Exploiting the Inherent Photophysical Properties of the Major

Article Cite This: J. Org. Chem. 2018, 83, 3126−3131

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Exploiting the Inherent Photophysical Properties of the Major Tirapazamine Metabolite in the Development of Profluorescent Substrates for Enzymes That Catalyze the Bioreductive Activation of Hypoxia-Selective Anticancer Prodrugs Xiulong Shen,† Charles H. Laber,† Ujjal Sarkar,† Fabio Galazzi,§ Kevin M. Johnson,† Nathaniel G. Mahieu,† Roman Hillebrand,† Tarra Fuchs-Knotts,† Charles L. Barnes,† Gary A. Baker,† and Kent S. Gates*,†,‡ †

Department of Chemistry, ‡Department of Biochemistry, and §Molecular Interaction Core, University of Missouri 125 Chemistry Building Columbia, Missouri 65211, United States S Supporting Information *

ABSTRACT: Hypoxia-selective cytotoxins (HSCs) seek to exploit the oxygenpoor nature of tumor tissue for therapeutic gain. Typically, HSCs require activation by one-electron bioreductive enzymes such as NADPH:cytochrome P450 reductase (CYPOR). Thus, successful clinical deployment of HSCs may be facilitated by the development and implementation of diagnostic probes that detect the presence of relevant bioreductive enzymes in tumor tissue. The work described here develops analogues of the well-studied HSC tirapazamine (3amino-1,2,4-benzotriazine 1,4-di-N-oxide, TPZ) as profluorescent substrates of the one-electron reductases involved in bioactivation of HSCs. Hypoxic metabolism of TPZ or 7-fluoro-TPZ by one-electron reductases releases inherently fluorescent mono-N-oxide metabolites that may serve as indicators, probes, markers, or stains for the detection of the enzymes involved in the bioactivation of HSCs. In particular, profluorescent compounds of this type can provide a foundation for fluorescence-based bioassays that help identify tumors responsive to HSCs.

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well-studied HSC tirapazamine (TPZ, 1a, Scheme 1).24,25 TPZ undergoes intracellular enzymatic one-electron reduction10 to yield an oxygen-sensitive drug radical 2 that selectively decomposes under hypoxic conditions to release a cytotoxic DNA-damaging drug radical or hydroxyl radical (HO•).26−31

INTRODUCTION Most tumors contain regions of poorly oxygenated (hypoxic) tissue.1,2 Hypoxia-selective cytotoxins (HSC) are a class of chemotherapeutic agents that seek to exploit tumor hypoxia for therapeutic gain.3−6 Typically, these compounds are activated by one-electron enzymatic reductases to generate a critical oxygen-sensitive drug radical anion.3−5,7,8 Many different enzymes can be involved in these single-electron reduction processes.3,9,10 In aerobic tissue, back-oxidation of the radical anion to the starting drug is kinetically favored, while in an oxygen-poor environment, the extended lifetime of the radical intermediate enables reactions that convert the parent drug to a cytotoxic agent.3−5,7,11 There are at least two absolute requirements for efficacious use of HSCs: (i) the tumor tissue must be hypoxic and (ii) the tumor tissue must express reductase enzyme(s) required for bioactivation of the drug. Successful clinical deployment of HSCs may be facilitated by the development and implementation of diagnostic probes that detect hypoxia and the bioreductive enzymes required for drug activation in tumor tissue.6,12−23 Here, we describe efforts toward the development of profluorescent enzyme substrates for the detection of the one-electron bioreductive enzymes involved in the activation of HSCs. The substrates developed here are derivatives of the © 2018 American Chemical Society

Scheme 1. Hypoxic Metabolism of TPZ (1a) and Analogues

Received: December 1, 2017 Published: February 20, 2018 3126

DOI: 10.1021/acs.joc.7b03035 J. Org. Chem. 2018, 83, 3126−3131

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The Journal of Organic Chemistry

results indicated that these analogues displayed hypoxiaselective cytotoxicity (and presumably cell permeability) against cancer cells in culture.32 Fluorescence Properties of 1,2,4-Benzotriazine Oxides 1a−c and 4a−c. We next undertook more complete photophysical characterization of the dioxides 1b and 1c and their putative mono-N-oxide metabolites 4b and 4c, in comparison to TPZ (1a) and its metabolite 4a (Table 1). The dioxides 1a−c were synthesized as shown in Scheme 2 via oxidation of the mono-N-oxide derivatives with a mixture of trifluoroacetic anhydride and H2O2.32,35,40

Many 1,2,4-benzotriazine 1,4-dioxides with analogous properties have been identified.32−34 Importantly, hypoxic metabolism of 1,2,4-benzotriazine 1,4-dioxides typically generates the corresponding 1-oxide (e.g., 4) as the major metabolite.26,27,35−37 Our work exploits the fortuitous fluorescence properties of the 1-oxide metabolite generated by TPZ.38,39 We explore the use of TPZ and other 1,2,4-benzotriazine 1,4dioxide analogues as profluorescent substrates for the selective detection of one-electron reductases involved in bioreductive activation of HSCs. We synthesized and studied the florescence properties of 25 1,2,4-benzotriazines and 1,2,4-benzotriazine 1-oxides leading to the identification of two analogues (7-fluoro, 4b and 7-OCH3, 4c) with brightness superior to the TPZ metabolite 4a. The photophysical properties including the fluorescence quantum yields, absorption/emission spectra, and Stokes shifts of 4b and 4c were determined and compared to 4a. LC−MS assays were used to characterize the enzymatic conversion of the corresponding dioxides 1a−c to the 1-oxide metabolites 4a−c under aerobic and hypoxic conditions. Fluorescence assays were used to detect the enzymatic generation of the fluorescent metabolites 4a−c. We found that hypoxic metabolism of TPZ (1a) or 7-fluoro-TPZ (1b) by one-electron reductases releases easily detected fluorescent 1-oxide metabolites that may serve as indicators, probes, markers, or stains for the detection of enzymes involved in the bioactivation of HSCs.

Table 1. Static Photophysical Properties of 1,2,4Benzotriazine Oxides

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RESULTS AND DISCUSSION Synthesis and Fluorescence Properties of 1,2,4Benzotriazine 1-Oxides 4. A collection of 25 1,2,4benzotriazine 1-oxides was synthesized by published routes and minor modifications of published routes (Scheme 2).32,35,40−43 The compounds in the collection contained

compd

λabs (nm)

log10 εa

λemb (nm)

4ad 4bd 4cd 4ae 4be 4ce 1ad 1bd 1cd 1ae 1be 1ce

407 415 430 415 422 441 477 489 432 457 468 440

3.747 3.807 3.646 3.751 3.749 3.655 3.867 3.853 3.684 3.786 3.807 3.765

478 491 509 521 526 547 505 573 510 545 562 542

Φflc (%) 16.86 29.94 47.22 3.67 2.44 8.68 0.60 0.77 2.27 0.33 0.30 1.24

± ± ± ± ± ± ± ± ± ± ± ±

1.40 2.09 5.07 0.21 0.21 1.00 0.01 0.02 0.35 0.01 0.01 0.16

ε in units of L·mol−1·cm−1. bFor excitation at 421 nm. cDetermined using coumarin 153 (C153) in ethanol (Φref = 38%) as the fluorescence quantum yield standard. dMeasured in acetonitrile. e Measured in water. a

The extinction coefficients for all compounds 1a−c and 4a− c varied by only about 30% (Table 1). The dioxides 1a−c displayed weak fluorescence in water and various nonaqueous solvents, with quantum yields in the range of 0.6−2.2% in acetonitrile and 0.3−1.2% in water (Table 1). The mono-Noxides displayed substantially stronger fluorescence quantum yields in the range of 16−47% in acetonitrile and 2−8% in water (Table 1). The mono-N-oxides 4a−c displayed large Stokes shifts, with separations between the absorption and emission maxima of 71, 76, and 79 nm in acetonitrile and 104, 106, and 106 nm in water for 4a−c, respectively (Table 1 and Figures S2−4). The absorption/emission maxima in water for the 7-fluoro analogue 4b (422/526 nm) and the 7-OCH3 derivative 4c (441/547 nm) were located at longer wavelengths than those for the TPZ metabolite 4a (415/521 nm). Longer emission wavelengths are preferred for biological and biochemical applications to avoid interference arising from autofluorescence of biological materials such as flavins and NADPH.44 The fluorescence brightness (Φε) of the 7-fluoro analogue 4b (1917 M−1 cm−1) is 2.0-fold larger and the 7OCH3 derivative 4c (2089 M−1 cm−1) 2.2-fold larger than the TPZ metabolite 4a (944 M−1 cm−1) in acetonitrile. Given the intended use of these compounds in biochemical systems, we examined the effects of added protein on the photophysical properties of 4b and 4c. We found that the fluorescence emission maxima and quantum yields for these two compounds are not markedly influenced by the presence of the protein bovine serum albumin (BSA), a plasma protein which displays ubiquitous nonspecific binding with small molecules (Figure 1).45

Scheme 2. Synthesis of TPZ and Analogues

various electron-donating and electron-withdrawing substituents. We performed preliminary measurements to determine whether any of the compounds displayed desirable photophysical properties such as high fluorescence quantum yields (Φ), large extinction coefficients (ε), and large brightness values (Φε) relative to the TPZ metabolite 4a. This survey revealed that most analogues had brightness values lower than that of 4a (Figure S1) We did, however, identify a small number of analogues with fluorescence brightness comparable to 4a (Figure S1). Of these, 4b and 4c were selected for further study based on their brightness, the anticipated synthetic accessibility of the requisite 1,4-dioxide analogues, the retention of fluorescence properties in water, and the fact that previous 3127

DOI: 10.1021/acs.joc.7b03035 J. Org. Chem. 2018, 83, 3126−3131

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The Journal of Organic Chemistry

Figure 1. Fluorescence quantum yields and emission maxima measured for 4b and 4c in water in the presence of increasing levels of bovine serum albumin (BSA). Error bars depict standard deviation in the measurements.

LC−MS Analysis of in Vitro Hypoxic Metabolic Conversion 1a−c to the Corresponding Mono-N-oxides 4a−c. Having determined that 4b and 4c possess promising photophysical properties, it was important to examine whether the corresponding di-N-oxides 1b and 1c undergo the expected hypoxia-selective, enzymatic conversion to the fluorescent monoxide metabolites 4b and 4c. We expected clean conversion to the mono-N-oxides but recognized that this expectation was based on results from only a handful of 1,2,4benzotriazine 1,4-dioxide analogues for which metabolism had been studied.26,27,35−37,46 For these assays, we used NADPH:cytochrome P450 oxidoreductase (CYPOR) and xanthine oxidase (XO) because these enzymes are commonly involved in one-electron bioreductive activation of HSCs both in vivo and in vitro.10,13,26,28,31,47−50 We used HPLC and LC−MS analysis to characterize the in vitro metabolism of 1b and 1c under aerobic and hypoxic conditions, in comparison with the parent compound 1a (Figures 2 and 3 and Figures S5 and S6). Hypoxic assays were carried out in an inert atmosphere glovebag using degassed solutions. We found that incubation of 1a−c (500 μM) with CYPOR (0.13 units/mL) and NADPH (2 mM, 4 equiv) in sodium phosphate buffer (50 mM, pH 7) under hypoxic conditions for 4 h at 24 °C generated substantial yields of the monoxide metabolites 4a−c (Figure 2). Representative LC−MS data for the in vitro metabolism of 1b are shown in Figure 3. The retention times and mass spectra of the metabolites 4 matched those of authentic monoxides prepared by chemical synthesis. In an identical assay, except under aerobic conditions, in vitro metabolism of 1a produced an 18-fold lower yield of the mono-N-oxide 4a compared to the hypoxic assay (Figure 2). The metabolism of 1b was similarly inhibited under aerobic conditions, giving an 8-fold lower yield of the mono-N-oxide 4b. In contrast, the metabolism of 1c proved to be poorly selective for hypoxia. Indeed, 1c underwent large amounts of aerobic metabolism, relative to 1a and b. Similar results were observed for the enzyme XO (Figure 2). Possible reasons for the aerobic metabolism of 1c are considered in the Conclusions. We also examined the in vitro metabolism of 1a−c by the obligate two-electron reductase NADPH:quinone oxidoreductase (DT-diaphorase)51 and by NADPH alone. We found that 1a and 1b were not metabolized to a significant extent under these conditions. On the other hand, NADPH alone and in concert with DT-diaphorase mediated significant conversion of 1c to 4c (Figure 2). Control experiments showed that the DTdiaphorase system was effective in reducing the archetypal

Figure 2. Yields of mono-N-oxides (4a−c) generated by in vitro metabolism of 1a, 1b, and 1c under various conditions. The standard deviation was calculated from two replicates.

substrates 1,4-naphthoquinone and 2,3-dimethoxy-1,4-naphthoquinone. Fluorimetric Analysis of the in Vitro Metabolic Conversion of 1a−c to 4b−c. Compounds 1a−c were incubated with CYPOR, XO, or DT-diaphorase enzyme systems under aerobic or hypoxic conditions, the reaction mixtures extracted with ethyl acetate, and the fluorescence of the extracts determined at the excitation/emission maxima of the corresponding mono-N-oxide metabolite (4a-c) anticipated for each reaction. Compounds 1a and 1b gave easily detected fluorescence signals in the presence of the one-electron reducing enzymes CYPOR or XO under hypoxic conditions. Relatively weak fluorescence output was produced by incubation of 1a and 1b with CYPOR and XO under aerobic conditions or when DT-diaphorase or NADPH alone was used. The fluorescence signals produced by the 7-fluoro analogue 1b were 16-fold and 24-fold above background in the presence of XO and CYPOR enzyme systems, respectively (Figure 4). Qualitatively similar results were obtained by direct fluorimetric analysis in aqueous buffer (Figure S7). In addition, the dioxide 1b can be metabolized in a hypoxic cell extract and fluorescence from the resulting mono-N-oxide can be detected against background autofluorescence (Figure S8). In vitro metabolism of the 7-OCH3 derivative 1c by CYPOR and XO produced strong fluorescence signals under hypoxic conditions but also generated rather high fluorescence responses under aerobic conditions. More significantly, in vitro metabolism of 1c by NADPH alone and in the presence of DT-diaphorase produced substantial fluorescence under both hypoxic and aerobic conditions. The fluorescence results are 3128

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Figure 3. LC−MS analysis of the in vitro hypoxic metabolism of 1b catalyzed by NADPH/CYPOR: (A) total ion chromatogram of the aerobic assay; (B) total ion chromatogram of the hypoxic assay; (C) LC−MS data for 4b; (D) LC−MS data for 1b.

consistent with the HPLC and LC−MS analyses described above.

substitutent may favor protonation of the activated intermediate 2/3 and/or increase the fragmentation rate constant (kf, Scheme 1), leading to diminished oxygen-sensitivity in the production of the 1-oxide metabolite 1c.11,30,31 Reasons for the different hypoxia selectivities observed in our assays compared to the cell culture assays reported previously deserve further study.32 Overall, we found that hypoxic metabolism of the substrates 1a and 1b generated easily detected fluorescent metabolites that have the potential to serve as markers for the one-electron reductases required for the hypoxia-selective bioactivation of HSCs. Probes of this type enable simultaneous functional determination of the total one-electron enzymatic reducing capacity in a biological sample.18 This may provide a more direct and perhaps simpler approach to the assessment of oneelectron reductase activity than transcriptomic measurements of gene expression or proteomic measurements protein expression. The benzotriazine-based probes clearly are well positioned to detect the enzymes relevant to the one-electron bioactivation of 1,2,4-benzotriazine 1,4-dioxide HSCs.9,13 Importantly, compounds such as 1b could be generally applicable because the enzymes involved in bioactivation of 1,2,4-benzotriazine 1,4-dioxides are likely to be major players in the bioactivation of many HSCs.8,9,13 Thus, profluorescent compounds such as 1b may provide a foundation for fluorescence-based bioassays that aid in the identification of tumors responsive to HSCs.18

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CONCLUSIONS Profluorescent enzyme substrates are important in chemical biology and clinical diagnostics.52,53 Our work exploits the fortuitous fluorescence properties of the 1-oxide metabolite generated by hypoxic metabolism of the HSC agent TPZ.38,39 Indeed, we found that hypoxic metabolism of TPZ (1a) and its fluoro analogue 1b generates easily detectable fluorescent metabolites 4a and 4b. The fluoro-TPZ analogue 1b developed here generates substantially brighter fluorescence output than the parent compound TPZ (1a). In addition to its superior brightness, the metabolite 4b emits fluorescence at longer (green) wavelengths (491 nm in CH3CN and 529 nm in H2O), a feature which may be advantageous in biochemical and biological applications.44 Importantly, 1a and 1b are not substrates for the two-electron reductase DT-diaphorase. This observation is consistent with inferences drawn previously from studies of TPZ in cell culture.13 It is critical that probes of this sort respond selectively to the one-electron reductases involved in the oxygen-sensitive bioactivation of HSCs. The behavior of the 7-OCH3 analogue 1c was surprising. This compound was previously reported to display hypoxiaselective cytotoxicity similar to the 7-fluoro analogue 1b.32 Thus, we expected that both 1b and 1c would undergo hypoxia-selective metabolism by one-electron reductases and would be refractory to processing by a two-electron reductase like DT-diaphorase. As described above, the 7-fluoro derivative 1b met these expectations. In stark contrast, the 7-OCH3 analogue 1c underwent significant amounts of aerobic metabolism by the one-electron reductases CYPOR and XO and also was a substrate for the two-electron reductase DTdiaphorase. As a result, compound 1c will likely not be a particularly useful probe for the selective detection of oneelectron reductases. We posit that the electron-donating OCH3

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EXPERIMENTAL METHODS

Materials. Materials were of the highest purity available and were obtained from the following sources: NADPH:cytochrome P450 reductase, DT-diaphorase, NADPH, xanthine, 1,4-naphthoquinone, 2,3-dimethoxy-1,4-naphthoquinone, sodium phosphate, DMSO, silica gel (0.04−0.063 mm pore size), and silica gel plates for thin layer chromatography from Sigma Chemical Co. (St. Louis, MO); xanthine oxidase from Roche Applied Science (Indianapolis, IN); HPLC-grade solvents (acetonitrile, methanol, ethyl acetate, hexane, trifluoroacetic 3129

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These values are comparable to the brightness of these compounds in acetonitrile (4b, 1917 M−1 cm−1, and 4c, 2089 M−1 cm−1). LC−MS Analysis of the in Vitro Enzymatic Metabolism of 3Amino-1,2,4-benzotriazine 1,4-Dioxides. Following incubation, the samples were dried in a speed-vac concentrator under vacuum at 24 °C, dissolved in DMSO (1 mL), and analyzed by LC−MS employing a C18 reversed-phase Betabasic column (5 μm particle size, 150 Å pore size, 25 cm length, and 4.6 mm i.d.) eluted with a gradient starting from 5% B (0.1% trifluoacetic acid in acetonitrile) and 95% A (0.1% trifluoacetic acid in water) for 5 min, followed by linear increase to 35% B over 60 min, a linear increase to 80% of B over 5 min, and finally a wash with 80% of B for 10 min. A flow rate of 1.0 mL/min was used, and the products were monitored by their UV absorbance at 254 nm. LC/ESI−MS experiments were carried out using an ion-trap mass analyzer on a LCQ FLEET instrument (Thermo Fisher Scientific). Positive-ion electrospray was used as the means of ionization. The heated inlet capillary temperature was 375 °C and electrospray needle voltage was 5 kV. Nitrogen sheath gas was supplied at 45 psi, and the LC/ESI−MS analysis was performed in the positive-ion mode. Quantum Yield Determination. Fluorescence quantum yields (Φu) were measured and calculated by the Parker−Rees method54 employing the relationship

Φu = Φref (A r /Lr)(Lu /A u )(ηu 2 /ηr 2) In the expression above, Φref denotes the quantum yield of a wellknown fluorophore, which in this case was coumarin 153 (Φref = 38% in ethanol). Au represents the absorbance of the unknown sample at the desired excitation wavelength, Ar is the reference fluorophore’s absorbance at the same excitation wavelength, Lu designates the total integrated luminescence intensity of the unknown sample when excited at the same excitation wavelength, and Lr is the total integrated luminescence intensity of the reference fluorophore when excited at the same excitation wavelength. The refractive indices of the solvents in which the unknown and reference emissions are measured in are denoted as ηu and ηr, respectively.55 HPLC Calibration Curves. Standard solutions of each analyte (10, 20, 30, 50, 100 μM in DMSO) were prepared using serial dilution from higher concentration stock solutions and analyzed using a C18 reversed-phase column (particle size 5 μm, 150 Å pore size, 25 cm length, and 4.6 mm i.d.) eluted with a mixture of water/methanol/ acetic acid (74:25:1) for 45 min at a flow rate of 0.9 mL/min, while products were detected by their UV absorbance at 254 nm. Quinone compounds were analyzed using the same system eluted with an isocratic solvent composed of water/methanol/acetic acid (59:40:1) for 45 min at a flow rate of 0.9 mL/min, while the compounds were detected by their absorbance at 260 nm.

Figure 4. Fluorimetric analysis of metabolites generated in the aerobic and hypoxic metabolism of 1a, 1b, and 1c by CYPOR, XO, and DTdiaphorase. The results of aerobic assays are shown with blue bars; the results of anaerobic assay are shown with red bars: (A) results for 1a; (B) results for 1b; (C) results for 1c. Assays were extracted with ethyl acetate and fluorescence of the organic extract measured. Reaction conditions: X/XO assays: di-N-oxide (500 μM), xanthine (2 mM), xanthine oxidase (0.04 U/mL), sodium phosphate buffer (50 mM, pH 7.0); CYPOR assays: di-N-oxide (500 μM), NADPH (2 mM), cytochrome P450 reductase (0.13 U/mL), sodium phosphate buffer (50 mM, pH 7.0); no-enzyme control assays: di-N-oxides (500 μM), NADPH (2 mM), sodium phosphate buffer (50 mM, pH 7.0); DTdiaphorase assays: di-N-oxides (500 μM), NADPH (2 mM), DTdiaphorase (0.01 U/mL), sodium phosphate buffer (50 mM, pH 7.0). Error bars represent the standard deviation.

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ASSOCIATED CONTENT

S Supporting Information *

acid and acetic acid) from Fischer (Pittsburgh, PA); and deuterated NMR solvents from Cambridge Isotope Laboratories (Andover, MA). General Procedure for in Vitro Metabolism Reactions. In a typical hypoxic assay, the 1,2,4-benzotriazine 1,4-dioxide (500 μM) was incubated with NADPH (2 mM) or xanthine (2 mM) and cytochrome P450 reductase (0.13 U/mL) or xanthine oxidase (0.04 U/mL, or DT-diaphorase (0.005 U/mL) or no enzyme in sodium phosphate buffer (50 mM, pH 7.0) containing DMSO (0.5% v/v) at 24 °C for 4 h. All components of the reactions except enzymes and NADPH were degassed by three freeze−pump−thaw cycles. Enzymes and NADPH were diluted with degassed water in an argon-filled glovebag to prepare stock solutions. Reactions were initiated by the addition of enzymes, wrapped in aluminum foil to prevent exposure to light, and incubated in an argon-filled glovebag. The aerobic controls were performed in an identical manner except with aerobic stock solutions and solvents. Following incubation, the reactions were opened to air and diluted to 1 mL by the addition of 800 μL of aerobic sodium phosphate buffer (50 mM, pH 7.0) and extracted with ethyl acetate (1 mL). The materials in the organic layer were analyzed using fluorescence spectroscopy or HPLC analysis. The brightness (Φ•ε) of 4b and 4c in ethyl acetate is 1475 and 1742 M−1 cm−1, respectively.

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.joc.7b03035. Brightness of 25 different benzotriazine and benzotriazine 1-oxides, photophysical properties of 4a−c, LC−MS analysis of hypoxic metabolism, fluorimetric analysis of hypoxic metabolism, and HPLC analysis of 4a−c (PDF)

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Tel: (573) 882-6763. ORCID

Nathaniel G. Mahieu: 0000-0002-0469-9934 Gary A. Baker: 0000-0002-3052-7730 Kent S. Gates: 0000-0002-4218-7411 Notes

The authors declare no competing financial interest. 3130

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(32) Hay, M. P.; Gamage, S. A.; Kovacs, M. S.; Pruijn, F. B.; Anderson, R. F.; Patterson, A. V.; Wilson, W. R.; Brown, J. M.; Denny, W. A. J. Med. Chem. 2003, 46, 169−182. (33) Hay, M. P.; Hicks, K. O.; Pchalek, K.; Lee, H. H.; Blaser, A.; Pruijn, F. B.; Anderson, R. F.; Shinde, S. S.; Wilson, W. R.; Denny, W. A. J. Med. Chem. 2008, 51, 6853−6865. (34) Hay, M. P.; Pruijn, F. B.; Gamage, S. A.; Liyanage, H. D. S.; Kovacs, M. S.; Patterson, A. V.; Wilson, W. R.; Brown, J. M.; Denny, W. A. J. Med. Chem. 2004, 47, 475−488. (35) Fuchs, T.; Chowdhury, G.; Barnes, C. L.; Gates, K. S. J. Org. Chem. 2001, 66, 107−114. (36) Hicks, K. O.; Siim, B. G.; Jaiswal, J. K.; Pruijn, F. B.; Fraser, A. M.; Patel, R.; Hogg, A.; Liyanage, H. D. S.; Dorie, M. J.; Brown, J. M.; Denny, W. A.; Hay, M. P.; Wilson, W. R. Clin. Cancer Res. 2010, 16, 4946−4957. (37) Laderoute, K.; Rauth, A. M. Biochem. Pharmacol. 1986, 35, 3417−3420. (38) Shi, X.; Poole, J. S.; Emenike, I.; Burdzinski, G.; Platz, M. S. J. Phys. Chem. A 2005, 109, 1491−1496. (39) Poole, J. S.; Hadad, C. M.; Platz, M. S.; Fredin, Z. P.; Pickard, L.; Guerrero, E. L.; Kessler, M.; Chowdhury, G.; Kotandeniya, D.; Gates, K. S. Photochem. Photobiol. 2002, 75, 339−345. (40) Johnson, K. M.; Parsons, Z. D.; Barnes, C. L.; Gates, K. S. J. Org. Chem. 2014, 79, 7520−7531. (41) Mason, J. C.; Tennant, G. J. Chem. Soc. B 1970, 911−916. (42) Sarkar, U.; Hillebrand, R.; Johnson, K. M.; Cummings, A. H.; PHung, N. L.; Rajapakse, A.; Zhou, H.; Willis, J. R.; Barnes, C. L.; Gates, K. S. J. Het. Chem. 2017, 54, 155−160. (43) Jiang, F.; Yang, B.; Fan, L.; He, Q.; Hu, Y. Bioorg. Med. Chem. Lett. 2006, 16, 4209−4213. (44) Lavis, L. D.; Raines, R. T. ACS Chem. Biol. 2008, 3, 142−155. (45) Vallner, J. J. J. Pharm. Sci. 1977, 66, 447−465. (46) Chowdhury, G.; Kotandeniya, D.; Barnes, C. L.; Gates, K. S. Chem. Res. Toxicol. 2004, 17, 1399−1405. (47) Fitzsimmons, S. A.; Lewis, A. D.; Riley, R. J.; Workman, P. Carcinogenesis 1994, 15, 1503−1510. (48) Patterson, A. V.; Barham, H. M.; Chinje, E. C.; Adams, G. E.; Harris, A. L.; Stratford, I. J. Br. J. Cancer 1995, 72, 1144. (49) Munro, A. W.; Noble, M. A.; Robledo, L.; Daff, S. N.; Chapman, S. K. Biochemistry 2001, 40, 1956−1963. (50) Ganley, B.; Chowdhury, G.; Bhansali, J.; Daniels, J. S.; Gates, K. S. Bioorg. Med. Chem. 2001, 9, 2395−2401. (51) Gutierrez, P. L. Free Radical Biol. Med. 2000, 29, 263−275. (52) Reymond, J.-L.; Fluxá, V. S.; Maillard, N. Chem. Commun. 2009, 34−46. (53) Yolken, R. H.; Stopa, P. J. J. Clin. Microbiol. 1979, 10, 317−321. (54) Parker, C. A.; Rees, W. T. Analyst 1962, 87, 83−111. (55) Baker, G. A.; Bright, F. V.; Detty, M. R.; Pandey, S.; Stilts, C.; Yao, H. J. Porphyrins Phthalocyanines 2000, 4, 669−683.

ACKNOWLEDGMENTS Dr. Lixin Ma and Dr. Hang Xu for their help in preparation of cell extracts.

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REFERENCES

(1) Mottram, J. C. Br. J. Radiol. 1936, 9, 606−614. (2) Vaupel, P. Oncologist 2008, 13, 21−26. (3) Wilson, W. R.; Hay, M. P. Nat. Rev. Cancer 2011, 11, 393−409. (4) Chen, C.-Y.; Willard, D.; Rudolph, J. Biochemistry 2009, 48, 1399−1409. (5) Phillips, R. M. Cancer Chemother. Pharmacol. 2016, 77, 441−457. (6) Hunter, F. W.; Wouters, B. G.; Wilson, W. R. Br. J. Cancer 2016, 114, 1071−1077. (7) Wardman, P. Curr. Med. Chem. 2001, 8, 739−761. (8) Hunter, F. W.; Young, R. J.; Shalev, Z.; Vellanki, R. N.; Wang, J.; Gu, Y.; Joshi, N.; Sreebhavan, S.; Weinreb, I.; Goldstein, D. P.; Moffat, J.; Ketela, T.; Brown, K. R.; Koritzinsky, M.; Solomon, B.; Rischin, D.; Wilson, W. R.; Wouters, B. G. Cancer Res. 2015, 75, 4211−4223. (9) Wang, J.; Guise, C. P.; Dachs, G. U.; Phung, Y.; Hsu, A. H.-L.; Lambie, N. K.; Patterson, A. V.; Wilson, W. R. Biochem. Pharmacol. 2014, 91, 436−446. (10) Patterson, A. V.; Saunders, M. P.; Chinje, E. C.; Patterson, L. H.; Stratford, I. J. Anti-Cancer Drug Des. 1998, 13, 541−573. (11) Wardman, P.; Dennis, M. F.; Everett, S. A.; Patel, K. B.; Stratford, M. R. L.; Tracy, M. Biochem. Soc. Symp. 1995, 61, 171−194. (12) Wardman, P.; Clarke, E. D.; Hodgkiss, R. J.; Middleton, R. W.; Parrick, J.; Stratford, M. R. L. Int. J. Radiat. Oncol., Biol., Phys. 1984, 10, 1347−1351. (13) Wang, J.; Foehrenbacher, A.; Su, J.; Patel, R.; Hay, M. P.; Hicks, K. O.; Wilson, W. R. Clin. Cancer Res. 2012, 18, 1684−1695. (14) Rajapakse, A.; Gates, K. S. J. Org. Chem. 2012, 77, 3531−3537. (15) Rajapakse, A.; Linder, C.; Morrison, R. D.; Sarkar, U.; Leigh, N.; Barnes, C. L.; Daniels, J. S.; Gates, K. S. Chem. Res. Toxicol. 2013, 26, 555−563. (16) Cui, L.; Zhong, Y.; Zhu, W.; Xu, Y.; Du, Q.; Wang, X.; Qian, X.; Xiao, Y. Org. Lett. 2011, 13, 928−931. (17) Jamieson, S. M. F.; Gu, Y.; Manesh, D. M.; El-Hoss, J.; Jing, D.; MacKenzie, K. L.; Guise, C. P.; Foehrenbacher, A.; Pullen, S. M.; Benito, J.; Smaill, J. B.; Patterson, A. V.; Mulaw, M. A.; Konopleva, M.; Bohlander, S. K.; Lock, R. B.; Wilson, W. R. Biochem. Pharmacol. 2014, 88, 36−45. (18) Su, J.; Guise, C. P.; Wilson, W. R. Biochem. J. 2013, 452, 79−86. (19) Hodgkiss, R. J.; Begg, A. C.; Middleton, R. W.; Parrick, J.; Stratford, M. R. L.; Wardman, P.; Wilson, G. D. Biochem. Pharmacol. 1991, 41, 533−541. (20) Zhu, W.; Dai, M.; Xu, Y.; Qian, X. Bioorg. Med. Chem. 2008, 16, 3255−3260. (21) Tanabe, K.; Hirata, N.; Harada, H.; Hiraoka, M.; Nishimoto, S.-i. ChemBioChem 2008, 9, 426−432. (22) Rajendran, J. G.; Krohn, K. A. Semin. Nucl. Med. 2015, 45, 151− 162. (23) Elmes, R. B. P. Chem. Commun. 2016, 52, 8935−8956. (24) Brown, J. M. Br. J. Cancer 1993, 67, 1163−1170. (25) Brown, J. M. Cancer Res. 1999, 59, 5863−5870. (26) Junnotula, V.; Sarkar, U.; Sinha, S.; Gates, K. S. J. Am. Chem. Soc. 2009, 131, 1015−1024. (27) Shen, X.; Rajapakse, A.; Gallazzi, F.; Junnotula, V.; FuchsKnotts, T.; Glaser, R.; Gates, K. S. Chem. Res. Toxicol. 2014, 27, 111− 118. (28) Daniels, J. S.; Gates, K. S. J. Am. Chem. Soc. 1996, 118, 3380− 3385. (29) Shinde, S. S.; Hay, M. P.; Patterson, A. V.; Denny, W. A.; Anderson, R. F. J. Am. Chem. Soc. 2009, 131, 14220−14221. (30) Wardman, P.; Priyadarsini, K. I.; Dennis, M. F.; Everett, S. A.; Naylor, M. A.; Patel, K. B.; Stratford, I. J.; Stratford, M. R. L.; Tracy, M. Br. J. Cancer 1996, 74, S70−S74. (31) Laderoute, K. L.; Wardman, P.; Rauth, M. Biochem. Pharmacol. 1988, 37, 1487−1495. 3131

DOI: 10.1021/acs.joc.7b03035 J. Org. Chem. 2018, 83, 3126−3131