Antimicrobial, Herbicidal and pesticidal potential of Tunisian eucalyptus species: Chemoprofiling and biological evaluation

Abstract

The Eucalyptus genus, characterized by its imposing stature and fragrant foliage, has been a source of fascination for humanity over the centuries. The focus of the present investigation was directed towards the essentials oils (EOs) of five Eucalyptus trees cultivated in Tunisia. The GC-MS analysis unveiled unique compositional profiles, a finding substantiated by both Hierarchical Clustering Analysis (HCA) and Principal Component Analysis (PCA) conducted on the leaves EOs. These analyses resulted in the formation of discrete HCA clades, delineating 23 significant components. Notably, the percentage of eucalyptol emerged as the pivotal factor demarcating the separation between three distinct groups. The statistical analysis revealed a dose-dependent relationship in both phytotoxicity evaluation and antibacterial activity. The EOs from Eucalyptus loxophleba and E. salubris exhibited the highest phytotoxicity, inhibiting radical elongation and germination of various seeds, especially Sinapis arvensis and Raphanus sativus. The antimicrobial assessment demonstrated significant inhibitory effects of the EOs on bacterial strains, with MIC values spanning from 14 to exceeding 50 mg/ml. The EOs also affected biofilm formation and cellular metabolism, displaying varied efficacy among different Eucalyptus species against some bacterial strains. The EOs exhibited selective inhibition against acetylcholinesterase (AChE), butyrylcholinesterase (BChE), α-amylase, and α-glucosidase. E. campaspe EO showed the highest AChE activity, while E. loxophleba and E. salubris EOs were most potent toward α-amylase. E. loxophleba EO demonstrated notable activity against α-glucosidase. Overall, these findings provide important data about the diverse biological activities of Eucalyptus EOs, suggesting potential applications in agriculture, medicine, and pharmacy.

1. Introduction

The use of eucalyptus essential oils (EOs) is increasingly gaining prominence in research, sparking sustained interest due to the chemical richness and varied biological activities associated with these aromatic extracts [1,2]. Their compositions rich in monoterpenes and sesquiterpenes give these EOs a distinctive olfactory signature while offering a broad range of biological properties [3].

The interest in eucalyptus species stems from their chemical diversity, providing a myriad of industrial applications ranging from construction to paper production, honey harvesting, and rubber extraction [4]. However, recent attention has shifted towards exploring the biological potential of their EOs, paving the way for in-depth studies on their anti-inflammatory [5], antioxidant [6], antitumoral properties [7], among others.

Recently, several Eucalyptus species wereinvestigated for their potential phytotoxic potential against weed plants and crops: Danna and coworkers showed that E. gunnii EO inhibited germination of Portulaca oleracea, Raphanus sativus, and Solanum lycopersicum seeds and affect radical growth in S. lycopersicum and E. pulverulenta EO was active against germination of P. oleracea and R. sativus and radical elongation of Lepidium sativum and Lolium multiflorum [8]). Polito and colleagues demonstrated phytotoxicity of E. griffithsii, E. hemiphloia, E. lesouefii, E. longicornis, E. pyriformis, E. viminalis, and E. wandoo EOs against germination and radical elongation of R. sativus, L. multiflorum, and Sinapis arvensis seeds [9]. Instead Flores-Macías et al. studied phytotoxic potential of E. globulus essential oil against Avena fatua and Amaranthus hybridus two herbicide-resistant weeds [10]. Moreover, E. falcata, E. sideroxylon and E. citriodora EOs phytotoxicity was studied by Amri and coworkers against plants weed (S. arvensis, Phalaris canariensis) and durum wheat crop (Triticum durum) [11].

E. brevifolia, E. camaldulensis, E. extensa, E. globulus, E. lehmannii, E. leptophylla, E. patellaris, and E. woollsiana EOs were studied for their antidiabetic propriety evaluating their activity against amylase and glucosidase [[12], [13], [14]], but only in our previous study on EOs from E. gunni and E. pulverulenta, α-amylase activity was studied in relation to phytotoxic activity as enzyme involved in germination process [8].

Instead, for that concern activity against cholinesterases E. brevifolia, E. extensa, E. lehmannii, E. leptophylla, E. patellaris, and E. woollsiana EOs were studied as a possible coadjutant in treatment of Alzheimer disease [12]. Instead, only E. globulus and E. citriodora EOs were analyzed for their anti-cholinesterases activity related to their insecticide property.

Among the many intriguing properties of eucalyptus EOs, their ability to combat biofilms, notoriously resistant to antimicrobial agents, has emerged as a promising area of research [12,15]. Their chemical diversity provides an arsenal of substances that can play an important role in the fight against biofilms, opening innovative prospects in various sectors, including pharmacy, cosmetics, and agri-food.

In this context, the aim of this study was to thoroughly determine the chemical composition and enzymatic activity, phytotoxic effects, and antibacterial potential of the EOs extracted from some Tunisian Eucalyptus species, including E. amplifolia Naudin, E. bicolor A. Cunn. ex T. Mitch., E. campaspe S. Moore., E. loxophleba Benth., and E. salubris F. Muell. Phytotoxic activity was assessed on the germination and radical growth of seeds from Sinapis arvensis L., Lolium multiflorum Lam., Raphanus sativus L., and Cucumis sativus L. Moreover, the EOs were assayed for their activity against α-amylase and α-glucosidase, two key enzymes in germination process, and against cholinesterases, target of many pesticides. Lastly, the inhibitory potential in against the biofilm formation in both Gram-positive (Staphylococcus aureus subsp. aureus, Pseudomonas aeruginosa and Listeria monocytogenes) and Gram-negative bacteria (Acinetobacter baumannii and Escherichia coli) was evaluated.

2. Materials and methods

2.1. Chemical reagent

All chemical reagents used for assays were bought from Merck, Darmstadt, Germania.

2.2. Botanical resources

The plant leaves of E. amplifolia Naudin, E. bicolor A. Cunn. ex T. Mitch., E. campaspe S. Moore., E. loxophleba Benth., and E. salubris F. Muell. were collected from different arboretums as mentioned in Table 1. Plants were identified by Prof. Vincenzo De Feo. Voucher specimens (DF/725/2021 for E. amplifolia, DF/824/2021 for E. bicolor and DF/765/2021 for E. campaspe, DF/753/2021 for E. loxophleba, DF/814/2021 for E. salubris) were deposited in the Herbarium of the Pharmaceutical Botany Chair at the University of Salerno. To obtain representative samples, a selection of almost five individual trees were made for each species, and were amalgamated. Subsequently, the homogenized samples from diverse species were transferred to a greenhouse, where they underwent a controlled shade drying process until a uniform weight was attained.

Table 1.

Arboretum, Harvest period and bioclimatic condition.

	Arboretum(governorate)	Period of harvesting	Bioclimatic condition
E. loxophleba	Hajeb Layoun (Kairouan)	April 2021	Semi-arid upper (with moderate winters)
E. bicolor	Djebel Mansour (Zaghouan)	Mars 2021	semi-arid Upper and middle
E. amplifolia	Mrifeg (Bizerte)	May 2022	Upper humid
E. campaspe	Henchir El Naam (Siliana)	February 2021	Semi-arid Upper and middle
E. salubris	El Hnaya (Sousse)	May2022	semi-arid Lower

2.3. Isolation and analysis of eucalyptus essential oils

Three hundred grams of each Eucalyptus species were ground in a Waring blender and then subjected to hydrodistillation, utilizing a Clevenger apparatus for 3 h, according to the European Pharmacopoeia [16]. The resulting oils were collected, dehydrated with anhydrous sodium sulfate and preserved in amber vials at 4 °C until subsequent studies.

The EO yield (w/v, %) was calculated according to the following equation:

$$Yield(\%)=\frac{Wo*100}{V_{EO}}$$

where W0 is the dried plant weight distilled and V_EO is the EO volume obtained.

A PerkinElmer Sigma-115 gas chromatograph (PerkinElmer, Waltham, MA, USA) endowed with a flame ionization detector (FID) and a data handling processor was used for analytical gas chromatography. The separation was carried out using a HP-5 MS fused-silica capillary column (30 m, 0.25 mm i.d., 0.25 μm film thickness, Agilent, Roma, Italy). Column temperature: 40 °C, with 5 min initial hold, and then to 270 °C at 2 °C/min, 270 °C (20 min); injection mode, splitless (1 μL of a 1:1000 n-hexane solution); injector and detector temperatures 250 °C and 290 °C, respectively. The analysis was also conducted using a fused silica HP Innowax polyethylene glycol capillary column (50 m, 0.20 mm i.d., 0.25 μm film thickness, Agilent, Roma, Italy). Helium was the carrier gas (1.0 ml/min).

GC/MS analyses were achieved on an Agilent 6850 Ser. II apparatus (Agilent, Roma, Italy), endowed with a fused silica DB-5 capillary column (30 m, 0.25 mm i.d., 0.33 m film thickness, Agilent, Roma, Italy), coupled to an Agilent Mass Selective Detector MSD 5973; ionization energy voltage 70 eV; electron multiplier voltage energy 2000 V. Column temperature: 40 °C, with 5 min initial hold, and then to 270 °C at 2 °C/min, 270 °C (20 min); injection mode, splitless (1 μL of a 1:1000 n-hexane solution); injector and detector temperatures 250 °C and 290 °C, respectively. Mass spectra (MS) were scanned in the range 40–500 amu, scan time 5 scans/s.

Many constituents were identified by GC by comparing their Kovats retention indices (Ri), which were calculated relative to the retention times (tR) of n-alkanes (C₁₀–C₃₅), to either those in the literature [[17], [18], [19]] and mass spectra on both columns, or those of single compounds available in our laboratories via the NIST 17 and Wiley libraries [20]. Peak area normalization was used to determine the component relative concentrations. No response factors were calculated.

2.4. Phytotoxic activity

Phytotoxic activity was investigated concerning the germination and radical growth of Lolium multiflorum Lam., Sinapis arvensis L., Raphanus sativus L., and Cucumis sativus L. seeds selected for their known and reliable germination characteristics.

S. arvensis seeds were collected from wild populations in Sidi Ismail, Beja, Tunisia on June 2021. L. multiflorum seeds were bought from Fratelli Ingegnoli s.p.a. (Milano, Italy), whereas R. sativus and C. sativus seeds were purchased from Blumen group s.r.l. (Bologna, Italy).

The seeds were sterilized in 95 % ethanol for 15 s and then placed in Petri dishes (Ø = 90 mm) lined with three layers of Whatman filter paper.

Following sterilization, the Petri dishes received 7 ml treatments: double distilled water (utilized as control to verify seed viability), a water–acetone mixture (99.5:0.5, v/v) as the other control (due to essential oils being dissolved in this solution owing to their lipophilicity), or essential oils at concentrations of 1000, 500, 250, and 100 μg/ml.

Control experiments exclusively utilizing water–acetone exhibited no notable distinctions compared to the water control.

Germination conditions were maintained at 20 ± 1 °C, with a natural photoperiod. Seed germination was monitored in Petri dishes every 24 h, considering a seed germinated when root became evident [21]. After 120 h for Raphanus sativus and Sinapis arvensis seeds, and after 168 h for Lolium multiflorum and Cucumis sativus seeds, radicle growth effects were measured in centimeters. Each experiment was replicated three times, employing Petri dishes containing 10 seeds each.

2.5. Anti-enzymatic activities

2.5.1. Cholinesterases inhibition

The cholinesterase activity was determined by Ellman's assay [22] with some modifications. Briefly, in a total of 1 ml, 415 μL of Tris-HCl buffer 0.1 M (pH 8), 10 μL of a buffer solution of EOs (in methanol) at different concentrations (100, 10, 1, and 0.1 mg/ml, and 25 μL of a solution containing 0.28 U/mL of AChE (or BChE) were incubated for 15 min at 37 °C. Then, a solution of acetylthiocholine iodide (AChI) or butyrylthiocholine iodide.

(BChI) 1.83 mM (75 μL) and 475 μL of 5,5 -dithiobis (2-nitrobenzoic acid) (DTNB) was added, and the final mixture was placed for 30 min at 37 °C. The absorbance was read at 405 nm in a spectrophotometer (Thermo Scientific Multiskan GO, Monza, Italy). Galantamine was used as the positive control.

2.5.2. α-Amylase inhibition assay

Amylase activity was carried out using a modified version of the Bernfeld method [23] Essential oil (EO) at varying concentrations (100 μL) was mixed with amylase (10 U/mL) and sodium phosphate buffer (pH = 6.9). Then, the solutions were placed at 37 °C for 10 min at the end soluble starch solution (1 %) was added, and placed for additional 20 min at 37 °C. The reaction was blocked with 3,5-dinitrosalicyclic acid (DNSA) solution (96 mM), and boiled at 100 °C for 10 min and then cooling with 600 μL of distilled water. The absorbance was measured at 540 nm using a UV Spectrophotometer (Thermo Fischer Scientific, Vantaa, Finland).

3. 3α-glucosidase inhibition assay

The evaluation of α-glucosidase inhibitory activity was carried out according to a previously reported method [24] with some modifications. In brief, a 96-multiwell plate was utilized, where 0.1 M phosphate buffer at pH 7.0 (150 μL) was introduced into each well. Subsequently, 10 μL of EOs, dissolved in MeOH to achieve various concentrations, were sequentially added.

The reaction started with 15 μL of α-glucosidase (1 U/mL) in each well, followed by placement at 37 °C. After 5 min, 75 μL of 4-nitrophenyl α-d-glucopyranoside substrate (2.0 mM) was added, and the plate was incubated for an additional 10 min at 37 °C. Absorbance was read at 405 nm using a UV Spectrophotometer (Thermo Fischer Scientific, Vantaa, Finland).

Acarbose served as the positive control, and the negative control recorded absorbance using phosphate buffer instead of the sample. Enzyme inhibition was calculated, with results expressed as IC₅₀ values. The experiments were triplicated. The percentage of enzyme activity inhibition for cholinesterases, α-amylase, and α-glucosidase was determined using the formula:

$$\%=\frac{A_0-A_1}{A_0}*100$$

A₀ is the absorbance of the control without the sample, and A₁ is the absorbance of the sample. The concentration providing 50 % inhibition (IC₅₀) was obtained by plotting the inhibition percentage against sample concentrations.

3.1. Antimicrobial activity

3.1.1. Microbial strains and cultivations conditions

The cultures of bacterial strains were obtained from the Leibniz Institute DSMZ (Germany), and were Acinetobacter baumannii ATCC 19606 and Escherichia coli DSM 8579, as Gram-negative strains and Staphylococcus aureus subsp. aureus Rosebach ATCC 25923, Pseudomonas aeruginosa DSM 50071 and Listeria monocytogenes ATCC 7644 as Gram-positive strains. The cultivation was performed in Luria broth at 37 °C for 18 h, except for A. baumannii, which was cultured at 35 °C.

3.1.2. Minimum inhibitory concentration (MIC)

To maintain sterility, EOs and dimethyl sulfoxide (DMSO) were subjected to ultrafiltration before being included in the study Stock of EOs, were dissolved in sterile DMSO to achieve a concentration of 500 mg/ml. MIC determination, performed in 96-multiwell microplates, employed a modified resazurin method [25]. Final serially descending concentration of EOs (diluted in DMSO) used in this study were 50.00 (dilution factor: 10), 25.00, (dilution factor: 20) and 12.50 mg/ml (dilution factor: 40), which were used for each strain. A resazurin solution (270 mg in 40 ml sterilized deionized water) was prepared. Thirty-five μL of 3.3 × strength iso-sensitized broth, 10 μL of bacterial suspension (1.5 × 10⁷ CFU/ml), 10 μL of resazurin solution, and diluted samples of EOs were supplemented, to reach the specific EOs concentration and a final volume/well of 250 μL with different volumes of sterile Luria Bertani broth. The initial row received iso-sensitized broth, bacterial suspension, resazurin solution, and DMSO to reach a final volume/well of 250 μL with sterile Luria Bertani broth. The plates were subsequently sealed. Incubation at 37 °C (35 °C for A. baumannii) for 24 h preceded visual observation for color changes. MIC was the lowest EO concentration preventing a color shift from dark purple to pink.

3.1.3. Biofilm inhibitory activity

To evaluate the inhibitory impact of essential oils on mature biofilm, 96-well microtiter plates with flat bottoms were employed. Bacterial cultures adjusted to a 0.5 McFarland standard [26] were dispensed into each well (10 μL), incubated for 24 h at 37 °C (35 °C for A. baumannii), and subsequently exposed to 5–10 or 20 μl/mL of EOs in a final volume of 250 μL Luria-Bertani broth. Following an additional 24-h incubation, planktonic cells were eliminated, sessile cells were fixed, stained with crystal violet, and the absorbance at λ = 540 nm was read using a Cary Varian spectrophotometer. Biofilm inhibitory activity was determined as a percentage relative to the control. Triplicate experiments were performed for reliability [27].

3.1.4. Metabolic activity evaluation using MTT colorimetric method

The metabolic activity of bacterial cells within the EOs-influenced biofilm was assessed utilizing the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) colorimetric method [27].

After a 24-h bacterial incubation with removal of planktonic cells, two EO concentration ranges, namely (5 and 10 μl/mL) and (10 and 20 μl/mL), were introduced. Following an additional 24-h incubation, planktonic cells were eliminated, and a solution of 150 μL phosphate-buffered saline (PBS) and 30 μL 0.3 % MTT was added. Microplates were placed for 2 h at 37 °C (35 °C for A. baumannii). Subsequently, the MTT solution was eliminated, and two washing steps with 200 μL sterile physiological solution were carried out. Finally, 200 μL of dimethyl sulfoxide (DMSO) were added to suspend the formazan crystals, and absorbance was read at λ = 570 nm using a Cary Varian spectrophotometer.

3.2. Statistical analysis

Experiments were triplicated. Phytotoxic assay data were analyzed using GraphPad Prism 6.0 (two-way ANOVA, Dunnett's multiple comparisons test, p < 0.05). Anti-enzymatic and antibacterial activities were assessed using SPSS 26 (one-way ANOVA, Tukey's post-hoc test, p < 0.05).

4. Results

4.1. Yield and chemical composition of essential oils

The EOs yields from the leaves of different Eucalyptus species are presented in Table 1, ranging from 1.47 % to 4.5 %. The order of representation, from highest to lowest, is as follows: E. salubris (4.51 %) > E. loxophleba (3.14 %) > E. campaspe (2.8 %) > E. amplifolia (1.63 %) > E. bicolor (1.47 %).

The exploration of essential oils (EOs) unveiled a varied composition among different Eucalyptus species (Table 2).

Table 2.

Percent composition of the EOs.

	E. amplifolia	E. bicolor	E. campaspe	E. loxophleba	E. salubris	KIa	KIb	Identificationc
1-Pentanol,2-ethyl-	–	–	–	0.1	–	830		1, 2
4-Methyl-1,6-heptadien-4-ol	–	14.1	–	–	–	832		1, 2
α-Pinene	4.3	6.4	7.0	2.9	1.8	857	1012	1, 2, 3
Octane,4,5-diethyl-	–	0.2	–	–	–	859		1, 2
Camphene	0.4	0.2	0.3	0.2	–	868	1075	1, 2, 3
Thuja-2,4(10)-diene	0.1	–	–	–	–	875	1122	1, 2
β-Pinene	–	0.2	0.2	4.8	–	890	1120	1, 2
Myrcene	–	0.1	–	–	–	893	1166	1, 2
α-Phellandrene	–	0.5	0.1	0.4	0.1	918	1177	1, 2, 3
α-Terpinene	–	0.1	–	0.3	0.5	930	1170	1, 2, 3
p-Cymene	–	–	–	2.8	6.0	938	1250	1, 2
Eucalyptol	68.5	32.4	72.1	43.3	70.8	943	1210	1, 2, 3
1,3,8-p-Menthatriene	3.3	–	–	0.4	4.8	948	1411	1, 2
γ-Terpinene	–	0.1	0.1	0.8	–	971	1221	1, 2, 3
Fenchone	–	–	0.1	–	–	994	1340	1, 2
p-Mentha-2,4(8)-diene	–	–	–	0.3	–	996	1639	1, 2
6-Camphenone	–	0.1	0.1	–	–	998		1, 2
p-Cymenene	–	–	–	0.6	–	999	1438	1, 2
Linalool	–	–	–	0.4	–	1009	1506	1, 2, 3
3-methylbutyl 3-methylbutanoate	–	0.2	–	0.7	–	1014		1, 2
Thujone	–	–	–	0.6	–	1018	1430	1, 2, 3
exo-Fenchol	0.7	0.3	1.0	–	0.4	1019	1570	1, 2, 3
trans-p-Ment-2-en-1-ol	–	–	–	0.5	–	1025		1, 2
6-Campholenal	–	0.1	0.2	0.9	–	1028	1496	1, 2
Isogeraniol	–	–	0.1	–	–	1036		1, 2
trans-Pinocarveol	1.1	2.3	2.9	5.4	2.3	1040	1664	1, 2
Camphor	–	0.2	0.3	–	–	1044	1519	1, 2, 3
neo-Isopulegol	–	0.2	0.3	–	–	1046		1, 2
Sabina ketone	–	–	–	1.0	–	1052	1651	1, 2
Pinocarvone	0.2	0.6	0.7	1.3	–	1065	1576	1, 2
Borneol	2.2	0.4	2.1	0.7	0.7	1067	1715	1, 2, 3
Terpinen-4-ol	–	0.3	0.6	3.7	–	1078	1636	1, 2, 3
cis-Sabinene hydrate	–	–	–	–	2.5	1083	1460	1, 2
Cryptone	–	–	0.3	7.2	–	1085	1659	1, 2
trans-p-Mentha-1(7),8-dien-2-ol	–	0.5	0.1	–	–	1088	1810	1, 2
α-Terpineol	0.5	1.0	3.3	2.2	–	1090	1662	1, 2, 3
p-Cymen-8-ol	–	–	–	0.9	–	1092	1828	1, 2
Safranal	–	–	–	1.2	–	1096	1648	1, 2
Myrtenol	–	–	–	1.1	–	1098	1791	1, 2
Verbenone	–	–	0.1	0.3	–	1100	1720	1, 2
Iso-Dihydro carveol	–	–	0.2	0.3	–	1105		1, 2
p-Cumic aldehyde	–	–	–	5.1	–	1133	1753	1, 2
Pulegone	–	–	0.6	1.6	–	1144	1662	1, 2
Phellandral	–	–	–	2.1	–	1164	1720	1, 2
p-Cymen-7-ol	–	–	–	2.4	–	1195	2113	1, 2
δ-Elemene	–	0.1	0.1	–	–	1208	1479	1, 2
Benzyl butanoate	–	0.2	0.5	–	–	1225		1, 2
α-Terpinyl acetate	–	1.0	–	–	–	1232	1685	1, 2
Isoledene	–	0.1	–	–	–	1251		1, 2
α-Copaene	0.1	0.2	–	–	–	1262	1477	1, 2
β-Patchoulene	0.1	–	–	–	–	1271		1, 2
α-Panasinsene	0.5	0.1	–	–	–	1272		1, 2
β-Longipinene	–	–	–	–	0.1	1276		1, 2
Longifolene	–	–	–	–	0.2	1283	1575	1, 2
trans-Caryophyllene	–	0.3	–	–	0.2	1284	1617	1, 2
α-Gurjunene	0.2	0.2	–	–	–	1288	1529	1, 2
β-Gurjunene	–	0.7	–	–	0.1	1304	1597	1, 2
α-Guaiene	0.4	–	0.2	–	–	1307	1600	1, 2
Aromadendrene	–	10.6	–	–	1.7	1308	1631	1, 2
cis-Muurola-3,5-diene	0.2	–	–	–	0.4	1311		1, 2
allo-Aromadendrene	7.7	–	0.1	–	0.4	1312	1660	1, 2
γ-Muurolene	–	–	–	–	0.3	1337	1725	1, 2
γ-Himachalene	–	–	–	–	0.2	1339	1709	1, 2
cis-β-Guaiene	0.6	–	–	–	–	1349	1664	1, 2
Viridiflorene	2.6	–	–	–	1.2	1350	1696	1, 2
9-epi-β-Caryophyllene	–	2.0	0.1	–	–	1352		1, 2
dehydro-Aromadendrene	–	0.1	–	–	–	1366		1, 2
trans-β-Guaiene	3.3	–	–	–	–	1371	1532	1, 2
γ-Cadinene	–	–	–	–	0.2	1384	1763	1, 2
7-epi-α-Selinene	0.3	–	–	–	–	1389		1, 2
Cadina-3,9-diene	–	0.1	–	–	–	1391		1, 2
δ-Cadinene	–	–	–	–	1.2	1397	1756	1, 2
trans-Cadina-1(6),4-diene	–	0.1	–	–	–	1400		1, 2
γ-Gurjunene	–	0.1	–	–	–	1410	1668	1, 2
Viridiflorene	–	–	–	0.1	–	1414	1696	1, 2
α-Amorphene	–	1.9	0.1	–	–	1419	1693	1, 2
trans-β-Guaiene	–	0.8	0.2	–	–	1425	1651	1, 2
Germacrene B	–	1.1	0.7	0.3	–	1437	1795	1, 2
Caryophyllene oxide	–	–	–	1.0	–	1440	2000	1, 2
Epiglobulol	–	5.6	1.4	–	–	1444	2100	1, 2
α-Cadinene	–	1.6	0.2	–	–	1450	1769	1, 2
Rosifoliol	–	1.3	0.2	–	–	1461		1, 2
Khusimone	–	0.4	–	–	–	1474		1, 2
γ-Eudesmol	–	0.8	–	–	–	1477	2176	1, 2
5-epi-7-epi-α-Eudesmol	–	–	0.2	–	–	1480		1, 2
Hinesol	–	0.9	–	–	–	1482	2228	1, 2
β-Eudesmol	–	2.5	0.4	–	–	1485	2248	1, 2
α-Eudesmol	–	3.0	0.9	–	–	1489	2247	1, 2
Total	97.3	96.3	98.1	97.9	96.1
Monoterpene hydrocarbons	8.1	7.6	7.7	13.5	13.2
Oxygenated monoterpenes	73.2	39.4	85.1	82.2	76.7
Sesquiterpene hydrocarbons	16.0	20.1	1.7	0.4	6.2
Oxygenated sesquiterpenes	–	14.5	3.1	1.0	–
Others	–	14.7	0.5	0.8	–

^{a, b}: Calculated Kovats retention indices on apolar HP-5 MS and polar HP Innowax capillary columns, respectively; c: identification method—1 = comparison of Kovats retention indices with published data, 2 = comparison of mass spectra with NIST 17 and Wiley 275 libraries and published data, 3 = coinjection with authentic compounds; - = absence.

A representative GC profile of E. bicolor was present in Fig. 2. Chemical structure of identified compounds was reported in Fig. 3.

Fig. 2.

A representative gas chromatography (GC) profile of E. bicolor.

Fig. 3.

Chemical structure of identified compounds.

The analysis disclosed 21 components in E. amplifolia Naudin EO (97.3 % of the total), 39 in E. bicolor EO (96.3 %), 37 in E. campaspe EO (98,1 %), 36 in E. loxophleba EO (97,9 %) and 22 in E. salubris EO (96.1 %). Oxygenated monoterpenes predominated in all EOs: 73.2 % for E. amplifolia, 39.4 % for E. bicolor, 85.1 % for E. campaspe, 82.2 % for E. loxophleba and 76.7 % for E. salubris. The EO of E. amplifolia also contained hydrocarbon sesquiterpenes (16.0 %) and hydrocarbon monoterpenes (8.1 %). Eucalyptol (68.5 %) was the main compound while the other main components are allo-aromadendrene (7.7 %), α-pinene (4.3 %), 1,3,8-p-menthatriene, trans-β-guaiene (3.3 %), viridiflorene (2.6 %), borneol (2.2 %) and trans-pinocarveol (1.1 %). The EO of E. bicolor also contained sesquiterpene hydrocarbons (20.1 %), oxygenated sesquiterpenes (14.5 %), monoterpenes hydrocarbons (7.6 %) and other compounds (14.7 %). Eucalyptol (32.4 %) was the main compound while the other main components were 4-methyl-1,6-heptadien-4-ol (14.1 %), aromadendrene (10.6 %), α-pinene (6.4 %), epiglobulol (5.6 %), α-eudesmol (3.0 %), β-eudesmol (2.5 %), trans-pinocarveol (2.3 %), 9-epi-β-caryophyllene (2.0 %), α-amorphene (1.9 %), α-cadinene (1.6 %), rosifoliol (1.3 %), germacrene B (1.1 %), α-terpineol and α-terpinyl acetate (both 1.0 %). The EO of E. campaspe also contained monoterpene hydrocarbons (7.7 %), oxygenated sesquiterpenes (3.1 %), sesquiterpene hydrocarbons (1.7 %) and other compounds (0.5 %). Eucalyptol (72.1 %) was the main compound while the other main components were α-pinene (7.0 %), α-terpineol (3.3 %), trans-pinocarveol (2.9 %), borneol (2.1 %), epiglobulol (1.4 %) and exo-fenchol (1.0 %). The EO from E. loxophleba also contained monoterpene hydrocarbons (13.5 %), oxygenated sesquiterpenes (1.0 %), sesquiterpene hydrocarbons (0.4 %) and other compounds (0.8 %). The major component was eucalyptol (43.3 %) while the other main components were cryptone (7.2 %), trans-pinocarveol (5.4 %), p-cumic aldehyde (5.1 %), β-pinene (4.8 %), terpinen-4-ol (3.7 %), α-pinene (2.9 %), p-cymene (2.8 %), p-cymen-7-ol (2.4 %), α-terpineol (2.2 %), phellandral (2.1 %), pulegone (1.6 %), pinocarvone (1.3 %), safranal (1.2 %), myrtenol (1.1 %), sabina ketone (1.0 %) and caryophyllene oxide (1.0 %). The EO from E. salubris also contained monoterpene hydrocarbons (13.2 %) and sesquiterpene hydrocarbons (6.2 %). The major component wais eucalyptol (70.8 %) while the other main components were p-cymene (6.0 %), 1,3,8-p-menthatriene (4.8 %), cis-sabinenehydrate (2.5 %), trans-pinocarveol (2.3 %), α-pinene (1.8 %), aromadendrene (1.7 %), viridiflorene and δ-cadinene (1.2 %).

Principal Component Analysis (PCA) (Fig. 4) was applied on 23 major compounds of the chemical composition from tested species in order to process the structural and compositional descriptors of the samples.

Fig. 4.

Principal Component Analysis (PCA) results carried out with chemical composition. Projection of 23 (>1 %) major compounds of eucalyptus species in the plane formed by the two main components. Abbreviations of compound β-Pinene (β-Pin), α-pinene (α-Pin), trans-Pinocarveol (trans-Pino), Terpinen-4-ol (Terpin-4-ol), p-Cumic aldehyde (p-Cum aldehy), Viridiflorene (Viridif), p-Cymen-7-ol (p-Cyme-7-ol), Cryptone (Crypt), eucalyptol (Eucaly), β-Eudesmol (β-Eud), α-Eudesmol (α-Eud), Aromadendrene (Aromade), Epiglobulol (Epiglobul), trans-β-Guaiene (trans-β-Guai), allo-Aromadendrene (allo-Aromaden), Phellandral (Phellan), 4-Methyl-1,6-heptadien-4-ol (4-Methyl), p-Cymene (p-Cym), 1,3,8-p-Menthatriene (Menthat), Borneol (Bor), cis-Sabinenehydrate (cis-Sabinenehy), α-Terpineol (α-Terpi).

The application of Principal Component Analysis (PCA) (Fig. 1) represent the horizontal component (F1) explained 41.21 % of the variance and the second (F2) 33.14 %, with β-Pinene (β-Pin), trans-Pinocarveol (trans-Pino), Terpinen-4-ol (Terpin-4-ol), p-Cumic aldehyde (p-Cum aldehy), Viridiflorene (Viridif), p-Cymen-7-ol (p-Cyme-7-ol), Cryptone (Crypt) being the variables with the greatest weight in horizontal component F1 and eucalyptol (Eucaly); α-pinene (α-Pin); β-Eudesmol(β-Eud); α-Eudesmol (α-Eud); Aromadendrene (Aromade); Epiglobulol (Epiglobul) being those in vertical component 2.

Fig. 1.

Schematic diagram of methodology used.

Concurrently, Hierarchical Cluster Analysis (HCA) (Fig. 5), utilizing Pearson correlation coefficients (r), delineated three distinct groups denoted as Group A, B, and C.

Fig. 5.

Hierarchical cluster analysis (HCA) of the EOs.

Group A (r = −0.332), predominantly comprising E. loxophleba, emerged prominently, establishing a distinctive dichotomy in the HCA. This group was characterized by elevated levels of cryptone and β-pinene, accompanied by a relatively low amount of eucalyptol. Simultaneously, this EO shares the presence of p-cymene with E. salubris EOs.

Group B (r = 0.567), consisting of E. salubris and E. amplifolia, was noteworthy for its highest concentration of eucalyptol.

Group C (r = 0.069) comprised E. bicolor and E. campaspae, distinguished by elevated levels of epiglobulol and aromadendrene. Intriguingly, E. campaspae exhibitrf a positive correlation with the second group, potentially attributed to the shared high level of eucalyptol.

An evident differentiation among the five eucalyptus species is discernible through the utilization of two statistical methodologies (PCA and HCA analysis). Consequently, it is deduced that these species exhibit distinctive chemical compositions, suggesting potential variations in their biological effects during subsequent testing procedures.

4.2. Phytotoxic activity

This research assessed the phytotoxic activities of the Eucalyptus EOs on the germination and radical growth of seeds from L. multiflorum, S. arvensis, R. sativus and C. sativus.

Table 3, Table 4 represented the radicle length in cm and the number of germinated seeds after treatment with the EOs.

Table 3.

Phytotoxic activity of the EOs against germination of L. multiflorum, S. arvensis, R. sativus, and C. sativus.

Germination
	L. multiflorum	S. arvensis	R. sativus	C. sativus
E. amplifolia
CTR	9.7 ± 0.6	10.0 ± 0.0	10.0 ± 0.0	10.0 ± 0.0
CTR + C3H6O	9.3 ± 0.6	10.0 ± 0.0	10.0 ± 0.0	10.0 ± 0.0
1000	1.0 ± 0.0d	0.0 ± 0.0d	3.3 ± 0.3d	5.3 ± 0.3d
500	2.0 ± 0.0d	0.0 ± 0.0d	6.0 ± 0.0d	7.0 ± 0.0d
250	3.0 ± 0.0d	2.0 ± 1.0d	7.7 ± 0.3d	7.7 ± 0.3d
125	6.3 ± 0.6d	5.7 ± 0.6d	9.0 ± 0.0a	9.0 ± 0.0a
E. bicolor
CTR	8.6 ± 0.6	9.7 ± 0.0	9.7 ± 0.3	10 ± 0.00
CTR + C3H6O	8.0 ± 0.0	9.6 ± 0.0	9 ± 0	10 ± 0.00
1000	1.3 ± 1.5b	0.0 ± 0.0d	2 ± 0d	5.3 ± 0.3d
500	5.3 ± 3.2	3.7 ± 0.9d	2.0 ± 0.6d	7.0 ± 0.0d
250	6.0 ± 2.0	6.0 ± 1.0b	3.3 ± 0.9d	8.0 ± 0.0d
125	9.0 ± 1.7	7.0 ± 0.0a	4.7 ± 0.3d	9.0 ± 0.0b
E. campaspe
CTR	8.7 ± 0.6	9.3 ± 1.5	8.3 ± 1.5	9.3 ± 0.6
CTR + C3H6O	7.3 ± 1.1	9.7 ± 0.6	5.7 ± 1.8	8.7 ± 0.6
1000	6.7 ± 0.6	0.7 ± 0.6d	0.3 ± 0.6d	5.0 ± 3.0b
500	7.7 ± 1.5	1.7 ± 1.1d	0.3 ± 0.6d	9.0 ± 0.0
250	8.7 ± 0.6	5.7 ± 2.3a	2.7 ± 0.6c	9.3 ± 0.6
125	8.7 ± 1.1	9.7 ± 0.6	3.0 ± 1.0c	10.0 ± 0.0
E. loxophleba
CTR	7.0 ± 2.0	9.3 ± 1.1	7.3 ± 1.5	9.0 ± 1.0
CTR + C3H6O	7.3 ± 0.6	10.0 ± 0.0	5.3 ± 1.5	9.3 ± 0.6
1000	0.0 ± 0.0b	0.0 ± 0.0d	0.0 ± 0.0d	8.7 ± 1.5
500	3.3 ± 2.5	0.0 ± 0.0d	0.0 ± 0.0d	8.5 ± 0.7
250	4.0 ± 1.7	0.0 ± 0.0d	0.0 ± 0.0d	9.3 ± 1.1
125	6.7 ± 2.5	1.3 ± 2.3d	0.3 ± 0.6d	9.0 ± 1.0
E. salubris
CTR	10.0 ± 0.0	10.0 ± 0.0	10.0 ± 0.0	10.0 ± 0.0
CTR + C3H6O	10.0 ± 0.0	9.7 ± 0.3	10.0 ± 0.0	10.0 ± 0.0
1000	5.7 ± 0.3d	0.0 ± 0.0d	0.0 ± 0.0d	6.0 ± 0.0d
500	6.3 ± 0.7d	0.0 ± 0.0d	0.0 ± 0.0d	6.7 ± 0.3d
250	7.7 ± 0.3b	0.0 ± 0.0d	6.3 ± 0.3d	8.0 ± 0.0d
125	8.0 ± 0.00b	8.3 ± 0.7b	8.0 ± 0.0d	8.3 ± 0.3c

Results, mean ± SD of three experiments.

^ap < 0.05.

^bp < 0.01.

^cp < 0.001.

^dp < 0.0001 vs. control (ANOVA, Dunnett's test).CTR: deionized water; CTR + C₃H₆O: water–acetone mixture (99.5:0.5, v/v).

Table 4.

Phytotoxic effects of the tested essential oils against radicle elongation of L. multiflorum, S. arvensis, R. sativum, and C. sativus.

Radical elongation (cm)	L. multiflorum	S. arvensis	R. sativus	C. sativus
E. amplifolia
CTR	2.7 ± 0.3	2.1 ± 0.1	2.4 ± 0.3	2.6 ± 0.1
CTR + C3H6O	2.7 ± 0.2	2.0 ± 0.1	2.3 ± 0.2	2.5 ± 0.0
1000	0.1 ± 0.0d	0.0 ± 0.0d	0.5 ± 0.7d	0.8 ± 0.0d
500	0.3 ± 0.0d	0.0 ± 0.0d	1.3 ± 0.1b	1.5 ± 0.1d
250	0.6 ± 0.0d	0.3 ± 0.0d	1.6 ± 0.0b	1.7 ± 0.1d
125	1.8 ± 0.1a	0.3 ± 0.1d	2.8 ± 0.1	2.1 ± 0.1b
E. bicolor
CTR	4.5 ± 2.1	2.1 ± 0.1	2.3 ± 0.2	2.8 ± 0.1
CTR + C3H6O	4.0 ± 1.8	2.4 ± 0.2	2.1 ± 0.0	2.7 ± 0.1
1000	0.0 ± 0.0b	0.1 ± 0.0d	0.0 ± 0.0d	0.5 ± 0.0d
500	0.9 ± 0.3a	0.2 ± 0.0d	0.3 ± 0.1d	1.8 ± 0.1d
250	1.6 ± 0.7a	0.3 ± 0.1d	0.8 ± 0.2d	1.6 ± 0.1d
125	2.2 ± 0.8	0.8 ± 0.0d	0.9 ± 0.1d	2.1 ± 0.0d
E. campaspe
CTR	5.6 ± 1.1	2.0 ± 1.3	2.0 ± 0.7	3.1 ± 0.9
CTR + C3H6O	5.5 ± 1.1	0.8 ± 0.5	1.3 ± 0.3	2.9 ± 1.0
1000	1.0 ± 0.6b	0.2 ± 0.0a	0.0 ± 0.0c	0.7 ± 0.5a
500	1.9 ± 0.5b	0.3 ± 0.1a	0.0 ± 0.0c	2.3 ± 0.9
250	2.6 ± 0.8a	0.3 ± 0.1a	0.8 ± 0.3a	3.1 ± 0.5
125	2.5 ± 0.9a	0.6 ± 0.3a	1.2 ± 0.5	3.8 ± 0.9
E. loxophleba
CTR	5.7 ± 1.4	2.5 ± 1.1	2.9 ± 0.8	3.6 ± 0.7
CTR + C3H6O	1.7 ± 0.7	1.0 ± 0.3	1.3 ± 0.7	1.5 ± 0.5
1000	0.0 ± 0.0d	0.0 ± 0.0d	0.0 ± 0.0d	0.2 ± 0.1d
500	0.3 ± 0.1d	0.0 ± 0.0d	0.0 ± 0.0d	0.8 ± 0.4d
250	0.3 ± 0.2d	0.0 ± 0.0d	0.0 ± 0.0d	0.8 ± 0.3d
125	1.4 ± 0.6d	0.1 ± 0.0d	0.4 ± 0.0d	1.5 ± 0.3d
E. salubris
CTR	2.7 ± 0.1	2.2 ± 0.1	2.7 ± 0.1	2.9 ± 0.1
CTR + C3H6O	2.6 ± 0.1	2.2 ± 0.1	2.6 ± 0.1	2.9 ± 0.2
1000	0.7 ± 0.1d	0.0 ± 0.0d	0.0 ± 0.0d	0.9 ± 0.1d
500	1.7 ± 0.3b	0.0 ± 0.0d	0.0 ± 0.0d	1.6 ± 0.5b
250	2.1 ± 0.2	0.0 ± 0.0d	0.9 ± 0.1d	1.5 ± 0.1b
125	2.2 ± 0.1	1.4 ± 0.2b	1.8 ± 0.1d	1.4 ± 0.1b

Results, mean ± SD of three experiments.

^ap < 0.05.

^bp < 0.01.

^cp < 0.001.

^dp < 0.0001 vs. control (ANOVA, Dunnett's test). CTR: deionized water; CTR + C₃H₆O: water–acetone mixture (99.5:0.5, v/v).

The finding indicated that the E. amplifolia EO effectively hindered the radical elongation of all tested seeds at concentration of 1000,500 and 250 μg/ml. Conversely, at the lowest concentration evaluated (125 μg/ml), this EO primarily impeded the growth of S. arvensis seeds, and was not active against radical growth of R. sativus seeds. Moreover, this EO was active in the same way against germination of all studied seeds at all concentrations used, except against the crops R. sativus and C. sativus: in fact, for these seeds fact data highlighted the lowest activity.

E. bicolor EO inhibited radical elongation and germination of S. arvensis, R. sativus and C. sativus seeds at all concentrations tested, instead the same EO was phytotoxic in different way for L. multiflorum. In fact, at concentrations of 1000, 500 and 250 μg/ml inhibited radical elongation, while only at the higher concentration (1000 μg/ml) inhibited seed germination.

The EO from E. campaspe exhibited activity in inhibiting the radical elongation of both L. multiflorum and S. arvensis at all concentrations tested. Moreover, this EO inhibited radical elongation of R. sativus seeds at concentrations of 1000,500 and 250 μg/ml, while was active against radical growth of C. sativus seeds only at the higher concentration tested. This EO hindered the germination of R. sativus seeds across all concentrations tested, while it did not exhibit activity against the germination of L. multiflorum.

E. loxophleba EO was the most phytotoxic essential oil, inhibiting radical elongation of all seeds studied at all concentrations tested. Moreover, this EO was active at lowest concentration against germination of S. arvensis and R. sativus.

E. salubris EO was active in similar way against radical elongation and germination of S. arvensis and R. sativus. Instead, for L. multiflorum and C. sativus seeds data showed a major activity against germination than radical elongation.

4.3. Anti-enzymatic activities

Table 5 showed the enzymatic inhibitory effects of the EOs. In particular, E. campaspae EO was the most active against AChE with an IC₅₀ value of 51 μg/ml. The other EOs inhibition are as follow: E. salubris < E. amplifolia < E. loxophleba < E. bicolor. All EOs were less active against BChE than AChE. E. campaspae EO and E. loxophleba EO showed the higher activity against BChE with an IC₅₀ value of 1.065 mg/ml followed by E. amplifolia and E. salubris EOs. Instead, E. bicolor EO was the least active with an IC₅₀ value of 14.920 mg/ml.

Table 5.

Inhibition of AChE, BChE, α-Amylase, and α-Glucosidase activities by tested EOs.

Essential oils	IC50 (mg/ml)
	AChE	BChE	α-amylase	α-glucosidase
E. amplifolia	0.26 ± 0.02c	2.78 ± 0.41a	17.12 ± 1.25c	26.32 ± 1.51b
E. bicolor	0.371 ± 0.002d	14.92 ± 3.787b	8.097 ± 0.172b	>30
E. campaspe	0.051 ± 0.007a	1.065 ± 0.098a	20.125 ± 3.201c	20.548 ± 1.236a
E. loxophleba	0.341 ± 0.013d	2.266 ± 0.024a	0.909 ± 0.128a	18.058 ± 2.451a
E. salubris	0.22 ± 0.0b	4.27 ± 0.26a	0.57 ± 0.01a	19.47 ± 1.25a
Galantamine	0.009 ± 0.003	0.06 ± 0.02	–	–
Acarbose	–	–	0.003 ± 0.001	0.7 ± 0.2

Results (mean ± SD, n = 3) with different letters indicate significant differences at p < 0.05 (one-way ANOVA, Tukey's post-hoc test).

The EOs from E. salubris and E. loxophleba exhibited the highest activity against α-amylase with an IC₅₀ value < 1 mg/ml followed by E. bicolor, E. amplifolia and E. campaspe EOs.

Instead, E. loxophleba, E. salubris and E. campaspe demonstrated the highest activity against α-glucosidase even if with high IC₅₀ values. E. bicolor EO was inactive against α-glucosidase with an IC₅₀ value > 30 mg/ml.

All samples showed IC₅₀ values higher than those of standards used as positive control.

4.4. Antimicrobial activity

The in vitro antibacterial efficacy of the EOs was assessed through identification of the minimum inhibitory concentration (MIC) (Table 6). Additionally, the potential effects on biofilm formation and bacterial cell metabolism were elucidated and presented in Table 7, Table 8.

Table 6.

MIC value (mg/mL) of five eucalyptus EOs against assessed pathogens.

Species	E. bicolor	E. loxophleba	E. campaspae	E. salubris	E. amplifolia
A. baumannii	>50a	38 ± 4a	>50a	16 ±3a	14 ±1a
E. coli	>50a	35 ± 2b	35± 4b	18 ±2b	14 ±2a
L. monocytogenes	30 ± 3b	35 ± 3b	35 ± 2b	14 ±2c	15 ±2 ab
P. aeruginosa	38 ± 3c	38 ± 2a	38 ± 4c	14 ±2c	15 ±1 ab
S. aureus	35 ± 2d	30 ± 3c	28 ± 2d	14 ±1c	16 ±2b

The tests were conducted three times and the results are presented as the average ± SD) With different letters, denote significant differences at p < 0.05, following one-way ANOVA and SNK post hoc test.

Table 7.

Inhibitory activity of the EOs on the mature biofilm.

	E. amplifolia		E. bicolor		E. compaspae		E. loxophleba		E. salubris
Concentrations (mg/mL)	10	20	10	20	10	20	10	20	10	20
A. baumannii	33.44 ± 4.32a	50.79 ± 3,67a	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00	21.82 ± 1.98a	30.66 ± 2.09a	3.41 ± 0.13a	57.24 ± 4,44a
E. coli	62.5 ± 3.96b	78.27 ± 5,61b	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00	42.65 ± 3.86a	40.22 ± 3.67b	45.85 ± 5.04b	33.38 ± 2.41b	35.35 ± 1,25b
L. monocytogenes	63.86 ± 3.87b	69.31 ± 5,01c	0.00 ± 0.00	18.49 ± 1.04a	12.32 ± 1.01a	57.42 ± 5.04b	23.92 ± 1.25a	44.85 ± 4.54c	68.54 ± 4.76c	71.21 ± 3,23c
P. aeruginosa	10.93 ± 0.87c	61.45 ± 3,78c	0.00 ± 0.00	34.14 ± 3.05b	19.03 ± 2.01b	23.35 ± 2.87c	0.00 ± 0.00	24.39 ± 1.87d	72.39 ± 2.51d	79.45 ± 3,55c
S. aureus	53.21 ± 3.53d	57.86 ± 3.67a	8.71 ± 0.92a	45.53 ± 3.99c	38.86 ± 2.44c	76.61 ± 3.52d	25.29 ± 1.55a	70.94 ± 2.69e	71.41 ± 3.01d	88.24 ± 1.67d

Results (mean ± SD, n = 3) with different letters denote significant differences at p < 0.05, following one-way ANOVA and SNK post hoc test.

Table 8.

Inhibation effects of EOs on bacterial sessile cell metabolism in mature biofilm.

	E. amplifolia		E. bicolor		E. compaspae		E. loxophleba		E. salubris
Concentrations (mg/mL)	10	20	10	20	10	20	10	20	10	20
A. baumannii	60.55 ± 1.25a	94.69 ± 2.08a	30.3 ± 2.02a	54.46 ± 4.87a	85.96 ± 2.12a	91.03 ± 3.11a	93.47 ± 1.32a	94.09 ± 2.86	53.04 ± 1.44a	64.13 ± 1.17a
E. coli	0.00 ± 0.00	78.81 ± 1.76b	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00	1.88 ± 0.02b	63.36 ± 2.61b	77.65 ± 3.51	53.58 ± 2.21a	81.72 ± 1.89b
L. monocytogenes	0.00 ± 0.00	81.21 ± 4.02c	14.2 ± 2.21b	16.22 ± 1.55b	0.00 ± 0.00	81.92 ± 2.21c	74.04 ± 2.54c	81.23 ± 1.88	28.89 ± 2.07b	81.62 ± 3.92b
P. aeruginosa	0.00 ± 0.00	80.88 ± 4.97c	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00	0.00 ± 0.00	71.82 ± 3.44	14.60 ± 1.01c	57.81 ± 4.07c
S. aureus	57.74 ± 2.01b	93.40 ± 1.04a	41.8 ± 3.13c	44.48 ± 3.32c	67.16 ± 1.67b	81.78 ± 1.13c	92.78 ± 2.09a	94.35 ± 2.67	74.15 ± 2.25d	92.62 ± 1.45d

Results (mean ± SD, n = 3) with different letters denote significant differences at p < 0.05, following one-way ANOVA and SNK post hoc test.

The EOs manifested pronounced inhibitory effects, characterized by MIC values spanning from 14 mg/ml (E. salubris) to values surpassing 50 mg/ml (E. bicolor), as delineated in Fig. 3. The variability in MIC values was contingent upon the specific EOs and bacterial strains under scrutiny. Notably, each EO demonstrated unique inhibitory effects on biofilm formation, with efficacy varying according to bacterial species and concentrations.

The MIC reveal diverse sensitivities across the tested bacterial strains, with Gram-positive strains exhibiting greater sensitivity than Gram-negative ones which the MIC recorded exceeded 50 mg/mL. As for Gram-positive strains, the maximum values were observed as follows: P. aeruginosa (38 ± 4, E. compaspae), L. monocytogenes (35 ± 3, E. loxophleba), and S. aureus (35 ± 2, E. bicolor).

With the exception of the EOs of E. bicolor and E. campaspae, mature biofilms exhibited sensitivity to the effects of EOs, emphasizing their responsiveness. Notably, no inhibition was detected in the case of A. baumannii when exposed to E. bicolor and E. campaspae. However, a significant reduction in cellular metabolism was observed at the highest concentration, resulting in percentages of 54.46 ± 4.87 and 91.03 ± 3.11 %, respectively.

Similarly, E. coli demonstrated resistance when treated with E. campaspae EO; no inhibition was observed at 10 mg/ml for both mature biofilm and cellular metabolism. Yet, with an increased concentration, a 42.65 ± 3.86 % inhibition of mature biofilm and only 1.88 ± 0.02 % inhibition of cellular metabolism were observed. This highlights the distinctive effectiveness of these EOs in influencing microbial activity. After the application of E. amplifolia EO, similar percentage of inhibition was observed on mature biofilm (78.27 ± 5.61) and on cell metabolism (78.81 ± 1.76) at 10 mg/ml.

Among Gram-positive strains, S. aureus demonstrated the highest sensitivity. The observed inhibition of mature biofilm varied from 8.71 ± 0.92 % (E. bicolor) to 88.24 ± 1.67 % (E. salubris). In the case of cell metabolism, the inhibition ranged from 41.83 ± 3.13 (E. bicolor) to 94.35 ± 2.67 % (E. loxophleba).

The effectiveness of EOs from E. salubris, E. amplifolia, and E. loxophleba was evident in reducing biofilm formation across various bacterial strains. Additionally, these EOs exhibited a hindering effect on the cellular metabolism of these pathogens. Interestingly, the inhibition caused by the EOs did not consistently correlate with a comparable influence on the metabolic activity of bacterial cells within the biofilm. For example, E. loxophleba showed a notably higher impact on P. aeruginosa cellular metabolism, reaching 71.82 ± 3.14 %, compared to its effect on mature biofilm at 24.29 ± 1.87 %, at the identical concentration of 20 mg/ml. Similarly, E. compaspae EO demonstrated a substantial 81.92 ± 2.21 % inhibition of L. monocytogenes cellular metabolism. However, this value diminished to 57.42 ± 5.04 % when the same EO was applied to mature biofilm, at of 20 mg/ml.

5. Discussion

In the majority of cases, the quantities of EO obtained closely match the data reported in the literature. For instance, the EO from E. bicolor demonstrated similarity to EO yields from Iraq (1.45 %) as reported by Dehghani et al. (2023) [28]. A separate study conducted in Tunisia indicated 1.50 ± 0.3 % EO yield [29], while another study from the Province of Isfahan, Iran reported 1.85 % EO yield [30]. However, for species such as E. amplifolia, E. salubris, and E. campaspe, the yields observed in this study surpass previously documented values. E. amplifolia EO was reported to yield 0.9–1.2 % from trees growing in South Wales, Australia [31], and 0.36 % from trees growing in Uruguay [32].

E. salubris EO was reported to yield 1.5–1.65 % from trees growing in Morocco [33] and 4–3.8 % from trees growing in Tunisia [34]. E. campaspe EO was reported to yield 1.90 % from trees growing in Arboretum de Waite, South Australia l [35]. Conversely, for E. loxophleba EO, the yields are lower than previous references. A yield of 7.1 % was reported from trees growing in southwest Iran [36], and a similar yield to our result, 3.8 %, was reported from trees growing in Narrogin, southwest Western Australia [37].

The disparities in yield compared to existing data may be ascribed to various factors, encompassing climate, soil characteristics [38] timing of collection [38] and geoclimatic conditions [39].

By exploring the biological potential of Eucalyptus EOs, this study contributed valuable insights into their diverse applications and effects.

The analysis of their chemical composition highlighted interspecific diversity which is confirmed by PCA and HCA analyses.

Notably, the chemical composition of E. amplifolia EO has been previously documented by Hellyer and McKern [31], reporting a composition rich in isovaleric aldehyde, α-pinene, eucalyptol, limonene, α-terpineol, aromadendrene, α- and β-eudesmol. However, the quantity of the most present components was not extensively detailed in the cited work, except for eucalyptol, which was present at 19.5 %. A comparison with subsequent works revealed variations in the concentrations of shared compounds, such as α-pinene, eucalyptol, and aromadendrene. In 1990, Dellacassa and collaborators [32] presented a different composition of E. amplifolia essential oil, with eucalyptol as the main component (40.8 %), accompanied by limonene, α-pinene, globulol, and aromadendrene. Comparisons with earlier works highlighted differences in the concentration of eucalyptol and other shared compounds, underscoring the dynamic nature of the chemical composition of E. amplifolia EO. A similar trend was observed in study by Bignell and coworkers [35], who reported an oil composition similar to the one examined in this study. Although both EOs shared eucalyptol as the main component, significant differences in concentration were noted for eucalyptol and allo-aromadendrene, reinforcing the importance of considering variations in chemical profiles.

The examination of E. bicolor EO by Elaissi and coworkers [29] highlighted similarities in composition with our study, presenting shared compounds such as eucalyptol, aromadendrene, α-pinene, and trans-pinocarveol. While the quantities differed, the commonality in major components highlights consistency in the chemical composition. Similarly, Al-Snafi [40], emphasized the prevalence of eucalyptol in, aligning with the findings of our study. Although variations in concentrations were noted, the shared presence of eucalyptol signifies its significance in defining the chemical profile.

Examining E. campaspe EO, Dhaouadi et al. [41] identified eucalyptol as the main component.

The investigation into E. loxophleba EOs by Aldoghaim [42], revealed differences in the concentration of eucalyptol when compared to a 2011 study by Rahimi-Nasrabadi and Batooli [36]. While eucalyptol was the only shared main component, the divergent quantities highlight the inherent variability in E. loxophleba EO. Similarly, the study of aTunisian E. loxophleba EO by Elaissi in 2010 [43] exhibited variations in the concentration of eucalyptol compared to other works. Despite differences, the common presence of eucalyptol reinforces its significance in defining the chemical composition.

Exploring E. salubris EOs, Elaissi and collaborators [34] demonstrated common compounds like limonene, eucalyptol, cryptone, and α-pinene, albeit with different concentrations. While variations exist, the presence of eucalyptol as a shared major component underscores its significance in defining the chemical profile. Zhang and collaborators [44] reported an EO of E. salubris whose main components were eucalyptol, α-pinene, p-cymene and trans-pinocarveol. The paper does not report the quantity of these components but all four are present as majority compounds in the EO reported in this work. In 2020 Abd-ElGawad and collaborators [45] report an EO of E. salubris which had eucalyptol, α-pinene and p-cymene as main components. This work does not report the quantity of these components either but all three are present as majority compounds in the OE reported in this work.

Evidently, the chemical analyses underscore the necessity to discern the potential influence of environmental factors on their compositional dynamics. This assertion underscores the intricacies involved in elucidating the complex chemical constitution of Eucalyptus EOs and the concomitant environmental variables that may exert discernible effects on their overall composition [[46], [47]].

In existing literature, there is an absence of prior research elucidating the phytotoxic effects on the seeds of L. multiflorum, S. arvensis, R. sativus, and C. sativus induced by various Eucalyptus species.

Numerous studies have underscored the capacity of Eucalyptus EOs to impede or delay the germination processes of seeds such as Triticum aestivum L., Zea mays L. Cassia occidentalis L., Amaranthus viridis L., Echinochloa crus-galli (L.) P. Beauv., Parthenium hysterophorus L., and Bidens pilosa L [[48], [49], [50]].

Our preceding investigations have also identified the phytotoxicity of several Eucalyptus EOs against R. sativus, L. multiflorum, and S. arvensis [9,51]

The phytotoxicity associated with Eucalyptus EOs is likely attributed to the elevated presence of the monoterpene eucalyptol (1,8 cineole), known for inhibiting mitochondrial respiration, mitosis, and DNA synthesis, thereby influencing the germination process [52]. This inhibitory activity is closely tied to the chemical structure of the compound, as exemplified by the structural similarity between the first commercialized allelopathic herbicide, cinmethylin and natural 1,8-cineole [53].

Conversely, the variation in phytotoxic activities observed among different EOs may stem from potential synergistic interactions between their constituents, influencing enzymes crucial to the germination process [52].

Key enzymes in the germination process include α-amylase and α-glucosidase, responsible for hydrolyzing starch reserves into maltose and further converting maltose into glucose, respectively [54,55].

Consequently, following the determination of phytotoxic activities from EOs, an assessment of their potential impact on the regulation of these enzymes was conducted. The results align with the phytotoxic assays, indicating that E. loxophleba and E. salubris EOs, exhibiting the highest phytotoxicity at the lowest concentrations tested (125 μg/ml), also demonstrated the lowest IC₅₀ values for both α-amylase and α-glucosidase. Furthermore, although E. bicolor EO did not display activity against α-glucosidase (IC₅₀ > 30 mg/ml), it exhibited an IC₅₀ < 10 mg/ml against α-amylase and proved more phytotoxic at the lowest concentration (125 μg/ml) compared to E. amplifolia and E. campaspe EOs, which showed IC₅₀ values > 20 mg/ml for one or both enzymes.

Existing literature only encompasses two prior studies that hint at a potential correlation between the phytotoxic and anti-α-amylase activities of Eucalyptus pulverulenta Sims, E. gunnii Hook. f., and O. heracleoticum L. EOs [8,56].

Remarkably, no previous studies have reported a plausible correlation between phytotoxicity and the inhibition of α-glucosidase activity.

Building upon the recognized advantages of natural pesticides over chemical counterparts, biopesticides, such as those derived from Eucalyptus EOs, present a promising alternative to conventional pest control methods [57]. The diverse and desirable properties of Eucalyptus EOs for pest management have been extensively explored in the literature, encompassing various species like E. camaldulensis Dehnh., E. citriodora Hook., E. globulus, E. grandis W. Hill., E. robusta Sm., and E. saligna Sm [[58], [59], [60], [61], [62], [63], [64]].

Despite the well-established pesticidal potential of Eucalyptus EOs, the exploration of their impact on cholinesterases, crucial enzymes targeted by pesticides, has been limited in previous studies [65,[66], [67]]. Addressing this research gap, our study delves into the pesticidal capabilities of five Eucalyptus EOs. The results unequivocally demonstrate the inhibitory effects on acetylcholinesterase (AChE) and activity against butyrylcholinesterase (BChE) for all EOs, except for E. bicolor, which exhibited relatively lower activity against BChE. These findings not only underscore the multifaceted utility of Eucalyptus EOs in pest management but also emphasize their potential as biopesticides with dual efficacy against key enzymatic targets.

In examining the antimicrobial potential of Eucalyptus EOs, our investigation extends to diverse geographical sources, revealing unique properties of each species. The E. loxophleba EO sourced from western Australia exhibited moderate antimicrobial potential, with zone of inhibition values ranging from 14.7 ± 0.6 to 16.7 ± 0.6 against E. coli ATCC 25922 and 15.3 ± 0.6 to 16.7 ± 0.6 against S. aureus ATCC 29213 when exposed to 25 and 50 μL of EOs. Additionally, the MIC was determined to be 8 % v/v for E. coli and 4 % v/v for S. aureus [42].

The examination of Tunisian E. salubris EOs revealed noteworthy effectiveness against various microorganisms, except for S. aureus and E. coli. Significant activity was observed against L. monocytogenes (18 mm) and K. pneumoniae (12 mm) [68].

In the case of E. salubris EOs from Jordanian trees, significant antibacterial effectiveness was observed against both Gram-negative and Gram-positive bacteria, with MIC values ranging from 98 μg/ml for S. aureus to 140 μg/ml for P. aeruginosa, and 119 μg/ml against E. coli [[69], [70]].

An E. bicolor EO from Iran demonstrated moderate to high antimicrobial impact against various bacteria, fungi, and yeast, with no effect against P. aeruginosa or E. coli [30].

Subsequent studies confirmed the absence of antibacterial effects on P. aeruginosa, while revealing a significant impact on E., and an MIC range of 62.4–61.4 μg/ml against S. aureus [[71], [72]].

Notably, while previous reports have documented the antibacterial activities of E. bicolor, E. salubris, and E. loxophleba, a comprehensive literature search revealed a dearth of information regarding the antibacterial activities of the EOs from the two additional species under examination, namely E. campaespe and E. amplifolia. This underscores the need for further exploration and understanding of the antimicrobial potential of these Eucalyptus species.

In conclusion, this study not only contributes valuable insights into the diverse applications of Eucalyptus EOs but also emphasizes the need for continued exploration and understanding of their complex biological interactions. The multifaceted nature of Eucalyptus EOs, as revealed through this research, underscores their potential for various applications in agriculture, pest management, and healthcare, paving the way for future studies and practical applications in these domains.

Funding

This research received no external funding.

Data availability statement

Data will be made available on request.

CRediT authorship contribution statement

Sana Khedhri: Writing – original draft, Writing – review & editing, Formal analysis, Data curation. Flavio Polito: Writing – review & editing, Investigation, Data curation. Lucia Caputo: Writing – review & editing, Software, Methodology, Data curation. Marwa Khammassi: Investigation. Ferjani Dhaouadi: Investigation. Ismail Amri: Project administration. Lamia Hamrouni: Resources, Conceptualization. Yassine Mabrouk: Investigation, Data curation. Florinda Fratianni: Data curation. Filomena Nazzaro: Validation, Methodology, Formal analysis, Data curation. Vincenzo De Feo: Writing – review & editing, Visualization, Validation, Supervision, Resources, Project administration, Funding acquisition, Conceptualization.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.