Chemical Composition, Antioxidant and Antimicrobial Activities of Moroccan Equisetum ramosissimum Essential Oil and Butanolic Extract

ABSTRACT

For the first time, the chemical profile and biological activities of Moroccan Equisetum ramosissimum essential oil (EO) were investigated, alongside comparative antioxidant and antimicrobial evaluation of a previously characterized butanolic extract (BE) and molecular docking of selected constituents. Hydrodistillation yielded 0.067% (v/w, dry weight) EO, characterized by gas chromatography‐mass spectrometry (GC‐MS) and gas chromatography–flame ionization detection (GC–FID) and dominated by oxygenated monoterpenes and sesquiterpenes, with (+)‐pulegone, γ‐eudesmol, cyperotundone, viridiflorol, and carvone as major constituents. Antioxidant activity was evaluated using 2,2‐diphenyl‐1‐picrylhydrazyl (DPPH), reducing power, and β‐carotene bleaching assays. The EO exhibited an effective concentration producing an absorbance of 0.5 (EC₅₀) of 50.0 µg/mL in the reducing power assay, whereas the BE showed lower IC₅₀ (50% inhibitory concentration) values than the EO in the DPPH and β‐carotene bleaching assays. Antimicrobial testing against 15 microbial strains revealed minimum inhibitory concentration (MIC) values of 0.19–12.5 mg/mL, with the lowest values against Candida albicans and Candida glabrata. Molecular docking provided complementary insights into potential interactions of selected major compounds with microbial targets. Overall, these in vitro and in silico findings support further investigation of Moroccan E. ramosissimum through toxicological and in vivo studies.

Moroccan Equisetum ramosissimum essential oil showed a distinct pulegone‐eudesmol‐rich profile and strain‐dependent antimicrobial activity, including against resistant isolates. The butanolic extract displayed stronger antioxidant activity in DPPH and β‐carotene bleaching assays, while docking suggested plausible interactions with selected microbial targets.

1. Introduction

Antimicrobial resistance has become a major global public health concern, compromising the effectiveness of conventional antimicrobial therapies and increasing the burden of difficult‐to‐treat infections. The emergence and spread of multidrug‐resistant microorganisms, including carbapenem‐resistant Gram‐negative bacteria, methicillin‐resistant Staphylococcus aureus, and vancomycin‐resistant enterococci, further emphasize the need to explore alternative sources of antimicrobial compounds [1, 2, 3]. In this context, medicinal plants and their secondary metabolites have attracted considerable interest as sources of structurally diverse bioactive compounds with potential antimicrobial and antioxidant properties [4, 5, 6].

Equisetum ramosissimum Desf. (Equisetaceae), locally known in Morocco as “Danab elkhayl,” is a perennial rhizomatous pteridophyte distributed across temperate and subtropical regions of the Old World, including Morocco [7, 8]. Botanically, the species is characterized by erect, longitudinally ribbed aerial stems, with highly reduced leaves fused into nodal sheaths and branches arising from the nodes; the reproductive structures are borne in terminal strobili [7]. The aerial parts of E. ramosissimum have a long history of traditional use for the management of urinary tract disorders, inflammatory and rheumatic conditions, musculoskeletal pain, respiratory complaints, and infectious diseases [9, 10]. These ethnomedicinal applications have prompted increasing interest in the phytochemical composition and biological properties of this species.

Previous phytochemical investigations have demonstrated that E. ramosissimum contains a diversity of secondary metabolites, particularly phenolic acids and flavonoid derivatives. Sureshkumar et al. characterized the chemical profile of E. ramosissimum stem extracts and reported antioxidant and antimicrobial properties [8]. In Moroccan plant material, our previous high‐performance liquid chromatography‐mass spectrometry (HPLC‐MS) investigation of the methanolic and butanolic extracts (BEs) revealed several phenolic constituents, including kaempferol glycosides and ferulic acid, while the extracts also exhibited antioxidant, analgesic, and anti‐inflammatory activities [11]. Antioxidant activity has also been reported for methanolic, aqueous, and ethyl acetate extracts of E. ramosissimum, although considerable variations in activity have been observed according to extraction solvent and geographical origin [12, 13, 14]. More recently, Korkmaz et al. evaluated ethanolic and methanolic extracts against a panel of bacterial and fungal reference strains, including methicillin‐resistant S. aureus, Acinetobacter baumannii, Pseudomonas aeruginosa, Candida albicans, and Candida glabrata, confirming the antimicrobial potential of non‐volatile extracts of the species [15]. In addition, liquid chromatography‐tandem mass spectrometry (LC‐MS/MS) profiling of an ethanolic extract from Turkish E. ramosissimum identified 13 phenolic compounds, including kaempferol, vanillic acid, and resveratrol, and demonstrated antioxidant and enzyme‐inhibitory properties [16].

In contrast to the increasing number of studies devoted to non‐volatile extracts, information on the essential oil of E. ramosissimum remains scarce. Alebous et al. characterized the hydrodistilled essential oil obtained from Jordanian plant material and identified 37 volatile constituents, with α‐bisabolol oxide A and cuminaldehyde as the predominant compounds [17]. However, that study focused on chemical characterization and did not investigate the antimicrobial activity of the essential oil. Moreover, essential‐oil composition may vary substantially according to geographical origin, climatic and edaphic conditions, phenological stage, genetic background, and processing conditions [18, 19, 20, 21]. Consequently, Moroccan populations of E. ramosissimum may exhibit a distinct volatile chemical profile and potentially different biological properties.

Despite the available evidence concerning the biological activities of E. ramosissimum extracts, important gaps therefore remain regarding its volatile fraction. In particular, the chemical profile and biological properties of the essential oil obtained from Moroccan E. ramosissimum have not been characterized, and its antimicrobial potential against a broad panel including clinically characterized multidrug‐resistant isolates remains unexplored. Accordingly, the novelty of the present study lies in the chemical characterization of Moroccan E. ramosissimum essential oil and in the integrated evaluation of its antioxidant and antimicrobial properties against reference microorganisms and clinically relevant resistant isolates, alongside comparison with a previously characterized active butanolic extract. Molecular docking was further employed as a complementary hypothesis‐generating approach to explore potential interactions between selected constituents and microbial targets.

2. Results

2.1. Extract Yield and Chemical Composition of the Essential Oil

The essential oil obtained by hydrodistillation from the air‐dried aerial parts of Moroccan E. ramosissimum was dark brown in color and had a strong, persistent odor. The extraction yield was 0.067% (v/w), calculated relative to the dry weight of the plant material. Overall, 89.18% of the volatile constituents were identified by GC‐MS analysis (Figure S1 and Table 1). The oil was predominantly composed of oxygenated monoterpenes (36.29%) and oxygenated sesquiterpenes (35.24%). The major constituents were (+)‐pulegone (15.23%), γ‐eudesmol (8.43%), cyperotundone (6.99%), viridiflorol (6.23%), carvone (5.74%), cadin‐4‐en‐10‐ol (5.64%), and spathulenol (4.15%).

TABLE 1.

Chemical compounds of Equisetum ramosissimum essential oil.

Chemical class/compound	Percentage (%)	RI exp.	RI lit.
Oxygenated monoterpenes	36.29
Eucalyptol	0.49	1032	1033
Camphor	1.89	1149	1150
Menthone	0.49	1158	1158
(+)‐Pulegone	15.23	1241	1241
Carvone	5.74	1246	1245
Piperitone	0.86	1267	1267
Thymol	1.99	1293	1293
Isoascaridole	2.17	1306	1306
Cumin alcohol	4.15	1310	1313
Piperitenone	1.33	1340	1345
Piperitenone oxide	1.95	1372	1372
Oxygenated sesquiterpenes	35.24
α‐Elemol	1.49	1546	1546
Spathulenol	4.15	1576	1576
Viridiflorol	6.23	1594	1596
γ‐Eudesmol	8.43	1598	1610
epi‐γ‐Eudesmol	0.67	1624	1624
Valerianol	0.50	1657	1655
Cadin‐4‐en‐10‐ol	5.64	1659	1660
Shyobunol	0.57	1686	1686
Cyperotundone	6.99	1693	1694
Nootkatone	0.57	1806	1805
Sesquiterpene hydrocarbons	1.95
γ‐Muurolene	1.67	1485	1485
(Z)‐α‐Bisabolene	0.28	1503	1502
Aldehydes (non‐terpenoid)	5.64
3‐Furfural	0.30	816	815
Benzaldehyde	0.53	960	961
Benzene acetaldehyde	4.46	1036	1037
Vanillin	0.35	1394	1396
Alcohols (non‐terpenoid)	5.35
4‐Methylpentanol	0.20	831	835
Furfuryl alcohol	3.70	849	845
Phenethyl alcohol	0.48	1113	1112
(2E,4E)‐Hexadienol	0.64	1164	1165
Nona‐(2E,6Z)‐dienol	0.33	1170	1172
Ketones (non‐terpenoid)	1.56
2,3‐Dihydro‐2,5‐dimethylfuran‐3‐one	0.16	930	930
Maltol	1.15	1108	1106
Pyruvophenone	0.25	1169	1167
Organic acids	0.32
n‐Pentanoic acid	0.32	911	913
Phenylpropanoids/phenolics	2.34
p‐Vinylguaiacol	2.34	1309	1310
Chromene derivatives	0.4
Desmethoxyencecalin	0.4	1645	1645
Total identified compounds	89.18

Note: RI exp., Experimental retention index relative to C8–C25 n‐alkanes on a TG‐5 MS column; RI lit., Literature retention index reported in the NIST database.

2.2. Antioxidant Activity

Table 2 summarizes the antioxidant activity of E. ramosissimum essential oil (EO) and butanolic extract (BE), expressed as IC₅₀ values for the DPPH and β‐carotene bleaching assays and EC₅₀ values for the reducing power assay. The antioxidant effects varied depending on both the tested sample and the assay used. The BE exhibited significantly greater activity than the EO in the DPPH and β‐carotene bleaching assays, as indicated by its lower IC₅₀ values (p < 0.05). In the DPPH assay, the BE exhibited an IC₅₀ of 64.33 ± 0.384 µg/mL, compared with 780 ± 15.945 µg/mL for the EO. Similarly, in the β‐carotene bleaching assay, the BE showed an IC₅₀ of 253 ± 17.521 µg/mL, whereas the EO reached 1238 ± 52.368 µg/mL. Conversely, in the reducing power assay, the EO displayed significantly greater activity than the BE, with EC₅₀ values of 50.00 ± 0.463 and 83.11 ± 0.583 µg/mL, respectively (p < 0.05). The reference antioxidants, quercetin and butylated hydroxytoluene (BHT), exhibited significantly lower IC₅₀ and EC₅₀ values than the EO and BE in the corresponding assays (p < 0.05).

TABLE 2.

Antioxidant activity of Equisetum ramosissimum essential oil (EO) and butanolic extract (BE), expressed as IC₅₀ values for the DPPH and β‐carotene bleaching assays and EC₅₀ values for the reducing power assay.

	DPPH (µg/mL)	Reducing power (µg/mL)	β‐Carotene (µg/mL)
Essential oil	780 ± 15.945d (740.39–819.61)	50 ± 0.463c (48.85–51.15)	1238 ± 52.368d (1107.90–1368.10)
BE	64.33 ± 0.384c (63.38–65.28)	83.11 ± 0.583d (81.66–84.56)	253 ± 17.521c (209.47–296.53)
Quercetin	2.7 ± 0.02b (2.65–2.75)	2.89 ± 0.029b (2.82–2.96)	1.07 ± 0.05a (0.95–1.19)
BHT	1.56 ± 0.01a (1.54–1.58)	1.16 ± 0.01a (1.14–1.18)	4.75 ± 0.93b (2.44–7.06)

Note: Values are expressed as mean ± SD of three technical replicates (n = 3) obtained from the same sample preparation. IC₅₀ and EC₅₀ values are presented with their corresponding 95% confidence intervals (95% CI). EC₅₀ refers to the effective concentration producing an absorbance of 0.5 in the reducing‐power assay. Different superscript letters within the same column indicate significant differences among the tested samples (p < 0.05), as determined by one‐way ANOVA followed by Tukey's post hoc test.

2.3. Antimicrobial Activity

As shown in Tables 3 and 4, the antimicrobial activity of E. ramosissimum essential oil (EO) and butanolic extract (BE) varied according to the microbial group tested, including yeasts, Gram‐positive bacteria, and Gram‐negative bacteria.

TABLE 3.

Inhibition zones (mm) of Equisetum ramosissimum essential oil (EO) and butanolic extract (BE) against tested fungal strains determined by the disk diffusion method.

Tested organisms	EO	BE	Fluconazole
Candida albicans BU7098	31 ± 2	11 ± 0.1	18 ± 0.12
Candida glabrata BU7170	25 ± 0.85	13 ± 0.22	20 ± 0.26

Note: Values are expressed as mean ± SD of three technical replicates (n = 3) obtained from the same sample preparation. Fluconazole (5 µg/disc) was used as a positive control for Candida species.

TABLE 4.

Inhibition zones (mm) of Equisetum ramosissimum essential oil (EO) and butanolic extract (BE) against tested bacterial strains determined by the disk diffusion method.

Tested organisms	EO	BE	Gentamicin
Gram‐positive bacteria
Vancomycin‐resistant Enterococcus spp.	14 ± 0.31	10 ± 0.11	NI
Enterococcus faecium ATCC 4602	9 ± 0.15	7 ± 0.33	29.67 ± 0.7
Methicillin‐sensitive Staphylococcus aureus P1085	13 ± 0.5	10 ± 0.09	25.0 ± 0.6
Methicillin‐resistant Staphylococcus aureus NCTC 12493	15 ± 0.68	12 ± 0.23	NI
Gram‐negative bacteria
Multidrug‐resistant Acinetobacter baumannii P1415	12 ± 0.11	9 ± 0.33	NI
Pseudomonas aeruginosa ATCC 27853	8 ± 0.2	6 ± 0.15	17.0 ± 0.31
Colistin‐resistant Pseudomonas aeruginosa P623	12 ± 0.66	10 ± 0.31	22.67 ± 0.4
Ertapenem‐resistant Salmonella spp. H18	12 ± 0.25	9 ± 0.5	NI
Klebsiella pneumoniae ATCC 700603	9 ± 0.09	8 ± 0.13	NI
Ertapenem‐resistant Klebsiella pneumoniae H2121	10 ± 0.1	8 ± 0.09	NI
Escherichia coli ATCC 25922	11 ± 0.45	8 ± 0.33	18.67 ± 0.61
Imipenem‐resistant Escherichia coli H1764	11 ± 0.33	9 ± 0.12	NI
Carbapenem‐resistant Enterobacter cloacae P611	8 ± 0.15	8 ± 0.09	16 ± 0.1

Note: Values are expressed as mean ± SD of three technical replicates (n = 3) obtained from the same sample preparation. Gentamicin (15 µg/disc) was used as a positive control for bacteria. NI: no inhibition observed.

Among yeasts, C. albicans BU7098 was the most susceptible strain, showing the largest inhibition zone with the EO (31 ± 2.00 mm), followed by C. glabrata BU7170 (25 ± 0.85 mm). Representative disk‐diffusion plate illustrating the activity of the EO against C. glabrata, together with the positive and negative controls, is provided in Figure S2. Both yeast strains showed smaller inhibition zones when treated with the BE.

For Gram‐positive bacteria, the largest inhibition zones produced by the EO were observed against methicillin‐resistant S. aureus NCTC 12493 (15 ± 0.68 mm) and vancomycin‐resistant Enterococcus spp. (14 ± 0.31 mm). Among the non‐resistant/reference Gram‐positive strains, methicillin‐sensitive S. aureus P1085 showed an inhibition zone of 13 ± 0.50 mm, whereas Enterococcus faecium ATCC 4602 showed the smallest inhibition zone (9 ± 0.15 mm).

Among the resistant Gram‐negative isolates, the EO produced inhibition zones of 12 ± 0.11 mm against multidrug‐resistant A. baumannii P1415, 12 ± 0.66 mm against colistin‐resistant P. aeruginosa P623, 12 ± 0.25 mm against ertapenem‐resistant Salmonella spp. H18, 10 ± 0.10 mm against ertapenem‐resistant Klebsiella pneumoniae H2121, 11 ± 0.33 mm against imipenem‐resistant Escherichia coli H1764, and 8 ± 0.15 mm against carbapenem‐resistant Enterobacter cloacae P611. Among the reference strains, the EO produced inhibition zones of 8 ± 0.20 mm against P. aeruginosa ATCC 27853, 9 ± 0.09 mm against K. pneumoniae ATCC 700603, and 11 ± 0.45 mm against E. coli ATCC 25922. Overall, the BE produced smaller inhibition zones than the EO under the tested disk diffusion conditions, particularly against Gram‐negative bacteria.

Tables 5 and 6 present the minimum inhibitory concentrations (MICs) and minimum bactericidal/fungicidal concentrations (MBCs/MFCs) of E. ramosissimum essential oil (EO) and butanolic extract (BE) against yeasts, Gram‐positive bacteria, and Gram‐negative bacteria, together with the corresponding MBC/MIC and MFC/MIC ratios. The antimicrobial response varied according to the tested microorganism and sample.

TABLE 5.

Minimum inhibitory concentration (MIC), minimum fungicidal concentration (MFC), and MFC/MIC ratio of Equisetum ramosissimum essential oil (EO) and butanolic extract (BE) against the tested yeast strains.

	MIC				MFC			Ratio
Test organism	EO	BE		Fluconazole	EO	BE	Fluconazole	EO	BE
Candida albicans BU7098	0.19		0.39	0.031	0.19	0.78	0.031	1	2
Candida glabrata BU7170	0.19		0.78	0.5	0.19	1.56	0.5	1	2

Note: Values are expressed in mg/mL. Fluconazole was used as a reference for yeasts. The MFC/MIC ratio was calculated for yeast strains. Ratios ≤ 4 were considered indicative of fungicidal activity, whereas ratios > 4 were considered indicative of fungistatic activity.

TABLE 6.

Minimum inhibitory concentration (MIC), minimum bactericidal concentration (MBC), and MBC/MIC ratio of Equisetum ramosissimum essential oil (EO) and butanolic extract (BE) against the tested bacterial strains.

	MIC			MBC			Ratio
Test organism	EO	BE	Gentamicin	EO	BE	Gentamicin	EO	BE
Gram‐positive bacteria
Vancomycin‐resistant Enterococcus spp.	0.78	3.12	NI	1.56	3.12	NI	2	1
Enterococcus faecium ATCC 4602	1.56	6.25	NT	3.12	6.25	NT	2	1
Methicillin‐sensitive Staphylococcus aureus P1085	1.56	3.12	0.008	3.12	6.25	0.015	2	2
Methicillin‐resistant Staphylococcus aureus NCTC 12493	0.39	1.56	NI	0.39	1.56	NI	1	1
Gram‐negative bacteria
Multidrug‐resistant Acinetobacter baumannii P1415	0.78	3.12	NI	1.56	3.12	NI	2	1
Pseudomonas aeruginosa ATCC 27853	1.56	6.25	NT	3.12	12.5	NT	2	2
Colistin‐resistant Pseudomonas aeruginosa P623	3.12	12.5	NT	6.25	12.5	NT	2	1
Ertapenem‐resistant Salmonella spp. H18	0.78	0.39	NI	0.78	0.78	NI	1	2
Klebsiella pneumoniae ATCC 700603	12.5	6.25	NI	12.5	6.25	NI	1	1
Ertapenem‐resistant Klebsiella pneumoniae H2121	6.25	6.25	NI	6.25	12.5	NI	1	2
Escherichia coli ATCC 25922	0.39	1.56	0.005	0.39	3.12	0.005	1	2
Imipenem‐resistant Escherichia coli H1764	1.56	3.12	NI	6.25	6.25	NI	4	2
Carbapenem‐resistant Enterobacter cloacae P611	6.25	3.12	NT	12.5	3.12	NT	2	1

Note: Values are expressed in mg/mL. Gentamicin was used as a reference antibiotic for bacteria. NI, no inhibition observed at the highest tested concentration; NT, not tested. The MBC/MIC ratio was calculated for bacterial strains. Ratios ≤ 4 were considered indicative of bactericidal activity, whereas ratios > 4 were considered indicative of bacteriostatic activity.

Among yeasts, C. albicans BU7098 and C. glabrata BU7170 showed the lowest MIC values for the EO (0.19 mg/mL), with MFC values identical to the corresponding MICs, resulting in MFC/MIC ratios of 1 for both strains. The BE exhibited MIC values of 0.39 and 0.78 mg/mL against C. albicans and C. glabrata, respectively, with an MFC/MIC ratio of 2 for both strains. Fluconazole showed a lower MIC against C. albicans, whereas its MIC against C. glabrata was higher than that recorded for the EO.

Among Gram‐positive bacteria, the resistant strains showed comparatively low MIC values. Methicillin‐resistant S. aureus NCTC 12493 exhibited MIC values of 0.39 and 1.56 mg/mL for the EO and BE, respectively, while vancomycin‐resistant Enterococcus spp. showed corresponding MIC values of 0.78 and 3.12 mg/mL. In comparison, methicillin‐sensitive S. aureus P1085 required higher concentrations of both samples (1.56 mg/mL for the EO and 3.12 mg/mL for the BE), and E. faecium ATCC 4602 showed MIC values of 1.56 and 6.25 mg/mL, respectively. The corresponding MBC for methicillin‐resistant S. aureus NCTC 12493 was identical to the MIC, resulting in an MBC/MIC ratio of 1 and indicating a bactericidal effect under the tested conditions. Vancomycin‐resistant Enterococcus spp. showed an MBC of 1.56 mg/mL for the EO, corresponding to an MBC/MIC ratio of 2.

Among Gram‐negative bacteria, the response varied according to the tested strain. Among the resistant isolates, multidrug‐resistant A. baumannii P1415 and ertapenem‐resistant Salmonella spp. H18 showed relatively low EO MIC values of 0.78 mg/mL, whereas colistin‐resistant P. aeruginosa P623, ertapenem‐resistant K. pneumoniae H2121, imipenem‐resistant E. coli H1764, and carbapenem‐resistant E. cloacae P611 showed EO MIC values of 3.12, 6.25, 1.56, and 6.25 mg/mL, respectively. Among the reference strains, the EO MIC values were 0.39 mg/mL for E. coli ATCC 25922, 1.56 mg/mL for P. aeruginosa ATCC 27853, and 12.5 mg/mL for K. pneumoniae ATCC 700603. The MBC/MIC ratios ranged from 1 to 4 for the EO and from 1 to 2 for the BE. Based on the commonly used threshold of ≤4, the antibacterial effects against these Gram‐negative strains were classified as bactericidal under the tested conditions. The BE exhibited comparable or lower MIC values than the EO against some Gram‐negative strains, particularly ertapenem‐resistant Salmonella spp. H18, K. pneumoniae ATCC 700603, and carbapenem‐resistant Enterobacter cloacae P611.

Comparison of resistant isolates and reference strains revealed marked strain‐dependent differences in the concentrations required for growth inhibition. For example, colistin‐resistant P. aeruginosa P623 required twofold higher MICs than P. aeruginosa ATCC 27853 for both the EO and BE. Similarly, imipenem‐resistant E. coli H1764 required a fourfold higher EO concentration and a twofold higher BE concentration than E. coli ATCC 25922. However, this trend was not consistent across all strains, as some resistant isolates, such as multidrug‐resistant A. baumannii P1415 and ertapenem‐resistant Salmonella spp. H18, remained susceptible at relatively low concentrations.

Overall, the antimicrobial effects of the EO and BE were strain‐dependent. Low MIC values were obtained against Candida species and selected bacterial strains, whereas MIC values of 6.25–12.5 mg/mL were recorded for the less susceptible microorganisms. Gentamicin and fluconazole generally exhibited lower MIC values than the plant‐derived samples.

2.4. Molecular Docking Study

The molecular docking results presented in Table 7 revealed clear differences in docking scores among the tested compounds across the selected protein targets. The docking protocol was validated by redocking the co‐crystallized ligands into their respective active sites. The resulting root‐mean‐square deviation (RMSD) values ranged from 0.61 Å to 1.98 Å, remaining below the commonly accepted threshold of 2.0 Å and supporting the ability of the docking protocol to reproduce the crystallographic binding poses.

TABLE 7.

Molecular docking scores (kcal/mol) of major compounds identified in Equisetum ramosissimum essential oil and butanolic extract against selected protein targets.

Ligand/Protein		CYP51	PBP2a	GyrB	LpxC
PDB Code		5V5Z	3ZFZ	5MMN	3P3G
RMSD (Å)		1.77	1.98	0.61	1.03
Co‐crystallized ligand		−12	−6.5	−8.9	−7.4
EO	Cyperotundone	−9.1	−7.0	−6.1	−6.6
	γ‐Eudesmol	−8.6	−6.5	−6.6	−6.3
	Pulegone	−7.5	−5.7	−5.4	−5.3
BE	Ferulic acid	−6.5	−5.5	−6.3	−5.5
	Kaempferol 3,7‐diglucoside	−8.5	−7.7	−8.2	−7.9
	Kaempferol 3‐O‐sophoroside‐7‐O‐glucoside	−7.1	−6.7	−7.3	−7.8

Abbreviations: BE, butanolic extract; EO: essential oil.

Note: The co‐crystallized ligands were used as reference ligands for validation of the docking protocol.

Overall, the kaempferol glycosides exhibited the most favorable docking scores among the tested natural compounds. Kaempferol 3,7‐diglucoside showed the most favorable and consistent docking scores toward penicillin‐binding protein 2a (PBP2a) (−7.7 kcal/mol), DNA gyrase subunit B (GyrB) (−8.2 kcal/mol), and UDP‐3‐O‐(R‐3‐hydroxymyristoyl)‐N‐acetylglucosamine deacetylase (LpxC) (−7.9 kcal/mol), with values comparable to or more favorable than those of the corresponding co‐crystallized ligands for PBP2a and LpxC. The second kaempferol derivative also demonstrated favorable docking scores, particularly toward GyrB and LpxC. Among the essential oil constituents, cyperotundone displayed the most favorable docking score for lanosterol 14α‐demethylase (CYP51) (−9.1 kcal/mol), together with less favorable scores toward the bacterial targets, whereas γ‐eudesmol showed intermediate docking scores. In contrast, (+)‐pulegone exhibited the least favorable docking scores across all targets. The co‐crystallized ligands showed the most favorable docking scores toward CYP51 (−12.0 kcal/mol) and GyrB (−8.9 kcal/mol), while the best‐performing natural compounds showed comparable or more favorable scores than the reference ligands for PBP2a and LpxC.

Analysis of the 2D interaction diagrams (Figures 1, 2, 3, 4) showed that the highest‐scoring ligands shared several key interactions with the co‐crystallized ligands within the active sites. In penicillin‐binding protein 2a (PBP2a), kaempferol 3,7‐diglucoside formed conventional hydrogen bonds and π–cation interactions with residues including LYS A:153, ASN A:164, and SER A:403, which were also involved in the interaction network of the reference ligand. In DNA gyrase subunit B (GyrB), both kaempferol derivatives and the co‐crystallized ligand established hydrogen‐bond interactions with conserved residues, including ASP A:73, ASN A:46, and THR A:165. In UDP‐3‐O‐(R‐3‐hydroxymyristoyl)‐N‐acetylglucosamine deacetylase (LpxC), the best‐scoring kaempferol glycosides formed hydrogen bonds with ASP A:242 and THR A:191, together with a metal‐acceptor interaction involving the catalytic Zn^{2 +} ion (ZN A:301), similarly to the reference ligand. For lanosterol 14α‐demethylase (CYP51), cyperotundone adopted a binding mode mainly stabilized by hydrophobic and π–sigma interactions near the heme group (HEM A:601) and surrounding residues, including PHE A:126 and THR A:122, comparable to those observed for the co‐crystallized ligand. These interaction diagrams illustrate predicted ligand–residue contacts within the selected binding sites and do not demonstrate enzyme inhibition.

FIGURE 1.

Predicted docking interactions of the co‐crystallized ligand (3P3) (A) and kaempferol 3,7‐diglucoside (B) within the active site of LpxC.

FIGURE 2.

Predicted docking poses of the co‐crystallized ligand (AI8) (A) and kaempferol 3,7‐diglucoside (B) within the active site of PBP2a.

FIGURE 3.

Predicted docking poses of the co‐crystallized ligand (O54) (A) and kaempferol 3,7‐diglucoside (B) within the active site of GyrB.

FIGURE 4.

Predicted docking interactions of the co‐crystallized ligand (1YN) (A) and cyperotundone (B) within the active site of CYP51.

3. Discussion

The present study provides an integrated characterization of the chemical profile and biological properties of Moroccan E. ramosissimum, combining essential‐oil analysis with antioxidant, antimicrobial, and molecular docking investigations. The findings revealed marked differences between the volatile and non‐volatile fractions, as well as assay‐ and strain‐dependent biological responses. Importantly, the results also highlight a distinct chemical profile for the Moroccan essential oil and antimicrobial activity against several clinically relevant resistant isolates.

To the best of our knowledge, this is the first report on the yield and chemical composition of E. ramosissimum essential oil from Morocco. This yield is lower than that reported by Alebous et al. [17] for E. ramosissimum collected in Jordan, which yielded 0.125% under similar hydrodistillation conditions. This difference may be related to geographical origin, climatic conditions, harvesting period, plant chemotype, and extraction parameters, all of which are known to influence essential oil biosynthesis and accumulation [19, 20, 21].

GC‐MS analysis identified 89.18% of the total volatile constituents, with oxygenated monoterpenes (36.29%) and oxygenated sesquiterpenes (35.24%) representing the predominant chemical classes. (+)‐Pulegone, γ‐eudesmol, cyperotundone, viridiflorol, carvone, cadin‐4‐en‐10‐ol, and spathulenol were the principal constituents. The remaining 10.82% corresponded to low‐abundance compounds that could not be assigned under the analytical conditions used. Although present at low levels, a possible contribution of these unidentified minor constituents to the biological activities cannot be excluded.

Published information specifically addressing the hydrodistilled essential oil of E. ramosissimum remains very limited. Alebous et al. [17] reported a Jordanian EO also dominated by oxygenated constituents, with oxygenated monoterpenes and oxygenated sesquiterpenes accounting for 36.1% and 23.6%, respectively. However, marked differences were observed in the major constituents. While the Moroccan EO was dominated by (+)‐pulegone, γ‐eudesmol, cyperotundone, viridiflorol, and carvone, the Jordanian oil was mainly characterized by α‐bisabolol oxide A (12.3%) and cuminaldehyde (9.8%). Thus, the identification of a pulegone–eudesmol‐rich profile represents a distinctive chemical feature of the Moroccan E. ramosissimum EO compared with the previously reported Jordanian oil. Such variability may be associated with geographical origin, environmental conditions, phenological stage, and genetic background [18, 20, 21].

The present study represents the first comparative evaluation of the antioxidant activity of E. ramosissimum EO and BE using multiple in vitro assays. Both samples exhibited measurable and assay‐dependent antioxidant effects, although their activities remained lower than those of the reference antioxidants, particularly for the EO in the DPPH and β‐carotene bleaching assays.

Previous studies on E. ramosissimum have mainly focused on polar extracts. Paulsamy et al. [12] reported a DPPH IC₅₀ of 78.58 µg/mL for a methanolic extract, which is close to that obtained for the BE in the present study. Similarly, Li et al. [13] reported an IC₅₀ of 43.41 ± 7.68 µg/mL for an ethyl acetate extract, while Štajner et al. [14] obtained an IC₅₀ of 63.2 ± 2.2 µg/mL for an aqueous extract. In contrast, Sureshkumar et al. [8] reported a weaker DPPH scavenging activity for an Indian methanolic extract, with an IC₅₀ of 123.89 ± 0.73 µg/mL. These differences highlight the influence of extraction solvent and geographical origin on the antioxidant capacity of E. ramosissimum.

The greater antioxidant activity of the BE in the DPPH and β‐carotene bleaching assays may be related to its phenolic and flavonoid composition, previously characterized by HPLC–MS [11]. Constituents such as kaempferol 3,7‐O‐diglucoside, kaempferol 3‐O‐sophoroside‐7‐O‐glucoside, ferulic acid, kaempferol di‐hexoside, tannins, and saponins may contribute to its radical‐scavenging properties.

In contrast, the EO showed greater activity than the BE in the reducing‐power assay, indicating that the relative antioxidant performance of the two samples depended on the assay employed. The antioxidant activity of the EO may be related to its high content of oxygenated monoterpenes and sesquiterpenes. Major constituents such as (+)‐pulegone, γ‐eudesmol, cyperotundone, viridiflorol, carvone, and cumin alcohol have been associated with antioxidant activity, either individually or within complex essential oils [22, 23]. Nevertheless, the contribution of individual constituents and possible synergistic interactions cannot be established from the present experiments.

The antimicrobial findings extend current knowledge on E. ramosissimum by evaluating the EO and BE against yeasts, Gram‐positive and Gram‐negative bacteria, including clinically relevant resistant isolates. A major contribution of the present findings is the antimicrobial evaluation of the chemically characterized Moroccan EO against this diversified microbial panel, alongside comparison with the BE.

Previous investigations have mainly focused on solvent extracts. Sureshkumar et al. [8] reported an MIC of 0.25 mg/mL for a methanolic extract against E. coli, which is comparable to the lowest MIC values obtained against selected bacterial strains in the present study. More recently, Korkmaz et al. [15] also demonstrated antimicrobial activity of ethanolic and methanolic extracts of E. ramosissimum against bacterial and fungal strains. However, their study concerned solvent extracts rather than the essential oil and used a different antimicrobial testing approach, limiting direct quantitative comparison with the present results. In addition, the essential oil of Equisetum arvense, a closely related species, has shown antimicrobial activity against E. coli, S. aureus, K. pneumoniae, and C. albicans [24].

Nevertheless, inhibition‐zone diameters should be interpreted with caution, particularly when comparing the EO with reference antimicrobial agents, because the hydrophobic and volatile constituents of essential oils may behave differently in agar‐based diffusion systems [25].

Another important observation was that susceptibility to the EO and BE did not consistently parallel the conventional antibiotic‐resistance phenotype. Some resistant isolates required higher concentrations than the corresponding reference strains, whereas others, including multidrug‐resistant A. baumannii P1415 and ertapenem‐resistant Salmonella spp. H18, remained susceptible at relatively low concentrations. Such variability may reflect strain‐specific differences in cell‐envelope composition, efflux mechanisms, membrane permeability, and susceptibility to the multiple bioactive constituents present in the EO and BE. Therefore, conventional antibiotic resistance did not necessarily translate into reduced susceptibility to these chemically complex plant‐derived samples.

Taken together, the antimicrobial effects of the EO and BE were clearly strain‐dependent. Low MIC values were obtained against Candida species and selected bacterial strains, whereas MIC values of 6.25–12.5 mg/mL against the less susceptible microorganisms reflected moderate activity under the tested conditions. Gentamicin and fluconazole were generally active at substantially lower concentrations than the plant‐derived samples. However, direct comparisons of potency should be interpreted cautiously because the EO and BE are complex mixtures tested at mg/mL concentrations, whereas the reference drugs are purified antimicrobial agents.

The antimicrobial activity of E. ramosissimum EO may be associated with its high content of oxygenated terpenoids [26, 27]. Pulegone has been particularly well documented for its antimicrobial properties. Oumzil et al. reported that pulegone was the most active constituent of Mentha suaveolens essential oil against Gram‐positive and Gram‐negative bacteria and fungi [28]. Pulegone was also reported as the only major constituent of Calamintha nepeta EO showing antimicrobial activity and was particularly active against Salmonella species [29].

Carvone has demonstrated antibacterial activity against S. aureus, including methicillin‐resistant strains, with MIC values ranging from 500 µg/mL to 1000 µg/mL [30]. It has also been reported to enhance gentamicin activity against MRSA and to affect bacterial membrane integrity in S. aureus and E. coli [31]. Viridiflorol may also contribute to the observed activity, as an essential oil containing viridiflorol as a major constituent exhibited activity against S. aureus, E. coli, A. baumannii, Salmonella Typhi, and several yeast species, including C. glabrata [32]. However, because viridiflorol was evaluated within a complex essential oil, its individual contribution cannot be established. These comparative data support a possible contribution of the major oxygenated terpenoids to the activity of the EO, although the effect of the complete oil may result from additive or synergistic interactions among major and minor constituents.

The antimicrobial activity of the BE may similarly be related to its phenolic and flavonoid composition previously characterized by HPLC–MS [11]. Ferulic acid has demonstrated antibacterial activity against E. coli, P. aeruginosa, and S. aureus, and Borges et al. reported alterations in bacterial surface properties, membrane permeability, and leakage of intracellular constituents [33]. Antimicrobial activity has also been reported for kaempferol derivatives. Özçelik et al. demonstrated activity of kaempferol‐3,7‐O‐α‐l‐dirhamnoside against several microorganisms relevant to the present study, including S. aureus, E. faecalis, E. coli, K. pneumoniae, A. baumannii, and C. albicans [34]. The antimicrobial activity of the BE may therefore reflect the combined contribution of ferulic acid, kaempferol derivatives, and other phenolic constituents.

Mechanistically, essential‐oil constituents and phenolic compounds may affect microbial cells by disturbing membrane integrity, increasing membrane permeability, and altering membrane‐associated proteins [35]. In fungal cells, such compounds may also interfere with ergosterol biosynthesis, inhibit mycelial growth, and suppress spore germination [36]. These mechanisms may help explain the observed antimicrobial activity; however, they were not directly investigated in the present study and should therefore be considered possible mechanisms rather than experimentally demonstrated ones.

Molecular docking was used as a complementary approach to explore potential interactions between selected constituents and microbial targets in relation to the experimentally observed antimicrobial activity. While docking cannot establish a definitive mechanism of action, it provides a useful framework for generating hypotheses concerning possible molecular interactions.

The flavonoid derivatives identified in the BE showed favorable predicted interactions with bacterial targets including PBP2a, GyrB, and LpxC. However, these predictions do not establish a mechanistic relationship with the MIC and MBC results. Indeed, the BE was less active than the EO against methicillin‐resistant S. aureus NCTC 12493 (MIC: 1.56 vs. 0.39 mg/mL, respectively), despite favorable docking scores for selected BE constituents. This finding indicates that docking scores for individual compounds do not directly predict the antimicrobial efficacy of complex plant‐derived samples.

For the EO, selected oxygenated terpenoids also showed favorable predicted interactions with microbial targets, notably PBP2a and the fungal enzyme CYP51. These in silico findings indicate plausible interactions with biologically relevant targets but do not demonstrate that these interactions are responsible for the experimentally observed antimicrobial activity.

Importantly, the lack of a strict correlation between individual docking scores and antimicrobial potency suggests that the biological activity of the EO and BE cannot be attributed to a single compound or target. Instead, multiple constituents and mechanisms may contribute to the overall activity. Accordingly, the docking findings should be considered hypothesis‐generating rather than evidence of a confirmed antimicrobial mechanism.

Several limitations should be considered when interpreting the present findings. The plant material originated from a single geographical area and harvesting period, and the number of isolates representing each resistance phenotype was limited. In addition, the EO and BE are complex mixtures, preventing definitive attribution of the observed biological activities to individual constituents or interactions among them. Finally, the proposed molecular interactions were based on docking predictions and were not experimentally validated. Further studies involving additional clinical isolates, isolated constituents, mechanistic assays, and appropriate in vivo models are therefore warranted.

4. Conclusions

This study reports, for the first time, the chemical composition and biological activities of the essential oil of Moroccan E. ramosissimum, together with the antioxidant and antimicrobial properties of its butanolic extract. The EO was characterized by a predominance of oxygenated monoterpenes and oxygenated sesquiterpenes, with (+)‐pulegone, γ‐eudesmol, and cyperotundone as major constituents. Both samples exhibited assay‐dependent in vitro antioxidant and strain‐dependent antimicrobial activities, with the EO showing low MIC values against Candida species and selected bacterial strains, whereas the BE displayed greater antioxidant activity than the EO in the DPPH and β‐carotene bleaching assays and inhibitory effects against selected bacterial strains. Molecular docking provided complementary insights into the potential interactions of selected constituents with microbial targets. Future studies should focus on validating these predicted mechanisms, identifying the compounds responsible for the observed activities, evaluating possible synergistic effects with conventional antimicrobials, and assessing safety and efficacy through cytotoxicity and in vivo studies.

5. Experimental Section

5.1. Plant Material

The aerial parts of E. ramosissimum were collected at the flowering stage in April 2020 from the Ourika region (31°21′04.55″ N, 7°45′53.70″ W). The plant material was taxonomically identified by Prof. Ouhammou in accordance with the Flora of Morocco [7]. A voucher specimen was deposited at the herbarium of the Faculty of Sciences Semlalia, Cadi Ayyad University, under the reference code MARK‐10118.

5.2. Butanolic Extraction

Using a Soxhlet apparatus, 150 g of powdered aerial parts of E. ramosissimum were initially extracted with dichloromethane (800 mL). After 2 h, the plant residue was successively re‐extracted with 80% methanol (1000 mL) and subsequently with saturated n‐butanol (1000 mL), each extraction being carried out for 24 h.

The resulting butanolic extract was filtered and concentrated under reduced pressure at 45°C using a rotary evaporator and then stored at 4°C until further biological assays. The butanolic extract was selected for further investigation because it exhibited the highest activity among the different solvent extracts evaluated during preliminary screening. Its chemical composition was previously characterized by our research group using HPLC–MS, as reported by Sissi et al. [11]. The same extract was used in the present study without further re‐extraction or modification.

5.3. Essential Oil Extraction

Two hundred grams of air‐dried aerial parts of E. ramosissimum were subjected to hydrodistillation using a Clevenger‐type apparatus for 4 h. The hydrodistillation procedure was performed three times independently under the same experimental conditions. The obtained essential oil was then stored in sealed amber vials at 4°C until chemical analysis and biological assays.

The essential oil yield was calculated according to the following equation:

$$\mathrm{Yield}\left(\%\right)=\frac{V_{\mathrm{EO}}}{D_{\mathrm{W}}}\times 100$$

where V _EO represents the volume of essential oil obtained (mL) and D _W corresponds to the dry weight of the plant material used for extraction (g).

5.4. Gas Chromatography

The chemical composition of the essential oil obtained from the aerial parts of E. ramosissimum was analyzed by GC–FID and GC‐MS. Analyses were performed using a Thermo Scientific Trace GC Ultra system equipped with two split/splitless injectors and an FID detector, coupled to a Thermo Scientific ISQ single quadrupole mass spectrometer, and fitted with a TG‐5 MS column (30 m × 0.25 mm × 0.25 µm). The analytical conditions were as follows: helium was used as the carrier gas at a flow rate of 1.5 mL/min; the injector and detector temperatures were set at 230°C and 250°C, respectively; the electron impact ionization energy was 70 eV; and mass spectra were acquired in full‐scan mode over an m/z range of 40–500. The essential oil sample (1 µL) was injected at a split ratio of 1:100. The column temperature was initially held at 60°C for 1 min, then increased to 230°C at a rate of 3°C/min.

The constituents were identified based on their retention indices (RI), calculated relative to a homologous series of n‐alkanes (C8–C25) analyzed under the same experimental conditions, and by comparison of their mass spectra with those from the NIST 08 Mass Spectra Library (Version 2.0f), Wiley Library, and Adams Library. The relative percentage of each compound was calculated from GC–FID peak areas, without correction factors [37].

5.5. Antioxidant Activity

For each antioxidant assay, every concentration of the EO, BE, and reference antioxidants was tested in three technical replicates using the same sample preparation. Results were expressed as mean ± standard deviation (SD), with n = 3 representing technical replicates.

5.5.1. Free Radical–Scavenging Activity Using 2,2‐Diphenyl‐1‐picrylhydrazyl (DPPH)

The antioxidant activity of the essential oil and butanolic extract obtained from the aerial parts of E. ramosissimum was evaluated using the DPPH radical scavenging assay. An aliquot of 50 µL of each concentration was transferred into test tubes, followed by the addition of 2 mL of a methanolic DPPH solution (60 µM).

The reaction mixtures were incubated in the dark for 20 min at room temperature, after which the absorbance was measured at 517 nm using a UV–visible spectrophotometer [38]. Butylated hydroxytoluene (BHT) and quercetin were used as reference antioxidants.

The percentage of DPPH radical scavenging activity was calculated using the following equation:

$$\mathrm{DPPH}\mathrm{scavenging}\mathrm{effect}\left(\%\right)=\frac{\left(\mathrm{A}\left(\mathrm{blank}\right)-\mathrm{A}\left(\mathrm{sample}\right)\right)}{\mathrm{A}\left(\mathrm{blank}\right)}\times 100$$

The IC₅₀ value was defined as the concentration of the sample required to scavenge 50% of the DPPH radicals, as previously reported [38].

5.5.2. ‐Carotene/Linoleic Acid Bleaching Assay

The β‐carotene/linoleic acid bleaching assay was performed according to the method described by Miraliakbari and Shahidi [39]. Briefly, 0.5 mg of β‐carotene was dissolved in 1 mL of chloroform, after which 25 µL of linoleic acid and 200 mg of Tween 40 were added. The chloroform was completely removed under reduced pressure, and 100 mL of distilled water was subsequently added. The resulting mixture was vigorously stirred to form a stable β‐carotene/linoleic acid emulsion.

Aliquots of 350 µL of different concentrations of the EO, BE, or reference antioxidants (BHT and quercetin) were transferred into test tubes containing 2.5 mL of the freshly prepared emulsion. The tubes were vortexed and incubated in a water bath at 50°C for 2 h. A negative control was prepared by replacing the sample with methanol under the same conditions. A blank without β‐carotene was also prepared to calibrate the spectrophotometer.

After incubation, the absorbance of each sample was measured at 470 nm using a UV–visible spectrophotometer. The percentage of β‐carotene bleaching inhibition was calculated using the following equation:

where A _s,_2h and A _b,2h correspond to the absorbance values of β‐carotene remaining after 2 h in the sample and blank, respectively, and A _ib represents the absorbance measured at the beginning of the experiment. The IC₅₀ value was determined from the dose–response curve by plotting the percentage of inhibition against the sample concentration.

5.5.3. Reducing Power Assay

The reducing power of the essential oil, butanolic extract, and reference antioxidants was evaluated according to the method described by Thaipong et al. [40]. Briefly, 1 mL of different concentrations of the EO, BE or positive controls (BHT and quercetin), prepared in methanol, was mixed with 2.5 mL of phosphate buffer (0.2 M, pH 6.6) and 2.5 mL of potassium ferricyanide solution [K₃Fe(CN)₆, 1%].

The reaction mixtures were incubated at 50°C for 20 min, then cooled to room temperature, and 2.5 mL of trichloroacetic acid (10%) was added to stop the reaction. The mixtures were subsequently centrifuged at 3000× g for 10 min. An aliquot of 2.5 mL of the supernatant was then mixed with 2.5 mL of distilled water and 0.5 mL of ferric chloride (FeCl₃, 0.1%).

The absorbance was measured at 700 nm against a blank prepared under the same conditions by replacing the sample with methanol [40]. The EC₅₀ value, defined as the concentration required to achieve an absorbance of 0.5 at 700 nm, was determined from the corresponding absorbance curve plotted as a function of sample concentration.

5.6. Antimicrobial Activity

5.6.1. Microbial Strains and Culture Preparation

A total of 15 pathogenic microorganisms were used in this study, comprising reference strains and clinical isolates with different antimicrobial susceptibility profiles, including multidrug‐resistant isolates: methicillin‐resistant S. aureus NCTC 12493, colistin‐resistant P. aeruginosa P623, imipenem‐resistant E. coli H1764, ertapenem‐resistant K. pneumoniae H2121, C. albicans BU7098, K. pneumoniae ATCC 700603, vancomycin‐resistant Enterococcus spp., E. faecium ATCC 4602, E. coli ATCC 25922, P. aeruginosa ATCC 27853, methicillin‐sensitive S. aureus P1085, multidrug‐resistant A. baumannii P1415, carbapenem‐resistant E. cloacae P611, ertapenem‐resistant Salmonella spp. H18 and C. glabrata BU7170. Microbial inocula were prepared using standard microbiological procedures. Briefly, a pure colony of each microorganism was suspended in 5 mL of sterile saline, vortexed thoroughly, and adjusted to a turbidity equivalent to a 0.5 McFarland standard (≈ 1 × 10⁸ CFU/mL). The standardized suspensions were subsequently inoculated onto appropriate agar media and incubated at 37°C for 24 h prior to antimicrobial testing.

Detailed information on the origin, identification procedures, and documented antimicrobial resistance/susceptibility characteristics of the microbial strains is provided in Table S2.

5.6.2. Agar Disc Diffusion Method

The antimicrobial activity of E. ramosissimum essential oil and butanolic extract was evaluated using the agar disc diffusion method, in accordance with the guidelines of the Clinical and Laboratory Standards Institute (CLSI), document M02‐A12 [41]. Standardized microbial suspensions were prepared from fresh cultures and adjusted to a turbidity equivalent to a 0.5 McFarland standard.

Each standardized microbial suspension was evenly spread onto Mueller–Hinton agar plates using a sterile cotton swab. Sterile filter paper discs (6 mm diameter) were impregnated either with 10 µL of pure essential oil (density: 997.4 mg/mL) or with 10 µL of butanolic extract solution prepared at a concentration of 100 mg/mL. The impregnated discs were then placed onto the inoculated agar surfaces.

Gentamicin (15 µg/disc) was used as a positive control for antibacterial activity, while fluconazole (5 µg/disc) was used as a reference antifungal agent for Candida species, according to CLSI recommendations. Prior to incubation, the plates were kept at 4°C for 2 h in the dark to allow diffusion of samples into the agar medium without microbial growth. The plates were then incubated at 37°C for 24 h, after which antimicrobial activity was assessed by measuring the diameter of the inhibition zones (mm) surrounding each disc. The disk diffusion assay was used as a preliminary qualitative assessment of antimicrobial activity. Because agar diffusion may be influenced by the physicochemical properties and diffusion behavior of plant‐derived samples, broth microdilution was used as the quantitative method for determining MIC and MBC/MFC values for all tested microorganisms. Each assay was performed using three technical replicates within the same experimental run, and the results were expressed as mean ± SD.

5.6.3. Determination of the Minimum Inhibitory Concentration (MIC)

The minimum inhibitory concentration (MIC) of E. ramosissimum essential oil and butanolic extract was determined using the broth microdilution method, in accordance with the guidelines of the Clinical and Laboratory Standards Institute (CLSI) M07‐A10 for bacteria and M27‐A3 for yeasts [42, 43].

Briefly, the EO and BE were initially solubilized in dimethyl sulfoxide (DMSO) and subsequently diluted in the appropriate culture broth. No additional emulsifying agent was used for the EO. Ten microliters of each serial dilution were added to microplate wells containing 100 µL of microbial suspension. The inocula were obtained by diluting fresh cultures in Mueller–Hinton broth (MHB) for bacteria (5 × 10⁵ CFU/mL) and Sabouraud dextrose broth (SDB) for yeasts (1–2 × 10³ CFU/mL). The final DMSO concentration was 1% (v/v) in all test wells. Vehicle‐control wells containing 1% (v/v) DMSO and the corresponding microbial inoculum, but no EO or BE, were included under the same experimental conditions. No inhibition of microbial growth was observed in the vehicle controls. After gentle mixing, the microplates were incubated at 37°C for 24 h. The MIC was defined as the lowest concentration of the tested sample showing no visible microbial growth. Gentamicin and fluconazole were used as positive controls for bacteria and yeasts, respectively, according to CLSI recommendations [42, 43].

5.6.4. Determination of Minimum Bactericidal and Fungicidal Concentrations (MBC and MFC)

Following MIC determination, 100 µL aliquots from wells showing no visible microbial growth were aseptically subcultured onto Mueller–Hinton agar plates for bacteria and Sabouraud dextrose agar plates for yeasts. The plates were incubated at 37°C for 24 h [42, 43].

The MBC and MFC values were defined as the lowest concentrations of the tested samples at which no microbial growth was observed on agar plates, indicating a bactericidal or fungicidal effect.

For bacterial strains, the MBC/MIC ratio was calculated by dividing the MBC by the corresponding MIC, whereas for yeast strains, the MFC/MIC ratio was calculated by dividing the MFC by the corresponding MIC. Ratios ≤ 4 were interpreted as bactericidal or fungicidal, while ratios > 4 were interpreted as bacteriostatic or fungistatic.

5.7. Molecular Docking Study

Molecular docking simulations were performed to investigate potential binding interactions between the major compounds identified in the essential oil and the butanolic extract of Equisetum ramosissimum and selected microbial target proteins. For each extract, the three major compounds were selected for docking analysis.

The three‐dimensional structures of the target proteins were retrieved from the RCSB Protein Data Bank (PDB), including penicillin‐binding protein 2a (PBP2a, PDB ID: 3ZFZ), lanosterol 14α‐demethylase (CYP51, PDB ID: 5V5Z), DNA gyrase subunit B (GyrB, PDB ID: 5MMN), and UDP‐3‐O‐(R‐3‐hydroxymyristoyl)‐N‐acetylglucosamine deacetylase (LpxC, PDB ID: 3P3G). The corresponding co‐crystallized ligands, AI8 (3ZFZ), 1YN (5V5Z), O54 (5MMN), and 3P3 (3P3G), were used as reference ligands for validation of the docking protocol.

Protein preparation was carried out using PyMOL, where the co‐crystallized ligands, water molecules, and other non‐essential heteroatoms were removed, while essential components involved in protein activity were retained. Specifically, the catalytic Zn^{2 +} ion in LpxC (3P3G) and the heme prosthetic group containing Fe in CYP51 (5V5Z) were retained in their crystallographic positions during receptor preparation because they are integral components of the respective active sites. These cofactors were included as part of the rigid receptor structures during docking. The active sites were defined based on the binding positions of the original co‐crystallized ligands. Grid boxes were generated accordingly to cover the corresponding binding pockets. The grid‐center coordinates, grid dimensions, co‐crystallized ligand identifiers, and exhaustiveness values used for each target are provided in Table S1.

The three‐dimensional structures of the selected ligands were downloaded from PubChem and prepared prior to docking. Molecular docking simulations were performed using AutoDock Vina implemented in PyRx, following energy minimization and conversion of ligand and protein files into PDBQT format. No explicit pH‐dependent protonation‐state assignment was performed; the structures were prepared using the standard PyRx/AutoDockTools workflow prior to PDBQT conversion. Partial charges and hydrogen atoms were assigned during PDBQT preparation using the default settings of the docking workflow. The exhaustiveness parameter was set to 8 for all docking calculations.

Docking scores were reported in kcal/mol. For each ligand–target complex, nine docking poses were generated. The top‐ranked pose (pose 1), corresponding to the most favorable AutoDock Vina docking score, was selected for subsequent interaction analysis. The docking protocol was evaluated by redocking each co‐crystallized ligand into its corresponding binding site using the same grid‐box dimensions and docking parameters. The RMSD values calculated between the crystallographic and redocked poses are reported in Table 7. Visualization and analysis of protein–ligand interactions were performed using BIOVIA Discovery Studio Visualizer 2025.

5.8. Statistical Analysis

Statistical analysis was performed only for the antioxidant assays. For each assay, the results were expressed as mean ± standard deviation (SD) of three technical replicates (n = 3) obtained from the same sample preparation. Differences among the EO, BE, and reference antioxidants were analyzed separately for each assay using one‐way analysis of variance (ANOVA), followed by Tukey's post hoc multiple‐comparison test. Differences were considered statistically significant at p < 0.05.

Author Contributions

Saida Sissi: writing – original draft, data curation, formal analysis, methodology, investigation. Ilham Dilagui: investigation, visualization. Soukaina Ouhaddou: writing – original draft, investigation, methodology. Sara Bellioua: methodology, investigation. Nabila Soraa: project administration, supervision. Khalid Bekkouche: validation, writing – review and editing, supervision. Mustapha Larhsini: conceptualization, supervision, validation. Mohamed Markouk: methodology, validation, supervision, project administration.

Funding

This work was supported by a national grant from the Ministry of Higher Education, Scientific Research and Professional Training (Morocco) (ENSSUP).

Conflicts of Interest

The authors declare no conflicts of interest.

Declaration of Generative AI and AI‐Assisted Technologies in the Writing Process

During the preparation of this manuscript, the authors used OpenAI to assist with language editing, rephrasing, and improving the clarity of selected sections. The authors reviewed and edited all AI‐assisted content and take full responsibility for the accuracy, integrity, and final content of the manuscript. No generative AI tools were used to generate or analyze experimental data, results, or figures.

Supporting information

Supporting File: cbdv71853‐sup‐0001‐SuppMat.docx.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.