Modulatory Effects of Cirsimarin on Lung Cancer A549 Cells Migration in 2D and 3D Cultures Involves Transcriptional Regulation of Metalloproteinases
Anna Gabriele Prado dos Santos, Celina Yung‐Ai Lin Lee, Érica Romão Pereira, Andresa Hiromi Sakai, Diego Luís Ribeiro, Victor Antônio Silva Lima, Marcos Bispo Pinheiro Camara, Cláudia Quintino da Rocha, Sabine A. S. Langie, Ilce Mara de Syllos Cólus, Juliana Mara Serpeloni
TL;DR
This study shows that cirsimarin, a flavone, inhibits lung cancer cell migration and viability in 2D and 3D models by affecting genes related to cell growth and matrix remodeling.
Contribution
The novel contribution is the identification of cirsimarin's antimetastatic potential through transcriptional regulation of metalloproteinases in lung cancer cells.
Findings
Cirsimarin reduced A549 cell viability and migration in 2D and 3D models.
It induced apoptosis and DNA damage at specific concentrations.
Downregulation of genes like MMP-2, MMP-9, and TP53 was observed.
Abstract
Current toxicology and cancer biology investigations have focused on developing alternative models that better recapitulate the in vivo architecture of tissues and organs. The present study evaluated the anticancer effects of the flavone cirsimarin, which presented successful antitumor activity on breast tumor cells. We assessed the impact of flavone on cell viability, proliferation, and migration, as well as on DNA integrity and modulation of related cellular pathways. In the 2D model, cirsimarin reduced cell viability at concentrations ≥ 80 μM after 24 h of treatment (resazurin assay), selectively in A549 cells compared to MRC‐5 non‐tumor cells. Apoptosis was induced at concentrations ≥ 40 μM, and clonogenicity was reduced by approximately 50% only at 160 μM. In the wound healing assay, cirsimarin (1–80 μM) completely inhibited cell migration and induced DNA damage (comet assay).…
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Taxonomy
TopicsSilymarin and Mushroom Poisoning · Curcumin's Biomedical Applications · Flavonoids in Medical Research
Introduction
1
Lung cancer, the most commonly diagnosed cancer in 2022, accounted for approximately 2.5 million new cases, or one in every eight cancer cases worldwide [1]. Furthermore, it was recognized as the leading cause of cancer‐related deaths worldwide, with an estimated 1.8 million fatalities [1]. Smoking, the primary risk factor for this type of cancer, is responsible for around 85% of diagnoses [2]. However, non‐smokers can also be affected by this neoplasm due to passive smoking [3, 4] and factors such as a sedentary lifestyle, obesity, poor diet, and genetic and/or occupational factors [5, 6]. The urgency of the rising incidence of lung cancer demands immediate attention and more effective treatments.
The histologic types of lung cancer are non‐small cell lung carcinoma (NSCLC), representing around 80%−85% of diagnoses, and small cell lung carcinoma (SCLC), which accounts for 15%−20% of cases [7]. NSCLC is subdivided into adenocarcinoma (the most prevalent), squamous, and large‐cell carcinomas [8]. The tumor type and stage determine the methods currently used to treat these tumors, including surgery, chemotherapy, radiotherapy, and immunotherapy [9, 10, 11]. However, the limitations of these methods underscore the pressing need for new therapeutic approaches. Cisplatin (cDDP) is the gold standard of first‐line chemotherapy for lung cancer treatment. However, it often leads to chemoresistance, resulting in treatment failure and limiting its application and efficacy [9, 12]. In addition, its use can cause adverse effects, such as nephrotoxicity, hepatotoxicity, gastrointestinal problems, and neurotoxicity [13].
Given the increasing incidence of lung cancer and the need for selective therapies to combat chemoresistance, the search for new therapeutic approaches is essential [14]. Natural products have been a rich source of bioactive compounds, and more than 60% of current anticancer drugs are derived from natural sources. In this regard, plant‐derived chemotherapeutics, such as vinblastine, vincristine, paclitaxel, docetaxel (DTX), and irinotecan, have emerged as valuable alternatives in the ongoing search for new cancer treatments. For example, the vinca alkaloids (vinblastine and vincristine), extracted from the Madagascar periwinkle (Catharanthus roseus), taxanes (paclitaxel and DTX), sourced from the Pacific yew tree (Taxus brevifolia), and topotecan and irinotecan, from the Chinese tree Camptotheca acuminata, are plant‐derived compounds currently used in cancer chemotherapies [15, 16, 17].
The flavone cirsimarin, which can be easily isolated from the plant Scoparia dulcis Linn, is an herb with a rich regional history of medicinal use for various diseases, including respiratory ones [18], which has sparked our interest. Recent research from our laboratory has unveiled its potential as an anticancer agent, as it reduced the cell migration of breast (MCF‐7) tumor cells in vitro by downregulating metalloproteinases (MMPs) [19]. If this finding is also observed in lung tumor cells, it could be a beacon of hope in the battle against lung cancer, as high expression of MMPs has been linked primarily with poor survival, high clinical stages, and metastasis of lung cancer [20]; furthermore, the development of individualized MMP inhibitors could be an exciting strategy in lung cancer therapy [21].
Thus, in the current study, we investigate the antitumor effects of cirsimarin on cell viability, proliferation, migration, and DNA stability, as well as the modulation of gene expression in A549 non‐small cell lung cancer (NSCLC) cells. These effects are examined in vitro in two‐dimensional (2D) and three‐dimensional (3D) cell culture models. Our goal is to determine whether cirsimarin can serve as a basis for developing a new therapeutic approach that is more effective and less toxic for treating lung cancer.
Materials and Methods
2
Collection, Extraction, Isolation, and Identification of Flavone Cirsimarin
2.1
The aerial parts of S. dulcis Linn were collected at the Cidade Universitária Dom Delgado of the Federal University of Maranhão (UFMA), city of São Luís, MA, Brazil (Latitude: 02° 33′8″ S, Longitude: 44° 18′ 25″ W, and Altitude: 24 m) in September 2021, under similar weather and temperature conditions. The species was identified in the herbarium “Atico Seabra” of the UFMA under exsicata N°. 1528. The plant name was verified on http://www.theplantlist.org, and the samples were collected in accordance with Brazilian legislation for the protection of biodiversity (SisGenN° AABBCA4). The samples were dried in an oven with forced air circulation at 45°C under constant weight and pulverized in a knife mill to obtain a moderately thick powder. The obtained powder, extracted by maceration, was subjected to a drug/solvent ratio of 1:3 (w/v) with ethanol (99.5%) for 48 h. The extractive solution was filtered and concentrated in a rotary evaporator under vacuum (40°C) to obtain a 70% ethanol extract. The cirsimarin (C_23_H_24_O_11_) was obtained and identified as previously described by Serpeloni et al. [19]. The cirsimarin was diluted 1:1 (v/v) with dimethyl sulfoxide (DMSO) and phosphate‐buffered saline (PBS; pH 7.4). The work concentrations (1−160 μM) were chosen based on the results obtained by Serpeloni et al. [19] and the definition of the stock solution (64,000 μM) that was kept frozen.
Cell Culture
2.2
The A549 (NSCLC) cell line was obtained from the American Type Culture Collection (ATCC; Cat. N° CCL‐185; Manassas, VA, USA). The authenticity of the cell line was validated by the Paternity Investigation Program through DNA Analysis (PIPAD) (Supporting Information S1: Figure 1), coordinated by Prof^a^. Dr. Karen Brajão de Oliveira. The MRC‐5 cell line obtained from the lung biopsy of a healthy individual (human fibroblasts) was kindly donated by Prof. Dr. Carlos Frederico Menck. It was used only to calculate the selectivity index in the resazurin assay. Both cell lines were cultured in Dulbecco's Modified Eagle Medium (DMEM; Cat. N° 31600‐034, Thermo Fisher Scientific) supplemented with 10% fetal bovine serum (FBS; Cat. N° 10‐bio500, Nova Biotecnologia, Cotia, SP, Brazil) and 1% antibiotic‐antimycotic solution 100X (15240062, Gibco, USA). The cultures were maintained in a humidified incubator (PHCBI MC0‐170AICUVL‐PA) at 37°C with an atmosphere of 5% CO_2_ and 95% relative humidity, following the protocols recommended by Aschner, Suñol, and Bal‐Price [22]. The solvent control (SV) consisted of a 1:1 solution of DMSO and PBS, with final concentrations of 0.5% for 2D cultures and 0.25% for 3D cultures. DTX was used as a positive control (PC) at concentrations of 50 µM for 2D cultures and 92.835 µM for 3D cultures.
Experiments in 2D Monolayer Culture Model
2.3
A basic representation of the workflow used to generate and analyze 2D and 3D in vitro experiments is represented in Figure 1.
*Overview of the various 2D and 3D experiments in A549 cells. Briefly, cirsimarin was evaluated in 2D and 3D culture models. The 2D model evaluated the effects of cirsimarin on cell viability, death induction, horizontal migration, survival, genotoxicity, and gene expression. The resazurin assay was also performed at 24 h in MRC‐5 cells to calculate the selectivity index. In 3D spheroids, the tests evaluated cell migration in the extracellular matrix, cell viability, and assessment of the spheroid's growth, morphology, and integrity.
Cell Viability Assays
2.3.1
A549 (1 × 10^4^) cells were seeded in 96‐well plates (Cat. N° 011096; JetBiofil, Guangzhou, China) for cell viability assays. After a stabilization period of 24 h, the cells were washed with PBS and treated with cirsimarin at concentrations ranging from 1 to 160 µM for 24, 48, and 72 h in a culture medium without FBS. The exposure period should be at least the duration of one cell cycle since the sensitivity of cell stages to the test substances may differ [23]. Thus, we chose to use one, two, and three cell cycle times, considering that, according to ATCC, the A549 doubling time is around 22−24 h. The resazurin assay was performed according to Walzl et al. [24]. For this assay, the MRC‐5 lineage was also used, with 1 × 10^4^ cells seeded in 96‐well plates, treated with the same concentrations for 24 h to compare CC_50_ values. After the treatments, 40 µL of a 0.15 mg/mL resazurin solution diluted in PBS was added per well, and the plate was incubated for 4 h. The absorbance was then measured using a spectrophotometer (Biotek Elx800, Winooski, USA) at λ = 570 nm (resorufin) and λ = 600 nm (resazurin). The absorbance values were adjusted using the oxidation factors at each wavelength (117,216 for 570 nm and 80,586 for 600 nm). Cell viability was calculated as the difference between the absorbance values at 570 and 600 nm.
The APH assay was performed according to the protocol of Friedrich et al. [25]. After treatments, wells were washed with PBS and 100 µL of APH buffer (0.7 mL of 0.1 mol/L sodium acetate, 0.02 mL of Triton X‐100, 21.4 mL of distilled water, and 44 mg of immunopure p‐nitrophenyl phosphate; Sigma‐Aldrich). The plates were incubated for 90 min at 37°C in a humidified incubator with 5% CO_2_. Subsequently, 10 µL of 1 N NaOH was added to each well, and the absorbance was measured at 405 nm using a spectrophotometer (Biotek Elx800, Winooski, USA). Data were normalized to the SV for both assays and set as 100% cell viability.
Other Cell Viability/Death Assays
2.3.2
Lactate Dehydrogenase (LDH) Release Assay
2.3.2.1
The Pierce LDH Cytotoxicity Assay Kit (Thermo Scientific, Cat. N° 88953) was used for the LDH assay. The plates with the A549 cells were assembled as described for the viability assays. After adding the treatments (cirsimarin 1−160 µM) for 48 and 72 h, 50 µL of the supernatant was removed and added to another plate for analysis, following the manufacturer's recommendations. The PC group treated the cells with 10 μL lysis solution from the kit. The absorbances were measured at λ = 440 nm and λ = 680 nm using a spectrophotometer (Biotek Elx800, Winooski, USA). The absorbance values measured at 680 nm were subtracted from those obtained at 490 nm to determine cytotoxicity. The data were then normalized, considering the SV 100% cell viability.
Triple Fluorescent Staining Assay
2.3.2.2
For cell death analysis, A549 (5 × 10^4^) cells were seeded in 24‐well plates (Cat. N° K12‐024; Kasvi, Guangzhou, China), stabilized for 24 h, and treated with cirsimarin (1−160 µM) for 24 h. After treatments, the culture medium was retained, the monolayer cells were washed with PBS, and this volume was transferred to a 15 mL tube (Falcon) containing the retained culture medium. The Falcon tube containing the cells was then centrifuged at 2500 rpm for 5 min to obtain the pellet of dead cells. The cells adhered to the plate were then trypsinized, inactivated, and combined with the previously obtained pellets. The cells were centrifuged at 2500 rpm for 5 min, and then the supernatant was discarded. Cell death was analyzed by triple staining the cells with the dyes propidium iodide (PI; Cat. N° 4170: Sigma‐Aldrich), Hoescht 33342 (Ho; Cat. N° 14533, Sigma‐Aldrich), and fluorescein diacetate (FDA; Cat. N° 191661000, Acros Organics). The suspension was homogenized, and 8 μL of the freshly prepared fluorochrome mixture was added to 30 μL of the suspension (final concentration in culture of 4.0 μg/mL Ho, 7.5 μg/mL FDA, and 1.0 μg/mL of PI). At analysis, 100 cells per slide (300 cells in total) were analyzed under an Olympus BX 43 fluorescence microscope (Olympus Microscopy, Europe) using a ×40 objective, recording the number of viable cells (FDA+ and Ho+, without apoptotic bodies), necrotic cells (PI+), and apoptotic cells (FDA+, with apoptotic bodies and Ho+), and PI− (early apoptosis) and PI+ (late apoptosis) [26].
Cell Survival (Clonogenic Assay)
2.3.3
The clonogenic assay was performed according to Franken et al. [27] to assess cell survival. First, A549 (5 × 10) cells were seeded in 24‐well plates (Cat. N° K12‐012; Kasvi, Guangzhou, China) and stabilized in a complete culture medium for 24 h. Subsequently, the cells were treated with cirsimarin (1−160 µM) for 24 h without FBS. After treatment, the cells were trypsinized, and 300 viable cells were transferred to 6‐well plates (Cat. N° K12‐006; Kasvi, Guangzhou, China) containing a complete medium. The plates were incubated for 10 days. After this period, the cells were washed with 1 mL of PBS and fixed with acetic acid, methanol, and distilled water (1:1:8) for 30 min. The cells were stained with 5% Giemsa for 5 min, washed with distilled water, and air‐dried at room temperature. Colonies were counted using a Leica ES2 Microsystems (Schweiz) binocular stereo microscope, with a colony defined as containing approximately 50 cells. The results were normalized to the SV, set as 100% cell survival.
Cell Migration (Wound Healing Assay)
2.3.4
The cell migration using the wound healing assay was performed according to the protocols of Liang et al. [28] and Martinotti and Ranzato [29]. Briefly, A549 (6 × 10^5^) cells were seeded in 12‐well plates (Cat. N° K12‐012; Kasvi, Guangzhou, China) and incubated for 24 h to form a confluent monolayer. A groove was created by scraping the monolayer with a 200 µL pipette tip, and the wells were washed with PBS to remove detached cells. Treatments with cirsimarin (1−40 μM) and controls, including a PC of cytochalasin (2 mg/mL; Cat. N° C6762; Sigma‐Aldrich), were added in complete medium. Images were captured at 0, 24, 48, and 72 h post‐treatment using the Q Color 3 image capture system (Olympus, Canada) coupled to an inverted Olympus CKX41 microscope (Olympus, Tokyo, Japan) with a ×4 objective. The resolution specifications of the Q Color 3 image capture system are 2080 × 1542 active pixels and 3.2 megapixels. Image analysis was conducted using ImageJ software [30] with the Wound_healing_size_tool [31]. The results were expressed as the percentage of wound closure (%), with the initial groove width at 0 h considered 100% within the same treatment.
Genotoxicity (Comet Assay)
2.3.5
The comet assay was performed according to Collins et al. [32] in two gels/slide format and adherence to the recommendations of the Minimum Information for Reporting on the Comet Assay [33]. Initially, A549 (1 × 10^5^) cells were seeded in 24‐well plates and incubated with culture medium supplemented with 10% FBS for 24 h for cells to adhere and stabilize. After this, the culture medium was removed and replaced with serum‐free medium plus the respective treatments: SV, PC (10 s in ultraviolet C light), and cirsimarin concentrations (1−80 µM). Ionizing radiation is a standard PC because it induces single‐strand breaks [34], cyclobutane pyrimidine dimers, or bulky adducts [35, 36]. After 4 h of treatment, the medium was removed, and the cells were washed with PBS, trypsinized, and centrifuged for 5 min at 1000 rpm. The supernatant was discarded, the pellet was resuspended in 100 µL of PBS, and 20 µL of this solution was resuspended with 80 µL of low‐melting‐point agarose (final concentration 0.5%). Subsequently, two drops of 70 µL were placed onto a 1% NMP agarose pre‐coated glass microscope slide (previously dried). The drops of gel were covered with 20 × 20 mm coverslips, and the slides were kept at 4°C for 10 min to solidify the gel. The coverslips were removed, and the slides were placed into a Coplin jar with cold lysis work solution (100 mL stock solution with 2.5 M NaCl, 100 mM EDTA, 10 mM Tris, 10 mL DMSO, 1 mL Triton X‐100, pH 10) for 24 h at 4°C. The slides were quickly immersed in distilled water and submerged in an alkaline solution (300 mM NaOH, 1 mM EDTA, pH > 13) in an electrophoresis tank for 20 min at 4°C, followed by electrophoresis in the same solution for 20 min (1 V/cm). After electrophoresis, the slides were immersed in a neutralization solution (0.4 M Tris and pH 7.5) for 15 min (3 × 5 min) and fixed in absolute ethanol (99%) for 5 min. For analysis, the slides were stained with GelRedTM (Biotium, Fremont, CA, USA), and 100 comets per slide were analyzed using an Olympus BX 43 fluorescence microscope (Olympus Microscopy, Europe) under a ×40 objective. The number of classes 0, 1, 2, 3, and 4 was noted, according to Møller et al. [37], ensuring a comprehensive analysis. All results on visual scores in the present paper are reported in the 0–400 arbitrary units (a.u.) range, using the following equation:
DNA damage index = 0.(n class 0) + 1.(n class 1) + 2.(n class 2) + 3.(n class 3) + 4.(n class 4).
Gene Expression (RT‐qPCR)
2.3.6
For gene expression analysis, A549 (2 × 10^5^) cells were seeded in 12‐well plates, stabilized for 24 h, and treated with SV, cirsimarin 40, or cirsimarin 80 µM for 12 h. Following the treatments, the supernatant was carefully removed, and the cells were trypsinized and transferred to 1.5 mL microtubes for RNA extraction using the PureLink RNA Mini Kit (Cat. N° 12183018 A; Thermo Scientific, Carlsbad, CA, USA), adhering strictly to the manufacturer's protocol. The total RNA was then quantified using the Nanodrop 2000 spectrophotometer (Thermo Fisher Scientific, USA). The reverse transcriptase technique used for complementary DNA (cDNA) synthesis was the Superscript III Kit (Cat. N° 56575; Invitrogen; Carlsbad, CA, USA), a method known for its thoroughness, using 500 ng of RNA, oligo‐DT, and random primers and RT‐qPCR using GoTaq qPCR Master Mix (Cat. N° A6001; Promega, USA) and StepOnePlus System thermocycler (Applied Biosystems). The target mRNAs for modulation and the primers for cDNA amplification were defined according to the results obtained in the functional assays. The final reaction volume was 11.5 µL, using 5 pmol of each primer oligonucleotide, 10 ng of template cDNA, and 5 µL of GoTaq qPCR Master Mix, and the volume was completed with water. The reaction mixture was subjected to the following amplification program: 95°C for 5 min; 40 cycles of 15 s at 95°C (denaturation), 15 s at 60°C (annealing), and 30 s at 72°C (extension). All reactions were performed in technical triplicate. Relative expression was calculated using the 2‐ΔΔCt method, according to Schmittgen and Livak [38]. Primers for genes associated with cell death (TP53, TNF‐α, and CASP9) and migration (MMP‐2, MMP‐9, and MMP‐11) were purchased from Sigma‐Aldrich (KiCqStart SYBR Green Primers) and normalized with the reference genes (GAPDH and HPRT1), which were chosen for their stability in the experimental conditions, according to the NormFinder program [39]. The primer sequences are represented in Supporting Information S1: Table 1.
Experiments in 3D Multicellular Tumor Spheroids (MCTS)
2.4
MCTSs were obtained using the method proposed by Friedrich et al. [40]. Briefly, spheroid initiation was performed in 96‐well plates pre‐coated with 1.5% agarose. To prepare the coating, 0.45 g of normal melting point (NMP) agarose (Cat. N° 16500‐100; Invitrogen, Carlsbad, CA, USA) was dissolved in 33 mL of culture medium (without FBS). The solution was autoclaved for 20 min at 121°C, then transferred to a laminar flow hood while still hot. Subsequently, 50 μL of this solution was pipetted into 96‐well plates using a Multipette M4 multipipettor (Eppendorf, Hamburg, Germany). After agarose solidification, 200 μL of complete DMEM medium containing A549 cells (4 × 10^3^) were seeded into each well. The plates were centrifuged in a microplate centrifuge (Mini P25) for 5 min, then transferred to a culture incubator with 5% CO_2_ at 37°C and 96% humidity and left undisturbed for 2 days to form 3D tumor spheroids.
Assessment of Growth, Morphology, and Integrity of 3D Tumor Spheroids
2.4.1
The growth, morphology, and integrity of A549 3D tumor spheroids were analyzed following the procedures outlined by Friedrich et al. [40] and Vinci et al. [41]. The photomicrographs were taken at 0 h (Day 2 after plating), 72 h, 144 h, and 216 h using an Olympus CKX41 inverted microscope with a ×4 objective and Qimaging Pro 7.1 software (Teledyne, Canada). Treatments with SV (DMSO 0.25%), PC (DTX 92.835 uM), and cirsimarin (1−160 µM) were refreshed at each time point. After capturing the images, 100 μL of the medium was removed from each well, and 100 μL of 2X concentrated treatments diluted in DMEM were added.
The integrity and morphology of the 3D tumor spheroids were evaluated by detecting irregular spheroids (non‐circular shapes), cell disaggregation, or irregular cell agglomeration. The circumference of each spheroid was measured using Zen 2.3 software (Zeiss, Germany) with the “Measure” tool to determine the area, presented in square micrometers (μm²). At the end of 216 h, cell viability was assessed using the resazurin assay, as described in Section 2.3‐1. Data were normalized as described for the 2D model.
Extracellular Matrix (ECM) 3D Migration Assay
2.4.2
The cell migration assay was conducted according to Vinci et al. [41]. Briefly, 50 µL of a 0.1% (w/v) bovine skin gelatin solution (Cat. N° G9391‐100G; Sigma‐Aldrich) were pipetted into 96‐well plates and allowed to fix at room temperature in the dark for 3 h. After this period, excess nonadherent gelatin was removed, and the wells were washed twice with PBS. A 1% (w/v) bovine serum albumin‐blocking solution (BSA, Sigma‐Aldrich) in PBS was then added (100 μL per well) and incubated for 1 h. Subsequently, the pre‐formed MCTS were transferred to the freshly prepared plates and treated with cirsimarin (1−160 µM). Photomicrographs were taken at 0, 24, and 48 h post‐treatment using an Olympus CKX41 inverted microscope with a ×4 objective and Qimaging Pro 7.1 software. Migration analysis was performed by measuring the area occupied by cells that migrated from the spheroid using AxioVision 3.1 software (Zeiss, Germany) with the “Measure” tool. The results were presented in square micrometers (μm²). Ho and PI dyes (30 μg/mL Ho and 40 µg/mL PI) were added after 48 h of treatment and incubated for 1.5 h in the dark to confirm cell death in the migration experiments. Finally, the A549 3D spheroids and cell migration areas were photographed and analyzed by the EVOS FL fluorescence microscope (Thermo Fisher Scientific, Waltham, USA) in a ×10 objective for representative images.
Bioinformatics Analysis
2.5
All data acquisition for the MMP‐11 mRNA expression investigation and clinical data correlation in lung adenocarcinoma (LUAD) samples were extracted from the UALCAN public tumor database (http://ualcan.path.uab.edu) [42] and The Cancer Genome Atlas (TCGA). We used the mRNA expression profile in TPM (transcripts per million) of LUAD from the UALCAN and TCGA databases for differential expression analysis, including normal and LUAD samples with different parameters. All data obtained from UALCAN were further processed and visualized using GraphPad Prism software. The survival data from patients were extracted from Kaplan–Meier Plotter [43] and were used to evaluate the effect of MMP‐11 gene expression with a meta‐analysis of published microarray data sets (Affy ID: 203876_s_at and 203878_s_at) on overall survival (OS) and first progression (PF; time when the cancer first gets worse or progresses) in patients with LUAD [44, 45]. The patients were divided into high‐expression and low‐expression groups based on the median expression value.
Statistical Analysis
2.6
All assays described were performed in three biological triplicates (n = 3) and at least three technical replicates. The statistical analyses were performed using GraphPad Prism 8.4.3 and 10.0 software (La Jolla, USA). Initially, the results obtained from biological assays were subjected to normality analysis of data distribution using the Shapiro−Wilk test. For samples with a parametric distribution, data were analyzed using the ANOVA test followed by the Dunnett, Tukey, or Sidak post‐test, and *p *≤ 0.05 was considered significant. The differences in gene expression (RT‐qPCR) were evaluated using the Student's t‐test. Treatments were considered to have altered gene expression when the fold change (FC) of the target gene differed from the solvent group (Student's t‐test, p ≤ 0.05). To explore the correlation between high levels of MMP‐11 expression and the survival rates of LUAD patients, the UALCAN uses the Log‐rank test, with a p‐value threshold ≤ 0.05. Similarly, the UALCAN portal uses the Student's t‐test with a p ≤ 0.01 to compare gene expression levels between different sample groups.
Results
3
2D Culture Assays
3.1
Cell Viability
3.1.1
Figure 2 presents the results of the resazurin and APH assays, illustrating the effects of cirsimarin treatment on cell viability. In the resazurin assay (Figure 2a), at all treatment times, cirsimarin at 80 and 160 μM concentrations significantly reduced cell viability compared to the SV. It was not possible to calculate the CC_50_ or the selectivity index in the resazurin assay after treating MRC‐5 cells for 24 h because none of the concentrations evaluated were cytotoxic. This result indicates that CIR exhibits preferential cytotoxicity toward A549 cells (Supporting Information S1: Figure 2).
*Cytotoxic effects of cirsimarin on A549 cells were assessed using resazurin (a) and acid phosphatase (b) assays. (a1) and (b1) show cells treated for 24 h; (a2) and (b2) show cells treated for 48 h; and (a3) and (b3) show cells treated for 72 h. A549 cells were exposed to various concentrations of cirsimarin (1−160 µM), along with positive controls (PC, docetaxel 50 µM) and solvent controls (SV, dimethyl sulfoxide 0.5%). Data are presented as mean ± standard deviation (n = 3). Statistical analysis was performed using ANOVA followed by Dunnett′s test. Symbols indicate statistically significant differences from the SV: *(p < 0.05), **(p < 0.01), ***(p < 0.001), and ***(p < 0.0001).
In the APH assay (Figure 2b), the results obtained at 24 h (Figure 2b1) showed a significant reduction in cell viability at cirsimarin concentrations starting from 80 μM, indicating a substantial decrease in viable cells. Furthermore, after 48 h (Figure 2b2), no significant changes in cell viability were detected across the tested concentrations. However, cirsimarin treatment significantly reduced cell viability at all evaluated concentrations after 72 h (Figure 2b3).
Cell Death and Cell Survival Assays
3.1.2
Figure 3a presents the results of the LDH release assay. At both 48 and 72 h treatment times, cirsimarin did not induce cell membrane disruption or the release of the LDH enzyme into the cytoplasm. This finding suggests that cirsimarin treatment does not compromise cell membrane integrity. Notably, significant results were only observed in the PC group provided by the kit.
*Assessment of cell death in A549 cells using lactate dehydrogenase (LDH) assay (a) and triple staining assay (b). Cells were treated with various concentrations of cirsimarin (1−160 µM), positive control (PC, docetaxel 50 µM), and solvent control (SV, dimethyl sulfoxide 0.5%) for 24 h (b), 48 h (a1), and 72 h (a2). (c) Shows representative images of viable, apoptotic, and necrotic cells stained with propidium iodide, Hoechst 33342, and fluorescein diacetate, analyzed using a fluorescence microscope at ×400 magnification. Data are presented as mean ± standard deviation (n = 3). Statistical analysis was performed using ANOVA followed by Dunnett's test, with significant differences from the SV indicated by *(p < 0.05) and ***(p < 0.0001).
The triple‐fluorescent staining assay, depicted in Figure 3c, differentiates viable, early apoptotic, late apoptotic, and necrotic cells using three fluorochromes: Ho, PI, and FDA. The results of this assay, shown in Figure 3b, indicate that cirsimarin induced apoptosis after 24 h of treatment at a concentration starting from 40 μM. Consequently, the percentage of viable cells decreased at the same concentration. The mean and standard deviation values are provided in Supporting Information S1: Table 2.
Finally, Figure 4 presents the clonogenic assay results, in which cells were treated with various concentrations of cirsimarin (1−160 μM). The clonogenic capacity of the cells was reduced by half at a concentration of 160 μM. The PC, treated with 50 μM DTX, absent in the graph, did not permit colony formation (as shown in the representative image).
*Clonogenic cell survival assay. (a) Surviving fraction after treatments with cirsimarin (1−160 μM) and solvent control (SV, dimethyl sulfoxide 0.5%). Data presented as mean ± standard deviation (n = 3). ANOVA followed by Dunnett's test. Statistically different from SV. *(p < 0.01). (b) Representative images of the clonogenic assay in A549 cells treated with the indicated treatments for 24 h. Cell colonies were stained with Giemsa, and the number of colonies was quantified after 10 days. Positive control (docetaxel 50 µM) formed no colony. Images of two additional biological replicates of each concentration are presented in Supporting Information S1: Figure 3.
Wound Healing Assay
3.1.3
Figure 5 presents the wound healing assay results, which assess the horizontal migration of A549 cells. A549 cells treated with the SV nearly closed the wound, achieving almost 100% closure within 24 h. In contrast, cells treated with the PC (cytochalasin) and various concentrations of cirsimarin (1−40 μM) demonstrated significantly impaired wound closure. Even after 72 h of treatment, the wound remained open in these groups, indicating that cirsimarin effectively inhibits cell migration at these concentrations. This suggests a strong potential for cirsimarin to impede cancer cell migration, an essential factor in metastasis.
*Wound healing assay. (a) Bar graph illustrating the percentage of wound closure at the indicated time points during the wound healing assay. A549 cells were treated for 72 h with various concentrations of cirsimarin (1−40 μM), positive control (PC, cytochalasin 2 mg/mL), and solvent control (SV, dimethyl sulfoxide 0.5%). Data are presented as mean ± standard deviation (n = 3). Statistical analysis was performed using ANOVA followed by Dunnett's test, with significant differences from SV indicated by ***(p < 0.0001). Wound closure was evaluated by measuring the remaining cell‐free area and expressed as a percentage of the initial cell‐free area (100%). (b) Representative images from the in vitro wound healing assays. Cells were photographed every 24 h, and the cell‐free area was measured. The white line delimits the migration area. Scale bar: 400 μm.
Comet Assay
3.1.4
Figure 6 presents the comet assay results, which measure DNA damage in A549 cells. The SV group had a damage index of 33.34 A.U. In contrast, the PC (ultraviolet C light) exhibited a significantly higher damage index of 73.41 A.U., demonstrating a clear statistical difference between these controls. For the cells treated with various concentrations of cirsimarin (1−80 μM), the damage indices were as follows: 77.33, 68.64, 73.54, 82.54, 83.25, and 74.00 A.U., respectively. All cirsimarin treatments showed statistically significant increases in DNA damage compared to the SV group. Notably, the 160 μM concentration was not evaluated in this assay as a substantial reduction in cell viability and induction of cell death had previously been observed, indicating potential cytotoxicity at this higher concentration. These findings suggest that cirsimarin induces considerable DNA damage, which is dose‐dependent, highlighting its potential genotoxic effects.
*DNA damage measured by alkaline comet assay. (a) Damage index in arbitrary units (A.U) in A549 cells treated with solvent (SV, dimethyl sulfoxide 0.5%), positive control (PC, 10 s in UVC), and cirsimarin (1−80 μM) for 4 h. Data presented as mean ± standard deviation (n = 3). ANOVA followed by Dunnett's test. Statistically different from the SV. *(p < 0.05), **(p < 0.01), and ***(p < 0.0001). (b) Representative images of different comet classes obtained by visual classification, as suggested by Moller et al. [37]. The comets represent classes 0–4 as used for visual scoring. The white bar delimits the size of the tail.
Modulation of Gene Expression
3.1.5
Figure 7 presents the results of the gene expression analysis. Following 12 h of treatment with 80 μM cirsimarin, all investigated genes (TP53, TNF‐α, CASP9, MMP‐2, MMP‐9, and MMP‐11) showed negative modulation. Additionally, only the MMP‐11 gene exhibited negative modulation with a lower concentration of cirsimarin (40 μM).
Gene expression analysis of marker genes in A549 cells. (a–f) Show the fold change in gene expression for (a) TP53, (b) TNF‐α, (c) CASP9, (d) MMP‐2, (e) MMP‐9, and (f) MMP‐11. A549 cells were treated with solvent control (SV, 0.5% dimethyl sulfoxide) or cirsimarin at 40 and 80 µM concentrations for 12 h. Data presented as mean ± standard deviation of the fold change in gene expression. Statistical analysis was performed using the Student's t‐test. () Statistically significant values of gene expression compared to the SV: *(p < 0.05) and ***(p < 0.0001).
3D Culture Assays
3.2
Growth, Morphology, and Viability of MCTS
3.2.1
Figure 8a1 presents the analysis results for the spheroids' area, morphology, and integrity. After 72 of treatment, concentrations ≥ 40 µM showed a noticeable decrease in the tumor spheroid area. Figure 8a2 presents photomicrographs of representative spheroids exposed to various cirsimarin and solvent control concentrations, demonstrating a general reduction in the 3D spheroid area with increasing concentration and treatment duration.
Figure 8b displays the results of the resazurin reduction assay performed after 216 h of treatment. The assay revealed that cirsimarin did not significantly affect cell viability at lower concentrations (1−20 µM). However, at concentrations of 40 µM and above, cirsimarin significantly reduced cell viability, with a viability value of 61.80% observed at 160 µM.
*Assessment of growth and integrity of A549 tumor spheroids. (a1) Graph depicting the area of spheroids (initial sizes ranging from 590 to 650 μm) after treatment with solvent control (SV, 0.25% dimethyl sulfoxide), positive control (PC, 92.835 µM docetaxel), and cirsimarin (1−160 μM). (a2) Time‐lapse series of representative brightfield images of spheroids, with medium and treatments refreshed every 3 days (0−216 h). (b) The means and standard errors of the mean for at least three independent repeat experiments (n = 3), indicating cell viability as assessed by the resazurin reduction assay. *Values denote statistical significance compared to SV: *(p < 0.05), **(p < 0.01), ***(p < 0.001), and ***(p < 0.0001).
3D Migration in the ECM
3.2.2
The results obtained in the ECM 3D migration assay are depicted in Figure 9. After 48 h under the concentration of 160 μM, the cell migration area was reduced (Figure 9a). In contrast, no reduction in 3D spheroid migration was observed at the other cirsimarin concentrations evaluated. However, there was a disparity in the compaction of A549 tumor spheroids exposed to the SV compared to spheroids treated at concentrations ≥ 40 μM after 48 h (Figure 9b). Considering that our research group previously observed a strong anti‐migratory effect of cirsimarin in MCF‐7 cells cultured in a 3D model, we compared the migration patterns of the two cell lines (Figure 9c). As seen in the images, on the same scale and starting from 3D spheroids with the same initial size, A549 cells exhibited a migration area similar to that of MCF‐7 cells 48 h earlier, highlighting the more aggressive migration potential of the lung lineage. Furthermore, in A549 spheroids, the visible necrotic center is smaller; however, there is a greater quantity of dead cells in the intermediate region of the 3D tumor spheroid.
*3D Cell migratory profile of A549 spheroids. (a) 3D cell migration from tumor spheroids in gelatin from bovine skin at 24 and 48 h after treatment with solvent control (SV, 0.25% dimethyl sulfoxide) and cirsimarin (1−160 μM). The relative migration area was quantified using Zen 2.3 software (Zeiss, Germany). * Statistically significant differences from the SV are indicated: ***p < 0.0001. (b) Representative images of A549 tumor spheroids after 48 h of treatment with cirsimarin. Scale bar: 400 μm; The white line delimits the migration area. (c) Fluorescence dye tracing of A549 spheroids after 48 h of treatment and MCF‐7 spheroids after 96 h of treatment to compare 3D cell migration. Propidium iodide (40 μg/mL) marks necrotic cells in red, and Hoechst (30 μg/mL) stains living cells in blue. Scale bar: 400 μm.
Prognostic Role of MMP‐11 Gene on LUAD
3.2.3
We used bioinformatics tools to examine MMP‐11 gene expression in LUAD patients and normal controls, considering that cirsimarin significantly downregulated MMP‐11 gene expression. First, we analyzed the MMP‐11 expression in LUAD using TCGA samples in the UALCAN web server and found significant increases in expression in various clinical contexts compared to normal samples (Figure 10). Specifically, MMP‐11 expression was significantly higher in primary tumor samples across all cancer stages (Stage 1−4) and in all smoking habit categories. Additionally, elevated MMP‐11 expression was observed in several histological subtypes, including NOS, Mixed, Clear Cell, LCB‐N, and SSP (p ≤ 0.0001 for each), as well as in both TP53‐mutant and TP53‐non‐mutant groups. Furthermore, samples with nodal metastasis showed significantly higher MMP‐11 expression.
*MMP‐11 expression in various clinical contexts in lung adenocarcinoma (LUAD) TCGA samples. (a) Comparison of MMP‐11 expression between normal (n = 59) and primary tumor samples (n = 515). (b) MMP‐11 expression across different cancer stages. (c) MMP‐11 expression based on smoking habits. (d) MMP‐11 expression across histological subtypes. (e) MMP‐11 expression based on TP53 status. (f) MMP‐11 expression based on nodal metastasis. The statistical analysis was performed using the Student's t‐test, and asterisks indicate statistical significance compared to the normal group (*p ≤ 0.01, ***p ≤ 0.0001).
Next, we used the Kaplan−Meier plotter platform to determine patient prognoses, such as OS and progression‐free (PF), with the MMP‐11 gene expression (Figure 11). The results demonstrated that LUAD patients with higher MMP‐11 gene expression (red line; n = 580) had an OS survival mean equal to 71 months, while the low expression group (black line; n = 581) had a mean of 110 months (*p *= 0.0015; Figure 11a). Furthermore, in FP survival (Figure 11b), the group with higher MMP‐11 expression (red line; n = 453) showed a mean survival of 15 months, while the low expression group (black line; n = 453) had a mean of 25 months. This difference was highly significant (*p *= 0.0000023; Figure 11b). These results suggest that high MMP‐11 expression is associated with poor outcomes in LUAD patients. The hazard ratio (HR) for OS was 1.32, indicating a 32% higher risk of death in the high‐expression cohort compared to the low‐expression group. Similarly, the HR for first progression (FPS) was 1.7, suggesting a 70% increased risk of disease progression or relapse in the high‐expression MMP‐11 group.
Survival analysis of MMP‐11 gene expression in lung adenocarcinoma (LUAD) patients using the Kaplan–Meier Plotter database. (a) Overall survival (OS; n = 581) and (b) first progression survival (FPS; n = 453) are shown for patients with high (red) and low (black) MMP‐11 mRNA expression. The analysis was based on clinical survival data in months. Statistical differences between the high‐ and low‐expression cohorts were calculated using Cox regression, with a p ≤ 0.05. The figure illustrates the association between elevated MMP‐11 expression and patient survival outcomes. HR = hazard ratio.
The bioinformatic data (Figures 10 and 11) demonstrated a significant association between elevated MMP‐11 expression and various clinical parameters in LUAD. These parameters include primary tumors, advanced cancer stages, smoking habits, specific histological subtypes, TP53 status, and nodal metastasis, indicating the role of MMP‐11 in disease progression and metastasis. Complementarily, our RT‐qPCR results (Figure 7) showed that cirsimarin treatment significantly downregulated MMP‐11 expression in A549 cells at 40 and 80 μM after 12 h. This downregulation correlates with decreased tumor cell migration in LUAD (A549) cells cultured in 2D monolayer and 3D tumor spheroid models.
Discussion
4
NSCLC is a significant public health challenge, as lung cancer remains the leading cause of cancer‐related mortality worldwide for both men and women [46]. Despite therapeutic advances, current approaches such as surgery, radiotherapy, chemotherapy, and targeted therapies remain limited by drug resistance and adverse effects. Chemotherapeutics such as vinorelbine (a semisynthetic vinblastine derivative), DTX (a taxane from T. brevifolia), and etoposide (from Podophyllum peltatum) are commonly used in NSCLC, either alone or in combination [46]. However, searching for novel, more effective compounds with fewer toxic side effects remains critical. In this context, our study provides valuable insights into the anticancer potential of cirsimarin, particularly highlighting its antimetastatic properties.
In the first parameter analyzed, resazurin assay, cell toxicity, cirsimarin ≥ 80 μM reduced cell viability in the 2D model, consistent with findings by Serpeloni et al. [19] in breast tumor cells (MCF‐7). In addition, we also observed a selective effect of cirsimarin for the A549 cells when compared to the non‐tumor cell line MRC‐5. In the APH assay, cirsimarin (1–160 μM) further reduced cell viability after treatment for 72 h, suggesting that this assay is more sensitive to detecting cytotoxicity. LDH assay showed that cirsimarin (1–160 μM) did not compromise A549 cell membrane integrity, suggesting that reduced viability at ≥ 80 μM is likely due to metabolic disruption rather than membrane damage. This result was confirmed using a triple‐staining assay, where cirsimarin at concentrations ≥ 40 μM triggered cell death by apoptosis, evidenced by the presence of DAF‐labeled apoptotic bodies. Apoptosis, a form of programmed cell death, maintains membrane integrity while forming apoptotic bodies, preventing the release of intracellular contents during early stages [47].
Chemotherapeutic drugs currently used in the clinic exert cytotoxicity through inducing DNA damage, mutations, and genome instability [48]. Cirsimarin (1–80 μM) increased the DNA damage after 4 h treatment. Considering the results of the cell death assays (after 24 h) and the concentrations analyzed in the comet assay, it is likely that apoptosis is triggered by DNA damage. Serpeloni et al. [19] observed that cirsimarin induces DNA damage in MCF‐7 cells in 3D after 24 h but not after 4 h, reflecting the delayed penetration of the compound due to the architecture of the 3D tumor spheroids. Similarly, other flavonoid glycosides, such as quercetin‐3‐glucoside, promote cell cycle arrest, increase DNA fragmentation, and activate apoptosis pathways through mitochondrial dysfunction [49].
Cirsimarin (80 μM) downregulated most of the cell death genes evaluated, corroborating previous findings in MCF‐7 breast tumor cells [19] and raising intriguing new questions. The p53 is a critical tumor suppressor protein involved in DNA repair, apoptosis, and cell cycle checkpoints to safeguard genomic integrity [50, 51, 52]. In our study, cirsimarin reduced both TP53 and CASP9 expressions in wild‐type A549 cells, even in the presence of DNA damage, confirmed by the comet assay. Considering that DNA damage activates the intrinsic apoptotic pathway through caspase‐9 (CASP9), our data suggest a CASP9‐independent pathway, such as mitochondrial dysfunction or ER stress‐related apoptosis. Similar behavior has been reported for the flavonoid glycoside Kuwanon C, which decreased the expression of CASP3 and CASP9 in HeLa cells while still inducing apoptosis and DNA damage through mitochondrial disruption [53]. This suggests flavonoid glycosides such as cirsimarin may bypass conventional apoptotic pathways to induce tumor cell death.
Considering that several DNA repair genes are direct targets of p53 or are indirectly regulated by it [54], our findings are particularly intriguing given that TP53 is typically upregulated in LUAD [55]. Downregulating TP53 in A549 cells could offer therapeutic advantages by disrupting repair pathways and promoting apoptosis. The suppression of TP53 expression by cirsimarin may impair the tumor's ability to escape programmed cell death, particularly in p53‐regulated cancers such as LUAD. In the clonogenic assay, only cirsimarin at 160 μM significantly reduced clonogenic capacity, supporting the hypothesis that lower concentrations of cirsimarin might not halt the cell cycle sufficiently to allow DNA repair processes.
Tumor cells with migratory ability tend to exhibit a more aggressive phenotype due to their metastasizing capacity. Inhibiting cell migration is crucial to preventing recurrence and improving patient survival, as metastasis is the leading cause of cancer‐related deaths [56]. Our results demonstrated that cirsimarin (1–40 μM) effectively inhibits migration, comparable to the PC cytochalasin. This anti‐migratory effect is likely related to the transcriptional downregulation of metalloproteinases 2, 9, and 11, along with TNF‐α. MMPs are essential proteolytic enzymes responsible for degrading the ECM by breaking down structural proteins such as collagen, laminin, elastin, and fibronectin [57, 58, 59]. The degradation of the ECM facilitates cell migration and invasion, which are critical for tumor progression, as tissue restructuring is necessary for tumor cells to invade other tissues [58, 60].
TNF‐α plays a central role in cancer biology by promoting proliferation, migration, invasion, inflammation, apoptosis, and necrosis. It can also modulate the expression of MMPs, which further contributes to tumor cell invasiveness [61, 62]. In A549 cells, Lee et al. [63] demonstrated that TNF‐α induces MMP‐9 expression through the activation of the TNFR1/TRAF2/PKCα‐dependent JNK1/2/c‐Jun and c‐Src/EGFR/PI3K/Akt pathways. A similar anti‐migration effect was observed with the flavone apigenin, which inhibited the migration of lung cancer cells by reducing MMP‐9 levels and blocking PI3K/Akt pathway signaling [64]. These findings highlight that flavone glycosides like cirsimarin and apigenin derivatives can suppress metastasis‐associated mechanisms, offering promising therapeutic potential.
MMPs contribute to drug resistance in lung tumors and are closely linked to reduced tumor immunity, weakening the innate immune response [20]. TCGA data analysis revealed significantly higher MMP‐11 expression in tumor tissues compared to normal tissues, with even greater expression in metastatic tumors, especially in individuals with more than 10 metastases. Therefore, the downregulation of MMP‐2, MMP‐9, and especially MMP‐11 by cirsimarin may benefit patients with aggressive tumors. Similar modulation of MMP‐2 and MMP‐9 has been observed with other flavonoids, such as luteolin [65] and the isoflavone Biochanin A [66]. These findings and the TCGA data suggest that cirsimarin's modulation of MMP‐11 contributes to its anti‐metastatic effect, underscoring its potential as a chemotherapeutic agent for inhibiting LUAD progression and metastasis.
This study also evaluated the ability of A549 cells to form tumor spheroids and explored the biological effects of cirsimarin in a 3D model. The formation of 3D spheroids relies on interactions between neighboring cells and the ECM, which promotes the establishment of the 3D structure [67]. The ECM plays a critical role by inducing integrin expression, facilitating cell adhesion, and spheroids' initial aggregation. Simultaneously, E‐cadherins mediate firm compaction and help define the structural zones within the 3D tumor spheroid [68].
After A549 spheroid formation, we observed a reduction in the spheroids in the SV group, potentially indicating greater compaction of the structure. Yun et al. [69] demonstrated that A549 spheroids overexpress adhesion genes, including claudin‐2. The downregulation of claudin‐2 enhances paracellular permeability and promotes doxorubicin accumulation in A549 spheroids [70, 71]. Similar reductions in the spheroid area were also observed in studies by Baek et al. [72] and Garnique and Machado‐Santelli [73]. Additionally, A549 spheroids express high levels of claudin‐1, contributing to increased chemoresistance within the 3D structure [74]. These features highlight the utility of the 3D lung cancer model, which closely mimics in vivo tumor conditions. Cirsimarin decreased cell proliferation in spheroids, a promising result considering their high chemoresistance. Comparing the 3D structure of breast (MCF‐7) and lung (A549) cells, Ruiz et al. [75] showed that MCF‐7 cell spheroids had a CC_50_ of 129.5 μM for cDDP, while the CC_50_ for A549 cells was > 500 μM. Maruhashi et al. [76] demonstrated that claudin‐2 positively regulates the expression of the multidrug resistance‐associated protein ABCC2 by reducing the sensitivity of A549 spheroids.
Shabalina et al. [77] demonstrated that A549 spheroids exhibit a distinct migration pattern when assessed on plastic and collagen as extracellular matrices. These authors identified three key regions: S0 as the spheroid's center, S1 as the intermediate region, and S2 as the outer ring representing the area of active migration. Our results confirmed this pattern, as shown in Figure 9c, for both MCF‐7 and A549 cell lines. In the migration assay, the S0 region of MCF‐7 spheroids corresponded to a necrotic core, while in A549 MCTS, necrotic cells also extended into the S1 region. At higher concentrations (≥ 40 μM), cirsimarin treatment led to the detachment of cells from the necrotic center, weakening the spheroid's structure, as shown in Figure 9b.
A key strength of this study is confirming that A549 spheroids are a reliable in vitro model for toxicological assays, addressing the limited information available on them. Moreover, phytochemicals such as flavonoids offer significant advantages due to their natural, plant‐derived origin and their low production and acquisition costs, which enhance both their accessibility and therapeutic potential [78]. However, the present study also has some limitations that may guide future studies. Zhang et al. [79] showed that cirsimarin has a short half‐life in vivo. In fact, according to Kin et al. [80], increased bioavailability and targeted delivery strategies may enhance the translational relevance of natural compounds such as cirsimarin. In addition, experiments that evaluate protein expression could be a better mechanistic validation for cirsimarin's antimigratory effects.
Conclusion
5
Taken together, the 2D results suggest that cirsimarin plays a significant role in lung tumor progression by modulating cell death and migration/metastasis. These findings are reinforced by the 3D model, which offers more profound insights into the compound's anticancer potential. This work provides a foundation for future research into other cellular pathways involved in the anticancer potential/activity of cirsimarin.
Author Contributions
Anna Gabriele Prado dos Santos and Juliana Mara Serpeloni: conceptualization. Anna Gabriele Prado dos Santos, Celina Yung‐Ai Lin Lee, Érica Romão Pereira, Andresa Hiromi Sakai, Victor Antônio Silva Lima, and Marcos Bispo Pinheiro Camara: methodology. Anna Gabriele Prado dos Santos, Sabine A. S. Langie, Ilce Mara de Syllos Cólus, and Juliana Mara Serpeloni: formal analysis. Juliana Mara Serpeloni, Cláudia Quintino da Rocha, and Ilce Mara de Syllos Cólus: resources. Juliana Mara Serpeloni and Diego Luís Ribeiro: data curation. Anna Gabriele Prado dos Santos, Juliana Mara Serpeloni, Sabine A. S. Langie, Celina Yung‐Ai Lin Lee, and Andresa Hiromi Sakai: writing – original draft preparation. Juliana Mara Serpeloni, Cláudia Quintino da Rocha, Ilce Mara de Syllos Cólus, and Diego Luís Ribeiro: writing – review and editing. Juliana Mara Serpeloni, Cláudia Quintino da Rocha, Ilce Mara de Syllos Cólus: supervision. Anna Gabriele Prado dos Santos, Juliana Mara Serpeloni: project administration. Juliana Mara Serpeloni and Cláudia Quintino da Rocha: funding acquisition. All authors have read and agreed to the published version of the manuscript.
Ethics Statement
The authors have nothing to report.
Consent
The authors have nothing to report.
Conflicts of Interest
The authors declare no conflicts of interest.
Supporting information
Supplementary Figure 1: Authenticity of the cell line assessed by the Paternity Investigation Program by DNA Analysis ‐ (PIPAD). Supplementary Figure 2: Cytotoxic effects of Cirsimarin on A549 lung tumor cells and MRC‐5 non‐tumor cells (lung fibroblasts) after 24 h of treatment evaluated by resazurin reduction assay. Supplementary Figure 3: Images of two additional biological replicates of each concentration for the clonogenic assay. Supplementary Table 1: Sequence of primers TNF‐α, CASP‐9, TP53, MMP‐2, MMP‐9, MMP‐11, GAPDH and HPRT1. Supplementary Table 2: Mean and standard deviation values of the triple fluorescent staining assay.
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