Shade dried A. nervosa leaf powder was extracted using water, ethyl acetate, and ethanol. The extracts were studied for immunomodulatory activity. The promising ethyl acetate extract was further analyzed by GC-MS for its components, where GIV was reported to be one of its major constituents. GIV used for this study was procured from Sigma-Aldrich (109-20-6), and various concentrations ranging from 500 µm to 5 mM were used to understand its antiproliferative activity and mechanism of action.

Cytotoxicity of GIV was checked against the HCT116 and HT29 cells using the MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay. Cells were harvested and seeded in a 96-well culture plate with a density of 1 × 10⁴ cells per well and incubated for 24 h. The cells were treated with different concentrations of GIV ranging from 0.5 to 8 mM for 48 h. After 48 h of treatment, cells were incubated with 50 µg of MTT (5 mg/ml, HIMEDIA, REF-TC191-1G) for 2 h. The formazan crystals formed were solubilized with 100 µl of dimethyl sufoxide (Fisher Scientific Prod. No. 12435), and the absorbance was recorded at 595 nm (Senthilraja and Kathiresan, 2015). The percent viability of the cells was calculated with respect to the untreated control cells. The standard procedure was used to calculate the IC₅₀ of the cells.

Additionally, to corroborate the MTT experiment results, 6 × 10⁴ cells/well of the HCT116 and HT29 cell lines were harvested and seeded in a six-well cell culture plate for 24 h. The cells were then treated with different concentrations of GIV from 0.5 to 5 mM. After the treatment of cells for the respective incubation period, the cells were stained using propidium iodide (PI, HIMEDIA, REF ML067) at a concentration of 10 µg/µl and incubated for 15–20 min. The cells were washed with phosphate-buffered saline (PBS) and then imaged at ×20 magnification using a Leica fluorescence microscope (Vyas et al., 2021).

To understand the mechanism of cell death, changes in mitochondrial membrane potential (MMP) were studied using JC-1 staining (Invitrogen, REF M34152) of GIV-treated cells. Cells treated with GIV for 24 and 48 h were stained with JC1 at a final concentration of 2 µM and incubated for 15–20 min. Cells were washed with PBS, and the image was taken by using a Leica fluorescence microscope at ×20 magnification. The stained cells were also used for flow cytometric analysis. Carbonyl cyanide m-chlorophenyl hydrazine (CCCP, Invitrogen, REF M34152) was used as a positive control to check the MMP loss during apoptosis (Sivandzade et al., 2019).

Cellular reactive oxygen species (ROS) generation was evaluated by dichlorofluorescein (DCFH-DA) dye (G-Biosciences Cat. #RC1066). Cells treated with GIV for 48 h were stained with dichlorofluorescein dye to a final concentration of 1 µM and incubated for 20 min. The cells were then washed with PBS and imaged at ×20 magnification using Leica fluorescence microscope (Wu and Yotnda, 2011).

The HCT116 cells were seeded in a T25 flask and treated with GIV. RNA was isolated from the cells using total RNA isolation kit (Promega, REF Z6111). RNA was quantified using Nanodrop, and 1 µg of the total RNA was used to synthesize cDNA using the cDNA synthesis kit (Promega, REF A3800). The cDNA obtained was used as a template to check the expression of apoptosis-associated genes, such as Caspase3, Caspase9, Bcl2, and Parp (primers were purchased from Imperialls, ILS) using SYBR Green Master Mix (Promega, REF A6001). 18s rRNA was used as an internal control. The Ct values attained through q-PCR were used to analyze the expression of the genes (Jozefczuk and Adjaye, 2011).

All the data are the representations of mean ± SD of at least three different experiments. Student' t-test was performed to derive p values, and p < 0.05 was considered as statistically significant.

Cytotoxicity of GIV was evaluated against two different CRC cell lines (HCT116 and HT29) using the MTT assay (Figure 1 and Supplementary Figure S2), which is a calorimetric assay to calculate the metabolic activity of the cell (Stockert et al., 2018). NADPH-dependent oxidoreductase enzyme in a healthy cell has the capability to reduce the tetrazolium dye—MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide)—to the insoluble purple dye called formazan (Senthilraja and Kathiresan, 2015). The higher the amount of formazan formed, the more viable the cells become. The IC₅₀ value of GIV was found to be around 1.8 mM for HCT116 cells and approximately 4 mM for HT29 cells.

Further to confirm the cytotoxicity observed by the MTT assay, PI staining was performed for which images were captured under fluorescence microscope. PI is a membrane impermeant red fluorescent DNA intercalating dye. This dye can cross the membranes only when the membranes are compromised (Lukowiak et al., 2001). It is used to differentiate live/dead cells on the basis of membrane integrity. The loss of membrane integrity is one of the hallmark features of cell death. PI when bound to DNA excites at 493 nm and emits a fluorescent red light of 636 nm (Cummings and Schnellmann, 2004; Lukowiak et al., 2001). Cells treated with GIV exhibited an increase in red fluorescence in comparison to the untreated control cells. As shown in Figure 2 and Supplementary Figure S3, this increase in red fluorescence was observed to be concentration dependent. All these results of PI microscopy supported the results of cytotoxicity observed by using the MTT assay.

To further confirm the process of cell death and contribution of the mitochondria in cell death, JC-1 staining was performed. The loss of MMP is one of the hallmark features of apoptosis. To evaluate the loss of MMP, JC-1—a lipophilic cationic dye—was used. In healthy cells, this dye gets accumulated in the negatively charged mitochondria, thus forming J-aggregates that emit red fluorescence. However, in unhealthy apoptotic cells, there is less negative charge inside the mitochondria due to membrane potential loss and therefore JC-1 dye enters the mitochondria but is unable to form stable J-aggregates. It exists in a monomeric form which emits green fluorescence (Sivandzade et al., 2019). For JC-1 stain imaging by fluorescence microscopy, TRITC and FITC channels were used, and both the images were overlayed to examine the concentration of J-aggregates verses J-monomers. As shown in Figure 3 and Supplementary Figure S4, cells treated with GIV clearly showed an increase in J-monomers, with increasing GIV concentration showing MMP loss. This indicates that GIV induces apoptosis through the mitochondria-mediated pathway. It has been reported that chemotherapeutic drugs are shown to induce MMP loss, thereby inducing apoptosis (Vyas et al., 2021).

This depolarization of the mitochondria and the induction of apoptosis were also quantified by the flow cytometry experiment. As shown in Figure 4, there is an increase in cells in the FITC channel upon treatment with GIV, representing the loss of MMP. CCCP is used as a positive control that depolarizes the mitochondria in almost all the cells. The bar diagram in Figure 4 shows the extent of depolarization with different concentrations of GIV.

ROS generally have a short life span and are highly reactive. At a moderate level, ROS are good for regulating normal physiological functions, and for maintenance of the redox balance and immune system. However, an excess of ROS damages nucleic acids, proteins, lipids, membranes, and organelles, leading to apoptosis-related cell death. ROS have an essential role in cell signaling and also regulate key pathways related to apoptosis, which are mediated by death receptors, the mitochondria, and the endoplasmic reticulum (Redza-Dutordoir and Averill-Bates, 2016). ROS generation has been reported to be a part of the activity of many chemotherapeutic drugs. There are various methods to measure free radical production in mammalian cells, but the use of 2ʹ-7ʹ-dichlorodihydrofluorescein diacetate (DCFH-DA) is the most common for this purpose. DCFH-DA is basically a cell-permeable, intracellular nonfluorescent precursor of DCF that is used as a probe to measure oxidative stress of cells. DCFH-DA gets hydrolyzed to DCFH that is impermeable and therefore remains inside the cell, which is finally oxidized to dichlorofluorescein that emits green fluorescence (Eruslanov and Kusmartsev, 2010). This DCF accumulation is measured by the increase in fluorescence at 530 nm by the FL-1 green channel of fluorescence microscopy. In the experiment of this study, HCT116 and HT29 cells treated with GIV showed an increase in green fluorescence in a concentration-dependent manner. As shown in Figure 5 and Supplementary Figure S5, the number of cells exhibiting green fluorescence increased with higher concentrations of GIV than did control cells. This indicates an increase in ROS generation upon treatment with GIV.

To check the apoptosis-related genes at the transcriptome level, gene expression profiling was performed in control HCT116 cells and 48-h drug-treated cells at concentrations of 1, 2, and 3 mM. BCl2, Caspase 3, Caspase 9, and PARP were used for expression profiling, and 18S RNA was used as a housekeeping gene. we continued our experiments with HCT116 cells alone as HT29 had an IC50 value at higher concentration IC₅₀ value at higher concentration, so we continued our experiments with HCT116 cells alone.

Bcl2 protein family is a well characterized protein family which has an important role in regulating the intrinsic pathway of apoptosis. This family of proteins contain both proapoptotic and antiapoptotic members. Bcl2 is an antiapoptotic member of this protein family (Tsujimoto, 1998). Bcl2 protein has been shown to promote an increase in cancer cell population by preventing cell death and promote resistance to chemotherapy in cancer cells (Reed et al., 1996). It conserves the integrity of the mitochondrial membrane to inhibit apoptosis, as it contains a hydrophobic carboxyl-terminal domain that is linked to the outer mitochondrial membrane. Bcl2 interacts with proapoptotic BAX, thereby preventing oligomerization of BAX and BAK and their translocation to the mitochondrial membrane. The translocation of BAX–BAK oligomer to the mitochondrial membrane punches pores in the membrane, releasing various proapoptotic factors such as cytochrome C from the mitochondria. Thus increased Bcl-2 expression prevents the apoptosome complex formation, preventing the activation of downstream caspase cascade. Bcl-2 has been also shown to control the initiation of some initiator caspases such as caspse2, thereby preventing apoptosis (Guo et al., 2002).

Caspases are endopeptidases, which are widely divided into three groups, namely, initiator (caspase 2, 8, 9, and 10), executioner (caspase 3, 6, and 7), and inflammatory (caspase 1, 4, 5, 11, and 12). The initiator caspases activate the apoptotic signaling cascade; however, the executioner does the proteolysis of important proteins that is important for apoptosis (Role of caspases in apoptosis, 2021). Caspase 9 is an initiator caspase that activates downstream executioner caspases, thus initiating apoptosis. By forming an apoptosome, the seven APAF-1 subunits activate one caspase-9; the activated caspase-9 then cleaves-3,-6 and -7, Which in turn cleaves various cellular targets, resulting in (Kuida, 2000). Caspase-3 is an executioner caspase, which upon activation performs an effective function of/in apoptosis by degrading intracellular protein (Porter et al., 2021). Tetrandrine citrate shows antitumor activity in glioma cells by increasing the expression of caspase-3 and ROS generation, while decreasing Bcl2 expression (Sun et al., 2019). The data from earlier JC-1 staining and ROS generation point toward the involvement of the intrinsic pathway of apoptosis. As a result, the levels of caspase-9, the intrinsic pathway intiator caspase, and executionary caspase-3 were measured.

PARPs (poly ADP–ribose polymerases) are the enzymes that catalyse the transfer of ADP–ribose to target proteins. PARPs also have an important role in various biological processes inside a cell, such as chromatin structure modulation, replication, transcription, recombination, and DNA repair (Morales et al., 2014). PARP helps damaged cells to repair themselves, thereby playing an important role in cancer. Therefore, PARP inhibitors stop PARP from performing repair work, thereby making cancer cells die (Cancer Research UK, 2021). PARP cleavage is found to be associated with cell death. Therefore, the cleavage product enhances apoptosis and prevents cell survival (D’Amours et al., 2001).

RT-QPCR experiments after GIV treatment show a significant increase in caspase-3 and caspase-9 levels in a concentration-dependent manner. However, a slight change in expression of Bcl-2 (antiapoptotic gene) and PARP (damaged cells repair gene) has been also observed, as shown in Figure 6. The increased expression of proapoptotic caspases and decreased expressions of antiapoptotic Bcl-2 and DNA repair–associated PARP further confirm apoptosis induction by GIV treatment.

After investigating the expression of apoptosis-related genes at the transcriptional level, their expressions at the protein level were checked by western blot analyses. Caspases have been reported to exist in an inactive zymogen state in healthy cells, but upon induction of apoptosis, these inactive zymogens cleaved to form an active caspase. As shown in Figure 7, caspases3 and 9 show cleaved products, which is a feature of apoptosis. The western blot analyses further confirmed the decrease in Bcl-2 expression as observed in the Rt-PCR experiments. Cleavage of PARP is another important event during apoptosis, which was also reported after GIV treatment. The western blots further confirmed that GIV induces apoptosis by activation of caspases, decreasing the expression of Bcl-2. These results confirmed the involvement of an intrinsic pathway of apoptosis in cell death of GIV-treated cells. Furthermore, the morphological features of dying cells along with the expression changes of the Bcl-2, Parp, and Caspase genes support apoptosis rather than any other form of cell death.