Molecular perspective on starfish tissue extracts: Targeting human carcinoma KB cells for anticancer therapy

2.1 Ethics declaration

In adherence to Indian ethics regulations, all animal experiments involving L. maculata were conducted, considering this species is neither endangered nor protected. The specimens were thoughtfully released into their natural habitat after completing the experimental procedures.

2.2 Animal regeneration experiments

Specimens of L. maculata (diameter 10–15 cm) were collected at a depth range of 10 to 15 m from Rameswaram Island in the Indian Ocean with the assistance of scuba divers (Fig. S1a-b). Starfish with no previous symptoms of regeneration were collected and kept at 15 °C in aquaria with seawater throughout the experimental duration. The starfish were fed small pieces of blue mussels (Mytilus edulis) twice a week. To induce regeneration, the starfish were anaesthetized with 4 % magnesium chloride in 50 % artificial seawater, and approximately 0.5 cm of the distal third of three arms for each specimen was amputated using a scalpel. The specimens were allowed to regenerate in the aquarium. The laboratory animal care committee of Aarupadai Medical College and Hospital, Puducherry, India, approved the experimental methods.

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Fig. 1

HPLC a) and GC–MS/MS b) chromatogram of the bioactive fraction of L. maculata.

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2.3 Preparation, purification, and fractionation of starfish extract

The extraction scheme for bioassay-guided fractionation from amputated L. maculata extracts using different solvents is shown in Fig. S2. Briefly, three-arm tips of each starfish specimen were amputated approximately 1.5 cm in length. The entire amputated starfish arm was macerated for 24 h at room temperature and refluxed in 6L of 70 % ethanol for 60 min (2.5 kg). Active fractions were pooled and condensed at 40 °C using a rotary evaporator, yielding 140 g of crude extract (LM-CE). The glassy brown suspension was dissolved in water and extracted three times with an equivalent amount of ether (petroleum) to obtain non-polar fraction compounds, such as lipids. The defatted water process was divided into ethyl acetate (EtOAc) and n-butanol (BuOH) fractions, yielding the ethyl acetate-derived portion (LM-EA-E, 49 g), the n-butanol fraction (LM-nB-E, 38 g), and the water-soluble fraction (LM-WS-E, 54 g). Furthermore, LM-nB-E was further fractionated using a chromatography column of Diaion HP-20 with H2O-MeOH solvent (100:0, 80:20, 70:30, 60:40, 40:60, 30:70, 20:80, 10:90, 0:100) to yield ten acetone fractions (LM-nB-E-Fr.1–10).The fraction eluted with H2O/MeOH (10:90) (LM-nB-E-Fr8) was further fractionated using a preparative C18 HPLC column with 80 % aqueous methanol and flow rates of 1 ml/min to obtain five active fractions. The optimization of this preparative C18 HPLC column separation involved adjustments in solvent composition, flow rate, and gradient conditions. The goal was to efficiently elute five active fractions representing distinct bioactive compounds for subsequent analysis and experimental investigations. Active fractions representing each peak were eluted and pooled together from HPLC (LM-nB-E-Fr8.1–5, 1.35 g) as a white amorphous powder (hereafter referred to as the bioactive fraction), which was analyzed for GC–MS/MS study. The isolated fractions were dissolved at 100 mg/ml concentrations in 10 % dimethyl sulfoxide (DMSO) for further experimental investigations.

2.4 Identification of the purified compounds

The active peak fraction collected from the HPLC was pooled and further analyzed to identify pharmacologically active constituents using the A7890 GC fuel chromatography MS (GC–MS) system. A quadrupole MS 5975C detector (30 m long, internal 0.25-mm diameter, 0.25 μm thickness; Agilent technology, Germany) with helium as a carrier gas was used to quantify the detected compounds. The injector size and temperature were 2 μl and 250 °C, respectively. The samples were injected into the column after 1 min at 50 °C and then heated at a rate of 25℃/min to 170℃ and 5℃/min to 280℃, respectively. The spectra were scanned in the 33–600 m/z range. The MS was run in electron ionization mode, and the collected mass spectra were compared to the MS software used in the NIST library databases.

2.5 In vitro screening of crude extracts and active fractions

Human oral epidermoid carcinoma KB cells were cultivated in RPMI1640 medium supplemented with 10 % fetal bovine serum and 1 % glutamine and incubated at 37 °C with 5 % CO². The KB cells were seeded at a density of 1 × 10⁶ cells per well in 96-well plates and incubated for 24 h. Cells were treated with crude extracts and active fractions at different concentrations of L. maculata tissue extracts, ranging from 0.1 μg/ml to 10 μg/ml for 24 h, and DMSO was used as a control. After achieving 90 % confluence, attached cells were washed twice with phosphate buffer solution, and 20 µl of MTT solution was added to each well. After overnight incubation at 37 °C, the MTT solution was removed, and DMSO was added to dissolve the formazan crystals produced by viable cells. Plates were agitated for 10 min, and absorbance was measured at 570 nm using a microplate reader. Cell viability was determined by comparing the absorbance of samples treated with cells to the absorbance of control cells. The IC⁵⁰ value was calculated using linear regression analysis and repeated in triplicate for each concentration.

2.6 Cell migration assay

An in vitro cell wound healing assay was performed to study the migratory behavior of KB cells. Confluent KB cell monolayers (approximately 90 %) were manually wounded by scratching the cells with a sterile P200 pipette tip. Following the manual scratching, there was an incubation period of 24 h to allow the cells to respond and migrate. The wounded cells were cultured in a new medium containing L. maculata active fractions (1 µg/ml), crude, and controlled after washing the cellular debris twice with PBS. Healing of wounds was captured using an Olympus optical camera with a light microscope, and direct measurements of repaired regions were documented to measure cell migration (Mariadoss et al., 2021).

2.7 DPPH radical scavenging assay

The extract scavenging activity was assessed using the DPPH assay (Zhu et al., 2011). 1.5 ml of DPPH solution (0.1 mM methanolic solution) was mixed with 1 ml of the sample at various concentrations. The reaction mixture was stirred for 30 min in the dark and then absorbed by a spectrophotometer at 517 nm. A control group with 0.1 ml (10 mg/ml) of ascorbic acid was used. The DPPH radical scavenging percentage was calculated using the following formula: $$\left(\%Inhibition\right)=\left(\mathit{Absorbance\; of\; control}-Absorbanceofsample\right)\times 100/\left(\mathit{Absorbance\; of\; control}\right)$$

2.8 ABTS⁺ radical scavenging activity

The radical scavenging activity of ABTS + was measured according to the procedures (Ilyasov et al., 2020). The ABTS⁺ solution was prepared and stored at room temperature for 16 h. Then, 1 ml of the ABTS + solution was mixed with 30 ml of deionized water to obtain a workable solution of 0.02 ± 0.70 at 734 nm. To 150 µl of the work solution, 1 ml of samples at various levels was added, and the reaction mixture was kept in a dark room for about 10 min before measuring absorbance at 734 nm. A standard control with 0.1 ml of ascorbic acid (10 mg/ml) was used. The ABTS scavenging activity was calculated using the following formula:

(% Inhibition) = (Absorbance of control - Absorbance of sample) × 100 / (Absorbance of control).

2.9 Superoxide radical scavenging activity

The superoxide radical scavenging activity of samples was determined according to the procedures (Zhu et al., 2006). The reaction mixture consisted of 150 µM nitro tetrazolium, 1 ml of aliquots (prepared in 0.1 M Tris HCL buffer, pH 7.4 of 60 µM phenazine methosulphate), and 468 µM nicotinamide adenine dinucleotide added to 1 ml of the sample at varying concentrations. The reaction mixture was then incubated for 5 min at 25 °C, and absorbance was measured at 560 nm. A standard control with 0.1 ml of ascorbic acid (10 mg/ml) was used. The superoxide radical scavenging activity was calculated using the following formula: $$\left(\%Inhibition\right)=\left(\mathit{Absorbance\; of\; control}-Absorbanceofsample\right)\times 100/\left(\mathit{Absorbance\; of\; control}\right)$$

2.10 Hydrogen peroxide radical scavenging activity

The hydrogen peroxide scavenging activity of the samples was determined according to the method of (Gülçin et al., 2004). A 40 mM hydrogen peroxide solution was prepared in PBS, and 1 ml of extracts was mixed gently with 2 ml of 40 mM hydrogen peroxide solution. The sample absorbance reaction mixtures were measured at 230 nm, this specific wavelength, chosen for its characteristic absorption peak related to hydrogen peroxide, enables us to assess the ability of the sample to neutralize this reactive oxygen species (ROS). Hydrogen peroxide is an important ROS involved in oxidative stress and cellular damage, rendering it a relevant target for antioxidant assessment. A standard control with 0.1 ml ascorbic acid (10 mg/ml) was used. The hydrogen peroxide scavenging activity was calculated using the following formula: $$\left(\%Inhibition\right)=\left(\mathit{Absorbance\; of\; control}-Absorbanceofsample\right)\times 100/\left(\mathit{Absorbance\; of\; control}\right)$$

2.11 HPLC: Amino acid composition analysis

The active fraction's amino acid composition was determined by the HPLC method (Merck Hitachi Lachrome D-7000 HPLC System) using a Shimadzu C-18 column connected with two solvent systems: (a) 0.1 % TFA solutions and (b) 0.1 % TFA in 90 % acetonitrile (Baker and Han, 1994). The samples were hydrolyzed in 6 M HCL for 24 h in vacuum-sealed ampoules at 110 °C, and then hydrolysates were analyzed on a Hitachi liquid chromatography system at room temperature. The HPLC system was used to identify and quantify amino acids by comparing retention time and peak areas to those of a standard amino acid composition (Sigma). The absorbance of HPLC column elutes was measured at 280 nm.

2.12 GC–MS: Fatty acid profile analysis

The fatty acid compositions of the active fraction extract were determined by GC–MS using a Varian Saturn 2000R gas chromatograph (Hewlett Packard 5890 model) programmed from 50 °C to 225 °C (40 °C/min), then kept constant for 30 min. FAMEs were categorized based on their electron effects MS spectra and retention time (Rt), which were comparable to a standard (Sigma), and quantified based on relative peak areas (Bligh and Dyer, 1959).

2.13 FT-IR (Fourier transformed infrared) spectroscopy

The FT-IR spectrometer (Thermo Nicolet, USA) was used to quantify the functional properties of the samples (Mariadoss et al., 2019a). Diffuse reflectance was applied in the mid-IR spectral region (400–4000 cm^-1). The extracted sample (2 mg) was compressed into a salt disc using a compression gauge (10 mm diameter) by combining it with 100 mg of potassium drying bromide powder (KBr). ORIGIN 8.0 software was used to investigate the peak absorption of light intensity.

2.14 Measurement of intracellular ROS levels in cells

The oxidation level of 2,7-diacetyl dichlorofluorescein (DCFH-DH; Sigma-Aldrich) was measured to determine the intracellular reactive oxygen (ROS) concentration, as described by (Sivaraj et al., 2018). The cells were seeded at a density of 5 × 10⁴ cells/well into dark 96-well tissue culture plates and treated for 3 h with 1 µg/ml of crude and active fractions. Then, the cells were stained for 30 min at 37 °C in the dark with 10 M DCFH-DA and rinsed with PBS thrice. Images were photographed using a fluorescence microscope (Nikon, Japan) at a magnification of 400. Using a microplate reader, Fluorescence was measured at excitation and emission wavelengths of 485 and 335 nm, respectively.

2.15 Morphological evaluation of apoptotic cells by acridine orange and ethidium bromide (AO/EtBr)

The AO/EtBr double staining test examined the apoptosis-related morphological alterations(Qingming et al., 2010). In a 24-well plate, 2 × 10⁵ cells were plated in each well and incubated for 24 h before being treated with crude and active fractions for another 24 h at 37 °C. The cells were washed twice with 1 × PBS and stained with 5 µl of AO/EB. Finally, fluorescent microscopy (Olympus) examined the cell morphology at 20 × magnification. Apoptotic cells were distinguished by morphological characteristics such as constricted cell bodies, condensed, consistently delimited, heavily coloured chromatin, and membrane-bound apoptotic bodies.

2.16 DNA damage analysis

DNA fragmentation, indicative of apoptosis, was assessed through alkaline single-cell gel electrophoresis, commonly called the comet assay, as described by (Gunaseelan et al., 2017). In this assay, cells were embedded in agarose gel on microscope slides, electrophoresed under alkaline conditions, and stained with a fluorescent DNA-binding dye. Subsequently, comet photographs were captured using an epifluorescence microscope (Nikon, Japan). The movement of the DNA tail and its length damage were calculated using CASP software. The images were used to measure individual nuclei's DNA content and calculate the percentage of DNA loss in the comet tail.

2.17 Western blot analysis

The cells (1x10⁶ cells/well) were treated with 1 µg/mL of control, crude, and active fractions. Samples were washed with PBS lysis buffer and centrifuged at 12,000 × g for 15 min. The NanoDrop was used to measure the protein concentrations of the samples. Aliquots of the lysates (30 µg of protein) were boiled for 5 min at 95 °C and transferred onto nitrocellulose membranes for blotting. Primary antibodies Bax (B3428) (Sigma), Bcl-2 (B9804) (Sigma), Caspase-3 (C5737) (Sigma), PCNA (SAB4100556) (Sigma), Cyclin-D1 (C7464) (Sigma), and β-actin (A2228) (Sigma) were used to probe the membranes for 1 h in TTBS. After that, the membranes were rinsed three times with TTBS at 10-minute intervals before being incubated with secondary antibodies immunoglobulin-G-horseradish peroxidase for two hours at 37 °C. Chemiluminescence was used to detect the protein bands, which were then visualized using the luminescent image Studio device (LI-COR).

2.18 Statistical analysis

Statistical analysis of the results was performed using Version 5.0 of GraphPad Prism. Samples were collected in triplicate for each experiment, and one-way analysis of variance (ANOVA) was used to assess the variations between the control and active fractions. ANOVA analysis was used to compare the data from various samplings to evaluate the variations between the control and fraction.

3.1 HPLC evaluation of L. maculata active fractions

The HPLC chromatograms of L. maculata are displayed in Fig. 1a. The partially purified fraction observed five peaks with retention times of 2.50, 4.36, 5.74, 7.02, and 13.17 min. The presence of the active compound in the fractions was indicated by the most prominent and broadest peak at 4.36 min, while peaks at 5.74 and 7.02 min suggested the presence of other biologically active compounds. These retention times serve as key indicators of the compounds presence and are vital for the identification and characterization of bioactive components in L. maculata tissue extracts.

3.2 Analysis of GC–MS/MS

To confirm the presence of the compounds in the bioactive fraction, we performed an analysis, as shown in Fig. 1B and detailed in Table S1. In this analysis, we identified compounds such as 5-cholest-7-en-3-ol, hexadecanoic acid, myo-inositol, 9,12,15-Octadecatrienoic acid, and 9,12-Octadecadenoic acid within the active fraction. Importantly, these compounds have demonstrated anticancer activity in various cell lines, including KB. While our study primarily focused on their identification and biological activities, the specific biosynthetic pathways responsible for these compounds within L. maculata were not investigated in detail in this study. We further verified these bioactive compounds' identities and biological activities using Dr. Dukes' phytochemical and elucidation techniques (Duke and Bogenschutz, 1994). This comprehensive approach confirmed their presence and established their specific chemical identity and ability to exhibit anticancer effects on KB cell lines. Our combined analytical and biological validation ensures the reliability of our findings and supports the bioactive nature of these compounds in the active fraction of L. maculata extract.

3.3 Cell proliferation and anti-migration effects induced by L. maculata bioactive fraction

The effect of the L. maculata bioactive fraction on cell growth and proliferation was assessed using the MTT assay. Fig. 2a illustrates a significant inhibition of KB cell proliferation by the active fraction after a 24-hour incubation period compared to the control and crude groups. Concentrations of 25, 50, or 100 µg/ml of the extracts did not significantly affect L. maculata. A dose-dependent response was observed with the active fractions effectively inhibiting KB cell growth at concentrations of 0.3, 0.78, 1.56, 3.12, and 6.25 µg/ml. The IC50 value of cell growth was determined to be 0.62 µg/ml, indicating a potent anti-proliferative effect of the active fraction. Moreover, treatments with active fractions at 6.25 µg/ml and 12.5 µg/ml demonstrated approximately 80 to 81 % inhibition of cell proliferation, respectively.

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Fig. 2

The cytotoxicity assay shows the impact of L. maculata active fraction on KB cells (a). The wound healing test was conducted on KB cells treated with 1 µg/ml control, crud, and active fractions for 24 h (b). T-Scratch software was used to capture and analyze wound edge photographs of cells treated with individual triplicates 24 h after treatment, allowing calculation of the percentage of migration.

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Apoptotic morphological changes were also observed, including nuclear chromatin attention, edge accumulation, cytoplasmic hypervacuolization, cellular shrinkage, and apoptotic bodies. Based on the MTT effects, working concentrations for further experiments were determined to be 0.3, 0.78, 1.56, and 3.12 µg/ml for active fractions. Furthermore, the potential impact of active fractions and crude extracts on KB cell migration was observed (Fig. 2b). The MTT assay confirmed that cells treated with active fractions migrated slower than untreated and crude extract-treated cells. Morphologically, the KB cells treated with the active fraction exhibited reduced adhesion, resulting in cell shrinkage and the appearance of apoptotic bodies. They eventually merged into massive cellular aggregates. In contrast, no distinct phenotypes or directional migrations were observed in cells treated with untreated and crude extracts. These results indicate that active fractions of L. maculata are crucial in regulating cell proliferation and behavior.

3.4 Evaluation of antioxidant activities by L. maculata bioactive fraction

The scavenging of DPPH radicals by the extracts significantly increased DPPH concentrations (Fig. 3a). The most substantial DPPH radical scavenging activities were observed in the active fractions, control, and crude extracts, indicating a higher concentration of potent antioxidants than the crude and control samples. Similarly, the ABTS⁺ radical scavenging activity aligned with the DPPH results, with the active fraction exhibiting the highest ABTS⁺ radical activity, followed by the dose-dependent control and crude activities (Fig. 3b). The study also evaluated the radical scavenging activity of the three different extracts in KB cells, considering that most cancer cell lines produce superoxide radicals, leading to oxidative stress and cellular damage (Fig. 3c). The active fraction displayed the highest scavenging activity of 80 % at 30 µg/ml and the lowest activity of 20 % at 5 µg/ml, demonstrating superior superoxide (O₂) radical scavenging activities compared to the control and crude extracts.

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Fig. 3

The antioxidant activities were assessed through DPPH radical scavenging (a), ABTS radical scavenging (b), superoxide radical scavenging (c), and hydroxyl radical scavenging activity (d). The results are presented as the mean ± standard deviation of six experiments from each sample, with values significantly different (P < 0.05) from the control values.

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Finally, the study assessed the hydroxyl radical scavenging activity, crucial for preventing cellular damage caused by reactive hydroxyl radicals (Fig. 3d). The crude extract exhibited the highest hydroxyl radical scavenging activity, followed by the active fraction and the control. The findings regarding antioxidant activities highlight the robust scavenging potential of the active fraction of L. maculata against various free radicals. This potent antioxidant capacity contributes to its potential therapeutic efficacy as an efficient antioxidant agent.

3.5 Amino acid composition analysis of L. maculata active fraction

The amino acid composition of the active fraction was determined by conducting an HPLC analysis, comparing it with 21 amino acid standards (Fig. 4a). The active fraction exhibited enrichment in essential amino acids (EAAs) such as Histidine, followed by Phenylalanine, Methionine, Valine, Lysine, Tryptophan, Isoleucine, and Leucine. Conversely, non-essential amino acids (NEAAs) like asparagine, Alanine, Glycine, Aspartic acid, Glutamine, Glutamic acid, Proline, Tyrosine, and Serine were present in lower quantities in the active fraction (Fig. 4b, Table 1). HPLC analysis facilitated identifying and quantifying specific amino acids in the active fraction of L. maculata. The higher proportions of essential amino acids imply that the active fraction holds significant nutritional and therapeutic potential, warranting further scientific investigation.

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Fig. 4

Standard Amino Acid Graph (a). Amino Acid Content of the Active Fraction (b), Gas Chromatography FAME: Profiling Fatty Acid Composition (c). Fatty Acid Content of the Active Fraction: The gas chromatography analysis identified fatty acid components in L. maculata active fraction, showing richness in PUFAs and significant SFAs (d). The Characteristic peaks corresponding to various functional groups were observed in the FT-IR Analysis of L. maculata Active Fraction, suggesting the presence of diverse compounds, including hydrophobic components, in the active fraction (e).

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3.6 Fatty acid composition analysis of L. maculata active fraction

The fatty acid composition of the active fraction was determined using gas chromatography FAME. Polyunsaturated fatty acids (PUFAs) were more abundant in the active fraction than saturated fatty acids. The primary components identified among the PUFAs were α-Linoleic acid C18:3 (545.6 mg) and Linoleic acid C18:2 (306.2 mg). Additionally, Oleic acid C18:1 (112.6 mg) was identified as the dominant monounsaturated fatty acid (MUFA) based on the standard's Rf value (Fig. 4c-d and Table 2). In addition to PUFAs and MUFAs, significant amounts of saturated fatty acids (SFAs) were present in the active fraction. Notably, Stearic acid C18:0 (325.70 mg), Margaric acid C17:0 (103.3 mg), and Palmitic acid C16:0 (34.7 mg) were identified as the most important components of SFAs. The predominance of PUFAs and MUFAs in the active fraction suggests potential health benefits and functional properties, warranting further research in nutrition and health sciences.

3.7 FTIR characterization of the active fraction

The active fraction was subjected to FTIR evaluation to characterize its functional groups (Fig. 4e and Table 3) and compare them with standard FTIR data (Krimm and Bandekar, 1986). Free anti-symmetry O–H hydrogen-bonded stretching frequencies were indicated by a strong and sharp peak between 3738.64 cm⁻¹ and 3938.64 cm⁻¹. Moreover, amides and primary amine groups with N–H stretching frequency at 3369.64 cm⁻¹ and nitrile compounds with N-O asymmetric deformation at 2358.94 cm⁻¹ were identified in the active fraction. Additionally, C–H stretching vibrations of free sugars, a saturated aliphatic CO₂ stretching band between 1828.52 cm⁻¹ and 1905.67 cm⁻¹, and an aliphatic CH₂ stretching band at 1454 cm⁻¹ with aliphatic-CH bending frequency were observed in the active fraction, pointing to the presence of hydrophobic compounds. Furthermore, the observation of alkene groups at 1653 cm⁻¹ indicated the existence of low C–C ring stretching frequency. The presence of hydrophobic compounds in the active fraction was further supported by the fingerprint range of carbohydrates, which was positioned between 665 and 1220 cm⁻¹(Al-Assaf et al., 2007).

3.8 Intracellular ROS induction by L. maculata active fraction

Intracellular reactive oxygen species (ROS) accumulation is a hallmark of the apoptotic process. Following the treatment of KB cells with control, crude, and active fractions, a significant elevation in ROS levels was observed, with the active fraction demonstrating the most substantial ROS generation (95 %), as depicted in Fig. 5a-b. These findings suggest that the induction of apoptosis may be attributed to the generation of ROS by the active materials present in L. maculata.

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Fig. 5

Intracellular ROS levels were assessed using DCFH-DA staining fluorescence microscopy in KB cells treated with 1 µg/ml L. maculata active fraction, revealing a pronounced increase in DCFH fluorescence, indicating elevated intracellular ROS (a). Intracellular ROS was quantified using spectrofluorometer analysis, confirming a significant rise in ROS levels upon treatment with L. maculata active fraction (b). Apoptosis induction in KB cells was assessed using AO/EB staining and visualization under a fluorescence microscope (c). The active fraction-treated cells displayed distinct morphological changes and the presence of apoptotic bodies, indicating the initiation of apoptosis (20x magnification).

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3.9 Apoptotic morphological changes in cancer KB cells induced by the active fraction of L. maculata

To elaborate on the mechanisms by which L. maculata extracts induce apoptosis in KB carcinoma cells, our study employed multiple approaches. First, we assessed apoptotic morphological changes using acridine orange/ethidium bromide (AO/EB) double staining. Additionally, intracellular ROS accumulation was monitored (Fig. 5c). When not treated with the active fraction, the cancer KB cells displayed red/orange (EtBr stained) and green (AO stained) fluorescence. In contrast, when exposed to the active fraction, the cells exhibited typical apoptotic features, including cellular shrinkage, chromatin condensation, nuclear fragmentation, and the formation of apoptotic bodies. Remarkably, even at a concentration as low as 1 μg/ml, the active fraction induced apoptosis in 80 percent of the cells, showcasing its superior efficacy compared to other extracts. These observations strongly support the mechanisms by which L. maculata extract induces apoptosis in KB carcinoma cells highlighting the potential of the active fraction as a potent agent in combating cancer cells.

3.10 Induction of DNA damage by the active fraction of L. maculata: A comet assay analysis in cancer KB cells

The presence of intra-nucleosome DNA fragmentation indicated apoptosis. To quantify DNA damage, the Comet assay was performed. The cells treated with the active fraction showed a substantial increase in DNA fragmentation, as evidenced by highly fragmented DNA compared to the large, non-fragmented DNA observed in the control group (Fig. 6a). Virtual images were analyzed using the CASP software to calculate various endpoints, with a particular emphasis on the DNA percentage in the comet tail, a commonly used parameter for evaluating DNA damage in the Comet assay (Fig. 6b). Remarkably, treatment with the active fraction at a concentration of 1 µg/ml consistently resulted in a significant increase in DNA percentage in the tail (35 %) and an elevation in tail movement (17 %) in KB cells.

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Fig. 6

Oxidative DNA damage was assessed using the comet assay, and fluorescence microscopy images of DNA fragmentation were captured in various treatment groups (a). The percentage of tail DNA, with standard deviation values from six experiments for each group was quantified as part of the comet parameter analysis (b).

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3.11 Impact of L. maculata active fraction on cell cycle-related and apoptosis-related protein expressions in KB cells

The precise mechanisms by which the bioactive fraction inhibits cell growth and promotes apoptosis in KB cells were investigated in our study. Firstly, we examined cell cycle-related protein expressions, including PCNA and Cyclin D1, during cell damage. The results in Fig. 7a-b demonstrated that the active fraction significantly inhibited cell proliferation by downregulating PCNA and Cyclin D1 expression levels. This indicates that the active fraction induces cell cycle arrest, preventing the uncontrolled growth of KB cells. Additionally, we investigated apoptosis-related proteins, specifically Bax, Bcl-2, and Caspase 3, to understand the mechanisms of cell death induced by the active fraction. As shown in Fig. 7c-d, the pro-apoptotic protein Bax was significantly upregulated in KB cells treated with the active fraction, while the anti-apoptotic protein Bcl-2 was markedly downregulated. Moreover, the active fraction increased caspase-3 levels in KB cells. These findings collectively suggest that the L. maculata active fraction promotes apoptosis by regulating the expression of Bcl-2 family proteins and activating caspase-3, leading to cellular damage and reduced cellular proliferation. Our study reveals that the active fraction of L. maculata exerts its effects on KB cells by influencing both cell cycle progression and apoptosis pathways, contributing to its anticancer properties. These mechanisms provide insights into how the bioactive fraction inhibits cell growth and encourages apoptosis in KB cells.

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Fig. 7

Western blot analysis of PCNA and cyclin D1 (a-b) and Bcl-2, Bax, and caspase-3 (c-d) expression in control, crude, and active fraction-treated KB cells. The lysates of KB cells were subjected to immunoblotting, revealing elevated levels of Bax and caspase-3 and decreased expression of Bcl-2 in the active fraction-treated cells, suggesting the induction of apoptosis.

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