Polarity-guided phytochemical extraction, polyphenolic characterization, and multimode biological evaluation of Seriphidium kurramense (Qazilb.) Y. R. Ling

2.4.1 Extract recovery

Weight of dried extracts was taken properly, and percent recovery of crude extracts was determined by applying following formula:

Extract recovery (%w/w) = WB/WE × 100.

Where WB = Weight of plant biomass and WE = Weight of crude extract after drying.

2.5.1 Determination of total phenolic content (TPC)

Total phenolic content (TPC) was calculated employing Folin-Ciocalteu (FC) reagent corresponding to an established protocol, previously illustrated (Ul-Haq et al., 2012). Test sample (20 μl) from the stock extracts (4 mg/ml) and 90 μl of FC reagent were transferred to the corresponding wells of microplate and incubated for 5 min at room temperature. sodium carbonate (6% w/v), 90 μl of was transferred to wells containing mixture. Incubation was done at 37 °C for 30  min and the absorbance was evaluated at 630 nm utilizing microplate reader (Biotech USA, microplate reader Elx 800). DMSO was used as negative control and gallic acid as positive control. Calibration curve was drawn using (y = 0.165x + 0.1631, R² = 0.9955) at varying concentrations (3.125–25 μg/ml) and total phenolic content was measured in microgram gallic acid equivalent per milligram of extract (μg GAE/mgE). Assay was carried out in triplicate.

2.5.2 Determination of total flavonoid content (TFC)

To quantify total flavonoids content, aluminum chloride based colorimetric method was followed (Ul-Haq et al., 2012). A mix containing 20 μl of the sample from the stock (4 mg/ml DMSO), 10 μl of 1.0 M potassium acetate, 10 μl of Alcl₃ (10% w/v in H₂O) and 160 μl of distilled water were poured in a 96-well plate. At room temperature, plate was incubated for half an hour and absorbance was measured at 415 nm using microplate reader. DMSO was treated as negative control and the experiment was run as triplicate. A calibration curve (y = 0.0508x + 0.0602, R² = 0.9947) of positive control (quercetin) was drawn at final concentration (2.5–40 μg/ml) and resultant flavonoid content was expressed in microgram equivalent of quercetin per milligram of extract (μg QE/mgE).

2.5.3 RP-HPLC quantitative analysis

High performance liquid chromatography system combined with DAD (diode array detector) was applied as formerly defined (Qasim et al., 2017, Ahmed et al., 2017) for detecting and quantifying of polyphenols in S. kurramense. HPLC system (Agilent Chem station Rev. B. 02–01-SRI 260) was provided with a Zorbex-C8 analytical column (5 µm particle size, 4.6 cm and 250 nm) associated with DAD, (Agilent technologies Germany). Binary gradient system comprises mobile phase A (methanol: water: acetic acid: acetonitrile in 10:85:1:5 ratio) and mobile phase B (acetonitrile: methanol: acetic acid in 40:60:1 ratio) was employed to achieve the detection of polyphenols. Flow rate was adjusted at 1 ml per minute and column was injected with 20 µl of the sample solution prepared in methanol. Before injecting new sample, column was tuned for 10 min. A gradient volume of (mobile phase B) was 0–50% in (0–20 min), 50–100% in (20–25 min) and 100% in last 25–30 min. Stock solutions of all the standards were freshly prepared within methanol and then further diluted to final concentrations of 10, 20, 50, 100, and 200 µg/ml. For each standard, the calibration curve was generated by utilizing the final concentration and peak area. Standard solutions and mobile phases were degassed first and then filtered out using 0.45 µm membrane filter (Millex-HV), whereas all samples were centrifuged and filtered prior to injecting into the HPLC system. The absorption was taken at certain wavelengths. Polyphenols were recognized by comparison of UV–Vis spectra and retention time of chromatographic peaks in comparison to reference standards at 257 nm for plumbagin, vanillic acid and thymoquinone, 279 nm for coumaric acid, catechin, syringic acid and gallic acid, 325 nm for apigenin, gentisic acid, caffeic acid, luteolin and ferulic acid and 368 nm for kaempferol, quercetin and myricetin. A wavelength of 360 nm was used as a reference wavelength so that in case of luminescence, the quantification of compounds showing inverted peaks may be double checked through their retention times and peak pattern with reference to 360 nm. Quantity of each polyphenol was quantified using calibration curves of standards i.e., for rutin (y = 9.547x + 22.217, R² = 0.997), vanillic acid (y = 6.357x + 12.113, R² = 0.998), ferulic acid (y = 19.51x-16.67, R² = 0.998), catechin (y = 7.878x-19.532, R² = 0.996), gallic acid (y = 23.573x-43.167, R² = 0.995), syringic acid (y = 9.532x + 13.754, R² = 0.998), myricetin (y = 5.2278x-6.3043, R² = 0.998), coumaric acid (y = 9.7644x + 14.281, R² = 0.999), caffeic acid (y = 25.093x + 92.465, R² = 0.995), gentisic acid (y = 12.21x-20.348, R² = 0.996) and apigenin (y = 18.111x + 25.565, R² = 0.997).

2.5.4 Antioxidant assays

2.6.1 Antibacterial susceptibility assay

Antibacterial potential of various extracts of S. kurramense, was evaluated according to an earlier described protocol (Peerzade et al., 2018). Pure cultures of both reference strains and resistant clinical isolates were evaluated. Reference bacterial strains included Escherichia coli (ATCC-25922), Salmonella typhi (ATCC-14028), Staphylococcus aureus (ATCC-6538), Staphylococcus epidermis (ATCC-12228), Pseudomonas aeruginosa (ATCC-27853), Bacillus subtilis (ATCC-6633), Klebsiella pneumoniae (ATCC-10031) and Salmonella enterica (ATCC-14028) while resistant clinical isolates included Pseudomonas aeruginosa, Escherichia coli, Methicillin Resistant Staphylococcus aureus (MRSA) and Staphylococcus hemolyticus. The standardized inoculum (5 × 10⁶ CFU/ml) of the refreshed bacterial cultures were swabbed onto the surface of respective plates containing Muller Hinton Agar (MHA) growth media utilizing sterile cotton swab under aseptic conditions. Afterwards, a well of 6 mm diameter was punched in each well using sterile cork borer. The bottom of each well was sealed by adding small amount of the medium to avoid leakage of the samples. Then 100 µl of the extract (20 mg/ml in DMSO) was poured in their respective wells which were previously labeled. After pouring the extracts the plates were left undisturbed at room temperature for the diffusion of extracts before the incubation. Cefixime at the final concentration of 5 µg (100 µl) and DMSO were used as positive and negative controls respectively in this assay. Inoculated petri plates were then kept at 37 °C in incubator for 24 h. After incubation, diameters of inhibitory zones were measured by vernier calipers in millimeters.

2.6.2 Antifungal susceptibility assay

For evaluation of antifungal potential of S. kurramense extracts, agar well diffusion method was performed in triplicate as explained earlier (Kalidindi et al., 2015) (Sales et al., 2016). The inoculum of test fungal strains Mucor species (FCBP-0300), Aspergillus niger (FCBP0198), Candida albicans (ATCC-10231), Fusarium solani (FCBP-0291) and Aspergillus fumigatus (FCBP-66), were harvested within Tween 20 solution (0.02% v/v) in distilled water and their turbidness was regulated with reference to 0.5 McFarland standard. Afterwards, 100 μl of standardized inoculum of all fungal strains was swabbed on previously labeled respective plates holding Sabouraud Dextrose agar (SDA). The inoculated plates were left undisturbed at room temperature for 20 min and then 6 mm diameter wells were created using a sterile cork borer. The bottom of each well was sealed by adding small amount of the medium to avoid leakage of the samples. Then each well was filled with 100 µl of respective extract (20 mg/ml in DMSO). DMSO and clotrimazole (4 mg/ml DMSO) were used as negative and positive controls in this assay. The seeded plates were then incubated at 28 °C for 24–48 h. Finally, the inhibitory zones around the samples and standard were observed and their diameters were recorded in millimeters.

To estimate the in-vitro antifungal activity of crude extracts against Aspergillus flavus (FCBP-0064); clinical isolates of Trichophyton rubrum, Trichophyton tonsurans, Aspergillus terreus, Alternaria alternata, Rhizopus arrhizus, and Fusarium dimerum, microbroth dilution method was utilized. The protocol developed by Clinical and Laboratory Standard Institute (CLSI) document M−38−A2 was followed (Wayne, 2008). RPMI-1640 was used as a culture medium for fungal growth. For inoculum preparation, 0.85% sterile saline was used. Spores were transferred through Pasteur pipette to saline solution (containing Tween-20) and the turbidity was corrected in accordance with McFarland standard 0.5. Firstly, 5 µl from each sample stock was transferred into the corresponding wells of microtiter plate, to which 195 µl of RPMI was added (500 µg/ml RPMI). Then two-fold serial dilutions were prepared containing 250, 125, 62.5, and 31.25 µg/ml RPMI in next wells. After sample addition, 100 µl of fungal suspension of respective strain was added to each well. Terbinafine at a concentration ranging from 0.5 to 0.001 µg/ml RPMI and Amphotericin B (16–0.313 µg/ml of RPMI) were used as positive controls and DMSO diluted within RPMI-1640 medium, (<1%) was used as negative control. Plates were then incubated at the temperature of 35 °C for varying periods for different fungi or till the growth appeared in control well. The results were recorded by visual aid using a magnifying glass. Assay was performed in triplicate and the lowest concentration inhibiting 100% growth was expressed as MIC.

2.6.3 Antiviral assay

By double layer agar technique, plaque reduction assay was accomplished as formerly depicted (Danesh et al., 2015). Concisely, 20 mg of each tested extract, at first, was dissolved within 1 ml of ethyl acetate, from which 100 µl was transferred to Eppendorf tubes and allowed to evaporate. To the dried extract 150 µl of known plaque forming unit PFU phage suspension was included and then at room temperature tubes were incubated for 50 min. After incubation, 250 µl of E. coli overnight culture was included to phage suspension and incubated at the temperature, 37 °C for about 20 min. Afterwards, mixture was mixed with 3 ml of softened agar and poured on a nutrient agar (NA) plate. Ethyl acetate combined with phage suspension and phage lysate treated with similar process (without plant extract) were used as two negative controls. Trifluridine was used as positive control. Afterwards, the plates were incubated for 24 h, at 37 °C and plaque was counted. Plaque reduction assay results were expressed in form of PFU ratio. $$\mathrm{P}\mathrm{F}\mathrm{U}\mathrm{r}\mathrm{a}\mathrm{t}\mathrm{i}\mathrm{o}=\frac{\mathrm{n}\mathrm{u}\mathrm{m}\mathrm{b}\mathrm{e}\mathrm{r}\mathrm{o}\mathrm{f}\mathrm{p}\mathrm{l}\mathrm{a}\mathrm{q}\mathrm{u}\mathrm{e}\mathrm{f}\mathrm{o}\mathrm{r}\mathrm{m}\mathrm{i}\mathrm{n}\mathrm{g}\mathrm{u}\mathrm{n}\mathrm{i}\mathrm{t}\mathrm{o}\mathrm{f}\mathrm{t}\mathrm{r}\mathrm{e}\mathrm{a}\mathrm{t}\mathrm{e}\mathrm{d}\mathrm{s}\mathrm{a}\mathrm{m}\mathrm{p}\mathrm{l}\mathrm{e}}{\mathrm{N}\mathrm{u}\mathrm{m}\mathrm{b}\mathrm{e}\mathrm{r}\mathrm{o}\mathrm{f}\mathrm{p}\mathrm{l}\mathrm{a}\mathrm{q}\mathrm{u}\mathrm{e}\mathrm{f}\mathrm{o}\mathrm{r}\mathrm{m}\mathrm{i}\mathrm{n}\mathrm{g}\mathrm{u}\mathrm{n}\mathrm{i}\mathrm{t}\mathrm{o}\mathrm{f}\mathrm{n}\mathrm{e}\mathrm{g}\mathrm{a}\mathrm{t}\mathrm{i}\mathrm{v}\mathrm{e}\mathrm{c}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{o}\mathrm{l}}$$ $$\mathrm{P}\mathrm{F}\mathrm{U}(\%)=ValueofPFUratio\times 100$$

2.7.1 Alpha amylase inhibition assay

Starch iodine based colorimetric assay was employed to gauge the α-amylase inhibitory capability of test extracts following the standard protocol (Zhao et al., 2019). Reaction mixture comprising 25 μl of enzyme (0.14 U/ml), 15 μl of phosphate buffer (pH 6.8), 40 μl of starch solution (2 mg/ml in potassium phosphate buffer) and 10 μl of the extract (4 mg/ml DMSO) was incubated for half an hour at 50 °C in 96 well plate then 1 M HCl (20 μl) was added as a stopping solution. Subsequently, 90 μl iodine reagent (5 mM potassium iodide and 5 mM of iodine) was moved to each well. Negative control having no extract while blank did not contain amylase enzyme and buffer was added to replace each of them. Positive control of this assay was Acarbose (250 μM). Following incubation, the absorbance was taken at 540 nm. Results were expressed as percent of α-amylase inhibition per mg of extract using the given equation: $$Percent\alpha -amylaseinhibition=(A{b}_s-A{b}_n)/(A{b}_b-A{b}_n)\times 100$$

Where Ab_n = Absorbance of the negative control, Ab_s = Absorbance of the sample and Ab_b is Absorbance of blank.

2.7.2 Alpha-glucosidase inhibition assay

Protocol reported earlier (Nair et al., 2013) was used for execution of this assay. Initially, (20 mM) of PNG (25 μl), 5 μl the tested sample and 69 µl 50 mM, phosphate buffer (pH 6.8) was mixed in the corresponding wells of 96 well plate and then addition of 1 µl of the enzyme solution was done. After then, immediately the absorbance was measured employing microplate reader at 405 nm. The mix was then incubated at 37 °C for a period of 30 min. 100 µl of sodium bicarbonate (0.5 mM) solution was transferred to stop the reaction. DMSO was used instead of test samples as a negative control and tested sample was replaced by Acarbose solution as a positive control, following the same procedure as described. Whole assay was run as triplicate analysis and absorbance was measured at 405 nm. Percent enzyme inhibition was calculated by employing the following formula: $$\mathit{Percent}\mathit{enzyme}\mathit{inhibition}=A_c-A_s/A_c\times 100$$

Ac = Absorbance of control, As = Absorbance of sample.

2.7.3 Urease inhibition assay

Jack bean urease inhibition assay was utilized as described (Bai et al., 2015), 20 µl from 5 Units per ml of the stock solution was mixed with 30 µl of the assay buffer (comprising K₂HPO_4, LiCl, urea and EDTA, at the concentration of 100 mM, 1 mM, 0.01 M, and 0.01 M respectively, pH 8.2), approx. 20 µl of the extract in 96 well plate and then this mixture was incubated for 30 min at 37 °C. This was followed by the addition of 50 µl of alkali reagent (comprising 0.5% NaOH and 0.1% w/v of sodium hypochlorite (NaOCl)) and phenol reagent (sodium nitroprusside, 0.005% w/v and 1% w/v phenol) and the incubation was done for about 20 min at the temperature 25 °C. Absorbance of the mixture was calculated at 625 nm using a microplate reader (OptiMax, Tunable) and all the extracts were analyzed at the concentration range 62.5–500 µg/ml.

In this experiment, thiourea was used as a standard for urease inhibitor. Assay was repeated thrice, and inhibitory activity was determined using the following formula: $$\mathit{Percent}\mathit{Inhibition}\mathit{of}\mathit{urease}\mathit{activity}=A{b}_{control}-A{b}_{sample}/A{b}_{control}\ast 100$$

Where, Ab_sample is the absorbance of sample and Ab_control is the absorbance in presence or absence of the sample. Graph pad Prism version 5.0 was used for calculation of IC₅₀.

2.8.1 Hemolytic assay

Hemolytic was performed by following protocol as described earlier (Singh and Katoch, 2020). Various concentrations of S. kurramense extracts (250 and 500 µg/ml) were used to determine their hemolytic potential against human erythrocytes. The hemolytic potential i.e., the concentration that can induce 50% (HC₅₀) or 10% (HC₁₀) hemoglobin release from RBCs due to exposure to extracts were determined using table curve 2D v5.01 software.

Erythrocytes were isolated after centrifugation of fresh human blood and rinsed three times employing sterile phosphate buffer saline solution (pH 7.4). Erythrocytes pellet was resuspended in phosphate buffer solution to achieve a 5% solution. RBCs suspension of 50 μl was moved to 96 well plate having 150 μl of samples at different concentrations. Incubation was done for 30 min at room temperature and then centrifuged for 20 min at 2000 rpm. Supernatant (100 μl) from the mixture was transported to a fresh 96 well plate and the absorbance was estimated at wavelength, 360 nm. 0.1% Triton X100 was employed as positive control and hemolysis was computed in percent using the formula. $$\%Hemolysis=\frac{\mathit{Absorbance}\mathit{of}\mathit{the}\mathit{sample}-Absorbanceofnegativecontrol}{\mathit{Absorbance}of\mathit{positive}\mathit{control}-Absorbanceofnegativecontrol}\ast 100$$

(Wani et al., 2017).

The experiment was done in triplicate.

2.8.2 Brine shrimp lethality assay (Cytotoxicity Testing)

For the assessment of primary biocompatibility profile of S. kurramense, 24 h lethality test was accomplished against brine shrimp larvae (Artemia salina) as illustrated earlier (Ul-Haq et al., 2012). Eggs of A. salina (Ocean star, USA) were incubated in a specifically constructed two-compartment plastic tray for hatching period of 24–48 h within simulated sea water (38 g/ liter augmented with 6 mg/liter of the dried yeast) under illumination and heat at 30–32 °C). Afterwards, using Pasteur pipette, ten mature phototropic nauplii were shifted to the wells of microtiter plate. Extract containing ≤ 1% DMSO in the sea water at concentrations of 200, 100, 50 and 25 μg/ml was added to each well carrying the shrimp larvae. Final volume of 300 μl was kept in each well. Positive control (1.25–10 µg/ml of doxorubicin) and negative controls (1% DMSO in sea water) were used. After time duration of 24 h, dead shrimps were tallied, and percentage death was confirmed. Median lethal concentration (LC₅₀) was processed using table curve 2D v5.01 software. The whole experiment was performed thrice.

4.2.1 Total phenolic content (TPC)

The total gallic acid equivalent phenols in S. kurramense extracts showed a range of 24.44 ± 0.34 to 1.20 ± 0.13 μg GAE/mgE. The highest TPC (24.44 ± 0.34 μg GAE/mgE) was recorded for Sk-Aq whereas lowest quantity (1.20 ± 0.13 μg GAE/mgE) was found to be present in Sk-nH. The detected quantity of total phenolic content in aqueous extract is also in accordance with another report published on Seriphidium oliverianum (Abbas et al., 2021). The phenolic content was observed to decrease in the subsequent order: Sk-Aq > Sk-M > Sk-EA > Sk-nH (Fig. 1). A broad array of solvents are needed for their extraction since the polyphenol polarities range from polar to non-polar, thus. Thus, polarity performs a vital role in enhancing the phenols solubility. A substantial relationship among antioxidant capacity and total phenolic content was observed in earlier investigations, suggesting phenolic compounds as the major sponsors to antioxidant potential of plants (Fernandes de Oliveira et al., 2012). Polyphenols have antioxidant (Kumar et al., 2013), antifungal (Chen et al., 2013), anticancer and antibacterial (Móricz et al., 2018) potential. Plant phenolics combat oxidative stress and the cell death by scavenging or chelating free radicals and trace elements (Cheynier, 2012) therefore the phenolic content was assessed to extrapolate the possible role of S. kurramense in infectious diseases. To the best of our knowledge, determination of TPC and TFC was carried out for the first time in current study.

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Fig. 1

Total phenolic content and total flavonoid content of S. kurramense extracts.Values are presented as mean ± standard deviation from the triplicate investigation. The columns with different superscript (^a-c) letters show significantly (P < 0.05) different means.

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4.2.2 Total flavonoid content (TFC)

Total flavonoid content of S. kurramense extracts in terms of μg QE/mgE are presented in Fig. 1. Among tested extracts, highest flavonoid content of 6.87 ± 0.25 μg QE/mgE was recorded in Sk-Aq. Same quantity of TFC (3.07 ± 0.29 and 3.07 ± 0.21 μg QE/mgE) was detected in Sk-M and Sk-EA. Lowest content of TFC (1.17 ± 0.22 μg QE/mgE) was observed in Sk-nH. Compounds with hydroxyl groups such as flavonoids, act through scavenging or chelating processes and are accountable for the free radical scavenging activity of plants (Ahmed et al., 2017). The potential mechanisms of antioxidant actions by flavonoids involve scavenging ROS, protecting and regulating antioxidant defenses and inhibition of ROS formation either by inhibiting enzymes or chelating the trace components (Zahra et al., 2017). Therefore, the present flavonoid determination in the extracts of S. kurramense suggests that it would be a beneficial antioxidant and would come forward as a successful candidate against infectious diseases by bolstering immune system through palliation of oxidative stress.

4.2.3 RP-HPLC analysis

Polyphenols were quantified by RP-HPLC profiling, using 15 standards. The chromatographic fingerprints of analytes and standards are shown in Fig. 2 and Table 2 depicts quantified values of 10 detected phenolic constituents. Sk-EA contained maximum concentration of Luteolin (3.82 ± 0.11 µg/mgE) followed by Ferulic acid (3.05 ± 0.03 µg/mgE). Gentisic acid (6.44 ± 0.01 µg/mg) and Quercetin (4.39 ± 0.01 µg/mgE) were found substantially in Sk-Aq whereas Sk-M was found to be rich in Luteolin (3.90 ± 0.03 µg/mgE) and Apigenin (3.72 ± 0.03 µg/mgE) The detected polyphenols such as apigenin and ferulic acid have established bioactive profile which further potentate the medicinal value of S. kurramense. Phenolic acids are known to have antioxidant (Cheynier, 2012), anti-inflammatory (Cory et al., 2018) and antimicrobial properties and are stated to be more potent against Gram negative bacteria as compared to Gram positive bacteria (Cetin-Karaca, 2011). In the present study considerable antibacterial activity against gram negative as compared to gram positive bacteria has been observed. Previously apigenin’s antibacterial (Kim et al., 2020), antifungal (Lee et al., 2018) and anti-parasitic capability has been reported (Wang et al., 2019b). Furthermore, ferulic acid was found to exert an immediate and constant inhibition of Coronobacter sakazakii (Shi et al., 2016). It has also been indicated in various studies that polyphenols have strong antiviral activities through various mechanisms (Musarra-Pizzo et al., 2019) (Bai et al., 2016) (Agrawal et al., 2020).

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Fig. 2

High performance liquid chromatography fingerprints of S. kurramense extracts. A = standards, B = Sk-EA, C = Sk-M, D = Sk-Aq1. Vanillic acid 2. Plumbagin 3. Thymoquinone 4. Gallic acid 5. Catechin 6. Syringic acid 7. Coumaric acid 8. Gentisic acid 9. Caffeic acid 10. Ferulic acid 11. Luteolin 12. Apigenin 13. Myricitin14. Quercetin 15. Kaempferol.

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4.3.1 Antioxidant assays

4.3.1.1

4.3.1.1 Free radical scavenging activity (FRSA)

Reactive oxygen species (ROS) and free radicals are produced as a result of biological processes. (Vaghasiya et al., 2011). They play an imperative part in oxidative stress linked to the pathogenesis of many bacterial, viral and chronic diseases (Aiyegoro and Okoh, 2010) (Kumar et al., 2008). Plants owing to their ROSs scavenging ability represent themselves as valuable suppliers of organic antioxidants. (Riaz et al., 2012). The percent free radical scavenging activity (%FRSA) of S. kurramense extracts was evaluated by measurement of their capability to scavenge stable free radical DPPH. Results of this assay are exhibited in Fig. 3.

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Fig. 3

Percent free radical scavenging activity of S. kurramense extracts. Values are presented as mean ± standard deviation from the triplicate investigation. The columns with different superscript (^a-e) letters show significantly (P < 0.05) different means.

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Highest free radical scavenging activity was exhibited by Sk-Aq (85.87 ± 1.00%) followed by Sk-M (54.01 ± 1.02%) and Sk-EA (13.53 ± 0.59%). The least FRSA (10.33 ± 0.55%) was displayed by Sk-nH. The plant polyphenols and flavonoids, among the plant components are deemed strong free radical scavengers, (Farah et al., 2018) and serve as prospective alternate to faux antioxidants owing to the safety plus efficacy (Tzima et al., 2018). As mentioned earlier free radicals are responsible for oxidative stress linked to various diseases, therefore, scavenging free radicals can be linked to mitigation of oxidative stress in these infections. Thus, the antioxidant activity can be correlated with antimicrobial activity as well. Free radical scavenging activity in this research could be attributed to polyphenols detected such as quercetin (OZGEN et al., 2016), gentisic acid (JOSHI et al., 2012) in Sk-Aq which displayed strong activity, apigenin in Sk-Aq (Kumar et al., 2013) and Sk-M and luteolin (ROY et al., 2015) in Sk-EA and Sk-M which have exhibited moderate activities. The antioxidant agents always show complex and variable mode of action hence a single method cannot be relied to determine antioxidant capability of test samples (Sini et al., 2011).

4.3.1.2

4.3.1.2 Total antioxidant capacity

Total antioxidant capacity of S. kurramense extracts was established by a colorimetric assay based on phospho-molybdenum which involves development of green complex (Fatima et al., 2015). The results are summarized in Fig. 4. Highest antioxidant potential was exhibited by Sk-EA (243.5.0 ± 1.12 µg AAE/mgE), followed by Sk-Aq (238.3 ± 0.91 µg AAE/mgE) and Sk-M (140.4 ± 0.76 µg AAE/mgE) while lowest TAC was recorded for Sk-nH (1.2 ± 0.34 µg AAE/mgE). Prominent total antioxidant capacity denotes that test extracts would play the role of promising antioxidants by reducing oxidative stress which is the foundation of many diseases by damaging the immune functions of biological systems (Apak et al., 2016).

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Fig. 4

Graphical representation of total antioxidant capacity and total reducing power of S. kurramense extracts. Values are presented as mean ± standard deviation from the triplicate investigation. The columns with different superscript (^a-d) letters show significantly (P < 0.05) different means.

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4.4.1 Antibacterial assay

Improper and misuse of antibiotics lead to the progression of microbial resistance, rendering the treatment of bacterial infections a major challenge for health professionals. Bacteria also have evolved certain mechanisms which contribute to resistance by either chemical or enzymatic modifications (Todar, 2011). Plants provide protection against infective diseases and resistance due to maximum safety and minimum side effects. (Ruddaraju et al., 2020). Therefore, screening natural sources especially plants for finding alternative antimicrobials is becoming a dire need at the present time (Malik et al., 2022).

Results of antibacterial activity of extracts are shown in Table 3. Sk-EA was found to be more effective against all tested strains of bacteria. The inhibitory zone diameter values ranged from 8.33 ± 1.52 to 21.33 ± 0.76 mm. The highest value (21.33 ± 0.76 mm) was recorded for Sk-EA against K. pneumoniae whereas least against S. enterica (8.33 ± 1.52 mm) by Sk-M. For resistant clinical isolates inhibitory zone diameter values ranged from 7.0 ± 0.00 to 11.83 ± 0.76 mm. The tested extracts with better antibacterial action against gram negative bacterial strains in comparison to gram positive however noteworthy activity against gram positive bacteria is also apparent. Along with that S. kurramense extracts have shown significant activity against resistant clinical isolates against resistant clinical isolates that have shown resistance against contemporary antibiotics proving their superlative activity.

Results of the current study strongly correlate with already documented reports where extracts of the plant understudy displayed significant antibacterial activity versus gram positive and gram negative bacterial strains (Ahmad et al., 2018). The results of antimicrobial potential of S. kurramense can be correlated with phenolic acids such as apigenin, ferulic acid, luteolin and quercetin detected in different extracts of the plant. Phenolic acids are known to have antimicrobial activity (Cetin-Karaca, 2011). Various degrees of efficacy are reported on apigenin’s antifungal, anti-parasitic and antibacterial potential, antibacterial mechanism being perturbance of cell membrane (Wang et al., 2019b) (Kim et al., 2020). Furthermore, ferulic acid was found to exert an immediate and constant inhibition of Coronobacter sakazakii by altering the cell membrane permeability (Shi et al., 2016). Luteolin exerts antibacterial action through affecting membrane permeability and by inhibiting the activity of DNA topoisomerases I and II (Wang and Xie, 2010). Another polyphenol quantified is quercetin which exhibits antibacterial action by altering the cell permeability, destroying bacterial cell wall, reducing enzyme activities, influencing protein synthesis and expression, and inhibiting synthesis of nucleic acids (Yang et al., 2020). In line with the results of antibacterial assay, it is apparent that S. kurramense extracts and/or it’s isolated phytochemicals may play a significant role in lead development to combat antibacterial drug resistance.

4.4.2 Antifungal assay

S. kurramense was assessed against twelve different fungal strains and the Table 4a. and Table 4b) depict the results of the assay. Noteworthy antifungal activity was recorded against C. albicans (31.33 ± 0.57 mm) by Sk-EA followed by Sk-nH (30.83 ± 0.76 mm) and moderate activity against A. fumigatus (13.66 ± 0.58 mm) by Sk-Aq. Significant activity against F. dimerum (MIC value of 62.5 µg/mL) was recorded for Sk-EA and Sk-M. Previous study reported noteworthy antifungal activity of EA extract against A. flavus and A. niger at 5 mg/ml concentration (67.2 and 32.2% inhibition respectively) however in the present study Sk-EA showed no activity against both mentioned fungal strains Therefore, the result of antifungal activity of ethyl acetate extract against A. flavus and A. niger of the previous study conducted (Ahmad et al., 2018) is not in agreement with the results of the present study. Several studies showed that secondary metabolites produced by plants have inhibitory effects on several pathogenic fungi (Castillo-Reyes et al., 2015) (Hossein and Maldonado, 1982) (Arif et al., 2009). The antifungal activity shown by various extracts of S. kurramense may be attributed to the polyphenols detected within extracts particularly quercetin in Sk-M and luteolin in both Sk-EA and Sk-M. Quercetin improves fluconazole-resistant C. albicans stimulated apoptosis by regulation of quorum sensing and causing mitochondrial dysfunction (Yang et al., 2020) (Rocha et al., 2019). Likewise, luteolin produces ultrastructural and morphological changes in the fungus (Báidez et al., 2006). The structure of the phenolics allows them to disperse through microbial membranes and penetrate the cell, where they intervene in metabolic pathways of fungi by intruding in the ergosterol synthesis, glucan, proteins,chitin and glucosamine in fungi (Ansari et al., 2013). (See Table 4a. and Table 4b).

4.4.3 Antiviral assay

The antiviral potential of S. kurramense extracts was tested against phage virus. Among the extracts, Sk-EA was found as highly potent that reduced PFU to zero percent, followed by Sk-nH and Sk-Aq. Results are shown in Table 5. Several studies have confirmed the antiviral potential of plants (Denaro et al., 2020, Jaime et al., 2013). The antiviral activity of the extracts can also be attributed with the presence of phenolic acids in extracts with documented antiviral activity such as luteolin quantified in Sk-EA which in recent studies has shown that it inhibits the secretion of Hepatitis B e-antigen (HBeAg) in HepG2.2.15 cells that bear the HBV genome and Hepatitis B surface antigen (HBsAg) suggests that luteolin acts as a potential anti-HBV agent (Bai et al., 2016), apigenin quantified in Sk-EA and Sk-Aq, inhibits the replication of EV71 which causes hand, foot and mouth disease (HFMD) by disturbing association of viral RNA with trans-acting factors and regulating cellular JNK pathway (Dai et al., 2019) and quercetin quantified in Sk-Aq, is effective against various viral infections by either blocking the virus entry or inhibiting viral replication enzymes such as viral polymerases (Agrawal et al., 2020).

4.5.1 α-amylase inhibition assay

Diabetes mellitus, a metabolic disorder of worldwide prevalence is characterized by hyperglycemia. The inhibitors of alpha amylase and alpha glucosidase enzymes which are responsible for metabolism and absorption of glucose are believed to be effective agents for controlling and managing diabetes mellitus (Tadera et al., 2006). Adverse effects linked with insulin and oral hypoglycemic agents, used to manage blood glucose levels, compelled the researchers to find alternate sources of new anti-diabetic therapeutics (Liyanagamage et al., 2020). Hence, there is a dire need to explore organic sources for enzyme inhibitors.

The results of alpha amylase inhibition assay are displayed in Table 6. Highest inhibition was shown by Sk-nH followed by Sk- M with IC₅₀ values of 96.26 ± 0.44 and 109.10 ± 0.24 μg/ml respectively. IC₅₀ value of Sk-Aq was 113.50 ± 0.37 μg/ml and that of Sk-EA was 117.0 ± 0.53 μg/ml.

The alpha amylase inhibiting potential of S. kurramense can be linked with presence of quercetin and ferulic acid as they have been detected in Sk-EA, Sk-M and Sk-Aq. Phenolics and flavonoids have the ability to inhibit alpha amylase enzyme (Etxeberria et al., 2012). Plant based phenolic compounds like quercetin and ferulic acid, catechin, are accountable for regulation of glycaemia by secreting insulin, increasing glucose uptake, α-glucosidase, and α-amylase inhibition and further inhibiting lipid peroxidation (Barone et al., 2009). The α-amylase inhibitory proficiency of S. kurramense combined with its antimicrobial and antioxidant attributes propose it to be a suitable candidate particularly where infections and metabolic disorders along with oxidative stress are implicated in disease progression such as diabetes.

4.5.2 α-glucosidase inhibition assay

Like alpha amylase inhibition another strategy developed to deal with type-2 diabetes is alpha-glucosidase’s inhibition. Generally, nutritional starch is processed by α-amylase into maltose and dextrin, while α-glucosidase in digestive tract hydrolyzes these carbohydrates into glucose increasing the blood glucose level (Zheng et al., 2020). Synthetic inhibitors are generally linked with digestive side effects. Thus, developing the inhibitors from natural sources present an alternate option for the management of hyperglycemia. In recent times, several studies have been performed to detect inhibitors of alpha glucosidase from natural resources (Assefa et al., 2019).

Glucose modulatory potential of S. kurramense was additionally evaluated by α-glucosidase inhibition assay. Results of this assay are summarized in Table 6. Sk-nH exhibited highest α-glucosidase inhibition i.e., 60.51 ± 0.37% with an IC₅₀ value of 48.35 ± 0.22 μg/ml whereas, remarkable enzyme inhibition potential was also shown by Sk-EA (57.13 ± 0.23%) and Sk-M (56.66 ± 0.43%) with IC₅₀ values of 58.02 ± 0.38 and 30.44 ± 0.47 μg/ml respectively at 100 μg/ml concentration. At the same concentration Sk-Aq exhibited comparatively low enzyme inhibition potential (47.10 ± 0.27%). No literature is available on alpha glucosidase inhibition of the plant understudy.

A number of studies indicated the alpha glucosidase inhibitory potential of polyphenols (Assefa et al., 2019) (Seo et al., 2015) (Benalla et al., 2010). Alpha glucosidase inhibitory effects of tested extracts, therefore, can be credited to polyphenols which have been detected and have documented glucosidase inhibitory activities such as luteolin (Yan et al., 2014), apigenin (Zeng et al., 2016) and ferulic acid (Zheng et al., 2020) quantified in Sk-EA and Sk-M. The results are consistent with the antioxidant activities of Sk-EA and Sk-M. Antioxidant action along with enzyme inhibition potential of S. kurramense prospects its use in infectious diseases.

4.5.3 Urease inhibition assay

The results for assessment of urease inhibition assay and IC₅₀ of the extracts are presented in Table 6. All the extracts exhibited anti-urease activity. Sk-M and Sk-Aq were found to be more potent by showing a percent inhibition value of 84.15 ± 0.18 and 87.99 ± 0.19 respectively. Furthermore, Sk-EA and Sk-nH exhibited 75.08 ± 0.28 and 60.62 ± 0.39 percent inhibition respectively. Maximum urease inhibitory potential of Sk-M and Sk-Aq could possibly be attributed to high content of polyphenols owing to the recorded reports of their urease inhibition activity (Kafarski and Talma, 2018) such as apigenin (Nile et al., 2018), quercetin (Shabana et al., 2010) and gentisic acid (Malik et al., 2019). In this study, the manifestation of anti-urease activity by all the four extracts indicates the presence of urease inhibitors in them, another study reported strong anti-urease activity of pure compounds isolated from Seriphidium quettense (Imran Tousif et al., 2017).

Urease hydrolyzes urea into ammonia and carbon dioxide. In humans, it assists Helicobacter pylori to survive inside the stomach, which can cause gastric, duodenal ulceration, gastritis and even gastric cancer (Tarsia et al., 2018). It can colonize in acidic environment by secreting urease which facilitates the conversion of urea to ammonia and this liberated ammonia guards it from stomach acids (Amin et al., 2013). Therefore, inhibition of urease enzyme that would ultimately halt the production of ammonia can be regarded as an efficacious strategy to eradicate the microorganism by making it susceptible to stomach acid (Rego et al., 2018).

Various treatment options are there for the treatment of urease producing bacteria such as H. pylori, but limitations include various complications such as high toxicity, high cost of pretreatment, instability and bacterial resistance. Therefore, discovery of novel urease inhibitors developed from medicinal plants having low toxicity, few side effects, high bioavailability, and great stability, is gaining importance as an alternative treatment against H. pylori and other urease producing bacteria and would also address the issue of rapid development of antimicrobial resistance and would also be cost-effective (Pervaiz et al., 2019). Thus, S. kurramense, considering the results of this study, can be a prospective antimicrobial.