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Table of Contents
ORIGINAL ARTICLE
Year : 2019  |  Volume : 6  |  Issue : 2  |  Page : 76-84

Modulation of oxidative stress by doxorubicin loaded chitosan nanoparticles


Department of Zoology, K. M. College, University of Delhi, Delhi, India

Date of Submission18-Jul-2018
Date of Decision23-Nov-2018
Date of Acceptance20-Dec-2018
Date of Web Publication31-May-2019

Correspondence Address:
Dr. Anita Kamra Verma
Department of Zoology, K. M. College, University of Delhi, Delhi - 110 007
India
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Source of Support: None, Conflict of Interest: None


DOI: 10.4103/JCRP.JCRP_18_18

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  Abstract 


Purpose of the Research: Chitosan nanoparticles (CHNP) are being used to modulate the generation of reactive oxygen species (ROS), as unwarranted generation of ROS can damage proteins, lipid membranes, and DNA of host cells. CHNP possess exceptional abilities to modulate antioxidants and suppress oxidative stress damage caused by the CHNP themselves in normal cells. Methods and Results: CHNP were prepared by ionic gelation in the size range of ~115 nm, with a polydispersity index of 0.365. Doxorubicin (DOX) was encapsulated in CHNP with entrapment efficiency ~48%. The modulation of free radicals and antioxidative enzymes by DOX-loaded CHNP (DLCHNP) was evaluated. The glutathione s-transferase and glutathione levels induced by DLCHNP were lower in Ehrlich ascites carcinoma cells(EACs) cells (6.60 ± 0.02 nM/min/mg protein and 0.92 ± 0.05 nM/min/mg protein, respectively) compared to void CHNP and DOX per se decreased levels of nitric oxide and superoxide dismutase (0.03 ± 0.001 nMoles and 28.84 ± 0.016 Unit/mg protein), elevated levels of GSSG (11.69 ± 0.004 nM/min/mg protein), marginally reduced levels of GSH reductase (1.87 ± 0.002 Unit/mg protein), reduced levels of GPx (31.35 ± 0.022 Unit/mg protein) and significantly enhanced levels of LPO (1.56 ± 0.01 nMoles/mg protein) indicated cellular damage. As observed in DNA fragmentation assay, void nanoparticles did not show any DNA damage whereas DLCHNP caused significant damage. Enhanced gene expressions of Cyt. C and p21 on EACs cells was observed in DLCHNP-treated cells compared to DOX per se. Conclusion: CHNP were not efficient in generating remarkable oxidative stress, but when coupled with a drug (i.e., DLCHNP) severe damage was caused to the cancer cells compared to the free drug. This indicated the potential of our encapsulated nanoparticles in drug delivery.

Keywords: Chitosan nanoparticles, DNA fragmentation, doxorubicin, Ehrlich ascites carcinoma, oxidative stress


How to cite this article:
Leekha A, Kumar V, Moin I, Gurjar BS, Verma AK. Modulation of oxidative stress by doxorubicin loaded chitosan nanoparticles. J Cancer Res Pract 2019;6:76-84

How to cite this URL:
Leekha A, Kumar V, Moin I, Gurjar BS, Verma AK. Modulation of oxidative stress by doxorubicin loaded chitosan nanoparticles. J Cancer Res Pract [serial online] 2019 [cited 2019 Jul 22];6:76-84. Available from: http://www.ejcrp.org/text.asp?2019/6/2/76/259489




  Introduction Top


Paradoxically, the depletion of antioxidants or increased production of reactive oxygen species (ROS)/reactive nitrogen species are reportedly the major sources of oxidative stress. ROS generally include the superoxide anion (O2•−), peroxide (O2−2.), hydroxyl radical (OH), and singlet oxygen (O2) excited forms of oxygen. Oxygen-derived radicals are constitutively present as an integral part of normal aerobic living and have extremely high chemical reactivity that may help to explain their damaging effects on cells. When produced in excess, ROS may damage proteins, lipids, and also DNA. If the damage is excessive, cell death can be induced.

Natural polymers such as cellulose, starch, chitosan, gelatin, and alginate have been extensively used in sustained drug delivery systems,[1],[2] and their acceptance as safe delivery vehicles is primarily owing to their biocompatibility and biodegradability.[3],[4],[5] Chitosan is a versatile natural polymer that has been used in many applications,[6],[7] and is commercially available in various forms and used for drug delivery because of its mucoadhesive properties.[8] Doxorubicin (DOX), an important member of the anthracycline family, is known to immediately regenerate its parent quinine by reducing oxygen to ROS such as hydrogen peroxide (H2O2) and the superoxide anion (O2•−). These free radicals are primarily responsible for the damage to DNA and cellular membranes.[9] DOX is a broad-spectrum anticancer drug, with toxicity against a variety of malignant cells. It has also been shown to induce toxicity in normal cells through the production of ROS through redox cycling reactions.[10],[11] Cancer cells not only escape immune surveillance but also evolve ways to survive, in diverse exogenous and endogenous oxidative circumstances. Even solid tumors are known to adapt to harsh conditions by developing antioxidant detoxifying mechanisms, using endogenous ROS for survival and proliferation. Oxidative stress results from an imbalance in the various levels of toxic ROS that may overcome endogenous antioxidant defenses leading to nonselective damage of tissues and organs in the host.[12] Minimizing oxidative stress may prevent cellular death, reduce inflammation, and prevent morbidity and mortality.[13] Many studies have suggested that ROS may exert contrasting cellular effects by promoting either cell proliferation and tumor progression or cell death and tumor regression. Hence, ROS can behave as a “double-edged weapon,” on the one hand, stimulating many other diseases and on the other hand as a therapeutic tool against cancer cells.[14] To induce ROS-associated tumor cell death, ROS levels must be maintained above a minimum threshold value. Therefore, therapy-triggered ROS generation to kill cancer cells specifically can be a forceful mechanism.[14] Anti-cancerous agents such as DOX that are frequently used in the treatment of several types of tumors work by targeting some of these apoptotic pathways. Consequently, it remains a challenge to evolve DOX-based therapy to generate “oxidative stress” in transformed cells and elucidate the mechanism of cell death. Therefore, the present study was undertaken to assess the (i) antioxidative activity, (ii) molecular mechanism of apoptosis, and (iii) DNA fragmentation by treatment with chitosan nanoparticles (CHNP) in an Ehrlich ascites carcinoma (EACs) cell line.


  Materials And Methods Top


DOX hydrochloride and chitosan 85% deacetylated were purchased from Sigma-Aldrich (St. Louis, MO, USA). Proteinase K, 1-chloro, 2, 4-dinitrobenzene (CDNB), and sodium bicarbonate were purchased from Sigma-Aldrich (Bengaluru, India) and paraformaldehyde was purchased from Merck India Ltd., (Mumbai, India). N-1-naphthyl ethylenediamine dihydrochloride (NEDD), ethylenediaminetetraacetic acid (EDTA), nitro-blue tetrazolium (NBT), trichloroacetic acid (TCA), and dinitrobenzoic acid (DTNB) were procured from SRL, India. All other chemicals were of analytical grade and obtained from Merck India Ltd. (Mumbai, India).

Determination of oxidative stress

EACs were maintained in RPMI-1640 supplemented with 10% fetal bovine serum. All of the cells were grown at 37°C in a humidified atmosphere of 5% CO2 in a CO2 incubator (Thermoscientific). The EACs were trypsinized at 80% confluency and centrifuged at 1500 rpm for 10 min. Cell pellets were washed twice in chilled PBS before the suspension for 30 min at 4°C in lysis buffer (10 mM Tris HCL (pH 7.4) and 250 mM sucrose) containing protease inhibitors (0.2 mM phenylmethylsulfonyl fluoride, 0.2 mM leupeptin, 0.3 mM aprotinin, 0.5 mM EDTA, and 0.1 mM pepstatin). After incubation, samples were sonicated and centrifuged at 10,000 rpm at 4°C for 30 min. The resulting supernatants were used for subsequent experiments.

Lipid peroxidation assay

The stock solution of TCA-TBA-HCl reagent containing 15% w/v trichloroacetic acid, 0.375% w/v thiobarbituric acid and 0.25 N hydrochloric acid was prepared. This solution was mildly heated to assist in the dissolution of the thiobarbituric acid. A 400 μl sample was vigorously mixed with 1600 μl of TCA-TBA-HCl. The solution was kept in a water bath at 100°C for 1 h. After cooling, the flocculent precipitate was removed by centrifugation at 3000 rpm for 10–15 min at room temperature (RT). The absorbance of the sample was read at 535 nm against a blank. Percentage (%) decrease in optical density (OD) was directly proportional to the decrease in the levels of lipid peroxidation (LPO). Malondialdehyde (MDA) levels (nM/mg protein) were calculated as per previously published protocols.[15]

Glutathione S-Transferase

Glutathione s-transferase (GST) activity was measured according to the method of Habig. The reaction was measured by observing the conjugation of CDNB with reduced glutathione (GSH). GST activity was measured spectrophotometrically using the substrate CDNB. This was done by observing an increase in absorbance at 340 nm. One unit of enzyme was defined as the amount required to conjugate 1 μmol of CDNB with reduced GSH per minute at 25°C. 20 μl of mitochondrial fraction and 180 μl of reaction cocktail were added to the final assay mixture of 200 μl per well. The absorbance was measured at 340 nm for 20 min against a blank containing phosphate buffer instead of the sample.[16]

Reduced glutathion assay

Reduced GSH estimation assay was performed according to the method of Moron et al. 1979. Briefly, 100 μl of mitochondrial fraction was precipitated with 20 μl of 5% trichloroacetic acid (TCA). The precipitated proteins were removed by centrifugation at 1200 rpm for 5 min at RT. A volume of 45 μl of mitochondrial fraction treated with TCA was added to the final assay mixture of 110 μl, 45 μl of sodium phosphate buffer (0.2 M, pH 8), and 20 μl of DTNB (10 mM) reagent. After 10 min, the absorbance was read at 412 nm against a blank containing TCA instead of the sample. The amount of reduced GSH was expressed as nM/min/mg protein of the sample.[17]

Glutathione peroxidase assay

The total GSH peroxidase activity was measured according to the method of Paglia and Valentine, 1967.[18] The reaction mixture contained 50 mM potassium phosphate buffer (pH 7), 1 mM EDTA, 1 mM sodium azide, 0.2 mM NADPH, 1 U GSH reductase (GR), and 1 mM reduced GSH. To the final assay mixture of 150 μl, 50 μl of mitochondrial fraction and 80 μl of the reaction mixture were added, and the reaction was initiated by adding 20 μl of 0.042% (w/w) H2O2 in each well of the 96-well plate. The absorbance was recorded at 340 nm for 5 min against a blank containing phosphate buffer instead of the sample. The values were expressed as μmol of NADPH oxidized to NADP using the extinction coefficient of 6.2 × 10/M/cm at 340 nm. The activity of GPx was expressed in terms of Units/mg protein of the sample.

Glutathione reductase assay

Glutathione reductase activity was measured according to the method of Miwa, 1972. GR is a ubiquitous enzyme that catalyzes the reduction of oxidized GSH (GSSG) to GSH (GSH). Glutathione reductase is essential for the GSH redox cycle that maintains adequate levels of reduced cellular GSH. For the Glutathione reductase assay, a cocktail was prepared by adding 2 ml of 9 mM of GSSG, 0.02 ml of 12 mM NADPH, and 2.6 ml of 1/15 M phosphate buffer pH 6.6. To the final assay mixture of 100 μl, 80 μl of cocktail and 20 μl of mitochondrial fraction were added per well. The activity of this enzyme was determined by monitoring the decrease in absorbance at 340 nm for 5 min against a blank containing phosphate buffer instead of the sample. The activity of GR was expressed in terms of μmol NADPH consumed/min/ml of the sample.[19]

Glutathione oxidized assay

GSSG levels in the samples were measured after derivation of GSH with 2-Vinylpyridine.[20] Briefly, with 500 μl samples, 1 μl of 2-vinylpyridine was mixed and incubated for 50–60 min at 37°C. The mixture was further centrifuged at 1500 g for 10 min after deproteination with 4% sulfosalicylic acid. The GSSG levels (nM/min/mg protein) were estimated in the supernatant after reaction with DTNB. The absorbance was recorded spectrophotometrically at 412 nm.

Super oxide dismutase assay

Superoxide dismutase activity was estimated according to the method of Kakkar et al., 1984. The reaction mixture consisted of 0.025 M sodium pyrophosphate (pH 8), 180 μM phenazine methosulfate and 300 μM NBT. To the final assay mixture of 100 μl, 20 μl of sample and 60 μl of reaction mixture were added, and the reaction was initiated by the addition of 20 μl of 780 μM NADH. The absorbance was measured kinetically at 560 nm for 5 min against a blank containing phosphate buffer instead of the sample. Superoxide dismutase (SOD) activity was expressed as Units/mg of protein.[21]

Nitric oxide assay

This assay was based on a diazotization reaction to detect nitric oxide (NO),[22] in which the Griess Reagent System is based on the chemical reaction between sulfanilamide and NEDD under acidic (orthophosphoric acid - 88% purity) conditions. Briefly, Reagent A-1% sulfanilamide in 2.5% orthophosphoric acid and Reagent B-0.1% NEDD in 2.5% orthophosphoric acid were mixed in equal amounts (1:1). To the final assay mixture of 200 μl, 100 μl of cell supernatent was mixed with Griess Reagent in an equal ratio of (1:1). The absorbance was read at 540 nm immediately against a blank containing phosphate buffer instead of the sample.

Immunocytochemistry

EAC cells were grown in a humidified CO2 incubator and plated on 12-mm round glass coverslips in 24-well cell culture plates. Cells were 60%–80% confluent and treated with DOX and DLCHNP for 24 h. The cells were then washed with PBS (pH 7.4) and fixed in 4% (v/v) paraformaldehyde for 5–10 min at RT. The cells were later permeabilized by 0.1% Triton-X in PBS for 10 min and again washed with PBS (pH 7.4). The cells were finally incubated with p21 and cytochrome C monoclonal goat antibody overnight at 4°C. An FITC-labeled secondary antibody was also added and left at RT for 2 h. The coverslips were then washed twice with PBS (pH 7.4) and mounted on glass slides using DPX and visualized in a fluorescent upright microscope (Nikon Eclipse 90i).

DNA fragmentation assay

The effects of void CHNP and DLCHNP on HEK and EAC cell lines by DNA fragmentation were determined as per a previously published protocol.[23] Briefly, HEK and EAC cells (1 × 106 cells/ml) were seeded in a 6-well plate and treated with different concentrations of DOX per se, void CHNP, and DLCHNP. The cells were trypsinized and pelleted at 1100 rpm for 10 min at RT. Cell pellets were resuspended in 500 μl PBS (pH 7.4), lysed by the addition of 60 μl lysis buffer (1 mM EDTA, 0.2% Triton X-100, 10 mM Tris HCl, pH 8.0) and treated with 2 μl ribonuclease A (20 mg/ml) at 56°C for 2 h, followed by digestion with 2 μl proteinase K (2.5 μg/ml) for 1 h at 37°C. This was further centrifuged at 12000 rpm for 30 min at RT, and the supernatant was collected. Finally, phenol-chloroform-isoamyl alcohol (25:24:1) extraction was performed, and DNA was precipitated with two volumes of 70% ethanol and one-tenth volume of 3 M sodium acetate (pH 4.8). The DNA pellets were then eluted in elution buffer (TE buffer, pH 8.0) and electrophoresed in 1.5% agarose gel stained with ethidium bromide, and DNA fragments were visualized under UV light.

Statistical analysis

The results were expressed as mean ± standard deviation. Comparisons among groups were analyzed using one-way ANOVA, and means were separated by Tukey's test using Prism (5.0) software (Prism Software Inc., CA, USA). Levels of significance were accepted at the ≤0.05 level.


  Results Top


We reported the preparation and characterization of CHNP in detail in our previous communication.[24] Briefly, CHNPs were ~115.4 nm with a polydispersity index (PDI) of 0.365 and a zeta potential + 19.5 ± 1.0. The morphology appeared round as observed by scanning electron microscopy (size ~150 nm) [Figure 1]a, [Figure 1]b, [Figure 1]c. The entrapment efficiency of DOX-loaded nanoparticles was around ~48% that is considered good for any cancer drug.
Figure 1: Physical characterization of chitosan nanoparticles. (a) The morphology of chitosan nanoparticles appeared round as observed in Scanning Electron Microscopy (size ~150 nm). (b) As measured by dynamic light scattering, the size was ~115.4 nm with a polydispersity index of 0.365 and (c) Zeta potential of + 19.5 ± 1.0

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The current study further extended our previous study by evaluating the free radicals and ROS generated by treatment of cells with DOX per se, CHNP, and DLCHNP. LPO is an important determinant when assessing cellular damage. LPO measured in terms of MDA levels observed in untreated (PBS) cells was 1.11 ± 0.005 nMoles/mg protein. An increased level was found in the DOX group (1.92 ± 0.007 nMoles/mg protein) followed by the DLCHNP group (1.56 ± 0.01 nMoles/mg protein) compared to the controls. However, the levels were only marginally increased in the CHNP (1.24 ± 0.003 nMoles/mg protein) group.

GSH is a family of five detoxifying enzymes that have a protective role in the survival of cells. The ratio of GSH S-transferase, reduced GSH exhibited a delicate balance indicating the oxidative state of cells. The GST level in untreated (PBS) cells was 8.44 ± 0.03 nM/min/mg of protein, compared to a higher level of 10.799 ± 0.03 nM/min/mg protein in the CHNP group. A marginal decrease in the level of GST was noted in the DOX per se treated cells (7.42 ± 0.02 nM/min/mg protein). However, DLCHNP-treated cells showed a decreased level (6.60 ± 0.02 nM/min/mg protein). The GSH level observed in untreated (PBS) cells was 6.95 ± 0.03 nM/min/mg of protein. Slightly decreased levels were found in the CHNP and DOX per se treated cells (5.31 ± 0.01 nM/min/mg and 5.27 ± 0.02 nM/min/mg, respectively). However, DLCHNP-treated cells exhibited a significantly lowered level (0.92 ± 0.05 nM/min/mg) compared to the controls. The observed level of GSSG in the cells treated with PBS was 9.46 ± 0.001 nM/min/mg protein. Enhanced levels were reported in all the experimental groups. The highest levels were observed in the DLCHNP group (11.69 ± 0.004 nM/min/mg protein) followed by the DOX group (11.07 ± 0.033 nM/min/mg protein). However, the CHNP-treated group showed only a marginal increase (11.99 ± 0.033 nM/min/mg protein) compared to the controls.

The level of GR in the cells treated with PBS was 2.34 ± 0.005 Units/mg/protein. A higher level (3.94 ± 0.004 Units/mg/protein) was found in the CHNP group compared to the controls. However, marginally decreased levels were noted in the DOX per se and DLCHNP treated cells (1.98 ± 0.005 Units/mg/protein and 1.87 ± 0.002 Units/mg/protein, respectively).

The level of GPx [Figure 2]c in the controls (PBS) was 67.70 ± 0.07 Units/mg/protein. The level in the CHNP group was similar (57.57 ± 0.11 Units/mg/protein); however, the levels were significantly lower in the DOX per se and DLCHNP groups (28.76 ± 0.06 Units/mg/protein and 31.35 ± 0.022 Units/mg/protein, respectively). The observed levels of GR [Figure 2]d in the cells treated with PBS were 2.34 ± 0.005 Units/mg/protein. Enhanced levels (3.94 ± 0.004 Units/mg/protein) were reported in CHNP group when compared to control. However, marginally decreased levels were manifested by DOX per se and DLCHNP treated cells, i.e., 1.98 ± 0.005 Units/mg/protein and 1.87 ± 0.002 Units/mg/protein, respectively.
Figure 2: Estimation of glutathione, glutathione s-transferase, glutathione reductase, glutathione peroxidase levels in cytosol in phosphate buffered saline, void chitosan nanoparticles, doxorubicin per se and doxorubicin-loaded chitosan nanoparticles-treated MCF- 7 cell lines (conc. 0.1 μg/ml). doxorubicin-loaded chitosan nanoparticles-treated cell induced low levels of glutathione and high levels of glutathione s-transferase. Glutathione peroxidase levels were significantly enhanced in the phosphate buffered saline controls followed by the void chitosan nanoparticles, doxorubicin per se and doxorubicin-loaded chitosan nanoparticles groups. Negligible glutathione reductase levels were observed in all of the groups. Data are expressed as mean ± standard error of mean (n = 3). *Indicates significant difference between doxorubicin and doxorubicin-loaded chitosan nanoparticles. * P =Probably significant)

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NO is a useful indicator of free radical generation. Several reports have indicated the dual effect of NO in cancer with regards to tumor growth and progression, as well as tumoricidal effects on certain cell types.[25],[26] The level of NO observed in untreated (PBS) cells was 0.10 ± 0.005 nMoles, and the highest level was found in the CHNP group (0.19 ± 0.007 nMoles) compared to the controls. A significantly lower level was observed in the DOX per se group (0.06 ± 0.003 nMoles), but an even lower level was noted in the DLCHNP group (0.03 ± 0.001 nMoles).

SOD activity was also an indicator of induced oxidative stress in all of the experimental groups. In PBS-treated cells, the level of SOD was 49.05 ± 0.027 Units/mg protein, which was similar to the level in the CHNP group (47.39 ± 0.001 Unit/mg protein). However, the levels were lower in the other two groups (DOX per se 35.60 ± 0.04 Unit/mg protein; and DLCHNP 28.84 ± 0.016 Unit/mg protein) [Figure 3]. The levels were significantly lower in the DLCHNP group.
Figure 3: Estimation of (a) superoxide dismutase and (b) Nitric oxide (NO) in phosphate buffered saline, void chitosan nanoparticles, doxorubicin per se and doxorubicin-loaded chitosan nanoparticles-treated cells (conc. 0.1 μg/ml). The lowest superoxide dismutase level in the cytosol was found in the doxorubicin-loaded chitosan nanoparticles-treated cells, followed by doxorubicin per se, chitosan nanoparticles and phosphate buffered saline treatment. In the nitric oxide assay, a minimum loss of membrane integrity was observed in doxorubicin-loaded chitosan nanoparticles -treated cells, with a maximum loss in the void chitosan nanoparticles-treated cells. Data are expressed as mean ± standard error of mean (n = 3). *Indicates significant difference between doxorubicin and doxorubicin-loaded chitosan nanoparticles. **P < 0.01,

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Expression of Cyt C and p21 in Ehrlich ascites carcinoma cells

The expressions of Cyt. C and p21 in the EAC cells were significantly increased (~three times) in DLCHNP-treated cells compared to the DOX per se group as observed by fluorescent immunocytochemistry [Figure 4].
Figure 4: Immunocytochemistry of Cyt C and p21 in Ehrlich ascites carcinoma cells. Deposition of Cyt C and p21 protein were highly expressed (~three times) in doxorubicin-loaded chitosan nanoparticles treated cell line than doxorubicin per se group

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DNA fragmentation assay

DNA damage was observed in both HEK and EAC cell lines by DNA fragmentation assay [Figure 5]. DOX per se and DLCHNP caused enhanced damage compared to void CHNP in both cell lines. Furthermore, the EAC cells were susceptible to DLCHNP, but negligible damage was observed in noncancerous cells, thereby indicating the potential role of nanoparticles in inhibiting the growth of cancer cells.
Figure 5: DNA fragmentation observed in HEK and Ehrlich ascites carcinoma cells treated with chitosan nanoparticles per se, doxorubicin, doxorubicin-loaded chitosan nanoparticles and incubated for 24 h. Ladder used: 250–25K bp. doxorubicin-loaded chitosan nanoparticles treated Ehrlich ascites carcinoma cells showed the characteristic DNA fragmentation indicating apoptotic cells. Slight fragmentation was observed in the group treated with doxorubicin per se Negligible cytotoxicity was observed by void chitosan nanoparticles clearly indicating the potential of doxorubicin-loaded chitosan nanoparticles on growth inhibition of cancer cells. The non-cancerous cell line (HEK) was apparently unaffected by the treatment

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  Discussion Top


In the present study, CHNP was prepared using the ionotropic gelation method[27] using sTPP as a cross-linker to achieve high drug entrapment efficiency. The hydrodynamic radius of nanoparticles was estimated by dynamic light scattering, and the size of the CHNP was ~115.4 nm with a PDI of 0.365 [Figure 1]. The surface charge of the nanoparticles was + 19.5 mV ± 1.0 as measured by zeta sizer. DOX was physically entrapped in the nanoparticles by bath sonication, with a calculated entrapment efficiency of ~48%. Since the formation of nanoparticles is a result of the interaction between the negative groups of sTPP and the positively charged amino groups of chitosan in acetic acid, it was imperative to evaluate the toxicity in terms of long-term use. We earlier reported negligible cytotoxicity by void CHNP after 24 h which gradually increased to ~35% in 72 h.

Chitosan is a multifunctional hydrogel polymer and is thus expected to have minimal interactions with cell membranes. However, CHNP reacted with the cells and induced pro-oxidant effects via intracellular ROS generation involving mitochondrial respiration and activation of NADPH-like enzyme systems.[28] Unfortunately, the antioxidant activity of chitosan is inversely proportional to the concentration and molecular weight of chitosan due to intricate intramolecular and intermolecular hydrogen bonds.[29],[30]

Any imbalance in the equilibrium between oxidant and endogenous anti-oxidant defense mechanisms can induce oxidative stress.[31],[32] The enhanced antioxidant capacity of tumor cells, caused by consistently high intracellular levels of antioxidants (e.g., GSH, vitamin E) or ROS detoxifying enzymes (e.g., superoxide dismutase), are important reasons for the resistance to ROS-generating treatments.[33],[34] Therefore, the present study was conducted to investigate the extent of ROS generation in cancer cell by a free drug (DOX) compared to the drug entrapped in the nanoparticles to possibly reduce serious side effects. Various biochemical assays were performed on EAC that were treated with void CHNP, DOX per se and DLCHNP to examine changes in the oxidant-antioxidant status of the cancer cells.

MDA is a widely used biomarker of oxidative stress[35] reflecting the extent of cell damage caused by LPO.[36] In the current study, significantly higher MDA levels were observed in the DOX per se and DLCHNP-treated groups compared to the controls, indicating higher oxidative stress. However, we found a negligible increase in MDA levels in the void CHNP that did not cause any significant damage to the cells, thereby confirming its biocompatibility.

An increase in ROS or decrease in free radical scavengers such as GSH is thought to be toxic to tumor cells. These findings are consistent with our experimental results in that decreased GSH levels were noted in all of the experimental groups. This decrease may have been due to increased levels of lipid oxidation products that have been reported to be associated with reduced availability of NADPH requisite for the activity of GSH reductase (GR) to convert GSSG to GSH [Figure 2].[37] Depletion of total GSH and decreased GSH/GST ratio is considered to be indicators of oxidative stress in various types of cancer.[38] We also observed low GSH levels and a significant increase in GST levels induced by treatment with DLCHNP, indicating that the induced GST participated in the detoxification and limited the oxidative damage. Several publications have reported abnormally high SOD activity in transformed cells.[39],[40] A decrease in SOD activity could lead to the accumulation of superoxide free radicals in cells, that may ultimately cause cell injury or cell death. In the present study, a significant decrease in SOD activity was observed in all of the treated groups, with a maximum decrease in the DLCHNP group. The depletion in SOD activity may have been due to the saturation of free radicals produced by the treatment with DLCHNP. Several studies have suggested that NO has potential mutagenic and carcinogenic activity in cervical cancer at significantly higher concentrations.[25],[26] In contrast, tumoricidal properties have also been reported in the treatment of various types of cancer.[41],[42],[43] We found a decrease in NO levels [Figure 4] in the DOX per se and DLCHNP-treated cells. This may be related to an alteration in pro-oxidant-antioxidant potential.

Taking into account the increased levels of MDA and highly diminished levels of the enzymes involved in the redox cycle, i.e., GR, GPx, GST, it would appear that the antioxidant defense mechanism of the cells may not be able to efficiently counteract the oxidative stress induced on exposure to free DOX and DLCHNP in EAC cancer cells.[44] Moreover, reduced SOD levels [Figure 3] compared to the control group further confirmed the impairment in anti-oxidant defense in the transformed cells. In addition, the DLCHNP-treated cells showed a significant decrease in SOD levels compared to the DOX group, representing their pivotal role in cancer therapeutics. Enhanced SOD activity can assist in scavenging excessive ROS.[45] A mechanistic approach can be used to suppress the growth of tumor cells through modulation of the oxidant/anti-oxidant equilibrium of cancer cells.

In our experiments, we observed a completely different trend in the levels of oxidants and anti-oxidants with the CHNP-treated cells with respect to DOX per se and DLCHNP. Levels of MDA were similar to the controls; however, SOD levels were marginally reduced in the CHNP group, suggesting the absence of oxidative stress. In contrast to the GSH levels observed in the other two groups (DOX per se and DLCHNP groups), we observed low GSH levels and negligible changes in GSSG levels, but increased GR and GST levels and marginally decreased GPx levels in the CHNP group. GSH peroxidase is a seleno-enzyme found in the cytoplasm of nearly all mammalian tissues, whose preferred substrate is H2O2. It serves as an antioxidant and catalyzes the reduction of harmful peroxides by GSH and protects cellular components against oxidative damage. Glutathione reductase further reduces GSSG to reduced GSH to complete the cycle. It is evident from our results that only CHNP were not efficient in generating remarkable oxidative stress and may have been unable to cause significant damage to cancer cells. However, when CHNP was coupled with a drug (i.e., DLCHNP) severe damage was noted to the cancer cells compared to the free drug. This indicated the potential of our encapsulated nanoparticles in drug delivery. Alternatively, oxidative stress has also been reported to be involved in the biological incompatibility of various polymers, which does not conform to the definition of biocompatibility.[46]

Cancer cells have enhanced levels of mitochondrial DNA mutations that may result in the accumulation of abnormal proteins of the electron transport chain leading to increased ROS production.[47],[48],[49] This enhanced ROS production further results in an increase in the number of mutations, oxidation of important proteins and other alterations finally culminating in cell death. Apoptosis or programmed cell death is crucial for the normal development and homeostasis of all multicellular organisms,[50] and it involves both extrinsic and intrinsic pathways. One of the most important stimuli in triggering intrinsic pathways is ROS. It has been reported that mitochondrion is the major site of ROS production and that the accumulation of ROS leads to the initiation of apoptosis.[51] H2O2 can cause the release of cytochrome c (marker protein for mitochondrial-associated apoptosis) and other pro-apoptotic factors from the mitochondria, often referred to as mitochondrial outer membrane permeabilization leading to the activation of caspases resulting in apoptosis.[52]

We used immunocytochemistry to assess the presence or absence of known pre-apoptotic proteins, p21 and Cyt. C [Figure 4]. In EAC cells, lower expressions of p21 and Cyt. C in DOX per se-treated cells were observed compared to DLCHNP. The maximum expressions of p21 and Cyt. C in DLCHNP-treated EAC cells indicated that apoptosis had been initiated in the cells. DOX, a broad spectrum anticancer drug, lacks tumor-targeting ability leading to reduced biodistribution and subsequently poor effectiveness.[53] Moreover, it has restricted transport through the cellular membrane leading to minimal drug internalization owing to the hydrophilic nature of DOX.[54] DOX is a DNA-intercalating agent that inhibits the progression of the enzyme topoisomerase II and stabilizes the topoisomerase II complex after it has damaged the DNA chain to progress to replication, thereby averting DNA resealing and finally obstructing the process of replication. The resulting DNA fragmentation leads to cell death.[55] [Figure 5] indicates the DNA fragmentation in EAC cells posttreatment with DLCHNP. There was no evidence of DNA damage in either void CHNP or DOX per se groups.


  Conclusion Top


CHNPs were successfully prepared using the ionotropic gelation method using sTPP as a crosslinker. The size of the CHNP was in the range of ~115 nm with a PDI of 0.345 and a zeta potential of + 19.5 ± 1.0 mV. Increased LPO and decreased antioxidant levels in the cancer cells indicated a state of oxidative stress. This imbalance in pro-oxidant–antioxidant status was observed in DLCHNP-treated cells and may possibly explain the cell death of these malignant cells. DLCHNP affected the differential cell death mechanism at the oncogene levels as indicated in EAC versus HEK cell lines. The expressions of two apoptotic proteins, Cyt. C and p21, were enhanced by approximately three times in the EAC line treated with DLCHNP compared to DOX per se, but there was no significant enhancement in the expressions of these proteins in the HEK cell line. DNA damage was assessed by performing DNA fragmentation assay on the EAC cells. Higher fragmentation of DNA was observed in the EAC cells treated with DLCHNP compared to the normal HEK cells [Figure 5].

The sustained activation of these signaling cascades has larger clinical implications. The induced redox imbalance caused by engineered nanoparticles can have adverse pathophysiological consequences resulting in inflammation, genotoxicity, fibrosis, and sometimes carcinogenesis. Understanding the cellular and molecular mechanisms of nanoparticle-induced oxidative stress may yield innovative approaches to assuage the toxicity of engineered nanoparticles. Furthermore, further studies to evaluate the oxidative potential of nanoparticles prior to their commercialization are necessary.

Acknowledgment

Ankita Leekha, Vijay Kumar, Imran Moin, and Bahadur Singh Gurjar are thankful to UGC, DBT, CSIR for their fellowship, respectively.

Financial support and sponsorship

Nil.

Conflicts of interest

There are no conflicts of interest.



 
  References Top

1.
Verma AK, Chanchal A, Maitra A. Co-polymeric hydrophilic nanospheres for drug delivery: Release kinetics, and cellular uptake. Indian J Exp Biol 2010;48:1043-52.  Back to cited text no. 1
    
2.
Senel S, Ikinci G, Kaş S, Yousefi-Rad A, Sargon MF, Hincal AA. Chitosan films and hydrogels of chlorhexidine gluconate for oral mucosal delivery. Int J Pharm 2000;193:197-203.  Back to cited text no. 2
    
3.
Kreuter J. Colloidal Drug Delivery Systems. New York: Marcel Dekker; 1994. p. 219-342.  Back to cited text no. 3
    
4.
Song C, Labhasetwar V, Cui X, Underwood T, Levy RJ. Arterial uptake of biodegradable nanoparticles for intravascular local drug delivery: Results with an acute dog model. J Control Release 1998;54:201-11.  Back to cited text no. 4
    
5.
Vinogradov SV, Bronich TK, Kabanov AV. Nanosized cationic hydrogels for drug delivery: Preparation, properties and interactions with cells. Adv Drug Deliv Rev 2002;54:135-47.  Back to cited text no. 5
    
6.
Agarwal S, Leekha A, Tyagi A, Kumar V, Moin I, Verma AK. Versatility of chitosan: A short review. J Pharm Res 2015;4:125-34.  Back to cited text no. 6
    
7.
Tyagi A, Agarwal S, Leekha A, Verma AK. Effect of mass and aspect heterogeniety of chitosan nanoparticle on bactericidal activity. Int J Adv Res 2014;2:357-8.  Back to cited text no. 7
    
8.
Takeuchi H, Yamamoto H, Niwa T, Hino T, Kawashima Y. Mucoadhesion of polymer-coated liposomes to rat intestine in vitro. Chem Pharm Bull (Tokyo) 1994;42:1954-6.  Back to cited text no. 8
    
9.
Gajewski E, Rao G, Nackerdien Z, Dizdaroglu M. Modification of DNA bases in mammalian chromatin by radiation-generated free radicals. Biochemistry 1990;29:7876-82.  Back to cited text no. 9
    
10.
Simůnek T, Stérba M, Popelová O, Adamcová M, Hrdina R, Gersl V. Anthracycline-induced cardiotoxicity: Overview of studies examining the roles of oxidative stress and free cellular iron. Pharmacol Rep 2009;61:154-71.  Back to cited text no. 10
    
11.
Menna P, Recalcati S, Cairo G, Minotti G. An introduction to the metabolic determinants of anthracycline cardiotoxicity. Cardiovasc Toxicol 2007;7:80-5.  Back to cited text no. 11
    
12.
Bulger EM, Maier RV. Antioxidants in critical illness. Arch Surg 2001;136:1201-7.  Back to cited text no. 12
    
13.
Christman JW, Blackwell TS, Juurlink BH. Redox regulation of nuclear factor kappa B: Therapeutic potential for attenuating inflammatory responses. Brain Pathol 2000;10:153-62.  Back to cited text no. 13
    
14.
Acharya A, Das I, Chandhok D, Saha T. Redox regulation in cancer: A double-edged sword with therapeutic potential. Oxid Med Cell Longev 2010;3:23-34.  Back to cited text no. 14
    
15.
Sekhara RD, Vrushabendra Swamy BM, Archana Swamy P. Protective effect of Cissampelos pareira linn. on paracetamol induced nephrotoxicity in male albino rats. Res J Pharm Biol Chem Sci 2012;3:695-705.  Back to cited text no. 15
    
16.
Habig WH, Pabst MJ, Jakoby WB. Glutathione S-transferases. The first enzymatic step in mercapturic acid formation. J Biol Chem 1974;249:7130-9.  Back to cited text no. 16
    
17.
Moron MS, Depierre JW, Mannervik B. Levels of glutathione, glutathione reductase and glutathione S-transferase activities in rat lung and liver. Biochim Biophys Acta 1979;582:67-78.  Back to cited text no. 17
    
18.
Paglia DE, Valentine WN. Studies on the quantitative and qualitative characterization of erythrocyte glutathione peroxidase. J Lab Clin Med 1967;70:158-69.  Back to cited text no. 18
    
19.
Miwa S. Hemotology. Mod Med Technol 1972;3:306-10.  Back to cited text no. 19
    
20.
Griffith OW. Determination of glutathione and glutathione disulfide using glutathione reductase and 2-vinylpyridine. Anal Biochem 1980;106:207-12.  Back to cited text no. 20
    
21.
Kakkar P, Das B, Viswanathan PN. A modified spectrophotometric assay of superoxide dismutase. Indian J Biochem Biophys 1984;21:130-2.  Back to cited text no. 21
    
22.
Griess P. Bemerkungen zu der abhandlung der H.H. Weselsky und Benedikt “Ueber einige azoverbindungen.” Chem Ber 1879;12:426-8.  Back to cited text no. 22
    
23.
Jena P, Mohanty S, Mallick R, Jacob B, Sonawane A. Toxicity and antibacterial assessment of chitosan-coated silver nanoparticles on human pathogens and macrophage cells. Int J Nanomedicine 2012;7:1805-18.  Back to cited text no. 23
    
24.
Agarwal S, Fatima S, Verma AK.In vitro antineoplastic potential of Chitosan nanoparticle against cervical cancer. IJPRPS 2014;3:444-56.  Back to cited text no. 24
    
25.
Naidu MS, Suryakar AN, Swami SC, Katkam RV, Kumbar KM. Oxidative stress and antioxidant status in cervical cancer patients. Indian J Clin Biochem 2007;22:140-4.  Back to cited text no. 25
    
26.
Beevi SS, Rasheed MH, Geetha A. Evidence of oxidative and nitrosative stress in patients with cervical squamous cell carcinoma. Clin Chim Acta 2007;375:119-23.  Back to cited text no. 26
    
27.
Verma AK, Pandey RP, Chanchal A, Sharma P. Immuno-potentiating role of encapsulated proteins of infectious diseases in biopolymeric nanoparticles as a potential delivery system. J Biomed Nanotechnol 2011;7:63-4.  Back to cited text no. 27
    
28.
Verma AK, Kumar V, Agarwal S, Leekha A, Tyagi A, Moin I, et al. Interplay of immune response and oxidative stress induced by Chitosan nanoparticle to possibly combat inflammation. World Res J Biosci 2013;1:28-34.  Back to cited text no. 28
    
29.
Feng T, Du YM, Li J, Hu Y, Kennedy JF. Enhancement of antioxidant activity of chitosan by irradiation. Carbohyd Polym 2008;73:126-32.  Back to cited text no. 29
    
30.
Xing R, Liu S, Guo Z, Yu H, Wang P, Li C, et al. Relevance of molecular weight of chitosan and its derivatives and their antioxidant activities in vitro. Bioorg Med Chem 2005;13:1573-7.  Back to cited text no. 30
    
31.
Vishal RT, Sharma S, Mahajan A, Bardi GH. Oxidative stress: A novel strategy in cancer treatment. JK Sci 2005;7:1-3.  Back to cited text no. 31
    
32.
Carmody RJ, Cotter TG. Signalling apoptosis: A radical approach. Redox Rep 2001;6:77-90.  Back to cited text no. 32
    
33.
Choi HJ, Jang YJ, Kim HJ, Hwang O. Tetrahydrobiopterin is released from and causes preferential death of catecholaminergic cells by oxidative stress. Mol Pharmacol 2000;58:633-40.  Back to cited text no. 33
    
34.
Hour TC, Chen J, Huang CY, Guan JY, Lu SH, Hsieh CY, et al. Characterization of chemoresistance mechanisms in a series of cisplatin-resistant transitional carcinoma cell lines. Anticancer Res 2000;20:3221-5.  Back to cited text no. 34
    
35.
Cini M, Fariello RG, Bianchetti A, Moretti A. Studies on lipid peroxidation in the rat brain. Neurochem Res 1994;19:283-8.  Back to cited text no. 35
    
36.
Surapaneni KM, Venkataramana G. Status of lipid peroxidation, glutathione, ascorbic acid, Vitamin E and antioxidant enzymes in patients with osteoarthritis. Indian J Med Sci 2007;61:9-14.  Back to cited text no. 36
[PUBMED]  [Full text]  
37.
Sarkar S, Yadav P, Trivedi R, Bansal AK, Bhatnagar D. Cadmium-induced lipid peroxidation and the status of the antioxidant system in rat tissues. J Trace Elem Med Biol 1995;9:144-9.  Back to cited text no. 37
    
38.
Dalle-Donne I, Rossi R, Colombo R, Giustarini D, Milzani A. Biomarkers of oxidative damage in human disease. Clin Chem 2006;52:601-23.  Back to cited text no. 38
    
39.
Hu Y, Rosen DG, Zhou Y, Feng L, Yang G, Liu J, et al. Mitochondrial manganese-superoxide dismutase expression in ovarian cancer: Role in cell proliferation and response to oxidative stress. J Biol Chem 2005;280:39485-92.  Back to cited text no. 39
    
40.
Taniguchi N, Ishikawa M, Kawaguchi T, Fujii J, Suzuki K, Nakata T. Expression of Mn-superoxide dismutase in carcinogenesis. Tohoku J Exp Med 1992;168:105-11.  Back to cited text no. 40
    
41.
Shang ZJ, Li JR. Expression of endothelial nitric oxide synthase and vascular endothelial growth factor in oral squamous cell carcinoma: Its correlation with angiogenesis and disease progression. J Oral Pathol Med 2005;34:134-9.  Back to cited text no. 41
    
42.
Li LM, Kilbourn RG, Adams J, Fidler IJ. Role of nitric oxide in lysis of tumor cells by cytokine-activated endothelial cells. Cancer Res 1991;51:2531-5.  Back to cited text no. 42
    
43.
Lechner M, Lirk P, Rieder J. Inducible nitric oxide synthase (iNOS) in tumor biology: The two sides of the same coin. Semin Cancer Biol 2005;15:277-89.  Back to cited text no. 43
    
44.
Radu M, Munteanu MC, Petrache S, Serban AI, Dinu D, Hermenean A, et al. Depletion of intracellular glutathione and increased lipid peroxidation mediate cytotoxicity of hematite nanoparticles in MRC-5 cells. Acta Biochim Pol 2010;57:355-60.  Back to cited text no. 44
    
45.
Naka K, Muraguchi T, Hoshii T, Hirao A. Regulation of reactive oxygen species and genomic stability in hematopoietic stem cells. Antioxid Redox Signal 2008;10:1883-94.  Back to cited text no. 45
    
46.
Wattamwar PP, Mo Y, Wan R, Palli R, Zhang Q. Antioxidant activity of degradable polymer poly (troloxester) to suppress oxidative stress injury in the cells. Adv Funct Mater 2010;20:147-54.  Back to cited text no. 46
    
47.
Kang D, Hamasaki N. Mitochondrial oxidative stress and mitochondrial DNA. Clin Chem Lab Med 2003;41:1281-8.  Back to cited text no. 47
    
48.
Carew JS, Huang P. Mitochondrial defects in cancer. Mol Cancer 2002;1:9.  Back to cited text no. 48
    
49.
Spitz DR, Sim JE, Ridnour LA, Galoforo SS, Lee YJ. Glucose deprivation-induced oxidative stress in human tumor cells. A fundamental defect in metabolism? Ann N Y Acad Sci 2000;899:349-62.  Back to cited text no. 49
    
50.
Gorman A, McGowan A, Cotter TG. Role of peroxide and superoxide anion during tumour cell apoptosis. FEBS Lett 1997;404:27-33.  Back to cited text no. 50
    
51.
Murphy MP. How mitochondria produce reactive oxygen species. Biochem J 2009;417:1-3.  Back to cited text no. 51
    
52.
Indran IR, Tufo G, Pervaiz S, Brenner C. Recent advances in apoptosis, mitochondria and drug resistance in cancer cells. Biochim Biophys Acta 2011;1807:735-45.  Back to cited text no. 52
    
53.
Hofland KF, Thougaard AV, Sehested M, Jensen PB. Dexrazoxane protects against myelosuppression from the DNA cleavage-enhancing drugs etoposide and daunorubicin but not doxorubicin. Clin Cancer Res 2005;11:3915-24.  Back to cited text no. 53
    
54.
Madhusudhan A, Reddy GB, Venkatesham M, Veerabhadram G, Kumar DA, Natarajan S, et al. Efficient pH dependent drug delivery to target cancer cells by gold nanoparticles capped with carboxymethyl chitosan. Int J Mol Sci 2014;15:8216-34.  Back to cited text no. 54
    
55.
Reinert KE. Anthracycline-binding induced DNA stiffening, bending and elongation; stereochemical implications from viscometric investigations. Nucleic Acids Res 1983;11:3411-30.  Back to cited text no. 55
    


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