Bee venom and thymoquinone combination inhibits cancer cells by inducing cell cycle arrest and apoptosis

Abstract

Natural products have gained significant interest in cancer therapy. Thymoquinone (TQ), a bioactive compound mainly derived from Nigella sativa seeds, and bee venom (BV), a complex mixture of bioactive components secreted by honeybees (Apis mellifera), are notable examples. This study aimed to evaluate the synergistic anticancer effects of TQ-BV combination on selected cancer cell lines—HeLa, MCF-7, HCT—and normal human skin fibroblasts (HSF). BV was collected using a novel remote-controlled extraction device designed to maintain venom purity by isolating it from environmental contaminants and minimizing light exposure. HPLC was used to quantify the main components of the venom, thereby detecting the sample’s purity. MTT assays assessed cell viability. Apoptotic activity was analyzed through Annexin V-FITC/PI staining. Flow cytometry was used to evaluate the cell cycle distribution, focusing on the Sub-G1, G1, and S phases in HeLa cells. Results from the MTT assay showed that the TQ-BV combination showed markedly increased potency against HeLa cells with an IC₅₀ value of (1.495 ± 0.198 µg/mL). Combination Index (CI) analysis confirmed a synergistic effect in all cell lines. The apoptosis assay revealed an increase in both early and late apoptotic cells with the combination treatment in HeLa cells, which exhibited a notable rise in late-stage apoptosis, indicating enhanced apoptotic activity. Cell cycle analysis revealed that the combination appeared to induce arrest at the G1 and S phases, which may have contributed to reduced proliferation. Overall, the TQ-BV combination exhibited strong anticancer potential by inducing cell cycle arrest and promoting apoptosis. The high purity of BV, achieved through an optimized extraction method, may have enhanced its efficacy. Further in vivo studies are needed to confirm these findings and explore potential clinical applications.

Supplementary Information

The online version contains supplementary material available at 10.1038/s41598-025-28733-9.

Introduction

Cancer remains one of the most serious health challenges globally, accounting for millions of deaths each year and posing a significant burden on healthcare systems¹. According to the World Health Organization (WHO), cancer is the second leading cause of death worldwide and is responsible for approximately 10 million deaths in 2022 alone^2,3. The growing cancer burden underscores the pressing need for efficient and available treatment options. Standard therapies, particularly chemotherapy, have become the foundation of cancer treatment. However, these treatments often lead to severe side effects, including nausea, fatigue, and hair loss, which significantly reduce patients’ quality of life and overall well-being^4,5. More importantly, many cancer cells develop tolerance to chemotherapy drugs, making these treatments less effective over time and highlighting a significant unfulfilled need for innovative therapeutic approaches⁶.

The shortcomings associated with conventional cancer therapies have motivated researchers to investigate substitute treatment options that are both effective and well-tolerated. Natural products have long been used in traditional medicine and have become an area of strong interest in oncology^7,8. Phytochemicals and other biologically active compounds derived from nature have shown potential as anticancer agents^9–11. Due to their varied pharmacological properties and less severe side effects compared to synthetic drugs. Studies of natural products have shown that many of these compounds can regulate key cellular pathways involved in cancer development, such as apoptosis, proliferation, and angiogenesis, making them potential candidates for anticancer therapy¹².

Thymoquinone (TQ), a bioactive compound primarily extracted from the seeds of Nigella sativa and commonly associated with black cumin, has recently garnered significant attention due to its wide range of pharmacological effects, including antioxidant, anti-inflammatory, and antimicrobial activities^13,14. TQ has demonstrated notable anticancer properties, making it a promising candidate for cancer treatment¹⁵. Studies indicate that TQ induces apoptosis, inhibits cell proliferation, and reduces cancer-related angiogenesis in various cancer cell lines. These effects are mediated through multiple mechanisms, including the regulation of key signaling pathways such as NF-κB, PI3K/AKT, and MAPK¹⁶. Additionally, thymoquinone exhibits antiangiogenic properties, leading to the inhibition of new blood vessel formation required by tumors for growth and metastasis. By downregulating VEGF and other angiogenic factors, thymoquinone effectively limits angiogenesis and, consequently, tumor growth and spread¹⁷.TQ’s capacity to modulate these pathways positions it as a versatile agent capable of acting at different stages of tumor development.

Bee venom (BV), also known as apitoxin, is a complex mixture of bioactive components secreted by honeybees (Apis mellifera). Historically used in apitherapy for its anti-inflammatory and analgesic properties^18,19. BV has recently emerged as a potential anticancer agent. Its main components, including melittin, phospholipase A2, and apamin, contribute to its wide range of biological activities. Among these, melittin is especially notable for its powerful anticancer effects. Studies have shown that BV and melittin can induce apoptosis, disrupt cell membranes, and inhibit metastasis in various cancers, including breast, prostate, and liver cancers²⁰. These findings suggest that BV has mechanisms of action that complement those of TQ, possibly boosting the overall anticancer effect when used together.

The search for potential molecular targets in cancer cells is enhanced by combining natural compounds that exhibit synergistic interactions and established anticancer properties²¹. The TQ-BV combination presents a novel therapeutic approach with promising synergistic antitumor effects. Preliminary studies suggest that using multiple bioactive compounds concurrently can yield improved outcomes through synergy, enabling greater efficacy at lower doses and potentially minimizing side effects. The combination is likely to amplify their antitumor activities, including the induction of apoptosis, inhibition of cell proliferation, and suppression of metastasis. Since they act via distinct yet complementary mechanisms, this combination therapy may address several challenges in cancer treatment, particularly in overcoming drug resistance and reducing adverse effects. Ultimately, this research aims to enhance our understanding of these natural agents in oncology, thereby contributing to the development of more effective and better-tolerated cancer therapies.

Materials and methods

Materials

Bee venom

BV extraction was carried out using a custom-built, remote-controlled device designed to be placed directly inside the beehive. This setup enables extraction without exposing the BV sample to light, which is crucial because BV is highly sensitive to light and can degrade when exposed. Additionally, the device stays within the hive environment, effectively shielding the sample from external dust and pollutants, thus maintaining its purity. Once placed inside the hive, the device is operated remotely, minimizing disturbances to both the bees and the sample. It uses a low-intensity electric current that encourages bees to sting the collection surface. A fine nylon layer covers the glass of the collection plate, enabling the stinger to pierce through and deposit venom onto the glass without harming the bee, as the stinger remains intact. Nylon also acts as a protective barrier, preventing contamination from hive materials and ensuring a cleaner sample. The extraction process generally takes between one and two hours, during which bees release venom in response to the electric stimulus. The device is gently removed from the hive at the end of the procedure. The collected venom on the glass plates is then transferred to a chilled, temperature-controlled box, which shields it from light and temperature fluctuations during transport. The venom is immediately stored in a deep freezer at -80 °C to preserve its biochemical properties for future research and applications. This method was tested on farms in Upper Egypt using the Apis mellifera carnica bee strain, known for its adaptability and resilience. Additionally, BV was extracted from specialized farms in Jordan to evaluate the new technique. The combination of remote operation, hive isolation, and strict post-extraction handling ensures the venom remains as pure and potent as possible, making it ideal for scientific and therapeutic uses.

Thymoquinone

TQ was obtained from Sigma-Aldrich (St. Louis, MO, USA). Cell culture medium, fetal bovine serum (FBS), and other related cell culture reagents were purchased from Gibco™ (Thermo Fisher Scientific, Grand Island, NY, USA).

Cell lines

Cell lines used in this study include HCT (colorectal cancer), HeLa (cervical cancer), and MCF-7 (breast cancer). Breast adenocarcinoma and human skin fibroblasts (HSFs) were procured from Nawah Scientific Inc., Cairo, Egypt. Human skin fibroblasts (HSFs) were included as a normal, non-cancerous cell line to evaluate the selectivity and cytotoxicity of BV, TQ, and their combination toward normal cells. HCT and HeLa cells were cultured in RPMI-1640 medium, whereas MCF-7 and HSF cells were maintained in DMEM. Both media were supplemented with 100 µg/mL streptomycin, 100 units/mL penicillin, and 10% (v/v) heat-inactivated fetal bovine serum (FBS) to support optimal cell growth and proliferation. Cultures were incubated in a humidified atmosphere containing 5% (v/v) CO₂.

Methods

MTT assay

The cytotoxic effects of BV and TQ on selected cell lines were assessed using the MTT assay. For this test, cells were initially seeded into 96-well plates at a density of 5 × 10³ cells per well in 100 µL of their respective complete growth medium. The plates were then incubated at 37 °C in a controlled environment with 5% CO₂ for 24 h, allowing the cells to adhere and reach a stable growth phase before treatment. After the pre-incubation period, 100 µL of fresh complete medium containing either BV or TQ at specified concentrations was added to each well. BV was prepared at concentrations of 400, 200, 100, 50, 25, 12.5, 6.25, and 3.12 µg/mL. TQ was prepared at concentrations of 131.36, 65.68, 32.84, 16.42, 8.21, 4.105, 2.0525, and 1.02625 µg/mL. For combination treatments, different concentration protocols were used across the cell lines. In MCF-7, HCT, and HSF cells, bee venom (BV) was fixed at 19 µg/mL, while serial dilutions of thymoquinone (TQ) were prepared at 6.568, 5.9112, 5.2544, 4.5976, 3.9408, 3.284, 2.6272, and 1.9704 µg/mL. Conversely, TQ was fixed at 6.568 µg/mL, with serial dilutions of BV at 20, 18, 16, 14, 12, 10, 8, and 6 µg/mL. For HeLa cells, different concentration series were used: BV was fixed at 9 µg/mL, while serial dilutions of TQ included 3.284, 2.956, 2.627, 2.299, 1.970, 1.642, 1.314, and 0.985 µg/mL. Additionally, TQ was fixed at 3.119 µg/mL, and serial dilutions of BV were prepared at 10, 9, 8, 7, 6, 5, 4, and 3 µg/mL.

After treatment, cells were incubated for an additional 48 h under the same conditions (37 °C, 5% CO₂) to allow the effects to manifest. After 48 h of drug exposure, the medium was carefully aspirated from each well to remove any leftover treatment. To assess cell viability, 20 µL of MTT reagent was added to each well. MTT, which stands for 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide, is a yellow, water-soluble compound that is turned into purple formazan crystals by the mitochondrial enzymes of metabolically active cells. The MTT reagent was prepared as a stock solution at a concentration of 1 mg/mL. Following the addition of MTT, 100 µL of phosphate-buffered saline (PBS) was added to each well to maintain isotonic conditions and support cellular activity during the assay. The plates were then incubated at 37 °C for an additional 4 h, allowing sufficient time for viable cells to convert MTT into insoluble formazan crystals. Once the incubation was complete, PBS was gently removed from each well, leaving the formazan crystals intact. To dissolve the formazan crystals, 100 µL of dimethyl sulfoxide (DMSO) was added to each well. DMSO is an organic solvent commonly used in MTT assays to dissolve formazan and produce a colored solution. The absorbance of this solution was measured at a wavelength of 570 nm using a multi-well plate reader (FLUOstar Omega, BMG LABTECH, a German manufacturer specializing in laboratory technology). The absorbance reading, taken at the peak absorbance wavelength for formazan, indicates the viability of cells in each well, with higher absorbance corresponding to greater cell viability. Experiments were conducted in three independent replicates to ensure the accuracy and reproducibility of the results²¹.

Combination index

The combination index (CI) was calculated using IC₅₀ values obtained from MTT cytotoxicity assays described in Sect. 2.2.1. The same concentration ranges and 48-hour treatment durations used in the MTT experiments were applied for CI analysis. CI values were computed using the standard equation (Eq. 1), where A CI value below 1 indicates synergy, while values equal to 1 indicate additivity, and values above 1 indicate antagonism²².

Data analysis

The dose-response relationships of the tested materials were evaluated using the E_max model, expressed by the formula:

Where R represents the residual unaffected fraction, indicating drug resistance; [D] is the applied drug concentration; K_d is the concentration at which 50% of the maximum inhibitory effect is achieved; and m is the Hill coefficient describing the steepness of the dose-response curve. The IC₅₀ value is defined as the drug concentration required to reduce cell absorbance to 50% of the control level, where K_d equals IC₅₀ when R is zero and the maximum effect (E_max) equals 100 minus R.

Apoptosis assay

The Annexin V-FITC/propidium iodide (PI) apoptosis detection kit was used to assess apoptosis and necrosis in treated cell populations. This kit, obtained from Abcam Inc. (Cambridge Science Park, Cambridge, UK), enabled the identification of apoptotic and necrotic cells via flow cytometry through dual fluorescent labeling. Following treatment, the cells were exposed to BV and TQ at concentrations of 9.96 µg/mL and 3.261 µg/mL, respectively. The same concentrations were applied for the combination treatment. After 48 h, the cells were collected by trypsinization. Approximately 1 × 10⁵ cells were harvested and washed twice with ice-cold phosphate-buffered saline (PBS) at physiological pH 7.4 to ensure the removal of any residual medium or compounds. After washing, the cells were resuspended in 0.5 mL of Annexin V-FITC/PI staining solution and incubated in the dark for 30 min at room temperature, following the manufacturer’s protocol. This incubation facilitates the binding of Annexin V-FITC to phosphatidylserine residues exposed on the surface of apoptotic cells, while propidium iodide (PI) stains the DNA of cells with compromised membrane integrity, allowing differentiation between apoptotic and necrotic cells.

Following the staining step, cells were analyzed using an ACEA Novocyte™ flow cytometer (ACEA Biosciences Inc., San Diego, CA, USA). The flow cytometer was configured to detect fluorescence signals in two channels: FL1 (488 nm excitation / 530 nm emission) for FITC-labeled Annexin V, and FL2 (535 nm excitation / 617 nm emission) for propidium iodide (PI). For each sample, data from 12,000 cellular events were acquired, and quadrant analysis was performed to distinguish populations of viable, early apoptotic, late apoptotic, and necrotic cells. Quantification and analysis of FITC- and/or PI-positive cells were performed using ACEA NovoExpress software (ACEA Biosciences Inc., San Diego, CA, USA), which enabled precise calculation of the proportions of apoptotic and necrotic cells in each sample. Experiments were conducted in triplicate to ensure accuracy and reproducibility of the results²³.

Cell cycle distribution assay

Cell cycle distribution analysis was performed using flow cytometry after treating cells with the test compounds at the following concentrations: BV, 9.96 µg/mL; and TQ, 3.261 µg/mL. The same concentrations were used for the combination treatment. After 48 h, roughly 1 × 10⁵ cells were harvested by trypsinization and detached from the culture surface. The cells were then rinsed twice with ice-cold phosphate-buffered saline (PBS) at pH 7.4 to remove any remaining medium or compounds. The collected cells were fixed by resuspending them in 2 mL of 60% ice-cold ethanol, then incubating at 4 °C for 1 h. This step preserves cellular structure for DNA analysis. After fixation, the cells were washed twice with PBS (pH 7.4) to remove any remaining ethanol. They were then resuspended in 1 mL of PBS containing 50 µg/mL RNase A and 10 µg/mL propidium iodide (PI). RNase A breaks down RNA, ensuring only DNA is stained, while PI binds to DNA, allowing for DNA content measurement.

After a 20-minute incubation in the dark at 37 °C to facilitate staining, the DNA content of the cells was analyzed using an ACEA Novocyte™ flow cytometer (ACEA Biosciences Inc., San Diego, CA, USA). Propidium iodide (PI) fluorescence was detected in the FL2 channel, with excitation and emission wavelengths of 535 nm and 617 nm, respectively. For each sample, data from 12,000 events were collected, allowing detailed analysis of the cell cycle phases. Cell cycle distribution was analyzed using ACEA Novo Express software (ACEA Biosciences Inc., San Diego, CA, USA), which quantitatively measures the proportion of cells in each phase of the cell cycle (G0/G1, S, and G2/M) based on DNA content. Experiments were performed in triplicate to ensure accuracy and reproducibility²⁴.

HPLC assay

Melittin, apamin, and phospholipase A2 in the BV sample were measured using high-performance liquid chromatography with ultraviolet detection (HPLC-UV). The separation was performed using an HPLC system (VWR International, Radnor, USA) equipped with a Supelco Supelcosil LC-318 column (4.6 × 250 mm, 5 μm; Sigma-Aldrich Co. LLC, Darmstadt, Germany). Absorbance was detected at 220 nm. The mobile phase consisted of two components: mobile phase A, high-purity water with 0.1% trifluoroacetic acid (TFA); and mobile phase B, a mixture of acetonitrile (ACN) and ultra-distilled water (80:20) with 0.1% TFA. A linear gradient program was applied, starting with 5% mobile phase B and progressively increasing to 80% over 24 min. The system was run at a flow rate of 1 mL/min, with an injection volume of 40 µL for each run. The analysis was conducted using the modified DIN 10,758 method. Standard stock solutions were prepared at a concentration of 1 mg/mL by dissolving phospholipase A2 and melittin in ultrapure water, while apamin was dissolved in ultrapure water containing 0.05 M acetic acid. These stock solutions were serially diluted to produce calibration standards at concentrations of 2%, 5%, 10%, 20%, 40%, and 50%. These standards were then used to construct calibration curves based on peak areas and heights.

For sample preparation, 5 mg of BV was accurately weighed and dissolved in 10 mL of ultrapure water. The solution was filtered through a syringe filter, transferred to vials, and injected into the HPLC system. Each sample was analyzed in triplicate to ensure consistency and accuracy. Quantification was performed using the external standard method by comparing the peak areas and heights of the BV components to those of the calibration standards, enabling precise determination of melittin, apamin, and phospholipase A2 concentrations in the samples. All experiments were conducted in triplicate to guarantee the accuracy and reproducibility of the results²⁵.

Statistical analysis

The data were analyzed for statistical significance using analysis of variance (ANOVA), followed by the LSD post hoc test. Results are expressed as mean ± RSD and were visualized using Prism^® version 5.00 (GraphPad Software Inc., La Jolla, CA, USA). Statistical tests were conducted using SPSS^® version 26, with a p-value of less than 0.05 considered the threshold for significance.

Results

MTT assay

The cytotoxic properties of TQ and BV were evaluated by determining the half-maximal inhibitory concentration (IC₅₀) values for BV, TQ, and their combination treatments across four different cancer cell lines: HCT, MCF-7, HSF, and HeLa. IC₅₀ values are presented in µg/mL for each treatment. The fixed TQ concentration, combined with BV treatment, was 6.568 µg/mL in MCF-7, HSF, and HCT cells, while in HeLa cells, it was 3.119 µg/mL. Conversely, the fixed BV concentration combined with TQ treatment was 19 µg/mL in MCF-7, HSF, and HCT cells, and 9 µg/mL in HeLa cells. The IC₅₀ values of BV alone were 16.29, 22.18, 20.21, and 9.96 µg/mL against HCT, MCF-7, HSF, and HeLa cells, respectively. The IC₅₀ values of TQ alone were 6.743, 5.587, 7.144, and 3.261 µg/mL for HCT, MCF-7, HSF, and HeLa cells, respectively. Notably, lower IC₅₀ values were observed for combination treatments. The combination of TQ with a fixed BV concentration yielded IC₅₀ values of 1.865, 2.73, 2.1, and 0.86 µg/mL against HCT, MCF-7, HSF, and HeLa cell lines, respectively. Additionally, the combination of BV with a fixed TQ concentration had IC₅₀ values of 5.7, 8.3, 6.4, and 4.55 µg/mL against the same cell lines. These data indicate increased cytotoxic activity of combination treatment compared to single-agent treatments. IC₅₀ values for each treatment and cell line are summarized in Fig. 1; complete concentration-viability curves are provided in the supplementary file (Supplementary Figs. 1–16).

Fig. 1.

The IC₅₀ values of BV, TQ, and their combination against HCT, MCF-7, HSF, and HeLa cell lines. Data represented as mean values, Error bars represent relative standard deviation (RSD%) from three independent experiments (n = 3).

Calculation of combination index

The MTT assay was used to determine the IC₅₀ values, which allowed for the calculation of the combination index (CI) using the previously described equation. Results illustrate the CI for a fixed concentration of TQ combined with serial dilutions of BV and vice versa across four cancer cell lines: HCT, MCF-7, HSF, and HeLa. CI values below 1 indicate synergistic effects between the two compounds in reducing cell viability. The observed trends across different cell lines demonstrate varying degrees of synergy depending on which compound was held constant (Table 1).

Table 1.

Summary of IC₅₀ values for single and combination treatments of TQ and bee venom BV across HCT, MCF-7, HSF, and HeLa, including combination index (CI) values and interpretation.

Cell- line	IC50 BV (µg/mL)	IC50 TQ (µg/mL)	IC50 (TQ + Fixed BV) (µg/mL)	IC50 (BV+ Fixed TQ) (µg/mL)	CI (TQ Fixed)	CI (BV Fixed)	Interpretation
HCT	16.29 ± 0.43	6.743 ± 0.13	1.865 ± 0.87	5.7 ± 0.125	0.94	0.83	synergism
MCF-7	22.18 ± 0.68	5.587 ± 0.83	2.73 ± 0.189	8.3 ± 0.26	0.93	0.86	synergism
HSF	20.21 ± 0.24	7.144 ± 0.29	2.1 ± 0.28	6.4 ± 0.11	0.91	0.84	synergism
HeLa	9.96 ± 0.19	3.261 ± 0.169	1.495 ± 0.198	4.55 ± 0.148	0.86	0.86	synergism

Phytochemical analysis using HPLC

Chromatographic analysis identified three main components in the BV sample: melittin, phospholipase A₂, and apamin (Fig. 2; Table 2). Melittin comprised approximately 82.57% ± 0.05 of the total peak area, while phospholipase A₂ and apamin accounted for 14.57% ± 0.31 and 2.87% ± 0.59, respectively. These results demonstrate high accuracy and consistency across measurements, confirming the typical composition of Apis mellifera venom.

Fig. 2.

HPLC chromatograms for the three consecutive injections show the reproducibility of retention times and peak profiles for the major components of BV. Peaks corresponding to Apamin, phospholipase, and melittin are consistently detected across injections.

Table 2.

HPLC detected phytochemicals and their relative retention time (RT) and area under the curve (AUC).

Component	Mean RT (min)	Mean AUC (mAU·s)	Mean % Area
Apamin	3.929 ± 0.127	248.577 ± 0.585	2.87 ± 0.59
Phospholipase A2	9.176 ± 0.082	1260.52 ± 0.35	14.57 ± 0.31
Melittin	12.96 ± 0.04	7167.18 ± 0.043	82.57 ± 0.05

Cell cycle distribution

The cell cycle distribution of HeLa cells was examined after 48 h of treatment with TQ, BV, and their combination. Data from three replicates were averaged, and ranges were determined for each cell cycle phase, including G1, S, G2/M, and Sub-G1 phases. These findings illustrate the effects of each treatment on cell proliferation and apoptosis, as well as their influence on cell cycle progression. Figure 3 shows a representative flow cytometric analysis of HeLa cells across the different treatment groups. Each panel features dot plots of forward scatter (FSC-A) and side scatter (SSC-A) parameters (left), along with histograms illustrating the distribution of cell cycle phases (G1, S, and G2/M) on the right. Refer to the supplementary data for complete flow cytometry plots from each experiment (Supplementary Figs. 17–20).

Fig. 3.

Results of representative flow cytometry for HeLa cells under different treatments. The related cell cycle histogram (right) and a forward scatter (FSC-A) versus side scatter (SSC-A) dot plot (left) are displayed in each panel. A: control; B: TQ; C: BV; and D: BV-TQ combination.

Flow cytometry analysis showed that untreated HeLa cells had a typical distribution, with most cells in the G1 phase (60.85%), followed by the S phase (21.16%) and the G2/M phase (20.10%), along with a low level of apoptosis indicated by Sub-G1 cells (3.87%). Treatment with TQ caused a decrease in the G1 (60.40%), S (16.65%), and G2/M (17.39%) phases, while the proportion of Sub-G1 cells increased to about 9.79%, indicating apoptosis. BV treatment resulted in a significant increase in cells in the Sub-G1 phase (76.80%), along with notable reductions in the G1 (14.90%), S (3.43%), and G2/M (7.54%) phases, demonstrating a strong apoptotic effect. When BV and TQ were combined, apoptosis was further increased, with Sub-G1 cells reaching 81.10%, and minimal presence in the G1 (12.55%), S (5.20%), and G2/M (3.52%) phases. Overall, these results illustrate that BV and TQ, especially when used in a combination, effectively promote apoptosis and disrupt the cell cycle in HeLa cells (Figs. 3 and 4).

Fig. 4.

Cell cycle phase distribution of HeLa cells under different treatments. Data represented as mean values, Error bars represent relative standard deviation (RSD%) from three independent experiments (n = 3).

These findings demonstrate that BV and TQ, both individually and in combination, significantly affect the cell cycle distribution of HeLa cells, primarily by triggering apoptosis. The combination treatment was the most potent, exhibiting the highest levels of apoptosis and the greatest suppression of cell cycle progression, particularly in the G1 and S phases. These results indicate a synergistic effect between TQ and BV, enhancing cell death and making the combination a promising therapeutic candidate for cancer treatment.

Apoptosis assay

Apoptosis was evaluated in HeLa cervical cancer cells after 48 h of treatment with TQ, BV, and their combination. Each trial showed a consistent distribution of cell populations under untreated conditions, establishing the baseline status of HeLa cells (Fig. 5).

Fig. 5.

HeLa cell apoptosis investigation using Annexin V-FITC/PI labeling (right) and forward scatter (FSC-A) vs. side scatter (SSC-A) for cell population gating (left) is depicted in representative flow cytometry dot plots. The treatments are as follows: A is the control; B is TQ; C is BV; and D is the BV-TQ combination. Each quadrant displays the percentage of living (Q3), early apoptotic (Q4), late apoptotic (Q2), and necrotic (Q1) cell populations.

In the control groups, cell populations remained predominantly viable, with mean survival rates of 96.28% (Control 1) and 95.29% (Control 2). The percentage of early apoptotic cells (Q4) was minimal, at 1.30% and 2.26%, respectively. Late apoptotic cells (Q2) accounted for 1.80% and 2.08%, while necrotic cells (Q1) comprised only 0.62% and 0.37%. These results demonstrate that untreated HeLa cells remained healthy, exhibiting minimal apoptotic activity.

In HeLa cells treated with TQ, the data indicated a moderate induction of apoptosis. The percentage of viable cells (Q3) averaged 95.01%, while early apoptotic cells (Q4) averaged 0.79%. Late apoptotic cells (Q2) represented 3.08%, indicating a notable increase in apoptosis. The necrotic cell population (Q1) was low, at 1.12%. These findings suggest that TQ has the potential to induce moderate apoptosis while maintaining a notable proportion of viable cells.

Treatment with BV caused a significant increase in apoptotic cells. The percentage of viable cells decreased to 77.66%, while early apoptotic cells (Q4) increased to 4.86%. Late apoptotic cells (Q2) averaged 16.71%, indicating that a substantial portion of the cell population was undergoing programmed cell death. The necrotic cell population (Q1) remained low, at approximately 0.77%. These findings suggest that bee venom induced a pro-apoptotic effect on HeLa cells, resulting in considerable cell death.

When TQ and bee venom were administered in combination, the results demonstrated the most pronounced apoptotic response. The average percentage of viable cells decreased to 61.08%, with early apoptotic cells (Q4) at 4.23%, and late apoptotic cells (Q2) significantly increasing to 32.52%. The necrotic population (Q1) was 2.18%, indicating a moderate level of necrosis. This combination treatment led to a pronounced increase in apoptotic cells, suggesting enhanced programmed cell death (Figs. 5 and 6). Refer to the supplementary data for complete results from all three experiments (Supplementary Figs. 21–24).

Fig. 6.

Apoptotic and necrotic cell events in HeLa cells under different treatments. Data represented as mean values, Error bars represent relative standard deviation (RSD%) from three independent experiments (n = 3).

Discussion

Recent research has increasingly focused on the potential of combination therapies that integrate natural compounds with conventional cancer treatments to enhance efficacy, minimize side effects, and overcome drug resistance. The strategy of using natural products in conjunction with conventional anticancer drugs has shown promising results, particularly because natural compounds often act through multiple mechanisms, targeting various pathways essential for cancer cell survival and proliferation²⁶. BV and TQ have been reported to exhibit diverse bioactivities, including anticancer effects^27,28. Given the limited data on their combined use, the present study aimed to evaluate the anticancer potential of TQ and BV, both individually and in combination, in selected cancer cell lines: HCT (colon cancer), HeLa (cervical cancer), and MCF-7 (breast cancer), as well as in a normal human skin fibroblast (HSF) cell line to asses selectivity and toxicity of each treatment. Cytotoxic effects, alterations in cell cycle distribution, and the ability to induce apoptosis were assessed using the MTT assay, flow cytometric cell cycle analysis, and Annexin V-FITC/propidium iodide (PI) apoptosis detection kit, respectively. Additionally, HPLC analysis was performed on BV to accurately quantify its major bioactive components, underscoring the potency and purity of the sample.

Quantitative analysis revealed that melittin constitutes approximately 82.57% of BV, with phospholipase A2 and apamin contributing approximately 14.57% and 2.87%, respectively (Table 2), consistent with previous studies^29,30. The high concentrations of melittin and other bioactive compounds are significant, suggesting a potential therapeutic benefit of BV in combination therapies targeting cancer cells. This elevated purity is attributed to the improved extraction method developed in this study, which was carefully designed to preserve sample integrity.

The MTT assay demonstrated dose-dependent reductions in cell viability for both agents across all cancer cell lines, with an apparent enhancement of efficacy observed in the combination treatment. For example, in HCT cells, the IC₅₀ of TQ combined with a fixed concentration of BV was (1.865 ± 0.87 µg/mL), substantially lower than the IC₅₀ of TQ alone (6.743 ± 0.13 µg/mL). This improved potency was also observed in HeLa and MCF-7 cell lines, where the combination treatment consistently produced lower IC₅₀ values, indicating a synergistic effect. This synergy was quantified by calculating the Combination Index (CI), with CI values less than 1 indicating a synergistic interaction. Specifically, the CI values for the fixed concentration of BV combined with serial dilutions of TQ were 0.94, 0.86, and 0.93 in HCT, HeLa, and MCF-7 cells, respectively. These CI values indicate that the combination of TQ and BV not only induces cytotoxicity but does so at lower doses of each agent, potentially reducing side effects while increasing therapeutic efficacy. The most pronounced synergistic effect was observed in HeLa cells, with a CI of 0.86, indicating a possible cooperative interaction between TQ and BV in promoting cell death (Table 1).

The cytotoxic potential of TQ and BV has been demonstrated in previous studies involving HeLa, MCF-7, and HCT cell lines^31–33. The enhanced antiproliferative effect of the TQ-BV combination may be attributed to their complementary and synergistic mechanisms of action.

TQ exerts its antiproliferative effects by inhibiting survival pathways. In cholangiocarcinoma models, both in vitro and in vivo, TQ reduced phosphorylated Akt, NF-κB activation, XIAP, Bcl-2, COX-2, and VEGF levels, leading to decreased proliferation and increased apoptosis³⁴. In breast cancer cell lines (MDA-MB-468 and T47-D), TQ induced G1-phase arrest and cell death. It suppressed the expression of cyclins D1 and E, survivin, Bcl-2/Bcl-xL, and key Akt signaling molecules, including PDK1, PTEN, GSK-3β, and Bad. Additionally, translation regulators such as 4E-BP1, eIF4E, S6 ribosomal protein, and p70S6K were also disrupted³⁵. Concurrently, melittin, the principal bioactive peptide in BV, inhibited cell growth and migration by targeting the PI3K/Akt/mTOR and MAPK pathways, which are critical for cell survival and proliferation³⁶. Melittin further increased endoplasmic reticulum (ER) stress and unfolded protein response (UPR) markers, including CHOP and spliced XBP-1 (XBP-1s), in HCT116 colorectal cancer cells. It also activated autophagic markers such as Beclin-1 and LC3-βII, while inhibiting NF-κB and death receptor 4/5 (DR4/DR5) signaling pathways³⁷.

The synergistic anticancer effect of TQ and BV likely results from complementary mechanisms involving oxidative stress and mitochondrial dysfunction. TQ enhances intracellular ROS through interference with the electron transport chain and suppression of antioxidant defenses, whereas melittin—the main BV component—induces ROS via membrane disruption and activation of redox-sensitive pathways³⁸. Their combined treatment amplifies ROS accumulation beyond cellular antioxidant capacity, leading to lipid, protein, and DNA damage, mitochondrial depolarization, and apoptosis.

Mitochondrial targeting is central to this synergy: TQ inhibits complexes I and III and disrupts ΔΨm, while melittin permeabilizes mitochondrial membranes, promoting cytochrome c, AIF, and Smac/DIABLO release³⁹. The resulting mitochondrial calcium overload intensifies apoptotic signaling, explaining the sub-G1 increase observed in co-treated HeLa cells.

Additionally, cross-talk between survival and apoptotic pathways reinforces this effect. TQ suppresses NF-κB activation by inhibiting IκB kinase, augmenting melittin-induced cytotoxicity. Both agents converge on the PI3K/Akt axis—TQ by blocking Akt phosphorylation and melittin by disrupting membrane signaling—thereby downregulating Bcl-2 and survivin and promoting Bax-mediated apoptosis⁴⁰. Collectively, these multi-targeted actions produce potent and irreversible apoptotic responses surpassing those of single-agent treatments.

In the current study, BV and TQ increased apoptotic activity and alterations in cell cycle progression, particularly cell cycle arrest in the G1 and S phases of the HeLa cell populations, suggesting enhanced apoptosis in cancer cells. The apoptosis assay in HeLa cells demonstrates a clear enhanced effect of combining BV and TQ, indicating an increased apoptotic response that exceeds that of either agent alone. Notably, the TQ-BV combination led to a significant increase in late-stage apoptosis, reflecting an intensified and irreversible commitment to cell death in these cancer cells, which is related to the complementary effects of both agents. This suggests a complex interplay between TQ and BV, likely involving both intrinsic and extrinsic apoptotic pathways.

TQ primarily induces apoptosis through partial caspase-independent mechanisms, including mitochondrial pathways (cytochrome c release and AIF translocation) as well as caspase-8 activation⁴¹. In HCT116 colon cancer cells, TQ decreases STAT3 and its downstream targets (survivin, c-Myc, cyclin D1/D2), upregulates p21 and p27, activates caspases-9, -7, and − 3, downregulates Bcl-2 and Bcl-xL, and promotes PARP cleavage. TQ also facilitates p53 binding to the CHEK1 promoter, reducing CHEK1 expression and thereby increasing cellular susceptibility to apoptosis and ROS-mediated DNA damage^42,43. Consistent with our findings, TQ interrupts the cell cycle at the G1/S phase in HCT116 colon cancer cells⁴⁰. Prior reports showed that TQ inhibits the PI3K/Akt/mTOR and NF-κB pathways while inducing mitochondrial-mediated apoptosis and G1/S arrest in colorectal cancer cells¹⁶. TQ also modulates survival signaling pathways such as PI3K/Akt and NF-κB, suggesting its ability to sensitize cancer cells to pro-apoptotic stimuli⁴⁴. In contrast to our results, it was demonstrated that TQ causes G2/M arrest and death in doxorubicin-resistant breast cancer cells by upregulating PTEN, decreasing Akt phosphorylation, and elevating p21, indicating that TQ may exert cell-type-specific effects on cell cycle regulation⁴⁵.

In colon cancer cells, BV increased the expression of death receptors DR4 and DR5, as well as p53, p21, Bax, and cleaved caspases⁴⁶. In PANC-1 and AsPC-1 pancreatic cancer cells, BV induced S-phase arrest, modulated cyclins and CDKs, activated the p53-p21 pathway, and promoted apoptosis⁴⁷. By inhibiting the NF-κB pathway, increasing Bax expression, and decreasing Bcl-2 levels, BV reduced cell viability and induced apoptosis in A549 cells⁴⁸.

At the same time, melittin promotes apoptosis by activating caspase-3 and downregulating Akt and Bcl-2, as cells are partially rescued by Bcl-2 overexpression or caspase inhibition⁴⁹. It also induces apoptosis and inhibits proliferation in non-small-cell lung cancer cells by phosphorylating ERK and downregulating TGF-β, which increases the levels of Apaf-1 and caspase-3⁵⁰. Recent work reveals that melittin can also generate endoplasmic reticulum (ER) stress and activate the unfolded protein response (UPR) and autophagy, processes that may collaborate with ROS-driven apoptosis to promote cancer cell death³⁷. It also reduces tumor cell survival via suppression of the PI3K/Akt/mTOR axis and has been demonstrated to promote apoptosis and inhibit proliferation in numerous cancer models⁵¹.

Additionally, BV melittin disrupts cell membranes, leading to mitochondrial destabilization and the release of cytochrome c, thereby enhancing apoptosis at the mitochondrial level⁵².

Our findings are further supported by previous studies demonstrating that melittin-rich BV induces G1 cell cycle arrest and promotes apoptosis in bronchogenic carcinoma (Chago-K1) and hepatocellular carcinoma (BEL-7402) cell lines^47,53.

The substantial presence of phospholipase A₂ at 14.57% further supports the venom’s bioactivity, as this enzyme plays a key role in damaging cancer cell membranes and enhancing the apoptotic potential of melittin⁵⁴. Apamin, a neurotoxic peptide found in BV, selectively inhibits SK-type Ca2+-activated K + channels. It acts by increasing intracellular calcium and apoptotic signaling, thereby improving anti-cancer activity and reducing systemic adverse effects, even at low concentrations^55,56. Through molecular docking studies, apamin has also been demonstrated to interact with tumor necrosis factor-α (TNF-α), indicating a possible function in regulating inflammatory and apoptotic signaling pathways associated with the advancement of cancer⁵⁷. Additionally, apamin has been demonstrated to significantly inhibit leukemia (K562) cell motility and invasion at picomolar doses, indicating that it has an impact on metastatic processes apart from direct cytotoxicity⁵⁸.

Together, these effects exert a dual influence on HeLa cells by promoting apoptosis and disrupting cell cycle progression. This disruption, particularly the arrest in the G1 and S phases, is critical because it prevents HeLa cell proliferation and may reduce the likelihood of tumor growth and metastasis. These results suggest that the hypothesis that the TQ-BV combination may provide an improved anticancer strategy than single-agent treatments by simultaneously inhibiting cell growth and inducing cell death.

Our study revealed that BV composition includes melittin at approximately 82.57% of the dry weight, apamin at 2.87%, and phospholipase A₂ at 14.57%. These findings closely align with previous research. For example, Tanuğur-Samancı and Kekecoğlu (2021) reported phospholipase A₂ ranging from 9.26% to 17.83%, apamin from 1.40% to 2.85%, and melittin content from 26.76% to 51.85%⁵⁹. Similarly, Carpena et al. (2020) found that melittin constitutes 50–60% of the dry weight of BV, while phospholipase A₂ accounts for 10–12%⁶⁰. These consistent results suggest that the composition of our sample is typical of BV. However, seasonal variations, bee species, and geographic origin can influence the concentrations of specific components. In conclusion, our findings support the general composition of BV as documented in the literature, with minor deviations likely attributable to biological and environmental factors.

The successful isolation of a highly concentrated melittin fraction demonstrates the effectiveness of the extraction method and highlights the therapeutic potential of this BV sample. The elevated levels of these components suggest that this advanced extraction approach could serve as a model for producing high-purity venom suitable for precision therapeutic applications, where maintaining bioactivity and minimizing contamination are crucial. These findings indicate the potential of combined TQ and BV treatment as a promising strategy to overcome the limitations of conventional therapies.

Study limitations

While this study demonstrates an apparent anticancer potential of the TQ-BV combination in vitro, several limitations should be acknowledged. First, all experiments were conducted using immortalized cancer cell lines, which may not fully recapitulate the complex tumor microenvironment, including immune cell interactions, stromal components, and three-dimensional architecture. Second, the lack of in vivo validation means that pharmacokinetic properties, bioavailability, tissue distribution, and potential systemic toxicity remain unexplored. Third, although our novel BV extraction method yielded high-purity samples, slight compositional variability may occur due to environmental factors such as temperature, seasonal variations, geographic location, and bee colony health. Fourth, we tested only a limited number of cancer cell lines and one normal cell line; broader screening across diverse cancer types and multiple normal cell lines would better establish therapeutic selectivity. Finally, the specific molecular targets and signaling cascades affected by the TQ-BV combination require further elucidation through transcriptomic, proteomic, and metabolomics analyses. Despite these limitations, this study establishes a solid foundation for future research and the development of natural compound-based anticancer therapies. Addressing these limitations through further studies is essential to fully understand the therapeutic potential of the TQ-BV combination and optimize its application in oncological treatment.

Future research directions

In vivo efficacy studies

• Xenograft tumor models using HeLa, MCF-7, and HCT cells in immunocompromised mice to assess tumor growth inhibition, survival rates, and optimal dosing regimens.

• Syngeneic tumor models to evaluate immune system interactions.

• Investigation of antimetastatic effects in metastasis-prone models.

• Comparison with standard chemotherapy agents (e.g., cisplatin, doxorubicin).

Toxicity and safety profiling

• Further studies using additional normal cell types (e.g., epithelial or liver cells) are recommended to assess toxicity and selectivity comprehensively.

• Acute and chronic toxicity studies to determine the maximum tolerated dose (MTD).

• Comprehensive organ toxicity assessment (liver, kidney, cardiac function).

• Hematological parameters and immune function monitoring.

• Investigation of potential allergic reactions to BV components.

• Comparison of the toxicity profile between combination and individual treatments.

Formulation development

• Nanoparticle-based delivery systems (liposomes, PLGA nanoparticles) to improve bioavailability and tumor targeting.

• Sustained-release formulations to maintain therapeutic concentrations.

• Topical or localized delivery systems for specific cancer types.

• Stability studies under various storage conditions.

• Scale-up considerations for clinical-grade BV extraction.

Conclusion

Despite these limitations, this study establishes a solid foundation for TQ-BV combination therapy as a promising natural product-based anticancer approach. The observed synergistic effects, as demonstrated by CI values < 1 across all tested cell lines, and the dramatic increase in apoptosis (81.10% Sub-G1 cells in HeLa), warrant further investigation.

Supplementary Information

Below is the link to the electronic supplementary material.

Author contributions

B.A. and H.T. wrote the main manuscript text and W.T. supervised and revised experimental part. M.A., H.A., M.H., P.A. reviewed and corrected the manuscript.

Funding

This article was funded by Applied Science Private University, Amman, Jordan (grant number DRGS-2024-1).

Data availability

The datasets used and/or analyzed during the current study available from the corresponding author on reasonable request.

Declarations

Competing interests

The authors declare no competing interests.