Biogenic metal nanoformulations induce Bax/Bcl2 and caspase mediated mitochondrial dysfunction in human breast cancer cells (MCF 7)

Abstract

Nanostructures of noble metals have been extensively studied recently because of their impressive physiochemical properties and wide range of applications in biology and medicine. In this study, the anticancer efficacy of green synthesized silver and gold nanoparticles (AgNPs and AuNPs) was assessed against human breast carcinoma cells (MCF-7). Treatment with different concentrations of NPs triggers the cellular toxicity in a dose and time dependent manner. Morphological features of apoptosis were measured using cell wall integrity, acridine orange/ethidium bromide and Hochest staining methods which clearly distinguishes the viable cells and the cells undergoing apoptosis. Flow cytometry and DNA fragmentation analysis were also used to substantiate that NPs induced cytotoxicity was primarily mediated by G2/M cell cycle arrest and apoptosis. NPs provoke intracellular reactive oxygen species that cause damage to various cellular components. Furthermore, the gene expression studies such as reverse transcription – polymerase chain reaction, quantitative polymerase chain reaction and western blot analysis shows the upregulation of Bax, Bcl2, caspases-6 and -9, PARP, p53 and downregulation of Bcl-2 depicts the induction of apoptosis upon exposure to NPs. The overall results clearly shows that green synthesized metal NPs can potentially inhibit the proliferation of MCF-7 cells and trigger apoptosis through Bax/Bcl2 and caspase–cascade mediated mitochondrial dysfunction. This research concludes that biogenic metal nano-drug formulations can be utilized for cancer chemotherapy.

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1. Introduction

In earlier reports, biological synthesis and assembly of metal nanoparticles (NPs) have been discussed extensively. Now it is very important to extend this active nanomaterial research towards drug development for various deadly diseases such as cancer, malaria, AIDS and tuberculosis. Among these cancer has always been one of the most potent scourges of mankind, that causes six million deaths and more than ten million new cases every year worldwide (WHO). It is also thought that by the end of 2020 the incidence of new cancer cases will increase two fold.¹ In particular breast cancer is the most common malignancy found among women in the most developed and developing regions of world, accounting for 23% of all female cancers. Incidence rates of breast cancer vary between countries according to age, socio-economic background, geographic distribution, stage at presentation and biological characteristics.²

Many cancer patients worldwide are receiving some form of chemotherapy in hope of prolonging their life. However, the two major limitations of traditional chemotherapy are dose-limiting systemic toxicity, which induce toxic responses in rapidly dividing cells and prevalence of multiple drug resistance (MDR).³ To conquer this situation there is urgent need of new tools and technologies with more specificity, cost-effective and have limited side effects to improve the quality of life in cancer patients.

Nanotechnology-based combinational therapy offers an extremely effective mechanism for cancer treatment which limits the detrimental side effects and the prevalence of MDR.⁴ There are different types of nanoformulations such as liposomes, dendrimers, polymeric micelles, carbon nanotubes and magnetic NPs which have been successfully developed for early diagnosis and treatment of cancer.⁵ Among the other nanomaterials, metallic NPs and especially, noble metals such as silver (Ag) and gold (Au) nanoparticles have emerged as being possible candidates for the highest degree of commercialization and extensive biomedical applications.

However, a combination of nanotechnology and molecular biology provides a new model for cancer treatment through the identification of novel molecular targets at various stages of carcinogenesis. Apoptosis, especially, remains as a key denominator for many of these approaches in the elimination of cancer cells. The execution of apoptosis is mediated by a class of cysteine proteases called caspases that are activated either by death receptor activation or mitochondrial membrane permeabilization.⁶ Several other protein families such as Bcl-2 and Bax are also influences the regulation of programmed cell death, thus, the up and downregulation these proteins ultimately determines the cell death.^7,8

The cytotoxicity effects of inorganic NPs are strongly influenced by their size, shape, surface chemistry and ability to bind with biological macromolecules.^9–11 Studies on different cell lines proved that metal NPs can easily cross through the cellular barriers and cause DNA damage, chromosomal aberrations and finally cell cycle arrest.¹² Biological synthesis of metal NPs using pharmacologically important plant entities have shown enhanced biological activities because of their surface functionalization with bioactive metabolites. In this study the molecular mechanism involved in biogenic metal NPs induced cytotoxicity against human breast carcinoma cells (MCF-7) was investigated.

2. Experimental section

2.1. Materials

Silver nitrate, chloroauric acid, cisplatin, Hoechst 33258, 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide (MTT), 2-7-diacetyl dichlorofluorescein (DCFH-DH), heat inactivated fetal bovine serum (FBS), minimum essential medium (MEM), glutamine, ethylenediaminetetraacetic acid (EDTA) and trypsin were purchased from Sigma-Aldrich (St. Louis, USA). Breast cancer cell line (MCF-7) was obtained from National Centre for Cell Science (NCCS), Pune, India. The MCF-7 cells were grown as monolayer in MEM, supplemented with 10% FBS, 1% glutamine, and 100 U ml⁻¹ penicillin–streptomycin and incubated at 37 °C in a 5% carbon dioxide atmosphere. Stocks were maintained in a 75 cm² tissue culture flask.

2.2. Synthesis and characterization of biogenic AgNPs

Podophyllum hexandrum Royle (Himalayan Mayapple) leaf extract was employed as a reducing and stabilizing bioagent for the synthesis of silver nanoparticles (AgNPs) and gold nanoparticles (AuNPs). The physiochemical properties of the synthesized NPs were characterized using ultraviolet-visible spectrophotometry (UV-vis), transmission electron microscopy (TEM), X-ray diffraction (XRD) and Fourier transform infrared spectroscopy (FTIR) as reported in earlier papers.^13,14

2.3. In vitro anticancer effect of biogenic NPs

2.3.1. Cytotoxicity assay. The cytotoxicity of the NPs were measured using a MTT assay as described earlier.¹⁵ Briefly, cells were seeded at a density of 5 × 10⁴ cells per well in 96-well plates. After 24 h, the cells were treated with NPs at various concentrations (50–250 μl ml⁻¹) and then incubated for 24 h. At the end of the incubation, 10 μl of MTT (5 mg ml⁻¹) per well was added and the plate was further incubated in dark at 37 °C for 4 h. The formazan crystals formed after 4 h were solubilized in 100 μl of dimethylsulfoxide after aspirating the medium. The absorbance was monitored at 570 nm (measurement) and 630 nm (reference) using a 96-well plate reader (Bio-Rad, Hercules, CA, USA). Each growth curve showed the means and standard deviation (SD) of at least three independent experiments. The growth inhibition was determined using: growth inhibition = (control optical density (OD) − sample OD)/control OD. The IC₅₀ value was defined as the concentration of NPs that produced a 50% reduction of cell viability.

2.3.2. Cytomorphological evaluation using phase contrast microscopy. After the incubation with NPs at different concentrations for 24 h, the gross morphological changes in the cells were observed using an inverted phase contrast microscope (Nikon, Japan) and photographed using a Nikon digital camera (Nikon, Japan).

2.4. Evaluation of apoptosis

2.4.1. Acridine orange (AO) and ethidium bromide (EB) staining. The extent of apoptosis induced by the NPs was identified by the morphological changes that occurred in the MCF-7 cells after 24 h of incubation.¹⁶ Cells were harvested and suspended in phosphate buffered saline solution (PBS). Cells were stained with 5 μl of a cocktail mix containing AO (100 μg ml⁻¹) and EB (100 μg ml⁻¹) in PBS. A cell suspension of 10 μl was applied to a glass slide, covered with a cover slip and cells were viewed and photographed under a fluorescent microscope (Nikon 80i Eclipse, Japan). Viable cells were identified by a bright green nucleus with intact structure. Early apoptotic cells show a bright green nucleus with condensed chromatin, while nuclei of apoptotic cells in the late phase were stained orange and showed a condensed chromatin structure. Whereas necrotic cells displayed an orange nucleus without condensed chromatin. At least 200 cells were counted for each condition tested.

2.4.2. Hoechst 33258 staining. The nuclear changes and apoptotic bodies induced by the NPs were characterized using Hoechst 33258 staining. Briefly, cells in 24-well plates were trypsinized, washed twice with PBS buffer (pH 7.4) and 25 μl of the cell suspension was stained with Hoechst 33258 for 10 min at room temperature in the dark. The condensed or fragmented nuclei of apoptotic cells were observed using fluorescence microscopy with a 4′,6-diamidino-2-phenylindole (DAPI) filter (excitation 365 nm and emission, 480 nm).

2.4.3. DNA fragmentation. Cells were collected after 24 h of treatment with NPs at the specified concentrations. DNA was extracted according to the standard protocol for DNA isolation.¹⁷ Briefly, cells were treated with different concentrations of NPs for 24 h, cells were harvested, counted and washed with PBS at 4 °C. The cells were pelleted using centrifugation at 200 g at 4 °C. The pellet was suspended in DNA lysis buffer [1 M Tris (pH 8.0), 0.5 M EDTA and 75% sodium lauryl sarcosine] and incubated overnight with proteinase K (0.5 mg ml⁻¹) at 50 °C. After overnight incubation RNase was added and the mixture was further incubated for an hour at 50 °C. DNA was extracted using phenol : chloroform (1 : 1 v/v) and then the extract was electrophoresed in 2% agarose gel for 2 h at 50 V. The gel was stained with EB (0.5 μg ml⁻¹) followed by UV exposure and photographed in the Gel Doc XR+ system (Bio-Rad, Hercules, CA, USA).

2.4.4. Determination of intracellular ROS. Cellular oxidative damage induced by NPs was quantified using a cell permeable dye, 6-carboxy-2′,7′-dichlorodihydrofluorescein diacetate (DCFDA) (Invitrogen). This compound is oxidized by reactive oxygen species (ROS) into the fluorescent carboxydichlorofluorescein (DCF) inside the cells. Briefly, experimental cells were incubated with 10 μM of DCFHDA for 30 min at 37 °C in the dark, washed twice with PBS and the fluorescence resulting from the oxidation of the dye in the cells was measured with an excitation wavelength of 480 nm and an emission wavelength of 530 nm.

2.4.5. Semi-quantitative RT-PCR and quantitative real time PCR analysis. Total RNA was extracted from experimental cells using TRIzol® reagent (Invitrogen, USA). RNA isolated from the cells was reverse-transcribed and amplified using the one-step reverse transcription – polymerase chain reaction (RT – PCR) System (Fermentas, USA). Forward and reverse primers for Bax (NM_001188.3) were 5′-GCC ACC AGC CTG TTT GAG-3′ and 5′-CTG CCA CCC AGC CAC CC-3′, for Bcl2 (NM_000657.2) were 5′-TAT AAG CTG TCG CAG AGG GGC TA-3′ and 5′-GTA CTC AGT CAT CCA CAG GGC GAT-3′ and for GAPDH (NM_002046.3) were 5′-AAT CCC ATC ACC ATC TTC CA-3′ and 5′-CCT GCT TCA CCA CCT TCT TG-3′, respectively. The PCR conditions were set as follows: 94 °C for 5 min; 35 cycles of 94 °C for 1 min, 55–62 °C for 1 min, 72 °C for 1 min and a final extension step of 72 °C for 10 min. The products were also verified using agarose gel electrophoresis. The level of GAPDH gene expression served as an internal control. From the prepared RNA, cDNA was prepared by using a Qiagen cDNA preparation kit. The mRNA expression of p53 was detected after treatment of the NPs using p53 forward and reverse primers as 5′-AATCATCCATTGCTTGGGACG-3′; 5′-CCGCAGTCAGATCCTAGCG-3′ in a 7500 real-time PCR System (Applied Biosystems) with the SYBR® Green Master mix (Invitrogen). Relative p53 mRNA levels were determined by comparing the PCR cycle thresholds between the cDNA of a specific gene and histone (ΔC_t).

2.4.6. Western blotting. MCF-7 cells were cultured in 6-well plates and then treated with or without NPs for 24 h. The cells were then lysed in a buffer containing 50 mM Tris–HCl (pH 8.0), 150 mM sodium chloride, 0.02% sodium azide, phenylmethane sulfonyl fluoride, aprotinin, and 1% Triton X-100, and centrifuged at 12 000 rpm for 30 min at 4 °C. The quantified proteins were electrophoresed using sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). The separated proteins were transferred to nitrocellulose membranes. Membranes were washed with tris-buffered saline Tween-20 (TBST), blocked with 5% skimmed milk for 1 h at 37 °C, incubated overnight at 4 °C with either goat anti-rabbit caspase 3, 8 and 9 or β-actin antibodies at the manufacturer's recommended dilutions. After incubation, the membrane was washed with TBST buffer. The membranes were then incubated with secondary mouse anti-goat peroxidase conjugated antibodies (Cell Signalling technologies, USA) for 1 h at 37 °C. After washing the membranes with TBST, they were developed with the diaminobenzidine (DAB) chromogenic detection method and then scanned.

2.4.7. Analysis of cell cycle disruption by flow cytometry. After their exposure to NPs, cells (1 × 10⁶ in 10 ml) were trypsinized and fixed in ethanol and stored at 4 °C until assayed. Samples were pelleted at 2000 rpm for 5 minutes, pellets were washed twice with ice-cold PBS and centrifuged for a further 5 minutes. Subsequently, the pellets were resuspended in 0.5 ml DNA staining solution (25 μg ml⁻¹ propidium iodide, 100 μg ml⁻¹ RNase A in PBS) and incubated at 37 °C for 30 minutes in the dark. Cell cycle analysis (10 000 events for each sample) was performed using a FACSCalibur flow cytometer with CellQuest software (BD Biosciences, San Jose, CA, USA) and expressed as fractions of cells in different cell cycle phases. Samples were run in triplicate and each experiment was repeated three times.

3. Results and discussion

The use of existing drugs for cancer chemotherapy are limited with poor specificity, high cost, high toxicity, side effects and emergence of drug resistance. To overcome this situation nano sized materials have been intensively studied because of their unique physiochemical properties, stability and biological fate.^18,19 In particular, biogenesis of nanomaterials was found to be a viable and easy alternative strategy because of its green chemistry approach. Apart from the fact that it is an eco-friendly method, interaction of biocompounds with noble metals offers multifunctional hybrid nanomaterials for cancer diagnosis and therapy.^20,21 This study was performed with a view to revealing the molecular mechanism involved in biogenic NPs induced cytotoxicity against human breast carcinoma cells. The cell viability assay clearly explains the cellular response to biogenic metal NPs, and the size, morphology and surface functionalization have been shown to have a major influence on biokinetics and toxicity.²²

3.1. Synthesis and characterization of biogenic NPs

As reported in our previous studies,^13,14 synthesized AgNPs and AuNPs using the leaf extract of P. hexandrum was initially confirmed with the development of a yellowish brown and a ruby red colour, respectively, because of the excitation of surface plasmonic vibrations. UV-Vis spectrophotometry of synthesized AgNPs produces an absorbance spectrum at 420 nm whereas AuNPs have shown an surface plasmon resonance (SPR) peak at 530 nm. TEM micrographs of synthesized AgNPs and AuNPs have been shown to be stable, crystalline and polydispersed with a size range of 12–40 nm and 5–35 nm respectively. A thin-layer of biomolecule coating on the surface of the nanoparticles which prevents the aggregation of NPs was also noticed. Interestingly, the XRD diffractogram of synthesized NPs gives Bragg's reflections at (1 1 1), (2 0 0), (2 2 0) and (3 1 1) which strongly show that synthesized NPs were face centred cubic (Fcc) crystalline in nature. Furthermore, FTIR analysis revealed that the water soluble phenolic constituents of P. hexandrum are mainly involved in the reduction and stabilization of NPs. Furthermore, the transmittance has clearly exposed the interaction of phenolic compounds with O–H functional groups, which may complete the reduction process.

3.2. Assay for cell viability

MTT data clearly reveals that the treatment with NPs partially decreased the MCF-7 cell viability in a dose and time dependent manner. From Fig. 1 it is evident that a 100% cell mortality rate was found at the maximum concentration after 48 h. Consequently, the IC₅₀ inhibitory concentration of AgNPs and AuNPs were fixed at 100 μl ml⁻¹ (Fig. 1a) and 200 μl ml⁻¹ (Fig. 2b), respectively. As described previously by various investigators, the toxicity of AgNPs could be based on their surface morphology and size.²³ In a recent study, it was noted that biogenic AgNPs synthesized using the leaf extract of Dendrophthoe falcata with a size range of 5–45 nm has shown enhanced cytotoxicity against human breast carcinoma cells (MCF-7) compared to an aqueous plant extract.²⁴ Likewise AuNPs also displayed size dependent cytotoxicity in treated fibroblasts, epithelial cells and melanoma cells.²⁵ Another report also showed that AuNPs induce cytotoxicity against human breast epithelial (MCF-7) cells in a dose dependent manner with a concentration range between 25 and 200 μg ml⁻¹.²⁶

Fig. 1 Inhibitory effects of the biosynthesized AuNPs and AgNPs on MCF-7 cells treated with different doses (50, 100, 150, 200 and 250 μl ml⁻¹) for 24 and 48 h. All the data are expressed as the mean ± SD of the three experiments with duplicate wells.

Fig. 2 Morphological characterization under phase contrast microscopy of cell death induced by the AuNPs and AgNPs in MCF-7 cells for 24 hours. Magnification at 200×.

3.3. Measurement cytomorphological changes in MCF-7

Phase contrast microscopy demonstrated dose dependent detachment of non-viable cells from the surface of culture plates (Fig. 2). MCF-7 cells exposed to NPs showed morphological changes including cell shrinkage and formation of apoptotic bodies. This result corroborated well with results from an earlier study where AgNPs synthesized from the leaf extract of Vitex negundo L. induced cellular shrinkage in treated human colon cancer cell line (HCT15).²⁷ These morphological changes are because of the activation of caspase cascades, which cleave the specific substrates responsible for the DNA repair activation. When NPs come in to contact with the cells they are taken up by a variety of mechanisms such as clathrin-dependent endocytosis and macropinocytosis.^28,29 These can lead to activation of cellular signalling processes producing ROS, inflammation and finally cell cycle arrest or cell death.³⁰

3.4. Detection of NPs induced apoptosis in MCF-7 cells

Apoptosis is the key event in cancer therapy that can be measured with the activation caspase–cascade, chromatin aggregation, partition of the cytoplasm and nucleus into membrane-bound vesicles (apoptotic bodies) that contain ribosomes, morphologically intact mitochondria, and nuclear material.³¹ In this study NPs treated MCF-7 cells lose their viability in a time and dose dependent manner. The ability of NPs (AgNPs and AuNPs) to induce apoptosis was determined by AO-EB staining. After treatment with the concentration of NPs, the cells were harvested and stained with AO-EB as described in the materials and methods section. Cells exposed to NPs showed greater apoptotic effect (Fig. 3a) when compared to control cells. A similar phenomenon was reported with Acorus calamus rhizome extract AgNPs that activated apoptosis in treated HeLa cells.³² Furthermore, the effects of NPs on the gross nuclear morphology of the MCF-7 cells were observed using fluorescence microscopy after Hoechst 33258 staining. After the treatment with different concentrations of NPs for 24 h, MCF-7 cells began to exhibit apoptotic characteristics such as nuclear blebbing, nuclear condensation and fragmentation. In the control group, the cells were regular in morphology and grew fully in patches and were confluent, and rarely sloughing off (Fig. 3b). Fig. 3c shows the percentage distribution of apoptotic cells (viable, early, late and necrotic cells).

Fig. 3 AuNPs and AgNPs induced apoptosis in MCF-7 cells. (a) Photomicrographs of AO/EB stained MCF-7 cells incubated for 24 h with either AuNPs or AgNPs. Viable (light green), early apoptotic (bright green fluorescence), late apoptosis (red to orange fluorescence) and necrosis (red fluorescence) cells were observed. (b) Representative fluorescent micrographs of MCF-7 cells stained with Hoechst 33258 fluorescent dye after 24 h incubation with either AuNPs or AgNPs. Magnification at 200×. (c) Percentage distribution of apoptotic cells (viable, early, late and necrotic cells). (d) AgNPs induced inter-nucleosomal DNA fragmentation. MCF-7 cells were treated with 0, 100 and 200 μl ml⁻¹ of AgNPs for 24 h. Cells were harvested and DNA was extracted as described in the Materials and methods section. Fragmented DNA was analyzed by agarose gel electrophoresis. Representative gels from one of the three experiments.

Fig. 4 AuNP and AgNP induced ROS in MCF-7 cells. MCF-7 cells were treated with different concentrations of AuNPs or AgNPs for 24 h and ROS was determined as mentioned in the Materials and methods section. All the data are expressed as the mean ± SD of the three experiments with duplicate wells. Statistically significant difference as compared to the controls (*p < 0.05 for each).

3.5. Effect of NPs induced DNA damage on MCF-7 cells

NPs have induced apoptosis in treated MCF-7 cells through DNA damage that have been confirmed with the DNA fragmentation assay. The induction of a DNA single strand break is often used to predict oxidative damage of tumour cells. As shown in Fig. 3d NPs pre-treatment at specific concentrations (AgNPs – 200 μl) and (AuNPs – 200 μl and 400 μl), cause the formation of a DNA ladder which shows that the NPs have induced cell death through the apoptotic pathway. However, a minimal dosage of AgNPs (100 μl) does not cause the formation of DNA laddering. It is now well established that excessive generation of ROS triggers signal transduction pathways leading to apoptosis.^29,30 In corroborate with several studies our data clearly proved that NPs induced cytotoxicity was initially triggered by oxidative stress mediated DNA damage. An important aspect of DNA damage is the formation free radicals that mainly cause a single strand break. The DNA laddering observed in the present study reveals that the generation of ROS (Fig. 4) is favoured over DNA damage and leads to cell death. An earlier study proved that highly reactive hydroxyl radicals released by the AgNPs attack cellular components including DNA, lipids, and proteins which causes various kinds of oxidative damage.³¹

3.6. NPs induction of apoptosis via mitochondria- and caspase-dependent pathway

To study the possible molecular mechanisms of NPs mediated cell death, the apoptotic regulators have been measured using mRNA and protein expression patterns. In order to confirm that NPs induced ROS which mediated the apoptotic pathway, the mRNA levels of Bcl-2 and Bax were semi-quantitatively determined using RT-PCR. Interestingly, after the treatment with NPs, the level of Bax and GAPDH were upregulated whereas the level of Bcl-2 was found to be decreased in a dose dependent manner (Fig. 5a). However, the activation of mitochondrial apoptotic pathway in caspase-6 and caspase-9 showed a significant increase after 24 h of treatment with NPs (Fig. 5b). Increase in the synthesis level of the poly(ADP-ribose) polymerase (PARP) shows that NPs induced apoptotic and necrotic cell death was mediated via Bcl-2/Bax proteins and the caspase–cascade. All the expression was normalized to the levels of the β-actin expression. These results clearly indicate that the mitochondrial signalling pathway was playing a crucial role in NPs-induced apoptosis in MCF-7cells. According to the present study NPs trigger the apoptotic protein Bax and downregulates Bcl-2 which results in activation of programmed cell death. It is well known that anti-apoptotic members of the Bcl-2 family specifically inhibit the release of certain apoptogenic factors.³² Downregulation of Bcl-2 is crucial in caspase activation and the induction of apoptosis because it releases cytochrome C from the mitochondria.⁶ However, in Bcl-2 heterodimerization with Bax, when the Bcl-2 expression level was downregulated, there was upregulation of Bax expression, and homodimers of Bax will always be formed and apoptosis will be stimulated. Earlier studies have proved that the ratio between pro- and anti-apoptotic proteins determine the susceptibility of cells to a death signal.³³ The caspases–cascade (cysteine-requiring aspartate proteases) was the most important characteristic of apoptosis, which can be triggered by several groups of so-called initiators (e.g., caspase-8, capase-9) and effectors (e.g., caspase-3, caspase-7, caspase-6). Among these caspase-3 and caspase-6 play a crucial role in apoptosis, whose activation is mediated by inhibitor caspases such as caspase-8 and caspase-9. Several studies have pointed out that the activation of caspases has shown a direct effect on the mitochondrial membrane potential.^34,35 However, further investigations are required to elucidate the potential cytotoxicity of different cell lines to NPs in vitro as well as to assess the biocompatibility and biosafety of such particles in vivo based on their size.

Fig. 5 a) Effect of AuNPs or AgNPs on mRNA levels of Bcl₂ and Bax in MCF-7 cells. b). Effect of AgNPs on caspase-6 and caspase-9 and PARP protein expression as determined by western blot analysis in the MCF-7 cell line. β-Actin was used as the control. Representative western blots of experiments performed in triplicate.

3.7. Cell cycle arrest

Generally, apoptosis and the cell cycle arrest are closely related.³⁶ In our study NPs treated MCF-7 cells, the proportion of cells was found to be decreased in the G1 and increased in subG1 phases of the cell cycle (Fig. 6). Furthermore as shown in Fig. 7, the mRNA levels of p53 in NPs treated cells was upregulated which confirms that the p53 induced DNA damage. These data clearly prove that metal nanoformulations developed in this research, actively inhibit the progression of the MCF-7 cell cycle at the G1 phase and the accumulation of cells at the subG1 phase which confirms the activation of the apoptotic pathway after NPs exposure.

Fig. 6 Effect of AuNPs or AgNPs on cell cycle distribution of MCF-7 cells. All the data is expressed as the mean ± SD of the three experiments.

Fig. 7 Quantitative real-time PCR analysis of p53 mRNA levels in MCF-7 cells exposed to AuNPs and AgNPs. Data represented are mean ± SD of three identical experiments with three replicate. Statistically significant difference as compared to the controls (*p < 0.05 for each).

4. Conclusion

To summarize the present study, biogenic metal NPs exhibits prominent anticancer activity against MCF-7 cells. At a minimal dosage NPs trigger mitochondrial-driven apoptosis via activation of procaspase-3 and procaspase-9. In addition, decreased expression of Bcl-2, increased expression of Bax and cleaved PARP proves the enhanced anticancer activity of bionanoparticles. The overall results suggested that biogenic metal nanoformulations can be potentially developed into a cost-effective, commercially viable drug candidate to combat breast cancer. In future, the synthesized NPs have to be assessed against different types of cancer cells. Furthermore, it is very important to assess the preclinical tests and the other molecular signalling pathways including angiogenesis need to be investigated and these studies are in progress.

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Footnote

† Authors contributed equally.

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