Anticancer Potential of Aqueous Ethanol Seed Extract of Ziziphus mauritiana against Cancer Cell Lines and Ehrlich Ascites Carcinoma

Abstract

Ziziphus mauritiana (Lamk.) is a fruit tree that has folkloric implications against many ailments and diseases. In the present study, anticancer potential of seed extract of Ziziphus mauritiana in vitro against different cell lines (HL-60, Molt-4, HeLa, and normal cell line HGF) by MTT assay as well as in vivo against Ehrich ascites carcinoma bearing Swiss albino mice was investigated. The extract was found to markedly inhibit the proliferation of HL-60 cells. Annexin and PI binding of treated HL-60 cells indicated apoptosis induction by extract in a dose-dependent manner. The cell cycle analysis revealed a prominent increase in sub Go population at concentration of 20 μg/ml and above. Agarose gel electrophoresis confirmed DNA fragmentation in HL-60 cells after 3 h incubation with extract. The extract also exhibited potent anticancer potential in vivo. Treatment of Ehrlich ascites carcinoma bearing Swiss albino mice with varied doses (100-800 mg/kg b.wt.) of plant extract significantly reduced tumor volume and viable tumor cell count and improved haemoglobin content, RBC count, mean survival time, tumor inhibition, and percentage life span. The enhanced antioxidant status in extract-treated animals was evident from decline in levels of lipid peroxidation and increased levels of glutathione, catalase, and superoxide dismutase.

Figure 1

Proliferation inhibition in different human cell lines upon ZMS treatment. ZMS at indicated concentrations were evaluated for its in vitro cytotoxicity against different cancer cell lines (HL-60, HeLa, and Molt-4) and normal cell line (HGF) employing MTT reduction assay. The results are expressed as percent of cell growth inhibition determined relative to untreated control cells. Data are mean value ± S.E.M. (n = 8) and representative of one of the three similar experiments.

Figure 2

Flow cytometric analysis of ZMS-induced apoptosis and necrosis in HL-60 cells using annexinV-FITC and PI double staining. HL-60 cells (1 × 106 /mL) were incubated with indicated concentrations of ZMS for 12 hours and stained with Annexin V-FITC/PI. Quadrant analysis of fluorescence intensity of ungated cells in FL-1 versus FL-2 channels was from 10000 events. Cells in the lower right quadrant represented apoptosis while in the upper right quadrant indicated postapoptotic necrosis and representative of one of the three similar experiments.

Figure 3

Cell cycle analysis in HL-60 cells after ZMS treatment. HL-60 cells (1 × 106 /mL) were treated with different concentration of ZMS for 24 hours. Cells were stained with PI to determine DNA fluorescence by flow cytometer. Sub-GO population indicative of DNA damage was analyzed from the hypodiploid sub-Go fraction (<2n DNA) of DNA cell cycle analysis. The cells for hypodiploid (sub GO/G1, <2n DNA) population were analyzed from FL2-A versus cell counts shown in % and representative of one of the three similar experiments.

Figure 4

Dose-dependent and time-dependent induction of DNA fragmentation in HL-60 cells after ZMS treatment. (a) 2 × 106 HL-60 cells treated with different concentrations of ZMS extract for 24 hours. DNA was electrophoresed on 1.8% agarose gel and stained with ethidium bromide. M-Marker; C-Untreated HL-60 cells; 10-10 μg/mL of ZMS treated HL-60 cells; 20-20 μg/mL of ZMS treated HL-60 cells; 40-40 μg/mL of ZMS treated HL-60 cells; 80-80 μg/mL of ZMS treated HL-60 cells; +Ct− Camptothecin treated HL-60 cells. (b) 2 × 106 HL-60 cells were treated with 20 μg/mL of ZMS extract and were incubated for 3 hours, 6 hours, 9 hours, 12 hours, and 24 hours. DNA was electrophoresed at respective time interval on 1.8% Agarose gel and stained with ethidium bromide.

Figure 5

Change in body weight in EAC bearing mice after ZMS treatment. The results are presented as Mean ± S.E.M. (n = 10). (a) P < .001 in comparison to untreated control; (b) P < .05 in comparison to EAC control group.

Figure 6

Reduction in lipid peroxidation in EAC-bearing mice after ZMS treatment. The results are presented as Mean ± S.E.M. (n = 10). (a) P < .001 in comparison to normal control; (b) P < .05 in comparison to EAC control group.

Figure 7

Augmentation of glutathione content in EAC bearing mice after ZMS treatment. The results are presented as Mean ± S.E.M. (n = 10). (a) P < .001 in comparison to normal control; (b) P < .05 in comparison to EAC control group.

Figure 8

Augmentation of superoxide dismutase activity in EAC bearing mice after ZMS treatment. The results are expressed as Mean ± S.E.M. (n = 10). (a) P < .001 in comparison to normal control; (b) P < .05 in comparison to EAC control group.

Figure 9

Augmentation of catalase activity in EAC bearing mice after ZMS treatment. The results are expressed as Mean ± S.E.M. (n = 10). (a) P < .001 in comparison to normal control; (b) P < .05 in comparison to EAC control group.

Figure 10

Presumptive mechanism of apoptosis induction by ZMS. ZMS may affect the genes and transcription factors or may affect proteosome to induce apoptosis or may trigger some signals that directly change the mitochondrial permeability transition pore. This induces changes in the members of Bcl-2 family. It may interact with BAX (Bcl-2 associated X protein) and BAK (Bcl-2-proapoptotic member) and stimulate the release of cyt. C (Cytochrome C). Cytochrome C released from mitochondria binds to APAF-1 (Apoptotic protease activating factor-1) and procaspase-9 forming apoptosome and activating caspase-9 which in turn activates executor caspases (Caspase 3, 6, and 7) leading to cell death via apoptosis.