Microbial Silver Nanoparticles Enhance the Performance of Maize Plants Cultivated in Naturally Occurring Saline Soil

Abstract

Soil salinity is a major abiotic stress that limits agricultural productivity worldwide. The aim of this study was to evaluate whether biogenic silver nanoparticles (AgNPs) can mitigate salt stress in maize while preserving soil biological health under realistic soil conditions. Biogenic AgNPs were synthesized using biomolecules from the actinobacterium Streptomyces sp. Z38 and characterized, confirming spherical morphology, colloidal stability, and surface functionalization. Maize plants grown under greenhouse conditions were treated with biogenic or chemically synthesized AgNPs, and plant performance, oxidative stress responses, and soil biological properties were evaluated. Under saline conditions (6 mS cm⁻¹), biogenic AgNPs markedly improved plant growth, almost fully restoring leaf dry weight (165.08 ± 23.68 mg) to values comparable with non-saline controls (171.81 ± 15.00 mg), while chemical AgNPs induced only partial recovery. Biogenic AgNPs also enhanced antioxidant defenses, increasing catalase activity by ~15% above non-saline levels and reducing lipid peroxidation from 232.34 ± 31.74 to 102.63 ± 5.75 Eq. MDA g⁻¹. In parallel, chlorophyll a content increased by ~29% relative to non-saline plants, indicating improved photosynthetic performance. Transmission electron microscopy of leaves confirmed AgNPs internalization, with nanoparticles primarily sequestered in vacuoles. Analyses of experimental soils showed that biogenic AgNPs enhanced microbial enzymatic activity and respiration, while chemical AgNPs had inhibitory effects. Ecotoxicological assays further indicated low soil toxicity following biogenic AgNPs plant treatment, as reflected by high lettuce germination rates. Overall, these findings highlight the potential of biogenic AgNPs obtained from actinobacteria as sustainable nanobiotechnological tools to mitigate salt stress in crops while improving soil health. Future field-scale studies will be required to validate their agronomic applicability.

1. Introduction

Soil salinization is one of the most critical environmental constraints threatening agricultural productivity worldwide. More than 20% of total cultivated lands are affected by salinity, and this percentage is expected to rise due to unsustainable irrigation practices, climate change, and sea-level rise [1,2]. From a physiological standpoint, salinity imposes an initial osmotic stress that restricts water uptake, followed by ionic toxicity driven mainly by Na⁺ and Cl⁻ accumulation, which disrupts K⁺ nutrition and cellular ion homeostasis. These processes converge on oxidative stress, promoting reactive oxygen species (ROS) overproduction, membrane lipid peroxidation, protein dysfunction, and photosynthetic impairment, ultimately reducing growth and yield [3,4].

Maize (Zea mays L.), a staple cereal of global importance, is moderately sensitive to salt stress, particularly during early stages of development. To counteract salt stress, plants deploy complementary strategies including ion exclusion and long-distance transport control (e.g., Na⁺ efflux and xylem retrieval), vacuolar sequestration of toxic ions, osmotic adjustment via compatible solutes, and reinforcement of antioxidant defenses (CAT, POD, SOD, APX) coupled to hormonal and signaling networks (e.g., ABA and ROS signaling) that balance stress tolerance with growth [4,5,6]. However, these natural defense responses are often insufficient to withstand prolonged/intense salt exposure or field-relevant salinity levels, motivating the development of additional, agronomically feasible strategies. In this context, nanotechnology has emerged as a promising approach to enhance plant stress tolerance. Specifically, metallic nanoparticles (NPs), including silver nanoparticles (AgNPs), have shown potential to improve plant performance under abiotic stress conditions by modulating growth, physiology, and biochemical responses [4,5,7,8].

Silver nanoparticles possess unique physicochemical properties that make them useful in a wide range of biological and environmental applications. In plants, AgNPs have been shown to enhance seed germination, promote root elongation, improve photosynthesis, and activate antioxidant defense systems under stress conditions [9,10]. Nevertheless, concerns remain regarding the ecotoxicity of chemical AgNPs, particularly their negative effects on beneficial soil microorganisms and non-target species, largely due to the uncontrolled release of silver ions (Ag⁺) [11].

An alternative and environmentally friendly approach to NPs synthesis involves the use of biological systems, such as bacteria, fungi, or plant extracts, to produce NPs through green synthesis routes. Biogenic AgNPs not only reduce the use of toxic reagents but also exhibit surface capping with biomolecules (e.g., proteins, polysaccharides) that can influence their stability, functionality, and interactions with biological systems [9,11,12]. In particular, actinobacteria of the Streptomyces genus are known for their rich secondary metabolism and have been reported as effective microbial factories for the biosynthesis of metallic NPs with bioactive properties [13,14].

Despite increasing interest in biogenic NPs, their application in agriculture, particularly under salt stress conditions, remains insufficiently explored. Moreover, few studies have compared the effects of biological vs. chemical AgNPs on both plant performance and soil microbial communities under salinity conditions. Considering the central role of the rhizosphere microbiome in plant health and nutrient cycling, assessing the impact of nanomaterials on soil quality is essential for the development of sustainable nano-enabled agro-technologies.

Although AgNPs have been widely investigated for their effects on plant growth and stress tolerance, most previous studies have primarily focused on chemically synthesized AgNPs and have largely evaluated plant responses under simplified or artificial conditions. In contrast, the present study integrates the use of Streptomyces-derived biogenic AgNPs, naturally occurring saline soil, and a combined assessment of plant physiological responses and soil biological health, including enzymatic activity, respiration, ecotoxicity, and microbial community structure. Therefore, a clear knowledge gap exists regarding the synergistic effects of Streptomyces-derived biogenic AgNPs on salt-stressed plants and soil microbial functioning within a unified experimental framework. The integrated plant–soil approach used in this study allows us to identify not only stress mitigation effects at the plant level, but also the broader ecological implications of AgNPs application, highlighting clear differences between biogenic and chemical nanoparticles.

Based on these considerations, we hypothesized that biogenic AgNPs synthesized by Streptomyces sp. Z38 enhance plant tolerance to salinity through their surface-associated biomolecules, while simultaneously exerting a positive influence on soil microbial activity. Accordingly, the aim of this study was to (i) biosynthesize and characterize AgNPs using Streptomyces sp. Z38, (ii) evaluate their effects on the growth, physiological, and biochemical responses of maize plants under moderate salt stress, and (iii) assess their impact on soil microbial activity, respiration, ecotoxicity, and microbial diversity. Comparisons were made with chemical AgNPs to elucidate differences in performance and environmental compatibility. The findings of this research provide valuable insights into the use of biogenic NPs as a sustainable strategy to enhance crop resilience and soil health in saline environments.

2. Results and Discussion

2.1. Characterization of Biogenic AgNPs

The successful biosynthesis of AgNPs from Streptomyces sp. Z38 was confirmed by the appearance of a characteristic surface plasmon resonance (SPR) absorption peak at 410 nm (Figure 1a), in accordance with previous findings [14,15]. An increase in absorbance over time was observed, suggesting a progressive accumulation of AgNPs in the solution. Based on these results, an incubation time of 72 h was selected as optimal and these AgNPs were used for subsequent characterizations and applications.

Given that the size and morphology of AgNPs significantly influence their physicochemical properties and biological activity, an initial morphological assessment was performed by SEM. The micrographs revealed that the biogenic AgNPs exhibited a predominantly spherical shape (Figure 1b). Size distribution analysis indicated that the particles had an average diameter of 40 nm, with sizes ranging between 10 and 70 nm (Figure 1b). These observations were corroborated by DLS analysis, which showed a mean hydrodynamic diameter of 44 nm (Figure 1c). It is well established that extracellularly synthesized AgNPs tend to be polydisperse and exhibit broader size variability compared to those formed intracellularly [16,17]. Notably, Streptomyces spp. have been shown to produce AgNPs within a wide size range: for example, sizes from 10 to 50 nm were previously reported [14], while other authors observed sizes between 11 and 21 nm [16], indicating that NPs dimensions can vary depending on the strain and synthesis conditions employed.

The zeta potential of the AgNPs was −44.5 ± 1.6 mV, indicating strong electrostatic repulsion between particles and, consequently, high colloidal stability. Zeta potential is a key parameter for evaluating NPs stability, since higher absolute values, such as that obtained in this study, reduce the likelihood of aggregation [18]. In addition, the chemically synthesized AgNPs exhibited an even more negative zeta potential (−56.6 mV), suggesting superior colloidal stability. This behavior is likely related to the presence of a specific stabilizing agent used during synthesis (in this case, citrate), which generates a more uniform surface charge distribution and stronger electrostatic repulsion between particles [19].

To elucidate the chemical groups involved in NPs stabilization and surface functionalization, FTIR and Raman spectroscopic analyses were performed on both the AgNPs and the extract used in their synthesis, corresponding to the bioactive water (BW) (Figure 2). FTIR spectra (Figure 2a) showed prominent peaks around 3420 cm⁻¹ and 1600 cm⁻¹, corresponding to –OH stretching and –C=O stretching, respectively, both associated with –CONH– amide linkages. The peak at ~1600 cm⁻¹ is indicative of amide I groups, commonly found in proteins [20], and was present in both the BW and AgNP spectra, suggesting that proteins from the BW may be involved in AgNP stabilization, as similarly reported for Streptomyces sp. M7 [14]. Proteins and other biomolecules are known to act as capping agents, providing stability, and possibly bioactivity to metal NPs [21].

Additional weak signals were observed at 1390 cm⁻¹ and 1200 cm⁻¹, corresponding to aromatic C–H bending and C=C stretching, respectively [22]. A distinct peak at 833 cm⁻¹ was detected in the BW but absent in the biogenic AgNP spectrum, suggesting that not all organic compounds present in the BW became associated with the NP surface.

Raman spectra recorded at excitation wavelengths of 532 nm and 785 nm provided further insights into the molecular interactions at the NP surface (Figure 2b). At 532 nm, a broad peak near 1000 cm⁻¹ was observed in the AgNP spectrum, attributed to amide groups in proteins. This implies that proteins may be attached to the AgNP surface via free amino groups or electrostatic interactions involving negatively charged carboxylates [23]. In the BW spectrum, multiple peaks between 1000 and 1600 cm⁻¹ were observed, likely corresponding to various organic groups weakly associated with the NPs (Figure 2b).

In the Raman spectra acquired at 785 nm (Figure 2b), a well-defined peak around 211 cm⁻¹ was identified and associated with Ag-O stretching vibrations [24]. Another peak at 1400 cm⁻¹, also attributed to amide groups, was detected in both AgNP and BW samples, reinforcing the involvement of proteinaceous compounds in NP synthesis and stabilization. Taken together, Raman and FTIR findings reinforce the hypothesis that proteins from the BW act as dual-function agents reducing and stabilizing AgNPs, as previously proposed [17]. Moreover, the thermal decomposition (TGA) of biosynthesized AgNPs obtained from Streptomyces sp. M7 and their chemical counterpart were studied by Paterlini et al. [14]. The authors informed thermal decomposition occurred in gradual five-step meanwhile AgNPs obtained by chemical synthesis showed just three thermal decomposition steps. Thus, it was possible to observe how the organic component bound to biogenic AgNPs affected their thermal stability [14].

Finally, the silver content of the biogenic NPs was quantified using ICP-MS, revealing that the AgNPs contained 10.40 ± 0.06% elemental silver. This value is crucial for understanding their chemical composition and potential applications [25]. The remaining mass likely corresponds to organic matter adsorbed on the NP surface, in agreement with the spectroscopic analyses. The presence of this organic capping layer plays a significant role in determining the physicochemical behavior of AgNPs, including their solubility, reactivity, and interactions with biological systems [26].

2.2. Implications of AgNPs Treatments on the Growth of Maize Plants Developed on Saline Soil

Soil salinity is typically assessed through electrical conductivity (EC), which reflects the concentration of dissolved salts. In addition, EC is widely used for classifying salt stress severity in agricultural systems [27]. According to FAO [28], maize is moderately salt-tolerant and can withstand EC values of up to approximately 8 mS/cm. Based on this information, a soil EC of 6 mS/cm soil was selected for subsequent experiments, as it is expected to induce significant salt stress while still allowing for plant development. Moreover, in order to evaluate contrasting conditions, a non-saline soil of 1 mS/cm was used for standard plant development (control). In addition to soil salinity levels, another key aspect considered in the experiment was the NPs application scheme. The selected concentration (75 mg L⁻¹) was based on previous studies reporting positive physiological effects of AgNPs on plants without inducing phytotoxicity, particularly under abiotic stress conditions. Raza et al. [29] applied different concentrations of biosynthesized AgNPs (25, 50, 75, and 100 mg L⁻¹) to Huanglongbing-diseased plants, a disease that severely affects citrus species, and their findings showed that 75 mg L⁻¹ was the most effective in enhancing the plants’ physiological profiles. Furthermore, the single foliar application at 15 days used in our experiment was designed to coincide with an early yet stable developmental stage of maize, when plants begin to exhibit clear physiological responses to salt stress but have not yet undergone irreversible damage. This strategy allowed us to assess the impact of a single, well-defined intervention, avoiding potential effects associated with cumulative responses from multiple applications.

As shown in Figure 3, salinity markedly affected maize shoot development. Leaf length decreased from 24.84 ± 3.75 cm under non-saline conditions to 20.09 ± 3.28 cm in saline soil, representing a 19% reduction. This effect was accompanied by a strong decline in leaf dry weight, which dropped from 171.81 ± 15.00 mg (non-saline soil) to 66.41 ± 6.24 mg (saline soil), corresponding to a 61% decrease. Foliar application of both AgNPs types mitigated the negative impact of salinity on leaves. Plants treated with biogenic AgNPs reached a leaf length of 28.06 ± 2.39 cm, while those treated with chemical AgNPs attained 29.95 ± 2.55 cm, both values being comparable to or higher than those observed under non-saline conditions. Regarding biomas, biogenic AgNPs almost fully restored leaf dry weight (165.08 ± 23.68 mg) to levels similar to the non-saline control (171.81 ± 15.00 mg), whereas chemical AgNPs induced a partial recovery (98.85 ± 15.03 mg), remaining approximately 42% lower than plants grown in 1 mS/cm soil (non-saline soil).

Root growth was also significantly impaired by salinity (Figure 3). Root length decreased from 48.78 ± 7.95 cm under non-saline conditions to 32.59 ± 5.03 cm in saline soil, representing a 33% reduction, while root dry weight declined from 52.01 ± 4.59 mg to 32.64 ± 0.81 mg (37% decrease). Application of both AgNPs treatments partially alleviated these effects. Root length increased to 37.73 ± 2.67 cm and 37.16 ± 2.29 cm in plants treated with biogenic and chemical AgNPs, respectively, although these values did not fully reach those of the non-saline control (48.78 ± 7.95 cm). In contrast, root dry weight showed a marked response to AgNPs application, increasing to 88.51 ± 4.59 mg in the biogenic AgNPs treatment and to 65.10 ± 9.74 mg in the chemical AgNPs treatment. These values represent increases of approximately 171% and 99%, respectively, relative to the saline control (32.64 ± 0.81 mg), indicating a strong stimulation of root biomass accumulation under salt stress conditions under the treatment with both types of AgNPs.

Stomatal uptake has been proposed as one of the primary entry routes for foliar-applied AgNPs [30]. Regarding this approach, TEM images was employed to examine leaf tissue from AgNPs-treated plants. Micrographs confirmed the internalization of both chemical and biogenic AgNPs into maize leaf cells (Figure 4). In samples treated with chemical AgNPs (Figure 4a), clusters of AgNPs were clearly detected within vacuoles (black arrows), indicating cellular uptake and compartmentalization. Meanwhile, in leaf sections from plants treated with biogenic AgNPs (Figure 4b), similar intracellular distribution patterns were observed. As shown in Figure 4b, three adjacent cells exhibit NPs aggregates localized in the central vacuole (black arrows).

It is well known that vacuole serve as intracellular reservoir capable of isolating potentially harmful or foreign substances, thereby minimizing damage to essential organelles such as nuclei and chloroplasts [31,32]. Thus, the absence of NPs within chloroplasts and other critical structures suggests a degree of biocompatibility for both NPs types under the conditions tested in this study. The intracellular sequestration of AgNPs within vacuoles also has implications for their internal transport within the plant. Current evidence indicates that, following foliar application, AgNPs are predominantly retained in leaf tissues, where vacuolar compartmentalization and cell wall associations limit their long-distance translocation [8,33]. In particular, the phloem-mediated transport of intact AgNPs appears to be highly restricted, whereas any potential downward redistribution is thought to occur mainly in ionic form (Ag⁺) or as ultrasmall nanoparticulate species generated through partial dissolution processes [33]. In this context, the localization patterns observed by TEM in the present study are consistent with a limited systemic mobility of AgNPs.

2.3. Physiological Status of Plants Treated with NPs

Exposure to abiotic stressors, including drought, salinity, and toxic metals often triggers an overproduction of ROS in plants, which may comprise growth and induce cellular damage or even cell death. In this context, metal-based NPs have been reported to enhance antioxidant defense mechanisms by upregulating key enzymes that detoxify ROS [7]. Silver nanoparticles possess intrinsic antioxidant properties that enable them to act as ROS scavengers [34]. Excessive ROS accumulation during salt stress can denature proteins, induce lipid peroxidation, and cause DNA damage, ultimately compromising cellular integrity [35]. Several studies have demonstrated that AgNPs enhance antioxidant enzyme activity and protect plant cells from oxidative damage. For instance, chemical AgNPs increased catalase and peroxidase activity in salt-stressed plants, improving their overall antioxidant capacity [36].

In the present study, salt stress imposed by cultivation in 6 mS/cm soil significantly altered the antioxidant status of maize plants (Figure 5). Catalase (CAT) activity decreased from 0.0350 ± 0.0013 ΔA240 g⁻¹ protein min⁻¹ under non-saline conditions (1 mS/cm) to 0.0226 ± 0.0023 ΔA240 g⁻¹ protein min⁻¹ in saline soil, representing a 35% reduction. A similar trend was observed for guaiacol peroxidase (G-POX), whose activity declined from 0.127 ± 0.007 to 0.083 ± 0.009 ΔA420 g⁻¹ protein min⁻¹, corresponding to a 34% decrease.

Foliar application of both biogenic and chemical AgNPs effectively counteracted the inhibitory effects of salinity on antioxidant enzyme activity. G-POX activity was fully restored in plants treated with biogenic (0.128 ± 0.005) and chemical AgNPs (0.127 ± 0.001), reaching values comparable to those observed under non-saline conditions (0.127). Regarding CAT activity, chemical AgNPs induced a partial recovery (0.0295 ± 0.0030), remaining approximately 16% below the non-saline control (0.0350). In contrast, biogenic AgNPs not only restored CAT activity but increased it to 0.0405 ± 0.0040, corresponding to an enhancement of approximately 15% above the non-saline condition.

AgNPs are often perceived by plants as mild stressors, triggering physiological responses that activate defense systems. The induction of antioxidant enzymes such as CAT and G-POX plays a central role in ROS detoxification and cell protection. For instance, biogenic AgNPs obtained from extracts of eucalyptus leaves significantly enhance the activity of these enzymes in salt-stressed plants [10], contributing to redox homeostasis. Furthermore, AgNPs may also stimulate the synthesis of secondary metabolites involved in stress resistance. For example, AgNPs increased the production of flavonoids and phenolic compounds in tomato plants, enhancing their stress tolerance [37,38].

Among the physiological indicators of oxidative stress, lipid peroxidation is one of the most reliable, with MDA being its most abundant byproduct [39]. Therefore, MDA content was quantified in leaves of maize plants belonging to the different treatments applied. As expected, salt stress caused a pronounced increase in MDA levels, which rose from 107.19 ± 8.90 Eq. MDA g⁻¹ in non-stressed plants to 232.34 ± 31.74 Eq. MDA g⁻¹ under saline conditions, representing a 117% increase (Figure 5). This confirmed the oxidative damage caused by salinity. However, treatment with both types of AgNPs markedly reduced lipid peroxidation, restoring MDA concentrations to 102.63 ± 5.75 and 112.74 ± 15.95 Eq. MDA g⁻¹ in plants treated with biogenic and chemical AgNPs, respectively. These values were comparable to those observed in plants grown under non-saline conditions (107.19 ± 8.90 Eq. MDA g⁻¹).

Overall, AgNPs application significantly improved the oxidative status of maize plants exposed to salt stress, as evidenced by the enhancement of antioxidant enzyme activities and the reduction in lipid peroxidation. Notably, biogenic AgNPs consistently exhibited a stronger protective effect across all evaluated physiological parameters, indicating a more efficient activation of antioxidant defense mechanisms under saline conditions.

Another common physiological response of plants to abiotic stress is a decline in photosynthetic capacity, which is frequently associated with alterations in photosynthetic pigment content. To evaluate the impact of salinity and AgNPs treatments on photosynthetic performance, the concentrations of chlorophyll a, chlorophyll b, and total carotenoids were quantified in maize leaf tissue (Figure 5).

Salt stress imposed by cultivation in 6 mS/cm soil significantly reduced chlorophyll a content, which decreased from 18.72 ± 2.55 µg mL⁻¹ under non-saline conditions (1 mS/cm) to 12.62 ± 1.79 µg mL⁻¹, corresponding to a 33% reduction. Foliar application of chemical AgNPs partially alleviated this effect, restoring chlorophyll a level to 17.51 ± 2.97 µg mL⁻¹, which remained approximately 6% lower than those observed in non-saline plants (18.72 ± 2.55 µg mL⁻¹). In contrast, treatment with biogenic AgNPs markedly enhanced chlorophyll a accumulation, reaching 24.19 ± 3.58 µg mL⁻¹, representing an increase of approximately 29% above the non-saline control suggesting an improvement in photosynthetic efficiency even beyond normal growth conditions (Figure 5).

A similar but less pronounced trend was observed for chlorophyll b. Salinity caused a moderate decrease from 5.18 ± 0.91 µg mL⁻¹ (non-saline soil) to 4.65 ± 0.78 µg mL⁻¹ (saline soil), corresponding to a 10% reduction. Both AgNPs treatments significantly increased chlorophyll b content beyond non-saline levels. Plants treated with chemical AgNPs reached 7.72 ± 1.22 µg mL⁻¹, while those treated with biogenic AgNPs exhibited the highest values (8.85 ± 0.80 µg mL⁻¹), corresponding to increases of approximately 49% and 71%, respectively, relative to the non-saline condition (5.18 ± 0.91 µg mL⁻¹).

In contrast to chlorophylls, carotenoid content increased under saline conditions, rising from 2.71 ± 0.38 µg mL⁻¹ in non-stressed plants to 3.47 ± 0.55 µg mL⁻¹ in salt-stressed plants, representing a 28% increase, likely as a compensatory response to reduced chlorophyll levels, enhancing light capture across different wavelengths. Application of AgNPs further modulated carotenoid accumulation. Chemical AgNPs increased carotenoid levels to 3.13 ± 0.32 µg mL⁻¹, corresponding to an increase of approximately 15% relative to the non-saline control (2.71 ± 0.38 µg mL⁻¹), whereas biogenic AgNPs induced a stronger response, reaching 4.36 ± 0.43 µg mL⁻¹, equivalent to an increase of approximately 61% (Figure 5).

Enhancement of photosynthetic efficiency is one of the most significant effects observed in AgNP-treated plants. Increased pigment concentration improves light absorption and CO₂ assimilation, which in turn promotes photosynthetic activity and biomass accumulation. This effect is especially valuable under salt stress conditions, which typically impair photosynthetic function by disrupting the activity of photosystems. AgNPs may help maintaining or even stimulating the activity of these complexes. A recent study by [40] showed that AgNPs enhanced the photosynthetic rate in salt-stressed plants, resulting in improved growth and biomass accumulation, a finding consistent with our results.

The physiological responses observed in maize plants treated with biogenic AgNPs under saline conditions are consistent with previous reports describing nanoparticle-mediated mitigation of salt-induced oxidative stress in crops. Several studies have shown that AgNP application can enhance antioxidant enzyme activities, reduce lipid peroxidation, and preserve photosynthetic pigment content in salt-stressed plants, including maize and other cereals [8,9]. Notably, the stronger response elicited by biogenic AgNPs in the present study aligns with evidence that biologically synthesized nanoparticles often exhibit improved biocompatibility and stress-alleviating capacity compared with chemically synthesized counterparts, likely due to their surface-associated biomolecules [26,41]. These similarities support the general relevance of our findings while highlighting the added value of biogenic AgNPs under saline conditions.

Mechanistically, the protective effect of biogenic AgNPs may be linked to the organic corona naturally inherited from Streptomyces metabolites. Biogenic AgNPs typically bear a mixed surface layer of proteins, peptides, and carbohydrate-like moieties that stabilizes the colloidal system and modulates its interfacial reactivity, including silver ion release and ROS generation [26,41]. This biologically derived surface corona can reduce nonspecific toxicity while favoring compatibility with plant tissues, potentially enabling a more efficient priming or elicitation of plant defense pathways, such as antioxidant enzyme activation and secondary metabolite production, compared with purely citrate-stabilized chemically synthesized AgNPs [12]. In addition, the nature of the surface corona influences the identity of the eco-corona formed upon interaction with biological fluids and the rhizosphere environment, which can in turn shape downstream microbial interactions [26]. Although the composition of the protein/polysaccharide corona were not directly characterized here, the presence of surface-associated biomolecules evidenced by FTIR and Raman analyses supports this explanation and suggests a plausible route by which Streptomyces-derived capping could participate in plant signaling. Future studies integrating corona proteomics/polysaccharidomics will be necessary to establish causal links between specific capping components and biological outcomes.

Given the central role of plant physiological status in regulating rhizosphere processes, the enhanced antioxidant defenses and photosynthetic pigment levels observed in AgNP-treated plants prompted the evaluation of whether these plant responses indirectly influenced the biological health of the surrounding saline soil.

2.4. Effects of AgNPs Treatments on Saline Soil

Plants play a central role in shaping the biological activity of the surrounding soil, particularly in the rhizosphere, mainly through the release of root exudates that modulate microbial metabolism and community structure [42,43]. Therefore, changes in plant physiological status may indirectly translate into alterations in soil microbiological activity. Microbial biomass, in turns, plays a critical role in decomposing soil organic matter and contributes essential nutrients (N, P, S) to plants [44]. In the present study, soils collected after plant growth under saline conditions (6 mS/cm) were analyzed to evaluate whether AgNPs treatments applied to plants influenced soil health indirectly through plant-mediated effects, rather than through direct nanoparticle-microorganism interactions. Nevertheless, we acknowledge that foliar-applied nanoparticles may not be fully confined to plant tissues. A fraction of the sprayed material could reach the soil via droplet drift during application and/or wash-off during irrigation events. Therefore, although the experimental design aimed to minimize direct soil exposure, we did not quantify Ag accumulation/speciation in soil or plant litter, which limits our ability to fully exclude a direct contribution of Ag-containing particles to the observed soil responses. Future work should explicitly address this point by tracking Ag in bulk soil and rhizosphere (e.g., ICP-MS) and, when possible, distinguishing particulate versus ionic forms (e.g., SP-ICP-MS), alongside mass-balance approaches following foliar application.

Soil enzymatic activity was first evaluated using the fluorescein diacetate (FDA) hydrolysis assay, a widely used indicator of overall microbial metabolic activity and soil health [45]. This assay is based on the hydrolysis of FDA by extracellular or membrane-bound enzymes. The higher the amount of fluorescein released, the greater the enzymatic activity, which reflects better soil health. Soils associated with plants treated with biogenic AgNPs showed a marked increase in FDA hydrolysis, reaching 238.5 ± 22.18 µg fluorescein mL⁻¹, compared to 160.17 ± 18.03 µg mL⁻¹ in saline control soils, representing an increase of approximately 49% (Figure 6), suggesting a positive effect on soil health. In contrast, soils associated with plants treated with chemical AgNPs exhibited significantly lower FDA activity (130.44 ± 13.65 µg mL⁻¹), corresponding to a 19% decrease relative to the saline control (160.17 ± 18.03 µg mL⁻¹). These results indicate that the physiological status of plants treated with biogenic AgNPs favored higher soil enzymatic activity, whereas chemical AgNPs were associated with reduced microbial metabolic potential and, consequently, suggesting a decline in soil health.

Soil respiration, assessed by quantifying CO₂ release from soil microbial biomass, was used as an additional indicator of overall biological activity [46]. Higher CO₂ release reflects increased microbial activity. No significant differences were observed between saline control soils (410.38 ± 30.25 µg CO₂ mL⁻¹) and soils associated with plants treated with chemical AgNPs (407.72 ± 19.25 µg CO₂ mL⁻¹) (Figure 6). In contrast, soils from plants treated with biogenic AgNPs exhibited significantly higher respiration rates, reaching 492.46 ± 29.10 µg CO₂ mL⁻¹, which represents an increase of approximately 20% compared to the saline control (410.38 ± 30.25 µg CO₂ mL⁻¹). This enhanced respiratory activity is consistent with the increased FDA hydrolysis and suggests a higher overall microbial metabolic activity in soils associated with biogenic AgNP-treated plants.

The observed improvements in soil biological activity are unlikely to result from a direct effect of AgNPs on soil microorganisms, as nanoparticles are known to be largely sequestered within plant cellular compartments. Instead, these effects are more plausibly explained by indirect, plant-mediated mechanisms. Plants experiencing reduced oxidative stress and improved physiological performance often exhibit qualitative and quantitative changes in root exudation, including increased release of carbohydrates, organic acids, amino acids, and signaling molecules which serve as substrates for soil microorganisms and stimulate microbial activity in the rhizosphere [47,48]. In this context, the enhanced plant performance observed in response to biogenic AgNPs may have promoted a more favorable rhizospheric environment, stimulating microbial respiration and enzymatic activity. Conversely, the poorer physiological status of plants treated with chemically synthesized AgNPs, which lack surface-associated microbial molecules, could have resulted in reduced or altered root exudation patterns, thereby limiting microbial metabolic activity. Although root exudate composition was not directly analyzed in this study, these findings support the hypothesis that biogenic AgNPs modulate soil biological processes through improved plant–soil interactions, an aspect that warrants further investigation.

To evaluate potential ecotoxicological implications, a germination bioassay was performed using Lactuca sativa seeds. Germination percentages in soils associated with saline control plants (66.67 ± 7.06%) and biogenic AgNP-treated plants (67.78 ± 7.13%) were statistically comparable (Figure 6). In contrast, soils associated with plants treated with chemical AgNPs showed a significantly lower germination rate (41.67 ± 4.50%), suggesting a potential ecotoxic effect. This response may be linked to changes in soil biological activity or to residual phytotoxic compounds although these results should be interpreted with caution. In this study, soil samples were not analyzed for total or bioavailable silver contents; therefore, we cannot rule out the presence of residual Ag in the soil. Such residues could contribute to phytotoxic responses. However, it is noteworthy that soils from plants treated with biogenic AgNPs did not exhibit reduced germination, suggesting that factors other than silver concentration per se (e.g., differences in NPs coatings, transformations, or soil–microbe interactions) may underlie the contrasting responses.

Overall, these results demonstrate that AgNPs treatments modulate soil biological activity indirectly by affecting plant health, which in turn influences the metabolic activity of the surrounding soil. Notably, biogenic AgNPs promoted a more favorable plant–soil interaction under saline conditions, resulting in enhanced soil enzymatic activity and respiration without detectable ecotoxic effects, whereas chemical AgNPs were associated with detrimental outcomes.

To further evaluate whether AgNPs treatments applied to maize plants under saline conditions were associated with changes in soil microbial communities, a metabarcoding analysis was performed on soils.

Taxonomic profiling at the phylum level (Figure 7a) revealed a conserved core microbiota across all treatments, dominated by Pseudomonadota, Verrucomicrobiota, and Bacillota, followed by Bacteroidota and Actinomycetota. Although relative abundances varied moderately among treatments, soils associated with AgNPs-treated plants showed a slight increase in phyla commonly linked to stress-adapted or oligotrophic environments, such as Chloroflexota and Thermoproteota, compared to the saline control. These changes occurred without major restructuring of the community, consistent with the modest variation observed in alpha diversity metrics.

Alpha diversity analyses revealed no pronounced differences among treatments (Figure 7b). Faith’s phylogenetic diversity ranged from 153.44 ± 6.66 in saline control soils to 158.82 ± 7.48 and 158.00 ± 11.92 in soils associated with plants treated with biogenic and chemical AgNPs, respectively. Similarly, the number of observed features showed comparable values across treatments, averaging 2747.91 ± 50.46 in the saline control, 2739.19 ± 72.04 in the biogenic AgNPs group, and 2784.95 ± 124.44 in the chemical AgNPs group. Shannon diversity index values were also highly similar among treatments, ranging between 9.51 and 9.55, indicating that overall bacterial diversity was largely conserved under all conditions (Figure 7b).

Despite the absence of strong differences in richness and diversity, Pielou’s evenness index exhibited a consistent increasing trend in soils associated with AgNPs-treated plants. Evenness increased from 0.824 ± 0.002 in saline control soils to 0.831 ± 0.009 and 0.838 ± 0.001 in soils corresponding to biogenic and chemical AgNPs treatments, respectively. This pattern suggests a more homogeneous distribution of taxa within the microbial community, potentially reflecting subtle shifts in resource availability or niche occupation.

Consistently, beta diversity analysis based on Bray–Curtis distances indicated no clear separation among communities in the original ordination, with a non-significant PERMANOVA result (F = 0.953; p = 0.589).

Overall, the metabarcoding results indicate that AgNPs treatments did not drastically alter the global diversity of saline soil bacterial communities. Instead, they were associated with subtle but coherent shifts in community structure and evenness, likely driven by changes in plant physiological status. This interpretation aligns with the soil enzymatic activity, respiration, and germination assays, which collectively indicate more favorable biological activity indicators, despite only subtle shifts in bacterial community structure in soils previously associated with plants treated with biogenic AgNPs.

This study was conceived as a proof-of-concept exploration of the potential of biogenic AgNPs to mitigate salt stress in maize and improve soil biological quality. Future research will include detailed analyses of AgNP stability, Ag⁺ dissolution kinetics, and post-application size and aggregation dynamics to deepen our understanding of nanoparticles fate and to optimize their agronomic use.

In contrast to most previous studies addressing AgNP-mediated stress mitigation in plants, this work provides an integrated evaluation of plant physiological responses and soil biological indicators under naturally saline conditions. The comparative analysis between biogenic and chemically synthesized AgNPs demonstrates that surface functionalization inherited from Streptomyces biomolecules plays a key role in enhancing plant performance while preserving soil biological quality. This combined plant–soil perspective represents a relevant advancement toward the sustainable use of nanomaterials in agriculture.

In this context, environmental and agronomic considerations are essential before extrapolating these findings to real field conditions. In the present study, AgNPs were evaluated under greenhouse conditions and over a limited experimental timeframe, which is appropriate for mechanistic assessment but does not capture the cumulative effects that may arise from repeated AgNP applications across seasons. In agricultural settings, nanosilver inputs could accumulate in soils over time depending on application rate, frequency, soil properties (e.g., organic matter, soil texture, cation exchange capacity), and transformation processes that govern Ag bioavailability and persistence [49,50]. Although sulfidation and complexation can decrease the short-term bioavailability of Ag, they may also increase persistence in soils, highlighting the need for long-term monitoring under realistic exposure scenarios [50].

Repeated AgNPs applications may also pose potential risks to non-target soil biota and key microbial functions. The antimicrobial activity of AgNPs and/or released Ag⁺ has been reported to affect soil microbial communities and nitrogen-cycle processes (e.g., nitrification/denitrification) in a concentration-, coating-, and soil-dependent manner [51,52,53]. Therefore, even when short-term assays indicate low ecotoxicity, chronic exposure could lead to functional trade-offs, shifts in sensitive taxa, or altered nutrient cycling under certain field conditions. Importantly, these risks cannot be fully resolved by short-term greenhouse experiments and should be addressed through multi-season field trials combined with fate and effects assessments.

From an agronomic perspective, these considerations support a precautionary framework in which biogenic AgNPs, despite their favorable performance in this study, should be evaluated using the minimum effective dose, optimized application timing to minimize wash-off/runoff, and soil-type-specific risk assessment. Recent work has also advanced hazard-threshold approaches for silver nanomaterials in soils, emphasizing the value of integrating exposure estimates with standardized ecotoxicological and functional endpoints [54]. Overall, our results provide a basis for further development of biogenic AgNPs as nanobiotechnological tools, but their field-scale deployment should be guided by long-term validation and environmental risk evaluation under realistic agronomic practices.

3. Materials and Methods

The potential of biogenic AgNPs as biostimulants under salt stress relies heavily on their physicochemical properties. Thus, we first confirmed the successful synthesis of AgNPs using Streptomyces sp. Z38 and conducted comprehensive characterization analyses to evaluate their morphology, size distribution, surface functionalization, and elemental composition.

3.1. Biosynthesis of Silver Nanoparticles

Biogenic AgNPs were synthesized by green synthesis using biomolecules produced by a rhizosphere actinobacterium isolated from plants grown on soils polluted with heavy metals and organochlorine pesticides [55]. Streptomyces sp. Z38 was cultivated on Tryptic Soy Broth (TSB) at 30 °C, 150 rpm for 72 h. The cells were harvested by centrifugation and washed twice with PBS buffer pH 7. Silver nanoparticles were synthesized as described before [15]. Briefly, washed pellets were resuspended in triple-distilled water and incubated at 30 °C, 180 rpm for 120 h. Following centrifugation, the cell-free supernatant containing bioactive compounds [bioactive water (BW)] was used for the biosynthesis of AgNPs. AgNO₃ solution was added to the BW to achieve a final concentration of 1 mM. The solution was then incubated in the dark at 30 °C and 150 rpm for 72 h. A visual color change from colorless to brown was considered indicative of AgNPs formation. The mixture was centrifuged at 12,500× g for 15 min at room temperature. The supernatant was discarded, and the AgNPs obtained were washed with distilled water and stored at 4 °C for further studies.

Chemical AgNPs were prepared by the well-known Turkevich method, using citrate as a reducing agent [56,57]. Briefly, 500 mL of a 1 mM AgNO₃ (Merck KGaA, Darmstadt, Germany) solution in deionized water was heated until it started to boil. After that, 50 mL of an aqueous solution of 10 mM sodium citrate (Merck KGaA) was added dropwise to the silver nitrate solution. The heating was continued for 10 min, and then cooled to room temperature. Then, the mixture was centrifuged at 12,500× g for 15 min at room temperature. The supernatant was discarded, and the chemical AgNPs obtained were washed with distilled water and stored at 4 °C for further studies.

3.2. Characterization of Silver Nanoparticles

3.2.1. UV–Visible Spectroscopy

The production of biogenic AgNPs was monitored through UV-visible spectroscopy (Libra S80, Biochrom Ltd., Cambridge, UK) in the range of 350–600 nm (1 nm interval) to identify the characteristic plasmon resonance band of AgNPs, as described previously [14]. Absorbance measurements were performed at intervals of 24, 48, and 72 h for biogenic AgNPs.

3.2.2. Scanning Electron Microscopy (SEM), Dynamic Light Scattering (DLS) and Zeta Potential

The presence, morphology and size of AgNPs were assessed by SEM with a JEOL JSM-35 CF instrument (Oberkochen, Germany) located in CIME (CCT-CONICET). The diameter of AgNPs (n = 52) was measured from the obtained images by using the free image-processing software, ImageJ (version 1.52a) [58].

The hydrodynamic diameter and surface charge (zeta potential) of the AgNPs were measured using a SZ-100 Horiba instrument located in CIBAAL (CONICET-UNSE). The samples were dispersed in distilled water (0.5 mg mL⁻¹) and sonicated for 10 min to avoid aggregation. Zeta potential measurements were performed at a detection angle of 170°, and at least 100 measurements were recorded over 30 s per sample.

3.2.3. Fourier Transform Infrared (FTIR) and Raman Spectroscopy

Molecular structures and chemical bonds of the AgNPs were investigated using attenuated total reflectance (ATR)-FTIR. Spectra were recorded using a Thermo Scientific 6700 spectrometer (Thermo Fisher Scientific, Waltham, MA, USA) with an ATR accessory and a deuterated triglycine sulfate (DTGS) detector, connected to a dry air circulation system to avoid interference from water vapor and carbon dioxide. Raman spectroscopy was performed with a Confocal Raman Thermoscientific DXR microscope (Thermo Fisher Scientific, Waltham, MA, USA) located in FBQyF (UNT).

3.2.4. Quantification of Silver

The silver content in the AgNPs was determined using inductively coupled plasma-mass spectrometry (ICP-MS). For this purpose, AgNPs were submitted to acid digestion in a 1000 W microwave oven (MSP microwave oven, CEM, Matthews, NC, USA) using closed vessels containing 1 mL of concentrated HNO₃ and 0.5 mL of 30% (v/v) H₂O₂ [59]. The analysis was conducted at INQUISAL (CONICET, UNSL). The resulting solutions were cooled, diluted with MilliQ water and analyzed for the Ag concentration with an Agilent 7700-collision/reaction cell ICP-MS (Agilent Technologies, Santa Clara, CA, USA). Hydrogen gas was employed as collision gas for Ag determination.

3.3. Plant Assay

3.3.1. Soil

Two natural soils were used to reach two contrasting saline conditions: one with a normal conductivity (1 mS/cm) collected from southern Tucumán, Argentina (27°11′13.2′′ S, 65°14′01.5′′ W), and another with extreme conductivity (75.4 mS/cm) sourced from Santiago del Estero, Argentina (26°17′00.2′′ S, 64°10′06.4′′ W). The soils were air-dried at 30 °C, manually sieved to remove debris, and homogenized.

Mixtures of both soils were prepared in different proportions to obtain a final conductivity of 6 mS/cm. The high-conductivity soil (75.4 mS/cm) was ground to break apart salt-induced aggregates. Electrical conductivity (EC) was measured using the 1:2 soil-to-bidistilled-water method [60]. Briefly, the mixtures were allowed to stand for 30 min, filtered through paper to remove soil particles, and the resulting extracts were used to determine pH and EC with a pH meter and a conductivity meter. All measurements were performed in quintuplicate.

The physicochemical characterization of the 6 mS/cm soil was conducted by an external certified laboratory (Tecnosuelo, Tucumán, Argentina). The soil was classified as silty clay loam based on capillary conductivity analysis and showed a pH of 6.87 (potentiometry), absence of detectable carbonates (calcimetry), and organic matter content of 3.34% (Walkley–Black method), total nitrogen of 0.233% (Kjeldahl method), available phosphorus of 16.7 mg Kg⁻¹ (Bray–Kurtz method), and exchangeable potassium of 1.04 cmol_c kg⁻¹ (Morgan method). Soil salinity (6 mS/cm), determined as the EC of the saturation extract, was also confirmed by the same external laboratory.

3.3.2. Seeds Preparation and Growth Conditions for Zea mays

Zea mays seeds (white sweet variety) were obtained from Fecoagro Cooperative, San Juan, Argentina. Seeds were sterilized superficially using the standardized protocol ethanol-sodium hypochlorite solution [61]. Seeds were dried in sterile conditions and stored in the dark at 4 °C until use.

For the plant growth experiments, 200 g of saline soil (6 mS/cm) was placed in individual pots. A single seed was sown per pot, and 50 mL of Hoagland nutrient solution [62] was added to ensure uniform initial moisture. Plants were maintained in a controlled environment chamber [12:12 (L:D photoperiod), 30 °C] and watered with tap water every three days for 30 days. On day 15, a single foliar application of the following solutions/treatments was performed: (1) Biogenic AgNPs (75 mg L⁻¹), (2) chemical AgNPs (75 mg L⁻¹) suspended both in sterile distilled water, and (3) Sterile distilled water (saline control). Additionally, a fourth treatment consisting of plants grown on 1 mS/cm non-saline soil was added; these plants were sprayed with sterile distilled water (non-saline control). All AgNP treatments (biogenic and chemical) were applied to plants grown exclusively on saline soil (6 mS/cm), whereas the non-saline soil (1 mS/cm) treatment was included only as a physiological control and did not receive nanoparticle application. Each treatment consisted of 15 plants, and was sprayed with 5 mL of NPs or sterile distilled water, resulting in a total of 60 plants. All plant- and soil-related analyses described below were conducted on 30-day-old plants, corresponding to the end of the experimental period.

3.4. Parameters Used to Evaluate the Effects of Nanoparticles on Plants

3.4.1. Plant Growth

At the end of the 30-day growth period, plants were harvested, and roots were carefully washed to remove residual soil. The lengths of the longest root and leaf were measured for each plant using a millimeter scale. Samples of roots and leaves from 10 plants per treatment were oven-dried at 80 °C until constant weight. Dry weights were measured using an analytical balance.

3.4.2. Transmission Electron Microscopy (TEM) Images of Leaves

To evaluate the uptake and distribution of AgNPs in plants, leaf sections were examined by TEM. For this purpose, fresh leaf tissues were collected and prepared following standard protocols. Analysis was performed using a Zeiss Libra 120 TEM (Carl Zeiss AG, Oberkochen, Germany) at the CIME (CCT-CONICET).

3.4.3. Physiological Status of Plants: Analysis on Leaf Tissues

Catalase (CAT) Activity

Catalase activity, which decomposes hydrogen peroxide into water and oxygen, was measured following a method previously described [63]. Fresh leaf tissue (1 g) was homogenized in 3 mL of 25 mM Tris-HCl buffer (pH 7.2) containing 5 mM dithiothreitol (DTT) and 5 mM EDTA. The homogenate was centrifuged at 12,000× g for 10 min, and the supernatant was used as the enzyme extract. The reaction mixture (2.9 mL) consisted of 50 mM sodium phosphate buffer (pH 7.0) and 10 mM H₂O₂, to which 100 µL of enzyme extract was added. The reaction was carried out at 25 °C, and the decrease in absorbance at 240 nm was monitored for 4 min. Catalase activity was expressed as the change in absorbance per gram of protein per minute (ΔA240 g⁻¹ prot. min⁻¹). All measurements were performed in quintuplicate.

Guaiacol Peroxidase (G-POX) Activity

The activity of guaiacol peroxidase, an enzyme involved in oxidative stress response, was determined using the method of Dhir [63]. Fresh leaf tissue (1 g) was homogenized in 3 mL of 25 mM Tris-HCl buffer (pH 7.2) with 5 mM DTT and 5 mM EDTA. The homogenate was centrifuged at 12,000× g for 10 min, and the supernatant was used as the enzyme extract. The reaction mixture (2 mL) consisted of 50 mM sodium phosphate buffer (pH 6.0), 60 mM guaiacol, and 300 µL of 10-volume H₂O₂ (~0.9 M). The reaction was initiated by adding 100 µL of the enzyme extract. The increase in absorbance at 420 nm was recorded over 5 min. G-POX activity was expressed as the change in absorbance per gram of protein per minute (ΔA420 g⁻¹ prot. min⁻¹). All measurements were performed in quintuplicate.

Malondialdehyde (MDA) Content

The malondialdehyde (MDA) content, an indicator of lipid peroxidation and oxidative stress, was determined using the method of Du and Bramlage [64]. Fresh leaf tissue (1 g) was homogenized in 3 mL of 0.1% trichloroacetic acid (TCA) using a chilled porcelain mortar. The homogenate was centrifuged at 12,000× g for 5 min, and the supernatant was collected. For MDA quantification, 1 mL of the supernatant was mixed with 1 mL of thiobarbituric acid (TBA) reagent (0.5 g of TBA dissolved in 100 mL of 20% TCA). The mixture was incubated in a water bath at 90 °C for 20 min, cooled on ice, and centrifuged at 12,000× g for 10 min. The absorbance of the supernatant was measured at 440, 532, and 600 nm using a UV-visible spectrophotometer (Hitachi U-2800A, Hitachi High-Tech Corporation, Tokyo, Japan). The MDA concentration was calculated using the molar extinction coefficient (1.57 × 10⁵ M⁻¹ cm⁻¹) and expressed as equivalents (Eq) of MDA per gram of fresh weight (FW). All measurements were performed in quintuplicate.

3.4.4. Pigment Quantification

Chlorophyll and carotenoid pigments were extracted from 20 mg of fresh leaf tissue using dimethyl sulfoxide (DMSO) following the method of Chappelle [65]. Samples were incubated in 2 mL DMSO at 45 °C for 12 h in the dark. Absorbance was measured at 665, 649, and 480 nm against a DMSO blank using a UV-visible spectrophotometer (Hitachi U-2800A, Tokyo, Japan). The concentrations of chlorophyll A, chlorophyll B, and total chlorophyll were calculated using the Wellburn equations [65]. All measurements were performed in triplicate.

3.5. Parameters Used to Evaluate Indirect Effects of Nanoparticles on the Soils Remaining from the Plant Experiment

As previously mentioned, at the end of the 30-day growth period, maize plants were harvested and the roots were carefully washed to remove residual soil. Prior to washing, both root-adhering soil (rhizosphere) and the surrounding root soil were collected. The soil samples were sieved (<4 mm) and stored at 4 °C until further processing.

3.5.1. Total Microbial Activity in the Soil

Total enzymatic activity in the remaining soils was quantified using the fluorescein diacetate (FDA) hydrolysis method, as described by Adam and Duncan [45]. For each soil sample (1 g, triplicated), 10 mL of 60 mM sodium phosphate buffer (pH 7.6) and 100 µL of 2 mg mL⁻¹ FDA solution were added. The mixture was incubated at 25 °C for 2 h with continuous shaking. The reaction was stopped by adding 10 mL of acetone, and the mixture was filtered. The filtrate’s absorbance was measured at 490 nm using a spectrophotometer (Hitachi U-2800A, Tokyo, Japan). A calibration curve prepared with fluorescein standards was used to calculate the concentration of fluorescein released, which served as an indicator of total microbial activity.

3.5.2. Respiration

Soil respiration, indicative of microbial metabolic activity, was measured using the CO₂ release method as described by Anderson (1982) [46]. Soil samples (20 g) were placed in plastic containers and moistened with 5 mL of distilled water. The containers were placed inside glass jars containing 30 mL of 0.1 N sodium hydroxide (NaOH) to trap the CO₂ released during a 10-day incubation at 30 °C. After incubation, the NaOH solution was titrated with 0.1 N hydrochloric acid (HCl) to quantify the CO₂ absorbed. All measurements were performed in triplicate.

3.5.3. Ecotoxicity

The ecotoxicity of soils cultivated with maize plants treated with biogenic AgNPs and their respective controls was assessed using Lactuca sativa (lettuce) as a standard bioindicator species. Lettuce seeds were purchased in Fecoagro Cooperative, San Juan, Argentina. Prior to the assay, soil samples were homogenized and passed through a 2 mm sieve, ensuring uniformity among replicates. For each treatment, 10 g of soil was placed in sterile Petri dishes (90 mm diameter). Soil moisture was adjusted to approximately 60% of the water-holding capacity with tap water to ensure adequate conditions for seed germination. Thirty L. sativa seeds of uniform size and appearance were evenly distributed on the soil surface in each dish. Seeds were gently pressed into the soil to ensure adequate contact. Dishes were capped, then covered with aluminum foil to prevent moisture loss and incubated in a chamber with environmentally controlled conditions (darkness, 25 °C) for five days. Each treatment included three independent replicates. At the end of the incubation period, germination was assessed by counting the number of seeds with radicle emergence ≥ 5 mm, following standard germination criteria. The germination percentage (G%) was calculated as G% = Ng/Nt × 100, where Ng is the number of germinated seeds and Nt is the total number of seeds [62].

3.5.4. Metabarcoding Analysis of Saline-Treated Soils

Total DNA was extracted from 0.25 g of soil from three independent biological replicates per saline soil treatment (6 mS/cm) using the DNA PowerSoil PRO KIT (QIAGEN, Hilden, Germany) according to the manufacturer’s instructions. DNA quality was measured using a nanodrop (Multiskan™ GO Microplate Spectrophotometer, Thermo Fisher Scientific, Waltham, MA, USA).

PCR amplification targeting the V3-V4 regions of the 16S rRNA gene and sequencing were performed by Novogene Corporation Ltd. (Beijing, China) using primers 341F (5′-CCTAYGGGRBGCASCAG-3′) and 806R (5′-GGACTACNNGGGTATCTAAT-3′) [66]. Sequencing libraries were generated using TruSeq^® DNA PCR-Free Sample Preparation Kit (Illumina, San Diego, CA, USA) following manufacturer’s recommendations, and index codes were added. Library quality was assessed using a Qubit@ 2.0 Fluorometer (Thermo Scientific) and an Agilent Bioanalyzer 2100 system. Libraries were sequenced on an Illumina NovaSeq platform, generating 250 bp paired-end reads.

Sequencing reads were analyzed using the QIIME2 pipeline [67]. Initially, the quality of the reads was screened using the demux plugin, and low-quality reads with Q value < 25 were trimmed. After this, the denoised reads were grouped into amplicon sequence variants (ASVs). The resulting ASVs were classified using the naive Bayesian classifier provided by QIIME2 and the taxonomic identification was carried out using the Silva database (v138.2) as reference.

Alpha diversity metrics (Observed features, Faith’s phylogenetic diversity, and Shannon index) and beta diversity metrics (unweighted and weighted UniFrac distances) were calculated in QIIME2. Statistical differences in alpha diversity among treatments were evaluated using the Kruskal–Wallis test, while differences in beta diversity were assessed using PERMANOVA.

3.6. Statistics

All results are expressed as mean ± SD. Prior to statistical analysis, data were tested for normality and homogeneity of variances using the Shapiro–Wilk and Levene’s tests, respectively. When these assumptions were met, statistical analyses were conducted using one-way analysis of variance (ANOVA) followed by Tukey’s post hoc test, implemented in MINITAB 21 Statistical Software (Minitab, State College, PA, USA). Statistical significance was accepted at p < 0.05.

4. Conclusions

The results of this study demonstrate that foliar application of AgNPs modulates the response of maize plants to salinity at multiple, interconnected levels. Improvements in shoot and root growth, together with enhanced photosynthetic pigment content and reinforced antioxidant defenses, indicate that AgNPs (particularly biogenic AgNPs) effectively alleviated salt-induced physiological constraints. These plant-level benefits were not accompanied by evidence of direct nanoparticle–microorganism interactions, but instead translated into indirect effects on the surrounding saline soil. Soils associated with AgNP-treated plants, especially those receiving biogenic AgNPs, exhibited higher enzymatic activity, increased microbial respiration, and no detectable ecotoxic effects, reflecting a more favorable biological status. Consistently, metabarcoding analysis revealed that these improvements were associated with subtle shifts in bacterial community structure and evenness rather than with major changes in overall diversity, supporting a model in which enhanced plant health under AgNPs treatment promotes a more balanced and functionally resilient rhizosphere. Together, these findings highlight the potential of biogenic AgNPs as a plant-centered strategy to mitigate salinity stress while preserving soil biological integrity. From an applied perspective, the present findings suggest that biogenic AgNPs may represent a promising nanobiotechnological tool for improving plant performance under saline conditions within a sustainable agriculture framework. Due to their biological origin and favorable compatibility with plant physiological processes and soil biological activity observed in this study, biogenic AgNPs could potentially contribute to salinity management strategies, particularly in low-input or organic-oriented agricultural systems. Nevertheless, their practical implementation should be approached cautiously and supported by field-scale validation, dose optimization, and comprehensive environmental risk assessment to ensure long-term agronomic efficacy and ecological safety.