Skip to main content
PMC home page
Toxicology Research logoLink to Toxicology Research
. 2018 Feb 6;7(2):156–171. doi: 10.1039/c7tx00269f

Genotoxicity inhibition by Syzygium cumini (L.) seed fraction and rutin: understanding the underlying mechanism of DNA protection†

Mohammad Shavez Khan a, Faizan Abul Qais a, Iqbal Ahmad a,✉, Afzal Hussain b, Mohamed F Alajmi b
PMCID: PMC6062347  PMID: 30090571

graphic file with name c7tx00269f-ga.jpgConsidering the ethnopharmacological importance of Syzygium cumini's seed and the lack of information on the antimutagenic and DNA-protecting mechanisms, a fraction-based study was conducted.

Abstract

Considering the ethnopharmacological importance of Syzygium cumini's seed and the lack of information on the antimutagenic and DNA-protecting mechanisms, a fraction-based study was conducted. Four different (hexane, chloroform, ethyl acetate, and aqueous) fractions were obtained from the sequential extraction of the methanolic extract of the seed. The most active antioxidant fraction (ethyl acetate) contained significant amount of phenolics and flavonoids. LC-qTOF-MS analysis of the ethyl acetate fraction revealed the presence of rutin, myricetin, naringin, cuscohygrin, and epoxycarryophyllone as constituent phytocompounds. The ethyl acetate fraction (100 μg ml–1) and a selected compound (rutin, 40 μg ml–1) showed remarkable decrease in the revertants frequency range from 74–77% and 66–84%, respectively, against both the mutagens (sodium azide (NaN3) and methyl methane sulfonate (MMS)) in the Salmonella typhimurium tester strains. All the statistical analyses were at a significance level of 0.05 between the different treatment groups. Moreover, the underlying mechanism of antimutagenicity using different treatment regime for rutin was explored. MMS-mediated DNA fragmentation and oxidation in lymphocytes were also shown to be decreased significantly when treated with the ethyl acetate fraction and rutin. Oxidative damage to pBR322 plasmid DNA was also reduced when incubated with different concentration of the ethyl acetate fraction and rutin. Biophysical (UV, fluorescence, ITC, etc.) and computational methods were employed to obtain a closer look at the DNA–rutin interaction. The data obtained clearly revealed that the ethyl acetate fraction exhibited promising antimutagenic and DNA-protective activity and its flavonoid constituents, including rutin, contribute significantly to the observed activity.

1. Introduction

In living cells, oxidative stress resulting from a physiological imbalance between the generation of reactive oxygen species (ROS) and the cellular machinery (including various antioxidant enzymes) to scavenge them can severely affect the structural integrity of macromolecules, including proteins, lipids, and DNA.1 Among other deleterious effects, ROS are also found to be closely associated with genotoxicity and mutagenicity and may give rise to complex pathophysiological conditions.2 The mechanism of ROS-induced DNA damage has been well characterized and shown to proceed via multiple pathways.3 Moreover, several genotoxic chemicals interact directly with DNA, proceeding through chemical modification of the native structure of DNA at a molecular level. Certain alkylating agents, such as methylmethane sulfonate (MMS), ethylmethane sulfonate (EMS), and methylnitrosourea (MNU), alter the primary structure of DNA molecules at specific positions and form intermediates or DNA adducts.4 The DNA adducts, when remain unattended or when the cellular repair machinery fails to remove these modifications, as in the case of compromised physiological conditions, could result in DNA damage, mutations, and the blocking of replication.5 Therefore nutritional supplements having antioxidant activity and DNA-protective potentials are necessary to overcome the deleterious effects of these oxidative processes to maintain genetic stability in the cell.

Dietary and traditionally used medicinal plants are known to possess great pharmacological importance, attributed to the presence of diverse phytochemical constituents. These plant-based chemical compounds convey their biological effects either by acting exclusively or synergistically, through one or more different mechanisms.6

Syzygium cumini L. Skeels (Syn: Eugenia jambola Lam. or Eugenia cuminii Druce.) commonly known as Jamun, black plum, or Indian blackberry, belongs to the family Myrtaceace and are large evergreen tropical trees native to the Indian sub-continent, but nowadays found naturalized throughout a major part of world.7 Different parts of the plants, including the leaves, barks, roots, and fruits, are used in various herbal formulations for the treatment of various disorders as diuretic, stomachic, carminative, hypo glycemic, antidiarrheal, and antidysentric.8 Laboratory investigations have provided information about the antioxidant efficacy of seeds9 along with other bioactivities, including anti-inflammatory,10 detoxifier,11 antimicrobial,12 gastro protective,13 protection against radioactivity,14 and antidiabetic;15 however, their genoprotective abilities against oxidation and alkylating damage still need to be systematically explored. Moreover, research involving fraction-based phytochemical screening and assigning the mode(s) of action to the constituents in a complex mixture (extract) is still warranted.

Thus, our aim was to obtain the most antioxidant active fraction of Syzygium cumini seed's extract and one of its representative compounds, like rutin, and to test then for their antigenotoxic potential through various approaches, such as Ames's test, Comet assay, and plasmid protection assay. Further biophysical insights of DNA–rutin interaction were also determined for obtaining a better understanding of the protective effect.

2. Materials and methods

2.1. Bacterial strains and chemicals

Salmonella typhimurium TA100 and TA102, both histidine auxotrophic mutants, were originally provided by Prof. B. N. Ames (Berkley, USA) and maintained in our laboratory. Gallic acid, ascorbic acid, quercitin, ethidium bromide (EB), and RPMI 1640 were procured from HiMedia Pvt. Ltd, India. Mutagens sodium azide (NaN3) and methyl methanesulfonate (MMS) were purchased from SRL Pvt. Ltd, India. 1,1-Diphenyl-2-picryl hydrazyl (DPPH), DNA sodium salt from calf thymus (ctDNA), Hoechst 33258, rutin standard, acridine orange (AO), histopaque 1119, and sodium phosphate monobasic salt were purchased from Sigma-Aldrich (USA). pBR322 plasmid DNA was purchased from Geni, Bangalore, (India). Methanol and acetonitrile of HPLC grade were obtained from Fisher Scientific UK. All the other chemicals and reagents were of analytical grade.

2.2. Plant material and extraction method

Fruits of Syzygium cumini (L.) were purchased from the local market in Aligarh in June and July. Subsequent identification of the plant material was done in the Department of Botany, Aligarh Muslim University, and a voucher specimen (AGM-JS-05/15) was submitted to the Department of Agricultural Microbiology, Aligarh Muslim University. The pulp was carefully removed and the seeds were washed in tap water and shade-dried for 20 days. The crude methanolic extract was prepared by soaking 500 g of dried and ground seed powder in 2.5 L of methanol with intermitting shaking for 5 days.16 The methanolic extract (10% w/w) was filtered and concentrated to dryness under reduced pressure using a rotatory evaporator. Further, the methanolic extract was dissolved in hot distilled water and successively extracted in n-hexane, chloroform, and ethyl acetate.17 Each fraction was concentrated under reduced pressure to give: a hexane fraction (HJs: 5.88% w/w), chloroform (CJs: 2.2% w/w), ethyl acetate (EAJs: 17.57%), and finally an aqueous layer (AJs: 54.55%). The obtained fractions were stored at 4 °C in refrigerator until further use. To achieve the desire concentrations, the fractions were reconstituted in different solvents or in the minimum amount of DMSO (0.5% v/v final concentration).

2.3. Qualitative phytochemical analysis

Preliminary phytochemical investigations of the fractions were performed using standard protocols to identify the chemical constituents, including the alkaloids, terpenoids, tannins, phenols, flavonoids, glycosides, and saponins, as described by Ahmad and Beg18 and Harborne.19

2.4. Estimation of the total phenolic and flavonoid contents

The total phenolic contents of the fractions were assessed by using Folin–Ciocalteau reagent as described by Zahin et al.16 The total phenolic content was expressed as gallic acid equivalents (mg per g dry weight). Estimation of the total flavonoid content was carried out using the aluminum chloride method, as described by Chang et al.20 The standard curve for flavonoids estimation was made using quercetin. The results were expressed in quercetin equivalents in mg per gram of dry weight (QE mg per g dry weight).

2.5. Antioxidant assays

The DPPH radical scavenging action of the different fractions of Syzygium cumini seed's extract was determined using the method of Brand–Williams et al.21 Each fraction's concentration exhibiting 50% inhibition (IC50) was obtained by plotting the fraction's concentration and percent inhibition. Low IC50 values represented a higher scavenging potential. The reduction potentials of the test samples were evaluated by measuring the absorbance of Perl's Prussian Blue complex formed by the reduction of ferricyanide in a stoichiometric excess in comparison to an antioxidant using the method of Oyaizu,22 as described by Gulcin.23 The efficacy of each fraction against lipid peroxidation was measured using the ammonium thiocyanate method, as described by Takao et al.24

2.6. LC-qTOF/MS analysis

The ethyl acetate fraction (EAJs) was subjected to LC-qTOF-MS analysis for identification of the phytoconstituents. For chromatographic separation, an Agilent 1290 infinity UPLC system was used and the samples were separated on a C18 column (Agilent Technologies, USA). The following elution program was applied at a flow rate of 0.5 ml min–1, where eluent A was 0.1% formic acid (v/v) and eluent B was 90% acetonitrile plus 0.1% formic acid: 0–28 min A 95% and B 5%, 33 min A 5% and B 95%, 35 min A 95% and B 5%, and 40 min A 95% and B 5%. The mass data were acquired on an MS Q-TOF mass spectrophotometer (Agilent, component model 6550) with the samples introduced automatically through the UPLC system. Spectra were recorded using a quadrupole time-of-flight (Q-TOF) unit with the following parameters: ion source: dual AJS ESI, MS Min range: 102 (m/z), MS Max range: 1000 (m/z), MS/MS scan rate: 1 spectra per second, VCap: 3500, Nozzle voltage: 1000 V, Fragmentor voltage: 175 V, Skimmer potential: 65 V. The stream parameters were as follows: Gas temperature: 250 °C, Gas flow: 13 l min–1, Nebulizer: 35 psi, Sheath gas temperature: 300, Sheath gas flow: 11 l min–1. The obtained mass spectra were processed using Agilent MassHunter Q-TOF B.05.01 (B5125.1) software.

2.7. Quantification of rutin in the EAJs fraction

Ultra-performance liquid chromatography (UPLC) equipped with a photodiode array detector (PDA) was used for the analysis and quantification of rutin in the ethyl acetate fraction of a methanolic extract of Syzygium cumini seed. The UPLC-PDA Agilent Technologies (1290 Infinity) machine was equipped with an infinity binary pump (G4220A), autosampler (G4226A), thermostat column compartment (G1316C), and photodiode array detector (G4212A). The above configuration of UPLC-PDA was programmed with Chem Station software. The elution of rutin was performed on an Eclipse C18 analytical column (4.6 × 100 mm, 3.5 μm) (Agilent, California, USA). The mobile phase consisted of solvent A, which was 100% acetonitrile, and solvent B, which was 100% water. The stepwise gradient was 10–90% B (0 min), 30–70% B (0–2 min), 35–65% B (2–5 min), 40–60% B (5–7 min), and 100% B (7–10 min) and the post time was 2 min. Finally, the rutin was eluted at a retention time (Rt) of 4.1 min. This developed UPLC-PDA method for rutin was applied for the qualitative and quantitative estimation of rutin in the EAJs.

2.8. DN-protection activity

2.8.1. Ames Salmonella histidine point mutation assay (Ames's test)

The antimutagenic activity of the EAJs fraction was determined using the method of Maron and Ames, 198325 with some modification as described earlier.16 Using a pre-incubation method, an aliquot of 0.1 ml of different concentrations of the EAJs fraction (25, 50, 100 μg per plate) reconstituted in a minimum amount of DMSO was mixed with 0.1 ml mutagen (NaN3 1 μg per plate or MMS 1.5 μg per plate) and incubated at 37 °C for 30 min. After incubation, 0.1 ml of bacterial culture (107 CFU ml–1) was added to the mixture, followed by the addition of top agar (0.5% NaCl, 0.6% agar, 0.5 mM Histidine-Biotine). After vortexing, the combined solution was poured on minimal media glucose plates (50 ml 40% glucose solution, 20 ml Vogel-Bonner medium, 20 g agar and 930 ml water). The plates were incubated for 48 h at 37 °C, after which the histidine-independent revertant colonies were scored. To assess the toxicity of the tested samples, parallel controls were included with different concentrations of the sample (EAJs fraction) and tester strains (TA 100 and TA 102). The non-toxicity of the tested concentrations was characterized from well-developed lawn, with no significant difference in the number of spontaneous revertant colonies between the control (without the test sample) and test plates (with the test sample). Scoring of the revertant colonies was carried out in triplicate plates for each concentration. The antimutagenic activity of the fraction was expressed in percent inhibition in the revertant frequency.

2.8.2. Determination of the antimutagenic mechanism of action of the compound

The antimutagenic activity and simultaneous toxicity assessment of the pure compound at different concentrations (20 and 40 μg per plate) were also evaluated using the above-described pre-incubation method. Additionally, to get enhanced mechanistic insight into the antimutagenic activity, the following method was adopted and briefly modified, as described in the literature.26

All components simultaneously (co-incubation): 0.1 ml of mutagen (NaN3 1 μg per plate or MMS 1.5 μg per plate), 0.1 ml of the compound (20 or 40 μg per plate), and 0.1 ml cell suspension (TA 100 or TA 102) were mixed simultaneously and incubated at 37 °C for 30 min. After the addition of soft agar and proper vortexing, the solution was poured onto minimal media glucose plates.

All components then the mutagen (pre-treatment): 0.1 ml of the compound (20 or 40 μg per plate) was mixed with 0.1 ml bacterial cell suspension (TA 100 or TA 102) and incubated at 37 °C for 30 min. Afterwards, 0.1 ml of mutagen (NaN3 1 μg per plate or MMS 1.5 μg per plate) was added and the whole content was poured onto minimal glucose plates.

2.8.3. Comet assay

First, the lymphocytes were isolated from the heparinized blood of a healthy volunteer by venipuncture. The blood was the diluted in normal saline followed by isolation of the lymphocytes with the help of histopaque using the standard protocol, and the cells were cultured in RPMI 1640 prior to the treatment. Single cell alkaline gel electrophoresis was performed according to the method of Singh et al.27 with minor changes. First layering of the frosted microscopic slides was done using 1.0% normal melting agarose (NMA) one day before the experiment to solidify it completely. Isolated lymphocytes were treated with different concentrations of EAJs fraction (50 and 100 μg ml–1) and rutin (20 and 40 μg ml–1) in the absence and presence of MMS, and then incubated at physiological temperature for 3 h. The treated cells were mixed with an equal volume of 0.5% low melting point agarose (LMPA), and 100 μl of cell suspension was overlaid on the base layer. Coverslips were paced to uniformly spread the cells, and slides were placed over an ice-pack to allow it to solidify, followed by a final layering with 0.5% LMPA. Slides were placed in an electrophoretic tank to immerse in freshly prepared cold lysis solution (pH 10.0) at 4 °C for 1 h for lysis of the cells. Washing of the slides was performed three times with normal saline and then the DNA of the lymphocytes was allowed to unwind in electrophoretic buffer (pH 13) for 30 min. The electrophoresis was performed at 300 mA current for 30 min at a field strength of 0.74 volts per cm. The slides were then neutralized by placing in neutralizing buffer for 10 min and then washed with saline. Finally, the slides were stained for 5 min with 100 μl ethidium bromide (20 μg ml–1) for 5 min and again washed to remove the excess of amount of stains. Coverslips were placed over the slides and these were then stored at 4 °C. The scoring was performed with Komet 5.5 software attached to an Olympus fluorescent microscope (Olympus Co., Japan) with an integrated camera (COHU, San Diego, USA). Finally, 25 cells from each replicate slide were scored and the data for DNA damage is presented here in in terms of the average tail length.

The study complied with all institutional and national guidelines. The experiment involving the use of human blood was approved by the Bio-ethical Committee of the Faculty of Agricultural Sciences, AMU, India (BE/FAgSc/01/18). Written informed consent from blood donor (Mr Mohammad Shavez Khan) was obtained.

2.8.4. Intracellular ROS quenching

DCF-DA dye was employed to measure the intracellular ROS activity induced by MMS. Lymphocytes were pre-incubated with 10 μM DCF-DA for 1 h at 37 °C and then centrifuged at 3500g for 5 min. The pellets of the lymphocytes were washed and suspended in normal saline to give a 5% (v/v) cell suspension. The suspension was then treated with different concentrations of EAJs fraction (50 and 100 μg ml–1) and rutin (20 and 40 μg ml–1) in the absence and presence of MMS and incubated at physiological temperature for 15 min. The fluorescence intensity was recorded on a spectroflourometer with excitation and emission filters set at 485 and 525 nm, respectively, and expressed in percent relative fluorescence.

2.8.5. Plasmid nicking assay

The protective ability of the EAJs fraction and rutin against oxidative DNA damage was evaluated by plasmid nicking assay using supercoiled pBR322 DNA using the method of Lee et al.28 Plasmid DNA (0.5 μg) was exposed to Fenton's reagent (30 mM H2O2, 50 μM ascorbic acid and 80 μM FeCl3) with different concentrations of EAJs fraction (25–100 μg) in a final reaction volume of 20 μl made up by adding Tris buffer. The reaction mixture was then incubated at 37 °C for 30 min. Thereafter, the samples were mixed with 1% tracking dye (0.25% bromophenol blue in 50% glycerol) and loaded onto 1% agarose gel (0.5 g of agarose in 50 ml 1× TBE buffer), and finally electrophoresis was performed followed by ethidium bromide staining and visualization.

2.9. Biophysical analysis

2.9.1. UV-Vis analysis

UV-vis absorption studies were carried out using a UV-1800 instrument (Shimadzu, Japan) at room temperature. Briefly, an increasing concentration of rutin (10–50 μM) was titrated to a fixed amount (40 μM) of DNA and the absorption spectra were recorded in 230–350 nm. The experiment was performed in Tris-HCl buffer (pH 7.4) and baseline correction was done using the same buffer.

2.9.2. Dye displacement assay

Dye displacement assays are preformed to investigate the precise binding biding site of any ligand to DNA using a site-specific probe whose binding site is already well established. The experiment was performed on a spectrofluorometer using specific dyes (both intercalator and minor groove binders). For the ethidium bromide (EB) displacement experiment, a solution containing DNA (30 μM) and EB (5 μM) was excited at 476 nm and the emission spectra was recorded in the range of 480 nm to 700 nm. To this solution, rutin was titrated (10–90 μM) and any change in the fluorescence emission signal was monitored. A similar experiment was performed for an acridine orange (AO) displacement assay in which the fluorescence emission spectra of the DNA–AO complex (330 μM and 5 μM respectively) was recorded in the 490–700 nm range by exciting at 480 nm followed by titration of the same concentration of rutin as mentioned above. Similarly, in a Hoechst 33258 displacement assay, the DNA–Hoechst complex was excited at 343 nm and the fluorescence emission spectra were recorded from 350 nm to 700 nm. The concentration of DNA and Hoechst 33258 were 30 μM and 5 μM, while rutin was varied from 10 μM to 90 μM. In the DAPI displacement experiment, the DNA–DAPI complex was excited at 338 nm and the emission was recorded in the range 350–600 nm. The concentration of DNA and DAPI were 30 μM and 5 μM, respectively, while rutin was titrated to 90 μM at 10 μM intervals.

2.9.3. DNA melting study

A DNA melting experiment was performed to examine the effect of rutin on the thermal stability of DNA. In the experiment, the optical density of DNA (50 μM) was recorded at temperature intervals of 5 °C ranging from 25 °C to 100 °C at 260 nm. In a similar way, the absorbance of DNA was recorded for the rutin–DNA complex (50 μM each). The ratio of absorbance at a temperature of absorbance at 25 °C (A/A25) was plotted as a function of temperature and the transition midpoint of this curve gave the melting temperature of the DNA.

2.9.4. Isothermal titration calorimetric analysis

ITC was carried out using a VP-ITC instrument (Microcal Inc., Northampton, MA) at 25 °C. Prior to loading, the DNA, rutin, and reference buffer were degassed thoroughly in a thermovac (140 mbar, 15 min). Rutin (1250 μM) was introduced into the sample cell by means of a syringe and the amount of each injection was set at 10 μl. DNA (50 μM) was maintained in the sample cell in 20 mM phosphate buffer (pH 7.4), while the reference cell contained only phosphate buffer. Then, 29 successive injections of rutin (10 μl each) were titrated for the sample cell containing DNA, with an initial delay of 60 s. The time duration for each injection was set to 20 s, with the spacing between two consecutive injections at 180 s. The reference power and the stirring speed were set at 16 μcal s–1 and 307 rpm, respectively. The calorimetric data were analyzed using MicroCal Origin 7.0 software provided with the instrument.

2.9.5. Molecular docking

Compounds detected by LC-qTOF-MS analysis were screened for DNA binding affinity. Their interaction mode with DNA was evaluated by in silico molecular docking using AutoDock-vina program,29 which uses the Broyden–Fletcher–Goldfarb–Shanno algorithm to search for the best binding site in terms of the binding energy, with this program reported to perform more accurate calculations. The three-dimensional (3D) structures of selected phytoconstituents (rutin, CID: 5280805) were obtained from ; https://pubchem.ncbi.nlm.nih.gov. Receptor files were downloaded in sdf format, which were later converted to the pdb format with Chimera 1.10.2. The flexible bonds were detected to allow flexibility in the ligand to guide the program to search for the best conformation, and the coordinate file was saved in the pdbqt format using MGL Tools-1.5.6. The 3D structure of B-DNA dodecamer d(CGCGAATTCGCG)2 was downloaded from the Protein Data Bank (PDB ID: ; 1BNA). With MGL Tools-1.5.6, the water molecules surrounding the receptor were deleted as they hinder proper docking. Merging of all the non-polar hydrogen molecules was done followed by the addition of Kollman charges. The size of the grid of receptor molecules was kept at 58 × 72 × 112 Å with the maximum spacing to cover entire molecule having a center of grid as x = 14.759, y = 20.984, z = 8.812. The output file was saved in the pdbqt format and other docking parameters were kept as default. The docked-out conformation with the lowest binding energy was selected for analysis, performed with PyMol and Discovery Studio 2016. For further verification of the in silico results, the docking studies were also performed using Hex 8.0 software (ESI 1†).

2.10. Statistical analysis

Data are expressed as the mean ± SD of three experimental replicates. Statistical significances were evaluated using IBM SPSS 22.0 for the Duncan Tukey test at a significance level of 0.05 and variance analysis by t-test in Microsoft excel (2013) for comparing the treated group with the untreated control group.

3. Results and discussion

3.1. Preliminary phytochemical analysis

Preliminary phytochemical investigation of the hexane (HJs), chloroform (CJs), ethyl acetate (EAJs), and aqueous (AJs) fractions of the Jamun seed methanolic extract showed positive results for the presence of alkaloids, terpenoids, tannins, phenols, flavonoids, and glycosides in the different fractions. The EAJs fraction was found to be positive for all the tested phytochemical classes, except saponins. The total phenolic content in the HJs (102.43 ± 8.28), CJs (173.06 ± 5.96), EAJs (409.87 ± 8.23), and AJs (195.02 ± 9.29) fractions were determined and expressed as gallic acid equivalents (mg g–1 of fractions). The total flavonoids content in the HJs (28.84 ± 3.07), CJs (372.69 ± 10.52), EAJs (476.29 ± 19.87), and AJs (67.94 ± 6.07) fractions were determined with respect to quercetin equivalents (mg g–1 of fractions).

3.2. Antioxidant activity

The free radical scavenging activity of the fractions were evaluated by DPPH assay and expressed in their respective IC50 values. The lowest IC50 value of the EAJs fraction (1.89 μg ml–1) was comparable to the standard antioxidant, ascorbic acid (1.17 μg ml–1). The IC50 values representing the antioxidant activity of the different fractions were in order EAJs > HJs > AJs > CJs, and were 1.89, 17.98, 24.54, and 27.50 μg ml–1, respectively. The observed relatively high radical scavenging activity of the ethyl acetate fraction (EAJs, IC50 = 1.89 μg ml–1) was in good accordance with the previous results as observed in different plants.30,31

The reductive potential of the different fractions were evaluated by potassium ferricyanide reducing assay at different concentrations and the results presented in Fig. 1A. A concentration-dependent increase in the reducing activity was observed in all the fractions. At the highest tested concentration (50 μg ml–1), the activity trend was observed as EAJs > AJs > CJs > HJs. Similarly, the EAJs fraction demonstrated the maximum diminution, signifying an effective peroxidation inhibitory potential in linoleic acid oxidation assay, as shown in Fig. 1B.

Fig. 1. Antioxidant activity of different fractions of Syzygium cumini's seed. (A) Ferric reducing power and (B) Linoleic acid oxidation inhibition.

Fig. 1

Open in a new tab

The findings reveal the broad spectrum of antioxidant activity of the EAJs fraction and suggest a correlation between the antioxidant activity and total phenolic/flavonoid content of the fractions. High phenolic and flavonoid contents were previously shown to be positively correlated with DPPH-based scavenging activity,32 ferric reduction,33 and a protective ability against primary and secondary products of lipid peroxidation.34

3.3. UPLC-qTOF-MS analysis EAJs fraction

Considering the high phenolic content and strong antioxidant activity of the EAJs fraction, it was subjected to UPLC-qTOF-MS for tentative identification of its phytoconstituents. As described in Table 1, five compounds, including rutin, myricetin, naringin, cuscohygrine, and epoxycarryophillone, along with other untargeted plant metabolites were detected by comparing the retention time, mass data and fragmentation, molecular formula, and corresponding [M–H]– ions with the database.

Table 1. Detected phytocompounds in UPLC-qTOF-MS analysis of EAJs fraction of Jamun's seed methanolic extract.

Retention time (tR) HR Mass (m/z) Formula Compound name Substance class
0.168 224.1889 C13H24N2O Cuscohygrine Alkaloid
13.643 580.1748 C27H32O14 Naringin Flavonoid
14.582 610.1513 C27H30O16 Rutin Flavonoid
15.265 318.0366 C15H10O8 Myricetin Flavonoid
17.981 236.1772 C15H24O2 Epoxycaryophyllanone Terpenoid
Open in a new tab

On the basis of the comprehensive protective activity of the EAJs fraction against free radicals, evidently by means of different antioxidant assays, in vitro evaluation of the fraction (EAJs) for antimutagenic and antigenotoxic activity was further undertaken. Out of the detected compounds, rutin was selected as a flavonoid representative. The role of rutin in the DNA-protection mechanism along with its quantification in the EAJs fraction was performed further. Previous studies by various workers demonstrated a close relationship between the antioxidant activity with the antimutagenic and antigenotoxic potential of plant extract and phytocompounds.35–39

3.4. Qualitative and quantitative analysis of rutin in the EAJs fraction

The identification of the rutin in EAJs fraction was determined by comparing their retention time with those of the standards of different concentrations under identical and optimized chromatographic conditions. The qualitative determination of rutin from plant extract was also determined by comparing the retention time with those of the standard. Analyte (rutin) detection from plant extract at the same retention time indicated the specificity of the developed UPLC-PDA method, as shown in Fig. 2. The quantitative analysis of the rutin in the fraction was carried out using identical and optimized chromatographic conditions using a calibration curve. The calibration curve of rutin was linear in the concentration range of 0.1–10 μg mL–1, with correlation coefficients of 0.9981 and an average recovery of 99.00%. The limits of detection (LOD) and limit of quantification (LOQ) were found to be 0.10 and 0.30 μgmL–1, respectively. The quantity of rutin in plant extract was found to be 480 ng mg–1 (w/w) of the dried weight extracts. The analysis confirmed the presence of rutin in the fraction in an appreciable amount.

Fig. 2. UPLC-PDA chromatogram for rutin (Rt at 4.1 min) in plant extract (A) using the identical conditions as in the case of the standard (B).

Fig. 2

Open in a new tab

3.5. DNA-protection studies

3.5.1. Antimutagenic activity

The ethyl acetate fraction was investigated for antimutagenic activities by Ames's Salmonella histidine revertant assay against direct acting mutagens (NaN3 and MMS). At the respective tested concentrations of the fraction (25, 50, and 100 μg per plate), no toxicity was observed against the S. typhimurium tester strains (TA100 and TA102). The data are presented in Table 2 showing the concentration-dependent increase percent inhibition of mutagenicity by the EAJs fraction against NaN3-induced mutagenicity. The EAJs fraction exhibited a significant decrease in mutagenicity percentage in TA 102 (76.13%), while a slightly lower antimutagenic response was observed in TA100 (74.60%) when tested at the highest dose per plate (100 μg). A dose-dependent inhibition of MMS mutagenesis was also shown by the EAJs fraction (Table 3) in both the tester strains, ranging from 35.57% to 77.13% and 29.81% to 74.60% for TA102 and TA100, respectively. Our results are in good accordance with previous reports, highlighting the antimutagenic activity of plant extracts containing an appreciable amount of phenolic/flavonoids.36,40,41 Results from the present study demonstrated for the first time that the ethyl acetate (EAJs) fraction can exert significant antimutagenic activity against oxidative (sodium azide) and alkylating (MMS) mutagenic chemicals.

Table 2. Effect of ethyl acetate fraction of Syzygium cumini's seed on the mutagenicity induced by sodium azide (NaN3) using Salmonella typhimurium strains.
Treatment Dose (μg per plate) Number of His+ revertants colonies/plate
TA 100 TA 102
Spontaneous 124.66 ± 7.17a 274.00 ± 11.04a
Positive control (NaN3) 1.5 1166.33 ± 22.86e 362.33 ± 14.16d
£Fraction 25 139.66 ± 11.59 289.00 ± 15.71
50 146.33 ± 16.56 305.33 ± 09.50
100 157.66 ± 19.03 319.66 ± 14.91
Fraction + mutagen 25 855.33 ± 19.22d (29.81%)*** 333.33 ± 15.28c (33.57%)
50 592.33 ± 16.81c (55.05%)*** 310.33 ± 06.94bc (59.71%)*
100 388.66 ± 12.2b (74.60%)*** 295.33 ± 08.04ab (77.13%)*
Open in a new tab
Table 3. Effect of ethyl acetate fraction of Syzygium cumini's seed on the mutagenicity induced by methyl methanesulphonate (MMS) using Salmonella typhimurium strains.
Treatment Dose (μg per plate) Number of His+ revertants colonies/plate
TA 100 TA 102
Spontaneous 124.66 ± 7.17a 274.00 ± 11.04a
Positive control (MMS) 1.00 978.00 ± 40.10e 1494.00 ± 37.34e
£Fraction 25 139.66 ± 11.59 289.00 ± 15.71
50 146.33 ± 16.56 305.33 ± 09.50
100 157.66 ± 19.03 319.66 ± 14.91
Fraction + mutagen 25 713.33 ± 22.88d (31.01%)* 1059 ± 27.90d (35.70%)**
50 493.66 ± 21.74c (56.75%)** 726.33 ± 22.93c (62.95%)***
100 324.00 ± 17.37b (76.64%)*** 558.33 ± 22.86b (76.71%)***
Open in a new tab

The different mechanism underlying the protective potential against mutagenic/carcinogenic chemicals by plant extracts and phytocompounds has been explained42 as including an inhibitory effect of the constituent phytochemicals on the mutagens or by interacting non-covalently with DNA molecules, which in turn restricts the DNA-oxidative intermediate reaction.43 Flavonoid enrich extracts were reported to exert enhanced antimutagenic activity. Ammar et al.44 showed distinct modulating effects from the flavonoid-rich extract of Rhamnus alaternus against direct acting mutagens. Similarly, Vattem et al.45 showed the synergistic activity of different flavonoids with phenolic juice of cranberry against the mutagenic potential of sodium azide and N-methyl-N′-nitro-N-nitrosoguanidine in Ames tests. Most of the phytocompounds detected in the EAJs fraction belong to the flavonoid subclass of the plant's secondary metabolites. Thus, we speculate that the antimutagenic activity of the EAJs fraction against direct acting mutagens could be attributed to its major polyphenolic constituents, primarily flavonoids.

To further augment this hypothesis and to explore the mechanism of rutin in antimutagenic action, the selected compound (rutin) previously identified and quantified in the EAJs fraction was subjected to antimutagenicity assay. Methodological variations in addition to the conventional method (pre-incubation) were adapted to obtain a detailed mechanistic preview, and the results are depicted in Tables 4 and 5. No sign of mutagenicity was observed at the tested concentrations (20 and 40 μg per plate) in either of S. typhimurium strains. With the conventional pre-incubation method (at the highest tested concentration), the percent inhibition was observed as 67.61% and 66.28% against NaN3 and 73.39% and 71.41% against MMS-induced mutagenesis in TA100 and TA102, respectively. Rutin when co-incubated with the mutagens and tester strains showed strong inhibition in revertant frequency. At the highest tested concentration, the percent inhibition of NaN3 mutagenesis was recorded as 73.47% and 71.21% for TA100 and TA102, respectively. Similarly, with MMS-induced mutagenesis, the percent inhibition in revertant frequency was 75.97% and 77.25% for TA100 and TA102, respectively. However, on adding mutagens after incubation of the bacterial cultures along with the different concentrations of the rutin, the decrease in revertant frequencies were the highest recorded (Tables 4 and 5). These results are in accordance with the antimutagenic activity of plant flavonoids reported previously.43,46,47 However, this study is probably the first to describe the differential action of rutin during different treatment regimes, and thus provides an enhanced understanding of its antimutagenic potential against both the mutagens NaN3 and MMS. The antimutagenic properties of flavonoids and other phenolics are largely ascribed to their ability to inhibit the mitochondrial enzymes P450, critically responsible for the physiological activation of indirect mutagens and carcinogens.48 Neutralization of the oxidative intermediates that are responsible for DNA damage is also considered as one of the protection mechanism based on the free radical scavenging feature of flavonoids.49 Additionally, the complexation of DNA and protective molecules (flavonoids) via covalent/non-covalent interactions is also considered responsible for the protection against DNA damage.50 Recently, it has been corroborated utilizing 3D pharmacophore modeling and molecular docking that certain plant phenolics, including rutin, interact with DNA and mask the primary amines from undergoing alkylation. Further, it was also deduced that rutin competes with different DNA-reactive intermediates produced by monofunctional alkylating agents (ethyl methanesulphonate), and has a greater binding affinity.51 However, the highest decrease in revertant frequency during the pre-treatment of the cells with rutin against direct acting mutagens suggested that in addition to its antioxidant activity, rutin can either block the entry of mutagens into cytosol or physically interact with DNA, providing a protective barrier to prevent DNA–mutagen interaction.

Table 4. Effect of rutin on the mutagenicity induced by sodium azide (NaN3) using Salmonella typhimurium strains.
Treatment Dose (μg per plate) Number of His+ revertants colonies/plate
TA 100 TA 102
Spontaneous 124.66 ± 7.17a 274.00 ± 12.76a
Positive control (NaN3) 1.5 1166.33 ± 22.86g 362.33 ± 14.16d
£Rutin β20 135.33 ± 12.89 296.66 ± 9.07
β40 144 ± 16.09 313 ± 12
Pre-incubation β20 797.33 ± 18.97f (35.48)*** 462.33 ± 15.52c (39.39)
β40 327.33 ± 12.28d (67.61)*** 303.66 ± 11.02bc (66.28)*
Co-incubation β20 766 ± 21.74f (38.43)*** 325.66 ± 12.39c (41.28)
β40 401.66 ± 19.43c (73.47)*** 299.33 ± 12.11b (71.21)*
Pre-treatment β20 711.33 ± 17.63e (43.71)*** 324.66 ± 8.17c (42.43)
β40 341.33 ± 13.07b (79.19)*** 293.66 ± 6.94ab (77.65)*
Open in a new tab
Table 5. Effect of rutin on the mutagenicity induced by methyl methanesulphonate (MMS) using Salmonella typhimurium strains.
Treatment Dose (μg per plate) Number of His+ revertants colonies/plate
TA 100 TA 102
Spontaneous 124.66 ± 7.17a 274.00 ± 12.76a
Positive control (MMS) 1.0 978 ± 40.10f 1495 ± 37.34f
£Rutin β20 135.33 ± 12.89 296.66 ± 9.07
β40 144 ± 16.09 313 ± 12
Pre-incubation β20 626 ± 20.11e (41.24)** 1019.66 ± 24.07e (38.92)**
β40 351.66 ± 17.98c (73.39)*** 623 ± 23.03d (71.41)***
Co-incubation β20 590.33 ± 22.42de (45.42)*** 931.66 ± 24.90d (46.13)***
β40 329.66 ± 15.15c (75.97)*** 551.66 ± 20.41c (77.25)***
Pre-treatment β20 569.66 ± 18.78d (47.85)** 905.66 ± 18.80d (48.26)***
β40 280 ± 8.04b (81.79)*** 464.33 ± 17.15b (84.33)***
Open in a new tab

3.5.2. DNA protection in human peripheral lymphocytes

The comet assay is a simple and precise technique to study DNA damage. The ability of the fraction and rutin to protect against DNA damage was studied by inducing DNA damage by MMS. The average tails length of lymphocytes was assessed for estimation of the relative DNA damage, which is shown in Fig. 3. It is evident from the figure that there was a significantly small tail length (8.64 μm) in the untreated lymphocytes, while the cells treated with MMS showed an average tail length of 28.73 μm. A many fold increase in the tail length of MMS-treated cells clearly depicts that there was a remarkable DNA damage caused by MMS. MMS-treated lymphocytes in the presence of 50 μg ml–1 and 100 μg ml–1 of plant extract showed reduced tail lengths of 25.964 μm and 18.208 μm, respectively. Similarly, the addition of rutin also showed a similar but pronounced effect as compared to plant extract. Rutin at 20 μg ml–1 and 40 μg ml–1 was found to protect against DNA damage by 20.57% and 47.14%, respectively. Moreover, no sign of significant genotoxicity was observed for the compound and fractions when compared to the negative control (DMSO). The decrease in tail length in the presence of the fraction/compound and lymphocytes showed that they protected the DNA from being fragmented or damaged by MMS. The results further strengthen our findings that both the fraction and rutin exhibit a protective effect. The genotoxic/antigentoxic potential of a crude extract of Syzygium cumini was also investigated previously.52,53 However, to date, no detailed study has been performed to evaluate the genotoxic/antigenotoxic effects of the antioxidant active Syzygium cumini seed's fraction in human peripheral lymphocytes against MMS-induced DNA damage.

Fig. 3. Antigenotoxic effect of EAJs fraction and rutin in human peripheral lymphocytes when treated with MMS using the comet assay. Control: Cells treated with 1% DMSO; MMS: cell treated with 200 mM MMS; E1: cells co-incubated with 50 μg ml–1 EAJs; E2: cells co-incubated with 100 μg ml–1 EAJs; R1: cells co-incubated with 20 μg ml–1 rutin; R2: cells co-incubated with 40 μg ml–1 rutin. * is p value equal to or less than 0.05 with respect to the control group and # is p value equal to or less than 0.05 with respect to MMS treatment group.

Fig. 3

Open in a new tab

As evident from the antigenotoxic activity of rutin in the present study, the chemoprotective effects of the EAJs fraction can be assumed to be associated with the presence of phenolic components. The direct antioxidant activity relationship in modulating the effect of the alkylating agent is not clear, instead alternative mechanisms, such as the transfer of an alkyl group to the plant extract/phytocompounds or competition for alkylation labile sites, have been proposed.54

3.5.3. Intracellular ROS quenching

The measurement of ROS using the DCF-DA assay can give an indication of the levels of oxidative stress in lymphocytes. Treatment of the lymphocytes with the fraction and rutin suppressed the intracellular oxidation induced by MMS in a concentration-dependent manner. Oxidative stress was found to decrease by 25.18% and 49.64% when the cell suspensions were treated with the highest concentrations of the fraction (100 μg ml–1) and rutin (40 μg ml–1), respectively, as represented in Fig. 4. Intracellular ROS inhibition by the antioxidant extract and compounds could be explained on the basis of their scavenging capacity.55 Alternatively, it has also been proposed that the treatment of antioxidant active compounds/extracts enhances the activities of CAT, SOD, and glutathione oxidase enzymes.56

Fig. 4. Inhibition of intracellular ROS generation by EAJs fraction and rutin in human peripheral lymphocytes induce by MMS. Control (MMS): Treated with MMS; E1: cells co-incubated with 50 μg ml–1 EAJs; E2: cells co-incubated with 100 μg ml–1 EAJs; R1: cells co-incubated with 20 μg ml–1 rutin; R2: cells co-incubated with 40 μg ml–1 rutin. * is p value equal to or less than 0.05 with respect to MMS treatment group.

Fig. 4

Open in a new tab

3.5.4. Plasmid nicking assay

The DNA-protective activity of the EAJs fraction against oxidative damage was also studied on the pBR322 plasmid DNA model (Fig. 5). Exposure of Fenton's reagent to the plasmid DNA caused a conversion of the supercoiled form to linearized and open circular form (Fig. 5, Lanes 1 and 2). However, the addition of different of concentrations of the EAJs fraction (25–100 μg ml–1) to the reaction mixture increased the formation of the supercoiled form of plasmid DNA (Fig. 5, Lanes 3, 4, 5, and 6). Additionally, when rutin (40 μg ml–1) was incubated with nicking mixture, it also showed an enhanced native form when compared to with Fenton's reagent alone (Fig. 5, Lane 7). These results can be correlated with the plasmid DNA-protecting activity of the phenolic-rich extracts/fractions.57 Polyphenols, such as rutin, have been previously reported to interact with transition metals and effectively reduce ferric iron to the ferrous iron state.58 As the antioxidant activity is largely studied in terms of the structural relationship of the flavonoids, the moieties of different flavonoids interact among each other in a complex mixture-like extract, which further augments their antioxidant potential.59

Fig. 5. Protective effect of EAJs fraction on OH radical-induced pBR322 plasmid DNA damage. Lane 1. Plasmid DNA + Tris buffer; Lane 2. Plasmid DNA + Fenton's reagent; Lane 3. Plasmid DNA + 25 μg ml–1 EAJs; Lane 4. Plasmid DNA + 50 μg ml–1 EAJs; Lane 5. Plasmid DNA + 75 μg ml–1 EAJs; Lane 6. Plasmid DNA + 100 μg ml–1 EAJs; Lane 7. Plasmid DNA + rutin (40 μg ml–1).

Fig. 5

Open in a new tab

3.6. Biophysical analysis

3.6.1. UV-Vis spectroscopic analysis

UV-Vis spectroscopy was used for the preliminary investigation of drug–DNA complex formation. The extent of interaction is determined by a shift in the peak position (λmax) along with changes in the absorbance. The absorption spectra of rutin in the absence and presence of varying concentrations of DNA are shown in Fig. 6A. It is obvious from the figure that DNA exhibited an absorbance maximum at 260 nm. With an increasing concentration of rutin, there was a hyperchromic effect with a negligible shift in the peak position. Such changes in absorption spectra correspond to the presence of various non-covalent interactions between rutin and DNA.60 The quantitative evaluation of the UV-vis spectroscopic data was done using the following equation:61Inline graphicwhere A0 and A are the absorbance of DNA in the absence and presence of rutin, εDNA and εB are the molar extinction coefficients of DNA and the bound complex, [C] is the concentration of rutin, and K is the binding constant. This inverse plot of 1/(A – A0) vs. 1/[C] is a linear fit, which is shown in Fig. 6B. The value of the binding constant was obtained from the ratio of the intercept to the slope, which was found to be 8.82 × 103 M–1.

Fig. 6. UV spectra of DNA in the absence and presence of different concentrations of rutin (10–50 μM) in Tris-HCl buffer (pH 7.4) at 37 °C (A). Plot of 1/(A – A0) vs. 1/[rutin] (B).

Fig. 6

Open in a new tab

3.6.2. Dye displacement assay

In the dye displacement experiments, the binding of any ligand will displace a dye having the same binding site as that of the dye. Any change in the native fluorescent emission behavior of the DNA–dye system can be traced by varying the concentration of the ligand to elucidate the exact binding site. EB is a well-known fluorescence probe that binds to DNA in an intercalative fashion. EB produces a very fluorescent signal when excited at 476 nm, which increases extensively in the presence of DNA. When rutin was titrated to DNA–EB complex, there was no noticeable change in the DNA–EB system, as shown in Fig. 7A. This indicated that rutin has a non-intercalative mode of binding to DNA. To further confirm this, another intercalating probe (AO) was used. AO also showed weak fluorescence in the free form in solution, but when intercalated in DNA, the fluoresce intensity increased many fold.62 The increasing concentration of rutin did not result in any remarkable effect on the fluorescent emission signal of DNA–AO complex, confirming that rutin did not intercalate into DNA (Fig. 7B). Another dye displacement assay was performed in the presence of a minor groove binder, i.e., Hoechst 33258. It is evident from Fig. 7C that there was a tremendous decrease in the fluorescence emission signal of the DNA–Hoechst complex with the progressive addition of rutin, which was due to the displacement of Hoechst from DNA. This observation was further validated by using another minor groove binding dye (DAPI). It was found that with increasing the concentration of rutin, there was a subsequent decrease in the fluorescence signal of DAPI–DNA complex, as shown in Fig. 7D.

Fig. 7. Competitive dye displacement assays. (A) Fluorescence titration of EB–DNA complex with rutin. No effect of rutin was seen on the EB–DNA system. EB–DNA complex was excited at 476 nm and emission spectra were recorded from 530 to 700 nm. (B) Fluorescence titration of AO–DNA complex with rutin. AO–DNA complex was excited at 480 nm and emission spectra were recorded from 490 to 600 nm. No change in fluorescence intensity was seen with the successive addition of rutin (C) Fluorescence spectra of Hoechst–DNA complex with rutin. Fluorescence intensity decreased gradually with the subsequent addition of rutin (0–90 μM). Hoechst–DNA complex was excited at 343 nm and emission spectra were recorded from 360 to 600 nm. (D) Fluorescence spectra of DAPI–DNA complex with rutin. Fluorescence intensity decreased gradually with the subsequent addition of rutin (0–90 μM). DAPI–DNA complex was excited at 338 nm and emission spectra were recorded from 350 to 600 nm.

Fig. 7

Open in a new tab

The comparative quantification of dye displacement data was performed by plotting the Stern–Volmer plot using following Stern–Volmer equation:F0/F = 1 + Ksv [Q]where, F0 and F are the fluorescence emission signal of DNA–dye complex in the absence and presence of rutin, Ksv is the Stern–Volmer constant, and [Q] is the concentration of rutin. The plot is shown in Fig. 8 and the values obtained are listed in Table 6. It is apparent from the data that the Ksv value for EB and AO displacement is negligible compared to Hoechst and DAPI displacement. This again validates that rutin was capable of displacing Hoechst as well as DAPI from DNA and therefore has a minor groove mode of binding. Thus the minor groove binding non-intercalative flavonoid or its derivatives could protect DNA macromolecules from undergoing oxidation or alkylation leading to mutations or DNA damage.63

Fig. 8. Stern–Volmer plot for quenching of the fluorescence intensity of different fluorescent dyes–DNA systems by the successive addition of rutin 10 to 100 μM.

Fig. 8

Open in a new tab
Table 6. Comparison of Ksv values for the quenching of fluorescence intensity by the displacement of different fluorescent dyes from DNA by rutin.
Dye K sv (×104 M–1) R 2
Ethidium bromide 0.07 0.8951
Acridine orange 0.04 0.8263
Hoechst 3325 4.57 0.9300
Open in a new tab

3.6.3. Thermal denaturation

The double helical structure of DNA is stabilized by hydrogen bonds between opposite strands and base pair stacking. At elevated temperature, these bonds stabilizing the helix are disrupted, causing the separation of strands, which is known as the thermal denaturation of DNA.64 The Tm value was determined by plotting A/A25 at different temperatures, and the transition midpoint of the sigmoidal-fitted curve yields Tm. It is known that many molecules interact with DNA via different modes causing an increment in the stability of DNA.65 We studied the effect of rutin on the thermal denaturation profile of DNA. The thermal denaturation curve of DNA in the absence and presence of DNA is shown in Fig. 9. The melting temperature for free DNA was obtained as 69.54 °C, while for the rutin–DNA complex, it was 74.12 °C. The obvious enhancement in Tm value clearly substantiates the tendency of rutin to interact physically with DNA macromolecules.

Fig. 9. Effect of rutin on the melting temperature of DNA. Melting curves of DNA (50 mM) in the absence and presence of rutin (50 mM).

Fig. 9

Open in a new tab

3.6.4. Isothermal titration calorimetric analysis

ITC is a sensitive technique that has been extensively used in studying the thermodynamic as well as kinetic parameters of DNA–ligand interactions.66 Therefore, to gain a further insight into various thermodynamic parameters of DNA–rutin interaction, we performed ITC experiments. Along with information about the thermodynamic quantities, such as the entropy change during binding (ΔS), enthalpy change (ΔH), and Gibb's free energy change (ΔG), it also determines the binding affinity (K) and the number of binding sites (n).67 In Fig. 10, the ITC profiles show the heat of reaction plotted against the molar ratio of rutin to DNA after baseline correction.

Fig. 10. Isothermal titration calorimetry of DNA and rutin interaction.

Fig. 10

Open in a new tab

The Gibbs energy change (ΔG) was calculated using the formula:ΔG = –RT ln Kb = ΔH – TΔSwhere T is the absolute temperature (298 K) and R = 8.315 J mol–1 K–1.

The thermodynamic parameters, binding constant, and stoichiometry of the rutin–DNA complex were predicted by fitting the integrated heats according to an independent binding model. The ITC data yielded a binding constant of 5.48 × 103 M–1 for the rutin–DNA interaction, which is in line with the results obtained from the spectroscopic studies. The negative value of ΔG (–3.168 kcal mol–1) obtained showed that the process of interaction between rutin and DNA was spontaneous, while the negative value of ΔH (–4.148 kcal mol–1) indicated the role of hydrogen bonds and van der Waal's interactions in the binding process.68 The negative value of ΔH also confirmed that the formation of the rutin–DNA complex was exothermic. Moreover, the large negative enthalpy changes along with calculated favorable free energy changes from the rutin–DNA binding constant suggested that the binding was enthalpy driven. It is an established fact that interactions in the case of intercalative binding are entropically driven, whereas in groove binding they are enthalpically driven.69 The positive value of ΔS (3.29 cal mol–1 K–1) indicated that the complexation of rutin to DNA was spontaneous. Moreover, the value of n (0.995), being close to unity, depicted the single binding site per nucleotide of DNA. It can thus be concluded from the thermodynamic outcomes of the isothermal calorimetric titration that the interaction of rutin to DNA was consistent with the groove mode of binding.

In general, one of the less explored mechanisms of DNA protection includes the physical interaction of antigenotoxic compounds with DNA. It has been proposed that certain DNA–compound interactions lead to the formation of a protective barrier against damaging chemicals more prominently effective against alkylating agents.51 Exploring different aspects of rutin–DNA interaction via biophysical methodologies provides strong evidence that rutin binds with DNA via non-covalent affinities preferably at grooves region. Hence, it can be speculated on the basis of DNA binding studies that rutin can mask the labile sites of DNA and protect it from undergoing modifications.

3.6.5. Molecular docking

Computational tools describing the molecular interaction of phytocompounds/drugs with DNA are often used to gain an insight into the mechanism underlying their biological activities, including antimutagenic and antigenotoxic effects.51,70 In silico techniques, such as molecular docking, have proven to be a useful tool to decipher the bio-macromolecular interaction mechanisms, including drug–DNA interaction, DNA–protein interaction, and protein–protein interaction. These tools provide more pronounced interaction insights of ligand–receptor binding and take less time.71 Molecular docking results showed that H29 of rutin was found to interact with oxygen of the phosphate group of the dA17 nucleotide via hydrogen bonding at a bond length of 2.30 Å. Similarly, the rutin–DNA complex was also stabilized by three hydrogen bods with dG14 at distances of 2.20 Å, 2.18 Å, and 2.82 Å, as depicted in Fig. 11. There was one classical and one non-classical hydrogen bond formed with dC15 with bond lengths of 2.16 Å and 3.79 Å, respectively. Molecular docking results found that the binding energy of rutin with DNA is –7.9 kcal mol–1, signifying a strong interaction. In another study, Mladenovic et al.51 observed that the binding energy of rutin with DNA was the highest among other docked phenolic acid and flavonoids, including myricetin and quercetin. Further, through molecular dynamic simulation, Chandran et al.72 showed that minor groove binding molecules enhance the stability of DNA through their non-covalent interactions. Such non-intercalating interactions may protect DNA from chemical-induced mutations by decreasing the sites available for chemical mutagens to cause mutagenesis.

Fig. 11. Molecular docked structures of rutin complexed with DNA [d(CGCGAATTCGCG)2(PDB ID: ; 1DNA)] (A), rutin interacts with dG10, dG14. dC15, and dA17 (B) and interacting atoms of rutin and DNA along with their bond lengths (C).

Fig. 11

Open in a new tab

4. Conclusions

In conclusion, the antioxidant active fraction of Syzygium cumini's seeds possess antigenotoxic properties against oxidative and alkylating DNA damage in different test systems. The observed activity of the fraction and rutin itself could be associated with antioxidant and DNA-protective potential. Biophysical interaction studies and molecular docking confirmed the possible mode of interaction of rutin with DNA and further contributed to our understanding of the mechanism of action. Synergistic interactions among the phytocompounds of the same group and with different groups in a bioactive plant extract/fraction should be attended to in future studies. On the other hand, standardized extracts/fraction may be evaluated in a suitable animal model for therapeutic efficacy.

Conflicts of interest

The authors declare no conflict of interest.

Supplementary Material

Click here for additional data file. (156.2KB, pdf)

Acknowledgments

We are grateful to SAIF, IIT Bombay, Mumbai, India for LC-qTOF-MS analysis. UGC, New Delhi, is acknowledged for providing research fellowship to MSK and FAQ through AMU, Aligarh.

Footnotes

†Electronic supplementary information (ESI) available. See DOI: 10.1039/c7tx00269f

References

  1. Sies H., Berndt C., Jones D. P. Annu. Rev. Biochem. 2017;86(1):715–748. doi: 10.1146/annurev-biochem-061516-045037. [DOI] [PubMed] [Google Scholar]
  2. Valko M., Leibfritz D., Moncol J., Cronin M. T., Mazur M., Telser J. Int. J. Biochem. Cell Biol. 2007;39(1):44–84. doi: 10.1016/j.biocel.2006.07.001. [DOI] [PubMed] [Google Scholar]
  3. Cooke M. S., Evans M. D., Dizdaroglu M., Lunec J. FASEB J. 2003;17(10):1195–1214. doi: 10.1096/fj.02-0752rev. [DOI] [PubMed] [Google Scholar]
  4. Drablos F., Feyzi E., Aas P. A., Vaagbø C. B., Kavli B., Bratlie M. S., Peña-Diaz J., Otterlei M., Slupphaug G., Krokan H. E. DNA Repair. 2004;3(11):1389–1407. doi: 10.1016/j.dnarep.2004.05.004. [DOI] [PubMed] [Google Scholar]
  5. Fu D., Calvo J. A., Samson L. D. Nat. Rev. Cancer. 2012;12(2):104–120. doi: 10.1038/nrc3185. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Modern phytomedicine: Turning medicinal plants into drugs, ed. I. Ahmad, F. Aqil and M. Owais, John Wiley & Sons, 2006. [Google Scholar]
  7. Aqil F., Munagala R., Jeyabalan J., Joshi T., Gupta R. C. and Singh I. P., The Indian Blackberry (Jamun), Antioxidant Capacity, and Cancer Protection, in Cancer: Oxidative Stress and Dietary Antioxidants, Amsterdam, 2015. [Google Scholar]
  8. Ayyanar M., Subash-Babu P., Ignacimuthu S. Complement. Ther. Med. 2013;21(3):232–243. doi: 10.1016/j.ctim.2013.03.004. [DOI] [PubMed] [Google Scholar]
  9. Aqil F., Gupta A., Munagala R., Jeyabalan J., Kausar H., Sharma R. J., Singh I. P., Gupta R. C. Nutr. Cancer. 2012;64(3):428–438. doi: 10.1080/01635581.2012.657766. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Chaudhuri A. K., Pal S., Gomes A., Bhattacharya S. Phytother. Res. 1990;4(1):5–10. [Google Scholar]
  11. Abdalla F. H., Bellé L. P., Bitencourt P. E., De Bona K. S., Zanette R. A., Boligon A. A., Athayde M. L., Pigatto A. S., Moretto M. B. Biometals. 2011;24(2):349–356. doi: 10.1007/s10534-010-9402-5. [DOI] [PubMed] [Google Scholar]
  12. Kothari V., Seshadri S., Mehta P. Res. Biotechnol. 2011;2(6):53–63. [Google Scholar]
  13. Chaturvedi A., Kumar M. M., Bhawani G., Chaturvedi H., Kumar M., Goel R. K. Indian J. Physiol. Pharmacol. 2007;51(2):131. [PubMed] [Google Scholar]
  14. Jagetia G. C., Shetty P. C., Vidyasagar M. S. Integr. Cancer Ther. 2012;11(2):141–153. doi: 10.1177/1534735411413261. [DOI] [PubMed] [Google Scholar]
  15. Farswan M., Mazumder P. M., Parcha V., Upaganlawar A. Pharmacogn. Mag. 2009;5(18):127. [Google Scholar]
  16. Zahin M., Ahmad I., Aqil F. Toxicol. in Vitro. 2010;24(4):1243–1249. doi: 10.1016/j.tiv.2010.02.004. [DOI] [PubMed] [Google Scholar]
  17. Berahou A., Auhmani A., Fdil N., Benharref A., Jana M., Gadhi C. A. J. Ethnopharmacol. 2007;112(3):426–429. doi: 10.1016/j.jep.2007.03.032. [DOI] [PubMed] [Google Scholar]
  18. Ahmad I., Beg A. Z. J. Ethnopharmacol. 2001;74(2):113–123. doi: 10.1016/s0378-8741(00)00335-4. [DOI] [PubMed] [Google Scholar]
  19. Harborne A. J., Phytochemical methods a guide to modern techniques of plant analysis, Springer science & business media, 1998. [Google Scholar]
  20. Chang C. C., Yang M. H., Wen H. M., Chern J. C. J. Food Drug Anal. 2002;10(3):178–182. [Google Scholar]
  21. Brand-Williams W., Cuvelier M. E., Berset C. L. LWT–Food Sci. Technol. 1995;28(1):25–30. [Google Scholar]
  22. Oyaizu M. Jpn. J. Nutr. Diet. 1986;44(6):307–315. [Google Scholar]
  23. Gulcin İ. Chem.-Biol. Interact. 2009;179(2):71–80. doi: 10.1016/j.cbi.2008.09.023. [DOI] [PubMed] [Google Scholar]
  24. Takao T., Kitatani F., Watanabe N., Yagi A., Sakata K. Biosci., Biotechnol., Biochem. 1994;58(10):1780–1783. [Google Scholar]
  25. Maron D. M., Ames B. N. Mutat. Res., Environ. Mutagen. Relat. Subj. 1983;113(3–4):173–215. doi: 10.1016/0165-1161(83)90010-9. [DOI] [PubMed] [Google Scholar]
  26. Hour T. C., Liang Y. C., Chu I. S., Lin J. K. Food Chem. Toxicol. 1999;37(6):569–579. doi: 10.1016/s0278-6915(99)00031-9. [DOI] [PubMed] [Google Scholar]
  27. Singh N. P., McCoy M. T., Tice R. R., Schneider E. L. Exp. Cell Res. 1988;175(1):184–191. doi: 10.1016/0014-4827(88)90265-0. [DOI] [PubMed] [Google Scholar]
  28. Lee J. C., Kim H. R., Kim J., Jang Y. S. J. Agric. Food Chem. 2002;50(22):6490–6496. doi: 10.1021/jf020388c. [DOI] [PubMed] [Google Scholar]
  29. Trott O., Olson A. J. J. Comput. Chem. 2010;31(2):455–461. doi: 10.1002/jcc.21334. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Lim D. W., Kim H., Park J. Y., Kim J. E., Moon J. Y., Park S. D., Park W. H. BMC Complementary Altern. Med. 2016;16(1):155. doi: 10.1186/s12906-016-1121-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Rashed K., Potočnjak I., Giacometti J., Škoda M., Domitrović R. J. Funct. Foods. 2014;8:319–330. [Google Scholar]
  32. Jabeur I., Martins N., Barros L., Calhelha R. C., Vaz J., Achour L., Santos-Buelga C., Ferreira I. C. Food Funct. 2017;8(3):975–984. doi: 10.1039/c6fo01778a. [DOI] [PubMed] [Google Scholar]
  33. Köksal E., Bursal E., Gülçin İ., Korkmaz M., Çağlayan C., Gören A. C., Alwasel S. H. Int. J. Food Prop. 2017;20(3):514–525. [Google Scholar]
  34. Bursal E., Köksal E., Gülçin İ., Bilsel G., Gören A. C. Food Res. Int. 2013;51(1):66–74. [Google Scholar]
  35. Makhafola T. J., Elgorashi E. E., McGaw L. J., Verschaeve L., Eloff J. N. BMC Complementary Altern. Med. 2016;16(1):490. doi: 10.1186/s12906-016-1437-x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Zahin M., Ahmad I., Aqil F. Drug Chem. Toxicol. 2017;40(2):146–153. doi: 10.1080/01480545.2016.1188397. [DOI] [PubMed] [Google Scholar]
  37. Bhouri W., Sghaier M. B., Kilani S., Bouhlel I., Dijoux-Franca M. G., Ghedira K., Ghedira L. C. Food Chem. Toxicol. 2011;49(5):1167–1173. doi: 10.1016/j.fct.2011.02.011. [DOI] [PubMed] [Google Scholar]
  38. Zahin M., Aqil F., Ahmad I. Mutat. Res., Genet. Toxicol. Environ. Mutagen. 2010;703(2):99–107. doi: 10.1016/j.mrgentox.2010.08.001. [DOI] [PubMed] [Google Scholar]
  39. Celikler S., Vatan O., Yildiz G., Bilaloglu R. Food Chem. Toxicol. 2009;47(4):796–801. doi: 10.1016/j.fct.2009.01.010. [DOI] [PubMed] [Google Scholar]
  40. Zahin M., Ahmad I., Gupta R. C., Aqil F. BioMed Res. Int. 2014:467465. doi: 10.1155/2014/467465. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Ajith T. A., Janardhanan K. K. Food Chem. Toxicol. 2011;49(10):2676–2680. doi: 10.1016/j.fct.2011.07.022. [DOI] [PubMed] [Google Scholar]
  42. Słoczyńska K., Powroźnik B., Pękala E., Waszkielewicz A. M. J. Appl. Genet. 2014;55(2):273–285. doi: 10.1007/s13353-014-0198-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Gulluce M., Orhan F., Adiguzel A., Bal T., Guvenalp Z., Dermirezer L. O. Toxicol. Ind. Health. 2013;29(6):534–540. doi: 10.1177/0748233712442732. [DOI] [PubMed] [Google Scholar]
  44. Ammar R. B., Sghaier M. B., Boubaker J., Bhouri W., Naffeti A., Skandrani I., Bouhlel I., Kilani S., Ghedira K., Chekir-Ghedira L. Chem.-Biol. Interact. 2008;174(1):1–0. doi: 10.1016/j.cbi.2008.04.006. [DOI] [PubMed] [Google Scholar]
  45. Vattem D. A., JaNG H. D., Levin R., Shetty K. J. Food Biochem. 2006;30(1):98–116. [Google Scholar]
  46. de Melo E. B., Martins J. P., Jorge T. C., Friozi M. C., Ferreira M. M. Eur. J. Med. Chem. 2010;45(10):4562–4569. doi: 10.1016/j.ejmech.2010.07.017. [DOI] [PubMed] [Google Scholar]
  47. Snijman P. W., Swanevelder S., Joubert E., Green I. R., Gelderblom W. C. Mutat. Res., Genet. Toxicol. Environ. Mutagen. 2007;631(2):111–123. doi: 10.1016/j.mrgentox.2007.03.009. [DOI] [PubMed] [Google Scholar]
  48. Ruiz-Pérez N. J., Arriaga-Alba M., Sánchez-Navarrete J., Camacho-Carranza R., Hernández-Ojeda S., Espinosa-Aguirre J. J. Sci. Rep. 2014;4:6743. doi: 10.1038/srep06743. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Edenharder R., Grünhage D. Mutat. Res., Genet. Toxicol. Environ. Mutagen. 2003;540(1):1–8. doi: 10.1016/s1383-5718(03)00114-1. [DOI] [PubMed] [Google Scholar]
  50. Resende F. A., da Silva Almeida C.P., Vilegas W., Varanda E. A. Food Chem. 2014;155:251–255. doi: 10.1016/j.foodchem.2014.01.071. [DOI] [PubMed] [Google Scholar]
  51. Mladenovic M., Matić S., Stanić S., Solujić S., Mihailović V., Stanković N., Katanić J. Biochem. Pharmacol. 2013;86(9):1376–1396. doi: 10.1016/j.bcp.2013.08.018. [DOI] [PubMed] [Google Scholar]
  52. Tripathi P., Patel R. K., Tripathi R., Kanzariya N. R. Drug Chem. Toxicol. 2013;36(4):396–402. doi: 10.3109/01480545.2012.749271. [DOI] [PubMed] [Google Scholar]
  53. Arun R., Prakash M. V., Abraham S. K., Premkumar K. J. Ethnopharmacol. 2011;134(2):329–333. doi: 10.1016/j.jep.2010.12.014. [DOI] [PubMed] [Google Scholar]
  54. Barcelos G. R., Grotto D., Angeli J. P., Serpeloni J. M., Rocha B. A., Bastos J. K., Barbosa F. Phytother. Res. 2011;25(9):1381–1388. doi: 10.1002/ptr.3436. [DOI] [PubMed] [Google Scholar]
  55. Lee M. S., Lee I. I., Kim Y., Kim Y. J., Heo H. J., Kim D. O. Food Sci. Biotechnol. 2014;23(4):1247–1252. [Google Scholar]
  56. Van Reyk D. M., King N. J., Dinauer M. C., Hunt N. H. Free Radicals Biol. Med. 2001;30(1):82–88. doi: 10.1016/s0891-5849(00)00449-4. [DOI] [PubMed] [Google Scholar]
  57. Yasir M., Sultana B., Amicucci M. J. Funct. Foods. 2016;26:645–656. [Google Scholar]
  58. Mira L., Tereza Fernandez M., Santos M., Rocha R., Helena Florêncio M., Jennings K. R. Free Radical Res. 2002;36(11):1199–1208. doi: 10.1080/1071576021000016463. [DOI] [PubMed] [Google Scholar]
  59. Dean J. D., Flavone: the Molecular and Mechanistic Study of How a Simple Flavonoid Protects DNA from Oxidative Damage, Electronic Theses and Dissertations, 2003, Paper 849. http://dc.etsu.edu/etd/849. [Google Scholar]
  60. Rahban M., Divsalar A., Saboury A. A., Golestani A. J. Phys. Chem. C. 2010;114(13):5798–5803. [Google Scholar]
  61. Qais F. A., Alam M. M., Naseem I., Ahmad I. RSC Adv. 2016;6(70):65322–65337. [Google Scholar]
  62. Qais F. A., Ahmad I., Long J., Wang X. M., Xu D. L., Ding L. S. J. Pharm. Biomed. Anal. J. Mol. Recognit. 2018;2014;14927(3):193–205. 131–137. doi: 10.1016/j.jpba.2017.10.016. [DOI] [PubMed] [Google Scholar]
  63. Jakhar R., Paul S., Park Y. R., Han J., Kang S. C. J. Photochem. Photobiol., B. 2014;131:96–103. doi: 10.1016/j.jphotobiol.2014.01.003. [DOI] [PubMed] [Google Scholar]
  64. Qais F. A., Abdullah K. M., Alam M. M., Naseem I., Ahmad I. Int. J. Biol. Macromol. 2017;97:392–402. doi: 10.1016/j.ijbiomac.2017.01.022. [DOI] [PubMed] [Google Scholar]
  65. Xia K., Zhang G., Gong D. Luminescence. 2017;32:988–998. doi: 10.1002/bio.3281. [DOI] [PubMed] [Google Scholar]
  66. Buurma N. J., Haq I. Methods. 2007;42:162–172. doi: 10.1016/j.ymeth.2007.01.010. [DOI] [PubMed] [Google Scholar]
  67. Yasmeen S., Riyazuddeena, Qais F. A. Int. J. Biol. Macromol. 2017;105:92–102. doi: 10.1016/j.ijbiomac.2017.06.122. [DOI] [PubMed] [Google Scholar]
  68. Husain M., Ishqi H., Sarwar T., Rehman S. MedChemComm. 2017;8:1283–1296. doi: 10.1039/c7md00094d. [DOI] [PMC free article] [PubMed] [Google Scholar]
  69. Mati S. S., Roy S. S., Chall S., Bhattacharya S., Bhattacharya S. C. J. Phys. Chem. B. 2013;117(47):14655–14665. doi: 10.1021/jp4090553. [DOI] [PubMed] [Google Scholar]
  70. Buraka E., Chen C. Y., Gavare M., Grube M., Makarenkova G., Nikolajeva V., Bisenieks I., Brūvere I., Bisenieks E., Duburs G., Sjakste N. Chem.-Biol. Interact. 2014;220:200–207. doi: 10.1016/j.cbi.2014.06.027. [DOI] [PubMed] [Google Scholar]
  71. Alam M. M., Qais F. A., Ahmad I., Alam P., Khan R. H., Naseem I. J. Biomol. Struct. Dyn. 2017;36(3):795–809. doi: 10.1080/07391102.2017.1298470. [DOI] [PubMed] [Google Scholar]
  72. Chandran A., Ghoshdastidar D., Senapati S. J. Am. Chem. Soc. 2012;134(50):20330–20339. doi: 10.1021/ja304519d. [DOI] [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. (156.2KB, pdf)

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

RESOURCES