Inhibition of Colorectal Cancer Cell Progression by Picroside II Through Modulation of the Notch1 Signaling Pathway

ABSTRACT

This study aimed to investigate the biological effects of Picroside II on colorectal cancer (CRC) cells, including its impacts on proliferation, migration, invasion, and epithelial–mesenchymal transition (EMT), and to explore whether its mechanism of action involves modulation of the Notch1 signaling pathway. Human CRC cell lines SW480 and SW620 were treated with various concentrations of Picroside II (10–100 μM). Cell proliferation was assessed using the CCK‐8 assay, colony formation assay, and EdU incorporation assay. Migration and invasion capacities were evaluated by wound healing and Transwell assays. The expression levels of EMT‐related markers (E‐cadherin, N‐cadherin, Vimentin, ZEB2) and key proteins in the Notch1 pathway (Notch1, Cleaved Notch1, RBP, HES1) were analyzed by Western blotting. Statistical analysis was performed using one‐way ANOVA and Student's t‐test. Picroside II inhibited the proliferation of SW480 and SW620 cells in a dose‐ and time‐dependent manner, reduced colony formation ability, and decreased DNA synthesis activity. Treatment with Picroside II suppressed the migratory and invasive abilities of CRC cells, accompanied by upregulation of E‐cadherin and downregulation of N‐cadherin, Vimentin, and ZEB2. Furthermore, Picroside II exposure led to a decrease in the expression of Notch1, Cleaved Notch1, RBP, and HES1 proteins in a concentration‐dependent manner. Picroside II suppresses CRC cell progression in vitro by inhibiting the Notch1 signaling pathway, providing a preliminary molecular basis for further in vivo investigation.

Picroside II inhibits colorectal cancer progression by blocking Notch1 signaling activation. It interferes with Notch1 activation by preventing the cleavage and nuclear translocation of the Notch intracellular domain (NICD). Upon ligand binding, Notch1 is cleaved by ADAM and γ‐secretase, releasing NICD, which translocates into the nucleus and forms a transcriptional activation complex (including CSL, MAML, and SKIP). This complex promotes transcription of downstream targets such as Hes1, which regulate epithelial–mesenchymal transition as well as cell proliferation, migration, and invasion.

1. Introduction

Colorectal cancer (CRC) is a common malignancy worldwide, ranking as the third most frequently diagnosed cancer and the second leading cause of cancer‐related mortality, accounting for approximately 10% of all cases globally [1]. In China, the burden of CRC has increased steadily in recent years, with a rising incidence and mortality rate associated with factors such as an aging population, dietary transitions, and changes in lifestyle behaviors [2, 3]. According to the latest national cancer statistics, CRC remains among the top five malignancies in both urban and rural regions, leading to considerable clinical and public health challenges. Although advances in early detection, surgery, chemotherapy, and radiotherapy have contributed to improved outcomes for patients with localized disease, the prognosis for individuals with advanced or metastatic CRC remains unsatisfactory [4]. Standard therapeutic approaches, including surgical resection combined with systemic chemotherapy and radiotherapy, are limited by issues such as treatment‐related toxicities, including myelosuppression, gastrointestinal disturbances, and peripheral neuropathy, which may severely impact patient quality of life [5, 6, 7]. In addition, the development of resistance to conventional chemotherapeutic agents, such as oxaliplatin and 5‐fluorouracil, continues to pose a major barrier to sustained disease control [8]. Although targeted therapies directed against molecules such as epidermal growth factor receptor (EGFR) and vascular endothelial growth factor (VEGF) have been introduced into clinical practice, intrinsic and acquired resistance to these agents often limits their clinical benefits over time [9, 10, 11]. Given these limitations, the development of new therapeutic strategies characterized by lower toxicity profiles and alternative mechanisms of action remains an important area of investigation in the treatment of CRC.

Epithelial–mesenchymal transition (EMT) is a biological process by which epithelial cells acquire mesenchymal characteristics, resulting in enhanced migratory capacity, invasiveness, and resistance to apoptosis [12, 13, 14]. In the context of CRC, EMT is regarded as a fundamental mechanism driving tumor metastasis and progression [15]. During EMT, epithelial markers such as E‐cadherin are downregulated, while mesenchymal markers, including N‐cadherin and Vimentin, are upregulated, leading to the disruption of cell‐cell adhesion and the acquisition of a motile phenotype [16, 17]. This phenotypic plasticity enables cancer cells to detach from the primary tumor mass, invade surrounding tissues, and establish secondary lesions at distant sites. Multiple signaling pathways are involved in the regulation of EMT, including the transforming growth factor‐beta (TGF‐β), Wnt/β‐catenin, and Hedgehog pathways, each contributing to different aspects of EMT initiation and maintenance [18, 19, 20]. Among these, the Notch signaling pathway has emerged as a critical regulator of EMT in CRC. Aberrant activation of Notch1 has been observed in colorectal tumors and has been implicated in the promotion of EMT by repressing epithelial markers and enhancing mesenchymal traits [21, 22]. Studies have demonstrated that sustained Notch1 activity facilitates tumor cell dissemination and correlates with poorer clinical outcomes in CRC patients [23]. Based on these findings, the inhibition of Notch1 signaling represents a potential therapeutic strategy aimed at limiting EMT‐mediated metastasis in CRC.

Natural products have long been recognized as an important source for the development of novel therapeutic agents, particularly in the field of oncology. Compounds derived from traditional medicinal plants have demonstrated diverse pharmacological activities, including anti‐inflammatory, antioxidative, and anticancer effects, with generally lower systemic toxicity compared to conventional chemotherapeutic agents [24]. In the context of CRC, various plant‐derived molecules such as curcumin, resveratrol, and berberine have been investigated for their ability to inhibit tumor growth, migration, and metastasis through the modulation of key oncogenic pathways, including those regulating EMT and tumor microenvironment remodeling [25, 26, 27]. Among the bioactive components extracted from traditional Chinese medicinal herbs, Picroside II, a major iridoid glycoside isolated from Picrorhiza scrophulariiflora, has attracted attention due to its demonstrated anti‐inflammatory and antioxidative properties [28, 29]. Preliminary studies have suggested that Picroside II exerts protective effects in models of organ injury and inflammation by modulating oxidative stress and apoptotic pathways [30]; however, its potential anticancer activity in particular CRC remains largely unexplored. The underlying mechanisms by which Picroside II may affect tumor cell behavior, including proliferation, migration, and invasion, have not been systematically elucidated. Given the critical role of EMT and the Notch1 signaling pathway in CRC metastasis, it is of interest to investigate whether Picroside II can interfere with these processes. Therefore, the present study aims to evaluate the effects of Picroside II on CRC cell proliferation, migration, invasion, and EMT, and to explore whether its actions involve modulation of the Notch1 signaling pathway.

2. Methods

2.1. Cell Culture and Treatment

Human CRC cell lines SW480 and SW620 were cultured in Dulbecco's Modified Eagle Medium (DMEM; Gibco, USA) supplemented with 10% fetal bovine serum (FBS; Gibco, USA) and 1% penicillin‐streptomycin (HyClone, USA) at 37°C in a humidified atmosphere containing 5% CO₂. Cells were routinely passaged at 70%–80% confluency using 0.25% trypsin‐EDTA (HyClone, USA) and were used for experiments during the logarithmic growth phase. Picroside II (Pic II) was purchased from Chengdu Must Bio‐Technology Co. Ltd. and dissolved in DMSO to prepare a 10 mM stock solution, which was stored at –20°C in the dark. Prior to treatment, the stock solution was diluted in complete DMEM to obtain working concentrations ranging from 10 to 100 μM, with the final DMSO concentration not exceeding 0.1%. 5‐Fluorouracil (5‐FU; Tianjin Jinyao Pharmaceutical Co. Ltd.) was used as a positive control at a final concentration of 2 μM. Cells were seeded into appropriate culture vessels according to the requirements of each assay and incubated overnight for adherence prior to drug treatment.

2.2. Cell Viability and Proliferation Assays

Cell viability and proliferation were assessed using the Cell Counting Kit‐8 (CCK‐8; APExBIO), colony formation assay, and EdU‐555 incorporation assay. For the CCK‐8 assay, SW480 and SW620 cells in the logarithmic growth phase were seeded into 96‐well plates at a density of 1 × 10⁴ cells/well and incubated for 12–24 h to allow for adherence. Cells were then treated with varying concentrations of Picroside II (1–90 μM) or 2 μM 5‐fluorouracil (5‐FU, positive control) and incubated for 24, 48, or 72 h. At each time point, 10 μL of CCK‐8 reagent was added to each well, and the cells were incubated for an additional 2.5–3 h at 37°C. Absorbance was measured at 450 nm using a microplate reader (Molecular Devices Corporation). The IC50 values and dose‐response curves were calculated using GraphPad Prism 8 software. For the colony formation assay, cells were seeded into six‐well plates at a density of approximately 300 cells/well and cultured for 24 h. After treatment with Picroside II at concentrations of 10, 25, and 100 μM for 72 h, the medium was replaced with fresh drug‐free medium, and cells were cultured for an additional 10–14 days. Colonies were then fixed with 4% paraformaldehyde and stained with crystal violet. The number of visible colonies was counted manually, and the colony formation rate was calculated. For the EdU‐555 incorporation assay, cells were seeded into six‐well plates and treated with Picroside II (10, 25, and 100 μM) for 48 h. Cells were then incubated with 2× EdU working solution (BeyoClick EdU‐555 Cell Proliferation Kit) for 2 h at 37°C. After fixation with 4% paraformaldehyde and permeabilization with 0.3% PBST, the click reaction was performed according to the manufacturer's protocol. Cell nuclei were counterstained with Hoechst 33342. Images were acquired using a fluorescence microscope (Olympus), and the percentage of EdU‐positive cells was calculated to quantify cell proliferation.

2.3. Migration and Invasion Assays

Cell migration ability was assessed using the wound healing assay and Transwell migration assay, while invasion capacity was evaluated using the Matrigel‐coated Transwell invasion assay. For the wound healing assay, SW480 and SW620 cells in the logarithmic growth phase were seeded into 6‐well plates at a density of approximately 1.5–2.0 × 10⁶ cells per well and cultured until 90%–100% confluence. A sterile 10 μL pipette tip was used to create a vertical scratch across the center of each well. The wells were washed twice with PBS to remove detached cells, and serum‐free DMEM containing different concentrations of Picroside II (10, 25, and 100 μM) or control medium was added. Wound closure was monitored at 0, 24, 48, and 72 h using an inverted microscope (Olympus), and migration was quantified by measuring the change in scratch width over time using ImageJ software. For the Transwell migration assay, 24‐well Transwell chambers with 8.0 μm pore polycarbonate membranes (Corning, #3422) were used. Cells were pretreated with Picroside II (10, 25, and 100 μM) for 24 h, then resuspended in serum‐free DMEM at a concentration of 2 × 10⁵ cells/mL. A total of 150 μL of the cell suspension was added to the upper chamber, while the lower chamber was filled with 600 μL of 20% FBS DMEM as a chemoattractant. After 48 h of incubation at 37°C, non‐migrated cells on the upper surface were removed with a cotton swab, and migrated cells on the lower surface were fixed with 4% paraformaldehyde and stained with crystal violet. Cells were counted in five randomly selected fields under a light microscope. For the invasion assay, the upper chamber of the Transwell insert was coated with 80 μL of Matrigel (Corning, diluted 1:8 with serum‐free DMEM) and incubated at 37°C until solidified. The subsequent procedures were identical to those of the migration assay. The number of invaded cells was determined in the same manner, and quantitative analysis was performed using ImageJ.

2.4. Western Blotting

Western blot analysis was performed to evaluate the expression levels of EMT markers and proteins associated with the Notch signaling pathway. SW480 and SW620 cells were treated with Picroside II (10, 25, and 100 μM) or vehicle control for 48 h. For the NICD overexpression rescue experiment, cells were transfected with an NICD overexpression plasmid or the corresponding empty vector using Lipofectamine 3000 according to the manufacturer's protocol. After transfection, cells were treated with Picroside II, and total proteins were collected for Western blot analysis. The expression levels of NICD, Notch1, Cleaved Notch1, HES1, E‐cadherin, N‐cadherin, Vimentin, and ZEB2 were examined. β‐actin was used as the internal loading control. After treatment, cells were washed with cold PBS and lysed in RIPA lysis buffer containing protease inhibitors (PMSF, 1 mM). The lysates were incubated on ice for 30 min and centrifuged at 15,000 rpm for 20 min at 4°C to collect the supernatant. Protein concentrations were quantified using the BCA protein assay kit, and equal amounts of protein (20–40 μg) were mixed with loading buffer, boiled for 10 min at 100°C, and separated on SDS‐PAGE gels (10% separation and 5% stacking gels). Proteins were transferred to PVDF membranes (0.45 μm, Millipore) using a wet transfer system, and membranes were blocked in 5% non‐fat milk prepared in TBST for 2 h at room temperature. The membranes were then incubated overnight at 4°C with primary antibodies against E‐cadherin, N‐cadherin, Vimentin, ZEB2, Notch1, Cleaved Notch1, RBPSUH, HES1, and β‐actin (used as the internal loading control). After washing, membranes were incubated with HRP‐conjugated secondary antibodies (anti‐rabbit or anti‐mouse, 1:4000) for 2 h at room temperature. Chemiluminescent detection was performed using ECL reagent, and bands were visualized using a chemiluminescence imaging system (AI600, GE Healthcare). Quantitative Western blot data are presented as mean ± SD from three independent biological replicates. Band intensities were quantified using ImageJ and normalized to β‐actin. Error bars represent SD. Statistical significance was determined by one‐way ANOVA followed by Tukey's post hoc test. Exact p‐values, F‐statistics, and degrees of freedom are provided in Supplementary Table S1. A schematic diagram of the canonical Notch1 signaling activation and the proposed inhibitory mechanism by Picroside II is shown in Figure 1.

Figure 1.

Picroside II inhibits colorectal cancer progression by blocking Notch1 signaling activation. It interferes with Notch1 activation by preventing the cleavage and nuclear translocation of the Notch intracellular domain (NICD). Upon ligand binding, Notch1 is cleaved by ADAM and γ‐secretase, releasing NICD, which translocates into the nucleus and forms a transcriptional activation complex (including CSL, MAML, and SKIP). This complex promotes transcription of downstream targets such as Hes1, which regulate epithelial–mesenchymal transition as well as cell proliferation, migration, and invasion.

2.5. Statistical Analysis

All data were analyzed using GraphPad Prism 8.0 software. Quantitative results are presented as mean ± standard deviation (SD) from at least three independent biological experiments. Data distribution was assessed using the Shapiro–Wilk test, and homogeneity of variance was evaluated using Levene's test. Comparisons between two groups were performed using unpaired two‐tailed Student's t‐tests when data met the assumptions of normality and equal variance. Multiple group comparisons were conducted using one‐way analysis of variance (ANOVA) followed by Tukey's post hoc test. For data that did not meet these assumptions, appropriate non‐parametric tests were applied. Exact p‐values, F‐statistics, degrees of freedom, and replicate numbers are reported in the corresponding figure legends or supplementary tables. A value of p < 0.05 was considered statistically significant.

3. Results

3.1. Picroside II Inhibits CRC Cell Proliferation

The CCK‐8 assay revealed that Picroside II treatment led to a dose‐ and time‐dependent reduction in cell viability in both SW480 and SW620 cells. The OD₄₅₀ readings increased with cell density and incubation time, with the maximum absorbance observed at 3 h. In SW480 cells, the OD₄₅₀ at 1 × 10⁵ cells/mL was 0.68 ± 0.04 at 1 h, 1.28 ± 0.06 at 2 h, and 2.14 ± 0.07 at 3 h (Figure 2A). Similarly, in SW620 cells, the OD₄₅₀ at 1 × 10⁵ cells/ml was 0.55 ± 0.03 at 1 h, 1.05 ± 0.04 at 2 h, and 1.84 ± 0.05 at 3 h (Figure 2B). Picroside II treatment resulted in a dose‐dependent reduction in cell viability. At 25 μM, the inhibition rates in SW480 and SW620 cells were 50.2% ± 3.1% and 55.2% ± 2.8%, respectively (Figure 2C). IC₅₀ values of Picroside II in SW480 cells were 27.1 μM at 24 h, 17.4 μM at 48 h, and 13.6 μM at 72 h. In SW620 cells, the IC₅₀ values were 28.9, 19.5, and 14.2 μM at the same time points, respectively (Figure 2F,I). In the colony formation assay, treatment with 10, 25, and 100 μM Picroside II reduced the number of colonies in SW480 cells to 66.1% ± 5.4%, 47.5% ± 4.2%, and 32.4% ± 3.9% of control levels, respectively (p < 0.05 for 10 μM, p < 0.01 for 25 and 100 μM). In SW620 cells, the colony numbers decreased to 73.8% ± 4.9%, 55.2% ± 3.8%, and 35.6% ± 3.1%, respectively (p < 0.05 for 10 and 25 μM, p < 0.01 for 100 μM) (Figure 3). EdU‐555 incorporation assays revealed that the proportion of EdU‐positive cells decreased in a concentration‐dependent manner. In SW480 cells, EdU‐positive rates were 49.3% ± 2.5% in control, 35.4% ± 2.1% at 10 μM, 31.2% ± 1.9% at 25 μM, and 25.6% ± 2.2% at 100 μM (p < 0.01 for all groups compared to control). In SW620 cells, rates were 56.7% ± 2.6%, 39.7% ± 1.7%, 33.4% ± 1.9%, and 26.3% ± 1.5%, respectively (p < 0.05 for 10 μM, p < 0.01 for 25 and 100 μM) (Figure 4).

Figure 2.

Effects of Picroside II on the viability of colorectal cancer cells. (A, B) Optimization of cell density and incubation time for the CCK‐8 assay in SW480 and SW620 cells, respectively. (C) Inhibition ratio of cell viability in SW480 and SW620 cells after treatment with different concentrations of Picroside II. (D, E) Dose–response curves of Picroside II in SW480 and SW620 cells at 24, 48, and 72 h, respectively. (F) IC₅₀ values of Picroside II in SW480 and SW620 cells at different treatment time points. Data are presented as mean ± SD from three independent experiments.

Figure 3.

Picroside II inhibits the colony formation ability of colorectal cancer cells. Representative images of colony formation assays in SW480 (A) and SW620 (C) cells treated with Picroside II at 0 (NC), 10, 25, or 100 μM for 72 h, followed by 10–14 days of culture in drug‐free medium. Picroside II treatment resulted in a dose‐dependent reduction in colony number and size. Quantitative analysis of colony formation in SW480 (B) and SW620 (D) cells.

Figure 4.

Picroside II reduces DNA synthesis activity in colorectal cancer cells as determined by EdU incorporation assay. Representative fluorescence microscopy images showing EdU (red), Hoechst (blue), and merged staining in SW480 (A) and SW620 (B) cells treated with Picroside II at 0, 10, 25, or 100 μM for 48 h. A dose‐dependent decrease in EdU‐positive nuclei was observed. Quantification of EdU‐positive cell percentages in SW480 (C) and SW620 (D) cells. The EdU incorporation rate significantly declined with increasing Picroside II concentrations.

3.2. Picroside II Suppresses CRC Cell Migration and Invasion

In the wound healing assay, the relative migration rate of SW480 cells at 48 h was 69.4% ± 2.6% in the control group, 43.7% ± 2.1% at 10 μM, 15.2% ± 1.8% at 25 μM, and 9.6% ± 1.5% at 100 μM. For SW620 cells, the corresponding values were 64.1% ± 2.3%, 36.4% ± 2.0%, 17.3% ± 1.7%, and 8.1% ± 1.3%, respectively. Statistically significant differences were observed at 100 μM compared with 0 μM in both cell lines (p < 0.01) (Figure 5). Additional time‐point observations showed limited wound closure at 12 h and clearer separation among treatment groups at 24–48 h. In the Transwell migration assay, the percentage of migrated SW480 cells was 100.0% ± 3.8% in control, 68.1% ± 3.2% at 10 μM, 41.6% ± 2.7% at 25 μM, and 26.3% ± 2.5% at 100 μM. In SW620 cells, the corresponding values were 100.0% ± 3.5%, 54.2% ± 3.0%, 36.7% ± 2.3%, and 20.4% ± 2.1%, respectively (Figure 6A,B). Preliminary dose‐range observations showed minimal effects at 5 μM and marked morphological changes at 200 μM; therefore, 10–100 μM was used for subsequent migration and invasion assays. In the Matrigel‐based invasion assay, the relative invasion rates of SW480 cells were 100.0% ± 4.1% in the control group, 74.6% ± 3.5% at 10 μM, 38.5% ± 2.9% at 25 μM, and 19.8% ± 2.2% at 100 μM. For SW620 cells, the corresponding values were 100.0% ± 3.9%, 66.9% ± 3.4%, 29.1% ± 2.8%, and 15.7% ± 2.0%, respectively (Figure 6C,D). The difference between 0 and 100 μM was statistically significant (p < 0.01). In the 0.1% DMSO vehicle control group, no measurable reduction in invasion was observed compared with the 0 μM Picroside II group. The 100 μM concentration used in the migration and invasion assays was higher than the 24–72 h IC50 values measured in the CCK‐8 assay. Reductions in wound closure, Transwell migration, and Matrigel invasion were also observed at 10 and 25 μM in both SW480 and SW620 cells.

Figure 5.

Wound healing assay showing the inhibitory effects of Picroside II on the migration of colorectal cancer cells. Representative phase‐contrast images of wound closure in SW480 (A) and SW620 (C) cells at 0 and 48 h after treatment with Picroside II at 0, 10, 25, and 100 μM. The width of the wound area was measured using yellow dashed lines. Quantitative analysis of relative migration rates in SW480 (B) and SW620 (D) cells demonstrated a significant inhibition of cell motility with increasing concentrations of Picroside II.

Figure 6.

Picroside II inhibits migration and invasion of colorectal cancer cells in Transwell assays. Representative images (A) and quantitative analysis (B) of Transwell migration assays in SW480 and SW620 cells treated with Picroside II (0, 10, 25, and 100 μM). Representative images (C) and quantification (D) of Matrigel‐coated Transwell invasion assays under the same treatment conditions. A concentration‐dependent reduction in migratory and invasive cell numbers was observed in both cell lines.

3.3. Picroside II Modulates EMT and Notch Signaling Pathways

Western blotting was used to measure the expression of EMT markers and proteins associated with the Notch signaling pathway in SW480 and SW620 cells treated with Picroside II at 0, 10, 25, and 100 μM for 48 h (Figure 7). In SW480 cells, E‐cadherin expression increased from 0.91 ± 0.06 at 0 μM to 1.03 ± 0.08, 1.18 ± 0.07, and 1.37 ± 0.06 at 10, 25, and 100 μM, respectively. In contrast, ZEB2, N‐cadherin, and Vimentin levels decreased with increasing Picroside II concentrations. ZEB2 decreased from 1.11 ± 0.07 to 0.56 ± 0.04, N‐cadherin from 1.02 ± 0.06 to 0.59 ± 0.03, and Vimentin from 1.05 ± 0.06 to 0.48 ± 0.03 at 100 μM. Significant differences were observed for E‐cadherin and Vimentin at 100 μM and for N‐cadherin at 25 μM compared with the 0 μM group. In a parallel DMSO vehicle control group (0.1%), the expression levels of EMT markers were similar to those in the 0 μM group, indicating that the vehicle did not measurably affect protein expression. In SW620 cells, E‐cadherin expression also increased from 0.59 ± 0.05 at 0 μM to 1.35 ± 0.07 at 100 μM, whereas ZEB2, N‐cadherin, and Vimentin decreased from 1.04 ± 0.06 to 0.73 ± 0.03, from 1.05 ± 0.05 to 0.61 ± 0.04, and from 1.21 ± 0.07 to 0.62 ± 0.03, respectively. For the Notch signaling pathway, Picroside II treatment reduced the expression of Notch1, Cleaved Notch1, RBP, and HES1 in both cell lines. In SW480 cells, Notch1 expression decreased from 0.97 ± 0.06 at 0 μM to 0.54 ± 0.03 at 100 μM, while Cleaved Notch1 decreased from 0.72 ± 0.04 to 0.22 ± 0.03. RBP and HES1 also declined from 1.08 ± 0.06 to 0.64 ± 0.03 and from 1.12 ± 0.06 to 0.69 ± 0.04, respectively. In SW620 cells, Notch1 expression decreased from 1.23 ± 0.07 to 0.75 ± 0.05, Cleaved Notch1 from 0.97 ± 0.05 to 0.29 ± 0.02, RBP from 1.16 ± 0.06 to 0.55 ± 0.04, and HES1 from 1.10 ± 0.05 to 0.61 ± 0.03. The reduction in Cleaved Notch1 was significant at 100 μM in both cell lines. All values represent means ± standard deviation from three independent experiments. β‐actin was used as the internal reference for normalization, and detailed densitometric values and statistical outputs are provided in Supplementary Table S1.

Figure 7.

Picroside II modulates the expression of EMT‐associated and Notch1 pathway–related proteins in colorectal cancer cells. (A) Representative Western blot images showing the expression of E‐cadherin, ZEB2, N‐cadherin, Vimentin, Notch1, Cleaved Notch1, RBPJ (RBPSUH), and HES1 in SW480 and SW620 cells following treatment with Picroside II at 0, 10, 25, and 100 μM for 48 h. β‐actin was used as the internal loading control. (B) Quantitative analysis of protein expression levels normalized to β‐actin, showing upregulation of E‐cadherin and downregulation of mesenchymal and Notch pathway proteins in a concentration‐dependent manner.

3.4. NICD Overexpression Partially Reverses Picroside II‐Induced Protein Changes

To examine the effect of NICD overexpression on Picroside II‐regulated Notch1‐ and EMT‐related protein expression, Western blotting was performed in the control, Picroside II, NICD overexpression, and Picroside II plus NICD overexpression groups (Figure 8A). Compared with the control group, Picroside II treatment reduced NICD expression from approximately 1.00 to 0.35‐fold and HES1 expression from approximately 1.00 to 0.30‐fold. NICD overexpression increased NICD expression to approximately 2.10‐fold and HES1 expression to approximately 2.00‐fold. In the Picroside II plus NICD overexpression group, NICD and HES1 expression levels were approximately 1.40‐fold and 1.20‐fold, respectively, which were higher than those in the Picroside II group but lower than those in the NICD overexpression group. For EMT‐related proteins, Picroside II increased E‐cadherin expression from approximately 1.00 to 2.05‐fold and reduced Vimentin expression from approximately 1.00 to 0.30‐fold. NICD overexpression decreased E‐cadherin expression to approximately 0.50‐fold and increased Vimentin expression to approximately 1.70‐fold. In the Picroside II plus NICD overexpression group, E‐cadherin expression was approximately 1.10‐fold, and Vimentin expression was approximately 0.95‐fold. Similar trends were observed for N‐cadherin and ZEB2 in the Western blot images. Quantitative analysis showed statistically significant differences between the control and Picroside II groups for NICD, HES1, E‐cadherin, and Vimentin expression. Significant differences were also observed between the NICD overexpression and Picroside II plus NICD overexpression groups for these proteins (Figure 8B).

Figure 8.

NICD overexpression partially reverses the inhibitory effects of Picroside II on Notch1 signaling and EMT‐related protein expression. (A) Western blot analysis of NICD, Notch1, Cleaved Notch1, HES1, E‐cadherin, N‐cadherin, Vimentin, and ZEB2 expression in CRC cells after treatment with Picroside II, NICD overexpression, or combined Picroside II and NICD overexpression. β‐actin was used as the internal loading control. (B) Quantitative analysis of NICD, HES1, E‐cadherin, and Vimentin protein expression normalized to β‐actin. Data are presented as mean ± SD from three independent experiments. *p < 0.05 compared with the indicated control group; #p < 0.05 compared with the NICD overexpression group.

4. Discussion

CRC continues to represent a substantial clinical and public health challenge worldwide due to its high incidence, metastatic potential, and limited long‐term therapeutic outcomes [31]. In this study, the biological activities of Picroside II on CRC cells were systematically evaluated through multiple experimental approaches, including cell viability assays, colony formation assays, EdU incorporation analysis, wound healing assays, Transwell migration and invasion assays, and Western blotting. The results demonstrated that Picroside II inhibited cell proliferation in a concentration‐ and time‐dependent manner, suppressed clonogenic survival, and reduced DNA synthesis activity. In addition, Picroside II exposure attenuated the migratory and invasive capabilities of SW480 and SW620 cells, accompanied by modulation of EMT‐related markers, characterized by the upregulation of E‐cadherin and the downregulation of N‐cadherin and Vimentin. Parallel analyzes revealed that Picroside II treatment led to a decrease in the expression of key components of the Notch1 signaling pathway, including Notch1, Cleaved Notch1, RBP, and HES1 proteins. These findings provide preliminary in vitro evidence that Picroside II suppresses CRC cell proliferation, migration, invasion, and EMT, at least partly through modulation of the Notch1 signaling pathway. However, further in vivo studies are needed to validate these effects and assess their pharmacokinetic and safety profiles.

Uncontrolled cellular proliferation represents a defining characteristic of malignancies, facilitating not only primary tumor expansion but also resistance to therapeutic interventions in CRC [32, 33]. In the current study, Picroside II was observed to suppress CRC cell proliferation in a dose‐ and time‐dependent manner, as indicated by CCK‐8 assays across multiple incubation periods. The progressive reduction of OD₄₅₀ values in SW480 and SW620 cells following Picroside II exposure suggests that the compound interferes with cellular metabolic activity, particularly mitochondrial dehydrogenase function, a proxy often used to reflect overall cell viability [34, 35]. This suppression of metabolic activity was accompanied by a decrease in colony‐forming ability, as demonstrated by colony formation assays, where the number and size of colonies declined substantially at concentrations of 25 and 100 μM. Since colony formation reflects not only cell survival but also long‐term reproductive integrity, these results imply that Picroside II compromises the clonogenic potential of CRC cells, a feature closely associated with tumor initiation and recurrence [36, 37]. In addition to impairing viability and clonogenicity, Picroside II reduced DNA synthesis, as evidenced by a decrease in EdU‐555 incorporation. Given that EdU labels newly synthesized DNA during the S phase of the cell cycle, the observed reduction implies that Picroside II may perturb cell cycle progression, potentially arresting cells at critical checkpoints or interfering with replication machinery [38, 39, 40]. These findings are consistent with reports on other natural compounds, such as geniposide and catalpol, which have been shown to modulate cyclin expression and induce cell cycle arrest in gastrointestinal tumors [41, 42]. Furthermore, the concentration range that produced these inhibitory effects did not overlap with concentrations known to cause acute cytotoxicity in normal cells, suggesting a potential therapeutic window that warrants further exploration. Although the molecular mediators underlying Picroside II's antiproliferative activity remain to be fully elucidated, the involvement of pathways regulating cell cycle control, mitochondrial integrity, and possibly DNA damage responses cannot be excluded [43].

The ability of tumor cells to migrate and invade through extracellular matrices is a critical prerequisite for metastasis, accounting for the majority of cancer‐related mortality in CRC [44]. In the present study, Picroside II was found to attenuate the migratory and invasive capacities of SW480 and SW620 cells, as demonstrated by wound healing and Transwell assays. In the wound healing model, Picroside II reduced the rate of wound closure in a concentration‐dependent manner, with a pronounced suppression observed at 25 and 100 μM. Since wound healing assays measure two‐dimensional collective migration, the reduction in gap closure suggests that Picroside II affects cytoskeletal dynamics, adhesion remodeling, or chemotactic signaling pathways, all of which are essential for coordinated cell movement [45, 46]. In Transwell migration and invasion assays, where cells must traverse a porous membrane or degrade a Matrigel barrier, respectively, Picroside II‐treated groups exhibited a substantial decline in the number of migrating and invading cells compared to controls. The inhibition of Transwell migration indicates that Picroside II may interfere with chemotaxis and mechanical deformability, whereas the suppression of Matrigel invasion suggests potential downregulation of matrix metalloproteinases (MMPs) or impairment of invadopodia formation, mechanisms well‐documented in metastatic CRC progression [47, 48]. Notably, lower concentrations of Picroside II did not elicit observable effects on migration, indicating a threshold‐dependent response, whereas at 200 μM, severe alterations in cell morphology and adhesion were noted, likely reflecting cytotoxic effects beyond specific antimigratory action. In addition, the interpretation of the 100 μM group requires caution. This concentration was higher than the IC50 values obtained from the CCK‐8 assay, and the reductions in wound closure, Transwell migration, and Matrigel invasion at this dose may partly reflect cytotoxic effects rather than migration‐ or invasion‐specific regulation. Therefore, the results at 10 and 25 μM are important for evaluating the effects of Picroside II under conditions with less extensive inhibition of cell viability. These observations align with studies on other phytochemicals, such as baicalin and scutellarin, which similarly inhibit CRC migration and invasion by targeting key regulators of cell motility and matrix degradation [49, 50, 51]. Although the precise molecular mediators underlying Picroside II's antimigratory effects were not delineated in this study, the concomitant suppression of EMT markers suggests that alterations in cellular phenotypes, such as reduced mesenchymal traits, may contribute substantially to the impaired migratory and invasive behavior.

The Notch signaling pathway plays a pivotal role in cell fate determination, stemness maintenance, and regulation of EMT during tumor progression [52]. In CRC, aberrant activation of Notch1 has been implicated in promoting tumor proliferation, invasion, metastasis, and chemoresistance, often correlating with poorer clinical outcomes [53, 54]. In this study, Western blot analysis revealed that Picroside II treatment was associated with a progressive decrease in the expression of Notch1 and its active form Cleaved Notch1 in both SW480 and SW620 cells, with the greatest reduction observed at the 100 μM concentration. This inhibition extended downstream to key transcriptional effectors of the Notch pathway, including RBPJ and HES1, both of which play critical roles in mediating canonical Notch signal transduction and sustaining mesenchymal characteristics in cancer cells [52, 55, 56]. The observed decrease in RBPJ and HES1 protein levels suggests that Picroside II may suppress not only the activation of Notch1 itself but also the subsequent transcriptional programs that facilitate EMT and tumor cell dissemination. Notably, Cleaved Notch1 expression exhibited a sharper decline compared to total Notch1 levels, indicating that Picroside II might preferentially interfere with the proteolytic cleavage and activation process mediated by γ‐secretase, a key step in Notch signaling [57, 58]. This potential inhibition of Notch1 cleavage warrants further investigation, as pharmacological blockade of γ‐secretase has been shown to impair CRC cell invasion and metastasis in preclinical models [54, 59]. The suppression of Notch signaling observed in Picroside II–treated CRC cells is particularly relevant given the multifaceted role of Notch1 in regulating not only EMT but also cellular stemness and resistance to therapy. Cancer stem‐like cells (CSCs) in CRC have been characterized by elevated Notch1 activity, which enhances their ability to initiate tumors, evade immune surveillance, and survive under cytotoxic stress [15, 21, 60]. By downregulating Notch1 and its downstream effectors, Picroside II may impair the maintenance of CSC‐like properties, thereby contributing to the overall reduction in cell proliferation, migration, and invasion observed in functional assays. Moreover, Notch1 signaling has been shown to engage in extensive crosstalk with other oncogenic pathways, including Wnt/β‐catenin, TGF‐β, and PI3K/AKT, amplifying its pro‐tumorigenic effects through complex regulatory networks [61, 62, 63]. The extent to which Picroside II might modulate these interacting pathways indirectly through Notch1 inhibition remains an important question for future research. Another consideration is the differential sensitivity of SW480 and SW620 cells to Picroside II in the context of Notch1 suppression. Although both cell lines exhibited downregulation of Notch1 and its effectors, the magnitude of inhibition appeared somewhat greater in SW620 cells, which are derived from metastatic lymph node lesions. This observation suggests that Picroside II may exert more pronounced inhibitory effects on cells with a higher baseline level of Notch1 activity, aligning with previous reports that metastatic CRC cells often exhibit elevated Notch1 signaling relative to primary tumors [53, 64, 65]. It would be valuable to extend these findings by analyzing Notch1 expression in a broader panel of CRC cell lines with varying degrees of differentiation and metastatic potential. Given the central role of Notch1 in tumor progression, the observed suppression of this pathway by Picroside II may warrant further mechanistic investigation. However, as the present study was limited to in vitro experiments, its in vivo efficacy, pharmacokinetic properties, safety profile, and translational potential remain to be determined [66, 67, 68].

Importantly, the NICD overexpression experiment provided additional evidence regarding the relationship between Notch1 signaling and the Picroside II‐induced changes in EMT‐related proteins. In the present study, Picroside II reduced the expression levels of NICD and HES1, whereas NICD overexpression increased the expression of these proteins. When NICD overexpression was combined with Picroside II treatment, the levels of NICD and HES1 were higher than those in the Picroside II group but lower than those in the NICD overexpression group. In parallel, NICD overexpression partially reversed the Picroside II‐induced changes in EMT‐related markers, including the increase in E‐cadherin and the decrease in Vimentin and ZEB2. These results suggest that Notch1 signaling is involved in the regulation of EMT‐related protein changes following Picroside II treatment. Nevertheless, the rescue effect was partial rather than complete. This finding indicates that Picroside II may also affect EMT and CRC cell progression through additional signaling pathways besides Notch1. Given the extensive crosstalk between Notch1 signaling and other pathways such as Wnt/β‐catenin, TGF‐β, and PI3K/AKT, further studies are needed to determine whether these pathways also contribute to the effects of Picroside II.

The present study has several limitations. First, all experiments were conducted in vitro using two CRC cell lines, SW480 and SW620. Although these models offer useful mechanistic insights, they cannot fully recapitulate tumor behavior in vivo, including the effects of the tumor microenvironment, immune interactions, drug metabolism, and tumor heterogeneity. In addition, this study did not include xenograft or other animal models, therefore, the in vivo antitumor activity of Picroside II remains unknown. Second, although the NICD overexpression experiment showed that Notch1 signaling is involved in the Picroside II‐induced regulation of EMT‐related proteins, whether Picroside II acts directly on the Notch signaling cascade or indirectly through upstream regulators remains unclear [69]. Third, the potential cytotoxicity and off‐target effects of Picroside II require further evaluation [70]. Because Picroside II was tested at concentrations up to 100 μM, additional cytotoxicity assays, such as LDH release or live/dead staining, are needed to distinguish specific anti‐migratory or anti‐invasive effects from non‐specific cytotoxicity. Finally, pharmacokinetic properties, biodistribution, effective tissue concentrations, and normal tissue toxicity were not assessed in this study. Therefore, no conclusions regarding therapeutic efficacy, systemic safety, or clinical applicability can be made based on the present in vitro data. Future studies should include in vivo tumor models, pharmacokinetic analysis, and normal tissue toxicity evaluation before the translational relevance of Picroside II can be determined. In conclusion, this study demonstrates that Picroside II inhibits CRC cell viability, migration, invasion, and EMT‐related changes in vitro. These effects are associated with downregulation of the Notch1 signaling pathway. Further in vivo validation and safety assessment are required to determine its therapeutic relevance.

Author Contributions

Rui Wang: writing – original draft, resources, methodology, writing – review and editing. Wei Zhang: software, formal analysis, methodology. Fang Wang: validation, data curation, software. Yong Zhang: formal analysis, data curation. Jiayi Li: writing – review and editing, funding acquisition, conceptualization.

Supporting information

Supporting File

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.