Skip to main content
NCBI home page
Toxicological Research logoLink to Toxicological Research
. 2022 Sep 2;38(4):591–600. doi: 10.1007/s43188-022-00150-4

Induction of synergistic apoptosis by tetramethoxystilbene and nutlin-3a in human cervical cancer cells

Hong-Gyu An 1, Sangyun Shin 1, Boyoung Lee 1, Yeonju Kwon 1, Tae-Uk Kwon 1, Yeo-Jung Kwon 1, Young-Jin Chun 1,✉
PMCID: PMC9532473  PMID: 36277372

Abstract

2,4,3’,5′-Tetramethoxystilbene (TMS) is a selective inhibitor of cytochrome P450 1B1 to block the conversion from estradiol to 4-OH-estradiol. Several studies suggested that TMS may act as a potent anti-cancer agent for hormone-related cancer including cervical cancer. Nutlin-3a is a cis-imidazoline analog that interferes with the interaction between mouse double minute 2 homolog (MDM2) and the tumor suppressor p53. The purpose of the study was to compare the cytotoxic effect of TMS and nutlin-3a treatment individually and in combination in HeLa cells. To assess the potential synergistic effects between TMS and nutlin-3a, low concentrations of TMS and nutlin-3a were simultaneously treated in HeLa cells. Based on cell viability, apoptosis assays, and the increase in cleaved caspase-3 and poly (ADP-ribose) polymerase cleavage, it was demonstrated that the combination with TMS and nutlin-3a exerts a synergistic effect on cancer cell death. Isobologram analysis of HeLa cells noted synergism between TMS and nutlin-3a. The combined treatment increased the expression of mitochondrial pro-apoptotic factors such as Bax and Bak, and decreased the expression of the XIAP. In addition, combination treatment significantly enhanced the translocation of AIF to the nucleus in HeLa cells. In conclusion, the results demonstrate that the combination of TMS and nutlin-3a induces synergistic apoptosis in HeLa cells, suggesting the possibility that this combination can be applied as a novel therapeutic strategy for cervical cancer.

Keywords: Tetramethoxystilbene, CYP1B1, nutlin-3a, p53, Combination treatment

Introduction

Cervical cancer is one of the most common gynecological malignancies in world and represents a global health challenge with 604,000 cases and 342,000 deaths in 2020, representing a critical threat to the health of women [1]. However, drug resistance and adverse effects are issues that need to be addressed in cervical cancer strategies [2]. To overcome the limitations of conventional treatment strategies, advanced treatment strategies including combined treatment are needed.

The protein p53 is well known as a tumor-suppressing transcription factor. Under carcinogenic conditions such as oncogene activation, hypoxia, or DNA damage, regulation of p53 expression leads to various cellular programs including cell cycle arrest, senescence, DNA repair, or apoptosis [3]. In cervical cancer, reinstating p53 has also become an essential but challenging object of study in anti-cancer strategies [4]. MDM2 is an E3 ubiquitin ligase that binds p53 together with its homolog MDMX (also known as MDM4). MDM2 inhibits the trans-activation of p53 through repression of nuclear translocation, targeting p53 with proteasomal degradation [5]. It is estimated that around 50% of cancers include wild-type TP53 genes encoding a functional p53 protein, thus inactivation of p53 is an important step in cancer progression [6]. However, p53 is often not activated predominantly due to the overexpression of the MDM2 [7]. Furthermore, p53-mediated cell cycle arrest and apoptosis are often inhibited due to increased MDM2 activity induced by viral oncoproteins E6 and E7 in cervical cancer. Thus, one of the conventional therapeutic strategies for cervical cancer is the activation of wild-type p53 by inhibition of MDM2 [8].

Nutlin-3a (Fig. 1A) is one of the MDM2 antagonists which inhibits p53-MDM2 interaction, preventing MDM2-mediated p53 degradation in wild-type p53 expressed cancer cells [9]. In this reason, induction of cell cycle arrest and apoptosis by treatment with MDM2 antagonists such as nutlin-3a is restricted to only wild-type p53 cancer [10]. Therefore, MDM2 antagonists have been tested in combinations with several other chemical agents in a search for synergy that would improve the anti-cancer activity, especially in cervical cancer [4]. Various studies examining the efficiency of combination therapy using nutlin-3a with DNA damaging drugs such as cisplatin and carboplatin have been conducted [4, 11].

Fig. 1.

Fig. 1

Open in a new tab

Chemical structures. A nutlin-3a. B 2,4,3′,5’-Tetramethoxy-trans-stilbene (TMS)

The cytochrome P450 (CYP) enzymes are involved in the metabolic activation of a diverse range of endogenous compounds and xenobiotics [12]. CYP1B1, observed in the cancer cells, belongs to the CYP1 family and is mainly implicated in the hydroxylation of estrogen and the activation of carcinogens [13]. 2,4,3′,5′-Tetramethoxystilbene, TMS (Fig. 1B), a methoxy derivative of oxyresveratrol, has been found to function as a selective inhibitor of CYP1B1 [14]. It has been reported that TMS is able to enhance apoptosis in MCF-7 and HL-60 cells, increasing apoptotic cells and chromosomal DNA fragmentation [15]. TMS induces a rise in the level of cell cycle inhibitor, p27kip1, through the reduction of Akt-mediated skp2 expression [16]. However, the detailed mechanisms of TMS-induced mitochondrial apoptosis have not been elucidated.

The aim of the present study was to investigate the synergistic effects of nutlin-3a and TMS on cell death in human cervical cancer cells, HeLa expressing wild-type p53. Intracellular mechanistic analysis was performed to reveal the effect of combination treatment with nutlin-3a. The results of this study might be suggested that an advanced strategy of combined treatment with the use of MDM2 antagonist, nutlin-3a, and CYP1B1 inhibitor, TMS could enhance cancer cell cytotoxicity with minimal adverse effects.

Materials and methods

Chemicals and reagents

2,4,3′,5′-Tetramethoxystilbene (TMS) was generously given by Dr. Sanghee Kim (Seoul National University, Seoul, Korea). Nutlin-3a was purchased from Sigma-Aldrich (St. Louis, MO, USA). α-MEM medium and fetal bovine serum (FBS) were obtained from HyClone (South Logan, UT, USA). The EZ-CyTox cell viability assay kit was purchased from Daeil Lab Service (Seoul, Korea). The enhanced chemiluminescence (ECL) kit and bicinchoninic acid (BCA) protein assay kit was obtained from Thermo Fisher Scientific (Waltham, MA, USA). Rabbit polyclonal antibody for Bak, mouse monoclonal antibodies for Bax and XIAP were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA), and rabbit polyclonal antibodies for caspase3, cleaved caspase3, and poly(ADP-ribose) polymerase (PARP) were purchased from Cell Signaling Technology (Beverly, MA, USA). All chemicals and reagents were of the highest quality commercially available.

Cell culture

Human cervical cancer cells HeLa were obtained from Korea Cell Line Bank (Seoul, Korea). The cells were cultured in α-MEM medium supplemented with 10% (v/v) heat-inactivated FBS, 100 µg/mL streptomycin, and 100 U/mL penicillin and retained at 37 °C in a humidified atmosphere of 5% CO2.

Cell viability assay

The cells (8 × 103 cells/well) were plated in 96-well microplates and incubated at 37 °C. The cells were treated with varying concentrations of TMS and nutlin-3a for 48 h. Following treatment, the cells were added to 10 µL of EZ-CyTox solution and incubated for 1 h at 37 °C. The produced formazan dyes were quantified by measuring the absorbance at 450 nm using the Tecan Sunrise™ microplate reader (Männedorf, Switzerland). All experiments were independently performed at least three times.

Western blot analysis

The cells were harvested by using scrapper and solubilized in ice-cold lysis buffer containing 50 mM Tris-HCl (pH 8.0), 1% Triton X-100, 150 mM NaCl, 0.5% sodium deoxycholate, 2 mM EDTA, 0.1% sodium dodecyl sulfate (SDS), 1 mM PMSF, and 10 mM NaF. Protein concentration was determined using the BCA Protein Assay Reagent. Extracted proteins (20 ~ 30 µg) were resolved on 8 ~ 12% SDS-PAGE and electrochemically transferred onto PVDF membranes. Nonspecific binding to membranes was then blocked with skimmed milk 5% in Tris-buffered saline containing 0.1% Tween-20 (TBS-T) for 2 h at 4 °C and incubated overnight with specific primary antibodies (1:1000 dilution). Subsequently, the membranes were incubated with secondary antibodies (1:5000 dilution) for 2 h at 4 °C. Protein bands were visualized by using the ECL method and detected using the ChemiDox XRS densitometer (Bio-Rad, CA, USA). Quantitative data were analyzed using Quantity One software (Bio-Rad).

Subcellular fractionation

After treatment, cells were harvested and washed with ice-cold PBS. Subcellular fractionation was performed using the mitochondria extraction kit, NE-PER® nuclear and cytoplasmic isolation kit for cultured cells (Thermo Scientific, Rockford, USA). Western blotting was carried out using the following antibodies against the indicated marker proteins as controls: COX-4 antibody for mitochondrial fraction, GAPDH antibody for cytosolic fraction, and Lamin A/C antibody for nuclear fraction.

Combination index (CI) values

The CI values were calculated using CompuSyn software (ComboSyn Inc., NJ, USA) to examine the interactions between TMS and nutlin-3a. CI values indicate synergistic (CI < 1), additive (CI = 1), or antagonistic (CI > 1) effects of diverse drug-drug concentrations [17]. The synergistic effects between TMS and nutlin-3a were evaluated via the isobologram method. Briefly, the IC50 value of each drug was separately plotted on the x- and y-axes to form a straight line. The data points in the isobologram correspond to the actual IC50 value of the combination of these chemicals. Data points on or near the line indicate an additive effect, those above the line represent antagonism, and those below the line indicate synergism.

Apoptosis assay

The cells were treated with the designated concentrations of TMS and nutlin-3a for 48 h, harvested with 0.05% trypsin-EDTA, and washed with PBS. The cells were then resuspended in PBS containing 5% BSA and stained with Luminex annexin V and dead cell reagent (Austin, TX, USA). Cells were detected by a MUSE cell analyzer (Merk Millipore).

Immunofluorescence

The cells seeded on poly D-lysine-coated coverslips were treated with the indicated chemical concentrations. The cells were fixed with 4% (w/v) formaldehyde in PBS for 30 min at 20 °C. After washing, the cells were blocked for 30 min in PBS containing 5% goat serum and 0.2% Triton X-100, and then incubated overnight with primary antibody for AIF (1:200). The cells were washed thoroughly and stained with Texas Red-conjugated goat anti-rabbit IgG (1:500) overnight. After the additional washing step, the coverslips were mounted onto glass slides using 4 µL UltraCruz™ Mounting Medium containing DAPI. Fluorescent signals were analyzed using an LSM800 Confocal Laser Scanning Microscope (Carl Zeiss, Jena, Germany).

Statistical analysis

Statistical analysis was performed using one-way analysis of variance (One-way ANOVA), followed by Dunnett’s Multiple Comparison t-test on Graph-Pad Prism 7 software (GraphPad Software Inc., CA, USA). Differences were considered statistically significant when *p < 0.05.

Results

TMS and nutlin-3a synergistically enhance cytotoxicity in HeLa cells

Prior to investigating the potential synergistic effect of TMS and nutlin-3a, the cytotoxicity of each drug in HeLa cells was determined by independent individual treatments. The results showed that IC50 values of TMS and nutlin-3a were 6.8 µM and 13.0 µM for HeLa cells, respectively (Fig. 2A , 2B). In these data, both chemicals indicated cancer cell cytotoxicity in a concentration-dependent manner. To investigate the synergistic effect of combining these agents in HeLa cells, 4 µM of TMS and 4 µM of nutlin-3a were selected as the optimal concentrations for combined treatment and each concentration inhibited cell growth by approximately 30 ± 10%. The cell viability of HeLa was reduced to 59.0 ± 3.3% by TMS (4 µM) and 76.8 ± 0.6% by nutlin-3a (4 µM) when each drug was utilized independently. The cell viability of HeLa was reduced to 28.1 ± 2.1% with the combination treatment. This was significantly higher compared to the cytotoxicity using each chemical individually (Fig. 2C).

Fig. 2.

Fig. 2

Open in a new tab

TMS and nutlin-3a synergistically enhance cytotoxicity in HeLa cells. A Inhibition of HeLa cell proliferation by TMS. The HeLa cells were treated with TMS (0, 1, 2.5, 5, 10, or 20 µM) for 48 h. The CCK assay was performed to determine cell viability. Data are representative of experiments performed in triplicate (*p ≤ 0.05). B Inhibition of HeLa cell proliferation by nutlin-3a. Cells were treated with nutlin-3a (0, 1, 5, 10, 20, or 40 µM) for 48 h. The CCK assay was performed to determine cell viability. All data are representative of experiments performed in triplicate (*p ≤ 0.05). C Cells were treated with a combination of TMS (4 µM) and nutlin-3a (4 µM) for 48 h. The CCK assay was performed to determine cell viability. All data are representative of experiments performed in triplicate (*p ≤ 0.05)

TMS and nutlin-3a synergistically increase apoptosis in HeLa cells

To determine whether enhanced apoptosis of combination treatment showed a synergistic effect, the CI value was calculated using IC50, and the slope value of the cell viability curve which was measured the effects of individual or combined treatments. Calculated CI values when cells were simultaneously treated with 4 µM of TMS and 4 µM of nutlin-3a were analyzed and found to be 0.4. Because this CI value was less than 1, the evidence indicated that combination treatment of TMS with nutlin-3a in HeLa cells was thus synergistic, as examined by a data point located far below the line in isobologram (Fig. 3A). Flow cytometry analysis also showed strong induction of apoptosis by the combination treatment in HeLa cells (Fig. 3B). These data suggested that the combination treatment of TMS and nutlin-3a induced synergistic growth inhibition in HeLa. Based on our findings, 4 µM of TMS and 4 µM of nutlin-3a were chosen as optimal concentrations to examine the induction of synergistic effects of these two drugs in HeLa cells. The concentration of these drugs induced moderate cytotoxicity when used alone but showed remarkable cytotoxicity when used in combination.

Fig. 3.

Fig. 3

Open in a new tab

Isobologram analysis of combination treatment with TMS and nutlin-3a. A Isobologram analysis of the anti-proliferative effects of combination treatment with TMS (4 µM) and nutlin-3a (4 µM) was performed and synergism was determined in the HeLa cells. B Cell apoptosis was detected by flow cytometry. The HeLa cells were co-treated with TMS (4 µM) and nutlin-3a (4 µM). The cells were stained Muse™ annexin V and dead cell reagent. After incubation for 30 min, cell viability and density were measured using the Muse™ cell analyzer. All data are representative of experiments performed in triplicate (*p ≤ 0.05)

The combination of TMS and nutlin-3a synergistically promotes PARP cleavage and caspase-3 activation

In order to examine alterations in apoptotic signaling attributed to the synergistic effect on cell growth inhibition, changes in PARP cleavage and caspase-3 activation were determined. The PARP is an important activator of caspase-independent apoptosis and acts as an AIF release inducer, eventually leading to high molecular weight DNA fragmentation [18]. Thus, a western blot was performed to examine the effect of the combination of TMS and nutlin-3a on proteolytic cleavage of PARP and caspase-3 activation. As shown in Fig. 4, individual treatment with TMS (4 µM) or nutlin-3a (4 µM) both showed a slight increase in cleavage of PARP and caspase 3. Combination treatment with TMS and nutlin-3a produced a significant elevation in PARP cleavage and caspase-3 cleavage in HeLa cells.

Fig. 4.

Fig. 4

Open in a new tab

The combination of TMS and nutlin-3a synergistically promotes PARP cleavage and caspase-3 activation. HeLa cells were co-treated with TMS (4 µM) and nutlin-3a (4 µM) for 48 h. After incubation, whole cell lysates were prepared. Extracted proteins were resolved by SDS-PAGE. Western blot analysis was conducted with antibodies against PARP and caspase 3. Uniform loading of proteins was confirmed by analysis of GAPDH in the protein extracts

TMS and nutlin-3a in combination synergistically induce apoptosis by increasing Bcl-2 family proteins and AIF translocation

Bax and its homolog, Bak are major regulators of the mitochondrial apoptosis pathway. When cells are subjected to stress conditions, Bax and Bak are accumulated in the mitochondrial surface and undergo structural changes, facilitating cytochrome c release which then stimulates apoptosis [19]. Additionally, Bax, interacting with VDAC, has been shown to be necessary for the release of AIF [20].

XIAP has received interest as a therapeutic target as it is probably the IAP member best characterized with respect to biochemical mechanism and construction [21–23]. Furthermore, XIAP is considered the only member of this family capable of directly inhibiting the initiation and execution phase of the caspase cascade, which is important for mediating the controlled death of malignant cells. Through its ability to inhibit caspases, overexpression of XIAP renders cells resistant to multiagent chemotherapy [24]. To investigate the association between combination treatment and mitochondrial-mediated apoptosis, protein expression of Bax, Bak, and XIAP was determined. As shown in Fig. 5, the combination of TMS and nutlin-3a significantly increased Bax and Bak protein expression whereas that of XIAP was significantly decreased in HeLa cells.

Fig. 5.

Fig. 5

Open in a new tab

TMS and nutlin-3a in combination synergistically induce apoptosis by increasing Bax, Bak and decreasing XIAP expression. HeLa cells were co-treated with TMS (4 µM) and nutlin-3a (4 µM) for 48 h. Western blot analysis was used to detect changes in protein levels using Bax, Bak, and XIAP antibodies. Uniform loading of proteins was confirmed by analysis of GAPDH in the protein extracts

Apoptosis-inducing factor (AIF) is a mitochondrial flavoprotein with NADH oxidase activity [25, 26]. AIF has been shown to be released from mitochondria to the cytosol, and then translocate to the nucleus when apoptosis is induced [25, 27]. To determine the specific molecular mechanisms underlying the effects of combining TMS with nutlin-3a, the translocation of AIF, a direct inducer of apoptosis associated with mitochondrial cell death and PARP cleavage, has been observed. Extra-mitochondrial AIF leads to cell death, whereas mitochondrial AIF is thought to be inactive, as far as apoptosis regulation is concerned [28]. The quantity of AIF in the mitochondria, cytoplasm, and nucleus was determined by western blot to clarify if combination treatment leads to nuclear translocation of AIF. Treatment with a combination of TMS and nutlin-3a confirmed that AIF localized in the mitochondria migrates to the cytoplasm and then to the nucleus (Fig. 6A). Confocal microscopy analysis also demonstrated that combination treatment promoted translocation of AIF into the nucleus, but TMS or nutlin-3a alone did not significantly alter the nuclear translocation of AIF (Fig. 6B,  7).

Fig. 6.

Fig. 6

Open in a new tab

The combination of TMS and nutlin-3a synergistically increases AIF translocation. A HeLa cells were co-treated with TMS (4 µM) and nutlin-3a (4 µM) for 48 h. After incubation, the mitochondrial, cytosolic, and nuclear fractions were separated. The isolated proteins were resolved by SDS-PAGE and western blot analysis was conducted with an AIF antibody. COX-4 level was determined as a loading control for mitochondria, GAPDH level was determined as a loading control for cytosol and total lysate, and lamin A/C level was determined as a loading control for the nucleus. B Confocal microscopy was performed to observe alterations in AIF expression. Microscopy scale bar = 20 μm

Fig. 7.

Fig. 7

Open in a new tab

Proposed mechanism by combined treatment with TMS and nutlin-3a

Discussion

To increase the therapeutic effect of existing cancer agents, several studies focused on the synergistic effect of cancer cell death through a combination of diverse drugs [4, 29]. The aim of combined treatment of anti-cancer drugs is to simultaneously inhibit different intracellular pathways that play an essential role in cell survival through concurrent inhibition, complementary inhibition, or sequential blockade, thereby significantly increasing cancer cell death [30]. There is a growing demand for an effective combination therapeutic strategy to acquire advantages such as increasing cytotoxicity and reducing adverse effects, and drug resistance [31, 32].

Nutlin-3a is a small-molecule inhibitor that disrupts the interaction between p53 and MDM2, known to promote the transcriptional activity of p53 by enhancing protein stability [33]. Previous studies have reported that nutlin-3a can be used in combination treatment as well as a single-dose regimen against wild-type p53-expressing cancers [34, 35]. It has been reported that MDM2 antagonists induce apoptosis only in a limited subset of wild-type p53 expressing cancer cells [36]. In many additional wild-type p53 cell lines, MDM2 antagonist induces cell cycle arrest, which is more of a form of reversible quiescence than irreversible senescence [37]. For this reason, several studies have focused on testing in combination with multiple other therapeutics in a search for a synergistic effect that would promote cancer cell death. However, in p53 mutant cells, cytotoxic effects of MDM2 antagonists including nutlin-3a showed limitations [4]. To overcome the limitations of MDM2 antagonist monotherapy, combined treatments have been studied. Combination treatment with cisplatin and nutlin-3a in sarcoma and non-small cell lung cancer cell lines revealed a clear synergistic effect that is more prone to apoptosis [38, 39]. In addition, the combination treatment of nutlin-3a with 1396 compound (XIAP inhibitor) induced strong activation of the apoptosis signaling pathway by increasing the p21 level [40]. Therefore, nutlin-3a is suitable for combination therapy for cancer. To develop an effective treatment strategy for hormone-related cancers including cervical cancer, combination partners to be administered with nutlin-3a were searched.

CYP1B1 showed a higher expression in hormone-related cancer such as breast cancer, prostate cancer, or ovarian cancer. Our previous studies reported that CYP1B1 promotes cell proliferation and metastasis by inducing EMT and Wnt/β-catenin signaling through induction of the sp1 transcription factor, and promotes cancer cell survival through the involvement of DNA methylation-mediated DR4 inhibition [41, 42]. In addition, CYP1B1 prevents proteasome-mediated XIAP degradation through the activation of PKCε signaling in cancer cells [43]. Thus, therapeutic strategies using inhibition of CYP1B1 could be promising for improving several limitations of the single-dose regime. After TMS was developed as a specific inhibitor of CYP1B1, it was shown that TMS inhibited microtubule formation, led to a cell cycle block at the G2-M phase, and induced apoptosis [44]. Furthermore, 8 weeks of treatment with TMS reduced tumor volume of tamoxifen-resistant MCF-7 breast cancer xenografts by 53% [45]. Thus, TMS is a promising therapeutic agent because of its specific ability to block several pathways involved in the development of hormone resistance. Additionally, TMS induces apoptosis in MCF-7 and HL-60 cancer cells in a concentration- and time-dependent manner and is able to induce mitochondrial apoptosis through increasing annexin A5 expression and translocation into mitochondria [15, 45]. Interestingly, our data showed potential synergistic effects induced by the combination of nutlin-3a and TMS. Current evidence suggested that the cytotoxicity of nutlin-3a to HeLa was significantly amplified by concurrent administration of TMS. In addition, co-administration of nutlin-3a with TMS significantly increased expression of Bax/Bak, cleaved caspase-3, and PARP cleavage, and decreased XIAP, suggesting the possibility of a synergistic action on mitochondrial-mediated apoptosis.

Our data also clearly showed that treatment with a combination of TMS and nutlin-3a promotes nuclear translocation of AIF from the mitochondria. Various studies have demonstrated that mitochondrial dysfunction and AIF release are associated with ROS overproduction [46]. A change in mitochondrial membrane permeability is required for AIF to be translocated in the nucleus and promotes cleavage of PARP, inducing mitochondria dysfunction [47]. Moreover, the down-regulation of AIF in various cancer cell lines, including MCF-7, decreased ROS levels [48]. p53 also promotes ROS-induced ferroptosis through a p21-dependent manner [49]. Increases in p21 expression also induce nuclear translocation of AIF [50]. In addition, previous studies have shown that suppression of CYP1B1 expression enhances ROS generation and inhibits cancer progression [51, 52]. Based on previous studies, combination treatment with nutlin-3a and TMS may induce apoptosis by ROS overproduction and subsequent AIF nuclear translocation. To elucidate the detailed molecular mechanism of combined treatment, further studies on ROS regulation are needed.

In this study, the data suggest a possible strategy using a combination of nutlin-3a and TMS for the treatment of cervical cancer. Such a combination strategy may be expected to improve the therapeutic efficacy of patients with limited susceptibility to single agents and other options.

Acknowledgements

This research was supported by the National Research Foundation of Korea (NRF) funded by the Korean government (MSIP) (Grant No.2021R1A2C201239512) and the Chung-Ang University Research Scholarship Grants in 2021. The funding agency had no role in the study design, data collection or analysis, the decision to publish, or the preparation of the manuscript.

Declarations

Conflict of interest

The authors declare no competing interests.

Footnotes

Hong-Gyu An and Sangyun Shin contributed equally to this study.

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

References

  • 1.Sung H, Ferlay J, Siegel RL, et al. Global cancer statistics 2020: GLOBOCAN estimates of incidence and mortality worldwide for 36 cancers in 185 countries. CA Cancer J Clin. 2021;71:209–249. doi: 10.3322/caac.21660. [DOI] [PubMed] [Google Scholar]
  • 2.Marth C, Landoni F, Mahner S, McCormack M, Gonzalez-Martin A, Colombo N. Cervical cancer: ESMO Clinical Practice Guidelines for diagnosis, treatment and follow-up. Ann Oncol. 2017;28:72–83. doi: 10.1093/annonc/mdx220. [DOI] [PubMed] [Google Scholar]
  • 3.Wang X, Simpson ER, Brown KA. p53: protection against tumor growth beyond effects on cell cycle and apoptosis. Cancer Res. 2015;75:5001–5007. doi: 10.1158/0008-5472.CAN-15-0563. [DOI] [PubMed] [Google Scholar]
  • 4.Kocik J, Machula M, Wisniewska A, Surmiak E, Holak TA, Skalniak L (2019) Helping the released guardian: drug combinations for supporting the anticancer activity of HDM2 (MDM2) antagonists. Cancers (Basel) 11:1014. 10.3390/cancers11071014 [DOI] [PMC free article] [PubMed]
  • 5.Davis JR, Mossalam M, Lim CS. Controlled access of p53 to the nucleus regulates its proteasomal degradation by MDM2. Mol Pharm. 2013;10:1340–1349. doi: 10.1021/mp300543t. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Joerger AC, Fersht AR. The p53 pathway: origins, inactivation in cancer, and emerging therapeutic approaches. Annu Rev Biochem. 2016;85:375–404. doi: 10.1146/annurev-biochem-060815-014710. [DOI] [PubMed] [Google Scholar]
  • 7.Khoo KH, Verma CS, Lane DP. Drugging the p53 pathway: understanding the route to clinical efficacy. Nat Rev Drug Discov. 2014;13:217–236. doi: 10.1038/nrd4236. [DOI] [PubMed] [Google Scholar]
  • 8.Nakamura M, Obata T, Daikoku T, Fujiwara H. The association and significance of p53 in gynecologic cancers: The potential of targeted therapy. Int J Mol Sci. 2019;20:5482. doi: 10.3390/ijms20215482. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Barbarotto E, Corallini F, Rimondi E, et al. Differential effects of chemotherapeutic drugs versus the MDM-2 antagonist nutlin-3 on cell cycle progression and induction of apoptosis in SKW6.4 lymphoblastoid B-cells. J Cell Biochem. 2008;104:595–605. doi: 10.1002/jcb.21649. [DOI] [PubMed] [Google Scholar]
  • 10.Tsao CC, Corn PG. MDM-2 antagonists induce p53-dependent cell cycle arrest but not cell death in renal cancer cell lines. Cancer Biol Ther. 2010;10:1315–1325. doi: 10.4161/cbt.10.12.13612. [DOI] [PubMed] [Google Scholar]
  • 11.Tonsing-Carter E, Bailey BJ, Saadatzadeh MR, et al. Potentiation of carboplatin-mediated DNA damage by the mdm2 modulator nutlin-3a in a humanized orthotopic breast-to-lung metastatic model. Mol Cancer Ther. 2015;14:2850–2863. doi: 10.1158/1535-7163.MCT-15-0237. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Guengerich FP. A history of the roles of cytochrome P450 enzymes in the toxicity of drugs. Toxicol Res. 2020;37:1–23. doi: 10.1007/s43188-020-00056-z. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Murray GI (2000) The role of cytochrome P450 in tumour development and progression and its potential in therapy. J Pathol 192:419-426. 10.1002/1096-9896(2000)9999:9999::AID-PATH7503.0.CO;2-0 [DOI] [PubMed]
  • 14.Chun YJ, Kim S, Kim D, Lee SK, Guengerich FP. A new selective and potent inhibitor of human cytochrome P450 1B1 and its application to antimutagenesis. Cancer Res. 2001;61:8164–8170. [PubMed] [Google Scholar]
  • 15.Chun YJ, Lee SK, Kim MY. Modulation of human cytochrome P450 1B1 expression by 2,4,3’,5’-tetramethoxystilbene. Drug Metab Dispos. 2005;33:1771–1776. doi: 10.1124/dmd.105.006502. [DOI] [PubMed] [Google Scholar]
  • 16.Kim SW, Jung HK, Kim MY. Induction of p27kip1 by 2,4,3’,5’- tetramethoxystilbene is regulated by protein phosphatase 2A-dependent Akt dephosphorylation in PC-3 prostate cancer cells. Arch Pharm Res. 2008;31:1187–1194. doi: 10.1007/s12272-001-1287-1. [DOI] [PubMed] [Google Scholar]
  • 17.Tallarida RJ. Drug synergism: its detection and applications. J Pharmacol Exp Ther. 2001;298:865–872. [PubMed] [Google Scholar]
  • 18.Yu SW, Wang H, Poitras MF, et al. Mediation of poly (ADP-ribose) polymerase-1-dependent cell death by apoptosis-inducing factor. Science. 2002;297:259–263. doi: 10.1126/science.1072221. [DOI] [PubMed] [Google Scholar]
  • 19.Cosentino K, García-Sáez AJ. Bax and bak pores: are we closing the circle. Trends Cell Biol. 2017;27:266–275. doi: 10.1016/j.tcb.2016.11.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Kuwana T, Mackey MR, Perkins G, et al. Bid, Bax, and lipids cooperate to form supramolecular openings in the outer mitochondrial membrane. Cell. 2002;111:331–342. doi: 10.1016/s0092-8674(02)01036-x. [DOI] [PubMed] [Google Scholar]
  • 21.Schimmer AD. Inhibitor of apoptosis proteins: translating basic knowledge into clinical practice. Cancer Res. 2004;64:7183–7190. doi: 10.1158/0008-5472.CAN-04-1918. [DOI] [PubMed] [Google Scholar]
  • 22.Shi Y. Caspase activation, inhibition, and reactivation: a mechanistic view. Protein Sci. 2004;13:1979–1987. doi: 10.1110/ps.04789804. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Wrzesień-Kuś A, Smolewski P, Sobczak-Pluta A, Wierzbowska A, Robak T. The inhibitor of apoptosis protein family and its antagonists in acute leukemias. Apoptosis. 2004;9:705–715. doi: 10.1023/B:APPT.0000045788.61012.b2. [DOI] [PubMed] [Google Scholar]
  • 24.Schimmer AD, Dalili S, Batey RA, Riedl SJ. Targeting XIAP for the treatment of malignancy. Cell Death Differ. 2006;13:179–188. doi: 10.1038/sj.cdd.4401826. [DOI] [PubMed] [Google Scholar]
  • 25.Susin SA, Lorenzo HK, Zamzami N, et al. Molecular characterization of mitochondrial apoptosis-inducing factor. Nature. 1999;397:441–446. doi: 10.1038/17135. [DOI] [PubMed] [Google Scholar]
  • 26.Lipton SA, Bossy-Wetzel E. Dueling activities of AIF in cell death versus survival: DNA binding and redox activity. Cell. 2002;111:147–150. doi: 10.1016/s0092-8674(02)01046-2. [DOI] [PubMed] [Google Scholar]
  • 27.Daugas E, Susin SA, Zamzami N, et al. Mitochondrio-nuclear translocation of AIF in apoptosis and necrosis. FASEB J. 2000;14:729–739. doi: 10.1096/fasebj.14.5.729. [DOI] [PubMed] [Google Scholar]
  • 28.Loeffler M, Daugas E, Susin SA, et al. Dominant cell death induction by extramitochondrially targeted apoptosis-inducing factor. FASEB J. 2001;15:758–767. doi: 10.1096/fj.00-0388com. [DOI] [PubMed] [Google Scholar]
  • 29.Mondesire WH, Jian W, Zhang H, et al. Targeting mammalian target of rapamycin synergistically enhances chemotherapy-induced cytotoxicity in breast cancer cells. Clin Cancer Res. 2004;10:7031–7042. doi: 10.1158/1078-0432.CCR-04-0361. [DOI] [PubMed] [Google Scholar]
  • 30.DeVita VT Jr, Young RC, Canellos GP (1975) Combination versus single agent chemotherapy: a review of the basis for selection of drug treatment of cancer. Cancer 35:98-110. 10.1002/1097-0142(197501)35:198::aid-cncr28203501153.0.co;2-b [DOI] [PubMed]
  • 31.Stanley A, Ashrafi GH, Seddon AM, Modjtahedi H. Synergistic effects of various Her inhibitors in combination with IGF-1R, C-MET and Src targeting agents in breast cancer cell lines. Sci Rep. 2017;7:3964. doi: 10.1038/s41598-017-04301-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Fan C, Zheng W, Fu X, Li X, Wong YS, Chen T. Enhancement of auranofin-induced lung cancer cell apoptosis by selenocystine, a natural inhibitor of TrxR1 in vitro and in vivo. Cell Death Dis. 2014;5:e1191. doi: 10.1038/cddis.2014.132. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Turner N, Moretti E, Siclari O, et al. Targeting triple negative breast cancer: is p53 the answer? Cancer Treat Rev. 2013;39:541–550. doi: 10.1016/j.ctrv.2012.12.001. [DOI] [PubMed] [Google Scholar]
  • 34.Kojima K, Konopleva M, Samudio IJ, et al. MDM2 antagonists induce p53-dependent apoptosis in AML: implications for leukemia therapy. Blood. 2005;106:3150–3159. doi: 10.1182/blood-2005-02-0553. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Hong B, van den Heuvel AP, Prabhu VV, Zhang S, El-Deiry WS. Targeting tumor suppressor p53 for cancer therapy: strategies, challenges and opportunities. Curr Drug Targets. 2014;15:80–89. doi: 10.2174/1389450114666140106101412. [DOI] [PubMed] [Google Scholar]
  • 36.Tovar C, Rosinski J, Filipovic Z et al (2006) Small-molecule MDM2 antagonists reveal aberrant p53 signaling in cancer: implications for therapy. Proc Natl Acad Sci USA 103:1888–1893. 10.1073/pnas.0507493103 [DOI] [PMC free article] [PubMed]
  • 37.Huang B, Deo D, Xia M, Vassilev LT. Pharmacologic p53 activation blocks cell cycle progression but fails to induce senescence in epithelial cancer cells. Mol Cancer Res. 2009;7:1497–1509. doi: 10.1158/1541-7786.MCR-09-0144. [DOI] [PubMed] [Google Scholar]
  • 38.Deben C, Wouters A, Op de Beeck K, et al. The MDM2-inhibitor nutlin-3 synergizes with cisplatin to induce p53 dependent tumor cell apoptosis in non-small cell lung cancer. Oncotarget. 2015;6:22666–22679. doi: 10.1158/1541-7786.MCR-09-0144. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Ohnstad HO, Paulsen EB, Noordhuis P, et al. MDM2 antagonist nutlin-3a potentiates antitumour activity of cytotoxic drugs in sarcoma cell lines. BMC Cancer. 2011;11:1–11. doi: 10.1186/1471-2407-11-211. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40.Carter BZ, Mak DH, Schober WD, et al. Simultaneous activation of p53 and inhibition of XIAP enhance the activation of apoptosis signaling pathways in AML. Blood. 2010;115:306–314. doi: 10.1182/blood-2009-03-212563. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 41.Kwon YJ, Baek HS, Ye DJ, Shin S, Kim D, Chun YJ. CYP1B1 enhances cell proliferation and metastasis through induction of EMT and activation of Wnt/β-catenin signaling via sp1 upregulation. PLoS One. 2016;11:e0151598. doi: 10.1371/journal.pone.0151598. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 42.Kwon YJ, Cho NH, Ye DJ, Baek HS, Ryu YS, Chun YJ. Cytochrome P450 1B1 promotes cancer cell survival via specificity protein 1 (Sp1)-mediated suppression of death receptor 4. J Toxicol Environ Health A. 2018;81:278–287. doi: 10.1080/15287394.2018.1440186. [DOI] [PubMed] [Google Scholar]
  • 43.Baek HS, Kwon YJ, Ye DJ, Cho E, Kwon TU, Chun YJ. CYP1B1 prevents proteasome-mediated XIAP degradation by inducing PKCε activation and phosphorylation of XIAP. Biochim Biophys Acta Mol Cell Res. 2019;1866:118553. doi: 10.1016/j.bbamcr.2019.118553. [DOI] [PubMed] [Google Scholar]
  • 44.Park HS, Aiyar SE, Fan P, et al. Effects of tetramethoxystilbene on hormone-resistant breast cancer cells: biological and biochemical mechanisms of action. Cancer Res. 2007;67:5717–5726. doi: 10.1158/0008-5472.CAN-07-0056. [DOI] [PubMed] [Google Scholar]
  • 45.Hong M, Park N, Chun YJ. Role of annexin a5 on mitochondria-dependent apoptosis induced by tetramethoxystilbene in human breast cancer cells. Biomol Ther (Seoul) 2014;22:519–524. doi: 10.4062/biomolther.2014.112. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Yan C, Huang D, Zhang Y. The involvement of ROS overproduction and mitochondrial dysfunction in PBDE-47-induced apoptosis on Jurkat cells. Exp Toxicol Pathol. 2011;63:413–417. doi: 10.1016/j.etp.2010.02.018. [DOI] [PubMed] [Google Scholar]
  • 47.Yu SW, Andrabi SA, Wang H et al (2006) Apoptosis-inducing factor mediates poly(ADP-ribose) (PAR) polymer-induced cell death. Proc Natl Acad Sci USA 103:18314–18319. 10.1073/pnas.0606528103 [DOI] [PMC free article] [PubMed]
  • 48.Urbano A, Lakshmanan U, Choo PH, et al. AIF suppresses chemical stress-induced apoptosis and maintains the transformed state of tumor cells. EMBO J. 2005;24:2815–2826. doi: 10.1038/sj.emboj.7600746. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49.Kuganesan N, Dlamini S, Tillekeratne LMV, Taylor WR. Tumor suppressor p53 promotes ferroptosis in oxidative stress conditions independent of modulation of ferroptosis by p21, CDKs, RB, and E2F. J Biol Chem. 2021;297:101365. doi: 10.1016/j.jbc.2021.101365. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 50.Delavallée L, Mathiah N, Cabon L, et al. Mitochondrial AIF loss causes metabolic reprogramming, caspase-independent cell death blockade, embryonic lethality, and perinatal hydrocephalus. Mol Metab. 2020;40:101027. doi: 10.1016/j.molmet.2020.101027. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 51.Palenski TL, Sorenson CM, Jefcoate CR, Sheibani N. Lack of Cyp1b1 promotes the proliferative and migratory phenotype of perivascular supporting cells. Lab Invest. 2013;93:646–662. doi: 10.1038/labinvest.2013.55. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52.Kwon YJ, Shin S, Chun YJ. Biological roles of cytochrome P450 1A1, 1A2, and 1B1 enzymes. Arch Pharm Res. 2021;44:63–83. doi: 10.1007/s12272-021-01306-w. [DOI] [PubMed] [Google Scholar]

Articles from Toxicological Research are provided here courtesy of Korean Society of Toxicology

RESOURCES