Focal molography for assessing protein-ligand interactions

The list of technologies used to find and validate fragments is long, with more than a dozen in our 2024 poll. But it’s always fun to learn about new approaches. At the Novalix conference earlier this year I learned of “focal molography,” and this is the topic of a recent paper in J. Med. Chem. by Nicholas Favalli, Andreas Frutiger, Sebastian Oehler, and colleagues at Philochem and lino Biotech.

 

The approach is described in detail in this Wikipedia article. Briefly, it is a biophysical technique based on changes in diffraction. A sensor chip contains a matrix of “molograms,” each of which is a lens constructed of micropatterned ridges and grooves. The ridges of each mologram are functionalized with unique single-stranded oligonucleotides. These capture complementary strands, each attached to a molecule to be screened. The grooves are coated with oligonucleotides without small molecules to act as a control in the subsequent readout. Proteins are then flowed over the chip. Protein binding to the small molecules of a specific mologram changes the topography of the ridge, and so the diffraction. This change can be measured in units of “coherent mass density” (pg/mm²). Because only specific binding to the ridges is detected (the grooves provide background controls for each mologram), the technique is suitable for even crude samples such as cell lysates.

 

An obvious application for focal molography is to confirm the binding of hits identified from DNA-encoded libraries (DEL), since the small molecules already have an oligonucleotide tag. This is the focus of the new paper.

 

The researchers started by assessing three known binders to the model protein carbonic anhydrase IX (CAIX). Surface plasmon resonance (SPR), the most frequently-used chip-based biophysical method, was used as a benchmark in two formats, one in which the small molecule was bound to the SPR chip, which is most comparable to focal molography, and one in which the protein was bound to the SPR chip.

 

SPR and focal molography agreed in rank order for the three compounds, but focal molography sometimes suggested tighter binding. For the highest-affinity compound, SPR gave a K_d of 28 nM for compound bound to the chip and 450-750 nM for protein bound to the chip, while focal molography gave a K_d of 0.82 nM. Agreement was better for the weaker binders, one of which was mid-nanomolar and the other low micromolar. The kinetic constants k_on and k_off also differed between focal molography and SPR by as much as two orders of magnitude.

 

Next, the researchers examined hits from a DEL screen against CAIX, some of which were selective against the related protein CAII. A total of two dozen molecules were tested by both focal molography and fluorescence polarization (FP) as an orthogonal technique. Again, the rank orders of affinities for the compounds to CAIX and CAII agreed while the absolute values varied, with focal molography usually suggesting ten-fold tighter binding. (Though the FP assay often bottomed out above the K_d values measured by focal molography.)

 

Finally, the researchers examined the binding of a known high-affinity binder to the oncology target fibroblast activation protein (FAP). In this case the measured affinities were close to the reported value.

 

This is an interesting approach, and according to the researchers “focal molography can support up to ~500 duplicate kinetic measurements per day on a single regenerable chip.” The observation that affinity measurements seem to be tighter than other methods does raise concerns. I wonder if the density of the molecules on the molograms leads to rebinding, as is sometimes seen when SPR chips are too heavily functionalized. Alternatively, since CAIX is a homodimer, perhaps avidity effects played a role. Also, the researchers suggest that the technique is best used for compounds with affinities better than 10 µM, which reduces utility for many fragment programs.

 

If you’ve used focal molography, how has it worked for you?

Surface Plasmon Resonance (SPR)

Fragment-based drug discovery took off with NMR in the 1990s and went mainstream with X-ray crystallography in the 2000s. Now surface plasmon resonance (SPR) is becoming increasingly popular as a primary means of identifying hits. The technique has been mentioned more than a dozen times on Practical Fragments, but we’ve never devoted an entire post to it until now.

This post follows up on two recent publications. The first is an excellent summary of SPR by our friends at FBDD-Lit. Peter Kenny gives an overview of the technique and reports on a workshop given by SPR mavens Dave Myszka and Rebecca Rich. He also covers some of the seminal papers in the field.

The second report is in the brand new journal ACS Medicinal Chemistry Letters. In it, Iva Navratilova and Andrew Hopkins of the University of Dundee provide practical advice on using SPR for fragment-screening.

The authors describe their work on using SPR to identify fragments that bind to carbonic anhydrase II, a popular target for proof-of-concept studies. They screened a library of 656 fragments with molecular weights between 94 to 341 Da, with an average of 187 Da, or about 13 non-hydrogen atoms. The entire screen, which was done at three concentrations (16.6, 50, and 150 micromolar) took 4 weeks from assay development to hit confirmation on a Biacore T100, and consumed a total of 27 micrograms of protein.

Importantly, Navratilova and Hopkins were keenly aware of the potential for false positives or nonspecific binders (of which there were 230 at the highest concentration!) One way they controlled for such artifacts was to include an unrelated reference protein; data could be corrected by subtracting the response to the reference protein from the response to the target protein. Another analytical method to reduce the number of false positives was to only consider compounds that exceeded a minimum threshold for ligand efficiency (a metric invented by Hopkins and co-workers), a decision justified here given the often high affinities observed for carbonic anhydrase inhibitors. After these filters, an examination of the stoichiometry of binding revealed a dozen specific binders and four non-specific binders, a hit rate of 1.8%.

My one reservation with this paper is that carbonic anhydrase is a particularly easy test case, unlikely to fairly represent many of targets that people screen. Indeed, the confirmed hits (all of which contain sulfonamides), have affinities from 0.13 to 14 micromolar – far better than a typical fragment screen, and comparable to many HTS screens. Still, the tools and analyses described should apply to more challenging targets.

Finally, it is worth noting that if you want access to SPR technology but don’t have the resources or expertise to do it yourself, at least a couple companies (Beactica and Graffinity) specialize in applying SPR to FBDD.

Enthalpy versus entropy

The earliest stages of lead discovery usually focus on obtaining a molecule with decent affinity for a given target. Affinity, or binding energy, can be dissected into two components: enthalpy and entropy. On a (very) simplistic level, enthalpic binding comes via specific molecular interactions, such as hydrogen bonds, while entropic binding results from nonspecific hydrophobic interactions. Optimizing enthalpy is usually more difficult than optimizing entropy: engineering a polar interaction requires more precision than adding a bit of grease. In a new paper in ChemMedChem, Andrew Scott and colleagues at Pfizer show how fragments that owe more of their binding affinity to enthalpy make better starting points for optimization than do fragments whose binding is more entropic, even if the entropic fragment is more potent.

The researchers used human carbonic anhydrase (hCA II), a venerable work-horse of biophysical studies. Benzenesulfonamide (compound 1, below) is a known binder, and the researchers studied the thermodynamics of 20 derivatives of this molecule using isothermal titration calorimetry (ITC), taking care to generate high-quality binding data. Adding a fluorine group to the 2-position of benzenesulfonamide (compound 2) improves the potency almost three-fold but lowers the ligand efficiency. In contrast, adding a fluorine to the 3-position (compound 3) improves the potency by seven-fold and also improves the ligand efficiency.



If you were choosing between these fragments solely on the basis of affinity or ligand efficiency, it would be reasonable to choose compound 3, and in fact a search of the literature turned up 15 carbonic anhydrase inhibitors that contained the 3-fluorobenzenesulfonamide substructure and none that contained the 2-flurobenzenesulfonamide substructure. However, a look at the thermodynamic parameters reveals that the affinity of compound 2 is driven by a sizable improvement in enthalpic binding, partially offset by lowered entropy. In contrast, compound 3 has a similar enthalpy of binding as compound 1 but increased entropy. What’s going on?

The researchers determined high-resolution crystal structures for all three of these molecules bound to hCA II. Interestingly, the structure of compound 2 shows a specific interaction between the fluorine atom and a main-chain NH of the protein. In compound 3, the fluorine points towards the hydrophobic wall of the protein.

Adding a 4-benzylamide substituent onto each of these molecules led to improvements in activity. However, this was a relatively modest boost for the more entropic compound 3 to compound 20, but considerably larger for the enthaplically-driven compound 2 to compound 19. Compound 19 shows a highly favorable binding enthalpy, and is the most potent and ligand-efficient of any of the three elaborated molecules.

Obtaining thermodynamic parameters for small-molecule protein interactions has historically been challenging, but in recent years miniaturization and improvements in technology have brought ITC into more non-specialist labs. If you have the resources, it may be worthwhile characterizing the thermodynamic profiles of your fragment hits, and – perhaps – looking more closely at those that show enthalpically-driven binding.

Fragments on Glass

I’m always a fan of new fragment technologies, and Hioryuki Osada and colleagues at RIKEN have just published a very intriguing one which they call a “fragment combination array,” or FCA.

The approach involves immobilizing fragments onto a specially prepared glass slide using a photogenerated carbene reaction; this can be done as an array of microscopic spots. Next, the glass slide is treated with a fluorescently labeled protein and washed. If the protein sticks to the small molecule, it will show up as a fluorescent spot. Appealingly, if the protein of interest is genetically fused to a fluorescent protein, crude cell lysates can be used, simplifying the assay. The researchers previously demonstrated that natural products and drugs could successfully be immobilized to a treated glass chip using this method, and that the molecules retained their ability to bind to protein targets, despite the covalent linkage to the chip.

Of course, binding interactions between drugs and their targets are generally much stronger than between fragments and their targets. Also, because fragments are so small, there is a higher probability that the part of the fragment used to attach to the glass will be critical to but inaccessible for binding. Fortunately, the carbene chemistry is fairly non-selective, inserting into C-H and O-H bonds at random; thus, the likelihood is that at least some of the fragments will bind in a productive fashion. The technique is conceptually similar to the SPR-based methodology used by Graffinity, though to my knowledge Graffinity screens individual fragments as opposed to binary pairs.

But does it actually work? An initial proof-of-concept used FKBP12 ligands, the same ones previously described in the first famous “SAR by NMR” paper. In that example, a low micromolar pipecolinic acid derivative was linked to a high micromolar benzanilide derivative to generate a nanomolar binder to FKBP12. In the current case, the immobilized pipecolinic acid derivative was able to capture fluorescently-labeled FKBP12, while the benzanilide was not. However, co-spotting the two fragments led to a much stronger signal (more fluorescence) than the pipecolinic acid spot alone, suggesting synergy between the two immobilized fragments.

Having shown this, the researchers next turned to the protein carbonic anhydrase II (CAII), which has a predilection for sulfonamides. They created an array from four aromatic sulfonamide-containing fragments (and one negative control) and ten diverse non-sulfonamide-containing fragments. A screen of these 50 different mixtures against fluorescently-labeled CAII revealed a number of hits, and by merging elements of one of the diverse compounds onto a sulfonamide “anchor” fragment, the researchers were able to improve the potency of the sulfonamide from 435 nM to 29 nM.

Of course, there are limits: clearly the technique is not as sensitive as many other fragment-detection methods, as illustrated by the inability to detect benzanilide binding to FKBP12, an interaction with a Kd in the mid to high micromolar range. In fact, both test cases involve protein targets that have been shown to be highly amenable to fragment-based methods, and both start with known fragments with relatively high affinities. Moreover, the covalent immobilization methodology won’t work for all fragments; acetazolamide, a fragment-like high-affinity binder of CAII, didn’t work in this assay, likely due to poor geometry or sterics of the immobilized fragment.

Still, FCA is a neat and potentially very rapid method for finding a second fragment once a first has been identified. It will be fun to watch how the technique evolves.