Design, synthesis, and preliminary antitumor mechanism of benzimidazole derivatives

2.1. Chemistry

4-Aminoacetophenone A (1.35 g, 0.01 mol), anhydrous K₂CO₃ (1.38 g, 0.01 mol), and chloroacetyl chloride B (0.8 mL, 0.01 mol) were combined in dimethylformamide (DMF) and stirred at ambient temperature for 24 h. The reaction mixture was then quenched with ice-cold water, and the resulting precipitate was collected by filtration, washed thoroughly, and vacuum-dried to yield intermediate C (N-(4-acetylphenyl)-2-chloroacetamide). Intermediate C (0.01 mol) was reacted with 2-mercaptobenzothiazole (0.01 mol) and K₂CO₃ (0.02 mol) in anhydrous acetone (20 mL) for 6 h at 25°C. The solid product was isolated via filtration, air-dried, and subsequently recrystallized from ethanol to afford compound D. A solution of D (0.001 mol) and a substituted formylhydrazide (0.001 mol) in ethanol (20 mL) was agitated at room temperature for 8 h. The precipitated crude product was filtered, washed, and subjected to ethanol recrystallization to obtain the target derivatives E.

2.2. Cell culture

Human cancer cell lines K562, A549, 5637, PC-3, and HEK-293 were acquired from the Cell Bank of the Chinese Academy of Sciences (Kunming). Cells were maintained in appropriate medium (RPMI-1640/DMEM) supplemented with 10% fetal bovine serum and 1% penicillin-streptomycin, under standard culture conditions (37°C, 5% CO₂ humidified atmosphere) [10].

2.3. Cell viability assay

The antiproliferative effects of target compounds against A549, 5637, PC-3, K562, and HEK-293 cell lines were evaluated through MTT assay according to established protocols [11]. In brief, cells pre-cultured in 96-well plates for 24 h were exposed to test compounds for 48 h, with 5-fluorouracil and imatinib serving as clinical reference agents. Initial screening at a concentration of 10 μM was followed by dose-response assessment using two-fold serial dilutions of compound E12 (1.25-20 μM). Post-treatment, cells were incubated with MTT solution (5 mg/mL, 10% v/v) for 4 h prior to supernatant removal and subsequent dissolution of formazan crystals in DMSO (150 μL/well) under dark-phase orbital agitation. Optical density measurements at 490 nm were recorded using a Tecan Infinite M200 Pro microplate reader, with triplicate determinations ensuring experimental reproducibility.

2.4. Cell apoptosis analysis

Quantitative assessment of programmed cell death was conducted via FITC-Annexin V/propidium iodide dual-staining methodology as previously described [12]. After a 24-h attachment period of exponentially growing cells in 6-well plates (2 × 10⁵ cells/well), cells were treated with E12 at various concentrations (10-20 μM) for 24 h. Cells were detached by trypsinization, washed twice with PBS, and resuspended in 100 μL binding buffer. Subsequent dual-labeling involved sequential addition of FITC-conjugated Annexin V (5 μL) and PI (5 μL) with gentle vortexing, followed by incubation in the dark at room temperature for 15 min and 5 min, respectively. The volume was then adjusted to 500 μL with binding buffer, and apoptotic subpopulations were quantified through multicolor flow cytometry (BD FACS Calibur™). Computational analysis of quadrant statistics was executed using FlowJo™ software (v10.8.1).

2.5. Mitochondrial membrane potential (ΔΨm) analysis

Mitochondrial membrane potential (ΔΨm) alterations in E12 treated K562 cells were monitored using the JC-1 fluorescent cationic dye (Solarbio, JC-1 Kit #M8650) according to manufacturer specifications. This potential-dependent probe undergoes J-aggregate (590 nm, red) to monomer (530 nm, green) spectral shift upon mitochondrial depolarization, with red/green fluorescence intensity ratio serving as a quantitative indicator. Cells were seeded in 6-well clusters (2 × 10⁵ cells/well) and allowed to adhere for 24 h before exposure to E12 for 48 h. Post-treatment, cells were stained with JC-1 working solution (10 μg/mL) in serum-free medium at 37°C (5% CO₂) for 30 min. Following two cold PBS washes (4°C, 1000 rpm, 5 min), cells were resuspended in detection buffer. Flow cytometric acquisition was performed on a BD FACS Calibur™ system with FL1 (Fluorescence channel 1) (530/30 nm band-pass filter) and FL2 (Fluorescence channel 2) (585/42 nm band-pass filter) channels for green/red signal quantification, respectively. Data processing utilized FlowJo™ software (v10.8.1).

2.6. Cell cycle assay

Cell cycle progression was analyzed using a DNA Content Quantitation Assay Kit (Solarbio, #CA1510) according to established methodologies [13]. Synchronized K562 cells (2 × 10⁵ cells/well in 6-well plates) underwent 24 h pre-culture prior to 24 h E12 exposure (10–20 μM gradient concentrations). Post-exposure cellular suspensions underwent cryofixation in 75% ethanol (-4°C, 12 h), followed by PBS rehydration and RNase A digestion (100 μL, 37°C, 30 min) to eliminate RNA interference. Nuclear DNA was subsequently stained with propidium iodide working solution (50 μg/mL in PBS, 400 μL) under dark-phase conditions (4°C, 30 min). Multiparametric flow cytometry analysis was performed on a BD FACS Calibur™ system with 488 nm excitation, utilizing FL2-A/FL2-W parameters for doublet discrimination. ModFit LT™ v5.0 software was used for data analysis

2.7. Intracellular reactive oxygen species (ROS) detection

Intracellular oxidative stress levels were quantified using the 2’,7’-dichlorodihydrofluorescein diacetate (DCFH-DA) fluorogenic probe according to established protocols [14]. E12-treated K562 cells (2 × 10⁵ cells/well in 6-well plates, 24 h pre-culture) were loaded with 10 μM DCFH-DA in serum-free RPMI-1640 (37°C, 5% CO₂, 20 min) to permit probe internalization and esterase-mediated deacetylation. Post-staining cellular suspensions underwent triple washing cycles with ice-cold PBS (300 × g, 5 min) to remove extracellular dye, followed by immediate flow cytometric analysis on a BD FACS Calibur™ platform. Fluorescent signal acquisition through FL1 channel was conducted within 30 min post-harvest to minimize auto-oxidation artifacts. Geometric mean fluorescence intensity (MFI) quantification was executed in FlowJo™ software (v10.8.1).

2.8. Western blotting

Cellular protein extraction was performed via ice-cold lysis buffer incubation (RIPA supplemented with 1× protease/phosphatase inhibitor cocktail) at 4°C for 60 min with periodic gentle inversion. Clarified lysates obtained by 12,000 rpm centrifugation (4°C, 15 min) were quantified using Solarbio BCA Kit (#PC0020) against BSA calibration curves. Equal protein loads (30 μg) were resolved on discontinuous SDS-PAGE gels (10%, 12.5%, or 15% acrylamide concentration based on target molecular weight) under constant voltage (120 V, 90 min). Electrophoretic transfer to 0.22-μm PVDF membranes was conducted in Towbin buffer (25 mM Tris, 192 mM glycine, 20% methanol) using a wet tank system at 200 mA constant current (90-120 min). Membranes were blocked with 5% nonfat milk in TBST (2 h at room temperature) prior to sequential immunodetection. Primary antibody probing was conducted at 4°C overnight in antibody diluent (Biosharp #BL506A) with the following specificities: BCR-ABL (Affinity AF7716, 1:500), cell cycle regulators Cyclin B1 (Affinity AF6188, 1:500) and CDK1 (Affinity DF8024, 1:500), apoptotic modulators Bax (Proteintech 5099-2-Ig, 1:2000)/Bcl-2 (Huabio H651066060, 1:2000), and p21 (Proteintech 10355-1-AP, 1:1000), using β-actin (Affinity AF7018, 1:3000) as loading control. After three TBST washes (5 min each), membranes were incubated with HRP-conjugated secondary antibodies (Absin abs20040, 1:10,000) for 1 h at room temperature. Immunoreactive bands were visualized using a Vilber Bio Imaging PulseTS6 chemiluminescent documentation system equipped with a cooled CCD camera; ImageJ software was used for quantitative analysis of gray values.

2.9. Molecular docking

The structure of compound E12 was drawn in ChemDraw, optimized with the MM2 force field, and exported to 3D format. The BCR-ABL crystal structure (PDB ID:3OZX) was acquired from RCSB PDB. Receptor preparation involved water/ligand removal in PyMOL 2.3.4, followed by hydrogen addition and charge neutralization using AutoDockTools. Both receptor and ligands were formatted in PDBQT files for docking simulations conducted in AutoDock Vina 1.1.2. Protein-ligand interactions were analyzed using PLIP and visualized with PyMOL.

2.10. Statistical analysis

Quantitative data were expressed as mean ± standard deviation (SD) and were analyzed using GraphPad Prism version 9.0. Intergroup comparisons were performed by one-way or two-way ANOVA with statistical significance set at P < 0.05.

3.1. Chemical synthesis

A series of benzimidazole derivatives (E1-14) was successfully synthesized through a three-step process (Scheme 1). The initial step involved the condensation of 4-aminoacetophenone (A) with chloroacetyl chloride (B) in the presence of K₂CO₃ and DMF, yielding intermediate N-(4-acetylphenyl)-2-chloroacetamide (C) in high yield. In the second step, intermediate C underwent nucleophilic substitution with 2-mercaptobenzimidazole under reflux in DMF to give intermediate D. This step was key for introducing the benzimidazole ring, which is known for its potential biological activity. The final step involved the reaction of intermediate D with various substituted formylhydrazides in ethanol under reflux, allowing for the introduction of diverse functional groups on the phenyl ring, which were designed to modulate the anticancer activity of the compound. Successful implementation of the synthetic strategy was verified through multinuclear magnetic resonance spectroscopy (¹H and ¹³C NMR) coupled with high-resolution mass spectral characterization.

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Scheme 1.

Synthesis of benzimidazole derivatives E1-14.

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Figure 1.

(a). Chemical structure formula of compound E12. (b). Dose-dependent growth trajectories of K562 and HEK-293 following 48 h E12 exposure.

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3.2. Cytotoxic activity

The antitumor activities of all target compounds (E1-E14) were evaluated via the MTT assay against four human cancer cell lines (A549, 5637, PC-3, and K562), with 5-fluorouracil as a positive control. As summarized in Table 1, the derivatives exhibited distinct activity profiles across cell lines: most compounds showed limited activity against K562 cells (<30% inhibition at 10 μM), with E12 (62.71%) being exceptions, while moderate effects were observed in A549 (e.g., E5: 52.20%; E12: 54.08%) and PC-3 (e.g., E1: 53.11%; E12: 55.87%). In contrast, all derivatives, including E12 (33.20%), displayed low potency against 5637 cells (<35% inhibition), likely due to intrinsic resistance mechanisms. Among the series, E12 emerged as the most promising candidate. Compound E12 (structure in Figure 1a) exhibited pronounced selectivity toward K562 leukemia cells, demonstrating dose-dependent suppression (0-40 μM, 48 h; Figure 1b, Table 2). Notably, E12 displayed a 3.6-fold preferential cytotoxicity in K562 cells (IC₅₀ = 11.07 μM) over normal counterparts (IC₅₀ > 40 μM), suggesting tumor-specific bioactivity.

3.3. Compound E12 induced apoptosis of K562 cells

Programmed cell death serves as a fundamental mechanism for maintaining cellular homeostasis and eliminating oncogenic transformations. The inability to execute apoptosis represents a critical oncogenic driver, facilitating tumor survival and therapeutic resistance [15,16]. To investigate the apoptotic potential of E12, we employed Annexin V-FITC/PI dual-staining coupled with flow cytometric analysis. Quantitative assessment revealed dose-dependent apoptotic induction in K562 cells, with 10, 15, and 20 μM E12 treatments increasing apoptotic fractions by 3.88%, 6.16%, and 7.86%, respectively, comparable to 1 μM imatinib (6.98%) versus untreated controls (Figure 2a).

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Figure 2.

Mechanistic evaluation of E12-induced apoptosis in K562 cells. (a) Annexin V/PI-based apoptosis profiling. (b) Bax/Bcl-2 ratio determination by immunoblotting. Data: mean ± SD of triplicates; significance levels: *P<0.05, ***P<0.001, ****P<0.0001 (ANOVA with Dunnett’s multiple comparison).

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Complementary western blot analysis demonstrated that 20 μM E12 treatment upregulated the pro-apoptotic protein Bax expression while downregulating anti-apoptotic protein Bcl-2, significantly altering the Bax/Bcl-2 equilibrium (Figure 2b). This molecular shift indicates activation of mitochondrial-mediated apoptosis, a key pathway in oncological therapeutics [17]. The concordance between flow cytometric quantification and the immunoblotting results substantiates the capacity of E12 to trigger intrinsic apoptosis in leukemia cells. Collectively, these mechanistic insights suggest that E12 exerts its antileukemic effects through targeted modulation of apoptotic regulators, highlighting its potential as a chemotherapeutic candidate.

3.4. Compound E12 induced mitochondrial membrane potential collapse in K562 cells

Mitochondria are central to cellular bioenergetics and the regulation of apoptotic cascades [18,19]. The depolarization of mitochondrial membrane potential (ΔΨm) represents an early hallmark of apoptosis, making it a key therapeutic target in oncology due to its capacity to selectively eliminate malignant cells while preserving normal tissue integrity. This selectivity positions ΔΨm disruption as a strategic approach to trigger tumor-specific apoptosis and inhibit cancer progression. To assess the mitochondrial-targeting activity of E12, K562 cells were exposed to escalating concentrations of the compound (10-20 μM) and analyzed using the potential-sensitive probe JC-1. Flow cytometric quantification (Figure 3) demonstrated a dose-dependent fluorescence shift from red to green in E12-treated cells. Specifically, the proportion of green-fluorescent cells increased from 5.24% (untreated control) to 17.3%, 28.1%, and 32.4% at 10, 15, and 20 μM E12, respectively. These findings not only confirm the ability of E12 to induce mitochondrial dysfunction but also suggest its role in initiating K562 cells’ apoptotic pathways.

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Figure 3.

JC-1 staining and flow cytometric assessment of mitochondrial membrane potential (ΔΨm) in K562 cells treated with E12 (10–20 μM, 24 h). DMSO and imatinib served as negative and positive controls, respectively. Fluorescence shifts from red (aggregated JC-1, intact ΔΨm) to green (monomeric JC-1, ΔΨm loss) indicate mitochondrial depolarization. Data are expressed as mean ± SD (n=3). Statistical significance (****P < 0.0001) was determined by one-way ANOVA with Dunnett’s post hoc test.

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3.5. Compound E12 arrested the K562 cell cycle at G2\M phase

Cell cycle control is essential for maintaining genomic stability, and its aberrant activation is a defining feature of oncogenesis. Pharmacological targeting of cell cycle checkpoints has gained traction as a therapeutic paradigm to constrain neoplastic growth and trigger apoptosis in malignancies [20,21]. To evaluate the impact of E12 on cell cycle dynamics, K562 cells were treated with a concentration gradient (0-20 μM) of the compound for 24 h, followed by cell cycle profiling via PI staining and flow cytometry. Quantitative analysis (Figure 4a) demonstrated a concentration-dependent accumulation of cells in the G2/M phase, with proportions rising from 7.76% (untreated) to 13.12%, 14.57%, and 19.16% at 10, 15, and 20 μM E12, respectively.

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Figure 4.

(a) Flow cytometric cell cycle profiling of K562 cells treated with E12 (10-20 μM, 24 h) using propidium iodide (PI) staining. DMSO and imatinib served as negative and positive controls, respectively. (b) Immunoblot detection of G2/M phase regulatory proteins (Cyclin B1, CDK-1, p21) in K562 cells exposed to escalating concentrations of E12 (10-20 μM). β-actin was used as a loading control. Data represent mean ± SD (n=3). Statistical significance (*P < 0.05, ***P < 0.001, ****P < 0.0001) was determined by two-way ANOVA with Dunnett’s post hoc test.

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Given the central role of p21, Cyclin B1, and CDK1 in governing G2/M transition, we further examined their expression patterns through immunoblotting. As illustrated in Figure 4(b), E12 treatment markedly upregulated p21 while downregulating Cyclin B1 and CDK1 in a dose-responsive manner. These coordinated changes in checkpoint regulators provide mechanistic evidence for E12-induced G2/M phase arrest, positioning it as a modulator of cell cycle progression in leukemia cells.

3.6. Compound E12 induces redox imbalance through ROS elevation in K562 cells

Reactive oxygen species (ROS) exhibit dual roles in cancer biology, functioning as signaling molecules supporting tumorigenesis at moderate levels while triggering cytotoxic effects when exceeding cellular tolerance thresholds [22,23]. This redox paradox underlies conventional anticancer strategies, including radiation and chemotherapy. DCFH-DA fluorescence assay was used to assess the changes in intracellular ROS levels in K562 cells after E12 treatment. As shown in Figure 5, the observed significant ROS accumulation implies that E12 effectively disrupts redox homeostasis in these leukemia cells. Mechanistically, such oxidative overload induces macromolecular damage to critical cellular components (DNA, proteins, and lipids), subsequently activating programmed cell death pathways as documented in oxidative stress responses [24]. Notably, the high proliferative capacity of K562 cells makes them particularly susceptible to redox manipulation, as rapidly dividing cells inherently display heightened sensitivity to the accumulation of oxidative damage. This mechanism aligns with therapeutic approaches targeting cancer cell-specific vulnerabilities while sparing normal cells with lower metabolic demands.

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Figure 5.

Flow cytometric quantification of intracellular ROS accumulation in K562 cells. DCFH-DA fluorescent staining profiles of cells treated with compound E12 (10-20 μM, 24 h). Vehicle control (0.1% DMSO) and imatinib mesylate (1 μM) were included as negative and reference controls, respectively. Quantitation of ROS levels expressed as mean fluorescence intensity (MFI). Data represent mean ± SD from three biological replicates (n=3). Statistical significance (***P < 0.001, ****P < 0.0001) versus vehicle control was determined by one-way ANOVA with Dunnett’s post hoc test

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3.7. Compound E12 suppresses BCR-ABL fusion protein expression

BCR-ABL, a primary therapeutic target in CML, promotes leukemogenesis by constitutively activating intracellular signaling cascades such as Ras/Raf/MAPK, JAK/STAT3, and PI3K/AKT via its tyrosine kinase activity. This dysregulation disrupts hematopoietic stem/progenitor cell differentiation and drives CML progression [25-27]. Given its central role, the inhibition of BCR-ABL-mediated signaling represents a strategic focus for CML drug development. To evaluate the impact of compound E12 on BCR-ABL expression, total cellular proteins were isolated from K562 cells exposed to E12 (10 and 20 μM) and analyzed via Western Blot. As illustrated in Figure 6, E12 treatment markedly decreased BCR-ABL protein levels. Since BCR-ABL drives aberrant proliferation and apoptosis resistance in CML cells, its suppression by E12 likely disrupts oncogenic signaling in K562 cells. This modulation of BCR-ABL expression correlates with impaired cell viability and enhanced apoptotic activity, underscoring the compound’s potential therapeutic mechanism.

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Figure 6.

Expression of BCR-ABL protein in K562 cells following compound treatment (*P < 0.05, ***P < 0.001, ****P < 0.0001 vs. control; n=3; analyzed by one-way ANOVA with Dunnett’s post hoc test).

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3.8. Molecular docking

The molecular docking analysis revealed a strong interaction between the target protein BCR-ABL and the small molecule E12, as indicated by a binding energy of -9.3 kcal/mol. This value suggests a favorable binding affinity, highlighting the potential of E12 to act as a potent inhibitor of BCR-ABL. As shown in Figure 7, several hydrophobic interactions contribute significantly to the stability of the complex. Key hydrophobic contacts were observed between E12 and amino acid residues LEU248 (leucine), VAL256 (valine), ALA269 (alanine), VAL289 (valine), MET318 (methionine), ASP381 (aspartic acid), and PHE382 (phenylalanine), which collectively form a hydrophobic pocket within the active site of BCR-ABL. Furthermore, hydrogen bonding interactions were identified as critical contributors to the specificity of the binding. Notably, THR315 (threonine) and ASP381 (aspartic acid) formed hydrogen bonds with E12, reinforcing its stable attachment to the active site of the protein. The interaction with THR315 is particularly significant, as this residue is involved in the ATP-binding domain of BCR-ABL and plays a pivotal role in the catalytic activity of the protein [28]. These findings suggest that E12 effectively occupies the active site of BCR-ABL, disrupting its kinase activity through robust hydrophobic interactions and hydrogen bonding. The binding mode of E12 demonstrates its potential as a selective inhibitor. This insight provides a molecular basis for further development of E12 as a targeted therapy for chronic myeloid leukemia.

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Figure 7.

Molecular docking model of E12 with BCR-ABL. (Grey dotted line: hydrophobic interactions; Blue solid line: hydrogen bond).

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