Next Article in Journal
Relationship between Biomass Burning Emissions and Deforestation in Amazonia over the Last Two Decades
Previous Article in Journal
Predicting the Potential Habitat of Three Endangered Species of Carpinus Genus under Climate Change and Human Activity
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Forests
Volume 12
Issue 9
10.3390/f12091218
forests-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Impact of Soil-Applied Microbial Inoculant and Fertilizer on Fungal and Bacterial Communities in the Rhizosphere of Robinia sp. and Populus sp. Plantations

by
Zoltán Mayer
1,
Andrea Gógán Csorbainé
2,
Ákos Juhász
1,
Attila Ombódi
2,
Antal Pápai
3,
Boglárka Kisgyörgy Némethné
3 and
Katalin Posta
1,*
1
Department of Microbiology and Applied Biotechnology, Institute of Genetics and Biotechnology, Hungarian University of Agriculture and Life Sciences, Páter Károly Street 1, 2100 Gödöllő, Hungary
2
Institute of Horticulture, Hungarian University of Agriculture and Life Sciences, Páter Károly Street 1, 2100 Gödöllő, Hungary
3
National Food Chain Safety Office, Agricultural Genetic Resources Directorate, Forestry Reproductive Material Department, Kis Rókus Street 15/A, I.19, 1024 Budapest, Hungary
*
Author to whom correspondence should be addressed.
Forests 2021, 12(9), 1218; https://doi.org/10.3390/f12091218
Submission received: 12 August 2021 / Revised: 1 September 2021 / Accepted: 5 September 2021 / Published: 7 September 2021
(This article belongs to the Section Genetics and Molecular Biology)
Browse Figures
Review Reports Versions Notes

Abstract

:
The impact of chemical fertilization on rhizosphere soil communities is a growing concern due to the changes they cause on microbes in soil ecosystems. The present study aims to compare mycorrhizal inoculation and fertilizer applications on bacterial and fungal communities in rhizosphere soil of intensively cultivated Robinia pseudoacacia and Populus × euramericana plantations using the Illumina Miseq sequencing platform. Our results revealed that the different host plants and applied treatments did not significantly affect soil bacterial diversity, but interfered with native rhizosphere bacterial communities in plantation sites. In contrast, host plants and inorganic fertilizer had a strong effect at the family and genus level on the composition of soil fungal communities. In conclusion, our findings suggest that the structure and composition of the fungal community are more sensitive to the nutrient sources in soil than bacteria.

1. Introduction

Black locust (Robinia pseudoacacia L.) is widely distributed through arid and semiarid regions of North America, Europe and Asia; it is used for high-quality firewood and timber production around the world [1]. In 2019, it was the most used tree species on the continent and covered an area of more than 454,531 hectares from the 1,867,557 hectares of forest surface in Hungary, this represents 54,676 thousand m3 of living timber. Black locust is present in 24.33% of the domestic forest area, but only represents 13.89% of living timber biomass [2]. It was declared as Hungaricum in 2014. The main advantage of black locust plantations is their fast and intensive growth. Nevertheless, it produces different growth intensities in different nutrient-supplied soils. As a result of cultivation on less favourable, i.e., moderately good quality or eroded soils it is only able to produce firewood quality. The species can provide high-quality timber on nutrient-rich and well-aerated soils [3]. Due to these characteristics, it is the most important tree species cultivated in Hungary. With its outstandingly intensive pace of development, the black locust reaches its peak height in 5 years, while it needs 10 years to reach its maximum in diameter [4,5]. Worldwide, black locust contributes greatly to soil quality by changing the biodiversity of soil, improving soil chemical properties and fertility [6,7,8,9,10], restoring degraded soils [11,12], increasing the root biomass, and sequestering the organic carbon in soil [13,14]. Biodiversity enhancement and soil improvement are the results of its ability to form symbiotic associations with both nitrogen-fixing rhizobia and phosphorus-acquiring mycorrhizal fungi. Binding 75–150 kg ha−1 of atmospheric nitrogen year−1 can significantly increase the available nitrogen content of the soil and affect pH [15,16]. Rising costs of fertilizers and environmental efforts are contributing to a reduction in their use for improving plant growth and yields. Populus x euramericana L. represents the majority of poplar plantations in European countries [17,18]. Part of the European Union’s policy is to produce energy through the use of biomass to mitigate the effects of climate change by reducing greenhouse gas emissions and securing energy supply through diversification of energy sources [19]. Agroforestry has been proposed as an alternative land-use system and is earmarked as a target area for the productive growth of trees such as poplar. In their natural habitats, as well as in plantations, poplars are colonized by ectomycorrhizal fungi, this interaction is important for tree nutrition and can profoundly modulate plant responses to unfavourable environmental conditions [20,21,22]. Previous studies discussed the mechanism of plant–microbe interactions which affects plant health and soil fertility [23,24]. Mycorrhizal fungal inoculation has the potential to be a useful biotechnological tool that benefits plant development and health, that increases plant defence mechanisms and alleviates different stress effects [25,26,27,28,29,30,31,32,33].
In this study, we established black locust and poplar plantation at the same study site in Hungary, to explore the response of the rhizosphere soil bacterial and fungal community after mycorrhizal inoculation and inorganic fertilization.

2. Materials and Methods

2.1. Study Site

The plantation study was conducted in Monorierdő, Pest County, Hungary (N 47°30′, E 19°48′) (Figure 1). The study of the plantation field experiments area has a moderately cold–dry continental climate with 10.5 °C annual mean temperature and 500–750 mm annual mean precipitation [34].

2.2. Intensively Cultivated Plantations Experiment Setup

Black locust and poplar plantation experiments were conducted in two separate parcels with the same soil properties and climate. The parcels were used for experimentation with forestry varieties over the past 20 years.
Black locust (one-year-old R. pseudoacacia L. cuttings; OBE01; OBE26; OBE34; OBE53; OBE54; OBE69 varieties) was inoculated at planting time (March 2018) with arbuscular mycorrhizal fungi inoculant labelled in the experiment as ‘III’ (50 g plant−1, 80 propagules g−1 Symbivit® produced by Symbiom Ltd., Lanskroun, Czech Republic; www.symbiom.cz, accessed on 4 September 2017), fertilized labelled ‘II’ (70 g plant−1, Osmocote OSM, NPK, 14-7-21, Israel Chemicals, Tel-Aviv, Israel, Pétisó (27% N, 7% CaO, 5% MgO), Superphosphate (18% P2O5); NPK 9-6-7.5 g plant−1) and non-treated plants were used as control ‘I’. We used a randomized block design of three treatments (840 plants treatment−1).
Poplar (one-year-old P. x euramericana L. cuttings; SV-778; SV-879; SV-890; I-214; AF-13; AF-28 varieties) was inoculated at planting time (October 2017) with ectomycorrhizal fungi inoculant, labelled as ‘C’ (3 g plant−1, Tuber brumale L.) and fertilized ‘B’ (278 g plant−1, Pétisó (27% N, 7% CaO, 5% MgO)) and non-treated control plants labelled in the experiment as ‘A’. The plantation experiment was arranged in a randomized block design with 450 plants treatment−1.
One hundred grams of soil sample in five replications of each treatment was collected (soil cores of 5 cm diameter and 25 cm length were collected. The top 5 cm of the cores was removed and the rest was mixed thoroughly) and were used to determine the microbiome of the soil. Sampling was conducted on 12 October 2020 (Figure S1). The height and stem diameter of trees were measured (9 October 2020) (Figure S1).

2.3. Metagenomic Sequencing

For a total of 30 soil samples (two plantations × three treatments × five soil samples), DNA was extracted using the Quick DNA Fecal/Soil Microbe Miniprep (ZymoResearch, Irvine, CA, USA) following the manufacturer’s instructions. The quality of the DNA extracted was determined using a nanospectrophotometer (Nanophotometer 2210, Implen, Germany). For sequencing, the DNA extractions from each treatment (five samples) were pooled into one sample. Thus, three pooled samples were sequenced for each plantation (six samples in total). The abundance of the bacterial and fungal communities of soil samples was estimated using high-throughput sequencing of the 16S rRNA gene on the Illumina Miseq platform at UD-GenoMed Ltd. (Debrecen, Hungary). The V3–V4 region of bacterial 16S rRNA gene and the fungal ITS1 region was amplified using universal primers 16S and ITS Amplicon PCR universal primers (Sigma-Aldrich, St. Louis, MI, USA) (Table S1), following the recommendations of the 16S and Fungal Metagenomic Sequencing Library Preparation guides (Illumina, San Diego, CA, USA). The KAPA HiFi Hot Start Ready Mix (KAPA Biosystems, Wilmington, MA, USA; Roche AG, Basel, Switzerland) was used to perform PCR amplification. These samples were denatured at 95 °C for 3 min and then amplification was performed using three steps of PCR for 25 cycles, denaturing at 95 °C for 30 s, annealing at 55 °C for 30 s, extending at 72 °C for 30 s. Post-amplification quality control was performed on an Agilent Bioanalyzer (Agilent Technologies, Santa Clara, CA, USA). Purification of 16S and ITS rDNA amplicons were achieved using MagSi-NGSPrep Plus (Magtivio B.V., Nuth, The Netherlands) magnetic beads. Index primers: 502, 503, 504, and 701, 702, 703, 704, 705, 706 were used to add Illumina index tags to the ends of the amplicons Nextera XT Index Kit (Illumina, San Diego, CA, USA). The thermal cycling protocol was 8 cycles of denaturation at 95 °C for 30 s, annealing at 55 °C for 30 s and extension at 72 °C for 30 s was performed by the KAPA HiFi Hot Start Ready Mix (KAPA Biosystems, Wilmington, MA, USA; Roche AG, Basel, Switzerland). PCR products were cleaned by magnetic beads and were subjected to library quantification using MagSi-NGSPrep Plus (Magtivio B.V., Nuth, The Netherlands). For the library validation, 1 µL of the diluted final library was run on a Bioanalyzer DNA 100 chip on the Agilent Bioanalyzer (Agilent Technologies, Santa Clara, CA, USA). Libraries were normalized, pooled and loaded onto the Illumina MiSeq platform for 2 × 250 bp paired-end sequencing. 16S and ITS rRNA gene paired-end amplicon reads were processed using the Frogs pipeline [35]. Briefly, forward and reverse reads were filtered and merged using vsearch [36] with the parameters: min amplicon size: 44; max amplicon size: 550; mismatch rate: 0.15). Merged sequences were clustered using swarm [37]. Chimera sequences were removed using remove_chimera.py from the Frogs pipeline. Taxonomic assignment was performed using BLAST [38] against SILVA_SSU_r132_March2018 database [39].

2.4. Statistical Analysis

Statistical analysis was performed with R Statistical Software 4.0.2 [40]. The differences of plant growth parameters, microbial α-diversity, and relative abundances of microbial taxa were determined by one-way analysis of variance (ANOVA), followed by Tukey’s post hoc test. In all tests, a p-value < 0.05 was considered to indicate statistical significance. Venn diagram-based analyses were performed in the ‘VennDiagram’ package. A heatmap of relative abundance by the corresponding z-score was plotted using the ‘pheatmap’ package. Z-score, calculated with the formula z = (x − µ)/σ, where x is the abundance of the taxonomic profiles in each sample, µ is the mean value of the abundances, and σ is the standard deviation of the abundances.

3. Results

3.1. Biomass Production and Plant Growth Rate

The effect of inoculation treatment on plantations did not show significant differences in black locust height and stem diameter. In the case of inoculation and fertilization, only the stem diameter of the poplar plantation showed a modestly significant change close to the margin of error. The largest stem diameter was detected in control plants while the smallest was at the treatment of fertilizer (Table S2).

3.2. Diversity and Composition of Microbial Communities

After the quality check, sequencing and chimera removal the diversities of the rhizosphere soil bacterial and fungal communities of black locust and poplar plantations using the 16S rRNA and ITS rRNA primer sets across per sample were obtained and cluster analysis to read operational taxonomic units (OTU) at 100% identity by BLAST (Figure 2, Tables S3 and S4). Venn diagrams were employed to compare the rhizosphere soil bacterial and fungal communities based on unique and shared OTUs among the different treatments. Bacterial communities in both plantations contained more OTUs than fungal communities. However, the fungal communities, compared to the bacterial communities, contained higher proportions of unique OTUs in the same treatments in rhizosphere soil of black locust (10.2%, 7.2% and 5.4%) and poplar (10%, 10.6% and 10.6%) plantations.
According to all sequences of R. pseudoacacia plantations, the dominant bacterial phyla (relative abundance greater than 1%) were as follows: Actinobacteria, Proteobacteria, Acidobacteria, Planctomycetes, Verrucomicrobia, Chloroflexi, Bacteroidetes, Firmicutes and Gemmatimonadetes with an average proportion of 30.39%, 29.43%, 12.99%, 8.33%, 4.46%, 4.36%, 3.70%, 3.16% and 1.80% respectively (Figure 3). The relative abundances of Actinobacteria, Proteobacteria, Acidobacteria, and Planctomycetes were significantly higher than other phyla. Above all, Actinobacteria and Acidobacteria were more abundant in fertilizer treated than in control and AM fungi inoculated soils. Conversely, the abundance of the second and fourth most dominant phyla, Proteobacteria and Planctomycetes decreased in fertilized treatment soils. The abundance of Bacteriodetes was the highest in arbuscular mycorrhiza fungi inoculated and the lowest in fertilized soil. Meanwhile, the relative abundance of Firmicutes behaved oppositely in treated soils (Figure 3).
The rhizosphere bacterial community of poplar plantations was dominated by Actinobacteria (30.71%), Proteobacteria (27.74%) followed by Acidobacteria (12.19%), Planctomycetes (9.50%), Chloroflexi (6.03%), Verrucomicrobia (4.47%), Bacteroidetes (3.04%), Firmicutes (2.86%) and Gemmatimonadetes (2.02%). The relative abundances of Actinobacteria and Proteobacteria were higher than the abundances of Acidobacteria and other phyla. Conversely to the relative abundance of phyla in the soil of black locust, Actinobacteria and Proteobacteria were not abundant among the treatments. Meanwhile, the relative abundance of Verrucomicrobia was lower in treated soils than in control (Figure 3).
All fungal sequences of black locust plantations were subjected to 14 phyla of control and mycorrhizal treatments and twelve phyla of fertilized treatment, of which 95.04%, 95.64% and 94.86% came from the top three phyla including Ascomycota, Basidiomycota and Mortierellomycota (Figure 4). Relative abundance of Ascomycota was the lowest (58.33% contrary to 76.44% and 72.79%), Basidiomycota was the highest (33.22% contrary to 13.31% and 20.53%) in the soil of arbuscular mycorrhizal treatment (Figure 4). Although not part of the top 10 phyla, the relative abundance of Glomeromycota was significantly higher (0.93%) in arbuscular mycorrhizal fungi inoculated black locust soil than in control (0.60%) and in fertilized (0.02%) treated soils. The fungal community of poplar plantations was dominated by Ascomycota, Basidiomycota and Mortierellomycota phyla of control (96.77%), fertilized (95.52%) and ectomycorrhizal inoculated (89.05%) treatments. Chytridiomycota (1.23%) and Kickxellomycota (0.77%) were present in much higher abundance in the ecto-mycorrhizal inoculated poplar rhizosphere, as compared to control (0.54% and 0.31%) and fertilized (0.05% and 0.05%) soil (Figure 4).
The highest bacterial and fungal species richness was observed in the rhizosphere soil of control treatment in black locust plantation (1881.14 and 562.11, based on Chao1 index). The control soil of black locust plantation had the highest diversity (Shannon’s diversity range at 6.79), while the fertilizer treatment had the lowest bacterial species richness and diversity. For the poplar plantation, the bacterial and fungal community richness and bacterial diversity increased in soil of fertilizer treatment. The fungal diversity in rhizosphere soil of ectomycorrhizal treatment was the highest in poplar plantations (Table 1).
The heat maps based on the relative abundance showed that bacterial the community was weakly affected by fertilizer treatment as well as mycorrhizal inoculation (Figure 5a and Figure S2, Figure 6a and Figure S3). In the soil of the black locust plantation, at the family level, the soil of control treatment was dominated by the Xanthobacteraceae family (6.08%), lagging behind, but Micrococcaceae (3.54%) and Sphingomonadaceae (3.60%) were present with higher relative abundance than other families (Beijerinckiaceae 2.81%, Chthoniobacteraceae 2.18%, Burkholderiaceae 2.71%, Micromonosporaceae 2.67%, etc.). The presence of these families was changed by AM fungi inoculation. The relative abundance of dominant Xanthobacteraceae decreased, while an increase was observed in Micrococcaceae (4.63%). However, Beijerinckiaceae (2.81%), which was detected low relative abundance in the control, was present at almost double the size (5.16%) in mycorrhizal treated soil (Figure 55a, Figure 6a and Figure S2). Micrococcaceae was present in higher numbers (5.59%) in the control soil of poplar plantation, followed by Xanthobacteraceae (4.02%), Sphingomonaceae (3.98%) and Beijerinckiaceae (3.49%). In fertilizer-treated soils, Pirellulaceae (3.91%) was present in soils with higher relative abundance (3.91%) than in control (2.09%) and ectomycorrhizal fungi treatment (2.52%) (Figure 6a). As a result of mycorrhizal inoculation, Sphingomonadaceae (4.43%) and Nocardioidaceae (3.90%) showed higher relative abundance (Figure 5a). At the genus level, Pseudarthrobacter (4.38%) and Microvirga (4.47%) were present with higher relative abundances in the rhizosphere soil of black locust plantation inoculated with arbuscular mycorrhizal fungi. Pseudarthrobacter genus dominated in the control soil of poplar plantation (5.49%) as well. It is noteworthy that Candidatus Udaeobacter was present in the control soil with 2.28% relative abundance compared to 0.79% and 0.66% of the treated soils (Figure S3). Candidatus Udaeobacter in the soil of the black locust plantation did not show a difference in the soils of control (1.80%) and fertilizer (1.92%) treatment, while it decreased in the arbuscular mycorrhizal fungi inoculated soil (0.68%) (Figure S2). Luteolibacter appeared in the control soil of the poplar plantation with an outstanding relative abundance (2.55%) compared to soil inoculated with ectomycorrhizal fungi (0.13%) and treated with fertilizer (0.12%). Nocardioides was present in higher relative abundance in the soils of arbuscular mycorrhizal (2.07%) and ectomycorrhizal (2.57%) treatments than in the soils of control and fertilizer treatments in black locust and poplar plantations as well (Figures S2 and S3).
The dominant fungal phyla across all soil samples were Ascomycota and Basidiomycota. In the soil of the black locust and poplar plantation, at the family level, the soil fungal community of all treatment was dominated by families belonging to Ascomycota phyla. In the soil of black locust plantations, no difference was detected (11.37% to 13.24%) in the most dominant Nectriaceae family among the treatments. The soil of the fertilized treatment was dominated by the Phallaceae family (16.67%), Aspergillaceae (12.37%) with mycorrhizal treatment (12.05%) and Nectriaceae (12.01%). The presence of the families in control and fertilized treatments were changed in the soil of arbuscular mycorrhizal fungi inoculation treatment. The relative abundance of Chaetomiaceae (1.92%) and Ceratobasidiaceae (8.07%) strongly increased, while a decrease was observed in Cordycipitaceae (0.36%) and Helotiaceae (1.21%) compared to control and fertilized treatments (Figure 5c,d). At the genus level, the dominant genera Fusarium, Phallus, Penicillium, Mortierella, Beauveria and Thanatephorus were present with the same tendency of relative abundance in the rhizosphere soil of black locust plantation as the families to which they belong (Figure S2). In the soil of poplar plantations, the relative abundance of the top 20 families and genera were different compared to the soil of black locust plantations. The same tendency of relative abundance in genus level was observed in the rhizosphere soil of poplar plantation as the families to which they belong. In control soil, Phallus (Phallaceae), Fusarium (Nectriaceae), Geospora (Pyronemataceae) with a unique case, and Mortierella (Mortierellaceae) were present with the highest relative abundance (Figure 6C,D and Figure S3). In the soil of fertilized treatment, Verticillium showed higher relative abundance (12.95%) compared to control (0.12%) and ectomycorrhizal treatment (2.36%). Ochroconis (Sympoventuriaceae) showed low relative abundance in black locust soils (0.04% to 0.06%), while in poplar plantations ectomycorrhizal treatment strongly affected (9.07%) its relative abundance compared to control (0.39%) and fertilized (0.53%) treatments (Figure S3).

4. Discussion

The selected plots had similar physiographic conditions and slope gradients. While in large-scale biomass production, as has been the case with plantations, neither the effect of inoculation nor the effect of added fertilizer could be observed during this time. However, it may appear as a noticeable sign that contrary to our expectations mycorrhizal inoculation is not definitely advantageous for the plant height and stem development in the plantation either. In the case of poplar ectomycorrhizal, inoculation and fertilized treatments were not synergistic and caused lags in stem development compared to the control, which is in agreement with previous results [41].
This study represents the first comparative description of the bacterial and fungal community structures of rhizosphere soil of black locust and poplar plantations using the next-generation Illumina-based sequencing approach. Our findings suggested that fertilizer use or inoculation by mycorrhizal fungi had a minor impact on soil bacterial composition in the rhizosphere both of black locust and poplar plantations. Our results show that the bacterial communities of field samples, regardless of treatment, are dominated by four phyla (Figure 3). This is consistent with the reports of Zhang and Xu [42] who showed that Proteobacteria and Acidobacteria are ubiquitous and are the most dominant in almost all soil types. These findings are further confirmed by Wei et al. [43] and Wu et al. [44], who found dominance of Proteobacteria, Actinobacteria, Acidobacteria, Chloroflexi, Gemmatimonadetes, Verrucomicrobia, Bacteroidetes, Planctomycetes, Saccharibacteria, and Nitrospirae in agricultural rhizosphere soils. Soil nutrients play an important role in the bacterial community composition as Proteobacteria belong to copiotrophic groups, due to their extracellular membranes consisting of lipopolysaccharides that are involved in the carbon conversion, and Acidobacteria, Verrucomicrobia and Bacteroidetes belonging to oligotrophic groups [45,46]. The presence of Acidobacteria and Proteobacteria with high relative abundance in the rhizosphere of black locust and poplar is probably attributed to the nutrient-rich conditions [47,48]. Actinobacteria participate in biological phosphorus removal and are an important participant of the rhizosphere microbial community. The presence of the other top nine dominant phyla—Planctomycetes, Verrucomicrobia, Chloroflexi, Firmicutes, and Gemmatimonadetes, are in agreement with the findings of previous studies [49,50]. Cederlund et al. [51] observed a decrease in Verrucomicrobia to nitrogen fertilization which can be paralleled with our results where the highest abundance of Verrucomicrobia was found in control soil of poplar plantation. The most represented family for free-living aerobic nitrogen-fixating bacteria was Beijerinckiaceae (Proteobacteria) and for denitrification the genus Pseudarthrobacter (family Micrococcaceae, phyla Actinobacteria), which has also been reported as a plant growth-promoting bacteria [52]. The family of Micrococcaceae, Micromonosporaceae, and Nocardioidaceae belong to Actinobacteria phyla and free-living micro-organisms in soil as Chthoniobacteraceae (Verrucomicrobia) and Pirellulaceae (Planctomycetes). Bacteria of these families are widespread in soils and terrestrial environments [53,54]. As a dominant family, Micromonosporaceae is described as an important participant in the turnover of organic plant material as some species belonging to this family have the ability to degrade chitin, cellulose, lignin, pectin and produce useful secondary metabolites and enzymes [55]. The species economy, and high enzyme activity of the rhizosphere soil of plantations, decreased compared to an agriculturally cultivated soil where rapidly alternating plant vegetation affects the composition of the microbial community and activity of metabolic functions of soil [56]. The amount of microorganisms in soil is often associated with a species economy but may differ in functionality. However, the used techniques do not allow us to know the exact genera that are involved in these metabolic processes. The genus Microvirga (Proteobacteria) showed a wide spectrum of metabolic activities and are adapted to various environments [57], as the genera Luteolibacter, Nocardioides and Candidatus Udaeobacter are abundant soil bacteria isolated from a variety of habitats [58]. Contrary to our study, Lin et al. [59] found a correlation in colonization and enrichment for Rhizobiales in successional environments. Despite the highest dominance of Rhizobiales order in both soils of the plantations, correlation among inoculation, treatment of fertilizer and the presence of Rhizobiales was not detectable. We expected soil bacterial diversity would be higher in treated soils compared to control as well as affected by different host plants as were observed in other studies [60,61,62]. However, our data do not support this notion; we found little difference in soil bacterial diversity between black locust and poplar plantation and among different treatments.
Compared to the soil bacterial community, the fungal community showed a much higher diversity among different treatments. The impact of chemical fertilization on soil fungal communities is a growing concern due to the changes it has on microbe composition in soil ecosystems. We found different effects of treatments and host plants on soil fungal Chao1 index, Shannon index and observed species. In the respective study site, the fungal community in chemical fertilizer-treated soil differed from mycorrhizal and control soils (Figure 2, Table 1). We found the highest fungal diversity in the control rhizosphere soil of black locust and the lowest one in fertilized soil. Meanwhile, the chemical fertilizer-treated soil was characterized with the highest diversity in poplar plantations, probably due to the positive stimulatory effects of chemical fertilizer on fungal populations [63]. These findings may be more dependent on the effect of tree species and less attributable to the effect of the treatment [64]. However, it was also reported that foliar application of mineral and organic fertilizers on ectomycorrhizal fungi abundance and diversity were near neutral [65]. Our results showed that the soil fungal community was dominated by Ascomycota in the same way in black locust and poplar plantations (Figure 4), which is similar to several other studies [66,67,68,69]. Ascomycota is the largest and most diverse phyla [70] and may be considered as the most influential and primary indicator for the rhizosphere fungal community. Fungi strains of the applied inoculant in the black locust plantation belong to the Glomeraceae family (Glomeromycota) and was present with low relative abundance (<0.2%) in rhizosphere soil in this study. In previous studies, the members of this family could only be identified from the root and by spore isolation in the soil of black locust [71,72]. These results indicate that the applied mycorrhizal inoculation has less effect on the structure of the fungal rhizosphere community under this condition. Mycorrhizal inoculation in poplar plantation was applied by Tuber brumale (Tuberaceae, Pezizales). The Pezizales order was identifiable with a 1.5% relative abundance in rhizosphere soil of ectomycorrhizal treatment. In the soil of ectomycorrhizal treatment of poplar plantation on the family level, Pyronemataceae, known as paraphyletic and ectomycorrhizal [73] former, was detected with lower relative abundances, while on the genus level Ocroconis belonging to the Pyronemataceae family and ectomycorrhizal symbionts containing the Pezizaceae family was represented with significantly higher abundance (Figure 6c,d). Thus, the relatively higher abundance of fungi belonging to Pezizaceae and Pyronemataceae in our study may be promoted by ectomycorrhizal inoculation [74,75,76]. In the present study, the data of control soil of plantations confirmed the findings of previous studies that reported the soil fungal community adaptation to different host tree plantations [77,78,79]. The differences in the patterns of increases at the family and genus level in control soil, shown by fungi, may indicate the differences in the response of the two different hosts found in the plantations despite the short time period.

5. Conclusions

Overall, our findings revealed that the composition and diversity of the rhizosphere soil microbiome are influenced by the plant species. The applied treatments (mycorrhiza, fertilization) only slightly affected the rhizosphere soil bacterial community in the rhizosphere of both plants. The diversity and composition of the soil fungal community are more affected by chemical fertilizer than mycorrhizal inoculation. In future studies, several factors, such as climatic variation, root exudates and weed vegetation, which could affect the bacterial and fungal communities, should be considered. Soil microbiome diversity and function monitoring also remain crucial in forest and plantation ecosystems.

Supplementary Materials

The following are available online at https://www.mdpi.com/article/10.3390/f12091218/s1, Figure S1. Auger for planting cuttings (a), black locust plantation in October 2020 (b), poplar plantation in October 2020 (c), rhizosphere soil sampling in October 2020 (d); (photos: National Food Chain Safety Office, Agricultural Genetic Resources Directorate, Forestry Reproductive Material Department)., Figure S2. Relative abundance by the corresponding z-score of top 20 bacterial (a) and fungal (c) genera; relative abundance (%) of top 20 bacterial (b) and fungal (d) genera in rhizosphere soil of black locust plantation by treatments; control (I), fertilizer (II) and arbuscular mycorrhizal (III) treatment., Figure S3. Relative abundance by the corresponding z-score of top 20 bacterial (a) and fungal (c) genera; relative abundance (%) of top 20 bacterial (b) and fungal (d) genera in rhizosphere soil of poplar plantation by treatments; control (A), fertilizer (B) and ectomycorrhizal (C) treatment., Table S1. 16S and ITS primers used in this study., Table S2. Plant height and stem diameter of black locust and poplar in the plantation experiment., Table S3. BLAST affiliation by treatment and taxonomy distribution of bacterial metagenomic analyses., Table S4. BLAST affiliation by treatment and taxonomy distribution of fungal metagenomic analyses.

Author Contributions

Conceptualization, Z.M., A.G.C. and K.P.; methodology, Z.M., A.G.C., A.P. and K.P.; software, Z.M., Á.J. and A.O.; validation, Z.M., A.G.C., A.O., A.P. and K.P.; formal analysis, Z.M., Á.J. and K.P.; investigation, Z.M., A.G.C., A.P. and B.K.N.; resources, K.P.; writing, Z.M.; writing—review and editing, Z.M. and K.P.; visualization, Z.M., Á.J.; supervision, Z.M.; project administration, A.G.C.; funding acquisition, K.P. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the Development and Innovation Fund of Hungary, grant number 2017-1.3.1-VKE-2017-00022 and by the Ministry of Innovation and Technology within the framework of the Thematic Excellence Programme 2020, Institutional Excellence Subprogramme, grant number TKP2020-IKA-12.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The data presented in this study are available on request from the corresponding author.

Acknowledgments

The authors wish to thank Beatrix Pethőné Rétháti for administrative support, Imréné Gódor for laboratory assistance and György Molnár for assistance with the field survey.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Sitzia, T.; Cierjacks, A.; de Rigo, D.; Caudullo, G. Robinia pseudoacacia in Europe: Distribution, habitat, usage and threats. In European Atlas of Forest Tree Species; Publication Office of the European Union: Luxembourg, 2016; pp. 166–167. [Google Scholar]
  2. Hungarian Central Statistical Office. Available online: https://www.ksh.hu/docs/hun/xstadat/xstadat_eves/i_ome002b.html (accessed on 23 March 2021).
  3. Straker, K.C.; Quinn, L.D.; Voigt, T.B.; Lee, D.K.; Kling, G.J. Black locust as a bioenergy feedstock: A review. Bioenergy Res. 2015, 8, 1117–1135. [Google Scholar] [CrossRef]
  4. Rédei, K.; Keserű, Z.; Antal, B. Improved clonal approaches to growing black locust (Robinia pseudoacacia L.) in Hungary: A case study. Int. J. Hortic. Sci. 2015, 21, 53–56. [Google Scholar] [CrossRef] [Green Version]
  5. Rédei, K.; Osvath-Bujtas, Z.; Veperdi, I. Black locust (Robinia pseudoacacia L.) improvement in Hungary: A review. Acta Silv. Lignaria Hung. 2008, 4, 127–132. [Google Scholar]
  6. Bolat, I.; Kara, Ö.; Sensoy, H.; Yüksel, K. Influences of black locust (Robinia pseudoacacia L.) afforestation on soil microbial biomass and activity. iForest-Biogeosci. For. 2015, 9, 171. [Google Scholar] [CrossRef]
  7. Papaioannou, A.; Chatzistahis, T.; Papaioannou, E.; Papadopoulos, G. Robinia pseudoacacia as a valuable invasive species for the restoration of degraded croplands. Catena 2016, 137, 310–317. [Google Scholar] [CrossRef]
  8. Gentili, R.; Ferrè, C.; Cardarelli, E.; Montagnani, C.; Bogliani, G.; Citterio, S.; Comolli, R. Comparing negative impacts of Prunus serotina, Quercus rubra and Robinia pseudoacacia on native forest ecosystems. Forests 2019, 10, 842. [Google Scholar] [CrossRef] [Green Version]
  9. Gillespie, A.R.; Pope, P.E. Rhizosphere acidification increases phosphorus recovery of black locust: II. Model predictions and measured recovery. Soil Sci. Soc. Am. J. 1990, 54, 538–541. [Google Scholar] [CrossRef]
  10. Tateno, R.; Tokuchi, N.; Yamanaka, N.; Du, S.; Otsuki, K.; Shimamura, T.; Xue, Z.; Wang, S.; Hou, Q. Comparison of litterfall production and leaf litter decomposition between an exotic black locust plantation and an indigenous oak forest near Yan’an on the Loess Plateau. China. For. Ecol. Manag. 2007, 241, 84–90. [Google Scholar] [CrossRef]
  11. Wang, Y.L.; Liu, G.B.; Kume, T.; Otsuki, K.; Yamanaka, N.; Du, S. Estimating water use of a black locust plantation by the thermal dissipation probe method in the semiarid region of Loess Plateau, China. J. For. Res. 2010, 15, 241–251. [Google Scholar] [CrossRef]
  12. Mantovani, D.; Veste, M.; Böhm, C.; Vignudelli, M.; Freese, D. Spatial and temporal variation of drought impact on black locust (Robinia pseudoacacia L.) water status and growth. Forest 2015, 8, 743–747. [Google Scholar] [CrossRef] [Green Version]
  13. Quinkenstein, A.; Boehm, C.; da Silva Matos, E.; Freese, D.; Huettl, R.F. Assessing the carbon sequestration in short rotation coppices of Robinia pseudoacacia L. on marginal sites in Northeast Germany. In Carbon Sequestration Potential of Agroforestry Systems; Springer: Dordrecht, The Netherlands, 2011; pp. 201–216. [Google Scholar] [CrossRef]
  14. Wang, J.J.; Hu, C.X.; Bai, J.; Gong, C.M. Carbon sequestration of mature black locust stands on the Loess Plateau, China. Plant Soil Environ. 2015, 61, 116–121. [Google Scholar] [CrossRef] [Green Version]
  15. Cierjacks, A.; Kowarik, I.; Joshi, J.; Hempel, S.; Ristow, M.; von der Lippe, M.; Weber, E. Biological flora of the British Isles: Robinia pseudoacacia. J. Ecol. 2013, 101, 1623–1640. [Google Scholar] [CrossRef]
  16. Song, X.; Shi, S.; Lu, S.; Ren, R.; He, C.; Meng, P.; Zhang, J.; Yin, C.; Zhang, X. Changes in soil chemical properties following afforestation of cropland with Robinia pseudoacacia in the southeastern Loess Plateau of China. For. Ecol. Manag. 2021, 487, 118993. [Google Scholar] [CrossRef]
  17. Dupraz, C.; Liagre, F. Agroforesterie: Des Arbres et Des Cultures; France Agricole Editions: Paris, France, 2008. [Google Scholar]
  18. Barrio-Anta, M.; Sixto-Blanco, H.; Canellas-Rey, D.V.I.; Castedo-Dorado, F. Dynamic growth model for I-214 poplar plantations in the northern and central plateaux in Spain. For. Ecol. Manag. 2008, 255, 1167–1178. [Google Scholar] [CrossRef]
  19. Rytter, R.M. The potential of willow and poplar plantations as carbon sinks in Sweden. Biomass Bioenergy 2012, 36, 86–95. [Google Scholar] [CrossRef]
  20. Danielsen, L.; Lohaus, G.; Sirrenberg, A.; Karlovsky, P.; Bastien, C.; Pilate, G.; Polle, A. Ectomycorrhizal colonization and diversity in relation to tree biomass and nutrition in a plantation of transgenic poplars with modified lignin biosynthesis. PLoS ONE 2013, 8, e59207. [Google Scholar] [CrossRef]
  21. Habib, M.T.; Heller, T.; Polle, A. Molecular physiology of tree ectomycorrhizal interactions. In Plant Roots: The Hidden Half; CRC Press: Boca Raton, FL, USA, 2013. [Google Scholar] [CrossRef]
  22. Luo, Z.B.; Wu, C.; Zhang, C.; Li, H.; Lipka, U.; Polle, A. The role of ectomycorrhizas in heavy metal stress tolerance of host plants. Environ. Exp. Bot. 2014, 108, 47–62. [Google Scholar] [CrossRef]
  23. Jeffries, P.; Gianinazzi, S.; Perotto, S.; Turnau, K.; Barea, J.M. The contribution of arbuscular mycorrhizal fungi in sustainable maintenance of plant health and soil fertility. Biol. Fertil. Soils 2003, 37, 1–16. [Google Scholar] [CrossRef]
  24. Toljander, J. Interactions between Soil Bacteria and Arbuscular Mycorrhizal Fungi; Swedish University of Agriculture Sciences, Acta Universitatis Agriculturae Sueciae: Uppsala, Sweden, 2006. [Google Scholar]
  25. Barman, J.; Samanta, A.; Saha, B.; Datta, S. Mycorrhiza. Resonance 2016, 21, 1093–1104. [Google Scholar] [CrossRef]
  26. Chen, J.; Zhang, H.; Zhang, X.; Tang, M. Arbuscular mycorrhizal symbiosis alleviates salt stress in black locust through improved photosynthesis, water status, and K+/Na+ homeostasis. Front. Plant Sci. 2017, 8, 1739. [Google Scholar] [CrossRef]
  27. He, F.; Sheng, M.; Tang, M. Effects of Rhizophagus irregularis on photosynthesis and antioxidative enzymatic system in Robinia pseudoacacia L. under drought stress. Front. Plant Sci. 2017, 8, 183. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  28. Dagher, D.J.; de la Providencia, I.E.; Pitre, F.E.; St-Arnaud, M.; Hijri, M. Arbuscular mycorrhizal fungal assemblages significantly shifted upon bacterial inoculation in non-contaminated and petroleum-contaminated environments. Microorganisms 2020, 8, 602. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  29. Lewis, D.E.; White, J.R.; Wafula, D.; Athar, R.; Dickerson, T.; Williams, H.N.; Chauhan, A. Soil functional diversity analysis of a bauxite-mined restoration Chronosequence. Microb. Ecol. 2010, 59, 710–723. [Google Scholar] [CrossRef] [PubMed]
  30. Diagne, N.; Ngom, M.; Djighaly, P.I.; Fall, D.; Hocher, V.; Svistoonoff, S. Roles of arbuscular mycorrhizal fungi on plant growth and performance: Importance in biotic and abiotic stressed regulation. Diversity 2020, 12, 370. [Google Scholar] [CrossRef]
  31. Liu, J.; Ha, V.N.; Shen, Z.; Zhu, H.; Zhao, F.; Zhao, Z. Characteristics of bulk and rhizosphere soil microbial community in an ancient Platycladus orientalis forest. Appl. Soil Ecol. 2018, 132, 91–98. [Google Scholar] [CrossRef]
  32. Jia, X.; Zhang, N.; Zhao, Y.; Wang, L.; Zhang, C.; Li, X.; Cao, K.; Gao, Y. A consecutive 4-year elevated air temperature shaped soil bacterial community structure and metabolic functional groups in the rhizosphere of black locust seedlings exposed to lead pollution. Sci. Total Environ. 2020, 732, 139273. [Google Scholar] [CrossRef]
  33. Giannoulis, K.D.; Skoufogianni, E.; Bartzialis, D.; Solomou, A.D.; Danalatos, N.G. Growth and productivity of Salvia officinalis L. under Mediterranean climatic conditions depends on biofertilizer, nitrogen fertilization, and sowing density. Ind. Crop. Prod. 2021, 160, 113136. [Google Scholar] [CrossRef]
  34. Hungarian Meteorological Service. Available online: https://www.met.hu/eghajlat/magyarorszag_eghajlata/ (accessed on 29 August 2021).
  35. Escudie, F.; Auer, L.; Bernard, M.; Mariadassou, M.; Cauquil, L.; Vidal, K.; Maman, S.; Hernandez-Raquet, G.; Combes, S.; Pascal, G. FROGS: Find, Rapidly, OTUs with Galaxy Solution. Bioinformatics 2018, 34, 1287–1294. [Google Scholar] [CrossRef]
  36. Rognes, T.; Flouri, T.; Nichols, B.; Quince, C.; Mahé, F. VSEARCH: A versatile open source tool for metagenomics. PeerJ 2016, 4, e2584. [Google Scholar] [CrossRef]
  37. Mahé, F.; Rognes, T.; Quince, C.; de Vargas, C.; Dunthorn, M. Swarm: Robust and fast clustering method for amplicon-based studies. PeerJ 2014, 2, e593. [Google Scholar] [CrossRef] [Green Version]
  38. McGinnis, S.; Madden, T.L. BLAST: At the core of a powerful and diverse set of sequence analysis tools. Nucleic Acids Res. 2004, 32 (Suppl. S2), W20–W25. [Google Scholar] [CrossRef]
  39. Quast, C.; Pruesse, E.; Yilmaz, P.; Gerken, J.; Schweer, T.; Yarza, P.; Peplies, J.; Glöckner, F.O. The SILVA ribosomal RNA gene database project: Improved data processing and web-based tools. Nucleic Acids Res. 2012, 41, D590–D596. [Google Scholar] [CrossRef]
  40. R Core Team. R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria. 2020. Available online: https://www.r-project.org/ (accessed on 10 February 2020).
  41. Bzdyk, R.M.; Olchowik, J.; Studnicki, M.; Nowakowska, J.A.; Oszako, T.; Urban, A.; Hilszczańska, D. Ectomycorrhizal Colonisation in Declining Oak Stands on the Krotoszyn Plateau, Poland. Forests 2019, 10, 30. [Google Scholar] [CrossRef] [Green Version]
  42. Zhang, L.; Xu, Z. Assessing bacterial diversity in soil. J. Soils Sediments 2008, 8, 379–388. [Google Scholar] [CrossRef]
  43. Wei, H.; Peng, C.; Yang, B.; Song, H.; Li, Q.; Jiang, L.; Wei, G.; Wang, K.; Wang, H.; Liu, S.; et al. Contrasting Soil Bacterial Community, Diversity, and Function in Two Forests in China. Front. Microbiol. 2018, 9, 1693. [Google Scholar] [CrossRef]
  44. Wu, S.J.; Deng, J.; Yin, Y.; Qin, S.J.; Zhu, W.X.; Zhou, Y.B.; Wang, B.; Ruan, H.; Jin, L. Bacterial Community Changes Associated with Land Use Type in the Forest Montane Region of Northeast China. Forests 2019, 11, 40. [Google Scholar] [CrossRef] [Green Version]
  45. Fierer, N.; Bradford, M.A.; Jackson, R.B. Toward an ecological classification of soil bacteria. Ecology 2007, 88, 1354–1364. [Google Scholar] [CrossRef]
  46. Jangid, K.; Williams, M.A.; Franzluebbers, A.J.; Sanderlin, J.S.; Reeves, J.H.; Jenkins, M.B.; Endale, D.M.; Coleman, D.C.; Whitman, W.B. Relative impacts of land- use, management intensity and fertilization upon soil microbial community structure in agricultural systems. Soil Biol. Biochem. 2008, 40, 2843–2853. [Google Scholar] [CrossRef]
  47. Gottel, N.R.; Castro, H.F.; Kerley, M.; Yang, Z.; Pelletier, D.A.; Podar, M.; Karpinets, T.; Uberbacher, E.; Tuskan, G.A.; Vilgalys, R.; et al. Distinct microbial communities within the endosphere and rhizosphere of Populus deltoides roots across contrasting soil types. Appl. Environ. Microbiol. 2011, 77, 5934–5944. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  48. Zhang, C.; Liu, G.; Xue, S.; Wang, G. Soil bacterial community dynamics reflect changes in plant community and soil properties during the secondary succession of abandoned farmland in the Loess Plateau. Soil Biol. Biochem. 2016, 97, 40–49. [Google Scholar] [CrossRef]
  49. Castañeda, L.E.; Barbosa, O. Metagenomic analysis exploring taxonomic and functional diversity of soil microbial communities in Chilean vineyards and surrounding native forests. PeerJ 2017, 5, e3098. [Google Scholar] [CrossRef] [PubMed]
  50. Liu, H.; Chen, L.; Wang, E.T.; Liu, Y.; Zhang, L.; Zhao, K.; Gu, Y.; Yu, X.; Ma, M.; Penttinen, P.; et al. Combined microbial consortium inoculation and black locust planting is effective in the bioremediation of waste drill cuttings. Front. Microbiol. 2020, 11, 536787. [Google Scholar] [CrossRef]
  51. Cederlund, H.; Wessén, E.; Enwall, K.; Jones, C.M.; Juhanson, J.; Pell, M.; Philippot, L.; Hallin, S. Soil carbon quality and nitrogen fertilization structure bacterial communities with predictable responses of major bacterial phyla. Appl. Soil Ecol. 2014, 84, 62–68. [Google Scholar] [CrossRef]
  52. Park, M.K.; Park, Y.J.; Kim, M.; Kim, M.C.; Ibal, J.C.; Kang, G.U.; Lee, G.Y.-D.; Tagele, S.B.; Kwon, H.-J.; Kang, M.-S.; et al. Complete genome sequence of a plant growth-promoting bacterium Pseudarthrobacter sp. NIBRBAC000502772, isolated from shooting range soil in the Republic of Korea. The Microbiol. Soc. Korea 2020, 56, 390–393. [Google Scholar] [CrossRef]
  53. Evtushenko, L.I.; Ariskina, E.V. Nocardioidaceae. Bergey’s Man. Syst. Archaea Bact. 2015, 14, 1–18. [Google Scholar] [CrossRef]
  54. De Vries, H.J.; Beyer, F.; Jarzembowska, M.; Lipińska, J.; van den Brink, P.; Zwijnenburg, A.; Timmers, P.H.A.; Stams, A.J.M.; Plugge, C.M. Isolation and characterization of Sphingomonadaceae from fouled membranes. npj Biofilms Microbiomes 2019, 5, 1–9. [Google Scholar] [CrossRef]
  55. Trujillo, M.E.; Hong, K.; Genilloud, O. The family Micromonosporaceae. The Prokaryotes 2014, 499–569. [Google Scholar] [CrossRef]
  56. Mayer, Z.; Sasvári, Z.; Szentpéteri, V.; Pethőné Rétháti, B.; Vajna, B.; Posta, K. Effect of long-term cropping systems on the diversity of the soil bacterial communities. Agronomy 2019, 9, 878. [Google Scholar] [CrossRef] [Green Version]
  57. Bobay, L.M.; Ochman, H. The evolution of bacterial genome architecture. Front. Genet. 2017, 8, 72. [Google Scholar] [CrossRef] [Green Version]
  58. Brewer, T.E.; Handley, K.M.; Carini, P.; Gilbert, J.A.; Fierer, N. Genome reduction in an abundant and ubiquitous soil bacterium ‘Candidatus Udaeobacter copiosus’. Nat. Microbiol. 2016, 2, 1–7. [Google Scholar] [CrossRef]
  59. Lin, Y.T.; Jangid, K.; Whitman, W.B.; Coleman, D.C.; Chiu, C.Y. Soil bacterial communities in native and regenerated perhumid montane forests. Appl. Soil Ecol. 2011, 47, 111–118. [Google Scholar] [CrossRef]
  60. Slabbert, E.; Jacobs, S.M.; Jacobs, K. The soil bacterial com- munities of South African fynbos riparian ecosystems invaded by Australian Acacia species. PLoS ONE 2014, 9, e86560. [Google Scholar] [CrossRef]
  61. Le Roux, J.J.; Ellis, A.G.; van Zyl, L.M.; Hosking, N.D.; Keet, J.H.; Yannelli, F.A. Importance of soil legacy effects and successful mutualistic interactions during Australian Acacia invasions in nutrient-poor environments. J. Ecol. 2018, 106, 2071–2081. [Google Scholar] [CrossRef]
  62. Yu, X.Y.; Zhu, Y.J.; Wang, B.; Liu, D.; Bai, H.; Jin, L.; Wang, B.T.; Ruan, H.H.; Mao, L.; Jin, F.J.; et al. Effects of nitrogen addition on rhizospheric soil microbial communities of poplar plantations at different ages. For. Ecol. Manag. 2021, 494, 119328. [Google Scholar] [CrossRef]
  63. Zhou, J.; Jiang, X.; Zhou, B.; Zhao, B.; Ma, M.; Guan, D.; Li, J.; Chen, S.; Cao, F.; Shen, D.; et al. Thirty four years of nitrogen fertilization decreases fungal diversity and alters fungal community composition in black soil in northeast China. Soil Biol. Biochem. 2016, 95, 135–143. [Google Scholar] [CrossRef]
  64. Aleklett, K.; Hart, M. The root microbiota—a fingerprint in the soil? Plant Soil 2013, 370, 671–686. [Google Scholar] [CrossRef]
  65. Bzdyk, R.M.; Olchowik, J.; Studnicki, M.; Oszako, T.; Sikora, K.; Szmidla, H.; Hilszczańska, D. The Impact of Effective Microorganisms (EM) and Organic and Mineral Fertilizers on the Growth and Mycorrhizal Colonization of Fagus syl-vatica and Quercus robur Seedlings in a Bare-Root Nursery Experiment. Forests 2018, 9, 597. [Google Scholar] [CrossRef] [Green Version]
  66. Wang, S.; Wang, X.; Han, X.; Deng, Y. Higher precipitation strengthens the microbial interactions in semi-arid grassland soils. Glob. Ecol. Biogeogr. 2018, 7, 1–11. [Google Scholar] [CrossRef]
  67. Ma, M.; Jiang, X.; Wang, Q.; Ongena, M.; Wei, D.; Ding, J.; Guan, J.; Cao, F.; Zhao, B.; Li, J. Responses of fungal community composition to long-term chemical and organic fertilization strategies in Chinese Mollisols. MicrobiologyOpen 2018, 7, e00597. [Google Scholar] [CrossRef]
  68. Yang, Y.; Cheng, H.; Dou, Y.; An, S. Plant and soil traits driving soil fungal community due to tree plantation on the Loess Plateau. Sci. Total. Environ. 2020, 708, 134560. [Google Scholar] [CrossRef]
  69. Wu, N.; Li, Z.; Meng, S.; Wu, F. Soil properties and microbial community in the rhizosphere of Populus alba var. pyramidalis along a chronosequence. Microbiol. Res. 2021, 250, 126812. [Google Scholar] [CrossRef]
  70. James, T.Y.; Stajich, J.E.; Hittinger, C.T.; Rokas, A. Toward a fully resolved fungal tree of life. Annu. Rev. Microbiol. 2020, 74, 291–313. [Google Scholar] [CrossRef]
  71. Yang, Y.; Song, Y.; Scheller, H.V.; Ghosh, A.; Ban, Y.; Chen, H.; Tang, M. Community structure of arbuscular mycorrhizal fungi associated with Robinia pseudoacacia in uncontaminated and heavy metal contaminated soils. Soil Biol. Biochem. 2015, 86, 146–158. [Google Scholar] [CrossRef] [Green Version]
  72. He, F.; Tang, M.; Zhong, S.L.; Yang, R.; Huang, L.; Zhang, H.Q. Effects of soil and climatic factors on arbuscular mycorrhizal fungi in rhizosphere soil under Robinia pseudoacacia in the Loess Plateau, China. Eur. J. Soil Sci. 2016, 67, 847–856. [Google Scholar] [CrossRef]
  73. Perry, B.A.; Hansen, K.; Pfisterm, D.H. A phylogenetic overview of the family Pyronemataceae (Ascomycota, Pezizales). Mycol. Res. 2007, 111, 549–571. [Google Scholar] [CrossRef]
  74. Fujimura, K.F.; Smith, J.E.; Horton, T.R.; Weber, N.S.; Spatafora, J.W. Pezizalean mycorrhizas and sporocarps in ponder- osa pine (Pinus ponderosa) after prescribed fires in eastern Oregon, USA. Mycorrhiza 2005, 15, 79–86. [Google Scholar] [CrossRef]
  75. Tedersoo, L.; Hansen, K.; Perry, B.A.; Kjoller, R. Molecular and morphological diversity of pezizalean ectomycorrhiza. New Phytol. 2006, 170, 581–596. [Google Scholar] [CrossRef]
  76. Tedersoo, L.; May, T.W.; Smith, M.E. Ectomycorrhizal lifestyle in fungi: Global diversity, distribution, and evolution of phylogenetic lineages. Mycorrhiza 2010, 20, 217–263. [Google Scholar] [CrossRef]
  77. Sheng, M.; Chen, X.; Zhang, X.; Hamel, C.; Cui, X.; Chen, J.; Chen, H.; Tang, M. Changes in arbuscular mycorrhizal fungal attributes along a chronosequence of black locust (Robinia pseudoacacia) plantations can be attributed to the plantation-induced variation in soil properties. Sci. Total. Environ. 2017, 599, 273–283. [Google Scholar] [CrossRef]
  78. Yang, T.; Adams, J.M.; Shi, Y.; He, J.S.; Jing, X.; Chen, L.; Tedersoo, L.; Chu, H. Soil fungal diversity in natural grasslands of the Tibetan Plateau: Associations with plant diversity and productivity. New Phytol. 2017, 215, 756–765. [Google Scholar] [CrossRef] [Green Version]
  79. Yang, W.; Zhang, D.; Cai, X.W.; Xia, L.; Luo, Y.Q.; Cheng, X.L.; An, S. Significant alterations in soil fungal communities along a chronosequence of Spartina alterniflora in a Chinese Yellow Sea coastal wetland. Sci. Total. Environ. 2019, 693, 133548. [Google Scholar] [CrossRef] [PubMed]
Forests 12 01218 g001 550
Figure 1. Study site of black locust (A) and poplar plantation (B) (https://earth.google.com/web/@47.30475687,19.48656325,140.81392265a,1194.88813009d,35y,0.00005561h,0.1766107t,-0r, accessed on 29 August 2021).
Figure 1. Study site of black locust (A) and poplar plantation (B) (https://earth.google.com/web/@47.30475687,19.48656325,140.81392265a,1194.88813009d,35y,0.00005561h,0.1766107t,-0r, accessed on 29 August 2021).
Forests 12 01218 g001
Forests 12 01218 g002a 550Forests 12 01218 g002b 550
Figure 2. Venn diagram showing the unique and shared OTUs obtained from MiSeq sequencing. Bacterial OTUs in soil of black locust (a) and poplar (b), fungal OTUs in soil of black locust (c) and poplar (d).
Figure 2. Venn diagram showing the unique and shared OTUs obtained from MiSeq sequencing. Bacterial OTUs in soil of black locust (a) and poplar (b), fungal OTUs in soil of black locust (c) and poplar (d).
Forests 12 01218 g002aForests 12 01218 g002b
Forests 12 01218 g003 550
Figure 3. The relative abundance of top 10 bacterial phyla in the soil of black locust and poplar plantation by treatments; control (I—black locust; A—poplar), fertilizer (II—black locust; B—poplar) and arbuscular mycorrhizal (III—black locust) and ectomycorrhizal (C—poplar) treatment.
Figure 3. The relative abundance of top 10 bacterial phyla in the soil of black locust and poplar plantation by treatments; control (I—black locust; A—poplar), fertilizer (II—black locust; B—poplar) and arbuscular mycorrhizal (III—black locust) and ectomycorrhizal (C—poplar) treatment.
Forests 12 01218 g003
Forests 12 01218 g004 550
Figure 4. The relative abundance of top 10 fungal phyla in the soil of black locust and poplar plantation by treatments; control (I—black locust; A—poplar), fertilizer (II—black locust; B—poplar) and arbuscular mycorrhizal (III—black locust) and ectomycorrhizal (C—poplar) treatment.
Figure 4. The relative abundance of top 10 fungal phyla in the soil of black locust and poplar plantation by treatments; control (I—black locust; A—poplar), fertilizer (II—black locust; B—poplar) and arbuscular mycorrhizal (III—black locust) and ectomycorrhizal (C—poplar) treatment.
Forests 12 01218 g004
Forests 12 01218 g005 550
Figure 5. Relative abundance by the corresponding z-score of top 20 bacterial (a) and fungal (c) family; relative abundance (%) of top 20 bacterial (b) and fungal (d) families in rhizosphere soil of black locust plantation by treatments; control (I), fertilizer (II) and arbuscular mycorrhizal (III) treatment.
Figure 5. Relative abundance by the corresponding z-score of top 20 bacterial (a) and fungal (c) family; relative abundance (%) of top 20 bacterial (b) and fungal (d) families in rhizosphere soil of black locust plantation by treatments; control (I), fertilizer (II) and arbuscular mycorrhizal (III) treatment.
Forests 12 01218 g005
Forests 12 01218 g006 550
Figure 6. Relative abundance by the corresponding z-score of the top 20 bacterial (a) and fungal (c) families; relative abundance (%) of top 20 bacterial (b) and fungal (d) families in rhizosphere soil of poplar plantation by treatments; control (A), fertilizer (B) and ectomycorrhizal (C) treatment.
Figure 6. Relative abundance by the corresponding z-score of the top 20 bacterial (a) and fungal (c) families; relative abundance (%) of top 20 bacterial (b) and fungal (d) families in rhizosphere soil of poplar plantation by treatments; control (A), fertilizer (B) and ectomycorrhizal (C) treatment.
Forests 12 01218 g006
Table
Table 1. α-diversity of bacterial and fungal community in rhizosphere soil of black locust and poplar plantations.
Table 1. α-diversity of bacterial and fungal community in rhizosphere soil of black locust and poplar plantations.
Treatment 1Observed Species (OTU)Chao1 Indexse.chao1Shannon Index
Black locust bacterial diversityI18641881.1411.476.79
II17841814.1910.686.61
III18281848.6510.486.65
Black locust fungal diversityI529562.1113.444.87
II508525.447.364.62
III477505.7411.164.70
Poplar bacterial diversityA18301848.708.516.57
B18821883.672.046.82
C18601863.7972.976.76
Poplar fungal diversityA460482.8810.164.98
B500544.3814.674.69
C510536.1012.165.04
1 Control (I—black locust; A—poplar), fertilizer (II—black locust; B—poplar) and arbuscular mycorrhizal (III—black locust) and ectomycorrhizal (C—poplar) treatment.
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Mayer, Z.; Csorbainé, A.G.; Juhász, Á.; Ombódi, A.; Pápai, A.; Némethné, B.K.; Posta, K. Impact of Soil-Applied Microbial Inoculant and Fertilizer on Fungal and Bacterial Communities in the Rhizosphere of Robinia sp. and Populus sp. Plantations. Forests 2021, 12, 1218. https://doi.org/10.3390/f12091218

AMA Style

Mayer Z, Csorbainé AG, Juhász Á, Ombódi A, Pápai A, Némethné BK, Posta K. Impact of Soil-Applied Microbial Inoculant and Fertilizer on Fungal and Bacterial Communities in the Rhizosphere of Robinia sp. and Populus sp. Plantations. Forests. 2021; 12(9):1218. https://doi.org/10.3390/f12091218

Chicago/Turabian Style

Mayer, Zoltán, Andrea Gógán Csorbainé, Ákos Juhász, Attila Ombódi, Antal Pápai, Boglárka Kisgyörgy Némethné, and Katalin Posta. 2021. "Impact of Soil-Applied Microbial Inoculant and Fertilizer on Fungal and Bacterial Communities in the Rhizosphere of Robinia sp. and Populus sp. Plantations" Forests 12, no. 9: 1218. https://doi.org/10.3390/f12091218

APA Style

Mayer, Z., Csorbainé, A. G., Juhász, Á., Ombódi, A., Pápai, A., Némethné, B. K., & Posta, K. (2021). Impact of Soil-Applied Microbial Inoculant and Fertilizer on Fungal and Bacterial Communities in the Rhizosphere of Robinia sp. and Populus sp. Plantations. Forests, 12(9), 1218. https://doi.org/10.3390/f12091218

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop