Phytochemical Profiling and Antioxidant Properties of Ziziphus lotus (L.) Fruits Supported by Xanthine Oxidase Inhibition and Molecular Docking

Abstract

Ziziphus lotus (L.) Lam., an extremophyte shrub native to the Mediterranean basin, yields underexplored fruits as a source of therapeutic agents. This study combined in vitro and in silico approaches to evaluate the antioxidant potential of Z. lotus fruits and predict their potential to inhibit xanthine oxidase (XO), a key enzyme in reactive oxygen species generation and oxidative stress-related pathologies. The ethyl acetate extract from the hydroalcoholic macerate was enriched in total phenolics (281.33 ± 1.5 μg GAE/mg) and flavonoids (127.26 ± 5.89 μg RE/mg) and displayed remarkable effects against the ABTS^•+ radical cation (IC₅₀ = 18.49 ± 1.47 μg/mL) and phenanthroline reducing power (A_0.5 = 8.38 ± 0.69 μg/mL), together with measurable xanthine oxidase inhibition (IC₅₀ = 170.4 ± 5.90 μg/mL). The compounds tentatively identified by full-scan UHPLC-QtoF-HRMS were docked against XO (PDB ID: 3NVY), with phytosphingosine (−8.5 kcal/mol) and rutin (−8.3 kcal/mol) exhibiting the strongest binding affinities, forming favorable predicted interactions with critical catalytic residues, followed by 6‴-feruloylspinosin, 3′,5′-di-C-β-glucopyranosylphloretin and hexadecasphinganine (ranging from −7.8 to −7.6 kcal/mol). Predictive structure–activity relationships were also observed. These results provide insights into the antioxidant potential of Z. lotus phytochemicals and highlight the value of this extremophile plant as sustainable resource for phytotherapy and the management of oxidative stress-related diseases.

1. Introduction

Medicinal plants remain an inexhaustible reservoir of structurally diverse natural products with high therapeutic potential, providing leads for drug discovery, nutraceutical development, health-promoting properties, and functional food innovation [1,2]. Among them, wild edible fruits are increasingly recognized for their complex phytochemical matrices, which often include vitamins, flavonoids, phenolic acids, tannins, and triterpenoids with antioxidant, anti-inflammatory, and antimicrobial properties [3,4]. In the current context of increasing interest in plant-derived bioactive molecules, the systematic exploration of underutilized species has become a scientific priority. A special emphasis is given to those adapted to extreme environments, where the stress-driven production of secondary metabolites may yield novel compounds of pharmacological value.

Ziziphus lotus (L.) Lam., commonly known as “Sedra” or “wild jujube,” is a thorny shrub native to the Mediterranean basin, North Africa, and parts of the Middle East (Figure 1). This species is remarkable for its extremophilic traits, thriving under arid conditions, intense heat, and nutrient-poor soils where most cultivated plants cannot survive. Such environmental resilience is often associated with the accumulation of high-value metabolites that confer protection against oxidative and abiotic stresses. In the face of accelerating climate change, the Mediterranean basin has emerged as one of the most vulnerable global hotspots, with average annual temperatures already 1.4 °C above pre-industrial levels and projected increases of up to 2–4 °C by the end of the century under high-emission scenarios [5,6]. These warming trends, combined with prolonged droughts, declining winter rainfall, and recurrent heatwaves [7,8], underscore the need to study plants like Z. lotus that can thrive under such pressures. Within this context, investigating the phytochemical composition and antioxidant properties of Z. lotus fruits provides a rational framework to link ecological adaptation with molecular signatures of stress tolerance, while also identifying resilient sources of bioactive compounds of interest under future climate constraints.

The genus Ziziphus has long been studied for its pharmacological properties, with a marked surge in research publications since 2020 according to bibliometric data from SciFinder database (Figure 2A), largely driven by interest in Z. jujuba and Z. mauritiana. In contrast, Z. lotus, despite its unique adaptations and ethnopharmacological value, has been comparatively underexplored, though its scientific visibility has irregularly increased since 2017 (Figure 2B). This growing trend highlights the need for deeper phytochemical and bioactivity studies on this species. Traditionally, various parts of Z. lotus (roots, leaves, and fruits) have been used in North African and Mediterranean folk medicine to treat gastrointestinal disorders, liver dysfunctions, urinary infections, skin diseases, insomnia, and diabetes [9,10,11]. The fruits, locally known as “N’Beg,” are consumed fresh, dried, or processed and also serve as valuable animal feed for sheep, goats, and camels [12]. Phytochemical studies have reported a high content of flavonoids, sterols, tannins, triterpenoid saponins, and polyphenols with notable antioxidant, antimicrobial, and immunomodulatory activities [13,14,15]. Several of these metabolite classes are well recognized for their capacity to modulate redox balance and to counteract oxidative stress through complementary antioxidant mechanisms, providing a biochemical rationale for further investigation of their activity in oxidative stress-related contexts.

Oxidative stress, resulting from the imbalance between reactive oxygen species (ROS) and antioxidant defenses, is a central mechanism in the onset and progression of chronic diseases. One major enzymatic source of ROS is xanthine oxidase (XO), a molybdenum-dependent enzyme catalyzing the oxidation of hypoxanthine to xanthine and subsequently to uric acid, with concomitant production of superoxide anions and hydrogen peroxide [16,17]. Excessive XO activity is implicated in cardiovascular and metabolic disorders as well as neurodegenerative conditions [18,19]. Accordingly, plant-derived XO inhibitors are of considerable interest as potential therapeutic agents and functional food components.

This study aimed to valorize Z. lotus fruits as a source of antioxidants and putative XO-inhibitory phytoconstituents through an integrated in vitro and in silico approach. The in vitro antioxidant properties of the ethyl acetate (EtOAc) extract were first assessed, followed by a chemical profiling to establish a molecular basis for the biological activities of this extract. Complementary molecular docking against XO was then performed to explore potential binding affinities and interactions with catalytically relevant residues, integrating predictive structure–activity considerations. By combining experimental and computational data, this work elucidates the molecular basis of the antioxidant potential of Z. lotus and emphasizes the broader relevance of extremophile Mediterranean species as sustainable reservoirs of bioactive molecules. This research aligns with the United Nations Sustainable Development Goals (SDG 3: Good Health and Well-being; SDG 12: Responsible Consumption and Production; and SDG 15: Life on Land), highlighting the intersection between phytochemistry, climate resilience, and global health priorities. By integrating phytochemistry, computational biochemistry, and climate science, this work positions Z. lotus not only as a promising reservoir of therapeutic agents but also as a strategic model for understanding how plant extremophiles can support human well-being in a changing world.

2. Results and Discussion

The fractionation of the hydroalcoholic macerate allowed the removal of undesirable lipid and other less-polar compounds while concentrating polar bioactive compounds in the EtOAc extract, which were recovered as a solid, dark extract after evaporation of the residual solvent in a 0.40% yield. This yield could be considered low if compared to those reported in the literature for ethanolic extracts obtained directly by maceration without fractionation. Rais et al. [20], working on three populations of Z. lotus L., reported yields ranging from 11.94% to 15.96%. Similarly, Abcha et al. [21] observed ethanolic extract yields from Z. lotus fruits ranging from 20.02% to 49.21% (w/w) relative to the plant material. These significant differences can be attributed not only to the harvesting period and the genetic characteristics of the samples analyzed, but also to the extraction method, as the previous studies used direct solvent maceration, while the present work involved liquid–liquid fractionation.

2.1. Determination of Total Phenolic Content (TPC) and Total Flavonoids Content (TFC)

The results of the TPC assay (

Table 1. TPC and TFC in the EtOAc extract from Z. lotus fruits.

TPC (µg GAE/mg)	TFC (µg RE/mg)
281.33 ± 1.5	127.25 ± 5.88

) showed that the EtOAc extract was rich in polyphenols (281.33 ± 1.5 μg GAE/mg of extract). This suggests that the defatting process first using hexane as solvent for the extraction, followed by extraction with dichloromethane, contributed to concentrating polyphenols in the EtOAc extract. This result is consistent with the findings of Khouchlaa et al. [22], who reported that an aqueous extract from Z. lotus fruits was rich in total polyphenols (285.19 mg GAE/mg of dry extract). In contrast, Cadi et al. [23] reported a markedly lower polyphenol content, with only 3.0 ± 0.10 mg GAE/g of dry matter in the EtOAc fraction. Moreover, Rais et al. [20] observed a maximum polyphenol content of 30.36 ± 0.30 mg GAE/g of extract when using methanol alone to extract bioactive compounds, while noting a significantly lower content when ethanol was used as the extraction solvent. Indeed, in addition to the extraction protocol, geographical and climatic conditions, genetic factors, the maturation stage, and storage duration have a significant influence on polyphenol content [24,25].

Regarding the TFC of the EtOAc extract, the results showed a value of 127.257 ± 5.887 μg RE/mg of extract, which is significantly higher than those reported in the literature. Indeed, Cadi et al. [23] reported a content of 2.0 ± 0.10 mg/g of extract, while Bouzid et al. [26] obtained values of 8.25 ± 0.20 and 12.66 ± 0.29 mg QE/g of dry matter. Similarly, Ghazghazi et al. [27] found a concentration of 1.22 ± 0.006 mg/g in Z. lotus fruit extracts. These findings further emphasize the critical influence of both the extraction methodology and the botanical origin on the efficiency of secondary metabolite recovery. Taken together, the notably elevated levels of polyphenols and flavonoids in the EtOAc extract highlight its promising value as a rich reservoir of bioactive compounds.

2.2. Identification of the Fruit Phytoconstituents and Occurrence in Z. lotus

The full-scan UHPLC-QToF-HRMS analysis of the EtOAc extract allowed the tentative identification of eight metabolites (1–8, Figure 3) that represent a diverse variety of structural classes, including highly polar carbohydrates and glycosylated flavonoids and less polar alkaloids and sphingoid bases. This chemically heterogeneous profile is consistent with the extraction workflow, which started from a hydroalcoholic macerate rather than a purely aqueous matrix or direct maceration of the plant material with EtOAc. Residual ethanol may act as a co-solvent during liquid–liquid partitioning, shifting distribution coefficients and enabling partial transfer of relatively polar constituents into the EtOAc extract, as reported in comparable phytochemical fractionations [28]. Moreover, because extraction was performed under neutral conditions to preserve labile metabolites, weakly basic alkaloids and amphiphilic sphingoid bases may have remained less ionized and thus showed appreciable affinity for EtOAc [29], supporting the plausibility of the metabolite classes annotated below.

Identification was first performed using positive-ion mode, followed by a negative-ion mode analysis to corroborate the assignments for metabolites showing sensitivity in this polarity (1–5). Additionally, UV/PDA chromatograms and UV–Vis spectra were examined to provide support for the proposed identifications. Total ion chromatograms (TICs), UV/PDA chromatograms at 355, 330 and 280 nm wavelengths, and extracted ion chromatograms (EICs) are provided in the Supporting Information (Figures S1–S4). The identified compounds are listed in

Table 2. List of the metabolites tentatively identified in the Z. lotus fruit extract following positive and negative ionizations by UHPLC-QToF-HRMS; “t_R”: retention time (min); “Δ ppm”: mass error; “-”: not detected.

Proposed Annotation	Molecular Formula	tR	Data from Positive Ionization			Data from Negative Ionization
			Measured m/z	Precursor Ion (m/z)	Δ ppm	Measured m/z	Precursor Ion (m/z)	Δ ppm
Sucrose (1)	C12H22O11	0.640	365.1057	C12H22O11Na [M + Na]+	−1.4	341.1084	C12H21O11 [M − H]−	−0.6
Rutin (2)	C27H30O16	1.760	611.1622	C27H31O16 [M + H]+	1.6	609.1451	C27H29O16 [M − H]−	−0.5
3′,5′-Di-C-β-glucopyranosylphloretin (3)	C27H34O15	1.957	599.1976	C27H35O15 [M + H]+	−0.3	597.1817	C27H33O15 [M − H]−	−0.3
Kaempferol O-rutinoside (4)	C27H30O15	2.560	595.1673	C27H31O15 [M + H]+	1.7	593.1501	C27H29O15 [M − H]−	−0.8
6′′′-Feruloylspinosin (5)	C38H40O18	2.948	785.2293	C38H41O18 [M + H]+	−2.8	783.2136	C38H39O18 [M − H]−	0.0
Amphibine B (6)	C39H47N5O5	3.879	666.3655	C39H48N5O5 [M + H]+	−1.2	-	-	-
Hexadecasphinganine (7)	C16H35NO2	5.174	274.2758	C16H36NO2 [M + H]+	4.4	-	-	-
Phytosphingosine (8)	C18H39NO3	5.242	318.3012	C18H40NO3 [M + H]+	1.3	-	-	-

in order of retention time.

Compound 1 was identified as sucrose by the dominant sodium adduct [M + Na]⁺ at m/z 365.1057 (with a supporting [M + K]⁺ at m/z 381.0793) and a characteristic in-source fragment at m/z 203.0531 ([Hexose + Na]⁺). Following negative ionization, the deprotonated ion [M − H]⁻ at m/z 341.1084 for the same peak, together with the formate adduct [M + HCOO]⁻ at m/z 387.1138, was also observed. Its identification is consistent with previous reports identifying sucrose as the most abundant sugar in Z. lotus fruits [30].

Compound 2 was identified as rutin (quercetin-3-O-rutinoside) based on its protonated molecular ion [M + H]⁺ at m/z 611.1622 and two diagnostic in-source fragment ions: m/z 465.1029, reflecting loss of 146.0579 Da (rhamnosyl moiety), and m/z 303.0505, corresponding to the quercetin aglycone. Following negative ionization, the deprotonated ion [M − H]⁻ at m/z 609.1451 was also observed for the same peak, whereas the experimental UV–Vis spectrum was consistent with that reported for rutin [31] (Figure 4A). This metabolite has been previously identified in Z. lotus [32].

Compound 3 was identified as 3′,5′-di-C-β-glucopyranosylphloretin based on its protonated ion at m/z 599.1976, with the deprotonated ion [M − H]⁻ at m/z 597.1817 matching for the same peak following negative ionization. This compound is a C-glycosylated chalcone and, to date, the only metabolite reported for Z. lotus [33] with a molecular weight of 598 Da. The experimental UV–Vis spectrum (Figure 4B) agreed with data from the literature for this compound [34].

Compound 4 was identified as kaempferol 3-O-rutinoside based on its [M + H]⁺ ion at m/z 595.1673 and the [M + Na]⁺ adduct at m/z 617.1498, consistent with a flavonoid O-rutinoside. The presence of a fragment ion at m/z 449.1085, corresponding to the loss of a rhamnosyl moiety (C₆H₁₀O₄), and a diagnostic aglycone fragment at m/z 287.0559 (kaempferol) strongly support this annotation. Distinction from the isomeric kaempferol 3-O-robinobioside (differing only in the hexose component; glucose vs. galactose) was achieved by examining sugar-derived oxonium ions: a strong peak at m/z 128.9526 (rhamnose oxonium) and a signal at m/z 147.0453 (dehydrated corresponding hexose), and the absence of the m/z 163 oxonium ion (galactose marker) [35], strongly suggested a rhamnose + glucose disaccharide rather than galactose + glucose. Compound 4 has been previously reported as a metabolite produced by Z. lotus [36].

Compound 5 was identified as 6‴-O-feruloylspinosin based on its protonated ion at m/z 785.2293, and the deprotonated ion [M − H]⁻ at m/z 783.2136 observed for the same peak following negative ionization. This feruloylated C-glycosylflavone has been reported within the Ziziphus genus [37] and the present study provides the first tentative identification of this compound in Z. lotus.

Compound 6 was identified as amphibine B based on its protonated ion at m/z 666.3655, matching the only known Ziziphus-derived metabolite with the formula C₃₉H₄₇N₅O₅. This cyclopeptide alkaloid has been previously detected in leaves and seeds from Z. lotus [33].

Compounds 7 and 8 were identified as long-chain sphingoid bases (LCBs) based on their protonated ions and characteristic in-source fragments. Compound 7 gave [M + H]⁺ at m/z 274.2747, consistent with the molecular formula of hexadecasphinganine (also named as C16 sphinganine), and displayed sequential water-loss peaks at m/z 256.2650 and 238.2542. Compound 8 produced [M + H]⁺ at m/z 318.3012, matching phytosphingosine, with similar water losses at m/z 300.2904 and 282.2796.

Although compounds 7 and 8 (putatively hexadecasphinganine and phytosphingosine) have not been previously reported in the genus Ziziphus, their occurrence in plant fruits is biochemically plausible. Inês et al. [38] documented roles of sphingolipids in fruit development and ripening, finding that the concentration of diverse LCBs increases during fruit development. Hexadecasphinganine (7) has been scarcely reported as a plant phytoconstituent, with a recent study describing its putative identification in a hydroalcoholic extract (from Alstonia angustiloba leaves), whereas phytosphingosine (8) has been more widely documented as a plant metabolite, being one of the most abundant LCBs in plants, whose biosynthesis is thought to occur through hydroxylation of sphinganine by hydroxylases [38,39]. Therefore, the putative identification of compounds 7 and 8 in Z. lotus fruits provides an extension of the natural occurrence of these LCBs, consistent with the sphingolipid metabolism.

2.3. Antioxidant Ability

The antioxidant effect of the EtOAc extract was evaluated using four different methods. The results, summarized in

Table 3. In vitro antioxidant activity via DPPH^•, ABTS^•+, phenantroline and FRAP methods.

	IC50 (µg/mL)		A0.5 (µg/mL)
	DPPH•	ABTS•+	Phenantroline	FRAP
EtOAc	80.53± 1.27 a	18.49 ± 1.47 a	8.38 ± 0.69 a	27.11 ± 1.37 a
Trolox	5.14 ± 0.12 b	3.27 ± 0.24 b	5.21 ± 0.09 b	5.43 ± 0.44 b
Ascorbic acid	4.40 ± 0.15 b	3.07 ± 0.06 b	3.08 ± 0.07 c	3.76 ± 0.33 b

Values with different letters are significantly different at p < 0.05 according to Tukey’s HSD test.

, show the inhibitory concentration expressed by the corresponding IC₅₀ values (μg/mL) for DPPH^• and ABTS^•+, as well as the A_0.5 values (μg/mL) for the phenanthroline and FRAP assays. The differences in activity between the extract and the reference standards can be attributed to the fact that, in the extract, the active compounds occurred within a complex mixture and were present at lower individual concentrations than ascorbic acid and Trolox when used as pure standards.

The results of the DPPH^• free radical scavenging assay revealed a moderate but significant antioxidant activity for the EtOAc extract, with an IC₅₀ value of 80.53 ± 1.27 μg/mL. This experimental result is consistent with previous in vitro studies that have demonstrated the antioxidant potential of Z. lotus, particularly in scavenging free radicals and protecting against oxidative stress [27,40]. Regarding the ABTS^•+ radical cation scavenging activity, the EtOAc extract exhibited a considerably stronger effect, with an IC₅₀ of 18.49 ± 1.47 μg/mL, which reflected a promising antioxidant effect. Similarly, Bouzid et al. [26] reported ABTS^•+ scavenging activity for the same species from fruit kernels (the non-edible part), further supporting its relevance as a source of natural antioxidants. In contrast, the metal chelation assay using the Fe²⁺-phenanthroline complex revealed a remarkably stronger chelating activity for the EtOAc extract, with an A_0.5 of 8.38 ± 0.69 μg/mL, closer to the values obtained for Trolox (A_0.5 = 5.21 ± 0.09 μg/mL) and ascorbic acid (A_0.5 = 3.08 ± 0.07 μg/mL). Thus, the extract demonstrated a significant capacity to interfere with iron-mediated oxidative processes, suggesting a potential protective effect against metal-induced oxidative damage. In the FRAP assay, the extract showed a moderate but still considerable reducing power, with an A_0.5 value of 27.11 ± 1.37 μg/mL. This result, while less potent than those of ascorbic acid (A_0.5 = 3.76 ± 0.33 μg/mL) and Trolox (A_0.5 = 5.43 ± 0.44 μg/mL), highlights the extract’s ability to act as an electron donor and reduce ferric ions (Fe³⁺) to ferrous form (Fe²⁺). This finding is consistent with previous reports by Bouzid et al. [26], who also observed substantial FRAP activity in Z. lotus fruit kernels, suggesting that environmental factors may influence the antioxidant potential of the plant. The antioxidant potential observed in the various in vitro assays may be attributed to the role of specific bioactive compounds in the extract. Full-scan UHPLC-QToF-HRMS analyses revealed the presence of eight distinct potential phytoconstituents (Table 2), including sucrose (1, identified in previous studies as a major constituent [28]), and rutin (2), shown in previous studies to exhibit a wide range of pharmacological properties, including antioxidant activity via binding to Fe²⁺ ions and thereby preventing interaction with hydrogen peroxide [41]. 3′,5′-Di-C-β-glucopyranosylphloretin (3) was also identified at a relative abundance of 7.47%. This C-glycosylated flavonoid, previously reported in several Ziziphus species, is known for its remarkable antioxidant potential. According to Pawłowska et al. [42], it ranks among the principal phenolic compounds isolated from this genus. Its occurrence in the analyzed extract likely plays a key role in the antioxidant effects observed, such as free radical neutralization and metal ion chelation. Other identified compounds, such as 6‴-feruloylspinosin (5) and hexadecasphinganine (7), have also been linked to antioxidant effects. According to Yang et al. [43], 6‴-feruloylspinosin (5) exhibits antioxidant properties in the seeds of Z. jujuba Mill. var. spinosa, while hexadecasphinganine (7) has been reported by Mufti et al. [44] to reduce oxidative stress and enhance antioxidant responses. Their combined presence may contribute synergistically to the overall antioxidant activity of the extract. In contrast, there is currently no evidence in the literature to the best of our knowledge supporting antioxidant activity for kaempferol-O-rutinoside, amphibine B, or phytosphingosine (4, 6 and 8), which limits any conclusions about their potential involvement. Altogether, these findings are consistent with earlier reports highlighting the richness of Z. lotus in antioxidant compounds [30]. This chemical diversity reinforces the therapeutic potential of the species as a natural source of antioxidants.

2.4. Xanthine Oxidase Inhibitory Activity and Molecular Docking Analysis

The EtOAc extract was assayed in vitro for XO inhibition in the range 300–18.8 μg/mL, yielding a dose–response curve with an IC₅₀ value of 170.4 ± 5.90 μg/mL (R² = 0.9977) and 66.5 ± 1.5% inhibition at the highest tested concentration (300 μg/mL). Based on this moderate but relevant activity, which is in a range coherent for plant extracts where bioactive phytoconstituents are diluted by other inactive matrix constituents, molecular docking was performed to assess the potential interaction of the eight phytoconstituents tentatively identified in the EtOAc extract (1–8, Table 2) against XO. The crystal structure of XO in complex with quercetin (PDB ID: 3NVY) was used, and quercetin was adopted as the reference ligand [45]. The structures of compounds 1–8 were docked against the XO binding site. It should be emphasized that molecular docking provides qualitative insight into potential binding modes and interaction hotspots within the XO active site but does not by itself establish functional inhibition or quantitative potency. Accordingly, the XO-inhibitory effect discussed below should be regarded as a predicted mechanism that requires further experimental validation.

Six of these molecules exhibited binding energies equal to or greater than that of quercetin (–7.0 kcal/mol). The remaining two compounds, kaempferol O-rutinoside (4) (–6.9 kcal/mol) and amphibine B (6) (–6.2 kcal/mol), displayed slightly weaker affinities compared to quercetin. These results are summarized in

Table 4. Binding affinities and molecular interaction profiles of the phytocompounds identified in Z. lotus fruits against docked xanthine oxidase (XO; PDB ID: 3NVY), in decreasing order of binding affinity (kcal/mol).

	Binding Energy (kcal/mol)	Hydrogen Interactions (Distance Å)	Hydrophobic Interactions	Electrostatic Interactions
Co-crystallized ligand (Quercetin)	−7.0	Glu802 (1.60), Phe1009 (2.72), Arg880 (1.91), Thr1010 (2.47)	Leu873, Leu648, Val1011	-
Phytosphingosine (8)	−8.5	Glu802 (2.20), Glu802 (2.66), Gln767 (2.52), Glu1261 (1.97)	Ala1079 (2), Ala1078, Phe914 (3), Phe1009, Val1011 (2), Leu873, Leu648, Phe649	Glu1261
Rutin (2)	−8.3	Arg880 (3.08), Phe914 (2.98), Ala1079 (2.58), Glu802 (2.69), Asn768 (2.33), Ser1075 (2.43), Met770 (2.20), Ser876 (2.28), Ser876 (2.45), Ser876 (2.29)	Phe914 (2), Val1011, Pro1076, Leu1014 (2), Leu648	Met770, Lys771
6‴-Feruloylspinosin (5)	−7.8	Arg880 (2.91), Glu1261 (2.63), Glu1261 (2.84), Leu648 (2.58), Lys771 (2.18), Lys771 (2.81)	Phe1009, Ala1079, Ala1078 (2), Phe914 (2)	-
3′,5′-Di-C-β-glucopyranosylphloretin (3)	−7.7	Glu802 (2.38), Ala1079 (2.98), Leu873 (2.28), Leu873 (2.70), His875 (2.97), Asn768 (1.96), Lys771 (2.25), Ser876 (2.28), Thr1010 (2.20), Thr1010 (1.90)	Leu648, Val1011, Leu873	-
Hexadecasphinganine (7)	−7.6	Arg880 (2.54), Ala1079 (2.98)	Phe649 (2), Leu873, Leu648 (2), Leu1012, Val1011 (2) Phe914	Phe914, Glu1261
Sucrose (1)	−7.0	Leu873, Ser876, Ser867, Thr1010, Phe1009, Phe914	-	-
Kaempferol O-rutinoside (4)	−6.9	His875, Asp872, Lys771, Val1011	Leu648, Phe649, Met770, Leu1014	-
Amphibine B (6)	−6.2	Phe1013	Leu873, Leu648, Val1011, Leu1014, Phe649	Met770, Lys771

and visualized in Figure 5, highlighting binding affinities and specific molecular interactions. Given the inherent limitations and uncertainty of docking scoring functions, small differences in binding energies should not be overinterpreted, and rank-order comparisons among compounds should be considered as tentative for prioritizing phytoconstituents 1–8 and structurally related compounds in further research.

Among all docked compounds, phytosphingosine (8) displayed favorable predicted binding mode (−8.5 kcal/mol). This compound formed multiple strong hydrogen bonds with key catalytic and stabilizing residues such as Glu802, Gln767, and Glu1261, with interaction distances ranging from 1.97 to 2.66 Å. Additionally, hydrophobic contacts with Phe914, Val1011, Leu873, and Ala1079 further contributed to the stabilization of the ligand within the binding pocket. An electrostatic interaction with Glu1261 was also noted, highlighting phytosphingosine as a ligand with a favorable predicted binding mode and interaction with key XO residues. However, sphingoid bases are not currently recognized as a class of established XO inhibitors in experimental studies, and therefore these docking results should be regarded as exploratory and hypothesis-generating rather than indicative of confirmed enzymatic inhibition. Rutin (2), with a binding energy of −8.3 kcal/mol, formed an extensive network of ten hydrogen bonds, notably with Arg880, Glu802, Ser876, Met770, and Phe914, highlighting its strong anchoring ability within the active site. Its interactions were not only numerous but also strategically distributed across the binding cavity, involving both polar and hydrophobic regions. Moreover, electrostatic interactions with Met770 and Lys771, and hydrophobic contacts with Val1011 and Leu648, emphasize rutin’s predicted binding interactions within the XO pocket, in partial agreement with reports suggesting that certain flavonoid scaffolds can interact with XO, although glycosylated derivatives generally exhibit lower inhibitory potency than their aglycones [46,47]. 6‴-Feruloylspinosin (5) and 3′,5′-Di-C-β-glucopyranosylphloretin (3) also exhibited high docking scores (−7.8 and −7.7 kcal/mol, respectively). The former showed notable electrostatic and hydrogen interactions with Arg880, Glu1261, and Lys771, along with π-stacking interactions involving Phe1009 and Phe914. The latter, a glycosylated flavonoid derivative, engaged in hydrogen bonding with critical residues such as Glu802, Asn768, and Thr1010, while simultaneously forming hydrophobic contacts with Val1011 and Leu648. It should be noted that flavonoid glycosides are generally recognized as weaker direct inhibitors of XO than their corresponding aglycones, largely due to steric hindrance and excessive polarity [46,48]. The favorable docking scores observed for rutin and related glycosylated flavonoids should not be therefore interpreted as evidence of strong enzymatic inhibition, but rather as an indication of their ability to interact with the large and solvent-accessible XO binding cavity through extensive hydrogen-bonding networks. Importantly, several experimental studies have consistently demonstrated that glycosylation of flavonoids reduces their direct XO inhibitory potency compared to the corresponding aglycones, mainly due to steric hindrance and reduced ability to access the molybdenum-containing catalytic center [49,50]. Therefore, although rutin and other glycosylated flavonoids exhibited favorable docking scores in the present study, these findings should be interpreted as indicative of potential binding within the extended XO cavity rather than as evidence of strong enzymatic inhibition, and do not override the established biochemical evidence regarding the generally weaker inhibitory activity of flavonoid glycosides.

Hexadecasphinganine (7) scored −7.6 kcal/mol and showed fewer hydrogen bonds but significant hydrophobic interactions with key nonpolar residues, particularly Leu873, Phe914, and Leu648, reinforcing the idea that hydrophobic stabilization is a major contributor to its binding mode. Electrostatic contact with Phe914 and Glu1261 was also observed. Sucrose (1) scored −7.0 kcal/mol and completed the lower-affinity end of the docked set, included as a highly polar reference constituent rather than as a bioactive XO inhibitor. Although its affinity was at the lower end of the selected range, it still demonstrated specific interactions with different catalytic residues like Phe1009, which may indicate weak but specific interactions with XO that could contribute indirectly to the overall antioxidant profile of the extract. Although sucrose formed several hydrogen bonds with residues such as Ser876, Thr1010, and Phe1009, the absence of hydrophobic and aromatic interactions limited its stabilization within the XO pocket, resulting in a binding score at the cutoff threshold. In general, the docked compounds share common interaction hotspots with XO, especially Glu802, Arg880, Phe1009, Val1011, and Leu873. These residues appear consistently involved in hydrogen bonding and hydrophobic stabilization across the most potent ligands and are therefore suggested to play a central role in ligand recognition and potential modulation of XO activity.

2.5. Predictive Druglikeness and Structure–Activity Relationships

Structure–activity relationships (SARs) and druglikeness of the identified phytoconstituents were explored using the physicochemical parameters calculated for compounds 1–8 shown in

Table 5. Physicochemical descriptors of the phytocompounds tentatively identified in Z. lotus fruits.

	Clog p	TPSA (Å2)	Fraction of Csp3	H-Bond Acceptors	H-Bond Donors	Rotatable Bonds
Sucrose (1)	−3.09	189.53	1.00	11	8	5
Rutin (2)	−1.36	269.43	0.44	16	10	6
3′,5′-Di-C-β-glucopyranosylphloretin (3)	−1.93	278.29	0.52	15	12	8
Kaempferol O-rutinoside (4)	−0.76	249.20	0.44	15	9	6
6‴-Feruloylspinosin (5)	1.51	284.73	0.37	18	9	12
Amphibine B (6)	5.54	120.08	0.38	6	3	12
Hexadecasphinganine (7)	5.17	66.48	1.00	3	3	14
Phytosphingosine (8)	5.51	86.71	1.00	4	4	16

as complementary in silico insights. Docking results and physicochemical trends converge on two recurring binding solutions. One represented by highly polyhydroxylated glycosides that anchor through dense hydrogen-bond networks, while the other involves long-chain sphingoid bases that exploit hydrophobic stabilization while maintaining a small number of directional hydrogen bonds to catalytic residues. These complementary strategies explain why both strongly polar scaffolds and amphiphilic chains can attain favorable docking scores within the same active-site environment. The interplay between hydrogen-bond capacity and polarity on the one hand, and hydrophobic surface area on the other, is evident when contrasting the top-scoring ligands. Phytosphingosine (8; −8.5 kcal/mol) couples moderate polarity (HBA/HBD 4/4; TPSA 86.7 Ų) with a fully saturated, conformationally flexible chain (fraction of Csp³ = 1.00) that fits the lipophilic channel and donates short hydrogen bonds to Glu802, Gln767, and Glu1261. Rutin (2; −8.3 kcal/mol), 6‴-feruloylspinosin (5; −7.8 kcal/mol), and 3′,5′-di-C-β-glucopyranosylphloretin (3; −7.7 kcal/mol) rely primarily on extensive polar contact networks, as reflected by their high numbers of hydrogen-bond donors and acceptors and large TPSA values, which likely contribute to stabilizing interactions within the XO binding pocket. At the margin, sucrose (1; −7.0 kcal/mol) illustrates the limitation of polarity without lipophilic complementarity, and despite numerous potential hydrogen bonds, weaker hydrophobic packing keeps its affinity at the cutoff. Hydrophobic contribution correlates with cLog p and with the presence of extended aliphatic surfaces. Hexadecasphinganine (7) and phytosphingosine (8) share high cLog p values (5.17 and 5.51, respectively) and maximize van der Waals contacts with Leu873, Phe914, and Val1011 while using a few polar interactions to orient the ligand. By contrast, flavonoid glycosides cluster at low or negative cLog p values due to multiple sugars. Their binding could be therefore dominated by polar recognition rather than lipophilic complementarity.

The highest-scoring ligands also have the most rotatable bonds (up to 16 in compound 8), which likely imposes an entropic cost. Within closely related structures, some structure–activity relationships (SARs) could be suggested. Rutin (2) and kaempferol-O-rutinoside (4) are isostructural except for an additional hydroxyl group on the aglycone of compound 2. This additional donor/acceptor increases TPSA (269 vs. 249 Ų) and could expand the hydrogen-bonding network to residues such as Arg880, Glu802, Ser876, Met770, and Phe914, coinciding with a stronger score for 2 and the exclusion of 4 from the active identified compounds in the extract. A similar pattern appears for the sphingoid bases. Phytosphingosine (8) differs from hexadecasphinganine (7) by one additional hydroxyl and a slightly different chain, modestly increasing TPSA (86.7 vs. 66.5 Ų) while preserving high lipophilicity and hydrophobic fit. The extra polar functional group in compound 8 could engage acidic residues (notably Glu802 or Glu1261), delivering a notable gain in docking energy (−8.5 vs. −7.6 kcal/mol) without compromising packing.

Across the top-scoring metabolites, two physicochemical features recur and appear favorable. First, the presence of at least one strong donor–acceptor pair positioned to contact Glu802, Glu1261, or Gln767 is consistently beneficial, even for predominantly hydrophobic scaffolds. Second, sufficient hydrophobic surface to engage Leu873, Phe914, and Val1011 improves stabilization in the pocket. In practical terms, a compact polar head paired with a hydrophobic tail, exemplified by 8, seems more efficient than distributing polarity across the entire framework, as occurs in heavily glycosylated flavonoids. Drug-likeness filters provide additional context for translatability. The highly polar glycosides, i.e., rutin (2), di-C-glucopyranosylphloretin (3), kaempferol-O-rutinoside (4), 6‴-feruloylspinosin (5) and sucrose (1), exceed Veber and Egan TPSA thresholds (≤140 and ≤131.6 respectively) by a wide margin, indicating that their polarity is likely too high for efficient passive transcellular diffusion across biological membranes. Amphibine B (6) sits at borderline to high lipophilicity (cLog p 5.54) with high flexibility (12 rotors), likely failing Veber (≤10 rotatable bonds) and offering no compensatory docking advantage. In contrast, the sphingoid bases 7 and 8 occupy a drug-like polarity window (TPSA 66–87 Ų; HBA/HBD ≤ 4) and satisfy Lipinski on polarity while straddling or slightly exceeding logP cutoffs. However, their very high flexibility violates Veber, suggesting low oral exposure unless rotors are reduced by conformational restriction. Overall, phytosphingosine (8) and hexadecasphinganine (7) emerge as mechanistically informative scaffolds that merit biochemical validation to assess whether their predicted binding translates into measurable XO inhibition or other functional effects.

3. Materials and Methods

3.1. Plant Material

This study was carried out on Z. lotus fruits collected in September 2023 from the El Khroub area in eastern Algeria. After collection, the fruits were cleaned and air-dried in the shade at ambient temperature for 15 days. The dried material (Figure 6) was then ground into a fine powder and stored in glass containers until use [51].

3.2. Preparation of the Bioactive Extract

The bioactive fruit extract was obtained by EtOAc liquid–liquid extraction of a hydroalcoholic macerate, following a modified method [52]. For this purpose, 20 g of plant powder was cold-macerated in 200 mL of an ethanol/water mixture (80:20, v/v) for 24 h under moderate stirring. Following filtration, liquid–liquid partitioning was carried out using three solvents of increasing polarity: hexane, dichloromethane, and EtOAc. The EtOAc extract was then dried over anhydrous Na₂SO₄, filtered, concentrated under reduced pressure and stored until further analysis. The extraction yield (Y) was calculated according to Equation (1).

$$\mathrm{Y}(\%)={\displaystyle \frac{\mathrm{w}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t}\mathrm{o}\mathrm{f}\mathrm{s}\mathrm{o}\mathrm{l}\mathrm{i}\mathrm{d}\mathrm{e}\mathrm{x}\mathrm{t}\mathrm{r}\mathrm{a}\mathrm{c}\mathrm{t}\mathrm{c}\mathrm{o}\mathrm{l}\mathrm{l}\mathrm{e}\mathrm{c}\mathrm{t}\mathrm{e}\mathrm{d}}{\mathrm{w}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t}\mathrm{o}\mathrm{f}\mathrm{s}\mathrm{t}\mathrm{a}\mathrm{r}\mathrm{t}\mathrm{i}\mathrm{n}\mathrm{g}\mathrm{d}\mathrm{r}\mathrm{y}\mathrm{p}\mathrm{l}\mathrm{a}\mathrm{n}\mathrm{t}\mathrm{m}\mathrm{a}\mathrm{t}\mathrm{e}\mathrm{r}\mathrm{i}\mathrm{a}\mathrm{l}}}\times 100$$

(1)

3.3. Determination of TPC and TFC

3.3.1. Determination of TPC

The TPC was estimated using the Folin–Ciocalteu reagent (FCR), following an adapted method [53]. A volume of 200 μL of extract was mixed with 1.5 mL of distilled water, followed by the addition of 100 μL of FCR and 500 μL of sodium carbonate solution (7%). After incubation for 2 h at room temperature in the dark, absorbance was measured at 760 nm. Results were expressed as micrograms of gallic acid equivalents (μg GAE) per milligram of extract, based on a calibration curve established with gallic acid concentrations ranging from 20 to 200 μg/mL.

3.3.2. Determination of TFC

The TFC was determined using the aluminum chloride method, following that reported by Aryal et al. [54] with slight modifications. A volume of 1 mL of extract (1 mg/mL) was mixed with 200 μL of AlCl₃ (10%), 200 μL of sodium acetate, and 5.6 mL of distilled water. After 20 min of incubation in the dark, absorbance was measured at 415 nm. Flavonoid concentrations were calculated using a calibration curve established with rutin (25–200 μg/mL). Results were expressed as micrograms of rutin equivalents per milligram of extract (μg RE/mg extract).

3.4. LC-MS Analysis

The Z. lotus fruit extract was analyzed using ultra-high-performance liquid chromatography coupled with high-resolution quadrupole time-of-flight mass spectrometry (UHPLC-QToF-HRMS) on a Xevo G2-S QTof system (Waters Corporation, Milford, MA, USA), following a full scan. The extract was previously filtered through a 0.22 μm hydrophobic membrane filter (in methanol) and injected (5 μL) at a concentration of 100 ppm, using methanol as the injection solvent. Chromatographic separation was achieved on an Acquity UPLC HSS T3 column (1.8 μm, 2.1 mm × 100 mm; Waters Corporation, Milford, MA, USA) maintained at 45 °C. The mobile phase consisted of (A) 0.1% v/v formic acid in water and (B) 0.1% v/v formic acid in acetonitrile, with a flow rate of 0.4 mL/min. The gradient was programmed as follows: 0–1.5 min, 20% B; 1.5–3 min, 20–40% B; 3–6 min, 40–60% B; 6–7 min, 60–95% B; 7–7.5 min, 95% B; 7.5–8 min, 95–20% B; 8–10 min, 20% B. High-resolution mass spectra were acquired in positive and negative electrospray ionization modes over an m/z range of 100–1200 Da, with a resolving power of 22,000 FWHM. The instrument was operated with a capillary voltage of 0.70 kV, a LockSpray capillary voltage of 3.00 kV, a cone voltage of 30 V, a collision energy of 6.0 eV, a source temperature of 120 °C, a desolvation temperature of 450 °C, a cone gas flow of 10 L/h, a desolvation gas flow of 850 L/h, and a scan time of 0.5 s. Tentative compound identification was performed based on high-resolution accurate mass measurements of precursor ions. Proposed molecular formulas were assigned by comparison with reported metabolites from the Ziziphus genus, using PubChem and SciFinder databases as main references. Additional peaks detected in the MS1 spectra were carefully examined to reinforce the tentative assignments. UV–Vis detection was performed using a photodiode array (PDA) detector, acquiring data from 210 to 500 nm. Chromatograms were subsequently extracted at selected wavelengths to provide complementary UV spectra for compound characterization.

3.5. Antioxidant Ability

A sequence of four different bioassays were performed for assessing the in vitro antioxidant potential of the Z. lotus fruit extract. Trolox and ascorbic acid were used as reference standards in all four assays as positive controls. Absorbance was measured using an EnSpire microplate reader (Perkin Elmer, Waltham, MA, USA).

3.5.1. DPPH^• Scavenging Assay

The DPPH assay is based on the reduction of the stable violet DPPH^• radical in the presence of antioxidants, leading to a discoloration measured at 517 nm. For this assay, a volume of 160 μL of methanolic DPPH solution was mixed with 40 μL of sample at different concentrations (6.25–800 µg/mL) in a microplate. After 30 min of incubation in the dark at room temperature, absorbance was measured at 517 nm [55]. The percentage of inhibition (%) was calculated using Equation (2).

$$\mathrm{D}\mathrm{P}\mathrm{P}\mathrm{H}\mathrm{s}\mathrm{c}\mathrm{a}\mathrm{v}\mathrm{e}\mathrm{n}\mathrm{g}\mathrm{i}\mathrm{n}\mathrm{g}\mathrm{e}\mathrm{f}\mathrm{f}\mathrm{e}\mathrm{c}\mathrm{t}\left(\%\right)={\displaystyle \frac{{\mathrm{A}\mathrm{b}\mathrm{s}\mathrm{o}\mathrm{r}\mathrm{b}\mathrm{a}\mathrm{n}\mathrm{c}\mathrm{e}}_{\mathrm{C}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{o}\mathrm{l}}-{\mathrm{A}\mathrm{b}\mathrm{s}\mathrm{o}\mathrm{r}\mathrm{b}\mathrm{a}\mathrm{n}\mathrm{c}\mathrm{e}}_{\mathrm{S}\mathrm{a}\mathrm{m}\mathrm{p}\mathrm{l}\mathrm{e}}}{{\mathrm{A}\mathrm{b}\mathrm{s}\mathrm{o}\mathrm{r}\mathrm{b}\mathrm{a}\mathrm{n}\mathrm{c}\mathrm{e}}_{\mathrm{C}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{o}\mathrm{l}}}}\times 100$$

(2)

The IC₅₀ values (μg/mL), corresponding to the concentration required to inhibit 50% of the DPPH^• radical, were determined from the dose–response curves.

3.5.2. ABTS^•+ Radical Cation Decolorization Assay

The ABTS^•+ assay is based on the reduction of the ABTS^•+ radical cation by antioxidants, resulting in a discoloration measured at 734 nm. The radical was generated by mixing ABTS (7 mM) with potassium persulfate (2.45 mM). The solution was adjusted to an absorbance of 0.70 at 734 nm. Then, 160 μL of this solution was mixed with 40 μL of sample. After 10 min of incubation, absorbance was measured and the percentage of inhibition calculated using the same formula as for the DPPH assay (Equation (2)) [56].

3.5.3. Ferric Reducing Antioxidant Power (FRAP)

This method evaluates the ability of antioxidants to reduce the ferric (Fe³⁺/ferricyanide) complex to ferrous (Fe²⁺), producing a bluish-green color measured at 700 nm. The assay involved mixing 10 μL of sample with phosphate buffer and potassium ferricyanide, with subsequent incubation at 50 °C for 20 min. After addition of trichloroacetic acid, water, and FeCl₃, absorbance was measured. The color intensity reflects the reducing power [57].

3.5.4. Phenanthroline Assay

This method evaluates the ability of antioxidants to reduce Fe³⁺ to Fe²⁺, forming a colored complex with 1,10-phenanthroline, measured at 510 nm. The reaction mixture contained 30 μL of 1,10-phenanthroline (0.5%), 50 μL of FeCl₃ (0.2%), 110 μL of methanol, and 10 μL of sample. After 20 min of incubation at 30 °C, absorbance was measured at 510 nm. The color intensity reflects the reducing activity [58].

3.5.5. Statistical Analysis

Experiments were repeated three times per treatment. Data were analyzed using SPSS (version 25.0). Statistical significance between groups was assessed using one-way analysis of variance (ANOVA), followed by Tukey’s HSD test for post hoc comparisons. Results with p-values below 0.05 were considered statistically significant.

3.6. Xanthine Oxidase Inhibition

XO inhibition was determined spectrophotometrically by monitoring the formation of uric acid from xanthine at 295 nm (25 °C) in 1 mL quartz cuvettes, following a published method with slight modifications [59]. Measurements were performed on a Cary 50 Bio UV–Vis spectrophotometer (Varian Australia Pty. Ltd., Mulgrave, Australia). Reaction mixtures contained 400 μL of phosphate buffer (120 mM at pH of 7.5), 330 μL of xanthine (150 μM), 250 μL of the extract (in distilled water to achieve the test concentrations of 300, 150, 75, 37.5, and 18.8 μg/mL), and 20 μL of XO solution (0.5 U/mL in buffer). The increase in absorbance at 295 nm was recorded during the initial linear phase (first 3 min) and initial rates (ΔAbs/min) were calculated. A control without extract and a positive control (allopurinol [59]) were assayed under the same conditions. Each analysis was performed in triplicate, with inhibition values calculated using Equation (3). Graphpad Prism v.5.0 (GraphPad Software Inc., San Diego, CA, USA) was used for IC₅₀ calculation by fitting the inhibition data to a sigmoidal dose–response model.

$$\mathrm{X}\mathrm{O}\mathrm{i}\mathrm{n}\mathrm{h}\mathrm{i}\mathrm{b}\mathrm{i}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n}\left(\%\right)=(1-{\displaystyle \frac{{\mathsf{\Delta }\mathrm{A}\mathrm{b}\mathrm{s}}_{\mathrm{S}\mathrm{a}\mathrm{m}\mathrm{p}\mathrm{l}\mathrm{e}}}{{\mathsf{\Delta }\mathrm{A}\mathrm{b}\mathrm{s}}_{\mathrm{C}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{o}\mathrm{l}}}})\times 100$$

(3)

3.7. Molecular Docking Analysis

3.7.1. Ligand Preparation

A total of eight phytoconstituents previously identified in the EtOAc extract of Z. lotus were selected for molecular docking analysis. The 2D chemical structures of the compounds were retrieved from the PubChem database (https://pubchem.ncbi.nlm.nih.gov) in SDF format. Each molecule was converted into its 3D conformation and minimized using Open Babel integrated in UCSF Chimera (version 1.16). Partial charges were assigned using the Gasteiger method [60,61]. The optimized structures were then saved in PDB format and subsequently converted to PDBQT format using AutoDockTools (version 4.2.6).

3.7.2. Protein Preparation

The 3D crystal structure of XO in complex with the flavonoid quercetin was downloaded from the Protein Data Bank (PDB ID: 3NVY). The protein was preprocessed using UCSF Chimera, where all water molecules, ions, and non-essential heteroatoms were removed. Hydrogen atoms were added to satisfy valences at physiological pH, and the resulting structure was saved in PDB format [62,63]. The protein was then prepared for docking in AutoDockTools, where polar hydrogens and Kollman charges were added. The final structure was saved in PDBQT format.

3.7.3. Redocking Validation

To validate the docking protocol, the co-crystallized ligand (quercetin) was first removed from the binding pocket and then re-docked into the same site using AutoDock Vina (v1.1.2). The docking grid was defined to fully encompass the active site using the coordinates of the native ligand. The best docked pose was aligned with the crystallographic conformation using Chimera, and the root-mean-square deviation (RMSD) was calculated. The resulting RMSD value of 0.842 Å indicated a good overlap between the docked and experimental positions, confirming the reliability of the docking settings [64].

3.7.4. Docking Procedure

Molecular docking was performed using AutoDock Vina. The search space was centered around the binding site of the co-crystallized ligand. Exhaustiveness was set to 8 to ensure a reasonable sampling of binding conformations [65]. Each ligand was docked individually into the XO binding pocket. The docking output consisted of nine poses ranked by predicted binding energy. The pose with the lowest binding energy was selected for further analysis.

3.7.5. Interaction Analysis

Docked complexes were visualized and analyzed using Discovery Studio Visualizer 2021 to identify key molecular interactions. Both 2D interaction diagrams and 3D binding poses were generated to highlight hydrogen bonds, hydrophobic contacts, and electrostatic interactions between the ligands and amino acid residues in the XO active site. Particular attention was paid to critical residues such as Glu802, Arg880, Phe1009, Val1011, Leu873, and Glu1261, which are known to contribute significantly to the catalytic activity and inhibitor binding of XO [66,67].

3.8. Calculation of Physicochemical Parameters

An array of physicochemical descriptors of the identified metabolites was employed to contextualize docking outcomes and assess drug-likeness against common rulesets (mainly Lipinski, Veber and Egan): cLog p values, topological polar surface area (TPSA), fractions of sp³ carbons, numbers of H-bond acceptors and donors, and rotatable bonds. Lipophilicity was calculated by the partition coefficient (Clog p) using the corresponding tool in the ChemBioDraw Ultra 21.0 software, while the rest of physicochemical parameters were calculated by the SwissADME software (https://www.swissadme.ch/index.php) (accessed on 22 July 2025) [68,69,70].

4. Conclusions

The present study highlights Z. lotus fruits as a valuable source of bioactive metabolites with antioxidant properties. The EtOAc extract exhibited high levels of total phenolics (281.33 ± 1.5 μg GAE/mg) and flavonoids (127.26 ± 5.89 μg RE/mg), together with significant in vitro antioxidant performance, as demonstrated by its effectiveness against the ABTS^•+ radical (IC₅₀ = 18.49 ± 1.47 μg/mL), phenanthroline-based reducing power (A_0.5 = 8.38 ± 0.69 μg/mL) and xanthine oxidase (IC₅₀ = 170.4 ± 5.90 μg/mL). Chemical profiling by UHPLC–QTOF–HRMS allowed the tentative annotation of eight metabolites, expanding current knowledge of the phytochemical composition of Z. lotus fruits, including the first report of hexadecasphinganine and phytosphingosine within the genus, a finding consistent with the known occurrence of sphingoid bases during fruit development. The experimental findings of this study are limited to phytochemical characterization, antioxidant assays, and XO inhibition at the crude extract level, while the docking analysis represents an exploratory approach for generating hypotheses on potential ligand–protein interactions relevant to oxidative stress and XO inhibition to examine possible ligand–protein interactions within the XO active site, underscoring the need for cautious interpretation of the docking results supported by existing experimental literature. Predictive SAR within close pairs (rutin > kaempferol-O-rutinoside; phytosphingosine > hexadecasphinganine) may suggest a potential benefit of an additional hydroxyl group when it can engage acidic residues without disrupting hydrophobic fit. Future XO inhibition assays at higher scales, kinetic analyses, and cellular ROS studies will be essential to confirm XO modulation, SAR considerations, and to establish their contribution to the antioxidant effects of Z. lotus fruits and their metabolites. Overall, these results strengthen the case for Z. lotus fruits as a promising source of antioxidant metabolites and provide a foundation for subsequent targeted quantification and focused bioactivity assays.