Phytochemical Profiling, Biological Evaluation, and Molecular Insights Into the Urease Inhibitory Potential of Fraxinus angustifolia Crude Extract and Fractions

ABSTRACT

Fraxinus angustifolia Vahl. has long been used in traditional medicine for the management of inflammatory and gastrointestinal disorders. This study investigated the anti‐urease, anti‐Helicobacter pylori, and anti‐inflammatory activities of the ethanolic leaf extract and its phenolic fractions, with a focus on the mechanisms underlying these effects. The extracts exhibited inhibition of urease activity and bacterial growth, with the aqueous fraction of chloroform (AC) showing the most pronounced effect (urease IC₅₀ = 384.55 ± 4.31 µg/mL; MIC = 1000 µg/mL against H. pylori). HPLC‐DAD analysis identified 11 phenolic compounds, including flavones and flavonols. Quercetin‐3‐O‐glucoside (3585.25 µg/g) and rutin (936.11 µg/g), were the most abundant phenolic compounds in the crude extract (CE), whereas chlorogenic acid (824.51 µg/g), rosmarinic acid (586.47 µg/g), and apigenin (148.25 µg/g) predominated in the AC fraction. Enzyme kinetic analysis revealed a mixed mode of urease inhibition, while molecular docking predicted stable binding of phenolic compounds within the urease catalytic pocket and adjacent regions. These findings provide new evidence supporting F. angustifolia leaves as a relevant source of urease inhibitors, suggesting its potential for the management of H. pylori infection and related gastric disorders. Further studies are underway to isolate the active compounds and evaluate their in vivo efficacy.

A phenolic‐rich fraction from Fraxinus angustifolia exhibits urease inhibitory, anti‐Helicobacter pylori, and anti‐inflammatory activities. Chemical profiling and molecular docking suggest that phenolic compounds contribute to these effects, offering new insights into the potential role of F. angustifolia phenolic in urease inhibition and the management of H. pylori‐related gastric disorders.

1. Introduction

Infection with Helicobacter pylori is a leading cause of chronic gastritis and peptic ulcers, which, if left untreated, may progress to gastric cancer. It accounts for approximately 95% of duodenal ulcers and 70% of gastric ulcers [1]. To survive in the acidic environment of the stomach, H. pylori, a spiral‐shaped, Gram‐negative, and flagellated bacterium, produces a critical enzyme, urease, which hydrolyzes urea into ammonia, thus neutralizing gastric acid. Urease plays a pivotal role in the pathogenesis of H. pylori [2], enabling bacterial colonization, thereby contributing to the inflammatory response in the gastric mucosa [3].

Beyond its role in peptic ulcer formation, H. pylori infection also disrupts the redox balance of the gastric mucosa, leading to increased production of reactive oxygen species (ROS) and reactive nitrogen species (RNS), such as nitric oxide (NO), which contribute to oxidative stress and exacerbate mucosal inflammation [4, 5]. Conventional treatments, including antibiotics, such as macrolides (e.g., clarithromycin) and nitroimidazoles (e.g., metronidazole), are often associated with adverse effects and an increase in bacterial resistance [6], underscoring the need for safer and more effective alternatives. Targeting molecular determinants of virulence, particularly urease, emerges as a potential therapeutic strategy to address these challenges. According to the World Health Organization (WHO), medicinal plants are recognized as valuable sources of bioactive compounds with potential therapeutic applications [7]. Among natural products, polyphenols, alkaloids, and terpenoids, derived from plant secondary metabolism, constitute the major classes of bioactive compounds. Their structural diversity, arising from complex biosynthetic pathways, underpins a wide range of pharmacological properties [8]. Many exert their effects through interactions with cellular targets, such as membrane receptors, ion channels, intracellular enzymes, and transport proteins [8].

Fraxinus angustifolia (Oleaceae), commonly known as narrow‐leaved ash, has long been used in traditional medicine in the Mediterranean Basin and North Africa. In Algerian folk medicine, leaf infusions are traditionally employed for their analgesic, anti‐inflammatory, and antibacterial properties [9, 10]. In Morocco, F. angustifolia is used for other traditional purposes, including the relief of gastro‐digestive problems [11]. Its leaves and winged fruits, called samaras, are used as a decoction and infusion to treat rheumatism [12], intestinal parasites [12], and diabetes [13]. In Southern Italy, manna—an exudate obtained from its bark—is traditionally employed as a mild laxative [14].

Previous studies have reported that F. angustifolia exhibits a wide range of biological activities, including antioxidant [15, 16], anti‐inflammatory [17], wound healing [18], hepatoprotective [19], antimicrobial [20, 21], as well as anti‐xanthine oxidase [22], antiproliferative, and anticancer effects [23]. Moreover, its samaras have been reported to improve glucose tolerance [24]. To the best of our knowledge, the potential of F. angustifolia against H. pylori and its urease enzyme has not yet been investigated.

The present study aimed to investigate the anti‐urease, anti‐H. pylori, and anti‐inflammatory activities of F. angustifolia extracts. These biological assays were complemented by HPLC phytochemical profiling and molecular docking analysis to gain insight into the mechanisms underlying urease inhibition and to explore the molecular interactions of selected phenolic compounds with the urease enzyme.

2. Results

2.1. Total Phenolic and Flavonoid Contents in F. angustifolia Extracts

The quantitative chemical analysis revealed that crude ethanolic extract and its fractions from F. angustifolia contain varying amounts of polyphenols and flavonoids (Table 1). The aqueous fraction of ethyl acetate (AA) exhibited the highest total phenolic content (175.65 ± 2.98 mg GAE/g E), followed by the crude ethanolic extract (CE) and the aqueous fraction of chloroform (AC), which showed comparable values (141.79 ± 1.40 mg GAE/g E and 156.90 ± 3.69 mg GAE/g E, respectively). The highest total flavonoid contents were recorded in the AC and AA fractions (70.24 ± 1.14 and 64.16 ± 0.85 mg RE/g E, respectively), with no significant difference between them (p > 0.05).

TABLE 1.

Phenolic and flavonoid contents of F. angustifolia crude extract and its fractions.

Extract and fractions/ standard	Total phenolic compounds (mg GAE/g Extract)	Total flavonoids (mg RE/ g Extract)
CE	141.79 ± 1.40 a	36.67 ± 0.73 a
EA	104.55 ± 2.15 b	44.91 ± 0.05 b
AA	175.65 ± 2.98 c	64.16 ± 0.85 d
CH	90.76 ± 1.19 b	58.78 ± 0.11 d
AC	156.90 ± 3.69 a	70.24 ± 1.14 d

All assays were performed in triplicates. Values with different superscripts (a, b, c, d) within the same column are significantly different (p < 0.05). Data are expressed as mean ± SD (n = 3). AA, aqueous fraction of ethyl acetate; AC, aqueous fraction of chloroform; CE, Crude Ethanolic; CH, organic fraction of chloroform; EA, organic fraction of Ethyl acetate.

2.2. Inhibitory Effect on H. pylori

As shown in Table 2, the CE extract and AC fraction exhibited the highest inhibitory effect on H. pylori growth, both displaying a minimal inhibitory concentration (MIC) of 1000 µg/mL, followed by the organic fraction of chloroform (CH), with an MIC of 2000 µg/mL. However, no inhibitory effect was observed for the remaining fractions on the in vitro growth of H. pylori. On the other hand, quercetin, kaempferol, and rutin showed MIC values of 25, 50, and 100 µg/mL, respectively. Comparatively, the reference antibiotic amoxicillin demonstrated substantially higher potency, with an MIC value of 0.39 µg/mL.

TABLE 2.

Urease inhibitory and anti‐H. pylori activities of F. angustifolia crude extract and its fractions.

	Urease inhibitory activity		Anti‐H. pylori activity
Extract and fractions/standard	Inhibition percentage (%) at 1050 µg/mL	IC50 (µg/mL)	MIC (µg/mL)
CE	68.09 ± 1.89 ***	588.70 ± 9.06 ***	1000 ****
EA	78.49 ± 2.32 ***	504.93 ± 7.75	>2000 ****
AA	79.16 ± 3.18 ***	480.40 ± 22.2	>2000 ****
CH	63.89 ± 1.47 ***	539.95 ± 6.85 *	2000 ****
AC	87.46 ± 1.44 ***	384.55 ± 4.31 ***	1000 ****
Amx	—	—	0.39
BA (1050 µg/mL)	44.70 ± 0.70	499.5 ± 29.3	—
Rutin (120 µM)	67.43 ± 0.26 ***	50.92 ± 1.49 ***	100 ****
Quercetin (120 µM)	94.58 ± 0.98 ***	8.32 ± 0.34 ****	25 **
Kaempferol (120 µM)	—	—	50 ****

All assays were designed in triplicate. Asterisks (*) indicate statistically significant differences compared to the BA standard. *p < 0.05, ** p < 0.01, *** p < 0.001; **** p < 0.0001. Values without asterisks are not significantly different (P ≥ 0.05). IC50: Half maximal Inhibitory Concentration. IC₅₀ value is given as µg/mL. AA, aqueous fraction of ethyl acetate; AC, aqueous fraction of chloroform; Amx, amoxicillin; BA, Boric acid; CE, Crude Ethanolic; CH, organic fraction of chloroform; EA, organic fraction of Ethyl acetate.

2.3. Urease Inhibitory Activity

Urease catalyzes the hydrolysis of urea into carbamic acid, which spontaneously decomposes into carbon dioxide and ammonia (NH₃), the latter being quantified spectrophotometrically at 630 nm. The urease inhibitory activity of F. angustifolia is summarized in Table 2. The different fractions from F. angustifolia extract displayed concentration‐dependent urease inhibitory potential. The highest urease inhibitory activity was exhibited by the AC fraction with 87.46% ± 1.44% at a concentration of 1050 µg/mL and an IC₅₀ of 384.55 ± 4.31 µg/mL, followed by the EA fraction, which showed 78.49% ± 2.32 % inhibition at the same concentration and IC₅₀ of 504.93 ± 7.75 µg/mL. The AC fraction demonstrated greater inhibitory potential than the reference inhibitor boric acid (IC₅₀ of 499.5 ± 29.3 µg/mL). Among the tested phenolic standards, rutin and quercetin showed strong urease inhibitory activity, with IC₅₀ values of 50.92 ± 1.49 µg/mL and 8.32 ± 0.34 µg/mL, respectively. Extracts and fractions exhibiting the most potent inhibition (lowest IC₅₀ values) were further subjected to kinetic analysis to elucidate their mechanisms of inhibition.

2.4. Kinetic Parameters

Kinetic studies were conducted using Lineweaver–Burk plots to determine the inhibition mode of the most active samples. For the AC fraction, the maximal velocity (Vmax) decreased and the lines intersected at a point away from the y‐axis, suggesting a mixed type of inhibition (Figure 1A). In the case of quercetin (Figure 1C), the lines intersected at the same point on the x‐axis, indicating a decrease in the maximum reaction rate (Vmax) and suggesting a non‐competitive type of inhibition. As for rutin (Figure 1D), it inhibited urease in a competitive manner, as evidenced by the lines intersected at the same point on the y‐axis in the Lineweaver‐Burk plot.

FIGURE 1.

Lineweaver‐Burk plots of urease inhibition by (A) AC subfraction of F. angustifolia; boric acid (B); quercetin (C), and rutin (D) versus reciprocal substrate concentrations (0.025—0.4 mM).

2.5. Anti‐Inflammatory Activity

Protein denaturation is a well‐established trigger for inflammatory responses and related diseases. As shown in Table 3, the AA fraction stood out for its significant inhibition of Bovine serum albumin (BSA) denaturation (IC₅₀ = 239.52 ± 11.72 µg/mL), whereas Acetylsalicylic acid (ASA) exhibited a stronger inhibition effect (IC₅₀ = 24.08 ± 0.39 µg/mL).

TABLE 3.

Anti‐inflammatory activity of F. angustifolia crude extract and its fractions [IC₅₀ (µg/mL)].

Extract and fractions/ standard	COX inhibition	BSA denaturation inhibition	NO inhibition
CE	50.29 ± 1.37	>500 ***	99.05 ± 3.32 *
EA	86.37 ± 3.16 ***	> 500 ****	77.91 ± 1.06
AA	65.96 ± 3.46 *	239.52 ± 11.72 **	88.97 ± 2.27
CH	103.4 ± 0.38 ***	> 500 ****	173.57 ± 7.84 ***
AC	189.8 ± 9.57 ***	> 500 ****	108.59 ± 1.26 ***
ASA	45.2 ± 0.52	24.08 ± 0.39	—
CA	—	—	83.56 ± 4.02

All assays were performed in triplicates. IC₅₀ values are given as µg/mL, mean ± standard deviation; Asterisks (*) indicate significant differences compared to the ASA standard and CA standard. * p < 0.05, ** p < 0.01, *** p < 0.001; **** p < 0.0001. Values without asterisks are not significantly different (P ≥ 0.05).

AA, aqueous fraction of ethyl acetate; AC, aqueous fraction of chloroform; ASA, acetylsalicylic acid, CH, organic fsraction of chloroform; CA, caffeic acid, CE, crude ethanolic; EA, organic fraction of Ethyl acetate.

In the COX inhibition assay, all extracts demonstrated a dose‐dependent inhibitory effect on enzyme activity. The CE extract displayed the strongest activity (IC₅₀ = 50.29 ± 1.37 µg/mL), which was statistically comparable to the ASA standard (IC₅₀ = 45.2 ± 0.52 µg/mL; p > 0.05). On the other hand, F. angustifolia crude extract and its factions showed marked inhibition of NO production. Among them, the organic fraction of ethyl acetate (EA) (IC₅₀ = 77.91 ± 1.06 µg/mL) and AA (IC₅₀ = 88.97 ± 2.27 µg/mL) fractions exhibited comparable activities to the caffeic acid used as standard (IC₅₀ = 83.56 ± 4.02 µg/mL; p > 0.05).

2.6. HPLC Analysis

The CE and the most active AC fraction against urease of F. angustifolia leaves were subjected to HPLC‐DAD analysis. Phenolic compounds were identified by comparison of the chromatogram peaks with synthetic standards based on retention time and UV–vis absorption spectra (Representative chromatograms of the F. angustifolia phenolic compounds are shown in Figure 2 and 3), while the corresponding UV spectra of the identified compounds and authentic standards are provided in the Supporting Information. Compound contents were quantified using external calibration curves established with reference standards, and the results are summarized in Table 4 (see Supporting Information for calibration curves and equations).

FIGURE 2.

HPLC profile of CE extract of F. angustifolia. 2,3,4‐THBA: 2,3,4‐Trihydroxybenzoic acid; Q‐3‐O‐glucoside: Quecetin‐3‐O‐glucoside. K‐O‐glucoside: Kaempferol‐O‐glucoside.

FIGURE 3.

HPLC profile of AC subfraction of F. angustifolia. Chlr acid: Chlorogenic acid; P‐Cmr acid: Para‐coumaric acid; Q‐3‐O‐glucoside: Quecetin‐3‐O‐glucoside; K‐O‐glucoside: Kaempferol‐O‐glucoside, and Ros acid: rosmarinic acid.

TABLE 4.

Identification of phenolic compounds in crude ethanolic and aqueous fraction of chloroform of leaf extract of F. angustifolia using HPLC‐DAD retention times.

Peak	Phenolic compound	Rt (min)	λ(nm)	Compound content (µg/g)
				CE Extract	AC fraction
1	Gallic acid	4.213	280	439.53 ± 18.76	—
2	2,3,4‐trihydroxybenzoic acid	7.717	280	86.27 ± 0.89	—
3	P‐Coumaric acid	9.317	280	—	96.34 ± 0.33
4	Syringic acid	20.88	280	385.38 ± 13.07
5	Apigenin	12.8	312; 360		148.25 ± 1.04
6	Gentisic acid	30.13	312	9.97 ± 0.50	—
7	Rosmarinic acid	24.99	312	221.96 ± 7.79	586.47 ± 13.87
8	Rutin	17.067	280; 312; 360	936.11 ± 4.59	316.34 ± 5.78
9	Kaempferol‐O‐glucoside	20.30	280; 312; 360	348.57 ± 28.69	121.39 ± 1.31
10	Quercetin‐3‐O‐glucoside	17.67	280; 312; 360	3585.25 ± 61.33	136.14 ± 2.89
11	Chlorogenic acid	6.98	280	—	824.51 ± 14.89

CE, Crude Ethanolic; AC, aqueous fraction of chloroform, Rt, retention time; λ, detection wavelength.

The results are given as mean ± standard deviation.

Analysis of the chromatograms in full‐scan mode revealed the presence of numerous compounds, which were identified through comparison with authentic reference standards.

In total, 11 different phenolic compounds were identified: eight in the CE extract and seven in the AC fraction, four of which were shared between the two. In the CE extract, two benzoic acid derivatives (gallic acid and 2,3,4‐trihydroxybenzoic acid), two hydroxycinnamic acids (gentisic acid and rosmarinic acid), and one phenylpropanoid (syringic acid) were identified. In addition, three flavonols, that is, rutin, kaempferol‐O‐glucoside, and quercetin‐3‐O‐glucoside, were detected (Figure 2). Major peaks were assigned to quercetin‐3‐O‐glucoside (3585.25 ± 61.33 µg/mg extract) and rutin (936.11 ± 4.59 µg/mg extract). In the AC fraction, three hydroxycinnamic acids (chlorogenic acid, p‐coumaric acid, and rosmarinic acid) were detected, along with one flavone (apigenin) and three flavonols (rutin, quercetin‐3‐O‐glucoside, and kaempferol‐O‐glucoside), as depicted in Figure 3. The most abundant compounds were chlorogenic acid (824.51 ± 14.89 µg/mg extract) and rosmarinic acid (586.47 ± 13.87 µg/mg extract).

2.7. Docking Analysis

The docking protocol was validated by re‐docking the co‐crystallized ligand, which yielded an RMSD value of 0.67 Å (<2 Å) (Figure 4). The docking outcomes demonstrated that the tested compounds exhibited considerable stability when contrasted with the co‐crystallized ligand, as reflected by their docking score values (Table 5). The obtained binding energy for Acetohydroxamic acid (AHA), the standard reference inhibitor of urease, was −4.3 kcal/mol. its key interactions with Ni^{2 +} ions, and HIS221, ASP362, and ALA365 residues (Table 5) were consistent with those reported in the crystallographic pose. Compared to this reference inhibitor, several tested compounds, including apigenin, chlorogenic acid, rutin, and rosmarinic acid, exhibited stronger binding affinities (−8.6; −7.3; −8.4, and −7.2 kcal/mol, respectively). Flavonoid compounds showed enhanced binding affinity in their aglycone forms. In particular, apigenin exhibited a strong docking score (−8.6 kcal/mol), forming five hydrogen bonds with HIS138, HIS322, ARG338, and ASP362, one π–anion interaction with ASP223, and three π–alkyl interaction involving CYS321, and ALA365 residues (Figure 5F).

FIGURE 4.

Superposition of the experimental co‐crystallized AHA reference ligand structure (yellow) and the computationally re‐docked pose (RMSD = 0.67 A°).

TABLE 5.

Binding affinity (kcal/mol), inhibition constant (µM), and interacting residues of identified compounds in comparison with the Acetohydroxamic acid (AHA) standard reference with the target enzyme.

Compounds	ΔG (kcal/mol)	KI (µM)	Hydrogen Bonds (Distance A°)	Hydrophobic Bonding	Electrostatic Bonding
Gallic acid	−6.4	20.3	Gly279 (2.21); Arg338 (2.40); Asp362 (2.67); Ala365 (2.29)	—	Ni3001
P‐Coumaric acid	−5.6	79.0	His138 (2.57); His322 (3.02); Asp362 (2.41)	Ala365	—
2,3,4‐trihydroxybenzoic acid	−6.9	8.7	His138 (2.58); His248 (2.18); Cys321 (2.94); His322 (1.91)	—	Arg338
Syringic acid	−6.2	28.4	His138 (2.41); His322 (2.94); Arg338 (2.11)	Cys321; Ala365	His221
Apigenin	−8.6	0.49	His138 (2.65); His322 (2.07); A338 (2.66); Asp362 (2.36)	Cys321; Ala365	Asp223
Gentisic acid	−6.6	14.5	His138 (2.65); His 221 (2.24); His322 (2.06); Asp362 (2.64)	Ala365	Arg338
Rosmarinic acid	−7.2	5.3	His248(2.66); Asp223 (2.14); Gly279 (2.05)	Ala169; Met366	Cys321; Arg338
Rutin	−8.4	0.70	Asp165 (2.98); Ala169 (2.66); His248 (2.33); His322 (2.14)	—	Cys321; Ala365
Kaempferol‐O‐glucoside	−5.6	78	Asn168 (1.98); His248 (2.35); Gly279 (2.83); Met366 (2.62)	Cys321;	Asp223; Arg338
Quercetin‐3‐O‐glucoside	−5.9	48	Glu222 (2.36); Gly279 (2.59); Met317 (2.54)	Ala365	Cys321
Chlorogenic acid	−7.3	4.45	Ala169 (2.23); Asn168 (2.47); Asp223 (2.13); Asp362 (2.2); Ala365 (2.13)	—	Ni3001
Acetohydroxamic acid	−4.3	700	His221 (3.57); His248 (3.52); Ala362 (2.28); Ala365 (2.08)	—	Ni3001; Ni3002

FIGURE 5.

2D dimension interactions of Quercetin‐3‐O‐glucoside (5A) Chlorogenic acid (5B), Rosmarinic acid (5C), Gallic acid (5D), Kaempferol‐O‐glucoside (5E), and Apigenin (5F) with the target urease enzyme.

Rosmarinic acid showed a favorable binding score (−7.2 kcal/mol) and established multiple stabilizing interactions within the urease active site. It formed three conventional hydrogen bonds with HIS248, ASP223, and GLY279, as well as a π‐cation interaction with ARG338. In addition, it interacted with the key residues CYS321 located on the mobile flap through sulfur interaction, although no direct interaction with the Ni^{2 +} atoms was observed. Furthermore, π‐alkyl contacts with residues ALA169 and MET366 contributed to the stabilization of the ligand–protein complex (Figure 5C). The combination of hydrogen bonding, π–cation and π‐sulfur interactions highlights the strong affinity of rosmarinic acid for the urease active site.

Gallic acid (−6.4 kcal/mol) established interaction patterns similar to those of 2,3,4‐trihydroxybenzoic acid, forming hydrogen bonds with ALA169, GLY279, and ASP362 and coordinating with the Ni3001 metal center. However, unlike 2,3,4‐trihydroxybenzoic acid, gallic acid formed additional hydrogen bonds with ARG338 and ALA365 instead of the two hydrogen bonds observed with HIS221 and HIS322 (Figure 5D).

Chlorogenic acid also achieved a notable docking score (−7.3 kcal/mol), forming four hydrogen bonds with ASP165, ASN168, ASP223, and ASP362, and one metal coordination with the nickel atom (Ni3001) through the carbonyl oxygen (Figure 5B).

Kaempferol‐O‐glucoside, with binding energy of −5.6 kcal/mol, showcased hydrogen bonds with ASN168, GLY279, and MET366, along with electrostatic interactions involving ASP223 and ARG338. On the other hand, quercetin‐3‐O‐glucoside (−5.9 kcal/mol) engaged in hydrogen bonds with GLU222, GLY279, and MET317, and an electrostatic interaction with CYS321 (Figure 5A).

Syringic acid and gentisic acid showed binding energies of −6.2 and −6.6 kcal/mol, respectively, and shared the same binding interactions with urease, involving hydrogen bonds with HIS138 and HIS322 and hydrophobic interaction with ALA365 residue.

3. Discussion

Urease inhibition has emerged as a promising therapeutic approach against H. pylori, a pathogen strongly implicated in gastritis, peptic ulcers, and even gastric cancer [25, 26]. Although several natural products have been reported as urease inhibitors, relatively few studies have explored species traditionally used in North Africa. We report here the inhibition of urease activity and H. pylori growth by crude extract and its fractions from F. angustifolia, a medicinal plant widely employed in Algerian folk medicine for the treatment of inflammatory disorders.

Our findings revealed significant variability in the total phenolic content of the crude extract and its fractions from F. angustifolia, with the AA fraction exhibiting the highest content (175.65 ± 2.98 mg GAE/g dry extract). This variability can be explained by differences in solvent polarity, which influence the solubility and recovery of phenolic compounds, as previously reported by Xu and Chang [27].

The crude ethanolic extract and its fractions exhibited notable urease inhibitory activity, with the AC fraction showing the strongest effect (IC₅₀ value of 384.55 ± 4.31 µg/mL). This fraction also inhibited H. pylori growth, with a MIC of 1000 µg/mL. Previous work by Zouaghi et al. [20] highlighted the antibacterial potential of F. angustifolia against diverse Gram‐negative and Gram‐positive microorganisms. Similarly, Ourabah et al. [21] demonstrated that F. angustifolia exhibited antimicrobial effects by inhibiting the formation of biofilm. In addition, phenolic constituents of F. mandshurica Rupr., a related species, have been reported to exert anti‐H. pylori effects. In this regard, Akter et al. [28] reported MIC values ranging from 12.5 to 100 µM, with fraxetin being the most effective compound, exhibiting an MIC of 12.5 µM against both H. pylori strains 43504 and 26695.

In relation to urease inhibition, several studies have emphasized the role of phenolic compounds, particularly flavonoids such as flavanones and isoflavones, as potent urease inhibitors in diverse plant species [29, 30, 31, 32]. Their inhibitory activity has been largely attributed to their hydroxyl groups, which can interact with thiol, carbonyl, or carboxyl residues within the urease active site, as highlighted by previous reports [25, 33, 34]. This mechanism likely contributes to the strong activity observed in the AC fraction. Consistently, several extracts from plants belonging to the Oleaceae family have been reported to inhibit H. pylori urease [35, 36].

Given the established role of H. pylori in chronic gastric inflammation and ulcer development [4], we evaluated the anti‐inflammatory potential of F. angustifolia crude extract and its fractions. Significant anti‐inflammatory effects were observed (Table 3), as evidenced by the inhibition of BSA denaturation, nitric oxide (NO) production, and COX activity. Among the tested samples, the AA fraction exhibited the strongest inhibition of protein denaturation (IC₅₀ = 239.52 ± 11.72 µg/mL) and NO production (IC₅₀ = 88.97 ± 2.27 µg/mL). A recent study has substantiated the anti‐inflammatory effect of F. angustifolia through inhibition of inflammatory cytokine production [37]. Likewise, another study conducted by Moulaoui et al. [18] on this plant's ethanolic extract demonstrated marked anti‐inflammatory effect in animal models by inhibiting the pro‐inflammatory MPO enzyme. These anti‐inflammatory findings are consistent with previous studies on Fraxinus species, which have been reported to possess anti‐inflammatory properties [17, 38]. Moreover, individual phenolic constituents of the genus Fraxinus, such as fraxidin‐O‐β‐D‐glucoside [9], rutin [16], chlorogenic acid [39], quercetin [23], and gallic acid [38] have also been shown to exert significant anti‐inflammatory effects, further supporting the role of phenolic compounds in the observed bioactivity. The ability of F. angustifolia extract and fractions to inhibit COX activity, scavenge NO, and prevent BSA denaturation underscores their potential to modulate key processes associated with inflammation, consistent with the bioactivity expected from their phenolic constituents.

The crude ethanolic extract (CE), as well as the most active AC fraction against urease, was subjected to HPLC analysis. Phytochemical profile of F. angustifolia leaf extracts revealed a rich diversity of polyphenolic compounds, including phenolic acids, hydroxycinnamic acids, flavones, flavonols, and phenylpropanoids.

The AC fraction was found to contain quercetin‐3‐O‐glucoside, rutin, apigenin, p‐coumaric acid, and other phenolic derivatives, with chlorogenic acid identified as the major constituent. Chlorogenic acid, one of the most abundant dietary polyphenols, has been extensively reported for its anti‐ulcer [40, 41] and anti‐inflammatory activities [42]. Kataria et al. [43] further demonstrated its pronounced urease inhibitory activity (IC₅₀ = 22.68 ± 0.06 µg/mL), suggesting that it largely accounts for the anti‐urease potency of the AC fraction. Docking simulations provided molecular support for these observations, as chlorogenic acid exhibited a binding energy of −7.3 kcal/mol and formed stable hydrogen bonds with residues ASP165, ASN168, ASP223, and ASP365 at the urease active site (Figure 5B). These interactions may contribute to the observed inhibitory activity in the AC fraction.

Rosmarinic acid, also detected in this fraction, might have contributed to the observed effects, as its multiple hydroxyl groups enhance interactions within the urease active site [33]. Docking analysis supported this hypothesis, revealing a favorable binding score (−7.2 kcal/mol), stabilized by hydrogen bonds with HIS248, ASP223, and GLY279, a π‐cation contact with ARG338, and a π‐sulfur interaction with CYS321, located on the mobile flap of urease. Notably, Cys321 is one of the key residues involved in the catalytic activity of this enzyme [33]. Beyond its urease inhibition, rosmarinic acid has been reported to exhibit a broad spectrum of biological activities, including anti‐inflammatory [44], antioxidant, nitric oxide inhibitory [45], and antiviral effects [46]. The mixed‐type inhibition observed for the AC fraction can be rationalized by the docking results. While chlorogenic acid and rosmarinic acid both interact within the catalytic region, the mixed pattern observed in kinetic assays indicated that additional interactions may occur at other binding sites, likely involving other phenolic constituents. Quercetin‐3‐O‐glucoside, another significant compound, exhibited a binding energy of −5.9 kcal/mol, forming multiple hydrogen bonds via its hydroxyl groups with GLU222, GLY279, and Met317 together with a π‐sulfur interaction with CYS321, indicating a favorable binding pattern. Notably, one of the hydrogen‐bond interactions involved the 5‐OH group of the flavonoid scaffold and GLU222. This observation is consistent with the structure–activity relationship study of Xiao et al. [47], which identified the 5‐OH group as one of the key structural features contributing to urease inhibitory activity. Other phenolic acids detected exclusively in the crude extract, such as gentisic and syringic acids, also showed favorable in silico binding (−6.6 and −6.2 kcal/mol, respectively) to the urease active site, providing additional molecular insight into their potential interactions with urease. Furthermore, apigenin was identified here for the first time in F. angustifolia, although it has previously been reported in other Fraxinus species, such as F. ornus [48] and F. excelsior [9]. Additionally, rutin, quercetin‐3‐O‐glucoside, and rosmarinic acid have been reported in F. angustifolia [16, 19]. In summary, the present study revealed that F. angustifolia extract and its fractions possess urease inhibitory and anti‐H. pylori activities, along with significant anti‐inflammatory effects. These bioactivities may be largely attributed to their rich phenolic profile, particularly quercetin‐3‐O‐glucoside, chlorogenic acid, rosmarinic acid, kaempferol‐O‐glucoside, and apigenin. Docking analysis provided molecular support for their urease inhibitory activity. Overall, this study expands current knowledge on the pharmacological potential of the Fraxinus genus and supports the potential role of phenolic constituents from F. angustifolia in urease inhibition and the management of H. pylori–related gastric disorders.

4. Conclusion

The present study demonstrated that F. angustifolia is a valuable source of multi‐target bioactive compounds with anti‐urease and anti‐inflammatory activities supported by both in vitro and in silico analyses. For the first time, we report the anti‐urease, anti‐ H. pylori and anti‐inflammatory activities of F. angustifolia crude extract and its fractions, with the aqueous fraction of chloroform exhibiting the most pronounced effects. This fraction effectively targeted a key virulence factor of the H. pylori pathogen by inhibiting urease activity, while mitigating inflammatory damage through modulation of NO and COX pathways. These biological activities are consistent with the high phenolic content of the CE and AC, as confirmed by HPLC‐DAD analysis. Additionally, molecular docking studies suggested favorable interactions of the major identified compounds with the urease target, providing mechanistic insights that may underlie the observed urease inhibitory effect. Overall, these findings suggest F. angustifolia as a promising source of bioactive compounds for the development of therapeutic strategies against H. pylori‐associated gastric diseases. While the antibacterial and urease inhibitory effects were assessed separately in vitro, it is plausible that in vivo, urease inhibition may enhance the antibacterial effect by maintaining physiological gastric pH, with concurrent anti‐inflammatory actions helping to prevent ulcer formation. Further studies, including targeted isolation of active constituents and in vivo evaluation in animal models, are warranted to fully elucidate the anti‐ulcer potential of F. angustifolia and its phenolic compounds.

5. Materials and methods

5.1. Chemicals and Plant Material

5.1.1. Chemicals

Ethanol, methanol, ethyl‐acetate, and chloroform (analytical grade) were obtained from Sigma‐Aldrich. Folin‐Ciocalteu reagent, sodium chloride (NaCl), gallic acid, quercetin, rutin, aluminium chloride (AlCl₃), nutrient agar, Mueller‐Hinton agar, Mueller‐Hinton broth, sodium carbonate (Na₂CO₃), bovine serum albumin (BSA) were purchased from Biochem (Montreal, Quebec, Canada). Sodium chloride (NaCl) was purchased from BIOCHEM Chemopharma (USA). Tris(hydroxymethyl)aminomethane hydrochloride (Tris–HCl), was purchased from Sigma‐Aldrich.

Standard compounds including rutin, cinnamic acid, chlorogenic acid, p‐coumaric acid, tannic acid, benzoic acid, kaempferol, rosmarinic acid, vanillin, quercetin, chrysin, quercetin 3‐O‐rhamnoside, epicatechin, quercetin 3‐O‐glucoside, and kaempferol 3‐O‐glucoside were purchased from Sigma‐Aldrich (St. Louis, Missouri, USA). HPLC grade solvents including water, acetonitrile and hydrochloric acid (HCl) were purchased from Merck (Darmstadt, Germany).

5.1.2. Plant Material

The leaves of F. angustifolia were harvested in July 2018 from the municipality of Chemini (Bejaia, Algeria). The Global Positioning Systems (GPS) location is 36.35°N 4.36°E. Plant material was identified by Professor Hacène Abdelkrim according to a listed voucher specimen (O/n°59) in the herbarium of Department of Botany, ENSA (Ecole Nationale des Sciences Agronomiques), El‐Harrach (Algiers, Algeria). The collected material was air‐dried in the dark at room temperature and then finely ground into fine powder (∼63 µm) using a commercial grinder (KIKA Labortechnik, Staufen, Germany).

5.1.3. Extraction and Fractionation Procedures

The extraction was performed according to the method described by Atmani et al. (2009) [15]. The plant powder (150 g) was macerated in ethanol 96% (1:4 w/v) for 24 h under agitation. After decantation, the combined extracts were concentrated until dryness under reduced pressure using rotavapor (Heidolph, Germany), yielding 9 g of crude ethanolic extract (CE).

A portion of the CE extract was first partitioned between ethyl acetate and water (1:3:1, w/v/v), yielding an organic ethyl acetate fraction (EA) and an aqueous fraction (AA). The EA fraction was then concentrated to dryness. A portion of the dried EA fraction was subsequently suspended in a chloroform/water mixture (1:3:1, w/v/v) and subjected to liquid–liquid partitioning. After phase separation, two fractions were obtained: an organic chloroform fraction (CH) and the corresponding aqueous fraction (AC), both of which were concentrated to dryness. The crude extract (CE) and all dried fractions (EA, AA, CH, and AC) were stored at −20 °C until use.

5.2. Bacterial Strains and Culture Conditions

The reference strain H. pylori ATCC 700824 (J99), originally isolated from the gastric biopsy of a patient with a duodenal ulcer, was obtained from the Pasteur Institute of Algeria (IPA, Algiers, Algeria) and used for the antibacterial assay. The identity of the strain was confirmed based on its growth under microaerophilic conditions, Gram‐staining characteristics, cell morphology, and positive urease activity. Bacterial cultures were maintained on Mueller–Hinton agar (MHA) supplemented with 5% horse blood. Plates were incubated at 37 °C for 72 h under microaerophilic conditions (5% O₂, 10% CO₂, and 85% N₂) [49].

5.3. Methods

5.3.1. Determination of Total Phenolic and Flavonoid Contents

Folin–Ciocalteu method was employed for the estimation of total phenolic content in F. angustifolia extracts following the method of Djeridane et al. [50]. Briefly, 100 µL of extract was mixed thoroughly with 500 µL of Folin–Ciocalteu reagent, followed by adding 1000 µL of distilled water. The mixture was pre‐incubated at room temperature for one minute. Then 1500 µL of 20% (w/v) sodium carbonate was added. The final mixture was incubated for 2 h at room temperature, in the dark. Absorbance was measured at 765 nm. Total phenolic content was expressed as milligram of gallic acid equivalent per gram of dry weight (mg GAE/g), extrapolated from a standard curve using gallic acid as standard.

On the other hand, F. angustifolia extract and its fractions were evaluated for their flavonoid content through the aluminum chloride colorimetric method described by Djeridane et al. [50]. Briefly, 1 mL of extract/fraction was added to 1 mL of 2% AlCl₃ methanol solution. After incubation in the dark for 15 min, the absorbance was read at 430 nm. The flavonoid content was calculated from a calibration curve using rutin as a standard. Results were expressed as milligram rutin equivalent per gram dry weight (mg RE/g).

5.3.2. Inhibitory Effect on H. pylori

The inhibitory effect of F. angustifolia against H. pylori was assessed by the determination of the minimal inhibitory concentration (MIC). MICs were determined using the micro dilution method according to CLSI guidelines [51]. In brief, 160 µL of Mueller‐Hinton broth supplemented with 5% horse blood was sterilely added to each well, 40 µL of extract was added, and 100 µL of the previously prepared inoculum (adjusted to 2 McFarland) was added to each well, resulting in a final concentration of 10⁷ CFU/mL. Final extract concentrations ranged from 0.0039 to 2 mg/mL. The reference antibiotic used was amoxicillin, with final concentrations ranging from 0.19 to 100 µg/mL. The inoculum (100 µl) was then transferred into 96‐well plates already containing 100 µl of twofold serially diluted sample or antibiotic standard in Mueller Hinton growth medium supplemented with 5% horse blood. The micro plate was incubated at 37 °C under microaerophilic conditions for 48 h. Amoxicillin served as a positive control, while broth without inoculum was used as a negative control. The minimum inhibitory concentration (MIC) was defined as the lowest concentration inhibiting visible bacterial growth. It is determined when no turbidity and / or cell growth was observed in the micro plate well after incubation [51].

5.3.3. Urease Inhibition Assay

Urease inhibitory activity was determined according to the indophenol blue spectrophotometric method described by Weatherburn [52], with some modifications. Briefly, in a 96‐well microplate, 25 µL of jack‐bean urease enzyme solution (4 units) was added to 15 µL of extract solution. The mixture was pre‐incubated at 30°C for 15 min. After that, 40 µL of urea solution (100 mM; 100 mM phosphate buffer; pH 7.4) was added. The mixture was incubated at 30°C for 30 min. After incubation, 70 µL of alkaline reagent (0.5% w/v NaOH and 0.3% active NaOCl) and 50 µL of phenol reagent (1% w/v phenol and 0.005% w/v sodium nitroprusside) were added to each well. The final mixture was incubated for 50 min at 37°C in the dark. Urease inhibitory activity was determined by measuring the production of ammonia. Absorbances were then measured at 630 nm. All reactions were carried out in triplicate. The percentage of urease inhibition was calculated using the formula:

$$\mathrm{Inhibition}\%=\left[\left({\mathrm{A}}_0-{\mathrm{A}}_1\right)/{\mathrm{A}}_0\right]\times 100$$

where, A₀ represents the absorbance of the control (enzyme without extract) and A₁ the absorbance in the presence of extract.

5.3.4. Determination of Inhibition Mode

The most active extracts and fractions were selected to determine their inhibition mode using Lineweaver–Burk plots. Initial reaction velocities (V) were determined by monitoring ammonia production at 630 nm over 50 min, with urea as the substrate at concentrations ranging from 25 to 400 µM. Assays contained a fixed urease concentration and inhibitor at its IC₅₀ value. The reciprocal of velocity (1/V) was plotted against the reciprocal of substrate concentration (1/[S]). The mode of inhibition was inferred from the characteristic intersection pattern of the regression lines compared with the control. All assays were carried out in triplicate.

5.3.5. Anti‐Inflammatory Assays

The anti‐inflammatory activity was evaluated using three assays: cyclooxygenase (COX) inhibition, nitric oxide (NO) scavenging, and the inhibition of bovine serum albumin (BSA) denaturation. The COX inhibition assay was performed following the method described by Schultz et al. [53]. Meanwhile, the no scavenging assay by using the method of Ajayi et al. [54]. The BSA denaturation assay was assayed according to the method of Harrabi et al. [55]. Acetylsalicylic acid (ASA) and caffeic acid (CA) were used as positive controls. All samples were evaluated in triplicates at different concentrations (6.25–500 µg/mL). The results are expressed as 50% inhibitory concentration (IC₅₀) values (µg/mL).

5.3.6. HPLC Conditions

F. angustifolia active extracts were subjected to HPLC analysis with a photodiode array detector (HPLC‐DAD). The phenolic profiles of samples were identified using the method of Capanoglu et al. [56]. Sample extracts were filtered with a 0.45 µm membrane filter and analyzed using a Waters 2695 HPLC system equipped with a PDA (Waters 2996) detector. A Supelcosil LC‐18 (25 cm × 4.60 mm, 5 µm column Sigma‐Aldrich, Steinheim, Germany) was used. The mobile phases were composed of Milli‐Q water with 0.1 percent (v/v) trifluoroacetic acid (TFA) (Mobile A) and acetonitrile with 0.1 percent (v/v) TFA (Mobile B). A linear gradient was utilized with 95% solvent A and 5% solvent B at 0 min, 65% solvent A and 35% solvent B at 45 min, and 25% solvent A and 75% solvent B at 47 min, returning to original conditions at 50 min. The flow rate was maintained at 1 mL/min. Detection was carried out at 280, 312, and 360 nm. Identification was based on retention times and characteristic UV spectra. Compound contents were quantified using external calibration curves established with authentic reference standards.

5.3.7. Molecular Docking Study

To gain insight into binding conformations and interaction between target enzyme (urease) and main compounds from F. angustifolia, the molecular docking study was performed using Autodock software version 4.2 (The Scripps Research Institute, USA [57]. The three‐dimensional crystal structure used as a target corresponded to the entry Crystal structure of H. pylori urease in complex with acetohydroxamic acid (PDB ID: 1E9Y), experimentally determined by x‐ray diffraction at a resolution of 3.0 Å and obtained from the RCSB Protein Data Bank (https://www.rcsb.org/).

The 3D structures of gallic acid (PubChem CID: 370), chlorogenic acid (PubChem CID: 1794427), quercetin 3‐O‐glucoside (PubChem CID: 5280804), apigenin (PubChem CID: 5280443), rutin (PubChem CID: 5280805), rosmarinic acid (PubChem CID: 5281792), kaempferol‐O‐glucoside (PubChem CID: 25203515), p‐coumaric acid (PubChem CID: 637542), syringic acid (PubChem CID: 10742), and gentisic acid (CID: 3469) were retrieved from the PubChem database (https://pubchem.ncbi.nlm.nih.gov/). Before the docking simulation, all ligands were optimized using Avogadro [58]. The protein was prepared using AutoDock Tools by removing water molecules, adding any missing atoms, adding polar hydrogens, and assigning charges. The center of the active site was set at 127.121, 129.092, and 86.81 Å for x, y, and z centers, respectively. The size of the grid box was set to 20 × 20 × 20 Å³ (x, y, z) with a spacing of 0.375 Å. Redocking of the native ligand acetohydroxamic acid (AHA), co‐crystallized with urease (PDB ID: 1E9Y), was performed to validate the docking parameters. AHA was also used as the reference standard inhibitor (positive control) in the docking study. The binding conformations and molecular interaction analysis of the docked compounds with the urease enzyme were performed using the Discovery Studio 2020 client. (Dassault Systèmes BIOVIA, 2021).

5.3.8. Statistical Analysis

All assays were performed in triplicate. Data were expressed as mean ± standard deviation (SD). The differences were considered statistically significant at p<0.05. Multiple comparisons between more than two groups were performed with one‐way ANOVA supplemented with Tukey's test using the Graph Pad Prism software version 8.1.

Author Contributions

Sarra Belkhir: investigation, methodology, data curation, formal analysis, funding acquisition, writing – original draft. Meriem Rahmani‐berboucha: methodology, data curation, investigation, formal analysis, writing – original draft. Sarah Sihem Zemam: methodology, investigation, data curation. Nadjet Debbache‐benaida: writing – review and editing, supervision. Fatma Duygu Ceylan: methodology, investigation, data curation, validation, writing – original draft. Nabil Adrar: methodology, investigation, writing – original draft, writing – review and editing, data curation. Amina Atia: methodology, investigation, writing – original draft, data curation. Naima Saidene: methodology, data curation, investigation, writing – original draft. Nabila Benamrouche: methodology, visualization, writing – review and editing. Dina Atmani‐kilani: conceptualization, visualization, writing – review and editing, supervision. Esra Capanoglu: methodology, writing – review and editing.

Conflicts of Interest

The authors declare that this manuscript is original, unpublished, that there are no plans to publish it elsewhere, and that there are no conflicts of interest to disclose.

Supporting information

Supporting File 1: cbdv71583‐sup‐0001‐SuppMat.pdf

Data Availability Statement

The data are available from the corresponding author upon reasonable request.