Cancer Letters 372 (2016) 89–100

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Cancer Letters j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / c a n l e t

Original Articles

Astemizole–Histamine induces Beclin-1-independent autophagy by targeting p53-dependent crosstalk between autophagy and apoptosis Rekha Jakhar 1, Souren Paul 1, Monika Bhardwaj, Sun Chul Kang * Department of Biotechnology, Daegu University, Kyoungsan, Kyoungbook 712-714, Republic of Korea

A R T I C L E

I N F O

Article history: Received 26 October 2015 Received in revised form 15 December 2015 Accepted 15 December 2015 Keywords: Apoptosis Non-canonical autophagy H1 receptor Astemizole ER stress p53

A B S T R A C T

Apoptosis and autophagy are genetically regulated, evolutionarily conserved processes that can jointly seal cancer cell fates, and numerous death stimuli are capable of activating either pathway. Although crosstalk between apoptosis and autophagy is quite complex and sometimes contradictory, it remains a key factor determining the outcomes of death-related pathologies such as cancer. In the present study, exposure of MCF-7 breast cancer cells to HIS and the H1 receptor antagonist AST both alone and together with HIS (AST–HIS) led to generation of intracellular ROS, which induced massive cellular vacuolization through dilation of the ER and mitochondria. Consequently, apoptosis by Bax translocation, cytochrome c release, and caspase activation were triggered. In addition, AST–HIS caused ER stress-induced autophagy in MCF-7 cells, as evidenced by an increased LC3-II/LC3-I ratio, with surprisingly no changes in Beclin-1 expression. Non-canonical autophagy was induced via p53 phosphorylation, which increased p53-p62 interactions to enhance Beclin-1-independent autophagy as evidenced by immunocytochemistry and immunoprecipitation. In the absence of Beclin-1, enhanced autophagy further activated apoptosis through caspase induction. In conclusion, these findings indicate that AST–HIS-induced apoptosis and autophagy can be regulated by ROS-mediated signaling pathways. © 2015 Elsevier Ireland Ltd. All rights reserved.

Introduction Breast cancer is the most frequently diagnosed neoplasia and second leading cause of cancer-related death in women [1]. There are various conventional methods available for cancer treatment, but these are often limited by toxicity or acquired resistance. Thus, there is an urgent need for novel agents in order to cure this neoplastic form of cancer. New approaches to fighting breast cancer

Abbreviations: AST, Astemizole; HIS, Histamine; H1R, Histamine H1 receptor; ROS, Reactive oxygen species; ER, Endoplasmic reticulum; PERK, Protein kinase R-like endoplasmic reticulum kinase; IRE1α, Inositol-requiring enzyme 1; ATF, Activating transcription factor; JNK, c-Jun N-terminal kinases; UPR, Unfolded protein response; GRP-78, Glucose-regulated protein 78; eIF2α, Eukaryotic initiation factor 2 α; CHOP, CCAAT/enhancer-binding protein (C/EBP) homologous protein; cAMP, Cyclic adenosine monophosphate; ATG, Autophagy protein; NAC, N-acetyl-L-cysteine; RAP, Rapamycin; p62, Sequestosome 1; Bcl-2, B cell lymphoma 2; BH3, Bcl-2 homology domain-3; BID, BH3 interacting domain; PARP-1, Poly (ADP-ribose) polymerase 1; LC3, Microtubule-associated protein light chain; AMPK, 5’ AMP-activated protein kinase; mTOR, Mammalian target of rapamycin; TSC, Tuberous sclerosis complex; ULK-1, Unc-51-like kinase; MAPK, Mitogen-activated protein kinases; PKC, Protein kinase C; TUNEL, Terminal deoxynucleotidyl transferase dUTP nick end labeling; H2DCFDA, 2’,7’-dichlorodihydrofluorescein diacetate acetyl ester; EMSA, Electrophoretic mobility shift assay; DRAM, Damage-regulated autophagy modulator; B2M, Beta-2-microglobulin; HPRT, Hypoxanthine-guanine phosphoribosyltransferase. * Corresponding author. Tel.: +82 53 850 6553; fax: +82 53 850 6559. E-mail address: [email protected] (S.C. Kang). 1 These authors equally contributed to this work. http://dx.doi.org/10.1016/j.canlet.2015.12.024 0304-3835/© 2015 Elsevier Ireland Ltd. All rights reserved.

include combined targeted systemic adjuvant therapies appropriate for heterogeneous tumors expressing different sets of molecular signatures. Therefore, we analyzed the synergistic effects of HIS and potent histamine H1 receptor antagonist AST (Fig. 1a) on breast cancer cell lines and elucidated their effects on key neoplastic pathways. Many reports have investigated that histamine plays a significant role in breast cancer progression, since functional histamine receptors and histidine decarboxylase activity are present in breast tissue [2]. Previous reports have documented that H1R antagonists induce cell death in human leukemia, myeloma [3,4], breast, colon, and liver cancer cells [5–7]. Development of cancer is associated with programmed cell death (PCD), of which there are two forms; type 1 (apoptotic) and type 2 (autophagic) cell death [8,9]. Among these two forms, apoptosis is the most prevalent form of PCD. Apoptosis is tightly controlled by a balance between pro-apoptotic and anti-apoptotic signals, which in turn are regulated by Bcl-2-family members [10]. Autophagy is an evolutionary conserved cellular defense process that maintains cellular turnover of proteins and organelles through bulk degradation of cellular content via autophagosome formation under stress conditions [11,12]. Apoptosis and autophagy have been shown to be interconnected by several molecular crosstalk nodes, and co-regulation of both affects the tumor suppressor pathway [13]. In fact, these two processes may even be simultaneously regulated by the same trigger and thus act in synergy as well as counter to each other. Two primary

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Fig. 1. Effect of AST–HIS on MCF-7 breast cancer cell viability. (a) Chemical structure of AST. (b) MCF-12A and MCF-7 cells were treated with indicated concentration of HIS, AST, or AST–HIS for 12 h, after which cytotoxicity was determined using LDH assay. (c) Representative microscopic image of MCF-12A and MCF-7 cell morphology after treatment as described in a (scale bar 0.5 mm). (d) Nile Red / DAPI staining of MCF-7 cells with or without RAP, HIS, AST, or AST–HIS treatment to measure cellular lipid level (scale bar 0.1 mm). (e) h1r mRNA levels was analyzed by real-time PCR. (f) Relative intensity (H1R/β-Actin) of data from Western blot analysis was analyzed by densitometry analysis using ImageJ software. β-actin was used as an internal control. (g) h2r, h3r, and h4r mRNA levels were analyzed by real-time PCR. The results represent the mean ± S.D. of three independent experiments, *P < 0.05, **P < 0.01.

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molecules that affect both apoptosis and autophagy are the tumor suppressors Beclin-1 and p53 [14]. p53 is a potent inducer of apoptosis and plays a primary role in limiting oncogenesis through cell cycle arrest for repair, whereas Beclin-1 participates primarily in autophagy and is monoallelically deleted in human breast and ovarian cancers [15,16]. Under various conditions, autophagy can be regulated by p53 either for a protective or death-inducing effect through the induction of multiple proteins [17]. However, understanding of the precise signal transduction pathway and downstream pathophysiological functions of apoptosis and autophagy via H1R remain unresolved. In order to analyze H1R signaling, we chose a human breast cancer cell line (MCF-7) to perform experiments. Therefore, in the present work, we aimed to study the synergistic effects of HIS and histamine H1 receptor antagonist AST on breast carcinoma. We have also established scientific criteria for a combined adjuvant therapy in order to identify regulators of ER stress and elucidate their effects on signaling mechanisms and cellular killing through abortive autophagy and apoptosis based on p53Beclin-1 crosstalk.

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ml) for 30 min in a 5% CO2 incubator at 37 °C. Coverslips were then rinsed with PBS and slides prepared, which were analyzed under a fluorescence microscope. Flow cytometry analysis To detect and quantify apoptotic cells, flow cytometry analysis was performed according to the manufacturer’s instructions using a TUNEL apoptosis detection kit (Roche Applied Sciences, Germany). Cells were stained for 30 min at 37 °C, removed from the plate with trypsin-EDTA, and collected in phenol red-free growth medium. To quantify the development of acidic vesicular organelles (AVOs), MCF-7 cells were stained with a Cyto-ID™ autophagy detection kit (Enzo Life Sciences, Plymouth Meeting, PA). Green (510–530 nm) fluorescence emission from 1 × 104 cells illuminated with blue (488 nm) excitation light was measured with a fluorescent activated cell sorter (FACS), (Beckman coulter, Fullerton, CA, USA) using Kaluza flow cytometry software. Quantifying autophagy with AO staining MCF-7 cells were seeded on a tissue culture grade coverslip (SPL, Republic of Korea). Cells were treated with or without AST, HIS, AST–HIS, or RAP as an autophagy inducer for 12 h, followed by washing with PBS and staining with 100 μg/ ml of AO in serum-free medium at 37 °C for 15 min. Then, cells were again washed with PBS and fluorescent micrographs obtained using a fluorescence microscope (Nikon Eclipse TS200, Nikon Corp., Tokyo, Japan).

Materials and methods Protein kinase C assay Reagents RPMI 1640, HEPES, Histamine, Astemizole, Dithiothreitol (DTT), Rapamycin, and DAPI were purchased from Sigma-Aldrich, St. Louis, USA. Fetal bovine serum was supplied by Gibco, USA. All solvents were of cell culture grade and supplied by SigmaAldrich, St. Louis, USA. Detailed list of antibodies used in this study are included in Table S1.

MCF-7 cells (1 × 105 cells/ml in 6-well plate) were incubated with DTT, AST, HIS, or AST–HIS for the indicated time. Cytoplasmic proteins were isolated using lysis buffer according to the manufacturer’s instructions. PKC assay was carried out using a non-radioactive ELISA-based method as detailed in the kit’s instruction manual (Enzo Life Sciences, Farmingdale NY, USA). PKC activity was determined from the standard curve and expressed as ng/min/ml.

Cells and in vitro treatment

Ca2+ assay

MCF-7 (ATCC, Rockville, MD) human breast cancer cells were cultivated in RPMI 1640 supplemented with 25 mM HEPES buffer, 25 mM sodium bicarbonate, 300 mM L-glutamate, and 10% heat-inactivated fetal bovine serum (Gibco, USA) in a CO2 incubator (5% CO2 in air) at 37 °C. MCF-12A (ATCC, Rockville, MD) human normal mammary epithelial cells were cultured according to the instructions of ATCC. For the experiment, cells were cultured in 12-well plates at a density of 1 × 105 cells/ well. Cells were treated with 5 μM HIS and 6 μM AST alone and together (AST–HIS) for 12 h. Cells treated with 5 mM DTT and 500 nm RAP were used as a positive control for apoptosis and autophagy, respectively. Cells were centrifuged (600 × g; 3 min) and harvested.

Calcium measurements were performed on attached populations of MCF-7 cells. Cells were loaded with 5 μM fura 2-AM for 60 min at 37 °C in HBSS (Hank’s balanced salt solution) buffer and then washed three times with HBSS at 37 °C to remove extracellular fura 2-AM. Fluorescence level of fura 2-loaded MCF-7 cells was monitored with a Nikon Eclipse TS100 Epi-fluorescence microscope, Japan. Fluorescence intensity of cells was measured using ImageJ software (NIH, USA) by subtracting the values of background fluorescence from total fluorescence.

LDH assay The viability of MCF-7 and MCF-12A cells after treatment was estimated by measuring LDH leakage using a cytotoxicity detection kit (Sigma-Aldrich, St. Louis, USA) according to the manufacturer’s protocol. LDH activity was measured as optical density at 450 nm using an ELISA reader (Bio-Tek Instrument Co., WA, USA). Cell morphological assessment Cell morphological changes were assessed after exposure of MCF-7 and MCF12A cells to AST, HIS, or AST–HIS for indicated time. Cells were observed under a compound microscope (Nikon Eclipse TS200, Nikon Corp., Tokyo, Japan). Nile red staining After 12 h of treatment, MCF-7 cells were fixed with 3% paraformaldehyde, incubated for 5 min with 0.1 μg/ml of Nile red (9-diethylamino-5H-benzo [α] phenoxazine-5-one) in PBS, washed with PBS, and examined by fluorescence microscopy (Nikon Eclipse TS100 Epi-fluorescence microscope, Japan). Detection of reactive oxygen species To detect intracellular ROS production, cells were incubated with the peroxidesensitive fluorescent probe H2DCFDA (Molecular probe) at 10 μM for 30 min in the dark. In the presence of intracellular H2O2, non-fluorescent membrane-permeable H2DCFDA is converted to impermeable fluorogenic 2’,7’-dichlorofluorescein. ROS production was thus analyzed with a fluorescence microscope.

cAMP assay cAMP assay was performed using cell-conditioned media using a cAMP assay kit (Enzo Life Sciences, Farmingdale, NY, USA) in accordance with the manufacturer’s protocol. The concentration of cAMP was observed by measuring the absorbance at 450 nm with an ELISA reader. Immunocytologic staining For immunocytochemistry, cells were cultured on a cell culture cover glass (SPL, Republic of Korea) for 24 h before treatment. After treatment with AST, HIS, AST– HIS, or RAP, cells were fixed with 4% paraformaldehyde in phosphate-buffered saline (PBS) for 20 min, blocked with 3% normal goat serum, and incubated with p53 and LC3 primary antibody overnight at 4 °C. Cells were then incubated with FITClabeled goat anti-rabbit IgG secondary antibody for 1 h. The coverslips were washed with PBS, mounted on glass slides with DAPI antifade solution (Invitrogen, USA), and viewed under a fluorescence microscope. EMSA EMSA analyses were performed using a LightShift® Chemiluminescent EMSA Kit (Thermo Scientific, USA). EMSA consensus sequence was labeled with biotin at the 3’ position according to the manufacturer’s protocol (Thermo Scientific, USA). Nuclear extracts (1 μg) were examined for their binding capacity to 20 fmol of the biotinlabeled standard p53 binding motif (F: 5′-TACAGAACATGTCTAAGCATGCTGGGGACT3′, R: 5′-AGTCCCCAGCATGCTTAGACATGTTCTGTA-3′). Binding reaction and 0.75% agarose gel electrophoresis were performed as recommended by the manufacturer. Complexes were then transferred onto Biodyne® precut nylon membranes (Thermo Scientific, USA). For supershift analysis, 2 μg of anti-p53 rabbit polyclonal antibody was used. Biotin-labeled DNA was detected by chemiluminescence as described by the manufacturer.

Measurement of mitochondrial membrane potential siRNA transfection MCF-7 cells were placed on coverslips in 12-well plates and then treated with or without AST, HIS, AST–HIS, or DTT for 12 h to measure changes in MMP level. Treated cells were then washed with PBS and incubated with rhodamine-123 (1 μg/

Cells were seeded on a 6-well plate and transfected with 25 pM siRNA against Beclin-1 (BECN1 no.s16537, Ambion, Life Technologies, Darmstadt, Germany) using

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7.5 μl of Lipofectamine® RNAiMAX Transfection Reagent (Invitrogen, USA). After 24 h of HIS, AST, or AST–HIS treatment, cells were lysed and expression levels of Beclin-1 and β-actin assayed by Western blotting. Quantitative real-time PCR analysis Total RNA was isolated from cells with a RNA-spin™ Total RNA Extraction Kit (Intron Biotechnology, Korea). Samples were then processed according to the manufacturer’s instructions. Concentrations of RNA were analyzed using a Qubit® 2.0 fluorometer RNA Assay Kit (Life technologies, USA). cDNA synthesis was carried out using a Maxime RT Premix cDNA synthesis kit (Intron Biotechnology, Korea) according to the manufacturer’s protocol. RT-PCR controls were β-actin, hprt, and b2m amplified under the same PCR parameters and used to normalize quantitative data. Quantitative real-time PCR was performed using the Agilent technology QPCR System (CA, USA). PCR primers used for h1r and p53 are mentioned in Table S3. Western blot analysis For isolation of whole cell proteins, cells were lysed in RIPA lysis buffer, incubated on ice for 30 min, and centrifuged at 12,000 rpm at 4 °C for 15 min. The supernatant extracts were quantified for proteins by BCA assay (Sigma-Aldrich, St. Louis, USA). Proteins were resolved on SDS–polyacrylamide gel and transferred to a polyvinyldenefluoride (PVDF) membrane (Roche Diagnostics, USA) by electroblotting. The membranes were subsequently incubated with primary antibodies at 4 °C for overnight. Antibodies were then incubated with the respective secondary antibody and visualized by enhanced chemiluminescence (Amersham, Velizy-Villacoublay, France) according to the recommended procedure. Densitometry analysis of all blots was performed using ImageJ (NIH, USA) software. Subcellular fractionation Cytosolic and nuclear fractions were isolated by using an NE-PER nuclear protein extraction kit (Thermo scientific) per the manufacturer’s instructions. The supernatant extracts were quantified for protein by BCA assay. Immunoprecipitation The immunoprecipitation of p53 was performed by pull-down of p53 from total protein lysates according to the manufacturer’s protocol (Invitrogen, Karlsruhe, Germany). Briefly, 5 mg of antibody-coupled magnetic Dynabeads® M-270 Exposy was prepared by using a Dynabeads Co-Immunoprecipitation Kit (Novex, USA). The p53-pulldown products were subjected to 10% denaturing polyacrylamide gel electrophoresis followed by Western blotting to determine p53-p62 interaction. Statistical analysis Comparisons between groups were evaluated by Student’s t-test using the Excel program (Microsoft Co., Redmond, WA, USA). Differences were considered statistically significant at *P < 0.05, **P < 0.01, ***P < 0.001.

Results AST–HIS induces cytoplasmic vacuolation and cell death by inhibiting H1R in MCF-7 cells To assess the cytotoxic effect of AST–HIS treatment on MCF-7 tumorigenic breast cells and MCF-12A non-tumorigenic breast cells, LDH assay was used. As shown in Fig. 1b we evaluated the potential synergistic cytotoxic effects of 5 μM HIS and 6 μM AST alone and together (AST–HIS) on MCF-7 breast cancer cells. At these concentrations, AST–HIS induced significant LDH release compared to HIS or AST treatment alone. Release of LDH was more pronounced in MCF-7 cells compared to the MCF-12A cells following AST–HIS treatment (Fig. 1b). AST–HIS treatment also caused a majority of cells to shrink, round up, and display numerous vacuoles in the cytoplasm, which are typical morphological alterations induced by apoptosis and autophagic cell death. These observations were either absent or less prominent in MCF-12A cells (Fig. 1c).These results signify that tumorigenic MCF-7 cells were specifically sensitive to AST–HIS treatment compared to the normal MCF-12A cells. We also examined the formation of lipid droplets as an indicator of breast cancer cell differentiation using the fluorescent dye Nile Red [18]. This dye associates with lipids in cells that are produced as a part of milk production. During nutrient depriva-

tion or under autophagic conditions, cellular lipids stored as triglycerides in lipid droplets are hydrolyzed into fatty acids for energy [19]. In our study, lipid accumulation was suppressed upon treatment with RAP and AST, and minimal droplet formation was observed in AST–HIS treated cells, signifying induction of lipolysis (Fig. 1d). In contrast, formation of lipid droplets increased in the cytoplasm of untreated and HIS-treated cells. To confirm the roles of AST–HIS and H1R expression in MCF-7 breast cancer cell cytotoxicity, we evaluated H1R expression by RT-PCR and Western blot analysis. The results show that AST–HIS treatment blocked H1R expression at both the mRNA and protein levels (Fig. 1e and f). We have also evaluated the effect of AST–HIS combination treatment on expression of other histamine receptors like h2r, h3r and h4r through RT-PCR. Fig. 1g demonstrates that AST–HIS combination treatment has no significant effect on the mRNA expression of these receptors. AST–HIS induces ER stress and JNK activation via Ca2+-mediated ROS generation in breast cancer cells Blocking of H1R by AST–HIS treatment caused disruption of Ca2+ mobilization, leading to significant Ca2+ release (P < 0.001) from ER to the cytosol (Fig. 2a). The ER and mitochondria form a highly dynamic interconnected network in which mitochondria uptake Ca2+ released from the ER [20]. In addition, perturbation of Ca2+ release from the ER lumen leads to Ca2+-dependent activation of cAMP to maintain calcium homeostasis, which was higher in the AST–HIS treatment group (Fig. 2b). Quite surprisingly, the level of cAMP was reduced after AST treatment (Fig. 2b). We are assuming that this decrease in cAMP level after AST-treatment may be dependent on cellular calcium pool. Previous reports discussed that calcium plays a dual role in regulation of cAMP balance in cells, as it can either increase or decrease cAMP level depending on cell types [21,22]. We are claiming that regulation of cAMP level by calcium is not only dependent on cells, rather dependent on stimulus also. As HIS is linked to the intracellular Ca2+ homeostasis through activation of H1R [23,24], therefore, it reduced Ca 2+ accumulation and increased cAMP level. On the other hand, treatment with AST, a known antagonist of H1R, increases cellular calcium level; therefore, it decreases cAMP level. Interestingly, AST-HIS treatment increases the levels of both Ca2+ and cAMP, which could be attributed to their synergistic efficacy. Accumulated Ca2+ ions in the mitochondria then generate stress through ROS production, which was measured by H2DCFDA staining assay in this study (Fig. 2c, d). Unstimulated MCF-7 cells were found to generate a low level of basal ROS, and this was used as a baseline to which all other results were compared. AST caused a significant increase in ROS generation of 135.8 ± 37.8 over basal (n = 10), AST together with HIS generated maximum of 143.1 ± 14.02 over basal (n = 10), and HIS alone generated only a minor amount of ROS of 59.9 ± 12.4 over basal (n = 10). DTT caused a higher level of ROS generation with a maximum of 167.9 ± 12.8, which was used as a positive control (Fig. 2d). ER stress plays an important role in induction of the UPR in response to multiple stressors such as ROS. To assess whether or not AST–HIS causes ER stress in MCF-7 cells, we measured GRP-78 expression and the phosphorylation status of eIF-2α, a putative marker of ER stress, by Western blot analysis. After treatment with AST– HIS, GRP-78 expression and eIF-2α phosphorylation were significantly elevated compared to other treated groups, indicating induction of ER stress by ROS production (Fig. 2e,h). In addition, the involvement of oxidative stress induced by AST–HIS was also confirmed using anti-oxidant NAC. As shown in Fig. 2f, increase in GRP-78 and UPR target proteins PERK and IRE1α were attenuated by co-treatment with NAC (5 mM) and AST–HIS in MCF-7 cells. These findings suggest that increased expression of GRP-78 upon AST– HIS treatment resulted in activation of CHOP and phosphorylation

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Fig. 2. AST–HIS-induced ER stress in breast cancer cells. (a) Quantification of intracellular calcium accumulation (Ca2+), (b) intracellular cAMP level, and (c) cellular ROS level were visualized by fluorescence microscopy (scale bar 0.1 mm), and (d) corresponding quantification data were measured in MCF-7 cells after DTT, HIS, AST, or AST–HIS treatment for 12 h. (e) MCF-7 cells were treated as in d, respectively, and cell lysates were prepared and subjected to Western blot analysis. (f-g) MCF-7 cells were treated with AST–HIS in combination with ROS and JNK inhibitors respectively, and the activation of GRP-78, PERK, IRE1α, JNK and Bcl-2 was detected by Western blotting. (h) All Western blots were analyzed by densitometry analysis using ImageJ software. β-actin was used as an internal control. The results represent the mean ±S.D. of three independent experiments, *P < 0.05, **P < 0.01, ***P < 0.001.

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Fig. 3. AST–HIS-induced apoptosis in MCF-7 cells. MCF-7 cells were treated with the indicated concentration of DTT, RAP, HIS, AST, or AST–HIS for 12 h, after which (a) cell apoptosis was analyzed by flow cytometry. (b) MCF-7 cells were treated as in a, respectively, after which cell lysates were prepared and subjected to Western blot analysis. (c) All Western blots were analyzed by densitometry analysis using ImageJ software. (d) Mitochondrial membrane potential (ΔΨm) was analyzed by fluorescence microscopy by rhodamine 123 staining (scale bar 0.1 mm) (e) and corresponding quantification data analysis in MCF-7 cells treated as indicated in a. The results represent the mean ±S.D. of three independent experiments, *P < 0.05, **P < 0.01, ***P < 0.001.

of stress-activated kinases JNK and p38 MAPK through increase in expression of ER stress markers PERK and IRE1α. Activated JNK reduced expression of Bcl-2 by phosphorylating Bcl-2 (Fig. 2e,h), which was confirmed by use of JNK inhibitor, SP600125 (20 μM), that potently inhibited the JNK and Bcl-2 phosphorylation induced by AST–HIS (Fig. 2g).

AST–HIS induces apoptotic cell death via mitochondria-mediated signaling pathway and activation of caspases Multiple pathways may be involved in ER stress-initiated cell death, including direct activation of initiator caspases, and apoptosis through crosstalk with mitochondria. To elucidate the mechanism underlying the cytotoxic effect of AST–HIS treatment on MCF-7 cells, apoptotic cell death was measured by TUNEL assay and Western blot analysis. AST–HIS (47.84%) significantly increased the population of late-stage apoptotic cells as compared to HIS or AST alone (25.69 or 33.24%, respectively) (Fig. 3a). For this,

caspases were investigated to elucidate whether or not AST–HISinduced cell death involves caspase activation. BID cleavage is a hallmark of mitochondrial apoptotic stimuli that activate initiator caspases such as caspase-9, which subsequently activates downstream executioner caspase (caspase-3) leading to cell death. AST– HIS treatment significantly increased BID cleavage and activated caspase-9 as well as caspase-3, resulting in PARP-1 cleavage in breast cancer cells compared to control, HIS and AST treated groups (Fig. 3b). Although MCF-7 cells are known to lack caspase-3, they still undergo mitochondria-dependent apoptosis and PARP-1 cleavage, which was confirmed by significantly increased expression of Bax, PUMA, and cytochrome c proteins (Fig. 3b,c). Mitochondrial integrity was examined by fluorescent dye rhodamine123, whose uptake depends on mitochondrial membrane potential (MMP). In our results, MMP was lost after DTT and AST–HIS treatment as compared to control cells (Fig. 3d,e), whereas AST or HIS alone induced very few changes in MMP level. Thus, cytochrome c redistribution from mitochondria is an early apoptotic event that precedes mitochondrial membrane depolarization.

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AST–HIS induces early stage autophagy in breast cancer cells through ER stress In light of the known association between accumulation of unfolded proteins and swelling of the ER, we undertook a series of studies using MCF-7 cells to determine whether or not AST–HIS induces a prototypical ER stress response concurrent with autophagy induction. To investigate the ability of AST–HIS to induce autophagy, cells were stained with acridine orange (AO) dye, a weak base that accumulates in acidic spaces and fluoresces a bright red color, and monitored visually by fluorescence microscopy [25]. As shown in Fig. 4a, AST–HIS induced autophagy in breast cancer cells by forming cytoplasmic vacuoles that were acidic in nature and identified by AO staining. At 12 h post-treatment, the percentages of acidic vacuolar organelles were low in cells treated with RAP, AST, or HIS alone, whereas a marked increase was observed upon AST– HIS treatment. Consistent with AO staining, p62 expression decreased while LC3II formation increased in AST–HIS-treated cells, as identified by Western blot analysis (Fig. 4b–d). We examined LC3 distribution as a hallmark of autophagy induction since LC3-I is degraded during the final stage of productive autophagy, whereas LC3-II accumulates in acidic vacuoles to form a punctate or dotted pattern when analyzed by immunostaining (Fig. 4e). Interestingly, redistribution of overexpressed LC3 into punctate structures appeared to depend on the presence of p62. Very few cells treated with AST or HIS alone to deplete endogenous p62 contained punctate LC3 structures (Fig. 4e). In contrast, overexpression of LC3 strongly increased upon treatment with RAP or AST–HIS (Fig. 4e). Consistent with a direct or indirect association between LC3 and p62, p62 is a multifunctional protein characterized initially by its ability to bind to PKC via its N-terminal PB1 (Phox and Bpem1) domain, thereby attenuating autophagy through phosphorylation of LC3 [26]. In order to determine whether or not AST–HIS indeed directly inhibits PKC via p62 inhibition and induces autophagy, we measured PKC activity in cellular cytoplasmic homogenate using ELISA-based PKC Kinase Activity Assay (Enzo Life Sciences). In our results, PKC activity was significantly (P < 0.05) inhibited by AST–HIS treatment (Fig. 4f). These data support the direct inhibition of PKC by AST– HIS as well as induction of autophagy through increased conversion of LC3-I to LC3-II (LC3 lipidation) as compared to AST or HIS alone. Furthermore, we tested the capacity of AST–HIS treatment to modify autophagy using a 488 nm excitable fluorescent reagent (CytoID™ Autophagy detection kit), which allows for the convenient quantification of autophagic cells by flow cytometry. As shown in Fig. 4g, the maximum numbers of autophagic cells were present in the AST–HIS-treated group (34.32%) as compared to the HIS or AST alone group (16.96 or 19.42%, respectively). AST–HIS-induced autophagy is mediated via activation of p53 related pathway

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tion [29]. EMSA revealed an increase in p53/DNA binding after AST– HIS treatment as compared to AST or HIS alone (Fig. 5f). Transcriptionally activated p53 inhibits mTOR signaling via activation of AMPK, which interacts with TSC2 and ULK1 to induce autophagy [30]. The levels of p-AMPK, TSC2, and ULK1 also increased gradually after RAP and AST–HIS treatment (Fig. 6a). Further, expression levels of other autophagic signals such as Atg5 and Atg7 increased, which corroborates observation that AST–HIS treatment induced autophagy through the p53/AMPK/mTOR axis (Fig. 6a). Next, we investigated the physical interactions of p53 and p62 using immunoprecipitation assay in MCF-7 cells (Fig. 6b). We observed a strong interaction between p53 and ubiquitin-binding protein p62 after AST–HIS treatment, which supports strong activation of autophagy through p53. However, when MCF-7 cells were treated with HIS or AST alone, the interaction between these proteins was attenuated (Fig. 6b). AST–HIS-induced Beclin-1-independent autophagy contributes to apoptosis induction We next assessed expression of Beclin-1, an important autophagy marker involved in autophagosome formation [31,32]. Western blotting data showed that the level of Beclin-1 remained unchanged after AST–HIS treatment, whereas it increased after RAP, HIS, or AST treatment (Fig. 6a). This result suggests the presence of Beclin-1-independent or non-canonical autophagy after AST–HIS treatment. To confirm that Beclin-1 is not required for AST–HISinduced autophagy, we knocked down Beclin-1 in MCF-7 cells using Beclin-1-specific siRNA. Surprisingly, inhibition of Beclin-1 using siRNA did not affect LC3-I conversion in AST–HIS-treated cells, whereas it strongly inhibited LC3-I conversion induced by either AST or HIS, as evidenced by Western blotting (Fig. 6d and e). AST–HIS treatment after Beclin-1 knockdown stimulated redistribution of LC3 fusion protein from a ubiquitous and diffuse pattern towards visible and punctate autophagosomes in the cytoplasm of MCF-7 cells (Fig. 6f). These results indicate that AST–HIS-induced autophagy in MCF-7 cells is Beclin-1-independent. After knockdown of Beclin-1, AST–HIS treatment also resulted in increased p53 expression in the nucleus rather than cytoplasm, which is an indicator of autophagy induction (Fig. 6c). Recently, Grishchuk et al. [33] showed that Beclin-1-independent autophagy leads to induction of caspase-dependent apoptosis. This prompted us to analyze levels of the apoptotic markers caspase-3, caspase9, Bax, Bcl-2, and BID by Western blotting. We observed that knockdown of Beclin-1 was accompanied by up-regulation of caspase-3, which subsequently activates caspase-9, Bax, BID, and down-regulate Bcl-2 in AST–HIS-treated cells, suggesting possible involvement of caspase-induced apoptosis (Fig. 6d). Thus, our data show that both autophagy and apoptosis occur in MCF-7 cells in response to AST–HIS treatment. Discussion

In order to determine the autophagic pathway induced by AST– HIS in MCF-7 cells, activation of p53 and autophagy-related genes along with mTOR inhibition were assessed by Western blotting. According to recent reports, p53 phosphorylation, which is mediated by JNK activation, is critical for autophagic and apoptotic death induction in MCF-7 cells. Nuclear or cytosolic localization of p53 defines its function [27]. As shown by Western blotting and immunofluorescence data (Fig. 5a–d), RAP and AST–HIS treatment caused significant increases in nuclear p53 expression, which transactivates a number of autophagic target genes, whereas decrease in cytoplasmic p53 level [28]. The RT-PCR data also shows the decrease of p53 at gene level after AST–HIS treatment (Fig. 5e). Nuclear localization of p53 was determined by EMSA binding assay to confirm its transcription-dependent effect on autophagy induc-

Substantial advances have been made in breast cancer research and treatment, but there remain significant gaps in translating this newly acquired knowledge into clinical improvements. Thus, there is a need to assess the value of several new approaches. In this study, we investigated the synergistic effect of AST–HIS, which triggered ER stress-induced apoptotic cell death in breast cancer cells followed by autophagy induction through a p53-dependent pathway. Moreover, this induced autophagy was shown to be Atg5- and Atg7dependent while simultaneously Beclin-1-independent, contributing to further apoptosis induction through caspase activation. According to several reports, ER stress develops as a result of harsh environmental cues such as nutrient deprivation, oxidative stress, and metabolic dysregulation [34]. The major protective and

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Fig. 4. Autophagy induction and autophagic flux after AST–HIS treatment. (a) Acridine orange images were taken at 12 h post-initial treatment of MCF cells with or without RAP, AST, HIS, or AST–HIS treatment using a fluorescence microscope. Autophagosomes are indicated by the red dots (Scale bar is 0.1 mm). (b) Representative immunoblots for p62 and LC3, (c) and (d) corresponding quantification showing that AST–HIS treatment significantly increased p62 ubiquitination and LC3-II/I level. (e) Fluorescence images of LC3 immunolabeling (green) in MCF-7 cells after treatment as mentioned in a, respectively. RAP and AST–HIS exposure induced a progressive increase in LC3-positive dots (green). DAPI-stained nuclei are blue (Scale bar is 0.05 mm). (f) PKC activity in MCF-7 cell was analyzed after treatment as described in a, and the data are represented as relative PKC activity. (g) Cell autophagy was analyzed by flow cytometry. Data represent averages ± S.D. of three independent experiments (*P < 0.05, **P < 0.01, ***P < 0.001).

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Fig. 5. Role of p53 in AST–HIS-mediated cell death. MCF-7 cells were treated with the indicated concentration of DTT, RAP, HIS, AST, or AST–HIS for 12 h, and cell lysates were prepared and subjected to Western blot analysis to analyze (a) cytoplasmic and (b) nuclear localization of p53 as well as (c) corresponding quantification showing significant increase in nuclear p53 expression. Data represent averages ± S.D. of three independent experiments (**P < 0.01, ***P < 0.001 for cytoplasmic p53, and ##P < 0.01, ###P < 0.001 for nuclear p53, when compared with control). (d) Immunocytochemical analysis showing nuclear and cytoplasmic localization of p53 (Scale bar is 0.1 mm). (e) p53 mRNA level was analyzed by real-time PCR. Data represent averages ± S.D. of three independent experiments (*P < 0.05, **P < 0.01). (f) EMSA was carried out using biotinylated DNA oligos containing p53-binding sites (Lanes 1, probe labeled with biotin; Lane 2-5, probe + nuclear extract from untreated, HIS, AST, and AST–HIS-treated MCF-7 cells; Lane 6, probe + p53 antibody + nuclear extract from untreated MCF-7 cells) as described in Materials and Methods.

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Fig. 6. Autophagy induced by p53-reactivating molecules after AST–HIS treatment and effect of Beclin-1 knockdown on AST–HIS-induced autophagy. (a) MCF-7 cells were treated with the indicated concentration of DTT, RAP, HIS, AST, or AST–HIS for 12 h, after which cell lysates were prepared and subjected to Western blot analysis. (b) MCF-7 cells were treated as indicated in a, respectively, after which cell lysates were prepared and subjected to immunoprecipitation using p53 antibody and immunoblotted with p62 antibody. (c) Cells were transfected with non-targeting siRNA (Control siRNA) or Beclin-1 siRNA and then treated with the indicated concentration of DTT, RAP, HIS, AST, or AST–HIS for 12 h. Immunoblots were also analyzed for (i) nuclear and (ii) cytoplasmic localization of p53. (d) Relative protein expression level for autophagy-apoptosis markers were analyzed by Western blotting. (e) Representative densitometry analysis of LC3 conversion, and (f) other autophagy-apoptosis marker proteins. (g) Immunocytofluorescence showed increased expression of LC3 in cells transfected with Beclin-1 siRNA compared with control siRNA after treatment with AST–HIS. Green fluorescence represents expression of LC3 puncta. Blue fluorescence of DAPI nuclear staining is also shown (scale bar 0.05 mm). Data represent averages ± S.D. of three independent experiments (*P < 0.05, **P < 0.01, ***P < 0.001).

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compensatory mechanism during ER stress is the UPR, which evolved to facilitate adaptation to changing environments and re-establish ER function [35]. However, if stress is excessive, compensatory mechanisms may not be able to fully sustain ER function, and ER decompensation leads to cell death by activating autophagy and apoptosis [36,37]. Our results show that AST combined with HIS can induce cytotoxicity in MCF-7 breast cancer cells compared with either agent alone by binding to H1R, resulting in induction of apoptosis or autophagic cell death. According to Lilly et al. [2], H1R antagonists modulate HIS-induced mobilization of Ca2+ by binding with H1R. The ER is one of the most important intracellular Ca2+ storage organelles, and depletion of ER Ca2+ stores and Ca2+ influx from the extracellular environment mediate AST–HIS-induced apoptosis. Mitochondria are situated in close proximity to the ER, and thus Ca2+ released from the ER are rapidly taken up by mitochondria to stimulate oxidative phosphorylation, enhanced ROS generation [20], and increased cAMP activation [38]. Importantly, AST–HIS induced both ROS generation and cAMP release, implying that AST–HIS can enhance ER stress in breast cancer cells. The current study supports the hypothesis that ROS signaling can act as a feedback signal to ER stress and lies upstream of the ER-sensing proteins PERK and IRE1α [39,40]. ROS induces phosphorylation of eIF-2α [41], which is a component of the integrated response to ER stress activating GRP-78, an ER-resident chaperone that plays critical roles in stimulating IRE1α, PERK, and ATF6 via physiological and pathological stress [42]. However, in our study, AST–HIS induced eIF-2α phosphorylation along with up-regulation of PERK expression accompanied by enhanced expression of CHOP. According to Oyadomari and Mori [43], overexpression of CHOP promotes apoptosis in several cell lines, whereas CHOP-deficient cells are resistant to ER stress-induced apoptosis, which supports the important role of CHOP in the induction of apoptosis. Moreover, MAPK family members, including JNK and p38, have been reported to be involved in ER stress-induced cell death [44]. In agreement with these reports, we also observed increased phosphorylation of JNK and p38, which is an early sign of ER stress generation. Increased phosphorylation of JNK regulates fundamental cellular pathways, autophagy, and apoptosis by phosphorylating Bcl-2 [45]. JNKmediated phosphorylation of Bcl-2 has been shown to interfere with binding of Bcl-2 to proapoptotic BH3 domain-containing proteins such as Bax and PUMA, thereby mediating collapse of mitochondrial membrane potential and leading to mitochondrial apoptosis [46]. Based on these reports, AST–HIS-induced mitochondrial membrane permeabilization causes release of the apoptotic molecules Bax, PUMA, and cytochrome c to the cytosol, leading to caspase-dependent apoptosis [47]. The present data also suggest that reduction of Bcl-2 expression caused direct modulation of caspase-3 and caspase-9 activation, leading to cell death. Apart from mitochondrial disruption, we also observed that PARP-1, a major regulator of apoptotic cell death, was cleaved by caspases and the proapoptotic Bcl-2 family member BID [47,48]. Proteolytic cleavage of BID converts cytosolic BID into its active form, t-BID, which can cooperate with other proapoptotic Bcl-2 members such as Bax and/or Bak to enhance mitochondrial membrane permeabilization [48]. Furthermore, MCF-7 cells were found to elicit ER stress-mediated autophagy via IRE1α-induced JNK and p38 activation in response to AST–HIS treatment. Our hypothesis that autophagy is induced during ER stress is supported by the following points. First, all ER stressors we examined activated autophagosome formation, as determined by AO staining and LC3 puncta formation via modulation of p62. Second, LC3 was converted from LC3-I to LC3-II in response to ER stress. Third, ER stress can stimulate proteosomal degradation of the cytosolic tumor suppressor p53, resulting in autophagy [49].

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Previous reports have also documented that JNK1-mediated phosphorylation of Bcl-2 interferes with binding of Bcl-2 to the apoptotic protein p53, which participates in ER stress-mediated autophagy and apoptosis [14]. Thus, p53 may be a potential molecular player in the crosstalk among apoptosis, autophagy, and ER stress. Indeed, whereas nuclear p53 can act as an autophagy-promoting transcription factor [50], its cytoplasmic localization inhibits autophagy [27]. AST–HIS treatment caused translocation of p53 from the cytosol to nucleus, as confirmed by immunocytochemistry and Western blotting. Nuclear p53 stimulates autophagy either by transcriptional up-regulation of AMPK, TSC-2, and DRAM, stimulation of autophagic vacuole formation, and/or inhibition of the antiautophagic mTOR pathway [50,51]. As shown in our results, AST– HIS treatment induced p53/DNA binding and thus autophagy through AMPK and TSC-2 up-regulation, resulting in inhibition of mTOR. Indeed, previous studies have shown that p53 can increase the conversion rate of LC3-I to LC3-II by interacting with p62, where p62 acts as a cargo protein by binding to ubiquitinated proteins and LC3-II on the membranes of autophagosomes [52]. AST–HIS treatment also increased p53-p62 interaction, as evidenced by immunoprecipitation. Besides p53, Beclin-1 is another important regulator molecule that participates in both autophagy and apoptosis [15]. Strikingly, the expression level of Beclin-1 remained unchanged or low even after AST–HIS treatment, hence confirming Beclin-1-independent or non-canonical autophagy. Although Beclin-1 is a key regulator of autophagy and its knockdown blocks autophagic cell death [53,54], AST–HIS treatment induced Beclin1-independent autophagy along with apoptosis as confirmed by Western blotting data after Beclin-1 knockdown. We also observed that Beclin-1 knockdown was accompanied by activation of caspase-3 in the presence of AST–HIS, thus sensitizing MCF-7 cells to death. Down-regulation of Beclin-1 also resulted in strong activation of BID and Bax, as shown by the Western blot data. Based on the aforementioned points, AST–HIS-induced apoptosis and autophagy could be connected during induction of cancer cell death via a mitochondria-mediated ROS-JNK-p53 pathway independent of Beclin-1 expression. As a result, p53 may switch the availability or presence of proteins that can influence tumor cell death fate. In summary, our results indicate that the combined use of AST and HIS, at a low toxicity profile, may synergize AST anti-proliferative effects by upregulating p53 phosphorylation and targeting p53p62 interaction. A clear evidence-based rationale is provided to test AST in combination with HIS as an adjuvant in the management of Beclin-1 independent autophagy by inducing ER stress, enhancing ROS generation, disrupting mitochondrial membrane potential, that leads to cell death in breast cancer cells. These findings support the continued exploration of AST–HIS as an alternative agent for breast cancer treatment.

Conflict of interest The authors declare that they have no conflict of interest.

Acknowledgement This research was supported by the Daegu University Research Grant, 2015.

Appendix: Supplementary material Supplementary data to this article can be found online at doi:10.1016/j.canlet.2015.12.024.

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