Fermented Nettles: Bioactive Profile and Seasonal Variability

Abstract

In traditional horticulture, fermented nettles (FN) enhance plant growth and resilience. However, their precise mode of action remains unclear. This study aims to characterize the bioactive profile of FN and to evaluate their potential as biostimulants beyond organic fertilizers. For this purpose, FN samples were prepared from Urtica dioica L. harvested in different seasons and analyzed by mass spectrometry (ICP-MS, LC-MS/MS, and GC×GC-MS), electrophoresis, and spectrophotometry to quantify macro- and micronutrients, nitrogen compounds, phytohormones, antioxidant capacity, enzyme activities, and microbial viability. The results show that FN are rich in essential nutrients (N, K, Ca, Fe, and Zn), hydrolytic enzymes (proteases, glycosidases and phosphatases), and phytohormones (auxins, cytokinins, gibberellins, abscisic acid, and salicylic acid). FN contain volatile compounds with potential antimicrobial effects, in addition to strong antioxidant properties. The monitored parameters support the dual role of FN as fertilizers and biostimulants, suggesting that FN act synergistically through nutrient enrichment, enzymatic degradation of macromolecules, hormonal signaling, and microbial priming. Based on our data, particularly because of the highest microbial viability and enzyme activities, the summer FN seem like the most suitable option. Moreover, the seasonal variability in composition highlights the importance of timing the harvest to optimize FN efficacy in sustainable agriculture.

1. Introduction

In a period of climate change, tackling arable land loss due to anthropogenic activities and demographic pressure requires concerted efforts to improve the quality and nutritional value of crops. There is a pressing need to reduce reliance on the excessive use of inorganic fertilizers and pesticides, which can lead to salinization, acidification, adverse effects on microbial communities, and development of pest resistance. Therefore, sustainable agriculture increasingly emphasizes the adoption of environmentally friendly cultivation practices, including the use natural fertilizers, green manure, and biostimulants to enhance crop growth, health, and resilience [1,2,3].

In traditional medicine, nettles (Urtica dioica L.) have long been used for their high content of many useful substances. Nettle is rich in a number of minerals, such as Ca, Fe, Mg, K, and P, but also contains, to a lesser extent, Mn, Cu, Zn, B, and Se [4,5]. The biochemical profile of nettles includes phenolics, flavonoids, and coumarins, in addition to proteins and amino acids, whose levels vary with the plant and cultivation site, while the choice of extraction method influences the accuracy and completeness of their detection [6,7].

Among phenolic acids, chlorogenic, quinic, p-coumaric, and caffeic acids are the most abundant, while flavonoids that are detected in lower concentrations include quercetin, kaempferol, kaempferol 3-O-glucoside, naringin, isorhamnetin-3-O-glucoside, isorhamnetin-3-O-rutinoside, apigenin-7-O-glucoside, daidzein, and chrysin [6,7,8]. Nettles also contain high amounts of pigments, such as chlorophyll a and b, as well as nine carotenoids, mainly lutein and β-carotene [5]. Fresh nettle leaves, in particular, show significant levels of vitamins A, C, D, E, K, and B complex [7,9], with polysaccharides, cellulose, and hemicellulose accounting for most carbohydrates. Because many of these substances have antioxidant properties, nettles display considerable antioxidant capacity [5], with additional bioactive effects. For example, the antibacterial activity of nettles against Gram-positive and Gram-negative bacteria, such as Escherichia coli, Salmonella spp., Staphylococcus aureus, Klebsiella pneumoniae, and Enterococcus faecalis, was observed.

Nettles are used in a variety of ways, from direct culinary processing to water extracts used in teas or alcohol extracts such as tinctures [4,9,10]. Nettle tea, similar to mint tea, was used in a fermentation process involving symbiotic bacteria and yeast (kombucha), resulting in a new mixture of phenolic compounds with potentially new properties [11,12]. Other applications include cosmetics and textile fiber due to the high quality of cellulose [10]. Nettles are also used as an additive in animal feed and in veterinary medicine [4,10].

Nettles are widely employed in organic farming as well, in the form of green manure, as fermented nettles (FN) [13,14,15]. The method of preparation and naming of FN (nettle water, aqueous nettle extract, nettle slurry, nettle manure, nettle fertilizer) varies among authors [14,15,16,17]. The first report on FN comes from Peterson and Jensén (1985) [15], who conducted the fermentation process for 14 days at 20 °C and compared fresh and pre-dried nettles. At the same time, they monitored the effect of the age of the nettles on the final product, in which they found N, P, K, Ca, Mg, S, B, Mn, Zn, Cu, and Mo and a relatively high content of Fe. They also documented high content of nitrogen, mostly as ammonium [15]. These substances indicate that FN may function as an organic fertilizer and a source of other nutrients in soils where such elements are deficient. In addition to that, FN contain significant amounts of bacteria, which can also promote plant growth. The presence of the phytohormone auxin was detected as well. Together, these results pointed to the biostimulatory properties of FN [15]. Maričić et al. (2021) [14] prepared a short-term extract macerated for 24 h and a long-term extract macerated for 14 days, which they applied to green beans with resulting positive effects on vegetative parameters. FN are also commercially available. Despite considerable interest in FN, the underlying mechanism of action remains poorly understood, limiting our ability to leverage their full potential in agricultural applications. To bridge this knowledge gap, it is essential to clarify the composition of nutrients and specific bioactive compounds in FN, including their effects on the plant metabolic pathways, to understand the role of microorganisms and to evaluate the overall effect on the environment. The composition of nettles can vary significantly depending on factors like harvest time, soil quality, temperature, water regime, sunshine, and probably also the content of microorganisms, and thus the composition of the final product after fermentation. We hypothesize that the composition of nettle plants will depend on the season, so the resulting FN may vary throughout the year. In this context, the present study reports a detailed analysis of FN by harvest season, assessing their content of (i) nitrogen in its various forms, (ii) microbial activity, (iii) degradation enzymes converting proteins, polysaccharides and other nutrients into simpler compounds, (iv) volatile compounds with antimicrobial properties, (v) phytohormones, (vi) antioxidant properties, and (vii) macro- and micronutrients. The resulting data improve our understanding of the role of FN as both an organic fertilizer and a biostimulant.

2. Materials and Methods

2.1. Preparation of Fermented Nettles

Approximately two-month-old shoots of nettles (Urtica dioica L.) naturally grown at Albertov, Prague 2, Czech Republic (50.069507, 14.426623) and with approximately the same height were cut and steeped in tap water at a 1:10 ratio (fresh nettle weight to water volume, e.g., 500 g of nettles in 5 L of tap water). The mixture was left to ferment for 2–3 weeks in an open container in a plant growth chamber under controlled conditions (24 °C, 16/8 h day/night cycle). Every day of fermentation, the mixture was thoroughly stirred. Fermentation was deemed complete once the leaves were decomposed and foaming had stopped. The mixture was then filtered through gauze, and FN were lyophilized overnight on a Lyovac GT2E (Finn-Aqua, Tuusula, Finland). A portion of FN was stored at 4 °C for further analysis. Unless specified otherwise, all experiments presented in the main text were performed with FN lyophilizate.

2.2. Total Nitrogen, Carbon, and Sulfur Content

The nitrogen (N), carbon (C) and sulfur (S) concentrations were determined by controlled sample combustion and analysis of the resulting products by gas chromatography with a thermal conductivity detector on Flash FA 1112 CHNS/O analyzer (Thermo Scientific, Waltham, MA, USA).

2.3. Free Amino Acids, Primary Amines and Inorganic Ions

Free amino acids were extracted in hydrochloric acid and determined by capillary electrophoresis with a contactless conductivity detector, enabling the separation of 20 proteinogenic amino acids in an acidic background electrolyte [18]. The experiments were conducted in a fused-silica capillary (Polymicro Technologies, Phoenix, AZ, USA) using a G7100A Capillary Electrophoresis System (Agilent Technologies, Santa Clara, CA, USA) with a contactless conductivity detector. The detector consisted of two cylindrical electrodes, each of which is 4 mm in length, separated by a 1 mm insulation gap. The inner diameter of the electrodes was 400 μm.

Primary amines were quantified in 96-well plates using a standard addition method based on Darrouzet-Nardi et al. [19]. FN (50 mg) were extracted with 1.0 mL 2 M KCl and incubated for 1 h at 28 °C under orbital shaking at 1200 min⁻¹ (ThermoMixer^® C, Eppendorf, Hamburg, Germany). Prepared in different dilutions, FN samples (10 μL) were mixed with 10 μL of alanine standard solution at concentrations of 0, 10, or 100 μM before adding 180 μL of the OPAME reagent (25 mg o-phthalaldehyde and 50 μL of β-mercaptoethanol in 200 mM borate buffer, pH 9.0). After 1 min of incubation, fluorescence was measured (excitation at 340 nm, emission at 455 nm) on an Infinite M200 plate reader (Tecan, Zürich, Switzerland). The fluorescence values of the samples were subjected to linear regression with varying standard concentrations, and the concentration of primary amino groups was determined from the x-intercept of the extrapolated line. For each sample, the lowest dilution yielding a regression slope comparable to that of the blank was selected to ensure accurate quantification.

Inorganic ions were analyzed on an Agilent 7100 capillary electrophoresis system (Agilent Technologies, Waldbronn, Germany), equipped with a TraceDec contactless conductivity detector (Innovative Sensor Technologies, Strasshof an der Nordbahn, Austria), using an unmodified fused-silica capillary (Polymicro Technologies, Phoenix, AZ, USA), 20 µm i.d., 375 µm o.d., 80.0 cm total, and 65.0 cm effective length. Samples were injected at a pressure of 5 kPa for 20 s. The cassette temperature was 25 °C. Ammonium cations were determined using 1.5 M HCOOH with 4 mM 18-crown-6 ether as a background electrolyte and 2 mM CsCl as an internal standard and a voltage of 30 kV. Nitrate and sulfate were separated in a 1 M HCOOH background electrolyte at −30 kV voltage; 2 mM KClO₃ was added as an internal standard.

2.4. Protein and Peptide Content

In appropriately diluted FN samples, total saccharide and protein concentrations were determined spectrophotometrically at 490 nm and at 595 nm to 450 nm ratio, respectively [20,21]. To this end, FN (50 mg) were dissolved in 1 mL distilled water and incubated for 1 h at room temperature under continuous shaking at 275 min⁻¹ (Incubator shaker ES-60, Hangzhou Miu Instruments Co. Ltd., Hangzhou, China) at room temperature. Then, to determine the distribution of proteins’ molecular weights, the solution was centrifuged (5000× g, 10 min). The resulting supernatant or liquid FN samples were diluted with sample buffers to perform 12 and 16% (w/v) Tris-glycine-SDS-PAGE and Tricine-SDS-PAGE [22,23], respectively. After electrophoresis (70/140 V, 8 °C), the proteins were visualized by silver staining. Unless otherwise specified, all steps were conducted at room temperature. The gels were immersed in fixation solution (50% (v/v) ethanol, 12% (v/v) acetic acid, 0.05% (v/v) formaldehyde) for 2 h. Then, the gels were washed twice with 20% (v/v) ethanol (20 min), briefly sensitized with 0.02% (w/v) sodium thiosulfate solution (2 min), and soaked in chilled impregnating solution (0.2% (w/v) silver nitrate, 0.076% (v/v) formaldehyde, 20 min). Subsequently, the gels were rinsed with distilled water and briefly immersed in development solution (6% (w/v) sodium carbonate, 0.0004% (w/v) sodium thiosulfate, 0.05% (v/v) formaldehyde). This solution was replaced with a fresh solution, in which the gels were incubated until the desired staining intensity was achieved. Development was terminated with 12% (v/v) acetic acid solution.

2.5. Microbial Activity Detection

Cell viability was determined using the resazurin test. Briefly, FN (10 mg) were incubated at 30 °C for 10 min in 200 µL phosphate-buffered saline (pH 7.2) with 20 µL 1 mM resazurine under orbital shaking (900 min⁻¹). After centrifugation (2 min at 12,100× g), the fluorescence of the supernatant, with excitation at 540 nm, emission at 590 nm, was measured in 96-well black plate fluorescence on Infinite M200 (Tecan, Zürich, Switzerland), according to Petiti et al. [24].

Total dehydrogenase activity was measured using a modified assay based on the ISO 23753-2:2019 standard method [25] and Babson and Phillips [26]. In test tubes, FN (10 mg) were incubated with 5 μL of a 1 mg·mL⁻¹ solution of 5-methylphenazinium methyl sulfate (PMS) in 100 mM Tris-HCl buffer (pH 7.6) and with 1.0 mL of a 15 mM substrate suspension of 2-(4-iodophenyl)-3-(4-nitrophenyl)-5-phenyltetrazolium chloride (INT) in the same buffer. For blank samples, the INT solution was replaced by 1.0 mL of 100 mM Tris-HCl buffer (pH 7.6). After incubation for 5 min at 30 °C under orbital agitation (180 min⁻¹) in the dark, the resulting product, 1-(4-iodophenyl)-5-(4-nitrophenyl)-3-phenylformazan (INTF), was extracted by shaking the mixture with 4.0 mL of acetone for 5 min at 30 °C, at 180 min⁻¹ in the dark. Following 5 min centrifugation (3000× g, 10 °C, Universal 32R, Hettich, Tuttlingen, Germany), the supernatant absorbance was measured at 485 nm against the corresponding blank. The control sample of method validity was measured with a sterilized FN, which showed no total dehydrogenase activity. Measurements were conducted in a glass cuvette using a Helios Alpha UV/Vis Spectrophotometer (Thermo Scientific, Waltham, MA, USA).

2.6. Hydrolytic Enzyme Activities

The activity of the exoglycosidases α-glucosidase (EC 3.2.1.20), β-glucosidase (EC 3.2.1.21), α-galactosidase (EC 3.2.1.22), β-galactosidase (EC 3.2.1.23), α-mannosidase (EC 3.2.1.24), and β-N-acetylhexosaminidase (EC 3.2.1.52) was determined using the end-point spectrophotometric method with specific p-nitrophenyl substrates at 405 nm in McIlvaine buffer, pH 6.0, according to Bělonožníková et al. [27]. The activity of the endoglycosidases β-1,3-glucanase, chitinase, cellulase, and α-amylase was determined using a slightly modified version of the 3-methyl-2-benzothiazolinone hydrazone (MBTH) test [28], as described by Bělonožníková et al. [27].

Total proteolytic activity was spectrophotometrically determined at 450 nm using azocasein substrate [29] and detected in gel [30]. Briefly, FN (30 mg) were extracted for 30 min at 37 °C with 1.0 mL of extraction buffer (63 mM Tris-HCl (pH 6.8), 10% (v/v) glycerol, 2% (w/v) sodium dodecyl sulfate, and 0.0013% (w/v) bromophenol blue), under orbital shaking (900 min⁻¹). Liquid FN samples were mixed 1:1 with the extraction buffer and incubated under the same conditions. Electrophoresis was performed in a 10% resolving acrylamide gel containing 0.12% (w/v) gelatine and a 4% (w/v) stacking gel mixture [22,30]. The initial voltage was set to 70 V and then increased to 140 V until the bromophenol blue reached the end of the resolving gel. Subsequently, the gels were rinsed in deionized water and then immersed and shaken in 2.5% (v/v) Triton X-100 for 1 h before rinsing with reaction buffer (50 mM Tris-HCl, pH 8.4, with 5 mM calcium chloride) twice. Following overnight incubation at 37 °C in the same buffer, the gels were briefly washed with deionized water and then stained in Coomassie Brilliant Blue solution (0.1% (w/v) Coomassie Brilliant Blue R-250 in 40% (v/v) methanol and 10% (v/v) acetic acid) for 2 h. Finally, the gels were soaked in a destaining solution (10% (v/v) acetic acid in 25% (v/v) ethanol) until bands were destained [30,31]. The extent of proteolysis was determined from a gel photograph using GelAnalyzer version 19.1 [32].

Acid (EC 3.1.3.2) and alkaline phosphatase (EC 3.1.3.1) activity was determined with p-nitrophenyl-phosphate in 200 mM citrate buffer, pH 4.5, and 200 mM borate buffer, pH 9.0, respectively, according to Maseko and Dakora [33].

Urease (EC 3.5.1.5) was determined using the phenol-hypochlorite assay with ammonium sulfate as a standard [34].

2.7. Phytohormone Analysis

Lyophilizates (5 mg) were resuspended in 100 μL 1 M formic acid, internal stable isotope-labeled standards were added, and samples were vortexed thoroughly. Isotope-labeled standards were added at 1 pmol per sample: ¹³C₆-IAA (Cambridge Isotope Laboratories, Tewksbury, MA, USA); ²H₄-SA (Sigma-Aldrich, Saint Louis, MO, USA); ²H₃-PA, ²H₃-DPA (NRC-PBI, Gatineau, QC, Canada); ²H₆-ABA, ²H₅-JA, ²H₅-tZ, ²H₅-tZR, ²H₅-tZRMP, ²H₅-tZ7G, ²H₅-tZ9G, ²H₅-tZOG, ²H₅-tZROG, ¹⁵N₄-cZ, ²H₃-DZ, ²H₃-DZR, ²H₃-DZ9G, ²H₃-DZRMP, ²H₇-DZOG, ²H₆-iP, ²H₆-iPR, ²H₆-iP7G, ²H₆-iP9G, ²H₆-iPRMP, ²H₂-GA₁₉, (²H₅)(¹⁵N₁)-IAA-Asp, and (²H₅)(¹⁵N₁)-IAA-Glu (Olchemim, Olomouc, Czech Republic). All samples were centrifuged for 10 min, at 4 °C, 30,000× g. The supernatant was applied to an SPE Oasis HLB 96-well plate pre-washed with 100 μL of 100% acetonitrile, followed by 100 μL distilled water and 100 μL 1 M HCOOH. The pellet was re-extracted with 100 μL 1 M HCOOH, re-homogenized in FastPrep-24 and centrifuged, the supernatant was applied to the same 96-well, and the wells were washed with 100 μL distilled water. To a collection plate, the samples were eluted twice by 50 μL 50% (v/v) acetonitrile and analyzed on a LC/MS system consisting of a UHPLC 1290 Infinity II (Agilent, Santa Clara, CA, USA) coupled to a 6495 Triple Quadrupole Mass Spectrometer (Agilent, Santa Clara, CA, USA), operating in MRM mode. The data were quantified by the isotope dilution method and processed with Mass Hunter software B.08 (Agilent, Santa Clara, CA, USA).

2.8. Volatile Metabolite Detection

Comprehensive two-dimensional GC×GC–MS measurements were performed on a GCMS QP2010 Ultra gas chromatograph coupled to a quadrupole mass spectrometer (Shimadzu, Kyoto, Japan) using a two-column system. The first DB-5 column (30 m × 0.25 mm, 0.25 μm; Restek, Bellefonte, PA, USA) was connected to the second SolGel-Wax column (3 m × 0.1 mm, 0.1 μm; Restek, Bellefonte, PA, USA) via a 3 m × 0.1 mm capillary. Helium, at a linear velocity of 20.0 cm·s⁻¹, served as a carrier gas. Lyophilized FN samples were extracted with methanol for 1 h at 30 °C under continuous shaking 900 min⁻¹. After centrifugation (16,000× g) for 10 min (Centrifuge Eppendorf, Hamburg, Germany), the upper phase was filtered through a 0.22 µm filter. For these extracts, 1 μL injections were performed in split mode (1:50). Liquid FN samples were extracted by solid-phase microextraction (SPME) of the headspace. For SPME, 200 μL of samples was placed in a 5 mL glass vial and heated in a 65 °C water bath for 5 min, and the volatiles were then extracted from the headspace for 10 min using 85 μm Carboxen/PDMS SPME fiber (Supelco, St. Louis, MO, USA). SPME injections were performed in splitless mode for 1 min. The injector temperature was 250 °C. The column temperature program was as follows: initial isotherm at 40 °C for 10 min, ramp to 180 °C at 5 °C per min and, then to 280 °C at 20 °C per min, with a final isotherm for 30 min. Mass spectrometry parameters were 70 eV electron ionization, 200 °C ion source temperature, 200 °C interface temperature, and 33–400 mass range m/z, acquired in scan mode. Cryogenic modulation with liquid nitrogen was applied, with a modulation period of 5 s throughout the whole measurement. Chromatograms were processed using GC Image software (version R3, 2022), and the measured spectra were compared with the NIST 20 Mass Spectral Library.

2.9. Chlorophyll and Carotenoid Contents

FN lyophilizates (5 mg) were extracted with pure ice-cold methanol (1 mL) under continuous shaking (900 min⁻¹) for 15 min at 10 °C. After centrifugation for 2 min at 4 °C (Centrifuge Eppendorf, 16,000× g), chlorophyll a and b were determined based on their absorption coefficients and absorbances at 470 nm, 652 nm, and 665 nm, while carotenoids were estimated by absorbance measurement at specific wavelengths—β-carotene (449 nm), lutein (444 nm), violaxanthin (438 nm), and neoxanthin (436 nm) [35,36]. All measurements were carried out on a Multiskan GO (Thermo Scientific, Waltham, MA, USA). Chlorophyll and total carotenoids contents were calculated based on Lichtenthaler [36].

2.10. Phenolics and Flavonoids

The total phenolic content was determined in a microtiter plate. A 20 µL aliquot of appropriately diluted FN was added to 100 µL of 10% (v/v) Folin–Ciocalteu reagent. After 4 min of incubation at room temperature, 80 µL of 75 g·L⁻¹ (w/v) sodium carbonate solution was added and absorbance at 760 nm was measured after 30 min on Multiskan GO (Thermo Scientific, Waltham, MA, USA) [37]. The flavonoids were determined by monitoring the formation of a complex with AlCl₃ according to Liu et al. [38]. In a microtiter plate, diluted FN (25 µL) were incubated with 110 µL of a 0.45% (w/v) sodium nitrite solution for 6 min. Then 15 µL of 100 mM AlCl₃ and 100 µL of 0.5 M NaOH were added, and absorbance was measured at 510 nm after 5 min on the same spectrophotometer.

2.11. Antioxidant System

Ferric reducing antioxidant power assay (FRAP) and antioxidant capacity using either 2,2-diphenyl-1-picrylhydrazyl (DPPH) or ABTS (2,2′-azinobis(3-ethylbenzothiazoline-6-sulfonate)) were measured according to Tupec et al. [39]. The activity and isoform content of peroxidase (EC 1.11.1.7), ascorbate peroxidase (EC 1.11.1.11), superoxide dismutase (EC 1.15.1.1), and laccase (EC 1.10.3.2) in liquid FN were detected after 12% (w/v) native PAGE [40,41].

2.12. Determination of Macro- and Microelements

Approximately 0.3 g of homogenized samples of FN in Teflon digestion vessels (DAP-60) were treated with 6 mL of HNO₃ at room temperature. After 24 h, the vessels were sealed and transferred to a microwave digestion system (SpeedWave^® XPERT, Berghof, Berlin, Germany). The digestion program consisted of heating to 200 °C at a rate of 5 °C per min, with a hold time of 30 min at the maximal temperature before cooling down. Prior to analysis, the digests were diluted with distilled water to the desired volume.

The concentrations of selected elements (Na, K, Mg, Ca, Mn, Fe, Cu, Zn, Mo, B, Si, and Se) were determined by ICP-MS on an Agilent 7900 ICP-MS (Agilent Technologies, USA) equipped with a Micromist glass nebulizer. The instrument was operated at 1550 W RF power and 1.03, 0.90, and 15.0 L per min gas flow for the nebulizer, auxiliary, and plasma gases, respectively. All measurements were performed in “no-gas” mode, using Sc, Y, and Bi as internal standards.

Calibration standards were prepared by the appropriate dilution of two multi-element certified reference solutions (AN9094MFN: Cu, Fe, Mn, Mo, Se, Zn; MC9091MN: Ca, K, Na, Mg; both 100 mg·L⁻¹ per element) and two single-element certified reference solutions (AN8053F: Si; AN8005H: B; both 100 mg·L⁻¹; Analytika, Prague, Czech Republic). The final calibration range for all elements was 0.0016–5.0 mg·L⁻¹.

Phosphates were determined according to Parre et al. [42]. Briefly, 100 μL of FN, or Na₂HPO₄ as a standard, were added to the 700 μL of distilled water, 200 μL of 100 mM ammonium heptamolybdate in 16% (v/v) H₂SO₄, and 200 μL of 1 mM malachite green in 2% (w/v) polyvinyl alcohol. After 30 min incubation at room temperature, absorbance was read at 623 nm on Multiskan GO (Thermo Scientific, Waltham, MA, USA).

2.13. Statistics and Data Processing

The results presented are based on the analysis of FN samples that were prepared and thoroughly analyzed within a single year. Nettles were harvested in large quantities (ca 200–700 g) in April (Spring), May (Spring/summer), July (Summer), and September (Autumn). Each analysis was performed in triplicate. All data were subjected to the Shapiro–Wilk normality test. One-way analysis of variance (ANOVA) was performed, followed by the Holm–Sidak post hoc test for multiple comparisons (α = 0.05). When the data did not meet the assumptions of normality, a Kruskal–Wallis one-way ANOVA on ranks was applied. Statistical analysis was performed in SigmaPlot 12.5. For all gels, band intensities were determined using GelAnalyzer 19.1 software [32]. Principal component analysis and correlation analysis was processed using the Seaborn library in Python (v0.13.2) [43].

3. Results and Discussion

Biostimulants, including seaweed extracts, humic and fulvic acids, protein hydrolysates, and microorganisms, are key components of sustainable agriculture and horticulture, enhancing plant growth, vitality, and stress resistance. These natural products are particularly valued for their ecologically friendly plant treatment [44,45]. In this study, we analyzed FN, traditionally used in gardening, for their content of nitrogenous substances, hydrolytic enzymes, microorganisms, phytohormones, volatile compounds, and macro- and microelements, discussing the role of these components in promoting plant growth. Additionally, FN were prepared from plants collected in different seasons (spring, spring/summer, summer, and autumn). After fermentation, FN were lyophilized for subsequent analysis. So, unless stated otherwise, the source material was FN lyophilizate. The results which were also obtained for liquid FN are presented in Supplementary Information (Figures S1–S4).

3.1. Fermented Nettles Contain Several Types of Nitrogen Compounds

Nettles are well known for their ability to thrive in nitrogen-rich environments and accumulate nitrogen (N) in their biomass. Later, they can serve as an effective organic N source, contributing to improved soil fertility and nutrient cycling [46]. In our experiment, the total nitrogen (TN) content (Figure 1A) in FN ranged from 3.7 to 6.4 g N/100 g dry weight (D.W.), which is comparable to that of manure (1.2–6.3 g N/100 g D.W.) and exceeds the N levels found in composted leaves [47]. With a carbon-to-nitrogen (C/N) ratio ranging from 6 to 11 (Figure 1B), FN function as a low C/N amendment (with C/N below 15) allowing rapid nitrogen mineralization, which makes nitrogen readily available to plants, supports early plant growth, and enhances microbial activity. In contrast, high C/N soil amendments (with C/N above 15) tend to immobilize N in microbial biomass due to their demand for N to decompose carbon-rich materials [47].

In nettles, N is present in various molecular forms, and during fermentation, which typically lasts for 14 days or more, some of these compounds undergo transformation. The resulting FN solution composed of both organic (e.g., proteins, free amino acids, pigments, and cofactors) and inorganic N forms (e.g., NH₄⁺, NO₃⁻) can serve as an effective organic fertilizer. Its N-containing components are readily available for plant uptake and can be utilized in the synthesis of essential biomolecules such as chlorophyll, proteins, nucleic acids, and secondary metabolites [14,15]. Although the fermentation of nettles was carried out in the presence of air, the resulting solution exhibited a high concentration of ammonium ions (Figure 1C) and a notably low concentration of nitrate ions—less than 0.005 mmol·g⁻¹ (0.33 mg·g⁻¹). These findings suggest that ammonification was the dominant nitrogen transformation, converting organic nitrogen compounds into NH₄⁺ and NH₃. The minimal presence of nitrate indicates that nitrification, which typically follows ammonification under aerobic conditions, occurred only to a negligible extent [15].

Fertilizers rich in ammonium ions (NH₄⁺) offer distinct agronomic advantages, particularly during the early stages of plant development and in cooler seasons. Ammonium binds to negatively charged soil particles, reducing nitrogen leaching and providing a continuous release of nitrogen through microbial nitrification. This form of N also acidifies the rhizosphere, enhancing the availability of micronutrients such as phosphorus and iron, which are critical for root establishment [48]. Although careful management is required to avoid excessive soil acidification and to ensure balanced nutrient availability, ammonium nutrition may improve crop quality—manifested as increased protein content, e.g., in wheat grain—and may also help protect plants such as tomato and potato from pathogens [49].

Further, we focused on organic N, specifically primary amines, free amino acids, and proteins (Figure 1D–F). For primary amine quantitation, we used a cheap, low-labor fluorescence method, while amino acids—an abundant subgroup of primary amines—were analyzed using a more in-depth but instrumentally demanding capillary electrophoresis (CE)-based approach. Both approaches yielded values of the same order of magnitude; however, only in summer and autumn samples, primary amines showed higher concentrations than free amino acids (Figure 1D,E). While the fluorescence method has proven reliable in soil extract analyses [19], its use with FN extracts required over 100-fold dilution to mitigate interference from coexisting substances, which may have compromised measurement accuracy. For this reason, we consider the method less suitable for this specific purpose and consequently decided to rely more on CE-derived amino acid data.

CE not only separates contaminants prior to quantification but also provides detailed profiles of individual amino acids. In all our FN samples, alanine, leucine, glycine, and glutamic acid were the most abundant (

Table 1. Seasonal variation in the amino acid content in fermented nettles (FN). Each value represents the mean ± SD. Different letters in the upper index denote significant differences (p ≤ 0.05) between groups according to one-way ANOVA (Holm–Sidak test). Abbreviations: D.W., dry weight; <LOQ, under limit of quantification; SD, standard deviation.

Amino Acids [μmol·g (D.W.)−1]	Spring	Spring/Summer	Summer	Autumn
Ala	56.1 ± 0.6 b	13.3 ± 0.1 a	9.0 ± 0.2 a	11.2 ± 1.4 a
Arg	<LOQ	4.4 ± 0.3 b	1.2 ± 0.1 a	1.3 ± 0.1 a
Asn	<LOQ	<LOQ	<LOQ	<LOQ
Asp	2.3 ± 0.7 a	5.9 ± 0.1 b	2.2 ± 0.2 a	3.3 ± 0.7 a
Cys	<LOQ	2.7 ± 0.2 b	<LOQ	1.5 ± 0.1 a
Gln	3.8 ± 0.3 b	1.0 ± 0.2 a	1.6 ± 0.1a	0.8 ± 0.0 a
Glu	16.3 ± 1.6 b	8.0 ± 0.2 a	6.2 ± 0.3 a	4.3 ± 0.6 a
Gly	28.0 ± 0.04 b	2.5 ± 0.2 a	2.8 ± 0.1 a	2.0 ± 0.3 a
His	2.4 ± 0.3 b	0.8 ± 0.02 a	<LOQ	<LOQ
Ile	15.1 ± 1.7 c	5.2 ± 0.3 b	0.8 ± 0.1 a	3.4 ± 0.7 b
Leu	28.5 ± 2.4 d	14.6 ± 0.6 c	2.4 ± 0.1 a	9.0 ± 1.1 b
Lys	10 ± 0 c	5.6 ± 0.2 b	1.8 ± 0.1 a	1.4 ± 0.2 a
Met	8.5±0.1 c	3.3 ± 0.1 b	<LOQ	2.1 ± 0.3 a
Phe	12.6 ± 0.2 c	4.5 ± 0.2 a	1.0 ± 0.1 a	5.1 ± 0.8 b
Pro	<LOQ	<LOQ	<LOQ	<LOQ
Ser	5.0 ± 2.7 b	3.2 ± 0.1 b	0.7 ± 0.1 a	2.6 ± 0.5 b
Thr	8.4 ± 2.7 b	3.9 ± 0.2 a	1.1 ± 0.1 a	2.4 ± 0.4 a
Trp	3.2 ± 0.0 b	0.8 ± 0.1 a	<LOQ	0.8 ± 0.1 a
Tyr	8.7 ± 0.1 c	3.1 ± 0.1 b	1.6 ± 0.1 a	3.1 ± 0.4 b
Val	22.3 ± 0.8 c	6.3 ± 0.4 b	2.1 ± 0.1 a	3.7 ± 0.8 a

), aligning with the amino acid composition previously observed in fresh Urtica dioica L. leaves [9]. Interestingly, glycine and glutamic acid—both when applied individually or in combination—have been shown to enhance vegetative and reproductive traits in guava trees [50]. Although not all amino acids were tested in this study, the observed effects suggest that they may also serve as energetically favorable alternatives to inorganic N sources [50]. Surprisingly, proline and asparagine, which were expected to be present in moderate amounts based on Urtica dioica L. leaf profiles, were below the detection limit in our FN samples (Table 1). The absence likely reflects microbial activity, where amino acids are selectively consumed based on their utility. Proline, for instance, may be rapidly taken up and retained due to its role as an osmoprotectant and ROS scavenger in some bacteria [51,52]; other amino acids also fulfill roles beyond serving as sources of C, N, and energy, including contributions to stress mitigation, signaling, secondary metabolism, metabolic regulation, and pH homeostasis [53]. When FN are used as a fertilizer, amino acids that they contain can, in a similar way, enhance plant stress tolerance by acting as stress-reducing agents and hormone precursors [54].

Next, we quantified the protein content in the FN (Figure 1F) and visualized their molecular weight profiles using denaturing electrophoresis (Figure 2). The protein profiles showed no noticeable differences between fresh and lyophilized FN samples (Figure 2A,B), with most proteins falling within the 10–40 kDa range. Given their total concentration, average molecular weight, and the limited number of amino groups per molecule (one N-terminal and none or a few on the side chains of lysine residues), proteins contribute minimally to the pool of primary amines (Figure 1D), unlike free amino acids. However, proteins can be enzymatically or chemically broken down into peptides and eventually into free amino acids, thereby supplementing the existing amino acid pool. In this context, the summer FN—with the highest protein content (Figure 1F)—has the potential to significantly increase amino acid levels, which are otherwise the lowest among all samples (Figure 1E).

3.2. Microbial Activity Peaks in the Summer Harvest

Microorganisms play a crucial role in the breakdown of plant material and the release of nutrients during nettle fermentation. In our FN samples, the microbial viability (Figure 3A) and total dehydrogenase activity reflecting general microbial activity (Figure 3B) follow the same seasonal trend—they are lower in the spring, peak in the summer, and decline again in the autumn. The increased microbial activity in summer FN may be driven by higher temperatures that accelerate microbial proliferation, metabolism, and enzymatic processes. Additionally, the greater availability of organic matter provides more nutrients, enhancing microbial growth and fermentation.

Microorganisms of various phyla inhabit soils, plant roots, the rhizosphere, the intercellular spaces of roots (endophytes), and above-ground surfaces of plants in densities of 10⁶ to 10⁷ per square centimeter [54]. Beneficial microorganisms—e.g., arbuscular mycorrhizal fungi, (rhizo) bacteria belonging to Rhizobium, Azotobacter, Pseudomonas, or Bacillus genus, and their consortia—were shown to function as microbial biostimulants [55,56,57]. Two endophytic Firmicutes—Bacillus cereus and Bacillus mycoides—were isolated from stinging nettles [58]; notably, certain B. cereus strains have demonstrated the ability to promote plant growth and alleviate abiotic stress, either individually or in synergy with other beneficial microorganisms [59].

The mechanisms accounting for the beneficial effects of microorganisms on plants include enhancing nutrient bioavailability (e.g., by enzymatic breakdown of organic residues, mineral solubilization, chelation of micronutrients), synthesizing volatile organic compounds with antimicrobial effects against pests, and producing phytohormones. In line with Figure 3 showing the highest content of microorganisms in summer FN, the highest activity of several studied enzymes is also observed during this season (Figure 4).

3.3. Degradation Processes Are Catalyzed by Various Hydrolytic Enzymes

FN represent a complex matrix that is almost “ever changing” due to a wide range of degradation enzymes of both microbial and plant origins. Through this diverse pool of hydrolytic enzymes, FN enhance nutrient availability for plants. In FN, at least ten hydrolases capable of degrading polysaccharides, including structural polysaccharides such as cellulose, hemicellulose, pectin, chitin, and starch, a representative storage polysaccharide, were detected (Figure 4A,B). Summer FN showed the highest activity of several exoglycosidases as well as the highest content of total saccharides (Figure 4B,C), which is also in line with (Figure 3 showing) the highest viability and activity of microorganisms in the FN from this season. Polysaccharide-hydrolyzing enzymes contribute to nutrient release and plant cell wall remodeling, while others (e.g., chitinases, β-1,3-glucanases) assist in plant defense by degrading phytopathogen cell walls [60]. Phosphatase activity (Figure 4D) supports phosphate mineralization in FN, an essential nutrient cycling process. Across all of our samples, acid phosphatase consistently exhibited higher activity than alkaline phosphatase. Urease activity (Figure 4E), which breaks down urea originating mainly from the metabolic pathways of amino acid and nucleotide degradation, remained unchanged in the FN samples across different seasons. The activity of proteases (Figure 4F), which are involved in breaking down proteins into smaller and more easily absorbable peptides and eventually individual amino acids, did not show significant seasonal differences. We did not observe correlation with the amount of proteins as the reactants, nor amino acids as the ultimate products of protein hydrolysis.

Zymographic analysis of FN samples revealed seasonal variation in protease molecular weight distribution, with higher activity associated with high-molecular-weight proteases (Figure 4G). Consistent with observations in Figure 2, no substantial differences were detected between lyophilized and liquid FN samples, except in the autumn, where protease activity in the liquid sample was markedly reduced compared to its lyophilized counterpart. Degradation enzymes play a pivotal role in nutrient release and biostimulant activity during fermentation and soil application. Enzymes such as xylanase and β-glucosidase not only break down complex polysaccharides in plant material, enhancing nutrient bioavailability, but also exert direct biostimulatory effects. For instance, xylanase and, to a lesser extent, β-glucosidase have been shown to significantly increase carotenoid, total phenolic, and vitamin C content in lettuce, thereby improving its antioxidant capacity [61]. The availability of commercial biostimulants (such as Canna Cannazym, Plagron Pure Zym, and Hesi Power Zyme)—each containing a diverse mix of enzymes designed to enhance cell biomass without causing phytotoxicity—reflects a growing interest in enzyme-based biostimulants among plant biofactories as well as their importance within the broader context of sustainable agriculture. Therefore, FN could serve both as a source of nutrients (e.g., amino acids, proteins, and saccharides; Figure 1E,F, Figure 2A,C, and Figure 4C; Table 1) as well as a supplier of enzymes (Figure 4A,B,D–F) that release additional nutrients or modulate the composition of microorganisms in the soil. The increased microbial activity observed in summer FN can be interpreted through several interconnected environmental and biochemical factors. Firstly, elevated temperatures during summer months tend to accelerate microbial metabolism, leading to faster growth rates and more active enzymatic processes. Secondly, the nutrient content in FN may be higher in summer due to increased organic matter availability, which provides microbes with more substrates to metabolize, thus supporting microbial growth and enzyme secretion. Correlation analysis of several FN parameters showed that microbial viability as well as total dehydrogenase activity positively correlated especially with exoglycosidase, α-galactosidase, β-galactosidase, β-glucosidase, and α-mannosidase activities and slightly with proteolytic activity (Figure S6). Thus, we suggest that these enzymes are mainly of microbial origin.

3.4. Phytohormones May Contribute to Plant Growth, Seed Germination, and Defense

Plant biostimulants may contain phytohormones, which play a key role in regulating plant growth, development, and resistance. Whereas auxin was the only phytohormone, which was previously detected in FN [15], a wide range of FN phytohormones and many of their metabolites were found in this study (

Table 2. Seasonal variation in the phytohormonal content in fermented nettles (FN). Each value represents the mean ± SD. Different letters in the upper index denote significant differences (p ≤ 0.05) between groups according to one-way ANOVA (Holm–Sidak test). Active cytokinins (CKs) are a sum of dihydrozeatin (DZ) and isopentenyl adenine (iP). Another active CK, cis-zeatin (cZ), is displayed in a separate row. Abbreviations: ABA, abscisic acid; D.W. dry weight; GAs, gibberellins; JA, jasmonic acid; JA-Ile, JA-izoleucine; IAA indole-3-acetic acid; OxIAA, oxindole-3-acetic acid; PAA, phenylacetic acid; SA, salicylic acid; SD, standard deviation.

Phytohormones	Spring	Spring/Summer	Summer	Autumn
IAA [nmol·g(D.W.)−1]	175.69 ± 30.77 a	371.94 ± 54.08 a	4797.34 ± 272.07 b	315.39 ± 104.41 a
PAA [nmol·g(D.W.)−1]	4095.19 ± 511 a	8542.34 ± 474.07 b,c	9233.88 ± 41.9 c	6962.94 ± 870.1 b
OxIAA [nmol·g(D.W.)−1]	26.82 ± 8.42 a	307.02 ± 87.82 b	63.34 ± 36.17 a	20.22 ± 6.85 a
cZ [pmol·g(D.W.)−1]	145.23 ± 4.54 c	24.35 ± 10.7 a	74.76 ± 0.58 b	38.7 ± 13.15 a
DZ and iP [pmol·g(D.W.)−1]	154.81 ± 11.89 c	103.86 ± 3.59 b	158.71 ± 7.36 c	50.33 ± 11.75 a
Active GAs [pmol·g(D.W.)−1]	1299.06 ± 21.38 a	1156.62 ± 16.49 a	1125.43 ± 26.84 a	1882.2 ± 108.53 b
ABA [pmol·g(D.W.)−1]	20.94 ± 10.87 a	44.61 ± 0.27 a	78.3 ± 17.22 a	61.2 ± 16.49 a
JA [nmol·g(D.W.)−1]	20.71 ± 7.87 a	19.26 ± 4.5 a	25.93 ± 7.03 a	20.65 ± 6.77 a
JA-Ile [nmol·g(D.W.)−1]	17.26 ± 5.42 a	10.33 ± 1.6 a	7.83 ± 1.5 a	7.19 ± 1.41 a
SA [nmol·g(D.W.)−1]	8.84 ± 0.1 a	4.91 ± 0.31 a	76.75 ± 2.69 a	32.51 ± 10.9 b

). It appears that the original plants and the rich mixture of microorganisms actively synthesize but also metabolize and inactivate the phytohormones under investigation. Active auxins were present in significant amounts, namely indole-3-acetic acid (IAA) and phenylacetic acid (PAA). PAA, which is evolutionarily older and could also be found in microorganisms [62], was detected in higher amounts than IAA in FN. Summer FN showed more than a ten times higher amount of IAA than the other FN seasons (Table 2). Auxins can be absorbed by plant roots, promoting growth and contributing to the interaction between plants and microorganisms, but they can also be used by microorganisms as a source of nutrients [63,64]. The elevated IAA content in summer may be caused by enhanced microbial activity, as in summer the microbial viability was much higher than in other sample points (see Figure 3). The biosynthesis of auxin indole-3-acetic acid by microorganisms was reported to have a major impact on plant–microbe interactions [65]. In summer, endogenous auxin content in nettles may also be high, promoting plant photosynthesis and metabolism, as indicated by high total saccharide contents (Figure 4C) or high protein content (Figure 1F). And auxin can be considered as a key molecule to improve plant performance [66]. Among the IAA metabolites, conjugates with amino acids and glucosyl ester, which may serve as storage forms, but also large amounts of oxidized form OxIAA and its metabolites, were found (Table S1). Spring/summer FN were the richest in OxIAA. In addition to auxin, cytokinin is another phytohormone regulating plant growth. Highly abundant active cytokinin, cis-zeatin (cZ), may be produced during the degradation of some transfer RNAs [67], which can occur to a significant extent during nettle fermentation (Table 2). In contrast to the relatively high levels of another active cytokinin found in the shoots isopentenyladenine (iP), we detected very low concentrations of trans-zeatin (tZ; Table S1) probably because tZ is predominantly synthesized in the roots, which were excluded from the fermentation mixture. Cytokinins promote cell division, delay senescence, and are related to nutrient availability. Spring FN showed the highest content of cZ and second highest level of other active cytokinins (Table 2).

Active gibberellins GA₃, GA₄, and GA₇ have also been found in FN samples, but there were no significant changes among the seasons (Table 2 and Table S1). Gibberellins generally promote stem growth and flowering [68], but in soil, they may be particularly important for promoting germination.

Phytohormones, primarily involved in stress responses and stimulating defense, were found in FN, as well. The key hormone in abiotic stress responses [69], abscisic acid (ABA) and its storage form, ABA glucosyl ester (ABA-GE), were present in significantly lower amounts than the inactive metabolites such as phaseic acid and dihydrophaseic acid (Table 2 and Table S1). These substances may originate from the nettle plants. The hormone associated predominantly with biotic stresses [69] is jasmonate (JA) and, above all, its active form, the isoleucine conjugate (JA-Ile), and methyl jasmonate (Me-JA) as a volatile form used for signaling within and between the plants (Table 2 and Table S1). Salicylic acid (SA) is the stress hormone associated mainly with the responses to biotrophic pathogens [69,70]; in soil, it can increase the resistance of seeds to phytopathogenic bacteria. Summer FN contained the highest amount of IAA, ABA, and SA (Table 2). Foliar applications of SA and ABA promoted plant growth, increased antioxidant enzyme activity, and reduced oxidative damage in the hybrid rice cultivar [71]. Khan et al. (2020) [72] demonstrated that a thermotolerant strain of Bacillus cereus, isolated from the roots of Echinochloa crus-galli L., was effective in alleviating heat stress in soybean plants. This was evidenced by the increased activity of antioxidant enzymes, elevated levels of photosynthetic pigments and amino acids, changed phytohormone levels, and reduced lipid peroxidation. Additionally, the strain was found to produce organic acids and phytohormones [72]. The phytohormone content may be one of the reasons why FN can positively promote plant growth and stimulate defense mechanisms, thereby increasing plant resistance to stress. Therefore, summer FN could be the most advantageous for use as a biostimulant based on the phytohormonal content.

3.5. Presence of Volatile Compounds with Antimicrobial Effects

Volatile substances were identified by GC×GS-MS after extraction with methanol from FN lyophilizate (Figure 5A) and by headspace solid-phase microextraction from liquid FN (Figure 5B). These are mainly oxygen-containing hydrocarbons (alcohols, aldehydes, ketones, acids), aromatic hydrocarbons, terpenoids, and sulfur compounds (Figure 5). It is well established that plant roots influence surrounding microorganisms through the release of exudates, which in turn secrete compounds that promote plant growth. Volatile organic compounds may also induce plant growth and enhance stress resistance. To date, alcohols, ketones, sulfur compounds, and terpenoids have been identified as bioactive volatile organic compounds [73]. Similar substances have also been detected by both extraction methods from FN (Figure 5), including common fatty acids, aldehydes, ketones, and alcohols as well as an amide and an alkyne. Another group of volatile substances identified in the FN samples includes terpenoids, such as limonene, phytol, and α-bisabolene epoxide (Figure 5). Phytol is a partly hydrogenated diterpene alcohol, a component of chlorophyll and a common degradation product of plant material, which is why phytol is widely regarded as a biogeochemical marker of aquatic environments. Its derivative without an alcoholic group, 3,7,11,15-tetramethyl-2-hexadecene, was also identified in the FN samples and isolated from the endophytic filamentous fungus Diaporthe schini, demonstrating antibacterial and antifungal activity [74]. Biological activity has also been reported for other compounds found. The 1,4-benzenedicarboxylic acid, bis(2-ethylhexyl) ester, has inhibitory effects against the fungi Aspergillus flavus and Candida krusei, and against several bacteria, such as Staphylococcus aureus, Escherichia coli, Shigella dysenteriae, and Klebsiella pneumoniae [74]. Pentadecanoic acid, 14-methyl-, methyl ester is produced by the strain MT597434.1 of the endophytic fungus Aspergillus niger and, together with other metabolites, successfully combats multidrug-resistant microorganisms [75]. Along with other metabolites, 3-Octadecen-1-ol acetate was identified in avocado seeds, which are known for their antibacterial activity and antioxidant capacity [76].

When applied as a soil drench, hexanoic acid is absorbed and accumulated in the roots, induces plant resistance, and functions as an elicitor against Pseudomonas syringae and Alternaria alternate. Plants subsequently increase callose deposition and caffeic and chlorogenic acid synthesis [77].

Isolated from compost, the Bacillus velezensis strain Q-426 produces a number of volatile compounds, mainly ketones, such as 2-heptanone, 6-methylheptan-2-one, 5-methylheptan-2-one, 2-nonanone, 2-decanone, 2-undecanone, 2-dodecanone, 2-tridecanone, 2-tetradecanone, and 2-nonadecanone, with excellent antifungal activity against plant pathogens [78]. We also detected several ketones among the volatile compounds of the FN samples (Figure 5). Many of the ketones with growth promotion effects produced by Burkholderia ambifaria strains were also found in FN [79]. Some of the volatile compounds detected in our study were also identified in edible macroalgae [80].

Sulfur compounds such as dimethyl sulfide and dimethyl disulfide were also identified in FN (Figure 5). These substances occur in freshwater environments, where they are mainly produced by sulfide methylation or the degradation of sulfur-containing amino acids [81]. Dimethyl disulfide promotes Arabidopsis growth and has antifungal effects [73,79]. Terpenoids such as alpha-terpineol, limonene, and dehydroedulan display a wide range of biological activities, including insecticidal properties [82]. In the principal component analysis based on the profiles of volatile compounds (Figure S5), the FN variations differ from each other. The summer FN negatively correlates with spring and spring/summer variants, and is mostly distinguished by several organic acids (Figure S5B; Table S2).

However, several of the volatile compounds found in fresh nettles, such as caryophyllene, linalool, carvone, carvacrol, and naphthalene [8], were not found in our FN samples, suggesting that these strong antimicrobial compounds are rapidly degraded by microbial communities. Some volatile substances originate from nettles, whereas others are formed during fermentation by microorganisms, primarily bacteria, with potential contributions from fungi and algae. Thus, the application of FN to the soil can introduce additional microorganisms that can affect the composition of initial microbial communities, e.g., by producing antimicrobial substances that may combat phytopathogens. In summary, volatile compounds in FN can promote plant growth by acting as signaling molecules, enhance resistance to pathogens through antimicrobial and antifungal properties, and support beneficial soil microbial interactions as well.

3.6. High Contents of Phenolics and Flavonoids May Serve as Beneficial Antioxidants

Another potential mechanism of action of FN may be their high phenolic content and antioxidant capacity (Figure 6). According to our results, both properties were especially prominent in the case of the FN harvested in summer (Figure 6C,D). The antioxidant capacity of summer FN matched (Figure 6D) that of aqueous extracts of dried herbs, namely Chelidonium majus L. and Lamium album L. (ranging in 150–175 µmol of Trolox equivalent·g (D.W.)⁻¹ as measured by the ABTS method) [39]. The antioxidant properties of FN are likely associated with various phenolic compounds and flavonoids. Additionally, vitamins such as thiamine, riboflavin, vitamin B6, niacin, and vitamins C, A, and K—previously identified in nettle leaves according to the published literature—may also contribute to its antioxidant potential. [83]. In nettles, the most abundant phenolic compounds are chlorogenic acid, rutin, isoquercetin, and quercetin, recognized for their beneficial effects [8,83]. Commonly found in plant decomposition products, phenolic compounds may act as key precursors of humic substances in soils and potentially as allelochemicals, exerting either beneficial or adverse effects of one plant on another [84]. In line with these findings, a phenolic-based biostimulant consisting of a mixture of protocatechuic acid, quercetin, chlorogenic acid, coumaroyl quinic acid, and gentistic acid has been shown to support potato yield and quality [85]. Based on our results, summer FN shows the highest biostimulant potential for plants and microorganisms given the total content of phenolics and flavonoids. Also, generally, correlation analysis revealed a strong positive correlation between microbial viability (and total dehydrogenase activity as well) and the total content of phenolics and flavonoids and antioxidant capacity (Figure S6).

Numerous factors, including UV radiation, pH, Fe content, redox conditions, organic matter content, and soil microorganisms, influence the production of harmful ROS and the occurrence of Fenton reactions in soils [86]. FN contain various photosynthetic pigments with antioxidant and ROS scavenging activity (Figure 6A,B) and antioxidant enzymes, such as superoxide dismutase and peroxidases (Figure 7A–C), whose activity was detected in liquid FN. Through their antioxidant properties, these molecules may help to increase resistance in plants treated with FN, especially under conditions involving oxidative stress and subsequent tissue damage.

Laccase, another enzyme involved in stress responses, plant growth, and cell wall remodeling [87], was also detected in liquid FN (Figure 7D). Laccases catalyze the oxidation of various aromatic compounds, contributing to lignin and polyphenol transformation in soils. While traditionally associated with fungi, some bacteria also produce laccase-like enzymes, which may play roles in natural degradation processes and the transformation of xenobiotic pollutants [88].

3.7. Fermented Nettles Are a Rich Source of Several Macro- and Micronutrients

FN can provide plants with a whole range of biogenic macro- and micronutrients (Figure 1;

Table 3. Seasonal variation in nutrient concentrations in fermented nettles (FN). Each value represents mean ± SD. Different letters in upper index denote significant differences (p ≤ 0.05) between groups according to one-way ANOVA (Holm–Sidak test). Macronutrients are expressed as mg·g (D.W.)⁻¹ and micronutrients in μg·g (D.W.)⁻¹. Abbreviations: D.W., dry weight; SD, standard deviation.

Nutrient	Spring	Spring/Summer	Summer	Autumn
K [mg·g(D.W.)−1]	93.7 ± 3.9 d	72.9 ± 1.3 c	59.7 ± 2.2 b	32.4 ± 1.7 a
Ca [mg·g(D.W.)−1]	55.9 ± 3.2 a	50.8 ± 1.0 a	65.6 ± 2.8 a	87.8 ± 5 b
Mg [mg·g(D.W.)−1]	8 ± 0.4 b	7 ± 0.9 a,b	5.5 ± 0.7 a	6.3 ± 0.4 a,b
S [mg·g(D.W.)−1]	2.8 ± 0.4 a	5.1 ± 0.4 a	3.5 ± 0.4 a	4.5 ± 1.3 a
Phosphates [mg·g(D.W.)−1]	30.7 ± 0.5 a	64.8 ± 3.2 a	479.8 ± 33.9 b	91.4 ± 2.9 a
Na [µg·g(D.W.)−1]	2633 ± 129 c	1811 ± 222 b	1464 ± 187 b	805 ± 52 a
Si [µg·g(D.W.)−1]	8959 ± 811 b	5384 ± 229 a	4482 ± 252 a	5442 ± 599 a
Fe [µg·g(D.W.)−1]	1647 ± 144 c	1293 ± 454 b	754 ± 101 a	1347 ± 338 b
B [µg·g(D.W.)−1]	562 ± 53 a	1713 ± 1268 b	1770 ± 558 b	1429 ± 383 b
Zn [µg·g(D.W.)−1]	302 ± 26 c	103 ± 9 b	12 ± 4 a	501 ± 82 d
Mn [µg·g(D.W.)−1]	273 ± 9 b	307 ± 6 c	218 ± 5 a	351 ± 40 d
Cu [µg·g(D.W.)−1]	161 ± 5 d	108 ± 16 b	48 ± 2 a	142 ± 18 c
Mo [µg·g(D.W.)−1]	58 ± 4 b	51 ± 7 b	32 ± 1 a	53 ± 6 b

). In this study, their content of macronutrients (K, Ca, Mg, and S) was quantitatively consistent with data from previous studies [15]. But FN are also a source of various micronutrients (Na, Si, Fe, B, Zn, Mn, Cu, and Mo) (Table 3). These micronutrients serve as structural components, cofactors, osmotic solutes, among other roles [15,89]. Phosphate is the second most abundant nutrient in inorganic fertilizer supplements because it is critical for plant growth but often deficient in soil ecosystems [90,91,92]. FN was relatively rich in phosphates, especially in the summer variant (Table 3). Primarily, soil provides the plants with at least 17 elements required for their life cycle [93]. For these reasons, micro- and macronutrient-rich FN can supplement plants, especially in the case of soils poor in key nutrients. In nettle leaves, mineral concentrations are known to vary widely with factors such as soil quality and vegetative stage [5,83,94]. Furthermore, increasing the doses of N fertilizers in soils directly alters the concentrations of Mn, Zn, and Cu in nettle leaves, albeit without affecting the Fe content [46]. In our experiments, spring FN contained the highest concentrations of Fe, Zn, Mn, and Cu (Table 3). Moreover, Si, which is a component of the defensive structures of trichomes in the form of SiO₂, was also detected in nettles [95] as well as here in FN (Table 3). Si also promotes crop growth, especially those of the Poaceae family. Rice is a major accumulator of Si, and higher rice yields per unit area are correlated with Si depletion from soils [96,97]. Another vital micronutrient for plants is Boron (B), which is also found in FN (Table 3). Although excess B can induce toxicity symptoms [98], the combination of B and Si upregulates antioxidant enzymes and significantly improves both biomass and root length in soybeans exposed to Al toxicity [99]. Nettles are nowadays being studied from the perspective of phytoremediation; they were able to uptake 8% of Zn, Pb, and Cd as well as organic pollutants after four months [100]. Their fast growth can be advantageous for fertilizer preparation given their rapid accumulation of nutrients, but also disadvantageous due to the possibility of uptake of heavy metals or organic pollutants, which may lead to the contamination of plants treated with FN. Even though the application of FN could supply the plant and soil with several macro- and micronutrients, for their usage as fertilizers, nettles should be collected from clean environments. Therefore, according to our results, the concentrations of macro- and micronutrients in FN likely depend more upon the place of growth than on the period of nettle harvest (Table 3).

Several monitored FN parameters were ranked from the highest to the lowest based on the average (SD not included) values measured for each harvest season (Figure S7). Based on our results, spring FN showed an increased content of N compounds; summer FN showed enhanced microbial viability, total saccharide, phenolic and flavonoid content, antioxidant properties, and exoglycosidase activity; and autumn FN showed increased chlorophyll content. However, these properties of FN may also vary due to the exact characteristics of the soil and to other factors and conditions under which nettles grow. Considering their properties, FN from different seasons seem to be beneficial for both plants and the soil microbiome.

4. Conclusions

This study provides a detailed biochemical characterization of fermented nettles, highlighting their multifaceted role in sustainable agriculture as both organic fertilizers and biostimulants. The results demonstrate that fermented nettles are rich in nitrogenous compounds, including ammonium ions, free amino acids, and proteins, with a favorable C/N ratio that supports microbial activity and nutrient availability. The application of fermented nettles may also supply essential macro- (K, Ca, Mg, S, P) and micronutrients (Na, Fe, Zn, Mn, Na, Cu, Mo, Si, B) to plants and soil. In addition, nettles contain a number of bacteria which, together with plant enzymes, are responsible for the degradation of plant tissue. Fermented nettles contained active endo- and exoglycosidases, proteases, urease, and phosphatases.

Additionally, fermented nettles contain phytohormones that promote plant growth (auxins, cytokinins, gibberellins) or are related to stress response (salicylic acid, jasmonate, abscisic acid). Moreover, fermented nettles are a source of volatile organic compounds (oxygenated hydrocarbons and terpenoids) with antimicrobial properties, and their antioxidant capacity, especially in the summer harvest, is also supported by high levels of phenolics, flavonoids, and antioxidant enzymes. These attributes suggest that fermented nettles not only enrich soil fertility but also enhance plant health and resistance to biotic and abiotic stressors. However, fermented nettles, while promising as a natural fertilizer or supplement, may present certain limitations. Their composition can vary significantly depending on factors like harvest time, soil quality, temperature, water regime, and last but not least, fermentation conditions, which affect consistency and efficacy. Product stability is another concern, as ongoing microbial activity or poor storage may lead to spoilage or reduced potency. Additionally, nettles are known to absorb heavy metals from contaminated environments, raising potential safety issues if not properly sourced. These factors should be considered to ensure safe and reliable application. We must view fermented nettles as a natural product that is variable in principle, which is why we monitored seasonal influences in this study. Further research is needed, as no tests have been here conducted on actual crops. Additional studies would help verify its effectiveness under real conditions, such as in greenhouses or open fields.

In conclusion, fermented nettles exhibit a synergistic combination of nutritional, enzymatic, hormonal, (anti)microbial, and antioxidant properties, so its application may be beneficial for plant fitness, soil vitality, and the proliferation of beneficial microorganisms. The seasonal variability underscores the importance of harvest timing, with a summer variant emerging as particularly potent. Overall, fermented nettles represent a promising natural solution for improving soil vitality and crop performance, reinforcing their dual function as fertilizers and biostimulants in sustainable agricultural practices.