Evidence of Cooperative Interactions between Rhizobacteria and Wood-Decaying Fungi and Their Effects on Maize Germination and Growth

Abstract

Advances in soil microbial communities are driving agricultural practices towards ecological sustainability and productivity, with engineering microbial communities significantly contributing to sustainable agriculture. This study explored the combined effects of two white-rot fungi (Trametes sp. and Pleurotus sp.) and six rhizobacterial strains belonging to four genera (Acinetobacter sp., Enterobacter sp., Flavobacterium sp., and Pseudomonas sp.) on maize growth and soil enzymatic activity over a 14-day period. At the plant level, germination, fresh and dry mass of the aerial and root parts, length, and stage of development of the stem, as well as the chlorophyll content, were evaluated. Furthermore, soil dehydrogenase, acid and alkaline phosphatases, pH, and electrical conductivity were evaluated. Rot fungi induced distinct effects on maize germination, with Pleurotus sp. strongly suppressing maize germination by 40% relative to that of the control. The isolated bacterial strains, except Enterobacter sp. O8, and 8 of the 12 fungus + bacterial strain combinations induced germination rates higher than those of the control (≥40%). Combinations of Flavobacterium sp. I57 and Pseudomonas sp. O81 with the rot fungus Pleurotus sp. significantly improved plant shoot length (from 28.0 to 37.0 cm) and developmental stage (fourth leaf length increase from 10.0 to 16.8 cm), respectively, compared with the same bacteria alone or in combination with the rot fungus Trametes sp. In the soil, the presence of both fungi appeared to stabilize phosphatase activity compared to their activity when only bacteria were present, while also promoting overall dehydrogenase enzymatic activity in the soil. Integrating all parameters, Trametes sp. rot fungus + Enterobacter sp. O8 may be a potential combination to be explored in the context of agricultural production, and future studies should focus on the consistency of this combination’s performance over time and its effectiveness in the field.

1. Introduction

Soil ecosystems are currently facing escalating pressures, such as soil degradation, water scarcity, and the persisting chemical input from agriculture, as outlined in European directives [1]. These pressures originate from multiple sources, including low soil recycling rates, intensified agricultural practices, and the overarching impact of climate change [2,3]. Recent reports have revealed a concerning statistic: between 60% and 70% of soil across Europe is in a state deemed unhealthy [1,2,3]. This degradation stems from a complex interplay of several factors, such as organic matter depletion, erosion, salinization, compaction, and contamination, as well as soil biodiversity reduction [1,4].

Modern agriculture is undergoing a significant transition to practices that prioritize ecological balance and productivity, methods that are primarily supported by a deeper understanding of soil microbiota. Harnessing the potential of soil organisms is emerging as an important strategy to mitigate these problems. Within the soil microbiota, bacteria and fungi are the most common organisms, being fundamental to soil health and fertility. The use of microorganisms in agriculture offers a diverse spectrum of benefits, including the possibility of reducing chemical inputs regularly used in conventional agricultural practices [4]. Contemporary agricultural practices still rely on chemical inputs that further disrupt soil microbial communities and harm ecosystem functions. Bioformulations can play a significant role not only in plant growth but also in suppressing pests and diseases throughout their life cycle. Thus, microbiota holds great potential for soil remediation and the production of better-quality and nutritionally valuable consumer items [5].

Bacteria constitute vital components of the soil microbiota, playing key roles in several ecological processes crucial to soil health and productivity, such as nutrient cycling, mineral solubilization, and disease suppression in soil ecosystems. Of particular importance is the ability of bacteria to fix atmospheric nitrogen and convert it into biologically available forms, thereby improving soil fertility [6]. However, bacteria may have reduced dispersion capacity compared to rot fungi, which may render them a very localized action; however, their utilization in the host is systemic in nature [7,8]. These disadvantages can be neutralized if combined with other microorganisms, namely wood-decaying fungi. Saprotrophic fungi, the main ones responsible for converting recalcitrant carbon in the soil (such as lignin and cellulose), may not act directly on plant growth or vitality but may complement the functions of bacteria through their hyphal network extension. The hyphal network facilitates the rapid colonization of new areas of soil, the efficient exchange of nutrients, and to the aggregation of soil particles, improving soil stability and water retention [9,10]. Moreover, an important interdependence between fungi and bacteria stands out, with fungi creating favorable conditions for bacterial mobility and activity and hyphal networks of rot fungi functioning as dispersing routes for bacteria [4,11]. The bacteria may benefit from the liquid film surrounding the hyphae to bridge air-filled or dry spaces [4,11]. Taking advantage of this symbiotic relationship between rot fungi and soil bacteria allows for the optimization of crop productivity while minimizing environmental impact, thus aligning with the most recent agricultural strategies for soil regeneration. Despite this, the understanding of the combined action of rhizobacteria with rot fungi remains a field requiring a large input of knowledge [4].

The effective implementation of symbiosis-based approaches demands further scientific investigation to address the prevailing knowledge gaps in this field. Insights are needed into the potential for ideal combinations of rot fungus and bacteria, as well as their ideal proportions. One rot fungi function explored so far is that of white rot fungi to be used as a carrier to introduce and stabilize the plant growth-promoting bacteria in the rhizosphere [12,13]. The study of Rojas-Higuera [13] with four white-rot fungi (Ganoderma lucidum, Pleurotus ostreatus, Trametes versicolor, and Phanerochaete chrysosporium) allowed us to understand the role of rot fungi as potential striking factors for bacteria survival and resilience over time. When in a consortium, all four fungi shaped organic matter and provided suitable conditions for Enterobacter sp. and Stenotrophomonas maltophilia populations to maintain their biological activity at 23 °C for at least 60 days [13]. Furthermore, the authors detected through scanning electron microscopy that, despite the decrease in bacterial concentration by almost five logs, the bacterial cells remained within the hyphal network even after six months [13]. Other factors, such as formulation stability, compatibility with existing agricultural practices, and long-term effects on soil health, require thorough investigation.

The development of new bioformulations that harness the potential of a wide range of microorganisms is extremely important for advancing agricultural sustainability and aligning with European directives for a greener future [1]. In light of pressing global challenges, such as climate change and resource scarcity, it is imperative to discover innovative solutions that increase agricultural productivity while minimizing environmental impact. Thus, this study aimed to explore the growth-enhancing action of soil bacteria and rot fungi in agricultural crops when applied in combination. To this end, the application of four genera of soil bacteria and two rot fungi was evaluated, either alone (each bacterial and fungal genus) or in combination (fungus and bacteria), for the growth of a crop of global importance, such as maize. The selection of these bacterial strains and rot fungi was based on previous studies. These studies were dedicated to the extensive characterization of the growth-promoting and plant-protective properties of bacteria [14] and to the study of the effects of applying rot fungal exudates to soils under osmotic stress [15,16]. Zea mays was used as a surrogate crop species to assess the potential promotion of growth induction by bacteria and rot fungi alone and in combination since it is one of the three most produced cereals worldwide [17]. Compared to the other main grains, maize had the largest output (1.2 billion metric tons in 2021) and the quickest increase since 2000 (about 104% increase), owing, in addition to the human food supply, to its larger use in the biofuel and animal feed industries [17].

2. Materials and Methods

2.1. Bacteria Cultures

Six bacterial strains belonging to four genera (Acinetobacter sp., Enterobacter sp., Flavobacterium sp., and Pseudomonas sp.) were used in this study. These bacterial strains were isolated in a previous study [14] with the following accession numbers: ON419247 (Flavobacterium sp., I57), ON419156 (Enterobacter sp., O8), ON419161 (Enterobacter sp., I2), ON419276 (Pseudomonas sp., O81), ON419180 (Pseudomonas sp., N26), and PP990354 (Acinetobacter sp. S2). Identification of the bacterial strains was performed in [14] using 16S rRNA gene amplification.

Bacterial cultures were maintained on tryptic soy agar (TSA), a general nutritious growth medium, at 26 °C in the dark for 24 h [18]. After that, a loopful of each bacterial strain was used to inoculate 10 mL of tryptic soy broth (TSB at pH 7–7.5, molecular biology grade, Merck, Darmstadt, Germany). Tubes were incubated at 26 °C in the dark for 24 h at 180 rpm (GFL orbital shaker, Burgwedel, Germany; Figure S1) [18]. For later applications (see Section 2.3), the bacterial cultures were centrifuged at 4000 rpm for 10 min (Thermo Scientific, Waltham, MA, USA), and the supernatant was discarded. The cells were then washed with sterile phosphate-buffered saline (PBS at pH 7.4–7.6; molecular biology grade, Merck, Darmstadt, Germany) solution, the centrifugation process was repeated, and the cells were finally resuspended in sterile PBS solution (Figure S1).

2.2. Fungi Cultures

Two terrestrial species of basidiomycete fungi, Trametes sp. and Pleurotus sp., were used in this study and were obtained from culture collections at UNESP, Universidade Estadual Paulista, Brazil, and the BCCMTM/MUCL Culture Collection, Belgium, respectively. Cultures of both species were maintained on potato dextrose agar (PDA; molecular biology grade, Merck, Darmstadt, Germany) at 28 °C in the dark for 5–8 days and weekly renewed [16]. To obtain substantial amounts of biomass to be used in the following assays, actively growing cultures in PDA were transferred to liquid medium cultures (potato dextrose broth—PDB at pH 5.6–6.0; molecular biology grade, Merck, Darmstadt, Germany). To this end, the previously sterilized PDB was aliquoted into several 500 mL Erlenmeyer flasks and inoculated with 2/3 7 mm-diameter discs of each species. Freshly inoculated liquid cultures were incubated at 28 °C and 120 rpm for 8 d in the dark [16]. Afterwards, the mycelium was collected from the liquid medium, stored at −20 °C, and lyophilized for 48 h (−80 °C, 1 bar; Telstar Lyoquest, Terrassa, Spain; Figure S1). Maceration of the freeze-dried powder of each fungal species allowed for more homogeneous incorporation of the mycelia into the soil (see Section 2.3).

2.3. Emergence and Growth Assays of Zea mays

Belgrano variety Zea mays seeds were used in this study (OECD Seed Scheme, France). The assays followed ISO guideline 11269-2 [19] for monospecific tests on the emergence of seeds and the development of higher plants. Five replicates were assembled per treatment, including that of the control. Each replicate was filled with 100 g of artificial OECD soil in a 200 cm³ recipient ([20]). The preference for artificial soil focused on avoiding natural soils where the presence of micropollutants could influence the test results. The artificial soil was prepared by adding and mixing peat (10% blond sphagnum peat with >90% organic matter and bulk density of 140–170 kg/m³; Siro^®, Mira, Portugal), river quartz sand (70% air dried; MaxMat^®, Porto, Portugal), and natural kaolinite clay (20%, Próvida^®, Mem Martins, Portugal), with adjustment of soil pH to 6.0 ± 0.5, by addition of calcium carbonate (≤1%; CaCO₃, chemically pure, Merck, Germany; Figure S1). The soil water-holding capacity (WHC_max) was also determined prior to the assays to estimate the amount of water to be added during the assay to achieve a soil moisture content of up to 45% of its WHC_max. Each replicate contained a single seed.

The bacteria were applied to the seeds by soaking the seeds in a solution of bacteria at a concentration of 1.0 × 10⁸ colony-forming units (CFU)/mL for 20 min, stirring at 100 rpm in sterile Erlenmeyer flasks. In addition, 1 mL of the same bacterial solution was inoculated into the soil of each replicate prior to seed placement. The seeds from the control condition were soaked for 20 min in the same sterile PBS solution, and one mL of the PBS solution was pipetted into the soil (Figure S1). In the case of wood-decaying fungi, dry mycelia were incorporated into dry soil at a proportion of 1 mg/g of soil and mixed (Figure S1).

Every test was conducted under controlled conditions at a temperature of 28 ± 2 °C (day), 22 ± 2 °C (night), and 16L: 8D photoperiod. Every day, the number of emerging seeds was counted in accordance with ISO guidelines [19]. Counting from the emergence of 50% of the control group’s seeds, the assay was carried out for 14 days. At the end of this period, plants were collected, and the aboveground length measured (reported in cm) and weighed (reported as mg fresh weight; Kern, EW1500-2M, resolution of 0.01 g). Afterwards, plants were stored at −20 °C until lyophilization (−80 °C, 1 Bar; Telstar, Lyoquest). Lyophilized plants were reweighed to assess dry weight (reported as mg of dry weight; −20 °C; Kern, EW1500-2M, resolution of 0.01 g) and their chlorophyll content analyzed (see Section 2.5).

Soil physicochemical parameters were also determined at the conclusion of the assays. Soil-deionized water suspensions at a ratio of 1:5 w:v were magnetically stirred for 15 min and left to rest for 60 min after measurements (WTW pH 330i meter; [21]). For electrical conductivity measurements (EC, µS/cm), soil-deionized water suspensions at a ratio of 1:5 w:v were mechanically shaken for 15 min with 50 mL of deionized water and left to rest overnight for the bulk sediment to settle (LF 330/SET meter; [22]).

The experiment that aimed to test the isolated or combined action of genera of soil bacteria and rot fungi was followed in parallel by a similar experiment in which no maize seeds were placed. This parallel experiment served as a positive control and was intended to help interpret the data obtained, with the plants serving as a reference point. This experiment made it possible to verify, under controlled conditions, how each bacterial strain or fungus shaped the physicochemical properties of the soil and its viability over the course of the experiment. Determinations in the experiment without plants included pH and electrical conductivity, phosphatases, and dehydrogenase activities (see Section 2.6).

2.4. Chlorophyll Determination

Photosynthetic pigment determination was performed through whole-pigment extraction to evaluate possible effects on the photosynthetic apparatus or changes in plant fitness, following the protocol outlined by Lichtenthaler [23]. In short, a known weight of previously freeze-dried plant tissue was transferred into a test tube and left to hydrate for 10 min with 100 µL of distilled water. Thereafter, 1 mL of 96% ethanol was added, and the sample was vortexed. The tubes were then wrapped in aluminum foil and incubated overnight at room temperature. The next day, samples were centrifuged (8000 rpm, 8 min), and the absorbance of the extract (supernatant) was read at three wavelengths of 648, 664, and 470 nm (denoted as A648, A664, and A470, respectively). The absorbance values should be in the range of 0.2 and 0.8 [23]. Chlorophylls a and b [Chla or (Chlb), respectively], total Chla + Chlb, ratio of Chla/Chlb, and total of carotenoids (Cx+c) were determined as a function of dry weight (DW) as follows [23]:

$$\mathrm{C}\mathrm{h}\mathrm{l}\mathrm{a}={\displaystyle \frac{13.3\times \mathrm{A}664-5.19\times \mathrm{A}648}{\mathrm{D}\mathrm{W}}}, (\mathrm{m}\mathrm{g}/\mathrm{g}·\mathrm{d}\mathrm{w})$$

$$\mathrm{C}\mathrm{h}\mathrm{l}\mathrm{b}={\displaystyle \frac{27.43\times \mathrm{A}648-8.1\times \mathrm{A}664}{\mathrm{D}\mathrm{W}}}, (\mathrm{m}\mathrm{g}/\mathrm{g}·\mathrm{d}\mathrm{w})$$

$$\mathrm{C}\mathrm{h}\mathrm{l}\mathrm{a}+\mathrm{C}\mathrm{h}\mathrm{l}\mathrm{b}={\displaystyle \frac{5.24\times \mathrm{A}664-22.24\times \mathrm{A}648}{\mathrm{D}\mathrm{W}}}, (\mathrm{m}\mathrm{g}/\mathrm{g}·\mathrm{d}\mathrm{w})$$

$$\mathrm{C}\mathrm{h}\mathrm{l}\mathrm{a}/\mathrm{b}={\displaystyle \frac{\mathrm{C}\mathrm{h}\mathrm{l}\mathrm{a}}{\mathrm{C}\mathrm{h}\mathrm{l}\mathrm{b}}} , (\mathrm{m}\mathrm{g}/\mathrm{g}·\mathrm{d}\mathrm{w})$$

$$\mathrm{C}\mathrm{h}\mathrm{l}\mathrm{x}+\mathrm{c}={\displaystyle \frac{(1000\times \mathrm{A}470-2.13\times \mathrm{C}\mathrm{h}\mathrm{l}\mathrm{a}-97.64\times \mathrm{C}\mathrm{h}\mathrm{l}\mathrm{b})/209}{\mathrm{D}\mathrm{W}}}, (\mathrm{m}\mathrm{g}/\mathrm{g}·\mathrm{d}\mathrm{w})$$

2.5. Enzymatic Determination

Phosphatases (alkaline, Ph = 11, and acid, pH = 6) were determined in accordance with the methodology outlined by Eivazi and Tabatabai [24]. Three replicates of each soil sample were analyzed. A stock solution of 4-nitrophenyl phosphate disodium salt hexahydrate (CAS Number: 4264-83-9, Merck, Darmstadt, Germany) was prepared for each enzyme in universal buffer with pH adjusted according to the optimal enzyme [24]. One gram of each soil sample was combined with 1 mL of the respective stock solution and 125 µL of toluene and vortexed for 10 s. After 60 min of incubation at 37 °C in the dark at 150 rpm, the samples were centrifuged at 10,000× g for 8 min. The p-nitrophenol (pNP) concentration was read at 420 nm in the sample’s supernatant using universal buffer as a blank (Multiskan Microplate Photometer, Thermo Scientific, Waltham, MA, USA).

The dehydrogenase activity was measured by mixing 20 g of soil with 0.2 g of CaCO₃ (reagent grade, Merck, Darmstadt, Germany) [25]. One milliliter of a 3.3% aqueous solution of 2,3,5-triphenyltetrazolium chloride (CAS Number 298-96-4; Merck, Darmstadt, Germany) and distilled water (2.5 mL) were added to each sample. After incubation for 24 h at 37 °C on an orbital shaker set to 150 rpm, the samples were centrifuged for 10 min at 10,000× g. Then, triphenylformanzan (the result of the dehydrogenase activity over the specified substrate) was extracted using methanol (99.8%, 67-56-1, Merck, Darmstadt, Germany; volume used until no red/pink color was observed) and measured at 485 nm (Multiskan Microplate Photometer, Thermo Scientific, Waltham, MA, USA) [25]. Metanol was used as the blank. The concentration of formazan was calculated in mg.

2.6. Data Analysis

All tested parameters were normalized, and a resemblance matrix was generated using the Euclidean distance metric. The Euclidean data matrix was used to perform a main test with a type III sum of squares, subjected to 9999 permutations using PRIMER 7 and PERMANOVA+. Regarding the sole application of rot fungi or rhizobacteria, it was assumed that there were no differences between any of the treatments for any of the tested parameters. Significant differences were considered only when p < 0.05 and were represented as homogenous groups. Regarding the results of combinations, differences were assessed within each bacterial genus and its combination with both rot fungi for each parameter, considering as a null hypothesis no differences between the treatments. Significant differences were considered only when p < 0.05 and were represented as homogenous groups. For enzymatic soil activity, the null hypothesis was that, for each parameter, no significant differences existed between the soil inoculated with and without plants. Significant differences were considered only when p < 0.05 and were identified using an asterisk.

3. Results

3.1. Soil Physico-Chemical Parameters

The soil physicochemical parameters, with and without plants, are presented in Supplementary Table S1. In the plant assay, the highest EC values were recorded for the Enterobacter sp. O8 alone or in combination with the two rot fungi (values ranging between 99.4 and 55.2 μS/cm), followed by the Pseudomonas sp. O81 bacterial strain combined with both rot fungi [values of 59.2 µS/cm for Pleurotus sp. + Pseudomonas sp. O81 and 71.8 μS/cm for Trametes sp. + Pseudomonas sp. O81; Table S1]. Furthermore, treatments with Pleurotus sp. alone and the bacterial strain Enterobacter sp. I2 alone increased the soil EC above 50 µS/cm (Table S1). When EC was evaluated in soil where no plants were grown, the EC parameter was, in general, always higher than that in soil with plants, except for Enterobacter sp. O8 alone or in combination with Pleurotus sp. and in Enterobacter sp. I2 alone (Table S1). The greatest difference between soil with and without plants was observed for the bacterial genera Flavobacterium sp. I57 and Acinetobacter sp. S2, and the combinations Trametes sp. + Flavobacterium sp. I57 and Trametes sp. + Enterobacter sp. I2 (>2.3-fold difference) (Table S1).

In relation to pH, in soil with plants, the parameter fluctuated by a single unit, with values ranging between 5.48 and 6.48 (Table S1). In the parallel test without plants, the soil pH was slightly higher but also fluctuated around one unit (5.87 ≤ pH ≤ 6.80; Table S1). When comparing soil with and without plants, increments of approximately one unit should be highlighted for the bacterial strain Enterobacter sp. O8 and a combination of Trametes sp. + Pseudomonas sp. O81 (Table S1).

3.2. Germination Rates

The germination percentages in relation to the control conditions are shown in Figure 1. Comparing the fungi first in relation to the control and comparing both fungi, it is worth highlighting the differential action of the two wood-decaying fungi: Trametes sp. showed germination rates 40% higher than the control, reaching 100% after 3 days, although they were not significantly different from the control. In soil supplemented with Pleurotus sp., germination was inhibited by 40%, though significant differences were not found in relation to the control group (Figure 1). Regarding maize germination in soil supplemented with bacteria only, it was found that all bacterial genera induced higher germination rates than control conditions at the end of 72 h (equal to or higher than 20%), except for Enterobacter sp. O8, where 20% suppression rates were registered compared with the control (Figure 1).

In the combinations, over the three days of germination recording, it was found that of the 12 combinations, eight registered higher germination rates than the control (Figure 1). Four combinations of Pleurotus sp. [Flavobacterium sp. I57, Enterobacter sp. I2, Acinetobacter sp. S2, and Pseudomonas sp. O81] and four with Trametes sp. [Flavobacterium sp. I57, Acinetobacter sp. S2, and Pseudomonas sp. N26; Figure 1]. Of these combinations compared to the isolated action of fungi, it is worth highlighting the higher yield of Pleurotus sp. when in combination with all bacterial genera, and in relation to bacteria, it is worth highlighting the suppressive effect of Pseudomonas sp. N26 and Enterobacter sp. I2 when combined with Pleurotus sp. and Trametes sp., respectively. However, there was an increase in the action of Enterobacter sp. O8 and Acinetobacter sp. S2 when combined with both fungi. Despite this, no significant differences from the control were found (Figure 1).

3.3. Zea mays Morphometric Parameters

To obtain a more detailed view of the benefits that bacteria or fungi alone or their combinations can have on maize growth, the data were divided as follows: the effects of bacteria alone on plant growth parameters (Figure 2), the effect of fungi alone on plant growth parameters (Figure 3), and the results of the bacteria + fungi combination on these same parameters (Figure 4).

Plants grown in soil supplemented with Pseudomonas sp. O81 were found to be significantly taller than plants from the other groups (p < 0.05; Figure 2A), except for the control group and soil supplemented with Acinetobacter sp. S2, where plants grew significantly less (p < 0.05; Figure 2A). Regarding the development of the fourth leaf, only Enterobacter sp. I2 induced a significantly higher development stage compared to the control (p < 0.05; Figure 2B). In relation to the fresh mass of maize roots, the bacterial genera Flavobacterium sp. I57 and Enterobacter sp. I2 induced the lowest and highest significant plant development, respectively (p < 0.05; Figure 2C), whereas in the soil amended with these two genera [Flavobacterium sp. I57 and Enterobacter sp. I2], the dry weight of maize roots was significantly lower in these two groups than in the control group (p < 0.05; Figure 2D). In relation to the weight of the aerial parts, plants grown in soil supplemented with Flavobacterium sp. I57 and Enterobacter sp. I2 had significantly higher fresh and dry weights of aerial parts, in addition to Acinetobacter sp. S2 and Pseudomonas sp. O81 only in the fresh mass of the aerial part (p < 0.05; Figure 2E,F).

Regarding Z. mays growth in soil enriched with rot fungal mycelia, it was not possible to retrieve data regarding the upper parts of the plant when the soil was amended with Pleurotus sp., as insufficient plants emerged (Figure 3A,B,E,F). In plants exposed to Trametes sp., the dry weight of plant roots was significantly lower under Trametes sp. exposure compared to the control (p < 0.05; Figure 3D), whereas plant shoot fresh weight was significantly higher than that of the control. (p < 0.05; Figure 3E). No other significant differences were found (Figure 3).

Regarding maize growth parameters measured after 14 days of exposure to combinations of six bacterial genera with two rot fungi, the results are shown in Figure 4. Statistical differences were found mainly for shoot length, shoot fresh weight, and shoot dry weight. Combinations of Pleurotus sp. with Flavobacterium sp. I57, Enterobacter sp. I2, and Pseudomonas sp. N26, and combinations of Trametes sp. with Enterobacter sp. O8, Acinetobacter sp. S2, Pseudomonas sp. O81, and Pseudomonas sp. N26 significantly increased shoot length when compared to the control condition (Figure 4A). Concerning the developmental stage (Figure 4B), the combinations that significantly promoted the development of the fourth leaf were Pleurotus sp. with Enterobacter sp. O8 and Pseudomonas sp. O81, and Trametes sp. with Acinetobacter sp. S2 and Pseudomonas sp. O81. Regarding root parameters, just three combinations showed significant differences in root dry weight (Figure 4D), Pleurotus sp. with Pseudomonas sp. O81 and Pseudomonas sp. N27, and Trametes sp. with the strain Acinetobacter sp. S2. Regarding the shoot fresh and dry weights (Figure 4E,F), except for Pleurotus sp. with Acinetobacter sp. S2, Pseudomonas sp. O81, and Trametes sp. with Flavobacterium sp. I57, all combinations significantly increased the weight of the plants when compared to the control.

3.4. Zea mays Photosynthetic Pigments

Determination of photosynthetic pigments (

Table 1. Zea mays photosynthetic pigment determination after a 14-day germination and seedling growth assay in standard OECD soil amended with mycelia of two different wood-decaying fungi (Trametes sp. and Pleurotus sp.) in combinations with six different bacterial strains [Flavobacterium sp. I57, Enterobacter sp. I2, Enterobacter sp. O8, Acinetobacter sp. S2, Pseudomonas sp. O81, Pseudomonas sp. N26]. The data are presented as the mean ± standard deviation (n = 5). Shared letters (a,b) indicate homogenous groups within the same parameter and the bacterial strain versus combination with fungi. N/A, not assessed because no plants emerged.

Photosynthetic Pigments (Combinations)
Conditions	Chlorophyll a (mg g−1 dw)	Chlorophyll b (mg g−1 dw)	Chlorophyll a+b (mg g−1 dw)	Chlorophyll x+c (mg g−1 dw)	Ratio a/b
Control	9.54 ± 1.89 a	2.48 ± 0.55 a	12.0 ± 2.42 a	2.55 ± 0.57 a	3.87 ± 0.29 a
Pleurotus sp.	N/A	N/A	N/A	N/A	N/A
Trametes sp.	6.87 ± 2.64 a	1.96 ± 0.57 a	8.83 ± 3.17 a	2.16 ± 0.54 a	3.48 ± 0.48 a
Flavobacterium sp. I57	10.6 ± 1.47 a	2.57 ± 0.36 a	13.1 ± 1.83 a	2.88 ± 0.17 a	4.10 ± 0.09 a
Pleurotus sp. + Flavobacterium sp. I57	6.46 ± 2.50 b	1.94 ± 0.42 a	8.41 ± 2.82 a	1.68 ± 0.65 b	3.29 ± 0.82 a,b
Trametes sp. + Flavobacterium sp. I57	8.26 ± 1.43 a,b	2.20 ± 0.40 a	10.5 ± 1.81 a	2.24 ± 0.37 b	3.76 ± 0.23 b
Enterobacter sp. I2	7.76 ± 1.97 a	1.87 ± 0.56 a	9.63 ± 2.52 a	2.16 ± 0.40 a	4.21 ± 0.24 a
Pleurotus sp. + Enterobacter sp. I2	8.29 ± 3.27 a	2.21 ± 0.80 a	10.5 ± 4.07 a	2.20 ± 0.84 a	3.71 ± 0.12 b
Trametes sp. + Enterobacter sp. I2	7.70 ± 0.18 a	2.13 ± 0.05 a	9.83 ± 0.13 a	1.96 ± 0.21 a	3.61 ± 0.17 a,b
Enterobacter sp. O8	10.4 ± 1.24 a	2.71 ± 0.30 a	13.2 ± 1.54 a	2.80 ± 0.50 a	3.96 ± 0.03 a
Trametes sp. + Enterobacter sp. O8	5.70 ± 0.09 b	1.45 ± 0.08 b	7.15 ± 0.08 b	1.79 ± 0.10 a	3.94 ± 0.27 a
Pleurotus sp. + Enterobacter sp. O8	9.70 ± 1.15 a	2.62 ± 0.09 a	12.3 ± 1.21 a	2.56 ± 0.37 a	3.69 ± 0.37 a
Acinetobacter sp. S2	9.48 ± 1.62 a	2.34 ± 0.44 a	11.8 ± 2.05 a	2.35 ± 0.31 a	4.06 ± 0.18 a
Pleurotus sp. + Acinetobacter sp. S2	8.12 ± 2.04 a	2.20 ± 0.53 a	10.3 ± 2.52 a	2.00 ± 0.34 a	3.68 ± 0.47 a
Trametes sp. + Acinetobacter sp. S2	8.68 ± 2.81 a	2.12 ± 0.61 a	10.8 ± 3.41 a	2.41 ± 0.46 a	4.03 ± 0.19 a
Pseudomonas sp. O81	9.32 ± 1.86 a	2.35 ± 0.48 a	11.7 ± 2.34 a	2.46 ± 0.44 a	3.96 ± 0.08 a
Pleurotus sp. + Pseudomonas sp. O81	8.35 ± 2.23 a	2.09 ± 0.49 a	10.4 ± 2.72 a	2.16 ± 0.55 a	3.98 ± 0.18 a
Trametes sp. + Pseudomonas sp. O81	9.52 ± 2.78 a	2.53 ± 0.40 a	12.0 ± 3.16 a	2.34 ± 0.53 a	3.69 ± 0.67 a
Pseudomonas sp. N26	10.2 ± 2.23 a	2.67 ± 0.32 a	12.9 ± 2.54 a	2.65 ± 0.54 a	3.77 ± 0.46 a
Pleurotus sp. + Pseudomonas sp. N26	8.82 ± 2.12 a	2.39 ± 0.58 a	11.2 ± 2.68 a	2.10 ± 0.47 a	3.70 ± 0.24 a
Trametes sp. + Pseudomonas sp. N26	9.34 ± 2.60 a	2.47 ± 0.59 a	11.8 ± 3.19 a	2.46 ± 0.62 a	3.75 ± 0.28 a

) showed that the concentrations of Chl a, carotenoids, and chl a/b ratio decreased significantly in Pleurotus sp. + Flavobacterium sp. I57 combinations compared to the other combination of Flavobacterium sp. I57 with Trametes sp. or Flavobacterium sp. I57 alone (p < 0.05; Table 1). The combination Enterobacter sp. O8 + Pleurotus sp. also resulted in a significant decrease in all photosynthetic pigments when compared to the other combinations and the treatment with Enterobacter sp. O8 only, except for carotenoids and the chlorophyll a/b ratio (p < 0.05; Table 1).

3.5. Soil Enzymatic Activity

The soil enzymatic activities of phosphatases (acid and alkaline) and dehydrogenase are summarized in

Table 2. Soil phosphatase activity (µg p NP/soil g/hour) in OECD soil amended with mycelia of two different wood-decaying fungi (Trametes sp. and Pleurotus sp.) in combinations with six different bacterial strains [Flavobacterium sp. I57, Enterobacter sp. I2, Enterobacter sp. O8, Acinetobacter sp. S2, Pseudomonas sp. O81, Pseudomonas sp. N26]. The data are presented as the mean ± standard deviation (n = 5). Shared letters (a,b for acid phosphatase or A,B for alkaline phosphatase) indicate homogenous groups between time periods and the bacterial strain versus combination with fungi.

Phosphatase (µg p NP/Soil g/hour)
Conditions	Day 0		Day 7		Day 14
Phosphatase	Acid	Alkaline	Acid	Alkaline	Acid	Alkaline
Control	98.88 ± 11.1 A	31.91 ± 9.04 a	67.85 ± 4.85 A	20.78 ± 5.35 a	120.7 ± 57.8 A	25.06 ± 4.48 a
Pleurotus sp.	78.75 ± 5.57 B	25.45 ± 4.00 a	82.04 ± 18.6 A,B	29.15 ± 3.24 b	148.4 ± 48.1 A	27.86 ± 6.84 a
Trametes sp.	98.84 ± 12.3 A	25.98 ± 3.03 a	103.7 ± 20.9 A	22.88 ± 0.62 a	137.7 ± 18.7 A	21.93 ± 6.32 a
Flavobacterium sp. I57	94.08 ± 11.4 A	25.65 ± 1.58 a	90.23 ± 12.3 A	20.97 ± 4.19 a	127.1 ± 12.3 A	28.15 ± 1.44 a
Pleurotus sp. + Flavobacterium sp. I57	113.2 ± 38.7 A	26.43 ± 2.90 a	131.1 ± 47.6 A	24.55 ± 3.58 a	111.9 ± 54.7 A	31.32 ± 8.83 a
Trametes sp. + Flavobacterium sp. I57	107.3 ± 9.00 A	29.55 ± 4.51 a	118.7 ± 34.9 A	36.89 ± 20.3 a	94.46 ± 41.4 A	28.58 ± 1.83 b
Enterobacter sp. I2	94.40 ± 4.35 A	24.79 ± 2.66 a	66.28 ± 11.0 A	20.37 ± 2.13 a	122.6 ± 23.5 A	26.22 ± 5.34 a
Pleurotus sp. + Enterobacter sp. I2	151.9 ± 37.3 B	30.81 ± 7.94 a	142.1 ± 31.9 B	24.51 ± 1.57 b	168.9 ± 34.9 A	48.44 ± 12.9 b
Trametes sp. + Enterobacter sp. I2	133.8 ± 20.6 B	21.36 ± 4.59 a	121.4 ± 11.7 B	26.84 ± 1.24 b	52.78 ± 25.7 B	21.83 ± 2.74 a
Enterobacter sp. O8	90.22 ± 6.31 A	26.19 ± 1.90 a	84.13 ± 3.70 A	23.79 ± 3.82 a	148.1 ± 44.2 A	22.29 ± 4.36 a
Pleurotus sp. + Enterobacter sp. O8	94.68 ± 6.46 A	33.93 ± 6.17 a	132.8 ± 34.9 A	30.15 ± 2.40 a	165.0 ± 84.3 A	38.66 ± 14.1 a
Trametes sp. + Enterobacter sp. O8	139.8 ± 19.9 B	53.63 ± 41.6 a	80.79 ± 13.6 A	26.60 ± 3.61 a	100.3 ± 21.4 A	32.66 ± 8.74 a
Acinetobacter sp. S2	97.79 ± 11.1 A	27.08 ± 3.89 a	86.78 ± 17.3 A	21.70 ± 2.33 a,b	188.1 ± 90.1 A	29.03 ± 3.65 a
Pleurotus sp. + Acinetobacter sp. S2	110.0 ± 22.9 A	23.41 ± 2.03 a	87.71 ± 15.6 A,B	35.99 ± 8.61 b	130.4 ± 28.5 A	34.28 ± 4.68 b
Trametes sp. + Acinetobacter sp. S2	120.1 ± 34.8 A	23.86 ± 0.82 a	132.9 ± 23.7 B	34.63 ± 1.82 b	93.27 ± 19.3 A	36.35 ± 8.84 b
Pseudomonas sp. O81	99.36 ± 9.08 A	38.22 ± 19.9 a	91.54 ± 6.84 A	23.93 ± 3.73 a	199.1 ± 53.1 A,B	28.21 ± 5.99 a
Pleurotus sp. + Pseudomonas sp. O81	96.32 ± 10.5 A	24.89 ± 3.64 a	203.8 ± 139 A	30.66 ± 7.91 a,b	60.04 ± 13.4 A	23.84 ± 3.04 b
Trametes sp. + Pseudomonas sp. O81	137.5 ± 41.2 A	20.42 ± 4.58 a	169.9 ± 107 A	33.17 ± 1.65 a	124.2 ± 26.2 B	41.70 ± 22.0 a
Pseudomonas sp. N26	106.7 ± 14.9 A	23.12 ± 2.09 a	87.54 ± 7.93 A	21.45 ± 2.83 a	212.4 ± 55.0 A	33.98 ± 8.05 a
Pleurotus sp. + Pseudomonas sp. N26	137.6 ± 19.1 A	25.41 ± 2.00 a,b	92.13 ± 15.4 A,B	29.22 ± 2.85 b	110.3 ± 44.9 A	23.68 ± 1.01 b
Trametes sp. + Pseudomonas sp. N26	135.8 ± 16.9 A	30.11 ± 3.70 a	120.8 ± 8.20 A	24.94 ± 2.07 a,b	109.5 ± 51.5 A	19.22 ± 0.74 a

and

Table 3. Soil dehydrogenase activity (ng TPF/g of soil/hour) in OECD soil amended with mycelia of two different decaying fungi (Trametes sp. and Pleurotus sp.) in combination with six different bacterial strains [Flavobacterium sp. I57, Enterobacter sp. I2, Enterobacter sp. O8, Acinetobacter sp., S2, Pseudomonas sp. O81, Pseudomonas sp. N26]. The data are presented as the mean ± standard deviation (n = 5). Shared letters (a–c) indicate homogenous groups between time periods and the bacterial strain versus combination with fungi.

Dehydrogenase Activity (ng TPF/g Soil/hour)
Conditions	Day 0	Day 7	Day 14
Control	4.32 ± 1.11 a	3.89 ± 0.55 a	9.11 ± 4.53 b
Pleurotus sp.	6.13 ± 1.45 a	3.75 ± 0.35 b	11.8 ± 1.26 c
Trametes sp.	5.24 ± 1.67 a	4.10 ± 0.59 a	13.4 ± 6.48 b
Flavobacterium sp. I57	3.63 ± 0.14 a	3.68 ± 0.35 a	3.94 ± 0.43 a
Pleurotus sp. + Flavobacterium sp. I57	4.17 ± 0.28 a	4.41 ± 0.58 a	8.81 ± 1.19 b
Trametes sp. + Flavobacterium sp. I57	8.51 ± 0.87 a	4.66 ± 0.35 b	9.79 ± 4.91 a,b
Enterobacter sp. I2	3.95 ± 0.30 a	4.41 ± 0.82 a	6.03 ± 0.99 b
Pleurotus sp. + Enterobacter sp. I2	7.00 ± 2.69 a	9.57 ± 4.53 a	11.7 ± 4.13 a
Trametes sp. + Enterobacter sp. I2	4.66 ± 0.60 a	4.74 ± 0.46 a	7.00 ± 2.31 a
Enterobacter sp. O8	3.82 ± 0.31 a	3.82 ± 0.05 a	6.10 ± 1.04 b
Pleurotus sp. + Enterobacter sp. O8	4.58 ± 0.60 a	6.46 ± 1.54 a	16.1 ± 7.82 a
Trametes sp. + Enterobacter sp. O8	9.28 ± 5.99 a	7.49 ± 2.31 a	10.0 ± 2.82 a
Acinetobacter sp. S2	3.75 ± 0.30 a	3.90 ± 0.23 a	10.2 ± 0.94 b
Pleurotus sp. + Acinetobacter sp. S2	9.94 ± 3.54 a,b	4.59 ± 0.21 a	7.52 ± 0.59 b
Trametes sp. + Acinetobacter sp. S2	6.19 ± 1.31 a	4.62 ± 0.63 a	11.4 ± 3.69 a
Pseudomonas sp. O81	4.05 ± 0.67 a	3.46 ± 0.51 a	11.6 ± 4.00 b
Pleurotus sp. + Pseudomonas sp. O81	6.87 ± 0.93 a	5.80 ± 1.15 a	14.5 ± 9.56 a
Trametes sp. + Pseudomonas sp. O81	4.01 ± 0.48 a	4.80 ± 1.35 a	13.6 ± 7.13 a
Pseudomonas sp. N26	4.17 ± 0.75 a	3.52 ± 0.22 a	12.3 ± 5.33 b
Pleurotus sp. + Pseudomonas sp. N26	5.59 ± 0.18 a	5.80 ± 1.15 b	8.23 ± 1.23 c
Trametes sp. + Pseudomonas sp. N26	3.77 ± 0.47 a	4.44 ± 0.41 a	10.8 ± 1.89 b

, respectively.

The acid phosphatase activity pattern in soils supplemented with fungi and mycelia solely was found to increase over time (Table 2). However, the same enzyme activity was found to decrease in all bacterial genera tested from day 0 to day 7, and then increase from day 7 to day 14 (Table 2). Regarding the activity of soil acid phosphatase in the bacteria + fungi combinations, it was observed that in the combination of Enterobacter sp. I2 with both rot fungi, acid phosphatase activity on days 0 and 7 was significantly higher than that in the soil amended only with Enterobacter sp. I2; however, the differences tended to disappear over time, remaining solely in the combination Trametes sp. + Enterobacter sp. I2 (Table 2). In addition, it was observed that in the combination of Trametes sp. + Enterobacter sp. O8 on day 0, whereas in the combination of Trametes sp. + Acinetobacter sp. S2 from day 0 to day 7, acid phosphatase activity was significantly higher than that of Enterobacter sp. O8 or Pleurotus sp. + Enterobacter sp. O8 and Acinetobacter sp. S2 or Pleurotus sp. + Acinetobacter sp. S2 but decreased to initial levels until the end of the experiment (Table 2). In the Trametes sp. + Pseudomonas sp. N26 combination, acid phosphatase activity decreased significantly from day 0 to day 7, although no differences were found at the end of the experiment (Table 2).

The activity of alkaline phosphatase varied slightly when soil was supplemented with rot fungi, increasing in the case of Pleurotus sp. from day 0 to day 7 and then decreasing until day 14, while in Trametes sp. Soil, alkaline phosphatase activity decreased over time (Table 2). Under the conditions of bacterial strain application alone, alkaline phosphatase activity showed the same response pattern in all genera, except for Enterobacter sp. O8 and Pseudomonas sp. O81: decreasing from sampling day 0 to day 7, while increasing up to day 14 to similar initial levels. In Enterobacter sp. O8 and Pseudomonas sp. O81, alkaline phosphatase activity decreased over time (Table 2). In the combinations of Pleurotus sp. + Enterobacter sp. I2, Trametes sp. + Enterobacter sp. I2, and Pleurotus sp. + Pseudomonas sp. N26, alkaline phosphatase activity increased until day 7 and remained at least when combined with Pleurotus sp. while decreasing in Pleurotus sp. + Pseudomonas sp. N26 (Table 2). In Acinetobacter sp. S2 combined with both rot fungi, the activity of this enzyme was significantly higher than in the treatment of the bacterial strain alone over time (Table 2).

The activities of acid and alkaline phosphatases in soil with and without plants are summarized in Supplementary Figures S2 and S3, respectively. Only a few differences were found between the two conditions (plant versus no plant), namely in the control conditions. The acid phosphatase was significantly lower in vessels containing plants in the control, Pseudomonas sp. N26 alone, and the combination Pleurotus sp. + Enterobacter sp. O8 (p < 0.05; Figure S2). The alkaline phosphates were significantly higher in vessels with plants under control conditions, Enterobacter sp. I2 alone, and the combinations of Trametes sp. + Flavobacterium sp. I57 and Trametes sp. + Enterobacter sp. I2 (p < 0.05; Figure S3).

The dehydrogenase activity during the plant assay was found to significantly increase overall among all treatments, including the control, except for the bacterial strain Flavobacterium sp. I57 alone and in combination with Trametes sp. and the bacterial strains Enterobacter sp. I2, Enterobacter sp. O8, and Pseudomonas sp. O81 when combined with both rot fungi (Table 3). Nevertheless, the dehydrogenase activity showed the highest differences between vessels with and without plants (Figure S4). In vessels where soil was amended solely with rot fungi, there was a differential result between the fungi, with Pleurotus sp. presenting a significantly higher activity of dehydrogenase in soils with plants, whilst in Trametes sp., the opposite trend was observed (p < 0.05; Figure S4). In all bacterial strains, dehydrogenase activity was found to be significantly higher in the absence of the plants compared to the same treatment with plants (p < 0.05; Figure S2). Combinations of the bacterial genera with Pleurotus sp. [except for Pseudomonas sp. O81 and Pseudomonas sp. N26] showed significantly higher dehydrogenase activity in vessels without plants (p < 0.05; Figure S4). No changes were observed in combinations with Trametes sp. and the control vessels (p > 0.05; Figure S4).

4. Discussion

The development of novel bioformulations that harness the potential of a wide range of microorganisms is of utmost importance for advancing agricultural sustainability and aligning with European directives for a greener future [1]. Given the growing need for solutions that promote sustainable agriculture and reduce environmental harm, it is crucial to explore innovative approaches to enhance crop productivity while addressing the pressing issues of climate change and resource depletion. This study aimed to trigger research in this field by presenting the results of combinations of two major groups of soil microbiota (fungi and bacteria) and their potential enhancing effects on crops such as maize.

Comparing the two isolated groups, the bacterial genera tested alone seemed to have an overall positive effect on the growth of maize, whereas the fungi had different results. There is extensive evidence of the positive impact of plant growth-promoting bacteria on plant fitness, and it is therefore not surprising that in this study, its application resulted in improvements in germination, with a tendency (although not significant) towards a stage of the plant’s superior development (verified by the size of the fourth leaf) and in the weights of the aerial parts of the plants. The reported effects of plant-growth bacteria application span from increased photosynthetic apparatus efficiency to higher nitrogen-phosphorous-potassium use efficiency, increased seed yield, overall plant and/or fruit growth, and a higher nutritional status of the end-product [26,27,28,29,30]. For instance, Mondani et al. [27] demonstrated that the application of Bacillus licheniformis in soybean crops (Glycina max) increased grain yield and protein content by approximately 23% and 19%, respectively [27], whereas Katsenios et al. [29] found that the mean fruit weight of tomato plants increased by up to 30% after the application of Bacillus subtilis, Bacillus amyloliquefaciens, Priestia megaterium, and Bacillus licheniformis [29].

As for the exclusive application of fungi, their differential action on maize germination dictated the results obtained later for the other physiological parameters tested. While Trametes sp. induced germination rates 40% above the control, Pleurotus sp. suppressed germination rates by the same amount. This result agrees with other findings in the literature for the same species of rot fungi. Borges et al. [16], when studying the application of T. versicolor and P. sajor caju exudates to improve the functionality of a salinized soil (26 mS/cm), found out that the application of T. versicolor exudate led to a 30% increase in Lens culinaris germination rates compared to seed germination rates in soil exposed to the same level of conductivity but without the exudate; while the use of Pleurotus sp. exudates in the same study did not result in any improvement in the germination [16]. This result can be explained by the use of dry mycelia, which may foster the removal of water from the vicinity for growth and expansion of the fungal hyphal network. Fungi are highly hygroscopic, and once in favorable conditions, they may easily and quickly absorb water molecules [31]. However, the water withdrawal rate and the need for rot fungi may be quite dissimilar, which is species-specific [32,33,34]. For example, in a study where the mycelium of another Pleurotus sp. was studied and adjusted for use as a potential ecological substitute for expanded polystyrene, it was discovered that this fungus, just in 24 h, considerably increased its water content by more than 114%, a process that was related to the high porosity of the fungal network of this fungus genus [32]. In turn, the structure of Trametes sp. has been reported to be denser and more compact, with a lower affinity for water [34]. Thus, this result can be supported by the competition for water between Pleurotus sp. and maize seeds. Water is essential for seed imbibition, especially in grain seeds, where approximately 35% to 45% of their relative weight at germination corresponds to water [35]. Therefore, the ability of Pleurotus sp. to act as a sponge could have hindered maize germination, while the greater water availability when the soil was supplemented with Trametes sp. may be well related to the higher root fresh weight recorded in the present study.

Notwithstanding the results obtained with Pleurotus sp. alone, several improvements must be highlighted when this species is combined with rhizobacteria. It is important to decipher these combination results because, under natural conditions, very intricate microbial complexity is most likely to exist, and therefore the combination results demonstrate once again the potential of Pleurotus sp. to be explored in more sustainable agricultural practices. For example, when combined with all bacterial genera, germination rates matched those of the control (in two of the six combinations), whereas the other four combinations exceeded control germination rates. Furthermore, Pleurotus sp., when combined with Flavobacterium sp. I57 or Pseudomonas sp. O81, significantly increased shoot length and plant development stage, which was not observed for combinations with Trametes sp. or just for the bacterial genera. However, it must be noted that these combinations with Pleurotus sp. caused significant decreases in the concentration of photosynthetic pigments. This could be related to the potential osmotic stress caused by Pleurotus sp., which may lead to stomatal closure in the plant and a subsequent reduction in carbon dioxide intake for photosynthesis. Notwithstanding, the presence of both fungi appeared to stabilize the activity of both acid and basic phosphatases in the soil, which appeared to fluctuate over time only in the case of the bacterial genera. Here, the presence of fungi appeared to have a greater effect on soil phosphatase activity than did the presence or absence of plants. Furthermore, the combined application of microbiota seemed to favor the soil pH at slightly acidic levels, which are described as being ideal for maize growth as they may facilitate the absorption of phosphorus from the surroundings [36], trigger calcium phosphate solubilization [37] or influence the bioavailability of other growth-decisive elements such as zinc [38].

Despite this, dehydrogenase activity (in the presence of the plant) appeared to be very high in Pleurotus sp. alone as well as in Trametes sp. combined with bacteria. In contrast, dehydrogenase activity remained very low and stable in Trametes sp. alone, in bacterial strains alone, or in combinations of bacteria with Pleurotus sp. The general enzymatic activity of the soil in the combinations may be related to the different wood decomposition environments of each fungus [39]. These wood-decaying fungi have different substrate selectivities, with Pleurotus sp. preferring soft wood and being more versatile, while Trametes sp. prefers a more recalcitrant substrate. Therefore, the composition and quality of the secretome are different; hence, each fungus will certainly create a distinct form of interaction with the bacterial strain it is combined with [40]. For example, when Pleurotus sp. was combined with rhizobacteria without plants, dehydrogenase activity was very high. In this case, it is possible that Pleurotus sp. may have turned the action of the bacteria toward its own growth and activity. For example, an investigation of the bacterial community associated with the growth stages of another Pleurotus species (P. eryngii) revealed that the fungus took advantage of the plant growth-promoting and biocontrol capabilities of Bacillus cereus to accelerate growth [41]. However, combinations of Pleurotus sp. and bacteria in the presence of plants may have been overwhelmed by the needs of the plants during their own growth, which led to the depletion of nutrients in the soils that would otherwise have been used by the fungi. It should be noted that the lack of understanding of this evolution over time does not allow us to understand whether the activity of the plant was always lower or decreased with an increase in the plant’s growth time.

In contrast, the general soil enzyme activity in Trametes sp. combined with bacteria was very high, both in the presence and absence of plants. It has been previously demonstrated that the growth and development of Trametes sp. can also benefit from the presence of rhizobacteria, as shown in the study by Borràs et al. [42]. The biomass of Trametes sp. in soil (using ergosterol as a proxy measure of biomass) was found to be the highest after 5 and 10 weeks of incubation in sterile soils amended with a bacterial inoculum composed of Pseudomonas aeruginosa and Rhodococcus erythropolis (10⁸ CFU/mL) compared to the other two treatments in the absence of inoculum (non-sterile soil and sterile soil; [42]). The differences with and without plants in this study, for both fungi combined with bacteria, may also be related to the ecological role of each fungus, with Pleurotus sp. being an early-stage colonist and Trametes sp. being an early- to mid-stage colonist (potential time delay effect; [43,44]). The assays carried out in this study may be considered short-term if they are paralleled with a field case study. Some results may be masked by the period that could be expressed in other endpoints, as shown by previous studies that ended at grain production (e.g., [27]).

Looking at the bigger picture, it is possible to show that some combinations (such as Trametes sp. + Enterobacter sp. O8) showed significant improvements. This genus (Enterobacter sp.) is well documented for its plant growth-promoting abilities and bioremediation potential (reviewed by [45]). The interactive mechanisms between white rot fungi and Enterobacter sp. have been highlighted elsewhere, promising not only to revolutionize agricultural practices but also to be promoted as agents of remediation [46,47]. Despite the pressures imposed by current challenges in agriculture, the high abundance of bacterial genera and fungal species, the multiplicity of combinations, and the time needed to invest in their study, the identification and optimization of bioformulations represent a critical step towards achieving a greener and more sustainable future for agriculture. It should also be borne in mind that only a few of the many other combinations that can be made were tested here, and the selection was based on some characteristics of each group (e.g., siderophores, indole acetic acid production, or tolerance to osmotic stress [14,15,16]). There are several factors (e.g., environmental) that could limit the applicability of these consortia in the field, and therefore, their use must be carried out in an integrated management practice, that is, combined with other sustainable agricultural practices designed to improve overall soil health and plant productivity, as argued elsewhere [48].

Resorting to this type of sustainable, eco-friendly measure may bring several benefits in addition to increasing soil productivity. Increasing or stimulating soil microbiota can help contribute to the biogeochemical cycle by promoting carbon sequestration in soils or restructuring degraded soils, indirectly related to the human component [49]. Therefore, the development of new bioformulations that harness the potential of a wide range of microorganisms is extremely important for advancing agricultural sustainability. This study aligns perfectly with European sustainability lighthouses to lead the transition towards healthy soils by 2030, further ensure food security, and catalyze a transition to a more sustainable and resilient agricultural system [1].

5. Conclusions

The work presented here is innovative in that it proposes a consortium between rhizobacteria and rot fungi, assuming that each part may address the weaknesses of the other group. Our results provide evidence that rhizobacteria, in combination with wood-decaying fungi, may be able to induce the developmental stage and weight of plants but also shape soil properties and activity in a positive manner. Therefore, studying the combination of wood-decomposing fungi and rhizobacteria could be a promising approach to achieving greater agricultural production while also promoting agricultural practices that help maintain soil health and promote ecosystem services. The engineering of microbial consortia with the potential to increase agricultural production is widely recognized, and these results could be extremely important in changing the way agriculture is thought of today. For example, if well established, the relationship between rhizobacteria and wood-decaying fungi may encourage the optimization of less-used agricultural practices such as direct sowing, which is a method that minimally disturbs the soil and its surroundings, preserving its structure and integrity. This is fundamental for the preservation of organic matter (to prevent erosion) and the future resilience of soils to pests and environmental changes.