Discovery and Characterization of Novel Nonsubstrate and Substrate NAMPT Inhibitors

In addition to activation of oncogenes and loss of tumor suppressor genes, cancer cells depend on normal, nononcogenic processes to cope with the stresses resulting from dysregulated cell signals. This heightened dependence on normal processes, also known as nononcogene addiction, represents an opportunity for targeting of cancer cells (1). Cancer metabolism, often summarized as the Warburg effect, is characterized by high utilization of glucose for glycolysis, which depends on NAD⁺. The glycolytic intermediates that are produced enter the pentose phosphate pathway, resulting in conversion of NADP⁺ to NADPH (2). In addition to the roles of NAD⁺ and NADP⁺ in metabolic redox cycles, NAD⁺ is a consumed substrate of enzymes such as PARPs that are involved in DNA damage recognition and repair, cyclic ADP ribose synthetases that generates calcium-mobilizing second messengers, and sirtuins that regulate transcription and metabolic processes (2, 3). The latter process of NAD⁺ consumption that results in nicotinamide production represents a form of nononcogene addiction, making many cancer cells highly dependent on salvage synthesis of NAD⁺ (4).

Depending on which genes are expressed and the availability of substrates, mammalian cells can utilize tryptophan through the de novo NAD⁺ synthesis pathway or salvage nicotinamide, nicotinic acid (NA), or nicotinamide riboside (NR) to maintain NAD⁺ levels (Supplementary Fig. S1; refs. 2, 3, 5, 6). As long as it is not methylated (7), the nicotinamide produced by NAD⁺-consuming enzymes can be recycled back to NAD⁺ in a pathway initiated by nicotinamide phosphoribosyltransferase (NAMPT; ref. 8–10). This enzyme utilizes phosphoribosyl pyrophosphate (PRPP) as a phosphoribose donor to convert nicotinamide to nicotinamide mononucleotide (NMN). NA, which is obtained from the diet or produced by bacterial hydrolysis of NAD⁺ metabolites in the gut (11), is utilized in a pathway initiated by NA phosphoribosyltransferase (NAPRT1). In a significant percentage of cancers, the NAPRT1 gene is epigenetically silenced, providing an opportunity to select patients whose cancers depend on the NAMPT-mediated pathway to regenerate NAD⁺ (12, 13). In addition, several cancer cell lines lack expression of key enzymes within the de novo pathway and are unable to generate NAD⁺ from tryptophan (14). Given the increased reliance on regenerating NAD⁺ from nicotinamide, inhibition of NAMPT represents a potential therapeutic intervention point to preferentially target cancer cells. Indeed, coadministration of an NAMPT inhibitor plus NA might constitute a way to target the heightened NAD⁺ requirements of tumor while protecting patients from toxicities in nontumor tissues (15). In addition, provision of NR has been shown to protect rats from paclitaxel-induced peripheral neuropathy (16).

The concept of inhibiting NAMPT as an anticancer strategy was first evaluated clinically in 2007 with FK866 (17) and GMX1778 (18). These first-generation NAMPT inhibitors exhibited potent antitumor activity in murine xenograft models but experienced limitations when evaluated clinically. Poor and variable oral bioavailability coupled with short plasma half-life necessitated the administration of FK866 and GMX1777 (19), a soluble prodrug of GMX1778, by 24- to 96-hour intravenous infusion (17). The lack of efficacy observed with first-generation NAMPT inhibitors was hypothesized to result from insufficient target engagement prior to reaching doses that caused toxicity (10). As such, there has been a resurgence of activities to develop better NAMPT inhibitors with improved pharmaceutical properties.

Several second-generation NAMPT inhibitors have been described in the literature and in patent applications (10, 20–24). Similar to FK866 and GMX1778, GNE-617 and other second-generation NAMPT inhibitors with cellular activity have an aromatic nitrogen positioned for phosphoribosylation by NAMPT (Fig. 1A; refs. 10, 25, 26). Phosphoribosylation of NAMPT inhibitors has been confirmed by mass spectrometry or cocrystal structures (25, 27–29). According to published work, the substrate activity of NAMPT inhibitors is required for potent cellular activity in vitro and preclinical efficacy in vivo (27).

Here, we identified novel nonsubstrate NAMPT inhibitors with robust preclinical efficacy and pharmacokinetics properties that enable oral dosing. The lead molecule in this series, A-1293201, contains an isoindoline “head group,” which lacks the aromatic nitrogen of the nicotinamide and nicotinamide-mimetic compounds. Work presented establishes the degree of NAMPT inhibition required to kill cancer cells with A-1293201 in vitro and in vivo. Our data demonstrate that formation of a phosphoribose adduct by NAMPT is not a requirement for activity within tumor cells.

PC3, HCT116, and NCI-H1975 cells were obtained from the ATCC in 2001, 1999, and 2006, respectively. Cells were cultured in RPMI1640 containing 50 mmol/L HEPES (Gibco 22400) supplemented with 10% FBS for HCT116 and NCI-H1975 cells and with 1% sodium pyruvate, 1% nonessential amino acids, and 10% FBS for PC3 cells. Cells were incubated at 37°C in 5% CO₂ and 80% relative humidity. Cells were authenticated by STR analysis in July 2016 for PC3 and HCT116 cells and in August 2013 for NCI-H1975 cells.

Cytotoxicity was determined using the CellTiter Glo cell viability assay that detects total intracellular ATP (Promega). Inhibitor affinity capture was performed as described in ref. 30 using molecular probes based on the FK866 or A-933414 structures. Captured proteins were separated by SDS-PAGE and SYPRO staining and identified by LC/MS-MS. FK866 and GMX1778 were synthesized at AbbVie. Additional compounds were synthesized using methods described in U.S. Patent 2016/9302989 B2 (31).

The full-length DNA encoding human NAMPT, residues 1 to 491 (GenBank accession number NM_005746.2) with the FLAG-tag (DYKDDDDK) introduced at the C-terminus, was synthesized and cloned into the pLVX-IRES-puro expression vector. Full-length NAMPT-FLAG was transiently expressed in HEK 293-6E cells (NRC-Canada) and purified in a two-step process using anti-FLAG affinity chromatography and size exclusion chromatography. Enzyme assays were performed using the direct NMN detection method (32).

Purified NAMPT at 10 to 13 mg/mL was incubated in the presence of 10 mmol/L nicotinamide for 2 hours on ice. Crystals grew at 4°C in hanging drop vapor diffusion setup containing 1:1 ratio of the protein and reservoir solution (25% PEG 3350, 0.2 mol/L ammonium sulfate, and 0.1 mol/L HEPES, pH 7.5). Inhibitor complexes were formed by soaking the crystals in the presence of the compound and prepared for X-ray studies by brief transfer into reservoir solution with 20% (v/v) glycerol and rapid plunge into liquid nitrogen. Diffraction data for the complex with A-1293201 were collected under gaseous nitrogen at 100 k at the Canadian Light Source Beamline 08ID-113, while data for the complex with A-1326133 were similarly collected at the Advanced Photon Source Beamline 17-ID. Diffraction intensities were processed using autoPROC (33), and the structure was solved by sequential molecular replacement with coordinates from pdb code 2GVJ using MOLREP (34) within the CCP4 program suite (35). The model was rebuilt using COOT (36) and refined against structure factors using the programs REFMAC5 (37) and autoBUSTER (38). Figures were prepared using the program PyMOL (Schroedinger, LLC). Atomic coordinates for the complexes with A-1293201 and A-1326133 have been deposited in the public wwPDB with accession codes 5U2M and 5U2N, respectively. NAMPT crystal structure data and refinement statistics are listed in Supplementary Table S1.

For PC3 cell viability assays, cells were treated in 96-well plates with compounds ± 0.3 mmol/L nicotinamide for 5 days, followed by a CellTiter Glo assay (Promega). At the 5-day time point, ATP was already depleted and the signal correlated with cell viability as measured by other methods. For shorter time course assays, HCT116, PC3, or NCI-H1975 cells were treated for the indicated times, then cellular levels of NAD⁺ + NADH (NADt) were measured using the NADH-Glo assay (Promega), and ATP levels were measured using the CellTiter Glo assay. Cell numbers were measured by quantifying cells stained with Vybrant Green (Invitrogen) and imaged in an IncuCyte FLR (Essen Bioscience).

NAD⁺ and related metabolites were analyzed as described previously (39, 40). Briefly, 1 μL of one of two internal standard solutions (solution A and solution B) were added to each sample (NCI-H1975 pellet containing 1 × 10⁶ cells) before extraction. Solution A was composed of extract from Fleischmann yeast grown in the presence of U-¹³C glucose diluted to produce a final concentration of 1:80 heavy extract to sample solution. Solution B contained 60 μmol/L ¹⁸O nicotinamide riboside and ¹⁸O nicotinamide. Samples were deproteinized using buffered ethanol [75%/25% (v/v) ethanol/10 mmol/L HEPES pH 7.1] heated to 80°C for 3 minutes with vortexing. Particulate was pelleted with centrifugation (16,100 × g, 10 minutes, 4°C). The supernatant was transferred to fresh tubes and then dried overnight via speed vacuum. Reconstituted extracts were separated and analyzed via LC/MS-MS as described previously (39, 40). Metabolites were detected using a Waters TQD operated in positive ion mode by multiple reaction monitoring (MRM). Metabolite peak areas were converted to mole amounts by regression to internal standard (IS) peak areas. Intracellular concentrations were calculated by dividing the mole amount by the total intracellular volume, assuming an intracellular volume of 2.5 pL per cell. One-way ANOVA followed by Dunnett multiple comparisons test was performed using GraphPad Prism version 7.00 for Windows, GraphPad Software (www.graphpad.com).

NCI-H1975 cells were harvested and aliquoted into 1 × 10⁶ cells in duplicate. Samples were spun and washed with cold PBS, followed by metabolite extraction in 500 μL of 80% (v/v) methanol in water containing 2.5 μmol/L U-¹³C glucose-6-phosphate as the IS. Samples were vortexed and incubated at −80°C for 20 minutes then spun for 30 minutes at 13,000 × g and 4°C. Supernatants were transferred into cold Eppendorf tubes, lyophilized to dryness, and stored at −80°C until analysis. Lyophilized samples were reconstituted with 50 μL of water, agitated for 15 minutes at 4°C, then diluted with 50 μL of acetonitrile, and agitated for 15 minutes at 4°C. Extracts were separated and analyzed as described previously (41); however, only positive mode MRM transitions for select metabolites were included in the method. Linear dynamic range for each metabolite was defined with standards, and samples were diluted and reinjected when necessary. Changes in metabolite concentrations are reported as fold change to control.

All experiments were conducted in accordance with IACUC guidelines. Compounds were formulated in the following vehicle 2% (vol) EtOH, 5% (vol) Tween 80, 20% (vol) PEG-400, 73% (vol) 0.2% HPMC and were administered orally in the pharmacology studies. HCT116 cells were grown to passage 2 in vitro. A total of 0.5 × 10⁶ cells were inoculated into the right flank of female C.B.-17 SCID mice on day 0 in a volume of 0.1 mL. Tumors were size matched on day 11 postinoculation with a mean tumor volume of 231 ± 28 (SD) mm³ with dosing beginning on the next day. Tumor volume was calculated twice weekly. Measurements of the length (L) and width (W) of the tumor were taken via electronic caliper, and the volume was calculated according to the following equation: V = L × W²/2 using Study Director version 3.1 (Studylog Systems, Inc.). For pharmacodynamic studies, HCT116 tumor–bearing mice were size matched and given oral dose(s) of A-1307138. Tumor samples were collected at various time points following dosing and were flash frozen in liquid nitrogen. Tumors were homogenized and NADt was measured using the NAD⁺/NADH Quantification Kit from MBL International Corporation.

To identify potential novel anticancer targets, a cell proliferation screen was performed to simultaneously identify oncology-relevant biologic activity (i.e., antiproliferative activity) and small molecules responsible for the activity of interest. The compounds could then be used to identify the target through affinity capture of the inhibitor/target complex. One cluster of compounds exhibited activity in five of the eight cell lines tested (Supplementary Fig. S2A). Inhibitor affinity capture studies of a novel compound, the isoindoline urea A-933414, from the hit cluster revealed NAMPT as the major biochemical target (Supplementary Fig. S2B and S2C). Inhibition of PC3 cell viability by A-933414 was rescued by coincubation of cells with the product of NAMPT, NMN, which overcomes intracellular NAMPT inhibition by virtue of extracellular dephosphorylation to nicotinamide riboside and intracellular conversion to NAD⁺ via the nicotinamide riboside kinase pathway (42). Rescue by NMN indicated that NAMPT was solely responsible for the antiproliferative activity of this chemotype (Supplementary Fig. 2D; Supplementary Table S2). These experiments also suggest that these compounds are exquisitely specific for NAMPT, as little to no antiproliferative activity was observed at concentrations 1,000-fold higher in the presence of NMN.

Extensive SAR work on this series indicated that structural modifications to several portions of the molecule provide improved potency, physical properties, and reduced clearance, resulting in significant murine exposure after oral dosing (Supplementary Table S3) without compromising selectivity toward NAMPT (manuscript in preparation; ref. 31). A lead compound from this series, A-1293201 (Fig. 1A), was found to potently inhibit recombinant, human NAMPT in an enzyme activity assay (Fig. 1B). Examination of the interactions between A-1293201 and NAMPT by X-ray crystallography revealed several distinct sites for cooperative interactions (Fig. 1C). The sites include the PRPP-binding subsite A, which is hydrophilic in nature and unoccupied by nicotinamide; the nicotinamide binding subsite B, where important pi-stacking and hydrogen bond interactions with nicotinamide are made; the tether subsite C, which is narrow, lipophilic, and not occupied by bound nicotinamide; and the distal opening subsite D, which widens and provides several bound waters and hydrophobic surfaces. Similar to FK866, the isoindoline urea portion of A-1293201 is engaged in important pi-stacking interactions in subsite B as well as hydrogen bonds and hydrophobic interaction in subsite C. In contrast to FK866 as well as recent second-generation NAMPT inhibitors, the isoindoline urea compounds are the first known potent nonphosphoribosylated NAMPT inhibitors.

To further characterize the ramifications of having inhibitors that do not serve as substrates to NAMPT, a nucleophilic nitrogen was added to the nicotinamide-mimetic portion of these compounds that occupied subsite B, producing potent and selective azaisoindoline urea inhibitors that could be phosphoribosylated by NAMPT (Supplementary Table S2). A-1267211, the azaisoindoline analogue of A-933414, and a racemic mixture of A-1331597, the azaisoindoline analogue of A-1293201, were both shown to be phosphoribosylated after incubation with NAMPT enzyme under conditions in which nicotinamide is also phosphoribosylated (Supplementary Fig. S3A). Phosphoribosylation of A-1267211 and another azaisoindoline, A-1307138, also occurred following treatment of PC3 cells with the compounds for 24 hours (Supplementary Fig. S3B).

A set of matched pairs of inhibitors that differ only in the presence or absence of the nitrogen at position 5 of the isoindoline ring was used for further analysis (Supplementary Table S4). The pairs of isoindoline and azaisoindoline ureas were tested in PC3 cells for inhibition of cell viability (Fig. 2). In most cases, the azaisoindoline version of each compound pair exhibited greater cellular potency than the isoindoline counterpart, ranging from 1.2- to 48-fold improvement in activity (Fig. 2). Although the isoindolines tended to be less potent in cells, many inhibited NAMPT and caused cytotoxicity with IC₅₀s below 50 nmol/L, and about a quarter of the isoindolines had similar cellular potencies as their azaisoindoline counterpart. In contrast, removal of the nitrogen from the pyridyl ring of FK866 completely abolished cellular potency (Fig. 2; Supplementary Tables S2 and S4).

Representative compounds from the isoindoline and azaisoindoline series were further characterized in biochemical and cellular assays to identify other distinguishing features. Compounds tested included the azaisoindoline/isoindoline pairs of A-1326133/A-1292945 and A-1331597/A-1293201 (Supplementary Table S2). The isoindoline of each pair showed similar potency in NAMPT enzyme assays, and A-1331597 and A-1293201 also had comparable IC₅₀s in PC3, HCT116, and NCI-H1975 cell viability assays. In contrast, the substrate compound A-1326133 was more potent than its corresponding isoindoline in the viability assays. Inhibition of PC3 cell viability by all of the compounds was rescued by coincubation of cells with NMN (Supplementary Table S2).

Effects of A-1293201, A-1331597, A-1292945, and A-1326133 on depletion of NAD⁺/NADH and ATP in HCT116 colorectal cancer cells over time were also measured. Inhibition of NAMPT by all four compounds resulted in maximal NADt (levels of NADH plus NAD⁺) depletion within 24 hours and ATP depletion within 48 to 72 hours (Fig. 3A–D). Following NADt and ATP depletion, HCT116 cell viability was severely compromised within 72 hours. ATP depletion and inhibition of cell viability only occurred at compound concentrations that resulted in greater than 90% reduction in NADt levels at 48 hours. If there is less than 90% depletion of NADt, the NADt levels recover and cells survive. These data suggest that cytotoxicity elicited by NAMPT inhibition is dependent on the kinetics of ATP depletion, which occurs subsequent to NADt depletion.

Comparison of NADt depletion dose–response curves after 24-hour treatment for the azaisoindoline/isoindoline pairs is shown in Fig. 3E. Although the IC₅₀ for NADt depletion for A-1293201 was 1.8 times lower than for A-1331597, both compounds depleted NADt by >90% at 37 nmol/L. In contrast, the IC₅₀ and the maximal NADt depletion for A-1326133 were both lower than for A-1292945. The maximal NADt depletion occurred at 4.1 nmol/L for A-1326133, but for A-1292945, NADt was not depleted >90% until 12.3 nmol/L. NADt recovery following washout of the compounds was also determined by treating HCT116 cells with the NAMPT inhibitors for 24 hours, removing the compound and washing the cells, and measuring NADt levels over time. The NADt recovery was plotted for all four compounds at the concentrations at which NADt was depleted by >90% at 24 hours before washout (Fig. 3F). The kinetics of NADt recovery was similar for A-1293201 and its partner, A-1331597, as well as for the isoindoline A-1292945. For these compounds, NADt had recovered by 50% around 4 hours after washout. However, recovery of NADt following washout of A-1326133 was delayed and did not reach 50% until 8 hours. By 16 hours following washout, NADt levels had fully recovered for all four of the compounds.

To determine the effects of A-1293201 and A-1326133 on the NAD⁺ metabolome and glycolysis, NCI-H1975 cells were treated with either vehicle (0.1% DMSO) or three concentrations (IC₂₀, IC₅₀, or IC_>90) of the compounds or FK866 for 8, 24, or 48 hours. The compound concentrations and incubation times were chosen based on the kinetics of NADt and ATP depletion in NCI-H1975 cells (Supplementary Fig. S4). After 24 hours and at concentrations matching the respective IC₅₀ and IC_>90, each compound significantly depressed NMN and NAD⁺ (Fig. 4). At 24 hours, ADP ribose (ADPr) was decreased significantly by treatment with each compound at IC_>90, while NADP⁺ tended to be lowered (Fig. 4). However, the data indicate that NMN, NADP⁺, and ADPr all strongly correlated with NAD⁺ in each experiment and all trended down together. The pattern observed at 24 hours continued at 48 hours (Fig. 4). The effects of FK866, A-1293201, and A-1326133 on glycolytic metabolites were also measured, and all three compounds caused similarly increased levels of fructose-6-phosphate/glucose-6-phosphate, fructose-1,6-bisphosphate, and dihydroxyacetone phosphate/glycedraldehyde-3-phosphate (Fig. 5), which is consistent with inhibition of glycolysis via NAD⁺ depletion previously reported for FK866 (43). Together, these data confirm that A-1293201 and A-1326133 target NAD⁺ metabolism, which subsequently depresses glycolysis.

The evaluation of the time of inhibition requirement for cytotoxicity identified by the in vitro time course studies was extended to an in vivo setting using A-1307138, an analogue of A-1326133 (Supplementary Table S2). Following a single oral dose of 15 mg/kg in an HCT116 xenograft tumor model, A-1307138 treatment reduced tumor NADt levels by greater than 90% after 24 hours, but NADt concentrations quickly recovered to approximately 50% of pretreatment levels 72 hours postdose (Fig. 6A). Notably, transient depletion of NADt levels by >90% for less than 24 hours once a week did not elicit any suppression of tumor growth (Fig. 6C). Extending the duration of NADt depletion by 2 to 3 consecutive days of dosing resulted in substantial inhibition of tumor growth in vivo (Fig. 6C), which corresponds to sustained NADt depletion, with >90% depletion maintained for over 72 hours (Fig. 6B). These data are consistent with the in vitro observations that inhibition of NAMPT for at least 48 hours is required to elicit NADt depletion–dependent cytotoxicity. It was determined that maintaining the time-over-threshold (IC₉₀) for 48 to 70 hours was required for robust antitumor efficacy. This is exemplified by the potent antitumor efficacy of A-1293201 and A-1326133 in the HCT116 xenograft model (Fig. 6D). Body weight loss did not exceed 10% for any of the compounds, and no deaths occurred during the in vivo studies (Supplementary Fig. S5).

We have identified and characterized novel potent and selective nonsubstrate and substrate NAMPT inhibitors. Both types of compounds exhibit potent in vitro and in vivo antiproliferative activity following oral administration and display similar kinetics of NADt and ATP depletion and induction of cytotoxicity, which is rescued by NMN. On the basis of mechanistic studies, inhibition of NAMPT by >90% for at least 48 hours is required for antiproliferative effects in HCT116 cells, in vitro and in vivo. As has been shown for previously published NAMPT inhibitors, within hours after treatment, NAD⁺ levels are decreased, but 24 to 48 hours are required for complete NAD⁺ depletion within the mitochondria and blockade of ATP generation. Once ATP is depleted, cell death occurs through apoptosis, autophagy, or oncosis (10, 44). The kinetics of NAD⁺ depletion can differ between cell types, possibly due to differences in NAD⁺ consumption rates by PARPs or other enzymes that utilize NAD⁺ as a substrate. The kinetics of NAD⁺ and ATP depletion in NCI-H1975 cells are slower than in HCT116 cells. As a result, NAD⁺ levels are not depleted >90% until between 36 and 48 hours and inhibition of glycolysis and ATP depletion are also delayed. A-1293201 and A-1326133 produce similar effects on the NAD⁺ metabolome and glycolytic metabolites as FK866. All of these properties indicate that the isoindoline and azaisoindoline compounds cause cell death by inhibiting NAMPT in cells and xenografts.

In contrast to previously described NAMPT inhibitors, including FK866, GMX1778, and GNE-617, A-1293201 does not contain an aromatic nitrogen in its nicotinamide-mimetic group (Fig. 1A). Crystallographic studies indicate that A-1293201 participates in similar interactions with NAMPT as the other inhibitors (45, 46). A-1293201 and A-1326133 occupy the nicotinamide-binding site and engage in a critical pi-stacking interaction with tyrosine 18 in the same manner as nicotinamide and FK866 (Fig. 1C; Supplementary Fig. S6). The central regions of all the compounds traverse a narrow, lipophilic tunnel out to an opening distal to the active site. The nicotinamide-mimetic moiety, where the presence of the aromatic nitrogen allows for phosphoribosyl adduct formation, is the area of divergence between A-1293201 and the other compounds. Our data indicate that phosphoribosylation of inhibitors is not required for potent inhibition of NAMPT and induction of cell death in cells or in xenograft models.

It is notable that most azaisoindolines display improved potency in cellular assays compared with their isoindoline analogues, even though their permeability and protein-binding properties are similar. One hypothesis is that the aromatic nitrogen in the azaisoindoline results in improved pi-stacking interactions with NAMPT and/or serves as a substrate, leading to increased target potency. However, similar IC₅₀s in enzyme assays for the azaisoindoline/isoindoline pairs of A-1326133/A-1292945 and A-1331597/A-1293201 suggests that improved pi-stacking interactions are not occurring, and these data are corroborated by the overlaid crystal structures of A-1293201 and A-1326133 (Supplementary Fig. S6). In addition, there does not appear to be a difference in dissociation rates between A-1293201 and A-1331597 or between A-1326133 and A-1292945 in TR-FRET assays (Supplementary Fig S7), even when the compounds are preincubated with the enzyme and PRPP for 24 hours to allow phosphoribosylation. These results suggest that for this set of compounds, phosphoribosylation does not lead to increased residence time on the enzyme. In these assays, both substrate and nonsubstrate compounds have slower off rates when PRPP is bound, indicating that both classes of compounds bind cooperatively with PRPP. These data suggest that the increased cellular potency of azaisoindolines relative to isoindolines is not the result of increased target affinity upon phosphoribosylation.

Another potential reason for the increased cellular potency of the azaisoindolines is that phosphoribosylated compounds are retained longer in cells due to decreased transit of the modified compound through the plasma membrane. Increased cellular retention was reported to contribute to cellular potency for GMX1778, which undergoes phosphoribosylation (28). In our washout studies, recovery of NADt levels is similar between A-1293201 and A-1331597, suggesting that A-1331597 is not retained in cells. Thus, although phosphoribosylation can improve the cellular potency for some compounds within the isoindoline series, in general, there is not a requirement for modification by NAMPT for potent inhibition, and enhanced binding can be attained through additional interactions distal to the nicotinamide-binding site.

Our discovery of potent and selective NAMPT inhibitors that do not contain an aromatic nitrogen in the nicotinamide moiety challenges the current dogma that this feature is required for cellular efficacy. A-1293201 and other isoindolines are efficacious despite the lack of a phosphoribosylation site, which could lead to cellular retention or other unexpected effects in cells. As a result, treatment with A-1293201 or other isoindolines may allow better control of NAMPT inhibition in normal cells.

J.L. Wilsbacher has ownership interest (including patents) in AbbVie, Inc. Y. He is an employee at AbbVie, Inc. D. Maag is a full-time employee at and has ownership interest (including patents) in AbbVie. G.G. Chiang is the senior group leader/principal research scientist at AbbVie, is a director at eFFECTOR Therapeutics, reports receiving a commercial research grant from AbbVie, and has ownership interest (including patents) in AbbVie. S.H. Rosenberg is an employee at and has ownership interest (including patents) in AbbVie. C. Brenner is in the scientific advisory board at ChromaDex, Inc., is the co-founder of ProHealthspan, and has ownership interest (including patents) in intellectual property around uses of nicotinamide. No potential conflicts of interest were disclosed by the other authors.

Conception and design: J.L. Wilsbacher, M. Cheng, D. Cheng, S.A.J. Trammell, Y. Shi, J. Guo, B.K. Sorensen, R.F. Clark, T.M. Hansen, M.L. Curtin, M.R. Michaelides, D. Maag, F.G. Buchanan, G.G. Chiang, C. Tse

Development of methodology: J.L. Wilsbacher, M. Cheng, D. Cheng, Y. Shi, J. Guo, S.L. Koeniger, P.J. Kovar, Y. He, A.V. Korepanova, P.L. Richardson, S.M. McLoughlin, D. Maag, C. Brenner, C. Tse

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): J.L. Wilsbacher, M. Cheng, D. Cheng, S.A.J. Trammell, Y. Shi, S.L. Koeniger, P.J. Kovar, Y. He, S. Selvaraju, K.L. Longenecker, A.V. Korepanova, S. Cepa, D.L. Towne, V.C. Abraham, H. Tang, S.M. McLoughlin, I. Badagnani, D. Maag, C. Brenner

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): J.L. Wilsbacher, M. Cheng, D. Cheng, S.A.J. Trammell, Y. Shi, J. Guo, S.L. Koeniger, P.J. Kovar, Y. He, K.L. Longenecker, A.V. Korepanova, S. Cepa, V.C. Abraham, S.M. McLoughlin, I. Badagnani, D. Maag, G.G. Chiang, W. Gao, C. Brenner

Writing, review, and/or revision of the manuscript: J.L. Wilsbacher, M. Cheng, D. Cheng, S.A.J. Trammell, Y. Shi, J. Guo, S.L. Koeniger, P.J. Kovar, Y. He, R.F. Clark, K.L. Longenecker, D. Raich, A.V. Korepanova, V.C. Abraham, H. Tang, P.L. Richardson, I. Badagnani, M.L. Curtin, M.R. Michaelides, D. Maag, G.G. Chiang, W. Gao, S.H. Rosenberg, C. Brenner, C. Tse

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): J.L. Wilsbacher, M. Cheng, D. Cheng, Y. Shi, P.J. Kovar, H. Tang, G.G. Chiang

Study supervision: G.G. Chiang, C. Tse

Other (chemical design and synthesis of NAMPT inhibitors): H.R. Heyman

The authors would like to thank Jameel Shah for his contributions to the initial screen design and validation of hits, Keith Glaser for experimental advice, and Niru Soni and Junling Li for their expert technical assistance. The authors would also like to acknowledge Stella Doktor, Patricia Stuart, Amanda Olson, Donald Osterling, and DeAnne Stolarik from the DMPK-BA Department at AbbVie for the in vitro ADME and in vivo pharmacokinetics measurements. Research described in this article was performed using beamline 08ID-1 at the Canadian Light Source, which is supported by the Canada Foundation for Innovation, Natural Sciences and Engineering Research Council of Canada, the University of Saskatchewan, the Government of Saskatchewan, Western Economic Diversification Canada, the National Research Council Canada, and the Canadian Institutes of Health Research. Use of the IMCA-CAT beamline 17-ID at the Advanced Photon Source was supported by the companies of the Industrial Macromolecular Crystallography Association through a contract with Hauptman-Woodward Medical Research Institute. This research used resources of the Advanced Photon Source, a U.S. Department of Energy (DOE) Office of Science User Facility operated for the DOE Office of Science by Argonne National Laboratory under contract no. DE-AC02-06CH11357.

This work was financially supported by AbbVie, Inc.

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