bind hundreds of times less tightly. (Derek Lowe recently wrote about one of them, and I highlighted a talk last year.)

All three papers focus on a method called reverse micelle encapsulation, in which an aqueous solution of protein and ligand is encapsulated in nanoscale reverse micelles measuring less than 100 Å in diameter. At this size, each micelle will contain at most just a single protein and a few thousand water molecules. Because of the small volume, the protein concentration – and that of any fragments – will be extraordinarily high. The micelles have polar groups pointed inwards towards their watery interior, and their hydrophobic tails point out towards solvent, typically pentane. The overall water content of the sample is typically around 2%.

Various NMR techniques can be used to study the proteins. Although the reverse micelles are larger than the proteins themselves and thus would be expected to tumble more slowly, the low viscosity of the pentane solvent makes up for this, providing high-quality spectra.

The primary paper, in ACS Chem. Biol., focuses specifically on fragments. To establish that the technique can detect weak interactions, the researchers show that they can measure the 26 mM dissociation constant of adenosine monophosphate to the enzyme dihydrofolate reductase.

Next, they turned to the protein interleukin-1β (IL-1β), an inflammatory target with no reported small-molecule binders. One challenge of the method is that hydrophobic fragments could partition into the micelles or even diffuse into the pentane, thus reducing their concentration. To avoid this, the researchers assembled a library of 233 very polar, water-soluble fragments with cLogP values < 0.5. A 2-dimensional NMR screen (¹⁵N-TROSY) using standard conditions (100 µM protein and 800 µM fragment) yielded no hits.

In contrast, NMR screening using reverse micelles with the protein at an effective concentration of 5 mM and fragments at 40 mM yielded 31 hits. Chemical shift perturbations (CSPs) were used to determine where they were binding. Ten of the fragments didn’t show clear binding to specific sites on the protein, but the remaining 21 did, with all but one binding to multiple sites. Of these, 13 also showed non-specific interactions with other regions of the protein. Altogether, the fragment binding sites covered 67% of the protein surface, with the receptor-binding interface particularly well-represented.

Concentration-dependent CSPs were used to determine dissociation constants, which ranged from 50 mM to over 1 M. An SAR-by-catalog exercise was able to improve the affinity of one fragment from 200 mM to 50 mM at one site, though it also binds three other sites with slightly weaker affinity.

The second paper, also in ACS Chem. Biol., uses IL-1β but focuses on the interaction of even smaller molecules such as pyrimidine, methylammonium, acetonitrile, ethanol, N-methylacetamide, and imidazole. Not surprisingly, the dissociation constants are even weaker, averaging 1.5 – 2.5 M.

Finally, a Methods in Enzymology paper goes into depth on how to actually run the experiments, including details on choosing detergents and making the micelles. At high fragment concentrations, for example, pH needs to be carefully controlled.

Five years ago we asked “how weak is too weak” for a fragment. In terms of practicality, I’d say that these fragments qualify. Indeed, the ligand efficiency for the best fragment mentioned above is just 0.15 kcal mol^-1 atom^-1.

But the findings do raise the almost philosophical question of what exactly constitutes a small molecule binding site. Astex researchers reported several years ago that most proteins have more than one, and their more recent work with MiniFrags suggest on average 10 sites at high enough concentrations. Similar results were also reported earlier this month from Monash. Whether or not the fragments from such screens turn out to be immediately useful, they could certainly advance our understanding of molecular recognition.

Fragment-based Drug Design Down Under 2019

The last major fragment meeting of 2019 took place at the Monash Institute of Pharmaceutical Sciences, Monash University, in marvelous Melbourne last week. This was the third Australian meeting devoted to fragments; you can read about the first, in 2012, here. With some 125 participants from four continents, two dozen talks, and nearly as many posters I’ll just try to capture major themes.

Biophysics played a starring role – if you haven’t already voted (right side of page) on which fragment-finding techniques you use please do so. Sarah Piper (Monash) discussed cryo-electron microscopy and showed some lovely high-resolution structures of proteins with bound ligands, though not yet with fragments. Sally-Ann Poulsen (Griffith University) described using native-state ESI mass spectrometry to discover new carbonic anhydrase binding fragments (see here). She uses a 96-well “nanoESI” chip to generate 5 µm droplets as opposed to the ~100 µm droplets typically fed into the instrument. Smaller droplets contain fewer molecules of salt and buffer, and thus generate cleaner spectra.

NMR screening is the go-to method for screening at Monash University, as highlighted by Martin Scanlon and multiple other speakers. Indeed, Monash has built their own version of Astex’s MiniFrag library – their MicroFrags include 92 compounds with 5-8 non-hydrogen atoms. Rebecca Whitehouse has screened these at 300 mM (yes, millimolar) by ¹⁵N-¹H HSQC against the E. coli protein DsbA (EcDsbA) and found numerous hits, including at an internal cryptic site previously identified by Wesam Alwan (Monash). Encouragingly, the results were consistent with a crystallographic screen of the same library done at 1 M.

SPR was highlighted by Nilshad Salim (ForteBio) and in a separate Biacore user day, and is an essential tool for off-rate screening (ORS). ORS facilitates screening of crude, unpurified reaction mixtures, since the off-rate of a compound bound to a protein is not dependent on compound concentration (see here). Compound purification is a major time-sink, and avoiding it is a key component of REFiL, or Rapid Elaboration of Fragments into Leads.

As Bradley Doak (Monash) discussed, REFiL entails the parallel synthesis of compound libraries around a selected fragment in 96-well plates using diverse reagents and high-yielding chemistries such as amide bond formation, alkylation, and reductive amination. Reaction mixtures are evaporated, resuspended in DMSO, and screened using ORS; this has led to affinity improvements of ten-fold or better compared with the original fragment for four projects tested thus far.

Beatrice Chiew (Monash) presented a case study against the oncology target 53BP1. Screening 1198 fragments led, after catalog-mining and rescreening, to 25 hits, all quite weak. Applying REFiL improved affinities by up to 15-fold, with the best molecules around 10 µM. Beatrice noted that because SPR provides “on-chip purification,” active compounds could be identified even when the reaction yields were less than 10%. She did note that examining the raw data (sensorgrams in SPR-speak) is important to recognize and avoid false positives.

Similarly, Luke Adams (Monash) applied REFiL to the bromodomain BRD3-ET. After two cycles, he was able to improve a 230 µM fragment to a 1.5 µM binder. Importantly, the off-rates were similar for the purified molecules and the crude reaction mixtures.

And Mathew Bentley (Monash) is exploring the potential of REFiL using crystallography, or REFiLX. This led to a 60 µM binder against the notoriously difficult EcDsbA. That affinity is more impressive given that the previous structure-based design and synthesis of more than 100 compounds – aided by 25 crystal structures – had failed to break 250 µM.

Vernalis pioneered off-rate screening, and Alba Macias described the company’s latest developments in this area. In the case of tankyrase, a 700 µM fragment was used to generate 80 compounds, which took one chemist a couple days. This yielded a 350 nM binder, the structure of which bound to the enzyme was solved using the crude reaction mixture for soaking.

Following up on this success, Vernalis is exploring the limits of crude reaction mixtures for high-throughput crystallography. Although promising, Alba noted caveats for the two proteins tested. Unlike off-rates, crystallographic success is dependent on compound concentration, so low-yielding reactions can lead to false negatives. And as anyone who has spent time working with fragments can attest, a beautiful co-crystal structure is no guarantee of high affinity, so false positives (ie, no improvement in affinity over the starting fragment) can be a problem too.

Alba also gave a brief summary of the discovery of S64315/MIK665, a fragment-derived MCL-1 inhibitor discovered by Vernalis, Servier, and Novartis that is currently in phase 1.

MCL-1 is a member of the BCL-2 of family proteins, and BCL-2 itself is targeted by the second fragment-derived drug to be approved. Guillaume Lessene (Walter & Eliza Hall Institute) spoke about both of these proteins, as well as BCL-x_L. Long-time readers may remember this selective BCL-x_L inhibitor, discovered using second-site NMR screening. Blocking this protein leads to platelet cell death, but AbbVie researchers are ingeniously side-stepping this liability by conjugating a related small molecule to an antibody to reduce systemic exposure. The resulting ABV-155 may be the first antibody drug conjugate derived from fragments, and was said to be in phase 1.

There was quite a bit more, though in the interest of time (and readers’ patience!) I’ll stop here. But I must note before closing that this meeting launched the Australian Research Council-funded Centre for Fragment-Based Design. This is in some ways an Antipodean version of FragNet, though with a longer (five-year) funding period and the opportunity to include a few postdocs as well as graduate students. If you’re interested, please contact them.

A new tool for detecting aggregation

Historically the most popular method for finding fragments has been ligand-detected NMR. Preliminary results of our current poll (to the right) suggest crystallography has pulled ahead. (Please do vote if you haven’t already done so.) However, NMR has many uses beyond finding fragments, as illustrated in a recent J. Med. Chem. paper by Sacha Larda, Steven LaPlante, and colleagues at INRS-Centre Armand-Frappier Santé Biotechnologie, NMX, and Harvard.

Among the many artifacts that can occur in screening for small molecules, one of the most insidious is aggregation. A distrubing number of small molecules form aggregates in water, and these aggregates give false positives in multiple assays. Unfortunately, determining whether aggregation is occurring is not always straightforward. The new paper provides a simple NMR-based tool to do just that.

All molecules tumble in solution, but small fragment- or drug-sized molecules tumble more rapidly than large molecules such as proteins. The “relaxation” of proton resonances is faster in slower tumbling molecules, and in the NMR experiment called spin-spin relaxation Carr-Purcell-Meiboom-Gill (T₂-CPMG) various delays are introduced and slower tumbling molecules show loss of resonances. Indeed, this technique has frequently been used in fragment screening: if a fragment binds to a protein, it will tumble more slowly, resulting in loss of signal.

The researchers recognized that an aggregate could behave like a large molecule, and they confirmed this to be the case for known aggregators, while non-aggregators did not. The experiment is relatively rapid (~30 seconds), and has been used to profile a 5000-compound library to remove aggregators.

One of the frustrations of aggregators is that it is currently impossible to predict whether a molecule will aggregate, and indeed, the researchers show several examples of closely related compounds in which one is an aggregator while the other is not. Even worse, the phenomenon can be buffer-dependent: the researchers show a fragment that aggregates in one buffer but not in another, even under the same pH.

Many fragment screens are done with pools of compounds, and the researchers find that molecules can show a “bad apple effect”, whereby previously well-behaved molecules appear to be recruited to aggregates.

The limit of detection for T₂-CPMG is said to be single-digit micromolar concentration of small molecule, though the researchers note that double- or triple-digit micromolar concentrations are more practical, which is more typical of fragment screens anyway. And some compounds may show rapid relaxation due to non-pathological mechanisms, such as tautomerization or various conformational changes.

Still, this approach seems like a powerful means to rapidly assess hits, and pre-screening a library makes sense. Another NMR technique using interligand nuclear Overhauser effect (ILOE) has also been used to test for aggregation, though not to my knowledge so systematically. For the NMR folks out there, which methods do you think are best to weed out aggregators?

Second harmonic generation (SHG) vs KRAS

Practical Fragments is currently running a poll on fragment-finding methods used by readers – please vote on the right-hand side. One biophysical method that perhaps we should have included is second harmonic generation (SHG). A recent paper in Proc. Nat. Acad. Sci. USA by Josh Salafsky, Frank McCormick, and collaborators at Biodesy, University of California San Francisco, and elsewhere describes the technique and its application to find fragments that bind to the oncogenic protein KRAS.

In SHG, two photons of the same energy are absorbed by a material which then emits a single photon with twice the energy. In the commercial instrument developed by Biodesy, a powerful 800 nm laser irradiates a dye, and the 400 nm photon it emits is detected. The intensity of the signal is exquisitely sensitive to the precise orientation of the dye. If a protein is labeled with an SHG-active dye and then immobilized on a glass surface, even subtle changes in conformation will be detected.

The researchers chose the G12D mutant form of KRAS, which is one of the most common variants and is associated with particularly aggressive tumors. They labeled the protein with a lysine-reactive SHG dye under conditions in which each protein would, on average, have one covalently-bound dye molecule (though some would have none and others would have more than one). Proteolysis and mass-spectrometry analysis revealed that the dye molecule labeled three different lysine residues, which the researchers viewed as a feature since a ligand causing a conformational change to any of the lysine residues would generate a signal. The researchers also demonstrated that the dye modification did not interfere with the ability of KRAS to bind to the RAS-binding domain of RAF.

Labeled KRAS was then immobilized and tested against several proteins known to bind it, including antibodies and the nucleotide exchange factor SOS. These produced SHG signals, presumably by causing conformational changes to KRAS, while non-binders such as tubulin did not.

Having established that the assay could detect binders, the researchers screened 2710 fragments at 250 and 500 µM, and obtained a whopping 490 hits. These were then triaged by screening at lower concentrations and performing dose-titrations, and 60 were then characterized by SPR.

Fragment 18, 4-(cyclopent-2-en-1-yl)phenol, showed binding by both SHG and SPR, and was further studied by 2-dimensional NMR (¹H-¹⁵N HSQC). This technique allowed measurement of the weak 3.3 mM dissociation constant. More importantly, it allowed the researchers to establish the binding location as being near the so-called “switch 2” region where SOS normally binds. This is the same region where a previous NMR screen had identified the slightly more potent fragment DCAI. The current paper confirmed that finding, though the researchers found evidence that DCAI may bind to other sites too. Docking studies using SILCS suggested that fragment 18 likely binds in a similar orientation as DCAI. Not surprising given the low affinity, the new fragment did not show functional activity in a biochemical screen.

SHG is an interesting approach, and the ability to rapidly assess protein conformational changes distinguishes it from other biophysical techniques. Site-specific labeling would produce more informative data on which regions of a protein move. However, I wonder if SHG is perhaps too sensitive, as evidenced by the large number of hits. Indeed, the researchers demonstrated that the promiscuous lipophilic amine mepazine also generated a strong SHG signal with KRAS. It would be interesting to do a head-to-head comparison with other similarly rapid techniques such as DSF or MST. Have you tried using SHG, and if so, how did it perform for you?