2.5.1.1 Free radical scavenging assay (% FRSA)

The scavenging activity of each solvent extract was assessed by previously described method (Khan et al., 2015). Briefly, an aliquot of 10 µl of test sample (4 mg/ml DMSO) and 190 µl of DPPH solution (9.2 mg/100 ml in methanol) was transferred into each well of 96 well plate followed by incubation at 37 °C for 60 min in dark environment. After incubation, absorbance was recorded at 630 nm. The percent free radical scavenging activity (% FRSA) of test samples was estimated by following equation: $$\%\mathrm{F}\mathrm{R}\mathrm{S}\mathrm{A}=(1-\mathrm{O}\mathrm{D}\mathrm{s}/\mathrm{O}\mathrm{D}\mathrm{c})\times 100$$

Where ODs is absorbance of sample solution containing DPPH reagent and ODc is absorbance of negative control containing DMSO and DPPH reagent. The test samples having % FRSA ≥ 50% were tested at final concentrations of 200, 66.66, 22.22 and 7.4 µg/ml to determine their 50% inhibitory concentration (IC₅₀). Ascorbic acid was used as positive control in this assay. IC₅₀ was calculated by graph pad prism 5 software and assay was run thrice.

2.5.1.2 Determination of total antioxidant capacity (TAC)

Total antioxidant capacity was determined by previously described protocol (Khan et al., 2015). The eppendorf tubes comprising of 100 µl of test sample and 900 µl of TAC reagent (0.6 M sulphuric acid, 28 mM monosodium phosphate, 48 mM ammonium molybdate solution in water) were incubated at 95 °C for 90 min in water bath and then cooled to room temperature. Finally, 200 µl of test sample was transferred to 96 well plate and absorbance was measured at 630 nm. Ascorbic acid (1 mg/ml DMSO) at final assay concentration of (0–50 µg/ml) was used as positive control and DMSO as negative control (blank). Total antioxidant capacity was computed using ascorbic acid calibration curve (y = 0.0188x − 0.0189, R² = 0.9983) and results were expressed as number of microgram equivalents of ascorbic acid per milligram dry plant weight (µg AAE/mg DW). The assay was performed thrice.

2.5.1.3 Determination of total reducing power (TRP)

Previously described protocol (Khan et al., 2015) was used for assessment of total reducing power of D. viscosa extracts. The eppendorf tubes containing 0.2 ml of test sample (4 mg/ml DMSO), 0.4 ml of phosphate buffer (0.2 mol/l, pH 6.6) and 0.5 ml of potassium ferricyanide (1% w/v) were incubated at 50 °C for 20 min. Then 0.4 ml of trichloroacetic acid (10% w/v) was incorporated and centrifuged at 3000 rpm for 10 min. From the centrifuged mixture, 150 µl of supernatant was transferred to 96 well plate containing 50 µl of ferric chloride (0.1% w/v). Absorbance was recorded at 630 nm. DMSO and ascorbic acid (1 mg/ml) at final concentrations of (0-25µg/ml) were used as negative and positive control. Total reducing power was computed using ascorbic acid calibration (y = 0.0559x − 0.014, R² = 0.994) and results were expressed as number of microgram equivalents of ascorbic acid per milligram of dry plant weight (µg AAE /mg DW) after triplicate analysis.

2.5.2.1 Antibacterial assay

Antibacterial potential of each solvent extract was determined by previously described disc diffusion method (Khan et al., 2015) against five different strains; Staphylococcus aureus ATCC-6538 Bacillus subtilis ATCC-6633, Escherichia coli ATCC-25922, Klebsiella pneumoniae ATCC-1705 and Pseudomonas aeruginosa ATCC-15442. A 50 µl aliquot of 24 hrs refreshed culture was used to prepare lawn on nutrient agar plates. Sterile discs instilled with 5 µl test samples (20 mg/ml in DMSO) were placed on agar plates. Cefixime and roxithromycin were employed as positive standards while DMSO was used as blank. The incubation was done at 37 °C for 24 hrs and antibacterial potential was determined by measuring average diameter of zone of inhibition around sample. The assay was performed thrice. Microbroth dilution method (Fatima et al., 2015) was used for determination of minimum inhibitory concentration (MIC) of test samples with ≥10 mm zone of inhibition. Density of bacterial inoculum was maintained at (5 × 10⁴ CFU/ml). Three-fold serial dilution of test sample with final well concentrations of (100, 33.33, 11.11, and 3.70 µg/ml) was prepared with sterile nutrient broth. Subsequently, 195 µl of bacterial culture was added followed by an overnight incubation at 37 °C. The lowest concentration at which the extract exhibited visible growth inhibition was designated as its MIC. The assay was performed in triplicate.

2.5.2.2 Antifungal assay

The sensitivity of test samples against five fungal strains i.e. Aspergillus fumigatus (FCBP- 66), Mucor sp. (FCBP-0300), A. niger (FCBP-0198), Fusarium solani (FCBP- 0291) and A. flavus (FCBP-0064) was determined by agar disc diffusion protocol (Fatima et al., 2015). The fungal spores were harvested in Tween 20 solution (0.02% v/v in water) and turbidity was adjusted to 0.5 McFarland turbidity standard. The SDA plates were swabbed by 100 µl of fungal spore suspension and then 5 µl of test sample from 20 mg/ml DMSO stock solution was infused on sterile filter disc and placed on plates. Sterile filter discs each infused with 5 µl of clotrimazole (4 mg/ml in DMSO) and DMSO served as positive and negative control respectively. All the sample and control treated discs were incubated for 24–48 hrs at 28–30 °C. The antifungal activity of test extracts was recorded by measuring zone of inhibition around samples to nearest (mm). The assay was run thrice.

2.5.3.1 Brine shrimp lethality assay

The preliminary cytotoxicity of crude extracts against brine shrimps (Artemia salina) larvae was determined by 24 hrs lethality test by previously described protocol (Haq et al., 2011). Simulated sea water (38 g/l with 6 mg/l yeast) was used to hatch Artemia salina eggs (Ocean star, USA) in specially designed bi-compartment perforated tray under light and warmth 30–32 °C. After hatching period of 24–48 hrs, the hatched nauplii were gathered and then collected with Pasteur pipette in small beaker containing sea water. To each well, 10 mature nauplii were transferred and 150 µl of sea water was added. Two-fold serial dilution of test extracts was made with final concentrations of (25–200 µg/ml). Corresponding volume of each test sample containing (not more than 1% DMSO in sea water) was added to the wells containing sea water and shrimp larvae. The volume of each well was made up to 300 µl with sea water. Serial concentrations of doxorubicin (1.25–10 µg/ml) and 1% DMSO served as positive and negative control respectively. After 24 hrs incubation period at 37 °C, dead nauplii were counted using inverted microscope. The whole experiment was run thrice. The LC₅₀ was calculated by graph pad prism 5 software.

2.5.3.2 THP-1 human leukemia cell line cytotoxicity assay

Standard MTT protocol was used for assessment of in vitro cytotoxic potential of D. viscosa crude extracts (Fatima et al., 2015). THP-1 human leukemia cells (ATCC# TIB-202) were cultured in RPMI-1640 growth medium (Gibco BRL, Life Technologies, Inc) buffered with 2.2 g/l NaHCO₃ and supplemented with 10% v/v heat inactivated fetal bovine serum (HIFBS); pH 7.4] in a humidified 5% carbon dioxide incubator (Panasonic, Japan MCO-18AC-PE) at 37 °C. An aliquot of 10 µl of test sample with final concentration of 20 µg/ml (having 1% DMSO in PBS, pH 7.4) was transferred to 96 well plate followed by addition of 190 µl of leukaemia cells (seeding density 5 × 10⁵ cells/ml) and kept for 72 hrs at 37 °C in 5% CO₂ incubator. Sterilized pre filtered MTT solution 4 mg per ml in distilled water (20 µl) was added to each well and incubated at 37 °C for 4 hrs (in humidified CO₂ incubator). After incubation, supernatant was carefully removed without disturbing coloured formazan sediments. Afterwards, 100 µl of DMSO was added to dissolve formazan sediments and the plate was kept aside for 1 hr to allow complete dissolution and then absorbance was noted at 540 nm. The vincristine and 5-fluorouracil at final concentration of 20 µg/ml served as positive controls while 1% DMSO in PBS employed as negative control. Test samples with ≥50% inhibition were further screened at four different concentrations (1.25–10 µg/ml). Experiment was run thrice and IC₅₀ was calculated using graph pad prism 5 software.

2.5.3.3 Hep G2 cell line cytotoxicity assay

The SRB colorimetric assay previously described by (Vichai and Kirtikara, 2006) was used to evaluate the cytotoxicity of test extracts against Hep G2 cancer cells (RBRC-RCB1648). The Hep G2 cells were grown in the Dulbecco's Modified Eagle Medium (DMEM) supplemented with 10% FBS, 100 µg/ml streptomycin sulphate, 100 IU/ml penicillin G sodium, and 0.25 µg/ml amphotericin B. The plate was then incubated at 37 °C in humidified atmosphere enriched with 5% CO₂ for 72 h (confluence of approximately 70–80%). The old media was changed with the fresh one and incubated again for another 24 h. Afterwards, the cells were trypsonised and diluted to get an assay density of 1x10⁵ cells/ml. An aliquot of 20 µl of each test sample [1% DMSO in PBS (pH 7.4)] was added to 96 well microplate followed by the addition of 180 µl of harvested (Hep G2) cells. The final assay concentration of test sample was 20 µg/ml. The positive and negative control wells contained Doxorubicin (20–0.8 µg/ml) and DMSO (1% v/v) in PBS instead of test sample. The reaction mixture was incubated at 37 °C for 72 hrs in humidified atmosphere (5% CO₂). Then 20 µl of cold TCA solution (20% w/v) was added for cell fixation proceeded by washing of fixed cells with tap water 4 times. The reaction plate was air dried and stained with 50 µl of (0.057% w/v SRB in 1% w/v acetic acid) for 30 min at room temperature. Wells were then washed (4 times) with 1% (v/v) acetic acid and the plates were dried overnight. The 200 µl of 10 mM Tris base: pH 10 was used to solubilize the bound dye for 1 hr. The optical density of reaction plate was measured at 515 nm using micro plate reader. Then percent survival was calculated. A zero-day control was run by transferring equivalent number of cells to sixteen wells of 96 well plate and incubated at 37 °C for 1 hr. After incubation cells were processed by above-described procedure.

Percent cell growth inhibition was calculated using following equation: $$\%\text{inhibition}=100-[({\text{O}}_{\text{cells}+\text{samples}}--{\text{O}}_{\text{day}0})/\left({\text{O}}_{\text{cells}+1\%\text{DMSO}}-{\text{O}}_{\text{day}0}\right)\times 100]$$

3.3.1.1 %FRSA assay

Many free radicals and reactive oxygen species are generated by living cells as by-products of physiological and biochemical processes. Free radicals are responsible for various chronic diseases including diabetes mellitus, aging, cancer, ischemic heart disease, atherosclerosis and many other degenerative diseases due to oxidative damage of proteins, DNA, amino acids and lipids (Aiyegoro and Okoh, 2010). Normally a low level of different forms of free radicals exists to regulate physiological functions that are scavenged by the inherited antioxidant system of body. The excessive production of these radicals cause damage to body cells (Vaghasiya et al., 2011). The concerns over the potential side effects of frequently used synthetic antioxidants such as butylated hydroxyanisole (BHA) and butylated hydroxytoluene (BHT) have focused research interests towards finding antioxidants from natural sources. The plants being reservoir of bioactive compounds constitute one of the most valuable sources for natural antioxidants. The plant polyphenols serve as potential alternative to the synthetic antioxidants due to safety and efficacy (Tzima et al., 2018). A large number of antioxidant compounds have been isolated from plants with ability to scavenge ROSs (Riaz et al., 2012). A single method is not reliable to determine antioxidant capability of test sample due to complex nature and variable mode of action of antioxidant agent, so different antioxidant assays should be performed (Sini et al., 2011).

The antioxidant activity of D. viscosa extracts was determined by measuring their ability to quench the stable free radicle i.e., DPPH the results of which are presented in Fig. 3. The percent free radical quenching ability was assessed by discoloration of purple colour of DPPH solution (Sharma and Bhat, 2009). The highest scavenging activity in flower and leaf part was exhibited by WA extracts with an IC₅₀ of 116.7 µg/ml and 47.99 µg/ml respectively. While in stem and root, maximum scavenging potential was displayed by MEA and EEA extracts showing IC₅₀ = 23.8 µg/ml and IC₅₀ = 24.95 µg/ml respectively. The results are in accordance with previous findings where polar solvents such as methanol and chloroform combination and aqueous extracts of leaf and stem showed highest scavenging potential (Akhtar and Mirza, 2015; Mothana et al., 2010). The scavenging mediated antioxidant activity might be due to the polyphenols detected in various solvent extracts of D. viscosa.

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Fig. 3

%FRSA (radical scavenging activity) of D. viscosa flower, leaf, stem and root parts in different solvents. Values are presented as mean ± Standard deviation from triplicate investigation. The columns with different superscript ^(a-g) letters show significantly (P < 0.05) different means.

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3.3.1.2 TAC

The total antioxidant capacity of D. viscosa extracts was determined by phosphomolybdenum based colorimetric method based on formation of green phosphate molybdenum complex (Khan et al., 2015). The results are presented in Fig. 4. The highest antioxidant activity was displayed by WA (22.55 ± 0.07 µg AAE/mg DW) of leaf, E extract of flower (7.59 ± 0.19 µg AAE/mg DW), WA extract of stem (5.41 ± 0.01 µg AAE/mg DW) and EAA extract of root (4.08 ± 0.06 µg AAE/mg DW). Our results strengthen previous findings where polar solvent extracts exhibited significant activity (Akhtar and Mirza, 2015; Riaz et al., 2012). A linear correlation was found between TPC and TAC with correlation coefficient R² of 0.9137 (flower), 0.7454 (leaf), 0.9208 (stem), 0.761 (root) respectively. Similarly, a linear correlation was also found between TAC and TFC with correlation coefficient R² of 0.7972 (flower), 0.6669 (leaf), 0.6059 (stem) and 0.8658 (root) respectively. A significant relationship among the phytochemicals and antioxidant activities further confirmed that phenolics and flavonoids are responsible for antioxidant capabilities (Djeridane et al., 2006; Kumar et al., 2013).

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Fig. 4

TAC (Total antioxidant capacity µg AAE/mg DW) and TRP (Total reducing power µg AAE/mg DW) of various solvent extracts of flower, leaf, stem and root part of D. viscosa. Values are presented as mean ± Standard deviation from triplicate investigation. *IC₅₀ > 100 µg/ml. The columns with different superscript ^(a-h) letters show significantly (P < 0.05) different means.

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3.3.1.3 TRP

The ferric reducing power of various plant parts are presented in Fig. 4. The assay is based on reduction of ferric ion of potassium ferricyanide (Fe³⁺) to ferrous form (Fe²⁺) which results in formation of blue coloured complex that gives maximum absorbance at 593 nm (Komes et al., 2011). The highest reductive potential in flower part was exhibited by E extract (11.16 ± 0.46 µg AAE/mg DW). While in leaf, stem and root, WA extracts displayed maximum reducing power with 7.67 ± 1.29, 9.49 ± 0.62 and 5.84 ± 0.56 µg AAE/mg DW. Our results are in agreement with previous findings where polar solvent extracts of D. viscosa i.e. methanol, chloroform and water displayed reducing potential (Akhtar and Mirza, 2015; Shahzadi et al., 2012). In the current study, a significant correlation was also found between phenolic contents and reducing power with excellent correlation coefficient R² of (0.973 for flower), (0.946 for leaf), (0.9907 for stem) and (0.9803 for root). Our findings strengthen previous documented results that phenolics are associated with reductive potential of plants by donating electrons or hydrogen atom that leads to termination of chain reaction by transforming free radical or active oxygen species to stable form (Brewer, 2011; Nasir et al 2020; Pal et al 2013; Zahra et al., 2017).

3.3.2.1 Antibacterial assay

The development of resistance against indiscriminate use of existing antibacterial drugs and treatment challenges against the bacterial infections has become major concern. Therefore, there is need to find alternative antimicrobials from the natural sources especially plants. (Ahmad and Beg, 2001). The antibacterial activity of plants largely depends on the solvent employed for extraction, part of plant used, organism tested, collection period and geographical location (Esmaeel and AL-Jobori, 2011; Kaushik and Goyal, 2008). The extracts with zone of inhibition (ZOI) ≥ 8 mm and ≤ 14 mm are considered moderately active, extracts with ZOI ≥ 15 mm and ≤ 21 are considered active, while the highly active extracts exhibited ZOI ≥ 22 mm and ≤ 29 mm. The EA extract of leaf was active against S. aureus with 19 ± 0.75 mm ZOI. Against B. subtilis nH and C extracts of root and flower were highly active with 22 ± 1.50 and 29 ± 0.84 mm of growth inhibition zone respectively. The chloroform and ethanol extracts of flower showed high activity against K. pneumoniae with zone of inhibition 22 ± 0.35 and 22 ± 0.98 mm respectively. The ethanol extract of leaf displayed highly active against P. aeruginosa with 22 ± 1.1 mm of growth zone inhibition. The C and E extracts of root displayed the best inhibitory potential against B. subtilis with 22 ± 0.75 and 22 ± 1.0 mm ZOI (Table 3). The results are strongly correlated with already documented reports where both polar and non-polar solvent extracts exhibited remarkable antibacterial activity against gram positive and gram negative strains (Esmaeel and AL-Jobori, 2011; Getie et al., 2003; Prakash et al., 2012; Ramamurthy et al., 2013). Hydroxylated phenolic compounds such as caffeic acid and rutin have shown toxicity to microorganisms (Lupaşcus et al., 2010). The antibacterial activities can be associated with phenolic and flavonoid contents of plant as they have reported antimicrobial properties and their possible mechanism involves the ability of these compounds to inhibit growth of bacteria by complexing with cell wall (Mehmood et al., 2013). The other possible mechanism involves blockade of cytoplasmic membrane functions, nucleic acid synthesis and metabolism (Cushnie and Lamb, 2005). The observed activity can be correlated to caffeic acid and rutin detected in HPLC analysis of subject plant. The antibacterial results suggested that D. viscosa has the potential to be used as herbal remedy for bacterial infections and can be a potential candidate for bioactivity guided isolation of novel antimicrobial agents.

3.3.2.2 Antifungal assay

The antifungal potential of D. viscosa was evaluated against five different fungal strains as shown in Table 4. The results have shown that almost all the plant parts have moderate antifungal activity against the tested fungal strains with a zone of growth inhibition ranging from 7 mm to 12 mm. The flower extracts showed sensitivity only against A. flavus with maximum inhibition zone of 11 mm displayed by WA extract. The leaf part indicated that maximum inhibition zone was observed in MC extract (11 ± 0.88 mm) against A. fumigatus. While against F. solani MC, EC, A, WA and WM extracts have shown maximum inhibition zone of 12 mm. The A extract showed maximum inhibition zone of 12 ± 0.77 mm against Mucor sp. Similarly, the stem extracts were active against A. flavus, A. niger and A. fumigatus. The root part revealed that maximum activity was demonstrated by WM extract (10 ± 0.69 mm) against A. flavus, A (12 ± 0.98 mm) and C (12 ± 0.43 mm) extracts against A. fumigatus, MC extract (12 ± 1.24 mm) against F. solani, EC (12 ± 0.92) and WM extracts (12 ± 0.23) mm against Mucor sp. The extracts were least active against A. niger. The antifungal activities of plants can be associated with secondary metabolites including phenols, flavonoids, phenolic glycosides, saponins and cyanogenic glycosides (Quiroga et al., 2001). It might be attributed that the observed antifungal activities are due to the phenols and flavonoids quantified in this plant. Moreover, in this study, the variable trend in activity was observed both in polar and non-polar solvent extracts. The results are in agreement with previous studies that report antifungal activities in both polar and non-polar solvent extracts (Pirzada et al., 2010; Ramamurthy et al., 2013).

3.3.3.1 Brine shrimp cytotoxicity

The preliminary cytotoxicity profiling of D. viscosa was assessed by employing brine shrimp lethality assay that is commonly employed to detect the antitumor, antimicrobial, insecticidal and anticancer properties of test samples (Ramamurthy et al., 2013). It is proposed that shrimp larvae behave similar to mammalian cells so the toxicity against these larvae can be inferred as cytotoxicity to mammalian cells (Ullah et al., 2012). The results of brine shrimp lethality assay are represented in the Table 5. The leaf extracts of D. viscosa possess toxicity against shrimp larvae with maximum lethality shown by EEA extract with 70% mortality (LC₅₀ = 95.46 µg/ml). Both polar and non-polar extracts exhibited lethality against brine shrimps. The concentration dependent toxic effect of test extracts was observed in this assay which is in accordance with previous reports (Prakash et al., 2012). The cytotoxicity of test extracts might be due to defensive secondary metabolites that include flavonoids, phenols, saponins or other compounds (Aziz et al., 2013). The activity might be attributed to the polyphenols detected by HPLC analysis. Our results are contradictory to previous findings where stem and root extracts also exhibited toxicity against Artemia salina larvae and results were more prominent in polar solvent extracts than non-polar solvent extracts (Prakash et al., 2012).

3.3.3.2 In vitro cytotoxicity against THP-1 human leukemia cell line

The cancer represents a complex disease generally related with an array of increasing effects both at cellular and molecular levels. Cancer is the major cause of death after cardiovascular diseases. The anticancer agents from natural sources are of particular interest due to decreased toxicity and more therapeutic efficacy (Kaur et al., 2011). Over 3000 plants have been used in cancer treatment. The anticancer drug discovery started in 1950 with the discovery of vinca alkaloids and podophyllotoxins, the first anticancer agents discovered and isolated from plant (Cragg and Newman, 2005). The plants abound in limitless reserves of secondary metabolites (flavonoids, alkaloids and terpenes) that possess significant anticancer properties (Scheck et al., 2006). Following the results of brine shrimp assay the plant was further evaluated to determine its in vitro cytotoxic potential against THP-1 human leukemia cell line. It can be seen from the Table 5, that the leaf part displayed prominent results with maximum inhibition revealed by W extract i.e., 71.0 ± 0.47% inhibition (IC₅₀ = 8.5 µg/ml) followed by WM extract with 67.53 ± 1.63% inhibition (IC₅₀ = 11.5 µg/ml). In current investigation, the flower extracts displayed no activity against leukemia cells. The stem part displayed the maximum inhibition by EEA and MEA extracts with 91.50 ± 1.70 and 90.0 ± 1.70% inhibition (IC₅₀ = 3.4 and 3.8 µg/ml respectively). While in root extracts MC was found to be the most potent with 70.0 ± 1.25% inhibition (IC₅₀ = 9.4 µg/ml). The other extracts also displayed cytotoxicity with an IC₅₀ > 20 µg/ml. The activity was more in moderately polar solvents as compared to highly polar solvents. The results are in accordance with previous studies in which polar solvent extract (methanol and water) have shown least activity against human amniotic epithelial cell line i.e. FL-cells (Mothana et al., 2010). The cytotoxicity of MEA and EEA can be correlated to apigenin detected by HPLC analysis of plant. Apigenin (4′,5,7-trihydroxyflavone), a dietary flavonoid was reported to have strong cytotoxic effects in various cancer cell lines (breast cancer, colon cancer, lung cancer and neuroblastoma) (Valdameri et al., 2011) and causes apoptosis in monocytic leukaemia cells of human (Arango et al., 2012). According to Jayasooriya et al. (2012), apigenin causes direct toxicity to human leukaemia cells through caspase pathway activation. As the crude extract contains multiple varieties of compounds so it is not justified to assign observed activity to a single compound. The activity might be due to other polyphenols (such as rutin, gallic acid, and myricetin) which have reported anticancer properties. Moreover, the activity in non-polar extracts might be due to the compounds other than those detected by HPLC analysis. This is the first report (to the best of our knowledge) that reports the in vitro cytotoxicity against THP-1 human leukemia cell line. The results suggest that the stem extracts of D. viscosa have significant cytotoxic potential which demands further investigation to isolate bioactive compounds responsible for the observed activity.

3.3.3.3 In vitro cytotoxicity against Hep G2 hepatoma cell line

The significant activity displayed by D. viscosa extracts against human leukaemia cell line, enabled us to check the in vitro cytotoxicity against Hep G2 human hepatoma cell line. The hepatocellular carcinoma (hepatoma) is one of the primary liver cancers. The hepatoma is the fifth most common cancer and third major cause of cancer related deaths (Forner et al., 2006). The hepatoma occurs due to hepatocellular damage caused by reactive oxygen species and generation of chronic inflammation. The main constraint in treatment of hepatocellular carcinoma is the increased tendency to develop multidrug resistant proteins and reduction of apoptotic proteins. Hence it demands the discovery of new chemotherapeutic agents from natural sources (Machana et al., 2012). The results revealed that moderate antiproliferative activity was displayed by all parts with the highest inhibition shown by nH (55.43 ± 0.55%) and WM (54.61 ± 1.7%) extract of flower, AEA extract of leaf (61.75 ± 0.66%) and WA extract of stem (47.8 ± 0.66%) inhibition (Table 5). The EEA extract was found to be the most potent with an average inhibition of 69.46 ± 1.67% (IC₅₀ = 20 µg/ml). The activity was observed both in non-polar and highly polar extracts which suggests that the plant contain a wide variety of phytochemicals having a diverse structural composition. The results are in relevance with the previous findings where M (polar solvent) extract of D. viscosa leaf exhibited activity with an IC₅₀ greater than 50 µg/ml against MDBK (Madin-Darby bovine kidney), MCF-7 (human breast adenocarcinoma), WEHI-164 (mouse fibrosarcoma), Hep G2 (human hepatocellular carcinoma) and A-549 (non-small cell lung carcinoma) cell lines (Esmaeili et al., 2014). The observed anticancer activity might be due to apigenin, rutin, quercetin or other polyphenols detected in plant. According to Cai et al. (2011), the apigenin inhibits the growth of hepatoma cells by arresting the G2 or M phase of cell cycle and induce apoptosis of hepatoma cells. So apigenin might be responsible for the observed activity. This demands the further assessment of active extracts by employing bioactivity guided isolation of active compound responsible for cytotoxic potential. This is the foremost report (to the best of our knowledge) on in vitro cytotoxicity of flower, stem, and root extracts of D. viscosa.