Causes and Significance of Increased Compound Potency in Cellular

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Causes and Significance of Increased Compound Potency in Cellular or Physiological Contexts Miniperspective Adam G. Schwaid* and Ivan Cornella-Taracido Merck & Co., Inc., Boston, Massachusetts 02115, United States ABSTRACT: Compound potency is a key metric that is often used to drive medicinal chemistry programs. Compound potency is also taken into account when identifying the mechanism of action of compounds whose pharmacological target is unknown, particularly when these compounds are identified in phenotypic screens. Often compound potency is determined from assays using recombinantly generated, purified protein. It is well understood in the medicinal chemistry community that potency measured with recombinant enzyme and potency measured in cell may not entirely coincide. Decreases in cellular vs recombinant potency are often anticipated or explainable. What is less often realized is that compound potency can increase in a cellular environment due to several factors including cellular metabolism of compounds, protein−protein interactions, post-translational modifications, and asymmetric intracellular localization of compound. Here we discuss these factors and highlight examples where increases in cellular compound potency were critical to the development of probes or drugs.

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IMPORTANCE OF UNDERSTANDING POTENCY

recombinant protein. This rightward shift in potency is often understood to be the consequence of compound serum binding, permeability, competition with endogenous substrates, compensatory cellular mechanisms and, in the case of in vivo experiments, pharmacokinetics and metabolism.5−8 These topics are thoroughly discussed in other reviews and will not be considered in depth here. A far less discussed but critically important topic are leftwards shifts in potency in complex systems, such as cells, lysates, or protein complexes, relative to purified recombinant enzyme (Figure 1). Compared to decreases in potency upon transitioning to a cellular system, increases in potency are relatively rare. Such shifts are difficult to measure, as cellular target occupancy is not routinely ascertained, which may contribute to a reporting bias regarding these events. Nonetheless, it is important to consider leftward potency shifts in drug discovery and in molecular mechanism discovery because it can profoundly change the interpretation of a molecule’s selectivity and mechanism of action. There are numerous reasons potency could increase when moving from recombinant enzyme to more complex systems. In particular, purified recombinant systems ignore cellular localization, lack endogenous contacts with other proteins or nucleic acids, and do not take into account posttranslational modifications or the formation of active metabolites.

In drug discovery, biochemical potency is often a key metric used to drive compound optimization. Maximizing potency is important to ensure target engagement and efficacy in vivo. Increasing compound potency can also avoid off-targets by creating a therapeutic window between target and off-target engagement.1 Potency is also important for pharmacokinetics and increases the chance that a compound can be administered at a level that permits peak to trough exposure and avoids toxicity related to metabolic byproducts or hepatotoxicity.1,2 In addition to being important for lead design, compound potency is often utilized during mechanism of action deconvolution. For instance, when considering hits from a phenotypic screen compound, potency against a recombinant panel of enzymes may be considered to triage compounds based on consistent potency in in situ and cellular assays. This may be particularly true when annotated compound sets are applied to phenotypic screen deconvolution, where the annotation derives principally from recombinant enzyme assays. Frequently, tool compounds are chosen based on their annotated potency for a particular enzyme or protein based on data generated from recombinant enzyme assays in the absence of cellular target engagement data.3,4 Consequently, potency is a metric that is used both to drive lead design and drive our understanding of biology.

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LEFTWARD POTENCY SHIFTS A commonly observed phenomenon is a decrease in compound potency in cellular or in vivo systems relative to assays with © XXXX American Chemical Society

Received: May 23, 2017

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DOI: 10.1021/acs.jmedchem.7b00762 J. Med. Chem. XXXX, XXX, XXX−XXX

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Figure 1. Compound binding potency is often determined primarily by binding to recombinant protein. However, numerous factors can result in leftward or rightward shifts in potency upon moving to a cellular or in vivo setting. Correctly interpreting and understanding compound potency in physiologically relevant settings is critical to compound development and understanding the compound’s mechanism of action.

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CELLULAR LOCALIZATION Typically, the cellular concentration of a compound is considered to be equivalent to the free fraction of compound the cells were treated with and uniform throughout the cells. However, variations within the cell, such as pH can affect the accumulation of compound in different parts of the cell (Figure 2A). For instance, Zuhl et al. found that numerous β-secretase 1 (BACE1) inhibitors bearing basic amines accumulate in the lysosome due to the low lysosomal pH and protonation of the basic amine. Accumulation of BACE1 inhibitors results in the apparently potent cellular inhibition of cathepsin D (CatD). Notably, the principle BACE1 inhibitor examined in this study, 1 (PF-9283), inhibited recombinant CatD with a potency of 12 μM but inhibited CatD in cells with a potency of 140 nM, an 85-fold shift (Figure 3).9 Neutralization of the lysosome, which prevents intralysosomal trapping of 1, prevented CatD target occupancy. In vivo treatment of 1 or other BACE1 inhibitors resulted in the accumulation of autofluorescent granules in the retinas of rats. Cellular potency for CatD and exposure of these BACE1 inhibitors was found to correlate with granule formation. Additionally, it is known that specific CatD inhibition also results in granule formation, supporting the hypothesis that these BACE1 inhibitors induced ocular toxicity through CatD. Notably, inhibitor potency versus recombinant enzyme did not track with ocular toxicity, highlighting the importance of measuring cellular potency. This example demonstrates how asymmetric subcellular localization of compound can affect potency and consequently selectivity. Likewise, lysosomal sequestration of sunitinib, a receptor tyrosine kinase inhibitor originally approved for renal cell carcinoma, can either increase or decrease the potency of sunitinib for its protein interaction partners.10,11 Sunitinib is a weakly basic compound with a pKa of 8.95. It has been shown by confocal microscopy to accumulate in the lysosome.11 Because its primary targets, vascular endothelial growth factor receptor 2 (VEGFR2), and platelet derived growth factor receptor (PDGFR) are cytosolic, one method of tumor cell resistance to sunitinib is to increase the level of acidic lysosomes in the cell. This results in increased concentrations of intracellular sunitinib yet decreased efficacy as measured by the effect of sunitinib on cell proliferation.11 On the other hand, lysosomal accumulation of sunitinib results in inhibition of lysosomal proteins and effects on the lysosome compartment.10 Sunitinib has been shown to induce cell death through

Figure 2. Several mechanisms can result in leftward shifts in compound potency in cells or in vivo. (A) Asymmetric compound localization (hexagons) within the cell can result in apparently potent binding to off-targets constrained to cellular organelles (yellow), whereas the intended targets (red) may be exposed to lower compound concentrations. (B) Compound potency may be increased by binding to protein complexes rather than isolated proteins. For instance, a compound may have only a shallow protein binding site in the absence of other proteins (left), whereas protein−protein complexes may form larger binding pockets either through allosteric interactions with the target protein or by forming direct contacts with the compound (middle and right). (C) Post-translational modifications can induce changes in protein conformation. These changes can lead to the exposure of compound binding pockets. This may result in a compound having increased potency for the post-translationally modified form of a protein found within the cell as opposed to a recombinantly expressed and purified protein.

lysosomal cell death pathways. Sunitinib was also shown to inhibit lysosomal sphingomyelinase activity and subsequently results in the permeabilization of lysosomes.10 It should also be noted that because some drugs can affect the characteristics of cellular organelles, for example, chloroquine treatment increases lysosomal pH, it has been hypothesized that the subcellular sequestration of compounds, and the consequent apparent effect on potency and efficacy could be influenced by drug−drug interactions.12,13 Although the above examples focus on lysosomal sequestration, there are also examples of drugs that localize to other cellular organelles.14 For instance, while in daunorubicin sensitive cells, daunorubicin accumulates in the cytoplasm and nucleus as expected in daunorubicin resistant K562 cells, Gong et al. determined that a fluorescently labeled daunorubicin accumulates in the mitochondria due to the action of intracellular ATP-binding cassette (ABC) transporter permeB

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Figure 3. Compounds referenced in the text.

ability glycoprotein (Pgp).15 Likewise, fluorescently tagged trimethoprim has been shown to localize to the cell membrane rather than diffusing throughout the cytoplasm.16 Drug localization has been hypothesized to be mediated by pH, intracellular transporters, and protein−small-molecule interactions.17,18 It is probable that accumulation in organelles other than the lysosome could result in an increase in the apparent binding affinity of small molecules.

ancy between inhibition and binding. Further studies with the antibiotics chloramphenicol and linezolid have shown that ribosome inhbition can be dependent upon the sequence of the polypeptide the ribosome is translating supporting the role of the protein complex in compound binding.27 A clearer example of this phenomena is evernimicin.28 Evernimicin binds directly to the 50S ribosome, but binding to purified 50S ribosome from Escherichia coli is 160 nM by radioligand binding assay, whereas binding to 70S ribosome, which consists of 30S and 50S ribosome, is 84 nM.28 Although the difference in affinity is relatively small, this instance demonstrates that recapitulating the protein complex that is present in the cellular environment is important to determining a relevant binding affinity. More recent examples of this phenomenon are available as well. Compound 2 (PF-06446846) is a molecule discovered from a phenotypic screen for compounds that reduce secretion of proprotein convertase subtilisin/kexin type 9 (PCSK9)(Figure 3).29,30 Compound 2 is reported to prevent translation of PCSK9 by binding to the ribosome in the presence of the PCSK9 mRNA 5′-UTR. Notably, 2 inhibits PCSK9 secretion in cells at a concentration of 300 nM, whereas in in vitro translation systems, it inhibited PCSK9 secretion with a potency of only 2 μM. Although this discrepancy cannot definitively be attributed to a difference in binding affinity to the protein−RNA complex, the large number of examples of similar phenomena with bacterial ribosome inhibitors suggests this is a plausible possibility. In general, many examples of a leftward shift in potency between recombinant and in cell systems emerge from molecules that were found via classical phenotypic screening based drug discovery. This is intuitively sensible because target based approaches frequently preclude the identification of these interactions by focusing on purified protein. Often, the first step in a target based screening funnel is to screen against recombinantly expressed purified protein. Evidently, limiting the drug discovery process in this manner risks missing important protein−compound interactions as is evidenced by the compounds discussed above which could not have been found if investigators had screened against a single protein alone. Nonetheless, there are some instances of increases in cellular potency due to protein complex formation emerging from compounds discovered through target based programs. For instance, chemoproteomic profiling of known histone deacetylase (HDAC) inhibitors in cell lysates found that the potency of inhibitors in lysates often varied compared to their potency in assays with recombinant protein.31 The potency of

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PROTEIN COMPLEXES Compound potency can be affected by intramolecular interactions between the target protein and endogenous proteins or nucleic acids. This can occur either by altering the conformation of the protein or by creating a new binding site at the interface of the protein−protein/nucleic acid binding site (Figure 2B). In these cases, binding potency can be underestimated or missed entirely when screening against purified protein. This is often observed in the case of inhibitors of DNA binding enzymes. For instance, binding of DNA gyrase to DNA induces the formation of a fluoroquinoline binding site.19 In the absence of this ternary complex, fluoroquinoline shows poor binding to both DNA gyrase and DNA. Likewise, the drug sofosbuvir, which inhibits synthesis of hepatitis C viral RNA, binds to nonstructural protein 5B (NS5B) polymerase as it is transcribing viral RNA. The metabolically activated trinucleotide of sofosbuvir makes electrostatic contacts with NS5B and engages in Watson−Crick base pairing interactions with a complementary adenine from RNA.20,21 The 2′-methyl and 2′-fluorine blocks chain elongation and inhibits RNA polymerization. Contacts from both NS5B protein and viral RNA contribute to the binding affinity of sofosbuvir. The dependence on protein complex formation for compound binding has also been observed for antibiotics that bind the ribosome. For example, radioligand binding assays with the antibiotics lincomycin and eperezolid and the 50S ribosome show a potency of ∼34 and ∼20 μM, respectively.22−24 However, functional assays show these molecules are more potent than would be predicted by 50S binding. In the case of eperezolid, in vitro translation shows a potency of 1.8 μM, a potency below the apparent binding potency of eperezolid to purified 50S ribosome.25 Furthermore, the MIC90s for lincomycin and eperezold are lower than would be anticipated based on a potency of 20−34 μM.22−24,26 These data suggest that small molecules may have different potency for purified subunits of the ribosome than for the complete ribosomal complex. This could explain the observed discrepC

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inhibition between post-translationally modified and unmodified protein was not apparent from recombinant protein assays as in vitro the inhibitor was found to label both glycosylated and partially deglycosylated LIPA equally.38 Post translational modifications can be labile and are dynamically regulated. Consequently they can be very difficult to study. In addition, there are many different ways a given protein can be post translationally modified and often a protein may be modified differently depending on the system in which it is expressed. When studying compound binding in recombinant systems it is almost impossible to account for all potential PTMs. Therefore, it becomes extremely important to monitor target occupancy in cells as well because PTMs may change compound potency depending on the biological system studied. The same challenges that make PTMs difficult to study in general make it difficult to determine if compound potency is affected by PTMs. Cases where there are known compound binding potency shifts are typically found once a chemoproteomic toolset to interrogate a particular enzyme class is discovered. Because most proteins do not have a toolset targeted to their particular protein family. It is likely there are more such instances of compounds binding to proteins only upon post-translational modification than is recognized in the literature. Likely, as more chemoproteomic tools are developed, more instances of compounds binding only to activated or inactivated protein conformations will come to light.

these inhibitors was found to vary in this experiment due to the presence of other proteins that formed protein complexes with HDACs. These protein−protein interactions could be copurified with affinity techniques using immobilized HDAC inhibitors.31 Likely, there are more instances of the dependence of compound affinity on the environment of the protein target than have been realized in the literature. Ascertaining direct binding affinity in systems representative of the endogenous environment of a protein is difficult. Fortunately, more and more tools to examine protein−small-molecule interactions in their native environment are being developed and awareness for the importance of measuring target occupancy in cells or lysates is increasing.32,33

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POST-TRANSLATIONAL MODIFICATIONS Many proteins undergo post-translational modifications (PTMs) during or after protein biosynthesis prior to exerting their biological roles. The most noted of these alterations may be phosphorylation, however, there are numerous other types of covalent post-translational protein changes. These include citrullination, acetylation, methylation, nitrosylation, glycosylation, and many more.34 These PTMs are often ignored when generating binding affinity data with recombinant protein for drug discovery purposes, mainly due to heterogeneous nature of the native protein pool and intrinsic difficulty of accessing the different isoforms. For instance, when an endogenously glycosylated protein is generated recombinantly in bacteria and then used for screening it is unlikely it will contain the same glycosylation pattern as endogenously expressed protein.35 Post-translational modifications can also affect the binding affinity of proteins and small molecules (Figure 2C). Some examples are provided by kinase inhibitors. Many kinases are phosphorylated, and their phosphorylation status subsequently effects their conformation and can affect the potency of inhibitors that bind them. Binding affinities of inhibitors for phosphorylated and nonphosphorylated kinases are not routinely compared using recombinant protein. However, chemoproteomic experiments using kinobeads have shed light on this. Kinobeads are made up of several kinase inhibitors immobilized on a solid support.36 Kinobeads are used to affinity purify kinases from lysates and competition experiments performed with free kinase inhibitor can be used to determine binding potency. Combined phosphoproteomic− chemoproteomic studies have shown that, although the majority of kinases that bind to kinobeads do so irrespective of kinase phosphorylation status, there are instances of kinases that bind to inhibitors more potently upon phosphorylation. For instance, Ruprecht et al. found that mitogen-activated protein kinase 1 (MAPK1), mitogen-activated protein kinase 3 (MAPK3), and ribosomal protein S6 kinase α-1 (RPS6KA1) were enriched preferentially by certain kinase inhibitors depending on whether or not cell lysate was pretreated with pervanadate.37 In fact, while some inhibitors only bound kinases after pervanadate treatment, and presumably phosphorylation, others only bound in the absence of pervanadate treatment. Bantscheff et al. made a similar discovery that dasatinib bound focal adhesion kinase (FAK) only in the active, phosphorylated state.36 In cells, inhibitors have also been shown to preferentially bind enzymes after post-translational modifications other than phosphorylation. Lysosmal acid lipase (LIPA) is a highly glycosylated protein, which is shuttled to the lysosome. In cells, covalent inhibitors were found to label only the glycosylated form of LIPA. The difference in cellular

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ACTIVE METABOLITES In vivo metabolism of drugs can form metabolites that also have biological activity. The formation of active metabolites from drugs in cells or in vivo can skew the potency predicted by recombinant protein binding assays. In some cases, the active metabolite of a drug is more active and is more potent than the parent molecule. While many prodrugs are designed to form an active metabolite in vivo from an inactive precursor in order to optimize pharmacokinetics, bioavailability, or tissue distribution of the drug, it is also the case that potent active metabolites are found in vivo by serendipity. The drug leflunomide is an immunosuppressive agent that is used to treat rheumatoid arthritis.39 In vivo, the oxazole ring of leflunomide is opened to a β-keto amide.40 The β-keto amide, teriflunomide, has micromolar potency against recombinant tyrosine kinases and was also shown to inhibit recombinant dihydroorotate dehydrogenase (DHODH) with nanomolar potency, in line with the cellular and in vivo potency of teriflunomide. Leflunomide has very poor affinity for recombinant DHODH.40,41 Likewise metabolism of ezetimibe results in an active metabolite that is more potent in vivo than the parent molecule. Upon administration ezetimibe is reversibly glucuronidated. Glucuronidation has not been shown to increase potency of ezetimibe for its target, Niemann−Pick C1-like 1 (NPC1L1), rather glucuronidation effects tissue localization of the molecule.42 In bile duct cannulated rats, a time course of ezetimibe tissue distribution after intraduodenal administration shows accumulation of ezetimibe predominantly in the bile, whereas the administration of the glucuronidated metabolite accumulates predominantly in the intenstinal lumen and wall, the site of action of ezetimibe.43 Consequently, administration in this manner of the glucuronidated metabolite has a greater effect on cholesterol absorption.43 D

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potency of 14 nM vs recombinant enzyme that exhibited a potency of 1.3 nM in human hepatocytes.49 Again, no clear explanation to account for the difference in potency between recombinant enzyme and cellular inhibition was presented. In addition to the other explanations presented here, it is also possible that enzymes may not maintain their proper conformations in lysate conditions, which could affect their capacity to bind small molecules. Dissecting the reasons for differences in binding between lysates and cells is experimentally challenging and can be even more challenging to predict a priori. Consequently, it is important to measure target occupancy in the most biologically relevant system possible. In vitro potency is often used to predict target occupancy in vivo and predict necessary exposure and dosing. Underestimating the potency of a molecule poses several unique risks. If a molecule is more potent than predicted, it may be possible to dose less, and in turn the therapeutic window may be larger than appreciated. It is possible that otherwise promising lead matter is being discarded for this reason. Off targets can also be missed for the same reason. Measuring selectivity in cells or tissues that best represent the in vivo site of action could help de-risk compounds before lengthy toxicology studies. Emerging chemoproteomic techniques make this possible.50,51 More and more often, scientists in the pharmaceutical industry are realizing the importance of measuring target engagement in vivo in order to derisk pharmacology. Understanding compound potency is also important for correctly deconvoluting phenotypic screens. If a compound appears to be potent in a phenotypic assay but binds a particular recombinant protein with poorer affinity than observed in cells, it is possible to erroneously discard a hit. This type of caveat must be kept in mind, particularly when working with annotated compound sets whose “targets” have been annotated through previous in-house experiments. While “annotated mechanism of action” compound collections are extremely valuable for target deconvolution, predictions must be taken with a grain of salt and it must be kept in mind that the compound potency was measured in a system that may not represent the cellular system. It is likely that as techniques to measure target occupancy in cells continue to be developed and as scientists gain a better appreciation of the importance of measuring binding potency in cells or in vivo, additional examples of leftward shifts in potency will emerge. Ideally, these instances will lead to better understanding of the causes of these discrepancies and improvements in prediction of in vivo binding potency.

More generally, active transport mechanisms should be considered when measuring the potency of a compound in cells. For instance, some statins, which are actively transported through organic anion transporters (OATPs), are much more potent in hepatocytes than myocytes and have even been documented to be more potent in hepatocytes than would be predicted by recombinant enzyme assays.44 Compound 3 (YM155) also illustrates this principle. Compound 3 is a DNA intercalator that is dependent upon the expression of solute carrier family 35 member F2 (SLC35F2) for cellular uptake and efficacy (Figure 3). After clinical trials with this compound were concluded, it was found that poor expression in patient tissues of SLC35F2 may have been a contributing factor to the failure of this molecule in the clinic.45 Active uptake mechanisms can impart cell type specificity and are important to keep in mind when exploring the mechanism of action of compounds. Similar to the example of ezetimibe, the metabolism of methotrexate, a dihydrofolate reductase (DHFR) antagonist that is used as an antiproliferative and antineoplastic agent, increases its intracellular concentration resulting in higher potency.46,47 Methotrexate is polyglutamylated within the cell which decreases cellular efflux. The degree of polyglutamylation can vary from cell type to cell type. For instance, 55.4% and 87.6% of methotrexate was found to be polyglutamylation in MCF-7 and ZR-75-B cells respectively after 24 h,47 whereas under the same treatment conditions only 32% of methotrexate was polyglutamylated in MD-231 cells.47 During washout experiments, MCF-7 and ZR-75-B cells continued to show inhibition of proliferation whereas MD-231 cell proliferation recovered post washout.47 Recently, cellular thermal stability shift assays (CETSA) were used to clearly demonstrate the increase in methotrexate potency in cells versus in lysates.46 In this case, the increase in compound potency in cells is dependent on the cell type. This highlights the need to consider compound potency not only in a cellular model but in a model that closely represents the physiological site of action. As demonstrated, these types of cellular or in vivo transformations can result in unanticipated increases in potency. Because most drugs form several different metabolites, it is not trivial to determine whether a metabolite with an increased potency is formed or what that metabolite may be. Nevertheless it is important to keep these transformations in mind when optimizing lead matter or attempting to rationalize observed in situ and cellular potency of small molecules identified in phenotypic screens.

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DISCUSSION AND CONCLUSIONS In addition to the reasons listed above for leftward shifts in compound potency, it is also the case that increases in compound potency in more biologically relevant contexts can happen for reasons that are, as of yet, unclear. For instance, Evans et al. screened a protein reactive library of compounds in cells for molecules that inhibited proliferation of breast cancer cells. They identified 4 (MJE3), which covalently binds to phosphoglycerate mutase 1 (PGAM1) and appeared to inhibit cancer cell proliferation by inhibiting glycolysis (Figure 3).48 While 4 clearly bound to PGAM1 in MDA-MB-231 cells, no binding was observed in lysates from MDA-B-231 cells or in cell lysates from COS-7 cells recombinantly expressing PGAM1. This is an instance where a compound is clearly more potent in cells, however, there is no clear explanation as to why. In the same vein, Pfizer reported a selective diacylglycerol O-acyltransferase 2 (DGAT2) inhibitor with a

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AUTHOR INFORMATION

Corresponding Author

*Phone: 617-992-3306. E-mail: [email protected]. ORCID

Adam G. Schwaid: 0000-0003-2434-6148 Notes

The authors declare no competing financial interest. Biographies Adam G. Schwaid completed his Ph.D. studies in the Department of Chemistry and Chemical Biology at Harvard University with Dr. Alan Saghatelian. Subsequently, he worked at Pfizer as a chemical biologist investigating cardiovascular, metabolic, and endocrine disease before joining the Chemical Biology group at Merck in 2016. E

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and inhibit multidrug resistance. Mol. Cancer Ther. 2013, 12 (10), 2018−2030. (11) Gotink, K. J.; Broxterman, H. J.; Labots, M.; de Haas, R. R.; Dekker, H.; Honeywell, R. J.; Rudek, M. A.; Beerepoot, L. V.; Musters, R. J.; Jansen, G.; Griffioen, A. W.; Assaraf, Y. G.; Pili, R.; Peters, G.; Verheul, H. M. W. Lysosomal sequestration of sunitinib: a novel mechanism of drug resistance. Clin. Cancer Res. 2011, 17 (23), 7337− 7346. (12) Michihara, A.; Toda, K.; Kubo, T.; Fujiwara, Y.; Akasaki, K.; Tsuji, H. Disruptive effect of chloroquine on lysosomes in cultured rat hepatocytes. Biol. Pharm. Bull. 2005, 28 (6), 947−951. (13) Pascolo, S. Time to use a dose of chloroquine as an adjuvant to anti-cancer chemotherapies. Eur. J. Pharmacol. 2016, 771, 139−144. (14) Zheng, N.; Tsai, H. N.; Zhang, X.; Shedden, K.; Rosania, G. R. The subcellular distribution of small molecules: a meta-analysis. Mol. Pharmaceutics 2011, 8 (5), 1611−1618. (15) Gong, Y.; Wang, Y.; Chen, F.; Han, J.; Miao, J.; Shao, N.; Fang, Z.; Yang, R. O. Identification of the subcellular localization of daunorubicin in multidrug-resistant K562 cell line. Leuk. Res. 2000, 24, 769−774. (16) Phetsang, W.; Pelingon, R.; Butler, M. S.; KC, S.; Pitt, M. E.; Kaeslin, G.; Cooper, M. A.; Blaskovich, M. A. T. Fluorescent trimethoprim conjugate probes to assess drug accumulation in wild type and mutant escherichia colia. ACS Infect. Dis. 2016, 2, 688−701. (17) Bour-Dill, C.; Gramain, M.-P.; Merlin, J.-L.; Marchal, S.; Guillemin, F. Determination of intracellular organelles implicated in daunorubicin cytoplasmic sequestration in multidrug-resistant MCF-7 cells using fluorescence microscopy image analysis. Cytometry 2000, 29, 16−25. (18) Gong, Y.; Duvvuri, M.; Krise, J. Separate roles for the golgi apparatus and lysosomes in the sequestration of drugs in the multidrug-resistant human leukemic cell line HL-60. J. Biol. Chem. 2003, 278 (50), 50234−50239. (19) Shen, L. L.; Mitscher, L. A.; Sharma, P. N.; O’Donnell, T. J.; Chu, D. W. T.; Cooper, C. S.; Rosen, T.; Pernet, A. G. Mechanism of inhibition of DNA gyrase by quinoline antibacterials: a cooperative drug-DNA binding model. Biochemistry 1989, 28 (9), 3886−3894. (20) Appleby, T. C.; Perry, J. K.; Murakami, E.; Barauskas, O.; Feng, J.; Cho, A.; Fox, D., III; Wetmore, D. R.; McGrath, M. E.; Ray, A. S.; Sofia, M. J.; Swaminathan, S.; Edwards, T. E. Structural basis for RNA replication by the hepatitis C virus polymerase. Science 2015, 347 (6223), 771−775. (21) Kulkarni, A. S.; Damha, M. J.; Schinazi, R. F.; Mo, H.; Doehle, B.; Sagan, S. M.; Götte, M. A complex network of interactions between S282 and G283 of hepatitis C virus nonstructural protein 5B and the template strand affects susceptibility to sofosbuvir and ribvarin. Antimicrob. Agents Chemother. 2016, 60 (4), 2018−2027. (22) Lin, A. H.; Murray, R. W.; Vidmar, T. J.; Marotti, K. R. The oxazolidinone eperezolid binds to the 50S ribosomal subunit and competes with binding of chloramphenicol and lincomycin. Antimicrob. Agents Chemother. 1997, 41 (10), 2127−2131. (23) Rybak, M. J.; Cappelletty, D. M.; Moldovan, T.; Aeschlimann, J. R.; Kaatz, G. W. Comparative in vitro activities and postantibiotic effects of the oxazolidinone compounds eperezolid (PNU-100592) and linezolid (PNU-100766) versus vancomycin against staphylococcus aureus, coagulase-negative staphylococci, Enterococcus faecalis, and Enterococcus faecium. Antimicrob. Agents Chemother. 1998, 42, 721− 724. (24) Fernandez-Muñoz, R.; Monro, R. E.; Torres-Pinedo, R.; Vazquez, D. Substrate and antibiotic-binding sites at the peptidyltransferase centre of escherichia coli ribosomes. Eur. J. Biochem. 1971, 23, 185−193. (25) Shinabarger, D. L.; Marotti, K. R.; Murray, R. W.; Lin, A. H.; Melchior, E. P.; Swaney, S. M.; Dunyak, D. S.; Demyan, W. F.; Buysse, J. M. Mechanism of action of oxazolidinones: effects of linezolid and eperezolid on translation reactions. Antimicrob. Agents Chemother. 1997, 41 (10), 2132−2136.

Iván Cornella-Taracido Ph.D., Director of Chemical Biology at Merck & Co., is responsible for creating a framework to leverage chemical biology towards building novel early discovery competencies, with a particular emphasis on integrating chemoproteomics, chemogenomics, and chemical probe development with molecular biology, pharmacology, screening, and informatics to enable molecular mechanism of action studies, target identification, and validation. Throughout his career, Iván has contributed to advance discovery projects across multiple therapeutic areas such as oncology, infection, and immunology. Before moving to industry, Iván was a U.S. National Cancer Institute Initiative for Chemical Genetics Postdoctoral Research Fellow at the Institute of Chemistry and Cell Biology of Harvard Medical School and NIH Postdoctoral Research Fellow at Boston College. He received his Ph.D. from the Universidade da Coruña, Spain.

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ABBREVIATIONS USED BACE1, β-secretase 1; CatD, cathepsin D; VEGFR2, vascular endothelial growth factor receptor 2; PDGFR, platelet derived growth factor receptor; Pgp, ATP-binding cassette (ABC) transporter permeability glycoprotein; NS5B, nonstructural protein 5B; PCSK9, proprotein convertase subtilisin/kexin type 9; HDAC, histone deacetylase; PTM, post-translational modification; MAPK1, mitogen-activated protein kinase 1; MAPK3, mitogen-activated protein kinase 3; RPS6KA1, ribosomal protein S6 kinase α-1; FAK, focal adhesion kinase; LIPA, lysosomal acid lipase; DHODH, dihydroorate dehydrogenase; NPC1L1, Niemann−Pick C1-like 1; OATP, organic anion transporter; SLC35F2, solute carrier family 35 member F2; DHFR, dihydrofolate reductase; CETSA, cellular thermal shift assay; PGAM1, phosphoglycerate mutase 1 (PGAM1); DGAT2, diacylglycerol O-acyltransferase 2

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