Skip to main content
NCBI home page
Toxicology Research logoLink to Toxicology Research
. 2016 Jan 7;5(2):547–556. doi: 10.1039/c5tx00391a

Low-dose chemotherapeutic drugs induce reactive oxygen species and initiate apoptosis-mediated genomic instability†

Renganathan Arun a, Sridaran Dhivya a, Suresh K Abraham b, Kumpati Premkumar a,✉
PMCID: PMC6062221  PMID: 30090369

graphic file with name c5tx00391a-ga.jpgLow-dose chemotherapy drugs initiate defective apoptosis, allowing a small population of cells to escape from cell cycle check points and survive with accumulated genetic damage.

Abstract

Prolonged cancer cell survival, acquiring drug resistance, and secondary cancer development despite chemotherapy are the major challenges during cancer treatment, whose underlying mechanism still remains elusive. In this study, low-doses of chemotherapeutic drugs (LDCD) – doxorubicin (DOX), etoposide (ETOP), and busulfan (BUS) were used to ascertain the effect of residual concentrations of drugs on breast cancer cells. Our results showed that exposure to LDCD caused significant induction of ROS, early signs of apoptosis and accumulation of cells in S and G2-M phases of the cell cycle in MCF-7 and MDA-MB-231 cell lines. Under drug-free recovery conditions, a decrease in the number of apoptotic cells and an increase in the number of colonies formed were observed. Analysis of the molecular mechanism showed lower expression of cleaved products of caspase 3, 9, PARP and occurrence of DNA strand breaks in recovered cells compared to LDCD-treated cells, suggesting incomplete cell death activation and survival of cells with genomic damage after therapeutic insult. Thus, LDCD induces defective apoptosis in cancer cells allowing a small population of cells to escape from cell cycle check points and survive with accumulated genetic damage that could eventually result in secondary cancers that warrants further studies for better therapeutic strategies.

1. Introduction

Chemotherapy is a feasible and effective therapeutic modality for treatment of various cancers, which is performed alone or in combination with surgery or radiotherapy.1,2 Apoptosis is a major molecular mechanism targeted by every chemotherapeutic agent to curb cancer progression.3,4 Most of the chemotherapeutic drugs such as etoposide, doxorubicin, methotrexate, camptothecin, busulfan, and vincristine are administered at a maximal tolerated dose (MTD) for a certain period of time followed by a drug-free period.5 However, the development of therapy-related secondary tumors has been observed during the drug-free period.6,7 Continuous administration of low-doses of drugs without drug-free breaks, referred to as metronomic treatment, was chosen as an alternate treatment strategy.8–10 Although chemotherapeutics have led to improvement in patient survival, certain confounding factors cause frequent failure of classical and metronomic chemotherapy leading to numerous drug-related toxicities.11 But, the critical molecular events underlying tumor killing and associated secondary tumor development still remain obscure. While most of the chemotherapeutic drugs directly bind to DNA and initiate apoptosis,12 still cancer cells fight to overcome this pharmacological insult.13

Research over the past decade on evasion of apoptosis by cancer cells has shown that differences in caspase cascade activation by transcriptional factors, histone deacetylases,14,15 mutational variations in pro- and anti-apoptotic proteins resulted in increased chemoresistance, insufficient apoptosis, and growth advantage for malignant cells14 against several chemotherapeutic agents.16 Although metronomic therapy has been an alternative to overcome the limitations of MTD, insufficient or low-doses of chemotherapeutic drugs (LDCD) administered during chemotherapy at levels sufficient enough to destroy proliferating cancer cells17 also have an indirect effect on malignant cells and might influence the relapse of tumors. Our current understanding of the effect of LDCD on cancer progression and the underlying mechanism is still in its infancy and warrants further research to gain a thorough insight into the alternate mechanisms defending cancer cells in response to chemotherapeutic drugs. Thus, this study aims to understand the survival events upon LDCD administration for three widely used drugs – doxorubicin (DOX), etoposide (ETOP), and busulfan (BUS), with different mechanisms of action. DOX acts on transcription factor p53 and DNA topoisomerase II,18,19 while ETOP is a direct topoisomerase II inhibitor20,21 and BUS is a bifunctional DNA-alkylating drug inducing DNA intra- and inter-strand crosslinks22,23 that trigger DNA damage responses leading to apoptosis.

The results of this study show that low-doses of DOX, ETOP, and BUS increase the production of reactive oxygen species (ROS) sufficient enough to induce defective apoptosis in breast cancer cells. This incomplete apoptosis might allow a minor population of cancer cells to escape from apoptosis along with genomic alterations, which could possibly lead to cancer recurrence and therapy-related secondary tumor formation.

2. Materials and methods

2.1. Cell culture

Human breast cancer cell lines MCF-7 and MDA-MB-231 were procured from National Centre for Cell Sciences, Pune, India and cultured in DMEM (HiMedia, India) containing 10% heat-inactivated fetal bovine serum (Gibco, USA), 100 units per ml penicillin G, 100 mg per ml streptomycin (HiMedia, India), 2 mM glutamine (Gibco, USA), 25 mM HEPES, and 2 mM sodium bicarbonate at a cell density of 1–3 × 106 cells per ml. Cells growing in the exponential phase were harvested at approximately 80–90% confluency and subsequently incubated with chemotherapeutic drugs for different time points (12, 24, 48, and 72 h) under standard conditions (37 °C in 5% CO2 with 95% humidity) in an Eppendorf – New Brunswick Galaxy CO2 incubator. The cells were grown up to three to four passages before the treatments.

2.2. Chemicals and drugs

Chemotherapeutic drugs DOX, ETOP, and BUS were purchased from Sigma Aldrich, USA. ETOP and BUS were dissolved in DMSO at 50 mM and 100 mM. DOX was dissolved in deionised water at 50 mM. Throughout the study the final concentration of DMSO in the controls and each sample did not exceed 0.1% to eliminate cytotoxicity of DMSO in both the cell lines.

2.2.1. Dose selection and cell viability assay

Breast cancer cells were treated with varying concentrations of DOX (0, 1, 2, 3, 4, 5, and 6 μM), ETOP (0, 0.5, 1.0, 1.5, 2.0, 2.5, and 3.0 μM), and BUS (0, 20, 40, 60, 80, and 100 μM) for 12 and 24 h to identify the time- and dose-dependent response of cells with chemotherapeutic drugs. Briefly, the breast cancer cells (MCF-7 and MDA-MB-231) were plated at a density of 1 × 104 cells per well on a 96-well plate in 100 μl medium. DOX, ETOP, BUS, or 0.1% DMSO (the drug solvent) were added at selected concentrations to the cell lines after being plated for 24 h. After the different treatment periods, cell viability was analyzed by the MTT assay. All the experiments were performed in triplicates to confirm the consistency of the results obtained.

2.3. Morphological assessment by phase contrast microscopy

Exponentially growing MCF-7 and MDA-MB-231 cells were seeded on 24-well plates at a density of 5 × 104 cells per ml per well. After 24 h, the cells were treated with low, non-cytotoxic doses of chemotherapeutic drugs as confirmed by the MTT assay. After the treatment period of 12 h, digital photographs were taken by using a phase contrast inverted microscope (TSF100, Nikon, Japan) to assess the morphology of the treated cells.

2.4. Clonogenic survival assay

Approximately 0.5–1 × 103 cells were plated in 2.0 ml complete medium onto 6-well culture plates. Cells were allowed to attach for 24 h before adding 2.0 ml of complete medium containing different concentrations of LDCD. After 12 h, the cells were washed twice with PBS and a fresh medium was added and the plates were incubated for 8–10 days for colonies to be formed. The colonies were fixed in methanol and stained with crystal violet and the number of colonies per well were counted to analyze the long-term effects of LDCD on proliferation efficiency.

2.5. Apoptosis measurements

2.5.1. Double staining – Hoechst 33258 and propidium iodide

The extent of apoptosis triggered by LDCD was determined by propidium iodide (PI, Sigma) and Hoechst 33258 (Sigma) double fluorescent staining. Briefly, at each time point of treatment, cells were trypsinized and washed twice with PBS (phosphate-buffered saline, pH 7.4). 100 μl of the cell suspension was stained with Hoechst 33258 (10 μg ml–1) and PI (1 μg ml–1) and analyzed for apoptosis and/or necrosis under a fluorescent microscope. Apoptotic cells displayed a distinct morphology characterized by the condensation of chromatin, nuclear shrinkage, and the formation of a few apoptotic bodies.

2.5.2. Western blot analysis for apoptotic markers

Cells treated with LDCD for specific time periods were harvested and lysed using 1× lysis buffer containing 50 mM Tris-Cl (pH 8.0), 150 mM NaCl, 0.02% sodium azide, 100 μg per ml phenyl-methane sulfonyl fluoride (PMSF), 1 μg per ml aprotinin, and 1% Triton X 100, and centrifuged at 10 000g for 30 min at 4 °C. Proteins were quantified by the Bradford assay and equal amounts of protein (30 μg) from the total cell lysate were separated and electro-transferred onto nitrocellulose membranes which were probed with anti-rabbit procaspase 3, 9, PARP; cleaved caspase 3, 9, PARP; and β-actin antibodies as per the manufacturer's recommended dilutions. After incubation, the membrane was washed with TBST (tris-buffered saline tween-20) buffer. The membranes were then incubated with a goat anti-rabbit secondary antibody conjugated to HRP (cell signalling technologies, USA) for 1 h at room temperature, washed, and developed using the DAB chromogenic detection method and scanned for expression of specific markers.

2.6. Intracellular ROS measurement

The fluorescent probe 2′,7′-dichlorofluorescein diacetate (CM-H2DCFDA) was used for assessment of intracellular ROS production. Briefly, cells were seeded (1 × 105 cells per well) in 6-well culture plates and treated with different concentrations of LDCD. After the experimental period, the cells were trypsinized, centrifuged and washed three times with ice-cold PBS. The cells were resuspended in PBS and incubated with CM-H2DCFDA (5 μM) at 37 °C for 20 min. The fluorescence intensity was measured at 530 nm after excitation at 480 nm using a spectrofluorimeter. The increase in fluorescence intensity was used to assess the generation of net intracellular ROS.

2.7. Cell cycle analysis

Analysis of cell cycle regulation upon LDCD treatment was performed as described previously.24 After the drug exposure for 12 h, the cells (1 × 106 in 10 ml) were trypsinized and centrifuged (2000g, 25 °C, 2 min). The pellet was fixed in 70% ethanol and incubated at room temperature for 30 min. After staining the fixed cells with DNA staining solution (25 μg per ml PI, 100 μg per ml RNase A in PBS), cell cycle analysis was performed using a FACSCalibur (BD Biosciences, San Jose, CA). The fraction of the total cell population present in the G1, S, and G2-M phases was obtained from DNA histograms by using Cell Quest software.

2.8. Comet assay

The alkaline comet assay was performed as described previously.25 Briefly, experimental cells after specific incubation periods were mixed with low melting-point agarose at 37 °C to a final concentration of 0.7%. The mixture (15 μl) was spread on 0.5% normal-melting-point agarose pre-treated slides. The slides were then placed in ice-cold lysis buffer (2.5 M NaCl, 100 mM EDTA, 10 mM Trizma base, 10% DMSO, 1% Triton-X) and incubated overnight. After lysis, the slides were placed in horizontal electrophoresis tanks filled with alkaline electrophoresis buffer (300 mM NaOH, 1 mM EDTA; pH 13.0) for another 20 min, and subjected to electrophoresis at 25 V/300 mA for a further 20 min. The lysis and electrophoresis for both neutral and alkaline assays were performed at 4 °C. After electrophoresis, the slides were washed (0.4 M Tris-HCl, pH 7.5; three washes of 5 min each), air-dried, and stained with ethidium bromide (2 μg ml–1) prior to scoring. For the neutral comet assay, the above procedure was followed with minor modifications. The slides were incubated in proteinase-K for 60 min at 37 °C and the use of TBE buffer. Olive tail moment was calculated to determine the DNA fragmentation on the experimental cells.26 Comets in the alkali assay revealed DNA single strand + double strand breaks and those in the neutral non-denaturing assay exhibited DNA dsb. The values were expressed as the mean ± standard error of mean with at least three independent experiments. Differences in the olive tail moment were compared using Dunnett's multiple comparison test based on mean values from individual experiments.

2.9. Mitotic index

To determine the mitotic index, after the experimental period, the cells were trypsinized and harvested. Then washed with ice cold PBS (pH 7.4) and re-suspended in 10 ml of pre-warmed hypotonic solution (0.075 M KCl) and kept at 37 °C for 10–15 min. The cells were then centrifuged and fixed in Carnoy's fixative (methanol : acetic acid, 3 : 1). The slides were prepared and stained with 4% Giemsa for 8–10 min. The frequency of mitotic cells per culture condition was determined by calculating the number of mitotic cells out of a population of 1000 consecutive cells, ignoring broken cells, clumped cells, and cellular debris. Polyploid cells (i.e. 8 N DNA mitotic cells) were included in the MI analysis.

2.10. Statistical analysis

The results are presented as mean ± S.D. of replicate analyses accompanied by the number of independent experiments. Statistical analyses were carried out using analysis of variance (ANOVA) by GraphPad Ver 5.03 statistical software and Microsoft Excel. Differences at p < 0.05 or less were considered statistically significant.

3. Results

3.1. Dose- and time-dependent effects of LDCD on breast cancer cells

Human breast cancer cells, MCF-7 (estrogen positive) and MDA-MB-231 (estrogen negative) were treated with different doses of DOX, ETOP and BUS to identify the lowest concentration that could initiate apoptotic events in the cells. The low-dose range for each chemotherapeutic drug was chosen based on our preliminary studies and the previous literature. Fig. 1 shows the viability of breast cancer cells treated with LDCD for 12 and 24 h. Treatment with low-dose concentrations of DOX (1 μM), ETOP (0.5 μM) and BUS (40 μM) for 12 h resulted in 90% of cell viability. At the maximum of a low-dose concentration of DOX (6 μM), ETOP (3 μM) and BUS (100 μM), we observed 80% of cell viability after 12 h. There was no remarkable change in the cell viability at low-doses of ETOP and BUS treated cells after 24 h. However, we observed 65% of cell viability after 24 h in cells treated with low-doses of DOX. From the results obtained, 1 and 2 μM of DOX, 0.5 and 1 μM of ETOP, and 40 and 80 μM of BUS were considered as ideal low-doses for the study and further experiments were carried out for an incubation time of 12 h. For recovery experiments, the cells were incubated under drug free conditions for 24 h after LDCD exposure.

Fig. 1. Response of MCF-7 and MDA-MB-231 cells to a range of low-dose chemotherapeutic drugs after 12 and 24 h. (A) Doxorubicin – 0, 1, 2, 3, 4, 5, and 6 μM; (B) etoposide – 0, 0.5, 1.0, 1.5, 2.0, 2.5, and 3.0 μM and (C) busulfan – 0, 20, 40, 60, 80, and 100 μM. Data expressed as mean ± S.D. of three independent experiments.

Fig. 1

Open in a new tab

3.2. Effect of LDCD on cellular morphology

The morphological assessment of cells subjected to LDCD treatment showed no gross change. Fig. S1† shows that higher doses of DOX (10 μM), ETOP (5 μM), and BUS (400 μM) induced cell shrinkage, accumulation of cell debris and detachment of cells from the monolayer indicating the occurrence of complete apoptosis/necrosis, while the LDCD treated cells showed a minimal number of apoptotic cells.

3.3. Effect of LDCD on apoptosis in breast cancer cell lines

Hoechst 33258/PI staining of experimental cells showed that DOX (1 and 2 μM) and ETOP (0.5 and 1 μM) treated cells had condensed nuclear material and bright blue fluorescence illustrating early apoptotic features (Fig. 2A), while BUS (40 and 80 μM) treated cells exhibited a comparatively lower condensation in both the cell lines. The high-dose treated cells showed a highly condensed and fragmented nucleus and signs of necrosis in both the cell lines with a more pronounced effect observed in ETOP (5 μM) treated cells.

Fig. 2. Hoechst 33258 and propidium iodide double staining of MCF-7 and MDA-MB-231 cells at 12 h incubation (A) with LDCD and after 24 h recovery (B). Apoptotic bodies (yellow arrow) and swollen enlarged cells typical for necrosis (purple) were observed in experimental cells (magnification – 400×).

Fig. 2

Open in a new tab

Repetition of dual staining in experimental cells after incubation in drug-free medium following LDCD treatment to check for events of recovery showed apoptotic characteristics in DOX (2 μM), ETOP (1 μM), and BUS (80 μM) treated cells while 0.5 μM ETOP and 40 μM BUS did not induce any apoptosis in both cell lines unlike cells before recovery (Fig. 2B). DOX (1 μM) treatment showed very minimal cells with apoptotic signs when compared to cells before recovery. The high-dose treated cells were observed to have typical necrotic characteristics.

3.4. Effects of LDCD on the clonogenic survival of breast cancer cells

Evaluation of enhanced survival ability acquired by breast cancer cells after initiation of apoptotic trigger and consequent formation of clones by employing the clonogenic survival assay showed a decrease of clonogenicity with the increase in dosage. Colonies were observed in all the LDCD treated cells. Higher doses of DOX (10 μM) and ETOP (5 μM) treated cells almost had no colonies, whereas BUS (400 μM) treated cells developed nearly 15% colonies in MCF-7 cell lines (Fig. 3A). However, the same doses resulted in the development of an increased number of colonies in MDA-MB-231 cell lines (Fig. 3B) compared to MCF-7.

Fig. 3. Influence of LDCD on the colony formation of MCF-7 (A) and MDA-MB-231 (B) cells. Data expressed as mean ± S.D. of three independent experiments. *p < 0.05, control versus drug-treated cells.

Fig. 3

Open in a new tab

3.5. Effect of LDCD on ROS generation

The extent of ROS generation during defective apoptosis in LDCD-treated cells was assessed using the DCFH-DA method. Fig. 4 shows that 2 μM DOX and 1 μM ETOP caused a significant fold change in ROS generation in both the cell lines compared to the control. BUS-treated cells had comparatively less ROS generated in both cell lines at all doses under investigation.

Fig. 4. Intracellular ROS generation of MCF-7 (A) and MDA-MB-231 (B) cells after LDCD treatment for 12 h. data expressed as mean ± S.D. of three independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001, control versus drug-treated cells.

Fig. 4

Open in a new tab

3.6. Effect of LDCD in triggering apoptotic cascade

To examine the alternations in the intracellular signaling molecules responsible for apoptosis, immunoblot analysis for caspase 3, 9, PARP, and β-actin (endogenous control) was performed. Decreased levels of cleaved caspase 9 and PARP were observed in the LDCD-treated MCF-7 cells compared to the cells exposed to higher doses (Fig. 5A). Upon cell recovery, mild expression of cleaved caspase-9 was observed in LDCD-treated cells (Fig. 5C) The ratio of cleaved caspase-9/procaspase-9 was significantly higher in DOX treated cells when compared to low-doses of ETOP and BUS during treatment which significantly reduced post recovery (Fig. 5B). There was no significant difference in the ratio of cleaved PARP/PARP within the LDCD treated cells (Fig. 5D). In the case of MDA-MB-231 cells, there was a mild to an increased expression of cleaved caspase-3, 9 and cleaved PARP (Fig. 5E) during 12 h treatment depending on the dose of drug and decreased expressions were observed in post recovery (Fig. 5F). The ratio of cleaved caspase/procaspase-3 and 9 was higher in DOX treated cells when compared to low-doses of ETOP and BUS (Fig. 5G and H), while the cleaved PARP/PARP ratio was decreased in DOX treated cells when compared to low-doses of ETOP and BUS (Fig. 5I). All cells produced pro-caspase 3, 9 and PARP in both treatment period and post-recovery.

Fig. 5. Effect of LDCD on the expression of apoptotic proteins in MCF-7 (A & B) and MDA-MB-231 (E & F) cells. Total protein from the cells treated with LDCD (A & E) and post recovery in drug-free medium (C & F), assessed for expression of apoptotic proteins. β-Actin – endogenous control. Densitometry analysis of cleaved caspase-3, 9 and PARP with full-length procaspase 3, 9 and PARP and relative changes are indicated (B, D, G, H & I). *p < 0.05, control versus drug-treated cells.

Fig. 5

Open in a new tab

3.7. Effect of LDCD on cell cycle distribution

Breast cancer cells exposed to LDCD were analysed for cell cycle perturbations using flow cytometry after staining the experimental cells with PI. Compared to the vehicle controls, DOX treatment showed a significant arrest of MCF-7 and MDA-MB-231 cells in the sub G1 and S phases of the cell cycle. In MCF-7, DOX treatment showed 28.13% cells in the S phase and 12% cells in the sub G1 phase at 1 μM (Fig. 6A and C). The extent of sub-G1 accumulation due to DOX treatment was more pronounced in MCF-7 cells than in MDA-MB-231 cells that exhibited G2-M arrest (24.44% cells) (Fig. 6B). Treatment with ETOP (1 μM) resulted in 22.14% cells in the S-phase and accumulation of cells in the G2-M phase was more pronounced at higher doses. MDA-MB-231 cells were least responsive to cell cycle arrest at low-doses of ETOP and BUS, with a maximum 13.96% and 15.83% of cells in the S phase and 34.31% and 37.84% of cells in the G2-M phase respectively when compared with control cells, which had 20.59% of cells in the S-phase and 30.96% of cells in the G2-M phase (Fig. 6D). There was no significant change in BUS-treated cells when compared to the control in both cell lines.

Fig. 6. Differences in the cell cycle distribution of MCF-7 (A & C) and MDA-MB-231 (B & D) cells exposed to LDCD. Cell cycle analysis by flow cytometry 12 h after LDCD treatment and percentage distribution of cells in the sub-G1, G1, S and G2-M phases from the resulting histograms and the mean values are shown.

Fig. 6

Open in a new tab

3.8. Effect of LDCD in altering mitotic index

Analysis of the influence of LDCD on cell division in breast cancer cells (Fig. 7) showed a non-significant decrease of the mitotic index after exposure of cells to low-doses of ETOP and BUS in both cell lines while low-doses of DOX caused a minimal decrease in the mitotic index in MDA MB-231 cells alone. There was a significant decreasing trend in the proliferation potential upon treatment with higher doses when compared to the control and cells treated with LDCD.

Fig. 7. Frequencies of the mitotic index in MCF-7 (A) and MDA-MB-231 cells (B) treated with LDCD. Data expressed as mean ± S.D. of three independent experiments. *p < 0.05, vehicle control versus drug-treated cells.

Fig. 7

Open in a new tab

3.9. Effect of LDCD on genomic integrity

The results of the comet assay showed that LDCD formed significant single and double strand DNA breaks in the experimental cells compared to the vehicle alone treated cells under both alkaline and neutral conditions (Table 1). However, LDCD-treated cells showed significantly lower strand breaks during the recovery experiment under the alkaline conditions. Under the neutral conditions, cells recovering after treatment with low-doses of DOX and ETOP showed higher incidence of strand breaks. BUS-treated cells had very minimal strand breaks after recovery.

Table 1. DNA damage evaluated under alkaline conditions (ssb + dsb) and under neutral conditions (dsb) in MCF-7 cells after 12 h of LDCD treatment (A) and post recovery for 24 h (B) a .

Treatment A B
(i) Alkaline condition (ssb + dsb)
Vehicle control 1.5 ± 0.4 1.7 ± 0.5
DOX
1 μM 9.1 ± 0.6* 6.5 ± 1.2
2 μM 13.0 ± 1.5* 8.2 ± 0.6*
10 μM 27.3 ± 1.0* 17.6 ± 2.6*
ETOP
0.5 μM 9.4 ± 0.8* 7.1 ± 0.5
1 μM 12.1 ± 0.9* 7.0 ± 0.8*
5 μM 22.8 ± 1.3* 17.5 ± 1.6*
BUS
40 μM 8.2 ± 0.4* 5.8 ± 0.4
80 μM 10.8 ± 0.6* 7.7 ± 0.9
400 μM 19.6 ± 0.4* 14.6 ± 1.4*
 
(ii) Neutral condition (dsb)
Vehicle control 0.9 ± 0.2 1.0 ± 0.2
DOX
1 μM 6.3 ± 1.0* 6.9 ± 0.9*
2 μM 11.2 ± 0.6* 5.5 ± 0.3*
10 μM 15.5 ± 2.0* 11.7 ± 1.2*
ETOP
0.5 μM 5.6 ± 0.2* 6.3 ± 1.4*
1 μM 8.5 ± 0.9* 6.8 ± 0.8*
5 μM 13.0 ± 1.2* 13.8 ± 1.2*
BUS
40 μM 5.0 ± 0.6* 2.9 ± 0.3
80 μM 8.8 ± 0.6* 4.9 ± 0.8
400 μM 14.5 ± 1.2* 13.8 ± 1.8*
Open in a new tab

aAll values are (mean of three independent experiments) ± (standard error of the mean).

4. Discussion

It is well established that chemotherapeutic agents induce tumor cell death by apoptosis. However, defects in the apoptosis pathway are known to play a crucial role in the chemoresistance and development of secondary cancers. Low-dose metronomic chemotherapy has been practiced as an alternative strategy due to the toxic side effects observed when anticancer drugs are administered at MTD to enhance therapeutic benefits in patients. It has been reported that low-doses of radiation or genotoxic compounds could turn on the protective apoptotic-mediated process, thus leading to mutations, neoplastic transformation, and secondary cancer. However, the mechanism has been far from clear. In this study, we have investigated the effect of low-dose chemotherapy and its impact on cancer cell survival. Earlier studies demonstrate that mutation prone clones defective in apoptosis escaped from cell death27,28 resulting in drug resistance and therapy-related cancer. Low-dose chemotherapy exposure to normal cells, such as endothelial and epithelial cells, has been explored previously.29,30 However, there is a lack of information on the same against the cancer cells and its elucidation is much warranted to identify the mechanism of prolonged survival of cancer cells. Thus, this study focussed on analysing changes in the survival of cancer cells upon treatment with LDCD using three widely used anticancer drugs, DOX, ETOP, and BUS, with different mechanisms of action against the two breast cancer cell lines.31

An initial determination of the least concentration of drug sufficient to induce the apoptotic trigger in the cells showed that the very minimal dose of DOX and ETOP used in this study had higher cytotoxicity compared to BUS. The difference of effect in the cell lines observed may be attributed to variation in the sensitivity of MCF-7 and MDA-MB-231 cells towards chemotherapeutic drugs.32,33 A brief exposure (12 and 24 h) of cells to all the three drugs did not affect the growth of monolayer cells. However, higher doses of all three drugs showed inhibitory effects. Thus, further experiments were restricted to an incubation period of 12 h owing to an insignificant change after 24 h incubation. The absence of any morphological change upon treatment with LDCD compared to higher doses suggested that the low concentration used in this study might have just triggered the initial molecular events in the cell which could consequently lead to apoptosis.

Though the chemotherapeutic drug DOX is an established apoptosis inducer that influences cell cycle progression and targets DNA integrity of the cell to induce apoptosis during classical chemotherapy,34,35 only fewer studies have focussed on the impact of DOX during the remission period. Analysis of the effect of chemotherapeutic drugs on cancer remission assessed by Hoechst 33258 and PI double staining and the clonogenicity assay showed that low-doses of DOX and ETOP induced many early morphological signs of apoptosis such as chromatin condensation and formation of apoptotic bodies, whereas low-dose of BUS showed mild or low response to apoptosis. The decrease in the number of apoptotic cells and the presence of significant number of colonies after the drug-free incubation in the LDCD-treated cells suggested that there is the possibility for cells to continue with their survival even after an initial trigger of apoptosis owing to the acquired ability of cancer cells to overcome cell death. Similar reports in the past showed that cancer cells acquired resistance to apoptosis activated by DNA-damaging chemotherapeutic agents36 and showed prolonged survival, a mechanism not completely understood. These observations demonstrated by LDCD-treated cells with respect to cell death and survival warrant a further study to delineate the underlying mechanism.

Further analysis of the cellular physiology showed a significant increase in ROS generation in cells treated with low-doses of DOX and ETOP similar to previous reports.34,37,38 ROS has been shown to be a key factor in controlling diverse cellular functions such as cellular transformation, proliferation, apoptosis, and necrosis.39 However, the exact role of ROS in these cellular functions still remains uncertain and controversial.

Research on the molecular pathway adapted by the LDCD to induce apoptosis involves analysis of changes in the expression of caspase cascade in the cells.40 Reports have shown that activated caspases lead to morphological hallmarks of apoptosis, including DNA fragmentation. The results of this study showed induction of cleaved caspase 3 by a low-dose of DOX (0.5 μg ml–1), there by implicating the activation of mitochondria-mediated apoptosis. The absence of a prominent band of cleaved caspase 3 in cells treated with ETOP and BUS at low-dose levels shows that the tested low-doses might have been sufficient enough to just activate early apoptotic events or cells might have adapted alternate mechanisms of cell death, such as senescence induced by BUS.41 The absence of execution of caspase expression in cells after drug-free periods implicated that the cells must have adapted mechanisms to repair the changes induced by the tested drugs. These observations suggest that LDCD treatment might cause defective apoptosis in the cells which could be overcome leading to survival during drug-free breaks.

Continuous treatment failure in the remission period and the results obtained from this study warranted further analysis of the cell cycle in experimental cells to elucidate the influence of LDCD on prolonged cell survival and therapy-related cancers. Moreover, cell cycle checkpoints aid in ensuring the accuracy of DNA replication and division and characterization of genomic instability leading to breast cancer progression.42–44 The results of flow cytometry analysis showed that DOX and ETOP at low-doses significantly accumulated more cells in the S and G2-M phases which implies that cell division could have halted to repair the DNA damage induced during drug treatment, while BUS does not alter the cell cycle distributions. Similar results of flow cytometry have been reported in previous studies with DOX, ETOP, and BUS.45,46 The mean mitotic index values were also decreased in the experimental cells compared to controls which could be due to slower progression of cells from the S (DNA synthesis) phase to the M (mitosis) phase of the cell cycle, as a result of LDCD exposure.

The occurrence of DNA strand breaks in the cells recovering from LDCD treatment can be attributed to the emerging consensus that induction of genomic instability could be the crucial early event in carcinogenesis, which enables an initiated cell to evolve into a cancer cell. Thus, the tested LDCD too might have induced subtle genomic alterations that might have not been recognized by the checkpoints of the cell enabling prolonged survival of defective cells.

Thus, the results of this study show a likely impairment in the induction of apoptosis and cell cycle progression in LDCD-treated cells. Mitotic indices of LDCD-treated cells exhibited a normal value compared to vehicle only cells, which suggest that low-doses of all three drugs might have triggered a defective apoptosis with an increase in ROS. Moreover, the defect in the cell cycle check points might have led the cells to escape from death and enter the mitosis stage leading to survival of cells with DNA damage. In conjunction with this view, DSBs were seen to persist in the cells treated with low-doses of DOX and ETOP after the recovery period which proves to be the probable population that can potentially be involved in tumor progression, possibly accumulating genomic variations and escaping apoptosis. However, further elucidation of the cellular mechanisms is essential to strategize an effective therapy in overcoming the current drawbacks and reduce the recurrence of tumors. In addition, future studies are necessary to clarify the extent of activation of apoptosis and epigenetic change, if any, involved in defective apoptosis and vulnerability to associated secondary malignancies.

In conclusion, this study provides evidence of low-dose chemotherapeutic effects on cancer cells. Traditional chemotherapeutic drugs kill cancer cells by activation of the apoptotic pathway. However, in clinical practice, the majority of human tumors, especially solid tumors, remain resistant to most therapeutic drugs, even when used in combination. It is becoming increasingly clear that the most important determinant of tumor resistance may be a generalized resistance to induction of apoptosis, rather than resistance based on specific alterations in the drug–target interaction. The overall results of this study reveal that LDCD initiates defective apoptosis. Furthermore, LDCD allows for survival of a small population of cells escaping from apoptosis possibly leading to genetic rearrangements which eventually result in secondary malignancies. Hence, it is important to evaluate the optimal chemotherapeutic and anti-angiogenic drug doses and scheduling characteristics in these tumors. Studies addressing the same would reveal more unresolved questions related to the underlying mechanisms in cancer progression and relapse and provide scope for the development of more efficient strategies for complete cure of cancer.

Conflict of interest

The authors declare that there are no conflicts of interest.

Supplementary Material

Click here for additional data file. (166B, txt)
Click here for additional data file. (3.9MB, pdf)

Acknowledgments

This work was supported by DST-SERC (SR/FT/LS-004/2009) fast track young scientist project DST-SERB (no. SB/EMEQ-082/2013) and ICMR (no. 54/15/CFP/GER/2011-NCD-II) Project India. Arun was supported by a CSIR Senior Research Fellowship 2013–2014. Dhivya was supported by Lady Tata Memorial Research Scholarship 2014.

Footnotes

†Electronic supplementary information (ESI) available. See DOI: 10.1039/c5tx00391a

References

  1. Yu H. K., Lee H. J., Yun S. J., Lee S. J., Langley R. R., Yoon Y., Yi L. S., Bae D. S., Kim J. S., Kim S. J. Transl. Oncol. 2014;7:368–376. doi: 10.1016/j.tranon.2014.04.005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Buckner J. C., Brown L. D., Cascino T. L., Gerstner J. B., Krook J. E., Westberg M. W., Wiesenfeld M., O'Fallon J. R., Scheithauer B. J. Neuro-Oncol. 1990;9:249–254. [PubMed] [Google Scholar]
  3. Fulda S., Debatin K. M. Oncogene. 2006;25:4798–4811. doi: 10.1038/sj.onc.1209608. [DOI] [PubMed] [Google Scholar]
  4. Kaufmann S. H., Earnshaw W. C. Exp. Cell Res. 2000;256:42–49. doi: 10.1006/excr.2000.4838. [DOI] [PubMed] [Google Scholar]
  5. Maddocks K., Ruppert A. S., Browning R., Jones J., Flynn J., Kefauver C., Gao Y., Jiang Y., Rozewski D. M., Poi M., Phelps M. A., Harper E., Johnson A. J., Byrd J. C., Andritsos L. A. Leuk. Res. 2014;38:1025–1029. doi: 10.1016/j.leukres.2014.05.011. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Chakraborty S., Tarantolo S. R., Batra S. K., Hauke R. J. Am. J. Clin. Oncol. 2013;36:132–142. doi: 10.1097/COC.0b013e3182438ddf. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Meadows A. T., Friedman D. L., Neglia J. P., Mertens A. C., Donaldson S. S., Stovall M., Hammond S., Yasui Y., Inskip P. D. J. Clin. Oncol. 2009;27:2356–2362. doi: 10.1200/JCO.2008.21.1920. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Wu J., Waxman D. J. Cancer Lett. 2014;353:272–280. doi: 10.1016/j.canlet.2014.07.033. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Morrison S. A., Hill S. L., Rogers G. S., Graham R. A. J. Surg. Res. 2014;192:235–241. doi: 10.1016/j.jss.2014.06.030. [DOI] [PubMed] [Google Scholar]
  10. Pasquier E., Kavallaris M., Andre N. Nat. Rev. Clin. Oncol. 2010;7:455–465. doi: 10.1038/nrclinonc.2010.82. [DOI] [PubMed] [Google Scholar]
  11. Wakabayashi T., Kawashima T., Matsuzawa Y. Multidiscip. Respir. Med. 2014;9:44. doi: 10.1186/2049-6958-9-44. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Palchaudhuri R., Hergenrother P. J. Curr. Opin. Biotechnol. 2007;18:8. doi: 10.1016/j.copbio.2007.09.006. [DOI] [PubMed] [Google Scholar]
  13. Geva-Zatorsky N., Dekel E., Cohen A. A., Danon T., Cohen L., Alon U. Cell. 2010;140:643–651. doi: 10.1016/j.cell.2010.02.011. [DOI] [PubMed] [Google Scholar]
  14. Evan G. I., Vousden K. H. Nature. 2001;411:342–348. doi: 10.1038/35077213. [DOI] [PubMed] [Google Scholar]
  15. Reed J. C. J. Clin. Oncol. 1999;17:2941–2953. doi: 10.1200/JCO.1999.17.9.2941. [DOI] [PubMed] [Google Scholar]
  16. Simonian P. L., Grillot D. A., Nunez G. Blood. 1997;90:1208–1216. [PubMed] [Google Scholar]
  17. Budman D. R., Berry D. A., Cirrincione C. T., Henderson I. C., Wood W. C., Weiss R. B., Ferree C. R., Muss H. B., Green M. R., Norton L., Frei 3rd E. J. Natl. Cancer Inst. 1998;90:1205–1211. doi: 10.1093/jnci/90.16.1205. [DOI] [PubMed] [Google Scholar]
  18. Qiu Q. C., Hu B., He X. P., Luo Q., Tang G. H., Long Z. F., Chen Z. C., He X. S. Genet. Mol. Biol. 2012;35:18–26. doi: 10.1590/s1415-47572012005000009. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Luanpitpong S., Chanvorachote P., Nimmannit U., Leonard S. S., Stehlik C., Wang L., Rojanasakul Y. Biochem. Pharmacol. 2012;83:1643–1654. doi: 10.1016/j.bcp.2012.03.010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Christmann M., Tomicic M. T., Roos W. P., Kaina B. Toxicology. 2003;193:3–34. doi: 10.1016/s0300-483x(03)00287-7. [DOI] [PubMed] [Google Scholar]
  21. Nakada S., Katsuki Y., Imoto I., Yokoyama T., Nagasawa M., Inazawa J., Mizutani S. J. Clin. Investigation. 2006;116:80–89. doi: 10.1172/JCI25716. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Iwamoto T., Hiraku Y., Oikawa S., Mizutani H., Kojima M., Kawanishi S. Cancer Sci. 2004;95:454–458. doi: 10.1111/j.1349-7006.2004.tb03231.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Mertins S. D., Myers T. G., Holbeck S. L., Medina-Perez W., Wang E., Kohlhagen G., Pommier Y., Bates S. E. Mol. Cancer Ther. 2004;3:849–860. [PubMed] [Google Scholar]
  24. Zhou H., Zheng C., Zou G., Tao D., Gong J. Int. J. Biochem. Cell Biol. 2002;34:678–684. doi: 10.1016/s1357-2725(01)00176-5. [DOI] [PubMed] [Google Scholar]
  25. Singh N. P., McCoy M. T., Tice R. R., Schneider E. L. Exp. Cell Res. 1988;175:184–191. doi: 10.1016/0014-4827(88)90265-0. [DOI] [PubMed] [Google Scholar]
  26. Kumaravel T. S., Jha A. N. Mutat. Res. 2006;605:7–16. doi: 10.1016/j.mrgentox.2006.03.002. [DOI] [PubMed] [Google Scholar]
  27. Yan B., Wang H., Peng Y., Hu Y., Zhang X., Chen Q., Bedford J. S., Dewhirst M. W., Li C. Y. Proc. Natl. Acad. Sci. U. S. A. 2006;103:1504–1509. doi: 10.1073/pnas.0507779103. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Zhivotovsky B., Kroemer G. Nat. Rev. Mol. Cell Biol. 2004;5:752–762. doi: 10.1038/nrm1443. [DOI] [PubMed] [Google Scholar]
  29. Bello L., Carrabba G., Giussani C., Lucini V., Cerutti F., Scaglione F., Landre J., Pluderi M., Tomei G., Villani R., Carroll R. S., Black P. M., Bikfalvi A. Cancer Res. 2001;61:7501–7506. [PubMed] [Google Scholar]
  30. Bocci G., Nicolaou K. C., Kerbel R. S. Cancer Res. 2002;62:6938–6943. [PubMed] [Google Scholar]
  31. Mross K., Steinbild S. J. Cancer Ther. Res. 2012;1:1–32. [Google Scholar]
  32. Aroui S., Ram N., Appaix F., Ronjat M., Kenani A., Pirollet F., De Waard M. Pharm. Res. 2009;26:836–845. doi: 10.1007/s11095-008-9782-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Park Y. Y., Jung S. Y., Jennings N. B., Rodriguez-Aguayo C., Peng G., Lee S. R., Kim S. B., Kim K., Leem S. H., Lin S. Y., Lopez-Berestein G., Sood A. K., Lee J. S. Carcinogenesis. 2012;33:1843–1853. doi: 10.1093/carcin/bgs167. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Kotamraju S., Konorev E. A., Joseph J., Kalyanaraman B. J. Biol. Chem. 2000;275:33585–33592. doi: 10.1074/jbc.M003890200. [DOI] [PubMed] [Google Scholar]
  35. Bian X., McAllister-Lucas L. M., Shao F., Schumacher K. R., Feng Z., Porter A. G., Castle V. P., Opipari Jr. A. W. J. Biol. Chem. 2001;276:48921–48929. doi: 10.1074/jbc.M108674200. [DOI] [PubMed] [Google Scholar]
  36. Masui K., Gini B., Wykosky J., Zanca C., Mischel P. S., Furnari F. B., Cavenee W. K. Carcinogenesis. 2013;34:725–738. doi: 10.1093/carcin/bgt086. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Rojas E., Mussali P., Tovar E., Valverde M. BMC Cancer. 2009;9:398. doi: 10.1186/1471-2407-9-398. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Tyurina Y. Y., Serinkan F. B., Tyurin V. A., Kini V., Yalowich J. C., Schroit A. J., Fadeel B., Kagan V. E. J. Biol. Chem. 2004;279:6056–6064. doi: 10.1074/jbc.M309929200. [DOI] [PubMed] [Google Scholar]
  39. Pan J. S., Hong M. Z., Ren J. L. World J. Gastroenterol. 2009;15:1702–1707. doi: 10.3748/wjg.15.1702. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Alnemri E. S., Livingston D. J., Nicholson D. W., Salvesen G., Thornberry N. A., Wong W. W., Yuan J. Cell. 1996;87:171. doi: 10.1016/s0092-8674(00)81334-3. [DOI] [PubMed] [Google Scholar]
  41. Probin V., Wang Y., Zhou D. Free Radicals Biol. Med. 2007;42:1858–1865. doi: 10.1016/j.freeradbiomed.2007.03.020. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Sherr C. J. Cancer Res. 2000;60:3689–3695. [PubMed] [Google Scholar]
  43. Richard S. M., Bailliet G., Paez G. L., Bianchi M. S., Peltomaki P., Bianchi N. O. Cancer Res. 2000;60:4231–4237. [PubMed] [Google Scholar]
  44. Senderowicz A. M., Sausville E. A. J. Natl. Cancer Inst. 2000;92:376–387. doi: 10.1093/jnci/92.5.376. [DOI] [PubMed] [Google Scholar]
  45. Luo G., Guan X., Zhou L. Cancer Biol. Ther. 2008;7:966–973. doi: 10.4161/cbt.7.6.5967. [DOI] [PubMed] [Google Scholar]
  46. Ritter C. A., Sperker B., Grube M., Dressel D., Kunert-Keil C., Kroemer H. K. Br. J. Pharmacol. 2002;137:1100–1106. doi: 10.1038/sj.bjp.0704972. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Click here for additional data file. (166B, txt)
Click here for additional data file. (3.9MB, pdf)

Articles from Toxicology Research are provided here courtesy of Oxford University Press

RESOURCES