NMR Methods for Identification of False Positives in Biochemical

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NMR Methods for Identification of False Positives in Biochemical Screens Miniperspective Anamarija Zega* Faculty of Pharmacy, University of Ljubljana, Aškerčeva 7, 1000 Ljubljana, Slovenia ABSTRACT: Over the past few decades, NMR spectroscopy has become an established tool in drug discovery. This communication will highlight the potential of NMR spectroscopy as a method for identification of problematic compounds and as a valuable aid toward revealing some mechanisms of promiscuous behavior. NMR methods for detecting false positives will be analyzed on the basis of their performance, strengths, limitations, and potential pitfalls. Additionally, this communication aims to provide an insight into the limitations of NMR-based methodologies applied to ligand screening in the context of false-positive hits.

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these filters still do not identify all interference compounds, and some “good actors” may slip through and cause problems. One of the most insidious problems in the early stage of drug design is aggregation, as reported by the Shoichet group in 2002.2 Several small molecules can form microscopic aggregates in aqueous buffers, and these aggregates can inhibit diverse drug targets nonspecifically.20−24 Aggregation can occur with a variety of molecules, including fragments. Indeed, fragmentbased drug discovery (FBDD) is most susceptible to this phenomenon, as in fragment screening, high concentrations are necessary for the detection of binding.25,26 To eliminate active molecules that act via aggregation, the use of detergents to disrupt aggregates is now common in screening campaigns. However, detergents are often not tolerated in assays. Additionally, detergents might not fully eliminate aggregation, as they usually only promote rightward shifts in concentration− response curves.20−24 The potential of NMR spectroscopy to be used to reveal information about molecular interactions at the atomic level makes this a particularly effective tool for insight into processes in screening mixture solutions and consequently for the differentiation between specific and nonspecific target binding. The simplest NMR parameter, the chemical shift, is highly sensitive to the precise environment of the atom, and it can reveal whether a hit molecule binds to the target and what parts of the molecule are involved in any interaction.10,12 Additionally, NMR techniques, such as saturation transfer difference (STD), are sensitive to the overall motion of the test compound, which is very different for free and bound ligands.12,27

INTRODUCTION Beginning around 20 years ago, medicinal chemists started to note that high-throughput screening hit lists are plagued by artifacts or promiscuous bioactive compounds.1−3 Since then, multiple sources for promiscuous behavior have been described.4 Together with the growing awareness of problematic molecules and studies of promiscuous mechanisms, software filters5−8 and biochemical methods to identify false positives started to emerge.9 NMR spectroscopy is among the biophysical tools that can be used to address this issue. Nuclear magnetic resonance spectroscopy is a powerful method for probing molecular interactions. It can be used to detect protein−ligand interactions across a large range of affinities, and it is widely used in drug discovery to identify hits in screening of compound libraries.10−12 As NMR can be used to detect very weak interactions, it helped pioneer the entire field of fragment-based drug discovery (FBDD), and today it still remains one of the major fragment-screening technologies.13−18 In the early 2000s, researchers started to realize that NMR can be used not just to screen compound libraries but also to validate screening hits.12,19 Here, NMR spectroscopy became important among the methods used to reveal promiscuous behaviors of hits and to explain promiscuous mechanisms. Compounds that aggregate or react are the most prevalent type of promiscuous “pollutants” in the chemical literature. Since the review of Rishton in 1997,3 awareness of potentially reactive functionalities that have to be avoided in screening libraries that are not intended for the design of covalent drugs has grown and more recently particularly as a result of the pan assay interference compounds (PAINS) initiative.4,6 New promiscuous structural classes have been discovered and included in readily available filtering software. However, © 2017 American Chemical Society

Received: October 14, 2016 Published: June 28, 2017 9437

DOI: 10.1021/acs.jmedchem.6b01520 J. Med. Chem. 2017, 60, 9437−9447

Journal of Medicinal Chemistry

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reference molecule are acquired. Nonspecific interactions between the reference molecule and the hit molecule aggregates result in partial transfer of the water magnetization stored by the aggregates to the reference molecule. This transfer is then detected in the chemical shift of the free reference molecule, as positive NMR signals. Conversely, in the absence of aggregates, the WaterLOGSY signal is negative.28 In the example of a SPAM experiment shown in Figure 1, aggregation of the known aggregator quercetin (1) was

This review will highlight both well-established methods and new advances in the use of NMR spectroscopy for the identification of problematic compounds while also providing examples of its use. Methods for detecting false positives will be analyzed on the basis of their performance, strengths, limitations, and potential pitfalls. Additionally, this communication is designed to provide some insights into the limitations of NMR-based methodologies as they are currently applied to drug design. For the medicinal chemist, this review is meant to provide a practical review of the area and allow more effective interpretation, quality control, and trouble-shooting of highthroughput assay results as well as facilitating a more informed collaboration with physical/structural chemistry groups.

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AGGREGATION Promiscuous inhibition caused by small-molecule aggregation is a complicated process, and as such it still remains one of the major sources of false-positive results. When a colloidal aggregate is formed, proteins adsorb to its surface and are partially denaturated, which can lead to nonspecific inhibition. A colloid will form reproducibly under specific conditions of buffer composition, temperature, and concentration, and its promiscuous inhibition will depend on the components of the buffer and the concentration of the protein.2,7,20−24 To address this challenge, Dalvit and co-workers (2006)28 proposed a set of NMR techniques that they referred to in terms of the “solubility, purity, and aggregation of the molecule” (SPAM), which enabled rapid characterization of the identity, purity, solubility, and aggregation state of a hit compound. This method requires acquisition of the onedimensional (1D) reference spectrum, a “water ligand observed via gradient spectroscopy” (WaterLOGSY) spectrum, and/or the selective longitudinal relaxation filter or transverse relaxation filter spectrum for the identified hit in aqueous solution and in the presence of a water-soluble reference molecule. WaterLOGSY is a commonly used 1D ligand-observed method for detection of molecules that interact with proteins.29−31 It is based on nuclear Overhauser effect spectroscopy (NOESY) and on intermolecular magnetization transfer via the bulk water. The inverted water magnetization can be transferred to the bound ligand via different pathways, where the ligand interacts with water via water−ligand−protein or protein−ligand complexes, the rotational correlation times of which yield negative cross-relaxation rates and show negative NOEs with water. In contrast, small molecules that only interact with the bulk water (i.e., nonbinders) undergo much more rapid tumbling, which translates into positive NOEs. Therefore, opposite signs for the signals from free versus protein-bound ligands are observed in the WaterLOGSY spectrum, which allows easy discrimination between binders and nonbinders.29−31 By use of the WaterLOGSY technique, compounds that aggregate will give responses similar to those seen in the presence of protein binding, as aggregates contain trapped water molecules or water is involved in the formation of hydrogen bonds. Selective excitation of the bulk water magnetization in the case of aggregator molecules results in magnetization transfer to the aggregate protons via chemical exchange with the labile protons of the aggregate and via bound, although exchangeable, water molecules.28 In SPAM experiments, WaterLOGSY spectra of the hit compound in the presence of a water-soluble nonaggregating

Figure 1. (A) Chemical structures of quercetin 1 and reference compound 2. (B) WaterLOGSY of reference compound 2. (C) WaterLOGSY of reference compound 2 in the presence of 1. (D) Expanded region of the 1H NMR spectrum of compounds 1 and reference compound 2. Adapted with permission from Current Drug Discovery Technologies (Dalvit et al.).28 Copyright 2006 Bentham Science Publishers.

detected. Figure 1D shows the expanded aliphatic region of the 1D proton spectrum of 1 in the presence of reference compound 2, where only the signals for dimethylsulfoxide (DMSO) and the methyl group of reference compound 2 can be seen. The WaterLOGSY spectrum of reference compound 2 (Figure 1B) shows a weak negative effect for the CH3 signal, which is typical of hydration of a small nonaggregating molecule. In the presence of 1, the CH3 signal changes sign and becomes positive (Figure 1C), which is an indication of aggregation, whereas the NMR signal of DMSO, which does not interact with the aggregate, still remains negative.28 La Plante et al.32,33 introduced an assay for the direct monitoring of molecular aggregation in aqueous media. In aqueous solutions, compounds adopt equilibria between three states that can be distinguished in the 1H NMR spectra: soluble single molecules, soluble aggregate entities, and the solid form. Solid precipitates tumble too slowly to be observed by solution NMR. Soluble druglike compounds that behave as fasttumbling single molecules result in sharp 1H resonances. The 1 H spectra of soluble compounds that can self-aggregate typically show unusual features due to the attractive tendencies of the aggregator compounds at higher concentrations which dissociate upon dilution. For detection of aggregates, La Plante et al.32,33 proposed a strategy that involves the acquisition and monitoring of 1H NMR spectra as a function of the compound 9438



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Figure 2. NMR dilution assays involve the acquisition and monitoring of 1H NMR spectra as a function of the compound concentration: (A) nonaggregator behavior in solution as a function of dilution; (B) unusual NMR spectra of an aggregator upon dilution. Adapted from Journal of Medicinal Chemistry, LaPlante et al., 2013.33

traditional functional group filters that are designed to capture problematic molecules.34 Many biologically relevant nucleophiles are thiols, such as glutathione, coenzyme A, and protein cysteines. The nucleophilicity of protein cysteine thiols can have significantly different reactivity profiles compared to small-molecule thiols, including glutathione, cysteamine, and coenzyme A, which are usually used to test compounds for chemical reactivity. Therefore, it is possible that some compounds might not form adducts with small-molecule probes but would still react with protein cysteine residues.34,35 To detect protein-reactive compounds in high-throughput screening (HTS) campaigns, Hajduk and co-workers designed an NMR assay that they called the La assay to detect reactive molecules by NMR (i.e., ALARM NMR), which measures whether a compound covalently modifies cysteines in the human La antigen (Figure 4).36,37 In the search for new antiviral agents, Huth et al. (2005)36 performed NMR-based screening of a La human antigen clone labeled with 13C that yielded high-resolution heteronuclear single quantum correlation (HSQC) spectra. They noted that the La antigen had an exceptionally high frequency of reaction with a variety of small compounds; however, their NMR studies of screening hits in the presence of the reducing agent dithiothreitol (DTT) did not yield a single valid hit. Huth et al. (2005)36 concluded that the protein clearly lacked a pocket that was suitable for small molecule inhibitors. However, as the La antigen contains two cysteines, the shift changes and line broadening in the 2D NMR spectra might have been due to covalent modification of cysteine, and the addition of DTT to the sample buffer prevented this covalent modification, as electrophilic compounds should react with the excess DTT and protect the protein cysteine residues. They proposed that by measuring the chemical shift changes of the 13C-labeled La protein in the presence of the test compound in both the absence and presence of DTT, the nonspecific reactivity of hits can be detected. ALARM NMR assays have been extensively validated,

concentration. Nonaggregating compounds will be expected to give sharp NMR resonances at all concentrations, and there should be no changes in the numbers and shapes of their resonances, which will also not shift left or right as the compound is diluted. Finally, the peak intensities correlate with the changes in concentration (Figure 2). On the other hand, 1H NMR spectra of aggregators can show broad, or both broad and sharp, resonances (i.e., dependent on the size of the aggregate), changes in resonance number (i.e., number of entities), resonances that shift left or right (i.e., due to molecular proximity), and intensities that do not correlate with the dilutions (i.e., due to concentration changes) (Figures 2 and 3).33 As the resonances of very large aggregates might be too broad to observe, La Plante et al. (2013)33 upgraded their system to propose the use of the detergent Tween 80 to break the aggregates up into smaller entities that should be observable by NMR. To avoid potential effects of the detergent on compound solubility, the additional step of mild centrifugation was included in the procedure to separate out any solid compound (if there is any) before detergent addition. In conclusion, following the detailed procedure that La Plante et al.32,33 described, the method is simple to carry out on equipment that is routinely available in medicinal chemistry laboratories, and it serves as a practical tool for basic hit evaluation and detection of aggregators.

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REACTIVE COMPOUNDS Reactive hits can promote chemical modifications of reactive protein residues, or they can chemically react with nucleophilic residues of assay reagents; this can confound the assay readout by producing what might be construed as biological activity. Typical reactive moieties such as aldehydes and acyl halides are suggestive of reactivity and can be removed from screening campaigns without further consideration. However, for some compounds, it is not always obvious that they will be prone to react with nucleophiles; hence, such compounds can escape 9439



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Figure 4. (A) Expanded region of a 2D 1H−13C HSQC spectra showing cross-peaks for four methyl groups in the human La antigen in the absence (red) and presence (black) of thiol-reactive compound 2-chloro-1,4-naphthoquinone (7) without DTT. (B) The same spectra for samples to which DTT has been added. (C) Structure of the Cterminal recognition motif domain of the human La antigen protein where the Cys and Leu residues for which the methyl chemical shifts are shown in (A) and (B) are indicated. Adapted from Journal of the American Chemical Society, Huth et al., 2005.36

identify inhibitors of Rtt109.39 They screened approximately 225K small molecules that were filtered computationally using filters for pan-assay interference compounds at the beginning of their triage.39 As one of three compounds that passed their initial HTS triage using substructural filters to remove panassay interference compounds and also several counter screens, their compound 8 was selected for more in-depth studies of its potential promiscuous mechanism. Among other triage tools, ALARM NMR was performed and compound 8 perturbed the La protein conformation, as shown by signal decreases relative to DMSO controls (Figure 5). The inclusion of DTT in the assay buffer did not attenuate the line broadening for compound 8, which is consistent with a non-thiol-reactive mechanism of action. This finding supports another (simultaneous) benefit of ALARM NMR use in HTS triages, as it can help to identify compounds that appear to nonspecifically modulate the conformation of a protein that is completely unrelated to the target, the La antigen.39 The ALARM NMR assay has several benefits. First, cysteines in a proteinaceous environment can have significantly different reactivity profiles than as a small molecule. As the La antigen is a protein, it represents a good surrogate for protein targets used in drug screening. This method is sensitive, it has medium throughput, and it is robust, as NMR spectra of the La protein are stable to air oxidation after several days at room temperature. Throughput can be increased with the use of an autosampler, a cryoprobe, efficient 13C enrichment, and an optimized pulse sequence. However, even if this method has high reliability and is relatively easy to perform, it requires the expense of labeled amino acid precursors and comparatively higher concentrations of proteins and reagents. Also, its sensitivity and how to best quantify the data (especially for equivocal findings) represent unanswered questions.34,36 Avonto et al. (2011)40 described a simple 1H NMR method for identification of Michael acceptors that used cysteamine as a probe and proposed it as prescreening for more complex ALARM NMR. They initially applied the method to identify

Figure 3. Schematic representation of the spectra of the nonaggregating compound riluzole (3) (A) in comparison with the unusual NMR spectra of aggregators methylene blue (4) (B), Evans blue (5) (C), and pranlukast (6) (D) upon dilution. Adapted from Journal of Medicinal Chemistry, LaPlante et al., 2013.33

and ALARM NMR is now an established NMR method for the detection of false positives.36 Recently, two post-HTS case analyses were reported by Walters and colleagues38,39 that support the use of ALARM NMR in the HTS evaluation process. In the first study they applied ALARM NMR to confirm promiscuous mechanisms of thiol reactive chemotypes that they encountered in a HTS campaign against the epigenetic target histone acetyltransferase Rtt109.38 ALARM NMR, consistent with other methods that they used, detected thiol reactive compounds indicated by evidence of the expected shift changes in the absence of DTT while being prevented in the presence of DTT.38 A second case report described the follow-up of a 4-aroyl-1,5-disubstituted-3hydroxy-2H-pyrol-2-one active compound from HTS to 9440



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Figure 6. 1H NMR spectra of the representative thiol-reactive 5benzylidene barbiturate in DMSO-d6 before (compound 10) (A) and after the addition of cysteamine probe (compound 11) (B). The NMR signal of the vinyl proton of 5-benzylidene barbiturate, which disappears during the course of the reaction, is marked with an arrow. Adapted from Arsovska et al., 2014 (Acta Chimica Slovenica).42 Figure 5. Two-dimensional 1H−13C HMQC spectra of selected 13Clabeled methyl groups for compound 8, as tested using ALARM NMR. The compounds were incubated with the La protein in either the absence (red) or presence (blue) of DTT. Compound 7 and fluconazole are shown as positive and negative controls, respectively. A scaled-up view of compound 8 is shown (“4 zoom”). Note that the spectra without and with DTT are nearly identical. Compound 9, which is an inactive analog, was also included as a negative control. Signals were normalized to DMSO controls. Reproduced from Bioorganic & Medicinal Chemistry Letters (http://www.sciencedirect. com/science/journal/0960894X?sdc=1), Vol. 25 (Dahlin, J. L.; Nissink, J. W.; Francis, S.; Strasser, J. M.; John, K.; Zhang, Z.; Walters, M. A. Post-HTS case report and structural alert: promiscuous 4-aroyl-1,5-disubstituted-3-hydroxy-2H-pyrrol-2-one actives verified by ALARM NMR, pp 4740−4752),39 Copyright 2015, with permission from Elsevier.

intensity were exhibited in the reactions with thiohydantoins, hydantoins, and thiazolidine-2,4-diones, which suggests that the distinct binding profile of these scaffolds is not related to reactivity. While the limitations of using cysteamine as a surrogate for protein thiols have been discussed above and cannot categorically exclude the possibility of reactions occurring with proteins, this simple 1H NMR method does provide a rapid and clear assessment of the potential reactivity of compounds with protein thiols. Finally, since the chemist is provided with exact location of the reaction site, monitoring for more than one Michael-type acceptor function in the same molecule is also possible.

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MISLEADING ASPECTS OF NMR It is also necessary to be aware that NMR itself is prone to produce false-positive results that are mostly related to aggregation. This problem is most pronounced in fragmentbased drug discovery (FBDD), where among the NMR-based fragment screening methods, saturation transfer difference (STD) is the most commonly used. With STD, the protein target is irradiated and transfers some of its magnetization to its bound ligands. When these dissociate, they retain some of the magnetization, and so the NMR signals of the ligands decrease. This ligand-based technique is easy to implement, does not require labeled protein, is amenable to high molecular weight targets, and is very sensitive, and thus it can be used to identify weak fragments. Unfortunately, the method has one major drawback: the technique identifies both specific and nonspecific binders, due to its accumulation of binding signals from multiple binding sites.44−47 Consequently, after a STD-based screening campaign, careful control experiments are required to discriminate between specific and nonspecific binders, for example, by repeating the experiments in the presence of a competitor ligand of known binding mode. However, even when such appropriate competitive references are available, displacement of the ligand by a competitor does not systematically imply that the ligand binds with a privileged binding mode. When such competitive reference compounds are not available, an orthogonal

Michael acceptor sites in complex multifunctional compounds and to sort them into reversible and irreversible thiol sinks. In the assay in DMSO, cysteamine reacts with electrophilic double bonds to provide Michael adducts, whereas no trapping reaction takes place in apolar solvents. It is important to note that cysteamine is a simple molecule and cannot be used as a direct surrogate for all protein thiol targets. However, with a pKa of 8.3, cysteamine has similar reactivity to many surface thiols of proteins, and to some extent it can be used as an approximation of thiol reactivity for screening purposes.41 Arsovska et al. (2014)42 applied the Avonto NMR method40 to the evaluation of Michael-type acceptor reactivity of 5benzylidene barbiturates, 5-benzylidene rhodanines, and related heterocycles that have gained a reputation as problematic promiscuous scaffolds. These compounds were treated with 1 molar equivalent of cysteamine in DMSO-d6 as a solvent, and their reactivity was estimated via monitoring 1H NMR signal intensity of the vinyl proton. Addition of cysteamine to known Michael acceptors 5-benzylidene barbiturates resulted in instantaneous elimination of the 1H NMR signal for the vinyl proton (Figure 6). This was then compared to the behavior of 5-benzylidene rhodanines under identical conditions. Disappearance of the 1H NMR signal in this case was relatively slow and incomplete. Similar low reaction rates and incomplete reaction behavior as interpreted from the 1H NMR signal 9441



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biophysical technique is required to validate a fragment hit.43−47 To overcome potential false STD hits due to aggregation, Vom et al.48 proposed recording the STD in the presence of a detergent. In their case study, a fragment screening campaign of a commercially available fragment library (of 500 compounds) was performed against ketopantoate reductase, a potential target for antimicrobial agents. Hits (as 196 out of 500 fragments) were validated by recording 1H/15N HSQC NMR spectra and by biochemical assays. By performing a dynamic light scattering (DLS) experiment and carrying out the biochemical assays in the presence of the detergent, they demonstrated that aggregation was responsible for at least two of the most potent hits. The initial library was screened again in the presence of Tween-20 (0.01%) and a lower fragment concentration (from 1.0 to 0.3 mM), which gave 71 hits that were also followed up by 1H/15N HSQC NMR (samples with no added detergent). The HSQC perturbations that clustered around the active site gave the final evidence that the resulting compounds were not aggregation-based false positives. However, as mentioned before, detergent does not always fully eliminate aggregation. Cala and Crim (2015)47 recently reported the use of STDbased epitope mapping to discriminate between specific and nonspecific ligands without competition experiments, through the identification of fragments that showed a privileged orientation upon interaction with the protein target. The basis for STD epitope mapping is that within a ligand that binds to a protein, a proton that binds closer to the protein will show a stronger STD effect than one that is exposed to the solvent.49 These data allow the rough modeling of how a ligand binds to a protein, or at least which parts of a ligand are closest to the protein. Cala and Crim (2015)47 argued that simply observing differences in STD effects between different protons in a ligand can distinguish whether or not that ligand is binding through a single binding mode. They performed screening of 500 fragments against the glycogen phosphorylase protein using STD. To discriminate between specific and nonspecific binders, they measured the STD effects (ISTD/I0) for the protons of each ligand. All of the fragments that showed significant epitope mapping (i.e., significant STD effect variations) could be competed with a known reference molecule, and this was less often the case for fragments that did not show epitope mapping (Figure 7). When a ligand binds to a binding site without any preferred binding mode, this will translate into STD effects that are similar for all ligand protons, which suggests nonspecific binding. As Cala and Crim (2015)47 noted, there are limitations. For example, if a fragment can bind in two different, but nonetheless specific, binding modes, it might show uniform STD effects and thus be a false negative. Here, the technique was relatively robust across a large range of experimental conditions, including the affinity of the fragments (KD of 50 μM to >1 mM), the saturation time (from 0.5 to 4.0 s), and the ligand/protein ratios (as 66 to 1, and 400 to 1). In addition, Cala and Crim (2015)47 showed that this allowed differentiation between hydrophobic and polar driven binding interactions. They performed and compared spectra of the privileged fragment 15 (Figure 8), 7-azaindole, in the presence of two different targets, glycogen phosphorylase and ERK1 kinase. The compound was shown to bind to the two different proteins with different epitope maps, which suggested different (but specific) binding modes for each protein (Figure 8).47

Figure 7. Spectra of three fragments (12, 13, 14) binding to the glycogen phosphorylase protein, showing the STD effects for each proton. Only fragment 12 (A) showed significant STD effect variations (>10%), indicating that it binds the protein target with a particular orientation. Adapted from Journal of Medicinal Chemistry, Cala et al., 2015.47

Figure 8. STD spectra of fragment 15 showing the binding driven by polar and hydrophobic interactions. The 1D (STD-off) spectrum of fragment 15 is shown in black. The STD spectrum in the presence of glycogen phosphorylase is shown in blue. The STD spectrum in the presence of ERK1 kinase is shown in red. STD effects for the ligand protons are shown, which demonstrate that fragment 15 bound with preferred orientations to both proteins and that the binding was driven by polar interactions only in the case of the kinase. Adapted from Journal of Medicinal Chemistry, Cala et al., 2015.47

Nonetheless, comparing STD effects across a ligand allows direct identification of specific binders based on the fragment showing specific molecular recognition toward the protein target.47 Another NMR technique that is prone to produce false positives and is also used in FBDD is the interligand nuclear Overhauser effect, or ILOE.50−53 This 2D NOESY NMR technique is used to detect when two small-molecule ligands bind simultaneously and with similar residence times in adjacent sites on a protein surface. A signal, strong negative 9442



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Figure 9. Typical example of 2D NOESY spectrum representing nonspecific binding: NOESY spectrum of compounds 16 (5-methoxyindole) and 17 (2-carboxybenzofuranoic acid) in perdeuterated pantothenate synthetase (dPtS), where negative peaks are colored in green and positive peaks are colored in blue. ILOE peaks between ligands 16 and 17 were observed; however, uniform negative peaks (green) between all of the possible combinations of protons were also seen, which indicated nonspecific binding. Adapted from Journal of the American Chemical Society, Sledz et al., 2010).54

the free ligand. If two ligands bind simultaneously in adjacent sites on the protein surface, forming a ternary complex, strong negative ligand−ligand NOEs (ILOEs) will also be observed.55 Abel and colleagues54 detected interligand NOEs between fragments 16 and 17; however, the signals were seen between all the protons of the fragment 16 and all the protons of the fragment 17, which suggested nonspecific binding to the protein (Figure 9). If the two fragments were to bind in a specific orientation and in close proximity to one another, one would expect that some protons from 16 would be closer to some protons in 17 than others, and thus there would be differences in signal intensities. Addition of the methyl group to 5-methoxyindole 16 to give compound 18 did not help to solve the problem, as ILOE signals were seen in the spectra from both the methyl groups of the new compound to all of the aromatic protons of 17 (Figure 10).54,17 Because it was felt that the hydrophobic nature of compound 18 could be causing the aggregation observed at high concentrations, Abell and colleagues54 modified this fragment by adding an N-CH2COOH group to give compound 19 (Figure 11). NMR experiments with compound 19 in the presence of 2-carboxybenzofuranoic acid and the protein then revealed specific ILOE signals between the 2-methyl group of compound 19 and H2 of 2-carboxybenzofuranoic acid 17.54,17 The fragments were then linked to give compound 20 (Scheme 1), which bound to the enzyme 3 orders of magnitude more tightly than each of the starting fragments. This compound was also well-behaved mechanistically and showed competitive inhibition with ATP, while the crystal structure revealed that it binds as expected given the structures of the individual fragments. ILOE-driven step-by-step reasoning was

ligand−ligand NOE will be generated only when ligands bind in relatively close proximity to each other, which allows the identification of fragments that could be productively linked together. Additionally, the method enables determination of the ligand binding orientations relative to each other. This technique has several benefits: as mentioned before for STD, ILOE can also be applied even in the complete absence of a 3D protein structure, and it can be used with proteins that are too large to study by other NMR methods. However, because of their potential aggregation and nonspecific binding, the application of this method to screening high concentrations of relatively hydrophobic fragments is more prone to artifacts. Abell and colleagues (2010)54,17 applied the ILOE approach to FBDD of inhibitors against pantothenate synthetase (PtS) from Mycobacterium tuberculosis. Here 5-methoxyindole (Figure 9, 16) and 2-carboxybenzofuranoic acid (Figure 9, 17) had been identified from fragment screening. WaterLOGSY and STD competition with known ligands revealed that the binding of both fragments was competitive with ATP and that binding of the fragment 17 was also competitive with the substrate pantoate, whereas competition between fragment 16 and fragment 17 was not observed. Furthermore, an attempt was made to use NMR to determine the relative binding modes of 16 and 17, and because the enzyme was too large for direct protein observation by NMR, ILOE was attempted. During a NOESY experiment, small molecules tumble rapidly in solution, and the dominant 1H relaxation mechanisms lead to weak positive NOEs. In contrast, when a small molecule is bound transiently to a target protein, it has long correlation time. In the case of a fast exchange between free and bound state (weak binders in the μM to mM range), this translates into very strong negative NOEs in the 2D NOESY spectrum of 9443



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ular mass and drives the τc upward, which could give a false positive result.56,57 Protein-observed techniques (such as 1 H−15N HSQC, 1H−13C HSQC, 1D protein-observed fluorine NMR (PrOF)) are inherently less prone to false positives.58,59 Because there are NMR resonances in a specific pattern due to the local chemical environment of each observed nucleus, the spectrum will reveal whether or not the protein is in solution and properly folded.59 If the protein is aggregated or precipitated, the resonances coalesce, disappear, or sharpen, so protein resonances provide an in situ quality control reducing potential false positives.58,59 It is worth noting here that besides problems that can occur with compounds and target proteins, each individual screening NMR method has its own collection of methodological artifacts that can result in false positives and false negatives. Gossert and Janhke recently published a practical guide58 to identifying and validating protein−ligand interactions using NMR, and in this review they also address some aspects of the identification and control of these methodological artifacts. Besides NMR, other established biophysical techniques are currently used to monitor the binding of a ligand to a target, such as X-ray crystallography, surface plasmon resonance (SPR), thermal shift analysis (TSA), isothermal titration calorimetry (ITC), and fluorescence polarization (FP).60 Several comparative studies of biophysical techniques have been made particularly on the field of FBDD where ligandobserved NMR was confirmed as a reliable screening technique.61−64 For example, Hubbard and Murray summarized the 10 years’ experience at Vernalis in applying different biophysical techniques for identification of fragments across a range of different targets. Among the three techniques used for initial screening, SPR, differential scanning fluorimetry (DSF), and ligand-observed NMR, the last emerged as the most robust technique for reliably identifying fragments that bind to a range of proteins.61 In a study by Meiby et al., a number of biophysical techniques, SPR, TSA, FP, WAC (weak affinity chromatography), and a consensus of three ligand observed NMR experiments (STD, WaterLOGSY, and Carr−Purcell−Meiboom−Gill (CPMG) echo train acquisition), were compared against each other using a carefully curated library on a wellknown tractable protein target. A subset of hits from these techniques was taken forward into lower throughput techniques, ITC and X-ray crystallography, for “validation” of

Figure 10. Part of the 2D NOESY spectrum of fragments 17 (2carboxybenzofuranoic acid) and 18 in dPtS. The ILOE peaks are marked with red arrows. Adapted from Journal of the American Chemical Society, Sledz et al., 2010.54

Figure 11. Part of the 2D NOESY spectrum with fragments 17 and 19 in dPtS. The ILOE peak is marked with a red circle. Adapted from Journal of the American Chemical Society, Sledz et al., 2010.54

successfully applied here overall; however, this example suggests that the ILOE method should be carefully controlled for aggregation artifacts.54,17 To confirm the existence of ILOEs and avoid artifacts from spin diffusion, compound aggregation, or nonspecific interactions, ILOEs should be observed in experiments performed with perdeuterated protein.55 Additionally, to prove that the detected ILOEs are target dependent and therefore indicators of true second-site binding, experiments should always be repeated in the absence of target protein.26 Other screening NMR methods, especially ligand-observed, are not immune to false positives due to nonspecific binding and aggregation. In most ligand-observed NMR experiments, slowing of the molecular motion of the ligand, which is revealed with increasing effective overall rotational correlation time τc, is used as an indication of ligand binding.55 As with ligand binding, compound aggregation increases the effective molec-

Scheme 1. Fragment Linking Leading to Potent Inhibitor 20 of M. tuberculosis PtS, Using ILOE-Driven Step-by-Step Reasoning54,17

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Table 1. Summary of Parameters of the NMR Methods Used for Detection of False Positives type of promiscuity aggregators

NMR experiment SPAM: WaterLOGSY; ref molecule needed (e.g., pteridin derivative)

advantages/strengths of the method

limitations (weakness)

NMR detection

ref

-medium throughput

500 MHz + cryoprobe

Dalvit et al.28

-medium throughput (relatively complicated sample preparation)

400 MHz

LaPlante et al.32,33

-proteinaceous environment

-13C labeled protein required

500 MHz + cryoprobe

Huth et al.;36,37 Dahlin et al.38,39

-probe stable to air oxidation

-robust

-2D NMR -high concn of protein and compounds -medium throughput

-simple experiment

-probe prone to oxidation

400 MHz

Avonto et al.;40 Arsovska et al.42

-high throughput -inexpensive

-nonproteinaceous environment

-inexpensive -measurement under the same exptl conditions as the screening campaign -robust -allows identification of false negatives

LaPlante’s method -simple NMR experiment -inexpensive thiol reactive compounds

ALARM NMR: 2D HSQC; La protein probe

-extensively validated method Avonto’s method: 1D NMR; cysteamine probe

NMR screening methods in FBDD: STD and ILOE. Therefore, researchers should aim to perform these assays with an awareness that artifacts can and do occur. When doubtful results emerge from these methods, there is the need to implement multiple, in some cases redundant, investigations to decrease the risk of nonspecific hits.

the binding observed in the primary technique. With 93% of 28 hits from the ligand-observed NMR screening yielding X-ray structures, NMR was the most effective predictor of crystallography success, which is still considered the gold standard for progressing ligands in the drug design process.62 It perhaps should not be expected that any of the methods for detecting binding are 100% reliable. All methods are prone to some artifacts resulting from their specific methods of detection and interpretation, and thus a key to success in screening campaigns seems to be to use a robust, highthroughput, low-sample consuming technique for primary screening and to confirm hits using lower-throughput but higher-content orthogonal techniques.60

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AUTHOR INFORMATION

Corresponding Author

*Phone: +386-1-4769673. Fax: +386-1-4258031. E-mail: anamarija.zega@ffa.uni-lj.si. ORCID

Anamarija Zega: 0000-0003-4065-0019

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Notes

PERSPECTIVES, CONCLUSION The success or failure of drug discovery programs is very much influenced by the screening hits. Despite the existence of computational filters and biophysical methods available to detect false hits, nonspecific compounds still represent a challenge to the pharmaceutical industry and especially in academia. In the identification of false positives NMR plays an important role. However, NMR under certain conditions can also generate its own particular false positives. The potential of NMR spectroscopy to reveal molecular interactions in a solution environment enables its use as a method to detect nonspecific binding and therefore to exclude ambiguously behaved compounds from a hit list (Table 1). Most of the assays presented above are easy to apply; they do not require special NMR equipment, and with the exception of ALARM NMR (where the labeled protein is needed), the probes or reference compounds used are small, commercially available molecules. As well as this potential, NMR is unlike some other biophysical methods, as it is accessible to, and routinely used by, almost all medicinal chemists. Last, but not least, there is the need to be aware that NMR spectroscopy itself is prone to the generation of false-positive hits. This problem is especially pronounced in FBDD, where high concentrations of fragments are needed for binding detection, which increases the probability of the formation of aggregates. This problem affects the two most commonly used

The author declares no competing financial interest. Biography Anamarija Zega received her Ph.D in Medicinal Chemistry from the University of Ljubljana, Faculty of Pharmacy, Slovenia in 2002. She did her postdoctoral research in Dr. Michel Arthur’s group at Laboratoire de Recherche Moléculaire sur les Antibiotiques, Inserm in Paris. She is currently Associate Professor for Medicinal Chemistry at the Faculty of Pharmacy in Ljubljana, and she continues her research interest in structure-based drug design with a particular emphasis on antimicrobials, NMR spectroscopy, and investigation of compound promiscuity.

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ABBREVIATIONS USED ALARM, La assay to detect reactive molecules by NMR; CPMG, Car−Purcell−Meiboom−Gill echo train aquisition; DLS, dynamic light scattering; DSF, differential scanning fluorimetry; DTT, dithiolthreitol; FBDD, fragment-based drug discovery; FP, fluorescence polarization; HSQC, highresolution heteronuclear single quantum correlation; HTS, high-throughput screening; ILOE, interligand nuclear Overhauser effect; ISTD/I0, saturation transfer difference effect; ITC, isothermal titration calorimetry; NMR, nuclear magnetic resonance; NOESY, nuclear Overhauser effect spectroscopy; NOE, nuclear Overhauser effect; PAINS, pan assay interference compounds; PrOF, protein-observed fluorine NMR; SPAM, solubility, purity, and aggregation of the molecule; SPR, surface 9445



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(19) Folmer, R. H. A. Integrating biophysics with HTS-driven drug discovery projects. Drug Discovery Today 2016, 21, 491−498. (20) Feng, B. Y.; Simeonov, A.; Jadhav, A.; Babaoglu, K.; Inglese, J.; Shoichet, B. K.; Austin, C. P. A high-throughput screen for aggregation-based inhibition in a large compound library. J. Med. Chem. 2007, 50, 2385−2390. (21) McGovern, S. L.; Helfand, B. T.; Feng, B.; Shoichet, B. K. A Specific mechanism of nonspecific inhibition. J. Med. Chem. 2003, 46, 4265−4272. (22) Sassano, M. F.; Doak, A. K.; Roth, B. L.; Shoichet, B. K. Colloidal aggregation causes inhibition of G protein-coupled receptors. J. Med. Chem. 2013, 56, 2406−2414. (23) Coan, K. E.; Maltby, D. A.; Burlingame, A. L.; Shoichet, B. K. Promiscuous aggregate-based inhibitors promote enzyme unfolding. J. Med. Chem. 2009, 52, 2067−2075. (24) Coan, K. E.; Shoichet, B. K. Stoichiometry and physical chemistry of promiscuous aggregate-based inhibitors. J. Am. Chem. Soc. 2008, 130, 9606−9612. (25) Davis, J. D.; Erlanson, D. A. Learning from our mistakes: the “unknown knowns” in fragment screening. Bioorg. Med. Chem. Lett. 2013, 23, 2844−2852. (26) Harner, M. J.; Frank, A. O.; Fesik, S. W. Fragment-based drug discovery using NMR spectroscopy. J. Biomol. NMR 2013, 56, 65−75. (27) Mayer, M.; Meyer, B. Characterization of ligand binding by Saturation Transfer Difference NMR Spectroscopy. Angew. Chem., Int. Ed. 1999, 38, 1784−1788. (28) Dalvit, C.; Caronni, D.; Mongelli, N.; Veronesi, M.; Vulpetti, A. NMR-based quality control approach for the identification of false positives and false negatives in high throughput screen. Curr. Drug Discovery Technol. 2006, 3, 115−124. (29) Dalvit, C.; Cottens, S.; Ramage, P.; Hommel, U. Half-filter experiments for assignment, structure determination and hydration analysis of unlabeled ligands bound to 13C/15N labelled proteins. J. Biomol. NMR 1999, 13, 43−50. (30) Dalvit, C.; Fogliatto, G.; Stewart, A.; Veronesi, M.; Stockman, B. WaterLOGSY as a method for primary NMR screening: practical aspects and range of applicability. J. Biomol. NMR 2001, 21, 349−359. (31) Dalvit, C.; Pevarello, P.; Tatò, M.; Veronesi, M.; Vulpetti, A.; Sundström, M. Identification of compounds with binding affinity to proteins via magnetization transfer from bulk water. J. Biomol. NMR 2000, 18, 65−68. (32) LaPlante, S. R.; Aubry, N.; Bolger, G.; Bonneau, P.; Carson, R.; Coulombe, R.; Sturino, C.; Beaulieu, P. L. Monitoring drug selfaggregation and potential for promiscuity in off-target in vitro pharmacology screens by a practical NMR strategy. J. Med. Chem. 2013, 56, 7073−7083. (33) LaPlante, S. R.; Carson, R.; Gillard, J.; Aubry, N.; Coulombe, R.; Bordeleau, S.; Bonneau, P.; Little, M.; O’Meara, J.; Beaulieu, P. L. Compound aggregation in drug discovery: implementing a practical NMR assay for medicinal chemists. J. Med. Chem. 2013, 56, 5142− 5150. (34) Dahlin, J. L.; Baell, J.; Walters, M. A.; Assay interference by chemical reactivity. In Assay Guidance Manual; Sittampalam, G. S., Coussens, N. P., Nelson, H., Arkin, M., Auld, D., Austin, C., Bejcek, B., Glicksman, M., Inglese, J., Iversen, P. W., Li, Z., McGee, J., McManus, O., Minor, L., Napper, A., Peltier, J. M., Riss, T., Trask, O. J., Jr., Weidner, J., Eds.; Eli Lilly & Company and the National Center for Advancing Translational Sciences: Bethesda, MD, 2015. (35) Wilson, J. M.; Wu, D.; Motiu-DeGrood, R.; Hupe, D. J. A Spectrophotometric method for studying the rates of reaction of disulfides with protein thiol groups applied to bovine serum albumin. J. Am. Chem. Soc. 1980, 102, 359−363. (36) Huth, J. R.; Mendoza, R.; Olejniczak, E. T.; Johnson, R. W.; Cothron, D. A.; Liu, Y.; Lerner, C. G.; Chen, J.; Hajduk, P. J. ALARM NMR: a rapid and robust experimental method to detect reactive false positives in biochemical screens. J. Am. Chem. Soc. 2005, 127, 217− 224. (37) Huth, J. R.; Song, D.; Mendoza, R. R.; Black-Schaefer, C. L.; Mack, J. C.; Dorwin, S. A.; Ladror, U. S.; Severin, J. M.; Walter, K. A.;

plasmon resonance; STD, saturation transfer difference; TSA, thermal shift assay; WAC, weak affinity chromatography; WaterLOGSY, water ligand observed via gradient spectroscopy

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NMR Methods for Identification of False Positives in Biochemical

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