Promotion of HeLa cells apoptosis by cynaropicrin involving inhibition of thioredoxin reductase and induction of oxidative stress

Free Radical Biology and Medicine 135 (2019) 216–226

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Original article

Promotion of HeLa cells apoptosis by cynaropicrin involving inhibition of thioredoxin reductase and induction of oxidative stress

T

Tianyu Liua, Junmin Zhangb,∗, Xiao Hana, Jianqiang Xuc, Yueting Wua, Jianguo Fanga,∗∗ a

State Key Laboratory of Applied Organic Chemistry and College of Chemistry and Chemical Engineering, Lanzhou University, Lanzhou, 730000, China School of Pharmacy, Lanzhou University, Lanzhou, 730000, China c School of Life Science and Medicine & Panjin Industrial Technology Institute, Dalian University of Technology, Panjin Campus, Panjin, 124221, China b

A R T I C LE I N FO

A B S T R A C T

Keywords: Thioredoxin reductase Cynaropicrin Reactive oxygen species (ROS) Redox regulation Oxidative stress Apoptosis

Cancer is considered as one of the highly mortal diseases globally. This is largely due to the lack of efficacious medicines for tumors, and thus development of potent anticancer agents is urgently needed. The thioredoxin (Trx) system is crucial to the survival ability of cells and its expression is up-regulated in many human tumors. Recently, increasing evidence has been established that mammalian thioredoxin reductase (TrxR), a selenocysteine-containing protein and the core component of the thioredoxin system, is a promising therapeutic target. The sesquiterpene lactone compound cynaropicrin (CYN), a major component of Cynara scolymus L., has shown multiple pharmacological functions, especially the anticancer effect, in many experimental models. Most of these functions are concomitant with the production of reactive oxygen species (ROS). Nevertheless, the target of this promising natural anticancer product in redox control has rarely been explored. In this study, we showed that CYN induces apoptosis of Hela cells. Mechanistic studies demonstrated that CYN impinges on the thioredoxin system via inhibition of TrxR, which leads to Trx oxidation and ROS accumulation in HeLa cells. Particularly, the cytotoxicity of CYN is enhanced through the genetic knockdown of TrxR, supporting the pharmacological effect of CYN is relevant to its inhibition of TrxR. Together, our studies reveal an unprecedented mechanism accounting for the anticancer effect of CYN and identify a promising therapeutic agent worthy of further development for cancer therapy.

1. Introduction Based on the data from World Health Organization, cancer is the second leading cause of death worldwide after cardiovascular diseases in 2018. One reason for the high mortality of cancer is the lack of efficacious medicines. One promising approach is to escalate the specificity of medicine by targeting the cancer-specific vulnerabilities. The approach could be more desirable and efficacious to lessen the incidence of cancer at the early stage. Alternatively, it could help to alleviate the symptoms of cancer or even to impede further progression of extant tumor. Hence, it is crucial to precisely recognize the risk points

in the pathogenic features of carcinoma in detail. One of the outstanding risk points, often accompanied with carcinogenesis and tumor progression, is the elevated level of reactive oxygen species (ROS) [1,2]. Generally, cancer cells are particularly sensitive to the perturbation of intracellular ROS level [3], and accordingly up-regulate antioxidant pathways to prevent excessive oxidative damage [4,5]. One type of the novel anti-cancer agents lies in targeting these protective pathways in already-sensitized cancer cells to generate the excessive ROS [6]. The excessive ROS in already-sensitized cancer cells is regarded as augmentation of the potency in some chemotherapies [7]. However, compared with giving the general inhibition to these pathways, a more

Abbreviations: Ac-DEVD-pNA, N-acetyl-Asp-Glu-Val-Asp-p-nitroanilide; AIF, apoptosis inducing factor; Annexin V-FITC, fluorescein-5-isothiocyanate-conjugated Annexin V; ASK1, apoptosis signal regulating kinase 1; BSA, bovine serum albumin; BSO, L-buthionine-(S,R)-sulfoximine; CYN, cynaropicrin; DCFH-DA, 2',7’dichlorfluorescein diacetate; DHE, dihydroethidium; DMEM, Dulbecco’s modified Eagle’s medium; DMSO, dimethyl sulfoxide; DTNB, 5,5′-Dithiobis-2-nitrobenzoic acid; FBS, fetal bovine serum; GPx, glutathione peroxidase; GR, glutathione reductase; HeLa-shNT, HeLa cells transfected with shNT plasmid; HeLa-shTrxR1, HeLa cells transfected with shTrxR1 plasmids; MTT, 3-(4, 5-dimethylthiazol-2-yl)-2, 5-diphenyltetrazolium bromide; NAC, N-acetyl-L-cysteine; NADPH, nicotinamide adenine dinucleotide phosphate; PI, propidium iodide; PMSF, phenylmethylsulfonyl fluoride; Ref-1, redox factor 1; ROS, reactive oxygen species; SecTRAP, Seccompromised thioredoxin reductase-derived pro-apoptotic protein; SOD, superoxide dismutase; TCEP, phosphine; Trx, thioredoxin; TrxR, thioredoxin reductase ∗ Corresponding author. ∗∗ Corresponding author. E-mail addresses: [email protected] (J. Zhang), [email protected] (J. Fang). https://doi.org/10.1016/j.freeradbiomed.2019.03.014 Received 19 January 2019; Received in revised form 11 March 2019; Accepted 11 March 2019 Available online 14 March 2019 0891-5849/ © 2019 Elsevier Inc. All rights reserved.

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dysfunction of diverse signal pathways, the significant antitumor activity of CYN has been reported, and the mechanisms of this natural product has been proposed, such as regulation of β1-integrin CD29 and CD98 functions [33], suppression of STAT3 activation [34], induction of cell cycle arrest at G1/S phase [35], down-regulation of the p210BCR/ ABL oncoprotein [36], reduction of NF-κB expression and elevation of lipid peroxidation [37]. Interestingly, CYN exerts therapeutic effect frequently concomitant with the production of ROS [24,27,29,34,37], but the potential molecular targets of this small molecule in redox control still have rarely been explored. Based on previous research, the chemically-reactive α, β-unsaturated carbonyl moiety is a desirable structural feature for the TrxR inhibitory effect [15,19,38,39]. Considering that this function group also exists in CYN, thus it is a reasonable hypothesis that CYN may interact with TrxR. In this study, we showed that CYN impinges on the thioredoxin pathway via inhibition of TrxR, and promotes oxidized Trx accumulation and ROS-mediated apoptosis in HeLa cells. Particularly, a genetic silence of TrxR expression enhances the cytotoxicity of CYN, indicating the pharmacological function of CYN is related to its inhibition of TrxR. Together, our results unveil a novel mechanism underlying the anticancer effect of CYN and demonstrate that CYN is a promising therapeutic agent deserving further development.

targeted agent, aiming at leading to selective oxidative stress in tumor while sparing normal tissue, is needed. The thioredoxin system, composed of thioredoxin (Trx), thioredoxin reductase (TrxR) and NADPH, occupies a significant position in regulating intracellular redox homeostasis [8,9]. This system promotes several crucial processes for tumor formation and progression, and has increased in human tumor cells [10,11]. In the physiological environment, TrxR has been believed the principal enzyme that catalyzes the reduction of oxidized Trx, so its activity is a key factor to regulate the function of this system. Structurally, TrxR is a selenium-dependent enzyme with a unique but essential selenocysteine (Sec) residue at the penultimate C-terminal position [9,12]. Nowadays, the mammalian TrxR enzymes are divided into three types, i.e., TrxR1 (cytoplasm/nucleus), TrxR2 (mitochondrion) and TrxR3 (testis), according to their cellular localization or tissue distribution [9,13,14]. Together, TrxR is progressively considered as a hot target for anticancer drug development, and increasing attention has been attracted to discover effective inhibitors of this enzyme [15–21]. Functional foods (like herbs, grains and spices) comprise considerable amounts of bioactive compounds contributed to promotion of health. With cancer propagation worldwide, researchers’ attention has been increasingly drawn by some functional foods due to their advantages in counteracting against various cancers [22]. So it is becoming important to find some way to locate small bioactive molecules in these functional foods and exploit their antitumor mechanisms. Cynara scolymus L., commonly called an artichoke, is farmed in several countries and noted for its pharmacological and physiological effects [23]. Such health benefits of artichokes have been attributed to the high sesquiterpene lactone content, especially the content of cynaropicrin (CYN) [24]. CYN was first isolated from artichoke in 1960, and now is regarded as a marker on the chemotaxonomy of artichoke plants [25]. Structurally, CYN has a 5-7-5 fused tricyclic skeleton with two hydroxyl groups, four exo-olefins and six stereocenters (Fig. 1A). In recent years, it has been reported to possess phenomenal pharmacologic abilities including anti-hepatitis C virus activity [26], anti-parasitic activity [27,28], anti-photoaging activity [29], anti-hyperlipidemic activity [30], anti-inflammatory activity [31], antibacterial activity [32] and so on. Considering that cancer is a complicated process involving the

2. Materials and methods 2.1. Reagents and enzymes CYN (97% purity) was obtained from Chengdu Biopurify Phytochemicals Ltd. (Chengdu, China) and a stock solution (100 mM) in dimethyl sulfoxide (DMSO; Sigma-Aldrich, St. Louis, USA) was stored at −20 °C. Final concentrations of DMSO in all experiments were lower than 0.1% (V/V). Most chemicals and bioreagents were obtained from Sigma-Aldrich (St. Louis, USA) except for NADPH (Roche, Mannheim, Germany), sephadex G-25 and fetal bovine serum (FBS; Sijiqing, Hangzhou, China), dihydroethidium (DHE; Santa Cruz Biotech, Santa Cruz, USA), 5,5′-Dithiobis-2- nitrobenzoic acid (DTNB; J&K Scientific, Beijing, China), 3-(4,5-dimethyl-2-thiazolyl)-2,5- diphenyl- 2-H-tetrazolium bromide (MTT; Amresco, Solon, USA), cytochrome c (BBI Life Fig. 1. Cytotoxicity of CYN. (A) Chemical structure of CYN. (B) Cytotoxicity of CYN against HeLa, HepG 2, SMMC-7721, BEAS-2B and HEK 293T cell lines. After exposure to indicated concentrations of CYN for 24 h, the cell viability was measured by MTT assay. (C) Dose- and time-dependent cytotoxicity of CYN against HeLa cells. After exposure to indicated concentrations of CYN for 24 h or 48 h, the cell viability was measured by the MTT assay. (D) Cell viability measured by trypan blue exclusion assay after HeLa cells were treated with CYN for 24 h. Data are expressed as mean ± SE from triplicates. **, P < 0.01 vs. the control groups in (B), (C) and (D). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

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containing GSSG (1 mM) and NADPH (400 μM) in TE buffer was added before measuring absorbance at 340 nm.

Sciences, Shanghai, China), trypan blue and bovine serum albumin (BSA; Beyotime, Nantong, China). The primary antibody against TrxR1 and the horseradish peroxidase-conjugated secondary antibody were from Santa Cruz Biotech (Santa Cruz, USA). The primary antibodies against Trx1 and β-actin were from Sangon Biotech (Shanghai, China). The recombinant rat TrxR1, mutant TrxR1 (U498C), and recombinant Escherichia coli Trx were prepared as described in our previous publications [40,41]. With DTNB as the substrate, the recombinant TrxR1 and U498C TrxR1 exhibited enzyme activities of ∼666 and ∼63 mol of NADPH oxidized/min/mol, respectively [42].

2.5. Cellular enzyme activity assay 2.5.1. Assay of TrxR activity Cellular TrxR activity was determined in 96-well plates by a microplate reader. HeLa cells were treated with different concentrations of CYN in 100 mm dishes (2 × 106 cells/dish) for 24 h, while the cells incubated only with DMSO served as a control. Following treatment, the collected cells were washed twice with PBS and the cellular protein was extracted with RIPA buffer (50 mM Tris-HCl: pH 7.5, 2 mM EDTA, 0.5% deoxycholate, 150 mM NaCl, 1% TritonX-100, 0.1% SDS, 1 mM Na3VO4 and 1 mM PMSF). The protein concentration was quantified by the Bradford procedure [47]. The Trx-mediated insulin reduction method was used to examine the cellular TrxR activity as previous described [44].

2.2. Cell culture The human cancer cell lines (HepG2 cells, HeLa cells and SMMC7721 cells) and normal cell lines (HEK 293T cells and BEAS-2B cells) were obtained from the Shanghai Institute of Biochemistry and Cell Biology, Chinese Academy of Sciences. Cells were routinely cultured in DMEM supplemented with 10% FBS, 2 mM glutamine, 100 units/mL penicillin and streptomycin (Amresco, Solon, USA) and incubated at 37 °C with 5% CO2 in humidified atmosphere. The shRNA plasmids targeting TrxR1 (shTrxR1) and a non-targeting control (shNT) were generously provided by Prof. Constantinos Koumenis from University of Pennsylvania School of Medicine [43], and the transfected cells were generated and maintained as previously reported [44,45].

2.5.2. Assay of GR and glutathione peroxidase (GPx) activity HeLa cells were treated with 10 μM, 20 μM or 40 μM CYN in 100 mm dishes (2 × 106 cells/dish) for 24 h, while the cells incubated only with the same amount of DMSO without CYN served as a control. Following treatment, the collected cells were washed twice with PBS and the cellular protein was extracted with RIPA buffer. The protein concentration was quantified by the Bradford procedure. The enzyme activities were determined by measuring the linear change of absorbance in 96-well plates by a microplate reader every 15 s for the initial 7.5 min (30 times totally). Blank reactions with cellular protein sample replaced by distilled water were subtracted from each assay. The enzyme activities were eventually shown as the percentage of control. The GR activity was determined as previous described with modifications [48,49]. Briefly, cellular protein samples (30 μg) or the same amount of distilled water were incubated with the reaction mixture consisted of 2 mM EDTA, 0.2 M potassium phosphate buffer (pH 7.0) and 0.2 mM NADPH in a final volume of 100 μL for 30 min. After addition of 100 μL of GSSG (2 mM) to initiate the reaction, the oxidation of the NADPH was measured spectrophotometrically at 340 nm for GR assay. The GPx activity was determined as previous described with modifications [50]. Briefly, cellular protein samples (20 μg) or the same amount of distilled water were incubated with the reaction mixture consisted of 50 mM potassium phosphate buffer (pH 7.0), 1 mM EDTA, 0.2 mM NADPH, 1 mM NaN3, 2 U/mL GR and 1 mM GSH in a final volume of 100 μL for 5 min. After addition of 100 μL of H2O2 (0.5 mM) to initiate the reaction, the oxidation of the NADPH was measured spectrophotometrically at 340 nm for GPx assay.

2.3. Determination of cell viability 2.3.1. MTT assay As described by us previously [42,46], cells were treated with different concentrations of CYN in 96-well plates (5 × 103 cells/well) for indicated times, while the cells incubated only with DMSO served as a control and the culture medium in the absence of cells served as a blank. Next, 10 μL of MTT (5 mg/mL) was added and cells were further grown for 4 h. Following treatment, 100 μL of the extraction buffer containing 10% SDS, 5% iso-butanol and 0.1% HCl was added. After overnight incubation the absorbance at 570 nm was measured by a microplate reader (Thermo Scientific Multiskan GO, Finland). Cell viability was calculated through the following calculation: Cell viability (%) = (ACYN-ABlank) / (AControl-ABlank) × 100%

2.3.2. Trypan blue exclusion assay HeLa cells were treated with different concentrations of CYN in 12well plate (1 × 105 cells/well) for 24 h, while the cells incubated only with DMSO served as a control. Following staining with trypan blue (0.4%, w/v), viable (non-stained) and dead (stained) cells were counted under a microscope. Cell viability was calculated as the percentage of viable cells to total cells.

2.6. Image-based TrxR activity assay As described by us previously [46,51,52], we applied the selective TrxR probe, TRFS-green, to image TrxR activity in live HeLa cells. Briefly, HeLa cells were treated with different concentrations of CYN in 6-well plate (2 × 104 cells/well) for 20 h, while the cells incubated only with DMSO served as a control. Following staining with TRFS-green (10 μM) in a fresh medium for 4 h, the cell images were captured by fluorescence microscope under a green channel (Leica DMI4000).

2.4. Enzyme activity assay in vitro The inhibitory effects of CYN on the purified enzymes were examined spectrophotometrically in 96-well plates by a microplate reader every 10 s for the initial 4 min (25 times totally) [44,45]. Samples only incubated with the same amount of DMSO without CYN served as a control and the enzyme activities were shown as the percentage of control. First, the recombinant TrxR1 (85 nM), U498C TrxR1 (350 nM) and yeast glutathione reductase (GR; 0.25 U/ml) were treated with NADPH for 5 min to generate fully reduced enzymes. Then the NADPHreduced enzymes were treated with different concentrations of CYN in a final volume of 50 μL for indicated times. Finally, the reaction was initiated by adding 50 μL of assay solution containing DTNB (2 mM) and NADPH (200 μM) in TE buffer (50 mM Tris-HCl; pH 7.5, 1 mM EDTA) and the absorbance at 412 nm was determined for TrxR assay. Accordingly, for GR activity determination, 50 μL of assay solution

2.7. Determination of intracellular ROS HeLa cells were treated with different concentrations of CYN in 12well plate (1 × 104 cells/well) for different time intervals (2 or 6 h), while the cells incubated only with DMSO served as a control. Subsequently, after removal of the medium, the cells were continued to incubate with 2', 7’-dichlorfluorescein diacetate (DCFH-DA; 10 μM) or DHE (10 μM) in a fresh medium for 0.5 h in the dark. Following incubation, the cell images were captured by fluorescence microscope. 218

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2.8. Induction of superoxide anion

were captured by a fluorescence microscope.

The NADPH-reduced recombinant TrxR1 (1.6 μM) was incubated with 400 μM CYN at 37 °C for 1 h in TE buffer. The remaining enzyme activity was less than 10% of the control as monitored by the DTNB reduction assay. After removal of the excessive CYN by a Sephadex G25 desalting column, the modified enzyme or control enzyme (55 μL) was added to 245 μL of TE buffer containing 250 μM NADPH. The production of superoxide anion was determined by the cytochrome c reduction assay as previous described [44]. The control enzyme without incubation with CYN was treated in the same manner.

2.12.2. Determination of caspase-3 activity Using N-acetyl- Asp-Glu- Val-Asp-p-nitroanilide (Ac-DEVD-pNA) as a substrate, we employed a colorimetric assay to measure the activity of caspase-3. Briefly, HeLa cells were treated with different concentrations of CYN in 100 mm dishes (1 × 106 cells/dish) for 48 h, while the cells incubated only with DMSO served as a control. Following treatment, the collected cells were washed twice with PBS and the cellular protein was extracted with RIPA buffer. The protein concentration was quantified by the Bradford procedure, and the caspase-3 activity was determined as described [49,51].

2.9. Determination of intracellular free thiols 2.12.3. Apoptosis by flow cytometry The fluorescein-5-isothiocyanate-conjugated Annexin V (Annexin VFITC) and propidium iodide (PI) double staining apoptosis detection kit (Zoman Biotech, Beijing, China) was employed to determine the apoptosis-inducing effect of CYN by flow cytometry. Briefly, HeLa cells were treated with different concentrations of CYN in 6-well plates (2 × 104 cells/well) for 48 h, while the cells incubated only with DMSO served as a control. Following treatment, the collected cells were washed twice with PBS and resuspended in 500 μL binding buffer based on the manufacture’s instruction. Following staining with 5 μL Annexin VFITC and 10 μL PI for 30 min, cells were filtered and analyzed by a FACSCantoTM flow cytometer (BD Biosciences, USA) with the Cell Quest software.

The DTNB titration was employed to determine the intracellular free thiols. HeLa cells were treated with different concentrations of CYN in 100 mm dishes (2 × 106 cells/dish) for 24 h, while the cells incubated only with DMSO served as a control. Following treatment, the collected cells were washed twice with PBS and the cellular protein was extracted with RIPA buffer. The protein concentration was quantified by the Bradford procedure. The level of intracellular free thiols was determined as described previously [42,44]. 2.10. Determination of the intracellular GSH/GSSG ratio The intracellular GSH/GSSG ratio was determined by an enzymatic recycling method. Briefly, HeLa cells were treated with different concentrations of CYN in 100 mm dishes (2 × 106 cells/dish) for 24 h, while the cells incubated only with DMSO served as a control. Following treatment, the collected cells were washed twice with PBS and next subjected to ultrasonication in ice-cold KPE buffer (0.1% Triton X-100, 5 mM EDTA, 0.1 M potassium phosphate buffer (pH 7.5), and 6 mg/mL sulfosalicylic acid). The protein concentration was quantified by the Bradford procedure. The intracellular GSH/GSSG ratio was determined according to the published protocol [46,53].

2.13. Statistics All experiments were assayed in triplicate and the results were presented as mean ± SE. Comparisons among multiple groups were statistically assessed by the one-way analysis of variance (ANOVA), followed by a post hoc Scheffe test. Statistical differences between two groups were analyzed by the Student’s t-test. P < 0.05 was considered as the criterion for statistical significance.

2.11. Determination of the Trx redox status in HeLa cells

3. Results

The redox status of Trx in HeLa cells was examined by the PAOsepharose pull-down assay described in our previous publications [41,51]. Briefly, HeLa cells were treated with different concentrations of CYN in 100 mm dishes (2 × 106 cells/dish) for 24 h, while the cells incubated only with DMSO served as a control. Following treatment, the whole cell protein was extracted with RIPA buffer. The protein concentration was quantified by the Bradford procedure. Next, control cell extracts were divided into three groups, two of which were selected to prepare completely oxidized and reduced samples through incubating samples with TCEP (5 mM) and diamide (5 mM) for 30 min respectively. Subsequently, all samples were loaded onto the PAO-sepharose beads and incubated in a rotate shaker for 30 min. After that, the oxidized Trx remained in the solution while the reduced Trx was fully pulled down on the PAO-sepharose beads. Following further centrifugation, the supernatant and sepharose beads were collected. Next, the sepharose-bound sample was washed twice and the reduced Trx was knocked out by TE buffer containing 20 mM DMPS. Finally, all samples were electrophoresed by SDS-PAGE and further identified by Western blot with an antibody against Trx1.

3.1. Cancer cell sensitivity to CYN CYN has shown potential anticancer effect on certain cancer cells in previous studies [33–37], and herein we confirmed the cytotoxicity of CYN against different cell lines. First, several cancer and normal cell lines were incubated with indicated concentrations of CYN for 24 h and then the cytotoxicity of CYN was detected by the MTT assay. The results showed that CYN exhibited significant inhibitory potency against HeLa, HepG 2 and SMMC-7721 cells. Relatively, normal cell lines, including BEAS-2B and HEK 293T cells, were less sensitive to CYN treatment (Fig. 1B). As CYN displayed the highest cytotoxicity to HeLa cells, we then chose HeLa cells to study the underlying mechanisms. As shown in Fig. 1C, CYN exhibited the dose- and time-dependent cytotoxic effect to HeLa cells and its half inhibitory concentration (IC50) were measured as ∼20 μM and ∼10 μM for the 24 h-treatment and 48 h-treatment, respectively. We further employed the trypan blue exclusion assay to determine the cell viability after CYN treatment. As shown in Fig. 1D, results from the trypan blue exclusion assay were consistent with those from the MTT assay. Collectively, our results demonstrated that CYN possesses pronounced cytotoxicity to various cancer cells.

2.12. Apoptosis assays 3.2. TrxR inhibition by CYN 2.12.1. Hoechst 33342 staining HeLa cells were treated with different concentrations of CYN in 6well plates (1 × 105 cells/well) for 24 h, while the cells incubated only with DMSO served as a control. Subsequently, after removal of the medium, the cells were continued to incubate in fresh FBS-free medium containing Hoechst 33342 (5 μg/mL) for 0.5 h in the dark. Cell images

The α, β-unsaturated carbonyl group at the lactone ring has been documented as the important pharmacophore of CYN [24]. The structural feature is capable of forming covalent bonds with sulfhydryl groups in biomolecules via a nucleophilic Michael addition [15,39]. As TrxR contains a highly reactive Sec residue [12], we thus hypothesized 219

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Fig. 2. Inhibition of TrxR by CYN. (A) Dose-dependent inhibition of the purified TrxR1, GR and U498C TrxR1. The NADPH-reduced recombinant TrxR1, U498C TrxR1 and GR were treated with different concentrations of CYN for 30 min, and the activities were measured. (B) Time-dependent inhibition of the purified TrxR1 by CYN. The NADPHreduced recombinant TrxR1 was treated with CYN (40 μM) for different times, and the activity was measured by the DTNB reduction assay. (C) Image of the TrxR activity in live HeLa cells by TRFS-green. After exposure to indicated concentrations of CYN for 24 h, HeLa cells were stained with 10 μM TRFS-green. Scale bars: 20 μm. (D) Determination of TrxR activity in CYN-treated HeLa cells. HeLa cells were treated with varying concentrations of CYN for 24 h, and the activity was measured by the Trx-mediated endpoint insulin reduction assay. (E) Alteration of protein level of TrxR1 in HeLa cells after CYN treatment. The cells were treated with CYN for 24 h, and the proteins’ expression was determined by Western blots. The arrow indicated the bands between 100 kDa and 130 kDa seen in western blots. The β-actin served as a loading control. Data are expressed as mean ± SE from triplicates, and all activities were expressed as the percentage of the control. *, P < 0.05 and **, P < 0.01 vs. the control groups in (A), (B) and (D). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

classical TrxR1 band at 55 kDa in the cell lysates. According to the previous publications [20,54], these bands are likely the dimer form of TrxR. Taken together, CYN might inhibit the activity of TrxR, and the Sec residue is essential for the inhibition by CYN.

that CYN could work as a TrxR inhibitor. First, we explored the effects of CYN on the purified TrxR to verify this assumption (Fig. 2A). Notably, CYN could significantly decrease the activity of the recombinant rat TrxR1, and the inhibitory effect was in a dose-dependent manner. Meanwhile, there was negligible inhibition of CYN to GR and U498C TrxR1, a point mutation of TrxR1 where the 498th Sec residue is substituted by a Cys residue. These data suggested that CYN selectively inhibits the activity of TrxR1 and the Sec residue in TrxR1 is required for the inhibition. Besides, we observed that CYN had the time-dependent inhibition of TrxR (Fig. 2B). For further study, we next set out to determine whether the TrxR activity in intact cells would be inhibited by CYN. Initially, we determined the TrxR activity in live HeLa cells with the TRFS-green (Fig. 2C), a selective TrxR probe developed by us previously [52]. The gradual attenuation of intracellular fluorescence intensity indicated that TrxR activity was inhibited by CYN in a dosedependent manner. To verify the image-based data, we also performed the classical Trx-mediated insulin reduction method to evaluate the inhibition of TrxR by CYN. There was the clear dose-dependent inhibition of the enzyme activity, which confirmed the inhibitory effect of CYN on the cellular TrxR activity (Fig. 2D). Besides, we assessed the protein level of TrxR1 in HeLa cells after the cells were treated with CYN for 24 h. The results from western-blot analysis demonstrated that there was no apparent effect on the protein level of TrxR1 (Fig. 2E), which suggested that the attenuated TrxR activity was a direct result of the inhibition. Interestingly, we also observed the bands between 100 kDa and 130 kDa (where the arrow indicates) apart from the

3.3. TrxR participation in the cellular action of CYN Our results have identified that CYN impaired the enzyme activity of both the purified TrxR and the cellular TrxR, and displayed significant cytotoxicity against several cancer cell lines. The TrxR activity in HeLa cells was suppressed by 33.16%, 55.81% and 81.79% respectively after 10, 20 and 40 μM CYN-treatment for 24 h. The corresponding cell viability was 65.01%, 48.43% and 33.32% under the same conditions. Plotting the cell viability data versus the enzyme inhibition data gave a good correlation (Fig. 3A), suggesting the cytotoxicity of CYN is related to its ability to inhibit TrxR. In order to confirm that the observed cytotoxicity of CYN is relevant to its inhibition of TrxR, we generated HeLa-shTrxR1 and HeLa-shNT cells, and compared the cytotoxicity of CYN to these two cell lines. The HeLashTrxR1 cells are HeLa cells that were stably transfected with a shRNA plasmid specifically targeting TrxR1, and the HeLa-shNT cells are HeLa cells that were stably transfected with a non-targeting shRNA plasmid [44,45]. The knockdown efficiency of TrxR1 in the HeLa-shTrxR1 cells and HeLa-shNT cells was evaluated by detecting the expression of TrxR1 protein (Fig. 3B). CYN showed elevating cytotoxicity on the HeLa-shTrxR1 cells compared with the HeLa-shNT cells (Fig. 3C). Our 220

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Fig. 3. Involvement of TrxR in the cellular action of CYN. (A) Inhibition of the TrxR activity in HeLa cells parallels with the death of cell. The graph showed the correlation of cellular TrxR activity (left column) and cell viability (right column). (B) The protein level of TrxR1 in two transfected cell lines. The β-actin served as a loading control. (C) Knockdown of TrxR1 enhances cytotoxicity of CYN. The transfected cells treated with indicated concentrations of CYN for 24 h, and the cell viability was determined by the MTT assay. Data are expressed as mean ± SE from triplicates. *, P < 0.05 and **, P < 0.01 vs. the HeLa-shTrxR1 cells in (C).

diamide group was largely present in the supernatant, which confirmed an effective way to detect the reduced and oxidized Trx (Fig. 4B). As shown in Fig. 4C, Trx was mainly presented in the reduced form in the control group (non-CYN treated cells). In contrast, CYN treatment led to a dose-dependent increase of oxidized Trx. Cells treated with 20 μM of CYN had an evident increase in the oxidized Trx, and the content of oxidized Trx exceeded reduced Trx when the cells were treated with 40 μM of CYN. The blots were quantified by Image J, and the ratio of reduced to oxidized Trx declined significantly in response to the increasing concentration of CYN (Fig. 4D). According to the blots, the total Trx after 40 μM of CYN treatment decreased significantly, which was in parallel with the results shown in Fig. 4A. These results showed that CYN could block the reduction of Trx in HeLa cells. Combined with the previous data, CYN at such concentrations could effectively inhibit the cellular TrxR activity. Therefore, the accumulation of oxidized Trx further supported that CYN targets TrxR in HeLa cells.

results parallel with the known TrxR-targeting agents including plumbagin [42], securinine [46] and alantolactone [51], which demonstrated that the knockdown of TrxR can enhance the cytotoxicity of TrxR inhibitors. In summary, our studies showed that eliminating TrxR activity is closely linked to the cytotoxicity of CYN. 3.4. Alteration of Trx redox status Under physiological conditions, TrxR is the principal reductase of Trx to maintain Trx in a reduced state [8,9]. Therefore, examining the intracellular Trx redox status could provide direct evidence for the change of TrxR activity. In regard of the level of Trx1 in HeLa cells treated with CYN for 24 h, neither 10 μM nor 20 μM CYN did significantly influence Trx1 levels; whereas 40 μM CYN-treatment remarkably decreased Trx1 expression (Fig. 4A). Next, we adopted the PAO-sepharose pull-down assay developed by us previously to determine the redox status of Trx in HeLa cells treated with CYN [41,51]. The fully reduced Trx and oxidized Trx were generated by incubating samples with TCEP and diamide, respectively. As expected, Trx in the TCEP group was predominantly bound to the beads, while that in the

3.5. Induction of oxidative stress The thioredoxin system occupies a significant position in regulating cellular redox homeostasis. Hence, it is meaningful to investigate whether CYN affects the ROS level and causes oxidative stress in HeLa cells. First, we assessed the level of ROS in HeLa cells using general fluorescence probe DCFH-DA. As a result, we detected weak fluorescence signal in the control group and a concentration-dependent increase of bright green fluorescence in CYN-treated cells, while the fluorescence decreased significantly by the pretreatment of NAC, a ROS scavenger (Fig. 5A). Due to the limitation of DCFH-DA [55], we used another fluorescence probe, DHE, to assess the level of ROS in the CYNtreated HeLa cells. Similarly, the DHE fluorescence was promoted by CYN treatment and decreased by NAC (Fig. 5B). To investigate whether accumulation of ROS was the preceding contribution to the cell death caused by TrxR inhibition, the less incubation times of CYN were employed (the incubation times were 2 h and 6 h for DCFH-DA and DHE respectively), prior to the significant cytotoxicity observed at 24 h. In addition, we made a quantitative analysis of fluorescence signal (Fig. 5C and D). These data suggested that CYN boosts the level of ROS preceding the death of HeLa cells. Furthermore, previous studies have indicated that certain compounds can irreversibly induce TrxR1 to become a potent superoxide-producing NADPH oxidase upon inhibition [44,56], and we therefore analyzed whether CYN has this effect. First, the reversibility of TrxR inhibition was examined by a Sephadex G-25 desalting column and the result showed that TrxR activity could not be recovered after removal of CYN from the CYN-incubated TrxR1 (Fig. 5E), suggesting that CYN is an irreversible inhibitor of TrxR. Compared with unmodified TrxR1, we further found that cytochrome c could be reduced constantly by the CYN-modified TrxR1 in a NADPHdependent manner and this redox reaction can be terminated by SOD

Fig. 4. Alteration of Trx1 redox status in HeLa cells after CYN treatment. (A) Alteration of protein level of Trx1 in HeLa cells. The cells were treated with CYN for 24 h, and the proteins’ expression was determined by Western blots. The β-actin served as a loading control. (B) The HeLa cell extracts incubated with TCEP (5 mM) and diamide (5 mM) for 30 min were used to validate the PAO-sepharose pull-down assay. O: oxidized form; R: reduced form; S: samples in the supernatant; P: samples eluted from the PAO-sepharose beads. (C) Measurement of the Trx1 redox status in HeLa cells by the PAO-sepharose pulldown assay. The HeLa cells were treated with varying concentrations of CYN for 24 h, and the cell extracts were prepared and subjected to the PAO-sepharose pull-down assay. (D) Histogram analysis of the ratio of reduced Trx1 to oxidized Trx1 by ImageJ. Data are expressed as mean ± SE from triplicates. **, P < 0.01 vs. the control groups in (D). 221

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Fig. 5. Induction of oxidative stress in HeLa cells. HeLa cells were pretreated in the absence or presence of NAC (0.5 mM) for 1 h followed by incubation with indicated concentrations of CYN for 2 h (DCFH-DA) or 6 h (DHE), and then the cells were stained with (A) DCFH-DA (10 μM) or (B) DHE (10 μM). Scale bars: 20 μm. The fluorescence intensity analysis of individual cells from (A) and (B) by ImageJ was shown in (C) and (D). (E) Irreversible inhibition of TrxR by CYN. The NADPHreduced recombinant TrxR1 (1.6 μM) was treated with CYN (400 μM) for 1 h. After removal of the excessive CYN by a Sephadex G-25 desalting column, the activity was measured by the DTNB reduction assay. The control enzyme without incubation with CYN was treated in the same manner. (F) Induction of superoxide anion production by CYN-modified TrxR. Superoxide anion generation was monitored by the cytochrome c reduction assay. The inset (right) shows the change in absorbance at 550 nm after addition of cytochrome c and SOD. SOD was added where the arrow indicates in the inset. (G) The decrease of free thiols in CYN-treated HeLa cells. After the cells were treated with CYN for 24 h, the intracellular free thiols were measured by DTNB titration method. (H) The decrease of GSH/GSSG in CYN-treated HeLa cells. After the cells were treated with CYN for 24 h, the GSH/GSSG was measured by the enzymatic recycling assay. Data are expressed as mean ± SE from triplicates. **, P < 0.01 vs. the control groups in (C), (D), (E), (G) and (H).

treatment of CYN (Fig. 5G). In parallel, we measured the cellular GSH/ GSSG status, another common index of the cellular redox state. There was a dose-dependent decline of GSH/GSSG in HeLa cells under CYN treatment (Fig. 5H). Taken together, our results, i. e., elevation of ROS level, drop of free thiols and decrease of GSH/GSSG, indicated that CYN disturbs the cellular redox balance and causes oxidative stress in HeLa cells.

(Fig. 5F), which indicated the inhibition of TrxR1 by CYN could directly contribute to the production of ROS. Cellular redox homeostasis is mainly dependent on the mutual transformation of sulfhydryl and disulfide bond [57]. Therefore, we quantified the level of cellular free thiols with the DTNB titration method to further assess the redox status in HeLa cells. There was a dose-dependent decrease of thiol content in HeLa cells under the

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Fig. 6. Effects of CYN on the GSH system. HeLa cells were treated with indicated concentrations of CYN for 24 h, and the intracellular GPx activity (A) and GR activity (B) were assessed. (C) Supplement of NAC lowers the cytotoxicity of CYN. HeLa cells were pretreated with indicated concentrations of NAC for 24 h, followed by CYN treatment for additional 24 h, and the cell viability was determined by the MTT assay. (D) Depletion of GSH enhances the cytotoxicity of CYN. HeLa cells were pretreated in the presence or absence of 50 μM BSO for 24 h, followed by CYN treatment for additional 24 h, and the cell viability was determined by the MTT assay. Data are expressed as mean ± SE from triplicates. **, P < 0.01 vs. the groups without NAC in (C). **, P < 0.01 vs. the groups without BSO in (D).

3.6. Role of GSH system in cellular response to CYN

3.7. Apoptosis induction by CYN

In mammalian cells, thioredoxin and glutathione systems are the two major redox-regulating systems which are dependent on the sulfhydryl groups [57,58]. CYN contains the potential reactive moiety, the Michael acceptor scaffold, for thiol modification. Above results have demonstrated that CYN caused the decline of intracellular enzyme activity of TrxR, accumulation of oxidized Trx, drop of total thiols and decrease of GSH/GSSG. Therefore exploring how the GSH system acts during CYN treatment would undoubtedly provide a valuable insight into understanding the molecular mechanism of CYN. In GSH system, GPx could convert H2O2 into water accompanied by the oxidation of GSH to GSSG, which is then recycled back into GSH by GR and NADPH [59,60]. Therefore, we determined the activity changes of GPx and GR in CYN-treated HeLa cells. The observed results suggested that CYN did not significantly impede the GPx and GR under conditions of inhibiting TrxR and inducing ROS (Fig. 6A and B). Importantly, considering the decrease of GSH/GSSG in the CYN-treated Hela cells, we held that buffering the excessive ROS caused by CYN could significantly consume the GSH rather than the direct inhibition of the GSH system. Due to the core status of GSH in the glutathione system [61], thus we next checked the survival ability of HeLa cells by manipulation of cellular GSH level. First, we carried out cytotoxicity tests using HeLa cells pretreated with NAC, the biosynthetic precursor of GSH. As shown in Fig. 6C, addition of NAC could reduce the number of dead cells induced by CYN. Moreover, the cytotoxicity of CYN was effectively impaired by the high concentration of NAC. To further investigate whether inhibition of GSH synthesis would affect the cytotoxicity of CYN, we employed L-buthionine-(S, R)-sulfoximine (BSO), a common inhibitor of GSH synthesis in cells, to pretreat HeLa cells. Treating the cells with 50 μM BSO for 24 h could markedly lower the content of GSH in HeLa cells below 20% of the control group. As expected, when HeLa cells were pretreated with BSO, the effect of CYN on cell death was augmented (Fig. 6D). We interpret our results to mean that CYN lead to no enzymatic alteration of the GSH system but decrease the ratio of GSH/GSSG because of the TrxR inhibition. Importantly, the level of GSH could significantly affect the cytotoxicity of CYN.

Since the reduced form of Trx could interact with various apoptosisrelated enzymes and signal factors to suppress apoptosis, the abrogation of apoptosis can be observed easily in many kinds of tumor cell lines because of the increase of thioredoxin system [62]. Furthermore, although the studies above have demonstrated that CYN selectively impairs cellular TrxR activity and induces the oxidative stress through blocking the reduction of Trx in HeLa cells, we could not work out that the observations occurred in HeLa cells treated with CYN would contribute to the cell death by inducing apoptosis. Thus, the apoptosisinducing effect of CYN was evaluated by Hoechst 33342 staining, flow cytometry analysis and caspase-3 activity test. Hoechst 33342 could bind to double-strand DNA and appear blue fluorescence. We found that cells in the control group were regularly arranged and the nuclei were circular with low fluorescence signal, while the apoptotic rate was increased with typical features of apoptosis such as intensive fluorescence and nuclear condensation with increasing concentrations of CYN (Fig. 7A). Additionally, apoptosis was observed quantitatively with flow cytometry by Annexin V-FITC/PI double-staining assay. Followed by the treatment with CYN at concentrations of 10 μM, 20 μM and 40 μM for 48 h, the apoptotic cells were detected to be 28%, 78% and 88% of total cells, respectively (Fig. 7B). Most importantly, the apoptosis would be attenuated markedly by pretreatment of NAC, suggesting that oxidative stress is involved in the CYN-induced apoptosis of HeLa cells. In our experiments, cell necrosis was rarely observed, and the quantification results were shown in Fig. 7C. Activation of caspase-3 is a characteristic hallmark of apoptosis [63], so caspase-3 activity was further determined. The caspase-3 activity was significantly elevated with the increase of CYN concentrations (Fig. 7D). These findings presented here suggest that CYN could induce apoptosis of HeLa cells. 4. Discussion Targeted therapy has optimized the typical treatment for cancer [64]. Thus, the ability to identify new anticancer agents and understand their molecular targets is of paramount importance. CYN is a main ingredient of the edible portions in nutritious plant artichoke, which has been used as safe dietary for centuries. In recent years, the significant 223

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Fig. 7. Induction of apoptosis by CYN in HeLa cells. (A) Morphological changes of nuclei in the CYN-treated HeLa cells. HeLa cells were stained with Hoechst 33342 (5 μg/mL) after exposed to the indicated concentrations of CYN for 24 h. The small red arrows indicated the typical apoptotic morphological changes of nuclei. Scale bars: 20 μm. (B) Flow cytometry analysis of HeLa cells. After the cells were exposed to different concentrations of CYN for 48 h, the cells were stained with Annexin V-FITC and PI. Then cells were examined for apoptosis by FACS analysis for 1 × 104 cells. For NAC treatment, the cells were pretreated with NAC (0.5 mM) for 24 h, and then the cells were subjected to CYN treatment. (C) Statistical analysis of live cells (Q3), apoptotic cells (Q2 and Q4), and necrotic cells (Q1) after CYN treatment. (D) The activation of caspase-3 in CYN-treated HeLa cells. The intracellular caspase-3 activities were measured by a colorimetric assay after the cells were treated with indicated concentrations of CYN for 48 h. Data are expressed as mean ± SE from triplicates. **, P < 0.01 vs. the control groups in (D). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

cancer cell lines and normal cell lines (Fig. 1). In addition, the death cell parallels with the inhibition of TrxR activity in HeLa cells (Fig. 3A). It is notable that genetic knockdown of TrxR augments the cytotoxic activity of CYN, which implies that the inhibition of TrxR involves in the death of HeLa cell (Fig. 3C). We demonstrated that CYN-mediated TrxR inhibition causes induction of oxidative stress in HeLa cells (Fig. 5), and the oxidative stress could be attributed to several putative interpretations. First, the redox regulating of Trx system is largely dependent on the reduced state of Trx, maintained by the physiological function of TrxR [8,9], so the content of reduced Trx could be directly affected by the change of TrxR activity. According results obtained from the PAO-sepharose pull-down assay (Fig. 4B), accumulation of oxidized Trx upon CYN-mediated TrxR inhibition could aggravate the oxidative stress. Second, we have indicated that CYN has the capacity to inhibit purified TrxR1 irreversibly (Fig. 5E), which could promote the production of superoxide anion (Fig. 5F). This TrxR1 conversion from antioxidant to prooxidant has been investigated in previous studies and the prooxidant enzyme was termed as “SecTRAP” (Sec-compromised thioredoxin reductase-derived pro-apoptotic protein), which can lead to rapid induction of oxidative stress in cells [65,66]. It is intriguing to consider that CYN-mediated oxidative stress not only as a consequence of inhibition of cellular TrxR activities, but also as a consequence of the conversion of the inhibited enzyme to an NADPH oxidase. Consistent with the findings mentioned above, our observations suggested that CYN treatment results in apoptosis in HeLa cells (Fig. 7). We propose that the pro-apoptotic function of CYN in HeLa cells can be triggered via the following pathways. First, cell apoptosis might be promoted by the oxidative stress induced by both SecTRAPs and TrxR inhibition. Inhibition of TrxR by CYN directly induces oxidative stress

antitumor activity of CYN has attracted increasing attention, regarding this natural product as a potential anticancer agent. Importantly, the small size and water-solubility could facilitate its cell membrane permeability and the formulation as therapeutic injections [24]. Herein we provided experimental evidence to demonstrate the inhibitory effect of CYN on TrxR both in vitro and in the cellular context. The purified enzyme analyses of TrxR1 inhibition identified intriguing differences between the recombinant rat TrxR1, U498C TrxR1 and GR (Fig. 2A). GR has a similar structure with TrxR but less sensitivity to CYN under our experimental conditions, suggesting the specific interaction of CYN with TrxR. Meanwhile, the negligible inhibition of U498C TrxR1 compared with recombinant TrxR1 indicated that the Sec residue in TrxR is required for the selective inhibition of TrxR by CYN. Effects of CYN on TrxR, GR and GPx in the cellular context further supported the selective inhibition (Figs. 2D, 6A and 6B), as only TrxR was inhibited by CYN among these enzymes. The highly reactive and surface-exposed Sec residue at the C-terminus may give reason for the preferential inhibition of TrxR1. There is now a growing body of evidence that various electrophiles possess biological function through working as Michael acceptors to directly target the Sec residue in TrxR [17,19,39,45]. As CYN irreversibly inhibited TrxR, this suggested that CYN likely follows the same mechanism to covalently bind to the nucleophilic Sec residue in TrxR. Except that part of our studies has focused on the inhibition of TrxR by CYN, the effects triggered by the inhibition in HeLa cells are striking and have conducted further research. Generally, cancer cells are particularly sensitive to perturbations of the intracellular antioxidant system because of the higher level of ROS and chronic signs of oxidative stress [3], which can also explain the cytotoxicity of CYN. Herein we identified intriguing cytotoxic differences of CYN between several 224

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and promotes ROS-mediated apoptosis. Besides, as discussed above, the CYN-modified TrxR is likely to promote the formation of SecTRAPs in cells and trigger rapid cell apoptosis [65,66]. Moreover, the thioredoxin system is well known to be able to modulate a large number of signaling pathways [62,67]. The reduced Trx, rather than the oxidized Trx, suppresses apoptosis signaling pathways through reduction of the disulfides in redox-regulated factors including apoptosis inducing factor (AIF) [68], apoptosis signal regulating kinase 1 (ASK1) [69], redox factor 1 (Ref-1) [70] and so on. The CYN-mediated TrxR inhibition prevents the substrate, Trx from being reduced, leading to a buildup of oxidized Trx with a concomitant increase in oxidized downstream protein targets, resulting ultimately in cellular apoptotic death. We postulated that CYN could also exert influence on the Trx-related factors, but further evidence will be required. In conclusion, we have demonstrated that TrxR is a target of CYN, and revealed that CYN could promote apoptosis of HeLa cells involving inhibition of TrxR and induction of oxidative stress. The discovery of the interaction between CYN and TrxR unveils a novel mechanism underlying the anticancer effect of CYN and supports that CYN is a promising therapeutic agent deserving further development.

[14] [15]

[16]

[17]

[18]

[19]

[20]

[21]

Conflicts of interest [22]

The authors declare no conflicts of interest.

[23]

Acknowledgements

[24]

This work was supported by grants from the National Natural Science Foundation of China (21572093 & 21778028), Natural Science Foundation of Gansu Province (18JR3RA296), Lanzhou University (the Fundamental Research Funds for the Central Universities, lzujbky2018-39) and the 111 Project. We deeply appreciated Prof. Constantinos Koumenis (University of Pennsylvania) for the shRNA plasmids of TrxR1.

[25]

[26]

[27]

References [28]

[1] A. Glasauer, N.S. Chandel, Targeting antioxidants for cancer therapy, Biochem. Pharmacol. 92 (2014) 90–101. [2] V. Helfinger, K. Schroeder, Redox control in cancer development and progression, Mol. Aspect. Med. 63 (2018) 88–98. [3] C. Gorrini, I.S. Harris, T.W. Mak, Modulation of oxidative stress as an anticancer strategy, Nat. Rev. Drug Discov. 12 (2013) 931–947. [4] G.M. DeNicola, F.A. Karreth, T.J. Humpton, A. Gopinathan, C. Wei, K. Frese, D. Mangal, K.H. Yu, C.J. Yeo, E.S. Calhoun, F. Scrimieri, J.M. Winter, R.H. Hruban, C. Iacobuzio-Donahue, S.E. Kern, I.A. Blair, D.A. Tuveson, Oncogene-induced Nrf2 transcription promotes ROS detoxification and tumorigenesis, Nature 475 (2011) 106–109. [5] I.S. Harris, A.E. Treloar, S. Inoue, M. Sasaki, C. Gorrini, K.C. Lee, K.Y. Yung, D. Brenner, C.B. Knobbe-Thomsen, M.A. Cox, A. Elia, T. Berger, D.W. Cescon, A. Adeoye, A. Brüstle, S.D. Molyneux, J.M. Mason, W.Y. Li, K. Yamamoto, A. Wakeham, H.K. Berman, R. Khokha, S.J. Done, T.J. Kavanagh, C.W. Lam, T.W. Mak, Glutathione and thioredoxin antioxidant pathways synergize to drive cancer initiation and progression, Cancer Cell 27 (2015) 211–222. [6] A.J. Montero, J. Jassem, Cellular redox pathways as a therapeutic target in the treatment of cancer, Drugs 71 (2011) 1385–1396. [7] K.A. Conklin, Chemotherapy-associated oxidative stress: impact on chemotherapeutic effectiveness, Integr. Cancer Ther. 3 (2004) 294–300. [8] J. Lu, A. Holmgren, The thioredoxin antioxidant system, Free Radic. Biol. Med. 66 (2014) 75–87. [9] J. Zhang, X. Li, X. Han, R. Liu, J. Fang, Targeting the thioredoxin system for cancer therapy, Trends Pharmacol. Sci. 38 (2017) 794–808. [10] M. Berggren, A. Gallegos, J.R. Gasdaska, P.Y. Gasdaska, J. Warneke, G. Powis, Thioredoxin and thioredoxin reductase gene expression in human tumors and cell lines, and the effects of serum stimulation and hypoxia, Anticancer Res. 16 (1996) 3459–3466. [11] D.T. Lincoln, E.E. Ali, K.F. Tonissen, F.M. Clarke, The thioredoxin-thioredoxin reductase system: over-expression in human cancer, Anticancer Res. 23 (2003) 2425–2433. [12] T. Tamura, V. Gladyshev, S.Y. Liu, T.C. Stadtman, The mutual sparing effects of selenium and vitamin E in animal nutrition may be further explained by the discovery that mammalian thioredoxin reductase is a selenoenzyme, Biofactors 5 (1995) 99–102. [13] E.S.J. Arnér, Focus on mammalian thioredoxin reductases—important

[29]

[30]

[31]

[32]

[33]

[34]

[35]

[36]

[37]

[38]

225

selenoproteins with versatile functions, Biochim. Biophys. Acta Gen. Subj. 1790 (2009) 495–526. A. Bindoli, M.P. Rigobello, Principles in redox signaling: from chemistry to functional significance, Antioxidants Redox Signal. 18 (2013) 1557–1593. F.F. Gan, K.K. Kaminska, H. Yang, C.Y. Liew, P.C. Leow, C.L. So, L.N. Tu, A. Roy, C.W. Yap, T.S. Kang, W.K. Chui, E.-H. Chew, Identification of Michael acceptorcentric pharmacophores with substituents that yield strong thioredoxin reductase inhibitory character correlated to antiproliferative activity, Antioxidants Redox Signal. 19 (2013) 1149–1165. X. Zheng, W. Ma, R. Sun, H. Yin, F. Lin, Y. Liu, W. Xu, H. Zeng, Butaselen prevents hepatocarcinogenesis and progression through inhibiting thioredoxin reductase activity, Redox biology 14 (2018) 237–249. E.-H. Chew, J. Lu, T.D. Bradshaw, A. Holmgren, Thioredoxin reductase inhibition by antitumor quinols: a quinol pharmacophore effect correlating to antiproliferative activity, FASEB J. 22 (2008) 2072–2083. E.-H. Chew, A.A. Nagle, Y. Zhang, S. Scarmagnani, P. Palaniappan, T.D. Bradshaw, A. Holmgren, A.D. Westwell, Cinnamaldehydes inhibit thioredoxin reductase and induce Nrf2: potential candidates for cancer therapy and chemoprevention, Free Radic. Biol. Med. 48 (2010) 98–111. J. Zhang, B. Zhang, X. Li, X. Han, R. Liu, J. Fang, Small molecule inhibitors of mammalian thioredoxin reductase as potential anticancer agents: an update, Med. Res. Rev. 39 (2019) 5–39. E. Hedstrom, S. Eriksson, J. Zawacka-Pankau, E.S.J. Arner, G. Selivanova, p53-dependent inhibition of TrxR1 contributes to the tumor-specific induction of apoptosis by RITA, Cell Cycle 8 (2009) 3584–3591. W.C. Stafford, X. Peng, M.H. Olofsson, X. Zhang, D.K. Luci, L. Lu, Q. Cheng, L. Trésaugues, T.S. Dexheimer, N.P. Coussens, M. Augsten, H.M. Ahlzén, O. Orwar, A. Östman, S. Stone-Elander, D.J. Maloney, A. Jadhav, A. Simeonov, S. Linder, E.S.J. Arnér, Irreversible inhibition of cytosolic thioredoxin reductase 1 as a mechanistic basis for anticancer therapy, Sci. Transl. Med. 10 (2018) eaaf7444. A. Rodriguez-Casado, The health potential of fruits and vegetables phytochemicals: notable examples, Crit. Rev. Food Sci. Nutr. 56 (2016) 1097–1107. N. Ceccarelli, M. Curadi, P. Picciarelli, L. Martelloni, C. Sbrana, M. Giovannetti, Globe artichoke as a functional food, Mediterr. J. Nutr. Metabol. 3 (2010) 197–201. M.F. Elsebai, A. Mocan, A.G. Atanasov, Cynaropicrin: a comprehensive research review and therapeutic potential as an anti-hepatitis C virus agent, Front. Pharmacol. 7 (2016) 472. D. Chaturvedi, Sesquiterpene lactones: structural diversity and their biological activities, Opportunity, Challanges and Scope of Natural Products in Medicinal Chemistry (Trivandrum: Natural Products Chemistry Division, North-East Institute of Science and Technology (CSIR)), 2011, pp. 313–334. M.F. Elsebai, G. Koutsoudakis, V. Saludes, G. Pérez-Vilaró, A. Turpeinen, S. Mattila, A.M. Pirttilä, F. Fontaine-Vive, M. Mehiri, A. Meyerhans, J. Diez, Pan-genotypic hepatitis C virus inhibition by natural products derived from the wild Egyptian artichoke, J. Virol. 90 (2015) 1918–1930. S. Zimmermann, M. Oufir, A. Leroux, R.L. Krauth-Siegel, K. Becker, M. Kaiser, R. Brun, M. Hamburger, M. Adams, Cynaropicrin targets the trypanothione redox system in Trypanosoma brucei, Bioorg. Med. Chem. 21 (2013) 7202–7209. F. Emendörfer, F. Emendörfer, F. Bellato, V.F. Noldin, V. Cechinel-Filho, R.A. Yunes, F. Delle Monache, A.M. Cardozo, Antispasmodic activity of fractions and cynaropicrin from Cynara scolymus on Guinea-pig ileum, Biol. Pharm. Bull. 28 (2005) 902–904. Y.T. Tanaka, K. Tanaka, H. Kojima, T. Hamada, T. Masutani, M. Tsuboi, Y. Akao, Cynaropicrin from Cynara scolymus L. suppresses photoaging of skin by inhibiting the transcription activity of nuclear factor-kappa B, Bioorg. Med. Chem. Lett 23 (2013) 518–523. H. Shimoda, K. Ninomiya, N. Nishida, T. Yoshino, T. Morikawa, H. Matsuda, M. Yoshikawa, Anti-hyperlipidemic sesquiterpenes and new sesquiterpene glycosides from the leaves of artichoke (Cynara scolymus L.): structure requirement and mode of action, Bioorg. Med. Chem. Lett 13 (2003) 223–228. J.Y. Cho, K.U. Baik, J.H. Jung, M.H. Park, In vitro anti-inflammatory effects of cynaropicrin, a sesquiterpene lactone, from Saussurea lappa, Eur. J. Pharmacol. 398 (2000) 399–407. A. Bachelier, R. Mayer, C.D. Klein, Sesquiterpene lactones are potent and irreversible inhibitors of the antibacterial target enzyme MurA, Bioorg. Med. Chem. Lett 16 (2006) 5605–5609. J.Y. Cho, A.R. Kim, H.-G. Joo, B.-H. Kim, M.H. Rhee, E.S. Yoo, D.R. Katz, B.M. Chain, J.H. Jung, Cynaropicrin, a sesquiterpene lactone, as a new strong regulator of CD29 and CD98 functions, Biochem. Biophys. Res. Commun. 313 (2004) 954–961. E. Butturini, A.C. de Prati, G. Chiavegato, A. Rigo, E. Cavalieri, E. Darra, S. Mariotto, Mild oxidative stress induces S-glutathionylation of STAT3 and enhances chemosensitivity of tumoural cells to chemotherapeutic drugs, Free Radic. Biol. Med. 65 (2013) 1322–1330. J.Y. Cho, A.R. Kim, J.H. Jung, T. Chun, M.H. Rhee, E.S. Yoo, Cytotoxic and proapoptotic activities of cynaropicrin, a sesquiterpene lactone, on the viability of leukocyte cancer cell lines, Eur. J. Pharmacol. 492 (2004) 85–94. A. Russo, M. Perri, E. Cione, M.L. Di Gioia, M. Nardi, M.C. Caroleo, Biochemical and chemical characterization of Cynara cardunculus L. extract and its potential use as co-adjuvant therapy of chronic myeloid leukemia, J. Ethnopharmacol. 202 (2017) 184–191. S.M. Lepore, V. Maggisano, G.E. Lombardo, J. Maiuolo, V. Mollace, S. Bulotta, D. Russo, M. Celano, Antiproliferative effects of cynaropicrin on anaplastic thyroid cancer cells, Endocr. Metab. Immune Disord. - Drug Targets 19 (2019) 59–66. S. Urig, K. Becker, On the potential of thioredoxin reductase inhibitors for cancer therapy, Semin. Canc. Biol. 16 (2006) 452–465.

Free Radical Biology and Medicine 135 (2019) 216–226

T. Liu, et al.

[39] W. Cai, L. Zhang, Y. Song, B. Wang, B. Zhang, X. Cui, G. Hu, Y. Liu, J. Wu, J. Fang, Small molecule inhibitors of mammalian thioredoxin reductase, Free Radic. Biol. Med. 52 (2012) 257–265. [40] J. Xu, Q. Cheng, E.S.J. Arnér, Details in the catalytic mechanism of mammalian thioredoxin reductase 1 revealed using point mutations and juglone-coupled enzyme activities, Free Radic. Biol. Med. 94 (2016) 110–120. [41] Y. Liu, D. Duan, J. Yao, B. Zhang, S. Peng, H. Ma, Y. Song, J. Fang, Dithiaarsanes induce oxidative stress-mediated apoptosis in HL-60 cells by selectively targeting thioredoxin reductase, J. Med. Chem. 57 (2014) 5203–5211. [42] J. Zhang, S. Peng, X. Li, R. Liu, X. Han, J. Fang, Targeting thioredoxin reductase by plumbagin contributes to inducing apoptosis of HL-60 cells, Arch. Biochem. Biophys. 619 (2017) 16–26. [43] P. Javvadi, L. Hertan, R. Kosoff, T. Datta, J. Kolev, R. Mick, S.W. Tuttle, C. Koumenis, Thioredoxin reductase-1 mediates curcumin-induced radiosensitization of squamous carcinoma cells, Cancer Res. 70 (2010) 1941–1950. [44] D. Duan, B. Zhang, J. Yao, Y. Liu, J. Fang, Shikonin targets cytosolic thioredoxin reductase to induce ROS-mediated apoptosis in human promyelocytic leukemia HL60 cells, Free Radic. Biol. Med. 70 (2014) 182–193. [45] D. Duan, B. Zhang, J. Yao, Y. Liu, J. Sun, C. Ge, S. Peng, J. Fang, Gambogic acid induces apoptosis in hepatocellular carcinoma SMMC-7721 cells by targeting cytosolic thioredoxin reductase, Free Radic. Biol. Med. 69 (2014) 15–25. [46] J. Zhang, J. Yao, S. Peng, X. Li, J. Fang, Securinine disturbs redox homeostasis and elicits oxidative stress-mediated apoptosis via targeting thioredoxin reductase, Biochim. Biophys. Acta (BBA) - Mol. Basis Dis. 1863 (2017) 129–138. [47] M.M. Bradford, A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding, Anal. Biochem. 72 (1976) 248–254. [48] B. Mannervik, Measurement of glutathione reductase activity, Current protocols in toxicology 00 (1999) 7.2.1-7.2.4. [49] J. Zhang, D. Duan, J. Xu, J. Fang, Redox-dependent copper carrier promotes cellular copper uptake and oxidative stress-mediated apoptosis of cancer cells, ACS Appl. Mater. Interfaces 10 (2018) 33010–33021. [50] R.A. Lawrence, R.F. Burk, Glutathione peroxidase activity in selenium-deficient rat liver, Biochem. Biophys. Res. Commun. 71 (1976) 952–958. [51] J. Zhang, Y. Li, D. Duan, J. Yao, K. Gao, J. Fang, Inhibition of thioredoxin reductase by alantolactone prompts oxidative stress-mediated apoptosis of HeLa cells, Biochem. Pharmacol. 102 (2016) 34–44. [52] L. Zhang, D. Duan, Y. Liu, C. Ge, X. Cui, J. Sun, J. Fang, Highly selective off-on fluorescent probe for imaging thioredoxin reductase in living cells, J. Am. Chem. Soc. 136 (2014) 226–233. [53] I. Rahman, A. Kode, S.K. Biswas, Assay for quantitative determination of glutathione and glutathione disulfide levels using enzymatic recycling method, Nat. Protoc. 1 (2006) 3159–3165. [54] J. Xu, S. Eriksson, M. Cebula, T. Sandalova, E. Hedstrom, I. Pader, Q. Cheng, C.R. Myers, W.E. Antholine, P. Nagy, U. Hellman, G. Selivanova, Y. Lindqvist,

[55]

[56]

[57] [58]

[59] [60]

[61] [62] [63] [64] [65] [66]

[67] [68]

[69]

[70]

226

E.S.J. Arnér, The conserved Trp114 residue of thioredoxin reductase 1 has a redox sensor-like function triggering oligomerization and crosslinking upon oxidative stress related to cell death, Cell Death Dis. 6 (2015) e1616. B. Kalyanaraman, V. Darley-Usmar, K.J. Davies, P.A. Dennery, H.J. Forman, M.B. Grisham, G.E. Mann, K. Moore, L.J. Roberts II, H. Ischiropoulos, Measuring reactive oxygen and nitrogen species with fluorescent probes: challenges and limitations, Free Radic. Biol. Med. 52 (2012) 1–6. N. Cenas, S. Prast, H. Nivinskas, J. Sarlauskas, E.S.J. Arner, Interactions of nitroaromatic compounds with the mammalian selenoprotein thioredoxin reductase and the relation to induction of apoptosis in human cancer cells, J. Biol. Chem. 281 (2006) 5593–5603. Y.-M. Go, D.P. Jones, Thiol/disulfide redox states in signaling and sensing, Crit. Rev. Biochem. Mol. Biol. 48 (2013) 173–181. Y. Du, H. Zhang, J. Lu, A. Holmgren, Glutathione and glutaredoxin act as a backup of human thioredoxin reductase 1 to reduce thioredoxin 1 preventing cell death by aurothioglucose, J. Biol. Chem. 287 (2012) 38210–38219. M. Deponte, Glutathione catalysis and the reaction mechanisms of glutathione-dependent enzymes, Biochim. Biophys. Acta Gen. Subj. 1830 (2013) 3217–3266. S. Toppo, L. Flohé, F. Ursini, S. Vanin, M. Maiorino, Catalytic mechanisms and specificities of glutathione peroxidases: variations of a basic scheme, Biochim. Biophys. Acta Gen. Subj. 1790 (2009) 1486–1500. H. Zhang, H.J. Forman, Glutathione synthesis and its role in redox signaling, Semin. Cell Dev. Biol. 23 (2012) 722–728. C.H. Lillig, A. Holmgren, Thioredoxin and related molecules-from biology to health and disease, Antioxidants Redox Signal. 9 (2007) 25–47. H.R. Stennicke, G.S. Salvesen, Biochemical characteristics of caspases-3,-6,-7, and8, J. Biol. Chem. 272 (1997) 25719–25723. J.W. Neal, G.W. Sledge, Decade in review-targeted therapy: successes, toxicities and challenges in solid tumours, Nat. Rev. Clin. Oncol. 11 (2014) 627–628. K. Anestål, S. Prast-Nielsen, N. Cenas, E.S.J. Arner, Cell death by SecTRAPs: thioredoxin reductase as a prooxidant killer of cells, PLoS One 3 (2008) e1846. K. Anestål, E.S.J. Arnér, Rapid induction of cell death by selenium-compromised thioredoxin reductase 1 but not by the fully active enzyme containing selenocysteine, J. Biol. Chem. 278 (2003) 15966–15972. M. Dagnell, E.E. Schmidt, E.S.J. Arner, The A to Z of modulated cell patterning by mammalian thioredoxin reductases, Free Radic. Biol. Med. 115 (2018) 484–496. S.B. Shelar, K.K. Kaminska, S.A. Reddy, D. Kumar, C.-T. Tan, C.Y. Victor, J. Lu, A. Holmgren, T. Hagen, E.-H. Chew, Thioredoxin-dependent regulation of AIFmediated DNA damage, Free Radic. Biol. Med. 87 (2015) 125–136. M. Saitoh, H. Nishitoh, M. Fujii, K. Takeda, K. Tobiume, Y. Sawada, M. Kawabata, K. Miyazono, H. Ichijo, Mammalian thioredoxin is a direct inhibitor of apoptosis signal-regulating kinase (ASK) 1, EMBO J. 17 (1998) 2596–2606. K. Hirota, M. Matsui, S. Iwata, A. Nishiyama, K. Mori, J. Yodoi, AP-1 transcriptional activity is regulated by a direct association between thioredoxin and Ref-1, Proc. Natl. Acad. Sci. Unit. States Am. 94 (1997) 3633–3638.