Influence of Agronomic Practices on the Antioxidant Activity of Three Mediterranean Officinal Wild Plants: Silybum marianum, Achillea millefolium, and Trifolium pratense

Abstract

The aim of this study was to evaluate the influence of various agronomic factors on plant growth and the accumulation of secondary metabolites with antioxidant properties. The three Mediterranean Officinal Wild Plants selected for this investigation were Silybum marianum, Achillea millefolium, and Trifolium pratense whose extracts, enriched in phenolic compounds, are well documented for their potential therapeutic effects. Three agronomic factors were evaluated, each with two treatment options, resulting in eight experimental combinations: (1) inoculation with plant growth-promoting rhizobacteria (PGPR) versus control (no inoculation); (2) high versus low fertilization rates of K₂O and P₂O₅ to modulate nutrient availability; (3) water stress at 40% of field capacity compared to the control with full field capacity. Plant growth was monitored using the BBCH (Biologische Bundesanstalt, Bundessortenamt and CHemical industry) scale to delineate key phenological phases, with treatments applied until the flowering stage was reached. Only the leaves of the plants were collected, and hydroalcoholic extracts were prepared for the evaluation of total antioxidant capacity (TAC) using the FRAP, DPPH, and ABTS assays. These assays were selected due to their complementary insights into the chemical mechanisms underlying TAC, as well as their ability to assess the physicochemical characteristics of the phytochemical constituents.

1. Introduction

According to the World Health Organization (WHO), medicinal plants are defined as any plants containing substances in one or more of their organs that can be used for therapeutic purposes or serve as precursors in the synthesis of beneficial pharmaceutical compounds [1]. Plants biosynthesize a diverse range of secondary metabolites (in particular, phenolic species) that do not appear to be directly involved in primary metabolic processes [2]. Although these secondary metabolites are not essential for sustaining fundamental physiological functions, they provide significant benefits by promoting plant growth, aiding adaptation to changing environmental conditions, and protecting against environmental stressors and damage [3,4]. Several agronomic factors influence plant growth and the accumulation of secondary metabolites, which are critical for the antioxidant properties of herbal extracts due to their pharmaceutical and nutraceutical importance [5].

The literature suggests that fertilization, particularly with nitrogen, potassium, and phosphorus, plays a crucial role in influencing both crop yield and quality. For example, increasing soil potassium levels has been linked to a higher concentration of phenolic compounds, which contribute to the antioxidant activity of medicinal plants [6]. Similarly, manipulating nitrogen fertilization during plant growth significantly impacts phenolic compound production [7]. Some studies propose a “physiological trade-off” between plant growth and secondary metabolite synthesis, indicating that, phenolic concentrations tend to increase when nutrient availability is limited, particularly under lower nitrogen treatments [8,9].

Assessing the antioxidant capacity of plant extracts is key to understanding their health benefits [10,11,12]. Antioxidants act through various mechanisms, such as radical scavenging, metal ion chelation, and electron transfer. Since compounds may perform differently across assays due to varying conditions, using multiple assays offers a more thorough evaluation [10,11,12,13,14]. This approach also improves data reliability and strengthens conclusions about antioxidant health benefits.

Three common methods for measuring antioxidant capacity are the Ferric Reducing Antioxidant Power (FRAP), 2,2-diphenyl-1-picrylhydrazyl (DPPH), and 2,2′-azinobis (3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) assays. Each assay has unique advantages and limitations, providing complementary insights into the antioxidant properties of plant extracts. Using multiple assays also enables cross-validation of results, reinforcing hypotheses about plant extracts’ efficacy and health benefits. Different compounds respond uniquely depending on their structural characteristics. For instance, FRAP favors phenolic compounds and metal chelators, while DPPH is more sensitive to electron-donating compounds like flavonoids and carotenoids. The ABTS assay accommodates a broader range of antioxidants, including phenolics, flavonoids, and vitamins.

In this manuscript, the phenological development of the three selected Mediterranean Officinal Wild Plants (MOWPs) Silybum marianum L. Gaertn (milk thistle), Achillea millefolium L. (yarrow), and Trifolium pratense L. (red clover) was observed and described in detail through photographs of various vegetative and reproductive key phenological stages, with focus on the plant portions growth of the leaves, stem, and flowers. The three selected MOWPs are recognized for their nutraceutical properties in the Mediterranean region. Notably, the phytocomplexes derived from these plants are rich in phenolic compounds, which are well documented for their potential therapeutic effects [15,16,17,18].

Silybum marianum is a spiny herbaceous species of the Asteraceae family. Under natural conditions, it typically remains in the vegetative stage during the first growing season following seed germination and is therefore commonly classified as a biennial plant [19]. It is considered a medicinal plant, cultivated for many important secondary metabolites expressing antioxidant activity, hepatoprotective effect, and antifibrotic properties [20,21].

Achillea millefolium is a perennial herbaceous plant belonging to the Asteraceae family. In traditional medicine, it is used as a tonic for blood circulation, antispasmodic, and hemostatic. The plant is rich in terpenic compounds such as cineole, pinene, and azulene, which enhance its phytochemical and pharmacological value [22].

Trifolium pretense belongs to the Fabaceae family and is a perennial herbaceous plant, although generally of limited longevity, typically no more than two years. It is mainly used as a forage plant, widespread throughout Europe, and known for its ability to fix atmospheric nitrogen in the soil. Recently, it has gained significant interest due to its richness in isoflavones [23].

The primary objective of this study was to evaluate the content of bioactive compounds in the selected MOWPs, as well as to assess the growth of the plants and the duration of their phenological stages. The effects of various combinations of agronomic factors on the presence of antioxidants were investigated through multiple assays conducted on different plant parts (leaves, stems, and flowers) of the cultivated species. This approach aimed to identify the most effective agronomic combinations for maximizing the yield of secondary metabolites.

2. Materials and Methods

2.1. Study Area

The experiment was carried at the Terminillo Apennine Center “Carlo Jucci” in the Province of Rieti, a University of Perugia agricultural experimental center in a controlled environment to avoid water inputs from rainfall but with temperatures identical to the outdoor environment. Rieti is situated at an altitude of about 405 m above sea level in central Italy’s Lazio region. It lies within the Rieti plain surrounded by mountains such as Mount Terminillo to the east. This geographical setting influences its climatic conditions by moderating temperature extremes compared to coastal areas. The climate of the study area is characterized by a “humid temperate climate” with a Köppen classification of Cfb, with cool winters and hot summers with significant precipitation throughout the year [24]. Overall, Rieti’s climate combines elements typical of both temperate inland regions and Mediterranean influences due to its location near mountainous terrain but still within a broader Mediterranean climatic zone. Temperature during the study period was evaluated by calculating 10-day mean temperature values and through the GDD amounts with a base temperature of 0 °C for analyzing the relationships of climatic effects on phenological stages for each species.

2.2. Experimental Design

Seed vernalization was initially performed by immersing the seeds in 20 mL of sterile distilled water, followed by a 48 h incubation period at 4 °C in the dark. Subsequently, the seeds were sown in multi-pots (40 pots, each with a 2.5 cm diameter) and maintained under controlled temperature conditions. Upon germination, the seedlings were transplanted into larger pots (20 cm × 20 cm × 15 cm) and placed outdoors under natural environmental conditions. The experimental design, a three-factorial pot experiment incorporating three binary factors, was conducted to examine the cultivation of selected species under varying environmental and agronomic conditions (

Table 1. Experimental factors (PGPR application, fertilization, and irrigation) combined in the different cultivation protocols.

Combination	PGPR	Fertilization	Water Supply
	Inoculation	(K2O-P2O5)	(Field Capacity)
0,0,0	0	0	0
0,1,0	0	1	0
0,1,1	0	1	1
0,0,1	0	0	1
1,1,0	1	1	0
1,0,0	1	0	0
1,0,1	1	0	1
1,1,1	1	1	1

PGPR: 0 = absence; 1 = presence. Fertilizer: 0 = low; 1 = high. WS: 0 = 40%; 1 = 100% field capacity.

). The study was performed under controlled conditions, utilizing eight distinct combinations of factors, resulting in eight possible cultivation protocols (2³ combinations). Each factor was assigned a binary value, with (1) representing the maximum level and (0) representing the minimum level. Each treatment combination was replicated five times. The study evaluated the effects of drought stress by comparing full irrigation at field capacity (1) with irrigation at 40% relative soil moisture (0) representing no stress and severe stress conditions, respectively. Furthermore, the influence of inorganic fertilizers, specifically P₂O₅ and K₂O, was assessed, with treatments applying either minimum or maximum quantities. In addition, the contribution of plant growth-promoting rhizobacteria (PGPR) is regarded as a key anti-stress factor, playing a crucial role in mitigating environmental stresses and enhancing plant resilience. The cultivation medium consisted of an organic substrate with a pH of 5.5 and nitrogen fertilization at 180 mg N/L. Each treatment combination was tested with five plant pot replications, each containing 0.78 dm³ of substrate.

Agronomical Factors

Drought stress was applied before and during early flowering stage. Soil moisture was measured every day by an electronic depth probe (Aicevoos Digital Soil Tester, Aicevoos, Wuhan, China). Samples treated with water stress were maintained for 5 weeks at 40% of relative soil moisture. For convenience, the relative soil moisture value was correlated with the field capacity through the method of pot weighing and measuring with the probe in the soil. Full soil water capacity was considered at around 70% moisture value.

A lower-nutrient soil was used for the experiment and leaves fertilization with phosphorus pentoxide (P₂O₅) and potassium oxide (K₂O) was carried out three times, the first after 44 days after sowing, the second after 69 days, and the third after 78 days. To prepare the fertilizer solution, 2.5 g of product (bottos–Pregade 30–20) was weighed and diluted in 2 L of water, and this solution was distributed by a pressure sprayer onto the leaves of the plants grown in accordance with the protocol that included the presence of the fertilizer.

The plant growth-promoting rhizobacteria (PGPR) utilized in the experiment was a commercial product containing a mixture of endomycorrhizal bacteria and fungi, including Bacillus spp., Streptomyces spp., and Pseudomonas spp. (with a rhizosphere bacterial content of 1.6 × 10⁸ CFU/g), and Trichoderma spp. (with a mycorrhizal fungal content of 5 × 10⁵ CFU/g). The product was inoculated into the growing substrate at a rate of 10 g per 100 seedlings. The first application was made immediately after transplanting (30 days after sowing), and the second application was performed 20 days later.

2.3. Evaluation of Growth Trend Using BBCH Scales

In this manuscript, the phenological development of three selected MOWPs is described using the extended BBCH (Biologische Bundesantalt, Bundessortenamt, and Chemische Industrie) scale, with growth stages categorized using both the 2-digit BBCH coding systems [25]. Representative photographs of the key phenological stages are included to supplement the discussion. Observations were recorded weekly, indicatively at late morning mid-week, starting from sowing to full vegetative development for Silybum marianum and to the beginning of flowering for the other two species.

2.4. Plant Material

The plants of Silybum marianum, Achillea millefolium, and Trifolium pratense were collected in October and immediately oven-dried at 45 °C for 3 days to obtain dry matter. IKA^® Tube Mill Control (IKA^®-Werke GmbH & co. KG, Staufen, Germany) was used to pulverize the dried plants (20,000 rpm for 45 s).

Reagents Employed for the Appraisal of the Total Antioxidant Capacity (TAC) with the Three Spectrophotometric Assays

Methanol (MeOH), ethanol (EtOH), potassium persulfate (K₂S₂O₈), 2,2′-Azino-bis-(3-ethylbenzothiazoline-6-sulphonate) diammonium salt (ABTS), 2,4,6-tris(2-pyridyl)-s-triazine (TPTZ), 6-hydroxy-2,5,7,8-tetramethyl-2-carboxylic acid (Trolox), 2,2-diphenyl-1-picrylhydrazyl (DPPH), ferric chloride (FeCl₃), and sodium acetate (NaOAc) were purchased from Merck Life Science S.r.l. (Milan, Italy).

2.5. Extraction of the Phenols Containing Phytocomplex

The powdered plants (30 mg) were extracted using 4 mL of an EtOH/water (50/50, v/v) solution through sonicating for 30 min at room temperature (r. t.). The mixture was then centrifuged (5000 rpm for 20 min at 4 °C) and the supernatant was collected and stored at 4 °C until analyzed.

2.5.1. Evaluation of the TAC Using the FRAP Method

The FRAP assay was performed with slight modifications to the method described in [13,26,27,28]. For the preparation of the FRAP reagent, 2.5 mL of a 10 mM TPTZ solution in 40 mM aqueous HCl and 2.5 mL of a 20 mM FeCl₃ aqueous solution were combined with 25 mL of 300 mM aqueous NaOAc (pH 3.6). The assay was conducted by mixing 100 μL of the ethanol/water extract, 100 μL of bi-distilled water, and 1.5 mL of the FRAP reagent. The solution was then incubated at room temperature in the dark for 4 min. Absorbance was measured at 570 nm. The total antioxidant capacity (TAC) values were determined using a calibration curve prepared with Trolox (as the standard), where the Trolox solutions were treated following the same procedure used for the samples (calibration curve range: 0.01–0.25 mg/mL; R² = 0.996). For the measurements, 0.2 mL of the solutions were used. All experiments were performed in triplicate for each extract, and the results are expressed as mg of Trolox equivalents (TEs) per gram of dry extract.

2.5.2. Evaluation of the Radical Scavenging Capacity by the DPPH Method

The DPPH assay was performed with slight modifications to the method described in [13,26]. DPPH was dissolved in ethanol to obtain a solution with an absorbance of 0.65 (±0.02) at 517 nm, and the absorbance value was allowed to stabilize for 2 h. Subsequently, 50 µL of the ethanol/water extract was added to 2.95 mL of the stabilized DPPH solution, and the mixture was kept at room temperature in the dark for 30 min. Absorbance was measured at 517 nm. The radical scavenging capacity was determined from a calibration curve prepared using Trolox (as the surrogate standard) solutions, which were treated in the same manner as the studied samples (calibration curve range: 0.005–0.25 mg/mL; R² = 0.993). For the measurements, 0.2 mL of the solutions were used. All experiments were performed in triplicate for each extract, and the results are expressed as mg of Trolox equivalents (TEs) per gram of dry extract.

2.5.3. Evaluation of the Radical Scavenging Capacity Using the ABTS Method

Two volumes of an aqueous ABTS•+ solution (0.36% w/v) were mixed with one volume of a 0.2% aqueous K₂S₂O₈ solution. The flask was covered with aluminum foil and left overnight at room temperature in the dark. The resulting ABTS•+ solution was then diluted with ethanol until the absorbance reached 0.70 (±0.05) at 690 nm. To 0.06 mL of the ethanol/water extract, 4.0 mL of the ABTS•+/ethanol solution was added, and the mixture was allowed to stand in the dark for 6 min. Absorbance was measured at 690 nm. The radical scavenging capacity was determined from a calibration curve prepared with Trolox (used as the surrogate standard), with solutions treated in the same manner as the studied samples (calibration curve range: 0.01–0.75 mg/mL; R² = 0.998). For the measurements, 0.2 mL of each solution was used. All experiments were performed in triplicate for each extract, and the results are expressed as mg of Trolox equivalents (TEs) per gram of dry extract.

2.6. Instrumentation

UV/Vis spectrophotometric assays were performed using a Sunrise™ Absorbance microplate reader (TECAN, Männedorf, Switzerland). All the spectrophotometric experiments were performed using disposable optical Corning^® 96-well plates from Merck Life Science (Merck KGaA, Darmstadt, Germany) and a Sunrise microplate reader (Tecan Italia S.r.l., Milan, Italy).

Liquid chromatographic separation and mass spectrometric analysis were performed on a UHPLC-MS/MS system consisting of an Agilent 1290 Infinity II combined with the Agilent 6560 mass spectrometer (Agilent Technologies Inc., Santa Clara, CA, USA). The chromatographic separation was performed using a ZORBAX RRHD Eclipse Plus C18 column (50 mm × 2.1 mm, 1.8 µm, 95 Å, Agilent Technologies Inc.). UHPLC eluent A was water (LC-MS grade, LiChrosolv, Supelco, Merck KGaA, Darmstadt, Germany) with 0.1% (v/v) formic acid (LC-MS grade, LiChropur, Supelco, Merck KGaA, Darmstadt, Germany) while eluent B was acetonitrile (LC-MS grade, LiChrosolv, Supelco, Merck KGaA, Darmstadt, Germany). The optimized gradient program was the following: 0–3.5 min, 5% (v/v) B; 3.5–5 min, 5–25% (v/v) B; 5–9 min, 25–35% (v/v) B; 9–11.5 min, 35–97% (v/v) B; 11.5–12.5 min, 97% (v/v) B; 12.5–13 min, 97–5% (v/v) B; 13–17 min, 5% (v/v) B (column equilibration/conditioning). The column temperature was set at 25 °C and the flow rate at 0.3 mL/min. The injection volume was 5 µL. The UV–Vis chromatograms were recorded at 250, 280, 320, and 350 nm. For MS detection, the Dual AJS ESI source operated in positive ion mode. The gas temperature was set at 300 °C with a flow of 5 L min⁻¹ while the sheath gas temperature was 350 °C with a flow of 11 L/min. The nebulizer pressure was set at 35 psi and the Capillary and Fragmentor voltages were 3500 V and 300 V, respectively. In the case of MS² analysis, multiple experiments using the iterative algorithm of the instruments were carried out. The fragmentation patterns of the compounds were recorded at a fixed collision energy (20 eV) with an isolation width of 4 m/z. The Masshunter Workstation Data Acquisition 10.0 (Agilent Technologies Inc., Santa Clara, CA, USA) program was used for data acquisition while the Masshunter Qualitative Analysis 10.0 (Agilent Technologies Inc., Santa Clara, CA, USA) software was used for data processing.

2.7. Statistics

All the analyses were performed in triplicate, and the values were reported as the mean ± standard deviation (SD) of the three independent analyses. The Statistica 12.0 software (StatSoft GmbH, Hamburg, Germany) was used to perform the statistical analyses. The significant differences among the studied parameters were analyzed using a one-way ANOVA, followed by a post-hoc Tukey’s Honestly Significant Difference (HSD) test at a significance level of p ˂ 0.01. The correlation between the total polyphenol content, DPPH antioxidant capacity, and FRAP-reducing power was evaluated through a Pearson correlation test.

3. Results and Discussion

3.1. Study Area Climate Conditions

The 10-day mean temperature values calculated in 2024 evidenced higher values in comparison to those of the precedent 30-year period. In Figure 1, the mean temperature and the GDD from the sowing period to the harvest were shown, and high differences in 2024 were recorded above all during July and August when the mean temperature showed about 3 °C higher than the historical data. Moreover, in 2024, a rapid reduction of mean temperature was recorded at the beginning of September with a drop of about 7 °C in a few days, which consequently determined a lower GDD accumulation during the last phenological phase development.

3.2. Phenological Observations

The seeds of the study plants were sown on 20 June 2024 in multi-pots and maintained under controlled temperature conditions. After germination, the shoots were transplanted into larger pots and placed outdoors for about 12 weeks under natural environmental conditions. In Silybum marianum, the beginning of leaf development, corresponding to BBCH 12 in Figure 2a, occurred just seven days following sowing (DFS). After the emergence of the first pair of true leaves, leaf development progressed with an increase in leaf number until a rosette stage was established at ground level (BBCH 31), which took place at 42_DFS, as shown in

Table 2. BBCH phenological scale of Silybum marianum with days following sowing (DFS).

DFS	BBCH Code	Description
0	0	Germination—Dry seeds
7	12	Leaf development—First pair of elliptic true leaves visible
21	15	Leaf development—Five leaves visible
34	18	Leaf development—Eight leaves visible
42	31	Rosette growth—10% of ground covered by leaves
67	37	Rosette growth—70% of ground covered by leaves
76	39	Rosette growth—90% of ground covered by leaves
90	42	20% of the maximum biomass reached
97	47	70% of the maximum biomass reached
105	49	90% of the maximum biomass reached
110	50	Flower central head present (3–5 mm ∅) but completely enclosed between developing leaves
116	51	Flower central head clearly visible between leaves, vertical bract tips, and internodes of main stem clearly elongating

. By the 90th day following sowing, vegetative growth resumed as the plant transitioned into stage BBCH 4, marking the onset of vegetative biomass development. This process continued until reaching the phenological stage 49 by the 105_DFS, where peak biomass accumulation was observed.

As regards Silybum marianum, we recorded a growth cycle duration of 100–130 days, which is consistent with findings from other similar studies [29,30] where under severe winter conditions, the species can be defined as an annual summer crop [31]. The phenological analysis of the selected species showed significant differences in their vegetative development. After a first growth phase similar to the other species, the Silybum marianum reached the rosette phenological stage (BBCH 3) at a later time and maintained this stage for a longer duration than the other species. This extended period was because, differently from the other two perennial species, Silybum marianum follows a biennial cycle, with buds located at ground level.

Moreover, the response to temperature trends during the cultivation period showed different effects on the selected species. Silybum marianum transitioned from phase 3 to phase 4 at the onset of significant cooling recorded during September, shifting from horizontal vegetative growth (rosette) to vertical growth by developing the bud.

In the Achillea millefolium, the cotyledons unfolding (BBCH 10) was observed after 7_DFS, as shown in Figure 2b. The vegetative phenological development was similar to the other species of the same family (milk thistle) with the complete leaf development reached at 34_DFS and the stem elongation (BBCH 3) at 42_DFS, as shown in

Table 3. BBCH phenological scale of Achillea millefolium with days following sowing (DFS).

DFS	BBCH Code	Description
0	0	Germination—Dry seeds
7	10	Leaf development—Cotyledons completely unfolded
21	12	Leaf development—2 true leaves
34	18	Leaf development—8 or more true leaves
42	32	Steam elongation—Stem 20% of final length
67	47	Harvestable vegetative plant parts have reached 70% of final size
76	49	Harvestable vegetative plant parts have reached final size
90	59	Inflorescence emergence—First flower petals visible
97	60	Flowering—First flowers open
105	61	Flowering—10% of flowers open
110	62	Flowering—20% of flowers open
116	64	Flowering—40% of flowers open

. The harvestable vegetative plant reached its final size at 76 DFS (BBCH 49). The reproductive phenological stage started at 90_DFS with inflorescence emergence while in the successive 4 weeks, the flowering process advanced to about 50% of flowers open. Generally, the Achillea millefolium takes less than 100 days from seed germination to flowering, although this can vary based on environmental conditions and care practices. As reported in the literature [32], the competitive ability of A. millefolium is mostly due to its relatively rapid growth.

In the Trifolium pratense, full germination was completed at 7_DFS, as shown in Figure 2c. The full stem elongation phase (BBCH 3) was completed at 67_DFS, followed by the development of harvestable vegetative parts that reached maximum biomass at 97_DFS (BBCH 49), as shown in

Table 4. BBCH phenological scale of Trifolium pratense with days following sowing (DFS).

DFS	BBCH Code	Description
0	0	Germination—Dry seeds
7	9	Germination—Emergence of cotyledons break through soil surface
21	16	Leaf development—Six true leaf
34	30	Steam elongation- Start of steam elongation
42	33	Steam elongation—Three visible nodes
67	39	Steam elongation—nine or more visible nodes
76	43	30% of maximum biomass reached
90	47	70% of maximum biomass reached
97	49	90% of maximum biomass reached
105	53	Inflorescence emergence—Flower buds enlarged; petals visible
110	60	Flowering—Beginning of anthesis: First open flower
116	62	Flowering—20% of flowers open

. The reproductive phenological phase was observed after a delay of 2 weeks with respect to yarrow plants, showing inflorescence emergence at 105_DFS and the first open flowers after only 5 days (110_DFS). Trifolium pratense exhibited a constant trend, remaining in phase 4 of vegetative growth and thus prolonging the vegetative phase. In contrast, the species Achillea, despite being exposed to a temperature drop, did not show any slowdown in the progression of developmental stages. It completed its reproductive structures and initiated the flowering phase with the emergence of the first visible flowers. As for Trifolium pratense, an intermediate growth period was recorded compared to the other two studied species, lasting approximately 120 days from sowing to flowering and consistent with other studies on Trifolium [33], although this period is influenced by climatic conditions, which vary by year, climatic region, and sowing season.

The growth followed different patterns as each species reached its phenological stages at different times. In Figure 3, all the phenological trends of the three species studied can be observed. We observed that the two species belonging to the Asteraceae family (i.e., yarrow and milk thistle) exhibited similar behavior during the first two stages of their phenological development: Stage 0 (germination), which lasted for 7 days, and Stage 1 (leaf development), which lasted for 35 days for both species. A temperature accumulation of 848 GDD was required to complete phenological stage 1 for both Asteraceae species, whereas a lower accumulation of 573 GDD was sufficient for Trifolium pratense. For phenological stage 3, Achillea millefolium exhibited the lowest temperature accumulation, requiring 464 GDD, whereas Trifolium pratense necessitated a considerably higher GDD amount (1000 units), a very similar condition to that of Silybum marianum, where 1115 GDD were required. However, Trifolium pratense reached the stem elongation phenological stage (BBCH 3) earlier than the two Asteraceae species (characterized by composite flowers) and maintained this stage for 42 days. However, the Silybum marianum reached the rosette phenological stage (BBCH 3) at a later time, around day 42, and maintained this stage for a longer duration than the two other species, lasting 48 days as shown in Figure 3. During the phenological stage (BBCH 4), to complete the development of harvestable vegetative biomass, Silybum marianum required only a temperature accumulation of 325 GDD, while Achillea millefolium and Trifolium pratense necessitated higher accumulations amounting to 525 and 680 GDD, respectively.

3.3. Evaluation of the Total Antioxidant Capacity (TAC) of Hydroalcoholic Extracts

Based on the considerations outlined in Section 2, it was decided that the three assays FRAP, ABTS, and DPPH be applied to the eight extracts obtained from the corresponding samples of the three species under study. Each of the 24 extracts was analyzed in triplicate. The main insight from the analysis is that each plant species studied has an optimal set of cultivation conditions that can maximize the total antioxidant capacity of its hydroalcoholic extract. In Table S1, the results from ANOVA test were shown to assess the statistically significant differences among the parameters studied.

For Silybum marianum (SM), the hydroalcoholic extract with the highest total antioxidant capacity was obtained under conditions that included the presence of PGPR, no fertilization with P₂O₅ and K₂O, and full irrigation at field capacity (conditions SM101: FRAP = 39.39 ± 1.59 mg TE/g extract; DPPH = 31.50 ± 0.80 mg TE/g extract; ABTS = 45.55 ± 0.91 mg TE/g extract,

Table 5. Results of the measured FRAP, DPPH, and ABTS values of the extracts of plants obtained under different cultivation conditions. SM: Silybum marianum; AM: Achillea millefolium; TP: Trifolium pratense. The triplets of numbers refer to the applied cultivation conditions as reported in the text (Table 1). All data are reported as the mean value of three independent measurements, along with the corresponding standard deviation. Different letters (_{a b c d e f g}) indicate statistically different mean values (p ≤ 0.01; ANOVA one-way, Tukey’s HSD tests) from highest to lowest. For each assay and plant source, the highest and lowest values are reported in bold and italics, respectively.

Sample	Type of Assay
	FRAP (mg TE/g Extract)	DPPH (mg TE/g Extract)	ABTS (mg TE/g Extract)
SM000	22.94 ± 0.82 d	15.87 ± 0.27 f	40.37 ± 0.21 c
SM111	34.08 ± 1.46 b	27.20 ± 1.33 c	39.46 ± 1.91 c
SM011	34.27 ± 0.58 b	29.48 ± 0.08 b	41.67 ± 0.21 cb
SM001	31.50 ± 2.04 b	23.23 ± 0.04 d	40.64 ± 0.17 c
SM110	33.28 ± 1.80 b	26.15 ± 0.53 b	44.32 ± 1.54 ab
SM100	16.07 ± 1.28 e	9.34 ± 0.38 g	34.55 ± 1.62  d
SM101	39.39 ± 1.59 a	31.50 ± 0.80 a	45.55 ± 0.91  a
SM010	27.67 ± 0.83 c	19.49 ± 0.68 e	44.20 ± 0.08 bc
AM000	86.47 ± 3.85 bc	24.64 ± 0.43 d	49.90 ± 2.89 c
AM111	107.31 ± 3.66  a	29.46 ± 1.34 ba	64.74 ± 7.80  a
AM011	95.76 ± 5.31 b	26.79 ± 2.00 dcb	60.60 ± 0.04 ba
AM001	82.19 ± 3.72 cd	28.06 ± 0.18 ba	57.85 ± 1.23 ba
AM110	71.67 ± 0.06  e	27.65 ± 1.15 ba	60.09 ± 4.16 ba
AM100	88.52 ± 5.31 bc	24.77 ± 0.54 dc	55.92 ± 1.27 cb
AM101	75.79 ± 3.00 ed	27.44 ± 0.59 cba	60.60 ± 2.34 ba
AM010	81.04 ± 2.43 cd	30.02 ± 1.41  a	53.85 ± 5.07 cb
TP000	44.02 ± 4.67 a	9.57 ± 1.37 a	84.11 ± 0.67 a
TP111	29.14 ± 0.91 cb	7.51 ± 0.79 ba	65.13 ± 3.13 cb
TP011	26.39 ± 2.51 cb	6.81 ± 0.68 b	56.45 ± 5.11 dc
TP001	31.54 ± 1.73 b	8.30 ± 1.05 ba	34.28 ± 2.65  e
TP110	28.86 ± 1.44 cb	6.95 ± 0.61 b	63.40 ± 3.92 c
TP100	40.93 ± 1.15 a	9.60 ± 1.15  a	73.34 ± 6.73 b
TP101	28.93 ± 0.38 cb	6.51 ± 0.16  b	52.00 ± 2.69 d
TP010	24.83 ± 1.59  c	6.55 ± 1.14 b	37.95 ± 3.64 e

). In contrast, the extract with the lowest total antioxidant capacity was obtained from plants grown under similar conditions but with irrigation at 40% relative soil moisture (conditions SM100: FRAP = 16.07 ± 1.28 mg TE/g extract; DPPH = 9.34 ± 0.38 mg TE/g extract; ABTS = 34.55 ± 1.62 mg TE/g extract, Table 5). Notably, the generally higher values observed in the ABTS assay compared to the DPPH assay suggest that both polar and non-polar compounds contribute to the total antioxidant capacity of the hydroalcoholic extract, acting through both the single electron transfer and hydrogen atom transfer mechanisms. The significant contribution of compounds operating via the single electron transfer mechanism is further supported by the relatively high values obtained in the FRAP assay (Figure 4). Somewhat converging information regarding the optimization of the total antioxidant capacity of extracts from Achillea millefolium (AM) was obtained using the three spectrophotometric assays (Figure 4). Specifically, it appears that the antioxidant power of hydroalcoholic extracts of this species can be mostly enhanced in the presence of PGPR, with fertilization using P₂O₅ and K₂O, and under full irrigation at field capacity. For only the DPPH assay, the best combination was the absence of PGPR, the presence of fertilization, and a restricted irrigation regime at 40% of field capacity (condition AM111: FRAP = 107.31 ± 3.66 mg TE/g extract; ABTS = 64.74 ± 7.80 mg TE/g extract; AM010: DPPH = 30.02 ± 1.41 mg TE/g extract, Table 5). Moreover, findings indicate that the antioxidant capacity of the extract is predominantly driven by polar compounds, which exhibit their reducing power through a single electron transfer mechanism, as indicated by the exceptionally high values obtained with the FRAP assay. The significant contribution of hydrophilic species to the total antioxidant capacity of the extract is also supported by the higher values observed in the ABTS assay compared to those in the DPPH assay.

In analogy with what has already been stated for the other two species, the three spectrophotometric assays consistently suggested the optimal cultivation conditions for Trifolium pratense (TP) (condition TP000: FRAP = 44.02 ± 4.67 mg TE/g extract; DPPH = 9.34 ± 0.38 mg TE/g extract; ABTS = 84.11 ± 0.67 mgTE/g extract, Table 5); for the DPPH assay, the slightly better combination was the presence of PGPR, absence of fertilization, and a restricted irrigation regime (TP100: DPPH = 9.60 ± 1.15 mg TE/g extract; TP000: DPPH = 9.57 ± 1.37 mg TE/g extract). In this case (conditions TP100), the hydroalcoholic extract showed that the highest total antioxidant capacity was obtained by cultivating the plant in the absence of PGPR, without fertilization with P₂O₅ and K₂O, and with irrigation at 40% relative soil moisture. A more detailed analysis of the results reveals that the species contributing to the antioxidant power of the hydroalcoholic extracts of Trifolium pratense are primarily hydrophilic in nature, as suggested by the higher values provided by the ABTS assay compared to those of the DPPH assay, as shown in Figure 4. Supporting this hypothesis further are the values obtained from the FRAP assay, which, as mentioned earlier, particularly responds to hydrophilic species. In terms of the mechanism of action, the results for the Trifolium pratense extracts seem to indicate that the species present act through both single electron transfer and hydrogen atom transfer mechanisms.

It is noteworthy that in Silybum marianum, all three spectrophotometric assays showed that the key parameter for optimizing the antioxidant properties of the hydroalcoholic extract is the irrigation level, indicating the presence/absence of water stress. Also, in Achillea millefolium species, the results from the spectrophotometric assays consistently indicate that water stress negatively affects the quality (in terms of antioxidant power) of the hydroalcoholic extract. The findings of this study do not reveal significant effects on antioxidant capacity due to fertilizer application in two out of the three species examined. However, in the case of Achillea millefolium, the addition of fertilizer resulted in an increase in antioxidant capacity. This observation aligns with the existing literature, which suggests that potassium application significantly influences the phenolic composition and antioxidant capacity of medicinal plants by enhancing potassium levels in plant leaves [6]. Potassium functions as an activator for enzymes involved in both photosynthesis and the biosynthesis of starch and proteins. Furthermore, higher potassium fertilization is associated with improved plant growth [34].

The analysis carried out in the Silybum marianum assays with only PGPR showed low values of total antioxidant capacity, probably due to an induction of systemic tolerance in plants towards drought stress through a variety of mechanisms like improvement of the antioxidant system [35,36]. A significant portion of plant damage under abiotic stress results from oxidative injury at the cellular level, caused by the production of reactive oxygen species (ROS) and their insufficient detoxification [37,38]. To counteract this, plant cells produce various antioxidant enzymes, such as catalase (CAT), peroxides (POX), superoxide dismutase (SOD), which neutralize reactive free radicals or inhibit their formation [39]. Notably, PGPR inoculation has been shown to enhance the activity of these antioxidant enzymes under severe drought conditions, suggesting its potential use in inoculants to mitigate oxidative damage induced by drought stress [40].

3.4. Ultra High-Performance Liquid Chromatography Coupled to Mass Spectrometry (UHPLC-MS/MS) Analysis of the Three Extracts SM101, AM111, and TP000

The three investigated extracts may potentially exhibit a broad spectrum of biological activities that deserve a more comprehensive investigation. Indeed, flavonoids such as apigenin, luteolin, and kaempferol are well documented for their potent antioxidant, anti-inflammatory, and anticancer properties [41], while rutin and quercitrin contribute to vascular protection [42] and immune modulation [43]. Phenolic acids, including gallic and caffeic acid, demonstrate antimicrobial, hepatoprotective, and neuroprotective effects [44,45]. Chlorogenic acid and cynarin (and, more generally, dicaffeoylquinic acid derivatives) are frequently associated with hypolipidemic and hepatoprotective activities [46,47], notably through the enhancement of bile flow and lipid metabolism.

The concomitant presence of these compounds suggests the potential for synergistic effects in mitigating oxidative stress, modulating inflammatory responses, and preventing chronic diseases. Based on these findings, it is plausible to hypothesize that this phytochemical profile underpins the extracts’ potential application in nutraceutical and therapeutic contexts aimed at promoting overall health and preventing metabolic or degenerative disorders.

Detailed information on the results of the UHPLC-MS/MS analysis are shown in

Table 6. Results of the MS and MS2 analysis of the extract SM101. Compounds are listed in alphabetical order. All compounds were identified as [M-H]- ions.

Possible Candidate	Formula	Rt (min)	m/z	MS/MS Fragments (m/z)	Exp Mass	Detected Mass	Score	Error (in ppm)
Apigenin	C15H10O5	12.07	269.03	117.03	270.0528	270.0528	89.56	1.88
Chlorogenic Acid	C16H18O9	10.58	353.0883	191.0548, 137.02	354.3100	354.0939	98.81	2.19
Chrysin	C15H10O4	11.29, 12.20	253.059		254.2400	254.0583	88.52	−3.10
Cinnamic Acid	C9H8O2	8.28	147.04		148.1586	148.0524	99.77	0.90
Coumaric Acid	C9H8O3	5.44, 6.53	163.0405	119.05	164.1600	164.0473	99.84	0.41
Cynarin	C25H24O12	10.58	515.27		516.4500	516.1280	96.53	2.43
Ferulic Acid	C10H10O4	7.15	193.01	117.03, 134.03, 175.78	194.1800	194.0579	86.00	2.48
Fumaric Acid	C7H6O3	2.75	137.02	93.03	138.1200	138.0318	85.96	0.45
Hydroxybenzoic Acid	C7H6O3	3.19	137.02	93.0329	138.1200	138.0317	99.34	−0.79
Kaempferol (or luteolin)	C15H10O6	11.38	285.0411	184.23	286.2300	286.0477	98.44	1.01
Malic Acid	C4H6O5	5.56	133.1442	115.0013	184.8400	134.0215	96.53	5.07
Naringenin	C15H12O5	12.03	271.062	151.41	272.2500	272.0685	99.57	0.22
Protechatecuic Acid	C7H6O4	5.01	153.0181	109.02	154.1200	154.0266	86.06	−2.14
Quercetin	C15H10O7	10.18	301.0362	151.0037	302.2360	302.0427	82.88	3.31
Quercitrin	C21H20O11	10.68	447.0946	284.0326, 255.02	448.3800	448.0070	98.85	1.07
Quinic Acid	C7H12O6	5.48, 7.53	191.0563		192.1700	192.0635	99.46	1.38
Rutin	C27H30O16	10.21	609.14	300.1	610.5200	610.1543	89.52	−3.46

,

Table 7. Results of the MS and MS2 analysis of the extract AM111. Compounds are listed in alphabetical order. All compounds were identified as [M-H]- ions.

Possible Candidate	Formula	Rt (min)	m/z	MS/MS Fragments (m/z)	Exp Mass	Detected Mass	Score	Error (in ppm)
Apigenin	C15H10O5	12.27	269.03	225	270.0528	270.0528	96.79	1.27
Caffeic Acid	C9H8O4	6.34	179.034	135.179	180.1600	180.0423	99.48	1.44
Chlorogenic Acid	C16H18O9	7.74, 9.74, 10.63	353.0883	191.0548	354.3100	354.0939	98.55	1.66
Chrysin	C15H10O4	11.29, 12.19	253.059		254.2400	254.0583	99.53	0.53
Cinnamic Acid	C9H8O2	10.73, 11.65	147.04		148.1586	148.0524	97.93	3.54
Coumaric Acid	C9H8O3	5.44, 6.53	163.0405	119.05	164.1600	164.0473	98.54	0.07
Fumaric Acid	C7H6O3	3.26	137.02	93.03	138.1200	138.0318	99.83	0.30
Gallic Acid	C7H6O5	1.18	169.0138		170.1200	170.0215	95.83	−0.91
Hydroxybenzoic Acid	C7H6O3	3.19	137.02	93.0329	138.1200	138.0317	98.85	0.59
Kaempferol	C15H10O6	11.53	285.0411	184.23	286.2300	286.0477	85.64	0.30
Malic Acid	C4H6O5	5.92	133.1442	115.87	184.8400	134.0215	97.50	3.07
Naringenin	C15H12O5	10.67	271.062	317.0667, 331.0823	272.2500	272.0685	96.99	2.53
Protechatecuic Acid	C7H6O4	4.97	153.0181	109.02	154.1200	154.0266	86.64	−0.44
Quercitrin	C21H20O11	10.30	447.0946	284.0326, 255.02	448.3800	448.0070	97.02	2.93
Quinic Acid	C7H12O6	5.56	191.0563	85.03	192.1700	192.0635	99.81	0.78
Rutin	C27H30O16	10.16	609.14	300.1	610.5200	610.1543	97.07	2.11

and

Table 8. Results of the MS and MS2 analysis of the extract TP000. Compounds are listed in alphabetical order. All compounds were identified as [M-H]- ions.

Possible Candidate	Formula	Rt (min)	m/z	MS/MS Fragments (m/z)	Exp Mass	Detected Mass	Score	Error (in ppm)
Apigenin	C15H10O5	12.06	269.15	117.03	270.2300	270.0528	98.52	2.13
Apigetrin	C21H20O10	12.10	431.0995		432.3810	432.1267	97.02	2.41
Caffeic Acid	C9H8O4	8.58	179.034	135, 179	180.1600	180.0423	99.84	0.56
Chrlorogenic Acid	C16H18O9	9.74	353.0883	191.05, 137.02	354.3100	354.0939	98.81	−2.78
Chrysin	C15H10O4	11.28	235.059		254.2400	254.0583	98.24	1.69
Cinnamic Acid	C9H8O2	10.73, 11.65	147.04		148.1586	148.0524	99.04	0.52
Coumaric Acid	C9H8O3	5.44, 10.13	163.0405	119.05	164.1600	164.0473	99.37	0.41
Ferulic Acid	C10H10O4	10.24	193.01	105.46, 106.04, 117.03, 134.03, 175.78	194.1800	194.0579	99.30	1.70
Fumaric Acid	C7H6O3	2.75	137.02	93.03	138.1200	138.0318	99.91	0.43
Gallic Acid	C7H6O5	1.63	169.0138	127.002	170.1200	170.0215	95.83	−0.91
Hydroxybenzoic Acid	C7H6O3	3.18	137.02	93.0329	138.1200	138.0317	99.91	0.43
Kaempferol (or lueolin)	C15H10O6	11.38	285.0411	184.23	286.2300	286.0477	85.64	0.30
Malic Acid	C4H6O5	8.78	133.1442	115.0013	184.8400	134.0215	99.86	1.10
Naringenin	C15H12O5	12.63	271.062	151.41	272.2500	272.0685	97.61	2.39
Protechatecuic Acid	C7H6O4	4.92	153.0181	109.02	154.1200	154.0266	99.13	−1.00
Quercetin	C15H10O7	11.60	301.0362	151.0037, 135.65	302.2360	302.0427	93.68	2.14
Quercitrin	C21H20O11	10.61	447.0946	284.0326, 255.02	448.3800	448.0070	96.46	0.28
Quinic Acid	C7H12O6	6.18	191.0563		192.1700	192.0635	81.90	−6.78
Rutin	C27H30O16	10.14	609.14	300.1	610.5200	610.1543	97.91	1.50
Vanillic Acid	C8H8O4	6.87	167.0348		168.1400	168.0420	86.42	−0.87

.

4. Conclusions

The results of the present study clearly demonstrate the importance of implementing specific cultivation management practices for MOWP to enhance the TAC of plant extract, with potential inherent health benefit implications. Accordingly, this study highlighted that, for Silybum marianum, the hydroalcoholic extract exhibiting the highest TAC was obtained under conditions involving the application of PGPR, no fertilization with P₂O₅ and K₂O, and full irrigation at field capacity (condition SM101). In the case of Achillea millefolium, the antioxidant activity of its hydroalcoholic extracts was maximized under conditions that included PGPR inoculation, fertilization with P₂O₅ and K₂O, and full irrigation at field capacity (condition AM111). Regarding Trifolium pratense, the highest total antioxidant capacity of the hydroalcoholic extract was observed when the plants were cultivated without PGPR, without P₂O₅ and K₂O fertilization, and under limited irrigation at 40% relative soil moisture (condition TP000).

Further improvements could be attained by adjusting the harvesting periods to different plant development phases or by exploring alternative extraction techniques, such as ultrasound or microwave-assisted extraction, instead of the traditional maceration method. In this context, the application of Design of Experiments (DoE) protocols could help identify optimal extraction conditions and pinpoint critical process parameters. As a further progression of the current study, the three most promising extracts will be evaluated for their specific biological properties, with a particular focus on their potential effects in relation to lipid dysmetabolism.