Preparation of the 70% Ethanol Hydroalcoholic Extract of Onopordum leptolepis

Plant specimens of Onopordum leptolepis were collected from the mountainous regions of Kermanshah, Iran (GPS coordinates: 34° 18' 51.01" N, 47° 03' 54.00" E) during the optimal flowering season. Botanical authentication was confirmed by an expert plant taxonomist, and a voucher specimen was deposited in the institutional herbarium (Voucher No. KU-OV-2020-001). The aerial parts (leaves and flowering stems) were cleaned and shade-dried at ambient room temperature to prevent the thermal degradation of thermolabile phytoconstituents. The dried material (40 g) was pulverized into a fine powder using an electric grinder to optimize surface area contact and enhance solvent penetration during extraction. Hydroalcoholic extraction was performed using the maceration technique with 70% ethanol (v/v), a solvent system established for efficiently extracting a broad spectrum of polar and semi-polar bioactive phytoconstituents. The pulverized plant material was immersed in 300 mL of 70% ethanol and agitated on an orbital shaker at moderate speed for 72 h at room temperature. The mixture was filtered through Whatman No. 1 filter paper to remove insoluble particulate residues. The filtrate was concentrated under a chemical fume hood with gentle airflow at room temperature and subsequently dried in a temperature-controlled oven at 45 °C for 24 h to ensure complete removal of residual ethanol. The final dried crude extract was stored in airtight, light-protected containers at −20 °C until experimental use. Stock solutions were prepared by dissolving the dried extract in dimethyl sulfoxide (DMSO) and subsequently diluted with culture medium to achieve designated working concentrations. The final DMSO concentration in all treatment wells was maintained at ≤ 0.1% (v/v), a level confirmed to exert negligible cytotoxicity. Vehicle control wells received an equivalent concentration of DMSO (0.1%) without plant extract12.

Cell Culture Conditions

Human colorectal carcinoma cells (SW480) and non-tumorigenic mouse subcutaneous fibroblasts (NCTC clone 929 / L929) were obtained from established cell repositories and cultured under standard sterile conditions in 25-cm² tissue culture flasks. SW480 cells were maintained in RPMI-1640 medium supplemented with 10% heat-inactivated fetal bovine serum (FBS) and 1% penicillin–streptomycin (100 U/mL penicillin, 100 µg/mL streptomycin). Experiments were conducted using cells between passages 4 and 10. L929 fibroblasts were cultured in Dulbecco’s Modified Eagle Medium (DMEM) enriched with 10% FBS and 1% penicillin–streptomycin. Both cell lines were maintained at 37 °C in a humidified atmosphere containing 5% CO, with growth media replenished every 48–72 h. Cell morphology and confluence were routinely monitored using an inverted phase-contrast microscope, and cells were subcultured using 0.25% trypsin–EDTA upon reaching 80–90% confluence13.

Cell Viability Assessment (MTT Assay)

Cell viability was evaluated using the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) colorimetric assay. Briefly, SW480 cells and L929 fibroblasts were seeded into 96-well plates at a density of 1.5 × 10 cells/well and incubated overnight to facilitate attachment and equilibration. Cells were subsequently treated with increasing concentrations of the 70% ethanol hydroalcoholic extract (12.5, 25, 50, 100, 200, 400, and 800 µg/mL) or vehicle control (0.1% DMSO) for 24 h. The concentration range was selected to encompass non-cytotoxic, moderately cytotoxic, and high-dose cytotoxic thresholds for comprehensive dose-response modeling. Following 24 h of incubation, 30 µL of MTT solution (5 mg/mL in PBS) was added to each well, and plates were incubated for an additional 3 h at 37 °C in the dark to facilitate the reduction of MTT into insoluble formazan crystals by active mitochondrial dehydrogenases. The supernatant was carefully aspirated, and 100 µL of DMSO was added to each well to solubilize the formazan precipitates. Plates were gently agitated on a microplate shaker for 20 min at room temperature to ensure complete dissolution, and absorbance was measured at 570 nm using an ELISA microplate reader. Percentage cell viability was calculated as follows: Cell Viability (%) = (OD / OD) × 100. The extract remained visually soluble throughout the 24-h incubation period without observable precipitation14.

Determination of IC Values

Cell viability data obtained after 24 h of extract exposure were fitted to a four-parameter logistic (4PL) nonlinear regression model using GraphPad Prism (version 8.0) and SPSS (version 16.0). Concentration–response curves were generated by plotting cell viability (%) against the log of extract concentration (µg/mL). The half-maximal inhibitory concentration (IC) values, corresponding 95% confidence intervals (95% CI), and goodness-of-fit coefficients (R²) were derived from the fitted regression curves.

Apoptosis Quantification (Diphenylamine-Based DNA Fragmentation Assay)

Apoptotic cell death was evaluated by quantifying low-molecular-weight DNA fragmentation using the diphenylamine (DPA) colorimetric method. Following 24 h of treatment with designated extract concentrations, approximately 5 × 10 SW480 cells were harvested, washed with cold PBS, and lysed in 1 mL of TTE lysis buffer (10 mM Tris-HCl, pH 7.4, 1 mM EDTA, 0.2% Triton X-100) to selectively release fragmented apoptotic DNA. Cell lysates were centrifuged at 20,000 × g for 10 min at 4 °C to separate fragmented DNA (supernatant, Sample A) from intact, high-molecular-weight chromatin (pellet, Sample B). The supernatant was collected into a fresh tube, while the pellet was resuspended in 1 mL of TTE buffer. Both fractions were combined with an equal volume of 25% trichloroacetic acid (TCA) and incubated overnight at 4 °C to precipitate DNA. Following centrifugation at 20,000 × g for 10 min, the resulting precipitates were hydrolyzed by adding 160 µL of 5% TCA and heating at 90 °C for 15 min. Subsequently, 320 µL of freshly prepared DPA reagent was added to each sample, followed by incubation at 37 °C for 4 h to allow color development. Optical density was measured at 600 nm using a spectrophotometer. The percentage of apoptotic DNA fragmentation was calculated using the formula: DNA Fragmentation (%) = [OD / (OD + OD)] × 10015.

Gene Expression Analysis by Quantitative Real-Time PCR (qRT-PCR)

Total cellular RNA was extracted from treated and untreated SW480 cells using TRIzol reagent according to the manufacturer’s instructions and stored at −80 °C. RNA concentration and purity were assessed using a NanoDrop spectrophotometer, with A/A absorbance ratios between 1.8 and 2.0 considered acceptable for downstream applications. Complementary DNA (cDNA) was synthesized from 1 µg of total RNA using the RevertAid First Strand cDNA Synthesis Kit (Thermo Fisher Scientific, USA; Cat. No. K1622). Quantitative real-time PCR was performed using PowerUp™ SYBR™ Green Master Mix (Applied Biosystems, USA; Cat. No. A25742) on a real-time PCR detection system. The amplification protocol consisted of an initial denaturation at 95 °C for 10 min, followed by 40 cycles of denaturation at 95 °C for 15 s, annealing at 60 °C for 30 s, and extension at 72 °C for 30 s. A terminal melting curve analysis (65–95 °C) was conducted to verify amplification specificity and confirm the absence of primer-dimer artifacts. The specific primer sequences were as follows:

• Notch116: Forward 5′-CGTTCCAGCAGTCTCCGTC-3′, Reverse 5′-GTGGGCCAGTCTCAAAGG-3′

• Jagged116: Forward 5′-AGTGCCTGAATGGACGGA-3′, Reverse 5′-TGGAGACTGGAAGACCGA-3′

• c-Myc17: Forward 5′-TCTCCATCCTATGTTGCGGTC-3′, Reverse 5′-TCCAAGTAACTCGGTCATCATCT-3′

• GAPDH18: Forward 5′-GTCTCCTCTGACTTCAACAGCG-3′, Reverse 5′-ACCACCCTGTTGCTGTAGCCAA-3′

GAPDH served as the internal housekeeping control. Relative mRNA expression levels were calculated using the comparative 2 method19.

Quantification of Notch1, Jagged1, and c-Myc Protein Levels by ELISA

Cellular protein levels of Notch1, Jagged1, and c-Myc were determined using commercially available, enzyme-linked immunosorbent assay (ELISA) kits according to the manufacturers’ protocols: human Notch1 ELISA kit (Invitrogen, Thermo Fisher Scientific; Cat. No. EHNOTCH1), human Jagged1 ELISA kit (R&D Systems; Cat. No. DY599), and human c-Myc ELISA kit (MyBioSource; Cat. No. MBS724950). Following 24 h of treatment, SW480 cells were gently detached using a non-enzymatic cell dissociation solution, washed twice with ice-cold PBS, and lysed in cell lysis buffer containing protease inhibitors. Protein extracts were added to pre-coated microplates and incubated with target-specific capture and biotinylated detection antibodies, followed by incubation with streptavidin-horseradish peroxidase conjugate. Chromogenic substrate solution was added, and absorbance was measured at 450 nm (with wavelength correction at 570 nm) using a microplate reader. Target protein concentrations were calculated from standard curves generated with serial dilutions of purified recombinant standards and normalized to total protein content. Untreated cells served as controls, and blank wells containing assay reagents without cell lysate were included to determine background absorbance20.

Scratch Wound-Healing Assay (Cell Migration)

Two-dimensional collective cell motility was evaluated using the scratch wound-healing assay. SW480 cells were seeded into 6-well plates at a density of 1.6 × 10 cells/mL and cultured until reaching a confluent monolayer. A uniform, linear cell-free scratch was generated across the center of each well using a sterile 1-mL pipette tip. Wells were washed gently with PBS to remove dislodged cells and debris. Cells were then incubated in DMEM supplemented with 2% FBS (to minimize confounding effects of cellular proliferation) in the presence or absence of O. leptolepis extract for 24 h at 37 °C. Wound closure was monitored by photographing identical coordinates immediately after scratch generation (0 h) and after 24 h using an inverted phase-contrast microscope equipped with a digital camera. Quantitative measurement of the denuded area was performed using automated threshold-based image segmentation in TScratch software (MathWorks Inc.). The percentage of wound closure was calculated as: Wound Closure (%) = [(Wound Area at 0 h − Wound Area at 24 h) / Wound Area at 0 h] × 10021.

Matrigel Transwell Invasion Assay

Cellular invasive capacity was evaluated using 24-well Transwell chambers containing 8-µm pore polycarbonate membrane inserts (6.5-mm diameter; Corning Inc.). The upper surface of each insert membrane was coated with 50 µL of growth-factor-reduced Matrigel (diluted in serum-free medium) and incubated at 37 °C for 30 min to allow gel polymerization. SW480 cells in the logarithmic growth phase were harvested and resuspended in serum-free medium at a density of 1.0 × 10 cells/mL. A 100-µL aliquot of cell suspension was placed into the upper chamber, while 600 µL of medium containing 2% FBS (serving as a chemoattractant), with or without O. leptolepis extract, was added to the lower chamber. Following 24 h of incubation at 37 °C in 5% CO, non-invaded cells and residual Matrigel on the upper membrane surface were carefully removed with cotton swabs. Cells that invaded through the membrane to the lower surface were fixed with 4% paraformaldehyde for 20 min at room temperature and stained with 0.5% crystal violet solution for 20 min. After washing thoroughly with distilled water and air-drying, representative microscopic images were captured. For quantitative analysis, the bound crystal violet was dissolved in 70% ethanol, and optical density was measured at 570 nm using a microplate reader. Absorbance values served as an indirect measure of the relative number of invaded cells22.

Statistical Analysis

All experiments were performed independently in triplicate (n = 3), with each assay conducted in technical triplicates. Quantitative data are presented as the mean ± standard error of the mean (SEM). The normality of data distribution was assessed using the Kolmogorov–Smirnov test, and all datasets satisfied normality assumptions (p > 0.05). Differences among multiple experimental groups were evaluated using one-way analysis of variance (ANOVA) followed by Tukey’s post hoc multiple-comparison test. Statistical analyses were performed using SPSS software (version 16.0; SPSS Inc., Chicago, IL, USA) and GraphPad Prism (version 8.0). A two-tailed p < 0.05 was considered statistically significant23.

Cytotoxic Dose–Response of Onopordum leptolepis Extract in SW480 Cells and L929 Fibroblasts

To characterize the cytotoxic activity and selectivity of the 70% ethanol hydroalcoholic extract of O. leptolepis, concentration–response profiles were established from 24-h MTT assay data using four-parameter logistic (4PL) nonlinear regression analysis. As depicted in Figure 1A, the extract produced a pronounced, concentration-dependent reduction in the viability of SW480 colorectal cancer cells. The fitted logistic model demonstrated strong concordance with experimental data (R² > 0.95), yielding an estimated IC of 290.11 µg/mL (95% CI: 259.8–324.0 µg/mL). Cell viability steadily declined with increasing extract concentrations, reaching approximately 32% at the highest tested concentration (800 µg/mL), confirming significant cytotoxic efficacy against SW480 cells. In contrast, non-tumorigenic L929 mouse fibroblasts exhibited marked resistance across the same concentration range (12.5–800 µg/mL; Figure 1B). L929 cell viability remained above 50% across all evaluated concentrations, retaining approximately 75% viability even at 800 µg/mL. Consequently, the IC for L929 fibroblasts could not be experimentally achieved within the tested range and is reported as IC > 800 µg/mL. Comparison of the two dose-response profiles indicates that O. leptolepis extract exhibits preferential cytotoxicity toward malignant SW480 cells relative to normal fibroblasts (Figure 1).

Comparative Effects of Onopordum leptolepis Extract on CRC Cell and Fibroblast Viability

Direct comparison of cell viability between SW480 CRC cells and L929 fibroblasts following 24 h of extract exposure across increasing concentrations (12.5–800 µg/mL) is presented in Figure 2. In SW480 cells, low extract concentrations (12.5–50 µg/mL) induced a modest but statistically significant decrease in viability (88–92% vs. 100% in control, p < 0.05). Substantial cytotoxicity occurred at intermediate concentrations of 100 and 200 µg/mL, where viability dropped significantly to approximately 70% and 55%, respectively (p < 0.001 vs. control). At 400 and 800 µg/mL, cell survival further declined to 48% and 32%, respectively (p < 0.001 vs. control), representing more than a two-thirds reduction in metabolic activity. Conversely, L929 fibroblasts maintained near-baseline viability at concentrations up to 200 µg/mL (p > 0.05 vs. control), with only a slight reduction at 400 µg/mL (approximately 88% viability, p < 0.01) and a moderate reduction at 800 µg/mL (around 75% viability, p < 0.001) (Figure 2).

Induction of Apoptotic DNA Fragmentation in SW480 Cells

To determine whether extract-induced cytotoxicity was associated with programmed cell death, apoptotic DNA fragmentation was quantified using the DPA colorimetric assay (Figure 3). Treatment of SW480 cells with O. leptolepis extract for 24 h led to a robust, dose-dependent increase in DNA fragmentation. Baseline fragmentation in untreated control cells was minimal (<5%). Exposure to low concentrations (12.5 and 25 µg/mL) resulted in a modest elevation of fragmentation to 8–12% (p < 0.05). A marked and statistically significant increase was observed at 50 and 100 µg/mL, where fragmentation rose to approximately 18% and 30%, respectively (p < 0.001). At 200 µg/mL, apoptosis reached nearly 42% (p < 0.001), representing more than an eight-fold increase relative to controls. Higher concentrations (400 and 800 µg/mL) elicited the most pronounced apoptotic responses, with fragmentation levels reaching approximately 48% and nearly 60%, respectively (both p < 0.001) (Figure 3).

Suppression of Notch Pathway Gene Expression in SW480 Cells

To examine the molecular mechanisms associated with the anticancer effects of O. leptolepis, relative mRNA expression levels of key Notch signaling components (Notch1, Jagged1, and c-Myc) were analyzed by qRT-PCR following 24 h of treatment (Figure 4). In untreated control cells, all three genes displayed stable baseline expression levels (normalized to 1.00 ± 0.10). Exposure to O. leptolepis extract resulted in significant transcriptional downregulation of Notch1 to 0.68 ± 0.12 (p < 0.001 vs. control) and Jagged1 to 0.63 ± 0.11 (p < 0.001 vs. control). Concurrently, expression of the downstream oncogenic effector c-Myc was significantly decreased to 0.52 ± 0.14 (p < 0.001 vs. control) (Figure 4).

Downregulation of Notch Pathway Protein Expression in SW480 Cells

Consistent with the observed transcriptional changes, quantitative ELISA demonstrated a significant reduction in cellular protein levels of Notch1, Jagged1, and c-Myc in SW480 cells following 24 h of extract treatment (Figure 5). Compared with untreated control cells (normalized to 1.0-fold), exposure to O. leptolepis extract significantly decreased Notch1 protein levels to approximately 0.80-fold, Jagged1 to 0.65-fold, and c-Myc to 0.60-fold (all p < 0.001 vs. control) (Figure 5).

Inhibition of SW480 Cell Migration by Onopordum leptolepis Extract

The effect of O. leptolepis extract on cell motility was evaluated using the scratch wound-healing assay (Figure 6). Untreated control SW480 cells exhibited vigorous collective migration, achieving 45–50% scratch area closure after 24 h. In contrast, treatment with O. leptolepis extract significantly impeded cell migration, reducing wound closure to 20–25% (p < 0.001 vs. control), demonstrating potent antimigratory activity in vitro (Figure 6).

Inhibition of SW480 Cell Invasion by Onopordum leptolepis Extract

The impact of O. leptolepis extract on cell invasiveness was assessed using Matrigel-coated Transwell invasion chambers (Figure 7). In untreated controls, robust cellular invasion through the reconstituted basement membrane was observed, corresponding to an optical density (OD) of 0.95–1.0. Exposure to O. leptolepis extract significantly decreased the absorbance of extracted crystal violet to approximately 0.80 (p < 0.001 vs. control), reflecting a substantial reduction in the number of invading cancer cells (Figure 7).