Chemistry
Melting points were determined on a Mettler FP1 melting point apparatus and are uncorrected. IR spectra were recorded neat on a Bruker Vector 33FT-infrared spectrometer. 1H-NMR and 13CNMR spectra were measured on a Bruker DRX-500 (500 MHz). Chemical shifts are given in parts per million (ppm), and TMS was used as an internal standard for spectra obtained in CDCl3. All coupling constants (J) are given in Hz. ESI (electrospray ionization) mass spectra were determined on a TSQ quantum (Thermo Electron Corporation) instrument. High-resolution mass spectra were measured using an LTQ Orbitrap (Thermo Electron Corporation) with positive ESI.Ketones and precursors of ketones used as starting materials were commercially obtained.
Method A: Triflation
To a solution of 2,6-di-tert-butyl-4-methylpyridine (7.23 mmol) and the corresponding ketone (6.57 mmol) in CH2Cl2 (20 mL) was added trifluoromethanesulfonic anhydride (7.16 mmol) dropwise at 0uC under nitrogen. A white precipitate was formed thereafter. The reaction mixture was warmed to room temperature and further stirred for 4 h. The solvent was removed in vacuo, and the resulting residue was diluted with petroleum ether (20 mL). After removal of the solid by filtration, the filtrate was concentrated in vacuo to give a crude product which was used in the next step without further purification.
Method B: Suzuki Coupling
A suspension of enol triflate (4.56 mmol), pyridine-3-boronic acid (5.92 mmol), sodium carbonate (22.8 mmol) and tetrakis(triphenylphosphine)palladium (0) (0.23 mmol) in dimethoxyethanol (45 mL) and water (15 mL) was stirred at 90uC under nitrogen for 2 h. The reaction mixture was cooled to room temperature slowly and diluted with water (10 mL). The aqueous layer was extracted with ethyl acetate (2630 mL) and the combined organic layers were washed with brine twice and dried over MgSO4. After evaporation in vacuo, the resulting residue was purified by flash chromatography to afford the corresponding product.Method C: Hydrogenation A mixture of 3-pyridyl derivative with an a-double bond (0.50 mmol) and 5% Pd/C (45 mg) in methanol (15 mL) was flushed with hydrogen for 2 d at room temperature The mixture was filtered through a Celite cake and concentrated in vacuo to afford the corresponding product.
Abstract
The major human apurinic/apyrimidinic endonuclease APE1 plays a pivotal role in the repair of base damage via participation in the DNA base excision repair (BER) pathway. Increased activity of APE1, often observed in tumor cells, is thought to contribute to resistance to various anticancer drugs, whereas down-regulation of APE1 sensitizes cells to DNA damaging agents. Thus, inhibiting APE1 repair endonuclease function in cancer cells is considered a promising strategy to overcome therapeutic agent resistance. Despite ongoing efforts, inhibitors of APE1 with adequate drug-like properties have yet to be discovered. Using a kinetic fluorescence assay, we conducted a fully-automated high-throughput screen (HTS) of the NIH Molecular Libraries Small Molecule Repository (MLSMR), as well as additional public collections, with each compound tested as a 7-concentration series in a 4 mL reaction volume. Actives identified from the screen were subjected to a panel of confirmatory and counterscreen tests. Several active molecules were identified that inhibited APE1 in two independent assay formats and exhibited potentiation of the genotoxic effect of methyl methanesulfonate with a concomitant increase in AP sites, a hallmark of intracellular APE1 inhibition; a number of these chemotypes could be good starting points for further medicinal chemistry optimization. To our knowledge, this represents the largest-scale HTS to identify inhibitors of APE1, and provides a key first step in the development of novel agents targeting BER for cancer treatment.
Citation: Dorjsuren D, Kim D, Vyjayanti VN, Maloney DJ, Jadhav A, et al. (2012) Diverse Small Molecule Inhibitors of Human Apurinic/Apyrimidinic Endonuclease APE1 Identified from a Screen of a Large Public Collection. Editor: Gayle E. Woloschak, Northwestern University Feinberg School of Medicine, United States of America Received May 2, 2012; Accepted September 25, 2012; Published October 23, 2012 This is an open-access article, free of all copyright, and may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose. The work is made available under the Creative Commons CC0 public domain dedication. Funding: This research was supported in part by the Molecular Libraries Initiative of the National Institutes of Health (NIH) Roadmap for Medical Research (grants 1 R03 MH092154-01 (DMWIII) and U54MH084681 (AS)) and the Intramural Research Program of NIA, NIH. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. Competing Interests: The authors have declared that no competing interests exist.
Introduction
The genome of mammalian cells is under constant threat from both endogenous (namely reactive oxygen species, such as the superoxide anion, hydroxyl radical, hydrogen peroxide, and nitrogen-reactive species) and exogenous (e.g., sunlight, ionizing radiation, chemical compounds and genotoxic drugs) DNA damaging agents that can introduce mutagenic and cytotoxic DNA lesions [1,2]. For example, it has been estimated that spontaneous depurination events result in more than 10,000 abasic lesions per mammalian cell per day [3,4]. Left unrepaired, DNA damage can result in detrimental biological consequences to the organism, including cell death and mutations that drive transformation to malignancy. Cells use various DNA repair systems as defenses to protect their genomes from DNA damaging agents and to maintain genome stability [5,6,7]. Not surprisingly, cells with a defect in one of their DNA repair mechanisms are typically more sensitive to certain genotoxic agents and suffer increased mutagenesis. Most antitumor drugs (e.g., alkylating, cross-linking and intercalating agents, topoisomerase inhibitors, and certain antimetabolites) induce DNA lesions that ultimately block or interfere with DNA replication in rapidly dividing cancer cells, resulting in increased susceptibility to activation of various programmed cell death responses [8]. An elevated DNA repair capacity in tumor cells results in anticancer drug and radiation resistance, severely limiting the efficacy of these agents. Recent basic and clinical studies have demonstrated emerging concept designs to block the functions of various proteins in specific DNA repair pathways, which would sensitize cancer cells to DNA damaging agents and potentially lead to an improved therapeutic outcome [9,10]. The base excision repair (BER) pathway is responsible for correcting damage to single DNA bases or to the sugar moiety of the phosphodiester backbone. Typically, the BER process starts with the enzymatic removal of a damaged base by either a monoor a bi-functional DNA glycosylase, which creates an abasic (AP) site or in some instances a DNA strand break. The AP site is incised by an essential enzyme known as apurinic/apyrimidinic endonuclease-1 (APE1) [11], which generates a single-stranded gap in DNA with 39-hydroxyl and 59-deoxyribosephosphate termini. This gap is filled in and ultimately sealed by the concertedaction of DNA polymerases and ligases [4]. In mammalian cells, APE1 is responsible for at least 95% of the endonuclease activity that incises at abasic sites as part of the short-patch and long-patch BER subpathways.
